February, 2020

NIOSH Manual of Analytical

Methods (NMAM), 5th Edition 

Editors: Ronnee Andrews, Ph.D. and Paula Fey O’Connor, NIOSH 

[This document is current as of the publication date above and will be periodically updated. For

the most current listing of methods and guidance chapters, please visit the NMAM website at 

www.cdc.gov/niosh/nmam.] 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

February, 2020

Foreword

The National Institute for Occupational Safety and Health (NIOSH) is the U.S. federal agency 

responsible for conducting research and making recommendations for the prevention of workrelated injury and illness. NIOSH is part of the Centers for Disease Control and Prevention (CDC) 

in the U.S. Department of Health and Human Services. The NIOSH Manual of Analytical Methods 

(NMAM) is a compilation of validated sampling and analytical methods that are used globally for 

occupational exposure assessment in the industrial (occupational) hygiene field and related 

professions. The methods that are published in NMAM are evaluated and validated in 

consideration of their fitness-for-purpose for exposure monitoring in work areas. NIOSH methods 

primarily address workplace air sampling and analysis, but NMAM also includes protocols for 

biological, surface, dermal, and bulk samples. Within NMAM, but separate from the methods 

themselves, are assorted chapters providing background and guidance covering a number of 

subjects. Explanatory chapters on quality assurance, sampling guidance, method development and 

evaluation, aerosol collection, etc., provide valuable information to users of NIOSH methods. 

NMAM chapters provide a convenient resource that augments technical information often (but not 

always) available elsewhere in texts and monographs. Now in its fifth edition, NMAM is 

continuously updated as new or revised methods are evaluated and their performance verified. 

This document is a compilation of its guidance chapters and methods, current as of the date shown 

on the front page. NMAM is published online on the NIOSH web page (www.cdc.gov/niosh/nmam) 

and is available worldwide free of charge. Users are encouraged to visit the NMAM 5th edition 

website for the most current methods and guidance chapters.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHAPTERS

PS - Purpose and Scope 

ME - Development and evaluation of methods 

UA - Measurement uncertainty and NIOSH method accuracy range 

SA - General considerations for sampling airborne contaminants 

AE - Factors affecting aerosol sampling

BA - Sampling and characterization of bioaerosols

FP - Filter pore size and aerosol sample collection 

FI - Measurement of fibers 

SM - Sampling and analysis of soluble metal compounds 

DL - Monitoring diesel particulate exhaust in the workplace 

CN - Analysis of carbon nanotubes and nanofibers on mixed cellulose ester filters by transmission electron microscopy

GL - Glossary 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHODS

0501 - PARTICULATES N.O.R., Total

1453 - VINYL ACETATE

2005 - NITROAROMATIC COMPOUNDS

2008 - CHLOROACETIC ACID

2014 - p-CHLOROPHENOL

2016 - FORMALDEHYDE

2027 - KETONES

2514 - ANISIDINE

2520 - METHYL BROMIDE

2531 - GLUTARALDEYHDE

2536 - VALERALDEHYDE

3800 - ORGANIC & INORGANIC GASES by Extractive FTIR

Spectrometry 

3900 - Volatile Organic Compounds, C1 to C10,

Canister Method 

5004 - HYDROQUINONE

5005 - THIRAM

5007 - ROTENONE

5008 - PYRETHRUM

5009 - BENZOYL PEROXIDE

5016 - STRYCHNINE 

5022 - ARSENIC, organo5033 - p-NITROANILINE

5040 - DIESEL PARTICULATE MATTER (as Elemental Carbon) 

5100 - CARBON BLACK

5504 - ORGANOTIN COMPOUNDS (as Sn)

5509 - BENZIDINE and 3,3’-DICHLOROBENZIDINE

5524 - METALWORKING FLUIDS (MWF) ALL CATEGORIES

5526 - METHYLTIN CHLORIDES

5600 - ORGANOPHOSPHORUS PESTICIDES

5601 - ORGANONITROGEN PESTICIDES

5700 - FORMALDEHYDE ON DUST (TEXTILE OR WOOD)

6001 - ARSINE

6002 - PHOSPHINE

6010 - HYDROGEN CYANIDE

6012 - SULFURYL FLUORIDE

6016 - AMMONIA by IC

6604 - CARBON MONOXIDE

7082 - LEAD by Flame AAS

7302 - ELEMENTS by ICP (Microwave Digestion) 

7304 - ELEMENTS by ICP (Microwave Digestion)

7306 - ELEMENTS by Cellulosic Internal Capsule Sampler 

7400 – ASBESTOS and OTHER FIBERS by PCM

7405 – ALKALI METAL CATIONS Na+, K+, Li+

7502 - ZINC OXIDE

7600 - CHROMIUM, HEXAVALENT

7602 – SILICA, Respirable Crystalline, by IR (KBr pellet) 

7603 - QUARTZ in Respirable Coal Mine Dust, by IR

(Redeposition)

7605 - CHROMIUM, HEXAVALENT by Ion Chromatography 

7701 - LEAD BY PORTABLE ULTRASONIC EXTRACTION/ASV

7703 - CHROMIUM, HEXAVALENT by Field-Portable 

Spectrophotometry

7704 - BERYLLIUM in Air by Fluorometry

7906 - PARTICULATE FLUORIDES and HYDROFLUORIC ACID

by Ion Chromatography

7907 - VOLATILE ACIDS by Ion Chromatography

7908 - NON-VOLATILE ACIDS - (Sulfuric Acid and Phosphoric 

Acid)

8007 - TOLUENE in Blood 

8200 – ELEMENTS in Tissues

8308 - FLUORIDE in Urine

8318 - 2,5-HEXANEDIONE in Urine

8319 - ACETONE and METHYL ETHYL KETONE in Urine

8320 – BUTYLTIN TRICHLORIDE in Urine

8321 - o-CRESOL in Urine

8322 - TRICHLOROACETIC ACID IN Urine

8324 - 3-BROMOPROPIONIC ACID in Urine

8326 - S-BENZYLMERCAPTURIC ACID AND SPHENYLMERCAPTURIC ACID IN Urine

9000 - ASBESTOS, CHRYSOTILE by XRD

9106 - METHAMPHETAMINE and Illicit Drugs, Precursors and 

Adulterants on Wipes by Liquid-Liquid Extraction

9109 - METHAMPHETAMINE and Illicit Drugs, Precursors and 

Adulterants on Wipes by Solid-Phase Extraction

9110 - BERYLLIUM in Surface Wipes by Fluorometry 

9111 - METHAMPHETAMINE on Wipes by LC/MS
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHAPTERS
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Purpose, Scopeand Use of the

NIOSH Manual of Analytical

Methods

by Kevin Ashley, Ph.D. and Paula Fey O’Connor, NIOSH

1 Purpose and scope PS-2

2 How to use NMAM PS-3

3 References PS-8
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-2 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

1 Purpose and scope

The health of working people in myriad industries and occupations is potentially at risk 

through workplace exposure to airborne chemical and biological agents [Hathaway and 

Proctor 2004; Rose and Cohrssen 2011; Eduard et al. 2012; Jakubowski 2012]. Commonly it is 

the responsibility of occupational hygienists and often other public health professionals to 

determine the effectiveness of measures taken to minimize and control worker exposures to 

airborne toxins and toxicants, and this is normally achieved by monitoring workplace air 

quality [DiNardi 2003; Vincent 2007, 2012; Kulkarni et al. 2011]. Air monitoring is vital 

because inhalation is ordinarily the most likely route of exposure in occupational settings. 

Frequently other routes of workplace exposure, notably dermal contact with chemical and 

biological agents, must also be considered [Semple and Cherrie 2003; Brisson and Ashley 

2011; Behroozy 2013]. Complementary biomonitoring methods are also often used to assess 

occupational exposures to toxic chemical compounds through measurement of specific 

analytes, e.g., metabolites and/or biomarkers, in body fluids (normally blood and urine) and 

tissues [Angerer and Greim 2006].

The NIOSH Manual of Analytical Methods (NMAM) is a compilation of analytical methods 

for air, biological, surface (including dermal) and bulk samples that have been evaluated and 

validated in consideration of their fitness for purpose for workplace exposure monitoring 

[NIOSH 1995]. NIOSH sampling and analytical methods are intended to promote accuracy, 

sensitivity, and specificity in industrial hygiene analyses and related applications. NMAM, 

which is published online (available at: www.cdc.gov/niosh/nmam), is constantly updated as 

new methods are developed and validated and as revised methods are evaluated and their 

performance verified. The methods published in NMAM are relied upon by authoritative 

bodies such as accrediting organizations and regulatory agencies. Besides sampling and 

analytical methods, NMAM also includes chapters on quality assurance, portable 

instrumentation, measurement of fibers, aerosol sampler design, and other guidance on 

specific areas of interest.

Often there are situations during use where certain NIOSH methods may require 

modification, for instance, to accommodate interfering compounds from a particular 

workplace, to take advantage of unique laboratory capabilities, to make use of equivalent 

sample preparation or analysis techniques, or to make possible the analysis of a single sample 

for multiple contaminants. When method modifications are made, quality control data 

demonstrating the reliability of the modified method must be obtained, recorded and 

reported. Examples where method modifications might be required include the following:
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-3 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

▪ The volume of air sampled on solid sorbents should be reduced in cases of high vapor

concentration or high humidity and, in some cases, may be increased if such

concentrations are relatively low.

▪ Automation of sample preparation and measurement procedures usually requires

modification of the manual procedure on which the modified method is based.

▪ Chromatographic conditions, including choice of column and detector, can be

modified to eliminate interferences or increase sensitivity during measurement.

▪ Acid mixtures used for sample dissolutions for elemental analysis may require

modification for certain sample matrices that are difficult to dissolve.

For the measurement of each analyte or group of analytes of concern in workplace 

environmental samples or in biological specimens obtained from workers, it is desired to 

produce sampling and analytical methods that will meet the needs of field investigators (e.g., 

industrial hygienists, control engineers or occupational physicians) as well as laboratory 

personnel (e.g., analytical chemists, biochemists, epidemiologists or toxicologists). Many 

NIOSH methods are developed in parallel with related voluntary consensus standards [Ashley 

2015]. The ultimate goal of the formalized NIOSH method development, evaluation and 

validation protocol is to make available sampling and analytical methods for applications in 

the occupational hygiene arena that are fit for purpose, analytically rigorous, and adequately 

ruggedized.

2 How to use NMAM

NIOSH methods are grouped alphabetically by method name, and some method names may 

refer to a group of related substances. It is also possible to locate methods through their 

arrangement by method number. Methods for particular analytes or groups of analytes can 

additionally be accessed by searching their Chemical Abstracts Service (CAS) number(s) 

through the online link.

a. Locating a NIOSH method

Often the easiest and fastest way to locate a method is to refer to the online method index,

which contains an alphabetical listing of analytes and listing by method number. Each

method’s cover page contains information on alternate chemical names and information

on: Compound(s), Method Number, Method Name, Sampling Rate, Minimum Volume,

Maximum Volume, Reagents, Analytical Technique and Sampler (for a quick reference).

It is also possible to search electronically by method number and/or CAS number (if

known).
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-4 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

b. Method numbering system

The general NMAM method numbering system is outlined in the table below. Substances

having the same sampling device, sample preparation procedure and measurement

technique are often grouped together in one method (e.g., organic vapors; metals).

Method No. Substances

0001-0799 General air samples

0800-0999 Bioaerosols 

1000-1999 Organic vapors on charcoal sorbents

2000-3499 Organic vapors on other solid sorbents

3500-3999 Organic vapors on other samplers (e.g., liquids; direct-reading 

instruments)

4000-4999 Organic vapors on diffusive samplers

5000-5999 Organic aerosols

6000-6999 Inorganic gases and vapors

7000-7999 Inorganic aerosols

8000-8999 Biological samples

9000-9999 Bulk samples; wipe samples

c. Indexes and Appendixes

Within the NMAM website there is an online link to indexes that can be used to locate

methods published in previous editions of the Manual:

1) Fourth Edition Methods

An index of fourth edition methods in order of method number. Note that the same

method numbering system is used for third, fourth and fifth edition NIOSH methods.

Also denoted is the current disposition of historical or discontinued methods.

2) First and Second Edition Method Numbers

An index of the first and second edition “P&CAM” and “S” methods, from which

many of the subsequent methods were derived. This index shows the disposition of all

of these earlier methods, whether they were later revised / updated or not.

3) Names and Synonyms

An alphabetical listing of chemical names and synonyms used in current (and many

previous edition) methods, including CAS numbers.
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-5 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

An online “Appendixes” link is also available for obtaining unit equivalents or for 

carrying out air concentration calculations for comparisons to Occupational Safety and 

Health Administration (OSHA) standards.

d. Method format

NIOSH methods consist of three major parts:

1) Front page

The first page of each method concisely summarizes sampling and measurement

parameters and gives estimates of limit of detection, working range, overall and

measurement precision, and interferences. References to other relevant methods are

given. Also provided are Method Classification, NIOSH Registry of Toxic Effects of

Chemical Substances (RTECS) number, and an estimate of method accuracy (see

Figure 1).

2) Instructions

The second page of each method begins with lists of required reagents and equipment.

Please note that these reflect the conditions under which the methods were evaluated

and that there may still be some latitude for variation. The user of the methods is

responsible for assuring the accuracy of the results (e.g., to determine that

breakthrough and recovery are acceptable for each lot of samplers used). For example,

typical tolerances for sorbent tubes are illustrated:

▪ Glass tubing used to contain solid sorbents: Inside diameter is usually not critical

within the range of 4 to 6 mm; length should be sufficient to contain the specified

mass of sorbent.

▪ Contents of sorbent tubes: Mass of sorbent within ±10% of specification; separators

of either glass wool or cleaned polyurethane foam (unless otherwise indicated);

sorbent mesh size of 20/40 unless sampling efficiency dictates otherwise. Filled

sorbent tubes should be sealed to protect them from contamination.

The Special Precautions section gives guidance on safe practices to be observed during 

sampling, sampler preparation and measurement. Next are the step-by-step 

instructions for Sampling, Sample Preparation, Calibration and Quality Control, 

Measurement, and Calculations. Any lengthy instructions for sampler preparation and 

standardization of stock solutions appear in method appendixes. Nomenclature is 

consistent with the NMAM Glossary (chapter) of Abbreviations, Definitions and 

Symbols. (Note that additional general information relating to sampling and 

measurement is contained in other NMAM chapters.)
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-6 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

3) Supporting information

Laboratory and field data relating to the method are summarized in the Evaluation of 

Method section and on the summary page, along with pertinent references.

Figure 1. Layout of front page of NIOSH methods
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-7 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

e. Method classification 

Methods in previous (fourth) edition of NMAM are classified into evaluation categories: 

Full, Partial, Unrated and Not Applicable. Classification is based on the results of 

laboratory testing and evaluation criteria as described in a NIOSH guidelines document 

[NIOSH 1995] and in Chapter ME (Development and Evaluation of Methods). Most 

methods in the fifth edition are classified as ’fully validated.’

The performance data from these evaluations are summarized in the Evaluation of 

Method section in each method. This section may also contain other corroborating data, 

e.g., results from collaborative testing, Proficiency Analytical Testing (PAT) data, or field 

data from NIOSH studies. For partially evaluated methods, this section will state which 

evaluation points were not tested, thus providing the user with information on which to 

make a reasonable judgment on the quality of the data obtained.

Evaluation – Full: Fully evaluated methods are those that have been tested and found to 

have met all of the factors of the NIOSH evaluation protocol [NIOSH, 1995]. 

Evaluation – Partial: Partially evaluated methods are those that have been subjected to 

some of the evaluation experiments but have not received a full evaluation (e.g., shortterm method development). These may also include methods that were fully tested but 

did not meet one or two of the evaluation criteria specified in the NIOSH protocol 

[NIOSH, 1995]; for example, some of the earlier-developed methods that do not meet the 

current ±25% accuracy criterion.

Evaluation – Unrated: Unrated methods have not been tested by NIOSH, but may have 

been developed by a recognized independent source such as OSHA.

Evaluation – N/A: The designation, Not Applicable (N/A), is applied to methods where no 

quantitative data are collected, such as:

▪ Procedures for sample collection only. The collected samples are analyzed 

 subsequently by an appropriate analytical method. 

▪ Qualitative methods that indicate results as a positive or negative (or inconclusive).

f. User experience with NIOSH methods

NIOSH strives to make the methods published in NMAM useful and fit for purpose in 

industrial hygiene analyses. Therefore, feedback on the experiences of people using the 

methods is important to us. Suggestions for improvement and questions relating to 
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-8 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

NMAM are welcome and should be directed to the editors of the Manual. Their contact 

information is provided on the NMAM webpage.

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement 

of the sponsoring organizations or their programs or products. Furthermore, NIOSH is 

not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.

3 References

Angerer J, Greim H, eds. [2006]. Essential biomonitoring methods. Weinheim, Germany: 

Wiley-VCH.

Ashley K [2015]. NIOSH manual of analytical methods 5th edition and harmonization of 

occupational exposure monitoring. Gefahrstoffe Reinh Luft 75(1/2):7-16. 

Behroozy A [2013]. On dermal exposure assessment. Int J Occup Environ Med 4:113-127. 

Brisson M, Ashley K, eds. [2011]. Surface and dermal Sampling (ASTM STP No. 1533). West 

Conshohocken, PA: ASTM International.

DiNardi SR, ed. [2003]. The occupational environment – Its evaluation, control, and 

management, 2nd ed. Fairfax, VA: American Industrial Hygiene Association (AIHA).

Eduard W, Heederik D, Duchaine C, Green BJ [2012]. Bioaerosol exposure assessment in the 

workplace – The past, present and recent advances. J Environ Monit 14:334-339. 

Hathaway GJ, Proctor NH, eds. [2004]. Chemical hazards of the workplace, 5th ed. New 

York: Wiley.

Jakubowski M [2012]. Biological monitoring versus air monitoring strategies in assessing 

environmental – occupational exposure. J Environ Monit 14:348-352. 

Kulkarni P, Baron PA, Willeke K, eds. [2011]. Aerosol measurement – Principles, techniques, 

and applications, 3rd ed. New York: Wiley.
NIOSH Manual of Analytical Methods  5th Edition  Chapter PS April 2016 Page PS-9 of PS-9

Purpose, Scopeand Use of the NIOSH Manual of Analytical Methods

NIOSH [1995]. Guidelines for air sampling and analytical method development and 

evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: 

National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 95-

117. 

Rose VE, Cohrssen B, eds. [2011]. Patty’s industrial hygiene and toxicology, 6th ed. New 

York: Wiley.

Semple S, Cherrie, JW [2003]. Dermal exposure assessment. In: Nieuwenhuijsen MJ, ed. 

Exposure assessment in occupational and environmental epidemiology. Oxford, UK: Oxford 

University Press.

Vincent JH [2007]. Aerosol sampling – Science, standards, instrumentation, and applications. 

New York: Wiley.

Vincent JH [2012]. Occupational and environmental aerosol exposure assessment – A 

scientific journey from the past, through the present and into the future. J Environ Monit 

14:340-347. 
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Developmentand Evaluation

of Methods 

by Eugene R. Kennedy, Ph.D.,Thomas J. Fischbach, Ruiguang Song, Ph.D., Peter M. Eller, Ph.D., 

Stanley A. Shulman, Ph.D., and R. DeLon Hull, Ph.D., NIOSH

1 Method Development ME-2

2 Method Evaluation ME-5

3 Field Evaluation ME-13

4 Documentation ME-14

5 Appendix - Accuracy and its Evaluation ME-15

6 References ME-17
NIOSH Manual of Analytical Methods  5th Edition  Chapter ME April 2016 Page ME-2 of ME-19

Development and Evaluation of Methods 

1 Method development

The development and evaluation of analytical methods that are useful, reliable and accurate 

for industrial hygiene monitoring problems require the application of some general guidelines 

and evaluation criteria. The guiding objective in this work requires that, over a specified

concentration range, the method provide a result that differs no more than ±25% from the 

true value 95 times out of 100. The application of consistent evaluation criteria and guidelines 

is particularly important when methods are developed by different individuals and 

organizations (e.g., contractors or outside laboratories) and compiled into a single manual. 

Adherence to guidelines should minimize overlooking potential problems in the methodology 

during its development, as well as provide cohesiveness and uniformity to the method that is 

developed. This chapter provides an outline of a generalized set of evaluation criteria prepared 

by NIOSH researchers for the evaluation of sampling and analytical methodology [NIOSH 

1995]. 

In the development of a sampling and analytical method, there is a logical progression of 

events that cover a search of the literature to gather pertinent information and the preliminary 

experimentation for selection of analysis technique and sampling medium. To initiate the 

development of a method, the identity of the analyte must be as fully defined as possible. 

Physical and chemical properties of the analyte should be defined so that procedures for 

proper handling and use of the analyte can be prepared. These also aid in establishment of 

analyte purity. Potential sources of this information include chemical reference books, health 

hazard evaluation reports, bulk sample analyses, material safety data sheets, chemical process 

information, etc.

Since innovation is a key element in the sampling and analytical method development 

process, detailed experiments for the initial development of the sampling approach and 

optimization of the analytical procedure are better left to the discretion of the researcher. 

During development, it should be recognized that appropriate, statistically designed 

experiments will optimize the amount of information obtained. Therefore, consultation with a 

statistician about appropriately designed experiments will be of value during this phase of the 

research. 

a. Preliminary experimentation 

Several key points, including calibration and selection of measurement technique and 

sampling media, should be studied during the initial method development experiments. 

The selection of sampling medium and procedure is a decision that usually is made early 

in the method development process. The physical state of the analyte (i.e., gas, aerosol, 

vapor, or combination thereof) plays an important factor in the selection of an appropriate 

sampler. Analytes which can exist in more than one physical state may require a 
NIOSH Manual of Analytical Methods  5th Edition  Chapter ME April 2016 Page ME-3 of ME-19

Development and Evaluation of Methods 

combination of sampling media in one sampler for efficient collection [NIOSH 1995]. 

Where possible, commonly available and easily used samplers should be investigated 

initially. As the preliminary testing of a sampling method progresses, further modification 

in the sampling medium or sampler design may be required and may affect the 

measurement procedure. Sampler design and media selection considerations should 

include U.S. Department of Transportation regulations and restrictions for shipment back 

to a laboratory for analysis.

Since industrial hygiene analytical methods are geared toward measuring personal 

exposure, the size, weight, and convenience of the sampler are important elements in 

sampler design. The personal sampler should allow freedom of movement and should be 

unobtrusive, unbreakable, and not prone to leakage. The pressure drop across the sampler 

should not be so great as to limit sample collection times to 10 h with personal sampling 

pumps. For situations where only a short term sample will be required (i.e., 15 min for 

ceiling determinations), this 10 h recommendations can be reduced to 1 h. The use of 

potentially toxic reagents should be avoided unless they can be used safely. Reagents used 

should not pose any exposure hazard to the worker wearing the sampler or to the 

industrial hygienist taking the samples.

b. Recovery of the analyte from the medium

During the course of method development experiments, the ability to recover the analyte 

from the sampling medium should be determined. A suggested experiment to accomplish 

this entails the fortification of sets of 6 samplers with amounts of analyte equivalent to 

sampling concentrations of 0.1, 0.5, 1.0, and 2.0 (or higher) times the exposure limit for a 

minimum of 4 h at the typical sampling rate used for that type of sampler. If the analyte 

has a ceiling or short-term exposure limit, the amount of analyte fortified should be 

adjusted for the shorter sampling time required for this type of exposure limit. If the 

sampler has a backup section, then a like number of separate backup sections should be 

fortified with amounts of analyte equivalent to 25% of the amount fortified on the front 

sections of the samplers, since this amount has been used to characterize the breakthrough 

limit of useful samples [Streicher et al. 1994]. Samples (and backup sections) should be 

prepared for analysis and analyzed according to previously determined procedures. 

Results of these analyses should be expressed in terms of estimated percent recovery 

according to the following formula:

Percent Recovery (est.) =� AAAAAAAAAAAA AAoo aaAAaaaaaaAAaa ooAAAAAAff AAAA ssaaAAssaaaass

AAAAAAAAAAAA AAoo aaAAaaaaaaAAaa ooAAssAAffooffaaff AAAA ssaaAAssaaaass� × 100 %

After initial analyses of the samples, the samples should be resealed and analyzed on the 

following day, if possible. If the sample workup procedure results in a solution of the 
NIOSH Manual of Analytical Methods  5th Edition  Chapter ME April 2016 Page ME-4 of ME-19

Development and Evaluation of Methods 

sample, these solutions should be recapped after the initial analysis, if possible, and 

reanalyzed on the following day using fresh standards.

The recovery of the analyte should be calculated for the primary and backup media in the 

sampler. Although complete recovery of the analyte from the sampler is most desirable, at 

a minimum, the estimated recovery of the analyte from the primary collection medium 

should be greater than or equal to 75% for concentrations equivalent to sampling 0.1, 0.5, 

1.0, and 2.0 times the exposure limit. If recovery varies with analyte loading, results should 

be graphed as recovery versus loading during calibration of the method, so that 

appropriate correction can be made to sample results, as long as recovery is greater than 

75% [Melcher et al. 1978]. If estimated recovery does not exceed 75%, the method is not 

suitable for monitoring at this limit.

Estimated recovery from any backup media should be noted so that appropriate 

corrections can be applied if breakthrough of the sampler has occurred during sampling. 

The recovery of the analyte from the medium in the backup section of a sampler may be 

different from that of the front section, since the backup section of a sorbent-based 

sampler usually contains only half of the sorbent of the primary section. If the same 

volume of desorption solvent is used for both the primary and backup sections of the

sampler, the desorption equilibrium can be shifted, since the backup section is being 

desorbed by twice the volume (i.e., on a mL solvent/mg sorbent basis) [Saalwaechter et al. 

1977]. 

Reanalysis of the samples on the day after initial analysis indicates if immediate analysis 

after sample preparation is required. Often when processing a large number of samples, it 

may be necessary to prepare the samples for analysis as a batch. In these instances, the last 

samples may not be analyzed for up to 24 h or more after preparation because of the time 

required for analysis. If samples prepared for analysis exhibit time-dependent stability 

after desorption, analyses must be conducted within acceptable time constraints. Analysis 

and reanalysis results should agree within 5% of each other.

c. Stability of the analyte on the medium

An extension to the experiment described above may be performed to investigate potential 

stability problems early in the experimentation. An additional set of fortified samples at 

each of the 4 concentrations should be prepared and analyzed after 7-days' storage at room 

temperature. Recovery should be similar to the above results within experimental error. 

Discrepancies larger than those expected by experimental error indicate sample stability 

problems that will need correcting by additional developmental effort (e.g., refrigerated 

storage). Comparison of results can be performed with statistical tests, such as an analysis 
NIOSH Manual of Analytical Methods  5th Edition  Chapter ME April 2016 Page ME-5 of ME-19

Development and Evaluation of Methods 

of variance (ANOVA) [Posner and Okenfuss 1981] test of the “Day” difference or a paired 

t-test [Box et al. 1978] of the means of the Day 1 and Day 7 storage results.

2 Method evaluation

After the initial development experiments for the method have been completed and a method 

has been proposed, the sampling and analysis approach should be evaluated to ensure that the 

data collected provides reliable, precise, and accurate results. Specifically, the goal of this 

evaluation is to determine whether, on the average, over a concentration range of 0.1 to 2 

times the exposure limit, the method can provide a result that is within ±25% of the true 

concentration 95% of the time. For simplification, the true concentration is assumed to be 

represented by an independent method. An experimental approach for collecting the data 

necessary for this determination is described below.

As part of the evaluation of a method, the sampling of a generated atmosphere is needed to 

more adequately assess the performance of a method [NIOSH 1984; Nelson 1971; Nelson 

1992]. This allows the determination of 1) the capacity of the sampler; 2) the efficiency of 

analyte collection by the sampler; 3) the repeatability of the method; 4) the bias in the method; 

5) interferences in the collection of the sample. Concentration ranges to be used in the 

evaluation of the method should be based on several factors. These ranges, at a minimum, 

should cover 0.1 to 2.0 times the exposure limit. In some instances, higher multiples of the 

exposure limit can be added if needed (e.g., 10 times the exposure limit). In situations where 

multiple exposure limits (i.e., from different authorities) exist for an analyte, the lowest 

exposure limit should be used to set the lower limit of the evaluation range (0.1 times lowest 

exposure limit) and the highest limit used to calculate the upper limit of evaluation range (2 

times the highest exposure limit). Intermediate evaluation concentrations should be within 

these exposure limits. The toxicity of an analyte (e.g., suspected carcinogenicity) may indicate 

that a concentration lower than that calculated by the exposure limit should be included in the 

measurement and evaluation ranges. Previous monitoring information from other methods 

may indicate that typical concentrations of the analyte may be below or above a concentration 

range based on the exposure limit. In this case, this lower or upper level may be included in 

the method evaluation.

a. Feasibility of analyte generation 

In order to provide a realistic test of the method under study, air concentrations covering 

the range from 0.1 to 2 times the exposure limit of the analyte should be generated. The 

generated atmospheres should be homogeneous in concentration and representative of the 

environment encountered when sampling for the analyte in the workplace.
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Development and Evaluation of Methods 

When attempting to generate a concentration of an analyte, the impact of environmental 

conditions, such as temperature, pressure, humidity, and interferences, on sampler 

performance and/or generation should be considered. The effect of elevated temperature 

on the collection medium of a sampler may decrease the capacity of the sampler or may 

decompose the analyte during generation and sampling. Reduced pressure may also 

reduce the capacity of a sampler. High relative humidity in many instances has been 

observed to reduce sampler capacity [Melcher et al. 1978]. In other instances it has 

increased sampler capacity [Cassinelli 1991]. A typical interference(s) should be generated

along with the analyte to approximate a typical workplace sampling environment.

Generation of particulate material can be extremely complex [Willeke 1980; Hinds 1982], 

especially if particles of a required size range must be generated for the evaluation of a 

specified sampler inlet design. The aerodynamic performance of the generator is a factor 

in the generation of this type of atmosphere and should be evaluated carefully. 

Appropriate, independent methods should be available to verify particle size, if this is a 

critical element in the generation.

The concentration of the generated atmosphere should be verified either by well 

characterized gravimetric/volumetric means or by analysis of replicate samples (if 

possible) by an independent method at each concentration used. Further details on this 

verification are included in the literature [NIOSH 1995; Ashley 2015]. A statistician should 

be consulted for advice on the design and sample sizes to accomplish this validation. 

Ideally, the independent method should not be biased and should provide an accurate 

estimate of the concentration generated, assuming error is randomly distributed around 

the mean. Also the precision and bias of the independent method should be homogeneous 

over the concentrations investigated. (See NIOSH [1995] for the definitions of these 

attributes.) In instances where the concentration of the generator can be based only on 

calculations using flow rates in the generator and the amount of analyte injected, the 

generation system should be well characterized so that analyte losses are minimized.

In some instances, generation of an analyte may be difficult and even hazardous. As an 

alternative to direct generation in these cases, samplers may be fortified with an amount of 

analyte expected to be sampled over a specified period of time at a specific flow rate. When 

this is necessary, fortification of the sampler by vaporization of a known amount of analyte 

onto the sampling medium is a more appropriate method, since this approach more 

closely approximates a generated atmosphere. The alternative of direct application of a 

solution of analyte onto the collection medium is less desirable but may be necessary in 

some instances. After fortification, air, conditioned at both high and low humidity, should

be drawn through samplers at the flow rate and time period used in the calculations for the 
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Development and Evaluation of Methods 

amount of analyte expected to be collected. In the method report, the fact that samples 

were not collected from a generated atmosphere should be discussed.

b. Capacity of the sampler and sampling rate 

To determine the applicability of the sampling method, the capacity of the sampler should 

be determined as a function of flow rate and sampling time. This is particularly important 

if the analyte has both a short-term exposure limit (STEL) and a time-weighted average. 

Flow rates typical for the media selected should be used. These may range from 0.01 to 4 

L/min, depending on sampler type. At extremely low flow rates (ca. 5 mL/min), the effect 

of diffusion of the analyte into the sampler must be considered. Flow rates should be kept 

at a high enough rate to prevent diffusion from having a positive bias in the sampler. 

Sampling should be performed at three different flow rates covering the range appropriate 

for the particular sampler type, unless the sampler is designed to operate at only one flow 

rate.

Sampling times should range from 22.5 min for STELs to 900 min (15 h) for timeweighted averages. Shorter sampling times (e.g., 7.5 to 22.5 min) may be used for ceiling

(C) measurements. Flow rates should be based on accurately calibrated sampling pumps 

or critical orifices. The amount of analyte collected at the lowest flow rate and shortest 

sampling time should be greater than the limit of quantitation of the method. The 

generated concentration used for capacity determination should be at least 2 times the 

highest published exposure limit and verified by an independent method.

Sampling should be conducted at ambient, elevated (>35 °C), and low (<20 °C) 

temperatures to assess the effect of temperature on sampling. To assess the effect of 

humidity on capacity, sampling should be performed at both low and high humidity (20% 

and 80%), since both have been observed to affect capacity [Cassinelli 1991; Melcher et al. 

1978]. Triplicate samplers at three different flow rates should be included to verify capacity 

at each of the six different humidity and temperature levels. For samplers which contain 

backup sampling media, only the front section of the sampler should be used. A means is 

required to quantitate analyte in the effluent from the sampler. This may involve the use of 

a backup sampler, continuous monitor or other appropriate means which can provide a 

measure of analyte concentration in the sampler effluent (ca. 1 to 5% of the influent 

concentration). If the mass of analyte found on a backup sampler totals 5% of the mass 

found on the front sampler or if the effluent concentration of the sampler contains 5% of 

the influent concentration, breakthrough has occurred and the capacity of the sampler has 

been exceeded.
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Development and Evaluation of Methods 

If the analyte is a particulate material and collected with a filter, the capacity of the filter is 

defined by the pressure drop across the sampler or by the loading of the filter. For 37-mm 

filter-based samplers, pressure drop should be less than 1000 mm of water for total loading 

less than 2 mg. Larger filters and especially filter capsules [Harper and Ashley 2013; 

O’Connor et al. 2014] will tolerate higher loadings (e.g, up to 5 mg).

If the collection process is based primarily on adsorption, breakthrough time should be 

proportional to the inverse of the flow rate [Jonas and Rehrmann 1973]. This relationship 

can be checked by plotting the 5% breakthrough time versus the inverse of the flow rate. If 

the resulting plot is a straight line, then this relationship should hold for all flow rates in 

the flow rate range studied. Some nonlinearity in the plot may be noted due to 

experimental variability and assumptions made to simplify the relationship of 

breakthrough time and flow rate. Results from these experimental trials should provide a 

prediction of the capacity of the sampler at various flow rates and sampling times. If the 

flow rates and sampling times used in the experiment do not provide for sufficient 

capacity, a lower flow rate range may have to be studied and the experiment repeated.

With samplers which use reagents for collection of the analyte, the amount of the reagent 

in the sampler will also be a limiting factor in the capacity of the sampler, based on the 

stoichiometry of the reaction. Other factors, such as residence time in the sampler and 

kinetics of reaction between analyte and reagent, may affect the capacity of this type of 

sampler.

The combined temperature and humidity conditions that reduce sampler capacity to the 

greatest extent should be used in all further experiments. The Maximum Recommended 

Sampling Time (MRST) for a specific flow rate is defined as the time at which sampler 

capacity was reached, multiplied by 0.667. This adds a measure of safety to this 

determination. The relationship of breakthrough time with flow rate can be used to adjust 

flow rates to optimize specific sampling times.

c. Sampling and analysis evaluation

To assess the performance of a method, certain additional experimental parameters should 

be evaluated through a series of defined experiments. The effect of environmental 

conditions (e.g., pressure, interferences) on sampling efficiency of the sampling medium 

can be evaluated by a factorial design [Box et al. 1978]. The temperature, relative humidity, 

flow rate, and sampling times, determined in the experiment described above to have most 

severely limited sampler capacity, should be used in these experimental runs [Ashley 

2015]. At a minimum, the effect of concentration on method performance should be 

investigated. Three sets of 12 samples should be collected from an atmosphere containing 
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Development and Evaluation of Methods 

concentrations of 0.1, 1.0, and 2.0 times the exposure limit at the humidity determined 

above to have reduced sampler capacity for the MRST determined in the preceding 

experiment.

If the analyte has a short-term or ceiling exposure limit in addition to an 8-hour timeweighted average, an additional 12 samplers should be collected at the STEL or C limit for 

the recommended sampling period at the appropriate flow rate. Potential interferences in 

the work environment should be included in the generation experiments to assess their 

impact on method performance. Concentrations up to 2 times the exposure limit value for 

the interference should be included. Other environmental factors may be studied, but will 

require a more comprehensive experimental design.

The effects of environmental conditions on analyte recovery should be assessed. A 

factorial design can be used to evaluate these factors to determine which exert a significant 

effect on analyte recovery. Those factors which are found to influence analyte recovery 

should be investigated further to determine if their impact is predictable. If these effects 

are not predictable, the utility of the method will be limited, based on the conditions 

defined by this experiment. If only concentration is evaluated, the analyte recovery should 

be the same at all concentrations after correctable biases have been included, such as 

desorption efficiency.

d. Sample stability

To assess sample stability, samples should be collected from a generated atmosphere, 

stored under defined conditions (i.e., ambient or refrigerated, light or dark), and analyzed

at specified time periods. A concentration of 0.5 times the lowest exposure limit should be 

sampled with 30 samplers for a minimum of 1⁄2 the MRST. The humidity and temperature 

of the generator should be at the same level as defined in the sample capacity experiment 

to reduce sample capacity. The samplers should be divided randomly into one group of 12, 

one group of 6, and four groups of 3, with the group of 12 analyzed as soon after collection 

as possible (Day 0). The group of 6 samplers should be analyzed after 7 days. The four 

remaining sets of 3 samplers should be analyzed after 10, 14, 21, and 30 days. The 

conditions of storage are determined by the nature of the analyte. If there is an indication 

of analyte instability on the sampling medium, refrigeration of the samplers may be 

required. However, storage for the first 7 days should be at room temperature.

Samples should normally be stable for a minimum of 7 days under ambient conditions to 

simulate shipping to a laboratory for analysis. If the average analysis results of the 

samplers analyzed on day 7 differs from the set analyzed on day 0 by more than 10%, the 

method does not meet the sample stability criterion. Either additional precautions, such as 
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Development and Evaluation of Methods 

shipment on ice and refrigerator storage, may be required or the method may have to be 

modified to address this problem. (Note: In practice, reactive or unstable samples should 

be shipped by overnight mail and analyzed as quickly as possible.) If a plot of recovery 

versus time indicates that recovery decreased by more than 10% after the initial 7-day 

storage period, sample instability is a problem. If samples need to be stored for longer 

periods, more restrictive storage conditions are required. Remedial action, such as cold 

storage may solve this longer term storage problem. After remedial precautions have been 

instituted in the method, the sample stability of the method must be determined anew.

e. Precision, bias, and accuracy

Results from four sets of samplers used in the analyte recovery experiment, the sampling 

and analysis experiments (e.g., the environmental parameters experiments), and the 

sample stability experiment can be used for the estimation of precision, bias, and accuracy 

of the method. A more exacting treatment of this is described elsewhere [NIOSH 1995]. 

Sampler results from the multi-level factorial design at the 0.1, 1.0, and 2.0 times the 

exposure limit value; the sampler stability experiment (at 0.5 times the exposure limit); 

and the environmental factors experiment are used in the calculations of method 

precision. The calculations for the estimated method precision, ŜrT, have been described 

previously [NIOSH 1995; Anderson et al. 1981; Busch and Taylor 1981; NIOSH 1980]. 

Before obtaining a pooled estimate of method precision from the four sets of samplers 

listed above, the homogeneity of the precision over the range of concentrations studied 

should be checked using a test, such as Bartlett's test [NIOSH 1995; Anderson et al. 1981; 

Busch and Taylor 1981]. If the precision is not found to be constant over concentrations, 

the sample set collected at 0.1 times exposure limit should be removed and Bartlett's test 

recalculated. Homogeneity of the method precision at all concentration levels is an 

assumption required to obtain pooled estimate of method precision.

Bias is assumed to be homogeneous over the evaluation range. This assumption should be 

tested by estimating the bias at each concentration and testing these for homogeneity 

using the procedures described in the literature [NIOSH 1995]. Method bias should be less 

than 10%. A test for this is also described [NIOSH 1980].

The bias and precision estimates can be used with the graph presented in Figure 1 or in 

Table I to estimate accuracy [NIOSH 1995]. The bias and precision estimates are plotted 

on the x- and y-axes of the graph. The intersection of these points on the parabolic grid in 

the graph can be used to estimate the accuracy of the method. This procedure gives an 

estimate of method accuracy but does not yield the statistic required to test compliance of 

the method with the ±25% accuracy criterion. Techniques for the latter determination are 

discussed in the Appendix and elsewhere [NIOSH 1995]. 
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Development and Evaluation of Methods 

If the results for 4 concentrations fail the 25% accuracy criterion, then the set of samples

collected at the lowest concentration level should be excluded from the data set. The 

pooled ŜrT and the bias should be recalculated on this reduced data set before performing 

the accuracy analysis described in the previous paragraph.

For the 12 samplers collected at the ceiling limit, the accuracy analysis described above 

should be repeated using only the data collected at the ceiling limit.

Figure 1. Nomogram relating accuracy to precision and bias. Accuracy (A), in 

percentage units, is a function of the bias (B) and the precision (SrT). Each curve is the 

locus of all points (B, SrT) that yield the value of A indicated on the curve.
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Development and Evaluation of Methods 

Table I. Values of the bias (B) and the precision (SrT) required to obtain designated 

values of accuracy (A) in percentage units#

A (%) B (%) Srt (%)

5 -3.5 0.945*

5 -2.5 1.559*

5 0.0 2.551*

5 2.5 1.483*

5 3.5 0.881*

10 -7.5 1.643*

10 -5.0 3.200

10 0.0 5.102

10 5.0 2.895*

10 7.5 1.414*

15 -10.0 3.378*

15 -5.0 6.381

15 0.0 7.653

15 5.0 5.774

15 10.0 2.764*

20 -10.0 6.755

20 -5.0 9.448

20 0.0 10.20

20 5.0 8.548

20 10.0 5.527

25 -10.0 10.13

25 -5.0 12.39

25 0.0 12.75

25 5.0 11.21

25 10.0 8.287

30$ -15.0& 10.73

30$ -7.5 14.55

30$ 0.0 15.31

30$ 7.5 12.52

30$ 15.0& 7.930

35$ -15.0& 14.31

35$ -7.5 17.59

35$ 0.0 17.86

35$ 7.5 15.14

35$ 15.0& 10.57

# Note: the values shown in this table are population or theoretical values.

* Below the minimum attainable precision with a 5% pump correction.

$ Does not fulfill the Accuracy Criterion (±25% of the true value).

& Does not fulfill the bias criterion (±10%).
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Development and Evaluation of Methods 

3 Field evaluation

While field evaluation is not required in method evaluation, it does provide a further test of 

the method, since conditions which exist in the field are difficult to reproduce in the 

laboratory. Also unknown variables may affect sampling results when field samples are taken. 

This type of evaluation is recommended to further study the performance of the method in 

terms of field precision, bias, interferences and the general utility of the method. Both the 

collection of area samples and personal samples should be included in the field evaluation of 

the method. 

Area samples should provide an estimate of field precision and bias. Personal samples may 

confirm these values and also provide a means to assess the utility of the method. A statistical 

study design should be prepared, based on the variability of the method and the statistical 

precision required for estimates of the differences in analyte concentrations yielded by the 

independent method and the method under evaluation [CEN 2015].

If this type of statistically designed study is not feasible, a minimum of 20 pairs of samples of 

the method under study and an independent method should be used for personal sampling. 

Placement of the samplers on the workers should be random to prevent the biasing of results 

due to the "handedness" of the worker. Workers sampled should be in areas where both low 

and high concentrations of the analyte may be present.

As a minimum, sets of 6 area samplers paired with independent methods should be placed in 

areas of low, intermediate, and high analyte concentration. If the atmosphere sampled is not 

homogeneous, precautions may have to be taken to ensure that all samplers are exposed to the 

same concentrations. This can be done by using field exposure chambers, such as those 

described in the literature [Cassinelli et al. 1985; Kennedy et al. 1985].

Field precision and bias of the area sampler results of the method under study should compare 

with laboratory evaluation results, provided that precautions have been taken to ensure that 

all samplers have been exposed to the same homogeneous atmosphere. Differences in 

precision and bias should be investigated. Sources of variation should be studied and 

corrections implemented where necessary. Evaluation of personal sampler results should be 

done cautiously, because observable differences may be due to work practices or other 

situations which are beyond the control of the method.

A field evaluation of a method also allows the developer of the method to determine its 

ruggedness. Although this may be a subjective judgement, first-hand experience with the 

method in the field may suggest changes in the sampler or method that may make the method 

more easily used in the field and less subject to variability.
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Development and Evaluation of Methods 

4 Documentation

Development and evaluation research on a sampling and analytical method should be 

documented in a final report. The report should describe what was determined about the 

method. If the results of the statistical analysis of the data indicate there is not 95% confidence 

that the accuracy of the method is less than or equal to ±25%, the report should state this fact. 

In some instances, the method may actually have an accuracy of less than 25%, but a larger 

sample size must be used to prove this statistically (See Appendix 1 of NIOSH 1995). 

The final report can be either a technical report or a failure report. The technical report 

(acceptable method developed) documents the successful development of the method. This 

report may be prepared in a format appropriate for submission to a peer-reviewed journal for 

publication. The failure report (no acceptable method developed) documents the research 

performed on an attempted method development for an analyte or analytes. The report 

should describe the failure of the method, as well as other areas of the method research that 

were successful. Recommendations to solve the failure of the method may be included.

If an acceptable method is developed, a sampling and analytical method should be prepared in 

appropriate format. The format of the resulting method should provide clear instructions for 

the use of the method. Sampling, sample workup, and analysis procedures should be clearly 

described. The necessary equipment and supplies for the method should be listed clearly in 

the method. A summary of the evaluation of the method should be included, as well as a 

discussion of method applicability and lists of interferences and related references. As a check 

on the clarity and performance, new methods should be reviewed and submitted to a user 

check (i.e., the method is used to analyze spiked or generated samples of known concentration 

by someone other than the researcher who developed it) and to a collaborative test, if feasible.
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Development and Evaluation of Methods 

5 Appendix - Accuracy and its evaluation

In the development of a sampling and analytical method, one of the goals is to minimize the 

measurement error to the lowest feasible and practical levels. It is assumed that all feasible 

corrections to reduce error have been made in the laboratory experimentation process. 

Method evaluation requires adequate characterization of the magnitude and distribution of 

the uncorrectable error that cannot be prevented. One might consider a hypothetical 

experiment in which a method is used repeatedly to measure the same concentration, T, 

under the same conditions. These measurements would tend to exhibit a pattern or statistical 

distribution, here assumed to be normal, with a mean, μ, and standard deviation, Š. The 

distribution can be characterized in terms of two components: its location relative to T, which 

is the systematic error termed bias (B), is given by (μ-T)/T; and its spread, which is the 

random error termed imprecision (ŜrT), is given by Š/μ. The bias and imprecision are used to 

determine the inaccuracy of the method but they are also important characteristics of the 

error in and of themselves, as will be discussed below.

Accuracy refers to the closeness of the measurements to T but it is defined in terms of the 

discrepancy of the measurements from T. Inaccuracy (I) is defined as the maximum error, 

regardless of sign, expressed as a percentage of T that occurs with a probability of 0.95. Thus, 

an inaccuracy (or accuracy) of 20% means that on the average 95 of every 100 measurements 

will differ from T by no more than 0.2T. The accuracy criterion for single measurements 

mentioned at the beginning of this chapter, often termed the “NIOSH Accuracy Criterion,” 

requires inaccuracy to be less than or equal to 25%.

Accuracy, bias, and imprecision have the following relationship:

0.95 = φ((1-B)/((1+B)SrT))- φ((-1-B)/((1+B)SrT)) 

where (φ) denotes the probability that a standard normal random variable is less than or equal 

to φ. A practically exact numerical solution to Equation (1) can be readily programmed using 

statistical software packages [Press et al. 1986]. A DOS program, ABCV.EXE, is also available 

which solves for I (denoted by A in the program), ŜrT (denoted by CV in the program), or B 

when the values for the other two quantities are input. An estimate of I can be obtained in 

either case by entering estimates of B and ŜrT. An approximate solution, which is accurate to 

about 1.1 percent, is given as follows [NIOSH 1995]:

I= 1.57 (B+1)SrT + �((0.39 (B + 1) ∙ SrT)2 + B2) for theoretical or true I 

Î= 1.57 (BB� +1)ŜrT + " �((0.39 (B� + 1) ∙ S�rT)2 + B�2) for estimates of I
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Development and Evaluation of Methods 

Also, the nomogram in Figure 1 can be used to solve for I or an estimate of I by entering B and 

SrT or their estimates. Procedures for obtaining “best” single point and 95% confidence 

interval estimates of B, and SrT and a 90% confidence interval estimate for I are given in 

NIOSH [1995].

The 90% confidence interval for I can be used to infer whether the method passes or fails the 

25% accuracy criterion (AC) for single measurements with 95% confidence as follows:

1) The method passes with 95% confidence if the interval is completely less than 25%.

2) The method fails with 95% confidence if the interval is completely greater than 25%.

3) The evidence is inconclusive if the interval includes 25% (there is not 95% confidence that 

the AC is true or that it is false).

When researchers interpret the results from analyses of the type described above, it is 

important to consider that most methods have many uses in addition to individual 

measurement interpretation. Because accuracy is very important whenever any quantity is to 

be estimated, the ideal (“other things being equal”) is to use the most accurate estimator 

regardless of its bias or imprecision. However, it is crucial to distinguish between the accuracy 

of the source or “raw” measurements and that of the final estimator, which might involve 

many intermediate analyses or operations. Unfortunately, the most accurate input or raw 

measurements do not always produce the most accurate final result unless the latter is a single 

measurement. The bias and imprecision of the source measurements can be differentially 

affected by intermediate operations in producing the final estimate. For example, if the final 

estimate is a function of a single average of many source measurements, its bias is not affected 

by the averaging while imprecision is reduced as a function of the square root of the number 

of measurements. Thus, a lower biased method might be preferable to another even if the 

inaccuracy of the latter is less. On the other hand, in comparative studies, the desired estimate 

is either a difference or ratio of means of measurements in which there can be partial or 

complete cancellation of the bias in the source measurements. Thus, the bias of the method 

used for the source measurements may be of little importance. If there are several methods 

applicable for a given user’s project (regardless of whether all fulfill the accuracy criterion for 

single measurements), the analyst would be well-advised to consult with the user (preferably 

in advance of measurement) to determine which of those methods would produce the optimal 

accuracy for the final results or estimates needed by that particular user. Accuracy, bias, and 

imprecision jointly form a complete or sufficient set for the efficient description of the 

measurement error characteristic of any method.
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Development and Evaluation of Methods 

Disclaimer 

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

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Press WH, Flannery BP, Teukolsky SA, Vettering WT [1986]. Numerical recipes – the art of 

scientific computing. Cambridge, UK: Cambridge University Press.

Saalwaechter A, McCammon C, Roper C, Carlberg K [1977]. Performance of the NIOSH 

charcoal tube technique for the determination of air concentrations of organic vapors. Am 

Ind Hyg Assoc J 38:476-486. 
NIOSH Manual of Analytical Methods  5th Edition  Chapter ME April 2016 Page ME-19 of ME-19

Development and Evaluation of Methods 

Streicher R, Kennedy E, Lorderau C [1994]. Strategies for simultaneous collection of vapours 

and aerosols with emphasis on isocyanate sampling. Analyst 119:89-97. 

Willeke K, ed. [1980]. Generation of Aerosols and Facilities for Exposure Experiments. Ann 

Arbor, MI: Ann Arbor Science Publishers.
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Measurement Uncertaintyand

NIOSH Method Accuracy Range

by David L. Bartley, PhD, Stanley A. Shulman, PhD, and Paul C. Schlecht, MS, NIOSH

1 Introduction UA-2

2 ISO GUM UA-4

3 The symmetric accuracy range A as used by NIOSH UA-6

4 Uncertainty and analytical lab procedures UA-9

5 Discussion UA-13

6 Technical notes UA-14

7 References UA-20

8 Terminology UA-22
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Measurement Uncertainty and NIOSH Method Accuracy Range

1 Introduction

Recently, the ISO Guide to the Expression of Uncertainty in Measurement (GUM) has come 

close to being universally adopted as the standardized way to characterize and document 

measurement uncertainty [ISO 2002, 2005, 2009, 2010; Ellison and Williams 2012]. Since the 

mid-1970s, accuracy criteria have been an integral part of the evaluations of the sampling and 

analytical methods used by the National Institute for Occupational Safety and Health 

(NIOSH), the Occupational Safety and Health Administration (OSHA), the Mine Safety and 

Health Administration (MSHA), and others. NIOSH has previously published extensive 

discussions addressing the issue of accuracy as a factor in the development, evaluation, and 

characterization of analytical methodology. Both traditional method accuracy and new 

measurement uncertainty concepts are intended to communicate measurement limitations to 

laboratory clients. Naturally, laboratories are interested in how NIOSH accuracy requirements 

[Busch 1977; NIOSH 1995] relate to measurement uncertainty.

This chapter provides guidance for achieving consistency in determining measurement 

uncertainty by those laboratories using NIOSH methods. Minor modifications to NIOSH 

accuracy measures, and an expansion of ISO GUM to cover situations unique to workplace 

atmospheric measurement can improve consistency and utility. See Bartley [2004] for 

additional information.

ISO GUM proposes pooling estimated variance components from diverse error sources. The 

square root of the pooled variance estimate is termed the combined uncertainty uc. 

Multiplication of uc by a coverage factor k (generally in the range of 2 to 3) results in an 

expanded uncertainty U. The purpose of the expanded uncertainty is for each measurement to 

provide an interval bracketing the measurand (the true value of what is to be measured) to 

account for errors in both the measurement and the determination of the uncertainty 

components themselves.

ISO GUM is somewhat unclear about the coverage factor k. Furthermore, the coverage factor 

can be interpreted in several ways. Most straightforward is the limited case where the 

uncertainty components can be re-evaluated each time the method is used (resulting in k

proportional to a Student-t quantile). In this case, the covering intervals bracket the 

measurand for (for example) 95% of the measurements.

Alternatively, the coverage factors based on the Student-t quantile specify intervals containing 

measurand values at levels of evaluation confidence in the mean (i.e., averaging over many 

method evaluations). In other words, for roughly 50% of method evaluations, intervals used at 

each measurement contain the measurand value greater than (for example) 95% of the time. 

The concept is consistent with the statistical theory of tolerance or prediction intervals.
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Measurement Uncertainty and NIOSH Method Accuracy Range

This approach is important to industrial hygiene since workplace air concentrations vary 

spatially and over time to such a degree that a method cannot be evaluated by simply taking 

replicate measurements [Vaughan et al. 1990]. However, industrial hygiene measurement 

methods have traditionally required confidence levels greater than 50% in the method 

evaluation. Generally, 95% confidence in a method validation is required. The different types 

of confidence levels are reflected simply in the numerical value and interpretation of the 

coverage factor.

Of equal importance in the industrial hygiene field are details needed to handle systematic 

error (bias) relative to reference concentration measurements found during method 

evaluation. For example, the sampling rate of a given diffusive sampler for gases or vapors is 

generally measured once by the diffusive sampler manufacturer prior to use by multiple 

clients. As the samplers are not re-calibrated for each use, residual bias exists in the 

measurements due to uncertainty in sampling rates used [ASTM 2013a]. (NIOSH methods 

typically do not cite performance for passive samplers because agreement among diffusive 

monitor manufacturers on test protocols has not yet been achieved, and a system of third 

party evaluation of diffusive monitor manufacturers sampling rates is not available.) Similarly, 

the calculation of desorption efficiencies may be performed only once or infrequently and can, 

therefore, introduce residual bias in measurements that use sorbent-captured samples, e.g., 

charcoal tubes.

In aerosol sampling, detailed knowledge of the particle size-dependent bias of a sampler 

relative to a sampling convention, such as adopted by ISO/CEN/ACGIH/ASTM [ISO 1995; 

CEN 1993; ACGIH 2015; ASTM 2013b] for defining respirable dust, is often necessary to 

judge the usefulness of a given sampler. Each type of aerosol sampler is characterized by 

specific particle collection characteristics, and some analytical methods (e.g. silica) may also 

exhibit particle size effects. Typically the issue of aerosol sampler bias is avoided or minimized 

in the industrial hygiene field by narrowing use to a specific aerosol sampler. For example, 

common industrial hygiene practice establishes a single sampler type, such as the 1.7 L/min 

10-mm nylon Dorr-Oliver cyclone, for respirable dust sampling in a particular application.

Sensitivity to other environmental factors, referred to in ISO GUM as influence variables, 

must be acknowledged. Suppose a sampler is sensitive to temperature changes that are 

impractical to measure in the field; i.e., sampler estimates are not temperature corrected. 

Then, suppose during method evaluation in the laboratory, measurement of this sensitivity is 

combined with knowledge of the expected temperature variation for a given field application. 

Putting together both would determine the uncertainty associated with the effect. Examples of 

the important effects of influence variables - such as wind velocity, temperature, pressure, and 

fluctuating workplace concentrations - on diffusive monitor uptake rates are common.
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Measurement Uncertainty and NIOSH Method Accuracy Range

2 ISO GUM

ISO GUM presents several concepts. One of these calls for the identification of sources 

(labeled j = 1, 2,...) of uncertainty uj (standard deviation estimate components) in a 

measurement method and for their classification into Type A or Type B uncertainties. Type A 

uncertainty is one that has been characterized by a statistically sound approach. In this

case, uujj

2 is given by ssjj2, an unbiased estimate (with vvjj degrees of freedom) of variance σσjj2. On 

the contrary, Type B uncertainty generally requires professional judgment. See Table 1 for 

examples of possible uncertainty components.

A common example of Type B uncertainty is the conservative assignment of a 5% relative 

standard deviation component (without error, i.e., with infinite degrees of freedom) as the 

random sampling pump uncertainty. As described in ISO GUM, such an assignment would be 

a result of sampling pump random errors that had a uniform distribution and fell within 

±√3 × 5% of zero with a probability “for all practical purposes equal to one”. Therefore, if it 

is judged that sampling pump variations are within these bounds, then the assignment of 5% 

as the relative standard deviation component is conservative. Other similar ways of handling 

Type B uncertainties are found in ISO GUM.

Table 1. Examples of potential uncertainty component sources.

Sampling

personal sampling pump flow rate: setting the pump and subsequent drift sampling rate 

of diffusive sampler

sampler dimension (aerosol and diffusive sampling)

Sample handling

sample preparation (e.g., handling silica quasi-suspensions) sample loss during 

transport or storage

Analytical

aerosol weighing

recovery (e.g, GC-based methods)

Poisson counting (e.g., in XRD methods)

Sensor variation

operator effects giving inter-lab differences (if data from several labs are to be used)
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Measurement Uncertainty and NIOSH Method Accuracy Range

Evaluation

calibration material uncertainty

evaluation chamber concentration uncertainty

other bias-correction uncertainty

Environmental influence parameters

temperature (inadequacy of correction, if correction is made as with diffusive samplers) 

atmospheric pressure

humidity

aerosol size distribution (if not measured by a given aerosol sampling method)

ambient wind velocity

sampled concentration magnitude itself (e.g., sorbent loading)

Within the field of industrial hygiene, the quantities uj are often standard deviation 

component estimates obtained from a single measurement-method evaluation, rather than 

from replicates. When the estimates are independent, a combined uncertainty uc may be 

computed (through the propagation of uncertainty approximation) as:

uucc = �uu1

2 + uu22 + ⋯ (1) 

Through a coverage factor k, generally approximated conservatively (e.g., see Technical Note 2 

at end of chapter), as equal to 3 for a single method evaluation, an expanded uncertainty U 

may be computed as:

UU = kk × uucc (2) 

The purpose of the expanded uncertainty U is to provide intervals, which generally contain 

measurand values (often referred to as the true values). In particular, given a concentration 

estimate cĉ (hats, as here, indicate estimates), the measurand value C is bracketed at better 

than 95% confidence by intervals of the type:

cĉ − UU < CC < cĉ+ UU (3) 

at 95% confidence in the method evaluation. The coverage factor k is intended to account for 

both (1) the fluctuation of the measurement about the measurand value and (2) the 

uncertainty in the assessment of this fluctuation.

Note: Requiring only mean confidence in the evaluation leads to k given in terms of a Studentt quantile. Here, however, in fixing the method evaluation confidence (e.g., at 95%), the chi-
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Measurement Uncertainty and NIOSH Method Accuracy Range

square distribution takes the place of the Student-t distribution. Double confidence levels (in 

the measurement and evaluation) directly relate to a well-developed [Bartley 2001; Bartley and 

Irwin 2002; Hald 1952; Wald 1942, 1943; Wald and Wolfowitz 1946; Wilks 1941, 1942; 

Aitchison and Dunsmore 1975] statistical theory of tolerance or prediction intervals. 

Another point of ISO GUM is semantic. Uncertainty, as in common usage, covers only what 

is unknown about a measurement. The known but uncorrected systematic deviation or bias

relative to reference concentrations does not enter into measurement uncertainty.

A related concept, accuracy, is defined qualitatively within ISO GUM as the “closeness of 

agreement between the result of a measurement and a true value of the measurand”. Accuracy 

can have both random and systematic components. It is not surprising then that if a bias

correction is made and if accuracy is quantified reasonably, the expanded uncertainty and an 

accuracy confidence limit can be equivalent.

As mentioned above, another aspect of ISO GUM deals with influence factors. If measurement 

results are expected to be sensitive to an environmental factor (e.g., ambient temperature), 

then the effect of such a factor on the measurement method must be measured in the 

laboratory. Given estimates of the environmental variations expected during method 

application, influence components of the combined uncertainty can be estimated for inclusion 

in the uncertainty budget of Eq. 1. Table 1 lists several influence factors, which may or may not 

be significant.

3 The symmetric accuracy range A as used by 

NIOSH

a. Definition and its approximation 

The symmetric accuracy range A is defined as the fractional range, symmetric about the 

true concentration C, within which 95% of sampler measurements ĉ are to be found. 

Another way of saying this is:

CC xx (1 − AA) < cĉ< CC xx (1 + AA) for 95% of measurements ĉ (4)

It is clear from this simple definition that the accuracy range function A must increase 

with both random effects and bias magnitude and therefore, is one means of quantifying 

accuracy as defined above according to ISO GUM.

More specifically, suppose that estimates ĉ are normally distributed about population 

mean c with standard deviation σ. Then we may characterize random measurement effects 
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Measurement Uncertainty and NIOSH Method Accuracy Range

in terms of the (true) relative standard deviation TRSD and bias of the mean concentration 

estimate c relative to the true concentration C as:

bbbbbb = cc − CC

CC

 (5) 

 TTTTTTTT = σσ

CC

 . 

The descriptive definition of Eq. 4 implies that the symmetric accuracy range A increases 

with both TRSD and bias magnitude |bias|. This feature can be seen directly in the 

following close approximation to the accuracy range function A, which follows [See 

Bartley 2001; Bartley and Irwin 2002 for derivation] from the definition in Eq. 4:

AA = 1.960 xx √bbbbbbss2 + TTTTTT 2, ii |bbbbbb | < TTTTTTTT

1.645 ; 

AA = |bbbbbb | + 1.645 × TTTTTTTT, ooooheeeeee ee. (6) 

This expression is simple enough for calculation by most hand-held calculators, and it is 

also a useful starting point for estimating the 95% confidence limit A95% on the accuracy 

range as measured during a method evaluation, accounting for evaluation errors.

b. Uses of the symmetric accuracy range 

1) Method validation 

One application of the symmetric accuracy range is for evaluating measurement 

methods. As mentioned in NMAM guidance chapters, a method evaluation consists of 

a number of measurements taken from replicate samplers at each of several controlled 

and known concentrations covering the range of expected method application. This 

type of experiment gives information about the samplers’ random errors and also the 

bias relative to reference concentrations. A confidence limit on the accuracy range can 

then be computed. One objective in a method suitable for NIOSH application is that 

the 95% confidence limit A95% not exceed 25%. A includes both the uncertainty (as the 

term is used by ISO GUM) and the systematic deviation or bias, so that correction of 

the bias by the sampler vendor or developer is encouraged by the very statement of this 

objective. See Eqs. 9-11 below for computing A95% when bias is negligible.

2) Measurement uncertainty 

Suppose then that bias correction has been made. For example, suppose that following 

evaluation, the sampler is used for future measurement with bias corrected on the basis 

of its measurement during the evaluation itself. Then computation of the confidence 
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Measurement Uncertainty and NIOSH Method Accuracy Range

limit A95% is possible accounting for the residual bias which is uncorrectable due to 

evaluation limitations, but nevertheless will be present in all future measurements. The 

quantity ĉ x A95% forms the counterpart to the expanded uncertainty U of ISO GUM for 

specifying evaluation confidence at 95%.

 

The relationship between ĉ x A95% (with corrected bias) and expanded uncertainty U

can be seen most clearly in the case that A95% is significantly smaller than 100%. In this 

case, Eq. 4 can be rewritten as the approximation:

cĉ − cĉ× AA95% < CC < cĉ+ cĉ× AA95%, (7) 

which means that at 95% confidence in the method evaluation, the inequality 

bracketing the measurand value C holds at probability > 95%. A study using 10,000-

point simulations indicates that A95% can be as large as 25%, with method evaluation 

confidence close to 95% using the approximation of Eq. 7. As can be seen directly, Eq. 

7 is the analogue to Eq. 3 when ĉ x A95% is adopted as the expanded uncertainty U: 

UU = cĉ× AA95%. (8) 

In the case that bias is known to equal zero (Technical Note 1 at the end of this 

chapter), A95% (at 15 degrees of freedom in the evaluation experiment) is simply:

AA95% = 2.8 × TTTT�TTTT. (9) 

Eq. 8 then gives: 

UU = 2.8 × TTTT�TTTT × cc�

UU = 2.8 × uucc . (10) 

Therefore, the coverage factor k is

k = 2.8, (11) 

consistent with the use of k = 3 as a conservative but not excessive value.

The user of a method then may report the expanded uncertainty U in a concentration

Estimate ĉ using Eq. 8, knowing the accuracy range confidence limit A95% as reported in 

the method. Of course, this approach relies on the sense of double confidence—in the 

evaluation and also in the subsequent application.
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Measurement Uncertainty and NIOSH Method Accuracy Range

Note: The European Assessment of Workplace Exposures Technical Committee, CEN 

TC 137, has adopted a similar viewpoint regarding method performance [CEN 2015]. 

In this case, an overall uncertainty, defined as |bias| + 2 × RSD, is used to quantify 

accuracy. When compared to Eq. 6, the overall uncertainty can be regarded as an 

approximation to the symmetric accuracy range.

4 Uncertainty and analytical lab procedures

Interest in measurement uncertainty and ISO GUM is currently finding its way into the 

criteria for the accreditation of analytical labs [ISO 2005]. The result will no doubt be high 

confidence in understanding one component of the combined or expanded uncertainty—

namely the analytical component. Several general approaches to controlling and 

characterizing analytical uncertainty in routine lab practices seem reasonable.

a. Validated method adoption 

One possibility is for a lab to adopt a published, evaluated method. Such an adoption 

would require an initial establishment of the method within the lab’s capabilities. 

Equivalence to the published method would be established during this initial phase. 

Thereafter, the method’s uncertainty as documented in the original publication would be 

claimed for the lab results. Ongoing analysis of a limited number of quality control 

samples would provide evidence that the method as implemented in the lab remains 

stable.

An example of this approach is the current practice in some labs that handle sorbent tubes 

to analyze about 4 lab blanks per set of field samples analyzed. The variability in the blank 

results are then continually compared to past lab performance so as to detect problems 

which may occur in analysis. Though the small number of degrees of freedom (= 3) does 

not give a tight figure on the uncertainty, it nevertheless gives assurance that the method is 

stable.

As a specific example of method evaluation data and documentation of an uncertainty 

budget, data from n = 16 exposures of diffusive samplers in a controlled environment are 

shown in Table 2. The evaluation is somewhat simplified for this example; a more 

comprehensive evaluation would also measure effects of wind velocity, humidity, 

temperature, and concentration time-dependence (potentially significant to diffusive 

monitoring). Analysis of these data can be handled by an ordinary calculator capable of 

computing means and standard deviations.
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Measurement Uncertainty and NIOSH Method Accuracy Range

Note that the uncertainty

the bias is the value that accounts for residual bias due to imperfect correction. Very 

similarly, the uncertainty in the reference concentration is pooled to arrive at a combined 

uncertainty. Interestingly, neither of these two contributions corresponds to quantities 

that vary during sampler application subsequent to its initial evaluation. The background 

for documenting residual (uncorrectable) bias can be seen in Technical Note 2 at the end 

of this chapter.

(TTTT�TTTT/√nn, where n = 16 is the number of measurements) in 

Table 2. Example of method evaluation and uncertainty budget

Evaluation of experimental results

The following are results from a simplified evaluation of a specific diffusive sampler for oxylene. There were four experimental runs with four samplers each. The reference 

concentration set within the exposure chamber is denoted as Ĉ having an assigned (Type 

B) relative uncertainty = 1%.

Run Ĉ (ppm) replicates(ppm) ⇒ TTRR�SSDDiiiiiiiiii�ss/CC� � biâs

1 123 

139.2 

2.6% 14.0% 

138.2 

138.6 

145 

2 101.1 

108.3 

1.8% 9.2% 

110 

110.7 

112.8 

3 12.7 

14.2 

7.9% 12.2% 

15.3 

12.9 

14.6 

4 91.3 

109 

2.1% 17.8% 

109.2 

107.1 

105 

Averaging the above bias estimates and pooling the inter-sampler estimates TTTT�TTTT2

iiiiiiiiii

resultsin: Mean bias estimate: biâs = 13.3% from average of 4 × 4 = 16 data points. The

TRSD estimate is TTTT�TTTTiiiiiiiiii = 4.4% having 4 × 3 = 12 degrees of freedom.
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Measurement Uncertainty and NIOSH Method Accuracy Range

Table 2. Continued

Uncertainty budget

The following includes bias correction by dividing future concentration estimates by (1 + 

biâs) as in Technical Note 2. 

Source Component Category

Inter-sampler 3.9% [= 4.4% / (1+.133)] Type A 

BiasCorrection Uncertainty 0.97% [= 3.9%/√16] Type A 

RefConcentration Uncertainty 0.5% [1%/√4, but not /(1 + biâs)] Type B 

Combined (Relative) Uncertainty uucc = √3.9%2 + 0.97%2 + 0.5%2 = 4.0% 

Expanded (Relative) Uncertainty UU = kk × uucc = 12.1% (kk = 3). 

Notes: Here k = 3.0. A more accurate determination based on the chi-square quantile at

12 degrees of freedom and prob = 0.05, gives k = 2.97, which is consistent with

conventional use of 3 as a conservative value.

Again, an expanded (relative) uncertainty U means that with greater than 95% of future 

bias- corrected estimates ĉ, true concentrations C are bracketed by:

 cĉxx (1 − UU) < CC < cĉxx (1 + UU), 

at 95% confidence in the above evaluation experiment. Generally, a quality control 

program is required to ensure that the method remains stable following evaluation.

Note that many methods (e.g., those based on sorbent tubes) employ personal sampling 

pumps, in which case normally a 5% (Type B) component representing sampling pump 

uncertainty would be included in the uncertainty budget.

Note also that the inter-sampler component includes both analytical and sampling subcomponents. Further refinement of the inter-sampler component may perhaps be useful 

for improving a method, but is not needed for establishing confidence intervals around 

(true) measurand values.

Note further that storage effects require estimating and inclusion in the budget if 

considered significant.

b. Pooled quality control results 

Another approach utilizes a large number (e.g., 50) of the most recent quality control 

sample results. By pooling uncertainty values, a running method evaluation can be 
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Measurement Uncertainty and NIOSH Method Accuracy Range

effected. The result is (1) a direct measure of the analytical uncertainty of the method as 

implemented in the lab, and (2) a means of detecting any problems that may creep into a 

method during routine use. Note that a running uncertainty average is similar to a partial 

method evaluation and not to a method re-evaluation at each measurement because 

consecutive running averages are strongly correlated.

This approach is adopted within a current MSHA procedure for the analysis of silica. A 

sampling filter is dissolved and re-deposited onto an analysis filter where the silica is 

quantified by infrared absorption. From each batch of samples to be analyzed, an analysis 

filter is retained for re-dissolution, re-deposition, and re-analysis within a subsequent 

batch. The result is a large number of pairs of nearly identical samples, which can give a 

running estimate of the method’s analytical uncertainty.

c. Continual method re-evaluation 

A third possibility, the closest to the original measurement approach of GUM uses a large 

number (e.g., 30) of independent control samples for each application measurement. This 

is the most expensive approach, but also may give the best estimate of the analytical 

uncertainty, especially in cases where uncertainties may be measurement-dependent. 

Because many more evaluative measurements per application measurement are needed, 

this approach is not easily implemented for most industrial hygiene applications.

As an example of this approach [ISO 2002], suppose that a lab estimates only a 30-day 

average concentration of a given gas or vapor. Further, every day a measurement is taken 

of a known calibration gas concentration. Then, if the method is expected to behave 

similarly for measurements of gas and field samples calibrations, the 30 control samples 

give analytical uncertainty estimates that differ month-to-month and from field 

measurement to field measurement.

When the concentration is low, approaching the method uncertainty uc, concepts of the 

limit of detection (LOD) and a related detection limit LD may be useful. LOD is used for 

controlling false positives when asserting the presence of a substance. On the other hand, 

the detection limit specifies what measurand value (e.g., concentration) is required so that 

the false negative rate is negligible when the substance is actually present. The limits can 

refer to the analytical measurement only or, as in this section, to the entire sampling and 

analytical measurement method.

In the following examples, several often realistic assumptions are made. The standard 

deviation in concentration estimates is assumed constant (i.e., independent of the sampled 

d. Limit of detection and detection limit
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Measurement Uncertainty and NIOSH Method Accuracy Range

concentration), unlike the commonly occurring constant relative standard deviation at 

larger concentrations. Also, bias (or uncertainty in its correction) is assumed to be 

negligible. More complicated cases generally have specific difficulties that are best be 

approached by a statistician.

With these assumptions, in terms of the combined uncertainty uc the limit of detection is 

traditionally [Keith et al. 1983; Currie 1997] taken to be:

 LLLL = 3 × uucc , (12) 

and the detection limit may be defined as

 LL = 2 xx LLLL . (13) 

Note that uc includes the uncertainty associated with correction, if any, with blank 

subtractions. See ISO [2009] for a detailed example.

After LOD and LD have been determined for a method they may be used as follows. A 

substance may be asserted as present if an estimate xx� exceeds LOD. Moreover, if unknown 

(true) concentration X exceeds LD, an estimate xx� is likely to exceed LOD. Given the above 

definitions and assumptions, the false positive rate r on asserting presence is closely equal 

to the non-detection rate.

Note: If the combined uncertainty uc is determined from a method evaluation providing 

an effective number υυ (as in Technical Note 2) degrees of freedom, then at 95% confidence 

in the method evaluation, the false positive rate r is limited by:

< 1 − φφ �3 × �χχ0.05,υυ 2

υυ �, (14) 

where φφ is the cumulative normal function. For example, if υ=15, then r < 3.5%.

5 Discussion

The approach presented here to document method accuracy range and uncertainty relates to 

the statistical theory [Bartley 2001; Wald and Wolfowitz 1946; Wilks 1941, 1942; Aitchison 

and Dunsmore 1975; CEN 2015; Smith 1936; Saterthwaite 1946] of tolerance or prediction 

intervals. This theory was originally developed in simplified form to predict the range of 

future measurements of a normally distributed random variable on the basis of n initial 
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Measurement Uncertainty and NIOSH Method Accuracy Range

measurements. The initial measurements are analogous to the method evaluation, whereas the 

future measurements represent method application subsequent to evaluation.

Because of measurement cost, workplace assessments cannot at present be conducted in such 

a way that continual re-evaluation is done at each measurement. The prediction interval 

approach given here shows a less costly way to document measurement uncertainty in those 

cases where a method has been initially evaluated and then used many times without reevaluation. The approach closely follows ISO GUM. Of course, for such an approach to 

actually make sense, an adequate quality control program must be instituted so that the 

measurement method remains stable during the time of its application following evaluation.

Several generalizations and variations of the material presented in this chapter are possible. 

The relative standard deviation and relative bias sometimes depend on the concentration 

sampled in a complicated way, requiring special attention. See, for example Currie [1997]. 

Also, asymmetric confidence intervals are sometimes required. Single-sided intervals are 

useful in some instances, e.g., alarm systems, as well as in quantifying limits of detection or 

quantitation, described briefly above. Ways to handle environmental influence parameters 

may also be complicated. See, for example ASTM [2013a]. In any case, despite the 

complexities possible, the examples given in this chapter may help to characterize method 

uncertainty in a reasonable manner. For additional examples and explanations, see 

Appendices A and D of Components for Evaluation of Direct-Reading Monitors for Gases 

and Vapors [NIOSH 2012] and ASTM standards on accuracy and uncertainty [ASTM 2014, 

2015]. 

6 Technical notes

a. Note 1: Example of accuracy range confidence limit:

Suppose it is known that the bias is zero. For example, an exposure standard may be set 

that specifies a given sampling and analytical method. In this case, the hazardous 

concentration may be said to be operationally defined. Operationally defined methods 

include NIOSH Methods 7400 and NIOSH 5040.

If the bias is zero, Eq. 6 simplifies to: 

 AA = 1.960 xx TTTTTTTT. 

Furthermore, if the relative standard deviation is estimated as TȒSD with υ degrees of 

freedom (computed using the Smith-Satterthwaite approximation [ISO 2010; Hald 1952; 
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Measurement Uncertainty and NIOSH Method Accuracy Range

Wald 1942] if TRSD has more than one component), then the 95% confidence limit on 

TRSD is: 

TTTTTTTT95% = TTTTTT̂TT × υυ

�χχ2 , υυ,0.05

where χχ2

υυ,0.05 is a 5% quantile value for the chi-square distribution, which can be read from 

a table in most elementary statistics texts. This determines the 95% confidence limit on the 

accuracy range itself as:

 AA95% = AÂ × υυ

�χχ2 . υυ,0.05

If υ = 15, υυ

�χχ2 giving coverage factor k = 2.8 (Eq. 11). υυ,0.05

Note that TRSD95% can be interpreted as a conservative estimate of TRSD and therefore can 

be treated as a Type B uncertainty with infinite number of degrees of freedom as described 

following Table 1.

b. Note 2: Single-evaluation correction of bias: 

Details are given here illustrating the tolerance interval approach, bias correction, 

imprecise reference concentrations, and the use of the symmetric accuracy range function. 

The derivation is not entirely general, but is given here for guidance in handling the 

myriad possibilities in measurement uncertainty. Though the derivation is slightly 

complicated, the result obtained is simple.

Suppose that estimates ĉ having an as-yet-unknown constant bias relative to true 

concentrations C (not necessarily constant) may be modeled as:

 cc = CC ∙ [1 + bbbbbbss + εε] , 

where the random variable εε is approximately normally distributed about zero with 

variance TRSD2

. For evaluating the method, assume that reference concentration 

measurements Ĉ can be made simultaneously and modeled by:

 CĈ = CC ∙ �1 + εεiiiirr�,

̂ ̂

̂

̂
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-16 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range

where εε 2

iiiirr has variance TTTTTTTTiiiirr assumed known accurately. Measure n values of the 

ratio θθ�: 

 θθ� ≡ cc = 1 + bbbbbbss + εεcc cc

and compute estimates biâs and TȒSTT2

cc at υ= n−1 degrees of freedom, where the 

approximately normally distributed random variable εεcc has variance TȒSTT2

cc given by:

 TTTTTTTT2

cc = TTTTTTTT2 + (1 + bbbbbb�ss)2xx TTTTTTTT2

iiiirr

(to the order of TRSD, neglecting Cauchy effects of reciprocals of random variables).

Future bias-corrected measurements xx�′

of unknown concentration Χ can be defined in

terms of raw measured values xx� as:

 xx

�′ ≡ xx�

1+bbiiaa�ss

 . 

The residual corrected bias' is then given by:

 bbbbbbss′ = bbiiaass−bbiiaa�ss

1+bbiiaa�ss

If n is large enough, |bbbbbbss′| will be small enough that the corrected symmetric accuracy 

range A' can be accurately approximated (Eq. 6) as:

 AA′2 = aa21.9602

(1+bbiiaa�ss)2 , 

where the unknown ɑ2 is:

 bb2 = (bbbbbbss − bbbbbb�ss)2 + TTTTTTTT2, 

whose confidence limit is now required.

̂

̂

̂ ̂

̂
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Measurement Uncertainty and NIOSH Method Accuracy Range

First note that the expected value of the first term is:

 EE[(bbbbbb − bbbb �ss)2] = TTTTTT cc

2

 . 

Therefore, an estimate �2 for ɑ2 can be constructed as:

 �2 ≡

TT � cc

2

+ TT � 2

 TT � 2 ≡ TT � cc

2 − (1 + bbbb �ss)2 xx TTTTTT rr

2 . 

Expressed in terms of TT � 2, �2 is: 

 

 �2 = TT � 2

 + 1 TT � 2

 + 1 ∙ (1 + bbbb �ss)2 xx TTTTTT rr

2 . 

Each term can now be identified, forming the basis for an uncertainty budget: the first is 

the (uncorrected) method uncertainty (squared); the second and third reflect the biascorrection uncertainty owing to finiteness of the validation experiment and the 

uncertainty in the reference concentration (as here measured n times).

A confidence limit ββ

2 at confidence level β (e.g., 95%) on ɑ2 is now constructed using ɑ�2: 

 ββ

2 = KK2 ∙ �2, 

where the constant K is to be determined so that

 pp � ββ

2 < 2� = 1 − ββ. 

First of all, the distribution of ɑ�2 is approximated as chi-square:

 υυ aa�2

EE[aa�2] ≈ χχ2 , 

where υυ is determined as with the Smith-Satterthwaite [Keith et al. 1983; Currie 1997] 

approximation, forcing variances to agree; often υυ ≈ nn − 1. 

Now,

 ββ

2 < 2⇔ �2 < KK−2 2, 
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-18 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range

or, in other words:

 χχ2 < υυeeeeee�KK−2aa2�

EE[aa2] . 

[Note that, EE[ �2] = EE[ 2], bb ≠ 2unlike Technical Note 3.] 

Remembering that ɑ2 depends on the estimate biɑ�s, K is given as a solution of the 

following integral equation: 

1 − ββ = � dd �ss PP⌈bbbb �ss⌉ � ddχχ2PPυυeeeeee[χχ2]

υυeeeeee∙KK−2∙ɑ−2 EE�ɑ2 � �

0

+∞

−∞

 = ∫ ddχχ2PPυυeeeeee[χχ2] × (1 + ΟΟ[1 nn2 ⁄ ]) υυeeeeee∙KK−2

0 , 

where the correction O[1/n2

] is easily proved by expanding the integrand about TTTTTTTTcc

2

in(bbbb �ss − bbbbbb )2. Therefore, the following simple asymptotic expression for K2 results:

 KK2 ≈ υυeeeeee

χχ1−ββ,υυeeeeee

2 . 

Thus, the coverage factor k is approximated as:

 kk ≈ 1.960 ∙ � υυeeeeee

χχ1−ββ,υυeeeeee

2 , 

which is less than and close to 3.0, if the effective number of degrees of freedom υυ ≥12, 

and β=0.95. 

In summary, the bias uncertainty is pooled together with the uncertainty components in 

TȒSD2

. It should be remembered, however, that only TȒSD refers to quantities, which vary 

at each of the future measurements following the initial evaluation. 

c. Note 3: Characterizing effects of uncorrected bias: 

If the systematic error (bias) is non-zero, confidence limits on the accuracy range A may 

be approximated as follows. The Smith-Satterthwaite approximation is generalized in 

approximating estimates AÂ in terms of a chi-square random variable χχυυ

2 for the two cases 

in Eq 6 by:

 AA�

AA = �χχυυ

2

υυ , ii|bbbbbb | <

1.645 ;

 AA�

AA = χχυυ

2

υυ , ooooheeeeee ee
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-19 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range

The effective number of degrees of freedom υ is determined by forcing the variance of 

to reproduce the estimated variance of or in their respective cases:

4

 υυ = 2AA

�2 , bbii |bbbbbbss| < TTTTTT

; vvaaii[AA ] 1.645

2

 υυ = 2AA oooo vv [AA� , heeeeeebbssee aaii ]

Calculation of vvbbee �AÂ2� ooee vvbbee �AÂ� is generally straightforward and depends on specifics 

of the evaluation experiment and on significant influence parameters. The confidence 

limit A95% is then determined as in Eq 9:

 AA95% = AÂ × υυ

� 2 , bbii |bbbbbbss| < TTTTTT

χχ0.05,υυ 1.645; 

 AA95% = AÂ × υυ

2 , ooooheeeeeebbssee

χχ0.05,υυ

This expression has been found [Bartley and Irwin 2002] quite accurate, exhibiting 

negligible effects from the discontinuity: The chi-square approximation is expected to be 

worst when |bbbbbbss|is large relative to TRSD. As an example, suppose the uncertainty has 

the following components: 5% from pump error and also a 5% analytical relative standard 

deviation. Suppose bias = 20%. Suppose bias and the analytical uncertainty are measured 

with υ = 15 degrees of freedom. Then 10,000-point simulations indicate that the calculated 

A95% is slightly conservative, giving 96% confidence.

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-20 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range 

7 References

ACGIH [2015]. Appendix C: Particle size-selective sampling criteria for airborne particulate 

matter. In: 2015 TLVs® and BEIs®. Cincinnati, OH: American Conference of Governmental 

Industrial Hygienists.

Aitchison J, Dunsmore IR [1975]. Statistical prediction analysis, Cambridge, England: 

Cambridge University Press.

ASTM [2013a]. D6246-08(2013) Standard practice for evaluating the performance of diffusive 

samplers. West Conshohocken, Pennsylvania: American Society for Testing and Materials 

International (ASTM).

ASTM [2013b]. D6062-07(2012) Standard guide for personal samplers of health-related 

aerosol fractions. West Conshohocken, Pennsylvania, American Society for Testing and 

Materials International.

ASTM [2014]. D3670 Standard guide for determination of precision and bias of methods. 

West Conshohocken, Pennsylvania, American Society for Testing and Materials International.

ASTM [2015]. D7440 Standard practice for characterizing uncertainty in air quality 

measurements. West Conshohocken, Pennsylvania, American Society for Testing and 

Materials International.

Bartley DL [2001]. Definition and assessment of sampling and analytical accuracy.

Ann Occup Hyg 45(5):357-364. 

Bartley DL and Irwin KC [2001]. Accuracy in measurements using diffusive samplers. In: 

Proceedings of the International Conference on Diffusive Samplers, Montpellier, France, pp. 

99-107. 

Bartley DL [2004]. Reconciling traditional accuracy assessment with the ISO guide to the 

expression of uncertainty in measurement (ISO/GUM). J Occup Environ Hyg 1:D37-D41.

Busch KA [1977]. Appendix A: SCP Statistical Protocol. In: Taylor DG, Kupel RE, Bryant JM, 

eds. Documentation of the NIOSH validation tests. Cincinnati, OH: U.S. Department of 

Health, Education, and Welfare, Center for Disease Control, National Institute for 

Occupational Safety and Health, DHEW (NIOSH) Publication. No. 77-185, pp. 7-11. 
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-21 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range

CEN [1993]. CEN EN481:1993 Workplace atmospheres – size fraction definitions for 

measurement of airborne particles. Brussels, Belgium: European Standardization Committee.

CEN [2015] EN 482:2012+A1:2015 Workplace exposures - General requirements for the 

performance of procedures for the measurement of chemical agents. Brussels, Belgium: 

Comité Européen de Normalization.

Currie LA [1997]. Detection: international update, and some emerging dilemmas involving 

calibration, the blank, and multiple detection decisions. Chemom Intell Lab Syst 37(1):151-

181. 

Ellison SLR, Williams A eds. [2012]. Eurachem/CITAC guide: Quantifying Uncertainty in 

Analytical Measurement, 3rd edition, Eurachem.

Hald A [1952]. A statistical theory with engineering applications. New York: John Wiley & 

Sons.

ISO [1995]. Air quality–particle size fraction definitions for health-related sampling. ISO 

7708:1995(en). Geneva, Switzerland: International Organization Standardization.

ISO [2002]. ISO 11222:2002 Air quality - determination of the uncertainty of the time average 

of air quality measurements. Geneva, Switzerland: International Organization for 

Standardization.

ISO [2005]. ISO/IEC 17025:2005 General requirements for the competence of testing and 

calibration laboratories. Geneva, Switzerland: International Organization for Standardization.

ISO [2009]. ISO 15767:2009 Workplace atmospheres - controlling and characterizing 

uncertainty in weighing collected aerosols. Geneva, Switzerland: International Organization 

for Standardization.

ISO [2010]. Evaluation of measurement data - guide to the expression of uncertainty in 

measurement. Geneva, Switzerland: International Organization for Standardization (ISO).

Keith LH, Crummett W, Deegan J Jr., Libby RA, Taylor JK, Wentler G [1983]. Principles of 

environmental analysis. Anal Chem 55(14):2210-2218. 
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-22 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range

NIOSH [2012]. Components for Evaluation of Direct-Reading Monitors for Gases and 

Vapors. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 2012-162. 

 

NIOSH [1995]. Guidelines for air sampling and analytical method development and 

evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM and Shulman SA. Cincinnati, 

Ohio: U.S. Department of Health and Human Services, Centers for Disease Control and 

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) 

Publication No. 95-117. 

Satterthwaite FE [1946]. An approximate distribution of estimates of variance components. 

Biometrics Bulletin 2(6):110-114. 

Smith HF [1936]: The problem of comparing the results of two experiments with unequal 

errors. Journal of the Council of Scientific and Industrial Research 9:211-212. 

Vaughn NP, Chalmers CP, Botham RA. [1990]. Field comparison of personal samplers for 

inhalable dust. Ann Occup Hyg, 34(6):553-573. 

Wald A [1942]. Setting tolerance limits when the sample is large, Ann Math Statist 13(4):389-

399. 

Wald A [1943]. An extension of Wilk’s method for setting tolerance limits. Ann Math Statist 

14(1):45-55. 

Wald A, Wolfowitz J [1946]. Tolerance limits for a normal distribution. Ann Math Statist 

17(2):208-215. 

Wilks SS [1941]. Determination of sample sizes for setting tolerance intervals. Ann Math 

Statist 12(1):91-96. 

Wilks SS [1942]. Statistical prediction with special reference to the problem of tolerance 

limits. Ann Math Statist 13(4):400-409. 

8 Terminology

A — symmetric accuracy range, relative (%) range of 95% of a method’s measurements about 

the (true) measurand

A95% — 95% confidence limit on the symmetric accuracy range
NIOSH Manual of Analytical Methods  5th Edition  Chapter UA April 2016 Page UA-23 of UA-23

Measurement Uncertainty and NIOSH Method Accuracy Range

bias — mean concentration estimate bias relative to the (true) measurand

biâs — bias estimate

ĉ — concentration estimate

C— true concentration

Ĉ — reference concentration (estimate)

k — coverage factor, a constant containing confidence information for obtaining the 

expanded uncertainty U as a factor of the combined uncertainty uc 

LD— detection limit (for controlling false negatives)

LOD — limit of detection (for controlling false positives)

n — number of measurements in a method evaluation

ssjj

2 — unbiased estimate of variance σσjj2

σσjj

2 — jth population variance component

TRSD— (true) relative standard deviation

uj — jth uncertainty component, an estimated standard deviation

uc— combined uncertainty, pooled uncertainty components

U— expanded uncertainty, a value giving intervals bracketing the (true) measurand at given 

confidence in the measurement and method evaluation

Ʋj— degrees of freedom in an estimate

χχυυ,0.05

2 — chi-square quantile. This quantity by definition exceeds the chi-square

 variable at probability = 5%. Note that many tables use the notation χχυυ,0.05

2 

 for this quantity.

In general, hats represent estimates. Primes indicate bias-corrected quantities.
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

General Considerations for

Sampling Airborne Contaminants

by Charles S. McCammon, Ph.D., CIH and Mary Lynn Woebkenberg, Ph.D.; recension by Kevin 

Ashley, Ph.D., NIOSH

1 Choosing measurement methods and sampling media SA-2

2 Types and uses of solid sorbents SA-3

3 Types and uses of aerosol samplers SA-5

4 Factors affecting the collection of gases, vapors, and aerosols SA-5

5 Establishing sampling parameters SA-7

6 Bulk samples SA-10

7 Blanks SA-11

8 Direct-reading methods SA-12

9 Sampling Strategy SA-13

10 Sampling and calibration techniques SA-13

11 References SA-20
NIOSH Manual of Analytical Methods  5th Edition  Chapter SA April 2016 Page SA-2 of SA-23

General Considerations for Sampling Airborne Contaminants 

1 Choosing measurement methods and sampling

media

In choosing methods for sampling of chemical and biological agents related to occupational 

exposures, thorough advance planning is required. Proper planning minimizes sampling and 

measurement costs while enabling the collection of high-quality data. Many criteria must be 

considered before collecting field samples from workplaces [ASTM International 2014]. The 

first step is to define the sampling objectives. These may include: a) documenting exposures in 

particular work settings; b) assessing compliance/non-compliance with existing regulatory or 

recommended occupational exposure limits (OELs); and/or c) determining sources of 

airborne contaminants. Sampling parameters that should be considered might include: a) type 

of sample (area vs. personal); b) contaminant(s) to be sampled; c) physical nature of airborne 

sample (vapor and/or aerosol); d) duration of sample collection; e) potential interferences; 

and f) estimated contaminant concentrations. Once these parameters are established, the 

suitable analytical method(s) and sampling media can be selected. For instance, when 

sampling for aerosols, the relevant particle size fraction to be sampled must be taken into 

account. Other general information needed to properly plan a sampling campaign include: a) 

the number of employees; b) the sampling strategy plan (discussed later); c) process flow 

diagram; d) safety data sheets on all process materials; and e) potential hazards involved in 

collecting and shipping field samples.

An accredited analytical laboratory should be used to conduct analysis of collected samples, 

and it is essential to consult with the analytical laboratory before sampling to ensure that the 

measurement methods available can meet the defined measurement needs. Consultation with 

the laboratory should be an early part of survey planning. The laboratory can also assist in 

choosing sampling media that are compatible with the sampling needs and the measurement 

methods available. The APPLICABILITY sections of the individual methods in NMAM can 

be helpful in choosing which of the available methods is best for a particular situation. Apart 

from NIOSH methods, methods from other organizations such as OSHA, ASTM 

International and ISO may be appropriate [Ashley 2015]. 

Whether through consultation with the laboratory or as cited in the relevant measurement 

method, the sampling media will be specifically identified. Methods will specify parameters 

such as: a) pore size and type of filter (for aerosol collection); b) concentration and amount of 

liquid media required (e.g., for impinger sampling or impregnation of filters); and c) type and 

amount of solid sorbent (for collection of gases / vapors); see below for common types and 

characteristics of various sampling media. 
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General Considerations for Sampling Airborne Contaminants 

The physical state(s) of the contaminant(s) being sampled may also be a factor in determining 

the media required. In the case of polyaromatic hydrocarbons (PAHs), for example, the 

correct sampler consists of a membrane filter to collect particulate matter and a solid sorbent 

tube to trap the vapors of certain PAHs so that total collection is assured. Also, for sampling 

of gases and vapors, it is generally necessary to mount a prefilter in front of the gas/vapor 

collection media in order to trap aerosols that might otherwise interfere with subsequent 

analysis of target analytes.

The sampling pump used to collect the sample must also be compatible with the sampling 

needs and the media used. Specifically, the pump must be capable of maintaining the desired 

flow rate over the time period needed using the sampling media specified. Some pumps may 

not be able to handle the large pressure drop of the media. This may be the case for fine mesh 

solid sorbent tubes, small pore size filters or when attempting to take a short-term sample on a 

sorbent tube of a higher than normal pressure drop at higher flow rates. As a rule of thumb, 

higher-flow pumps (>1 L/min) can handle at least 3 kPa pressure drop at 1 L/min for 8 h. 

Some pumps can handle up to 7.5 kPa pressure drop at flows up to 5 L/min. Most low flow 

pumps (0.01 to 0.2 L/min) can handle the pressure drops of available sorbent tubes without 

problems, except that the nominal flow rate may decrease for certain models. All pumps 

should be calibrated with representative sampling media prior to use. It is good practice to 

check the pump calibration before and after use each day. As a minimum, calibration should 

be done before and after each use.

2 Types and uses of solid sorbents* [Melcher 1987]

a. Activated charcoal

By far the most commonly used solid sorbent is activated charcoal. This sampling medium 

is characterized by a very large surface area to weight ratio. It has a reactive surface and 

high adsorptive capacity. This surface reactivity means that activated charcoal is not useful 

for sampling reactive compounds (e.g., mercaptans, aldehydes) because of poor desorption 

efficiency. The high capacity, however, makes it the sorbent of choice for those 

compounds that are stable enough to be collected and recovered in high yield. 

Breakthrough capacity is a function of type (source) of the charcoal, its particle size and 

packing configuration in the sorbent bed. Humidity may affect the adsorption 

characteristics as well.

*NOTE: Solid sorbents are used for the collection of vapors only. Aerosols are 

notcollected effectively by most sorbent beds, but may be collected by other components 

of the sampler (e.g., a prefilter).
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General Considerations for Sampling Airborne Contaminants 

b. Silica gel

Silica gel is less reactive than charcoal. In part because of its polar nature, coated silica gel 

has been shown to be effective for sampling airborne ketones and aldehydes [Tejada 1986; 

García-Alonzo and Pérez-Pastor 1998].

c. Porous polymers

Porous polymers feature lower surface area and much less reactive surface than charcoal. 

Adsorptive capacity is, therefore, generally lower, but reactivity is much lower as well.

d. Synthetic sorbents

Synthetic carbonaceous sorbents demonstrate properties midway between charcoal and 

porous polymers.

e. Coated sorbents

Coated sorbents are those upon which a layer of a reagent has been deposited. The 

adsorptive capacity of such systems usually approaches the capacity of the reagent to react 

with the particular analyte of interest [Kennedy 1988].

f. Molecular sieves

Zeolites and carbon molecular sieves retain adsorbed species according to molecular size. 

A limiting factor is that the water molecule is of similar size to many small organic 

compounds and is usually many orders of magnitude higher in concentration than the 

species of interest. This unfavorable situation may result in the displacement of the analyte 

by water molecules. Drying tubes may be used during sampling to eliminate the effects of 

humidity [Langhorst 1983].

g. Thermal desorption

Thermal desorption tubes may contain several different sorbents in order to collect a wide 

range of different chemical agents [Hodgson et al. 1988]. These tubes are generally used in 

situations where unknown chemicals or a wide variety of organics are present, e.g., in 

indoor environmental air quality investigations. Subsequent analysis is often by gas 

chromatography/mass spectrometry (GC/MS).
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General Considerations for Sampling Airborne Contaminants 

3 Types and uses of aerosol samplers

[Baron 1998]

a. Membrane filters

By far the most frequently used as sampling media for aerosols are membrane filters. This 

class of filters includes those made from polyvinyl chloride, polytetrafluoroethylene 

(PTFE), copolymers, and mixed cellulose esters (MCE). Filters from this class are used for 

sampling airborne particles such as asbestos, crystalline silica, particulate PAH’s, particles 

not otherwise regulated, and elements for atomic spectrometric analysis.

b. Glass and quartz fiber filters

Quartz filters have replaced glass fiber in many applications. They are used in applications 

such as sampling for mercaptans and diesel exhaust. Impregnated quartz filters are often 

used to sample reactive aerosols.

c. Polycarbonate straight pore filters

Because of their characteristics, polycarbonate filters are good for the collection of 

particles to be analyzed by electron microscopy and x-ray fluorescence.

d. Respirable dust samplers

Cyclone samplers are used to collect airborne respirable dust. For instance, a high-flow 

cyclone attached to a sampler containing a polyvinyl chloride filter is used to collect 

respirable crystalline silica.

e. Inhalable dust samplers

Various samplers have been designed and/or evaluated for collecting samples of inhalable 

airborne particles. For example, the Institute of Occupational Medicine’s (IOM) sampler 

is used, in conjunction with a polyvinyl chloride filter, for sampling formaldehyde on dust 

[NIOSH 1994a].

4 Factors affecting the collection of gases, vapors, 

and aerosols [Hebisch et al. 2009; Jones 1994; Baron 1994; Kulkarni et al.

2011]

a. Temperature

Since adsorption is typically exothermic, adsorption is generally reduced at higher 

temperatures. Additionally, if there is a reaction between an adsorbed species and the 

surface, or between two or more adsorbed species (e.g., hydrolysis or polymerization), the 
NIOSH Manual of Analytical Methods  5th Edition  Chapter SA April 2016 Page SA-6 of SA-23

General Considerations for Sampling Airborne Contaminants 

rate of such reactions increases at higher temperatures. The temperature stability of a filter 

must be considered when sampling in hot environments.

b. Humidity**

Water vapor is adsorbed by polar sorbents; their breakthrough capacity for the analyte is 

thereby reduced for most organic compounds. However, for water-soluble compounds, 

the breakthrough capacity is increased, e.g., for chlorine and bromine [Cassinelli 1991] 

and formaldehyde [NIOSH 1994b]. This effect varies from substantial for more polar 

sorbents, such as charcoal and silica gel, to a smaller effect for synthetic carbonaceous and 

porous polymers. 

Filter media may also be affected by humidity; moisture may affect a filter’s collection 

efficiency. Very low humidities (≤10% RH) may make some filters (e.g., MCE) develop 

high charge levels, causing non-uniform deposits and repulsion of particles [Chen and 

Baron 1996]. Water absorption by some filters (e.g., MCE) can cause difficulty in 

obtaining tare weights for gravimetric analysis; thus weight-stable filter materials must be 

used for this application.

c. Sampling flow rate**/face velocity

Breakthrough volume of a solid sorbent bed tends to be smaller at higher sampling flow 

rates, particularly for coated solid sorbents. For sorbents such as charcoal, whose 

breakthrough capacity for most organic compounds can be significantly reduced by high 

humidity, lower sampling flow rates may actually result in smaller breakthrough volumes 

[Foerst 1979]. The collection efficiency of membrane filters will change with face velocity.

d. Concentration**

As the concentration of contaminant in air increases, breakthrough capacity (mg 

adsorbed) of a solid sorbent bed increases, but breakthrough volume (L of air sampled) 

decreases [Foerst 1979]. The effect of concentration is similar for filters.

**NOTE: It is important to distinguish between equilibrium (saturation) adsorptive 

capacity and kinetic (breakthrough) adsorptive capacity of the solid sorbent. Breakthrough 

capacity is the important characteristic in actual sampling situations; it may be affected

significantly by sampling flow rate and relative humidity of the air being sampled and may 

be significantly less than saturation capacity, which is not dependent on sampling flow rate 

or relative humidity.
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General Considerations for Sampling Airborne Contaminants 

e. Particle characteristics 

Filter collection efficiency is a function of effective pore size [Lee and Ramamurthi 1993]. 

Particles smaller than about 0.2 μm are collected primarily by diffusion, while particles 

larger than about 0.2 μm are collected primarily by impaction and interception. Most 

sampling filters are highly efficient (>95%) for all particle sizes, with the minimum 

efficiency in the 0.2 μm size range. Polycarbonate straight pore filters exhibit poor 

collection by diffusion, so particles smaller than the pores are not collected efficiently.

f. Filter considerations

The pressure drop of a filter can limit the sampling time because of the load on the 

personal sampling pump [Breuer 2012]. In addition, pressure drop increases with dust 

loading on the filter [Lippman 1995]. Fine particles (<0.5 μm) will increase the pressure 

drop much faster than coarse particles (>10 μm). Heavy loading (> about 2 mg) may result 

in poor adhesion of collected particles to the filter surface. 

5 Establishing sampling parameters

Once the sampling media and measurement method are chosen, the specific sampling 

parameters need to be determined [Eller 1986; ASTM International 2013]. For most methods, 

this will not pose a problem as the flow rate recommended in the method can be used for the 

desired sampling period, e.g., 1 to 5 L/min for 8 h for most aerosols or 10 to 200 mL/min for 

8 h for most sorbent tube samples. It is necessary to consider the applicable OEL for shortterm (e.g., 15 min) or long-term (e.g., 8 h) time-weighted average (TWA) sampling. 

Generally, the parameters which must be considered are flow rate, total sample volume, 

sampling time (tied into the two previous parameters), and limit of quantitation (LOQ). Some 

of these variables will be fixed by sampling needs, e.g., sampling time or by the measurement 

method of choice (LOQ or maximum sampling volumes). The choice of these variables can 

best be explained through the use of the following examples.

a. Sampling for gases and vapors using solid sorbents 

NIOSH Method: 1453 for Vinyl Acetate [NIOSH 2013]

Recommended Sample Volume: 1.5 – 24 L 

Applicable Range: 0.07 to 46 mg/m3 (0.02 to 13.1 ppm) 

NIOSH REL: 14 mg/m3 (4 ppm) - Ceiling 

17.5 mg/m3 (5 ppm) – (European OEL) TWA 

Recommended Flow Rate: 0.05 to 0.2 L/min 

Breakthrough Time: 30 min @ 0.2 L/min and 150 mg/m3

 

Breakthrough Capacity: 9 mg
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General Considerations for Sampling Airborne Contaminants 

Suppose it is desired to determine both ceiling and TWA exposures of workers exposed to 

vinyl acetate and the concentrations are unknown.

1.) Ceiling Determination:

If sampling were done at 0.2 L/min for 30 min and a total sample volume of 6 L 

collected, would this present a breakthrough problem? Probably not, since 

breakthrough tests were carried out in concentrations ca. 10x above the NIOSH 

Ceiling value [NIOSH 2013]. In the breakthrough test, a concentration of 10 times the 

NIOSH Ceiling value (150 mg/m3

) was sampled at 0.2 L/min for 30 min (6 L) before 

breakthrough occurred, collecting a total weight of 9 mg of vinyl acetate. This test was 

conducted in a humid environment with only vinyl acetate present. A safety factor of 

50% should normally be allowed to account for humidity effects. Thus, if sampling is 

done for about 15 min at 0.2 L/min, levels of vinyl acetate up to 40 ppm could still be 

collected without sample breakthrough. 

Also to be considered are the other organics present. If a concentration of 200 ppm 

acetone exists in this environment, then an additional safety factor should be added. 

An arbitrary 50% reduction in sampling rate at 0.1 L/min might be done. With the 

safety factors built in, collecting a 6-L sample should be acceptable.

2.) TWA Determination: 

In a similar situation, the goal is to collect 8-h samples for comparison to the 5 ppm 

TWA [NIOSH 2013]. If sampling were done at 0.1 L/min, then the total sample 

volume would be 48 L, substantially above the 24-L recommended sample volume. If 

the flow was dropped to 0.05 L/min, then the sample volume would be halved (to 24 L, 

the maximum recommended in the method). This sample volume might be acceptable 

if the vinyl acetate concentrations are around 10 ppm and no other competing 

organics are present, e.g., acetone. However, the safer approach would be to collect two 

consecutive samples at 0.05 L/min for 4 h (total sample volume of 12 L each).

b. Pushing a method to the limit ̶ limit of quantitation

NIOSH Method 1009 for Vinyl Bromide (VB) [NIOSH 1994c]

Recommended Sample Volume: <10 L @ 0.20 L/min or less

Working Range: 0.3 to 33 ppm (1.3 to 145 mg/m3

) for a 6-L air sample; 

this equals 8 to 355 μg VB per tube

Limit of Detection: 3 μg VB per tube

In this particular example, let us say that the object is to estimate exposure down to 0.1 

ppm (0.44 mg/m3

), which is below the working range. In order to collect 8 μg of vinyl 
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General Considerations for Sampling Airborne Contaminants 

bromide (the limit of quantitation) at this concentration, 20 L of air will have to be 

collected. This volume is substantially above the maximum recommended sample volume 

of 10 L. Since the recommended sample volume is generally a conservative value used to 

protect against breakthrough under worst case conditions (i.e., high humidity and high 

concentrations), considerable leeway exists for the size of the air sample. In this example, 

the 20-L air samples should be taken at 0.2 L/min or lower, and the possibility of 

breakthrough should be monitored by observing the relative amounts of analyte on the 

backup sections of the samples. 

The best approach is to consult with the analytical laboratory and then to take a sufficient 

number of samples to determine the useful limits of the sampler in the particular 

application. The presence of high relative humidity and other organic solvents will 

severely reduce the number of active sites available on the sorbent for collection of the 

contaminant of interest (with concomitant breakthrough a concern). In pushing a method 

to the limit, it is often necessary to sample beyond the breakthrough volume, normally 

while observing recommended maximum sampling flow rate, in order to obtain the 

sensitivity to determine the concentration of interest. If this is done, then the risk must be 

accepted that the method may not apply outside the limits tested.

c. Sampling for aerosols using a filter

NIOSH Method 7908: Non-volatile Acids (Sulfuric and Phosphoric Acids) 

[NIOSH 2014]

Recommended Sample Volume: 15 ̶ 2000 L

Applicable Range of the Method: 0.010 to > 10 mg/m3

NIOSH REL: 1 mg/m3 (H3PO4) (TWA); 3 mg/m3 (STEL)

Recommended Flow Rate: 1 to 5 L/min

Suppose it is desired to determine both an exposure taking place during a specific 15-

minute operation as well as a TWA exposure of workers exposed to phosphoric acid, and 

the concentrations are unknown.

1.) 15-Minute Process Sample:

This sample would meet the method conditions by sampling for the 15 minutes at 

5 L/min, since this would collect 75 L. Sampling at 1 L/min for 15 minutes would 

probably not allow for the collection of sufficient sample required for analysis.
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General Considerations for Sampling Airborne Contaminants 

2.) TWA Determination:

In this situation, it is necessary to collect an 8-h sample to compare with the 1 mg/m3

NIOSH REL. Since an 8-h TWA sample covers 480 minutes, sampling can no longer 

be done at 5 L/min since this would collect >2000 L, in excess of the upper 

recommended sample volume. Sampling at 1 L/min would collect a 480 L sample, and 

sampling at 2 L/min would collect a 960 L sample: both acceptable per the conditions 

of the method.

6 Bulk samples

The collection of bulk samples can often assist with air sampling efforts. This is especially true 

where there is mixed solvent exposure or unknown dust exposure, and for determining silica 

content of dusts. The primary purpose of obtaining bulk samples is to provide the analytical 

laboratory with a large enough sample for qualitative and sometimes quantitative analysis. 

The two main types of bulk samples are bulk air and mass bulk (liquid or solid) samples.

a. Bulk air samples 

Generally, a bulk air sample is defined as a large volume area sample collected for the 

purpose of qualitative analysis. A good example is a multiple solvent exposure where the 

exact identity of the airborne solvents is unknown, e.g., painting operations. For most 

organic solvents, a bulk air sample consists of a sorbent tube collected at 1 L/min for an 

hour or more. Although the sample is likely to exhibit breakthrough, this does not matter 

since one is primarily interested in identifying which substances are present rather than 

their exact concentrations (the latter aim is accomplished through the separate collection 

of air samples in accordance with defined method parameters). Any questions concerning 

how or whether or not a bulk air sample is needed should be addressed to the analytical 

laboratory prior to sampling. In the case of silica sampling, either a bulk air or solid bulk 

sample (e.g., a deposited sample) or both are suggested so that enough material will be 

available to determine free silica content.

b. Bulk liquids and solids 

Collection of bulk materials may be needed to identify the substances present in the 

workplace and, in some cases, to estimate the relative levels of certain substances present 

in the raw material. A good example of the latter is the case of mixed solvent exposure 

when determining whether a certain contaminant of interest is present, e.g., benzene. In 

some cases, up to 30 solvents may be present, but their identities and proportions are not 

certain. This example is also true for dusts, as was discussed previously for silica, and for 

metals, which may exist in trace quantities.
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General Considerations for Sampling Airborne Contaminants 

In choosing bulk samples for collection, the end goal must be considered: qualitative 

and/or quantitative analysis. Any information that can be given to the laboratory on what 

may or may not be present will help with the analysis. Advance consultation with the 

laboratory is desirable. In choosing bulk dust samples, the sample should be representative 

of the airborne dust to which the workers are being exposed. Usually this is a settled dust 

sample collected from locations near the workers' job site. In other cases, a process dust 

sample may be chosen to determine the composition of the material before it becomes 

airborne. 

When shipping bulk samples, care must be taken to preserve the integrity of the samples 

and to follow established shipping regulations. If applicable, hazardous shipping 

procedures must be followed. Consult with an experienced hazardous goods shipper to 

determine appropriate protocol. Only 5 to 10 mL of the liquid or 10 to 100 mg solid is 

typically needed, so generally bulk sample sizes are kept small. It is important to consult 

with the laboratory before collecting bulk samples to ensure proper sample size and 

containers. In general, leak-proof glass containers are best since they will not react with 

most chemicals; however, polyethylene containers can be used in the majority of cases. A 

convenient container is a 20-mL scintillation vial with PTFE-lined cap. Specific chemicals 

for which polyethylene containers should not be used include aromatic compounds, 

chlorinated hydrocarbons and strong acids. The lids of the containers should be sealed 

with shrink bands or tape for further assurance against leakage. Specific restrictions and 

labeling requirements should be checked prior to shipping any samples.

In the case of volatile bulk samples (and some air samples), consideration should be given 

to shipping the samples on dry ice or with bagged refrigerant (e.g., "blue ice"). Do not ship 

volatile compounds together with air samples. Specific labeling is usually required when 

dry ice is used in shipping.

7 Blanks

Certain numbers of blanks are required by the analytical laboratory for each set of samples to 

be analyzed. The specific method being used should be consulted concerning the number and

type of blanks required. There are two types of sampler blanks: field and media blanks. Field 

blanks are clean samplers taken to the sampling site, handled in the same way as the air 

samples, except that no air is drawn through them. Media blanks are simply unopened, new 

samplers which are sent to the laboratory with the field samples (these blanks are not usually 

taken to the field). It is also recommended that additional blind field blanks be sent along with 

the field samples, and labeled as field samples, as a further check on the analysis. Blanks are 

good insurance to deal with contamination, but the best approach is to avoid sample 

contamination by using careful sampling protocols. The general recommendation for the 
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General Considerations for Sampling Airborne Contaminants 

number of field blanks is three field blanks for each batch of samples. Media blanks should 

also be included. These unexposed, unopened samplers are used to give an estimate of media 

background. The laboratory should analyze at least three (3) media blanks from the same lot 

as the field samples. This number should be increased for media which are coated or 

impregnated with reagent. Again, it is crucial to consult the specific method for the number 

and type of blanks as these numbers will vary. Another recommended practice is to include 

blind spiked samples as quality control checks of the analytical laboratory.

8 Direct-reading methods [Pearce and Coffey 2011]

The variety of types of direct-reading methods available is large and expanding, including 

detector tubes (both short- and long-term), aerosol monitors, passive monitors for certain 

gases and portable instrumentation for gas chromatography or infrared spectrometry [Todd 

1997]. Many direct-reading instruments now used for personal or area measurements have 

evolved from laboratory or process control instruments [Woebkenberg and McCammon 

1995; Todd 1997]. Some direct-reading instruments are screening devices while others give 

quantitative result and can be used for compliance purposes [Song et al. 2001; Ashley et al. 

2002]. 

Some of the considerations (e.g., specificity and sensitivity) for the use of direct-reading 

methods for quantitative determinations are similar to those already given for filter or sorbent 

sampling and analytical methods. In many cases, direct-reading instruments, which are 

physically small and portable, qualify as personal sampling devices.*** These offer the 

additional advantages over classical methods by reducing labor and analytical costs and may 

be the methods of choice when instantaneous results are important, even at the expense of 

some degree of sensitivity or specificity. In general, manufacturers' instructions should be 

followed in the calibration and use of these devices. Because of the severe conditions to which 

direct-reading instruments may be subjected, performance checks and preventive 

maintenance on a periodic basis or before each use are very important. Many direct-reading 

instruments are powered by batteries which can fail to provide a full charge over the full 

sampling period unless frequently or fully discharged and recharged several times just prior to 

use. An additional responsibility, i.e., that of field calibration of the direct-reading instrument, 

falls on the field sampling personnel.

***NOTE: Portable instrument are generally described as weighing less than 5 kg and 

powered by self-contained batteries [Ashley 2003]. For personal monitoring, the instrument 

configuration should be such that the breathing zone can be monitored.
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General Considerations for Sampling Airborne Contaminants 

9 Sampling Strategy

To obtain the maximum amount of information during the course of a sampling campaign 

with a minimum number of samples, a statistical sampling strategy should be developed 

before conducting any study [Leidel et al. 1977; ASTM International 2013]. Several pieces of 

information must be known in advance to plan a sampling strategy, including the size of the 

workforce to be sampled, the accuracy of the sampling and measurement method to be used 

and the confidence one wishes to have in predicting the occupational exposures. 

For example, to determine with 90% confidence that at least one worker from a workplace 

subgroup will be in the top 10% of the exposures occurring in the group, the number of 

employees to sample would be chosen from the scheme below. (Other figures are applicable 

for confidence limits of 95% and for the top 20% of exposures.) Again, judicious use of 

sampling statistics will optimize the number of samples needed.

Table 1. Minimum sample size (n) for including (@ 90% confidence level) at least one high 

risk employeeA [Leidel et al. 1977]

Size of employee group (N)

1 2 3 4 5 6 7 8 9 10 11- 13- 15- 18- 21- 25- 30- 39- 40- 50+

12 14 17 20 24 29 37 29 49

Minimum number of measured employees (n)

1 2 3 4 5 6 7 7 8 9 10 11 12 13 14 15 16 17 18 22

A Exposure in highest 10% of N.

10 Sampling and calibration techniques

The following are suggested general techniques for active sampling using some of the more 

common samplers. These instructions elaborate on those given in NIOSH methods. Consult 

individual methods for details regarding sample size.

a. Calibration of personal sampling pumps 

The accuracy of determining the concentration of a toxic substance in air is no greater 

than the accuracy with which the air volume is measured. Therefore, accurate calibration 

of the airflow rate through the sampling train is necessary. Ordinarily, pumps should be 

calibrated in the laboratory and the field, both before field use and after each field 

sampling campaign. 
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General Considerations for Sampling Airborne Contaminants 

The choice of a reference instrument will depend on where the calibration is to be 

performed. For laboratory use, primary standards, such as a spirometer or soap-bubble 

meter, are recommended [Okladek 1988]. Several electronic soap-bubble calibrators and 

dry-cell calibrators are commercially available as primary calibrators. Other instruments, 

such as a wet-test, mass-flow or a dry-gas meter, may be used. The following instructions 

are for the soap-bubble meter. If another calibration device is used, equivalent procedures 

should be followed.

1.) Set up the apparatus as shown in Figure 1.

2.) Ensure that the rechargeable batteries will power the pump for the entire sampling 

interval by one of the following methods: 1) run the pump for that length of time, 

checking for satisfactory operation; or 2) test the battery independently of the pump 

using a current capacity tester [Kovein and Hentz 1988]. Fully recharge the batteries.

3.) Turn the pump on and moisten the inner surface of the soap-bubble meter with the 

soap solution. Draw bubbles upward until they travel the entire length of the buret 

without breaking.

4.) Adjust the pump to the desired nominal flow rate. Check the water manometer. The 

pressure drop across the sampler should not exceed 2.5 cm Hg of water.

5.) Start a soap bubble in the buret and, with a stopwatch, measure the time that it takes to 

traverse two calibration marks. For a 1000-mL buret, a convenient calibration volume 

is 500 mL. Repeat the determination at least twice more. Average the results and 

calculate the flow rate by dividing the calibration volume by the average time.

6.) Record the following data:

a. volume measured

b. elapsed time

c. pressure drop

d. air temperature

e. atmospheric pressure

f. serial number of the pump

g. pump model

h. date and name of operator.
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General Considerations for Sampling Airborne Contaminants 

7.) If the sampling pump used for sample collection uses a variable area flow meter 

(rotameter) for flow rate indication, the calibrated flow rate must be adjusted for the 

actual air pressure and temperature during sampling [Okladek 1988]. The expression 

for this correction is as follows.

NOTE: This correction is not used for non-rotameter sampling pumps.

V (Corrected volume, L) = Q t (PcTs /PsTc) 0.5 where:

Q = indicated flow rate (L/min) 

t = sampling time (min) 

Pc = pressure during calibration of sampling pump (kPa)

Ps = pressure of air sampled (same units as Pc) 

Tc = temperature during calibration of sampling pump (K) 

Ts = temperature of air sampled (K).

Figure 1. Calibration Apparatus

b. Sampling instructions for solid sorbent tube sampler 

Use these instructions for active personal sampling (i.e., pumped sample airflow) for 

substances which are retained on solid sorbents such as activated charcoal, silica gel, 

porous polymers, etc.
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General Considerations for Sampling Airborne Contaminants 

1.) Calibrate each personal sampling pump at the desired flow rate with a representative 

solid sorbent tube in line. Use a bubble meter or equivalent flow measuring device.

2.) Break the ends of the solid sorbent tube immediately before sampling to provide an 

opening at least one-half of the internal diameter at each end.

3.) Connect the solid sorbent tube to a calibrated personal sampling pump with flexible 

tubing with the smaller sorbent section (backup section) nearer to the pump. Do not 

pass the air being sampled through any hose or tubing before entering the solid 

sorbent tube. Position the solid sorbent tube vertically during sampling to avoid 

channeling and premature breakthrough.

4.) Prepare the field blanks at about the same time as sampling is begun. These field 

blanks should consist of unused solid sorbent tubes from the same lot used for sample 

collection. Handle and ship the field blanks exactly as the samples (e.g., break the ends 

and seal with plastic caps) but do not draw air through the field blanks. A minimum of 

three field blanks are normally required for each batch of samples.

5.) Take the sample at an accurately known flow rate as specified in the method for the 

substance and for the specified air volume. Typical flow rates are in the range 0.01 to 

0.2 L/min. Check the pump during sampling to determine that the flow rate has not 

changed. If sampling problems preclude the accurate measurement of air volume, 

discard the sample. Take a minimum of three replicate samples for quality control for 

each set of field samples.

6.) Record pertinent sampling data including location of sample, times of beginning and 

end of sampling, initial and final air temperatures, relative humidity and atmospheric 

pressure.

7.) Seal the ends of the tube immediately after sampling with plastic caps. Label each 

sample and blank clearly with waterproof identification.

8.) Pack the tubes tightly with adequate padding to minimize breakage for shipment to the 

laboratory. In addition to the sample tubes and field blanks, ship at least six unopened 

tubes to be used as media blanks and three additional tubes so that desorption 

efficiency studies can be performed on the same lot of sorbent used for sampling.
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General Considerations for Sampling Airborne Contaminants 

9.) Ship bulk samples in a separate package from the air samples to avoid contamination 

of the samples. If applicable, hazardous shipping procedures must be followed. 

Consult with an experienced hazardous goods shipper to determine appropriate 

protocol. Suitable containers for bulk samples are glass with a polytetrafluoroethylene 

(PTFE)-lined cap, e.g., 20-mL glass scintillation vials. It is important to consult with 

the laboratory before collecting bulk samples to ensure use of proper sample size and 

containers.

c. Sampling instructions for filter sampler 

Use these instructions for personal sampling of inhalable aerosols. Methods requiring 

these instructions specify FILTER as the sampling method. These instructions are not 

intended for respirable aerosol sampling.

1.) Calibrate the personal sampling pump with a representative filter in line using a 

bubble meter or equivalent flow measuring device.

2.) Assemble a filter or internal capsule in a cassette filter holder. Support the filter by a 

cellulose backup pad or stainless steel screen. Close the filter holder to ensure that its 

parts mate evenly and securely to prevent leakage [Frazee and Tironi 1987]. Seal the 

filter holder with plastic tape or a shrinkable cellulose band. 

3.) Remove the filter holder plugs and attach the filter holder to the personal sampling 

pump with a piece of flexible tubing. Position the filter holder in the worker's personal 

breathing zone, with the sampler inlet pointed downwards.

4.) Prepare the field blanks at about the same time as sampling is begun. These field 

blanks should consist of unused filters (or internal capsules) and filter holders from the 

same lot used for sample collection. Handle and ship the field blanks exactly as the 

samples, but do not draw air through the field blanks. Three field blanks are required 

for each batch of samples.

5.) Sample at the prescribed flow rate (usually 1 to 5 L/min) until the recommended 

sample volume is reached. Set the flow rate as accurately as possible (e.g., within ± 5%) 

using a calibrated flowmeter. Take three replicate samples (minimum) for quality 

control for each set of field samples.
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General Considerations for Sampling Airborne Contaminants 

6.) Observe the sampler frequently and terminate sampling at the first evidence of 

excessive filter loading or change in personal sampling pump flow rate. (It is possible 

for a filter to become plugged by heavy particulate loading or by the presence of oil 

mists or other liquids in the air.)

7.) Disconnect the filter holder after sampling. Cap the inlet and outlet of the filter holder 

with plugs. Label the sample. Record pertinent sampling data, including times of 

beginning and end of sampling, initial and final air temperatures, relative humidity 

and atmospheric pressure. Record the type of personal sampling pump used and 

location of the sampler.

8.) Ship the samples to the laboratory as soon as possible in a suitable container designed 

to prevent damage in transit. Ship bulk material to the laboratory in a chemically inert 

container. Never store, transport or mail bulk samples in the same container as the 

samples or field blanks. In addition to the samples and field blanks, ship at least four 

unused filters or filter capsules from the same lot for use as media blanks.

d. Sampling instructions for filter + cyclone sampler 

Use these instructions for personal sampling of respirable (or thoracic) aerosols [Frazee 

and Tironi 1987]. Methods requiring these instructions specify CYCLONE + FILTER as 

the sampling method.

1.) Calibrate the pump to the rate specified by the cyclone, with a representative cyclone 

sampler in line using a bubble meter (or a secondary flow measuring device which has 

been calibrated against a bubble meter). The calibration of the personal sampling 

pump should be done close to the same altitude where the sample will be taken.

2.) Assemble the pre-weighed filter in the cassette filter holder. Use a conductive or staticdissipative cassette. Support the filter with a cellulose backup pad or stainless steel 

screen. Close firmly to prevent sample leakage around the filter. Seal the filter holder 

with plastic tape or a shrinkable cellulose band.

3.) Remove the cyclone's grit cap and vortex finder before use and inspect the cyclone 

interior. If the inside is visibly scored, discard this cyclone since the dust separation 

characteristics of the cyclone might be altered. Clean the interior of the cyclone to 

prevent reentrainment of large particles.
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General Considerations for Sampling Airborne Contaminants 

4.) Assemble the two-piece filter holder, coupler, cyclone and sampling head. The 

sampling head rigidly holds together the cyclone and filter holder. Check and adjust 

the alignment of the filter holder and cyclone in the sampling head to prevent leakage. 

Connect the outlet of the sampling head to the personal sampling pump by a 1-m piece 

of 6-mm ID flexible tubing.

5.) Position the cyclone assembly in the worker's personal breathing zone and attach the 

personal sampling pump to a belt or harness. Ensure that the cyclone hangs vertically 

with the inlet pointed downwards. Explain to the worker that the cyclone must not be 

inverted.

6.) Prepare the field blanks at about the same time as sampling is begun. These field 

blanks should consist of unused filters and filter holders from the same lot used for 

sample collection. Handle and ship the field blanks exactly as the samples, but do not 

draw air through the field blanks. A minimum of three field blanks are required for 

each batch of samples.

7.) Turn on the pump and begin sample collection. If necessary, reset the flow rate to the 

pre-calibrated value, using the manufacturer's adjustment procedures. Since it is 

possible for a filter to become plugged by heavy particulate loading or by the presence 

of oil mists or other liquids in the air, observe the filter and personal sampling pump 

frequently to keep the flow rate within ± 5% of the target flow rate. Sampling should 

be terminated at the first evidence of a problem.

8.) Disconnect the filter after sampling. Cap the inlet and outlet of the filter holder with 

plugs. Label the sample. Record pertinent sampling data, including times of beginning 

and end of sampling, initial and final air temperatures and atmospheric pressure or 

elevation above sea level. Record the type of personal sampling pump, filter, cyclone 

used and the location of the sampler.

9.) Ship the samples and field blanks to the laboratory in a suitable container designed to 

prevent damage in transit. Ship bulk samples in a separate package.

10.) Take a minimum of three replicate samples for every set of field samples to assure 

quality of the sampling procedures. The set of replicate samples should be exposed to 

the same dust environment, either in a laboratory dust chamber or in the field. The 

quality control samples must be taken with the same equipment, procedures and 

personnel used in the routine field samples. The relative standard deviation, sr, 
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General Considerations for Sampling Airborne Contaminants 

calculated from these replicates, should be recorded on control charts and action taken 

when the precision is out of control.

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

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for Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 

[www.cdc.gov/niosh/nmam/]. 

NIOSH [1994b]. Formaldehyde: Method 1206. In: Eller PM, Cassinelli ME, eds. NIOSH 

manual of analytical methods (NMAM). 4th ed. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 

[www.cdc.gov/niosh/nmam/].

NIOSH [1994c]. Vinyl Bromide: Method 1009. In: Eller PM, Cassinelli ME, eds. NIOSH 

manual of analytical methods (NMAM). 4th ed. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 

[www.cdc.gov/niosh/nmam/].

NIOSH [2013]. Vinyl Acetate: Method 1453. In: Ashley K, O’Connor PF, eds. NIOSH 

manual of analytical methods (NMAM), 5th ed. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2014-151 

[www.cdc.gov/niosh/nmam/].

NIOSH [2014]. Non-Volatile Acids: Method 7908. In: Ashley K, O’Connor PF, eds. NIOSH 

manual of analytical methods (NMAM), 5th ed. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for 
NIOSH Manual of Analytical Methods  5th Edition  Chapter SA April 2016 Page SA-23 of SA-23

General Considerations for Sampling Airborne Contaminants 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2014-151 

[www.cdc.gov/niosh/nmam/].

Okladek J [1988]. An overview of flow metering devices. Am Lab 20:84-90.

Pearce T, Coffey C [2011]. Integrating direct-reading exposure assessment methods into 

industrial hygiene practice. J Occup Environ Hyg 8:D31-D36. 

Song R, Schlecht PC, Ashley K [2001]. Field screening test methods – Performance criteria 

and performance characteristics. J Hazard Mater 83:29-39. 

Tejada SB [1986]. Evaluation of silica gel cartridges coated in situ with acidified 

2, 4-dinitrophenylhydrazine for sampling aldehydes and ketones in air. Int J Environ Anal

Chem 26:167-185. 

Todd LA [1997]. Direct-reading instruments for gases, vapors, and aerosols. In: DiNardi SR, 

ed. The occupational environment – its evaluation and control. Fairfax, VA: American 

Industrial Hygiene Association, pp. 177-208. 

Woebkenberg ML, McCammon CS [1995]. Direct-reading gas and vapor Instruments. In: 

Cohen BS, Hering SV, eds. Air sampling instruments. 8th ed. Cincinnati, OH: ACGIH, 

pp 439-510. 
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Factors AffectingAerosolSampling 

by Paul A. Baron, Ph.D., NIOSH 

The NMAM team gratefully acknowledges Bon-Ki Ku, Kevin Ashley and Pramod Kulkarni 

for insightful review of this chapter. 

1 Introduction AE - 2

2 Inlet efficiency of the sampler AE - 8

3 Classifier accuracy AE - 11

4 Sampler assembly AE - 15

5 Electrostatic losses AE - 16

6 Sampler deposition uniformity AE - 18

7 Sampler wall losses AE - 19

8 Collection media and analytical issue AE - 20

9 Sampler field comparisons AE - 22

10 Conclusions AE - 24

11 References AE - 25
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Factors Affecting Aerosol Sampling 

1 Introduction

The need for aerosol sampling is driven by research or regulatory needs to understand or 

quantify the properties of airborne particles in the workplace or ambient environments. The 

property of most common interest is the airborne concentration of particulate mass defined as 

the aerosol mass per unit volume of air, usually expressed in units of micrograms or 

milligrams per cubic meter. Alternatively, concentrations of other related properties such as 

surface area or number, or particle size distributions are also of interest in certain cases, 

especially where exposure to nanoparticles or ultrafine aerosols are involved. In many 

applications, airborne concentration of a certain chemical or analyte, usually expressed in 

terms of micrograms per cubic meter, is more important. Aerosol sampling is the process of 

collecting a representative sample of airborne particles of interest from the air environment by 

physically separating them from the sampled air of known volume. The degree to which the 

physically separated sample represents the in situ aerosol depends on the design of the 

physical separation device, often known as the aerosol sampler. Other factors that affect the 

representativeness of the particulate sample include environmental conditions (e.g., wind, 

temperature, humidity), particle characteristics (particularly if they are highly irregular or 

nonspherical), and subsequent analytical methods used for particle analysis. This chapter 

focuses mainly on the key characteristics of aerosol samplers that may influence the 

representativeness of the sampled aerosol. Direct-reading aerosol samplers are not discussed 

in this chapter. 

Most samplers use size-selective inlets that conform to certain health-based conventions. The 

American Conference of Governmental Industrial Hygienists (ACGIH) [Vincent 1999a; 

ACGIH 2015], the International Organization for Standardization (ISO) [ISO 1995], and the 

European Standardization Organization (Comité Europeén de Normalisation, CEN) [CEN 

1993] have adopted identical particle size-selective sampling conventions for inhalable, 

thoracic, and respirable aerosols (Figure 1). The purpose of these conventions is to provide a 

scientific basis for a new generation of particle size-selective occupational exposure limits 

(OELs) for aerosols. Such OELs can therefore be matched to the relevant sites of aerosol 

deposition after inhalation into the respiratory tract, and in turn to the health effects of 

interest in a given exposure assessment. These sampling conventions are used throughout this

manual unless otherwise specified.
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Factors Affecting Aerosol Sampling 

Figure 1. ISO/ACGIH/CEN sampling conventions [ISO 1995]. An ideal sampler should 

have a sampling efficiency curve that matches one of these curves as closely as possible 

under all wind directions and velocities. The 50% cut points for the respirable and thoracic 

conventions are 4 and 10 μm, respectively.

The criteria presented in this manual are used to determine the most appropriate aerosol 

sampling equipment. The type of sampling to be performed determines which criteria are 

important for estimating the adequacy of the sampler and determining aerosol concentration 

levels. For example, “total dust” samplers generally do not have a size selective particle 

classifier preceding the filter media and fall under the inhalable sampling convention. 

Alternatively, sampling for regulatory or voluntary compliance with aerosol exposure 

standards usually requires greater accuracy, increased efficiency, size-specific selectivity, and 

good analytical precision. Furthermore, regulations may require the use of a specific sampler 

and sampling conditions to standardize sampling results (eliminate bias) and reduce 

uncertainty among laboratory reports. See the chapter on measurement uncertainty and 

NIOSH method accuracy for further discussion on standardization and aerosol measurement 

error.

Open-face filter cassettes do not use constricted opening or tubing and expose the filter 

directly to the aerosol to minimize the losses. They provide relatively uniform particle 

deposition on the filter. On the other hand, closed-face filter cassettes are often necessary to 

connect to upstream tubing or size-selective inlets. 
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Factors Affecting Aerosol Sampling 

Over the past decades, researchers have pointed out strengths and weaknesses with several 

types of aerosol samplers (Figures 2A-J). Some of these samplers were adapted from existing 

devices used for other purposes, e.g., the 10-mm nylon cyclone and the 37-mm cassette, 

without the benefit of current testing technology and understanding of particle behavior. 

Figure 2A. 37-mm (top) and 25-mm (bottom) filter cassettes. Open faced cassettes are 

shown on left. Closed-faced cassettes include placement of clear piece (on right) over filter. 

These are shown in acrylic copolymer (clear, non-conducting) material and are more 

prone to electrostatic losses. Other construction materials are available, including 

conducting plastic. Sampling flow rates range from 0.5 to 10 L/min. 

Figure 2B. IOM inhalable sampler. This is the first sampler designed specifically to match

the inhalable sampling convention. The sampling cartridge is shown on the right with the 

inlet, filter, and support grid. All dust entering the cartridge is collected and analyzed.

Sampler is made of conductive plastic and operates at 2 L/min. 
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Factors Affecting Aerosol Sampling 

Figure 2C. Seven hole sampler used for inhalable dust sampling in the UK. Sampler is 

made of conductive plastic and operates at 2 L/min. 

Figure 2D. Button sampler. So-named because the inlet, a hemispherical screen with ~380 

μm diameter holes, resembles a large button. It was developed as an inhalable samplerwith

reduced wind direction response and improved filter deposit uniformity. The sampler uses

metal construction and operates at 4 L/min.

Figure 2E. Asbestos sampling cassette. The long (50 mm) inlet was designed to prevent 

incidental contactwith the filter surface and,whenfacing downward, actsto a certain extent

as an elutriator, preventing larger particlesfrom reaching the filter surface. It is made of

conductive plastic and is operated at a flowrate between 0.5 L/min to 16 L/min. 
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Factors Affecting Aerosol Sampling 

Figure 2F. Bell mouth cowl sampler. An alternative to the standard asbestossampling

cassette. The inlet is flared to reduce the effect of external air motion on sample

uniformity. It is made of conductive plastic and is operated at a flow rate between 

0.5 L/min to 16 L/min.

Figure 2G. Dorr-Oliver 10-mm nylon cyclone. It is used as a respirable sampler most

often at 1.7 L/min. A version of this sampler is used at 2 L/min in coal mines with a 1.38

correction factor appliedto the resultantmass. The holder encasesbodyof the cyclone so

thatonly the inletsection is visible just below the connection to the 37 mm cassette. The

coal mine dust sampler version uses a cassette with an aluminum cartridge encasing the 

collection filter.
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Factors Affecting Aerosol Sampling 

Figure 2H. Higgins-Dewell cyclone. This sampler was developed and used primarily as a 

respirable sampler in the UK. This version provides aninterfacedirectlyto a 37-mm

cassette. Cyclone construction issteel and operates at 2.2 L/min.

Figure 2I. GK2.69 cyclone. This cyclone was designed to match the slope of the thoracic

and respirable conventions more closely than other sampling cyclones. Cyclone

construction is stainless steel and operates at 4.2 L/min as a respirable sampler and 1.6

L/min as a thoracic sampler. GK4.162 RASCAL Cyclone is also available for higher flow 

rates of 8.5-9 L/min.
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Factors Affecting Aerosol Sampling 

Figure 2J. Personal cascade impactor sampler (model Marple 290, Thermo Fisher 

Scientific). The sampler has five stagesin the version shown and an additional three stages

are available to provide cutpoints from 0.4 μm to 21 μm. The flap over the inlet reduces 

direct projection of large particles into the inlet. This sampler allows measurement of 

aerosol size distribution and calculationof respirableand thoracicfractions. It is 

constructed of aluminumand operatesat 1.7 L/min.

More recently developed aerosol samplers were designed to either maximize the information 

gained regarding aerosol concentration levels or to minimize any inherent losses associated 

with the sampler design. Thus, each sampler design may have been based on some of the 

following criteria: inlet or aspiration efficiency, classifier accuracy, cassette assembly (bypass 

leakage), electrostatic losses, particle deposition uniformity, collection media stability, sampler 

surface losses, and sampler field comparisons. These criteria are discussed in the following 

sections of this chapter.

2 Inlet efficiency of the sampler

An important review of sampling theory and practice was compiled in a book by Vincent 

[2007]. The inlet efficiency of several samplers has been evaluated including thin-walled 

tubular inlets [Grinshpun et al. 1993], a cyclone [Cecala et al. 1983], an asbestos sampler 

[Chen and Baron 1996], total aerosol sampling cassettes [Fairchild et al. 1980], and inhalable 

aerosol samplers [Kenny et al. 1997; Aizenberg et al. 2001]. All samplers have an inlet 

efficiency, also called aspiration efficiency, which varies as a function of particle aerodynamic 

diameter, inlet velocity, inlet shape and dimensions, dimensions of the body it is attached to, 

external wind velocity, and external wind direction. The aspiration efficiency of an aerosol 

sampler can be defined as

AA = CCSS

CC0

where CS is the concentration of particles passing directly through the plane of the sampling

orifice and C0 is the ambient concentration. This is true for aerosol samplers for which the

entire amount of aerosol that enters the plane of the sampling orifice is quantified, as is the
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Factors Affecting Aerosol Sampling 

case for the IOM sampler (SKC, Inc., Eighty Four, PA). However, it is important to note that

many commercially available aerosol samplers use only the aerosol collected on the filter that

is housed in the sampler, while any particles deposited on the inner surfaces upstream of the

filter are disregarded. Performance of this type of sampler is therefore best characterized by its 

sampling efficiency, in which the aspiration efficiency is modified by the particle sizedependent wall losses prior to the filter. The aspiration or sampling efficiency of a particular 

aerosol sampler, whichever is the most relevant, expressed as a function of particle 

aerodynamic diameter, is the primary index of sampler performance. The overall sizedependent transmission efficiency of the sampler must match the appropriate health-based 

criterion (e.g., the inhalability criterion) to allow its use for a health-based assessment of 

personal aerosol exposure.

As an aerosol is being sampled, the large-particle trajectories are more affected by external

flow fields than those of small particles. Thus, the shape, orientation, and inlet flow field will

be most critical for inhalable aerosol sampler inlets; they will be less important for thoracic

samplers and unlikely to be important for respirable samplers, except at very high wind

velocities. The flow field near the inlet of a sampler is different when the sampler is mounted

on a person (or in laboratory simulations using a mannequin) than when it is freestanding.

Therefore, it has been recommended that measurement of inhalability be determined from

mannequins in wind tunnels [Vincent 1999a; Kennedy et al. 1995; Kennedy and Hinds 2002; 

Aitken et al. 1999]. Flow field studies using mannequins in wind tunnels indicate that the air 

is slowed down by the body, resulting in an enrichment of large particles in the upstream side 

of the body [Rodes et al. 1995]. When intended for use at low wind speeds (less than about

1 m/s [200 ft/min] (representing most indoor workplaces), it may be possible to test the 

samplers as freestanding devices if it can be shown there is no effect from the mannequin 

body [CEN 1997]. However, if the sampler is to be used at higher windspeeds, which 

frequently occur outdoors, personal aerosol samplers should be evaluated on a mannequin in 

a wind tunnel [CEN 1997].

Respirable aerosol samplers generally do not have problems with inlet effects because the

particles being sampled have low enough inertia and settling velocity. However, Cecala et al.

[1983] found that the 10-mm nylon cyclone operated free-standing in a wind tunnel 

oversampled at external air velocities greater than 4 m/s when the inlet faced the wind and 

undersampled at 90° and 180° to the wind at velocities greater than 1 m/s. The maximum 

sampling error of about 40% was observed at 10 m/s. It is expected that these errors would be 

reduced if the sampler were located on a person, because the air velocity decreases near the 

body surface. It should be noted that the Cecala et al. work was conducted in the context of 

aerosol sampling in underground mining environments. Here, windspeeds of the magnitude 

quoted are not uncommon. However, in industrial workplaces more generally, windspeeds are 

much lower. Two surveys of a wide range of workplaces [Baldwin and Maynard 1998; Berry 
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Factors Affecting Aerosol Sampling 

and Froude 1989] revealed that actual indoor windspeeds rarely exceeded 0.2 to 0.3 m/s and 

more typically were less than 0.1 m/s.

The EPA PM10 standard for environmental sampling specifies a sampler that has a 50%

cutpoint at 10.6 μm particle diameter, approximately the same as that for the thoracic sampler 

[Baron and John 1999]. Although the requirements for environmental PM10 samplers stipulate 

wind tunnel testing, similar work on personal PM10 and thoracic samplers is yet to be 

performed. It is expected that these samplers will be more susceptible to wind effects than 

respirable samplers because larger particles are more susceptible to inertial and gravitational 

effects.

The most extensive comparison of available inhalable aerosol samplers was that carried out

under the auspices of the European Commission (EC) [Kenny et al. 1997]. Eight samplers 

were tested: CIP-10 (foam-based, French); 37-mm closed-face cassette (Spain and US) (Figure 

2A); 37-mm open-face cassette (Sweden) (Figure 2A); PAS-6 (Netherlands); PERSPEC (Italy); 

GSP (Germany, sold as CIS sampler in US; BGI, Inc. Waltham MA); IOM (United Kingdom; 

(Figure 2B); and the Seven-Hole Sampler (United Kingdom; Casella CEL, Inc., UK) (Figure 

2C). Conditions of the experiment included measurement of sampler collection efficiencies on

a mannequin for aerosol particles with diameters as large as 100 μm at a wind speed of 

0.5m/s, 1.0 m/s, and 4 m/s. Samplers were positioned on a mannequin rotating within a wind 

tunnel. The samplers were all conductive; the 37-mm cassette samplers were painted with an 

external conductive coating. The aerosol was, however, not neutralized. The results of this 

experiment indicated high inter-sampler variability, but permitted estimates of bias relative to 

the inhalable convention. The EC study also indicated that most samplers work reasonably 

well at low wind speeds (<1 m/s) for particle median diameters below 25 μm [Kenny 1995]. 

The study indicated that experiments of this type were difficult, expensive, and generally had 

poor precision. Perhaps better understanding of the flow field near the body may lead the way 

to improved and simplified sampler testing. Recent work suggests ways of making the wind 

tunnel testing of inhalable samplers simpler and less expensive, e.g., by using a compact body 

to simulate the chest of a mannequin [Witschger et al. 1997] and by using miniaturized 

mannequins and samplers that are calculated to be aerodynamically equivalent 

[Ramachandran et al. 1998].

The orientation and diameter of an inhalable sampler inlet may affect the collection of very 

large particles (generally >100 μm), since these may be thrown into the inlet as projectiles. The

current definition of inhalable aerosol only covers particles up to100 μm aerodynamic 

diameter.

In situations where large particles can be generated (e.g., abrasive blasting, wood working, and

grinding operations) excessive collection of particles up to the millimeter range is likely to 
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Factors Affecting Aerosol Sampling 

occur. There have been some attempts to modify inlets with shields to provide a barrier

against the collection of large particles, but these modified inlets have not been demonstrated 

to provide the same agreement with the inhalable convention as the unmodified ones.

Another potential problem with inhalable samplers is the collection of passively sampled

particles. Measurements when the sampler airflow is turned off indicate that IOM samplers,

which pointed outward from the body and had a large inlet diameter (15-mm), can collect 

quite significant amounts of dust, with median values of 9 to 32 percent of the mass collected 

during active sampling [Lidén et al. 2000b]. Open-faced cassettes had only 2 to 11 percent of 

the mass passively collected. These samplers have a larger inlet (37-mm), but point downward, 

reducing the likelihood of particle settling onto the collection surface. The mechanism of 

collection is unclear, but the dust may be transported into the inlet by turbulence and 

deposited by settling or turbulent diffusion. How this passively collected dust modifies the 

amount collected during active sampling remains under investigation.

A comparison of measurements obtained with the 37-mm closed-face cassette (4-mm inlet

diameter) to the IOM sampler (15-mm inlet diameter) in several workplaces gave similar 

results when the material on the interior walls of the 37-mm cassette were added to the analyte

deposited on the filter [Demange et al. 2002; Harper and Demange 2007]. This suggests that 

the two samplers can have similar inlet efficiencies in spite of differences in inlet size and 

orientation if the median particle size sampled is not too large. Therefore, if the total 

aspiration (which includes the mass from filter and the wall deposits) of the IOM sampler 

conforms with the ISO inhalable size-selection criterion, then so does the total aspiration of 

the closed cassette filter. Several studies have now shown that in metals industries the total of 

mass from particulate filter and the wall deposit are comparable for both the closed cassette 

filter and the IOM samplers [Harper and Demange 2007].

3 Classifier accuracy

The theory of classifier separation is based on particle aerodynamic diameter, which is defined 

as the diameter of a 1 g/cm3 density sphere having the same gravitational settling velocity as 

the particle in question. If the particle is markedly nonspherical or irregularly shaped, the 

aerodynamic diameter may depend on particle’s orientation and other factors, possibly 

contributing to sizing errors. For example, fibers and plate-like particles settle slightly 

differently depending on orientation [Kulkarni et al. 2011]. Thus, the sampling conventions, 

based on aerodynamic diameter of particles reaching specified parts of the respiratory system, 

become somewhat ambiguous for these types of particles. For such nonspherical particles, 

further testing of classifiers to simulate particle behavior in the respiratory tract may be 

necessary. For instance, Maynard [1996] found that plate-like particles may orient differently 

in elutriators, impactors, and cyclones. This preferred orientation in a cyclone produced a 
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Factors Affecting Aerosol Sampling 

collection efficiency 15% below that estimated to occur in the respiratory system. In addition, 

Baron et al. [2008] showed that the overall enveloping physical size of airborne single-walled 

carbon nanotube (SWCNT) agglomerates is much larger than their aerodynamic size, by a 

factor of up to 10. Ku and Kulkarni [2015] measured both aerodynamic and mobility (or 

diffusion) diameters of airborne carbon nanotubes (CNTs) and other nanomaterials to show 

that aerodynamic diameter is smaller by a factor of 2 to 4 than mobility diameter for SWCNT 

and multi-walled CNT particles. These studies indicate that relevant equivalent diameters 

must be used to obtain reliable estimation of lung deposition fraction. Improved 

understanding of fiber [Esmen and Erdal 1991], nanotube [Baron et al. 2008; Ku and 

Kulkarni, 2015] and plate-like particle [Maynard 1996] behavior in the respiratory tract is 

needed to aid in development of more accurate samplers for these types of particles. The 

phase (i.e. liquid or solid) of the aerosol particles also influences sampling errors. Koehler et 

al. [2012] examined the sampling efficiency as a function of particle phase of three personal 

aerosol samplers, including the IOM and button sampler. They found that large liquid 

droplets have low transmission efficiencies through the screened inlets and that the bounce of 

solid particles significantly affects the aspiration efficiencies of screened inlets.

Various types of classifiers have been constructed to meet the ACGIH/ISO conventions. For 

example, respirable samplers have used cyclones [Caplan et al. 1977], impactors [Marple 1978; 

Kimura 1978; John 1994], elutriators [Lynch 1970], and porous foam [Brown 1980; Courbon 

et al. 1988] to remove non-respirable particles from the aerosol prior to filter collection. The 

technology for testing these samplers has improved in recent years through use of a real-time 

aerodynamic sizing instrument and resulted in quicker and more precise measurements 

[Baron 1993; Gudmundsson and Lidén 1998]; this technique has allowed the accuracy of these 

samplers to be investigated more carefully [Bartley et al. 1994]. However, a round-robin 

comparison of 50% cut-point measurements from six laboratories using an aerodynamic 

sizing instrument to test the same cyclone agreed within a range of 11% [Lidén 2000a]. 

Further work on the testing protocol is needed to improve interlaboratory agreement. Many 

current classifiers do not match the shape of the respirable convention exactly and produce 

biases that depend on size distribution. Two comparisons of several respirable samplers have 

been performed using the aerodynamic sizing technique [Chen et al. 1999b; Görner et al. 

2001]. 

The introduction of the thoracic fraction in the ACGIH/ISO conventions has spurred interest 

in thoracic classifiers for certain types of aerosols, e.g., cotton dust, asbestos and sulfuric acid 

[Baron and John 1999]. The performance characteristics of the vertical elutriator (operated at 

7.4 L/min) used for cotton dust approximately meets the thoracic definition [Robert 1979]. 

Laboratories in many countries perform asbestos fiber measurement using the technique of 

counting only fibers with diameters of 3 μm or less; this size selection was shown to be 

approximately equivalent to thoracic sampling [Baron 1996]. Further tests indicate that 
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Factors Affecting Aerosol Sampling 

several thoracic samplers may be appropriate for asbestos sampling [Jones et al. 2001; 

Maynard 1999]. Thoracic sampling is also recommended for sulfuric acid [Lippmann et al. 

1987] and metal working fluids [NIOSH 1998].

Several samplers based on inertial, cyclone and foam separators have been specifically 

developed to meet the thoracic definition [Fabriès et al. 1989; Fang and Lippmann 1995; Mark 

et al. 1988; Kenny and Gussman 1997]. The CIP-10 sampler has been used for thoracic 

sampling in Europe [Gorner et al. 1994], but is not applicable to aerosols with a significant 

submicrometer fraction [Fabriès et al. 1989]. Several of these samplers have been tested to 

compare with the thoracic convention [Jones et al. 2001; Maynard 1999]. The GK2.69 cyclone 

(Figure 2I) has been used for metal working fluids [NIOSH 1998] and GK4.162 cyclone has 

been used for measurement of crystalline silica [Qi et al., 2015]. Several developmental 

samplers have also been developed. Koehler and Volckens [2013] have developed multistage 

regional deposition sampler that allows estimation of regional deposition of aerosol in the 

human respiratory system. This sampler is not suitable for gravimetric analysis but is well 

suited for measurement using variety of chemical analyses. A personal nanoparticle 

respiratory deposition sampler was developed by Cena et al. [2011] for particles smaller than 

300 nm diameter, whose aspiration efficiency curves matches the fractional International 

Commission on Radiological Protection (ICRP) deposition curve for human respiratory tract 

below ~300 nm. Tsai et al. [2012] have developed a personal nanoparticle sampler which 

simultaneously collects both respirable and nanoparticles fraction (<100 nm aerodynamic 

diameter).

The PM10 standard for environmental sampling is very similar to the thoracic convention and 

impactors with a 10 μm cutoff size have been used for personal PM-10 sampling [Buckley et 

al. 1991]. A cascade impactor, e.g., the Andersen personal cascade impactor (Figure 2J), can be 

used to calculate the thoracic fraction of an aerosol. Although a thoracic sampler is 

commercially available (Figure 2I), further work is needed to determine its applicability for 

specific types of aerosol. For example, a thoracic sampler for fibers must result in a uniform 

deposit of the particles on the filter for accurate analysis results.

The overall accuracy of a classifier with respect to sampling in accordance with the one of the 

sampling conventions can be estimated using a bias map (Figure 3). The bias map displays the 

percent difference between the predicted mass collected by the sampler and the mass expected 

according to the convention as a function of the parameters of a lognormal particle diameter 

distribution for a range of likely workplace distributions. Such a bias map can be used for 

selecting a sampler for a workplace having a certain range of particle sizes or for developing 

samplers that agree more closely with the sampling conventions.
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Factors Affecting Aerosol Sampling 

Figure 3. Bias map for the 10-mm nylon cyclone (operated at 1.7L/min) compared with the 

respirable convention [Courbon et al. 1988]. The contour lines represent percent bias for 

specific lognormal size distributions. The calculation of this map is based on laboratory 

measurement of cyclone penetration and can be used with field size distributions to 

estimate sampling bias.

The bias map in Figure 3 was created by: (a) fitting the penetration curve for the 10-mm nylon 

cyclone (Figure 2G) [Gudmundsson and Lidén 1998] at 1.7 L/min with a lognormal curve (a 

logistic curve also can be used), (b) calculating the bias between the respirable convention and 

the curve from the previous step for a range of lognormal size distributions, and (c) plotting 

the bias contour lines as a function of the size distribution mass median aerodynamic 

diameter (MMAD) and geometric standard deviation (GSD). The 10-mm nylon cyclone 

shows significant negative biases, especially at large MMADs and small GSDs, because the 

cyclone penetration curve drops off more rapidly with size than the curve for the respirable 

convention. The “best” flow rate to use in a workplace when sampling according to one of the 

conventions becomes a matter of judgment, depending on the size distribution typically 

encountered in that workplace. A cyclone that fits the convention more exactly will exhibit 

smaller biases throughout the entire size distribution range. Bias maps are available for several 

respirable samplers [Chen et al. 1999b; Görner et al. 2001]. It should be mentioned that in 

some cases, e.g., coal mine dust sampling, a single sampler is specified by regulation. This 

sampler specification eliminates the question of bias for that type of measurement.

Investigation of the effect of changing the physical dimensions of a commercial cyclone 

resulted in modifications that improved the match to the respirable sampling convention 
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Factors Affecting Aerosol Sampling 

[Lidén and Gudmundsson 1996]. Chen and coworkers developed a virtual cyclone that 

appeared to give excellent compliance with the respirable curve [Chen et al. 1999a]. It is 

possible to make samplers that have predicted biases less than 10% over the entire range of 

likely workplace size distributions. While the behavior of certain samplers, such as impactors, 

can be predicted theoretically, it is still important to measure penetration curves 

experimentally to ensure correct application of the theory. Bias maps based on these data then 

allow estimation of accuracy for a specific workplace application. As improved samplers are 

tested and become commercially available, more accurate thoracic and respirable aerosol 

measurement on a routine basis will be possible.

As mentioned above, several inhalable samplers were investigated in a wind tunnel to evaluate

their sampling efficiency compared to the inhalable convention [Kenny et al. 1997]. Based on 

these data, sampler performance (maximum bias confidence limit) was ranked [Bartley 1998] 

and the IOM, GSP, and CIP-10 samplers were rated the best.

As interest in the new particle size-selective conventions by standards setting bodies has

grown, efforts have been made to define protocols to guide the testing and validation process

for available samplers. One approach was developed by the CEN [Lidén 1994; CEN 1998]. In 

the CEN model, for any given sampler to be tested, the first step is a critical review of the 

sampling process for the instrument in question. This is intended to identify factors that may 

influence the performance of the sampler, including particle size, windspeed, aerosol 

composition, filter material, etc. This is essential in the process of sampler evaluation, 

determining under what conditions the sampler will need to be tested. Three options are then 

presented for the testing of samplers: (a) the laboratory testing of samplers to compare 

performance with the sampling conventions, (b) the laboratory comparison of instruments, 

and (c) the field of comparison of instruments. Research projects have been conducted in 

recent years to define testing protocols (option a), funded both by the European Community 

and by NIOSH, to consolidate the scientific basis for such protocols and to identify improved 

and more cost-effective methods.

4 Sampler assembly

Some samplers are designed such that improper assembly can result in internal leakage, i.e., 

aerosol particles bypassing the filter. This bypass leakage has been noted in the 37-mm closedface cassette [Frazee and Tironi 1987; Van den Heever 1994]. Although at least one study 

found no problem with hand assembly of these cassettes [Puskar et al. 1991], NIOSH and 

others have occasionally observed, after sampling black or colored dusts, streaks of dust on the 

filter’s compression seal region or an incomplete compression mark, indicating aerosol 

leakage bypassing the filter. An airtight seal in these cassettes is achieved by compression of 

two plastic parts that must be parallel and joined with the proper force. If this seal is not 
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Factors Affecting Aerosol Sampling 

compressed with sufficient force, the vacuum behind the filter may pull the filter from the 

seal, especially at high flow rates. Too high a compression force results in cracking the cassette 

or cutting the filter, also producing leakage.

Prudence dictates that a check of cassette integrity and the seal area on each filter should be 

made after sampling to ensure that the cassette was properly assembled; otherwise, the sample 

may underestimate the actual exposure.

At least two approaches to eliminating the sealing problem with press-fitted cassettes have 

been taken. One was to assemble the cassette using a press. Pressing the cassette together by 

hand often produces misalignment of the cassette parts, resulting in bypass leakage. Frazee 

and Tironi [1987] designed a mechanical press that held the two cassette pieces in proper 

alignment, while applying just enough pressure to effect a seal, but not so much as to cause 

cracking of the plastic. This press was designed to allow motion of the cassette pieces to 

compress to a certain distance. A commercial pneumatic press (Accu-PressTM, Omega 

Specialty, Chelmsford, MA) used a selected pressure to compress the cassette components. 

For additional information on bypass leakage and bypass leak test procedures see [Baron 

2002]. The second approach was to redesign the cassette to provide a more positive filter seal 

[Van den Heever 1994]. In a well-designed sampler, opening the seal should not cause tearing 

and loss of the filter or collection medium during removal from the sampler.

The 37-mm closed-face cassette is usually sealed with tape or shrink bands around the 

outside. There is a common misconception that this seal prevents bypass leakage in the 

cassette. These external seals primarily cover the joint between the cassette components to 

prevent deposited particles on the external surface of the cassette from contaminating the 

sample during filter removal. The tape or shrink band also aids in holding the cassette 

together and preventing external air leakage. However, Puskar, et al. [1991] found that even 

by using this precaution, a significant amount of dust was found downstream of the filter. The 

authors hypothesized that this dust was deposited during filter removal.

Three other commercial samplers, the IOM (Figure 2B), the CIS, and the coal mine dust 

sampler (MSA, Inc., Pittsburgh, PA) use a cartridge to hold the filter. For the first two 

samplers, an external threaded cover applies pressure to the cartridge to ensure a good seal 

around the filter. This prevents twisting at the filter surface while creating positive, even 

contact around the filter edge.

5 Electrostatic losses

Most aerosol particles generated in workplaces have electrostatic charge levels considerably 

higher than the steady-state or equilibrium charge level [Johnston et al. 1985]. The 
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Factors Affecting Aerosol Sampling 

equilibrium charge (Boltzmann equilibrium) levels are usually achieved after particles are

suspended in the ambient atmosphere for approximately an hour in the presence of naturally 

occurring atmospheric ions of positive and negative polarity. When freshly generated particles 

are sampled in the presence of an electric field, as when a sampler walls are highly charged 

[Baron and Deye 1990b], the particle trajectories can be modified to such an extent that the 

particles are inefficiently sampled. No electrostatically induced particle motion occurs when 

either the particle charge or electric field during sampling is zero. When both the sampler 

walls and particles are both highly charged, external force on the particles from electrical field 

is much greater than that caused by gravity, inertia, diffusion or other mechanisms.

Samplers can achieve a high charge level when they are electrically insulated from ground and 

are triboelectrically charged (i.e., by contacting or rubbing against other surfaces); this 

sampler charging, as well as particle charging, tends to occur more frequently at low (<20% 

RH) humidity levels. Certain plastic materials, such as polycarbonate, 

polytetrafluoroethylene, polyvinyl chloride (PVC), and polystyrene readily retain high charge 

levels; others, such as Tygon® or conductive silicone rubber tubing retain relatively little

charge [Liu et al. 1985]. The PVC/polystyrene copolymer used in the 37-mm closed-face 

cassette is an excellent electrical insulator and can retain high charge levels on its surface. 

These charges can be incorporated in the bulk plastic during manufacture or accumulated on 

the surface by handling or contact with other objects; the charge levels and polarity are highly 

localized and variable. Such samplers can exhibit particle losses to the internal walls of the 

cassette and negative sampling biases [Baron and Deye 1990a]. Non-conductive plastic 

asbestos samplers were shown to produce large negative biases and variable results [Baron and 

Deye 1990a,b; Baron et al. 1994].

Conductive samplers have demonstrably lower losses when sampling charged particles. Metal 

samplers obviously have high conductivity. Samples collected using nylon cyclones were 

shown to exhibit higher variability [Almich and Carson 1974; Briant and Moss 1984] and 

negative biases [Briant and Moss 1984] when sampling charged dusts. However, the degree of 

conductivity required is not high; as long as charges can move over the sampler surface and 

reach equilibrium in seconds, the effect of charges transferred to the sampler is likely to be 

minimized. Materials with this low level conductivity (surface resistivity <108 ohms/square) 

are often termed “static-dissipative.” Graphite-loaded plastics were developed that have 

adequate conductivity to distribute charges over the surface of the cassette (e.g., the 25-mm 

asbestos sampler). A simple test to ensure adequate conductivity of these samplers can be 

performed by attaching a good quality multimeter at any two points on the sampler surface. 

Resistance readings in the range of tens of megohms or less indicate sufficient conductivity for 

sampling purposes.
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Factors Affecting Aerosol Sampling 

Some metals are coated with a thin, non-conductive layer, e.g., anodized aluminum. These 

coatings may retain a surface charge, but this charge will induce an opposite charge in the 

conductive layer beneath the surface, effectively canceling out the field produced by the 

surface charge. Recent measurements at NIOSH using a non-contacting electrostatic 

voltmeter (Model 300, Trek Inc., Medina NY) indicated that no significant external field 

(<50 volts) could be produced near an anodized surface by rubbing the surface with various 

plastics or other materials. Plastic or cellulose-based materials rubbed in a similar manner 

produced electrostatic potentials measured in the hundreds to thousands of volts. Thus, 

metals with a thin, insulating surface layer are not likely to produce significant external fields 

that would affect aerosol sampling.

Digestible cassette inserts or capsules, consisting of a static-dissipative plastic dome directly 

sealed to a filter, have been developed for metals analysis that substantially reduce the error 

associated with wall losses [Ashley et al. 2013].

A further electrostatic problem not specifically associated with the cassette is the use of filters 

made of highly nonconductive materials, such as PVC, polytetrafluoroethylene, or 

polycarbonate. In addition to having desirable chemical properties, these filters have the 

advantage of not absorbing water from atmosphere, leading to improved weight stability 

[Lowrey and Tillery 1979; Bowman et al. 1984; NIOSH 1994]. However, these filters can retain 

a high electrostatic charge level, resulting in non-uniform particle deposition and even 

repulsion of particles from the filter surface. Such filters are also more difficult to handle 

during weighing because of charge effects. Even filters that are normally more conductive, 

such as cellulose-based filters, can become non-conductive and exhibit non-uniform particle 

deposition and particle losses at very low humidity levels (<10% RH) [Chen and Baron 1995]. 

A treatment was developed to make filters more conductive without significantly affecting 

weighing accuracy or moisture absorption [Mark 1974]. In one study, it was found that 

applying this treatment to the filter decreased particle losses from 14% to 2% [Blackford et al. 

1985]. Anti-static sprays are available that leave a temporary static-dissipative coating on 

surfaces.

6 Sampler deposition uniformity

Some analytical methods require that sampled particles be deposited uniformly on the filter 

surface. For instance, asbestos fiber analysis by microscopy requires uniform deposition of 

fibers on the filter for accurate results. Direct silica analysis of collected filter samples also is 

improved with uniform particle deposition. Classifiers using inertial or gravitational forces 

tend to stratify the aerosol stream. A small, high velocity inlet in a sampler, such as the 4-mm 

opening in the 37-mm closed-face cassette, can also result in the larger particles being 
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Factors Affecting Aerosol Sampling 

deposited in a small central area on the filter. Even sampling at high flow rates through more 

open inlets can cause a non-uniform deposit [Feigley et al. 1992]. This results in particle 

deposits that vary in uniformity as a function of particle size. Such deposition patterns are 

visible when sampling colored particles [Sass-Kortsak et al. 1993]. Open-pore foam classifiers 

may improve the uniformity of particle deposits on the filter, but have not been thoroughly 

evaluated [Aitken et al. 1993; Vincent et al. 1993]. Careful design of classifiers to ensure 

mixing of the aerosol prior to deposit on the filter may result in adequate uniformity [Fang 

and Lippmann 1995]. Even inhalable samplers or samplers that have no classifier may be 

prone to non-uniform deposits under certain conditions of sampler orientation relative to 

gravitational settling, orientation relative to external winds, or when sampling charged 

particles [Baron and Deye 1990b; Liu et al. 1985; Baron et al. 1994; Chen and Baron 1995]. 

Flaring the inlet of such a sampler, as in the commercial “bell-mouth cowl,” (Figure 2F, 

Envirometrics, Charleston, SC), is one approach to improving sample uniformity under 

anisokinetic conditions [Feigley et al. 1992]. In another study, a sampler having an inlet screen 

(button sampler, Figure 2D, SKC, Inc. Eighty Four, PA) exhibited improved filter deposit 

uniformity when compared to a closed-face cassette [Hauck et al. 1996].

The filters in some samplers require support to prevent tearing or distortion of the filter. The 

support device may cause occlusion of parts of the filter surface, resulting in non-uniform 

particle deposits [Hook et al. 1983].

On occasion, it was observed that poorly-sized tubing connectors protruded into the 37-mm 

cassette and touched the filter surface. This caused all the airflow to pass through the filter 

adjacent to the small area of the connector opening. When undetected, this caused low 

sampling efficiency and pump failure because of the high pressure drop.

7 Sampler wall losses

Particle deposits on internal surfaces (i.e., wall losses) of the 37-mm closed-face cassette for 

several hundred field measurements were found to be large and highly variable (2 - 100% of 

dust collected in the cassette) [Demange et al. 1990]. Another study found only 22% of the 

dust on the filter, 65% on the upstream portion of the cassette, and 22% downstream of the 

filter [Puskar et al. 1991]. In a study of an in-line cassette of similar shape, it was found that 

the internal wall deposition of particles could be largely eliminated by: (a) making the cassette 

conductive, (b) creating an aerodynamically smooth surface having no corners for eddies to 

form, and (c) decreasing the diameter of the filtration area so that dust does not deposit on the 

filter adjacent to the upstream walls of the cassette [Blackford et al. 1985]. By incorporating 

these three corrective measures, the wall losses in the latter cassette were reduced from 

25-30% to 5%. These losses appear to be caused by a combination of electrostatic, inertial, 

gravitational and diffusion mechanisms.
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Factors Affecting Aerosol Sampling 

Another solution to the problem of not capturing 100% of the sampled particles on the filter is 

to use an internal capsule sealed to the filter. All the particles collected in the combined filter 

capsule are analyzed. The air stream entering the cassette is surrounded by the cartridge and 

any deposition on the walls of the capsule is retained for analysis. The IOM sampler for 

inhalable dust uses this approach by having a cartridge form the inlet of the sampler (Figure 

2B). This approach also has been used in the in-line cassette of the coal mine dust personal 

sampling unit (MSA, Pittsburgh, PA) where an aluminum foil cover is crimped onto the filter. 

A similar cartridge was designed for the 37-mm closed-face cassette in measurements of 

pharmaceutical dust [Puskar et al. 1992]. Capsules made of “static dissipative” plastic for 

gravimetric analysis and capsules composed of cellulosic media for elemental analysis are 

commercially available (Accu-CapTM, Omega Specialty Instruments, Chelmsford, MA; 

WoodchekTM, MSA Inc. Pittsburgh, PA). It is important that capsule material be compatible 

with the analytical method. For instance, the plastic material used in the first version of the 

IOM sampler cartridge (Figure 2B, SKC, Eighty Four, PA) was found to absorb milligrams of 

water over periods of days, making the accuracy of gravimetric measurements problematic 

[Smith et al. 1997; Li and Lundgren 1999; Lidén and Bergman 2001]. Demange et al. [2002] 

more recently demonstrated significantly improved agreement between inhalable sampling 

using the IOM sampler and the 37-mm cassette by including all deposits inside the cassette. 

This suggests that the accuracy and precision of the 37-mm cassette can be improved by 

including internal sampler deposits by wiping or washing, or by using an internal capsules 

[Ashley and Harper 2013; Harper and Ashley 2013; Andrews et al. 2016].

8 Collection media and analytical issues

Interaction of particulate filter with the sampled aerosol and the flow can lead to certain 

measurement errors, which are sometimes referred to as filter artifacts. These artifacts can 

include adsorption of gases and vapors from the air stream, the adsorption or desorption of 

moisture by the filter media, evaporation of volatile or semi-volatile organic matter from the 

filter media, and particle bounce from the filter media. All these factors can contribute to the 

measurement bias.

The filter medium should be compatible with the analytical method. Some analytical methods 

require specific filter media or properties. For instance, atomic absorption and inductively 

coupled plasma analyses typically require complete ashing of the filter material; organic 

compound analyses require that no reaction or adsorption of the compounds occur at the 

filter surface. Several studies have dealt with gravimetric stability of different filter types and 

recommended specific procedures [Lowrey and Tillery 1979; Bowman et al. 1984; NIOSH 

1994; ASTM 2000; Chow 1995; Raynor et al. 2011]. Generally, plastic materials that do not 

absorb water (polycarbonate, polyvinyl chloride, polytetrafluoroethylene) are more weight 
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Factors Affecting Aerosol Sampling 

stable than natural cellulose-based materials; uncoated glass fiber filters also may absorb water 

[Lowrey and Tillery 1979; Bowman et al. 1984; NIOSH 1994; ASTM 2000; Chow 1995]. 

Controlled environmental conditions in the weighing room, where temperature and humidity 

are strictly controlled, are essential to reduce measurement bias in gravimetric analysis. In a 

controlled study Tsai et al. [2012] have shown that the mass of MCE membrane filters was less 

stable than that of the glass fiber filters in both controlled and uncontrolled environmental 

conditions. Also, they found that under uncontrolled conditions (where humidity and 

temperatures were not controlled), glass fiber filter mass was much less stable than that of 

PTFE and PVC membrane filters. MCE and glass fiber filters demonstrated significantly 

better stability under controlled conditions; whereas the PVC and especially the PTFE filters 

were found to be extremely stable in both controlled and uncontrolled conditions [Raynor et 

al., 2011]. Other non-aqueous vapors can also adsorb to the filter media or previously 

collected particulate deposits. However, these artifacts are typically important only for semivolatile organic compounds.

It should be noted that weight stable materials also tend to be more highly charged, resulting 

in more charged particle repulsion and deposit non-uniformity. When a plastic (Tyvek®) 

backup pad is crimped into a cartridge together with a filter, the weight stability of the 

cartridge may suffer [Kogut et al. 1999]. To improve the weight stability of coal mine dust 

sampler cartridges, stainless steel backup pads have been used by MSHA. The IOM sampler 

can be purchased with either a plastic or a stainless steel cartridge. The plastic cartridge has 

been shown to exhibit poor weight stability and should not be used for gravimetric analysis 

[Smith et al. 1997; Lidén and Bergmann 2001].

Lawless and Rodes investigated the use of modern electronic balances to determine factors 

affecting the accuracy of gravimetric measurements and found that balance stability, balance 

leveling, vibration and thermal drafts, electrostatic charge reduction, positioning of the filter 

in the balance so that the filter did not hang over the edge of the pan, and temperature and 

humidity control were all important in achieving accurate results [Lawless and Rodes 2001].

Although not strictly a problem with the collection medium, the sampler construction 

material should not outgas vapors that can condense on the collection medium and affect the 

analysis. Early (circa 1970) versions of the closed-face cassette were made of a plastic called 

“tenite,” which resulted in weight gain of the filter over time. This currently does not appear 

to be a problem.

Impactors have been used as samplers and, especially with cascade impactors, the deposits on 

the impaction stages are measured. Particle bounce from the collection substrates on the 

impaction plates can be severe, especially for large solid particles impacting onto a smooth 
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Factors Affecting Aerosol Sampling 

metal plate [Marple and Olsen 2011]. Bounce can also be significant for highly nonspherical, 

low density particles, as was shown recently for nanotubes agglomerates [Birch et al. 2011; 

Maynard et al. 2004; Baron et al. 2008]. Several modifications to the collection substrate are 

available to improve collection efficiency of each stage. These modifications should be 

compatible with the analytical method. Oil can be placed on the collection substrate that wicks 

up over collected particles and continually provides an oiled surface. To avoid the interference 

or contamination of particles by oil, the following scheme has been used in recent studies: a 

pair of cascade impactors was prepared for sampling at a given location. Oiled filters were 

used on every other stage of each impactor. One of the impactors was loaded with oiled filters 

on Stages A and C, while Stages B and D were uncoated and used to sample particles onto the 

substrates. The second impactor contained oiled filters on Stages B and D, while Stages A and 

C were used for particle sampling. This approach provided data for all four stages (plus after 

filter) and minimized bounce to the adjacent lower stage [Baron et al. 2008; Birch et al. 2011]. 

A filter or sintered metal can be used to provide a reservoir for this oil. For gravimetric 

analysis, this oil must have a low vapor pressure and not migrate off the collection substrate. 

Alternatively, grease can be used, but after the surface is coated with collected particles, 

additional particles are more likely to bounce. Filters have also been used as substrates and 

provide a convenient substrate that is somewhat better than a smooth metal surface. Selection 

and use of an impactor is a complex issue and has been described in reviews [Lodge and Chan 

1986; Marple et al. 2001]. Accurate analysis of cascade impactor data can also be difficult and

simple regression analysis of the data may not provide the best answer [Marple et al. 2001; 

Kandlikar and Ramachandran 1999; Cooper 2001].

9 Sampler field comparisons

Direct field comparisons of various samplers are frequently reported in the literature. Because 

of the typical high variability of aerosol concentrations and size distributions in workplaces, it 

is difficult to use these situations for accurate assessment of sampler performance. However, 

field studies are important to verify the overall performance of a sampler and to indicate 

specific sampler issues. The problems with samplers as discussed above can be highlighted 

with some examples observed in field studies.

a. Sampler bias affected by internal deposits 

A study of wood dust sampling comparing collocated free-standing samplers indicated 

that an MSA cassette (having an aluminum cartridge crimped onto the filter) used as a 

sampler gave two times better precision and collected 2.6 to 3.5 times more dust than the 

standard 37-mm closed-face cassette [NACSI 1992]. Both these samplers have the same 

size and shape of inlet. The same study showed that the IOM sampler collected 1.3 times 

more dust than the MSA cassette, indicating that the particle size, inlet shape and inlet 

orientation are important factors in inhalable sampling. Among a number of inhalable 
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Factors Affecting Aerosol Sampling 

samplers in current use around the world, the IOM sampler appears to agree the best with 

the inhalable dust sampling convention [Kenny et al. 1997; Bartley 1998]. Several studies 

have shown that the IOM sampler collects anywhere from slightly more to 3.5 times more 

dust than the closed-face cassette [Vaughan et al. 1990; Burdorf et al. 1994; Notø et al. 

1996; Perrault et al. 1996; Wilsey et al. 1996]. However, Demange et al. showed that for 

several work sites with relatively small MMAD (about 15 μm diameter), measurements 

from 37-mm cassettes agreed well with the IOM results if the deposits on the internal 

surfaces of the cassette were added to the filter analyte [Demange et al. 2002; Harper and 

Demange 2007]. Measurements with the 37-mm cassette are not expected to agree as well 

with the IOM when the particle sizes are much larger because of differences in aspiration

efficiency. However, by including all aspirated material, i.e., all material entering the 

37-mm cassette inlet, in the analysis, agreement with the inhalable convention can be 

improved.

b. Sampler precision affected by internal deposits 

The issue of measurement bias from internal wall deposits in the sampler has gained 

increasing recognition over the past few decades [Ashley and Harper 2013]. Though it is 

now widely recognized that the wall deposits must be included in the analysis, many 

published methods have not been modified. OSHA currently recommends including wall 

deposits. 

In a field study of lead dust, it was found that the measurements from a closed-face 37-mm 

cassette gave a coefficient of variation (CV) of 1.0 and 0.33 when sampling at 2 L/min and 

10 L/min, respectively, while the button sampler gave a CV of 0.10 under the same 

conditions [Hauck et al. 1996]. The button sampler has few internal surfaces for wall 

deposition, suggesting that elimination of this type of loss would improve the precision of 

the 37-mm cassette. Demange et al. [2002] found improved precision for the 37-mm 

cassette data when the wall deposits were added to the analyte.

In spite of some of its drawbacks, the 37-mm cassette is likely to be used for some time. It 

appears that from the standpoint of improving agreement with the inhalable convention 

and improving precision, the inhalable sampler wall losses should be minimized through 

sampler design (or through use of a cartridge such as the AccuCap or in the MSA coal 

mine cassette) or the wall deposits should be included in the analysis.
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Factors Affecting Aerosol Sampling 

10 Conclusions

Clearly, when proper features are incorporated in the sampler design, significant 

improvements in bias and precision can be achieved for some currently used aerosol samplers. 

Several recommendations regarding the application of these samplers are listed:

 Classifiers used to select respirable, thoracic, or other fractions should be evaluated 

based on bias maps obtained from experimental data and combined with particle size 

distributions from workplace measurements to evaluate their applicability.

 Further research and development is needed to improve sampler design to better 

match ACGIH/ISO conventions and reduce inter-laboratory variability in conducting 

aerosol sampling. It is important to report the sampler and flow rate used to allow 

evaluation of potential biases due to sampling. It is also important to account for wall 

losses to reduce overall bias and allow better comparison across different samplers and 

ISO standards.

 The filter cassette and fittings should be air-tight and have no bypass leakage. A 

pneumatic or mechanical press should be used to assemble the cassette and a leak test 

should be used to establish appropriate pressure and proper assembly procedures. See 

[Baron 2002].

 The sampler should be made of conductive or static-dissipative materials.

 Internal deposits in sampling cassettes should be included in the analysis. One 

approach to improving the closed-face cassette measurements is to use an internal 

digestible cassette insert or cartridge that collects all the sampled dust entering the 

cassette. The cartridge must be compatible with the analytical method. Another 

approach is to wipe or wash the internal surfaces of the cassette and add this material 

to the filter analyte.

Disclaimer 

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.
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Factors Affecting Aerosol Sampling 

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NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Sampling and Characterization of 

Bioaerosols 

by William G. Lindsley, Brett J. Green, Francoise M. Blachere, Stephen B. Martin, Brandon 

F. Law, Paul A. Jensen and Millie P. Schafer, NIOSH

1 Introduction BA-2

2 Principles of bioaerosol collection BA-5

3 Devices used for bioaerosol sampling BA-10

4 Considerations for bioaerosol sampling BA-28

5 Selection of bioaerosol samplers BA-36

6 Sample preparation for culturable bioaerosols BA-39

7 Identification of culturable bioaerosols BA-42

8 Enumeration of culturable bioaerosols BA-46

9 Sample analysis methods for non-viable and non-culturable bioaerosols BA-49

10 Limitations of bioaerosol sampling and characterization BA-62

11 Safety considerations BA-65

12 Resources BA-65

13 References BA-66

14 Appendix 1- List of manufacturers/distributors of common bioaerosol 

 samplers and related products BA-101

15 Appendix 2 – Commonly used bioaerosol samplers BA-104
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Sampling and Characterization of Bioaerosols 

1 Introduction

Bioaerosols are airborne particles that originate from biological sources including animals, 

plants, fungi, bacteria, protozoa, and viruses. Examples of bioaerosols encountered in 

occupational environments include plant pollen, algae, fungal spores, bacteria such as 

actinomycetes, droplets produced during coughing and sneezing that may contain bacteria 

and viruses, dust containing insect excreta, animal dander, and fragments derived from each 

of these sources. Bioaerosols are ubiquitous and can be isolated from indoor, outdoor, and 

occupational environments using a variety of methods that either enumerate viable or a 

collection of viable and non-viable bioaerosols. Photomicrographs of example viral, bacterial, 

fungal, and plant bioaerosols are presented in Figure 1.

Bioaerosol monitoring is a rapidly emerging area of industrial hygiene due to the improved 

analysis methods such as polymerase chain reaction (PCR) and the impact that occupational 

exposures may have on worker respiratory health, particularly in microbial contaminated 

environments [Eduard et al. 2012; Environment Agency 2009; Haig et al. 2016; Hung et al. 

2005; Macher 1999; Morey 2007; Nazaroff 2016]. Some human diseases encountered in 

healthcare settings such as measles and tuberculosis can be spread by bioaerosols containing 

infectious microorganisms [Ijaz et al. 2016; Jones and Brosseau 2015]. Soil saprophytic fungi 

such as Coccidioides immitis can be aerosolized during occupational disturbance activities 

and, if inhaled, can result in an acute pulmonary infection [Das et al. 2012; Wilken et al. 2014; 

Wilken et al. 2015]. The measurement of these bioaerosols in industrial hygiene includes the 

measurement of viable (culturable and non-culturable) and nonviable bioaerosols in indoor 

settings (e.g., industrial, office, education, and residential buildings), industrial facilities (e.g., 

biotechnology, composting, waste disposal, manufacturing, textile, and food processing), and 

outdoor environments (e.g., farms, feed lots, and general air quality). Monitoring for 

bioaerosols in the occupational environment is one of the many tools the industrial hygienist 

uses in the assessment of indoor air quality, infectious disease outbreaks, agricultural 

exposures, and industrial health.

Bioaerosol monitoring may be appropriate during workplace health and exposure 

assessments, epidemiological investigations, research studies, or in situations deemed 

appropriate by an occupational physician or immunologist. Sampling can also be used to 

evaluate occupational environments before and after mitigation of microbial contaminants. 

When investigating bioaerosols as a possible source of workplace exposures and health issues, 

bioaerosol sampling should be part of an integrated assessment of work conditions. This 

should also include examining heating, ventilation and air conditioning (HVAC) systems; 

checking for water infiltration and moisture control; evaluating microbial contamination in 

evaporative cooling systems, metal working fluids, and waste water; evaluating possible 
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Sampling and Characterization of Bioaerosols 

 internal and external sources of bioaerosols; and other measures [Macher 1999]. In general, if 

visible growth or contamination (microbial growth on floors, walls, or ceilings, or in the 

HVAC system) is observed, this normally should be mitigated first before indoor bioaerosol 

sampling is conducted. If personnel remain symptomatic after remediation, air sampling may 

be appropriate, but the industrial hygienist should be aware that false negative results are 

possible and should be interpreted with caution.

The industrial hygienist has a variety of tools and methodologies available to conduct an 

environmental survey [ASTM 2014a; Flannigan et al. 2011; Hung et al. 2005]. However, many 

of these approaches have lacked standardization and this has made the interpretation and 

comparison between studies challenging [Flannigan et al. 2011]. In 2005, the American 

Industrial Hygiene Association (AIHA) published the second edition of the Field Guide for 

the Determination of Biological Contaminants in Environmental Samples [Hung et al. 2005]. 

This reference provides the industrial hygienist access to the most up to date methods to 

detect and quantify bioaerosols in the environment, and covers methods of how to conduct a 

survey, sample bioaerosols, and interpret the collected data [Hung et al. 2005]. Similarly, other 

reference sources have been published by Flannigan et al. [2011] and the American 

Conference of Governmental Industrial Hygienists (ACGIH) [Macher 1999] that extensively 

outline available methods to analyze collected bioaerosols as well as strategies to conduct an 

environmental survey. ASTM International has issued a wide range of standards on indoor air 

quality, including assessment of fungal growth and collection of bioaerosols and a guide to 

developing an air sampling strategy [ASTM 2009; ASTM 2014a; ASTM 2014b; ASTM 2014d]. 

The European Committee for Standardization has also published standards on sampling for 

bioaerosols and related topics [CEN 2000; CEN 2003; CEN 2004]. The sections presented 

below provide a very broad overview of the viable and non-viable methods available to detect 

bioaerosol sources that are described in the references listed above. 
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Sampling and Characterization of Bioaerosols 

 

Figure 1: Photomicrographs of acellular, prokaryotic and eukaryotic microorganisms that can 

be encountered in occupational or industrial environments. (A) Transmission electron 

micrograph of Influenza/flu (H1N1) virus particles (Photo courtesy of National Institute of 

Allergy and Infectious Diseases; CDC Public Health Image Library (PHIL) ID#: 18156); (B) 

Scanning electron micrograph of bacilli derived from the Gram-negative bacteria, Legionella 

pneumophila (Photo courtesy of National Institute of Allergy and Infectious Diseases; CDC 

Public Health Image Library (PHIL) ID#: 11150); (C) Scanning electron micrograph of 

Aspergillus species reproductive structures including chains of asexual spores (Photo courtesy 

of CDC/ Robert Simmons; CDC Public Health Image Library (PHIL) ID#: 13367); and (D) 

Scanning electron micrograph of tricolpate pollen derived from the angiosperm plant species, 

Oenothera fruticosa (Photo courtesy of CDC/ Janice Carr, Betsy Crane; CDC Public Health 

Image Library (PHIL) ID#: 8729). The CDC Public Health Image Library at 

http://phil.cdc.gov/Phil/home.asp has thousands of health-related images available to the 

public free of charge.
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Sampling and Characterization of Bioaerosols 

 

2 Principles of bioaerosol collection

a. Aerodynamic diameter

The aerodynamic diameter of an airborne particle (usually written as “da” or “dae”) is the 

single most important parameter that determines how the particle will behave in the air, 

including how long it will stay airborne and where it will deposit in the respiratory system 

if inhaled. If a particle is falling in still air, it will reach an equilibrium velocity where the 

gravitational force pulling it downward is balanced by the drag force on its surface. This 

velocity is called the terminal settling velocity, and it depends upon the size, shape and 

density of the particle. The aerodynamic diameter of a particle is defined as the diameter of 

a sphere with unit density (that is, a density of 1 g/cm3) that has the same terminal settling 

velocity as the particle. Consider, for example, the irregularly-shaped fungal fragment 

shown in Figure 2. Suppose this particle has a terminal settling velocity of 0.05 cm/sec. 

This is the same settling velocity as that of a spherical particle with a unit density that has a 

diameter of 4 μm. Thus, the fungal fragment is said to have an aerodynamic diameter of 4 

μm. Similarly, a different particle with a terminal settling velocity of 1.21 cm/sec has an 

aerodynamic diameter of 20 μm, since a 20 μm unit density sphere settles at that rate. It is 

important to note that the aerodynamic diameter may be very different from the physical 

size of a particle. A very dense and compact particle may have an aerodynamic diameter 

much larger than its actual dimensions, while a very light particle or one with fibrous 

branches may have an aerodynamic diameter that is much smaller than its physical size. It 

is possible for two particles to have very different shapes and physical sizes, but have the 

same aerodynamic diameter. Conversely, two particles may have similar physical sizes, but 

have very different aerodynamic diameters. A more detailed discussion of the 

aerodynamic diameter can be found in Hinds [1999] and Vincent [2007].
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Sampling and Characterization of Bioaerosols 

 

Figure 2: Aerodynamic diameter of an aerosol particle. In this case, the fungal fragment on 

the left is said to have an aerodynamic diameter of 4 μm, since it falls at the same terminal 

settling velocity as a 4 μm sphere with a unit density.

Aerodynamic diameter is used in aerosol science because particles with the same 

aerodynamic diameter tend to move and be collected in the same ways. For example, two 

particles with the same aerodynamic diameter will have the same likelihood of being 

collected by an impaction aerosol sampler even if they have different physical and 

morphological characteristics. For this reason, the performance of aerosol collection 

devices is usually described by giving the aerodynamic diameter of the particles that will be 

collected.

b. Collection efficiency and cut-off diameter

The collection efficiency of an aerosol sampler is the fraction of the aerosol particles of a 

particular aerodynamic diameter that will be collected by the sampler. For example, if 95% 

of the airborne particles with a 2 μm aerodynamic diameter that enter the sampler are 

deposited in the collection fluid or on the collection surface, then the sampler is said to 

have a 95% collection efficiency for 2 μm particles.

Most commonly-used aerosol filters have a high collection efficiency for particles of all 

sizes [NIOSH 2016b]. However, impactors, cyclones and impingers use the inertia of 

airborne particles to separate them from the air stream, and thus they have a high 

collection efficiency for particles with larger aerodynamic diameters and a low collection 

efficiency for smaller ones (Figure 3) [Hering 2001; Hinds 1999; Marple and Olson 2011]. 

These devices are said to have a “cut-off diameter”; that is, particles with an aerodynamic 
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Sampling and Characterization of Bioaerosols 

 diameter larger than the cut-off diameter are collected while particles with an aerodynamic 

diameter less than the cut-off diameter are not collected and pass through the device. A 

perfect collection device would have a 100% collection efficiency for particles larger than 

the cut-off diameter and 0% for smaller particles. In practice, this is not the case: the 

collection efficiency curve for an inertia-based sampler looks like the example curve shown 

in Figure 3. The aerodynamic diameter at which the collection efficiency is 50% is defined 

as the cut-off diameter (usually written as d50). A device with a more abrupt transition 

from 100% to 0% collection efficiency (that is, closer to the ideal device) is said to have a 

sharp cut-off.

For a given inertial collection device, the 50% cut-off diameter depends upon the air 

flowrate through the device. Increasing the flowrate will decrease the d50 and shift the 

collection efficiency curve to the left, while decreasing the flowrate will increase the d50 and 

shift the collection efficiency curve to the right. For example, the first stage of the NIOSH 

two-stage cyclone aerosol sampler has a d50 of 4.9 μm at 2 liters/minute of air flow, 4.1 μm 

at 3.5 liters/minute, and 2.1 μm at 10 liters/minute [Blachere et al. 2009]. For this reason, it 

is important to check the air flowrate before aerosol sampling and control it during 

sampling so that the particles are correctly segregated by size.

Figure 3: Example collection efficiency curve for an inertia-based aerosol sampler. Note 

that the collection efficiency is high for particles with large aerodynamic diameters and 

low for small particles. In this example, the 50% cut-off diameter (d50) for this device is 1 

μm.
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Sampling and Characterization of Bioaerosols 

 

c. Size-selective bioaerosol sampling in industrial hygiene

Size-selective bioaerosol sampling may be done for several reasons. Since the settling

velocity of aerosol particles is determined by the aerodynamic diameter, knowing the size 

distribution of an aerosol helps in predicting how long the particles are likely to remain 

airborne and how far they can travel. In health care settings, for example, various medical 

procedures can produce a spray of droplets containing infectious microorganisms. Large 

droplets tend to fall onto surfaces fairly close to the source, while smaller droplets can 

remain airborne and carry pathogens many feet away from a patient [Davies et al. 2009; 

Jones and Brosseau 2015]. Another application of size-selection is to isolate different types 

of bioaerosol particles, such as separating fungal fragments from intact fungal spores 

[Adhikari et al. 2013; Seo et al. 2014].

Size-selective sampling is most commonly used to help understand the potential health 

effects of bioaerosol particles, which often depend upon where the particles are deposited 

in the respiratory tract. In general, larger bioaerosol particles tend to deposit higher in the 

respiratory tract (that is, in the nasal or oral cavities or larger airways), while smaller 

particles are able to travel deeper into the lungs to the smaller airways [Hinds 1999; 

Vincent 2005]. Some pathogens such as Mycobacterium spp., Bacillus spp., and Aspergillus 

spp. are thought to be more likely to cause a pulmonary infection if they reach the deeper 

airways, and the response to bioaerosols containing immunogenic material such as 

endotoxins or fungal antigens may also vary depending upon the site of deposition. For 

this reason, size-selective sampling is often used in industrial hygiene to better understand 

the potential risks that workplace bioaerosols present.

The American Conference of Governmental Industrial Hygienists (ACGIH), the 

International Organization for Standardization (ISO) and the European Standardization 

Committee (CEN) have defined three particle collection efficiency curves for aerosol 

samplers used to conduct size-selective aerosol sampling (Figure 4) [ACGIH 2001; ISO 

2012; Vincent 2005]. The idea is that an aerosol sampler that conforms to one of the three 

criteria will collect aerosol particles in a way that approximates the fraction of particles 

that will reach different parts of the respiratory tract. These criteria are not specific to 

bioaerosols, but rather are applied to all types of aerosol particles.
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Figure 4: ACGIH/ISO sampling criteria for the inhalable, thoracic and respirable fractions 

of aerosol particles. The inhalable fraction contains all of the particles that are inhalable, 

which includes the particles in the thoracic and respirable fraction. Similarly, the thoracic 

fraction includes the particles in the respirable fraction. The 50% cut-off diameters are 100 

μm for the inhalable fraction, 10 μm for the thoracic fraction, and 4 μm for the respirable 

fraction [ACGIH 2001; ISO 2012; Vincent 2005].

A sampler that collects the inhalable fraction accumulates the fraction of aerosol particles 

of each size that would be expected to be drawn into the nose or mouth during normal 

breathing. This includes larger particles that would be expected to be deposited in the 

nasal or oral cavities as well as smaller particles that can be conveyed to the lower airways. 

An aerosol sampler that conforms to the inhalable sampling criteria collects 50% of the 

100 μm particles, 77% of the 10 μm particles, and 97% of the 1 μm particles in the ambient 

aerosol. The inhalable fraction is lower for larger particles because the greater inertia of 

these particles means they are less likely to be pulled into the body during inhalation. 

The thoracic fraction includes aerosol particles that are likely to travel into the trachea and 

bronchi. An aerosol sampler that conforms to the thoracic sampling criteria will collect 

50% of the 10 μm particles and 97% of the 1 μm particles in the ambient aerosol. This 

fraction includes fewer large particles because these particles tend to be removed from the 

airstream by the head airways.
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 The respirable fraction includes aerosol particles that are able to reach the deepest airways, 

which are the respiratory bronchioles and the alveoli. An aerosol sampler that conforms to 

the respirable sampling criteria will collect 50% of the 4 μm particles, 97% of the 1 μm 

particles, and 99% of the 0.3 μm particles in the ambient aerosol. The respiratory 

bronchioles and the alveoli are of particular concern because these airways do not have 

cilia. Non-soluble particles that land in the nasopharyngeal region or upper airways tend 

to collect in the airway mucus and are removed from the respiratory tract by the cilia 

relatively quickly. However, particles that deposit in the alveoli and respiratory 

bronchioles can remain in the lungs for longer durations (in some cases, for life) unless 

they can be broken down or removed by migrating pulmonary macrophages. This fraction 

includes only the smallest particles because the larger particles are removed from the 

airstream by the head and thoracic airways.

It should be noted that, even though larger bioaerosol particles will tend to deposit in the 

upper airways and be cleared more quickly, they can still trigger an allergic/inflammatory 

response in susceptible individuals. Particles containing viable pathogens also commonly 

cause infections after being deposited in the upper airways.

When describing size-selective sampling, particles are often said to “penetrate” to a 

particular region of the respiratory tract. This does not mean penetrate in the sense of 

entering the tissue, but rather simply being present in the air stream flowing into that 

region, as compared to particles which were deposited before reaching a particular 

location. For example, an aerosol particle that is able to remain in the air stream and reach 

the lung alveoli is said to have penetrated to the alveolar region, even if it does not 

necessarily deposit there. This is the same context as with filtration, where a particle is said 

to penetrate a filter if it flows through the filter material and remains in the air stream. It 

also should be noted that the ACGIH/ISO criteria give an approximation of the fraction of 

aerosol particles that can penetrate to different regions of the respiratory tract. However, 

they do not indicate what fraction of the aerosol particles will actually deposit in the 

airways and what fraction will be exhaled. The lung deposition of aerosol particles is 

complex and depends upon many factors. More information about this topic can be found 

in Hinds [1999] and Vincent [2005; 2007].

3 Devices used for bioaerosol sampling

Most aerosol sampling devices involve techniques that separate particles from the air stream 

and collect them in or on a preselected medium. Impactors, filters, impingers and cyclones are 

four common sampling techniques used to separate and collect bioaerosols [Haig et al. 2016; 

Macher et al. 1995; Reponen et al. 2011b; Willeke and Macher 1999]. A few systems that use 

electrostatic precipitation or condensation-based collection are also available [Haig et al. 
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Sampling and Characterization of Bioaerosols 

 2016], and some real-time bioaerosol monitoring systems are available that do not require 

that the bioaerosol particles be isolated before analysis. Below are some specific types of 

bioaerosol sampling devices employed by industrial hygienists.

a. Filters

Aerosol filters are commonly used to collect bioaerosol particles because of their simplicity 

and low cost. Filter-based sampling is particularly useful for personal bioaerosol sampling 

because filter-based collectors are small and lightweight and work well with personal 

sampling pumps. Filters can be preceded by a size-selective inlet, such as a cyclone or 

impactor, to remove larger particles and provide size-classification of the bioaerosol 

particles. Most aerosol filter media can be classified as fibrous, membrane, or capillary 

pore (also called straight-through pore) [Raynor et al. 2011]. Fibrous filters are usually 

made of a deep mesh of glass fibers. Membrane filters are manufactured in a variety of 

pore sizes from polymers such as cellulose ester, polyvinyl chloride, or 

polytetrafluoroethylene (PTFE). Capillary pore filters are made of polycarbonate. The 

choice of a filter medium depends on the contaminant of interest and the requirements of 

the analytical technique. For gravimetric analysis, non-hygroscopic materials such as glass 

fibers, silver, or polyvinyl chloride membranes are selected because their masses are less 

affected by changes in humidity. For analysis by microscopy, cellulose ester or 

polycarbonate membranes are common choices because cellulose ester membranes can be 

rendered transparent for easier visualization, while polycarbonate filters have a smooth 

collection surface that works well with light or electron microscopy. Samples also can be 

eluted from cellulose ester and polycarbonate filters, but in some cases the recovery 

efficiency can be low [Eduard et al. 1990; Rule et al. 2007]. Samples to be cultured can be 

collected on gelatin filters, and the filters can then be dissolved in water and spread on 

culture plates, dissolved in growth media, or placed directly on culture plates and allowed 

to melt. Gelatin filters are fragile and can crack or melt in use. For analysis using 

immunological assays or polymerase chain reaction (PCR), PTFE filters are a common 

choice because they do not interfere with the assays and because samples can be readily 

eluted from them.

Filters are frequently described or specified using the term “pore size” or “equivalent pore 

diameter”. It is important to note that the filter pore size does NOT indicate the minimum 

particle size that will be collected by the filter; in fact, aerosol filters generally will collect 

particles much smaller than the nominal pore size. The mechanisms by which aerosol 

filters work and the role of pore size in selecting filters is described is more detail 

elsewhere [NIOSH 2016b].
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 Aerosol filters are usually supplied as disks of 25, 37 or 47-mm diameter. Because the flow 

resistance (often called the pressure drop) of a filter increases with the air velocity through 

the filter, the use of a larger filter results in a lower flow resistance for a given volumetric 

flow rate. On the other hand, the use of a smaller filter concentrates the deposit of the 

contaminant onto a smaller total area, thus increasing the density of particles per unit area 

of filter. This may be helpful for direct microscopic examination of low concentrations of 

organisms, and reduces the amount of elution media needed for immunological or PCRbased assays. In areas of high concentration, the microorganisms may have to be eluted, 

diluted, and then refiltered for microscopic analysis. Breuer [2012] reported on the flow 

resistance of common aerosol filters and its relationship to sampling pump selection. Soo 

et al. [2016] measured the filtration characteristics and flow resistance of a variety of 

commonly-used aerosol filters.

In the USA, the most common method of aerosol sampling with filters is to place the 

filters in disposable two-piece or three-piece plastic filter cassettes with a support pad to 

add rigidity. The three-piece cassette may be used either in open- or closed-face modes. 

Open-face sampling is performed by removing the end plug and the plastic cover from the 

three-piece cassette and is used when the particulate matter must be uniformly deposited 

(i.e., for microscopic analysis). If a three-piece cassette is used in the open-face 

arrangement, the plastic cover is retained to protect the filter after sampling is concluded. 

It should be noted that the aspiration efficiencies of open-face and closed-face filter 

cassettes are reported to be somewhat different [Beaulieu et al. 1980; Kenny et al. 1997]. 

In addition to collecting on the filter, aerosol particles (especially large particles) may 

collect on the internal walls of the filter cassette. Depending upon the purpose of the 

collection, wall-deposited material may need to be included in the analysis. This can be 

done by using a filter with an attached capsule or by washing or wiping the internal 

surfaces of the cassette [Ashley and Harper 2013]. 

It is important to verify that the filter cassette and fittings are air-tight and have no bypass 

leakage around the filter. Cassettes should not be hand-assembled; they should be pressed 

together with a mechanical or hydraulic press. All plastic cassettes should be securely 

assembled and sealed with a cellulose shrink band or tape around the seams of the cassette 

to prevent external air leakage. The cassettes should be made of conductive or static 

dissipative materials to avoid losses due to electrostatic effects. More information on using 

filter cassettes for aerosol sampling can be found elsewhere [NIOSH 2003a; NIOSH

2016a].
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b. Impactors 

An impactor consists of a series of nozzles (circular- or slot-shaped) and an impaction 

surface [Hering 2001; Marple and Olson 2011; Marple and Willeke 1976]. Air is drawn 

into the impactor using a vacuum pump, and the air stream flows through the nozzles and 

toward the impaction surface, where particles are separated from the air stream by their 

inertia (Figure 5). Larger particles collect on the impaction surface, while small particles 

that do not impact follow the air stream. The impaction surface typically consists of a 

greased plate or tape, filter material, or growth media (agar) contained in Petri dishes. In 

some applications, impactors are not used as collection devices themselves, but rather to 

remove particles above a certain size before collection or characterization of the 

downstream aerosol.

Figure 5: Impaction. As the air stream exits the impactor nozzle, it quickly changes 

direction as shown by the arrows. Smaller particles such as those on the left flow with the 

air stream and are not collected. Larger particles cannot change direction as quickly due to 

their higher inertia and collide with the collection surface, where they accumulate.

A cascade impactor consists of a stack of impaction stages: each stage consists of one or 

more nozzles and a target or substrate. The nozzles may take the form of holes or slots. 

Each succeeding stage has smaller nozzles and thus collects smaller particles (that is, each 

succeeding stage has a smaller cut-off diameter). A filter may be used after the final 

impaction stage to collect any particles smaller than the final cut-off diameter. If the 

substrate is a greased plate or filter media, it may be weighed to determine the collected 

mass, or it may be washed and the wash solution analyzed. If the substrate is growth media 

in culture plates, they may be incubated and examined for microbial growth.
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 The most commonly used impactor for sampling airborne culturable bacteria and fungi is 

the Andersen impactor, which uses from one to six impactor stages containing Petri plates 

as seen in Figure 6 [Andersen 1958]. Since the bioaerosol particles impact directly onto the 

growth media, the samplers can be directly transferred to an incubator and observed for 

microbial growth. However, this method depends upon collecting viable microorganisms 

that are capable of growth on the specific nutrient media. 

Glass Petri plates are recommended for use with the Andersen impactor; plastic culture 

plates are often used, but this can result in loss of aerosol material due to electrostatic 

surface charges in the plastic [Andersen 1958; Kuo 2015]. NIOSH Method 0800 describes 

how to collect culturable airborne fungi and bacteria in buildings using an Andersen 

cascade impactor [NIOSH 2003b].

Figure 6: Schematic of a 6-stage Andersen cascade impactor [Andersen 1958]. Each stage 

contains a Petri plate (green) filled with nutrient agar (brown). The stages have 

progressively smaller nozzles, which create higher particle impaction velocities onto the 

agar. The aerosol particles (red) flow from the top into the first stage, where particles with 

aerodynamic diameters larger than 7 μm impact the agar. The remaining particles flow to 

the second stage, where particles with aerodynamic diameters between 7 μm and 4.7 μm 

are collected, and so on for the rest of the stages.
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One significant advantage of the Andersen impactor is that samples can be collected

directly onto culture plates and transferred to an incubator, which simplifies handling and

eliminates some losses that can occur in processing. However, there are also several

limitations. In low concentration environments, sampling time is limited to approximately

20 minutes to avoid drying the agar. The high flow rate (28.3 liters/minute) makes the 

sampler unsuitable for high concentration environments such as some agricultural sites 

(i.e. animal facilities) where a 1 minute sample may overwhelm the plates.

When using the Andersen impactor, it is also necessary to correct for “coincidence error” 

using a positive-hole correction factor. This occurs because it is possible for multiple

particles, each containing one or more organisms, to pass through a particular hole during

sampling and impact onto the growth medium, with one or more bacterial or fungal

colonies forming at the same impaction sites. The colonies formed by the multiple 

particles can then be inaccurately counted as a single colony. As the number of organismcontaining particles deposited onto the growth medium increases, the probability that the 

next organism-containing particle will impact an "occupied" hole increases. For example, if

75% of the holes have received at least one particle, the chance that the next particle will

impact a "clean" hole is one in four (25%). To account for this, a probability-based 

coincidence correction factor needs to be applied to the results for each impactor stage.

The basic formula for the coincidence correction is as follows [Andersen 1958; Macher

1989]: 

Where:

N = the total number of holes in the impactor stage

r = the number of colonies observed on the culture plate

Pr = the estimated culturable particle count

Andersen impactors have from one to 400 holes per stage. Macher [1989], Willeke and

Macher [1999] and Andersen [1958] provide tables of positive-hole correction factors.

Investigators often employ stationary cascade impactors either as the primary collection

mechanism, or as a preclassifier (for example, to remove nonrespirable particles from the

sampled air stream). Marple and Willeke [1976] have reported that high velocity, inlet 

losses, interstage losses, and particle reentrainment affect the performance characteristics

of an impactor. Particles larger than the cut-off diameter may bounce after impacting the

collection surface and travel to subsequent impaction stages. This is particularly a problem

with dry solid collection surfaces; for this reason, solid collection surfaces are usually

greased or oiled [Hering 2001]. Fungal spores have been shown to be prone to deaggregation and bounce when collected with an impactor, which can cause the spores to be

collected on stages with smaller cut-off diameters. This can make the spore aggregates

appear to have smaller aerodynamic diameters than is actually the case [Trunov et al.
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Sampling and Characterization of Bioaerosols 

 2001]. Although personal cascade impactors are available, these devices are not as widely 

used in personal sampling for bioaerosols as are filters [Macher and Hansson 1987].

The slit-to-agar impactor is a type of impactor in which the aerosol particles are deposited 

on a Petri plate that slowly rotates. The rotation of the plate means that particles which are 

collected at different times deposit in different locations, and thus provides an indication 

of changes in the bioaerosol concentration over time [Ho et al. 2005; Jensen et al. 1992; 

Smid et al. 1989; USP 1997]. Examples of slit-to-agar samplers include the Dycor Slit 

Sampler from Dycor and the Air Trace Environmental Slit-to-Agar Sampler from Particle 

Measuring Systems.

The Hirst/Burkard spore trap has been widely used to collect outdoor aerospora. It was 

first described by Hirst [1952] and consists of a unit that houses a vacuum pump and 

rotating drum that is lined with polyester tape. The drum rotates at 2 mm per hour and is 

continuously run for seven days. Bioaerosols pass through an orifice on the sampler and 

particles impact on the tape. Following the seven-day sampling interval, the tape is 

removed and cut in 48 mm intervals that correspond to individual sampling days. 

Bioaerosols deposited on the tape are stained and then resolved, identified, and quantified 

using bright field microscopy.

 

[Tovey et al. 2016] developed a personal aerosol sampler with a rotating surface that allows 

time-resolved collection of aerosol particles onto an electret strip or an adhesive film. They 

used the sampler to study personal exposures to dust mite allergens over time.

A novel example of an impaction-based personal bioaerosol sampler is the intranasal air 

sampler fabricated by Graham et al. [2000], which fits within the intranasal cavity of the 

subject. Bioaerosols enter the nasal cavity following inhalation and pass through slits 

where particles are deposited by impaction on either an adhesive backed tape or collection 

cup lined with silicon grease. This impaction sampler has been utilized in a number of 

studies that have evaluated exposure to indoor and occupationally relevant aeroallergen 

sources [Gore et al. 2002; Mitakakis et al. 2000; Renstrom et al. 2002]. 

Other impaction-based approaches have also been used in the assessment of outdoor 

bioaerosols, including the Rotorod, Air-o-cell and Allergenco samplers [Frenz 1999; Lee et 

al. 2004a; Pityn and Anderson 2013; Portnoy et al. 2000]. ASTM Standards D7391 and 

D7788 discuss the collection and analysis of airborne fungal structures by inertial 

impaction [ASTM 2009; ASTM 2014d].
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c. Cyclones 

A cyclone sampler consists of a circular chamber with the aerosol stream entering through 

one or more tangential nozzles as shown in Figure 7 [Hering 2001]. Like an impactor, a 

cyclone sampler depends upon the inertia of the particle to cause it to deposit on the 

sampler wall as the air stream curves around inside the chamber. Also like an impactor, a 

cyclone sampler has a collection efficiency curve like the one shown in Figure 3, and the 

collection efficiency curve depends upon the flow rate. Cyclones are less prone to particle 

bounce than impactors and can collect larger quantities of material. They also may provide 

a more gentle collection than impactors, which can improve the recovery of viable 

microorganisms. However, cyclones tend to have collection efficiency curves that are less 

sharp than impactors, and it is simpler to design a compact cascade impactor compared to 

a cascade of cyclone samplers.

In industrial hygiene, cyclone aerosol samplers are frequently used in conjunction with a 

filter to conduct size-selective aerosol sampling [Hering 2001]. For example, in NIOSH 

Method 0600, a cyclone is used to remove the non-respirable fraction from the aerosol 

(following the ACGIH/ISO criteria described earlier), and a filter is then used to collect the 

respirable fraction [NIOSH 2003c]. A sampler developed at NIOSH uses two cyclones 

followed by a filter; the first cyclone collects the non-respirable fraction of the particles, the 

second cyclone collects the respirable particles > 1 μm, and the filter collects particles < 1 

μm [Blachere et al. 2009]. The NIOSH cyclone aerosol samplers have been used in 

applications including measurements of airborne viruses in healthcare settings; airborne 

fungi and fungal fragments in residences; airborne dimorphic fungal pathogens such as 

Paracoccidioides brasiliensis in Brazil, and bioaerosols in agricultural operations [Arantes 

et al. 2013; Blachere et al. 2009; Blais Lecours et al. 2012; Kettleson et al. 2013; Lee and Liao 

2014; Lindsley et al. 2010a; Lindsley et al. 2010b; Martin et al. 2015; Seo et al. 2014; Singh 

et al. 2011a; Singh et al. 2011b].
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Figure 7: Cyclone aerosol collection. When the aerosol stream enters the body of the

cyclone through the inlet, the air flow follows the curved interior wall and flows in a spiral 

pattern. If aerosol particles are larger than the cut-off diameter, then the inertia of the 

particles causes them to collide with the wall of the cyclone and accumulate. After 

spiraling downward, the air flow comes up through the center of the cyclone and exits 

through the outlet (called a vortex finder) at the top. The illustration shows a tangential 

inlet reversed-flow cyclone, which is the most common type of cyclone sampler.

d. Impingers 

Many microorganisms can lose their viability if they are collected onto dry solid surfaces 

or filters because of impact damage and desiccation [Cox 1987; Jensen et al. 1992; Macher 

and First 1984; Verreault et al. 2008; Wang et al. 2001]. One way to avoid this is to collect 

culturable bioaerosols in liquids using an impinger [Henningson and Ahlberg 1994; 

Henningson et al. 1988; Lembke et al. 1981; Reponen et al. 2011b; Verreault et al. 2008]. A 

typical impinger is shown in Figure 8. The body of the impinger is filled with a collection 

liquid, and the aerosol stream flows down through a nozzle and enters the liquid at a high 

velocity. The aerosol particles are collected when they collide with the bottom of the 

collection vessel or disperse into the liquid. Impingers often have curved inlets to remove 

larger particles from the air stream before collection. Because impingers are essentially 

another type of inertial collection device, they have a collection efficiency curve and a cutoff diameter like impactors and cyclones. However, the collection efficiency curves tend to 

be less sharp. The high velocity air stream directed into the liquid also creates considerable 

agitation and can produce foaming if the collection liquid contains surfactants. Additives 

to the collection medium such as proteins, antifoam, or antifreeze aid in resuscitation of 

bacterial cells, prevent foaming and loss of the collection fluid, and minimize injury to the 

cells [Chang and Chou 2011; Cown et al. 1957; Dungan and Leytem 2015]. The presence 
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Sampling and Characterization of Bioaerosols 

 of proteins and other additives can also greatly influence the survival of airborne viruses 

during collection by impingers [Ijaz et al. 1985b; Schaffer et al. 1976; Verreault et al. 2008]. 

Water loss over time reduces the liquid level in the impinger and increases the 

concentration of the non-volatile components, which limits the available collection time 

[Lin et al. 1997]. Sample losses due to re-aerosolization and particle deposition inside the 

impinger can be significant [Grinshpun et al. 1997; Han and Mainelis 2012].

Figure 8: Impingement. Bioaerosol particles exit the nozzle of the impinger at high velocity 

and impact the liquid or the bottom surface of the collection vessel. Some types of 

impingers produce air bubbles in the collection media, which can enhance particle 

collection, but can damage some types of microorganisms.

Two common impingers used for bioaerosol sampling are the Greenburg-Smith impinger 

[Greenburg 1932] and the All-Glass Impinger with the nozzle 30 mm above the base of the 

collection vessel, called the AGI-30 [May and Harper 1957]. The Greenberg-Smith and 

AGI-30 samplers operate by drawing aerosols at nominal flow rates of 28.3 and 12.5 

L/min, respectively, through an inlet tube [Macher et al. 1995]. The AGI-30 inlet tube is 

curved to simulate particle collection in the nasal passage [Cox 1987]. Investigators have 

reported problems with low sampling efficiencies and high losses due to particles in the 

collection being re-aerosolized and lost [Grinshpun et al. 1997; Kesavan et al. 2010; Lin et 

al. 1997].

When the AGI-30 is used to recover total airborne organisms from the environment, the 

curved inlet tube is washed with a known amount of collecting fluid after sampling 

because larger particles (i.e., over 15 μm) are collected on the tube wall by inertial force. 

After sampling for the appropriate amount of time, 10 mL of the full-strength collection 
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 fluid is filtered through a 0.45-μm pore size membrane filter. Serial dilutions of the 

remaining collection fluid are handled similarly [Greenberg et al. 1992]. The membrane 

filters are placed in sterile plastic petri plates filled with the appropriate medium and 

incubated for later identification and enumeration.

e. Wetted-surface bioaerosol samplers

Several types of bioaerosol sampling devices have been developed in which the aerosol 

stream impacts onto a wetted surface or onto the wall of a cyclone wetted with collection 

media [Kesavan and Sagripanti 2015; Kesavan et al. 2011]. These systems largely avoid the 

bubbling and agitation associated with conventional impingers, which may be detrimental 

to some microorganisms [Lin et al. 2000], and can provide sharper collection efficiency 

curves. One of the simplest examples of a wetted-surface sampler is the SKC BioSampler 

[Lin et al. 2000; Willeke et al. 1998]. It is similar to an AGI-30, except that it has three 

nozzles that curve so that the aerosol stream is tangential to the wall of the collection 

vessel. This causes the collection liquid to swirl and greatly reduces the agitation, bubbling 

and consequent reentrainment seen with the AGI-30. The BioSampler collects particles 

with aerodynamic diameters of approximately 0.3 μm to 8 μm into the collection media, 

although the upper cut-off diameter is not sharp [Hogan et al. 2005; Kesavan et al. 2010; 

Willeke et al. 1998]. The BioSampler reportedly can be used with non-evaporating fluids 

such as mineral oil to eliminate the collection time limits imposed by water evaporation, 

provided that the microorganism can survive collection and processing [Lin et al. 2000]. 

Alternatively, fluid can be exchanged or added to the sampler as needed [Rule et al. 2005; 

Rule et al. 2007].

The CIP10-M, a modified version of the CIP10 aerosol sampler, collects airborne 

microorganisms in a liquid layer on the interior surface of a rapidly-rotating cup. As with 

the BioSampler, the CIP10-M can be used with mineral oil as the collection fluid to avoid 

fluid evaporation. It is reported to have collection efficiencies of >80% for particles >2.8 

μm, 50% for 2.1 μm particles, and <10% for particles of <1 μm [Görner et al. 2006; Simon 

et al. 2016].

May [1966] designed a three-stage sampler in which aerosol particles are collected by 

impaction onto a wetted fritted surface in the first two stages and the third stage is a 

swirling aerosol collector similar to the BioSampler. Both glass and stainless steel versions 

are available. In his original report, May [1966] used particles with a density of 1.5 g/cm3

and reported cut-off sizes of 6 μm, 3.3 μm and 0.7 μm, which correspond to aerodynamic 

diameters of about 7.3 μm, 4 μm, and 0.86 μm. The May sampler is reported to give 

comparable results to the Andersen impactor [Zimmerman et al. 1987].
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Sampling and Characterization of Bioaerosols 

 Several wetted-surface bioaerosol samplers recirculate the collection fluid and add 

additional fluid as needed to replace evaporative losses. This extends the collection time 

available and allows the concentration of the aerosol from a large volume of air at a high 

flow rate into a relatively small volume of liquid, which is of great advantage when 

searching for pathogens that may be present in very low concentrations. For this reason, 

such systems are often used for bioterrorism and homeland security applications. The 

Coriolis sampler [Carvalho et al. 2008], the OMNI-3000 [Zhao et al. 2014], the SASS 2000 

[Ravva et al. 2012], and the SpinCon [Yooseph et al. 2013] use a wetted wall cyclone for 

bioaerosol collection, while the BioCapture 650 [Ryan et al. 2009] collects particles onto a 

wetted rotating impactor. Kesavan and Sagripanti [2015] reported the results of 

performance tests for several of these types of bioaerosol samplers.

When conducting long-term bioaerosol collection into liquid media, it is important to 

note that if the collected bioaerosol particles remain in the collection media for an 

extended time and if steps are not taken to inhibit growth, spore germination and cell 

amplification of some fungi and bacteria can occur. This can result in the appearance of 

much higher bioaerosol concentrations than are actually present in the environment.

f. Condensation-based bioaerosol samplers

Some bioaerosol particles are too small to be readily collected by impactors or impingers. 

These particles can be collected using filters, but filter collection can reduce the viability of 

microorganisms. One solution is to humidify the aerosol stream and then cool it, which 

causes water vapor to condense on the aerosol particles and create a droplet surrounding 

the particle. This larger particle can then be collected by impaction or impingement, as 

shown in Figure 9. This is similar in principle to condensation-based particle counters, 

which are used to measure the concentration of small airborne particles. Some researchers 

showed that adding water vapor to an aerosol stream enhanced the recovery of airborne 

viruses and bacteriophages, which may work by this method (although this is unclear) 

[Hatch and Warren 1969; Trouwborst and Kuyper 1974; Warren et al. 1969]. More 

recently, Milton developed a condensation-based system to collect fine particles 

containing influenza virus from the exhaled breath of human subjects [McDevitt et al. 

2013; Milton et al. 2013]. A condensation-based bioaerosol sampler called a growth-tube 

collector has been used to collect MS2 bacteriophage and influenza virus in the laboratory, 

and is reported to be especially effective at recovering viable virus in sub-micrometer 

particles [Lednicky et al. 2016; Pan et al. 2016; Walls et al. 2016]. A version of this system 

called the Spot Sampler (Aerosol Devices, Inc.) is commercially available.
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Figure 9: Condensation-based aerosol particle collector.

g. Electrostatic samplers

Electrostatic precipitation works by using a strong electric field to create a high 

concentration of unipolar ions. The rapid motion of these ions causes them to collide with 

and charge airborne particles, and the resulting charge on the particles causes them to be 

attracted to the collection surface [Hinds 1999]. Electrostatic precipitation systems have 

been used to collect bioaerosol particles such as allergens, bacteria and viruses [Artenstein 

et al. 1968; Artenstein et al. 1967; Custis et al. 2003; Donaldson et al. 1982; Heitkamp et al. 

2006; Lee et al. 2004b; Parvaneh et al. 2000; Roux et al. 2013]. Such devices offer simplicity 

of design with few moving parts, and are generally effective at collecting small particles. 

One electrostatic bioaerosol sampling device is available commercially from Inspirotec 

[Gordon et al. 2015].

Some electrostatic bioaerosol samplers collect particles into liquid to concentrate the 

particles and help preserve the viability of microorganisms. The Large Volume Air 

Sampler (LVS) developed by Litton in the 1960’s washed the collection surface with 

recirculating fluid; this sampler was successfully used to collect pathogenic respiratory 

bacteria and viruses in a variety of settings [Artenstein et al. 1968; Artenstein et al. 1967; 

Donaldson et al. 1982]. Pardon et al. [2015] developed a system that collects particles 

directly on a microfluidic chip. The electrostatic aerosol collector devised by Han et al. 

[2015] collects the deposited aerosol into rolling water droplets, which greatly 
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concentrates the particles. The Aerosol-to-Liquid Particle Extraction System (ALPES) uses 

an electrostatic system to collect aerosol particles into recirculating liquid, which helps 

preserve the viability of microorganisms [Heitkamp et al. 2006]. 

Electrostatically-charged cloths are used to collect airborne particles that settle onto them, 

and also to wipe settled dust from surfaces. These are discussed in the next two sections.

h. Passive bioaerosol samplers

Passive bioaerosol sampling refers to the collection of bioaerosols by allowing them to

gravitationally settle onto a collection device, such as a culture plate, foil sheet, electretbased filter or electrostatically-charged cloth. Compared to active sampling, passive

bioaerosol sampling has several advantages, including simplicity, low cost, lack of

disturbance of the surrounding air, and the ability to collect for extended time periods

[Haig et al. 2016; Pasquarella et al. 2000; Vincent 2007].

Passive bioaerosol sampling can be limited by several variables including the air currents 

around the device and airborne particle size. As discussed earlier, large particles settle 

much more quickly than small particles. Thus, large particles are much more likely to be 

collected by passive samplers [Haig et al. 2016; Reponen et al. 2011b]. As a result of these 

limiting variables, results from passive bioaerosol sampling cannot be directly related to 

the concentration of airborne particles and may not correlate well with results from active 

sampling [Reponen et al. 2011b]. However, some authors have proposed that passive 

sampling may be useful in evaluating the likelihood that bioaerosol particles will 

contaminate surfaces such as open wounds in operating rooms, since they mimic the 

contamination event more closely than does an active sampler [Friberg et al. 1999; Haig et 

al. 2016; Pasquarella et al. 2000].

Passive bioaerosol collectors are often placed 1.5 to 2 meters above the ground to avoid 

collection of large dust particles from sources other than airborne particles, such shoes, 

clothing, skin and animals [Frankel et al. 2012; Lioy et al. 2002; Noss et al. 2008; Rintala et 

al. 2012]. Grills, screens or shields may also be placed around or over the collection device 

to screen out large debris [Brown et al. 1996; Wagner and Macher 2003; Whitehead and 

Leith 2008; Wurtz et al. 2005].

Settle plates

Settle plates (also called settling plates or sedimentation plates) are culture plates 

containing nutrient agar that are opened and placed collection-side up in a location of 

interest. Airborne particles are allowed to settle onto the plates for a specified time, 

and the plates are then closed, incubated and inspected for growth. Settle plates are 
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commonly used to assess airborne microbial contamination and are listed in methods 

and standards from the ISO, the American Public Health Association (APHA) and the 

United States Pharmacopeia (USP) [Dyer et al. 2004; ISO 2003; USP 1997]. However, 

because the results from settle plates cannot be directly compared to the amount of 

airborne microbes, they should only be used for qualitative, not quantitative, 

evaluations. The CDC recommends the use of high-volume air samplers rather than 

settle plates when investigating airborne fungal spore contamination in health care 

facilities [CDC 2003].

Settle plate methods suffer from a lack of standardization of methodology, which 

makes results difficult to compare. Pasquarella et al. [2000] reviewed the use of settle 

plates and proposed an Index of Microbial Contamination (IMA) to standardize the 

use of settling plates. To measure the IMA, 90 mm culture plates are placed 1 meter 

above the floor and 1 meter from any walls, and collect settled particles for 1 hour 

(called the 1/1/1 scheme). The number of colony-forming units (CFUs) detected on 

each plate is then used to calculate the IMA in CFUs/dm2/hour [Pasquarella et al. 

2000]. 

Electrostatic dust collectors

Noss et al. [2008] developed a method called the electrostatic dustfall collector (EDC) 

that collects settling airborne particles onto four electrostatically-charged cloths. 

EDC’s have been used in studies of culturable bacteria and fungi, endotoxin, glucan 

and inflammatory mediators in airborne particles [Adams et al. 2015; Frankel et al. 

2012; Huttunen et al. 2016; Kilburg-Basnyat et al. 2016; Kilburg-Basnyat et al. 2015; 

Noss et al. 2010; Noss et al. 2008]. Noss et al. [2008] and Frankel et al. [2012] reported 

good correlations between the EDC and active aerosol samplers. Adams et al. [2015] 

compared EDC’s to Petri dishes and other passive collection materials and found that 

the results correlated reasonably well, but that a rigorous extraction protocol was 

required to get consistent results from the EDC’s. Brown et al. [1996] developed a 

passive electrostatic-based personal aerosol sampler and reported that it gave a 

reasonable correlation with inhalable dust measurements at farms and a rubber plant.

Other passive bioaerosol samplers

The UNC Passive Aerosol Sampler consists of a 6.8 mm diameter collection substrate 

mounted on a scanning-electron microscope stub and shielded by a protective screen 

[Wagner and Macher 2003; Whitehead and Leith 2008]. Airborne particles settle or 

diffuse onto the substrate and can be analyzed by optical or electron microscopy. 

Other investigators have used aluminum sheets in boxes, Petri dishes, and sheets of 

various plastic materials as passive bioaerosol collectors [Adams et al. 2015; Meadow et 

al. 2015; Wurtz et al. 2005].
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i. Settled dust collection devices

The collection and analysis of dust that has settled onto floors, carpets, and other surfaces

is widely used as a means of identifying bioaerosols in buildings, especially allergens,

endotoxin and molds [Hung et al. 2005; Lioy et al. 2002; Martyny et al. 1999; Morey 2007;

Rintala et al. 2012]. Settled dust sampling allows for the collection of large quantities of

material, provides a long-term sample, and does not require a dedicated sampling device

for each location. Dust assays allow quantitative data to be generated per weight and

surface area of dust. Some investigators find it useful to compare different sites in a

building or to sample before and after remediation efforts to see if the source of a

bioaerosol has been eliminated.

Settled dust will vary within a building depending upon the location and collection surface 

[Lioy et al. 2002; Rintala et al. 2012]. In addition to settling from the air, dust can be 

produced by a variety of other mechanisms, making it difficult to distinguish the source. 

Floor and carpet dust, for example, will include outside material brought in by shoes, skin 

flakes, clothing fibers and animal dander. Sampling locations well above floor level are 

often chosen to minimize the amount of dust that is not from settled airborne particles 

[Frankel et al. 2012; Rintala et al. 2012].

Vacuums

The US Department of Housing and Urban Development has developed a protocol for 

the vacuum collection of home dust samples to test for allergens [HUD 2008]. 

Vacuum collection of settled dust from floors and carpets has been used to determine 

the Environmental Relative Moldiness Index (ERMI), which is a measure of mold 

contamination in homes [Kettleson et al. 2015; Reponen et al. 2012; Reponen et al. 

2011a; Taubel et al. 2016; Vesper et al. 2013; Vesper et al. 2007]. ERMI is discussed in 

more detail later in this chapter. Note that vacuuming can increase the levels of 

bioaerosols in a location. Thus, air sampling should be completed before collecting 

surface samples by vacuuming [Hung et al. 2005; Hunter et al. 1988].

Swabs

Swabs are widely used to collect airborne material that has settled onto surfaces. Swabs 

are also used to identify microbial contaminants that may be colonizing building 

materials within the indoor environment. However, obtaining consistent and reliable 

results from swab sampling is far more difficult than is often appreciated, and careful 

attention is needed to the choice of swab material, elution media, and method of 

swabbing. If swab samples are to be cultured, aseptic technique is needed to avoid 

contamination. ASTM International has a standard for collecting fungal material by 

swab [ASTM 2012]. The APHA has published a standard method for swab sampling of 
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 food-contact surfaces [Dyer et al. 2004], while the USP and ISO have standards that 

include swab sampling for microbiological contamination in clean rooms [ISO 2003; 

USP 1997]. 

An example of a validated protocol for swab sampling is that provided by NIOSH for 

surface sampling for Bacillus anthracis spores [Hodges et al. 2010; Hodges et al. 2006; 

NIOSH 2012b]. In this procedure, a defined area is first outlined using a template or a 

ruler and masking tape. A sterile macrofoam swab is then moistened using a buffer 

solution that neutralizes disinfectants. The surface is swabbed using horizontal strokes, 

followed by vertical strokes, and finally diagonal strokes, and the swab is then placed 

in a sterile tube for transport and analysis. Aseptic technique is used throughout the 

procedure.

The choice of swab material can have a significant impact on the collection of 

microorganisms from a surface. Moore and Griffith [2007] studied the recovery of 

Escherichia coli and Staphylococcus aureus from stainless steel squares using nylonflocked swabs and spatulas, cotton swabs and rayon swabs. They reported that nylonflocked and cotton swabs were equally effective at removing bacteria from dry 

surfaces, but that cotton swabs removed bacteria more effectively from wet surfaces 

than rayon or nylon-flocked swabs. However, nylon-flocked swabs and spatulas 

released the bacteria into the elution media more readily than rayon swabs, which in 

turn released more bacteria than cotton swabs. For viruses, polyester-tipped swabs 

were found to be more effective than cotton swabs or antistatic wipes at recovering 

MS2 bacteriophage from stainless steel and plastic [Julian et al. 2011], while 

macrofoam swabs performed best when recovering wet or dried norovirus from 

stainless steel surfaces, followed by cotton, rayon and polyester swabs [Park et al. 

2015]. 

The elution media used to wet the swabs and recover the bacteria from the swabs also 

can have a substantial effect on sampling. Moore and Griffith [2007] tested eleven 

different swab wetting solutions containing various combinations of salts, surfactants 

and nutrients. They found that the recovery efficiency varied widely depending upon 

the species of bacteria, type of swab, and whether the surface was wet or dry. For MS2 

bacteriophage, saline or Ringer’s solution (an isotonic salt solution) worked better 

than viral transport media or pure water [Julian et al. 2011]. It is important to note 

that the elution media must both remove the biological material from the surface and 

subsequently elute it from the swab in order to be effective.

Although they may be overlooked, storage conditions play an important role in swab 

sampling. After sample collection, room temperature storage of moist swabs may lead 
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to microbial growth if the elution media or swab contain nutrients, while the presence 

of chemicals such as Tween 80 may reduce viability over time. These problems can be 

alleviated by placing the swabs in cold storage as quickly as possible [Moore and 

Griffith 2007].

Wipes

All of the considerations and limitations of swab sampling also apply to wipe 

sampling. Swabs are typically more useful for small surfaces and hard-to-reach 

locations, while wipes are more effective at collecting dust from large non-porous 

surfaces [NIOSH 2012b]. Electrostatic wipes have been used to collect settled dust for 

studies of mold and endotoxin [Bolaños-Rosero et al. 2013; Thorne et al. 2005]. 

However, Thorne et al. [2005] found that wipes and gloves themselves were frequently 

contaminated with endotoxin and needed to be tested before use.

Adhesive tape

Adhesive tape can be used to collect dust samples from surfaces for microscopic 

examination (this is called tape lift or cellotape sampling) [ASTM 2014c; Martyny et al. 

1999; Morey 2007]. Typically, a section of adhesive tape is gently pressed onto a 

surface of interest, removed with a slow steady force, and then attached to a glass slide 

or placed in a vial. The samples are relatively simple to collect, but the results depend 

upon the ability of the examiner to identify microorganisms and their fragments, and 

do not provide a quantitative assessment of exposure.

Contact plates

Contact plates are typically round culture plates in which the agar is poured so that the 

top of the agar forms a meniscus slightly above the top rim of the plate. A surface 

sample is collected by inverting the plate and pressing the agar directly onto a flat 

surface of interest. The plate is then removed, incubated and inspected for microbial 

growth. This sampling method is often called the replicate organism direct agar 

contact (RODAC) procedure, and it is commonly used for biocontamination 

monitoring in the pharmaceutical and food industries [Dyer et al. 2004; ISO 2003; USP 

1997]. Because many of the surfaces of interest in these industries are routinely 

disinfected, contact plates are available with agars that contain neutralizers for 

disinfectants. One report indicated that nitrocellulose membranes were slightly more 

effective than RODAC plates at surface sampling, and are easier to use on curved 

surfaces [Poletti et al. 1999].
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j. Heating, ventilation and air conditioning (HVAC) filters

Building HVAC systems filter large quantities of outside and recirculated inside air as they 

maintain environmental conditions inside buildings. Researchers have taken advantage of 

these existing filtration systems as a way to study bioaerosols in a variety of structures 

[Goyal et al. 2011; Haaland and Siegel 2016; Noris et al. 2011]. Testing the collected 

particulate material on HVAC filters provides an inexpensive way of studying bioaerosols 

collected from large volumes of air over long time periods. However, some limitations 

must be kept in mind. Extracting bioaerosols from these filters can be difficult and the 

methods require validation [Farnsworth et al. 2006]. Many microorganisms lose viability 

after collection, so although PCR-based methods may be effective, culture-based methods 

likely will not work except for very hardy microbes [Farnsworth et al. 2006]. Finally, 

commonly-used HVAC filters can have relatively low collection efficiencies, especially for 

small particles [ASHRAE 2009]. Haaland and Siegel [2016] reviewed 60 studies in which 

HVAC filter analyses were used to study bioaerosols in buildings.

k. Real-time bioaerosol monitoring

Many biological molecules have an intrinsic autofluorescence, and this phenomenon has 

been used as the basis for continuous real-time bioaerosol detection systems [Pöhlker et al. 

2012]. This technique is most commonly employed for studies of atmospheric bioaerosol 

particles and for biodefense and biosecurity applications. These systems can distinguish 

biological from non-biological particles, and can usually provide information about the 

particle size and some characteristics of the bioaerosols. One device, the TSI Ultraviolet 

Aerodynamic Particle Sizer (UV-APS), was used in several studies [Bhangar et al. 2016; 

Hairston et al. 1997; Kanaani et al. 2008]; it has been replaced by an updated version called 

the Fluorescence Aerosol Particle Sensor (FLAPS) III. Other real-time bioaerosol detectors 

include the BioScout [Saari et al. 2014], the Wideband Integrated Bioaerosol Sensor 

(WIBS-4) [Toprak and Schnaiter 2013], and the Fido B2 (formerly called the 

Instantaneous Bioaerosol Analysis and Collection, IBAC) [Santarpia et al. 2013].

4 Considerations for bioaerosol sampling

a. Development of a bioaerosol sampling strategy

The first step in designing a sampling strategy for bioaerosol sampling is to determine the 

purpose of the sampling [ASTM 2014a]. For example, bioaerosol sampling may be 

conducted to estimate worker exposure to bioaerosols, or to select or evaluate engineering 

controls to reduce exposures, or to identify the source of a bioaerosol. A sampling strategy 

then should begin with an overview of the site of interest and development of initial 

hypotheses regarding the types, sources and distributions of bioaerosols. After this, the 

sampling methods, times, durations, and the analytical methods can be selected. Note that 
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 bioaerosol sampling is almost always done in conjunction with the collection of other 

types of data, such as worker health information, visual observations, air flow 

measurements, surface sampling, and information about possible sources.

b. Sampling locations

The sampling locations should be selected to assist in evaluation of the working 

hypotheses about possible exposures [ASTM 2014a]. If worker exposures are being 

evaluated, then the samplers should be placed in areas occupied by the workers. If 

contamination of a ventilation system is being examined, then sampling in the system and 

at the ventilation louvers would be appropriate. Care must be exercised to ensure that 

people do not tamper with the samplers and that microorganisms on surfaces or in duct 

work are not inadvertently aerosolized.

Bioaerosol samples should be drawn directly into the sampler rather than being 

transported to the sampler by tubing. If transport tubing must be used, it should be as 

short and straight as possible. Abrupt flow constrictions and bends in the tubing should be 

especially avoided, as considerable sample deposition can occur at these locations. The 

tubing diameter should be large enough that the flow is not turbulent and that the d50 of 

any bends is well above the size of the bioaerosol particles [Pui et al. 1987; Tsai and Pui 

1990]. The tubing should be made of a material that does not lead to losses through 

electrostatic deposition [Liu et al. 1985]. A review of the many issues surrounding the 

transporting of aerosols through sampling lines is provided by Brockmann [2011].

Personal aerosol sampling provides a much better representation of worker and resident 

exposure to aerosol particles than area (static) sampling [Cherrie et al. 2011; Kissell and 

Sacks 2002; Rodes and Thornburg 2005]. However, most samplers for viable bioaerosols 

do not lend themselves to personal sampling. Thus, a combination of personal and area 

sampling may be necessary to fully characterize the exposure [Toivola et al. 2002].

c. Concentrations of indoor and outdoor bioaerosols

Indoor bioaerosol sampling is conducted in occupational (industrial, education, and office 

environments) and non-occupational (residential and buildings) settings. Outdoor 

bioaerosol sampling is often performed to provide comparative data for indoor sampling 

and to help determine possible sources of contaminants. Outdoor bioaerosol sampling also 

is conducted in occupational environments such as agricultural settings, composting sites 

and sewage treatment plants [Environment Agency 2009; Lee and Liao 2014; Masclaux et 

al. 2014]. In addition, outdoor sampling may be performed for pollen and fungi to assist 

allergists in their treatment of patients by identifying taxa distribution and concentrations 

in air over time.
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 The concentrations of bioaerosol particles vary widely depending upon the meteorological

parameters, the location of sources, the time of year and the amount of ventilation. 

Shelton et al. [2002] studied 1,717 buildings in the United States. They found that outdoor 

levels of airborne fungi are usually higher than indoor levels, and that fungal levels were 

highest in the fall and summer and lowest in the winter and spring. Outdoor levels varied 

from 1 to more than 8,200 colony-forming units (CFU)/m3 of air, with a median of 540 

CFU/m3

. Indoor levels ranged from 1 to over 10,000 CFU/m3, with a median of 82 

CFU/m3. An examination of fungi in flood-damaged homes found fungal concentrations 

of 1,100 to 8,400 spores/m3 outside and 500 to 101,100 spores/m3 inside [Reponen et al. 

2007]. An investigation of 100 large office buildings by Tsai and Macher [2005] found that 

airborne bacterial concentrations tend to be higher outdoors than indoor (except for 

Gram-positive cocci). Outdoor concentrations tended to be higher in the winter (194 vs. 

165 CFU/m3), while indoor concentrations were higher in the summer (116 vs. 87 

CFU/m3). Forty-one percent of the bioaerosol samples were below the detection limit, and 

>95% of the culturable bacteria were mesophilic (grow at moderate temperatures). In a 

report on agricultural workers working in animal confinements, Lee et al. [2006] found 

breathing zone culturable bioaerosol exposures of 300 to 36,000 CFU/m3 for fungi, 3000 to 

3.3 x 108 CFU/m3 for bacteria, and up to 2,800 CFU/m3 for actinomycetes. During grain 

harvesting, workers were exposed to culturable bioaerosol levels of 82,000 to 7.4 × 106 

CFU/m3 for fungal spores, 40,000 to 1.4 × 106 CFU/m3 for bacteria, and up to 2.6 × 104 

CFU/m3 for actinomycetes.

If one or more genera of fungi or bacteria are found indoors in concentrations greater than 

outdoor concentrations, then the source of amplification may need to be found and 

remediated. When conducting indoor bioaerosol sampling, it is advisable to sample 

before, during, and after the sampling area is occupied, including times when the heating, 

ventilating, and air conditioning system is activated and inactivated.

d. Viable and nonviable bioaerosols

Viable microorganisms are metabolically active (living) organisms with the potential to 

reproduce, grow and colonize. Viruses are not metabolically active but are considered 

viable if they are capable of reproducing in an appropriate cellular host. Viable 

microorganisms may be culturable or non-culturable. Culturable organisms reproduce 

under controlled laboratory conditions. Non-culturable organisms do not reproduce in 

the laboratory because of intracellular stress or because the conditions (e.g., culture 

medium or incubation temperature) are not conducive to growth. Some bacteria can be 

very difficult or impossible to culture from bioaerosols. For example, although human 

Mycobacterium tuberculosis is readily transmitted among people and from people to 

Guinea pigs, it has never been successfully cultured from an environmental aerosol 
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 sample, probably because of its extremely low airborne concentrations and slow growth 

rate [Nardell 2016]. Other bioaerosols such as Histoplasma capsulatum or Pneumocystis 

carinii may take weeks to grow or may not even grow in culture at all [Dennis 1990; Ibach 

et al. 1954]. As the name implies, viable bioaerosol sampling involves collecting a 

bioaerosol and culturing the collected particles. Only culturable microorganisms are 

enumerated and identified, thus leading to an underestimation of bioaerosol 

concentration. Non-viable and viable but non-culturable microorganisms are often 

studied by collecting them with a dry aerosol sampler or a membrane filter. The 

microorganisms are then enumerated and identified using microscopy, classical 

microbiology, molecular biology, or immunochemical techniques [Hung et al. 2005; 

Macher 1999; Reponen et al. 2011b; Tortora et al. 2013].

Assessment of viable bacteria is also dependent on a number of variables including 

nutrient media, temperature and culture conditions. In indoor environments the 

collection of viable bacteria may be confounded by endogenous bacterial microflora such 

as Staphylococcus epidermis that sheds with skin flakes [Hung et al. 2005]. Concentrations 

of viable bacteria have been reported to be as high as 105 CFU/m3 in indoor environments; 

however, like fungi, the proportion of the total bacterial burden may be higher if nonviable bacteria are also included [Hung et al. 2005]. In addition, viable assessment of 

several bacterial species of clinical significance may not be the best approach as these 

bacteria do not remain viable in the air. Alternative methods such as immunoassays or 

molecular-based methods may provide suitable approaches for quantifying bacterial 

pathogens.

e. Bioaerosol particle sizes

As noted earlier, the aerodynamic diameter (dae) of an airborne particle is the most 

important factor determining how long it will remain in the air, how likely it is to be 

inhaled, and where it will deposit in the respiratory tract. The sizes of bioaerosol particles 

can range from tens of nanometers for small fragments to hundreds of micrometers for 

pollen, fungi or large agglomerations. However, most of the bioaerosol particles of interest 

in the indoor environment fall between about 100 nm and 10 μm [Nazaroff 2016]. For 

bacteria, vegetative cells typically have physical diameters of about 0.2 to 2 μm and are 2 to 

8 μm in length, while bacterial spores are somewhat smaller [Tortora et al. 2013]. Airborne 

particles containing bacteria were found to have aerodynamic diameters of about 1 to 3 

μm in indoor environments [Gorny et al. 1999; Kujundzic et al. 2006; Meklin et al. 2002]. 

Mycobacterium tuberculosis is a rod-shaped bacteria with a length of about 6.6 μm [Schafer 

et al. 1999]. When aerosolized from a liquid culture, M. tuberculosis DNA was found in 

particles with aerodynamic diameters of 0.6 to 1.8 μm [Schafer et al. 1999]. Aerosolized 

Mycobacterium bovis BCG (a commonly-used surrogate for M. tuberculosis) was found in 
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 particles with aerodynamic diameters of 0.5 to 9.9 μm [Schafer et al. 1998]. Air sampling 

around indoor whirlpools in a public facility found airborne mycobacteria DNA in 

particles with aerodynamic diameters of 0.5 to 9.9 μm [Schafer et al. 2003]. Actinomycete

spores tend to be smaller, with aerodynamic diameters of cultured spores ranging from 0.6 

to 1.5 μm [Madelin and Johnson 1992; Reponen et al. 1998]. Fungal spores have physical 

diameters of about 0.5 to 30 μm or larger, while the aerodynamic diameters of airborne 

fungal spores and spore clusters are reported to be from 0.9 to 5 μm [Eduard 2009; 

Hussein et al. 2013; Reponen et al. 2011b].

Airborne microorganisms are often present as parts of aggregations, droplets or 

agglomerations that can be much larger than the size of the native microorganism. In 

indoor environments with large amounts of other aerosol particles like cigarette smoke, 

bacteria have been found on particles with aerodynamic diameters up to 10 μm, which was 

larger than airborne bacterial particles in cleaner environments. This was thought to occur 

because the aerosol particles were forming agglomerates [Gorny et al. 1999]. In a farm 

study, airborne Actinomycetes and fungal spores were more likely to be found in 

aggregates in environments with higher spore concentrations [Karlsson and Malmberg 

1989]. In two studies of airborne influenza virus in health care facilities, about half of the 

airborne virus was found in particles with aerodynamic diameters of 4 μm or greater, even 

though the virus itself is only about 100 nm in diameter, because the virus was contained 

in aerosolized droplets of respiratory fluids [Blachere et al. 2009; Lindsley et al. 2010a]. 

Agglomerates of fungal spores can break apart upon impaction inside an impactor and be 

collected on subsequent stages with smaller cut-off diameters [Trunov et al. 2001].

Bioaerosols may also be present as cellular fragments that are much smaller than the 

source microorganisms. Endotoxins are fragments of the cellular walls of Gram-negative 

bacteria that have been implicated in a variety of illnesses [Eduard et al. 2012; Jacobs 1989; 

Olenchock 2002]. Fragments of fungal cell walls also are thought to be associated with 

several types of adverse respiratory health effects [Green et al. 2011; Green et al. 2006b; 

Olenchock 2002]. Very high levels of fungal fragments have been measured in flooddamaged homes contaminated with mold [Reponen et al. 2007]. Fungal fragments also 

contain a variety of secondary metabolites, mycotoxins, beta-glucan, antigens and 

allergens [Green et al. 2011; Green et al. 2006b]. In one study of indoor air in homes, the 

majority of the endotoxin and fungal wall material was found in particles with 

aerodynamic diameters of less than 1 μm [Adhikari et al. 2013]. Another study found 

considerable amounts of endotoxin in aerosol particles from metalworking fluids that 

were between 0.16 and 0.39 μm [Wang et al. 2007]. Indoor and outdoor measurements of 

endotoxin levels found that the largest proportion was detected in particles with 

aerodynamic diameters of less than 1 μm [Kujundzic et al. 2006].
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Sampling and Characterization of Bioaerosols 

 It is common to use an aerosol spectrometer in conjunction with bioaerosol sampling to 

better understand the size distribution of the airborne particles. One consideration when 

interpreting the data is, of course, that the large majority of these devices do not 

distinguish between biological and non-biological aerosols. Another less-obvious factor is 

that while a few aerosol spectrometers such as the TSI Aerodynamic Particle Sizer measure 

the aerodynamic diameter of the airborne particles, many aerosol spectrometers measure 

particles using light scattering and thus provide an approximate physical diameter instead 

[Hinds 1999; Sorensen et al. 2011]. The difference between the aerodynamic and optical 

diameters may be significant depending upon the shape and density of the particles.

f. Temperature and humidity

The temperature and humidity of the environment can affect the size of bioaerosol 

particles, the viability of airborne microorganisms, the growth of microorganisms on 

surfaces, and the amount of electrostatic charges on aerosols and surfaces. Because of these 

effects, the environmental temperature and humidity should be recorded during 

bioaerosol sampling.

Water evaporates rapidly from wet aerosol particles [Hinds 1999]. If an airborne particle is 

initially an aqueous solution containing non-volatile substances such as salts and organic 

material, and if the relative humidity is above the crystallization relative humidity (CRH, 

also called the efflorescence relative humidity), then some of the water will evaporate and 

the solution will become more concentrated, but the particle will remain liquid. If the 

relative humidity is below the CRH, then all of the water will evaporate (that is, the particle 

will desiccate) [Nicas et al. 2005]. Similarly, if an airborne particle is initially a dry 

combination of salts and organic material, and if the relative humidity is below the 

deliquescence relative humidity (DRH), then the particle will remain desiccated. However, 

if the relative humidity is above the DRH, then the particle will absorb water until it 

liquefies and becomes an aqueous solution. The DRH is always greater than the CRH 

[Nicas et al. 2005]. A particle in an environment above its CRH (or DRH if it was initially 

dry) will be larger and heavier and will settle faster than the same particle when the 

humidity is below the CRH, which can affect the size and amount of bioaerosol particles 

that are collected during sampling [Mikhailov et al. 2004]. This phenomenon was seen in a 

study of particles in human exhaled breath, where the particles detected in low humidity 

air were substantially smaller than those detected when the air was more humid 

[Holmgren et al. 2011].

Bioaerosol particles may also undergo an increase in size when the humidity increases due 

to water absorption and swelling of hygroscopic components. An increase in relative 

humidity has been shown to increase the aerodynamic diameter of fungal spores [Madelin 
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 and Johnson 1992; Reponen et al. 1996]. Similar results have been reported for 

Actinomycetes spores [Madelin and Johnson 1992].

For airborne viruses, survival decreases as air temperature increases [Ijaz et al. 2016; Tang 

2009]. Exposing most viruses to temperatures of 60°C or higher for 60 minutes will 

inactivate them, although the viruses can be somewhat protected if they are encased in 

organic material [Tang 2009]. For example, in one set of experiments, airborne particles 

containing vaccinia virus, influenza virus, and Venezuelan equine encephalomyelitis virus 

all showed higher survival rates at 7-12°C than at 21-24°C, and still lower survival at 32-

34°C [Harper 1961]. Aerosol transmission of influenza virus among Guinea pigs is 

blocked at air temperatures of 30°C [Lowen et al. 2008]. The effect of humidity on virus 

survival depends upon the virus; in general, viruses with lipid envelopes tend to survive 

better at low humidity, while non-enveloped viruses survive better at high humidity [Ijaz 

et al. 2016; Tang 2009]. For example, influenza viruses and coronaviruses have enveloped 

capsids, and both survive better at low humidities compared to high [Ijaz et al. 1985a; Ijaz 

et al. 2016; Noti et al. 2013; Schaffer et al. 1976]. On the other hand, rotaviruses and 

rhinoviruses have non-enveloped capsids and survive better at high humidities compared 

to low [Ijaz et al. 1985b; Ijaz et al. 2016; Karim et al. 1985].

The survival of airborne bacteria also decreases as air temperature increases; the survival of 

virtually all airborne bacteria declines when temperatures are above 24°C [Ijaz et al. 2016; 

Tang 2009]. However, as with viruses, the effects of humidity on bacterial survival are 

much more complex, and depend not only upon species but also upon the methods of 

culture and aerosolization [Cox 1989; Tang 2009]. In field experiments in a greenhouse, 

survival of certain bacteria was 35- to 65-fold higher at 80% RH than at 40% [Walter et al. 

1990]. In laboratory experiments, survival of certain bacteria was virtually complete at low 

RH but was reduced at RH values above 80% [Cox 1968]. Higher humidities can also 

significantly decrease the efficacy of ultraviolet germicidal irradiation (UVGI) for reducing 

levels of viable airborne bacteria [Peccia et al. 2001]. Cox [1987] believes the potential for 

the movement of the solvent water is an important environmental criterion in assessing 

survivability of bacteria, viruses, and phages.

Fungi and fungal spores generally are better able to withstand environmental stresses 

compared to vegetative bacteria and viruses [Ijaz et al. 2016; Tang 2009]. Warm 

temperatures, wet substrates and humid air conditions favor the growth of fungi on 

surfaces [Eduard 2009; Tang et al. 2015]. Temperature can induce morphological changes 

in dimorphic fungi such as the pathogen Histoplasma capsulatum [Salvin 1949]. It is not 

clear, however, how air temperature and humidity affect the viability of airborne fungi and 

fungal spores [Tang 2009].
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g. Electrostatic effects

Aerosol particles in the workplace can be highly charged, and the electrostatic charge can 

vary considerably depending upon the aerosol generation mechanism and the particle 

characteristics [Johnston et al. 1985]. Aerosol particles are especially prone to develop 

electrostatic charges in low humidity environments [Baron and Deye 1990]. Like most 

particles, freshly generated microbial aerosols are nearly always electrostatically charged 

unless steps are taken to neutralize them. Lee et al. [2004b] found that airborne fungi and 

bacteria carried a net negative charge in most of the laboratory and field environments 

that they studied. Mainelis et al. [2002] found that a strong positive electrostatic charge 

reduced the viability of Pseudomonas fluorescens bacteria but did not affect Bacillus subtilis

spores.

The effect of electrostatic charge on aerosol collection is often overlooked, resulting in the 

possible bias of sampling results [NIOSH 2016a; Vincent 2007]. Aerosol samplers made of 

non-conductive plastics can develop substantial electrostatic charges, which can degrade 

their performance significantly [NIOSH 2016a; Baron and Deye 1990]. The use of 

polyethylene or polytetrafluoroethylene (PTFE) tubing to transport air streams to a 

sampler can remove a sizeable amount of aerosol particles by electrostatic deposition [Liu 

et al. 1985]. As noted above, the use of plastic Petri dishes in an Andersen impactor can 

result in bioaerosol particle losses [Andersen 1958; Kuo 2015]. Whenever possible, it is 

better to use aerosol samplers made of conductive materials such as metals or speciallytreated plastics [NIOSH 2016a].

h. Flow calibration

Accurate airflow rates are very important in calculating the concentration of 

microorganisms in the air. All samplers should be calibrated before and after sampling to 

ensure that the flow rate is within the manufacturer's specifications and does not change 

from the initial calibration. Calibration may be performed using a primary standard such 

as a spirometer or bubble calibrator. Where it is not possible to calibrate using a primary 

standard, a calibrated secondary standard such as a dry gas meter may be used. The 

calibration of such a secondary standard should be traceable to a primary standard. A 

detailed explanation of the calibration of airflow rates is given by McCammon Jr. and 

Woebkenberg [NIOSH 2016c].

i. Blanks 

Laboratory media blanks are unexposed, fresh samples of media, such as agar plates, filters 

and impinger fluids. These samples are generally not taken into the field. Before using any 

batch of media, incubate at least three culture plates under the same conditions as planned 
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Sampling and Characterization of Bioaerosols 

 for the field samples, in order to check for sterility of the media. Approximately five media 

blanks should be included with each sample set. If the samples are to be analyzed by an 

outside laboratory, consult the specific laboratory procedure for the number of blanks to 

be submitted. Similarly, blank filters should be processed in the same manner as planned 

for field samples in order to check for contamination. 

Field blanks are simply unopened, fresh media samples that are handled in the same way 

as field samples, including labeling, except that no air is drawn through the sampler. The 

generally recommended practice for the number of field blanks is to provide at least two 

field blanks for every 10 samples with a maximum of 10 field blanks for each sample set.

5 Selection of bioaerosol samplers

The first step in selecting a bioaerosol sampling device is to establish the purpose of the 

sampling. Once the goal of the bioaerosol sampling is determined, the appropriate sampling 

methods may be chosen. The selected bioaerosol sampler must be capable of high efficiency 

particle collection within the physical and biological conditions required by the 

microorganisms to be sampled. The most appropriate sampling methods will be dictated in 

part by the techniques that will be used to analyze the sample. Methods for bioaerosol sample 

analysis are discussed in the next section. A list of some manufacturers and suppliers of 

bioaerosol sampling equipment and supplies is shown in Appendix I. The characteristics of 

several commonly used bioaerosol samplers are shown in Appendix II.

a. Sampling for airborne bacteria and fungi 

Choosing a bioaerosol sampler for bacteria and fungi begins by deciding how the 

bioaerosol will be analyzed, and in particular whether the viability of the bacteria or fungi 

will be evaluated. Culturable bioaerosol sampling instruments must minimize injury 

during the collection process and maintain the culturability of the collected 

microorganisms. If the sample will not be cultured, then the samples usually can be 

collected dry using a membrane filter, cyclone, impactor, or a combination of these. Dry 

collection is typically simpler and less expensive to perform, and filters and cyclones can 

handle a wide range of particle concentrations. Organisms that are difficult or impossible 

to grow in culture are often collected using dry techniques and assessed using polymerase 

chain reaction (PCR) based methods, which have the advantage of speed and specificity. 

PCR has been used for rapid detection of Histoplasma capsulatum and mycobacteria [Reid 

and Schafer 1999; Schafer et al. 1999; Schafer et al. 2003]. A DNA-based mold specific 

quantitative PCR (msQPCR) method is widely used to evaluate indoor fungal bioaerosols 

in the academic, government and commercial sectors, and is the basis for the 

Environmental Relative Moldiness Index (ERMI) used to quantify mold contamination in 
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 homes [Kettleson et al. 2015; Vesper et al. 2013]. The ERMI and other PCR-based assays 

are discussed in greater detail later in this chapter.

If viability is to be studied, then the samples usually will need to be collected with an 

impinger or in an Andersen impactor loaded with agar plates, because many 

microorganisms will lose viability due to damage or desiccation if collected dry [Cox 1987; 

Hung et al. 2005]. For example, a membrane filter sampler is not appropriate for sampling 

culturable Escherichia coli because the cells desiccate and become either nonviable or 

viable but not culturable under these conditions [Jensen et al. 1992]. Similar results have 

been reported for other bacteria and fungi [Macher and First 1984; Wang et al. 2001]. 

Depending upon the target microorganism, impingers may be filled with distilled water or 

a buffered isotonic solution, sometimes with antifoaming agents to reduce foaming and 

proteins to enhance survival. Mineral oil has also been used in impingers instead of 

aqueous solutions to avoid evaporation [Lin et al. 2000]. Impactors are loaded with agar 

plates; the choice of agar depends upon the microorganisms of interest and the desired 

selectivity (discussed in the next section).

As noted previously, depending upon the investigation that is being conducted, the 

particle size distribution of the bioaerosol may be very important in the evaluation of the 

data obtained. If particle size information is needed to, for example, determine how much 

of the bioaerosol is in the respirable size fraction, then a size-selective sampler should be 

used for at least some of the collections if possible. For example, if an SAS-Compact 

sampler was the selected sampler for collection of culturable Escherichia coli, an Andersen 

6-Stage sampler could be used to determine the particle size distribution at each location 

sampled. The expected size of the bioaerosol particles is also an important factor in 

choosing a sampler. For example, an impactor with a d50 of 4 μm should not be used to 

collect Aspergillus niger spores (dae 1-3 μm) because most spores would remain entrained 

in the air and pass through the instrument.

NIOSH Method 0800 discusses sampling for culturable airborne bacteria and fungi with 

an Andersen cascade impactor [NIOSH 2003b]. Standard methods for the collection of 

airborne fungi by inertial impaction are presented in ASTM Standards D7788 [ASTM 

2009; ASTM 2014d]. ASTM Standard D7391 also discusses the aspects related to the 

laboratory analysis.

b. Sampling for airborne viruses

Airborne viruses are more difficult to study in bioaerosols than bacteria and fungi for a 

variety of reasons [Prussin et al. 2014; Verreault et al. 2008]. Viruses are more difficult to 

culture because they are obligatory intracellular parasites that require a host cell for 
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 reproduction [Tortora et al. 2013]. Bioaerosols of pathogenic viruses have been found in 

many settings to be present in low concentrations that can be difficult to detect [Blachere 

et al. 2009; Bonifait et al. 2015; Lindsley et al. 2010a; Tseng et al. 2010; Yang et al. 2011]. 

Viruses also are generally more susceptible to damage during aerosol collection than are 

bacteria or fungi, although their sensitivity varies widely with the collection method and 

species [Appert et al. 2012; Turgeon et al. 2014; Zuo et al. 2013]. Aerosol sampling 

methods for viruses have been reviewed by Verreault et al. [2008].

Bacteriophages are viruses that infect bacteria rather than multicellular organisms. They 

are used in laboratory aerosol studies as tracers for aerosol particles and as surrogates for 

airborne viruses that infect humans [Fisher et al. 2012; Tseng and Li 2005; Turgeon et al. 

2014]. Bacteriophages are not known to be hazardous to humans but are of interest to 

industries that rely on bacteria such as cheese manufacturers [Verreault et al. 2011].

Polymerase chain reaction (PCR) based methods are often used to study viral bioaerosols. 

PCR has the advantages of being very sensitive and very specific, and considerably easier 

to perform than viral cultural assays. For this reason, most recent field studies of airborne 

viruses have used PCR as the detection method. Examples include studies of viruses in 

healthcare facilities [Blachere et al. 2009; Booth et al. 2005; Lindsley et al. 2010a; 

Thompson et al. 2013; Tseng et al. 2010], influenza at poultry and pig farms [Corzo et al. 

2013; Jonges et al. 2015], airborne viruses in a sewage treatment plant [Masclaux et al. 

2014], and respiratory viruses in human coughs and exhaled breath [Gralton et al. 2013; 

Lindsley et al. 2010b; Milton et al. 2013].

PCR has both the advantage and disadvantage of not requiring that the virus be viable in 

order to be detected. This eliminates the need to preserve viability during and after 

collection and allows the use of dry collection methods such as cyclone samplers, dry 

impactors and filters, which are simpler and easier to carry out. On the other hand, this 

also means that it is unclear whether the airborne virus is infectious or not, which makes 

interpretation of data more difficult. This is a common criticism of PCR-based bioaerosol 

studies.

If the virus in a bioaerosol sample is to be cultured, in most cases the sample will need to 

be collected into an aqueous media using an impinger or wetted surface aerosol collector. 

Fabian et al. [2009] showed that collecting airborne influenza virus in aqueous media 

using an SKC BioSampler preserved infectivity much better than dry collection using 

filters or an impactor. A less-common method is to collect viable viruses using an 

Andersen impactor. Gustin et al. [2011] collected airborne influenza virus using an 

Andersen impactor by placing a filter and a thin layer of gelatin on top of the agar in the 

culture plates. After collection, the gelatin was removed and melted at 37°C to allow 
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 subsequent culture of the virus. Note that the collection media must be compatible with 

the cell culture system used to host the virus.

6 Sample preparation for culturable bioaerosols

Collecting and culturing viable airborne microorganisms is the most common technique used 

by industrial hygienists to assess bioaerosols [Macher 1999]. However, the appropriate sample 

preparation method is highly dependent upon the microorganism(s) of interest, sample 

source, and down-stream analysis. These sampling approaches are further confounded as 

viable bioaerosols have been estimated to account for approximately 1% of the total bioaerosol 

load, and non-viable bioaerosols are often overlooked [Hung et al. 2005]. In contrast, nonculturable bacteria and fungi cannot be grown in conventional lab-based conditions, but their 

presence is still important from a health perspective [Green et al. 2011; Mitakakis et al. 2003]. 

Non-viable bioaerosols can be determined through other detection methodologies such as 

microscopy, proteomic, immunological, and molecular analysis methods, and some of these 

approaches are discussed in section 9. A list of the common bioaerosols encountered in 

indoor and outdoor environments, as well as the fungi that are common contaminants of 

indoor building materials, can be found in Flannigan et al. [2011].

a. Sample preparation for bacteria and fungi

Viable bacterial and fungal bioaerosol identification is made through the collection, 

deposition, and growth of a viable propagule or intact cell on a selected nutrient agar 

medium contained in a sterile petri dish or liquid culture suspension [Macher 1999]. 

These methods are similar for both fungal and bacterial bioaerosols [Flannigan et al. 2011; 

Hung et al. 2005]. Selection of the nutrient media, incubation conditions (time and 

temperature), and potential damage to the culturable bioaerosol during sampling are 

among several critical variables to review before the collection, growth and proliferation of 

a viable propagule [Eduard et al. 2012; Hung et al. 2005; Macher 1999]. These parameters 

have been reviewed elsewhere, but should be taken into consideration when planning an 

environmental survey [Hung et al. 2005; Macher 1999].

Growth media can be defined as either broad or selective [Macher 1999]. As the term 

implies, broad nutrient media supports the growth of a diverse number of 

microorganisms. In contrast, a selective growth medium, with appropriate energy sources, 

nutrients, and pH, is used to enrich growth of the specific microorganism in question and 

inhibit the growth of competitive organisms [Macher 1999]. A variety of broad and 

selective nutrient media for bacteria and fungi are available to the industrial hygienist and 

can be found in Hung et al. [2005] and Macher [1999].
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 Following sample collection, liquid or agar cultures are incubated at a suitable temperature 

and atmosphere (facultative versus aerobic) for an appropriate time. Fast-growing bacteria 

may develop microcolonies in hours, while fungi may take days to develop into a visible 

colony and perhaps sporulate. Organisms such as Mycobacterium tuberculosis or the 

dimorphic fungal pathogens, Histoplasma capsulatum or Blastomyces dermatitidis may 

require weeks of incubation to produce visible colonies [ATS 1990; Babady et al. 2011]. 

For fungi, plates are typically incubated at room temperature (18°C-25°C) or, if it is a 

clinically relevant isolate, at 35°C [ACGIH 1989; Baron and Finegold 1990; Hung et al. 

2005; Macher 1999]. In contrast, environmental bacteria are grown between 18°C and 

28°C, while thermophilic bacteria are grown between 50°C and 58°C [Hung et al. 2005; 

Macher 1999]. 

After allowing for vegetative growth of all viable propagules on the selected nutrient 

medium, the number of colonies is identified, quantified and presented as colony forming 

units (CFUs) [Eduard et al. 2012]. Media blanks (laboratory and field) should be processed 

using the same methods as samples to control for environmental or laboratory 

contaminants. A collection of bioaerosol identification manuals is presented in both the 

AIHA and ACGIH manuals [Hung et al. 2005; Macher 1999]. Color micrographs of 

common fungal contaminants are also presented in Flannigan et al. [2011]. Along with the 

quantification of viable microorganisms, taxonomic data and an interpretation of the 

datasets are generally reported [Hung et al. 2005].

The interest in detecting and quantifying fungi has increased following consensus 

documents that reported associations between fungi in damp indoor environments and 

adverse respiratory health effects [IOM 2004; Mendell et al. 2011; WHO 2009]. Compared 

to bacteria, additional variables need to be taken into consideration by the industrial 

hygienist when evaluating viable fungal bioaerosols including water activity, colony 

competition, and carbohydrate nutrient sources [Hung et al. 2005; Macher 1999]. Broad 

viable culture approaches favor species belonging to the phylum Ascomycota, as well as 

species that outcompete slower-growing species. Several different types of media and 

physiological conditions (e.g. temperature) may also need to be employed to assess 

complete fungal diversity using this approach. For fungi, selection of the nutrient media 

may potentially bias the growth of specific viable fungal bioaerosols. Common nutrient 

media include malt extract agar (MEA) supplemented with chloramphenicol or rose 

bengal agar to suppress bacterial growth [Hung et al. 2005; Macher 1999]. Cellulose agar 

can also be used for the selection of indoor fungal contaminants such as Stachybotrys 

chartarum [Hung et al. 2005]. Dichloran glycerol (DG18) can be used to select for those 

fungi that are xerotolerant [Flannigan et al. 2011; Hocking and Pitt 1980; Macher 1999]. 

Temperature and incubation time can also be used to select for specific fungal bioaerosols 

such as Aspergillus fumigatus which are capable of growth within human hosts [Flannigan 
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 et al. 2011; Hung et al. 2005]. A selection of nutrient media and growth conditions that can 

be used for viable fungal culture can be found in Hung et al. [2005], Flannigan et al. [2011] 

and Macher [1999].

b. Sample preparation for viruses

Because viruses are obligate intracellular parasites, special precautions must be taken in an 

effort to minimize damage to the collected virus-laden aerosol. Environmental factors 

such as humidity, temperature and gas composition of the air can significantly impact the 

infectiousness of a virion and should be monitored closely [Ijaz et al. 2016]. Several studies 

have shown that the inactivation of an airborne virus is directly related to the relative 

humidity and temperature [Weber and Stilianakis 2008]. In one study, high humidity 

levels caused a loss of infectious influenza virus from simulated coughs [Noti et al. 2013]. 

Similarly, using a ferret animal model, Lowen et al. [2007] demonstrated a correlation 

between airborne transmission of influenza and the relative humidity and temperature. 

Through the use of an ozone-oxygen delivery system, researchers were able to show that 

ozone-mediated reactive oxygen species (ROS) caused lipid peroxidation and subsequent 

damage to the lipid envelope and viral capsid [Murray et al. 2008]. Such studies highlight 

the importance of collecting viral aerosols under optimal environmental conditions and, 

when possible, minimizing the detrimental effects of environmental factors on collected 

samples.

Air sampling techniques also may cause damage to the virus and compromise analysis. 

Before collecting viral aerosols, the hardiness of the target virus must be taken into 

account. Currently, there are over 200 known respiratory viruses that fall under one family 

of DNA viruses (Adenoviridae) and four families of RNA viruses (Orthomyxoviridae, 

Paramyxoviridae, Picornaviridae and Coronaviridae) [Abed and Boivin 2006]. While all 

viruses package their genome in a protective protein coat known as the capsid, some 

viruses also possess a lipid bilayer envelope that, as the name implies, surrounds the viral 

capsid. Once outside the host, the viral envelope is highly sensitive to desiccation, 

temperature fluctuations and readily undergoes degradation. Variations in temperature 

can greatly affect viral enzymatic activity and nucleic acid stability [Tang 2009]. As noted 

earlier, viruses with lipid envelopes tend to survive better at low humidity, while nonenveloped viruses survive better at high humidity [Tang 2009]. Also, RNA viruses are 

inherently more unstable than DNA viruses due to the presence of the 2’-hydroxyl group 

on the ribose sugar molecule of RNA that is susceptible to base-catalyzed hydrolysis and 

degradation. Therefore it is critical that collection methods do not disrupt the lipid 

membrane and/or compromise the integrity of nucleic acids. Impairment of either the 

lipid envelope or nucleic acids can significantly impact detection of the viral aerosol and 

lead to false negatives.
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Sampling and Characterization of Bioaerosols 

 To optimize collection efficiency while maintaining infectiousness of the viral aerosol, 

researchers must be discriminating when deciding on what type of aerosol sampler to use 

and how long the sampling collection period should be. As discussed in Section 3, some 

commercially available bioaerosol sampling devices exist, each of which possesses unique 

collection properties. In a review by Verreault et al. [2008], liquid impingers were found to 

be the most effective sampling devices for capturing small viral particles while maintaining 

virion integrity and infectiousness [Verreault et al. 2008].

While liquid impingement preserves and maintains viral integrity (in comparison to dry 

impaction), the type of collection medium must also be considered. With culture-based 

identification methods, it is of utmost importance to maintain the stability of the collected 

viruses while using cell-culture compatible media. The type of liquid medium, as well as 

the volume used, are important variables to consider. Virus collection and transport media 

are typically isotonic solutions with a buffer to control the pH, protein to protect the virus, 

and antibiotics to prevent microbial growth. If used in an impinger, the viral collection 

media may also include an antifoaming agent. Specimen handling, storage time and 

storage temperature can significantly impact the integrity of sample analysis. Collection 

and storage under suboptimal conditions can result in viral inactivation and degradation 

of nucleic acids. Bioaerosol samples containing viruses should be processed as soon as 

possible after collection. They should be refrigerated or frozen and transported as quickly 

as possible. The stability and retention of viability depend upon the virus [Johnson 1990].

7 Identification of culturable bioaerosols

Identification of the microbial taxa is a critical element in the determination of the viable 

bioaerosol load in an industrial or occupational environment. The science of classification, 

especially the classification of living forms, is called taxonomy. The objective of taxonomy is 

to classify living organisms to establish the relationship between one group of organisms and 

another, and to differentiate between them based on phenotypic and genotypic characteristics. 

The identification of viable fungal bioaerosols has been challenging due to the confusion of 

current nomenclature [Flannigan et al. 2011]. As a result, investigators often use synonymous 

names that over time have been placed in another group. Familiarity with taxonomy and 

nomenclature is critical when undertaking assessments of the viable microbial burden in an 

environment [Flannigan et al. 2011]. Several criteria and methods for the classification of 

culturable microorganisms are briefly discussed in the following subsections. Besides using 

these methods, the nonviable and non-culturable methods of identification discussed in 

Section 9 may also be used in combination with these viable methods.

Classical microbiology includes general methods for classifying or identifying 

microorganisms. The least specific of these is the observation of growth characteristics. 
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 Growth characteristics include the appearance of the microorganisms in a liquid medium, 

colony morphology on solid medium, pigmentation, and arrangement of reproductive 

structures such as fungal sexual or asexual spores.

a. Bacteria 

Bacteria are prokaryotes with distinguishing morphological characteristics that include the 

cell shape, cell size, arrangement of cells, and the presence or absence of flagella, capsules, 

or endospores. Simple and differential staining may be performed on bacteria to enhance 

visualization and to aid in grouping and identification [Tortora et al. 2013]. In simple 

staining, a single basic dye is used that highlights the cellular morphology. Stains such as 

methylene blue, carbolfuchsin, crystal violet, or safranin may be used for bacteria. 

A differential stain distinguishes among structures or microorganisms based on varying 

reactions to the staining procedure. Two examples of differential stains are the Gram stain 

and the acid-fast stain. In Gram staining, bacteria are stained and then washed with 

alcohol. Gram-positive bacteria possess a cell wall composed of a relatively thick 

peptidoglycan layer and teichoic acids, which retains the dye complex. Gram-negative 

bacteria possess a cell wall composed of a thin peptidoglycan layer and an outer membrane 

which consists of lipoproteins, lipopolysaccharides, and phospholipids, and do not retain 

the dye complex when washed [Tortora et al. 2013]. A few of the commercially available 

identification kits require a Gram-stain prescreening to assure that the correct reagents are 

used. Acid-fast stains are used for some species of bacteria, particularly those of the genus 

Mycobacterium, which do not stain readily. In the acid-fast staining process, the 

application of heat facilitates the staining of the microorganism [Tortora et al. 2013].

b. Legionella 

Bacteria that are placed in the genus Legionella, are the etiological agents of pulmonary 

infections called Legionellosis [Fields 2002]. Legionella pneumophila (Figure 1B) is the 

most widely known species that has been implicated in Legionnaires’ disease, which can 

result in pneumonia. Milder illness with fever and body aches is referred to as Pontiac 

fever. Bacteria placed in this genus consist of Gram negative rods and are associated with 

freshwater in the environment [ASTM 2015; Macher 1999]. Exposure to warm 

temperatures (25-42°C) can result in the growth and proliferation of the bacteria, a 

problem that has emerged in water and air handling systems within the indoor built 

environment [Hung et al. 2005]. Growth and persistence within Protista have also been 

reported [Hung et al. 2005]. For health care facilities, ASHRAE Guideline 12–2000 

recommends storing and distributing cold water at <20°C (68°F), whereas hot water 

should be stored at >60°C (140°F) and circulated with a minimum return temperature of 

51°C (124°F). In other settings, hot water should be stored at ≥40°C (≥120°F) [ASHRAE 
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 2000]. Building air conditioning cooling towers, humidifiers, or structures used for 

bathing such as hot tubs are particularly susceptible to Legionella amplification. Legionella 

can be aerosolized within water droplets via abiotic disturbance mechanisms and 

disseminated into the breathing zone of the subject. Airborne levels lower than 10 

CFU/mL have been associated with Legionnaires’ disease [ASTM 2015; Demirjian et al. 

2015; Hung et al. 2005].

Since the identification of L. pneumophila and association with Legionnaires’ disease in 

1976, there have been many studies that have focused on a variety of approaches to detect 

and mitigate this bacterial species from the built environment. These approaches are 

broadly reviewed in Hung et al. [2005]. Along with industrial hygiene practices and 

building maintenance, environmental monitoring and surveillance programs are critical to 

ensure effectiveness of employed engineering controls and maintenance/disinfection 

programs [ASTM 2015]. ASTM International has published a standard for inspecting 

water systems and investigating outbreaks [ASTM 2015]. The CDC has also published a 

sampling procedure [CDC 2015].

In 2015, ASHRAE published a consensus standard for the primary prevention of 

Legionnaires’ disease in building water systems [ASHRAE 2015]. Similar environmental 

assessment methods are utilized in maintenance programs and outbreak cases. In this 

approach, bulk water samples are typically collected (250 mL to 1 L for non-potable and 

1000 mL for potable water), concentrated via filtration, resuspended, and then plated on a 

growth medium (such as buffered charcoal yeast extract media) to enable the propagation 

and identification of Legionella spp. [CDC 2015]. Samples may also be direct plated, acid 

treated, or heat treated to enhance the recovery of the bacterium. Colonies represent the 

viable fraction of Legionella in a water sample and these colonies are quantified and 

reported as CFU/mL

 

In addition to viable culture-based approaches, molecular-based methods such as PCR as 

well as antibody-based methods have been developed to enable the detection of Legionella. 

Air sampling is not considered a reliable method for Legionella surveillance in the built 

environment [Hung et al. 2005]. 
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 Using an environmental microbiology laboratory with expertise in propagating Legionella

spp. is important when evaluating Legionella contamination of water systems within the 

built environment. A number of laboratories participate in the CDC’s Environmental 

Legionella Isolation Techniques Evaluation (ELITE) Program. Participation in the 

program is voluntary and enables laboratories to test their proficiency in Legionella

isolation and identification techniques against standardized samples. A list of ELITE 

member laboratories can be accessed at

https://wwwn.cdc.gov/elite/Public/MemberList.aspx. 

c. Fungi 

In general, culturable fungi are classified by colony features including the septation of 

hyphae and colony morphological phenotypes, including pigmentation and the 

presentation of asexual and sexual spores on hyphae. Stains such as Calberla’s solution, 

lactophenol cotton blue, periodic acid-Schiff stain, Grocott’s methenamine silver stain, 

and calcofluor white may be used in combination with potassium hydroxide (10% KOH) 

to resolve these colony structures using microscopic-based approaches [Hung et al. 2005]. 

The identification and classification of fungal colonies should be performed by an 

examiner that is skilled in microbiology and mycology. A number of commercial labs that 

employ examiners skilled in the identification of microorganisms encountered in indoor 

and occupational environments are accredited by the AIHA Environmental Microbiology 

Laboratory Accreditation Program (EMLAP). 

In addition to viable culture-based approaches, biochemical, physiological, and nutritional 

tests for bacteria and fungi can be used [Flannigan et al. 2011]. These testing strategies 

offer identification based on numerous variables including cell wall constituents, pigment 

biochemicals, storage inclusions, antigens, optimum temperature and temperature range, 

the effect of oxygen on growth, pH tolerance, osmotic tolerance, salt requirement and 

tolerance, antibiotic sensitivity, energy sources, carbon sources, nitrogen sources, 

fermentation products, and modes of metabolism (autotrophic, heterotrophic, 

fermentative, respiratory). As a rule, batteries of such tests, rather than any one individual 

test, are used to identify or classify microorganisms. A few commercially available test 

batteries are discussed briefly in the section on biochemical approaches.
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8 Enumeration of culturable bioaerosols

a. Enumeration of bacteria and fungi

The total concentration of culturable airborne microorganisms in a sample is determined 

by collecting the bioaerosol sample on a culture plate or plates (or inoculating culture 

plates with a bioaerosol sample), incubating the plates, and dividing the number of 

colonies observed on the culture plates by the volume of air sampled. Note that, as 

discussed in section 2, the number of colonies counted on an agar plate from a bioaerosol 

impactor must be adjusted using a positive-hole correction factor to correct for multiple 

microorganisms depositing beneath an impactor hole [Andersen 1958; Leopold 1988; 

Macher 1989]. A colony is defined as a macroscopically visible growth of microorganisms 

on a solid nutrient medium. Concentrations of culturable bioaerosols collected during air 

sampling are normally reported as colony forming units (CFU) per unit volume of air 

[Eduard et al. 2012]. CFUs also can be determined from samples collected in a swab or 

dust sample collected from the floor or area of contamination [Hung et al. 2005].

Often, it is difficult to identify multiple colonies at one location on a plate because of the 

lack of differential colony morphology [Burge et al. 1977]. In addition, some organisms 

produce large, spreading colonies while others produce microcolonies. Analysis of plates 

containing multiple types of microorganisms can be difficult because the chemicals 

secreted by one microorganism might inhibit the growth of other microorganisms at that 

same location [Burge et al. 1977]. The morphology of the colony of one microorganism 

also may completely obscure that of another, and a fast-grower might obscure a slowgrower.

b. Enumeration of viruses

Before the advent and mainstreaming of molecular-based detection methodologies, cell 

culture-based methods and serological assays were considered the gold standard for the 

detection of viral pathogens. Typically, through the use of commercially available 

immortal cell lines, researchers can screen collected bioaerosols by inoculating cells and 

looking for common cytopathic effects (CPEs) such as rounding of infected cells, fusion 

with adjacent cells and lysis of cells. Examples of well-known cell lines that are routinely 

used in viral diagnostics include primary rhesus monkey kidney (RhMK) cells, primary 

rabbit kidney cells, human lung fibroblasts (MRC-5), human epidermoid carcinoma cells 

(HEp-2), human lung carcinoma cells (A549) and Madin Darby kidney cells (MDCK) 

[Leland and Ginocchio 2007]. Selection of the appropriate cell line is based on the 

specimen source and the suspected causal viral pathogen. Certain viral pathogens may 

require several cell passages before CPEs can be observed. More information on 

enumeration assays for viable viruses can be found in standard virology reference texts, 

such Principles of Virology [Flint et al. 2009b] and Fields Virology [Fields et al. 2007].
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Viral plaque assay

A widely used approach for detecting viral pathogens and quantifying viral titers is the 

viral plaque assay (VPA) [Condit 2007]. Under biological safety controls, the 

appropriate cell line is propagated and plated, usually in a 6-well format, at a 

concentration at which cells form a monolayer. The cell monolayer is next treated for a 

defined period (30-60 minutes) with a specific volume (0.1 mL to 1.5 mL) of the 

collected bioaerosol and incubated at the specified temperature and CO2 levels for 24 

to 72 hours. Throughout the incubation period, cells are routinely inspected and CPEs 

are documented. Upon completion of the incubation period, cells are chemically fixed, 

stained and plaques (zones of cellular clearing) are enumerated. The final 

concentration of the collected viral aerosols is calculated based on the number of 

plaques, dilution of the inoculum and volume plated, and is expressed in plaque 

forming units per mL (PFU/mL]). While the VPA is a cost-effective method of 

assessing sample viral loads, results can take anywhere from 3-5 days. Other 

limitations such as the inability to detect low viral titers and inactivated 

(noninfectious) virus and the failure of some viruses to form plaques, may 

compromise detection and underestimate the viral loads in an aerosol sample. 

Likewise, common indoor and outdoor contaminants (such as fungi and bacteria) can 

impair the VPA by disrupting the cellular monolayer or outcompeting for nutrients in 

the cell culture medium.

Tissue culture infectious dose assay

Another cell culture-based approach to identifying viral aerosols is the Tissue Culture 

Infectious Dose assay (TCID50), also known as an endpoint dilution assay [Condit 

2007]. As with the VPA, select cells are plated at a desired concentration in a 96-well 

format and inoculated with serial dilutions of the collected sample. Following a 

specified incubation period, cells are examined for CPEs. The TCID50 is defined as the 

dilution of virus required to infect 50% of the cell culture wells [Reed and Muench 

1938]. Based on the number of cells that are infected at the designated virus dilution, 

viral titers are mathematically calculated. Limitations of the TCID50 are similar to 

those observed with the VPA.

Immunofluorescence antibody (IFA) assays

To enhance viral detection and quantify viral loads, immunofluorescence antibody 

(IFA) assays (direct or indirect) are frequently used in combination with cell culturebased methods [Flint et al. 2009a; Tortora et al. 2013]. By combining infected cells 

with a fluorescently-labeled, antigen-specific antibody, it is possible to increase 

detection levels without the lengthy incubation periods that are typically necessary 

with VPAs and TCID50 assays.
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c. Interpretation of data

In industrial hygiene surveys that evaluate bioaerosols, indoor bioaerosol levels are usually 

compared to outdoor levels or to a control area. In general, indoor bioaerosol levels are 

lower than outdoor levels, and the taxa are similar [ACGIH 1989; Burge et al. 1977; Hung 

et al. 2005; Macher 1999; Solomon et al. 1980]. However, elevated indoor bioaerosol levels 

may be a sign of dampness, water infiltration, or microbial contamination [Hung et al. 

2005]. In 2010, The ACGIH published a variety of occupational exposure limits for 

aerosols that are derived from biological material in specific industries and include 

subtilisins derived from Bacillus subtilis, as well as cotton, grain, flour, wood, and organic 

dusts [ACGIH 2015; Eduard et al. 2012]. To date, no occupational exposure limits for 

specific fungal bioaerosols exist and these typically fall under particulate matter not 

otherwise regulated (10 mg/m3 for inhalable dust; [ACGIH 2015; Eduard et al. 2012]. 

Proposed limits developed in other regions of the world are provided in Eduard et al. 

[2012]. 

Although the quantification of viable fungal propagules can provide helpful datasets to 

evaluate differences between indoor and outdoor fungal diversity, the interpretation of 

results should be evaluated closely. Total fungal exposure will be underestimated as nonviable fungal bioaerosols are not captured in the analysis [Eduard et al. 2012]. Fungal 

genera, including Cladosporium, Alternaria, and Epicoccum, and Basidiomycetes are 

predominantly localized in outdoor environments and the presence of elevated 

concentrations may be an indicator of indoor fungal contamination [Hung et al. 2005]. 

Similarly, the presence of certain hydrophilic species including Stachybotrys chartarum

and Aspergillus versicolor, and Chaetomium globosum may be signs of indoor fungal 

contamination and may require immediate inspection [Flannigan et al. 2011; Hung et al. 

2005]. 

Where local amplification and dissemination of bacteria have not occurred in an occupied, 

indoor environment, Gram-positive cocci (e.g., Micrococcus and Staphylococcus) are 

normally dominant [Morey et al. 1986]. Airborne human skin scales and respiratory 

secretions may contain Gram-positive cocci [ACGIH 1989; Hung et al. 2005]. Detection of 

high levels of these microorganisms may be an indication of over-crowding and 

inadequate ventilation. Indoor air that tests high for Gram-negative bacteria indicates a 

need to identify and eliminate the source of contamination. Concentrations ranging from 

4,500-10,000 CFU/m3 have been suggested as the upper limit for ubiquitous bacterial 

aerosols [ACGIH 1989; Nevalainen 1989]. These exposure limits, however, do not apply to 

pathogenic microorganisms. Actinomycetes (mesophilic and thermophilic) are commonly 

found in agricultural areas. Their presence in indoor environments is an indicator of 

contamination [ACGIH 1989; Banaszak et al. 1970; Lacey and Crook 1988]. Thermophilic 
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 Actinomycetes at concentrations above 70 CFU/m3 in an affected person's work area have 

been regarded as the threshold for triggering remedial action [Otten et al. 1986].

9 Sample analysis methods for non-viable and 

non-culturable bioaerosols

The collection and classification of nonviable and non-culturable microorganisms cannot be 

performed by using viable culture methods. A large proportion of fungal bioaerosols are nonviable and would not grow and proliferate on nutrient media [Eduard et al. 2012]. This 

fraction of the bioaerosol load is equally important to assess in industrial hygiene surveys that 

investigate the role of personal bioaerosol exposure on respiratory health [Brasel et al. 2005; 

Green et al. 2011; Mitakakis et al. 2003]. These bioaerosols can also contain antigens, 

allergens, microbial volatile organic compounds and even mycotoxins [Brasel et al. 2005; 

Eduard et al. 2012; Green et al. 2011; Green et al. 2006b]. Identification of nonviable or nonculturable microorganisms or components of microorganisms (such as cell wall fragments) 

can be performed using a variety of other available assessment strategies such as microscopy, 

immunoassays and, more recently, molecular biology techniques [Afanou et al. 2015; Brasel et 

al. 2005; Eduard et al. 2012; Flannigan et al. 2011; Green et al. 2011; Hung et al. 2005; Macher 

1999; Rittenour et al. 2012]. 

Microscopy includes a variety of approaches that utilize bright-field, light, phase contrast, 

fluorescence or even electron-based approaches [Eduard et al. 2012; Macher 1999]. These 

methods enable the enumeration of both viable and nonviable microorganisms [Macher 1999] 

as well as other non-culturable bioaerosols including cell wall fragments, plant pollen and 

pteridophyte and bryophyte spores [Green et al. 2011; Rittenour et al. 2012]. These 

approaches provide a platform to visualize particle morphology and to identify reproductive 

fungal structures of individual genera that are based on a combination of propagule 

phenotypes [Flannigan et al. 2011; Macher 1999]. However, these approaches can be 

confounded by observer bias, especially when it comes to differentiating bioaerosols that 

contain similar morphological phenotypes such as amerospores (e.g. Aspergillus conidia) 

[Flannigan et al. 2011; Hung et al. 2005]. The ASTM has published a standard method for the 

use of optical microscopy to categorize and quantify fungal structures in samples collected by 

inertial impaction [ASTM 2009].

To overcome these methodological challenges, alternative methods based on the 

quantification of bioaerosol biomarkers (proteins or DNA) have been developed that enable 

the quantification of these bioaerosol sources [Eduard et al. 2012]. These include a variety of 

assessment methods that can be used to qualitatively and quantitatively assess bioaerosol 

exposure by detecting cell wall components, proteins, carbohydrates, or oligonucleotides (e.g. 
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endotoxin or β-glucan) [Eduard et al. 2012]. Other chemical and proteomic methods 

including HPLC, flow cytometry, and mass spectrometry-based approaches can also be used 

to detect and quantify cell wall components such as microbial volatile organic compounds and 

mycotoxins. These approaches have been reviewed more extensively by Flannigan et al. 

[2011]. 

Several modifications of classical biochemical procedures have been used in recent years to 

facilitate inoculation of media, decrease the incubation time, automate the procedure, and 

systematize the determination of species based on reaction patterns. Historically, clinical 

microbiological techniques have been used for analysis of environmental samples. However, 

clinical strains and environmental isolates may differ, requiring modification of clinicallybased techniques.

a. Microscopy

Bright-field or light

In bright-field or light microscopy, an ordinary microscope is used for the

morphological observation and sizing of sampled bioaerosols. Visible light from an

incandescent source is used for illumination and the specimen appears against a bright

backfield. Objects smaller than 0.2 μm cannot be resolved. The image contrast

(visibility) decreases as the refractive index of the substance/microorganism under

observation and the mounting medium become similar. To maximize the contrast, the

mounting medium should have the same refractive index as glass or the immersion oil.

Membrane filters are often "cleared" by using the appropriate immersion oil or acetone

vapor/triacetin combination. This method is commonly used to observe various

stained specimens and to identify and count viable and non-viable bioaerosols. In

addition, pollen grains are often identified and enumerated in this manner [Eduard et

al. 1990].

Collection of fungal bioaerosols onto an adhesive surface followed by microscopic

identification based on the morphological characteristics of the spores (size, shape,

septation etc.) is another common method of assessment. This non-viable method

overcomes limitations introduced in viable analyses and many genera can be

differentiated based on differences in spore morphology [Eduard et al. 2012; Macher

1999]. Microscopic examination of fungi captured on filters or an adhesive tape are

divided into seven spore morphological characteristics and include amero-, didymo-,

helico-, stauro-, dictyo-, phragmo-, and scoleco- spores [Kendrick 2000]. Amerospores

are the most common spore morphology encountered in air samples and are the most

challenging to differentiate taxonomically [Kendrick 2000]. Amerospores are usually

placed in a group represented as Aspergillus/Penicillium group [Hung et al. 2005].
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Many other common environmental fungal bioaerosols share similar morphologies 

which can make taxonomic placement challenging for the untrained or inexperienced 

observer [Eduard et al. 2012]. Typical magnification used in the assessment of fungal 

propagules ranges from 400-1000X. A standard operating procedure for the 

assessment of microscopic non-viable samples is presented in Hung et al. [2005] and 

by ASTM [2009]. 

The confounding factors associated with traditional fungal exposure assessment 

methods have limited our understanding of the spectrum of fungal bioaerosols in 

industrial and occupational environments. Measures using these approaches also 

cannot be acquired in real time. Furthermore, identifying and quantifying the 

complete diversity of fungal bioaerosols using a standardized methodology is critical 

in the determination of fungal bioaerosols within occupational environments [ASTM 

2009; Hung et al. 2005].

Phase contrast

Phase-contrast microscopy is used when the microorganism under observation (e.g., 

Escherichia coli) is hyaline and an alternative mounting medium is not possible. As 

light passes through the specimen, variations in the index of refraction of the 

components cause phase shifts in the light. A phase-contrast microscope uses a special 

condenser and diffraction plate that cause these phase shifts to appear as differences in 

brightness and contrast. One cannot see an object exactly matching the refractive 

index of the mounting liquid; however, very slight differences produce visible images. 

This type of microscope is commonly used to provide detailed examination of the 

internal structures of living specimens; no staining is required.

Fluorescence

Fluorescence microscopy uses an ultraviolet or near-ultraviolet source of illumination 

that causes fluorescent compounds in a specimen to emit light. Fluorescence 

microscopy for the direct count of microorganisms has been described in a number of 

studies [Eduard et al. 2012]. Direct-count methods to enumerate microorganisms 

(especially bacteria) have been developed using fluorescence microscopy and some 

stains such as acridine orange, fluorescein isothiocyanate (FITC) and 4’,6-diamidino2-phenyl-indole (DAPI) [Macher 1999; Thermo Fisher Scientific 2014]. Utilization of

stains such as calcofluor white can resolve fungal spores and hyphal structures

[Haghani et al. 2013]. This stain binds to chitin; however, other plant-derived and

insect bioaerosol sources (e.g. dust mite, plant pollen) may also be resolved using this

stain. Viability stains also have been developed and are available commercially for the

detection of viable fungi and bacteria bioaerosols in collected air samples [Thermo

Fisher Scientific 2014].
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Electron

Electron microscopy consists of a beam of electrons that enable structures smaller than 

0.2 μm, such as viruses, to be resolved. Scanning electron microscopy (SEM) and, 

more recently, field emission scanning electron microscopy (FESEM), are approaches 

used to study the surface features of prokaryote and eukaryote cells as well as viruses 

(usually magnified 1,000-10,000X). These bioaerosols are immobilized onto a semisolid filter submicron membrane or in the form of a liquid suspension and a threedimensional image of the area is generated. Images from SEM can provide vital 

information about the size, morphology and concentration of the collected bioaerosol 

[Afanou et al. 2014; Eduard et al. 2012]. However, SEM does not provide information 

on viability of the collected bioaerosol. FESEM has been recently used to resolve fungal 

fragments that are produced from fungal colonies following abiotic disturbance 

[Afanou et al. 2015; Afanou et al. 2014]. Transmission electron microscopy can also be 

used to examine viruses or the internal ultrastructure in thin sections of cells (usually 

magnified 10,000-100,000X), although the image produced is not three- dimensional. 

Compared to other methods of assessment described in this chapter, SEM and 

FESEM-based approaches require a highly trained technician to obtain images from 

bioaerosols captured on filter membranes.

b. Endotoxin assays

The lipopolysaccharide endotoxin is a virulence factor possessed by all Enterobacteriaceae

(as well as other Gram-negative bacteria) that is found in the outer membrane of the cell

wall. Airborne endotoxin has been found in high concentrations in agricultural, industrial,

and office environments [Eduard et al. 2012; Milton et al. 1990; Rylander and Vesterlund

1982; Singh et al. 2011b]. Individuals may experience disseminated intravascular

coagulopathy, respiratory tract problems, cellular and tissue injury, fever, and other

debilitating problems. Endotoxin can be detected in air samples collected on glass filters

using the Limulus amebocyte lysate (LAL) assay [Eduard et al. 2012]. This assay uses

amebocytes from the blood-like circulating fluid of the Limulus polyphemus (horseshoe

crab). After exposure to the lysed amebocyte cells, the chromogenic version of the LAL

enables endotoxins to be quantified [Eduard et al. 2012]. Laboratories use this assay to test

for contamination by Gram-negative bacteria [Baron and Finegold 1990].

Although widely used, endotoxin aerosol measurement techniques lack comparability 

between results obtained in different laboratories because of differing sampling, 

extraction, and analytical methods [Jacobs 1989; Milton et al. 1990; Olenchock et al. 1983; 

Rylander and Vesterlund 1982] and water-insoluble endotoxins are not detected [Eduard 

et al. 2012]. A monoclonal antibody assay has also been developed but it is less sensitive 
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 than the LAL method. Similarly, chemical-based approaches are available to detect 

endotoxins including gas chromatography-mass spectrometry [Eduard et al. 2012].

c. Biochemical analysis methods

Because of the high frequency of isolation of Gram-negative rods in clinical settings, 

several commercial multi-test systems have been developed for identification of members 

of the family Enterobacteriaceae and other pathogenic microorganisms. These 

microorganisms are indistinguishable except for characteristics determined by detailed 

biochemical testing. These systems require that a pure culture be examined and 

characterized. A list of some commercially available identification kits is provided in Table 

III. All of these multitest systems have documented accuracies greater than 90% in clinical 

settings [Baron and Finegold 1990; Koneman 1988]. For fungi, API (Analytab Products, 

Plainview, NY) and BIOLOG can also be used to differentiate yeasts based on the 

respective biochemical and physiological profiles [Flannigan et al. 2011].

d. Chemotaxonomic approaches 

Cellular fatty acids (CFA) of bacteria are structural in nature, occurring in the cell 

membrane or cell wall of all bacteria. When the bacteria are grown under standardized 

growth conditions, the CFA profiles are reproducible within a genus, down to the 

subspecies or strain level in some microorganisms. The Sherlock Microbial Identification 

System (MIS), developed by MIDI (Newark, DE), provides a chromatographic technique 

and software libraries capable of identifying various microorganisms based on their CFA 

composition [Sasser 1990a; Sasser 1990b]. The chromatographic technique is also known 

as gas chromatography fatty acid methyl ester analysis (GC-FAME). MIS has a database 

containing the analysis libraries for culturable Gram-negative and Gram-positive bacteria, 

and yeasts. In a comparison study [Amy et al. 1992], only 8 of 18 isolates, identified by 

either API multitest or MIDI MIS, were identified accurately using BIOLOG multitest. A 

prototype method for extracting and analyzing fungi is currently being distributed by 

MIDI.

e. Chemical-based approaches

A variety of chemical-based approaches are available for the detection and quantification 

of bacteria and fungi in environmental samples [Flannigan et al. 2011; Hung et al. 2005]. 

Common approaches include thermal desorption - gas chromatography - mass 

spectrometry (TD-GC-MS), high performance liquid chromatography, gas 

chromatography-tandem mass spectrometry, and matrix-assisted laser 

desorption/ionization time of flight (MALDI-TOF) mass spectrometry [Flannigan et al. 

2011]. These methods require the formation of a library of markers or spectral signatures 

that are used to discriminate between various prokaryote and eukaryote species. Examples 
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of spectral signatures that have been used for the detection of fungi include microbial 

volatile organic compounds (mVOCs), mycotoxins, ergosterol, 3-hydroxy fatty acids, 

muramic acid as well as intracellular and extracellular proteins. NIOSH Method 2549 is a 

TD-GC-MS based approach that allows for the detection of mVOCs in environmental 

samples [NIOSH 2003d].

f. High performance liquid chromatography

High performance liquid chromatography (HPLC) is commonly used for bioaerosol

“fingerprinting” and biomass determination. Techniques such as proteomics and

identifying variations in chromatographic patterns can be used to determine the source of

airborne material. For example, ergosterol has been used for decades to detect fungal

contamination [Seitz et al. 1979] and even determine taxonomy [Axelsson et al. 1995;

Pasanen et al. 1999; Schnurer 1993]. Keratin analysis can be used to identify bioaerosols

derived from vertebrates and possibly the habitat within an environment [Staton et al.

2013]. Detection can be as straightforward as using UV absorbance or as complex and

specific as employing an ion trap mass spectrometer.

HPLC can be adventitious compared to other methods of analysis; it is an established 

technology, fairly inexpensive after initial equipment costs, fast and accurate. Its 

disadvantages include a lack of specificity inherent with detectors using UV absorbance 

(especially at lower wavelengths), and that complex matrices associated with bioaerosols 

can prove to be troublesome and may require multi-step enhancement procedures such as 

solid-phase extraction. Buffered solvent systems are sometimes required, which can be 

technically difficult to use.

Mycotoxins

Fungal contamination is a concern in food production because it can modify the 

nutritional content of feed and cereal grains and introduce potentially adverse

mycotoxins. Before the use of HPLC, methods of identifying fungal contamination 

were time consuming or missed non-viable organisms which still contributed to the 

biomass [Seitz et al. 1979]. HPLC can be used to detect ergosterol, which is a structural 

sterol nearly universally present in fungi but not naturally present in grains [Pasanen 

et al. 1999]. HPLC has also been used to detect ergosterol, mannitol and arabitol in 

bioaerosols [Buiarelli et al. 2013].

Ergosterol is extracted using a liquid-liquid extraction and concentrated. The clean 

samples are then analyzed by HPLC with UV detection. The ergosterol UV spectrum 

varies significantly from the UV spectrum of higher plant sterols, making it specific to 

fungal contaminants. Though the method is still fairly time consuming, it eliminates 
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the need for fungal culturing and can detect the presence of non-viable fungi. When 

combined with other sterols this information can be used to help determine fungal 

species [Schnurer 1993]. 

In addition to using sterols to identify fungal contamination, mycotoxins can also be 

analyzed by liquid chromatography with mass spectrometry (LC-MS). Identification of 

specific mycotoxins can help identify the species of fungi present [Bennett and Klich 

2003; Castillo et al. 2016]. 

Mycotoxins can play a role in indoor air quality (IAQ), food safety and possibly 

bioterrorism. The use of LC-MS as a screening tool to identify mycotoxins can reduce 

the use of more intensive molecular techniques for identification and quantitation. 

When using mass spectral analysis it is important to have a reliable and accurate 

database for identification and a qualified analyst to correctly interpret data.

Other biomolecules

More recently, HPLC has been proposed as a forensic tool to identify and track 

vertebrate species using keratin profiles. Like ergosterol in fungi, keratin is found 

mainly in dander left by vertebrates [Plowman 2007]. If patterns can be established, it 

may be possible to identify what species had been present in a specific dwelling and 

possibly even track movements [Staton et al. 2013]. Bacterial contamination can also 

be tracked using endotoxin analysis and bacterial peptidoglycan fingerprinting [Staton 

et al. 2013].

Sample preparation and enhancement

Bioaerosols can have complex matrices with many interfering constituents. Ultraviolet 

absorption detectors are fairly inexpensive and straightforward to use, but they can 

suffer from a lack of specificity and sensitivity. Although mass spectrometry detectors 

do not suffer from a lack of specificity or sensitivity, a dirty sample can still present 

challenges. 

Another potential pitfall is the presence of large particles in bioaerosol samples. In 

general, analytical HPLC systems and detectors use small diameter tubing and small 

orifice injectors that are easily clogged or contaminated by particles. Most analytical 

systems have some sort of less-expensive trapping or pre-column that can be sacrificed 

in order to spare the more expensive analytical columns. However, trap columns are 

still very much an expense and should only be considered if other options are not 

available. A multitude of cleanup procedures are available to lessen or eliminate 

problems due to particles, the most common of which are simple centrifugation and 
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filtration. Both methods will lessen the likelihood of clogging or damaging the system, 

but neither offers target analyte enrichment.

Liquid-liquid extraction

In order to enrich analytes, samples can be concentrated, chemical interferences can be 

removed, or a combination of both methods can be used. One of the oldest methods of 

enrichment is a liquid-liquid extraction, which separates analytes based on relative 

solubility in a given solvent [Koncsag and Barbulescu 2011]. In general, two 

immiscible solvents are mixed, one solvent containing the whole extract (called the 

“feed”) and one that ideally solubilizes the analyte of interest preferentially. Based on 

solubility, the chemical constituents will either stay in the feed solvent or partition into 

the other solvent. Once the solvents are allowed to phase-separate, the solvent 

containing the enriched analyte (called the raffinate) is removed. 

Liquid-liquid extraction is usually done using an aqueous solvent and an organic 

solvent. Solvent selection is critical and can be difficult. The goal of the extraction is to 

choose a solvent that leaves behind as much of the interfering matrix as possible in the 

feed solvent while also being able to preferentially solvate the analyte of interest. The 

solvent also must be compatible with the analytical instrumentation. Liquid-liquid 

extractions tend to use large amounts of solvent, which can result in the analyte of 

interest being below analytical levels of detection and/or levels of quantitation. Many 

organic solvents can be easily concentrated by evaporation, but this can pose problems 

with labile or volatile analytes.

Solid-phase extraction

Another option that can help avoid these pitfalls is the use of solid-phase extraction 

(SPE) [Sigma-Aldrich 1998]. SPE exploits the analyte affinity (or lack thereof) for a 

solid material packed inside a column. Sample enrichment can occur by the column 

retaining interfering compounds and the analyte of interest passing through, or by the 

column retaining the analyte and the interfering chemicals passing through. 

Concentration of the analyte of interest can be achieved by eluting the analyte in a 

smaller volume of solvent, and filtration can be achieved simultaneously. SPE may 

make it possible to achieve analyte concentration and avoid potential losses that could 

arise from concentrating a large volume of solvent. Appropriate solvent selection is 

also critical to successful SPE.
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g. Immunoassays

The immunoassay is an analytical technique for measuring a targeted antigen, which is

also referred to as an analyte. A critical component of the immunoassay is the antibody or

ligand, which binds a specific antigen or binding site. The binding of the antibody or

ligand forms the basis for the immunoassay, and numerous formats have been devised

which permit visual or instrumental measurements of this reaction. Antibodies include

either monoclonal or polyclonal antibodies and these are commonly employed to detect

organisms by binding to antigens, usually proteins, polysaccharides or other cell wall

components [Hung et al. 2005; Macher 1999]. The analysis is usually performed following

extraction of the analytes to form a heterogeneous matrix. In most immunoassays, there is

little need for extensive sample cleanup. Following the development of

radioimmunoassays, many immunoassays that use monoclonal antibodies are now readily

available from commercial sources, permitting laboratories to use standardized

immunoassays or rapidly develop in-house immunochemical assays. In addition,

commercially available immunoassays or multiplex platforms are available to quantify a

variety of indoor or occupational bioaerosol sources [King et al. 2013]. Some of the more

widely used immunoassay formats are as follows:

Enzyme immunoassays (EIA)

Enzyme Immunoassays (EIA) are composed of a variety of assay formats that can be 

used to quantify bioaerosols in an air or dust sample. The binding of an antibody or 

antigen to an enzyme, such as horseradish peroxidase (HRP) or alkaline phosphatase 

(AP), is the basis of EIA techniques. Enzymatic activity, in the presence of a 

chromogen, results in a colored end-product that is quantified using a 

spectrophotometer. Many, if not most, commercially available EIAs are enzyme-linked 

immunosorbent assays (ELISAs). There are four types of ELISAs which include direct 

ELISA, indirect ELISA, sandwich ELISA and competitive ELISA. 

The sandwich ELISA method is typically used for the detection for airborne viruses 

and aeroallergens [Hung et al. 2005]. In this assay, a capture antibody is bound to a 

solid surface, usually a 96-well plate. The bioaerosol extract is added to the plate 

containing the capture antibody and incubated for a specified length of time, washed 

with a phosphate buffer solution and probed with an enzyme-labeled antibody that 

enables detection and quantification, either through colorimetric changes or 

fluorescence emissions. The advantages to using a sandwich-based ELISA method are 

that it is highly specific and can be used on complex samples such as aerosols. Some 

disadvantages to using a sandwich-based ELISA are poor antibody recognition and/or 

minimal detection due to low sample concentration. Lastly, it should be noted that 

ELISA, like any protein-based assay, does not distinguish between viable and non-
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viable viral aerosols. Multiplexed technologies that enable the detection and 

quantification of multiple allergen sources in one sample are also available from a 

variety of commercial sources [King et al. 2013].

EIA methods to assess fungal bioaerosols based on the detection of fungal cell wall 

components, enzymes, antigens and allergens have become commercially available. β1,3-D glucan, extracellular polysaccharides, mycotoxins and a variety of fungal 

antigens can now be quantified in air and dust samples using immunochemical, 

enzymatic and chemical detection platforms [Chew et al. 2001; Douwes et al. 1997; 

Eduard et al. 2012]. Although many of these biomarkers and methods serve as a proxy 

measure for total fungal biomass, these approaches can be confounded by limitations 

associated with component extraction biases. In addition, complex extraction, 

washing, amplification or immunochemistry steps are required that may add hours or 

even days before a dataset is finalized for analysis and interpretation.

To date, there are a number of commercial companies that have developed ready to 

use EIA kits for the detection and quantification of a variety of indoor and 

occupational biomarkers including dog, cat, dust mite, fungal and rodent allergens 

[Filep et al. 2012]. These EIA approaches enable the collection and quantification of 

these biomarkers in the work environment and provide a ready to use platform for the 

industrial hygienist.

Fluorescent immunoassays (FIA)

Utilization of fluorescent-labeled antibodies to detect bacterial antigens was 

introduced by Coons et al. [1941; 1942]. Various FIA techniques have now evolved 

and are commonly utilized in laboratories. These include: (1) direct FIA, to detect cellbound antigens using a fluorescent antibody; (2) indirect FIA to detect cell-bound 

antigens using a primary antibody and a fluorescent secondary antibody; and (3) 

indirect FIA to detect serum antibodies using an antigen, serum, and a fluorescent 

antibody. Various fluorescent dyes, such as fluorescein, fluorescein isothiocyanate, and 

rhodamine isothiocyanate, may be employed [Thermo Fisher Scientific 2014]. A 

fluorescent or confocal microscope is used to evaluate the samples and to count the 

number of fluorescently stained organisms [Garvey et al. 1977; Popp et al. 1988]. 

Similarly, flow cytometry-based approaches using fluorescent-labelled antibodies have 

also been employed to evaluate a variety of bioaerosol sources including bacteria, 

pollen and fungi [Rittenour et al. 2012; Rule et al. 2007; Rydjord et al. 2007]. 

Multiplexed approaches have been developed for the detection of multiple allergens in 

the same extracted sample [King et al. 2013].

FIA can be used to detect viruses. An example of a direct FIA assay for the detection of 

virus-laden aerosols is the Focus Forming Assay (FFA) [Flint et al. 2009a]. With the 
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FFA, fluorescent microscopy is used to visualize immunostained cells and viral titers 

are quantified as focus forming units per milliliter, or FFU/mL. While the FFA is more 

sensitive than culture-based methods alone, samples with low viral titers may weakly 

fluoresce and possibly be considered undetectable. To overcome weak detection levels, 

an indirect IFA assay may be more appropriate [Leland and Emanuel 1995; Madeley 

and Peiris 2002]. With indirect IFA assays, a primary, unconjugated antibody is used 

in combination with a fluorophore-conjugated, secondary antibody directed against 

the primary antibody. Because the secondary antibody is able to bind to multiple 

epitopes on the primary antibody, it increases fluorescence and enhances overall 

detection. While IFA is a trusted method of viral detection and quantification, it is 

fraught with limitations including excessive cost and the necessity of a skilled 

technician experienced in the reading of immunofluorescence. Likewise, because 

viruses are constantly undergoing antigenic drift and occasionally antigenic shift, 

changes in viral antigens can affect the binding affinity of the primary antibody and 

may result in false negatives.

Ligand-based assays

As an alternative to cell culture and immunofluorescent-based assays, there are several 

protein-based methods of detection that can be used to quantify viral loads in an 

aerosol sample. The hemagglutination (HA) assay is a non-fluorescence quantitative 

assay that is based upon the ability of certain viral pathogens to agglutinate speciesspecific erythrocytes [Condit 2007]. In a serial twofold dilution, viral samples are 

mixed with a 1% solution of erythrocytes and incubated at room temperature for 30-

60 minutes. Viral samples which form an agglutinated lattice are able to prevent red 

blood cells from precipitating out of solution by the binding of the hemagglutinin 

protein (present on the surface of the viral pathogen) to the sialic acid receptors 

(present on the surface of red blood cells). The titer of the sample is based on the well 

with the last agglutinated appearance, immediately before the well in which the red 

blood cells have settled out of solution. Hemagglutination units (HAUs) are typically 

used to quantify the viral concentration. 

One of the major limitations of the HA assay is that it does not distinguish between 

infectious and non-infectious viral particles. Likewise, certain bacteria and fungi 

possess hemolytic activity and when present in a collected bioaerosol, can alter the HA 

assay and result in false positive readings. To circumvent this issue, a variation of the 

HA assay, known as the Hemagglutination-Inhibition test, can be performed [Stewart 

et al. 1967]. The HA inhibition test measures serum antibodies that are directed 

against the viral pathogen. When present in sufficient concentration, the serum 

antibodies are able to prevent agglutination of red blood cells thereby providing an 

alternative means of quantifying viral loads.
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Direct and indirect immunostaining

Direct and indirect immunostaining methods have been previously described for the 

detection of bioaerosol sources. Popp et al. [1988] developed a staining technique to 

enumerate bioaerosol samples directly captured on microscope slides. These 

approaches have enabled the identification of specific bioaerosol sources, especially 

those that do not contain morphological phenotypes that would be used by a trained 

microbiologist to resolve and identify a specific microorganism [Popp et al. 1988]. 

Alternatives to this approach have been developed and utilized in a variety of indoor 

and occupational environments. A press blotting approach that included immobilizing 

proteins from collected bioaerosols captured on an adhesive tape provided insight into 

the bioaerosols that contain allergen in the outdoor environment [Takahashi et al. 

1993; Takahashi and Nilsson 1995]. An alternative method, called the Halogen 

Immunoassay, enables the immunostaining of allergen and antigen around bioaerosols 

captured on a protein binding membrane such as PVDF or mixed cellulose ester 

[Green et al. 2006c; Tovey et al. 2000]. This immunoassay approach has been used in a 

variety of indoor and occupational settings to evaluate allergen sources including, cat, 

dog, latex, rodent, plant, and fungi [Green et al. 2006a; Green et al. 2003; Green et al. 

2005a; Green et al. 2011; Green et al. 2005b; Green et al. 2005c; Green et al. 2006b; 

Green et al. 2006c; Mitakakis et al. 2001; Poulos et al. 2002; Poulos et al. 1999; 

Razmovski et al. 2000; Renstrom 2002; Tovey and Green 2004]. Recently these 

approaches have been adapted to FESEM applications and have been used to detect 

morphologically indiscernible fungal fragments [Afanou et al. 2015].

Biosensors

To overcome some of the technical challenges associated with traditional methods to 

assess bioaerosol exposure, real-time sensor technologies are being developed for the 

detection of bioaerosols [Fronczek and Yoon 2015; Hook-Barnard et al. 2014]. The 

sensor technologies are based on a variety of signal detection strategies that include 

optical, mechanical, electrical, or magnetic sensing approaches. These methods have 

resulted in platforms that have enabled the rapid detection of microbial pathogens and 

in an automated format. Using this technology requires little technological skill and 

can be developed into an automated handheld device. These developments have 

resulted in the fabrication of remote monitoring units in the agricultural sector that 

can process data remotely and apply it to geographical information systems or 

forecasting models.

Biosensors have been developed for a variety of applications including the detection of 

microbial pathogens [Fronczek and Yoon 2015; Hook-Barnard et al. 2014]. Typically, 

a biologically derived analyte (such as fungal spores, hyphae, antigens, peptides or 

nucleotides) is collected and interacts with a selected bioreceptor immobilized on a 
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 sensor surface. Examples of bioreceptors include oligonucleotides, 

monoclonal/polyclonal antibodies, enzymes, cells, and even phages. The bioreceptor 

system consists of a physiochemical transducer that produces a measurable signal. 

Transducers can be optical and include surface plasmon resonance (SPR), which is a 

method that measures changes in the refractive index during molecular binding events 

[Unser et al. 2015; Usachev et al. 2014]. In contrast, electrochemical transducers 

measure changes in current, potential, impedance, and conductance across an 

electrode surface for detection events [Patolsky et al. 2006]. Examples of 

electrochemical detection include the measurement of electrical conductance 

produced by antibody-antigen binding events [Patolsky et al. 2004]. Both optical and 

electrochemical transducers have been developed for the detection of a variety of 

pathogens in the biosecurity, medical and agricultural sectors.

h. Gene-based assays

Cell culture, protein-based, and immunological-based assays are invaluable diagnostic 

tools. However, the evolution of nucleic acid-based molecular diagnostics are rapidly 

becoming the preferred method for detecting and quantifying bioaerosols [West et al. 

2008]. Nucleic acid-based molecular diagnostics can be divided into two categories, (1) 

Direct sample analysis and (2) Indirect sample analysis, such as the Viral Replication 

Assay (VRA), which requires cultivation of the target microorganism prior to molecular 

analysis [Blachere et al. 2011]. Regardless of which approach is taken, there are three steps 

involved: extraction and purification of nucleic acids; amplification of the gene target; and 

detection of the amplicon. For viral, bacterial, plant and fungal nucleic acid extraction and 

purification, a number of kits are commercially available. Such kits generally rely upon 

either silica adsorption (spin-column) or affinity purification (magnetic separation) 

methodologies. Because bioaerosols are typically dilute in nature, investigators should 

determine which method yields the greatest amount of nucleic acids while minimizing 

sample handling, contamination and degradation. These are important variables to 

consider and can vary depending on the selected approach.

With the advent of molecular assays such as the Polymerase Chain Reaction (PCR) and 

Real-Time Quantitative PCR (RT-qPCR), which detect specific genetic sequences in the 

sample DNA or RNA, it has been possible to provide standardized assays, reduce 

turnaround time, and enhance assay sensitivity and detection specificity [Cella et al. 2013; 

Life Technologies 2014; Mahony 2008]. Using gene-specific oligonucleotides coupled with 

either an intercalating fluorescent dye (e.g. SYBR green) or a fluorogenically labeled gene 

probe (e.g. VIC, 6FAM), industrial hygienists are able to monitor indoor and outdoor 

bioaerosols for the presence of microorganisms. Likewise, multiplexing PCR and beadbased multiplexing PCR, which couples PCR and flow-cytometry, can be used for high-
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 throughput screening of multiple respiratory viruses in a single reaction mixture. It should 

be noted that poor assay design, primer and probe base-pair mismatches and degraded 

template nucleic acids can lead to false negatives or reduced detection sensitivity. 

Investigators should ensure optimal assay design, validate limits of detection, and run 

valid PCR controls in parallel.

Within the last two decades, a variety of molecular technologies have been used to 

quantify eukaryotic biomass including fungi and plant pollen in occupational, health, 

residential, and industrial samples [Rittenour et al. 2012; Scott et al. 2011; Summerbell et 

al. 2011]. Examples of these technologies include molecular based methods to evaluate 

specific or conserved gene loci (internal transcribed spacer region of ribosomal RNA) such 

as Sanger [Rittenour et al. 2014] or next generation sequencing [Kettleson et al. 2015], 

denaturing gradient gel electrophoresis [Johansson et al. 2014] and quantitative PCR 

[Eduard et al. 2012; Vesper et al. 2007]. The latter approach includes examples such as a 

DNA-based mold specific quantitative PCR (msQPCR) method that enables the detection 

and quantification of 36 indicator fungal species [Vesper et al. 2007]. This msQPCR 

method has been used to develop an Environmental Relative Moldiness Index (ERMI) to 

quantify the mold burden in homes. Originally developed by the U.S. Environmental 

Protection Agency (EPA) [Vesper et al. 2007], this approach has been licensed to a variety 

of companies in the commercial sector and is widely used to evaluate indoor fungal 

bioaerosol particles in settled dust during investigations of indoor air quality [BolañosRosero et al. 2013; Kettleson et al. 2015; Reponen et al. 2012; Reponen et al. 2011a; Taubel 

et al. 2016; Vesper et al. 2013; Vesper et al. 2007]. The development of this methodology 

has provided the first step towards a standardized approach to quantify fungal bioaerosol 

sources within the indoor environment. Other metagenomic molecular methods including 

Sanger, 454, and Illumina miSeq sequencing platforms have also provided new insights 

into the complete diversity of bacterial and fungal bioaerosols in indoor, outdoor and 

occupational environments.

10 Limitations of bioaerosol sampling and 

characterization

Bioaerosol sampling can be a useful tool to study occupational exposures, potential health 

hazards, and the transmission of infectious diseases. However, bioaerosol sampling has 

significant limitations, and these need to be kept in mind when deciding whether or not to 

collect bioaerosol samples, preparing a sampling plan, and interpreting the results.

The first and most important limitation is the lack of standards and guidelines for acceptable 

bioaerosol exposure limits. NIOSH and other organizations have set recommended exposure 
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 limits for several organic materials which may contain microorganisms and their fragments, 

such as cotton dust, grain dust, starch and wood dust [NIOSH 2010]. Although numerous 

studies have suggested a connection between exposure to various bioaerosols and respiratory 

illnesses, exposure limits do not currently exist in the US for airborne pollen, fungi, protozoa, 

bacteria, viruses, or their fragments. These limits have not been established largely because it 

is not possible to definitively state that a particular bioaerosol concentration will or will not 

lead to adverse health outcomes [Eduard et al. 2012; Heederik 2013; Morey 2007; Nevalainen 

et al. 2015; NIOSH 2012a]. This is true for several reasons: bioaerosols are often a complex 

mixture of microorganisms and organic materials; thousands of species of microorganisms 

exist, and most have not been studied; microorganisms and their fragments can cause illnesses 

in a variety of ways, including allergic reactions, infections and toxicity; the health effects of 

biological materials can vary substantially from person to person; and sampling and analytical 

procedures are not standardized, which makes it difficult to compare results [Eduard et al. 

2012; Heederik 2013; Morey 2007; Nevalainen et al. 2015; Taubel et al. 2016]. Although 

research is ongoing, no standards for acceptable levels of bioaerosols in the environment have 

been established by the US government or organizations such as the ACGIH or the AIHA.

In addition to the lack of occupational exposure limits for bioaerosols, measuring and 

interpreting bioaerosol concentrations are more complex than is often appreciated. Bioaerosol 

concentrations can vary significantly from location to location within a building, especially if 

the bioaerosol has one or a few localized sources. A study of bioaerosol exposure in a large 

engine plant found that levels of airborne fungi, bacteria and endotoxin varied from location 

to location within the plant [Thorne et al. 1996]. A study of airborne fungi in two residences 

found significant differences between two rooms sampled at the same time [Hyvarinen et al. 

2001]. In healthcare settings, patients with certain respiratory infections expel bioaerosol 

particles containing infectious pathogens. Because of the dispersion of the aerosol and the 

settling of larger droplets, the bioaerosol concentration decreases rapidly as the distance from 

the patient increases [Jones and Brosseau 2015]. A study of airborne influenza in a healthcare 

clinic found that the concentrations were much higher in examination rooms containing 

patients with influenza than other locations, and that the airborne influenza concentration 

also varied from location to location within the waiting room [Lindsley et al. 2010a].

Most bioaerosol collection methods provide a snapshot of the environmental bioaerosols at a 

specific time. Temporal variations in bioaerosol concentrations are commonly observed, 

especially if the bioaerosol generation occurs during episodic events rather than continuously. 

One study of indoor airborne mold in a residence found that day-to-day concentrations of 

airborne fungi varied considerably, and that levels were 26 times higher in the summer than in 

the winter [LeBouf et al. 2008]. Another residential study found that more airborne fungi were 

present in the morning than the afternoon and earlier in the winter compared to later 

[Hyvarinen et al. 2001]. In the influenza study mentioned above, day-to-day levels of airborne 
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 influenza virus varied considerably depending upon the number of influenza patients present 

[Lindsley et al. 2010a].

Other factors also influence bioaerosol concentrations. Building airflow patterns and the 

operation of the HVAC system can affect bioaerosol levels, particularly if the HVAC system is 

a source of bioaerosol particles [Macher 1999]. Areas occupied by people show increased 

levels of bioaerosols compared to empty spaces, both because people themselves shed 

bioaerosol particles and because human activities such as walking and sitting can re-suspend 

dust from floors and furniture [Buttner and Stetzenbach 1993; Ferro et al. 2004; Hung et al. 

2005; Qian et al. 2012]. Outdoor air is an important source of airborne fungi in many indoor 

environments due to fresh air being drawn in by HVAC systems and infiltration through 

cracks and openings [Eduard 2009].

Because of these issues, if bioaerosol sampling is to be conducted, it needs to be a part of a 

well-planned and comprehensive sampling strategy. The development of a sampling plan 

should begin with a thorough inspection and understanding of the workplace, including the 

building, HVAC system, and possible sources of bioaerosols. The sampling plan should 

integrate other types of data collection with the bioaerosol sampling, such as bulk sampling of 

possible source materials, surface sampling of settled dust, and health surveys of workers. 

Collections will need to be carried out at multiple locations and multiple time points, and 

even then it must be kept in mind that such samples may not fully characterize the exposure 

and that false negative results are quite possible [ASTM 2014a; Hung et al. 2005; Macher 1999; 

Morey 2007].

Bioaerosol sampling can be beneficial when done in the appropriate context [Hung et al. 2005; 

Macher 1999; Morey 2007]. It can be helpful to compare indoor and outdoor levels of 

bioaerosols to identify possible indoor problem microorganisms. Sampling for specific 

microorganisms of concern can be useful, especially if there is a known source, such as a 

composting operation or an aeration tank in a sewage treatment facility [Environment Agency 

2009; Masclaux et al. 2014]. Bioaerosol sampling is also a valuable research tool for better 

understanding sources and exposures. On the other hand, sampling for bioaerosols will not be 

helpful if there is not some basis for interpreting the resulting data. For example, because of 

the lack of dose-response information and the variability associated with bioaerosol sampling, 

NIOSH does not recommend routine air sampling when investigating possible respiratory 

illness due to exposures in damp buildings. Instead, NIOSH recommends inspections of the 

building and its HVAC systems to locate moisture and microbial growth problems, followed 

by remediation [NIOSH 2012a].
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11 Safety considerations

Investigators should use appropriate personal protective equipment (PPE) and practice good 

personal hygiene when conducting indoor environmental quality, disease outbreaks, and 

agricultural health investigations that have resulted in medically diagnosed symptoms. PPE 

may include respiratory protection to prevent inhalation of microbes and microorganismresistant clothing to prevent transmission to investigators by bodily contact with 

microorganisms. Good personal hygiene practices include washing exposed skin and clothing 

thoroughly and refraining from eating, drinking, or smoking in a contaminated area. These 

simple steps will help minimize the ingestion, inhalation, or uptake of microorganisms.

All samplers, culture plates, and other equipment should be handled aseptically to prevent 

contamination of the samples and, more importantly, to prevent the spread of potential 

human pathogens to the worker or the work environment. All surfaces, including washed 

hands, may harbor microorganisms or spores unless they are specifically sterilized. Practically 

speaking, however, not all objects can be sterilized. While disinfection with an oxidizing 

chemical or alcohol destroys most vegetative cells, these agents do not destroy all spores. 

Samplers should be disinfected or, if possible, sterilized after each sample collection. Special 

care should be given to samplers with convoluted inlets or air pathways where 

microorganisms may accumulate.

Information on the safe handling of biological specimens can be found in in the free online 

manual “Biosafety in Microbiological and Biomedical Laboratories” from the Centers for 

Disease Control and Prevention at 

(http://www.cdc.gov/biosafety/publications/bmbl5/index.htm) [CDC 2009]. Information on 

the handling of some specific pathogens can also be found at the CDC website (www.cdc.gov).

12 Resources

The NIOSH Manual of Analytical Methods has several chapters discussing other aspects of 

aerosol sampling, including general considerations and factors affecting aerosol sampling, an 

explanation of filter pore size, sampling airborne fibers, sampler wall losses, and avoiding 

bypass leakage in filter cassettes [NIOSH 2003e]. NIOSH also maintains a web page on indoor 

environmental quality with more information and links to additional resources at 

http://www.cdc.gov/niosh/topics/indoorenv/. The American Industrial Hygiene Association 

(AIHA; https://www.aiha.org) has reference materials and resources on indoor air quality and 

bioaerosols, as does the American Conference of Governmental Industrial Hygienists 

(ACGIH; http://www.acgih.org). ASTM International (http://www.astm.org) publishes 

numerous standards and guides on the evaluation of indoor air quality, including developing 
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Sampling and Characterization of Bioaerosols 

 an air sampling strategy and the collection and evaluation of bioaerosols [ASTM 2009; ASTM 

2014a; ASTM 2014b; ASTM 2014d; ASTM 2014e]. The Centers for Disease Control and 

Prevention has guidelines on environmental infection control in healthcare facilities that 

include recommendations on environmental sampling [CDC 2003].

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

13 References

Abed Y, Boivin G [2006]. Treatment of respiratory virus infections. Antiviral Res 70(2):1-16. 

ACGIH [1989]. Guidelines for the assessment of bioaerosols in the indoor environment. 

Cincinnati, OH: American Conference of Governmental Industrial Hygienists.

ACGIH [2001]. Documentation of the Threshold Limit Values and Biological Exposure 

Indices. Appendix C: Particle size-selective sampling criteria for airborne particulate matter. 

Cincinnati, Ohio: American Conference of Governmental Industrial Hygienists.

ACGIH [2015]. TLVs and BEIs based on the documentation of the Threshold Limit Values 

for chemical substances and physical agents and Biological Exposure Indices. Cincinnati, OH: 

American Conference of Governmental Industrial Hygienists.

Adams RI, Tian Y, Taylor JW, Bruns TD, Hyvarinen A, Taubel M [2015]. Passive dust 

collectors for assessing airborne microbial material. Microbiome 3:46. 

Adhikari A, Reponen T, Rylander R [2013]. Airborne fungal cell fragments in homes in 

relation to total fungal biomass. Indoor Air 23(2):142-147. 

Afanou KA, Straumfors A, Skogstad A, Nayak AP, Skaar I, Hjeljord L, Tronsmo A, Eduard W, 

Green BJ [2015]. Indirect immunodetection of fungal fragments by field emission scanning 

electron microscopy. Appl Environ Microbiol 81(17):5794-5803. 
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Sampling and Characterization of Bioaerosols 

 

14 Appendix 1- List of manufacturers/distributors of 

common bioaerosol samplers and related 

products

This list is not inclusive, and the inclusion of a specific product or company does not 

constitute endorsement by the National Institute for Occupational Safety and Health 

(NIOSH). If your bioaerosol equipment manufacturing or supply company is not listed or the 

information is incorrect or out-of-date, please contact us and we will review your information 

for inclusion as the list is updated.

Manufacturers/Distributors of Common Bioaerosol Samplers and Related Products

A.P. Buck Inc.

7101 Presidents Drive, Suite 110

Orlando, FL 32809 USA 

Phone: (800) 330-BUCK [2825] 

Phone: (407) 851-8602 

Fax: (407) 851-8910 

http://www.apbuck.com

Ace Glass Incorporated

1430 North West Boulevard

P.O. Box 688

Vineland, NJ 08362 USA 

Phone: (800) 223-4524 

Phone: (856) 692-3333 

Fax: (800) 543-6752 

Fax: (856) 692-8919 

http://www.aceglass.com

Aerosol Devices Inc.

2614 S. Timberline Road, #109-125 

Fort Collins, CO 80525 USA 

Phone: (970) 744-3244 

http://aerosoldevices.com

Aquaria srl

Via della Levata, 14

Lacchiarella (Milan) 20084 Italy 

Phone: +39 02-90091399 

Fax: +39 02-9054861 

http://www.aquariasrl.com

Barramundi Corporation

6449 South Tex Point 

PO Drawer 4259 

Homosassa, FL 34448 USA 

Phone: (800) 382-1817 

Phone: (352) 628-0200 

Fax: (352) 628-0203 

http://barramundicorp.com

BD Biosciences 

2350 Qume Drive 

San Jose, CA 95131 

Phone: 877-232-8995 

http://www.bdbiosciences.com

Beijing SENNON Technology

Development Company, Ltd.

North Building No. 2

Dongdajie xili, Fengtai District

Beijing 100071 P.R. China

Phone: +86 10-6381 8024 

Fax: +86 10-6380 6170 

http://www.sennon.net/eng

Bertin Corporation

9700 Great Seneca Highway 

Suite # 662 

Rockville, MD 20850 USA 

Phone: (240) 428-1047 

http://www.coriolis-airsampler.com

Bi-Air Corporation

1349 Montevideo Ave 

Placentia, CA 92870 USA 

Phone: (714) 985-9659 

Fax: (714) 528-5429 

http://expertonmold.com

bioMérieux, Inc.

595 Anglum Road 

Hazelwood, MO 63042 USA 

Phone: (800) 634-7656 

Fax: (800) 657-3053 

http://www.biomerieux-usa.com

Bioscience International

11333 Woodglen Drive 

Rockville, MD 20852 USA 

Phone: (301) 231-7400 

Fax: (301) 231-7277 

http://www.biosci-intl.com

Burkard Manufacturing Company 

Ltd.

Woodcock Hill Industrial Estate

Rickmansworth, Hertfordshire WD3 

1PJ England 

Phone: +44 (0) 1923 773134 

Fax: +44 (0) 1923 774790 

http://www.burkard.co.uk

Climet Instruments Company

1320 W. Colton Avenue 

Redlands, CA 92374 USA 

Phone: (909) 793-2788 

http://www.climet.com

Droplet Measurement Technologies

2545 Central Avenue 

Boulder, CO 80301 USA 

Phone: (303) 440-5576Fax: (303) 440-

1965 

http://www.dropletmeasurement.com
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-102 of BA-115

Sampling and Characterization of Bioaerosols 

Manufacturers/Distributors of Common Bioaerosol Samplers and Related Products -

Continued

Dycor Technologies, Ltd.

1851 94th Street 

Edmonton, AB T6N 1E6 Canada 

Phone: (800) 663-9267 

Phone: (780) 486-0091 

Fax: (780) 486-3535 

http://www.dycor.com

EMD Millipore

290 Concord Road 

Billerica, MA 01821 USA 

Phone: (800) MILLIPORE [645-5476] 

Fax: (781) 533-6000 

http://www.emdmillipore.com

EMSL Analytical, Inc 

200 Route 130 North 

Cinnaminson, NJ 08077 

Phone: (800) 220-3675 

http://www.emsl.com Environics Oy

Environmental Monitoring Systems, Inc. 

3864 Leeds Avenue 

Charleston, SC 29405 USA

Phone: (800) 293-3003 

Phone: (843) 724-5708 

Fax: (866) 724-5702 

Fax: (843) 724-5702 

http://www.emssales.net

Evogen, Inc.

10513 W. 84th Terrace 

Lenexa, KS 66214 USA 

Phone: (888) 450-4321 

Phone: (913) 948-5640 

Fax: (913) 948-5664 

http://evogen.com

F.W. Parrett Limited

7 Coppergate Close

Bromley, Kent BR1 3JG England 

Phone: +44 020-8460-2116 

Fax: +44 020-7504-3536 

http://www.parrett.uk.com

FLIR Systems, Inc.

70 Castilian Drive 

Goleta, CA 93117 USA 

Phone: (888) 747-FLIR [3547] 

http://www.flir.com

GE Healthcare Life Sciences

(Whatman)

800 Centennial Avenue 

P.O. Box 1327 

Piscataway, NJ 08855 USA 

Phone: (800) 526-3593 

Fax: (877) 295-8102 

http://www.gelifesciences.com

Indoor Biotechnologies Inc

700 Harris Street

Charlottesville, VA 22903 USA 

https://inbio.com

InnovaPrep

132 East Main Street

Drexel, MO 65742 USA 

Phone: (816) 619-3375 

http://innovaprep.com

InnovaTek, Inc. 

3100 G. Washington Way 

Suite 108

Richland, WA 99354 USA 

Phone: (509) 375-1093 

Fax: (509) 375-5183 

http://www.innovatek.com

Inspirotec

2319 West Wabansia Avenue #1

Chicago, IL 60647

Phone: 847 302 1839 

Fax: 847 234 2089 

http://www.inspirotec.com 

MSP Corporation

5910 Rice Creek Parkway

Suite 300

Shoreview, MN 55126 USA 

Phone: (651) 287-8100 

Fax: (651) 287-8140 

http://www.mspcorp.com

Pall Corporation

25 Harbor Park Drive

Port Washington, NY 11050 USA 

Phone: (800) 521-1520 

Phone: (516) 484-3600 

Fax: (516) 801-9754 

http://www.pall.com

Particle Measuring Systems

5475 Airport Boulevard

Boulder, CO 80301 USA 

Phone: (800) 238-1801 

Phone: (303) 443-7100 

Fax: (303) 449-6870 

http://pmeasuring.com

Research International, Inc.

17161 Beaton Road SE 

Monroe, WA 98272 USA 

Phone: (800) 927-7831 

Phone: (360) 805-4930 

Fax: (360) 863-0439 

http://www.resrchintl.com

RJ Lee Group, Inc.

350 Hochberg Road

Monroeville, PA 15146 

Manufacturers/Distributors of 

Common Bioaerosol Samplers and 

Related Products

(724) 325-1776

Fax: (724) 733-1799

http://www.rjlg.com 

Sammonkatu 12

P.O. Box 349

FI-50101 Mikkeli Finland 

Phone: +358 201 430 430 

Fax: +358 201430 440 

http://www.environics.fi/

Sartorius Stedim Biotech GmbH

August-Spindler-Strasse 11

Goettingen 37079 Germany 

Phone: (800) 368-7178 

Phone: +49 551-308-0 

Fax: +49 551-308-3289 

https://www.sartorius.com

SKC, Inc.

863 Valley View Road 

Eighty Four, PA 15330 USA 

Phone: (800) 752-8472 

Phone: (724) 941-9701 

Fax: (724) 941-1369 

http://www.skcinc.com
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-103 of BA-115

Sampling and Characterization of Bioaerosols 

 Manufacturers/Distributors of Common Bioaerosol Samplers and Related Products -

Continued

Tecora 

211-215 Rue la Fontaine 

94134 Fontenay sous Bois Cedex, France 

Phone: Tel: +33 1 48 75 82 82 

Fax: +33 1 48 75 82 96 

http://www.tecora.com/en/

Thermo Scientific

27 Forge Parkway

Franklin, MA 02038 USA 

Phone: (866) 282-0430 

Phone: (508) 520-0430 

Fax: (508) 520-1460 

http://www.thermoscientific.com

TSI Incorporated

500 Cardigan Road

Shoreview, MN 55126 USA 

Phone: (800) 874-2811 

Phone: (651) 483-0900 

Fax: (651) 490-3824 

http://www.tsi.com

Veltek Associates, Inc.

15 Lee Boulevard

Malvern, PA 19355 USA 

Phone: (888) 4-STERILE [478-3745] 

Phone: (610) 644-8335 

http://sterile.com

Zefon International, Inc. 

5350 SW 1st Lane 

Ocala, FL 34474 USA 

Phone: (800) 282-0073 

Phone: (352) 854-8080 

Fax: (352) 854-7480 

http://www.zefon.com
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-104 of BA-115

Sampling and Characterization of Bioaerosols 

15 Appendix 2 – Commonly used bioaerosol samplers

This list is not inclusive, and the inclusion of a specific product or company does not constitute endorsement by the National Institute for 

Occupational Safety and Health (NIOSH). If a bioaerosol sampler is not listed or the information is incorrect or out-of-date, please contact 

us and we will review your information for inclusion as the list is updated.

Table I. Common Commercially-Available Filter Samplers for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

d50

(μm) Application¥

Bi-Air Bi-Air Filter Cassette Filter 1 – 5 Varies C,M,O

Burkard High Throughput 'Jet' Spore and Particle Sampler Filter 850 Varies C,M,O

Dycor CSU-1 Low Volume Air Sampler Filter 10 Varies C,M,O

EMD Millipore Various filter and membrane media Filter Varies Varies C,M,O

GE (Whatman) Various filter and membrane media Filter Varies Varies C,M,O

InnovaPrep ACD-200 Bobcat Air Sampler Filter 100 or 200 C,M,O

Pall Corp. Various filter and membrane media Filter Varies Varies C,M,O

Research 

International SASS® 3100 Dry Air Sampler Filter 50 – 310 0.3 – 0.5 C,M,O

Sartorius AirPort MD8 Air Sampler Filter 30, 40, or 50 - C

SKC Button Aerosol Sampler Filter 4 - M,O

Zefon Various filter and membrane media Various Varies Varies C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
Table II A. Common Commercially-Available Single-stage Impactor Samplers for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Collection 

Media

Flowrate

(L/min)

d50

(μm) Application¥

A.P. Buck

BioAireTM B6 Single Stage Microbial Sampler Agar 28.3 - C 

Bio-CultureTM Microbial Air Sampler Agar 30 – 120 - C 

BioSlideTM Microbial Air Sampler Slide 10 – 20 - M 

Aquaria

MICROFLOW 60 Microbiological Air Sampler Agar 30 – 120 - C 

MICROFLOW 60-90/C Microbiological Air Sampler Agar 30 – 120 - C 

MICROFLOW 90/C Microbiological Air Sampler Agar 30 – 120 - C 

Barramundi Mattson-Garvin Model 220 Air Sampler (240V is Model 270) Agar 28.3 - C 

Beijing

SENNON

Handy Microbial Air Sampler® Agar/Filter - - C,M,O

JWL-IIA Mini® Microbial Air Sampler Agar 20 - C 

JWL-IIB202 Universal® Microbial Air Sampler Agar 20 - C 

JWL-IIC Professional® Microbial Air Sampler Agar 20 - C 

bioMérieux airIDEAL® 3PTM Traceability Air Sampler Agar 100 - C 

Samp'airTM Microbial Air Sampler Agar 100 - C 

Bioscience 

International

SAS Duo 360 High Volume Microbial Air Sampler Agar 360 - C 

SAS Isolator Microbial Air Sampler Agar 180 - C 

SAS Super 100 Microbial Air Sampler Agar 100 - C 

SAS Super 180 Microbial Air Sampler Agar 180 - C 

Burkard

24-Hour Recording Volumetric Spore Trap Slide 10 - M 

Continuous Recording Air Sampler Slide 10 - M 

Personal Volumetric Air Sampler Slide 10 - M 

Portable Air Sampler for Agar Plates Agar 10 or 20 - C 

Recording Air Sampler Side 10 - M 

Seven-Day Recording Volumetric Spore Trap Slide 10 - M 

Climet

CI-90 & CI-90+ Airborne Microbial Sampler Agar 100 - C 

CI-95 & CI-95+ Airborne Microbial Sampler Agar 100 - C 

CI-99 Microbial Air Sampler Agar 100 - C 

Dycor Dycor Slit Sampler Agar 15 – 50 - C 

EMD Millipore

MAS-100 Iso MH® Air Sampler Agar 100 - C 

MAS-100 Iso NT® Air Sampler Agar 100 - C 

MAS-100 NT® and MAS-100 NT Ex® Air Sampler Agar 100 - C 

MAS-100 VF® Active Air Sampler Agar 100 - C 

RCS® Isolator Microbial Air Sampler Agar 100 - C 

RCS® Plus Ex Explosion-Proof Microbial Air Sampler Agar 100 - C 

NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-105 of BA-115

Sampling and Characterization of Bioaerosols 

 
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-106 of BA-115

Sampling and Characterization of Bioaerosols

Table II A. Common Commercially-Available Single-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/ 

Distributor Sampler Name Collection 

Media

Flowrate 

(L/min)

d50 

(μm) Application¥

EMD Millipore RCS® High Flow Touch Portable Air Sampler Agar 100 - C

Environmental 

Monitoring 

Systems

Allergenco MK-III Slit-Impaction Sampler Slide 15 - M

Allergenco-D Disposable IAQ Air Monitoring Cassettes Slide 15 - M

Allergenco-D Posi-Track Full Slide, IAQ Impactor Cassette Slide 15 1.7 M

BioSIS Slit Impaction Air Sampler Slide 5 – 50 - M

cyclex-d Cassettes Slide 20 < 1.0 M

E6 Single-Stage Bioaerosol Impaction Sampler Agar 28.3 - C

Micro5 MicroCell Cassettes Slide 5 < 1.0 M

Micro5 Posi-Track Full Slide, IAQ Impactor Cassette Slide 5 <1.0 M

F.W. Parrett MicroBio MB1 Air Sampler Agar 100 - C

MicroBio MB2 Air Sampler Agar 100 - C

Particle 

Measuring

Systems

Air Trace® Environmental Slit-to-Agar Sampler Agar 28.3 - C

BioCaptTM Impactor Active Microbial Air Sampling Atrium Agar 25 - C

MiniCaptTM Portable Microbial Air Sampler Agar 50 or 100 - C

Sartorius AirPort MD8 Air Sampler Agar 125 - C

SKC

BioStage® Standard Single-Stage Viable Cascade Impactor Agar 28.3 - C

BioStage® 200 Single-Stage Viable Cascade Impactor Agar 14.15 - C

VersaTrap® Spore Trap Cassette Slide 15 - M

Thermo Scientific

IUL Basic Air Air Sampler Agar 60 – 100 - C

IUL Spin Air Air Sampler Agar 60 – 100 - C

IUL Spin Air Basic Air Sampler Agar 60 – 100 - C

N6 Single-Stage Viable Andersen Impactor Agar 28.3 0.65 C

Oxoid Air Sampler Agar 100 - C

Veltek SMA MicroPortable® Air Sampler Agar 1 or 5 - C

Zefon 

International

A-6 Bioaerosol Impactor Agar 28.3 0.65 C

Air-O-Cell® Sampling Cassette Slide 15 2.6 M

Via-Cell® Bioaerosol Sampling Cassette Slide 15 1.56 C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-107 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Collection 

Media

Flowrate # of 

(L/min) Stages

d50

(μm) Application¥

MSP 

Corporation

Model 100-NR (non-rotating)/100-R (rotating) MOUDITM 

Impactor Filter 30 8 

0.18 

0.32 

0.56 

1.0

1.8

3.2

5.6

10

M,O

Model 110-NR (non-rotating)/100-R (rotating) MOUDITM 

Impactor Filter 30 10

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 100-S4 MOUDITM Impactor Filter 30 3 

1.0

2.5

10

M,O

Model 115 Nano-MOUDITM Impactor Filter 10 3 

0.010

0.018

0.032

M,O

Model 116 Nano-MOUDITM Impactor Filter 30 3 

0.010

0.018

0.032

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-108 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection 

Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

MSP 

Corporation

continued

Model 120 R (rotating) Moudi-IITM Impactor Filter 30 10

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 122-NR/122-R Moudi-IITM and NanoMoudi-IITM Impactor Filter 30 13

0.010

0.018

0.032

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-109 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

MSP 

Corporation

continued

Model 125-NR/125-R Moudi-IITM and NanoMoudi-IITM Impactor Filter 10 13

0.010

0.018

0.032

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 135-6 MiniMOUDITM (Marple Personal II) Impactor Filter 2 6 

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 135-8 MiniMOUDITM (Marple Personal II) Impactor Filter 2 8 

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-110 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

MSP 

Corporation 

continued

Model 135-10 MiniMOUDITM Impactor Filter 2 10

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 135-13 MiniMOUDITM Impactor Filter 2 13

0.010

0.018

0.032

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Eight Stage Non-Viable Cascade Impactor Filter 28.3 8 

0.43

0.65

1.1

Thermo 2.1

Scientific 3.3

4.7

5.8

9.0

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-111 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection 

Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

Thermo 

Scientific 

continued

3.5

Marple 294 Personal Cascade Impactor Filter 2 4 9.8

14.8

21.3

M,O

0.52

0.93

Marple 296 Personal Cascade Impactor Filter 2 6 1.55

3.5

6.0

9.8

M,O

0.52

Marple 298 Personal Cascade Impactor Filter 2 8 

0.93

1.55

3.5

6.0

9.8

14.8

21.3

M,O

0.65

1.1

Six Stage Viable Andersen Cascade Impactor Agar 28.3 6 2.1

3.3

4.7

7.0

C 

Two-Stage Viable Andersen Cascade Impactor Agar 28.3 2 0.8

8 C 

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - 

C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-112 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table III. Common Commercially-Available Cyclones and Impinger Samplers for Bioaerosol Collection

Cyclones

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

Burkard Cyclone Sampler for Airborne Particles Dry Vial 20 1 - C,M,O

Cyclone Sampler for Field Operation Dry Vial 16.5 1 - C,M,O

Evogen Sceptor DryCloneTM Dry Vial 400 1 - C,M,O

FLIR Systems C100 Modular Tactical Collector Dry Vial 150 1 - C,M,O

Impingers

Ace Glass

AGI-30 Impinger Liquid 12.5 1 - C,M,O

Greenburg-Smith Impinger Liquid 28.3 1 - C,M,O

Midget Impinger Liquid 1 - C,M,O

4

Burkard Multistage Liquid Impinger Liquid 20 3 10

> 10

C,M,O

Dycor

XMX/102 High Volume Bioaerosol Sampling System Serum Tube 530 1 - C,O

XMX/2L-MIL Bioaerosol Sampler – Military Liquid 530 1 - C,M,O

XMX-CV Microbial Air Sampler – Civilian Liquid 530 1 - C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - 

C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-113 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table IV. Common Commercially-Available Wetted-Surface and Condensation-Based Samplers for Bioaerosol Collection

Wetted-Surface Bioaerosol Samplers

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

Bertin Coriolis® μ Microbial Air Sampler Liquid 100 – 300 1 < 0.5 C,M,O

Coriolis® RECON Portable Air Sampler Liquid 600 1 ≈ 0.5 C,M,O

Bioscience 

International SAS Cyclone Air Sampler Liquid 1200 1 - C,M,O

Evogen Sceptor SpinConTM Advanced Air Sampler Liquid 450 1 - C,M,O

FLIR Systems Fido® B1 (BioCapture® 650) Portable Air Sampler Liquid 

Cartridge 200 1 - C,M,O

InnovaPrep SpinCon® II Advanced Air Sampler Liquid 450 1 - C,M,O

InnovaTek BioGuardian® Air Sampler Liquid 100, 350, 

or 1000 1 - C,M,O

Research 

International

BioHawk® 8-Channel Collector/Bioidentifier Liquid 325 1 - O 

SASS® 2300 Wetted-Wall Air Sampler Liquid 325 1 - C,M,O

SASS® 2400 Low-Volume Wetted-Wall Air Sampler Liquid 40 1 - C,M,O

SKC BioSampler® Liquid 12.5 1 - C,M,O

Tecora CIP10-M personal bioaerosol sampler Liquid 10 1 2.1 C,M,O

Condensation-Based Bioaerosol Samplers

Aerosol Devices LSS100 Series Liquid Spot Sampler Liquid 1.0 – 1.5 1 n/a C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol 

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-114 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table V. Common Commercially-Available Electrostatic, Passive Aerosol and Settled Dust Collection Samplers for Bioaerosol Collection

Electrostatic Samplers

Manufacturer/

Distributor

Sampler Name Collection 

Media

Flowrate

(L/min)

# of 

Stages

d50

(μm)

Application¥

Inspirotec Inspirotec sampler Cartridge 130 1 n/a O 

Passive electrostatic dust collectors are listed under Passive Aerosol Samplers. Electrostatic cloths used for wipe sampling are listed under 

Settled Dust Collection Devices.

Passive Aerosol Samplers

BD Biosciences Ready-to-use settle plates Agar n/a n/a n/a C 

EMD Millipore Ready-to-use settle plates Agar n/a n/a n/a C 

Thermo-Scientific Ready-to-use settle plates Agar n/a n/a n/a C 

Department of Occupational and 

Environmental Health, University of Iowa

Electrostatic Dust Collector (EDC) Electrostaticallycharged clothsn/a n/a n/a O 

RJ Lee Group UNC Passive Aerosol Sampler Various 

substrates

n/a n/a n/a M, O

Settled Dust Collection Devices

Indoor Biotechnologies DUSTREAM® Collector (DU-ST-1) 40 μm nylon 

mesh filter

n/a n/a n/a O 

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-115 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table VI. Common Commercially-Available Real-Time Bioaerosol Monitors for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Detection Method Flowrate

(L/min) Application¥

Bioscience 

International SAS-PCR Pathogenic Microorganisms Air Sampler PCR - R 

Droplet 

Measurement 

Technologies

Wideband Integrated Bioaerosol Sensor (WIBS) Fluorescence 0.3 R 

Dycor C-FLAPS Biological Detection System Fluorescence 350 R 

Environics ENVI BioScoutTM Fluorescence 2 R 

FLIR Systems Fido® B2 Instantaneous Biological Aerosol Detector Fluorescence 3.8 R 

Particle 

Measuring

Systems

BioLazTM Real-Time Microbial Monitor Fluorescence 3.6 R 

Research 

International

BioHawk® 8-Channel Collector/Bioidentifier Fluorometric Bioassay 325 R 

TacBioTM Biological Aerosol Detector Fluorescence 1 R 

TSI BIOTRAK® Real-Time Viable Particle Counter 9510-BD Fluorescence 28.3 R 

Fluorescence Aerosol Particle Sensor (FLAPS) 3317 (FLAPS III)TM Fluorescence 1 R 

Application¥ - 

R= Real-time or near-real time bioaerosol detection

PCR= Polymerase chain reaction
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Filter Pore Size and Aerosol Sample 

Collection

by William G. Lindsley, PhD, NIOSH

1 Introduction FP-2

2 Physical structures of filters FP-2

3 Determination of equivalent pore diameter FP-3

4 How an aerosol filter collect particles FP-6

5 Aerosol filter efficiency and pore size FP-8

6 Significance of pore size FP-10

7 Filter selection FP-11

8 Conclusion FP-12

9 References FP-13
NIOSH Manual of Analytical Methods  5th Edition  Chapter FP April 2016 Page FP-2 of FP-14

Filter Pore Size and AerosolSample Collection

1 Introduction

Aerosol sampling filters are commonly used in industrial hygiene and environmental 

monitoring to collect airborne particles for analysis. The filter characteristics provided by the 

manufacturer frequently include the term “pore size” or “equivalent pore diameter,” and pore 

size is also specified in many particle-sampling methods written by government agencies and 

standards organizations. Unfortunately, the pore size of a filter is often misunderstood, which 

can lead to the misinterpretation of test results and the selection of filters with much higher 

flow resistances than are needed for a particular application. The purpose of this article is to 

discuss how aerosol filters actually work and what the equivalent pore diameter really means, 

and then to explain how this information should be used when selecting filters and 

interpreting data. Much of the information and terminology presented here were drawn from 

Hinds [1999], Brock [1983], Lippmann [2001] and Raynor et al. [2011]. All of these sources 

provide a more in-depth discussion of filter theory and use and are highly recommended if 

more information is desired.

2 Physical structures of filters

To understand what the term “pore size” does and does not indicate for filters, we begin by 

looking at the physical structures of some different types of filters. Most filters used in aerosol 

sampling fall into one of three categories. Fibrous filters like the glass fiber filter shown in 

Figure 1A consist of a deep mesh of fibers with random orientations. Porous membrane 

filters, such as those made from mixed cellulose esters (MCE) or polytetrafluoroethylene 

(PTFE), have a complex structure with tortuous routes through the filter material as shown in 

Figures 1B and 1C. A capillary pore filter consists of a thin, smooth polycarbonate (PC) or 

polyethylene terephthalate (PET) film with circular pores, as shown in Figure 1D. These are 

also called straight-through pore filters or track-etch membrane filters (because of the 

manufacturing method), or Nucleopore filters after the original manufacturer.
NIOSH Manual of Analytical Methods  5th Edition  Chapter FP April 2016 Page FP-3 of FP-14

Filter Pore Size and Aerosol Sample Collection

Figure 1: Scanning Electron Micrographs (SEM) of four filter types. The vertical tick 

marks above “10.0 μm” in the lower right-hand corner of each SEM are 1 μm apart; the 

entire scale is 10 μm in length.

A: Glass fiber filter with a 1-μm equivalent pore diameter. 

B: Mixed-cellulose esters (MCE) filter with 0.8-μm equivalent pore diameter.

C: Polytetrafluoroethylene (PTFE) filter with 3-μm equivalent pore diameter.

D: Polycarbonate capillary pore filter with 1-μm pore size.

3 Determination of equivalent pore diameter

So what is the pore size of a filter? For the capillary pore filter, the pore size is relatively 

straightforward: the pores are circular and reasonably uniform and run straight through the 

filter material, so the pore size is the diameter of the pores. This is what many people imagine 

when they think of the pore size. However, the other types of filters do not have these simple 

pore structures. The filter material forms intricate paths, and the airstream lines twist and turn 

as they pass through the filter. Thus, because these filters do not have an obvious, simple 

dimension that characterizes their pores, an “equivalent pore diameter” is used to describe the 

filters. This provides a useful way to categorize filters with different sized openings and to 

ensure consistent performance characteristics. When a manufacturer specifies the pore size of 
NIOSH Manual of Analytical Methods  5th Edition  Chapter FP April 2016 Page FP-4 of FP-14

Filter Pore Size and Aerosol Sample Collection

a filter, they are giving the actual pore diameter for capillary pore filters and the equivalent 

pore diameter for other types of filters.

The equivalent pore diameter is commonly measured by a “bubble-point test” [ASTM 2011]. 

This test is fairly simple, is non-destructive, and provides a good quality-control check for the 

filter. A bubble-point test works like this: Imagine that you have an ideal capillary pore filter 

with smooth holes that are of a uniform diameter, as shown in Figure 2. Now imagine that 

there is air on one side of this filter and a liquid that wets the filter on the other side. If the air 

pressure is low, the surface tension of the liquid will stop the air bubble from being pushed 

through the filter. If you slowly increase the air pressure, at some point it will be high enough 

to overcome the surface tension, and a visible stream of air bubbles will be produced. This 

pressure is called the bubble point. The pore diameter can be calculated from the bubble-point 

pressure with this formula [Brock 1983]: 

DD = 

4γγ cos θθ

PP

 × 106 (Equation 1) 

Where:

D = pore diameter (micrometers)

P = bubble-point air pressure (Pa)

γ = surface tension of the liquid (N/m)

θ = contact angle between the liquid and the filter material

Figure 2: Principle of the bubble-point test.
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Filter Pore Size and Aerosol Sample Collection

Note that as the pore diameter gets smaller, more air pressure is required for air to bubble 

through the ideal filter (Figure 3). Thus, you could take your actual filter and see how much 

air pressure is needed to bubble air through it. You could then calculate the pore size of an 

ideal filter that requires the same amount of air pressure to form bubbles as does your actual 

filter using Equation 1. The pore size of an ideal filter with the same bubble point as your 

actual filter is the “equivalent pore diameter” of your filter. 

Figure 3: Pore diameter vs. air pressure for ideal filter in bubble-point test. The curve was 

calculated with use of water (which has a surface tension of 72.8 mN/m) as the liquid. It is 

assumed that the water completely wets the filter material, and thus the contact angle θ = 

0°. Calculated using Equation 1.

Two things should be observed at this point. First, because less air pressure is required to push 

bubbles through larger openings than through smaller ones, the bubble-point test indicates 

the size of the largest pores in the filter, not the average pores. For this reason, the bubblepoint test is useful for quality control checks of filters, since it will indicate if defects or 

excessively large pores are present. However, if the filter has a wide range of pore sizes, most 

of the pores will be smaller than the equivalent pore diameter determined by this test. 

Second, the equivalent pore diameter provides a convenient reference point for describing and 

comparing filters of the same type, but not of different types. For example, the openings in a 

porous membrane filter with a 5-μm equivalent pore diameter will be somewhat larger than 

the openings in a porous membrane filter with a 1-μm equivalent pore diameter. However, the 

pore sizes of different types of filters cannot be meaningfully compared; a capillary filter with 

a 1-μm pore size bears little resemblance to a porous membrane filter with a 1-μm equivalent 

pore diameter.
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Filter Pore Size and Aerosol Sample Collection

4 How an aerosol filter collects particles

Now that you understand how the equivalent pore diameter is determined, let’s discuss how 

this relates to aerosol sampling. First, we need to review how aerosol sampling filters collect 

airborne particles. People often assume that a filter works like a sieve—that is, that a filter is 

like a sheet or mesh with holes of a particular size, and that particles larger than the holes 

collect on the filter while particles smaller than the holes pass through it. In fact, aerosol 

filtration is far more complex than this simple model would suggest, and one consequence is 

that aerosol filters can efficiently collect particles much smaller than would be expected on the 

basis of the pore size of the filter.

When an airstream containing airborne particles passes through a filter, the particles are 

collected by five mechanisms (Figure 4):

1) Interception: Interception occurs when a particle moving with the airstream contacts the 

filter material. Intercepted particles include those that are bigger than the filter pores 

(sieving), and also particles that are smaller than the pores but are carried close enough to 

touch the surface of the filter as they follow the airstream. The closer the diameter of the 

particle is to the diameter of the opening in the filter, the more likely interception is to 

occur. Interception can be very important in the collection of fibers and other irregularly 

shaped aerosol particles because an elongated particle is more likely to come in contact 

with the filter, especially if it is sideways to the flow or if it is tumbling [Issacs et al. 2005].

2) Impaction: Impaction occurs when the airstream changes direction abruptly and the 

inertia of a particle causes it to continue in its original direction and collide with the filter 

material. Impaction is analogous to an insect hitting the windshield of a car driving on a 

highway: the air molecules can quickly change direction and move up and over the car, 

but the inertia of the insect causes it to change direction more slowly and impact the 

windshield. The likelihood that a particle will deposit by impaction increases 

proportionally with the density, velocity and diameter2 of the particle. Impaction usually 

is most important for larger particles (around 1 μm and larger) because of their greater 

inertia. 

3) Diffusion: Brownian motion causes small aerosol particles to move randomly and disperse 

within an airstream. If the particles collide with the filter material, they can deposit on it. 

Diffusion is most important for particles of around 0.1 μm and smaller.

4) Electrostatic attraction: Aerosol sampling filters may carry an electrostatic charge, which 

can attract charged airborne particles. Charged filter materials can also attract neutral 

particles by inducing a dipole within the particle, and charged particles can be attracted to 
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Filter Pore Size and Aerosol Sample Collection

neutral filter materials by image forces (forces created when a charged particle induces an 

opposite charge in the filter material). This mechanism is especially important for 

electret-treated filters (filters treated to have permanent electrostatic charges).

5) Sedimentation: Sedimentation (or settling) occurs when particles fall onto filter materials 

because of gravitational forces. Sedimentation is generally significant only for very large 

particles, very slow flow velocities, or if the air is flowing downward into the filter. 

Because of this, few particles are collected by sedimentation during most workplace 

aerosol sampling.

Figure 4: Aerosol particle collection mechanisms. Different types of filters have different 

structures but they all collect particles using the mechanisms shown here. The relative 

importance of the various collection mechanisms depends upon the size, shape, density 

and electrostatic charge of the aerosol particles and the velocity of the air flow through the 

filter.

An example of the collection efficiencies due to each mechanism for different aerosol particle 

sizes is shown in Figure 5. The effectiveness and relative importance of these mechanisms 

depend upon many factors. For example, a higher flow velocity favors impaction by increasing 

the momentum of the particles, whereas a lower velocity allows more time for particles to 

diffuse to the filter surface. A highly charged filter and/or aerosol will encourage electrostatic 

deposition. Fibers and particles with irregular shapes or branching structures are more likely 

to be intercepted. The filter collection efficiency remains high for nanoparticles from 10 nm 

down to at least 2 nm; it is thought that the collection efficiency for nanoparticles smaller than 

2 nm may decrease due to thermal rebound, but this is still being investigated [Givehchi and 

Tan 2014; Wang and Tronville 2014].
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Filter Pore Size and Aerosol Sample Collection

Figure 5: Theoretical collection efficiencies of aerosol particle collection mechanisms for a 

fibrous filter 1 mm thick with 2 μm fibers and an air velocity of 10 cm/sec. Diffusioninterception interaction is the particle collection due to an enhancement of interception by 

particle diffusion. The filter surface is assumed to be horizontal with air flowing 

downward into it, which enhances sedimentation. Total shows the collection efficiency of 

the filter due to all mechanisms combined. Electrostatic collection is not included because 

it is very difficult to model. These calculations were based on “single-fiber efficiency” for 

filters, which is explained in more detail in Hinds [1999]. Figure is adapted from Hinds 

[1999].

5 Aerosol filter efficiency and pore size

Now, with these collection mechanisms in mind, think about the structures of the different 

filter types shown in Figure 1. The fibrous and porous membrane filters do not have simple, 

well-defined pores like a sieve or a simple mesh. Instead, these filters have pathways with a 

broad range of sizes and a variety of irregular shapes. Thus, particles entering these filters are 

forced to follow a meandering path, which greatly increases the likelihood that the particles 

will be intercepted, impact on the filter, or diffuse onto it. For this reason, the probability that 

a particle will be collected by one of these filters is much higher than one might think based 

simply on the stated pore size (which is the equivalent pore diameter of the filters). The 

capillary pore filters provide a more direct pathway through the filter, but even in this case 

interception, impaction, and diffusion act to collect particles smaller than the sizes of the 

pores because of the deposition mechanisms discussed in the previous section and shown in 

Figure 4. 
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Filter Pore Size and Aerosol Sample Collection

The effect of the structures of these filters and the aerosol particle collection mechanisms that 

we have discussed can be seen in Figure 6. This plot shows how well particles of different sizes 

are collected by porous membrane filters, which have tortuous flow paths with equivalent 

pore diameters of 0.3 and 3 μm, and capillary pore filters with pore diameters of 1 and 3 μm. 

Note that for all particle sizes, the collection efficiency was ≥99.7% for the 0.3-μm porous 

membrane filter and ≥98.4% for the 3-μm porous membrane filter, even though the test 

particles were much smaller than the equivalent pore diameters of the filters. The collection 

efficiencies of the capillary pore filters were substantially lower, but these filters were also able 

to collect particles much smaller than their pore sizes. This can also be seen in Figure 7, which 

shows submicron NaCl aerosol particles collected using a 3-μm porous membrane filter. 

These results clearly illustrate that the equivalent pore diameter of a filter does not indicate the 

size of the airborne particles that the filter will collect and that the structure of the filter has a 

much greater effect on the collection characteristics.

Figure 6: Aerosol particle diameter vs. collection efficiency for polytetrafluoroethylene 

(PTFE) porous membrane filters with 0.3-μm and 3-μm equivalent pore diameters, and 

polycarbonate (PC) capillary pore filters with 1-μm and 3-μm pore sizes. The differences 

in performance are not due to the different materials used for the filters, but rather 

because the porous membrane filters have tortuous paths which greatly increase the 

likelihood of particle deposition, while the capillary pore filters have pores that are 

straighter and smoother. The collection efficiency is the percentage of the particles in the 

airstream that are collected by the filter. The face velocity (average flow velocity of air into 

the filter) was 3.5 cm/s for the 0.3-μm PTFE filter and 16 cm/s for the others. The aerosol 

particles were NaCl. Figure is adapted from Burton et al. [2007].
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Filter Pore Size and Aerosol Sample Collection

Figure 7: NaCl aerosol particles collected using a PTFE porous membrane filter with a 

3-μm equivalent pore diameter at a face velocity of 8.3 cm/s. As can be seen, particles much 

smaller than 3 μm were captured by the filter.

It is also of interest to note that the collection efficiencies of the capillary pore filters decreased 

as particle size decreased down to 0.047 and 0.063 μm, and then increased as the particle sizes 

decreased further. This phenomenon is seen with other types of filters as well. As seen in 

Figure 5, this occurs because impaction decreases as the particle size decreases, which causes 

the overall collection efficiency curve to dip downward. However, as particles become even 

smaller, diffusion becomes a more important collection mechanism, and the collection 

efficiency increases. The particle size for which the collection efficiency is lowest is called the 

“most penetrating particle size”, or MPPS. The MPPS for a given filter will vary depending 

upon the air flow rate, the electrostatic charges of the particles and the filter, the amount of 

particles that are deposited on the filter, and other factors [Lee and Liu 1980; Martin and 

Moyer 2000].

6 Significance of pore size

So why is it important to understand pore size? First, an investigator may assume incorrectly 

that an aerosol sample collected by a filter includes only particles larger than the stated pore 

size of the filter, when in fact the filter collected smaller particles as well. This can lead to a 

misinterpretation of test results and a misunderstanding of the actual size characteristics of 

the aerosol being sampled. An investigator also might wrongly try to use a filter with a 

particular stated pore size as a pre-filter to remove larger particles before collecting a sample; 

in this case, many smaller aerosol particles would be removed as well, and the true exposure to 

small particles could be badly underestimated. 
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Filter Pore Size and Aerosol Sample Collection

Second, it may be mistakenly thought that two filters with the same stated pore size have the 

same particle collection characteristics. In fact this is not at all true for filters of different types, 

as seen in Figure 6: the collection efficiency of a porous membrane filter with a given 

equivalent pore diameter can be much higher than a capillary pore filter with the same stated 

pore size. 

Third, a filter with a smaller pore size usually has a higher resistance to flow (and therefore a

higher pressure drop across the filter) than does a filter of the same type with a larger pore size 

[Breuer 2012]. Thus, an aerosol sampling pump has to create a stronger vacuum to pull air at 

the same flow rate through a filter with a smaller pore size. If a filter with a very small pore 

size is selected on the erroneous belief that the small pore size is needed to collect all of the 

airborne particles, then the pump may not be able to reach the desired flow rate or may not be 

able to maintain the desired flow rate as the filter becomes loaded with particles and the flow 

resistance increases. This may cause the collected sample to be smaller than expected. If the 

sampling pump is battery powered, its running time may be greatly reduced, and the pump 

may even shut down prematurely.

7 Filter selection

Given all of this, what is the best way to select an aerosol filter for a particular application? The 

first step is to consider the purpose of the sampling and how the samples will be processed. 

For example, polycarbonate capillary pore filters are often used when samples are to be 

examined using scanning electron microscopy (SEM). Fibers are typically collected using 

mixed cellulose ester filters, which can be rendered transparent for counting by phase-contrast 

microscopy. Alkaline dusts are collected using PTFE filters, which allow for analysis by 

titration. For gravimetric analysis, filters that are not hygroscopic and that have stable weights, 

such as PVC, are needed. The characteristics of the aerosol particles to be collected also 

influence the choice of filter. Bioaerosols, for example, may lose viability due to damage or 

desiccation when collected onto filters. Liquid aerosol droplets behave in much the same 

manner as solid particles while airborne, but once they are collected the liquids can coat the 

fibers and coalesce into larger droplets, which can reduce the collection efficiency of a filter 

[Charvet et al. 2010; Contal et al. 2004]. In addition, oils can mask the charged regions of 

electret-treated filters, which can greatly reduce the collection of particles by electrostatic 

mechanisms [Barrett and Rousseau 1998]. 

The next step is to see if a recommended test method has been published for the aerosol 

particles of interest. Organizations such as the National Institute for Occupational Safety and 

Health (NIOSH), the Occupational Safety and Health Administration (OSHA), the 

Environmental Protection Agency (EPA), the American Conference of Governmental 

Industrial Hygienists (ACGIH), and ASTM International (formerly known as the American 
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Filter Pore Size and Aerosol Sample Collection

Society for Testing and Materials) publish test methods for a variety of aerosols that include 

the characteristics of the filters to be used. For example, in the NIOSH Manual of Analytical 

Methods, Method 0600 for respirable particle sampling specifies the use of a size-selective 

cyclone and a “5.0-μm pore size, polyvinyl chloride filter or equivalent hydrophobic 

membrane filter supported by a cassette filter holder (preferably conductive.)” The NIOSH 

Manual of Analytical Methods also has several chapters discussing different aspects of aerosol 

sampling, including general considerations and factors affecting aerosol sampling, sampling 

bioaerosols, sampling airborne fibers, sampler wall losses, and avoiding bypass leakage in filter 

cassettes [NIOSH 2003; NIOSH 2014].

If a test method is not available, the collection characteristics of different types of filters can be 

found in reference texts such as those by Lippmann [2001] and Raynor et al. [2011]. A search 

of the scientific literature also can produce the results from the testing of various filters to 

collect different kinds of airborne particles. For example, information on the flow resistance of 

many types of filters and sampling tubes can be found in Breuer [2012], and Soo et al. [2016] 

recently tested 29 commercially available aerosol filters and reported their flow resistances and 

collection efficiencies. Filter manufacturers often provide data on the collection characteristics 

of their filters and on recommended filters for various applications. Finally, it is important to 

note that filter collection performance can vary with the flow rate and aerosol particle 

characteristics as well as the filter type and manufacturer. Thus, care should be taken when 

applying results from one sampling situation to a different set of conditions.

8 Conclusion

The equivalent pore diameter provides a helpful way to categorize filters and to test for 

consistency in filter characteristics. However, it should not be construed as an indication of 

the sizes of aerosol particles that will be collected by the filters. A better understanding of the 

meaning of the term pore size, the structures of the different types of filters, and the 

mechanisms by which aerosol particles are collected will help in selecting a filter for a 

particular application and to correctly interpret the results of aerosol sampling.

Acknowledgements

The author thanks Giles Barton of GE Healthcare and Leonid Turkevich, Nancy Burton, 

Stephen Martin and Seleen Collins of NIOSH for their advice and assistance. The filter images 

were taken by Diane Schwegler-Berry of NIOSH.

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 
NIOSH Manual of Analytical Methods  5th Edition  Chapter FP April 2016 Page FP-13 of FP-14

Filter Pore Size and Aerosol Sample Collection

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

9 References

ASTM [2011]. Standard test methods for pore size characteristics of membrane filters by 

bubble point and mean flow pore test. ASTM F316 - 03(2011). West Conshohocken, PA: 

ASTM.

Barrett LW, Rousseau AD [1998]. Aerosol loading performance of electret filter media. AIHA 

J 59(8):532-539. 

Breuer D [2012]. Flow resistance of samplers for personal monitoring in work areas and 

requirements for sampling pump performance. J Occup Environ Hyg 9(2):D25–D32. 

Brock TD [1983]. Membrane filtration: a user's guide and reference manual. Madison, WI: 

Science Tech.

Burton NC, Grinshpun SA, Reponen T [2007]. Physical collection efficiency of filter materials 

for bacteria and viruses. Ann Occup Hyg 51(2):143–151. 

Charvet A, Gonthier Y, Gonze E, Bernis A [2010]. Experimental and modelled efficiencies 

during the filtration of a liquid aerosol with a fibrous medium. Chemical Engineering Science 

65(5):1875-1886. 

Contal P, Simao J, Thomas D, Frising T, Callé S, Appert-Collin JC, Bémer D [2004]. Clogging 

of fibre filters by submicron droplets. Phenomena and influence of operating conditions. J 

Aerosol Sci 35(2):263-278. 

Givehchi R, Tan Z [2014]. An overview of airborne nanoparticle filtration and thermal 

rebound theory. Aerosol Air Qual Res 14(1):45-63.

Hinds WC [1999]. Aerosol technology: properties, behavior, and measurement of airborne 

particles. New York: John Wiley & Sons.

Isaacs KK, Rosati JA, Martonen TB [2005]. Mechanisms of particle deposition. In: Ruzer LS, 

Harley NH, eds. Aerosols Handbook Measurement, Dosimetry, and Health Effects. Boca 

Raton, FL: CRC Press, pp. 75-99. 
NIOSH Manual of Analytical Methods  5th Edition  Chapter FP April 2016 Page FP-14 of FP-14

Filter Pore Size and AerosolSample Collection

Lee KW, Liu BYH [1980]. On the minimum efficiency and the most penetrating particle size 

for fibrous filters. Journal of the Air Pollution Control Association 30(4): 377-381. 

Lippmann M [2001]. Filters and Filter Holders. In: Cohen BS, McCammon CS Jr., eds. Air 

Sampling Instruments for Evaluation of Atmospheric Contaminants. Cincinnati, OH: 

American Conference of Governmental Industrial Hygienists, pp. 281-314. 

Martin SB Jr., Moyer ES [2000]. Electrostatic respirator filter media: filter efficiency and most 

penetrating particle size effects. Appl Occup Environ Hyg 15(8):609-617. 

NIOSH [2003]. NIOSH manual of analytical methods (NMAM). 4th ed., 3rd supplement. 

Schlecht PC, O’Connor PF, eds. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication 2003-154 

[http://www.cdc.gov/niosh/docs/2003-154/]. 

NIOSH [2014]. NIOSH manual of analytical methods (NMAM). 5th ed. Ashley K, O’Connor 

PF, eds. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 2014-151 [http://www.cdc.gov/niosh/nmam/].

Raynor PC, Leith D, Lee KW, Mukund R [2011]. Sampling and analysis using filters. In: 

Kulkarni P, Baron PA, Willeke K, eds. Aerosol measurement : principles, techniques, and 

applications. Hoboken, NJ: John Wiley & Sons, pp. 107-128. 

Soo J-C, Monaghan K, Lee T, Kashon M, Harper M [2016]. Air sampling filtration media: 

Collection efficiency for respirable size-selective sampling. Aerosol Sci Technol 50(1):76-87. 

Wang J, Tronville P [2014]. Toward standardized test methods to determine the effectiveness 

of filtration media against airborne nanoparticles. Journal of Nanoparticle Research 16(6):1-

33.
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Measurement of Fibers

by Paul A. Baron, Ph.D., NIOSH

Adapted from Baron [2001] 

The NMAM team gratefully acknowledges Chen Wang, Joe Fernback and Alan Dozier for 

insightful review of this chapter.

1 Introduction FI-2

2 Fiber dimensions FI-5

3 Phase contrasting light microscope counting (PCM) FI-6

4 Polarizing light microscopy (PLM) of bulk materials FI-14

5 Electron microscopy FI-17

6 Scanning electron microscopy (SEM) FI-18

7 Transmission electron microscopy (TEM) FI-18

8 Optical detection (light scattering) FI-20

9 Fiber classification FI-21

10 Conclusions FI-22

11 References FI-23
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Measurement of Fibers

1 Introduction 

Fiber-related disease has provided much of the impetus for fiber research in recent years. 

Asbestos has been the fiber type most commonly associated with disease. The name “asbestos” 

is a commercial term applied to the fibrous forms of several minerals that have been used for 

similar purposes and includes chrysotile, amosite, crocidolite, and the fibrous forms of 

tremolite, anthophyllite, and actinolite. The three primary diseases associated with asbestos 

exposure are asbestosis, the result of inflammation and collagen formation in lung tissue; lung 

cancer; and mesothelioma, an otherwise rare form of cancer associated with the lining 

surrounding the lungs. A current theory describing the toxicity of fibers indicates that fiber 

dose, fiber dimension, and fiber durability in lung fluid are the three primary factors 

determining fiber toxicity [Lippmann 1990].

The dose, or number of fibers deposited in the lungs, is clearly an important factor in 

determining the likelihood of disease. Both fiber diameter and length are important in the 

deposition of fibers in the lungs and how long they are likely to remain in the lungs. Figure 1 

indicates some of the factors that determine fiber deposition and removal in the lungs. Fiber 

length is thought to be important because the macrophages that normally remove particles 

from the lungs cannot engulf fibers having lengths greater than the macrophage diameter.

 

Thus, longer fibers are more likely to remain in the lungs for an extended period of time. The 

macrophages die in the process of trying to engulf the fibers and release inflammatory 

cytokines and other chemicals into the lungs [Blake et al. 1997]. This and other cellular 

interactions with the fibers appear to trigger the collagen buildup in the lungs known as 

fibrosis or asbestosis and, over a longer period, produce cancer as well. Fiber diameter is also 

important because fiber aerodynamic behavior indicates that only small diameter fibers are 

likely to reach into and deposit in the airways of the lungs. The smaller the fiber diameter, the 

greater its likelihood of reaching the gas exchange regions. Finally, fibers that dissolve in lung 

fluid in a matter of weeks or months, such as certain glass fibers, appear to be somewhat less 

toxic than more insoluble fibers. The surface properties of fibers are also thought to have an 

effect on toxicity. Asbestos is one of the most widely studied toxic materials and there have 

been many symposia dedicated to and reviews of its behavior in humans and animals [Selikoff 

and Lee 1978; Rajhans and Sullivan 1981; WHO 1986; ATSDR 1990; Dement 1990].
NIOSH Manual of Analytical Methods  5th Edition  Chapter FI April 2016 Page FI-3 of FI-31

Measurement of Fibers

Figure 1. Schematic of mechanisms that affect fiber deposition and retention in the lungs. 

The deposition depends on all the indicated parameters in a complex fashion. However, 

larger diameter particles are affected more by gravitational settling, impaction, and 

interception, resulting in greater deposition further up in the respiratory tract. The saddle 

points, or carinae, in the branching respiratory tree are often a focal point for deposition of 

larger diameter fibers. Smaller diameter particles are affected more by diffusion and can 

collect in the smaller airways and gas exchange region (alveoli). Particle removal from the 

lungs is primarily effected by the cilia coating the non-gas exchange regions of the lungs; the 

cilia push mucus produced in the lungs and any particles trapped in the mucus out of the 

lung and into the gastrointestinal tract in a matter of hours or days. Some fibers are 

sufficiently soluble in lung fluid that they can disappear in a matter of months. Finally, white 

blood cells or macrophages roam the gas exchange regions and ingest particles deposited 

there for removal through the lymph system. Human macrophages are approximately 17 μm 

in diameter and can only ingest particles smaller than they are. Therefore, thin fibers are 

likely to deposit in the gas exchange region and, of these, the long insoluble fibers can remain 

in the lungs indefinitely.
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Measurement of Fibers

Several techniques were used for asbestos measurement up until the late 1960s [Rajhans and 

Sullivan 1981]. Earlier than this, it was not widely recognized that the fibrous nature of 

asbestos was intimately related to its toxicity, so many techniques involved collection of 

airborne particles and counting all large particles at low magnification by optical microscopy. 

Thermal precipitators, impactors (konimeters), impingers, and electrostatic precipitators were 

all used to sample asbestos. Perhaps the primary technique in the United States (US) and the 

United Kingdom (UK) during this early period was the liquid impinger, in which particles of 

dust larger than about 1-μm aerodynamic diameter were sampled at 2.7 L/min and impacted 

into a liquid reservoir [Rajhans and Sullivan 1981]. After sampling, an aliquot of the liquid 

was placed on a slide in a special cell, particles larger than 5-μm size were counted, and the 

results were reported in millions of particles per cubic foot. Dissatisfaction with this approach 

stemmed from lack of correlation between measured particle concentration and disease in the 

workplace. Various indices of exposure have been developed that attempt to relate a portion 

of the fiber size distribution to the toxic effects. The appropriate indices for each of the 

asbestos related diseases as a function of fiber length and diameter (Figure 2) were suggested 

by Lippmann [Lippmann 1988].

Figure 2. Comparison of proposed size ranges of asbestos fibers causing specific diseases 

compared with the fiber sizes detected using TEM and PCM techniques. Lung cancer and

mesothelioma are more likely to occur at current occupational and environmental levels

than asbestosis. PCM can cover only a portion of the total fiber distribution; PCM is used

as an indicator of total exposure. TEM can cover the entire size range, but most methods

emphasize one size range over another through selection of magnification and counting

rules.
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Measurement of Fibers

2 Fiber dimensions

Fibers are particles that have one dimension significantly larger than the other two. Fibers are 

often characterized or selected according to their aspect ratio, i.e., the ratio of the large 

dimension to one of the small dimensions. If no other criteria are used, then materials that 

might not normally be considered fibrous may contain a fraction of particles that meet the 

criteria for fibers. The distribution of fiber dimensions in a sample can usually be 

characterized by assuming a cylindrical geometry (i.e., the two small dimensions are identical) 

and measuring the length and diameter of individual fibers. The distribution of airborne fiber 

sizes generated by grinding bulk material or by mechanically releasing particles into the air 

often results in a two-dimensional (bivariate) lognormal distribution. Such a distribution is 

characterized by five parameters: the geometric mean length, the geometric mean diameter, 

the length and diameter geometric standard deviations, and a correlation term that relates 

length to diameter [Schneider et al. 1983]. In addition, several other parameters that are a 

function of length and diameter, such as aerodynamic diameter, can also be characterized by a 

lognormal distribution [Cheng 1986].

 

Often the discussion of fibers assumes that fibers are straight objects that can be well defined 

by several parameters as indicated above. However, many real-world particles are not so 

simple to describe. In fact, the detailed features of many fibers can aid in their identification 

[McCrone 1980]. Fibers are often curved, have splayed ends, or differ in other ways from a 

cylindrical shape. Asbestos mineral is composed of fibrils (about 0.03-μm diameter) that are 

packed together. This fibrillar structure is characteristic of asbestiform minerals. When the 

mineral is broken apart mechanically, the material separates primarily between fibrils and the 

resulting fibers are usually bundles of fibrils. The ends of the fibers can be broken apart, with 

smaller bundles or individual fibrils spread apart, yet still be part of the fiber. Fibers can be 

contaminated by attachment of other dust particles, creating a complex structure with 

aerodynamic behavior not matching that of cylindrical fibers. The complexity of fiber shapes 

affects all of the measurement and separation techniques described below and frequently 

makes it difficult to compare one method to another.

In addition to asbestos fibers, there are many types of fibrous materials being produced for 

commercial purposes. These include fibrous glass, mineral wool, refractory ceramic fibers, 

wood and other plant fibers, and synthetic organic fibers. Most of these fibers tend to have 

larger diameters than asbestos fibers. On the other hand, carbon nanotubes (CNTs) 

(<0.005um diameter) have recently been produced in small-scale commercial quantities and 

because of their high tensile strength, high conductivity, and other special properties, show 

great promise as a commercial material [Liu et al. 1998]. Unlike asbestos fibers, which have 

discrete lengths and diameters (i.e., aspect ratios), CNTs occur mainly as entangled particle 

agglomerates and may contain varying amounts of amorphous carbon and residual catalyst 
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Measurement of Fibers

metal. The complexity and variety of structures makes CNT particle counting a challenge. 

Measurement techniques must be tailored to the size distribution and physicochemical 

properties of these materials.

This review primarily relates to measurement of fibers in air. There are several techniques that 

address concentration of asbestos and other fibers in bulk material and measurement of mass 

concentration of fibers [Beard and Rook 2001]. One of these bulk methods, polarizing light 

microscopy, will be discussed below.

3 Phase contrasting light microscope counting 

(PCM)

As asbestos-induced disease became widely studied in the 1960s, cellulose-based membrane 

filter sampling was applied to asbestos sampling in combination with high magnification 

phase contrast light microscopy (PCM) for counting fibers. This technique involved collection 

of fibers uniformly over the surface of a cellulose ester filter, placing the filter or a segment of 

the filter on a microscope slide and making it transparent, and observing the fibers in the 

sample with a high magnification (~450X) phase contrast light microscope. Over the years, 

many researchers have endeavored to improve and standardize the PCM method. One 

researcher, Walton, discussed many aspects of this technique in a review [Walton 1982]. The 

high variability of the analysis results and the method’s dependence on operator technique 

made method improvement and research difficult. The PCM method does not measure all 

fibers; typically only those >0.25 μm diameter are visible and counted and only those >5 μm 

length are counted by protocol. Therefore, the PCM method is only an index of exposure and 

uses the assumption that what is detected is correlated with the fibers actually causing disease 

(Figure 2). The PCM method does not allow identification of asbestos fibers. This is an 

important limitation when the method is used in settings where fiber concentrations with a 

significant non-asbestos fraction may occur. This should be remembered when considering 

some of the parameters discussed below. The aim of evaluating changes to the PCM technique 

may depend on whether consistency with other laboratories within a country or throughout 

the world is more important than making measurements that are more closely related to 

health effects. A number of factors which influence analysis results have been investigated, 

including the following: 
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a. Microscope-related parameters 

1.) Microscope magnification

The exact level of microscope magnification depends on microscope design, but most 

current methods use 450X (±10%) total magnification. Pang and coworkers 

investigated 1250X magnification to improve fiber detectability, but this has not been 

adopted in any established methods [Pang et al. 1989]. Pang also investigated the effect 

of using lower magnification (400X) and found that counts were lower for chrysotile 

asbestos by 25%, but that amosite fiber counts were unaffected [Pang 2000].

2.) Phase contrast optics

This contrast enhancement technique allows detection of asbestos fibers down to

about 0.25 μm diameter for chrysotile and about 0.15 μm for amphiboles. Other 

techniques such as dark field microscopy may offer improved detectability, but also

increase the background from non-fibrous particles.

3.) Test slide to check optics

A test slide was developed to allow a check of proper alignment and magnification in 

the microscope [LeGuen et al. 1984]. This ensures a reasonable level of uniformity in 

microscope setup and operation, including the operator’s visual perception. Improper 

setup can reduce detectability of fibers. There have also been cases where the optics 

were “too good,” and results were obtained that were higher than the reference count.

4.) Counting area in microscope field

Some early measurements with the phase contrast microscope were made using a 

rectangular graticule for defining the counting area, while others were made using the 

entire microscope viewing area. It was found that larger viewing areas resulted in 

lower counts, so the Walton-Beckett graticule [Walton and Beckett 1977] was 

developed that nominally gave a 100-μm diameter counting area (the area is calibrated 

more precisely for each microscope) and has been incorporated in all current methods.

b. Sample preparation techniques 

1.) Filter type 

Virtually all measurements are made using 0.8-μm pore size mixed cellulose ester 

(MCE) filters. Some measurements are made using 1.2-μm pore size filters when 

sampling low concentrations to allow higher flow rate through the filter. Smaller pore 

size filters are used to ensure that fibers are deposited as near the surface of the filters 

as possible. This results in fibers ending up in the same plane so that they can be 

readily viewed with a minimum change of focus during fiber counting. Pore sizes 
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smaller than 0.8 μm are only used with line-operated pumps because of limited suction 

power available with personal sampling pumps.

2.) Selection of the liquid for making filter transparent

A liquid is placed on the filter that closely matches the filter refractive index, yet has an 

index that is as far as possible from that of the fibers being detected. Rooker et al. 

showed that refractive index difference between cleared filter and fibers translated 

directly into detectability of small diameter fibers [Rooker et al. 1982]. A viscous 

solution of dimethyl phthalate and diethyl oxalate mixed with cellulose filter material 

was commonly used in the 1970s and early 1980s. However, it did not result in a 

permanent sample, with crystallization of the mount and movement of fibers often 

occurring several days after sample preparation. Permanent slides were needed for 

quality assurance purposes and the sample preparation technique was also slow and 

required some skill. A rapid acetone-based filter clearing technique was developed that 

could be used safely in field situations [Baron and Pickford 1986]. After clearing, filters 

were coated with triacetin to surround the fibers. This resulted in a longer lasting 

sample (typically months to years) and is currently specified in most methods. 

Another technique uses a resin called Euparal to surround the fibers and results in a 

permanent slide preparation [Ogden et al. 1986].

3.) Filter loading

The number of fibers on the filter is usually specified to be within a certain loading 

range to ensure consistent counting. Cherrie et al. demonstrated using a serial dilution 

technique that counting efficiency was a function of concentration of fibers on the 

filter [Cherrie et al. 1986]. At very low filter loadings (<100 fibers/mm2

) there was a 

tendency to count high relative to an intermediate range of concentrations (100-1300 

fibers/mm2), where the counts were a linear function of loading. This “overcounting” 

was apparently due to greater visibility of fibers in a clean visual field. This effect was 

noted for both human counters and an image analysis system. At high filter loadings 

(>1300 fibers/mm2

), undercounting occurred due to overlap of fibers with other fibers 

and with nonfibrous particles. Most published methods indicate that optimum 

counting occurs within the 100-1300 fibers/mm2 range, while some restrict the range 

further to less than 650 fibers/mm2

. 

4.) Fiber counting rules

The basic fiber counting rules for most current methods indicate that a countable fiber 

should be longer than 5 μm, narrower than 3 μm, and have an aspect ratio greater than 

3:1. These rules were selected because shorter fibers were difficult to detect by optical 

microscopy and the 3:1 aspect ratio was used to discriminate between fibrous and non-
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fibrous particles in occupational settings. There has been a great deal of controversy 

over these rules. The use of a longer fiber cutoff, e.g., 15-20 μm, has been suggested, 

based on two separate arguments: first, that most asbestos fibers are relatively long and 

thin (with high aspect ratio) and the longer fiber cutoff would discriminate better 

toward fibers that were truly asbestos fibers according to mineralogical definitions 

[Wylie 1979]; and second, that fibers that enter the lungs are removed readily by 

macrophages if they are shorter than about 15 μm [Blake et al. 1997]. Longer fibers 

cannot readily be engulfed by macrophages, thus staying in the lungs for a long period 

and causing continuing fibrosis.

The aspect ratio criterion has also been questioned because many non-asbestiform 

particles have shape distributions that include particles with aspect ratios greater than 

3:1. Since asbestos and other minerals often contain single crystal particles not in the 

asbestiform habit, it has been argued that these single crystals, or cleavage fragments, 

should not be counted. However, the Occupational Health and Safety Administration 

(OHSA) has supported the 3:1 minimum aspect ratio through legal precedent. The 

National Institute for Occupational Safety and Health (NIOSH) has noted that because 

of the great difficulty in differentiating whether individual high aspect ratio particles 

are cleavage fragments or asbestiform fibers, all such particles should be counted. 

These high aspect ratio particles may cause disease whether or not they are 

asbestiform.

Other aspects of fiber counting have been investigated, including how to count nonstandard fiber shapes, overlapping fibers, overlapping compact particles on fibers, and 

bundles of fibers. Each of these factors can have a noticeable effect on the final count. 

Cowie and Crawford investigated the effect of some of these factors and estimated 

most of them made a difference in the final count on the order of 20% [Cowie and 

Crawford 1982]. Many of the methods currently in use have slight variations in their 

interpretation of which fibers to count and thus can contribute to variation in results 

between countries and organizations.

NIOSH Method 7400 contains two sets of counting rules, the A and the B rules. The A 

rules are used for asbestos and are consistent with counting rules in previous NIOSH 

methods. The A rules are required for asbestos counting by OSHA because of legal 

precedent in regard to the 3:1 aspect ratio rule. The A rules do not have an upper 

diameter limit for fibers to be counted. The B rules were introduced as an alternative 

to the A rules when Cowie and Crawford found that these rules agreed best with 

previous PCM counts, yet had improved precision [Cowie and Crawford 1982]. The B 

rules have been informally adopted for use with fibers other than asbestos because 

these rules include the upper diameter limit of 3 μm. This upper diameter limit 
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significantly reduces the counting of typically large-diameter fibers, e.g., glass and 

cellulose, which are unlikely to deposit in the lungs [Breysse et al. 1999].

c. Quality assurance schemes 

1.) Sample recounts

Most methods require individual counters to recount about 10% of the field samples to 

ensure consistent counting procedures and alert the analyst in the case of problem 

samples. It is also recommended that counters have samples that are routinely 

recounted to ensure consistent counting within a laboratory over time.

One of the difficulties in analyzing errors made by analysts during PCM counting is 

that individual fields are difficult to relocate after the analyst has finished counting a 

slide. Differences in counts between analysts have often been ascribed to local 

variations in loading on the filter. Pang’s development of a slide coverslip that defines 

counting areas on the sample solves this problem [Pang 2000]. Areas on the coverslip 

are vacuum coated with a thin layer of gold and platinum using an electron 

microscope grid as a mask. This leaves defined areas on the coverslip that can be 

located by grid index marks. Thus, specific fields in a sample can be readily located. 

Using this grid mapping approach, the location, orientation and shape of each fiber 

can be noted and differences in counts can be reconciled on a fiber-by-fiber basis. The 

coverslips have been used to study fiber counting accuracy by comparing routine 

counting of specified fields to counts agreed upon by a group of competent counters. It 

was found that the principal errors for chrysotile fiber samples were due to missing 

fibers close to the visibility limit, while the principal errors for amosite fiber samples 

were caused by incorrectly sizing fiber length near the 5-μm limit. The chrysotile 

samples were therefore typically undercounted (negative bias), while the amosite 

samples had increased variability with individual counters being biased either high or 

low. Both these errors can be reduced by training counters with pre-counted reference 

slides prepared using Pang’s coverslips [Pang 2000] (Omega Specialty Instrument Co. 

Chelmsford MA). In addition, these reference slides can be used on a routine basis to 

ensure consistency in counting. These coverslips or modified versions show great 

promise for training analysts and perhaps for improving quality assurance schemes.

2.) Interlaboratory sample exchanges

Crawford et al. found that use of sample exchange programs was more important in 

ensuring agreement between laboratories than similarity in details of the counting 

rules [Crawford et al. 1982]. Thus, exchange of field samples between laboratories is

commonly performed to improve consistency of counting. A description of several 

quality assurance techniques for asbestos fiber counting is described by Abell et al. 
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[Abell et al. 1989]. To fulfill Method 7400 requirements for an interlaboratory sample 

exchange, Tombes and Calpin have described a simple approach using appropriate 

statistical tests [Tombes and Calpin 2002].

3.) Quality check samples

In order to get agreement between laboratories within a country or internationally, 

several programs send out identical samples to participating laboratories to assess their 

relative performance [Schlecht and Shulman 1986; Kauffer 1989; Crawford et al. 1992; 

Arroyo and Rojo 1998]. These programs provide feedback, often tied to laboratory 

accreditation, which provides incentive for laboratories to ensure that their 

performance is similar to that of other laboratories.

d. Qualitative fiber analysis 

In addition to simply counting the fibers, there are techniques available for providing at 

least tentative identification of fiber type; use of these techniques is commonly called 

differential counting. Fiber shape can be used to limit the type of fiber counted. For 

instance, glass fibers tend to be straighter, with smoother sides than chrysotile fibers. 

Polarizing light techniques can also be used to identify larger diameter (> 1 μm) fibers. 

These are based on the optical properties of the materials, including refractive index and 

crystallinity. These techniques can provide quite positive identification for the presence of 

certain types of fibers, but are limited in application to airborne fibers because they only 

work for the larger diameter fibers. These techniques are often used in analysis of bulk 

materials [NIOSH 1994a]. The use of identification techniques is not allowed in reporting 

fiber counts using Method 7400 so that the results are consistent between laboratories. 

Considerable confusion has been caused in the past by individual laboratories using some 

of these identification techniques to change the counting procedure and, hence, the final 

results. 

Several PCM fiber counting methods have been published by national [NHMRC 1976; 

HSE 1990] and international organizations [Asbestos International Association 1979; 

WHO 1997]. Most countries have methods very similar to the ones referenced here.

e. Sampling volume for asbestos abatement applications 

Sampling for asbestos after abatement requires the selection of a sampling volume so that 

one can have high confidence that the air meets acceptable concentration standards. The 

following is an example of how to calculate this sampling volume.
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The approach assumes that one wishes to select sampling parameters in order to have a 

high degree of confidence that a target exposure standard (e.g. NIOSH REL, OSHA PEL, 

EPA clearance standard) is met.

Several factors need to be established in order to perform this calculation if the target 

exposure standard involves clearance monitoring. The U.S. Environmental Protection 

Agency (EPA) authorizes the use of PCM for some clearance monitoring applications and 

specifies that a level of 0.01 fibers/mL be met. On the method synopsis page, Method 7400 

indicates that the limit of detection (LOD) for PCM analysis is 5.5 fibers/100 fields. This is 

based on intralaboratory variability. A major difference between Method 7400 and other 

analytical methods in the NIOSH Manual of Analytical Methods (NMAM) is that there is 

no reference method for Method 7400. Therefore, the consensus mean is the “true” value 

and the interlaboratory results effectively define the method accuracy. Under the heading 

“Evaluation of Method, B. Interlaboratory comparability,” Method 7400 provides a means 

of calculating the confidence limits on a single analysis result (Equations 3 and 4). From 

Equation 3, the interlaboratory variability at the LOD is such that the upper 95% 

confidence limit on a measured value is 300% greater than (or 4 times) the measured 

value.

Using the upper confidence limit, the equation in Section 21 in Method 7400 can be used 

to estimate the sampling volume.

nnnnnnnnnnnn oooo ff ff

aa aa oooo 100 ff ffffff ∗ tt tt ffff aa aa

ss ss vv = tt tt4

With the appropriate values inserted, the equation becomes

5.5 ff ff

0.785 2 ∗ 385 2

ss ss vv = 0.01 ff

mm

4

Solving this equation for sampling volume gives 1080 L. This is the minimum volume that 

will give a result allowing a single sample to indicate compliance with the 0.01 fiber/mL 

limit with 95% confidence. It requires that the sample give a result less than or equal to the 

LOD or 5.5 fibers per 100 fields. A higher fiber count may still indicate that the 

concentration meets the target level, but not with the same level of confidence. This is 

likely to be a conservative estimate of concentration and additionally ensure compliance 

with the standard because the fiber concentration is low and, as indicated above, low fiber 
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Measurement of Fibers

loadings are usually overestimated. However, the background concentration of nonfibrous dust on the filter also must be low to ensure that fibers are not obscured.

f. Other techniques 

Since fiber counting by human analysts produces relatively high biases and variability, 

several researchers have attempted to develop automated counting systems. With the 

increases in computer power over the last 25 years, it has been tempting to assume that 

fiber counting is a solvable problem and significant efforts have been made to develop 

such a system. The most intensive effort to produce a fiber counting system was carried 

out by Manchester University in collaboration with the Health and Safety Executive in the 

UK [Kenny 1984]. The Manchester Asbestos Program (MAP) was able to give reasonably 

good agreement with human counters for certain types of samples. It was used as a 

reference analyst for the US and UK reference sample programs for several years. 

Eventually, the MAP was dropped as the reference because it was not sufficiently 

consistent for all types of samples.

 

The principle problems with image analysis of asbestos fibers include: the complexity of 

many fiber shapes, including bundles, agglomerates, and split fibers; the fibers often go in 

and out of the plane of focus; the background includes many particles and other nonfibrous shapes; the phase contrast optics produces haloes around particles in the sample 

that can be detected as fibers; and finally, and perhaps most importantly, the contrast 

between the fibers and background is poor and many fibers are near the detection 

threshold. An evaluation of the MAP program indicated that a significant fraction of the 

fibers were misidentified as multiple fibers, not detected at all, and groups of compact 

particles or edges of large particles were detected as fibers [Baron and Shulman 1987].

Inoue and coworkers have more recently developed image analysis software using a 

microprocessor-based PC [Inoue et al. 1998]. Initial tests indicate that it works 

approximately as well as human counters. Inoue also evaluated how well human counters 

and the image analyzer did in detecting the same fibers in a sample and found that only 

about 50% of the fibers were consistently counted by all counters, so the image analysis 

system did approximately as well as the human counters [Inoue et al. 1999]. Further 

testing of the image analysis system is needed.

In addition to image analysis, optical microscopy can be enhanced using a personal 

computer to more easily observe the image and to mark and measure fiber dimensions, 

with automatic recording of the fibers counted [Lundgren et al. 1995]. This does not 

appear to improve the counting accuracy since the analyst still decides which fibers are to 

be counted.
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4 Polarizing light microscopy (PLM) of bulk 

materials (Adapted from Baron [1993])

The asbestos fibers in bulk material can be released and become airborne when the bulk 

material is disturbed. For this reason, it is desirable to measure the asbestos content of bulk 

samples. PLM is often used to determine the percent asbestos in bulk material. The EPA 

[Asbestos-containing materials, 1987] has defined asbestos containing material (ACM) as 

material containing more than 1% asbestos using the PLM method, which effectively 

estimates concentration by area observed. Some confusion exists regarding the units of 

asbestos percentage. EPA originally indicated that the limit for ACM was 1% by mass 

[Asbestos-containing materials, 1987], but because of the difficulties in determining 

corrections for differences in material density and in determining particle volumes, the limit 

was changed to 1% by area as determined by the PLM method [EPA 1990b]. OSHA does not 

specify units for percent asbestos in its regulations [OSHA 1994].

Several PLM techniques are used for identifying fiber type as well as semi-quantifying the 

percent fibrous material (usually asbestos) in a sample [McCrone et al. 1978; Middleton 1979; 

Asbestos-containing materials, 1990; Perkins and Harvey 1993; NIOSH 1994a]. These 

techniques depend on particle shape, the refractive index, and other optical properties of 

individual particles. Many of these PLM techniques require visual observation of color in the 

fiber and become less reliable for fibers thinner than about 1 μm [Vaughan et al. 1981]. 

a. Sampling 

Several procedures have been suggested for obtaining representative bulk samples of ACM 

in a fashion that prevents unnecessary exposure to asbestos aerosol [EPA 1985a,b; 

Jankovic 1985]. Representative sampling of commercial ACM materials is often 

problematic; these materials may vary significantly in asbestos concentration between 

nearby locations and even at different depths at the same location. Sampling from 

multiple locations and compositing samples helps improve the likelihood of obtaining a 

representative sample.

The material should be wetted or sealed during sample removal. A small coring device, 

such as a cork borer, can be used to obtain a sample from the full depth of the material. At 

least three samples per 1000 ft2 of ACM should be taken [Asbestos-containing materials, 

1987]. The sample should be placed in a well-sealed, rugged container. Finally, the 

sampled area should be repaired or sealed to minimize further fiber release.
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Surface sampling has been proposed by several groups, but there is no relationship 

between airborne fibers and those found on surfaces [Chatfield 2000]. Therefore, surface 

sampling for fibers is not recommended.

b. Sample preparation analysis 

Sample preparation for a PLM analysis involves grinding the material to the optimum 

particle size range (1-15-μm diameter) and dispersing the particles in a liquid of known 

refractive index on a glass slide [Perkins and Harvey 1993]. Particle size uniformity in the 

prepared sample is extremely important. A few large chunks of material may contain more 

asbestos than hundreds of much smaller particles. Friable material, i.e., that which is 

crumbly or can be crushed by hand, may readily release fibers and is considered more 

hazardous. Friable materials are generally easier to prepare for analysis than some other 

ACMs, such as vinyl asbestos floor tiles, which may require dissolution or ashing of the 

matrix material so that the fibers are separated and visible in the microscope. Before and 

after preparation, the sample is observed with a stereomicroscope at 10-100X 

magnification to evaluate sample uniformity and observe whether fibrous material is 

present.

Some materials that interfere with accurate fiber identification either by their similarity or 

by covering up the fibers can be removed by physical treatment of the sample. For 

instance, organic materials, such as cellulose fibers or diesel soot can be removed by low 

temperature, oxygen-plasma ashing [Baron and Platek 1990]. Leather fibers and chrysotile 

have a similar appearance and refractive index. The leather can be removed by ashing at 

400°C [Churchyard and Copeland 1988].

Fiber morphology, i.e. the structure and shape of the fiber, can be used to assist in its 

identification. Morphology of fibers can give some indication of fiber type. For instance, 

chrysotile fibers tend to be curly, while amphibole fibers are straight, especially when they 

are shorter than 50 μm. Asbestos fibers often have frayed or split ends, while glass or 

mineral wool fibers are typically straight or slightly curved with fractured or bulbous ends. 

Many plant fibers are flattened and twisted, with diameters between 5-20 μm. Note that it 

is not recommended to base identification solely on morphology.

Fiber refractive index and other crystalline properties can be used to identify fiber type 

with reasonable certainty. Several techniques for determining these properties can be used 

in a polarizing light microscope. When viewed in the microscope with crossed polarizing 

filters, isotropic (isometric or amorphous) fibers appear consistently bright when rotated, 

while anisotropic (uni- or biaxial crystal structure) fibers appear bright, but disappear 

when rotated to their extinction angle, which is a function of crystal structure. Thus, 
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amorphous materials such as glass or mineral wool fibers can easily be discriminated from 

asbestos.

 

During PLM analysis, fibers are immersed in a fluid selected to have a known refractive 

index. When a fiber has a larger refractive index than the surrounding fluid medium, the 

bright halo (Becke line) around that fiber appears to move into it as the microscope focus 

is raised; when the fiber has a smaller refractive index, the Becke line moves out of it. 

Placing the fibrous material into several different refractive index fluids allows the fiber 

refractive index to be bracketed.

Dispersion, or refractive index change with wavelength, of a fiber can be used for 

identification. When particles are placed in a liquid whose dispersion is different from that 

of the particle, the particle may exhibit a color caused by the refraction of light. This 

technique requires the use of special "dispersion staining" optics. By using several 

refractive index liquids in series, the refractive index and the dispersion of the fiber can be 

established and compared with those of standard materials or published data [McCrone 

1980].

Once the sample has been uniformly dispersed on a slide in the appropriate refractive 

index liquid, specific fiber types, e.g., asbestos, can be identified and the percent fibers 

estimated. Two approaches are typically used: visual comparison with prepared reference 

slides or pictures and point counting. When attempting to estimate whether a material is 

ACM (i.e., > 1% asbestos), the visual comparison technique is adequate when more than 

about 10% of the particles observed are asbestos. Point counting is used for lower 

concentration samples to provide higher accuracy [EPA 1990a]. It involves observing 400 

or more randomly selected "points" (identified with a reticle crosshair) in the sample. The 

number of points containing asbestos is divided by the total number of points observed to 

give the percent asbestos. A combination of these approaches balances the analysis time 

and accuracy of the results [Webber et al. 1990].

PLM also can be used for qualitative analysis of air sample filters by collapsing the filter 

and using low temperature plasma etching of the surface to expose the fibers. Various 

refractive index liquids can then be placed on the etched surface to surround the fibers, 

allowing techniques noted above to be used [Vaughan et al. 1981]. The smallest fibers that 

can be identified by this method are about 1-μm diameter.

c. Accuracy 

PLM analysis is primarily used for qualitative identification of fiber type. Accurate 

identification of asbestos and other fibers requires proper training in the crystallographic 
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Measurement of Fibers

properties of particles as well as training and familiarization with the PLM. As with fiber 

counting, a laboratory quality assurance program is necessary to ensure consistently 

accurate results. The National Voluntary Laboratory Accreditation Program (NVLAP) 

operated by the National Institute for Standards and Technology (NIST) inspects 

laboratories for proper practice as well as providing unknown samples four times a year to 

check their performance in fiber identification. Under a predecessor to this program, 

approximately 350 laboratories correctly classified 98.5% of the samples as asbestos and 

correctly identified the specific asbestos types in approximately 97% of the samples. A 

blind test of 51 laboratories resulted in 97.5% correct classifications and 79.1% correct 

identifications [EPA 1986]. The American Industrial Hygiene Association Proficiency 

Analytical Testing Program provides similar PLM audit samples to laboratories. Some 

common interferences for bulk analysis by PLM include sepiolite, vermiculite, and 

cleavage fragments of non-asbestos amphiboles.

 

PLM has been cast in a quantitative measurement role by the EPA requirement of 

determining whether a school building material meets the 1% asbestos level defining 

ACM. Many variables including particle size, density and shape are not adequately 

controlled or measured in the analysis and contribute to errors in the percent mass 

estimate. Thus, PLM analysis is at best a semi-quantitative technique.

Chatfield indicated that the accuracy of PLM for low concentrations of asbestos was poor 

and described a set of procedures that concentrated the asbestos into a weighable fraction 

[Chatfield 2000]. An EPA report describes, in addition to the PLM and Chatfield’s 

gravimetry methods, a TEM and an x-ray diffraction method for bulk analysis of asbestos 

[Chatfield 2000]. NIOSH Method 9000 describes an x-ray diffraction method for 

chrysotile [NIOSH 1994c].

5 Electron microscopy

Scanning electron microscopy (SEM) has not been the focus of as much method development 

as either light microscopy or transmission electron microscopy (TEM). PCM found favor 

because of the low equipment cost and lower training level required for analysis. TEM is 

preferred for environmental and research studies because it offers the highest resolution and 

the most positive identification capabilities. TEM allows visibility of all asbestos fibers down 

to the individual fibrils, electron diffraction for crystal structure identification, and energy 

dispersive x-ray analysis for elemental measurement. SEM has intermediate resolution, with 

many instruments of this type not able to see all asbestos fibers. However, many modern 

SEMs have the capability of detecting asbestos fibrils, though contrast with background may 

be poor for some fiber types, especially if a high contrast substrate is not used. Energy 

dispersive x-ray analysis is also available for many SEMs, providing some qualitative 
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Measurement of Fibers

information of fiber type. However, since electron diffraction typically cannot be performed 

by SEM, this often leaves open the question of positive identification of fibers.

6 Scanning electron microscopy (SEM)

Particles are observed in the SEM when a beam of electrons is focused onto the sample surface 

and scanned over an area. The electrons are scattered from the surface and detected above the 

surface synchronously with the beam scan rate and an image of the scanned surface is created. 

Thus, the SEM measures the surface of particles on a substrate. The best image can be 

obtained on conducting objects deposited on a smooth, conducting substrate. Particles are 

often deposited on aluminum or carbon planchets that fit directly into the SEM or onto 

polycarbonate membrane (track-etched, Nuclepore®) filters. The samples are usually coated 

with gold or carbon to increase conductivity.

There have been some SEM methods developed for fiber counting [Asbestos International 

Association 1984; WHO 1985; ASTM 1996; ISO 2002]. These methods are primarily used for 

inorganic man-made fibers that have larger diameter fibers than can occur with asbestos. 

Thus, all the fibers are potentially visible using the SEM.

7 Transmission electron microscopy (TEM)

The transmission electron microscope (TEM) allows detection of particle shape and structure 

down to the smallest asbestos fibers (Figure 2) and can be used to determine crystal structure 

from electron diffraction as well as determining elemental composition from energy

dispersive x-ray analysis. Although TEM analysis is potentially very powerful and accurate, 

the process of sample collection and preparation and details involved in sample analysis can 

degrade the quantitative accuracy of the technique. Several more specialized techniques, such 

as electron energy loss spectroscopy and secondary ion mass spectrometry, have been used for 

analyzing particles and can also be applied to fibers [Fletcher et al. 2001].

Airborne fiber samples for TEM analysis are typically collected onto a filter, usually a 

polycarbonate membrane or MCE membrane filter. For the latter filter type, the filter is 

chemically collapsed to form a smooth upper surface on which collected fibers are trapped. 

Sometimes the surface is etched using a low temperature asher to expose the fibers collected 

on or near the surface of the original filter. The filter is coated with a carbon film that entraps 

fibers exposed on the filter surface and the filter material is then dissolved away. The carbon 

film is transferred to a TEM grid (usually 3-mm diameter) and the sample can be placed in the 

TEM for analysis.
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Measurement of Fibers

For Method 7402, the surface is not ashed because some fibers, e.g., cellulose, may be removed 

and give an inaccurate total fiber count [Baron and Platek 1990]. Ashing can thus affect the 

measurement of the asbestos fiber fraction.

The above approach to preparing MCE filters for TEM analysis is called the direct-transfer 

approach, since fibers are transferred to the carbon film with minimum disturbance to the 

way they were collected. An alternative technique is to dissolve the entire filter in liquid, 

ultrasonicate the suspension to disperse the particles, and deposit an aliquot of the particle 

suspension onto a polycarbonate filter for final transfer to the carbon film. This is called the 

indirect-transfer technique. With the indirect technique, the optimum particle loading of the 

TEM sample can be obtained and soluble particles can be removed from the sample. However, 

the suspension process can change the apparent size distribution of the particles and fibers by 

breaking apart agglomerates or even breaking apart asbestos fibers into smaller fibers or fibrils 

[Sahle and Laszlo 1996]. The breakup problem can be especially severe for chrysotile, causing 

a large increase in fiber count. Quality assurance is especially important with TEM analysis of 

fibers. The NVLAP program provides quality assurance accreditation for laboratories 

performing TEM analysis using the Environmental Protection Agency’s Asbestos Hazard 

Emergency Response Act (AHERA) method. Note that data provided under the AHERA 

method, because of significant differences in counting rules, the types of structures counted as 

asbestos, and the size range of fibers, cannot be directly compared with counts by Methods 

7400 or 7402.

The process of sample collection and preparation is a complex one that can introduce biases 

into the final measurement. Since only small portions of the filter are measured during TEM 

analysis, sampled fibers that deposit non-uniformly onto the filter due to inertial, 

gravitational, and electrostatic effects will be measured inaccurately [Chen and Baron 1996]. 

Fibers that penetrate the filter surface and are not transferred to the carbon film will be lost. If 

the filter is incompletely dissolved away from the carbon film, the sample will be difficult to 

analyze.

Many of the sources of bias and variability noted in sampling and counting by PCM also apply 

to TEM analysis. Fiber counting in a TEM can also introduce biases and variability in the final 

result. There is a tendency to use the high magnification of the TEM to look for the smallest 

fibers, while ignoring some of the larger ones. Even so, fibers shorter than 0.5 μm tend to be 

missed because they are difficult to see in the background clutter of the sample [Steel and 

Small 1985]. Taylor et al. found that TEM counting gave poorer precision than counting the 

same sample by PCM and recommended that the fraction of asbestos fibers counted by TEM 

be applied to the PCM count as indicated in Method 7402 [Taylor et al. 1984]. This combined 

PCM/TEM approach gave better precision than counting by TEM alone.
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Measurement of Fibers

In addition to recognizing fibrous shape and structure of the several asbestos minerals, 

qualitative analysis of fibers by TEM primarily involves two techniques, energy dispersive xray analysis and electron diffraction. X-ray analysis produces responses for each of the 

elements (typically atomic number > 6, but is instrument dependent) present in a particle; the 

responses occur as peaks in an energy spectrum. Specific asbestos minerals can be identified 

using peak intensity ratios observed in standard samples and as specified in the method.

The crystal structure of individual fibers is evaluated using electron diffraction. Focusing the 

TEM electron beam on a single fiber produces a diffraction pattern consisting of a number of 

spots. The spot locations depend not only on the particle crystal structure, but also on the 

geometry of the electron beam optics and other instrumental parameters. The diffraction spot 

locations relative to one another give a very specific identification of crystal structure. For 

easily recognized minerals, such as chrysotile, the visual identification of the diffraction 

pattern is often sufficient. However, to identify fibers not fitting the x-ray analysis pattern for 

standard asbestos minerals, careful measurement, or indexing, of the diffraction spots is 

important.

The combination of x-ray analysis and electron diffraction gives a highly definitive 

identification of specific minerals. However, as with any analytical methods, there are 

exceptions that require greater expertise to recognize potential interferences. Some minerals 

that are difficult to differentiate from regulated asbestos minerals include non-regulated 

amphiboles and fibrous talcs. There are several established methods for analyzing fibers, 

especially asbestos fibers, by TEM [Asbestos-containing materials, 1987; NIOSH 1994b; ISO 

1995, 1999; ASTM 1998].

8 Optical detection (light scattering)

Two types of light scattering detectors are commonly used for measuring airborne dust 

concentrations: the optical particle counter (OPC), which detects and counts individual 

particles, and the photometer (sometimes called a nephelometer), which detects the scattering 

from all particles in a defined detection volume. A standard OPC was used to detect asbestos 

concentrations in a workplace where the aerosol was primarily fibrous and good correlation 

with fiber counts was obtained [Rickards 1978]. A nephelometer may also be used, but may 

have an even greater interference from non-fibrous dusts.

The fibrous aerosol monitor (Model FM-7400, MIE, Inc. Bedford MA) used an electrostatic 

alignment technique by applying a field that aligns and rotates individual fibers in a laser 

beam. The light scattered from the fibers uniquely identified the presence of individual fibers. 

This allowed specific detection of fibers [Lilienfeld et al. 1979] and was even used to measure 

fiber length [Marijnissen et al. 1996].
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Measurement of Fibers

 

Several field tests have indicated that the fibrous aerosol monitor agrees reasonably well with 

field measurements of fibers by phase contrast microscopy, though mostly at concentrations 

above ambient levels. It has been used at abatement sites to provide rapid feedback and ensure 

acceptable containment of airborne fibers during asbestos removal.

9 Fiber classification

Several devices have been used to measure or separate fibers by diameter. A spiral centrifuge 

was used to separate fibers and reference spherical particles to estimate fiber aerodynamic 

diameter [Stöber 1972]. It was found that the aerodynamic diameter was directly proportional 

to physical diameter, proportional to the square root of the fiber density, and proportional to 

fiber length to the 1/6th power. For mineral fibers having a density of about 3 g/cm3

, the 

aerodynamic diameter was approximately three to five times the physical diameter of the 

fiber. Behavior of glass fibers in a cascade impactor was investigated by Burke and Esmen 

[Burke and Esmen 1978]. A small correction to the aerodynamic diameter was developed to 

take into account interception of longer fibers with the impaction surface. An inertial 

spectrometer was used to measure fiber aerodynamic diameter and good diameter separation 

was achieved [Morigi et al. 1999]. Baron and Deye developed a technique for separating fibers 

by length using dielectrophoresis [Baron et al. 1994; Deye et al. 1999]. This technique was also 

shown to be useful for measuring fiber length and diameter distributions [Baron et al. 2000].

As with most airborne dusts, fiber settling will reduce the number of larger diameter fibers in 

a distribution as the distance from the source of the dust increases. Esmen et al. showed that 

average fiber concentration in workplaces decreased exponentially with an increase of fiber 

diameter, indicating that the larger diameter fibers settled out more quickly than smaller 

diameter fibers [Esmen et al. 1979]. Cyclones, impactors and porous foam classifiers were 

evaluated for efficiency of removing airborne fibers not likely to deposit in the lungs 

[Maynard 1996].

The aerodynamic diameter of fibers is dependent primarily on fiber physical diameter and 

fiber density, with a minor dependence on fiber length [Baron 1996]. The diseases caused by 

asbestos fibers are lung diseases and so it makes sense to measure only fibers that can enter the 

lungs, i.e., thoracic fibers. Identical conventions for thoracic samplers have been published by 

ISO, ACGIH [ACGIH 2002], and CEN. Baron [Baron 1996] showed that sampling fibers with 

a thoracic sampler was approximately equivalent to counting only mineral fibers with a 

physical diameter smaller than 3 μm. Jones et al. [Jones et al. 2001] reported that there 

appeared to be no impediment to using a thoracic sampler for fiber sampling; they found that 

several samplers matched the thoracic convention, the sample collected by these samplers 

could be analyzed by standard methods, and that field studies indicated equivalence to the 
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Measurement of Fibers

current method. Maynard [Maynard 1999] also found that there appeared to be no variation 

in penetration through these samplers as a function of fiber length. The advantage to using a 

thoracic sampler, apart from adhering to conventional sampling practice, is that it would 

remove larger compact particles and fibers from the sample and result in a cleaner sample. 

Although current US practice does not use an upper diameter limit for asbestos fibers, such a 

limit is commonly used for man-made fibers. Except for the United States, all national and 

international organization methods use an upper diameter limit of 3 μm for fiber counting of 

asbestos fibers.

 

It is likely that thoracic sampling will eventually be in routine use for measurement of asbestos 

and other fibers. This approach has several advantages. It places the fiber method in line with 

other dust sampling conventions. It removes some of the larger particles in the sample, 

resulting in a cleaner sample for the analyst. It removes the need for determining fiber 

diameter during counting and it is consistent with previous practice of using an upper 

diameter limit of 3 μm for fiber counting in some methods. Thoracic sampling has the 

disadvantage of requiring the flow rate for a specific sampler to be fixed. This reduces the 

flexibility to target the loading of the filter by adjusting the flow rate. However, several 

classifiers can be designed to operate at selected flow rates to allow some flexibility in 

sampling.

10 Conclusions 

The capability for measurement of fiber size distributions is available through microscopy 

and, to a much lesser extent, through direct-reading instrumentation. Because of differences 

in counting rules, resolution capability, and ability to distinguish asbestos from interfering 

particles or other fibers, PCM, PLM, SEM, and TEM methods often do not produce results 

which are directly comparable. The traditional methods of microscopy are relatively 

inaccurate when compared to chemical analysis methods for most other analytes because of 

the many sources of error in the sampling and analysis procedure. To improve laboratory-tolaboratory agreement, counter training and quality control, including the exchange of samples 

among laboratories and proficiency testing, are important. Implementation of training 

through the use of Pang’s coverslips allows investigation of counting errors and potential 

improvement of PCM counting accuracy. Thoracic sampling could eliminate interfering 

particles and thereby improve measurement methods in the future.
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Measurement of Fibers

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

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NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Sampling and Analysis of Soluble 

Metal Compounds

by Kevin Ashley, Ph.D., NIOSH and Richard Fairfax, CIH, OSHA

[Much of this chapter was adapted from Fairfax and Blotzer 1994; and Ashley 2001.]

1 Introduction SM-2

2 Soluble and insoluble metal compounds SM-2

3 Health effects SM-4

4 Sampling considerations SM-5

5 Analytical considerations SM-5

6 ISO procedure for soluble metals and metalloids SM-7

7 Summary SM-10

8 References SM-11

9 Appendix - Solubilities of selected metals and metal compounds SM-14
NIOSH Manual of Analytical Methods  5th Edition  Chapter SM April 2016 Page SM-2 of SM-17

Sampling and Analysis of Soluble Metal Compounds

1 Introduction

Workplace exposure standards have been established for several soluble metals and metalloids 

to take into account the increased bioavailability of some metal compounds. Exposure 

standards for soluble compounds can be up to 500 times lower than the exposure standards 

for less soluble compounds for the same metal. However, there is often confusion among 

chemists, industrial hygienists, and laboratories over what is meant by “soluble” when the 

metal species, extraction fluid, or solubility conditions are not specified in the exposure 

standard nor in the supporting exposure standard documentation [Fairfax and Blotzer 1994]. 

In addition, the metals and metal compounds may interact chemically or physically with the 

sampling media or with each other [Ashley 2001]. Such complications can affect the stability 

and speciation of the metals and their compounds, and must be addressed in order to obtain 

meaningful results. These issues are becoming more important in workplace airborne metals 

exposure monitoring.

The solubility of a metal will depend on the chemical form of the metal, the fluid used to 

extract the metal, and the conditions under which the extraction occurs (e.g., temperature, 

volume, time). Unfortunately, the degree of method specificity needed to obtain 

measurements that are reproducible among laboratories is generally either missing or is 

subject to a variety of interpretations from exposure standards and supporting 

documentation. The need for a better definition of what is meant by the term “soluble” in 

relation to exposure standards was first raised in the 1990s, but as of the new millennium no 

significant improvement had occurred within exposure standard-setting organizations in the 

United States. Therefore, to meet the needs of analysts, laboratories, and laboratory clients for 

better definition of the analyte of interest, and to improve measurement reproducibility 

among laboratories, various organizations are working to achieve international consensus on 

extraction of soluble metal compounds. Consensus guidelines have been promulgated in an 

International Standard [ISO 2012a], and this will serve to fill the void and improve the 

situation. 

2 Soluble and insoluble metal compounds

The American Conference of Governmental Industrial Hygienists and several countries (e.g., 

France, Germany, UK, USA) have established occupational exposure limits (OELs) for soluble 

metal and metalloid compounds [ACGIH 2015; IFA 2014]. Some examples of elements for 

which soluble OELs have been promulgated are listed in Table 1 (see Appendix for additional 

details). For many of these elements (e.g., Tl, Ag, Cr[VI], Ni, Pt), the OELs for the soluble 

compounds are lower than for the corresponding insoluble forms [ACGIH 2001; CRC 2015]. 
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Sampling and Analysis of Soluble Metal Compounds

Table 1. Example metallic elements for which soluble compounds have been assigned 

OELs [IFA 2014; ACGIH 2015]

Aluminum – Soluble compounds, as Al

Barium – Soluble compounds, as Ba

Chromium – Water-soluble Cr[VI] compounds 

Iron – Soluble salts, as Fe

Molybdenum – Soluble compounds, as Mo

Nickel – Soluble compounds, as Ni

Platinum – Soluble salts, as Pt

Rhodium – Soluble compounds, as Rh

Silver – Soluble compounds, as Ag

Thallium – Soluble compounds, as Tl

Tungsten – Soluble compounds, as W

Uranium – Soluble compounds, as U

a. Solvent 

The term “solubility,” as used by analytical chemists, ordinarily pertains to the dissolution 

of a material in pure water [CRC 2015]. The subject of water solubility of metal 

compounds is covered in several references [CRC 2015; ACGIH 2001; Beliles 1994; 

O’Neil 2006]. According to Patty’s Industrial Hygiene and Toxicology [Beliles 1994] and 

the individual ACGIH Threshold Limit Value® (TLV) documentation for these metals 

[ACGIH 2001], the solubilities of metals and metal compounds are quite variable 

depending upon the solvent. Other pertinent references sustain the notion that solubility, 

regarding metals and their compounds, is generally identified in terms of their solubility in 

water [ACGIH 2001; O’Neil 2006]. The solubility and insolubility of numerous inorganic 

substances are presented in the Appendix. What is meant by “soluble” depends on the 

operational definition employed for the extraction conditions desired by the investigator. 

b. Temperature 

Temperature is another variable that directly affects solubility. Most current analytical 

methods specify deionized water, but not water temperature (some procedures call for hot 

water (37 °C), but others use water at room temperature). Some important questions thus 

arise: 

1.) If using deionized water, should chemists assure that water temperature has been 

heated to body temperature (i.e., 37 °C)? For occupational exposure assessment 

purposes, should solubility be based upon body temperature?
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Sampling and Analysis of Soluble Metal Compounds

2.) Should the OEL value documentation specify that analytical procedures use extraction 

media heated to body temperature?

3 Health effects

From a health perspective, the solubility of a metal or metal compound is not the only 

consideration of interest. Ultimately, the most important consideration is the extent to which 

such soluble metals accumulate in body fluids or target organs, leading to toxic levels of the 

metal ion. This is of more concern than solubility in water, acids, or alkalis per se. Further 

complicating the solubility issue is the fact that the term “soluble” may have different 

meanings among industrial hygienists and chemists. Chemists generally use the term 

“soluble” as defined by the CRC Handbook of Chemistry and Physics [CRC 2015]. According 

to the CRC Handbook, a material is considered soluble if a saturated solution in water (at 

25 °

C) contains more than 1% (m/v); any material in which 1 percent or less is dissolved is 

considered insoluble. Unfortunately, a material listed as insoluble, using the CRC definition, 

could still dissolve in body fluids and produce a significant tissue concentration which is 

biologically detrimental. The point is that an OEL for an “insoluble” compound may not be 

sufficient to protect exposed workers. When asked in an informal poll to choose a solvent in 

which to measure the relative solubility of metals, industrial hygienists chose, in order of 

preference, water, body fluids, and a petroleum solvent [Fairfax and Blotzer 1994]. 

a. Body fluids

When considering the biological effect of the solubility of a material, we should ideally 

first consider body fluids. However, body fluids vary considerably in pH. For example, the 

pH of the stomach is acidic, the pH in the intestine is alkaline, the pH of blood serum is 

approximately neutral, some macrophages are highly acidic, and the pH of saliva is slightly 

acidic. Furthermore, body fluids contain a variety of solutes, including salts and 

polypeptides (proteins). Polypeptide molecules can bind to metal ions in solution and 

often contain functional groups that can chelate metals. Polypeptides have strong 

chelating ability in body fluids and will account for the considerable difference between 

the solubility of a metal in body fluids versus that same metal in water. Metals, in turn, are 

bound to different proteins, depending upon where in the body they are located at a given 

time. In passing through the body, a metal ion is bound by different polypeptides. For 

each of them, a different reaction may be involved. Some reactions may increase or 

decrease the toxicity of the metal ion. 

Because of the effect of proteins, pH, and other solutes in body fluids, the solubility of a 

metal compound in body fluids will be quite different than the metal’s solubility in water 

[ACGIH 1987]. 
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Sampling and Analysis of Soluble Metal Compounds

4 Sampling considerations

a. Filter reactivity 

The filter medium used must not react with the airborne particulate collected by the 

sampler so as to change the chemical form of the captured sample. This can occur if a 

soluble compound reacts with the filter material or a contaminant therein to produce an 

insoluble or less soluble compound. An example of this problem has been observed with 

silver, where a soluble silver compound, AgNO3, can react with chloride in some mixed 

cellulose ester (MCE) or polyvinyl chloride (PVC) filters to form AgCl, which is much less 

soluble in water. Thus, low recoveries of “soluble” silver will result unless an alternative 

filter medium, such as polytetrafluoroethylene (PTFE), is used. Another problem is 

illustrated by soluble forms of Cr[VI], which can react with the filter medium to form 

insoluble compounds and/or undergo reduction to Cr[III] and, therefore, be lost to Cr[VI] 

analysis. Hence, chemical compatibility issues must be investigated before samples meant 

for “soluble” extraction procedures are taken.

Membrane filters are appropriate for sampling aerosols for subsequent determination of 

soluble metal compounds. Such filters are manufactured from a variety of polymeric 

materials by a number of different processes. Choice of polymer material comprising the 

filter (e.g., MCE, PTFE, PVC) will depend on chemical reactivity issues discussed in the 

preceding paragraph. The metal content of the filters must be as low as possible, since it 

can make a significant contribution to the blank value. 

5 Analytical considerations

A number of analytical methods for soluble metal compounds in occupational hygiene 

samples have been published by various organizations [NIOSH 1994; HSE 1998; BIA 1989; 

INRS 2014]. Efforts to harmonize sample preparation approaches have led to the 

promulgation of related consensus standards [Ashley 2015]. 

a. European standard

Guidance on sample preparation methods for soluble metals and metalloids in workplace 

air has been promulgated recently in a European Standard [CEN 2009]. In these 

published methods and guidelines [NIOSH 1994; HSE 1998; BIA 1989; INRS 2014], two 

methodologies are generally favored for the extraction of “soluble” metal species: 

(1) extraction in pure water, or (2) extraction in diluted (~0.1 M) hydrochloric acid (HCl). 

In the European Standard [CEN 2009], both strategies are given as options for the 

dissolution of “soluble” metals in workplace air samples. Extraction of metals and 

metalloids in water is meant to reflect the chemical definition of “soluble” (as mentioned 

above), while extraction in diluted HCl is designed to mimic the dissolution of “soluble” 
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Sampling and Analysis of Soluble Metal Compounds

metal compounds in stomach acid, which is highly acidic (pH 1). A temperature of 37 °C 

is recommended since this is normal body temperature.

b. Extraction solvent 

A researcher in Germany [Hahn 2000] argued for the establishment of 0.1 M HCl in an 

operational definition for soluble metals in occupational hygiene samples and, also, 

specified mechanical agitation at 37 °C for two hours prior to sample analysis. An 

exception is made for thallium, which can form insoluble TlCl, in which case HCl is 

replaced by diluted nitric acid (HNO3). (A similar problem would arise if HCl were used 

as the extraction acid for soluble silver compounds.) The German extraction method for 

several soluble metal species [BIA 1989] is consistent with the strategy outlined [Hahn 

2000], which attempts to address bioavailability by choosing HCl as the extraction acid. 

However, in standard methods promulgated by the United States [NIOSH 1994], the 

United Kingdom [HSE 1998], and France [INRS 2014], deionized water is the solvent 

chosen in the operational definition of “soluble” for numerous metallic elements in 

workplace air samples. An exception to extraction in water is made for nickel [HSE 1998], 

where an ammonium citrate solution is specified as the leachate for soluble compounds of 

this element. Ammonium citrate provides buffering and chelating properties that are 

desirable for leaching soluble nickel compounds [HSE 1998]. 

c. Operational definitions

Operational definitions of “soluble” metal species have been promulgated for consumer 

products such as toys, paper products, paints and art materials [ASTM 2014; CEN 2013; 

ISO 2011]. Standard procedures for the extraction of metal compounds from consumer 

products are based on sample treatments in 0.07 - 0.14 M HCl (depending on the sample) 

for an hour at a temperature of 37 °C [Hahn 2000] or at room temperature [CEN 2013]. 

An ASTM International procedure (formerly the American Society for Testing and 

Materials) [ASTM 2014] is meant to provide an estimate of the bioavailability of several 

metals in art materials, using ~0.1 M HCl and extraction at body temperature.

d. Quantity of solvent

Another uncontrolled variable is the quantity of solvent used in laboratory analytical 

procedures for soluble metals at different laboratories. Different laboratories may (and 

do) use different amounts of deionized water for extraction. For example, one laboratory 

might use 10 mL of deionized water to extract the metal from a sample, while another lab 

may use anywhere from 25 to 100 mL to extract the compound. Depending upon the 

amount of material present in the sample, the procedure using 25 to 100 mL can dissolve a 

larger mass of solute than that using 10 mL. A conservative analytical method for metals 

used 15 mL of deionized water for extraction. This volume was chosen as a convenience 
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Sampling and Analysis of Soluble Metal Compounds

and may not bear any relationship to what the body might absorb. Thus, two identical 

samples could produce two different results depending on the volume of solvent and other 

analytical parameters. A further factor affecting solubility is the particle size distribution 

of the sample: smaller particles are ordinarily more easily dissolved than larger ones. Thus 

for two samples having the same mass, the sample comprised of small particles may be 

more easily dissolved than a sample having large particles.

6 ISO procedure for soluble metals and metalloids

Various procedures for the extraction of soluble metals and metalloids have been used for 

years, based on different operational definitions of solubility. The International Organization 

for Standardization, Technical Committee 146 on Air Quality, Subcommittee 2 on Workplace 

Atmospheres, Working Group 2 on Inorganic Particulate Matter (ISO/TC 146/SC 2/WG 2) 

has attempted to standardize extraction procedures for “soluble” metal compounds by 

offering an operational definition in terms of a sample preparation method for metallic 

elements in industrial hygiene measurements.

a. Bioavailability

In the 1990s, it was argued that the solubility in body fluids should be considered in the 

development of a new definition for soluble TLVs [Fairfax and Blotzer 1994]. But since 

different body fluids have different solubility characteristics (e.g., pH, salts, polypeptides), 

such an operational and uniform definition for “bioavailable” cannot realistically be 

decided. Indeed, the meaning of “bioavailability” has been debated nationally and 

internationally for years, and it was not deemed practicable nor defensible to attempt to 

operationally define solubility based on biochemical arguments. Hence, it was decided by 

consensus within the ISO working group (ISO/TC 146/SC 2/WG 2) to describe procedures 

for soluble metal compounds in terms of strictly chemical, and not biochemical, criteria 

[Ashley 2001]. 

b. Laboratory consistency

With regard to analytical methods for the extraction of soluble metals and their 

compounds, it was suggested that the extraction media, temperature, and extraction 

volume should be consistent among all laboratories [Fairfax and Blotzer 1994]. For an 

operational definition of “soluble” to be offered, delineation of these analytical parameters 

is necessary in order to fully standardize the extraction procedure for soluble metal 

species. Moreover, the apparatus used, as well as chemical compatibility issues, must be 

amply described. Matters that are outside of laboratory control, notably sampling, cannot 
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Sampling and Analysis of Soluble Metal Compounds

always be adequately influenced; however, recommendations as to sampling media, 

sample handling, and transport requirements should be provided to the field industrial 

hygienist.

c. Development of ISO procedure

The ISO working group responsible for the development of ISO 15202-2, ISO/TC 146/SC 

2/WG 2, began its task in September 1995, not long after the publication of the 

aforementioned article [Fairfax and Blotzer 1994]. Shown in Table 2 is a list of countries 

that participated in voting on the technical content of the draft international standard 

when it was circulated for balloting, from 1996 to 2000.

Because of the various operational definitions for “soluble” metal compounds, a significant 

challenge was presented to the ISO working group responsible for the development of an 

international standard method to describe a procedure for extracting soluble metals and 

metalloids for subsequent atomic spectrometric analysis. Two choices were available 

based on the standard methods mentioned above: (1) extraction in pure water, or (2) 

extraction in 0.1 M HCl. It was decided by consensus of ISO delegates present at the 

earlier working group meetings to follow the former course, where solubility of metal 

compounds in occupational hygiene samples is defined in chemical terms. This decision 

was upheld during the later international voting process, which involved those countries 

listed in Table 2. Nevertheless, text within the International Standard [ISO 2012a] states 

that individual countries may specify alternative procedures for the measurement of 

soluble metal species in workplace air samples. This, then, leaves open the option to use 

other extraction media, such as 0.1 M HCl.

Table 2. “Participating Member”* Countries of ISO/TC 146/SC 2 (During the period 

of development of ISO 15202-2 [1996-2000])

Belgium

Germany

India

Italy

Japan

Korea

Netherlands

Poland

Spain

Sweden

Turkey

United Kingdom

United States

* “Participating,” or P-Member, countries are those nations able to vote on Draft 

International Standards, and therefore may provide comments on the technical content of 

the documents during voting. “Observing,” or O-Member, nations (not listed) may also

offer comments and can participate in the development of ISO standards. However, in the 

formal ISO voting process, O-member nations can vote only on Final Draft International 

Standards, which allows only for editorial, and not technical, changes at this stage.
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Sampling and Analysis of Soluble Metal Compounds

The ISO procedure for soluble metals and metalloids assumes that samples were collected 

using the International Standard for the collection of workplace air samples for subsequent 

multi-element analysis, ISO 15202-1 [ISO 2012b]. However, the choice of filter material 

used for the collection of samples targeted for the “soluble” metals procedure is important. 

Annex A of ISO 15202-1 [ISO 2012b] and Annex B of ISO 15202-2 [ISO 2012a] provide 

useful guidance on this subject.

The “soluble” metals procedure described in ISO 15202-2 [ISO 2012a] lists all of the 

elements in Table 1 except for Cr[VI] and Fe. Nevertheless, the sample preparation 

protocol described in this International Standard is certainly applicable to these other two 

metallic elements as well. The ISO “soluble” metals method [ISO 2012b] calls for 

treatment of collected workplace filter samples in 5 mL of deionized water (or ammonium 

citrate leach solution in the case of Ni) and mechanical agitation in a water bath at 37 ± 

2 °C for 60 min. Undissolved material is thereafter separated from the sample solution 

using a suction filtration apparatus or a syringe filter, and ensuring use of filtration 

materials that are unreactive towards the soluble metal compounds of interest. After 

filtration, the sample solution is acidified with nitric acid in order to stabilize the dissolved 

metallic elements within the extracted sample. This test sample is then ready for analysis 

by ICP-AES (or ICP-mass spectrometry [ICP-MS] if very low detection limits are 

required). Of course other analytical techniques, e.g., atomic absorption spectrometry 

[Wang et al. 2000; Draper et al. 1999] or electrochemical analysis [Ashley 1994; Draper et 

al. 1999], can be used as analytically equivalent alternatives.

d. Method performance 

The performance of soluble extraction methods has been evaluated for several soluble 

metal species, e.g., those of nickel, silver and hexavalent chromium. Soluble extraction 

and atomic spectrometric analysis of soluble nickel in cellulosic air filter samples and in 

bulk reference samples, using 0.1 M ammonium citrate for extraction (as part of a 

sequential extraction method), was ruggedized and validated both within a single 

laboratory and via interlaboratory trials [Zatka et al. 1992]. This ammonium citrate 

leaching procedure forms the basis of the ISO 15202-2 (Annex B) methodology 

[ISO 2012a] for soluble nickel compounds. In other research, soluble silver compounds 

on PTFE filters were subjected to leaching in deionized water [ISO 2012a] within opaque 

sampling cassettes, with subsequent analysis by ICP-MS [Drake et al. 2006]. The use of 

opaque samplers was necessary to prevent photoreduction of silver ions in solution, and 

ICP-MS was required due to the need for lower method detection limits for the soluble 

silver fraction in real samples. The soluble silver procedure was validated in-house by 

using silver nitrate spikes on PTFE filters, demonstrating >90% recoveries. Field studies 

were also carried out in silver refineries by using a multiport sampler, where it was found 
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Sampling and Analysis of Soluble Metal Compounds

that the soluble silver fraction was less than 2% of the mass of total silver in collected air 

samples [Drake et al. 2006]. In further work, the use of deionized water for leaching of 

soluble hexavalent chromium compounds was evaluated [Ashley et al. 2009] as part of a 

study to validate a standardized sequential extraction procedure for Cr[VI] preceding ion 

chromatographic analysis [ASTM 2013]. Laboratory experiments on soluble Cr[VI] 

compounds spiked onto PVC filters resulted in quantitative recoveries, yet interference 

from Fe[II] (if present in samples) was unavoidable. Water leaching of paint pigment 

samples and welding fume samples was also evaluated as part of the investigation. It was 

found that soluble Cr[VI] compounds were prevalent in welding fumes, but their contents 

were variable in the different paint pigments that were tested [Ashley et al. 2009]. 

7 Summary

The exposure standards for some metals vary up to a factor of 500 to take into account the 

increased solubility and bioavailability of some compounds. Even compounds generally 

considered by chemists as being “insoluble” may have sufficient solubility in body fluids to be 

of biological importance. Exposure standards for soluble metals such as ACGIH TLVs and 

other OELs are not specific with regard to extraction fluid, fluid temperature, agitation and 

other factors affecting solubility. Since these factors significantly affect solubility, some 

standardization or adoption of an operational definition is necessary if there is to be 

reproducibility among laboratories conducting soluble metal analyses. In some countries 

(e.g., nations in the European Union), the national requirements provide this specificity. In 

other instances, including exposure monitoring standards in the United States, formal 

national guidelines are not available.

Therefore, adherence to international extraction guidelines or methods such as those 

described in ISO standards is necessary to produce measurements that are reproducible with 

other laboratories, and have utility to the laboratory client when exposure standards or 

national guidelines are vague. ISO 15202 has the advantage over other guidelines and 

standards in that it has had input from more than 13 participating countries (Table 2). Thus 

the use of ISO 15202 is encouraged until either exposure standards or national guidelines 

provide better specificity. 

Laboratories cannot recover soluble metal data if inappropriate sample media are used. 

Sample stability is a problem that must be addressed when sampling for silver (Ag), 

chromium (Cr) and other soluble metal compounds. Although the ISO procedure has not 

been validated for all soluble metal species, the standard recommends that method validation 

be carried out using representative soluble metal compounds for target elements. ISO 15202 

provides guidance on sample media selection for soluble metals that can be a useful guide for 

industrial hygienists and other laboratory clients.
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Sampling and Analysis of Soluble Metal Compounds

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

8 References

ACGIH [1987]. American Conference of Governmental Industrial Hygienists (ACGIH) 

Committee Activities and Reports – TLVs for Chemical Substances. Appl Ind Hyg 2:R4-R5.

ACGIH [2001]. Documentation of the Threshold Limit Values and Biological Exposure 

Indices, 7th ed. Cincinnati, OH, USA: American Conference of Governmental Industrial 

Hygienists (ACGIH).

ACGIH [2015]. Threshold Limit Values for Chemical Substances and Physical Agents and 

Biological Exposure Indices. Cincinnati, OH, USA: American Conference of Governmental 

Industrial Hygienists (ACGIH).

Ashley K [1994]. Electroanalytical Applications in Occupational and Environmental Health. 

Electroanalysis 6:805-820. 

Ashley K [2001]. International Standard Procedure for the Extraction of Metal Compounds 

Having Soluble Threshold Limit Values. Appl Occup Environ Hyg 16:850- 853. 

Ashley K [2015]. Harmonization of NIOSH Sampling and Analytical Methods with Related 

International Voluntary Consensus Standards. J Occup Environ Hyg 12:D107-D115. 

Ashley K, Applegate GT, Marcy AD, Drake PL, Pierce PA, Carabin N, Demange M [2009]. 

Evaluation of Sequential Extraction Procedures for Soluble and Insoluble Hexavalent 

Chromium Compounds in Workplace Air Samples. J Environ Monit 11:318-325. 

ASTM [2013]. ASTM D6832, Standard Test Method for the Determination of Hexavalent 

Chromium in Workplace Air by Ion Chromatography ad Spectrophotometric Measurement 

Using 1,5-Diphenylcarbazide. West Conshohocken, PA, USA: ASTM International; 

[www.astm.org]. Date accessed: April 6, 2016.
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ASTM [2014]. ASTM D5517, Standard Test Method for Determining Extractability of Metals 

from Art Materials. West Conshohocken, PA, USA: ASTM International.

Beliles RP [1994]. The Metals. In: Patty’s Industrial Hygiene and Toxicology, 4th ed., Vol. 2c. 

New York, NY, USA: Wiley, pp. 1879-2317. 

BIA [1989]. Kennzahl 6015, Methodik zur Bestimmung “löslicher Metallverbindungen.” 

In: BIA – Arbeitsmappe Messung von Gefahrstoffen, Vol. 15 (Lfg. IX/95). Sankt Augustin, 

Germany: Berufsgenossenschaftliches Institut für Arbeitssicherheit (BIA).

CEN [2009]. EN 13890, Workplace Atmospheres - Procedures for Measuring Metals and 

Metalloids in Airborne Particles – Requirements and Test Methods. Brussels, Belgium: 

Comité Européen de Normalisation (CEN).

CEN [2013]. EN 71-3, Safety of Toys. Brussels, Belgium: Comité Européen de Normalisation 

(CEN).

CRC [2015]. Handbook of Chemistry and Physics, 96th ed.; Lide, D. R., ed. CRC Press, Boca 

Raton, FL, USA.

Drake PL, Marcy AD, Ashley K [2006]. Evaluation of a Standardized Method for 

Determining Soluble Silver in Workplace Air Samples. J Environ Monit 8:134-139. 

Draper WM, Ashley K, Glowacki CR, Michael PR [1999]. Industrial Hygiene Chemistry– 

Keeping Pace with Rapid Change in the Workplace. Anal Chem 71:33R-60R.

Fairfax R, Blotzer M [1994]. TLVs – Soluble and Insoluble Metal Compounds. Appl Occup 

Environ Hyg 9:683-686. 

Hahn J-U [2000]. Aufarbeitungsverfahren zur analytischen Bestimmung “löslicher” 

Metallverbindungen – Ein pragmatischer Vorschlag (Preparation Method to Analyze 

“Soluble” Metal Compounds – A Pragmatic Suggestion). Gefahrstoffe Reinh Luft 60:241-243. 

HSE [1998]. MDHS Method 42/2, Nickel and Inorganic Compounds of Nickel; and Method 

46/2, Platinum Metal and Soluble Platinum Compounds. In: Methods for the Determination 

of Hazardous Substances. London, United Kingdom: Health and Safety Executive (HSE).
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IFA [2014]. GESTIS – Database on Hazardous Substances – International Limit Values for

Chemical Agents. Sankt Augustin, Germany: Institut für Arbeitsschutz der Deutschen 

Gesetzlichen Unfallversicherung (IFA); [https://www.dguv.de/ifa/index-2.jsp]. Date 

accessed: April 6, 2016. 

INRS [2014]. Fiche 003, Métaux – Métalloïdes. In: Métrologie des Polluants, Evaluation de 

l’Exposition Professionnelle, Méthodes de Prélèvement et d’Analyse de l’Air. Paris, France: 

Institut National de Recherche et de Sécurité (INRS); [www.inrs.fr]. Date accessed: April 6, 

2016. 

ISO [2011]. ISO 3856, Paints and Varnishes – Determination of “Soluble” Metal Content 

(7 Parts). Geneva, Switzerland: International Organization for Standardization (ISO); 

[www.iso.org]. Date accessed: April 6, 2016. 

ISO [2012a]. 15202-2, Workplace Air - Determination of Metals and Metalloids in Airborne 

Particulate Matter by Inductively Coupled Plasma Atomic Emission Spectrometry; Part 2, 

Sample Preparation – Annex B: Sample Dissolution Method for Soluble Metal and Metalloid 

Compounds. Geneva, Switzerland: International Organization for Standardization (ISO). 

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Airborne Particulate Matter by Inductively Coupled Plasma Atomic Emission Spectrometry; 

Part 1, Sampling. Geneva, Switzerland: International Organization for Standardization (ISO). 

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O’Connor, PF, eds. Cincinnati, OH: U.S. Department of Health and Human Services, 

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Health DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/]. 

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Sampling and Analysis of Soluble Metal Compounds

9 Appendix - Solubilities of selected metals and 

metal compounds [Fairfax and Blotzer 1994]

Aluminum and compounds

Aluminum metal reacts with dilute hydrochloric acid, sulfuric acid, potassium hydroxide, and

sodium hydroxide.

The salts of aluminum, including sodium aluminate, aluminum fluoride, aluminum chloride, 

and cryolite, are all soluble in water.

Arsenic and compounds

Arsenic — Insoluble in hot and cold water; soluble in nitric acid.

Arsenic trioxide and pentoxide — Soluble in cold and hot water, alcohol, alkalies, and 

hydrochloric acid; arsenic pentoxide is soluble in acids.

Lead arsenate — Insoluble in cold water; soluble in hot water, nitric acid, and caustic alkalies.

Calcium arsenate(s) — Insoluble in water and acids.

Sodium arsenate(s) — Very soluble in water.

Arsenic acid — Soluble in water and alcohol.

Arsenic trisulfide — Practically insoluble in water.

Barium and compounds

Barium metal — Insoluble in water; soluble in alcohol.

Most of the compounds of barium are soluble in (cold or hot) water, for example, barium 

chloride, barium oxide, barium acetate, and barium cyanide. Barium hydroxide is slightly 

soluble in water. Barium carbonate is insoluble to slightly soluble in water, and is soluble in 

acids.

Beryllium and compounds

Beryllium — Slightly soluble in hot water; insoluble in cold water; soluble in dilute alkalies 

and acids.

Beryllium oxide — Insoluble in water; soluble in some acids and alkalies.

Beryllium hydroxide — Insoluble in water; soluble in acids and alkalies.

Beryllium fluoride — Soluble in cold and hot water, alcohol, and sulfuric acid.

Beryllium sulfate — Soluble in water and concentrated sulfuric acid.

Chromium and compounds

Chromium reacts with dilute hydrochloric acid and sulfuric acid, but not with nitric acid.

Chromium metal — Insoluble in hot and cold water.
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Hexavalent chromium compounds, including chromium trioxide, the anhydride of chromic 

acid, chromates, dichromates, and polychromates, tend to be of low solubility in water and 

can be subdivided into two subgroups: 

1.) Water-soluble hexavalent chromium compounds include chromic acid, its anhydride, and 

the monochromates and dichromates of sodium, potassium, ammonium, lithium, cesium, 

and rubidium.

2.) Water-insoluble hexavalent chromium compounds include zinc chromate, lead chromate, 

barium chromate, and sintered chromium trioxide. NOTE: Depending on the reference 

[CRC 2015, O’Neil 2006] both calcium chromate and strontium chromate are listed as 

soluble and insoluble in water. 

Iron and compounds

Iron, ferrous oxide, ferric oxide, and iron oxide — Insoluble in hot and cold water.

Ferric chloride, ferric nitrate, ferric sulfate, ferrous sulfate and ferrous chloride — Soluble in 

hot and cold water.

Ferric chloride — Soluble in ethanol, methanol, and ether.

Ferric nitrate and ferrous chloride — Soluble in ethanol and acetone.

Ferric sulfate — Sparingly soluble in ethanol; insoluble in acetone.

Ferrous sulfate — Insoluble in ethanol.

Molybdenum and compounds

Molybdenum — Insoluble in hot or cold water; soluble in nitric, sulfuric, and hydrochloric 

acids.

Molybdic oxide — Sparingly soluble in water; soluble in acids and alkalies.

Molybdenum disulfide — Insoluble in hot or cold water and dilute acids; soluble in hot 

sulfuric acid, aqua regia, and nitric acid.

Ammonium molybdate — Soluble in hot or cold water, acids, and alkalies.

Calcium molybdate — Insoluble in cold water; soluble in hot water.

Lead molybdate — Insoluble in water and alcohol; soluble in acid and potassium hydroxide.

Sodium molybdate — Soluble in hot and cold water.

Nickel and compounds

Nickel — Insoluble in hot and cold water; soluble in nitric, sulfuric, and hydrochloric acids.

Nickel oxide — Insoluble in hot and cold water; soluble in ammonium hydroxide and acids.

Nickel acetate — Soluble in cold water; insoluble in alcohol [CRC 2015]; soluble in alcohol 

[O’Neil 2006]. 

Nickel carbonate — Soluble in cold water; insoluble in hot water.
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Sampling and Analysis of Soluble Metal Compounds

Nickel hydrates — Some forms soluble, others insoluble (in water).

Nickel sulfate — Soluble in hot and cold water.

Platinum and compounds

Platinum — Insoluble in hot or cold water and single mineral acids; soluble in aqua regia; 

attacked by halogens, alkali cyanides, and caustic alkalies.

Platinum forms are series of complex chloroplatinate salts that are water soluble.

Rhodium and compounds

Rhodium — Insoluble in hot or cold water; soluble in hot sulfuric acid plus hydrochloric acid; 

slightly soluble in acids and aqua regia.

Water-soluble rhodium compounds include rhodium trichloride, sodium chlororhodite, and 

rhodium carbonyl acetylacetonate.

Silver and compounds

Silver — Insoluble in water and inert to most acids; reacts readily in dilute nitric acid or hot 

concentrated sulfuric acid; soluble in fused alkali hydroxides in the presence of air.

Silver oxide — Soluble in hot and cold water, acids, and alkalies.

Silver acetate — Soluble in hot or cold water and nitric acid.

Silver bromide — Insoluble in hot or cold water and nitric acid.

Silver chloride — Soluble in hot water; slightly soluble in cold water and ammonium 

hydroxide.

Silver cyanide — Soluble in cold water, nitric acid, and ammonium hydroxide.

Silver nitrate — Soluble in hot and cold water.

Thallium and compounds

Thallium — Insoluble in hot and cold water; soluble in nitric, sulfuric, and hydrochloric acids.

Thallous oxide — Soluble in water, acids, and alcohols.

Thallic oxide — Insoluble in hot or cold water; soluble in acids.

Thallous acetate and thallic chloride — Soluble in cold water and alcohol.

Thallous bromide and chloride — Slightly soluble in water.

Thallous sulfate — Soluble in hot or cold water.

Thallous sulfide — Soluble in cold water.

Tungsten and compounds

Tungsten — Insoluble in hot or cold water, hydrofluoric acid, and potassium hydroxide; 

soluble in mixtures of hydrofluoric and nitric acid; slightly soluble in sulfuric acid.

Tungsten trioxide — Insoluble in hot or cold water and acids; soluble in hot alkalies and 

hydrofluoric acid.
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Sampling and Analysis of Soluble Metal Compounds

Tungstic acid — Insoluble in cold water and most acids; soluble in hot water, alkalies, 

hydrofluoric acid, and ammonia.

Sodium tungstate — Soluble in hot and cold water; slightly soluble in ammonia; insoluble in 

acids and alcohol.

Tungsten carbide — Insoluble in water; soluble in mixtures of hydrofluoric and nitric acid, 

and in aqua regia.

Tungsten diboride — Insoluble in hot or cold water; soluble in aqua regia.

Tungsten hexachloride — Soluble in hot water.

Tungsten oxytetrachloride — Soluble in hot or cold water.

Tungsten hexafluoride — Soluble in hot or cold water and alkalies.

Tungsten disulfide — Insoluble in cold water and in alcohol; soluble in mixtures of 

hydrofluoric and nitric acid.

Phosphotungstic acid — Soluble in cold water, alcohol, and ether.

Ammonium paratungstate — Soluble in water; insoluble in alcohol.

Uranium and compounds

Uranium — Insoluble in hot or cold water, alcohol, and alkali.

Uranium dioxide — Insoluble in hot or cold water; soluble in nitric acid and concentrated 

sulfuric acid.

Triuranium octoxide — Insoluble in hot or cold water; soluble in nitric acid and sulfuric acid.

Uranium tetrafluoride — Insoluble in cold water, dilute acids, and alkalies; soluble in 

concentrated acids and alkalies.
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Monitoring Diesel Exhaust in the 

Workplace

by M. Eileen Birch, Ph.D., NIOSH

1 Introduction DL-2

2 Analytical method DL-6

3 Interlaboratory comparisons DL-23

4 Occupational exposure criteria (U.S.) DL-27

5 Summary DL-28

6 References DL-30
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Monitoring Diesel Exhaust in the Workplace

1 Introduction

a. Health effects 

Over a million U.S. workers (e.g., trucking, mining, railroad, construction, agriculture) are 

occupationally exposed to diesel exhaust [NIOSH 1988]. The widespread use of dieselpowered equipment is a recognized health concern. Exposure to diesel exhaust is 

associated with an increased risk of lung cancer [Attfield et al. 2012; Garshick et al. 2004; 

HEI 1995; IARC 2012; Silverman et al. 2012]. Diesel exhaust is pervasive, and 

environmental exposure is a public health concern; but workplace exposures pose higher 

risk because they are generally much higher than those encountered by the general 

population. 

In 1988, the National Institute for Occupational Safety and Health (NIOSH) reported 

diesel exhaust as a potential occupational carcinogen and recommended that employers 

reduce workers’ exposures [NIOSH 1998]. This recommendation was based on five 

independent animal studies, in which rats exposed to unfiltered exhaust showed an 

increased incidence of benign and malignant lung tumors [IARC 1989]. Other 

organizations, including the International Agency for Research on Cancer (IARC) [IARC 

1989], the World Health Organization (WHO) [WHO 1996], the California 

Environmental Protection Agency [CalEPA 1998], the U.S. Environmental Protection 

Agency (EPA) [EPA 2000a], and the National Toxicology Program [NTP 2000] reviewed 

the animal and human evidence, and each classified diesel exhaust as a probable human 

carcinogen or similar designation. In 2012, based on epidemiological studies, IARC 

[IARC, WHO 2012] reclassified diesel exhaust as carcinogenic to humans (Group 1). In 

particular, a major study of U.S. miners, conducted by NIOSH and the National Cancer 

Institute (NCI), found increased risk of death from lung cancer in exposed workers 

[Attfield et al. 2012; Silverman et al. 2012].

 

Noncancer health effects also are associated with diesel exhaust exposure, including 

immunologic, respiratory, and cardiovascular effects. Diesel exhaust particles can act as 

nonspecific airway irritants at relatively high exposures. At lower levels, they can trigger 

release of mediators (cytokines, chemokines, immunoglobulins, and oxidants) of allergic 

and inflammatory responses [Pandya et al. 2002]. Diesel particles may promote expression 

of the immunologic response phenotype (Th2) associated with asthma and allergic disease 

and may have greater immunologic effects in the presence of environmental allergens. 

Internationally, the prevalence of asthma (and related hospitalizations and mortality) 

continues to rise in adults and children. Children may be more vulnerable than adults 

[Edwards et al. 1994; Weiland et al. 1994; Wjst et al. 1993; van Vlient et al. 1997]. Studies 

indicate children living along major trucking thoroughfares are at increased risk for 

asthmatic and allergic symptoms. In the United States, the number of individuals with 
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Monitoring Diesel Exhaust in the Workplace

self-reported asthma increased by 75% from 1980 to 1994 [Mannino et al. 1998]. The 

immunologic evidence is consistent with results of epidemiologic studies that associate

traffic-related air pollution, especially diesel exhaust particles, with an increase in 

respiratory diseases.

Studies have consistently found positive associations between particulate air pollution and

daily mortality [Brown et al. 2000; Dockery et al. 1993; EPA 1999, 2000b; Pope et al. 1995a; 

Pope et al. 1995b; Pope et al. 2002; Samet 2000; Schwartz 1997; Schwartz et al. 1996]. The 

traditional U.S. air quality standard for particulate matter is based on particles having 

diameters ≤10 μm (PM10) [52 Fed. Reg. 24634 (1987)]. In 1997, EPA proposed a new 

standard [62 Fed. Reg. 38652 (1997)] (see www.epa.gov) based on particles having 

diameters ≤2.5 μm (PM2.5). These smaller particles originate mainly from combustion 

sources. The new standard was proposed because recent studies had found higher 

correlation between fine particle pollution and adverse health effects. In an analysis 

[Schwartz et al. 1996] of data from six U.S. cities, fine particles were consistently associated

with increased risk of death from chronic obstructive pulmonary disease, pneumonia, and

ischemic heart disease. Positive associations between fine particle pollution and hospital

admissions due to respiratory and cardiovascular illness also have been found [Schwartz

1994; Burnett et al. 1995; Schwartz and Morris 1995]. Particles produced by combustion

sources were implicated in these findings. In addition to asthma, chronic inhalation of

diesel exhaust particles may play a role in these adverse health outcomes. Reviews on the 

health effects of diesel exhaust have been published [CalEPA 1998; EPA 2000a; HEI 1995, 

2002; IARC 1989; IARC (WHO) 2012; NIOSH 1988; NTP 2000; Solomon et al. 1998; 

WHO 1996]. 

b. Composition

Diesel engine exhaust is a highly complex and variable mixture of gases, vapors, and fine

particles. The amount and composition of the exhaust vary greatly, depending on factors

such as fuel and engine type, maintenance schedule, tuning, workload, and exhaust gas 

treatment. The gaseous constituents have included hydrocarbons and oxides of carbon,

sulfur, and nitrogen. Particulate components consisted of liquid droplets and soot particles

bearing organic compounds, sulfates, metals, and other trace elements. The organic

fraction (droplets and particle adsorbed) was mainly unburned fuel and oil, but thousands 

of compounds (e.g., aldehydes, polycyclic aromatic hydrocarbons [PAH]) have been found

in the organic fraction—some of which are genotoxic [HEI 1995, 2002; IARC 1989; 

NIOSH 1988; WHO 1996].
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Monitoring Diesel Exhaust in the Workplace

c. Analyte choice: elemental carbon 

Because diesel exhaust is a highly complex mixture, a surrogate of exposure must be 

selected. In the early 1990’s, NIOSH researchers considering potential surrogates sought 

an overall measure of the particulate fraction because animal studies associated lung 

tumor induction with unfiltered diesel exhaust [IARC 1989; NIOSH 1988], and most (90% 

in one study) [Schuetzle 1983] of the exhaust’s mutagenic potency was associated with the 

particulate fraction. At the time, gravimetric methods for respirable combustible and 

submicrometer dusts were being used in mines, but gravimetric methods lack selectivity 

and are not suitable for low-level (e.g., <200 μg/m3) measurements. Methods for 

characterization of the soluble organic fraction of diesel particulate matter (DPM) also 

were available; others have since been developed. Although measurement of specific 

organic compounds, particularly genotoxins, may be relevant in characterizing the 

potential toxicity of diesel exhaust, a single compound or compound class would not 

reflect exposure to the particulate fraction—even if unique markers are found—because 

the composition of the exhaust is highly variable.

Carbon is a logical exposure surrogate for traditional diesel engines because DPM is 

predominantly (typically more than 80%) carbon [Japar et al. 1984; Lies 1989; Pierson et 

al. 1983]. However, carbon in the organic fraction (i.e., organic carbon, or OC) of DPM is 

not a selective measure because other sources of OC (e.g., cigarette smoke and other 

combustion aerosols, asphalt fumes) are present in many workplaces. Elemental carbon

(i.e., carbon in the soot particle core, or EC) is a better surrogate [Birch and Cary 1996a] to 

monitor because it is a more selective measure of particulate diesel exhaust and still 

constitutes a sizable fraction (30%–90%) (see [HEI 2002], Part I, Section 1) of the 

particulate mass. Fine EC particles are derived primarily from the combustion of fossil 

fuels, and diesel engines have been major sources of these particles. Carbonaceous aerosols 

such as cigarette and wood smokes contain little, if any, EC [Birch 1998a; Birch and Cary 

1996a]. Gasoline engines emit far less EC than diesels, so the contribution of this source is 

relatively small. Other sources such as coal combustion, incinerators, and tire debris can 

contribute to the background (environmental) levels of EC, but diesel engines were the 

primary emitters [Cass and Gray 1995; Sawyer and Johnson 1995]. In occupational

settings, where diesel equipment is used in relatively close proximity to workers, the 

contribution of these remote sources is negligible, especially when EC levels are well above 

background. Environmental EC concentrations are typically in the 1–3 μg/m3 range [Birch 

and Cary 1996a], depending on the local air pollution, while in workplaces with diesel 

equipment (operating), EC concentrations are generally much higher [e.g., Haney and 

Fields 1996; NIOSH 1992; NIOSH 1993; NIOSH 1994c; Stanevich et al. 1997; Verma et al. 

1999; Whittaker et al. 1999; Zaebst et al. 1991]. Higher environmental background (e.g., 3-

-5 μg/m3) has been reported for more polluted U.S. cities (e.g., Los Angeles), but the higher 
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Monitoring Diesel Exhaust in the Workplace

EC levels were attributed to nearby diesel vehicles [Cadle and Mulawa 1990; Gray et al. 

1984] (see also [Cass and Gray 1995]). At one monitoring site (Glendora, CA), 

examination of the data collected at 1-minute intervals revealed that emission plumes 

from diesel vehicles located 50 meters from the site contributed contamination up to 

5 μg/m3 above the background level [Hansen and Novakov 1990].

Carbonaceous dusts such as coal dust (EC content depends on coal rank), carbon blacks, 

and carbon nanomaterials contain EC, but particles in powders (dispersed as particle 

agglomerates) and mechanically generated dusts are much larger (generally >1 μm 

diameter) than combustion-based particles. Therefore, these dusts can be effectively 

excluded from the diesel sample on the basis of size. Only low levels of EC (≤15 μg/m3) 

were found in electric-powered (i.e., nondieselized) coal mines when impactors with 

submicrometer cutpoints were used for air sampling [Birch and Cary 1996b]. Guidance on 

air sampling is discussed in a following section. In addition to selectivity, potential health 

effects were considered when an EC surrogate was proposed [55 Fed. Reg. 110 (1990); 

Birch 1991; Birch and Cary 1996a]. Diesel particles and other types of insoluble fine 

particles are inhaled deeply into the lungs, where they can induce an inflammatory 

response. Further, EC particles were shown to increase the long-term retention [Sun et al. 

1982; Wolff et al. 1986] of adsorbed genotoxins and other chemical toxins because the 

particles have a high affinity for them [Niessner and Wilbring 1989]. The adsorbed 

organic fraction results from rapid cooling of the exhaust mixture, which causes 

enrichment of some species on the particle surface [Natusch 1978; Thrane et al. 1985; 

Yamasaki et al. 1982]. Enrichment by this mechanism is associated with compounds of 

moderate to low volatility [Thrane et al. 1985; Yamasaki 1982]. For example, PAHs having 

four or more aromatic rings are generally associated with particulate matter [Bjorseth and 

Becher 1986]; this is important because these higher-ring condensates are expected to be 

the most carcinogenic or mutagenic [Grimmer et al. 1983; Pott 1985]. In combination 

with an inflammatory response induced by the particles, genotoxic agents may promote 

tumorigenesis. Ultrafine particles (<0.10 μm) may pose an even greater health risk. Results 

of toxicological studies on solid particles having aerodynamic diameters in this size range 

indicate ultrafine particles are especially toxic, even those not having an organic fraction 

and consisting of materials considered relatively nontoxic (e.g., carbon black, titanium 

dioxide). In a study of rats [Donaldson et al. 2000], a 10-fold increase in inflammation was 

seen with exposures to ultrafine particles, relative to the same mass of fine particles. This is 

significant because, by mass, the majority of diesel particles are in the fine particle range, 

and most are in the ultrafine range by number [Kittelson et al. 2002]. Given the physical 

and chemical nature of EC particles emitted by diesel engines, monitoring and controlling 

exposures to these particles is prudent.
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Monitoring Diesel Exhaust in the Workplace

2 Analytical method

a. Background 

A monitoring method for DPM was published as Method 5040 in the NIOSH Manual of 

Analytical Methods (NMAM). The method is based on a thermal-optical analysis 

technique for particulate carbon. Both OC and EC are determined, but EC is a better 

exposure surrogate. Rationale for selection of an EC surrogate is summarized in the 

previous section (Analyte Choice: Elemental Carbon). Method updates and an NMAM 

Chapter (Q) have since been published [NIOSH 1994a (1998 supplement); NIOSH 1994b 

(2003 supplement); NIOSH 2003] to include interlaboratory data (e.g., round robin 

results) and other diesel-related information obtained since its initial publication (in 

1996). This 5th edition chapter is not a review of relevant literature published since the 4th 

edition. Its purpose is to update references and information on the following topics: 

classification of diesel exhaust as a human carcinogen, a study of miners exposed to diesel 

exhaust, results of a subsequent round robin by NIOSH investigators, and application of 

the Method to carbon nanomaterials. NIOSH 5040 has been used in numerous industrial 

hygiene surveys on diesel exhaust [e.g., Haney and Fields 1996; NIOSH 1992: NIOSH 

1993; NIOSH 1994c; Stanevich et al. 1997; Verma et al. 1999; Whittaker et al. 1999; Zaebst 

et al. 1991], and it was applied to an epidemiological study (NIOSH/NCI) of miners 

[Attfield et al. 2012; Silverman et al. 2012]. Details on Method operation and performance 

are provided in this chapter. Exposure criteria also are discussed.

b. Instrumentation 

Of the possible approaches for OC-EC analysis, a thermal-optical technique was 

investigated because it offered greater selectivity (pyrolysis correction for char) and 

flexibility (automated analysis, programmable parameter files) than previously used 

methods. Prior to its proposed use for monitoring occupational exposure to diesel exhaust, 

thermal-optical analysis (or OC-EC methods in general) had not been applied to 

occupational monitoring, but the technique had been routinely applied to environmental 

monitoring of particulate carbon air pollution.

The thermal-optical analyzer (Figure 1) has been described previously [Birch and Cary 

1996a; NIOSH 1994a (1998 supplement)]. Design changes (e.g., reflectance monitoring 

added, software upgraded) have since been made, but the operation principle remains 

unchanged. OC-EC quantification is accomplished through temperature and atmosphere 

control. In addition, the analyzer is equipped with an optical feature that corrects for char 

formed through sample pyrolysis (thermal breakdown in inert atmosphere). Some samples 

contain components (e.g., cigarette and wood smokes) that carbonize (convert to char 

carbon) when the sample is heated in helium during the first part of the analysis. Like the 
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Monitoring Diesel Exhaust in the Workplace

EC typical in fine particle pollution, char strongly absorbs light, particularly in the 

red/infrared region, resulting in a decrease in the filter transmittance/reflectance. Both 

volatile products and char are formed during the decomposition process, which may begin 

near 300 °C and continue until the maximum temperature (860-880 °C) is reached. 

Optical correction for char is made through use of a pulsed diode laser and photodetector 

that continuously monitor the filter transmittance/reflectance.

Figure 1. Schematic of thermal-optical transmittance instrument (V = valve) for 

determining OC and EC in carbonaceous aerosols.

In the thermal-optical analysis, a filter portion (punch) of known area (typically 1.5 cm2

) is 

placed in the sample oven, and the oven is tightly sealed. Quartz-fiber filters are required 

because temperatures in excess of 850 °C are employed. The analysis proceeds in inert and 

oxidizing atmospheres. In both, the evolved carbon is catalytically oxidized to carbon 

dioxide (CO2). The CO2 is then reduced to methane (CH4), and CH4 is quantified with a 

flame ionization detector (FID).

OC (and carbonate, if present) is first removed in helium, as the temperature is increased 

to a preset maximum (usually 850 °C or higher). If charring occurs, the filter transmittance 

decreases as the temperature is stepped to the maximum. After OC is removed, an oxygenhelium mix is introduced to effect combustion of the remaining carbon. As oxygen enters 

the oven, light-absorbing carbon is oxidized and a concurrent increase in filter 

transmittance/reflectance occurs. The split (vertical line prior to EC peak in Figure 2) 

between the OC and EC is assigned when the initial (baseline) value of the filter 

transmittance is reached. All carbon removed before the OC-EC split is considered 
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Monitoring Diesel Exhaust in the Workplace

organic, and that removed after the split is considered elemental. If no char is formed, the 

split is assigned prior to removal of light-absorbing carbon. If the sample chars, the split is 

not assigned until enough light-absorbing carbon is removed to increase the 

transmittance/reflectance to its initial value. In general, char is more readily oxidized than 

diesel-particle EC. The delay (i.e., the transit time from sample to FID) between the laser 

and FID signals is considered in the split assignment. Ordinarily, the split is assigned in 

the oxidative mode of the analysis.

Figure 2. Thermogram for filter sample containing organic carbon (OC), carbonate 

(CC), and elemental carbon (EC). PC is pyrolytically generated carbon or “char.” Final 

peak is methane calibration peak. Carbon sources: pulverized beet pulp, rock dust 

(carbonate), and DPM.

EC and OC results are reported in micrograms per square centimeter (μg/cm2

) of the 

sample deposit. The total OC and EC on the filter are calculated by multiplying the 

reported values by the deposit area. In this approach, a homogeneous deposit is assumed. 

For triplicate analyses, the precision (relative standard deviation) is normally under 5%, 

and it is typically 2% or better [NIOSH 1994b (2003 supplement)]. The total carbon (TC) 

in the sample is the sum of OC and EC. If carbonate is present, the carbon in it is 
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Monitoring Diesel Exhaust in the Workplace

quantified as OC, unless a carbonate-subtracted value is requested. Additional details 

about carbonates are given in a following section.

c. Accuracy 

Reference materials are not available for determining the accuracy of OC-EC 

measurements on filter samples of complex carbonaceous aerosols. For this reason, only 

the accuracy of the method in the determination of TC could be examined. No discernable 

differences in the responses of five different organic compounds were found. Linear 

regression of the data (43 analyses total) for all five compounds gave a slope and 

correlation coefficient (r) near unity [slope = 0.99 (±0.01), r2 = 0.999, n = 43]. In addition 

to the OC standards, eight different carbonaceous materials were analyzed. Three different 

methods (including the thermal-optical method) were used, and laboratories reported the 

TC contents of the samples. The samples analyzed included DPM and other types of 

carbonaceous matter (coals, urban dust, humic acid). Thermal-optical results agreed well 

with those reported by the two other laboratories. The variability in TC results for the 

three laboratories ranged from about 1%–7%. These findings [Birch and Cary 1996a; 

NIOSH 1994a (1998 supplement)] indicate that TC is accurately quantified, irrespective of 

sample type. 

d. Limit of detection 

To estimate the method’s limit of detection (LOD), a set of low-level calibration standards 

(ethylenediaminetetraacetic acid [EDTA]) was analyzed [Birch and Cary 1996a; NIOSH 

1994a (1998 supplement)]. The standards covered a loading range from 0.23 to 2.82 μg C 

(or from 0.15 to 1.83 μg C per cm2 of filter). Results of linear regression of the low-level 

calibration data were then used to calculate a LOD as 3 σy/m, where σy is the standard error 

of the regression and m is the slope of the regression line. The LOD estimated through the 

linear regression results was 0.24 μg C, or 0.15 μg/cm2. This value showed good agreement 

with the LOD estimated as 3σblank (three times the standard deviation for blanks), which 

gave a value of about 0.3 μg C. The mean (n = 40) instrumental blank was 0.03 ±0.1 μg C. 

With a 960-L air sample collected on a 37-mm filter and use of a 1.5 cm2 sample portion, 

this LOD translates to an air concentration of about 2 μg/m3. As with all sampling and 

analytical methods, the LOD is a varying number that depends on the instrument, 

sampling parameters, and means by which the LOD is calculated. 

NIOSH Method 5040 was developed for monitoring workplace exposure to DPM, 

especially in mines, where EC concentrations are relatively high (e.g., hundreds of μg/m3). 

However, the thermal-optical technique on which 5040 is based has application to other 

types of carbonaceous aerosols. Thermal-optical analysis also has been applied to studies 

of U.S. workers exposed to carbon nanotubes and nanofibers (CNT and CNF) [Birch et al. 
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Monitoring Diesel Exhaust in the Workplace

2011; Dahm et al. 2012; Dahm et al. 2015; NIOSH 2013]. For these studies, a lower LOD 

was needed because workplace concentrations of CNT/CNF are generally low. As 

recommended in Method 5040, a smaller (25-mm) filter and a higher flow rate were used 

to obtain a lower LOD (about 1 μg EC/m3 or lower). Manual assignment of the OC-EC 

split also was made. Application of Method 5040 to carbon nanomaterials is discussed 

below (see Carbon Nanomaterials).

e. Air sampling 

In the initial evaluation of the thermal-optical method, a set of laboratory-generated air 

samples was analyzed. A dilution tunnel equipped with a dynamometer was used for 

generation of diesel particulate samples. Four EC concentrations, ranging from 23 to 

240 μg/m3 (EC loadings from 2.7 to 27 μg/cm2), were generated. The analytical results 

[NIOSH 1994b (1998 supplement)] indicated that the method met the NIOSH accuracy 

criterion [Kennedy et al. 1995]. The variance was roughly proportional to the mean 

concentration; therefore, the relative standard deviation (RSD) decreased with increasing 

concentration. The accuracy was calculated accordingly. The accuracy was ±16.7% at the 

lowest loading (2.7 μg/cm2), with an overall precision (RSD) of 8.5%. On the basis of a

method evaluation, the NIOSH accuracy criterion requires a confidence limit on the 

accuracy less than 25% at the 95% confidence level. Restated, the criterion dictates that 

greater than 95% of the measurements fall within ±25% of the true value at 95% 

confidence in the method’s validation experiments. The method was considered unbiased 

(i.e., considered the reference method), and the overall precision reflected method 

accuracy. In this initial test, the sample generation and collection system was the main 

source of variability, not the analysis.

 

When only combustion-source EC is present, different samplers can be expected to give 

comparable EC results because particles from combustion sources are generally less than 

1 μm (diameter). As such, the particles are evenly deposited on the filter and collected with 

the same efficiency (near 100%). To confirm this assumption, seven different sampler 

types (open-faced 25-mm and 37-mm cassettes; 298 personal cascade impactor [7 stages, 

0.9-μm cutpoint]; 4 prototype impactors) were used to collect diesel aerosol at the loading 

dock of an express mail facility. The RSD for the mean EC concentration was 5.6% [Birch 

and Cary 1996a]. Based on the 95% confidence limit (19%; 13 degrees of freedom, n = 14) 

on the accuracy, results of this experiment also indicated that the NIOSH accuracy 

criterion [Kennedy et al. 1995] was fulfilled. The amount of EC collected (240 μg per 

sample) would have been equivalent to sampling an air concentration of 250 μg/m3 for 8 h 

at 2 L/min. Variability in the OC results was higher (RSD = 12.3%), which is to be 

expected when different samplers are used to collect aerosols that contain semivolatile 

(and volatile) components because these can have a filter face velocity dependence.
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Monitoring Diesel Exhaust in the Workplace

Similar performance was obtained from collected samples in an underground 

molybdenum mine. Five different sampler types were used (closed-face 25-mm and 37-

mm cassettes; 298 cascade impactor [7 stages, 0.9-μm cutpoint]; cyclone with filter; inhouse impactor). The RSD for the EC results (mean EC = 297 μg/m3

) was 7%. The EC 

deposits obtained with all five sampler types were homogeneous, even when the ore 

deposit was visually heavier in the center of the filter (e.g., with the closed-face 37-mm 

cassette). Although the dust loading was higher in the center of the filter, portions taken 

from the center gave equivalent EC results, indicating the ore contained no EC 

component. The TC results for the center portions were only slightly higher, so this 

particular ore was mostly inorganic. EC concentrations found with three different sampler 

types (nylon cyclone, open-faced cassette, and impactor with submicrometer cut) also 

were comparable in a study of railroad workers [Verma et al. 1999].

If high levels of other dusts are present, a size classifier (e.g., impactor and/or cyclone) 

should be used to prevent filter overloading, particularly if the dust is carbonaceous. In the 

latter case, a size classifier provides a more selective measure of the diesel-source OC. It 

also provides a better measure of the diesel-source EC if the dust contains an EC 

component, which is less common. A finely ground sample of the bulk material can be 

analyzed to determine whether a specific dust poses potential interference [Birch and Cary 

1996a]. Depending on the dust concentration, size distribution, and target analyte, an 

impactor may be required.

For mines, the Mine Safety and Health Administration (MSHA) recommends a specialized 

impactor to minimize collection of carbonates and other carbonaceous dusts [66 Fed. Reg. 

5706 (2001)]. An impactor can greatly improve the selectivity of the TC measurement in 

some cases, but it may exclude a small amount of the DPM. Then, too, some OC 

interferences cannot be excluded on the basis of size (e.g., condensation aerosols, fumes, 

wood and cigarette smokes). If present in the sampling environment, these materials can 

positively bias the OC (TC) results to some degree, depending on their relative 

concentrations and the sampling location. Although 37-mm or 25-mm cassettes are often 

suitable for general industry, the required sampler depends on the sampling environment.

f. Carbonates 

The presence of carbonate is indicated by a narrow peak during the fourth temperature 

step in helium (Figure 2). Its presence is verified by exposing a second portion of the filter 

to hydrogen chloride (HCl) vapor prior to analysis. When the acidified portion is 

analyzed, a diminished (or absent) peak during the fourth temperature step is indicative of 

carbonate in the original sample. (Note: Acid treatment may sometimes alter the 
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appearance of the EC profile in the thermogram [output signal of thermal-optical 

instrument], but the EC result itself should not be affected significantly.)

A desiccator containing concentrated HCl (added to the desiccator or a petri dish placed 

at the bottom of it) can be used to acidify the sample portions. The desiccator, or 

alternative vessel, should be used in a well-ventilated hood. The filter portions are placed 

on the desiccator tray, and the tray is placed in the desiccator. A wetted pH indicator stick 

can be used to check acidity. A wetted indicator stick inserted between the desiccator lid 

and base should give a pH near 2. Portions should be exposed to the acid vapor for about 

1 hour (large particles may require more time). After acidification, the tray is placed on a 

clean, inert surface inside the hood. The residual acid on the portions should be allowed to 

volatilize in the hood for at least an hour prior to analysis.

Environmental samples typically contain little (if any) carbonate, but levels in some 

occupational samples can be quite high. For example, respirable dust samples collected in 

limestone and trona mines can contain high levels of calcium carbonate and sodium 

sesquicarbonate, respectively. In such cases, acidified samples give a better measure of the 

diesel-source OC (TC). If the carbonate loading is relatively high (e.g., carbonate carbon 

>10% of the TC), the difference between the TC results (before and after acidification) 

gives an estimate of carbonate carbon (CC). Estimation of CC by difference assumes the 

carbonate is evenly deposited on the filter, which may not always be the case. It also 

assumes that OC loss through acidification is negligible. The latter assumption is generally 

true for workplace samples, but depends on sample composition and loading.

Alternatively, a carbonate-subtracted OC (TC) result can be obtained through separate 

integration of the carbonate peak. This applies only to carbonates (e.g., calcium carbonate) 

that can be removed as a single peak during the fourth temperature step in helium. The 

carbonate of interest should be analyzed to ensure this is the case. A minor adjustment 

(lower) to the third temperature step may be necessary to prevent partial loss during this 

step. Sodium sesquicarbonate (trona) is removed over multiple temperature steps. Samples 

containing trona should be acidified if a carbonate-subtracted result is desired. (Note: At 

elevated temperature, trona and other compounds containing sodium can etch the quartz 

oven wall and sample holder. An etched surface can reduce laser throughput. Avoid frequent 

analysis of high-sodium materials and direct contact of these materials with the oven wall 

and sample holder.) 

The presence of a small “peak” during the fourth temperature step is not necessarily due to 

carbonate. Other carbonaceous matter (e.g., char) is sometimes removed during the fourth 

step. Unlike carbonates, which produce a relatively sharp peak, other materials typically 

evolve as a small, broad peak. If determination of relatively low (e.g., CC <10% of TC) 
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amounts of CC is desired, a second portion of the sample should be acidified and analyzed 

to verify the presence of carbonate and reveal any underlying baseline features contributed 

by other materials. A more accurate estimate of CC can then be obtained by integrating 

the first (nonacidified) sample over the missing peak area (i.e., area removed through 

acidification).

g. Carbon nanomaterials 

NIOSH Method 5040 was developed for monitoring workplace exposure DPM, as EC, but 

the thermal-optical analysis technique on which it is based has application to other types 

of carbonaceous aerosols and materials. Method 5040 has been applied to field studies on 

carbon nanomaterials, wherein EC was used as a measure of exposure to carbon 

nanotubes and nanofibers (CNT and CNF) [Birch et al. 2011; Dahm et al. 2012; Dahm et 

al. 2015; Fatkhutdinova et al. 2016; NIOSH 2013]. Because CNT/CNF are composed of 

EC, EC is a quantitative measure of airborne CNT/CNF [Birch et al. 2011; NIOSH 2013]. 

In 2013, NIOSH set a recommended exposure limit (REL) for CNT/CNF: an 8-hr time 

weighted average (TWA) of 1 μg/m3 as respirable EC [NIOSH 2013].

For application of NIOSH 5040 to CNT/CNF, a manual OC-EC split is assigned. The 

larger size (μm versus nanoscale DPM) and agglomerate structure of CNT/CNF particles, 

combined with low air concentrations (i.e., filter loadings), make the 5040 auto-split 

unreliable [Dahm et al. 2015; NIOSH 2013]. Adjustments to the thermal program also may 

be needed depending on the material, size fraction collected, and environmental 

background [Birch et al. 2011; Dahm et al. 2015; Doudrick et al., 2012; NIOSH 2013]. 

Analysis of bulk and background samples is recommended [NIOSH 2013]. Determining 

the thermal profiles of bulk materials produced/processed in a given workplace, and using 

this information, along with results for background samples, assists manual assignment of 

the OC-EC split and can provide more accurate CNT/CNF measurement.

 

Because CNT/CNF oxidize over a range of temperatures, it is important to analyze bulk 

and background samples [Birch et al. 2011; Dahm et al. 2015; NIOSH 2013]. For example, 

Mitsui-7 MWCNT (Mitsui & Co., LTD, Tokyo, Japan), used in early NIOSH toxicity 

studies, is more difficult to oxidize (onset > 700 °C). A NIOSH (Cincinnati, OH, USA) 

laboratory normally uses a final temperature (in oxygen-helium) of 920 °C, which is 

adequate provided that the loading is not too high and the hold time (at final temperature) 

is long enough. A higher (up to about 940 °C) or lower maximum can be used, and the 

heating period at maximum temperature can be adjusted according to the type and 

amount of material analyzed. If the signal returns to baseline, EC in the sample is 

completely oxidized. If not (i.e., there is signal ‘tailing’ into the calibration peak), the final 

temperature can be increased and an extended period can be used. In addition to Mitsui-7 
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(MWCNT), some graphite products are difficult to oxidize. On the other hand, if a 

material is completely oxidized at 875 °C, prolonged heating at 920 °C is unnecessary. In 

some cases (e.g., early EC loss), a lower maximum temperature in the helium also may be 

appropriate [NIOSH 2003]. These changes are minor adjustments that can be made to 

optimize the thermal analysis.

 

In general, most CNT/CNF materials can be oxidized from the filter. However, there may 

be instances where sample oxidation is incomplete, even with holding at 920 °C (or 

higher) for an extended period (e.g., high loadings of graphite products or Mitsui-7). For 

bulks analyses, only small amounts (e.g., about 10 μg or less) of material should be 

analyzed, to better match the loadings of the field samples, for comparison with the bulk. 

If residual material (CNT) remains (post analysis), incomplete oxidation is obvious as the 

signal does not return to baseline and a filter deposit is visible.

 

Even with the aid of bulks and judicious choice of background samples, it may not be 

possible to exclude EC background, posing an interference problem at low CNT/CNF 

loadings. Also, amorphous carbon in the CNT/CNF materials may cause positive bias 

[NIOSH 2013]. If the oxidation rates of the amorphous carbon and CNT/CNF are 

sufficiently different, it may be possible to remove the amorphous material to a greater 

extent, but the oxidation rates may be too similar for speciation.

h. Sampling artifacts: organic aerosol 

1.) Face Velocity

Quartz-fiber filters are routinely used to collect airborne particulate matter for carbon 

determination. Quartz filters have high collection efficiency for particulate carbon, but 

collection of organic aerosol is artifact prone [Cui et al. 1998; Kirchstetter et al. 2001; 

McDow and Huntzicker 1990; Turpin et al. 1994; Zang and McMurry 1992]. 

Particulate carbon (EC and nonvolatile OC) results should not depend on filter face 

velocity. This is true for EC, but OC (and TC) results can have a face velocity 

dependence due to sampling artifacts. Namely, adsorption of organic vapor can 

positively bias the results, while evaporative losses have the opposite effect. Although 

both processes may occur, studies indicate vapor adsorption is the dominant artifact 

[Kirchstetter et al. 2001; Turpin et al. 1994]. In one study [Turpin et al. 1994], the OC 

(TC) results for air samples collected at face velocities of 20, 40, and 80 cm/sec had a 

face velocity dependence before, but not after, correction for adsorbed vapor. Lack of 

dependence after correction supports adsorption as the dominant artifact. At the much 

lower face velocities typical of occupational monitoring (e.g., about 4 cm/sec with a 

37-mm filter and 2 L/min flow rate), OC losses induced by pressure drop across the 

filter are expected to be minor relative to adsorption. OC sampling artifacts were 
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acknowledged when the thermal-optical method was proposed [55 Fed. Reg. 110 

(1990); Birch 1991], but this issue was not investigated prior to publication of NIOSH 

5040 because an EC surrogate was recommended. In view of the MSHA-proposed air 

standard for TC (proposed after 5040 publication), and the fact that both OC and EC 

are determined, the issue was later addressed. 

2.) Adsorbed OC

Correction for adsorbed organic vapor through use of traditional blanks (media or 

field) may not be accurate because the amount of vapor adsorbed by them is variable 

[Birch et al. 1999a]. More importantly, traditional blanks collect vapor passively, while 

the samples collect it actively (during sampling). A more representative correction for 

adsorbed organic vapor can be made through use of two filters in tandem [Kirchstetter 

et al. 2001; Turpin et al. 1994]. The air sample is collected with a sampler containing a 

Teflon® or quartz upper filter and a bottom quartz filter. After sampling, the bottom 

filter is used to correct for adsorbed vapor. The vapor adsorbed on the bottom filter 

more closely represents that adsorbed on the sample filter (quartz) because both are 

collected actively. Use of two quartz filters is preferable to Teflon and quartz because 

the filter used to correct for adsorbed vapor is in the same sampler; however, the 

second (bottom) quartz filter may underestimate the amount of adsorbed OC relative 

to a quartz filter under Teflon [Kirchstetter et al. 2001; Turpin et al. 1994]. These 

results have been attributed to depletion of the vapor concentration by the top quartz 

filter, which presents a lower concentration to the bottom one. A shorter equilibration 

time (for partitioning between gas phase and adsorbed state) is expected with Teflon 

filters because they have less surface area and are more inert than quartz. Thus, less 

adsorption is expected on Teflon relative to quartz. Because differences in the amount 

of adsorbed OC have been reported for the two sampling configurations, both were 

examined.

Six sets of air samples were collected. Two pre-cleaned quartz-fiber filters were loaded 

into each of eight 37-mm cassettes. In four of these, a Teflon filter was then placed on 

top. The bottom quartz filters in all eight cassettes, as well as the four directly under the 

Teflon filters, provided measures of adsorbed OC. Filters in four additional cassettes 

served as passive field blanks, meaning no air was pulled through them. The second 

(bottom) quartz filters in the cassettes with the Teflon filters were compared to the 

bottom quartz filters in those having only a quartz pair. If negligible adsorption occurs 

on Teflon, the bottom quartz filters in the two different sampling configurations 

should give equivalent results. To avoid possible variability due to lot-to-lot 

differences, filters from the same lot were used to collect a given sample set.
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3.) Portable dust chamber

A portable dust chamber [Kogut et al. 1997] designed for sampling in mines was used 

for sample collection. The chamber allowed simultaneous collection of up to eighteen 

samples. Twelve cassettes were mounted inside the chamber in a symmetrical fashion. 

Samples were collected on six different days, five days in a loading dock area and a 

sixth near an outside smokers’ shelter. A diesel truck was operating in the loading 

dock, over different periods, on three of the days. Personal air pumps were 

programmed to run at 2 L/min over an 8-hour period. In one case, the pumps were 

stopped after 23 minutes. The short sampling period provided a low loading of diesel 

exhaust (a light deposit was visible) and was of interest with respect to the amount of 

adsorbed carbon collected in the short time frame.

Results for the six sample sets are shown in Table1 and Figure 3. The QQ2 results 

correspond to the bottom quartz filters in the four cassettes containing quartz pairs 

only. The TQQ1 results are for the four filters directly beneath Teflon filters; TQQ2 

results are for quartz filters beneath the TQQ1 filters (i.e., top Teflon, middle TQQ1, 

bottom TQQ2). The mean carbon on the bottom quartz filters ranged from 1.35 to 

3.44 μg/cm2, and the variability (RSD) ranged from about 4% to 12% on a given day. 

The loading found on the two sets of bottom filters (QQ2 and TQQ2) was not 

statistically different, so all bottom filters were pooled in the calculation of the mean. 

The pooled means for the bottom quartz filters (BQ), the means for the quartz under 

Teflon (TQQ1), and the corresponding OC and EC loadings are plotted in Figure 3. 

Results for the TQQ1 filters were only slightly higher than those for the bottom quartz 

filters, even with the 23-minute sampling period (set 4). Thus, using a separate cassette 

that has a top Teflon filter appears unnecessary. Although differences between the 

amount of carbon adsorbed on a quartz filter under another quartz and on one 

beneath a Teflon filter reportedly can be significant [Kirchstetter et al. 2001; Turpin et 

al. 1994], these studies concerned environmental monitoring at much higher flow 

rates. The negligible differences reported here most likely relate to the lower flow rates 

and filter face velocities used.
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Figure 3. Results for six sets of air samples collected in two outdoor locations. OC 

and EC results are the means for the top quartz filters (QQ1) of quartz-filter pairs. 

TQQ1 is the mean carbon (total) result for quartz filters placed directly beneath 

Teflon. BQ is the mean carbon result for all bottom quartz filters (i.e., QQ2 and 

TQQ2). See text for additional details.

Table 1. Results summary for total carbon (TC, μg/cm2

) from six sets of air samples 

collected in two outdoor locations (SD = standard deviation; RSD = relative 

standard deviation). See text for details.

Set Location QQ1 QQ1 QQ1 TQQ1 TQQ1 TQQ1 Bottom Bottom Bottom Bottom Bottom Adsorbed 

mean SD RSD mean SD RSD quartz quartz quartz quartz quartz fraction 

(%) (%) mean, mean, mean, mean, mean, (%)

QQ2 TQQ2 pooled SD RSD (%)

1 shelter 4.81 0.47 9.77 3.66 0.43 11.8 3.46 3.41 3.44 0.30 8.80 76

2 dock 3.52 0.25 7.10 2.82 0.36 12.8 2.48 2.33 2.43 0.23 9.57 80

3 dock 3.61 0.05 1.39 2.89 0.04 1.38 2.41 2.42 2.41 0.18 7.45 80

4 dock 4.59 0.18 3.92 1.79 0.24 13.4 1.41 1.36 1.38 0.09 6.46 30

5 dock 8.43 0.33 3.91 1.84 0.35 19.0 1.35 1.36 1.35 0.16 12.0 22

6 dock 22.8 0.35 1.54 3.62 0.18 4.97 3.10 3.18 3.14 0.13 4.09 16

The mean carbon loading (Table 1) on the sample filters (QQ1) was about 4 μg/cm2

 on 

four different days (sample sets 1–4). A diesel truck was operating briefly on one of 

these days (set 4). On two other days (sets 5 and 6), the truck was running for longer 

periods, as evidenced by the higher EC loadings (Figure 3). The corresponding TC 

loadings on these two days were about 8 μg/cm2 and 23 μg/cm2. Over the loading 

range, the amount of adsorbed OC had no dependence on the TC loading, and the OC 

results for the bottom quartz filters (1.35 to 3.44 μg/cm2

) were higher than those for the 

passive field blanks (0.17 to 0.74 μg/cm2). Two of the results (sets 4 and 5) were slightly 

lower than the others. In one case (set 4), the sampling period (23 minutes) was 

shorter, but the adsorbed OC results for the bottom filters in both sampling 

configurations (i.e., QQ2 and TQQ2) were equivalent and comparable to the upper 
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quartz filters (TQQ1), below the Teflon. This implied that partitioning [Turpin et al. 

1994] (between gas and adsorbed states) on the sample filters (QQ1) had reached 

equilibrium; thus, the lower result was not due to lack of equilibration. In large part, 

the results for passive blanks provided a likely explanation for the lower value. The 

mean blank (passive) for this set was 0.17 (±0.10) μg/cm2, which was the lowest value 

obtained. Passive blank results for the other sets (excluding set 5) ranged from 

0.25 (±0.07) to 0.74 (±0.14) μg/cm2

. It is not known whether the same explanation 

applied to set 5 because passive blanks for this set were accidentally contaminated 

(results voided). As in sets 1–3 and 6, set 5 was collected for an 8-hour period, but it 

was collected during the spring, while the other sets were collected during midsummer. 

Changed weather conditions may explain the lower value.

Over three days, the adsorbed OC fraction (calculated as the TQQ1 TC mean divided 

by QQ1 TC mean) constituted about 80% of the TC on the sample filters (QQ1). On 

three other days (sets 4, 5, and 6), the adsorbed OC fraction was 39%, 22%, and 16% of 

the total. A backup quartz filter provided a better correction for adsorbed carbon than 

a traditional blank would have. Traditional blanks (media and field) underestimated 

the adsorbed OC, causing overestimation of the true particulate OC (TC) 

concentration. The need for this correction, and the number of backup filters analyzed, 

depends on the sampling strategy and environment. The lower the OC loading, the 

greater the influence of adsorbed vapor on the particulate OC measurement. If the EC 

loading also is low, the same holds true for the TC result. Thus, when carbon loadings 

are low, correction for adsorbed OC is important for accurate measurement of 

particulate OC and TC concentrations. However, the correction addresses adsorbed 

vapor only—not interference of less volatile materials (e.g., components in cigarette 

and wood smokes, oils) collected primarily on the top filter. Depending on vapor 

pressure, sampling conditions (temperature, flow rate, sampling period), and filter 

loading, these materials also collect on the bottom filter to some extent. The bottom 

filter cannot correct for this interference. If high OC loadings are found on the bottom 

filter, less volatile OC interferences should be suspected.

i. EC oxidation in helium 

In the NIOSH 5040 analysis, oxidation of original EC (as opposed to char) is sometimes 

seen during the last temperature step in helium, but this is generally not common. If early 

removal occurs, it is important to ensure that oxygen contamination is not responsible. 

This check can be performed by analyzing a sucrose standard solution applied to a clean, 

unused filter punch. Although char loss is sometimes observed with sucrose, the filter 

transmittance normally does not reach its initial value until after oxygen is introduced. If 

oxygen contamination can be ruled out, the early EC loss is most likely due to oxidants in 
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the sample. In our laboratory, EC loss in helium has been observed with samples from a 

few mines [Birch 1998b]. With these samples, the filter transmittance exceeded its baseline 

(initial) value during the last temperature step in helium. No sample pyrolysis was 

apparent, so the increase in transmittance was not caused by char loss. Moreover, analysis 

of sucrose standards revealed only a minor increase in transmittance in helium, and no 

increase was seen with samples from other mines. For these reasons, oxygen 

contamination was ruled out. On average, about 10% of the EC was removed at 870 °C in 

helium, but the carbon removed was included in the EC result because the OC-EC split 

was assigned when the initial transmittance was reached (in helium). Reducing the 

maximum temperature to 750 °C was recommended [NIOSH 1994a, 1994b] for these 

types of samples (i.e., no sample charring and early splits in helium). Early EC loss was not 

seen at this temperature. A lower (<750 °C) temperature may be required, depending on 

the sample composition. Our analytical results and precision were not affected by the early 

splits, but a reduced temperature was recommended in the interest of interlaboratory 

precision, which might be adversely affected. Interlaboratory testing was not conducted to 

determine if this is the case, and, if so, whether a lower temperature would improve 

precision.

j. Reference materials 

As mentioned, a suitable reference material for OC and EC is not available. A Certified 

Value (17.68% g/g) for the TC mass fraction of an urban dust standard (SRM 1649a, 

formerly SRM 1649) was reported by the National Institute of Standards and Technology 

(NIST), but only Information (not Certified) Values were provided for the EC content. 

The EC content (as EC:TC fractions) was determined by a variety of methods. As 

expected, based on prior studies [Birch 1998a; Chow et al. 2001; Countess 1990; Norris et 

al. 1999; Schmid et al. 2001], the results were quite variable. The EC:TC fractions found by 

13 methods ranged from about 7% to 52%, and the data were distributed in three clusters. 

Method bias was not evaluated. The reported range is too broad to use this material as an 

analytical standard. According to the Certificate of Analysis (1649a), the reported values 

may be useful for comparison with results obtained by similar methods, but this may not 

be the case for methods with optical corrections because filter samples are not available. 

Depending on its placement, bulk dust on a filter can present different optical properties, 

which may increase variability. Sample composition is an important consideration in the 

production of an analytical reference material. As is true with many standards, no single 

OC-EC material can be representative of all samples because there are many sources of 

particulate carbon and many different monitoring sites—both occupational and 

environmental. Production of an OC-EC reference material is further complicated by the 

variability between methods (SRM 1649a provides only Information Values for different 

methods), and the need of a filter-based material for thermal-optical methods.
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Because sample components that char pose a potential interference in the determination 

of EC, and interlaboratory variability is greater when samples contain them, analysis of 

organic materials that char provides an important quality assurance check. Sucrose serves 

as both an analytical standard and a check of the method’s char correction, but sucrose is a 

simple carbohydrate (a disaccharide). Air samples can contain complex particulate matter, 

such as wood and cigarette smokes, plant debris (cellulose), and products of biomass 

burning. In some cases, biomass may contribute a sizable fraction of the sample carbon. 

About 38% of the particulate carbon in SRM 1649a (an urban dust) is derived from 

biomass (see Certificate of Analysis [NIST 2001] and related publication [Currie et al. 

2002]). For these reasons, a material more representative of the plant-derived components 

in air samples was sought.

Various plant-derived materials were examined to determine whether any would be useful 

as laboratory control samples. A well-characterized, stable material that chars also would 

be useful for proficiency testing, and it could potentially be used to produce a standard 

reference material. After a preliminary screening of candidate materials (e.g., agarose, 

alginic acid, starch, and cellulose), alginic acid was selected for further study. Alginic acid

is a polysaccharide derived from sea kelp [Morrison and Boyd 1973], it has adequate 

solubility in water, and it forms a significant amount of char during the analysis.

To prepare a solution, about 150 mg of alginic acid (Sigma-Aldrich, St. Louis, MO) was

added to a test tube containing 10 mL of purified water (ultrafiltered, type I). The tube was 

shaken vigorously and allowed to sit overnight at room temperature. The following day, 

the tube was shaken again and then centrifuged to settle the suspended material. The clear 

supernatant was removed and syringe filtered. Aliquots of the alginic acid solution were 

applied to clean filter punches. The solution was analyzed over a seven-week period. 

Results of these analyses are plotted in Figure 4. No evidence of solution degradation was 

observed. A mean carbon loading of 13.98 μg/cm2 (RSD = 3.06%, n = 22) was obtained for 

10-μL aliquots of the solution. This translates to a carbon concentration of 2.06 μg/μL. 

About half of the carbon in alginic acid remained on the filter as char before oxygen was 

introduced, but the mean EC result (0.04 ±0.06 μg/cm2

) was not statistically different from 

that for the media blanks (0.06 ±0.03). Therefore, the pyrolysis correction was accurate.
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Figure 4. Results for clean punches (blanks) spiked with 10-μL aliquots of alginic acid 

solution. Analyses done over a 7-week period. The middle horizontal line is the mean 

result (13.98 μg/cm2

). The upper and lower dashed lines are ±5% of the mean.

Sets of diesel soot samples spiked with the alginic acid solution also were examined. 

Multiple punches (1.5 cm2) were taken from a soot sample collected on an 8” x 10” filter. 

The punches were then analyzed before and after spiking with a 10-μL aliquot of the 

solution. Punches in a given set were taken from the same area of the filter to minimize 

variability, and the solution was applied in a consistent manner (dispersed evenly on one 

end of the punch where the laser penetrates it). Mean TC results for the unspiked and 

spiked samples are provided in Table 2. Results for three sample sets (A, B, and C) are 

reported. For each set, the difference between the mean results for the unspiked and 

spiked samples is reported; this difference is the alginic acid carbon. Based on the mean 

(13.98 μg/cm2) for 22 spiked blanks, recoveries of alginic acid carbon from the soot 

samples were 96%, 101%, and 110% (sets A, B, and C, respectively). The pooled recovery 

for all spiked samples was 97%.

Table 2. Recovery of alginic acid carbon (means and standard deviations, n=3; μg/cm2

) 

from unspiked diesel particulate samples and samples spiked with 10-μL alginic acid

solution.

Sample set Unspiked Spiked Difference Recovery* (%)

A 22.9 (1.49) 34.9 (0.66) 13.5 96

B 22.3 (0.28) 36.4 (0.65) 14.1 101

C 22.6 (0.33) 38.0 (0.80) 15.4 110

All 22.6 (0.84) 36.1 (1.32) 13.5 97

*Based on mean result (14.0 μg/cm2) for 22 analyses of spiked blank punches; analyses 

carried out over a 7-week period.
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Table 3. Results for elemental carbon (EC; means and standard deviations, n=3; 

μg/cm2) from spiked and unspiked diesel particulate samples.

Sample set Unspiked Spiked Difference

A 4.24 (0.20) 4.36 (0.42) 0.12 (0.47)

B 3.88 (0.01) 3.78 (0.12) -0.10 (0.12)

C 4.32 (0.14) 4.43 (0.12) 0.11 (0.18)

EC results for the three sample sets are listed in Table 3. Spiked sample results for two sets 

(A and C) were slightly higher, while those for a third set (B) were slightly lower. 

Differences between the mean EC loadings on the unspiked and spiked samples were not 

statistically different. Thus, an accurate correction was again made for the char formed 

through pyrolysis of alginic acid.

k. Quality assurance 

1.) NIOSH 

Quality assurance (QA) procedures followed at NIOSH and NIOSH contract 

laboratories have included repeat analyses, analysis of OC standards, and analysis of 

media and field blanks. Results for repeat analyses of over 200 filter samples at a 

contract laboratory (DataChem Laboratories, Inc., Salt Lake City, UT) gave a pooled 

RSD (95% confidence) of about 4% for EC and TC, and 5% for OC [Birch et al. 1999a]. 

The samples were collected in mines, and some contained other carbonaceous matter 

such as limestone. In the analysis of OC standards, recovery was typically within 6% of 

the expected value [Birch et al. 1999a]. For example, the mean recovery for more than 

400 sucrose standards analyzed at DataChem over a one-year period was 99.97% 

±6.07%. Nearly identical results (100.33% ±5.15%, n = 462) were obtained by Clayton 

Laboratory Services (Novi, MI). Different sets of field blanks (41 at DataChem and 129 

at Clayton) analyzed by the two laboratories also gave comparable results [Birch et al. 

1999a]. Mean blank results (μg carbon per filter) were as follows: DataChem OC = 9.96 

±7.05, Clayton OC = 12.80 ±5.76; DataChem EC = –0.54 ±1.89, Clayton EC = –0.08 

±2.14; DataChem TC = 9.26 ±7.30, Clayton TC = 12.70 ±6.18.

In lieu of an OC-EC reference material, a limited confirmation of results by a second 

laboratory is advised. For example, our laboratory (NIOSH, Cincinnati, OH) has 

routinely performed repeat analyses on samples previously analyzed at a contract 

laboratory. As an added check, subsets of representative field samples also have been 

analyzed by a third laboratory. In the analysis of fifty samples (analyzed at DataChem, 

NIOSH, and Clayton) from mines, the pooled RSD (95% confidence) was about 6% for 

TC, 10% for EC, and 12% for OC [Birch et al. 1999a]. These results are consistent with 

those found in a previous round robin [Birch 1998a].
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2.) EPA

The EPA conducted a special study [EPA 2001] as part of the QA oversight for the 

PM2.5 Speciation Trends Network (STN). Samples collected as part of this network 

were removed from refrigerated storage eight months after they were analyzed at 

Research Triangle Institute (RTI, Research Triangle Park, NC). The samples were then 

shipped to the EPA New England Regional Laboratory (NERL, Lexington, MA) for 

reanalysis. Good interlaboratory agreement was reported. The acceptance criteria for 

the archived samples were based on filter loading (μg/cm2). The three acceptance 

criteria are the following: a difference ≤1 μg/cm2 for loadings less than 5 μg/cm2, a 

relative percent difference (RPD) ≤20% for loadings from 5 to 10 μg/cm2, and an RPD 

≤15% for loadings above 10 μg/cm2. The OC, EC, and TC data had 85%, 96%, and 93% 

of the interlaboratory results, respectively, within the acceptance criteria. The lower 

percentages for OC and TC are thought to reflect contamination of lightly loaded 

samples from adsorbed vapors and handling. An OC blank (passive) correction was 

applied but may not have been representative of the entire sample set. Additional 

comparison results are discussed in the next section: Interlaboratory Comparisons.

3 Interlaboratory comparisons

a. NIOSH 5040

When results of the initial method evaluation were published [Birch and Cary 1996a], an 

interlaboratory comparison was not possible because the thermal-optical instrument was 

available in only one laboratory. Interlaboratory comparisons are especially important in 

this type of analysis because variable results have been obtained by different OC-EC 

methods [e.g., Birch 1998a; Chow et al. 2001; Countess 1990; Norris et al. 1999; Schmid et 

al. 2001;], and a reference material is not available. After additional laboratories acquired 

thermal-optical instruments, a round robin comparison [Birch 1998a] was conducted. 

Matched sets of filter samples containing different types of complex carbonaceous aerosols 

were distributed to eleven laboratories. Six of the eleven analyzed the samples according to 

NIOSH 5040, while five used purely thermal (i.e., no char correction) methods. Good 

interlaboratory agreement was obtained among the six laboratories that used NIOSH 

5040. In the analysis of samples containing DPM, the variability (RSD) for the EC results 

ranged from 6% to 9%. Only low EC fractions were found in wood and cigarette smokes. 

Thus, these materials pose minimal interference in the analysis of diesel-source EC. In 

addition, only minor amounts of EC were found in two OC standards that char: about 1% 

and 0.1% for sucrose and the disodium salt of ethylenediaminetetraacetic acid (EDTA), 

respectively. Two aqueous solutions of OC standards were included in the comparison as a 

check on the validity of the char correction and accuracy of the TC results. Variability 

(RSD) of the TC results for the two standard solutions and five filter samples ranged from 

3% to 6%.
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Note: In the round-robin study [Birch et al. 1996a] discussed above, the maximum 

temperature in helium ranged from about 850 °C to 900 °C. Comparable results were 

obtained because some differences in the thermal program do not affect the results 

significantly (e.g., also see Chai et al. 2012). For the purpose of standardization, use of the 

same maximum is recommended. 

Results of other interlaboratory comparisons on NIOSH 5040 have been reported. In one 

study [Schauer et al. 2001], seven environmental aerosol samples were analyzed in 

duplicate by eight laboratories. Four samples were collected in U.S. cities, and three were 

collected in Asia. Variability of the EC results ranged from 6% to 21% for six samples 

having EC loadings from 0.7 to 8.4 μg/cm2

. Four of the six had low EC loadings 

(0.7 μg/cm2 to 1.4 μg/cm2). The variability of the OC results ranged from 4% to 13% (OC 

loadings ranged from about 1 to 25 μg/cm2). Results for TC were not reported, but the 

variability reported for the OC results should be representative of that for TC because the 

samples were mostly OC (75% to 92%). For another comparison [Chai et al. 2012], 

matched-filter sets with known OC–EC contents were generated and distributed to six 

laboratories. In this study, the maximum temperature in helium varied from 650 °C to 

850 °C, but good agreement between the participating laboratories was found. Results 

indicated a uniform carbon distribution for the filter sets. Relative standard deviations for 

mean TC, OC, and EC results for seven laboratories were less than 10%, 11%, and 12% 

(respectively). Except for one EC result (RSD = 16%), RSDs reported by individual 

laboratories, for TC, OC, and EC, were under 12%. The method of filter generation is 

generally applicable and reproducible. Depending on the application, different filter 

loadings and types of OC materials can be employed. Matched filter sets can be used for 

determining the accuracy of OC–EC methods, which are operational.

b. Other methods 

1.) Thermal methods

Different thermal methods have given consistent agreement for TC, but OC-EC results 

have been quite variable [Birch 1998a; Chow et al. 2001; Countess 1990; Norris et al. 

1999; Schmid et al. 2001]. The degree of variability depends on the sample type. In 

general, there is greater disagreement among methods when samples contain materials 

that char [Birch 1998a; Countess 1990; Schmid et al. 2001] (e.g., wood and cigarette 

smokes). Methods that employ a lower maximum temperature and/or do not correct 

for char obtain results that are positively biased relative to NIOSH 5040 [Birch 1998a]. 

When a lower temperature (typically 550 °C) is used, less thermal breakdown of 

refractory organic components (and possibly carbonates) may occur. This, as well as 

lack of char correction, can positively bias the EC results. In the round robin study 

[Birch 1998a] discussed above, three laboratories employed four purely thermal 
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methods that specified a maximum in nitrogen (not helium) of about 550 °C. Unlike 

the NIOSH 5040 results, these laboratories reported high EC contents for the two OC 

standards (about 52% for sucrose and 70% for EDTA). Similar findings (i.e., positive 

bias) for thermal methods having no char correction were obtained in another 

international round robin [Schmid et al. 2001]. 

 

In contrast to round-robin results obtained previously [Birch 1998a], relatively good 

agreement was found in a comparison [Birch et al. 1999b] between NIOSH 5040 and a 

thermal method, ZH 1/120.44, used in Germany [Dahmann 1997; ZH 1/120.44]. The 

comparison was limited to two laboratories. Method ZH 1/120.44 specified a 550 °C 

maximum in nitrogen. The other European laboratories that participated in the 

previous round robin [Birch 1998a] used variations of this method. In the European 

laboratories, nitrogen was used as the inert gas, and carbon determination was based 

on coulometric titration of carbon dioxide. For the comparison [Birch et al. 1999b], 

samples were obtained in a mine where diesel equipment was being operated. The 

samples had a much higher EC content (about 50%) than did the round robin samples. 

No charring was noted in the thermograms, and only a minor amount of carbon was 

removed above 550 °C. Although differences in the OC-EC results were again seen, 

they were minor relative to those obtained in the round robin [Birch 1998a]. The mean 

EC fractions (EC:TC) found with methods ZH 1/120.44 and NIOSH 5040 were 

0.53 (σ = 0.19) and 0.46 (σ = 0.15), respectively. This relatively minor difference in the 

reported fractions is attributed to the different thermal programs employed.

2.) IMPROVE method

Another thermal-optical method, called the IMPROVE (Interagency Monitoring of 

Protected Visual Environments) method [Chow et al. 1993], also was included in the 

NIOSH 5040 round robin [Birch 1998a]. Relatively good agreement between NIOSH 

5040 and the IMPROVE method was obtained, although the IMPROVE EC was 

consistently higher. The carbon analyzers used for the two methods are based on 

similar measurement principles, but they differ with respect to design and operation. 

For example, the optical correction in NIOSH 5040 is based on filter transmittance, 

whereas that for the IMPROVE method is reflectance based. When the 

intercomparison study was conducted, the instrument (Sunset Laboratory, Inc., Forest 

Grove, OR) used for NIOSH 5040 incorporated a pulsed diode laser (670 nm) and 

photodetector positioned on opposite sides of the filter, while the Desert Research 

Institute (DRI, Reno, NV) instrument employed for the IMPROVE method used a 

quartz tube and fiber optic to measure helium–neon laser light (632.8 nm, 

unmodulated) reflected from the filter surface. In addition to instrumental differences, 

NIOSH 5040 specifies a higher maximum temperature (typically 850 °C or higher) in 

helium than the IMPROVE method (550 °C). As discussed in the preceding section, a 
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higher temperature is used to better remove refractory OC components and 

carbonates. In the NIOSH 5040 analysis, the transmittance of some samples continues 

to decrease as the temperature is stepped to 850 °C, indicating further carbonization 

above 550 °C. Additional thermal breakdown and charring result in lower EC results 

because char correction is larger, and volatile pyrolysis products are released to a 

greater extent. Although environmental samples contain only small amounts (if any) of 

carbonate, levels in some workplaces (e.g., mines, construction sites) can be relatively 

high. Collection of carbonate can be prevented (or minimized) through use of an 

impactor [66 Fed. Reg. 5706 (2001)].

Results of two other comparisons [Chow et al. 2001; Norris et al. 1999] between 

NIOSH 5040 and the IMPROVE method have been reported. In one [Chow et al. 

2001], the NIOSH 5040 EC was typically less than half the IMPROVE EC; however, a 

Sunset Laboratory instrument was not used. Instead, samples were analyzed with a 

DRI instrument, by running two different thermal programs [Chow et al. 2001]. 

Because differences in instrument design affect the OC-EC results to varying degrees, 

depending on sample type, the results reported for NIOSH 5040 (emulated on a DRI 

instrument) may not be representative of those obtained with a Sunset Laboratory 

instrument. Adjustment of the temperature program, according to another method’s 

specifications, does not necessarily produce the same results as that method. For 

example, lower EC results [Birch 1998a] were obtained with a Sunset Laboratory 

instrument when a sucrose standard was analyzed according to the temperature 

program specified for a method used in Europe. No pyrolysis correction was made, 

and the total analysis time was shorter, yet the EC result was much lower (about 3 μg 

with the Sunset instrument and 11 μg with a different instrument).

When the NIOSH 5040 thermal program was used on a DRI instrument, the filter 

transmittance reportedly exceeded its initial value before the addition of oxygen [Chow 

et al. 2001]. In an audit by a CARB (California Air Resources Board) laboratory, the 

same problem was seen with a DRI instrument, but not a Sunset Laboratory 

instrument [EPA 2003]. Char loss was extensive when the NIOSH 5040 program was 

used on a DRI instrument. Again, the filter transmittance exceeded its initial value in 

helium, but the split point was not assigned until after oxygen was introduced. These 

results contrast with those obtained by our laboratory and others, and such behavior 

was not seen in another comparison [Norris et al. 1999]; however, that comparison 

(and the CARB audit) was a direct comparison of the two methods (i.e., samples were 

analyzed with both Sunset and DRI instruments). Only 2 of 52 samples analyzed by 

Sunset Laboratory showed an increase in transmittance before the addition of oxygen, 

and the OC-EC split was near the point where oxygen was added [Norris et al. 1999]. 

Both samples had low carbon loadings (OC = 4.0 μg/cm2

, EC = 0.6 μg/cm2, 
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OC = 3.3 μg/cm2, EC = 0.5 μg/cm2) and were thought to be wood-smoke dominated. 

As pyrolysis was evident, the increase in filter transmittance was attributed to char loss. 

Comparable EC results would have been obtained if the char had not been removed 

until after oxygen was added, because char is assigned to the OC fraction. In the 

NIOSH 5040 analysis of organic compounds (e.g., sucrose), partial char loss in helium 

sometimes occurs. The varying degree of loss may relate to differences in filter purity.

4 Occupational exposure criteria (U.S.)

In 1995, the American Conference of Governmental Industrial Hygienists (ACGIH) proposed 

a Threshold Limit Value (TLV®) for diesel exhaust [ACGIH 1995] (see Notice of Intended 

Changes for 1995–1996). A TLV of 150 μg of submicrometer particulate matter (mass) per 

cubic meter of air was proposed. Four years later, a value of 50 μg/m3 was proposed [ACGIH 

1999]. An EC-based standard was later proposed, because EC is a demonstrated exposure 

marker and can be accurately quantified at low levels [ACGIH 2001a]. A TLV-TWA (timeweighted average) of 20 μg EC per cubic meter of air was recommended [ACGIH 2001b]. By 

comparison, the proposed TLV was high relative to an environmental standard supported by 

the California Office of Environmental Health Hazard Assessment (OEHHA). The California 

OEHHA classified particulate emissions from diesel-fueled engines as a toxic air contaminant 

(TAC) and supported 5 μg/m3 as the chronic inhalation reference exposure level (REL) 

[CalEPA 1998]. Because EC is a fraction of the diesel particulate emissions, the difference 

between the two air standards is greater than four fold, with the magnitude of the difference 

being dependent on the EC fraction of the diesel particulate mass. For example, if EC 

constituted 40% of the mass, the proposed TLV would be ten times higher than the OEHHA 

REL.

 

An exposure limit for DPM has been promulgated for metal and nonmetal mines, but there 

currently are no limits for other occupational settings (ACGIH withdrew a proposed limit 

[ACGIH 2003] for DPM [as EC] from its Notice of Intended Changes [NIC] list in 2003 and 

placed it under study). An EC (or TC) standard, rather than submicrometer particulate mass, 

would simplify interpretation of the analytical results because the target analyte and exposure 

standard would be the same. This would eliminate the problem of extrapolation (to 

submicrometer mass), which is not straightforward and introduces unacceptable error. As 

discussed previously, OC interferences are a problem with TC measurements; variable EC 

content is a problem when measuring EC. If EC concentrations are high and samples are 

relatively free of OC contaminates, the TC concentration is a reasonable measure of the diesel 

particulate mass, and the EC:TC ratio is representative of the EC fraction of the mass. 

However, this situation is uncommon outside of mines. In general industry, EC and OC levels 

are normally much lower. And when EC levels are low, the EC:TC ratios are not reliable 

estimates of the EC fraction of DPM because OC interferences can skew the ratios low and 
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Monitoring Diesel Exhaust in the Workplace

increase variability in the apparent ratio. In such cases, TC is an inaccurate measure of the 

diesel particulate concentration.

Exposures in the mining industry are of particular concern because diesel particulate 

concentrations in mines [63 Fed. Reg. 17491 (1998); 63 Fed. Reg. 58103 (1998); 66 Fed. Reg. 

5706 (2001)] have sometimes exceeded 2 mg/m3

, which is one thousand times higher than a 

typical environmental level. In its final rule [66 Fed. Reg. 5706 (2001)] on metal and nonmetal 

mines, MSHA proposed an interim exposure standard of 400 μg TC per cubic meter of air. 

Five years after publication of the rule, a final standard of 150 μg TC per cubic meter was to 

apply. In response to a legal challenge, MSHA initiated a limited new rulemaking [67 Fed. 

Reg. 60199 (2002)] to revise certain provisions of the final rule. Among other amendments, 

MSHA agreed to propose a change of exposure surrogate from TC to EC. Comments on 

appropriate interim and final limits for EC were requested in the new rulemaking. MSHA 

exposure standards consider health risks and technical and economic feasibility, based on 

available engineering controls and their cost.

5 Summary

NIOSH 5040 is based on a thermal-optical analysis technique for particulate carbon. Its 

intended application is assessment of workplace exposure to particulate diesel exhaust, but 

thermal-optical analysis has been routinely applied to environmental carbonaceous aerosols. 

Both OC and EC (and TC as OC + EC) are determined by the method. For reasons discussed 

in this chapter, EC was selected as an exposure surrogate for diesel exhaust. Over 30 

epidemiological studies are consistent in finding a positive association between exposure to 

diesel exhaust and lung cancer, but many studies lack quantitative exposure data. Accurate 

monitoring methods are necessary for quantifying the exposure risks.

The thermal-optical instrument incorporates an optical feature that corrects for the char 

formed during the analysis of some materials. The EC results of purely thermal methods show 

positive bias when samples contain such materials. Although minimal charring has been 

observed with most diesel soot samples from mines, workplace (and environmental) air often 

contains components that carbonize. Depending on the workplace and thermal protocol, 

these components can be significant contributors to the measured EC. For this reason, an 

analytical protocol that maximizes removal of OC components (and carbonates) and corrects 

for the char formed through thermal decomposition is important. In the NIOSH 5040 

analysis, the transmittance/reflectance of some samples continues to decrease as the 

temperature is stepped to 850 °C (and higher) in helium, which indicates incomplete 

carbonization at the lower temperatures used in some methods. Partial char loss may occur in 

helium at higher temperature, but removal of original EC is less common. Good 

interlaboratory agreement between OC-EC results has been obtained by laboratories using 
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NIOSH 5040. Because a certified reference material is not currently available, confirmation of 

results by a second laboratory using NIOSH 5040 is advised, particularly when samples char 

during the analysis because interlaboratory results for these types of samples are typically 

more variable. Organic materials that char are useful as quality control samples. Accurate 

pyrolysis correction was obtained in the NIOSH 5040 analysis of an alginic acid solution 

spiked onto blank filters and diesel soot samples. Sucrose has traditionally been used as a 

check on the method’s char correction, but other materials are needed that are more 

representative of the complex components in air samples.

To ensure data quality, participation in proficiency testing among laboratories involved in 

major studies also is advised. Several commercial laboratories offer the NIOSH 5040 analysis, 

and many instruments have been used globally for environmental and occupational 

monitoring. Method standardization is critical if results obtained by different laboratories are 

to be compared. Interlaboratory studies are useful in exposing differences among methods, 

but standards need to be included to identify method biases. Results of such comparisons 

must be interpreted with a clear understanding of each method’s limitations.

 

Continued research is needed to better assess the potential health effects of diesel exhaust and 

other types of fine/ultrafine particle pollution. Although the organic compounds associated 

with DPM have potential health effects, those traditionally measured are not unique to diesel 

exhaust. Nevertheless, characterization of this fraction may be useful in assessment of 

exposure risks, particularly if compounds enriched in diesel relative to other particulate 

emissions can be used as indicators of mutagenic potency (e.g., specific nitro-PAH). This 

information also may identify engine types and operating conditions that produce higher 

emission rates of genotoxic compounds.

As reasoned previously [Birch and Cary 1996a], regardless of whether the potential adverse 

health effects of diesel particles are due to their tiny, carbonaceous cores, adsorbed 

compounds, or a combination of both, monitoring and control of the particulate component 

are necessary if the effects are particle related. In most workplaces, diesel engines are the 

primary source of fine-particle EC. Other combustion sources may contribute to 

environmental levels of EC. These sources may be relevant from an emission control 

perspective, but if the potential toxicity of the particles is similar, their origin is not relevant 

from a health perspective.
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Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not 

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.

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NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Analysis of Carbon Nanotubes 

and Nanofibers on Mixed 

Cellulose Ester Filters by 

Transmission Electron 

Microscopy 

by M. Eileen Birch, Chen Wang, Joseph E. Fernback, H. Amy Feng, Quinn T. Birch, and Alan K. Dozier, 

NIOSH

1 Introduction CN- 2

2 Sample Preparation CN- 2

3 TEM Analysis and Counting Method CN- 6

4 Conclusions CN-18

5 References CN-18
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

1 Introduction

A transmission electron microscopy (TEM) method for the analysis of filter samples of 

carbon nanotubes and nanofibers (CNT and CNF) is described in this Chapter. The 

approach is a modification to NIOSH NMAM 7402, asbestos by TEM [NIOSH 1994], and 

was applied to personal breathing zone (PBZ) and area samples collected at CNT/CNF 

survey sites [Birch et al. 2011; Dahm et al. 2015]. Procedures specific to asbestos fiber 

identification and counting were eliminated from 7402. In contrast to asbestos, CNT 

materials used commercially are typically in the form of complex agglomerates, with 

highly entangled, curved tubes. This curvature can sometimes make even individual 

fibers difficult to size. As such, a modified method that considers these structural 

differences was needed for particle classification. 

TEM provides visualization of airborne CNT and CNF particles, indicating their size,

shape, and agglomeration state [Birch et al. 2016]. These materials have distinct features 

(e.g., graphene layers, cylindrical structures, and hollow cores) that distinguish them 

from other types of particles. Given the considerable differences between CNT/CNF 

particles and asbestos fibers, particle counting methods necessarily differ. In the case of 

CNT/CNF, all particles, agglomerates as well as single fibers, are counted as a CNT/CNF

‘structure.’ Unlike Method 7402, no minimum size cutoffs have been used to date 

because of a lack of knowledge on lung deposition and toxicological endpoints. 

Based on the number of CNT/CNF structures counted, and the collected air volume, a 

CNT/CNF concentration (i.e., structures/cm3) can be calculated. For samples collected at 

the survey sites, a significant positive correlation between structure counts and 

elemental carbon (EC) data was found, but there was considerable scatter in the data 

[Dahm et al. 2013, Dahm et al. 2015]. Given the method limitations discussed herein, the 

calculated values for structure counts are considered semi-quantitative indicators of air 

concentration.

2 Sample preparation 

For the field surveys on CNT/CNF, open-face, 25-mm, three-piece cassettes were used 

for air sampling, approximating the inhalable fraction [Dahm et al. 2013, Dahm et al. 

2015]. As specified in NMAM 7402, air samples were collected on mixed cellulose ester 

(MCE) filters having a nominal pore size of 0.8 μm (7402 specifies 0.45 to 1.2 μm), with 

sampling pumps operated at 5 L/min (0.5 to 16 L/min specified in 7402). Polycarbonate 

(PC) membrane filters also may be acceptable, but this media was not used in the field 

studies, and the filter preparation method differs from that described herein. 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

The sampling period depends on the air concentration, and it is important to avoid filter 

overload. Some trial and error may be necessary. In general, if a filter deposit is clearly 

visible, the sample is likely overloaded. An estimate of structure counts may still be 

possible, but overloading results in particle deposition on previously deposited material 

and may preclude an accurate structure count. 

Three TEM grids per field sample, and one grid each for field and media blanks (when 

provided), were prepared using procedures outlined in NMAM 7402, with the following 

modifications: 

 A scalpel is used to remove a wedge-shaped portion from the filter. Do not use a cork 

borer to remove filter sections. 

 Only one sample wedge should be placed on each slide. Do not affix the section to the 

slide with any type or adhesive, as specified in 7402. (Note: After carbon coating the 

wedge and removing small portions for transfer to TEM grids, the remaining portion 

can be secured [e.g., with tape] to the slide for long term storage, if desired.)

 The “hot block” technique [Baron and Pickford 1986] is used to clear (collapse) the 

filter wedge. Other methods also may be suitable.

A description of the procedures used for sample and grid preparation is provided in the 

following three sections. 

a. Filter 

Carefully open the sampling cassette with a cassette opener, with the particle 

collection side facing upwards (Figure 1, step 1). Remove about one fourth of the 25-

mm MCE filter with a clean scalpel. Place the wedge, particle side up, on a frosted 

glass microscope slide labelled with the sequence and sample numbers (Figure 1, 

step 2). Include a media (filter) blank with each set of slides to be coated. Use the 

“hot block” technique [Baron and Pickford 1986] to clear (collapse) the filter wedge 

(Figure 1, step 3). After clearing the filter, place the slides in a carbon coater to apply 

a thin carbon film (Figure 1, step 4); the filter surface should be evenly coated. After 

coating the filter wedges, the samples are ready for transfer to TEM grids. A Jaffe 

wick washer [Jaffe 1948] is used for final processing of the sample grids. Preparation 

of the washer and transfer of the sample sections to grids therein, for final 

processing, are described in the next two sections. 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

Figure 1. Filter preparation steps prior to loading sample portions on TEM grids: (1) 

remove 25-mm MCE filter from air sampling cassette; (2) remove about one fourth of 

the filter and place the filter wedge on a glass slide; (3) clear the MCE filter with 

acetone vapor in a hot block; (4) apply carbon coating; (5) cut section from coated 

filter and divide into three portions, one per grid. Each filter portion has particles 

sandwiched between the carbon film and collapsed filter. 

 

 

b. Jaffe Wick washer

Prepare a Jaffe wick washer using a petri dish (15-mm depth recommended, but 

other sizes may be suitable), stainless steel screen (20 mesh), and lens tissue (Figure

2, a) or a stainless steel screen disk (40 mesh) (Figure 2, b) as a wicking substrate 

(one disk per sample). Bend two edges of the 20-mesh screen to form a platform 

(Figure 2) that fits inside the petri dish and is no higher than half the dish height. The 

platform supports the tissue (Figure 2, a) or stainless disk (Figure 2, b) onto which 

the sample grids are placed and keeps them above the solvent level during sample 

processing (Figure 2). 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

Figure 2. Sample transfer and final processing: place a piece of lens tissue (a) or a 

stainless steel screen disk (b) on a stainless screen platform in a Jaffe wick washer 

filled with dimethylformamide (DMF). Sides of the platform are bent to keep sample 

grids above the solvent level. Depending on size, multiple disks can be used to hold 

grids. 

Use a pencil to label the lens tissue with the sample identifiers and place the tissue 

on the 20-mesh screen platform (Figure 2, a). The samples placed on the tissue 

should be the same set that was coated (together) in the carbon coater. Place the 

petri dish in a fume hood and carefully add dimethylformamide (DMF) to the dish 

until the lens tissue is saturated or screen disks are wetted, but ensure that no 

puddles form on surfaces of the tissue or disk. A combination of DMF and acetone 

also may be used for filter dissolution. (Note!: DMF [CAS 68-12-2] is flammable and 

hazardous in case of skin contact, eye contact, ingestion, and inhalation. Ensure 

adequate ventilation, proper personal protective equipment, and engineering 

controls in handling and storage. Consult SDS prior to use.) In a row, place three TEM 

grids per sample in the designated areas of the tissue or on a disk (Figure 2). If 

screen disks are used, place three TEM grids per sample on each (Figure 2, b). 

Depending on disk size, multiple disks can be used. Use one TEM grid each for the 

blanks (field and media). 

c. Sample transfer and final processing 

Using a scalpel and tweezers, cut and remove three small sections of the coated filter 

wedge as shown in Figure 1, step 5, starting at the center area (bottom left of wedge) 

of the filter and moving outward, across the bottom, towards the filter edge. The area 

of the cut sections should be similar and enough to cover a grid. Carbon side up, 

carefully place the excised filter sections on the three grids. If lens tissue is used, 
NIOSH Manual of Analytical Methods  5th Edition  Chapter CN June 2017 Page CN-6 of CN-19

Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

place the section from the center of the filter (Figure1, Step 5, A) on the leftmost grid, 

the middle section (Figure1, Step 5, B) on the center grid, and the outermost section 

(Figure1, Step 5, C) on the rightmost grid. The locations are labeled as shown in 

Figure 1, step 5. When all filter sections have been transferred to the grids, slowly 

add more solvent to raise the level as high as possible without disturbing the sample 

preparations. Cover the petri dish, if lens tissue is used, and elevate one side of the 

dish by placing a microscope slide under it (allowing drops of condensed solvent to 

form near the edge of the cover rather than in the center, where they can drip on the 

grid preparation). To dissolve the filter, the tissue method requires the grids to sit 

overnight, while the screen disk method requires only about 30 minutes. If screen 

disks are used, leave the petri dish uncovered, in a fume hood or other well 

ventilated area. Once the residual MCE polymer is completely dissolved in the 

solvent, only particles embedded in the carbon film remain on the grid, as illustrated 

in Figure 3. Ensure that the processed grids are solvent free prior to the TEM 

analysis.

Figure 3. Removal of residual MCE polymer from carbon-coated sample by 

dimethylformamide (DMF). After polymer removal, only particles embedded in the 

carbon film remain on the grid. 

3

a. Sample quality

.TEM analysis and counting method 

Align the microscope for TEM applications following the manufacturer’s instructions. 

Load the sample into a TEM grid holder and insert the holder into the TEM column. 

First, examine the grid at low magnification (500-1,000X) to determine the filter 

loading and preparation quality. In general, to be acceptable for analysis, the grid 

should have at least 75% intact grid openings and a particle loading less than about 

25% (area coverage). 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

If these criteria are met, randomly select at least four openings (with intact carbon 

film) from four quadrants of the grid and estimate the number of particles per 

opening to determine the distribution variability. If the particle loading is uneven or 

heavy, the sample might not be useful. Discuss any uneven distributions or loading 

issues with the requestor of the analysis. 

b. Definition of a CNT/CNF structure 

Count all particles that meet the following definition of a CNT structure:

 Any CNT or CNF individual fiber (Figure 4) with an aspect ratio (i.e., length to 

width) greater than 3:1. There are no minimum length or diameter cutoffs. If a 

fiber structure has the morphological characteristics of CNT/CNF and meets the 

3:1 aspect ratio criterion, it is counted. In our work, isolated CNT fiber or bundle 

structures were relatively rare. CNT agglomerates were far more common. 

Figure 4. Examples of CNT fiber structures with various lengths and widths. 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

 Any particle agglomerate with associated CNT/CNF: There are many types of 

structures that fit this criterion. Some structures, classified as matrix particles, 

contain relatively few CNT/CNF, being mainly amorphous carbon or other 

components. In contrast, structures that consist mainly of CNT/CNF are classified 

as cluster particles. 

In this chapter, ‘agglomerate’ refers to both cluster (Figure 5) and matrix (Figure 6) 

particles. Although matrix particles have associated CNT, they are often observed as 

fiber structures protruding from the particle edge. The structure of the CNT, 

presumably within the matrix, cannot be determined in the TEM analysis. Another 

classification problem occurs when agglomerates have cluster and matrix regions 

(Figure 7) of similar size. Structure counts are discussed in the next section. 

 Figure 5. Examples of CNT/CNF clusters: (a)-(f) collected from workplaces. 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

Figure 6. Examples of CNT matrix particles: (a)-(d) collected from workplaces. 

There are specific definitions of CNT and CNF; however, field samples have contained 

complex mixtures of structures that often do not fit these narrow definitions. For 

example, products described as CNT can contain CNF and vice versa, and a variety of 

irregularly shaped, nanoscale carbon particles have been observed. In practice, if the 

analytical request specifies a ‘CNT’ or ‘CNF’ facility, these terms are used in the 

analytical report, along with the corresponding TEM images. In addition, images and 

information on other types of particles are provided.
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c. Structure counts 

The counting method for individual fibrous structures generally follows that for 

asbestos, as in NMAM 7402 and EPA’s AHERA (Asbestos Hazard Emergency 

Response Act) method [40 CFR 763; NIOSH 1994]. That is, each fiber with an aspect 

ratio > 3:1 is counted and the diameter and length are recorded. At a minimum, the 

diameter and length ranges are recorded, and averages are determined, if 

appropriate. This procedure is negotiable with the requestor. 

As mentioned, in the field samples, isolated CNT fibers were rare relative to particle 

agglomerates. The classification of CNT/CNF agglomerates is accomplished by 

considering both particle size and structure. The size bin depends on the maximum 

crosswise dimension. At least one dimension, the maximum crosswise length, is 

recorded for each structure. Other characteristics also can be examined to meet 

investigator needs. Some agglomerates, especially larger ones, may contain a nonCNT/CNF component. In these instances, if the agglomerate looks to be mostly 

CNT/CNF, it is classified as a cluster. If non-CNT/CNF components appear more 

abundant, the matrix classification is used. However, as discussed, some 

agglomerates may have roughly equal cluster and matrix portions. For these, a 

matrix category is usually assigned but this classification is somewhat arbitrary. 

Given the difficulties of particle classification, it is important to provide 

representative images of all particles in the sample. 

If the particle loading and distribution are acceptable, an initial particle count can be 

performed at low magnification. Select the lowest magnification that shows the 

majority of the particulate of interest: 5,000-15,000X is usually sufficient. Smaller 

particles may not be visible at low magnification, but the purpose of this step is to 

assess the number of grid openings likely to be counted. If the sample composition is 

known (e.g., a laboratory generated CNT or CNF sample), an initial count can be 

made to count larger (micrometer sized) CNT (CNF) particles, which dominate the 

particle mass. After examination of the sample at low magnification, a switch to high 

magnification (e.g., 40,000X) is made to image particles/features not observable at 

low magnification. The magnification should be adjusted as required for counting 

and confirming the presence of CNT. 

As discussed, for the analysis, three 3-mm copper TEM grids are usually prepared 

from each sample. A total of 40 openings (stop counting at 100 structures) are 

examined at high magnification (13-15 openings for each grid). The number of grid 

openings counted per grid should be approximately equal for the three grid 

preparations, and the openings counted on each grid should be selected as randomly 

as possible. Only grid openings that have the carbon film intact should be counted. 

Any particles containing CNT/CNF are counted as CNT or CNF structures, which can 

range from single fibers (Figure 4) to structures of varying size and composition 

(Figs. 5-9). Counted CNT/CNF structures are categorized by size-bins based on the 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

maximum crosswise dimension. In our studies [Dahm et al. 2015; Birch et al. 2016], 

most of the CNT agglomerates had a roughly spherical shape (i.e., the longest 

dimension was no more than twice the length of the orthogonal dimension). 

Therefore, the maximum crosswise dimension was used to categorize structure size. 

The counted structures were placed into five discrete size-bins, based on healthrelevant size fractions and observations from field studies [Birch 2011; Dahm et al. 

2013]. The structure categories also included a separate bin for single fibers. Size 

bins for the CNT/CNF structures have the following maximum crosswise dimensions: 

< 1 μm, 1-2 μm, 2-5 μm, 5-10 μm, and > 10 μm. This sizing scheme was used to 

analyze CNT/CNF samples collected during field surveys [Dahm et al. 2015]. A 

statistical analysis of the TEM counts for a subset of the field samples is provided in 

the following section. 

Figure 7. CNT particle with both matrix (right side) and cluster (left) features. 

Structure categorized as a CNT matrix particle. 
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Figure 8. CNT particles in personal breathing zone samples collected from various 

sites: (a) MWCNT fiber (polymer composite site), b) SWCNT agglomerate from 

aerosolized, aqueous suspension (electronics site), c) MWCNT agglomerate 

(composites/thermoplastics site), and d) MWCNT agglomerate (primary 

manufacturer of CNT). 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

Figure 9. Examples of CNT particles embedded in polymer composite: (a)-(d). 

d. Statistics 

A total of 3000 observations (structures/opening) from 75 field samples with light 

loading (<5 structures per grid opening) were analyzed. The mean, standard 

deviation, and confidence intervals (95% confidence level) were calculated. Factors 

considered included grid (1-3), opening (1-14), and loading level (1-3). Grids were 

counted in the order of grid 1, 2, and then 3, going from the center to the edge of the 

filter (wedge portion). From Figure 10 and Table 1, the means and variabilities of 

counts from different loading levels follow the same trend. The variability of grid 1 

(from center area of filter) was slightly higher than the other 2 grids. The variability 

becomes smaller when the loading level is increased. The box plot in Figure 11 shows 

the statistics of counts by openings. The means and variabilities of counts from 

opening 1 and 2 were slightly higher than counts from opening 12, 13 and 14. 

Because opening 1 was the first opening counted, and 14 the last, it is possible that 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

the increased variability is due to higher analyst error during the initial and final 

counts. The test of half data (including openings 1-7) shows similar behavior as that 

for all data (Figure 12). Based on the results for this sample set, counts from one or 

two grids, with 7 openings per grid, may be adequate if the deposit is even. Larger 

data sets from samples with different loadings and collected from various sites are 

required to develop a statistical model for further analysis. 

Figure 10. Boxplot of CNT counts by categorized loading (1: mean ≤ 1, 2: 1 < mean ≤ 

2, and 3: mean > 2) from three grids. The trend of mean counts from grid 1 to 3 is 

consistent for different loading levels. Mean count of grid 1 is slightly higher than 

those of grid 2 and 3. 

 
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Table 1. Statistics of sample counts obtained from different grids and loading levels 

(M=Mean, SD=Standard Deviation, CI=Confidence Intervals).

Loadinga 

 Grid 1b Grid 2 b Grid 3 b

M (SD) 95% CI M (SD) 95% CI M (SD) 95% CI

Level 1 0.51 (0.84) [0.44, 0.58] 0.44 (0.64) [0.38, 0.49] 0.43 (0.66) [0.37, 0.49] 

Level 2 1.45 (1.12) [1.29, 1.60] 1.33 (0.91) [1.21, 1.46] 1.35 (0.90) [1.23, 1.48] 

Level 3 2.65 (1.32) [2.50, 2.80] 2.64 (1.10) [2.51, 2.77] 2.55 (1.07) [2.43, 2.68] 

All 1.33 (1.41) [1.24, 1.41] 1.26 (1.28) [1.18, 1.34] 1.24 (1.25) [1.16, 1.32] 

a Loading levels are defined by M, where Level 1 (M ≤ 1), Level 2 (1 < M ≤ 2) and Level 3 (M > 2). 

b Grids collected from different locations on a filter wedge (center to edge).

 

Figure 11. Boxplot of CNT counts by openings (1-14). The variability in openings 1 

and 2 significantly higher than that of openings 13 and 14. The means of openings 1, 2 

and 3 were also higher than the means of openings 12, 13, and 14. 
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

Figure 12. Boxplot of CNT counts by grids (1-3) from all data (counts from openings1-

14) and half data (counts from openings 1-7). 

 
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e. Limit of detection (LOD)

For the field samples [Dahm et al. 2015], the ‘no structures detected’ TEM samples

were those with counts below the LOD, defined as 1 CNT or CNF structure per

sample. The corresponding unit of structures per cm3 LOD (air concentration) was

then calculated (by the requestor) based on the total air volume sampled. 

f. CNT/CNF polymer composites

Dusts generated from CNT/CNF polymer composites that have been mechanically

worked (e.g., drilled, cut, sanded, blasted, etc.) contain particles with CNT/CNF

embedded in the polymer matrix. Our analyses of polymer composite dusts have 

shown little evidence of individual fiber release from the composite matrix, but 

respirable sized matrix particles with protruding CNT/CNF have been observed 

(Figure 9). These findings are consistent with other reports [Bello et al. 2012; 

Kingston et al. 2014] on potential CNT/CNF release from polymer matrices. Particle 

counts for these types of samples also can be performed, though only a limited 

number of surveys have been conducted. Because the CNT/CNF are embedded in a 

polymer matrix, the toxicity of these dusts may differ substantially from dusts with

free (unbound) agglomerates. Nevertheless, representative images of the particles

and information on particle shape and size can be documented.

g. Limitations

The composition of CNT powders varies substantially, especially those produced or

used in workplaces. These materials contain varying amounts of amorphous carbon 

and residual catalyst metals, and there are many types of structures, usually in

agglomerate form. As mentioned, in the case of composite dusts, CNT are embedded

in a polymer matrix, which complicates their measurement. This chapter concerns

the analysis of CNT powders.

In the case of ‘matrix’ particles, TEM images show mainly dark regions, with varying

amounts of CNT observed at the particle’s edge. The composition of material in the 

dark regions is unknown. For example, it may be mainly CNT, amorphous carbon, or

a combination of both. Thus, based on overall envelope size, a particle consisting

mainly of amorphous carbon is counted in the same bin as an agglomerate composed

mainly of CNT. Both matrix and cluster (structures with mainly CNT) type particles 

were common in field samples. A different counting method [Chen et al. 2012] was 

applied to a specific type of MWCNT (Mitsui MWCNT-7) used in an animal inhalation 

study but this particular product, having a more fibrous structure and more single 

fibers, differs from the CNT materials manufactured/used by U.S. companies. Images

(Figs. 4-9) of the different particle types found in our field surveys make obvious the 

difficulty in developing a standardized TEM structure count method for these 

materials.
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Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

4. Conclusions

The complexity and variety of structures makes CNT/CNF particle counting a challenge.

Unlike elemental carbon or other less selective measures, TEM can confirm the presence 

of CNT. However, with respect to quantification, it is important to recognize the 

limitations of a counting method. Because of the heterogeneity of CNT powders,

particles in the same size bin can vary greatly in their properties. Complex structures of

varying mass, density, shape, and composition (e.g., amorphous carbon, residual

catalyst) can fall into the same size bin. And dissimilar particles (e.g., cluster and matrix)

in a given bin may have different toxicological effects. Thus, even for a given material,

particle envelope size alone may not be an adequate risk indicator. Given the 

heterogeneity and variety of the CNT/CNF products to which workers are exposed, it is

important to apply several methods to assess worker exposure, through inhalation and

other routes.

Disclaimer 

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the

sponsoring organizations or their programs or products. Furthermore, NIOSH is not

responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.

5. References

Baron PA, Pickford GC[1986]. An Asbestos Sample Filter Clearing Procedure. Appl Ind 

Hyg 1(4):169-171. 

Bello D, Wardle BL, Yamamoto N, deVilloria RG, Hallock M[2011]. Exposures to 

nanoscale particles and fibers during handling, processing, and machining of 

nanocomposites and nanoengineered composites reinforced with aligned carbon 

nanotubes. Proceedings of the 17th International conference on composite materials 

(ICCM), Edinburgh, Scotland, July 27–31, 2011. 

Birch ME [2011]. Exposure and emissions monitoring during carbon nanofiber 

production—Part II: Polycyclic Aromatic Hydrocarbons. Ann Occup Hyg 

55(9):1037-1047. 

Birch ME, Ruda-Eberenz TA, Chai M, Andrews R, Hatfield RL [2013]. Properties that 

Influence the Specific Surface Areas of Carbon Nanotubes and Nanofibers. Ann Occup 

Hyg 57(9):1148-1166. 

Birch ME, Wang C, Fernback JE, Feng HA, Birch QT, Dozier AK [2016]. Workplace 

Monitoring of Airborne Carbon Nanomaterials by HRTEM. Microsc Microanal 22(S3): 

1910-1911. 
NIOSH Manual of Analytical Methods  5th Edition  Chapter CN June 2017 Page CN-19 of CN-19

Analysis of Carbon Nanotubes and Nanofibers on MCE Filters by TEM 

Chen BT, Schwegler-Berry D, McKinney W, Stone S, Cumpston JL, Friend S, Porter DW, 

Castranova V, Frazer DG [2012]. Multi-walled Carbon Nanotubes: Sampling Criteria and 

Aerosol Characterization. Inhalation Toxicology24(12):798–820. 

CFR. Code of Federal Regulations. Washington, DC: U.S. Government Printing Office, 

Office of the Federal Register. 

Dahm MM, Evans DE, Schubauer-Berigan MK, Birch ME, Fernback JE [2012]. 

Occupational Exposure Assessment in Carbon Nanotube and Nanofiber Primary and 

Secondary Manufacturers. Ann Occup Hyg56(5):542-556. 

Dahm MM, Evans DE, Schubauer-Berigan MK, Birch ME, Deddens JA [2013]. 

Occupational Exposure Assessment in Carbon Nanotube and Nanofiber Primary and 

Secondary Manufacturers: Mobile Direct-Reading Sampling. Ann Occup Hyg 

57(3):328-344. 

Dahm MM, Schubauer-Berigan MK, Evans DE, Birch ME, Fernback JE, Deddens JA 

[2015]. Carbon Nanotube and Nanofiber Exposure Assessments: An Analysis of 14 Site 

Visits. Ann Occup Hyg 59(6):705-723. 

Jaffe MS [1948]. Handling and washing of fragile replicas. J. Appl. Physics19:1189-1191. 

Kingston C, Zepp R, Andradyc A, Boverhofd D, Fehire R, Hawkins D, Roberts J, Sayreg P, 

Shelton B, Sultani Y, Vejinsj V, Wohlleben W [2014], Release Characteristics of Selected 

Carbon Nanotube Polymer Composites. Carbon68:33-57. 

NIOSH [1994]. Method 7402 asbestos by TEM. In: Eller PM, Cassinelli ME, eds. NIOSH 

Manual of Analytical Methods. 4th ed. Cincinnati, OH: U.S. Department of Health and 

Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 

[www.cdc.gov/niosh/nmam/]. 
NIOSH Manual of Analytical Methods (NMAM), 5th Edition

Glossary of Abbreviations, 

Definitions and Symbols

Compiled by Kevin Ashley, Ph.D., NIOSH 

A GL-2

B GL-4

C GL-7

D GL-9

E GL-12

F GL-13

G GL-13

H GL-14

I GL-14

L GL-15

M GL-16

N GL-17

O GL-17

P GL-18

Q GL-20

R GL-20

S GL-21

T GL-23

U GL- 24

V GL-25

W GL- 26

X GL- 26
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Glossary of Abbreviations, Definitions and Symbols

Terms & Definitions

A 

AAS - Atomic absorption spectrometry

Absorption Barrier - Any exposure surface that may retard the rate of penetration of an

agent into a target. Examples of absorption barriers are the skin, respiratory tract lining,

and gastrointestinal tract wall (cf. exposure surface). [Source: Zartarian V, Bahadori T,

McKone T [2005]: Adoption of an official ISEA glossary. J Expo Anal Environ Epidemiol

15:1–5.] 

Acceptable Range (biological) - The range of values of a biological monitoring analyte

that would be expected in workers with exposure to the chemical agent in the workplace

at or below regulatory or recommended levels. These ranges are often method-specific.

[Adapted from: NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. 

DHHS (NIOSH) Publication No. 94-113.] 

Accuracy – 

1. The degree of agreement between a measured value and the accepted reference value.

In this manual, accuracy is calculated from the absolute mean bias of the method plus

the overall precision, SS�rrrr at the 95% confidence level. For an individual measurement,

it includes the combination of precision and bias [Source: NIOSH [1977]:

Documentation of the NIOSH Validation Tests. DHEW (NIOSH) Publication No. 77-

185.]

2. Measure of confidence in a measurement. It is a qualitative term referring to whether

there is agreement between a measurement made on an object and its true (target or

reference) value. [Source: NIST/SEMATECH e-Handbook of Statistical Methods;

Gaithersburg, MD: National Institute of Standards and Technology,

http://www.itl.nist.gov/div898/handbook/]

3. The ability of a method to determine the “true” concentration of the environment

sampled. Accuracy describes the closeness of a typical measurement to the quantity

measured although it is defined and expressed in terms of the relative discrepancy of a

typical measurement from the quantity measured. The special sense of accuracy for a

method is embodied in the following definition and criterion: The accuracy of a

method is the theoretical maximum error of measurement, expressed as the

proportion or percentage of the amount being measured without regard for the

direction of the error that is achieved with 0.9 probability by the method. [Source:

NIOSH [1995]: Guidelines for air sampling and analytical method development and
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Glossary of Abbreviations, Definitions and Symbols

evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. DHHS

(NIOSH) Publication No. 95-117, http://www.cdc.gov/niosh/docs/95-117/pdfs/95-

117.pdf.]

4. The degree of conformity of a value generated by a specific procedure to the assumed

or accepted true value. It includes both precision and bias. [Source: ASTM [2014].

D1356, Standard terminology relating to sampling and analysis of atmospheres. West

Conshohocken, PA: ASTM International]

ACGIH - American Conference of Governmental Industrial Hygienists

Acute Exposure - A contact between an agent and a target occurring over a short time, 

generally less than a day. Note: Other terms, such as “short-term exposure” and “single dose,” 

are also used. [Source: Zartarian V, Bahadori T, McKone T [2005]: Adoption of an official 

ISEA glossary. J Expo Anal Environ Epidemiol 15:1–5.] 

Aerosol – 

1. Airborne particles and the gas (and vapor) mixture in which they are suspended.

Note: The airborne particles can be in or out of equilibrium with their own vapors.

[Source: CEN [2011]. EN 1540, Workplace atmospheres – terminology. Brussels:

European Standards Commission.]

2. Dispersion of solid or liquid particles in a gaseous medium. [Source: ASTM [2014].

D1356, Standard terminology relating to sampling and analysis of atmospheres. West

Conshohocken, PA: ASTM International.]

Agent - A chemical, biological, or physical entity that contacts a target. [Source: Zartarian V, 

Bahadori T, McKone T [2005]: Adoption of an official ISEA glossary. J Expo Anal Environ 

Epidemiol 15:1–5.] 

Analyte – 

1. Substance or chemical constituent that is determined in an analytical method [Source:

CEN [2011]. EN 1540, Workplace atmospheres – terminology. Brussels: European

Standards Commission.]

2. A specific chemical moiety being measured, which can be intact drug, biomolecule or

its derivative, metabolite, and/or degradation product in a biologic matrix. [Source:

FDA [2001]. Guidance for industry - bioanalytical method validation,

http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf.]
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Glossary of Abbreviations, Definitions and Symbols

Ashing - The decomposition, prior to analysis, of organic matrix constituents of the sample 

and sampler. The most common ashing techniques are solvent, acid, or alkali dissolution; 

alkaline fusion; and oxidation using either low-temperature oxygen plasma or muffle furnace. 

[Source: NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS 

(NIOSH) Publication No. 94-113.] 

ASV - Anodic stripping voltammetry

Atmospheric Concentration - The quantity of a constituent substance per unit volume of 

air [Adapted from definition of ‘concentration’ in: ASTM [2014]. D1356, Standard 

terminology relating to sampling and analysis of atmospheres. West Conshohocken, PA: 

ASTM International.] 

Atmospheric Deposition - The transfer of an atmospheric constituent to a surface due to 

gravity or another mechanism, or the material which is transferred [Adapted from definition 

of ‘deposition’ in: ASTM [2014]. D1356, Standard terminology relating to sampling and 

analysis of atmospheres. West Conshohocken, PA: ASTM International.] 

Atmospheric Dispersion - The most general term for a system consisting of a constituent 

suspended in air [Adapted from definition of ‘dispersion’: ASTM [2014]. D1356, Standard 

terminology relating to sampling and analysis of atmospheres. West Conshohocken, PA: 

ASTM International.] 

AW - Atomic weight

B 

B - Media blank result for a single-section sampler (e.g., sorbent tube.)

Bb - Media blank result for back section of a sampler.

Bf - Media blank result for front section of a sampler.

Background Level - The amount of an agent in a medium (e.g., water, soil) that is not 

attributed to the source(s) under investigation in an exposure assessment. Background level(s) 

can be naturally occurring or the result of human activities. (Note: Natural background is the

concentration of an agent in a medium that occurs naturally or is not the result of human 

activities.) [Source: Zartarian V, Bahadori T, McKone T [2005]: Adoption of an official ISEA 

glossary. J Expo Anal Environ Epidemiol 15:1–5.]
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Glossary of Abbreviations, Definitions and Symbols

Bias – 

1. A systematic (nonrandom) deviation of the method average value or the measured

value from an accepted value. [Source: ASTM [2014]. D1356, Standard terminology

relating to sampling and analysis of atmospheres. West Conshohocken, PA: ASTM

International.]

2. Difference between the average measured mass or concentration and reference mass

or concentration expressed as a fraction of reference mass or concentration. [Source:

NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS

(NIOSH) Publication No. 94-113.]

3. An estimate of a systematic measurement error [Source: ISO [2015]. ISO 18158

Workplace air – terminology. Geneva: International Organization for

Standardization.]

Bioaerosol - An aerosol consisting of (a) biological agent(s). Note: Airborne dusts of organic

origin, for example, cotton dust, flour dust and wood dust, are not considered to be 

bioaerosols and are therefore not covered by this definition [Source: ISO [2015]. ISO 18158

Workplace air – terminology. Geneva: International Organization for Standardization.]

Bioavailability – 

1. The rate and extent to which an agent can be absorbed by an organism and is

available for metabolism or interaction with biologically significant receptors.

Bioavailability involves both release from a medium (if present) and absorption by an

organism. [Source: Zartarian V, Bahadori T, McKone T [2005]: Adoption of an

official ISEA glossary. J Expo Anal Environ Epidemiol 15:1–5.]

2. The extent to which a chemical substance to which the body is exposed (by ingestion,

inhalation, injection, or skin contact) reaches the systemic circulation, and the rate at

which this occurs. It is recognized that the bioavailability (for gastrointestinal

absorption) of, for example, both essential and non-essential metals, depends on

various factors including the composition of the diet and the type of the chemical

compound and its state of dispersion. For instance, the absorption of lead and

cadmium is increased if the food is deficient in calcium or iron [Source: ILO/IPCS.

Glossary of terms on chemical safety (after WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]

Biological Agent - One of a number of agents such as bacteria, viruses, fungi and other 

micro-organisms or parts of them and their associated toxins, including those which have 
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Glossary of Abbreviations, Definitions and Symbols

been genetically modified, cell cultures or endoparasites which are potentially hazardous to 

human health. Note: Dusts of organic origin, for example, cotton dust, flour dust and wood 

dust, are not considered to be biological agents and are therefore not covered by this 

definition. [Source: ISO [2015]. ISO 18158 Workplace air – terminology. Geneva: 

International Organization for Standardization.] 

Biological Matrix - A discrete material of biological origin that can be sampled and processed 

in a reproducible manner. Examples are blood, serum, plasma, urine, feces, saliva, sputum, 

and various discrete tissues. [Source: FDA [2001]. Guidance for industry - bioanalytical 

method validation, http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf.] 

Biological Monitoring – 

1. The measurements of the absorption of an environmental chemical in the worker by

analysis of a biological specimen for the chemical agent, its metabolites or some

specific effect on the worker. [Source: NIOSH [1994]. NIOSH Manual of analytical

methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.]

2. The periodic examination of biological specimens (in accordance with the definition

of monitoring). It is usually applied to exposure monitoring but can also apply to

effect monitoring. [Source: ILO/IPCS. Glossary of terms on chemical safety (after

WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]

Biomarker of Effect/Response - A measurable biochemical, physiologic, behavioral, or 

other alteration in an organism that, depending on the magnitude, can be recognized as 

associated with an established or possible health impairment or disease [Source: National 

Research Council of the National Academies (NRC) [2006]. Human Biomonitoring for 

Environmental Chemicals. Washington DC: The National Academies Press, 

http://www.nap.edu/catalog/11700/human-biomonitoring-for-environmental-chemicals.] 

Biomarker of Exposure (e.g., Biological Indicator of Exposure) - A chemical, its metabolite, 

or product of an Interaction between a chemical or some target molecule or cell that is 

measured in and organism, such as humans [Source: National Research Council of the 

National Academies (NRC) [2006]. Human Biomonitoring for Environmental Chemicals. 

Washington DC: The National Academies Press, http://www.nap.edu/catalog/11700/humanbiomonitoring-for-environmental-chemicals.] 

Biomarker of Susceptibility - An indicator of an inherent or acquired ability of an organism 

to respond to exposure to a specific chemical substance. Such an indicator may be the result of 
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Glossary of Abbreviations, Definitions and Symbols

a genetic factor, nutritional status, lifestyle, or life stage that affect susceptibility to a chemical 

exposure. This kind of biomarker can be used to distinguish susceptible individuals or groups; 

for example, a cytochrome phenotype. [Source: U.S. Environmental Protection Agency (EPA) 

[2016]. Defining pesticide biomarkers, http://www.epa.gov/pesticide-science-and-assessingpesticide-risks/defining-pesticide-biomarkers.] 

Biomonitoring - A method used to assess human exposure to chemicals by measuring a 

chemical, its metabolite, or a reaction product in human tissues or specimens, such as blood 

and urine. [Source: National Research Council of the National Academies (NRC) [2006]. 

Human Biomonitoring for Environmental Chemicals. Washington DC: The National 

Academies Press, http://www.nap.edu/catalog/11700/human-biomonitoring-forenvironmental-chemicals.] 

Blank - See Field blank, Media blank, and Reagent blank.

Blank Sample - Unused collection substrate, taken from the same batch used for sampling,

processed so as to measure artifacts in the measurement (sampling and analysis) process.

[Source: CEN [2005]. EN 14902, Ambient air quality — Standard method for the measurement

of Pb, Cd, As and Ni in the PM10 fraction of suspended particulate matter. Brussels: European 

Standards Commission.] 

BP - Boiling point, °C.

Breakthrough Volume - Volume of air that can be passed through a sampler before the gas

or vapor exceeds the capacity of the sampler. [Source: ISO [2015]. ISO 18158 Workplace air – 

terminology. Geneva: International Organization for Standardization.] 

Breathing Zone - The space around a worker’s face from where he or she takes his or her 

breath. For technical purposes a more precise definition is as follows: A hemisphere of radius 

0.3 m extending in front of the human face, centered on the midpoint of a line joining the 

ears; the base of the hemisphere is a plane through this line, the top of the head and the larynx.

The definition is not applicable when respiratory protective equipment is used. [Source: CEN 

[2011]. EN 1540, Workplace atmospheres – terminology. Brussels: European Standards 

Commission.] 

C 

C – 

1. Concentration of gaseous, liquid, or solid substance in air, mg/m3;

2. Acceptable ceiling concentration (for a specified maximum time of exposure)
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Glossary of Abbreviations, Definitions and Symbols

when applied to personal permissible exposure limits.

Calibration Graph - Plot of analytical response vs. known mass or concentration of analyte.

CAS # - Chemical Abstracts Service Registry Number.

CE - Collection efficiency, expressed as a decimal fraction.

Chemical Agent - Chemical element or compound on its own or admixed as it occurs in the 

natural state or as produced, used, or released, including release as waste, by any work activity,

whether or not produced intentionally and whether or not placed on the market. [Source: ISO 

[2015]. ISO 18158 Workplace air – terminology. Geneva: International Organization for 

Standardization.] 

Chronic Exposure - A continuous or intermittent long-term contact between an agent and a 

target. (Other terms, such as "long-term exposure," are also used). [Source: Zartarian V, 

Bahadori T, McKone T [2005]. Adoption of an official ISEA glossary. J Expo Anal Environ 

Epidemiol. 15:1-5.] 

49 CFR 171-177 - Title 49 (Transportation), Code of Federal Regulations. U. S. regulations 

governing shipment of hazardous materials.

Conc. - Concentrated; concentration

Concentration – 

1. A general term referring to the quantity of a material or substance contained in unit

quantity of a given medium. [Source: ILO/IPCS. Glossary of terms on chemical safety

(after WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]

2. The quantity of a substance contained in a total unit quantity of sample. [Source:

ASTM [2014]. D1356, Standard terminology relating to sampling and analysis of

atmospheres. West Conshohocken, PA: ASTM International.]

Control (biological) - A value or group of values of a biological monitoring parameter 

collected from workers with little or no occupational exposure to the specific chemical agent. 

[Source: NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS 

(NIOSH) Publication No. 94-113.] 
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Glossary of Abbreviations, Definitions and Symbols

Cv - Concentration of gaseous substance in air, parts per million (V/V). In this manual, Cv is 

referred to NTP such that Cv = (C x 24.46)/MW.

Cumulative Exposure - The sum of exposures of an organism to a pollutant over a period of 

time. [Source: EPA [1997]: EPA Terms of Environment – Glossary of Exposure Assessment 

Related Terms: A Compilation. Prepared by the Exposure Terminology Subcommittee of the 

IPCS Exposure Assessment Planning Workgroup for the International Programme on 

Chemical Safety Harmonization of Approaches to the Assessment of Risk from Exposure to 

Chemicals, 2001.] 

CV – See SSrr : Estimate of the relative standard deviation, equal to S (sample standard 

deviation) divided by the mean of a series of measurements. A measure of precision; 

previously referred to as CV (coefficient of variation). [Source: NIOSH [1994]. NIOSH 

Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.] 

D 

D - Density, g/cm3

DE - Desorption efficiency; fraction of known quantity of analyte recovered from spiked solid 

sorbent media blank. DE may be a function of loading, and should be determined by the 

chemist for each lot of solid sorbent used for sampling, in the concentration range of interest. 

Plot (mass recovered minus average media blank)/ mass added vs. (mass recovered minus 

average media blank). [Source: NIOSH [1994]. NIOSH Manual of analytical methods 

(NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.] 

Detector - The part of the monitor that sees and/or measures and/or quantifies and/or 

ascertains the dimensions, quantity, or concentration of the gas or vapor of interest. [Source: 

NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) 

Publication No. 94-113.] 

Ds - Stokes diameter.

Detection Limit - See LOD; MDL. Lowest amount of an analyte that is detectable with a 

given confidence level. Note: The detection limit can be calculated as three times the standard 

deviation of blank measurements. This represents a probability of 50% that the analyte will 

not be detected when it is present at the concentration of the detection limit. [Source: ISO 

[2015]. ISO 18158 Workplace air – terminology. Geneva: International Organization for 

Standardization.] 
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Glossary of Abbreviations, Definitions and Symbols

Diffusive Sampler – 

1. Device which is capable of taking samples of gases or vapors from the atmosphere at a

rate controlled by a physical process such as gaseous diffusion through a static air

layer or permeation through a membrane, but which does not involve active

movement of air through the sampler. [Source: ASTM [2014]. D1356, Standard

terminology relating to sampling and analysis of atmospheres. West Conshohocken,

PA: ASTM International.]

2. Passive sampler that collects gases or vapors at a rate governed by diffusion through a

static air layer and/or permeation through a membrane. [Source: ISO [2015]. ISO

18158 Workplace air – terminology. Geneva: International Organization for

Standardization.]

Dose – 

1. The amount of agent that enters a target after crossing an exposure surface. If the

exposure surface is an absorption barrier, the dose is an absorbed dose/uptake dose

(see uptake); otherwise it is an intake dose (see intake). [Source: Zartarian V,

Bahadori T, McKone T [2005] Adoption of an official ISEA glossary. J Expo Anal

Environ Epidemiol, 15:1-5.]

2. The amount of a chemical administered to an organism. [Source: ILO/IPCS. Glossary

of terms on chemical safety (after WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]

Dosimeter - An instrument to measure dosage; many so called dosimeters actually 

measure exposure rather than dosage. [Source: EPA [1997] EPA Terms of

Environment – Glossary of Exposure Assessment Related Terms: A Compilation.

Prepared by the Exposure Terminology Subcommittee of the IPCS Exposure

Assessment Planning Workgroup for the International Programme on Chemical

Safety Harmonization of Approaches to the Assessment of Risk from Exposure to

Chemicals, 2001.] 

Dosimetry - Process or technology of measuring and/or estimating dosage. [Source:

EPA [1997] EPA Terms of Environment – Glossary of Exposure Assessment Related

Terms: A Compilation. Prepared by the Exposure Terminology Subcommittee of the 

IPCS Exposure Assessment Planning Workgroup for the International Programme on

Chemical Safety Harmonization of Approaches to the Assessment of Risk from

Exposure to Chemicals, 2001.] 
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Glossary of Abbreviations, Definitions and Symbols

E 

ECD - Electron capture detector

EPA - U.S. Environmental Protection Agency

Est - Estimated

Exposure - 

1. Contact between an agent and a target. Contact takes place at an exposure

surface over an exposure period. [Source: Zartarian V, Bahadori T, McKone T

[2005]. Adoption of an official ISEA glossary. Jour Expo Anal & Environ

Epidemiol, 15:1–5.]

2. The amount of an environmental agent that has reached the individual (external

dose) or has been absorbed into the individual (internal dose, absorbed dose).

[Source: ILO/IPCS. Glossary of terms on chemical safety (after WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/gl

ossary.htm]

3. (By inhalation) Situation in which a chemical agent or biological agent is

present in the air that is inhaled by a person. [Source: ISO [2015]. ISO 18158

Workplace air – terminology. Geneva: International Organization for

Standardization.]

Exposure Assessment - 

1. The process of estimating or measuring the magnitude, frequency and duration of

exposure to an agent, along with the number and characteristics of the population

exposed. Ideally, it describes the sources, pathways, routes, and the uncertainties in the

assessment. [Source: Zartarian V, Bahadori T, McKone T Adoption of an official

ISEA glossary. J Expo Anal Environ Epidemiol, 15:1–5.]

2. The quantification of the amount of exposure to a hazard for an individual or group

[Source: ILO/IPCS. Glossary of terms on chemical safety (after WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]
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Glossary of Abbreviations, Definitions and Symbols

Exposure Concentration - The exposure mass divided by the contact volume or the 

exposure mass divided by the mass of contact volume depending on the medium.

[Source: Zartarian V, Bahadori T, McKone T [2005]: Adoption of an official ISEA 

glossary. J Expo Anal Environ Epidemiol, 15:1–5.] 

Exposure Route - The way an agent enters a target after contact (e.g., by ingestion,

inhalation, or dermal absorption). [Source: Zartarian V, Bahadori T, McKone T [2005]: 

Adoption of an official ISEA glossary. J Expo Anal Environ Epidemiol, 15:1–5.] 

F 

F - Fiber(s)

FID - (Hydrogen-air) flame ionization detector

Field Blank – 

1. A sample (or sampler) handled exactly the same as the field samples, except no

air is drawn through it. Used to estimate contamination in preparation for

sampling, shipment and storage prior to measurement, but not actually

subtracted from sample readings (see media blank). [Source: NIOSH [1994].

NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH)

Publication No. 94-113.]

2. Blank (sample) that is transported to the sampling site, but not used for sample

collection. Discussion: A field blank is loaded in the sampler, where applicable,

and returned to the laboratory in the same way as a sample. The results from the

analysis of field blanks are used to identify contamination of the sample arising

from handling in the field and during transport. [Source: ISO [2015]. ISO 18158

Workplace air – terminology. Geneva: International Organization for

Standardization.]

FPD - Flame photometric detector

FTIR - Fourier transform infrared spectroscopy

G 

GC - Gas chromatography
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Glossary of Abbreviations, Definitions and Symbols

GFAAS - Graphite furnace atomic absorption spectrometry

GPO - U.S. Government Printing Office, Washington, DC 20402 

H 

Hemolysis - Rupture of red blood cells. [Adapted from Webster’s English dictionary online, 

http://www.merriam-webster.com/]. Discussion: hemolysis may occur due to improper 

collection and handling of whole blood samples.

HGAAS - Hydride generation atomic absorption spectrometry

Hydrolysis - Chemical process of decomposition involving the splitting of a bond and the 

addition of the hydrogen cation and the hydroxide anion of water [Source: Webster’s English

dictionary online, http://www.merriam-webster.com/] 

HPLC - High performance liquid chromatography

I 

IC - Ion chromatography; ion-exchange chromatography

ICP-AES - Inductively coupled plasma - atomic emission spectrometry, also called ICP.

Internal Capsule - Air sampler insert consisting of a plastic housing (with an air inlet) 

attached to a membrane filter. Discussion: The internal capsule is fabricated so as to fit inside 

the body of the sampling device (e.g., closed-face cassette sampler), enabling capture of 

airborne particles within the housing / filter construct. [Adapted from: Harper M, Ashley K 

[2013]: Acid-soluble internal capsules for closed-face cassette elemental sampling and analysis 

of workplace air. J Occup Environ Hyg 10:297-306.] 

Interference Equivalent - Mass or concentration of interfering substance (interferant) 

which gives the same measurement reading as unit mass or concentration of substance being 

measured. [Source: NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. 

DHHS (NIOSH) Publication No. 94-113.] 

Interferent - Constituent of the (air) sample or other aspect of the sampling or analytical 

procedure having an adverse effect on the accuracy of the measurement. Note: Interferents 

can include components of sampling or analysis equipment, reagents, etc. [Source: ISO 
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Glossary of Abbreviations, Definitions and Symbols

[2015]. ISO 18158 Workplace air – terminology. Geneva: International Organization for 

Standardization.] 

IR – Infrared

L 

LAQL - Lowest analytically quantifiable level; see LOQ.

LC - Liquid chromatography

LOD – 

1. Limit of detection (detection limit); smallest amount of analyte which can be

distinguished from background. A good estimate for unbiased analyses, with media

blanks not distinguishable from background, is three times the standard error of the

calibration graph for low concentrations, divided by the slope (instrument reading

per unit mass or per unit concentration of analyte). [Source: NIOSH [1994]. NIOSH

Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-

113.]

2. The lowest concentration of an analyte that the bioanalytical procedure can reliably

differentiate from background noise. [Source: FDA [2001]. Guidance for industry -

bioanalytical method validation,

http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf.]

3. The smallest amount, or lowest concentration, of a given substance that a given

procedure will detect [Source: ILO/IPCS. Glossary of terms on chemical safety (after

WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]

LOQ - Limit of quantitation; mass of analyte equal to 10 times the standard error of the 

calibration graph divided by the slope; approximately the mass of analyte for which relative 

standard deviation, SSrr, equals 0.10. [Source: NIOSH [1995]: Guidelines for air sampling and

analytical method development and evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller 

PM, Shulman SA. DHHS (NIOSH) Publication No. 95-117, 

http://www.cdc.gov/niosh/docs/95-117/pdfs/95-117.pdf.] 

Limit of Quantification - Synonymous with limit of quantitation, LOQ. Mass of analyte 

equal to 10 times the standard error of the calibration graph divided by the slope; 
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Glossary of Abbreviations, Definitions and Symbols

approximately the mass of analyte for which relative standard deviation, SSrr, equals 0.10. 

[Source: NIOSH [1995]: Guidelines for air sampling and analytical method development and 

evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. DHHS (NIOSH)

Publication No. 95-117, http:/www.cdc.gov/niosh/docs/95-117/pdfs/95-117.pdf.] 

LTA - Low temperature (oxygen plasma) ashing

M 

MCE; MCEF - Mixed cellulose ester; Mixed cellulose ester membrane filter

MDL - Method detection limit; mass of analyte equal to 3 times the standard error of the 

calibration graph divided by the slope; approximately the mass of analyte for which standard 

deviation, SSrr, equals 0.03. [Source: NIOSH [1995]: Guidelines for air sampling and analytical

method development and evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, 

Shulman SA. DHHS (NIOSH) Publication No. 95-117, http://www.cdc.gov/niosh/docs/95-

117/pdfs/95-117.pdf.] 

Metabolite – 

1. A substance produced directly by a biotransformation of a chemical. For example,

phenol in urine is a metabolite of benzene and is representative of benzene absorption

in the worker. [Source: NIOSH [1994]. NIOSH Manual of analytical methods

(NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.]

2. A substance resulting from chemical transformation in an organism. [Source:

ILO/IPCS. Glossary of terms on chemical safety (after WHO, 1979),

http://www.ilo.org/legacy/english/protection/safework/cis/products/safetytm/glossary

.htm]

Microbiome - The complete genetic content of all the microorganisms that typically inhabit a 

particular environment, especially a site on or in the body, such as the skin or the 

gastrointestinal tract. [Source: Medical Dictionary online, http://www.online-medicaldictionary.org/] 

Measurement Range - Range of substance, in mass per sample, from the LOQ (or from 10 

times the LOD, if LOQ is not known) to an upper limit characteristic of the analytical method, 

e.g., the limit of linearity or the mass at which precision of the method starts to become worse

than SSrr = 0.1. [Source: NIOSH [1995]: Guidelines for air sampling and analytical method

development and evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA.
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Glossary of Abbreviations, Definitions and Symbols

DHHS (NIOSH) Publication No. 95-117, http://www.cdc.gov/niosh/docs/95-117/pdfs/95-

117.pdf.]

Media Blank - An unexposed sampler, not taken or shipped to the field, used for background 

correction of sample readings or for recovery studies. [Source: NIOSH [1994]. NIOSH 

Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.] 

MP - Melting point, °C

MS - Mass spectrometry

MW - Molecular weight

N 

Nanoparticle - Material with all three dimensions in the size range from approximately 1 nm 

to 100 nm [Source: ISO [2015]. ISO 18158 Workplace air – terminology. Geneva: 

International Organization for Standardization.] 

NIOSH - National Institute for Occupational Safety and Health, Centers for Disease Control 

and Prevention, Public Health Service, U. S. Department of Health and Human Services.

Normal Range (biological) - The range of values for an analyte of interest in biological 

monitoring that would be expected in workers without exposure to the environmental 

chemical agent in the workplace. Note: Normal ranges are often method-specific. [Adapted 

from: NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS 

(NIOSH) Publication No. 94-113.] 

NTIS - National Technical Information Service, Springfield, VA 22161.

NTP - Normal temperature and pressure, 25 °C (298 K) and 101.33 kPa (760 mm Hg), at 

which the molar volume of an ideal gas is 24.46 L.

O 

OSHA - Occupational Safety and Health Administration, U. S. Department of Labor.
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Glossary of Abbreviations, Definitions and Symbols

P 

P – 

1. Peak (maximum permissible instantaneous) concentration;

2. Pressure, in kPa, at which sampling pump is calibrated or when air sample was taken.

PAH - Polynuclear aromatic hydrocarbons; PNAH

PCM - Phase contrast microscopy

PEL - OSHA PEL; OSHA permissible exposure limit, expressed as ppm or mg/m3 (of a 

substance in air).

Personal Exposure Monitor - Device used to measure an individual’s personal exposure to 

environmental contaminants or other stressors. A device worn on or near the contact 

boundary that measures concentration [Source: Zartarian VG, Ott WR, Duan N [1997]. A 

quantitative definition of exposure and related concepts. J Exposure Anal Environ Epidemol 

7(4):411-437. and EPA [1997] EPA Terms of Environment – Glossary of Exposure Assessment

Related Terms: A Compilation. Prepared by the Exposure Terminology Subcommittee of the 

IPCS Exposure Assessment Planning Workgroup for the International Programme on

Chemical Safety Harmonization of Approaches to the Assessment of Risk from Exposure to

Chemicals, 2001.]

Personal Sampler - Sampling device, attached to a person that collects gases, vapors, or 

airborne particles in the breathing zone for the purpose of measuring exposure to chemical 

agents and/or biological agents. [Source: ISO [2015]. ISO 18158 Workplace air – terminology. 

Geneva: International Organization for Standardization.] 

PID - Photoionization detector

Plasma, blood - The clear supernatant from whole blood collected with anticoagulants. 

Discussion: Blood is collected, mixed with the anticoagulant and centrifuged to separate the 

plasma from red blood cells. [Adapted from Medical Dictionary online, http://www.onlinemedical-dictionary.org/.] 

PLM - Polarized light microscopy
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Glossary of Abbreviations, Definitions and Symbols

Pool (biological) - A combination of biological specimens (i.e., urine or serum) from many 

workers that is used to prepare small aliquots to be run with each batch of analyses. NOTE: 

The analyte must be stable in the biological matrix and under the storage conditions used. 

Discussion: Aliquots of these pools are analyzed with each batch of samples and the data are 

used to develop quality control charts. [Source: NIOSH [1994]. NIOSH Manual of analytical 

methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.] 

Precision – 

1. The relative variability of measurements on replicate samples about the mean of the

population of measurements. Discussion: Precision is expressed by the relative

standard deviation of a series of measurements, and reflects the ability of a method to

replicate measurement results. [Source: NIOSH [1995]: Guidelines for air sampling

and analytical method development and evaluation. By Kennedy ER, Fischbach TJ,

Song R, Eller PM, Shulman SA. DHHS (NIOSH) Publication No. 95-117,

http://www.cdc.gov/niosh/docs/95-117/pdfs/95-117.pdf.]

2. The repeatability or reproducibility of individual measurements expressed as standard

deviation, S, or relative standard deviation, SSrr (q.v.). See Accuracy. [Source: NIOSH

[2003]. Glossary of abbreviations, definitions, and symbols. In: NIOSH Manual of

analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113,

http://www.cdc.gov/niosh/docs/2003-154/pdfs/glossary.pdf.]

3. The closeness of agreement (degree of scatter) between a series of measurements

obtained from multiple sampling of the same homogenous sample under the

prescribed conditions. [Source: FDA [2001]. Guidance for industry - bioanalytical

method validation,

http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf.]

Proficiency Testing - Any interlaboratory testing program where stable specimens are sent 

to participating laboratories for analysis. Discussion: Results from all participating 

laboratories are compared, pooled, and tabulated by the testing program operator with the 

purpose of improving laboratory performance. [Adapted from Medical Dictionary online,

http://www.online-medical-dictionary.org/.] 

PTFE - Polytetrafluoroethylene

PVC - Polyvinyl chloride
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Glossary of Abbreviations, Definitions and Symbols

Q 

Q - Sampling flow rate, L/min

Quantification Range - The range of concentrations, including upper and lower 

quantification limits (ULOQ and LLOQ, respectively), that can be reliably and reproducibly 

quantified with accuracy and precision through the use of a concentration-response 

relationship. [Source: FDA [2001]. Guidance for industry - bioanalytical method validation,

http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf.] 

R 

Reagent Blank - All reagents used in sample preparation, in the same quantities used to 

prepare blank and sample solutions. Note: The reagent blank is used to assess contamination 

from the laboratory environment and to characterize background from the reagents used in 

sample preparation. [Source: ISO [2015]. ISO 18158 Workplace air – terminology. Geneva: 

International Organization for Standardization.] 

Recovery, R - 

1. Fraction recovered (see DE); previously associated with Analytical Method Recovery

(AMR), a term which is no longer preferred. [Source: NIOSH [1994]. NIOSH Manual

of analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.]

2. The extraction efficiency of an analytical process, reported as a percentage of the

known amount of an analyte carried through the sample extraction and processing

steps of the method. [Source: FDA [2001]. Guidance for industry - bioanalytical

method validation,

http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf.]

Relative Standard Deviation (RSD) - Quotient of standard deviation over the mean; See SSrr

and Precision.

Repeatability - The variation in measurements taken under the same conditions. [Adapted 

from: ISO [2015]. ISO 18158 Workplace air – terminology. Geneva: International 

Organization for Standardization.] 

Reproducibility - A measure of the precision of a method under the same operating 

conditions on the same sample over short period of time. [Adapted from ASTM [2014]. 
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Glossary of Abbreviations, Definitions and Symbols

D1356, Standard terminology relating to sampling and analysis of atmospheres. West

Conshohocken, PA: ASTM International.] 

Respirable Dust - Dust deposited in the non-ciliated portions of the lungs. Discussion: 

Respirable dust is measured by using a respirable sampler when the respirable fraction of 

airborne dust is of interest. [Adapted from ISO [1995]. ISO 7708 Air quality — particle size 

fraction definitions for health-related sampling.’ Geneva: International Organization for 

Standardization.] 

RRff - In thin-layer chromatography, the ratio of distance travelled by the analyte from point of 

application to that of the solvent front.

RF - Radio frequency

Rotameter - A device, based on the principle of Stoke’s Law, for measuring rate of fluid flow, 

consisting of a tapered vertical tube having a circular cross section, and containing a float that 

is free to move in a vertical path to a height dependent upon the rate of fluid flow upward 

through the tube. [Source: ASTM [2014] D1356, Standard terminology relating to sampling 

and analysis of atmospheres. West Conshohocken, PA: ASTM International.] 

RTECS - Registry of Toxic Effects of Chemical Substances (NIOSH)

Ruggedness Test - Partial or complete analysis of variance using experiments in which 

operational parameters of a sampling and measurement method are varied within a small 

range to determine their effect on overall variance [Source: Youden W, Steiner EH [1975]: 

Statistical manual of the AOAC, Arlington, VA: Association of Official Analytical Chemists.] 

S 

SS –

1. Estimate of the standard deviation;

2. Specific mass, particles/mg.

SSbb - Estimate of the standard deviation of media blank

SSrr - Estimate of the relative standard deviation, equal to S divided by the mean of a series of 

measurements. A measure of precision. Equivalent to CV (coefficient of variation).

SS�rr - Pooled relative standard deviation
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Glossary of Abbreviations, Definitions and Symbols

SS�rrrr - Estimate of overall precision including pump error

Sample Dissolution - The process of obtaining a solution containing the analyte(s) of 

interest from a sample. This may or may not involve complete dissolution of the sample. 

[Source: ASTM [2010]. D7035, Standard test method for determination of metals and 

metalloids in airborne particulate matter by inductively coupled plasma atomic emission 

spectrometry (ICP-AES). West Conshohocken, PA: ASTM International.]

Selectivity - 

1. The ability of a bioanalytical method to measure and differentiate the analytes in the

presence of interfering components that may be expected to be present. These could

include metabolites, impurities, degradants, or matrix components. [Adapted from

Medical Dictionary online, http://www.online-medical-dictionary.org/.]

2. Extent of independence of a measuring procedure from interferences. [Source: ISO

[2015]. ISO 18158 Workplace air – terminology. Geneva: International Organization

for Standardization.]

Screening Test (biological) - An easily performed method, often relatively non-specific, to 

assess worker exposure to a class of compounds by use of biological monitoring. [Source: 

NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) 

Publication No. 94-113.] 

SEM - Scanning electron microscopy

Sensitivity – 

1. The slope of the calibration curve. If the curve is in fact a ‘curve’, rather than a straight

line, then of course sensitivity will be a function of analyte concentration or amount.

If sensitivity is to be a unique performance characteristic, it must depend only on the

chemical measurement process, not upon scale factors. [Source: Currie LA [1995]

Nomenclature in evaluation of analytical methods including detection and

quantification capabilities. Pure & Appl Chem 67(10):1699-1723.]

2. The smallest change in the measured analyte concentration that will produce a

reproducible change in a monitor’s readout. [Source: NIOSH [2012]. NIOSH

Technical report: Components for evaluation of direct-reading monitors for gases and

vapors. DHHS (NIOSH) Publication No. 2012-162,

http://www.cdc.gov/niosh/docs/2012-162/pdfs/2012-162.pdf.]
NIOSH Manual of Analytical Methods  5th Edition  Chapter GL April 2016 Page GL-22 of GL-25

Glossary of Abbreviations, Definitions and Symbols

Serum - The clear supernatant from whole blood collected without anticoagulants,

allowed to clot (30 minutes) and centrifuged to separate serum from the clotted blood. 

Serum does not contain clotting factors. [Adapted from Medical Dictionary online, 

http://www.online-medical-dictionary.org/.] 

Sorbent Tube - Sorbent or a support impregnated with reagent, through which sampled air 

passes. Note: Some sorbent tubes are intended for use as active samplers and some as passive 

samplers. [Source: ISO [2015]. ISO 18158 Workplace air – terminology. Geneva: International 

Organization for Standardization.] 

Spike - A known mass of analyte added to a sampler for the purpose of determining 

recovery (analyst spikes), or for quality control (blind spikes). Also see DE. [Source: NIOSH 

[1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication 

No. 94-113.] 

Sp. gr. - Specific gravity. Relative to water at the same temperature.

Spot Sample (urine) - Urine sample collected at a specified time. [Source: NIOSH [1994]. 

NIOSH Manual of analytical methods (NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-

113.] 

STEL - Short-Term (15-min) Exposure Limit.

T 

T - Retention time, min

t – 

1. Temperature, °C;

2. Time, min.

cc - Temperature, degrees kelvin (K), at which sampling pump was calibrated

TEM - Transmission electron microscopy

TLC - Thin-layer chromatography
NIOSH Manual of Analytical Methods  5th Edition  Chapter GL April 2016 Page GL-23 of GL-25

Glossary of Abbreviations, Definitions and Symbols

TLV - Threshold limit value, listed in TLVs® and BEIs®, Threshold Limit Values for

Chemical Substances and Physical Agents and Biological Indices (American

Conference of Governmental Industrial Hygienists, Cincinnati, OH, 2015; updated annually).

ss - Temperature, kelvins (K), at which air sample was taken

TWA - Time-weighted average; the concentration of a chemical or biological agent in the 

atmosphere, averaged over the reference period. [Adapted from: ISO [2015]. ISO 18158

Workplace air – terminology. Geneva: International Organization for Standardization.] 

U 

Ultrafine Particle - Particle with a nominal diameter (such as geometric, 

aerodynamic, mobility, projected-area or otherwise) of 100 nm or less, produced as a 

by-product of a process such as welding and combustion. [Source: ISO [2015]. ISO 

18158 Workplace air – terminology. Geneva: International Organization for 

Standardization.] 

Uncertainty - A limited knowledge of the agreement between data, information, or 

outcomes relative to an unknown truth. The uncertainty of a measurement is the 

parameter associated with the result of a measurement that characterizes the dispersion

of the values that could reasonably be attributed to the measurand (the quantity being 

measured). [Source: Taylor BN, Kuyatt CE [1994]. NIST Technical note 1297, Guidelines

for evaluating and expressing the uncertainty of NIST measurement results.

Gaithersburg, MD, National Institute of Standards and Technology (NIST).]

User Check - An evaluation of a written procedure for clarity and accuracy in which an

independent laboratory analyzes a small number of spiked samples following a draft

sample preparation and analysis exactly as written and reviews the draft method for

clarity. [Source: NIOSH [1994]. NIOSH Manual of analytical methods (NMAM), 4th ed. 

DHHS (NIOSH) Publication No. 94-113.] 

UV – Ultraviolet
NIOSH Manual of Analytical Methods  5th Edition  Chapter GL April 2016 Page GL-24 of GL-25

Glossary of Abbreviations, Definitions and Symbols

V 

V - Volume of air sample, in L, as taken at the sampling site, corrected if necessary for

rotameter calibration at a different temperature and pressure:

VV = (ffffffff rrrrrrrr)( tttt ) �

PPccTTss

PPssTTcc

�

0.5

Validated Method - A Method which meets or exceeds certain sampling and measurement

performance criteria; see for example, the criteria given in Chapter ME [Source: NIOSH 

[2016]. Development and evaluation of methods. In: NIOSH Manual of analytical methods, 

5th ed. DHHS (NIOSH) Publication No. 2014-151, http://www.cdc.gov/niosh/nmam.] 

Validation - Process of evaluating the performance of a measuring procedure and checking 

that the performance meets certain pre-set criteria. Discussion: Performance characteristics to 

be considered include confirmation of identity, selectivity/ specificity, limit of detection, limit 

of quantification, analytical recovery, working and linear dynamic ranges, accuracy, 

measurement repeatability, measurement reproducibility, ruggedness, and robustness. [Source: 

CEN [2011]. EN 1540, Workplace atmospheres – terminology. Brussels: European Standards 

Commission.] 

Vapor – 

1. The gaseous phase of matter that normally exists in a liquid or solid state [Source:

ASTM [2014]. D1356, Standard terminology relating to sampling and analysis of

atmospheres. West Conshohocken, PA: ASTM International.]

2. Gas phase of a substance in a state of equilibrium or disturbed equilibrium with the

same substance in a liquid or solid state below its boiling or sublimation point [Source:

CEN [2011]. EN 1540, Workplace atmospheres – terminology. Brussels: European

Standards Commission.]

Vm - Volume of 1 mole of ideal gas at the specified temperature and pressure (e.g., 24.45 L at 

25 °C and 1 atmosphere).

VOL-MAX - Maximum recommended air sample volume, L, based on sampler capacity or 

other limitation, at the OSHA PEL

VOL-MIN - Minimum recommended air sample volume, L, based on an atmosphere at the 

OSHA PEL concentration and collecting a mass of substance which is equal to the LOQ. See 

also Working range.
NIOSH Manual of Analytical Methods  5th Edition  Chapter GL April 2016 Page GL-25 of GL-25

Glossary of Abbreviations, Definitions and Symbols

VP - Vapor pressure

W 

W - Mass of analyte found on an exposed single-section sampler (e.g., membrane filter)

Wb - Mass of analyte found on the back section of an exposed sampler

Wf - Mass of analyte found on the front section of an exposed sampler

Working Range - Range of air concentrations, in ppm or mg/m3, at specified air sample 

volume, extending from the LOQ to a maximum determined by sampler capacity or 

measurement considerations. [Source: NIOSH [1994]. NIOSH Manual of analytical methods 

(NMAM), 4th ed. DHHS (NIOSH) Publication No. 94-113.] 

X 

XRD - X-ray diffraction

XRF - X-ray fluorescence

Disclaimer

Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not

responsible for the content of these websites. All web addresses referenced in this document 

were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHODS
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATES NOT OTHERWISE REGULATED, TOTAL 0501

DEFINITION: total aerosol mass CAS: NONE RTECS: NONE

METHOD: 0501, Issue 1 EVALUATION: FULL Issue 1: 8 May 2015

OSHA: 15 mg/m3

, total dust (inert or nuisance dust; particles 

not otherwise regulated) [1]

NIOSH: no REL

For other OELs and guidelines: See references [2,3]

PROPERTIES: Contains no asbestos and less than 1% quartz

SYNONYMS: Nuisance dusts; particles not otherwise specified (PNOS)

SAMPLING

SAMPLER: INTERNAL CAPSULE

(tared 37-mm, 2- to 5-μm PVC filter melded to 

PVC housing) in 37-mm 2-piece cassette

FLOW RATE: 1 to 2 L/min

VOL-MIN: 17 L @ 15 mg/m3 -

-MAX: 333 L @ 15 mg/m3

SHIPMENT: Routine

SAMPLE 

STABILITY: 28 days minimum

BLANKS: Minimum of 2 field blanks per batch

ACCURACY

RANGE STUDIED: 0.1 to 4 mg per sample

BIAS: 0.058 [4]

OVERALL 

PRECISION (  ): 0.059 [4

ACCURACY: ± 15.5%

MEASUREMENT

TECHNIQUE: GRAVIMETRIC 

(INTERNAL CAPSULE WEIGHT)

ANALYTE: Airborne particulate material

BALANCE: 0.001 mg sensitivity; use same balance

before and after sample collection

CALIBRATION: National Institute of Standards & 

 Technology Class S-1.1 weights or 

 ASTM Class 1 weights

RANGE: 0.25 to 5 mg per sample

ESTIMATED LOD: 0.075 mg per sample

PRECISION ( ): 0.031 @ ≈2 mg per sample [4

APPLICABILITY: The working range is 2.5 to 50 mg/m3 for a 100-L air sample. This method is nonspecific and determines the 

‘total’ dust concentration to which a worker is exposed. 

INTERFERENCES: Moisture and static electricity can affect gravimetric measurements. Humidity control and minimization of 

static effects are addressed in this procedure.

OTHER METHODS: The method is similar to Method 5100 for carbon black [5]. This method is preferred over 0500, Issue 2 [6]. 

OSHA method PV2121 describes a similar procedure (but for respirable sampling) using an alternative sampler design [7].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATES NOT OTHERWISE REGULATED, TOTAL: METHOD 0501, Issue 1, dated 8 May 2015 - Page 2 of 4

EQUIPMENT:

1. Sampler: Internal capsule, 37-mm polyvinyl chloride (PVC), 2- to 5-μm pore size membrane or 

equivalent hydrophobic filter attached to PVC housing and supporting pad in 37-mm 2-piece 

cassette filter holder 

NOTE: The cassettes should be fabricated so as to ensure complete sealing of the internal capsule 

after sample collection.

2. Personal sampling pump, 1 to 2 L/min, with flexible connecting tubing

3. Microbalance, capable of weighing to ±0.001 mg

4. Static neutralizer, e.g., 210Po; replace no more than nine months after the production date

5. Tool for handling internal capsules, e.g., forceps (preferably plastic)

6. Environmental chamber or room for balance (e.g., 20 °C ± 1 °C and 50% ± 5% RH)

SPECIAL PRECAUTIONS: None.

PREPARATION OF INTERNAL CAPSULES BEFORE SAMPLING:

1. Equilibrate the PVC filter capsules in an environmentally controlled weighing area or chamber for at 

least 24 hours.

NOTE: An environmentally controlled chamber is desirable, but not required.

2. Place backup pads in filter cassette bottom sections.

3. Weigh the filter capsules in an environmentally controlled area or chamber. Record the internal 

capsule tare weight, W1

 (mg).

a. Zero the balance before each weighing.

b. Handle the filter capsule with forceps. Pass the internal capsule over an antistatic radiation source. 

Repeat this step if the capsule does not release easily from the forceps or if it attracts the balance 

pan. Static electricity can cause erroneous weight readings.

4. Assemble the filter capsules in the filter cassettes and close firmly so that leakage around the internal 

capsule will not occur. Place a plug in each opening of the filter cassette. Place a cellulose shrink band 

around the filter cassette, allow to dry and label the cassette with indelible ink.

SAMPLING:

5. Calibrate each personal sampling pump with a representative sampler in line.

6. Sample at 1 to 2 L/min for a total sample volume of 17 to 333 L. Do not exceed a total filter capsule 

loading of approximately 5 mg total dust. Take two to four replicate samples for each batch of field 

samples for quality assurance on the sampling procedure.

SAMPLE PREPARATION:

7. Wipe dust from the external surface of the filter cassette with a moist paper towelette to minimize 

contamination. Discard the paper towelette.

8. Remove the top and bottom plugs from the filter cassette. Equilibrate for at least 24 hours in the 

balance room. 

9. Using forceps, open the cassette and remove the internal capsule gently to avoid loss of dust or 

damage to the capsule.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATES NOT OTHERWISE REGULATED, TOTAL: METHOD 0501, Issue 1, dated 8 May 2015 - Page 3 of 4

CALIBRATION AND QUALITY CONTROL:

10. Zero the microbalance before all weighings. Use the same microbalance for weighing filter capsules

before and after sample collection. Calibrate the balance with National Institute of Standards and

Technology Class S-1.1 or ASTM Class 1 weights.

11. Process laboratory blanks, spiked QC samples and field blanks at a minimum frequency of 1 per 20

field samples. Internal capsules used for QC samples should come from the same lot. Spiked QC

samples, loaded with 0.25-4 mg of material per internal capsule, should be prepared using weightstable material such as Arizona Road Dust [8].

MEASUREMENT:

12. Weigh each capsule, including field blanks. Record the post-sampling weight, W2

 (mg). Record

anything remarkable about a capsule (e.g., overload, leakage, wet, torn, etc.).

CALCULATIONS:

13. Calculate the concentration of total particulate matter, C (mg/m3

), in the air volume sampled, V (L):

  , mg/m

where: W1 = tare weight of capsule before sampling (mg)

W2

 = post-sampling weight of sample-containing capsule (mg)

 B1

 = mean tare weight of blank capsules (mg)

 B2

 = mean post-sampling weight of blank capsules (mg)

EVALUATION OF METHOD:

Lab testing was carried out using blank internal capsules and with capsules spiked with 0.1 – 4 mg of 

NIST SRM 1648 (Urban Particulate Matter) and Arizona Road Dust (Air Cleaner Test Dust) [4]. Precision and 

accuracy data are given on page 0501-1. Weight stability over 28 days was verified for both blanks and 

spiked capsules [4]. Independent laboratory testing on blanks and field samples have verified long-term 

weight stability as well as sampling and analysis uncertainty estimates [4,8].

REFERENCES:

[1] CFR. 29 CFR Part 1910.1000. Code of Federal Regulations. Washington, DC: U.S. Government Printing

Office, Office of the Federal Register.

[2] NIOSH [2005]. NIOSH pocket guide to chemical hazards. Barsan ME, ed. Cincinnati OH: U.S.

Department of Health and Human Services, Centers for Disease Control and Prevention, National

Institute for Occupational Safety and Health. DHHS (NIOSH) Publication no. 2005-149.

[3] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2014]. GESTIS-Database

on hazardous substances. (German Social Accident Insurance). Sankt Augustin, Germany: IFA. 

https://www.dguv.de/ifa/index-2.jsp (Date accessed: February, 2015).

[4] NIOSH [2014]. Backup Data Report-NIOSH 0501 and 5100. By O’Connor SP, O’Connor PF, Feng HA,

Ashley K. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control

and Prevention, National Institute for Occupational Safety and Health. [http://www.cdc.gov/niosh/

docs/2003-154/pdfs/bud/1453_bud.pdf].

[5] NIOSH [2015]. Carbon black: Method 5100. In: Ashley KE, O’Connor PF, eds. NIOSH manual of

analytical methods. 5th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATES NOT OTHERWISE REGULATED, TOTAL: METHOD 0501, Issue 1, dated 8 May 2015 - Page 4 of 4

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 2014-151. [www.cdc.gov/niosh/nmam/]. 

[6] NIOSH [1994]. Particulates not otherwise regulated, total: Method 0500. In: Eller PM, Cassinelli ME, 

eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and 

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/]. 

[7] OSHA [2003]. Gravimetric determination: PV2121. In: Monitoring methods used by OSHA. Salt Lake 

City, UT: U.S. Department of Labor, Occupational Safety and Health Administration. [https://www.

osha.gov/dts/sltc/methods/partial/pv2121/pv2121.html] (Date accessed: February, 2015).

[8] O’Connor S, O’Connor PF, Feng HA, Ashley K [2014]. Gravimetric analysis of particulate matter using 

air samplers housing internal filtration capsules. Gefahrsstoffe Reinh Luft 75: 403-410.

METHOD WRITTEN BY: Kevin Ashley, Ph.D., NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by 

NIOSH. In addition, citations to websites external to NIOSH do not constitute NIOSH 

endorsement of the sponsoring organizations or their programs or products. Furthermore, 

NIOSH is not responsible for the content of these websites. All web addresses referenced in 

this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

VINYL ACETATE 1453

CH3CO2CHCH2 MW: 86.09 CAS: 108-05-4 RTECS: AK0875000

METHOD: 1453, Issue 3 EVALUATION: FULL Issue 1: 15 August 1994

Issue 3: 15 March 2013 

OSHA: None

MSHA: None

NIOSH: Ceiling 4 ppm (15 min) 

OTHER OELs: [1,2] 

PROPERTIES: Liquid; BP 72.7 °C; d 0.934 g/mL @ 20 °C;

VP 11.8 kPa (89 mm Hg) @ 20 °C; 

vapor density (air = 1) 3.0

SYNONYMS: Acetic acid, vinyl ester; acetic acid, ethenyl ester; Vinyl A monomer; ethylene ethanoate; 1-acetoxyethylene; ethenyl 

acetate; vinyl acetate monomer; vinyl ethanoate.

SAMPLING 

SAMPLER: SOLID SORBENT TUBE 

(carbon molecular sieve; 160 mg/80 mg)

FLOW RATE: 0.05 to 0.2 L/min

VOL-MIN: 1.5 L @ 4 ppm

 -MAX: 24 L

SHIPMENT: Routine

SAMPLE 

STABILITY: At least 30 days @ 5 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY 

RANGE 

STUDIED: 3.7 to 46 mg/m3 (24-L sample) [3]

BIAS: -0.04

OVERALL

PRECISION (SS�rrrr): 0.064

ACCURACY: 14.1%

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, FID

ANALYTE: Vinyl acetate

DESORPTION: 1.0 mL 95:5% (v/v) methylene 

chloride/methanol

INJECTION 

VOLUME: 1 μL 

TEMPERATURE -INJECTION: 210 °C

-DETECTOR: 260 °C

-COLUMN: 35 °C, 5 min; 5 °C/min to 

50 °C; hold 1 min 

CARRIER GAS: Helium, split flow 21.0 mL/min

COLUMN: Capillary, fused silica, 30 m x 0.32 mm,

coated internally with 1 μm film (5% 

phenyl) methylpolysiloxane

CALIBRATION: Standard solutions of vinyl acetate in 

95:5% (v/v) methylene 

chloride/methanol

RANGE: 3 to 1120 μg per sample

ESTIMATED LOD: 1 μg per sample

PRECISION (SS�rr): 0.040

APPLICABILITY: The working range is 0.02 to 13.1 ppm (0.07 to 46 mg/m3

) for a 24-L air sample. The method is sensitive enough 

for ceiling measurements.

INTERFERENCES: Vinyl acetate that is not stabilized or has been depleted of inhibitor can polymerize. Any substance collected

with the vinyl acetate that is capable of reacting with it is a potential interference. Acids, bases, free radical initiators, etc., are 

capable of reacting with vinyl acetate during and after air sampling.

OTHER METHODS: This method is an update for NMAM 1453 first issued on August 15, 1994 [4]. This method is adapted from 

OSHA Method No. 51 [5] and NIOSH 1453 Issue 2 and replaces NIOSH method P&CAM 278 [6]. 
VINYL ACETATE: METHOD 1453, Issue 3, 15 March 2013- Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS: 

1. Vinyl acetate*, 99% purity, inhibited with

3 – 22 ppm hydroquinone.

2. Methanol*, distilled in glass.

3. Methylene chloride*, distilled in glass.

4. Desorption solvent, 95%/5% (v/v)

methylene chloride/methanol.

5. Helium, purified.

6. Hydrogen, prepurified.

7. Air, filtered, compressed.

8. Calibration stock solution, 74.7 mg/mL. Add

400 μL vinyl acetate into a 5-mL volumetric

flask and dilute to volume with methanol.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: glass tube, 7 cm long, 6-mm OD, 4-

mm ID, flame-sealed ends with plastic caps,

containing two sections of carbon molecular

sieve, (front = 160 mg, back = 80 mg)

separated by a 2-mm urethane foam plug. A

silylated glass wool plug precedes the front

section and a 3-mm urethane foam plug

follows the back section. Pressure drop across

the tube at 1 L/min airflow must be less than

25 mm/Hg. Tubes are commercially available.

2. Personal sampling pump, 0.1 to 0.2 L/min,

with flexible connecting tubing.

3. Gas chromatograph with flame ionization

detector (GC-FID), column (page 1453-1) and

data collector.

4. Vials, 2-mL, with PTFE-lined caps.

5. Syringe, 10-μL and other sizes as needed.

6. Volumetric flasks, 2- and 5-mL.

SPECIAL PRECAUTIONS: Methylene chloride is an irritant, can be absorbed through the skin, and is a 

potential occupational carcinogen [7]. Vinyl acetate can irritate the eyes, nose, and throat. Vinyl Acetate is 

flammable in air (percent by volume): lower, 2.6%; upper, 13.4% [8]. Vinyl acetate can become unstable if 

the polymerization inhibitor decreases to unsafe levels [8]. Methanol is flammable and a dangerous fire 

and explosion risk. It is moderately toxic by ingestion and inhalation. Wear appropriate protective clothing 

and work with these compounds in a well ventilated hood.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break the ends of the sampler immediately before sampling. Attach sampler to personal sampling

pump with flexible tubing.

3. Sample at an accurately known flow rate of 0.1 to 0.2 L/min for a total sample size of 1.5 to 24 L.

NOTE: If high concentrations are expected, multiple samplers may need to be used to collect over an 8

hour period (collect two 4-hour samples for example) in order to not exceed the capacity of the 

sorbent tube

4. Cap the samplers. Pack securely for shipment.

SAMPLE PREPARATION:

5. Place the front sorbent section along with the glass wool plug in a separate vial from the back sorbent

section of the sampler. Discard the foam plugs.

6. Add 1.0 mL of 95:5% (v/v) methylene chloride/methanol to each vial. Cap each vial.

7. Allow to stand 30 min with occasional agitation.

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards.

a. Add 400 μL vinyl acetate to methanol in a 5-mL volumetric flask and dilute to the mark. This is the

stock calibration solution. Use serial dilutions, as needed, to obtain the desired concentration

range.
VINYL ACETATE: METHOD 1453, Issue 3, 15 March 2013- Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

b. Analyze with samples and blanks (steps 11 and 12).

c. Prepare calibration graph (area vs. μg/mL).

9. Determine desorption efficiency (DE) at least once for each lot of sorbent used for sampling in the

range of interest (step 8). Prepare three tubes at each of five levels plus three media blanks.

a. Remove and discard back sorbent section of a media blank sampler.

b. Inject a known amount (2 to 20 μL) of a standard mixture of vinyl acetate directly onto front sorbent

section with a microliter syringe.

NOTE: Inject no more than 20 μL onto the sorbent. Dilute stock solution as appropriate.

c. Cap the tube. Allow to stand overnight.

d. Desorb (steps 5 through 7) and analyze with working standards (steps 11 and 12).

e. Prepare a graph of DE vs. μg vinyl acetate recovered.

10. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration graph

and recovery graph are in control.

MEASUREMENT:

11. Set gas chromatograph according to manufacturer’s recommendations and to conditions given on

page 1453-1. Inject sample aliquot manually using solvent flush technique or with autosampler.

NOTE: If peak area is above the linear range of the working standards, dilute an aliquot of the desorbed

liquid with desorption solvent, reanalyze, and apply the appropriate dilution factor in 

calculations.

12. Measure the peak area of the vinyl acetate signal.

CALCULATIONS:

13. Determine the mass, μg (corrected for DE) of vinyl acetate found in the sample front (Wf) and back (Wb)

sorbent sections, and in the average media blank front (Bf) and back (Bb) sorbent sections.

NOTE: If Wb > Wf /10, report breakthrough and possible sample loss.

14. Calculate concentration, C, of vinyl acetate in the air volume sampled, V (L):

CC = �WWff + WWbb� − �BBff + BBbb�

V , mmmm/mm3

NOTE: μg/L ≌ mg/m3

EVALUATION OF METHOD:

This method was validated originally by the Organic Methods Development Branch, OSHA Technical 

Center, Salt Lake City, UT, over the range 187 to 710 μg per sample using Ambersorb® XE-347 as sorbent. 

Breakthrough studies at 70% RH, sampling an atmosphere of 150 mg/m3 at 0.19 L/min, indicated a tube 

capacity of approximately 9 mg of vinyl acetate [5]. 

This media is no longer available and is replaced with Carboxen® 564 carbon molecular sieve tubes 

(OrboTM-92 or equivalent tubes). Using OrboTM-92 media, the average desorption efficiency for samples 

in this range was 98.5%. Recoveries from samples stored for 30 days at 5 °C were 90 to 110% for samples

spiked at 187 and 747 μg per sample. The pooled coefficient of variation obtained from replicate

determinations of analytical standards in this range was 0.020 [3,9]. 
VINYL ACETATE: METHOD 1453, Issue 3, 15 March 2013- Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] SCOEL [2005]. Recommendation from the scientific committee on occupational exposure limits for

vinyl acetate. Brussel, Belgium: Scientific Committee on Occupational Exposure Limits.

[https://ec.europa.eu/social/main.jsp?catId=148&intPageId=684&langId=en] October 2005.

[2] ACGIH [2014]. TLVs® and BEIs® based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and biological exposure indices. Cincinnati, OH: American

Conference of Governmental Industrial Hygienists [http://www.acgih.org/].

[3] NIOSH [2013]. Backup data report for vinyl acetate. Cincinnati, OH: U.S. Department of Health and

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health (unpublished).

[4] NIOSH [1994]. Vinyl acetate: Method 1453. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical

methods, 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease

Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH)

Publication No. 94-113 [www.cdc.gov/niosh/nmam/].

[5] OSHA [1985]. Vinyl acetate: OSHA Method ORG-51. Salt Lake City, Utah: U.S. Department of Labor,

Occupational Safety and Health Administration. [http://www.osha.gov/dts/sltc/methods/organic/

org051/org051.html]. Date accessed October 2012.

[6] NIOSH [1978]. Vinyl acetate: P & CAM 278. In: Taylor DG, ed. NIOSH manual of analytical methods,

2nd ed., Vol 4. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 78-175.

[7] NIOSH [2005]. NIOSH pocket guide to chemical hazards. Barsan ME, ed. Cincinnati OH: U.S.

Department of Health and Human Services, Centers for Disease Control and Prevention, National

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149.

[8] Gustin J-L [2005]. Understanding vinyl acetate polymerization accidents. Chem Health Saf

12(6):36-46.

[9] Yoon YH, Perkins JB [1998]. Vinyl acetate by GC-FID sample analysis backup data report. Salt Lake

City, UT: DataChem Laboratories. Developed under NIOSH contract CDC-200-2001-08000

(unpublished). 

METHOD REVISED BY:

3rd Issue (2013) Y. T. Gagnon and P. F. O’Connor, CDC/NIOSH, Cincinnati, Ohio.

2nd Issue (1998) Y. H. Yoon, Ph.D. and J. B. Perkins, DataChem Laboratories, Inc., Salt Lake City, Utah.

1st Issue (1994) A. A. Grote, CDC/NIOSH, Cincinnati, Ohio.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

NITROAROMATIC COMPOUNDS 2005

(1) Nitrobenzene: C6H5NO2 MW: (1) 123.11 CAS: (1) 98-95-3 RTECS: (1) DA6475000

(2) o-Nitrotoluene: CH3C6H4NO2 (2) 137.14 (2) 88-72-2 (2) XT3150000

 m-Nitrotoluene: 99-08-1 XT2975000

 p-Nitrotoluene: 99-99-0 XT3325000

(3) 4-Chloronitrobenzene: C6H4ClNO2 (3) 157.56 (3) 100-00-5 (3) CZ1050000

METHOD: 2005, Issue 4 EVALUATION: FULL Issue 1: 15 August 1990

Issue 4: 4 March 2016

OSHA: Table 1

NIOSH: Table 1

PROPERTIES: Table 1

SYNONYMS: (1) Nitrobenzol, oil of mirbane; (2) o-Methylnitrobenzene, 2-Methylnitrobenzene, 2-Nitrotoluene mMethylnitrobenzene, 3-Methylnitrobenzene, 3-Nitrotoluene p-Methylnitrobenzene, 4-Methylnitrobenzene, 4-Nitrotoluene; (3) pChloronitrobenzene, 1-Chloro-4-nitrobenzene, 4-Nitrochlorobenzene, PCNB, PNCB

SAMPLING

SAMPLER: SOLID SORBENT TUBE (silica gel, 150 

mg/75 mg)

FLOW RATE: (1) and (3) 0.01 - 1 L/min; 

(2) 0.01 to 0.02 L/min

(1) (2) (3)

VOL-MIN: 10 L 1L 1L

 -MAX: 150 L 30L 150L

SHIPMENT: routine

SAMPLE 

STABILITY: 30 days @ 0 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: Table 1

BIAS: Table 1

OVERALL

PRECISION (SS�rrrr): Table 1

 

ACCURACY: Table 1

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, FID

ANALYTE: nitrobenzene, nitrotoluene isomers, 

4-chloronitrobenzene

DESORPTION: 1 mL methanol in ultrasonic bath for 30

minutes mL acetonitrile

INJECTION

VOLUME: 1 μL

TEMPERATURE

 -INJECTION: 250 °C

 -DETECTOR: 300 °C 

 -COLUMN: 80 °C, 1 min, 8 °C/min to 180 °C 

CARRIER GAS: He, 2.5 to 3.0 mL/min

COLUMN: capillary, 30 m x 0.53-mm ID; μm film

crossbonded 5% diphenyl 95% 

dimethylpolysiloxane

CALIBRATION: analytes in methanol 

RANGE: Table 1

ESTIMATED LOD: Table 1

PRECISION (SS̅

rr): Table 1

APPLICABILITY: The working ranges for a 30-L air samples are 0.396 to 1.92 ppm (1.98 to 9.60 mg/m3

) for nitrobenzene; 0.346 to 

1.73 ppm (1.97 to 9.86 mg/m3

) for o-nitrotoluene; 0.344 to 1.72 ppm (1.96 to 9.81 mg/m3) for m-nitrotoluene; 0.303 to 1.52 ppm 

1.73 to 8.67 mg/m3) for p-nitrotoluene; and 0.308 to 1.54 ppm (1.98 to 9.92 mg/m3) for 4-chloronitrobenzene [1,2].

INTERFERENCES: Any compounds with retention times similar to the analytes of interest will interfere. During sampling, high 

humidity may greatly decrease breakthrough volume

OTHER METHODS: This method is an update of NMAM 2005, Nitrobenzenes, issued 15 August 1994, which combined and 

replaced methods S217, S218, and S223 [2,3].
NITROAROMATIC COMPOUNDS: METHOD 2005, Issue 4, dated 4 March 2016 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Methanol, HPLC chromatographic grade.

2. Nitrobenzene*, reagent grade.

3. o-,m-,p-nitrotoluene isomers,* reagent

grade.

4. 4-chloronitrobenzene*, reagent grade.

5. Calibration stock solution, 500 μg/mL.

Prepare each analyte in methanol.

6. Helium, purified and filtered.

7. Hydrogen, purified and filtered.

8. Air, purified and filtered.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: silica gel sampling tube; glass tube, 7 

cm long, 6-mm OD, 4-mm ID, flame-sealed 

ends with plastic caps, containing two sections 

(front=150 mg; back=75 mg) of 20/40 mesh 

silica gel separated by a 2-mm urethane foam 

plug. A silylated glass wool plug precedes the 

front section and a 3-mm urethane foam plug 

follows the back section.

2. Personal sampling pump, 0.01 to 1 L/min, with 

flexible connecting tubing.

3. Gas chromatograph, FID, integrator and 

column (page 2005-1).

4. Autosampler vials, glass, 2-mL, with PTFElined crimp caps.

5. Volumetric flasks, 10-mL.

6. Pipets, 5-mL and 3-mL, with pipet bulb.

7. Syringes, 10-μL, 100-μL, and 1-mL.

8. Ultrasonic bath.

 

SPECIAL PRECAUTIONS: These analytes are severe poisons and irritants. Prevent contact with eyes, 

skin, or clothing by wearing eye protection, chemically resistant gloves, and a lab coat. Avoid inhalation. 

Nitrobenzene and m-nitrotoluene are absorbed through contact with skin and can cause 

methemoglobinemia [5,6]. 4-Chloronitrobenzene is a carcinogen. Methanol is highly flammable.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break the ends of the sampler immediately before sampling. Attach sampler to personal sampling 

pump with flexible tubing.

3. Sample at an accurately known flow rate between 0.01 and 0.2 L/min for nitrotoluene isomers. Use a 

flow rate of 1 L/min or less for nitrobenzene and 4-chloronitrobenzene. Note the maximum and 

minimum sample volumes on page 2005-1.

4. Cap both ends of the sampler. Pack securely for shipment.

SAMPLE PREPARATION:

5. Place the front (include the glass wool plug) and back sorbent sections of each sample tube in 

separate vials. Discard the foam plugs.

6. Add 1.0 mL of methanol to each vial. Attach crimp cap securely to each vial.

7. Allow to desorb 30 min in an ultrasonic bath.
NITROAROMATIC COMPOUNDS: METHOD 2005, Issue 4, dated 4 March 2016 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards to cover the analytical range of the method. If 

necessary, additional standards may be added to extend the calibration curve.

a. Add known amounts of calibration stock to methanol in 10-mL volumetric flasks and dilute to 

the mark.

b. Analyze together with samples and blanks (steps 11 and 12).

c. Prepare calibration graph (peak area vs μg analyte).

9. Determine desorption efficiency (DE) at least once for each lot of silica gel used for sampling in the 

calibration ranges (step 8).

a. Prepare three tubes at each of five levels plus three media blanks.

b. Inject a known amount of calibration stock solution directly onto the front sorbent section of 

each silica gel tube with a microliter syringe.

c. Allow the tubes to air equilibrate for several minutes, then cap the ends of each tube and allow 

to stand overnight.

d. Desorb (steps 5 through 7) and analyze together with standards and blanks (steps 11 and 12).

e. Prepare a graph of DE vs μg analyte recovered.

10. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration 

graph and DE graph are in control.

MEASUREMENT:

11. Set gas chromatograph according to manufacturer’s recommendations and to conditions given on 

page 2005-1. Inject a 1-μL sample aliquot manually using the solvent flush technique or with an 

autosampler.

NOTE: If peak area is above the linear range of the working standards, dilute with methanol, 

reanalyze and apply the appropriate dilution factor in the calculations.

12. Measure peak areas.

CALCULATIONS:

13. Determine the mass, μg (corrected for DE) of analyte found in the sample front (Wf) and back (Wb) 

sorbent sections, and in the average media blank front (Bf) and back (Bb) sorbent sections.

NOTE: If Wb > Wf /10, report breakthrough and possible sample loss.

14. Calculate concentration, C, of analyte in the air volume sampled, V(L):

CC = WWff + WWbb − BBff − BBbb

VV , mmmm/mm3

NOTE: μg/mL≅mg/m3

EVALUATION OF METHOD: 

The 3rd issue update of this method included the use of capillary column chromatography (RtxTM-5 Amine) 

that lowered the LOD/LOQ values, a lower 5-level desorption efficiency study, and a 30-day storage 

stability study for each analyte [1]. The method evaluation data for these compounds are listed in Table 2. 

Methods S217, Nitrobenzene, and S218, 4-nitrochlorobenzene, were initially issued on November 21, 1975 

[4]. Method S223, o-nitrotoluene was issued on December 19, 1975 [4]. The analytes m-nitrotoluene and pnitrotoluene were added on May 15, 1984 [3]. In the original method development work, sample tube 

capacity, or breakthrough, was determined as 5% of the generated atmosphere concentration as

measured in the effluent of the sample tubes. Capacity was measured at >2.8 mg/sample for 

nitrobenzene; >2.5 mg/sample for nitrotoluene isomers; and >2.2 mg/sample for 4-chlorobenzene [2]. 
NITROAROMATIC COMPOUNDS: METHOD 2005, Issue 4, dated 4 March 2016 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] Pendergrass SM [1997]. Backup data report for nitroaromatic compounds method development. 

Cincinnati, OH: National Institute for Occupational Safety and Health. Unpublished.

[2] NIOSH [1977]. Methods S217, S218, and S223. In: Documentation of the NIOSH validation tests. 

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center of Disease Control and 

Prevention, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 

77-185.

[3] NIOSH [1994]. m-Nitrotoluene and p-Nitrotoluene: Method 2005. In: Eller PM, Cassinelli ME, eds. 

NIOSH manual of analytical methods, 4th ed. Cincinnati, OH: U.S. Department of Health and 

Human Services, CentersforDisease Control and Prevention, National Institute forOccupational

Safety and Health, DHHS (NIOSH) Publication No. 94-113.

[4] NIOSH [1978]. Nitrobenzene: Method S217; p-Nitrochlorobenzene:Method S218; and

Nitrotoluene: Method S223. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd ed.

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control,

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-157-C.

[5] NIOSH[1977]. Aromatic hydrocarbons and nitrobenzeme. In: Key MM, Henschel AF, Butler J, Ligo 

RN, Tabershaw IR, Ede L, eds. Occupational diseases: A guide to theirrecognition. revised ed.

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-181. 

[6] NIOSH [1981]. Occupational health guidelines for chemical hazards, nitrobenzene, 

nitrochlorobenzene and nitrotoluene, U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication

No. 81-123.

METHOD WRITTEN BY:

Stephanie M. Pendergrass, NIOSH

TABLE 1. General Information

Chemical OSHA PEL NIOSH REL Physical Properties Method 

Evaluation

Nitrobenzene

1 ppm

(5 mg/m3) 

(skin)

1 ppm

(5 mg/m3

) 

(skin)

Colorless oily liquid, almond odor; 

d=1.196 g/mL @ 20 °C; MP 6 °C; BP 

210- 211 °C; VP=37 Pa (0.30 mm Hg) 

@ 20 °C

Full

o-Nitrotoluene

5 ppm

(30 mg/m3) 

(skin)

2 ppm

(11 mg/m3) 

(skin)

yellowish liquid; d=1.163 g/mL @ 20 

°C; MP -4 °C; BP 222 °C; VP=20 Pa 

(0.15 mm Hg) @ 20 °C

Full

m-Nitrotoluene

5 ppm

(30 mg/m3) 

(skin)

2 ppm

(11 mg/m3

) 

(skin)

liquid; d=1.157 g/mL @ 20 °C; MP 16 

°C; BP 232 °C; VP=20 Pa (0.15 mm Hg) 

@ 20 °C

Partial

p-Nitrotoluene

5 ppm

(30 mg/m3) 

(skin)

2 ppm

(11 mg/m3) 

(skin)

yellow crystals; d=1.163 g/mL @ 20 

°C; MP 52 °C; BP 238 °C; VP=17 Pa 

(0.12 mm Hg) @ 20 °C

Partial

4-Chloronitrobenzene

0.16 ppm

(1. mg/m3

) 

(skin)

Ca*

(skin)

yellow crystals; d=1.298 g/mL @ 20 

°C; MP 83 °C; BP 242 °C; VP=28 Pa (0.2 

mm Hg) @ 30 °C

Partial

* - Cancer suspect agent
NITROAROMATIC COMPOUNDS: METHOD 2005, Issue 4, dated 4 March 2016 - Page 5 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 2. Method Summary

Chemical

Range 

Studied 

(mg/m3)

SS�rrrr Bias Accuracy 

(±%)

Analytical 

Range

LOD (μg/

sample) SS�rr

Desorption 

Efficiency 

(%)

30-Day 

Storage 

(% Rec)

Nitrobenzene 1.98-9.60 0.0590 0.0186 12.3 2 to 598 0.6 0.12 98.7 100.2

o-Nitrotoluene 1.97-9.86 0.0142 -0.120 21.1 3 to 582 0.8 0.028 98.2 101.2

m-Nitrotoluene Not studied ndb nd nd 3 to 579 1.0 0.042 97.5 99.4

p-Nitrotoluene Not studied nd nd nd 9 to 511 2.6 0.061 96.9 99.4

4-Chloronitrobenzene 1.98-9.92 0.1034 0.0869 27.3c 8 to 595 2.5 0.063 100.3 97.6

a

30-L air sample

b Not determined

c Exceeds the NIOSH accuracy criterion of ± 25% at the 95% confidence level

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

CHLOROACETIC ACID 2008

ClCH2COOH MW: 94.50 CAS: 79-11-8 RTECS: AF8575000

METHOD: 2008, Issue 3 EVALUATION: FULL Issue 1: 15 May 1989

Issue 3: 15 March 2016

OSHA: None

NIOSH: None

PROPERTIES: solid; MP 61 to 63 °C; BP 189 °C; 

VP 0.1 kPa (0.75 mm Hg) @ 20 °C; flash

point 126 °C; lower explosive limit in

air 8% v/v

SYNONYMS: chloroethanoic acid; monochloroacetic acid

SAMPLING

SAMPLER: SOLID SORBENT TUBE (silica gel, 100 

mg/50 mg)

FLOW RATE: 0.05 to 0.2 L/min

VOL-MIN: 1 L @ 0.25 ppm 

 -MAX: 100 L

SHIPMENT: routine

SAMPLE

STABILITY: at least 7 days @ 25 °C; 32 days 

refrigerated [1] 

BLANKS: 2 to 10 field blanks per set

ACCURACY 

RANGE 

STUDIED: 0.35 to 29 mg/m3 [1,2] (3-L samples)

BIAS: 2.0%

OVERALL

PRECISION (SS�rrrr): 0.08 [1]

ACCURACY: ± 17.7%

MEASUREMENT

TECHNIQUE: ION CHROMATOGRAPHY, CONDUCTIVITY 

DETECTION

ANALYTE: chloroacetate ion

DESORPTION: 2 mL deionized water

INJECTION 

LOOP VOLUME: 500 μL

ELUENT: 1.5 mM NaHCO3; 1.0 mL/min

COLUMNS: US Pharmacopeia (USP) L12 separator 

column and manufacturer’s compatible 

anion guard column

CALIBRATION: standard solutions of chloroacetic acid in

deionized water

RANGE: 1 to 80 μg per sample [1]

ESTIMATED LOD: 0.04 μg per sample [2]

PRECISION (SS�rr): 0.016 [1]

APPLICABILITY: The working range is 0.09 to >85 ppm (0.3 to 30 mg/m3

) for a 3-L air sample.

INTERFERENCES: Chloroacetyl chloride is a positive interferent since it is hydrolyzed to monochloroacetic acid by the 

measurement procedure and is efficiently collected by silica gel [3]. Particulate salts of the acid are positive interferents. The 

chromatographic conditions given will separate acetate, chloride, dichloroacetate, fluoride, glycolate, and trichloroacetate ions 

from chloroacetate ion. 

OTHER METHODS: This revises P&CAM 332 [2]. The columns used in P&CAM 332 are no longer available. The newer columns 

indicated here show improvements in the analytical range and sensitivity.
CHLOROACETIC ACID: METHOD 2008, Issue 2, dated 15 March 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Water, filtered, deionized. Specific 

conductance ≤10 μS/cm.

2. Sodium bicarbonate (NaHCO3), reagent 

grade.

3. Chloroacetic acid, ≥99%.*

4. Eluent: 1.5 mM NaHCO3. Dissolve 0.504 g 

NaHCO3 in 4 L filtered, deionized water.

5. Calibration stock solution, 1000 μg/mL. 

Dissolve 100 mg chloroacetic acid in 100 mL 

filtered, deionized water. 

*See SPECIAL PRECAUTIONS.

EQUIPMENT: 

1. Sampler: silica gel sorbent tube; glass tube, 7 

cm long, 6-mm OD, 4-mm ID, with plastic 

caps, containing two sections of 20/40 mesh 

silica gel (front = 100 mg; back =50 mg) 

contained and separated by three silanized 

glass wool plugs. Pressure drop across the 

tube at 0.2 L/min is ca. 0.6 kPa (2.6 in. H2O).

Tubes are commercially available.

NOTE: Chloroacetic acid is irreversibly 

adsorbed on urethane plugs. Use 

sorbent tubes with glass wool plugs.

2. Personal sampling pump, 0.05 to 0.2 L/min, 

with flexible connecting tubing.

3. Ion chromatograph (IC), anion separator (USP 

L12) and compatible guard column, anion 

suppressor (page 2008-1), conductivity 

detector and integrator. 

4. Ultrasonic bath.

5. Vials, 20-mL, glass, with aluminum-lined 

plastic screw caps.

6. Syringes, 3-mL, polyethylene with luer tip.

7. Filter holder, luer tip, 13-mm, with 

polytetrafluoroethylene (PTFE) filter, 5-μm 

pore size, or PTFE syringe filter.

8. Pipets, 10-μL to 2-mL.

9. Flasks, volumetric, 10- and 100-mL.

 

SPECIAL PRECAUTIONS: Chloroacetic acid is irritating to skin and mucous membranes [4]. Work with 

concentrated material only in a hood. 

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break ends of sampler immediately before sampling. Attach sampler to personal sampling pump with 

flexible tubing.

3. Sample at an accurately known flow rate between 0.05 to 0.2 L/min for a total sample size of 1 to 100 L.

4. Cap the samplers. Pack securely for shipment.

NOTE: Store samples in the dark. Refrigerate samples if stored longer than 7 days. 

SAMPLE PREPARATION:

5. Allow refrigerated samples to equilibrate to room temperature.

6. Transfer front sorbent section with front glass wool plug to vial. Place back sorbent section and other 

two glass wool plugs in separate vial.

7. Add 2.0 mL deionized water to each vial. Cap immediately.

8. Agitate vials in ultrasonic bath for 30 min at room temperature.

9. Draw sample extract through 13-mm PTFE filter with 3-mL syringe. 
CHLOROACETIC ACID: METHOD 2008, Issue 2, dated 15 March 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALIBRATION AND QUALITY CONTROL:

10. Calibrate daily with at least six working standards.

a. Add known aliquots of calibration stock solution to deionized water in 10-mL volumetric flasks and 

dilute to the mark. Use serial dilutions as needed to obtain chloroacetic acid concentrations in the 

range 0.02 to 40 μg/mL.

b. Analyze together with samples and blanks (steps 13 through 15).

c. Prepare calibration graph [peak height (mm or μS) vs. μg chloroacetic acid per sample].

11. Determine desorption efficiency (DE) for each batch of silica gel used for sampling in the calibration 

range. Prepare at least three tubes at each of five levels.

a. Place silica gel from unused front section in vial.

b. Inject a known amount (2 to 20 μL) of calibration stock solution, or a serial dilution thereof, onto 

front sorbent section with a microliter syringe.

c. Cap the vial. Allow to stand overnight.

d. Desorb (steps 7 through 9) and analyze together with working standards (steps 13 through 15).

e. Prepare graph of DE vs. μg chloroacetic acid recovered.

12. Analyze three quality control spikes and three analyst spikes to ensure that the calibration graph and 

DE graph are in control.

MEASUREMENT:

13. Set ion chromatograph according to manufacturer's recommendations and to conditions given on 

page 2008-1.

14. Inject sample aliquot manually or use autosampler.

a. Flush sample loop with 0.5 mL sample extract, then inject 0.5 mL sample.

b. Rinse sample loop with 1 to 2 mL deionized water between determinations of separate samples.

NOTE: All samples, eluents, and water flowing through the IC must be filtered to avoid plugging 

the system valves or columns.

15. Measure peak height.

NOTE: If sample peak height exceeds linear calibration range, dilute with deionized water, reanalyze, 

and apply appropriate dilution factor.

CALCULATIONS:

16. Determine mass, μg (corrected for DE), of analyte found in the sample front (Wf) and back (Wb) sorbent 

sections, and in the average media blank front (Bf) and back (Bb) sorbent sections.

17. Calculate concentration, C, of chloroacetic acid in the air volume sampled, V (L):

CC = WWff + WWbb − BBff − BBbb

VV mmmm mm3 ⁄

EVALUATION OF METHOD:

This method was developed and evaluated by Southern Research Institute [1] using dynamicallygenerated atmospheres of chloroacetic acid over the concentration range of 0.35 to 29 mg/m3 at 25 to 27 

°C and at relative humidity (RH) ≥80%. Average recovery based on 18 samples, six at each of three levels, 

was 98% representing a negligible bias. Precision at 0.35 mg/m3 was inhomogeneous with those of higher 

levels; therefore, precisions were not pooled. Using this poorest precision (SS�rr = 0.064), the overall precision 

(S�rT) was estimated to be ≤0.081. 

The breakthrough volume of the 100-mg sorbent section was found to be >100 L at 0.2 L/min when 

sampling chloroacetic acid concentrations of 60 mg/m3 at 42 °C and RH of 10 and 80% and 35 mg/m3 at 27 

°C and 10 and 90% RH. Samples stored at ambient temperature for 7 days had a mean recovery of 91% 
CHLOROACETIC ACID: METHOD 2008, Issue 2, dated 15 March 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

and a precision, SSrr, of 0.047. Samples refrigerated after day 7, and stored for 32 days exhibited a mean

recovery of 100% with a precision, SS�rr, of 0.085 based on samples analyzed on day 1.

�

REFERENCES:

[1] Dillon HK, Mason DW, Boyd KW [1980]. Development of air sampling and analytical methods for 

toxic chlorinated organic compounds: research report for monochloroacetic acid. Birmingham, AL: 

Southern Research Institute. NIOSH contract no. 210-78-0012. Unpublished.

[2] NIOSH [1980]. Chlorine: Method P&CAM 322. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for 

Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication 

No. 80-125.

[3] McCullough PR, Worley JW [1979]. Sampling of chloroacetyl chloride in air on solid support and 

determination by ion chromatography. Anal Chem 51:1120-1122.

[4] Budavari S, ed. [1989]. Merck Index, 11th ed. Rahway, NJ: Merck and Co. 

METHOD REVISED BY:

Mary Ellen Cassinelli, NIOSH

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

p-CHLOROPHENOL 2014

C6H4ClO MW: 128.56 CAS: 106-48-9 RTECS: SK2800000

METHOD: 2014, Issue 2 EVALUATION: FULL Issue 1: 19 September 1980

Issue 2: 25 February 2016

OSHA: None

NIOSH: None

PROPERTIES: crystals; d 1.224 g/mL @ 20 °C; MP 43.2-43.7 °C; 

BP 220 °C; VP 0.013 kPa (0.1 mm Hg) @ 20 °C;

flash point 121 °C (closed cup)

SYNONYMS: 4-chlorophenol; 4-chloro-1-hydroxybenzene

SAMPLING

SAMPLER: SORBENT TUBE (silica gel, 150 mg/75 mg)

FLOW RATE: 0.05 - 0.2 L/min

VOL-MIN: 1.5 L @ 1 ppm

 -MAX: 40 L

SHIPMENT: routine

SAMPLE

STABILITY: 7 days @ 25 °C at least 29 days @ 0 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 0.910 to 23.4 mg/m3 [1] (3-L samples)

BIAS: none identified

OVERALL

PRECISION (SS�rrrr): 0.061 for range studied [1]

ACCURACY: ± 15% (12-28%)

MEASUREMENT

TECHNIQUE: HPLC/UV

ANALYTE: p-chlorophenol

EXTRACTION: 1 mL acetonitrile

MOBILE 

PHASE: 30% acetonitrile, 70% water to 80%

acetonitrile/20% water in 20 minutes, 

1 mL/min

COLUMN: C18 (5 μm particle size, 4-mm ID by 30-

cm long, stainless steel)

DETECTOR: UV @ 280 nm

CALIBRATION: p-chlorophenol in 30% (v/v) acetonitrile in 

water

RANGE: 8 to 64 μg/sample [1]

ESTIMATED LOD: 2.5 μg/sample [1]

PRECISION (SS�rr): 0.024 [1]

APPLICABILITY: The working range is 0.15 to 53 ppm (0.8 to 280 mg/m3

) for a 10-L air sample.

INTERFERENCES: None identified. The chromatographic conditions described will separate phenol; o-chlorophenol; 2,3-, 2,4-, 

2,5-, 2,6-, 3,4-, and 3,5- dichlorophenol; o- and p-nitrophenol; 2,4-dimethylphenol; 2,4,5-trichlorophenol; 4-chloro-omethylphenol; 2,4-dinitrophenol; 4,6-dinitro-2-methylphenol; and pentachlorophenol.

OTHER METHODS: This method replaces P&CAM 337 [2]. The other columns for the analysis of p-chlorophenol have been 

reported in the literature [3-5].
p-CHLOROPHENOL: METHOD 2014, Issue 2, dated 25 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. p-Chlorophenol 99%.*

2. Acetonitrile, distilled in glass.

3. Hexane, distilled in glass.

4. Water, HPLC quality distilled, deionized.

5. p-Chlorophenol stock solution, 20 mg/mL.

Dissolve 500 mg p-chlorophenol in 30% (v/v)

acetonitrile in water to make 25 mL solution.

Stable at least 3 months in airtight container.

6. Desorption efficiency (DE) stock solution, 

5 mg/mL. Dissolve 125 mg p-chlorophenol 

in hexane to make 25 mL solution.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: borosilicate glass tubes, 7 cm long

with a 6-mm OD and a 4-mm ID, flame sealed 

at both ends. Each tube contains two sections 

of 20/40 mesh silica gel (a 150-mg sorbent

section and a 75-mg backup section 

separated and held in place with glass wool 

plugs). Tubes are commercially available. 

2. Personal sampling pump, calibrated, capable

of operating 8 hours at 0.05 to 0.2 L/min with 

flexible connecting tubing.

3. HPLC with UV detector (280 nm), C18 column,

injector, and electronic integrator.

4. Microliter syringes, various sizes.

5. Volumetric flasks, various sizes.

6. Centrifuge tubes, 12-mL, glass with screw

caps.

7. Pipette, 1- and 2-mL, and convenient sizes for

making dilutions.

8. Vials, 1-mL, with caps containing PTFE-lined

silicone septa.

9. Ultrasonic bath.

SPECIAL PRECAUTIONS: p-Chlorophenol is toxic by skin absorption, inhalation, or ingestion. It also is a 

strong irritant to tissue and is combustible with a flash point of 121 °C.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Immediately before sampling, break open the ends of the tube to provide openings that are at least 2-

mm in diameter. Attach sampler to personal sampling pump with flexible tubing.

3. Sample at an accurately known flowrate between 0.05 and 0.2 L/min for a total sample size of 1 to 40

liters.

4. Cap the tubes, record sample identity and all relevant sample data (duration, ambient temperature

and pressure). Pack securely for shipment.

NOTE: Refrigerate all samples at 0 °C when stored longer than 7 days.

SAMPLE PREPARATION:

5. If refrigerated, allow tube to equilibrate to room temperature.

6. Transfer each section of silica gel in a sorbent tube to a separate 12-mL centrifuge tube. Combine the

glass wool plug near the inlet with the front sorbent section. Combine the two urethane foam plugs

with the back section.

7. Add 1 mL of acetonitrile, cap, and desorb in an ultrasonic bath for 30 minutes.

8. Add 2 mL of distilled, deionized water to each tube, cap, and mix the solutions.

9. Centrifuge the samples and transfer about 1 mL of the supernatant in each tube to a separate vial and

seal with a PTFE-lined septum.
p-CHLOROPHENOL: METHOD 2014, Issue 2, dated 25 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALIBRATION AND QUALITY CONTROL:

10. Calibrate daily with at least six working standards in the range 2.5 to 64 μg per sample.

a. Dilute aliquots of p-chlorophenol stock solution with 30% (v/v) acetonitrile in water in volumetric 

flasks to encompass the range of interest. Prepare fresh daily.

b. Analyze working standards with samples and blanks steps.

c. Prepare calibration graph (peak area or peak height vs. μg of p-chlorophenol per sample.

11. Determine desorption efficiency (DE) for each lot of silica gel used for sampling in the calibration 

range. Prepare three tubes at each of five levels.

a. Remove backup section. Inject known amounts of DE stock solution (2 to 10 μL) onto the silica gel 

with a microliter syringe.

b. Cap the tubes and allow to stand overnight.

c. Desorb (steps 7 through 9) and analyze together with standards and blanks (steps 13 and 14).

d. Prepare a graph DE vs. μg p-chlorophenol recovered.

12. Analyze three quality control spikes and three analyst spikes to ensure that the calibration graph and 

DE graph are in control. 

MEASUREMENT:

13. Set HPLC according to manufacturer's recommendations and to conditions on page 2014-1. Inject 

sample aliquot manually or with autosampler.

NOTE: If peak is above the linear range of the working standards, dilute with 30% (v/v) acetonitrile in 

water, reanalyze, and apply the appropriate dilution factor.

14. Measure peak area or peak height.

CALCULATIONS:

15. Determine the mass, μg (corrected for DE), of analyte found on the sample front (Wf) and back (Wb) 

sorbent sections, and in the average media blank front (Bf) and back (Bb) sorbent sections.

16. Calculate concentration of p-chlorophenol in the air volume sampled, V (L):

CC = WWff + WWbb − BBff − BBbb

VV , mmmm/mm3

EVALUATION OF METHOD: 

The overall method was evaluated by collecting 3-L samples of test atmospheres containing pchlorophenol in the range of 0.91 - 23.4 mg/m3 at 29 °C and a relative humidity of greater than 80%. The 

amounts collected ranged from 2.6 - 64 μg per 150-mg bed of silica gel. The breakthrough volume of the 

sorbent tube was found to be approximately 60 L with a sampling rate of 0.2 L/min at a p-chlorophenol 

concentration of about 70 mg/m3

, a sampling temperature of 43 °C, and a relative humidity of greater than 

80%. Samples of p-chlorophenol on silica gel were found to be stable at 25 °C for 7 days and for 29 days if 

stored at 0 °C after the seventh day. Silica gel gave an average desorption efficiency of 96% with a SSrr 2.4% 

for loadings of 2.54 - 48.0 μg of p-chlorophenol on 150-mg beds of sorbent material. 

̅
p-CHLOROPHENOL: METHOD 2014, Issue 2, dated 25 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] Dillon HK, Emory MB [1980]. Development of air sampling and analytical methods for toxic 

chlorinated organic compounds: Research report for p-chlorophenol. NIOSH contract no. 210-78-

0012. Birmingham, AL. Southern Research Institute.

[2] NIOSH [1981]. P-Chlorophenol: P&CAM 337. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. Cincinnati, OH: U.S. Department of Health and Human Services, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 82-100.

[3] Korhonen IO [1980]. Separation of chlorophenol isomers on quartz columns. J Chromatogr 

303(1):197-205.

[4] Buisson RSK, Kirk PWW, Lester JN [1984]. Determination of chlorinated phenols in water, wastewater 

and sludge by capillary GC/ECD. J Chromatogr Sci 22(8):399-42.

[5] Lee HB, Hong-You RL, Fowlie PJ [1989]. Chemical derivatization analysis of phenols. Part VI. 

Determination of chlorinated phenolics in pulp and paper effluents. J Assoc Off Anal Chem 

72(6):979-984.

METHOD REVISED BY:

Charles Neumeister, NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

FORMALDEHYDE 2016

 H2C=O MW: 30.03 CAS: 50-00-0 RTECS: LP8925000

METHOD: 2016, Issue 3 EVALUATION: FULL Issue 1: 15 January 1998

Issue 3: 25 February 2016

OSHA: 0.75 ppm; 2 ppm STEL

NIOSH: 0.016 ppm; C 0.1 ppm; carcinogen 

(1 ppm = 1.23 mg/m3 @ NTP)

PROPERTIES: Gas; BP -19.5 °C; specific gravity 1.067 (air = 1); 

 explosive range 7 to 73% (v/v) in air

SYNONYMS: methanal; formalin (aqueous 30 to 60% w/v formaldehyde); methylene oxide

SAMPLING

SAMPLER: CARTRIDGE (Cartridge containing silica 

gel coated with 2,4-

dinitrophenylhydrazine)

FLOW RATE: 0.03 to 1.5 L/min

VOL-MIN: 1 L @ 0.25 mg/m3

 -MAX: 15 L @ 2.5 mg/m3

SHIPMENT: Place caps onto cartridge. Ship on ice.

SAMPLE

STABILITY: 34 days @ 5 °C [1]

BLANKS: 2 to 10 field blanks per set; 6 to 10 media 

blanks per set

ACCURACY

RANGE 

STUDIED: 0.025 to 2.45 mg/m3 (22-L samples) [2]

BIAS: +4.4%

OVERALL

PRECISION (ŜrT): 0.057 [1, 2]

ACCURACY: ±19.0%

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: 2,4-dinitrophenylhydrazone of 

formaldehyde

EXTRACTION: Elution with 10 mL of carbonyl-free 

acetonitrile

INJECTION 

 VOLUME: 20 μL

MOBILE 

 PHASE: 45% acetonitrile/55% water (v/v), 

1.3 mL/min

COLUMN: 3.9 x 150-mm, stainless steel, packed 

with 5-μm C18

DETECTOR: UV @ 360 nm

CALIBRATION: Samplers fortified with standard 

solutions of formaldehyde in water

RANGE: 0.23 to 37 μg per sample [1, 2]

ESTIMATED LOD: 0.07 μg/sample [1]

PRECISION (SS�rr): 0.032 @ 1.0 to 20.0 μg/sample [1]

APPLICABILITY: The working range is 0.015 to 2.5 mg/m3 (0.012 to 2.0 ppm) for a 15-L sample. This method can be used for the 

determination of formaldehyde for both STEL and TWA exposures [1, 2].

INTERFERENCES: Ozone has been observed to consume the 2,4-dinitrophenylhydrazine (2,4-DNPH) reagent and to degrade the 

formaldehyde derivative [3]. Ketones and other aldehydes can react with 2,4-DNPH; the derivatives produced, however, are 

separated chromatographically from the formaldehyde derivative. 

OTHER METHODS: NIOSH methods 2541 [4] and 3500 [5] and OSHA method 52 [6] are other methods for determination of 

formaldehyde in air. NIOSH method 5700 employs 2,4-DNPH and HPLC for determination of formaldehyde on textile or wood 

dust [7]. A journal method employs the same procedure for formaldehyde in automobile exhaust [8].
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 2 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Formaldehyde stock solution,* aqueous, 

standardized, 1 mg/mL (see APPENDIX A).

Alternatively, standardized formaldehyde 

solution, aqueous, 4 m g/mL (commercially 

available). 

2. Acetonitrile,* distilled in glass, low carbonyl 

content.**

3. Water, deionized and distilled.

4. Sulfuric acid, 0.02 N (pH standardization 

procedure) or 0.1 N (colorimetric procedure).

5. Sodium hydroxide, 0.01 N.

6. Sodium sulfite (Na2SO3), 1.13 M (pH

procedure) or 0.1 M (colorimetric procedure). 

Prepare fresh immediately before use. 

7. Thymophthalein indicator solution, 

0.04% (w/v) in 50:50 ethanol:water.

*See SPECIAL PRECAUTIONS. 

**Carbonyl content of acetonitrile can be 

determined by passing 10 mL of the solvent 

through a cartridge of DNPH-coated silica 

gel and analyzing by HPLC. Formaldehyde 

content should be below the LOD.

EQUIPMENT:

1. Sampler: Plastic holder containing 0.35 g of 

150-250 μm (60-100 mesh) silica gel coated 

with 1.0 mg of acidified 2,4- 

dinitrophenylhydrazine. Pressure drop across 

sampler should be less than 28 inches of 

water (7 kPa) at 1.5 L/min. Samplers are 

commercially available. (See APPENDIX B for 

additional information.) 

2. Personal sampling pump, 0 .03 to 1 .5 L/min 

with flexible connecting tubing.

3. Vials, 4-mL, glass with PTFE-lined rubber 

septa caps.

4. Vials, 20-mL, glass.

NOTE: Do not use vials with “polycone” liners 

(sources of high formaldehyde blanks) [5, 9].

5. Liquid chromatograph with UV detector, 

recorder, integrator, and column (page 2016-

1).

6. Syringes, 100-μL, 500-μL and 10-mL.

7. Volumetric flasks, 10-mL, 25-mL, and 1-L.

8. Burets, 50-mL.

9. pH meter.

10. Magnetic stirrer.

11. Beaker, 50-mL.

12. Flask, Erlenmeyer, 250-mL.

13. Ozone scrubber (optional).

14. Aluminum foil or black electrical tape 

(optional).

 

SPECIAL PRECAUTIONS: Formaldehyde is a suspect carcinogen and a proven human sensitizer; it should 

be handled in a fume hood [10-12]. Acetonitrile is toxic and is a fire hazard (flash point = 12.8 °C). 

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler (and ozone scrubber, if used) in 

line.

2. Open sampler packet and remove end caps.

3. Attach sampler to the sampling pump with flexible tubing. Bi-directional samplers can be connected 

at either end. 

NOTE: The sampler does not have a backup section for determination of breakthrough. If high 

concentrations of aldehydes and ketones are anticipated, connect two samplers in series. The 

back pressure of the sampling train will be higher and a lower flow rate may be required.

4. Sample 1 to 15 L of air at 0.03 to 1.5 L/min.

NOTE: To protect from intense light, such as bright sunlight, the sampler can be wrapped with 

aluminum foil or electrical tape.

5. Place end caps onto the sampler and seal sampler in an envelope. Protect samples from heat.

6. Ship samples on ice (0 °C).
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 3 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION: 

NOTE: Check acetonitrile for formaldehyde content by elution and analysis of a blank cartridge; the 

formaldehyde level should be below the detection limit. Since background levels of 

formaldehyde on the samplers may change during storage, compare samples with sampler 

blanks from the same lot. Samples and blanks should be stored under the same conditions.

7. Elute the formaldehyde derivative from the cartridge samplers with 10-mL quantities of acetonitrile.

a. Collect effluent from each sampler in a 10-mL volumetric flask.

b. Add acetonitrile to the mark for each sampler.

NOTE: The silica gel bed of the sampler will retain approximately 0.5 mL of the original 10 mL.

CALIBRATION AND QUALITY CONTROL: 

8. Calibrate daily with at least six media working standards over the range of interest.

a. Prepare a series of aqueous formaldehyde solutions for the fortification of samplers. Suggested 

concentrations include 1, 4, and 20 μg/mL. See APPENDIX A for standardization of formaldehyde in 

water.

b. Connect the outlet of a cartridge sampler to a personal sampling pump with flexible tubing. Turn 

on the pump and make sure there is a flow of air through the sampler.

c. Load a 100-μL syringe with a selected volume of aqueous formaldehyde solution in the range of 30 

to 90 μL. Suggested quantities of formaldehyde for spiking include 0.04, 0.10, 0.20, 0.30, 0.40, 0.80, 

1.0 and 2.0 μg/sample.

d. Place the tip of the syringe needle against the frit in the inlet of the sampler and eject the 

formaldehyde solution.

e. Prepare the media working standard (steps 7a and 7b).

f. Prepare additional working standards (steps 8b through 8e).

g. Transfer 3-mL aliquots of working standards to 4-mL vials, and analyze (steps 10, 12 and 13).

h. Prepare calibration graph, peak area or peak height vs. μg formaldehyde per sample.

9. Fortify and analyze three quality control spikes and three analyst spikes to ensure that calibration 

graph is in control.

MEASUREMENT:

10. Set liquid chromatograph according to manufacturer’s recommendations and to conditions given on 

page 2016-1.

11. Transfer a 3-mL aliquot of the sample solution from step 7 to a 4-mL vial. Cap the vial.

12. Inject a 20-μL sample aliquot.

13. Measure peak area or peak height.

NOTE 1: If sample peak is larger than the largest standard peak, dilute an aliquot of the remaining 

sample solution, reanalyze, and apply appropriate dilution factor in the calculations.

NOTE 2: To ensure validity of the samples, identify those samples which contain more than 37 μg of 

formaldehyde. The capacity of the samplers before breakthrough may have been exceeded 

for these samples, and collection of smaller samples would be warranted.

NOTE 3: The size of the 2,4-DNPH peak should be about 2.7 times the size of the formaldehyde-DNPH 

peak or larger. Otherwise, breakthrough from the sampler may have occurred.

CALCULATIONS:

14. Determine mass, μg, of formaldehyde, W, found in the sample and the average media blank, B, from 

the calibration graph.
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 4 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

15. Calculate concentration, C, of formaldehyde in the air volume sampled, V (L).

CC = 

WW − BB

VV , mmmm/mm3

NOTE: μg/L ≌ mg/m3

EVALUATION OF METHOD:

Issue 1

This method was originally evaluated with Waters Sep-Pak XPoSure Aldehyde samplers using data 

produced at NIOSH and at Waters Corporation [2]. Test atmospheres of formaldehyde were generated at 

Waters Corp. [2]. Overall measurement precision, ŜrT, was 0.057 based on NIOSH guidelines [13] including 

a 5% pump error factor and estimated bias of +4.4%. Sample storage stability was evaluated over the range 

of 0.5 to 55 μg formaldehyde/sample. Losses for Waters samplers were 4 to 8% when stored up to 14 days 

at 4 °C. An additional study with Waters samplers found that losses were 5% or less after 4 days of storage 

at ambient temperature. All calibration standards used at Waters Corporation were liquid standard 

solutions of formaldehyde-DNPH derivative in acetonitrile [2, 14]. 

The capacity of DNPH-coated silica gel samplers was found to vary with relative humidity (RH) in addition 

to concentration of formaldehyde. At a formaldehyde concentration of 1.2 m g/m3 and at 5% 

breakthrough, the Waters sampler had a capacity at <10% RH of 55 μg, and at >85% RH a capacity of 77 μg. 

At 2.4 mg/m3 and <10% RH, the 5% breakthrough capacity of the Waters sampler was 59 μg of 

formaldehyde. At 2.6 mg/m3 and >85% RH, the 5% breakthrough capacity was 106 μg. Thus, the smallest 

capacity at 5% breakthrough was 55 μg of formaldehyde; the upper limit of the range of the method is two 

thirds of 55 μg, or 37 μg. Capacity information for the Waters sampler is applicable to the Supelco sampler 

because (a) the Waters and Supelco samplers contain 0.9 and 1 mg of DNPH, respectively, and (b) each 

sampler contains 350 mg of silica gel.

Issue 2

In subsequent work on this method, additional formaldehyde samplers were evaluated, Supelco S10 

LpDNPH cartridges and SKC, Inc. Aldehyde samplers (DNPH-coated silica gel tubes No.226-119) [1]. The 

sorbent beds of Supelco and Waters cartridges and front sections of SKC samplers were treated with 

acetonitrile. Formaldehyde was not detected on any blank sampler (LOD = 0.01 μg/mL). The SKC sampler 

for aldehydes may be used for formaldehyde with modifications of this method (See APPENDIX B). 

However, evaluation of the SKC sampler at NIOSH has been limited.

Supelco samplers were fortified with known quantities of free formaldehyde in water, and calibration was 

performed with media standards prepared from Supelco samplers fortified with known quantities of free 

formaldehyde in water; average recoveries ranged from 96.3% to 99.3% for five levels at 1.00 to 20 μg of 

formaldehyde per sampler (pooled S�r = 0.0316; n = 6 at each level).

In a storage study, six Supelco samplers were fortified with 4-μg quantities of free formaldehyde in water. 

Samplers were stored 34 days at 5 °C in the dark; the average recovery based on media standards was 99% 

(S�r = 0.014).

Eight media standards were prepared by fortification of Supelco samplers with solutions of free 

formaldehyde in water. The solutions were drawn through the DNPH-coated silica gel beds with an air 

pump. The resulting LOD and LOQ were 0.07 and 0.23 μg/sample, respectively, according to a least squares 

calibration graph.
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 5 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Standard solutions of formaldehyde-DNPH at 0.10 and 1.0 μg/mL (formaldehyde equivalent 

concentrations) were stored in airtight vials at 5 °C in the dark and were analyzed periodically. The 

standard solutions were found to be stable (with no detectable loss) for at least 10 weeks and at least 12 

weeks, respectively.

It is suggested that the reader see the Backup Data Report for a comparison of media standards with liquid 

standards for calibration [1]. Air sampling for a 24-hour period can be performed with a single Supelco 

sampler. Thus, background levels at <1 ppb can be determined.

REFERENCES:

[1] Tucker SP [2002]. Formaldehyde backup data report for method 2016 issue 2. Cincinnati, OH: 

National Institute for Occupational Safety and Health. Unpublished.

[2] Iraneta PC, Collamati RA, Costello DN, Crowley RJ, Fernandes RL, Hopkins MR, Knowles CE, Martin DM 

[1995]. A validation study of a new active sampler for the analysis of formaldehyde in workplace and 

indoor air. Milford, MA: Waters Corp. Unpublished.

[3] Kleindienst TE, Corse EW, Blanchard FT [1998]. Evaluation of the performance of DNPH-coated silica 

gel and C18 cartridges in the measurement of formaldehyde in the presence and absence of ozone. 

Environ Sci Technol 32(1):124-130.

[4] NIOSH [1994]. Formaldehyde by GC: Method 2541. In: Eller PM, Cassinelli ME, eds. NIOSH manual of 

analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers 

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 94-113.

[5] NIOSH [1994]. Formaldehyde by VIS: Method 3500. In: Eller PM, Cassinelli ME, eds. NIOSH manual of 

analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers 

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 94-113.

[6] OSHA [1985]. Formaldehyde: Method 52. In: OSHA analytical methods manual. Salt Lake City, UT: 

U.S. Department of Labor, Occupational Safety and Health Administration, OSHA Analytical 

Laboratory. 

[7] NIOSH [1994]. Formaldehyde on dust (textile or wood): Method 5700. In: Eller PM, Cassinelli ME, eds. 

NIOSH Manual of Analytical Methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and

Health, DHHS (NIOSH) Publication No. 94-113.

[8] Lipari F, Swarin S [1982]. Determination of formaldehyde and other aldehydes in automobile exhaust 

with an improved 2,4-Dinitrophenylhydrazine method. J Chromatogr 247:297-306.

[9] Dollberg DD, Smith DL, Tharr DG [1986]. Lab/Field coordination. Appl Ind Hyg 1:F44-F46.

[10] ACGIH [2001]. 2001 Threshold limit values® and biological exposure indices. Cincinnati, OH. 

American Conference of Governmental Industrial Hygienists, Inc.

[11] ACGIH [1991]. Formaldehyde. In: Documentation of Threshold Limit Values and Biological Exposure 

Indices, 6th ed. Cincinnati, OH: American Conference of Governmental Industrial Hygienists, Inc.

[12] CFR. Code of Federal Regulations [1999]. Formaldhyde. In: Title 29, Part 1910.1048, 29 CFR Chapter 

XVII (7-1-99 Edition), 354-378. Washington, DC: U.S. Government Printing Office, Office of the 

Federal Register.
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 6 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[13] Kennedy ER, Fischbach TJ, Song R, Eller PE, Shulman SA [1995]. Guidelines for air sampling and 

analytical method development and evaluation. Cincinnati, OH: US Department of Health and 

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health. DHHS (NIOSH) Publication No. 95-117.

[14] Waters Corp. [1994]. Waters Sep-Pak XPoSure aldehyde sampler care and use manual. Milford, MA. 

[15] Walker JF [1964]. Formaldehyde. In: American chemical society monograph series. 3rd ed. Chapter 

18. Reinhold Publishing Corporation, New York. 

METHOD REVISED BY:

Samuel P. Tucker, Ph.D., NIOSH. 

Method originally written by P Iraneta, Waters Corp., MJ Seymour and ER Kennedy, Ph.D., NIOSH.
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 7 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPENDIX A -PREPARATION AND STANDARDIZATION OF FORMALDEHYDE STOCK SOLUTION (ca. 1 

mg/mL)

Preparation. Dilute 2.7 mL 37% aqueous formalin solution to 1 L with distilled, deionized water. This 

solution is stable for at least three months when stored at room temperature.

Standardization by pH Titration. Place 5.0 mL of freshly prepared 1.13 M sodium sulfite solution in a 50-

mL beaker and stir magnetically. Adjust pH to between 8.5 and 10 with base or acid. Record the pH. Add 3.0 

to 12.0 mL formaldehyde stock solution. The pH should now be greater than 11. Titrate the solution back to 

its original pH with 0.02 N sulfuric acid (1 mL acid = 0.600 mg formaldehyde; about 17 mL acid needed). If 

the endpoint pH is overrun, back-titrate to the endpoint with 0.01 N sodium hydroxide. Calculate the 

concentration, Cs (mg/mL), of the form aldehyde stock solution:

CCss = 30.0(NNaaVVaa − NNbbVVbb)

VVss

, mmmm/mmmm

Where:

30.0 = 30.0 g/equivalent of formaldehyde

Na = normality of sulfuric acid (0.02 N)

Va = volume of sulfuric acid (mL) used for titration

Nb = normality of NaOH (0.01 N)

Vb = volume of NaOH (mL) used for back-titration 

Vs = volume of formaldehyde stock solution (mL)

Standardization by Colorimetric Titration. Place 50 mL of freshly prepared 0.1 M sodium sulfite and 3 

drops of 0.04% thymophthalein indicator (w/v) in 50:50 ethanol:water into a 250-mL Erlenmeyer flask. 

Titrate the contents of the flask to a colorless endpoint with 0.1 N sulfuric acid (usually 1 or 2 drops is 

sufficient). The indicator is blue at pH values above the endpoint and is colorless at pH values below the 

endpoint. Transfer 3.0 to 12.0 mL of the formaldehyde solution to the same flask and titrate the mixture 

with 0.1 N sulfuric acid to a colorless endpoint. Calculate the concentration, C (mg/mL), of formaldehyde in 

solution.

CC = 30.0(NNaaVVaa)

VVss

, mmmm/mmmm

Where:

30.0 = g/equivalent of formaldehyde

Na = normality of sulfuric acid (0.1 N)

Va = volume of sulfuric acid used for titration (mL) 

Vs = volume of formaldehyde stock solution (mL)

NOTE: Sulfuric acid (0.1 N) is substituted for 0.1 N hydrochloric acid, which is specified in OSHA Method 52, 

in order to prevent possible formation of bis(chloromethyl)ether, a potent carcinogen, by reaction of 

formaldehyde with hydrochloric acid [12 ].

This colorimetric titration was adapted from OSHA Method 52 [5], which was based on the procedure of 

Walker [15].
FORMALDEHYDE: METHOD 2016, Issue 3, dated 25 February 2016 - Page 8 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPENDIX B - USE OF SKC SAMPLER FOR FORMALDEHYDE

DNPH-coated silica gel tubes may be used for sampling formaldehyde with modifications of this method. 

These modifications include the following:

(a) The maximum recommended air volume should be less than 15 L at an air concentration of 2.5 mg/m3

(indicated on page 2016-1), because the upper limit of the method for the SKC sampler is probably less 

than 37 μg.

(b)The procedure for recovery of analyte from the sorbent would be modified, i.e., placement of the 

sorbent sections in vials, addition of solvent, and possible use of an ultrasonic bath.

(c) A volume of solvent much less than 10 mL can be used for recovery. However, the minimum volume 

should be tested for adequate recovery.

(d)Consequences of using a much smaller volume of solvent for recovery include a lower LOD and LOQ, 

the need for a different range of calibration standards, and the need for a different range of fortification 

levels (step 8).

(e) The maximum volume of solution for fortification of the front sorbent bed must be less than 90 μL and 

should be determined (step 8).

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

KETONES 2027

Formula: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1 

METHOD: 2027, Issue 1 EVALUATION: FULL Issue 1: 19 July 2016

OSHA: Table 2

NIOSH: Table 2 

Other OELs: Refs [1,2}

PROPERTIES: Table 1 

SYNONYMS: See individual compounds in Table 1

SAMPLING

SAMPLER: SOLID SORBENT TUBE (silica gel, 500 

mg/1000 mg); min. of 2 field blanks per set

FLOW RATE: 0.05 – 0.1 l/min

VOL-MIN: 2 L

 -MAX: 10 L

SHIPMENT: routine

SAMPLE

STABILITY: Stable at least 28 days @ 20 °C [3,4]

ACCURACY

RANGE 

STUDIED: Table 3 

BIAS: Negligible

OVERALL

PRECISION (SS�rrrr): Table 3

UNCERTAINTY: Table 3 

MEASUREMENT

TECHNIQUE: gas chromatography, FID

ANALYTE: see Table 1

DESORPTION: ternary mixture of 

CH2Cl2/methanol/water (65:33:2)

INJECTION

 VOLUME: 1 μL

COLUMNS

(IN PARALLEL): 60 m low-polarity fused silica, ID 0.25 

mm and film thickness 0.25 μm; and 

60 m high-polarity polyethylene glycol, 

ID 0.25 mm and film thickness 0.25 μm. 

TEMPERATURE:

 INJECTION: 250 °C

 DETECTOR: 250 °C

 COLUMN: 50 °C (11 min) to 150 °C (4 °C/min)

CALIBRATION: solution of analytes in desorption 

solvent

RANGE: Table 3

ESTIMATED LOD: Table 3

PRECISION (SS̅

rr): Table 3 

APPLICABILITY: 2-2400 mg/m3 for air sample vol. up to 10 L.

INTERFERENCES: Organic compounds with similar retention times as the analytes of interest may interfere.

OTHER METHODS: NIOSH 2553 and 2555 [5] are partially validated methods for the determination of ketones using CS2 for 

desorption. OSHA methods based on sorbent tube sampling, solvent desorption and GC/FID analysis have been promulgated for 

several ketones [6].
KETONES: METHOD 2027, Issue 1, dated 17 July 2016 - Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Acetone, >99.8%, analytical grade.

2. 2-Butanone, >99.8%, analytical grade.

3. Cyclohexanone, >99.5%, analytical grade.

4. Cyclopentanone, >99.0% purity.

5. 2-Hexanone, >98 % purity.

6. 4-Methyl -2-pentanone, >99.0%, analytical 

grade.

7. 2,6-Dimethyl-4-heptanone, ≥99%, analytical 

grade.*

8. Methanol, >99.9 %, analytical grade.

9. Dichloromethane, analytical grade.

10. Water, ultrapure, >18.2 MΩ-cm resistivity.

11. 2-Methylheptane, chromatographic quality.

12. n-Undecane, chromatographic quality.

13. Air, prepurified and filtered.

14. Hydrogen, prepurified and filtered.

15. Helium, prepurified and filtered.

16. Nitrogen, prepurified and filtered.

17. Internal standard solution: Mix 500 μL of nundecane and 500 μL 2-methylheptane.

18. Ternary mixture: Mix 650 mL 

dichloromethane, 330 mL methanol and 20 

mL water in a 1000-mL- volumetric flask.

*NOTE: Each new bottle must be analyzed 

immediately after opening to accurately 

measure the concentration.

EQUIPMENT

1. Sampler: glass tube, length 125 mm, 7-mm 

OD, 5-mm ID, flame sealed ends, containing 

two sections of silica gel (front 500 mg, back 

1000 mg) separated by a 2-mm ceramic plug. 

The ceramic plug is in front and a silylated 

glass wool plug follows the back section. 

Tubes are commercially available.

2. Personal sampling pump capable of 

producing flow rates between 0.05 to 0.1 

L/min, with flexible connecting tubing. 

3. Gas chromatograph, equipped with split 

injector (capillary flow split), two separation 

columns with separate FIDs, and data 

processing unit.

4. Fused silica capillary columns (for parallel 

separation): 60 m low-polarity fused silica, 

0.25 mm ID and film thickness 0.25 μm. 60 m 

high-polarity polyethylene glycol, 0.25 mm 

ID and film thickness 0.25 μm.

5. Vessels, 15-mL, glass, with screw caps and 

PTFE septum.

6. Volumetric flasks, 5, 50 and 1000 mL.

7. Syringes, glass, 10 to 500 μL.

8. Pipettes, glass, 5 and 10 mL.

9. Bottles, polyethylene, 100 mL.

10. Syringes, polypropylene, 2 mL.

11. Syringe filter cartridges, with 0.45-μm pore 

size polytetrafluoroethylene (PTFE) 

membrane filters.

12. Micro-syringes, 50 μL, with 60 mm x 0.6 mm 

needles.

13. Auto-sampler vials, with PTFE septum, 1.5 mL 

capacity.

SPECIAL PRECAUTIONS: Analytes should be handled in a fume hood. Wear protective clothing and 

eyewear. Dichloromethane is a potential occupational carcinogen [12].

SAMPLING, SAMPLE TRANSPORT AND STORAGE

1. Calibrate each personal sampling pump with a representative sampler in the line.

2. Break the ends of the sampler immediately before sampling. Attach sampler to the personal 

sampling pump with flexible tubing.

3. Sample at an accurate known flow rate between 0.05 and 0.1 L/min for a total sample size of not 

more than 10 L.

4. Cap the samplers with plastic (not rubber) caps and pack securely for shipment.

5. Analyze samples within 28 days of receipt.
KETONES: METHOD 2027, Issue 1, dated 17 July 2016 - Page 3 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION 

6. Place the entire contents of the sorbent tube in a 15-mL screw-cap-vessel.

7. Add 5 mL ternary mixture and securely cap the vessel immediately.

8. Allow to stand at least 18 hours with occasional agitation.

9. Add 2.5 μl of internal standard solution.

10. Take sample solution with a 2-ml-polypropylene syringe and filter through a PTFE filter into an auto 

sampler vial.

CALIBRATION AND QUALITY CONTROL 

11. Calibrate with at least six standards for each substance in the range given in Table 3. 

a. Place the contents of a sorbent tube in a 15-mL screw-cap-vessel and add 5 mL ternary mixture 

solvent. 

b. Add known amount of analyte. 

c. Store at room temperature for 18 hours and then add 2.5 μL of internal standard solution. 

Note: Internal standards are used to aid in identification of retention times of unknowns. 

d. Analyze and prepare a calibration graph for the two columns separately (peak areas of analyte 

and internal standard (nonpolar column:2-methylheptane; polar column: n-undecane) vs 

concentration).

12. Prepare each day at least three quality control solutions in the same manner and range and analyze 

together with samples and blanks.

13. Determine desorption efficiency (DE) at least once for each batch of sorbent used for sampling in the 

calibration range. 

a. Prepare three tubes at each of five levels plus three media blanks. 

b. Inject a known amount of analyte solution directly onto the front sorbent section while drawing

purified air through the tube. 

c. Draw approximately 10 L air through the sorbent tube, then cap the ends of each tube and allow 

to stand overnight.

d. Desorb (steps 6 to 9) and analyze together with quality control samples and blanks (steps 14 to 

15). 

14. Analyze a minimum of three quality control spikes and three analyst spikes to insure that the 

calibration graph and the DE graph are in control.

MEASUREMENT 

15. Set gas chromatograph parameters to manufacturer’s recommendations and to conditions give on 

Page 1. Inject a 1-μL sample aliquot into the gas chromatograph. 

NOTE: If the peak area exceeds the linear calibration range, dilute with desorption solvent, reanalyze 

and apply the appropriate dilution factor in calculations. 

16. Measure analyte peak areas at applicable retention times (Table 4). 

CALCULATIONS:

17. Calculate the mass concentration of each analyte, C (mg/m3), in the air volume sampled, V (L): 

CC = (CC1 ∙ VV1 ∙ FFd) − (CC0 ∙ VV0)

VV ∙ ηη
KETONES: METHOD 2027, Issue 1, dated 17 July 2016 - Page 4 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

where: 

C0 = mean concentration, in mg/L, of analyte in the field blank test solutions;

C1 = concentration, in mg/L, of analyte in the sample test solution;

V = volume, in liters, of the air sample;

V0 = volume, in mL, of the field blank test solutions; 

V1 = volume, in mL, of the sample test solutions;

Fd = dilution factor for each sample test solution;

η = desorption efficiency.

EVALUATION OF METHOD:

The method recovery, at levels from the LOQ to at least 2 times the REL, was determined for each analyte 

by carrying out sampling and analytical experiments at a dynamic test gas facility [3,4]. Experiments were 

conducted at four concentration levels of each analyte and for relative humidities of 20%, 50% and 80% at 

20 °C. All analytes exhibited method recovery >95 % at the levels evaluated (see Table 3). At low relative 

humidity (<30%) the recoveries of cyclohexanone, cyclopentanone and 2-hexanone were reduced 

without water in the extraction solution. Use of a small content of water (2%) during extraction avoids 

losses based on low humidity during sampling. Each analyte, at levels below and above the REL, was 

evaluated for its storage stability [3,4]. The samples were prepared by sampling at a dynamic test gas 

device at a relative humidity of 50% (20 °C). The samples were then stored at room temperature for four 

weeks. All analytes demonstrated acceptable recoveries (>90%) after 28 days of storage [3]. The 

performance of the method was separately verified in independent laboratories [4,13].

REFERENCES:

[1] American Conference of Governmental Industrial Hygienists (ACGIH), Threshold Limit Values for

Chemical Substances and Physical Agents & Biological Exposure Indices. ACGIH: Cincinnati (2014);

updated annually.

[2] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA), GESTIS – Database on

International Limit Values for Hazardous Substances (German Social Accident Insurances). IFA: Sankt

Augustin, Germany (2014); https://www.dguv.de/ifa/index-2.jsp.

[3] Breuer, D.: Ketones, IFA Working Folder, Sheet No. 7708; Institute for Occupational Safety and Health

of the German Social Accident Insurances. Erich Schmidt Verlag: Bielefeld, Germany (2005).

[4] Breuer, D, Eisenhardt, A. Solvent mixtures, Method No. 4 (Ketones); in Air Monitoring Methods, The

MAK Collection for Occupational Health and Safety. Wiley-VCH: Weinheim, Germany (2013).

[5] NIOSH Manual of Analytical Methods, 4th ed., Method Nos. 2555 (Ketones I) and 2553 (Ketones II).

CDC/NIOSH: Cincinnati (2003).

[6] Occupational Safety and Health Administration (OSHA), Sampling and Analytical Methods (Method

nos. 1, 16, 69, PV2031). OSHA Salt Lake Technical Center, Sandy, UT;

www.osha.gov/dts/sltc/methods/index.html (accessed May 2015). 
KETONES: METHOD 2027, Issue 1, dated 17 July 2016 - Page 5 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[7] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA), GESTIS – substance

database (German Social Accident Insurances). IFA: Sankt Augustin, Germany (2014);

https://www.dguv.de/ifa/index-2.jsp.

[8] Sigma-Aldrich [2011]. 2012-2014 Handbook of Fine Chemicals. St. Louis, MO: Sigma-Aldrich Co.

[9] NIOSH Pocket Guide to Chemical Hazards, DHHS (NIOSH), Cincinnati; www.cdc.gov/niosh/npg;

(accessed June 2015).

[10] EN 482: Workplace exposure – General requirements for the performance of procedures for the

measurement of chemical agents. CEN: Brussels (2012).

[11] EN 1076: Workplace exposure – Procedures for measuring gases and vapours using pumped

samplers – Requirements and test methods. CEN: Brussels (2009).

[12] NIOSH [1986]. Current intelligence bulletin 46: Methylene Chloride, U.S. Department of Health and

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health, DHHS (NIOSH) Publication No. 86-114.

[13] Bureau Veritas North America [2016]. Ketones User Check Sequence 11762-CA (unpublished report).

BVNA: Novi, MI, 3 Feb. 2016. 

METHOD WRITTEN BY:

Dietmar Breuer, Dr. rer. nat. and Anna Eisenhardt, Institute for Occupational Safety and Health of the 

German Social Accident Insurances (IFA), Sankt Augustin, Germany. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
KETONES: METHOD 2027, Issue 1, dated 17 July 2016 - Page 6 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 1. Synonyms, Formulae, Molecular weights, Properties, CAS#, RTECS

Compound/

synonyms

CAS#

RTECS

Empirical 

formula

Molecular 

weight 

Boiling

pointa

(°C)

Melting 

pointa

(°C)

Vapor

pressure 

@ 20°C

(kPa)a

Vapor 

pressure 

@ 20°C 

(mm Hg)a 

Density

@ 20 °Ca

(g/mL)

Acetone /

2-Propanone

67-64-1 

AL3150000

C3H6O 58.08 56.2 -95 24.6 184 0.79

2-Butanone

Methylethyl ketone

78-93-3 

EL6475000

C4H8O 72.11 79.6 -86 10.5 78.8 0.81

Cyclohexanone /

Cyclohexyl ketone

108-94-1 

GW1050000

C6H10O 98.14 155.6 -26 0.455 3.41 0.95

Cyclopentanone /

Ketocyclopentane

120-92-3 

GY4725000

C5H8O 84.12 130.6 -58.2 1.14 8.55 0.95

2-Hexanone /

Butyl methyl ketone

591-78-6 

MP1400000

C6H12O 100.16 127.2 -56 1.28 9.60 0.81

4-Methyl-2-

pentanone / 

MIBK/Methyl 

isobutyl ketone 

108-10-1 

SA9275000

C6H12O 100.16 116.8 -80.3 1.88 14.1 0.80

2,6-Dimethyl-4-

heptanone/ 

Diisobutyl ketone

108-83-8 

MJ5775000

C9H18O 142.24 168 -46 0.23b 1.7b 0.81

a Physical and chemical property data from GESTIS substance database, except where noted otherwise [7]

b Vapor pressure for 2,6-dimethyl-4-heptanone [8]

Table 2. Occupational exposure limits, ppm [9]

Substance OSHA PELs NIOSH RELs

TWA TWA STEL mg/m3 per ppm 

Acetone 1000 250 2.41

2-Butanone 200 200 300 2.95

Cyclohexanone 50 25 4.08

Cyclopentanone 3.50

2-Hexanone 100 1 4.16

4-Methyl-2-pentanone 100 50 75 4.16

2,6-Dimethyl4-heptanone 50 25 5.91
KETONES: METHOD 2027, Issue 1, dated 17 July 2016 - Page 7 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 3. Measurement range, precision, uncertainty and recovery (N=6)

Substance LOD LOQ Measurement

(mg/sample) (mg/sample) Range Precision Ua

 Mean 

(mg/sample) (Sr) (%) Recovery 

(DE)

Acetone 0.006 0.02 0.02 – 26 0.0083-

0.024

11 – 12 0.97

2-Butanone 0.006 0.02 0.02 – 14 0.0078-

0.024

11 – 12 0.98

Cyclohexanone 0.006 0.02 0.02 – 12 0.010-

0.022

12 – 13 0.97

Cyclopentanone 0.006 0.02 0.02 – 2 0.008-

0.023

11 – 12 0.95

2-Hexanone 0.006 0.02 0.02 – 0.47 0.018-

0.022

11 – 12 0.98

4-Methyl2-

pentanone

0.006 0.02 0.02 – 2 0.011-

0.023

11 – 12 0.99

2,6-Dimethyl-4-

heptanone 

0.005 0.02 0.02 – 5 0.006-

0.023

11 – 12 1.01

a Expanded uncertainty calculated in accordance with EN 482 [10] and EN 1076 [11]

Table 4. Approximate retention times on polar and nonpolar separation columns

Substance Approximate retention time 

non-polar column

Approximate retention time 

polar column 

(min) (min)

Acetone 6.5 7.1

2-Butanone 9.2 a) 

Cyclohexanone 25.5 25.9

Cyclopentanone 20.3 21.2

2-Hexanone 20.0 17.3

4-Methyl-2-pentanone 17.2 13.8

2,6-Dimethyl4-

heptanone

28.4 22.0

a) The retention time of butanone is equal to that of the solvent on the polar column. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ANISIDINE 2514

CH3OC6H MW: 123.16 CAS: (o-) 90-04-0 RTECS: (o-) BZ5410000 (p-) 104-94-9 (p-) BZ5450000

METHOD: 2514, Issue 3 EVALUATION: FULL Issue 1: 15 May 1985 

Issue 3: 26 February 2016

OSHA: 0.5 mg/m3 (skin)

NIOSH: 0.5 mg/m3 (skin);

o-isomer suspect carcinogen

PROPERTIES: o-isomer: liquid; d 1.092 g/mL @ 15 °C;

BP 225 °C; MP 5 °C; VP 0.1 mm Hg @ 27 °C. 

p-isomer: solid; MP 57 °C; BP 246 °C

SYNONYMS: o-isomer: 2-aminoanisole; 2-methoxybenzenamine; o-methoxyaniline; 

p-isomer: 4-aminoanisole; 4-methoxybenzenamine; p-methoxyaniline

SAMPLING

SAMPLER: SOLID SORBENT TUBE

(XAD-2, 150 mg/75 mg) 

FLOW RATE: 0.5 – 1.0 L/min

VOL-MIN:

 MAX:

24 L

320 L

SHIPMENT: routine

SAMPLE

STABILITY: at least 1 week @ 25 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 0.13 to 0.58 mg/m3 [1] (225-L samples)

BIAS: 0.12% 

OVERALL

PRECISION (SS�rrrr): 0.068 [1] 

ACCURACY: ± 13.3% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: o-anisidine and p-anisidine

DESORPTION: 5 mL methanol; stand 15 min

INJECTION

 VOLUME: 10 μL

MOBILE PHASE: 5% acetonitrile/65% water @ 1.2 

mL/min; ambient temperature

COLUMN: 50 cm x 2-mm ID stainless steel packed 

with C18

DETECTOR: UV absorption @ 254 nm

CALIBRATION: analytes dissolved in methanol

RANGE: 12 to 360 μg per sample (each isomer) 

[2]

ESTIMATED LOD: 0.35 μg per sample [2]

PRECISION (SS�rr): 0.029 @ 30 to 240 μg per sample [1] 

HPLC/UV

APPLICABILITY: The working range is 0.06 to 0.8 mg/m3 (0.012 to 0.16 ppm) for a 200-L air sample

INTERFERENCES: None identified

OTHER METHODS: This revises Method S163 [2]. This method replaces P&CAM 168 [3] because XAD-2 has a much greater 

capacity than silica gel for o-anisidine at high humidity [1].
ANISIDINE: METHOD 2514, Issue 3, dated 26 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. p-Anisidine, reagent grade.*

2. o-Anisidine, reagent grade.*

3. Methanol, HPLC grade.*

4. Acetonitrile, HPLC grade.*

5. Water, distilled, deionized.

6. Methylene chloride.

7. Calibration stock solution, 15.0 mg/mL

p-anisidine and 15.3 mg/mL o-anisidine.

Dissolve 750 mg p-anisidine and 700 μL

o-anisidine in methanol to make 50 mL

solution.

8. HPLC mobile phase:

35% acetonitrile/65% water. Filter (5-μm

PTFE) and degas prior to use.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: glass tube, 7-cm long, 8-mm OD,

6-mm ID, with plastic caps, containing two

sections of 20/50 mesh XAD-2 (150 mg front

and 75 mg back) separated and held in place

by plugs of silylated glass wool. Pressure

drop across the tube must be <3.4 kPa

(2.5 cm Hg) at 1 L/min airflow.

2. Personal sampling pump, 0.5 to 1 L/min,

with flexible connecting tubing.

3. HPLC with UV absorption detector at 254

nm, integrator and column (page 2514-1).

4. Vials, 20-mL, scintillation.

5. Pipet, 5-mL.

6. Syringes, 10-μL, readable to 0.1 μL.

7. Volumetric flasks, 5- and 50-mL.

SPECIAL PRECAUTIONS: Anisidine can irritate the skin. Methanol and anisidine can be absorbed 

through the skin. Avoid inhalation of vapors of these compounds and of acetonitrile [4].

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Uncap the sampler immediately before sampling. Attach sampler to personal sampling pump with

flexible tubing.

3. Sample at an accurately known flow rate between 0.5 and 1.0 L/min for a total sample size of 24 to

320 L. Do not sample at less than 0.5 L/min.

4. Cap the samplers. Pack securely for shipment.

SAMPLE PREPARATION:

5. Place the front and back sections of the sampler in separate vials. Discard the glass wool plugs.

6. Add 5.0 mL methanol to each vial. Cap each vial and swirl vigorously.

7. Allow to stand for 15 min. Analyze within one day.

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards over the range 0.4 to 360 μg anisidine per sample for

each isomer.

a. Add a known volume of calibration stock solution, or a dilution thereof in methanol, to a 5-mL

volumetric flask and dilute to the mark with methanol.

b. Analyze together with samples and blanks (steps 11 and 12).

c. Prepare calibration graph (peak area vs. μg analyte) for each isomer.

9. Determine desorption efficiency (DE) at least once for each lot of XAD-2 used for sampling in the

concentration range of interest. Prepare four tubes at each of five levels.

a. Remove and discard the back sorbent section of a media blank sampler.

b. Inject a known amount (e.g., 1 to 20 μL) of calibration stock solution, or a dilution thereof in

methanol, directly onto the front section with a microliter syringe.

c. Cap the tube. Allow to stand overnight.
ANISIDINE: METHOD 2514, Issue 3, dated 26 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

d. Desorb (steps 5 through 7) and analyze with working standards (steps 11 and 12).

e. Prepare graphs of DE vs. μg isomer recovered.

10. Analyze three quality control blind spikes and three analyst spikes with each subsequent set from the

same lot to ensure that the calibration graph and DE graph are in control.

MEASUREMENT:

11. Set HPLC to conditions given on page 2514-1. Inject sample aliquot.

NOTE: Sensitivity is ca. 0.083 absorbance unit/mg of either isomer in a 10-mL injection volume with a

1-cm flow cell. Approximate retention times are 7.5 min for p-anisidine and 12 min for o-anisidine.

12. Measure peak area.

CALCULATIONS:

13. Determine the quantities (sum of quantities of the o-and the p-isomers corrected for DE), mg of

analytes found in the sample front (Wf) and back (Wb) sorbent sections, and in the average media blank

front (Bf) and back (Bb) sorbent sections.

NOTE: If Wb > Wf /10, report breakthrough and possible sample loss.

14. Calculate the sum of the concentrations, C, of the isomers in the air volume sampled, V (L):

CC = WWff + WWbb − BBff − BBbb

VV , mmmm/mm3

EVALUATION OF METHOD: 

Method S163 was issued on February 16, 1979 [2], and was validated over the range 0.13 to 0.58 mg/m3 for 

the o-isomer and 0.12 to 0.58 mg/m3 for the p-isomer using 225-L air samples [1, 5]. The generation system 

was constructed so that samples were generated for both isomers at the same time. Concentrations were 

verified by an independent method using bubblers containing methanol and HPLC analysis [1]. The overall 

precision, SŜ

rrrr, for the combined sampling and measurement of both isomers was 0.068 with an average 

recovery of 100.7% for the o-isomer and 99.7% for the p-isomer [1], which represent non-significant biases 

for the isomers. A separate breakthrough test was conducted for each isomer with XAD-2. Samples were 

taken at ca. 1.0 L/min. o-Anisidine was generated at 1.03 mg/m3 with a relative humidity of 81% at 21 °C. 

Breakthrough (3%) from 150 mg XAD-2 occurred after 236 min, but did not increase above this amount up 

to 479 min. p-Anisidine was generated at 1.04 mg/m3 with a relative humidity of 82% at 20 °C. 

Breakthrough did not occur during the 361-min test. Samples containing both isomers were stored for one 

week at room temperature and found to be stable. Desorption efficiencies averaged 0.95 and 0.91 for oand p-anisidine, respectively, in the range 30 to 240 μg per sample.

REFERENCES:

[1] NIOSH [1979]. Backup data report S163. Unpublished.

[2] NIOSH [1979]. Anisidine, o & p isomers: Method S163. In: Taylor DG, ed. NIOSH Manual of analytical

methods. 2nd ed (vol. 5). Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center

for Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication

No. 79-141.

[3] NIOSH [1977]. Aromatic amines in air: Method P&CAM168. In: Taylor DG, ed. NIOSH Manual of

analytical methods. 2nd ed (vol. 1). Cincinnati, OH: U.S. Department of Health, Education, and

Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, DHEW

(NIOSH) Publication No. 77-157-A.
ANISIDINE: METHOD 2514, Issue 3, dated 26 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[4] NIOSH [1981]. NIOSH/OSHA Occupational health guidelines for occupational hazards. Cincinnati, OH: 

U.S. Department of Health and Human Services, Centers for Disease Control, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 81-12.

[5] NIOSH [1980]. NIOSH research report - development and validation of methods for sampling and 

analysis of workplace toxic substances. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 80-133.

METHOD REVISED BY:

Edward Slick, NIOSH; S163 originally validated under NIOSH Contract 210-76-0123.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYL BROMIDE 2520

 MW: 94.94 CAS: 74-83-9 RTECS: PA4900000

METHOD: 2520, Issue 3 EVALUATION: FULL Issue 1: 15 May 1985

Issue 3: 20 January 2016

OSHA: C 20 ppm (skin)

NIOSH: Lowest feasible; carcinogen

(1 ppm = 3.95 mg/m3 @ NTP)

PROPERTIES: Gas; d 1.73 g/mL @ 0 °C; BP 4 °C; VP 189.34 

kPa (1420 mmHg) @ 20 °C

SYNONYMS: Monobromomethane; bromomethane

SAMPLING

SAMPLER: SOLID SORBENT TUBES

(two petroleum charcoal tubes, 400 mg and 

200 mg; drying tube, 9 g sodium sulfate, 

necessary at humidity >50%)

FLOW RATE: 0.01 L/min to 0.1 L/min

VOL-MIN: 1 L @ 5 ppm

-MAX: 5 L with drying tube, 1 L without drying 

tube [1]

SHIPMENT: Ship on dry ice at −10 °C [1]

SAMPLE

STABILITY: Six d at −10 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: 0.84 ppm to 32.0 ppm (5 L samples)

BIAS: −5.2%

OVERALL

PRECISION ( ): 0.089

ACCURACY: ±19.4%

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, ATOMIC 

EMISSION DETECTION (GC-AED) 

monitoring bromine, carbon, and 

hydrogen channels

ANALYTE: Methyl bromide

DESORPTION: 400 mg tube: 3 mL methylene chloride; 

200 mg tube: 2 mL methylene chloride

INJECTION: 1 μL

TEMPERATURE-INJECTOR: 250 °C

-TRANSFER LINE/

CAVITY BLOCK: 250 °C

-COLUMN: 30 °C, 3.5 min; 12 °C /min to 

130 °C; hold 1 min

CARRIER GAS: Helium

COLUMN: US Pharmacopeia (USP) G2 capillary, 

30 m × 0.32 mm ID, 1.0 μm film thickness; 

1 m × 0.53 mm deactivated fused silica 

pre-column

CALIBRATION: Brominated compounds in methylene 

chloride

RANGE: 33.0 μg to 2687 μg bromine per sample

ESTIMATED LOD: 16.6 μg per sample [1]

PRECISION ( ): 0.066 [1]

APPLICABILITY: The working range is 0.84 ppm to 32.0 ppm (3.3 mg/m3 to 126 mg/m3) for a 5 L sample. Ceiling 

measurement samples may require dilution when analyzed.

INTERFERENCES: Water vapor interferes with collection at relative humidities (RH) >50%. To eliminate the interference, 

precede the sampling train with a drying tube, and limit the sample volume to 5 L. If drying tubes are not available, limit the 

sample volume to 1 L under humid conditions.

OTHER METHODS: This is NIOSH method 2520 [2] revised to account for humidity effects, as well as instability of standards 

and samples. Other researchers [3] have prepared methyl bromide standards gravimetrically and used GC-ECD for 

analysis. The gravimetric calibration procedure did not give consistent results when compared with the procedure used 

in this revision of NIOSH method 2520. GC-ECD with a US Pharmacopeia (USP) S3 capillary column may be an alternative 

technique to GC-AED if other bromine standards are used to confirm the concentration of methyl bromide standards.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYL BROMIDE: METHOD 2520, Issue 3, dated 20 January 2016 - Page 2 of 5

REAGENTS:

1. Eluant: methylene chloride,* chromatographic

grade.

2. Methyl bromide,* 99.5%.

3. Calibration stock solution: To 4 mL

of methylene chloride, add 12 μL

dibromomethane.

NOTE: Other brominated compounds may be

used if liquid at room temperature.

4. Desorption stock solution: Bubble methyl

bromide gas slowly into chilled eluant.

Determine the methyl bromide concentration

by comparison with calibration standards.

5. Helium,* prepurified, 99.995%.

6. Oxygen,* ultra purified, as reagent gas for

plasma, 207 kPa (30 psi).

7. Air, filtered.

8. Dry ice, flaked, for chilling solvent.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: petroleum charcoal sampling tubes;

two glass tubes, each tube, 10 cm long, 8 mm

OD, 6 mm ID, containing 20/40 mesh activated

(600 °C) petroleum charcoal, first tube 400

mg, second tube 200 mg, held in place with

silylated glass wool plugs; drying tube, glass, 9

g sodium sulfate. Tubes are connected in series

with short pieces of plastic tubing. Pressure

drop across sampler <3.4 kPa (25 mmHg) at

1.0 L/min airflow. Tubes are commercially

available.

NOTE: If RH ≥50%, precede sampling train with

drying tube.

2. Personal sampling pump, 0.01 to 0.1 L/min,

with flexible connecting tubing.

3. Gas chromatograph, atomic emission detector

(helium plasma), integrator or computer, and

column (page 2520-1).

4. Vials, 4 mL and 10 mL, glass, with PTFE-lined

caps.

5. Syringe, gas-tight, 10 mL.

6. Microliter syringes, 10 μL, 50 μL, 100 μL, 250 μL,

500 μL for preparing standard solutions.

7. Pipettes, 2 mL, graduated in 0.1 mL

increments.

SPECIAL PRECAUTIONS: Methylene chloride is a suspect carcinogen. Methyl bromide is a suspect 

carcinogen and is toxic by ingestion, inhalation, and skin absorption [4]. Users must be familiar with 

the proper use of flammable and nonflammable gases, cylinders, and regulators.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break the ends of the sampling tubes immediately before sampling. Attach sampler to personal

sampling pump with flexible tubing.

NOTE: Place drying tube in front of the sorbent train.

3. Sample at an accurately known flow rate between 0.01 L/min and 0.1 L/min for a total sample size of

1.0 L to 5.0 L.

NOTE: Limit sample volume to 1.0 L if RH ≥ 50% and no drying tube is used.

4. Separate the front and back tubes immediately after sampling. Cap the tubes. Pack securely in dry

ice for shipment. Store at −10 °C until analysis.

SAMPLE PREPARATION:

5. Place the sorbent sections from each sampling tube in separate vials. Discard the glass wool plugs.

6. Add 3.0 mL chilled eluant (methylene chloride) to each 400 mg section, and 2.0 mL chilled eluant to

each 200 mg section. Immediately cap each vial.

7. Allow to stand at least 30 min at room temperature with occasional agitation. Rechill before

transferring to autosampler vials or diluting.

NOTE: Because of the volatility of the analyte, it is suggested that any dilutions be prepared at the
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYL BROMIDE: METHOD 2520, Issue 3, dated 20 January 2016 - Page 3 of 5

time of transfer to autosampler vials. Dilutions can be stored in the freezer until determined 

that they are needed.

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards over the range 0.14 μg to 272 μg methyl bromide

per sample from calibration stock solution.

NOTE: 0.085 μmol/μL bromine is equivalent to a methyl bromide concentration of 8.07 μg/μL.

a. Add known aliquots (2.0 μL, 20 μL, 45 μL, and 120 μL) of calibration stock solution to methylene

chloride in 10 mL vials with PTFE-lined caps. Take 1 mL of lowest standard and dilute to 10 mL

with methylene chloride. Transfer standards to autosampler vials and immediately cap each vial.

b. Analyze together with samples and blanks (steps 11 and 12).

c. Prepare calibration graph (peak area vs. concentration).

9. Determine desorption efficiency (DE) at least once for each lot of charcoal used for sampling in the

calibration range (step 8). Prepare three tubes at each of five levels plus three media blanks.

a. Inject a known amount (1 μL to 10 μL) of methyl bromide desorption stock solution

(concentration determined against other brominated compounds), or a serial dilution thereof,

directly onto media blank samplers with a microliter syringe.

b. Cap the tubes. Allow to stand overnight at 0 °C.

c. Desorb (steps 5, 6, and 7) and analyze together with working standards (steps 11 and 12).

d. Prepare a graph of DE vs. μg bromine recovered.

10. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration

graph and DE graph are in control.

MEASUREMENT:

11. Set gas chromatograph-atomic emission detector to manufacturer’s recommendations and to

conditions given on page 2520-1. Inject sample aliquot manually using solvent flush technique or

with autosampler.

NOTE 1: Methyl bromide elutes before the methylene chloride solvent peak. Vent the solvent peak

to avoid extinguishing the helium plasma. Vent time ranges from 3.2 min to 5.9 min; this 

may need to be adjusted for each system.

NOTE 2: If peak area is above the linear range of the working standards, dilute with eluant, reanalyze, 

and apply the appropriate dilution factor in calculations.

12. Measure peak area.

CALCULATIONS:

13. Determine the mass, μg (corrected for DE) of methyl bromide found in the 400 mg sample tube ( )

and 200 mg sample tube ( ), and in the average media blanks for the 400 mg ( ) and 200 mg ( )

sorbent tubes.

NOTE: If , report breakthrough and possible sample loss.

14. Calculate concentration, , of methyl bromide in the air volume sampled, (L):

, μg/L or mg/m3.

EVALUATION OF METHOD:

NIOSH method 2520 for methyl bromide, issued in May 1985 [2], was based on NIOSH method S372 

[5]. Issue 2 (dated 5/15/96) of NIOSH method 2520 was further revised to account for the effect of 

humidity and to address the instability of standards and samples [1]. The addition of a drying tube to 

the sampling train helped reduce the effects of >50% RH. The analytical technique was changed to GC 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYL BROMIDE: METHOD 2520, Issue 3, dated 20 January 2016 - Page 4 of 5

with atomic emission detection (GC-AED). The atomic emission detector monitors individual elements, 

in this case bromine, independent of the source compound. Calibration can be done with brominated 

compounds that are liquid at room temperature, thereby eliminating the need for methyl bromide 

standards. Methylene chloride replaced carbon disulfide as desorption solvent.

The revised NIOSH method 2520 (issued 5/15/96) for methyl bromide was evaluated at 7.8 mg/m3, 

20.0 mg/m3, and 125.0 mg/m3 (2 ppm, 5 ppm, and 32 ppm, respectively) [1]. Test atmospheres were 

generated by delivering methyl bromide gas from two diffusion tubes kept at −12 °C into an airstream 

flowing at 12 L/min. Humidity was added downstream when needed, and airflow passed through 

two mixing chambers before reaching a sampling manifold. The concentration was monitored with 

a total hydrocarbon analyzer and confirmed by calibrating against other brominated standards 

by GC-AED. Three compounds used as calibration standards (dibromomethane, 1-bromopropane, 

and 1-bromobutane) were chosen as closest in chemical structure to methyl bromide, although a 

compound-independent response was assumed [6]. A three-compound calibration was used during the 

development of the GC-AED analytical procedure. Since there was good agreement between the three 

compounds, the method was written with only one brominated standard, dibromomethane.

When challenged with methyl bromide at a calculated concentration of 27 ppm, the capacity of the 

sampler (a 400 mg petroleum charcoal tube plus a 200 mg petroleum charcoal tube) at 40% RH and 

20 °C was 1386.5 μg regardless of flow rate (10.5 mL/min, 40 mL/min, or 100 mL/min). However, at 100% 

RH and 39 °C, the average capacity fell to 298.6 μg. With a 9 g sodium sulfate drying tube in line, the 

capacity was increased to 651.8 μg; concentration averaged 130 mg/m3 (33 ppm). Even with a drying 

tube in line, severe breakthrough occurred at the 10 L sample volume (50% was found on the back 

tube). Without the use of a drying tube, a 1 L sample volume is recommended, based on a 170.6 μg 

capacity (1.6 L) found at the 40 mL/min rate multiplied by a 0.67 caution factor.

Recovery fell below 70% for sample loadings less than 58 μg when carbon disulfide was used for 

desorption (Figure 1). This would not allow accurate sampling at 7.8 mg/m3 (2 ppm), the exposure level 

most frequently encountered. Therefore, alternate desorption solvents were tested. Desorption with 

methylene chloride improved recovery at the 15 μg level to 76.7%. However, sample stability still fell 

below 70% after storage for six days at −10 °C regardless of sample level. This stability limitation remains 

a concern and rapid sample analysis is required.

Figure 1. Comparison of desorption solvents.

REFERENCES:

[1] NIOSH [1995]. Research into problems with NIOSH method 2520 for methyl bromide. By Gagnon YT,

Ringenburg VL, Fajen JM. Cincinnati, OH: U.S. Department of Health and Human Services, Centers
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYL BROMIDE: METHOD 2520, Issue 3, dated 20 January 2016 - Page 5 of 5

for Disease Control and Prevention, National Institute for Occupational Safety and Health, available 

as accession No. PB95-179842 from NTIS [www.ntis.gov]. Unpublished.

[2] NIOSH [1984]. Methyl bromide: Method 2520 (supplement issued 5/15/85). In: Eller PM, ed. NIOSH

manual of analytical methods. 3rd ed. Cincinnati, OH: U.S. Department of Health and Human

Services, Centers for Disease Control, National Institute for Occupational Safety and Health, DHHS

(NIOSH) Publication No. 84-100, available as accession No. PB86-116266 from NTIS [www.ntis.gov].

[3] Woodrow JE, McChesney MM, Sieber JN [1988]. Determination of methyl bromide in air samples by

headspace gas chromatography. Anal Chem 60:509–512.

[4] NIOSH [1984]. Current intelligence bulletin 43—monohalomethanes: methyl chloride, methyl

bromide, methyl iodide. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication

No. 84-117, available as accession No. PB85-178549 from NTIS [www.ntis.gov].

[5] NIOSH [1977]. Methyl bromide: Method S372. In: Taylor DG, ed. NIOSH manual of analytical

methods. 2nd ed. Vol. 3. Cincinnati, OH: U.S. Department of Health, Education, and Welfare,

Center for Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH)

Publication No. 77-157-C, available as accession No. PB276838 from NTIS [www.ntis.gov].

[6] Weile Y, Yieru H, Qingyu O [1992]. Quantitative characteristics of gas chromatography with

microwave-induced plasma detection. In: Uden PC, ed. Element-specific chromatographic

detection by atomic emission spectroscopy. Washington DC: American Chemical Society, ACS

Symposium Series No. 479, pp. 44–61.

METHOD REVISED BY:

Yvonne T. Gagnon, MRB/DPSE/NIOSH

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

GLUTARALDEHYDE 2531

 MW: 100.12 CAS: 111-30-8 RTECS: MA2450000

METHOD: 2531, Issue 3 EVALUATION: FULL Issue 1: 15 May 1989

Issue 3: 20 January 2016

OSHA: No PEL

NIOSH: C 0.2 ppm

(1 ppm = 4.09 mg/m3 @ NTP)

PROPERTIES: Oil; d 0.72 g/mL @ 20 °C; BP 188 °C; MP 

−14 °C; VP 2.2 kPa (17 mmHg) @ 20 °C

SYNONYMS: Glutaric dialdehyde; 1,5-pentanedial

SAMPLING

SAMPLER: SOLID SORBENT TUBE

(10% 2-(hydroxymethyl)piperidine on XAD-2, 

120 mg/60 mg)

FLOW RATE: 0.01 L/min to 0.08 L/min, or 0.2 L/min for 20 

min

VOL-MIN: 4 L @ 0.2 ppm

-MAX: 39 L

SHIPMENT: Routine

SAMPLE

STABILITY: At least 5 weeks @ 25 °C [1]

FIELD

BLANKS: 2 to 10 field blanks per set

MEDIA

BLANKS: 10 per set

ACCURACY

RANGE STUDIED: 0.8 mg/m3 to 9 mg/m3 [1] (22 L samples)

BIAS: 0.3%

OVERALL

PRECISION ( ): 0.087 [1]

ACCURACY: ±17.4%

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, FID

ANALYTE: Oxazolidine derivative of glutaraldehyde

DESORPTION: 2 mL toluene; 60 min ultrasonic

INJECTION: 1 μL splitless; split vent time 30 s

TEMPERATURE-INJECTOR: 250 °C

-DETECTOR: 280 °C

-COLUMN: 1 min @ 70 °C; 20 °C/min; hold 2 

min @ 290 °C

CARRIER GAS: Helium, 0.5 mL/min; makeup 2.9 mL/min

COLUMN: Capillary, 10 m × 0.25 mm, 5% 

phenyl, 95% methyl polysiloxane (US 

Pharmacopeia (USP) phase G27)

CALIBRATION: Standard glutaraldehyde solutions spiked 

on sorbent

RANGE: 3 μg to 180 μg per sample [1]

ESTIMATED LOD: 1 μg per sample [2]

PRECISION ( ): 0.093 [2] @ 5 μg to 50 μg per sample [1]

APPLICABILITY: The working range is 0.03 ppm to 2 ppm (0.14 mg/m3 to 8 mg/m3) for a 22 L air sample; the method is 

sensitive enough for ceiling determinations. The method is suitable for the simultaneous determination of furfural and 

glutaraldehyde.

INTERFERENCES: None have been observed.

OTHER METHODS: This is a new method. A wide-bore 10 m capillary column is an alternate chromatographic column.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

GLUTARALDEHYDE: METHOD 2531, Issue 2, dated 20 January 2016 - Page 2 of 5

REAGENTS:

1. Toluene,* chromatographic quality.

2. 2-(Hydroxymethyl)piperidine. Recrystallize

several times from isooctane until there is

one major peak (>95% of area) by GC analysis.

Store in desiccator.

3. XAD-2 resin.

4. Glutaraldehyde,* 250 g/L solution in water.

5. Glutaraldehyde stock solution, 10 μg/μL (see

Appendix A).

6. Glutaraldehyde oxazolidine (see Appendix B)

stock solution, 2 mg/mL. Add 20 mg to toluene

and dilute to 10 mL.

7. Sulfuric acid,* 0.01 mol/L.

8. Sodium hydroxide,* 0.01 mol/L.

9. Sodium sulfite, 1.13 mol/L. Dissolve 14.2 g

reagent grade sodium sulfite in deionized,

distilled water and dilute to 100 mL. Prepare

fresh immediately before use.

10. Water, deionized, then distilled.

11. Hydrogen,* prepurified.

12. Air, filtered.

13. Helium,* prepurified.

14. Magnesium sulfate.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: resin-filled sampling tube; glass tube,

10 cm long, 6-mm OD, 4-mm ID, flame-sealed

ends and plastic caps, containing two sections

of 40/60 mesh 2-(hydroxymethyl)piperidinecoated XAD-2 (see Appendix C). Sorbent

sections are retained and separated by small

plugs of silanized glass wool. Pressure drop

across the tube at 0.10 L/min airflow must

be less than 0.76 kPa (5.7 mmHg). Tubes are

commercially available.

2. Personal sampling pump, 0.01 L/min to 0.08

L/min, with flexible connecting tubing.

3. Gas chromatograph, flame ionization detector,

integrator, and column (page 2531-1).

4. Ultrasonic bath.

5. Vials, glass, 4 mL, with septum and plastic

screw caps.

6. Flasks, volumetric, 10 mL, 25 mL, and 50 mL.

7. Pipets, TD, 1 mL, 2 mL, and 10 mL with pipet

bulb.

8. Pipets, disposable, 2 mL.

9. Syringes, 10 μL (readable to 0.1 μL), 25 μL, and

50 μL.

10. File.

11. Beakers, 50 mL.

12. pH meter.

13. Magnetic stirrer.

14. Burets, 50 mL.

15. Flasks, round-bottomed, 100 mL.

16. Soxhlet extraction apparatus.

17. Vacuum oven.

18. Distillation apparatus.

SPECIAL PRECAUTIONS: Glutaraldehyde can irritate the mucous membranes and act on the central 

nervous system [3]. Toluene is flammable. Sulfuric acid is highly corrosive and sodium hydroxide is 

caustic. All work with these compounds should be performed in a well-ventilated hood. Use proper 

protective clothing including gloves, safety glasses, and laboratory coat. Users must be familiar with 

the proper use of flammable and nonflammable gases, cylinders, and regulators.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break ends of the sampler immediately before sampling. Attach sampler to personal sampling pump

with flexible tubing.

3. Sample at an accurately know flow rate between 0.01 and 0.08 L/min for a total sample size of 4 L to

39 L. For ceiling determination, sample at 0.2 L/min for 20 min.

NOTE: The aldehyde reacts with the 2-(hydroxymethyl)piperidine to form an oxazolidine derivative

in the sorbent bed during sampling (see equation below). Sampling rate is limited by the 

speed of this reaction. Sampling rates above 0.1 L/min for extended periods may cause 

breakthrough owing to incomplete reaction, possibly invalidating the sample.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

GLUTARALDEHYDE: METHOD 2531, Issue 2, dated 20 January 2016 - Page 3 of 5

SAMPLE PREPARATION:

4. Score each sampler with a file in back of the back sorbent section.

5. Break sampler at score line. Remove and place back glass wool plug and back sorbent section in a

vial.

6. Transfer front section with remaining glass wool plugs to a second vial.

7. Add 2.0 mL toluene to each vial. Screw cap tightly onto each vial.

8. Agitate vials in an ultrasonic bath for 60 min.

NOTE: Desorption efficiency is affected by the amount of time that the vials are allowed to spend in

the ultrasonic bath. A minimum of 60 min residence time in the ultrasonic bath is required to 

ensure adequate desorption.

CALIBRATION AND QUALITY CONTROL:

9. Prepare glutaraldehyde oxazolidine standard solutions.

a. Add known amounts of glutaraldehyde oxazolidine stock solution (equivalent to the range of the

samples) to toluene in 10 mL volumetric flasks and dilute to the mark.

b. Analyze (steps 12 and 13) with samples and blanks for qualitative identification of derivative

peaks.

10. Calibrate daily with at least five working standards covering the range of the samples.

a. Weigh 120 mg portions of unused sorbent from media blanks into vials.

b. Add aliquiots of glutaraldehyde stock solution or dilutions thereof. Cap vials and allow them to

stand overnight at room temperature.

c. Desorb (steps 7 and 8) and analyze (steps 12 and 13) with samples and blanks.

d. Prepare calibration graph (combined peak area vs. μg glutaraldehyde).

NOTE: Because the working standards are prepared on media blanks, no additional blank

correction or desorption efficiency correction is necessary. Check desorption efficiency 

occasionally in the range of interest (see Appendix D).

11. Analyze three quality control blind spikes to ensure that the calibration graph is in control.

MEASUREMENT:

12. Set gas chromatograph to manufacturer’s recommendations and to conditions given on page

2531-1. Inject 1 μL sample aliquot.

NOTE: If the amount of oxazolidine in the aliquot exceeds the capacity of the column, dilute the

sample with toluene and apply the appropriate dilution factor in calculations.

13. Measure total peak area of the two analyte peaks.

NOTE: On the recommended column, the oxazolidine derivative gives two peaks, since the

diastereoisomers are resolved; for the glutaraldehyde derivative = 9.4 min and 9.7 min; for 

2-(hydroxymethyl)piperidine = 2.6 min for these conditions.

CALCULATIONS:

14. Determine the mass, μg, of glutaraldehyde found in the sample front ( ) and back ( ) sorbent

sections.

NOTE: If , report breakthrough and possible sample loss.

15. Calculate concentration, , of glutaraldehyde in the air volume sampled, (L):

, μg/L or mg/m3.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

GLUTARALDEHYDE: METHOD 2531, Issue 2, dated 20 January 2016 - Page 4 of 5

EVALUATION OF METHOD:

Atmospheres were generated by injection of an aqueous solution of glutaraldehyde by syringe pump 

into a heated block injector and flash vaporization into a stream of air flowing at a fixed rate [1]. Relative 

humidity during generation was controlled at 80% ±5%. The generator and sampling manifold systems 

have been described previously [4]. Concentration of glutaraldehyde vapor was independently verified 

by the 2,4-dinitrophenylhydrazine procedure of Lipari and Swarin [5]. No bias with dynamicallygenerated atmospheres was observed with the method over the range 0.8 mg/m3 to 8 mg/m3 using 

22 L air samples. Desorption efficiencies on statically-spiked samples averaged 87% in the ranges 5 

μg to 50 μg per sample. Recovery averaged 1.10 with = 0.043 for twelve tubes spiked with 67 μg 

glutaraldehyde [6].

REFERENCES:

[1] Kennedy ER, Gagnon YT, Okenfuss JR, Teass AW [1988]. The determination in air of selected lowmolecular weight aldehydes as their oxazolidines by capillary gas chromatography. Appl Ind Hyg

3(10):274–279.

[2] NIOSH [1987, 1989]. Analytical reports: sequence No. 5827 (November 13, 1987); sequence No.

6565-J (February 27, 1989). Cincinnati, OH: U.S. Department of Health and Human Services, Centers

for Disease Control, National Institute for Occupational Safety and Health, Division of Physical

Sciences and Engineering. Unpublished.

[3] Windholz M, Budavari S, Blumetti RF, Otterbein ES, eds. [1983]. The Merck index: an encyclopedia of

chemicals, drugs, and biologicals. 10th ed. Rahway, NJ: Merck & Co., Inc.

[4] Kennedy ER, Hill RH [1982]. Determination of formaldehyde in air as an oxazolidine derivative by

capillary gas chromatography. Anal Chem 54(11):1739–1741.

[5] Lipari F, Swarin SJ [1982]. Determination of formaldehyde and other aldehydes in automobile

exhaust with an improved 2,4-dinitrophenylhydrazine method. J Chromatogr 247:297–306.

[6] NIOSH [1989]. User check, NIOSH sequence No. 6565-M. Cincinnati, OH: U.S. Department of Health

and Human Services, Centers for Disease Control, National Institute for Occupational Safety and

Health, Division of Physical Sciences and Engineering. Unpublished.

METHOD WRITTEN BY:

Julie R. Okenfuss and Eugene R. Kennedy, Ph.D., NIOSH/DPSE.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.

APPENDIX

A. Preparation and Standardization of Glutaraldehyde Stock Solution (about 10 μg/μL)

Dilute 1 mL of 250 g/L aqueous glutaraldehyde to 25 mL with distilled, deionized water to make 

the glutaraldehyde stock solution. Put 10.0 mL of 1.13 mol/L sodium sulfite solution in a beaker and 

stir magnetically. Adjust pH to between 8.5 and 10 with base or acid. Record the pH. Add 1.0 mL 

glutaraldehyde stock solution. The pH should be about 12. Titrate the solution back to its original pH 

with 0.01 mol/L sulfuric acid. If the endpoint pH is overrun, back-titrate to the endpoint with 0.01 mol/L 

sodium hydroxide. Calculate the concentration, , of the glutaraldehyde stock solution:

, μg/μL,
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

GLUTARALDEHYDE: METHOD 2531, Issue 2, dated 20 January 2016 - Page 5 of 5

where: 50.06 = MW of glutaraldehyde divided by 2, 

= concentration (mol/L) of sulfuric acid, 

= volume of sulfuric acid (mL) used for titration, 

= concentration (mol/L) of sodium hydroxide, 

= volume of sulfuric acid (mL) used for titration, and

= volume of glutaraldehyde stock solution (1.0 mL).

B. Synthesis of Glutaraldehyde Oxazolidine

Place a solution of purified 2-(hydroxymethyl)piperidine (0.57 g, 5 mmol) in 10 mL of toluene in a 50 mL 

round-bottomed flask. Several 2 mL portions of toluene can be used to rinse residual 2-(hydroxymethyl)

piperidine from the container used for weighing. Add magnesium sulfate (2.5 g) to the round-bottomed 

flask to dry the glutaraldehyde solution as it is added and to remove the water which forms during 

the reaction. Add a solution of 1 mL of 250 g/L aqueous glutaraldehyde (0.25 g, 2.5 mmol) in 10 mL of 

toluene to the 2-(hydroxymethyl)piperidine solution dropwise with stirring over 1 h. Stir the solution 

overnight, then filter to remove the magnesium sulfate. Remove the toluene from the solution at 

reduced pressure by rotary evaporation. The product is a yellow viscous oil, about 90% to 95% pure.

NOTE: Exact amounts of reagent are required for this synthesis since excess glutaraldehyde can cause 

appreciable formation of the mono-oxazolidine derivative of glutaraldehyde.

C. Sorbent Preparation (optional if commercially prepared tubes are used)

Extract XAD-2 sorbent 4 h in a Soxhlet extractor with a mixture of equal volumes of acetone and 

methylene chloride. Replace with fresh solvent and repeat. Vacuum dry overnight. Add 1 g purified 

2-(hydroxymethyl)piperidine in 50 mL toluene for each 9 g extracted XAD-2 sorbent. Allow this mixture 

to stand 1 h with occasional swirling. Remove the solvent by rotary evaporation at 37 °C and dry at 0.13 

kPa (1 mmHg) at ambient temperature for approximately 1 h. To determine the amount of background 

for each batch, extract several 120 mg portions of the coated sorbent with toluene and analyze (steps 7, 

8, 9, 10, 11, 12, and 13). No blank peak is expected for glutaraldehyde.

D. Desorption Efficiency

The determination of desorption efficiency (DE) is not necessary when using the calibration procedure 

in step 10. If desired, the following procedure can be used to determine DE:

1. Prepare and analyze a set of glutaraldehyde oxazolidine standard solutions (step 9.a) and a set of

working standards (step 10), including media blanks.

2. Treating the working standards as unknowns, read the mass (μg) of oxazolidine found in each

working standard ( ), and in the average media blank ( ).

3. Using the mass of glutaraldehyde, μg, spiked onto the working standard ( ) and the stoichiometric

conversion factor between glutaraldehyde and glutaraldehyde oxazolidine (2.94), calculate the

desorption efficiency .

4. Prepare a graph of DE vs. μg glutaraldehyde recovered per sample .
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

VALERALDEHYDE 2536

CH3(CH2)3CH=O MW: 86.13 CAS: 110-62-3 RTECS: YV3600000

METHOD: 2536, Issue 4 EVALUATION: FULL Issue 1: 15 May 1989 

Issue 4: 3 March 2016

OSHA: none

NIOSH: 50 ppm 

PROPERTIES: liquid; d 0.810 g/mL @ 20 °C; BP 103 °C; VP 6.7

kPa (50 mm Hg) @ 25 °C; vapor density (air = 1) 

3.0; flash point = 12.2 °C

SYNONYMS: pentanal; amyl aldehyde.

SAMPLING

SAMPLER: SOLID SORBENT TUBE (10% 2-

(hydroxymethyl)piperidine on XAD-2, (120 

mg/60 mg)

FLOW RATE: 0.01 - 0.04 L/min

VOL-MIN: 0.5 L @ 50 ppm

-MAX: 10 L

SHIPMENT: routine

SAMPLE

STABILITY: at least 4 weeds @ 25 °C [1]

FIELD BLANKS: 2 to 10 field blanks per set

MEDIA BLANKS: 18 per set

ACCURACY

RANGE 

STUDIED: 9 to 374 mg/m3 [1] (12-L samples)

BIAS: 0.12% 

OVERALL

PRECISION (SS�rrrr): 0.073 [1] 

ACCURACY: ± 14.4% 

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, FID

ANALYTE: valeraldehyde oxazolidine (9-butyl-1-

aza-8-oxabicyclo[4.3.0]nonane)

DESORPTION: 2 mL toluene, 60 min ultrasonic

INJECTION 

VOLUME: 1 μL splitless

TEMPERATURE

 INJECTION: 250 °C

- DETCTOR: 280 °C

-COLUMN: 0.5 min @ 70°C; 50 °C/min to 120°C, hold 

4 min; 20°C/min to 170°C, hold 7 min

CARRIER GAS: He, 27 cm/sec linear velocity

makeup flow 29 mL/min

COLUMN: capillary, 15 m x 0.32-mm, 5% phenyl, 

95% methyl polysiloxane, 1-μm film (US 

Pharmacopeia (USP) G-27)

CALIBRATION: standard solutions of valeraldehyde on 

sorbent

RANGE: 4 to 3900 μg/sample [1]

ESTIMATED LOD: 2 μg/sample [1]

PRECISION (SS̅

rr): 0.066 @ 2 to 508 μg per sample [1]

APPLICABILITY: The working range is 0.11 to 110 ppm (0.4 to 390 mg/m3

) for a 10-L air sample. The method is also suitable for 

determination of furfural and glutaraldehyde in a mixture [2].

INTERFERENCES: None have been observed; an alternate capillary column, 15 m x 0.32-mm cyanopropylphenyl 

dimethylpolysiloxane 1-μm film (USP G43) can be used.

OTHER METHODS: The method of Lipari and Swarin [3] uses 2,4-dinitrophenylhydrazine for the collection of valeraldehyde.
VALERALDEHYDE: METHOD 2536, Issue 4, dated 3 March 2016 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Toluene, chromatographic quality.

2. 2-(Hydroxymethyl) piperidine. Recrystallize 

several times from isooctane until there is 

one major peak (>95% of area) by GC 

analysis. Store in desiccator.

3. XAD-2. Extract 4 h in Soxhlet with 50/50 (v/v) 

acetone/methylene chloride. Replace with 

fresh solvent and repeat. Vacuum dry 

overnight. (Optional if commercial tubes are 

used.)

4. Valeraldehyde, * 99% purity.

5. Valeraldehyde stock solution, 40 μg/μL (see 

APPENDIX). Add 400 mg valeraldehyde to 

toluene and dilute to 10 mL.

6. Valeraldehyde oxazolidine stock solution, 10 

mg/mL (see APPENDIX A). Add 0.10 g 9-

butyl-1-aza-8-oxabicyclo-[4.3.0] nonane to 

toluene and dilute to 10 mL.

7. Hydrogen, prepurified.

8. Air, filtered, compressed.

9. Helium, purified.

10. Magnesium sulfate, anhydrous.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: resin-filled sampling tube; glass 

tube, 10 cm long, 6-mm OD, 4-mm ID, flamesealed ends with plastic caps, containing two 

sections of 2-(hydroxymethyl) piperidinecoated XAD-2 (front = 120 mg, back = 60 mg) 

(see APPENDIX B). Sorbent sections are 

retained and separated by small plugs of 

silanized glass wool. Pressure drop across the 

tube at 0.1 L/min must be less than 760 Pa 

(5.7 mm Hg). Tubes are commercially 

available.

2. Personal sampling pump, 0.01 to 0.04 L/min, 

with flexible connecting tubing.

3. Gas chromatograph, flame ionization 

detector, integrator and column (page 2536-

1).

4. Ultrasonic bath.

5. Vials, glass, 4-mL, with septum and plastic 

screw caps.

6. Flasks, volumetric, 10-, 25-, and 50-mL.

7. Pipets, volumetric, 1-, 2-, and 10-mL with 

pipet bulb.

8. Pipets, disposable, 2-mL.

9. Syringes, 10-μL (readable to 0.1 μL), 25-, and 

50-μL.

10. File or tube scorer.

11. Beakers, 50-mL.

12. Magnetic stirrer.

13. Flasks, round-bottomed, 100-mL.

14. Soxhlet extraction apparatus.

15. Vacuum oxen.

16. Distillation apparatus.

SPECIAL PRECAUTIONS: Valeraldehyde can irritate the mucous membranes [4]. It is flammable, a 

dangerous fire risk. Toluene is extremely flammable. All work should be performed in a well-ventilated 

fume hood.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break ends of the sampler immediately before sampling. Attach sampler to personal sampling 

pump with flexible tubing.

3. Sample at an accurately known flow rate between 0.01 and 0.04 L/min for a total sample size of 0.5 

to 10 L.

NOTE: Sampling rate is limited by the speed of the following reaction. Rates above 0.04 L/min may 

cause appreciable breakthrough owing to incomplete reaction, possibly invalidating the sample.
VALERALDEHYDE: METHOD 2536, Issue 4, dated 3 March 2016 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CH3(CH2)3CH=0 +

valeraldehyde

C5H9NHCH2OH

2-(hydroxymethyl)-

piperidine

 → 

toluene/

ultrasonic energy

C5H9NHC2H2O(CH2)3

CH3

valeraldehyde 

oxazolidine

+ H20

SAMPLE PREPARATION:

4. Score each sampler with a file or tube scorer in back of the back sorbent section.

5. Break sampler at score line. Remove and place back glass wool plug and back sorbent section in a

vial.

6. Transfer front section with remaining glass wool plugs to a second vial.

7. Add 2.0 mL toluene to each vial. Screw cap tightly onto each vial.

8. Agitate in an ultrasonic bath for 60 min.

CALIBRATION AND QUALITY CONTROL:

9. Identification of analytical peaks.

a. Add known amounts of valeraldehyde oxazolidine stock solution to toluene in 10-mL

volumetric flasks and dilute to the mark.

b. Analyze (steps 12 and 13) with samples and blanks for qualitative identification of derivative

peaks.

10. Calibrate daily with a least six working standards prepared in triplicate covering the range 2 to 3900

μg valeraldehyde per sample.

a. Weigh 120-mg portions of unused sorbent from media blanks into vials.

b. Add aliquots (1 to 10 μL) of valeraldehyde stock solution, or dilutions thereof, to the sorbent.

Cap vials and allow to stand overnight at room temperature.

c. Desorb (steps 7 and 8) and analyze (steps 12 and 13) with samples and blanks.

d. Prepare calibration graph (combined peak area vs. μg valeraldehyde).

NOTE: Because the standard samples are prepared on media blanks, no additional blank

correction or desorption efficiency correction is necessary. Check desorption efficiency 

in the range of interest and at least once over the entire range of the method with each 

lot of sorbent used. (see APPENDIX C). 

11. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration

graph is in control.

MEASUREMENT:

12. Set gas chromatograph to manufacturer's recommendations and to conditions given on page

2536-1. Inject 1-μL sample aliquot.

NOTE: If the amount of valeraldehyde oxazolidine in the aliquot exceeds the capacity of the

column, dilute with toluene, reanalyze and apply the appropriate dilution factor in 

calculations. The upper limit for the column on (page 2536-1) is equivalent to ca. 260 μg 

valeraldehyde per sample.

13. Measure total peak area of the two analyte peaks.

NOTE: Valeraldehyde oxazolidine gives two peaks, since the diastereoisomers are resolved with

retention times 5.4 and 6.3 min. Retention time for 2-(hydroxymethyl) piperidine is 2.2 min 

for these conditions. 
VALERALDEHYDE: METHOD 2536, Issue 4, dated 3 March 2016 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALCULATIONS:

14. Determine the mass, μg, of valeraldehyde found in the sample front (Wf) and back (Wb) sorbent

sections.

NOTE: If Wb > Wf /10, report breakthrough and possible sample loss.

15. Calculate concentration, C, of valeraldehyde in the air volume sampled, V (L):

CC = WWff + WWbb

VV , mmmm/mm3

EVALUATION OF METHOD:

Atmospheres were generated by injection of valeraldehyde with a syringe pump into a heated block 

injector and flash vaporizer into a stream of air at 80% ± 5% RH flowing at a fixed rate. The generator and 

sampling manifold system have been described previously [5]. Concentration of valeraldehyde vapor was 

independently verified by the 2,4-dinitrophenylhydrazine procedure of Lipari and Swarin [3] or by 

monitoring with an AID Model 590 organic vapor monitor. Breakthrough studies of valeraldehyde at 100 

ppm, conducted at 75 and 50 mL/min flow rates, gave 5% breakthrough at 170 min and 280 min, 

respectively.

The method was evaluated over the range of 9 to 374 mg/m3 using 12-L samples. Desorption efficiencies 

from statically-spiked samples average 102.5% (89.2-126.6%) for the range 2 to 508 μg/sample. No bias 

with dynamically-generated samples was observed with the method when samples were collected at 

40 mL/min and below. When samples were collected at ca. 60 mL/min, a negative bias of approximately 

20-30% was observed. Samples were found to be stable for at least 4 weeks when stored at room

temperature.

REFERENCES:

[1] Kennedy ER, Gagnon YT, Okenfuss JR, Teass AW [1988]. The determination in air of selected lowmolecular weight aldehydes as their oxazolidines by capillary gas chromatography. Appl Ind Hyg

3(10): 274-279.

[2] NIOSH [1984]. Glutaraldhyde: Method 2531. In: Eller PM, Cassinelli ME, eds. NIOSH manual of

analytical methods. 3rd. ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers

for Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication

No. 84-100.

[3] Lipari F, Swarin SJ [1982]. Determination of formaldehyde and other aldehydes in automobile

exhaust with an improved 2,4-dinitrophenylhydrazine method. J Chromatog 247:297-306.

[4] Budavari S, ed. [1989]. The Merck Index. 11th ed. Rahway, NJ: Merck & Co., Inc.

[5] Kennedy ER Hill RH, Jr. [1982]. Determination of formaldehyde in air as an oxazolidine derivative by

capillary gas chromatography. Anal Chem 54:1739-1741.

METHOD REVISED BY:

Yvonne T. Gagnon and Eugene R. Kennedy, Ph.D., NIOSH
VALERALDEHYDE: METHOD 2536, Issue 4, dated 3 March 2016 - Page 5 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPENDIX A: Synthesis of 9-Butyl-1-Aza-8-Oxabicyclo[4.3.0]nonane:

Place a solution of purified 2-(hydroxymethyl)piperidine (1.15 g; 10 mmol) in 20 mL of toluene in a 100- mL 

round-bottomed flask. Use several 2-mL portions of toluene to rinse residual 2- (hydroxymethyl)piperidine 

from the container used for weighing. Add anhydrous magnesium sulfate (2.0 g) to the flask to dry the 

valeraldehyde solution as it is added and to remove the water which forms during the reaction. Add a 

solution of 0.947 g valeraldehyde (11 mmole) in 20 mL of toluene to the 2- (hydroxymethyl)piperidine 

solution dropwise with stirring over 1 h. (NOTE: Excess aldehyde was added to ensure complete 

conversion of 2-(hydroxymethyl)piperidine to oxazolidine.) Stir the solution overnight, then filter to 

remove the magnesium sulfate. Remove the toluene from the solution at reduced pressure (1 mm Hg) by 

rotary evaporation. The product is a pale yellow viscous oil, ca. 90 to 95% pure by gas chromatography. 

Store the oxazolidine at 0 °C to prevent decomposition.

Mass spectral data for 9-butyl-1-aza-8-oxabicyclo[4.3.0]nonane: m/e with relative intensities in parenthesis, 

182 (7.0%), 152 (4.6%), 126 (100%), 110 (11.3%), 98 (37%). IR data (Vapor phase @ 280 °C) for this 

compound in cm-1 with relative intensity in parenthesis are: 2945 (s), 2874 (m), 2781 (m), 1455 (w), 1383 

(w), 1339 (w), 1265 (w), 1203 (w), 1133 (m), 1075 (w), 1028 (m).

APPENDIX B: Sorbent Preparation (optional if commercially-prepared tubes are used):

Add 1 g purified 2-(hydroxymethyl)piperidine in 50 mL toluene for each 9 g extracted XAD-2 sorbent. Allow 

this mixture to stand 1 h with occasional swirling. Remove the solvent by rotary evaporation at 37°C. Dry 

at 130 Pa (1 mm Hg) at ambient temperature for approximately 1 h. To determine the amount of 

background for each batch, desorb several 120-mg portions of the coated sorbent with toluene and 

analyze (steps 7 through 13). No blank peak is expected for valeraldehyde.

APPENDIX C: Desorption Efficiency: 

The determination of desorption efficiency (DE) is not necessary when using the calibration procedure in 

step 10, although the DE should be determined once for each lot of sorbent used, using the following 

procedure:

a. Prepare and analyze a set of valeraldehyde oxazolidine standard solutions (step 9.a) and a set of

working standards (step 10), including media blanks.

b. Treating the working standards as unknowns, read the mass (μg) of valeraldehyde oxazolidine found

in each working standard (W), and in the average media blank (B).

c. Using the mass of valeraldehyde, μg, spiked onto the working standard (Wo) and the stoichiometric

conversion factor of 2.13 between valeraldehyde and valeraldehyde oxazolidine, calculate the

desorption efficiency:

DDDD = WW − BB

WWoo ∙ 2.13

d. Prepare a graph of DE vs. μg valeraldehyde recovered per sample, (W - B)/2.13.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC AND INORGANIC GASES

by Extractive FTIR Spectrometry 

3800

FORMULA: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 3800, Issue 2 EVALUATION: FULL Issue 1: 15 March 2003

Issue 2: 13 January 2016

OSHA: Table 1

NIOSH: Table 1

PROPERTIES: Table 1

SYNONYMS: See example compounds in Table 1

SAMPLING

SAMPLER: PORTABLE DIRECT-READING 

INSTRUMENT (with filter, if required)

FLOW RATE: ~0.1 L/min to ~20 L/min 

(system-dependent)

VOL-MIN: Instrument dependent

-MAX: None

PRESSURE: Extracted gases between 96.6 kPa and 103 

kPa (725 mmHg and 795 mmHg) absolute

TEMPERATURE: Extracted gases between 10 °C and 30 °C

BLANKS: Nitrogen gas, or zero air

ACCURACY

RANGE STUDIED: See APPENDIX E

BIAS: See APPENDIX E

OVERALL

PRECISION ( ): See APPENDIX E

ACCURACY: See APPENDIX E

MEASUREMENT

TECHNIQUE: EXTRACTIVE FOURIER TRANSFORM 

INFRARED (FTIR) SPECTROMETRY

ANALYTE: See Table 1 (additional compounds 

may be identified/quantified according 

to data quality objectives and QA/QC 

requirements)

ANALYTICAL

FREQUENCIES: See Table 2 for example compounds

CALIBRATION: Calibration gas standards

IDENTIFICATION: Infrared spectra interpretation and 

computerized reference library searches

RANGE: See Table 2 (dependent on compound 

and absorption path length)

ESTIMATED LOD: See Table 2 for example values at 10 m 

absorption path length

PRECISION ( ): See APPENDIX E

APPLICABILITY: The usefulness of FTIR techniques has been demonstrated in ambient air and combustion gas mixtures 

[1,2]. With the participation of an experienced analyst (see APPENDIX A), the method can be used for the characterization of 

workspace air containing mixtures of volatile organic and inorganic compounds. See Table 1 for examples.

INTERFERENCES: Overlap of infrared absorption features may affect the quantification of each compound. By appropriate 

use of multivariable least squares analyses, the analyst may be able to obtain accurate compound concentrations for 

overlapping compounds.

OTHER METHODS: This method is based on portions of EPA method 320 and its addendum [3], which describe the 

determination of gaseous compound concentrations by extractive FTIR spectrometry. Several compatible ASTM standards 

describing infrared techniques and terminology are also available [4,5,6,7].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 2 of 44

REAGENTS:

1. Nitrogen gas or zero air, high purity or better.

2. Calibration transfer standard (CTS) gas, 2%

accuracy or better. The proper concentration

depends on both the compound used and the

system absorption path length. For ethylene in

nitrogen, a standard concentration leading to

a concentration-path length product (CPP) of

100 ppm · m to 400 ppm · m is recommended.

(For example, a standard of 10 ppm to 40 ppm

ethylene in nitrogen is recommended for a

10 m absorption cell).

3. Liquid nitrogen* for cooling the infrared

detector, if required.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. FTIR spectrometer system (source,

interferometer, sample absorption cell,

and detector) configured for absorption. A

minimum instrumental linewidth (MIL) of 2

 or lower is recommended.

NOTE: A lower or higher MIL may be required 

or suitable for specific gas matrices. 

Choose internal absorption cell 

materials to minimize surface/analyte 

interactions.

2. Computer system with hardware, software,

and required reference spectra for acquisition,

storage, and analysis of sample spectra. (A data

backup system is recommended.)

3. Sampling pump, 0.1 to 10 L/min, with

appropriate particulate filters.

4. Non-reactive gas regulators and sample

tubing.

5. Rotameters or other devices, 5% precision or

better, for measuring flow rates of sample and

calibration gases.

6. Temperature measurement and/or control

equipment for all sampling system elements

and IR absorption cell.

NOTE: Temperature control equipment

required if ambient temperature <10 °C 

or >30 °C.

7. Pressure gauge for measuring absolute gas

pressure in absorption cell, 5% accuracy

at 101.3 kPa (1 atm, 760 mmHg) absolute

pressure.

8. For system tests (but not for normal operation):

Vacuum pump and gauge capable of 13.3 kPa

(100 mmHg) absolute pressure; mid-infrared

attenuating filters (50% and 25%); impinger.

SPECIAL PRECAUTIONS: This method requires the use of compressed gases and/or cryogenic liquids 

and/or toxic chemicals. These materials are dangerous and should be handled only by experienced 

personnel and according to relevant safety standards. This method does not address all of the safety 

concerns associated with its use. It is the responsibility of the user to establish appropriate safety and 

health practices and to determine the applicability of regulatory limitations.

NOTE: Please refer to the appendices for explanations of the terminology used in this document 

(APPENDIX A) and several FTIR system tests, which must be completed before any testing is performed 

(APPENDIX B). Appendices C, D, and E provide (respectively) a general description of FTIR spectrometry, 

considerations for preparing reference libraries, and examples of calculations required for performance 

of this method.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 3 of 44

PRETEST PROCEDURES:

The following procedures (steps 1 and 2) may be performed only by an experienced “analyst” (see 

APPENDIX A).

1. Verify that the FTIR system has been tested for wavenumber reproducibility, MIL, absorption path 

length, system response time, residual squared area (RSA), and detector linearity as described in 

APPENDIX B. If the system is new and/or has been recently assembled and/or has been recently 

serviced, perform and document results of the tests described in APPENDIX B before proceeding.

2. Prepare a test plan. The plan must include the following:

a. The proposed system configuration, including the absorption path length and integration time 

for sample spectra.

b. The data quality objectives, analytical regions, and expected LOD values for each analyte. See 

Table 2 and APPENDIX E for example values and calculations.

c. The names of all “operators” and “analysts” to be involved in the test. Analysts must be 

experienced in all aspects of the test procedures listed below and may perform any or all of the 

test procedures. Operators must be experienced in the performance of steps 3, 4, 5, 6, 7, 8, 9, 10, 

11, 12, and 13 listed below and may perform only those portions of the test procedures.

d. Verification that the system configuration, existing RSA values, and related LOD values are 

consistent with the test’s data quality objectives. (See APPENDIX E.)

The following procedures (steps 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, and 13) may be performed by an 

experienced operator or analyst.

3. Activate the FTIR system according to the manufacturer’s instructions. Allow sufficient time for the 

infrared source, the infrared detector, and (if required) the temperature control systems to stabilize.

4. Verify that the computer system is programmed to store interferograms; if the available data storage 

capacity is insufficient for storage of all interferograms, store single-beam spectra instead. Verify 

adequacy and performance of the (recommended) data backup system.

5. Bypassing the sampling system, flow nitrogen or zero air directly into the infrared absorption cell 

until a stable infrared response and moisture levels are reached. Record a background spectrum 

using an integration time equal to or greater than that planned for subsequent sample spectra.

6. Use the entire FTIR system (including all sampling components) to record an absorbance spectrum 

of a sample stream of nitrogen or zero air. Use the integration time planned for subsequent sample 

spectra. Verify that the sample flow rate meets or exceeds that specified in the system response 

time documentation. Examine the resulting “system zero” (SZ) spectrum and verify the absence of 

contaminants in the sampling system and infrared absorption cell. If contaminants are detected, 

clean or replace the appropriate sampling system components and/or infrared absorption cell and 

record a new SZ spectrum. If contaminants cannot be eliminated, the test results and LOD values 

may require revision/correction during the quality control procedures (see steps 14, 15, 16, and 17).

CALIBRATION:

7. Using the entire sampling system, acquire two or more pretest CTS spectra and use them to 

calculate the system absorption path length (see APPENDIX B, section B1). Use the integration 

time planned for subsequent sample spectra. Verify that is within 5% of the value quoted in the 

test plan. Verify that the sample temperature and pressure are within 10 °C to 30 °C and 96.6 kPa to 

103 kPa (725 mmHg to 795 mmHg), respectively.

8. When possible, perform the following system checks before sampling. If necessary, they may be 

delayed until performance of the (post-testing) quality control procedures (see steps 14, 15, 16, 

and 17). If performed after sampling, the results of these procedures may require revision of the test 

results and LODs.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 4 of 44

a. LOD check. Using one of the SZ spectra, calculate the RSA values (see APPENDIX B, section 

B2) and LOD values (see Appendix E, section E1) in the analytical regions pertaining to the test 

analytes. Verify that the system path length (see APPENDIX B, section B1), current RSA values, and 

current LODs are consistent with the test’s data quality objectives (see Appendix E, section E1).

b. Wavenumber reproducibility check. Record a preliminary workspace air sample spectrum and 

perform the calculations described in Appendix B, section B4.

SAMPLING:

9. Using the integration time specified in the test plan, obtain samples and record infrared spectra 

of the desired workplace gases using the entire sampling system. The sampling location may be 

changed as desired. For tests of duration greater than 2 h, or if the FTIR system is moved during 

sampling, monitor the system’s single-beam response level. If changes greater than 5% in nonabsorbing regions of the single-beam spectrum occur, suspend sampling and record a new 

background spectrum (see step 5). Sample spectra must be acquired at each sampling location for a 

time period no less than the system response time (see the system test documentation).

10. Repeat step 6; acquire at least one posttest SZ spectrum of nitrogen or zero air; confirm the absence 

of sampling system contaminates.

11. Record at least one posttest CTS spectrum (see step 7); confirm that the system configuration and 

the system absorption path length (to within 5%) match those of the test plan.

12. (Optional.) Acquire a posttest background spectrum (see step 5).

SAMPLE ANALYSIS:

13. Using the analytical regions specified in the test plan, employ an appropriate mathematical analysis 

(see APPENDIX E) to determine preliminary analyte concentrations and their 3σ uncertainties from 

the sample spectra, reference spectra, absorption path length, and gas pressure.

NOTE: Reference spectra for all analytes must meet or exceed the QA/QC requirements of 

APPENDIX D. The reference library must include at least one wavenumber standard spectrum 

and at least one CTS spectrum. The sample absorption due to any analyte may not exceed the 

maximum CPP represented in the reference library for that compound.

QUALITY CONTROL:

The following procedures (steps 14, 15, 16, 17, and 18) may be performed only by an experienced 

analyst.

14. Using one of the SZ spectra, calculate the RSA and LOD values (see APPENDIX E) in the analytical 

regions pertaining to the test analytes. Verify that the system path length, current RSA values, and 

current LODs are consistent with the test’s data quality objectives. Using a suitable workspace air 

spectrum, perform the wavenumber reproducibility and resolution tests described in APPENDIX B, 

sections B3 and B4. If either the wavenumber reproducibility or resolution results fail to meet the 

specifications, corrective actions are necessary (see step 17).

15. Perform qualitative and/or quantitative analyses of the pretest and posttest SZ spectra and confirm 

the absence of sampling system contaminants. If contaminants are found that possess significant 

absorbance in any analytical region, corrective actions may be necessary (see step 17). Perform 

qualitative and/or quantitative analyses of the pretest and posttest CTS spectra and confirm that the 

system absorption path length is within 5% of the value quoted in the test plan.

16. Verify that the reference spectra and results meet the data quality objectives of the test plan (see 

APPENDIX D). If they do not, corrective actions may be necessary (see step 17). Examine the results 

of the quantitative sample analysis (step 13) and verify some subset of them (including those with 

relatively high and relatively low concentrations) manually to ensure proper operation of analytical 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 5 of 44

program. (Techniques for generating spectra representing the analytes at the concentration 

indicated by the analysis — “scaled spectra,” as defined in APPENDIX A — are described in 

APPENDIX E. The analyst should generate such spectra and compare them, either visually or 

mathematically, to the indicated sample spectra.)

17. Corrective actions: If the results of steps 14, 15, and 16 indicate that the data quality objectives of 

the test plan have not been achieved, the analyst may perform one or more of the following actions:

• Averaging of sequential sample spectra to reduce the RSA.

• Inclusion of contaminant reference spectra in the mathematical analysis.

• Development of more accurate analyte or interferant reference spectra.

• Inclusion of reference spectra for additional compounds in the mathematical analysis.

• Exclusion from the mathematical analysis of analyte and/or interferant compounds that are 

clearly absent in the samples.

• Deresolution or wavenumber adjustment of the reference spectral library to match that of the 

sample spectra (or vice versa).

• Revision of the original data quality objectives (those included in the test plan) to levels 

supported by the test data.

NOTE: After performing any such corrective actions, the analyst must repeat steps 13, 14, 15, and 16

and reevaluate the LOD value for each analyte compound.

REPORTING:

18. Reporting requirements include the analyte and interferant concentrations, the concentration 

uncertainties, the FTIR spectrometer configuration, the sampling locations and conditions, the 

source(s) of the reference spectra, the CTS spectral analysis results, the results of QA/QC procedures, 

and certificates of analysis for all standard gases. Any variations of the test procedures and original 

data quality objectives from those included in the test plan should also be documented and 

reported. (Several of the terms used here are defined in APPENDIX A and described in the following 

appendices. In particular, see Table D1 for a description of the FTIR spectrometer configuration 

parameters.)

EVALUATION OF METHOD:

Field evaluations of extractive FTIR methods have been performed for many compounds (see, for 

example, references 1 and 2) according to EPA method 301 [8].

REFERENCES:

[1] EPA [1993]. Fourier transform infared (FTIR) method validation at a coal fired boiler [www3.epa.gov/

ttnemc01/ftir/reports/r03.html].

[2] Reagen WK, Wright BD, Krueger DJ, Plummer GM [1999]. Fourier transform infrared method 

validation at a carbon bed solvent recovery unit for four gaseous hydrocarbons. Environ Sci Technol 

33(10):1752–1759.

[3] EPA [2014]. Method 320 — Measurement of vapor phase organic and inorganic emissions by 

extractive FTIR spectroscopy [www3.epa.gov/ttn/emc/methods/method320.html].

[4] ASTM [2012]. D6348-12e1: Standard test method for determination of gaseous compounds by 

extractive direct interface fourier transform infrared (FTIR) spectroscopy. West Conshohocken, PA: 

ASTM International [www.astm.org].

[5] ASTM [2015]. D1356-15b: Standard terminology relating to sampling and analysis of atmospheres. 

West Conshohocken, PA: ASTM International [www.astm.org].

[6] ASTM [2006]. E168-06: Standard practices for general techniques of infrared quantitative analysis 

(withdrawn 2015). West Conshohocken, PA: ASTM International [www.astm.org].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 6 of 44

[7] ASTM [2013]. E1252-98(2013)e1: Standard practice for general techniques for obtaining infrared 

spectra for qualitative analysis. West Conshohocken, PA: ASTM International [www.astm.org].

[8] EPA [2014]. Method 301 — Field validation of pollutant measurement methods from various waste 

media [www3.epa.gov/ttn/emc/methods/method301.html].

[9] EPA [1993]. EPA reference absorbance spectra; EPA reference spectra interferograms [www3.epa.

gov/ttn/emc/ftir/data.html].

[10] NIST [2016]. Quantitative infrared database [webbook.nist.gov/chemistry/quant-ir/].

[11] EPA [2016]. Software tools and viewers [www3.epa.gov/ttn/emc/ftir/tools.html].

[12] Griffiths PR, De Haseth JA [1986]. Fourier transform infrared spectrometry. New York: Wiley.

[13] Hamilton WC [1964]. Method of least squares and theory of linear hypotheses. In: Statistics in 

physical science; estimation, hypothesis testing, and least squares. New York: Ronald Press Co., pp. 

124–145.

[14] Plummer GM, Reagen WK [1996]. An examination of a least squares fit FTIR spectral analysis 

method. Unpublished paper (No. 96-WA65.03) presented at the 89th Annual Meeting of the Air and 

Waste Management Association, Nashville, TN, June 23–28.

[15] EPA [2012]. EPA traceability protocol for assay and certification of gaseous calibration standards. 

Research Triangle Park, NC: U.S. Environmental Protection Agency, Office of Research and 

Development, National Risk Management Research Laboratory, EPA/600/R-12/531 [cfpub.epa.gov/

si/si_public_record_report.cfm?dirEntryId=245292].

METHOD WRITTEN BY:

Grant M. Plummer, Ph.D. (Rho Squared), William K. Reagen, Ph.D. (3M), and Perry W. Logan (3M).

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

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Table 1. Representative volatile compounds and chemical data

Name Synonyms* Formula MW† CAS No. RTECS No.

BP 

(°C)†

VP at 

20 °C 

(kPa)†

Exposure Limits (OELs)*

NIOSH OSHA

Acetone Dimethyl ketone, 

Ketone propane, 

2-Propanone

58.1 67-64-1 AL3150000 56 24 TWA 250 ppm (590 

mg/m3)

TWA 1000 ppm 

(2400 mg/m3)

Ammonia Anhydrous ammonia, 

Aqua ammonia, 

Aqueous ammonia

17.0 7664-41-7 BO0875000 −33 1.01 

× 103 

(26 °C)

TWA 25 ppm (18 

mg/m3) ST 35 ppm 

(27 mg/m3)

TWA 50 ppm (35 

mg/m3)

Benzene Benzol, Phenyl 

hydride

78.1 71-43-2 CY1400000 80 10 Ca TWA 0.1 ppm ST 

1 ppm

[1910.1028] TWA 1 

ppm ST 5 ppm

2-Butanone Ethyl methyl ketone, 

MEK, Methyl acetone, 

Methyl ethyl ketone

72.1 78-93-3 EL6475000 80 10.5 TWA 200 ppm (590 

mg/m3) ST 300 ppm 

(885 mg/m3)

TWA 200 ppm (590 

mg/m3)

Carbon disulfide Carbon bisulfide 76.1 75-15-0 FF6650000 46 48 

(25 °C)

TWA 1 ppm (3 

mg/m3) ST 10 ppm 

(30 mg/m3) [skin]

TWA 20 ppm C 30 

ppm 100 ppm (30 

min maximum peak)

Ethylene oxide Dimethylene oxide; 

1,2-Epoxyethane; 

Oxirane

44.1 75-21-8 KX2450000 11 146 Ca TWA <0.1 ppm 

(0.18 mg/m3) C 5 

ppm (9 mg/m3) [10 

min per day]

[1910.1047] TWA 1 

ppm 5 ppm [15 min 

excursion]

Formaldehyde Methanal, Methyl 

aldehyde, Methylene 

oxide

30.0 50-00-0 LP8925000 −19.5§ 519‡

(3890 

mmHg, 

25 °C)

Ca TWA 0.016 ppm C 

0.1 ppm [15 min]

[1910.1048] TWA 

0.75 ppm ST 2 ppm

n-Hexane Hexane, Hexyl 

hydride, 

normal-Hexane

86.2 110-54-3 MN9275000 69 17 TWA 50 ppm (180 

mg/m3)

TWA 500 ppm (1800 

mg/m3)

Hydrogen fluoride Anhydrous hydrogen 

fluoride; Aqueous 

hydrogen fluoride 

(i.e., Hydrofluoric 

acid); HF-A

20.0 7664-39-3 MW7875000 20 122 

(25 °C)

TWA 3 ppm (2.5 

mg/m3) C 6 ppm (5 

mg/m3) [15 min]

TWA 3 ppm

See footnotes at end of table. (Continued)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

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Table 1 (Continued). Representative volatile compounds and chemical data

Name Synonyms* Formula MW† CAS No. RTECS No.

BP 

(°C)†

VP at 

20 °C 

(kPa)†

Exposure Limits (OELs)*

NIOSH OSHA

Methanol Carbinol, Columbian 

spirits, Methanol, 

Pyroligneous spirit, 

Wood alcohol, Wood 

naphtha, Wood spirit

32.0 67-56-1 PC1400000 65 12.3 TWA 200 ppm (260 

mg/m3) ST 250 ppm 

(325 mg/m3) [skin]

TWA 200 ppm (260 

mg/m3)

Methylene chloride Dichloromethane, 

Methylene dichloride

84.9 75-09-2 PA8050000 40 47.4 Ca TWA 25 ppm ST 125 

ppm

Nitrous oxide Dinitrogen monoxide, 

Hyponitrous acid 

anhydride, Laughing 

gas

44.0 10024-97-2 QX1350000 −88.5 5.06 

× 103

TWA 25 ppm (46 

mg/m3) (TWA over 

the time exposed) 

[Note: REL for 

exposure to waste 

anesthetic gas.]

Styrene Ethenyl benzene, 

Phenylethylene, 

Styrene monomer, 

Styrol, Vinyl benzene

104.1‡ 100-42-5 WL3675000 145 0.67 TWA 50 ppm (215 

mg/m3) ST 100 ppm 

(425 mg/m3)

TWA 100 ppm C 200 

ppm 600 ppm (5 min 

maximum peak in 

any 3 h)

Sulfur dioxide Sulfurous acid 

anhydride, Sulfurous 

oxide, Sulfur oxide

64.1 7446-09-5 WS4550000 −10 330 TWA 2 ppm (5 

mg/m3) ST 5 ppm (13 

mg/m3)

TWA 5 ppm (13 

mg/m3)

Tetrafluoroethylene Tetrafluoroethene; 

Perfluoroethylene; 

Perfluoroethene; TFE‡

100.0‡ 116-14-3 KX4000000 −75.9§ 3.27 

×103‡

(24500 

mmHg, 

25 °C)

Tetrahydrofuran Diethylene oxide; 

1,4-Epoxybutane; 

Tetramethylene oxide; 

THF

72.1 109-99-9 LU5950000 66 19.3 TWA 200 ppm (590 

mg/m3) ST 250 ppm 

(735 mg/m3)

TWA 200 ppm (590 

mg/m3)

See footnotes at end of table. (Continued)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

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Table 1 (Continued). Representative volatile compounds and chemical data

Name Synonyms* Formula MW† CAS No. RTECS No.

BP 

(°C)†

VP at 

20 °C 

(kPa)†

Exposure Limits (OELs)*

NIOSH OSHA

Toluene Methyl benzene, 

Methyl benzol, Phenyl 

methane, Toluol

92.1 108-88-3 XS5250000 111 3.8 

(25 °C)

TWA 100 ppm (375 

mg/m3) ST 150 ppm 

(560 mg/m3)

TWA 200 ppm C 300 

ppm 500 ppm (10 

min maximum peak)

Trichloroethylene Ethylene trichloride, 

TCE, Trichloroethene, 

Trilene

131.4 79-01-6 KX4550000 87 7.8 Ca TWA 100 ppm C 200 

ppm 300 ppm (5 min 

maximum peak in 

any 2 h)

Vinylidene fluoride Difluoro-1,1-ethylene; 

1,1-Difluoroethene; 

1,1-Difluoroethylene; 

Halocarbon 1132A; 

VDF; Vinylidene 

difluoride

64.0 75-38-7 KW0560000 −83 3.57 

×103*

(35.2 

atm)

TWA 1 ppm C 5 ppm 

[use 1910.1017]

m-Xylene 1,3-Dimethylbenzene; 

meta-Xylene; m-Xylol

106.2 108-38-3 ZE2275000 139 0.8 TWA 100 ppm (435 

mg/m3) ST 150 ppm 

(655 mg/m3)

TWA 100 ppm (435 

mg/m3)

o-Xylene 1,2-Dimethylbenzene; 

ortho-Xylene; o-Xylol

106.2 95-47-6 ZE2450000 144 0.7 TWA 100 ppm (435 

mg/m3) ST 150 ppm 

(655 mg/m3)

TWA 100 ppm (435 

mg/m3

p-Xylene 1,4-Dimethylbenzene; 

para-Xylene; p-Xylol

106.2 106-42-3 ZE2625000 138.4 0.9 TWA 100 ppm (435 

mg/m3) ST 150 ppm 

(655 mg/m3)

TWA 100 ppm (435 

mg/m3)

*Source: NIOSH Pocket Guide to Chemical Hazards [www.cdc.gov/niosh/npg/]. Date accessed: February 2016. †

Source: International Chemical Safety Cards (ICSC) [www.cdc.gov/niosh/ipcs/]. Date accessed: February 2016. ‡

Source: PubChem [pubchem.ncbi.nlm.nih.gov/]. Date accessed: February 2016. §

Source: Haynes WM, Lide DR, Bruno TJ [2013]. CRC handbook of chemistry and physics: a ready-reference book of chemical and physical data. 94th ed. Boca 

Raton, FL: CRC Press.
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Table 2. Representative infrared data for the compounds of Table 1

Compound

Analytical 

Region 

( )

Reference 

Spectrum 

Filename*

LOD at 

10 m†

(ppm)

Maximum 

concentration 

at 10 m‡

(ppm)

Maximum 

RSA§

( )

Reference 

Spectrum 

Source

Acetone 1163 to 1265 192mav01.spc 0.95 148 0.0211 EPA**

Ammonia 998 to 1131 nh3mav01.spc 0.77 470 0.0363 3M††

Benzene 3000 to 3150 015mav01.spc 0.32 149 0.036 EPA**

2-Butanone 1127 to 1235 mekmav01.spc 0.27 463 0.0233 3M††

Carbon disulfide 2109 to 2200 028mav01.spc 0.13 151 0.0123 EPA**

Ethylene oxide 3059 to 3070 084mav01.spc 0.11 138 0.0025 EPA**

Formaldehyde 2727 to 2844 087bb.spt 0.4 1125 0.0267 EPA** ‡‡

n-Hexane 2778 to 3051 095mav01.spc 0.1 150 0.0639 EPA**

Hydrogen fluoride 4034 to 4206 21hfrav 0.93 15.8 0.15 3M††

Methanol 941 to 1100 104mav01.spc 0.28 151 0.0447 EPA**

Methylene chloride 701 to 789 117mav01.spc 0.31 150 0.062 EPA**

Nitrous oxide 1226 to 1333 n2omav01.spc 0.36 904 0.0301 3M††

Styrene 738 to 944 147mav01.spc 1.84 150 0.0363 EPA**

Sulfur dioxide 1290 to 1410 so2.spc 0.35 ~200§§ 0.1394 NIST§§

Tetrafluoroethylene 1080 to 1215 tfemav05.spc 0.17 25.7 0.093 3M††

Tetrahydrofuran 2750 to 3085 thf405.spc 0.18 41 0.0782 3M††

Toluene 701 to 768 tolmav01.spc 1.16 463 0.0499 EPA**

Trichloroethylene 762 to 966 tcemav01.spc 0.43 464 0.1071 3M††

Vinylidene fluoride 1080 to 1215 dfemav05.spc 0.21 25.7 0.093 3M††

m-Xylene 782 to 805 172mav01.spc 1.36 146 0.0377 EPA**

o-Xylene 709 to 781 171mav01.spc 0.65 150 0.0444 EPA**

p-Xylene 749 to 840 173mav01.spc 1.17 151 0.0561 EPA**

*Used in LOD calculations; averaged spectra from the quoted libraries; data available from the authors.

†

Approximate LOD for a 10 m absorption path length. Typical values of the RSA, quoted analytical regions, and

quoted reference spectral data were used to calculate the LOD as described in Equation E1. Note: The LOD

may vary from laboratory to laboratory, analyst to analyst, instrument to instrument, and day to day. Therefore,

any determination of this value should be performed under the same conditions used for sample analysis and

reported only with those analyses. It is stressed that the values below are only conservative estimates of the

expected performance of this method.

‡

The maximum ppm · m value available for the compound in the quoted reference spectrum source. §

Maximum RSA in the specified analytical regions consistent with the quoted LOD values. See APPENDIX E,

section E1.

**See reference 9. Average absorbance and accepted standard concentration values for the low-concentration 

pairs of spectra were used. Interferograms were truncated to 0.5 resolution and triangular apodization 

was employed.

††Data available from the authors.

‡‡From EPA reference spectra recorded at 100 °C [9]. §§NIST quantitative infrared database [10]. The quoted maximum concentration for sulfur dioxide is based on

linearity studies of 0.5 resolution spectra with triangular apodization. Nonlinear absorbance leads to 

errors of less than 10% at 1000 ppm · m.
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APPENDIX A. TERMINOLOGY

absorbance — in terms of the incident intensity and transmitted intensity , the absorbance is given 

by . From a pair of FTIR single-beam spectra B (the background spectrum) and S (the 

sample spectrum), the sample absorbance for each wavenumber value (with index i) in the spectra is 

approximated by .

absorbance linearity — a characteristic of (ideal) absorbance spectrum; for such a spectrum, the 

measured absorbance is described by Beer’s law (Equation C1).

absorption band — a contiguous wavenumber region of a spectrum (equivalently, a contiguous set of 

absorbance spectrum data points) in which the absorbance passes through a maximum or a series of 

maxima.

absorption cell — a structure that contains a fluid sample, but allows light to pass through a sample at 

known temperature, pressure, and absorption path length.

absorption path length — the distance, measured in the direction of propagation of the beam of 

radiant energy, between the surface of the specimen on which the radiant energy is incident and the 

surface of the specimen from which it is emergent.

absorptivity — a measure of the fraction of the incident infrared radiation that is absorbed by a 

particular compound per molecule and per absorption path length; see Equation C1.

analyst — a person familiar with and experienced in performance of all aspects of this FTIR-based 

method. Analysts may perform any portion(s) of the method and must perform certain portions of 

the method (see also “operator”).

analyte — a compound whose concentrations in a sample is of interest and is to be accurately 

quantified (see also “interferant”).

analytical region — a contiguous wavenumber region (equivalently, a contiguous set of absorbance 

spectrum data points) used in the quantitative analysis for one or more analytes.

NOTE: The quantitative result for a single analyte may be based on data from more than one 

analytical region.

aperture — an optical device that physically restricts the diameter of the optical beam.

apodization — modification of the interferogram through its multiplication by a weighing function 

whose magnitude varies with the position of the interferometer’s moving element.

background spectrum — the single-beam spectrum obtained with all system components and without 

sample present (or in the presence of a non-absorbing gas replacing the sample).

baseline — any line (or smooth function of wavenumber) drawn on an absorption spectrum to establish 

a reference point that represents a function of the radiant power incident on a sample at a given 

wavelength.

Beer’s law — the direct proportionality of the absorbance of a compound in a homogeneous sample to 

its concentration. See Equation C1, which also describes the more general case of gas mixtures.

calibration transfer standard (CTS) gas — a gas standard of a compound used to measure the sample 

absorption path length; see step 7, step 11, APPENDIX B (section B1), and APPENDIX D (section D5).

 — see “wavenumber.”

compound — a substance possessing a distinct, unique molecular structure.

concentration — the quantity of a compound contained in a unit quantity of sample. The unit “ppm” 

(number, or mole, basis) is recommended and is equivalent to the volume basis for ideal gases.

concentration-path length product (CPP) — the mathematical product of concentration of the species 

and the absorption path length. For reference spectra, this is a known quantity; for sample spectra, 

it is the quantity directly determined from Beer’s law. The unit “parts per million · meters” (ppm · m) is 

recommended.

data quality objectives — parameters pertaining to a certain application of this method, including the 

estimated LOD values for each compound.

deresolve — to form spectra of lower resolution (higher full-width-at-half-maximum) from spectra of 

higher resolution (lower full-width-at-half-maximum); see reference 11 for a deresolution program.
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detector linearity — a characteristic of an (ideal) IR detector; for such a detector, the measured detector 

output voltage, when plotted against the total IR in a broadband IR signal incident on the detector, 

would form a straight line.

double-beam spectrum — a transmittance or absorbance spectrum derived by dividing the sample 

single-beam spectrum by the background spectrum.

NOTE: The term “double-beam” is used elsewhere to denote a spectrum in which the sample 

and background interferograms are collected simultaneously along physically distinct 

absorption paths. Here, the term denotes a spectrum in which the sample and background 

interferograms are collected at different times along the same absorption path.

extractive — the type of spectroscopy that includes extracting and transporting a sample stream from 

gases at a certain location to an absorption cell and isolating the sample in the absorption cell for 

analysis. Other types of spectroscopy in which the sample is not isolated in an absorption cell include 

“remote”, “open path”, and “local open path” techniques.

FFT (fast Fourier transform) — a discrete (digital) approximation to an FT (Fourier transform; see below) 

involving the factoring of the original data into sparse matrices containing mostly zeros.

filter — (1) A device, made of inert materials, that physically removes solid and liquid phase particles 

from a gas stream. (2) An optical device that transmits some fraction of the radiation incident on 

it; “neutral density” and “mesh” filters transmit an approximately constant fraction of the incident 

radiation at all wavelengths over a specified wavelength range.

frequency, ν — the number of cycles per unit time; for light, , where c is the speed of light 

and λ is the light’s wavelength. Unlike the speed and wavelength, which are medium-dependent, 

the frequency of light is independent of the medium through which the light travels. The term 

“frequency” is often used to denote the wavenumber (w, ) in FTIR spectroscopy because (in a 

given medium) the wavenumber is proportional to the frequency. (See APPENDIX C, section C4, and 

“wavenumber” in this appendix.)

FT (Fourier transform) — the mathematical process for converting an analytical (non-discrete) 

amplitude-time function to an amplitude-frequency function, or vice versa.

FTIR (Fourier transform infrared) spectrometer — an analytical system that employs a source of midinfrared radiation, an interferometer, an enclosed sample cell of known absorption path length, 

an infrared detector, optical elements that transfer infrared radiation between components, and 

a computer system. The time-domain detector response (interferogram) is processed by a Fourier 

transform to yield a representation of the infrared power vs. infrared frequency. See Figures C1 and 

C2.

FTIR spectrometry — use of an FTIR system to perform quantitative measurements.

FTIR system — the combination of an FTIR spectrometer and a sample interface.

FTIR system configuration — the set of parameters required to reproduce, as closely as possible, 

results from a particular FTIR system at a later time. This set includes (at least) the nominal MIL, the 

absorption path length, the apodization function, the gas temperature, the gas pressure, the zero 

filling factor, the measured wavenumbers of specific water absorption bands, the sources of the 

reference library spectra, the integration time, the detector type and serial number, the detector gain 

(including hardware and software settings).

FTIR system response time — the minimum time required for the output of an FTIR system to accurately 

reflect a sudden change in the sample gas composition; see APPENDIX B, section B5.

full-width-at-half-maximum (FWHM) — for a single, symmetric absorption band, the full width of the 

band in wavenumbers ( ) between its 50% relative maximum absorption levels.

impinger — a sample collection device, constructed of inert materials, that passes a gas stream through 

a liquid phase.

infrared detector — a device that (ideally) produces a voltage proportional to the total infrared power 

incident on it. Examples are 1) the mercury-cadmium-telluride (MCT) detector, which requires 

cooling (and is often cooled to liquid nitrogen temperature) and 2) the deuterated triglycine sulfate 

(DTGS) detector, often operated at ambient temperature.
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infrared source — a device that emits a pattern, stable in intensity and wavelength profile, of infrared 

radiation over a wide range of infrared wavelengths. High temperature filaments or ceramic 

elements, in conjunction with appropriate focusing optics, are often employed.

integration time — the total time over which the interferometric results of single scans are averaged 

to produce an interferogram (and its subsequent single- and double-beam spectra). Most software 

packages allow selection of the number of scans rather than the integration time. The integration 

time is approximately equal to (but is always less than) the time interval over which the selected 

number of scans is actually executed.

interferant — a compound whose presence in a sample spectrum must be taken into account to 

accurately determine one or more analyte concentrations, but whose concentration need not be 

accurately determined.

interferogram — record of the IR detector’s response to the modulated component of the interference 

signal measured as a function of retardation.

interferometer — device that divides a beam of radiant energy into two or more paths, generates an 

optical path difference between the beams, and recombines them in order to produce repetitive 

interference maxima and minima as the optical retardation is varied.

least squares fitting (LSF) algorithm — a calculation whereby one or more compound concentrations 

are estimated from a sample spectrum by minimizing the squared error in Beer’s law within a defined 

analytical region (see Equations C1, C2, C3, C4, C5, and C6).

limit of detection (LOD, ppm) — for a defined FTIR system configuration and sample matrix, an estimate 

of the lowest detectable concentration of a specific analyte based on the FTIR system’s RSA and the 

analyte’s integrated absorbance for a selected analytical region.

line — see “absorption band.”

linewidth — see “full-width-at-half-maximum” and “minimum instrumental linewidth.”

metering valve — a gas valve allowing reproducible adjustments of a gas stream flow rate on the order 

of 2% of the valves full flow rate.

mid-infrared — the region of the electromagnetic spectrum from approximately 400 to 5000 .

minimum instrumental linewidth (MIL) — for a given FTIR spectrometer and FTIR system configuration, 

the minimum measured FWHM for any absorption band. In wavenumbers, the MIL is often estimated 

as the reciprocal of the retardation expressed in cm. The MIL depends on the choice of apodization 

function and is often larger than the MIL estimated from the retardation.

multi-pass cell — an absorption cell that uses mirrors to pass the infrared radiation through the gas 

sample more than once; this leads to an absorption path length larger than the physical length of the 

cell (see also “White cell”).

operator — a person familiar with and experienced in performance of only some aspects of this FTIRbased method. Operators may perform many portions of this method, but specific portions of the 

method must be performed by an “analyst.”

peak — see “absorption band.”

qualitative analysis — examination of sample spectra to determine the presence or absence of 

particular compounds in a sample.

quantitative analysis — estimation of the actual concentrations of a specific set of compounds using a 

specific set of analytical regions.

reference spectra — absorption spectra of gases with known chemical compositions, recorded at a 

known absorption path length, which are used in the quantitative analysis of gas samples.

residual squared area (RSA) — a measure of the noise (random and systematic) and/or spectral 

artifacts for an absorbance spectrum in some analytical region; see APPENDIX B, section B2 for a 

mathematical definition. The RSA can be used to estimate the LOD for a given compound measured 

with a given FTIR system configuration.

retardation — optical path difference between two beams in an interferometer; also known as “optical 

path difference” or “optical retardation.” In the case of a standard Michelson interferometer, the 

retardation is simply twice the distance moved by a mirror in the interferometer during a scan.
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rotameter — a device indicating the volumetric flow rate of a gas by the vertical displacement of an 

object suspended by the gas stream.

sample interface — that part of the FTIR system that comes in contact with the sample and/or 

calibration gases. It includes the sample probe, sample filter, sample line, sample pump, gas valves, 

internal surfaces of the absorption cell, pressure gauge, sample rotameter, the vent lines, and the 

calibration components (gas cylinders, regulators, and rotameters).

sampling location — the point in space at which sample gases enter the sample interface.

scaling — application of a multiplicative factor to the absorbance values in a spectrum.

scan — digital representation of the detector output obtained during one complete motion of the 

interferometer’s moving assembly or assemblies.

single-beam spectrum — Fourier-transformed interferogram, representing the relative detector 

response vs. wavenumber.

NOTE: The term “single-beam” is used elsewhere to denote any spectrum in which the sample and 

background interferograms are recorded on the same physical absorption path; such usage 

differentiates such spectra from those generated using interferograms recorded along two 

physically distinct absorption paths (see “double-beam spectrum” above). Here, the term 

applies (for example) to the two spectra used directly in the calculation of transmittance and 

absorbance spectra of a sample.

system zero (SZ) spectrum — the absorbance spectrum of a non-absorbing gas (nitrogen or zero air) 

acquired using those portions of the sampling interface used to acquire actual sample gases.

transmittance, — the ratio of radiant power transmitted by the sample to the radiant power incident 

on the sample. Estimated in FTIR spectroscopy by forming the ratio of the single-beam sample and 

background spectra; often presented as % (100 × ) in spectral representations.

uncertainty — a mathematical quantity determined in an LSF procedure, used to estimate the likely 

error in the determination of the sample concentration in a procedure; see Equations C1, C2, C3, C4, 

C5, and C6.

wavelength λ, — the physical distance between successive maxima in the electromagnetic waves that 

comprise light. The wavelength and speed of light depend on the medium through which the light 

travels.

wavenumber, w — the reciprocal of the wavelength, also the number of wavelengths of light per unit 

length, usually expressed in the units . As is true of both the speed and wavelength of light, the 

wavenumber is dependent on the medium through which the light travels. (See APPENDIX C, section 

C4 and “frequency” in this appendix.)

wavenumber adjustment — reassignment of the values associated with single- and/or doublebeam spectra. Adjustments may be made locally by shifting or stretching the wavenumber scale, or 

globally stretched by changing the laser wavenumber during the FFT.

White cell — alternate name for a multi-pass absorption cell (see “multi-pass” above) indicating its 

inventor.

zero filling — the addition of zero-valued points to the end of a measured interferogram. In most 

computer programs, specification of a zero filling “factor” of results in an interferogram with 

times as many points as the original interferogram.
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APPENDIX B. SYSTEM TESTS

These procedures must be conducted at least once on new or significantly altered (by replacement of 

components, dis- and reassembly, etc.) systems. The tests described in sections B2 and B4 must also be 

repeated during either the pretest preparations or quality assurance procedures. In all cases, activate 

the FTIR system and allow sufficient time for the infrared source, the infrared detector, and (if required) 

the temperature control systems to stabilize before proceeding.

B1. Absorption Path Length

Obtain one or more absorbance spectra of a CTS gas (ethylene at 200 ppm · m to 300 ppm · m 

is recommended) at the gas temperature and pressure of a reference CTS spectrum of the same 

compound. For each spectrum, calculate the indicated absorption path length as

, (Equation B1)

where: = path length indicated by the sample CTS spectrum (m), 

 = path length of the reference CTS spectrum (m), 

 = area of the sample CTS spectrum ( ), 

 = area of the reference CTS spectrum ( ), 

 = pressure of the sample CTS spectrum (kPa), and 

 = pressure of the reference CTS spectrum (kPa).

When multiple CTS spectra are available, assign to the mean of the single-spectrum results. The 

reference CTS spectrum path length and concentration used must be based on multiple, high-quality 

gas standards and physical length measurements (see APPENDIX D, section D5). The analyst must 

document criteria for the selection of the analytical region and any baseline correction procedures 

employed.

B2. Residual Squared Area

NOTE: If the following calculations are performed during testing or as part of the QC procedures 

(steps 14, 15, 16, and 17), perform them using a workspace air spectrum instead of the “water vapor 

(absorbance) spectrum” described in the two paragraphs below.

Use the integration time selected for field testing in recording the spectra described below. Record a 

background spectrum of dry nitrogen gas or zero air. Using a suitable impinger, humidify the nitrogen 

or zero air stream and record a single-beam spectrum at an absolute pressure between 96.6 kPa and 

103 kPa (725 mmHg and 795 mmHg). Form the absorbance spectrum of this water vapor sample 

from the single-beam spectra. Assign the spectrum a unique filename and save it for the calculations 

described below.

From this water vapor spectrum, subtract a scaled spectrum (see APPENDIX A) formed from the 

water vapor reference spectrum to be used in subsequent quantitative analyses. The scaling factor may 

be varied to minimize the absorbance in the resulting difference spectrum in the various analytical 

regions to be used in the analyses. Subtract a constant offset, a linear function, or a quadratic function 

from the difference spectrum in each analytical region to form a residual spectrum R for each region. 

For each residual spectrum R with discrete absorbance value for wavenumber i = p to q, in the 

wavenumber range to , the RSA, , is defined as:

. (Equation B2)
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The RSA has the dimensions ( ) and serves as a measure of the integrated absorbance of spectral 

noise and water subtraction artifacts over the analytical region. The RSA is compared to the total 

absorbance of a compound in the same region to estimate the LOD for the compound in that region 

(see APPENDIX D, section D9 and APPENDIX E, section E1).

The calculation described above assumes that water is the only significant infrared absorber 

in the samples besides the analytes and that only one analyte absorbs in any analytical region. If 

other analytes or interferants are present, a more conservative RSA may be estimated by adding the 

absorbance of the additional compounds to the difference spectrum using a set of suitable reference 

spectra, then subtracting their absorbance using a different set of reference spectra.

B3. Minimum Instrumental Linewidth

Evacuate the absorption cell to a pressure below 13.3 kPa (100 mmHg) and record a background 

spectrum. Obtain a workspace air sample at an absolute pressure of approximately 40 kPa (300 mmHg). 

Record the absorbance spectrum of this low-pressure sample. Measure at the FWHM linewidth, in , 

of at least two isolated water vapor lines (for example, the lines near 1918 and 2779 ). The MIL 

is the mean of these FWHM measurements.

B4. Wavenumber Reproducibility

NOTE: If this calculation is performed during testing or as part of the QC procedures (see steps 6

and 10), perform these determinations using a workspace air spectrum instead of the water vapor 

absorbance spectrum described in section B2.

Using a water vapor spectrum recorded as described in section B2, determine the center 

wavenumber values and of two isolated water vapor absorption features; the peaks near 

1918 and 2779 are suggested, though any other pair of isolated lines separated by 500 

or more is suitable. Compare these results to those center wavenumber values and and for the 

same absorbance features in the water vapor wavenumber standard associated with the reference 

library to be used in quantitative analyses as follows: Calculate the wavenumber reproducibility, , in 

for each of the two absorption bands as

, (Equation B3)

for i = 1, 2.

Compare the maximum of these two values to the MIL for the FTIR system (see section B3). If the 

wavenumber reproducibility to MIL ratio exceeds 2%, adjustment of the wavenumber scale for the 

sample spectra may be required.

Mathematical wavenumber adjustments may be made locally by shifting or stretching the 

wavenumber scale, or globally stretched by changing the laser wavenumber during the FFT. However, 

large shifts (on the order of 5% or more of the MIL) indicate that the system requires physical 

adjustments, such as realignment of the laser system responsible for control of the interferometer’s 

moving element. In addition, mathematical wavenumber adjustments require some sort of 

interpolation procedure in conjunction with the quantitative spectral analysis, and those procedures 

may result in spectral mismatches whose effects on the accuracy of the analysis are not easily 

quantified.

The necessity of such wavenumber adjustments depends, in part, on the widths of the absorption 

peaks of the compounds involved in the spectral analysis. Because many of the absorption bands of 

water — a nearly ubiquitous interferant in workspace air IR analysis — are very narrow, an accurate 

analysis usually requires the relatively stringent limits placed above on the wavenumber reproducibility 

to MIL ratio. However, it is possible to obtain accurate results when this ratio exceeds the recommended 

limit, especially when only broad absorbance features are actually employed. The analyst may choose to 

approve analytical results obtained when this ratio exceeds the recommended limit.
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B5. System Response Time

Direct nitrogen or zero air through the entire sample interface and record spectra at approximately 

30 s intervals. Abruptly replace the nitrogen or zero air gas flow with CTS gas and continue to record 

spectra. The system response time is the subsequent time required for the FTIR system to generate an 

absorbance spectrum in which the CTS compound’s calculated concentration reaches 95% of the final 

(stable) concentration value indicated in later spectra.

B6. Detector Linearity

For the chosen optical configuration, attenuate the power incident on the detector by either 1) 

modifying the aperture setting or 2) placing filters (neutral density or mesh) in the infrared beam path 

(see Figure C1). At approximately 100%, 50%, and 25% of the system full IR power level, collect pairs of 

background and CTS spectra. Compare the areas of the CTS bands for the three spectra and verify that 

they are equal to within 5% of their mean value. If they are not, apply software linearization corrections 

to the interferometric data according to the manufacturer’s instructions. If this option is unavailable, it 

is necessary to either 1) characterize the system non-linearity and/or apply appropriate concentration 

corrections or 2) operate the system with attenuation sufficient to ensure linear detector response.
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APPENDIX C. GENERAL DESCRIPTION OF FTIR SPECTROMETRY

C1. FTIR Spectrometer Components

Figure C1 illustrates the basic FTIR spectrometer configuration required for gas phase analyses. The 

infrared radiation emitted by the infrared (IR) source contains energy at all wavelengths between 2.0 μm 

and 20 μm; this is the portion of the electromagnetic spectrum usually referred to as the “mid-infrared.” 

In the units typically employed in FTIR spectroscopy (wavenumbers, or ), this is the wavenumber 

range 5000 to 500 . The IR radiation passes through an interferometer, where the motion of an 

optical element, usually a mirror, optically modulates the infrared beam. The modulated IR beam then 

enters an absorption cell through a window (typically made of potassium bromide or zinc selenide) 

and interacts with the gases of interest. The physical length over which this interaction takes place is 

the “absorption path length.” In “multi-pass” (or “White”) absorption cells, mirrors within the cell direct 

the IR beam through the sample gas many times; in such cells, the absorption path length can be from 

4 to 50 (or more) times the cell’s physical length. (A larger absorption path length generally leads to 

greater sensitivity.) The IR beam then exits the sample cell via a second window and is refocused onto 

an IR detector. Because this extractive technique requires the transport of gas samples through the 

FTIR absorption cell, the design and integrity of the sampling system is of great importance. It is also 

important that the sampling system allow the operator to perform all the necessary calibration and 

sampling procedures without compromising the speed and flexibility of the analytical system.

Figure C1. FTIR spectrometer components and beam path.

Figure C2 illustrates a sampling system configuration that meets these requirements, though other 

configurations are possible. A computer controls the actions of the interferometer and records the 

voltage output of the IR detector throughout the motion of the interferometer’s optical element. Ideally, 

the detector voltage is proportional to the total power in the IR beam. The computer must accurately 

record the detector voltage as a function of the position of the moving element in the interferometer, 

so a secondary, laser-based optical system is usually used to measure the moving element’s position 

very precisely. In most circumstances, the motion of the mirror or other optical element is repeated 

many times and the resulting individual “scans” are “co-added” (averaged) to reduce the system’s RSA.
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Figure C2. Sampling components and sample paths.

A plot of the resulting IR power-vs.-position signal, referred to as an interferogram, is shown in 

Figure C3. This interferogram is the co-added result of 64 scans with only nitrogen gas (and some low 

level of water vapor) present in the absorption cell. Nitrogen is one of the few compounds that does 

not interact with infrared radiation, so this interferogram very closely represents the fundamental FTIR 

system response in the absence of a sample. Note that the signal is relatively large near the beginning 

of the interferogram, where the “zero phase difference (ZPD) burst” is located. The ZPD is often used to 

rapidly obtain an estimate of the IR signal strength during alignment of the optical system.

Figure C3. Mid-infrared interferogram.
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C2. The FFT

Every data point in the interferogram contains intensity information about every infrared wavelength 

transmitted from the source to the detector. It is possible to recover the intensity information as a 

function of wavelength through application of an FFT, from which the FTIR technique’s name is derived. 

This digital transformation of the interferogram can be thought of as the mathematical inverse of the 

optical modulation applied to the infrared beam as it passes through the interferometer. Its function 

is similar to that of the human brain and ear, which provide intensity information (loudness) versus 

wavelength (pitch) for complex signals (sound waves) incident on the eardrum. (Note that, as for an 

interferogram, each point in a complex sound wave contains intensity information about every pitch 

contained in the wave. Yet the ear and brain allow a symphony audience to immediately perceive, for 

instance, that the piccolo is playing very loudly while the tuba is playing very quietly.) Reference 12

(chapter 3) provides a complete mathematical description of the FFT.

C3. Instrument Resolution, Apodization Function, and MIL

Most software packages supplied with FTIR systems provide several options associated with the 

collection of data and application of the FFT. These typically include - at least - the nominal “instrument 

resolution” (specified in ) and the “apodization function” (e.g., “boxcar” and “triangular”). These 

parameters are very important in quantitative spectroscopy and are addressed in turn below.

The instrument resolution is the most fundamental and important instrument parameter. It specifies 

the nominal minimum FWHM, in , of any spectral “peak” (or “line”) in the final instrument output. 

Every FTIR instrument has a minimum FWHM determined by the maximum distance traversed by the 

interferometer’s moving element during a single scan. (For the basic Michelson interferometer, the 

FWHM in is equal to , where d is the distance in cm traversed by a moving mirror during a 

scan.) Clearly, instruments with low FWHM provide more spectral information than instruments with 

higher FWHM capability. However, this additional information comes at high costs associated with the 

design, construction, size, mechanical stability, portability, speed, and RSA of the instruments.

It is important to recognize the two uses of the word “resolution” in the nomenclature used to 

describe FTIR spectrometers: Instruments of high resolving power, or “high resolution,” provide 

spectral features of low FWHM; when the nominal resolution is specified in units of , a lower

specification corresponds to higher resolving power, or “higher resolution”. Most commercially available 

FTIR spectrometers suitable for field use provide FWHM values greater than or equal to 0.5 , that is, 

they are systems whose nominal spectral resolution is specified as a number higher than 0.5 . Most 

of the instruments capable of higher resolution (lower FWHM) are suitable for use only in very stable 

laboratory environments.

Standard FTIR operating software always provides options for recording spectra with FWHM values 

higher than the instrument’s actual lower FWHM limit. These options simply move the mirror (or other 

optical element) through only some fraction of its maximum possible travel. Operating the instrument 

in this manner results in larger FWHM values (“lower” resolution, and shorter interferograms) than the 

instrument is mechanically capable of providing. Spectra of lower resolution (higher FWHM) provide 

less information, but can be generated more quickly and, in most cases, with lower RSA than spectra of 

higher resolution.

The instrument operator can also choose the apodization function to be used in the generation 

of FTIR spectra. Apodization is a mathematical alteration of the interferogram that can be performed 

before application of the FFT. Several standard alteration functions have been devised and each affects 

the final absorption spectrum of the sample gas in a different way. As with the selection of instrument 

resolution, each choice has its advantages and drawbacks. The simplest choice, known as the “boxcar 

apodization” function, results in the lowest FWHM but also in relatively low signal to noise (S/N) ratio. 

(Spectra generated with the boxcar function are often referred to as “unapodized” spectra.) Other 

choices (triangular, Norton-Beer, and several other apodization functions) provide higher S/N ratio 
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at the cost of higher FWHM values and other trade-offs in quantitative spectroscopy. Reference 12

provides a more thorough description of the characteristics of various apodization functions.

For a given instrument configuration — which includes the nominal spectral resolution and the 

choice of apodization function — every FTIR system is capable of generating absorption bands with 

an MIL. Unlike the actual spectral resolution (which has several accepted physical definitions — see 

reference 12, chapter 1, section IV) and nominal spectral resolution parameters, the MIL is a parameter 

that is readily measured to the accuracy required for practical applications of FTIR spectrometry. It can 

be measured using the water absorption bands present in low-pressure workspace air samples (see 

APPENDIX B, section B3).

C4. Single-Beam Spectra

The mathematical result of the FFT (applied to an apodized IR interferogram) is a called a singlebeam spectrum. Single-beam spectra represent the infrared power transmitted through the FTIR 

spectrometer as a function of the infrared “wavenumber” w, which is usually expressed in the units of 

reciprocal centimeters ( ). The wavenumber is actually a measure of the frequency, rather than the 

wavelength, of the infrared radiation. In a vacuum, the wavelength and frequency are related through 

the equation , where λ is the wavelength (cm), ν is the frequency ( , or Hz), and c (cm/s) is the 

speed of light, equal to 2.99792954 × in a vacuum. In these units, the wavenumber in is given 

by the equation . Figure C4 presents the single-beam spectra for two samples consisting 

mainly of nitrogen gas (≥99%) but with different concentrations (≤1%) of water vapor. The vertical 

scales of the two spectra are nearly identical, but they have been offset for clarity. The detected infrared 

power in certain wavenumber regions is clearly lower in the high moisture sample, indicating both the 

qualitative nature and strength of water’s absorption of infrared radiation.

Figure C4. Single-beam spectra of nitrogen at different moisture levels.

C5. Double-Beam Spectra — Transmittance and Absorbance

Combinations of pairs of single-beam, such as the pair of spectra S and B shown in Figure C4, are 

referred to as double-beam spectra; they provide the quantitative basis for FTIR spectroscopy. One type 

of double-beam spectrum is the transmittance. The percent transmittance of a sample gas possessing 

the single-beam spectrum S — with respect to the background single-beam spectrum B — is defined 

as (%), where and are the respective transmitted intensities; a transmittance value is 
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defined for each wavenumber value of the two spectra. If the background spectrum B closely represents 

the response of the FTIR system to a transparent sample, then the percent transmittance closely 

approximates the percentage of the infrared radiation transmitted by the sample (represented by the 

spectrum S). Because water is the only absorbing compound present in the single-beam spectrum B, 

the spectrum transmittance (shown in Figure C5) closely approximates the percent transmittance 

spectrum of water.

Figure C5. Double-beam transmittance spectrum of water.

The same pair of spectra define the (double-beam) absorbance of the sample through the 

equation . The absorbance spectrum of water, as approximated by the two single-beam 

spectra S and B, is shown in Figure C6. The absorbance is the desired quantity because it appears in the 

general linear absorption model known as Beer’s law (see below).

Figure C6. Double-beam absorbance spectrum of water.
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C6. Beer’s Law

The absorbance spectrum of a sample gas is determined from single-beam spectra of an infraredtransparent gas (the background spectrum) and the sample gas (see section C5). The fundamental 

relationship connecting the absorbance spectrum of a sample gas to the concentrations of the 

compounds comprising the sample is referred to as Beer’s law. This relationship can be written as

, (Equation C1)

where: i = an index labeling the frequencies of the observed absorbance values in the sample’s 

absorbance spectrum,

 = the observed absorbance of a sample at the i

th wavenumber,

= the sample absorption path length (m),

j = an index labeling the absorbing compounds comprising the sample,

 = the absorptivity of the j

th compound at the ith wavenumber (ppm/m),

 = the volumetric concentration of the j

th compound (ppm), and 

= the number of absorbing compounds in the sample.

The procedures described in this method are all related to the parameters that comprise Beer’s law. 

The following list describes these procedures and relationships in a general context:

1. Record a reference absorbance spectrum for each interesting compound of a nitrogen-diluted 

sample of the pure compound, at a known concentration and path length, then divide the 

absorbance at each point in that spectrum by the CPP. This process yields the absorptivity spectrum 

( ), or reference spectrum, for each compound.

2. Measure the absorbance for the mixture of compounds (see steps 5 and 9 above).

3. Determine the path length for the current measurement of (see steps 5 and 7 above).

4. Select the analytical region — that is a set of frequencies, corresponding to the possible values 

of the index i — that are to be used to determine the concentration of each compound, and then 

mathematically invert Equation C1 to determine the desired concentrations . (APPENDIX E

addresses the topic of spectral analysis in detail.)

NOTE: The true absorptivity for a single gaseous compound is a characteristic only of the compound’s 

structure. However, details of the FTIR system performance and operation affect the observed 

absorptivity and its accuracy. Similarly, FTIR measurements provide only an approximation of the true 

absorbance spectrum of a mixture of gaseous compounds, though it is, under many circumstances, a 

sufficiently accurate approximation. It is the responsibility of the analyst to verify and ensure that the 

reference and sample spectra provide a sufficiently accurate quantitative analysis according to Beer’s 

law. The following sections of this appendix describe the mathematics of such an analysis. APPENDIX D

addresses the topics of developing and using reference spectral libraries. APPENDIX E provides an 

illustrative example of the design and evaluation of the quantitative analytical process.

C7. Determining Concentrations with LSF Algorithms

When a sample gas contains only one absorbing compound, Equation C1 simplifies to

. (Equation C2)

This means that in any analytical region where only one gas absorbs, any one (of the usually many) 

absorbance spectrum values can be used to yield the concentration .

The absorbance area for a single-component spectrum in an analytical region (from i = p to i = q) 

can be written as

, (Equation C3)
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where is the area in the reference spectrum for that compound in the same analytical region. (This is 

the basis of the absorption path length calculation described in step 7 and APPENDIX B, section B1.) 

Because calculation of the absorbance area involves many points in the sample spectrum, Equation C3

leads to much more accurate results than the single-point calculation represented by Equation C2.

However, when many absorbing compounds are present in a sample, the absorption patterns of the 

various compounds often overlap. In this case, there is usually not an isolated analytical region for each 

compound in which only that compound absorbs infrared radiation; no single absorbance point and no 

simple absorbance area is suitable for determining any of the component concentrations. In this case, 

the simplest method for determining concentrations is to use an LSF algorithm.

LSF algorithms use the fact that there is some set of estimated concentrations that minimizes 

the “squared error” in Beer’s law for any given analytical region, for any set of compounds. The only 

requirement on the chosen analytical region is that it must contain a sufficient number of data points; 

since each FTIR spectrum contains many thousands of absorbance values, this requirement is nearly 

always fulfilled.

If we use the estimated concentrations (rather that the true concentrations ) in Beer’s law, they 

will lead to some estimated error at each value of i (that is, at each point in the analytical region we 

choose). Equation C1 becomes:

. (Equation C4)

The estimated squared error (or “variance”) in Beer’s law using the estimated concentrations is:

, (Equation C5)

where represents the number of absorbance values in the analytical region. Reference 13

demonstrates that 1) for > there is a unique set of estimated concentrations that minimizes the 

estimated squared error; 2) this set of values is calculable from the known quantities in Equations C1, 

C2, C3, C4, and C5; and 3) estimates of the uncertainties in the quantities are also calculable from 

the same quantities. The value is generally accepted as a conservative estimate of the statistical 

uncertainty in the related estimated LSF concentration (see reference 4).

The estimated LSF error at each point in the analytical region,

, (Equation C6)

is usually stored following the analysis as a “residual spectrum,” which can provide an estimate of the 

LODs for other compounds. In addition, the residual spectrum and the concentration uncertainties 

can allow the analyst to detect and identify compounds that are actually present in the sample gas but 

which were not included in the mathematical analysis. APPENDIX E provides an example illustrative of 

these procedures.

The above description illustrates a simple and easily interpreted LSF analysis. More sophisticated 

LSF analytical techniques, possibly more accurate for particular types of samples, are described in the 

literature (see, for example, reference 14 and references therein).

C8. Calibration Transfer and Reference Libraries

Equations C1, C2, C3, C4, C5, and C6 demonstrate the importance of quantities (the absorption path 

length) and (the absorptivity) in FTIR spectrometry. Accurate determinations of these quantities allow 

the use of reference libraries for quantitiative analyses without the necessity of compound-specific 

field calibrations. The system tests described in the procedures and in APPENDIX B are intended to 

ensure suitability of the system configuration for such calibration transfers, as are the requirements 
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of obtaining CTS spectra in field. APPENDIX D describes procedures for recording and processing 

reference library spectra.

C9. Corrections to Deviations from Beer’s Law Exhibited by FTIR Spectra

Beer’s law is based on fundamental, well-established physical principles. It holds absolutely for gas 

samples that are at thermal equilibrium and dominated by induced (rather than spontaneous) emission 

and absorption processes. (See Note below.) However, this is not to say that the absorbance, as measured 

by an FTIR spectrometer, follows Beer’s law under all conditions. Deviations from Beer’s law in FTIR spectra 

are often observed; however, they indicate inaccuracies in the FTIR spectra, not “violations” of Beer’s law. 

For example, deviation from Beer’s law is commonly exhibited by sets of single-component reference 

spectra recorded over a range of absorbance levels. At large enough values of the absorbance, the 

points of stronger absorption bands of such spectra no longer increase linearly with the CPP ; 

this is why Table 2 specifies a maximum ppm · m value for the listed reference spectra. If the assumption 

of detector linearity does not hold (see APPENDIX B), similar effects are often present in reference and 

sample spectra; this is the basis for the system test described in APPENDIX B, section B6.

Mathematical correction of the concentration estimates derived from Beer’s law can often reduce 

the error in sample analyses when either type of non-linear effect occurs. Figure C7 provides an 

example of such a correction. The actual and calculated ppm · m values for a set of reference spectra 

are plotted against each other, a “piece-wise linear” approximation to the pattern is shown by the 

solid line, and the dashed line indicates the ideal linear behavior based on the spectrum of lowest 

absorbance. At any ppm · m value indicated in a Beer’s law sample analysis (that is, for any y-axis value 

up to approximately 900 ppm · m in the example), reasonably accurate values are available from the 

corresponding x-axis position of the solid line. If the analyst employs such corrections, he or she should 

also calculate the residual (Equation C6) using the reference spectrum that minimizes the squared error 

(Equation C5). FTIR analysts and manufacturers have devised other correction procedures to improve 

the sample analysis accuracy in such circumstances, and these are included in some commercially 

available software packages.

Figure C7. Example absorbance non-linearity for a set of single-component reference 

spectra.

NOTE: Intense infrared radiation, such as that produced by some lasers, can induce non-equilibrium 

populations of a molecule’s rotation-vibration energy states. However, the thermal infrared sources 
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used in commercially available FTIR spectrometers are much weaker than such lasers. In gases at 

atmospheric pressure, the commercially available thermal sources induce transition rates between 

quantum energy states that are small, compared to those of the competing collisional relaxation 

processes, and cannot induce non-equilibrium energy state populations. In addition, the induced 

transition rates related to absorption and emission at mid-infrared frequencies are much larger than 

the corresponding spontaneous (natural) emission rate. As a result, all accurate FTIR measurements at 

mid-infrared frequencies obey Beer’s law to within the uncertainty related only to the S/N ratio of the 

measured spectra.
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APPENDIX D. PURPOSES, PREPARATION, AND USE OF REFERENCE SPECTRAL LIBRARIES

D1. Purposes of Reference Spectral Libraries

FTIR analyses rely on the availability of libraries of spectral information on the compounds of interest. 

For gases, the measured absorbance of a single component is often completely independent of the 

concentrations of other gases comprising the sample, and single-component reference spectra are 

usually employed. (For condensed phases, there are often strong interactions between components, 

and reference spectral libraries of mixtures are usually required.) Reference libraries may be used for 

quantitative measurements of analyte concentrations, for the mathematical removal of spectral features 

of interferants in a mixture, or simply for the identification of compounds in a mixture. Clearly, the 

required level of quantitative accuracy of the library is different for these three tasks; the highest quality 

is required for analyte concentration determinations, while no quantitative information is required for 

interferant removal and compound identification.

A useful characteristic of extractive FTIR spectrometry is that it provides accurate field measurements 

for many compounds, but requires field calibration procedures involving only two compounds. The 

water vapor available in every ambient air sample serves to calibrate the wavenumber (x) axes of FTIR 

absorbance spectra; a single CTS gas serves to calibrate their concentration-related (y) axes. When these 

two field calibrations are combined with an appropriate reference spectral library describing additional 

compounds, the measurement capability of the technique is practically limited only by the quality 

and scope of the reference library. If the reference library is carefully prepared and properly employed, 

this characteristic can lead to greatly lowered field test costs, since the calibration materials need be 

handled only once and only in the laboratory.

D2. Reference and Field FTIR System Configurations

In the ideal case, the reference library can be prepared on the field instrument, but this is often 

impractical; the reference library is usually prepared on specific laboratory systems and employed in 

measurements made with many field systems. Reference libraries recorded on a specific instrument 

provide accurate quantitative analyses for spectra recorded on other instruments only when the 

configurations of the various systems are compatible. Table D1 lists compatibility considerations for the 

reference system configuration parameters.

D3. Wavenumber Standard Spectra

Except in extremely dry conditions, the IR absorption by water vapor in workspace air is detectable at 

even very short absorption path lengths. The wavenumber positions of water vapor indicated by the 

reference system allow the analyst to confirm the wavenumber accuracy of sample spectra recorded on 

a different FTIR system in the field. Reference spectra used for this purpose are referred to here as the 

wavenumber standard spectra.

Every spectrum in a reference library should be associated with a wavenumber standard spectrum 

collected under the identical system configuration; even when the reference FTIR system is in a stable 

environment, it is recommended that a wavenumber standard spectrum be recorded daily with the 

system.

Before recording reference spectra, the analyst should check the most recent wavenumber standard 

spectrum against previous wavenumber standard spectra in the library; see APPENDIX B, section B4

for recommended calculations. If the wavenumber reproducibility of the system is poor, the resulting 

reference spectra may not be suitable for quantitative use on other field systems.
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Table D1. Reference and field system configuration parameters

Parameter Requirements for Reference and Field Systems

Minimum 

instrumental 

linewidth

Reference MIL must be less than or equal to field MIL. See APPENDIX B, section B3 for 

MIL measurement technique.

Gas 

temperature

Reference temperature within 20 °C of field temperature. Density corrections based 

on ideal gas law are accurate over only this narrow temperature range and their exact 

accuracy is compound-dependent.

Gas pressure Reference pressure within 20% of absolute field pressure. Pressure corrections based 

on ideal gas law are accurate over at least this range, but their accuracy is compounddependent. Atmospheric pressure is recommended for all measurements.

Apodization 

function

Reference and field apodization functions must be the same. A single set of reference 

interferograms (background and sample) can be used to generate a multiple sets of 

absorbance spectra using different apodization functions.

Zero filling 

factor

Reference and field zero filling factors must be the same. A single set of reference 

interferograms (background and sample) recorded with no zero filling can be used to 

generate a multiple set of absorbance spectra using different zero filling factors.

Wavenumber 

accuracy

Characterized by the position of water absorption bands in a wavenumber standard 

spectrum (see below); if additions to an existing library are being made, care must be 

taken to match the x-axes of all spectra as closely as possible.

Integration 

time

Reference integration times (for both background and sample interferograms) should 

be greater than or equal to expected field integration times. For reference libraries, the 

longest practical integration time is recommended.

Detector 

type and 

serial 

number

If an MCT or other potentially non-linear detector is employed in the reference system, 

IR attenuation may be required to ensure linearity; see APPENDIX B, section B6; 

because each detector may exhibit linearity to a specific degree, the serial number of 

the detector should be included in any specification of the system configuration.

D4. Obtaining and Preparing Standard Reference Gases

The accuracy of any FTIR quantitative analysis is limited by the accuracy of the concentrations of the 

gas standards used in preparing the reference library. It is therefore important to use gas standards of 

the highest available quality. NIST-traceable gravimetric standards are available from many commercial 

sources and are quoted to 2% accuracy in many circumstances; EPA has also published guidance on 

the preparation of “EPA Protocol Gases” (see reference 15), though these are available for only a limited 

number of compounds. Users should obtain documentation from the gas vendors regarding the 

analytical techniques applied and the stability limits (concentration and time) of the compounds in the 

cylinders. When possible, an alternative analytical method should be used to verify the quoted cylinder 

concentrations, especially for compounds that are reactive, corrosive, or have relatively high boiling 

points.

If cylinder standards are not available for a compound, the analyst may prepare reference gases 

based on gravimetric, barometric, or dynamic methods. In any case, the related mass, pressure, and flow 

measurements should be made with NIST-traceable equipment whenever possible. In general, methods 

that provide a stream of standard gas (dynamic methods) provide more reliable results than static 

methods, since the concentrations of static gases are more prone to change when reactions between 

the standards and the sampling system or absorption cell surface can occur.
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D5. Determining the Reference Absorption Path Length

The accuracy of any FTIR quantitative analysis is limited by the accuracy to which the path length of 

the absorption cell used in preparing the reference library is known. For single-pass absorption cells 

through which a collimated IR beam is passed, the path length can be physically measured to high 

accuracy. For multi-pass cells, the nominal path length can be estimated from the base path length 

and the number of passes. However, because focused beams and curved mirrors are required in multipass cells, the path length estimated in this fashion can differ significantly from the actual path length. 

Because of this, combinations of physical and spectroscopic measurements with multiple CTS gas 

standards (see below) and single-pass absorption cells should be employed to determine the actual 

path length of multi-pass absorption cells.

D6. Recording CTS Spectra

The CTS gas is used to characterize the absorption path lengths of the reference and field FTIR 

systems. Ethylene in nitrogen, at concentrations leading to between 100 ppm · m and 300 ppm · m 

CPP values for the systems, is recommended; for ethylene, spectra with ppm · m values greater than 

300 begin to exhibit non-linear absorbance and must be carefully analyzed (see sections D8 and 

D9 below). Standards of this gas with 2% accuracy are readily available and both the EPA and NIST 

libraries contain ethylene spectra that can be used to link those libraries to measurements performed 

with field instruments. However, almost any other stable compound with appropriate IR absorption 

characteristics can be used in developing an independent reference library and for accurate field 

measurements.

In the field, CTS spectra must be recorded both before and after sample spectra are acquired using 

the same system configuration employed in recording the sample spectra (see steps 7 and 11 above).

When developing reference spectra, the analyst must record CTS spectra at least daily; each reference 

spectrum should be associated with at least one CTS spectrum. The analyst must use the same system 

configuration employed in recording the reference spectra, with one exception: The integration time 

used for the CTS spectra may be less than that used for the reference spectra, if the quality of the 

resulting CTS spectra is still sufficient for an accurate determination of the absorption path length. It 

is highly recommended that the analyst store all the interferograms from which the CTS absorbance 

spectra are generated, including all background interferograms. The interferometric data provide the 

most direct method of verifying the FFT calculations and/or adding reference absorbance spectra for 

other apodization function choices.

D7. Recording Reference Spectra

Before recording reference spectra, verify that the requirements specified for system checks, system 

configuration parameters, gas standards, absorbance path length determinations, and CTS spectra 

described in the pertinent sections of this appendix and Appendix B have been met. Additional checks 

of the sample gas pressure and temperature should also be made periodically as the spectra are 

recorded.

It is highly recommended that the analyst store all the interferograms from which the reference 

absorbance spectra are generated, including all background interferograms. The interferometric data 

provide the most direct method of verifying the FFT calculations and/or adding reference absorbance 

spectra for other apodization function choices.

Beer’s law (Equation C1) describes the fundamental linearity of the infrared absorbance versus 

concentration. However, the resolution (and other) limitations of instruments typically used to generate 

field FTIR spectra often lead to non-linear behavior for many compounds, even at low absorbance 

levels. The expected accuracy of FTIR-based measurements results for a particular compound can be 
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achieved only when 1) the reference absorbance is characterized at a sufficient number of CPP values, 

up to some maximum value and 2) the sample absorbance associated with any compound represents 

a value lower than that of the maximum CCP value represented in the reference library for that 

compound.

For each analyte, the analyst should record two reference spectra at each of two concentration values 

(that is, at least four spectra) at a single absorption path length; the two concentrations should be 

separated by no more than a factor of 10, and a maximum factor of 5 is recommended. After recording 

these spectra, the analyst must 1) permanently record the system parameters and the maximum 

measured CPP and 2) characterize the linearity of the absorbance across the measured CPP range (see 

section D8). It may be necessary to record additional reference spectra for the analyte if either the 

absorbance linearity or maximum CPP value proves unsuitable for later field measurements.

D8. Linearity Checks

The performance of linearity checks on a set of reference spectra is an important aspect of FTIR 

spectrometry and is best illustrated by a practical example. The following example is based on a series 

of reference measurements performed on the compound tetrafluoroethylene, hereafter referred to as 

TFE.

Five absorbance spectra for TFE are shown in Figure D1. These spectra were recorded over the 

entire mid-IR wavenumber range, but only that portion of the spectra showing the most intense TFE 

absorption bands is included in the figure. Each of the spectra was calculated by mathematically 

averaging a number of TFE absorbance spectra recorded with the system configuration parameters 

given in Table D2, and a linear baseline correction was applied to each of these spectra over the range 

illustrated in Figure D1.

Figure D1. Reference absorbance spectra of TFE (x-axis truncated).
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Table D2. System configuration parameters for TFE reference spectra (see Figure D1)

System Configuration Parameters Values/Notes

MIL ( ) 0.5

Integration time (s) 1080 (120 s per spectrum × 9 spectra)

Absorption path length (m) 10.23

Gas temperature (°C) 15

Gas pressure (kPa) 101.3 ± 1.3 (760 mmHg ± 10 mmHg)

Apodization function Triangular

Zero filling factor Zero

Detector type MCT

Detector gain (hardware) Jumpers B, C, and D

Detector gain (software) 1

Reference CTS spectra j2kety.spc

Wavenumber standard spectrum j2kety.spc

The reference gas samples were generated by diluting with nitrogen the output of a single standard 

gas cylinder of TFE. The TFE concentration of the standard cylinder was determined (by the gas vendor) 

according to NIST-traceable gravimetric measurements and the flow rates of the cylinder gas were 

measured with a NIST-traceable volumetric device.

One technique to check for the linearity of this set is to form a normalized average of these five 

spectra and then use it in a linear analysis of the five original spectra. The normalization consists of 

dividing each spectrum by its CPP; these values, based on the CTS-derived path length of 10.23 m, are 

given with the actual gas concentrations and new spectral filenames in Table D3. Figure D2 illustrates 

the normalized spectra, all of which represent spectra of CPP values 1.00 ppm · m. These spectra are 

nearly equal to one another, as predicted by Beer’s law. Only the spectrum tfe01n.spc, based on the 

original 25.53 ppm spectrum, is easily discernible from the other spectra in the figure.

Table D3. Normalization factors for TFE spectra

Original Spectrum

Concentration 

(ppm)

ConcentrationPath Length Product 

(ppm · m)

Normalized 

Spectrum

tfe01.spc 2.50 25.5 tfe01n.spc

tfe02.spc 6.63 67.8 tfe02n.spc

tfe03.spc 13.3 133 tfe03n.spc

tfe04.spc 20.4 208 tfe04n.spc

tfe05.spc 25.1 257 tfe05n.spc
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Figure D2. Normalized reference absorbance spectra for TFE.

Table D4 shows the results of a linear least-squares analysis using the average normalized spectrum 

as a single reference spectrum (with CPP 1.00 ppm · m) over the analytical range 1050 to 1400 . 

The bias in the linear method is clearly evident in the fourth column of the table, which indicates that 

the analysis yields concentrations that are slightly too high at the lowest concentration and slightly 

too low at the highest concentration. The average of the last column in the table, which represents 

the absolute percent difference between the actual and calculated concentrations, the “fractional 

calibration uncertainty” (FCU; see reference 3), is 3.2%. This value represents the average error over 

the concentration range for these reference spectra to be expected from the linear analysis up to the 

maximum CPP value used in the analysis (257 ppm · m). If additional accuracy is required, non-linear 

analyses or corrections such as those discussed above (Appendix B, section B6) can be employed.

Table D4. Results of linear least squares analysis for TFE reference spectra

TFE Reference 

Spectrum

Calculated 

Concentration 

(ppm)

Actual 

Concentration 

(ppm)

Actual − 

Calculated 

Concentration 

(ppm)

Absolute 

Percent 

Difference

tfe01.spc 2.69 2.50 0.19 7.7

tfe02.spc 6.64 6.63 0.01 0.2

tfe03.spc 12.8 13.0 −0.2 1.4

tfe04.spc 19.8 20.4 −0.6 2.9

tfe05.spc 24.2 25.1 −0.9 3.7

D9. Calculating the LOD

The estimated LODs are the concentrations — for the given system configuration, reference spectra, 

and analytical region — at which each compound’s absorbance area is equal to the RSA. For a specified 

system configuration and analytical region, an estimate of the LOD ( ) for a compound can be made 
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from the absorbance area of a reference spectrum in that region ( , ), the reference spectrum CPP 

( , ppm · m), the sample absorption path length ( , m), and the RSA values ( , ) as

. (Equation D1)

A simple trapezoidal approximation of over the analytical region of interest, including baseline 

corrections when necessary, is suitable for the LOD estimate. The RSA is formed from the product of the 

FTIR system’s typical root-mean-square noise level in absorbance and the width of the analytical region 

in (see APPENDIX B, section B2); the RSA estimate should reflect the error incurred in subtracting 

known spectral interferences. APPENDIX E, section E1 presents detailed examples of RSA and LOD 

calculations.

D10. Using Existing Reference Libraries

The analyst may use any of the publicly available reference libraries for quantitative analyses, but it is 

strongly recommended that quantitative results for analytes be based only on spectra from libraries 

that meet the requirements discussed in this appendix. If the reference library consists of spectra 

recorded at a spectral resolution different from that of the field system, the analyst must 1) deresolve 

the spectra mathematically to that of the field data and 2) ensure that the library data are suitably linear 

over the desired measurement range at the spectral resolution of the field system.
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APPENDIX E. EXAMPLE CALCULATIONS

This appendix uses an analysis of TFE and 1,1-difluoroethylene (DFE) in workplace air to illustrate the 

calculations of RSA and LODs. The topics of designing, applying and verifying, and correcting a spectral 

analysis are also addressed below.

The FTIR system configuration assumed for this set of calculations is given below. These parameters 

are consistent with the LOD values presented in Table 2.

System Configuration Parameters Values/Notes

MIL: 0.5 

Absorption path length: 10 m

Apodization function: Triangular

Gas temperature: 293 K

Gas pressure: 101.3 kPa

Zero filling factor: None

Wavenumbers of water bands: 1918 and 2779 

Reference library sources: NIST, EPA

Integration time: 70 s (64 scans)

Detector type: MCT

Detector gain - hardware: Gain jumpers A, D, and H

Detector gain - software: 1.0

E1. RSA, Analysis Design, and LODs

The analysis is to be carried out in workspace air, so the only potential spectral interferences are due to 

water and carbon dioxide. Figure E1 illustrates example reference spectra of TFE, DFE, and water. (The 

water spectrum shown is typical of workspace air samples; carbon dioxide does not absorb in any of the 

spectrum. As a result, the limited spectral region shown in Figure E2 is actually of the greatest interest 

for an analysis of DFE and TFE in workspace air.

Figure E1. Absorbance reference spectra of TFE, DFE, and water.
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Figure E2. Possible analytical regions.

Shown in greater detail in Figure E2 are the three reference spectra and two possible analytical 

regions. Analytical region 1, from 1370 to 1295 , includes water bands that interfere with the 

TFE absorbance in the sample spectra. Analytical Region 2 (1215 to 1100 ) may be the best 

choice for analyzing samples for both TFE and DFE (though the weaker features near 3000 remain 

an option for DFE); it avoids the TFE interference with water, but necessarily includes some spectral 

overlap between the two analytes. Until the relative TFE, DFE, and water concentrations in the samples 

are known, it is unclear which of the two regions will provide better concentration estimates for TFE, so 

calculations for both regions are described below.

Estimation of the LODs for TFE and DFE requires a calculation of the RSA in the actual sample spectra 

over the selected analytical regions. The RSA represents an estimate, based on the assumed sample 

matrix and available reference spectra, of the residual absorbance (see Equation C6) that will be used to 

calculate the concentration uncertainties from the actual sample spectra.

The reader should note that the RSA and derived LOD are only estimates. If the actual sample 

matrix differs substantially from that of the sample matrix assumed in the performance of these 

estimates, the actual concentration uncertainties and LODs may differ substantially from the RSA and 

LOD values obtained in the calculations described below. For workplace air samples, water is the only 

interfering compound that is certain to appear in any of the analytical regions considered below. A 

realistic estimate of the RSA is therefore available by forming the mathematical difference of two water 

spectra — of substantially different water concentrations — recorded using the same FTIR system 

configuration. For each analyte, this estimate assumes that only water and that analyte will absorb 

substantially in the actual sample spectra. Later analyses based on the actual sample matrix could lead 

to either lower or higher concentration, uncertainty, and LOD estimates; examples of such analyses are 

detailed below.

Figures E3 and E4 illustrate two absorbance spectra recorded using the system configuration 

described above. They represent spectra of workspace air at approximately 20% and 40% relative 

humidity. The upper trace in each figure is slightly offset for clarity.
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Figure E3. Water spectra 1215 to 1080 .

Figure E4. Water spectra 1370 to 1295 .

Figure E5 shows two residual spectra formed by subtracting scaled versions of the lower-absorbance 

spectrum from the higher-absorbance spectrum. The scaling factors were determined in independent 

LSF analyses over the two analytical regions. Linear baseline corrections (offset and slope) were 

also determined during the analyses, so the mean value of each residual spectrum is zero. (These 

calculations were performed by using the ANOVA data analysis tool in Microsoft Excel 97). The RSA 

values for the residual spectra are defined in Equation C2 and the related calculations are presented in 

Table E1. (The calculations also were performed in the program Excel by applying the function STDEV to 

the regression residuals.)
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Figure E5. Water residual spectra.

Table E1. Results of RSA analysis from water reference spectra

Analytical 

Region 

( )

Calculated 

Scaling 

Factor

Scaling 

Factor 

Uncertainty 

(1σ)

Residual RMS 

Standard Deviation 

(Absorbance)

RSA; 

Equation B2

( )

1370 to 1295 1.636 0.003 0.00575 0.431

1215 to 1080 1.870 0.008 0.00069 0.093

An estimate of the LODs ( , ppm) for the two compounds — for the specified system configuration 

— can be made from the absorbance areas of the reference spectra over these analytical ranges ( , 

), the reference spectra CPPs ( , ppm · m), the sample absorption path length (m), and the RSA 

values ( , ) derived above.

The absorbance areas were determined using a standard trapezoidal approximation without baseline 

correction. (For these spectra, baseline corrections lead to values that differ from those quoted by less 

than 3% and have been neglected below.) Figure E6 shows the spectral features of TFE and DFE used 

to calculate the for the reference spectra tfeav05.spc (CPP = 256.7 ppm · m) and dfeav05.spc (CPP = 

197.8 ppm · m).

The estimated LODs are the concentrations — for the given system configuration, reference 

spectra, and analytical region — at which each compound’s absorbance area is equal to the RSA. 

Mathematically, the LOD estimates are given by

. (Equation E1)

Table E2 lists the pertinent quantities and resulting LODs for DFE and TFE in the two analytical 

regions considered. The results indicate that the analytical region from 1215 to 1080 is likely to 

provide the most reliable TFE concentrations.
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Figure E6. TFE and DFE spectra for absorbance area calculations.

Table E2. LODs for TFE and DFE from RSA analysis of water reference spectra

Analytical 

Region 

( ) Compound

Reference Spectra System Configuration

File Name

CPP 

(ppm · m)

Absorbance 

Area 

( )

RSA 

( )

L 

(m)

Estimated 

LOD 

(ppm)

1370 to 1295 TFE tfe05.spc 256.7 16.03 0.431 10.0 0.69

1215 to 1080 TFE tfe05.spc 256.7 13.97 0.093 10.0 0.17

1215 to 1080 DFE dfe05.spc 197.8 8.72 0.093 10.0 0.21

E2. LSF Determinations of TFE and DFE Concentrations

Presented in this section is a description of an LSF analysis for the two compounds TFE and DFE in 

ambient air samples. As is always the case before actual measurements are taken, no actual sample data 

for mixtures of the compounds in the assumed matrix (ambient air) were available as these analyses 

were performed. Accordingly, the following description is based on synthetic spectra generated from 

available reference spectra for TFE, DFE, and water.

In order to not overestimate the effectiveness of the FTIR technique, the noise levels in the synthetic 

sample spectra were artificially raised and the reference spectra used are not those most likely to 

provide an optimal analysis. This is the most prudent course available to an analyst attempting to 

predict the performance of the FTIR method under essentially unknown conditions and it is the course 

recommended by the authors to analysts who find themselves in this common situation.

Estimates of the LOD for this sample matrix (see section E1) indicate that the analytical region from 

1215 to 1080 will likely provide the most reliable TFE and DFE concentrations. The system 

configuration parameters used in those estimates are assumed to hold in the following description.
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E2A. Generation and Analysis of Synthetic Sample Spectra of TFE and DFE

Table E3 describes the mathematical generation of synthetic samples for the following LSF analysis 

description. Each sample spectrum consists of the sum of scaled reference spectra for TFE (tfe4.spc, CPP 

= 208.3 ppm · m), DFE (dfeav05.spc, CPP = 197.8 ppm · m) and water (wat02bl.spc). The scaling factors 

indicated in Table E3 were used to generate synthetic sample spectra at the listed concentrations and 

the assumed absorption path length of 10.0 m.

Table E3. Parameters used to generate synthetic sample spectra 

containing TFE, DFE, and water

TFE 

Scaling 

Factor

TFE 

Synthetic 

Concentration 

(ppm)

DFE 

Scaling 

Factor

DFE 

Synthetic 

Concentration 

(ppm)

Water 

Scaling 

Factor

Synthetic 

Sample 

Spectrum 

File Name

0 0 0 0 0.6 S001.spc

0 0 0 0 1.2 S002.spc

1.200 25.0 1.264 25.0 0.6 S551.spc

1.200 25.0 0.253 5.00 0.6 S511.spc

0.240 5.00 1.264 25.0 0.6 S151.spc

1.200 25.0 1.264 25.0 1.2 S552.spc

1.200 25.0 0.253 5.00 1.2 S512.spc

0.240 5.00 1.264 25.0 1.2 S152.spc

A spectrum representing random absorbance noise was added to each synthetic spectrum. The 

noise spectrum was generated using the function RANDOM within the program GRAMS/32 V4.11 

(Galactic, Inc.; see the Array Basic User’s Guide, V4.1, page 316); similar functions are also available in 

other spreadsheet software programs. For the noise spectrum R, with discrete absorbance values 

for wavenumbers i = p to q, the root mean square (RMS) absorbance noise (see also Equation B2, 

which defines the RSA), calculated over the appropriate analytical region(s) according to

, (Equation E2)

was 0.00034 in the region 1438 to 1282 ; this is very close to the value obtained from actual 

absorbance spectra (provided by MIDAC Corporation) recorded with the quoted system configuration 

in the region 1005 to 932 . The synthetic spectra represent four different mixtures of TFE and 

DFE (including one “mixture” that contains neither compound), each at two different moisture levels.

Table E4 lists the results of a LSF analysis (program 4FTIMD V15, Rho Squared; see reference 14) for 

TFE, DFE, and water in the analytical region 1215 to 1080 using the reference spectral files 

TFE3.spc (CPP = 133.2 ppm · m), dfeav04.spc (CPP = 133.3 ppm · m), and wat01bl.spc. Note that these 

are not the same reference spectra that were used in the generation of the synthetic sample spectra 

and that the results for water are in arbitrary units. The analysis included the determination of two 

baseline correction parameters along with the compound concentrations.
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Table E4. Original LSF results for original synthetic sample spectra containing TFE, DFE, and water

Synthetic 

Sample 

Spectrum 

File Name

TFE*

Synthetic 

Concentration 

(ppm)

LSF 

Result 

(ppm)

LSF 3σ

Uncertainty 

(%)

DFE†

Synthetic 

Concentration 

(ppm)

LSF 

Result 

(ppm)

LSF 3σ

Uncertainty 

(%)

Water‡

LSF Result 

(arbitrary)

LSF 3σ

Uncertainty 

(%)

S001.spc 0 0.02 52.5 0 0.07 13.7 1.12 1.23

S002.spc 0 0.03 32.2 0 0.14 8.95 2.24 0.80

S551.spc 25.0 24.41 0.26 25.0 24.30 0.29 0.87 11.8

S511.spc 25.0 24.43 0.25 5.00 4.88 1.43 0.87 11.5

S151.spc 5.00 4.88 0.41 25.0 24.33 0.09 1.07 3.04

S552.spc 25.0 24.43 0.26 25.0 24.35 0.29 1.99 5.20

S512.spc 25.0 24.45 0.26 5.00 4.95 1.43 1.99 5.11

S152.spc 5.00 4.90 0.43 25.0 24.39 0.10 2.19 1.56

*For TFE, excluding samples S001 and S002, the average difference between the synthetic concentration and the LSF result 

was 2.24%. The average 3σ uncertainty was 0.31%. †

For DFE, excluding samples S001 and S002, the average difference between the synthetic concentration and the LSF result 

was 2.34%. The average 3σ uncertainty was 0.61%. ‡

For water, excluding samples S001 and S002, the average 3σ uncertainty was 6.38%.

The LSF results for the non-zero TFE and DFE spectra are consistently good, being different from the 

synthetic concentrations less than 3% in every case. The concentration uncertainties listed in Table E4

are the statistically determined 3σ values from the LSF analyses. These percent uncertainty parameters 

and the visual appearance of the LSF residual spectra are important indicators of the quality of the least 

squares analysis and are discussed further in the following section.

The TFE and DFE results for the two synthetic spectra that contain no absorption features of TFE and 

DFE (S001 and S002) are of interest and importance. The LSF concentration results are small. (Their 

maximum is 0.14 ppm). They are smaller in each case than the LOD estimates of Table E2 and their 

corresponding 3σ uncertainties from this LSF analysis are smaller still. Unfortunately, no consensus 

exists on the exact mathematical relationship between the LODs calculated as described in this 

document and the 3σ concentration uncertainties. This statement is supported by the contents of 

section A2 in reference 4 — a consensus document — that prescribes three different methods for 

calculating LODs (or, in the terminology of reference 4, “minimum detectable concentrations”). These 

three prescriptions include one similar to the LOD method prescribed in this document and one that is 

based on the concentration uncertainties derived from spectra similar to S001 and S002. In this limited 

example, two of the three prescriptions included in reference 4 provide very different results and they 

indicate that the LOD calculation described in this document provides the most conservative estimate 

— that is, the highest estimate — of the three LOD prescriptions of reference 4.

E2B. Analyses of Synthetic Sample Spectra with an Interfering Compound

Every compound-specific quantitative analytical technique, including FTIR spectrometry, can fail 

to provide accurate results when interfering compounds appear in a sample. However, the results 

of a mathematical FTIR spectral analysis designed for a particular set of compounds provides clues 

to its failure in the presence of interferants. An experienced analyst can often adjust the analysis to 

accommodate the interferants and provide accurate results.

To illustrate this important aspect of FTIR spectrometry, a synthetic interference was introduced to 

the spectra described in Table E3. Scaled versions of a reference spectrum (hfpav06.spc, 256.6 ppm · m) 

of the compound hexafluoropropylene (HFP), representing the compound at two concentrations (5.00 

and 10.0 ppm), were added to the original synthetic spectra. The resulting final synthetic sample spectra 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 41 of 44

are described in Table E5 below. The least squares analysis described above for TFE and DFE only, when 

applied to these spectra containing spectral features of HFP, gives the results shown in Table E6 below.

Table E5. Parameters used to generate synthetic sample spectra 

containing TFE, DFE, water, and HFP

Original Synthetic 

Sample Spectrum 

File Name

HFP 

Scaling 

Factor

HFP Synthetic 

Concentration 

(ppm)

Final Synthetic 

Sample Spectrum 

File Name

S001.spc 0.195 5.00 S0011.spc

S002.spc 0.390 10.0 S0022.spc

S551.spc 0.195 5.00 S5511.spc

S151.spc 0.195 5.00 S1511.spc

S512.spc 0.195 5.00 S5121.spc

S511.spc 0.390 10.0 S5112.spc

S552.spc 0.390 10.0 S5522.spc

S152.spc 0.390 10.0 S1522.spc

Table E6. Original LSF results for final synthetic sample spectra containing TFE, DFE, water, and HFP

Synthetic 

Sample 

Spectrum 

File Name

TFE*

Synthetic 

Concentration 

(ppm)

LSF 

Result 

(ppm)

LSF 3σ

Uncertainty 

(%)

DFE†

Synthetic 

Concentration 

(ppm)

LSF 

Result 

(ppm)

LSF 3σ

Uncertainty 

(%)

Water‡

LSF Result 

(arbitrary)

LSF 3σ

Uncertainty 

(%)

S0011.spc 0 1.00 39.6 0 −1.58 26.5 1.65 0.89

S0022.spc 0 2.01 39.5 0 −3.16 26.4 3.29 1.78

S5511.spc 25.0 25.06 1.51 25.0 22.65 1.89 1.43 43.3

S5112.spc 25.0 25.73 2.90 5.00 1.57 53.8 1.99 61.4

S1511.spc 5.00 5.53 6.64 25.0 22.67 1.84 1.63 37.0

S5522.spc 25.0 25.73 2.90 25.0 21.05 4.01 3.10 39.2

S5121.spc 25.0 25.09 1.52 5.00 3.29 13.1 2.55 24.4

S1522.spc 5.00 6.20 11.87 25.0 21.09 3.96 3.30 36.4

*For TFE, excluding samples S001 and S002, the average 3σ uncertainty was 4.56%. †

For DFE, excluding samples S001 and S002, the average 3σ uncertainty was 13.1%. ‡

For water, excluding samples S001 and S002, the average 3σ uncertainty was 40.3%.

The average 3σ uncertainties shown in Table E4 (those of the original LSF analysis without HFP 

interference) and Table E6 (those of the original LSF analysis with HFP interference) are clearly quite 

different. Averaged over the six synthetic spectra containing TFE and DFE in each case, the (absolute) 

percent concentration uncertainties for TFE, DFE, and water are greater for the spectra with HFP 

interference. For TFE and DFE, the average percent uncertainties of Table E6 are over ten times greater 

than those of Table E4. This exercise shows that:

• For sample spectra containing substantial TFE and DFE concentrations, the HFP interference 

causes large increases in the TFE and DFE relative absolute concentration uncertainties.

• For sample spectra containing low or zero concentrations TFE and DFE, the relative absolute 

uncertainties are NOT a reliable indicator of the HFP interference. However, for such spectra, the 

concentration results themselves are often clearly unreliable; note that the DFE concentration 
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ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 42 of 44

results are negative and large compared to the LOD estimate for this compound. (Note also that 

negative concentration results that are small compared to the LOD are statistically valid and do 

not necessarily indicate a failure of the analysis.)

If the concentration results show anomalies such as those noted above, the residual spectra from 

the analyses will also exhibit anomalous behavior. This is illustrated in Figure E7, which shows the 

residual spectra for four analyses. Compared to the residual spectra resulting from analyses of spectra 

without the HFP interference, those with HFP interference show large absorbance features that cannot 

be modeled by the three reference spectra employed. The analyst needs to identify the interfering 

compound before proceeding. This can often be done by visually comparing the suspect residual 

spectra to reference spectra of a number of likely interferants, as illustrated in Figure E8.

Figure E7. Residual spectra with and without HFP interferance.

Figure E8. Comparisons of residual, sample, and reference spectra.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANIC & INORGANIC GASES by Extractive FTIR Spectrometry: METHOD 3800, Issue 2, dated 13 January 2016 - Page 43 of 44

Figure E8 shows that the positions of three major relative maxima in the residual spectrum (top 

trace) correspond well to only one of the nine candidate reference spectra in the lower traces of the 

figure. (The nine spectra shown are all of light, partially or fully fluorinated hydrocarbons.) An additional 

absorbance band from this single reference spectrum also corresponds to a large peak in both of the 

sample spectra (second and third traces from top of the figure).

A closer examination of this reference spectrum — that of HFP, of course — is illustrated in Figure E9. 

It clearly shows a close relationship between the shape of the residual spectrum and the absorption 

features of HFP. The relationship is not exact because the sample spectrum has had incorrect amounts 

of the compounds TFE and DFE subtracted and this distorts the residual spectrum.

Figure E9. Original LSF residual and HFP reference spectra.

Though this identification is tentative, it can be tested by including HFP in the LSF analysis. Table E7

shows the TFE, DFE, and water concentrations resulting from an analysis identical to those described 

above except for the addition of HFP as a fourth compound. (As usual, an HFP reference spectrum 

different from the one used to add the synthetic interference was employed in the analysis.) To facilitate 

comparisons with the preceding tables, the HFP results have been excluded from Table E7; they were in 

all cases similar in quality to those obtained for the other compounds.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

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Table E7. Final LSF results for final synthetic sample spectra containing TFE, DFE, water, and HFP

Synthetic 

Sample 

Spectrum 

File Name

TFE*

Synthetic 

Concentration 

(ppm)

LSF 

Result 

(ppm)

LSF 3σ

Uncertainty 

(%)

DFE†

Synthetic 

Concentration 

(ppm)

LSF 

Result 

(ppm)

LSF 3σ

Uncertainty 

(%)

Water‡

LSF Result 

(arbitrary)

LSF 3σ

Uncertainty 

(%)

S0011.spc 0 1.00 39.6 0 −1.58 26.5 1.65 0.89

S0022.spc 0 2.01 39.5 0 −3.16 26.4 3.29 1.78

S5511.spc 25.0 24.43 0.27 25.0 24.27 0.33 0.87 12.1

S5112.spc 25.0 24.45 0.27 5.00 4.81 1.72 0.88 12.34

S1511.spc 5.00 4.91 0.44 25.0 24.27 0.11 1.08 3.23

S5522.spc 25.0 24.46 0.28 25.0 24.27 0.35 1.99 5.51

S5121.spc 25.0 24.45 0.27 5.00 4.92 1.63 1.99 5.25

S1522.spc 5.00 4.94 0.57 25.0 24.29 0.14 2.20 2.05

*For TFE, excluding samples S0011 and S0022, the average 3σ uncertainty was 0.35%. †

For DFE, excluding samples S0011 and S0022, the average 3σ uncertainty was 0.71%. ‡

For water, excluding samples S0011 and S0022, the average 3σ uncertainty was 6.74%.

The results show that inclusion of HFP in the analysis yields results of the original (high) quality 

shown in Table E3 for TFE, DFE, and water. The percent relative uncertainties are slightly higher in the 

final analysis than in the original analysis; this is to be expected because the same amount of spectral 

information is being used to determine an additional parameter in the final analysis. The quality of the 

analysis could probably be further improved by extending the analytical region to include the entire 

HFP absorbance band shown in Figure E9.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

Volatile Organic Compounds, C1 to C10, Canister Method 3900

Formula: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 3900, Issue 1 EVALUATION: FULL Issue 1: 30 August 2018

OSHA: Table 3

NIOSH REL: Table 3 

Other OELs: [1-3]

PROPERTIES: Table 1

ANALYTES: ethanol, 2-propanol, acetone, 2,3-butanedione, 2,3-pentanedione, 2,3-hexanedione, dichloromethane, 

trichloromethane, hexane, benzene, toluene, ethylbenzene, o-xylene, m,p-xylene, methyl methacrylate, α-pinene, d-limonene

SAMPLING

SAMPLER: FUSED-SILICA LINED STAINLESS STEEL 

CANISTER, 6 L, 450, or 400 mL

FLOW RATE: 0.06 to 50 mL/min. See SAMPLING section

VOL-MIN: Depends on canister volume. See 

SAMPLING section

SHIPMENT: Routine

SAMPLE

STABILITY: 58 days @ 25 °C (30 days for ethanol, 

2-propanol, and acetone @ 10 ppb; 21 days 

for α-pinene and d-limonene @ 0.8 ppm)

BLANKS: 1 field blank per set

ACCURACY

RANGE 

STUDIED: Tables 4a and 4b 

BIAS: Tables 4a and 4b

OVERALL

PRECISION (SS�rrrr): Tables 4a and 4b 

ACCURACY: Tables 4a and 4b 

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, MASS 

SPECTROMETRY

ANALYTE: Compounds listed in Table 1

PRECONCENTRATION 

VOLUME: 25 to 500 mL injection (ppblevels) for a 6L canister, 25 to 250 mL 

injection (ppb-levels) for a 400/450 mL 

canister, 1 mL loop (ppm-levels)

PRECONCENTRATION 

CONDITIONS: Module 1 (empty): 

Focused @ -20° C, desorbed @ 10° C, baked 

@ 150° C (7 min); Module 2 (glass beads): 

Focused @ -80° C, desorbed @ 180° C, 

baked @ 190° C; Module 3 (focuser): 

Focused @ -150° C.

GC/MS 

CONDITIONS: Injection: preconcentrator transfer line 100°C

Detector: 280° C 

Column: 35° C (2 min hold) to 170° C (ramp 

8°C/min), ramp 20° C/min to 220° C (3 min 

hold)

MS Source: 230° C 

Quadrupole: 150° C 

Solvent delay: 4.5 min

MS Scan: 35-350 amu

CARRIER GAS: Helium, 1 mL/min

COLUMN: Capillary, fused silica, 60 m x 0.32-mm ID; 1-

μm film 100% dimethylpolysiloxane

CALIBRATION: Gas phase analytes in canisters

RANGE: Tables 4a and 4b

ESTIMATED LOD: Tables 4a and 4b

PRECISION (SS�rr): Tables 4a and 4b

APPLICABILITY: The method was developed for measuring a range of volatile organic compounds in healthcare settings [4], but 

may be used in other occupational settings. The method was developed to measure the following analytes: ethanol, acetone, 2-

propanol, dichloromethane, hexane, trichloromethane, 2,3-butanedione, 2,3-pentanedione, and 2,3-hexanedione, benzene, 

methyl methacrylate, toluene, ethylbenzene, m,p-xylene, o-xylene, α-pinene, and d-limonene. The working range is 0.24 to 22 ppb
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 2 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

for ppb-level analysis and 0.07 to 2.0 ppm for ppm-level analysis depending on analyte (Table 4a). This method may be applicable 

for other volatile organic compounds that exhibit acceptable stability, bias, precision and accuracy. Alternate detectors (e.g., flame 

ionization detector, electron capture detector, thermal conductivity detector, time-of-flight or quadrupole time-of-flight mass 

spectrometer), canister volumes, GC columns, internal standards and equipment may be used after method performance has been 

assessed and shown to be within acceptable limits. Training on pressure dilution techniques and preconcentration systems as well 

as proficiency testing should be periodically conducted to ensure operating laboratories maintain optimal canister method 

performance.

INTERFERENCES: Other compounds with similar retention times and quantitation ions may interfere (e.g., alcohols, ketones, 

aromatics, beta-pinene). For example, vinyl acetate has the same molecular weight as 2,3-butanedione and has the same retention 

time elution on the recommended column. High humidity samples that are pressurized may form condensed water causing losses 

of water-soluble compounds. Water content should be managed during preconcentration of the sample by purging traps with 

helium, reducing the sample injection volume onto the preconcentrator, or splitting the sample stream prior to concentration 

and/or gas chromatograph injection. 

OTHER METHODS: This is a method for the determination of VOCs, C1 to C10, in mixed exposure environments. The method has 

been modified from the Environmental Protection Agency (EPA) method TO-15 to include specific chemicals and low-ppm 

concentration volatile analysis [5]. ASTM D5466 is also available as guidance for canister sampling and analysis [6].

REAGENTS:

1. Ethanol, reagent grade*

2. 2-Propanol, reagent grade*

3. Acetone, reagent grade*

4. 2,3-Butanedione, reagent grade*

5. 2,3-Pentanedione, reagent grade*

6. 2,3-Hexanedione, reagent grade*

7. Dichloromethane, reagent grade*

8. Trichloromethane, reagent grade*

9. Hexane, reagent grade*

10. Benzene, reagent grade*

11. Toluene, reagent grade*

12. Ethylbenzene, reagent grade*

13. o-Xylene, reagent grade*

14. m-Xylene, reagent grade*

15. p-Xylene, reagent grade*

16. Methyl methacrylate, reagent grade*

17. α-Pinene, reagent grade

18. d-Limonene, reagent grade*

19. Ultra-high purity air

20. Ultra-high purity nitrogen

21. Ultra-high purity helium

22. Internal standard cylinder @ 20 ppb 

(bromochloromethane; 1,4-difluorobenzene; 

chlorobenzene-d5; and 

bromofluorobenzene)

23. Distilled, deionized water, 18 MOhm, low VOC 

content

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Gas chromatograph/mass spectrometer 

system capable of attaching to 

preconcentrator system 

2. Whole-air preconcentrator system capable of 

variable injection volumes from 25 to 500 mL

3. Canister autosampler with 1-mL loop 

injection system

4. Gas dilution system capable of measuring 

pressure transfers

5. Heated canister cleaning system capable of 

evacuating canister to 0.0066 kPa prior to 

sampling and cycling between vacuum and 

humidified gas 

6. Capillary or diaphragm flow controller for 

restricted flow sampling, or fitting for 

instantaneous sampling with 2-μm, sintered, 

stainless steel particle filter 

7. 400 mL, 450 mL, or 6 L fused-silica lined 

coated canisters

8. Gas-tight syringes, 2.5 mL, 1 mL, 250 μL, 100 

μL, 25 μl and 10 μL

9. Disposable glass pipets

10. 2 L glass bulb

 

SPECIAL PRECAUTIONS: Dichloromethane and trichloromethane are confirmed animal carcinogens and 

benzene is a confirmed human carcinogen (see International Agency for Research on Cancer (IARC)). 

Terpenes are flammable and considered irritants. Perform all standards preparation in a well-ventilated 
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 3 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

hood. Wear appropriate personal protective equipment during sampling activities and analysis. It is 

essential that suitable gloves, eye protection, laboratory coat, etc. be used when working with the 

chemicals.

SAMPLING:

1. Evacuate canister to 0.0066 kPa prior to sampling. 

2. Sampling may be performed with a flow controller (e.g., diaphragm or capillary) into a large canister 

(e.g., 6 L) for area sampling or a small canister (e.g., 400 or 450 mL) for personal sampling. Selection of 

the appropriate flow controller and flow rate will depend upon the size of canister being used and 

sampling period that is necessary to characterize exposures.

3. For diaphragm flow controllers, follow the manufacturers procedures for calibration to an appropriate 

flow rate (e.g., 10 mL/min for a 6 L canister for an 8-hour work shift) prior to each day of sampling; the 

final pressure in the canister should be less than 84.1 kPa. 

4. For capillary flow controllers, the flow is governed by the length and internal diameter of the capillary. 

Use a flow controller that will fill the can volume to 30.4 kPa during your sampling period. Do not 

exceed a final sample pressure of 50.6 kPa. 

5. Use a 2-μm, sintered, stainless steel particle filter to protect the flow controllers and canisters when 

sampling in dusty atmospheres. 

SAMPLE PREPARATION:

6. For ppb-level analysis, attach the canister to the preconcentrator via an autosampler or direct transfer 

line. If sample needs to be pressurized to increase sample volume prior to analysis, perform pressure 

dilution with ultra-high purity (UHP) nitrogen or air and apply dilution factors in calculation. Final 

sample pressure after analysis should be greater than 50.6 kPa to limit potential for sample dilution due 

to system leakage. 

7. For ppm-level analysis, pressurize the canister (e.g., 152 kPa or follow instrument manufacturer’s 

recommendation) in order to obtain a sufficient excess pressure to flush the 1 mL loop. An alternative to 

using the loop for quantitation could be running a loop or low volume injection of the canister to 

investigate the analyte concentration range. Then, perform pressure dilutions into ppb-level calibration 

curve.

NOTE: Allow samples or calibration standards that are pressurized a minimum of 6 hours to equilibrate 

prior to analysis. Pressure dilutions greater than 50x should be performed with successive 

dilutions into additional canisters with equilibration time allowed at each step. 

CALIBRATION AND QUALITY CONTROL:

8. Tune the mass spectrometer and calibrate with standards over the range 1 to 20 ppb (25 to 500 mL 

nominal injection volume) and 0.2 to 2 ppm (1 mL loop flushed with 100 mL of sample). Five to seven 

calibration points within the dynamic range should be used. Internal standard (50 mL at 20 ppb) is 

added to the preconcentrator prior to standard or sample transfer. 

9. A mixture of the analytes must be produced before gas-phase standard preparation. 

10. Gas-phase standards may be prepared in 2 L glass bulbs by adding a known volume of a neat chemical 

mixture (see Calculation section) to the vessels. The mass injected into the bulb is determined from the 

concentration of analyte in the mixture and the volume of mixture delivered to the vessel. 

NOTE: Alcohols and ketones may react in the presence of heat. The heated-bulb preparation method is 

not recommended for alcohols and ketones when prepared in the same heated bulb as the 

reaction will change the gas concentration produced. Two alternatives to the heated-bulb 
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 4 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

preparation method are generation without heat in a large inert bag (e.g., tedlar) protected from 

light or purchasing a calibration cylinder from a reference material supplier. 

11. The vessel is heated to 60° C for 30 minutes and allowed to cool to room temperature before a small gas 

volume is transferred to evacuated-calibration canisters via a gas-tight syringe. An alternative to

standard preparation by a heated glass bulb is to purchase gas standards at known concentrations and 

use a gas dilution system to measure pressure transfers. Gas standards should be purchased from a 

reputable, accredited reference material producer (e.g., ISO 17034 [7] accreditation recommended). A 

gas standard at approximately 2 ppm of each of the 17 compounds (balance nitrogen) has been stable 

for 2 years in a pressurized aluminum cylinder with only a ± 2% deviation from the original certified 

value.

12. Add 80 μl of clean distilled water to an evacuated 6 L canister, add gas standard, and dilute with UHP

nitrogen or air to 172 kPa (i.e., ~40%RH). 

13. Develop response factors for each analyte to the closest internal standard using the mass spectrometer 

quantitation software and the appropriate quantitation (primary) ion. Response factors (RF) are 

calculated as follows [8]: 

RRRR = AAxx ∗ CCiiii

AAiiii ∗ CCxx

Where:

Ax = area of the quantitation ion for the analyte (counts);

Ais = area of the quantitation ion for the internal standard (counts);

Cis = concentration of internal standard (ppb); and

 Cx = concentration of the analyte (ppb). 

14. A humidified (e.g., ~40%RH) laboratory blank and humidified (e.g., ~40%RH) 10 ppb check standard 

must be analyzed within a 24 hour period of sample analysis. Perform a full calibration when the check 

standard falls outside established acceptance criteria as developed by the laboratory. Check standard 

recoveries of 90% to 110% are achievable but may be higher or lower depending on instrumentation. A 

replicate sample analysis should be conducted every 10 samples, when possible, to assess 

measurement repeatability. A check standard should be analyzed near the end of the analytical 

sequence to ensure recoveries of each compound are within 80% to 120%. 

MEASUREMENT:

15. Set preconcentrator/GC/MS according to manufacturer’s recommendations and to conditions given on 

page 3900-1. 

16. Inject 50 mL of internal standard at 20 ppb.

17. Inject between 200 and 500 mL of sample for ppb-level analysis and 1 mL of sample for ppm-level 

analysis. Alternative injection volumes may be used as long as performance is verified prior to sample 

analysis.

NOTE 1: If peak area is above linear range of the working standards, dilute with UHP nitrogen using a 

gas dilution system or inject smaller sample volumes, reanalyze and apply the appropriate 

dilution factor in calculations. 

NOTE 2: m- and p-Xylene isomers co-elute on the specified column and should be analyzed/reported 

together. 

CALCULATIONS:

18. Calculation for the concentration of analyte in calibration standard (Cbulb; ppm) in the glass bulb:

CCbbbbbbbb = CCaaVVmmYY ∗ 22.4

MMWWii ∗ VVbbbbbbbb

∗ TTbbaabb

273.15 ∗ 760

PPbbaabb
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 5 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Where: 

 Ca = concentration of analyte in the mixture (mg/mL); 

Vm = volume of mixture injected in bulb (mL); 

Y = purity of analyte (scale 0 to 1); 

MWs = molecular weight of analyte (g/mol); 

Vbulb = volume of glass bulb (m3

) determined experimentally; 

Tlab = temperature of the laboratory (K); 

Plab = pressure of the lab (mm Hg). 

NOTE: 760 is assuming standard atmospheric pressure. 273.15 is International Union of Pure and 

Applied Chemistry (IUPAC) standard temperature and 22.4 is molar volume at STP.

19. Transfer a small aliquot of the gas from the bulb to an evacuated canister for calibration standards. 

20. Calculate the final concentration in the canister (Cfinal; ppb) based on:

CCffiiffaabb = CCbbbbbbbb ∗ 1000 ∗ VViiiiii

VVccccii

∗ PPccaaaa

PPff

 

Where: 

Cbulb = concentration in glass bulb (ppm); 

Vinj = volume transferred to the canister (mL); 

Vcan = volume of the canister (mL); 

Patm=atmospheric pressure (kPa); and

Pf = final canister pressure (kPa).

21. Determine the analyte concentration (Cx) in ppb or ppm according to the following equation:

CCxx = AAxx ∗ CCiiii

AAiiii ∗ RRRR

Where: 

Ax = area of the primary ion for the analyte (counts); 

Ais = area of the primary ion for the internal standard (counts); 

Cis = concentration of internal standard (ppb); 

RF = response factor calculated using linear regression. 

Note: Most data analysis software packages will calculate analyte concentration provided you 

enter the calibration information in the quantification database. Blank correction may be 

necessary when instrument or field blank results are consistent and above the analyte detection 

limit.

22. Dilution factors (ratio of final volume to initial volume) must be applied to standards or samples as 

applicable. 

EVALUATION OF METHOD:

Method Accuracy

Bias, precision, accuracy and stability for 14 volatile organic compounds (ethanol, acetone, 2-propanol, 

dichloromethane, hexane, trichloromethane, benzene, methyl methacrylate, toluene, ethylbenzene, m,pxylene, o-xylene, α-pinene, and d-limonene) at ppb-level and ppm-level concentrations [4] were 

determined using Kennedy et al. [9]. Three additional analytes (2,3-butanedione, 2,3-pentanedione, and 
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 6 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

2,3-hexanedione) were added to the target list after this study and accuracy has been assessed during 

chamber studies (see section below). The upper and lower 95% confidence limits of the average bias were 

calculated (i.e., ±1.96*standard error). To be acceptable either both bias limits must have an absolute value 

less than 10%, or an absolute value of 10% must fall between the limits. For a method to meet the NIOSH 

“25% accuracy” criterion, the 95% confidence statistic for accuracy estimated with the hyperbolic 

approximation formula must be below 25%. The stability metric is a measure of bias comparing average 

concentration on day 30 or greater to average concentration on day zero; the absolute value of the 

stability metric must be less than 10% to be acceptable. Day 30 stability was obtained from LeBouf et al. 

[4]. Day 58 stability was obtained from NIOSH [10]. While method performance varied by analyte, the 

validation results demonstrate that this method is a viable air sampling and analytical methodology for 

measuring a wide range of air concentrations of select volatile organic compounds in mixed exposure 

environments. 

For the preconcentration method used to measure ppb-level concentrations, all accuracy criteria were 

within acceptable limits for 16 of the 17 target analytes (Table 4a). 2-Propanol failed the 95% accuracy 

criterion at ppb-level concentrations and accuracy validation results are considered inconclusive, perhaps 

because the water management step in the preconcentration may have affected the concentrations of this 

analyte due to its polar nature. Ethanol, however, passed the accuracy criterion despite its similar polarity. 

Bias was acceptable for all 17 analytes. The average bias values for 2-propanol and methyl methacrylate 

were greater than 10% but still within acceptable parameters when 95% confidence limits were calculated; 

further evaluation to confirm or refute this excessive bias result is warranted, particularly because bias 

values may have been influenced by error introduced during the preparation of standards (e.g., liquid 

injection into glass bulb or concentration dilution). Storage stability of the 17 analytes was confirmed for 

58 days at approximately 10 ppb for most analytes with the exception of ethanol, 2-propanol, and acetone 

(stable for 30 days) (Table 4a) [4, 10]. 

Because trichloromethane failed Bartlett’s test of homogeneity [9] (i.e., precision varied across 

concentrations) for ppb-level, the accuracy value for this analyte displayed in Table 4a was not calculated 

based on a value for precision that was pooled across all concentrations investigated. Rather, the accuracy 

value was calculated based on the worst-case concentration-specific precision and bias and a modified 

accuracy calculation that accounts for the reduced degrees of freedom. Further investigation of the 

precision for trichloromethane using the ppb-level method is warranted. In the meantime, the method can 

be used to quantify this analyte, but the analyst must be cognizant of the concentration-dependence of 

measurement variability.

For the loop method used to measure ppm-level concentrations, all accuracy and bias criterion were 

within acceptable limits for each analyte (Table 4b). Bartlett’s test demonstrated that measurement 

precision was non-homogeneous over concentrations for both dichloromethane and methyl methacrylate. 

The accuracy values for these two analytes were calculated based on the worst-case concentration-specific 

precision and bias and a modified accuracy calculation that accounts for the reduced degrees of freedom.

All analytes were stable for 30 days, except α-pinene and d-limonene, which remained stable for 21 days

(Table 4b). The shorter acceptable storage time for α-pinene and d-limonene may be due to losses from 

chemical reactions with other components of canister contents, particularly oxidizing species such as 

ozone or hydroxyl and nitrate radicals naturally found in indoor air [11, 12].

Inter-laboratory Study

An inter-laboratory study was conducted to estimate precision from nine laboratories [10]. Laboratories 

were recruited from American Industrial Hygiene Association (AIHA)-accredited or National Environmental 

Laboratory Accreditation Program (NELAP)-accredited laboratories which had the experience and 

equipment necessary to analyze canister samples. Precision estimates for repeatability ranged from 0.04 to 

0.55 at ppb concentrations and from 0.1 to 0.47 for ppm concentrations. Precision estimates for 
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 7 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

reproducibility ranged from 0.1 to 0.62 at ppb concentrations and from 0.19 to 0.58 at ppm concentrations, 

depending on analyte and nominal level. Plots of h and k statistics which measure between and within 

laboratory consistency indicated inconsistencies with reported results from laboratories 5, 6, and 7. These 

laboratories were investigated to determine the cause of the inconsistent results. Issues with storage time 

and adherence to the method protocol were encountered but results were retained due to the low 

number of laboratories participating. Training on pressure dilution techniques and preconcentration 

systems as well as proficiency testing should be periodically conducted to ensure operating laboratories 

maintain optimal canister method performance.

Chamber Studies

The canister method was challenged in laboratory chamber studies [10] under varying environmental 

conditions and concentrations of alpha-diketones to assess sampling and analytical accuracy following 

ASTM D6246 [13]. The following conditions were assessed: temperature, humidity, wind speed, and 

concentration. Bias pulse tests were performed to assess the bias associated with a known drop in flow rate 

(~13% over the sampling period) for capillary flow controllers. Flow rate bias (bias pulse) was assessed 

using the difference in measurements for peak exposures occurring at the beginning and end of the 

sampling period. Tests were also performed to assess inter-day and inter-sampler variation. A dynamic 

volatile organic compound generation and sampling system was used to produce known concentrations 

of challenge agents in a glass sampling chamber with 18 sampling ports. The system was placed in a large 

walk-in environmental control chamber to regulate temperature. Capillary flow controllers were 

constructed using Swagelok connections and deactivated fused-silica tubing to collect air samples at 15 

mL/min for 15 minutes into a 450 mL canister to test the maximum recommended sampling fill pressure of 

50.6 kPa. Canisters were pressurized with ultra-high purity nitrogen and analyzed using an 

autosampler/preconcentrator system attached to a gas chromatograph/mass spectrometer. Results were 

corrected for pressure dilution and compared to a theoretical concentration calculated from flow dilution 

of the gas standard used in the generation system. Accuracy was calculated for each target analyte for all 

conditions combined. Variance estimates for each factor influencing accuracy were used to apportion the 

relative influence of each test condition on the overall performance of the method. Upper confidence 

limits on accuracy (A95) were below 0.25 for all analytes: 0.085 for diacetyl, 0.142 for 2,3-pentanedione, and 

0.194 for 2,3-hexanedione. Overall precision was largest for 2,3-hexanedione at 0.061 with 27.4% of the 

total variance due to inter-day variability. The peak exposure condition (i.e., effect of a bias pulse given the 

flow rate drop over the sampling period) accounted for less than 6% of the variability regardless of analyte, 

meaning the known drop in flow rate from the capillary flow controller did not overly influence air 

sampling results. Additional VOCs (ethanol, acetone, isopropyl alcohol, dichloromethane, 

trichloromethane, methyl methacrylate, hexane, benzene, toluene, xylenes, α-pinene, d-limonene) were 

present during low and high humidity trials to assess the effect of humidity on method performance. For 

the high humidity trial (n=10), A95 for the additional VOCs were below 0.25 except for ethanol and methyl 

methacrylate. For the low humidity trial (n=5), A95 for the additional VOCs were below 0.25 except for 

ethanol and isopropyl alcohol. Canister method is a reliable, robust sampling and analytical method that 

may be used for a variety of analytes (e.g., C1 to C10) under varying environmental conditions. 

Flow Controllersfor Personal Sampling

Diaphragm flow controllers are heavy due to the stainless steel body holding the diaphragm making them 

inconvenient for personal sampling. The capillary flow controllers are light weight and provide a broad 

dynamic range of flow rates that enables full-shift (8-12 hours) sample collection in a 400/450mL canister 

[8]. With capillary flow controllers, the flow is governed by the length and internal diameter of the capillary 

and the pressure in the canister [8, 14]. As the canister fills with sample, the pressure differential between 

the atmosphere and the inside of the canister decreases. This decrease causes a slight, non-linear 

reduction in flow rate over time. The effect of the change can be mitigated by keeping the final sample 

volume in the canister at around 30% (but no greater than 50%). The effect of a known, decrease in flow 
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 8 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

rate on bias has been assessed and will not overly influence time-integrated measurements of analyte 

concentration even when a peak concentration at the beginning or end of the sampling period is collected 

[10, 14]. 

Additional Guidance

Follow EPA guidelines for cleaning canisters, sampling train cleanliness, canister cleanliness, leak testing, 

and canister pressurization [5, 15]. Do not clean fused silica lined canisters with humidified air and heat 

above 80°C. Use humidified nitrogen and heat to clean fused silica lined canisters. ASTM D5466 is also 

available as guidance for canister sampling and analysis [6]. 

REFERENCES:

[1] ACGIH [2018]. TLVs® and BEIs® based on the documentation of the Threshold Limit Values for 

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American 

Conference of Governmental Industrial Hygienists. 

[2] Permanent Senate Commission for the Investigation of Health Hazards of Chemical Compounds in 

the Work Area [2016]. List of MAK and BAT values 2016; Maximum Concentrations and Biological 

Tolerance Values at the Workplace. Weinheim, Germany: Wiley-VCH Verlag GmbH & Co., 

https://dx.doi.org/10.1002/9783527805983. 

[3] Institute for Occupational Safety and Health of the German Social Accident Insurance (IFA) [2017]. 

GESTIS international limit values, http://limitvalue.ifa.dguv.de/. 

[4] LeBouf RF, Stefaniak AB, Virji MA [2012]. Validation of evacuated canisters for sampling volatile 

organic compounds in healthcare settings. J Environ Monit 14(3):977-983.

[5] EPA [1999]. Compendium Method TO-15: Determination of volatile organic compounds (VOCs) in air 

collected in specially-prepared canisters and analyzed by gas chromatography/mass spectrometry 

(GC/MS). Cincinnati, OH, U.S. Environmental Protection Agency, EPA/625/R-96/010b.

[6] ASTM [2015]. Standard test method for determination of volatile organic compounds in atmospheres 

(canister sampling methodology). West Conshohocken, PA: ASTM International, ASTM D5466 – 15.

[7] ISO [2016]. General requirements for the competence of reference material producers. Geneva, 

Switzerland: International Organization for Standardization, ISO 17034. 

[8] Rossner A, Farant JP [2002]. Development of a flow controller for long-term sampling of gases and 

vapors using evacuated canisters. Environ Sci Technol 36:4912-4920.

[9] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. By

Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 95-117.

[10] NIOSH [2018]. NIOSH 3900 Backup Data Report. By LeBouf RF, Feng A, Shulman SA. Cincinnati, OH: 

U.S. Department of Health and Human Services, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, https://www.cdc.gov/niosh/nmam/datadoc.html. 

[11] Atkinson R, Arey J [2003]. Gas-phase tropospheric chemistry of biogenic volatile organic compounds: 

a review. Atmos Environ 37:S197-S219.

[12] Forester C, Wells J [2009]. Yields of carbonyl products from gas-phase reactions of fragrance 

compounds with OH radical and ozone. Environ Sci Technol 43:3561-3568.
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 9 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[13] ASTM [2013]. Standard practice for the evaluating the performance of diffusive samplers. West 

Conshohocken, PA: ASTM International, ASTM D6246 – 08.

[14] Rossner A, Wick DP [2005]. A field study to assess the long-term sampling feasibility of evacuated 

canisters and the development of a mathematical model to analyze potential sampling bias. J Occ 

Environ Hyg 2:474-480.

[15] EPA [1999]. Compendium Method TO-14A: Determination of volatile organic compounds (VOCs) in 

ambient air using specially prepared canisters with subsequent analysis by gas chromatography. 

Cincinnati, OH, US Environmental Protection Agency, EPA/625/R-96/010b.

[16] Code of Federal Regulations [2017]. Title 29 (Labor), Part 1910 (Occupational Safety and Health 

Standards); 29 CFR 1910.1000 Table Z-1; US Government Printing Office, Washington DC.

[17] NIOSH [2010]. Pocket guide to chemical hazards. Cincinnati, OH: U.S. Department of Health and 

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 2010-168c.

METHOD WRITTEN BY:

Ryan F LeBouf, PhD, CIH. CDC/NIOSH, Respiratory Health Division.

Disclaimer: Mention of any company or product does not constitute endorsement by the National 

Institute for Occupational Safety and Health (NIOSH), Centers for Disease Control and Prevention (CDC). In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 10 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 1. SYNONYMS, FORMULAE, MOLECULAR WEIGHTS AND PHYSICAL PROPERTIES

IUPAC Name / 

Synonym CAS # / RTECS 

Empirical 

formula

Molecular 

Weight

Boiling 

Point 

(°C)

Vapor 

Pressure 

@ 20° C 

(mm Hg)

Vapor 

Pressure 

@ 20° C 

(kPa)

ethanol / ethyl 

alcohol

64-17-5/ 

KQ6300000

C2H6O 46.07 78.3 44 5.87

2-propanol / 

isopropyl alcohol

67-63-0/ 

NT8050000

C3H8O 60.1 82.3 33 4.4

acetone / 2-

propanone

67-64-1/ 

AL3150000

C3H6O 58.08 56.1 180 24

2,3-butanedione / 

diacetyl

431-03-8/ 

EK2625000

C4H6O2 86.09 88 52.2 6.96

2,3-pentanedione

/ acetyl propionyl

600-14-6/ 

SA1850000

C5H8O2 100.12 110-

112

21.4 2.85

2,3-hexanedione / 

acetyl butyryl

3848-24-6 C6H10O2 114.14 128 10 1.33

dichloromethane / 

methylene 

chloride

75-09-2/ 

PA8050000

CH2Cl2 84.93 40 350 46.7

trichloromethane / 

chloroform

67-66-3 CHCl3 119.38 61.2 160 21.3

hexane / n-hexane 110-54-3/ 

MN9275000

C6H14 86.18 68.7 124 16.5

benzene / benzol 71-43-2/ 

CY1400000

C6H6 78.11 80.1 75 10

toluene / 

methylbenzene

108-88-3/ 

XS5250000

C7H8 92.14 110.6 22 2.93

ethylbenzene / 

phenylethane

100-41-4/ 

DA0700000

C8H10 106.17 136.2 7 0.93

o-xylene / 1,2-

dimethylbenzene

95-47-6/ 

ZE2450000

C8H10 106.17 144.5 7 0.93

m-xylene / 1,3-

dimethylbenzene

108-38-3/ 

ZE2275000

C8H10 106.17 139.1 9 1.2

p-xylene / 1,4-

dimethylbenzene

106-42-3/ 

ZE2625000

C8H10 106.17 138.2 9 1.2

methyl-2-methyl2-propenate / 

methyl 

methacrylate 

80-62-6/

OZ5075000

C5H8O2 100.12 100.5 29 3.87

4,6,6-

trimethylbicyclo[3.

1.1]hept-3-ene / αpinene / 2-pinene

80-56-8/ 

DT7000000

C10H16 136.23 156.2 4.75 

(25° C)

0.63

(4R)-1-methyl-4-

prop-1-en-2-

ylcyclohexene / dlimonene / pmentha-1,8-diene, 

("R")

5989-27-5/ 

OS8100000

C10H16 136.23 178 1.43 0.19
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 11 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 2. CHARACTERISTIC MASSES (M/Z) USED FOR QUANTITATION

Substance

Primary 

Ion

Secondary

Ion

ethanol 45 46, 43

2-propanol 45 43

acetone 43 58

2,3-butanedione 86 43

2,3-pentanedione 100 57, 43

2,3-hexanedione 71 43, 114

dichloromethane 84 49, 86

trichloromethane 83 85, 47

hexane 57 41, 43

benzene 78 77, 52

toluene 91 92

ethylbenzene 91 106

o-xylene 91 106, 105

m,p-xylene 91 106, 105

methyl methacrylate 69 41, 100

α-pinene 93 91, 92

d-limonene 68 93, 67

bromochloromethaneA 130 49, 93

1,4-difluorobenzeneA 114 63, 88

chlorobenzene-d5A 117 82, 119 Ainternal standards

TABLE 3. EXPOSURE LIMITS (PPM) [16,17] 

Substance

OSHA 

TWA

OSHA 

Peak

NIOSH 

TWA

NIOSH 

STEL

mg/m3 

per ppm

ethanol 1000 - 1000 - 1.89

2-propanol 400 400 500 2.46

acetone 1000 - 250 - 2.38

2,3-butanedione - - 0.005 0.025 3.52

2,3-pentanedione - - 0.0093 0.031 4.09

2,3-hexanedione - - - - 4.67

dichloromethane 25 125 N/AA (caB

) 3.47

trichloromethane - 50 - 2 (60 min) 4.88

hexane 500 - 50 - 3.53

benzene 1 5 0.1 (ca) 1 (ca) 3.19

toluene 200 500 100 150 3.77

ethylbenzene 100 - 100 125 4.34

o-xylene 100 - 100 150 4.34

m-xylene 100 - 100 150 4.34

p-xylene 100 - 100 150 4.34

methyl methacrylate 100 - 100 - 4.09

α-pinene - - - - 5.57

d-limonene - - - - 5.57 AN/A = not applicable

B

ca = carcinogen
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 12 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 4A. RANGE, OVERALL BIAS, PRECISION AND ACCURACY FOR PPB LEVELS [4, 10] 

Substance

Range of 

generated 

samples

Overall 

Bias (%)

Overall 

Precision 

(SrT) 

Overall 

Accuracy 

(%) 

58 day 

Stability (%)

ethanol 1.3 - 9.9 -9.3 0.058 ±22.2 -0.5 (30 days)

2-propanol 1.3 - 10.0 -12.3 0.098 ±28.9 7.8 (30 days)

acetone 1.3 - 10.0 -3.2 0.064 ±18.6 3.1 (30 days)

2,3-butanedione 21.5 - 381 2.2 0.032 ±8.5 4.1

2,3-pentanedione 20.8 - 377 4.8 0.048 ±14.2 2.7

2,3-hexanedione 20.4 - 368 -6.0 0.061 ±19.4 -2.9

dichloromethane 1.3 - 9.8 2.9 0.017 ±7.1 0.22

trichloromethane 1.3 - 9.8 -0.3 0.020* ±16.5* 3.1

hexane 1.3 - 9.9 -0.1 0.020 ±5.5 3.6

benzene 1.3 - 9.8 -3.0 0.022 ±8.3 3.6

toluene 1.3 - 9.9 -4.0 0.020 ±8.6 1.6

ethylbenzene 1.3 - 9.9 -4.4 0.020 ±9.1 -0.6

o-xylene 1.3 - 9.8 -6.0 0.027 ±12.3 -3.5

m,p-xylene 1.3 - 9.8 -6.7 0.025 ±12.6 -8.1

methyl methacrylate 1.3 - 9.9 -10.2 0.066 ±24.9 1.5

α-pinene 1.3 - 9.8 -4.7 0.028 ±11.4 -3.3

d-limonene 1.3 - 9.8 -4.6 0.069 ±21.1 -1.9

*worst-case precision estimate since failed Bartlett’s test of homogeneity of variance across concentrations

SrT = pooled precision as relative standard deviation across multiple concentration levels

Overall Accuracy=upper confidence limit on accuracy, A95

Stability (%) = ((Concentration at storage day)/(Concentration at day 0)-1)*100%
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 13 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 4B. RANGE, OVERALL BIAS, PRECISION AND ACCURACY FOR PPM-LEVELS [4] 

Substance

Range of 

generated 

samples

Overall 

Bias (%)

Overall 

Precision 

(SrT) 

Overall 

Accuracy 

(%) 

30 day Stability 

(%) 

ethanol 0.81 - 1.8 9.4 0.028 ±17.0 -9.2

2-propanol 0.82 - 1.8 8.9 0.027 ±16.1 -2.9

acetone 0.82 - 1.8 8.7 0.019 ±13.8 -1.6

dichloromethane 0.79 - 1.8 3.0 0.041* ±17.8* -4.7

trichloromethane 0.80 - 1.8 5.7 0.012 ±8.9 -1.9

hexane 0.81 - 1.8 6.1 0.012 ±9.4 0.1

benzene 0.80 - 1.8 6.3 0.011 ±9.3 7.3

toluene 0.81 - 1.8 5.5 0.015 ±9.3 6.4

ethylbenzene 0.81 - 1.8 4.7 0.017 ±9.2 0.8

o-xylene 0.80 - 1.8 5.4 0.019 ±10.3 -0.3

m,p-xylene 0.80 - 1.8 3.9 0.019 ±8.8 -2.1

methyl methacrylate 0.81 - 1.8 5.7 0.016* ±15.6* 1.4

α-pinene 0.81 - 1.8 6.7 0.022 ±12.5 0.1 (21 days)

d-limonene 0.80 - 1.8 13.1 0.039 ±24.0 -0.03 (21 days)

*worst-case precision estimate since failed Bartlett’s test of homogeneity of variance across concentrations

SrT = pooled precision as relative standard deviation across multiple concentration levels

Overall Accuracy=upper confidence limit on accuracy, A95

Stability (%) = ((Concentration at storage day)/(Concentration at day 0)-1)*100%

TABLE 5A. LIMITS OF DETECTION (LOD), MEASUREMENT RANGE, AND PRECISION FOR PPB-LEVELS [4, 10] 

Substance LOD (ppb) Range (ppb) Precision (RMSEA) Linearity R2

ethanol 0.4 0.25 – 21.4 0.272 0.999

2-propanol 0.5 0.27 – 22.0 0.077 0.998

acetone 0.4 0.27 – 20.6 0.080 0.999

2,3-butanedione* 0.3 0.26 – 21.0 0.033 0.999

2,3-pentanedione* 0.2 0.24 – 18.4 0.148 0.999

2,3-hexanedione* 0.4 0.24 – 18.0 0.238 0.999

dichloromethane 0.2 0.26 – 20.8 0.049 0.999

trichloromethane 0.1 0.27 – 21.6 0.044 0.999

hexane 0.2 0.27 – 21.8 0.030 0.999

benzene 0.2 0.27 – 21.6 0.032 0.999

toluene 0.3 0.27 – 21.4 0.023 0.999

ethylbenzene 0.3 0.26 – 21.0 0.019 0.999

o-xylene 0.3 0.26 – 21.0 0.019 0.999

m,p-xylene 0.4 0.27 – 22.0 0.015 0.999

methyl methacrylate 0.5 0.25 – 20.4 0.035 0.999

α-pinene 0.5 0.26 – 21.0 0.019 0.999

d-limonene 0.8 0.26 – 20.8 0.136 0.999

*Selected ion monitoring mode used to improve detection limits ARMSE= root mean square error
Volatile Organic Compounds, C1 to C10, Canister Method: METHOD 3900, Issue 1, dated 30 August 2018 - Page 14 of 14

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 5B. LIMITS OF DETECTION (LOD), MEASUREMENT RANGE, AND PRECISION FOR PPM-LEVELS [4] 

Substance LOD (ppm) Range (ppm) Precision (SEEA) Linearity R2

ethanol 0.2 0.07 - 1.9 0.058 0.998

2-propanol 0.2 0.07 - 2.0 0.066 0.998

acetone 0.3 0.08 - 2.0 0.071 0.998

dichloromethane 0.4 0.15 - 1.9 0.099 0.999

trichloromethane 0.3 0.11 - 1.9 0.093 0.999

hexane 0.2 0.11 - 2.0 0.072 0.999

benzene 0.2 0.11 - 1.9 0.064 0.999

toluene 0.3 0.11 - 1.9 0.101 0.998

ethylbenzene 0.3 0.11 - 1.9 0.108 0.998

o-xylene 0.1 0.11 - 1.3 0.031 0.998

m,p-xylene 0.3 0.11 - 1.9 0.110 0.998

methyl methacrylate 0.2 0.11 - 1.9 0.089 0.998

α-pinene 0.3 0.11 - 1.9 0.116 0.999

d-limonene 0.4 0.11 - 1.9 0.156 0.999 ASEE = standard error of estimate from regression curve
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

HYDROQUINONE 5004

HOC6H4OH MW: 110.11 CAS: 123-31-9 RTECS: MX3500000

METHOD: 5004, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984

Issue 3: 26 February 2016

OSHA: 2 mg/m3

NIOSH: C 2 mg/m3/15 min

PROPERTIES: solid; MP 170 °C; BP 285 °C @ 730 mm; 

VP 0.0024 Pa (1.8 x 10 -5 mm Hg; 0.11 mg/m3

) @ 

25 °C

SYNONYMS: 1,4-benzenediol; hydroquinol; quinol; 1,4-dihydroxybenzene

SAMPLING

SAMPLER: FILTER (0.8-μm cellulose ester membrane)

FLOW RATE: 1 - 4 L/min

VOL-MIN: 30 L @ 2 mg/m3

 -MAX: 180 L

FIELD

TREATMENT: transfer filter immediately to jar with

10 mL 1% acetic acid

SHIPMENT: ship sample solutions

SAMPLE

STABILITY: at least 7 days @ 25 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 0.8 to 4 mg/m3 [1] (90-L samples)

BIAS: 4.4%

OVERALL

PRECISION (SS�rrrr): 0.061

ACCURACY: ± 15.0% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: hydroquinone

EXTRACTION: 1% acetic acid, 10 mL

INJECTION 

 VOLUME: 100 μL

MOBILE 

PHASE: 1% acetic acid in H2O; 1 mL/min

COLUMN: 25 cm x 4.6-mm ID C18/USP L1 column; 

ambient temperature, 400 to 600 psi 

(2800-4100 kPa)

DETECTOR: UV @ 290 nm

CALIBRATION: solutions of hydroquinone in 1% aqueous 

acetic acid

RANGE: 0.06 to 0.8 mg per sample [2]

ESTIMATED LOD: 0.01 mg per sample [3]

PRECISION (SS�rr): 0.030 [1]

APPLICABILITY: The working range is 0.7 to 8 mg/m3 for a 90-L air sample or 2 to 25 mg/m3 for a 30-L air sample. This method can 

be used when significant concentrations of hydroquinone vapor are not present.

INTERFERENCES: None known. Hydroquinone is unstable on the collection media and must be stabilized immediately after 

collection by dissolution in 1% acetic acid.

OTHER METHODS: This is Method S57 [2] in a revised format. The method also appears in a NIOSH recommended standard [4].
HYDROQUINONE: METHOD 5004, Issue 3, dated 26 February 2016 - Page 2 of 3 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Hydroquinone, reagent grade.

2. Distilled water.

3. Acetic acid, glacial.

4. Acetic acid, 1%. Dilute 10 mL acetic acid to 

1 L with distilled water. 

NOTE: This solution is needed at the 

sampling site for field treatment of 

samples. 

5. Calibration stock solution, 3.6 mg/mL. 

Dissolve 0.0900 g hydroquinone in 25 mL 1% 

acetic acid. Prepare fresh daily, in duplicate.

EQUIPMENT:

1. Sampler: 37-mm cassette containing 0.8-μm 

cellulose ester membrane filter and cellulose 

backup pad.

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing.

3. High pressure liquid chromatograph with UV 

detector at 290 nm, integrator and column 

(page 5004-1).

4. Jars, 60-mL, ointment, low form, with PTFE 

film gaskets and screw caps.

5. Syringe, 100-μL, or autosampler for sample 

injection.

6. Microliter syringes for standard preparation.

7. Volumetric flasks, 10- and 25-mL, and 1-L.

SPECIAL PRECAUTIONS: None

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 1 and 4 L/min for a total sample size of 30 to 180 L.

NOTE: This method will collect hydroquinone aerosol only. The equilibrium vapor pressure is 

equivalent to 0.11 mg/m3 at 25 °C and may be a significant factor at elevated temperatures.

3. Immediately after sampling, transfer the filter (do not include backup pad) with tweezers to a 60-mL 

ointment jar. Add 10 mL 1% acetic acid. Process field blanks similarly.

4. Ship the filters in the ointment jars.

SAMPLE PREPARATION:

5. Transfer the sample solution from the ointment jar to a 25-mL volumetric flask.

6. Rinse the ointment jar twice with 5 mL 1% acetic acid. Add the washings to the volumetric flask. Make 

up to volume with 1% acetic acid.

CALIBRATION AND QUALITY CONTROL: 

7. Calibrate daily with at least six working standards over the range 0.01 to 0.8 mg hydroquinone per 

sample.

a. Add known amounts of calibration stock solution to 1% acetic acid in 25-mL volumetric flasks and 

dilute to the mark.

b. Analyze together with samples and blanks (steps 9 and 10).

c. Prepare calibration graph (peak area vs. mg hydroquinone).

d. Prepare recovery graph (recovery vs. mg hydroquinone).

8. Check recovery with at least three spiked media blanks per sample set.

a. Add aliquot of calibration stock solution with a microliter syringe directly to a representative filter. 

Transfer filter to 60-mL ointment jar, add 10 mL 1% acetic acid, and allow to stand overnight.

b. Prepare and analyze together with working standards (steps 5, 6, 9 and 10).

c. Calculate recovery [(mg recovered - mg blank)/mg added)].

MEASUREMENT:

9. Set HPLC to conditions given on page 5004-1. Inject 100-mL sample aliquot.
HYDROQUINONE: METHOD 5004, Issue 3, dated 26 February 2016 - Page 3 of 3 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

10. Measure peak area. Retention time is ca. 5.2 min under these conditions.

CALCULATIONS:

11. Determine the mass, mg (corrected for recovery) of hydroquinone, found in the sample (W) and in the 

average media blank (B).

12. Calculate concentration of hydroquinone, C, in the air volume sampled, V (L):

CC = (WW − BB)xx103

VV , mmmm mm3 ⁄

EVALUATION OF METHOD:

Method S57 [2] was issued on November 26, 1976, and validated over the range 0.8 to 4 mg/m3 at 20 °C 

and 762 mm Hg using 90-L air samples [1,5]. Overall precision, SS�rrrr, was 0.061 with average recovery 105%, 

representing a non-significant bias. The atmospheres were generated by atomization of an aqueous 

solution of hydroquinone into dry air; aerosol concentrations were independently verified by direct UV 

spectrophotometry on filter samples. Average collection efficiency was 100% at 4 mg/m3

. No loss of 

hydroquinone was seen from filters spiked with 720 μg hydroquinone, and then used to sample 180 L 

clean air, indicating the hydroquinone vapor pressure is not significant at these conditions. At elevated 

temperature, however, contribution from vapor may be significant. Storage studies were conducted by 

storing exposed filters in 1% acetic acid at ambient temperature for seven days. No change in 

hydroquinone concentration was seen.

REFERENCES:

[1] NIOSH [1976]. Backup data report No. S57. In: Ten NIOSH analytical methods, set 2. Unpublished. 

Available as Order No. PB 271-464, from NTIS, Springfield, VA. 

[2] NIOSH [1977]. Hydroquinone: Method S57. In: Taylor DG, ed. NIOSH manual of analytical methods. 

2nd ed. (vol 2) Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

157-B.

[3] A.D. Little, Inc. [1983]. User check. NIOSH Sequence #4121Q. Unpublished.

[4] NIOSH [1978]. Criteria for a recommended standard-occupational exposure to hydroquinone. 

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 78-155.

[5] NIOSH [1980]. NIOSH research report-development and validation of methods for sampling and 

analysis of workplace toxic substances. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 80-133.

METHOD REVISED BY:

Jerome Smith, Ph.D., NIOSH; S57 originally validated under NIOSH Contract 210-76-0123.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

THIRAM 5005

((CH3)2NC(=S)S-)2 MW: 240.43 CAS: 137-26-8 RTECS: JO1400000

METHOD: 5005, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984

Issue 3: 15 March 2016

OSHA: 5 mg/m3

NIOSH: 5 mg/m3

; Group I Pesticide

PROPERTIES: white crystalline powder; d 1.29 g/mL; 

MP 155 °C; VP not significant

SYNONYMS: bis(dimethylthiocarbamoyl)disulfide; tetramethylthiuram disulfide; tetramethylthioperoxydicarbonic diamide

SAMPLING

SAMPLER: FILTER (1-μm PTFE membrane)

FLOW RATE: 1 to 4 L/min

VOL-MIN: 10 L

 -MAX: 400 L

SHIPMENT: routine

SAMPLE

STABILITY: 7 days at 25 °C

BLANKS: 2 to 10 field blanks per set

BULK SAMPLES: desirable; 1 to 5g

ACCURACY 

RANGE 

STUDIED: 3 to 12 mg/m3 [1] (240-L samples) 

BIAS: -0.18%

OVERALL

PRECISION (SS�rrrr): 0.055 [1]

ACCURACY: ± 10.67%

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: Thiram

EXTRACTION: (filter) 10 mL CH3CN, 30 min; (cassette top) 

10 mL CH3CN rinse

INJECTION 

 VOLUME: 5 μL

MOBILE PHASE: 60% acetonitrile/40% water, 1 mL/min

COLUMN: C18 (30 cm x 3.9-mm-ID stainless steel); 

ambient temperature

DETECTOR: UV @ 254 nm, 1-cm cell

CALIBRATION: standard solutions of Thiram in acetonitrile

RANGE: 0.1 to 3 mg per sample [1]

ESTIMATED LOD: 0.005 mg per sample [1]

PRECISION (SS�rr): 0.012 [1]

APPLICABILITY: The working range is 0.5 to 15 mg/m3 for a 200-L air sample. NIOSH researchers have used this method at 

facilities that use Thiram as an insecticide. 

INTERFERENCES: None known. 

OTHER METHODS: This is Method S256 [2] in a revised format. An earlier spectrophotometric method, P&CAM 228 [3], has not 

been revised because of excessive analytical variability [4].
THIRAM: METHOD 5005, Issue 3, dated 15 March 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Acetonitrile, HPLC grade.*

2. Water, distilled, deionized.

3. Thiram, reagent grade.*

4. Air or nitrogen, compressed, for drying 

syringes.

5. Calibration stock solution, 0.75 mg/mL. 

Dissolve an accurately weighed 7.5 mg 

Thiram in acetonitrile and dilute to 10 mL. 

Prepare fresh daily in duplicate.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: 1-μm polytetrafluoroethylene (PTFE) 

membrane filter, 37-mm diameter, two-piece 

polystyrene cassette filter holder with backup 

pad, sealed with tape or a shrinkable band. 

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing.

3. Liquid chromatograph, UV detector at 254

nm, integrator and column (page 5005-1).

4. 13 mm x 5 μm PTFE filters and stainless steel 

filter holder to protect the LC column.

5. Vials, 20-mL, glass, PTFE-lined screw caps.

6. Syringe, 1-mL, with luer lock style fitting.

7. Pipets, 10-mL, with pipet bulb.

8. Tweezers.

9. Volumetric flasks, 10-mL. 

 

SPECIAL PRECAUTIONS: Acetonitrile is toxic and flammable; work with it only in a hood. Thiram is an 

irritant of skin and mucous membranes, a skin sensitizer, and suspected teratogen [5].

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at 1 to 4 L/min for a total sample size of 10 to 400 L. Do not exceed 2 mg total dust loading on 

the filter.

3. Collect a bulk sample (1 to 5 g) in a glass vial with PTFE-lined cap; ship separately from filters.

SAMPLE PREPARATION:

4. Remove filter from cassette with tweezers and place in 20-mL vial.

5. Add 10 mL acetonitrile. Cap the vial.

6. Rinse the inside top of cassette with 10 mL acetonitrile into a 20-mL vial. Cap the vial.

7. Agitate samples during the 30-min desorption period. 

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards over the range 0.005 to 3 mg Thiram per sample.

a. Add known amounts of calibration stock solution to acetonitrile in 10-mL volumetric flasks and 

dilute to the mark.

b. Analyze together with samples and blanks (steps 9 and 10).

c. Prepare calibration graph (peak area vs. mg Thiram).

MEASUREMENT:

9. Set liquid chromatograph to conditions on page 5005-1. Inject 10-μL sample aliquot. Rinse and dry 

syringe between injections.

10. Measure peak area.
THIRAM: METHOD 5005, Issue 3, dated 15 March 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALCULATIONS:

11. Read the mass, mg, of Thiram found in the sample filter (Wf) and top rinse (Wt) and in the average 

media blank (B) from calibration graph.

12. Calculate the concentration of Thiram, C (mg/m3

), in the air volume sampled, V (L): 

CC = �WWff + WWtt − BB� ∙ 103

VV , mmmm mm3 ⁄

EVALUATION OF METHOD:

Method S256 was issued on June 8, 1979 [2], and validated by collecting 18 samples (six each at 0.5, 1 and 

2 times the OSHA standard) from dynamically-generated test atmospheres using Thiram 65 (65% Thiram; 

Mayer Chemical Co.), as well as a set of six samples which was stored at room temperature for seven days 

to establish stability [1,4]. The stored sample results were within 2.1% of samples analyzed after one day, 

indicating adequate storage stability for seven days. Eighteen more samples were spiked directly (six each 

at 0.5, 1 and 2 times the OSHA standard). The pooled relative standard deviation for these three sets of 

samples was found to be 0.012. The average recovery for all three levels was 99.8%; therefore, there is no 

bias for this method. The pooled relative standard deviation for the three sets of samples collected from 

test atmospheres was 0.022. Test atmospheres at 12 mg/m3 Thiram were sampled with PTFE filters 

followed by bubblers containing acetonitrile; no detectable Thiram (LOD = 0.005 mg) was found in the 

bubblers indicating that vapor pressure of Thiram was insignificant.

REFERENCES:

[1] Stanford Research Institute [1979]. Backup Data Report: Method S256. Menlo Park, CA: Stanford 

Research Institute. NIOSH contract 210-76-0123. Available as order no. PB-81-244634 from NTIS.

[2] NIOSH [1979]. Thiram in air: Method S256. In: Taylor DG, ed. NIOSH manual of analytical methods. 

2nd. ed. (Vol 5). Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for 

Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) 79-141.

[3] NIOSH [1976]. Thiram in air: Method P&CAM 228. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. (Vol 1). Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center 

for Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication 

No. 77-157-A.

[4] IIT Research Institute [1976]. Failure Report: Method S256. Chicago, IL: IIT Research Institute. NIOSH 

contract no. 99-74-45. Unpublished.

[5] NIOSH [1978] NIOSH criteria for a recommended standard: occupational exposure during 

manufacture and formulation of pesticides, Cincinnati, OH: U.S. Department of Health, Education, 

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, 

DHEW (NIOSH) Publication No. 78-174.

[6] NIOSH [1980]. NIOSH research report – development and validation of methods for sampling and 

analysis of workplace toxic substances. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 80-133. 
THIRAM: METHOD 5005, Issue 3, dated 15 March 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD REVISED BY:

Yvonne T. Gagnon, NIOSH; S256 originally validated under NIOSH Contract 210-76-0123.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ROTENONE 5007

C2 3H2 2O6 MW: 394.43 CAS: 83-79-4 RTECS: DJ2800000

METHOD: 5007, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984 

Issue 3: 26 February 2016

OSHA: 5 mg/m3

 

NIOSH: 5 mg/m3

; Group II Pesticide

PROPERTIES: solid, MP 163 °C or 181 °C; BP 220 °C @ 0.5 mm

Hg; d ca. 1 g/cm3

; VP not significant

SYNONYMS: tubatoxin; cube 

SAMPLING

SAMPLER: FILTER (1-mm PTFE membrane)

FLOW RATE: 1 - 4 L/min

VOL-MIN: 8 L

 -MAX: 400 L 

SHIPMENT: routine

SAMPLE

STABILITY: at least 7 days @ 25 °C in dark

BLANKS: 2 to 10 field blanks per set

BULK SAMPLE: desirable; 1g

ACCURACY

RANGE 

STUDIED: 1 to 11 mg/m3 [1] (100-L sample)

BIAS: -0.6% 

OVERALL

PRECISION (SS�rrrr): 0.079 

ACCURACY: ± 13.5% 

MEASUREMENT

TECHNIQUE: HPLC; UV DETECTION

ANALYTE: Rotenone

EXTRACTION: 4 mL acetonitrile; 30 min

INJECTI ON

VOLUME: 10 μL

MOBILE 

PHASE: 60% methanol/40% water, 2 mL/min

DETECTOR: UV @ 290 nm; 0.1A full-scale; 1-cm cell

COLUMN: C18 (30 cm x 3.9-mm ID stainless steel); 

ambient temperature

CALIBRATION: solutions of Rotenone in acetonitrile

RANGE: 0.04 to 1 mg per sample 

ESTIMATED LOD: 4 μg per sample [1, 2]

PRECISION (SS�rr): 0.024 [1]

APPLICABILITY: The working range is 0.4 to 10 mg/m3 for a 100-L air sample and the method is applicable to commercial 

formulations.

INTERFERENCES: None known. Rotenone, a naturally occurring insecticide, is adequately separated by HPLC from other 

compounds (e.g., sumatrol, α-toxicarol, deguelin, elliptone, malaccol, and tephrosin [3]) present in commercial cube root 

extracts [4]. Rotenone is sensitive to photodecomposition. 

OTHER METHODS: This is Method S300 [2] in a revised format.
ROTENONE: METHOD 5007, Issue 3, dated 2 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Acetonitrile, HPLC grade.*

2. Methanol, HPLC grade.

3. Rotenone, 97% purity.

4. Water, distilled, HPLC grade.

5. Calibration stock solution, 3 mg/mL.

Dissolve 0.075 g Rotenone in 25 mL

acetonitrile. Prepare fresh daily in duplicate.

6. Recovery stock solution, 50 mg/mL.

Dissolve 0.500 g Rotenone in acetone.

Dilute to 10 mL. Prepare fresh daily.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: 37-mm, two-piece cassette

containing 1-μm PTFE membrane filter

with backup pad.

NOTE: Use an opaque cassette or

otherwise shield the filter from light 

to minimize photodecomposition 

of Rotenone during and after 

sampling.

2. Personal sampling pump, 1 to 4 L/min,

with flexible connecting tubing.

3. HPLC, UV detector, integrator and column

(page 5007-1).

4. Jars, ointment, 60-mL, with PTFE-lined

caps.

5. Vials, 4-mL, with PTFE-lined caps.

6. Syringes, 5-mL.

7. Filtration device, 13-mm with 1-μm PTFE

filters, or PTFE syringe filters.

8. Volumetrics, 10- and 25-mL.

9. Syringes, microliter, for sample injection

and standard preparation.

10. Pipet, 4-mL, with pipet bulb.

SPECIAL PRECAUTIONS: Avoid breathing acetonitrile vapors; may cause skin irritation.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 1 and 4 L/min for a total sample size of 8 to 400 L.

Do not exceed 2 mg total dust loading on the filter.

3. Collect a bulk sample (1 g) in a glass vial with PTFE-lined cap; ship separately from filters.

SAMPLE PREPARATION:

4. Open filter cassette; transfer filter to ointment jar.

5. Add 4.0 mL acetonitrile; gently swirl for 30 min.

6. Filter each sample using a 5-mL syringe with PTFE syringe filter or filtration device. Deliver filtrate to

a 4-mL vial.

CALIBRATION AND QUALITY CONTROL:

7. Prepare at least six working standards daily in the range 0.01 to 1 mg Rotenone per sample.

a. Add known amounts of calibration stock solution to acetonitrile in 10-mL volumetric flasks and

dilute to the mark.

b. Analyze together with samples and blanks (steps 9 and 10).

c. Prepare calibration graph (peak area vs. mg Rotenone).
ROTENONE: METHOD 5007, Issue 3, dated 2 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

8. Check recovery (R) with at least three spiked media blanks per sample set in the calibration range

(step 7).

a. Add aliquots of recovery stock solution to blank filters with a microliter syringe. Air dry.

b. Analyze together with working standards (steps 4 through 6, 9 and 10).

c. Calculate recovery [(mg recovered - mg blank)/mg added].

d. Prepare recovery graph (R vs. mg Rotenone).

MEASUREMENT:

9. Set HPLC system according to manufacturer's recommendations and to conditions given on page

5007-1. Inject 10-μL sample.

NOTE: If peak area is above linear range of calibration graph, dilute, reanalyze, and apply

appropriate dilution factor in calculations.

10. Measure peak area.

CALCULATIONS:

11. Read the mass, mg (corrected for recovery) of Rotenone found on the filter (W) and average media

blank (B) from the calibration graph.

12. Calculate the concentration, C (mg/m3

), of Rotenone in the air volume sampled, V (L):

CC = (WW − BB)xx103

VV , mmmm mm3 ⁄

EVALUATION OF METHOD: 

Method S300 [2] was issued on May 11, 1979, and validated over the range 1.16 to 11.1 mg/m3 at 25°C and 

760 mm, using 100-L samples [1, 5]. Overall precision, ŜrT, was 0.079 with average recovery 100.4%, 

representing a non-significant bias. The concentration of Rotenone (generated by Wright dust feeder 

using Ortho Rotenone Dust [1%; Chevron Chemical Co.] enriched to 10% Rotenone with analytical grade 

Rotenone [Aldrich Chemical Co.]) was independently verified by collection in dioxane and HPLC analysis.

Recovery was 0.98 in the range 250 to 1000 μg Rotenone per sample. Collection efficiency of the PTFE filter 

was found to be greater than 99% and no detectable Rotenone (LOD = 4 μg) was found on Chromosorb

102 tubes placed behind the PTFE filters at 11.8 mg/m3

. No loss was seen from spiked filters stored in the

dark at room temperature for seven days.

REFERENCES:

[1] NIOSH. Backup data report. Unpublished. Available as Order No. PB 82-114729 from NTIS, 

Springfield, VA.

[2] NIOSH [1979]. Rotenone: Method S300. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd 

ed. (Vol 5). Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 79-

141.

[3] Gunther FA, Blinn RG [1975]. Analysis of insecticides and acaricides. NY: Interscience, pp 419-420.

[4] Bushway RJ, Engdahl BS, Colvin BM, Hanks AR [1975]. Separation of rotenoids and the determination 

of rotenone in pesticide formulations by high-performance liquid chromatography. J Assoc Off Anal 

Chem 58(5):965-970.
ROTENONE: METHOD 5007, Issue 3, dated 2 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[5] NIOSH [1980]. NIOSH research report-development and validation of methods for sampling and 

analysis of workplace toxic substances. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 80-133.

METHOD REVISED BY:

Jerome Smith, Ph.D., NIOSH; S300 originally validated under NIOSH Contract 210-76-0123. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

PYRETHRUM 5008

C2 0H2 8O3 to C2 2H3 0O5 MW: 316.4-372.4 CAS: 8003-34-7 RTECS: UR4200000

(Table 1) (active constituents) 

METHOD: 5008, Issue 3 EVALUATION: FULL Issue 1: 15 May 1985 

Issue 3: 25 February 2016

OSHA: 5 mg/m3

NIOSH: 5 mg/m3; Group II pesticide 

PROPERTIES: viscous brown resin or solid; 

VP not significant 

SYNONYMS: Active constituents include pyrethrin I and II, jasmolin I and II, and cinerin I and II. 

SAMPLING

SAMPLER: FILTER (glass fiber, 37 mm)

FLOW RATE: 1 - 4 L/min

VOL-MIN: 20

 -MAX: 400 L

SHIPMENT: routine; ship bulk sample separately

SAMPLE 

STABILITY: at least 1 week @ 25 °C [1] 

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 1.4 to 8.5 mg/m3 [1] (132-L samples)

BIAS: -4.5% 

OVERALL

PRECISION (SS�rrrr): 0.070 [1] 

ACCURACY: ± 13.8% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: six active constituents of pyrethrum

EXTRACTION: 10 mL acetonitrile; stand 30 min

INJECTION

VOLUME: 25 μL 

COLUMN: C18, reverse phase, 10-μm packing, 

25 to 30 cm

MOBILE PHASE: 85% acetonitrile/15% water, isocratic, 

1.0 mL/min, room temperature, 2000 

kPa (400 psi)

DETECTOR: UV absorption @ 225 nm 

CALIBRATION: solutions of pyrethrum in acetonitrile 

RANGE: 0.1 to 1.8 mg/sample 

ESTIMATED LOD: 0.01 mg/sample

PRECISION (SS�rr): 0.040 [1]

APPLICABILITY: The working range is 0.5 to 10 mg/m3 for a 200-L air sample.

INTERFERENCES: Specific interferences have not been studied. Mass spectrometry or gas-liquid chromatography with electroncapture detection may be needed for confirmation [1].

OTHER METHODS: This method is S298 [2] in a revised format. For bulk samples, gas-liquid chromatography with electron-capture 

detection has been recommended [3]. A modified HPLC method (C8 column, programmed methanol/water or acetonitrile, 

isocratic) has been used [4].
PYRETHRUM: METHOD 5008, Issue 3, dated 5 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Pyrethrum*, analytical standard solution 

(commercially available). 

NOTE: Store away from direct light. Active 

constituents of pyrethrum oxidize in air and 

photo- decompose.

2. Acetonitrile, HPLC grade.*

3. Water, HPLC grade.

4. Isopropanol.

5. Calibration stock solution,* 60 mg/mL, in 

isopropanol.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: 37-mm glass fiber filter and 

cellulose backup pad in filter holder held 

together with tape or shrinkable band.

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing.

3. High pressure liquid chromatograph with 

UV absorption detector at 225 nm, 

integrator and column (page 5008-1).

4. Ointment jars, 2-oz, squat form, with 

polytetrafluoroethylene (PTFE)-lined screw 

caps.

5. Tweezers.

6. Pipet, TD, 10-mL, with pipet bulb.

7. Volumetric flasks, 10-mL.

8. Syringe, 10-mL, with syringe filter.

9. Syringes or pipets, 5- to 100-μL. 

SPECIAL PRECAUTIONS: p-Chlorophenol is toxic by skin absorption, inhalation, or ingestion. It also is a

strong irritant to tissue and is combustible with a flash point of 121 °C.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 1 and 4 L/min for a total sample size of 20 to 400 L.

SAMPLE PREPARATION:

3. Transfer the filter carefully to an ointment jar using tweezers.

4. Add 10.0 mL acetonitrile. Seal and gently swirl the jar to wet the filter. Let stand 30 min with 

occasional swirling.

5. Filter the sample solution through a syringe filter.

CALIBRATION AND QUALITY CONTROL:

6. Calibrate daily with at least six working standards over the range 0.01 to 1.8 mg pyrethrum per sample.

a. Add known amounts of calibration stock solution, or a dilution thereof, to acetonitrile in 10-mL 

volumetric flasks and dilute to the mark.

b. Analyze together with the samples and blanks (steps 9, 10 through 11).

c. Prepare calibration graph (peak area vs. mg pyrethrum/10 mL).

7. Determine recovery (R) at least once for each batch of filters used for sampling in the range of the 

samples. Prepare three filters at each of five levels plus three media blanks.

a. Deposit a known amount of calibration stock solution, or a dilution thereof, onto the filters. Allow 

filters to air-dry.

b. Store samples overnight in ointment jars.

c. Prepare (steps 4 and 5) and analyze together with working standards (steps 9 through 11).

d. Prepare a graph of R vs. mg pyrethrum recovered.

8. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration graph 

and R graph are in control.
PYRETHRUM: METHOD 5008, Issue 3, dated 5 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

MEASUREMENT:

9. Set HPLC system according to manufacturer's recommendations and to conditions on page 5008-1.

10. Inject sample aliquot using syringe, fixed volume sample loop, or autosampler.

11. Measure peak area.

NOTE: Pyrethrum is a mixture of at least six components which elute in two major peaks

(tr = 5 to 7 min under these conditions). The minor peaks have been shown by mass spectrometry not 

to be pyrethrums. The components may be separated by gas chromatography [1,4].

CALCULATIONS:

12. Determine the mass, mg (corrected for R) of pyrethrum found in the sample (W) and in the average 

media blank (B) from the calibration graph.

13. Calculate concentration, C, of pyrethrum in the air volume sampled, V (L):

CC = (WW − BB)xx103

VV , mmmm/mm3

EVALUATION OF METHOD:

Method S298 [2] was issued on August 3, 1979, and validated over the range 1.4 to 8.5 mg/m3 using 

atmospheres generated from Premium Pyrocide 175 (McLaughlin Gormley King Co.) [1]. Standards and 

collected filter samples were analyzed by GC/MS. Lab testing was done with spiked filters and 

atmospheres dynamically generated by atomization of a hydrocarbon solution; verified by PTFE 

filter/isooctane bubbler analyzed by gas chromatography with electron capture detector (absence of 

pyrethrum in the bubbler was established by mass spectrometry). Samples containing 0.7 mg pyrethrum 

collected from a test atmosphere were stable for seven days at ambient conditions (average recovery = 

98.2%). Collection efficiency equal 99.7% for 120-L samples collected at 1 L/min at 9 mg/m3

. Precision and 

accuracy are given on page 5008-1.

REFERENCES:

[1] NIOSH [1979]. Backup Data Report: Method S298. Unpublished.

[2] NIOSH [1980]. Pyrethrum: Method S298. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd 

ed. (Vol 6) Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 80-125.

[3] Association of Official Analytical Chemists (AOAC) [1982]. Changes in official methods of analysis. J 

Assoc Off Anal Chem 65: 455-456.

[4] Hultquist A, Izumi RM [1982]. NIOSH sequence reports 3481-J and 4151-J. Salt Lake City, UT: Utah 

Biomedical Test Laboratory. Unpublished.

[5] NIOSH [1981]. Occupational health guidelines for chemical hazards. Cincinnati, OH: U.S. Department 

of Health and Human Services, Centers for Disease Control, National Institute for Occupational Safety 

and Health, DHHS (NIOSH) Publication No. 81-123. 
PYRETHRUM: METHOD 5008, Issue 3, dated 5 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD REVISED BY:

James E. Arnold, NIOSH

Table 1. Active constituents of Pyrethrum.

Compound Formula M.W. CAS #

Cinerin I C20H28O3 316.44 25402-06-6 

Cinerin II C21H28O5 260.45 121-20-0 

Jasmolin I C21H30O3 330.47 4466-14-2 

Jasmolin II C22H30O5 374.48 1172-63-0 

Pyrethrin I C21H28O3 328.45 121-21-1 

Pyrethrin II C22H28O5 372.46 121-29-9 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

BENZOYL PEROXIDE 5009

(C6H5CO)2O2 MW: 242.22 CAS: 94-36-0 RTECS: DM85750000

METHOD: 5009, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984 

Issue 3: 26 February 2016

OSHA: 5 mg/m3

NIOSH: 5 mg/m3

PROPERTIES: solid; MP 103 °C; d 1.334 g/mL @ 25 °C; VP not

significant; auto ignition temperature 80 °C

SYNONYMS: dibenzoyl peroxide, benzoyl superoxide 

SAMPLING

SAMPLER: FILTER (0.8-μm cellulose ester membrane) 

FLOW RATE: 1 - 3 L/min

VOL-MIN: 40 L @ 5 mg/m3

 -MAX: 400 L 

SHIPMENT: refrigerated

SAMPLE 

STABILITY: 9% loss from filter after 1 week @ 25 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 3 to 9 mg/m3 [1]; (90-L samples)

BIAS: -0.52% 

OVERALL

PRECISION (SS�rrrr): 0.06 [1] 

ACCURACY: ± 11.82% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: benzoyl peroxide

EXTRACTION: ethyl ether, 10 mL

PRESSURE- 

 COLUMN: 9000 KPa (1300 psi)

MOBILE PHASE: 70/30 methanol/water; 1.6 mL/min

COLUMN: 250 mm x 3 mm-ID stainless steel; 

spherical silica particles with 5% bonded 

coating of octadecyl groups, US 

Pharmacopeia (USP) L1

DETECTOR: UV photometer @ 254 nm

CALIBRATION: benzoyl peroxide in ethyl ether

RANGE: 0.2 to 1.7 mg per sample [1] 

ESTIMATED LOD: 0.01 mg per sample [2] 

PRECISION (SS̅

rr): 0.024 [1]

APPLICABILITY: The working range is 2 to 19 mg/m3 for a 90-L air sample.

INTERFERENCES: None identified

OTHER METHODS: This is Method S253 [3] in a revised format. A non-specific gravimetric method and a non-specific colorimetric

method appear in the criteria document [4].
BENZOYL PEROXIDE: METHOD 5009, Issue 3, dated 26 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Benzoyl peroxide, 99% pure.*

2. Ethyl ether, purified, without stabilizer.*

3. Methanol, distilled in glass.

4. Water, deionized and distilled.

5. Calibration stock solution, 10 mg/mL. 

Dissolve 250 mg benzoyl peroxide in ethyl 

ether and dilute to 25 mL. Stable at least one 

week at 4 °C.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: cellulose ester membrane filter, 0.8-

μm pore size, 37-mm diameter, with backup 

pad in cassette filter holder.

2. Personal sampling pump, 1 to 3 L/min, with 

flexible connecting tubing.

3. Refrigerant, water solution, sealed, refreezable, 

reusable.

4. Vials, 20-mL, polytetrafluoroethylene (PTFE)-

lined caps.

5. High performance liquid chromatograph, 254-

nm UV detector, sample injection valve with a 

20-μL external sample loop, syringe filter, 

integrator and column (page 5009-1) or 

autosampler. 

6. Tweezers.

7. Microliter syringes, 10- and 100-μL.

8. Volumetric flasks, 10- and 25-mL, assorted 

sizes. 

9. Pipet, 10-mL, with pipet bulb.

SPECIAL PRECAUTIONS: Benzoyl peroxide is a flammable solid and may explode when heated; it will 

attack some plastics, rubber, and coatings [4]. Ethyl ether is highly flammable and forms explosive 

peroxides on exposure to air.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 1 and 3 L/min for a total sample size of 40 to 400 L. 

Do not exceed 2 mg particulate loading on the filter.

3. Ship samples in an insulated container with bagged refrigerant.

SAMPLE PREPARATION:

4. Immediately upon receipt at laboratory, refrigerate the samples.

5. As soon as possible after receipt, transfer each filter to a clean vial with tweezers.

6. Pipet 10 mL ethyl ether into each vial; screw on cap. Swirl to mix.

CALIBRATION AND QUALITY CONTROL:

7. Calibrate daily with at least six working standards covering the range 0.01 to 1.7 mg per sample.

a. Add calibration stock solution with a microliter syringe to ethyl ether in a 10-mL volumetric flask 

and dilute to the mark.

b. Analyze together with samples, blanks and control samples (steps 10 through 12).

c. Prepare calibration graph of peak area vs. mass (mg) of benzoyl peroxide per sample.

8. Determine recovery of benzoyl peroxide from filters at least once for each lot of samplers in the 

calibration range (step 7). Prepare three filters at each of five levels plus media blanks.

a. Add aliquot of calibration stock solution with a microliter syringe directly onto a media blank filter 

in a vial.

b. Prepare and analyze together with working standards (steps 5, 6 and 10 through 12).

c. Prepare a graph of recovery vs. mg benzoyl peroxide recovered.
BENZOYL PEROXIDE: METHOD 5009, Issue 3, dated 26 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

9. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration 

graph and recovery graph are in control.

MEASUREMENT:

10. Set high performance liquid chromatograph according to manufacturer's recommendations and to 

conditions given on page 5009-1.

11. Flush sample loop thoroughly with sample (0.1 mL) and inject sample or autosampler. 

12. Measure peak area.

CALCULATIONS:

13. Read mass, mg (corrected for recovery), of benzoyl peroxide found on sample filters, W (mg), and on 

average blank filters, B (mg), from calibration graph.

14. Calculate concentration, C (mg/m3

), of benzoyl peroxide in the air volume sampled, V (L):

CC = (WW − BB)xx103

VV , mmmm mm3 ⁄

EVALUATION OF METHOD

Method S253 was issued on January 21, 1977 [3], and validated over the range 3.1 to 19.1 mg/m3 at 26 °C 

and 764 mm Hg, using 90-L samples [1, 5]. The collection efficiency of the filter was determined to be 1.00, 

since no benzoyl peroxide was detected on a backup filter mounted directly behind the front filter. 

Storage stability studies on the filters held in the filter cassettes indicated a 9.3% decrease in the amount of 

benzoyl peroxide recovered after one week. Benzoyl peroxide was stable in ethyl ether at room 

temperature for at least one week. Overall precision, SS�rrrr, was 0.060. Recovery was 0.97 in the range 0.225 

to 0.900 mg per sample for 18 spiked samples.

REFERENCES:

[1] NIOSH [1976]. Backup data report: S253 for Benzoyl Peroxide. In: Ten NIOSH analytical methods (Set 

2). Available as order No. PB 271-464 from NTIS. Unpublished.

[2] Wade R [1980]. Memorandum: UBTL analytical laboratory report for benzoyl peroxide, Sequence 

#2182-J. Salt Lake City, UT: Utah Biomedical Test Laboratory. Unpublished.

[3] NIOSH [1978]. Benzoyl peroxide: Method S253. In: NIOSH manual of analytical methods. 2nd ed. (Vol 

4). Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 78-175.

[4] NIOSH [1977]. NIOSH criteria for a recommended standard: occupational exposure to benzoyl 

peroxide. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

166.

[5] Gunderson EC, Anderson CC [1980]. Development and validation of methods for sampling and 

analysis of workplace toxic substances. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 80-133.
BENZOYL PEROXIDE: METHOD 5009, Issue 3, dated 26 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD REVISED BY:

Charles Neumeister, NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

STRYCHNINE 5016 

C2 1H2 2N2O2 MW: 334.42 CAS: 57-24-9 RTECS: WL2275000

METHOD: 5016, Issue 3 EVALUATION: FULL Issue 1: 15 May 1985 

Issue 3: 26 February 2016

OSHA: 0.15 mg/m3

NIOSH: 0.15 mg/m3/10h; Group I Pesticide

PROPERTIES: solid; MP 268 °C; VP not significant

SYNONYMS: strychnidin-10-one

SAMPLING

SAMPLER: FILTER (glass fiber, 37-mm) 

FLOW RATE: 1 - 3 L/min

VOL-MIN: 70 L @ 0.15 mg/m3

 -MAX: 1000 L 

SHIPMENT: routine

SAMPLE

STABILITY: at least 7 days @ 25 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY 

RANGE 

STUDIED: 0.073 to 0.34 mg/m3

 [3] 180-L samples)

BIAS: 4.4%

OVERALL

PRECISION (SS�rrrr): 0.059 [1] 

ACCURACY: ± 14.6% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: strychnine

DESORPTION: 5 mL mobile phase

INJECTION

 VOLUME: 20 μL

MOBILE PHASE: aqueous 1-heptane sulfonic acid + 

CH3CN; pH 3.5; 1 mL/min @ ambient 

temperature

COLUMN: 25 cm x 4.2-mm ID, packed with C18/US 

Pharmacopeia (USP) L1, 10-μm particle 

size

DETECTOR: UV absorption @ 254 nm

CALIBRATION: solutions of strychnine in mobile phase 

RANGE: 10 to 70 μg per sample 

ESTIMATED LOD: 0.8 μg per sample [1,2] 

PRECISION (SS̅

rr): 0.042 @ 13 to 62 μg per sample [1]

APPLICABILITY: The working range is 0.05 to 10 mg/m3 for a 200-L air sample.

INTERFERENCES: None known

OTHER METHODS: This revises Method S302 [2].
STRYCHNINE: METHOD 5016, Issue 3, dated 26 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Strychnine, 98%.*

2. Acetonitrile, chromatographic quality.

3. Water, chromatographic quality, distilled.

4. Acetonitrile:water, 1:1 (v/v). Mix equal

volumes of acetonitrile and distilled water.

5. Acetic acid, glacial.*

6. Acetic acid, 0.01 N. Dilute 0.6 mL glacial acetic

acid to 1 L with distilled water.

7. 1-Heptanesulfonic acid sodium salt, 95%.

8. Calibration stock solution, 1 mg/mL

strychnine in 0.01 N acetic acid. Use an

ultrasonic bath to aid dissolution.

9. Mobile phase: Dissolve 1.1014 g 1-

heptanesulfonic acid sodium salt in 980 mL

1:1 acetonitrile:water. Adjust pH to 3.5 with

glacial acetic acid. Dilute to 1 L with 1:1

acetonitrile:water. Filter through 0.45-μm

PTFE filter and refrigerate. Prepare fresh

weekly and if cloudiness appears.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: glass fiber filter, 37-mm diameter, in

cassette filter holder.

2. Personal sampling pump, 1 to 3 L/min, with

flexible connecting tubing.

3. HPLC, UV absorption detector at 254 nm,

recorder, integrator and column (page 5016-

1).

4. Syringe filter, polypropylene housing, nylon

membrane, 0.2-μm pore size.

5. Syringe, 20-μL.

6. Micropipets or syringes, 10- to 100-μL.

7. Jars, glass, squat-form ointment, with

polytetrafluoroethylene (PTFE)-lined screw

caps.

8. Volumetric flasks, glass, 10-mL.

9. Pipet, 5-mL delivery, with pipet bulb.

10. Tweezers.

11. Ultrasonic bath.

SPECIAL PRECAUTIONS: Strychnine is a potent convulsant. Use impervious clothing, gloves and face 

shields when handling it [3,4]. Avoid skin contact with acetic acid as it may cause severe burns.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 1 and 3 L/min for a total sample size of 70 to

1000 L.

SAMPLE PREPARATION:

3. Transfer the filter carefully using tweezers to an ointment jar.

4. Add 5.0 mL mobile phase. Seal and gently swirl the jar to wet the filter.

5. Filter the sample solution through a syringe filter.

CALIBRATION AND QUALITY CONTROL:

6. Calibrate daily with at least six working standards over the range 1 to 70 μg strychnine per sample

(0.2 to 14 μg/mL).

a. Add known amounts of calibration stock solution to mobile phase in 10-mL volumetric flasks and

dilute to the mark.

b. Analyze together with the samples and blanks (steps 9, 10, and 11).

c. Prepare calibration graph (peak area vs. μg strychnine).

7. Determine recovery (R) at least once for each lot of filters used for sampling in the calibration range.

Prepare three filters at each of five levels plus three media blanks.

a. Deposit a known amount of calibration stock solution onto the filter. Allow filters to air dry.

b. Store samples overnight in ointment jars.

c. Prepare for analysis (steps 3 through 5) and analyze together with working standards (steps 9

through 11).
STRYCHNINE: METHOD 5016, Issue 3, dated 26 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

d. Prepare a graph of R vs. μg strychnine recovered.

8. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration

graph and R graph are in control.

MEASUREMENT:

9. Set HPLC system according to manufacturer's recommendations and to the conditions given on page

5016-1.

10. Inject sample aliquot using syringe, fixed volume sample loop or autosampler.

11. Measure peak area.

CALCULATIONS:

12. Determine the mass, μg (corrected for R) of strychnine found in the sample (W) and in the average

media blank (B) from the calibration graph.

13. Calculate concentration, C, of strychnine in the air volume sampled, V (L):

CC = WW − BB

VV , mmmm mm3 ⁄

EVALUATION OF METHOD:

Method S302 was issued on February 17, 1978 [2], and evaluated over the range 0.073 to 0.34 mg/m3 using 

a 180-L sample [1,5]. Overall precision, SS�rrrr, was 0.059 with an average recovery of 100.9%, representing a 

non-significant bias. The concentration of strychnine was independently verified by analysis of filter 

samples with a UV spectrophotometer at 254 nm. Recovery was 0.98 in the range of 13.5 to 54.1 μg per 

sample. Collection efficiency was determined to be at least 98.8% for 180-L sample at a concentration to 

be 0.15 μg/mL. Average recovery for one-day old samples was 105% vs. 98.8% for seven-day old samples 

stored at room temperature. Since the filter samples for the recovery and stability studies were prepared 

with strychnine acetate rather than the free base, the recovery of the free base and the stability of such 

samples is unknown.

REFERENCES:

[1] NIOSH [1976]. Back-up data report for Strychnine. In: Ten NIOSH analytical methods. Available as 

order No. PB288-629 from NTIS. Unpublished.

[2] NIOSH [1979] Strychnine: Method S302. In: Taylor, DG ed. 2nd ed. (Vol 5). NIOSH manual of 

analytical methods. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for 

Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication 

No. 79-141.

[3] NIOSH [1978]. NOSH criteria for a recommended standard: occupational exposure during the 

manufacture and formulation of pesticides. Cincinnati, OH: U.S. Department of Health, Education, 

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, 

DHEW (NIOSH) Publication No. 78-174. Available as order No. PB81-227001 from NTIS.

[4] NIOSH [1981]. Occupational health guidelines for chemical hazards. Cincinnati, OH: U.S. 

Department of Health and Human Services, Centers for Disease Control, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 81-123.
STRYCHNINE: METHOD 5016, Issue 3, dated 26 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[5] NIOSH [1980]. Development and validation of methods for sampling and analysis of workplace and 

toxic substances. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 80-133.

METHOD REVISED BY:

M.J. Seymour, NIOSH

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

ARSENIC, organo- 5022

(1) CH3AsO3H2 MW: (1) 139.96 CAS: (1)124-58-3 RTECS: (1) PA1575000

(2) (CH3) 2AsO2H (2) 137.99 (2)75-60-5 (2) CH7525000

(3) H2NC6H4AsO3H (3) 217.07 (3) 98-50-0 (3) CF7875000

METHOD: 5022, Issue 3 EVALUATION: FULL Issue 1: 15 May 1985 

Issue 3: 3 March 2016

OSHA: 0.5 mg/m3 (as As)

NIOSH: None

PROPERTIES: (1) Solid; MP 161 °C

(2) Solid; MP 195 °C

(3) Solid; MP 232 °C

SYNONYMS: (1) Methylarsonic acid: methanearsonic acid. (2) Dimethylarsinic acid: cacodylic acid; hydroxydimethyl arsine 

oxide. (3) p-Aminophenyl arsonic acid: p-arsanilic acid; atoxylic acid.

SAMPLING

SAMPLER: FILTER

(1-μm PTFE)

FLOW RATE: 1 - 3 L/min

VOL-MIN: 50 L @ 0.1 mg/m3

 -MAX: 1000 L 

SHIPMENT: routine

SAMPLE 

STABILITY: stable

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 0.005 to 0.2 mg/m3 [1,2] 

BIAS: none significant 

OVERALL

PRECISION (SS�rrrr): 0.047 @ 0.02 mg/m3 [1];

0.14 @ 0.005 mg/m3 [1]

ACCURACY: ± 20% @ 0.02 mg/m3

MEASUREMENT

TECHNIQUE: ION CHROMATOGRAPHY/HYDRIDE 

ATOMIC ABSORPTION

ANALYTE: anions (IC); AsH3 (AAS) 

EXTRACTION: borate-carbonate buffer, 25 mL

ION CHROMATOGRAPHY: 

INJECTION

LOOP VOLUME: 0.8 mL 

COLUMNS: two, 3 x 150-mm anion

ELUENT: borate-carbonate buffer; 2.5 

mL/min; 3450 kPa (500 psi); 

ambient temperature

AAS: 

QUARTZ 

FURNANCE: 800 °C

WAVELENGTH: 193.7 nm (no D2) 

CALIBRATION: organoarsenicals in water

RANGE: 0.5 to 2 μg As per sample

ESTIMATED LOD: 0.2 μg As per sample [1]

PRECISION (SS�rr): Table 1

APPLICABILITY: The working range is 0.005 to 10 mg/m3 (as As) for a 100-L air sample. The method is designed to quantitate 

particulate organo-arsenic compounds.

INTERFERENCES: Inorganic arsenic (III) co-elutes with dimethylarsenic acid using Eluent A but the two may be separated with 

Eluent B. Other ions at high concentrations in the sample can interfere with the chromatographic separation of the arsenicals. 

As2O3 is not efficiently sampled by this sampler; for quantitation of that compound see Method 7901. 

OTHER METHODS: This is P&CAM 320 in revised format [2]. Method 7200 measures total As by hydride/AAS. Method 7901 

measures As2O3, which can exist as a vapor and aerosol. 
ARSENIC, organo-: METHOD 5022, Issue 3, dated 3 March 2016 - Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Deionized water.

2. Hydrochloric acid, conc.

3. Eluent A (2.4 mM HCO3

-

/1.9 mM CO3

2-/1.0 

mM B4O7

2-). Dissolve 0.8067 g NaHCO3, 

0.8055g Na2CO3, and 1.5257 g Na2B4O7

•10H2O in 4 L deionized water.

4. Eluent B (5 mM B4O7

2-). Dissolve 7.6284 g 

Na2B4O7 • 10 H2O in 4L deionized waster.

5. Potassium persulfate solution,* K2S2O8, 

saturated in 15% (v/v) HCl.

6. Sodium borohydride, 1% NaBH4 (w/v) in 

0.2% KOH (w/v). Add 5 g NaBH4 and 1 g 

KOH to deionized water; dilute to 500 mL. 

Prepare fresh weekly.

7. Stock standards, 1000 μg As/mL:

a. Methylarsonic acid.* Dissolve 0.9341 g 

CH3AsO3H2, in deionized water; dilute 

to 500 mL.

b. Dimethylarsenic acid.* Dissolve 

0.9210 g (CH3)2AsO2H in deionized 

water; dilute to 500 mL.

c. p-Aminophenylarsonic acid.* Dissolve 

1.4485 g p-H2NC6H4AsO3H2 in 5 mL 1 N 

NaOH. Dilute to 500 mL with 

deionized water. Protect from light.

d. Arsenic trioxide.* Dissolve 0.6602 g 

As2O3 in 5 mL 1 N NaOH. Dilute to 500 

mL with deionized water.

e. Arsenic pentoxide.* Dissolve 0.7669 g 

As2O5 in 5 mL 1 N NaOH. Dilute to 500 

mL with deionized water.

8. Calibration stock solution, 1 μg/mL mixed 

analyte. Dilute 0.1 mL of each stock 

standard (REAGENTS, 7.) with Eluent A in a 

100-mL volumetric flask. Prepare fresh 

daily.

9. Argon.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: PTFE polyethylene-backed 

membrane filter, 1-μm pore size, 37-mm 

diameter with backup pad; in cassette filter 

holder.

2. Personal sampling pump, 1 to 3 L/min, with 

flexible connecting tubing. 

3. Ion chromatograph with suppressor and 

detector bypassed. Route column effluent 

via PTFE tubing (0.3mm ID x 0.6mm OD) 

directly into arsine generator (Figure 1). 

4. Syringes, plastic, 10-mL, with male luer lock 

style fittings.

5. Arsine generator: proportioning pump with 

flow-rated pump tubes and 1.5-mm ID x 3-

mm OD manifold mixing coils, 5-turn and 20-

turn, 1.5-mm ID x 3.5-mm OD glass "T" 

connectors; gas-liquid separator and 

expansion chamber (Figure 2); 1 m of 1/4" 

OD PTFE tubing; three PTFE 1/4" ID Swagelok 

fittings; and rotometer (100 to 900 mL/min).

6. Atomic absorption spectrophotometer 

(reciprocal linear UV dispersion 0.65

nm/mm); As electrodeless discharge lamp 

and power supply; and atomization cell (16-

cm x 13-mm ID windowless quartz tube with 

18-cm x 4-mm ID inlet tube fused in the 

center), wound with Nichrome wire (14 Ω/m, 

spaced 2 to 3 mm between turns and 

wrapped with heat resistant tape) (Figure 3). 

Temperature in the cell is measured by a 

thermocouple (800 °C). Mount the cell on top 

of a single-slot AAS burner head and align 

with burner alignment controls.

7. Beakers, 50-mL.** 

8. Ultrasonic waterbath.

9. Volumetric flasks, 10-, 100- and 500-mL.**

10. Pipets, 25-μL and 0.1- to 1-mL.

**Soak all glassware in mild detergent, rinse with 

deionized water, 10% HNO3, and deionized water.

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling chemicals. All work 

should be performed in a fume hood. Potassium persulfate is a powerful oxidizing agent. Arsine gas is 

extremely toxic and can be fatal. The arsenic compounds used in the stock standards are poisonous [3].
ARSENIC, organo-: METHOD 5022, Issue 3, dated 3 March 2016 - Page 3 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 1 and 3 L/min for a total sample volume of 50 to 

1000 L.

3. Cap the cassettes and pack securely for shipment.

SAMPLE PREPARATION:

4. For each sample, pipet 25 mL Eluent A into a clean 50-mL beaker.

5. Open the cassette, remove the PTFE filter with clean forceps, and transfer it to the beaker. Place the 

exposed side of the filter in contact with the solution. Cover beaker.

6. Agitate contents of the beaker for 30 min in an ultrasonic water bath. If the extracts will not be analyzed 

immediately, store at ca. 4 °C until measurement. 

CALIBRATION AND QUALITY CONTROL:

7. Calibrate daily with at least six working standards over the range 0.2 to 2 μg As per sample (0.008 to 0.08 

μg As/mL).

a. Add known amounts of calibration stock solution to Eluent A in 10-mL volumetric flasks and dilute 

to the mark.

b. Analyze together with the samples and blanks (steps 8 through 12).

c. Prepare calibration graph for each arsenic species (peak area or height vs. μg As). 

MEASUREMENT:

8. Set the ion chromatograph to the conditions given on page 5022-1. Allow the columns to equilibrate 

with eluent >1 h before connecting effluent to the arsine generator.

NOTE: Eluent A allows the separation of methylarsonic acid (retention time (tr) = 2 min), 

p-aminophenylarsonic acid (tr = 4 min), and As(V) (tr = 7.5 min); As(III) and dimethylarsenic acid 

(tr = 1 min) are not resolved. If a signal is obtained at the approximate retention time of the 

latter two compounds, or if both compounds are known to be present in the sample, perform a 

second analysis using Eluent B (lower ionic strength). If either of the two compounds is known 

not to be present, Eluent A will effectively determine the remaining compounds. With Eluent B 

the other species have very long retention times and will accumulate on the column, tying up 

active resin sites. Therefore, flush the column with Eluent A after each 10 to 15 samples and 

reequilibrate with Eluent B before further analysis. 

9. Connect the IC effluent to the arsine generator into which the following flow:

Saturated K2S2O8 solution: 0.8 mL/min 

NaBH4 solution: 2.0 mL/min

Ar carrier gas: 300 mL/min

NOTE: The gaseous arsines formed in the arsine generator are first separated from liquid solution using 

the gas-liquid separator (Figure 2) and then transferred by argon carrier gas through PTFE 

tubing to the heated quartz furnace.

10. Set the AAS according to manufacturer's recommendations and to the conditions given on page 

5022-1. Align the quartz cell in the optical path. Heat the quartz cell gradually to 800 °C using a 

variable transformer and thermocouple.

11. Using a syringe, inject a sample aliquot (ca. 2 to 3 mL) into the chromatograph, flushing the injection 

loop to avoid contamination from the previous injection. Rinse the syringe with deionized water and 

dry it between samples, or use disposable syringes.

12. Identify the component peaks. Measure peak height or area.
ARSENIC, organo-: METHOD 5022, Issue 3, dated 3 March 2016 - Page 4 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Calculations

13. From the calibration graphs, calculate the amount (μg) of arsenic for each species in the sample (W) 

and in the average media blank (B).

14. Calculate the arsenic concentrations, C, (mg/m3) in the air volume sampled, V (L):

CC = WW − BB

VV , mmmm/mm3

EVALUATION OF METHOD: 

The measurement precision obtained under the conditions recommended in this procedure is presented in 

Table 1 [1]. The overall precision of the method was tested using filters loaded in a dynamic aerosol 

generation/sampling system with particulates of the three organoarsenical compounds. The 

concentration levels tested for each species were 5, 10, and 20 μg As/m3 of air. Depending on the 

concentration and species, the relative standard deviation ranged from 14.4% at the lowest level to 4.7% at 

the highest level.

The collection efficiency of the method for organoarsenicals in the range of 5 to 20 μg/m3 using a 300-L 

sample was found to be >99%. The collection efficiency of the method for inorganic arsenic was not 

determined.

The accuracy of the overall method was determined by analyzing additional aerosol samples from each set 

using Neutron Activation (NAA) and X-ray Fluorescence (XRF) analyses. Since NAA and XRF techniques 

provide only the total elemental arsenic, the total arsenic obtained from the IC-AAS analysis was used for 

comparison. The accuracy ranged from 90 to 120% of the values obtained by NAA and XRF.

REFERENCES:

[1] Colovos G, Hester N, Ricci GR, Shepard, LS. [1980]. Development of a method for the determination of 

organoarsenicals in air. NIOSH Contract #210-77-0134, NTIS No. PB83-180794.

[2] NIOSH [1980]. Particulate arsenicals: Method P&CAM 320. In: Taylor, DG, ed. NIOSH manual of 

analytical methods, 2nd ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers 

for Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 80-125.

[3] Budavari S, [1989]. Merck Index. 11th ed. Rahway, NJ: Merck & Co., Inc.

METHOD REVISED BY:

Mary Ellen Cassinelli, NIOSH; 

P&CAM 320 originally developed under NIOSH Contract 210-77-0134.
ARSENIC, organo-: METHOD 5022, Issue 3, dated 3 March 2016 - Page 5 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 1. Sensitivity, detection limit and working range data for analysis of particulate 

arsenicals [1].

Arsenical

Sensitivity 

(ng/mL/

1% Abs)

Detection limit 

(as As) for 300 L 

sample volume 

(μg/m3) 

Detection 

limit (as As) 

for solution 

(ng/mL)

Range* for 

300 L sample 

volume 

(μg/m3) 

Range* 

for 

solution 

(ng/mL)

Measurement 

precision

(% SS�r)

Dimethylarsenic 

acid 1.3 0.62 7 1.7-6.7 20-80 11.2

Arsenic (III) 2.1 0.71 8 1.7-6.7 20-80 11.2

Methylarsonic acid 2.1 0.72 9 1.7-6.7 20-80 8.1

pAminophenylarsonic

acid

6.3 0.64 8 1.7-6.7 20-80 6.0

Arsenic (V) 13.0 0.46 6 1.7-6.7 20-80 10.8

*The upper limit of the range can be increased by using higher concentration standards which are

injected via loops of smaller volume. Although not tested with air samples, the useful range can be

extended from 5 μg/m3 down to 1.7 μg/m3 based upon the measurement range.

FIGURE 1. IC/AAS Analytical System.
ARSENIC, organo-: METHOD 5022, Issue 3, dated 3 March 2016 - Page 6 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

FIGURE 2. Gas-Liquid Separator and Expansion Chamber.

FIGURE 3. Quartz Furnace Atomization Cell.
ARSENIC, organo-: METHOD 5022, Issue 3, dated 3 March 2016 - Page 7 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

p-NITROANILINE 5033

H2NC6H4NO MW: 138.14 CAS: 100-01-6 RTECS: BY7000000

METHOD: 5033, Issue 2 EVALUATION: FULL Issue 1: 15 August 1994

Issue 2: 29 February 2016

OSHA: 6 mg/m3 (skin) 

NIOSH: 3 mg/m3 (skin)

PROPERTIES: yellow crystals; MP 146 °C; BP 332 °C; VP 0.02 

Pa (1.5 x 10 -4 mm Hg) @ 20 °C

SYNONYMS: 4-nitrobenzenamine, p-aminonitrobenzene, p-nitrophenylamine 

SAMPLING

SAMPLER: FILTER (0.8 μm mixed cellulose ester 

membrane)

FLOW RATE: 1 - 3 L/min

VOL-MIN: 16 L @ 6 mg/m3

 

 -MAX: 350 L 

SHIPMENT: routine

SAMPLE 

STABILITY: 7 days @ 25 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 3.9 to 12.9 mg/m3

 (90-L samples) [1]

BIAS: -1.7% 

OVERALL

PRECISION (SS�rrrr): 0.054 for range studied [1] 

ACCURACY: ± 11% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: p-nitroaniline 

EXTRACTION: 5 mL isopropanol

MOBILE PHASE: 40% isopropanol/60% hexane isocratic, 

1 mL/min

COLUMN: silica (25-cm x 4.6-mm x 6.4- mm 

stainless steel, US pharmacopeia (USP) 

L18) 

DETECTOR: UV @ 375 nm 

CALIBRATION: solutions of p-nitroaniline in 

isopropanol

RANGE: 100 to 2000 μg per sample [1]

ESTIMATED LOD: 20 μg/sample [1]

PRECISION (SS̅

rr): 0.020 [1]

APPLICABILITY: The working range is 1.1 to 22 mg/m3 for a 90-L air sample. This method is applicable to p-nitroaniline aerosol but 

not for environments where vapor may be present.

INTERFERENCES: None identified

OTHER METHODS: This revises Method S7 [2].
p-NITROANILINE: METHOD 5033, Issue 2, dated 25 February 2016 - Page 2 of 3 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. p-Nitroaniline, 99%.*

2. Isopropanol, distilled in glass.*

3. Hexane, distilled in glass.*

4. Calibration stock solution, 10 mg/mL. 

Dissolve 0.100 g p-nitroaniline in 10 mL 2-

propanol.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: mixed cellulose ester (MCE) 

membrane filter (37-mm) and cellulose 

backup pad (37- mm) in a 2-piece filter 

holder held together by shrinkable band.

2. Personal sampling pump, calibrated, capable 

of operating 8 h at 1 to 3 L/min, with flexible 

connecting tubing.

3. HPLC with UV detector (375 nm), silica 

column (USP L18), injector, and electronic 

integrator (5033-1). 

4. Microliter syringes, various sizes.

5. Volumetric flasks, various sizes.

6. Squat form ointment jars with 

polytetrafluoroethylene (PTFE) gaskets and 

screw caps.

7. Pipet, 5-mL, with pipet bulb.

8. Tweezers.

 

SPECIAL PRECAUTIONS: p-Chlorophenol is toxic by skin absorption, inhalation, or ingestion. It also is a 

strong irritant to tissue and is combustible with a flash point of 121 °C. Work should be performed in 

hood

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Remove the front and rear plugs immediately before sampling and attach sampler to personal 

sampling pump with flexible tubing. Sample at a known flowrate between 1 and 3 L/min and collect 

a total sample of 16 to 350 L of air.

3. Cap the filters, record sample identity and all relevant sample data (duration, ambient temperature 

and pressure).

4. Prepare blank sample filters by handling in the same manner (open, seal, and transport) as samples 

except that no air is sampled through the blanks.

5. Ship in suitable container with blanks. 

SAMPLE PREPARATION:

6. Open the filter holder and using tweezers, transfer each mixed cellulose ester filter to an ointment jar.

7. Add 5 mL isopropanol, cap, and swirl the jar to ensure that the filter is thoroughly wetted.

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards.

a. Dilute aliquots of p-nitroaniline stock solution with isopropanol in volumetric flasks to encompass 

the range of interest.

b. Prepare calibration graph (peak area vs. μg p-nitroaniline per sample).

9. Determine the recovery (R) of the filters at least once for each lot of filters used in the range of 

interest.

a. Using a microliter syringe, spike 6 filters at each of 3 concentration levels with p-nitroaniline in 

isopropanol. Allow the filters to dry overnight for solvent evaporation.

b. Analyze the spiked filters. Prepare graph of μg recovered vs. μg spiked.
p-NITROANILINE: METHOD 5033, Issue 2, dated 25 February 2016 - Page 3 of 3 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

10. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration 

graph and recovery graph are in control.

MEASUREMENT:

11. Set HPLC according to manufacturer's recommendations and to conditions on page 5033-1.

12. Separation is achieved on a silica column using a mobile phase of 40% isopropanol/60% hexane at 

1 mL/min. Inject 20-μL sample aliquot.

NOTE: If peak area is above the linear range of the working standards, dilute with isopropanol and 

apply the appropriate dilution factor in calculations.

13. Measure peak areas.

CALCULATIONS:

14. Determine the mass of p-nitroaniline, μg (corrected for recovery), for the sample (W) and average 

media blank (B).

15. Calculate the concentration of p-nitroaniline, C (mg/m3

), in the air volume sampled, V (L):

CC = WW − BB

VV , mmmm mm3 ⁄

EVALUATION OF METHOD: 

The method is not applicable to environments where p-nitroaniline vapor is present. However, 

experiments indicate that less than 0.04 mg/m3 of p-nitroaniline would exist as a vapor at 40 °C [1]. 

Extraction efficiencies of spiked filters containing 0.278, 0.556, or 1.112 mg of p-nitroaniline were 96.6%, 

94.7%, and 94.6%, respectively [1]. Samples generated to measure the filter's collection efficiency at 

1.5 L/min for concentration of 3.9, 6.75, and 12.88 mg/m3 showed recoveries of 95.9%, 98.8%, and 100.9% 

respectively [1]. The average recovery for 7-day-old samples was 98.6% [1].

REFERENCES:

[1] NIOSH [1977]. Back-up data report for p-Nitroaniline: Method S7. Unpublished. 

[2] NIOSH [1977]. p-Nitroaniline: Method S7. In: Taylor DG, ed. NIOSH manual of analytical methods. 

2nd ed. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

157-B.

[3] NIOSH [1981]. Occupational health guidelines for chemical hazards. Cincinnati, OH: U.S. Department 

of Health and Human Services, Centers for Disease Control, National Institute for Occupational Safety 

and Health, DHHS (NIOSH) Publication No. 81-123. Available as #PB 83-154609 from NTIS.

METHOD REVISED BY:

Charles Neumeister, NIOSH.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

DIESEL PARTICULATE MATTER

(as Elemental Carbon) 5040

C MW: 12.01 CAS: none RTECS: none

METHOD: 5040, Issue 4 EVALUATION: FULL Issue 1: 15 May 1996

Issue 2: 10 March 2016

OSHA: None 

NIOSH: None

PROPERTIES: nonvolatile solid

SYNONYMS: diesel particulate matter, diesel exhaust, diesel soot, diesel emissions

SAMPLING

SAMPLER: FILTER: quartz-fiber, 37-mm or 25-mm; 

size-selective sampler may be required [1]

FLOW RATE: 2 to 4 L/min (typical)

VOL-MIN: 142 L @ 40 μg/m3

 -MAX: 19 m3 (for filter load of ~ 90 μg/cm2

)

SHIPMENT: Routine.

SAMPLE

STABILITY: Stable. 

BLANKS: 2 to 10 field blanks per set

ACCURACY 

RANGE 

STUDIED: 23 to 240 μg/m3 (See also ref. [1].)

BIAS: None (See also ref. [1].)

OVERALL

PRECISION (SS�rrrr): 0.07

ACCURACY: ± 16.7% at 23 μg/m3 (See also ref. [1].)

MEASUREMENT

TECHNIQUE: Thermal-optical analysis; flame ionization

detector (FID)

ANALYTE: Elemental carbon (EC). Total carbon is 

determined, but an EC exposure marker 

was proposed. See [1] for details.

FILTER 

PUNCH SIZE: 1.5 cm2 (or other [1])

CALIBRATION: Methane injection

RANGE: 1 to 105 μg per filter portion (See also [1].)

ESTIMATED LOD: 0.3 μg per filter portion

PRECISION (SS�rr): 0.19 @ 1 μg C, 0.01 @ 10 to 72 μg C

APPLICABILITY: The working range is approximately 6 to 630 μg/m3, with an LOD of ~ 2 μg/m3 for a 960-L air sample collected on 

a 37-mm filter with a 1.5 cm2 punch from the sample filter. If a lower LOD is desired, a larger sample volume and/or 25-mm filter 

may be used (e.g., a 1920-L sample on 25-mm filter gives an LOD of 0.4 μg/m3). The split between organic carbon (OC) and EC may 

be inaccurate if the sample transmittance is too low. The EC loading at which this occurs depends on laser intensity. In general, the 

OC-EC split may be inaccurate when EC loadings are above 20 μg/cm2

. High loadings can give low (and variable) EC results because 

the transmittance remains low and relatively constant until some of the EC is oxidized. The split should be reassigned (prior to EC 

peak) in such cases [2]. An upper EC limit of 800 μg/m3 (90 μg/cm2) can be determined. Low EC loadings (e.g., near the LOD) also 

may require a manual split to improve accuracy [1].

INTERFERENCES: Total carbon (as OC and EC) is determined by the method, but EC was recommended as a measure of workplace 

exposure because OC interferences may be present [1,2]. Cigarette smoke and carbonates ordinarily do not interfere in the EC 

determination. Less than 1% of the carbon in cigarette smoke is elemental. If heavy loadings of carbonate or organic dusts are 

anticipated, a size-selective sampler (impactor and/or cyclone) should be used [1]. For measurement of diesel-source EC in coal 

mines, a cyclone and impactor with a submicrometer cutpoint are required to minimize collection of coal dust. A cyclone and/or 

impactor may be necessary in other workplaces if EC-containing dusts are present.

OTHER METHODS: Other methods for determination of EC and OC have been employed, but these are not equivalent to the 

method described herein. Information on other methods is summarized elsewhere [1]. The method procedures are unchanged 

from the 4th edition supplement to NMAM, but the corresponding diesel guidance chapter was updated for the 5th edition.
DIESEL PARTICULATE MATTER (as Elemental Carbon) METHOD 5040, Issue 4, dated 10 March 2016 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Aqueous solutions of reagent grade (99+%) 

sucrose, 0.1 to 3 mg C per mL solution. Ensure 

filter spike loading range brackets that of 

samples.

2. Ultrapure H20, Type I, or equivalent.

3. UHP helium (99.999%), scrubber also required for 

removal of oxygen.

4. Hydrogen, purified (99.995%), cylinder or 

hydrogen generator source.

5. Ultra Zero air (low hydrocarbon).

6. 10% oxygen in helium balance, both gases UHP, 

certified mix.

7. 5% methane in helium balance, both gases UHP, 

certified mix. 

EQUIPMENT:

1. Sampler: Quartz-fiber filter, precleaned (in low 

temperature asher 2 to 3 h, or muffle furnace for 1 

to 2 h at ~ 800 °C), 37-mm, in a 3-piece cassette 

with filter support (stainless steel screen, cellulose 

pad, or a second quartz filter). Alternative samplers 

may be required in dusty environments. See ref. [1] 

for details.

NOTE 1: High purity, high efficiency, binder- free 

quartz-fiber filters must be used (e.g., Pall 

Gelman Sciences Pallflex Tissuequartz 

2500QAT-UP. Precleaned filters are 

available from several laboratories. Filters 

also can be purchased and cleaned inhouse. Filters should be cleaned in a 

muffle furnace operated at 800-900 °C for 

1- 2 hours. Check (analyze) filters to 

ensure removal of OC contaminants. A 

shorter cleaning period may be effective. 

OC results immediately after cleaning 

should be below 0.1 μg/cm2

. OC vapors 

readily adsorb on to clean filters. Even 

when stored in closed containers, OC 

loadings may range from 0.5 μg/cm2 after 

several weeks.

NOTE 2: Cellulose supports give higher OC blanks 

than screens and quartz filters. Bottom 

quartz filters can be used to correct for 

adsorbed vapor; see ref. [1].

2. Personal sampling pump with flexible tubing.

3. Thermal-optical analyzer; see ref. [1].

4. Metal punch for removal of 1.5 cm2 rectangular 

portion of filter.

NOTE: A smaller portion (e.g., taken with cork 

borer) may be used, but the area must be 

large enough to accommodate the entire 

laser beam (i.e., beam should pass through 

the sample, not around it). The area of the 

portion must be accurately known, and the 

sample must be carefully positioned (the 

filter transmittance will decrease 

dramatically when the sample is properly 

aligned). A filter portion ≥ 0.5 cm2 with 

diameter or width ≤ 1 cm is recommended.

5. Syringe, 10-μL.

6. Aluminum foil.

7. Needle (for lifting filter punch portion).

8. Forceps

9. Volumetric flasks, Class A.

10. Analytical balance. 

 
DIESEL PARTICULATE MATTER (as Elemental Carbon) METHOD 5040, Issue 4, dated 10 March 2016 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SPECIAL PRECAUTIONS: Hydrogen is a flammable gas. Users must be familiar with the proper use of 

flammable and nonflammable gases, cylinders, and regulators. According to the instrument manufacturer, 

the instrument is a Class I Laser Product. This designation means there is no laser radiation exposure 

during normal operation. Weakly scattered laser light is visible during operation, but does not pose a 

hazard to the user. The internal laser source is a Class IIIb product, which poses a possible hazard to the 

eye if viewed directly or from a mirror-like surface (i.e., specular reflections). Class IIIb lasers normally do 

not produce a hazardous diffuse reflection. Repairs to the optical system, and other repairs requiring 

removal of the instrument housing, should be performed only by a qualified service technician.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

NOTE: Both open and closed-faced cassettes have been used. Both configurations generally give even 

deposits. At higher flow rates (e.g., 4 L/min), small spots occasionally have been observed in the 

center of the filters when closed-faced cassettes are used. This material likely consisted of 

impacted diesel agglomerates and/or non-diesel particulate matter. EC results for multiple 

portions of the filters were in good agreement, so the spots had little analytical impact. Other

samplers also can be used (see ref. [1]) provided an even deposit of diesel particulate results. An 

even deposit is necessary because the sample portion analyzed must be representative of the 

entire deposit. If the deposit is not homogeneous, the entire sample must be analyzed. An 

impactor/cyclone may be needed in some cases [1]. 

2. Attach sampler outlet to personal sampling pump with flexible tubing.

3. Sample at an accurately known flow rate. Typical rates are 2-4 L/min (note: Lower flows (e.g., 1 L/min) 

have been used in mines to prevent overloading).

4. After sampling, replace top piece of cassette, if removed, and pack securely for shipment to laboratory. 

NOTE: Diesel particulate samples from occupational settings generally do not require refrigerated 

shipment unless there is potential for exposure to elevated temperatures (that is, well above 

collection temperature). Filter samples normally are stable under laboratory conditions. Some

OC loss may occur over time if samples contain OC from other sources (for example, cigarette 

smoke). Sorption of OC vapor after sample collection has not occurred, even with samples 

having high (e.g., 80%) EC content.

SAMPLE PREPARATION:

5. Place sample filter on a freshly cleaned aluminum foil surface. Isopropyl alcohol or acetone can be 

used to clean the foil. Allow residual solvent to vaporize from the surface prior to use. Punch out a 

representative portion of the filter. Take care not to disturb deposited material and avoid hand contact 

with sample. A needle inserted at an angle is useful for removal of the filter portion from the punch 

body. Newer instruments have an externally mounted bracket to support the quartz sample holder 

while the previous sample is removed and a new one is loaded. Through a hole in the side of the 

standard punch, a needle can be used to push the filter portion from the punch onto the sample 

holder. Alternative approaches also can be used, depending on the user’s preference, as long as 

contamination is avoided.

CALIBRATION AND QUALITY CONTROL:

6. Analyze at least one replicate sample. For sets of up to 50 samples, replicate 10 % of the samples. For 

sets over 50 samples, replicate 5 % of the samples. If a filter deposit appears uneven (this should not 

be the case if the cassette is sealed properly), take a second portion (step 5) for analysis to check 

evenness of deposition.

NOTE: Precision of replicate analyses of a filter is usually better than 5% (1 to 3% is typical).

7. Analyze three quality control blind spikes and three analyst spikes to ensure that instrument 

calibration is in control. Prepare spike as follows:
DIESEL PARTICULATE MATTER (as Elemental Carbon) METHOD 5040, Issue 4, dated 10 March 2016 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

a. With 10-μL (or other) syringe, apply an aliquot of OC standard solution directly to filter portion 

taken (step 5) from a precleaned filter. For best results, the precleaned filter punch should be 

cleaned again in the sample oven prior to application of the aliquot.

NOTE: With small aliquots (e.g., ≤ 10 μL), disperse standard solution over one end of filter portion 

to ensure standard is in laser beam. To prevent possible solution loss to surface, hold the 

portion off the surface. Larger volumes can easily penetrate to the underside of the filter 

portion.

b. Allow water to evaporate and analyze spikes with samples and blanks (steps 9 and 10).

NOTE: A pronounced decrease in filter transmittance during the first temperature step of the 

analysis indicates water loss. Allow portions to dry longer if this occurs. Spiked punches also 

can be dried in the oven, if desired. For quick drying, the ‘clean oven’ command on the 

menu can be selected and canceled after about 4 seconds. The time allowed may depend 

on instrument, but oven temperatures should be below 100 °C to avoid boiling the 

solution. This approach is convenient and prevents potential adsorption of organic vapors 

in laboratory air. Some software versions may not have this feature.

8. Determine instrument blank (results of analysis with freshly cleaned filter portion) for each sample set. 

MEASUREMENT:

9. Adjust analyzer settings according to manufacturer's recommendations (see instrument operation 

manual and background information in ref. [1]). Place sample portion into sample oven.

NOTE: Forms of carbon that are difficult to oxidize (e.g., graphite) may require a longer period and 

higher temperature during the oxidative mode to ensure that all EC is removed (the EC peak 

should never merge with the calibration peak.) Adjust time and temperature accordingly. A 

maximum temperature above 940 °C should not be required.

10. Determine EC (and OC) mass, μg. Analyzer results are reported in units μg/cm2 of C. The reported

values are normally based on a sample portion of about 1.5 cm2

, which is the area of the standard 

punch provided by the manufacturer. If the portion area used differs from the value entered in the 

ocecpar.txt file, multiply the result by 1.5 (or value in ocecpar.txt file) and divide the product by the 

actual area analyzed to obtain the area-corrected result (i.e., reported result x 1.5 /portion area = 

corrected result in μg/cm2

). This is most easily done in the data spreadsheet. Alternatively, the correct 

results will be obtained with the data calculation program if the portion area is entered in the 

parameter file (ocecpar.txt), but this approach is error prone and tedious when punches of different 

areas are used for a sample set because correct results will not be obtained for all punch sizes. 

Updated software avoids this potential problem by prompting entry of the sample punch area prior to 

the analysis.

CALCULATIONS

11. Multiply the reported (or area-corrected) EC result (μg/cm2

) by filter deposit area, cm2, (typically 8.5 cm2

for a 37-mm filter) to calculate total mass, μg, of EC on each filter sample (WEC). Do the same for the 

blanks and calculate the mass found in the average field blank (Wb). The mass of OC is calculated 

similarly, but the mean OC field blank may underestimate the amount of OC contributed by adsorbed 

vapor. A quartz filter placed beneath the sample filter can provide a better estimate of the adsorbed 

OC [1].

12. Calculate the EC concentration (CEC) in the air volume sampled, V (L):

CCEEEE = WWEE − WWbb

VV , mmmm/mm3
DIESEL PARTICULATE MATTER (as Elemental Carbon) METHOD 5040, Issue 4, dated 10 March 2016 - Page 5 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

EVALUATION OF METHOD: 

Details on the evaluation of this method are provided in an NMAM guidance chapter on diesel [1]. The 

chapter includes a summary of interlaboratory comparison work conducted since the initial publication of 

the method. Background information and guidance on method use, including sampling requirements, also 

are provided. In general industry, 37-mm or 25-mm cassettes are normally suitable for air sampling, but 

there are exceptions. A cyclone in series with an impactor having a submicrometer cutpoint must be used 

in coal mines, and the Mine Safety and Health Administration (MSHA) has recommended use of a cycloneimpactor sampler in metal and nonmetal mines [3]. The impactor is commercially available through SKC, 

Inc. A size-selective sampler (either impactor and/or cyclone) also may be required in other dusty 

environments [1], particularly if the dust is carbonaceous. If a sample contains carbonate, the carbonate 

carbon (CC) will be quantified as OC. A carbonate-subtracted result can be obtained through acidification 

of the sample portion or through separate integration of the carbonate peak [1] (Note: Trona and other 

compounds containing sodium can etch the quartz oven wall during the analysis. Avoid spillage in the 

sample oven and frequent analysis of these materials.) These procedures are described in a Chapter of this 

Manual [1]. The thermal-optical method is applicable to nonvolatile carbon species (i.e., particulate OC, CC 

and EC). The method is not appropriate for volatile or semivolatiles, which require sorbents for efficient 

collection.

REFERENCES:

[1] NIOSH [2016]. Chapter DE: Monitoring of diesel particulate exhaust in the workplace. In: O’Connor 

PF, Ashley K, eds. NIOSH manual of analytical methods. 5th ed. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2014-151.

[2] Birch ME, Cary RA [1996]. Elemental carbon-based method for monitoring occupational exposures to 

particulate diesel exhaust. Aerosol Sci Tech 25:221-241.

[3] 66 Fed. Reg. 13 [2001]. Department of Labor, Mine Safety and Health Administration: Diesel 

particulate matter exposure of underground metal and nonmetal miners; final rule. (To be codified 

at 30 CFR Part 57).

METHOD REVISED BY:

M. Eileen Birch, Ph.D., NIOSH

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CARBON BLACK 5100

 C MW: 12.01 CAS: 1333-86-4 RTECS: FF5800000

METHOD: 5100, Issue 1 EVALUATION: FULL Issue 1: 30 April 2015

OSHA: 3.5 mg/m3

[1]

NIOSH: 3.5 mg/m3

 (in presence of PAHs: carcinogen/PAHs to 

0.1 mg/m3

, cyclohexane extractable fraction) [2]

 For other OELs 

and guidelines: See references [3,4]

PROPERTIES: Solid; may contain polynuclear aromatic hydrocarbons (PAHs)

SYNONYMS: Acetylene black; amorphous carbon; furnace black; lamp black

SAMPLING

SAMPLER: INTERNAL CAPSULE, (tared 37-mm, 2- to 5-μm 

PVC filter melded to PVC housing) in 37-mm 

2-piece cassette

FLOW RATE: 1 to 2 L/min

VOL-MIN: 75 L @ 3.5 mg/m3

-MAX: 1400 L @ 3.5 mg/m3

SHIPMENT: Routine

SAMPLE 

STABILITY: 28 days minimum

BLANKS: Minimum of 2 field blanks per batch

ACCURACY

RANGE STUDIED: 0.1 to 4 mg/sample

BIAS: 0.058 [5]

OVERALL 

PRECISION (ŜrT): 0.059 [5]

ACCURACY: ± 15.5%

MEASUREMENT

 TECHNIQUE: GRAVIMETRIC (INTERNAL CAPSULE WEIGHT)

ANALYTE: Carbon black

BALANCE: 0.001 mg sensitivity; use same balance 

before and after sample collection

CALIBRATION: National Institute of Standards and 

Technology (NIST), Class S-1.1 weights or 

ASTM Class 1 weights

RANGE: 0.25 to 5 mg per sample

ESTIMATED 

LOD: 0.075 mg per sample

PRECISION ( ): 0.031 at 2 mg per sample [5

APPLICABILITY: The working range is 1.5 to 25 mg/m3 for a 200-L air sample. This method is not applicable for the determination of “cyclohexane-solubles” [3]. The method is nonspecific; information on any other particulate materials that may be present 

should be assessed.

INTERFERENCES: Moisture and static electricity can affect gravimetric measurements. Humidity control and minimization of 

static effects are addressed in this procedure. The presence of co-sampled airborne particulate material is a positive interference 

since this is a gravimetric method.

OTHER METHODS: This method is preferred over Method 5000, Issue 2 [6], and is similar to Method 0501 for particulates not 

otherwise regulated [8]. OSHA method PV2121 describes a similar procedure (but for respirable sampling) using an alternative 

sampler design [9].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CARBON BLACK: METHOD 5100, Issue 1, dated 30 April 2015 - Page 2 of 4

EQUIPMENT:

1. Sampler: Internal capsule, 37-mm PVC, 2- to 5-μm pore size membrane or equivalent hydrophobic 

filter attached to PVC housing and supporting pad in 37-mm 2-piece cassette filter holder

NOTE: The cassettes should be fabricated so as to ensure complete sealing of the internal capsule 

after sample collection.

2. Personal sampling pump, 1 to 2 L/min, with flexible connecting tubing

3. Microbalance capable of weighing to ±0.001 mg

4. Static neutralizer; e.g. 210Po; replace no more than nine months after production date

5. Tool for handling internal capsules, e.g., forceps (preferably plastic)

6. Environmental chamber or room for balance (e.g. 20 ± 1 °C and 50 ± 5% RH)

SPECIAL PRECAUTIONS:  Carbon black containing polynuclear aromatic hydrocarbons (cyclohexane - 

extractable materials) in excess of 0.1% (w/w) should be treated as a suspect carcinogen [3].

PREPARATION OF FILTER CAPSULES BEFORE SAMPLING:

1. Equilibrate the PVC filter capsules in an environmentally controlled weighing area or chamber for at 

least 24 h.

NOTE: An environmentally controlled chamber is desirable, but not required.

2. Place backup pads in filter cassette bottom sections.

3. Weigh the filter capsules in an environmentally controlled area or chamber. Record the capsule tare 

weight, W1 (mg).

a. Zero the balance before each weighing.

b. Handle the filter capsule with forceps. Pass the capsule over an antistatic radiation source. Repeat 

this step if the capsule does not release easily from the forceps or if it attracts balance pan. Static 

electricity can cause erroneous weight readings.

4. Assemble the filter capsules in the filter cassettes and close firmly so that leakage around the internal 

capsule will not occur. Place a plug in each opening of the filter cassette. Place a cellulose shrink band 

around the filter cassette, allow to dry, and label the cassette with indelible ink.

 SAMPLING:

5. Calibrate each personal sampling pump with a representative sampler in line.

6. Sample at 1 to 2 L/min for a total sample volume of 75 to 1400 L. Do not exceed a filter capsule loading 

of approximately 5 mg total dust. Take two to four replicate samples for each batch of field samples for 

quality assurance on the sampling procedure.

SAMPLE PREPARATION:

7. Wipe dust from the external surface of the filter cassette with a moist paper towelette to minimize 

contamination. Discard the towelette.

8. Remove the top and bottom plugs from the filter cassette. Equilibrate for at least 24 h in the balance 

room.

9. Using forceps, open the cassette and remove the internal capsule gently to avoid loss of dust or 

damage to the capsule.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CARBON BLACK: METHOD 5100, Issue 1, dated 30 April 2015 - Page 3 of 4

CALIBRATION AND QUALITY CONTROL:

10. Zero the microbalance before all weighings. Use the same microbalance for weighing filter

capsules before and after sample collection. Calibrate the balance with National Institute of

Standards and Technology Class S-1.1 or ASTM Class 1 weights.

11. Process laboratory blanks, spiked QC samples and field blanks at a minimum frequency of 1 per

20 field samples. Internal capsules used for QC samples should come from the same lot. Spiked

QC samples, loaded with 0.25 to 4 mg of material per internal capsule, should be prepared using

weight-stable material such as Arizona Road Dust [10].

MEASUREMENT:

12. Weigh each capsule, including field blanks. Record the post-sampling weight, W2

 (mg). Record

anything remarkable about a capsule (e.g., overloading, leakage, wet, torn, etc.).

CALCULATIONS:

13. Calculate the concentration, C (mg/m3

), of carbon black in the air volume sampled, V (L):

  , mg/m

where: W1 = tare weight of capsule before sampling (mg)

 W2

 = post-sampling weight of sample-containing capsule (mg)

 B1

 = mean tare weight of blank capsules (mg)

 B2

 = mean post-sampling weight of blank capsules (mg)

EVALUATION OF METHOD:

Lab testing was carried out using blank internal capsules and with capsules spiked with 0.1 – 4 mg of 

NIST SRM 1648 (Urban Particulate Matter) and Arizona Road Dust (Air Cleaner Test Dust) [5]. Precision 

and accuracy data are given on page 5100-1. Weight stability over 28 days was verified for both blanks 

and spiked capsules [5]. Independent laboratory testing on blanks and field samples have verified 

long-term weight stability as well as sampling and analysis uncertainty estimates [5].

REFERENCES:

[1] CFR. 29 CFR Part 1910.1000. Code of Federal Regulations. Washington, DC: U.S. Government

Printing Office, Office of the Federal Register.

[2] NIOSH [2005]. NIOSH pocket guide to chemical hazards. Barsan ME, ed. Cincinnati OH: U.S.

Department of Health and Human Services, Centers for Disease Control and Prevention, National

Institute for Occupational Safety and Health. DHHS (NIOSH) Publication no. 2005-149.

[3] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio:

American Conference of Governmental Industrial Hygienists. [www.acgih.org]. [Date accessed:

February, 2015.]

[4] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2014]. GESTISDatabase on hazardous substances. (German Social Accident Insurance). Sankt Augustin,

Germany: IFA. https://www.dguv.de/ifa/index-2.jsp [Date accessed: February, 2015.] 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CARBON BLACK: METHOD 5100, Issue 1, dated 30 April 2015 - Page 4 of 4

[5] NIOSH [2014]. Backup Data Report-NIOSH 0501 and 5100. By O’Connor SP, O’Connor PF, Feng HA, 

Ashley K. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control 

and Prevention, National Institute for Occupational Safety and Health. [http://www.cdc.gov/niosh/

docs/2003-154/pdfs/bud/1453_bud.pdf]. 

[6] NIOSH [1994]. Carbon black: Method 5000. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical 

methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) 

Publication No. 94-113 [www.cdc.gov/niosh/nmam/].

[7] NIOSH [1994]. Particulates not otherwise regulated, total: Method 0500. In: Eller PM, Cassinelli ME, eds. 

NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and 

Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/]. 

[8] NIOSH [2015]. Particulates not otherwise regulated, total: Method 0501. In: Ashley KE, O’Connor PF, 

eds. NIOSH manual of analytical methods. 5th ed. Cincinnati, OH: U.S. Department of Health and 

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 2014-151. [www.cdc.gov/niosh/nmam/].

[9] OSHA [2003]. Gravimetric determination: PV2121. In: Monitoring methods used by OSHA. Salt Lake 

City, UT: U.S. Department of Labor, Occupational Safety and Health Administration. [https://www.

osha.gov/dts/sltc/methods/partial/pv2121/pv2121.html] (Date accessed: February, 2015).

[10] O’Connor S, O’Connor PF, Feng HA, Ashley K [2014]. Gravimetric analysis of particulate matter using air 

samplers housing internal filtration capsules. Gefahrsstoffe Reinh Luft 75: 403-410.

METHOD WRITTEN BY: Kevin Ashley, Ph.D., NIOSH/DART

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for 

the content of these websites. All web addresses referenced in this document were accessible as of the 

publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

ORGANOTIN COMPOUNDS (as Sn) 5504

Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 5504, Issue 3 EVALUATION: FULL Issue 1: 15 August 1987

Issue 3: 4 February 2016

OSHA: 0.1 mg/m3

NIOSH: 0.1 mg/m3 (skin)

 

PROPERTIES: Table 1 

SYNONYMS: Table 1

SAMPLING

SAMPLER: FILTER + SORBENT 

(glass fiber + XAD-2, 80 mg/40 mg)

FLOW RATE: 1 to 1.5 L/min

VOL-MIN: 50 L

 -MAX: 500 L

SHIPMENT: ship assembled sampler in dry ice

SAMPLE

STABILITY: stable 7 days @ 0 °C [1]

FIELD

BLANKS: 2 to 10 field blanks per set

MEDIA

BLANKS: 12 per sample set

ACCURACY

RANGE 

STUDIED: 0.07 to 0.2 mg/m3 [1] (300-L samples)

BIAS: see EVALUATION OF METHOD

OVERALL

PRECISION (SS�rrrr): 0.07 to 0.10 [1]

ACCURACY: see EVALUATION OF METHOD

MEASUREMENT

TECHNIQUE: HPLC/ATOMIC ABSORPTION, GRAPHITE 

FURNACE

ANALYTE: tin

DESORPTION: 10 mL 0.1% acetic acid/H3CN; ultrasonic, 

30 min

SEPARATION: HPLC (C18 or cation exchange column)

GRAPHITE 

FURNACE: dry 30 sec @ 80 °C; atomize 5 sec @ 

2750 °C (gas interrupt mode)

INJECTION 

 VOLUME: 20 μL

WAVELENGTH: 286.3 nm, with background correction

CALIBRATION: standard solutions of organotin 

compounds in acetic acid/CH3CN

RANGE: 5 to 50 μg Sn per sample

ESTIMATED LOD: 1 μg Sn per sample [1]

PRECISION (SS�rr): 0.07 to 0.08 [1]

APPLICABILITY: The working range is 0.015 to 1 mg/m3 (as Sn) for a 300-L air sample. The method was validated using TeBT, TBTC, 

TCHH, and BuIOMA as surrogates for the most important classes of organotin compounds [1]. If speciation of organotin 

compounds is not required and if inorganic tin compounds are absent, the HPLC separation may be deleted. Use special care to 

avoid losses of TeBT and other volatile tetra-substituted organotin compounds

INTERFERENCES: Organotin compounds not separated chromatographically will mutually interfere. Other compounds with 

similar retention times will not interfere unless they contain tin.

OTHER METHODS: This replaces the colorimetric criteria document method [2]. 
ORGANOTIN COMPOUNDS (as Sn): METHOD 5504, Issue 3, dated 4 February 2016 - Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Zirconium acetate oxide, reagent grade.

2. Ammonium acetate, reagent grade.

3. Diammonium citrate, reagent grade.

4. Acetonitrile, chromatographic grade.

5. Deionized water.

6. Acetic acid, glacial, reagent grade.

7. Methanol, chromatographic grade.

8. Acetic acid, 0.1% (v/v) in acetonitrile.

9. Acetate buffer solution (v/v) 70% methanol, 

27% deionized water, 3% aqueous 1 M 

ammonium acetate.

10. Citrate buffer solution (v/v) 70% methanol, 

26.8% deionized water, 3% aqueous 1 M 

diammonium citrate in 0.2% (v/v) glacial 

acetic acid.

11. Organotin standard solutions, 1000 μg/mL 

(as Sn), prepared from pure organotin 

compounds in 0.1% (v/v) acetic acid in 

acetonitrile.

12. Calibration stock solution, 10 μg/mL (as Sn). 

Prepare a standard mixture of the organotin 

compounds of interest. Pipet 0.1 mL of each 

organotin standard solution into a 10-mL 

volumetric flask. Dilute to the mark with 

0.1% (v/v) acetic acid in acetonitrile. Prepare 

fresh daily.

 

EQUIPMENT:

1. Sampler: glass fiber filter, 37-mm 

(commercially available) in cassette filter 

holder followed by XAD-2 sorbent tube, 80 

mg front section/40 mg back section 

separated and retained by silanized glass 

wool (commercially available).

2. Personal sampling pump, 1 to 1.5 L/min, with 

flexible connecting tubing.

3. Shipping container, refrigerated, with dry ice.

4. High performance liquid chromatograph 

(HPLC), interfaced with autoinjection system 

(Fig. 1), with binary solvent capability, solvent 

gradient capability and columns:

a. non-tetraorganotin species: cation 

exchange column.

b. tetraorganotin compounds: C18 column 

5. Atomic absorption spectrophotometer (AAS) 

having recorder output proportional to 

absorbance units, graphite furnace accessory 

(pyrolytic with Zr coating, coated L'vov 

platform may be required; see APPENDIX), 

sample autoinjection system with moving 

sample tube rack or carrousel (Fig. 1), 

automatic micropipettor for accurately 

injecting 20-μL sample aliquots into graphite 

furnace, background correction (e.g., D2 or H2

lamp) capability, and tin electrodeless 

discharge lamp or hollow cathode lamp.

6. Bath, ultrasonic.

7. Volumetric flasks, 10-mL.

8. Syringe, 20-μL, readable to 0.5 μL.

9. Beakers, Phillips, 125-mL.

10. Pipets, 5- and 10-mL; 10- and 100-μL.

11. Oven or muffle furnace, 200 ̊C.

12. Plastic film.

 

SPECIAL PRECAUTIONS: None.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break the ends of the sorbent tube immediately before sampling and connect to the filter with a short 

piece of tubing. Attach sampler to personal sampling pump with flexible tubing.

3. Sample at an accurately known flow rate between 1 and 1.5 L/min for a total sample size of 50 to 500 L.

4. Cap the samplers. Pack securely for shipment in dry ice.

NOTE: The cassette and sorbent tube should remain connected for storage. Store samples at less than 

0 °C. Analyze within seven days after collection. 
ORGANOTIN COMPOUNDS (as Sn): METHOD 5504, Issue 3, dated 4 February 2016 - Page 3 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION:

5. Open the cassette filter holder. With tweezers, carefully transfer the filter to a 125-mL beaker.

6. Place the front sorbent section and front glass wool plug into a second beaker. Place the back sorbent 

section and remaining glass wool plugs into a third beaker.

7. Pipet 10.0 mL acetonitrile and 10 μL acetic acid into each beaker. Cover with plastic film.

8. Agitate beakers in ultrasonic bath for 30 min.

CALIBRATION AND QUALITY CONTROL:

9. Calibrate daily with at least six working standards.

a. Add known amounts of calibration stock solution to 0.1% (v/v) acetic acid in acetonitrile in 10-mL 

volumetric flasks and dilute to the mark. Use serial dilutions as needed to obtain concentrations of 

each organotin compound in the range 0.1 to 5 μg/mL (as Sn).

b. Analyze with samples and blanks (steps 12 through 15), alternating samples and standards with 

similar responses.

c. Prepare calibration graphs (peak height vs. μg Sn) for each organotin compound.

10. Determine recovery (R) in the range of interest. Prepare three samplers at each of three levels plus 

three media blanks.

a. Remove and discard back sorbent section of a media blank sampler.

b. Inject a known amount (2 to 20 μL) of organotin standard solution directly onto front sorbent 

section and a separate aliquot onto the filter, with a microliter syringe.

c. Cap the sampler. Allow to stand overnight.

d. Desorb (steps 5 through 8) and analyze with working standards (steps 12 through 15).

e. Prepare a graph of R vs. μg Sn recovered for each organotin compound.

11. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration 

graphs and recovery graphs are in control. 

MEASUREMENT:

12. Set the AAS and graphite furnace according to manufacturer's recommendations and to conditions on 

page 5504-1. Adjust the sample injection/dry/atomize cycle so that it occurs exactly once every 60 sec. 

13. Operate the HPLC according to manufacturer's recommendations and the following conditions:

a. For non-tetraorganotin compounds:

(1) Column: strong cation exchange (EQUIPMENT, 4a).

(2) Flush column with 50 to 60 mL acetate buffer prior to sample injection.

(3) Eluent: flowrate = 2 mL/min

(4) Eluent Gradient: 

Time (min) % Acetate Buffer % Citrate Buffer

0-15 100 0 

15-18 100-0 0-100

18-40 0 100

(5) After the chromatogram is complete, re-equilibrate the HPLC system to initial conditions by 

pumping acetate buffer through the column for 15 min.

(6) Inject 100-μL sample aliquot.
ORGANOTIN COMPOUNDS (as Sn): METHOD 5504, Issue 3, dated 4 February 2016 - Page 4 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

b. For tetraorganotin compounds:

(1) Column: C18. 

(2) Flush column with 50 to 60 mL of 100% acetonitrile prior to sample injection.

(3) Eluent: flowrate = 2 mL/min, isocratic, 100% acetonitrile.

(4) Inject a 100-μL aliquot of sample solution.

14. Collect HPLC eluent at the rate of 1 fraction (2 mL) per minute in AAS autosampler (Fig. 1).

NOTE: In this example system, the column effluent is fed directly into one of the sample cups. After 1 

min, the sample holder rotates and the eluted sample is in a position to be sampled for AA 

measurement. The furnace injection device withdraws a portion of the sample and places it 

in the furnace while the next sample is being eluted and collected.

15. Measure total AAS peak area for each organotin compound.

NOTE 1: The recorder output will consist of AA peaks which form a chromatographic peak for each 

organotin compound if a line is drawn connecting the highest points of the AA peaks. 

Determine total absorbance for a species by the sum of the absorbencies of the 

corresponding AA peaks.

NOTE 2: The characteristics of the graphite tube can influence the results drastically. Pay careful 

attention to the response of the standards and replace the graphite tube if erratic results and 

non-reproducible peak areas occur. 

CALCULATIONS: 

16. Read the mass, μg Sn (corrected for R), for each organotin compound found on the filter (W) and front 

sorbent (Wf) and back sorbent (Wb) sections, and on the average media blanks [filter (B) and front 

sorbent (Bf) and back sorbent (Bb) sections] from the calibration graphs.

17. Calculate concentration, C (mg/m3

), for each organotin compound as Sn in air as the sum of the 

particulate concentration and the vapor concentration in the air volume sampled, V (L):

CC = WW − BB + WWff + WWbb − BBff − BBbb

VV , mmmm mm3 ⁄

NOTE: Wf and Wb will include any analyte originally collected on the filter as particulate, then 

volatilized during sampling or storage.

EVALUATION OF METHOD:

The method was validated with tetrabutyltin (TeBT), tributyltin chloride (TBTC), tricyclohexyltin hydroxide 

(TCHH), and dibutyltin bis(isooctylmercaptoacetate) (BuIOMA) [1]. The working ranges, validation ranges, 

and estimated linear working ranges (as tin) for 300-L air samples of these organotin species at an 

atmospheric temperature and pressure of 20 °C and 756 mm Hg, respectively, appear below:

Species Validation 

Range 

(mg/m3) 

Estimated linear 

Working Range 

(mg/m3) 

Estimated 

linear Working 

Range (μg/mL)

Measurement 

Precision (%)

Bias 

(%·)

Overall 

Precision 

(±%)

Accuracy

TeBT 0.027-0.112 0.02-0.17 0.05-5.0 8.1 +1.8 10.0 21.4

TBTC 0.042-0.191 0.01-0.34 0.3-10 5.9 ·6.7 9.9 26.1

TCHH 0.071-0.218 0.01-0.34 0.3-10 6.9 ·2.3 7.1 16.2

BulOMa 0.070-0.220 0.01-0.34 0.3-10 7.7 ·1.2 7.4 15.7

The methods were developed using a Partisil-10 strong cation exchange column and a Solvecon 

precolumn kit (Whatman, Inc.) for the analysis of the non-tetraorganotin compounds. For the 

tetraorganotin compounds, a LiChrosorb RP-18 HPLC column (Whatman, Inc.) was used.
ORGANOTIN COMPOUNDS (as Sn): METHOD 5504, Issue 3, dated 4 February 2016 - Page 5 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] Gutknecht WF, Grohse PM, Homzak CA, Tronzo C, Ranade MH, and Damle A [1982]. Development of 

a method for the sampling and analysis of organotin compounds. Research Triangle Park, NC: 

Research Triangle Institute. NIOSH contract no. 210-80-0066. Available as order no. PB83-180737 

from NTIS.

[2] NIOSH [1976]. NIOSH criteria for a recommended standard: occupational exposure to organotin 

compounds. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

115. 

METHOD REVISED BY:

Eugene R. Kennedy, Ph.D., NIOSH
ORGANOTIN COMPOUNDS (as Sn): METHOD 5504, Issue 3, dated 4 February 2016 - Page 6 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPENDIX: Preparation of Zr-Coated Graphite Furnace Tube and Platform

1. Zirconium coating: soak pyrolytic graphite tubes and platforms overnight in 4.5% (w/v) zirconium

acetate oxide solution; then dry in a muffle furnace for 2h at 200 °C.

2. Pyrolytic zirconium-coated graphite furnace tubes are acceptable for measurements of all organotin

compounds. However, a pyrolytic zirconium-coated graphite platform (e.g., L'vov platform) is

recommended for better precision at low levels and improved atomization response for volatile species 

such as TeBT. Platforms may be purchased commercially or prepared from pyrolytic graphite tubes, as 

shown in Fig. 2. Special care must be exercised during placement of the platform in the tube (Fig. 3) 

and in optical alignment

Table 1. Formulae and Physical Properties 

Compound/ RTECS Formula M.W. % Sn Synonym

Dibutyltinbis(isooctyl

mercaptoacetate)

WH6719000

C28H56O4S2Sn 639.57 18.6 BuIOMA;

CAS #25168-24-5 

Tetrabutyltin WH8605000 C16H36Sn 347.16 34.2 Stannane,tetrabutyl-; TeBT;

CAS #1461-25-2 

Tributyltinchloride

WH6820000

C12H27ClSn 325.49 36.5 Stannane, chlorotributyl-;

TBTC; CAS #1461-22-9 

Tricyclohexyltin

hydroxide

WH8750000

C18H34OSn 385.16 30.8 Stannane,

tricyclohexylhydroxy-; 

TCHH; CAS #13121-70-5 

Figure 1 HPLC/AAS interface system Figure 2 Construction of a laboratory made 

platform
ORGANOTIN COMPOUNDS (as Sn): METHOD 5504, Issue 3, dated 4 February 2016 - Page 7 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 3 Placement of commercial and “laboratorymade” graphite L’vov platforms in furnace tubes.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

BENZIDINE and

3,3’-DICHLOROBENZIDINE 5509

(1) (C6H4NH2)2 MW: (1) 184.23 CAS: (1) 92-87-5 RTECS: (1) DC9625000

(2) (C6H3ClNH2)2 (2) 253.13 (2) 91-94-1 (2) DD0525000

METHOD: 5509, Issue 3 EVALUATION: FULL Issue 1: 15 May 1989 

Issue 3: 29 February 2016

OSHA: lowest feasible; carcinogen (29 CFR 1910.1010)

NIOSH: lowest feasible; carcinogen (29 CFR 1910.1010)

PROPERTIES: (1)solid; MP 127 °C; BP 400 °C

(2) solid; MP 132 °C

SYNONYMS: (1) [1,1'-biphenyl]-4,4'-diamine; p-diaminodiphenyl and (2) 3,3’-Dichloro[1,1’-biphenyl]-4-4’-diamine

SAMPLING

SAMPLER: FILTER (13-mm glass fiber) 

FLOW RATE: 0.2 L/min

VOL-MIN: 20 L @ 10 μg/m3

 -MAX: 100 L 

SHIPMENT: ship benzidine samples in dry ice

SAMPLE 

STABILITY: (1) 11 days @ 15 °C [1]

(2) 12 days @ 23 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: (1) 21 to 63 μg/m3 [1]

(2) 20 to 130 μg/m3 [1]

BIAS: (1) -3%

(2) -4.0%

OVERALL

PRECISION (SS�rrrr): 0.07 [1] 

ACCURACY: (1) ± 13.7% 

(2) ± 15.2%

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: benzidine or 3,3'-dichlorobenzidine 

DESORPTION: 0.5 mL 0.17% (v/v) trimethylamine in 

methanol; stand 60 min

INJECTION

VOLUME: (1) 10 μL 

(2) 15 μL

MOBILE PHASE: (1) 60% methanol/40% water 

(2) 70% acetonitrile/30% water

FLOW RATE: 1.5 mL/min; ambient temperature

COLUMN: C18 US Pharmacopeia (USP) L1, 10 μm 

particles, 4-mm ID by 30-cm long

DETECTOR: UV @ 254 nm 

CALIBRATION: solutions of analyte(s) in eluent

RANGE: 0.2 to 7 μg/sample 

ESTIMATED LOD: 0.05 μg/sample [1]

PRECISION (SS̅

rr): ≤0.07 [1] 

APPLICABILITY: The working range for benzidine or 3,3'-dichlorobenzidine is 4 to 200 μg/m3 for a 50-L air sample. Benzidinium 

sulfate and 3,3'-dichlorobenzidine dihydrochloride will be collected and converted to benzidine and 3,3'-dichlorobenzidine, 

respectively, during sample preparation.

INTERFERENCES: Aniline interferes in the determination of benzidine but may be resolved [2]. 4,4'-Methylenebis(2-chloroaniline) 

interferes in the determination of 3,3'-dichlorobenzidine [1]. A number of compounds were shown not to interfere [1, 2] (see step 

12, NOTE 2).

OTHER METHODS: This combines and replaces P&CAM 243 and P&CAM 246 [3].
BENZIDINE and 3,3’-DICHLOROBENZIDINE: METHOD 5509, Issue 3, dated 29 February 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Methanol, HPLC grade.

2. Acetonitrile, HPLC grade.

3. Triethylamine.

4. Water, distilled, deionized.

5. Benzidine.*

6. 3,3'-Dichlorobenzidine.*

7. Eluent: 0.17% (v/v) triethylamine in methanol. 

Dilute 170 μL triethylamine to 100 mL with 

methanol.

8. Calibration stock solution, 0.5 μg/μL. Dissolve 50 

mg analyte in 100 mL eluent.

9. Recovery (R) stock solution, 0.5 μg/μL. Dissolve 50 

mg analyte in 100 mL methanol.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: 13-mm, Type AE, glass fiber filter in

a 13-mm filter holder.

2. Personal sampling pump, 0.2 L/min, with

flexible connecting tubing.

3. High-performance liquid chromatograph, UV

detector,integratorandcolumn(page5509-1).

4. Test tubes, 1-mL, with polyethylene stoppers.

5. Syringes, glass, 10- and 25-μL, readable to

0.1 μL.

6. Pipets, delivery, 0.5- and 5-mL, graduated in

0.1 mL.

7. Flasks, volumetric, 10- and 100-mL.

8. Centrifuge.

9. Test tube shaker, vortex type.

SPECIAL PRECAUTIONS: Benzidine is a recognized human carcinogen and can be absorbed through the 

skin [4,5,6]. 3,3'-Dichlorobenzidine is a carcinogen [4,6]. Take appropriate precautions to avoid personal and 

area contamination.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at 0.2 L/min for a total sample size of 20 to 100 L.

3. Cap the sampler.

4. Ship and store samples at -15 °C if benzidine may be present.

NOTE: Samples may be stored at room temperature if only 3,3'-dichlorobenzidine is present.

SAMPLE PREPARATION:

5. Place glass fiber filter in a test tube.

6. Add 0.5 mL eluent to each test tube. Seal each test tube and shake them on a test tube shaker.

7. Allow samples to stand for 1 h with intermittent shaking.

8. Centrifuge each sample for 10 min.

CALIBRATION AND QUALITY CONTROL:

9. Calibrate daily with at least six working standards over the range 0.05 to 7 μg analyte per sample.

a. Deliver aliquots of calibration stock solution with microliter syringe to eluent in 10-mL volumetric 

flasks and dilute to the mark.

b. Analyze together with the samples and blanks (steps 12 through 14).

c. Prepare calibration graph (peak area vs. μg analyte).

10. Determine recovery (R) at least once per year for each lot of filters. Prepare four filters at each of five 

levels, plus three media blank filters.

a. Place sample filters into separate test tubes.

b. Inject an aliquot of R stock solution, or a dilution thereof in methanol, directly onto the filter.

c. Cap the test tubes. Allow to stand overnight.

d. Prepare (steps 5 through 8) and analyze (steps 12 through 14) with working standards.

e. Prepare a graph of R for each filter vs. μg analyte recovered.
BENZIDINE and 3,3’-DICHLOROBENZIDINE: METHOD 5509, Issue 3, dated 29 February 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

11. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration and R 

graphs are in control.

MEASUREMENT:

12. Set the liquid chromatograph to conditions given on page 5509-1 for analyte of interest.

NOTE 1: If aniline is present, use a C18 column and follow the procedures found in reference [2].

NOTE 2: The following compounds were found not to interfere with the determination of either 

compound: o-, p- and m-chloroaniline; 4,4'-methylenedianiline and ß-naphthylamine. 2-

Chloro-4-methylaniline; 3,3'-dichlorobenzidine; 4,4'-methylenebis(2-chloroaniline); 

hydrazobenzene; and 1,2- and 1,4-naphthoquinone do not interfere in the determination of 

benzidine. Benzidine, aniline, N-methylaniline, 2-toluidine and 3,3'-dimethylbenzidine will 

not interfere in the determination of 3,3'-dichlorobenzidine [1].

13. Inject an aliquot (see page 5509-1 for appropriate size).

14. Measure peak area.

CALCULATIONS:

15. Read the mass, μg (corrected for R), of analyte found on the sample filter (W) and on the media blank 

filter (B) from the calibration graph.

16. Calculate concentration, C, of analyte in the air volume sampled, V (L):

CC = WW − BB

VV , mmmm mm3 ⁄

EVALUATION OF METHOD: 

This method was evaluated over the range 21 to 63 μg/m3 for benzidine and the range 20 to 130 μg/m3 for 

3,3'-dichlorobenzidine. The generated atmospheres for both compounds were at 30 °C and 80% relative 

humidity. The sampling rate was 0.8 L/min. The pooled overall precision (SŜ

rrrr) was 0.07 for 29 benzidine 

samples and 0.07 for 28 3,3'-dichlorobenzidine samples. The sampling methods were evaluated for effects of 

temperature (25, 30, and 35 °C) and relative humidity (20 and 80%). No detectable quantities of either 

benzidine or 3,3'-dichlorobenzidine were found on backup silica gel tubes [1]. At 180 °C, vapors of these 

carried by a stream of dry nitrogen at 0.3 L/min did not break through the backup silica gel in 3 h [1].

The average recovery of benzidine from filters was determined to be 97% over the range 0.2 to 2.0 μg when 

stored at -15 °C for 11 days. Recoveries dropped to 89% and 75% after 15 days and 21 days, respectively. 

Recoveries of benzidine and benzidinium sulfate from filters and silica gel indicated that the compounds 

were unstable in these matrices at ambient temperature. Recoveries over the range of 0.5 to 5 μg 3,3'-

dichlorobenzidine and its dihydrochloride were 96% after 21 days from both filters and silica gel stored at 

-15 °C and at ambient temperature [1].

REFERENCES:

[1] Morales R, Rappaport SM, Weeks, Jr. RW, Campbell EE, Ettinger HJ [1977]. Development of sampling and 

analytical methods for carcinogens, January 1 - September 30, 1976, Los Alamos Scientific Laboratory, 

Los Alamos, NM. Available as LA-7058-PR, from NTIS. Unpublished.

[2] Kennedy ER, Seymour MJ [1981]. Development of an analytical method for benzidine-based dyes. In: 

Chemical hazards in the Workplace, ACS symposium series: Vol. 149. Washington, DC: American 

Chemical Society, pp 21-35.
BENZIDINE and 3,3’-DICHLOROBENZIDINE: METHOD 5509, Issue 3, dated 29 February 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[3] NIOSH [1977] P&CAM 243 and P&CAM 246. In: Taylor DG, ed. NIOSH manual of analytical methods, 2nd 

ed. (Vol. 1) Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Centers for Disease 

Control, National institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-157-B. 

[4] NIOSH [1980]. NIOSH technical report: carcinogenicity and metabolism of azo dyes, especially those 

derived from benzidine. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health.

[5] NIOSH/NCI [1978]. Current intelligence bulletin 24: Direct Blue 6, Direct Black 38, and Direct Brown 95 

benzidine-derived dyes. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for

Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 

78-148.

[6] NIOSH [1981]. Occupational health guidelines for chemical hazards. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control, National Institute for Occupational Safety and 

Health, DHHS (NIOSH) Publication No. 81-123.

METHOD REVISED BY:

M. J. Seymour, NIOSH.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of these 

websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES 5524

DEFINITION: Metalworking fluids CAS: None RTECS: None

METHOD: 5524, Issue 2 EVALUATION: FULL 

Issue 1: 15 March 2003 

Issue 2: 29 December 2014

OSHA : No PEL

NIOSH: 0.4 mg/m3 as thoracic fraction 

(0.5 mg/m3 as ‘total’ aerosol)

Other

OELs: [1,2]*

PROPERTIES: Not defined. Fluids may contain varying 

amounts of mineral oil, emulsifiers, water, 

alkanolamines, polyethoxyethanols, biocides, 

surfactants, pressure additives and boron 

compounds.

SYNONYMS: Metalworking fluids (MWF), metal removal fluids, machining fluids, mineral oils, straight fluids, soluble fluids, 

synthetic fluids and semi-synthetic fluids

SAMPLING

SAMPLER: Thoracic particles: FILTER + CYCLONE (tared 

37-mm, 2-μm polytetrafluoroethylene (PTFE) 

filter + thoracic cyclone) 

Total particulate: tared 37-mm, 2-μm PTFE 

filter

FLOW RATE: Thoracic – 1.6 L/min,

Total – 2 L/min

VOL-MIN: 768 L at 0.4 mg/m3

 or 0.5 mg/m3

 MAX: Not determined

SHIPMENT: Ship overnight in a refrigerated container

SAMPLE 

STABILITY: Refrigerate upon receipt; analyze within 2 

weeks of collection

BLANKS: At least 5 field blanks per set

BULK

SAMPLE: One for each fluid at each site for solubility 

testing

ACCURACY

RANGE STUDIED: 0.05 to 0.9 mg/sample

BIAS: Not determined

OVERALL

PRECISION (ŜrT): Total weight- 0.06

Extracted weight - 0.07

ACCURACY (Estimated): Total weight - 0.12

Extracted weight - 0.14

MEASUREMENT

TECHNIQUE: Gravimetric

ANALYTE: Airborne metalworking fluid aerosol 

EXTRACTION: Ternary solvent: dichloromethane: 

methanol:toluene (1:1:1)

Binary solvent: methanol: water (1:1)

BALANCE: 0.001 mg sensitivity; use same balance 

before and after sample collection

CALIBRATION: National Institute of Standards and 

Technology Class S-1.1 weights or ASTM 

Class 1 weights

RANGE: 0.05 to 2 mg per sample

ESTIMATED LOD: Total weight - 0.03 mg per sample [3] 

Extracted weight - 0.03 mg per sample 

weight [3]

PRECISION ( ): Total weight - 0.04 (≥ 0.2 mg/sample) [4]

Extracted weight - 0.05 (≥ 0.2 mg/ 

sample) [4]

APPLICABILITY: The method is applicable to all metalworking fluids-straight, soluble, synthetic and semi-synthetic as 

long as they are soluble in the extraction solvent regimen [4,5]. Only one MWF (Glacier, Solutia Inc.) has thus far been found 

to be insoluble in the ternary extraction solvent; however, that MWF is soluble in the binary blend and tests have shown 

that the binary solvent in combination with the ternary solvent is effective in extracting this fluid [6]. Thoracic samplers

meeting the International Standard ISO 7708 thoracic convention within the performance specifications of the European 

Standard EN13205, parts 1 through 6 [2] may be considered for this method if they have been validated for the collection of 

MWF. Depending on the type of thoracic sampler used, the recommended flow rate may differ from that specified above. 

Sampling at flow rates exceeding those recommended here may result in increased evaporative loss of sample. Welding 

fume may significantly interfere with proper operation of certain impactor-style samplers.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 2 of 10

INTERFERENCES: None identified. However, any material that is collected on the filter and is soluble in 

the extraction solvents may interfere (positively) with the analysis.

OTHER METHODS: This method is similar to NMAM Method 0500 for Particulates Not Otherwise 

Regulated [7] and replaces NMAM Method 5026 as a general technique for analysis of MWF. NMAM 

Method 5026, which employs infrared analysis, may be used solely for the analysis of (straight) MWF 

that produce mineral oil mists; that method is not recommended for use with water-based fluids [8]. 

ASTM D7049 is another method that may be used to analyze metalworking fluids [9].

REAGENTS:

1. Dichloromethane, distilled-in-glass (DIG) (See

Note)‡

2. Methanol, distilled-in-glass (See Note)‡

3. Toluene, distilled-in-glass (See Note)‡

4. Water, filtered, double deionized (18 MΩ∙cm

resistivity)

5. Calcium sulfate, desiccant

6. Ternary solvent blend‡

: Mix equal volumes of

dichloromethane, methanol and toluene in a

clean dust-free container. Use a bottle with a

screw cap (e.g., a clean, empty solvent bottle):

Mix the solvents by gentle swirling, not by

violent shaking.

7. Binary solvent blend‡

: Mix equal volumes

of methanol and water in a clean dust-free

container. Use a bottle with a screw cap (e.g., a

clean, empty solvent bottle): Mix the solvents

by gentle swirling, not by violent shaking.

NOTE: Lower grade solvents have not been

evaluated for this method. If it is desired 

to use ACS or liquid chromatographic 

grade solvents, in the interest of 

economy, the user must demonstrate 

that these solvents perform equally to 

the DIG grade ( blanks < those obtained 

with the DIG grade).

‡

 See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Sampler: 37-mm PTFE, 2-μm pore size

membrane filter (see Appendix 1 for PTFE filter

cleanup procedure) and PTFE supporting pad

in 37-mm cassette filter holder. Use a 2-piece

(closed face) cassette for sampling ‘total’ 

aerosol. For sampling the thoracic fraction, use

a 3-piece cassette with thoracic cyclone that

samples at 1.6 L/min at the thoracic cutpoint.

See Evaluation of Method. Also, see page

5524-1 for discussion of alternative thoracic

sampling.

2. Personal sampling pump, 1.6 to 2 L/min, with

flexible connecting tubing

3. Cassette shrink bands, cellulose, 37-mm size

4. Microbalance, capable of weighing to 0.001mg

5. Static neutralizer: e.g., 210Po; replace nine

months after the production date

6. Forceps (preferably nylon or chrome-plated

steel)

7. Extraction funnel, See Figure 1 for

specifications p. 5524-10

8. Desiccator

9. Wash Bottle, PTFE

10. Vials, 20-mL and 10-mL, glass, with leak-proof

PTFE-lined caps

11. Syringe, gas-tight with large bore needle, e.g.,

16-gauge needle

12. Graduated cylinder, glass, 20 mL

13. Paper towels

14. Metal screen for drying filters following

extraction, approximately 550 cm2

 or other

convenient size. Grid size approximately 12 to

18 mm. Pre-wash screen with ternary blend

solvent and allow to dry.

SPECIAL PRECAUTIONS: Dichloromethane is an inhalation hazard and is a suspect carcinogen. Handle 

all solvents in a fume hood. The solvents are flammable and have associated adverse health effects. 

Avoid breathing vapors. Avoid skin contact. Use extreme caution when blending the solvents together. 

The heat of mixing can cause pressure to develop as the solvents are blended, e.g., blowing a stopper 

from a glass-stoppered container. Use clean containers sealed with PTFE-lined screwcaps. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 3 of 10

CALIBRATION AND QUALITY CONTROL:

1. Zero the microbalance before all weighings. Use the same microbalance for weighing filters before

and after sample collection. Maintain and calibrate the balance with National Institute of Standards

and Technology Class S-1.1 or ASTM Class 1 weights.

2. Process at least three tared media blanks through the measurement process for ‘total’ aerosol and

the extractable materials.

PREPARATION OF FILTERS BEFORE SAMPLING:

3. Number the backup pads with indelible ink and place them, numbered side down, in the filter

cassette bottom sections.

4. Pre-weigh the filters by the weighing procedure given in step 5. Record the mean tare weight of

sample filters, W1 and field blanks, B1 (mg).

5. Weighing procedure:

a. Equilibrate the filters in an environmentally controlled weighing area or chamber for 1 hour

(minimum.)

b. Zero the balance before each weighing. Using forceps, pass each filter over a static neutralizer.

Repeat this step if the filter does not release easily from the forceps or attracts the balance pan.

(Static electricity can cause erroneous weight readings.)

c. Weigh each filter until a constant weight is obtained (i.e., two successive weighings within 10 μg.)

d. Record the mean of the last two weights to the nearest microgram.

6. Assemble the filter in the 2- or 3-piece filter cassettes and close firmly so that leakage around the

filter will not occur. Place a plug in each opening of the filter cassette. Place a cellulose shrink band

around the filter cassette and allow to dry. Alternatively, use heavy duty elastic tape instead of the

shrink band. Mark with the same number as the backup pad.

SAMPLING:

Bulks: For solubility testing, obtain liquid samples of pure uncut bulk metal-working fluids (MWF) 

that are expected to be sampled in worker breathing zone. Place these samples in small (e.g., 

10 mL) leak-proof glass container(s) that are sealed with a leak-proof PTFE-lined screwcap. 

Then place them inside of a resealable plastic bag and ship these samples to the laboratory 

along with the sample filter cassettes. 

Air: For collection of a thoracic sample, insert the cyclone into the inlet of a 3-piece cassette. For 

collection of a “total” sample, do not use the cyclone. 

7. Calibrate each personal sampling pump with a representative sampler in line.

8. For thoracic measurements, sample at 1.6 L/min for 8 hours. For ‘total’ aerosol measurements, sample

at 2 L/min for 8 hours. Do not exceed a total filter loading of approximately 2 mg.

9. Submit at least 5 blank filter samples as field blanks and 3 filters for media blanks for each set of

samples collected per day. Handle the field blanks in the same way as the field samples; i.e., open

each in a non-contaminated environment, then close the sampler and ship it to the lab along with

the remaining samples. Media blanks are not opened.

10. Refrigerate all samples that are to be stored overnight (or longer) prior to shipment to the

laboratory. Ship all samples in refrigerated containers to the laboratory via overnight express

delivery service.

11. Refrigerate the samples immediately upon receipt at the lab until ready for analysis.

12. Analyze the samples within two weeks of receipt at the laboratory.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 4 of 10

SAMPLE PREPARATION AND MEASUREMENT:

13. Solubility test of bulk MWF:

a. Shake the container of bulk MWF to ensure that a homogeneous sample is obtained.

b. Place 10 mL of the ternary solvent blend in a 20-mL glass vial.

c. Using a large-bore gas-tight syringe, inject 50 μL of the bulk MWF into the ternary solvent blend.

Cap the vial and shake as necessary to dissolve the MWF. The fluid is soluble if the resulting

solution is clear and free of precipitates and phase separation.

d. If the MWF is soluble in the ternary blend, the samples can be extracted with the ternary blend.

14. Wipe dust from the external surface of each filter cassette (containing either samples or blanks) with

a moist paper towel to minimize contamination. Discard the paper towel.

15. Remove the top and bottom plugs from the filter cassette. Equilibrate the filters (in the cassettes) for

no more than 2 hours in a desiccator that uses calcium sulfate.

16. Remove from the desiccator. Equilibrate for at least 1 hour in the balance room or chamber.

17. Remove the cassette band, pry open the cassette, and remove the filter gently to avoid loss of

sample.

NOTE: If the filter adheres to the underside of the cassette top, very gently lift it away by using the

dull side of a scalpel blade. This must be done carefully or the filter will tear.

18. Weigh and record (steps 5b through 5d) the post-sampling weight of each filter, W2 (mg) and

blank, B2 (mg). Record anything remarkable about the filter (e.g., overload, leakage, wet, or torn). 

Perform the extraction as soon as possible. Store in a clean dust free environment until ready to 

perform the extraction, etc.

EXTRACTION:

19. Perform all extractions in an exhaust hood. General guidelines (see NOTE below):

NOTE: Samples weighing less than 0.4 to 0.5 mg (for a 1 m3 sample) may be extracted as desired.

The reason that the cutoffs of 0.4 and 0.5 mg (per 1000 L sample) have been specified is to 

assure compliance with the occupational exposure limit (OEL). If the gross sample weight 

indicates that the OEL has not been exceeded, there may be no reason to extract the sample. 

Otherwise, the usefulness of any extraction data obtained at levels less than 0.4 to 0.5 mg per 

sample is guided by the limit of quantitation (LOQ) of the extraction procedure. Extraction 

data obtained at levels between the limit of detection (LOD) and the LOQ of the extraction 

procedure should be used with appropriate caution due to the imprecision associated with 

such data. 

20. Do not presume that a fluid that is soluble only in the binary-blend should be extracted using only

the binary blend. The possibility of mixed exposures always dictates that the extraction procedure 

with both the binary and ternary solvent blends should be followed. If the weights of samples 

exceed the amount expected to be collected at the REL, e.g., 0.4 mg (thoracic fraction) or 0.5 mg 

(‘total’ aerosol) for a 1 m3 air sample, then extract the samples and blanks as follows: 

a. Place each filter (membrane side up) in the filter funnel assembly connected to the vacuum

source.

b. Pour one 10-mL aliquot of the ternary solvent down the inside wall of the funnel over the filter.

Allow the solvent to contact the filter for no more than 5 to 10 minutes. Remove the solvent 

under slight vacuum.

c. Pour one 10-mL aliquot of the binary solvent down the inside of the funnel over the filter. Allow

the solvent to contact the filter for no more than 5 to 10 minutes. Remove the solvent under 

slight vacuum.

d. Pour a second 10-mL aliquot of the ternary solvent down the inside of the funnel over the filter.

Allow at least 30 seconds of contact time. Remove the solvent under slight vacuum. Wash the 

inner wall of the filter funnel with 1 to 2 mL of the ternary blend contained in a PTFE wash bottle. 

Remove the solvent under slight vacuum.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 5 of 10

e. Turn off the vacuum to the filter funnel.

f. Carefully remove the filter from the filter funnel, place it on the clean metal screen, and allow to

dry on the metal screen for at least 2 hours in a fume hood. Do not remove the filter from the

funnel while vacuum is applied or the filter may delaminate.

NOTE: One metalworking fluid, Glacier (Solutia Chemical, St Louis), was insoluble in the ternary 

blend but was soluble in the binary blend. Tests have shown that this fluid is extracted 

efficiently from the filters using steps 20a through 20e [5].

21. Weigh each filter, including field blanks, following steps 5a through 5d. Record the post-extraction

weight, W3 (mg), of the extracted sample filters and B3

 (mg), for the extracted blank filters. Record 

anything remarkable about the extracted filter (e.g., torn, wet, delamination, etc.).

CALCULATIONS:

22. Calculate the concentration of ‘total’ aerosol or thoracic fraction, C (mg/m3

), in the air volume

sampled, V (L):

 where: W1 = mean tare weight of filter before sampling (mg) (step 5)

W2 = mean post-sampling weight of sample-containing filter (mg) (step 18)

B1

 = mean tare weight of all blank filters (mg) (step 5)

B2

 = mean post-sampling weight of all blank filters (mg) (step 18)

23. Calculate the concentration of extracted MWF aerosol CMWF (mg/m3

), in the air volume sampled, V (L):

 where: W2 = mean post-sampling weight (pre-extraction weight) of sample-containing 

 filter (mg) (step 18)

W3

 = mean post-extraction weight of sample-containing filter (mg) (step 21)

B2

 = mean post-sampling weight of all blank filters (mg) (step 18)

B3

 = mean post-extraction weight of all blank filters (mg) (step 21)

24. Report the concentration C as ‘total’ aerosol or thoracic fraction weight; report the concentration

CMWF as the weight of the MWF aerosol.

EVALUATION OF METHOD:

The ‘total’ weight procedure permits an estimate of the ‘total’ collected particulate aerosol, including 

nuisance dust, airborne metal particulate and metal working fluid. If the extraction procedure is used, 

the technique permits an estimate of the ‘total’ collected metalworking fluid to which the worker is 

exposed. The development of the ternary solvent used in this method is described in reference 3. 

This method was initially tested with representative samples of straight, soluble, semi-synthetic, and 

synthetic metalworking fluids (MWFs). Samples were spiked onto tared polytetrafluoroethylene (PTFE) 

membrane filters, stored overnight, and analyzed the following day. The samples were weighed, then 

the MWF was extracted from the filter with a 1:1:1 blend of dichloromethane:methanol:toluene. The 

fractions extracted (FE or mass recovered/mass spiked) exceeded 94% for all fluids extracted from the 

filters over the range from 200 μg to 815 μg for the straight fluid, from 223 μg to 878 μg for the soluble 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 6 of 10

fluid, from 51 μg to 189 μg for the semi-synthetic fluid, and from 102 μg to 420 μg for the synthetic 

fluid. For those weights of all four fluids spiked at levels ≥ 200 μg, the relative standard deviation was 

estimated to be 4% for the total weight procedure and 5% for the extraction procedure. If the sampling 

imprecision of 5% is included, these estimates become 6% and 7% respectively for the total weight 

and extraction procedures. Limits of quantitation, estimated from blanks carried through the entire 

analytical procedure, were 30 μg for the weighing technique and 60 μg for the extraction technique. No 

estimate of the bias was available [4]. The filters are desiccated to remove excess water from water-based 

MWF samples.

In a more rigorous test of this method for a 79-plant survey [2], the average limits of quantitation were 

estimated to be 0.1 mg for both the total and extracted weight procedures. However, there was high 

variability in these estimates for the sites sampled. The upper 95% confidence limit for the LOQs for both 

the total weight and extracted weight measurements was 0.3 mg. In order to assess the effectiveness of 

the extraction step, a secondary extraction of the most heavily-loaded filters obtained in this survey was 

conducted; on average, less than 5% of the sample weight was removed during the second extraction, 

indicating that the majority of extractable material had been removed during the first extraction. 

Samples were refrigerated upon receipt at the laboratory [2,10].

During the 79-plant survey, all thoracic sampling was conducted with a BGI Mdl 2.69 Thoracic sampler. 

This sampler has a thoracic cut point of 10 μm at 1.6 L/min. The stability of quality assurance (QA) 

samples, spiked separately with a straight, a soluble, a semi-synthetic, and a synthetic fluid, indicated 

that the QA samples all lost weight according to simple linear decay equations. This loss in weight was 

likely due to evaporation of the spiking solvent and water (for soluble, semi-synthetic and synthetic 

MW fluids.) These decay equations were used to estimate the amounts expected to be reported for QA 

filters by the performing laboratory. For storage periods ranging from 17 to 26 days, the total weight 

of samples recovered for all QA samples were greater than or equal to 80% of those expected from the 

decay equations. For these QA samples, the fractions extracted of all four fluid types were greater than 

or equal to 90%.

The binary solvent extraction step has been added to assure complete extraction of MWF components 

that may be incompletely removed by the ternary blend. In addition, the binary solvent extends the 

procedure to samples that contain ternary blend-soluble fluids co-mingled with ternary blend-insoluble 

fluids, e.g., Glacier (Solutia Inc.) Tests of the extraction of five MWF (including Glacier) showed that 

extraction efficiencies using the ternary blend in combination with the binary blend were comparable 

to those reported in reference 1 using the ternary blend alone (FE greater than 90 %; CV less than 0.10). 

The binary solvent extractant liquor obtained from the Glacier samples generally contained potassium 

and phosphorous at levels approximately expected for the mass spiked onto the filters. The binary 

solvent extracts of the four other test fluids were analyzed for sodium, potassium or boron marker 

elements. Sodium was present in the extract of the soluble fluid at greater than background levels. The 

boron marker was not detected in the extract from the semi-synthetic fluid. The potassium marker was 

not detected in the extract from the synthetic fluid [5].

This method was further evaluated in a six-laboratory round robin study using synthetically generated 

atmospheres of an aerosol of a soluble MWF at the 0.5 mg/sample level. The data were evaluated 

according to ASTM standard E691-99. Pooled estimates of the total coefficients of variation were 0.13 for 

both the total and extracted weight samples. Overall there was no significant bias in the results. LOQs 

were comparable to those reported above [11].

REFERENCES:

[1] Health and Safety Executive (HSE), United Kingdom [2003]. MDHS 95/2, Measurement of personal 

exposure of metalworking machine operators to airborne water-mix metalworking fluid. [http://

www.hse.gov.uk/pubns/mdhs/pdfs/mdhs95-2.pdf] Website accessed March 25, 2013.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 7 of 10

[2] BSI (British Standards Institute) [2014]. BS EN 13205:2014.  Workplace exposure. Assessment of 

sampler performance for measurement of airborne particle concentrations. Parts 1 through 6. 

Brussels, Belgium: European Committee for Standardization.

[3] Glaser RA, Shulman S, Kurimo R, Piacitelli G [2002]. Data supporting a provisional American Society 

for Testing and Materials (ASTM) Method for metalworking fluids, Part 3: evaluation of a provisional 

ASTM Method for metalworking fluids in a survey of metalworking facilities. Journal of Testing and 

Evaluation 20(5): 439-451.

[4] Glaser RA, Shulman S, Klinger P [1999]. Data supporting a provisional American Society for Testing 

and Materials (ASTM) Method for metalworking fluids, Part 2: preliminary report of evaluation of 

a ternary extraction solvent in a provisional ASTM Test Method for metalworking fluids (PS-42-97). 

Journal of Testing and Evaluation 17(3):131-136.

[5] Glaser RA [1999]. Data supporting a provisional American Society for Testing and Materials (ASTM) 

Method for metalworking fluids, Part 1: a solvent blend with wide-ranging ability to dissolve 

metalworking fluids. Journal of Testing and Evaluation 17(3):171-174.

[6] Glaser RA, Kurimo R, Neumeister C, Shulman S [2005]. Data supporting ASTM Method D7049-04 for 

determination of metalworking fluids, ASTM Journal of Testing and Evaluation, 33(5):323-330.

[7] NIOSH [1994]. Particulates not otherwise regulated: Method 0500. In: Eller PM, Cassinelli ME, eds. 

NIOSH Manual of Analytical Methods, 4th ed. Cincinnati OH: U. S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and 

Health, DHHS (NIOSH) Publication No. 94-113. [www.cdc.gov/niosh/nmam/].

[8] NIOSH [1994]. Oil Mist, Mineral: Method 5026. In: Eller PM, Cassinelli ME, eds. NIOSH Manual of 

Analytical Methods, 4th ed. Cincinnati OH: U. S. Department of Health and Human Services, Centers 

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 94-113. [www.cdc.gov/niosh/nmam/].

[9] ASTM D7049 [2010]. Standard Test Method for Metal Removal Fluid Aerosol in Workplace 

Atmospheres.  ASTM International, West Conshohocken, PA.

[10] Piacitelli G, Hughes R, Catalano J, Sieber W, Glaser RA, Kent M [1997]. Exposures to Metalworking 

fluids in small size machine shops. Presented at The Industrial Metalworking Environment: 

Assessment and Control of Metal Removal Fluids Symposium, Detroit, Michigan, September 1997.

[11] Glaser RA, Kurimo R, Shulman S [2007]. Performance testing of NIOSH Method 5524/ASTM method 

D-7049-04 for determination of metalworking fluids. J Occup Environ Hyg, 4:583-595.

METHOD WRITTEN BY:

R. Glaser, NIOSH/DART and edited by Y. T. Gagnon, NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for 

the content of these websites. All web addresses referenced in this document were accessible as of 

the publication date.

* NOTE: Because exposure limits and guidelines may change over time, NIOSH recommends referring to

the following sources for updated limits and guidelines. [1,2] 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 8 of 10

Appendix 1. Procedure to remove PTFE filter contaminants that are soluble 

 in and extracted with the ternary blend. 

Use this procedure to prevent high blanks from being obtained following extraction of PTFE filters. The 

filters are rinsed with ternary solvent (1:1:1 dichloromethane:methanol:toluene) described in NMAM 

Method 5524. The filters are air-dried and protected from airborne contamination prior to weighing 

and assembly into cassettes for field sampling. See evaluation of procedure which follows these 

instructions. 

EQUIPMENT, SUPPLIES, REAGENTS AND STANDARDS 

1. 250 mL glass beaker(s)

2. 500 mL of ternary solvent 1:1:1 ratio of dichloromethane:methanol:toluene in a stoppered container

3. A fume hood

4. Watch glasses to fit the beakers

5. Glass stir rod

6. One box (or as many as required) of 2 micron PTFE filters to be cleaned; recommend cleaning no

more than 1 package of 50 filters per 100 mL of solvent

7. Stainless steel forceps to manipulate filters

8. Large, lint-free paper towels

9. Clean, stainless steel metal trays or screen approximately 45 cm x 60 cm (18 inch x 24 inch)

10. Nitrile gloves

11. Clean, wide mouth glass container e.g., ointment jar with Teflon cap or French square bottle

PROCEDURE 

Perform all of the following tasks inside a fume hood! 

1. Wear nitrile gloves throughout this procedure to protect yourself from the solvent and to protect

the filters from skin oil during handling. 

2. Pour approximately 100 mL of ternary solvent into a 250 mL glass beaker.

3. Remove the PTFE filters from the package and remove the plastic spacers from between the filters.

Place the filters individually (sample side up) into the beaker filled with ternary solvent. Be careful

that the filters stay separated from each other and do not clump together.

4. Carefully stir the filters with the glass stir rod; do not allow the filters to turn over. Cover the beaker

with the watchglass. Allow the filters to extract in the solvent in the beaker for 10 minutes.

5. Decant the ternary solvent out of the beaker and into a waste bottle. Use the stirring rod to

compress and retain the filters in the beaker as the solvent is slowly decanted off.

6. Refill the beaker with 100 mL of fresh ternary solvent after decanting following step 5. Repeat steps

4 and 5, two times for a total of three solvent washes.

7. After the three rinses are complete, drain off as much solvent as possible. Remove the filters from

the beaker using forceps and place them onto a clean stainless steel metal tray that has been

covered with a large, lint-free paper towel. Alternatively, place them on a clean stainless steel screen.

Place them onto the tray or screen, sampling side up. Allow the filters to dry overnight.

Warning: Keep the hood sash approximately at or below the sash height level as marked on the

hood. If the sash is pushed lower, the higher air flow may blow the filters off the drying

tray or screen. Place a sign on the sash indicating that it is to be left at this height 

overnight and not to be moved.

8. Place the dried filters in a clean French square glass bottle or a wide-mouthed TeflonTM capped

ointment jar. Label the container “PTFE 2-μm filters rinsed with MWF solution,” giving the date, 

initials, and number of filters. Do not store filters in plastic containers. Filters are now ready to be 

used for sampling. Since the filters are not separated by spacers, use care to remove them from the 

storage container for use.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 9 of 10

Cleanup Procedure Evaluation

This cleanup procedure has been incorporated into NMAM Method 5524 in order to deal with reported 

spurious weight gains and losses before and after analysis of the PTFE filters used with this method. 

It is believed that trace levels of dust or extractable material are entrained in these filters during the 

manufacturing process. This cleanup procedure has been evaluated using 60 filters from three different 

batches of PTFE filters (20 filters/batch) [1]. Prior to cleaning, the filters were weighed (untreated 

filters), then washed with the ternary blend according to the procedure in Appendix 1, dried, and then 

reweighed (treated filters).

Results: The differences in each of the 3 batches were compared by subtracting the treated filter 

weights from their untreated weights. For all 60 samples, the overall mean difference in weights (+/-the 

standard error of the mean) was 0.2 μg (± 1.4 μg). For each of the 3 batches, the differences in weight 

were reported as mean (+/-standard error): 5 μg (± 2.5 μg) , -2 μg (± 2.0 μg), and -3 μg(± 2.7 μg) for 

batches 1, 2 and 3, respectively. The weight differences were not statistically significant from zero overall 

or by batch using a paired t-test (p = 0.05), and allowing for multiple comparisons. 

To determine if the washing procedure affected the filter’s performance for analysis of metalworking 

fluids, each of the filters was analyzed according to the procedure of NMAM Method 5524, which 

includes extraction with the binary and ternary solvent blends. The following differences were 

computed: post-analysis weights of the filters minus their treated or untreated weights. The average 

difference in the weights of the 60 untreated filters and their post-analysis weights reported as mean 

(± standard error of the mean) was 34 μg (± 1.9 μg). For each of the 3 batches, the differences in the 

untreated and post-analysis weights were: 30 μg (± 3.9 μg), 38 μg (± 2.0 μg) and 34 μg (± 3.3 μg) for 

batches 1, 2 and 3, respectively. These differences are statistically significantly different from zero, both 

overall and individually by batch, using a paired t-test (p = 0.05), and allowing for multiple comparisons. 

The average differences in the weights of the 60 treated filters and their post-analysis weights were: 

34 μg (± 1.4 μg). Again the differences in weight are reported as mean (± standard error). For each of 

the 3 batches, the differences, (post-analysis minus treated) weights, were: 35 μg (± 3.4 μg), 36 μg 

(± 2.0 μg), and 31 μg (± 1.8 μg) for batches 1, 2 and 3, respectively. These differences are statistically 

significantly different from zero, both overall and individually by batch, using a paired t-test (p = 0.05), 

and allowing for multiple comparisons.

In summary, the weight differences were statistically different from zero by batch and overall using a 

paired t-test for both the post-analysis minus treated weights and for the post analysis-untreated

weights (p = 0.05). However, the untreated/treated differences were not statistically different using the 

same tests.

These experiments indicate that cleaning the filters lowered the overall LOQ of the analytical method. 

The LOQ determined from the differences in weights between the untreated and analyzed filters was 

140 μg and was greater than the LOQ of 110 μg determined from the differences in weights between 

the treated and analyzed filters. The extraction procedure produced a more consistent blank and 

therefore a lower average standard deviation from which lower LOD s and LOQs were determined.

[1] NIOSH [2014]. CEMB Analytical Services Report: Sequence 11600-CA. Novi, MI: Bureau Veritas North 

America, unpublished.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METALWORKING FLUIDS (MWF) ALL CATEGORIES: METHOD 5524, Issue 2, dated 29 December 2014 - Page 10 of 10

Figure 1. Schematic diagram of funnel. May be ordered from Case Custom Environmental Equipment, 

 Erlanger, Kentucky (859-250-8558); www.casecustomenvironmentalequipment.com) or 

 equivalent source. Dimensions are given in inches.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYLTIN CHLORIDES 5526

Formula: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 5526, Issue 2 EVALUATION: FULL Issue 1: 15 March 2003

Issue 2: 13 January 2016

OSHA: See Table 1

NIOSH: See Table 1

PROPERTIES: See Table 1

SYNONYMS: Methyltin trichloride: monomethyltin trichloride, trichloromethylstannane 

Dimethyltin dichloride: dichlorodimethylstannane, dichlorodimethyltin 

Trimethyltin chloride: chlorotrimethylstannane, chlorotrimethyltin, trimethylstannyl chloride

SAMPLING

SAMPLER: FILTER + SORBENT TUBE

(OVS-2 tube: 13 mm glass fiber filter; XAD-2, 

270 mg/140 mg)

FLOW RATE: 0.25 L/min to 1 L/min

VOL-MIN: 15 L

-MAX: 75 L

SHIPMENT: Ship assembled sampler cold

SAMPLE

STABILITY: 14 d @ 4 °C

FIELD

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: Table 2

BIAS: Table 2

OVERALL

PRECISION ( ): Table 2

ACCURACY: Table 2

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, FPD (tinspecific filter)

ANALYTE: Sodium tetraethylborate derivatives of 

the analytes

DESORPTION: 2 mL 0.17 mol/L acetic acid in acetonitrile

INJECTION: 1 μL

TEMPERATURE-INJECTOR: Cool on-column

-DETECTOR: 250 °C

-COLUMN: 50 °C (3 min), 12 °C/min to 200 

°C (1 min)

CARRIER GAS: Helium, 6 mL/min

COLUMN: Capillary, fused silica, 30 m × 0.53 

mm ID, 1.5 μm film, (5% phenyl)-

methylpolysiloxane; US Pharmacopeia 

(USP) G27

CALIBRATION: Analytes in hexane

RANGE: 0.01 μg to 15 μg per sample (as tin) [2]

ESTIMATED LOD: 0.01 μg [2]

PRECISION ( ): 0.065 [2]

APPLICABILITY: This method was developed for air monitoring of methyltin chlorides [2].

INTERFERENCES: None were identified.

OTHER METHODS: NIOSH method 5504 is another method for the measurement of organotin compounds using a filter 

with a sorbent tube for collection and HPLC/AA for analysis [3].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYLTIN CHLORIDES: METHOD 5526, Issue 2, dated 13 January 2016 - Page 2 of 5

REAGENTS:

1. Acetic acid, 99+%.*

2. Acetonitrile, HPLC grade.*

3. Diethyl ether, 99+%.*

4. Hexane, HPLC grade*.

5. Methyltin trichloride, 97%.

6. Dimethyltin dichloride, 97%.

7. Trimethyltin chloride.

8. Acetic acid, 0.17 mol/L in acetonitrile. Dilute 

1.00 mL of glacial acetic acid to 100 mL with 

acetonitrile.

9. Sodium tetraethylborate.*

NOTE: Purity of sodium tetraethylborate 

is critical. It should appear as a fine 

white powder. Clumping or yellowing 

indicates that the compound must be 

replaced.

10. Sodium acetate-acetic acid buffer, pH 4.0 ± 0.2. 

11. Derivatizing solution: Prepare a solution 

containing 10 g/L of sodium tetraethylborate 

in diethyl ether. Prepare the solution fresh 

daily, preferably immediately before use.

12. Calibration stock solution: Prepare a stock 

calibration solution by accurately weighing 

(to ± 0.1 mg) about 0.1 g each of methyltin 

trichloride, dimethyltin dichloride, and 

trimethyltin chloride into a 50 mL volumetric 

flask. Dilute to volume with 0.17 mol/L acetic 

acid in acetonitrile. The stock solution contains 

about 1000 μg/mL (as tin) of each component. 

Stored in a freezer, this solution remains stable 

for several weeks.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: OSHA versatile sampler (OVS-2 tube), 

13 mm OD inlet, 6 mm OD outlet. Front section 

contains 270 mg 20/60 mesh XAD-2 sorbent 

held in place by an 11 mm diameter glass fiber 

filter and PTFE ring, separated from the back 

section of 140 mg XAD-2 sorbent by a short 

plug of polyurethane foam. The back section is 

held in place with a long plug of polyurethane 

foam. The tube is available commercially.

2. Personal sampling pump: 0.1 L/min to 1 L/min 

with flexible and inert connecting tubing.

3. Gas chromatograph equipped with a cool 

on-column injection port, automated injector, 

capillary column, flame-photometric detector 

(FPD) with a tin-specific filter (610 nm), and 

data collection system (page 5526-1).

4. Balance capable of weighing to ±0.1 mg.

5. Mechanical shaker.

6. Vials, 10 mL, with PTFE-lined screwcaps.

7. Transfer pipettes or eyedroppers.

8. GC autosampler vials with PTFE-lined caps.

9. Volumetric flasks, 5 mL, 10 mL, and 50 mL.

10. Pipettes capable of accurately delivering 10 μL, 

1.0 mL, and 2.0 mL.

11. Refrigerant packs for shipping.

SPECIAL PRECAUTIONS: Concentrated acetic acid is corrosive and an irritant. Sodium 

tetraethylborate is sensitive to air and moisture. This chemical should be stored and handled under 

a nitrogen atmosphere. Acetonitrile, diethyl ether, and hexane are all flammable. Wear appropriate 

protective clothing and work with these compounds in a well-ventilated hood.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Uncap the sorbent tube immediately before sampling and connect to a personal sampling pump 

with flexible tubing.

3. Sample at an accurately known flow rate between 0.25 L/min for full shift sampling and 1 L/min for 

15 min samples (STEL) for a maximum of 75 L.

4. Cap the samplers. Pack securely for shipment with cold refrigerant packs.

NOTE: After receipt at the laboratory, samples should be stored at 4 °C and analyzed within 14 d of 

collection.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYLTIN CHLORIDES: METHOD 5526, Issue 2, dated 13 January 2016 - Page 3 of 5

SAMPLE PREPARATION:

5. Remove the retaining ring from the sorbent tube and discard. Transfer the glass fiber filter and the 

main resin bed to a 10 mL sample vial.

6. Remove and discard the foam plug that separates the two resin sections in the sorbent tube. 

Transfer the backup resin bed to a second 10 mL sample vial.

7. Into each vial, add 2 mL of 0.17 mol/L acetic acid in acetonitrile to desorb the methyltin chlorides. 

Put the vials on a mechanical shaker for 30 min.

8. Into each vial, add 2 mL of pH 4 buffer and 1 mL of derivatizing solution. Place the vials on the 

mechanical shaker for 15 min.

9. Extract the solution with three 1 mL portions of hexane. Combine the hexane fractions in a 5 mL 

volumetric flask and dilute to volume with hexane.

CALIBRATION AND QUALITY CONTROL:

10. Calibrate daily with at least six working standards over the range of interest.

a. Accurately pipette 10 μL of the calibration stock solution into a vial that contains 5 mL of 0.17 

mol/L acetic acid in acetonitrile. Derivatize like a sample following steps 7, 8, and 9.

b. Serially dilute this standard derivatized solution with hexane to produce solutions over the range 

of 0.2 μg/mL to 0.02 μg/mL (as tin) of each methyltin chloride.

c. Transfer to autosampler vials with PTFE-lined caps.

d. Analyze together with samples and blanks (steps 13, 14, and 15).

e. Prepare calibration graph (peak areas vs. μg tin for each methyltin chloride).

11. Determine desorption efficiency (DE) at least once for each lot of OVS tubes used for sampling in the 

calibration range (step 10). Prepare three samplers at each of six levels plus three media blanks.

a. Transfer the front sorbent section of the OVS sampler to a 4 mL vial.

b. Inject a known volume of calibration stock solution, or serial dilution, directly onto the front 

sorbent bed of each OVS tube.

c. Cap vial and allow spiked sorbent bed to stand overnight.

d. Desorb (steps 7, 8, and 9) and analyze together with standards and blanks (steps 13, 14, and 15).

e. Prepare a graph of DE vs. μg for each methyltin chloride recovered.

12. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration 

graph and DE graphs are in control.

MEASUREMENT:

13. Set gas chromatograph according to manufacturer’s recommendations and to conditions given on 

page 5526-1.

14. Inject a 1 μL sample aliquot manually using solvent flush technique or with an autosampler.

NOTE: If peak area is above the linear range of the working standards, dilute with hexane, reanalyze, 

and apply the appropriate dilution factor in the calculations.

15. Measure the areas of the peaks of the methyltin chlorides. A sample chromatogram is shown in 

Figure 1.

CALCULATIONS:

16. Determine the mass, μg (corrected for DE), for each methyltin chloride found in the sample front 

( ) and back ( ) sorbent sections, and in the average media blank front ( ) and back ( )sorbent 

sections.

NOTE: If , report breakthrough and possible sample loss.

17. Calculate concentration, , of each methyltin chloride in the air volume sampled, (L):
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYLTIN CHLORIDES: METHOD 5526, Issue 2, dated 13 January 2016 - Page 4 of 5

, μg/L or mg/m3.

EVALUATION OF METHOD:

This method, developed to collect air samples and analyze the methyltin chlorides, was adapted 

from existing methodology for measuring organotin compounds in air [3,4]. Reference 2 details 

the validation of this method for sampling and quantifying of trimethyltin chloride, dimethyltin 

dichloride, and methyltin trichloride in air. The validated method was needed to provide air monitoring 

capabilities during methyltin chloride and stabilizer production. Table 2 lists the results of the validation 

experiments, which followed the guidelines established by NIOSH [5]. However, fewer replicates and 

concentration levels were used than recommended. This method met the NIOSH criteria for accuracy, 

bias, and sample stability. The recommended sampling conditions are 250 mL/min for 5.5 h maximum 

for time-weighted average (TWA) sampling and 1000 mL/min for 20 min maximum for short-term 

exposure limit (STEL) sampling.

REFERENCES:

[1] Agency for Toxic Substances and Disease Registry (ATSDR) [2005]. Chemical and physical 

information. In: Toxicological profile for tin and tin compounds. Atlanta, GA: U.S. Department of 

Health and Human Services, Public Health Service [www.atsdr.cdc.gov/toxprofiles/tp55-c4.pdf].

[2] Yoder RE [2000]. Validation of a method for the collection and quantification of methyltin chlorides 

in the air. Philadelphia, PA: Atofina Chemicals. Unpublished.

[3] NIOSH [1994]. Organotin compounds (as ): Method 5504 (supplement issued 3/15/2003). In: 

Schlecht PC, O’Connor PF, eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. 

Department of Health and Human Services, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/

niosh/nmam/].

[4] Shangwei H [1997]. Development of a method for the sampling and analysis of dimethyltin 

dichloride and trimethyltin chloride. New Orleans, LA: Tulane University, Department of 

Environmental Sciences, School of Public Health and Tropical Medicine. Unpublished.

[5] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. 

By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 95-117 [www.cdc.gov/niosh/

docs/95-117/pdfs/95-117.pdf].

METHOD WRITTEN BY:

R.E. Yoder, C. Boraiko, Atofina Chemicals; P.F. O’Connor, NIOSH/DART.

DISCLAIMER: Mention of any company or product does not constitute endorsement by the National Institute for 

Occupational Safety and Health (NIOSH). In addition, citations to Web sites external to NIOSH do not constitute NIOSH 

endorsement of the sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for 

the content of these Web sites. All Web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHYLTIN CHLORIDES: METHOD 5526, Issue 2, dated 13 January 2016 - Page 5 of 5

Table 1. General information [1]

Analyte Formula MW CAS No. RTECS No. Properties

OSHA 

PEL*

(mg/m3)

NIOSH 

REL*

(mg/m3)

Methyltin 

trichloride

240.08 993-16-8 WH8585500 Colorless crystals; 

MP 43 °C; BP 171 °C

0.1 0.1

Dimethyltin 

dichloride

219.67 753-73-1 WH7245000 Colorless crystals; 

MP 90 °C (107 °C); 

BP 185 °C to 190 °C

0.1 0.1

Trimethyltin 

chloride

199.26 1066-45-1 WH6850000 Colorless crystals; 

MP 37.5 °C; 

BP 154 °C to 156 °C

0.1 0.1

*Measured as the amount of tin for organotin compounds in air.

Table 2. Method evaluation [2]

Compound

Range Studied 

(μg tin per sample) Bias (%)

Precision ( ) 

(%)

Accuracy 

(%)

Trimethyltin chloride 0.46 to 9.29 0.3 5.2 ±11

Dimethyltin dichloride 0.42 to 9.17 −2.5 5.8 ±12

Methyltin trichloride 0.07 to 10.48 −5.2 7.6 ±16

Figure 1. Sample chromatogram.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

ORGANOPHOSPHORUS PESTICIDES 5600

Analytes: Table 1 Formula: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 5600, Issue 2 EVALUATION: FULL Issue 1: 15 August 1993

Issue 2: 2 March 2016

OSHA: Table 2

NIOSH: Table 2

PROPERTIES: Table 3

SYNONYMS: Table 4 

SAMPLING

SAMPLER: FILTER + SOLID SORBENT TUBE

 (OVS-2 tube: 13-mm quartz filter; XAD-2, 

270 mg/140 mg)

FLOW RATE: 0.2 to 1 L/min

VOL-MIN: 12 L 

 -MAX: 480 L; 60 L (Malathion, Ronnel) 

SHIPMENT: Routine

SAMPLE

STABILITY: at least 10 days @ 25 °C

at least 29 days @ 0 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY [9]

RANGE 

STUDIED: Table 5

BIAS: Table 5

OVERALL

PRECISION (SS�rrrr): Table 5 

ACCURACY: Table 5

MEASUREMENT

TECHNIQUE: GC, flame photometric detection (FPD)

ANALYTE: Organophosphorus pesticides, Table 1 

EXTRACTION: 2 mL 90% toluene/10% acetone solution

INJECTION 

VOLUME: 1 to 2 μL

TEMPERATURE

-INJECTION: 240 °C

-DETECTOR: 180 °C to 215 °C (follow manufacturer’s 

recommendations)

CARRIER GAS: Helium @ 15 psi (104 kPa)

COLUMN: Fused silica capillary column; Table 6

DETECTOR: FPD (phosphorous mode) 

CALIBRATION: Standard solutions of organophosphorus 

compounds in toluene

RANGE: Table 8

ESTIMATED LOD: Table 9

PRECISION (SS̅

rr): Table 5 

APPLICABILITY: The working ranges are listed in Table 5. They cover a range of 1/10 to 2 times the OSHA PELs. This method also is 

applicable to STEL measurements using 12-L samples. This method may be applicable to the determination of other 

organophosphorous compounds after evaluation for desorption efficiency, sample capacity, sample stability, and precision and 

accuracy. 

INTERFERENCES: Several organophosphates may co-elute with either target analyte or internal standard causing integration 

errors. These include other pesticides (see Table 7), and the following: tributyl phosphate (a plasticizer), tris-(2-butoxy ethyl) 

phosphate (a plasticizer used in some rubber stoppers), tricresyl phosphate (a petroleum oil additive, hydraulic fluid, plasticizer, 

flame-retardant, and solvent), and triphenyl phosphate (platicizer and flame-retardant in plastics, laquers, and roofing paper). 

OTHER METHODS: This method may be used to replace previous organophosphorous pesticide methods. See Table 11 for partial 

listing. The sampler recommended here is similar in concept to the device of Hill and Arnold [11], but offers greater convenience 

and lower flow resistance. 
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 2 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Organophosphorous analytes listed in 

Table 1; and triphenyl phosphate, analytical 

standard grade.*

2. Toluene, pesticide analytical grade.*

3. Acetone, ACS reagent grade or better.*

4. Desorbing solution. Add 50 mL acetone to a 

500-mL volumetric flask. Dilute to volume 

with toluene. (Do not keep longer than 30 

days at 0-4 °C).

NOTE: For optional internal standard, add 1 

mL of a 5 mg/mL solution of 

triphenyl phosphate in toluene to 

500 mL desorbing solution. 

5. Organophosphorous stock solutions, 10 

mg/mL. Prepare individual standard stock 

solutions of each pesticide of interest in 

90/10 toluene/acetone (v/v). All pesticides 

in Table 1 were found to be soluble to at 

least 10 mg/mL.

6. Spiking solutions for calibration (step 9) and 

media fortification (steps 10, 11) 

NOTE: Spiking solutions may contain more 

than one analyte.

a. Spiking solution SS-1: Dilute the volume 

of stock solution indicated in Table 12 to 

10 mL with toluene or 90/10 

toluene/acetone (v/v).

NOTE: Spiking solutions may contain 

more than one analyte.

b. Spiking solution SS-0.1: Dilute 1 mL of 

SS-1 solution with toluene to 10 mL. 

7. Purified gases: Helium, Hydrogen, Dry air, 

Nitrogen, and Oxygen (if required by 

detector) 

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: resin filled sampling tube; glass 

tube, 11-mm i.d. x 13-mm o.d. x 50-mm 

long, with the outlet end drawn to a 6-mm

o.d. x 25-mm long tube. The enlarged part 

of the tube contains a 270-mg front section 

of 20/60 mesh XAD-2 sorbent held in place 

by a 9 to10-mm o.d. quartz fiber filter and 

polytetrafluoroethylene (PTFE) retaining 

ring. The front section is separated from the 

back section of 140 mg XAD-2 sorbent with 

a short plug of polyurethane foam. The back 

section is held in place by a long plug of 

polyurethane foam. The tube is available 

commercially. See Figure 2.

NOTE: Some commercially available 

samplers contain glass fiber filters. 

These are specified in the OSHA 

methods (see Table 11). These tubes, 

however, did not perform as well for 

the more polar analytes (amides, 

phosphoramides, and sulfoxides; 

(see Table 10). Low or erratic 

recoveries for Malathion may be 

encountered with glass fiber filters.

2. Personal sampling pump, 0.2 to 1 L/min. 

with flexible connecting tubing, preferably 

silicon, polyethylene, or PTFE tubing.

3. Vials, 4-mL with PTFE-lined cap; 2-mL GC 

autosampler vials with PTFE-lined crimp 

caps.

4. Gas chromatograph, flame photometric 

detector with 525-nm bandpass filter for 

phosphorous mode, integrator, and column 

(Table 6).

5. Syringes, 5-mL and 100-, 50-, and 10-mL for 

making standard solutions and GC 

injections.

6. Volumetric flasks, 500-, 10-, and 2-mL.

7. Tweezers.

8. GC vial crimper.

9. Small ultrasonic cleaning bath.

 
 ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 3 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SPECIAL PRECAUTIONS: Organophosphorous compounds are highly toxic. Special care must be taken 

to avoid inhalation or skin contact through the wearing of gloves and suitable clothing when handling 

pure material [13,14,15,16,17]. Toluene is flammable and toxic. Acetone is highly flammable. Prepare all 

samples in a well ventilated hood.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Connect the sampler to personal sampling pump with flexible tubing. The sampler should be placed 

vertically with the large end down, in the worker’s breathing zone in such a manner that it does not 

impede work performance [12]. 

3. Sample at an accurately known flowrate between 0.2 and 1 L/min for a total sample size of 12 to 240 L.

4. Cap both ends of the sampler with plastic caps and pack securely for shipment.

SAMPLE PREPARATION:

5. Remove cap from large end and remove PTFE retainer ring; transfer filter and front XAD-2 section to a 

4-mL vial. Transfer the short polyurethane foam plug with back-up XAD-2 section to a second 4-mL 

vial.

6. Add 2-mL of desorbing solvent with internal standard to each vial using a 5-mL syringe or 2-mL 

pipette. Cap each vial.

7. Allow to stand 30 minutes, immerse vials approximately 1/2 inch in an ultrasonic bath for 30 minutes. 

Alternatively, place the vials in a shaker or tumbler for 1 hour.

8. Transfer 1 to 1.5 mL from each 4-mL vial to a clean 2-mL GC vial, cap and label.

CALIBRATION AND QUALITY CONTROL:

9. Calibrate daily with at least six working standards covering the analytical range of the method for

individual analytes. See Table 12 for recommended preparation of spiking solutions.

a. Add known amounts of calibration spiking solution (SS-1 or SS-0.1 according to schedule

in Table 12) to desorbing solution in 2-mL volumetric flasks and dilute to the mark.

NOTE: If an internal standard is included in the desorbing solution, then exactly 2 mL of

desorbing solution in a volumetric flask must be concentrated slightly under a 

gentle stream of nitrogen in order to accommodate the specified volume of the 

spiking solutions. After adding the spiking solutions to the slightly concentrated 

desorbing solution, dilute to the 2-mL mark with toluene or 90/10 

toluene/acetone. 

b. Include a calibration blank of unspiked desorption solution.

c. Analyze together with field samples, field (trip) blanks, and laboratory control samples (step 

12 and 13).

d. Prepare calibration graph (peak area vs. μg analyte), or if internal standard (IS) is used

(peak area of analyte/peak area of IS vs. μg analyte).

10. Prepare Laboratory Control Samples (LCS) with each sample set, in duplicate.

a. Remove cap from large end of sampler tube, apply 30 μL of spiking solution SS-1 (refer to 

Table 13) to face of quartz fiber filter. Cap and allow to stand for a minimum of 1 hour. Preferably, 

these should be prepared as soon as samples arrive and should be stored with the field samples 

until analyzed.

b. Include an unspiked sampler as a media (method) blank.

c. Analyze along with field samples and blanks, and liquid calibration standards (steps 12 through 16).
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 4 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

11. When extending application of this method to other organophosphorous compounds, the following 

minimal desorption efficiency (DE) test may be performed as follows:

a. Determine the NIOSH REL or OSHA PEL, in mg/m3

. 

b. Prepare spiking solution SS-1 (refer to Table 12); or use the following formulae, which are 

specific for the calculation of the weight of analyte to add to 10 mL 90/10 toluene/acetone).

For REL > 1 mg/m3 (assuming 12-L collection vol.), let W = REL x 4 m3

For REL ≤ 1 mg/m3 (assuming 120-L collection vol.), let W = REL x 40 m3

Where W = weight (mg) of analyte to dissolve into 10 mL of desorbing solvent.

Let [SS-1] = W/10 mL where [SS-1] = concentration of spiking solution SS-1 in mg/mL.

c. Prepare spiking solution SS-0.1 by diluting 1 mL of SS-1 to 10 mL in a volumetric flask. Let [SS0.1] = [SS-1] x 0.1 where [SS-0.1] = concentration of spiking solution SS-0.1.

d. Prepare three tubes at each of five levels plus three media blanks. Concentration at each level 

may be calculated using formulae in last entry of Table 12. 

i. Remove plastic cap from large end of sampler, apply appropriate volume of spiking solution to 

face of quartz fiber filter following schedule in Table 13. 

ii. Cap and allow sampler to stand overnight.

e. Prepare tubes for analysis (steps 5 through 8).

f. Analyze with liquid standards (steps 12 and 13).

g. Prepare a graph of desorption efficiency (DE) vs. μg of analyte.

h. Acceptable desorption criteria for 6 replicates is >75% average recovery with a standard deviation 

of < ±9%.

MEASUREMENT:

12. Set gas chromatograph according to manufacturer’s recommendations and to conditions listed in

Table 6 and on page 5600-1. Inject sample aliquot manually using solvent flush technique or with 

autosampler. See Table 7 for retention times of selected analytes.

NOTE: If peak area is greater than the linear range of the working standards, dilute with desorbing 

solution containing internal standard and reanalyze. Apply the appropriate dilution factor in 

calculations.

13. Measure peak area of analyte and of internal standard. 

CALCULATIONS:

14. Determine the mass in μg (corrected for DE) of respective analyte found in the sample front (Wf) and 

back (Wb) sorbent sections, and in the media blank front (Bf) and back (Bb) sorbent sections.

NOTE: The filter is combined with front section. If Wb > Wf/10 report breakthrough and possible 

sample loss.

15. Calculate concentration, C, of analyte in the air volume sampled, V (L): 

CC = �WWff + WW bb − BBff − BBbb�

VV ,

mmmm

mm3

CONFIRMATION:

16. Whenever an analyte is detected, and its identity is uncertain, confirmation may be achieved by 

analysis on a second column of different polarity. If primary analysis was performed using a non-polar 

or weakly polar column (DB-1 or DB-5), confirmation should be accomplished by reanalysis on a polar 

column (DB-1701 or DB-210). See Table 7 for approximate retention times for each column type. Fewer 

analytes co-elute on DB-210 than on DB-1701. Relative retention times are more convenient for the 

identification of unknown analytes. If triphenyl phosphate is not used as the retention time reference 

compound, then another related compound such as tributyl phosphate, Ronnel, or Parathion may be 

substituted.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 5 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

EVALUATION OF METHOD: 

This method was evaluated over the ranges specified in Table 5 at 25 °C using 240-L air samples. Sampler 

tubes were tested at 15% and 80% relative humidity and at 10 °C and 30 °C. In these tests, test 

atmospheres were not generated; instead, analytes were fortified on the face of the sampler filters. This 

was followed by pulling conditioned air at 1 L/min for 4 hours. No difference in sampler performance was 

noted at any of these temperature/humidity combinations. Evaluations of sampler precision and stability 

were conducted at 30 °C and 15% relative humidity. Overall sampling and measurement precisions, bias, 

accuracy, and average percent recovery after long-term storage are presented in Table 5. No breakthrough 

was detected after 12 hours of sampling at 1 L/min with a sampler fortified with the equivalent of 4x the 

NIOSH REL. Malathion and Ronnel were tested at 1/40 x REL, Sulprofos at 1/20 x REL (See Table 5, note 4). 

All criteria [9] were met.

REFERENCES:

[1] NIOSH [1987]. Registry of toxic effects of chemical substances. Sweet DV, ed. Cincinnati, OH: U.S. 

Department of Health and Human Services, Centers for Disease Control, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 87-114.

[2] Budavari, S. ed. [1989]. Merck Index. 11th ed. Rahway, NJ: Merck and Company. 

[3] Sine C, ed. [1991]. Farm chemicals handbook. Willoughby, OH: Meister Publishing Co.

[4] OSHA [1987]. Methods: PV2015, PV2027, PV2037, PV2045, PV2071, PV2087, PV2099, PV2105, 

PV2112, and PV2134. In: OSHA Sampling and analytical methods. Salt Lake City, UT: U.S. Department 

of Labor, Occupational Safety and Health Administration. 

[5] NIOSH [1992]. Recommendations for occupational safety and health. Cincinnati, OH: U.S. 

Department of Health and Human Services, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 92-100.

[6] NIOSH [2016]. NIOSH Pocket guide to chemical hazards. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 2010-168c, http://www.cdc.gov/niosh/npg/.

[7] NIOSH [1977]. Methods: P & CAM 158, 295, 336; S208, S209, S210, S280, S285, S295, S296, S299, and 

S370. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd ed. Cincinnati, OH: U.S. Department 

of Health, Education, and Welfare, Center for Disease Control, National Institute for Occupational 

Safety and Health, DHEW (NIOSH) Publication No. 77-157-C.

[8] NIOSH [1984]. Methods 2503, 2504, 5012, and 5514. In: Eller PM, Cassinelli ME, eds. NIOSH manual of 

analytical methods. 3rd ed. U.S. Department of Health and Human Services, Centers for Disease 

Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 84-100. 

[9] Kennedy ER, Abell MT, Reynolds J, Wickman D [1993]. A sampling and analytical method for the 

simultaneous determination of multiple organophosphorus pesticides in air. Am Ind Hyg Assoc J 

55(12):1172-1177. 

[10] J & W Scientific [1991]. J & W Scientific catalog of high resolution chromatography products. Folsom, 

CA: J & W Scientific.

[11] Hill RH Jr., Arnold JE [1979]. A personal air sampler for pesticides. Arch Environ Contam Toxicol

8:621-628.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 6 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[12] OSHA [1986]. Method 62. In: OSHA Sampling and analytical methods. Salt Lake City, UT: U.S. 

Department of Labor, Occupational Safety and Health Administration. 

[13] NIOSH [1976]. NIOSH criteria for a recommended standard: occupational exposure to malathion. 

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 76-205.

[14] NIOSH [1976]. NIOSH criteria for a recommended standard: occupational exposure to parathion. 

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 76-190.

[15] NIOSH [1976]. NIOSH criteria for a recommended standard: occupational exposure to methyl 

parathion. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

106.

[16] NIOSH [1978]. NIOSH criteria for a recommended standard: occupational exposure during the 

manufacture and formulation of pesticides. Cincinnati, OH: U.S. Department of Health, Education, 

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, DHEW 

(NIOSH) Publication No. 78-174.

[17] Federal Working Group on Pest Management [1974]. Occupational exposure to pesticides: report to 

the federal working group on pest management from the task group on occupational exposure to 

pesticides. Washington, D.C.: Federal Working Group on Pest Management. Available as from 

www.ntis.gov as accession no. PB244259

METHOD WRITTEN BY:

John Reynolds and Donald Wickman, DataChem, Salt Lake City.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 7 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 1. Formula and registry numbers

Compound

(alphabetically)

MW(1)

(Daltons) Empirical Formula CAS No.(2,3,4) RTECS(2)

Azinphos Methyl 317.32 C10H12N3O3PS2 86-50-0 TE1925000

Chlorpyrifos 350.58 C9H11Cl3NO3PS 2921-88-2 TF6300000

Diazinon 304.34 C12H21N2O3PS 33-41-5 TF3325000

Dicrotophos 237.19 C8H16NO5P 141-66-2 TC3850000

Disulfoton 274.39 C8H19O2PS 298-04-4 TD9275000

Ethion 384.46 C9H22O4P2S4 563-12-2 TE4550000

Ethoprop 242.33 C8H19O2PS2 13194-48-4 TE4025000

Fenaminphos 303.36 C13H22NO3PS 22224-92-6 TB3675000

Fonofos 246.32 C10H15OPS2 944-22-9 TA5950000

Malathion 330.35 C10H19O6PS2 121-75-5 WM8400000

Methamidophos 141.12 C2H8O2PS 10265-92-6 TB4970000

Methyl Parathion 263.20 C8H10NO5PS 298-00-0 TG0175000

Mevinphos 224.15 C7H13O6P 7786-34-7 GQ5250000

Monocrotophos 223.17 C7H14NO5P 6923-22-4 TC437500

Parathion 291.26 C10H14NO5PS 56-38-2 TF4550000

Phorate 260.36 C7H17O2PS3 298-02-2 TD9450000

Ronnel 321.54 C8H8Cl3O3PS 299-84-3 TG0525000

Sulprofos 322.43 C12H19O2PS3 35400-43-2 TE4165000

Terbufos 288.42 C9H21O2PS3 13071-79-9 TD7740000

(1) Molecular weights are calculated from the empirical formula using 1979 IUPAC Atomic Weights of the 

Elements

(2) RTECS = NIOSH Registry of Toxic Effects of Chemical Substances [1]

(3) Merck Index [2]

(4) Farm Chemicals Handbook [3]
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 8 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 2. Toxicity and maximum exposure limits

Compound 

(alphabetically) LD50, mg/kg(1) OSHA PEL(3)

(mg/m3) 

NIOSH REL(4)

(mg/m3) 

NIOSH REL(4)

(ppm)

Azinphos Methyl 11 f 0.2 0.2 0.015

Chlorpyrifos 145 0.2 0.2 0.014

Diazinon 250 m, 285 f 0.1 0.1 0.008

Dicrotophos 16 f, 21 m 0.25 0.25 0.026

Disulfoton 2.3 f, 6.8 m 0.1 0.1 0.009

Ethion 27 f, 65 m 0.4 0.4 0.025

Ethoprop 61.5(2) 

Fenamiphos 19.4 0.1 0.1 0.008

Fonofos 3-17(2) 0.1 0.1 0.010

Malathion 1000 15 10 0.740

Methamidophos 25 m, 27 f

Methyl Parathion 14 m, 24 f 0.2 0.2 0.019

Mevinphos 3.7 f, 6.1 m 0.1 skin 0.1 0.01

Monocrotophos 17 m, 20 f 0.25 0.25 0.027

Parathion 3.6 f, 13m 0.1 0.05 skin 0.004

Phorate 1.1 f, 2.3 m 0.05 0.05 skin 0.005

Ronnel 1250 m, 2630 f 15 10 0.760

Sulprofos 227 1 1 0.076

Terbufos 1.6-1.7 m, 1.3-1.57 f

(1) Rat-oral; from Merck Index, unless otherwise noted, f = female, m = male. [2]

(2) Farm Chemicals Handbook [3]

(3) OSHA Final Rule, 1989 (unenforceable, 1992); only Malathion and Parathion had previous PELs

(4) NIOSH Recommendations for Occupational Safety and Health [5]
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 9 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 3. Physical properties (1)

Compound

(Alphabetically)

Liquid 

Density 

(g/mL)

mp (°C) bp

(°C @ 1 atm)

Vapor 

Pressure 

(Pascal)

Vapor 

Pressure 

(mm Hg)

Solubility in 

Water (2,3), % 

by weight

(g/100mL @ 

20 °C)

Azinphos Methyl 1.44 73-74 decomposes 0.024 8x10-9 0.003%

Chlorpyrifos 1.40 41-42 160 

(decomposes) 0.0027 2x10-5 0.0002%

Diazinon 1.116-

1.118 liquid >120 

(decomposes) 0.019 1.4x10-4 0.004%

Dicrotophos 1.216 liquid 400 0.013 0.0001 miscible

Disulfoton 1.144 oil >180 °C flash pt. 0.0267 0.0002 0.003% (22.7 °C)

Ethion 1.220 -12 to -13 >150 

(decomposes) 0.0002 1.5x10-6 0.0001%

Ethoprop 1.094 oil 86-91 0.047 3.5x10-4 0.075%

Fenaminphos 1.15 49 decomposes 0.00012 5x10-5 0.03%

Fonofos 1.16 30 130 0.03 (25 °C) - 0.001%

Malathion 1.23 2.9 60 

(decomposes)(4) 0.005 4x10-5 0.02%

Methamidophos 1.31 44 - 0.002 3x10-4 - 

Methyl 

Parathion 1.358 37-38 <120(decomposes) 0.0002 7.5x10-6 0.006% (25 °C)

Mevinphos 1.25(5) 20.6(5) 300(5) 

(decomposes) 0.4 3x10-3 miscible

Monocrotophos 1.3 53-54(6) 125 0.0003 7x10-6 miscible

Parathion 1.26 6 375 0.005 3.78x10-5 0.001%

Phorate 1.156 liquid 118-120 0.11 8.4x10-4 0.005%

Ronnel 1.49(7) 41 decomposes 0.1 8x10-4 0.004% (25 °C)

Sulprofos 1.20 liquid 210 <0.0001 <10-6 low

Terbufos 1.105 -29.2 88 °C flash pt. 0.0346 2.6x10-4 0.0005

(1) From Merck Index, unless otherwise noted [2].

(2) Farm Chemicals Handbook [3].

(3) NIOSH Pocket Guide [6]. 

(4) NIOSH 4th Edition Method 5012 (EPN, Malathion, Parathion) [8].

(5) NIOSH 4th Edition Method 2503 for Mevinphos [8].

(6) 54-55 °C for pure material, 25-30 °C for commercial mixture. 

(7) NIOSH 2nd Edition Method for Ronnel, S299 [7]
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 10 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 4. Synonyms 

Compound(1)

(alphabetically) Other name(2) CAS name(3,4)

Azinphos Methyl Guthion* O,O-dimethyl S-[(4-oxo-1,2,3-benzotriazin-3(4H)-yl)methyl] phosphorodithioate

Chlorpyrifos Dursban* O,O-diethyl 0-(3,5,6-trichloro-2-pyridinyl) ester

Diazinon Spectracide* O,O-diethyl 0-[6-methyl-2-(1-methylethyl)-4-pyrimidinyl] phosphorodithioate

Dicrotophos Bidrin* 2-dimethyl-cis-2-dimethylcarbamoyl-1-methylvinyl phosphate

Disulfoton Di-Syston* O,O-diethyl S-[2-(ethylthio)ethyl] phosphorodithioate

Ethion S,S'-methylene, O,O,O,'O'-tetraethyl ester, O,O,O'-O'-Tetra ethyl S,S'-methylene

di-phosphorodithioate (4)

Ethoprop Prophos* Phosphorodithioic acid, 0-ethyl S,S-dipropyl ester

Fenaminphos Nemacur*,Phenamiphos (1) Ethyl-3-methyl-4-(methylthio) phenyl (1-methylethyl) phosphoramidate

Fonofos Dyfonate* O-ethyl, S phenyl ethyl phosphorodithioate

Malathion Cythion* S-[1,2 bis(ethoxycarbony1)ethyl] O,O-dimethyl-phosphorodithioate; Diethyl(dimethoxyphosphinothioylthio) succinate

Methamidophos Monitor* Phosphoramidothioic acid, O,S-dimethyl ester

Methyl Parathion Parathion Methyl (1) Phosphorothioic acid, O,O-dimethyl 0-[4-nitrophenyl] ester

Mevinphos Phosdrin* Methyl-3(dimethyoxyphosphinyloxy)but-2-enoate 2-methyoxy carbonyl-1-methylvinyl dimethyl phosphate

Moncrotophos Azodrin* Monocron Dimethyl [1-methyl-3-(methylamino)-3-oxo-1-propenyl] ester(4)

Parathion Ethyl Parathion (1) O,O-diethyl 0-(4-nitrophenyl) phosphorodithioate

Phorate Thimet* Phosphorodithioic acid, O,O-diethyl S-[(ethylthio) methyl] phosphorodithioate

Ronnel Fenchlorphos (1) O,O-dimethyl O-(2,4,5-trichlorophenyl) ester

Sulprofos Bolstar* O-ethyl 0-[4-(methylthio) phenyl] S-propyl phosphorodithioate(4)

Terbufos Counter* O,O-diethyl S-[[(1,1-dimethylethyl) thio] methyl] ester(4)

(1) Common name as given in Farm Chemicals Handbook [3].

(2) *=Trade name (Trademark or Registered Name) as given in Farm Chemicals Handbook [3].

(3) Source, Merck Index [2].

(4) NIOSH RTECS [1] or alternate CAS name in Merck Index [2]. 
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 11 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 5. Method Evaluation (1)

Compound 

(alphabetically)

Range 

Studied(2) 

(mg/m3) 

Range 

Studied(2) 

(mg/sample)

Accuracy Bias 

Average

Overall 

Precision SS�rrrr

Measurement 

Precision SS�rr

% Recovery at 30 

days @ 25 °C

(0 °C)

Azinphos Methyl 0.02-0.4 0.0048-0.096 +0.178 -0.038 0.070 0.030 97 (105)

Chlorpyrifos 0.02-0.4 0.0048-0.096 ±0.163 -0.027 0.068 0.018 92 (90)

Diazinon 0.01-0.2 0.0024-0.048 ±0.162 -0.032 0.065 0.020 94 (93)

Dicrotophos 0.025-0.5 0.006-0.120 ±0.169 -0.037 0.66 0.025 89 (92)

Disulfoton 0.01-0.2 0.0024-0.048 ±0.196 -0.064 0.066 0.024 87 (89)

Ethion 0.04-0.8 0.0096-0.192 ±0.165 -0.29 0.068 0.18 96 (95)

Ethoprop(3) 0.01-0.2 0.0024-0.048 ±0.157 -0.025 0.066 0.024 97 (93)

Fenaminphos 0.01-0.2 0.0024-0.048 ±0.155 -0.029 0.063 0.022 94 (96)

Fonofos 0.01-0.2 0.0024-0.048 ±0.168 -0.036 0.66 0.023 95 (92)

Malathion(4) 0.025-0.5 0.006-0.120 ±0.172 -0.038 0.067 0.019 93 (93)

Methamidophos(5) 0.02-0.4 0.0048-0.096 ±0.156 -0.018 0.069 0.026 88 (95)

Methyl Parathion 0.02-0.4 0.0048-0.096 ±0.160 -0.034 0.063 0.018 95 (95)

Mevinphos 0.01-0.2 0.0024-0.048 ±0.176 -0.42 0.067 0.028 89 (91)

Moncrotophos 0.025-0.5 0.006-0.12 ±0.185 -0.043 0.071 0.026 88 (92)

Parathion 0.005-0.1 0.0012-0.024 ±0.163 -0.012 0.071 0.019 92 (92)

Phorate 0.005-0.1 0.0112-0.024 ±0.202 -0.070 0.066 0.025 91 (91)

Ronnel(4) 0.025-0.5 0.006-0.120 ±0.172 -0.040 0.066 0.018 95 (94)

Sulprofos(4) 0.01-0.2 0.0024-0.048 ±0.181 -0.047 0.067 0.017 94 (94)

Terbufos(3) 0.01-0.2 0.0024-0.048 ±0.188 -0.054 0.067 0.022 92 (91)

(1) Back-up Data Report [9]

(2) The ranges studied were 1/10 to 2x the NIOSH REL (except as noted) using a flowrate of 1 L/min. over 4 hours sampling time.

(3) No NIOSH REL or OSHA PEL available; used 0.1 mg/m3

. 

(4) Malathion and Ronnel were studied at 1/400 to 1/20 the NIOSH REL, Sulprofos at 1/200 to 1/10 the NIOSH REL.

(5) No NIOSH REL or OSHA PEL available; used 0.2 mg/m3

. 

(6) Data in parentheses are for % recovery at 30 days at 0 °C
 ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 12 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 6. Recommended gas chromatographic columns and conditions (1) 

DB-1(2) DB-5(2) DB-1701(2) DB-210(2)

Stationary Phase(3) Polarity Non- Weakly Moderately Moderately 

Length (meters) 30 30 30 30

I.D. (millimeters) 0.32 0.32 0.32 0.32

Film thickness (mm)(4) 0.25 1.0 1.0 0.25

Injection volume (μL)(5) 1 1 1 1 

Injection mode(6) SPL DIR DIR SPL

Initial oven temp (°C) 100 125 125 100

Final oven temp (°C) (6) 275 275 275 250

Recommended max oven temp (°C)(7) 325 325 280 240/260

Oven temp program ( °C/min) 3.0 4.0 4.0 3.0

He Carrier gas head pressure (psi) 15 15 15 15

(1) Actual conditions may vary depending on column and analytical objectives.

(2) Wide Bore Fused Silica Capillary Column 

(3) DB-1, 100% methyl silicone; DB-5, 5% phenyl, methyl silicone; DB-1701, 14% cyanopropylphenyl, 

methyl silicone; DB-210, 50% trifluoropropyl, methyl silicone. DB-1 is non-polar, DB-5 is weakly polar, 

and DB-1701 and DB-210 are moderately strong polar phases. Equivalent phases are acceptable. Other 

phase types may also work well.

(4) Film thickness: Thinner films give faster separations at lower temperatures promoting analyte stability.

(5) Injection (Vol.): Use 2 mm i.d. injection port liners for 0.5 mL injection and 4 mm i.d. injection port liners 

for 1 to 2 mL injections with 0.32 mm i.d. capillary columns.

(6) Injection (mode): SPL = splitless mode, initial oven temp. 5 to 10 °C less than b.p. of desorption solvent; 

DIR = direct mode, initial oven temperature > b.p. of desorption solvent; OC = on-column, sample 

injected within the lumen of the column rather than within the injection port liner. In the splitless and 

direct injection modes, split-vent off time should be 60 seconds for 1 to 2 μL injections with 4 mm i.d. 

injection port liners, and 20 to 30 seconds for 0.5 μL injections with 2 mm i.d. injection port liners.

(7) J & W Scientific Catalog of High Resolution Chromatography Products, p. 21. [10]
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 13 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 7. Approximate retention times (RT) of selected organophosphorous compounds on capillary 

columns (1,2)

Compound (by RT on DB-1) DB-1 RT

(min)

DB-1 

RRT(3)

DB-1 Elution

T, °C(4)

DB-5 RT 

(min)

DB-1701 RT 

(min)

DB-210 RT 

(min)

TEPP 3.71 0.128 111 5.47 7.18(B) 7.88

Triethylphosphorothioate 4.37 0.107 113 6.34 7.14(B) 4.93

Methamidophos 5.12 0.125 115 7.64 13.61 12.03

Dichlorvos 5.81 0.142 117 8.24 10.67 10.54

Mevinphos 10.45 0.256 131 12.92 16.69 19.20

Ethoprop 17.15 0.420 151 19.09 21.52 20.10

Naled 17.61 0.431 153 (6) 23.17(C) 21.46(H)

Dicrotophos 18.00 0.440 154 19.94 25.84(E) 31.43

Monocrotophos 18.27 0.447 155 20.12 28.11 31.60

Sulfotepp 19.06 0.466 157 (6) 23.09(C) 21.11

Phorate 19.18 0.469 158 20.94 23.10(C) 18.92

Dimethoate 19.44 0.476 158 21.84 (6) 29.33(I)

Demeton-S 20.15 0.493 160 21.70 25.06(D) 24.97

Dioxathion 21.30 0.521 164 23.04 26.33(F) 23.46

Fonofos 22.04 0.539 166 23.57 25.87(E) 22.20

Terbufos 22.22 0.544 168 23.80 25.02(D) 21.52(H)

Disulfoton 23.09 0.565 169 24.19 26.43(F) 22.78

Diazinon 23.37 0.572 170 23.75 25.00(D) 20.99

Methyl Parathion 25.37 0.621 176 26.48 31.37 33.21

Oxydemeton Methyl 26(5) 0.63(5) 179 (6) (6) (6)

Ronnel 26.86 0.657 181 27.39 29.30 26.27

Pirimiphos Methyl 28.13 0.688 184 27.90 29.72 26.77

Malathion 28.53 0.698 186 28.33 31.78(G) 33.08(J)

Fenthion 28.74 0.703 186 28.93 31.78(G) 29.35(I)

Parathion 28.98 0.709 187 29.10(A) 33.28 35.60

Chlorpyrifos 29.11 0.712 187 29.10(A) 30.79 27.72

Crufomate 29.64 0.725 189 29.54 34.00 35.34

Isofenphos 31.91 0.780 196 31.17 33.81 33.02(J)

Tetrachlorvinphos 33.26 0.814 200 32.60 35.96 37.01

Fenamiphos 34.09 0.834 202 33.03 37.14 38.95

Merphos 35.19 0.861 206 (6) 30.57 23.89

Fensulfothion 36.61 0.896 210 35.78 42.41 46.98

Ethion 37.88 0.927 214 36.30 39.30 37.96

Sulprofos 38.49 0.942 216 36.96 39.54 37.11

Triphenyl Phosphate 40.88 1.000 223 39.06 (6) (6)

EPN 42.64 1.043 228 41.06 47.83 47.13

Azinphos Methyl 44.16 1.080 232 43.67 (7) 49.24

Leptophos 45.12 1.104 235 43.91 47.38 41.68

Azinphos Ethyl 46.55 1.139 240 46.50 47.43 50.40

Coumaphos 49.31 1.206 248 50.10 67.86 60.88

(1) Actual retention times (RT) will vary with individual columns and chromatographic conditions. See Table 10 for

chromatographic performance notes. Data from Backup Data Report [9].

(2) Capillary Column conditions given in Table 6. Sets of co-eluting or nearly co-eluting peaks are identified by

letters: (A), (B), (C), (D), (E), (F), (G), (H), (I), and (J)

(3) Relative Retention Times, relative to Triphenyl Phosphate.

(4) Elution temperature (°C) for DB-1 column (see Table 6 for column conditions.)

(5) Broad, tailing peak.

(6) No data.

(7) Did not elute.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 14 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 8. Applicable working range

Compound Atmospheric 

working range 

(mg/m3)(1)

Atmospheric 

working range 

(ppm)(2)

Sample working 

range 

(μg/sample)(3)

Instrument 

working range (ng 

on column)(4)

Azinphos Methyl 0.02-0.6 0.0015-0.046 2.4 to 72 1.2-36

Chlorpyrifos 0.02-0.6 0.0014-0.042 2.4 to 72 1.2-36

Diazinon 0.01-0.3 0.0008-0.024 1.2 to 36 0.6-18

Dicrotophos 0.025-0.75 0.0026-0.077 3.0 to 90 1.5-45

Disulfoton 0.01-0.3 0.0009-0.027 1.2 to 36 0.6-18

Ethion 0.04-1.2 0.0025-0.076 4.8 to 144 2.4-72

Ethoprop 0.01-0.3 0.0010-0.030 1.2 to 36 0.6-18

Fenamiphos 0.01-0.3 0.0008-0.024 1.2 to 36 0.6-18

Fonofos 0.01-0.3 0.0010-0.030 1.2 to 36 0.6-18

Malathion 1.0-30 0.074-2.2 12. to 360(5) 6.-180(5)

Methamidophos 0.02-0.6 0.0035-0.10 2.4 to 72 1.2-36

Methyl Parathion 0.02-0.6 0.0019-0.056 2.4 to 72 1.2-36

Mevinphos 0.01-0.3 0.0011-0.033 1.2 to 36 0.6-18

Monocrotophos 0.025-0.75 0.0027-0.082 3.0 to 90 1.5-45

Parathion 0.005-0.15 0.0004-0.013 0.6 to 18 0.3-9 

Phorate 0.005-0.15 0.0005-0.014 0.6 to 18 0.3-9 

Ronnel 1.0-30 0.076-2.3 12. to 360(5) 6.-180(5)

Sulprofos 0.1-3.0 0.0076-0.23 12. to 360 6.-180

Terbufos 0.01-0.3 0.0008-0.026 1.2 to 36 0.6-18

(1) To cover range of 1/10 to 3x NIOSH REL.

(2) Calculated for 25 °C and 760 mm Hg (NTP).

(3) Calculated for a collection volume of 120 L (2 hrs @ 1 L/min., 4 hrs @ 0.5 L/min., or 10 hrs @ 0.2 L/min.).

(4) Desorbing sample in 2.0 mL solvent and injecting 1 μL into gas chromatograph.

(5) Calculated for a collection volume of 12 L (12 min. @ 1 L/min., 24 min. @ 0.5 L/min., or 1 hour @ 0.2 L/min.).
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 15 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 9. Limits of detection and margin of sensitivity

Compound Instrument 

Estimated 

LOD (ng on 

colum)

Sample 

estimated LOD 

(μg/sample)(1)

Atmospheric 

estimated LOD 

(mg/m3)

(1)

Margin of 

sensitivity 

(REL/LOD)(2)

Azinphos Methyl 0.06 0.12 0.0012 167

Chlorpyrifos 0.02 0.04 0.0004 500

Diazinon 0.02 0.04 0.0004 250

Dicrotophos 0.1 0.2 0.002 125

Disulfoton 0.02 0.04 0.0004 250

Ethion 0.02 0.04 0.0004 1000

Ethoprop 0.02 0.04 0.0004 (3) 

Fenamiphos 0.07 0.14 0.0014 71

Fonofos 0.02 0.04 0.0004 250

Malathion 0.05 0.1 0.001 10000

Methamidophos 0.3 0.6 0.005 (3) 

Methyl Parathion 0.02 0.04 0.0004 500

Mevinphos 0.06 0.12 0.0012 83

Monocrotophos 0.2 0.4 0.004 63

Parathion 0.02 0.04 0.0004 125

Phorate 0.02 0.04 0.0004 125

Ronnel 0.02 0.04 0.0004 25000

Sulprofos 0.03 0.06 0.0005 2000

Terbufos 0.02 0.04 0.0004 (3) 

(1) Calculated for a collection volume of 120 L (2 hrs @ 1 L/min., 4 hrs @ 0.5 L/min., or 10 hrs @ 0.2 L/min.).

(2) REL in mg/m3 (Table 2) ÷ Atmospheric LOD (Table 9).

(3) No REL
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 16 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 10. Notes on analytical characteristics of organophosphorous compounds(1,2)

Compound (alphabetically) A. Chemical 

and Physical

B. Desorption 

and Dissolution

C. Gas 

Chromatography 

Azinphos Methyl (Guthion*) - - 3,5,6

Azinphos Ethyl (Guthion Ethyl) - - 5 

Chlorpyrifos (Dursban*) - - - 

Coumaphos (Co-Ral*) - - 5 

Crufomate (Ruelene*) 1 1,4 1 

Demeton (Systox*) 2,6 5 3 

Diazinon (Spectracide*) - - - 

Dichlorvos (DDVP, Vapona*) 7 - 4 

Dicrotophos (Bidrin*) - - - 

Dimethoate (Cygon*) 1 1,4 1 

Dioxathion (Delnav*) - - - 

Disulfoton (Di-Syston*) 2 - 2 

EPN (Santox*) - - 5 

Ethion - - - 

Ethoprop (Prophos*) - - - 

Fenamiphos (Nemacur*) 1 1,4 1 

Fensulfothion (Dasanit*) 3 4 - 

Fenthion (Baytex*) - 5 - 

Fonofos(Dyfonate*) - - - 

Isofenphos(Oftanol*) 1 1 1 

Leptophos (Phosvel*) - 5 5 

Malathion (Cythion*) - 1,3,4 - 

Merphos (Folex*) 4 2 

Methamidophos(Monitor*) 1 1,2,4 1,4

Methyl Parathion (Parathion Methyl) - 5 - 

Mevinphos(Phosdrin*) 6,7 1,5 3,4

Monocrotophos (Azodrin*) 1 - 1 

Naled (Dibrom*) 5 - 2 

Oxydemeton Methyl (Metasystox-R) 3 - 1,2

Parathion (Ethyl Parathion) - - - 

Phorate (Thimet*) 2,7 - 2 

Pirimiphos Methyl (Actellic*) - - 4 

Ronnel (Fenchlorphos) - - - 

Sulfotepp (TEDP) - - - 

Sulprofos (Bolstar*) - - - 

TEPP 7 5 4 

Terbufos(Counter*) 2 - 2 

Tetrachlorvinphos (Gardona*) - - - 

Tributyl Phosphate - - 7 

Triphenyl Phosphate - - 7 

* = Trade name, Registered name, or Trademark (Farm Chemicals Handbook [3]).

(1) Observations made during selection and validation of selected analytes. [9]

(2) Refer to notes on the following pages.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 17 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 10 continued. Notes on Analytical Characteristics

A. Chemical and Physical

1. Amide or phosphoramide, slightly acidic, very polar chemically.

2. Alkyl thio-ether, easily oxidized to sulfone and sulfoxide.

3. Sulfoxides, easily oxidized to sulfone. Also very polar chemically.

4. Phosphite, easily air oxidized to phosphate (Merphos to DEF).

5. Vicinal dibromide, easily debrominated (Naled to Dichlorvos).

6. Two or more isomers commonly exist (e.g. Demeton-O and Demeton-S; cis- and transmevinphos).

7. Relatively volatile, can be lost if media or vials are left uncapped for even a short period of time.

General: Organophosphorous compounds are easily destroyed at mildly alkaline conditions 

(pH ≥8). Loses can occur for trace levels of compounds on alkaline glass surfaces. 

Glassware should be neutralized after washing, if alkaline detergent is used.

B. Desorption and Dissolution

1. Solubility of concentrated solutions in toluene enhanced by the addition of 1% Methanol or 10% 

acetone. Solubility in hexane very unfavorable even for dilute solutions.

2. Changing from 100% toluene to 90/10 toluene/acetone, desorption from glass fiber filters 

improved from 62% to 98%, desorption from quartz fiber filters improved from 30% to 101%.

3. Changing from glass fiber filters to quartz fiber filters, desorption in toluene improved from 16% to 

88% and desorption in 90/10 toluene/acetone improved from 70% to 99%.

4. These compounds are more chemically polar than the other listed organophosphorous 

compounds; desorption from XAD-2 or from glass or quartz fiber filters in hexane was incomplete 

or non-existent. Desorption in toluene was adequate except as noted in 2 and 3 above. The use of 

toluene containing 10% acetone improved recoveries for all analytes to satisfactory levels.

5. The desorption characteristics of these compounds were not evaluated.

General: 1. The presence of acidic hydrogen or double bonded oxygen anywhere in the 

molecular structure greatly decreases solubility in non-polar solvents and increases 

the difficulty of desorption from polar surfaces and sorbents.

2. While glass fiber filters and toluene desorbant were adequate for most compounds in

preliminary tests, the method was given wider application for the more polar 

compounds by the use of quartz fiber filters and 90/10 toluene/acetone desorbant. 

3. Greater flame photometric detector response was observed for organophosphorous

compounds when injections were made in toluene or 90/10 toluene/acetone. 

Solvents with lower boiling points (e.g. methylene chloride, chloroform, methyl-tbutyl ether, and ethyl acetate) possessed fair to good desorption power, but rendered 

less satisfactory gas chromatographic responses for the analytes. This effect may be 

due to better analyte mass-transfer from the injection port to the capillary column 

with higher boiling solvents using splitless or direct injection techniques.

C. Gas Chromatography

1. Poor chromatography may be encountered with dirty or undeactivated columns or injection ports. 

Clean quartz wool plugs stuffed in the injection port liner are better than silanized glass wool at 

reducing losses within the injection port.

2. Multiple, shifted, irregular, or severely tailing peaks may be observed in the chromatogram if 

degradation or oxidation of the analytes occur prior to injection, within the injection port, or 

during chromatographic separation on-column.

3. Multiple peaks may be observed due to presence of isomers.

4. Short elution time, compound may co-elute with solvent if oven temperature is too high.

5. Long elution time, compound may be lost if run time is too short, column or injection port is too 

cool, or split-vent-valve opens too soon when injected in splitless or direct injection mode.

6. Azinphos Methyl did not elute from DB-1701 even though Azinphos ethyl did elute.

7. Potential internal standards: Triphenyl phosphate is more favorable if multiple analytes are 

expected because it is less volatile and elutes in an area of the chromatogram having fewer 

competing analytes.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 18 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 11. Other methods of analysis for organophosphorous compounds in air

Document Method Number Organophosphorous 

Compound(s)

AEC&T(1) Chlorpyrifos, Demeton-O, 

Demeton-S, Diazinon, 

Dimethoate, Malathion, 

Paraoxon, and Parathion

NMAM, 2nd ed (2) v. 1 P&CAM 158

v. 5 P&CAM 295

v. 6 P&CAM 336

v. 3 S 208

v. 3 S 209

v. 3 S 210

v. 6 S 280

v. 3 S 285

v. 3 S 295

v. 6 S 296

v. 6 S 299

v. 3 S 370

Parathion

Dichlorvos (DDVP)

TEPP

Tributyl phosphate

Triorthocresyl phosphate

Triphenyl phosphate

Demeton

EPN

Parathion

Mevinphos

Ronnel

Malathion

NMAM, 4th ed. (3) 2503

2504

5012

5514

Mevinphos

TEPP

EPN, Malathion, and Parathion

Demeton

OSHA A.M.M. (4) 62 Chlorpyrifos, Diazinon, 

Parathion, DDVP, and 

Malathion

OSHA Partially Validated

Methods(5)

PV2087

PV2045

PV2134

PV2015

PV2027

PV2071

PV2099

PV2037

PV2105

PV2112

Azinphos methyl 

Monocrotophos

Coumaphos 

Crufomate 

Fonofos

Pirimiphos Methyl

Dicrotophos

Sulprofos

Disulfoton 

Methyl Parathion

(1) Hill and Arnold in Arch. Environ. Contam. & Toxicol . [11]

(2) NIOSH Manual of Analytical Methods, 2nd ed., Volumes 1-7 [7]

(3) NIOSH Manual of Analytical Methods, 4th ed. [8]

(4) OSHA Analytical Methods Manual [12]

(5) OSHA Partially Validated Methods (refer to by name) [4]
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 19 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 12. Spiking concentrations for organophosphorous compounds

Compound Total μg 

spiked(1) at 

1/30 x REL

Total μg 

spiked(1) at 

1/10 x REL

Total μg 

spiked(1) at 

1/3 x REL

Total μg 

spiked(1) at 

1 x REL

Total μg 

spiked(1) at 

3 x REL

Volume (mL) of 

10 mg/mL stock 

solution needed to 

prepare 10 mL of 

SS-1 

Azinphos Methyl 0.8 2.4 8 24 72 0.8

Chlorpyrifos 0.8 2.4 8 24 72 0.8

Diazinon 0.4 1.2 4 12 36 0.4

Dicrotophos 1.0 3 10 30 90 1.0

Disulfoton 0.4 1.2 4 12 36 0.4

Ethion 1.6 4.8 16 48 144 1.6

Ethoprop 0.4 1.2 4 12 36 0.4

Fenamiphos 0.4 1.2 4 12 36 0.4

Fonofos 0.4 1.2 4 12 36 0.4

Malathion(3) 4 12 40 120 360 4.0

Methamidophos 0.8 2.4 8 24 72 0.8

Methyl Parathion 0.8 2.4 8 24 72 0.8

Mevinphos 0.4 1.2 4 12 36 0.4

Monocrotophos 1.0 3 10 30 90 1.0

Parathion 0.2 0.6 2 6 18 0.2

Phorate 0.2 0.6 2 6 18 0.2

Ronnel(3) 4 12 40 120 360 4.0

Sulprofos 4 12 40 120 360 4.0

Terbufos 0.4 1.2 4 12 36 0.4

General (for 120L)(2,3) x/30 x/10 x/3 x 3x 4y

(1) Total μg per sample, for spiked media, or per 2 mL desorption solution for liquid calibration standards.

(2) Where x, μg/sample = REL, μg/L x 120 L/sample; and y, mg/mL = REL, mg/m3 x 4 m3

/mL.

(3) For all REL > 1 mg/m3

, use 1/10 x REL in the calculations (assumes that collection volume in these cases 

would be 12 L instead of 120 L).

TABLE 13. Spiking of media

Spiking Levels(1)

(fraction of REL)

Spiking 

Solution

Preferred Syringe 

Size Spiking Volume(2)

1/30x REL SS-0.1 50 μL 10 μL

1/10x REL SS-0.1 50 μL 30 μL

1/3x REL SS-1 50 μL 10 μL

1x REL SS-1 50 μL 30 μL

3x REL SS-1 100 μL 90 μL

(1) For a collection volume of 120 L. Range corresponds to values within “Working Range” column, 

Table 8.

(2) For liquid calibration standard preparations, add specified volume to 2 mL desorption solution in 

2-mL volumetric flask. For laboratory control samples spiked at the REL, apply volume specified 

in “Spiking Volume” column to front section of sampler; do in duplicate. For Desorption Efficiency 

determination, apply specified volume to front section of sampler; do each of five levels in 

triplicate.
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 20 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 1. Structures of organophosphorus compounds
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 21 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 2. OVS-2 Sampler
ORGANOPHOSPHORUS PESTICIDES: METHOD 5600, Issue 2, dated 2 March 2016 - Page 22 of 22

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 3. Typical chromatogram of organophosphorus compounds

Column: DB-1 fused silica capillary column, 30 meters x 0.32 mm I.D. x 0.25 μm film thickness

Temperature program: 80 to 275 °C at 3.0 °C per minute. 

Concentration of analytes: 0.6 x NIOSH REL, except Sulprofos (0.06x REL), and Malathion and Ronnel (both at 0.006x)

1.8 μg/mL Parathion and Phorate

3.6 μg/mL All other compounds

7.2 μg/mL Azinphos methyl, Chlorpyrifos, Methamidophos, and Methyl parathion

9.0 μg/mL Dicrotophos and Monocrotophos

14 μg/mL Triphenyl phosphate

14.4 μg/mL Ethion
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ORGANONITROGEN PESTICIDES 5601

Formula: Figure 1 MW: Table 1 CAS: Table 1 RTECS: Table 1 

METHOD: 5601, Issue 2 EVALUATION: FULL Issue 1: 15 January 1998

Issue 2: 25 February 2016

OSHA: Table 1

NIOSH: Table 1

PROPERTIES: Table 1 

SYNONYMS: Aldicarb, Benomyl, Captan, Carbaryl, Carbendazim, Carbofuran, Chlorprophan, Diuron, Formetanate, Methiocarb, 

Methomyl, Oxamyl, Propham, Propoxur, Thiobencarb (Table 1)

SAMPLING

SAMPLER: FILTER/SOLID SORBENT TUBE 

(OVS-2 Tube: 13-mm quartz fiber filter; 

XAD-2, 270 mg/140 mg) 

FLOW RATE: 0.1 - 1 L/min [1]

VOL-MIN: variable (see Table 2)

 -MAX: 480 L

SHIPMENT: routine

SAMPLE at least 30 days @ -12 °C [1]

STABILITY: at least 7 days @ 24 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: Table 2 

BIAS: Table 2 

OVERALL

PRECISION (SS�rrrr): Table 2 

ACCURACY: Table 2

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION

ANALYTE: organonitrogen pesticides (Table 1)

EXTRACTION: 2 mL extraction solution (0.2% V/V 0.1 M 

aqueous triethylamine phosphate buffer 

in acetonitrile, pH 6.9 to 7.1)

INJECTION

 VOLUME: 5 μL

MOBILE

PHASE A: 2% 1-propanol in aqueous 0.02 M 

triethylamine phosphate (pH 6.9 to 7.1)

MOBILE

PHASE B: 2% 1-propanol in acetonitrile

PROGRAM: mobile phase B, 3 to 95% in 30 min, hold 

95% 5 min

COLUMN: C18, 30 cm x 3.9-mm ID; ambient 

temperature (Table 3) 

DETECTOR: UV absorption at 200 and 225 nm

CALIBRATION: solutions of analytes in extraction fluid

RANGE: Table 2

ESTIMATED LOD: Table 2

PRECISION (SSrr): Table 2 ̅

APPLICABILITY: The working ranges (Table 2) for aldicarb, carbofuran, and oxamyl range from 0.5 to 10 times the OSHA PEL. 

Others cover 0.1 to 2 times the OSHA PELs with appropriate dilutions [1].This method may be applicable to the determination of 

other organonitrogen compounds after evaluation, and to a broad range of pesticides having UV chromophores, e.g., acetanilides, 

acid herbicides, organophosphates, phenols, pyrethroids, sulfonyl ureas, sulfonamides, triazines, and uracil pesticides.

INTERFERENCES: Because of the broad response of the UV detector at shorter wavelengths, there are many potential 

interferences. Those tested include solvents (chloroform and toluene), antioxidants (BHT), plasticizers (dialkylphthalates), nitrogen 

compounds (nicotine and caffeine), impurities in HPLC reagents (e.g., in triethylamine), other pesticides (2,4-D, atrazine, parathion, 

etc.), and pesticide hydrolysis products (1-naphthol). Retention times are given in Table 4. Confirmation techniques are 

recommended when analyte identity is uncertain.

OTHER METHODS: This method may be used to replace previous related pesticide methods: S273 Carbaryl [2]; 5006 Carbaryl [3]; 

OSHA 63 Carbaryl and 74 Aldicarb [4]; OSHA Stopgap methods for several pesticides [5]; and EPA TO-10 for Captan, Folpet, and 

Mexacarbate [6]. The OVS-2 Tube is similar in concept to the device of Hill and Arnold [7], but offers convenience and lower flow 

resistance.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 2 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Carbamate, urea, and sulfenimide analytes 

listed in Table 1; internal standards 

acetanilide and acetophenone, analytical 

grade.*

2. Acetonitrile, UV grade.*

3. Methanol, HPLC grade.*

4. Deionized water, ASTM Type II.

5. 1-Propanol, UV grade.*

6. n-Butyl isocyanate.*

7. Triethylamine (TEA), HPLC grade.* Keep 

refrigerated (0 to 4 °C) and store under 

nitrogen for longer shelf life [1, 8].

8. Ortho-phosphoric acid, >85% by weight, 

ACS grade or better.*

9. Extraction solution. Prepare separate 

trimethylamine phosphate (TEA-PO4) 

preservative and internal standard 

solutions.

a. TEA-PO4 preservative, 0.1 M. Dissolve1.4 

mL of TEA in 90 mL of deionized water. 

Add phosphoric acid to lower pH to 7.0 

(± 0.1) as indicated by a calibrated pH 

meter. Bring volume to 100 mL. Keep 

tightly capped and refrigerated. 

NOTE: Do not use chloroacetic acid as a 

preservative [9]. Formetanate, at 

least, is unstable with chloroacetic 

acid.

b. Internal standard stock solution, 

5 mg/mL. Add 100 mg of each internal 

standard of choice for each 20 mL of 

solution required. Dissolve in 

acetonitrile. Store capped at –12 ± 1 °C.

c. Final extraction solution. Add 1 mL of the 

TEA-PO4 solution and 12 mL of the 

internal standard stock solution to a 500-

mL volumetric flask. Dilute to volume 

with acetonitrile. Concentration of TEA = 

0.2 mM, water = 0.2%, and internal 

standards = 120 μg/mL. Stable up to 30 

days at 0 to 4 °C.

10. Individual analyte stock solutions, 5 

mg/mL. Add each analyte to acetonitrile in 

separate volumetric flasks. Use methylene 

chloride for benomyl and carbendazim. Use 

50/50 v/v methanol/acetonitrile or formetanate. 

Store at –12 ± 1 °C. (Solutions are stable up to 30 

days.)

EQUIPMENT

1. Sampler: OSHA Versatile Sampler (OVS-2 

tube), resin filled sampling tube; glass tube, 

11-mm i.d. x 13-mm o.d. x 50-mm long, with 

the outlet end drawn to a 6-mm o.d. x 25-

mm long tube. The enlarged part of the tube 

contains a 270-mg front section of 20/60 

mesh XAD-2 sorbent held in place by a 9-10-

mm o.d. quartz fiber filter and 

polytetrafluoroethylene (PTFE) retaining ring. 

The front section is separated from the back 

section of 140 mg XAD-2 sorbent with a 

short plug of polyurethane foam. The back 

section is held in place by a long plug of 

polyurethane foam. The tube is available 

commercially. OVS-2 tubes with glass fiber 

filters have equivalent desorption efficiencies 

and are commercially available.

2. Personal sampling pump: 0.1 to 1 L/min with 

flexible and inert connecting tubing.

3. High Performance Liquid Chromatograph 

capable of mixing two mobile phases in a 

linear gradient. Must be capable of pumping 

up to 4000 psi, to accommodate 300 mm 

long columns.

4. Autosampler, low dead-volume, capable of 

5-μL injections. Preservative (TEA-PO4) in the 

desorbing solution may be eliminated if a 

refrigerated autosampler tray is available**

5. Analytical columns.

a. Primary column: Base-deactivated 

octadecylsilyl (C18) column, 3.9-mm ID X 

300 mm, 5-μm particle size.

b. Secondary column: Cyanopropyl silica 

column, 4.6 X 250-mm, 5-μm particle size.

6. Guard column, low dead-volume, containing 

analytical column packing material**

7. Ultraviolet detector, low dead-volume, with 

1-cm path length cell capable of monitoring 

two wavelengths (200- and 225-nm) 

simultaneously.

8. Vials: 4-mL with PTFE-lined caps; 2-mL HPLC 

autosampler vials with PTFE- or 

polyethylene-lined snap caps.

9. Syringes: 0.01-, 0.05-, 0.1-, 1.0- and 2.5-mL;

luer lock style, 1- or 2.5-mL for sample 

filtering.

10. Volumetric flasks: 2-, 5-, 10-, 25-, 50-, 100-, 

500-, and 1000-mL.

11. PTFE syringe filter: 0.45-μm. 
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 3 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS: (continued)

11. Calibration stock solution. Combine stock 

solutions of analytes of interest in a 

volumetric flask to produce the highest 

concentration standard (suggest 120 to 480 

μg/mL).

NOTE: Do not combine benomyl and 

carbendazim in the same standard 

solution. [10-12]. See APPENDIX.

12. Quality control spiking solutions: Add 

analyte stock solutions to acetonitrile at 

concentrations in the analytical range of the 

samples. Store in the freezer at –12 ± 1 °C 

until immediately before spiking.

NOTE: Spiking solutions must not contain 

internal standard.

13. Mobile phase A. Combine 20 mL of 1-

propanol and 2.8 mL of TEA in a 1-L 

volumetric flask and bring to volume with 

deionized water. Adjust pH to 7.0 (± 0.1) 

with phosphoric acid using a pH meter. 

Final concentrations: 2% 1-Propanol, 0.02 M 

TEA-PO4. Degas prior to use.

14. Mobile phase B. Add 20 mL of 1-propanol to 

acetonitrile in a 1-L volumetric flask and 

bring to volume. Degas prior to use.

*See SPECIAL PRECAUTIONS.

EQUIPMENT: (continued)

12. Forceps.

13. Small vial/tube tumbler capable of 5 to 10 

RPM.

14. pH meter.

15. Graduated cylinders, 10-mL, 25-mL.

16. Pippettes, glass, disposable.

**Low dead volumes will give lower dwell 

volumes [13, 14] and better resolution [15]

SPECIAL PRECAUTIONS: Pesticides: Avoid inhaling vapors or dust; avoid skin contact. Wear gloves and 

suitable clothing when handling pure material. Solvents: Avoid skin contact and open flame. Use in a 

hood. Phosphoric acid: Avoid skin contact. n-Butyl isocyanate may act as a sensitizer. Avoid skin contact.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Connect the sampler to the personal sampling pump with flexible tubing. Place sampler vertically, 

with the large end down, in the worker’s breathing zone.

3. Sample at an accurately known flow rate between 0.1 and 1 L/min for a total sampling volume up to 

480 L. Record volume, and document presence of any known or potential interferences.

4. Cap both ends of the sampler with plastic caps and pack securely for shipment. 

CALIBRATION AND QUALITY CONTROL 

5. Remove cap from large end. Transfer filter, PTFE retainer ring, and front resin section to a 4-mL vial. 

Transfer the polyurethane foam divider plug along with the back-up resin bed to a second 4-mL vial.

6. Add 2.0 mL of desorbing solvent with internal standards to each vial using a 2.5- or 5-mL syringe or 

2-mL pipette; cap each vial.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 4 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

7. Mix by rotating the vials end-over-end at 5 to 10 RPM for approximately 45 minutes.

8. Filter an aliquot into a 2-mL autosampler vial through a 0.45-μm PTFE filter.

CALIBRATION AND QUALITY CONTROL 

9. Determine retention times for the analytes of interest using the column and chromatographic 

conditions of choice for the analysis.

10. Calibrate daily with at least six working standards covering the analytical range for individual analytes.

a. Prepare working standards by diluting aliquots of the high-level calibration standard with 

desorbing solution containing internal standard in a volumetric flask. Include an unspiked 

desorption solution calibration blank.

b. Filter aliquots of standards and blanks for analysis (See Step 8).

c. Analyze together with samples, blanks, and laboratory control samples (Steps 12 through 14).

d. Prepare a calibration graph (ratio of peak area of analyte over peak area of internal standard vs. μg 

analyte).

NOTE: Use of an internal standard is recommended [1, 6], but optional if the precision of the 

injection device and HPLC system are known to be adequate.

11. Prepare desorption efficiency (DE) samples and Laboratory Control Samples (LCS) with each sample 

set at a rate of 10% of samples.

a. Remove cap and the PTFE retainer ring from large end of sampler tube (to prevent wicking behind 

the ring). Apply known volume of calibration solution to face of quartz fiber filter.

NOTE: Spike no more than 15 to 30 μL at a time. If more needs to be applied, connect the sampler 

to a vacuum pump with a flow ≤1 L/min, then apply spiking solution in 15- to 30-μL 

aliquots. Allow several minutes for the solvent to evaporate between each aliquot, to 

prevent wicking along the sides of the tube into the back-up section (5% or more may 

deposit on the walls of the tube).

b. Cap and allow to stand a minimum of one hour.

NOTE: Prepare LCS when samples arrive and store with field samples until analyzed.

c. Include an unspiked sampler as a media (method) blank.

d. Analyze with the field samples, blanks, and the liquid standards (Steps 12 through 14). 

MEASUREMENT:

12. Set liquid chromatograph according to manufacturer’s recommendations and to conditions listed in 

Table 3. Select two wavelengths for detection with 200 and 225 nm for general-purpose screens. For 

selected analytes, chose a more specific wavelength UV spectra where available.

13. Inject sample aliquot with autosampler. See Table 4 for approximate retention times of selected 

analytes.

NOTE: If peak area is greater than the area of the highest standard, dilute with desorbing solution 

containing internal standards and reanalyze. Apply the appropriate dilution factor in 

calculations.

14. Measure peak area of analyte(s) and internal standard(s). Divide peak area of analyte by peak area of 

internal standard on same chromatogram. 

CALCULATIONS:

15. Determine the mass, μg, (corrected for DE) of analyte found in the sample filter and front sorbent 

section (Wf), back sorbent section (Wb), and the media blank front (Bf ) and back (Bb) sorbent sections 

from a standard curve.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 5 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

16. Calculate concentration, C (mg/m3), of each analyte in the air volume sampled, V (L)

CC = WWff + WWbb − BBff − BBbb

VV , mmmm mm3 ⁄

NOTE: μg/mL ≅ mg/m3

CONFIRMATION:

Retention Times with Alternate Conditions. Whenever the identity of an analyte is uncertain, 

confirmation may be achieved by analysis on an alternate column. If primary analysis was performed on a 

base-deactivated octadecylsilyl (C18) column, identity may confirmed by reanalysis on a cyanopropyl silica 

column, or by changing to a water/methanol mobile phase (see Table 9 for recommended alternative 

conditions). See Table 4 for approximate retention times for each column type and condition. Relative 

retention times (retention indices for a particular set of conditions) are more convenient for the 

identification of unknown analytes.

UV or Mass Spectra. Confirmation may be achieved through comparison of unknown spectra with 

reference spectra where available. Relative response ratios will give a moderate level of confirmation. 

Some analytes (O-aryl carbamates especially) can be confirmed by GC/MS using highly deactivated 

injection ports and analytical GC columns, or by HPLC/MS.

EVALUATION OF METHOD:

This method was evaluated over the ranges specified in Table 2 at 25 °C with 240-L air samples. Samplers 

were tested at 15 and 80% relative humidity and 10 and 30 °C. In these experiments, test atmospheres 

were not generated; instead, analytes were fortified on the face of the sampler filters. The conditioned air 

was pulled through the samplers at 1 L/min for four hours. No significant difference in sampler 

performance was noted at any of these temperature/humidity combinations. Evaluation of sampler

precision and stability was conducted at ambient conditions of temperature and relative humidity. Overall 

sampling and measurement precision, bias, accuracy, and average percent recovery after long-term 

storage are presented in Table 2. No breakthrough was detected with samplers fortified with 480 μg per 

analyte per tube after sampling eight hours at 1 L/min. For the estimation of LOD/LOQ, a series of mediaspiked standards were prepared in triplicate, analyzed, and responses fitted to a quadratic curve. The Limit 

of Detection (LOD) and Limit of Quantitation (LOQ), given in Table 2, were estimated according to NIOSH 

SOP 018 [1,17]. Criteria established by NIOSH were met [1].

REFERENCES:

[1] NIOSH [1995]. Back-up data report: carbamate, urea, and sulfenimide pesticides. Unpublished.

[2] NIOSH [1977]. Carbaryl: Method S273. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd 

ed. (Vol 3) Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH), Publication No. 77-

157-C.

[3] NIOSH [1994]. Method 5006. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical methods. 4th 

ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and 

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-

113.

[4] OSHA [1985]. OSHA Methods 63 and 74. In: OSHA analytical methods manual. Salt Lake City, UT: 

Carcinogen and Pesticide Branch, OSHA Analytical Laboratory.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 6 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[5] OSHA. OSHA Stopgap methods for individual pesticides. Salt Lake City, UT: Carcinogen and Pesticide 

Branch, OSHA Analytical Laboratory.

[6] EPA [1986]. EPA compendium of methods for the determination of toxic organic compounds in 

ambient air. US EPA, Publication No. EPA/600/4-87/006.

[7] Hill RH Jr., Arnold JE [1979]. A personal air sampler for pesticides. Arch Environ Contam Toxicol

8:621-628.

[8] Dolan JW [1993]. LC troubleshooting. LC-GC 11(7):500.

[9] EPA [1988]. EPA Method 531.1 (Rev. 3.0). In: Methods for the determination of organic compounds 

in drinking water. US EPA. Publication No. EPA 600 4-88 039.

[10] Chibia, Mikio, Doornbos F [1974]. Instability of benomyl in various conditions. Bulletin Environ 

Contami Toxicol 11(3):273-274.

[11] Calmon JP, Sayag DR [1976]. Kinetics and mechanism of conversion of methyl 1-(butylcarbamoyl)-2-

benzimidazolecarbamate(benomyl) to methyl 2-benzimidazolecarbamate (MBC). J Agric Food Chem 

24(2):311-314.

[12] Calmon JP, Sayag DR [1993]. Instability of methyl 1-(butylcarbamoyl)-2-benzimidazolecarbamate 

(benomyl) in various solvents. J Agric Food Chem 24(2):426-428.

[13] Dolan JW [1993]. LC troubleshooting. LC-GC 11(12):858-860.

[14] Dolan JW [1993]. LC troubleshooting. LC-GC 11(6):412-314.

[15] Dolan JW [1993]. LC troubleshooting. LC-GC 12(4):298.

[16] Gere DR [1993]. Column watch. LC-GC 11(10):710-712.

[17] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. By 

Kennedy ER, Fischbach TJ, Song R, Eller PM, Schulman SA. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 95-117.

[18] NIOSH [1987]. Registry of toxic effects of chemical substances. U.S. Department of Health and 

Human Services, Centers for Disease Control, National Institute for Occupational Safety and Health, 

DHHS (NIOSH) Publication No. 87-114.

[19] Budavari S, ed. [1989]. Merck Index. 11th ed. Rahway, NJ: Merck & Co., Inc. 

[20] Meister [1995]. Farm Chemicals Handbook. Willoughby, OH: Meister Publishing Co.

[21] Wise SA, May WE [1983]. Effect of C18 surface coverage on selectivity in reversed-phase liquid 

chromatography of polycyclic aromatic hydrocarbons. Anal Chem 55(9):1479-1485. 

[22] Cole LA, Dorsey JG [1992]. Temperature dependence of retention in reversed-phase liquid 

chromatography. 1. Stationary-phase considerations. Anal Chem 64(13):1317-1323.

[23] Wirth MJ [1994]. Column watch. LC-GC 12(9):656-664.

[24] Cole LA, Dorsey JG [1990]. Reduction of reequilibration time following gradient elution reversedphase liquid chromatography. Anal Chem 62(1):16-21.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 7 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[25] Foley JP, May WE [1987]. Optimization of secondary chemical equilibria in liquid chromatography: 

variables influencing the self-selectivity, retention, and efficiency in acid-base systems. Anal Chem 

59(1):110-115.

[26] Dolan JM [1993]. LC troubleshooting. LC-GC 11(2):94.

[27] Roos RW, Lau-Cam CA [1986]. General reversed-phase high-performance liquid chromatographic 

method for the separation of drugs using triethylamine as a competing base. J Chrom 370:403-418.

[28] Kirkland JJ, Boyes BE, DeStefano JJ [1994]. Changing band spacing in reversed-phase HPLC. Am Lab 

Sept:36-42.

[29] Seaver, Sadek P, Sadek C [1994]. LC troubleshooting. LC-GC 12(10):742-746.

[30] Zweig, Gunter, Ru-yu G [1983]. Determination of benomyl by reversed-phase liquid 

chromatography. Anal Chem 55(8):1448-1451.

[31] Dolan, JW [1992]. LC troubleshooting. LC-GC 10(10):746.

[32] Evans CE, Victoria L, McGuffin [1991]. Direct examination of the injection process in liquid 

chromatographic separations. Anal Chem 63(14):1393-1402.

[33] Steffeck RJ, Woo SL, Weigand RJ, Anderson JM [1995]. A comparison of silica-based C18 and C8 HPLC 

columns to aid column selection. LC-GC 13(9):720-726Body Text. 

METHOD REVISED BY:

Jun-Jie Lin, MSPH, and John M. Reynolds, DataChem Laboratories, Salt Lake City, UT.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 8 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 1. General Information

Name/Synonym Formula MW Properties Solubility in

H2O (g/L)

Exposure Limit 

(mg/m3) 

Aldicarb

CAS# 116-06-3 

RTECS UE2275000

C7H14N2O2S 190.3

MP 99-100 °C; vp 3.9 mPa

(2.9x10-5 mm Hg) @ 25 °C;

LD50 1 mg/kg

6 @ 25 °C

Benomyl 

CAS# 17804-35-2 

RTECS DD6475000

C14H18N4O3 290.36 MP decomposes; vp <1.3 mPa 

(<1x10-5 mm Hg) @ 20 °C; LD50

>9590 mg/kg

0.002 @ 25 °C OSHA 5.0 (resp)

Captan

CAS# 133-06-2 

RTECS GW5075000

C9H8Cl3NO2S 300.6 MP 178 °C; vp <1.3 mPa (<1x10-5 

mm Hg) @ 20 °C; LD50 9000 

mg/kg

<0.005 @ ~25 °C NIOSH 5

Carbaryl 

CAS# 63-25-2 

RTECS FC5950000

C12H11NO2 201.24 MP 142 °C; vp <5.3 mPa (<4x10-5 

mm Hg) @ 25 °C; LD50 250 mg/kg 0.12 @ 30 °C

OSHA 5 

NIOSH 5

Carbendazim 

CAS# 10605-21-7 

RTECS DD6500000

C9H9N3O2 191.21

MP 302-307 °C (decomposes); 

LD50 6400 mg/kg

0.008/pH 7 

@ 24 °C

Carbofuran 

CAS# 1563-66-2 

RTECS FB9450000

C12H15NO3 221.28

MP 150-153 °C; vp 0.031 mPa 

(2.3x10-7 mm Hg) @ 20 °C; LD50 5.3 

mg/kg

0.70 @ 25 °C NIOSH 0.1

Chlorpropham CAS# 

101-21-3 

RTECS FD8050000

C10H12ClNO2 213.68

MP 40.7-41.1 °C; vp 2.7 mPa 

(2x10-

5 mm Hg) @ 33 °C; LD50

1200 mg/kg

slightly soluble

Diuron

CAS# 330-54-1 

RTECS YS8925000

C9H10Cl2N2O 233.11

MP 158-159 °C; vp 0.41 mPa 

(3.1x10-

6 mm Hg) @ 50 °C; LD50

437 mg/kg

0.042 @ 25 °C NIOSH 10

Formetanate.HCl CAS# 

23422-53-9 

RTECS FC2800000

C11H16ClN3O2 257.75

MP 200-202 °C (decomposes); 

LD50 20 mg/kg

>50% as 

hydrochloride

Methiocarb

CAS# 2032-65-7 

RTECS FC5775000

C11H15NO2S 225.34 MP 121.5 °C; vp 0.036 mPa

(2.7x10-7 mm Hg) @ 25 °C; LD50 

60 mg/kg

insoluble

Methomyl

CAS# 16752-77-5 

RTECS AK2975000

C5H10N2O2S 162.24

MP 78-79 °C; vp 6.7mPa (5x10-

5

mm Hg) @ 25 °C; LD50 17 mg/kg 58 @ 25 °C NIOSH 2.5

Oxamyl

CAS# 23135-22-0 

RTECS RP2300000

C7H13N3O3S 219.3

MP 100-102 °C; vp 31 mPa 

(2.4x10-

4 mm Hg) @ 20 C; LD50 5 

mg/kg

280 @ 25 °C

Propham

CAS# 122-42-9 

RTECS FD9100000

C10H13NO2 179.24 MP 90 °C; vp 18 mPa (1.35x10-4

mm Hg); LD50 50 mg/kg 0.25 @ 25 °C

Propoxur

CAS# 114-26-1 RTECS 

FC3150000

C11H15NO3 209.27 MP 91.5 °C; vp 1.3 mPa

(9.75 mm Hg) @ 20 C; LD50 83 

mg/kg

2 @ 20 °C NIOSH 0.5

Thiobencarb CAS# 

28249-77-6 

RTECS EZ7260000

C12H16ClNOS 257.81 Not available; LD50 1130 mg/kg ~0.03 @20 °C

Abbreviations: MW=molecular weight (Daltons); RTECS=Registry of Toxic Effects of Chemical Substances 

[18]; LD50=lethal dose 50% [19,20]; mPa=milliPascals
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 9 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 2. Method Evaluation

Compound

Min.

Sample

Vol (L)

Range Studied 

(μg/samp)

LOD

(μg/ samp)

Mean

Bias

Overall

Precision 

(sŝrrrr)

Accuracy

Storage

Stability

31-Day 

% recovery

24 °C

Storage 

Stability 

31 day 

% recovery

-12 °C

Aldicarb 240 12.0-240 1.2 -0.009 0.066 ± 0.131 93.2 95.6

Benomyl 6 12.0-120 0.6 A A A A A 

Captan 30 48.0-960 4.8 -0.036 0.061 ±0.142 98.7 102.2

Carbaryl 6 12.0-240 0.06 +0.012 0.061 ±0.123 88.2 91.8

Carbendazim 6 B 0.6 +0.006 0.061 ± 0.121 92.1 89.3

Carbofuran 240 12.0-240 0.6 -0.020 0.060 ± 0.126 89.1 92.4

Chlorpropham 6 12.0-240 0.6 -0.017 0.068 ± 0.140 84.3 85.9

Diuron 3 12.0-240 0.6 -0.062 0.060 ± 0.167 86.0 87.1

Formetanate. 60 12.0-240 0.6 +0.032 0.056 ± 0.129 89.8 93.0

Methiocarb 60 12.0-240 0.6 +0.009 0.061 ± 0.122 85.1 89.1

Methomyl 12 12.0-120 0.6 -0.002 0.063 ± 0.124 90.5 95.2

Oxamyl 240 12.0-240 0.6 +0.037 0.055 ± 0.132 94.8 95.9

Propham 3 12.0-240 0.8 -0.053 0.066 ± 0.168 88.5 92.5

Propoxur 60 12.0-240 0.6 +0.007 0.079 ± 0.156 91.4 95.4

Thiobencarb 6 12.0-240 0.6 -0.068 0.073 ± 0.197 75.0 79.8

A Results calculated as carbendazim, the primary breakdown product of benomyl. B See range for benomyl, a precursor for carbendazim.
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 10 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 3. Recommended liquid chromatographic columns and conditions

C18 Column C18 Column Cyano Column Cyano Column

Solvent Acetonitrile Methanol Acetonitrile Methanol

Application Primary Analysis Confirmation Confirmation Confirmation

Stationary phase Octadecyl Octadecyl Cyanopropyl Cyanopropyl

Length (mm) 300 150 250 250

ID (mm) 3.9 3.9 4.6 4.6

Particle size (μm) 4 4 5 5 

Ligand densityA 2.7 2.7 5.2 5.2

Mobile phase A Solvent Water Water Water Water

Mobile phase A organic modifierB 2% 1-propanol None None None

Mobile phase A bufferC TEA-PO4 None TEA-PO4 TEA-PO4

Mobile phase A concentration (M) 0.02 M None 0.02 M 0.02 M

Mobile phase B solventD Acetonitrile Methanol Acetonitrile Methanol

Mobile phase B organic modifier 2% 1-propanol None None None

Initial hold time 0 0 0 0 

Program rate 3-95% B 10-80% B 3-60% B 3-95% B

Program time (min) 30 30 30 30

Program type Linear Linear Linear Linear

Final hold time (min) 5 5 5 5 

Flow rate (mL/min) 1.00 1.00 1.00 1.00

Column temperature (°C) Ambient (~24) Ambient (~24) Ambient (~24) Ambient (~24)

Dwell volume (mL) 0.6-0.8 3.5-3.8 0.6-0.8 0.6-0.8

Injection volume (μL) 5 30 5 5 

Injection solvent Acetonitrile 1:3 acetonitrile:H2O Acetonitrile Acetonitrile

A Ligand density (micromole/m2

) is a better description of surface coverage than % carbon loading [22, 23]. B

 Choice of alcohol modifier is not critical. Percentages may be varied to adjust retention times and peak shapes for 

early eluting analytes. Reequilibration time may be shorter with 1-propanol [24-26]. C Buffer is very essential for basic analytes such as Formetanate, Carbendazim, and Benomyl [12,27,28]. Formentate is 

cationic at about pH 7, and its actual elution time is sensitive to small changes in pH and ionic strength of the buffer 

in mobile phase A. D Acetonitrile is the better choice when monitoring compounds at UV absorptions below 210 nm [20,29].
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 11 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 4. Approximate retention times and indices for organonitrogen pesticides and potentially 

interfering compounds

CompoundA

(by retention time)

C18 Column 

Retention 

indexB 

C18 Column 

(MeCN 

solvent) 

retention time 

(min)

C18 Column 

(MeOH solvent) 

retention time 

(min)

Cyano Column 

retention 

index

Cyano column 

(MeCN solvent) 

retention time 

(min)

Cyano column 

(MeOH solvent) 

retention time 

(min)

Solvent void 

volume

0.000 2.3 1.4 0.000 3.0 3.2

Asulam 0.004 2.3

Imazapyr 0.372 3.2

Acetaminophen/IS 1.000 4.7 1.000

Oxamyl 1.269 6.1 9.9 1.865 7.0 6.4

Caffeine 1.397 6.8

Methomyl 1.445 7.0 10.7 1.915 7.4 6.6

Formetanate 1.573 7.7 c 2.574 12.8 11.3

Sulfometuron

methyl

1.868 9.3

Acetanilide/IS 2.000 9.9 2.00 8.1 7.7

Fenuron 2.053 10.2

2,4-D acid 2.064 10.2

Nicotine 2.179 10.8

Carbendazim 2.192 10.8 c 4.274 13.6 12.9

Chlorimuron ethyl 2.422 11.9

Aldicarb 2.755 13.5 19.9 3.00 10.5 10.1

Tebuthiuron 2.817 13.8 23.9

m-Cresol 2.866 14.0

Bromacil 2.902 14.2

Hexazinone 2.921 14.3

Dinoseb 2.928 14.3

Simazine 2.938 14.3

Monuron 2.981 14.5

Acetophenone/IS 3.00 14.6 3.00 10.5 9.9

Cyanazine 3.003 14.6

Metribuzin 3.115 15.0

Thiodicarb 3.247 15.5

Aminocarb 3.301 15.7

Propoxur 3.317 15.8 22.9 3.675 13.1 11.9

Bendiocarb 3.376 16.0 23.3

Carbofuran 3.399 16.1 23.2 4.018 14.4 14.1

Fluometuron 3.551 16.6 25.2

Chloroform 3.601 16.8

Carbaryl 3.654 17.0 24.5 5.236 19.1 18.2

Atrazine 3.668 17.1

Metalaxyl 3.837 17.6

Diuron 3.843 17.6 27.0 5.751 21.1 19.9

DEET 3.851 17.7

Alpha-Naphthol 3.893 17.8

Propiophenone/IS 4.000 18.2 4.00 14.4

Propachlor 4.241 18.8

Thiophanate 4.241 18.8 27.6

Propham 4.267 18.9 25.9 5.092 17.7 17.3

Diethyl phthalate 4.367 19.2

Clomazone 4.459 19.4

Siduron 4.615 19.9

Desmedipham 4.696 20.1

Phenmedipham 4.700 20.1 >33

Methiocarb 4.744 20.2 29.3 6.680 22.5 21.8
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 4. Continued

CompoundA

(by retention time)

C18 Column 

Retention 

indexB 

C18 Column 

(MeCN 

solvent) 

retention time 

(min)

C18 Column 

(MeOH solvent) 

retention time 

(min)

Cyano Column 

retention 

index

Cyano column 

(MeCN solvent) 

retention time 

(min)

Cyano column 

(MeOH solvent) 

retention time 

(min)

Linuron 4.848 20.5 28.9

BDMC/IS 4.904 20.6

SWEP 4.919 20.7

Captan 4.926 20.7 27.7 6.172 20.9 21.6

Promecarb 4.981 20.8 20.4

Butyrophenone/IS 5.000 20.9 5.000 17.4

Mexacarbate 5.186 21.3

Toluene 5.269 21.5

Chlorpropham 5.504 22.1 30.1 6.700 23.4 23.3

Folpet 5.537 22.2 >33

Barban 5.566 22.3 >33

Malathion 5.731 22.7

Fenitrothion 5.802 22.8

Benomyl 5.822 22.9 c 7.391 25.8 23.9

Oryzalin 5.860 23.0

Metolachlor 5.876 23.0

Alachlor 5.979 23.3

Acetochlor 5.983 23.3

Valerophenone/IS 6.000 23.3 6.000 20.9

Captafol 6.018 23.4 30.1

Neburon 6.045 23.4

Parathion 6.640 24.7

Hexanophenone/IS 7.000 25.5 7.000 24.0

Thiobencarb 7.148 25.8 33.4 7.916 26.2 26.8

Heptanophenone/IS 8.000 27.6 8.000 26.4

Di-n-butyl 

phthalate

8.016 27.6

Chlorpyrifos 8.701 28.9

Pendimethalin 8.724 28.9

2,4-D butoxyethyl 

ester

8.892 29.2

Octanophenone/IS 9.000 29.4 9.000 29.0

BHT 9.488 30.2

Amitraz 9.886 30.9

Nonanophenone/IS 10.000 31.1

2,4-D ethylhexyl 

ester

10.545 31.9

Decanophenone/IS 11.000 32.5

A Organonitrogen pesticides are in bold letters.

B Estimated (~) retention times are extrapolated from shorter columns using relative retention times.

C Without TEA-PO4 buffer, basic compounds had irregular peak shapes and retention times, or were not 

detected. Abbreviations: MeCN=acetonitrile; MeOH=methanol; IS=internal standard.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 1. STRUCTURE OF CARBAMATE, UREA, AND SULFENIMIDE PESTICIDES
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPENDIX A. NOTES ON HPLC ANALYS IS OF CARBAMATE, UREA, AND SULFEN IMIDE PESTICIDES

A. ANALYTES

1. Aldicarb

a. UV response for aldicarb passes through a minimum at approximately 225 nm. If only two 

wavelength channels are used (225 nm and 200 nm), the response for aldicarb at 200 nm 

should be monitored. At 205 nm the signal will be smaller, but the signal-to- noise ratio may 

be better on particular instruments. An alternative but lesser maximum is at approximately 

245 to 246 nm. This latter maximum, though small, may have less background noise and 

interferences from coelutants.

b. Aldicarb is a highly toxic pesticide. Care should be exercised in handling pure stock material.

2. Benomyl (see also Carbendazim)

a. Benomyl breaks down rapidly by either hydrolysis in protic solvents (e.g., water or methanol) 

or solvolysis in nonproticsolvents (e.g., methylene chloride or acetonitrile) to carbendazim. 

The breakdown of benomyl can be so rapid that no benomyl will be detected at all after 

approximately 4 to 24 hours at room temperature.The rate of solvolysis is slower in the less 

polar methylene chloride andvery rapid in acetonitrile, even exceeding the rate of hydrolysis 

[10-12]. Benomyl standards in these nonprotic solvents can be stabilized with the addition of 

1% n-butyl isocyanate [30]. The preservative effect is lost as soon as the solution is diluted in 

solvents that contain or consist of protic solvents, since protic solvents also react with the 

isocyanates. It is generally unnecessary to add any preservative to the benomyl standard 

solution (it breaks down anyway). The carbendazim produced can precipitate out, if the 

solutions are too concentrated. In such an event, the addition of 1% n-butyl isocyanate is 

essential.

b. Since benomyl breaks down to carbendazim, do not include both benomyl and carbendazim 

in the same standard mixture.

c. When analyzing for benomyl, both benomyl and carbendazim must be determined. The 

results can be reported as either benomyl or carbendazim by converting the response of one 

to the equivalent response for the other at any particular wavelength. The relative response 

of benomyl to carbendazim at 225 nm has been determined to be approximately 1.0738, 

(adsorption-benomyl/ adsorption-carbendazim). This ratio should be determined for 

individual instruments with equimolar solutions analyzed separately, and with the benomyl 

injection solution being preserved with 1% n-butyl isocyanate. To convert carbendazim 

response to the equivalent benomyl response so that the values can be summed, multiply the 

carbendazim response by 1.0738 and add this to the benomyl response. Report results as 

benomyl. Alternately, to report results as carbendazim, divide benomyl response by 1.0738 

and then add this to the carbendazim response. Report as total carbendazim.

d. See notes for carbendazim.

3. Captan

a. Captan is not stable in methanol or in aqueous mixtures of methanol or acetonitrile at 

temperatures greater than –12 °C. Therefore, do not dilute desorbing solutions with water or 

methanol to weaken the injection solvent. Instead, use small injection volumes of acetonitrile 

solutions.

b. Use methylene chloride for making stock standard solutions. See Note 16 of this Appendix.

c. Captan has low absorption at 225 nm (see UV spectrum for captan). If only two wavelength 

channels are used (225 and 200 nm), the response for captan at 200 nm should be monitored. 

Although a lesser response may be obtained at approximately 205 or 210 nm, this can be 

accompanied by a proportionately better signal-to-noise response and may be an optional 

choice of wavelength if it is compatible with other analytical objectives.

4. Carbaryl (see also Diuron): Carbaryl tends to break down to 1-naphthol. Chloroacetic acid in 

methanol will inhibit this reaction [9], but this reagent is deleterious to other analytes. Carbaryl is 
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

stabilized in acetonitrile desorbing solutions containing the TEA- PO4 preservative for at least 24 

hours at room temperature, long enough to permit HPLC analysis. Any 1-naphthol that is 

detected may represent breakdown of carbaryl prior to desorption or may come from another 

source. Carbaryl has been stabilized with the TEA-PO4 buffer/preservative for up to three weeks at 

room temperature. 

5. Carbendazim

a. See notes for benomyl.

b. Carbendazimcan break down to 4-aminobenzimidazole and other compounds. In acetonitrile 

containing TEA-PO4 preservative, this breakdown appears to be inhibited for up to 24 hours at 

room temperature, long enough to permit analysis of desorbates.

c. Carbendazim is itself used as a pesticide. There are also other analytes such as thiophanate 

methyl that can break down to carbendazim. Therefore, any carbendazim should not 

automatically be assumed to represent benomyl.

6. Carbofuran

a. In most instances carbofuran and bendiocarb coeluted under all conditions tested. Their UV 

spectra are so nearly alike as to be practically indistinguishable. Any positive identification for 

carbofuran must be tempered by this knowledge and would require an alternate confirmation 

such as by mass spectrometry or a knowledge of the history of the sample.

b. Propoxur and carbofuran elute very close together. The separation of the two compounds 

with nearly baseline resolution is a good test of resolving power of the HPLC system for neutral 

compounds on a C18 HPLC column. If they cannot be resolved, there may be problems with 

large dead volumes in the system, analytical column quality, or injection quality.

7. Chlorpropham: No serious problems are expected to be encountered.

8. Diuron: The breakdown product of carbaryl, 1-naphthol, elutes immediately after diuron under the 

conditions given. On certain columns, it may coelute with diuron, giving false positive responses. 

The retention time of 1-naphthol relative to diuron must be determined for any particular column 

if carbaryl is anticipated to have been present in the sample.

9. Formetanate

a. At analytical concentrations formetanate is not stable either in methanol or in aqueous 

mixtures of methanol or acetonitrile with or without chloroacetic acid. Therefore, do not dilute 

desorbing solutions with methanol or water to weaken the injection solvent. Do not use 

chloroacetic acid as a preservative. Use only small injection volumes of acetonitrile solutions.

b. The breakdown of formetanate in analytical solutions is inhibited in acetonitrile containing 

TEA-PO4 preservative at room temperature for up to 24 hours, long enough to permit analysis.

c. At stock standard concentrations, formetanate dissolves better in 50:50 mixtures of methanol:

acetonitrile and is stable if kept in freezer at –12 ± 1 °C.

d. Formetanate is basic and possesses a single positive charge in solution at neutral pH (6.9 – 7.1). 

The use of TEA-PO4 buffer and/or base-deactivated HPLC columns is essential for the analysis 

for this compound.

e. The exact retention time of formetanate is more affected by exact pH and ionic strength of the 

buffer (or any other electrolytes) than are the noncharged analytes. Day-to-day inconsistencies 

in the preparation of mobile phases will cause differences in retention time. If the analyte is 

eluting too close to methomyl (which just precedes it under the specified conditions), its 

retention time may be delayed by either raising the pH slightly (by 0.1 to 0.2 units), or by 

lowering the ionic strength of mobile phase A. 

10. Methiocarb: Methiocarb has the same problems as carbaryl, only to a lesser extent. As methiocarb 

breaksdown, there are several other peaks that arise. These have been shown to be 

alkylthiophenol and/or oxidation products.

11. Methomyl: Attention should be placed on the problem of methomyl as it relates to its early elution 

time. Refer to Notes B1a and B4 of this Appendix.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

12. Oxamyl: Attention should be placed on the problem of oxamyl as it relates to its early elution time. 

Refer to Notes B1a and B4 of this Appendix.

13. Propham: No problems are expected with propham.

14. Propoxur

a. With prolonged standing in desorbing solutions at room temperature without preservative, 

propoxur has been observed to break down to some degree, although much less than 

carbaryl.

b. Propoxur elutes very close to carbofuran. See comments under carbofuran.

15. Thiobencarb: Thiobencarb is relatively nonpolar and has lower (but acceptable) recoveries from 

the XAD-2 resin than the other analytes.

16. Solvent for Analyte Standard Solutions: Most of the analytes (Table 1) are soluble in acetonitrile at 

stock standard concentrations. The exceptions are benomyl captan and formetanate. Appropriate 

solvents are described under headings for these compounds. In each of the solvents mentioned, 

analytes are stable at –12 ± 1 °C for at least 30 days.

17. Desorption Solvent and Preservative for Analytes: All analytes (except benomyl) were stable in 

acetonitrile at –12 °C for at least 48 hours. If refrigerated HPLC autosampler trays are available and 

all desorbing and tumbling operations are carried out in subambient conditions, no preservative 

should be needed. At room temperature, several analytes (e.g., carbaryl, methiocarb, and oxamyl 

especially) degraded unpredictably over 24 to 48 hours. The addition of as little as 0.2% v/v of 

aqueous 0.1 M triethylamine phosphate (TEA-PO4) buffer, pH 6.9 to 7.1, to the acetonitrile 

desorptionsolvent stabilizes all analytes (except benomyl) from hydrolysis or solvolysis for at least 

48 hours at room temperature. Methanol, isopropanol, aqueous methanol, and acetonitrile 

containing any of these alcohols promote degradation of several analytes (especially captan and 

formetanate) and poor desorption efficiencies. The use of chloroacetic acid, as is required for 

aqueous samples in EPA Method 531.1 [9], is destructive to at least formetanate. Since the 

desorption solvent is also the HPLC injection solvent, refer to note on Injection Solvent (B1) for 

additional comments. 

B. HPLC CONDITIONS

1. Injection Solvent

a. Normally a solvent producing equal or lower capacity factors than the mobile phase is 

desirable as an injection solvent in order to produce sharp peaks for early eluting analytes 

(e.g., oxamyl and methomyl) [31,32]. Since the initial mobile phase mixture of this method is 

mostly water, the only way to achieve this would be to dilute the samples with water. Water, 

however, is deleterious to several of the analytes specified in this method (see note A17 

above). Therefore, an acceptable alternative is to inject very small volumes, not greater than 

5 μL [31], of the desorption solvent (acetonitrile in this method) on a high-resolution HPLC 

column. If it is known that only analytes that are stable in aqueous solutions are to be 

determined (e.g., oxamyl and methomyl, but not formetanate or captan), the desorbates may 

be diluted with water and larger volumes injected. By diluting with water, sharper peaks may 

be obtained for early eluting compounds with accelerated elution conditions such as the use 

of shorter HPLC columns, higher percentages of organic modifier, or higher percentages of 

mobile phase B in the initial conditions.

b. All sample extracts must be filtered through individual 0.45-micron PTFE filters in order to 

prolong guard column lifetimes and to protect the injection system valving.

c. The accidental or intentional inclusion of significant amounts of solvents less polar than 

acetonitrile in the injection solvent, such as tetrahydrofuran or acetone, may shorten retention 

times and adversely affect the peak shapes of early eluites (e.g., oxamyl and methomyl) [31].

2. Guard Columns: Guard columns are essential to the long life and reproducibility of results on the 

main analytical column. There are several on the market. Those giving the lowest possible dead 

volume preserve good peak shapes and resolution of quality analytical columns and are preferred.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3. Analytical Columns

a. General: The main analytical column specified in this method is a C18 reverse-phase column. 

Other columns known to perform well also may be used, following manufacturer 

recommendations. There are many good columns available [28, 33].

b. Base Deactivation: The basic compounds (benomyl, carbendazim, and formetanate) present 

special challenges that are easily overcome with highly inert or base-deactivated columns [26]. 

The addition of TEA-PO4 buffer to mobile phase A also greatly improves performance for these 

compounds [16, 26, 27].

c. Dimensions: A longer column (300 mm) was used in this method in order to improve 

resolution for a large number of analytes and interferences, expected and unexpected. Long 

columns have higher operating pressures; therefore, necessary steps should be taken to 

provide for them, such as thicker-walled (narrower-bore) transfer lines (if polymer tubing is 

used, etc.) Shorter columns may work well if a limited number of analytes or interferences are 

expected. The diameter (3.9-mm) is not necessarily critical and should be governed by user 

preference and equipment, adjusting flow rates and other parameters as necessary. Diameters 

from 2.0 to 4.6 mm should be expected to perform similarly, as long as the columns are stable 

and rugged.

d. Packing Density: Packing (or ligand) density is a better parameter than carbon loading for 

comparing columns [21,22]. A column with a ligand density near that of the one used in this 

method should be expected to perform similarly.

4. Mobile Phase Composition

a. Modifier: Because of the high percentage of water in the initial mobile phase, a condition 

referred to as hydrophobic collapse of the C18 phase occurs [23,24], which results in poor 

reequilibration and irreproducible retention times and peak shapes for early eluting analytes 

such as oxamyl and methomyl. The addition of a constant amount of an alcohol to both 

mobile phases A and B has been reported and found to improve column performance under 

these conditions. The method specifies 2% 1-propanol to be added to both mobile phases A 

and B. A concentration of 1-propanol between 3% and 4% has been reported as optimum [24, 

25]; 2% is a compromise made in order to generate retention (capacity) factors for the earliest 

eluites, oxamyl and methomyl, of greater than 5,as suggested in the literature [26]. Also, 1-

propanol has been reported (and found) to reduce the time required for reequilibration at the 

end of the run [24]. Other alcohols may be used at higher concentrations (except for 

reducing reequilibration times) [1]. These are isopropanol (at 3% to 5%) and methanol (at 5% 

to 10%). If early-eluting analytes are to be determined, or if a column is found that gives 

sufficiently long retention times for the earliest expected eluite with an initial mobile phase 

composition as high as 5% to 10% acetonitrile in water, the alcohol could be eliminated (from 

both phases). Table 4 would not apply if such changes are made.

b. Mobile Phase B: Pure methanol as mobile phase B results in a steep rise in the baseline UV 

response, making automatic integrations difficult and UV scanning for confirmation spectra 

nearly impossible. For this reason acetonitrile has been chosen [28, 29]. Methanol is acceptable 

if these conditions are tolerable and is recommended as an alternate solvent system on a C18 

column for confirmation (Section F3); however, precision and LOD values reported in Table 2 

would not be applicable.

c. Solvent Programming: One of the most serious concerns that may be encountered in trying to 

make adaptations of this method is an attempt to shorten retention times by using higher 

concentrations of organic modifier in mobile phase A, higher percentages of mobile phase B in 

the initial solvent program condition, or faster solvent programming. In any case, these 

changes will seriously affect the peak shape, sensitivity, and retention time reproducibility of 

early eluting analytes. There is also a greater possibility of false positives from potential 

interferences because of poor resolution in the early region of the chromatogram. Retention 

times for the earliest eluting analyte should be kept at 3 to 6 times the retention time of the 
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

solvent from an unretained analyte, which is equivalent to a capacity factor of 2 to 5 [5]. The 

retention or capacity factor for any analyte can be calculated as follows:

Retention or Capacity factor = (ttrr − tt0)⁄tt0

Where tr = the retention time of the compound

And t0 = the retention time of an unretained analyte (the mobile phase holdup time)

The value of keeping anaylytes from eluting too close to the solvent front cannot be 

overemphasized.

d. Buffer: For the compounds evaluated, formetanate, carbendazim, and benomyl required a 

buffer in order to obtain good peak shapes. Of several pHs tested, pH 6.8 to 7.1 gave the best 

results. Buffer concentration should be 0.01 to 0.05M. The concentration and pH of the buffer 

had a great effect on exact retention time of formetanate.

5. Dwell Volumes: Dwell volumes can affect the retention times and peak shapes of analytes. This is 

internal volume of the system from the point of mixture of the mobile phases A and B to the head 

of the column. It includes the volume of the transfer lines; any in-line filters (which should be 

between the pump and the sample injector if at all, and definitely not after the sample injector); 

the sample injector; the guard column; and the head of the analytical column. This volume can 

range from approximately 0.6 to 3.8 mL. At a flow rate of 1 mL/minute, each mL dwell volume 

represents a minute delay between the time that the pump produces a given change in a mixture 

of solvent and the time when the analytical column experiences that change. In effect, it is 

equivalent to a solvent program delay [13,14]. For this method, lower dwell volumes and no 

program delays gave better peak shapes for the early eluites.

6. Dead Volumes: Dead volumes can cause the method to fail seriously. Important areas to look for 

are larger bore and longer-than-necessary transfer lines between the sample injector and the 

analytical column [15]. Another area of potential large dead volumes are poorly designed or 

connected guard columns and sample injectors.

7. UV Wavelength Selection

a. The wavelengths specified in this method are a best compromise for all of the analytes listed 

in Table 1. There are maxima for each of the pesticides that would give greater sensitivity 

and/or signal-to-noise ratio. If only a selected few of the pesticides in Table 1 are to be 

determined, other wavelengths may be considered. For example, if only ureas are to be 

determined, the selection of an absorbance band of 240 to 250 nm would give greater 

sensitivity for these compounds and less interference from others.

b. Spectra are also provided in the Backup Data Report [1] for many analytes and potential 

interferences. These were obtained under actual operating conditions. Because of unavoidable 

errors in background subtractions, the profiles of these spectra are subject to errors at the low 

wavelength ends of the spectra (190 nm to 210 nm). This could be attributed to the 

absorbance from the alcohol modifier to the mobile phases, which, in spite of efforts to add an 

equal amount to both phases, produces a slight baseline rise near the end of the 

chromatograms.

c. Many of the O-aryl carbamates and the sulfenimides absorb well only in the low UV range 

(<215 nm). This is generally an area of great background noise. Many contaminants 

(plasticizers and solvents) also absorb in this range better than at higher wavelengths. 

Selecting a longer wavelength that is not at the absorption maximum may give a better signalto-noise ratio and thus actually increase sensitivity. This should be determined by 

experimentation for selected analytes.

d. Bandwidth: A bandwidth of 15 nm was used for evaluation of this method.

C. INTERNAL STANDARDS

1. Calibration Internal Standards: Internal standards are essential for obtaining the precisions listed 

in Table 2 [1, 16]. Acetanilide was found to be unretained by the media during desorption, and 
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therefore, could be added to the extraction fluid. It was also found to be relatively stable and not 

to interfere with the retention times of other analytes. A second compound, acetophenone, may 

be added as a back-up internal standard. It is also unretained by all media except XAD-2, on which 

it has about a 95% recovery. It may be used whenever the acetanilide is interfered with by 

coeluting analytes or contaminants. It alsoserves to indicate, by monitoring relative response 

between the two internal standards, when the first internal standard has a coeluting interference.

2. Alternate Calibration Internal Standards: It may be wise, especially under isocratic conditions, to 

select other internal standards having capacity factors closer to particular analytes of interest. 

There is a wide range of alkyl phenones available for late-eluting analytes, and 4-hydroxyacetanilide (acetaminophen) for earlier eluting analytes. Their retention times are listed in Table 4. 

Since these longer alkyl chain phenones have increasingly lower recoveries from the XAD-2 resin, 

they should not be added to the extraction solutions until after the resin is removed.

3. Retention Index Reference Standards: These are optional standards and are used to establish 

relative retention times for qualitative purposes. These are highlighted in Table 4 and include a 

homologous series of alkyl phenones and acetanilides. Establishing retention times between two 

nearest-eluting reference standards gives a more consistent retention value than retention time 

alone or relative retention time using a single internal standard alone. This value, the Retention 

Index, varies according to column and analytical conditions, but should be relatively consistent for 

any one set of conditions and more reliable as a qualitative tool over a long period of time. This 

value is calculated as follow:

RI(A) = Retention Index of Analyte “A”

TTrr(AA) − TTrr(RRRR−PP)

TTrr(RRRR−FF) − TTrr(RRRR−PP)

+ NN(RRRR−PP)

Where: Tr(A) = retention time of analyte A

Tr(RS-P) = retention time of preceding reference standard

Tr(RS-F) = retention time of following reference standard

N(RS-P) = a number assigned to preceding reference standard, with zero assigned to the 

first peak in the series.

(Numerical assignments for the acetanilide and phenone series are suggested in Table 4.

In order to avoid confusion in the chromatograms, the retention index reference standards are 

analyzed periodically as external standards and not added to the analytical samples themselves. 

The use of these reference standards is optional but is suggested where application of this method 

is expected to encounter consistently a broad range of unknown analytes and to augment other 

confirmatory techniques.

D. INTERFERENCES:

The UV detector responds to many compounds. Retention times of some common potential 

interferences are provided in Table 4. Interfering compounds that may be encountered are 

discussed below. 

1. DF Impurities in Mobile Phases and Additives: Only HPLC-grade solvents should be used. 

Triethylamine (TEA) was found to develop unidentified impurities over time which contributed to 

significant irregularities in the baseline. This degradation was reduced or eliminated by storing the 

TEA under nitrogen in a small desiccator at 0 to 4 °C [8].

2. Organic Solvents or Fuels

a. Chloroform showed a response that nearly coelutes with carbaryl (Table 4). Benomyl and 

captan, therefore, are made up in methylene chloride, which is more UV transparent than 

chloroform.

b. Toluene (Table 4).
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c. Blends of solvents having ketones, ester, or any of the above compounds, such as lacquer 

solvents, gasoline, paint stripper, and cleaning solvents, are to be avoided or documented 

during sample collection and handling.

3. Industrial Chemicals (Plastic and Rubber Additives)

a. Several plasticizers might elute in the window of interest depending upon the selection of 

column and conditions. These include diethyl and dibutyl phthalates. Dibutyl phthalate is 

typically found in polyvinyl gloves, flexible tubing, and in coatings on bottles and tool handles. 

Contact with these materials should be avoided or documented. Other plasticizers, such as 

dioctyl phthalate and bis-ethylhexyl adipate, have late elution times and will elute after about 

30 minutes under the conditions specified. However, if the run times are shortened, these may 

be carried over on-column to subsequent analyses, causing interferences.

b. Common antioxidants such as BHT (2,6-di-tert-butyl-4-methyl phenol) also elute late and may 

be carried over on-column, creating interferences in subsequent analyses.

4. Other Pesticides: Spray mixtures very often contain a mixture of pesticides. It is not uncommon to 

find chlorpyrifos, an organophosphate, or a pyrethroid pesticide in combination with propoxur, for 

example. Both of these noncarbamate pesticides can be detected under the conditions of the 

method. The retention time and spectra of chlorpyrifos is included for qualitative purposes. Most 

pyrethroids elute later than most of the carbamates; if present, they may elute in a subsequent run 

if run times are too short. It is also not uncommon to mix herbicides of different classes such as 

diuron with bromacil, atrazine, or 2,4-D. Because of this possibility, retention times and spectra of 

other common herbicides are also provided in Table 4 for qualitative purposes.

5. Miscellaneous Chemicals: There are a number of chemicals which elute in the retention time 

window of interest for carbamates and urea pesticides and may, with particular columns or 

conditions, interfere with analytes of interest. The presence or the known use of these compounds 

or their sources should be documented as part of the sample history. Table 4 lists a few of the 

compounds, which include the following.

a. The common insect repellent, DEET (N,N-diethyl-meta-toluamide). Since DEET may be heavily 

used by outdoor workers, its presence on exposed areas of the skin or clothing may contribute 

to sampler contamination either through direct contact with the face of the sampler or 

through collection of vapors if the sampler is in close proximity to areas of application or is 

exposed to overspray during application from a spray can or bottle.

b. Inadvertent collection of tobacco side-stream smoke may introduce potentially interfering 

compounds, one of which is nicotine.

c. Compounds in beverages used during work periods, which include at least caffeine.

E. SAMPLER

1. The OVS-2 Sampler. The OVS-2 (OSHA Versatile Sampler with XAD-2) combines both filter and 

XAD-2 sorbent in one unit. The filter is necessary to trap submicron aerosols that would pass 

through the XAD-2 bed. Substitutions should not be made.

2. Quartz Fiber and Glass Fiber Filters (GFF). The OVS-2 tubes are available with either glass or quartz 

fiber filters. OSHA Methods specify GFF. This method specifies a quartz fiber filter. For analytes 

being desorbed with acetonitrile, no difference in desorption efficiency was observed between 

glass fiber and quartz fiber filters. Therefore, the tubes may be interchanged for the analytes 

specified in this method.

3. Flow Rates. The OVS-2 sampler is designed for a flow rate of 1 L/min. At slower flow rates, 0.1 to 0.2 

L/min, there may not be enough capture velocity for aerosols.

4. Applying Liquid Spikes. The Teflon® retainer ring should be removed when spiking the face of the 

OVS-2 tubes with more than approximately 10 μL, in order to prevent wicking of the carrier solvent 

behind the ring and consequent loss of standard. Volumes of spiking fluid greater than 15 to 30 μL 

will flood the XAD-2 section and possibly wick into the back-up section. Whenever more than 15 

to 30 μL is to be applied to the tubes, air must be drawn through the tubes at approximately 1 
ORGANONITROGEN PESTICIDES: METHOD 5601, Issue 2, dated 25 February 2016 - Page 21 of 21

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

L/min during the spiking procedure and the solvent added in 15- to 30-μL increments with a few 

minutes between each addition allowed for drying of the solvent. SD

F. CONFIRMATION 

1. By Relative Retention Times (Retention Index).The Retention Index (RI) may vary considerably from 

column to column and from one set of conditions to another. But it will be reasonably consistent 

once a set of conditions has been chosen and will be much more reliable for day-to-day 

comparisons than will absolute retention times. Actual RIs need to be established for each set of 

conditions. Compounds that are ionic under elution conditions or that interact strongly with polar 

sites on the column will have the most variable retention times and retention indices.

2. By Second Column. The cyanopropyl stationary phase strongly induces some exchanges in elution 

orders and alters relative spacings between adjacent analytes in the chromatogram that may be 

useful in confirmations.

3. By Alternate Solvent. As mentioned earlier (Section B1), methanol as the mobile phase B solvent 

on a C18 column can be used just as effectively to establish confirmations because methanol 

interacts differently with the stationary phase than acetonitrile, and so different molecular forces 

come into play. Significant alterations in retention order are thus obtained; for some analytes this 

is more dramatic than with a cyanopropyl column.

4. By Ratio of Two UV Absorption Bands. As long as the UV absorption channels are not saturated, 

there should be a consistent ratio between the background-corrected absorption bands that is 

characteristic of each analyte and should reflect the ratio of relative heights of the absorption 

spectra at their respective UV bands in the UV spectra. This ratio is dependent, however, on the 

bandwidth of the adsorption bands employed, and consistent bandwidths must be used. The 

consistency of the ratio of absorption across the HPLC peak is also indicative of the purity of the 

peak. A constantly changing ratio indicates that the peak may have multiple components.

5. By Matching to Reference UV Spectra. Unknown spectra should not be oversaturated in any 

portion. They need to be background-corrected properly. If the baseline is rising, a background 

selected from the backside of the peak may induce losses of absorption in the region below 210 

nm. Conversely, one selected from in front of the unknown HPLC peak may add to this region of 

the spectra. It is better to get an averaged background first. Spectra maxima should match within a 

few nanometers. Relative absorbance at each maxima may vary even after background 

subtraction, depending upon the concentration of the analytes and the characteristics of different 

scanning UV detectors.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

FORMALDEHYDE ON DUST (TEXTILE OR WOOD) 5700

 H2C=O MW: 30.03 CAS: 50-00-0 RTECS: LP8925000

METHOD: 5700, Issue 2 EVALUATION: FULL Issue 1: 15 August 1994

Issue 2: 29 February 2016

OSHA: 0.75 ppm; 2 ppm STEL

NIOSH: 0.016 ppm; C 0.1 ppm (15 min); carcinogen

PROPERTIES: gas; BP -19.5 °C; vapor density 1.067 (air = 1); 

explosive range 7 to 73 % in air

SYNONYMS: methanal; formalin (aqueous 30 to 60% w/v formaldehyde); methylene oxide

SAMPLING

SAMPLER: Institute of Occupational Medicine (IOM) 

inhalable dust sampler or equivalent 

containing a 25-mm PVC filter, 5 μm pore 

size

FLOW RATE: 2.0 L/min

VOL-MIN: 240 L @ 0.002 mg/m3

 -MAX: 1050 L

SHIPMENT: Place cassette with filter in 30-mL screwcap low density polyethylene (LDPE) 

bottle; keep upright. Ship cold.

SAMPLE 21 days (matrix dependent)

STABILITY: (cold storage advised)

BLANKS: 2 to 10 field blanks per set

ACCURACY 

RANGE 

STUDIED: 0.007 to 0.16 mg/m3 [1] (1050-L samples)

BIAS: -4%

OVERALL

PRECISION (ŜrT): 0.093 [1] 

ACCURACY: ± 22% 

MEASUREMENT

TECHNIQUE: HPLC, UV DETECTION 

ANALYTE: 2,4-dinitrophenylhydrazone derivative of 

formaldehyde

EXTRACTION: 10 mL distilled water @ 37 °C, 4 h; 1 mL to 

3 mL 2,4-dinitrophenyl - 

hydrazine/acetonitrile (1.3 mg/mL 

DNPH/ACN)

INJECTION 

VOLUME: 15 μL

MOBILE PHASE: 30% acetonitrile/66% methanol/water 

(1:1v/v), 1 mL/min

COLUMN: C18, 5-μm particle size, 10-cm x 8-mm ID 

(with a C18 guard column)

DETECTOR: UV @ 365 nm 

CALIBRATION: standard solutions of formaldehyde in 1.3 

mg/mL DNPH/ACN

RANGE: 0.4 to 4000 μg/sample [1]

ESTIMATED LOD: 0.8 μg/sample [1]

PRECISION (SS�rr): 0.078 @ 7 to 174 μg per sample [1]. 

APPLICABILITY: The working range is 0.0004 to 3.8 mg/m3 for a 1050-L air sample. This method has been used for the 

determination of formaldehyde in both textile dusts and wood dusts [1]. Caution should be exercised in the way that data 

collected with this method are interpreted. These results should be reported separately from vapor-phase formaldehyde exposure 

data until sufficient data has been collected to allow appropriate epidemiological interpretation of formaldehyde-containing 

particulate exposures.

INTERFERENCES: None identified

OTHER METHODS: In the absence of phenol or other substances known to interfere with the chromotropic acid analysis of 

formaldehyde, the analysis procedure described in NIOSH method 3500 [2] can also be used with this extraction technique (See 

Appendix). The analysis procedure used in this method has also been used for the determination of formaldehyde in automobile

exhaust [3]. An alternate analysis [4] may be used in conjunction with this method to determine the amount of "released" 

formaldehyde from the collected particulate material. NIOSH method 5700 determines both "released" formaldehyde and 

formaldehyde equivalents (e.g., small oligomeric pieces of formaldehyde-containing resin) present in the hydrolysis solutions. The 

use of these two analytical approaches may differentiate between the two forms of formaldehyde present in the sample by the 

difference in results. 
FORMALDEHYDE ON DUST (TEXTILE OR WOOD) 5700, Issue 2, dated 29 February 2016 - Page 2 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. 2,4-dinitrophenylhydrazine (2,4-DNP) (1.3 g) 

in 1 L acetonitrile; 1.3 mg/mL solution.

2. Formaldehyde stock solution, 1 mg/mL (see 

Appendix).

3. Methanol, distilled in glass.

4. Acetonitrile, distilled in glass.

5. Water, deionized and distilled.

6. Perchloric acid solution, 1 N. *

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: 25-mm PVC filter (5.0-μm pore 

size) in a 25-mm Institute of Occupational 

Medicine Inhalable (IOM) dust sampler. The 

sampler should meet the American 

Conference of Governmental Industrial 

Hygienists (ACGIH) definition for collection 

of the inhalable fraction of particulate mass 

[5]. 

2. Personal sampling pump, 2.0 L/min, with 

flexible polyethylene or 

polytetrafluoroethylene (PTFE) tubing.

3. Bottles, screw-cap, low-density 

polyethylene (LDPE), 30-mL. 

Note: Do NOT use bottles with 'polycone' 

liners (source of high formaldehyde 

blank).

4. Liquid chromatograph with a UV detector, 

recorder, integrator and column (page 

5700-1).

5. Tweezers.

6. Syringes, 10-, 25-, 50-, and 100-μL.

7. Volumetric flasks, 10-, 100-mL and 1 L.

8. Pipets, 0.1-, 0.5-, 1.0-, and 3-mL glass, 

delivery, with pipet bulb.

9. Graduated cylinders, glass, 25-mL.

10. Cotton gloves.

11. Equipment for standardizing formaldehyde 

stock solution, Burets, 50-mL.

12. pH meter.

13. Magnetic stirrer.

14. Beaker, 50-mL. 

15. Vials, 5-mL.

16. Filters, 0.45 μm.

17. Scintillation vials, 20-mL.

 

SPECIAL PRECAUTIONS: Perchloric acid is a strong oxidizing agent, toxic by ingestion and inhalation

and is a strong irritant. Use only in a well-ventilated fume hood. Formaldehyde is a suspect carcinogen 

[1,6] and should be handled in a hood.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. If gravimetric measurements are needed, handle the IOM cassettes only while wearing cotton gloves. 

Follow NIOSH method 0500 [7] for pre- and post-collection weighing procedure. Place the cassette 

containing a 25-mm PVC 5-μm pore filter in the filter holder.

3. Attach outlet of filter holder to the sampling pump.

4. Sample 240 to 1050 L of air at 2.0 L/min flow rate.

5. Transfer cassette from filter holder carefully to a 30-mL LDPE screw-cap bottle while wearing cotton 

gloves and seal the bottle. Ship in a suitable container in order to prevent damage during transit and 

keep bottles upright.
FORMALDEHYDE ON DUST (TEXTILE OR WOOD) 5700, Issue 2, dated 29 February 2016 - Page 3 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

6. Collect a bulk sample (ca. 1 g) of the dust/fiber in a glass vial and ship separately.

SAMPLE PREPARATION:

7. If gravimetric weighing is done, post-weigh the entire cassette (see NIOSH method 0500 [7]) then 

return the cassette to the screw-cap bottle.

8. Transfer 8-mL distilled water to the LDPE screw-cap bottle containing the cassette. Use tweezers (two) 

to disassemble cassette which is immersed in water, remove all cassette pieces (shake off excess water) 

leaving the filter and dust in the bottle. Transfer the resulting solution to a 20-mL scintillation vial and 

rinse the screw-cap bottle with an additional 2 mL of distilled water. Replace the cap on the vial and 

then place the vial in a 37 °C water bath for four hours. Remove from the bath and allow to cool 1/2 

hour. Filter through a 0.45 μm pore size filter into another bottle.

CALIBRATION AND QUALITY CONTROL:

9. Prepare a calibration stock solution by dilution of 1-mL of 1 mg/mL formaldehyde stock solution to 

100 mL with distilled water.

10. Pipet aliquots of calibration stock solution (e.g., 40, 200, 750, 2000, 5000, and 10000 μL) into 10-mL 

volumetric flasks to prepare working standards. Higher level standards, if needed, can be prepared by 

using different dilutions of the formaldehyde stock solution.

11. Add distilled water to bring the volume of each working standard to 10 mL.

12. Analyze working standards together with samples and blanks (steps 14 through 17).

13. Prepare calibration graph of area vs. amount (μg) of formaldehyde per sample (10-mL total volume).

MEASUREMENT:

14. Transfer a 1-mL aliquot of the solution from step 8 (or step 11) to a 5-mL vial containing 3-mL of an 2,4- 

dinitrophenylhydrazine/acetonitrile solution (1.3 mg/mL). Add one drop 1 N perchloric acid (to 

catalyze the reaction to form the corresponding hydrazone), cap the vial and shake it gently for several 

seconds.

15. Set liquid chromatograph to conditions given on page 5700-1.

16. Inject 15-μL sample aliquot.

17. Measure peak area.

NOTE 1: If peak area is greater than the highest standard, take a smaller aliquot of the 

remaining unreacted sample solution, dilute to 1 mL with distilled water and analyze (steps 

14 through 17) and apply appropriate dilution factor in the calculations.

NOTE 2: For optimum results, all samples containing over 400 μg/mL should be diluted and 

reanalyzed, since the maximum amount of formaldehyde that can react with 3.9 mg of 2,4-

dinitrophenylhydrazine is 590 μg.

CALCULATIONS:

18. Determine mass, μg, of formaldehyde (W) found in the sample and the average media blank (B) from 

the calibration graph. Use the appropriate aliquot factor (e. g., 1 mL aliquot/original volume from step 

14) and the total sample volume (10-mL) unless corrected for in calibration plot (step 13).

NOTE: Include dilution factor for any sample which exceeded the highest calibration standard.

19. Calculate concentration, C, of formaldehyde from dust in the air volume, V (L).

CC = WW − BB

VV , mmmm mm3 ⁄
FORMALDEHYDE ON DUST (TEXTILE OR WOOD) 5700, Issue 2, dated 29 February 2016 - Page 4 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

EVALUATION OF METHOD: 

This method was evaluated over the range of 7 to 174 μg/sample at 24 °C and pressure of 761 mm Hg [1] 

using the chromotropic analysis procedure. Overall measurement precision, ŜrT, was 0.093 including a 5% 

pump error factor. Similar precision was observed with the 2,4-dinitrophenylhydrazine analysis procedure. 

Sample stability during storage was evaluated at 7 μg formaldehyde/2 mg of wood dust treated with a 

urea/formaldehyde resin. Samples showed 101% recovery after 21 days of storage at ambient conditions 

compared to one-day old samples. However, studies with a different matrix (urea- formaldehyde treated 

wood fibers) indicated that samples may be unstable when stored at room temperature.

Comparison of results from 2,4-DNP analyses with those from acetylacetone analyses [4] indicated that 

there was a significant difference between the two methods. The acetylacetone analyses used much 

milder conditions and appears to be reacting only with released formaldehyde, whereas the 2,4-DNP 

analysis required the addition of perchloric acid and may be breaking up oligomeric pieces of the partially 

hydrolyzed resin in the sample. By analyzing samples with the two methods, a further characterization of 

the sample matrix and potential exposure may be obtained. In matrices where materials that can form 

complexes with formaldehyde are present, such as sulfites, the acetylacetone analysis may be biased low, 

since it is unable to break up these complexes [8].

Two comparisons of the IOM samplers with closed-face cassette samplers were conducted in field 

situations [1]. The IOM samplers, on average, collected more dust/fibers measured gravimetrically than the 

closed-face cassette samplers in both studies. Formaldehyde levels in the dust/fibers on a μg/mg basis 

measured with the extraction procedure used in this method were comparable for both the IOM and 

closed-face cassette samplers. On the average, the IOM sampler tended to collect more dust than the 

closed-face cassette sampler.

REFERENCES:

[1] Kennedy ER, Gagnon YT, Teass AW, Seitz T [1992]. Development and evaluation of a sampling and 

analytical method for the determination of formaldehyde content of dust. Appl Occ Environ Hyg 7(4): 

231-240.

[2] NIOSH [1994]. Formaldehyde: Method 3500. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical 

methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) 

Publication No. 94-113 [www.cdc.gov/niosh/nmam/]. 

[3] Lipari F, Swarin S. [1982]. Determination of formaldehyde and other aldehydes in automobile exhaust 

with an improved 2,4-dinitrophenylhydrazine method. J Chromatogr 247: 297-306.

[4] Elia VJ, Messmer RA [1992]. Evaluation of method for estimating formaldehyde released from resincontaining paper and wood product dusts. Am Ind Hyg Assoc J 53: 632-638. 

[5] ACGIH [1985]. Particle Size Selective Sampling in the Workplace: Report of the ACGIH technical 

committee on air sampling procedures. Cincinnati, OH. American Conference of Governmental 

Hygienists. 

[6] NIOSH [1981]. NIOSH/OSHA occupational health guidelines for occupational hazards. Cincinnati, OH: 

U. S. Department of Health and Human Services, Centers for Disease Control, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 81-123.
FORMALDEHYDE ON DUST (TEXTILE OR WOOD) 5700, Issue 2, dated 29 February 2016 - Page 5 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[7] NIOSH [1994]. Particulate not otherwise regulated, total: Method 0500. In: Eller PM, Cassinelli ME, eds. 

NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and 

Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/]. 

[8] Nash T [1953]. The colorimetric estimation of formaldehyde by means of the Hantzsch reaction. 

Biochem J 55:416.

METHOD WRITTEN BY:

Y.T. Gagnon and E.R. Kennedy, NIOSH
FORMALDEHYDE ON DUST (TEXTILE OR WOOD) 5700, Issue 2, dated 29 February 2016 - Page 6 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPENDIX: Preparation and Standardization of Formaldehyde Stock Solution (ca. 1 mg/mL)

Dilute 2.7 mL 37% aqueous formalin solution to 1 L with distilled, deionized water. This solution is stable 

for at least three months. Standardize by placing 5.0 mL of freshly prepared 1.13 M sodium sulfite solution 

in a 50-mL beaker and stir magnetically. Adjust pH to between 8.5 and 10 with base or acid. Record the 

pH. Add 10.0 mL formaldehyde stock solution. The pH should now be greater than 11. Titrate the solution 

back to its original pH with 0.02 N sulfuric acid (1 mL acid = 0.600 mg formaldehyde; about 17 mL acid 

needed). If the endpoint pH is overrun, back-titrate to the endpoint with 0.01 N sodium hydroxide. 

Calculate the concentration, Cs (mg/mL), of the formaldehyde stock solution:

CCss = 30.0(NNaaVVaa − NNbbVVbb)

VVss

where: 30.0 = 30.0 g/equivalent of formaldehyde

Na = normality of sulfuric acid (0.02 N) 

Va = volume of sulfuric acid (mL) used for titration

Nb = normality of NaOH (0.01 N) 

Vb = volume of NaOH (mL) used for back-titration

Vs = volume of formaldehyde stock solution (10.0 mL)

Alternate Analysis Procedure for the Determination of Formaldehyde Liberated from Dust/Fiber 

Samples

NOTE: This analysis procedure should only be used where interferences to the chromotropic acid analysis 

are not present. These interferences include phenol, oxidizable organic material, other aldehydes and 

alcohols.

For formaldehyde determination by the chromotropic acid method, a 4-mL aliquot of the solution resulting 

from the desorption and incubation of the filter (step 8.) was analyzed by the procedure described in 

NIOSH Method 3500 [2]. The amount of chromotropic acid added to the sample could react with a 

maximum of 42 μg of formaldehyde per sample aliquot. (Whenever the amount of formaldehyde 

approached 30 μg per aliquot, the sample was diluted and reanalyzed.) The absorbance of samples at 580 

nm was then compared to a calibration curve constructed from results obtained from analysis of 

calibration standards containing known amounts of formaldehyde. The amount of formaldehyde present 

in each sample was determined based on the calibration curve.

Calibration standards were prepared by dilution of a standard solution of formalin in distilled water 

(1 mg/mL) and analyzed in the same manner as the samples. The calibration range usually covered 1 to 25 

μg of formaldehyde/4-mL aliquot. A limit of detection of 0.44 μg formaldehyde/filter sampler was 

determined in laboratory evaluation of this analysis procedure.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ARSINE 6001

 MW: 77.95 CAS: 7784-42-1 RTECS: CG6475000

METHOD: 6001, Issue 3 EVALUATION: FULL Issue 1: 15 May 1985

Issue 3: 20 October 2015

OSHA: 0.05 ppm (0.2 mg/m3)

NIOSH: C 0.002 mg/m3 (15 min); carcinogen

PROPERTIES: Gas; d 3.484 g/L @ 20 °C; BP −62.5 °C; MP 

−116.3 °C

SYNONYMS: Hydrogen arsenide; arsenic trihydride

SAMPLING

SAMPLER: SOLID SORBENT TUBE 

(coconut shell charcoal, 100 mg/50 mg)

FLOW RATE: 0.01 L/min to 0.2 L/min

VOL-MIN: 0.1 L @ 0.05 ppm

-MAX: 10 L

SHIPMENT: Routine

SAMPLE

STABILITY: At least 6 d @ 25 °C [1]

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: 0.09 mg/m3 to 0.4 mg/m3 [1] (10 L 

samples); 0.001 mg/m3 to 0.01 mg/m3 [2]

BIAS: −6.13% at 0.01 L/min to 0.2 L/min flow 

rates [1]; −11% @ 0.876 L/min [2]

OVERALL

PRECISION ( ): 0.087 [2]

ACCURACY: ±23.2%

MEASUREMENT

TECHNIQUE: ATOMIC ABSORPTION, GRAPHITE 

FURNACE

ANALYTE: Arsenic

DESORPTION: 1 mL 0.01 mol/L nitric acid; 30 min in 

ultrasonic bath

MATRIX

MODIFIER: Nickel, 1000 μg/mL

WAVELENGTH: 193.7 nm; deuterium or hydrogen 

correction

GRAPHITE

FURNACE: DRY: 40 s @ 110 °C; CHAR: 15 s @ 1200 °C; 

ATOMIZE: 7 s @ 2540 °C

INJECTION: 50 μL

CALIBRATION: Arsenic in 0.01 mol/L nitric acid with 100 

mg charcoal present

RANGE: 0.01 μg to 0.3 μg per sample [2]

ESTIMATED LOD: 0.004 μg per sample

PRECISION ( ): 0.060 @ 0.012 μg to 0.11 μg per 

sample [2]

APPLICABILITY: The working range is 0.0003 ppm to 0.06 ppm (0.001 mg/m3 to 0.2 mg/m3) for a 10 L air sample. This is an 

elemental analysis and is not compound-specific.

INTERFERENCES: Use background correction to control molecular absorption. Other arsenic compounds (gases or 

aerosols) may be collected on the sampler and would be erroneously reported as arsine. A cellulose ester filter in front of 

the charcoal tube may be used to remove aerosols [3,4]. The effect of relative humidity on the capacity of charcoal for arsine 

has not been studied.

OTHER METHODS: This method combines and replaces NIOSH methods P&CAM 265 [5] and S229 [6] for arsine.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ARSINE: METHOD 6001, Issue 3, dated 20 October 2015 - Page 2 of 4

REAGENTS:

1. Water, distilled or deionized.

2. Nitric acid, concentrated.*

3. Nitric acid, 0.01 mol/L. Dilute 0.4 mL 

concentrated nitric acid to 1 L with water.

4. Nitric acid, 0.1 mol/L. Dilute 4 mL concentrated 

nitric acid to 1 L with water.

5. Arsenic stock solution, 1000 μg/mL arsenic.*

Commercial standard or dissolve 1.322 g dried, 

certified reagent arsenic trioxide in 100 mL 

of 0.1 mol/L nitric acid; dilute to 1 L with 0.1 

mol/L nitric acid.

6. Calibration stock solution, 1.0 μg/mL arsenic.*

Dilute 0.1 mL arsenic stock solution (1000 μg/

mL arsenic) to 100 mL with 0.01 mol/L nitric 

acid. Prepare fresh daily.

7. Nickel nitrate solution, 1000 μg/mL nickel. 

Commercial nickel atomic absorption standard 

or dissolve 3.112 g dried reagent nickel nitrate 

in 100 mL of 0.1 mol/L nitric acid; dilute to 1 L 

with water.

8. Argon, compressed.*

9. Arsine,* 99%, or certified mixture in nitrogen.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Activated coconut shell charcoal 

(100 mg/50 mg sections, 20/40 mesh), in a 

glass tube, 7 cm long, 6 mm OD, 4 mm ID, with 

flame-sealed ends. A silylated glass wool plug 

precedes the front section and urethane foam 

plugs separate the sorbent sections and follow 

the back section.

NOTE: Use a cellulose ester membrane filter 

in front of the sampler if particulate 

arsenic is present [3,4].

2. Personal sampling pump, 0.01 L/min to 0.2 L/

min, with flexible connecting tubing.

3. Graphite furnace atomic absorption 

spectrophotometer with non-pyrolytic tubes, 

background correction, and electrodeless 

discharge (and power supply) or hollow 

cathode lamp for arsenic.

4. Volumetric flasks, 1 L and 100 mL.†

5. Micropipets, 5 μL to 500 μL.†

6. Centrifuge tubes, 10 mL or 15 mL.†

7. Ultrasonic bath.

8. Centrifuge.

9. Syringe, gas, 0.1 mL, readable to 1 μL.

†

Clean all glassware with concentrated nitric 

acid and rinse thoroughly with distilled or 

deionized water before use.

SPECIAL PRECAUTIONS: Arsenic is a human carcinogen [7]. Wear gloves, lab coat, and safety glasses 

while handling acids. Perform all concentrated acid handling in a fume hood. Arsine is extremely 

poisonous by inhalation. Handle in well-ventilated hood and wear appropriate protective clothing 

and gloves. Users must be familiar with the proper use of flammable and nonflammable gases, 

cylinders, and regulators.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break the ends of the sampler immediately before sampling. Attach sampler to personal sampling 

pump with flexible tubing.

NOTE: Use a cellulose ester membrane prefilter if particulate arsenic compounds may be present 

[3,4].

3. Sample at an accurately known flow rate between 0.01 L/min and 0.2 L/min for a total sample size of 

0.1 L to 10 L.

4. Cap the sampler with plastic (not rubber) caps and pack securely for shipment.

SAMPLE PREPARATION:

5. Place the front and back sorbent sections of the sampler tube in separate centrifuge tubes. Discard 

the glass wool and foam plugs.

6. Add 1.0 mL of 0.01 mol/L nitric acid to each tube. Cap each tube.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ARSINE: METHOD 6001, Issue 3, dated 20 October 2015 - Page 3 of 4

7. Agitate for 30 min in an ultrasonic bath.

8. Centrifuge each tube.

CALIBRATION AND QUALITY CONTROL:

9. Calibrate daily with at least six working standards over the range 0.004 μg to 0.3 μg arsenic per 

sample.

a. Add known amounts of calibration stock solution and 0.01 mol/L nitric acid for a 1.0 mL final 

solution volume to centrifuge tubes containing 100 mg activated charcoal from a media blank 

sampler.

b. Analyze standards together with samples and blanks (steps 12 and 13). Analyze a working 

standard for every five samples to check for instrument drift.

c. Prepare a calibration graph (absorbance vs. μg arsenic).

10. Determine desorption efficiency (DE) at least once for each batch of charcoal used for sampling in 

the range 0.004 μg to 2 μg arsenic per sample. Prepare three tubes at each of five levels plus three 

media blanks.

a. Remove and discard back sorbent section of a media blank sampler.

b. Inject a known amount of pure arsine gas (or a certified gas mixture containing arsine) directly 

onto front sorbent section with a microliter syringe.

c. Cap the tube. Allow to stand overnight.

d. Desorb (steps 5, 6, 7, and 8) and analyze together with working standards (steps 12 and 13).

e. Prepare a graph of DE vs. μg arsenic recovered.

11. Analyze three quality control spikes to ensure that the calibration graph is in control.

MEASUREMENT:

12. Set the spectrophotometer and furnace to manufacturer’s recommendations and to conditions 

given on page 6001-1.

13. Inject a 50 μL aliquot of sample or standard followed by a 50 μL aliquot of nickel nitrate solution 

prior to initiating the analysis program. Measure peak area.

NOTE 1: If sample absorbance is above the linear range of the standards, dilute with 0.01 mol/L nitric 

acid, reanalyze and apply the appropriate dilution factor in calculations.

NOTE 2: Monitor the reproducibility of peak area for a working standard throughout the 

measurements. If erratic results occur, reoptimize instrumental parameters and replace the 

graphite tube.

CALCULATIONS:

14. Determine the mass, μg, of arsine found in the sample front ( ) and back ( ) sorbent sections, 

and in the average media blank front ( ) and back ( ) sorbent sections by multiplying the mass of 

arsenic found for each of these sections by 1.040 (MW of arsine/MW of arsenic).

NOTE: If , report breakthrough and possible sample loss.

15. Calculate concentration, , of arsine in the air volume sampled, (L):

, μg/L or mg/m3.

EVALUATION OF METHOD:

NIOSH method S229 [6] was evaluated over the range 0.094 mg/m3 to 0.404 mg/m3 using 10 L air 

samples collected on SKC Lot 105 activated coconut charcoal [1]. Breakthrough (onto the backup 

section) did not occur after 240 min of sampling at 0.227 L/min from an arsine concentration of 0.405 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ARSINE: METHOD 6001, Issue 3, dated 20 October 2015 - Page 4 of 4

mg/m3 (0.022 mg loading). The recovery was found to be 93.7%. Desorption efficiency was 0.90 at 1 μg 

arsine per sample and 1.00 at 2 μg and 4 μg arsine per sample.

NIOSH method P&CAM 265 [5] was evaluated over the range 0.001 mg/m3 to 0.01 mg/m3 using 15 L 

air samples [2]. These samples were collected on SKC Lot 106 activated coconut charcoal at a sampling 

flow rate of 0.875 L/min for 15 min. At this flow rate, a collection efficiency of 89.1% was found [3]. The 

effect of high humidity on the sampler capacity was not studied. Desorption efficiency was 0.90 in the 

range 0.015 μg to 0.2 μg arsine per sample.

REFERENCES:

[1] NIOSH [1977]. Backup data report: arsine, No. S229. By Taylor DG, Kupel RE, Bryant JM. In: 

Documentation of the NIOSH validation tests. Cincinnati, OH: U.S. Department of Health, Education, 

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, 

DHEW (NIOSH) Publication No. 77-185, available as accession No. PB274248 from NTIS [www.ntis.

gov].

[2] NIOSH [1977]. Evaluation and refinement of a personal sampling method for arsine. Cincinnati, OH: 

U.S. Department of Health, Education, and Welfare, Center for Disease Control, National Institute for 

Occupational Safety and Health, NIOSH research report, prepared under NIOSH Contract No. 210-

76-0142. Unpublished.

[3] Costello RJ, Eller PM, Hull RD [1983]. Measurement of multiple inorganic arsenic species. Am Ind 

Hyg Assoc J 44(1):21–28.

[4] NIOSH [1994]. Arsenic and compounds, as (except and ): Method 7900; Arsenic 

trioxide, as : Method 7901. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical methods. 

4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control 

and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 94-113 [www.cdc.gov/niosh/nmam/].

[5] NIOSH [1978]. Arsine in air: P&CAM 265. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd 

ed. Vol. 4. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 78-

175, available as accession No. PB83-105437 from NTIS [www.ntis.gov].

[6] NIOSH [1977]. Arsine: Method S229. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd ed. 

Vol. 3. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-157-C, 

available as accession No. PB276838 from NTIS [www.ntis.gov].

[7] NIOSH [1975]. Criteria for a recommended standard: occupational exposure to inorganic arsenic 

(new criteria - 1975). Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for 

Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication 

No. 75-149 [www.cdc.gov/niosh/docs/1970/75-149.html].

METHOD REVISED BY:

R. D. Hull, Ph.D., NIOSH/DBBS.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PHOSPHINE 6002

 MW: 34.00 CAS: 7803-51-2 RTECS: SY7525000

METHOD: 6002, Issue 3 EVALUATION: FULL Issue 1: 15 August 1994

Issue 3: 20 October 2015

OSHA: 0.3 ppm (0.4 mg/m3)

NIOSH: 0.3 ppm (0.4 mg/m3); 1 ppm (1 mg/m3) STEL

PROPERTIES: Gas, BP 87.8 °C; vapor density 1.17 (air 

= 1); spontaneously flammable in air if 

diphosphane is present

SYNONYMS: Hydrogen phosphide; phosphorus hydride; phosphorated hydrogen; phosphorus trihydride

SAMPLING

SAMPLER: SORBENT TUBE

(mercuric cyanide-coated silica gel, 300 

mg/150 mg)

FLOW RATE: 0.01 L/min to 0.2 L/min

VOL-MIN: 1 L @ 0.3 ppm

-MAX: 16 L

SHIPMENT: Routine

SAMPLE

STABILITY: 7 d @ 25 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: 0.195 mg/m3 to 0.877 mg/m3 [1] (16 L 

samples)

BIAS: 0.4%

OVERALL

PRECISION ( ): 0.091 @ 2.64 μg to 17.41 μg per sample 

[2]

ACCURACY: ±17.6%

MEASUREMENT

TECHNIQUE: UV-VIS SPECTROMETER

ANALYTE: Phosphate

EXTRACTION: 10 mL hot (65 °C to 70 °C) acidic 

permanganate reagent solution

DETECTOR: UV @ 625 nm

CALIBRATION: Standard solutions of potassium 

dihydrogen phosphate (1.00 mL contains 

phosphorus equivalent to 49.94 μg of 

phosphine)

RANGE: 0.3 μg to 10 μg per sample [2]

ESTIMATED LOD: 0.1 μg per sample [1]

PRECISION ( ): 0.074 @ 2.6 μg to 17.4 μg per sample [2]

APPLICABILITY: The working range is 0.013 ppm to 0.6 ppm (0.02 mg/m3 to 0.9 mg/m3) for a 16 L air sample.

INTERFERENCES: The colorimetric determination of phosphate is subject to interference by any species that also forms a 

molybdate complex under these conditions; possible interfering species include phosphorus trichloride and phosphorus 

pentachloride vapors and organic phosphorus compounds.

OTHER METHODS: This revises NIOSH method S332 [2]. OSHA method 1003, “Phosphine” [3], employing a mercuric 

chloride-treated polyester filter used to capture phosphine and ICP-AES to analyze for the total phosphorus, may be used as 

an alternative method.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PHOSPHINE: METHOD 6002, Issue 3, dated 20 October 2015 - Page 2 of 5

REAGENTS:

1. Potassium dihydrogen phosphate, anhydrous, 

ACS reagent grade.

2. Sulfuric acid,* concentrated, ACS reagent 

grade.

3. Ammonium molybdate tetrahydrate.

4. Ferrous ammonium sulfate, anhydrous.

5. Potassium permanganate.*

6. Stannous chloride.

7. Glycerol.

8. Toluene.*

9. Isobutanol.*

10. Methanol.*

11. Water, deionized or distilled.

12. Mercuric cyanide.*

13. Standard phosphate solution. Dissolve 200 mg 

anhydrous potassium dihydrogen phosphate 

in distilled water and dilute to 1 L (1.00 mL 

contains phosphorus equivalent to 49.94 μg of 

phosphine).

14. Molybdate solution. Dissolve 49.4 g of 

ammonium molybdate tetrahydrate and 

112 mL concentrated sulfuric acid in distilled 

water and dilute to 1 L.

15. Alcoholic sulfuric acid solution. Add 50 mL of 

concentrated sulfuric acid to 950 mL methanol.

16. Toluene-isobutanol solvent. Mix equal volumes 

of toluene and isobutanol.

17. Ferrous solution. Dissolve 7.9 g anhydrous 

ferrous ammonium sulfate and 1 mL 

concentrated sulfuric acid in distilled water 

and dilute to 100 mL.

18. Stannous chloride solution. Dissolve 0.4 g 

stannous chloride in 50 mL glycerol (heat to 

dissolve).

19. Acidic permanganate solution. Dissolve 

0.316 g potassium permanganate and 6 mL 

concentrated sulfuric acid in distilled water 

and dilute to 1 L.

20. Mercuric cyanide solution.* Dissolve 2 g 

mercuric cyanide in 100 mL water.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Sorbent tube, silica gel (mercuric 

cyanide), 300 mg/150 mg sorbent, 40/60 mesh; 

or, glass tube 12 cm long, 6 mm O.D., 4 mm I.D., 

flame-sealed ends with plastic caps, with two 

sections of mercuric cyanide-treated silica gel 

(45/60 mesh), (front = 300 mg, back = 150 mg), 

separated and retained by silylated glass wool 

plugs. (See APPENDIX.)

2. Personal sampling pump, 0.01 L/min to 0.2 

L/min, with flexible polyethylene or PTFE 

tubing.

3. Spectrometer capable of measuring 

absorbance or transmittance at 625 nm.

4. Two matched 5 cm absorbance cells, silica, 

with tight fitting caps.

5. Separatory funnel, 125 mL.

6. Beakers, 50 mL.

7. Pipets, 0.2 mL, 10 mL, and 25 mL, and other 

convenient sizes to make standard dilutions.

8. Volumetric flasks, 10 mL, 25 mL, 100 mL, and 

1000 mL.

9. Water bath (maintained at 65 °C to 70 °C).

10. Graduated cylinders, glass, 10 mL.

11. Syringes, 0.5 mL and 1.0 mL.

12. Balance.

13. Thermometer.

14. Stopwatch.

15. Barometer.

SPECIAL PRECAUTIONS: Caution should be exercised when preparing the sampling media because 

mercuric cyanide is toxic by inhalation, ingestion, and skin contact. Contact of mercuric cyanide with 

acid will produce hydrogen cyanide gas. Concentrated sulfuric acid is highly corrosive. All work with 

these compounds should be performed in a hood. Use proper protective clothing including gloves, 

safety glasses, and laboratory coat. Potassium permanganate is a strong oxidizer with risk of fire and 

explosion upon contact with combustible substances and reducing agents. Toluene, isobutanol, and 

methanol are flammable.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PHOSPHINE: METHOD 6002, Issue 3, dated 20 October 2015 - Page 3 of 5

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Immediately before sampling, break the ends of the silica gel tubes to provide an opening of at least 

one half the internal diameter of the tube. Attach the silica gel tube to the sampling pump with 

flexible tubing.

3. Sample at an accurately known flow rate between 0.01 L/min and 0.2 L/min for a total sample size of 

1 L to 16 L.

4. Seal tubes with plastic (not rubber) caps.

SAMPLE PREPARATION:

5. Place front and back sorbent sections in separate 50 mL beakers.

6. Add 10 mL of acidic permanganate solution to each beaker. Place in a water bath maintained at 

65 °C to 70 °C for 90 min.

7. Decant the acidic permanganate solution into a 10 mL volumetric flask, and dilute to volume with 

distilled water.

8. Wash the silica gel twice with 3 mL portions of distilled water and decant the contents into another 

10 mL volumetric flask containing 1 mL of ferrous solution. Dilute to volume with distilled water.

9. Add the contents of both 10 mL volumetric flasks (extract and washings) to a 125 mL separatory 

funnel.

10. Add 7.5 mL of molybdate reagent and 25 mL of toluene-isobutanol solvent to the funnel. Shake 

funnel for 60 s. Let the separatory funnel stand for 60 s to allow the aqueous and nonaqueous layers 

to separate. Discard the lower (aqueous) layer.

11. Pipet 10 mL of the nonaqueous layer into a 25 mL volumetric flask containing 10 mL of the alcoholic 

sulfuric acid solution.

CALIBRATION AND QUALITY CONTROL:

12. Calibrate daily with at least six working standards.

a. Add 10 mL of acidic permanganate solution and 1 mL of ferrous reagent to a 125 mL separatory 

funnel.

b. Add 2 μL to 400 μL of the standard phosphate solution to cover the range 0.1 μg to 10 μg of 

phosphine. Add 8 mL to 9 mL of water to make the total volume of the solution (permanganate 

solution, ferrous solution, phosphate solution and water) equal to 20 mL. Prepare at least six 

calibration standards and a blank containing no phosphate.

c. Add 7.5 mL of molybdate reagent and 25 mL of toluene-isobutanol solvent to the funnel. Shake 

funnel for 60 s. Let the separatory funnel stand for 60 s to allow the aqueous and nonaqueous 

layers to separate. Discard the lower (aqueous) layer. (step 10)

d. Pipet 10 mL of the nonaqueous layer into a 25 mL volumetric flask containing 10 mL of the 

alcoholic sulfuric acid solution. (step 11)

e. Analyze with samples and blanks (steps 15, 16, 17, and 18).

f. Prepare a calibration graph (absorbance versus μg of phosphine added).

13. Determine desorption efficiency (DE) at least once for each lot of sorbent used for sampling in the 

range of interest. Prepare three tubes at each of five levels plus three media blanks.

a. Remove and discard back sorbent section of a media blank sampler.

b. Inject a known amount (20 μL to 400 μL) of standard phosphate solution directly onto front 

sorbent section with a microliter syringe.

c. Cap the tube. Allow to stand overnight.

d. Desorb (steps 6, 7, and 8) and analyze with working standards (steps 15, 16, 17, and 18).

e. Prepare a graph of DE vs. μg recovered.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PHOSPHINE: METHOD 6002, Issue 3, dated 20 October 2015 - Page 4 of 5

14. Analyze three quality control blind spikes and three analyst spikes to ensure that the calibration 

graph and DE graph are in control.

MEASUREMENTS

15. Turn on the spectrophotometer and allow sufficient time for warmup. Adjust the wavelength to 625 

nm and set the zero and 100% transmittance scale using 5 cm cells filled with distilled water. Check 

these settings prior to making any measurement to check on instrument drift.

NOTE: Steps 16, 17, and 18 must be performed within 1 min.

16. Add 0.5 mL (25 drops) of stannous chloride reagent and dilute to volume using alcoholic sulfuric 

acid solution. Mix thoroughly.

17. Transfer the sample into a 5 cm cell and stopper immediately.

18. Measure the absorbance or transmittance using water as a blank.

CALCULATIONS:

19. Determine mass, μg (corrected for DE), of phosphine found in the sample front ( ) and back ( ) 

sorbent sections, and in the average media blank front ( ) and back ( ) sorbent sections.

NOTE: If , report breakthrough and possible sample loss.

20. Calculate concentration, , of phosphine in the air volume sampled, (L).

, μg/L or mg/m3.

EVALUATION OF METHOD:

This method was validated over the range 0.195 mg/m3 to 0.877 mg/m3 at 19 °C and 102.0 kPa (765.3 

mmHg) using 16 L samples [1]. Desorption efficiency must be determined over the range used. The 

upper range of the method depends on the adsorptive capacity of the mercuric cyanide-treated gel. 

This capacity may vary with the concentration of phosphine and other substances in the air. When an 

atmosphere at 90% relative humidity containing 0.957 mg/m3 of phosphine was sampled at a flow rate 

of 0.2 L/min, breakthrough was determined to occur at a sample volume of 20.75 liter (capacity = 19.86 

μg phosphine).

REFERENCES:

[1] NIOSH [1979]. Backup data report: phosphine, No. S332. In: Ten NIOSH analytical methods, set 7. 

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, available as accession No. PB84-243534 from 

NTIS [www.ntis.gov]. Unpublished.

[2] NIOSH [1979]. Phosphine: Method S332. In: Taylor DG, ed. NIOSH manual of analytical methods. 

2nd ed. Vol. 5. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 

79-141, available as accession No. PB83-105445 from NTIS [www.ntis.gov].

[3] OSHA [2000]. Phosphine: OSHA Method 1003 [www.osha.gov/dts/sltc/methods/mdt/

mdt1003/1003.html]. Date accessed: October 2015.

METHOD REVISED BY:

Charles Lorberau, NIOSH/DPSE.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PHOSPHINE: METHOD 6002, Issue 3, dated 20 October 2015 - Page 5 of 5

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.

APPENDIX: PREPARATION OF SAMPLING MEDIA

Coating of Silica Gel

1. Dry 100 g of silica gel (45/60 mesh) at 90 °C for 2 h.

2. Prepare a mercuric cyanide solution in distilled water (2 g mercuric cyanide in 100 mL water).

3. Add the dried silica gel to the mercuric cyanide solution and let set for 15 min with occasional

stirring.

4. Drain the excess mercuric cyanide solution and dry the remaining silica gel at 90 °C for 3 h.

5. Cool the silica gel to room temperature in a covered beaker.

6. Expose the silica gel to a humid atmosphere (>80% RH) for 24 h.

Preparing the Sampling Tubes

7. Place a plug of silylated glass wool at the end of a glass tube (6 mm OD and 4 mm ID) about 12 cm

long. Pour 300 mg of the treated silica gel into the tube. Place another plug of silylated glass wool

behind this front section. Add another 150 mg of the treated silica gel into the tube. Place a final

plug of silylated glass wool behind this backup section.

8. Check the pressure drop of a representative sampler. The pressure drop across the tube must be less

than 6.7 kPa (50 mmHg) at a flow rate of 0.2 L/min.

9. Flame seal both ends of the sampling tubes.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

HYDROGEN CYANIDE 6010

HCN MW: 27.03 CAS: 74-90-8 RTECS: MW6825000

METHOD: 6010, Issue 3 EVALUATION: FULL Issue 1: 16 May 1989

Issue 3: 30 June 2017

OSHA: 10 ppm (11 mg/m3

) (skin)

NIOSH: STEL 4.7 ppm (5 mg/m3) (skin)

IDLH: 50 ppm [1]

OTHER OELs: Refs. [2, 3] 

PROPERTIES: gas; BP 26 oC, vapor density 0.93 (air = 1.00); 

d(liq) 0.69 g/mL @ 20 oC, VP 82.7 kPa (620 mm Hg @ 20 oC); 

explosive range 5 to 40% v/v in air 

SYNONYMS: Hydrocyanic acid, prussic acid, formonitrile

SAMPLING

SAMPLER: SOLID SORBENT TUBE (soda lime, 600 

mg/200 mg), with glass fiber pre-filter

FLOW RATE: 0.025 to 0.05 L/min

VOL-MIN: 2 L @ 5 ppm

 -MAX: 90 L

SHIPMENT: routine

SAMPLE

STABILITY: at least two weeks at 25 oC [4]

BLANKS: 3 field blanks min. per set

ACCURACY

RANGE 

STUDIED: 2 to 15 mg/m3 (3-L samples) [4]

BIAS: negligible

OVERALL

PRECISION (SS�rrrr): 0.076 [4]

ACCURACY: ±15%

MEASUREMENT

TECHNIQUE: SPECTROPHOTOMETRY, visible absorption

ANALYTE: cyanide ion complex

DESORPTION: 10 mL deionized water; stand 60 min

COLOR 

DEVELOPMENT: N-chlorosuccinimide/succinimide 

oxidizing agent and barbituric 

acid/pyridine coupling agent; absorption 

@ 580 nm in 1-cm cuvette

 

CALIBRATION: standard solutions of KCN in 1 M NaOH

RANGE: 10 to 300 μg CN- per sample [4]

ESTIMATED LOD: 1 μg CN- per sample [4]

PRECISION (SS�rr): 0.041 @ 10 to 50 μg CN- per sample [4]

APPLICABILITY: The method is applicable to STEL measurements of HCN gas. The working range is 0.3 to 235 ppm (3 to 260 

mg/m3

) for a 3-L air sample. Particulate cyanides are trapped by an initial glass fiber filter (pre-filter) that precedes the sorbent tube. 

This method is more sensitive and subject to fewer interferences than NIOSH Method 7904 [5], which uses ion specific electrode 

analysis. The method has been used to measure HCN in firefighting environments [6].

INTERFERENCES: A high concentration of hydrogen sulfide (H2S) can cause a negative interference by reaction with cyanide to 

form thiocyanate. 

OTHER METHODS: The procedure is based on the method of Lambert et al. [7]. NIOSH Method 7904 relies on the use of ion 

selective electrodes for measurement of CN- [5]. A related method, which employs diffusive sampling onto soda lime sorbent, 

followed by desorption and ion chromatography with electrochemical detection, has been promulgated by OSHA [8].
HYDROGEN CYANIDE: METHOD 6010, Issue 3, dated 30 June 2017 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Potassium cyanide,* reagent grade.

2. Succinimide, reagent grade.

3. N-Chlorosuccinimide, reagent grade.

4. Barbituric acid, reagent grade.

5. Pyridine, spectroscopic quality.

6. Phenolphthalein, 1% (w/v) in ethanol or

methanol, reagent grade.

7. Hydrochloric acid, conc., reagent grade.

8. Sodium hydroxide, reagent grade.

9. Soda lime (CaO + 5-20% NaOH), reagent

grade. Crush and sieve to 10/35 mesh; store

in a sealed container.

10. Water, deionized (18 MΩ-cm).

11. NaOH solution, 0.1 M.

12. Calibration stock solution, 1 mg/mL CN-

:

Dissolve 0.125 g KCN in ≈40 mL 0.1 M NaOH

in a 50-mL volumetric flask. Dilute to the

mark with 0.1 M NaOH. Standardize by

titration with AgNO3 (see APPENDIX).

13. HCl solution, 0.15 M.

14. N-Chlorosuccinimide/succinimide oxidizing

reagent: Dissolve 10.0 g succinimide in ≈200

mL deionized (DI) water. Add 1.00 g Nchlorosuccinimide; agitate to dissolve.

Adjust volume to 1 L with DI H2O. Solution is

stable for 6 months when kept at ≈4 o

C.

15. Barbituric acid/pyridine reagent: Add ≈30

mL DI H2O to 6.0 g barbituric acid in a 100-

mL Erlenmeyer flask. Slowly add 30 mL

pyridine with stirring. Adjust volume to 100

mL with DI H2O. Solution is stable for 2

months when kept at ≈4 o

C.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: glass tube, length 9 cm, 7-mm

OD/5-mm ID, with plastic caps, containing

two sorbent sections of granular soda lime

10/35 mesh (front = 600 mg; back = 200 mg),

separated by and contained within silanized

glass wool plugs. A 5-mm dia. glass fiber filter

is placed before the inlet.

NOTE: Tubes are commercially available.

2. Spectrophotometer, Visible (580 nm), with 1-

cm light path cuvettes.

3. Personal sampling pump, 0.025-0.05 L/min,

with flexible connecting tubing.

4. Pipets, volumetric, 0.1-, 0.5-, 1.0-, 2.0-, 10.0-

mL.

5. Vials, glass or plastic, 15-mL, with

polytetrafluoroethylene (PTFE)-lined caps.

6. Flasks, volumetric, 25-, 50-, 100-, and 1000-

mL, with stoppers.

7. Pipets, transfer, disposable.

8. Laboratory wipes.

9. Syringes, 10-mL, polyethylene with Luer tip

readable to 0.1 μL.

10. Flask, Erlenmeyer, 100-mL.

11. Luer-lock PTFE membrane filters, 13-mm dia.,

0.45-μm pore size, to fit onto 10-mL syringes.

SPECIAL PRECAUTIONS: HCN gas and cyanide are highly toxic and may be fatal if swallowed, inhaled or 

absorbed through skin [1]. Soda lime and NaOH are very caustic [1]. Wear appropriate personal 

protection during sampling activities and analysis. It is essential that suitable gloves, eye protection, 

laboratory coat, etc., be used when working with these chemicals. Perform sample preparation and 

analysis in a clean, well-ventilated area that is well removed from any possible contamination. 

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in the line.

2. Break both ends of the sampling tube immediately prior to sampling. Attach the glass fiber filter to

the inlet of the sorbent tube. Attach sampler to personal sampling pump with flexible tubing.

3. Sample at an accurately known flow rate between 0.025 and 0.05 L/min (e.g., 0.033 L/min) for a total

sample size of 0.6 to 90 L.
HYDROGEN CYANIDE: METHOD 6010, Issue 3, dated 30 June 2017 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

4. Remove the sorbent tube from the sampling apparatus, cap both ends and pack securely for shipment

to the laboratory. Discard the pre-filter.

SAMPLE PREPARATION:

5. Score each sample tube with a file and break sampler at the score line.

6. Transfer front and back sorbent sections to separate 15-mL vials. Discard glass wool plugs. 

7. Add 10.0 mL DI H2O to each vial containing a sorbent section. Seal each vial.

8. Allow to stand 60 min. with occasional agitation. Transfer to a 10-mL plastic syringe fitted with a 0.45-

μm PTFE filter, and collect the filtrate in a clean vial.

CALIBRATION AND QUALITY CONTROL:

9. Calibrate daily with at least six working standard solutions over the range 1 to 300 μg CN- per sample.

a. Prepare a working standard solution, 1.00 μg/mL CN-

, by diluting 100 μL of calibration stock 

solution to 100 mL with 0.1 M NaOH.

b. Pipet 0.50, 1.00, 1.50, 2.00, and 2.50 mL of the working standard solution into 25-mL volumetric 

flasks to make up 0.50-, 1.00-, 1.50-, 2.00-, and 2.50-μg CN- standards.

c. Analyze together with field samples and blanks (steps 12 through 19).

d. Prepare calibration graph (absorbance vs. μg CN-

).

10. Determine desorption efficiency (DE) at least once for each lot of soda lime used for sampling. Prepare 

at least three tubes at each of five levels plus three media blanks.

a. Remove and discard back sorbent section of an unused (blank) sampler.

b. Inject a known amount of calibration stock solution directly onto the soda lime with a microliter 

syringe.

c. Cap and allow to stand overnight.

d. Desorb (steps 5 through 8) and analyze together with working standards and blanks (steps 12 

through 18).

e. Prepare a graph of DE vs. μg CN- recovered. 

11. Analyze at least three quality control blind spikes and at least three analyst spikes to ensure that the 

calibration graph and DE graph are in control.

MEASUREMENT:

12. Set spectrophotometer to record absorbance at 580 nm. 

13. Pipet a sample aliquot estimated to contain 0.5 to 2.5 μg CN- into a 25-mL volumetric flask. 

Alternatively, to cover an unknown sample range, pipet 0.50-, 1.00- and 3.00-mL aliquots into separate 

25-mL volumetric flasks for each field sample. Larger or smaller aliquots may be taken, based on prior 

knowledge of the expected analyte level.

14. Pipet 0.5 mL 0.1 M NaOH into a 25-mL volumetric flask for each standard or sample. 

15. Add one drop of phenolphthalein solution to each standard and sample.

NOTE: Add a small portion of DI H2O to increase volume for easier mixing. All solutions should be 

alkaline (pink) at this juncture.

16. Starting with the reagent blank, add 0.15 M HCl dropwise, with mixing, until reaching the point at 

which the pink color just disappears. CAUTION: HCN may be produced, so this step must be carried 

out in a fume hood. Immediately add 1.0 mL N-chlorosuccinimide/succinimide oxidizing reagent. Mix 

and let stand.

NOTE 1: To avoid possible loss of HCN, add the oxidizing agent before proceeding to the next sample.

NOTE 2: Do not prepare more samples than can be analyzed within the 30-minute maximum time for 

color development.

17. After at least 5 min. standing (but not longer than 15 min.), starting with the reagent blank, add 1.0 mL 

of barbituric acid/pyridine coupling reagent and mix.
HYDROGEN CYANIDE: METHOD 6010, Issue 3, dated 30 June 2017 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

18. Adjust sample volume to 25 mL with DI H2O and allow to stand for at least 12 min (but no longer than

30 min.) for color development.

19. Using the spectrophotometer, read the absorbance at 580 nm using a 1-cm light path cuvette. If

sample absorbance is outside the range of the calibration standards, remove an aliquot, dilute and

reanalyze. Apply appropriate dilution factor in calculations.

CALCULATIONS:

20. From the absorbance readings, calculate the mass of CN-

 (μg) in each aliquot analyzed. Apply the

appropriate dilution factor to calculate the mass of CN- (μg) in the original 10-mL solution.

21. Determine the mass of CN- (μg), corrected for desorption efficiency (DE), found in the front (Wf) and

back (Wb) sections of the sorbent tube. Also determine the average background mass of media blank

front (Bf) and back (Bb) sorbent sections. If Wb > Wf/10, report breakthrough and possible sample loss.

22. Calculate the concentration, C, of HCN in the air volume sampled, V(L):

CC = �WWff + WWbb − BBff − BBbb� ∗ 1.039

VV , mmmm/mm3

(where 1.039 = conversion factor for CN- to HCN). 

EVALUATION OF METHOD:

The method was initially evaluated by sampling test atmospheres of HCN generated from a compressed 

gas mixture of HCN in nitrogen [4]. The range of HCN concentrations was equivalent to 2 to 15 mg/m3 for a 

3-L air sample. For 22 samples, an overall precision S�rT of 0.076 was obtained with nearly 100% recovery.

Breakthrough was observed to occur at [HCN] ≈150 mg/m3 for sample volumes ≈10 L. Sample tubes

spiked with KCN standard solutions (n=22) in the range of 10 to 50 μg per sample indicated recoveries of

nearly 100% with a pooled precision S�r of 0.041. Sample tubes similarly spiked with KCN solutions were

analyzed after storage and demonstrated that the samples were stable for at least two weeks. Sample

collection of [HCN] in nitrogen of 130 mg/m3 at 50 mL/min for 7.3 minutes (0.36 L sample volume) resulted

in quantitative recoveries [9]. Further evaluation of the method was carried out in test atmospheres of

HCN generated in air of low (20%) and high (80%) relative humidity [9]. It was found that a sampling rate

of 50 mL/min resulted in quantitative recoveries for HCN concentrations at 0.1 – 2× the OSHA PEL, while

sampling at 200 mL/min gave recoveries <90% in this concentration range; similar results were obtained

for both low and high humidity conditions. In consideration of these observations, the maximum

sampling flow rate recommended for the method is 0.05 L/min.

REFERENCES:

[1] NIOSH [2016]. NIOSH Pocket guide to chemical hazards. Cincinnati, OH: U.S. Department of Health

and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health, DHHS (NIOSH) Publication No. 2010-168c, http://www.cdc.gov/niosh/npg/.

[2] ACGIH [2017] TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, OH: American

Conference of Governmental Industrial Hygienists.

[3] Institute for Occupational Safety and Health of the German Social Accident Insurance (IFA) [2017].

GESTIS international limit values, http://limitvalue.ifa.dguv.de/.

[4] Williamson G [1988]. Method development protocol and backup data report on hydrogen cyanide,

internal report. Cincinnati, OH: NIOSH. Unpublished.
HYDROGEN CYANIDE: METHOD 6010, Issue 3, dated 30 June 2017 - Page 5 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[5] NIOSH [1994]. Method 7904 – Cyanide, aerosol and gas. In: Schlecht PC, O’Connor PF, eds. NIOSH

manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services,

Centers for Disease Control and Prevention, National Institute for Occupational Safety and Health,

DHHS (NIOSH) Publication No. 94-113.

[6] Williamson G [1988]. Analysis of air samples on project 166 (Fire smoke) on HCN, sequence report

6366-A. Cincinnati, OH: NIOSH. Unpublished.

[7] Lambert JL, Ramasamy J, Paukstelis JV [1975]. Stable reagents for the colorimetric determination of

cyanide by modified König reactions. Anal Chem 47:916-918.

[8] OSHA [2010]. Method no. 1015 (HCN). In: OSHA sampling and analytical methods manual. Sandy, UT:

U.S. Department of Labor, Occupational Safety and Health Administration,

http://www.osha.gov/dts/sltc/methods.

[9] Simmons M [2010]. Use of active sorbent soda lime tube for the collection of hydrogen cyanide,

internal report. Sandy, UT: Department of Labor, Occupational Safety and Health Administration.

Unpublished.

METHOD AUTHORED BY:

George Williamson, NIOSH/DART (ret.)

REVISED BY:

Kevin Ashley, NIOSH and Mike Simmons, OSHA/SLTC

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.

APPENDIX: STANDARDIZATION OF CYANIDE CALIBRATION STOCK SOLUTION

Titrate an aliquot of the cyanide standard stock solution (KCN in NaOH) with standard silver nitrite (AgNO3) 

solution. The end point is the first formation of a white precipitate, Ag[Ag(CN)2]. Calculate the cyanide 

concentration by using the following equation:

MMcc = 52.04 ∗ VVaa ∗ �MMaa

VVcc

�

where: Mc = cyanide concentration (mg/mL)

Va = volume (mL) of standard AgNO3 solution

Ma = concentration (moles/L) of standard AgNO3 solution

Vc = volume (mL) of cyanide calibration stock solution titrated
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SULFURYL FLUORIDE 6012

 MW: 102.06 CAS: 2699-79-8 RTECS: WT5075000

METHOD: 6012, Issue 2 EVALUATION: FULL Issue 1: 15 August 1994

Issue 2: 12 January 2016

OSHA: 5 ppm

NIOSH: 5 ppm; STEL 10 ppm

(1 ppm = 4.17 mg/m3 @ NTP)

PROPERTIES: Gas; BP −55 °C; vapor density (air = 1) 3.5; 

VP 1.7 × 103 kPa; nonflammable, colorless, 

odorless [1]

SYNONYMS: Sulfonyl difluoride, sulfur difluoride dioxide, sulfuric oxyfluoride

SAMPLING

SAMPLER: SOLID SORBENT TUBE

(coconut shell charcoal, 800 mg/200 mg)

FLOW RATE: 0.05 L/min to 0.1 L/min

VOL-MIN: 1.3 L @ 5 ppm

-MAX: 10 L

SHIPMENT: Ship at 0 °C

SAMPLE

STABILITY: At least 12 d @ 0 °C

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: 20 mg/m3 to 420 mg/m3 (0.2 L to 6 L 

samples)

BIAS: −3.0%

OVERALL

PRECISION ( ): 0.070 [2]

ACCURACY: ±16.7%

MEASUREMENT

TECHNIQUE: ION CHROMATOGRAPHY

CONDUCTIVITY DETECTION

ANALYTE: Fluoride ion

EXTRACTION: 20 mL 40 mmol/L sodium hydroxide; 

sonicate 60 min

INJECTION: 50 μL

ELUENT: 40 mmol/L sodium hydroxide, 1.0 mL/

min

COLUMN: US Pharmacopeia (USP) L12 separator 

column, manufacturer’s compatible anion 

guard column, and micromembrane 

suppressor as recommended by the 

manufacturer. See OTHER METHODS.

DETECTOR: Conductivity, 30 μS full scale

CALIBRATION: Standard solutions of fluoride ion spiked 

onto sample media

RANGE: 10 μg to 80 μg fluoride per sample [3]

ESTIMATED LOD: 7 μg sulfuryl fluoride per sample [3]

PRECISION ( ): 0.052 (27 μg to 420 μg sulfuryl fluoride 

per sample) [2]

APPLICABILITY: The working range is 2.2 ppm to 17 ppm (9 mg/m3 to 75 mg/m3) for a 3 L air sample. This method is 

applicable to STEL measurements using a 1.5 L sample. The method has been used to sample for sulfuryl fluoride at 

dwelling fumigation sites [3,4].

INTERFERENCES: Other fluoride compounds may interfere.

OTHER METHODS: This method is based on the method of Bouyoucos, et al. [5]. NIOSH method S245 uses gas bag samples, 

gas chromatography-flame photometric detector (GC-FPD) [6].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SULFURYL FLUORIDE: METHOD 6012, Issue 2, dated 12 January 2016 - Page 2 of 4

REAGENTS:

1. Sodium hydroxide,* ACS reagent grade.

2. Sulfuric acid,* concentrated, ACS reagent 

grade.

3. Water, high-purity.

4. Desorbing/extracting solution and eluent: 

40 mmol/L sodium hydroxide. Dissolve 3.2 g 

sodium hydroxide in 2 L of degassed highpurity water.

5. Suppressor regenerant, 12.5 mmol/L sulfuric 

acid. Add 0.70 mL concentrated sulfuric acid to 

1 L of high-purity water.

6. Calibration stock solution, 1 mg/mL fluoride 

anion. Dissolve 0.2210 g sodium fluoride in 

high-purity water and dilute to the mark in a 

100 mL volumetric flask.

7. Sulfuryl fluoride* calibration gas standard(s) 

(optional).

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Activated coconut shell charcoal 

sampling tube; glass tube, 11 cm long, 10 mm 

OD, 7 mm ID, flame-sealed ends, containing 

two sections of activated (600 °C) coconut 

shell charcoal (front = 800 mg, back = 200 

mg), separated by a 2 mm urethane foam 

plug. A silylated glass wool plug precedes the 

front section and a 3 mm urethane foam plug 

follows the back section. Pressure drop across 

the tube at 1 L/min airflow must be less than 

3.4 kPa.

2. Personal sampling pump, 0.05 L/min to 0.10 

L/min, with flexible polyethylene or PTFE 

tubing.

3. Refrigerant, water solution, sealed, refreezable, 

reusable.

4. Filter, membrane, 0.45 μm pore size, 13 mm, 

with Luer fitting.

5. Ion chromatograph, with a conductivity 

detector, chart recorder, integrator, and 

columns (page 6012-1).

6. Vials, glass, 20 mL, with plastic caps.

7. Vials, polyethylene, 20 mL, with plastic caps.

8. Micropipettes, with disposable plastic tips.

9. Volumetric flasks, 100 mL.

10. Pipet, 10 mL, graduated in 0.1 mL intervals.

11. Pipet, volumetric, 20 mL.

12. Syringes, 10 mL, plastic, with Luer tip.

13. Sonicator.

14. Analytical balance, to ±0.0001 g.

SPECIAL PRECAUTIONS: Sulfuryl fluoride is a restricted use pesticide owing to its inhalation toxicity. 

It is extremely hazardous as vapor or liquid. Inhalation of vapors may be fatal. Read and follow all 

label precautions [7]. Sulfuric acid and sodium hydroxide are corrosive to skin, eyes, and mucous 

membranes. Use proper protective clothing including gloves, safety glasses, and laboratory coat. 

Handle all hazardous chemicals in a fume hood.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Break the ends of the sampler immediately before sampling. Attach a sampler to personal sampling 

pump with flexible tubing.

3. Sample at an accurately known flow rate between 0.05 L/min and 0.1 L/min for a total sample 

volume of 1.3 L to 10 L.

4. Cap the samplers with plastic (not rubber) caps and pack securely for shipment at 0 °C.

SAMPLE PREPARATION:

5. Place the front and back sorbent sections of the sampler tube in separate 20 mL plastic vials. Discard 

the glass wool and foam plugs.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SULFURYL FLUORIDE: METHOD 6012, Issue 2, dated 12 January 2016 - Page 3 of 4

6. Add 20 mL 40 mmol/L sodium hydroxide eluent to each plastic vial. Cap. Sonicate for 60 min.

7. Transfer a 5 mL to 7 mL aliquot to a tare weighted 20 mL glass vial using a plastic syringe fitted with 

a 0.45 μm membrane filter.

8. Reweigh each glass vial and contents so that the net weight of the aliquot can be calculated.

9. Take each sample to complete dryness in an uncapped glass vial on a hot plate. Cool, then 

reconstitute to the original net weight with high-purity water.

CALIBRATION AND QUALITY CONTROL:

10. Calibrate daily with at least six working standards.

NOTE: Standards should be spiked onto charcoal tubes as follows to avoid high recoveries seen with 

liquid standards [4].

a. Add known amounts of calibration stock solution onto charcoal tubes (5.0 μg to 80 μg fluoride) 

and desorb in the same manner as field samples (steps 5, 6, 7, 8, and 9).

b. Analyze working standards together with samples and blanks (steps 12, 13, and 14).

c. Prepare a calibration graph of peak height vs. amount (μg) of fluoride per 20 mL of sample.

11. (Optional). Determine recovery ( ) for each lot of tubes used for sampling in the concentration 

range of interest. Prepare four tubes at each of five levels plus three media blanks.

a. Collect a known amount of sulfuryl fluoride gas onto each charcoal tube (steps 1, 2, 3, 4, 5, 6, 7, 8, 

and 9).

b. Analyze samples in the same manner as field samples (steps 12, 13, and 14).

c. Prepare graph of recovery vs. μg sulfuryl fluoride.

MEASUREMENT:

12. Set ion chromatograph to conditions given on page 6012-1.

13. Refilter sample if necessary, then inject a sample aliquot into the ion chromatograph.

14. Measure peak height.

CALCULATIONS:

15. Determine mass (μg) of fluoride found on the front ( ) and back ( ) sections, and in the average 

media blank front ( ) and back ( ) sorbent section.

NOTE: If , report breakthrough and possible sample loss.

16. Calculate concentration, , of sulfuryl fluoride in the actual air volume, (L), applying the 

conversion factor 2.686 (molecular weight of sulfuryl fluoride divided by the atomic weight of 2 

fluoride anions; the reaction is ):

, μg/L or mg/m3.

EVALUATION OF METHOD:

This method was evaluated over the range 20 mg/m3 to 420 mg/m3. Overall sampling and 

measurement precision, , was 0.070 [2]. The average recovery of sulfuryl fluoride from charcoal 

was 99% when sampling atmospheres prepared in aluminized gas bags (Calibrated Instruments, Inc., 

Hawthorne, NY 10532). Recovery of fluoride from sampling media was 97% in the range 10 μg to 160 μg 

fluoride per sample. Sample stability during storage was evaluated at an air concentration of 417 mg/m3 

sulfuryl fluoride. Samples showed 101% recovery after 12 d of storage at 0 °C to 5 °C compared to oneday-old samples.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SULFURYL FLUORIDE: METHOD 6012, Issue 2, dated 12 January 2016 - Page 4 of 4

REFERENCES:

[1] CDPR [2006]. Exposure assessment document for pesticide products containing sulfuryl fluoride. By 

Cochran R, DiPaolo D. Sacramento, CA: California Environmental Protection Agency, Department of 

Pesticide Regulation [www.cdpr.ca.gov/docs/emon/pubs/tac/tacpdfs/sulfluor/final_rcd_vol2.pdf]. 

Date accessed: January 2016.

[2] NIOSH [1991]. Backup data report for sulfuryl fluoride. By Williamson GY. Cincinnati, OH: U.S. 

Department of Health and Human Services, Centers for Disease Control, National Institute for 

Occupational Safety and Health, Division of Physical Sciences and Engineering, Methods Research 

Support Branch. Unpublished.

[3] NIOSH [1991]. Analytical report: sequence No. 7161-A. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control, National Institute for Occupational Safety and 

Health. Unpublished.

[4] NIOSH [1993]. Analytical report: sequence No. 7691-D. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health. Unpublished.

[5] Bouyoucos SA, Melcher RG, Vaccaro JR [1983]. Collection and determination of sulfuryl fluoride in 

air by ion chromatography. Am Ind Hyg Assoc J 44(1):57–61.

[6] NIOSH [1980]. Sulfuryl fluoride: Method S245. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. Vol. 6. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 80-125, available as accession No. PB82-157728 from NTIS [www.ntis.gov].

[7] NIOSH/OSHA [1981]. Occupational health guideline for sulfuryl fluoride. In: NIOSH/OSHA 

Occupational health guidelines for chemical hazards. Cincinnati, OH: U.S. Department of Health and 

Human Services, Centers for Disease Control, National Institute for Occupational Safety and Health, 

DHHS (NIOSH) Publication No. 81-123 [www.cdc.gov/niosh/docs/81-123/].

METHOD WRITTEN BY:

George Y. Williamson, MRSB, DPSE.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

AMMONIA by IC 6016

NH3 MW: 17.03 CAS: 7664-41-7 RTECS: BO0875000

METHOD: 6016, Issue 2 EVALUATION: FULL Issue 1: 15 May 1996

Issue 2: 3 March 2016

OSHA: 50 ppm

NIOSH: 25 ppm; STEL 35 ppm

PROPERTIES: gas; MP -77.7 C; BP -33.4 C; VP 888 kPa (8.76 

atm) @ 21.1 C; vapor density 0.6 (air = 1); 

explosive range 16 to 25% v/v in air 

SYNONYMS: none

SAMPLING

SAMPLER: SOLID SORBENT TUBE (sulfuric acidtreated silica gel); 

a 0.8-μm MCE prefilter may be used to 

remove particulate interferences.

FLOW RATE: 0.1 - 0.5 L/min

VOL-MIN: 0.1 L @ 50 ppm

 -MAX: 96 L @ 50 ppm {1}

SHIPMENT: routine

SAMPLE 

STABILITY: at least 35 days @ 5 °C [2] 

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 17 to 68 mg/m3 [1] (30-L samples)

BIAS: -2.4% 

OVERALL

PRECISION (SS�rrrr): 0.071 [1] 

ACCURACY: ± 14.5% 

MEASUREMENT

TECHNIQUE: ION CHROMATOGRAPHY,

CONDUCTIVITY DETECTION

ANALYTE: ammonium ion (NH4

+)

EXTRACTION ION: 10 mL deionized water

INJECTION 

VOLUME: 50 μL

ELUENT: 48 mM HCl/4 mM DAP-HCl/4 mM 

L-histidine-HCl; 1 mL/min

alternate: 12 mM HCl/0.25 mM DAPHCl/0.25 mM L-histidine-HCl; 1 mL/min

COLUMNS: cation separator; cation guard; cation 

micromembrane suppressor

CONDUCTIVITY

 SETTING: 30 μS full scale 

CALIBRATION: standard solutions of NH4

+ in deionized 

water

RANGE: 8 to 100 μg/sample [3] 

ESTIMATED LOD: 2 μg/sample [3] 

PRECISION (SS̅

rr): 0.038 [2]

APPLICABILITY: The working range is 24 to 98 ppm (17 to 68 mg/m3

) for a 30-L sample [1]. This method is applicable to STEL 

measurements when sampled at 0.2 L/min.

INTERFERENCES: Ethanolamines (monoethanolamine, isopropanolamine, and propanolamine) have retention times similar to 

NH4

+. The use of the alternate (weak) eluent will aid in separating these peaks.

OTHER METHODS: This method combines the sampling procedure of methods S347 [4] and 6015 [5] with an ion 

chromatographic analytical procedure similar to Method 6701 [6] and OSHA Method ID-188 [3].
AMMONIA by IC: METHOD 6016, Issue 2, dated 3 March 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Water, deionized, filtered.

2. Sulfuric acid (H2SO4), 0.01 N:* Add 0.28 mL 

conc. H2SO4 to 500 mL deionized water in 1-L 

volumetric flask. Dilute to 1 L with deionized 

water.

3. Hydrochloric acid (HCl), 1 N:* 

Add 82.5 mL conc. HCl to 500 mL deionized

water in 1-L volumetric flask. Dilute to 1 L 

with deionized water.

4. 2,3-diaminopropionic acid 

monohydrochloride (DAP-HCl)

5. L-histidine monohydrochloride monohydrate 

(L-histidine-HCl)

6. Eluent (48 mM HCl/4 mM DAP-HCl/4 mM

L-histidine-HCl): Place 0.560 g DAP-HCl and

0.840 g L-histidine-HCl in a 1-L volumetric 

flask. Add 48 mL of 1 N HCl, dilute to volume 

with deionized water. Prepare monthly.

7. Alternate eluent (12 mM HCl/0.25 mM DAPHCl/0.25 mM L-histidine-HCl): Dilute

252 mL strong eluent and 36 mL 1 N HCl to 

4 L with deionized water. Prepare fresh for 

each use.

8. Tetramethylammonium hydroxide (TMAOH),

25% in water.

9. Regenerant solution: Dilute 57.4 mL of 25%

TMAOH to 4 L with deionized water.

10. Ammonia stock solution, 1000 μg/mL as NH3 

(1059 μg/mL as NH4

+

): Dissolve 3.1409 g 

ammonium chloride in deionized water.

Dilute to 1 L.

*See SPECIAL PRECAUTIONS. 

EQUIPMENT:

1. Sampler:

a. Prefilter: 37-mm mixed cellulose ester 

membrane filter, 0.8-μm pore size, 

stainless steel or porous plastic screen in 

two piece cassette filter holder.

b. Sulfuric acid-treated silica gel, glass tube,

unsealed and fire-polished, 6.0 cm long, 

6-mm OD, 4-mm ID, containing two 

sections of 20/40 mesh sulfuric acidtreated silica gel (200 mg front/100 mg 

back) separated and held in place with 

plugs of silylated glass wool, and capped 

with plastic caps. 

2. Personal sampling pump, 0.1 to 0.5 L/min, 

with flexible tubing.

3. Ion Chromatograph with conductivity 

detector, cation column and guard, and 

cation micromembrane suppressor (see 

Evaluation).

4. Syringes, 10-mL, polyethylene, Luer tip.

5. Centrifuge tubes, 15-mL, graduated, plastic 

with screw caps.

6. Volumetric flasks, 10-, 50-, 100-mL, and 1-L.

7. Syringe filters, 13-mm, 0.8-μm, membrane 

filter.

8. Micropipets, disposable tips.

9. Analytical balance (sensitivity to 0.01 mg). 

SPECIAL PRECAUTIONS: Concentrated acids are corrosive to skin. Handle acid in a fume hood. Wear 

protective gloves.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate between 0.1 and 0.5 L/min for a total sample size of 0.1 to 

96 L.

3. Cap the sampling tubes with plastic (not rubber) caps immediately after sampling.

4. Pack securely for shipment.
AMMONIA by IC: METHOD 6016, Issue 2, dated 3 March 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION:

5. Remove caps from sampling tubes. Transfer the front and back sections of sulfuric acid-treated silica 

gel to separate 15-mL graduated centrifuge tubes.

NOTE: Firm tapping of the tube may be necessary to effect complete transfer of the sulfuric 

acid-treated silica gel.

6. Add 10 mL of deionized water to each centrifuge tube. Cap and shake vigorously. Allow to stand 45 

minutes with occasional shaking. (Desorption is complete in 45 minutes.)

NOTE: Analyses should be completed within one day after the ammonia is desorbed.

7. Transfer samples to 10-mL syringes fitted with inline syringe filters for manual injection or transfer to 

autosampler vials.

CALIBRATION AND QUALITY CONTROL:

8. Calibrate daily with at least six working standards over the range of 1 to 110 μg NH3 per sample 

(about 0.11 to 12 μg/mL NH4

+

).

9. Add known aliquots of ammonia stock solution to 0.01 N H2SO4 in 10-mL volumetric flasks. 

NOTE: Prepare standards just before use.

10. Analyze working standards together with samples and blanks (steps 9 through 11).

11. Prepare calibration graph (peak height vs. μg NH3).

MEASUREMENT:

12. Set ion chromatograph to conditions given on page 6016-1, according to manufacturer’s instructions.

13. Inject 50-μL sample aliquot manually or with autosampler. For manual operation, inject 2 to 3 mL of 

sample from filter/syringe to ensure complete rinse of sample loop.

14. Measure peak height.

NOTE: If peak height exceeds linear calibration range, dilute with 0.01 N H2SO4, reanalyze and 

apply the appropriate dilution factor in calculations.

CALCULATIONS:

15. Determine the mass, μg, of ammonia found in the sample front (Wf) and back (Wb) sorbent sections, 

and in the average media blank front (Bf) and back (Bb) sorbent sections.

16. Calculate concentration, C, of NH3 in the air volume sampled, V (L):

CC = WWff + WWbb − BBff − BBbb

VV , mmmm/mm3

EVALUATION OF METHOD: 

This method combines the sampling procedure of NIOSH Methods S347 [4] and 6015 [5] with the ion 

chromatographic analytical procedure of NIOSH Method 6701 [6] and OSHA Method ID-188 [3]. This 

method used HPIC-CS3 cation separator, HPIC-CG3 cation guard and CMMS-1 cation micromembrane 

suppressor. This method will serve as an alternate analytical procedure to the automated 

spectrophotometric procedure of NIOSH Method 6015 [5]. Although the methods from which this method 

is derived are fully evaluated methods, the combination of the sulfuric acid-treated silica gel sampler and IC 

analysis has not received a full evaluation, as such. During the development of the passive monitor method 

for ammonia (6701), sulfuric acid-treated silica gel tubes were used as one of the reference methods [6]. 

The silica gel samples with IC analysis showed good agreement with the other reference methods, bubbler 

collection with colorimetric analysis using Nessler’s Reagent, and bubbler collection with IC analysis.

A storage stability study compared the sulfuric acid-treated silica gel tube and sulfuric acid-treated carbon 

beads used in OSHA Method ID-188 [3]. When stored at room temperature for five days and then 
AMMONIA by IC: METHOD 6016, Issue 2, dated 3 March 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

refrigerated for 21 days, silica gel samples had a mean recovery of 102 ± 3.8% (n = 8), while carbon beads 

had a mean recovery of 95 ± 1.6% (n = 8). The samples stored on carbon beads for 35 days showed 

significantly lower (although still acceptable) recovery compared to samples stored for 14 days: 103 ± 3.8% 

for silica gel (n = 12), and 108 ± 7.0% for carbon beads (n = 12) [2].

REFERENCES:

[1] NIOSH [1977]. Ammonia: Backup data report No. S347. Ten NIOSH analytical methods, set 6. SRI

International Contract No. 210-76-0123. (NTIS PB-288629) 

[2] Copenhafer RB, Dearden PA [1995]. User check of method 6015 for ammonia. Sequence 8321-J, K, L, 

M. Salt Lake City, UT: DataChem Labaratories. Unpublished.

[3] OSHA [1991]. Ammonia in workplace atmospheres - solid sorbent: Method ID-188. In: OSHA analytical 

methods manual, 2nd ed., Part 2, Inorganic substances. Salt Lake City, UT: U.S. Department of Labor, 

Occupational Safety and Health Administration.

[4] NIOSH [1979]. Ammonia: Method S347. In: Taylor DG, ed. NIOSH manual of analytical methods, 2nd 

ed., volume 5. Cincinnati, OH: U.S. Department of Health, Education and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 79-141.

[5] NIOSH [1994]. Ammonia: Method 6015. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical 

methods, 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS(NIOSH) 

Publication No. 94-113 [www.cdc.gov/niosh/nmam]. 

[6] NIOSH [1984]. Ammonia: Method 6701. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical 

methods, 3rd ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 84-100.

METHOD WRITTEN BY:

Mary Ellen Cassinelli, NIOSH

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CARBON MONOXIDE 6604

CO MW: 28.00 CAS: 630-08-0 RTECS: FG3500000

METHOD: 6604, Issue 2 EVALUATION: FULL Issue 1: 15 May 1996

Issue 2: 16 March 2016 

OSHA: 50 ppm

NIOSH: 35 ppm; C 200 ppm

PROPERTIES: colorless, odorless gas; BP -192 °C MP -207 °C;

vapor density (air=1) 0.97; flammable (explosive)

limits in air: 12.5 to 74.2%

SYNONYMS: monoxide; carbon oxide; carbonic oxide; flue gas

SAMPLING

SAMPLER: PORTABLE ELECTROCHEMICAL DIRECTREADING CO MONITOR

FLOW RATE: instrument dependent

VOL-MIN: 10 L

 -MAX: none

SHIPMENT: routine shipment of instrumentation

SAMPLE

STABILITY: at least 7 days @ 25 °C [1] 

(aluminized air bags)

BLANKS: fresh air or compressed CO-free air from

cylinder

ACCURACY 

RANGE 

STUDIED: 0 to 200 ppm

BIAS: - 1.7% [2]

OVERALL

PRECISION (SS�rrrr): 0.022 [2]

ACCURACY: ± 6.0%

MEASUREMENT

TECHNIQUE: ELECTROCHEMICAL SENSOR

ANALYTE: carbon monoxide (CO)

CALIBRATION 

-ZERO: CO-free air

-SPAN: standard cylinders of span gas in the

desired range

RANGE: 0 to 200 ppm

ESTIMATED LOD: 1 ppm

PRECISION (SS�rr): 0.035 @ 20 ppm

0.012 @ 50 ppm

0.008 @ 100 ppm [2]

APPLICABILITY: Portable, direct-reading carbon monoxide monitors are applicable to any work environment for personal or area 

monitoring. 

INTERFERENCES: Several gaseous pollutants (e.g., NO2, SO2) may cause an interference at levels over 5 ppm. If these or other 

pollutants are known or suspected to be present, use a monitor with a chemical interference scrubber over the sensor. Unknown

pollutants may require further experimentation to determine their effect on the sensor. As tested, SO2 (5 ppm), CO2 (5000 ppm), 

methylene chloride (500 ppm), diesel fuel (6 μL/L, about 0.3 ppm benzene), and gasoline vapor (1 μL/L, about 1 ppm benzene) had 

no impact on most monitor readings [2]. Some monitors are equipped with a chemical interference scrubber while others offer 

this as an option.

OTHER METHODS: OSHA methods ID 209 (CO by direct-reading monitor) [3] and ID 210 (CO by gas bag sampling) [4] are similar 

techniques. 
CARBON MONOXIDE: METHOD 6604, Issue 2, dated 16 March 2016 - Page 2 of 3 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. CO* calibration gas, 20 to 50 ppm, 

compressed gas cylinder, appropriate 

pressure regulator, and other items as 

recommended by manufacturer for field 

check of monitor response.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Electrochemical carbon monoxide

monitoring instrument designed and 

intended for industrial use (not a residential 

use alarm/monitor); fixed-location mountable 

for area alarm/monitoring, compact portable 

for person alarm/monitoring, or both; with 

integral concentration display.

2. Personal sampling pump, 0.250 L/min, with 

inlet and outlet, used for bag filling and 

sample analysis (e.g., when off-site analysis is 

needed).

3. Air bags, aluminized, 2-L, or other appropriate 

sizes (optional).

4. Replacement batteries or battery recharger, 

as appropriate for monitor. 

 

SPECIAL PRECAUTIONS: Carbon monoxide is a highly flammable, dangerous fire and explosive risk, and 

is toxic by inhalation. Shipments of compressed calibration gases must comply with 49 CFR 1992 

regulations.

SAMPLING:

1. Zero monitor with CO-free air at the same temperature and relative humidity as the work 

environment, if possible.

NOTE 1: Monitors are more sensitive to temperature variations than to humidity variations. Most 

monitors have temperature compensating circuitry. 

NOTE 2: If applicable, bag samples may be collected in aluminized bags (2-L or larger) and analyzed 

later by placing the calibration cap over the sensor and pumping the sample across the 

sensor at a nominal rate of 0.250 L/min with a personal sampling pump. 

2. For personal monitoring, locate the monitor as near the worker’s breathing zone as possible. 

3. For area monitoring, locate monitor in an area with good air circulation about 60 to 70 inches above 

the floor.

NOTE: Make sure the sensor is not obstructed in either application.

CALIBRATION AND QUALITY CONTROL:

4. Calibrate with a standard calibration mixture of CO in air (e.g., in gas bag samples of known 

concentrations) from a pressurized cylinder at the CO level recommended by the monitor 

manufacturer (normally, 20 to 50 ppm CO). The monitor should be calibrated at the temperature and 

relative humidity as near as possible to that of the work environment in which it will be used.

5. Check the calibration daily and recalibrate whenever the monitor reading varies from the span gas by 

5% or more, or as the manufacturer recommends.

MEASUREMENT: 

6. Read concentration directly (in ppm) from the monitor display.

Some monitors (data logger models) will maintain a continuous record of the data as it is accumulated 

and will calculate the average, TWA, peak, etc. concentrations. These data may be read from the 

display at any time. Some monitors will also store this information for downloading to a computer or 

printer at the end of the monitoring period. Other monitors only display the current reading, requiring 
CARBON MONOXIDE: METHOD 6604, Issue 2, dated 16 March 2016 - Page 3 of 3 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

the operator to manually record the data. All monitor models are equipped with alarms that will warn 

the user (audibly, visually or both) whenever the concentration of CO exceeds the preset level of the 

alarm. Many are equipped with two-level alarms [5].

EVALUATION OF METHOD: 

The performance of six direct-reading carbon monoxide monitors was evaluated over a period of 12 

months at CO concentrations up to 200 ppm and a range of ambient temperatures and relative 

humidity. Most of the tests were conducted at or near the PEL. For mean recovery studies, six different 

monitors were used and readings were taken approximately 1 h apart. Recovery at 20 ppm was 105% (n 

= 42); at 50 ppm, 99.6% (n = 36); and at 100 ppm, 99.9% (n = 30). Thus, the overall mean bias was 

calculated at -1.7%. The precision (S�r) at 20 ppm was 0.035 (35 readings from 5 monitors over a 7-h 

period). At 50 ppm, S�r was 0.012 (30 readings from 5 monitors over a 6-h period), and at 100 ppm, 

S�r was 0.008 (36 readings from 6 monitors over a 6-h period). Tests also were conducted to determine 

response time, zero and span drift, alarm decibel level, battery life, life of the sensors, as well as the 

effects of selected interferences (gases, vapors, and RF) and the effects of handling and transporting to 

remote sites. See [6] for preliminary work on carbon monoxide monitoring.

REFERENCES:

[1] Stanford Research Institute [1977]. Backup Data Report: Method S340. Menlo Park, CA: Stanford 

Research Institute. NIOSH contract no. 210-76-0123. 

[2] Woodfin WJ, Woebkenberg ML [1985]. Evaluation of portable direct-reading carbon monoxide 

monitors. Unpublished. 

[3] OSHA [1993]. Method ID 209, Carbon monoxide in workplace atmospheres (Direct-reading 

monitor); In: Sampling and analytical methods. Salt Lake City, UT: U.S. Department of Labor, 

Occupational Safety and Health Association.

[4] OSHA [1991]. Method ID 210, Carbon monoxide in workplace atmospheres (Gas bag sampling); In: 

Sampling and analytical methods. Salt Lake City, UT: U.S. Department of Labor, Occupational Safety 

and Health Association.

[5] Ashley K [1994]. Electroanalytical applications in occupational and environmental health. 

Electroanalysis 6:805-820.

[6] Woebkenberg ML [1983]. Monitoring methodology for gaseous hazards – Passive monitors and 

portable instruments. Scand J Work Environ Health 9:223-229. 

METHOD REVISED BY:

W. James Woodfin, NIOSH

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

LEAD by Flame AAS 7082

Pb MW: 207.19 (Pb) CAS: 7439-92-1 (Pb) RTECS: OF7525000 (Pb)

MW: 223.19 (PbO) CAS: 1317-36-8 (PbO)RTECS: OG1750000 (PbO)

METHOD: 7082, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984

Issue 3: 12 July 2017

OSHA: 0.050mg/m3

NIOSH: 0.050 mg/m3

OTHER OELs: [1-3]

PROPERTIES: soft metal; d 11.3 g/cm3; MP 327.5 °C 

valences +2, +4 in salts

SYNONYMS: elemental lead and lead compounds, except alkyl lead

SAMPLING

SAMPLER: FILTER (0.8-μm cellulose ester membrane) 

or INTERNAL CAPSULE, cellulose acetate 

dome with inlet opening attached to filter

FLOW RATE: 1 to 4 L/min

VOL-MIN: 200 L @ 0.05 mg/m3

 -MAX: 1500 L

SHIPMENT: routine

SAMPLE

STABILITY: stable at least 7 weeks [4] 

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 0.13 to 1.7 mg/m3 [8] 

BIAS: -3.1%

OVERALL

PRECISION (SS�rrrr): 0.07 [5,6] 

ACCURACY: ±17.6%

MEASUREMENT

TECHNIQUE: ATOMIC ABSORPTION 

SPECTROPHOTOMETER, FLAME

ANALYTE: lead

ASHING: conc. HNO3, 6 mL + 30% H2O2, 1 mL; 140°C

FINAL

SOLUTION: 10% HNO3, 10 mL

FLAME: air-acetylene, oxidizing

WAVELENGTH: 283.3 nm

BACKGROUND 

CORRECTION: D2 or H2 lamp, or Zeeman

CALIBRATION: Pb2+ in 10% HNO3

RANGE: 10 to 200 μg/sample [6,7]

ESTIMATED LOD: 2.6 μg/sample [8] 

PRECISION (SS�rr): 0.03 [5] 

APPLICABILITY: The working range is 0.05 to >1 mg/m3 for a 200-L air sample. The method is applicable to elemental lead, 

including Pb fume, and all other aerosols containing lead. This is an elemental analysis, not compound specific. Aliquots of the 

samples can be analyzed separately for additional elements. This method has been updated to include internal capsule samplers.

INTERFERENCES: Use D2 or H2 continuum or Zeeman background correction to control flame or molecular absorption. High 

concentrations of calcium, sulfate, carbonate, phosphate, iodide, fluoride, or acetate can be corrected for. 

OTHER METHODS: This method combines and replaces P&CAM 173 [7] and S341 [8] for lead. NIOSH Methods 7300 (ICP-AES), 

7701 (ASV) and 7105 (GFAAS) are alternative analytical methods. NIOSH Method 7505 is specific for lead sulfide. A consensus 

standard method, ASTM D6785, has been published [9].
LEAD by Flame AAS: METHOD 7082, Issue 3, dated 12 July 2017 - Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Nitric acid, conc., trace metal grade* 

2. Nitric acid, 10% (v/v). Add 100 mL conc. 

HNO3 to 500 mL water; dilute to 1 L 

3. Hydrogen peroxide, 30% H2O2 (w/w), 

reagent grade.* 

4. Calibration stock solution, 1000 μg/mL Pb. 

Commercial standard or dissolve 1.00 g Pb 

metal in minimum volume of (1+1) HCl and 

dilute to 1 L with 1% (v/v) HCl. Store in a 

polyethylene bottle. Stable > one year. 

5. Air, compressed, filtered. 

6. Acetylene 

7. Distilled or deionized water 

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Cellulose ester filter or cellulose 

acetate internal capsule attached to mixed 

cellulose ester membrane filter, 0.8-μm pore 

size, 37-mm diameter, in 2-piece cassette 

filter holder. 

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing. 

3. Atomic absorption spectrophotometer with 

an air-acetylene burner head and 

background correction.

4. Lead hollow cathode lamp or electrode 

dischargeless lamp.

5. Regulators, two-stage, for air and acetylene.

6. Beakers, Phillips, 125-mL, or Griffin, 50-mL 

with watch glass covers.**

7. Volumetric flasks, 10- and 100-mL.**

8. Assorted volumetric pipets, as needed.**

9. Hotplate, surface temperature 140 °C.

10. Bottles, polyethylene, 100-mL. 

** Clean all glassware with conc. nitric acid 

and rinse thoroughly with distilled or 

deionized water before use. 

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities and analysis. It 

is essential that suitable gloves, eye protection, laboratory coat, etc., be used when working with the 

chemicals. Concentrated nitric acid is an irritant and may burn skin. Perform all acid digestions in a fume 

hood. Hydrogen peroxide is a strong oxidizing agent, a strong irritant, and corrosive to the skin. Wear 

gloves and eye protection. 

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

NOTE: See NMAM guidance chapters for discussion on sampling.

2. Sample at an accurately known flow rate between 1 and 4 L/min (± 5%) for up to 8 h for a total sample 

size of 200 to 1500 L for TWA measurements. Do not exceed a filter loading of approximately 5 mg 

total dust. 

NOTE: Filter overloading can be assessed by periodic visual checks. See NMAM guidance chapters for 

additional discussion on filter capacity. 

SAMPLE PREPARATION:

3. Open the cassette filter holders and transfer the samples and blanks to clean beakers. 

NOTE: If internal capsules are not used, wipe the internal cassette surfaces with a polyvinyl alcohol 

wipe or cellulosic wipe wetted with deionized water, and add the wipe to the digestion vessel

(to transfer non-filter aerosol deposits into the digestion vessels) [10]. 
LEAD by Flame AAS: METHOD 7082, Issue 3, dated 12 July 2017 - Page 3 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NOTE: The following sample preparation gave quantitative recovery (see EVALUATION OF METHOD) 

[8]. Steps 4 through 9 of Method 7300 or other quantitative ashing techniques may be 

substituted, especially if several metals are to be determined on a single filter.

NOTE: The Appendix gives a microwave digestion procedure, which may be necessary for complete 

recovery of lead from some matrices, especially epoxy-based paint.

4. Add 3 mL conc. HNO3, and 1 mL 30% H2O2 and cover with a watch glass. Start reagent blanks at this 

step.

NOTE: If PbO2 is not present in the sample, the 30% H2O2 need not be added [6,8].

5. Heat on 140 °C hotplate until volume is reduced to about 0.5 mL.

6. Repeat two more times using 2 mL conc. HNO3 and 1 mL 30% H2O2 each time.

7. Heat on 140 °C hotplate until ca. 0.5 mL liquid remains.

8. When sample is dry, rinse the watch glass and walls of the beaker with 3 to 5 mL 10% HNO3. Allow the 

solution to evaporate to dryness.

9. Cool each beaker and dissolve the residues in 1 mL conc. HNO3. 

10. Transfer the solution quantitatively to a 10-mL volumetric flask and dilute to volume with distilled 

water.

NOTE: If the concentration (M) of any of the following is expected to exceed the lead concentration (M) 

by 10-fold or more, add 1 mL 1 M Na2EDTA to each flask before dilution to volume: CO3

2-, PO43-, 

I

-

, F-, CH3COO- [8]. If Ca2+ or SO4

2- are present in 10-fold or greater excess, make all standards and 

samples with 1% (w/w) La2+ [7].

CALIBRATION AND QUALITY CONTROL:

11. Prepare a series of working standards covering the range 0.25 to 20 μg/mL Pb (2.5 to 200 μg Pb per 

sample).

a. Add aliquots of calibration stock solution to 100-mL volumetric flasks. Dilute to volume with 10% 

HNO3. Store the working standards in polyethylene bottles and prepare fresh weekly.

b. Analyze the working standards together with the blanks and samples (steps 14 and 15).

c. Prepare a calibration graph of absorbance vs. solution concentration (μg/mL)

12. Aspirate a standard for every 10 samples to check for instrument drift.

13. Check recoveries with at least one spiked media blank per 10 samples. Use method of standard 

additions occasionally to check for interferences. 

MEASUREMENT:

14. Set spectrophotometer as specified by the manufacturer and to conditions on page 7082-1.

NOTE: An alternate wavelength is 217.0 nm [11]. Analyses at 217.0 nm have slightly greater sensitivity, 

but poorer signal-to-noise ratio compared to 283.3 nm. Also, non-atomic absorption is 

significantly greater at 217.0 nm, making the use of D2 or H2 continuum, or Zeeman background 

correction mandatory at that wavelength. 

15. Aspirate standards, samples, and blanks. Record absorbance readings.

NOTE: If the absorbance values for the samples are above the linear range of the standards, dilute with 

10% HNO3, reanalyze, and apply the appropriate dilution factor in the calculations. 

CALCULATIONS:

16. Using the measured absorbances, calculate the corresponding concentrations (μg/mL) of lead in the 

sample, Cs, and average media blank, Cb, from the calibration graph.

17. Using the solution volumes (mL) of the sample, Vs, and media blanks, Vb, calculate the concentration, 

C (mg/m3

), of lead in the air volume sampled, V (L): 

CC = CCssVVss − CCbbVVbb

VV ,

mmmm

mm3

NOTE: μg/mL ≈ mg/m3. 
LEAD by Flame AAS: METHOD 7082, Issue 3, dated 12 July 2017 - Page 4 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

EVALUATION OF METHOD:

The predecessor to NIOSH 7082, Method S341 [8], was issued on October 24, 1975, and validated over the 

range 0.13 to 0.4 mg/m3 for a 180-L air sample, using generated atmospheres of lead nitrate [5]. Recovery 

in the range 18 to 72 μg Pb per sample was 98%, and collection efficiency of 0.8-μm mixed cellulose ester 

filters (Millipore Type AA) was 100% for the aerosols at the detection limit of 0.013 mg/m3

. Subsequent 

studies on analytical recovery of 200 μg Pb per sample gave the following results [6,8]: 

Species Digestion method Analytical recovery, %

Pb metal HNO3 only 92.4 ± 4

Pb metal HNO3 + H2O2 103 ± 3

PbO HNO3 only 93 ± 4

PbS HNO3 only 93 ± 5

PbO2 HNO3 only 82 ± 3

PbO2 HNO3 + H2O2 100 ± 1

Pb in paint* HNO3 only 95 ± 6

Pb in paint* HNO3 + H2O2 95 ± 6

*Standard Reference Material #1579, U.S. National Institute of Standards and Technology

Additional collection efficiency studies were also done using Gelman GN-4 metrical cellulose acetate 

membrane filters for the collection of Pb fume, which had geometric mean diameter of 0.1 μm [5]. Mean 

collection efficiency for 24 sampling runs at flow rates between 0.15 and 4.0 L/min was >97 ± 2%. Overall 

precision, SrT, was 0.072 for lead nitrate aerosol [5,8] and 0.068 for Pb fume [6,8]. 

Evaluation information on internal capsule samplers may be found in NIOSH 7306 [12]. 

REFERENCES:

[1] ACGIH [2016]. TLVs® and BEIs® based on the documentation of the Threshold Limit Values for 

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American 

Conference of Governmental Industrial Hygienists. 

[2] Permanent Senate Commission for the Investigation of Health Hazards of Chemical Compounds in 

the Work Area [2016]. List of MAK and BAT values 2016; Maximum Concentrations and Biological 

Tolerance Values at the Workplace. Weinheim, Germany: Wiley-VCH Verlag GmbH & Co., http:// 

dx.doi.org/10.1002/9783527805983. 

[3] Institute for Occupational Safety and Health of the German Social Accident Insurance (IFA) [no date]. 

GESTIS international limit values, http://limitvalue.ifa.dguv.de/. Date accessed: July 12, 2017. 

[4] Marks GE, Knutsen EO [1975]. Complete testing of the NIOSH method for the determination of trace 

metals by atomic absorption spectrophotometry. Chicago, IL: IIT Research Institute, NIOSH contract 

no. CDC-99-74-48.

[5] Taylor DG, Kupel RE, Bryant JM. [1977]. Documentation of the NIOSH validation tests. Cincinnati, OH: 

U.S. Department of Health, Education, and Welfare, Center for Disease Control, National Institute for 

Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-185. Available from www.ntis.gov

as accession no. PB274248 

[6] Gutknecht WF, Ranade MH, Grohse PM, Damle A, O’Neal D [1981]. Heavy metal aerosols: Collection 

and dissolution efficiencies. Research Triangle Park, NC: Research Triangle Institute, Contract 210-79-

0058, Available from www.ntis.gov as accession no. PB83-106740.
LEAD by Flame AAS: METHOD 7082, Issue 3, dated 12 July 2017 - Page 5 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[7] NIOSH [1979]. Method P&CAM 173. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd ed. 

Cincinnati, OH: U.S. Department of Health, Education and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-157-A.

[8] NIOSH [1982]. Method S341. In: Taylor DG, ed. NIOSH manual of analytical methods. 2nd ed. 

Cincinnati, OH: U.S. Department of Health and Human Services, Center for Disease Control, National 

Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 82-100.

[9] ASTM International [2013]. ASTM D6785, Standard test method for determination of lead in 

workplace air using flame or graphite furnace atomic absorption spectrometry. West Conshohocken, 

PA: ASTM International.

[10] Ashley K, Harper M [2013]. Closed-face filter cassette (CFC) sampling - Guidance of procedures for 

inclusion of material adhering to internal sampler surfaces. J Occup Environ Hyg 10:D29-D33.

[11] Perkin-Elmer Corporation [1976]. Analytical methods for atomic absorption spectrophotometry. 

Norwalk, CT: Perkin-Elmer Corporation.

[12] NIOSH [2015]. Method 7306. In: Ashley K, O’Connor PF, eds. NIOSH manual of analytical methods. 5th

ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and 

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 14-

151. 

[13] DataChem Laboratories [no date]. In-house procedure for microwave sample digestion. Salt Lake 

City, UT: DataChem Laboratories. Unpublished.

[14] EPA [1986]. SW-846: Test methods for evaluating solid waste, Physical and chemical methods, 3rd ed. 

Washington, DC: U.S. Environmental Protection Agency.

[15] Kingston HM, Jassie LB, eds. [1988]. Safety guidelines for microwave systems in the analytical 

laboratory. In: Introduction to microwave acid decomposition: Theory and practice. Washington, DC: 

American Chemical Society.

[16] ASTM International [2011]. ASTM D1193, Standard specification for reagent water. West 

Conshohocken, PA: ASTM International.

[17] Kingston HM, Jassie LB, eds. [1988]. Introduction to microwave acid decomposition: Theory and 

practice. Washington, DC: American Chemical Society.

[18] Kingston HM [1988]. Quarterly Report, EPA IAG #DW1-393254-01-1.

[19] Binstock DA, Yeager WM, Grohse PM, Gaskill A [1989]. Validation of a method for determining 

elements in solid waste by microwave digestion, Draft technical report for RTI Project number 321U3579-24. Research Triangle Park, NC: Research Triangle Institute.

METHOD REVISED BY:

Mark Millson, R. DeLon Hull, PhD, and Ronnee N. Andrews, PhD, NIOSH. S341 originally validated under 

NIOSH contract CDC-94-74-45; additional studies under NIOSH contract 210-79-0058.

James B. Perkins, David L. Wheeler, and Keith Nicholson, Ph.D., DataChem Laboratories, Salt Lake City, UT, 

prepared the microwave digestion procedure described in the Appendix. 
LEAD by Flame AAS: METHOD 7082, Issue 3, dated 12 July 2017 - Page 6 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.

APPENDIX: MICROWAVE DIGESTION FOR LEAD IN PAINT CHIPS (AND OTHER MATRICES)

This procedure is an alternative to the procedure presented in the Sample Preparation section of this 

method. It provides a rapid, complete acid digestion prior to analysis by flame atomic absorption (FAA), 

heated graphite furnace atomic absorption (HGFAA), and inductively coupled plasma spectroscopy (ICP) 

[13]. 

Apparatus and Materials [14-19]

1. Microwave unit, to provide programmable power with a minimum of 574 W and programmable to 

within ± 10 W of the required power.

2. The microwave unit cavity shall be corrosion resistant as well as ventilated. All electronics are 

protected against corrosion for safe operation.

3. The system requires microwave transparent and reagent resistant vessels, such as perfluoroalkoxy 

alkane (PFA) digestion vessels (120-mL capacity), capable of withstanding pressures up to 7.5 ± 0.7 atm 

(760 ± 70 kPa). Vessels shall also be capable of controlled pressure relief at pressures exceeding 7.5 ± 

0.7 atm (760 ± 70 kPa). Other, equivalent types of vessels designed to operate at temperatures and 

pressures required and recommended by manufacturer can be used. 

4. A rotating turntable is employed to ensure homogeneous distribution of microwave radiation within 

the unit. The speed of the turntable should be a minimum of 3 rpm.

5. A safety concern relates to the use of sealed containers without pressure relief valves in the unit. 

Temperature is the important variable controlling the reaction. Pressure is needed to attain elevated 

temperatures but must be safely contained [15].

6. Polymeric volumetric ware in plastic (PTFE or polyethylene), 50- or 100-mL capacity.

7. Disposable polypropylene filter funnel.

8. Analytical balance, 300-g capacity, and minimum ± 0.001 g. 

Reagents

1. Nitric acid, concentrated, spectroscopy grade.

2. Reagent Water. Reagent water shall be interference free. All references to water in the method refer to 

reagent water that meets the ASTM Type 2 standard.

Procedure

1. Calibration of Microwave Equipment

a. Calibrate microwave equipment in accordance with manufacturer's instructions. If calibration 

instructions are not available, see EPA Method 3051 [14].

2. All digestion vessels and volumetric ware must be carefully acid washed and rinsed with reagent 

water. All digestion vessels should be cleaned by leaching with hot (1:1) nitric acid for a minimum of 

fifteen minutes, rinsed with reagent water, and dried in a clean environment

3. Sample Digestion

a. Tare the PFA digestion vessel.

b. Weigh out 0.1 g paint chip sample to the nearest 0.001 g into the tared PFA sample vessel. With 

large paint chip samples, measure out a 2 cm2 piece, weigh to the nearest 0.001 g, and 

quantitatively transfer it to the vessel.
LEAD by Flame AAS: METHOD 7082, Issue 3, dated 12 July 2017 - Page 7 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

c. Add 5.0 ± 0.1 mL concentrated nitric acid to the sample vessel in a fume hood. If a vigorous 

reaction occurs, allow the reaction to stop before capping the vessel. Cap the vessel and torque 

the cap to 16 N-m according to the manufacturer's directions. The sample vessel may be

connected to an overflow vessel using PFA connecting tubes. Place the vessels in the microwave

carrousel. Connect the overflow vessels to the center well of the unit.

d. Place the vessels evenly distributed in the turntable of the microwave unit. Any vessels containing 

5 mL of nitric acid for reagent blank purposes are counted as sample vessels. When fewer than the 

recommended number of samples are to be digested, i.e., three samples plus one blank, the 

remaining vessels should be filled with 5 mL of nitric acid to achieve the full complement of 

vessels. This provides an energy balance since the microwave power absorbed is proportional to 

the total mass in the cavity [17]. Irradiate each group of samples to achieve a temperature of 180 

°C. Temperature ramp times should be appropriate for the vessels used. A sample digestion 

program for 12 samples is presented in Table A1. Adjust power values depending upon the 

number of samples included in the microwave at one time.

TABLE A1. PROGRAM VARIABLES FOR PAINT CHIPS SAMPLE DIGESTION WITH NITRIC ACIDA

Stage 1 Stage 2 Stage 3

Power 90% 90% 0%

Pressure (kPa) 375 750 ---

Run time (min) 10 20 5

Time @ P (min) 5 15 ---

Temperature (°C) 180 180 None applied AFor 12 microwave vessels that contain 0.1 g of sample and 5 mL of liquid per vessel

e. At the end of the microwave program, allow the vessels to cool to a temperature below the boiling 

point of concentrated nitric acid (or that of the acid mixture used) before removing them from the 

microwave unit. If sample loss is detected (e.g., material in overflow collection vessel, liquid 

outside liner), determine the reason for the loss (e.g., loss of vessel seal integrity, use of a digestion 

time longer than 30 minutes, too large a sample, or improper heating conditions). Once the source 

of the loss has been corrected, prepare a new sample beginning at step 2. If insufficient material is 

available for reanalysis, dilute remaining digestate and note that some sample loss may have 

occurred. 

f. Uncap and vent each vessel in a fume hood. Transfer the sample to an acid-cleaned polyethylene 

bottle. Dilute to 25 mL using reagent water. If the digested sample contains particulates which 

may clog nebulizers or interfere with injection of the sample into the instrument, allow the 

sample to settle or filter it: 

Settling: Allow the sample to stand until the supernatant is clear (usually overnight is sufficient). If 

it does not filter the sample.

Filtering: Filter using disposable syringe filters, filter apparatus, etc. The filtering apparatus must be 

thoroughly precleaned and rinsed with dilute nitric acid. Filter the sample through 

quantitative filter paper into a second acid-cleaned container.

The digestate is now ready for analysis for elements of interest using the appropriate method.

4. Calculations: Report the concentrations based on the actual weight of the original sample.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 ELEMENTS by ICP (Microwave Digestion) 7302

AW: Table 1 CAS: Table 2 RTECS: Table 2

METHOD: 7302, Issue 1 EVALUATION: FULL Issue 1: 21 July 2014

OSHA: Table 2 

NIOSH: Table 2

Other OELs: [1,2]

PROPERTIES: Table 1

ELEMENTS: aluminum cadmium lead phosphorus strontium yttrium

antimony calcium lithium platinum tellurium zinc

arsenic chromium magnesium potassium thallium zirconium

barium cobalt manganese selenium tin

beryllium copper molybdenum silver titanium

boron iron nickel sodium vanadium

SAMPLING

SAMPLER: FILTER (mixed cellulose ester membrane (MCE), 

37-mm diameter, 0.8-μm pore size)

FLOW RATE: 1 to 4 L/min

VOL-MIN:

-MAX: Table 1

SHIPMENT: Routine

SAMPLE

STABILITY: Stable

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: See Table 4

ACCURACY: See Table 4

BIAS: See Table 4

OVERALL

PRECISION ( ): See Table 4 

MEASUREMENT

TECHNIQUE: INDUCTIVELY COUPLED ARGON PLASMA, 

ATOMIC EMISSION SPECTROSCOPY 

(ICP-AES)

ANALYTE: Elements listed above

REAGENTS: 10.0 mL of 1:1 nitric (HNO3) and ASTM Type II 

water

FINAL

SOLUTION: 20% HNO3, 25 mL

WAVELENGTH: Depends upon element (see Table 3)

BACKGROUND

CORRECTION: Spectral wavelength shift

CALIBRATION: Elements in 20% HNO3

RANGE: See Table 4

ESTIMATED LOD: Table 3

PRECISION ( ) : Table 

APPLICABILITY: This method is for the analysis of metal and nonmetal dust collected on MCE filters in the workplace. The 

working range varies from element to element. The method entails simultaneous elemental analysis using a microwave digestion approach to simplify and expedite the analysis.

INTERFERENCES: Spectral interferences are the primary interferences encountered in ICP-AES analysis. These are minimized 

by judicious wavelength selection, inter-element correction factors and background correction [3].

OTHER METHODS: This method complements NIOSH hotplate digestion methods 7300 and 7301 for trace elements. Flame 

atomic absorption spectroscopy (e.g., Methods 7013 through 7082) is an alternative analytical technique for many of these 

elements [4]. Graphite furnace AAS (e.g., 7102 for Be, 7105 for Pb) is usually more sensitive [4]. NMAM 7301 and 7303 contain 

alternative extraction procedures.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 2 of 9

REAGENTS:

1. Nitric acid, conc., trace metal grade*

2. Calibration stock solutions, 1000 μg/mL 

and 10,000 μg/mL commercially available, 

or prepared per instrument manufacturer’s 

recommendation (see step10)

3. Digestion acid*. 1:1 water, ASTM type II, and 

nitric acid*, trace metal grade

4. Argon, liquid

5. De-ionized Water, ASTM Type ll [5]

6. Dilution acid*, 20% nitric acid in ASTM Type II 

water

* See Special Precautions

EQUIPMENT:

1. Sampler: mixed cellulose ester membrane (MCE) 

filter, 0.8-μm pore size, 37-mm diameter; in 

2-piece cassette filter holder

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing

3. Inductively coupled plasma-atomic emission 

spectrometer, equipped as specified by the 

manufacturer for analysis of elements of interest

4. Regulator, two-stage for argon

5. Microwave, programmable power, active 

temperature control, minimum of 574 W, 

corrosion resistant ventilated oven and 

turntable

6. Microwave digestion vessels, high pressure, 

closed PTFE, 100-mL capacity

7. Volumetric flasks, 25 mL**

8. Assorted volumetric pipettes as needed**

** Acid wash all glassware and vessels before using.

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals. 

All work should be performed with adequate ventilation to personnel and equipment. Because this 

method involves the use of capped digestion containers, avoid the use of other acids such as perchloric 

acid in combination with nitric acid that could cause a violent reaction [6,7]. In the preparation of the 

digestion and dilution acids, it is imperative that acid be added to water in order to avoid a violent 

exothermic reaction.

SAMPLING

1. Calibrate each personal sampling pump with a representative sampler connected to the pump (in 

line).

2. Sample at an accurately known flow rate between 1 and 4 L/min. For estimated sampling volumes see 

Table 1. For TWA measurements see Table 2. Do not exceed a filter loading of approximately 2 mg total 

dust.

NOTE: Filter overloading can be assessed by periodic visual checks. See NMAM guidance chapters for 

discussion on sampling.

SAMPLE PREPARATION

NOTE: If total weights are desired, weighing should be done at this step. Follow NIOSH method 0500 

for gravimetric analysis [11].

3. Open the cassette filter holders and transfer the samples, blanks, and Quality Control (QC) filters to 

clean PTFE digestion vessels. Wipe the internal cassette surfaces with a 37-mm MCE filter, polyvinyl 

alcohol wipe or cellulosic wipe wetted with deionized water, and add to the digestion vessel to transfer 

non-filter aerosol deposits into the digestion vessels.

4. Add digestion acid up to 10 mL, and cap the vessels.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 3 of 9

NOTE: In order to avoid a violent exothermic reaction, do not add water to concentrated nitric acid. 

Acid should be added after the water has been placed in the vessel.

5. Place digestion vessels in microwave, and run the preprogrammed digestion procedure for 12- vessel

digestion: 1200 W power, ramp to 150 °C over 20 min, hold for 10 min at 150 °C followed by at least a 5

min cool down (power will be adjusted lower for fewer vessels).

6. Allow the samples to cool to room temperature.

7. Remove vessel lids and rinse contents into 25-ml volumetric flasks with ASTM Type ll water.

8. Dilute to the mark with ASTM Type ll water and mix.

9. Submit extracted and diluted samples for analysis. 

CALIBRATION AND QUALITY CONTROL

10. Calibrate the spectrometer according to the manufacturer recommendations.

NOTE: Typically an acid blank and multi-element working standards are used. The following multi-

 element combinations are chemically compatible in 20% HNO3

.

a. Al, As, Ba, Be, Ca, Co, Cr, Cu, Fe, Li, Mg, Mn, Mo, Na, Ni, Pb, Se, Sr, Ti, V, Y, Zn, Zr;

b. B, K, P, Sn, Te, Tl;

c. Ag, Cd, Sb;

d. Pt.

11. Analyze all applicable standards at least once every twenty (20) analyses (minimum frequency 5%).

12. Check recoveries with at least one media blank and two spiked media blanks per twenty samples. Use

a spike level that is within the range of 10 to 20 times the LOQ.

NOTE: Whenever possible, QA/QC samples should be prepared from certified reference materials in

a matrix similar to the bulk material sampled.  Liquid spiked filters are only surrogates for real 

world samples and QC data based upon certified samples are preferred.

MEASUREMENT

13. Set ICP-AES spectrometer to conditions specified by manufacturer.

14. Analyze standards and samples at applicable wavelengths for each element (target analytes are in

Table 3).

NOTE: If the values for the samples are above the linear range of the instrument, dilute the

 solutions with dilution acid, reanalyze, and apply the appropriate dilution factor in

 calculations.

CALCULATIONS

15. Obtain the solution concentrations for the sample, Cs

 (μg/mL), and the average media blank, Cb (μg/

mL), from the instrument.

16. Using the solution volume of sample, Vs

 (mL), and media blank, Vb (mL), calculate the concentration for

the sample, C (mg/m3

), of each element in the air volume sampled, V (L), as follows:

  , mg/m

NOTE: μg/Liter is essentially equal to mg/m3.

EVALUATION OF METHOD

Method 7302 was evaluated using multi-element filter spikes at six spiking levels, based on the estimated 

LOQ for each element [8]. Using microwave digestion is less time consuming and more convenient than 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 4 of 9

using the traditional mixed acid hot plate approach. The elimination of perchloric acid in the sample 

digestion procedure helps to improve the safety of the method. [7] 

Summary data are presented in Table 3 for levels 3X LOQ (lower level in Table 3) and 300X LOQ (higher 

level in Table 3) and for the ranges of loadings given in Table 4. Samples were subjected to microwave 

digestion using a CEM MDS-2100 device according to the conditions specified in the “sample preparation” 

section above (see Note of step #5). The values in Tables 3 and 4 were determined using several different 

ICP-AES instruments which were operated according to manufacturer’s instructions.The precision and 

recovery data, instrumental detection limits, sensitivity, and analytical wavelengths are listed in Table 3 

and Table 4. All of the precision data were evaluated for homogeneity for all six concentration levels tested 

using the Bartlett’s test and the results are listed in the method backup data report [8] and summarized 

in Tables 3 and 4. A statistical analysis found that the data were poolable and all elements had calculated 

method precision accuracies of less than 25%. This overall precision (ŜrT) and accuracy as given in Table 4 

is an upper limit predictor of precision. Accuracy data (Table 4) demonstrate the utility of the method for 

all of the elements listed.

A discussion of metals and metalloid analysis by ICP-AES is presented in an international voluntary 

consensus standard [3] and the microwave digestion procedure has been evaluated against other 

digestion procedures through an interlaboratory trial [10].

REFERENCES

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American

Conference of Governmental Industrial Hygienists. [www.acgih.org]. [Data accessed April 2014.]

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database on

hazardous substances (German Social Accident Insurance). Sankt Augustin, FRG: [http://www.dguv.

de/ifa/GESTIS]. [Data accessed April 2014.]

[3] ASTM [2010]. ASTM D7035-10, Standard test method for the determination of metals and metalloids

in airborne particulate matter by inductively coupled plasma atomic emission spectrometry. West

Conshohocken, PA: ASTM International [www.astm.org].

[4] NIOSH [1994] NIOSH Manual of analytical methods (NMAM). 4th ed. Schlecht PC, O’Connor PF, eds.

Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No.

94-113 [www.cdc.gov/niosh/nmam/].

[5] ASTM International [2011]. ASTM D1193-77, Standard specification for reagent water. West

Conshohocken, PA: ASTM International [www.astm.org].

[6] NIOSH [2005]. NIOSH Pocket guide to chemical hazards. Barsan ME, ed. Cincinnati, OH: US

Department of Health and Human Services, Centers for Disease Control and Prevention, National

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149.

[7] Furr AK, ed. [1995]. CRC Handbook of laboratory safety, 4th ed. Boca Raton, FL: CRC Press.

[8] Stone LC, Perkins JB, Rogers DM [2003]. Elements by ICP using microwave digestion, NMAM 7302,

Issue 1, Backup data report. Cincinnati OH: Datachem Laboratory developed under contract CDC-200-

2001-08000. Unpublished.

[9] Lide DR, ed. [1994] Handbook of chemistry and physics. 74th ed. Baca Raton, FL.: CRC Press.

[10] Butler OT, Howe AM [1999]. Development of an international standard for the determination of

metals and metalloids in workplace air using ICP-AES: Evaluation of sample dissolution procedures

through an interlaboratory trial. J. Environ. Monit. 1: 23-32.

[11] Code of Federal Regulations, 29 CFR Part 1910.1000 (Table Z-1) [https://www.osha.gov/pls/oshaweb/

owadisp.show_document?p_table=STANDARDS&p_id=9992]. Website accessed on December 13,

2013.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 5 of 9

METHOD WRITTEN BY:

David M. Rogers, Lee C. Stone, James B. Perkins, DataChem Laboratories, Salt Lake City, Utah.; Yvonne 

Gagnon, Ronnee Andrews, Ph.D., NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. 

In addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 6 of 9

TABLE 1. PROPERTIES AND SAMPLING VOLUMES

Element (Symbol)

Properties [9]

Atomic Weight MP, °C

Air Volume, L @

OSHA PEL [11] 

MIN MAX

Aluminum (Al) 26.98 660 5 100

Antimony (Sb) 121.8 630 10(2) 2000(2)

Arsenic (As) 74.92 817 5 2000

Barium (Ba) 137.3 727 5(2) 200(2)

Beryllium (Be) 9.01 1287 1250 2000

Boron (B)(1) 10.81 2300 5 2000

Cadmium 112.41 321 12 2000

Calcium (Ca)(1) 40.08 842 5 200

Chromium (Cr) 52.00 1907 5 1000

Cobalt (Co) 58.93 1495 25 2000

Copper (Cu) 63.54 1083 5 1000

Iron (Fe) 55.85 1535 5 100

Lead (Pb) 207.19 328 50 2000

Lithium (Li)(1) 6.94 181 100 2000

Magnesium (Mg) 24.31 651 5 67

Manganese (Mn) 54.94 1245 5 200

Molybdenum (Mo) 95.94 2623 5 67

Nickel (Ni) 58.71 1455 5 1000

Phosphorus (P) 30.97 44 25 2000

Platinum (Pt) 195.09 1769 1250 2000

Potassium (K)(1) 39.10 64 5 2000

Selenium (Se) 78.96 221 13 2000

Silver (Ag) 107.87 961 250 2000

Sodium (Na)(1) 22.99 98 13 2000

Strontium (Sr)(1) 87.62 777 5 2000

Tellurium (Te) 127.60 450 25 2000

Tin (Sn) 118.69 232 20(2) 2000(2)

Thallium (Tl) 204.37 304 25 2000

Titanium (Ti) 47.87 1668 5 100

Vanadium (V) 50.94 1910 5 2000

Yttrium (Y) 88.91 1522 5 1000

Zinc (Zn) 65.37 419 5 200

Zirconium (Zr) 91.22 1855 5 200

(1) No PEL, REL, or STEL data found [1,6,11].

(2) Air volumes estimated from TWAs and LOQs (see Tables 2, 3) [1].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 7 of 9

TABLE 2. EXPOSURE LIMITS, CAS #, RTECS [1,6,11]

Element 

(Symbol) CAS # RTECS

Exposure Limits in mg/m

(C = ceiling limit)

3

OSHA NIOSH

Aluminum (Al) 7429-90-5 BD0330000 15 (total dust)

5 (respirable)

10 (total dust), 2 (soluble)

5 (respirable, fume)

Arsenic (As) 7440-38-2 CG0525000 0.010 (inorganic) C 0.002(1), 0.5

Barium (Ba) 7440-39-3 0.5 (Soluble compounds, 

as Ba)

0.5 (Soluble compounds, 

as Ba)

Beryllium (Be) 7440-41-7 DS1750000 0.002, C 0.005 C 0.0005(1)

Boron (B) 7440-42-8 ED7350000 15 as oxide 10 as oxide

Cadmium (Cd) 7440-43-9 EU9800000 0.005 lowest feasible conc.(1)

Calcium (Ca) 7440-70-2 EV8040000 No OELs No OELs

Chromium (Cr) 7440-47-3 GB4200000 0.5 (II & III), 0.005 (VI)

1 (metals, insoluble salts)

0.5

Cobalt (Co) 7440-48-4 GF8750000 0.1 (dust, fume) 0.05 (dust, fume)

Copper (Cu) 7440-50-8 GL5325000 1(dust, mists)

0.1 (fume)

1 (dust, mists)

0.1 (fume) 

Iron (Fe) 7439-89-6 NO4565500 10 (fume) as oxide 5 (dust, fume) as oxide

Lead (Pb) 7439-92-1 OF7525000 0.05 0.05

Lithium (Li) 7439-93-2 OJ5540000

Magnesium (Mg) 7439-95-4 OM2100000 15 (dust) as oxide --

Manganese (Mn) 7439-96-5 OO9275000 C 5 1; STEL 3

Molybdenum (Mo) 7439-98-7 QA4680000 5 (soluble)

15 (total insoluble)

Nickel (Ni) 7440-02-0 QR5950000 1 0.015, Ca

Phosphorus (P) 7723-14-0 TH3500000 0.1 0.1

Platinum (Pt) 7440-06-4 TP2160000 0.002 (soluble) 1 (metal) 0.002 (soluble)

Potassium (K) 7740-09-7 TS6460000

Antimony (Sb) 7440-36-0 CC4025000 0.5 0.5

Selenium (Se) 7782-49-2 VS7700000 0.2 0.2

Silver (Ag) 7440-22-4 VW3500000 0.01 (dust, fume, metal) 0.01 (dust, fume, metal)

Sodium (Na) 7440-23-5 VY0686000

Strontium (Sr) 7440-24-6 WK7700000

Tellurium (Te) 13494-80-9 WY2625000 0.1 0.1

Tin (Sn) 7440-31-5 XP7320000 2 2

Titanium (Ti) 7440-32-6 XR1700000 15 (as TiO2) lowest feasible(1)

Thallium (Tl) 7440-28-0 XG3425000 0.1 (soluble) skin 0.1 (soluble) skin

Vanadium (V) 7440-62-2 YW1355000 C 0.5 (respirable) as V2

O5

C 0.1 (fume) as V2

O5

C 0.05 as V2

O5

Yttrium (Y) 7440-65-5 ZG2980000 1 1

Zinc (Zn) 7440-66-6 ZG8600000 5 (ZnO fume)

15 (ZnO dust)

5 (ZnO respirable)

5; STEL 10 (ZnO fume)

5; C 15 (ZnO dust)

Zirconium (Zr) 7440-67-7 ZH7070000 5 5, STEL 10

(1) Carcinogen
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 8 of 9

TABLE 3. MEASUREMENT WAVELENGTHS AND RECOVERY DATA [8]

 Lower Level Higher Level

Element(1) Wavelength

(nm)[6]

LOD

(μg/sample)

μg/sample % Recovery 

N=6)

% RSD μg/sample % Recovery

(N=6)

% RSD

Ag 328.1 0.1 1.50 95.5 1.01 150 99.0 0.497

Al 308.2 1 7.50 92.7 0.981 750 98.7 0.462

As 193.8 1 7.50 101 2.22 750 107 0.340

B 249.7 0.5 3.75 112 2.96 375 99.5 0.454

Ba 493.4 0.06 0.752 104 3.09 75.2 101 0.438

Be 313.0 0.009 0.076 95.8 2.36 7.60 103 0.714

Ca 315.9 2 22.5 107 2.87 2250 99.0 0.620

Cd 228.8 0.1 1.50 98.8 3.46 150 104 0.701

Co 228.6 0.3 3.75 99.7 1.72 375 104 0.566

Cr(2) 267.7 0.4 3.75 103 7.87 375 103 3.36

Cu 324.8 0.07 0.752 98.8 3.47 75.2 94.2 0.371

Fe 259.9 2 15.0 112 2.43 1500 101 0.263

K 766.5 2 15.0 98.3 5.70 1500 103 0.472

Li 670.8 0.03 0.752 92.4 2.98 75.2 98.8 0.749

Mg 279.1 0.5 7.50 89.3 3.52 750 95.1 0.309

Mn 257.6 0.02 0.752 86.2 2.38 75.2 98.2 0.389

Mo 202.0 0.2 2.25 96.8 5.41 225 103 0.373

Na 589.0 4 37.5 100 0.823 3750 110 0.457

Ni 231.6 0.2 2.25 98.3 5.21 225 97.7 0.592

P 214.9 2 15.0 100 5.67 1500 104 0.315

Pb 220.4 0.6 7.50 98.9 3.94 750 104 0.570

Pt(2) 265.9 8 75.0 98.3 0.282 10000 95.7 1.49

Sb 206.8 0.4 7.50 94.4 3.21 750 103 0.255

Se 196.1 3 37.5 104 3.21 3750 106 0.270

Sn(2) 189.9 0.8 37.5 105 5.04 3750 90.3 3.23

Sr 421.6 0.02 3.75 92.6 2.36 375 97.5 0.553

Te(2) 214.3 2 15.0 90.1 21.8 1500 103 0.614

Ti 337.3 0.2 1.50 101 1.70 150 98.8 0.575

Tl 190.9 0.9 7.5 103 4.14 750 99.3 0.352

V 292.4 0.1 0.752 93.7 4.74 75.2 103 0.341

Y(2) 371.0 0.02 0.376 107 4.44 37.6 102 3.33

Zn(2) 213.9 0.1 1.50 106 13.1 150 97.4 3.42

Zr 339.2 0.06 0.750 93.1 5.35 75.0 95.4 0.971

(1) Values reported were obtained with a Fisons ARL Accuris ICP-AES; performance may vary with instrument and should 

 be independently verified.

(2) Values reported were obtained with a Perkin Elmer Optima 3000 DV ICP-AES. Sample concentration was based on Fisons

 ICP LOD data.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP (Microwave Digestion): METHOD 7302, Issue 1, dated 21 July 2014 - Page 9 of 9

TABLE 4. PRECISION AND ACCURACY DATA BY ELEMENT [8]

Element

(μg/sample)

Range

(μg/sample) Bias  (%) Accurac

Ag 0.5 to 150 -0.0175 0.668 2.85

Al 2.5 to 750 0.0505 1.455 7.41

As 2.5 to 750 -0.2249 0.554 23.40

Ba 0.25 to 75.2 -0.0330 0.920 4.82

Be 0.025 to 7.60 0.0297 0.863 4.39

Ca 7.43 to 2250 -0.0081 0.836 2.18

Cd 0.50 to 150 -0.0082 0.729 2.02

Co 1.24 to 375 -0.0161 0.574 2.56

Cr 1.24 to 375 -0.0204 0.655 3.12

Cu 0.248 to 75.2 0.0160 0.984 3.21

Fe 5.00 to 1500 -0.0039 1.637 3.30

K 5.00 to 1500 0.1487 1.665 17.61

La 12.6 to 50.1 -0.0136 0.920 2.87

Li 0.25 to 75.2 0.2241 1.209 24.40

Mg 2.5 to 750 0.0180 0.844 3.19

Mn 0.25 to 75.2 -0.0348 0.865 4.91

Mo 0.75 to 225 0.0140 1.469 3.82

Ni 0.75 to 225 -0.0063 0.672 1.73

P 5.0 to 1500 0.0669 1.212 8.69

Pb 2.5 to 750 -0.0246 0.544 3.36

Sb 2.5 to 750 0.0172 0.722 2.91

Se 12.4 to 3750 0.0538 0.758 6.63

Sn 12.4 to 3750 0.0561 0.936 7.15

Sr 1.24 to 375 -0.0074 0.710 1.90

Te 5.0 to 1500 0.0161 0.892 3.08

Ti 0.5 to 150 0.0212 1.043 3.84

Tl 2.5 to 750 -0.0293 0.602 3.92

V 0.25 to 75.2 0.0175 1.223 3.76

Y 0.12 to 37.6 -0.0179 1.115 3.62

Zn 0.5 to 150 0.0075 1.343 3.02

Zr 0.25 to 75.0 0.0314 0.980 4.76
ELEMENTS by ICP Microwave Digestion 7304

MW: Table 1 CAS: Table 2 RTECS: Table 2

METHOD: 7304, Issue 1 EVALUATION: FULL Issue 1: 25 May 2014

OSHA: Table 2 

NIOSH: Table 2

Other OELs: [1,2]

PROPERTIES: Table 1

ELEMENTS: aluminum cadmium iron molybdenum selenium titanium

arsenic calcium lead nickel sodium vanadium

barium chromium lithium phosphorus strontium yttrium

beryllium cobalt magnesium platinum tellurium zinc

boron copper manganese potassium thallium zirconium

SAMPLING

SAMPLER: FILTER, (polyvinyl chloride (PVC), 37-mm 

diameter, 5.0 μm pore size)

FLOW RATE: 1 to 4 L/min

VOL-MIN:

-MAX: Table 1

SHIPMENT: Routine

SAMPLE

STABILITY: Stable

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: See Table 4

ACCURACY: See Table 4

BIAS: See Table 4

OVERALL PRECISION ( S

ˆ

rT ): See Table 4

MEASUREMENT

TECHNIQUE: INDUCTIVELY COUPLED ARGON 

PLASMA, ATOMIC EMISSION 

SPECTROSCOPY (ICP-AES)

ANALYTE: Elements listed above

REAGENTS: 12 mL of 5:1 concentrated nitric acid and 

ASTM Type ll water

FINAL SOLUTION: 20% HNO3, 50 mL

WAVELENGTH: Depends upon element; Table 3

BACKGROUND

CORRECTION: Spectral wavelength shift

CALIBRATION: Elements in 20% HNO3

RANGE: See Table 4

ESTIMATED LOD: Table 3

PRECISION ( Sr ): Table 3

APPLICABILITY: The working range of this method varies from element to element. This method is for the analysis of 

metal and nonmetal dust collected on PVC filters that are also used for gravimetric analysis. This is a simultaneous elemental analysis using a microwave digestion approach to simplify and expedite the analysis. Some elements such as antimony, 

silver, and tin do not form stable solutions in nitric acid when chloride from the PVC filters is present. In such cases a mixed 

cellulose ester (MCE) filter is necessary (See NMAM 7302). A different acid medium also helps but this technique is not 

described in this method.

INTERFERENCES: Spectral interferences are the primary interferences encountered in ICP-AES analysis. These are minimized by judicious wavelength selection, inter-element correction factors and background correction. [3,4,5,6]

OTHER METHODS: This method complements NIOSH hotplate digestion methods 7300 and 7301 for trace elements. 

Flame atomic absorption spectroscopy (e.g., Methods 70XX) is an alternative analytical technique for many of these elements. [7] Graphite furnace AAS (e.g., 7102 for Be, 7105 for Pb) is usually more sensitive. [7] NMAM 7301 and 7303 contain 

alternative extraction procedures.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 2 of 11

REAGENTS

1. Nitric acid, conc., trace metal grade*

2. Calibration stock solutions, 1000 ug/mL

and 10,000 ug/mL commercially available, 

or prepared per instrument manufacturer

recommendation (see step 10)

3. Argon, liquid

4. De-ionized W ater, ASTM Type ll [8]

5. Dilution acid: 20% nitric acid in ASTM Type ll 

water*

* See SPECIAL PRECAUTIONS

EQUIPMENT

1. Sampler: Polyvinyl chloride filter, 5.0-μm pore

size, 37-mm diameter; in 2-piece cassette filter

holder

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing

3. Inductively coupled plasma-atomic emission

spectrometer, equipped as specified by the 

manufacturer for analysis of elements of 

interest

4. Regulator, two-stage for argon

5. Microwave, programmable power, active 

temperature control, minimum of 574 W,

corrosion resistant ventilated oven and 

turntable

6. Microwave digestion vessels, high pressure,

closed PTFE, 100-mL capacity

7. Volumetric flasks, 50 mL**

8. Assorted volumetric pipettes as needed**

** Acid wash all glassware and vessels before using.

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals. All 

work should be performed with adequate ventilation to personnel and equipment. Because this 

method involves the use of capped digestion containers, avoid the use of other acids such as perchloric 

acid in combination with nitric acid that could cause a violent reaction [1,9]. In the preparation of 

the digestion and dilution acid, it is imperative that acid be added to water in order to avoid a violent 

exothermic reaction.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler connected to the pump 

(in line.)

2. Sample at an accurately known flow rate between 1 and 4 L/min. For estimated sampling volumes

see Table 1. For TWA measurements see Table 2. Do not exceed a filter loading of approximately

2 mg total dust.

NOTE: Filter overloading can be assessed by periodic visual checks. See NMAM guidance chapters

for additional discussion on filter capacity.

SAMPLE PREPARATION:

NOTE: If total weights are desired, it should be done at this step. Follow NIOSH method 0500 for 

gravimetric analysis [12].

3. Open the cassette filter holders and transfer the samples, blanks, and Quality Control (QC) filters to

clean PTFE digestion digestion vessels. Wipe the internal cassette surfaces with a 37 mm PVC filter 

wetted with deionized water and add to the digestion vessel to transfer non-filter aerosol deposits 

into the digestion vessels.

4. Add 2 mL of ASTM Type ll water followed by adding (slowly) 10 mL concentrated nitric acid, then

cap each vessel.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 3 of 11

NOTE: In order to avoid a violent exothermic reaction, do not add water to concentrated nitric acid. 

Acid should be added after the water has been placed in the vessel. 

5. Place digestion vessels in microwave and run preprogrammed PVC digestion procedure. Example

microwave conditions for 12-vessel digestion: 1200 W power, ramp to 215 °C over 20 min, hold for 

10 min at 215 °C followed by at least a 5 min cool down (power will be adjusted lower for fewer 

vessels). 

6. Allow the samples to cool to room temperature.

7. Remove vessel lids and rinse contents into 50-mL volumetric flasks with ASTM Type ll water.

8. Dilute to the mark with ASTM Type ll water and mix.

9. Submit samples for analysis.

NOTE: A residual solid may be present after digestion. Filter/centrifuge the samples before analysis,

as appropriate.

CALIBRATION AND QUALITY CONTROL:

10. Calibrate the spectrometer according to the manufacturers’ recommendations.

NOTE: Typically an acid blank and a single or multi-element working standard are used. The

following multi-element combinations are chemically compatible in 20% HNO3

.

a. Al, As, Ba, Be, Ca, Co, Cr, Cu, Fe, Li, Mg, Mn, Mo, Na, Ni, Pb, Se, Sr, Ti, V, Y, Zn, Zr;

b. B, K, P, Te, Tl;

c. Cd;

d. Pt.

11. Analyze all applicable standards at least once every twenty (20) analyses (minimum frequency 5%).

12. Check recoveries with at least one media blank and two spiked media blanks per twenty samples.

Use a spike level that is within the range of 10 to 20 times the Limit of Quantitation (LOQ.)

NOTE: Whenever possible, QA/QC samples should be prepared from certified reference materials

in a matrix similar to the bulk material sampled.  Liquid spiked filters are only surrogates for 

real world samples and QC data based upon certified samples would be ideal.

MEASUREMENT:

13. Set the ICP-AES spectrometer to conditions specified by manufacturer.

14. Analyze standards and samples at applicable wavelengths for each element (target analytes are in

Table 3).

NOTE: If the values for the samples are above the linear range of the instrument, dilute the

solutions with dilution acid, reanalyze, and apply the appropriate dilution factor in the 

calculations.

CALCULATIONS:

15. Obtain the solution concentrations for the sample, Cs

 (μg/mL), and the average media blank, Cb

(μg/mL), from the instrument.

16. Using the solution volume of sample, Vs

 (mL), and media blank, Vb (mL), calculate the concentration

for the sample, C (mg/m3

), of each element in the air volume sampled, V (L), as follows:

NOTE: μg/Liter is approximately equal to mg/m3.
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 4 of 11

EVALUATION OF METHOD:

This method is less time consuming and more convenient than using the acid hotplate approach. The 

elimination of perchloric acid in the sample digestion procedure helps to improve the safety of the

method. [9] Use of the PVC filters allows for the acquisition of total mass per filter in addition to total 

metals concentration.

The evaluation of this method, 7304, for PVC filters was determined at six concentration levels based on 

the LOQ for each element listed on page 1 [13]. All of the precision data was evaluated for homogeneity for 

all concentration levels tested using the Bartlett’s test and the results are listed in the method backup data 

report [12] and summarized in Tables 3 and 4. In many cases the highest concentration level (300 times 

the LOQ) was not poolable due in every case to the precision being so small relative to the other values, 

usually less than CV = 0.001 (<0.1%). Therefore, the overall precision (Ŝ ) and accuracy as given in Table 4 is rT

an upper limit predictor of precision; precision at concentration levels greater than 300 times the LOQ (see 

Table 3) will probably be much smaller.

For many of the metals, precision at the 3 times and/or 1 times the LOQ levels was reasonable (CV less than 

10%) but were not poolable due to the precisions at the higher concentration levels being so much smaller. 

In one case (strontium) the lowest level was not poolable because its CV was an inlier (less than 1%), being 

much smaller than those at the higher concentration levels. In most cases the precision appeared to be a 

function of concentration. This is observable in Table 3 where the CVs for the 10 times the LOQ (lower level) 

and 300 times the LOQ (higher) levels are compared.

Three elements, antimony, silver, and tin, had poor recoveries. It is believed that the chloride ions 

produced in the digestion of the PVC filters is causing the formation of precipitates. These metals are 

preferably sampled on MCE filters. The values in Tables 3 and 4 were determined using several different 

ICP-AES instruments and also several different microwave ovens. All were operated according to the 

manufacturer’s instructions.

REFERENCES:

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American

Conference of Governmental Industrial Hygienists. [www.acgih.org]. [Data accessed April 2014.]

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database on

hazardous substances (German Social Accident Insurance). Sankt Augustin, FRG: 

[https://www.dguv.de/ifa/index-2.jsp]. [Data accessed April 2014.]

[3] Hull, RD [1981]. Multi-element analysis of industrial hygiene samples, NIOSH internal report presented

at the American Industrial Hygiene Conference, Portland, Oregon, May.

[4] NIOSH [1994]. Elements: Method 7300 (Supplement Issued 3/15/2003). In: Schlecht PC, O’Connor

PF, eds. NIOSH Manual of analytical methods, 4th ed. Cincinnati, OH: U.S. Department of Health and

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health, DHHS Publication No. 94-113 [www.cdc.gov/niosh/nmam/].

[5] Harrison, GR [1969]. Massachusetts Institute of Technology wavelength tables, 2nd ed. West Hanover,

MA: Halliday Lithograph, Volume 1.

[6] ASTM International [2010]. ASTM D7035, Standard test method for the determination of metals

and metalloids in airborne particulate matter by inductively coupled plasma atomic emission

spectrometry. West Conshohocken, PA: ASTM International [www.astm.org].

[7] NIOSH [1994]. NIOSH manual of analytical methods (NMAM®). 4th ed. Schlecht PC, O’Connor PF, eds.

Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No.

94-113 [www.cdc.gov/niosh/nmam/].

[8] ASTM [1983]. ASTM D1193-77, Standard specification for reagent water. West Conshohocken, PA:

ASTM International [www.astm.org]. 
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 5 of 11

[9] Furr KA [1995]. CRC Handbook of Laboratory Safety, 4th ed. Boca Raton, FL: CRC Press.

[10] NIOSH [1973]. The industrial environment - its evaluation & control. 3rd ed. U.S. Department of 

Health Education and Welfare, Center for Disease Control, National Institute for Occupational 

Safety and Health. DHEW (NIOSH) Publication No. 74-117.

[11] NIOSH [1994]. Particulates not otherwise regulated: Method 0500. In: Eller PM, Cassinelli ME, 

eds. NIOSH Manual of Analytical Methods, 4th ed. Cincinnati, OH: U.S. Department of Health and 

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health. DHHS (NIOSH) Publication No. 94-113.

[12] Stone LC, Perkins JB, Rogers DM [2003]. Backup data report NMAM 7304: Elements by ICP on 

PVC filters using microwave digestion. Cincinnati OH: DataChem Laboratories, Inc. under NIOSH 

contract CDC-200-2001-08000. Unpublished.

[13] Lide DR [1993]. Handbook of chemistry and physics. 74th ed. Boca Raton FL: CRC PRess. 

[14] NIOSH pocket guide to chemical hazards. Barsan ME, ed. Cincinnati, OH: US Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149. [www.cdc.gov/niosh/

npg]. 

[15] Code of Federal Regulations, 29 CFR Part 1910.1000, Table Z-1[2013]. Limits for air standards. 

National occupational safety and health standards. [https://www.osha.gov/pls/oshaweb/

owadisp.show_document?p_table=STANDARDS&p_id=9992] Website accessed on December 13, 

2013.

METHOD WRITTEN BY:

David M. Rogers, Lee C. Stone, John M. Reynolds, James B. Perkins, DataChem Laboratories, Salt Lake 

City, Utah; Yvonne Gagnon, Ronnee Andrews, Ph.D., DART/CEMB.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their 

programs or products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses 

referenced in this document were accessible as of the publication date.
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 6 of 11

Table 1. PROPERTIES AND SAMPLING VOLUMES

Element (Symbol) Properties [13]

Atomic Weight MP, °C

Air Volume, L @ OSHA PEL [4]

MIN MAX

Aluminum (Al) 26.98 660 5 100

Arsenic (As) 74.92 817 5 2000

Barium (Ba) 137.3 727 5(2) 200(2)

Boron (B)(1) 10.81 2076 5 2000

Beryllium(Be) 9.01 1287 1250 2000

Calcium (Ca)(1) 40.08 842.5 5 200

Cadmium (Cd)(3) 112.40 321 13 2000

Cobalt (Co) 58.93 1495 25 2000

Chromium (Cr) 52.00 1907 5 1000

Copper (Cu) 63.54 1083 5 1000

Iron (Fe) 55.85 1538 5 100

Potassium (K)(1) 39.10 64 5 2000

Lithium (Li)(1) 6.94 181 100 2000

Magnesium (Mg) 24.31 651 5 67

Manganese (Mn) 54.94 1246 5 200

Molybdenum (Mo) 95.94 2623 5 67

Sodium (Na)(1)(3) 22.99 97.72 13 2000

Nickel (Ni) 58.71 1455 5 1000

Phosphorus (P) 30.97 44 25 2000

Lead (Pb) 207.19 327.2 50 2000

Platinum (Pt)(3) 195.1 1772.2 1250 2000

Selenium (Se) 78.96 221 13 2000

Strontium (Sr)(1) 87.62 777 5 2000

Tellurium (Te) 127.60 450 25 2000

Titanium (Ti) 47.90 1668 5 100

Thallium (Tl) 204.37 304 25 2000

Vanadium (V) 50.94 1910 5 2000

Yttrium (Y) 88.91 1522 5 1000

Zinc (Zn) 65.37 419 5 200

Zirconium (Zr) 91.22 1855 5 200

(1) No PEL, REL, or STEL data found [1,14].

(2) Air Volumes Estimated from TW A and LOQ’s (see Tables 2, 3). [10]

(3) These metals, as well as tin and antimony, form precipitates in nitric acid when chloride from 

 the PVC filters is present.
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 7 of 11

Table 2. EXPOSURE LIMITS, CAS #, RTECS [1,14,15]

Element

(Symbol) CAS # RTECS #

Exposure Limits, mg/m3

(C = ceiling limit)

OSHA NIOSH

Aluminum (Al) 7429-90-5 BD0330000 15 (total dust)

5 (respirable)

10 (total dust)

5 (respirable, fume) 

Arsenic (As) 7440-38-2 CG0525000 0.010 (inorganic) C 0.002(1)

Barium (Ba) 7440-39-3 CQ8370000 0.5 (soluble) 0.5 (soluble)

Beryllium (Be) 7440-41-7 DS1750000 0.002, C 0.005 C 0.0005(1)

Cadmium (Cd) 7440-43-9 EU9800000 0.005 lowest feasible conc.(1)

Calcium (Ca) 7440-70-2 No OEL No OEL

Cobalt (Co) 7440-48-4 GF8750000 0.1 0.05 (dust, fume)

Chromium (Cr) 7440-47-3 GB4200000 0.5 (II & III), 

0.005 (VI)

0.5 (II & III), 0.0002 (VI)

Cobalt (Co) 7440-48-4 GF8750000 0.1 0.05 (dust, fume)

Copper (Cu) 7440-50-8 GL5325000 1 (dust, mists)

0.1 (fume)

1 (dust, mists)

0.1 (fume)

Iron (Fe) 7439-89-6 NO4565500 10 (fume) as oxide 5 (dust, fume) as oxide

Magnesium (Mg) 7439-95-4 OM2100000 15 (dust) as oxide --

Manganese (Mn) 7439-96-5 OO9275000 C 5 1; STEL 3

Molybdenum (Mo) 7439-98-7 QA4680000 5 (soluble)

15 (total insoluble)

--

Nickel (Ni) 7440-02-0 QR5950000 1 0.015(1)

Phosphorus (P) 7723-14-0 TH3500000 0.1 0.1

Lead (Pb) 7439-92-1 OF7525000 0.05 0.05

Platinum (Pt) 7440-06-4 TP2160000 0.002 (soluble) 1 (metal)

Selenium (Se) 7782-49-2 VS7700000 0.2 0.2

Silver (Ag) 7440-22-4 VW3500000 0.01 (soluble, metal) 0.01 (soluble, metal)

Tellurium (Te) 13494-80-9 WY2625000 0.1 0.1

Titanium (Ti) 7440-32-6 XR1700000 15 (as TiO2

) lowest feasible(1)

Thallium (Tl) 7440-28-0 XG3425000 0.1 (soluble) 0.1(soluble)

Vanadium (V) 7440-62-2 YW1355000 C 0.5 (respirable) as 

V2O5

C 0.1 (fume) as V2

O5

C 0.05

Yttrium (Y) 7440-65-5 ZG2980000 1 1

Zinc (Zn) 7440-66-6 ZG8600000 5 (ZnO fume)

15 (ZnO dust)

5 (ZnO respirable)

5; STEL 10 (ZnO fume)

5; C 15 (ZnO dust)

Zirconium (Zr) 7440-67-7 ZH7070000 5 5, STEL 10

(1) Carcinogen
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 8 of 11

Table 3. MEASUREMENT WAVELENGTHS AND RECOVERY DATA

Element(1) Wavelength 

(nm)

LOD (μg/

sample)

Lower Level(4,5)

μg/

sample N = Percent Recovery

Precision 

Higher Level(5)

μg/

sample N = Percent Recovery

Precision 

(Sr)

Ag 328.07 0.1 3.00 5 63.01 0.0739 300 6 3.92 0.0865

Al 308.22 2 50.25(4) 6 89.78 0.0565 1500 6 100.71 0.0055

Al(2) 308.214 0.5 15.0 5 115.05 0.0199 1500 6 105.17 0.0056

As 193.76 2 15.0 5 93.29 0.0570 1500 6 115.84 0.0174

Ba 493.41 0.2 1.50 5 107.16 0.0295 150 6 102.22 0.0104

B 249.68 0.4 7.50 5 86.38 0.0277 750 6 101.19 0.0082

Be 313.04 0.008 0.152 6 102.38 0.0861 15.2 6 107.71 0.0091

Ca 315.89 151(4) 6 94.64 0.0512 4500 6 116.25 0.0153

Ca(2) 315.88 2 45.0 5 104.82 0.0090 4500 6 98.13 0.0066

Cd 228.80 0.2 3.00 5 109.65 0.0316 300 6 111.68 0.0152

Co 228.62 0.7 7.50 5 89.87 0.0338 750 6 114.15 0.0141

Cr 267.72 0.7 7.50 5 112.65 0.0233 750 6 118.65 0.0136

Cr(2) 267.71 0.3 7.50 5 102.60 0.0048 750 6 92.98 0.0066

Cu 324.75 0.08 1.50 5 106.84 0.0364 150 6 100.42 0.0058

Cu(2) 324.75 0.08 1.50 5 117.16 0.0361 150 6 103.13 0.0150

Fe 259.94 15 30 5 120.58 0.0405 3000 6 112.41 0.0083

Fe(2) 259.94 5 30 5 112.55 0.0489 3000 6 97.20 0.0085

K 766.49 3 100(4) 6 85.57 0.0254 3000 6 86.46 0.0260

K(2) 766.49 100(4) 6 99.40 0.0300 3000 6 90.02 0.0205

Li 670.78 0.06 1.50 5 97.51 0.0253 150 6 81.96 0.0378

Mg 279.08 0.9 15.0 5 105.25 0.0088 1500 6 97.47 0.0077

Mg(2) 279.07 0.4 15.0 5 107.33 0.0043 1500 6 101.75 0.0058

Mn 257.61 0.09 1.50 5 110.24 0.0150 150 6 115.56 0.0090

Mo 202.03 0.4 4.50 5 87.79 0.0433 450 6 120.57 0.0093

Mo(2) 202.029 0.3 4.50 5 89.75 0.0215 450 6 100.44 0.0154

Na 589.00 5 75.0 6 124.56 0.0859 7500 6 83.07 0.0248

Ni 231.60 0.3 4.50 5 102.93 0.0475 450 6 110.59 0.0080

Ni(2) 231.60 0.2 4.50 5 109.91 0.0047 450 6 101.77 0.0139

P 214.92 2 30.0 5 81.82 0.0511 3000 6 107.20 0.0103

P(2) 214.91 2 30.0 5 86.36 0.0077 3000 6 103.33 0.0174

Pb 220.35 1 15.0 5 95.85 0.0308 1500 6 100.54 0.0154

Pt 203.65 9 150 5 104.67 0.0182 15000 6 105.19 0.0088

Sb(3) 206.84 0.7 15.0 6 25.29 0.5861 1500 6 111.95 0.0086

(1) Values reported were obtained with a Fisons ARL Accuris ICP-AES unless otherwise noted; performance may vary with instrument

 and should be independently verified.

(2) Values reported were obtained with a Perkin Elmer Optima 3000 DV ICP-AES. 

(3) Elements that were evaluated and found not suitable for analysis by this method. 

(4) Values given (lower level) are for the 10xLOQ level due to low recoveries at the 3xLOQ level.

(5) LOQ = Estimated limit of quantitation
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 9 of 11

Table 3. MEASUREMENT WAVELENGTHS AND RECOVERY DATA

Element(1) Wavelength 

(nm)

LOD (μg/

sample)

Lower Level(4,5)

μg/

sample N = Percent Recovery

Precision 

(Sr)

Higher Level(5)

μg/

sample N = Percent Recovery

Precision 

(Sr)

Se 196.09 5 75.0 5 102.05 0.0531 7500 6 111.35 0.0063

Se(2) 196.02 2 75.0 5 99.93 0.0051 7500 6 99.72 0.0082

Sn 189.9 75.0 5 30.82 0.0502 7500 6 79.56 0.0124

Sn(2,3) 189.9 0.4 75.0 5 37.87 0.0816 7500 6 92.34 0.0129

Sr 421.55 0.04 7.50 5 100.00 0.0049 750 6 99.54 0.0055

Te 214.27 4 30.0 5 95.80 0.0624 3000 6 110.81 0.0094

Te(2) 214.28 2 30.0 5 97.18 0.0100 3000 6 99.64 0.0074

Ti 337.28 0.2 3.00 5 81.66 0.0392 300 6 103.42 0.0101

Ti(3) 334.94 0.1 3.00 5 82.68 0.0374 300 6 96.13 0.0121

Tl 190.86 2 15.0 5 96.38 0.0605 1500 6 97.25 0.0148

Tl(3) 190.79 1 15.0 5 97.75 0.0032 1500 6 92.04 0.0119

V 292.40 0.1 1.50 5 104.54 0.0528 150 6 111.15 0.0160

V(2) 292.40 0.09 1.50 5 100.99 0.0146 150 6 99.38 0.0232

Y 371.03 0.07 0.752 5 105.98 0.0245 75.2 6 105.03 0.0073

Zn 213.85 0.2 3.00 5 110.76 0.0327 300 6 116.84 0.0153

Zn(2) 213.86 0.4 3.00 5 93.45 0.0351 300 6 94.01 0.0055

Zr 339.20 0.2 1.50 5 102.61 0.0242 150 6 101.56 0.0144

(1) Values reported were obtained with a Fisons ARL Accuris ICP-AES unless otherwise noted; performance may vary with instrument

 and should be independently verified.

(2) Values reported were obtained with a Perkin Elmer Optima 3000 DV ICP-AES. 

(3) Elements that were evaluated and found not suitable for analysis by this method. 

(4) Values given (lower level) are for the 10xLOQ level due to low recoveries at the 3xLOQ level.

(5) LOQ = Estimated limit of quantitation
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 10 of 11

Table 4. OVERALL PRECISION AND ACCURACY DATA [13]

Element Instrument(1)

Range Studied 

(μg/sample)

From To

Bias Range of Bias

From To

Precision 

SrT

Accuracy 

(%)

Lowest 

Level(2)

Aluminum Fisons 5.025 1500 -0.0318 -0.1022 0.0240 0.0419 9.9 50.25

Aluminum P-E Optima 5.025 1500 0.0833 0.0567 0.1505 0.0379 15.1 15

Antimony Fisons 5.025 1500 Poor and variable recoveries across study range.

Arsenic Fisons 5.025 1500 0.0630 -0.0671 0.1584 0.0461 14.3 15

Barium Fisons 0.5038 150.4 0.0433 0.0222 0.0716 0.0182 7.6 0.5

Beryllium Fisons 0.0509 15.2 0.0652 0.0366 0.0980 0.0163 9.5 0.0509

Boron Fisons 2.514 750.4 -0.0387 -0.1362 0.0118 0.0164 6.4 7.504

Cadmium Fisons 1.005 300.0 0.0923 0.0718 0.1167 0.0307 14.8 1.005

Calcium Fisons 15.08 4500 0.0779 -0.0536 0.1624 0.0313 13.4 150.75

Calcium P-E Optima 15.08 4500 0.0453 0.0098 0.0963 0.0245 8.8 15.08

Chromium Fisons 2.514 750.4 0.1395 0.0974 0.1865 0.0214 18 2.514

Chromium P-E Optima 2.514 750.4 -0.0018 -0.0701 0.1245 0.0131 <5 2.514

Cobalt Fisons 2.514 750.4 0.0592 -0.1013 0.1508 0.0264 10.4 7.504

Copper Fisons 0.5038 150.4 0.0475 0.0272 0.0684 0.0240 8.9 0.5038

Copper P-E Optima 0.5038 150.4 0.0829 0.0313 0.1716 0.0217 12.1 1.504

Iron Fisons 10.05 3000 0.1101 0.0630 0.2057 0.0397 18.6 30

Iron Fisons 10.05 3000 0.0836 0.0630 0.0974 0.0396 15.4 100.5

Iron P-E Optima 10.05 3000 0.0445 -0.0205 0.1255 0.0404 11.4 30

Lead Fisons 5.025 1500 -0.0241 -0.0668 0.0124 0.0279 6.9 5.025

Lithium Fisons 0.5038 150.4 -0.0690 -0.1804 0.0132 0.0276 11.1 0.5038

Magnesium Fisons 5.025 1500 0.0156 -0.0253 0.0524 0.0171 <5 5.025

Magnesium P-E Optima 5.025 1500 0.0715 0.0421 0.1372 0.0249 11.5 5.025

Manganese Fisons 0.5038 150.4 0.1357 0.1005 0.1755 0.0201 17.3 0.5038

Fisons 1.509 450.4 -0.0388 -0.1597 0.1353 0.0795 16.7 1.509

P-E Optima 1.509 450.4 -0.0489 -0.2033 0.0969 0.0179 7.7 1.509

Nickel Fisons 1.509 450.4 0.0787 0.0293 0.1274 0.0338 13.8 4.504

Nickel P-E Optima 1.509 450.4 0.0645 0.0177 0.1406 0.0159 9.2 1.509

Phosphorus Fisons 10.05 3000 -0.0546 -0.1818 0.0011 0.0417 12 30

Phosphorus P-E Optima 10.05 3000 -0.0163 -0.1364 0.0333 0.0124 <5 10.05

Platinum Fisons 50 15000 0.0423 0.0097 0.0671 0.0226 8.2 150

Potassium Fisons 10.05 3000 -0.0909 -0.1443 -0.0316 0.0265 13.1 100.5

Potassium P-E Optima 10.05 3000 -0.0499 -0.0998 -0.0060 0.0249 8.8 100.5

Selenium Fisons 25.12 7500 0.0941 0.0675 0.1150 0.0150 12.1 25.12

Selenium P-E Optima 25.12 7500 0.0026 -0.0027 0.0115 0.0127 <5 25.12

Silver Fisons 1.005 300 Poor and variable recoveries across study range.

Sodium Fisons 25.12 7500 -0.0492 -0.1694 0.0718 0.0246 8.8 251.2

Strontium Fisons 2.514 750.4 0.0172 -0.00002 0.0373 0.0153 <5 2.514

Tellurium Fisons 10.05 3000 0.0295 -0.0420 0.1037 0.0404 9.8 30

(1) Values reported were obtained with a Fisons ARL Accuris ICP-AES or a Perkin Elmer Optima 3000 DV ICP-AES.

(2) Lowest level in range studied at which recoveries were between 81 and 121% recovery and relative standard deviation (Sr)

 less than 0.1100 on 5 or 6 replicates. Performance may vary with instrument and should be independently verified.
ELEMENTS by ICP Microwave Digestion: METHOD 7304, Issue 1, dated 25 May 2014 - Page 11 of 11

Table 4. OVERALL PRECISION AND ACCURACY DATA [13]

Element Instrument(1)

Range Studied 

(μg/sample)

From To

Bias Range of Bias

From To

Precision 

SrT

Accuracy 

(%)

Lowest 

Level(2)

Tellurium P-E Optima 10.05 3000 -0.0043 -0.0282 0.0163 0.0155 <5 10.05

Thallium Fisons 5.025 1500 -0.0081 -0.0362 0.0334 0.0407 8.2 15

Thallium P-E Optima 5.025 1500 -0.0505 -0.0688 -0.0048 0.0250 9 5.025

Tin Fisons 25.12 7500 Poor and variable recoveries across study range.

Tin P-E Optima 25.12 7500 Poor and variable recoveries across study range.

Titanium Fisons 1.005 300 -0.0827 -0.1834 0.0342 0.0269 12.3 3

Titanium P-E Optima 1.005 300 -0.1072 -0.1732 -0.0387 0.0321 15.3 1.005

Vanadium Fisons 0.5038 150.4 0.0704 0.0438 0.1114 0.0195 10.5 0.5038

Vanadium P-E Optima 0.5038 150.4 -0.0063 -0.0217 0.0099 0.0198 <5 0.5038

Yttrium Fisons 0.2519 75.2 0.0598 0.0466 0.0795 0.0164 8.9 0.2519

Zinc Fisons 1.005 300 0.1452 0.0630 0.2976 0.0340 22 1.005

Zinc Fisons 1.005 300 0.1190 0.0630 0.1683 0.0356 18.7 3

Zinc P-E Optima 1.005 300 -0.0502 -0.0655 -0.0388 0.0295 9.6 3

Zirconium Fisons 0.5025 150 0.0164 -0.0096 0.0350 0.0175 <5 0.5025

(1) Values reported were obtained with a Fisons ARL Accuris ICP-AES or a Perkin Elmer Optima 3000 DV ICP-AES.

(2) Lowest level in range studied at which recoveries were between 81 and 121% recovery and relative standard deviation (Sr)

 less than 0.1100 on 5 or 6 replicates. Performance may vary with instrument and should be independently verified.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler 7306

AW: Table 1 CAS: Table 2 RTECS: Table 2

METHOD: 7306, Issue 1 EVALUATION: FULL Issue 1: 10 September 2015

OSHA PELs: Table 2

NIOSH RELs: Table 2

OTHER OELs: [1,2]

PROPERTIES: Table 1 

ELEMENTS: aluminum

antimony 

arsenic 

barium 

 cadmium indium magnesium potassium tellurium yttrium

calcium iron manganese selenium thallium zirconium

chromium lanthanum molybdenum silver titanium zinc 

cobalt lead nickel strontium tungsten 

beryllium copper lithium phosphorus tin vanadium 

SAMPLING 

SAMPLER: Internal capsule, cellulose acetate dome with 

inlet opening, attached to 0.8-μm pore size 

mixed cellulose ester (MCE) membrane filter 

and housed within a 2-piece, closed-face 

cassette (CFC) filter holder, 37-mm diameter

FLOWRATE: 1 to 4 L/min

VOL-MIN: Table 1

-MAX: Table 1

SHIPMENT: Routine

SAMPLE 

STABILITY: Stable

BLANKS: Minimum of 2 field blanks per set

ACCURACY

RANGES STUDIED: Tables 3 and 4

BIAS: Table 4

OVERALL PRECISION (ŜrT) Table 4

ACCURACY: Table 4

MEASUREMENT

TECHNIQUE: INDUCTIVELY COUPLED PLASMA – 

ATOMIC EMISSION SPECTROMETRY 

(ICP-AES)

ANALYTES: Elements above

SAMPLE

DISSOLUTION: Hotplate digestion (NIOSH 7300 or 7301), 

microwave digestion (NIOSH 7302) or 

hot block extraction (NIOSH 7303)

SOLUTION: Dependent upon sample preparation 

method

WAVELENGTH: Depends upon element; See Table 3

BACKGROUND

CORRECTION: Spectral wavelength shift

CALIBRATION: Elements in acid matrix-matched to the 

sample; varies depending on sample 

preparation method

RANGE: Varies with element

ESTIMATED LOD: Table 3

PRECISION ( S ): Table 4 r

APPLICABILITY: The working range of this method is 4 x 10-5 mg/m3 to 10 mg/m3 for each element in a 500-L air sample. 

This is simultaneous elemental analysis, not compound specific. Verify that the types of compounds in the samples are 

soluble with the dissolution procedure selected. Some compounds of these elements require special sample treatment. 

INTERFERNCES: Spectral interferences are the primary interferences encountered in ICP-AES analysis. These are minimized 

by judicious wavelength selection, interelement correction factors and background correction [3,4].

OTHER METHODS: The internal capsule sampler used in this method is a recommended alternative to filter-only sampling [5] of NIOSH methods 7300 [6], 7301 [7], 7302 [8] and 7303 [9]. Use of an internal capsule sampler is an efficient means 

to account for sampler wall deposits that would otherwise be excluded by filter-only sampling. Unless other means are used 

to account for non-filter deposits inside the cassettes (e.g. within-cassette extraction, rinsing or wiping), internal capsule 

samplers should be used. OSHA method ID-125G [10] describes ICP-AES multielement analysis after hotplate digestion 

using nitric acid, sulfuric acid and hydrogen peroxide. ASTM D7035 [11] and ISO 15202 [12] are related voluntary consensus 

standard ICP-AES methods for multielement sampling and analysis of workplace atmospheres.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 2 of 11

REAGENTS:

1. Nitric acid (HNO3

)*, concentrated, Trace metal

grade

2. Hydrochloric acid (HCl)*, conc., Trace metal

grade (only required if hydrochloric acid

digestion is to be carried out)

3. Perchloric acid (HClO4

)*, conc., optima (only

required if perchloric acid digestion is to be

carried out)

4. Calibration stock solutions, 1000 μg/L:

Commercially available; or prepared per

instrument manufacturer’s recommendation

5. Dilution acid: 20% nitric acid* (7302);

4% nitric acid:1% perchloric acid* (7300);

5% aqua regia (7301)*; or 5% nitric acid:

5% hydrochloric acid* (7303) [dilution acid

is dependent upon sample preparation

method used]

6. Argon, as specified by ICP-AES manufacturer

7. Deionized water, ASTM Type II [13] or

equivalent

* See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Sampler: cellulose acetate internal capsule

attached to mixed cellulose ester membrane

filter, 0.8-μm pore size; 37-mm diameter, in

2-piece, closed-face cassette filter holder

2. Personal sampling pump, 1 to 4 L/min,

with flexible connecting tubing, capable of

maintaining constant flow

3. Inductively coupled plasma-atomic emission

spectrometer, equipped as specified by the

manufacturer for analysis of elements of

interest

4. Regulator, two-stage, for argon

5. Beakers, Phillips, 125-mL, or Griffin, 50-mL,

with watch glass covers**

6. Volumetric flasks, 10-, 25-, 100-mL, and 1-L**

7. Assorted volumetric pipets, as needed**

8. Forceps, plastic or plastic-tipped

9. Hotplate (NIOSH 7300 or 7301), microwave

oven (NIOSH 7302) or hot block (NIOSH

7303) [6-9].

10. Centrifuge tubes, 50-mL or sized

appropriately for hot block apparatus.

** Clean all glassware with conc. nitric acid and 

rinse thoroughly in deionized water before use

SPECIAL PRECAUTIONS: All perchloric acid digestions must be carried out in a perchloric acid fume 

hood. When working with concentrated acids, wear protective clothing, safety goggles and gloves. All 

work should be performed with adequate ventilation for personnel and equipment. It is imperative 

that acid be added to water in order to avoid a violent exothermic reaction.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

NOTE: See NMAM guidance chapters for discussion on sampling.

2. Sample at an accurately known flow rate between 1 and 4 L/min (± 5%) for a total sample

size of <1 to 2000 L (see Table 1) for TWA measurements. Do not exceed a sampler loading of

approximately 5 mg total dust.

NOTE: Filter overloading can be assessed by periodic visual checks. See NMAM guidance chapters

for additional discussion on filter capacity. 

SAMPLE PREPARATION:

3. Open the cassette filter holders and, using nonmetal forceps, transfer the samples and blanks to

clean digestion vessels.

NOTE: Samples may not easily fit into the digestion vessels. Care must be taken to ensure no

sample is lost during the placement of the samples in the digestion vessels. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 3 of 11

4. Carry out sample dissolution in accordance with one of the sample preparation procedures described

in NIOSH 7300, 7301, 7302, or 7303 [6-9].

NOTE: The dissolution acid level within the vessel should cover the internal capsule.

5. After allowing to cool to room temperature, transfer the solutions quantitatively to 25-mL volumetric

flasks.

6. Dilute to volume.

NOTE: If greater sensitivity is required, the final sample volume may be held to 10 mL.

CALIBRATION AND QUALITY CONTROL:

7. Calibrate the spectrometer according to the manufacturer recommendations.

NOTE: Typically, an acid blank and 1.0 μg/mL multielement calibration standards are used.

 The following multielement combinations are chemically compatible in 5% HNO3

:

a. Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, La, In, Na

b. Ag, K, Li, Mg, Mn, Ni, P, Pb, Se, Sr, Tl, V, Y, Zn, Sc

c. Mo, Sb, Sn, Te, Ti, W, Zr

8. Analyze at least one calibration standard per ten samples.

9. Check recoveries with at least one media blank and two spiked media blanks per twenty samples.

Media should be spiked with analytes of interest.

NOTE: Whenever possible, QA/QC samples should be prepared from certified reference materials in

a matrix similar to the bulk material sampled. Liquid spiked filters are only surrogates for real 

world samples and QC data based upon certified samples are preferred.

MEASUREMENT:

10. Set ICP-AES spectrometer to conditions specified by manufacturer.

11. Analyze standards and samples by ICP-AES in accordance with manufacturer recommendations.

NOTE: If the values for the samples are above the range of the standards, dilute the solutions (ensuring

the samples remain acid matrix-matched to the calibration standards), reanalyze and apply the 

appropriate dilution factor in the calculations.

CALCULATIONS:

12. Obtain the solution concentrations for the sample, Cs

 (μg/mL), and the average media blank ,

Cb

 (μg/mL), from the instrument.

13. Using the solution volumes of sample, Vs

 (mL), and media blank, Vb (mL), calculate the concentration,

C mg/m3

), of each element in the air volume sampled, V (L):

  , mg/m

NOTE: μg/L is approximately equal to mg/m3

EVALUATION OF METHOD:

A previous interlaboratory investigation of aerosol-loaded cellulosic capsules (provided by SKC, Inc., EightyFour, PA) provided the background information for the development of this method [14]. That investigation 

yielded data for Cd, Cr, Co, Cu, Fe, Mn, Ni and Pb [14]. Recoveries for these elements were quantitative and 

values of overall relative standard deviation were <0.20 [14], which compare favorably with the variability 

typically observed in interlaboratory multielement analysis of air samples [15,16]. 

To obtain performance data for additional elements, interlaboratory performance data were obtained using 

37-mm diameter cellulose acetate internal capsules attached to mixed-cellulose ester filters (Solu-sertTM from 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 4 of 11

Zefon International, Ocala, FL). Solu-sertTM capsules were dosed with 33 elements at three different 

spiking levels (spike levels certified by High-Purity Standards, North Charleston, SC), listed in Table 3. 

Sets of the spiked samples were conveyed to participating volunteer laboratories and analyzed by 

ICP-AES after sample dissolution. A variety of sample preparation procedures were used by the labs 

including NIOSH 7300, 7301, 7302, 7303, hot plate digestion utilizing nitric acid, sulfuric acid and 

hydrogen peroxide (modified NIOSH 7300), and microwave assisted digestion using nitric acid and 

hydrogen peroxide (modified NIOSH 7302). Results were received from 9 laboratories; however, not 

every spiked element was reported by each laboratory. Individual sample results may be found in the 

backup data report [17]. Statistical calculations were performed using SAS Software (version 9.2, SAS 

Institute Inc., Cary, NC). For each data subset, Grubbs’ test at 1% confidence level was used to identify 

outliers, which if identified, were removed prior to further statistical calculations. No statistically 

significant differences were found between the laboratory results for the spiked samples; therefore, all 

reported data (regardless of the sample preparation method) are included in Tables 3 & 4. Recoveries 

were quantitative (as defined in Kennedy, et al. [18]) and interlaboratory variability was <0.20 for most 

elements and loading levels (Table 3). Mean overall recoveries below 90% were found only for Cr, K, 

and W at low loadings and for Ag at medium and high spike levels and for In at the high spike level. 

RSD values > 0.20 were found only for Sn at the low spike level, Ag at medium and high spike levels, 

and In and K at all loadings. Results for precision, bias and accuracy are summarized in Table 4. 

With the domed top, the internal capsule can be difficult to fit into standard sample digestion 

vessels. One way to achieve this is to place the samplers into the vessels by bending them slightly 

inward (using coated forceps) and to push them into the bottom of the vessel to ensure they are 

covered with the digestion acids. Care must be taken to ensure that sample is not lost in this process. 

Additional guidance is available from the internal capsule manufacturer [19].

While no statistically significant differences were found based upon the sample preparation, it is 

important to note that interlaboratory variation is included in those calculations. Some differences in 

the sample preparation methods may have been statistically significant without that variability. Of 

particular importance are the less than quantitative recoveries for Sb, Sn, and Ti using NIOSH 7300. 

This sample preparation (as written) may not be amenable to the analysis of those elements. While this 

method lists several options for sample preparation, it is imperative that the suitability of the particular 

sample preparation method for the analytes of interest be considered.

Appreciable (>0.5 μg) media background levels were reported by Certified Reference Material (CRM) 

provider and/or the participating laboratories for several elements, notably Al, Ca, Cr, Fe, In, K, Mg, P, 

Sb, Se, and Tl. Trace media background levels of a few other elements, i.e., Ba, Cu and Zn, were also 

obtained. This background was effectively corrected for, as evidenced by the quantitative recoveries 

obtained for the vast majority of the elements and loading levels. Where the background levels may 

pose a greater influence is in the calculation of the method LOD. Method LODs (calculated using the 

standard deviation of blank responses) for internal capsule samples were greater than those calculated 

for MCE filters alone for many of the elements [17]. Care should be taken in choosing the appropriate 

media in concert with the expected sample concentrations.

REFERENCES:

[1] ACGIH [2014]. TLVs and BEIs based on the documentation of the Threshold Limit Values for 

chemical substances and physical agents and biological exposure indices. Cincinnati, Ohio: 

American Conference of Governmental Industrial Hygienists [www.acgih.org]. Date accessed 

May 2015.

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS 

database on hazardous substances (German Social Accident Insurance). Sankt Augustin, 

FRG: [http://www.dguv.de/ifa/GESTIS]. Date accessed May 2015. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 5 of 11

[3] Dean JA [1995]. Atomic spectroscopy. In: Analytical Chemistry Handbook; Section 7. New York,

NY: McGraw-Hill.

[4] Montaser A, Golightly DW, eds. [1992]. Inductively coupled plasmas in analytical atomic

spectrometry. 2nd ed. Hoboken, NJ: John Wiley & Sons, Inc.

[5] Ashley K, Harper M [2013]. Closed-face filter cassette (CFC) sampling – Guidance on procedures

for inclusion of material adhering to internal sampler surfaces. J Occup Environ Hyg 10:D29-D33.

[6] NIOSH [1994]. Elements by ICP (Nitric/Perchloric Acid Ashing): Method 7300. In: Schlecht PC,

O’Connor PF, eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department

of Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/

nmam/].

[7] NIOSH [1994]. Elements by ICP: Method 7301. In: Schlecht PC, O’Connor PF, eds. NIOSH manual

of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services,

Centers for Disease Control and Prevention, National Institute for Occupational Safety and

Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/].

[8] NIOSH [2014]. Elements by ICP (Microwave Digestion): Method 7302. In: Ashley K, O’Connor PF,

eds. NIOSH manual of analytical methods. 5th ed. Cincinnati, OH: U.S. Department of Health and

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health, DHHS (NIOSH) Publication No. 2014-151 [www.cdc.gov/niosh/nmam/].

[9] NIOSH [1994]. Elements by ICP (Hot Block/HCl/HNO3 Digestion): Method 7303. In: Schlecht PC,

O’Connor PF, eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department

of Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/

nmam/].

[10] OSHA [2002]. Metal and metalloid particulates in workplace atmospheres (ICP analysis).

In: OSHA Sampling and Analytical Methods. Sandy, UT: Occupational Safety and Health

Administration.

[11] ASTM International [2010]. ASTM D7035, Standard test method for the determination of metals

and metalloids in workplace air by inductively coupled plasma atomic emission spectrometry.

West Conshohocken, PA: ASTM International.

[12] ISO [2011]. ISO 15202, Workplace air – Determination of metals and metalloids in airborne

particulate matter by inductively coupled plasma atomic emission spectrometry (3 parts).

Geneva, Switzerland: International Organization for Standardization.

[13] ASTM [2011]. ASTM D1193, Standard specification for reagent water. West Conshohocken, PA:

ASTM International.

[14] Harper M, Ashley K [2013]. Acid-soluble internal capsules for closed-face cassette elemental

sampling and analysis of workplace air. J Occup Environ Hyg 10:297-306.

[15] Ashley K, Shulman SA, Brisson MJ, Howe AM [2012]. Interlaboratory evaluation of trace element

determination in workplace air samples by inductively coupled plasma mass spectrometry. J

Environ Monit 14:360-367.

[16] Stacey P, Butler OT [2008]. Performance of laboratories analysing welding fume on filter samples

– Results from the WASP Proficiency Testing Scheme. Ann Occup Hyg 52:287-295.

[17] Ashley K, Andrews RN, Feng HA [2014]. Elements by cellulosic internal sampler and ICP-AES,

NIOSH 7306, Backup Data Report. Cincinnati, OH, Unpublished.

[18] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation.

By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of

Health and Human Services, Centers for Disease Control and Prevention, National Institute for

Occupational Safety and Health, DHHS (NIOSH) Publication No. 95–117.

[19] Zefon International. Operating instructions Solu-SertTM soluble filter capsule. Ocala, FL: Zefon

International. [https://www.zefon.com/Content/Products/SamplingMedia/Solu-Sert_IFU.pdf] 

Date accessed: May 2015. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 6 of 11

[20] NIOSH [1994]. Measurement uncertainty and NIOSH method accuracy: Chapter P. In: Schlecht 

PC, O’Connor PF, eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. 

Department of Health and Human Services, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.

gov/niosh/nmam/].

[21] Lide DR [2014]. Handbook of Chemistry and Physics, 94th ed. Boca Raton: FL, CRC Press

[22] NIOSH [2005]. NIOSH pocket guide to chemical hazards. Barsan ME, ed. Cincinnati, OH: US 

Department of Health and Human Services, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149. [www.cdc. 

gov/niosh/npg].

METHOD WRITTEN BY:

Kevin Ashley, Ph.D., Ronnee N. Andrews, Ph.D., and H. Amy Feng, NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 7 of 11

TABLE 1. PROPERTIES AND SAMPLING VOLUMES [21]

Element (Symbol)

Atomic Weight

(AW) MP, °C

Volume, L @ OSHA PELA

MINB MAXC

Silver (Ag) 107.87 961 6 >2000D

Aluminum (Al) 26.98 660 <1 330

Arsenic (As) 74.92 817 32 >2000

Barium (Ba) 137.34 727 3 >2000

Beryllium (Be) 9.01 1287 10 >2000

Calcium (Ca) 40.08 842 -- --

Cadmium (Cd) 112.41 321 3 >2000

Cobalt (Co) 58.93 1495 <1 >2000

Chromium (Cr) 52.00 1907 1 >2000

Copper (Cu) 63.54 1083 <1 >2000

Iron (Fe) 55.85 1538 2 500

Indium (In) 114.82 156 8 >2000

Potassium (K) 39.10 64 -- --

Lanthanum (La) 138.91 920 -- --

Lithium (Li) 6.94 181 -- --

Magnesium (Mg) 24.31 651 <1 330

Manganese (Mn) 54.94 1246 <1 1000

Molybdenum (Mo) 95.94 2623 <1 330

Nickel (Ni) 58.71 1455 2 >2000

Phosphorus (P) 30.97 44 9 >2000

Lead (Pb) 207.19 328 4 >2000

Antimony (Sb) 121.75 631 1 >2000

Selenium (Se) 78.96 221 2 >2000

Tin (Sn) 118.69 232 <1 >2000

Strontium (Sr) 87.62 777 -- --

Tellurium (Te) 127.60 450 7 >2000

Titanium (Ti) 47.87 1668 -- --

Thallium (Tl) 204.37 304 1 >2000

Vanadium (V) 50.94 1910 -- --

Tungsten (W ) 183.85 3422 -- --

Yttrium (Y) 88.91 1522 <1 >2000

Zinc (Zn) 65.37 419 -- --

Zirconium (Zr) 91.22 1855 <1 1000

A Exposure limits listed in Table 2.

B Min calculated using the method quantitation limits following NIOSH 7302

 sample preparation.

C Max calculated using 5 mg of sample collected.

D Indicates that the calculated maximum volume is greater than the volume collected

at the maximum flow rate (4 L/min) of the method for an 8-hour shift.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 8 of 11

TABLE 2. EXPOSURE LIMITS, CAS#, RTECS [22]

Element 

(Symbol) CAS # RTECS #

Exposure Limits

OSHA PEL (mg/m3

) NIOSH REL (mg/m3)

Silver (Ag) 7440-22-4 VW3500000 0.01 (dust, fume, metal) 0.01 (metal, soluble) 

Aluminum (Al) 7429-90-5 BD0330000 15 (total dust)

5 (respirable)

10 (total dust)

5 (respirable fume) 

2 (soluble) 

Arsenic (As) 7440-38-2 CG0525000 0.010 CE 0.002 (15 min), CaF 0.5 

Barium (Ba) 7440-39-3 CQ8370000 0.5 

Beryllium (Be) 7440-41-7 DS1750000 0.002, C 0.005 C 0.0005, Ca 

Calcium (Ca) 7440-70-2 -- -- -- 

Cadmium (Cd) 7440-43-9 EU9800000 0.005 lowest feasible, Ca 

Cobalt (Co) 7440-48-4 GF8750000 0.1 (dust, fume) 0.05 (dust, fume) 

Chromium (Cr) 7440-47-3 GB4200000 1 (metals, insoluble salts) 

0.5 (Cr II & Cr III)

0.5 

Copper (Cu) 7440-50-8 GL5325000 1 (dust, mists) 

0.1 (fume)

1 (dust, mists) 

0.1 (fume)

Iron (Fe) 7439-89-6 NO4565500 10 (fume) as oxide 5 (dust, fume) oxide as Fe 

Indium (In) 7440-74-6 NL1050000 0.1 0.1

Potassium (K) 7440-09-7 TS6460000 -- --

Lanthanum 7439-91-0 -- – – 

Lithium (Li) 7439-93-2 OJ5540000 -- --

Magnesium (Mg) 7439-95-4 OM2100000 15 (dust) as oxide -- 

Manganese (Mn) 7439-96-5 OO9275000 C 5 C 1; STELG 3 

Molybdenum (Mo) 7439-98-7 QA4680000 5 (soluble) 

15 (total insoluble dust)

--

Nickel (Ni) 7440-02-0 QR5950000 1 0.015, Ca 

Phosphorus (P) 7723-14-0 TH3500000 0.1 0.1 

Lead (Pb) 7439-92-1 OF7525000 0.05 0.05

Antimony (Sb) 7440-36-0 CC4025000 0.5 0.5 

Selenium (Se) 7782-49-2 VS7700000 0.2 0.2 

Tin (Sn) 7440-31-5 XP7320000 2 2 

Strontium (Sr) 7440-24-6 WK7700000 – – 

Tellurium (Te) 13494-80-9 WY2625000 0.1 0.1 

Titanium (Ti) 7440-32-6 XR1700000 15 as dioxide Ca as the oxide

Thallium (Tl) 7440-28-0 XG3425000 0.1 (skin) (soluble) 0.1 (skin) (soluble) 

Vanadium (V) 7440-62-2 YW1355000 C 0.5 (respirable) as 

pentoxide

C 0.05 as pentoxide

Tungsten (W) 7440-33-7 YO7175000 -- 5; STEL 10

Yttrium (Y) 7440-65-5 ZG2980000 1 1 

Zinc (Zn) 7440-66-6 ZG8600000 5 (fume, respirable dust) 

as oxide

15 (total dust) as oxide

5 (dust, fume) as oxide

C 15 (dust) as oxide

STEL 10 (fume) as oxide

Zirconium (Zr) 7440-67-7 ZH7070000 5 5; STEL 10 

E C: ceiling

F

 Ca: carcinogen

G STEL: short-term exposure limit
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 9 of 11

TABLE 3. MEASUREMENT WAVELENGTHS (λ), DETECTION LIMITS (LOD) AND RECOVERY DATA FROM INTERLABORATORY RESULTS

Element

WavelengthH

(λ), nm

LODI 

(ug/sample)

Low Level 

(ug/sample)

% Recovery

(%RSD; NJ)

Medium 

level

(ug/ sample)

% Recovery

 (%RSD; NJ)

High Level 

(ug/ sample)

% Recovery

 (%RSD; NJ)

Ag 328.068 0.020 5.0 93.2 (17; 7) 10.1 83.5 (26; 7) 20.1 68.2 (52; 7)

Al 396.152 0.38 10.6 95.7 (6.1; 7) 30.9 95.7 (3.1; 7) 60.8 96.2 (2.4; 7)

As 189.042 0.099 5.0 99.4 (7.3; 7) 20.2 103 (5.3; 8) 40.1 102 (5.8; 8)

Ba 455.404 0.55 2.21 97.0 (8.4; 8) 7.3 96.9 (2.5; 7K

) 15.2 99.9 (5.0; 8)

Be 313.042 0.0064 2.01 101 (4.4; 9) 7.0 101 (6.8; 9) 14.9 100 (5.1; 9)

Ca 315.887 3.9 114 95.8 (16; 8) 165 97.4 (15; 8) 215 95.8 (13; 8)

Cd 226.502 0.0052 2.01 101 (3.6; 9) 7.0 101 (4.0; 9) 14.9 102 (1.9; 8)

Co 228.616 0.0090 2.01 103 (6.8; 9) 7.0 102 (5.7; 9) 14.9 102 (5.8; 9)

Cr 267.716 0.28 2.91 80.2 (16; 8) 7.9 96.3 (7.2; 9) 15.8 97.2 (3.5; 9)

Cu 324.754 0.15 3.16 100 (4.5; 8K

) 15.1 101 (4.0; 9) 29.9 100 (3.2; 9)

Fe 259.941 5.3 21.3 107 (15; 9) 41.0 104 (5.5; 9) 80.5 103 (5.4; 9)

In 230.606 0.26 5.0 92.1 (33; 5) 14.9 90.8 (27; 5K

) 39.7 86.2 (32; 5)

K 766.491 0.70 10.6 89.0 (36; 5) 15.7 98.0 (24; 6) 20.7 106 (20; 6)

La 408.672 0.026 3.01 101 (8.1; 4) 10.1 102 (7.2; 4) 20.1 99.9 (7.9; 4)

Li 670.780 0.010 2.01 93.0 (8.5; 7) 7.0 94.9 (7.0; 7) 14.9 98.6 (5.2; 7)

Mg 279.079 1.1 12.7 93.9 (16; 7) 27.9 98.7 (9.1; 7) 103 101 (7.2; 7)

Mn 294.921 0.031 2.01 99.8 (7.6; 9) 7.0 100 (6.1; 9) 14.9 100 (5.7; 9)

Mo 202.095 0.021 2.01 101 (6.2; 9) 7.1 102 (4.8; 9) 15.0 102 (5.2; 9)

Ni 231.604 0.56 2.01 108 (9.6; 9) 7.0 104 (7.4; 9) 14.9 103 (7.0; 9)

P 178.287 0.27 10.1 104 (9.0; 5) 24.9 103 (8.9; 6) 99 102 (1.7; 5K

)

Pb 220.353 0.062 10.0 101 (6.2; 9) 25.2 100 (5.3; 9) 100 100 (7.3; 9)

H Commonly used wavelength; choose wavelength appropriate for your instrument settings [3,4]

I

 LOD values calculated using the responses of 7 media blanks, prepared following the microwave sample preparation in NIOSH 7302

J

 Mean recovery & relative standard deviation for N = number of laboratories reporting results for each element

K

 Excludes outlier(s) (Grubbs’ test)
ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 10 of 11

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 3 continued from page 9

TABLE 3. MEASUREMENT WAVELENGTHS (λ), DETECTION LIMITS (LOD) AND RECOVERY DATA FROM INTERLABORATORY RESULTS

Element

WavelengthH

(λ), nm

LODI 

(ug/sample)

Low Level 

(ug/sample)

% Recovery

(%RSD; NJ)

Medium 

level

(ug/ sample)

% Recovery

 (%RSD; NJ)

High Level 

(ug/ sample)

% Recovery

 (%RSD; NJ)

Sb 206.833 0.11 5.0 99.1 (6.7; 8) 25.1 99.0 (5.7; 9) 40.2 100 (3.6; 8K)

Se 196.090 0.14 3.0 112 (15; 6) 15.1 109 (8.7; 8) 30.1 104 (10; 8)

Sn 189.991 0.065 2.01 98.7 (26; 4) 7.0 96.9 (18; 6) 14.9 103 (5.0; 5K

)

Sr 407.771 0.014 2.01 100 (1.9; 6K

) 7.1 102 (7.5; 7) 15.0 101 (6.1; 7)

Te 214.281 0.22 3.0 104 (18; 5) 12.6 103 (7.7; 5) 20.1 101 (12; 5)

Ti 334.941 0.042 2.01 105 (5.7; 7) 7.0 101 (10; 7) 14.9 103 (3.4; 6K

)

Tl 190.864 0.046 3.0 102 (7.1; 6) 10.1 99.6 (5.8; 7) 20.1 98.3 (7.5; 7)

V 311.071 0.0091 3.02 101 (5.7; 9) 7.0 102 (6.2; 9) 14.9 102 (5.8; 9)

W 207.911 0.055 10.1 83.0 (12; 5) 25.1 97.6 (12; 5) 40.2 96.4 (16; 5)

Y 371.030 0.0039 2.01 101 (3.8; 5) 7.1 102 (5.9; 5) 15.0 101 (4.8; 5)

Zn 213.856 0.69 5.2 101 (6.0; 9) 25.1 101 (4.9; 9) 59.7 101 (4.8; 9)

Zr 339.198 0.0099 2.01 101 (6.2; 5) 7.0 99.5 (9.2; 6) 14.9 93.1 (19; 6)

H Commonly used wavelength; choose wavelength appropriate for your instrument settings [3,4]

I

 LOD values calculated using the responses of 7 media blanks, prepared following the microwave sample preparation in NIOSH 7302

J

 Mean recovery & relative standard deviation for N = number of laboratories reporting results for each element

K

 Excludes outlier(s) (Grubbs’ test)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ELEMENTS by Cellulosic Internal Capsule Sampler: METHOD 7306, Issue 1, dated 10 September 2015 - Page 11 of 11

TABLE 4. RANGE, BIAS, PRECISION (  and ŜrT) AND ACCURACY DATA FROM

INTERLABORATORY RESULTS

Element nL Range, ug/sample Bias M

  Ŝr

N Accuracy (%)

Ag 21 5.0 to 20.1 -0.184 0.041 0.065 29.0

Al 21 10.6 to 60.8 -0.0414 0.006 0.050 12.4

As 23 5.0 to 40.1 0.0141 0.016 0.052 10.7

Ba 23 2.21 to 15.2 -0.0206 0.036 0.062 12.8

Be 27 2.01 to 14.9 0.00536 0.025 0.056 11.0

Ca 24 114 to 215 -0.0367 0.001 0.050 11.9

Cd 26 2.01 to 14.9 0.0133 0.022 0.055 11.1

Co 27 2.01 to 14.9 0.0238 0.036 0.062 12.9

Cr 26 2.91 to 15.8 -0.0281 0.046 0.068 14.4

Cu 26 3.16 to 29.9 0.00347 0.017 0.053 10.4

Fe 27 21.3 to 80.5 0.0476 0.011 0.051 13.2

In 15 5.0 to 39.7 -0.103 0.056 0.075 22.6

K 17 10.6 to 20.7 -0.0239 0.029 0.058 12.3

La 12 3.01 to 20.1 0.0119 0.025 0.056 11.2

Li 21 2.01 to 14.9 -0.0447 0.039 0.064 14.9

Mg 21 12.7 to 103 -0.0219 0.012 0.051 10.9

Mn 27 2.01 to 14.9 0.00127 0.039 0.063 12.4

Mo 27 2.01 to 15.0 0.0169 0.032 0.060 12.1

Ni 27 2.01 to 14.9 0.0498 0.055 0.074 17.2

P 16 10.1 to 99 0.0310 0.010 0.051 11.5

Pb 27 10.0 to 100 0.00439 0.006 0.050 9.9

Sb 25 5.0 to 40.2 -0.00631 0.013 0.052 10.2

Se 22 3.0 to 30.1 0.0864 0.055 0.075 20.9

Sn 15 2.01 to 14.9 -0.00541 0.117 0.128 25.0

Sr 20 2.01 to 15.0 0.00930 0.011 0.051 10.2

Te 15 3.0 to 20.1 0.0256 0.063 0.081 16.6

Ti 20 2.01 to 14.9 0.0305 0.032 0.059 13.1

Tl 20 3.0 to 20.1 0.00128 0.030 0.058 11.4

V 27 3.02 to 14.9 0.0143 0.020 0.054 10.9

W 15 10.1 to 40.2 -0.0240 0.010 0.051 11.0

Y 15 2.01 to 15.0 0.00971 0.020 0.054 10.7

Zn 27 5.2 to 59.7 0.00873 0.013 0.052 10.3

Zr 17 2.01 to 14.9 -0.0206 0.033 0.060 12.4

L n = total number of results reported for each element by up to 9 participating laboratories minus outliers

 (Grubbs’ test, 1% confidence level)

M 

  = precision [17

N ŜrT = overall precision =  ; same as total precision as defined in [18,20
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTOS and OTHER FIBERS by PCM 7400

Formula: Various MW: Various CAS: Table 3 RTECS: Various

METHOD: 7400, Issue 3 EVALUATION: FULL Issue 1: 15 May 1989

Issue 3: 14 June 2019 

OSHA: 0.1 asbestos fiber (>5 μm long and ≥3:1 aspect

ratio)/cc; 1 f/cc, 30 min excursion; carcinogen

NIOSH: 0.1 fiber (>5 μm long and ≥3:1 aspect ratio)/cc, for a

400 L sample; carcinogen

MSHA: As OSHA

PROPERTIES: solid, fibrous, crystalline, anisotropic 

SYNONYMS: actinolite or ferroactinolite; amosite; anthophyllite; chrysotile; crocidolite; tremolite; amphibole asbestos; refractory 

ceramic fibers; fibrous glass

SAMPLING

SAMPLER: FILTER (0. 45- to 1.2-μm mixed cellulose 

ester membrane, 25-mm; conductive cowl 

on cassette

FLOW RATE*: 0.5 to 16 L/min

VOL-MIN*: 400 L @ 0.1 fiber/cc

-MAX*: (step 4, Sampling)

*Adjust to give 100 to 1300 fiber/mm2 

SHIPMENT: routine (pack to reduce mechanical and 

static electrical shock) (step 6, Sampling)

SAMPLE

STABILITY: stable 

BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE 

STUDIED: 80 to 100 fibers counted

BIAS: see Evaluation of Method

OVERALL

PRECISION (SS�rrrr): 0.115 to 0.13 [1]

ACCURACY: see Evaluation of Method

MEASUREMENT

TECHNIQUE: LIGHT MICROSCOPY, PHASE CONTRAST

ANALYTE: fibers (manual count)

SAMPLE

PREPARATION: Treatment of filter by acetone or

dimethylformamide (DMF)/acetic acid, 

followed by triacetin or Euparal mounting 

medium [2-4]

COUNTING

RULES: Described in previous version of this

method as “A” rules [1,5]

EQUIPMENT: 1. positive phase-contrast microscope; 

2. graticule (100-μm field of view);

3. phase-shift test slide

CALIBRATION: Phase-shift test slide

RANGE: 100 to 1300 fibers/mm2 filter area

ESTIMATED LOD: 7 fibers/mm2 filter area

PRECISION (SSrr): 0.10 to 0.12 [1]; see Evaluation of Method

APPLICABILITY: The quantitative working range is 0.04 to 0.5 fiber/cc for a 1000-L air sample. The LOD depends on sample 

volume and quantity of interfering dust, and is <0.01 fiber/cc if free of interferences. The method gives an index of airborne fibers. 

This method can be used in conjunction with electron microscopy (e.g., Method 7402) for assistance in identification of fibers. For 

fibers with diameters >1 μm, polarizing light microscopy (as in NIOSH Method 7403) may be used to identify and eliminate 

interfering non-crystalline fibers [6]. Asbestos fibers thinner than about 0.05-0.15 μm diameter, depending on asbestos type, will 

not be detected by this method [7-10]. This method may be used for other materials with alternate counting rules.

INTERFERENCES: If the method is used to detect a specific type of fiber, any other fiber may interfere because all particles meeting 

the counting criteria are counted. Chain-like particles may appear fibrous. High levels of non-fibrous dust particles may obscure 

fibers in the field of view and increase the detection limit. 

OTHER METHODS: This revision replaces Method 7400, issue 2 (dated 08/15/1994).
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 2 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Acetone*, reagent grade.

NOTE: Dimethylformamide (DMF)*, reagent

grade/glacial acetic acid can be used 

as an alternative filter clearing 

reagent.

2. Triacetin (glycerol triacetate), reagent grade.

NOTE: Euparal (synthetic Canada Balsam)

can be used as an alternative 

mounting media.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Field monitor, 25-mm, 3-piece

cassette with 50-mm electrically conductive

extension cowl and mixed cellulose ester

(MCE) filter, 0.45- to 1.2-μm pore size, and

backup pad.

NOTE 1: Analyze representative filters for fiber

background before use to check for 

clarity and background. Discard the 

filter lot if mean is ≥ 5 fibers per 100 

graticule fields. These are defined as

laboratory blanks. Manufacturerprovided quality assurance checks on 

filter blanks are normally adequate as 

long as field blanks are analyzed as 

described below. 

NOTE 2: The electrically conductive extension 

cowl reduces electrostatic effects 

[11]. Ground the cowl when possible 

during sampling.

NOTE 3: 0.8- μm pore size filters are 

commonly used for personal 

sampling. However, 0.45-μm filters 

are recommended for sampling 

when performing TEM analysis on 

the same samples. Check personal 

sampling pumps before use with 

0.45-μm filters to ensure they can 

operate at the higher pressure drop. 

Perform calibration with same type 

of filter as used for sampling.

2. Sampling pump, battery or line-powered

vacuum, of sufficient capacity to meet flow

rate requirements and, for personal sampling

pumps, applicable ISO Standard [12], with

flexible connecting tubing.

NOTE: See Step 4 in Sampling section for flow

rate.

3. Wire, multi-stranded, 22-gauge; 1” hose

clamp to attach wire to cassette for

grounding, if needed.

4. Tape, shrink- or adhesive-, or cellulose bands.

5. Slides, glass, pre-cleaned, 25- x 75-mm.

6. Cover slips, 22- x 22-mm, No. 1 1⁄2, unless

otherwise specified by microscope

manufacturer.

7. Cover slips, as above, imprinted with a

relocatable grid, where required for quality

assurance or training purposes.

8. Lacquer or nail polish.
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 3 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

9. Knife, #10 surgical steel, curved blade, or

scissors.

10. Forceps (tweezers).

11. Flash vaporization system for clearing filters

on glass slides using acetone. (See ref. [11] for

specifications or see manufacturer’s

instructions for equivalent devices.). Use a

drying oven or warming plate located in fume 

cabinet if using DMF/glacial acetic acid.

12. Micropipettes or microsyringes, 5-μL and 100-

to 500-μL.

13. Microscope, positive phase (dark) contrast,

with green or blue filter, adjustable field iris,

8x to 10x eyepiece, and 40x to 45x phase

objective (total magnification ca. 400x);

numerical aperture (NA) = 0.65 to 0.75.

14. Graticule, Walton-Beckett type (Type G22 and

G24 are optimized for different counting rules

per Appendix C.) with 100-μm projected

diameter circular field (area = 0.00785 mm2) at 

the specimen plane. Alternative graticules

may be used where similar performance has

been demonstrated; e.g., the RIB graticule.

NOTE: The graticule is custom-made for each 

microscope. See APPENDIX A for the 

custom-ordering process. 

15. Phase contrast test slide. A slide with blocks

of visible ruled lines where at least one block

of lines is certified as invisible under the

microscope set up conditions given below.

16. Telescope, ocular phase-ring centering.

17. Stage micrometer (0.01-mm divisions).

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities and analysis. It 

is essential that suitable gloves, eye protection, laboratory coat, etc., be used when working with the 

chemicals. Acetone is toxic at high exposures and is extremely flammable. Take precautions not to ignite 

it. Heating of acetone in volumes greater than 1 mL must be done in a ventilated laboratory fume hood 

using a flameless, spark-free heat source. DMF is toxic by inhalation; heating in a drying oven or warming 

plate located in an operating fume exhaust hood will reduce potential exposure. DMF is also toxic via 

absorption through the skin. 

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line. Use a flow meter 

whose calibration is traceable to national or international standards.

NOTE: See NMAM guidance chapters for discussion on sampling.

2. Cassette assemblies shall be tested to ensure they are not likely to fall apart during sampling. (This

testing can be undertaken by the manufacturer.) A press may be useful in ensuring a tight fit, but shall

not cause the filter to be cut. To reduce contamination, seal the crease between the cassette base and

the cowl with a shrink band or light-colored adhesive tape. Commercial pre-assembled cassettes may

already include a taped seal. For personal sampling, fasten the uncapped open-face cassette to the

worker’s lapel. The open face shall be oriented downward. 
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 4 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NOTE: Electrically grounding the cowl is highly recommended during area sampling, where possible, 

but especially under conditions of low relative humidity. Use a hose clamp to secure one end of 

the wire (Equipment, Item 3) to the monitor’s cowl. Connect the other end to an earth ground 

(e.g., cold water pipe). It is recognized that circumstances do not always allow this procedure.

3. Submit at least two field blanks (or 10% of the total samples, whichever is greater) for each set of

samples. Handle field blanks in a manner representative of actual handling of associated samples in the

set. Open field blank cassettes at the same time as other cassettes just prior to sampling. Store top

covers and cassettes in a clean area (e.g., a closed bag or box) with the top covers from the sampling

cassettes during the sampling period.

4. Sample at 0.5 L/min or greater [13]. Adjust sampling flow rate, Q [L/min], and time, t (min), to produce a

fiber density, E, of 100 to 1300 fibers/mm2 (3.85×104 to 5×105 fibers per 25-mm filter with effective

collection area Ac = 385 mm2

) for optimum counting. These variables are related to the concentration of

fibers in air, L (fibers/cc), of the fibrous aerosol being sampled by:

tt = AAcc ∗ EE

QQ ∗ LL ∗ 103

NOTE 1: The purpose of adjusting sampling times is to obtain optimum fiber loading on the filter. The 

collection efficiency does not appear to be a function of flow rate in the range of 0.5 to 16 

L/min for asbestos fibers [14]. However, counting efficiency is a function of filter loading, with 

lower loadings typically resulting in higher proportional concentrations [14-16]. A sampling 

rate of 1 to 4 L/min for 8 h is appropriate in atmospheres containing about 0.1 fiber/cc in the 

absence of significant amounts of non-asbestos dust. Dusty atmospheres require smaller 

sample volumes (≤400 L) to obtain countable samples. In such cases take short, consecutive 

samples and average the results over the total collection time. For documenting episodic 

exposures, use high flow rates (7 to 16 L/min) over shorter sampling times. In relatively clean 

atmospheres, where targeted fiber concentrations are much less than 0.1 fiber/cc, use larger 

sample volumes (3000 to 10000 L) to achieve quantifiable loadings. Take care, however, not 

to overload the filter with background dust. If ≥50% of the filter surface is covered with 

particles, the filter may be too overloaded to count and will bias the measured fiber 

concentration. 

NOTE 2: OSHA regulations specify a minimum sampling volume of 48 L for an excursion measurement, 

and a maximum sampling rate of 2.5 L/min for all personal asbestos sampling [5].

5. At the end of sampling, shut off the pump, record the time, remove the cassette from the tube

attaching it to the pump, and replace top cover and end plugs. Capping the cassette before shutting off

the pump and removing the tubing will cause a vacuum within the cassette and damage to the filter.

6. Ship samples with conductive cowl attached in a rigid container with packing material to protect

cassettes from jostling or damage.

NOTE: Do not use untreated polystyrene foam in shipping container because electrostatic forces may

cause fiber loss from sample filter. 

SAMPLE PREPARATION:

Two acceptable procedures for clarifying either a whole filter or a portion of a filter are described below. 

These procedures are the Acetone clearance procedure and the DMF/acetic acid clearance procedure. In 

either procedure, the filter material is placed on a glass microscope slide. It is then made transparent 

(clarified). Then, either triacetin or Euparal are placed on the clarified filter and a cover slip is placed on top.

NOTE 1: The object is to produce samples with a smooth (non-grainy) background in a medium with 

refractive index ≤ 1.46. An early method (P&CAM 239 in APPENDIX F) used dimethyl phthalate 

and diethyl oxalate [17]. This method may still be used as an alternative to those described 

below, but because the preparations are only stable for one to two days, the procedure is not 

further discussed. 
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 5 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NOTE 2: Other procedures use a technique for clearing the filter, either acetone (section A) or 

DMF/glacial acetic acid (section B), followed by mounting the cleared preparation (section C 

or D). Acetone collapses the filter into a gel, and has the advantage of being the fastest 

procedure to prepare the filter for the mounting medium. However, acetone can leave 

residual polymeric structures in the filter that may be counted as fibers. DMF/acetic acid 

dissolves the filter and does not leave residual structures to the same extent as acetone 

collapse. 

NOTE 3: After the filter is clarified, either triacetin (section C) or Euparal (section D) are acceptable 

alternative mounting media. Triacetin provides reasonably permanent mounts when the 

amount used is restricted to ≤ 3.5 μL, otherwise fiber migration has been observed in slides 

made with larger quantities [4]. Euparal provides permanent mounts for long-term storage (> 

5 years) regardless of the amount used. Either mounting medium can be used with either 

filter preparation procedure, and all combinations have been demonstrated to provide 

equivalent fiber counts [4].

7. Prepare samples in a clean area away from any bulk samples which might contaminate the samples.

Ensure that the glass slides and cover slips are free of dust and fibers.

8. Determine and record the effective sample collection or filtration area and record the information

referenced against the sample ID number.

9. Cut wedges of about 1/4 for a 25-mm diameter filter using a curved-blade surgical steel knife using a

rocking motion to prevent tearing. Do not use a blade that was used to open the cassette. Scissors are

an option for cutting the filter. The entire filter can be used instead of a wedge, but with the

understanding that the whole sample is lost if there is a problem in preparation. Place the wedge or

entire filter, dust side up, on slide using forceps. To prevent cross-contamination of samples, blades or

scissors should be cleaned between samples.

NOTE 1: Filters can be cut while still in the base of the cassette, or removed and cut on a special cutstand. If a cut-stand is used it shall be cleaned between filters to ensure no crosscontamination. 

NOTE 2: Static electricity will usually keep the wedge on the slide.

A. ACETONE CLEARANCE PROCEDURE

NOTE 1: The aluminum “hot block” or similar flash vaporization techniques may be used outside the

laboratory [2].

NOTE 2: Excessive water in the acetone may slow the clearing of the filter. Also, filters that have been 

exposed to high humidity or water prior to clearing may have a grainy background.

NOTE 3: Rest the “hot block” on a ceramic plate and isolate it from any surface susceptible to heat 

damage unless it is designed with appropriate safety features to prevent fire or damage. 

A1. If the temperature can be varied, adjust the “hot block” to ca. 70 °C [2], otherwise switch on until ready.

A2. Mount a wedge cut from the sample filter on a clean glass slide. Insert slide with wedge into the 

receiving slot at base of the “hot block.” Immediately place the tip of a micropipette or microsyringe 

containing ca. 250 μL acetone (use the minimum volume needed to consistently clear the filter 

sections) into the inlet port of the PTFE cap on top of the “hot block” and inject the acetone into the 

vaporization chamber with a slow, steady pressure on the plunger button while holding the 

micropipette or microsyringe firmly in place. After waiting 3 to 5 seconds for the filter to clear, remove 

the micropipette or microsyringe and glass slide from their ports.

NOTE: Using excess acetone, or delivering it too fast into the “hot block” may cause material to be 

washed off the surface of the filter. Using insufficient acetone may cause the filter to curl. 

CAUTION: Although the volume of acetone used is small, use safety precautions. Work in a wellventilated area (e.g., laboratory fume hood). Take care not to ignite the acetone. Continuous 

use of this device in an unventilated space may produce unhealthful levels or even explosive 

acetone vapor concentrations.
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 6 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

B. DMF/ACETIC ACID CLEARANCE PROCEDURE

B1. Make a mixture of 1.4mL (35% volume) of DMF, 0.6mL (15% volume) of glacial acetic acid and 2mL 

(50% volume) of distilled water in a dark brown vial [3]. This should be done in a fume hood to 

minimize exposure to chemicals. This solution must be made at least once per week and the vial shall 

be stored in accordance with applicable laboratory safety procedures (e.g., kept in a vented chemical 

cabinet rated for flammable materials).

B2. Turn on the drying oven or warming plate (vented or located in a laboratory fume hood) and allow to 

come to working temperature. Place a cut filter wedge on a glass slide using forceps. Using 20 ± 5 μL 

DMF solution, place several drops along the edge of the wedge and allow the solution to wick onto 

the filter to avoid washing fibers off the filter. Warm the slide at 60 (+2/-5) °C for 30 minutes in the 

drying oven or on the warming plate.

C. TRIACETIN MOUNTS AFTER CLEARING AND COLLAPSE

C1. Using a 5-μL micropipette or microsyringe, immediately place 3.0 to 3.5 μL triacetin on the wedge. 

Gently lower a clean cover slip onto the wedge at a slight angle to reduce bubble formation. Avoid 

excess pressure and movement of the cover glass. Do not wait longer than 30 seconds before applying 

the coverslip.

NOTE: If too many bubbles form or the amount of triacetin is insufficient, the cover slip may become 

detached within a few hours. If excessive triacetin remains at the edge of the filter under the 

cover slip, fiber migration may occur [4].

C2. Mark the outline of the filter segment just inside the edge of the segment with a glass marking pen 

(such as a permanent ink marker) to aid in microscopic evaluation and to ensure the edge is avoided in 

the examination. Markings on the bottom of the slide are visible, but outside the depth of focus of 

fibers and this must be accounted for when positioning the microscope for counting. However, 

marking the bottom avoids any pressure on the coverslip. If marking the coverslip is preferred, it must 

be done very lightly and carefully.

C3. Glue the edges of the cover slip to the slide using lacquer or nail polish [18]. Counting may proceed 

immediately after clearing and mounting are completed.

NOTE: If clearing is slow, warm the slide on a hotplate (surface temperature 50 °C) for up to 15 min in 

order to hasten clearing. Heat carefully to prevent gas bubble formation.

D. EUPARAL MOUNTS AFTER CLEARING

D1. Add 1 drop of Euparal solution on the middle of the wedge and 1 drop to the center of a cover slip. In 

order to avoid trapping air bubbles, gently lower the cover slip onto the slide at a slight angle. Warm 

the slide at 60 (+2/-5) °C for 1 hour to polymerize the Euparal resin. 

D2. Mark the outline of the filter segment just inside the edge of the segment with a glass-marking pen 

(such as a permanent ink marker) to aid in microscopic evaluation and to ensure the edge is avoided in 

the examination. Markings on the bottom of the slide are visible, but outside the depth of focus of 

fibers and this must be accounted for when positioning the microscope for counting. However, 

marking the bottom avoids any pressure on the coverslip. If marking the coverslip is preferred, it must 

be done very lightly and carefully.

D3. It is not necessary to seal the edges of the cover slip with nail polish if Euparal is used. Counting may 

proceed immediately after clearing and mounting are completed.
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CALIBRATION AND QUALITY CONTROL:

10. Microscope adjustments. Follow the manufacturer’s instructions for centering and focusing the

objectives, condenser, stage, and lamp, if applicable. At least once daily, and preferably at regular

intervals throughout the day, use a phase telescope ocular (or Bertrand lens, for some microscopes) to

ensure that the phase rings (annular diaphragm and phase-shifting elements) are concentric. With each

microscope, keep a paper or electronic logbook in which to record the dates of microscope cleanings

and major servicing.

a. Each time a sample is examined, do the following:

i. Adjust the light source for even illumination across the field of view at the condenser iris.

Use Köhler illumination, if available. With some microscopes, the illumination may have to

be set up with bright field optics before aligning the phase contrast elements.

ii. Focus on the particulate material to be examined (do not focus on the marking line).

iii. Using the condenser focus, make sure that the field iris is in focus, centered on the sample,

and open only enough to fully illuminate the field of view.

b. Check the phase-shift detection limit of the microscope periodically for each

microscopist/microscope combination:

i. Center the certified phase-contrast test slide under the phase objective.

ii. Bring the blocks of grooved lines into focus in the graticule area.

NOTE: The slide contains seven blocks of grooves (ca. 20 grooves per block) in descending order of 

visibility. For asbestos counting, it is intended that some blocks of lines are completely visible 

and one or more are completely invisible when centered in the graticule area (blocks in 

between may be partially visible). The visibility of the blocks must match the statements in 

the accompanying certificate [19]. A microscope which fails to meet this requirement for a 

test slide has resolution either too low or too high for fiber counting. 

iii. If image quality deteriorates, clean the microscope optics. If the problem persists, consult

the microscope manufacturer.

c. Measure the projected diameter of the Walton-Beckett type graticule to assure that it is 100 μm ± 2

μm. Use the measured value to calculate the actual area of graticule. Check the measured value at

the minimum and maximum interpupillary distance of the eyepieces. If the diameter changes, then

the diameter may change any time a different microscopist uses the microscope and changes the

interpupillary distance. If service is performed on the microscope, or components of the microscope

are changed, measure the projected diameter again as changes may occur if the tube length or the

location of the graticule in the eyepiece is changed.

i. Center the stage micrometer under the 40X objective of the microscope.

ii. Bring the scale into focus in the graticule area

iii. The lines in the scale will appear wide. Using the mechanical stage of the microscope,

manipulate the slide until the graticule is at one edge of a major line of the stage

micrometer.

iv. Estimate the diameter of the graticule using edges of the lines of the stage micrometer. Do

not attempt to use the center of the micrometer lines.

v. If the diameter is greater than 102 μm or less than 98 μm, determine if the inter-pupillary

distance is correct for the analyst. If it is, then the microscope must be adjusted by

someone qualified to service the microscope, or the graticule replaced by one meeting the

projected diameter specification.

11. Determine a microscopist’s ability to observe, measure, and count fibers.

NOTE: Prior to examining samples, microscopists shall undergo training to observe, measure, and

count fibers and the results of such training shall be available for inspection. A procedure for 

such training, which allows proficiency to be demonstrated quantitatively, may be found in 

Appendix B. 

12. Document the laboratory’s precision for each microscopist for replicate fiber counts.
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a. Maintain as part of the laboratory quality assurance program a set of reference slides to be used on a

daily basis [20]. These slides shall consist of filter preparations including a range of loadings and

background dust levels from a variety of sources including both field and reference samples (e.g.,

PAT, AAR, or commercial samples). The Quality Assurance Officer shall maintain custody of the

reference slides and shall supply each microscopist with a minimum of one reference slide per

workday. Change the labels on the reference slides periodically so that the microscopist does not

become familiar with the samples.

b. From blind repeat counts on reference slides, estimate the laboratory intra- and inter-microscopist

precision. Obtain separate values of relative standard deviation (Sr) for each sample matrix analyzed

in each of the following ranges: 5 to 20 fibers in 100 graticule fields, >20 to 50 fibers in 100 graticule

fields, and >50 to 100 fibers in 100 graticule fields. Maintain control charts for each of these data

files.

c. Alternatively, a laboratory may develop an analytical uncertainty model from intra- and intermicroscopist precision measurements (but, in practice, most laboratories will choose option b).

NOTE: Certain sample matrices (e.g., asbestos cement) have been shown to give poor precision [23].

13. Prepare and count field blanks along with the field samples. Report counts on each field blank.

NOTE 1: The identity of blank filters shall be unknown to the microscopist until all counts have been

completed.

NOTE 2: If a field blank yields greater than 7 fibers per 100 graticule fields, report possible 

contamination of the samples.

14. Perform blind recounts by the same microscopist on 10% of filters counted (slides relabeled by a

person other than the microscopist). Use the following test to determine whether a pair of counts by

the same microscopist on the same filter shall be rejected because of possible bias: Discard the data if

the absolute value of the difference between the square roots of the two counts (in fiber/mm2) exceeds

2.77XXSSrr

′ where XX is the average of the square roots of the two fiber counts (in fiber/mm2) and SSrr′ =

(SSrr/2) where Sr is the intra-microscopist relative standard deviation for the appropriate count range (in

fibers) determined in step 12. For more complete discussions see reference [20].

NOTE 1: Fiber counting as the measurement of randomly placed fibers may be described by a Poisson

distribution, so that a square root transformation of the fiber count data will result in 

approximately normally distributed data [20].

NOTE 2: If a pair of counts is rejected by this test, recount all other samples in the set and test the new 

counts against the first counts. Discard all rejected paired counts. It is not necessary to use 

this statistic on blank counts.

NOTE 3: Do not use an intra-microscopist variation calculated from standard relocatable test slides 

(Appendix B) for this test.

15. The microscopist is a critical part of this analytical procedure. Care must be taken to provide a nonstressful and comfortable environment for fiber counting. Use an ergonomically designed chair, with

the microscope eyepiece situated at a comfortable height for viewing. Set external lighting at a similar

level to the illumination level in the microscope to reduce eye fatigue. In addition, microscopists shall

take 10- to 20-minute breaks from the microscope every one or two hours to limit fatigue [21]. During

these breaks, eye and upper back, and neck exercises can be performed to relieve strain.

16. All laboratories engaged in asbestos counting shall participate in a proficiency testing program such as

the AIHA Industrial Hygiene Proficiency Analytical Testing (IHPAT) Program for asbestos.

17. Each laboratory analyzing asbestos samples shall implement an interlaboratory quality assurance

program that as a minimum includes participation of at least two other independent laboratories. Each

laboratory shall participate in round robin testing at least once every 6 months with at least all the

other laboratories in its interlaboratory quality assurance group. Each laboratory shall submit slides

typical of its own work load for use in this program. The round robin shall be designed and results

analyzed using appropriate statistical methodology.
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MEASUREMENT:

18. Center the slide on the stage of the calibrated microscope under the objective lens. Focus the

microscope on the sample-containing plane of the filter.

19. Adjust the microscope (Step 10).

20. Counting rules: (same as P&CAM 239 rules [1,17,22]: see examples in APPENDIX C).

a. Count any fiber longer than 5 μm which lies entirely within the graticule area.

i. Count only fibers longer than 5 μm.

ii. Measure length of curved fibers along the curve. Count only fibers with a length-to-width

ratio equal to or greater than 3:1.

b. For fibers which cross the boundary of the graticule field:

i. Count as 1/2 fiber any fiber with only one end lying within the graticule area, provided that

the fiber meets the criteria of rule “a” above.

ii. Do not count any fiber which crosses the graticule boundary more than once.

iii. Reject and do not count all other fibers.

c. Count bundles of fibers as one fiber unless individual fibers can be identified by observing both ends

of a fiber.

d. Count enough graticule fields to yield 100 fibers. Count a minimum of 20 fields. Stop at 100 graticule

fields regardless of count.

21. Start counting from the tip of the filter wedge and progress along a radial line to the outer edge. Shift

up or down on the filter, and continue in the reverse direction. Select graticule fields randomly. Ensure

that, as a minimum, each analysis covers one radial line from the filter center to the outer edge of the

filter. When an agglomerate or bubble covers ca. 1/8 or more of the graticule field, reject the graticule

field and select another. [35] Do not report rejected graticule fields in the total number counted.

NOTE 1: When counting a graticule field, continuously scan a range of focal planes by moving the fine

focus knob to detect very fine fibers which have become embedded in the filter. The smalldiameter fibers will be very faint but are an important contribution to the total count. A 

minimum counting time of 15 s per field is appropriate for accurate counting.

NOTE 2: Do not count at the outside edge of the filter, or at edges where the filter was cut, as the fiber 

distribution may be disturbed in these areas. Move in at least 1 mm from the edge (i.e., inside 

the marked line). 

NOTE 3: Under certain conditions, electrostatic charge may affect the sampling of fibers. These 

electrostatic effects are most likely to occur when the relative humidity is low (below 20%), 

and when sampling is performed near the source of aerosol. The result is that deposition of 

fibers on the filter is reduced, especially near the edge of the filter. If such a pattern is noted 

during fiber counting, choose fields as close to the center of the filter as possible [11]. 

NOTE 4: Counts are to be recorded on a paper or electronic data sheet that provides, as a minimum, 

record of the counts for each field, filter identification number, analyst’s name, date, total 

fibers counted, total fields counted, mean count, fiber density, and commentary. The average 

count is calculated by dividing the total fiber count by the number of fields observed. Fiber 

density (fibers/mm2) is defined as the average count (fibers/field) divided by the field 

(graticule) area (mm2/field).

CALCULATIONS AND REPORTING OF RESULTS: 

22. Calculate and report fiber density on the filter, E (fibers/mm2

), by dividing the mean fiber count per

graticule field, F/nf, minus the mean field blank count per graticule field, B/nb, by the graticule field area,

Af:

EE =

� FF

nnff

� − � BB

nnbb

�

AAff

ffffffffffff/mmmm2 ,
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NOTE 1: Af=0.00785 mm2 is the area for a graticule with a projected diameter of 100 μm. The actual 

area shall be calculated using the measured diameter of the graticule. The measured 

diameter shall be 100 ±2 μm at the proper magnification. The diameter of the graticule shall 

be re-measured whenever the microscope is serviced or if necessary if there is a change in 

measured size with inter-pupillary distance (e.g., between different microscopists).

NOTE 2: Fiber counts above 1300 fibers/mm2 and fiber counts from samples with >50% of filter area 

covered with particulate shall be reported as “uncountable” or “probably biased.” Fiber 

counts outside the 100–1300 fiber/mm2 range shall be reported as having “greater than 

optimal variability” and as being “probably biased.”

23. Calculate and report the concentration, (C) in fibers/cc, of fibers in the air volume sampled, (V) in liters,

using the effective collection area of the filter, Ac (nominally 385 mm2 for a 25-mm filter):

CC = EEAAcc

VV ∗ 103

NOTE: The area varies according to the cassette inner diameter, where it contacts the filter, which 

could vary between manufacturers and even between production runs. For most accurate 

results the value of Ac may be checked and adjusted accordingly, but, in practice, the variation 

in this parameter is very small in comparison to other sources of uncertainty.

24. Report intra- and interlaboratory relative standard deviations with each set of results.

NOTE: Precision depends on the total number of fibers counted [1,24]. Relative standard deviation is

documented in references [1,24-25] for fiber counts up to 100 fibers in 100 graticule fields. 

Comparability of interlaboratory results is discussed below. As a first approximation, use 213% 

above and 49% below the count as the upper and lower confidence limits for fiber counts 

greater than 20 (Figure 1).

EVALUATION OF METHOD:

Summary of Method Revisions:

Each revision or release clarified minor points of grammar and procedure as well as major changes. This 

summary lists the major changes only.

The initial US Public Health Service/NIOSH method for counting fibers was published in 1968 [26]. It was 

based upon the British Hygiene standard [27] and procedures developed by the British Asbestos Research 

Council. The procedure was published in the NIOSH Criteria for a Recommended Standard [28] Section VIII, 

Appendix I (1972).

An informal draft version was used until 1976 when NIOSH issued it as P & CAM 239. It was published in the 

NIOSH Manual of Analytical Methods, 2nd ed., Vol. 1, P&CAM 239, U.S. Department of Health, Education, 

and Welfare Pub. (NIOSH) 11-157-A (1977) [17].

NIOSH Method 7400 Issued 15 February 1984

In 1984, the NIOSH P&CAM 239 Fiber counting method was renamed Method 7400 and rewritten to be in 

the standard format for inclusion in the NIOSH Manual of Analytical Methods. This revision included 

changes in three areas:

1. Sampling: The change from a 37-mm to a 25-mm filter improved sensitivity for similar air volumes. The

change in flow rates allowed for 2 m3 full-shift samples to be collected, providing that the filter is not

overloaded with non-fibrous particulates. The collection efficiency of the sampler is not a function of

flow rate in the range 0.5 to 16 L/min [17].

2. Sample preparation technique: The acetone vapor-triacetin preparation technique was included as a

faster, more permanent mounting technique than the dimethyl phthalate/diethyl oxalate method of
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P&CAM 239 [2,7,17]. The aluminum “hot block” technique minimizes the amount of acetone needed to 

prepare each sample. 

3. Measurement:

a. The Walton-Beckett graticule standardizes the area observed [21,29-30].

b. A phase shift test slide standardizes microscope optics [31].

c. Because of past inaccuracies associated with low fiber counts, the minimum recommended loading

was increased to 100 fibers/mm2 filter area (a total of 78.5 fibers counted in 100 fields, each with field

area = 0.00785 mm2). Lower levels generally result in an overestimate of the fiber count when

compared to results in the recommended analytical range [15]. The recommended loadings are

expected to yield intra-microscopist Sr in the range of 0.10 to 0.17 [32, 36].

d. An alternative set of counting rules (B rules) was introduced to improve precision of counting.

NIOSH Method 7400 Issued 15 February 1984, Revision #1 15 May 1985

Revision #1 included a discussion of interlaboratory variability and comparison of the A and B counting 

rules.

NIOSH Method 7400 Issued 02 February 1984, Revision #2 15 May 1986

Revision # 2 added an expanded discussion of interlaboratory variability and comparison of the A and B 

counting rules. It moved the “B” rules to an appendix as alternate counting rules (which should be used 

when counting man-made mineral fibers, e.g., fibrous glass, refractory ceramic fibers, etc.). It added three 

Appendices:

a. Appendix A: Calibration of the Walton Beckett Graticule

b. Appendix B: Examples of Counting Rules

c. Appendix C: Alternate Counting Rules

NIOSH Method 7400 Issued 02 February 1984, Issue 1: Revision #3 15 May 1989

Issue 1: Revision #3 corrected some technical issues and clarified the method language.

NIOSH Method 7400 Issued 02 February 1984, Issue 2: 15 August 1994

Issue 2 refined the method language, added some procedural notes and added an additional Appendix D

listing Equivalent Limits of Detection and Quantitation. 

NIOSH Method 7400 Issued 02 February 1984, Issue 3: 29 April 2019

Issue 3 of this Method incorporates:

1. More accurate values for the minimum width limits for asbestos fiber detection. A value of 0.15 μm for

chrysotile and 0.05 μm for amphibole asbestos is based on published research [7,8]. A recent user check

confirms that these values are achievable [10]. However, not all microscopists can see finer chrysotile

fibers between 0.15 μm and 0.2 μm without adequate training [33-34]. Note that PCM does not

distinguish between asbestos types.

2. DMF/acetic acid as an alternative filter clearing procedure to acetone, and Euparal as an alternative

mounting medium to triacetin. Either mounting medium is used with either clearing procedure. The

DMF/acetic acid procedure does not leave undissolved residues from the filter medium, which may

appear as fibers. Euparal produces permanent mounts regardless of the quantity of medium used [3].

These alternatives have been shown in a user-check to provide fiber counts that are not significantly

different from counts in slides made from the original dimethyl phthalate/diethyl oxalate procedure [4].

This revision also allows the continued use of dimethyl phthalate/diethyl oxalate, if preferred.
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3. A generic description of phase-shift test slides, since changes in the manufacture have led to confusion

[19]. The description now only requires the slides to have a certificate stating that there are fully visible

blocks and invisible blocks, so that the required phase-shift is bracketed. The microscopist is expected

to be able to visualize a slide according to its certificate. Newer slides have been shown in a user-check

to provide fiber counts that are not significantly different from counts in slides where the original

version of the phase shift test slide was used for calibration [10].

4. Standard relocatable test slides as an additional non-mandatory quality assurance Appendix, and as

documentation of inter- and intra-microscopist precision without the contribution from the selection of

counting fields. Several user-checks have demonstrated increases in average analyst performance after

training on these slides [33-34].

5. Guidance on the calculation of an expanded uncertainty budget. This guidance allows users to

determine expanded uncertainty as required under international standards [35].

6. Re-numbering of appendices.

INTERLABORATORY COMPARABILITY

An international collaborative study involved 16 laboratories using prepared slides from the asbestos 

cement, milling, mining, textile, and friction material industries [23]. The relative standard deviations (Sr) 

varied with sample type and laboratory. The ranges were:

Rules Intralaboratory Sr Interlaboratory Sr Overall Sr 

AIA (NIOSH A Rules)* 0.12 to 0.40 0.27 to 0.85 0.46

Modified CRS (NIOSH B 

Rules)*

0.11 to 0.29 0.20 to 0.35 0.25

*Under AIA (Asbestos International Association) rules, only fibers having a diameter less than 3 μm are

counted and fibers attached to particles larger than 3 μm are not counted. NIOSH A Rules are otherwise

similar to the AIA rules.

A NIOSH study conducted using field samples of asbestos gave intralaboratory Sr in the range 0.17 to 0.25 

and an interlaboratory Sr of 0.45 [20]. This agrees well with other recent studies [21,23-24,36].

At this time, there is no independent means for assessing the overall accuracy of this method. One 

measure of reliability is to estimate how well the count for a single sample agrees with the mean count 

from a large number of laboratories. The following discussion indicates how this estimation can be carried 

out based on measurements of the interlaboratory variability, as well as showing how the results of this 

method relate to the theoretically attainable counting precision and to measured intra- and 

interlaboratory Sr (NOTE: The following discussion does not include bias estimates and cannot be taken to 

indicate that lightly loaded samples are as accurate as properly loaded ones).

Theoretically, the process of counting randomly (Poisson) distributed fibers on a filter surface will give a Sr

that depends on the number, N, of fibers counted: SSrr = 1

√NN

Thus, Sr is 0.1 for 100 fibers and 0.32 for 10 fibers counted. The actual Sr found in a number of studies is 

greater than these theoretical numbers [15,25,30,36].

An additional component of variability comes primarily from subjective interlaboratory differences. In a 

study of ten microscopists in a continuing sample exchange program, Ogden [24] found this subjective 

component of intralaboratory Sr to be approximately 0.2 and estimated the overall Sr by the term: 

�NN+(0.2+NN)2

NN .

Ogden found that the 90% confidence interval of the individual intralaboratory counts in relation to the 

means were +2 Sr and −1.5 Sr . In this program, one sample out of ten was a quality control sample. For 
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laboratories not engaged in an intensive quality assurance program, the subjective component of 

variability can be higher.

In a study of field sample results in 46 laboratories, the Asbestos Information Association (AIA) also found 

that the variability had both a constant component and one that depended on the fiber count [18]. These 

results gave a subjective interlaboratory component of Sr (on the same basis as Ogden’s) for field samples 

of ca. 0.45. A similar value was obtained for 12 laboratories analyzing a set of 24 field samples [36]. This 

value falls slightly above the range of Sr (0.25 to 0.42 for 1984–85) found for 80 reference laboratories in the 

NIOSH PAT program for laboratory-generated samples [25].

A number of factors influence intralaboratory Sr for a given laboratory, such as that laboratory’s actual 

counting performance and the type of samples being analyzed. In the absence of other information, such 

as from an interlaboratory quality assurance program using field samples, the value for the subjective 

component of variability is chosen as 0.45. It is recommended that each laboratory establish an 

interlaboratory quality assurance program to improve their performance and thus reduce the

intralaboratory Sr. 

The above relative standard deviations apply when the population mean has been determined. It is more 

useful, however, for laboratories to estimate the 90% confidence for of the count distribution for both 

interlaboratory and intralaboratory results [24].

For example, if a sample yields a count of 24 fibers, Figure 1 indicates that the mean interlaboratory count 

will fall within the range of 227% above and 52% below that value 90% of the time. These percentages can 

be applied directly to the air concentrations as well. If, for instance, this sample (24 fibers counted) 

represented a 500-L volume, then the measured concentration is 0.02 fibers/cc (assuming 100 fields 

counted, 25-mm filter, 0.00785 mm2 counting field area). If this same sample were counted by a group of 

laboratories, there is a 90% probability that the mean would fall between 0.01 and 0.08 fiber/cc. These 

limits are to be reported in any comparison of results between laboratories.

Note that the interlaboratory Sr of 0.45 used to derive Figure 1 is used as an estimate for a random group of 

laboratories. If several laboratories belonging to a quality assurance group can show that their 

interlaboratory Sr is smaller, then it is more correct to use that smaller Sr. However, the estimated

interlaboratory Sr of 0.45 is to be used in the absence of such information. Note also that it has been found 

that interlaboratory Sr can be higher for certain types of samples, such as asbestos cement [23].

Quite often the estimated airborne concentration from an asbestos analysis is used to compare to a 

regulatory standard. For instance, if one is trying to show compliance with an 0.5 fiber/cc standard using a 

single sample on which 100 fibers have been counted, then Figure 1 indicates that the 0.5 fiber/cc 

standard must be 213% higher than the measured air concentration. This indicates that if one measures a 

fiber concentration of 0.16 fiber/cc (100 fibers counted), then the mean fiber count by a group of 

laboratories (of which the compliance laboratory might be one) has a 95% chance of being less than 0.5 

fibers/cc; i.e., 0.16 + 2.13 × 0.16 = 0.5.

It can be seen from Figure 1 that the Poisson component of the variability is not very important unless the 

number of fibers counted is small. Therefore, a further approximation is to simply use +213% and -49% as 

the upper and lower confidence values of the mean for a 100-fiber count.
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Figure 1. Interlaboratory precision of fiber counts

The curves in Figure 1 are defined by the following equations:

UUUUUU = 2XX + 2.25 + �(2.25 + 2XX)2 − 4(1 − 2.25SSrr

2)XX2

2(1 − 2.25SSrr

2)

and

2XX + 4 − �((4 + 2xx)2 − 4(1 − 4SSrr

2)XX2) UUUUUU =

2(1 − 4SSrr

2

)

where Sr = subjective interlaboratory relative standard deviation, which is close to the total interlaboratory 

Sr when approximately 100 fibers are counted, X = total fibers counted on sample, UCL = upper 95% 

confidence limit, and LCL = lower 95% confidence limit.

NOTE: The range between these two limits represents 90% of the total range. 

It is also possible to use a similar approach to determining expanded uncertainty within a single laboratory.

The formula:

SSrr = �NN + (0.2NN)2

NN

suggests that if n is the number of counts taken in a single reading, then the distribution of
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nn − NN

�nn + SSrr

2NN2

though unknown, depends only weakly on N, because the mean and variance are independent of N. In 

fact, from many readings by a number of microscopists, with N equal to the mean across the microscopists, 

confidence intervals were determined [24] as:

−1.8 ≤

nn − NN

�NN + SSrr

2NN2

≤ +2.6

By solving the two quadratic equations at the two extreme limits for N in terms of n, confidence limits 

enclosing the consensus mean N, given single measurements n are easily obtained: LLCCLL[nn] < NN < UUCCLL[nn]

(95% Confidence Limit), where LCL and UCL are given by:

LLCCLL[nn] = 2nn + 2.62 − �(2.62 + 2nn)2 − 4(1 − 2.62SSrr

2)nn2

2(1 − 2.62SSrr

2)

and

UUUUUU[nn] = 2nn + 1. 82 + �(1. 82 + 2nn)2 − 4(1 − 1. 82SSrr

2)nn2

2(1 − 1. 82SSrr

2)

NOTE: In these formulae the confidence level is expanded from 90% to 95%.

From these equations, the confidence limits accounting for both subjective microscopist and Poisson 

components for various fiber counts are calculated (Table 1) by taking Sr = 0.2. The intra- and intermicroscopist variability may be greater if quality control is poor.

Table 1. Intra-microscopist (Sr=0.2) 95% confidence interval bracketing the consensus mean for various 

numbers (n) of fibers in a single count

Number (n) for fibers Lower confidence limit 

(LCL) 

Upper confidence limit 

(UCL)

Expanded uncertainty 

(%) 

5 1.6 13 (−68;160)

7 2.6 16 (−63;129)

10 4.2 21 (−58;110)

20 10 37 (−50;85)

50 29 85 (−42;70)

100 62 163 (−38;63)

200 127 319 (−36;60)

NOTE: Also shown is the confidence interval expressed as expanded uncertainty asymmetric around n and 

stated relative to n.

UNCERTAINTY BUDGET EXCLUDING COUNTING ERRORS

Various factors aside from counting errors can introduce random variation in measured values of asbestos 

concentrations [35]. Such factors are identified in Table 2. 
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Table 2. Example of an uncertainty budget for non-counting variable

Variable Estimated uncertainty Uncertainty squared

Sampling flowrate 0.03 0.0008

Sampling time 0.02 0.0004

Master stage micrometer 0.01 0.0001

Calibration of submaster 0.01 0.0001

Calibration of graticule 0.02 0.0004

Area of exposed filter 0.05 0.0025

Sum of squares - 0.0043

Square root of sum of squares 

= Overall uncertainty

- 0.066

Expanded uncertainty 

(coverage factor k = 2)

- 0.13 or 13 %

The uncertainty in Table 2 contributes in quadrature to the expanded uncertainty. Therefore, as 13% is 

small relative to the values in Table 1, it may be reasonable to conclude that the contribution of noncounting errors to the total uncertainties is not significant in comparison to the Poisson and subjective 

errors.

NOTE 1: Non-counting errors are negligible only if consistently controlled.

NOTE 2: Inhomogeneous deposition of dust on the filter leads to gross errors, the magnitude of which 

cannot be estimated. Counting 20 or more fields ensures that minor divergence from 

randomness does not bias the result. 

REFERENCES:

[1] NIOSH [1979]. USPHS/NIOSH Membrane Filter Method for Evaluating Airborne Asbestos Fibers. By

Leidel NA, Bayer SG, Zumwalde RD, Busch KA. Cincinnati, OH: U.S. Department of Health, Education,

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, DHEW

(NIOSH) Publication No. 79-127.

[2] Baron P, Pickford PA, Pickford GC [1986]. An asbestos sample filter clearing procedure. Appl Ind Hyg

1:169–171, 199.

[3] LeGuen JMM, Galvin S [1981]. Clearing and mounting techniques for the evaluation of asbestos

fibers by the membrane filter method. Ann Occup Hyg 24:273–280.

[4] Lee EG, Pang TWS, Nelson J, Andrew M, Harper M [2011]. Comparison of mounting methods for the

evaluation of fibers by phase contrast microscopy. Ann Occup Hyg 55:644-657.

[5] OSHA [1994]. 29 CFR Part 1910.1001 Occupational Exposure to Asbestos. Washington, DC: U.S.

Department of Labor, Occupational Safety and Health Administration.

[6] McCrone W, McCrone L, Delly J [1978]. Polarized light microscopy. Ann Arbor, MI: Ann Arbor Science.

[7] Rooker SJ, Vaughn NP, LeGuen JM [1982]. On the visibility of fibers by phase contrast microscopy.

Am Ind Hyg Assoc J 43:505–515.

[8] Kenney LC, Rood AP, Blight BJN [1987]. A direct measurement of the visibility of amosite asbestos

fibres by phase contrast optical microscope. Ann Occup Hyg 33:261–264.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[9] Harper M, Lee EG, Doorn SS, Hammond O [2008]. Differentiating non-asbestiform amphibole and

amphibole asbestos by size characteristics. J Occup Envir Hyg 5:761–770.

[10] Lee EG, Nelson J, Kashon M, Harper M [2015]. Evaluation of the dark-medium objective lens in

counting asbestos fibers by phase-contrast microscopy. Ann Occup Hyg 59:616–628.

[11] Baron P, Deye G [1990]. Electrostatic effects in asbestos sampling: parts I and II. Am Ind Hyg Assoc J

51:51–69.

[12] ISO [2013]. ISO 13137-13 Workplace atmospheres -- Pumps for personal sampling of chemical and

biological agents -- Requirements and test methods. Geneva, Switzerland: International Organization

for Standardisation.

[13] Johnston AM, Jones AD, Vincent JH [1982]. The influence of external aerodynamic factors on the

measurement of the airborne concentration of asbestos fibers by the membrane filter method. Ann

Occup Hyg 25:309–316.

[14] Beckett ST [1980]. The effects of sampling practice on the measured concentration of airborne

asbestos. Ann Occup Hyg 23:259–272.

[15] Cherrie J, Jones AD, Johnston AM [1986]. The influence of fiber density on the assessment of fiber

concentration using the membrane filter method. Am Ind Hyg Assoc J 47(8):465–474.

[16] Lee EG, Nelson J, Hintz PJ, Joy G, Andrew ME, Harper M [2010]. Field performance of the CATHIA-T

sampler and two cyclones against the standard cowled sampler for thoracic fiber concentrations.

Ann Occup Hyg 54:545-556.

[17] NIOSH [1977]. P&CAM 239. In: Taylor DG, ed. NIOSH manual of analytical methods, 2nd ed. Cincinnati,

OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, National Institute

for Occupational Safety and Health, DHEW(NIOSH) Publication No. 77-157-A.

[18] AIA [1979]. AIA Health and Safety Recommended Technical Method #1: airborne asbestos fiber

concentrations at workplaces by light microscopy. London, England: Asbestos International

Association.

[19] Crane D, Harper M [2011]. The phase-shift test slide.

https://www2.aiha.org/Registries/aar/Documents/Phase%20Shift%20Interpretation%20OSHA%

20NIOSH_2011_04_11.pdf .

[20] Abell MS, Shulman SA, Baron P [1989]. The quality of fiber count data. Appl Ind Hyg 4:273–285.

[21] AIA [1983]. A study of the empirical precision of airborne asbestos concentration measurements in

the workplace by the membrane filter method. Arlington, VA: Asbestos Information Association, Air

Monitoring Committee.

[22] NIOSH [1976]. Revised Recommended Asbestos Standard. Cincinnati, OH: U.S. Department of Health,

Education, and Welfare, Center for Disease Control, National Institute for Occupational Safety and

Health DHEW (NIOSH) Publication No. 77-169.

[23] IOM [1982]. A comparison of the effects of different counting rules and aspect ratios on the level and

reproducibility of asbestos fiber counts. Part I: effects on level and Part II: effects on reproducibility.

By Crawford NP, Thorpe HL, Alexander W. Edinburgh, Scotland: Institute of Occupational Medicine.

[24] HSE [1982]. Health and Safety Executive research paper 18: the reproducibility of fiber counts. By

Ogden TL. Sheffield, England: Health and Safety Laboratory, Health and Safety Executive.
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 18 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[25] Schlecht PC, Schulman SA [1986]. Performance of asbestos fiber counting laboratories in the NIOSH

Proficiency Analytical Testing (PAT) Program. Am Ind Hyg Assoc J 47:259–266.

[26] Edwards GH, Lynch JR [1968]. The method used by the Public Health Service, for enumeration of

asbestos dust on membrane filters. Ann Occup Hyg 11:1-6.

[27] Lane RE, Barnes JM, Roach SA, Smith S, Addingley CG, Holmes S, Hunt R, Knox JF, King E [1968].

Hygiene standard for chrysotile asbestos dust. Ann Occup Hyg 11:47 -49.

[28] NIOSH [1972]. NIOSH criteria for a recommended standard: occupational exposure to asbestos.

Cincinnati, OH: U.S. Department of Health Education and Welfare, Center for Disease Control,

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 72-10267.

[29] Royal Commission on Matters of Health and Safety Arising from the Use of Asbestos in Ontario.

Measurement of asbestos fiber concentrations in workplace atmospheres: Study No. 9. By Chatfield,

EJ. Toronto, Ontario.

[30] Walton WH [1982]. The nature, hazards, and assessment of occupational exposure to airborne

asbestos dust: a review. Ann Occup Hyg 25:115–247.

[31] LeGuen JMM, Ogden TL, Shenton-Taylor T [1984]. The phase-shift test slide. Ann Occup Hyg 28:237-

247.

[32] Taylor DG, Baron PA, Shulman SA, Carter JW [1984]. Identification and counting of asbestos fibers.

Am Ind Hyg Assoc J 45:84–88.

[33] Pang TWS, Harper M [2008]. The quality of fiber counts using improved slides with relocatable fields.

J Environ Monit 10:89-95.

[34] Harper M, Slaven JE, Pang TWS [2009]. Continued participation in an asbestos fiber counting

proficiency test with relocatable grid slides. J Environ Monit 11:434-438.

[35] ISO [2014]. ISO 8672-14[E] Air quality — determination of the number concentration of airborne

inorganic fibres by phase contrast optical microscopy — membrane filter method. Geneva,

Switzerland: International Organization for Standardisation.

[36] Baron PA, Shulman SA [1987]. Evaluation of the Magiscan image analyzer for asbestos fiber counting.

Am Ind Hyg Assoc J 48:39-46.

[37] Pang TWS [2000]. Precision and accuracy of asbestos fiber counting by phase contrast microscopy.

Am Ind Hyg Assoc J 61:529–538.

[38] Harper M, Bartolucci A [2003]. Preparation and examination of proposed consensus reference

standards for fiber-counting. Am Ind Hyg Assoc J 64:283-287.

[39] WHO [1985]. Reference methods for measuring airborne man-made mineral fibers (MMMF). By: The

WHO/EURO Technical Committee for Monitoring an Evaluating Airborne MMMF. Copenhagen,

Denmark: World Health Organization.

[40] NIOSH [1977]. NIOSH criteria for a recommended standard: occupational exposure to fibrous glass.

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control,

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-152.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD WRITTEN BY:

Paul A. Baron, PhD, NIOSH; Revised by Martin Harper, PhD, CIH, FAIHA, CChem, FRSC, NIOSH (retired) and 

Rosa Key-Schwartz, PhD, NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by the National 

Institute for Occupational Safety and Health, Centers for Disease Control and Prevention. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.

Table 3. CAS numbers for analytes

Name CAS No.

Actinolite [77536-66-4]

Ferroactinolite [15669-07-5]

Amosite [12172-73-5]

Anthophyllite [77536-67-5]

Chrysotile [12001-29-5]

Crocidolite [12001-28-4]

Tremolite [77536-68-6]

Amphibole asbestos [1332-21-4]

Refractory ceramic fibers [142844-00-6]

Fibrous glass [14808-60-7] 

APPENDIX A: CALIBRATION OF THE GRATICULE

Before ordering any graticule, the following calibration must be done to obtain a counting area (D) 100 μm 

in diameter at the image plane. The diameter, dc (mm), of the circular counting area and the disc diameter 

must be specified when ordering the graticule. 

1. Insert any available graticule into the eyepiece and focus so that the graticule lines are sharp and clear.

2. Set the appropriate inter-pupillary distance and, if applicable, reset the binocular head adjustment so

that the magnification remains constant.

3. Install the 40x to 45x phase objective.

4. Place a stage micrometer on the microscope object stage and focus the microscope on the graduated

lines.

5. Measure the magnified grid length of the graticule, Lo (μm), using the stage micrometer.

6. Remove the graticule from the microscope and measure its actual grid length, La (mm). This can best

be accomplished by using a stage fitted with verniers.

7. Calculate the circle diameter, dc (mm), for the Walton-Beckett graticule: ddcc = LLaa

LLoo

× DD

EXAMPLE: if Lo = 112 μm, La = 4.5 mm, and D = 100 μm, then dc = 4.02 mm.

8. Check the field diameter, D (acceptable range 100 μm ± 2 μm) with a stage micrometer upon receipt of

the graticule from the manufacturer. Determine field area (acceptable range 0.00754 mm2 to 0.00817

mm2).

APPENDIX B: DETERMINATION OF A MICROSCOPIST’S ABILITY TO OBSERVE, MEASURE AND COUNT 

FIBERS USING RELOCATABLE GRIDDED COVERSLIPS (Non-mandatory)

1. Reference slides are commercially available, made from chrysotile or amosite proficiency test filters

obtained from the Industrial Hygiene Proficiency Analytical Testing (IHPAT) scheme of the American

Industrial Hygiene Association Proficiency Analytical Testing, LLC [37]. Each slide has been mounted
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with a permanent Euparal medium and covered with a relocatable gridded cover slip. The fibers visible 

in each grid opening have been identified and their locations marked on a drawing of each opening. 

The identity, number and position of each fiber have been verified by a second microscopist. The 

accuracy of this verification has been shown to be within 3% of consensus values determined by 

multiple microscopists, separately and together [38]. Such slides are referred to in this method as 

standard relocatable test slides, and they are applied in various ways to improve and assess the quality 

of fiber counts.

NOTE: It is also possible to make standard relocatable test slides in the laboratory from PAT or field 

samples by using commercially available relocatable gridded cover slips. Either filter clearance 

or slide mounting procedure can be used with these cover-slips, but the DMF/acetic acidEuparal procedure has been demonstrated to provide slides with long-term stability. The fibers 

in each grid are identified, marked and verified by at least two microscopists as above.

2. Count the fibers in each designated field and refer the counts to the accompanying slide descriptions.

Calculate a score from the number of absolute discrepancies between the reported and verified fibers

in each field as shown in:

SSSSSS = (1 − � nnnn dd dddd

nnnn vv ffffffffffff�) × 100

3. Microscopists shall obtain a discrepancy score of more than 70 for a slide containing amosite fibers

and more than 50 for a slide containing chrysotile fibers from the AIHA IHPAT scheme (or more than 70

for a slide prepared from chrysotile field samples) before proceeding. These scoring criteria are based

on the results that have been achieved in round-robin studies and proficiency test programs. If these

scores are not achieved, the Quality Assurance Officer is expected to work with the microscopist to

review the slide descriptions to determine the cause and then attempt to rectify the situation by

having the microscopist repeat the microscope set-up or by re-training.

NOTE: The number of absolute discrepancies has been shown to be linearly related to the sum of

counting errors [37]. The positive discrepancies are mainly due to sizing extra fibers (especially 

amphiboles). The negative discrepancies are mainly due to oversight of fibers (especially 

chrysotile). Chrysotile filters from the AIHA IHPAT program contain a large number of fibers 

with narrow widths near the limit of detection and thus provide a very robust test of visual 

acuity and concentration. In theory, it should be possible for microscopists to obtain high 

discrepancy scores using slides made from chrysotile PAT samples, but a discrepancy score 

above 50 is sufficient to ensure the microscopist will be able to see the majority of the wider 

fibers typical of field samples.

4. The repeated fiber counts obtained from standard relocatable test slides can be used to document the

laboratory’s precision for each microscopist, per step 13. If repeated counts from specific fields are

used, the resulting intra- and inter-microscopist precision will be improved because there will not be a

contribution from the selection of field areas to be counted.

APPENDIX C: COMPARISON OF COUNTING RULES

Figure 2 shows a Walton-Beckett graticule as seen through the microscope. This is a Type G22 graticule 

with 3:1 ratios, providing optimal assistance for these counting rules. The rules will be discussed as they

apply to the labeled objects in the figure.
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Figure 2 Walton-Beckett graticule with fibers
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These rules are sometimes referred to as the “A” rules:

Object Count Discussion

1 1 fiber Optically observable asbestos 

fibers are actually bundles of fine 

fibrils. If the fibrils seem to be 

from the same bundle, the object 

is counted as a single fiber. Note, 

however, that all objects meeting 

length and aspect ratio criteria 

are counted whether or not they 

appear to be asbestos.

2 2 fibers If fibers meeting the length and 

aspect ratio criteria (length >5 

μm and length-to-width ratio > 3 

to 1) overlap, but do not seem to 

be part of the same bundle, they 

are counted as separate fibers.

3 1 fiber Although the object has a 

relatively large diameter (>3 μm), 

it is counted as a fiber under the 

rules. There is no upper limit on 

the fiber diameter in the 

counting rules. Note that fiber 

width is measured at the widest 

compact section of the object.

4 1 fiber Although long fine fibrils may 

extend from the body of a fiber, 

these fibrils are considered part 

of the fiber if they seem to have 

originally been part of the 

bundle.

5 Do not count If the object is ≤ 5 μm long, it is 

not counted

6 1 fiber A fiber partially obscured by a 

particle is counted as one fiber. If 

the fiber ends emanating from a 

particle do not seem to be from 

the same fiber and each end 

meets the length and aspect ratio 

criteria, they are counted as 

separate fibers.

7 1⁄2 fiber A fiber which crosses into the 

graticule area one time is 

counted as 1⁄2 fiber.

8 Do not count Ignore fibers that cross the 

graticule boundary more than 

once.

9 Do not count Ignore fibers that lie outside the 

graticule boundary.
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NOTE 1: OSHA Method ID-160 has slightly different counting rules, e.g. Rule 5: If the object is ≤ 5 μm, it 

is not counted, would become: If the object is < 5 μm, it is not counted. In practice, these 

minor differences would not likely modify calculated concentrations to the extent that 

decisions based on those concentrations would be affected, but this opinion has not been 

tested. 

NOTE 2: Fibers wider than 3 μm are included in this method. Some other methods for counting 

asbestos fibers restrict the count to only those fibers of 3 μm width or less, as these are the 

fibers most likely to penetrate to the bronchial and lower airways. However, the inlet efficiency 

of the cowl has a similar effect in that it limits the maximum diameter of fibers entering the 

cassette, so the effect of this additional dimensional counting rule on the total fiber count is 

expected to be small. Any bias from not applying an upper width limit is also conservative (i.e. 

concentrations will be assessed higher).

APPENDIX D: ALTERNATE COUNTING RULES FOR NON-ASBESTOS FIBERS

Other counting rules may be more appropriate for measurement of specific non-asbestos fiber types, such 

as fibrous glass. These include the rules given below (“B” rules from NIOSH Method 7400, Revision #2, dated 

08/15/1987), the World Health Organization reference method for man-made mineral fiber [39], and the 

NIOSH fibrous glass criteria document method [40]. The upper diameter limit in these methods prevents 

measurements of non-thoracic fibers. It is important to note that the aspect ratio limits included in these 

methods vary. Graticules designed for counting according to these rules are available (e.g. Walton-Beckett 

Type G24).

OSHA recommends using these alternate, or “B” counting rules as given below for all man-made vitreous 

fibers (MMVF). It is emphasized that hybridization of different sets of counting rules is not permitted. 

Report specifically which set of counting rules are used with the analytical results.

“B” Counting Rules

1. Count only ends of fibers. Each fiber must be longer than 5 μm and less than 3 μm diameter.

2. Count only ends of fibers with a length-to-width ratio equal to or greater than 5:1.

3. Count each fiber end which falls within the graticule area as one end, provided that the fiber meets

rules 1 and 2 above. Add split ends to the count as appropriate if the split fiber segment also meets the

criteria of rules 1 and 2 above.

4. Count visibly free ends which meet rules 1 and 2 above when the fiber appears to be attached to

another particle, regardless of the size of the other particle. Count the end of a fiber obscured by

another particle if the particle covering the fiber end is less than 3 μm in diameter.

5. Count free ends of fibers emanating from large clumps and bundles up to a maximum of 10 ends (5

fibers), provided that each segment meets rules 1 and 2 above.

6. Count enough graticule fields to yield 200 ends. Count a minimum of 20 graticule fields. Stop at 100

graticule fields, regardless of count.

7. Divide total end count by 2 to yield fiber count.
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APPENDIX E: EQUIVALENT LIMITS OF DETECTION AND QUANTITATION

Fiber density on filter* 

(Fibers per 100 fields)

Fiber density on filter* 

(Fibers /mm2

) 

Fiber concentration in 

air, f/cc (400-L air 

sample)

Fiber concentration in 

air, f/cc (1000-L air 

sample)

200 255 0.25 0.10

100 127 0.125 0.05

LOQ 80 102 0.10 0.04

50 64 0.0625 0.025

25 32 0.03 0.0125

20 25 0.025 0.010

10 12.7 0.0125 0.005

8 10.2 0.010 0.004

LOD 5.5 7 0.00675 0.0027

*Assumes 385 mm2 effective filter collection area, and field area = 0.00785 mm2, for relatively “clean” (little

particulate aside from fibers) filters

APPENDIX F: P&CAM 239 ASBESTOS FIBER IN AIR [17]

Analyte: Asbestos fibers

Matrix: Air

Procedure: Filter collection, microscopic 

count 

Date Issued: 3/30/77

Method No.: P&CAM 239

Range: 0.1-60 fibers/cm3

Precision (CVT): 0.24 to 0.38

Classification: D (Operational)

1. Principle of the Method

1.1. This method describes the equipment and procedures for collecting, mounting, and counting

asbestos fibers on cellulose ester membrane filters in the evaluation of personal samples of airborne 

asbestos fibers. The purpose of the method is to determine an employee's index of exposure to 

airborne asbestos fibers. The method is primarily a personal monitoring technique, but can be used 

for area monitoring.

1.2. The sample is collected by drawing air through a membrane filter by means of a battery powered 

personal sampling pump. The filter is transformed from an opaque solid membrane to a 

transparent optically homogeneous gel. The fibers are sized and counted using a phase-contrast

microscope at 400-450X magnification.

1.3. Definitions. Asbestos fiber, for counting purposes, means a particulate which has a physical 

dimension longer than 5 micrometers and with a length to diameter ratio of 3 to 1 or greater. 

Asbestos includes chrysotile, cummingtonite-grunerite (amosite), crocidolite, fibrous tremolite,

fibrous anthophyllite, and fibrous actinolite.

1.4. Any laboratory attempting to use this procedure should have at least one counter attend a training 

course conducted by an experienced, proficient laboratory. Novice, untutored counters, using only 

published instructions, can easily obtain counts of half those performed by experienced, proficient 

counters. Large differences between laboratories can be caused by: 1) differences in technique and 

observing ability among counters and 2) small, but significant, differences between microscopes 

meeting the basic specifications of Section 6.2. The following procedures are recommended:
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1.4.1.All microscopists who perform asbestos counting should meet together for an "asbestos 

counting workshop" at least quarterly.· This is best accomplished with counters from several 

laboratories using their own microscopes.

1.4.2. Each microscopist should count the same series of slides and with the results being compared.

1.4.3.Differences between counters should be resolved with side-by-side counting of the fields by 

the different counters.

1.4.4. Individuals who are found to be persistent outliers over several sessions should be encouraged 

to seek other tasks in their respective laboratories.

2. Range and Sensitivity

2.1. The usable range is primarily a function of sample volume, microscope count field area, and

background airborne particulates. The influence of these variables is discussed in 8.1.3. For a 

microscope count field area of 0.003 mm2 (see Figure 1) and a pump flow rate of 1.7 lpm, the 

optimal fiber densities would be produced over the range of 0.4 fiber/cm3 (8-hour sample) to about 

60 fibers/cm3 (15-minute sample). For a field area of 0.006 mm2 (see Figure 2) and a pump flow rate 

of 1.7 lpm, the optimal range is 0.2 fiber/cm3 (8-hour sample) to about 30 fibers/cm3 (15-minute 

sample). In each case, the optimal detection limits are inversely proportional to pump flow rate.

The upper detection limit can be extended by using sample times less than 15 minutes or using

lower flow rates. The lower detection limit can be extended by increasing the flow rate up to about 

2.5 lpm. Filter surface fiber densities less than optimal (less than about 0.5 to 1.0 fiber per count 

field) are still adequate, but will lead to decreased precision for the method (increased coefficient of 

variation, see Section 4). The minimum total fiber count in 100 fields considered adequate for 

reliable quantitation is 10 fibers. Thus, the lower limit of reliable quantitation is 0.1 fiber/cm3

(100,000 fibers/m3

). For this level, a flow rate of about 2.5 lpm is recommended. For a field area of

0.003 mm2 , the minimum sample time would be about 2 hours. For a field area of 0.006 mm2

, the 

minimum sample time would be about 1 hour.

2.2. This method considers only fibers with a length to diameter ratio of 3 to 1 or greater and a length 

greater than 5 micrometers.

3. Interferences

In an atmosphere known to contain asbestos, all particulates with a length to diameter ratio of 3 to 

1 or greater, and a length greater than 5 micrometers should, in the absence of other information,

be considered to be asbestos fibers and counted as such.

4. Precision and Accuracy

4.1. In the past decade, there have appeared a number of articles examining sources of variation in the

asbestos sampling and counting procedure. These include: Lynch et al. (11.1), Weidner and Ayer 

(11.2), Conway and Holland (11.3), Leidel and Busch (11.4), Beckett and Attfield (11.5), and Rajhans 

and Bragg (11.6). The sources of variation will be discussed by stages in the membrane filter 

evaluation procedure.

4.2. Sources of Variation in the Sampling Process. These include variations in pump flow rate, 

proximity of the filter to the employee's body, and filter location (left to right) in the employee's

breathing zone.

4.2.1. Section 9.1 requires that the personal sampling pump be calibrated with sufficient accuracy 

such that the 95% confidence limits on the flow rate are ±10%. This is equivalent to a 

coefficient of variation (CV) of about 5%. However, this CV makes a negligible contribution to 

the total CV for the method due to the relatively large CV of the counting procedure.

4.2.2.Conway and Holland (11.3) concluded that positioning of the filter cassette on the wearer 

(regarding the angular portions of the filter and their proximity to the wearer) is not a 

significant factor in determining the fiber distribution on filters.

4.2.3.Weidner and Ayer (11.2) concluded that there is no appreciable difference between samples 

collected on either the right or left sides of a breathing zone or between samples collected 

side-by-side, especially for samples with concentrations less than 2.5 fibers/cm3

. 
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4.3. Sources of Variation in the Counting Procedure

4.3.1.Random variations exist in the fiber distribution on a filter wedge (intra-wedge variability). The 

industrial hygiene literature has seen considerable debate in the last 20 years concerning 

whether or not the distribution of mineral dust or asbestos fibers on a filter surface is 

adequately described by a Poisson distribution probability density function. Leidel and Busch 

(11.4) found excellent agreement between empirical error variance and theoretical variance 

calculated from the assumption of Poisson distributed true counts. They concluded that there 

was not excessive variation among count fields for a filter wedge and that clumping of fibers 

(non-random coalescence) did not occur.

4.3.2.Variations exist in the fiber distribution on the total filter surface (inter-wedge variability) due 

to the random or non-random distribution of fibers across the total surface of the filter. This 

type of variation is easily confused with intra-wedge variations. The count procedure does not 

require counting of multiple sectors of the filter. There may be significant differences between 

average counts for different wedges, or the fiber distribution variations for the total filter 

surface may be greater than the variations of the Poisson distribution. If either of these occur 

experimentally, one must use the experimental variations to estimate the minimum precision 

of the count procedure. The minimum precision is governed by the variations of the fiber 

distribution on the total surface of the filter.

Conway and Holhind (11.3) concluded the distribution of fibers on filters is not uniform and, 

the distribution of fiber counts is more disperse than Poisson. For their filters which had 

significant variations in fiber concentrations between sectors (as much as 50-60% of the total 

filter mean), they described the following relation for the standard deviation of the total 

number of fibers counted on a wedge (N)

ee (NN) = 1.6(NN)1/2

where N is about 100. The Poisson standard deviation would be:

PP σσ (NN) = (NN)1/2

Rajhans and Bragg (11.6) in Series I of their study found significant variation between filter 

segments and rejected the Poisson distribution for the total filter surface. However, in Series II 

of their study, utilizing various experimental modifications, they found no significant variation 

between filter segments and no reason to reject the assumption of Poisson distributed fiber 

counts.

4.3.3. Systematic variations due to differences between microscopes were studied by Leidel and 

Busch (11.4). In their study using five different brands of microscopes, they found no significant 

differences among four, but the fifth gave counts approximately 45% higher on the average 

than the other four.

4.3.4.Variations due to differences between counters should be examined at three levels:

experienced counters occasionally counting, experienced counters routinely counting, and 

inexperienced (new or untutored) counters. Leidel and Busch (11.4) studied five experienced 

counters, with one counting only occasionally. There were no significant differences among 

three of the counters, but a fourth was 16% lower than the first three. The fifth, who 

occasionally counted, averaged 27% higher than the first three. Conway and Holland (11.3) 

studied three experienced counters and three inexperienced counters. They found statistically 

significant differences between the means of both the experienced and inexperienced 

counters that typically were in the range plus or minus 5 to 15%. They concluded that 

experience as a fiber counter is not a significant parameter affecting intercounter variations.

Rajhans and Bragg (11.6) found no significant differences among means of five experienced

counters in Series I of their study. But in their carefully controlled Series II, an analysis of 

variance showed significant variations between counters that were plus or minus 1 to 15%.

4.3.5.Variations between laboratories are most likely due to systematic biases and are not a 

significant additional source of random variations. Any additional variations are most likely due 

to differences in counting technique. Beckett and Attfield (11.5) observed that standard 
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counters improved greatly after personal instruction; also new counters, after instruction, 

tended to overcompensate and get exceedingly high counts. Additionally, they found that 

counts from an experienced laboratory that had not had contact with other laboratories 

performing the same analysis were as far from the standard values as were the counts by new 

counters.

4.4. Sources of variations between samples taken at different times on one employee during one work 

shift can affect the exposure estimate for that employee. These are primarily due to a) differences in 

exposure concentrations during the day, b) differences in location of the employee within the plant, 

and c) differences in work operation performed by the employee during the day. These sources of 

variation can be controlled by proper choice of sampling strategy. Refer to Leidel and Busch (11.7) 

and Leidel, Busch, and Lynch (11.8) for an extended discussion of sampling strategies. Interday 

temporal variations can affect the exposure estimates obtained on different days. Refer to Leidel, 

Busch, and Crouse (11.9) for a discussion of this type ofvariation.

4.5. Until recently, the total coefficient of variation (CVT) for the sampling and counting procedure was 

best estimated from the work of Conway and Holland (11.3). The conclusions of their study 

included:

4.5.1. The precision of their procedure for filters not containing an abundance of fine fibers can be 

estimated by a coefficient of variation of 16.2%. This value includes variation among counters 

and observed interactioneffects.

4.5.2. The accuracy of the procedure for similar filters may be estimated for a 100-fiber count by a 

coefficient of variation of 21.4%. This assumes that the contribution of the overall variance 

from the nonuniform fiber distribution isadditive.

4.5.3.A high percentage of very fine fibers on the filter can significantly affect the standard deviation 

and confidence limits for counts by different counters. After combining variations in fiber 

concentrations over the entire filter with those for different counters, it was concluded:

1. For filters with a low concentration of fine fibers, the coefficient of variation is estimated at

21% and the 95% confidence interval is±43%.

2. For filters with a high concentration of fine fibers, the coefficient of variation is estimated

at 25% and the 95% confidence interval is±50%.

Lynch, Kronoveter, and Leidel (11.1) have also reported on variations of the method. Their 

intralaboratory study utilized the data from a large number of dust counts made by different

methods by experienced counters over a period of years in an epidemiologic study of the asbestos 

products industry. They concluded that the standard deviation of counts of fibers longer than 5

micrometers on membrane filters could be estimated from the relation σ=(N )

0.591

. Thus for counts 

of about 100 fibers, the coefficient of variation could be estimated at about 15.2% and the 95%

confidence limits at ±30.4%.

These values are lower than the values reported by Conway and Holland (11.3). Recently, the 

Johns-Manville Corporation conducted an in-house investigation of the asbestos count method.

(11.10). The study data contained total fiber countsfor over 100 filters with each filter counted by

two to five counters. From the Johns-Manville data, NIOSH calculated over 100 estimates of the

count CV for the method (11.11). The NIOSH CV estimates included random intrafilter variations

and intercounter variations, but did not include random pump flow rate variations. It was found

that the count coefficient of variation (all random variations except for pump variations) was a

function of the total fiber count. NIOSH then included a CV of 0.05 for random pump variations (see 

Section 9.1) in the CV-estimator equation to obtain a CVT-estimator. The CVT-estimator line is

plotted on Figure 3 for total fiber counts in the range 10 to 100 fibers. Or the following equation 

can be used: CVT=[antilog 10(-0.215-0.203(log10FB))+0.0025]2

where FB is total fiber count as discussed in Section 10. 

Figure 3 demonstrates that for a total fiber count of 100, the best CVT is attainable with the 

appropriate sampling times given in 8.1.3 and the count rules in 8.3.9. When making 
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 28 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

decisions regarding compliance with the OSHA asbestos exposure standards in 29 CFR 

1910.1001, the statistical procedures given in Leidel et al. (11.11) should be followed. The

procedures are based on statistical theory and assumptions given in References 11.12,

11.13.

Because of the possibility of systematic biases due to differences between microscopes,

counters, and laboratories as discussed above, it is strongly recommended that any

laboratory counting asbestos should participate in an interlaboratory quality control

program that includes the counting of standard reference filters. These standard filters

are available from NIOSH through the Proficiency Analytical Testing (PAT) Program. The 

PAT Program is used by the American Industrial Hygiene Association (AIHA) as part of its 

Laboratory Accreditation Program. Each laboratory's quality control program must 

include protocols for routinely adjusting, and calibrating sampling and counting 

equipment plus training and evaluation programs for counters.

5. Advantages and Disadvantages of the Method

5.1. The method is intended to give an index of employee exposure to airborne asbestos fibers of

specified dimensional characteristics.

5.2. It is not meant to count all asbestos fibers in all size ranges or to differentiate asbestos from other 

fibrous particulates.

6. Apparatus

6.1. Sampling Equipment

The personal sampling equipment train consists of 1) personal sampling pump, 2) tubing, 3) 

clothing spring clip, 4) tubing-to-field monitor metal adaptor, and 5) field monitor (filter and 

holder).

6.1.1.Personal Sampling Pump. The pump must be capable of sampling at 1.0 to 2.5 liters per minute 

(lpm) against a flow resistance of 7.5 inches of water (1.4 mm Hg) for 8 continuous hours on a 

fully charged battery.

6.1.2. Tubing. Laboratory tubing such as rubber or plastic with 6-mm bore and about 100 cm length.

6.1.3.Clothing Spring Clip. The clip attaches the rubber tubing to the lapel or shirt of the individual 

being monitored.

6.1.4. Tubing-to-field Monitor Adaptor. A short metal adaptor with ridges on one end to grip the 

inside of the tubing. The other end is designed for a pressure fit into the field monitor.

6.1.5. Field Monitor (Filter and Holder). The only field monitor currently considered acceptable by 

NIOSH is manufactured by the Millipore Corporation. The unit consists of 1) a three section 

styrene plastic case designated Millipore Aerosol Monitor Case, 2) a 37-mm diameter plain 

white cellulose ester membrane filter designated Millipore AA (pore size of 0.8 micrometer), 3) 

a support pad, and 4) two plastic sealing caps. If a large number of samples are to be taken, it 

may be less expensive to reuse the plastic cases. Great care must be taken in the cleaning and 

reassembly process. The outside mating surfaces of the field monitors may be covered with a

"shrink-fit" band to provide proper sealing and a writing surface for filter identification.

6.2. Optical Equipment and Microscope Features

6.2.1.Microscope body with binocular head.

6.2.2. 10X Huygenian eyepieces are recommended. Other eyepieces can be substituted if necessary. 

Wide field eyepieces can be used; however, wide field eyepieces may yield a count field area 

less than 0.003 mm2 with the Porton reticle. This is not always desirable from the standpoint of 

obtaining optimum sampling times (see Section 8.1.3). If wide field eyepieces are used, it is 

preferable to use the Patterson Globe and Circle reticle to obtain a larger count field area.

6.2.3.Koehler illumination (preferably built- in with provisions for adjusting light intensity).

6.2.4.A Porton reticle is recommended. Others such as the Patterson Globe and Circle can be 

substituted.

6.2.5.Mechanical stage.
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6.2.6. Phase-Contrast condenser with a numerical aperture (N.A.) equal to or greater than the N.A. of 

the objective.

6.2.7. 40-45X phase contrast achromatic objective (N.A. 0.65 to0.75).

6.2.8. Phase-ring centering telescope or Bertrandlens.

6.2.9.Green or blue filter, if recommended by microscopemanufacturer.

6.2.10. Stage micrometer with 0.01 mmsubdivisions.

6.2.11. For general guidance on phase contrast microscopy, consult Needham (11.12), Clark (11.15) 

and McCrone (11.14).

6.3. Filter Mounting Equipment. Experience has shown that certain equipment is useful for efficient 

sample mounting. The following items are recommended for extracting and mounting a portion of 

the filter for counting.

6.3.1. Microscope slides. 2.5 by 7.5 cm glass slides are most commonly used. Sample number, data, 

initials, etc., can be conveniently written on a frosted endslide.

6.3.2.Cover Slips. Cover slips are a necessary part of the slide mount and optical system. The shape 

should be appropriate for the size of the filter wedge. The appropriate cover slip depends upon 

the objective to be used. Ordinarily, objectives are optically corrected for a #11⁄2 (0.17 

millimeter) thickness cover slip. Improper cover glass thickness will detract from the final 

image quality.

6.3.3. Scalpel. A scalpel is needed to cut out a portion of the filter to be examined. A number-ten 

curved blade scalpel isrecommended.

6.3.4. Tweezers. A pair of fine-tipped tweezers is used to remove the membrane filter slice from the 

field monitor and place it upon the slide.

6.3.5. Lens Tissue. To insure cleanliness, a lint-free tissue is recommended. This tissue should also be 

used for wiping mounting tools and for cleaning slides and coverslips.

6.3.6.Glass Rod. A fire-polished glass rod may be used to spread the mounting solution on the slide.

6.3.7.Wheaton Balsam Bottle. This special glass container has a glass top which prevents 

contamination of the mounting solution. A glass rod isincluded for dispensing the solution.

7. Reagents

Chemicals should be reagent grade, free from particles and color, conforming to the specifications of the

Committee on Analytical Reagents of the American Chemical Society, where such specifications are

available.

7.1. Dimethyl phthalate

7.2. Diethyl oxalate

Avoid getting the mounting solution on the skin. Wash skin promptly with soap and water if skin contact

occurs.

8. Procedure

8.1. Sampling

8.1.1.General Information

Guidelines for the monitoring of employee exposures to industrial atmospheres are given in 

Reference 11.8. The Federal requirements for monitoring employee exposure to airborne 

asbestos are found in 29 CFR 1910.1001.

8.1.2. Mounting the Sampling Pump on the Worker

Fasten the sampling pump to the worker's belt and fasten the field monitor to the lapel or .shirt 

front (as close to the breathing zone as is practical). Remove the top cover of the plastic 

monitor, then invert the monitor making certain the exposed filter is facing downward. Turn 

the pump on and adjust to the calibrated flow rate (1.0 to 2.5 lpm). Record the following 

information in a logbook.

1. Filter number

2. Pump start time anddate

3. Flow rate

4. Subject's name and job title
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5. Type of operation or process

6. Ventilation controls and is the worker wearing a respirator approved for asbestos?

The pump should be checked periodically during the sampling period for proper operation

and flow rate.

8.1.3.Optimum Sampling Times

The requirement for the minimum count of 100 fibers or 20 fields in 8. 3. 9 was determined to 

be the best compromise to achieve adequate precision for the airborne fiber estimate and 

reasonable counting times. An optimum fiber density of about 1 to 5 fibers per microscope 

count field is recommended. To estimate appropriate sampling times for feasible counting and 

optimal counting, one must consider the following constraints:

1. microscope count field area (generally 0.003 to 0.006mm2

)

2. pump flow rate (typically 2.5 lpmmaximum)

3. average airborne fiber concentrations

4. counting rule range of 20 to 100 fields

5. adequate fiber density to obtain a minimum count of 10 fibers in 100 fields, which is the

least total fiber count that yields an acceptable countprecision

6. background airborne particulate levels that can reduce the count precision due to an

obscuring of fibers on the filtersurface

The preceding constraints were considered in drawing Figures 1 and 2. These figures were 

developed from the following relationship:

ffddmmddeeffnnaa ttffmmff = �FFLL� �

MMFFAA� FFBB EECCAA

(FFFF)(AACC)(1000)

mmffnnnnttffff

where:

FB/FL = 1 to 5 fibers/field

ECA = effective collecting area of filters (855 mm2 for 37-mm fiilter with effective diameter of 

33 mm)

MFA = microscope field area (generally 0.003 to 0.006 mm2

) 

FR = Pump flow rate (generally 1.0 to 2.5 lpm)

AC = Air concentration of fibers in fibers/cm3 . 

Figure 1 (microscope field area = 0.003 mm2

) and Figure 2 (microscope field area = 0.006 

mm2

) show optimum and feasible sampling times for a pump flow rate of 1.7 lpm. Each 

individual responsible for sampling asbestos should prepare a similar chart for his particular 

pump flow rate and microscope field area before sampling is performed to aid in estimating 

proper sampling times. On Figures 1 and 2, the areas with solid shading lines are generally 

the optimum conditions for counting. The broken shading lines are for conditions very close 

to optimal.

However, feasible counting conditions may extend down to about 0.1 fiber/field and above 

5 fibers/field. Recommended sampling times are most strongly influenced by background 

airborne particulate levels, once all the other constraints have been estimated. For heavy 

particulate levels, it may be necessary to limit each filter to about 60 to 180 minutes 

sampling duration. Each individual responsible for sampling should work closely with the 

microscopist to attain as high as possible filter surface fiber densities (up to about 5 

fibers/field), while avoiding filter surface background particulate levels that create very 

difficult or impossible counting conditions. If one has very little idea of airborne fiber and 

particulate levels, the best procedure is to take several long samples (as. one 8-hour or two 

consecutive 4-hour samples) in conjunction with several short samples (as four consecutive 

2-hour or eight consecutive 1-hour samples). If the longer samples prove very difficult to

count, the microscopist will have the shorter samples to fall back on.

From Figures 1 and 2, it can be seen that there are certain sampling times which will yield

optimum fiber densities on the filter for almost all airborne fiber concentrations from 1 to 10

fibers/cm3

. These optimum times have been calculated and are presented in Figure 4. Note
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

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that the optimum times given by Figure 4 are approximate and can be varied by as much as

±25%. The nomogram is intended as a guide to be used where no prior knowledge of the air 

concentration is available.

8.1.4. End of Sampling Period

Remove the field monitor, replace the plastic top cover and the small end caps, and store the 

monitor. Always shut off the pump when changing monitors to avoid contaminating or 

damaging the pump. Record the pump shutoff time and flow rate in the logbook.

8.1.5. Blanks

With each batch (25 to 50 filters) of samples sent for analysis, submit two unopened field

monitors which have been subjected to the same treatment as the samples except that they

were not exposed to the sampling environment. Label these as blanks. If the blanks yield fiber 

counts greater than 5 fibers/ 100 fields, then the entire sampling procedure should be 

examined carefully for the cause of contamination. The mounting solution of Section 8.2.1 

should also be examined for contamination and/or crystal growth. 

8.1.6. Shipping

The field monitors in which the samples are collected should be shipped in a rigid container 

with sufficient packing material to prevent crushing.

8.1.7.Numbers of Samples

When sampling for the Federal ceiling standard of 10 fibers (>5 μm)/cm3

, [29 CFR 1910-.1001(b) 

(3), effective July 7, 1972], only one sample (15 minutes maximum duration) is necessary, 

theoretically. However, several samples should be taken during expected periods of peak air 

concentrations to allow for detection of gross sampling or counting errors.

When sampling for determination of noncompliance with the Federal 8-hour TWA standard of 

2 fibers (>5 μm)/cm3

, [29 CFR 1910.100l(b) (2)], one should continuously sample as large a 

portion of the work day as is feasible for airborne concentrations of about 2 to 10 fibers/cm3

. 

However, for a lower airborne concentration such as 0.5 fiber/cm3

, one sample might require 4 

to 8 hours sampling time in order to get the proper filter fiber density (Section 8.1.3). For this 

situation, the 8-hour TWA exposure would be determined from one 8-hour or two 4-hour 

samples as appropriate.

8.2. Sample Preparation

8.2.1. Preparation of Mounting Solution

A very important part of the sample evaluation is the mounting process. This process involves a 

special mounting medium of prescribed viscosity. The proper viscosity is important in order to 

expedite filter dissolving and still minimize particle migration. After the sample has been 

mounted, an elapsed time of approximately sixty minutes is needed before the sample is ready 

for evaluation.

Combine the dimethyl phthalate and diethyl oxalate in a one to one ratio by volume and pour 

into a Wheaton balsam bottle. Add approximately 0.05 (±0.005) grams of new membrane filter 

per milliliter of solution to reach the necessary viscosity. The mixture must be stirred 

periodically until the-filters have dissolved and a homogeneous mixture is formed. The normal 

shelf life of the mounting solution is about three months. Twenty milliliters of mounting 

solution will prepare approximately 300 samples.

8.2.2. Sample Mounting

Cleanliness is important! A dirty working area may result in sample contamination and 

erroneous counts. The following steps should be followed when mounting a sample.

1. Clean the slides and cover slips with lens tissue. Lay each slide down on a clean surface

with the frosted end up. It is a good practice to rest one edge of the cover slip on the slide

and the other edge on the working surface. By doing this, you keep the bottom surface

(the one which contacts the filter) from becomingcontaminated.

2. Wipe all the mounting tools clean with lens tissue and place them on a clean surface

(such as lenstissue). All tools should be wiped clean prior to mounting each sample.
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3. Using the glass rod supplied with the Wheaton balsam bottle, apply a drop of mounting

solution onto the center of the slide. It may be necessary to adjust the quantity of

solution so that after the cover slip has been placed on top, the solution extends only

slightly beyond the filter boundary. If the quantity is greater than this, particle migration

may occur.

4. Using another glass rod, spread the mounting media into a triangular shape. The size of

this triangle should coincide with the dimension of the filterwedge.

5. Separate the middle and bottom sections of the field monitor case to expose the filter.

Cut a triangular wedge from the center to the edge of the filter using .the scalpel. The size

of the wedge should approximate one-eighth of the filter surface. The filter can be very

carefully removed from the cassette for cutting, but this should only be done with great

care.

6. Grasp the filter wedge with the tweezers on the perimeter of the filter which was

clamped between the monitor case sections. Do not touch the filter with your fingers.

Place the wedge, sample side up, upon the mountingmedium.

7. Pick up a clean coverslip with tweezers and carefully place it on the filter wedge. Once

this contact has been made, do not reposition the coverslip.

8. Label the slide with the sample number and current date before proceeding to the next

filter: On the bottom (backside) of the slide, trace the perimeter of the filter wedge with a

felt tip marking pen. This will ·enable the counter, after the filter has become transparent,

to stay within the filter perimeter whencounting.

9. The sample should become transparent within fifteen minutes. If the filter appears

cloudy, it may be necessary to press very lightly on the cover slip. This is rarely necessary;

however, counting should not be started until an hour after the mounting. This allows the

microscopic texture of the filter to become invisible to microscopeviewing.

10. Discard the sample mount after two days if it has not been counted. Crystals appearing

similar to asbestos fibers may begin to grow at the mounting media/air interfaces. They

seldom present any problems if the slide is examined before two days. In any case, stay

away from the filter's edges when counting andsizing.

8.3. Counting of Fibers 

8.3.1. Place the slide on the mechanical stage of the microscope and position the center of the 

wedge under the objective lens and focus upon the sample. Start counting from one end of 

the wedge and progress along a radial line to the other end (count in either direction from 

perimeter to wedge tip). Random fields are selected, without looking into the eyepieces, by 

slightly advancing· the slide in one direction with the mechanical stage control.

8.3.2. It is essential to continually scan over a range of focal planes (generally the upper 10 to 15 

micrometers of the filter surface) with the fine focus control during each field count. This is 

especially necessary for asbestos fibers due to their impaction into the filter matrix.

8.3.3.On most airborne samples, asbestos fibers will generally have fiber diameters less than one 

micrometer. Therefore, it is necessary to look carefully for faint fiber images.

8.3.4. Regularly check phase ringalignment.

8.3.5.When an agglomerate (mass of material) covers a significant portion of the field of view 

(approx 1/6 or greater)--reject the field and select another. (Do not include it in the number of 

fields counted.) However, report the fact asit may have meaning on other data collection.

8.3.6. Bundles of fibers are counted as one fiber unless both ends of the fiber can be clearly resolved.

8.3.7.Count only fibers with a length to width ratio greater than or equalto 3:1.

8.3.8.Count only fibers greater than 5 micrometers in length. (Be as accurate as possible in accepting 

fibers near thislength.) Measure curved fibers along the curve to estimate the total length.

8.3.9.Count as many fields as necessary to yield a total count of at least 100 fibers. Exceptions: a) 

count at least 20 fields even if you count more than 100 fibers, and b) stop at 100 fields even if 

you haven't reached 100 fibers.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

8.3.10. For fibers that cross either one or two sides of the counting field, the following procedure is 

used to obtain a representative count.

COUNT any fiber greater than 5 micrometers in length that lies entirely within the counting 

area. COUNT as "1/2 fiber" any fiber with only one end lying within the counting area. DO NOT 

COUNT any fiber crossing any two sides.

Reject and do not count all other fibers. Refer to Figures 5 through 10. Note that the fibers in 

Figures 5 through 10 are not representative of the appearance of most asbestos fibers. Most 

fibers have a very faint image.

9. Calibration and Standards

9.1. Sampling Train Calibration

The accurate calibration of the sampling pump is essential to the correct calculation of the air 

volume sampled. The frequency of calibration is dependent on the use, care, and handling to which 

the pump is subjected. Pumps must be recalibrated if they have just been repaired, misused, or 

received from the manufacturer. If the pump receives hard usage, more frequent calibration may be 

necessary. Ordinarily, pumps should be calibrated in the laboratory both before they are used in the 

field and after they have been used to collect a large number of field samples.

The accuracy of calibration is dependent upon the type of instrument used as a reference. The 

choice of a calibration instrument will depend largely on where the calibration is performed. For 

laboratory testing, a 1-liter buret used as a soap bubble flow meter or wet-test meter is 

recommended. Other standard calibrating instruments, such as a spirometer, Marriott's bottle, or 

dry gas meter can be used. The calibration should be of sufficient precision that the 95% confidence 

limits on the flow rate are ±10% (95% of the flow rates will fall within ±10% of the calibrated value).

Instructions for calibration with the soap bubble flow meter follow. The sampling train used (pump, 

hose, filter cassette) in the pump calibration should be the same as the one used in the field.

9.1.1.Check the voltage of the pump battery with a voltmeter both with the pump off and while it is 

operating to assure adequate voltage for calibration. If necessary, charge the battery to 

manufacturer's specifications.

9.1.2. Fill a beaker with 10 ml of soap solution.

9.1.3.Connect the filter cassette inlet to the top of the buret with a length of hose.

9.1.4. Turn the pump on and moisten the inside of the soap bubble meter by immersing the open 

end of the buret into the soap solution and drawing bubbles up the inside of the buret. 

Perform this task until the bubbles are able to travel the entire length of the buret without 

breaking.

9.1.5.Adjust the pump rotameter to provide a flow between 1.5 to 2.5 lpm.

9.1.6.With a water manometer, check that the pressure drop across the filter is less than 13 inches of 

water (about 1 inch of mercury).

9.1.7. Start a soap bubble up the buret and measure the time it takes for the bubble to travel a 

minimum volume of 1 liter.

9.1.8.Repeat the procedure in 9 .1. 7 at least three times, average the results, and calculate the 

calibrated flowrate by dividing the volume traveled by the soap bubble by the elapsed time. If 

the range between the highest and lowest of the three flow rates is greater than about 0.33 

lpm, then the calibration should be repeated since it is likely that the precision is not adequate.

9.1.9.Data required for the calibration include the volume measured, elapsed time, pressure drop, air 

temperature, atmospheric pressure (or elevation), pump serial number, date, and name of 

person performing the calibration.

9.1.10. Corrections to the flow rate for pumps with rotameters may be necessary if the pressure 

(elevation) or temperature where the samples are collected (actual flow rate) differs 

significantly from that where the calibration was performed (indicated flow rate). Actual flow 

rates at time of sampling may be calculated for a linear scale rotameter by using the following 

correction formula: 
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QQaaccaaaaaaaa = QQiiiiiiiiccaaaaiiii�� PPccaaaa

PPaaccaaaaaaaa

� ∗ �TTaaccaaaaaaaa

TTccaaaa

�

where both pressure (P) and temperature (T) are in absolute units such as: 

psia = psig + 14.7

deg Rankin = deg Fahrenheit + 460 

deg Kelvin = deg Celsius + 273

9.2. Microscope Setup

9.2.1. Porton Reticle and the Counting Field

The asbestos fiber count procedure consists of comparing fiber length to the diameters of 

calibrated circles of a Porton reticle, and counting all fibers greater than 5 micrometers in 

length lying within a given counting field area. The Porton reticle is a glass plate inscribed with 

a series of circles and rectangles. The left half of the reticle is divided into six rectangles 

constituting the counting field. The counting field is illustrated in Figures 5 through 10.

9.2.2. Placement in Eyepiece

The Porton reticle is placed inside the Huygenian eyepiece where it rests on the field limiting 

diaphragm. If other types of eyepieces are used, it may be necessary to insert a counting collar 

for retaining the reticle. The reticle should always be kept clean, since dirt on the reticle is in 

focus and could complicate the counting and sizing process.

9.2.3. Stage Micrometer

The Porton reticle cannot be used for counting until it has been properly calibrated with a 

stage micrometer. Most stage micrometer scales are approximately two millimeters long and 

are divided into units of one-hundredth of a millimeter (ten micrometers).

9.2.4. Microscope Adjustment

When adjusting the microscope, follow the manufacturer's instructions while observing the 

following guidelines.

1. The light source image must be in focus and centered on the condenser iris or annular

diaphragm.

2. The particulate material to be examined must be infocus.

3. The illuminator field iris must be in focus, centered on the sample, and opened only to

the point where the field of view isilluminated.

4. The phase rings(annular diaphragm and phase-shifting elements) must be concentric.

9.2.5. Porton Reticle CalibrationProcedure

Each eyepiece-objective-reticle combination on the microscope must be calibrated. Should 

any of the three be changed (disassembly, replacement, zoom adjustment, etc.), the 

combination must be recalibrated. Calibration may change if interpupillary distance is 

changed. For proper calibration, the following procedure should be followed closely.

With a 10X objective in place, place the stage micrometer on the mechanical stage, focus the 

millimeter scale, and center the image. Change to the 40-45X objective and adjust the first 

millimeter scale division to coincide with the left boundary of the Porton rectangle. Measure 

the distance between the left and extreme right boundaries of the Porton rectangle, 

estimating any portion of the final division. This measurement represents 200 L units. The 

rectangle is 100 L units on the short vertical dimension. The calculated "L" is inserted into the 

formula D = L(2N)

1/2 where "N" is the circle number (indicated on the reticle) and "D" is the circle 

diameter. Since the circle diameters vary logarithmically, every other circle doubles in 

diameter. For example, circle number three is twice the diameter of number one; number four 

is twice the diameter of number two. When the circle sizes have been determined, the 

counting field area which consists of the left six smaller rectangles can be calculated from the 

relation 10,000 L2

. This completes the reticle calibration for this specific objective-eyepiecereticle combination.
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NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Example for Porton Reticle

The following calibration was obtained for a pair of 10X Huygenian eyepieces and a 43X 

objective:

200 L = 0.148 mm = 148 micrometers

100 L = 0.074 mm = 74 micrometers

One L-unit = 0.74 micrometers

Thus Circle #1 has a diameter D = L(2N)

1/2 = 0.74(21

)

1/2 = 0.74 (1.414) = 1.05 micrometers.

Then our circle diameter calibration table looks like:

Diameter of Circle #1 = 1.05 micrometers 

 #2 = 1.48 

#3 = 2.09

#4 = 2.96

#5 = 4.19

#6 = 5.92

Field area = (10,000) (L

2

) = (100 L) (100 L) = (0.074) (0.074) = 0.0055 mm2

Thus fibers with a length greater than a distance halfway between the diameters of the #5 and

#6 circles would be counted.

If a Patterson Globe and Circle reticle is used, a different calculation procedure is required. The

circle diameters are related as follows. The #25 circle diameter is (0.1) (reticle length).

The circle diameters are proportional to the ratio of their numbers. Thus the #20 circle 

diameter is (20/25) or 0.8 times the #25 circle diameter. 

10. Calculations

10.1.The average airborne asbestos fiber concentration estimated by the filter sample may be calculated

from the following formula:

AA =

��

FF

FF � − �BB BB

BB �� (EE )

(1000)(FFFF)(TT)(MM )

where:·

AC = Airborne fiber concentration in (fibers >5 μm)/cm3

. 

BFB = Total number of fibers counted in the BFL fiels of the blank or control filters in fibers >5 μm.

BFL = Total number of fields counted on the blank or control filters.

ECA = Effective collecting area of filter (855 mm2 for a 37-mm filter with effective diameter of 33 

mm).

FR = Pump flow rate in liters/min (lpm).

FB = Total number of fibers counted in the FL fields in fibers >5 μm.

FL = Total number of fields counted on the filter.

MFA = Microscope count field area in mm2 (generally 0.003 to 0.006).

T = Sample collection time in minutes.

10.2. Recount criteria. It is very desirable for a counter to conduct a "blind recount" for about 1 in every 

10 filter wedges (slides) counted. Alternatively, a second counter could perform the blind recount. 

In training sessions for novice counters, the trainee should conduct a blind recount for filter wedges 

counted by an experienced, proficient counter. In all cases, we will observe differences between the 

first and second counts of the same filter wedge. Most of these differences will be due to chance 

alone, that is, due to the random variability (precision) of the count method. Statistical recount 

criteria enable us to decide whether observed differences can reasonably be explained due to 

chance alone or are probably due to systematic differences between counters or microscopes or 

due to some other biasing factor.

The following recount criterion is for a pair of counts that estimate some airborne fiber 

concentration (AC) in fibers/cm3

. The criterion is given at the type-1 error level. That is, there is a 5% 

maximum risk that we will reject a pair of counts for the reason that one might be biased, when the 

large observed difference is really due to chance.
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 36 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Reject a pair of counts because one might be biased if:

(AA 2 − AA 1) ee 2.77(AACC����)(CC FFFF)

where:

AC1 = lower estimated airborne fiber concentration

AC2 = higher estimated airborne fiber concentration

AA���� = average of the two airborne concentration estimates

CVFB = average CV for the two concentration estimates which are a function of the total fiber count 

(FB) in each case. Use the relation in Section 4 or Figure 3.

For a pair of counts on the same filter, reject the pair because one might be biased if:

(FF 2 − FFBB1) ee 2.77(FF����)(CC FFFF

− )

where:

FB1 = lower fiber count on the filter (total fibers)

FB2 = higher fiber count on the filter (total fibers)

FF���� = average of the two total fiber counts

CC FFFF

− = CVT for the value FB. Use the relation in Section 4 or Figure 3.

11. References

11.1.Lynch, J. R., K. J. Kronoveter, and N. A. Leidel, "Validity of the Poisson Distribution in Dust Counting",

unpublished.

11.2.Weidner, R. B. and H. E. Ayer, "Dust Exposure in Asbestos Processing", Transactions of the American

Conference of Governmental Industrial Hygienists, May 1972, pp. 103-121, San Francisco, California.

11.3.Conway, R. E. and W. D. Holland, "Statistical Evaluation of the Procedure for Counting Asbestos

Fibers on Membrane Filters", LFE Corporation, Richmond, California. Prepared for the Asbestos on 

Membrane Assoc/North America, New York, 1973.

11.4.Leidel, N. A. and K. A. Busch, "An Evaluation of Phase Contrast Microscopes for Asbestos Counting", 

presented at the 1974 American Industrial Hygiene Conference, Miami Beach, Florida, May 18, 1974, 

unpublished.

11.5.Beckett, S.T. and M.D. Attfield, "Inter-Laboratory Comparison of the Counting of Asbestos Fibers 

Sampled on Membrane Filters", Ann Occup Hyg 17:85-96, 1974.

11.6.Rajhans, G. S. and G. M. Bragg, "A Statistical Analysis of Asbestos Fiber Counting in the Laboratory 

and Industrial Environment", Am Ind Hyg Assoc J 36(12):909-915, 1975.

11.7.Leidel, N. A. and K. A. Busch, "Statistical Methods for the Determination of Noncompliance with 

Occupational Health Standards", NIOSH Technical Publication 75-159, 1975.

11.8.Leidel, N. A., K. A. Busch, and J. R. Lynch, "Occupational Exposure Sampling Strategy Manual", NIOSH 

Technical Publication 77-173, 1977.

11.9.Leidel, N. A.; K. A. Busch, and W. E. Crouse, "Exposure Measurement Action Level and Occupational 

Environmental Variability", NIOSH Technical Publication 76-131, 1975.

11.10. Comments of the Johns-Manville Corporation with Respect to the Notice of Proposed Rulemaking:

Occupational Exposure to Asbestos, Federal Register, October 9, 1975. Submitted to the public 

record at the U. S. Department of Labor, Occupational Safety and Health Administration, April 1976.

11.11. Leidel, N. A., S. G. Bayer, R. D. Zumwalde, and K. A. Busch, USPHS/NIOSH Membrane Filter Method 

for Evaluating Airborne Asbestos Fibers, to be published by NIOSH in 1977.

11.12. Needham, G. H., The Practical Use of the Microscope, Charles C. Thomas Publishing Corporation, 

Springfield, Illinois, 1958.

11.13. Clark, G. L., The Encyclopedia of Microscopy, Rheinhold Publishing Corporation, New York, 1961.

11.14. McCrone, W. C. and J. G. Delly, I. The Particle Atlas, Edition Two, Ann Arbor Science Publishers, Inc., 

Ann Arbor, Michigan, 1973.
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 37 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 1 Optimum Sampling Times for airborne asbestos where microscopic field area = 0.003 mm2
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 38 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 2 Optimum sampling times for airborne asbestos where microscopic field area = 0.006 mm2

Figure 3 Total coefficient of variation as a function of total fiber count
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 June 2019 - Page 39 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 4 Nomogram of optimum sampling times for airborne asbestos fibers in concentrations of 1 to 10 fibers/cm3 

Figure 5 DO NOT COUNT. Fiber crosses top and bottom sides.

Figure 6 COUNT. One fiber.
ASBESTOS and OTHER FIBERS by PCM: METHOD 7400, Issue 3, dated 14 june 2019 - Page 40 of 40 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Figure 7 COUNT. One-half fiber. Fiber crosses left side and one end lies within the count area.

Figure 8 COUNT. One-half fiber. Fiber crosses bottom side and one end lies within count area.

Figure 9 DO NOT COUNT. Fiber crosses two sides.

Figure 10 DO NOT COUNT. Fiber crosses two sides (bottom left corner). COUNT. One-half fiber. Fiber crosses bottom side and one end lies 

within count area. COUNT. One fiber (top right corner).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ALKALI METAL CATIONS Na+

, K+, Li+ 7405

NaOH, KOH, LiOH MW: 40.00 (NaOH); CAS: 1310-73-2 (NaOH) RTECS: WB4900000 (NaOH)

 56.11 (KOH); 1310-58-3 (KOH); TT2100000 (KOH); 

 23.95 (LiOH) 1310-65-2 (LiOH) OJ6307070 (LiOH)

METHOD: 7405, Issue 1 EVALUATION: FULL Issue 1: 20 March 2018 

OSHA: 2 mg/m3 (NaOH)

NIOSH: C 2 mg/m3 (NaOH, KOH); IDLH 10 mg/m3 (NaOH)

OTHER OELs: Refs. [1,2]

PROPERTIES: basic, hygroscopic, caustic solids and aerosols, 

VP not significant

SYNONYMS: sodium hydroxide; potassium hydroxide; lithium hydroxide; alkali (LiOH, NaOH, KOH); caustic soda (NaOH); caustic 

potash (KOH); lye (NaOH, KOH)

SAMPLING

SAMPLER: FILTER, 37-mm quartz fiber, in 2- or 3-piece 

filter holder

FLOW RATE: 2-4 L/min

VOL-MIN: 30 L

 -MAX: 1000 L

SHIPMENT: routine

SAMPLE

STABILITY: ≥7 days at 25 oC & ≥27 days at 4 oC [3].

BLANKS: 3 field blanks minimum per sample set

ACCURACY

RANGE 

STUDIED: NaOH: 0.08 to 3.7 mg/m3

KOH: 0.07 to 7.4 mg/m3; 

LiOH: 0.16 to 7.2 mg/m3 [3]

BIAS: negligible

OVERALL

PRECISION (SS�rrrr)*: NaOH: 0.023 to 0.066; KOH: 0.018 to

0.043; LiOH: 0.016 to 0.023 [3]

EXPANDED 

UNCERTAINTY*: <22% for all three cations [6] 

*Accuracy calculations were carried out in accordance with 

ISO guidelines [5] rather than NIOSH accuracy criteria.

MEASUREMENT

TECHNIQUE: Ion Chromatography

ANALYTE: Na+, K+, Li+

DISSOLUTION: 10 mL 0.005 M H2SO4, 15 minutes in 

ultrasonic bath

ELUENT: 0.005 M H2SO4

DETECTOR: Conductivity detector

CALIBRATION: ≈0.5 to ≈10 mg/L Na+, K+, Li+ in 0.005 M 

H2SO4

RANGE: <0.01 to >5 mg Na+, K+ per sample; <0.05 

to >2.5 mg Li+ per sample [3,4]

ESTIMATED LOD: 0.003 mg Na+, K+ per sample; 

0.002 mg Li+ per sample [3]

PRECISION (SS�rr): NaOH: 0.009 to 0.030; KOH: 0.008 to 0.019;

LiOH: 0.007 to 0.010 [3,4]

APPLICABILITY: The method is meant for the measurement of alkaline dusts. It is specific for Na+, K+, and Li+ but does not 

differentiate between the hydroxides and their corresponding salts if both are present in the air. If the cations are present only in 

the form of hydroxides, the method is specific for these basic compounds. In other circumstances, the results obtained represent 

the highest air concentration of the hydroxides that could be extant. The method can also be used to measure alkaline earth

hydroxides such as calcium hydroxide [3,4].

INTERFERENCES: Sodium and potassium are ubiquitous in the environment and, if present in the sampled air at high 

concentrations, other soluble salts of Na and K can positively interfere. High concentrations of ammonium can interfere with the 

sodium peak in the chromatographic signal.
ALKALI METAL CATIONS: METHOD 7405, Issue 1, dated 20 March 2018 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

OTHER METHODS: NIOSH Method 7401 for alkaline dusts is an acid-base titrimetric procedure that measures hydroxide 

concentration (alkalinity), [OH-] [7]. A harmonized international standard, ISO 17091 [4], has been published.

REAGENTS:

1. Sodium, potassium and lithium standard

solutions, 1000 ± 10 mg/L; traceable to

national standards.

2. Calibration stock solution: sodium and

potassium, 50 mg/L; lithium, 25 mg/L. Pipet

625 μL Li standard solution and 1250 μL of

each of the Na and K standard solutions into

25-mL volumetric flask. Dilute to volume

with deionized water. This solution can be

stored at 4°C for at least 2 months.

3. Sulfuric acid stock solution, 2.5 M H2SO4.

4. Sulfuric acid extraction / eluent solution,

0.005 M H2SO4. Pipet 2 mL 2.5 M sulfuric acid

into a 1-L volumetric flask containing

deionized water. Fill to volume using

deionized water.

5. Water, deionized, >18.2 MΩ-cm-1 resistivity.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Samplers: 37-mm diameter quartz fiber filters

in cassette filter holders.

NOTE: Sodium and potassium are ubiquitous

in the environment and their presence 

in filter materials can lead to high 

blanks. Thus it is essential to check 

blank levels of each batch of filters. 

2. Personal sampling pump, 2 to 4 L/min, with

flexible connecting tubing.

3. Flow meter, portable.

4. Ion chromatograph (IC) with autosampler,

cation exchange column (e.g., 250 mm x 4

mm) with guard column (e.g., 50 mm x 4

mm), column oven capable of maintaining

column temperature at 40°C, column

thermostat, suppressor (e.g., cation selfregenerating suppressor) and conductivity

detector.

5. Data analysis unit (e.g., computer).

6. Volumetric flasks, polypropylene, 25-mL, 50-

mL, 1-L.

NOTE: For this method, it is preferable to use

plastic labware rather than glassware.

7. Adjustable pipettors with plastic tips, 10-

10000 μL.

8. Screw-cap polyethylene vessels, 15-20 mL.

9. Autosampler vials, polypropylene, with

screw-caps and siliconpolytetrafluoroethylene (PTFE) septa.

10. Disposable syringes, 5-10 mL.

11. Disposable filters, 0.45 μm pore size PTFE, 25-

mm diameter, suitable for IC.

12. Ultrasonic bath.

13. Ultrapure water system.

14. Tweezers, plastic or plastic-tipped.

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities and analysis. It 

is essential that suitable gloves, eye protection, laboratory coat, etc., be used when working with the 

chemicals. Perform sample preparation and analysis in a clean, well-ventilated area that is well removed 

from any possible contamination. Any area of skin affected by the stock, dissolution or detection solutions 

must be immediately washed with plenty of water. Avoid exposure by contact with skin or eyes, or by 

inhalation of vapor.

SAMPLING, SAMPLE TRANSPORT AND STORAGE: 

1. Calibrate each personal sampling pump with a representative sampler in line.
ALKALI METAL CATIONS: METHOD 7405, Issue 1, dated 20 March 2018 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

2. Sample at an accurately known flow rate between 2 and 4 L/min for a sample size of 30 to 1000 L. Do 

not overload the samplers.

3. When sampling is completed, remove the samplers and cap them. 

4. Submit at least three blank filters as field blanks from the same lot as sample filters for each set of 

samples, and handle them in the same way as field samples.

5. Transport the samples to the laboratory in a manner that prevents contamination or damage to the 

filters, and in accordance with established chain-of-custody procedures [8]. 

6. The samples can be stored at room temperature for one week; for longer storage, refrigerate the 

samples at 4 o

C.

NOTE: These storage times relate to the evaluated conditions. Longer storage at room temperature 

may be possible, but was not tested. 

7. Samples should be analyzed no more than four weeks after sample collection.

SAMPLE PREPARATION:

8. Remove sample cassettes from storage and, if necessary, bring to room temperature. 

9. Using tweezers, remove the sample filters from the filter holder (cassette) and place them in 15-20 mL 

plastic (polyethylene) screw-cap vessels. 

10. Rinse the inside walls of the cassette with 2-3 mL of extraction / eluent solution (0.005 M H2SO4) and 

direct the rinsate(s) into the plastic vessels that contain the filters. 

11. Bring the final volume of extraction / eluent solution inside the vessels to 10 mL, ensuring that the 

filters are completely immersed.

12. Seal the 15-20 mL plastic vessels and place them in an ultrasonic bath. Sonicate for 15 minutes and 

then allow to stand for at least 30 minutes. 

13. For each prepared sample, using a disposable syringe, remove 5 mL of the extraction solution and 

filter into an autosampler vial through a disposable PTFE filter. 

CALIBRATION AND QUALITY CONTROL:

14. Prepare at least five calibration working solution standards to cover the required concentration range, 

e.g., from 0.6 to 20 mg/L for Na+ and K+ and 0.3 to 10 mg/L for Li+

. Accurately pipet appropriate 

volumes of calibration stock solution into individual, labeled volumetric flasks. Dilute to the mark with 

water, close, and mix. 

NOTE: These calibration working solutions can be stored at room temperature for at least 2 weeks. 

Before use, their stability should be confirmed. If stability is inadequate, these solutions should

be prepared fresh daily.

15. Calibrate the ion chromatograph with the above five calibration solutions by preparing a calibration 

graph of peak height (in e.g., μS) vs. concentration (mg/L) for each cation.

16. Analyze calibration working standards together with samples, reagent blanks and field blanks at a 

frequency of at least 1 per 20 samples (3 minimum of each). 

17. Check recoveries with at least two media blanks and 3 quality control spikes per 20 samples.

MEASUREMENT:

18. Set the ion chromatograph to the recommended eluent flow rate (e.g., 1.0 mL/min) and system 

pressure (ca. 10 MPa) and other conditions specified by the instrument manufacturer. 

19. Using the autosampler, a 25-μL sample aliquot of the contents of each autosampler vial is injected into 

the ion chromatograph. 

20. Measure the peak heights of the lithium, sodium and potassium peaks (at retention times of 

approximately 4, 4.5 and 6 min, respectively) (Figure 1). If the peak height exceeds the linear 

calibration range, dilute with eluent, reanalyze, and apply the appropriate dilution factor in 

calculations.
ALKALI METAL CATIONS: METHOD 7405, Issue 1, dated 20 March 2018 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALCULATIONS:

21. Calculate the mass concentration of each cation, C (mg/m3

), in the air volume sampled, V (m3):

CC = (CCii × VVii × FF) − (CC0 × VV0)

VV ,

where:

C0 = mean concentration, in mg/L, of cation in field blank test solution;

Ci = concentration, in mg/L, of the ith cation (i = Na+

, K+, Li+) in sample test solution;

V = volume of air sample (m3

), where 1 L = 0.001 m3 

V0 = volume of field blank test solution (L)

Vi = volume of sample test solution (L) for ith cation (i = Na+

, K+, Li+) 

F = dilution factor for each sample test solution

EVALUATION OF METHOD:

The method was evaluated in accordance with an applicable international consensus standard, ISO 20581 

[9]. The component of variation of the method arising from analytical variability, as determined from 

spiked and dried quartz fiber filters, was found to be in the range of 0.9% to 3.0% for NaOH, 0.8% to 1.9% 

for KOH, and 0.7% to 1.0% for LiOH [3,6]. Mean recoveries from these samples were 102 ± 2% for NaOH, 100 

± 2% for KOH and 101 ± 2% for LiOH [3]. The expanded uncertainty of the method, using a coverage factor

(k) of 2 [10], was estimated as <22% for all three hydroxides [6]. Method quantitation limits were calculated 

based on the use of low-spike calibration standards [3, 11]. For a sampled air volume of 420 L, the 

quantitation limits were 0.041 mg/m3 for NaOH and LiOH and 0.034 mg/m3 for KOH. Storage stability of 

four weeks was demonstrated by recoveries >95% for quartz fiber filters spiked with NaOH, KOH and LiOH 

and stored at room temperature for 7 days and 4 o

C thereafter [6]. Influence of humidity was investigated 

at a relative humidity (RH) of 70%, established by means of a dynamic test gas apparatus [3]. Amounts of 

0.1 – 0.2 mg of NaOH, KOH and LiOH were spiked onto quartz fiber filters and then dried. Over two hours, 

air with RH=70% was drawn through these filters at 3.5 L/min; humidity was not found to influence the 

analytical recoveries of the three hydroxides. 

REFERENCES:

[1] American Conference of Governmental Industrial Hygienists (ACGIH) [2016]. Threshold Limit Values 

for chemical substances and physical agents & Biological Exposure Indices. Cincinnati, OH: ACGIH; 

updated annually. 

[2] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA) [no date]. GESTIS 

international limit values, http://limitvalue.ifa.dguv.de/. Date accessed January 17, 2018. 

[3] Breuer D, Heinrich B [2001]. Alkali metal hydroxides and alkaline earth hydroxides, Method no. 1

(lithium hydroxide, sodium hydroxide, potassium hydroxide, calcium hydroxide); in Air monitoring 

methods, Vol. 8 – The MAK Collection for Occupational Health and Safety. Weinheim, Germany: 

Wiley-VCH. 

[4] International Organization for Standardization (ISO) [2013]. ISO 17091, Workplace air – 

Determination of lithium hydroxide, sodium hydroxide, potassium hydroxide and calcium 

dihydroxide – Method by measurement of corresponding cations by suppressed ion 

chromatography. Geneva: ISO.

[5] ISO [1995]. ISO Guide 98, Guide to the expression of uncertainty in measurement. Geneva: ISO. 
ALKALI METAL CATIONS: METHOD 7405, Issue 1, dated 20 March 2018 - Page 5 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[6] Breuer D [2009]. Hydroxides (LiOH, NaOH, KOH, Ca(OH)2), in BGIA working folder, Sheet no. 7683.

Berlin: Erich-Schmidt Verlag.

[7] NIOSH [1994]. Method 7401, Alkaline dusts. In Schlect PC, O’Connor PF, eds. NIOSH manual of

analytical methods, 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS

(NIOSH) Publication No. 94-113.

[8] ASTM International [2010]. ASTM D4840, Standard guide for sampling chain of custody procedures.

West Conshohocken, PA: ASTM International.

[9] ISO [2016]. ISO 20581, Workplace air quality – General requirements for the performance of

procedures for the measurement of chemical agents. Geneva: ISO.

[10] Comité Européen de Normalisacion (CEN) [2009]. EN 13890, Workplace exposure – Procedures for

measuring metals and metalloids in airborne particles – Requirements and test methods. Brussels:

CEN.

[11] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. By

Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati: U.S. Department of Health and

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health, DHHS (NIOSH) Publication No. 95-117.

METHOD CONTRIBUTORS: 

Dietmar Breuer, Dr. rer. nat., Petra Heckmann and Krista Gusbeth, IFA, Sankt Augustin, Germany; final 

editing by Kevin Ashley, Ph.D., NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by the National 

Institute for Occupational Safety and Health (NIOSH), Centers for Disease Control and Prevention (CDC). In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.

FIGURE 1. Chromatogram of standard solution containing Li, Na, and K.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ZINC OXIDE 7502

 MW: 81.38 CAS: 1314-13-2 RTECS: ZH4810000

METHOD: 7502, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984

Issue 3: 20 October 2015

OSHA: 5 mg/m3 (fume, resp dust); 15 mg/m3 (total dust)

NIOSH: 5 mg/m3, C 15 mg/m3 (15 min) (dust); 5 mg/m3, STEL 

10 mg/m3 (fume)

PROPERTIES: Solid; d 5.61 g/cm3 @ 25 °C; MP 1975 °C

SYNONYMS: China white; zinc white; zincite

SAMPLING

SAMPLER: FILTER

(0.8 μm PVC membrane, 25 mm diameter, in 

open-face cassette)

FLOW RATE: 1 L/min to 3 L/min

VOL-MIN: 10 L

-MAX: 400 L

SHIPMENT: Routine

SAMPLE

STABILITY: Stable

BLANKS: 2 to 10 field blanks per set

BULK

SAMPLE: High-volume air sample required

ACCURACY

RANGE STUDIED: 0.1 mg/m3 to 11 mg/m3 [1,2] (180 L 

samples)

BIAS: 2.7% [2,3,4]

OVERALL

PRECISION ( ): 0.09 [2]

ACCURACY: ±21.6%

MEASUREMENT

TECHNIQUE: X-RAY POWDER DIFFRACTION

ANALYTE: Crystalline zinc oxide; direct analysis on 

filter

XRD: Copper target X-ray tube; optimize 

for intensity; 1° slit; graphite 

monochromator; scintillation detector; 

slow step scan, 0.02 degrees per 10 

seconds; integrated intensity with 

background subtraction

CALIBRATION: Suspensions of zinc oxide in 2-propanol

RANGE: 50 μg to 2000 μg per sample

ESTIMATED LOD: 5 μg per sample

PRECISION ( ): 0.15 @ 1 mg/m3; 0.05 for greater than 2 

mg/m3

APPLICABILITY: The working range is 0.25 mg/m3 to 10 mg/m3 for a 200 L air sample. The method does not distinguish zinc 

oxide fume from zinc oxide dust.

INTERFERENCES: Major interferences include ferric oxide, zinc, diamminedichlorozinc, triammonium 

pentachlorozincate(3−), diammonium tetrachlorozincate(2−), and diammonium zinc disulfate hexahydrate; these are 

resolved by using alternate analyte peaks. Particle size affects intensity measurements.

OTHER METHODS: This method combines and replaces NIOSH methods P&CAM 222 [1] and S316 [5]. The criteria 

document contains an elemental analysis for zinc [6].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ZINC OXIDE: METHOD 7502, Issue 3, dated 20 October 2015 - Page 2 of 5

REAGENTS:

1. Zinc oxide, ACS reagent grade. Average 

particle size between 0.5 μm and 10 μm.

2. 2-Propanol.*

3. Desiccant.

4. Glue or tape for securing filters to XRD holders.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: polyvinyl chloride (PVC) or PVCacrylonitrile membrane filters, 25 mm 

diameter, 0.8 μm pore size; three-piece filter 

cassette.

NOTE: An extension cowl on the filter cassette 

is desirable to produce a more uniform 

deposit and to prevent contamination 

of the open-face filter during sampling.

2. Personal sampling pump, 1 L/min to 3 L/min, 

with flexible connecting tubing.

3. High-volume sampling pump, 10 L/min.

4. X-ray powder diffractometer equipped with 

copper target X-ray tube and scintillation 

detector.

5. Reference specimen (mica, Arkansas stone or 

other stable standard) for data normalization.

6. Filtration apparatus and side arm vacuum flask 

with 25 mm filter holders.

7. Analytical balance (0.01 mg), magnetic 

stirrer, ultrasonic bath or probe, volumetric 

pipettes and flasks, desiccator, reagent bottles 

with ground glass stoppers, drying oven, 

polyethylene wash bottle.

SPECIAL PRECAUTIONS: 2-Propanol is flammable.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample open-face at 1 L/min to 3 L/min for a total sample size of 10 L to 400 L. Do not exceed 2 mg 

total dust loading on the filter.

3. Take a high-volume (4000 L) air sample using a clean sampler and high-volume sampling pump in 

the same area as the personal sample(s) for qualitative identification.

SAMPLE PREPARATION:

4. Mount field samples and blanks on XRD sample holders using appropriate methods for securing the 

sample to the XRD holder.

CALIBRATION AND QUALITY CONTROL:

5. Prepare two suspensions of zinc oxide in 2-propanol by weighing 10 mg and 50 mg of the dry 

powder to the nearest 0.01 mg. Quantitatively transfer to a 1 L glass-stoppered bottle using 1 L 

2-propanol.

6. Suspend the powder in 2-propanol by using an ultrasonic probe or bath for 20 min. Immediately 

move the flask to a magnetic stirrer with thermally-insulated top and add a stirring bar to the 

suspension. Cool the solution to room temperature before withdrawing aliquots.

7. Prepare a series of standard filters using the 10 mg/L and 50 mg/L suspensions. Using appropriate 

pipets, prepare a sufficient number of standards in triplicate to cover the analytical range (or sample 

range if known). Standards at 20 μg, 30 μg, 50 μg, 100 μg, 200 μg, and 500 μg are usually sufficient.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ZINC OXIDE: METHOD 7502, Issue 3, dated 20 October 2015 - Page 3 of 5

8. Mount a filter on the filtration apparatus. Place several mL 2-propanol on the filter surface. Turn off 

the stirrer and shake vigorously by hand. Within a few seconds of setting the bottle down, remove 

the lid and withdraw an aliquot from the center of the suspension. Do not adjust the volume in the 

pipet by expelling part of the suspension. If more than the desired aliquot is withdrawn, return all of 

the suspension to the bottle, rinse and dry the pipet. Transfer the aliquot from the pipet to the filter 

funnel, keeping the tip of the pipet near the surface of the liquid but not submerged.

9. Rinse the pipet with several mL of 2-propanol, draining the rinse into the funnel. Repeat the rinse 

several more times.

10. Apply vacuum and rapidly filter the suspension. Leave vacuum on until filter is dry. Do not wash 

down the sides of the funnel after the deposit is in place to avoid disturbing it. Transfer the filter to 

the XRD sample mount.

11. Perform step scans on the standards using the same diffraction peaks and instrumental conditions 

as for samples (step 16). The resulting intensities for standards, , are normalized in the following 

procedure.

12. Determine the net count, , of the reference specimen before or after each unknown, standard, 

or blank is scanned. Use a diffraction peak of high intensity that can be measured rapidly 

but reproducibly (less than 1% ). Select a convenient normalization scale factor, , which is 

approximately equivalent to the net count for the reference specimen peak. This factor never 

changes and should be used for all measurements made on a particular diffractometer. Calculate 

and record the normalized intensity, , for the analyte or silver peaks on each sample, field blank, 

media blank, and standard:

.

NOTE: Normalizing to the reference specimen intensity compensates for long-term drift in X-ray 

tube intensity. If intensity measurements are stable, the reference specimen may be run less 

frequently. In this case, the net intensities of the analyte, , should be normalized to the most 

recently measured reference intensity.

13. Prepare a calibration graph ( vs. μg of analyte).

NOTE: Poor reproducibility at any given level indicates problems in the sample preparation 

technique and new standards should be made. The data should lie along a straight line. A 

weighted least squares ( weighting) is preferable. Curvature can be eliminated with 

absorption corrections based on the mass absorption coefficient of the analyte [7,8,9].

14. Determine the initial slope, m, of the linear portion of the calibration graph in counts per microgram. 

The intercept, b, of the line with the axis should be approximately zero.

NOTE: A large negative intercept indicates an error in determining the background. This may arise 

from incorrectly measuring the baseline or from interference by another phase at the angle 

of background measurement. A large positive intercept indicates an error in determining the 

baseline or that an impurity is included in the measured peak.

MEASUREMENT:

15. Obtain a qualitative X-ray diffraction scan (broad 2-theta range) of the high-volume respirable 

sample to determine the presence of zinc oxide and any matrix interference. The expected 

diffraction peaks are as follows:

Zinc Oxide Peak (2-Theta Degrees)

Primary Secondary Tertiary

36.26 31.75 34.44

16. Analyze the sample filters by step-scanning the most intense interference-free diffraction peak of 

zinc oxide and determine the integrated intensity. Measure the background on each side of the peak 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ZINC OXIDE: METHOD 7502, Issue 3, dated 20 October 2015 - Page 4 of 5

for one half the time used for peak scanning and add the counts from each side for a total (average) 

background. Determine the 2-theta position of the background for each sample. The net count or 

intensity, , is the difference between the peak integrated count and the total background count. 

The net intensity is normalized as in step 12 to obtain .

17. Scan each field blank over the same 2-theta range used for the analyte. These analyses serve only to 

verify that contamination of the filters has not occurred. The analyte peak should be absent.

CALCULATIONS:

18. The concentration, , of zinc oxide in the air sample is:

, μg/L or mg/m3,

where: = normalized intensity for sample peak, 

b = intercept of calibration graph ( vs. ), 

m = initial slope of calibration graph (counts per microgram), and 

 = air volume sampled (L).

In heavily-loaded samples, particularly those rich in heavy elements, X-ray absorption may 

cause reduced intensities and underestimation of zinc oxide. If this is suspected, an absorption 

correction can be made [8]. The collection filters and blanks are mounted on a smooth metal plate 

(substrate) for XRD quantitation. The substrate should have a non-interfering diffraction peak which 

is measured at the same time as the zinc oxide peak. By measuring the substrate peak on both 

samples and blanks, an absorption correction like that in the silica method (NIOSH method 7500) 

can be made. The absorption correction factor must be calculated for each sample from the formula 

in NIOSH method 7500, which takes into account the diffraction angles for the particular substrate 

chosen. Altree-Williams used a silver filter under the collection filter [7].

EVALUATION OF METHOD:

In a comparison of this method with atomic absorption spectrophotometry [3,4], zinc was determined 

on 15 Gelman DM-800 filters containing added zinc oxide in the range 250 μg to 1000 μg. The average 

percent difference for the 15 pairs was 2.7%. NIOSH method S316 was validated with generated 

samples in the range of 2.4 mg/m3 to 9.9 mg/m3 [2,5,10]. A pooled of 0.088 was found for 18 samples 

of 180 L collected from the aerosol.

REFERENCES:

[1] NIOSH [1977]. Zinc oxide in air: P&CAM 222. In: Taylor DG, ed. NIOSH manual of analytical methods. 

2nd ed. Vol. 1. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

157-A, available as accession No. PB274845 from NTIS [www.ntis.gov].

[2] NIOSH [1977]. Backup data report: zinc oxide fume, No. S316. In: Ten NIOSH analytical methods, 

set 3. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control, 

National Institute for Occupational Safety and Health, available as accession No. PB275834 from 

NTIS [www.ntis.gov]. Unpublished.

[3] Haartz JC, Bolyard ML, Abell MT [1975]. Quantitation of zinc chloride and zinc oxide in airborne 

samples. Unpublished paper presented at the American Industrial Hygiene Conference, 

Minneapolis, MN, June 1–6.

[4] Dollberg DD, Abell MT, Lange BA [1980]. Occupational health analytical chemistry: quantitation 

using x-ray powder diffraction. In: Dollberg DD, Verstuyft AW, eds. Analytical techniques in 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ZINC OXIDE: METHOD 7502, Issue 3, dated 20 October 2015 - Page 5 of 5

occupational health chemistry. Washington, DC: American Chemical Society, ACS Symposium Series 

No. 120, pp. 43–65.

[5] NIOSH [1978]. Zinc oxide fume: Method S316. In: Taylor DG, ed. NIOSH manual of analytical

methods. 2nd ed. Vol. 4. Cincinnati, OH: U.S. Department of Health, Education, and Welfare,

Center for Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH)

Publication No. 78-175, available as accession No. PB83-105437 from NTIS [www.ntis.gov].

[6] NIOSH [1975]. Criteria for a recommended standard: occupational exposure to zinc oxide.

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control,

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 76-104 [www.

cdc.gov/niosh/docs/1970/76-104.html].

[7] Altree-Williams S, Lee J, Mezin NV [1977]. Quantitative x-ray diffractometry on respirable dust

collected on Nuclepore filters. Ann Occup Hyg 20(2):109–126.

[8] Leroux J, Powers CA [1970]. Direct x-ray diffraction quantitative analysis of quartz in industrial dust

films deposited on silver membrane filters. Occup Health Rev 21(1):26–34.

[9] Williams PP [1959]. Direct quantitative diffractometric analysis. Anal Chem 31(11):1842–1844.

[10] NIOSH [1980]. Development and validation of methods for sampling and analysis of workplace

toxic substances. By Gunderson EC, Anderson CC, Smith RH, Doemeny LJ. Cincinnati, OH: U.S.

Department of Health and Human Services, Centers for Disease Control, National Institute for

Occupational Safety and Health, DHHS (NIOSH) Publication No. 80-133, available as accession No.

PB89-182042 from NTIS [www.ntis.gov].

METHOD WRITTEN BY:

D. D. Dollberg, Ph.D, and M. T. Abell, NIOSH/DPSE.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT 7600

AW: 52.00 CAS: 18540-29-9 RTECS: GB6262000 

MW: 99.99 CAS: 1333-82-0 RTECS: GB6650000

METHOD: 7600, Issue 3 EVALUATION: FULL Issue 1: 15 May 1989

Issue 3: 20 October 2015

OSHA: 0.005 mg/m3 (1910.1026); C 0.1 mg/m3 as 

(exceptions to 1910.1026)

NIOSH: 0.0002 mg/m3 (8 h); carcinogen

PROPERTIES: Oxidizing agent

SYNONYMS: Vary depending upon the compound

SAMPLING

SAMPLER: FILTER

(5.0 μm PVC membrane)

FLOW RATE: 1 L/min to 4 L/min

VOL-MIN: 34 L @ 0.005 mg/m3

-MAX: 400 L

SHIPMENT: Routine

SAMPLE

STABILITY: Analyze within 2 weeks [1]

FIELD BLANKS: 2 to 10 field blanks per set

ACCURACY

RANGE STUDIED: 0.05 mg/m3 to 0.2 mg/m3 [2] (22 L 

samples)

BIAS: −5.48%

OVERALL

PRECISION ( ): 0.084 [2]

ACCURACY: ±18.6%

MEASUREMENT

TECHNIQUE: VISIBLE ABSORPTION 

SPECTROPHOTOMETRY

ANALYTE: Chromium-diphenylcarbazone complex

EXTRACTION

SOLUTION: 0.25 mol/L sulfuric acid or solution of 

20 g/L sodium hydroxide and 30 g/L 

sodium carbonate (see steps 4 and 5)

WAVELENGTH: 540 nm; 5 cm path length

CALIBRATION: Standard solutions of potassium 

chromatein 0.25 mol/L sulfuric acid

RANGE: 0.2 μg to 7 μg per sample

ESTIMATED LOD: 0.05 μg per sample

PRECISION ( ): 0.029 @ 0.3 μg to 1.2 μg per sample [3]

APPLICABILITY: The working range is 0.00042 mg/m3 to 3.6 mg/m3 for a 400 L air sample. This method may be used for the 

determination of soluble hexavalent chromium (using the acidic extraction solution) or insoluble hexavalent chromium 

(using the basic extraction solution) [3].

INTERFERENCES: Possible interferences are iron, copper, nickel, and vanadium; 10 μg of any of these causes an absorbance 

equivalent to about 0.02 μg hexavalent chromium due to formation of colored complexes. Interference due to reducing 

agents (e.g., elemental iron, divalent iron) is minimized by alkaline extraction (step 5).

OTHER METHODS: This method combines and replaces NIOSH methods P&CAM 169 [1], S317 [2], and P&CAM 319 [3]; the 

hexavalent chromium criteria document [4] contains a summary of more recent methods for air analysis, wipe analysis, and 

biological monitoring.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT: METHOD 7600, Issue 3, dated 20 October 2015 - Page 2 of 5

REAGENTS:

1. Sulfuric acid,* concentrated (98% mass

fraction), reagent grade.

2. Sulfuric acid, 3 mol/L. Add 167 mL

concentrated sulfuric acid to water in a 1 L

flask; dilute to the mark.

3. Acidic extraction solution, sulfuric acid,

0.25 mol/L. Add 14.0 mL concentrated sulfuric

acid to water in a 1 L flask; dilute to the mark.

4. Sodium carbonate, anhydrous, reagent grade.

5. Sodium hydroxide,* reagent grade.

6. Potassium chromate,* reagent grade.

7. Diphenylcarbazide solution. Dissolve 500 mg

sym-diphenylcarbazide in 100 mL acetone and

100 mL water.

8. Hexavalent chromium standard,* 1000

μg/mL. Dissolve 3.735 g potassium chromate

in deionized water to make 1 L, or use

commercially available solution.

9. Calibration stock solution,* 10 μg/mL. Dilute

1000 μg/mL hexavalent chromium standard

1:100 with deionized water.

10. Basic extraction solution. Dissolve 20 g sodium

hydroxide and 30 g sodium carbonate in

deionized water to make 1 L of solution.

11. Glassware cleaning solution.* Add a volume of

concentrated nitric acid to an equal volume of

deionized water.

12. Nitrogen, purified.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Polyvinyl chloride (PVC) filter, 5.0 μm

pore size, 37 mm diameter in polystyrene

cassette filter holder.

NOTE: Some PVC filters promote reduction

of hexavalent chromium. Check each 

lot of filters for recovery of hexavalent 

chromium standard.

2. Personal sampling pump, 1 L/min to 4 L/min,

with flexible connecting tubing.

3. Vials, scintillation, 20 mL glass, PTFE-lined

screw cap.†

4. Forceps, plastic.

5. Spectrophotometer, UV-visible (540 nm), with

cuvettes, 5 cm path length.

6. Filtration apparatus, vacuum.†

7. Beakers, borosilicate, 50 mL.†

8. Watch glass.†

9. Volumetric flasks, 25 mL, 100 mL, and 1000

mL.†

10. Hotplate, 120 °C to 400 °C.

11. Micropipettes, 10 μL to 1 mL.

12. Centrifuge tubes, 40 mL, graduated, with

plastic stoppers.†

13. Büchner funnel.†

14. Pipettes, TD, 5 mL.†

†

Clean all glassware with the glassware 

cleaning solution and rinse thoroughly before 

use.

SPECIAL PRECAUTIONS: NIOSH considers all hexavalent chromium compounds to be suspect 

occupational carcinogens [4]. Concentrated acids are highly corrosive, and sodium hydroxide is 

caustic. All work with these compounds should be performed in a hood. Use proper protective 

clothing including gloves, safety glasses, and laboratory coat. Potassium chromate is a strong oxidizer 

with risk of fire and explosion upon contact with combustible substances and reducing agents.

SAMPLING:

1. Calibrate the sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate in the range 1 L/min to 4 L/min for a sample size of 34 L to

400 L. Do not exceed 1 mg total dust loading on the filter.

3. Remove the filter from the cassette within 1 h of completion of sampling and place it in a vial to be

shipped to the laboratory. Handle the filter only with forceps. Discard the backup pad.

SAMPLE PREPARATION:

NOTE: There are two sample preparation techniques outlined below. For soluble chromates or 

chromic acid, follow step 4; for insoluble chromate or hexavalent chromium in the presence 

of iron, divalent iron, or other reducing agents, follow step 5.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT: METHOD 7600, Issue 3, dated 20 October 2015 - Page 3 of 5

4. Sample preparation for soluble chromates and chromic acid.

a. Remove the blank and sample filters from the vials, then fold and place them into centrifuge

tubes.

b. Add 6 mL to 7 mL of acidic extraction solution to each tube, cap, and shake to wash all surfaces of

the filter. Allow filter to remain in tube 5 min to 10 min [5].

c. Remove the filter from the tube with plastic forceps, carefully washing all surfaces with an

additional 1 mL to 2 mL of acidic extraction solution. Discard the filters. Start reagent blanks at

this point.

d. Filter the solution through a moistened PVC filter in a Büchner funnel to remove interferences

from suspended dust. Collect the filtrate in a clean centrifuge tube. Rinse the bottle, which

contained the filter, with 2 mL to 3 mL of acidic extraction solution and pour into the funnel.

Rinse the funnel and filter with 5 mL to 8 mL of acidic extraction solution.

e. Add 0.5 mL diphenylcarbazide solution to each centrifuge tube. Bring the total volume in each

centrifuge tube to 25 mL with acidic extraction solution. Shake to mix and allow color to develop

(at least 2 min but no longer than 40 min [5]). Transfer the solution to a clean 5 cm cuvette and

analyze within 40 min of mixing (steps 9, 10, and 11).

5. Sample preparation for insoluble chromates and for hexavalent chromium in the presence of iron or

other reducing agents:

NOTE: If significant amounts of trivalent chromium are expected to be present, degas the sample

solution by bubbling nitrogen through it for 5 min before proceeding and purge the 

headspace above the solution during step 5.a.

a. Remove the PVC filter from the bottle, place it in a 50 mL beaker, and add 5.0 mL of basic

extraction solution. Start reagent blanks at this point. Purge the headspace above the solution 

with nitrogen throughout the extraction process to avoid oxidation of any trivalent chromium. 

Cover the beaker with a watch glass and heat it to near the boiling point on a hotplate with 

occasional swirling for 30 min to 45 min. Do not boil the solution or heat longer than 45 min. Do 

not allow the solution to evaporate to dryness because hexavalent chromium may be lost owing 

to reaction with the PVC filter. An indication that hexavalent chromium has been lost in this 

manner is a brown-colored PVC filter.

b. Cool the solution and transfer it quantitatively with distilled water rinses to a 25 mL volumetric

flask, keeping the total volume about 20 mL.

NOTE: If the solution is cloudy, filter it through a PVC filter in a vacuum filtration apparatus using 

distilled water rinses.

c. Add 1.90 mL of 3 mol/L sulfuric acid to the volumetric flask and swirl to mix.

CAUTION: Carbon dioxide will be evolved causing increased pressure in the flask. Let the solution 

stand for several minutes until vigorous gas evolution ceases.

d. Add 0.5 mL diphenylcarbazide solution, dilute to the mark with distilled water and invert several

times to mix thoroughly. Pour out about one-half of the contents of the flask, stopper the flask 

and shake it vigorously several times, removing the stopper each time to relieve pressure.

NOTE: This step releases bubbles of carbon dioxide which otherwise would cause high and 

erratic readings.

e. Transfer an aliquot of the solution remaining in the flask to a 5 cm cuvette and analyze (steps 9,

10, and 11).

CALIBRATION AND QUALITY CONTROL:

6. Calibrate daily with at least six working standards. Transfer 6 mL to 7 mL of acidic extraction

solution to each of a series of 25 mL volumetric flasks. Pipet 0 mL to 0.7 mL of 10 μg/mL calibration

stock solution into the volumetric flasks. Add 0.5 mL diphenylcarbazide solution to each flask and

sufficient acidic extraction solution to bring the volume to 25 mL. These working standards contain

0 μg to 7 μg hexavalent chromium.

7. Analyze the working standards together with blanks and samples (steps 9, 10, and 11).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT: METHOD 7600, Issue 3, dated 20 October 2015 - Page 4 of 5

8. Prepare a calibration graph [absorbance vs. μg hexavalent chromium].

MEASUREMENT:

9. Set wavelength on the spectrophotometer to 540 nm.

10. Set to zero using an acidic extraction solution reagent blank.

11. Transfer sample solution to a cuvette and record the absorbance.

NOTE 1: A sample containing 1.5 μg hexavalent chromium in 25 mL gives about 0.2 absorbance.

NOTE 2: If the absorbance values for the samples are higher than the standards, dilute using

acidic extraction solution, repeat this step, and multiply the resulting absorbance by the 

appropriate dilution factor.

CALCULATIONS:

12. From the calibration graph, determine the mass of hexavalent chromium in each sample, (μg),

and in the average blank, (μg).

13. Calculate the concentration, (mg/m3), of hexavalent chromium in the air volume sampled, (L).

, μg/L or mg/m3.

NOTE: If the hexavalent chromium concentration is to be reported as chromic acid ( ), multiply 

by 1.92 (MW of chromic acid divided by AW of chromium).

EVALUATION OF METHOD:

P&CAM 169 and S317 are essentially the same method and are suitable for soluble chromate and 

chromic acid. Method S317 was validated with generated samples of chromic acid mist [2,5], and 

P&CAM 169 was tested with field samples [1,6]. P&CAM 319 was developed because a method was 

needed to analyze for insoluble chromates [3]. This method was tested with insoluble chromates in 

matrices such as paints, primer, and ceramic powders [3].

Precision, analytical range, recovery data, etc., for the three methods pooled are as follows:

Total : 0.084

Measurement [1,2,3]: 0.02 to 0.04

Range [3]: 0.5 μg/m3 to 10 μg/m3

Collection efficiency [5]: 94.5%

Sampling rate [1,3]: 1.5 L/min to 2.5 L/min

Stability (two weeks) [1]: 96% recovery

REFERENCES:

[1] NIOSH [1977]. Chromic acid mist in air: P&CAM 169. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. Vol. 1. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, 

Center for Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) 

Publication No. 77-157-A, available as accession No. PB274845 from NTIS [www.ntis.gov].

[2] NIOSH [1977]. Chromic acid and chromates: Method S317. In: Taylor DG, ed. NIOSH manual of 

analytical methods. 2nd ed. Vol. 3. Cincinnati, OH: U.S. Department of Health, Education, and 

Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, DHEW 

(NIOSH) Publication No. 77-157-C, available as accession No. PB276838 from NTIS [www.ntis.gov].

[3] NIOSH [1980]. Hexavalent chromium: P&CAM 319. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. Vol. 6. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health,, DHHS (NIOSH) Publication 

No. 80-125, available as accession No. PB82-157728 from NTIS [www.ntis.gov].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT: METHOD 7600, Issue 3, dated 20 October 2015 - Page 5 of 5

[4] NIOSH [2013]. Criteria for a recommended standard: occupational exposure to hexavalent 

chromium. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) 

Publication No. 2013-128 [www.cdc.gov/niosh/docs/2013-128/].

[5] NIOSH [1977]. Backup data report: chromic acid and chromates, No. S317. By Taylor DG, Kupel 

RE, Bryant JM. In: Documentation of the NIOSH validation tests. Cincinnati, OH: U.S. Department 

of Health, Education, and Welfare, Center for Disease Control, National Institute for Occupational 

Safety and Health, DHEW (NIOSH) Publication No. 77-185, available as accession No. PB274248 from 

NTIS [www.ntis.gov].

[6] Abell MT, Carlberg JR [1974]. A simple reliable method for the determination of airborne hexavalent 

chromium. Am Ind Hyg Assoc J 35(4):229–233.

METHOD REVISED BY:

Martin T. Abell, NIOSH/DPSE; NIOSH method S317 validated under NIOSH Contract CDC-99-74-45.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

SILICA, Respirable Crystalline, by IR (KBr pellet) 7602

SiO2 MW: 60.08 CAS: 14808-60-7 RTECS: VV7330000 (quartz)

 14464-46-1 VV7325000 (cristobalite)

 15468-32-3 VV335000 (tridymite)

METHOD: 7602, Issue 4 EVALUATION: FULL Issue 1: 15 February 1984

Issue 4: 25 July 2017

OSHA: 0.05 mg/m3

NIOSH: 0.05 mg/m3; carcinogens

OTHER OELS: [1,2]

PROPERTIES: solid; d 2.65 g/cm3; crystalline transformation: 

α-quartz to β-quartz @ 573 °C; quartz to tridymite @ 867 °C; 

tridymite to cristobalite @ 1470 °C

SYNONYMS: free crystalline silica; silicon dioxide

SAMPLING

SAMPLER: CYCLONE + PRE-WEIGHED FILTER (PVC), 

with conductive filter holder

FLOW RATE: ~2 to ~4 L/min, specific for the cyclone 

used

VOL-MIN: 400 L

 -MAX: 1000 L (total dust < 4 mg)

SHIPMENT: Routine

SAMPLE

STABILITY: Stable

BLANKS: 3 field blanks minimum per sample set

ACCURACY

RANGE 

STUDIED: 5 to >160 μg

BIAS: Varies depending on matrix effects

OVERALL

PRECISION (SS�rrrr): 0.18 [3]

ACCURACY: ±19% at ≈40-160 μg per sample; ±36% at

≈15 μg per sample [4]

MEASUREMENT

TECHNIQUE: INFRARED SPECTROMETRY

ANALYTE: Quartz / Cristobalite

ASHING: Muffle furnace or RF plasma asher

PELLET: Mix residue with KBr; press 13-mm pellet

IR DETECTION: Scan absorbance between 400 cm-1 and 

1000 cm -1 

CALIBRATION: Certified reference material (CRM) 

respirable crystalline silica (RCS) quartz & 

cristobalite; e.g., NIST SRM 1878b quartz, 

NIST 1879a cristobalite, and/or other RCS

CRMs; diluted in KBr and pressed into 13-

mm pellet

RANGE: 10 to >160 μg 

ESTIMATED LOD: 5 μg 

PRECISION (SS�rr): < 0.15 @ 30 μg per sample

APPLICABILITY: The working range is 0.025 to 0.4 mg/m3 for a 400-L air sample or 0.02 to 0.25 mg/m3 for a 1000-L air sample. 

Quartz and cristobalite have a major IR absorbance band at ≈800 cm-1 and can be measured together if present in samples; 

tridymite can also be measured by IR, but its presence is rare in workplaces [5-9]. 

INTERFERENCES: Amorphous silica, kaolinite, muscovite, talc, albite, vermiculite and other minerals that absorb IR radiation in 

the 800 cm-1 region are potential interferences to RCS measurement [5,10] [Table 1]. Potentially interfering silicates can be 

removed using a phosphoric acid cleanup procedure [11]. Calcite at >20% dust loading can negatively interfere by reacting with 

quartz during muffle ashing [11, 12]. The sample preparation procedures described in this method enable minimization of

interferences.

OTHER METHODS: NIOSH Method 7603 describes IR measurement of respirable crystalline silica (RCS) after redeposition onto a 

filter for analysis (vs. use of a KBr pellet) [13]. X-ray diffraction (e.g., NIOSH Method 7500 [13]) is a complementary method for RCS 

measurement. An international voluntary consensus standard describing the measurement of RCS by IR spectrometry, ASTM 

D7439, has been promulgated [14]. 
RESPIRABLE CRYSTALLINE SILICA BY IR (KBr Pellet): METHOD 7602, Issue 4, dated 25 July 2017 - Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Respirable crystalline silica (RCS) primary 

certified reference materials (CRMs); e.g., 

NIST SRM 1878b (quartz), NIST SRM 1879a 

(cristobalite) [15].

2. Potassium bromide (KBr), IR quality.

3. Kaolinite (kaolin) CRM. 

4. Deionized water, ≥18 MΩ-cm resistivity.

5. 2-Propanol, reagent grade.

6. Ethanol, 95%, for cleaning of sample 

handling equipment.*

7. Hydrochloric acid (HCl), 9% w/w.* Add 25 mL 

conc. HCl (37% w/w) to 50 mL of deionized 

water, cool, and bring to 100 mL with 

deionized water.

8. Calibration stock standard, 0.5% w/w.

Accurately weigh and thoroughly mix 5 g 

KBr (dried overnight at 110  ̊C) with 25 mg 

quartz. Store in a bottle in a desiccator. 

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler:

a. Filter, 37-mm dia., 5-μm pore size, polyvinyl 

chloride (PVC), supported with backup pad

or screen in 2-piece, conductive / staticdissipative filter holder.

b. Cyclone, respirable, designed to operate at

a flow rate of ≈2 or ≈4 L/min, specific for 

each sampling device. 

c. Sampling head holder, for keeping the 

cassette, cyclone and coupler tightly 

together so that air enters only through

cyclone inlet.

2. Personal sampling pumps, for RCS sample 

collection at ≈2 or ≈4 L/min.

3. Infrared spectrometer, double-beam 

dispersive or Fourier transform device, with 4 

cm-1 resolution or better.

4. Laboratory press (10 tons pressure), for 

preparing KBr pellets, with evacuable 13-mm 

KBr pellet die.

5. Vacuum pump, for connecting to laboratory 

press.

6. Low-temperature (RF plasma) asher and 

aluminum weighing pan, or muffle furnace 

and porcelain crucibles.

7. Analytical balance, capable of weighing to 

the nearest 0.001 mg.

8. Mortar and pestle, 50-mm agate or mullite

9. Metal microspatula

10. Non-serrate, non-magnetic forceps

11. Desiccator

12. Camel’s hair brush

13. Glassine paper / weighing boats

14. Membrane filtration apparatus (47-mm)

15. Laboratory oven 

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities, sample 

preparation and analysis. It is essential that suitable gloves, eye protection, laboratory coat, etc., be used, 

especially when working with concentrated acids. Note that KBr is incompatible with strong acids. Ethanol 

and 2-propanol are flammable. Avoid exposure by inhalation of silica dust or acid fumes. Perform sample 

preparation and analysis in a clean, well-ventilated area that is well removed from any possible

contamination. Any skin affected by exposure to acids must be immediately washed with plenty of water. 
RESPIRABLE CRYSTALLINE SILICA BY IR (KBr Pellet): METHOD 7602, Issue 4, dated 25 July 2017 - Page 3 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line. Use a conductive filter 

holder.

2. Sample at ≈2 L/min [16] or ≈4 L/min [17] using a pump set at a specific flow rate for the selected 

cyclone, designed to collect the respirable aerosol fraction (in accordance with ISO 7708[18]), for a 

total sample size of 400 to 1000 L. Take care not to overload the filter.

NOTE 1: Do not allow the sampler assembly to be inverted at any time when using a cyclone. Turning 

the cyclone to anything more than a horizontal orientation may deposit oversized material 

from the cyclone body onto the filter.

NOTE 2: Higher sampling flow rates may be suitable for certain sampler designs [19].

3. After collection of samples, remove the filter holders from the cyclones. Seal the filter holders 

(cassettes) and transport them to the laboratory in a container designed to prevent damage in transit.

SAMPLE PREPARATION:

4. Use one of the following methods to prepare samples and blanks:

a. Low-temperature (RF plasma) asher: Remove the filters from filter holders and place them in clean, 

labelled aluminum dishes. Place the dishes in the low-temperature asher so that sample exposure 

to the plasma is optimized. Ash the samples according to manufacturer’s instructions. Carefully 

bring the asher to atmospheric pressure and remove the dishes.

b. Muffle furnace: For samples expected to contain a significant amount of calcite (>20% of the total 

dust loading), wash filters with 9% w/w HCl per procedure in 4.b.i. For other samples, proceed to 

step 4.b.ii. 

i. Place a 0.5-μm pore size, 47-mm dia. PVC filter in the filtration apparatus. Remove the 37-mm 

dia. sample filter from the filter holder and center atop the 47-mm PVC filter. Clamp a funnel 

over the frit so that the dust deposit on the filter is completely exposed to rinsates. Add 10 mL 

of 9% HCl and 5 mL of 2-propanol; allow to stand for 5 min. Apply vacuum and slowly aspirate 

the acid and alcohol in the funnel. Wash with three successive 10-mL portions of deionized 

water. Release vacuum.

ii. Place 37-mm filter samples and blanks in porcelain crucibles, loosely cover, and ash in muffle 

furnace for 2 h at 600  ̊C (800  ̊C if graphite present). Allow to cool to room temperature.

5. Add approximately 300 mg KBr, weighed to the nearest 0.1 mg and dried overnight at 110  ̊C, directly 

to each sample. Mix the sample residue (remaining from the above ashing procedure) with KBr using a 

pestle and grind / homogenize the mixture with mortar and pestle. Quantitatively transfer the mixture 

to a 13-mm evacuable pellet die using glassine paper and camel’s hair brush. Place the die into the 

laboratory press and apply requisite pressure to prepare the sample KBr pellet. Weigh the finished 

pellet to the nearest 0.1 mg and calculate the ratio of the weight of the finished pellet to the weight of 

KBr initially added. Clean sample handling equipment with ethanol between samples.

CALIBRATION AND QUALITY CONTROL:

6. Prepare at least 5 working standard KBr pellets. Calibration standards shall be prepared from primary

RCS CRMs. 

NOTE: Suitable RCS standards include primary reference materials such as NIST SRMs 1878b (quartz) 

and 1879a (cristobalite) [15]; the use of secondary reference materials as calibration standards is 

not appropriate for IR measurements [12, 20].

a. Weigh, to the nearest 0.001 mg, portions of primary CRMs containing 10 to 200 μg RCS of material.

b. Add an accurately weighed (to the nearest 0.001 mg) 300 mg portion of KBr. Prepare KBr pellet 

samples as in Step 5. Compute the ratio of the weight of finished pellet / weight of solids added. 

c. Measure the absorbance at 800 cm-1 for each KBr pellet calibration standard following the 

analytical procedure below (Step 9). Plot the absorbance vs. mass of SiO2 (μg).
RESPIRABLE CRYSTALLINE SILICA BY IR (KBr Pellet): METHOD 7602, Issue 4, dated 25 July 2017 - Page 4 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

7. If samples were low-temperature ashed (Step 4.a.) and kaolinite is known to be present in the matrix of 

the collected samples, prepare at least five different KBr pellets containing 100 to 600 μg of kaolinite. 

Measure the absorbance at 800 cm-1 (RCS) and at 915 cm-1 (kaolinite). Use the relationship between 

the RCS and kaolinite peak to correct the absorbance value at 800 cm-1 samples containing kaolinite

[21]. 

8. Carry media blanks and filters spiked with known amounts of RCS (e.g., quartz) through the sample 

preparation procedures (Steps 4 and 5) to monitor for contamination and losses. 

NOTE: Knowledge of and training in geochemistry and mineralogy is strongly recommended for users 

of this method. Although many analytical chemists are familiar with IR spectroscopy (e.g., as 

applied to organic analysis), mineralogical samples such as those containing RCS require 

additional knowledge of geochemistry and mineralogy to correctly interpret IR spectra and 

account for matrix interferences and mineral transformations.

MEASUREMENT:

9. Set the infrared spectrometer to absorbance mode and to the appropriate settings for quantitative 

analysis. For each IR scan, place a KBr pellet in the sample holder and obtain a spectrum at 4 cm-1

resolution or better from 400 cm-1 to 1000 cm-1. Rotate the KBr pellet 45 ̊ and scan again. Repeat twice 

more until 4 scans have been obtained. Draw an appropriate baseline under the absorbance band at 

800 cm-1 from approximately 820 to 670 cm-1. Measure the absorbance from the 800 cm-1 peak 

maximum to baseline in absorbance units. Average the four absorbance values for each KBr pellet

sample. 

NOTE: If the peak at 800 cm-1 is small, expand the ordinate to enhance the peak height. 

10. If the sample was ashed at low temperature (Step 4.a.), the presence of kaolinite will be indicated by an 

absorption band with maximum at 915 cm-1. Draw an appropriate baseline under the absorbance 

band at 915 cm-1 from approximately 960 to 860 cm-1. Measure the absorbance from the 915 cm-1 peak 

maximum to baseline.

CALCULATIONS:

11. To correct for the presence of kaolinite (if necessary), use the measured absorbance at 915 cm-1 (Step 

10) and refer to the kaolinite curve (Step 6) to establish the corrected absorbance for RCS at 800 cm-1. 

Use this corrected RCS absorbance value in Step 13. 

12. If correction for kaolinite is not required, use the measured (uncorrected) absorbance at 800 cm-1 to 

determine the weight (Ws, μg) of RCS (e.g., quartz) from the calibration graph. 

13. From the weight of RCS (Ws, μg), calculate its concentration, C (mg/m3

), in the volume of air sampled, V 

(L):

C = [Ws / V], mg/m3

14. If percent RCS (%Q) is of interest, divide the weight of RCS, Ws (μg), by the total sample weight, Wt (μg): 

%Q = [Ws / Wt] × 100

EVALUATION OF METHOD:

From various experimental studies, analytical figures of merit for the KBr pellet IR method for RCS have 

been summarized in Ref. [14]. Filter collection was carried out using respirable cyclone samplers with flow 

rate ≈2 L/min, with sampling volumes ranging from 300 to 1000 L. Following sample ashing with a plasma 

asher or muffle furnace, the remaining material was homogenized with KBr and pressed into a KBr pellet. 

IR spectrometric measurements were then carried out on the as-prepared KBr discs. By comparison of 

sample IR response to the signal from similarly-prepared CRM quartz standards, and with interference 
RESPIRABLE CRYSTALLINE SILICA BY IR (KBr Pellet): METHOD 7602, Issue 4, dated 25 July 2017 - Page 5 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

correction [21], the RCS content was measured. Applicable analytical ranges were from 3 to 900 μg per 

sample, with estimated method detection limits (MDLs) of 5 μg or less. Reported precision estimates, in 

terms of relative standard deviation, were 0.15 or below at RCS mass per sample of 30 μg. To further 

optimize the method, matching CRM particle size distribution to that of RCS samples [20] has been applied 

to minimize the effect of particle size on the IR response. The KBr pellet IR method has been used to 

measure the crystalline silica content of respirable dusts collected at various workplaces, including coal 

mines [7,8,22], construction sites [23], granite quarries [9] and gold mines [24]. Laboratories using the KBr 

disc method have performed successfully in interlaboratory proficiency analytical testing programs 

[3,25,26].

REFERENCES:

[1] ACGIH [2017] TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical 

substances and physical agents and Biological Exposure Indices. Cincinnati, OH: American 

Conference of Governmental Industrial Hygienists.

[2] Institute for Occupational Safety and Health of the German Social Accident Insurance (IFA) [2017]. 

GESTIS international limit values, http://limitvalue.ifa.dguv.de/. 

[3] Harper M, Sarkisian K, Andrew M [2014]. Assessment of respirable crystalline silica using Proficiency 

Analytical Testing results from 2003-2013. J Occup Environ Hyg 11:D157-D163. 

[4] NIOSH [2016]. Development and evaluation of methods (Chapter ME). By Kennedy ER, Fischbach TJ, 

Song R, Eller PM, Hull RD. In: NIOSH manual of analytical methods, 5th ed.; Ashley K, O’Connor PF, eds. 

Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 2014-151. 

[5] Ojima J [2003]. Determining crystalline silica in respirable samples by infrared spectrophotometry in 

the presence of interferences. J Occup Health 45:94-103. 

[6] NIOSH [1977]. Method P&CAM 100 - Quartz in coal dust by infrared spectroscopy. In: Taylor DG, ed. 

NIOSH manual of analytical methods, 2nd ed. Cincinnati, OH: U.S. Department of Health, Education 

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, 

DHEW (NIOSH) Publication No. 77-157.

[7] Larsen DJ, von Loenhoff LJ, Crable JV [1972]. The quantitative determination of quartz in coal dust 

by infrared spectroscopy. Am Ind Hyg Assoc J 33:367-372. 

[8] Dodgson J, Whittaker W [1973]. The determination of quartz in respirable dust samples by infrared 

spectrophotometry – 1: The potassium bromide disc method. Ann Occup Hyg 16:373-387. 

[9] Cares JW, Goldin AS, Lynch JJ, Burgess WA [1973]. The determination of quartz in airborne respirable 

granite dust by infrared spectrophotometry. Am Ind Hyg Assoc J 34:298-305. 

[10] Taylor DG, Nenadic CM, Crable JV [1970]. Infrared spectra for mineral identification. Am Ind Hyg 

Assoc J 31:100-108. 

[11] Talvitie NA [1951]. Determination of quartz in the presence of silicates using phosphoric acid. Anal 

Chem 23:623-626. 
RESPIRABLE CRYSTALLINE SILICA BY IR (KBr Pellet): METHOD 7602, Issue 4, dated 25 July 2017 - Page 6 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[12] NIOSH [2003]. Determination of airborne respirable silica (Chapter R). By Key-Schwartz RJ, Baron PA, 

Bartley DL, Rice FL, Schlecht PC. In: NIOSH manual of analytical methods, 4th ed.; Schlecht PC, 

O’Connor PF, eds. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 94-113. 

[13] NIOSH [2003]. Methods 7603 and 7500. In: NIOSH manual of analytical methods, 4th ed.; Schlecht PC, 

O’Connor PF, eds. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 94-113.

[14] ASTM International [2014]. ASTM D7948, Standard test method for measurement of respirable 

crystalline silica in workplace air by infrared spectrometry. West Conshohocken, PA: ASTM 

International. 

[15] NIST [no date]. Standard reference materials (SRMs). Gaithersburg, MD: U.S. Department of 

Commerce, National Institute of Standards and Technology, https://www.nist.gov. Date accessed: 

July 25, 2017. Technology (NIST), Standard Reference Materials (SRMs); www.nist.gov (Accessed 

August 2016). 

[16] Stacey P, Mecchia M, Verpaele S, Pretorius C, Key-Schwartz R, Mattenklott M [2014]. Differences 

between samplers for respirable dust and the analysis of quartz – An international study. In: Silica 

and Associated Respirable Mineral Particles (ASTM STP 1565); Harper M, Lee T, Eds. W. 

Conshohocken, PA: ASTM International; pp. 73-102. 

[17] Lee T, Lee EG, Kim SW, Chisholm W, Kashon M, Harper M [2012]. Quartz measurement in coal dust 

with high-flow rate samplers – Laboratory study. Ann Occup Hyg 56:413-425. 

[18] ISO [1995]. Air quality – Particle size fraction definitions for health-related sampling. Geneva, 

Switzerland: International Organization for Standardization.

[19] Verpaele S, Jouret J [2013]. A comparison of the performance of samplers for respirable dust in 

workplaces and laboratory analysis for respirable quartz. Ann Occup Hyg 57:54–62. 

[20] Stacey P, Kauffer EM, Moulut J-C, Dion C, Beauparlant M, Fernandez P, Key-Schwartz RJ, Friede B, 

Wake D [2009]. An international comparison of the crystallinity of calibration materials for the 

analysis of respirable α-quartz using x-ray diffraction and a comparison with results from the infrared 

KBr disc method. Ann Occup Hyg 53:639-649. 

[21] Lee T, Chisholm WP, Kashon M, Key-Schwartz RJ, Harper M [2013]. Consideration of kaolinite 

interference correction for quartz measurements in coal mine dust. J Occup Environ Hyg 10:425-434. 

[22] Bandyopadhyay AK [2010]. Determination of quartz content for Indian coals using an FTIR 

technique. Int J Coal Geol 81:73-87. 

[23] Virji MV, Bello D, Woskie SR, Liu XM, Kalil AJ [2002]. Analysis of quartz by FTIR in air samples of 

construction Appl Occup Environ Hyg 17:165-175. 

[24] Verma DK, Sebestyen A, Julian JA, Muir DCF [1992]. A comparison of two methods of sampling and 

analysis for free crystalline silica α-quartz. Appl Occup Environ Hyg 7:758-763. 

[25] Shulman SA, Groff JH, Abell MT [1992]. Performance of laboratories measuring silica in the 

Proficiency Analytical Testing program. Am Ind Hyg Assoc J 53:49-56. 
RESPIRABLE CRYSTALLINE SILICA BY IR (KBr Pellet): METHOD 7602, Issue 4, dated 25 July 2017 - Page 7 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[26] Stacey P, Tylee B, Bard D, Atkinson R [2003]. The performance of laboratories analyzing α-quartz in 

the Workplace Analysis Scheme for Proficiency. Ann Occup Hyg 47:269-277. 

METHOD REVISED BY:

Kevin Ashley, Ph.D., Rosa J. Key-Schwartz, Ph.D., and Martin Harper, Ph.D., CIH, CDC/NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.

Table 1. Minerals potentially encountered and their characteristic IR bands (450-1000 cm-1) 

Mineral Major/Interfering Peaks, cm-1 Identifying Peaks, cm-1

Quartz 800, 780 694, 512, 467

Cristobalite 798 623, 490

Tridymite 789 617, 476

Amorphous silica 800 464

Kaolinite 795, 754 915, 547, 474

Muscovite 800, 750 535, 481

Mullite 837, 748 556, 468

Pyrophyllite 830, 814 948, 477, 457

Albite 788, 746 726, 652, 598, 470

Montmorillonite 797 918, 668, 526, 470

Daphnite 798, 771 667, 610, 539, 467

Anorthite 760, 730 577, 538, 481

Orthoclase 765, 745, 730 645, 593, 540

Talc 797, 778 668, 641, 620

Vermiculite 810, 755 685, 510
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 QUARTZ in Respirable Coal Mine Dust, by IR (Redeposition) 7603

SiO2 MW: 60.08 CAS: 14808-60-7 RTECS: VV7330000 (quartz) 

METHOD: 7603, Issue 4 EVALUATION: FULL Issue 1: 15 May 1989

Issue 4: 25 July 2017

OSHA: 0.05 mg/m3

NIOSH: 0.05 mg/m3; suspect carcinogen

OTHER OELS: [1,2]

PROPERTIES: solid; d 2.65 g/cm3; crystalline transformation: 

α-quartz to β-quartz @ 573 °C; quartz to tridymite @ 867 °C

SYNONYMS: free crystalline silica; silicon dioxide

SAMPLING

SAMPLER: CYCLONE + PREWEIGHED FILTER (PVC), 

with conductive filter holder

FLOW RATE: ~2 to ~4 L/min, specific for the cyclone 

used

VOL-MIN: 300 L @ 0.1 mg/m3

 -MAX: 1000 L (total dust < 4 mg)

SHIPMENT: Routine

SAMPLE

STABILITY: Stable

BLANKS: 3 field blanks minimum per sample set

ACCURACY

RANGE 

STUDIED: <10 to ~500 μg per sample [3-6]

BIAS: Negligible when matrix effects are 

accounted for

OVERALL

PRECISION (SS�rrrr): 0.07 to 0.10 [3-7]

ACCURACY: ±17% to ±33% [5-7] 

MEASUREMENT

TECHNIQUE: INFRARED SPECTROMETRY

ANALYTE: Quartz

ASHING: Muffle furnace or RF plasma asher

REDEPOSIT: Polypropylene filter (47 mm, 0.45 μm pore 

size) [3]

IR DETECTION: Scan absorbance between 450 cm-1 and 

1000 cm -1

CALIBRATION: Primary certified reference material quartz, 

e.g., NIST SRM 1878b, suspended in 2-

propanol; or NIST SRM 2950a (quartz on

filter)

RANGE: 10 to 500 μg quartz per sample [3-5,7]

ESTIMATED LOD: 1 μg [3,4] 

PRECISION (SS�rr): 0.004 to 0.10 @ 50 to 250 μg per sample [3-

6]

APPLICABILITY: The working range is 0.01 to 0.50 mg/m3 for a 1000-L air sample [3,5-7]. The method was originally developed 

specifically for respirable coal mine dust samples [6,8-10]. IR is useful for a matrix such as coal mine dust, where the interferences 

can be predictably removed and minimized.

INTERFERENCES: Amorphous silica, kaolinite, muscovite, talc, albite, vermiculite and other minerals that absorb IR radiation in 

the 800 cm-1 region are potential interferences to RCS measurement [11,12] [Table 1]. Potentially interfering silicates can be 

removed using a phosphoric acid cleanup procedure [13]. Calcite at >20% dust loading can negatively interfere by reacting with 

quartz during muffle ashing [10,14]. The sample preparation procedures described in this method enable minimization of

potential interferences.

OTHER METHODS: NIOSH Method 7602 describes sample preparation and IR measurement of respirable crystalline silica (RCS) by 

means of a KBr pellet [15]. X-ray diffraction (e.g., NIOSH Method 7500 [15]) is a complementary method for quartz measurement by 

IR. MSHA Method P-7 [6] is an equivalent IR method for respirable quartz. An international voluntary consensus standard 

describing the measurement of RCS by IR spectrometry, ASTM D7439, has been promulgated [16].
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 2 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Quartz primary certified reference material 

(CRM); e.g., NIST SRM 1878b or SRM 2950a 

[17]. 

2. Kaolinite CRM; e.g., material meeting 

requirements from USP#

3. Deionized water, ≥18 MΩ-cm resistivity. 

4. 2-Propanol, reagent grade.

5. Ethanol, 95%, for cleaning of sampling 

equipment.*

6. Hydrochloric acid* (HCl) solution, 25% v/v: 

Add 25 mL conc. HCl (37% w/w) to 50 mL 

with deionized water, cool, and bring to 100 

mL with deionized water.

NOTE: Required if calcite is present and 

samples are prepared using a muffle 

furnace.

7. Calibration stock solution (quartz 

suspension in 2-propanol), 15 mg/L: 

Accurately weigh 7.5 mg of quartz and add 

to about 400 mL of 2-propanol in a 500-mL 

volumetric flask. Bring the suspension to the 

mark with 2-propanol. 

8. Kaolinite suspension, 100 mg/mL: 

Accurately weigh 50 mg of dried kaolinite

and add to about 400 mL of 2-propanol in a 

500-mL volumetric flask. Bring the 

suspension to the mark with 2-propanol.

NOTE: Not required if muffle furnace used 

for sample ashing.

9. Desiccant.

10. Oxygen, purified.

#

United States Pharmacopeia Convention

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler:

a. Filters, 37-mm dia., 5-μm pore size, 

polyvinyl chloride (PVC), supported with 

backup pad or screen in 2-piece, 

conductive / static-dissipative filter holder.

b. Cyclone, respirable, designed to operate at

a flow rate of ≈2 or ≈4 L/min, specific for 

each sampling device. 

c. Sampling head holder, for keeping the 

cassette, cyclone and coupler tightly 

together so that air enters only through

cyclone inlet.

2. Personal sampling pumps, for sample 

collection at ≈2 or ≈4 L/min.

3. Infrared spectrometer, double-beam 

dispersive or Fourier transform device, with 4 

cm-1 resolution or better.

4. Filtration apparatus (for redepositing sample 

after ashing), consisting of fritted support, 

side-arm vacuum flask and glass filtration 

funnel with bakelite base, 1.0 cm internal 

diameter. The base should seal tightly to the 

fritted support to prevent leakage.

5. Filters for standards and redeposition, 47-

mm, 0.45-μm pore size, polypropylene (PP).

6. Glass fiber filters, 25-mm dia., for backup 

during filtration. 

7. Filtration funnel for treating filters (required 

only if using a muffle furnace): same as in 6. 

above except with funnel i.d. about 1.6 cm 

and with 0.5-μm pore size 37-mm dia. PVC 

filters to recollect residue.

8. Low-temperature (RF plasma) asher and 

aluminum weighing pan, or muffle furnace 

and 10-mL porcelain crucibles (with covers). 

9. Analytical balance, capable of weighing to 

the nearest 0.001 mg.

10. Ultrasonic bath. 

11. Desiccator. 

12. Beakers, 50-mL. 

13. Magnetic stirrer with insulated top. 

14. Volumetric flasks with stoppers, 500-mL.

15. Tweezers and metal spatulas.

16. Petri dishes, for 47-mm dia. filters.

17. Wash bottles.

18. Pipets, various sizes as required.
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 3 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities, sample 

preparation and analysis. It is essential that suitable gloves, eye protection, laboratory coat, etc., be used, 

especially when working with concentrated acids. Ethanol and 2-propanol are flammable. Avoid exposure

by inhalation of silica dust or acid fumes. Perform sample preparation and analysis in a clean, wellventilated area that is well removed from any possible contamination. Any skin affected by exposure to 

acids must be immediately washed with plenty of water. 

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line. Use conductive filter 

holders.

2. Pre-weigh each filter to the nearest 0.01 mg.

3. Sample at ≈2 L/min [18] or ≈4 L/min [5] using a pump set at a specific flow rate for a selected cyclone,

designed to collect the respirable aerosol fraction (in accordance with ISO 7708[19]), for a total sample 

size of 400 to 1000 L. Take care not to overload the filter.

NOTE 1: Do not allow the sampler assembly to be inverted at any time when using a cyclone. Turning 

the cyclone to anything more than a horizontal orientation may deposit oversized material 

from the cyclone body onto the filter.

NOTE 2: Higher sampling flow rates may be suitable for certain sampler designs [20].

4. After collection of samples, remove the filter holders from the cyclones. Seal the filter holders 

(cassettes) and transport them to the laboratory in a container designed to prevent damage in transit.

SAMPLE PREPARATION:

5. Remove the filters from filter holders and, after preconditioning to ambient conditions, re-weigh the 

filters to the nearest 0.01 mg; the difference is the sample weight, Ws (μg).

6. Use one of the following methods to prepare samples and blanks:

a. Low-temperature (RF plasma) asher: Transfer the weighed filters to clean 50-mL beakers. Place the 

beakers in the low-temperature asher so that sample exposure to the plasma is optimized. Ash the 

samples for 2 h at 300 W RF power and oxygen flow rate of 75 mL/min, according to manufacturer’s 

instructions. Carefully bring the asher to atmospheric pressure, remove the beakers, and add 15 mL 

2-propanol to each beaker. 

b. Muffle furnace: For samples expected to contain a significant amount of calcite (>20% of the total 

dust loading), wash filters with 25% v/v HCl using the filtration apparatus with the 1.6-cm i.d. glass 

funnel. For other samples, proceed to step 6.b.ii. 

i. Place a 25-mm glass fiber filter over the frit area of the filtration funnel, then place a 0.45-μm 

pore size, 37-mm dia. PP filter atop the glass fiber filter. Clamp the funnel over the frit and 

ensure that there is no leakage. Using tweezers and gloved hands, take the weighed sample 

filter and fold it in half with the collection surface inside, then fold again into quarters, and place 

the folded filter into the funnel. Add 10 mL of 25% HCl (v/v), then 5 mL of 2-propanol; allow to 

stand for 5 min. Apply vacuum and slowly aspirate the acid and alcohol in the funnel. Press the 

folded sample filter onto the surface of the collection filter if necessary to remove all liquid. 

Release vacuum, remove both filters and place in a porcelain crucible. Allow to air-dry.

ii. Place filter samples and blanks in porcelain crucibles, loosely cover, and ash in muffle furnace for 

2 h at 600  ̊C (800  ̊C if graphite present). Allow to cool to room temperature.

iii. Add ~3 mL 2-propanol to the ash within each crucible, scrape the crucible to loosen all particles, 

and quantitatively transfer to contents to a 50-mL beaker. Wash the crucible several times with 

2-propanol and add the wash to the beaker. Add 2-propanol to the beaker to bring the volume 

to ~15 mL.

7. Redeposit the sample residue as follows, using the filtration apparatus with the 1.0 cm dia. funnel. 

With a slight vacuum applied, place a 25-mm glass fiber filter on the fritted base. Cut a 47-mm 

0.45-μm PP filter in half. Superimpose one half over the other, glossy sides down, and place on the 
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 4 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

glass fiber filter. (The lower half of the PP filter serves as a blank for use in the reference beam of the IR 

spectrometer.) Position the filter funnel, apply clamp, turn off vacuum and ensure that there is no

leakage. Place sample beakers from step 6.b.iii into ultrasonic bath for at least 30 sec to ensure 

homogeneous dispersion. Remove each beaker, wipe off excess water from the outside of the beaker, 

and transfer the slurry to the filtration funnel. Reapply vacuum and, during filtration, rinse the beaker 

twice with 2-propanol to ensure quantitative transfer of entire contents to the funnel. Control the 

filtration rate to keep the liquid near the top of the funnel during rinsing to avoid disturbing the 

deposit. When the depth of liquid in the funnel reaches about 4 cm above the filter, gently rinse the 

inside of the funnel with 2-propanol and complete filtration. Remove clamp and funnel, taking care 

not to disturb the deposit. Release the vacuum. Define the deposit area by marking around the 

circumference using a pencil or scriber. (This is especially important for standards or light-colored 

samples.) Place the PP filter halves in Petri dishes and allow to air dry.

CALIBRATION AND QUALITY CONTROL:

8. Prepare at least 5 working standard quartz filters. Calibration standards shall be prepared from 

primary CRMs of known purity, particle size range and homogeneity. 

NOTE: Suitable quartz standards include primary reference materials such as NIST SRM 1878b or 2950; 

the use of secondary reference materials as calibration standards is not appropriate for IR 

measurement [14]. 

a. Place the flask containing the calibration stock solution in an ultrasonic bath for 30-45 min. 

b. Move the flask to a magnetic stirrer and stir slowly while the flask cools to room temperature. 

Continue slow stirring while preparing calibration standards.

c. Mount a PP filter in the filtration apparatus in the same manner used to redeposit the samples. Add 

5 mL of 2-propanol to the funnel. Withdraw an aliquot of the quartz suspension from the center of 

the flask. Draw liquid to the mark but do not attempt to adjust volume by draining pipet. Carefully 

wipe the outside of the pipet, then drain the suspension into the filter funnel. Rinse down the 

inside wall of the pipet with a few mL of 2-propanol, draining the washings into the funnel. Apply 

vacuum to complete the filtration. Prepare quartz standards in this manner to cover the range 10 

to 250 μg per filter.

d. Carry an additional set of these calibration standards and media blanks through steps 6 and 7 to 

monitor for contamination and losses.

NOTE: Accuracy depends on obtaining uniform deposition of samples and calibration standards 

across the filter surface and obtaining reproducible aliquots from the quartz suspension. 

This requires skill and practice. The quartz calibration curve should be prepared before 

analyzing samples as a check on the analyst’s ability to prepare uniform deposits. 

Repeatability should be <10% on replicate calibration standards with >40 μg of quartz.

9. Perform IR scans for each standard filter using the other half of the filter in the reference beam (see 

Measurement section). Construct a calibration graph of absorbance at 800 cm-1 vs. μg of quartz per 

filter. The plot should be linear and pass through the origin.

10. If samples were low-temperature ashed and kaolinite is known to be present in the matrix of the 

collected samples, prepare at least five different loadings of PP filters containing 100 to 600 μg of 

kaolinite. Measure the IR absorbance of kaolinite standards from 1000 cm-1 to 650 cm-1. Prepare a 

graph with absorbance at 915 cm-1 as ordinate vs. absorbance at 800 cm-1 as abscissa. Use the 

relationship between the quartz and kaolinite peak to correct the absorbance value at 800 cm-1

samples containing kaolinite [20]. 

11. Carry media blanks and filters spiked with known amounts of quartz through the sample preparation 

procedures to monitor for contamination and losses. 

NOTE: Knowledge of and training in geochemistry and mineralogy is strongly recommended for users 

of this method. Although many analytical chemists are familiar with IR spectroscopy (e.g., as 

applied to organic analysis), mineralogical samples such as those containing RCS require 
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 5 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

additional knowledge of geochemistry and mineralogy to correctly interpret IR spectra and 

account for matrix interferences and mineral transformations.

MEASUREMENT:

12. Set the infrared spectrometer to double-beam absorbance mode and to the appropriate settings for 

quantitative analysis. For each IR scan, place a PP sample filter half in the sample beam and a PP blank 

filter half in the reference beam, and obtain a spectrum at 4 cm-1 resolution or better from 450 cm-1 to 

1000 cm-1. For each PP filter sample, center the IR beam on filter deposit.

13. Draw an appropriate baseline under the absorbance band at 800 cm-1 from approximately 820 to 670 

cm-1. Measure the absorbance from the 800 cm-1 peak maximum to baseline in absorbance units. 

NOTE: If the peak at 800 cm-1 is small, expand the ordinate to enhance the peak height. 

14. If the sample was ashed at low temperature, the presence of kaolinite will be indicated by an 

absorption band with maximum at 915 cm-1. Draw an appropriate baseline under the absorbance 

band at 915 cm-1 from approximately 960 to 860 cm-1. Measure the absorbance from the 915 cm-1 peak 

maximum to baseline.

15. Analyze blanks and, if necessary, correct baseline.

CALCULATIONS:

16. To correct for the presence of kaolinite (if necessary), use the measured absorbance at 915 cm-1 and 

refer to the kaolinite curve to establish the corrected absorbance for quartz at 800 cm-1 [21]. Use this 

corrected absorbance value at 800 cm-1 in calculating the airborne quartz concentration. 

17. If correction for kaolinite is not required, use the measured (uncorrected) absorbance at 800 cm-1 to 

determine the weight (Ws, μg) quartz from the calibration graph. 

18. From the weight of quartz (Ws, μg), calculate its concentration, C (mg/m3

), in the volume of air 

sampled, V (L):

C = [Ws / V], mg/m3

19. If percent quartz (%Q) is of interest, divide the weight of quartz, Ws (μg), by the total sample weight, Wt

(μg): 

%Q = [Ws / Wt] × 100

EVALUATION OF METHOD:

The method is based on MSHA method P-7, which was collaboratively tested [4] and optimized [6]. 

Interlaboratory evaluation included a ruggedization step to test the effects of the use of muffle furnace or 

plasma asher, amount of calcite or kaolinite in the sample, ashing time, pH of solvent used to remove 

calcite, shipment of samples, and other factors. None of these contributions was found to have a 

statistically significant effect. Filter collection has been carried out using respirable cyclone samplers with 

flow rate ≈2 L/min [4] and ≈4 L/min [5], with sampling volumes ranging from 300 to 1000 L; analysis was 

subsequently carried out using the redeposition method. Applicable analytical ranges were from 3 μg [5] 

to 500 μg quartz [4,6] per sample, with estimated method detection limits (MDLs) of 1-5 μg or even less [3]. 

Reported precision estimates, in terms of relative standard deviation, were 0.10 or below at quartz mass 

per sample of 20-400 μg [3-6]. The IR redeposition method has been used extensively to measure the 

quartz content of respirable dusts collected in coal mines [3,8-10,21,22]. Laboratories using the IR 

redeposition method have performed successfully in interlaboratory proficiency analytical testing 

programs [7, 23, 24].
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 6 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] ACGIH [2017] TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical 

substances and physical agents and Biological Exposure Indices. Cincinnati, OH: American 

Conference of Governmental Industrial Hygienists.

[2] Institute for Occupational Safety and Health of the German Social Accident Insurance (IFA) [2017]. 

GESTIS international limit values, http://limitvalue.ifa.dguv.de/. 

[3] Farcas D, Lee T, Chisholm WP, Soo J-C, Harper M [2016]. Replacement of filters for respirable quartz 

measurement in coal mine dust by infrared spectroscopy. J Occup Environ Hyg 13:D17-D22. 

[4] Anderson CC [1983]. Collaborative test of two methods for determining free silica in airborne dust. 

Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 83-124. 

[5] Lee T, Lee EG, Kim SW, Chisholm WP, Kashon M, Harper M [2012]. Quartz measurement in coal dust 

with high flow rate samplers – Laboratory study. Ann Occup Hyg 56:413-425. 

[6] Mine Safety and Health Administration (MSHA) [2013]. Infrared determination of quartz in respirable 

coal mine dust; Method No. P-7. Pittsburgh, PA: MSHA. 

[7] Harper M, Sarkisian K, Andrew M [2014]. Assessment of respirable crystalline silica using Proficiency 

Analytical Testing results from 2003-2013. J Occup Environ Hyg 11:D157-D163. 

[8] Larsen DJ, von Loenhoff LJ, Crable JV [1972]. The quantitative determination of quartz in coal dust 

by infrared spectroscopy. Am Ind Hyg Assoc J 33:367-372. 

[9] Freedman RW, Toma SZ, Lang HW [1974]. On-filter analysis of quartz in respiratory coal dust by 

infrared absorption and x-ray diffraction. Am Ind Hyg Assoc J 35:411-418. 

[10] NIOSH [1977]. Method P&CAM 100 - Quartz in coal dust by infrared spectroscopy. In: Taylor DG, ed. 

NIOSH manual of analytical methods, 2nd ed. Cincinnati, OH: U.S. Department of Health, Education 

and Welfare, Center for Disease Control, National Institute for Occupational Safety and Health, 

DHEW (NIOSH) Publication No. 77-157. 

[11] Taylor DG, Nenadic CM, Crable JV [1970]. Infrared spectra for mineral identification. Am Ind Hyg 

Assoc J 31:100-108. 

[12] Ojima J [2003]. Determining crystalline silica in respirable samples by infrared spectrophotometry in 

the presence of interferences. J Occup Health 45:94-103. 

[13] Talvitie NA [1951]. Determination of quartz in the presence of silicates using phosphoric acid. Anal 

Chem 23:623-626. 

[14] NIOSH [2003]. Determination of airborne respirable silica (Chapter R). By Key-Schwartz RJ, Baron PA, 

Bartley DL, Rice FL, Schlecht PC. In: NIOSH manual of analytical methods, 4th ed.; Schlecht PC, 

O’Connor PF, eds. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 94-113. 

[15] NIOSH [2003]. Methods 7603 and 7500. In: NIOSH manual of analytical methods, 4th ed.; Schlecht PC, 

O’Connor PF, eds. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication 

No. 94-113. 
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 7 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[16] ASTM International [2014]. ASTM D7948, Standard test method for measurement of respirable

crystalline silica in workplace air by infrared spectrometry. West Conshohocken, PA: ASTM

International.

[17] NIST [no date]. Standard reference materials (SRMs). Gaithersburg, MD: U.S. Department of

Commerce, National Institute of Standards and Technology, https://www.nist.gov. Date accessed:

July 25, 2017.

[18] Stacey P, Mecchia M, Verpaele S, Pretorius C, Key-Schwartz R, Mattenklott M [2014]. Differences

between samplers for respirable dust and the analysis of quartz – An international study. In: Silica

and associated respirable mineral particles (ASTM STP 1565); Harper M, Lee T, eds. W.

Conshohocken, PA: ASTM International; pp. 73-102.

[19] ISO [1995]. Air quality – Particle size fraction definitions for health-related sampling. Geneva,

Switzerland: International Organization for Standardization.

[20] Verpaele S, Jouret J [2013]. A comparison of the performance of samplers for respirable dust in

workplaces and laboratory analysis for respirable quartz. Ann Occup Hyg 57:54–62.

[21] Lee T, Chisholm WP, Kashon M, Key-Schwartz RJ, Harper M [2013]. Consideration of kaolinite

interference correction for quartz measurements in coal mine dust. J Occup Environ Hyg 10:425-434.

[22] Ainsworth, SM [2005]. Infrared analysis of respirable coal mine dust for quartz: Thirty five years. J

ASTM Int 2(4):1-13, DOI: 10.1520/JAI12231.

[23] Shulman SA, Groff JH, Abell MT [1992]. Performance of laboratories measuring silica in the

Proficiency Analytical Testing program. Am Ind Hyg Assoc J 53:49-56.

[24] Stacey P, Tylee B, Bard D, Atkinson R [2003]. The performance of laboratories analyzing α-quartz in

the Workplace Analysis Scheme for Proficiency. Ann Occup Hyg 47:269-277. 

METHOD REVISED BY:

Kevin Ashley, Ph.D., Rosa J. Key-Schwartz, Ph.D., and Martin Harper, Ph.D., CIH, CDC/NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
QUARTZ IN RESPIRABLE COAL DUST BY IR (Redeposition): METHOD 7603, Issue 4, dated 25 July 2017 - Page 8 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 1. Minerals potentially encountered and their characteristic IR bands (450-1000 cm-1) 

Mineral Major/Interfering Peaks, cm-1 Identifying Peaks, cm-1

Quartz 800, 780 694, 512, 467

Cristobalite 798 623, 490

Tridymite 789 617, 476

Amorphous silica 800 464

Kaolinite 795, 754 915, 547, 474

Muscovite 800, 750 535, 481

Mullite 837, 748 556, 468

Pyrophyllite 830, 814 948, 477, 457

Albite 788, 746 726, 652, 598, 470

Montmorillonite 797 918, 668, 526, 470

Daphnite 798, 771 667, 610, 539, 467

Anorthite 760, 730 577, 538, 481

Orthoclase 765, 745, 730 645, 593, 540

Talc 797, 778 668, 641, 620

Vermiculite 810, 755 685, 510
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT

by Ion Chromatography 7605

Cr(VI) MW: 52.00 (Cr); 99.99 (CrO3) CAS: 18540-29-9 RTECS: GB6262000

METHOD: 7605, Issue 2 EVALUATION: FULL Issue 1: 15March 2003

Issue 2: 10 March 2016

OSHA: 0.005 mg/m3 (1910.1026); C 0.1 mg/m3 as 

CrO3 (exceptions to 1910.1026) 

NIOSH: 0.0002 mg/m3 (8h); carcinogen

PROPERTIES: oxidizing agent

SYNONYMS: Vary depending upon the compound; chromate commonly used; “chrome six.” 

SAMPLING

SAMPLER: FILTER (5.0-μm PVC membrane)

FLOW RATE: 

VOL-MIN: 

 -MAX:

SHIPMENT: 

SAMPLE

STABILITY: 

BLANKS: 

1 to 4 L/min

1 L @ 0.05 mg/m3

400 L

Routine. Can be shipped cold when

deemed necessary

Stable for 2 weeks at room temperature. 

Stable for 4 weeks if stored in refrigerator. 

2 to 10 field blanks per set

ACCURACY 

RANGE 

STUDIED: 0.05 to 120 μg Cr(VI) [1,2]

BIAS: -1.6% [2]

OVERALL

PRECISION (SS�rrrr): 0.07

ACCURACY: ± 17.4% (0.6 - 960 g/m3

) [1]

MEASUREMENT

TECHNIQUE:

ANALYTE: 

EXTRACTION: 

INJECTION 

 VOLUME: 

COLUMN: 

MOBILE 

PHASE:

FLOW RATE: 

POST-COLUMN 

 REAGENT:

DETECTOR: 

Ion chromatography with post-column

derivatization and UV detection.

CrO4

2--diphenylcarbazide (DPC) complex 

5 mL 2% NaOH- 3% Na2CO3. Dilute to 25 

mL after heating.

100 μL

Dionex NG1 Guard, HPIC- AS7 Separator or 

equivalent

250 mM (NH4)2SO4 /100 mM NH4 OH

1.0 mL/min; 0.7 mL/min post column

2.0 mM DPC + 10% MeOH +1N H2SO4

UV @ 540 nm

CALIBRATION: Standard solutions of K2Cr2O7 in 

NaOH-Na2CO3 buffer

RANGE: 0.05 to 20 μg per sample

ESTIMATED LOD: 0.02 μg per sample [3]

PRECISION (SS�rr): 0.015 @ 0.5 to 5 μg/sample [3]

APPLICABILITY: The working range is 0.00025 to 0.1 mg/m3 for a 200-L air sample. This method can be used for the determination 

of Cr(VI) using 2% NaOH - 3% Na2CO3 for extraction.

INTERFERENCES: Possible interferences are iron, copper, nickel, and vanadium; 10 μg of any of these causes an 

absorbance equivalent to approximately 0.02 μg Cr(VI) due to formation of colored complexes. Interference due to reducing 

agents (e.g., Fe, Fe++) is minimized by alkaline extraction (step 5). 

OTHER METHODS: Method 7703 is a field portable method for Cr(VI). OSHA Method W4001 is for the measurement of Cr(VI) 

in wipe samples [4]. OSHA Method ID-215 is applicable to measurement of Cr(VI) and employs precipitation to reduce Cr(III) 

oxidation. [5]. A similar air method is ISO 16740 [6]. EPA Method 218.6 is for water matrices [7]. Method 7604 is also specific 

for hexavalent chromium, using ion chromatography, but has a higher detection limit than 7605. 
CHROMIUM, HEXAVALENT by Ion Chromatography: METHOD 7605, Issue 2, dated 10 March 2016 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Sulfuric acid, conc. (98 % w/w).*

2. Ammonium hydroxide, conc. (28 %).*

3. Ammonium sulfate monohydrate, reagent

grade.

4. Sodium carbonate, anhydrous.

5. Sodium hydroxide, reagent grade.*

6. Methanol, HPLC grade.*

7. 1,5-Diphenylcarbazide, reagent grade.

8. Potassium dichromate or potassium

chromate.* Dry at 100 °C and store in a

desiccator.

9. Post-Column Derivatizing Reagent:

Diphenylcarbazide solution. Dissolve 500 mg

1,5-diphenylcarbazide in 100 mL HPLC-grade

methanol. While stirring, add 500 mL water

containing 28 mL of conc. sulfuric acid.

Dilute to a final volume of one liter with

water. This reagent is stable for 4 - 5 days.

Prepare in one-liter quantities, as needed.

10. Cr(VI) standard, 1000 μg/mL. Dissolve 2.829 g

potassium dichromate in deionized water to

make one liter, or use commercially available

solution.

NOTE: 3.731 g K2CrO4 can also be used.

11. Calibration stock solution, 1.0 μg/mL. Dilute

1000 μg/mL Cr(VI) standard 1 :1000 with

deionized water.

12. Filter extraction solution, 2% NaOH-3%

Na2CO3. Dissolve 20 g NaOH and 30 g Na2CO3

in deionized water to make one liter of

solution.

13. Eluent (mobile phase); 250 mM ammonium

sulfate/200 mM ammonium hydroxide.

Dissolve 33 g ammonium sulfate in

approximately 500 mL distilled water and

add 6.5 mL conc. ammonium hydroxide.

Dilute to one liter with distilled water and

mix.

14. Nitrogen, pre-purified.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: polyvinyl chloride (PVC) filter, 5.0- μm

pore size, 37 -mm diameter in polystyrene

cassette filter holder.

NOTE: Some PVC filters promote reduction of

Cr(VI). Check each lot of filters for 

recovery of Cr(VI) standard.

2. Personal sampling pump, 1 to 4 L/min, with

flexible connecting tubing.

3. Vials, scintillation, 20-mL glass,

polytetrafluoroethylene (PTFE)-lined screw

cap.**

4. Forceps, nonmetallic.

5. Gloves, polypropylene or latex.

6. Liquid chromatography apparatus consisting

of autosampler; pump; columns ; postcolumn reagent delivery system with 2.2-m

high pressure tubing mixing/reaction loop

with 1 min a water bath at 32 °C ± 3 °C; and UV 

detector.

7. Columns: NG1 (Dionex Corp.) or equivalent

guard column; HPIC-AS7, 4 x 250-mm (Dionex

Corp.) separator column (or equivalent);

8. Filtration apparatus, PTFE luer lock style filter

syringe.

9. Beakers, borosilicate, 50 -mL.**

10. Watchglass.**

11. Volumetric flasks, 25-, 100-, and 1000-mL.**

12. Oven at 107°C, not to exceed 115°C.

NOTE: Hot plate or ultrasonic bath can be 

used.

13. Micropipettes, 10-μL to 0.5-mL.

14. Pipettes, TD 5-mL.**

15. Bagged refrigerant.

**Clean all glassware with 1:1 HNO3:H2O and

rinse thoroughly before use

SPECIAL PRECAUTIONS: Many chromate compounds are suspected human carcinogens [8]. All sample 

preparation should be performed in a hood. Concentrated acids and bases are toxic and corrosive. When 

working with concentrated acids and bases, wear protective clothing. Ammonium hydroxide is a 

respiratory irritant. Methanol is flammable and toxic.
CHROMIUM, HEXAVALENT by Ion Chromatography: METHOD 7605, Issue 2, dated 10 March 2016 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLING:

1. Calibrate the sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate in the range 1 to 4 L/min for a sample size of 1 to 400 L. Do

not exceed 1 mg total dust loading on the filter.

3. Filters can be left in the cassettes for shipping to the lab, but to minimize sample contamination

during shipping, remove the filter from the cassette within one hour of completion of sampling and

place it in a vial to be shipped to the laboratory. Handle the filter only with gloved hands and

nonmetallic forceps. Discard the backup pad. As a precaution, it is recommended to ship the samples

with bagged refrigerant.

SAMPLE PREPARATION:

4. Don a clean pair of disposable plastic gloves (to prevent sample contamination). Using forceps,

transfer the PVC filter to a 50-mL beaker, and add 5.0 mL filter extraction solution, 2% NaOH/3%

Na2CO3. Start media blank s at this point.

NOTE 1: If significant amounts of Cr[III] are expected to be present in the samples, either (a) degas the

sodium hydroxide/sodium carbonate extraction solution by bubbling nitrogen through it for 

5 min. before proceeding, or ( b) use a precipitation reagent [1].

NOT E 2: If only soluble chromates are of interest, use ammonium sulfate buffer in place of carbonate 

extraction solution [9,10].

5. Cover the beaker with a watchglass and heat it to near the boiling point (100°C to 115°C) in an oven

with occasional swirling for 45 min. Do not boil the solution. Longer heating times (up to 90 minutes)

may be necessary for some samples (e.g., paint spray). Do not allow the solution to evaporate to

dryness because hexavalent chromium may be lost due to reaction with the PVC filter and/or cocollected aerosol constituents. An indication that hexavalent chromium has been lost in this manner is

a brown-colored PVC filter.

NOTE: A hot plate, heater block, or ultrasonic bath can also be used for this step [9,11].

a. Cool the solution and transfer it quantitatively with distilled water rinses to a 25-mL volumetric

flask. Bring to volume with distilled water.

NOTE: If the solution is cloudy, filter an aliquot through a PTFE luer lock style filter attached to a

syringe.

b. Transfer an aliquot of the solution to the appropriate vial for the chromatograph’s autosampler and

analyze (steps 9 through 13).

CALIBRATION AND QUALITY CONTROL:

6. Calibrate daily with at least six working standards. Transfer 5 mL of extraction solution to each of a

series of 25-mL volumetric flasks. Pipet known volumes (0 to 5 mL) of calibration stock solution (1.0

μg/mL) into the volumetric flasks. For higher standards, pipet 10 - 20 μL of the 1000 μg/mL

concentrated stock and bring the volume to 25 mL with distilled water. These working standards

contain 0 to 20 μg Cr(VI) per sample.

7. Analyze the working standards together with blanks and samples (steps 9 through 13).

8. Prepare a calibration graph [instrument response vs. μg Cr(VI)].

MEASUREMENT:

9. Set wavelength on the detector to 540 nm.

10. Set the liquid chromatograph to manufacturer's recommendations and parameters given on page

7605-1. With a mobile phase flow rate of 1.0 mL/min., a post-column reagent flow rate of 0.7 mL/min.,

and a 2.2-m post-column tube, the derivative retention time should be approximately 3.7 - 4.7

minutes.
CHROMIUM, HEXAVALENT by Ion Chromatography: METHOD 7605, Issue 2, dated 10 March 2016 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NOTE: If the instrument response for the samples is higher than the standards, dilute using a 1:5 

dilution of extraction solution:water to maintain a constant ionic strength; repeat the analysis; 

and multiply the measured concentration by the appropriate dilution factor. Alternatively, 

inject a smaller volume and multiply by the appropriate factor.

11. After the analysis is complete, flush the entire system with ASTM Type II water for at least one hour at

mL/min. with all columns on line. Remove the columns and continue flushing for an additional two

hours. Flush the autosampler with several injections of water. Leaving the columns in line while the

system is idle is not recommended.

CALCULATIONS

12. From the calibration graph, determine the mass of Cr(VI) in each sample, W (μg), and in the average

blank, B (μg).

13. Calculate concentration, C (mg/m3

), of Cr(VI) in the air volume sampled, V (L):

CC = WW − BB

VV , mmmm/mm3

NOTE: μg/L ≅ mg/m3

EVALUATION OF METHOD: 

This method was evaluated in the laboratory with spiked filters and a certified reference material 

containing a known loading of Cr(VI). This certified reference material (CRM) is European Commission, 

Institute for Reference Materials and Measurements (EC/IRMM) CRM 545, Cr(VI) and Cr(total) in welding 

dust loaded on a glass fiber filter [12]. This method was evaluated for extraction efficiency over the 

concentration range of 0.15 - 5 μg/sample testing two brands of filters, SILICAL® and GLA-5000TM[3]. In these 

experiments, test atmospheres were not generated; instead, Cr(VI) as the dichromate was fortified on the 

face of the sample filters, then 240 L of air with 35% relative humidity was pulled through at 1 L/min. A 30-

day storage study using both types of filters was conducted at 1.5 μg/sample (30x LOQ) at ambient room 

temperature and 4 °C. The average recovery of the stored samples was 94 .8%. The estimation of the limit 

of detection and limit of quantitation (LOD/LOQ ) were determined analyzing a series of liquid standards. 

The LOD and LOQ, 0.02 μg/sample and 0.07 μg/sample, respectively were calculated by Burkart's method 

[13 ].

To fully evaluate this method, a field-study was conducted in which side- by-side samples were taken to 

measure exposures to Cr(VI) during spray-painting and electroplating operations. These samples were 

analyzed subsequently by 4 different methods (NIOSH Method 7605, 7703, 7300 and OSHA ID-215) [1]. 

NIOSH method 7300 was used to measure total chromium. The results from the other 3 methods correlated 

very well showing no statistical difference among the 3 methods. Recoveries of 98.4 (±3.4) % were 

obtained for CRM 545 (n = 6) [2].

REFERENCES:

[1] Boiano JM, Wallace ME, Sieber WK, Groff JH, Wang J, Ashley K [2000]. Comparison of three sampling

and analytical methods for the determination of airborne hexavalent chromium. J Environ Monit

2:329- 333.

[2] Ashley K, Andrews RN, Cavazos L, Demange M [2001]. Ultrasonic extraction as a sample preparation

technique for elemental analysis by atomic spectrometry. J Anal At Spectrom 16:1147-1153.

[3] Foote P, Wickman DC, Perkins JB [2002]. Back-up data report for determination of hexavalent

chromium by HPLC with post-column derivatization, prepared under NIOSH Contract

2000-95-2955. Unpublished.
CHROMIUM, HEXAVALENT by Ion Chromatography: METHOD 7605, Issue 2, dated 10 March 2016 - Page 5 of 5 

[4] Eide ME [2000]. Hexavalent Chromium, Method No. W 4001. Salt Lake City, Utah: U.S. Department of

Labor (USDOL), Occupational Safety and Health Administration (OSHA), Salt Lake Technical Center.

[5] Ku JC, Eide M [1998]. Hexavalent chromium in the workplace atmosphere, OSHA ID -215 . Salt Lake

City, Utah: US Department of Labor (USDOL), Occupational Safety and Health Administration (OSHA),

Salt Lake Technical Center.

[6] ISO [2005; reapproved 2010]. Method No. 16740 Workplace air - Determination of hexavalent

chromium in airborne particulate matter - Method by ion chromatography and spectrophotmetric

measurement using diphenyl carbazide. Geneva, Switzerland: International Organization for

Standardization (ISO).

[7] Arar EJ, Pfaff JD, Martin TD [1994]. Method 218.6, Revision 3.3. Cincinnati, Ohio: Environmental

Monitoring Systems Laboratory, Office of Research and Development, U.S. Environmental Protection

Agency.

[8] NIOSH [1975]. Criteria for a recommended standard: occupational exposure to chromium (VI).

Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease Control,

National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 76-129.

[9] Wang J, Ashley K, Kennedy ER, Neumeister C [1997]. Determination of hexavalent chromium in

industrial hygiene samples using ultrasonic extraction and flow injection analysis. Analyst

122:1307-1312.

[10] Ndung’u K, Djane N-K, Malcus F, Mathiasson L [1999]. Ultrasonic extraction of hexavalent chromium

in solid samples followed by automated analysis using a combination of supported liquid membrane

extraction and uv detection in a flow system. Analyst 124:1367-1372.

[11] Wang J, Ashley K, Marlow D, England EC, Carlton G [1999]. Field method for the determination of

hexavalent chromium by ultrasonic and strong anion exchange solid phase extraction. Anal Chem

71:1027-1032.

[12] EC/IRMM [1997]. Certificate of analysis, CR M545: Cr(VI) and total leacheable Cr in welding dust

loaded on a filter. Brussels: Commission of the European Communities, Institute for Reference

Materials and Measurements.

[13] Burkart JA [1986]. General procedures for limit of detection calculations in industrial hygiene

chemistry laboratory. Appl Ind Hyg, 1(3):153 -155. 

METHOD REVISED BY:

Kevin Ashley, NIOSH 

Penny A. Foote, James B. Perkins, Don C. Wickman, DataChem Laboratories, Inc, Salt Lake City, Utah 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

LEAD BY PORTABLE

ULTRASONIC EXTRACTION/ASV 7701

Pb MW: 207.19 (Pb) CAS: 7439-92-1 (Pb) RTECS: OF7525000 (Pb)

223.19 (PbO) 1317-36-8 (PbO) OG1750000 (PbO)

METHOD: 7701, Issue 3 EVALUATION: FULL Issue 1: 15 January 1998

Issue 3: 17 February 2016

OSHA: 0.05 mg/m3

NIOSH: 0.05 mg/m3

PROPERTIES: soft gray metal; d 11.3 g/cm3 @ 20 °C; MP

327.5 °C, BP 1749 °C; valences 2+, 4+ in salts

SYNONYMS: elemental lead, lead compounds (except alkyl lead)

SAMPLING

SAMPLER: FILTER (37-mm, 0.8-μm pore, mixed 

cellulose ester membrane)

FLOW RATE: 1 to 4 L/min

VOL-MIN: 20 L @ 0.05 mg/m3

-MAX: 1500 L 

SHIPMENT: Routine

SAMPLE

STABILITY: Stable

BLANKS: Minimum of 2 field blanks per set

ACCURACY

RANGE 

STUDIED: 0.025 to 0.150 mg/m3 (as Pb) 

(based on Pb mass loadings) [4]

BIAS: None identified in laboratory studies [3-5] 

<10% in field study [6] 

OVERALL

PRECISION (SS�rrrr): 0.087 (screen-printed electrodes);

0.094 (Hg film on glassy carbon

electrodes) [4]

ACCURACY: ± 17.2% 

± 19.3%

MEASUREMENT

TECHNIQUE: PORTABLE ANODIC STRIPPING

VOLTAMMETRY

ANALYTE: Lead

EXTRACTION: 10% HNO3 10 mL; ultrasonication

ANALYSIS

ALIQUOT VOL: 0.1 to 5 mL

DEPOSITION

POTENTIAL: -0.8 V to -1.0 V vs. Ag/AgCl

ANODIC SWEEP: Deposition potential to 0.0 V vs. 

Ag/AgCl sweep rate variable [1,2]

REFERENCE

ELECTRODE: Ag/AgCl or calomel

WORKING

ELECTRODE: Mercury film on glassy carbon or screen

printed

SUPPORTING

ELECTROLYTE: Mixture of NaCl/NaOH or KCl/KOH, 

L-ascorbic acid

CALIBRATION: Pb2+ in 5% HNO3

RANGE: 0.31 to >1000 μg Pb per sample [3,4]

ESTIMATED LOD: 0.05 μg/sample [1]

PRECISION (SS̅

rr): 0.068 @ 60 μg (as Pb) per sample [3]

APPLICABILITY: The working range is (at least) 0.20 to 5.00 mg/m3 (as Pb) for a 120-L air sample. Lead determination by ultrasonic 

extraction/ASV method is applicable to the on-site, field-based determination of lead in air filter samples, and also may be used for 

laboratory-based air filter sample preparation and analysis.

INTERFERENCES: Thallium is a known interference, but its presence is unlikely in the vast majority of samples. Extremely high 

concentrations of copper may cause a positive bias. Surfactants can poison electrode surfaces, so if the presence of surfactants is 

suspected they must be eliminated during sample preparation [7,8].

OTHER METHODS: Laboratory-based methods include atomic spectrometric methods following concentrated acid hotplate 

digestion: NIOSH methods 7082 (flame AAS), 7105 (graphite furnace AAS), and 7300-series methods (ICP-AES) [9]. ASTM 

standards based on NIOSH methods for sample collection, preparation and analysis have been published [10]. 
LEAD BY PORTABLE ULTRASONIC EXTRACTION/ASV: METHOD 7701, Issue 3, dated 17 February 2016 - Page 2 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Nitric acid,* 10% (v/v) (Prepared from concentrated

nitric acid, reagent grade; spectroscopic grade if

trace analysis).

2. Distilled or deionized water (ASTM Type I or better

[11]).

3. Calibration stock solution, 1000 μg/mL Pb.

Commercial standard, or dissolve 1.00 g Pb metal in

minimum of 10 mL of 50% HCl and dilute to 1 L

with 1% (v/v) HCl. Store in a polyethylene bottle.

Stable for at least one year.

4. Supporting electrolyte: Aqueous inert salt mixture

such as 2.5 M NaCl and 0.25 M NaOH * (reagent

grade or equivalent) [3,4].

5. Dissolved oxygen scavenger such as 0.25 M Lascorbic acid (tissue culture grade or equivalent)

[3,4].

6. Mercuric nitrate (reagent grade), if required (for Hg

film electrodes).

7. Certified Reference Materials (CRMs) for lead.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: Mixed cellulose ester filter, 0.8 μm

pore size, 37-mm diameter, in cassette filter 

holder or cellulosic internal capsule.

2. Personal sampling pump, 1 to 4 L/min, with 

flexible connecting tubing.

3. Field-portable anodic stripping voltammeter.

4. Disposable or renewable voltammetry

electrodes.

5. Ultrasonic bath, 50 watts minimum power.

6. Power source for ultrasonic bath.

7. Plastic centrifuge tubes, 50-mL, with screw

caps.

8. Test tube rack (size to fit in ultrasonic bath).

9. Plastic sample cell container.

10. Mechanical pipets (class A equivalent), 0.1-mL

to 10-mL, as needed.

11. Pipet tips for mechanical pipets.

12. Forceps.

13. Bottles, polyethylene, 100- to 1000-mL.

14. Volumetric flasks, 100-mL (for preparatory lab

work).

15. Plastic rods.

16. Wipes (ASTM E1792 [12])

NOTE: Clean all glassware and reusable

plasticware with diluted nitric acid and 

rinse thoroughly with distilled or 

deionized water before use.

SPECIAL PRECAUTIONS: Nitric acid and sodium hydroxide are irritants and may burn skin. Perform 

extractions in a well-ventilated area. Wear gloves and eye protection.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in the line.

2. Sample at an accurately known flow rate between 1 and 4 L/min for up to 8 h for a total sample size of

20 to 1500 L for TWA measurements. Do not exceed a filter loading of ca. 5 mg total dust.

NOTE: Filter overloading can be assessed by periodic visual checks. See NMAM guidance chapters for

additional discussion on filter capacity. 

SAMPLE PREPARATION:

3. Open the cassette filter holders and, with forceps, transfer the samples and blanks to separate 50-mL

centrifuge tubes. Use internal cellulosic capsules or wipe the internal cassette surfaces with a 37 mm

MCE filter or cellulosic wipe wetted with deionized water and add to the centrifuge tube to transfer

non-filter aerosol deposits into the tube. See [13] for additional information. Push the filters to the

bottom of the tubes with plastic rods.

NOTE: An alternative means to include non-filter aerosol deposits is to carry out extraction directly

within the cassette.

4. Add 10 mL of 10% HNO3 and cap the centrifuge tubes.
LEAD BY PORTABLE ULTRASONIC EXTRACTION/ASV: METHOD 7701, Issue 3, dated 17 February 2016 - Page 3 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

5. Place centrifuge tubes in ultrasonic bath, and agitate at room temperature for at least 30 min.

NOTE 1: The water level in the bath should be above the level of liquid within the centrifuge tubes. 

NOTE 2: Proper performance of the ultrasonic bath should be demonstrated before use. This can be

accomplished by checking recoveries of lead from performance evaluation materials.

6. Shake tubes for 5 to 10 sec, and allow to settle.

CALIBRATION AND QUALITY CONTROL:

7. Prior to field work, prepare a series of working standards covering the range 0.25 to 20 μg/mL Pb.

a. Add aliquots of calibration stock solution to 100-mL volumetric flasks, and dilute to volume

with 10% HNO3. Store and transport the working standards in polyethylene bottles, and

prepare fresh weekly.

b. Analyze the working standards together with the blanks and samples (steps 11 through 14).

c. Prepare a calibration graph of instrumental response vs. lead concentration (μg/mL Pb).

NOTE: Some portable instruments read concentration directly. Calibrate according to

manufacturer’s directions.

8. Analyze at least one standard for every 20 samples to check for instrument drift (steps 11 through 14).

9. Check recoveries with at least one spiked media blank per 20 samples (one per batch minimum). Use

certified reference materials to substantiate recoveries. Use method of standard additions to check for

matrix effects or interferences.

10. Check for lead contamination by analyzing at least one reagent and one media blank per 20 samples

(minimum of one per batch) (steps 11 through 14).

MEASUREMENT:

11. Set instrument parameters as specified by the manufacturer, or use conditions specified on page

7701-1.

NOTE: If renewable electrodes are used, clean the glassy carbon electrode and deposit a fresh mercury

film prior to conducting analyses.

12. Transfer sample aliquot (1 to 5 mL) to analytical cell, and dilute with 10% HNO3, if necessary.

NOTE: High concentrations of lead may require analysis of diluted analyte solutions.

13. Add supporting electrolyte and oxygen scavenger to sample cell. Ensure final volume of the cell is 5 mL

(disposable electrodes) or 10 mL (renewable electrodes) by diluting with distilled or deionized water.

14. Measure lead content of the sample aliquot (μg/sample or μg/mL), and record the result.

NOTE 1: For renewable electrodes, rinse and drain electrochemical sample cell at least three times with 

distilled or deionized water between sample runs. For disposable electrodes, use a fresh 

plastic sample cell container for each sample analyzed.

NOTE 2: If the measured value is above the linear range of the standards, dilute with 10% HNO3, 

reanalyze, and apply the appropriate dilution factor in the calculations.

CALCULATIONS:

15. Using measured lead contents, calculate the corresponding concentrations (μg/mL) of lead in the

original extracted sample, Cs, and average media blank, Cb, from the calibration graph.

NOTE: Be sure to account for dilution factors.

16. Using solution volumes (mL) of the samples, Vs, and media blanks, Vb, calculate the concentration, C

(mg/m3

), of lead in the air volume sampled, V (L):

CC = CCssVVss − CCbbVVbb

VV , mmmm mm3 ⁄

NOTE: μg/L ≅ mg/m3
LEAD BY PORTABLE ULTRASONIC EXTRACTION/ASV: METHOD 7701, Issue 3, dated 17 February 2016 - Page 4 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

EVALUATION OF METHOD:

This method was evaluated with lead aerosol samples generated in the laboratory (40 to 80 μg Pb per filter) 

[3], and with air particulate samples collected from workplaces where abrasive blasting of leaded paint on 

highway bridges was being conducted [4]. For the latter, lead loadings cover the range from below the 

detection limit of 0.09 μg Pb per filter to loadings in excess of 1500 μg Pb per filter [4]. The method also has 

been evaluated with performance evaluation materials and by interlaboratory testing [3-5]. Lead recoveries 

from Certified Reference Materials (CRMs) were found to be quantitative (≥90%) and equivalent to recoveries 

obtained using confirmatory analytical methods (NIOSH 7082, 7105, and 7300 [9]).

REFERENCES:

[1] Wang J [1985]. Stripping analysis. New York: VCH Publishers.

[2] Wang J [1996 ]. Electrochemical preconcentration. In: Kissinger PT, Heineman WR, eds. Laboratory 

techniques in electroanalytical chemistry. 2nd ed. New York: Marcel Dekker, pp. 719-737.

[3] Ashley K [1995]. Ultrasonic extraction and field-portable anodic stripping voltammetry of lead from 

environmental samples. Electroanalysis 7:1189-1192.

[4] Ashley K, Mapp KJ, Millson M [1998]. Ultrasonic extraction and field-portable anodic stripping 

voltammetry for the determination of lead in workplace air samples. Am Ind Hyg Assoc J 59:671-679.

[5] Ashley K, Song R, Esche CA, Schlecht PC, Baron PC, Wise TJ [1999]. Ultrasonic extraction and portable 

anodic stripping voltammetric measurement of lead in paint, dust wipes, soil, and air: An 

interlaboratory evaluation. J Environ Monit 1:459-464.

[6] Sussell A, Ashley K [2002]. Field measurement of lead in workplace air and paint chip samples by 

ultrasonic extraction and portable anodic stripping. J Environ Monit 4:156-161.

[7] Ashley K [1994 ]. Electroanalytical applications in occupational and environmental health. 

Electroanalysis 6:805-820.

[8] Ashley K, Wise TJ, Mercado W , Parry DB [2001]. Ultrasonic extraction and portable anodic stripping 

voltammetric measurement of lead in dust wipe samples. J Hazard Mater 83:41-50.

[9] NIOSH [1994]. Lead: Method 7082, Lead by GFAAS: Method 7105, Elements: Method 7300. In: Eller PM, 

Cassinelli ME, eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/].

[10] ASTM [2003]. Standard Nos. E1613, E1741, E1775, E1979, E2051. In: Annual Book of ASTM Standards, 

Vol. 04.11. W. Conshohocken, PA: ASTM International. 

[11] ASTM [2003]. Standard No. D1193: Specification for reagent water. In: Annual Book of ASTM Standards, 

Vol. 11.01. W. Conshohocken, PA: ASTM International. 

[12] ASTM [2011]. Standard No. E1792: Standard specification for wipe sampling materials for lead in 

surface dust. In: Annual Book of ASTM Standards, Vol. 04.11. W. Conshohocken, PA: ASTM 

International. 

[13] Ashley K, Harper M [2013]. Closed-face filter cassette (CFC) sampling-guidance on procedures for 

inclusion of material adhering to internal sampler surfaces. J Occup Environ Hyg 10:D29-D33. 
LEAD BY PORTABLE ULTRASONIC EXTRACTION/ASV: METHOD 7701, Issue 3, dated 17 February 2016 - Page 5 of 5 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD REVISED BY:

Kevin Ashley, Ph.D., NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of these 

websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT,

by Field-Portable Spectrophotometry 

7703

AW: 52.00 CAS: 18540-29-9 RTECS: GB6262000 

MW: 99.99 CAS: 1333-82-0 RTECS: GB6650000

METHOD: 7703, Issue 2 EVALUATION: FULL Issue 1: 15 March 2003

Issue 2: 20 October 2015

OSHA: 0.005 mg/m3 (1910.1026); C 0.1 mg/m3 as 

(exceptions to 1910.1026)

NIOSH: 0.0002 mg/m3 (8 h); carcinogen

PROPERTIES: Oxidizing agent

SYNONYMS: Vary depending on the compound; chromate commonly used

SAMPLING

SAMPLER: FILTER

(5.0 μm PVC membrane [1,2]; 0.8 μm MCE or 

1.0 μm PTFE acceptable for field analysis [3])

FLOW RATE: 1 L/min to 4 L/min

VOL-MIN: 54 L (2 L/min for 27 min)

-MAX: 1200 L (3 L/min for 400 min)

SHIPMENT: Refrigerant pack at 4 °C ± 2 °C (optional)

SAMPLE

STABILITY: Analyze within 24 h; if applicable, keep 

samples at 4 °C ± 2 °C

BLANKS: One per twenty field samples, minimum of 2 

per set

ACCURACY

RANGE STUDIED: 0.045 μg/m3 to 1146 μg/m3 (about 20 L 

to 200 L samples) [3,4]

BIAS: −1.00% [3]

OVERALL

PRECISION ( ): 0.080

ACCURACY: ±15.7%

MEASUREMENT

TECHNIQUE: FIELD-PORTABLE VISIBLE 

SPECTROPHOTOMETRY

ANALYTE: Chromium-diphenylcarbazone complex

EXTRACTION: 10 mL 0.05 mol/L ammonium sulfate / 

0.05 mol/L ammonium hydroxide (pH = 8 

± 0.5), ultrasonic extraction 30 min

ISOLATION: Strong anion exchange solid phase 

extraction

ELUTION

SOLUTION: 0.5 mol/L ammonium sulfate / 0.1 mol/L 

ammonium hydroxide

WAVELENGTH: 540 nm, 1 cm path length

CALIBRATION: Standard solutions of potassium 

chromate in elution solution

RANGE: 1 μg to 400 μg per sample

ESTIMATED LOD: 0.08 μg hexavalent chromium per 

sample [3]

PRECISION ( ): 0.035 @ 3 μg to 400 μg per sample [3]

APPLICABILITY: The working range is 0.22 μg/m3 to 333 μg/m3 for a 1200 L air sample. This method may be used for the 

determination of soluble forms of hexavalent chromium. Insoluble hexavalent chromium requires modification of the 

method using ultrasonic extraction with carbonate buffer.

INTERFERENCES: Interferences from reducing agents such as divalent iron are minimized to the extent possible by the 

alkaline ultrasonic and solid phase extraction procedures. Interferences from other metal cations are eliminated by solid 

phase extraction [5]. Some reduction can occur on the filter during sampling, and is usually due to the presence of divalent 

iron, organic material, and/or acidic conditions [6]. Reduction of hexavalent chromium can occur over time on any filter 

type, and is especially problematic on MCE filters [7]. However, the use of MCE and PTFE filters has been found to be acceptable for field use, where performance has been found to be equivalent to that of PVC filters [3]. During ultrasonic extraction, 

oxidation of trivalent chromium in solution to hexavalent chromium is prevented by the use of an ammonium buffer [8].

OTHER METHODS: This method is designed to be used in the field, but can also be utilized in the fixed-site laboratory. It 

is an alternative to laboratory methods such as NIOSH method 7605 or OSHA method ID-215 (hot plate digestion and ion 

chromatography). NIOSH method 7600 is a similar procedure, but no separation step is used. A field method not involving 

hexavalent chromium isolation, MDHS method 61, has been promulgated by the British Health and Safety Executive [9].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT (Field-Portable Spectrophotometry): METHOD 7703, Issue 2, dated 20 October 2015 - Page 2 of 6

REAGENTS:

1. Ammonium sulfate, reagent grade.

2. Ammonium hydroxide,* reagent grade.

3. Water, distilled or deionized.

4. Hydrochloric acid (37% mass fraction),*

reagent grade.

5. Acetonitrile,* reagent grade.

6. 1,5-Diphenylcarbazide (DPC), reagent grade.

7. Methanol,* reagent grade.

8. Extraction solution (extraction buffer): 

0.05 mol/L ammonium sulfate / 0.05 mol/L 

ammonium hydroxide, 1 L, aqueous in distilled 

or deionized water.

NOTE: Modification of method by using 

carbonate buffer (e.g., sodium 

carbonate) is required for extraction of 

insoluble hexavalent chromium.

9. Elution solution (elution buffer): 0.5 mol/L 

ammonium sulfate / 0.1 mol/L ammonium 

hydroxide, 250 mL, in distilled or deionized 

water.

10. Potassium chromate,* reagent grade.

11. Hexavalent chromium standard,* 1000 

μg/mL. Dissolve 3.735 g potassium chromate 

in deionized water to make 1 L, or use 

commercially available solution.

12. Calibration stock solution,* 100 μg/mL: dilute 

1000 μg/mL hexavalent chromium standard 

1:10 with extraction buffer. (Solution is stable 

for a month.)

13. Diphenylcarbazide complexation solution 

(20 mmol/L): Measure 0.48 g DPC powder 

and place in a 100 mL volumetric flask. Add 

about 80 mL of acetonitrile and dissolve the 

DPC. Bring up to the mark with additional 

acetonitrile and mix thoroughly.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Samplers: 5 μm pore size polyvinylchloride 

(PVC), 0.8 μm pore size mixed cellulose ester 

(MCE), or 1.0 μm polytetrafluoroethylene 

(PTFE) filters, 37 mm diameter, with backup 

pads, in polystyrene cassette filter holder, 2- or 

3-piece.

NOTE: MCE filters, and some PVC filters, 

promote reduction of hexavalent 

chromium on a timescale of a few days. 

However, either filter type is acceptable 

for field use if the samples are to be 

analyzed within 24 h of collection.

2. Personal sampling pump, 1 L/min to 4 L/min, 

with flexible connecting tubing.

3. Ultrasonic bath (sonicator), 100 W minimum 

power.

4. Solid phase extraction manifold, multi-port.

5. Portable vacuum pump with pressure 

metering valve.

6. Portable visible spectrophotometer, sample 

path length 1 cm with quartz cuvette(s).

7. Strong anion exchange solid phase extraction 

(SPE) cartridges, 10 mL, disposable; loaded 

with 500 mg or 1000 mg quaternary amine 

bonded silica, capacity about 1 mmol/g of 

singly charged anion.

8. Pipettors, mechanical, assorted volumes (e.g., 

1 mL to 10 mL) with disposable tips.

9. Micropipettors, mechanical, assorted volumes 

(e.g., 10 μL to 100 μL) with disposable tips.

10. Centrifuge tubes, plastic, 15 mL, with screw 

caps.

11. Scintillation vials, 20 mL, glass, with PTFE-lined 

screw caps.

12. Assorted beakers (and possibly Erlenmeyer 

flasks), various volumes.

13. Volumetric flasks, 25 mL, 100 mL, 250 mL, and 

1000 mL.

14. Forceps, PTFE-coated.

15. Glass or plastic rods.

16. Disposable gloves, plastic or latex.

17. Laboratory wipes.

18. Portable power generator (if necessary).

NOTE: If no power supply is available at 

the field site, electric power can be 

provided by means of a portable, 

gasoline (or other) generator.

SPECIAL PRECAUTIONS: NIOSH considers all hexavalent chromium compounds to be suspect 

occupational carcinogens [10]. Efforts must be made to prevent aerosolizing chromate-containing 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT (Field-Portable Spectrophotometry): METHOD 7703, Issue 2, dated 20 October 2015 - Page 3 of 6

compounds and solutions. All sample preparation should be carried out in a well-ventilated area 

(vacuum hood preferable); forced ventilation should be used if no hood is available. Methanol and 

acetonitrile solutions are flammable and must be handled carefully, i.e., wearing of impermeable 

gloves and avoidance of vapors. To the extent possible, solutions should be prepared in the 

laboratory before taking them to the field. Concentrated hydrochloric acid is highly corrosive 

and ammonium hydroxide is a respiratory irritant. All work with these compounds should be 

performed in a hood. Use proper protective clothing including gloves, safety glasses, and laboratory 

coat. Potassium chromate is a strong oxidizer with risk of fire and explosion upon contact with 

combustible substances and reducing agents.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line.

2. Sample at an accurately known flow rate in the range of 1 L/min to 4 L/min for a sample size of 54 L 

to 1200 L. Do not exceed 2 mg of particulate loading on the filter. Label the filter cassette.

3. Don a fresh pair of disposable plastic or latex gloves (to prevent sample contamination).

4. With PTFE-coated forceps, remove filters from cassettes after completion of sampling, and place 

in separate plastic 15 mL centrifuge tubes for subsequent sample preparation. Discard cellulose 

backup pads and gloves.

SAMPLE PREPARATION:

5. Add 10 mL of extraction solution (weak buffer) to each 15 mL centrifuge tube containing the filter 

sample. Ensure that the filter is covered by the extraction solution. If necessary, push the filter down 

with a clean glass or plastic rod to immerse the entire filter. Cap and label the tubes.

6. Place sample tubes in the ultrasonic bath (sonicator). The water level in the bath should be higher 

than the liquid level in the centrifuge tube. Sonicate for 30 min.

NOTE: Numerous centrifuge tubes containing sample filters can be subjected to sonication at one 

time, depending upon the size of the ultrasonic bath. Ensure that the bath is warm (but 

< 40 °C).

7. Set up the solid phase extraction manifold.

a. Place disposable solid phase extraction (SPE) cartridges in each port, and place scintillation vials 

beneath the cartridges. Label the cartridges.

b. Attach the vacuum pump to the SPE manifold.

c. To condition SPE cartridges, pipet 3 mL of methanol into each cartridge, and evacuate. Then 

pipet 3 mL of extraction solution into each cartridge, and evacuate. Repeat.

8. Extract hexavalent chromium from sample solution.

a. Pipet 3 mL to 5 mL of each ultrasonicated sample solution from the centrifuge tubes into the 

disposable SPE cartridge. Dispose of the pipet tip.

b. Adjust the vacuum to obtain an extraction rate of about one drop per second (approximately 

27 kPa or 203 mmHg; no more than 34 kPa or 254 mmHg). Manually tighten cartridges by 

twisting, if necessary, to slow down the rate of liquid dripping.

NOTE 1: For samples in which the expected hexavalent chromium concentration is high, 

smaller aliquots (1 mL to 2 mL) should be dispensed into the SPE cartridges to prevent 

breakthrough. High concentration of hexavalent chromium can be assessed visually by 

its orange color.

NOTE 2: For samples having low hexavalent chromium concentration, additional 3 mL to 5 mL 

aliquots of ultrasonicated sample solution can be loaded onto SPE cartridges (step 8.a). 

In this manner, the cartridge can be used to preconcentrate hexavalent chromium.

c. When it appears the solution has passed through all the cartridges, increase the vacuum to 

ensure that all solution passes through the cartridges. This step selectively binds hexavalent 

chromium to the stationary phase of each cartridge.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT (Field-Portable Spectrophotometry): METHOD 7703, Issue 2, dated 20 October 2015 - Page 4 of 6

d. To remove residue of trivalent chromium and other potential interferences, reduce the vacuum 

to 0. Add 1 mL distilled or deionized water to each cartridge, adjust vacuum to obtain 1 drop per 

second (about 27 kPa or 203 mmHg), then reduce vacuum to 0 when completed.

e. Remove the scintillation vials beneath the cartridges and discard.

NOTE: This solution contains unwanted fractions that should contain no hexavalent chromium.

9. Place clean, labeled scintillation vials beneath correct cartridges in the SPE manifold.

a. Add 9 mL of the elution solution (elution buffer) to each cartridge to elute hexavalent chromium, 

and repeat steps 8.b, 8.c, and 8.d.

b. Remove the scintillation vials, and cap them. Dispose of the used SPE cartridges.

NOTE: The scintillation vials now contain extracted and isolated hexavalent chromium, which is 

ready for subsequent analysis.

10. Uncap each scintillation vial containing extracted and isolated hexavalent chromium, and add 

100 μL hydrochloric acid.

11. Add 2 mL DPC complexation solution, recap vials, and mix thoroughly. Allow to stand for at least 5 

min for complete color development.

CALIBRATION AND QUALITY CONTROL:

12. Calibrate daily with at least 6 working standards over the range of 0 μg/mL to 2 μg/mL of hexavalent 

chromium per standard.

a. To 10 mL volumetric flasks containing about 5 mL of elution solution (strong buffer), pipet known 

volumes (10 μL to 200 μL) of hexavalent chromium calibration stock solution (100 μg/mL) to 

produce concentrations of 0.1 μg/mL, 0.2 μg/mL, 0.5 μg/mL, 1.0 μg/mL, and 2.0 μg/mL. Add 

100 μL of hydrochloric acid and 2 mL of DPC complexation solution to each. Dilute to the mark 

with elution solution and mix thoroughly.

NOTE: A minimum of two of the concentration levels (e.g., 0.1 μg/mL and 1.0 μg/mL) should be 

run at least in triplicate.

b. Prepare a blank by pipetting 100 μL of hydrochloric acid and 2 mL of DPC complexation solution 

into 10 mL volumetric flask containing about 5 mL of the elution solution (elution buffer); dilute 

to the mark with elution solution and mix thoroughly.

c. Analyze the calibration solutions and the blank (steps 15, 16, 17, 18, 19, and 20).

13. Analyze at least two field blanks, one field blank per twenty samples (steps 10, 11, 15, 16, 17, 18, 19, 

and 20). Also analyze at least three of the calibration solutions in triplicate.

14. Prepare a calibration graph of absorbance vs, hexavalent chromium concentration.

NOTE: As an alternative to steps 12, 13, and 14, the standard addition approach can be used [11].

MEASUREMENT:

15. Turn on the spectrophotometer, and allow for an appropriate warm-up period.

16. Set the spectrophotometer to 540 nm. Set portable spectrophotometer parameters according to the 

manufacturer’s instructions and the conditions on page 7703-1.

17. Rinse the quartz cuvette three times with distilled or deionized water, then rinse with blank solution.

18. Measure the blank. Adjust the spectrometer to zero absorbance.

19. Uncap the scintillation vial containing the sample solution to be analyzed.

a. Condition the cuvette by filling with the solution to be analyzed, and discard the solution.

b. Refill the cuvette with the sample solution to be analyzed.

c. Place the cuvette in the spectrophotometer.

NOTE: Wipe any extra moisture or liquid off the sides of the cuvette with a dry laboratory wipe, 

and take care to handle the cuvette only by the frosted sides.

20. Analyze samples, standards, and blanks. Record the absorbance.

NOTE: If the absorbance value is greater than 2 absorbance units, dilute the solution to be analyzed 

with elution solution (strong buffer) and reanalyze.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT (Field-Portable Spectrophotometry): METHOD 7703, Issue 2, dated 20 October 2015 - Page 5 of 6

CALCULATIONS:

21. From the calibration graph, determine the mass of hexavalent chromium in each sample, (μg), 

and in the average field blank, (μg).

NOTE: If standard addition method was used, make appropriate adjustments from the calibration 

graph obtained [11].

22. Calculate the concentration, , of hexavalent chromium in the air volume sampled, (L):

, μg/L or mg/m3.

NOTE 1: If samples were diluted during sample preparation, be sure to account for the dilution 

factor in the calculation.

NOTE 2: If the hexavalent chromium concentration is to be reported as chromic acid ( ), multiply 

 by 1.92 (MW of chromic acid divided by AW of chromium).

EVALUATION OF METHOD:

This method was evaluated in the laboratory with spiked filters [3,4,5] and a certified reference 

material containing a known loading of hexavalent chromium [4]. This certified reference material 

(CRM) is European Commission, Institute for Reference Materials and Measurements (EC/IRMM) CRM 

545, hexavalent chromium and total chromium in welding dust loaded on a glass fiber filter [12]. The 

method has also been evaluated in the field, where samples collected during aircraft maintenance 

operations were analyzed on-site [3,4]. The accuracy was estimated using the protocol summarized in a 

NIOSH technical report [13].

Alternative filter types can also be used, e.g., PTFE, binder-free glass fiber filters, or quartz fiber 

filters. Filter materials should be tested before use to ensure hexavalent chromium stability. Filters can 

be pretreated with base to minimize hexavalent chromium reduction during sampling in high-iron or 

acidic environments [6].

REFERENCES:

[1] NIOSH [1994]. Chromium, hexavalent: Method 7600. In: Eller PM, Cassinelli ME, eds. NIOSH manual 

of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, 

Centers for Disease Control and Prevention, National Institute for Occupational Safety and Health, 

DHHS (NIOSH) Publication No. 94-113 [www.cdc.gov/niosh/nmam/].

[2] ASTM [2013]. Standard test method for the determination of hexavalent chromium in workplace 

air by ion chromatography and spectrophotometric measurement using 1,5-diphenylcarbazide: 

method D6832-13. West Conshohocken, PA: ASTM International. DOI: 10.1520/D6832-13.

[3] Marlow D, Wang J, Wise TJ, Ashley K [2000]. Field test of a portable method for the determination of 

hexavalent chromium in workplace air. Am Lab 32(15):26–28.

[4] Wang J, Ashley K, Marlow D, England EC, Carlton G [1999]. Field method for the determination of 

hexavalent chromium by ultrasonication and strong anion-exchange solid-phase extraction. Anal 

Chem 71(5):1027–1032.

[5] Wang J, Ashley K, Kennedy ER, Neumeister C [1997]. Determination of hexavalent chromium 

in industrial hygiene samples using ultrasonic extraction and flow injection analysis. Analyst 

122(11):1307–1312.

[6] Foster RD, Howe AM, Cottrell SJ, Northage C [1996]. An investigation of the exposure of workers to 

hexavalent chromium in the air of plating works. Sheffield, UK: Health and Safety Executive, Health 

and Safety Laboratory, HSL Report IR/L/95/15.

[7] Molina D, Abell MT [1987]. An ion chromatographic method for insoluble chromates in paint 

aerosol. Am Ind Hyg Assoc J 48(10):830–835.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CHROMIUM, HEXAVALENT (Field-Portable Spectrophotometry): METHOD 7703, Issue 2, dated 20 October 2015 - Page 6 of 6

[8] Ndung’u K, Djane N-K, Malcus F, Mathiasson L [1999]. Ultrasonic extraction of hexavalent chromium 

in solid samples followed by automated analysis using a combination of supported liquid 

membrane extraction and UV detection in a flow system. Analyst 124(9):1367–1372.

[9] Health and Safety Executive [1988]. Total hexavalent chromium compounds in air—colorimetric 

laboratory method using 1,5-diphenylcarbazide: MDHS 61. In: Methods for the determination of 

hazardous substances. London: Health and Safety Executive.

[10] NIOSH [2013]. Criteria for a recommended standard: occupational exposure to hexavalent 

chromium. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) 

Publication No. 2013-128 [www.cdc.gov/niosh/docs/2013-128/].

[11] Skoog DA, West DM, Holler FJ [1988]. Fundamentals of analytical chemistry. New York: W.B. 

Saunders.

[12] EC/IRMM [1997]. Certificate of analysis, CRM 545: and total leachable in welding dust 

loaded on a filter. Brussels: Commission of the European Communities, Institute for Reference 

Materials and Measurements.

[13] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. 

By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 95-117 [www.cdc.gov/niosh/

docs/95-117/pdfs/95-117.pdf].

METHOD WRITTEN BY:

Kevin Ashley, Ph.D., NIOSH/DART; Jin Wang, Ph.D., NIOSH/HELD; David Marlow, NIOSH/DART; James 

Boiano, CIH, NIOSH/DSHEFS.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

BERYLLIUM in Air 

by Fluorometry 7704

Be MW: 9.0121 CAS: 7440-41-7 RTECS: DS1750000

METHOD: 7704, Issue2 EVALUATION: FULL Issue 1: 6 April 2007

Issue 2: 12 December 2015

OSHA: 2 μg/m3

, ceiling 5 μg/m3, peak 25 μg/m3

MSHA: 2 μg/m3 

DOE: 2 μg/m3 (action level 0.2 μg/m3) 

NIOSH REL: 0.5 μg/m3

OTHER OELs: see Table 5

PROPERTIES: solid, d 1.85 g/mL, MP 1,278 °C, VP 0 kPa

(0 mm Hg) @ 25 °C

SYNONYMS: beryllium metal, beryllia (BeO)

SAMPLING

SAMPLER: FILTER (mixed cellulose ester or nylon 

membrane, 0.8 μm pore size, 25- or 37-

mm diameter)

FLOW RATE: (1 to 4) L/min

VOL-MIN: 240 L

 -MAX: 2,000 L

SHIPMENT: routine

SAMPLE

STABILITY: stable

BLANKS: 3 field blanks min.

ACCURACY

RANGE 

STUDIED: 0.0001 to 6 μg per filter [1,2]

BIAS: negligible [1,2]

OVERALL

PRECISION (SS�rrrr): 0.094

ACCURACY: 18.9%

MEASUREMENT

TECHNIQUE: UV/VIS FLUOROMETRY

ANALYTE: complex of hydroxybenzoquinoline

sulfonate (HBQS) with beryllium

DISSOLUTION: ammonium bifluoride (aqueous), 10 g/L

DETECTION 

SOLUTION: contains 63.4 μmol/L HBQS, 2.5 mmol/L 

EDTA, and 50.8 mmol/L lysine 

monohydrochloride (optional); pH 

adjusted to 12.85 with 10 mol/L NaOH,

as necessary

DETECTOR: excitation, 360 nm to 390 nm; emission, 

integrated between 470 and 480 nm 

(λmax ≈ 475 nm)

CALIBRATION: beryllium standard solutions

RANGE: (0.005 to 6) μg per filter [1]

ESTIMATED LOD: 0.0001 μg per filter [Table 4]

PRECISION (SS�rr): 0.021 at ≈0.2 μg per filter, 0.076 at ≈1.5 

μg per filter, 0.052 at ≈3 μg per filter

APPLICABILITY: The working range of the method is 0.0005 μg/m3 to 0.6 μg/m3 for an air sample of 1,000 L. The analysis is for 

total beryllium and is not compound specific.

INTERFERENCES: Minor interference from iron can result if iron concentrations are high. Samples high in iron demonstrate a 

yellow or gold coloration. This interference can be minimized by allowing the solution to sit for at least two hours, during which 

time the solution clears, and then filtering the sample extract before use. An alternative method is to filter the solution after 30 

minutes of standing through a hydrophilic filter of pore size of 0.2 μm or smaller.

OTHER METHODS: Method 7300 (hot plate digestion and inductively coupled plasma atomic emission spectrometry) is an 

alternative procedure for the determination of elemental beryllium [3], but with higher detection limits. ASTM method D7202 is a 

similar procedure to detect elemental beryllium by fluorescence [4].
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 2 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Ammonium bifluoride.* 

2. Ethylenediaminetetraacetic acid (EDTA), 

disodium salt, dihydrate. 

3. 10-Hydroxybenzo[h]quinoline-7-sulfonate 

(HBQS) [5]. 

4. L-Lysine monohydrochloride. 

5. Sodium hydroxide.* 

6. Water, deionized. 

7. Dissolution solution:* aqueous ammonium 

bifluoride, 10 g/L (prepared by dissolving 

ammonium bifluoride in deionized water). 

8. Detection solution:* 63.4 μmol/L HBQS, 2.5 

mmol/L EDTA, and 50.8 mmol/L lysine 

monohydrochloride; pH adjusted to 12.85 

with 10 mol/L NaOH). An alternative 

preparation of dye solution without lysine 

(lysine-free) may be made by adding 1.104 g 

of EDTA and 64 μmoles of the 10-HBQS dye 

in 900 ml of water. After a clear solution is 

obtained, 114.5 ml of 2.5 N NaOH is added 

and mixed to obtain the final dye solution. 

The pH of the dye solution is 13.2. The lysinefree dye solution (commercially available) 

may be used for all analytical purposes and 

also provides superior detection limits.

9. Beryllium standard solution,* 1,000 mg/L 

(commercially available). 

10. Beryllium-spiked media* (commercially 

available).

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: membrane filter, mixed cellulose 

ester (MCE) or nylon, 0.8 μm pore size, 25- or 

37-mm diameter. 

2. Personal sampling pump, 1 L/min to 4 L/min, 

with clamps and flexible connecting tubing. 

3. Ultraviolet/visible (UV/Vis) fluorometer, with 

excitation lamp (λ = 380 nm) and timeintegrating visible detector (400 nm to 700 

nm, λmax ≈ 475 nm) or optical filters for 

appropriate wavelengths (excitation of 360 

nm to 390 nm; emission of ≈475 nm, with full 

width at half maximum of ± 5 nm). 

4. Mechanical agitator, shaker, or rotator. 

5. Hot block (for beryllium oxide extraction). 

6. Fluorescence cuvettes, disposable, 10 mm 

diameter, transparent to UV/Vis radiation. 

7. Centrifuge tubes, plastic, 15 mL. 

8. Syringe filters, hydrophilic polypropylene, 0.2 

μm pore size, 25-mm diameter, in plastic 

housings.

NOTE: Polytetrafluoroethylene (PTFE) filters 

are unsuitable for this method. 

9. Pipettors, mechanical, of assorted sizes. 

10. Pipet tips, plastic, disposable, of assorted 

sizes. 

11. Labware, plastic (e.g., beakers, flasks, 

graduated cylinders), of assorted sizes. 

12. Tweezers, plastic or plastic-coated. 

13. Laboratory wipes. 

14. Personal protective wear (e.g., respirators, 

gloves, lab coats, safety eyewear), as needed. 

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities and analysis. It 

is essential that suitable gloves, eye protection, laboratory coat, etc., be used when working with the 

chemicals. Perform sample preparation and analysis in a clean, well-ventilated area that is well removed 

from any possible beryllium contamination. Any area of skin affected by the dissolution or detection 

solutions must be immediately washed with plenty of water. Ammonium bifluoride will etch glass, so it is 

essential that all ammonium bifluoride solutions be contained in plastic labware. Avoid exposure by 

contact with skin or eyes, or by inhalation of vapor.

SAMPLING:

1. Calibrate each personal sampling pump with a representative sampler in line. 

2. Sample at an accurately known flow rate between 1 L/min and 4 L/min for a total sample size of 240 L 

to 2,000 L for TWA measurements. Do not exceed a filter loading of ≈2 mg total dust. 

3. After sampling, remove the filters from the cassettes using clean tweezers, and place into labeled

15 mL plastic centrifuge tubes.
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 3 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION: 

4. Add 5 mL of the dissolution solution (ammonium bifluoride, 10 g/L) to each 15 mL centrifuge tube 

containing an air filter sample, and cap each tube.

5. Place each tube into a mechanical rotator, and rotate for at least 30 min.

NOTE: Rotator may also be substituted by a shaker or an agitator as long as the dissolution solution 

wets the filter well. Sonication has also been shown to be effective. For dissolution of refractory 

materials such as high-fired beryllium oxide, agitation of the dissolution solution with the media 

must be replaced by heating to 85 °C for 60 minutes or more. Any standard dissolution process 

is particle-size dependent [6]. The two sources of BeO used to validate the method are 

described in the backup data report [7]. 

6. Filter each solution with a hydrophilic polypropylene syringe filter into a clean tube.

NOTE: This tube should be able to accept a cap so that the solution may be saved and used later for 

reanalysis if required. 

7. Pipet 0.1 mL of each sample filtrate into cuvettes containing 1.9 mL of the detection solution. Cap and 

mix briefly.

NOTE: The above procedure is typically used to analyze a range of 0.05 μg to 6 μg of beryllium on the 

sampling media. Alternative ratios of dissolution solution and detection solution may be used 

for analyzing alternative ranges of beryllium concentration. To test a range of 0.005 μg to 0.4 μg 

of beryllium on the sampling media, 0.4 mL of the sample filtrate is added to 1.6 mL of the 

detection solution in the cuvettes. The lysine-free dye solution may also be used for obtaining 

even lower detection limits at a dilution ratio of 3×, where 1.33 ml of the dye solution is mixed 

with 0.67 ml of the filtered solution extracts (Table 3) and beryllium in the range of 0.0005 μg to 

0.4 μg may be determined.

NOTE: If high iron or titanium concentration is suspected or is evident (owing to the appearance of 

suspended precipitate), allow the solution to settle and filter the solution using a hygroscopic 

syringe filter (e.g., polyethersulfone, or hydrophilic polypropylene).

NOTE: The stability of the detection and the dissolution solution is more than six months and of the 

mixed measurement solution comprising both is greater than 30 days. The solutions must be 

kept in sealed containers, and the detection and mixed solutions must be stored away from 

light.

NOTE: If the samples are suspected of having a contaminant that fluoresces and has excitation and 

emission spectra that overlap with that of the signal produced by the fluorescent dye bound to 

beryllium, then this contaminant needs to be removed. The presence of such a contaminant can 

be verified by subjecting the filtered sample to fluorescence excitation after the extraction step 

(without adding the fluorescent dye). If a fluorescence signal is detected, then that signal is 

ascribed to the presence of a fluorescent contaminant. To remove the contaminant, high-purity 

activated charcoal is added to the beryllium extraction solution (~10 mg/ml) and the extraction 

procedure is carried out at elevated temperature (80 to 90 °C for at least 45 minutes). If the 

beryllium extraction procedure has already been performed, then after the addition of activated 

charcoal, the extraction process is repeated at the elevated temperature. The solution is filtered 

to remove the activated charcoal before adding this to the detection solution to make the 

measurement solution. Details of this process have been published [8].
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 4 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALIBRATION AND QUALITY CONTROL:

8. Calibrate the fluorometer with beryllium stock standard solutions. Prepare a calibration graph of 

fluorescence intensity vs. beryllium concentration (ng/mL) in the stock standard.

NOTE: To test a range of 0.05 μg to 6 μg of beryllium on the sampling media, beryllium stock standard 

solutions are made up using beryllium spectrometric standards diluted with the ammonium 

bifluoride dissolution solution. A recommended series of stock standard solutions is (800, 200, 

40, 10, and 0) ng/mL. As with the samples, the stock standards are prepared for analysis by 

adding 0.1 mL of beryllium stock standard into 1.9 mL of detection solution (20-fold dilution). 

Please see Table 1. Either of the two detection solutions may be used. 

NOTE: To test a range of 0.005 μg to 0.4 μg of beryllium on the sampling media, a recommended series 

of stock standard solutions is (80, 20, 4, 1, and 0) ng/mL. These standards with lower beryllium 

concentration can be prepared by 10-fold dilution of the stock standards mentioned in the note 

above. As with the samples, these stock standards are prepared for analysis by adding 0.4 mL of 

beryllium stock standard into 1.6 mL of detection solution (5-fold dilution). Please see Table 2. 

Either of the two detection solutions may be used. 

NOTE: When using the lysine-free dye solution ONLY, To test a range can be tested of 0.0005 to 0.4 μg 

of beryllium on the media using a recommended series of stock solutions is (0, 0.15, 0.3, 0.6 and 

2.4) ng/ml. These standards with lower beryllium concentration can be prepared by dilution of 

the stock standards mentioned in the note above. The standards are prepared for analysis by 

adding 0.67 mL of beryllium stock standard into 1.33 mL of detection solution (3-fold dilution). 

This dilution will result in 0, 0.05, 0.1, 0.2 and 0.8 ppb of beryllium in these standards. Please see 

Table 3. MCE air filters and cellulosic capsules with MCE filters were spiked with a solution of 

beryllium acetate and analyzed in triplicate after extracting beryllium in 5ml of 1% ABF solution 

at 85oC for 60 minutes and then mixed with lysine-free dye solution in a 3-fold dilution. The 

results are shown in Table 4. The difference in the average fluorescent signals from blanks and 

the 0 ppb standard were subtracted from the fluorescent readings of the spiked filters.

NOTE: If alternative ratios of dissolution solution and detection solution are used for sample 

preparation, then a similar ratio for calibration solutions is required.

9. Analyze a stock standard, a reagent blank, and a media blank at least once every 20 samples. Ensure 

that the concentration range of the stock standards spans the beryllium levels found in the samples. 

10. Analyze one media spike and one quality control blind spike per 20 samples (minimum of three each 

per sample set) to insure that percent recovery is in control (e.g., 100 ± 15). Correct sample results for 

the average recovery if it differs significantly from 100 %.

NOTE: If it is suspected that beryllium oxide may be present, then it is recommended to use beryllium 

oxide for media and blind spikes.

MEASUREMENT:

11. For each sample, obtain the fluorescence intensity at λmax or with optical filter for appropriate wavelength. 

12. If the fluorescence response for any of the samples is above the range of responses for the stock 

standards, dilute the sample filtrate with dissolution solution, reanalyze, and apply the appropriate 

dilution factor (D) in subsequent calculations. 
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 5 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALCULATIONS:

13. Obtain the solution concentration for each sample filtrate, Cs (ng/mL), and the average media blank, Cb

(ng/mL) from the calibration graph. 

14. Using the dissolution volumes (normally 5 mL) of sample, Vs (mL), and media blank, Vb (mL), calculate 

the concentration, C (μg/m3

), of Be in the air volume sampled, V (L), while accounting for the dilution 

factor (D). 

C=D X (CsVs-CbVb)/V ng/L or μg/m3

NOTE: Tables 1, 2 and 3 can be used for correlating the amount of beryllium in the sampling media 

with the concentrations of beryllium in solution. Table 1 is for testing media with 0.2 μg to 4 μg 

of beryllium at 20-fold dilution; Table 2 is for testing media with 0.02 μg to 0.4 μg of beryllium at 

5-fold dilution; and Table 3 is for testing media with 0.0005 μg to 0.012 μg of beryllium at 3-fold 

dilution. Lysine-free dye solution may be used for any of these dilutions, but for 3x dilution, 

lysine-free dye solution must be used.

Table 1. Correlation of amount of Be in sampling media with Be concentration in stock standard and 

Be concentration as analyzed, assuming 0.1 mL of sample or stock standard is added to 1.9 mL of 

detection solution (20-fold dilution).

Be concentration in stock 

standard (ng/mL)

Be concentration as analyzed 

(ng/mL)

Amount of Be in the media* 

(ng)

0 0 0 

10 0.5 50

40 2 200

200 10 1000

800 40 4000

*Equals stock standard Be concentration (ng/mL) × volume (5 mL) of dissolution solution used to extract 

media.

Table 2. Correlation of amount of Be in sampling media with Be concentration in stock standard and 

Be concentration as analyzed, assuming 0.4 mL of sample or stock standard is added to 1.6 mL of 

detection solution (5-fold dilution).

Be concentration in stock 

standard (ng/mL)

Be concentration as 

analyzed (ng/mL)

Amount of Be in the media* 

(ng)

0 0 0 

1 0.2 5 

4 1 25

20 4 100

80 16 400

*Equals stock standard Be concentration (ng/mL) × volume (5 mL) of dissolution solution used to extract 

media.
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 6 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 3. Correlation of amount of Be in sampling media with Be concentration in stock standard and 

Be concentration as analyzed, assuming 0.67 mL of sample or stock standard is added to 1.33 mL of 

lysine-free dye solution (3-fold dilution). 

Be concentration in stock 

standard (ng/mL)

Be concentration as 

analyzed (ng/mL)

Amount of Be in the media* 

(ng)

0 0 0 

0.15 0.05 0. 75

0.3 0.1 1.5

0.6 0.2 3 

2.4 0.8 12

*Equals stock standard Be concentration (ng/mL) × volume (5 mL) of dissolution solution used to extract 

media.

Table 4: Analysis of beryllium spiked MCE filters and cellulosic filter capsule inserts using 3-fold 

dilution of the extraction solution with lysine-free dye solution. Beryllium concentration of standard 

solutions after mixing with the dye solutions were 0, 0.05, 0.1, 0.2 and 0.8ng/ml. Samples analyzed 

in triplicate, averages and standard deviations shown.

Nominal Be concentration 

on the spiked filter, μg

Be concentration as analyzed 

on the spiked MCE filter (μg)

Be concentration as analyzed 

on the spiked cellulosic

internal capsule insert (μg)

0 0.0000±1.2E-5 -0.0001±3.7E-5 

0.0005 0.00044±1.9E-5 0.0005±3.6E-5 

0.001 0.0009±2.4E-5 0.0009±2.6E-5 

0.002 0.0018±4.0E-5 0.0020±1.1E-4 

0.005 0.0043±1.3E-5 0.0045±6.4E-4 

0.05 0.045±5.5E-4 0.046±1.0E-3 

0.48 0.42±3.5E-3 0.45±4.0E-3 
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 7 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 5. Other published OELs and guidelines†

Country or organization 8 Hour inhalation 

exposure limit (μg/m3) 

Short term inhalation 

exposure limit (μg/m3) 

Germany 0.06 (respirable); 0.14 

(inhalable)

0.06 (respirable); 0.14 

(inhalable)

ACGIH TLV® * 0.05 

Australia, France, New Zealand, Singapore, 

Sweden, Switzerland, United Kingdom

2 

Canada (Quebec) 0.15 

Ireland, Poland, Spain 0.2 

Czech Republic, Japan, Latvia, Norway 1 

Finland 0.1 0.4

China 0.5 0.1

Austria 2 0.8

Denmark 1 2 

Hungary 2 2 

Belgium, Canada (Ontario), Korea (Republic of) 2 10

*Abbreviations: ACGIH = American Conference of Governmental Industrial Hygienists, TLV® = Threshold

Limit Value

†Occupational exposure limits and guidelines other than NIOSH’s recommended exposure limit (REL) have 

not been reviewed by NIOSH. Professional society and other country exposure limits and guidelines are 

provided as an aid to NMAM users seeking additional information. Inclusion of these standards and 

guidelines does not constitute endorsement by NIOSH.

EVALUATION OF METHOD:

The method was evaluated [1,2,7] in accordance with published guidelines [9]. Experiments were 

conducted using an Ocean Optics® portable fluorescence device with the following components:

USB 200 spectrometer with spectral grating #2 (UV/Vis 600), LS-1 lamp (380 nm) in LS-450 housing,

UV-2 casting, OFLV linear filter 200-850, L2 collection lens and slit-200.

Tests were carried out in relative irradiance mode using 2- or 5-second integration times.

The method was evaluated using beryllium oxide spiked onto mixed cellulose ester (MCE) filters at levels of 

(0, 0.02, 0.1, 0.2, 0.3, 0.4, 1.5, 3.0, and 6.0) μg (five samples at each level) [2,6].

Long-term stability of samples was verified from spikes (number [n] = 30) of 0.1 μg Be on MCE filters. Samples were analyzed at day one (n = 12) and then one week (n = 6), ten days (n = 3), two weeks (n = 3), three 

weeks (n = 3), and one month (n = 3) after spiking. No diminution of fluorescence signal was observed from 

samples prepared and analyzed after having been stored for up to thirty days.

Interference tests were carried out using solutions of 0 nmol/L, 100 nmol/L, and 1.0 μmol/L Be in the presence of 0.4 mmol/L Al, Ca, Co, Cu, Fe, Ti, Li, Ni, Pb, Sn, U, V, W, or Zn (separate experiments were carried out 

for each potential interferant). Interlaboratory evaluations of the method were also performed [10, 11].

The method using the lysine-free detection solution on MCE filters and SOLU-SERTTM internal capsules was 

compared and tested (with the detection solution containing lysine for comparison) in Table 4 and this was 

carried out on a GlomaxTM spectrometer (From Turner Biosystems, Sunnyvale, CA) with an emission filter of 

475 ± 5 nm and the excitation was at 360 nm.
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 8 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] Agrawal A, Cronin J, Tonazzi J, McCleskey TM, Ehler DS, Minogue EM, Whitney G, Brink C, Burrell AK, 

Warner B, Goldcamp MJ, Schlecht PC, Sonthalia P, Ashley K [2006]. Validation of a standardized portable fluoresence method for determining trace beryllium in workplace air and wipe samples. J 

Environ Monit 8: 619- 624. 

[2] Ashley K, Agrawal A, Cronin J, Tonazzi J, McCleskey TM, Burrell AK, Ehler DS [2007]. Ultra-trace determination pf beryllium in occupational hygiene samples by ammonium bifluoride extraction and 

fluorescence detection using hydroxybenzoquinoline sulfonate. Anal Chim Acta 584: 281-286. 

[3] NIOSH [1994]. Elements by ICP (nitric/perchloric acid ashing): Method 7300 (supplement issued 

3/15/03). In: NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health 

and Human Services, Public Health Service, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 94–113. 

[4] ASTM [2015]. D7202 – Standard test method for determination of beryllium in the workplace using 

field-based extraction and fluorescence detection. West Conshohocken, PA: ASTM International. 

[5] Matsumiya H, Hoshino H, Yotsuyanagi T [2001]. A novel fluorescence reagent, 10-hydroxybenzo[h]quinoline-7-sulfonate, for selective determination of beryllium(II) ion at pg cm−3 levels. 

Analyst 126:2082–2086. 

[6] Goldcamp MJ, Goldcamp DM, Ashley K, Fernback JE, Agrawal A, Millson M, Marlow D, Harrison K 

[2009]. Extraction of beryllium from refractory beryllium oxide with dilute ammonium bifluoride and 

determination by fluorescence – A multi-parameter performance evaluation. J Occup Environ Hyg 6: 

735-744.

[7] McCleskey TM, Agrawal A, Ashley K [2007]. Backup data—method nos. 7704 and 9110 / beryllium, 

Issue 1. NIOSH Docket Office, Cincinnati, Ohio, e-mail: NIOSHDocket@cdc.gov. 

[8] Agrawal A, Adams LL, Agrawal A, Cronin JP, Tonazzi JCL, Duran BL [2011]. Beryllium measurement by 

optical fluorescence in samples contaminated by fluorescent impurities. J ASTM Int 8(8), 14 pp. [JAI 

103411]. 

[9] Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA [1995]. Guidelines for air sampling and analytical method development and evaluation. Cincinnati, OH: U.S. Department of Health and Human 

Services, Public Health Service, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 95–117. 

[10] Ashley K, McCleskey TM, Brisson MJ, Goodyear G, Cronin J, Agrawal A [2005]. Interlaboratory 

evaluation of a portable fluorescence method for the measurement of trace beryllium in the 

workplace. J ASTM Int 2(9), 8 pp., Paper ID JAI13156. 

[11] Oatts TJ, Hicks CE, Adams AR, Brisson MJ, Youmans-McDonald LD, Hoover MD, Ashley K [2012]. 

Preparation, certification and interlaboratory analysis of workplace air filters spiked with high-fired 

beryllium oxide. J Environ Monit 14: 391-401.
 BERYLLIUM in Air by Fluorometry: METHOD 7704, Issue 2, 12 December 2015 - Page 9 of 9 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD REVISED BY:

T. Mark McCleskey, Los Alamos National Laboratory, Los Alamos, NM; Kevin Ashley, CDC/NIOSH, Division of 

Applied Research and Technology; Anoop Agrawal, Berylliant Inc., Tucson, AZ.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATE FLUORIDES and HYDROFLUORIC ACID 7906

by Ion Chromatography 

MW: F-, 18.998 (HF, 20.01) CAS (HF): 7664-39-3 RTECS: (HF) MW7875000

METHOD:  7906, Issue 2 EVALUATION: FULL Issue 2: 20 May 2014 

Issue 1: 15 August 1994 

U. S. OELs

 OSHA : 2.5 mg/m3

 (fluorides); 3 ppm (HF)

NIOSH: 2.5 mg/m3

 (fluorides); 

3 ppm (HF), STEL (HF): 6 ppm

Other OELs: [1,2]

PROPERTIES: HF: gas; mp = -83.1 °C; bp = 19.5 °C; sp. gr. 

0.987; VP = 783 mm Hg (104.4 kPa).

Fluoride salts liberate HF vapor in the presence of acids.

SYNONYMS: Hydrogen fluoride; hydrofluoric acid; Sodium fluoride (CAS #7681-49-4)

SAMPLING

SAMPLER: PRE-FILTER, cellulose nitrate, 0.8 μm pore size, 

37-mm diameter, + TREATED FILTER, cellulose 

nitrate impregnated with Na2

CO3, 0.8 μm 

pore size, 37-mm diameter, separated by a 

chemically inert spacer

FLOW RATE: 1 to 2 L/min 

VOL-MIN: 15 L 

-MAX: 1000 L

SHIPMENT: 4 °C

SAMPLE 

STABILITY: Stable for at least 14 days (fluorides) / 28 days 

(HF) if stored at @ 20 °C for 7 days and 4 °C 

thereafter [1]

BLANKS: 3 minimum field blanks per set

ACCURACY*

RANGE STUDIED: 0.04 to 5.9 mg/m3

 (fluorides);

0.30 to 6 ppm (HF)

BIAS*: Negligible [3]

OVERALL

PRECISION*( S

ˆ

rT ): Fluorides, 0.074; HF, 0.125 [5,6]

EXPANDED

UNCERTAINTY*: less than 20% for F-

; 

less than 22% for HF [4,5] 

*Accuracy calculations were determined using reference 4

 rather than the traditional NIOSH accuracy criterion.

MEASUREMENT 

TECHNIQUE: ION CHROMATOGRAPHY (IC) with 

conductivity detection

ANALYTE: Fluoride ion (F-

)

EXTRACTION: Aqueous solution of sodium carbonate / 

sodium hydrogen carbonate

INJECTION

VOLUME: 50 μL

ELUENT: 8 mM Na2

CO3 / 1 mM NaHCO3

 flow rate 1.0 mL/min

COLUMNS: Pre-column, anion-exchange column and 

suppressor column 

CALIBRATION 

RANGE: F-

; 0.4 to 8.0 mg/L [3,5]

ESTIMATED LOD: 0.1 mg/L / 0.009 mg/m3

 (120 L air volume; 

as F-

) [3] 

PRECISION ( S ): Fluorides, 0.033; HF, 0.056 [3] r

APPLICABILITY: The working range is (at least) 0.1 to 8 mg/sample for a 250 Liter air sample.

INTERFERENCES: Co-sampled formate and acetate compounds in the work environment can cause a positive interference; 

ve electrodes is a suitable alternative if airborne formate and acetate are expected. Cations that 

such as Ca2+, Fe3+, and Al3+ can cause negative interferences. Recovery of gaseous HF is reduced at high 

thus detection by ion selecti

form insoluble fluorides, 

humidity.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATE FLUORIDES and HYDROFLUORIC ACID: METHOD 7906, Issue 2, dated 20 May 2014 - Page 2 of 6

OTHER METHODS: This method updates NIOSH 7906 [7] by providing full method accuracy information, 

and specifies the use of an impregnated filter instead of a treated cellulose pad. The procedure, which 

is consistent with ISO 21438-3 [6], replaces NIOSH 7903 [8] for the determination of particulate fluorides 

and gaseous HF in workplace air samples. The advantages of this method are that it can allow for the 

collection of the inhalable fraction of particulate fluorides by means of the pre-filter (housed within an 

optional inhalable sampler) and it can provide for lower limits of detection and quantitation for gaseous 

HF due to higher sampling flow rates. NIOSH 7902 [9] and ASTM D4765 [10] are alternative methods that 

are recommended for insoluble fluorides, which rely on detection of F- by ion-selective electrode.

REAGENTS:

1. Water, deionized (DI), ≥18 MΩ-cm resistivity

2. Sodium carbonate (Na2

CO3), anhydrous,

American Chemical Society (ACS) analytical

grade

3. Sodium hydrogen carbonate (NaHCO3

), ACS

analytical grade

4. Filter impregnation solution (0.75 M Na2

CO3);

Dissolve 7.95 g Na2

CO3 in deionized water

and swirl to mix. Then bring to 100 mL in a

volumetric flask, stopper and mix thoroughly.

5. Eluent stock solution: 0.8 M Na2

CO3 / 0.1 M

NaHCO3

; Dissolve 16.96 g Na2CO3 and 1.68 g

NaHCO3

 in deionized water and swirl to mix.

Then bring to 200 mL in a volumetric flask,

stopper and mix thoroughly.

6. Eluent solution: 8 mM Na2

CO3 / 1 mM NaHCO3;

Transfer 10 mL of 0.8 M Na2

CO3 / 0.1 M

NaHCO3

stock solution to a 1 Liter volumetric

flask, dilute to the mark with deionized water,

stopper and mix thoroughly. Prepare fresh

eluent solution weekly.

7. Fluoride (F-

) standard solution, 1000 mg/L

8. Calibration stock solution, 100 mg/mL (as

the anion): Place a 10 mL aliquot of fluoride

standard solution into a 100 mL volumetric

flask, dilute to the mark and mix thoroughly.

The solution is stable for at least 4 weeks at

room temperature.

9. To prepare the impregnated filter, evenly

distribute 120 μL 0.75 M Na2

CO3 over the

surface of the filter and allow to dry at room

temperature for several hours.

*See Special Precautions

EQUIPMENT: 

1. Sampler: 37-mm diameter cellulose

nitrate pre-filter, 0.8 μm pore size, 37-mm

impregnated cellulose nitrate filter. The

untreated filter collects particulate fluorides,

while the impregnated filter collects HF

vapor. Arrange the filters serially (with the

pre-filter upstream of the impregnated filter)

in a 37-mm diameter 3-piece chemically inert

cassette filter holder.

2. Personal sampling pump, 1 to 2 L/min, with

flexible connecting tubing

3. Ion chromatograph (IC), with pre-column (50

mm by 4.0 mm), anion-exchange column (200

mm by 4.0 mm), suppressor column (4 mm)

and conductivity detector; and connected to

data processing unit

4. Ultrasonic bath

5. Vessels, 10-mL, inert plastic, with screw caps

6. Volumetric flasks, 25- to 1000 mL

7. Pipets, 50- to 10000 μL

8. Beakers, 25- to 100 mL

9. Filtration apparatus, with filter funnel

10. Disposable filters, cellulosic, 0.45 μm pore size

11. Styrene-based sulfonic acid resin cartridges

12. Water purification system, to prepare greater

than or equal to 18 MΩ-cm resistivity

deionized water

13. Bottles, polyethylene, 100 mL

14. Syringes, plastic, 5 mL and/or 10 mL

15. Syringe filter cartridges, with 0.8-μm pore size

polytetrafluoroethylene (PTFE) membrane

filters

16. Micro-syringes, 50 μL, with 60 mm x 0.6 mm

needles

17. Auto-sampler vials, polypropylene, 0.75 mL

capacity.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATE FLUORIDES and HYDROFLUORIC ACID: METHOD 7906, Issue 2, dated 20 May 2014 - Page 3 of 6

SPECIAL PRECAUTIONS: Acids, particularly HF, are extremely corrosive to skin, eyes, and mucous 

membranes. HF will attack glass. Plastic labware is recommended. Wear gloves, lab coat, and safety 

glasses while handling acids. All work should be performed with adequate ventilation for personnel 

and equipment. It is imperative that acid be added to water in order to avoid a violent exothermic 

reaction [11,12].

SAMPLING, SAMPLE TRANSPORT AND STORAGE:

1. Prior to sampling, load each clean sampler, first with a sampling (impregnated) filter, then with a prefilter, separating the filters with a spacer. Ensure that the configuration in which the filters are loaded

leads to the sampled air passing first through the pre-filter and then through the sampling filter.

2. Calibrate each personal sampling pump with a representative sampler in the line.

3. Sample at an accurately known flow rate between 1 to 2 mL/min for a total sample size of 15 to

1000 L. Avoid sampler overloading.

4. After sampling, remove the filters from the cassette and place them in screw-cap plastic vessels. For

the pre-filter portion: With approximately 2 mL eluent(extraction) solution, rinse material from the

inside surfaces of the pre-filter portion of the cassette into the vessel. Add eluent solution into the

vessels until a final volume of 10 mL is reached.

5. Submit at least three blank untreated filters and three blank impregnated filters as field blanks for

each set of samples collected per day. Handle these in the same way as the field samples; i.e., place

each filter into a vessel, add 10 mL of eluent solution and ship it to the lab along with the remaining

samples.

6. Refrigerate all samples that are to be stored overnight (or longer) prior to shipment to the

laboratory. Ship all samples to the laboratory in accordance with established chain-of-custody

procedures [13].

7. Refrigerate the samples immediately upon receipt at the lab until ready for analysis.

8. Analyze samples within 2 weeks of receipt. The samples can be stored at room temperature for one

week; for longer storage, refrigerate the samples (4 °C).

SAMPLE PREPARATION AND MEASUREMENT: 

9. Remove sample vessels from storage and bring them to room temperature.

10. Sonicate the samples in an ultrasonic bath for at least 15 minutes and allow to cool for at least 30

minutes.

11. Using 5- or 10-mL syringes, filter each sample extract solution through a PTFE filter and a styrenebased sulfonic acid resin cartridge (follow the manufacturer’s instructions), discard the first two

milliliters and place the remaining solution in clean plastic vessels.

CALIBRATION AND QUALITY CONTROL: 

12. With dilution of the calibration stock solution in eluent solution, prepare calibration working

standard solutions covering the range of approximately 0.4 to 8 μg/mL of fluoride. Store working

standards in tightly sealed polyethylene bottles. Prepare fresh working standards biweekly.

13. Calibrate the ion chromatograph with at least six working standards covering the range of (at least)

0.4 to 8 μg/mL of fluoride per sample by preparing a calibration graph of anion peak height (mm or

μS) vs. concentration (μg/mL).

14. Analyze working standards together with samples, reagent blanks and field blanks at a frequency of

at least 1 per 20 samples (3 minimum of each). Cellulose nitrate filters demonstrate variable batchdependent blank values, thus media blank correction is essential.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATE FLUORIDES and HYDROFLUORIC ACID: METHOD 7906, Issue 2, dated 20 May 2014 - Page 4 of 6

MEASUREMENT

15. Set the ion chromatograph to recommended eluent flow rate, (e.g. 1.0 mL/min or approximately

13 MPa pressure), and other conditions as specified by the instrument manufacturer.

16. Inject a sample aliquot (e.g. 50 μL) into the chromatograph and measure the peak height of the

fluoride peak. If the peak height exceeds the linear calibration range, dilute with eluent, reanalyze

and apply the appropriate dilution factor in calculations.

CALCULATIONS: 

17. Calculate the mass concentration of fluoride, C (mg/m3), in the air volume sampled, V (L):

C= (C V F )-(C V )

V* { }*F 1* 1* d 0* 0

c

h

where:

C0

 = mean concentration, in μg/mL, of fluoride in the field blank test solutions;

C1

 = concentration, in μg/mL, of fluoride in the sample test solution;

V = volume, in liters, of the air sample;

V0

 = volume, in mL, of the field blank test solutions; 

V1

 = volume, in mL, of the sample test solutions;

Fd

 = dilution factor for each sample test solution;

Fc = 1.053 = conversion factor to convert from fluoride to HF concentration (if applicable);

η = recovery.

EVALUATION OF METHOD: 

This updates Issue 1, NIOSH 7906 [7], by providing full method accuracy information, and specifies 

the use of an impregnated filter instead of a treated cellulose pad. The procedure described in 

issue 2 of NIOSH 7906, which is consistent with ISO 21438-3 [6], also replaces NIOSH 7903 [8] for the 

determination of particulate fluorides and gaseous HF in workplace air samples. 

Recovery of 100% ± 2% was found after sample collection for both F-

 and HF [3,5,6], and no 

breakthrough of HF was observed from impregnated filters for up to 5 mg fluoride [6,14]. The 

component of the coefficient of variation of the method that arises from analytical variability, 

determined from the analysis of spiked filters, was 2.4% to 5.6% for HF and 1.7% to 3.3% for fluorides 

[3,6]. Laboratory testing with test atmospheres of HF (0.5 to 5 mg/m3

) demonstrated quantitative 

sampling efficiencies (greater than 95%) at relative humidity (RH) of 20% to 60%, but recovery was 

around 60% at relative humidity of 80%. 

The recovery of HF as an effect of relative humidity (RH) greater than 60 percent can be calculated as 

follows (see also Figure 1): 

h= 

[226.5-(2.0914 * RH)]

100 

No effect of humidity (RH from 20% to 80%) was observed on recovery of particulate fluoride over the 

concentration range of 0.3 to 5 mg/m3

. Results for repeatability and reproducibility of the method for 

test atmospheres of hydrogen fluoride and fluoride aerosol (Relative humidity was approximately 50%) 

are shown in Table I. The method has been successfully applied in independent investigations [14] and 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATE FLUORIDES and HYDROFLUORIC ACID: METHOD 7906, Issue 2, dated 20 May 2014 - Page 5 of 6

has been promulgated as an International Standard [6]. The back-up data and user check reports are 

references 3 and 14 respectively.

REFERENCES: 

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for 

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: 

American Conference of Governmental Industrial Hygienists. [www.acgih.org]. Data accessed April 

2014.

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database 

on international limit values for hazardous substances (German Social Accident Insurance). Sankt 

Augustin, FRG: [http://www.dguv.de/ifa/GESTIS]. Data accessed April 2014.

[3] Breuer D, Gusbeth K [2005]. Hydrogen fluoride and fluorides. In: The MAK-Collection Part III: Air 

monitoring methods. (Deutsche Forschungsgemeinschaft) Vol. 9. Weinheim, FRG: Wiley-VCH. pp. 

105-118.

[4] ISO (International Organization for Standardization) [1995]. ISO Guide 98: Guide to the expression 

of uncertainty in measurement. Geneva, Switzerland: ISO).

[5] Breuer D [2006]. Fluoride und fluorwasserstoff (Fluorides and hydrogen fluoride), in BGIA Working 

Folder, Sheet No. 7512. Berlin, FRG: Erich-Schmidt Verlag. ISBN 3-503-020853.

[6] ISO (International Organization for Standardization) [2010]. ISO 21438-3, Workplace air — 

Determination of inorganic acids by ion chromatography — Part 3: Hydrofluoric acid and 

particulate fluorides. Geneva, Switzerland: International Organization for Standardization (ISO).

[7] NIOSH [1994]. Fluorides, aerosol and gas, by IC: Method 7906. In: Eller PM, Cassinelli ME, eds. NIOSH 

Manual of analytical methods, 4th ed., Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety 

and Health, DHHS (NIOSH). Publication No. 94-113. [www.cdc.gov/niosh/nmam].

[8] NIOSH [1994]. Acids, inorganic: Method 7903. In: Eller PM, Cassinelli ME, eds. NIOSH Manual of 

analytical methods, 4th ed., Cincinnati, OH: U.S. Department of Health and Human Services, Centers 

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH). Publication No. 94-113. [www.cdc.gov/niosh/nmam].

[9] NIOSH [1994]. Fluorides, aerosol and gas, by ISE: Method 7902. In: Eller PM, Cassinelli ME, eds. 

NIOSH Manual of analytical methods, 4th ed., Cincinnati, OH: U.S. Department of Health and Human 

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and 

Health, DHHS (NIOSH). Publication No. 94-113. [www.cdc.gov/niosh/nmam].

[10] ASTM International [2008]. ASTM D4765, Standard test method for fluorides in workplace 

atmospheres by ion-selective electrodes. West Conshohocken, PA: ASTM International.[11] 

NIOSH [2005]. NIOSH Pocket guide to chemical hazards. Barsan ME, ed. Cincinnati, OH: US 

Department of Health and Human Services, Centers for Disease Control and Prevention, National 

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149.

[12] Furr AK, ed. [1995] CRC Handbook of laboratory safety, 4th ed. Boca Raton, FL: CRC Press.

[13] ASTM International [2010]. ASTM D4840, Standard Guide for Sampling Chain of Custody 

Procedures. West Conshohocken, PA: ASTM International.

[14] Demange M, Oury V, Rousset D [2011]. Evaluation of sampling methods for measuring exposure 

to volatile inorganic acids in workplace air. Part 2 – sampling capacity and breakthrough tests for 

sodium carbonate-impregnated filters. J Occup Environ Hyg. 8:642-651.

[15] Comité Européen de Normalisation (CEN) [2012]. EN 482, Workplace exposure — General 

requirements for the performance of procedures for the measurement of chemical agents. Brussels, 

Belgium: Comité Européen de Normalisation (CEN).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

PARTICULATE FLUORIDES and HYDROFLUORIC ACID: METHOD 7906, Issue 2, dated 20 May 2014 - Page 6 of 6

METHOD WRITTEN BY: 

Dietmar Breuer, Dr. rer. nat., and Petra Heckmann, Institute for Occupational Safety and Health of the 

German Social Accident Insurances, Sankt Augustin, Germany; and Kevin Ashley, Ph.D., NIOSH/DART

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for 

the content of these websites. All web addresses referenced in this document were accessible as of 

the publication date.

FIGURE 1. HF recovery as function of relative humidity

TABLE 1. Performance evaluation for HF and F- ion measurements from test atmospheres [5]

Concentration 

(mg/m3

) Analyte S 1

r (n = 6) RSD2

(%) U3

HF 0.25 0.007 3.1 0.22

HF 1.2 0.044 2.8 0.21

HF 2.3 0.026 1.1 0.20

HF 5.0 0.099 2.0 0.11

Fluoride 0.04 0.002 3.8 0.19

Fluoride 0.52 0.006 1.3 0.12

Fluoride 2.95 0.009 0.3 0.11

Fluoride 5.9 0.065 1.1 0.11

1 Standard deviation for 6 test samples

2 Relative standard deviation

3 Expanded measurement uncertainty; k = 2 (calculated in accordance with EN 482 [15])
VOLATILE ACIDS by Ion Chromatography 7907

(Hydrogen Chloride, Hydrogen Bromide, Nitric Acid)

Formulae: 1. HCl; 

 2. HBr; MW: Table 1 CAS: Table 1 RTECS: Table 1 

 3. HNO3 

METHOD: 7907, Issue 1 EVALUATION: FULL 

 

 Issue 1: 20 May 2014

 OSHA : Table 2

NIOSH: Table 2

Other OELs: [1,2]

PROPERTIES: Table 3

SYNONYMS: 1) HCl: Hydrochloric acid, salt acid, muriatic acid; 2) HBr: Hydrobromic acid; 3) HNO3: azotic acid

SAMPLING

SAMPLER: Two (2) FILTERS in series separated with a 

chemically inert spacer: (1) pre-filter: 37-mm 

diameter quartz fiber filter; (2) sampling filter: 

37-mm diameter quartz fiber filter impregnated 

with 500 μl Na2

CO3 solution (1 M)

FLOW RATE: 2 L/min 

VOL-MIN: 30 L 

-MAX: 600 L

SHIPMENT: Routine

SAMPLE

STABILITY: Stable for 1 week @ 20 °C and 4 °C thereafter to 

at least 28 days [3]

BLANKS: 3 blanks minimum per set

ACCURACY*

RANGE STUDIED: HCl: 0.04 to 1.6 mg/sample; 

HBr: 0.03 to 2.3 mg/sample; 

 HNO3

: 0.04 to 1.5 mg/sample

BIAS*: Negligible [4]

OVERALL ˆ PRECISION ( SrT )*: HCl, 0.06; HBr, 0.06; 

 HNO3

, 0.07 [4]

EXPANDED 

UNCERTAINTY*: Less than 12% for HCl and HBr; less 

than 14 % for HNO3

 [3,5]

*Accuracy calculations were determined using ISO Guide 98 

[5] rather than the traditional NIOSH accuracy criterion.

MEASUREMENT

 TECHNIQUE: ION CHROMATOGRAPHY (IC) with 

conductivity detection

ANALYTES: Chloride (Cl-

), bromide (Br -) and nitrate 

(NO -

3 ) ion

EXTRACTION: Aqueous solution of sodium carbonate / 

sodium hydrogen carbonate

INJECTION

VOLUME: 25 μL

ELUENT: 3.1 mM Na2

CO3 / 0.35 mM NaHCO3, flow rate 

1.5 mL/min

COLUMNS: Pre-column, anion-exchange column and 

suppressor column 

CALIBRATION 

RANGE: Chloride, bromide and nitrate at 0.4 mg/mL 

to 4 mg/mL 

ESTIMATED LOD: 0.003 mg/sample for all three acids; or 

 0.0012 mg/m3

 for a 240 L air volume [4] 

PRECISION ( S ): 0.01 for all three acids [3] r

APPLICABILITY: The working range is (at least) 0.01 to 2.0 mg/sample for a 240 Liter air sample [3,4].

INTERFERENCES: Inorganic acids can react with co-sampled particulate matter on the pre-filter, leading to low results. One 

such example is in the galvanizing industry, where the presence of zinc oxide can be a major confounding factor in the measurement of HCl. Potentially interfering particulate chlorides and nitrates removed by the pre-filter can react with the sampled 

acids and liberate HCl and HNO3

 that is subsequently collected on the sampling filter, leading to high results [6].
VOLATILE ACIDS by Ion Chromatography: METHOD 7907, Issue 1, dated 20 May 2014 - Page 2 of 6

OTHER METHODS: This procedure, which is consistent with ISO 21438-2 [4], replaces NIOSH 7903 [7] 

for the determination of HCl, HBr and HNO3

 in workplace air samples by IC. The main advantage of this 

method is that it provides for lower limits of detection and quantitation for volatile acids due to higher 

sampling flow rates.

REAGENTS:

1. Water, deionized (DI), ≥18 MΩ-cm resistivity

2. Sodium carbonate (Na2

CO3), anhydrous, 

American Chemical Society (ACS) analytical 

grade

3. Sodium hydrogen carbonate (NaHCO3

), ACS 

analytical grade

4. Extraction & eluent stock solution: 0.62 M 

Na2

CO3 / 0.069 M NaHCO3; Dissolve 13.14 g 

Na2

CO3 and 1.15 g NaHCO3 in 50 mL of 

deionized water and swirl to mix. Then bring 

to 200 mL in a volumetric flask, stopper and 

mix thoroughly.

5. Extraction and eluent solution; 0.0031 M 

Na2

CO3/ 0.00035 M NaHCO3: transfer 10 mL 

0.62 M Na2

CO3 / 0.069 M NaHCO3 stock 

solution to a 2 liter volumetric flask, dilute to 

the mark with deionized water, stopper and 

mix thoroughly.

6. Chloride (Cl-

), bromide (Br -) and nitrate (NO -

3 ) 

standard solutions, 1000 mg/L.

7. Calibration stock solution, 100 μg/mL (as the 

anion): Transfer 10 mL aliquots of chloride, 

bromide and nitrate standard solution into 

a 100 mL volumetric flask, dilute to the 

mark with deionized water, stopper and mix 

thoroughly.

8. Sodium carbonate solution, for impregnation 

of 37 mm diameter quartz fiber filters, 

1 mol/L: dissolve 10.6 g Na2

CO3 in deionized 

water, quantitatively transfer the solution 

into a 100 ml volumetric flask, dilute to the 

mark with deionized water, stopper and mix 

thoroughly.

9. Preparation of the sampling filter: Apply 

500 μL of the sodium carbonate solution, 

1 mol/L, evenly on the 37 mm quartz 

fiber filter and allow to dry for 6 hours in a 

desiccator. (See NOTE 1, Step 1, p 7907-3.) 

Good for 14 days.

*See Special Precautions

EQUIPMENT: 

1. Sampler: filter, 37-mm diameter quartz fiber 

impregnated with 500 μl 1 M Na2

CO3; and 

pre-filter, 37-mm diameter quartz fiber filter 

separated by a spacer in a chemically inert 

cassette filter holder

2. Spacers, of 37-mm diameter suitable for use 

with the samplers, manufactured from a 

chemically inert material, e.g. polypropylene 

sleeves

3. Personal sampling pump, 1 to 5 L/min, with 

flexible connecting tubing

4. Ion chromatograph (IC), with pre-column 

(50 mm by 4.0 mm), anion-exchange column 

(200 mm by 4.0 mm), suppressor column 

(4 mm) and conductivity detector; and 

connected to data processing unit

5. Ultrasonic bath

6. Vessels, 15-mL, plastic, with screw caps

7. Volumetric flasks, 10- to 2000 mL

8. Pipets, 10 to 5000 μL

9. Beakers, 25 to 100 mL

10. Water purification system, to prepare ≥18 

MΩ-cm resistivity deionized water

11. Bottles, polyethylene, 100 mL

12. Syringes, plastic, 5 mL

13. Syringe filter cartridges, with 0.45-μm pore 

size polytetrafluoroethylene (PTFE) membrane 

filters

14. Micro-syringes, 50 μL, with 60 mm x 0.6 mm 

needles

15. Auto-sampler vials, with slotted septum, 2 mL 

capacity

16. Analytical balance, with capability of weighing 

to nearest 0.01 mg 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

VOLATILE ACIDS by Ion Chromatography: METHOD 7907, Issue 1, dated 20 May 2014 - Page 3 of 6

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling acids. All work 

should be performed with adequate ventilation for personnel and equipment. It is imperative that 

acid be added to water in order to avoid a violent exothermic reaction. Avoid direct contact since 

acids are both corrosive and irritants to eyes, skin, and the respiratory system [8,9].

SAMPLING, SAMPLE TRANSPORT AND STORAGE:

1. Prior to sampling, load each clean sampler, first with a sampling (impregnated) filter, then with a prefilter, separating the filters with a spacer. Ensure that the configuration in which the filters are loaded 

leads to the sampled air passing first through the pre-filter and then through the sampling filter.

Note 1: Three-piece polystyrene cassettes are appropriate, with the middle ring section of the cassette 

acting as the spacer. Impregnated filters that are stored in a desiccator can be prepared up to 

14 days prior to their use for sampling.

Note 2: Silica gel sorbent tubes [7] can be used in lieu of impregnated filters, but each sorbent tube 

must be preceded by a pre-filter.

2. Calibrate each personal sampling pump with a representative sampler in the line.

3. Sample accurately at 2 L/min for a total sample size of 30 to 500 L. Avoid sampler overloading [6].

4. Transport the samples to the laboratory in a manner that prevents contamination or damage to the 

filters.

5. Submit at least three blank impregnated filters as field blanks for each set of samples collected per day. 

Handle these in the same way as the field samples.

6. Ship all samples to the laboratory in accordance with established chain-of-custody procedures [10]. 

7. The samples can be stored at room temperature for one week; for longer storage, refrigerate the 

samples (4 °C).

8. Analyze samples between 4 days and 4 weeks of sample collection. 

SAMPLE PREPARATION: 

9. Prior to carrying out sample dissolution, store the sampling filter for at least four days (to avoid nitric 

acid losses).

10. Remove sample cassettes from storage and bring them to room temperature. Discard the pre-filters.

NOTE: If desired, the pre-filters can be analyzed for determination of particulate chlorides, bromides 

and nitrates. If this analysis is carried out, it is recommended to rinse the inside surfaces of the 

sampler with a few mL of DI water so as to include wall deposits along with material collected 

on the prefilter.

11. Place the sampling filters (i.e., impregnated filter samples) in 15-mL plastic screw-cap vessels and add 

10 mL of deionzied water to each sample. Securely cap the vessels.

12. Sonicate the samples in an ultrasonic bath for at least 15 minutes, and allow to cool for approx. 30 

minutes.

13. Using 5-mL syringes, filter 5-mL aliquots of each sample extract solution through a PTFE filter into 

clean auto-sampler vials.

CALIBRATION AND QUALITY CONTROL: 

14. With dilution of the calibration stock solution in eluent solution, prepare calibration working standard 

solutions covering the range of approximately 0.4 to 4 mg/L of chloride, bromide and nitrate. Store 

working standards in tightly sealed polyethylene bottles. Prepare fresh working standards weekly.

15. Calibrate the ion chromatograph with at least six working standards covering the range of 0.4 to 

4 mg/L of chloride, bromide and nitrate per sample by preparing a calibration graph of anion peak 

height (mm or μS [micro-siemens]) vs. concentration (mg/L).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

VOLATILE ACIDS by Ion Chromatography: METHOD 7907, Issue 1, dated 20 May 2014 - Page 4 of 6

16. Analyze working standards together with samples, reagent blanks and field blanks at a frequency of at

least 1 per 20 samples (3 minimum of each).

MEASUREMENT:

17. Set the ion chromatograph to recommended eluent flow rate (e.g., 1.5 mL/min) and recommended

pressure ( e.g., 1.1 x 105

 kPa), and other conditions as specified by the instrument manufacturer.

18. Inject a sample aliquot, e.g., 25-μL, into the chromatograph, and measure the peak heights of

the chloride, bromide and nitrate peaks (at retention times of approx. 5 min, 9 min and 12 min,

respectively). If the peak height exceeds the linear calibration range, dilute with eluent, reanalyze and

apply the appropriate dilution factor in calculations.

CALCULATIONS: 

19. Calculate the mass concentration of each anion, C (mg/m3

), in the air volume sampled, V (L):

where: 

C0

 = mean concentration, in mg/L, of anion in the field blank test solutions;

C1

 = concentration, in mg/L, of anion in the sample test solution;

V = volume, in liters, of the air sample;

V0

 = volume, in mL, of the field blank test solutions; 

V1

 = volume, in mL, of the sample test solutions

Fd

 = dilution factor for each sample test solution

Fc

 = conversion factor to convert from anion to acid concentration (Fc = 1.0284 for chloride, 

1.0126 for bromide, and 1.0163 for nitrate

EVALUATION OF METHOD:

On impregnated quartz fiber filters, greater than 95 % recovery of hydrochloric and nitric acid was found 

four weeks after sample collection, and no breakthrough was observed at sample loadings of up to 2.5 

mg HCl or 5 mg HNO3

 [3,4,11]. Mean analytical recovery determined from the analysis of spiked filters 

has been found to be in the range of 97 to 100% for HBr, HCl and HNO3

 [3,11]. The component of the 

coefficient of variation of the method that arises from analytical variability, determined from the analysis 

of filters sampled at a dynamic test gas apparatus, was 0.8% to 1.3% for hydrogen chloride and 1.1% to 

8.8% for nitric acid; and as determined from the analyses of spiked samples of hydrogen bromide, this 

value was 0.8% to 1.4% [3]. The method has been independently verified for all three acids, in accordance 

with applicable performance criteria [11,12]. An interlaboratory study with 5 participants found negligible 

biases and good agreement for hydrogen chloride and nitric acid at concentrations between 0.6 and 8 

mg/m3

 for HCl and 0.8 and 10 mg/m3 for HNO3 [6]. The back-up data and user check reports are references 

6 and 11, respectively.

REFERENCES: 

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical 

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American 

Conference of Governmental Industrial Hygienists. [www.acgih.org]. [Data accessed April 2014.]

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database 

on international limit values for hazardous substances (German Social Accident Insurance). Sankt 
VOLATILE ACIDS by Ion Chromatography: METHOD 7907, Issue 1, dated 20 May 2014 - Page 5 of 6

Augustin, FRG: [http://www.dguv.de/ifa/GESTIS]. Data accessed April 2014.

[3] Breuer D, Gusbeth K [2007]. Anorganische säuren, flüchtig: bromwasserstoff,

chlorwasserstoff,

salpetersäure (Volatile inorganic acids: hydrobromic, hydrochloric, and nitric acids), IFA Working 

Folder, Sheet No. 6172. Bielefeld, FRG: Erich Schmidt Verlag.

[4] ISO (International Organization for Standardization) [2007]. ISO 21438-2, Workplace atmospheres -

Determination of inorganic acids by ion chromatography - Part 2: Volatile acids, except hydrofluoric

acid (hydrochloric acid, hydrobromic acid and nitric acid). Geneva, Switzerland: International

Organization for Standardization.

[5] ISO (International Organization for Standardization) [1995]. ISO Guide 98: Guide to the expression of

uncertainty in measurement. Geneva, Switzerland: I International Organization for Standardization.

[6] Howe A, Musgrove D, Breuer D, Gusbeth K, Moritz A, Demange M, Oury V, Rousset D, Dorotte M [2011].

Evaluation of sampling methods for measuring exposure to volatile inorganic acids in workplace air.

Part 1: Sampling hydrochloric acid (HCl) and nitric acid (HNO3) from a test gas atmosphere. J Occup

Environ Hyg. 8:492-502.

[7] NIOSH [1994]. Acids, inorganic: Method 7903. In: Eller PM, Cassinelli ME, eds. NIOSH Manual of

analytical methods, 4th ed., Cincinnati, OH: U.S. Department of Health and Human Services, Center for

Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH).

Publication No. 94-113. [www.cdc.gov/niosh/nmam].

[8] NIOSH [2005]. NIOSH Pocket guide to chemical hazards. Barsan ME, ed. Cincinnati, OH: US Department

of Health and Human Services, Centers for Disease Control and Prevention, National Institute for

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149.

[9] Furr AK, ed. [1995] CRC Handbook of laboratory safety, 4th ed. Boca Raton, FL: CRC Press.

[10] ASTM International [2010]. ASTM D4840: Standard guide for sampling chain of custody procedures.

West Conshohocken, PA: ASTM International.

[11] Breuer D [1997]. Volatile inorganic acids (HCl, HBr, HNO3). In: Kettrup A, Greim H, eds. Analyses of

hazardous substances in air (Deutsche Forschungsgemeinschaft). Vol. 6. Weinheim, FRG: Wiley-VCH.

pp. 214-223.

[12] Comité Européen de Normalisation (CEN) [2012]. EN 482, Workplace exposure — General

requirements for the performance of procedures for the measurement of chemical agents. Brussels,

Belgium: Comité Européen de Normalisation (CEN). 

METHOD WRITTEN BY: 

Dietmar Breuer, Dr. rer. nat., and Petra Heckmann, Institute for Occupational Safety and Health of the 

German Social Accident Insurances, Sankt Augustin, Germany; and Kevin Ashley, Ph.D., NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date.
VOLATILE ACIDS by Ion Chromatography: METHOD 7907, Issue 1, dated 20 May 2014 - Page 6 of 6

TABLE 1. General Information

Acid MW CAS Number RTECS Number

HCl 36.46 7647-01-0 MW4025000

HBr 80.91 10035-10-6 MW3850000

HNO3 63.01 7697-37-2 QU5775000

TABLE 2. Occupational Exposure Limits

Acid OSHA NIOSH

HCl Ca 5 ppm, 7 mg/m3 Ca 5 ppm, 7 mg/m3

HBr 8 hr TWAb 3 ppm 10 mg/m3 Ca 3 ppm, 10 mg/m3

HNO3 8 hr TWAb 2 ppm 5 mg/m3 TWAc 2 ppm 5 mg/m3

STELd 4 ppm 10 mg/m3

 a Ceiling Limit

 b Time-Weighted Average

 c Up to 10 hr TWA in a 40 hr work week

 d Short Term Exposure Limit

TABLE 3. Properties

Acid Physical State MP (°C) BP (°C) Vapor Pressure (VP)

HCl gas

liquid

-114

--

-84.9

110

(38%) 28.3 kPa

(azeotropic, 20.2%)

HBr gas -86.9 -66.8 2026.4 kPa

HNO3 liquid -42 -- (68% Nitric) 101.3 kPa
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NON-VOLATILE ACIDS (Sulfuric Acid and Phosphoric Acid) 7908

Formulae: H2SO4 MW: 98.08 CAS: 7664-93-9 RTECS: WS5600000

 H3

PO4 MW: 98.0 CAS: 7664-38-2 RTECS: TB6300000

METHOD: 7908, Issue 1 EVALUATION: FULL Issue 1: 10 May 2014

U.S. OELs

OSHA : 1 mg/m3

 (H2SO4 & H3PO4)

NIOSH: 1 mg/m3

 (H2SO4 & H3PO4); STEL: 3 mg/m3 (H3PO4)

Other

OELs: [1,2]

PROPERTIES:

H2

SO4: Liquid, mp = 3.0 °C (98%); 10 °C (100%); bp = 338 

°C (98%); 330 °C (100%); VP = < 0.0001 kPa @ 

20 °C

H3

PO4: Solid (often used in an aqueous solution), mp = 42.4 

°C; bp = 158 °C; vp = 0.0038 kPa @ 20 °C

SYNONYMS: H2

SO4: hydrogen sulfate, oil of vitriol; H3PO4: ortho-phosphoric acid

SAMPLING

SAMPLER: FILTER, 37-mm diameter quartz fiber; or 

polytetrafluoroethylene (PTFE), 0.45 μm pore 

size

FLOW RATE: 1 to 5 L/min 

VOL-MIN: 15 L 

-MAX: 2000 L

SHIPMENT: 4 °C

SAMPLE 

STABILITY: Stable for 1 week at about 20 °C and at 4 °C 

thereafter to 28 days [3]

BLANKS: 3 field blanks minimum per set

ACCURACY*

RANGE STUDIED: 0.005 to 2.0 mg/sample

BIAS*: Negligible [3]

OVERALL

PRECISION ( 

ˆ )*: H2

SO4 = 0.086; H3PO4 = 0.106

EXPANDED

UNCERTAINTY*: less than 23% for both H2

SO4 and 

 H3

PO4 [6]

* Accuracy calculations were determined using references

 5 and 6 rather than the traditional NIOSH accuracy criteria.

MEASUREMENT

TECHNIQUE: ION CHROMATOGRAPHY (IC) with 

conductivity detection

ANALYTES: Sulfate (SO4

2-) ion, phosphate (PO43-) ion

EXTRACTION: Aqueous solution of sodium carbonate / 

sodium hydrogen carbonate

INJECTION

VOLUME: 50 μL

ELUENT: 2.7 mM Na2

CO3/0.3 mM NaHCO3, flow rate 

1.5 mL/min

COLUMNS: Pre-column, anion-exchange column and 

suppressor column 

CALIBRATION 

RANGE: H2

SO4; 0.2 mg/L to 8 mg/L; 

H3

PO4; 0.8 mg/L to 8 mg/L [6]

ESTIMATED LOD: H2

SO4 = 0.002 mg/m3; 

H3

PO4 = 0.003 mg/m3 (1 m3 air volume)[6] 

PRECISION ( ): H

SO4 = 0.043; 

H3

PO4 = 0.032 [3,7]

APPLICABILITY: The working range for H2SO4 is (at least) 0.002 to 1.0 mg/sample, for H3PO4 is 0.004 to 1.0 mg/sample for 

a 420 Liter air sample [3,6].

INTERFERENCES: Particulate salts of sulfate or phosphate will give a positive interference. 

OTHER METHODS: This procedure, which is consistent with ISO 21438-1 [6], replaces NIOSH 7903 [8] for the determination of sulfuric and phosphoric acid in workplace air samples by IC. The main advantage of NIOSH method 7908 is that it can 

allow for the collection of the inhalable fraction of sulfuric and phosphoric acid aerosols by means of the pre-filter (housed 

within an optional inhalable sampler.)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NON-VOLATILE ACIDS (Sulfuric Acid and Phosphoric Acid): METHOD 7908, Issue 1, dated 20 May 2014 - Page 2 of 5

REAGENTS:

1. Water, deionized (DI), ≥18 MΩ-cm resistivity

2. Sodium carbonate (Na2

CO3), anhydrous, 

American Chemical Society (ACS) analytical 

grade

3. Sodium hydrogen carbonate (NaHCO3

), ACS 

analytical grade

4. Extraction & eluent stock solution: 0.27 M 

Na2

CO3 / 0.03 M NaHCO3; dissolve 2.86 g 

Na2

CO3 and 0.25 g NaHCO3 in 25 mL of 

deionized water and swirl to mix. Then bring to 

100 mL in a volumetric flask, stopper and mix 

thoroughly.

5. Extraction & eluent solution: 0.0027 M Na2

CO3 / 

0.0003 M NaHCO3

; transfer 10 mL of 0.27 M 

Na2

CO3 / 0.03 M NaHCO3 stock solution to a 

1 L volumetric flask, dilute to the mark with 

deionized water, stopper and mix thoroughly.

6. Sulfate (SO4

2-) ion and phosphate (PO43-) ion 

standard solutions, each 1000 mg/L

7. Calibration stock solution, 100 mg/L (as the 

anion): Place 10 mL aliquots of sulfate and 

phosphate standard solution into a 100 mL 

volumetric flask, dilute to the mark and mix 

thoroughly.

*See Special Precautions

EQUIPMENT: 

1. Sampler: filter, 37-mm diameter quartz fiber; or 

polytetrafluoroethylene (PTFE), 0.45 μm pore 

size, in cassette filter holder manufactured 

from acid-resistant (chemically inert) material

NOTE: Quartz fiber filters should be binderless 

and heat-treated. 

2. Personal sampling pump, 1 to 5 L/min, with 

flexible connecting tubing

3. Ion chromatograph, with pre-column (50 mm 

by 4.0 mm), anion-exchange column (200 mm 

by 4.0 mm), suppressor column (4 mm) and 

conductivity detector; and connected to data 

processing unit

4. Ultrasonic bath

5. Vessels, 10-mL, plastic (e.g., polypropylene), 

with screw caps

6. Volumetric flasks, 25- to 1000 mL

7. Pipets, 50 to 10000 μL

8. Beakers, 25 to 100 mL

9. Water purification system, to prepare greater 

than or equal to18 MΩ-cm resistivity deionized 

water

10. Bottles, polyethylene, 100 mL

11. Syringes, plastic, 5 mL

12. Syringe filter cartridges, with 0.8-μm pore size 

polytetrafluoroethylene (PTFE) membrane 

filters

13. Micro-syringes, 50 μL, with 60 mm x 0.6 mm 

needles

14. Tweezers, PTFE-coated

15. Auto-sampler vials, 2 mL capacity

16. Analytical balance, with capability of weighing 

to nearest 0.01 mg 

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling acids. All work 

should be performed with adequate ventilation for personnel and equipment. It is imperative that acid 

be added to water in order to avoid a violent exothermic reaction. There is risk of fire and explosion 

when phosphoric acid has contact with bases, combustible substances, oxidizing agents, reducing 

agents or water. Avoid physical contact since both sulfuric and phosphoric acid are both corrosive and 

irritants to eyes, skin, and the respiratory system. These are caustic materials and can react with metals 

to form flammable hydrogen gas. Do NOT mix with solutions containing bleach or ammonia [9,10].

SAMPLING, SAMPLE TRANSPORT AND STORAGE:

1. Calibrate each personal sampling pump with a representative sampler in-line.

2. Sample at an accurately known flow rate between 1 and 5 L/min for a total sample size of 15 to 1000 

Liters. Avoid sampler overloading.

3. Immediately after sampling, remove the filter from the cassette with PTFE-coated 

tweezers and place it in a 10-mL screw-cap plastic vessel. With about 2 mL extraction 

solution (0.0027 M Na2

CO3 / 0.0003 M NaHCO3), rinse material from the inside surfaces of the 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NON-VOLATILE ACIDS (Sulfuric Acid and Phosphoric Acid): METHOD 7908, Issue 1, dated 20 May 2014 - Page 3 of 5

cassette into the vessel. Add additional extraction solution into the vessel until a final volume of 5 

mL is reached.

4. Submit at least three field blanks for each set of samples collected per day. Handle these in the same 

way as the field samples; i.e., place each filter into a vessel, add 5 mL of eluent solution and ship it to 

the lab along with the remaining samples.

5. Refrigerate all samples that are to be stored overnight (or longer) prior to shipment to the 

laboratory. Ship all samples to the laboratory in accordance with established chain-of-custody 

procedures [11]. 

6. Refrigerate the samples (4 °C) immediately upon receipt at the lab until ready for analysis. 

7. Analyze samples within 4 weeks of receipt. 

SAMPLE PREPARATION: 

8. Remove sample vessels from storage and bring them to room temperature.

9. Sonicate the samples in an ultrasonic bath for at least 15 minutes and allow to cool for at least 30 

minutes.

10. Using 5-mL syringes, filter each sample extract solution through a PTFE filter into clean plastic 

vessels or into autosampler vials (if autosampler used).

CALIBRATION AND QUALITY CONTROL: 

11. Through dilution of the calibration stock solution in eluent solution, prepare calibration working 

standard solutions covering the range of approximately 0.2 to 8 mg/L of sulfate and phosphate. 

Store working standards in tightly sealed polyethylene bottles. Prepare fresh working standards 

weekly.

12. Calibrate the ion chromatograph with at least six working standards covering the range of 0.2 to 8 

mg/L of sulfate and phosphate ion per sample by preparing a calibration graph of anion peak height 

(mm or μS [micro siemens]) vs. concentration (mg/L).

13. Analyze working standards together with samples, reagent blanks and field blanks at a frequency of 

at least 1 per 20 samples (3 minimum of each).

MEASUREMENT: 

14. Set the ion chromatograph to desired eluent flow rate, e.g.,1.5 mL/min, and column pressure, e.g., 

1.1 x 105

 kPa, and other conditions as specified by the instrument manufacturer.

15. Inject a sample aliquot, e.g., 50-μL, into the chromatograph, and measure the peak heights of the 

phosphate and sulfate peaks (at retention times of about 9 min and 11.6 min, respectively). If 

the peak height exceeds the linear calibration range, dilute with eluent, reanalyze and apply the 

appropriate dilution factor in calculations.

CALCULATIONS: 

16. Calculate the mass concentration of each anion, C (mg/m3

), in the air volume sampled, V (L): 

 

where: 

C0

= mean concentration, in mg/L, of anion in the field blank test solutions;

C1

= concentration, in mg/L, of anion in the sample test solution;

V = volume, in liters, of the air sample;

V0

= volume, in mL, of the field blank test solutions; 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NON-VOLATILE ACIDS (Sulfuric Acid and Phosphoric Acid): METHOD 7908, Issue 1, dated 20 May 2014 - Page 4 of 5

V1

= volume, in mL, of the sample test solutions

Fd = dilution factor for each sample test solution

Fc

 = conversion factor to convert from anion to acid concentration: 

Fc

 = 1.021 for sulfate; Fc = 1.031 for phosphate

EVALUATION OF METHOD: 

Laboratory testing with generated atmospheres of sulfuric acid mist yielded a collection efficiency of 

greater than 95% over the range 0.5 to 10 mg/m3

 of H2SO4 on 0.45 μm pore size PTFE filters [4]. Greater 

than 95% recovery of sulfuric acid and phosphoric acid was found four weeks after sample collection. 

On quartz fiber filters, 97 to 100% recovery of sulfuric acid was found four weeks after sample collection, 

and no breakthrough was observed at sample loadings of up to 1 mg [3]. Mean analytical recovery 

determined from the analysis of spiked quartz fiber filters has been found to be in the range of 97 to 

100% for both acids [3,4]. The component of the coefficient of variation of the method that arises from 

analytical variability, determined from the analysis of spiked quartz fiber filters, was 0.7% to 3.2% for 

phosphoric acid and 0.5% to 2.6% for sulfuric acid [3]. An interlaboratory study with 26 participants 

found negligible biases and interlaboratory relative standard deviations of 12 to 15% for sulfuric acid 

and phosphoric acid concentrations between 0.05 and 1 mg/m3

 [12]. The method has also been field 

tested for sulfuric acid measurements at sample volumes of up to nearly 2,000 Liters [13]. The analytical 

figures of merit for the method satisfy performance criteria specified in an applicable consensus 

standard [14]. The back-up data and user check reports are references 4 and 12 respectively.

REFERENCES: 

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio:

American Conference of Governmental Industrial Hygienists. [www.acgih.org]. [Data accessed April

2014.]

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database

on international limit values for hazardous substances (German Social Accident Insurance). Sankt

Augustin, FRG: [https://www.dguv.de/ifa/index-2.jsp]. [Data accessed April 2014.]

[3] Breuer D [2002]. Inorganic acid mists (H2SO4, H3PO4). In: Kettrup A, Greim H, eds. Analyses of

hazardous substances in air (Deutsche Forschungsgemeinschaft). Vol. 6. Weinheim, FRG: Wiley-VCH.

pp. 67-78.

[4] Breuer D [1999]. Measurement of vapor-aerosol mixtures. J Environ Monit. 1:299-305.

[5] ISO (International Organization for Standardization) [1995]. ISO Guide 98: Guide to the expression

of uncertainty in measurement. Geneva, Switzerland: ISO.

[6] ISO [2007]. ISO 21438-1: Workplace atmospheres – Determination of inorganic ions by ion

chromatography – Part 1: Non-volatile acids (sulfuric acid and phosphoric acid). Geneva,

Switzerland: International Organization for Standardization (ISO).

[7] Breuer D [2013]. Anorganische säuren, partikulär: phosphorsäure, schwefelsäure (Inorganic acid

mists: phosphoric acid, sulfuric acid), in IFA Working Folder, Sheet No. 6173. Berlin, FRG: ErichSchmidt Verlag. ISBN 978-3-503-13084-9.

[8] NIOSH [1994]. Acids, inorganic: Method 7903. In: Eller PM, Cassinelli ME, eds. NIOSH Manual of

analytical methods, 4th ed., Cincinnati, OH: U.S. Department of Health and Human Services, Center

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS

(NIOSH). Publication No. 94-113. [www.cdc.gov/niosh/nmam.]

[9] NIOSH [2005]. NIOSH Pocket guide to chemical hazards. Barsan ME, ed. Cincinnati, OH: US

Department of Health and Human Services, Centers for Disease Control and Prevention, National

Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 2005-149. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

NON-VOLATILE ACIDS (Sulfuric Acid and Phosphoric Acid): METHOD 7908, Issue 1, dated 20 May 2014 - Page 5 of 5

[10] Furr AK, ed. [1995] CRC Handbook of laboratory safety, 4th ed. Boca Raton, FL: CRC Press.

[11] ASTM International [2010]. ASTM D4840: Standard guide for sampling chain of custody procedures. 

West Conshohocken, PA: ASTM International.

[12] Breuer D, Howe A [2006]. Performance of methods for measurement of exposure to inorganic acids 

in workplace air. J Environ Monit. 8:120-126.

[13] Breuer D, Heckmann P, Gusbeth K, Schwab G, Blaskowitz M, Moritz A [2012]. Sulfuric acid at 

workplaces – applicability of the new indicative occupational exposure limit value (IOELV) to 

thoracic particles. J Environ Monit. 14:440-445.

[14] Comité Européen de Normalisation (CEN) [2012]. EN 482, Workplace exposure — General 

requirements for the performance of procedures for the measurement of chemical agents. Brussels, 

Belgium: Comité Européen de Normalisation (CEN).

METHOD WRITTEN BY: 

Dietmar Breuer, Dr. rer. nat., Institute for Occupational Safety and Health of the German Social Accident 

Insurances, Sankt Augustin, Germany; and Kevin Ashley, Ph.D., NIOSH/DART

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for 

the content of these websites. All web addresses referenced in this document were accessible as of 

the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TOLUENE in Blood 8007

 C7H8 MW: 92.14 CAS: 108-88-3 RTECS:XS5250000 

METHOD: 8007, Issue 1 EVALUATION: FULL Issue 1: 5 March 2013

NIOSH and OSHA: NA

Because data on exposure limits and guidelines may change 

over time, NIOSH recommends referring to the following 

sources for updated limits and guidelines [1,2].

 PROPERTIES: Clear liquid; bp 110.6 °C; VP 28.4 mm Hg at 

25 °C; d20 0.8669 g/mL.

 BIOLOGICAL INDICATOR OF: Exposure to toluene

SYNONYMS: Methylbenzene, Phenylmethane, Toluol, Methylbenzol

SAMPLING 

SPECIMEN: Blood (collected within 12 hours of last 

exposure)

VOLUME: Fill a 10-mL vacuum specimen tube as 

completely as possible (to reduce the amount of 

airspace)

PRESERVATIVE: Sodium citrate (light blue top)

SHIPMENT: Insulated container with bagged refrigerant

SAMPLE

STABILITY: Stable at least 30 days at 4 °C [3]

CONTROLS: Collect and refrigerate immediately 

ACCURACY

RANGE STUDIED: 0.0208-5.76 μg/mL [3]

ACCURACY: <5% [3]

BIAS: 0.0006 

OVERALL

PRECISION (ŜrT): 0.0157 [3]

SAMPLE STABILITY: 99.0% (Day 30) [3]

MEASUREMENT 

TECHNIQUE: HEADSPACE/GAS CHROMATOGRAPHY, FID 

ANALYTE: Toluene

HEADSPACE 

CONDITIONS: Transfer Temp,129 °C; Withdrawal, 0.80 min; 

Thermostat Time, 30 min; Needle, 120 °C; 

GC Cycle Time, 34 min; Sample Temp: 99 °C; 

Pressurize, 4.0 min; Inject, 0.40 min

INJECTION 

CONDITIONS: Direct injection onto column for 0.4 min,

 then split flow for the remainder of the 

run at 130 mL/min

TEMPERATURE-INJECTION: 129 °C

-DETECTOR: 250 °C

-COLUMN: 50 °C (hold for 2 min); 50 to 220 

°C @ 10 °C/min; 220 °C 

(hold for 5 min)

CARRIER GAS: Helium, at 21 psi head pressure

COLUMN: Capillary, fused silica, 75-m x 0.53 mm, 

coated internally with 3.0 μm film 6% 

cyano- propylphenyl, 94% dimethylpolysiloxane, bonded and cross-linked

CALIBRATION: Analyte in control blood; with isobutanol 

 internal standard (IS)

Estimated LOD: 0.006 μg/mL in whole blood [3]

APPLICABILITY: Can be used in monitoring the exposure of workers and drug abusers for toluene.

INTERFERENCES: None identified. The chromatographic separation conditions may be adjusted to correct separation 

problems.

OTHER METHODS: NIOSH Method 8002 is a partially-evaluated, packed-column GC/FID method with lower accuracy and a 

higher detection limit. [4] The National Center for Environmental Health (NCEH)/CDC has a method for 31 volatile organic 

compounds in blood that includes toluene. This NCEH method is somewhat more complicated, comparable in accuracy, and 

~1,000 times more sensitive. [5]
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TOLUENE in Blood: METHOD 8007, Issue 1, dated 5 March 2013 - Page 2 of 5

REAGENTS:

1. Whole blood*, citrate treated

2. Toluene* (reagent grade, >99.5%)

3. Isobutanol internal standard (IS) as a 20 mg/L 

solution in water*

4. Water, ASTM Type II

5. Helium, UHP or higher

6. Hydrogen, UHP or higher

7. Air, UHP or higher

8. Toluene calibration standards*

9. Ethylene glycol*

* See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Gas chromatograph, with FID, column, and data 

collector (page 8007-1)

2. Headspace sampler, holds up to 40 vials and 

can thermostat up to 12 vials simultaneously for 

automated headspace analysis (Any instrument 

that can achieve the headspace settings specified 

on p. 8007-1 is acceptable)

3. Bagged refrigerant, and refrigerator

4. Adjustable pipette, 10-mL, tips 10-mL, plastic or 

serological

5. Headspace vials, 20-mL, with caps and septa

6. Micro-liter syringes, 10-μL , 25-μL, 100-μL, 1-mL

7. Volumetric flasks, 10-mL, 25-mL, 1-L

8. Syringes, twist-on fitting (luer lock style or 

equivalent), 5-mL, and 20 gauge needles

9. Tubes, vacuum blood collection, 10-mL, citrate 

coated (light blue top)

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals and blood 

products. Disposable plastic, glass, and paper (pipet tips, gloves, etc.) that contact blood should be placed 

in a biohazard container. Contact with biological samples can have serious health consequences through 

exposure to hepatitis, HIV, and other diseases. All personnel collecting, handling, or analyzing samples 

should follow universal precautions [6] and comply with the OSHA bloodborne pathogens standard 

[7] which includes immunization for hepatitis B. Toluene is extremely flammable. All work should be 

performed in a fume hood.

SAMPLING:

1. Collect the blood by filling a 10-mL vacuum blood collection tube (light blue top, citrate) as completely 

as possible in order to keep the air space to a minimum. Be certain to invert the blood tube several (5-10) 

times to mix the anticoagulant.

NOTE: If desired, duplicate blood tubes could be drawn to allow for estimation of measurement 

precision, especially if collecting from numerous participants.

2. Immediately transfer the sample to a refrigerator or cooler (≤ 4 °C.)

3. Ship the sample vial in a well-insulated container equipped with blue ice or other cooling material.

SAMPLE PREPARATION:

4. Allow the blood sample to reach room temperature, then mix gently so as to not produce a froth by 

inverting the tubes 4 to 5 times.

5. Add 1.5 mL of the IS solution to a 20-mL headspace vial.

6. Extract 1.5 mL of sample from the vacuum vial using a 5-mL syringe with a twist-on fitting (luer lock style 

or equivalent) or micropipette and transfer the blood to a headspace vial.

7. Cap both sample and headspace vials immediately. 

8. Mix the contents of each headspace vial thoroughly.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TOLUENE in Blood: METHOD 8007, Issue 1, dated 5 March 2013 - Page 3 of 5

STOCK STANDARD PREPARATION:

9. Prepare a primary stock standard solution by diluting 20 μL of toluene to 50 mL with ethylene glycol (20 

μL/50 mL = 345.7 μg/mL.) Store in a tightly capped glass container having little or no head space.

NOTE: Aliquots of toluene should be introduced below the ethylene glycol to prevent loss of toluene. 

10. Prepare stock standards by diluting aliquots of the primary standards with ethylene glycol. Suggested 

levels: 34.6 μg/mL, 3.46 μg/mL, 0.346 μg/mL.

CALIBRATION AND QUALITY CONTROL:

11. Calibrate with at least five working standards in duplicate covering the concentration range of the 

samples.

a. Prepare each working standard by adding 1.5 mL of IS solution and 1.5 mL of blank blood to a 

headspace vial and cap the vial. Spike the mixture in the vial (through the septum using a microsyringe) with a stock standard solution to the desired concentration of toluene and mix by shaking.

b. Prepare at least two blanks by repeating step a. but omitting the toluene spike.

c. Measure the peak areas of toluene and isobutanol in the chromatograms. Subtract the average 

toluene peak area of the blank from the toluene peak areas of the standards (see NOTE 1.) Divide 

the peak area of the blank-corrected toluene by the peak area from the isobutanol peak. Prepare a 

calibration curve of the peak area Std/area IS versus the toluene concentration of the standards.

NOTE 1: A trace amount of toluene may be present in the blood of some donors. These levels 

 will vary depending upon environmental exposures. If the blanks show the presence 

 of toluene the standards need to be blank corrected, or else the sample results 

 will be biased low. Geometric means for the U.S. population as determined from the 

 National Health and Nutrition Examination Survey may be useful for comparison. [8]

NOTE 2: It is also highly recommended that a reagent blank or blanks be included in the analysis. 

 Atmospheric toluene in the lab may contribute errors to the measured values. A 

 reagent blank using water, a blank headspace vial, or both of these options will 

 show if the laboratory conditions are free from a quantifiable amount of toluene.

12. Prepare two levels of quality control (QC) samples by spiking toluene into whole blood. These levels 

could be at ~10 X LOQ and 200 X LOQ, but could be adjusted to better suit the anticipated levels of 

the sample set. Unspiked samples of the blood used to prepare the QC samples should be analyzed to 

determine the blank level and the true target level. QC samples should be analyzed with every batch 

such that they constitute 10% of the sample batch.

13. QC values should be within +/- 20% of the spiked values. If not, the batch is considered out of control, 

the batch data discarded, and corrective actions should be taken before more samples are analyzed. 

Alternatively, if the method has a long history in the lab allowing enough data for control charts to be 

constructed, the control charts could serve as the in or out of control decision guide, which could be 

looser or stricter than the recommended 20%.

MEASUREMENT:

14. Set the gas chromatograph according to manufacturer’s recommendations and to conditions given on 

page 8007-1.

15. Set the headspace sampler according to manufacturer’s recommendations and to conditions given on 

page 8007-1.

16. Inject and analyze samples, standards, QC samples, and blanks.

17. Measure the peak area of both toluene and isobutanol in the chromatograms (do not subtract the blank 

from the samples). Divide the peak area of the toluene peak by the area from the isobutanol peak.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TOLUENE in Blood: METHOD 8007, Issue 1, dated 5 March 2013 - Page 4 of 5

CALCULATIONS:

18. Determine the concentration (μg/mL) of the toluene in each sample of blood using the calibration curve

obtained in step 11c.

EVALUATION OF METHOD:

This method was evaluated over the range specified on page 8007-1. These ranges, 0.0208 to

5.76 μg/mL, represent from 1 x LOQ to 300 x LOQ. Six replicates were analyzed at each level. The average 

recoveries at the various levels ranged from 97% to 106% for toluene. The LOD and LOQ were determined 

by preparing a series of duplicate standards. Each series was made up and analyzed on a different day. The 

resulting data was then fitted to a quadratic curve. The LOD and LOQ were estimated according to Burkart’s 

Method. [9] A long-term storage study was carried out at the 10 x LOQ level. Citrated whole blood samples 

that were spiked with toluene were stored at 24 °C and 4 °C for 1, 4, and 7 days and for 7,10, 21, and 30 

days (respectively,) and then analyzed. Average recoveries were 96% at room temperature and 99% at 4 °C. 

Room temperature storage is not advised because if the sample clots before it is analyzed, the results will be 

compromised.

LIMITATION OF METHOD:

Concerning the use of isobutanol as an internal standard: While the boiling points of toluene and 

isobutanol are similar, their Henry’s Law constants and thus their partitioning coefficients are quite different. 

This could lead to biases if there are differences in polarity in the samples or between the samples and the 

standards. This is one reason the calibration standards in this method are prepared using blood and not 

just solvent. An alternative approach could entail the use of an internal standard that partitions similarly 

to toluene, such as fully-deuterated toluene (toluene-d8). This would cause an increase in the cost of the 

method per sample (and would need to be validated by the end user), but is mentioned here as an option.

REFERENCES:

[1] ACGIH [2014]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American Conference

of Governmental Industrial Hygienists. [www.acgih.org]. Date accessed: May 2014.

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area

[2014]. List of MAK and BAT values 2014; Maximum concentrations and biological tolerance

values at the workplace. DFG: Deutsche Forschungsgemeinschaft, [http://onlinelibrary.wiley.com/

book/10.1002/9783527682027]. Date accessed: January 2015.

[3] Stone LC, Perkins JB [2003]. Backup data report for toluene in blood. DataChem Laboratories, Inc. under

NIOSH contract CDC-200-2001-08000 (unpublished.)

[4] NIOSH [1994]. Methyl ethyl ketone, ethanol and toluene in blood: Method 8002. In Eller PM, Cassinelli

ME, eds. NIOSH Manual of Analytical Methods, 4th ed. Cincinnati, OH: U.S. Department of Health and

Human Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety

and Health, DHHS(NIOSH) Publication no. 94-113.

[5] Blount BC, Kobelski RJ, McElprang DO, Ashley DL, Morrow JC, Chambers DM, and Cardinali FL [2006].

Quantification of 31 volatile organic compounds in whole blood using solid-phase microextractions

and gas chromatography-mass spectrometry. J Chrom B 832(2): 292-301.

[6] NIOSH [2011]. Bloodborne infectious diseases: HIV/AIDS, Hepatitis B, Hepatitis C; universal precautions

for preventing transmission of bloodborne infections [2011]. [www.cdc.gov/niosh/topics/bbp/

universal.html]. Accessed September 2012.

[7] OSHA [2012]. Bloodborne pathogens and needlestick prevention. [www.osha.gov/SLTC/

bloodbornepathogens/index.html]. Accessed September 2012.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TOLUENE in Blood: METHOD 8007, Issue 1, dated 5 March 2013 - Page 5 of 5

[8] CDC [2012]. Fourth national report on human exposure to environmental chemicals, updated tables, 

September 2012. [www.cdc.gov/exposurereport/pdf/FourthReport_UpdatedTables_Sep2012.pdf]. 

Accessed October 2012.

[9] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene 

chemistry laboratory. Appl. Ind. Hyg. 1(3):153-155.

METHOD WRITTEN BY:

L. C. Stone and J. B. Perkins, DataChem Laboratories, Inc., Salt Lake City, Utah under NIOSH Contract CDC-200-

2001-08000. Final editing by D. A. Shoemaker, NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

ELEMENTS in Tissues 8200

AW: Table 1 CAS: Table 2 RTECS: Table 2

METHOD: 8200, Issue 1 EVALUATION: FULL Issue 1: 30 August 2018 

OSHA: None

NIOSH: None

Other OELs: [1-3]

PROPERTIES: Table 1 

ELEMENTS: calcium, copper, iron, potassium, magnesium, manganese, molybdenum, nickel, phosphorus, zinc

SAMPLING

SPECIMEN: TISSUE 

SPECIMEN SIZE: dependent on tissue; 25-40 mg for nails

PRESERVATIVE: none, if nails; freeze dried, if internal organ

SHIPMENT: routine, nails; see IATA regulations, freezedried tissue

SAMPLE

STABILITY: not established

BLANKS: collect at least 3 specimens from 

unexposed workers per set

ACCURACY

RANGE 

STUDIED: Tables 2 & 4

BIAS: Table 4

OVERALL

PRECISION (SS�rrrr)*: Table 4 

ACCURACY*: Table 4 

*The definitions of precision and accuracy in this method are 

those utilized by the U.S. Food and Drug Administration [4].

MEASUREMENT

TECHNIQUE: INDUCTIVELY-COUPLED PLASMA ATOMIC 

EMISSION SPECTROSCOPY (ICP-AES)

ANALYTE: Table 1

DIGESTION ACID: 4/1 (v/v) HNO3/HClO4

FINAL 

SOLUTION: 4%/1% (v/v) HNO3/HClO4; 10 mL

WAVELENGTH: varies with element; Table 2

CALIBRATION: elements in 4%/1% (v/v) HNO3/HClO4

RANGE: varies with element

ESTIMATED LOD: Table 2

PRECISION (SS�rr): Table 2

APPLICABILITY: This method was developed to analyze nail samples from animals exposed to welding fume in laboratorycontrolled inhalation studies. It is useful for monitoring nail tissues of workers exposed to several metals simultaneously. It may 

also be used for the analysis of freeze-dried tissue samples (liver, lungs, etc.) from laboratory studies. This is a simultaneous 

multielemental analysis, but is not compound-specific. Verify that the types of compounds in the samples are soluble with the 

dissolution procedure selected. Some compounds of these elements require special sample treatment. Other elements are 

amenable to this sample preparation and analysis, but due to availability of certified reference materials, only those elements listed 

have been validated.

INTERFERENCES: Spectral interferences are the primary interferences encountered in ICP-AES analysis. These are minimized by 

judicious wavelength selection, interelement correction factors and background correction [5,6]. 

OTHER METHODS: This method uses a measurement technique similar to that of Methods 7300 [7] (Elements; for air samples) 

and 8310 [8] (Metals in urine). The sample preparation is comparable to that in NIOSH 7300 [7].
ELEMENTS in Tissues: METHOD 8200, Issue 1, dated 30 August 2018 - Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Nitric acid, conc. (HNO3) (Optima grade).* 

2. Perchloric acid, conc. (HClO4) (Optima 

grade).* 

3. Digestion acid, 4:1 (v/v) HNO3:HClO4.* Mix 4 

volumes conc. HNO3 with one volume HClO4. 

4. Element standards, 100 μg/mL. 

Commercially available or prepared per 

instrument manufacturer’s 

recommendations. 

5. Reference materials of known elemental 

composition. 

6. Argon, as specified by ICP-AES manufacturer. 

7. Deionized water, ASTM Type II [9] or 

equivalent. 

8. Yttrium internal standard.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Bottles, glass or polyethylene with 

polytetrafluoroethylene (PTFE)-lined caps, 20-

mL.** 

2. Regulator, two-stage, for argon. 

3. Digestion vessel: Beakers, Griffin (50-mL) or 

Phillips (125-mL), PTFE or borosilicate glass** 

or digestion tubes (50-mL), polypropylene.

4. Watch glass covers.** 

5. Hotplate or hot block, for use at 120 to 150 °C. 

6. Volumetric flasks, 10-mL, 25-mL.** 

7. Assorted volumetric pipettes, as needed. 

8. Gloves, plastic, metal-free. 

9. Scissors, surgical, stainless steel. 

**All glassware should be detergent-washed, 

thoroughly rinsed with deionized water, soaked 

12 h in conc. HNO3 and thoroughly rinsed with 

deionized water.

 

SPECIAL PRECAUTIONS: All perchloric acid digestions must be carried out in a perchloric acid fume 

hood. When working with concentrated acids, wear protective clothing, safety goggles and gloves. All 

work should be performed with adequate ventilation for personnel and equipment. It is imperative that 

acid be added to water in order to avoid a violent exothermic reaction.

Samples of tissue collected from humans and animals pose a real health risk to laboratory workers who 

collect and handle these samples. These risks are primarily due to personal contact with infective 

biological samples and can have serious health consequences, such as infectious hepatitis, and other 

diseases. There is also some risk from the chemical content of these samples, but this risk is much less. 

Those who handle blood and tissue specimens should follow CDC Universal Precautions [10], wear 

protective gloves, and avoid aerosolization of the samples. Mouth pipetting, of course, must be avoided.

SAMPLING:

1. Collect tissue samples in pre-weighed screw top bottles or tubes. 

NOTE 1: For nail samples, the tips of the nails should be gently trimmed using stainless steel surgical 

scissors. Clippings from each subject should be pooled.

NOTE 2: If the intent of the analysis is to determine internal elemental content, surface contamination 

should be avoided and nail samples may need to be cleaned of exogenously bound metals 

before collection and/or sample preparation. 

NOTE 3: If shipping samples, follow current International Air Transport Association (IATA) regulations 

and use a certified IATA Dangerous Goods Shipper if necessary.

2. Weigh tissue samples. 
ELEMENTS in Tissues: METHOD 8200, Issue 1, dated 30 August 2018 - Page 3 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION:

3. Transfer tissue sample to digestion vessel. Rinse the sample container with a minimal amount of 

deionized water three times and transfer rinses to digestion vessel containing sample. 

4. Add 5.0 mL digestion acid to each tissue sample. Cover beakers with watch glasses. Heat at 120-130 °C 

for 2 h. 

NOTE: Start reagent blanks and QC samples at this step. 

5. Increase hotplate temperature to 150 °C and heat until sample digestion is complete (no visible solid).

NOTE: A typical time frame for rodent nail samples is less than 1 week. If needed, additional acid may 

be added due to acid volume loss with heating.

6. Remove watch glass covers and reduce acid volume to near-dryness (ca. 0.5 mL).

7. Add 0.5 mL digestion acid.

8. After allowing to cool to room temperature, transfer the solutions quantitatively using deionized 

water to 10-mL flasks.

9. Dilute to volume using deionized water.

NOTE 1: The final sample volume may be held to different volumes based upon the analyst’s needs. 

Use of final volumes other than 10 mL will require an adjustment to the amount of digestion 

acid added in Step 7. The final sample should contain a minimum of 4%/1% HNO3/HClO4. 

NOTE 2: If an internal standard will be used for the instrumental analysis, it should either be added 

before sample dilution to final volume or introduced during sample uptake into the instrument. 

CALIBRATION AND QUALITY CONTROL:

10. Calibrate the spectrometer according to manufacturer's recommendations.

NOTE: Typically, an acid blank, 0.1 μg/mL, and 1.0 μg/mL multielement calibration standards are used. 

It is important that the calibration standards match the acid matrix and acid concentration of 

the samples. If an internal standard will be used for instrumental analysis, it must be added to 

the standards at the same concentration as in the samples. The following multielement 

combinations are chemically compatible in 4%/1% HNO3/HClO4: 

a. Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, La, In, Na

b. Ag, K, Li, Mg, Mn, Ni, P, Pb, Se, Sr, Tl, V, Y, Zn, Sc

c. Mo, Sb, Sn, Te, Ti, W, Zr

11. Analyze at least one calibration standard per ten samples to check that the instrument calibration is 

still valid. If control tissue samples are available, check recoveries with at least one media blank and 

two spiked media blanks per twenty samples. Media should be spiked with analytes of interest. If no 

control tissue samples are available, check recoveries using a reagent blank and certified reference 

materials or liquid spikes.

NOTE: Whenever possible, QA/QC samples should be prepared from certified reference materials in a 

matrix similar to the bulk material sampled. Liquid spiked samples are only surrogates for real 

world samples and QC data based upon certified samples are preferred.

MEASUREMENT:

12. Set the spectrometer to conditions specified by the manufacturer. 

13. Analyze calibration standards and samples by ICP-AES in accordance with manufacturer 

recommendations. 

NOTE: If the values for the samples are above the range of the standards, dilute the solutions (ensuring 

the samples remain acid matrix-matched to the calibration standards), reanalyze and apply the 

appropriate dilution factor in the calculations.
ELEMENTS in Tissues: METHOD 8200, Issue 1, dated 30 August 2018 - Page 4 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

CALCULATIONS:

14. Obtain the solution concentration found for each element in the sample, Cs (μg/mL), and the average 

blank, Cb (μg/mL), from the measurement data. 

15. According to the method of standardization, calculate the concentration, C (μg/g), of each element in 

the mass of sample taken, M (g). Use the final solution volumes of sample, Vs (mL) and blank, Vb (mL): 

C =

CSVS − CbVb

M , μg/g

EVALUATION OF METHOD:

Acceptable and unacceptable levels for elements within tissue samples have not been determined by this 

method. Iyengar [11] reports metals concentrations in tissues and body fluids. Lauwerys [12] discusses 

metals and can be consulted for guidance and interpretation. For tissue, trace element concentrations will 

vary with the tissue type or organ. Iyengar [11] reports metals concentrations for different tissues of nonoccupationally exposed people. 

This method was validated using control rat nails spiked with elements of interest and NIST SRM 1577C 

Bovine Liver. The results for the validated elements may be found in Tables 2 & 3, respectively. Mo was not 

included in the spiked control rodent nails, but was present in the SRM. The variability in the control rat 

nails, along with the concentration of elements in the SRM, limited the number of elements that could be 

quantitatively recovered. For example, the variability in the control nails was greater than the spiked 

amounts for Ca and P (Table 2) and the amounts present in the SRM for Ag, As, Ca, Cd, Cr, Ni, P, Sb, Sr, & V. 

Additionally, the concentrations of Co & Li detected in the SRM were less than the method LOQ. 

Acceptable amounts of Ca, Cu, Fe, K, Mg, Mn, Mo, P, and Zn were recovered at each level using the bovine 

liver SRM (Table 3).

While acceptable amounts were recovered using SRM 1577C for Ca, the large variability (with respect to 

the amount present in SRM 1577C) in the control nail samples for Ca calls into question the applicability of 

SRM 1577C for the validation of rat nail samples for Ca.

REFERENCES:

[1] ACGIH [2018]. TLVs® and BEIs® based on the documentation of the Threshold Limit Values for 

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American 

Conference of Governmental Industrial Hygienists. 

[2] Permanent Senate Commission for the Investigation of Health Hazards of Chemical Compounds in 

the Work Area [2016]. List of MAK and BAT values 2016; Maximum Concentrations and Biological 

Tolerance Values at the Workplace. Weinheim, Germany: Wiley-VCH Verlag GmbH & Co., 

https://dx.doi.org/10.1002/9783527805983. 

[3] Institute for Occupational Safety and Health of the German Social Accident Insurance (IFA) [2017]. 

GESTIS international limit values, http://limitvalue.ifa.dguv.de/. 

[4] US FDA [2001]. Guidance for industry: Bioanalytical method validation. Washington, DC: U.S. Food 

and Drug Administration, http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf. 

[5] Dean JA [1995]. Atomic spectroscopy. In: Analytical chemistry handbook; Section 7. New York, NY: 

McGraw-Hill. 
ELEMENTS in Tissues: METHOD 8200, Issue 1, dated 30 August 2018 - Page 5 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[6] Montaser A, Golightly DW, eds. [1992]. Inductively coupled plasmas in analytical atomic 

spectrometry. 2nd ed. Hoboken, NJ: John Wiley & Sons, Inc. 

[7] NIOSH [2003]. Method 7300, Elements by ICP (nitric/perchloric acid ashing). In: Schlecht PC, 

O’Connor PF, eds. NIOSH manual of analytical methods. 4th ed. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2003-154.

[8] NIOSH [1994]. Method 8310, Metals in urine. In: Schlecht PC, O’Connor PF, eds. NIOSH manual of 

analytical methods. 4th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers 

for Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS 

(NIOSH) Publication No. 94-113.

[9] ASTM [2011]. ASTM D1193, Standard specification for reagent water. West Conshohocken, PA: ASTM 

International. 

[10] CDC [1988]. Perspectives in disease prevention and health promotion update: Universal precautions 

for prevention of transmission of human immunodeficiency virus, hepatitis b virus, and other 

bloodborne pathogens in health-care settings. MMWR 37(24):377-388.

[11] Iyengar GV, Kollmer WE, Bowen HJM [1978]. The elemental composition of human tissues and body 

fluids: A compilation of values for adults. New York: Verlag Chemie Weinheim.

[12] Lauwerys RR [1983]. Industrial chemical exposure: Guidelines for biological monitoring. Davis, CA: 

Biomedical Publications.

[13] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. By 

Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of Health 

and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational 

Safety and Health, DHHS (NIOSH) Publication No. 95–117.

[14] NIOSH [2016]. Measurement uncertainty and NIOSH method accuracy: Chapter UA. In: Ashley K, 

O’Connor PF, eds. NIOSH manual of analytical methods. 5th ed. Cincinnati, OH: U.S. Department of 

Health and Human Services, Centers for Disease Control and Prevention, National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 2014-151, 

https://www.cdc.gov/niosh/nmam/. 

METHOD WRITTEN BY:

Ronnee N. Andrews, Ph.D., NIOSH

Disclaimer: Mention of any company or product does not constitute endorsement by the National 

Institute for Occupational Safety and Health (NIOSH), Centers for Disease Control and Prevention (CDC). In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
ELEMENTS in Tissues: METHOD 8200, Issue 1, dated 30 August 2018 - Page 6 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 1. PROPERTIES AND GENERAL INFORMATION

Element (Symbol) Atomic Weight MP, °C CAS # RTECS

Calcium (Ca) 40.08 842 7440-70-2 ---

Copper (Cu) 63.54 1083 7440-50-8 GL5325000

Iron (Fe) 55.85 1538 7439-89-6 NO4565500

Potassium (K) 39.10 64 7440-09-7 TS6460000

Magnesium (Mg) 24.31 651 7439-95-4 OM2100000

Manganese (Mn) 54.94 1246 7439-96-5 OO9275000

Molybdenum (Mo) 95.94 2623 7439-98-7 QA4680000

Nickel (Ni) 58.71 1455 7440-02-0 QR5950000

Phosphorus (P) 30.97 44 7723-14-0 TH3500000

Zinc (Zn) 65.37 419 7440-66-6 ZG8600000

TABLE 2. MEASUREMENT WAVELENGTHS (λ), DETECTION LIMITS (LOD) AND RECOVERY DATA FROM SPIKED 

CONTROL RODENT NAILS

Element λA, nm LODB, μg/g

Spike 

concentration 

range, μg

Low level 

% Recovery 

(% RSD)C

Medium level 

% Recovery 

(% RSD)C

High level 

% Recovery 

(% RSD)C

Ca 315.887 39. 62.5-500 N/AD 38.7 (139) 50.8 (44.6) 

Cu 324.754 6.0 6.25-50 99.1 (1.9) 93.9 (2.6) 95.3 (1.3) 

Fe 259.941 12. 6.25-50 89.8 (3.7) 89.7 (11.3) 89.2 (1.8) 

K 766.491 34. 93.8-750 112. (8.3) 113. (6.1) 124. (3.0) 

Mg 279.079 16. 6.25-50 62.8 (50.3) 81.0 (12.2) 82.2 (4.9) 

Mn 257.611 1.4 1.25-10 88.2 (1.8) 83.4 (2.9) 86.0 (1.5) 

Mo 202.095 0.92 - - - - 

Ni 231.604 1.7 1.25-10 90.7 (1.9) 85.2 (2.6) 87.0 (1.6)

P 178.287 2.3 6.25-50 N/A N/A N/A

Zn 213.856 20. 18.8-150 96.9 (3.1) 95.8 (2.6) 91.0 (0.9) 

ACommonly used wavelength; choose wavelength appropriate for your instrument settings [5,6] BLOD values calculated using the responses of 7 reagent blanks and corrected for average sample weight of 0.0393 g. CMean recovery & relative standard deviation for n=6, results corrected for average reagent blank and control nail response, % RSD 

calculated from % recovery

DN/A indicates that the value could not be calculated; variability of elemental concentration in control nails greater than spiked 

amount for Ca & P 
ELEMENTS in Tissues: METHOD 8200, Issue 1, dated 30 August 2018 - Page 7 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 3. RECOVERY DATA FROM BOVINE LIVER CERTIFIED REFERENCE MATERIAL

Element True value, μg/g

Low level % 

Recovery (%RSD)A

Medium level % 

Recovery (%RSD)A

High level % 

Recovery (%RSD)A

Ca 131 102.3 (5.7) 95.9 (4.5) 90.5 (1.3)

Cu 275.2 96.4 (1.9) 94.1 (3.2) 94.7 (1.4)

Fe 197.94 85.6 (2.4) 92.8 (3.6) 98.7 (1.3)

K 10230 105.9 (3.4) 101.5 (3.4) 98.7 (1.3)

Mg 620 89.0 (2.2) 85.0 (3.2) 84.5 (1.4)

Mn 10.46 84.7 (2.1) 82.7 (3.5) 83.0 (1.2)

Mo 3.3 94.5 (2.0) 94.7 (4.0) 96.5 (1.3)

P 11750 92.8 (3.1) 89.4 (3.9) 89.1 (1.5)

Zn 181.1 94.9 (4.7) 91.6 (4.0) 91.9 (1.6)

AMean recovery & relative standard deviation for n=6

TABLE 4. ACCURACY, BIAS, AND PRECISION CALCULATED FROM BOVINE LIVER CERTIFIED REFERENCE 

MATERIAL

Element nA Range, μg/sample Bias B SrT Accuracy, % 

Ca 18 13.0 – 26.6 -0.0956 0.00032 9.6

Cu 18 27.4 – 55.9 -0.0493 0.00008 4.9

Fe 18 19.7 – 40.2 -0.0649 0.00013 6.5

K 18 1020 – 2080 0.0622 0.00000 6.2

Mg 18 61.6 – 126 -0.155 0.00003 15.5

Mn 18 1.04 – 2.13 -0.165 0.00193 16.9

Mo 18 0.328 – 0.671 -0.0474 0.00774 6.0

P 18 1170 – 2380 -0.0950 0.00000 9.5

Zn 18 18.0 – 36.8 -0.0712 0.00019 7.1

An = total number of results reported for each element, ND results not included BSrT = precision [13, 14] 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

FLUORIDE in URINE 8308

F MW: 19.00 CAS: 16984-48-8 RTECS: LM6290000

METHOD: 8308, Issue 3 EVALUATION: FULL Issue 1: 15 February 1984 

Issue 3: 14 March 2016

BIOLOGICAL INDICATOR OF: exposure to inorganic fluorides [1,2]

SYNONYMS: None

SAMPLING

SPECIMEN: urine, pre- and post-shift

VOLUME: 50 mL in chemically clean polyethylene

bottles

PRESERVATIVE: 0.2 g EDTA added to bottles before

collection

SHIPMENT: in insulated containers using bagged

refrigerant

SAMPLE

STABILITY: 2 weeks @ 4  ̊C, longer if frozen

CONTROLS: collect 3 sets of specimens from

unexposed workers (pre- and post-shift)

ACCURACY

RANGE 

STUDIED: not studied

BIAS: not determined

OVERALL

PRECISION (SS�rrrr): not determined

ACCURACY: ± 23.6%

MEASUREMENT

TECHNIQUE: ION SELECTIVE ELECTRODE (ISE)

ANALYTE: fluoride ion (F-

)

DILUTION: mix equal volumes of urine with TISAB

CALIBRATION: solutions of sodium fluoride in water

QUALITY

CONTROL: spiked urine pools; correct for creatinine 

content

RANGE: 1 to 100 mg/L urine

ESTIMATED LOD: 0.1 mg/L urine

RECOVERY: 0.95 [3]

PRECISION (SS�rr): 0.04

APPLICABILITY: Any fluorine-containing substances that can be metabolized to fluoride (F-) can be monitored using this 

procedure. Inorganic compounds of fluoride can be absorbed by the body resulting in the excretion of fluoride ions as sodium 

fluoride. Dietary and domestic water sources of fluoride must be considered, as well as dental treatments.

INTERFERENCES: Hydroxide, the only positive interference, is eliminated by use of the buffer. Negative interferences from 

complexation of fluoride by cations, such as calcium, are minimized by EDTA preservative and the high ionic strength buffer.

OTHER METHODS: This method is P&CAM 114 [4] in a revised format. Other methods that have been used are those described in 

the NIOSH criteria documents on inorganic fluorides [1] and hydrogen fluoride [2].
FLUORIDE in URINE: METHOD 8308, Issue 3, dated 14 March 2016 - Page 2 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Distilled or deionized water.

2. Sodium citrate tribasic dihydrate 

(Na3C6H3O7·2H2O), ACS reagent grade or 

better.

3. Ethylenediaminetetraacetic acid (EDTA) 

disodium salt, ACS reagent grade or better.

4. Acetic acid, glacial, ACS reagent grade or 

better.

5. Sodium chloride, ACS reagent grade or 

better.

6. Sodium hydroxide, 5 M. Dissolve 20 g NaOH 

in distilled water; dilute to 100 mL.

7. Sodium fluoride, ACS reagent grade or 

better.

8. Calibration stock solution, 100 μg F- /mL. 

Dissolve 0.2211 g dry sodium fluoride in 

distilled water. Make 1000 mL solution.

9. Total ionic strength activity buffer (TISAB), 

pH 5. Add 57 mL glacial acetic acid, 58 g 

sodium chloride, and 0.30 g sodium citrate 

to a 1-L beaker containing 500 mL distilled 

water. Stir to dissolve. Place beaker in water 

bath for cooling. Slowly add 5 M sodium 

hydroxide until the pH is between 5.0 and 

5.5. Cool to room temperature; dilute to 1 L 

with distilled water. 

10. Fluoride in Urine Certified Reference 

Materials (CRMs). 

EQUIPMENT:

1. Polyethylene bottles, 125-mL, widemouth.

2. Fluoride ion specific electrode (ISE), with 

reference electrode.

3. pH/millivolt meter, reading to ± 0.5 mV.

4. Stirrer, magnetic.

5. Stirring bars, PTFE-coated.

6. Beakers, plastic, 50-mL.

7. pH electrode.

8. Pipets, appropriate sizes for standards.

9. Volumetric flasks for standards.

10. Water bath.

11. Tissues, low-lint lab wipers. 

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals and 

human urine products. Disposable plastic, glass, and paper (pipet tips, gloves, etc.) that contact urine 

should be placed in a biohazard container. Standard precautions should always be used when handling 

bodily fluids and/or extracts of bodily fluids [5]. Handle urine samples and urine extracts using proper 

gloves. Glacial acetic acid is flammable and corrosive and should be handled in a fume hood.

SAMPLING:

1. Collect pre- and post-shift spot urine samples in polyethylene bottles containing 0.2 g EDTA. 

2. Ship samples in insulated container at about 4 °C using bagged refrigerant.

SAMPLE PREPARATION:

3. Perform a creatinine determination on an aliquot of the urine (e.g., [6]). 

CALIBRATION AND QUALITY CONTROL:

4. Prepare at least five working standards in the range 0.1 to 100 μg F- /mL by appropriate dilutions of 

the calibration stock solution with distilled water.
FLUORIDE in URINE: METHOD 8308, Issue 3, dated 14 March 2016 - Page 3 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

5. Analyze a set of working standards together with the samples and blanks (steps 9 through 12) 

starting with the lowest concentration.

NOTE: Working standards, samples, and blanks must be analyzed under the same conditions, 

including temperature, for accurate results.

6. Prepare a semi-log calibration curve plotting millivolts on the linear scale (y-axis) and fluoride 

concentration, μg/mL, on the log scale (x-axis). 

7. Maintain standardization by running a standard with every 10 specimens.

8. Run a spiked urine control specimen with every 10 specimens to maintain quality assurance.

NOTE: Urine used for spiked controls must be analyzed before use to determine background 

fluoride concentration. 

MEASUREMENT:

9. Add 10 mL well-mixed urine and 10 mL TISAB to a 50-mL plastic beaker.

10. Place a small stirring bar into beaker and mix continuously on a magnetic stirrer at room 

temperature.

11. Immerse electrodes. Allow sample to mix for 2 to 3 min and then record millivolt reading.

12. Rinse electrodes and stirring bar thoroughly with distilled water and wipe dry with tissue before next 

sample analysis. 

CALCULATIONS:

13. Convert the millivolt readings to fluoride concentration using the calibration curve from step 6.

14. Express fluoride concentration as mg F-

/g urinary creatinine.

GUIDES TO INTERPRETATION: 

Urine concentrations of fluorides in normal non-occupationally exposed workers have been reported to 

range from 0.2 to 3.2 mg/L depending on dietary intake [7]. Pre-shift levels of less than 4 mg/g creatinine 

and post-shift levels of less than 7 mg/g creatinine appears to protect workers against bony fluorosis [8]. 

NIOSH has recommended that post-shift urine specimens should not exceed 7 mg/L (corrected to a specific 

gravity of 1.024) and pre-shift specimens should not exceed 4 mg/L (corrected to a specific gravity of 1.024) 

[1,2].

The Biological Exposure Indices (BEI) for fluoride (as of the date of this method’s publication) are 2 mg/L 

prior to shift and 3 mg/L at end of shift [9]. This BEI changed in 2011.

EVALUATION OF METHOD

No formal method evaluation has been reported; however, Tusl [3] reported recoveries of added fluoride 

from 94 to 100%. Precision based on analysis of 25 specimens in triplicate is estimated to be better than

SS̅

rr = 0.04. This method employs standard methodology that has been shown to provide adequate 

performance data for decades. Additional evaluation data may be found in, but is not limited to, the 

following references [10-13].

REFERENCES:

[1] NIOSH [1976]. NIOSH criteria for a recommended standard: occupational exposure to inorganic 

fluorides. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 76-

103.

[2] NIOSH [1976]. NIOSH criteria for a recommended standard: occupational exposure to hydrogen 

fluoride. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 76-

143.
FLUORIDE in URINE: METHOD 8308, Issue 3, dated 14 March 2016 - Page 4 of 4 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[3] Tusl J [1970]. Direct determination of fluoride in human urine using fluoride electrode. Clin Chim 

Acta 27:216-218.

[4] NIOSH [1977]. Fluoride in urine: P&CAM 114. In: Taylor DG, ed. NIOSH manual of analytical methods. 

2nd ed., Vol 1. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, Center for Disease 

Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) Publication No. 77-

157A.

[5] CDC [2007]. 2007 Guidelines for isolation precautions: preventing transmission of infectious agents 

in healthcare settings [http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html]. Date 

accessed: March 2016.

[6] Tietz NW [1976]. Fundamentals of clinical chemistry. 2nd ed. Philadelphia, PA: W. B. Saunders Co., pp. 

994-999.

[7] Baselt RC [1980]. Biological monitoring methods for industrial chemicals. Davis, CA: Biomedical 

Publications, pp. 140-143.

[8] Lauwreys RR [1983]. Industrial chemical exposure: guidelines for biological monitoring. Davis, CA: 

Biomedical Publications, pp. 26-27, 134.

[9] ACGIH [2014]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical 

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American 

Conference of Governmental Industrial Hygienists [www.acgih.org]. Date accessed: May 2014.

[10] Singer L, Armstrong WD [1973]. Determination of fluoride in ultrafiltrates of sera. Biochem Med 

8:415-422.

[11] Chiba K, Tsunoda K, Haraguchi H, Fuwa K [1980]. Determination of fluoride in urine and blood serum 

by aluminum monofluoride molecular absorption spectrometry and with a fluoride ion selective 

electrode. Anal Chem 52:1582-1585.

[12] Milde D, Nováková K, Čermáková I [2004]. Fluoride determination in urine with fluoride ion selective 

electrode: within laboratory method and sample storage optimization. Acta Univ Palacki Olomu, Fac 

Rerum Nat, Chem 43:104-109.

[13] Singh B, Gaur S, Garg VK [2007]. Fluoride in drinking water and human urine in Southern Haryana, 

India. J Hazard Mater 144:147-151.

METHOD WRITTEN BY:

William P. Tolos, NIOSH. 

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
2,5-HEXANEDIONE 

in URINE 8318 

CH3COCH2CH2COCH3 MW: 114.14 CAS: 110-13-4 RTECS: MO3150000

METHOD: 8318, Issue 1 EVALUATION: FULL Issue 1: 15 August 2017

NIOSH and OSHA: NA

Because data on exposure limits and guidelines may 

change over time, NIOSH recommends referring to the 

following sources for updated limits and guidelines 

concerning 2,5-hexanedione as well as its use as a 

marker for other compounds [1-3].

PROPERTIES: Clear liquid; MP -5.5 °C; BP 191 °C; 

d2 5 0.973 g/mL; VP 0.43 mm Hg at 20 °C [4]

BIOLOGICAL INDICATOR OF: exposure to n-hexane and methyl n-butyl ketone

SYNONYMS: acetonyl acetone; diacetonyl; alpha, beta-diacetylethane; 1,2-diacetylethane; 2,5-diketohexane; 2,5-dioxohexane 

SAMPLING

SPECIMEN: two urine samples (one at the beginning of 

work week and one end of shift late in the 

work week)

VOLUME: complete void stored in a polyethylene 

screw-cap bottle 

PRESERVATIVE:none

SHIPMENT: refrigerated or frozen in well-insulated 

container

SAMPLE

STABILITY: stable 7 days @ 24 °C and at least 30 days @ 

4 °C [5]

CONTROLS: collect and pool urine from matched 

population of unexposed workers if 

possible then refrigerate immediately

ACCURACY

RANGE STUDIED: 2.1 - 212 mg/L [5]

BIAS: -0.0189

OVERALL

PRECISION (SS�rrrr): 0.0235 [5]

ACCURACY: ± 5.6%

RECOVERY: 102% (Day 32) [5]

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, FLAME 

IONIZATION DETECTOR

ANALYTE: 2,5-hexanedione (2,5-HD)

TREATMENT: acid hydrolysis; dichloromethane 

extraction

INJECTION VOLUME: 3μL; splitless for 0.7 min

TEMPERATURE

-INJECTION: 200 °C

-DETECTOR: 250 °C

-COLUMN: 45 °C (2 min hold); 45 to 60 °C @ 5 °C/min; 

60 to 140 °C @ 10 °C/min; 140 to 220 °C @

50 °C/min; 220 °C (5 min. hold); final hold 

time may be extended if necessary.

CARRIER GAS: Helium, at 15 psi head pressure

COLUMN: Capillary, fused silica, polyethylene glycol, 

30 m x 0.32 mm ID, 0.5 μm film

CALIBRATION: analyte in control urine; 2-methyl-3-

heptanone or other appropriate internal 

standard

ESTIMATED LOD: 0.2 mg/L in pooled urine [5]

APPLICABILITY: Can be used in monitoring the exposure of workers to n-hexane or methyl n-butyl ketone. This method measures 

“total” amount of 2,5-HD, not the “free” amount. See further discussion in Other Methods section.

INTERFERENCES: Other compounds besides n-hexane (e.g. methyl n-butyl ketone) can be metabolized into 2,5-HD. Acid hydrolysis can 

convert other metabolites (e.g. 4,5-dihydroxy-2-hexanone) into 2,5-HD, giving values higher than non-hydrolysis methods [6-8]. Coexposure 

to toluene and methyl ethyl ketone has been shown to inhibit n-hexane metabolism [9]. General population background levels of 2,5-HD vary 

by age and gender but average 0.4 mg/L [10]. 

OTHER METHODS: This method is based on the acid hydrolysis method of Iwata et al. [9], and Fedtke and Bolt [8] as described by Kawai et al. 

[11]. As discussed in greater detail in the Backup Data Report [5], this method uses an acid hydrolysis step to convert some of the intermediate 

metabolites to 2,5-HD, a measure of “total” 2,5-HD. A non-hydrolysis method, which measures the “free” 2,5-HD, was not investigated in this 

work. The “free” amount is associated with the Biological Exposure Index [1]. Other exposure values are more associated with the “total” amount 

found [2,3] and still others give the option of either approach [12]. Users should recognize the advantages and limitations of each approach when 

selecting a methodology. More on this can be found in the Evaluation of Method section and the backup data report [5].

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 
2,5 HEXANEDIONE in urine: Method 8318, Issue 1, dated August 15, 2017 Page 2 of 7 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

Control urine, collected and pooled from 

unexposed workers*

2,5-hexanedione stock solutions: Prepare 

by diluting the appropriate amounts of the 

pure analyte in methanol. Prepare working 

standards by diluting aliquots of the stock 

solution with pooled urine*

Extraction Solution: Dissolve 5 mg of the 

internal standard (2-methyl-3-heptanone) 

into dichloromethane; bring to 1.0 L total 

volume*

Hydrochloric acid (HCl), concentrated* 

(Trace metal grade or higher)

Methanol* (HPLC grade or higher)

Dichloromethane* (ACS reagent grade or 

higher)

Helium, purified

Hydrogen, prepurified

Air, filtered

* See SPECIAL PRECAUTIONS

EQUIPMENT:

Bottles, polyethylene screw-top, 125-mL

Gas chromatograph, with flame ionization 

detector, data system, and column (page 

8318-1)

Vials, autosampler, glass, 2-mL, PTFE-lined 

crimp caps

Bagged refrigerant or dry ice

Pipettes, 5-, 2-, and 1-mL, plastic or 

serological

Heated water bath

Centrifuge

Culture tubes, glass, 13 x 100 mm with 

PTFE-lined screw caps

Syringes, 10-, 100-mL 

Pipets, glass

pH meter

Magnetic stirrer

Volumetric flasks, glass, 1-L, 10-mL

SPECIAL PRECAUTIONS: Standard precautions should always be used when handling bodily fluids and/or

extracts of bodily fluids [13]. Handle urine specimens and urine extracts using powder-free latex or nitrile 

gloves. All work should be performed in a fume hood since all chemicals are respiratory irritants and some 

may cause narcotic effects. Inhalation of large amounts of methanol or dichloromethane can cause 

unconsciousness and even death. 2,5-HD is a neurotoxin at high exposure levels. Methanol is a fire hazard 

and causes blindness or death if ingested. Hydrochloric acid is an extremely corrosive chemical capable of 

severe tissue damage. 

SAMPLING:

1. Collect a complete void of urine in a 125-mL polyethylene bottle. Collect two urine specimens from

each worker: one specimen at the beginning of the work week and one specimen at end of shift late

in the work week.

2. Collect and pool urine from unexposed workers to be used for controls.

3. Tightly cap each bottle and ship refrigerated or frozen in a well-insulated container.

NOTE: Commercial shippers have special labeling requirements for packages containing biological

samples and dry ice.

SAMPLE PREPARATION:

4. Allow urine specimens to reach room temperature.

NOTE: If desired, remove an aliquot of urine to determine creatinine levels (g/L urine).
2,5 HEXANEDIONE in urine: Method 8318, Issue 1, dated August 15, 2017 Page 3 of 6

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

5. Pipet 5.0 mL of urine into a 13 x 100 mm culture tube or other appropriate container.

6. Utilizing a magnetic stirrer, add concentrated HCl until the pH of the urine is within the desired

range (0.5 to 1.0).

NOTE: An exact control of the sample’s pH is critical for consistent and reproducible results [11].

7. Cap tube and heat in a water bath at 100 °C for 30 min.

8. Remove from water bath and allow to cool completely before proceeding.

9. Pipet 2 mL of the extraction solution, containing the internal standard, into the tube.

10. Cap and shake vigorously for 1 min. Allow the phases to separate. A centrifuge is recommended to

aid in separation.

11. Remove a portion of the extraction solution (bottom layer) using a glass pipet and transfer it to a

2-mL glass GC autosampler vial. Cap and analyze.

CALIBRATION AND QUALITY CONTROL:

12. Calibrate daily with at least six working standards covering the concentration range of the

specimens.

13. Prepare each working standard by diluting a known amount of 2,5-HD stock solution into enough

pooled urine to make a total of 5.0 mL.

14. Prepare at least one pooled urine blank by transferring 5.0 mL of pooled urine (the same pooled

urine used for creating the working standards) into a culture tube.

15. Prepare at least two levels of quality control (QC) spikes of 2,5-HD in pooled urine to be analyzed

with each analysis batch. These levels should be at approximately 10 times the limit of quantitation

(LOQ) and 200 times the LOQ, but can be adjusted to better suit the anticipated levels of the

specimens. QC samples should be analyzed with every batch such that they constitute 10% of the

specimen batch.

16. Process the 5 mL of each working standard and each pooled urine blank using the same procedure

as for the specimens (steps 5 through 11).

17. Analyze the working standards, the pooled urine blanks, the QC samples, and the samples together.

18. QC values should normally be within ±20% of the spiked values. If not, the batch is considered out of

control, the batch data discarded, and corrective actions taken before more specimens are analyzed.

MEASUREMENT:

19. Set gas chromatograph according to manufacturer’s recommendations and to conditions given on

page 8318-1. Typical retention times for the two compounds under these conditions are 4.7 min for

the internal standard and 10.5 min for 2,5-HD.

20. Inject each of the standards, QC samples, samples, and blanks.

21. Measure peak area. Normalize analyte response by dividing the peak area of the analyte by the peak

area of the internal standard on the same chromatogram.

22. Prepare a calibration graph by plotting, for each working standard, the normalized analyte response

(peak area of analyte divided by the peak area of the internal standard on the same chromatogram)

on the y-axis vs. mg of analyte/L of urine on the x-axis. The simplest model that adequately describes

the data should be used but either a linear (mostly likely 1/x weighted because of the range of the

calibration curve) or a quadratic model may be utilized in processing the analytical data.

The standard curve should have a coefficient of determination (r2

) of equal to or greater than 0.98 to

be acceptable for use. Furthermore, when each standard is plugged back into the calibration
2,5 HEXANEDIONE in urine: Method 8318, Issue 1, dated August 15, 2017 Page 4 of 6

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

equation, the value should be within ±20% of the expected. Acid hydrolysis can create detectable 

amounts of 2,5-HD in pooled urine blanks. Before plotting the calibration graph, subtract the 

normalized analyte response of the pooled urine blank from the normalized analyte response of 

each working standard.

CALCULATIONS:

23. Determine the 2,5-HD concentration, Cs (mg/L) in the urine specimen from the calibration graph

created in step 16.

24. If desired, calculate the concentration of 2,5-HD in milligrams per gram of creatinine C (mg/g) in the

urine specimen by dividing by the creatinine value Cr (g creatinine/L urine) obtained from the

aliquot removed in step 4. Compare the results from before and after exposure. Also compare the

results of the “exposed” group to that of the control group.

CC = CCss

CCrr

EVALUATION OF METHOD

This method was evaluated over the ranges specified on page 8318-1. These ranges represent from 3 times 

the LOQ to 300 times the LOQ. Seven replicates were analyzed at each level. The average recoveries at the 

various levels ranged from 97% to 103%. The limit of detection (LOD) and Limit of Quantification (LOQ) 

were determined by preparing in duplicate and analyzing a series of standards with the data fitted to a 

quadratic curve. The LOD and LOQ were estimated according to Burkart’s Method [14]. A long-term storage 

study was carried out at the 10 x LOQ level. Pooled urine samples spiked with the analyte were stored at 

4 °C for 1, 4, 7, 10, 21, or 32 days and then analyzed. All recoveries were nearly 100%. Spiked pooled urine 

samples were stable after 7 days at room temperature.

The issue of measuring “total” (the 2,5-HD determined after a hydrolysis step) versus “free” (the 2,5-HD 

determined without a hydrolysis step) is a contentious one among world bodies that develop 

biomonitoring guidelines. As previously stated, this method uses a hydrolysis step and was not evaluated 

without it. Therefore, it measures the “total” amount. NIOSH does not desire to step into the debate over 

which may be better, but just to offer this method as one way to assess exposure. Some brief summaries of

the two approaches: “total” produces a larger number, requiring less sensitivity from the analytical 

methodology but is subject to higher variability resulting primarily from two sources: variability in 

hydrolysis procedure [11] and production of 2,5-HD from other metabolites in human urine that vary in 

concentration. The “free” approach gives a smaller number but tends to be more specific for n-hexane and 

methyl n-butyl ketone exposure as the other metabolites in urine do not interfere, so more sensitivity is 

required but background levels are greatly reduced.
2,5 HEXANEDIONE in urine: Method 8318, Issue 1, dated August 15, 2017 Page 5 of 6

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] AIHA [2014]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American

Conference of Governmental Industrial Hygienists, www.acgih.org.

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area [2014].

List of MAK and BAT values 2014; Maximum concentrations and biological tolerance values at the

workplace. DFG: Deutsche Forschungsgemeinschaft,

http://onlinelibrary.wiley.com/book/10.1002/9783527682027.

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail.

Valeurs limites d’exposition aux postes de travail. 3rd Ed., https://www.suva.ch/

[4] Sigma-Aldrich [2016]. 165131 ALDRICH, 2,5-Hexanedione,

http://www.sigmaaldrich.com/catalog/product/aldrich/165131

[5] Downs CC, Perkins JB [2002]. Back-up data report for 2,5-hexanedione method development.

DataChem Laboratories, Inc. under NIOSH contracts CDC-200-95-2955 and CDC-200-2001-08000.

[6] Fedtke N, Bolt HM [1986]. Detection of 2,5-hexanedione in the urine of persons not exposed to nhexane. Int Arch Occup Environ Health 57: 143-148.

[7] Fedtke N, Bolt HM [1987]. 4,5-Dihydroxy-2-hexanone: a new metabolite of n-hexane and of 2,5-

hexanedione in rat urine. Biomed Environ Mass Spectrom 14:563-572.

[8] Fedtke N, Bolt HM [1987]. The relevance of 4,5-dihydroxy-2-hexanone in the excretion kinetics of nhexane metabolites in rat and man. Arch Toxicol 61:131-137.

[9] Iwata M, Takeuchi Y, Hisanaga N, Ono Y [1983]. A study on biological monitoring of n-hexane

exposure. Int Arch Occup Environ Health 51:253-260.

[10] Tondel M, Murgia N, Persson B, Lindh J, Vrethem M, Fredrikson M [2011]. 2,5-Hexandione in the

general population: environmental exposure or endogenous production? Epidemiology 22(1):S34-

S35.

[11] Kawai T, Mizunuma K, Yasugi T, Uchida Y, Ikeda M [1990]. The method of choice for the determination

of 2,5-hexanedione as an indicator of occupational exposure to n-hexane. Int Arch Occup Environ

Health 62:403-408.

[12] JSOH [2016]. Recommendation of occupational exposure limits (2016-2017). The Japan Society for

Occupational Safety and Health. Journal of Occupational Health 58(5):489-518. 
2,5 HEXANEDIONE in urine: Method 8318, Issue 1, dated August 15, 2017 Page 6 of 6

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[13] CDC [2007]. 2007 Guidelines for isolation precautions: preventing transmission of infectious agents in

healthcare settings, http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html.

[14] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene

chemistry laboratory. Appl Ind Hyg 1(3):153-155.

METHOD WRITTEN BY:

Corey C. Downs and James B. Perkins, DataChem Laboratories, Inc., Salt Lake City, Utah under NIOSH 

Contracts CDC-200-95-2955 and CDC-200-2001-08000. Final editing by Dale A. Shoemaker, Ph.D., 

NIOSH/DART/CEMB.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ACETONE and METHYL ETHYL KETONE in urine 8319

(1) Acetone, CH3COCH3 MW: 58.08 CAS: 67-64-1 RTECS: AL3150000

(2) Methyl ethyl ketone, CH3COCH2CH3 MW: 72.11 CAS: 78-93-3 RTECS: EL6475000

METHOD: 8319, Issue 1 EVALUATION: FULL Issue 1: 28 October2014

OSHA & NIOSH: N/A

Because data on exposure limits and guidelines may 

change over time, NIOSH recommends referring 

to the following sources for updated limits and 

guidelines [1-4].

PROPERTIES: (1) bp 56.2 °C; d20 0.789 g/mL 

(2) bp 79.6 °C; d20 0.805 g/mL

BIOLOGICAL INDICATOR OF: 

 Exposure to (1) acetone

 (2) methyl ethyl ketone

SYNONYMS: (1) dimethyl ketone; 2-propanone; ketone propane; dimethyl formaldehyde; pyroacetic ether

(2) 2-butanone; MEK; butanone; methyl acetone; butan-2-one; oxobutane

SAMPLING

SPECIMEN: Two urine samples (before and after exposure)

VOLUME: Fill a 20-mL amber VOA vial leaving minimal 

headspace

SHIPMENT: Ship in an insulated container with bagged 

refrigerant

SAMPLE

STABILITY: Stable at least 30 days at 4 °C [5]

CONTROLS: Collect and pool urine from matched 

population of unexposed workers if possible, 

then refrigerate immediately

ACCURACY

 RANGE STUDIED: (1) 2.1 - 606 mg/L [5]

(2) 2.2 - 617 mg/L [5]

ACCURACY: (1) ± 11.5%, (2) ± 15.0%

BIAS: (1) -0.0444, (2) -0.0782

OVERALL

PRECISION (ŜrT): (1) 0.0468 [5], (2) 0.0507

SAMPLE

STABILITY: (1) 101% (Day 30) [5]

(2) 105% (Day 30) [5]

MEASUREMENT

 TECHNIQUE: GAS CHROMATOGRAPHY, FLAME 

IONIZATION DETECTOR, HEADSPACE 

ANALYTES: Acetone and methyl ethyl ketone

INJECTION

VOLUME: 500 μL or timed (see measurement 

section)

TEMPERATURE

-INJECTION: 180 °C

-DETECTOR: 250 °C

-COLUMN: 40 °C (4 min); 40 to 60 °C @ 3 °C/min; 60 to 

220 °C @ 20 °C/min; 220 °C (2 min)

CARRIER GAS: Helium, at 15 psi head pressure

COLUMN: Capillary, fused silica, 

6% cyanopropylphenyl, 

94% dimethylpolysiloxane, 75 m x 

0.53 mm ID, 3.0 μm film thickness

CALIBRATION: Analyte in control urine; 2-pentanone or 

other appropriate internal standard

ESTIMATED LOD: (1) 0.6 mg/L in pooled urine [5] 

(2) 0.6 mg/L in pooled urine [5]

APPLICABILITY: This method can be used in the analysis of acetone and methyl ethyl ketone in urine specimens. These compounds may be found in the urine of individuals exposed to acetone and methyl ethyl ketone [6,7].

INTERFERENCES: Acetone is a metabolite of 2-propanol; MEK is a metabolite of 2-butanol. Exposure to 2-propanol or 2-butanol 

may result in increased acetone or MEK excretion, respectively [8,9]. Diabetes and fasting also produce elevated urinary acetone levels [10]. Ethanol reduces MEK metabolism and thus increases the MEK concentration in urine [11]. Gender differences 

and use of hormonal contraceptives have been shown to affect the metabolism and excretion of MEK [12].

OTHER METHODS: There are several commercially-available, direct-reading, dipstick-type tests that are non-specific for 

ketones in urine. These are often used in hospitals or by diabetic patients. The Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) has published a headspace method for a variety of alcohols and ketones in urine and blood [13].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ACETONE and METHYL ETHYL KETONE in urine: METHOD 8319, Issue 1, dated 28 October 2014 - Page 2 of 6

REAGENTS:

1. Pooled control urine collected from unexposed 

workers*

2. Acetone (≥99%) and MEK (≥99%) stock 

solution: Prepare by diluting the appropriate 

amounts of the pure analyte in water.* 750 μL 

in 10 mL gives ~60 mg/mL of each. Be certain 

to use the densities and purity factors when 

calculating the exact concentrations. [5]

3. Internal standard solution: Dilute 80 mg of 

2-pentanone (≥99%) in enough water to make 

1.0 L (80 mg/L).* Alternatively, 100 μL can be 

added to 1 L which gives roughly the same 

concentration.

4. Water, ASTM Type II [14]

5. Helium, purified

6. Hydrogen, prepurified

7. Air, filtered, prepurified

* See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Bottles, polyethylene screw-top, 125-mL

2. Volatile organic analysis (VOA) vials, 20-mL, 

amber, polytetrafluoroethylene (PTFE) caps

3. Gas chromatograph, with flame ionization 

detector (FID), data system and column 

(p. 8319-1)

4. Headspace autosampler

5. Bagged refrigerant

6. Pipette, 10-mL, plastic or serological

7. Headspace vials, 20-mL, PTFE/Butyl septa, 

aluminum crimp cap

8. Microliter syringes, 10-μL, 100-μL, 1-mL

9. Volumetric flask, 10-mL

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals and 

human urine products. Disposable plastic, glass, and paper (pipet tips, gloves, etc.) that contact 

urine should be placed in a biohazard container. Standard precautions should always be used when 

handling bodily fluids and/or extracts of bodily fluids [15]. Handle urine samples and urine extracts 

using proper gloves. Acetone and MEK are highly flammable liquids. All work should be performed 

in a fume hood since both chemicals are respiratory irritants and may depress the central nervous 

system at high exposure levels.

SAMPLING:

1. Collect urine in a 125-mL polyethylene bottle or other suitable container. Collect at least two urine 

samples from each worker. Collect one sample before exposure and one sample after exposure.

NOTE: It is important to avoid contamination of the urine samples by making sure that samples are 

collected in a clean area away from the source(s) of exposure and under hygienic conditions 

(after washing hands.)

2. For each sample, immediately transfer from the 125-mL polyethylene bottle enough urine to fill a 

20-mL amber VOA vial such that a minimal headspace is left. Cap the containers tightly. When the 

VOA vial is inverted no air bubbles should be present. Refrigerate after collection.

3. Collect and pool urine from unexposed workers to be used for controls. Refrigerate after collection.

4. Ship the VOA vials and pooled control urine in a refrigerated, well-insulated container. Store 

refrigerated upon receipt.

SAMPLE PREPARATION:

5. Allow urine to reach room temperature.

6. Pipet 10.0 mL of urine from the VOA vial into a 20-mL headspace vial.

7. Add 0.5 mL of the internal standard solution.

8. Cap vial immediately.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ACETONE and METHYL ETHYL KETONE in urine: METHOD 8319, Issue 1, dated 28 October 2014 - Page 3 of 6

CALIBRATION AND QUALITY CONTROL:

9. Calibrate daily with at least six working standards covering the concentration range of the method 

(2 to 600 mg/L).

NOTE: If the range of concentrations of the samples is known (or expected), the calibration curve 

range can be adjusted accordingly.

a. Prepare a diluted stock solution by pipetting 1 mL of the concentrated stock solution into a 

10-mL volumetric flask and filling to the mark with pooled urine.

b. Prepare each working standard by diluting a known amount of the diluted stock solution 

prepared in Step 9a into enough pooled urine to make a total of 10 mL. 

NOTE: A second, more dilute stock solution in urine can be prepared, if desired, so that the lowest 

calibration standards are made using more easily-measured spiking volumes.

c. Prepare at least one pooled urine blank by transferring 10 mL of pooled urine (the same pooled 

urine used for creating the working standards) into a vial.

d. Process the 10 mL of each working standard and each pooled urine blank using the same 

procedure as for the samples (steps 5 through 8).

e. Analyze the working standards, the pooled urine blanks, and the samples together. FIGURES 1 

and 2 show representative chromatograms of blank and fortified urines.

f. Prepare a calibration graph by plotting, for each working standard, the normalized analyte 

response (peak area of analyte divided by the peak area of the internal standard on the same 

chromatogram) on the y-axis vs. μg of analyte/mL of urine on the x-axis. The simplest model 

that adequately describes the data should be used but either a linear (mostly likely 1/x weighted 

because of the range of the calibration curve) or a quadratic model may be utilized in processing 

the analytical data. Because humans can endogenously produce both acetone and methyl 

ethyl ketone, the compounds may be detected in the pooled urine blanks. Before plotting the 

calibration graph, subtract the normalized analyte response of the pooled urine blank from 

the normalized analyte response of each working standard. The standard curve should have 

a coefficient of determination (r2

) of equal to or greater than 0.98 to be acceptable for use. 

Furthermore, when each standard is substituted back into the calibration equation, the value 

should be within ±20% of the expected.

10. Prepare at least two levels of quality control (QC) samples by spiking both analytes in urine. These 

levels should be at approximately 10-fold the limit of quantitation (LOQ) and 200-fold the LOQ, but 

can be adjusted to better suit the anticipated levels of the sample set. Unspiked samples of the 

urine used to prepare the QC samples should be analyzed to determine the blank level and the true 

target level. QC samples should be analyzed with every batch such that they constitute 10% of the 

sample batch.

11. QC values should normally be within ±20% of the spiked values. If not, the batch is considered out 

of control, the batch data discarded, and corrective actions should be taken before more samples 

are analyzed.

MEASUREMENT:

12. Set gas chromatograph according to manufacturer’s recommendations and to conditions given on 

p. 8319-1.

13. Set headspace autosampler according to manufacturer’s recommendations and to the following 

conditions: (NOTE: different types of headspace samplers may require alternative conditions and 

some of these might not apply.)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ACETONE and METHYL ETHYL KETONE in urine: METHOD 8319, Issue 1, dated 28 October 2014 - Page 4 of 6

a. Suggested conditions for a balanced-pressure type of headspace system:

Transfer Temp: 129 °C Withdrawal: 0.2 min

Thermostat Time: 30 min Needle Temp: 120 °C

GC Cycle Time: 28 min Sample Temp: 80 °C

 Pressurize: 1.0 min Inject: 0.08 min

b. Suggested conditions for a syringe-injection type of headspace system:

Incubation Temp: 95 °C Injection volume: 500 μL

Incubation Time: 15 min Fill speed: 120 μL/sec

Agitation speed: 250 rpm Delay: 5 sec

Run time: 26 min Injection speed: 300 μL/sec

Syringe Temp: 95 °C Delay: 500 msec

14. Measure peak area. Normalize the analyte response by dividing the peak area of the analyte by the

peak area of the internal standard on the same chromatogram for each working standard, sample,

and pooled urine blank.

CALCULATIONS:

15. Determine both the acetone and MEK concentrations (mg/L) in the urine sample from the

calibration graph prepared in Step 9f.

EVALUATION OF METHOD:

This method was evaluated over the ranges of the two analytes specified on p. 8319-1. These ranges 

represent from 1 x LOQ to 300 x LOQ. Six to seven replicates were analyzed at each level. The average 

recoveries at the various levels ranged from 94% to 106% for acetone and 85% to 98% for MEK. The 

LOD and LOQ were determined by preparing and analyzing a series of standards in duplicate with the 

data fitted to a quadratic curve. The LOD and LOQ were estimated according to Burkart’s Method [16]. 

A long-term storage study was carried out at the 10 x LOQ level. Pooled urine samples spiked with the 

analytes were stored at 4 °C for 1, 4, 7, 10, 21, or 30 days and then analyzed. All recoveries were nearly 

100%. When stored at room temperature, a significant reduction in analyte recovery was observed after 

7 days [5].

REFERENCES:

[1] ACGIH [2013]. TLVs® and BEIs® based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio:

American Conference of Governmental Industrial Hygienists. www.acgih.org [Date accessed: August 

2014.]

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area

[2013]. List of MAK and BAT values 2013; Maximum concentrations and biological tolerance values

at the workplace. DFG: Deutsche Forschungsgemeinschaft, Wiley-VCH Verlag GmbH & Co. www.dfg.

de/en/dfg_profile/publications/senate_commissions/index.html#micro7535782 [Date accessed:

August 2014.]

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail.

Valeurs limites d’exposition aux postes de travail. https://www.suva.ch/ [Date accessed: August

2014.]

[4] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA). 

https://www.dguv.de/ifa/index-2.jsp [Date accessed: August 2014.] 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ACETONE and METHYL ETHYL KETONE in urine: METHOD 8319, Issue 1, dated 28 October 2014 - Page 5 of 6

[5] Downs CC, Perkins JB [2002]. Back-up data report for acetone and methyl ethyl ketone method 

development. DataChem Laboratories, Inc. under NIOSH contracts CDC-200-95-2955 and 

CDC-200-2001-08000.

[6] Pezzagno G, Imbriani M, Ghittori S, Capodaglio E, Huang J [1986]. Urinary elimination of acetone in 

experimental and occupational exposure. Scand J Work Environ Health 12:603-608.

[7] Perbellini L, Brugnone F, Mozzo P, Cocheo V, Caretta D [1984]. Methyl ethyl ketone exposure in 

industrial workers. Uptake and kinetics. Int Arch Occup Environ Health 54:73-81.

[8] Kawai T, Yasugi T, Horiguchi S, Uchida Y, Iwami O, Iguchi H, Inoue O, Watanabe T, Nakatsuka H, Ikeda 

M [1990]. Biological monitoring of occupational exposure to isopropyl alcohol vapor by urinalysis 

for acetone. Int Arch Occup Environ Health 62:409-413.

[9] Williams RT [1950]. Detoxication mechanisms: the metabolism and detoxication of drugs, toxic 

substances and other organic compounds. Chapman & Hall Ltd., London p. 66.

[10] Bales JR, Bell JD, Nicholson JK, Sadler PJ [1986]. HNMR Studies of urine during fasting: excretion of 

ketone bodies and acetylcamitine. Magn Reson Med 3:849-856.

[11] Liira J, Riihimaki V, Engstrom K [1990]. Effects of ethanol on the kinetics of inhaled methyl ethyl 

ketone in man. Br J Ind Med 47:325-330.

[12] Tomicic C, Berode M, Oppliger A, Castella V, Leyvraz F, Praz-Christinaz S-M, Danuser B [2011]. Sex 

differences in urinary levels of several biological indicators of exposure: A human volunteer study. 

Toxicol Lett 202:218-225.

[13] Angerer J, Gündel J [2012]. Alcohols and Ketones [Biomonitoring Methods, 1996]. The MAK 

Collection for Occupational Health and Safety. pp. 1-33.

[14] ASTM [2011]. ASTM Standard D1193, Standard specification for reagent water. ASTM International, 

West Conshohocken, PA. DOI 10.1520/D1193-06R11.

[15] CDC [2007]. 2007 Guidelines for isolation precautions: Preventing transmission of infectious agents 

in healthcare settings. http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html. [Date 

accessed: April 2013.]

[16] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene 

chemistry laboratory. Appl Ind Hyg 1(3):153-155.

METHOD WRITTEN BY:

Corey C. Downs and James B. Perkins, DataChem Laboratories, Inc., Salt Lake City, Utah under NIOSH 

Contracts CDC-200-95-2955 and CDC-200-2001-08000. Final editing by Dale A. Shoemaker, Ph.D., 

NIOSH/DART.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their 

programs or products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses 

referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ACETONE and METHYL ETHYL KETONE in urine: METHOD 8319, Issue 1, dated 28 October 2014 - Page 6 of 6

FIGURE 1. Representative overlaid chromatograms from a high and low urine standard and urine blanks with 

 and without internal standard (IS), full-scale

FIGURE 2. Representative overlaid chromatograms from a high and low urine standard and urine blanks with 

 and without internal standard (IS), reduced-scale
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

BUTYLTIN TRICHLORIDE in Urine 8320

Formula: C4H9CI3Sn MW: 282.18 CAS: 1118-46-3 RTECS: WH6780000

METHOD: 8320, Issue 1 EVALUATION: FULL Issue 1: 10 January 2019 

OSHA: N/A

NIOSH REL: N/A

OTHER OELs: Because exposure limits and guidelines may 

change over time, NIOSH recommends references [1-3] for 

updated limits and guidelines.

PROPERTIES: Colorless liquid; MP -63  ̊C; BP 93  ̊C @ 10 

mmHg; Density - 1.693 g/mL @ 25  ̊C; VP 0.083 mmHg @ 20  ̊C 

[4]

SYNONYMS: butyltrichlorotin, butyltrichlorostannane, monobutyltin trichloride (MBTC)

SAMPLING

SPECIMEN: Spot urine specimens (before and after 

exposure; end of shift or end of work 

week) [5]

VOLUME: 15 mL minimum

PRESERVATIVE: None

SHIPMENT: Freeze urine: ship frozen in an insulated 

container

SAMPLE

STABILITY: Stable at least 300 days at -20 oC [6]

ACCURACY [6,7]

RANGE 

STUDIED: 0.12-58 μg/5-mL sample

BIAS: 0.063

OVERALL

PRECISION (SS�rrrr): 0.041

ACCURACY: ± 13.0%

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY/MASS 

SPECTROMETRY-SELECTED ION 

MONITORING (GC/MS-SIM)

ANALYTE: Derivative of butyltin trichloride: 

butyltriethyltin 

EXTRACTION: Liquid-liquid extraction 

DERIVATIZATION: Grignard reaction with ethylmagnesium

bromide

INJECTION VOLUME: 2 μL 

TEMPERATURE: 

INJECTION: 230 oC splitless for 0.75 min

DETECTOR: 285 oC

OVEN: 75 oC (1 min); 75 oC to 225 oC @ 12 oC/min; 

hold 5 min

MASS SELECTIVE 

DETECTOR PARAMETERS: 

SIM IONS (m/z): 178.9, 234.9

DWELL TIME: 200 msec per m/z

CARRIER GAS: Helium at 1.3 mL/min

COLUMN: Capillary, fused silica, phenyl arylene 

polymer virtually equivalent to (5% phenyl)-

methylpolysiloxane (30 m x 0.32 mm ID, 0.5 

μm film)

CALIBRATION: Spikes of known concentrations prepared in 

pooled urine

ESTIMATED LOD: 0.01 μg/5-mL sample [6]

APPLICABILITY: This method can be used for the Determination of butyltin trichloride in urine samples. It may also be useful for 

other butyltin chloride compounds. See Evaluation of Method section for more information.

INTERFERENCES: Because of the separation and mass-selective capabilities of GC/MS, interferences are not anticipated to have 

any impact on the analysis.

OTHER METHODS: None
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 2 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS:

1. Butyltin trichloride, ≥95% purity

2. Ethylmagnesium bromide, 1.0 M solution 

in tetrahydrofuran

3. Helium, UHP or Grade 5 (99.999%) or 

better

4. Sulfuric acid, 1 M: Dilute 5.6 mL conc. 

sulfuric acid to 100 mL deionized water

5. Citric acid/sodium citrate buffer (~pH 2.3): 

Dissolve 20.554 g citric acid monohydrate 

and 0.652 g sodium citrate dihydrate in 

1 L deionized water

6. Toluene, ACS reagent grade or better

7. Tropolone extraction solution: 0.1% in 

toluene: Dissolve 100 mg tropolone (2-

hydroxy-2, 4, 6-cycloheptatrienone, CAS# 

[533-75-5]) in 100 mL toluene

8. Alumina, acidic, Brockman activity I, 80-

200 mesh

9. Sodium sulfate, granular, anhydrous; 

reagent grade or better 

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Centrifuge tubes, 15-mL 

polypropylene with screw caps

2. Centrifuge, capable of at least 

2400 RPM

3. Pipettes, Pasteur, 14.6 cm (53⁄4")

4. Culture tubes, glass, ~8-mL (13 mm x 

100 mm), screw top with PTFE-lined 

caps

5. GC-MS capable of selected ion 

monitoring, column, autosampler and 

data collection system (p. 8320-1)

6. Microliter syringes

7. Volumetric flasks, glass, 5-mL

8. Pipettor, adjustable from 1-5 mL

9. Vortex mixer

10. Tumbler for centrifuge tubes, 

approximately 20 RPM

11. Bottles, polyethylene, 30- or 125-mL

12. Glass wool 

 

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals and 

human urine products. Disposable plastic, glass, and paper (pipette tips, gloves, etc.) that contact urine 

should be placed in a biohazard bag. Standard precautions should always be used when handling bodily 

fluids and/or extracts of bodily fluids [8]. Handle urine samples and urine extracts using proper gloves. All 

work should be performed in a fume hood.

SAMPLING:

1. Collect urine in a 30-mL or 125-mL polyethylene bottle or other appropriate container.

NOTE: Collect two urine specimens for each worker, one prior to exposure and one after exposure. 

The NIOSH Health Hazard Evaluation found no significant difference between end of shift 

and end of work week [5]. Also, collect and pool urine from unexposed subjects for use as a 

standard and quality control sample matrix. 

2. Freeze the urine (-20 oC or lower) and ship frozen in an insulated container.
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 3 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION:

3. Allow urine to reach room temperature and mix thoroughly.

4. Place 5 mL of the urine specimen in a 15-mL polypropylene centrifuge tube using a pipettor.

5. To the urine specimen, add:

a. 3 mL of ~pH 2.3 citric acid/sodium citrate buffer

b. about 350 mg anhydrous sodium sulfate (to break up emulsions) 

c. 1.5 mL 0.1% tropolone in toluene solution

6. Cap and tumble for one hour at ~20 RPM.

7. Centrifuge for at least 10 min at 2400 RPM (1050 G).

8. Transfer the upper toluene layer to an 8-mL glass culture tube with a PTFE-lined cap using a Pasteur 

pipette.

9. Repeat steps 5c through 8 and combine the extracts.

10. Add about 350 mg anhydrous sodium sulfate to the combined extracts to remove any of the 

aqueous phase that may have been inadvertently transferred.

11. Decant the dried sample from step 10 into a second 8-mL glass culture tube with a PTFE-lined cap.

12. Rinse the sodium sulfate residue in the first tube twice with a small (~300 μL) quantity of toluene 

which is then combined with the dried extract in the second tube.

13. Add 250 μL of the ethylmagnesium bromide solution to the extract, mix, and allow the mixture to 

react for 15-20 minutes.

14. Add 1 mL of 1 M sulfuric acid (to react with the remaining ethylmagnesium bromide) and vortex the 

tube for approximately 30 seconds.

15. Prepare a clean-up column in a 14.6 cm disposable Pasteur pipette by adding, in order, 1) a glass 

wool plug, 2) alumina, acidic, Brockmann activity I, 80-200 mesh, enough to form a column of 

approximately 1 cm, and 3) ~100 mg of anhydrous sodium sulfate.

16. After the layers separate, elute the top toluene layer through this clean-up column.

17. Collect the eluate in a 5-mL volumetric flask.

18. Rinse the tube with the sulfuric acid solution (Step 14) several times (a minimum of two) with small 

amounts (~300 μL) of toluene and elute the rinses through the clean-up column.

19. Flush the remaining sample from the column by adding small portions (~300 μL, a minimum of 

twice) of toluene to the top of the column.

20. Adjust the final volume to 5.0 mL with toluene as needed.

21. Transfer an aliquot of the extract to a GC vial for analysis.

CALIBRATION AND QUALITY CONTROL:

22. Calibrate daily with at least six working standards covering the expected concentration range of the 

samples. Suggested working standard concentration range is 0.007-150 μg/5 mL [6].

a. Prepare each working standard by spiking a known amount of the butyltin trichloride stock 

solution into enough pooled urine to make a total of 5.0 mL.

b. Prepare at least one pooled urine blank by transferring 5.0 mL of pooled urine (the same 

pooled urine used for creating the working standards) into a culture tube.

c. Prepare quality control (QC) samples by spiking known amounts of a separately prepared 

butyltin trichloride stock solution into pooled urine (the same pooled urine used for creating 

the working standards and blanks). It is recommended that two QC levels be prepared; a 

lower level at 3-5X the limit of quantitation (LOQ) and a higher level nearer the top end of 

the calibration curve. Run enough QC samples to be greater than 5% of the sample set. QC 

values should normally be within ±20% of the spiked values. If not, the batch is considered 

out of control, the batch data discarded, and corrective actions taken before more 

specimens are analyzed.

23. Process each working standard, blank, and QC sample using the same procedure as for the samples 

(steps 3 through 21).
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 4 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

24. Prepare calibration curves by plotting peak area (or peak height, although area is recommended) for 

the standards on the y-axis vs. concentration of analyte on the x-axis. The standard curve should 

have a coefficient of determination (r2) of equal to or greater than 0.98 to be acceptable for use. 

Furthermore, when each standard is plugged back into the calibration equation, the value should be 

within ±20% of the expected.

MEASUREMENT:

25. Set the gas chromatograph according to manufacturer’s recommendations and to conditions given 

on the front page of the method. With the chromatographic conditions outlined on page 1, the 

retention time of the butyltin trichloride derivative (butyltriethyltin) was 7.3 min. A sample 

chromatogram is shown in Figure 1.

26. Set the mass spectrometer according to manufacturer’s recommendations and to conditions given 

on the front page of the method. The mass spectrum for the butyltriethyltin can be found in the 

backup data report [6]. The most abundant isotopic mass of this compound is 264. The monitored 

ions are M-85 and M-29.

27. Inject each sample, working standard, blank, and QC sample.

28. Measure peak area or peak height in each chromatogram.

NOTE: If the sample peak area or height is greater than that of the highest standard, dilute the 

sample extract with toluene and reanalyze. Apply the appropriate dilution factor in the 

calculations.

CALCULATION OF ANALYTE PER SAMPLE: 

29. Determine mass of analyte in μg per 5-mL sample using the calibration curve prepared in step 24. 

Apply any dilution factor if needed.

NOTE: If the creatinine value of the sample was obtained, the concentration could be reported as 

μg/g creatinine if desired. The mass of analyte in μg/L can be calculated by multiplying the 

result by 200. The result can also be expressed in μg tin/volume by multiplying the μg per 5-

mL sample or μg/L by the butyltin trichloride conversion factor (%tin/mole) of 0.4207. 

EVALUATION OF METHOD:

The first procedure considered for this method was a modification of the Chau, et al. protocol using in situ

derivatization of the alkyltin compounds by sodium tetraethylborate [9]. While this procedure was found 

to be generally acceptable, recoveries were found both by Chau and DataChem Laboratories (DCL) to be 

inferior to the tropolone extraction/ethylmagnesium bromide derivatization. A composite procedure made 

up of modifications of elements of both Chau’s protocol and a procedure used by Meinema [10] was 

subsequently used at DCL. Since tin concentrations were expected to be low, a GC/MS-SIM analysis was 

chosen over the GC/FPD procedure. This method was evaluated over the following range: butyltin 

trichloride: 0.12-4.0 μg/5 mL sample. This range represents from 3X the estimated LOQ to 100X the 

estimated LOQ. Six replicates were analyzed at each level. The average recovery at the various levels 

ranged from 103% at 3X LOQ to 110% at 30X LOQ for butyltin trichloride [6]. Recoveries were determined 

by quantitating against spiked and derivatized standards in urine. The analytical range of the method was 

extended during a secondary laboratory evaluation step (known as a User Check). Two higher 

concentration levels were evaluated (12.5 and 60.5 μg/5 mL) by analyzing five replicates at each level. 

These levels showed recoveries of 90.2 and 92.7% with relative standard deviations of 4.0 and 3.6% 

respectively [6]. The Limit of Detection (LOD) and LOQ were determined by analyzing a series of 

derivatized spiked standards with the data fitted to a quadratic or linear curve, then estimated according 

to the Burkart Method [11]. The LOD was determined to be 0.01 μg/5 mL and the LOQ to be 0.04 μg/5 mL. 

A long-term storage study was carried out at the 10X LOQ level. Urine spiked with the analyte was stored at 

-20 oC for 1, 7, 32, 137, and 315 days and then analyzed. No significant losses were observed over the entire 

period [6].
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 5 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Additional compounds studied:

This method was originally developed to include two other butyltin chlorides: dibutyltin dichloride and 

tributyltin chloride, even though butyltin trichloride is the most occupationally relevant one as it is used in 

the glass industry. The analytical parameters obtained during method development for these two 

additional compounds were also acceptable [6]. However, the data obtained during the secondary 

laboratory evaluation for these additional compounds were not of such a quality that they could be 

considered fully evaluated. Both the method development data and secondary laboratory data can be 

found in the method backup data report [6]. Some of the pertinent analytical parameters for these two 

additional compounds are listed below as well as changes/additions to the written method needed for 

their analysis in case there is a desire to analyze for these compounds. The properties of these additional 

compounds are given in Table 1.

The ranges studied for the additional compounds were 0.18-6.0 μg/5-mL sample (with additional levels of 

11.9 and 57.5 μg/5 mL added during the User Check) for dibutyltin dichloride and 0.15-5.0 μg/5-mL sample

(with additional levels of 12.0 and 58.5 μg/5 mL added during the User Check) for tributyltin chloride.

NOTE: the additional levels added during the User Check were not used in the following accuracy, bias, 

and precision calculations. These values may all be found in the method backup data report [6].

ACCURACY: Dibutyltin dichloride: ± 13.5%

 Tributyltin chloride: ± 8.62%

BIAS: Dibutyltin dichloride: -0.091

Tributyltin chloride: -0.051

OVERALL

PRECISION (Ŝrt): Dibutyltin dichloride: 0.0266

Tributyltin chloride: 0.0215

ESTIMATED LOD: Dibutyltin dichloride: 0.02 μg/5 mL

 Tributyltin chloride: 0.01 μg/5 mL

NOTE: The tributyltin chloride LOD experiment had an interfering ion that caused this number to be more 

approximate than that of the other two compounds. See backup data report [6] for more information.

Recoveries ranged from 88.1% (at 30X LOQ) to 93.9% (at 10X LOQ) for dibutyltin dichloride and from 94.2% 

(at 10X LOQ) to 95.9% (at 100X LOQ) for tributyltin chloride [6].

Changes and additions to the method needed to incorporate these two additional compounds:

The mass spectrometer parameters should be changed to: SIM ions (m/z): 4-9.6 min: 178.9, 234.9, 262.9, 

290.9; 9.6 min to end of run: 234.9, 262.9, 290.9.

Additional reagents required: dibutyltin dichloride, ≥95% purity and tributyltin chloride, ≥95% purity.

Retention times: dibutyltin dichloride derivative (dibutyldiethyltin), 9.2 min; tributyltin chloride derivative 

(tributylethyltin), 10.7 min (these peaks are shown in Figure 1.)

Conversion factors (%tin/mole) to be used if results are expressed in μg tin/volume are 0.3907 for 

dibutyltin dichloride and 0.3647 for tributyltin chloride.
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 6 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] ACGIH [2016]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American

Conference of Governmental Industrial Hygienists, www.acgih.org.

[2] DFG [2014]. List of MAK and BAT values 2014: maximum concentrations and biological tolerance

values at the workplace. Bonn, Germany: DFG: Deutsche Forschungsgemeinschaft (German Research

Foundation), http://onlinelibrary.wiley.com/book/10.1002/9783527682027.

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail.

Valeurs limites d’exposition aux postes de travail. 3rd Ed. Lucerne, Switzerland: Schweizerische

Unfallversicherungsanstalt, https://www.suva.ch/.

[4] Sigma-Aldrich [2018]. www.sigmaaldrich.com.

[5] NIOSH [2005]. Evaluation of monobutyltin trichloride exposure among glass workers. By Dowell CH,

Page EH, Mueller C, Mortimer V, Snawder J. Cincinnati, OH: U.S. Department of Health and Human

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and

Health, Health Hazard Evaluation Report #2003-0016-2959.

[6] Wickman DC, Perkins JB [2005]. Backup data report for butyltin chlorides in urine method

development, DataChem Laboratories, Inc. under NIOSH contract CDC-200-2001-08000.

[7] NIOSH [1995]. Guidelines for air sampling and analytical method development and evaluation. By

Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of Health

and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational

Safety and Health, DHHS (NIOSH) Publication No. 95-117.

[8] CDC [2007]. 2007 Guidelines for isolation precautions: preventing transmission of infectious agents in 

healthcare settings. By Siegel JD, Rhinehart E, Jackson M, Chiarello L, and the Healthcare Infection

Control Practices Advisory Committee. Atlanta, GA: U.S. Department of Health and Human Services,

Centers for Disease Control and Prevention,

http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html.

[9] Chau YK, Yang F, Brown M [1997]. Evaluation of derivatization techniques for the analysis of

organotin compounds in biological tissue. Anal Chim Acta 338:51-55.

[10] Meinema HA, Burger-Wiersma T, Versluis-de Haan G, Gevers EC [1978]. Determination of trace

amounts of butyltin compounds in aqueous systems by gas chromatography/mass spectrometry.

Environ Sci Technol 12:288-293.

[11] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene

chemistry laboratory. Appl Ind Hyg 1(3):153-155.

[12] Chemical Book [2018]. www.chemicalbook.com/ChemicalProductProperty_EN_CB0262161.htm.

[13] Synquest [2018]. www.synquestlabs.com/product/id/56422.html. 
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 7 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TABLE 1. GENERAL INFORMATION for additional compounds [4]

Dibutyltin dichloride Tributyltin chloride

Formula C8H18CI2Sn C12H27CISn

M.W. 303.84 325.51

CAS# [683-18-1] [1461-22-9]

RTECS WH7100000 WH6820000

Synonyms Dibutyldichlorotin

dibutyldichlorostannane

chlorotributyltin

tributylchlorotin

TBTC

Pestanal®

MP

37-40 °C -9 °C [12]

-19 °C [13]

BP @ mmHg 135 °C @10mm 171-173 °C @ 25mm

Density 1.36 g/cm3 1.20 g/mL

Appearance Beige or colorless solid Clear, light yellow 

liquid

Vapor pressure 0.0012 mmHg @ 25 °C <0.01 mmHg @ 25 °C 
BUTYLTIN TRICHLORIDE IN URINE: METHOD 8320, Issue 1, 10 January 2019 - Page 8 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

FIGURE 1. Chromatogram of the three butyltin derivatives: butyltriethyltin (the derivative of butyltin 

trichloride), dibutyldiethyltin (the derivative of dibutyltin dichloride), and tributylethyltin (the derivative of 

tributyltin chloride)

METHOD WRITTEN BY:

Don C. Wickman and James B. Perkins, DataChem Laboratories, Inc., Salt Lake City, Utah under NIOSH 

Contract CDC-200-2001-08000. Final editing by Dale A. Shoemaker, Ph.D., NIOSH.

Disclaimer: Mention of any company or product does not constitute endorsement by the National

Institute for Occupational Safety and Health, Centers for Disease Control and Prevention. In addition, 

citations to Web sites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these Web sites. All Web addresses referenced in this document were accessible at the time of publication.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

 

o-CRESOL in URINE 8321

CH3C6H4OH MW: 108.14 CAS: 95-48-7 RTECS: GO6300000

METHOD: 8321, Issue 1 EVALUATION: FULL Issue 1: 18 March 2016

OSHA: N/A

NIOSH: N/A

Because data on exposure limits and guidelines may change 

over time, NIOSH recommends referring to references 1-4 for 

updated limits and guidelines concerning o-cresol and its use 

as a marker for other compounds.

PROPERTIES: colorless solid; MP 31 °C; BP 191 °C

Density (35 °C) = 1.0327 g/mL [5]

SYNONYMS: ortho-cresol, 2-cresol, o-cresylic acid, 1-hydroxy-2-methylbenzene, 2-hydroxytoluene, 2-methylphenol 

SAMPLING

SPECIMEN: Urine

VOLUME: 50-100 mL in screw cap bottle; minimum 

of 10 mL

SHIPMENT: < 6 o

C 

SAMPLE 

STABILITY: Stable at least 30 days at 4 °C [6]

CONTROLS: Collect urine from unexposed workers, 

pool and refrigerate

ACCURACY

ESTIMATED

 LOD: 0.01 μg/mL [6]

RANGE 

STUDIED: 0.0214 to 30.4 μg/mL [6]

ACCURACY: ± 21% [6]

BIAS: -0.0952 [6]

OVERALL

PRECISION (SS�rrrr): 0.0771 [6]

RECOVERY: 92 –112 % [6]

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY/MASS 

SPECTROMETRY with SELECTED ION 

MONITORING (GC/MS-SIM)

ANALYTE: o-Cresol

TREATMENT: Acid hydrolysis and liquid/liquid 

extraction

INJECTION

VOLUME: 2 μL

TEMPERATURE

 -INJECTION: 265 °C

 -DETECTOR: 285 °C

 -COLUMN: 50 °C(hold 2 min); 50 to 150 °C @ 

10 °C/min; 150 to 310 °C @ 20 °C/min 

(hold 5 min)

CARRIER GAS: Helium, at 1.3 mL/min

COLUMN: Capillary, fused silica, phenyl arylene 

polymer virtually equivalent to (5% 

phenyl)-methylpolysiloxane (30 m x 0.32 

mm ID, 0.5 μm film); close equivalent to 

USP G27

MS PARAMETERS: SIM (ions m/z 108, 107, 77 and 123) 

Quantifying ions: o-cresol: 108, 

nitrobenzene: 123. Confirmation ions: ocresol: 107, nitrobenzene: 77

DWELL TIME: 30 msec

CALIBRATION: o-Cresol spiked in control urine and 

nitrobenzene as internal standard
o-CRESOL in URINE: METHOD 8321, Issue 4, dated 18 March 2016 - Page 2 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

APPLICABILITY: This method can be used for the determination of total o-cresol in urine specimens. Cresols are excreted in urine 

primarily as conjugates. This method uses an acid hydrolysis step to convert the conjugates to free o-cresol. (See Evaluation of 

Method Section.) Exposure to o-cresol will cause elevated urinary levels; however, o-cresol is a metabolite produced from toluene 

exposure. Not only would toluene exposure interfere with determining o-cresol exposure, but measuring o-cresol in urine is one of 

the recommended methods for determining toluene exposure. Recommended levels for using o-cresol to measure toluene 

exposure (at the time of publication of this method) include: ACGIH-TLV, 0.3 mg/g creatinine [1]; DFG-BAT, 1.5 mg/L [2]; and SUVAVBT, 0.5 mg/L [3].

INTERFERENCES: None noted in the analytical method, but background levels of 0.032 – 0.070 μg/mL o-cresol have been 

observed [7-9]. These values are near and slightly above the limit of quantitation. Exposure to toluene will cause higher levels of 

urinary o-cresol as discussed above. p-Cresol is a normal component of human urine [10] and may, along with m-cresol, also be a 

co-exposure [11-12]. These two compounds have similar mass spectra to o-cresol but are chromatographically resolved under the 

conditions of this method [6].

OTHER METHODS: There are numerous literature methods for the determination of o-cresol in urine [10], but standardized 

methods from governmental agencies or consensus standards organizations are not currently available. NIOSH 8305 is a similar but 

only partially-validated method for phenol and p-cresol [11]. An older validated method using steam distillation and HPLC-UV 

detection for ~30 related compounds that includes o-cresol can be found in the German MAK collection [12].

REAGENTS:

1. o-Cresol* (≥99% purity) primary stock 

solution (PSS) about 10,000 μg/mL (0.100 g 

of o-cresol in 10 mL of methanol). Store at -

10 °C ± 2 °C

2. o-Cresol intermediate solutions of 1,000 and 

100 μg/mL (0.5 and 0.05 mL of PSS diluted 

to 5.0 mL with methanol). Store at -10 °C ± 

2 °C 

3. Human urine* provided by non-exposed 

individuals, store at < 6 °C**

4. Methanol*, GC grade or better

5. Hydrochloric acid*, concentrated, 33-38%, 

ACS reagent grade or better

6. Nitrobenzene (≥99% purity) internal 

standard (IS) solution of 500 μg/mL (0.125 g 

of nitrobenzene in 250 mL of methanol). 

Store at 10 °C ± 2 °C 

7. Water, ASTM Type II [13]

8. Methyl tert-butyl ether* (MTBE), HPLC grade 

or better

9. Sodium sulfate, granular, anhydrous, ≥99% 

purity. 

10. Helium, UHP or Grade 5

*See SPECIAL PRECAUTIONS.

**Human urine recommended due to 

evidence of column problems caused by 

synthetic urine.

EQUIPMENT:

1. GC-MS capable of selected ion monitoring 

with data system, autosampler, and column 

(page 8321-1)

2. Flasks, glass, volumetric: 5, 10 and 250 mL

3. Syringes, glass: 10, 50, 100, 500 and 2,500 μL

4. Tubes, centrifuge, 15-mL graduated 

polypropylene, with caps

5. Pipetter 1-10 mL, with disposable tips

6. Pasteur pipettes, glass

7. Water bath, 95 °C 

8. Wash bottle

9. Vials, autosampler, amber glass

10. Cold storage for -10 °C and 4 °C 

11. Bottles, 125 mL polyethylene

12. Analytical balance, to ±0.0001 g. 

 

SPECIAL PRECAUTIONS: Wear gloves, lab coat, and safety glasses while handling all chemicals and human 

urine products. Disposable plastic, glass, and paper (pipet tips, gloves, etc.) that contact urine should be 

placed in a biohazard container. Standard precautions should always be used when handling bodily fluids 

and/or extracts of bodily fluids [14]. Handle urine samples and urine extracts using proper gloves. All work 

should be performed in a fume hood. Methanol and MTBE are both flammable. Hydrochloric acid is an 

extremely corrosive chemical capable of severe tissue damage.
o-CRESOL in URINE: METHOD 8321, Issue 4, dated 18 March 2016 - Page 3 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLING:

1. Collect 50-100 mL urine in a 125 mL polyethylene bottle. Collect 2 urine specimens for each worker: 

one specimen before exposure and one specimen after.

2. Collect and pool urine specimens from unexposed workers to be used for controls.

3. Tightly cap each bottle and ship refrigerated or frozen in an insulated container to maintain the 

temperature at 6 °C or below.

SAMPLE PREPARATION:

4. Allow urine to reach room temperature and mix thoroughly.

NOTE: If desired, remove an aliquot of urine to determine creatinine levels.

5. Dispense a 5-mL aliquot of the urine specimen into a 15-mL graduated centrifuge tube.

6. Using a Pasteur pipette, add 1 mL of concentrated HCI, up to the 6 mL calibration mark in the tube.

7. Cap the tube. Shake vigorously for one minute.

8. Place the tube in a water bath at 95 °C for 1.5 hr. 

NOTE: Covering the water bath may be required to maintain the temperature at 95 °C.

9. Remove the tube from the bath and let it cool to room temperature.

10. Add 250 μL of the nitrobenzene IS solution.

11. Add water (ASTM Type II) to the tube, filling it to the 10 mL calibration mark.

12. Pipet 2 mL of MTBE to the tube and cap it.

13. Shake the tube vigorously for 2 min.

14. Allow the phases to separate.

15. Transfer the top organic phase to an amber GC vial containing approximately 0.2 g of anhydrous 

sodium sulfate (to ensure the dryness of the organic fraction).

16. Cap the vials and store at <6 °C until GC-MS analysis.

CALIBRATION AND QUALITY CONTROL:

17. Calibrate daily with nine working standards containing o-cresol at approximately the following 

concentrations: 0.04, 0.1, 0.2, 1, 2, 10, 20, 40 and 60 μg/mL.

a. Prepare the standards by adding measured amounts of o-cresol intermediate stock solutions to 

centrifuge tubes containing 5 mL of control urine.

b. Process the standards following the same procedure used for the samples (steps 4-16).

c. Prepare at least one method blank for every 20 samples by taking an aliquot of 5 mL control 

urine and processing it in the same manner as the samples (steps 4-16).

d. Prepare at least two levels of quality control (QC) samples using a separate source of o-cresol and 

prepare in the same manner as the calibration standards within the analytical range. These levels 

should be at roughly 10 times the limit of quantitation (LOQ) and 100 X LOQ. Analyze at least 

two QC samples of different concentration for every 20 samples.

18. Prepare a calibration graph by plotting, for each working standard, the normalized analyte response 

(peak area of analyte divided by the peak area of the internal standard on the same chromatogram) 

on the y-axis vs. μg of analyte/mL of urine on the x-axis. The simplest model that adequately 

describes the data should be used, but either a linear (most likely 1/X weighted because of the range 

of the calibration curve) or a quadratic model may be utilized in processing the analytical data. 

Because there may be detectable levels of o-cresol in the pooled urine blanks, before plotting the 

calibration graph, subtract the normalized analyte response of the pooled urine blank from the 

normalized analyte response of each working standard. The standard curve should have a coefficient 

of determination (r2

) of equal to or greater than 0.98 to be acceptable for use. Furthermore, when 

each standard is substituted back into the calibration equation, the value should be within ±20% of 

the expected value.

19. QC values should be within ±20% of the spiked values. If not, the batch is considered out of control, 

the batch data discarded, and corrective actions taken before more samples are analyzed.
o-CRESOL in URINE: METHOD 8321, Issue 4, dated 18 March 2016 - Page 4 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

MEASUREMENT AND CALCULATIONS: 

20. Set gas chromatograph according to manufacturer’s recommendations and to conditions given on 

page 8321-1.

21. Set the mass spectrometer according to manufacturer’s recommendations, to conditions given on 

page 8321-1, and to SIM for ions m/z 77, 107, 108, and 123.

22. Inject 2 μL of each sample, blank and QC sample.

23. Measure peak areas for o-cresol and nitrobenzene in the chromatograms. Divide the peak area of ocresol by the peak area of nitrobenzene in the same chromatogram.

CALCULATIONS:

24. Determine concentration of o-cresol from the calibration curve produced in step 18.

EVALUATION OF METHOD: 

This method was evaluated at five concentration levels over the range 0.0214 – 2.14 μg/mL. This range 

represents from 1 x LOQ to 100 x LOQ. Six replicates were analyzed at each level. The average recoveries at 

the various levels ranged from 86.8 to 118.4%. The limit of detection (LOD) and LOQ study was performed 

by analyzing a series of standards ranging from 0.0107 to 64.1 μg/mL, fitting the data to a quadratic curve, 

and estimating the values using “Burkart’s Method” [15]. The value obtained for the LOD by this method 

(0.009 μg/mL) was lower than the lowest standard in the determination (0.0107 μg/mL), so the LOD used 

for the method was the value of the lowest standard, 0.01 μg/mL. A long-term storage study was carried 

out at the 30 x LOQ level: pooled urine specimens spiked with o-cresol at 0.854 μg/mL were stored at 4 °C 

for 1, 7, 10, 14, 21 and 30 days and then analyzed. The recoveries across the entire study (1-30 days) were 

between 92-112%. Another set of specimens stored at room temperature were analyzed after 1 and 7 

days, producing recoveries ranging from 90-102% [6]. The analytical range of the method was extended 

during a secondary laboratory validation step. Two concentration levels were evaluated (5.06 and 30.30 

μg/mL) by analyzing five replicates at each level. These levels showed recoveries of 100.2 and 92.8% with 

relative standard deviations of 0.63 and 1.40% respectively [6].

In order to minimize confusion among users of NIOSH Method 8321, the authors would like to reemphasize that p-cresol is an endogenous human metabolite produced from protein breakdown and will 

always be found in human urine [10]. The mass spectrum of p-cresol is quite similar to that of o-cresol and 

so there is some possibility of peak misidentification. Fortunately, while the peaks are near each other in 

the chromatogram under the method conditions, the p-cresol peak elutes later, is adequately resolved, 

and thus will not interfere with o-cresol analyses [6].

Discussion concerning acid hydrolysis step: The efficiency of the acid hydrolysis step in this procedure was 

not evaluated. A common method of converting these conjugates (glucuronides and sulfates) back to the 

parent compounds is heating the sample with a mineral acid. Several literature examples that employ 

hydrochloric acid under very similar conditions can be found [8,9,12,16-18]. A common thread among 

these methods is that none of them appear to investigate the efficiency of the acid hydrolysis step. 

Fustinoni et al. did investigate the hydrolysis step both in terms of amount of acid used and reaction time 

required [19]. They found that 50 μL of concentrated HCl per 300 μL of urine and reacting for 60 min at 

100 °C gave yields of greater than 97%. NIOSH Method 8321 uses 1 mL concentrated HCl per 5 mL urine, 

heating at 95 °C for 1.5 h, which are nearly identical conditions. These similar conditions should assure 

adequate efficiency of the acid hydrolysis step and have proven to not be detrimental to the o-cresol [6].
o-CRESOL in URINE: METHOD 8321, Issue 4, dated 18 March 2016 - Page 5 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REFERENCES:

[1] ACGIH [2014]. TLVs and BEIs based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio:

American Conference of Governmental Industrial Hygienists. [www.acgih.org]. Date accessed: May

2014.

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area

[2014]. List of MAK and BAT values 2014; Maximum concentrations and biological tolerance values

at the workplace. DFG: Deutsche Forschungsgemeinschaft,

[http://onlinelibrary.wiley.com/book/10.1002/9783527682027]. Date accessed: January 2015.

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail.

Valeurs limites d’exposition aux postes de travail. 3rd Ed. [https://www.suva.ch/]. Date accessed:

May 2014.

[4] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA)

[http://www.dguv.de/ifa/GESTIS/]. Date accessed: May 2014.

[5] CRC Handbook of Chemistry and Physics [2013]. 94

th Edition, CRC Press, Taylor and Francis Group,

Boca Raton, FL.

[6] Siso MC, Perkins JB [2005]. Backup data report for o-cresol in urine method development.

DataChem Laboratories, Inc., under NIOSH contract CDC-200-2001-08000.

[7] Bieniek G [1997]. Urinary excretion of phenols as an indicator of occupational exposure in the cokeplant industry. Int Arch Occup Environ Health 70(5):334-340.

[8] Inoue O, Kanno E, Kasai K, Ukai H, Okamoto S, Ikeda M [2004]. Benzylmercapturic acid is superior to

hippuric acid and o-cresol as a urinary marker of occupational exposure to toluene. Toxicol Lett

147(2):177-186.

[9] Bahrami A, Jonidi-Jafari A, Folladi B, Mahjub H, Sadri Q, Zadeh MM [2005]. Comparison of urinary ocresol and hippuric acid in drivers, gasoline station workers and painters exposed to toluene in

west of Iran. Pak J Biol Sci 8(7):1001-1005.

[10] ATSDR [2008]. Toxicological profile for cresols. [https://www.atsdr.cdc.gov]. Date accessed: 

February 2015.

[11] NIOSH [1994]. Phenol and p-cresol in urine: Method 8305. In Eller PM, Cassinelli ME, eds. NIOSH

manual of analytical methods, 4th ed. Cincinnati, OH: U.S. Department of Health and Human

Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety

and Health, DHHS (NIOSH) Publication No. 94-113.

[12] Lewalter J, Schucht T [2012]. Benzene derivatives, suitable for steam distillation (phenol; m-/pcresol; o-cresol; o-chlorophenol; o-nitrophenol; p-chlorophenol; nitrobenzene; 1,2-dinitrobenzene;

2-chloro-5-methylphenol; 2,5-dichlorophenol; 3,4-dichlorophenol; 2,3-dinitrotoluene)

[Biomonitoring Methods, 1985]. The MAK Collection for Occupational Health and Safety, pp 31–56.

[13] ASTM [2011]. ASTM D1193-06(2011), Standard specification for reagent water, ASTM International,

West Conshohocken, PA, 2011, www.astm.org. 
o-CRESOL in URINE: METHOD 8321, Issue 4, dated 18 March 2016 - Page 6 of 6 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[14] CDC [2007]. 2007 Guidelines for isolation precautions: preventing transmission of infectious agents 

in healthcare settings. [http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html]. Date 

accessed: April 2013.

[15] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene 

chemistry laboratory, Appl Ind Hyg 1(3):153-155.

[16] Hasegawa K, Shiojima S, Koizmui A, Ikeda M [1983]. Hippuric acid and o-cresol in the workers of 

urine exposed to toluene. Int Arch Occup Environ Health 52(3):197-208.

[17] Truchon G, Tardif R, Brodeur J [1999]. o-Cresol: A good indicator of exposure to low levels of 

toluene. Appl Occup Environ Hyg 14(10):677-681.

[18] de Paiva MJN, Martins I, de Siqueira MEPB [2007]. Analysis of ortho-cresol in urine by solid phase 

microextraction-capillary gas chromatography. J Braz Chem Soc 18(5):1034-1039.

[19] Fustinoni S, Mercadante R, Campo L, Scibetta L, Valla C, Foà V [2005]. Determination of urinary 

ortho- and meta-cresol in humans by headspace SPME gas chromatography/mass spectrometry. J 

Chromatogr B 817:309-317.

METHOD WRITTEN BY:

María Carolina Siso and James B. Perkins, DataChem Laboratories, Inc., Salt Lake City, Utah under

NIOSH Contract CDC-200-2001-08000. Final editing by Dale A. Shoemaker, Ph.D., NIOSH.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TRICHLOROACETIC ACID IN URINE 8322

FORMULA: C2HCl3O2 MW: 163.39 CAS: 76-03-9 RTECS: AJ7875000

METHOD: 8322, Issue 1 EVALUATION: FULL Issue 1: 17 April 2015

OSHA & NIOSH: N/A

Other OELS: Because data on exposure limits and 

guidelines may change over time, NIOSH 

recommends referring to the following 

sources for updated limits. [1 - 4]

PROPERTIES: White solid; mp 59.1 °C; bp 198.2 °C; 

 d = 1.61 g/cm3

 [5]

BIOLOGICAL

INDICATOR OF: Exposure to trichloroethylene, trichloroacetic acid, methyl chloroform, tetrachloroethene, chloral hydrate

SYNONYMS: Trichloroethanoic acid, Aceto-caustin

SAMPLING

SPECIMEN: Urine

VOLUME: At least 10 mL

PRESERVATIVE: None

SHIPMENT: Freeze urine; ship in dry ice in an insulated 

container.

SAMPLE 

STABILITY: Stable at least 30 days at -17 oC [6]

CONTROLS: Urine specimens from matched population of 

non-exposed persons

ACCURACY

RANGE STUDIED: 0.9 to 100 μg/mL (as trichloroacetic acid)

BIAS: 0.0113 

OVERALL 

PRECISION (ŜrT): 0.03656

ACCURACY: 7.5%

MEASUREMENT

TECHNIQUE GAS CHROMATOGRAPHY with ELECTRON 

CAPTURE DETECTOR (GC-ECD)

ANALYTES: Trichloroacetic acid (determined as the 

methyl ester)

PROCEDURE: Methylation of an aliquot with BF3

·methanol 

to form the methyl ester followed by 

extraction into toluene

INJECTION: 1 μL, splitless for 0.5 min

TEMPERATURES

-INJECTION: 250 oC

-DETECTOR: 300 oC

-OVEN: 80 oC (hold for 0.5 min), 80 to 180 oC at 20 oC/

min, hold for 7 min

CARRIER GAS: Helium, ~3.5 mL/min

MAKEUP GAS: Nitrogen, 40 mL/min

COLUMN: Capillary, fused silica, 6% cyanopropylphenyl-94% dimethylpolysiloxane, 75 m x 

0.53 mm ID, 3 μm film

CALIBRATION: Trichloroacetic acid prepared in water to 

cover range and derivatized with the samples

ESTIMATED LOD: 0.08 μg/mL (as trichloroacetic acid) [6]

APPLICABILITY: This method can be used for the determination of trichloroacetic acid (TCAA) in urine specimens. TCAA is one of 

several metabolites detected after exposure to a variety of chlorinated compounds (representative compounds listed above) or 

from contaminated drinking water [7-11].

INTERFERENCES: None observed in the analytical method apart from some carryover issues (see Evaluation of Method section.) 

TCAA is a non-specific metabolite of several compounds. Urinary TCAA levels reflect exposure to any and all of these precursors. 

Background TCAA was detected in 76% of urine samples in a US general population sample, with a median concentration of 3.3 

μg/L, approximately 300 times lower than the range of this method [12].

OTHER METHODS: There are numerous literature methods for the determination of TCAA in urine [13]. The National Center for 

Environmental Health/Centers for Disease Control and Prevention (NCEH/CDC) has a method that is more expensive but also significantly more sensitive [11]. This method is based on the procedure used by O’Donnell [7] with some modifications.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

TRICHLOROACETIC ACID IN URINE: METHOD 8322, Issue 1, dated 17 April 2015 - Page 2 of 5

REAGENTS: 

1. Sodium trichloroacetate [CAS #650-51-1], 

97% purity or greater*

2. Boron trifluoride-methanol solution, 14%*

3. Toluene, ACS reagent grade or better*

4. Sodium sulfate, anhydrous, granular; reagent 

grade or better

5. Acetic acid, glacial; reagent grade or better* 

6. Acetone, reagent grade or better*

7. Methanol, reagent grade or better*

8. Helium, purified

9. Nitrogen, Ultra High Purity or P5

10. Water, ASTM Type II [14]

SOLUTIONS:

1. 1:3 glacial acetic acid:deionized water

2. 1:1 acetone:methanol

* See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Centrifuge tubes, polypropylene, ~15-mL, with 

screw caps, or other suitable container for 

specimen collection and storage

2. Gas chromatograph with electron capture 

detector, autosampler, data collection system and 

column (page 8322-1)

3. Microliter syringes, various sizes

4. Volumetric flasks, glass, various sizes

5. Adjustable pipettor with disposable plastic tips, 

0.1 to 1-mL

6. Disposable Pasteur transfer pipettes, 15 and 

23  cm

7. Culture tubes, 13 mm x 100 mm (~8 mL), with 

PTFE-lined caps

8. Vortex mixer

9. Glass wool

10. Vials, autosampler, glass, 2-mL with caps

11. Oven, capable of maintaining 60 oC

SPECIAL PRECAUTIONS: Standard precautions should always be used when handling bodily fluids and/or 

extracts of bodily fluids [15]. Handle urine specimens and urine extracts using powder-free latex or nitrile 

gloves. Acetic acid, acetone, toluene, and methanol are flammable; handle with care and use in a chemical 

fume hood. Handle all chemicals using the required safety precautions. Reagents with manufacturer 

expiration dates should be observed. The Environmental Protection Agency (EPA) has designated TCAA as a 

known mouse carcinogen and a possible human carcinogen [16].

SAMPLING:

1. Collect at least 10 mL of urine in ~15-mL polypropylene tubes or other suitable container. 

NOTE: Because of the relatively lengthy half-life values of TCAA, ACGIH recommends sampling at the end 

of shift at the end of workweek [1].

2. Freeze the urine and ship in dry ice in an insulated container. 

Reminder: Commercial shippers have special labeling requirements for packages containing 

 biological samples and dry ice.

SAMPLE PREPARATION:

3. Thaw urine specimens, bring to room temperature, and mix thoroughly.

4. Place 200 μL of urine specimen in an 8-mL glass culture tube with a PTFE-lined cap.

5. Add 0.5 mL 14% boron trifluoride in methanol; cap and mix.

6. Heat in oven at 60 oC for a minimum of 1.5 hr (maximum 2.5 hr).

7. Cool to room temperature and then add 2.0 mL toluene.

8. Vortex or shake vigorously for 1 min.

9. After the layers separate, transfer the upper toluene layer to a drying column containing anhydrous 

sodium sulfate. The drying columns are prepared in 15-cm Pasteur pipettes with a glass wool plug and 

about 200 to 300 mg anhydrous sodium sulfate, sufficient to form a bed depth of ~1 cm. 

10. Collect the eluate in a 2-mL GC vial. Cap vial.
TRICHLOROACETIC ACID IN URINE: METHOD 8322, Issue 1, dated 17 April 2015 - Page 3 of 5

CALIBRATION AND QUALITY CONTROL:

11. Prepare a stock solution by accurately weighing a known quantity of sodium trichloroacetate into a

volumetric flask. Add a known volume of deionized water and mix. Convert the weight of the sodium

trichloroacetate to TCAA by multiplying by 0.8814 (MW TCAA divided by MW sodium trichloroacetate

= 0.8814). As an example, 34 mg of sodium trichloroacetate into a 10-mL flask makes a 3 mg/mL stock

solution to be used in preparing the calibration standards.

NOTE: Sodium trichloroacetate was used instead of trichloroacetic acid for all phases of this method

development as well as in the preparation of standards. Trichloroacetic acid is very hygroscopic; the 

salt is much less so.

12. Prepare working (calibration) standards by serial dilution to cover the analytical range. A suggested

working standard concentration range is 0.08 to 300 μg/mL. Withdraw 200 μL of each calibration standard 

and follow steps 4 through 10.

13. Determine the retention time for the analyte of interest.

14. Prepare at least one blank urine specimen without an analyte spike to verify whether the source (of blank

urine) contained no detectable quantity of TCAA.

15. Prepare at least two levels of quality control spikes of TCAA, sodium salt to be analyzed with each analysis

batch. These levels should be at ~10 X the limit of quantitation (LOQ) and 200 X LOQ, but can be adjusted

to better suit the anticipated levels of the set of specimens. QC samples must be analyzed with every

batch such that they constitute 10% of the sample batch.

16. QC values must be within ±20% of the spiked values. If not, the batch is considered out of control, the

batch data discarded, and corrective actions taken before more samples are analyzed.

17. Calibrate daily with at least six liquid working standards covering the expected concentration range of the

samples.

MEASUREMENT:

PRECAUTION: SYRINGE-RINSE SOLUTIONS: Toluene will extract material from some urine 

specimens that may eventually clog the syringe and cause injection errors unless the syringe is 

rinsed with the following solutions following each injection.

First rinse solution: 1:3 glacial acetic acid:deionized water

Second rinse solution: 1:1 acetone:methanol

18. Set the gas chromatograph according to manufacturer’s recommendations and to conditions given on

page 8322-1. With the chromatographic conditions listed, the retention time of the methyl ester of TCAA 

was 8.87 min [6].

19. Inject each of the samples, standards, blanks, and quality control samples.

20. Measure peak area or peak height; peak area is recommended.

NOTE: If the sample peak area or height is greater than that of the highest calibration standard, dilute with

toluene and reanalyze. Apply the appropriate dilution factor in the calculations.

21. Prepare a calibration curve by plotting instrument responses (usually peak area) for the standards vs.

concentration. The simplest model that adequately describes the data should be used, but either a linear 

(mostly likely 1/x weighted because of the range of the calibration curve) or a quadratic model may be 

utilized in processing the analytical results. The standard curve must have a coefficient of determination 

(r2

) of equal to or greater than 0.98 to be acceptable for use. Furthermore, when each standard is plugged 

back into the calibration equation, the measured value must be within ±20% of the expected value.

CALCULATION OF ANALYTE PER SAMPLE:

22. Determine the concentration of TCAA in μg/mL (mg/L) using the response of each sample and the

calibration curve prepared in step 21. Apply any dilution factor if applicable.

NOTE: If the creatinine value is available, the concentration may be reported as μg/g creatinine if desired.
TRICHLOROACETIC ACID IN URINE: METHOD 8322, Issue 1, dated 17 April 2015 - Page 4 of 5

EVALUATION OF METHOD: 

This method was evaluated over a range of 0.9-30 μg/mL. This range covers 3x, 10x, 30x, and 100x of 

the estimated LOQ. Six replicates were prepared and analyzed at each concentration level. The average 

recoveries for each of the concentration levels were 94.8% (3 x LOQ), 102.3% (10 x LOQ), 110.4% (30 x LOQ), 

and 97.1% (100 x LOQ). Recoveries were determined by comparison against spiked and derivatized liquid 

standards (standards prepared in deionized water). The upper concentration range was extended to 100 

μg/mL during testing by an independent laboratory. Five samples were analyzed at this concentration and 

the average recovery was 105.0%. Overall accuracy was calculated to be 7.5%; bias was 0.0113, and overall 

precision was 0.0366 [6]. The limit of detection (LOD) and LOQ were determined by analyzing a series of 

derivatized spiked standards, with the data fitted to a quadratic curve, then estimated according to the 

Burkart method [17]. A long-term storage stability study was carried out at the 10x, 30x, and 100x LOQ levels. 

Urine samples were spiked with trichloroacetate and stored at -17 oC for 7, 14, 21, 30, and 46 days and then 

analyzed. Recoveries at 30 and 46 days were all greater than 90% [6].

During the testing performed by the independent laboratory (“User Check”), broad, interfering carryover 

peaks from the urine matrix were noticed. The lab found that raising the final temperature of the GC program 

to 240 oC (instead of 180 oC) and adding a longer hold time 10 min (instead of 7 min) reduced the carryover 

problem, allowing more precise and accurate measurement of the peak of interest. This adjusted GC program 

would now be: 80 oC for 0.5 min, heat to 240 oC at 20 oC/min, and hold for 10 min. Either set of conditions 

may be used.

NOTE: While the overall accuracy and precision for the User-Check samples were within acceptable 

limits [6], there were spurious results in 10% of the samples (2 out of 20.) No reason is known 

for these outliers, nor were the samples able to be re-injected, re-extracted, or re-analyzed. To 

improve user confidence in the results obtained by this method, it is suggested to randomly run 

duplicate analyses of 10-20% of the samples and to randomly re-inject 10-20% of the samples. If 

the method is used in an on-going manner and no problems or spurious results are noted, this 

recommendation could be lowered or eliminated.

REFERENCES:

[1] ACGIH [2014]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American Conference

of Governmental Industrial Hygienists. [www.acgih.org] Date accessed May 2014.

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area

[2014]. List of MAK and BAT values 2014; Maximum concentrations and biological tolerance

values at the workplace. DFG: Deutsche Forschungsgemeinschaft. [http://onlinelibrary.wiley.com/

book/10.1002/9783527682027] Date accessed January 2015.

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail. 

Valeurs limites d’exposition aux postes de travail. 3rd Ed. [https://www.suva.ch/] Date accessed 

May 2014.

[4] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [IFA) http://www.dguv.de/ifa/

GESTIS] Date accessed January 2015.

[5] CRC Handbook of Chemistry and Physics [2013]. 94th Edition, CRC Press, Taylor and Francis Group, Boca

Raton, FL.

[6] Wickman DC, Perkins JB, Reynolds JM [2012]. Backup data report for trichloroacetic acid in urine, ALS

Environmental (formerly DataChem Laboratories Inc.) under NIOSH contract CDC-200-2001-08000. 
TRICHLOROACETIC ACID IN URINE: METHOD 8322, Issue 1, dated 17 April 2015 - Page 5 of 5

[7] O’Donnell GE, Juska A, Geyer R, Faiz M, Stalder S [1995]. Analysis of trichloroacetic acid in the urine of 

workers occupationally exposed to trichloroethylene by capillary gas chromatography. J Chromatogr A 

709:313-317.

[8] Bernauer U, Birner G, Dekant W, Henschler D [1996]. Biotransformation of trichloroethylene: Dosedependent excretion of 2,2,2-trichloro-metabolites and mercapturic acids in rats and humans after 

inhalation. Arch Toxicol 70:338-346.

[9] Raaschou-Nielsen O, Hansen J, Christensen JM, Blot WJ, McLaughlin JK, Olsen JH [2001]. Urinary 

concentrations of trichloroacetic acid in Danish workers exposed to trichloroethylene, 1947-1985. Am J 

Ind Med. 39:320-327.

[10] Vartiainen T, Pukkala E, Rienoja T, Strandman T, Kaksonen K [1993]. Population exposure to triand tetrachloroethylene and cancer risk: Two cases of drinking water pollution. Chemosphere, 

27(7):1171-1181.

[11] Kuklenyik Z, Ashley DL, Calafat AM [2002]. Quantitative detection of trichloroacetic acid in human urine 

using isotope dilution high-performance liquid chromatography-electrospray ionization tandem mass 

spectrometry. Anal Chem 74(9):2058-2063.

[12] Calafat AM, Kuklenyik Z, Caudill SP, Ashley DL [2003]. Urinary levels of trichloroacetic acid, a disinfection 

by-product in chlorinated drinking water, in a human reference population. Environ Health Perspect 

111:151-154.

[13] Delinksy AD, Bruckner JV, Bartlett MG [2005]. A review of analytical methods for the determination of 

trichloroethylene and its major metabolites chloral hydrate, trichloroacetic acid and dichloroacetic acid. 

Biomed Chromatogr 19:617-639.

[14] ASTM [2011]. ASTM D1193, Standard specification for reagent water. ASTM International, West 

Conshohocken, PA. [www.astm.org] Date accessed: April 2013. DOI 10.1520/D1193-06R11.

[15] CDC [2007]. 2007 Guidelines for isolation precautions: Preventing transmission of infectious agents in 

healthcare settings. [http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html] Date accessed: 

April 2013.

[16] US EPA [2011]. Trichloroacetic acid (CASRN 76-03-9). United States Environmental Protection Agency. 

[www.epa.gov/iris/subst/0655.htm] date accessed April 7, 2014.

[17] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene 

chemistry laboratory. Appl Ind Hyg 1(3):153-155.

METHOD WRITTEN BY: Don C. Wickman and James B. Perkins; DataChem Laboratories, Inc., Salt Lake City, 

Utah, under contract CDC 200-2001-08000.

METHOD EDITED BY: John M. Reynolds, ALS Environmental (formerly DataChem Laboratories, Inc.), Salt 

Lake City, Utah and Dale A. Shoemaker, Ph.D., NIOSH/DART/CEMB.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE 8324

metabolite of 1-bromopropane

FORMULA: C3

H5BrO2 MW: 152.97 CAS: 590-92-1 RTECS: UE7875000

METHOD: 8324, Issue 1 EVALUATION: FULL Issue 1: 12 September 2014

Exposure limits and guidelines:

 OSHA: None

NIOSH: None

Other OELs: Because data on exposure limits and guidelines 

may change over time, NIOSH recommends referring 

to the following sources for updated limits [1 - 4].

PROPERTIES: Solid; d 1.48 g/mL at 25 o

C; MP 58-62 oC

BIOLOGICAL 

INDICATOR OF: Exposure to 1-bromopropane

SYNONYMS: β-Bromopropionic acid; 2-carboxyethylbromide.

SAMPLING 

SPECIMEN: Urine 

VOLUME: At least 15 mL 

PRESERVATIVE: None added

 Refrigerate or freeze upon collection

SHIPMENT: Ship cold or frozen with ice or dry ice

Freeze upon receipt at the laboratory

SAMPLE

STABILITY: Stable in frozen urine for at least 30 days

CONTROLS: Urine specimens obtained from non-exposed 

individuals

ACCURACY

RANGE 

STUDIED: Table 1

BIAS: None established

PRECISION: Table 1

RECOVERY

ACCURACY: Full recovery (95% overall) was established by 

a spiked urine recovery, overall RSD was +3.1% 

(Table 1).

The definitions of precision and accuracy in this method are 

those utilized by the US Food and Drug Administration [5].

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY, MASS SPECTROMETRY 

with SELECTED ION MONITORING (GC/

MS-SIM)

ANALYTE: tert-butyldimethylsilane derivative of 

3-bromopropionic acid

EXTRACTION: Liquid-liquid extraction (LLE) 

INJECTION 

VOLUME: 0.5 μL, splitless

TEMPERATURES 

 - INJECTION: 200 o

C 

 - COLUMN: 60 o

C initial, 4 oC/min to 180 oC,15 oC/min to 

255 o

C, post-run at 270 oC for 5 min

 - DETECTOR: Source 230 o

C, Quadrupole 150 oC; Solvent 

delay, 15 min

RUN TIME: Approximately 50 minute cycle time

CARRIER GAS: Helium, 0.8 mL/min constant flow

COLUMN: Capillary, fused silica, 100% dimethyl-polysiloxane, 

50 m X 0.20 mm (ID), 0.33 μm film thickness

CALIBRATION: Analyte in control urine; with internal 

standard

QUALITY 

CONTROL: At least one level of spiked urine sample prepared 

from a separately weighed stock solution

RANGE: 2.0 to 100 μg/mL

ESTIMATED LOD: Approximately 0.01 μg/mL (IUPAC) [6]. 0.1 

μg/mL is the lowest calibration standard. 

APPLICABILITY: 3-Bromopropionic acid (3-BPA) has been reported to be a rat metabolite [7] and is a potential human biomarker 

for exposure to 1-bromopropane. This method measures the quantity of free 3-BPA in urine. 1-Bromopropane is used as an industrial solvent. In one limited study, 3-BPA was not detected in individuals exposed to low levels of 1-bromopropane [8].

INTERFERENCES: None found or identified.

OTHER METHODS: This method is from the one described by B’Hymer and Cheever [9] and further investigated by Mathias, et al. 

[8].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 2 of 9

REAGENTS:

1. 3-Bromopropionic acid (3-BPA) reference standard 

as a 1.0 mg/mL stock solution in deionized water. 

Store in a refrigerator.

2. 3-Chloropropionic acid (3-CPA) reference 

standard, 20 μg/mL in deionized water, internal 

standard solution. Store in a refrigerator.

3. Ethyl acetate, HPLC grade or better*

4. N-Methyl-N-(tert-butyldimethylsilyl)

trifluoroacetamide (MTBSTFA) with 1% tertbutyldimethylchlorosilane (TBDMCS)

5. Magnesium sulfate, anhydrous, powdered, ACS 

reagent grade

6. Hydrochloric acid, concentrated, ACS reagent 

grade

7. Water, deionized (ASTM type II)

8. Nitrogen, prepurified grade or better

9. Urine, non-exposed*

* See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Gas chromatograph with mass spectrometric 

detector, column, autosampler, and data collector 

(page 8324-1)

2. Bottles, polypropylene, 125-mL with caps

3. Analytical balance, to ±0.0001 g

4. Analytical evaporator with nitrogen gas sweep

5. Test-tube vortex mixer

6. Heating block or oven

7. Automatic pipettor with disposable tips

8. Repeating pipet dispenser, 1 Liter

9. Tubes, disposable screw-top culture (16 X 100 mm 

and 16 X 150 mm), with polytetrafluoroethylene 

(PFTE)-lined caps

10. Metal spatula

11. Flasks, volumetric; 10, 50 and 100-mL

12. Autosampler vials, 2 mL, silanized, with caps and 

septa

13. Glass funnels

14. Glass wool, silanized

15. Disposable glass pipets

16. Dry ice or bagged refrigerant

SPECIAL PRECAUTIONS: Standard precautions should always be used when handling bodily fluids and/

or extracts of bodily fluids [10]. Handle urine samples and urine extracts using powder-free latex or nitrile 

gloves. Ethyl acetate and MTBSTFA are flammable; handle with care and use in a chemical fume hood. 

Handle all chemicals using the required safety precautions. Manufacturer expiration dates for reagents 

should be observed.

SAMPLING:

1. Collect at least 15 mL urine in an appropriate polypropylene tube or bottle and cap. Refrigerate or freeze 

immediately after collection. Collect at least two urine specimens for each worker: one before the work 

shift and one after.

2. Ship the specimens stored in either wet or dry ice in an insulated container. Store frozen upon arrival 

at the laboratory. A reminder: commercial shippers have special labeling requirements for packages 

containing dry ice.

SAMPLE PREPARATION:

3. Thaw the urine specimen to room temperature.

4. Mix thoroughly to ensure urine homogeneity.

5. Transfer 2.0 mL of urine into a 16 X 100 mm (or larger) screw-capped culture tube. 

6. Acidify by adding 40 μL of concentrated hydrochloric acid.

7. Add 0.5 mL of deionized water.

8. Add 0.5 mL of the 20 μg/mL 3-CPA internal standard solution.

9. Ethyl acetate extraction: Dispense 4 mL of ethyl acetate into the culture tube, cap, and vortex (or 

vigorously shake) for 1 minute.

10. Allow layers to separate, collect the ethyl acetate (top) layer.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 3 of 9

11. Repeat the extraction (steps 9 and 10) three more times. Collect and combine all ethyl acetate 

extracts using a glass pipet into a 16 X 150 mm culture tube.

12. Dry the ethyl acetate extract by adding approximately 100 to 200 mg of anhydrous magnesium 

sulfate and swirl for about 15 seconds.

13. Transfer the extract solution into a 16 X 150 mm culture tube by means of a glass funnel with 

silanized glass wool patch to remove the wet magnesium sulfate. The glass wool patch must be 

packed tightly enough to prevent particles of magnesium sulfate from passing through.

14. Rinse the tube and the funnel with ethyl acetate to ensure complete transfer.

15. Concentrate the combined ethyl acetate extract for each sample to 1 mL using a nitrogen sweep at 

room temperature and transfer the solution to a 2-mL GC autosampler vial.

16. tert-Butyldimethylsilane derivatization: Add 50 μL of MTBSTFA with 1% TBDMCS silanizing reagent 

to each autosampler vial and cap immediately.

17. Heat the solution for 1.5 hours at 70 o

C in a heating block or oven.

CALIBRATION AND QUALITY CONTROL:

18. 3-Bromopropionic acid (3-BPA) standards are prepared in blank, non-exposed urine. The 1 mg/mL 

stock 3-BPA solution is diluted in deionized water to make 0.4, 1, 2, 4, 8, 20, 80, 200, 400, 600, and 

800 μg/mL 3-BPA solutions for spiking.

19. Transfer 2.0 mL of non-exposed urine into a 16 X 100 mm (or larger) screw-capped culture tube. 

20. Acidify by adding 40 μL of concentrated hydrochloric acid.

21. Add 0.5 mL of the 20 μg/mL 3-CPA internal standard solution.

22. Add 0.5 mL of the appropriate 3-BPA spiking solution described in step 18 to make urine samples 

equivalent to 0.1, 0.25, 0.5, 1, 2, 5, 20, 50,100, 150 and 200 μg/mL of 3-BPA in the original 2.0 mL 

volume of urine.

23. Prepare at least one blank urine without a 3-BPA spike to verify the source of blank urine contains no 

detectable quantity of 3-BPA.

24. Prepare at least two levels of quality control (QC) standard of 3-BPA fortified urine using a separately 

weighed and prepared 3-BPA stock solution. One level should be within the lower 25% of the 

calibration curve and one level within the upper 25% of the calibration curve. More than two QC 

levels can be used. QC samples should be analyzed with every batch such that they constitute at 

least 5% of the sample batch.

25. QC values should be within ±20% of the spiked values. If not, the batch is considered out of control, 

the batch data discarded, and corrective actions should be taken before more samples are analyzed. 

26. Ethyl acetate extraction: Prepare the spiked standard urine samples, the blank urine sample(s), and 

the QC standards the same as described in the preceding Sample Preparation section using Steps 9 

through 17.

MEASUREMENT:

27. Set the gas chromatograph according to the manufacturer’s recommendations and to the 

conditions listed on page 8324-1.

28. Set the mass selective detector to selected ion monitoring mode for ions m/z 211 (derivative of 

3-BPA) and 165 (derivative of 3-CPA). 

NOTE: The use/non-use of qualifier ions for this method is discussed in the literature [9].

29. Inject 0.5 μL of each sample, standard, blank, and QC standard extract from Steps 17 and 26.

30. Measure the peak areas of the tert-butyldimethylsilane derivatives of 3-BPA and 3-CPA in the 

chromatograms of the standards.

31. Divide the peak area of the derivative of 3-BPA by the peak area from the derivative of 3-CPA in the 

same chromatogram.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 4 of 9

32. Prepare a linear calibration curve of the peak Area Std./Area Int. Std. versus the concentration of the 

standard for 3-BPA.

33. Measure the peak areas of the derivatives of 3-BPA and 3-CPA in the remainder of the 

chromatograms.

34. Divide the peak area of the derivative of 3-BPA by the peak area from the derivative of 3-CPA in the 

same chromatogram.

CALCULATIONS:

35. Determine the concentration of 3-BPA in the extracts from the original urine (2.0 mL specimen) from 

the curve obtained in step 32. The results are expressed as μg/mL of 3-BPA in urine.

EVALUATION OF METHOD:

This method was evaluated and described by B’Hymer [6] and in detail by B’Hymer and Cheever [9]. A 

general summary of this published information is given below:

Accuracy and Precision: Two recovery studies using multiple GC columns over several days 

demonstrated the accuracy and precision of this test method. The first recovery study was performed 

over three separate experimental batch runs, and these data are presented in Table 1A. Average 

recovery was between 93 and 98% for the three 3-BPA spiked level urine samples investigated. For each 

batch run, the experimental trial consisted of three samples at three different concentration levels. The 

recovery for each level (n=9 samples) is displayed in Table 1A. The second recovery study (data shown 

in Table 1B) used spiked urine samples from 20 non-exposed volunteers and demonstrated that the 

procedure was accurate (95% average recovery) and precise (2.5% relative standard deviation.) No 

interferences were detected in the unspiked urine from the 20 volunteer specimens. Both recovery 

studies generated a total of 47 spiked urine samples at 2, 10, 20 and 50 μg/mL 3-BPA levels. Precision 

expressed as percent relative deviation (% RSD) was as high as 5.7% on the 2 μg/mL recovery samples 

(n = 9). Overall recovery was 95% and overall RSD was 3.1% (n = 47).

Linearity: All calibration curves used during the development of this method were linear and had 

correlation coefficients of 0.98 and greater. The concentration range was 0.1 to 200 μg/mL 3-BPA in 

urine with 2.0 mL urine sample size. Calibration curves were run at the beginning and end of all sample 

batch runs; calibration curve slope drift was found to be acceptable.

Specificity: The optimized chromatographic conditions developed for this procedure proved to be 

specific and have no major interferences. The mass spectrometric detector was useful in adding 

additional specificity to the method. The ion m/z 211 was chosen for monitoring the calibration curve 

used in the calculations because of its greater abundance, and it was a characteristic fragment for 

the tert-butyldimethylsilane (TBDMS) derivative of 3-BPA. This is the molecular ion less the tert-butyl 

group, m/z 57 (Figure 1). Ion m/z 165 was used to monitor the TBDMS derivative of 3-CPA, the internal 

standard, for the same reasons. Full-scan mass spectra of the TBDMS derivatives are presented in 

Figure 2. An example chromatogram for blank urine and urine spiked with 3-BPA and 3-CPA is shown in 

Figure 3.

Robustness: Multiple HP-1 (100% dimethylpolysiloxane) columns of different manufacturing lots were 

used during the recovery studies. Accuracy and precision did not appear to be affected; therefore, 

the method appears to be reproducible with any normal functioning HP-1 capillary column. Recovery 

results from individual urine samples spiked with 3-BPA indicate that the method is accurate and not 

significantly affected by individual urine sample matrix differences during analyte extraction.

Stability: While sample stability was not exhaustively evaluated, an aqueous stock standard solution of 

3-BPA stored for two weeks at 4 o

C gave full recovery assay values when compared to a freshly prepared 

3-BPA standard. The 3-BPA in the urine specimens appears to be stable in frozen urine for a much 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 5 of 9

longer time frame of two months or more. Derivatized sample extracts appeared to be stable during a 

one week time frame.

Range: This method should be considered accurate for the estimation of 3-BPA in human urine within 

the 2.0 to 100 μg/mL method validation range.

REFERENCES:

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio:

American Conference of Governmental Industrial Hygienists. [www.acgih.org] Date accessed: April

2014.

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area

[2013]. List of MAK and BAT values 2013; Maximum Concentrations and Biological Tolerance Values

at the Workplace. DFG: Deutsche Forschungsgemeinschaft, Wiley-VCH Verlag GmbH & Co. [http://

www.dfg.de/en/dfg_profile/publications/senate_commissions/index.html#micro7535782]. Date

accessed April 2014.

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail.

Valeurs limtes d’exposition aux postes de travail. [https://www.suva.ch/]. Date accessed: April 2014.

[4] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA) [http://www.dguv.

de/ifa/GESTIS]. Date accessed: April 2014.

[5] US FDA [2001]. Guidance for industry: Bioanalytical method validation. U.S. Food and Drug

Administration, Washington, DC. [http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.

pdf]. Date accessed April 2014.

[6] B’Hymer C [2004]. 3-Bromopropionic acid in urine, Method 8324. Backup data report.

[7] Jones AR, Walsh DA [1979]. The oxidative metabolism of 1-bromopropane in the rat. Xenobiotica

9(12): 763-772.

[8] Mathias PI, Cheever KL, Hanley KW, Marlow KL, Johnson BC, B’Hymer C [2012]. Comparison

and evaluation of urinary biomarkers for occupational exposure to spray adhesives containing

1-bromopropane. Toxicol Mech Methods 22(7): 526-532.

[9] B’Hymer C, Cheever KL [2004]. Development of a gas chromatographic test for the quantification of

the biomarker 3-bromopropionic acid in human urine. J Chromatogr B 802: 361-366.

[10] CDC [2007]. 2007 Guidelines for isolation precautions: preventing transmission of infectious agents

in healthcare settings. [http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html]. Date

accessed: April 2013. 

METHOD WRITTEN BY:

Clayton B’Hymer, PhD, NIOSH/DART/BHAB. Final editing performed by Dale A. Shoemaker, PhD, NIOSH/

DART/CEMB.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs 

or products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced 

in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 6 of 9

Figure 1. The major ions monitored for the tert-butyldimethylsilane derivatives were the molecular 

ions less the tert-butyl group (m/z = 57). The ions used for quantitation are m/z (mass to charge) 165 

for the internal standard derivative and m/z 211 for the 3-bromopropionic acid derivative.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 7 of 9

Figure 2a.

Figure 2b.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 8 of 9

Figure 3. A total ion chromatogram of a (A) blank non-exposed volunteer urine specimen and (B) 

20 μg/mL spiked 3-BPA urine solution with 5 μg/mL equivalent 3-CPA as the internal standard. No 

interfering peaks were evident in any of the group of 20 non-exposed volunteer specimens.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

3-BROMOPROPIONIC ACID in URINE: METHOD 8324, Issue 1, dated 12 September 2014 - Page 9 of 9

Table 1. Recovery studies of 3-bromopropionic acid:

 (A) multilevel recovery study of 3-BPA from spiked urine samples1,

 (B) recovery of 20 μg/mL 3-BPA spikes from individual urines of 20 non-exposed volunteers2

 Note: Overall recovery of all samples was 95% and overall RSD was 3.1% (n = 47.)

(A)

Spike level 

(μg/mL)

Mean 3-BPA 

recovered,

(n = 9, μg/mL)

Average % 

Recovery

Standard 

Deviation 

(μg/mL)

% Relative 

Standard 

Deviation

2 1.91 96 0.11 5.7

10 9.32 93 0.13 1.4

50 48.9 98 0.36 0.7

(B)

Volunteer 

urine Spike 

level

(μg/mL)

Mean 3-BPA 

recovered 

(n = 20, μg/mL)

Average % 

Recovery

Standard 

Deviation

(μg/mL)

% Relative 

Standard 

Deviation

20 19.0 95 0.48 2.5

1

 Three different spiked urine samples were prepared at each level and chromatographed on three 

separate experimental trial runs (a total of nine samples at each spike level were analyzed.)

2

 All non-spiked specimens showed no 3-BPA derivative peak in the chromatograms.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine 8326

Metabolites of toluene and benzene

1. S-Benzylmercapturic acid: FORMULA: C12H15NO3S MW: 253.3 CAS: 19542-77-9

2. S-Phenylmercapturic acid: FORMULA: C11H13NO3

S MW: 239.3 CAS: 20640-68-0

METHOD: 8326, Issue 1 EVALUATION: Full Issue 1: 20 May 2014

Published limits and guidelines using these compounds as markers: 

1. Toluene: OSHA and NIOSH: None

2. Benzene: OSHA and NIOSH: None

Other OELs: Because data on exposure limits and guidelines may change 

over time, NIOSH recommends referring to the following sources for 

updated limits and guidelines concerning the use of these compounds as 

markers for toluene and benzene [1- 4].

PROPERTIES: 1. Solid; density =1.246 g/cm3; mp 162-163 °C 

2. Solid; density = 1.28 g/cm3

; mp 155 °C

BIOLOGICAL 

INDICATOR OF: Exposure to 1. toluene and 

 2. benzene

SYNONYMS (not all inclusive): 

1. S-Benzylmercapturic Acid (BMA): S-benzyl-N-acetyl-L-cysteine; (2R)-2-Acetamido-3-(phenylmethylsulfanyl)propanoic acid; Alanine, N-acetyl-3-

(benzylthio)-; S-phenylmethyl-N-acetyl-L-cysteine

2. S-Phenylmercapturic acid (PMA): S-phenyl-N-acetyl-L-cysteine; (2R)-2-Acetamido-3-(phenylsulfanyl)propanoic acid; (2R)-2-Acetylamino-3-

(phenylthio)propionic acid

SAMPLING

SPECIMEN: Urine 

VOLUME: At least 8 mL 

PRESERVATIVE: None added. Refrigerate or freeze upon 

 collection.

SHIPMENT: Ship cold or frozen with ice or dry ice. Freeze upon 

receipt at the laboratory. 

SAMPLE

STABILITY: Stable in frozen urine for periods of a month or more 

and for several freeze/thaw cycles [5,6].

CONTROLS: Urine specimens obtained from non-exposed or low 

level exposed individuals. 

ACCURACY*

RANGE 

STUDIED: See Table 2

BIAS: Negligible

OVERALL

 PRECISION (ŜrT)*: See Table 2

ACCURACY*: Overall recoveries obtained from 

spiked urine samples (n=48) were 103% 

and 106% for S-benzylmercapturic 

acid and S-phenylmercapturic acid, 

respectively. The precision as relative 

standard deviation was no greater than 

5.0% at any concentration level (n=9, 

Table 2). 

* The definitions of precision and accuracy in this method are 

those utilized by the US Food and Drug Administration [7].

MEASUREMENT

TECHNIQUE: HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY - 

TANDEM MASS SPECTROMETRY (HPLC/MS/MS)

ANALYTES: S-Benzylmercapturic acid and S-phenylmercapturic 

acid

EXTRACTION: Solid-Phase Extraction (SPE) C18 

COLUMN: C18 [dimethyloctadecylsilane solid phase type, 3.5 

μm particle size] (150 mm by 3 mm)

MOBILE

PHASES: A = 5/95/0.1% (v/v/v) acetonitrile/water/acetic acid

B = 75/25/0.1% (v/v/v) acetonitrile/water/acetic acid

FLOW RATE: 0.3 mL/min (0.4 mL/min post run)

GRADIENT: Time (min) vs. Mobile Phase Composition

0 to 10 0 to 40% B

10 to 18 40 to 100% B

18 to 20 100% B

20 to 21 100% B Flow increased to 0.4 mL/min

21 to 28 100% B (0.4 mL/min flow)

28 to 30 100 to 0% B (re-equilibration, 0.3 mL/min)

30 to 37 0% B (re-equilibration)

INJECTION VOLUME: 8 μL

IONIZATION

SOURCE: Electrospray at 3500 Volts and negative scan mode, 

nebulizer gas at 35 psi and 10 L/min flow

DETECTOR (MS/MS): Dwell time = 200 msec; Fragmentor at 80 Volts; 

 Collision energy at 8 Volts; Collision gas: nitrogen at 

 0.06 L/min

MULTIPLE REACTION MODE: Quantification mass transitions; 

 BMA = m/z 252 → 123, PMA = 238 → 109, 

d5-BMA = 257 → 128, d5-PMA = 243 → 114 

TOTAL RUN TIME: Approximately 37 minute cycle time

CALIBRATION: BMA and PMA solutions with internal standards

QUALITY CONTROL: At least one level of spiked urine specimen 

prepared from a separately weighed stock solution

RANGE: 0.5 to 50 ng/mL for BMA and PMA

ESTIMATED LOD: Approximately 0.2 ng/mL for BMA and PMA; by lowest 

standard levels, BMA and PMA = 0.5 ng/mL

PRECISION ( ): See Table 2
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 2 of 10

APPLICABILITY: BMA and PMA are metabolites of toluene and benzene, respectively. PMA is a very specific bioe. BMA can form from exposure to other sources, such as benzyl acetate or benzyl alcohol which 

nal care products [8]. Both toluene and benzene are common solvents with multiple occupare exposure to toluene and benzene (less commonly) can occur from environmental and other 

 measures the quantity of the two target metabolite analytes in urine.

marker for benzen

can be found in perso

tional uses; furthermo

sources. This method

INTERFERENCES: None found or identified.

OTHER METHODS: This method is based on the research of B’Hymer [9,10]. There are numerous literature methods for 

these two compounds, but standardized methods from governmental agencies or consensus standards organizations 

are not currently available, though there is also a recent method developed at CDC/National Center for Environmental 

Health for these analytes and others [11].

REAGENTS:

1. S-Benzylmercapturic acid (BMA, N-acetyl-S-benzyl-DLcysteine, CAS no. 19542-77-9) reference standard 0.20 

mg/mL stock solution in methanol. Store at 4 °C in 

amber vials (or in the dark).

2. S-Phenylmercapturic acid (PMA, phenyl-N-acetyl-DLcysteine, CAS no. 20640-68-0) reference standard 0.20 

mg/mL stock solution in methanol. Store at 4 °C in 

amber vials (or in the dark).

3. Deuterated S-benzyl-d5

-mercapturic acid reference 

standard (d5

-BMA), 0.15 mg/mL stock solution in 

methanol. Store at 4 °C in amber vials (or in the dark). 

4. Deuterated S-phenyl-d5

-mercapturic acid reference 

standard (d5

-PMA), 0.15 mg/mL stock solution in 

methanol. Store at 4 °C in amber vials (or in the dark).

5. Internal standard spiking solution, deuterated stock 

solutions diluted to approximately 30 ng/mL in water

6. Acetone, HPLC grade or better*

7. Acetonitrile, HPLC grade or better*

8. Acetic Acid, glacial, ACS reagent grade or better*

9. Methanol, HPLC grade or better*

10. Water, doubly deionized, minimum resistivity of 18 

MΩ-cm

11. Water, HPLC grade

12. Acetonitrile/water (50/50%, v/v) injector rinse solution

13. Chromatographic matrix adjustment solution, 49/50/1% 

(v/v/v) acetonitrile/water/acetic acid

14. Mobile phase A (5/95/0.1%, v/v/v) acetonitrile/water/

acetic acid, filtered through 0.7 μm glass microfiber 

filters

15. Mobile phase B (75/25/0.1%, v/v/v) acetonitrile/water/

acetic acid, filtered through 0.7 μm glass microfiber 

filters

16. Synthetic urine substitute, if desired

17. Nitrogen, UHP

* See SPECIAL PRECAUTIONS*

EQUIPMENT:

1. High-Performance Liquid Chromatograph (HPLC) 

equipped with a tandem mass spectrometric detector 

with data collection system, page 8326-1

2. HPLC column (C18 [dimethyloctadecylsilane solid phase 

type], 150 mm X 3 mm, 3.5 μm particle size)

3. Autosampler

4. Analytical balance, 0.1 mg readability

5. Rotary vacuum concentrator with cold trap and vacuum 

pump or analytical evaporator with nitrogen

6. Automatic pipet with disposable tips in the 1000 μL 

volume delivery range

7. Solid-phase extraction (SPE) cartridges, C18, 500 mg, 3 

mL

8. SPE vacuum manifold apparatus

9. Culture tubes, disposable screw-top (16 X 150 mm) with 

PTFE-lined caps

10. Tubes, polypropylene, disposable screw-top, 15 mL. 

Polypropylene bottles and caps may also be used for 

sample collection.

11. Metal spatula

12. Volumetric flasks: 10, 50, 100 and 200 mL

13. Autosampler vials, amber, with caps and septa

14. Disposable glass pipets

15. Sintered glass filtration apparatus and 1-L side- armed 

Erlenmeyer flask

16. Microfiber filters, glass, 0.7 μm; diameter to fit the 

filtration apparatus

17. Wet or dry ice 

SPECIAL PRECAUTIONS: Standard precautions should always be used when handling 

bodily fluids and/or extracts of bodily fluids [12]. Handle urine specimens and urine extracts 

using powder-free latex or nitrile gloves. Acetic acid, acetone, acetonitrile and methanol are 

flammable; handle with care and use in a chemical fume hood. Handle all chemicals using the 

required safety precautions. Reagents with manufacturer expiration dates should be observed.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 3 of 10

SAMPLING:

1. Collect at least 8 mL of urine in an appropriate polypropylene tube or bottle and cap. Refrigerate 

at 4 °C or freeze after collection. Collect at least two urine specimens for each worker, one before 

the work shift and one after.

2. Ship the specimen stored in either wet or dry ice in an insulated container. Freeze specimens 

upon arrival at the laboratory and store frozen. A reminder: commercial shippers have special 

labeling requirements for packages containing dry ice.

SAMPLE PREPARATION:

NOTE: BMA and PMA are somewhat light sensitive (see Evaluation of Method section). Perform 

the sample preparation steps in a low light environment. Extreme measures are not 

required.

3. Thaw the urine specimen to room temperature.

4. Mix urine specimen thoroughly to ensure homogeneity.

5. Transfer 4.0 mL of urine into a 16 X 150 mm (or larger) screw-capped culture tube. 

6. Add 0.5 mL of deionized water to aid in the dissolution of solids.

7. Add 0.5 mL of the 30 ng/mL deuterated BMA/PMA internal standard solution.

8. Solid-phase extraction

NOTE: All SPE steps are performed using the vacuum manifold apparatus. The flow rate should 

not exceed 1 mL/min. The SPE cartridges should not be allowed to go to dryness until 

the end of Step 8d. Refer to manufacturers’ recommendations for use of specific SPE 

cartridges.

a. Pre-wash the C18 SPE cartridge with 2 mL of acetone.

b. Equilibrate the SPE cartridge with 2 mL of HPLC grade water.

c. Load the 5-mL urine mixture and draw the sample through the cartridge.

d. Wash the cartridge with 1 mL of HPLC grade water. Discard any liquid collected up to this 

point.

e. Apply (or increase) the vacuum to pull most of the water from the cartridge.

f. Elute the analytes with 3 mL of acetone three times, collecting all of the acetone washes into a 

15-mL plastic screw-capped tube.

9. Evaporate the 9 mL of acetone from the extracts to dryness by means of a vacuum rotary 

concentrator or by using a nitrogen sweep.

10. Cap the plastic tubes containing the dry extract and store in a refrigerator/freezer until ready for 

chromatographic analysis.

11. Prior to chromatographic analysis, dissolve the extract in 1 mL of HPLC mobile phase A and 

transfer the sample into an amber HPLC autosampler vial.

CALIBRATION AND QUALITY CONTROL:

12. Prepare a BMA/PMA standard mixture by combining 2.0 mL of each stock solution and diluting to 

100 mL with deionized water to make an approximately 4 μg/mL solution concentration for each.

13. BMA/PMA standard solutions: The 4 μg/mL BMA/PMA solution is diluted in water to make 4, 8, 16, 

40, 80, 320 and 400 ng/mL BMA/PMA solutions. 

14. Accurately transfer 0.500 mL of each BMA/PMA solution from step 13 into a separate HPLC 

autosampler vial. [This delivers 2, 4, 8, 20, 40, 160 and 200 ng of each analyte to each autosampler 

vial.] 

15. Add 0.5 mL of the deuterated internal standard spiking solution [30 ng/mL] to each vial. [This 

delivers 15 ng of the internal standards to each autosampler vial.]
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S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 4 of 10

16. Add 0.1 mL of the chromatographic matrix adjustment solution to each standard vial.

NOTE: These standard samples are equivalent to 0.5 to 50 ng/mL BMA/PMA urine samples [based on 

the original 4.0 mL urine volume extracted and placed in each autosampler vial.]

17. Prepare one blank urine sample without analyte spikes; alternatively, prepare one blank using a 

urine substitute by following steps 14-16 using 0.5 mL of the unspiked urine or substitute in step 14.

18. Prepare at least one quality control (QC) standard of urine or urine substitute fortified with PMA/

BMA using separately prepared stock solutions. A 10 ng/mL equivalent spike level is suggested and 

more than one level can be used if desired.

MEASUREMENT:

19. Set the high-performance liquid chromatograph according to the manufacturer’s recommendations 

and to the conditions listed on page 8326-1. A needle rinse with 50/50% (v/v) acetonitrile/water is 

required to eliminate sample carry-over by the autosampler.

20. Set the mass spectrometric detector to multiple reaction mode (MRM) according to the 

manufacturer’s recommendations and the conditions listed on page 8326-1. Example conditions 

are summarized below:

Table 1. MS/MS Conditions using negative electrospray ionization

Analyte Precursor 

Ion

MS1

Resolution

Product

Ion

MS2

Resolution

Dwell

Time 

(msec)

Fragmentor

Voltage

Collision

Energy

(volt)

d5

-BMA 257 unit 128 unit 200 80 8

BMA 252 unit 123 unit 200 80 8

d5

-PMA 243 unit 114 unit 200 80 8

PMA 238 unit 109 unit 200 80 8

21. Inject 8 μL of each sample extract, standard, QC standard, and blank. Sample chromatograms for 

each compound are illustrated in Figure 1.

22. Measure the peak areas of the two analytes (BMA and PMA) and those for the deuterated internal 

standards (d5

-BMA and d5-PMA) in the chromatograms. Divide the peak area of the analytes by the 

peak area from the matching deuterated internal standard.

23. Prepare calibration curves of the peak Area Std./Area Internal Std. (ratio calculated in step 22) versus 

the urine equivalent concentration of the standards for the two analytes.

CALCULATIONS:

24. Determine the concentration of the two analytes in the extracts from the original urine (4.0 mL 

specimen) from the curves obtained in step 23. The results can be expressed as ng/mL of each 

analyte in urine.

EVALUATION OF METHOD:

This method was evaluated and described in detail by B’Hymer [9,10]. A general summary of this 

published information is given below: 

Accuracy and Precision. Three recovery studies using multiple columns over several days 

demonstrated the accuracy and precision of this test method. The first recovery study was performed 

using fortified urine samples over three separate experimental batch runs, and these data are presented 

in Table 2. Average recoveries were between 102 and 106% for the two analytes over the four spiked 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 5 of 10

concentration levels investigated. For each batch run, the experimental trial consisted of three samples 

at four different concentration levels. The recovery and precision for each level (n = 9 samples) are 

displayed in Table 2. The relative standard deviations (RSD) observed ranged from 2.0 to 5.0%. These 

accuracy and precision numbers fall well within parameters established for bioanalytical methods [7].

The second recovery study was performed using a urine substitute, fortified over three separate 

experimental batch runs, and these data are presented in Table 3. Average recoveries were between 

99 and 109% for the two analytes over the four spiked concentration levels investigated. For each 

batch run, the experimental trial consisted of three samples at four different concentration levels. 

The recovery for each level (n = 9 samples) is displayed in Table 3. As seen in the RSD column of 

Table 3, precision was much worse for the urine substitute recovery study. This is due to a higher level 

of ion suppression in the electrospray source. This was especially pronounced at the lower spiked 

concentration levels (1 and 2 ng/mL). The highest result for the RSD of the 9 samples fortified at 1 ng/L 

BMA was 29%. While the accuracy numbers meet standard criteria, the precision values at the lower 

concentrations do not [7]. The use of a urine substitute for QC samples is not recommended for this 

method.

The third recovery study used urine specimens from twelve non-exposed volunteers; six were smokers 

and six were non-smokers. Most participants’ specimens had base levels for both BMA and PMA as 

are shown in Table 4. Again, reasonable accuracy and precision were demonstrated for a 6 ng/mL 

equivalent level spike; individual recoveries ranged from 95 to 109% for BMA and 101 to 123% for PMA 

(Table 4). No interferences were detected in the unspiked urine from the 12 volunteer specimens. This 

study illustrates two important points: 1. Levels of both metabolites can be found in the urine of nonexposed individuals, which is why aqueous standards are used in this method, and 2. It is important to 

ascertain smoking status and other non-occupational exposures that may cause elevated levels of these 

metabolites.

Linearity. All calibration curves used during the development of this method were linear and had 

correlation coefficients of 0.99 and greater. The concentration range was equivalent to 0.5 to 50 ng/mL 

BMA and PMA for the extraction of 4.0 mL urine. Calibration curves were run at the beginning and end 

of all sample batch runs; calibration curve slope drift was found to be minimal.

Specificity. The optimized chromatographic conditions developed for this method, along with the 

tandem mass spectrometric detector, proved to be specific and showed no major interferences. The 

mass transition ions of BMA and PMA chosen in this method had the greatest response and were the 

predominant daughter ions. 

Robustness. Two C18 HPLC columns (Zorbax Rx-C18, Agilent Technologies, Santa Clara, CA, USA) from 

the same manufacturer but from different manufacturing lots were used during the recovery studies. 

Accuracy and precision were not affected; therefore, the method was found to be reproducible with 

any normal functioning C18 HPLC column. Recovery results from individual volunteer urine specimens 

spiked with the analytes indicate that the method is accurate and not significantly affected by 

individual urine specimen matrix differences during analyte extraction. The urine substitute was found 

to cause problems with precision, especially at lower levels; this was attributed to an increase in ion 

suppression over human urine within the electrospray source.

Stability. Sample stability was evaluated, whereby a six-day stability study was conducted on the final 

chromatographic sample solution. BMA and PMA were stable at 8 °C (the autosampler temperature) 

and at room temperature in the absence of light. A light stability experiment was carried out by storing 

this solution in a clear glass vial at room temperature on the window sill, thus creating a worst-case 

scenario. After 1 day of storage in light under these conditions, BMA and PMA had mean assay values of 

75 and 72% (n = 3), respectively, when compared to solutions of freshly prepared reference standards. 

After three days of light exposure, extensive degradation was noticed; BMA mean assay values had 

fallen to 9% (n = 3) of the original level and PMA had fallen to 16% (n = 3). After six days of exposure to 

light, both analytes were nearly completely degraded. Although both analytes benefit from the use of 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 6 of 10

individual deuterated internal standards, the use of amber glass autosampler vials or other means of 

reducing light exposure is recommended to ensure sample stability during extended chromatographic 

batch runs. 

Range. This method should be considered accurate for the estimation of BMA and PMA in human urine 

within the 0.5 to 50 ng/mL standard curve range. Field samples at higher levels can be diluted to a 

concentration within that range for analysis.

REFERENCES:

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio:

American Conference of Governmental Industrial Hygienists. [www.acgih.org]. Date accessed: April 

2014.

[2] Commission for the Investigation of Health Hazards of Chemical Compounds in the Work Area

[2013]. List of MAK and BAT values 2013; Maximum Concentrations and Biological Tolerance Values

at the Workplace. DFG: Deutsche Forschungsgemeinschaft, Wiley-VCH Verlag GmbH & Co. [http://

www.dfg.de/en/dfg_profile/publications/senate_commissions/index.html#micro7535782]. Date

accessed April 2014.

[3] SUVA (Swiss National Accident Insurance Fund) [2013]. Protection de la santé au poste de travail.

Valeurs limtes d’exposition aux postes de travail. [https://www.suva.ch/]. Date accessed: April 2014.

[4] Institut für Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung (IFA) [http://www.dguv.

de/ifa/GESTIS]. Date accessed: April 2014.

[5] Boogaard PJ, Van Sittert NJ [1995]. Biological monitoring of exposure to benzene: A comparison

between S-phenylmercapturic acid, trans, trans-muconic acid and phenol. Occup Environ Med

52:611-620.

[6] Schettgen T, Musiol A, Kraus T [2008]. Fast determination of urinary S-phenylmercapturic acid

and S-benzylmercapturic acid by column-switching liquid chromatography-mass spectrometry. J

Chromatogr B 863:283-292.

[7] FDA [2001]. Guidance for industry: Bioanalytical method validation, U.S. Food and Drug

Administration, Rockville, MD. [http://www.fda.gov/downloads/Drugs/Guidances/ucm070107.pdf].

Date accessed April 2014.

[8] Lovreglio P, Barbieri A, Carrieri M, Sabatini L, Fracasso ME, Doria D, Drago I, Basso A, D’Errico MN,

Bartolucci GB, Violante FS, Soleo L [2010]. Validity of new biomarkers of internal dose for use in the 

biological monitoring of occupational and environmental exposure to low concentrations of

benzene and toluene. Int Arch Occup Environ Health 83:341-356.

[9] B’Hymer C [2011]. Backup data report for method no. 8326: S-benzylmercapturic acid and

S-phenylmercapturic acid in urine.

[10] B’Hymer C [2011]. Validation of an HPLC-MS/MS method for the determination of urinary

S-benzylmercapturic acid and S-phenylmercapturic acid. J Chromatogr Sci 49:547-553.

[11] Alwis KU, Blount BC, Britt AS, Patel D, Ashley DL [2012]. Simultaneous analysis of 28 urinary VOC

metabolites using ultra high performance liquid chromatography coupled with electrospray

ionization tandem mass spectrometry (UPLC-ESI/MS-MS). Anal Chim Acta 750:152-160.

[12] CDC [2007]. 2007 Guidelines for isolation precautions: Preventing transmission of infectious agents

in healthcare settings. [http://www.cdc.gov/hicpac/2007IP/2007isolationprecautions.html]. Date

accessed: April 2014. 
S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 7 of 10

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD WRITTEN BY:

Clayton B’Hymer, Ph.D., NIOSH/DART/BHAB. Final editing performed by Dale A. Shoemaker, Ph.D., 

NIOSH/DART/CEMB.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. 

In addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for the 

content of these websites. All web addresses referenced in this document were accessible as of the 

publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 8 of 10

Figure 1. Chromatograms obtained using the described procedure: (A) fortified urine containing 1 

ng/mL S-PMA, 7.2 ng/mL S-BMA, and 3.8 ng/mL of each of the deuterated internal standards (IS) and 

(B) non-fortified urine from above containing only the background level of 6.2 ng/mL S-BMA
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 9 of 10

Table 2. Multiple level recovery experiment of BMA and PMA from urine

Analyte

Analyte

added

(ng/mL)

Mean 

concentration 

(ng/mL, n = 9)

Mean 

background 

corrected 

concentration 

(ng/mL, n = 9)

Average 

recovery (%)

Standard 

deviation 

(ng/mL) %RSD1

BMA2 1 7.43 7.18 103 0.37 5.0

BMA2 2 8.44 8.18 103 0.19 2.3

BMA2 8 14.6 14.2 103 0.32 2.2

BMA2 30 38.2 36.2 106 0.94 2.4

PMA 1 1.02 1.00 102 0.05 4.9

PMA 2 2.10 2.00 105 0.09 4.3

PMA 8 8.23 8.00 103 0.27 3.3

PMA 30 31.9 30.0 106 0.65 2.0

Notes: 1. %RSD = percent relative standard deviation

 2. The non-fortified reference urine had a background level of 6.2 ng/mL BMA and no 

detectable level of PMA.

 Three different spiked samples were prepared at each level and analyzed during three 

separate experimental trial runs (a total of nine samples at each spike level were analyzed). 

The same C18 column was used for experimental batch trials 1 and 2; a second C18 column 

was used on trial run 3.

Table 3. Multiple level recovery experiment of BMA and PMA from a urine substitute

Analyte

Analyte 

added 

(ng/mL)

Mean 

concentration 

(ng/mL, n = 9)

Average 

recovery 

(%)

Standard 

deviation 

(ng/mL) %RSD1

BMA 1 1.07 107 0.31 29

BMA 2 2.13 106 0.34 16

BMA 8 8.31 104 0.53 6.4

BMA 30 30.7 102 0.76 2.5

PMA 1 1.09 109 0.27 25

PMA 2 1.98 99 0.16 7.9

PMA 8 7.96 100 0.48 6.0

PMA 30 31.1 104 0.91 2.9

Notes: 1. %RSD = percent relative standard deviation.

 Three different spiked samples were prepared at each level and analyzed during 

three separate experimental trial runs (a total of nine samples at each spike level 

were analyzed).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

S-Benzylmercapturic acid and S-phenylmercapturic acid in urine: METHOD 8326, Issue 1, dated 20 May 2014 - Page 10 of 10

Table 4. Individual smoker and non-smoker recovery experiment of BMA and 

PMA

Background Level Fortified Urine 

(background + 6 ng/mL)

Fortified Urine Recovery

Individual 

Sample

BMA (ng/

mL)

PMA (ng/

mL)

BMA (ng/

mL)

PMA (ng/mL) BMA [ng/mL 

(percent)]

PMA [ng/mL 

(percent)]

Smoker 1 2.7 0.2 8.7 6.2 8.6 (99%) 6.5 (104%)

Smoker 2 28.3 0.3 34.3 6.3 34.7 (101%) 6.9 (108%)

Smoker 3 15.9 0.9 21.9 6.9 21.3 (97%) 7.9 (114%)

Smoker 4 5.7 0.3 11.7 6.3 11.6 (99%) 7.7 (121%)

Smoker 5 1.3 nd 7.3 6.0 6.9 ( 95%) 6.4 (106%)

Smoker 6 9.2 0.7 15.2 6.7 15.5 (102%) 7.0 (104%)

Mean 10.5 0.4 16.5 6.4 16.4 (99%) 7.1 (110%)

Non-smoker 1 0.3 nd 6.3 6.0 6.2 ( 98%) 6.2 (103%)

Non-smoker 2 7.1 nd 13.1 6.0 14.3 (109%) 7.4 (123%)

Non-smoker 3 6.8 nd 12.8 6.0 13.1 (102%) 6.8 (112%)

Non-smoker 4 23.3 nd 29.3 6.0 28.6 ( 97%) 6.3 (105%)

Non-smoker 5 4.7 nd 10.7 6.0 11.0 (103%) 6.3 (105%)

Non-smoker 6 7.2 nd 13.2 6.0 13.5 (102%) 6.1 (101%)

Mean 8.2 - 14.2 6.0 14.5 (102%) 6.5 (108%) 

Notes: The instrumental limit of detection (LOD) was estimated to be approximately 0.2 ng/mL 

for both analytes. The lowest standard value was 0.5 ng/mL for both analytes. [Values 

near the LOD were reported to one significant figure.] 

 nd = none detected (< 0.2 ng/mL)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTOS, CHRYSOTILE by XRD 9000

MW: ~283 CAS: 12001-29-5 RTECS: CI6478500

METHOD: 9000, Issue 3 EVALUATION: FULL Issue 1: 15 May 1989

Issue 3: 20 October 2015

EPA Standard (Bulk): 1% by weight PROPERTIES: Solid, fibrous mineral; conversion to forsterite 

at 580 °C; attacked by acids; loses water 

above 300 °C

SYNONYMS: Chrysotile

SAMPLING

BULK

SAMPLE: 1 g to 10 g

SHIPMENT: Seal securely to prevent escape of asbestos

SAMPLE

STABILITY: Indefinitely

BLANKS: None required

ACCURACY

RANGE STUDIED: 1% to 100% in talc [1]

BIAS: Negligible if standards and samples are 

matched in particle size [1]

OVERALL

PRECISION ( ): Unknown; depends on matrix and 

concentration

ACCURACY: ±14% to ±25%

MEASUREMENT

TECHNIQUE: X-RAY POWDER DIFFRACTION

ANALYTE: Chrysotile

PREPARATION: Grind under liquid nitrogen; wet-sieve 

through 10 μm sieve

DEPOSIT: 5 mg dust on 0.45 μm silver membrane 

filter

XRD: Copper target X-ray tube; optimize for 

intensity; 1° slit; integrated intensity with 

background subtraction

CALIBRATION: Suspensions of asbestos in 2-propanol

RANGE: 1% to 100% asbestos

ESTIMATED LOD: 0.2% asbestos in talc and calcite; 0.4% 

asbestos in heavy X-ray absorbers such as 

ferric oxide

PRECISION ( ): 0.07 (5% to 100% asbestos); 0.10 (@ 3% 

asbestos); 0.125 (@ 1% asbestos)

APPLICABILITY: Analysis of percent chrysotile asbestos in bulk samples.

INTERFERENCES: Antigorite (massive serpentine), chlorite, kaolinite, bementite, and brushite interfere. X-ray 

fluorescence and absorption is a problem with some elements; fluorescence can be circumvented with a diffracted beam 

monochromator, and absorption is corrected for in this method.

OTHER METHODS: This is NIOSH method P&CAM 309 [2] applied to bulk samples only, since the sensitivity is not adequate 

for personal air samples. An EPA test method for the determination of asbestos in bulk insulation samples is similar to this 

one [3]. NIOSH method 7400 is an optical counting procedure for airborne fibers in personal samples. NIOSH methods 

7402 (Asbestos by Transmission Electron Microscopy) and 9002 (Asbestos by Polarized Light Microscopy) are also useful for 

positive identification of asbestos.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTON, CHRYSOTILE by XRD: METHOD 9000, Issue 3, dated 20 October 2015 - Page 2 of 7

REAGENTS:

1. Chrysotile,* certified reference material.

2. 2-Propanol.*

3. Desiccant.

4. Glue or tape for securing silver filters to XRD 

holders.

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Vials, plastic (for bulk sample).

2. Freezer mill, liquid nitrogen-cooled, grinding 

vials, and extractor.

3. Ultrasonic bath.

4. Sieve, 10 μm, for wet-sieving.

5. Filters, polycarbonate, 1.0 μm, 37 mm.

6. Filtration apparatus and side-arm vacuum flask 

with 25 mm and 37 mm filter holders.

7. Oven, drying, 110 °C.

8. Analytical balance, readable to 0.01 mg.

9. Beaker, Griffin, 50 mL, with watch glass cover.

10. Filters, silver membrane, 25 mm diameter, 

0.45 μm pore size.

11. Desiccator.

12. Bottles, glass, 1 L, with ground glass stoppers.

13. Wash bottle, polyethylene.

14. Magnetic stirrer.

15. X-ray powder diffractometer with copper 

target X-ray tube and scintillation detector.

16. Reference specimen (mica, Arkansas stone or 

other stable standard) for data normalization.

17. Volumetric pipettes and flasks.

SPECIAL PRECAUTIONS: Asbestos, a human carcinogen, should be handled in a hood [4]. 

2-Propanol is flammable.

SAMPLING:

1. Place several grams of the dust to be analyzed in a plastic vial, seal the vial securely, and ship in a 

padded carton.

SAMPLE PREPARATION:

2. Place about 0.5 g of sample dust in a grinding vial and grind in a liquid nitrogen-cooled mill for 2 

min to 10 min.

3. Wet sieve the ground dust using a 10 μm sieve and 2-propanol. Place the dust on the sieve and 

place the sieve directly in an ultrasonic bath or in a wide dish in the bath. Use enough 2-propanol 

to cover the dust (put water in the bath if a dish is used to contain the 2-propanol). Apply ultrasonic 

power to sieve the dust.

NOTE: It may take some time to obtain several mg of dust. Heating of the 2-propanol is likely and 

cooling periods may be required.

4. Recover the sieved sample dust from the 2-propanol by filtering the suspension through a nonfibrous filter (polycarbonate) or by driving off the 2-propanol on a hot plate. Dry the sieved sample 

in 110 °C oven for 4 h or more.

5. Weigh out about 5 mg of the sieved material onto a small square of tared weighing paper. Record 

the actual sample weight, , to the nearest 0.01 mg. Transfer the dust to a 50 mL beaker, washing 

the weighing paper with several mL of 2-propanol. Add 10 mL to 15 mL 2-propanol to the beaker.

6. Cover the beaker with a watch glass. Agitate in an ultrasonic bath at least 3 min until all 

agglomerated particles are dispersed. Wash the underside of the watch glass with 2-propanol, 

collecting the washings in the beaker.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTON, CHRYSOTILE by XRD: METHOD 9000, Issue 3, dated 20 October 2015 - Page 3 of 7

7. Place a silver filter in the filtration apparatus. Attach the funnel securely over the entire filter 

circumference. With no vacuum, pour 2 mL to 3 mL 2-propanol onto the filter. Pour the sample 

suspension from the beaker into the funnel and apply vacuum. During filtration, rinse the beaker 

several times and add rinsings to the funnel.

NOTE: Control the filtration rate to keep the liquid level in the funnel near the top during rinsing. Do 

not wash the walls or add 2-propanol to the funnel when the liquid level is lower than 4 cm 

above the filter. Leave the vacuum on after filtration for sufficient time to produce a dry filter.

8. Remove the filter with forceps and attach it to the sample holder for XRD analysis.

CALIBRATION AND QUALITY CONTROL:

9. Prepare and analyze working standard filters:

a. Prepare two suspensions of chrysotile asbestos in 2-propanol by weighing 10 mg and 100 mg 

of the dry powder to the nearest 0.01 mg. Quantitatively transfer each to a 1 L glass-stoppered 

bottle using 1.00 L 2-propanol.

NOTE: Depending on the particle size of the standard, it may need to be ground and wet sieved 

(step 3). Dry the standards in a 110 °C oven for 4 h or more. Store in a desiccator.

b. Suspend the powder in the 2-propanol with an ultrasonic probe or bath for 20 min. Immediately 

move the flask to a magnetic stirrer with thermally-insulated top and add a stirring bar to the 

suspension. Cool the solution to room temperature before withdrawing aliquots.

c. Mount a filter on the filtration apparatus. Place several mL 2-propanol on the filter surface. Turn 

off the stirrer and shake vigorously by hand. Within a few seconds of setting the bottle down, 

remove the lid and withdraw an aliquot from the center of the 10 mg/L or 100 mg/L suspension. 

Do not adjust the volume in the pipet by expelling part of the suspension. If more than the 

desired aliquot is withdrawn, return all of the suspension to the bottle, rinse and dry the pipet, 

and take a new aliquot. Transfer the aliquot from the pipet to the filter. Keep the tip of the pipet 

near the surface but not submerged in the delivered suspension.

d. Rinse the pipet with several mL 2-propanol, draining the rinse into the funnel. Repeat the rinse 

several more times. Prepare working standard filters, in triplicate, by this technique, at e.g., 0 μg, 

20 μg, 30 μg, 50 μg, 100 μg, 200 μg, and 500 μg.

e. Apply vacuum and rapidly filter the suspension. Leave vacuum on until filter is dry. Do not wash 

down the sides of the funnel after the deposit is in place since this will rearrange the material on 

the filter. Transfer the filter to the sample holder.

f. Analyze by XRD (step 12). The XRD intensities (step 12.d) are designated and are then 

normalized (step 12.e) to obtain . The intensities for standards greater than 200 μg should be 

corrected for matrix absorption (steps 12.f and 13).

g. Prepare a calibration graph by plotting , as a function of the deposited asbestos mass, , μg, of 

each standard.

NOTE: Poor repeatability (relative standard deviation greater than 10% above 40 μg chrysotile) 

indicates that new standards should be made. The data should lie along a straight line. It is 

preferable to use a weighted least squares with weighing, where is the variance of 

the data at a given loading.

h. Determine the slope, m, of the calibration curve in counts per microgram. The intercept on the 

abscissa should be 0 μg ± 5 μg.

NOTE: A large intercept indicates an error in determining the background, i.e., an incorrect 

baseline has been calculated or interference by another phase.

10. Select six silver membrane filters as media blanks (for determination of sample self-absorption, 

step 13) randomly from the same box of filters to be used for depositing the samples. Mount each of 

the media blanks on the filtration apparatus and apply vacuum to draw 5 mL to 10 mL of 2-propanol 

through the filter. Remove, let dry, and mount on sample holders. Determine the net normalized 

count for the silver peak, , for each media blank (step 12). Obtain an average value, , for the 

normalized silver peak intensities of the six media blanks.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTON, CHRYSOTILE by XRD: METHOD 9000, Issue 3, dated 20 October 2015 - Page 4 of 7

MEASUREMENT:

11. Obtain a qualitative X-ray diffraction scan (e.g., 10 degrees to 80 degrees 2-theta) of the sample 

to determine the presence of chrysotile and interferences. The expected diffraction peaks are as 

follows:

Mineral

Peak (2-Theta Degrees)

Primary Secondary

Chrysotile 12.08 24.38

Silver 38.12 44.28

12. Mount the filter (sample, standard, or blank) in the XRD instrument and:

a. Determine the net intensity, , of the reference specimen before each filter is scanned. Select a 

convenient normalization scale factor, , which is approximately equivalent to the net count for 

the reference specimen peak, and use this value of for all analyses.

b. Measure the diffraction peak area of a chrysotile peak that is free of interference. Scan times 

should be long, e.g., 15 min.

c. Measure the background on each side of the peak for one-half the time used for peak scanning. 

The sum of these two counts is the average background. Determine the position of the 

background for each sample.

d. Calculate the net intensity, (the difference between the peak integrated count and the total 

background count).

e. Calculate and record the normalized intensity, , for the sample peak on each sample and 

standard:

.

NOTE: Normalizing to the reference specimen intensity compensates for long-term drift in X-ray 

tube intensity. If intensity measurements are stable, the reference specimen may be 

run less frequently; net intensities should be normalized to the most recently measured 

reference intensity.

f. Determine the net count, , of an interference-free silver peak on the sample filter following 

the same procedure. Use a short scan time for the silver peak (for example, 5% of scan time for 

analyte peaks) throughout the method. Normalize (step 12.e) to obtain .

g. Scan each field blank over the same 2-theta range used for the analyte and silver peaks. These 

analyses serve only to verify that contamination of the filters has not occurred. The analyte peak 

should be absent. The normalized intensity of the silver peak should match that of the media 

blanks.

CALCULATIONS:

13. Calculate the percentage, , of chrysotile in the bulk dust sample:

, %,

where: = normalized asbestos peak intensity for sample peak, 

b = intercept of calibration graph ( vs. ), 

m = slope of calibration graph (counts per μg), 

 = = absorption correction factor (see Tables 1a and 1b), 

 = , 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTON, CHRYSOTILE by XRD: METHOD 9000, Issue 3, dated 20 October 2015 - Page 5 of 7

 = = transmittance of sample, 

 = normalized silver peak intensity from sample, 

 = average normalized silver peak intensity from media blanks (average of six values), and 

 = mass, μg, of deposited sample.

NOTE: For a more detailed discussion of the absorption correction procedure, see references 5, 6, 7, 

and 8.

EVALUATION OF METHOD:

This method is based on the work of B.A. Lange in developing P&CAM 309 [1,2]. Samples in the range 

of 1% to 100% chrysotile in talc were studied to establish the feasibility of an XRD method for airborne 

asbestos. Analytical precision was as follows:

% Chrysotile in Talc (%)

100 6.9

10 4.7

7 9.8

5 8.2

3 10.1

1 12.5

This work also showed that bias of results after absorption corrections are made is negligible.

REFERENCES:

[1] Lange BA, Haartz JC [1979]. Determination of microgram quantities of asbestos by x-ray diffraction: 

chrysotile in thin dust layers of matrix material. Anal Chem 51(4):520–525.

[2] NIOSH [1979]. Chrysotile asbestos (bulk): P&CAM 309. In: Taylor DG, ed. NIOSH manual of analytical 

methods. 2nd ed. Vol. 5. Cincinnati, OH: U.S. Department of Health, Education, and Welfare, 

Center for Disease Control, National Institute for Occupational Safety and Health, DHEW (NIOSH) 

Publication No. 79-141, available as accession No. PB83-105445 from NTIS [www.ntis.gov].

[3] Perkins RL, Harvey BW [1993]. Test method: method for the determination of asbestos in bulk 

building material. Washington, DC: US Environmental Protection Agency, Office of Research and 

Development, EPA/600/R-93-116 [www.nist.gov/nvlap/upload/EPA-600-R-93-116.pdf].

[4] NIOSH [1976]. Revised recommended asbestos standard. Cincinnati, OH: U.S. Department of Health, 

Education, and Welfare, Center for Disease Control, National Institute for Occupational Safety and 

Health, DHEW (NIOSH) Publication No. 77-169 [www.cdc.gov/niosh/docs/77-169/].

[5] Leroux J, Powers CA [1970]. Direct x-ray diffraction quantitative analysis of quartz in industrial dust 

films deposited on silver membrane filters. Occup Health Rev 21(1):26–34.

[6] Williams PP [1959]. Direct quantitative diffractometric analysis. Anal Chem 31(11):1842–1844.

[7] Abell MT, Dollberg DD, Lange BA, Hornung RW, Haartz JC [1981]. Absorption corrections in x-ray 

diffraction dust analyses: procedures employing silver filters. In: Russell PA, ed. Electron microscopy 

and x-ray applications to environmental and occupational health analysis. Vol. 2. Ann Arbor, MI: Ann 

Arbor Science Publishers, Inc., pp. 115–118.

[8] Dollberg DD, Abell MT, Lange BA [1980]. Occupational health analytical chemistry: quantitation 

using x-ray powder diffraction. In: Dollberg DD, Verstuyft AW, eds. Analytical techniques in 

occupational health chemistry. Washington, DC: American Chemical Society, ACS Symposium Series 

No. 120, pp. 43–65.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTON, CHRYSOTILE by XRD: METHOD 9000, Issue 3, dated 20 October 2015 - Page 6 of 7

METHOD REVISED BY:

M. T. Abell, NIOSH/DPSE.

DISCLAIMER: Mention of any company or product does not constitute endorsement by NIOSH. In addition, citations to 

websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring organizations or their programs or 

products. Furthermore, NIOSH is not responsible for the content of these websites. All web addresses referenced in this 

document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

ASBESTON, CHRYSOTILE by XRD: METHOD 9000, Issue 3, dated 20 October 2015 - Page 7 of 7

Table 1a. Absorption correction factor as 

a function of transmittance (1.00 to 0.70) 

for chrysotile primary and secondary peak 

combinations with silver primary peak

Transmittance

Absorption Correction Factor

Primary 

Peak

Secondary 

Peak

1.00 1.0000 1.0000

0.99 1.0157 1.0078

0.98 1.0317 1.0157

0.97 1.0480 1.0237

0.96 1.0647 1.0319

0.95 1.0817 1.0402

0.94 1.0991 1.0486

0.93 1.1168 1.0572

0.92 1.1350 1.0659

0.91 1.1535 1.0747

0.90 1.1724 1.0837

0.89 1.1917 1.0928

0.88 1.2114 1.1021

0.87 1.2316 1.1115

0.86 1.2522 1.1212

0.85 1.2733 1.1309

0.84 1.2948 1.1409

0.83 1.3168 1.1510

0.82 1.3394 1.1613

0.81 1.3624 1.1718

0.80 1.3859 1.1825

0.79 1.4100 1.1933

0.78 1.4346 1.2044

0.77 1.4598 1.2157

0.76 1.4856 1.2272

0.75 1.5120 1.2389

0.74 1.5390 1.2508

0.73 1.5666 1.2630

0.72 1.5949 1.2754

0.71 1.6239 1.2881

0.70 1.6536 1.3010

Table 1b. Absorption correction factor as 

a function of transmittance (0.69 to 0.39) 

for chrysotile primary and secondary peak 

combinations with silver primary peak

Transmittance

Absorption Correction Factor 

Primary 

Peak

Secondary 

Peak

0.69 1.6839 1.3142

0.68 1.7151 1.3277

0.67 1.7470 1.3414

0.66 1.7797 1.3555

0.65 1.8132 1.3698

0.64 1.8475 1.3845

0.63 1.8827 1.3995

0.62 1.9188 1.4148

0.61 1.9558 1.4305

0.60 1.9938 1.4465

0.59 2.0328 1.4629

0.58 2.0728 1.4797

0.57 2.1139 1.4969

0.56 2.1560 1.5145

0.55 2.1993 1.5325

0.54 2.2438 1.5510

0.53 2.2895 1.5700

0.52 2.3365 1.5895

0.51 2.3848 1.6095

0.50 2.4344 1.6300

0.49 2.4855 1.6510

0.48 2.5380 1.6727

0.47 2.5921 1.6950

0.46 2.6478 1.7179

0.45 2.7051 1.7414

0.44 2.7642 1.7657

0.43 2.8251 1.7907

0.42 2.8879 1.8165

0.41 2.9526 1.8431

0.40 3.0195 1.8705

0.39 3.0885 1.8989
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE and Illicit Drugs, Precursors 9106

and Adulterants on Wipes by Liquid-Liquid Extraction

FORMULA: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 9106, Issue 1 EVALUATION: Partial Issue 1: 17 October 2011

OSHA: none for surfaces

NIOSH: none for surfaces

Other OELs and guidelines: [1, 2, 3]

PROPERTIES: Table 2

SYNONYMS: Table 3

SAMPLING

SAMPLER: Wipe

SAMPLE AREA: 100 cm2 or 1000 cm2

SHIPMENT: Preferably ship refrigerated, <6 °C

SAMPLE

STABILITY: 30 days at <6 °C (See Table 4) 

FIELD BLANKS: 2 to 10 blanks per sample set

MEASUREMENT ACCURACY

LEVEL STUDIED: 3 μg/sample on smooth surfaces 

BIAS: Table 10a and 10b

OVERALL

PRECISION ( ): Table 7a and 7b [4] 

ACCURACY: Table 7a and 7b [4]

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY/MASS 

SPECTROMETRY

ANALYTES: Table 1

DESORPTION: 0.1 M sulfuric acid

CLEANUP/

EXTRACTION: Hexane cleanup followed by methylene 

chloride extraction

DERIVATIZATION: Chlorodifluoroacetic anhydride

INJECTION: 2 μL, splitless

TEMPERATURE

 - Injection: 265 °C

- Detector: 285 °C

 - Column: 90 °C (2 min), 310 °C (10 °C/min),

 hold for 11 min

MASS 

SPECTROMETER: Scan mode (29 – 470 AMU), 2 scan per 

sec, Selected ion monitoring (SIM) 

mode Table 5

CARRIER GAS: Helium, 1.5 mL/min 

COLUMN: Capillary, fused silica, 30 m × 

0.32 mm ID; 0.5 μm film phenyl arylene 

polymer

CALIBRATION: Standards from spiked wipes with 

internal standard, See Table 6

RANGE: Table 7a and 7b [4]

ESTIMATED LOD: Table 4

PRECISION ( ): Table 7a and 7b [4]

APPLICABILITY: For methamphetamine, the range is 0.05 to 60 μg/sample (sample = 100 cm2 or 1000 cm2). This method was 

developed for the analysis of selected drugs and precursors on surfaces in clandestine drug labs. [4, 5] Sampling methodology was tested using wipes on smooth, non-porous surfaces. The APPENDIX contains sampling information for other types of 

surfaces.

INTERFERENCES: No chromatographic interferences detected. Water, surfactants and polyols inhibit derivatization.

OTHER WIPE METHODS: NIOSH 9109 uses solid-phase extraction and gas chromatography/mass spectrometry (GC/MS) to 

measure multiple drugs [6]. NIOSH 9111 uses liquid chromatography/mass spectrometry (LC/MS) to measure methamphetamine [7]. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 2 of 30

REAGENTS:

NOTE: See APPENDIX A for special instructions on reagents.

1. Analytes listed in Table 1.*

2. Internal standards from those listed in Table 8.

3. Solvents, residue free analytical grades:

a. Hexane *

b. Isopropanol (IPA)*

c. Methanol *

d. Methylene chloride (CH2Cl2) *

e. Toluene *

f. Acetone*

4. Concentrated sulfuric and hydrochloric acids (AR or trace metals 

analysis grades).*

5. Sodium hydroxide, A.C.S. grade.*

6. Anhydrous granular sodium sulfate, AR grade.

7. Anhydrous granular potassium carbonate, AR grade.

8. Bromothymol Blue, ≥95%, A.C.S., phenolphthalein, A.C.S.; crystal 

violet (Gentian Violet), ≥95%, A.C.S.

9. Purified gases: helium for carrier gas, nitrogen for drying. 

10. Chlorodifluoroacetic anhydride, 98%* derivatizing agent. Moisture 

sensitive!

11. 4,4’-Dibromooctafluorobiphenyl, 99%, instrument internal standard 

(IIS).

12. Deionized water (ASTM type II).

SOLUTIONS:

NOTE: See APPENDIX A for special instructions on solutions.

1. Prepare solutions of analytes of interest. Calculate concentrations as 

the free base. Keep refrigerated (<6 °C). Protect solutions from light.

a. Stock solutions are prepared at about 1-2 mg/mL in methanol.

b. Analyte spiking solutions are prepared by diluting the stock 

solutions to about 200 μg/mL each in methanol.

2. Prepare internal standard spiking solution in methanol at about 

200 μg/mL. (Note: Add about 2 milligrams of crystal violet per 20 mL 

of internal standard spiking solution to help indicate which samples 

have been spiked.)

3. Desorption solution: 0.1 M H2

SO4 (sulfuric acid.) Add 22 mL conc. 

sulfuric acid to 4 L deionized water.

4. Bromothymol blue and phenolphthalein pH indicator solution: 1 mg/

mL each in 4:1 isopropanol: deionized water.

5. Sodium hydroxide*, 10 M: Dissolve 40 grams sodium hydroxide in 

enough deionized water to make 100 mL. Do not store in glassstoppered bottle.

6. Hydrochloric acid, 0.3 M, in methanol: Dilute 2.5 mL conc. hydrochloric 

acid in about 80 mL methanol; dilute to 100 mL with methanol.

7. Crystal violet indicator: 2-3 mg/mL in isopropanol.

8. Reconstitution solvent: 10% acetone in toluene with 4 μg/mL of 

4,4’-dibromooctafluorobiphenyl (optional).*

* See SPECIAL PRECAUTIONS

EQUIPMENT:

NOTE: See APPENDIX B for special instructions on equipment.

1. Wipe, (7.6 cm × 7.6 cm) 12-ply or equivalent.

2. Sample storage and shipping container: 50-mL polypropylene 

centrifuge tubes with PTFE-lined caps.

3. Extraction tubes and vials:

a. Glass test tubes, 25-mL (20 × 120 mm), with PTFE-lined caps;

b. Glass test tubes, 14-mL (16 × 100 mm), with PTFE- lined caps, 

(ASTM Specification E982 [5], or equivalent, suitable for repeated 

autoclaving);

c. Amber GC autosampler vials (2-mL) and caps.

4. Gas chromatograph/mass spectrometer detector, with column and 

integrator, see p. 9106-1.

5. Liquid Transfer:

a. Syringes: 10-, 25-, 100-, and 500-μL sizes.

b. Mechanical pipette with disposable tips or repeating dispenser: 

0.5-, 2.5-, and 10-mL sizes.

c. Repeating dispenser: 1- to 5-mL.

d. Three repeating dispensers: 10 to 20-mL each.

6. Volumetric flasks: 10-, 100-, and 250-mL.

7. Forceps.

8. Gloves, latex or nitrile. Avoid vinyl gloves (see 9106-3, Sampling 

step 1, NOTE 2).

9. Scoop for solid reagents.

10. Empty drying columns: 1 cm i.d. × 12-15 cm length polypropylene 

plastic columns having a fritted polyethylene disc or equivalent (e.g. 

10-mL pipette tip with small wad of silanized glass wool packed into 

the tip).

11. Nitrogen blow-down apparatus with water bath capable of 

maintaining 35 oC.

12. Vortex mixer.

13. Rotating mixer capable of 10-30 rpm.

14. Aspirator flask: 1-L, with aspirator tubing and a 12.5 cm long 16 

gauge needle.

15. Centrifuge: capable of up to 4000 x g and of holding 25-mL glass test 

tubes.

16. Oven capable of 70 to 90 oC ± 2 oC. 

17. Test tube racks, heat resistant to 90 oC.

18. Pasteur pipettes.

19. pH paper.

20. Template: 10 cm × 10 cm or 31.7 cm × 31.7 cm opening, made of 

relatively rigid disposable cardstock or sheet of PTFE.

21. Ice or other cold media for shipping.
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METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 3 of 30

SPECIAL PRECAUTIONS: The solvents are flammable and have adverse health effects. 

Phenethylamines target the nervous system at very low concentrations and are easily absorbed 

through the skin. Avoid breathing vapors. Avoid skin contact. Work should be performed in a hood 

with adequate ventilation. Analysts must wear proper eye and hand protection (e.g. latex gloves) 

to prevent absorption of even small amounts through the skin. Dissolving sodium hydroxide and 

concentrated hydrochloric or sulfuric acid in water is highly exothermic. Goggles must be worn. The 

derivatization reagents react violently with water.

Caution must also be exercised in the collecting, handling, and analysis of samples. Clandestine 

drug labs may produce unknown and seriously toxic by-products. For example, in the manufacture 

of designer drugs (e.g., MPPP, a homolog of Alphaprodine), at least one very neurotoxic by-product, 

1-methyl-4-phenyl-1,2,5,6-tetrahydropyridine (MPTP), has been identified that specifically and 

irreversibly causes Parkinson’s disease [8,9].

SAMPLING:

See APPENDIX C for special instructions on sampling.

1. Using a new pair of gloves, remove a gauze wipe from its protective package. Moisten the wipe

with approximately 3 to 4 mL of methanol (or isopropanol).

NOTE 1: Apply no more solvent than that needed to moisten approximately the central 80% of 

the area of the gauze wipe. Excess solvent may cause sample loss due to dripping from 

the wipe.

NOTE 2: Do not use vinyl gloves due to the potential for leaching of phthalate plasticizers and 

contamination of the samples.

2. Place the template over the area to be sampled (may tape in place along outside edge of

template). Wipe the surface to be sampled with firm pressure, using vertical S-strokes. Fold the 

exposed side of the pad in and wipe the area with horizontal S-strokes. Fold the pad once more 

and wipe the area again with vertical S strokes.

3. Fold the pad, exposed side in, and place in shipping container and seal with cap.

NOTE: Keep samples refrigerated (<6 oC). While methamphetamine and several related amines 

are stable on the recommended wipe media for at least 7 days at room 

temperature, refrigeration is recommended as soon as possible (see Table 4).

4. Either clean the template before use for the next sample or use a new disposable template.

5. Label each sample clearly with a unique sample identifier.

6. Prepare a minimum of two field blanks with one field blank for every ten samples.

NOTE: In addition, include at least 3 media blanks for the analytical laboratory to use for their

purposes. The wipes used for the media blanks should be from the same lot as the field 

samples.

SAMPLE PREPARATION:

See APPENDIX D for special instructions on sample preparation.

7. Desorption from media:

a. Remove cap from shipping container.

NOTE: Sample wipe should fit loosely in the container. If not, transfer sample to a larger

container.

b. Spike 60 μL of internal standard spiking solution onto each wipe sample.

c. Add 30 mL desorption solution (0.1 M sulfuric acid).

NOTE: If the samples were transferred to a larger container, rinse the original shipping

container with the desorption solution, shake, and decant the rinsate into the larger 

container.

d. Cap securely and mix contents by inverting the tubes end over end on a rotary mixer or

equivalent at 10-30 rpm for at least one hour.
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e. Check the pH which should be about ≤ 4. Adjust the pH with diluted (2.5 to 3 M) sulfuric acid

drop-wise, mixing the contents by shaking or inversion a few times after each addition of acid

before checking the pH.

f. After mixing, transfer 10 mL of supernatant to a 25-mL glass centrifuge tube.

NOTE: If cleanup is to be performed on a subsequent day, store samples in a refrigerator.

Analytes are stable in the desorption solution for at least one week refrigerated.

8. Cleanup: Potential contamination from oils, triglycerides, plasticizers and other hydrocarbons are

reduced through a hexane back-extraction step.

a. To each 10-mL aliquot of acid desorbate, add 10 mL of hexane, cap and mix on a rotary mixer

for one hour. Allow to stand for 15-30 minutes for the phases to separate. If an emulsion forms,

centrifuge the tubes at 1500-2000 rpm for a few minutes. If the emulsion persists, add about 0.5

mL of acetonitrile to the surface of the emulsion and gently mix the layers at the interface of the

emulsion. Centrifuge again if necessary.

b. Aspirate the (upper) organic layer to waste. Exercise care to not remove any of the aqueous layer.

9. Extraction of analytes into methylene chloride:

a. Add 1-2 drops (20-50 μL) of the mixed pH indicator (phenolphthalein + bromothymol blue)

solution to each sample. The color of the samples should be yellow, which indicates that the

samples are sufficiently acidic for desorption of the analytes from the wipe samples.

b. Add 0.5 mL of 10 M sodium hydroxide to each sample. The color of the samples should turn

brilliant purple or magenta, confirming that the pH is greater than 9-9.5 (necessary for the

extraction of the amines into methylene chloride). If the color remains yellow, or only turns green

or light blue, check the pH with pH paper to confirm that it is greater than 9.5. If it is not, add

another 0.5 mL of 10 M sodium hydroxide, mix, and check the pH again.

NOTE: The color of the solution will gradually fade from purple to deep blue within about 20-30

minutes. This is due to the known tendency of phenolphthalein to fade at high pH.

c. Add 10 mL of methylene chloride to each sample. Cap and mix on a rotary mixer for one hour.

Allow to stand for 15 to 30 minutes. If an emulsion forms, centrifuge as described above (step 8a).

d. Aspirate the aqueous (upper) layer to waste as described above, being careful to not remove any

of the lower methylene chloride layer.

10. Removal of water from the methylene chloride extract:

a. Prepare potassium carbonate-sodium sulfate drying columns.

Note: See APPENDIX E for preparation of drying columns.

b. Rinse the packed columns with about 6 mL of methylene chloride. Dry columns afterwards by

forcing dry nitrogen or clean air through the top for 10-20 seconds.

c. Arrange 14-mL collection tubes (16 × 100 mm test tube) in test tube racks. Add 6 μL of crystal

violet solution and 100 μL of 0.3 M hydrochloric acid in methanol to each collection tube.

NOTE: Crystal violet is not critical but helps later on as a visualizing aid for monitoring the

progress of drying. Hydrochloric acid is critical to prevent loss of the amphetamines 

during evaporative concentration.

d. Position the drying columns over the collection tubes.

e. Transfer (decant) the methylene chloride layer into the drying column reservoir. After the last

of the sample passes into the bed of the column, rinse the drying column twice with 1 mL of

methylene chloride each time and combine with sample eluate.

11. Derivatization:

See APPENDIX F for special instructions on derivitization.

a. Evaporate the methylene chloride eluates in a nitrogen blow-down apparatus with the water

bath set at 35 °C. Rinse the tips of the evaporation needles thoroughly with methanol or acetone

between samples to prevent cross-contamination. When the samples are dry, remove and cap the

tubes immediately.
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NOTE: The dark color of the crystal violet helps make the residue more visible when it is dried. 

If at least 0.1 mL of isopropanol was present in the eluates, the crystal violet will also go 

through a series of color changes that helps in monitoring the drying process.

b. To each dried sample, add 100 μL of chlorodifluoroacetic anhydride and recap tubes. Mix the

contents by vortexing briefly.

NOTE 1: It is recommended that the tubes be kept capped and to only uncap about 5 at a time 

for the addition of the derivatizing reagent. Do not leave the acid anhydride bottle open 

between taking aliquots since the reagent is moisture sensitive.

NOTE 2: If incomplete derivatization is routinely experienced, increase volume of reagent to 

150 or 200 μL. The color of the crystal violet will turn yellow or yellow-green with the 

addition of chlorodifluoroacetic anhydride.

c. Heat in an oven at 70-75 °C for 20-30 minutes.

d. After heating, allow the tubes to cool to room temperature. Remove caps and evaporate

the contents to dryness under a stream of nitrogen at room temperature. As the solution

concentrates it turns from a yellow or yellowish-green solution to a bluish-green just before

going to dryness. At the point of dryness the color of the residue normally turns rapidly to blue

or violet, depending upon the amount of coextractants (the more co-extractants, the more blue

the color and the less likely a violet color will develop). Remove the tubes just as soon as the blue

or violet color becomes apparent. Losses have been experienced if blowing is continued for more

than 2 minutes beyond the blue or violet color stage.

NOTE: If an oil-like residue or film persists, then the sample may have too many contaminants

that were not removed at the cleanup step or were introduced subsequent to cleanup. In 

such a case, return to step 7f and perform the clean-up (step 8) on another 10-mL aliquot 

of the sample desorbate using methylene chloride as the cleanup solvent instead of 

hexane. Discard the (lower) organic layer to waste before proceeding to steps 9 through 

11.

e. Reconstitute the dried residue with 1 mL of the reconstitution solvent. The reconstituted solution

normally will become deep blue in color. Mix by vortexing briefly a couple of times. Transfer the 

solutions to 2-mL amber-colored GC vials containing 200 to 250 mg anhydrous sodium sulfate. 

Cap vials, label, and analyze by GC/MS (See MEASUREMENT, steps 15-17).

NOTE: Derivatives of phenylpropanolamine (norephedrine) break down significantly over several 

days at room temperature. GC vials containing derivatives should be kept refrigerated 

until analysis.

CALIBRATION AND QUALITY CONTROL:

12. Determine retention times for the derivatives of the analytes of interest using the column and

chromatographic conditions specified on page 9106-1. Table 9 gives typical retention times for

various drugs, precursors, and adulterants.

13. Calibrate daily with at least six calibration standards plus a blank (CS0) selected from Table 6 to cover

the analytical range.

a. Prepare the analyte spiking solution as follows: Add known amounts of individual drug stock

solutions to a volumetric flask and dilute to volume with methanol. A recommended final

concentration for this solution is approximately 200 μg each per mL.

b. Prepare calibration standards and media blanks in clean shipping containers (e.g. 50-mL

polypropylene centrifuge tubes or equivalent).

NOTE: Liquid standards (standards without added blank wipe media) may be prepared in lieu of

media standards if cotton gauze was used for the samples.

c. Add 3 mL methanol (or isopropanol, if isopropanol was used with the samples in the field) to

each calibration standard and media blank.
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d. Spike a known volume of analyte spiking solution into each calibration standard by spiking

directly onto the media or into solution. Use the spiking volumes suggested in Table 6 to cover

the desired range.

e. Process each of these through steps 7 through 11 (same as the field samples.)

f. Analyze these along with the field samples. (See MEASUREMENT, steps 15-17.)

14. Prepare matrix-spiked (QC) and matrix-spiked duplicate (QD) quality control samples [12].

a. Cotton gauze from the same lot used for taking samples in the field should be provided to the

analytical laboratory to prepare these matrix-spiked quality control samples.

b. The quality control samples (QC and QD) must be prepared independently at concentrations

within the analytical range. (See Table 6 for applicable concentration ranges.)

c. One quality control media blank (QB) must be included with each QC and QD pair.

i. Transfer clean gauze wipes to new shipping containers.

ii. Add 3 mL of isopropanol (or methanol, if methanol was used in wiping) to each gauze wipe.

iii. Spike QC and QD with a known amount of analyte as suggested in Table 6.

d. Process each of these through steps 7 through 11 (same as the field samples).

e. Analyze these along with the field samples. (See MEASUREMENT, steps 15-17.)

MEASUREMENT:

See APPENDIX G for special instructions on measurement.

15. Analyze the calibration standards, quality control samples, blanks, and samples by GC/MS.

a. Set gas chromatograph according to manufacturer’s recommendations and to conditions listed

on page 9106-1.

b. Set mass spectrometer conditions to manufacturer’s specifications and those given on page

9106-1 for the scan mode or those in Table 5 for the SIM mode.

c. Inject sample aliquot with autosampler or manually.

NOTE: After the derivatives are prepared and just before analyzing any samples or standards,

inject the highest concentrated standard several times in order to prime or deactivate 

the GC column and injection port. This will help minimize any drift in the instrument’s 

response to target analytes relative to their internal standards.

d. After analysis, the vials should be recapped promptly and refrigerated if further analysis is

anticipated.

16. Using extracted ion current profiles for the primary (quantification) ions specific to each analyte,

measure GC peak areas of analyte(s) and internal standard(s) and compute relative peak areas 

by dividing the peak area of the analyte by the area of the appropriate internal standard. 

Recommended primary (quantification) ions and internal standards are given in Tables 5, 7, and 8. 

Prepare calibration graph (relative peak area vs. μg analyte per sample).

17. Samples from initial investigations of clandestine laboratories are likely to include highly

contaminated samples. If sample results exceed the upper range of the calibration curve, the sample 

in the GC vial may be diluted and reanalyzed or a smaller aliquot of the initial acid desorbate diluted, 

re-extracted, derivatized, and analyzed. Refer to APPENDIX H for instructions and limitations on 

making dilutions.

CALCULATIONS:

18. Determine the mass (in μg/sample) of respective analyte found in the wipe samples, and in the

media blank from the calibration graph.

19. Calculate final concentration, C, of analyte in μg/sample:

Where: c = concentration in sample (in μg/sample determined from the calibration curve). 
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  volume correction factor (needed only when the volume of internal standard

spiking solution used for spiking the samples – such as for composite samples 

requiring larger desorption solution volumes – is different from that used for 

spiking the calibration standards). (See Table 6, footnote 4.)

V1

 = volume in μL of internal standard spiking solution used to spike samples.

V2

 = volume in μL of internal standard spiking solution used to spike the standards.

  dilution factor, if applicabl

V3

 = 10 mL (volume of desorbate taken for cleanup in step 8).

V4

 = volume in mL of desorbate actually taken for cleanup and diluted to 10 mL with

 blank desorbing solution containing internal standard.

b = concentration in media blank (in μg/sample determined from the calibration 

curve). volume correction factor for the media blank (needed only if the volume of

internal standard spiking solution used for spiking the media blank is different 

from that used for spiking the calibration standards.)

V5

 = volume in μL of internal standard spiking solution used to spike media blank.

20. Report concentration, C’, in μg per total area wiped (in cm2

) as follows:

Where: C = μg/sample (step 19).

A = Total area wiped in cm2

 per sample.

NOTE: For example, if the sample was a composite sample and the area was 400 cm2

, report 

results as μg/400 cm2

 and not averaged to μg/100 cm2. In general, if the area wiped 

was greater than or less than 100 cm2

, do not convert value to μg/100 cm2. To avoid 

confusion, report separately both μg/sample (C) and the total area wiped in cm2

 per 

sample (A) for both discrete and composite samples.

EVALUATION OF METHOD:

This method was evaluated for those analytes listed in Tables 7a and 7b over a range of approximately 

0.1 μg/sample to 30 μg/sample. These concentration levels represent approximately the 1 through 300 

times the limit of quantitation (LOQ) level for most of the analytes [9]. Results are reported in the Backup 

Data Report for NIOSH 9106 [4].

The limits of detection (LOD and LOQ) were determined by preparing a series of liquid standards in 

desorption solution, processing them through the liquid-liquid extraction procedure of NIOSH 9106, 

and analyzing in both the scan and SIM modes. The LODs were estimated using the procedure of 

Burkart [12]. An LOD of 0.05 μg/sample for methamphetamine on wipes was achieved in either scan 

or SIM mode. The LOD was set at 0.05 μg/sample because that was the level of the lowest calibration 

standard for the LOD study. Lower LODs (e.g. 0.02 μg/sample) have been achieved in practice by 

including calibration standards at lower concentration levels. The cleanliness and performance of the 

mass spectrometer must be maintained such that at 0.1 μg/sample, a signal of at least 5 to 10 times the 

baseline noise is achievable. This is more easily accomplished in the SIM mode with the mass spectrometer. 

Six different wipe media were evaluated. These were 3”×3” 12-ply cotton gauze, 4”×4” AlphaWipes® 

(TX® 1004), 4”×4” 4-ply NU GAUZE®, 4”×4” 4-ply MIRASORB®, 4”×4” 6-ply SOF-WICK®, and 4”×4” 4-ply 

TOPPER® sponges. Results are given in the Backup Data Report [1]. No synthetic media performed 
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better than cotton gauze. Some media (TOPPER® and SOF-WICK®) did not perform well, possibly due to 

co-extracted nonionic (polyethoxyethylene type) surfactants that are not removed using hexane and 

incompletely removed using methylene chloride in the cleanup step.

Precision and accuracy were determined by analyzing 6 replicates at each of 6 concentration levels 

(nominally 0.1, 0.3, 1, 3, 10, and 30 μg/sample). Results are presented in Tables 7a and 7b for cotton 

gauze. The best precision and accuracies were dependent upon the use of carefully chosen internal 

standards, especially where there is steric hindrance of the amine (e.g. having N-ethyl and N-propyl 

groups).

Long term sample storage stability was determined for periods up to 30 days under refrigeration

(4 °C ± 2 °C) and for up to 7 days at room temperature (22-24 °C). Results are given in Table 4.

The precision and accuracy and long term storage stability evaluations were conducted using 

isopropanol as the wetting solvent. A second precision and accuracy study using methanol confirmed 

that methanol is an acceptable substitute for isopropanol.

Recovery of amphetamines from six different types of surfaces using cotton gauze was evaluated (see 

Tables 10a and 10b). The practice of serial wiping (wiping the same surface area a second time with 

a second gauze wipe and combining both wipes as a single sample) was evaluated. Four solvents for 

wetting the gauze were tested (distilled water, 5% distilled white vinegar, isopropanol, and methanol). 

Six replicate samples were made on a latex painted wall. Recoveries and precisions are given in Table 

10a. The recoveries with 5% distilled white vinegar were better than for distilled water, but not as 

good as for isopropanol. Methanol is superior to isopropanol. Recoveries with isopropanol are greatly 

improved with a repeat (serial) wipe (11% improvement compared to only about 6% improvement 

with methanol). The study and results are reported in the Backup Data Report for NIOSH 9109 [13]. 

Additional research on surface sample recovery and solvent effectiveness has been reported by 

Serrano et al. [15] and VanDyke et al. [16].

REFERENCES:

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for 

chemical substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: 

American Conference of Governmental Industrial Hygienists. [www.acgih.org]. Data accessed April 

2014.

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database 

on hazardous substances (German Social Accident Insurance). Sankt Augustin, FRG: [http://www. 

dguv.de/ifa/GESTIS ]. Data accessed April 2014.

[3] EPA [2013]. Voluntary guidelines for methamphetamine laboratory cleanup. [https://www.epa.gov/

sites/production/files/documents/meth_lab_guidelines.pdf]. Date accessed March 2016.

[4] Reynolds JM, Siso MC, Perkins JB [2004]. Backup data report for NIOSH 9106, Methamphetamine 

and illicit drugs, precursors, and adulterants on wipes by liquid-liquid extraction, Abridged, 

prepared under NIOSH Contract 200-2001-0800, Unpublished.

[5] Martyny JW, Arbuckle SL, McCammon CS Jr., Esswein EJ, Erb N [2003]. Chemical exposures 

associated with clandestine methamphetamine laboratories. [https://www.env.nm.gov/cdli/docs/

Chemical_Exposures.pdf].

[6] NIOSH [2011]. Methamphetamine and illicit drugs, precursors, and adulterants on wipes by solid 

phase extraction: Method 9109. In: Ashley KA, O’Connor PF, eds. NIOSH manual of analytical 

methods. 5th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for 

Disease Control and Prevention, National Institute for Occupational Safety and Health, [www.cdc.

gov/niosh/nmam/].

[7] NIOSH [2011]. Methamphetamine on Wipes by Liquid Chromatography/Mass Spectrometry: 

Method 9111. In: Ashley KA, O’Connor PF, eds. NIOSH manual of analytical methods. 5th ed. 

Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and 

Prevention, National Institute for Occupational Safety and Health, [www.cdc.gov/niosh/nmam/].
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[8] ASTM International [2015]. ASTM-E982-94(2015), Standard specification for laboratory glass test

tubes. West Conshohocken, PA: ASTM International.

[9] Baum RM [1985]. New variety of street drugs poses growing problem. Chemical and Engineering

News, 63(63):7-16.

[10] Buchanan JF, Brown CR [1988]. Designer drugs, a problem in clinical toxicology. Medical Toxicology

3:1-17.

[11] NIOSH [1994]. Chapter C: Quality Assurance. In: Eller PM, Cassinelli ME, eds. NIOSH Manual of

Analytical Methods, 4th ed. Cincinnati, OH: Department of Health and Human Services, Centers for 

Disease Control and Prevention, National Institute for Occupational Safety and Health, DHHS

(NIOSH) Publication No. 94-113. [www.cdc.gov/niosh/nmam/].

[12] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene

chemistry laboratory. Appl Ind Hygiene 1(3):153-155.

[13] Reynolds JM, Siso MC, Perkins JB [2004]. Backup data report for NIOSH 9109, Methamphetamine

and illicit drugs, precursors, and adulterants on wipes by solid phase extraction. Unpublished.

Prepared under NIOSH Contract 200-2001-0800.

[14] Budavari S, ed. [1989]. Merck Index, 11th ed. Rahway, NJ: Merck and Co., Inc.

[15] Serrano KA, Martyny JW, Kofford S, Contreras JR, Van Dyke MV [2012]. Decontamination of clothing

and building materials associated with the clandestine production of methamphetamine. J Occup

Environ Hyg 9(3):185-197.

[16] Van Dyke MV, Serrano KA, Kofford S, Contreras J, Martyny JW [2011]. Variability and specificity

associated with environmental methamphetamine sampling and analysis. J Occup Environ Hyg

8(11):636-41.

[17] Sweet DV, ed. [1997]. Registry of toxic effects of chemical substances. DHHS (NIOSH) Publ. No.

97-119. https://www.cdc.gov/niosh/.

[18] Sigma-Aldrich MSDS sheets, http://www.sigma-aldrich.com [May 10, 2004].

[19] Cerilliant Analytical Reference Materials. 811 Paloma Drive, Suite A, Round Rock, Texas 78664 (www.

cerilliant.com).

[20] The National Alliance for Model State Drug Laws (NAMSDL) [2007]. State Controlled Substance(s)

Environmental Issues Bill – Status Update [http://www.namsdl.org/]. Date accessed: March 11, 2008.

[21] Howard PH, Meylan WM [1997]. Handbook of physical properties of organic chemicals. Boca Raton,

Florida: CRC/Lewis Publishers. See also Syracuse Research Corporation’s website http://www.srcinc.

com/ (May 10, 2004) for demos and updates.

[22] Zhang JS, Tian Z, Lou ZC [1989]. Quality evaluation of twelve species of chinese ephedra (Ma

Huang). Acta Pharmaceutic Sinica 24(11):865-871.

[23] Cui JF, Zhou TH, Zhang JS, and Lou ZC. [1991]. Analysis of alkaloids in chinese ephedra species by

gas chromatographic methods. Phytochemical Analysis 2:116-119.

[24] Andrews KM [1995]. Ephedra’s role as a precursor in the clandestine manufacture of

methamphetamine. Journal of Forensic Sciences 40(4):551-560.

[25] OSHA [2001]. Evaluation guidelines for surface sampling methods. Industrial Hygiene Chemistry

Division, OSHA Salt Lake Technical Center, Salt Lake City, Utah. [http://www.osha.gov/dts/sltc/

methods/surfacesampling/surfacesampling.pdf] Date accessed: May 2011. 

METHOD DEVELOPMENT BY:

John M. Reynolds, Maria Carolina Siso, and James B. Perkins, DataChem Laboratories, Inc., Salt Lake City, 

Utah under NIOSH Contract CDC 200-2001-0800.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In 

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the 

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for 

the content of these websites. All web addresses referenced in this document were accessible as of 

the publication date. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 10 of 30

Table 1. Formula and registry numbers of analytes.

Compound

(alphabetically)

MW(1) (Daltons)

Free 

base HCl salt

Hemisulfate 

salt

Structural Formula

As free base CAS #(2) RTECS(6)

(DL)-Amphetamine 135.21 171.67 184.25 C6

H5·CH2·CH(CH3)·NH2 300-62-9(3)

60-13-9(5)

SH9450000

SI1750000

(D)-Amphetamine(7) 135.21 171.67 184.25 C6

H5·CH2·CH(CH3)·NH2 51-64-9(3)

51-63-8(5)

SI1400000

(L)-Amphetamine 135.21 171.67 184.25 C6

H5·CH2·CH(CH3)·NH2 156-34-3(3) SH9050000

Caffeine 194.19 (CH3

)

3

·[C5HN4O2] 58-08-2(3) EV6475000

(DL)-Ephedrine 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)·NH·CH3 90-81-3(3)

134-71-4(4)

(L)-Ephedrine(8) 165.24 201.70 214.28 C6H5·CH(OH)·CH(CH3)NH·CH3 299-42-3(3)

50-98-6(4)

134-72-5(5)

KB0700000

KB1750000

KB2625000

(D)-Ephedrine 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)NH·CH3 321-98-2(3)

24221-86-1(4)

KB0600000

KB1925000

(±)-MDEA 207.27 243.73 CH2

O2C6H3·CH2·CH(CH3)NH·C2H5 82801-81-8(3)

116261-63-2(4)

(±)-MDMA 193.24 229.71 CH2O2C6H3·CH2·CH(CH3)·NH·CH3 42542-10-9(3)

92279-84-0(4)

SH5700000

(+)-MDMA(7) 193.24 229.71 CH2

O2C6H3·CH2·CH(CH3)·NH·CH3 64057-70-1(4) SH5700000

(DL)-Methamphetamine 149.24 185.70 198.28 C6

H5·CH2·CH(CH3)·NH·CH3 4846-07-5(3)

(D)-Methamphetamine(7) 149.24 185.70 198.28 C6H5·CH2·CH(CH3)·NH·CH3 537-46-2(3)

51-57-0(4)

SH4910000

SH5455000

(L)-Methamphetamine 149.24 185.70 198.28 C6

H5·CH2·CH(CH3)·NH·CH3 33817-09-3(3) SH4905000

Phencyclidine 243.39 279.85 C6

H5·C[C5H10]·N[C5H10] 77-10-1(3)

956-90-1(4)

TN2272600

TN2272600

Phentermine 149.24 185.70 C6

H5·CH2·C(CH3)

2

·NH2 122-09-8(3)

1197-21-3(4)

SH4950000

(DL)-Norephedrine 151.21 187.67 200.25 C6

H5·CH(OH)·CH(CH3)·NH2 14838-15-4(3)

154-41-6(4)

RC2625000

DN4200000

1R,2S (-)-Norephedrine 151.21 187.67 200.25 C6

H5·CH(OH)·CH(CH3)·NH2 492-41-1(3) RC2275000

1S,2R (+)-Norephedrine 151.21 187.67 200.25 C6

H5·CH(OH)·CH(CH3)·NH2 37577-28-9(3)

1S,2S (+)-Norephedrine 151.21 187.67 200.25 C6H5·CH(OH)·CH(CH3)·NH2 36393-56-3

2153-98-2(4)

492-39-7(4)

RC9275000

(D)-Pseudoephedrine(8,9) 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)·NHCH3 90-82-4(3)

345-78-8(4)

UL5800000

UL5950000

(L)-Pseudoephedrine(10) 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)·NH·CH3 321-97-1(3)

(1) Molecular weights are calculated from the empirical formula using the 1987 IUPAC Atomic Weights of the Elements, Merck Index [9]. The molecular 

weight of the hemisulfate is 1⁄2 the weight of the 2:1 sulfate salt (2 moles amine + 1 mole H2

SO4).

(2) CAS from various sources: Merck Index [14], NIOSH RTECS [17], MSDS sheets from Sigma/Aldrich [18], Cerilliant [19], and other sources [21].

(3) Free base form.

(4) Hydrochloride salt.

(5) 2:1 Sulfate salt (2 moles amine + 1 mole H2

SO4.

(6) RTECS = NIOSH Registry of Toxic Effects of Chemical Substances [17].

(7) More active isomer.

(8) Naturally occurring isomer.

(9) The D form of pseudoephedrine is a decongestant.

(10) The L form of pseudoephedrine is a bronchodilator. Dehydroxylation forms the less active L-methamphetamine.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 11 of 30

Table 2. Physical properties of analytes(1)

Compound (alphabetically) CAS m.p.(°C)

Vapor Pressure

(mm Hg) pKa

(4) Log P(5)

Solubility in Water, 

g/100 mL

(DL)-Amphetamine 300-62-9 — — 10.1 @ 20°C 1.76 2.8 @ 25°C

(D)-Amphetamine 51-64-9 <25 — 9.9 1.76 —

(D)-Amphetamine sulfate 51-63-8 >300 — — 6.81 —

(L)-Amphetamine 156-34-3 — 0.201 @ 25 °C 10.1 @ 20°C 1.76 2.8 @ 25°C

Caffeine 58-08-2 238 15 @ 89°C 10.4 @ 40°C -0.07 2.16 @ 25°C

(DL)-Ephedrine 90-81-3 76.5 — — 0.68 —

(L)-Ephedrine 299-42-3 34 0.00083 @ 25°C 10.3 @ 0°C 1.13 6.36 @ 30°C

(L)-Ephedrine HCl 50-98-6 218 2.04E-10 @ 25°C pH 5.9 @ 1/200 dil.(3) -2.45 25(6)

MDEA 82801-81-8 — — — — —

MDMA HCl 42542-10-9 148-149(2) — — — —

(D)-Methamphetamine 537-46-2 — 0.163 @ 25°C 9.87 @ 25°C 2.07 1.33 @ 25°C

(D)-Methamphetamine HCl 51-57-0 170-175(2) — — — —

Phencyclidine 77-10-1 46.5 — 8.29 4.69 —

Phencyclidine HCl 956-90-1 233-235(2) — — — —

Phentermine 122-09-8 — 0.0961 @ 25°C — 1.90 1.86 @ 25°C

Phentermine HCl 1197-21-3 198(2) — — — —

(±) Phenylpropanolamine 14838-15-4 — 0.000867 @ 25°C 9.44 @ 20°C 0.67 14.9 @ 25°C

(±) Phenylpropanolamine HCl 154-41-6 194 — — -2.75 —

(L)-Norephedrine 492-41-1 51-53(3) — — — —

1S,2S (+)-Norephedrine 36393-56-3 77.5-78 0.000867 @ 25°C 9.44 @ 20 0.83 14.9 @ 25

1S,2S (+)-Norephedrine HCl 492-39-7 — — pH 5.9-6.1 in

aq. soln. (3) 0.22 2 @ 25

(D)-Pseudoephedrine 90-82-4 119 0.00083 @ 25°C 10.3 @ 0°C 0.89 10.6 @ 25°C

(D)-Pseudoephedrine HCl 345-78-8 181-182(2) — pH 5.9 @ 1/200 dil.(3) — —

(1) Handbook of Physical Properties of Organic Chemicals unless otherwise noted [21].

(2) Merck Index [14].

(3) Sigma-Aldrich MSDS [18].

(4) Negative log of the acid dissociation constant for the amine in aqueous solution.

(5) Log P = octanol-water partition coefficient.

(6) Temperature not given in source.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 12 of 30

Table 3. Synonyms of analytes

Generic names(1) Trade and street names(2) Additional names(3) 

(DL)-Amphetamine; 

(±)-Amphetamine

Benzedrine; Phenedrine; Bennies (±)-α-Methylbenzeneethanamine(4); dl-α-Methylphenethylamine(4); dl-1-Phenyl-2-

aminopropane; (±)-Desoxynorephedrine

(D)-Amphetamine; 

(+)-Amphetamine

Dextroamphetamine; Dexedrine; 

dexies

(S)-α-Methylbenzeneethanamine(4); d-α-Methylphenethylamine(4); 

d-1-phenyl-2-aminopropane; d-β-Phenylisopropylamine

(L)-Amphetamine; 

(-)-Amphetamine

Levoamphetamine; component of 

Adderall

(R)-α-Methylbenzeneethanamine(4); l-α-Methylphenethylamine(4); 

l-1-phenyl-2-aminopropane; (-)-1-phenyl-2-aminopropane

Caffeine Component (with ephedrine) of 

cloud 9 and herbal XTC 

3,7-Dihydro-1,3,7-trimethyl-1H-purine-2,6-dione(4); 

1,3,7-Trimethylxanthine

(DL)-Ephedrine;

(±)-Ephedrine

Ephedral; Racephedrine; Sanedrine (R*,S*)-(±)-alpha-[2-(Methylamino)ethyl]benzenemethanol; 

DL-alpha-[1-(Methylamino)ethyl]benzyl alcohol; dl-Ephedrine

(L)-Ephedrine;

(-)-Ephedrine;

(1R,2S)-(-)-Ephedrine;

l-Ephedrine

Primatene; Xenadrine; Ma Huang 

(Ephedra sinica and other species(5)); 

(with caffeine) cloud 9 and herbal 

ecstasy

(R-(R*,S*))-α-(1-Methylaminoethyl)benzenemethanol; L-erythro-2-

(Methylamino)-1-phenylpropan-1-ol; (1R,2S)-(-)-2-Methylamino-1-

phenyl-1-propanol; (-)-alpha-(1-Methylamino-ethyl)-benzyl alcohol;

(-)-1-hydroxy-2-methylamino-1-phenylpropane; L-(-)-Ephedrine

(D)-Ephedrine (1S,2R)-(+)-2-Methylamino-1-phenyl-1-propanol; (+)-Ephedrine

MDEA MDE; Eve (±)-3,4-Methylenedioxy-N-ethylamphetamine; 

N-ethyl-alpha-methyl-1,3-benzodioxole-5-ethanamine

MDMA Adam, ecstasy, X, XTC N,α-Dimethyl-3,4-1,3-benzodioxole-5-ethanamine; 

3,4-Methylenedioxymethamphetamine

(DL)-Methamphetamine;

(±)-Methamphetamine

N,α-Dimethylbenzeneethanamine(4); N,α-Dimethylphenethylamine;

dl-Desoxyephedrine; N-methyl-β-phenylisopropylamine

(D)-Methamphetamine;

(+)-Methamphetamine;

d-Methamphetamine

Methedrine; Desoxyn; chalk; crank; 

crystal; glass; ice; meth, speed; upper

(S)-N,α-Dimethylbenzeneethanamine; (S)-(+)-N,α-Dimethylphenethylamine(4); d-1-Phenyl-2-methylaminopropane;

d-Desoxyephedrine; d-N-methyl-β-phenyl-isopropylamine

(L)-Methamphetamine;

(-)-Methamphetamine

Component in decongestant vapor 

inhaler (Vick’s brand)

(R)-(-)-N,α-Dimethylphenethylamine; (-)-Deoxyephedrine; 

(-)-2-(Methylamino)-1-phenylpropane 

Phencyclidine Sernylan; Sernyl; angel dust; PCP; 

peace pill

1-(1-Phenylcyclohexyl) piperidine(4)

Phentermine Fastin; Normephentermine α,α-Dimethylbenzeneethanamine(4); α,α-Dimethylphenethylamine(4);

1,1-Dimethyl-2-phenylethylamine; α-Benzylisopropylamine

(DL)-Norephedrine;

(±)-Norephedrine

(±)-Phenylpropanolamine; Obestat; 

Phenedrine;

(R*,S*)-(±)-α-(1-Aminoethyl)benzenemethanol(4); -(±)-α-(1-Aminoethyl)benzyl alcohol(4); (±)-2-Amino-1-phenyl-1-propanol

(L)-Norephedrine;

(-)-Norephedrine

Natural form found in Ephedra sinica 

and other species(5)

(1R,2S)- 2-Amino-1-phenyl-1-propanol; (1R,2S)-Norephedrine;

l-erythro-2-Amino-1-phenylpropan-1-ol

(D)-Norephedrine;

(+)-Norephedrine

Metabolite of cathinone in urine of 

Khat users.

(1S,2R)- 2-Amino-1-phenyl-1-propanol; (1S,2R)-Norephedrine; 

d-erythro-2-Amino-1-phenylpropan-1-ol

(+)-Norpseudoephedrine;

Cathine

Amorphan; Adiposettin; Reduform; 

found naturally in Khat plant

(R*,R*)-α-(1-Aminoethyl)benzenemethanol(4); d-threo-α-2-Amino-1-

hydroxy-1-phenylpropane; 1S,2S-(+)-Norpseudoephedrine

L-(+)-Pseudoephedrine;

(+)-Pseudoephedrine; 

d-Pseudoephedrine

Afrinol; Novafed; Sinufed; Sudafed; 

natural form found in Ephedra sinica 

and other species(5)

(S-(R*,R*))- α-[1-(Methylamino)ethyl]benzenemethanol; (1S,2S)-

(+)-2-Methylamino-1-phenylpropanol; d-(alpha-(1-Methylamino)-

ethyl)benzyl alcohol; (1S,2S)-(+)-Pseudoephedrine; d-threo-2-

Methylamino-1-phenylpropan-1-ol; (+)-ψ-Ephedrine

D-(-)-Pseudoephedrine;

(-)-Pseudoephedrine

(1R,2R)-(-)-Pseudoephedrine; (-)-ψ-Ephedrine; l-threo-2-

Methylamino-1-phenylpropan-1-ol; (+)-ψ-Ephedrine

(1) Common or generic names. Salts forms are not given for simplicity.

(2) Trade and street names are exemplary, not exhaustive. Street names change over time and by locality. Salts and free base forms are not distinguished.

(3) Other names from Merck Index [14], NIOSH Registry of Toxic Effects of Chemical Substances [17], and MSDS sheets [19] and other reference materials [20]. NOTE: For 

amphetamine and methamphetamine the prefixes R-, D-, d-, and (+)- , although they mean different things, are essentially synonymous for the dextrorotatory 

stereoisomer and S-, L-, l-, and (-) are essentially synonymous for the levorotary stereoisomer. Many other synonyms exist.

(4) Uninverted CAS name as given in Merck Index [14].

(5) Extracts of Ephedra species contain various amounts of (+)-Norephedrine, (-)-N-methylephedrine, and (+)-N-methylpseudoephedrine. (+)-Norephedrine is 

reduced to amphetamine and N-methylephedrine and N-methylpseudoephedrine reduce to N,N-dimethylamphetamine [23, 24]. The presence of these latter two 

compounds in methamphetamine samples indicate that Ephedra spp. extracts may have been used in the synthesis [25].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 13 of 30

Table 4. Limit of detection (LOD), method detection limit (MDL), and sample storage stability on cotton 

 gauze. (1)

Compound Int. std.(2)

Estimated LOD(3)

Scan Mode 

(μg/sample)

SIM Mode 

(μg/sample)

Estimated MDL(4)

Scan Mode 

(μg/sample)

SIM Mode 

(μg/sample)

Storage Stability(5)

30 days 4 °C 7 days 22 °C

(D)-Amphetamine D11-Amp

D14-Met

0.07

0.06

0.05

0.06

0.04

0.03

0.02

0.03

100.5

99.7

94.5

87.9

Caffeine D11-Amp

D14-Met

1

1

0.2

0.2

0.4 (6)

0.4 (6))

0.02

0.03

99.3

98.5

98.8

91.9

(L)-Ephedrine D11-Amp

D14-Met

0.09

0.08

0.1

0.09

0.02

0.01

0.01

0.06

95.6

94.8

97.2

90.5

MDEA N-PAmp 0.05 0.07 0.1 0.02 98.9 102.1

MDMA D11-Amp

D14-Met

0.05

0.05

0.06

0.07

0.04

0.03

0.02

0.02

99.7

98.9

111.1

103.2

(D)-Methamphetamine D11-Amp

D14-Met

0.07

0.05

0.05

0.05

0.03

0.03

0.02

0.02

98.7

98.0

100.6

93.5

Phencyclidine D11-Amp

D14-Met

0.3

0.3

0.06

0.07

0.03

0.03

0.02

0.02

103.7

102.9

105.2

97.7

Phentermine D11-Amp

D14-Met

0.06

0.05

0.06

0.05

0.05

0.05

0.02

0.02

0.01

0.02

0.02

102.0

101.1

101.5

94.3

(±)-Norephedrine(7) D11-Amp

D14-Met

0.2

0.2

(8) 0.1 (9)

0.2 (10)

(8) 94.3

93.6

92.7

86.2 (8)

Pseudoephedrine

D11-Amp

D14-Met

NMPhen

0.08

0.07

0.06

0.07

0.09

0.09

0.03

0.05

0.05

0.02

0.03

0.02

100.4

99.6

-

97.9

91.1

-

(1) Backup Data Report [4].

(2) Internal standards: D11-Amp = Amphetamine-D11, D14-Met = Methamphetamine-D14, NMPhen = N-Methyl phenethylamine, N-PAmp = N-Propyl 

amphetamine.

(3) LODs vary according to individual GC columns, instrument conditions and cleanliness, media interferences, and internal standards used. The lowest 

calibration standard for these determinations was 0.05 μg/sample. Lower LODs are achievable with lower concentration calibration standards and 

operation of the mass spectrometer in the SIM mode. LODs were calculated on liquid standards using the procedure of Burkart [12].

(4) MDLs are provided as an alternate expression of sensitivity. These MDLs are calculated as the standard deviation of six replicates on spiked media 

analyzed at the 0.1 μg/sample level (except as noted) times the Student’s t value for 6 replicates (3.365). (Normally 7 replicates are used.)

(5) Cotton gauze samples were spiked at 3 μg/sample per analyte. Six samples were analyzed immediately after preparation. Six samples were stored at 

room temperature (about 22 °C) for 7 days and then analyzed. Eighteen samples were stored at +4 °C (±2 °C). Of the 18 samples stored at +4 °C, six 

each were analyzed at 7 and 21 days and three each were analyzed at 14 and 30 days. (Backup Data Report [4].) Apparent recoveries vary according to 

internal standard used.

(6) The 0.3 μg/sample level was undetectable in the scan mode. MDLs were calculated from the 1 μg/sample level.

(7) (±)-Norephedrine = (±)-phenylpropanolamine.

(8) (±)-Norephedrine was not evaluated in the SIM mode due to breakdown of derivative with room temperature storage for one week.

(9) MDL calculated from the 0.3 μg/sample level. (Recoveries were >120% at the 0.1 μg/sample level.)

(10) MDL calculated from the 1 μg/sample level. (Recoveries were >120% at the 0.1 and 0.3 μg/sample levels.)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 14 of 30

Table 5. Example of mass spectrometer operation parameters for selected ion monitoring mode.(1)

Chlorodifluoroacetyl derivatives Scan window(2) Acquisition ions (m/z) per group(3)

Acquisition Group 1 10.5 to 13.0 104 118 128 156 160 170 172 177

Acquisition Group 2 13.0 to 15.2 104 156 158 170 172 198 296

Acquisition Group 3 15.2 to 18.0 109 135 162 170 184 194 200 242

GC Peak 

No.(4) Target Analytes and Internal Standards:(5)

Retention Time(6)

(min)

Primary Ion (m/z)

(7) 

(Quantification Ion) 

Secondary ion and approximate relative abundance(8)

(relative to the Primary Ion)

Acquisition Group 1 

2 Amphetamine-D11 (I$)(9) 11.07 160 128 85%

3 Amphetamine 11.15 156 118 85%

5 Phentermine 11.34 170 172 33%

8 n-Methyl phenethylamine (I$)(9) 12.20 156 104 95%

9 Methamphetamine-D14 (I$)(9) 12.51 177 128 32%

10 Methamphetamine 12.61 170 118 32%

Acquisition Group 2:

18 Phenylpropanolamine 13.27 156 246 25%

20 Dibromooctafluorobiphenyl(10) 13.63 296 456 115%

21 N-n-Propylamphetamine (I$)(9) 13.8 198 156 75%

25 Ephedrine 14.27 170 172 33%

28 Pseudoephedrine 14.74 170 172 33%

Acquisition Group 3:

32 Caffeine 15.66 194 109 50%

40 Phencyclidine 16.41 200 242 35%

41 MDMA 16.48 170 162 95%

43 MDEA 16.87 184 162 75%

(1) In this example, 10 analytes and 5 internal standards are grouped into 3 acquisition groups having no more than 8 primary and secondary ions per acquisition 

group. For 6 analytes and internal standards or less, one acquisition group may be sufficient. 

(2) Scan window is in minutes. Actual times are dependant upon GC column and instrument conditions.

(3) Ions (m/z) in bold numbers are suggested primary (quantification) ions. For best signal to noise ratio, do not exceed 10 ions per acquisition group. Dwell times per 

ion (m/z) is 50 milliseconds.

(4) GC peak numbers are those in Figure 1 and Table 9.

(5) The list of analytes and internal standards shown is an example. Analyte(s) and internal standard(s) must be selected according to analytical objectives. 

(6) Retention times are dependant upon GC column and instrument conditions.

(7) The better ions for quantification are usually the base peak or those with masses >100 m/z and relative abundances >50% of the base peak. These minimize 

interference from co-eluting hydrocarbons. The suggested primary ions are not necessarily the base peaks in the mass spectra of the analytes, especially if the 

base peaks are ions common to aromatics (e.g. m/z 91) and paraffinic or olefinic hydrocarbons (e.g. m/z 42, 57, and 58). Suggested ions for other analytes and 

internal standards are given in Tables 8 and 9.

(8) Secondary ions may be used for quantification if the primary ion encounters interference. Secondary ions improve qualitative identification for SIM analyses. The 

relative abundances given are approximate (±10 to 20%) and depend upon specific instrument tuning and conditions. They are relative to the primary ion and 

not necessarily to the base peak in the mass spectrum of each analyte. The relative abundance of secondary ions for each analyte needs to be determined from a 

mass spectrum acquired on the instrument to be used.

(9) (I$) = internal standard. Internal standards must be paired with the appropriate analytes. Tables 7a and 7b give precision and accuracy data for various pairings. 

Other potentially useful internal standards are given in Table 8. Highly deuterated analogs of the target analytes are preferred, where available.

(10) Dibromooctafluorobiphenyl is an optional secondary internal standard useful for monitoring autosampler performance and instrument tuning. A shift in the mass 

axes or the relative abundance of m/z 296 to that of m/z 456 throughout an analytical sequence will help signal degraded tuning.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 15 of 30

Table 6. Suggested spiking schedule for calibration standards and quality control samples

Calibration Standards(10)

Add the following to clean shipping containers 

(e.g. 50-mL polypropylene centrifuge tubes) in the following order.

Number 

of 

Wipes(1,2)

Volume(2) of 

Isopropanol 

or Methanol(3)

Volume(2)

of Internal 

Standard 

Spiking 

Solution(4,5)

Volume 

of Target 

Analyte 

Spiking 

Solution(5,6)

Volume of 

Spiking Solution 

diluted 1/20(5,7)

Volume(2) of 

Desorption 

Solution(8)

Resulting 

μg/sample 

as Free 

Base(9)

CS0 0 3 mL 60 μL 0.0 μL 30 mL 0.00

CS1 0 3 mL 60 μL 2 μL 30 mL 0.02

CS2 0 3 mL 60 μL 5 μL 30 mL 0.05

CS3 0 3 mL 60 μL 10 μL 30 mL 0.1

CS4 0 3 mL 60 μL 20 μL 30 mL 0.2

CS5 0 3 mL 60 μL 60 μL 30 mL 0.6

CS6 0 3 mL 60 μL 10 μL 30 mL 2.0

CS7 0 3 mL 60 μL 30 μL 30 mL 6.0

CS8 0 3 mL 60 μL 100 μL 30 mL 20

CS9 0 3 mL 60 μL 300 μL 30 mL 60

CS10 0 3 mL 60 μL 1000 μL 30 mL 200

Quality Control Samples(11)

QB (media blank) 1 3 mL 60 μL 0.0 μL 30 mL 0.0

QC (matrix spike) 1 3 mL 60 μL 3-300 μL or 20-60 μL 30 mL 0.2-60

QD (matrix spike duplicate) 1 3 mL 60 μL 3-300 μL or 20-60 μL 30 mL 0.2-60

(1) Gauze wipes may be added to the calibration standards but are not necessary if cotton gauze is used. Blank gauze wipes must always be added to the 

quality control samples, QB, QC, and QD.

(2) a.) If a sample consists of 2 gauze wipes, the volume of desorption solution must be increased to 40 mL to accommodate the second wipe. The shipping 

container should be a 50-mL polypropylene centrifuge tube or equivalent to accommodate the extra volume of desorption solution for 2 wipes. It is not 

critical to know the exact volume of desorption solution and wetting alcohol used per sample. It only needs to be enough to cover the samples and to 

permit free percolation through the samples. See step 7.

b.) If a set of samples consists predominantly of 2 gauze wipes, the QB, QC, and QD should also consist of 2 wipes and treated as per the samples. The 

volume of isopropanol (or methanol) added to the QC samples should be increased to 4 mL for two gauze wipes to simulate samples containing two 

gauze wipes. 

(3) If methanol was used for wipe sampling, it should also be used in the calibration standards, blanks, and QCs instead of isopropanol. 

(4) Concentration of internal standards in the internal standard spiking solution is approximately 200 μg/mL as the free base. It is critical to know the exact 

volume of internal standard spiking solution that is added to the calibration standards, samples, blanks, and quality control samples. The volume spiked 

into the samples may vary with sample size but the volume spiked into each of the calibration standards must not vary. See step 7b.

(5) For quality control samples, spike onto wipe media within the shipping container. For liquid calibration standards (in lieu of media calibration standards), 

spike into the isopropanol (or methanol).

(6) Concentration of analytes in the target analyte spiking solution is approximately 200 μg/mL as the free base.

(7) Concentration of analytes in the diluted spiking solution for this table is approximately 10.0 μg/mL as the free base and can be prepared by diluting 

100 μL of the target analyte spiking solution to 2 mL in methanol.

(8) Desorption solution is 0.1 M sulfuric acid in deionized water.

(9) This is μg per total sample irrespective of the total desorption solution volume or the area wiped.

(10) Select 6 calibration standards from the list to cover the analytical range plus the blank.

(11) Prepare one set of quality control samples for every 20 samples or less.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 16 of 30

Table 7a. Precision and accuracy in scan mode.(1)

Compound

Internal 

Standard(2)

Range(3)

μg/sample Accuracy

Overall 

Precision 

Bias

Average Range

(D)-Amphetamine

D11-Amp 0.1-30 17.1 0.0670 -0.0613 -0.1048 - -0.0170

D14-Met 0.1-30 13.7 0.0610 +0.0338 -0.0151 - +0.1056

NMPhen 0.1-30 12.5 0.0559 -0.0310 -0.0651 - +0.0177

Caffeine

D11-Amp 1.0-30 20.0 0.0708 -0.0832 - 0.1476 - -0.0542

D14-Met 1.0-30 12.5 0.0636 -0.0014 - 0.0274 - +0.0381

NMPhen 1.0-30 15.6 0.0796 -0.0040 -0.0789 - +0.1321

(L)-Ephedrine

D11-Amp 0.1-10 15.4 0.0627 +0.0510 -0.0148 - +0.1128

D14-Met 0.3-10 17.8 0.0674 +0.0666 +0.0261 - +0.1660

NMPhen 0.3-30 15.0 0.0707 +0.0293 -0.0259 - +0.0973

MDEA N-PAmp 0.3-29 16.6 0.0817 -0.0224 -0.0656 - +0.0657

MDMA

D11-Amp 0.3-27 20.2 0.0778 -0.0739 -0.1011 - -0.0489

D14-Met 0.3-27 16.6 0.0652 +0.0589 -0.0947 - +0.0036

NMPhen 0.3-27 22.0 0.0722 -0.1017 -0.1486 - -0.0315

(D)-Methamphetamine

D11-Amp 0.1-30 14.7 0.0631 -0.0435 -0.0657 - -0.0060

D14-Met 0.1-30 12.5 0.0546 -0.0348 -0.1144 - +0.0188

NMPhen 0.1-10(5) 14.9 0.0503 -0.0665 -0.1179 - +0.0110

Phencyclidine

D11-Amp 0.1-10 18.2 0.0690 -0.0683 -0.1257 - -0.0136

D14-Met 0.3-3 13.4 0.0465 -0.0577 -0.0662 - -0.0493

NMPhen 0.3-10 16.8 0.0609 -0.0682 -0.1137 - +0.0091

Phentermine

D11-Amp 0.1-30 15.2 0.0486 -0.0720 -0.1010 - +0.0291

D14-Met 0.1-30 10.7 0.0509 +0.0190 -0.0395 - +0.0671

NMPhen 0.1-30 9.6 0.0420 -0.0269 -0.0612 - +0.0340

(±)-Norephedrine(4) D11-Amp 1-30 6.5 0.0328 +0.0061 -0.0070 - +0.0248

Pseudoephedrine

D11-Amp 0.3-30 17.2 0.0571 -0.0783 -0.1273 - -0.0560

D14-Met 0.3-30 14.9 0.0649 -0.0422 -0.0888 - +0.0395

NMPhen 0.3-30 18.7 0.0488 -0.1068 -0.1505 - -0.0422

(1) Backup Data Report [4]. Values are for chlorodifluoroacetyl derivatives and analysis by GC-MS in scan mode. Each sample consisted of a pair of 3” x 3” 12-ply cotton 

gauze pads. There were 6 replicate samples per concentration level and six concentration levels evaluated from approximately 0.1 to 30 μg/sample. 

(2) Internal Standards Deuterated: Non-deuterated:

 D11-Amp = Amphetamine-D11 NMPhen = N-Methyl phenethylamine

 D14-Met = Methamphetamine-D14 N-PAmp = N-Propyl amphetamine

(3) Range used for calculation of precision, accuracy, and bias. The entire range studied for all analytes was approximately 0.1 to 30 μg/sample (1xLOQ to 300xLOQ).

(4) (±)-Norephedrine = (±)-phenylpropanolamine.

(5) One or more higher level concentrations were omitted from the computations due to inlier CVs (<0.0200.)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 17 of 30

Table 7b. Precision and accuracy in SIM mode.(1)

Compound

Internal 

Standard(2)

Range(3) μg/

sample Accuracy(4)

Overall(4)

Precision 

Bias

Average Range

(D)-Amphetamine D11-Amp 0.1-30 14.3 0.0412 -0.0750 -0.1153 - -0.0351

D14-Met 0.1-30 10.1 0.0508 -0.0074 -0.0500 - +0.0389

NMPhen 0.1-30 13.3 0.0439 -0.0606 -0.1117 - -0.0318

Caffeine D11-Amp 0.1-30 21.3 0.0578 -0.1182 -0.1949 - -0.0697

D14-Met 0.1-30 14.4 0.0534 -0.0558 -0.1061 - -0.0170

NMPhen 0.3-30 19.8 0.0387 -0.1338 -0.1775 - -0.0820

(L)-Ephedrine D11-Amp 0.3-30 9.1 0.0421 -0.0199 -0.0423 - +0.0157

D14-Met 0.3-30 20.5 0.0503 0.1226 +0.0637 - +0.1883

NMPhen 0.3-30 10.2 0.0449 +0.0260 -0.0075 - +0.0769

MDEA N-PAmp 0.3-29 10.3 0.0264 -0.0597 -0.0879 - -0.0095

MDMA D11-Amp 0.1-27 16.2 0.0503 -0.0750 -0.1423 - -0.0292

D14-Met 0.1-0.9(5) 15.4 0.0503(6) -0.0712 -0.1247 - +0.0032

NMPhen 0.1-27 15.4 0.0496 -0.0722 -0.1136- -0.0108

(D)-Methamphetamine D11-Amp 0.1-10(5) 16.5 0.0379 -0.1030 -0.1414 - -0.0660

D14-Met 0.1-30 9.2 0.0351 -0.0343 -0.0767 - +0.0006

NMPhen 0.1-30 13.6 0.0322 -0.0827 -0.1221 - -0.0403

Phencyclidine D11-Amp 0.1-10(5) 17.7 0.0428 -0.1068 -0.1303 - -0.0586

D14-Met 0.1-3 11.3 0.0450 -0.0393 -0.0683 - -0.0205

NMPhen 0.1-3(5) 16.1 0.0449 -0.0871 -0.1279 - -0.0383

Phentermine D11-Amp 0.1-30 12.8 0.0394 -0.0637 -0.0982 - -0.0433

D14-Met 0.1-30 9.8 0.0495 -0.0051 -0.0375 - +0.0556

NMPhen 0.1-30 11.0 0.0394 -0.0451 -0.0766 - -0.0163

Pseudoephedrine D11-Amp 0.3-30 17.3 0.0402 -0.1073 -0.1496 - -0.0514

D14-Met 0.3-30 11.7 0.0519 -0.0294 -0.0559 - +0.0532

NMPhen 0.3-30 17.0 0.0450 -0.0956 -0.1197 - -0.0576

(1) Backup Data Report [4]. Values are for chlorodifluoroacetyl derivatives and analysis by GC-MS in SIM mode (see Table 5 for MS conditions). Each 

sample consisted of a pair of 3” × 3” (7.5 cm x 7.5 cm) 12-ply cotton gauze pads. There were 6 replicate samples per concentration level and six 

concentration levels evaluated from approximately 0.1 to 30 μg/sample. Norephedrine (phenylpropanolamine) was not evaluated in the SIM mode 

due to breakdown at room temperature storage for several days prior to analysis.

(2) Internal Standards, Deuterated: Non-deuterated:

 D11-Amp = Amphetamine-D11 NMPhen = N-Methyl phenethylamine

 D14-Met = Methamphetamine-D14 N-PAmp = N-Propyl amphetamine

(3) Range used for calculation of precision, accuracy, and bias. The entire range studied for all analytes was approximately 0.1 to 30 μg/sample (1×LOQ to 

300×LOQ).

(4) NIOSH [1995]. NIOSH Technical Report: Guidelines for Air Sampling and Analytical Method Development and Evaluation. By Kennedy ER, Fischbach TJ, 

Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S. Department of Health and Human Services, Public Health Services, Centers for Disease Control and 

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 95-117.

(5) One or more higher level concentrations were omitted from the computations due to inlier CVs (<0.0200).

(6) The overall precision, ŜrT , is an estimate due to inlier precisions (<0.02) at several higher concentration levels.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 18 of 30

Table 8a. Recommended internal standards and best application(1)

COMPOUND NAME CAS

MW as

free base

Quant. 

Ion

Secondary 

Ion COMMENTS

(±)-Amphetamine-D11 Not available 146.12 160 128 Preferred analog for amphetamine

(±)-Amphetamine-D8 145225-00-9 143.15 126(3) 159(3) Alternate for amphetamine-D11

(±)-Amphetamine-D6 Not available 141.16 160 123 Alternate for amphetamine-D11

(±)-Methamphetamine-D14 Not available 163.12 177 128 Preferred methamphetamine analog

(±)-Methamphetamine-D11 152477-88-8 160.15 176 126 Alternate for methamphetamine-D14

(±)-Methamphetamine-D9 Not available 158.16 177 123 Alternate for methamphetamine-D14

N-Methylphenethylamine 589-08-2 135.23 156 104 Alternate for methamphetamine-D14

Phencyclidine-D5 60124-86-9 248.35 205 247 Use only for phencyclidine.

MDEA-D6

(2) 160227-44-1 213.22 190 165 Use only for MDEA.

N-Propylamphetamine(2) Not available 177.29 198 156 Alternate for MDEA-D6

(1) Care must be exercised in the selection of internal standards for each analyte because of differences in derivatization efficiencies due to structural differences.

a. Deuterated analogs of each target analyte may be acceptable as internal standards if they are isotopically pure enough and their ions do not interfere with 

the quantification ions (usually base peaks) of the target analyte, especially at the limit of detection for the target analyte. Conversely it is also important 

that ions in the target analyte, especially at high concentrations, do not interfere with the quantification ion (usually base peaks) of any deuterated analog 

used as the internal standard.

b. The more highly deuterated an analog, the more it will chromatographically separate from the target analyte, reducing interference from common ions.

c. Phentermine and mephentermine have been used as internal standards. Such use is not advised in this method because of their reported occasional use as 

adulterants in certain illicit drugs such as MDMA.

(2) N-Propylamphetamine and MDEA-D6

 are only applicable to MDEA and other hindered amines (e.g. fenfluramine and MBDB) due to similar steric hindrance at 

the nitrogen (N-ethyl or N-propyl substitution) which affects derivatization efficiency.

(3) It is better to use m/z 126 because at high concentration levels unlabelled amphetamine contributes significant interference to m/z 159 of amphetamine-D8

.

Table 8b. Recommended best application of internal standards

Recommended Deuterated

Internal Standards

Recommended Alternate

Non-deuterated Internal Standards(3)

TARGET ANALYTE

Amphetamine-D11

(2)

Methamphetamine-D14

(2) MDEA-D6(1)

Phencyclidine-D5

N-Methylphenethylamine

4-Phenyl-1-

butyl-amine

N-Propyl-amphetamine(1)

Amphetamine X X X

Caffeine X X

Ephedrine X X X X

MDEA X X

MDMA X X

Methamphetamine X X X

Phencyclidine X X X

Phentermine X X

(±)-Norephedrine(4) X X

Pseudoephedrine X X

(1) N-Propylamphetamine and MDEA-D6 are only applicable to MDEA and other hindered amines (e.g. fenfluramine and MBDB) due to similar steric hindrance at the 

nitrogen (N-ethyl or N-propyl substitution) which affects derivatization efficiency.

(2) The alternate deuterated compounds listed in part A above may be used. Avoid ring-labeled amphetamine-D5

 (CAS 65538-33-2) since the primary (quantification) 

ion is the same as for amphetamine and GC peaks overlap significantly. Also avoid methamphetamine-D5

 (CAS 60124-88-1) since GC peaks significantly overlap 

and secondary ions for the chlorodifluoroacetyl derivative are not baseline resolved.

(3) The listed non-deuterated compounds are effective as internal standards for the listed target analytes. Non-deuterated internal standards might not be 

permissible.

(4) (±)-Norephedrine is the same as (±)-phenylpropanolamine.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 19 of 30

Table 9. Gas chromatographic retention times for chlorodifluoroacetyl derivatives of amphetamines, 

precursors, adulterants, and miscellaneous drugs of abuse.(1)

GC 

Peak 

No.(2) Compound

Recommended Quantification 

(1’) and Confirmation(2’, 3’) 

Ions (m/z)

(3)

1’ 2’ 3’ Form(4)

Retention 

Time in 

Minutes

Relative 

Retention 

Time(5)

Relative 

Retention 

Time(6)

1 Nicotine 84 133 162 parent 8.92 0.396 0.757

2 (DL)-Amphetamine-D11(I$)(7) 160 128 162 derivative 10.26 0.800 0.870

3 (DL)-Amphetamine 156 118 158 derivative 10.34 0.807 0.877

4 Phenethylamine(8) 104 91 - derivative 10.38 0.810 0.880

5 Phentermine(8) 170 172 132 derivative 10.52 0.821 0.892

6 N-Methyl pseudoephedrine(9) 134 162 75 derivative 10.54 0.822 0.894

7 N-Methyl pseudoephedrine(9) 72 - - parent abt 11 0.86 0.93

8 N-Methyl phenethylamine (I$)(7) 156 104 158 derivative 11.37 0.887 0.964

9 (DL)-Methamphetamine-D14 (I$)(7) 177 98 179 derivative 11.70 0.913 0.992

10 (DL)-Methamphetamine 170 172 118 derivative 11.79 0.920 1.000

11 Fenfluramine(8) 184 186 159 derivative 11.83 0.923 1.003

12 S-(-)-Cathinone (from Khat plant) 105 77 132 derivative 11.99 0.935 1.017

13 Bupropion (Wellbutrin®, Zyban®) 44 100 111 parent 12.14 0.947 1.030

14 N-Ethyl amphetamine 184 186 118 derivative 12.22 0.953 1.036

15 Ecgonine, methyl ester 182 82 311 derivative 12.36 0.964 1.048

16 S-(-)-Methcathinone (“Cat”) 170 105 172 derivative 12.38 0.966 1.050

17 Norpseudoephedrine (Cathine) 156 158 246 bis-derivative 12.46 0.972 1.057

18 (±)-Norephedrine 156 158 246 bis-derivative 12.49 0.974 1.059

19 Aminorex 107 79 232 derivative (-CN) 12.70 0.991 1.077

20 Dibromooctafluorobiphenyl (I$)(7) 296 456 454 parent 12.82 1.000 1.087

21 N-Propyl amphetamine (I$)(7) 198 156 200 derivative 12.97 1.012 1.100

22 4-Methoxyamphetamine 121 148 78 derivative 13.22 1.031 1.121

23 4-Phenyl-1-butylamine (I$)(7) 176 104 - derivative 13.27 1.035 1.126

24 1S,2R(+)-Ephedrine-D3

 (I$)(7) 173 175 85 derivative 13.44 1.048 1.140

25 (DL)-Ephedrine 170 172 260 bis-derivative 13.48 1.052 1.143

26 Acetaminophen(8) 108 221 263 derivative 13.67 1.066 1.159

27 Methyl phenidate (Ritalin®) 84 56 91 parent 13.81 1.077 1.171

28 Pseudoephedrine 170 172 260 bis-derivative 13.93 1.087 1.182

29 Meperidine (Demerol® etc.) 71 247 172 parent 13.99 1.091 1.187

30 Atropine 124 94 103 parent (-H2O) 14.25 1.112 1.209

31 (±)-MDA 135 162 291 derivative 14.36 1.120 1.218

32 Caffeine(8) 194 109 67 parent 14.84 1.158 1.259

33 N,N-Dimethyltryptamine (DMT) 58 129 102 derivative 14.97 1.168 1.270

34 (±)-BDB 135 176 170 derivative 15.11 1.179 1.282

35 Ketamine (“special K”)(8,11) 180 182 209 parent 15.20 1.186 1.289

36 Lidocaine(8) 86 58 120 parent 15.28 1.192 1.296

37 Trifluoromethylphenyl piperazine(11) 200 145 172 derivative 15.46 1.206 1.318

38 Benzyl piperazine(11) (“Legal XTC”) 91 197 175 derivative 15.54 1.202 1.318

39 Phencyclidine-D5

 (I$)(7) 205 96 246 parent 15.59 1.216 1.322

40 Phencyclidine (PCP) 200 242 243 parent 15.62 1.218 1.325

41 MDMA(11) 170 162 135 derivative 15.66 1.221 1.328

Table 9 continued,
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 20 of 30

Table 9, continued. Gas chromatographic retention times for chlorodifluoroacetyl derivatives of 

amphetamines, precursors, adulterants, and miscellaneous drugs of abuse.(1)

GC 

Peak 

No. (2) Compound

Recommended Quantification (1’) 

and Confirmation

(2’, 3’) Ions (m/z)

(3)

1’ 2’ 3’ Form(4)

Retention Time 

Minutes

Relative 

Retention 

Time(5)

Relative 

Retention 

Time(6)

42 MDEA-D6

 (I$)(7) 190 165 135 derivative 16.01 1.249 1.358

43 MDEA(11) 184 162 135 derivative 16.04 1.251 1.360

44 Phenylephrine(8) 156 158 374 tris-derivative 16.10 1.256 1.366

45 (±)-MBDB 184 176 135 derivative 16.29 1.271 1.382

46 Theophylline(8) 180 95 68 parent 16.34 1.275 1.386

47 Mescaline 181 194 179 derivative 16.43 1.282 1.394

48 Phenylephrine(8) 156 248 158 bis-derivative 16.65 1.299 1.412

49 Chlorpheniramine(8) 203 205 167 parent 16.73 1.305 1.419

50 Methyl phenidate 196 198 - derivative 17.20 1.322 1.459

51 4-Bromo-2,5-DMPEA(10) (Nexus) 242 244 229 derivative 17.57 1.370 1.490

52 cis-(±)-4-Methylaminorex (“U4Euh”) 203 160 117 derivative 17.89 1.396 1.517

53 Dextromethorphan(8) 271 59 150 parent 18.09 1.411 1.534

54 Methaqualone 235 250 233 parent 18.27 1.425 1.550

55 Cocaine 82 182 303 parent 18.62 1.452 1.579

56 Atropine(8) 124 82 94 derivative 19.10 1.490 1.620

57 Diazepam (Valium® etc.) 256 283 284 parent 20.76 1.619 1.761

58 Hydrocodone (Lortab® etc.) 299 242 284 parent 20.91 1.631 1.774

59 Hydromorphone (Dilaudid®) 285 228 229 parent 21.04 1.641 1.785

60 Hydrocodone (Lortab® etc.) 411 354 298 derivative 21.13 1.648 1.792

61 Morphine 268 397 269 derivative 21.20 1.654 1.798

62 Codeine 282 411 283 derivative 21.28 1.660 1.805

63 Oxycodone (OxyContin®) 315 230 316 parent 21.57 1.682 1.830

64 Hydromorphone (Dilaudid®) 397 341 398 derivative 21.78 1.699 1.847

65 Flunitrazepam (Rohypnol®, roofies)(11) 312 285 286 parent 22.19 1.731 1.882

66 Morphine 380 382 509 bis-derivative 22.26 1.736 1.888

67 Fentanyl (Sublimaze® etc.) 245 146 189 parent 22.96 1.791 1.947

(1) Actual retention times may vary depending on individual GC column and GC conditions. Gas chromatographic conditions used are on p. 9106-1. The mass 

spectrometer was operated under the conditions given on p 9106-1 (or see the Backup Data Report [4].)

(2) GC peak numbers represent peaks as numbered in Figure 1.

(3) Use extracted ion chromatograms of the primary ions (1’) for quantifying peaks in either the scan mode or the SIM mode. Use the secondary and tertiary ions (2’ 

and 3’) for qualitative identification when necessary. These ions are selected for nearness to the primary ion to minimize false negatives from skewed spectra and 

from low mass interference from hydrocarbons.

(4) Not all forms are presented. Parent compounds are not presented that have irregular or overly broad GC peak shapes under the GC conditions used. Spectra for 

chlorodifluoroacetyl derivatives are given in the Backup Data Report [4]

(5) Retention time relative to 4,4’-dibromooctafluorobiphenyl.

(6) Retention time relative to the chlorodifluoroacetyl derivative of methamphetamine.

(7) I$ = Internal standard.

(8) Intentional or unintentional adulterants. For example, phentermine may be added to MDMA and caffeine added to methamphetamine. Chlorpheniramine is an 

unintentional adulterant when pseudoephedrine containing chlorpheniramine is used as a methamphetamine precursor.

(9) Presence of (+)-Norephedrine, N-methylpseudoephedrine and/or N-methylephedrine in pseudoephedrine or ephedrine indicates extracts of Ephedra species 

(spp.) as source. Presence of amphetamine and N,N-dimethylamphetamine in methamphetamine final product also indicates the same source.[22, 23, 24]

(10) 4-Bromo-2,5-dimethoxyphenethylamine

(11) Typical “club drugs” (piperazine analogs as ecstasy substitutes, ketamine and flunitrazepan as predatory drugs).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 21 of 30

Table 10a. Recovery from latex-painted wall with various solvents; one gauze wipe compared with the 

sum of two gauze wipes(1,2)

Test Compound

Water(3)

First Gauze Wipe

Plus Second 

Wipe(4)

Percent %RSD Percent

Isopropanol

First Gauze Wipe

Plus Second 

Wipe(4)

Percent %RSD Percent

Methanol

First Gauze Wipe

Plus Second 

Wipe(4)

(5) Percent %RSD Percent

Amphetamine 51 14 56 67 6.0 78 90 4.0 96

Cocaine 36 22 36 69 22 80 89 9.1 94

Ephedrine 48 23 52 76 7.4 85 91 4.4 96

MDMA 40 20 44 61 9.0 70 88 5.3 94

MDEA 45 22 50 69 12 80 90 11 97

Methamphetamine 46 16 50 64 7.4 75 87 3.5 94

Phencyclidine 27 26 30 64 9.6 73 86 5.2 91

Phentermine 53 9.2 58 78 6.6 91 95 2.9 101

Phenylpropanolamine 58 21 62 80 9.3 95 85 5.0 94

Pseudoephedrine 49 20 53 73 7.0 85 95 3.3 101

(1) Backup Data Report for NIOSH 9109 [13]. Area of each sample was 100 cm2.

(2) Wall was an existing standard gypsum board wall painted with a latex based paint. Painted surface was at least one year old. There were six replicates for each 

solvent tested.

(3) Water was deionized water (ASTM type II). Note low recovery and high %RSD.

(4) For the serial wipe study, each 100-cm2

 area was wiped again with a fresh pre-wetted gauze wipe and the amount recovered was determined separately. In 

practice, a second (serial) wipe is included with the first gauze wipe; both gauze wipes constitute a single sample. The percent recoveries shown in the column 

represent the sum of the amounts recovered in both the first and second wipes.

(5) Each pre-measured area was spiked with 3 μg of each analyte in methanol and the methanol allowed to dry for several minutes prior to wipe sampling.

Table 10b. Recovery from various surfaces with various solvents; one gauze wipe compared with the 

sum of two gauze wipes(1)

Isopropanol Methanol

First Gauze Wipe Plus Second Wipe(2) First Gauze Wipe Plus Second Wipe(2)

Surface Material(3) Replicates Percent %RSD Percent Percent %RSD Percent

Enamel (lid of washing machine) 4(3) 58 5.7 68 81 2.4 87

Vinyl veneer on particle board 4(4) 60 5.2 68 81 4.8 89

Latex painted wall 6(3) 64 7.4 75 87 3.5 94

Refrigerator door 2(4) 65 2.9 76 91 4.0 92

Varnished hardwood panel 2(5) 72 5.4 76 82 3.7 86

Formica® countertop 4(4) 75 4.9 82 87 3.8 91

(1) Backup Data Report for NIOSH 9109 [13]. Area of each sample was 100 cm2.

(2) For the serial wipe study, each 100-cm2

 area was wiped again with a fresh pre-wetted gauze wipe and the amount recovered was determined separately. In 

practice, a second (serial) wipe is included with the first gauze wipe; both gauze wipes constitute a single sample. The percent recoveries shown in the column 

represent the sum of the amounts recovered in both the first and second wipes.

(3) The Refrigerator door and the washing machine lid were from used appliances. The vinyl-veneered particle board (a book shelf), the Formica® countertop, and 

the varnished hardwood paneling were all purchased new. All surfaces of used and new materials were pre-cleaned with multiple rinses of methanol prior to 

spiking. Each pre-measured 100-cm2 square was spiked with 3 μg methamphetamine.

(4) Samples were taken using the side-to-side and then top-to-bottom wiping technique. 

(5) Half of the samples were wiped using the side-to-side wiping technique and half were wiped using the concentric squares wiping technique. There were no 

significant differences in recoveries. Percent recoveries and %RSDs are for both techniques combined. 

(6) Samples were taken each time using only top-to-bottom wiping with the grain of the wood in an “N” pattern.
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Figure 1. Typical chromatograms of chlorodifluoroacetyl derivatives by GC-MS in scan mode

Figure 1a. Extracted ion chromatogram for m/z 156 (155.70 to 156.70). 

Figure 1b. Extracted ion chromatogram for m/z 170 (169.70 to 170.70).

Figure 1c. Total ion chromatogram (TIC).

GC Peak Identification: See Table 9 for identification of numbered GC peaks. (But note that retention times in Table 9 do not 

correspond to those in Figure 1 because a different 0.5 μm film phenyl arylene polymer capillary column was used.)

GC-MS Conditions: See p. 9106-1 for GC-MS conditions. 
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APPENDIX

A. REAGENTS and SOLUTIONS:

1. For derivatization, pentafluoropropionic anhydride (PFPA) may be substituted for 

chlorodifluoroacetic anhydride (CDFAA). Spectra, retention times, suggested quantification ions, and 

precision and accuracy data for PFPA derivatives are given in the Backup Data Report [4]. Spectra for 

CDFAA derivatives are also given in the Backup Data Report [4].

NOTE: 100 μL of pentafluoropropionic anhydride (PFPA) may be substituted for chlorodifluoroacetic 

anhydride, but the samples must be heated to 90 °C for 20-30 minutes in step 11c.

2. The instrumental internal standard, 4,4’-dibromooctafluorobiphenyl is optional. It is useful for 

monitoring instrument tuning and autosampler performance.

3. Primary amines form Schiff bases and enamines with ketones and aldehydes. These may in turn 

form derivatives with the acylating reagents. The use of acetone must be avoided strictly prior to 

the analytes being derivatized. Glassware and equipment rinsed with acetone must be thoroughly 

dried. Toluene should be avoided for making up standard solutions because it usually contains 

benzaldehyde, an oxidation product of toluene. Condensation products have been observed 

between primary amines and benzaldehyde. The only solvents recommended for the preparation of 

stock solutions and dilutions thereof are methanol (preferably) and isopropanol.

4. The reconstitution solvent should not contain methanol or other alcohol since the derivatized 

alcohol groups in ephedrine type compounds are hydrolyzed over time. Toluene containing 10 

percent acetone is recommended.

B. EQUIPMENT:

1. Wipe media: Besides cotton gauze, 4”×4” (10 cm x 10 cm) 4-ply MIRASORB® (Johnson and Johnson), 

and 4”×4” (10 cm x 10 cm) AlphaWipe® (TX® 1004, Texwipe Corp.) were acceptable wipe media and 

can be used in the absence of cotton gauze. MIRASORB®, a non-woven cotton/polyester blend, is 

discontinued but counterparts exist that claim to be of identical construction and fiber composition.

AlphaWipe® is a hydrophilic, highly adsorbent, tightly knitted continuous filament polyester wipe.

Precision and accuracy data for MIRASORB® and AlphaWipe® are given in the Backup Data Report [4].

2. Shipping containers: The 50-mL polypropylene centrifuge tubes with caps are preferred for one 

or two gauze wipes and are not as breakable as glass 40-mL VOA vials. The 40-mL VOA vials are 

acceptable for single gauze wipes. Larger containers (glass with a PTFE lined cap) should be used 

for combining more than two gauze wipes into a single sample. The size of the container for two or 

more wipes should be approximately 25 mL per gauze wipe (e.g. a minimum size of 100-mL for up 

to four gauze wipe samples). There needs to be enough extra headspace in the shipping container 

to allow the desorption solution to cover the gauze wipes and to percolate freely through the wipe 

sample(s) during mixing.

3. Each regulatory agency having legal jurisdiction over the contaminated site may require different 

but specific off-site preparation and on-site sampling procedures. It is important to consult local 

regulatory agencies or departments of health having legal jurisdiction over contaminated sites to 

determine specific sampling, quality control, analyses, and reporting requirements.

C. SAMPLING:

1. Follow specific requirements of surface area to be wiped (usually 100 cm2

 or 1000 cm2) and action 

threshold (or maximum allowable residual level) set by the state or specified by the client. Uptake 

rates depend upon the wipe sampling method used, so the specific wipe technique used must be 

specified, and any deviations from the required wipe sampling requirements noted.

NOTE: To ensure that samples have not been tampered with, the use of custody seals and a chainof-custody form is strongly recommended.

2. Prepare a rigid template from disposable cardstock or a sheet of PTFE having either a 10 cm × 10 cm 

or 32 cm × 32 cm square-cut hole. The template must be able to retain its shape during wiping to 

ensure that the areas wiped were either 100 cm2

 or 1000 cm2. Secure the template(s) to the area(s) 
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to be wiped (e.g. with tape along outside edge of template). If a single-use disposable template 

is not used, clean the template between samples to avoid cross-contamination, and provide the 

laboratory with a blank wipe of the cleaned template between samples to determine that no crosscontamination has occurred. 

3. A template might not always be applicable, as in curved or odd-shaped areas such as around burners 

on stove tops or a fan blade. In such cases sample an area as close to either 100 cm2

 or 1000 cm2 as 

feasible and provide the measurement to the regulatory agency and to the analytical laboratory for 

proper reporting. Tape can be used to delineate the sampling area.

4. It is recommended to provide extra wipe media from the same lot for required media blanks, field 

equipment blanks, samples, and quality-control samples.

5. Gauze in sterile packaging is recommended to minimize the chance for cross-contamination, which 

can more easily occur with open bulk packaged cotton gauze.

6. To prevent contamination in the field, another alternative is to pre-wet and insert the gauze 

wipes into the sample containers off-site. This avoids any possibility of the bottle of methanol or 

isopropanol becoming contaminated on-site with methamphetamine (or other analytes). If the wipes 

were prepared off-site, then remove pre-wetted gauze wipe from sample container, opening only 

one sample container at a time. In either case, squeeze out and discard any excess solvent from the 

gauze wipe. Use fresh latex or nitrile gloves for each separate sample and blank. Do not use vinyl 

gloves due to the potential for leaching of phthalate plasticizers and contamination of the samples.

7. Wipe techniques

a. Concentric Squares Wiping Technique (particularly suitable for smooth and non-porous surfaces): 

Fold the pre-wetted gauze in half and then fold in half again. Using firm pressure wipe the area 

within the template. Start at one of the inside corners of the template and wipe in concentric 

squares, progressing toward the center. End with a scooping motion. Without allowing the gauze 

to touch any other surface, reverse the last fold so that the exposed side of the gauze is facing 

inward and using a fresh surface of the gauze, wipe the same area in the same manner as before. 

Roll or fold the gauze again and insert into the shipping container.

Note: Wiping in concentric squares is described by OSHA [25]. It is especially suitable for large 

(e.g. 1000 cm2

) areas.

b. Side-to-side Wiping (or Blotting) Technique (particularly suitable for rough, porous, and/or soiled 

surfaces): Fold the pre-wetted gauze in half and then fold in half again. Using firm pressure wipe or 

blot the area within the template with at least five overlapping side-to-side horizontal passes (see 

NOTE) beginning at the top and progressing to the bottom in a “Z” pattern. End with a scooping 

motion. If blotting, blot at least five times on each horizontal pass (see NOTE). Without allowing 

the gauze to touch any other surface, reverse the last fold so that the exposed side of the gauze 

is facing inward. Using a fresh surface of the gauze, wipe or blot the area again with at least five 

overlapping top-to-bottom vertical passes beginning at the left side and progressing to the right 

in an “N” pattern. If blotting, blot at least five times on each vertical pass. Roll or fold the gauze 

again and insert into the shipping container. Blotting is suggested in areas so soiled or rough that 

the threads of the gauze media are continually snagged.

NOTE: On areas larger than 100 cm2

, more than five passes and blots will be needed.

c. Repeat or Serial Wiping: If isopropanol is used for wiping, a serial or repeat wipe sample of the 

same area with a fresh gauze wipe will improve sampling efficiency. (See recoveries for second 

wipe in Tables 10a and 10b.) For serial wiping, repeat the wiping procedure described above 

(APPENDIX steps 7a or 7b) with a fresh gauze wipe. Place the second gauze wipe into the same 

shipping container as the first gauze. The 50-mL polypropylene centrifuge tubes are large enough 

to contain up to two gauze wipes.

NOTE: If the area to be wiped remains substantially wet from the first gauze, the second gauze 

wipe might be used in the dry state to soak up the residual solvent from the first gauze 

wipe.

8. Composite sampling: Composite samples are allowed by some regulatory agencies. Their use for 

quantitative purposes may be subject to the permission and guidance of regulatory agencies. Refer 
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to guidelines of regulatory agency for directions on composite sampling. A basic default guideline 

for composite sampling is as follows: Do not mix inconsistent samples; that is, areas wiped must 

be equal in area, sampled areas must have the same high or low probability of contamination, and 

sampled areas must relate to a specific target appliance or site and not to several appliances or 

incongruous sites combined.

NOTE: Composite samples cannot meet specific action-threshold requirements for discrete sampling 

locations. Nor do composite samples consisting of four wipes, for example, improve the 

sensitivity by decreasing the LOD four fold; instead it raises the LOD by a factor related 

to the extra volume of desorption solution that is required to desorb a larger number of 

wipes. The following example illustrates these two points. Assume that the action level was 

0.1 μg/100 cm2

. If the analysis gave an LOD of 0.06 μg/sample for a single wipe or discrete 

sample covering an area of 100 cm2

, then the LOD for the analysis could be expressed as 0.06 

μg/100 cm2

, which is low enough to be able to determine whether any discrete sample is at 

or exceeds the action level. Now if a composite of four wipes was taken, each with an area 

of 100 cm2

 for a total area wiped of 400 cm2, the LOD for that composite sample is not 0.06 

μg/400 cm2

 nor is it 0.015 μg/100 cm2; it is actually several times larger than 0.06 μg/400 cm2. 

First of all it increases relative to the ratio of the volume of desorption solution used to desorb 

the sample compared to that used for the calibration standards. Secondly it has nothing to 

do with the area that was wiped, because the LOD for the calibration curve is determined 

in terms of μg per sample, independent of the area. To explain the first point, assume 

approximately 90 mL was used (for ease in calculation) to desorb the four wipes and 30 mL 

(the normal amount for a single wipe) was used to desorb each calibration standard. The 

calculation of the LOD for the four composited samples would be μg/sample × (desorption 

volume for 4 wipes) ∕ (desorption volume for the calibration standards), or 0.06 μg/sample 

× (90 mL/30 mL), or 0.18 μg/sample for the composited sample. Since the area wiped for 

the composite sample was 400 cm2

, the LOD for that sample could be expressed as 0.18 

μg/400 cm2

. Regarding the second point, this value, 0.18 μg/400 cm2, cannot be construed 

or mathematically reduced to 0.045 μg/100 cm2

 because it cannot be known whether three 

of the four wipes were blank and the fourth wipe just under the value of 0.18 μg. Hence, the 

effective LOD per individual wipe has to be regarded not only as 0.18 μg/400 cm2

 but also 

as 0.18 μg/100 cm2

 because any value determined for entire 400 cm2 might have come from 

just one of those 100 cm2

 areas. Thus, for composite samples, the LOD must be expressed in 

terms of the entire area wiped and not extrapolated to some portion thereof. In this example, 

an LOD of 0.18 μg/100 cm2

 is above the action threshold of 0.1 μg/100 cm2, meaning that this 

composite sample cannot satisfy the requirement that residual levels be below 0.1 μg/100 

cm2

. It remains for the regulatory agency and not the laboratory to determine how to apply 

results for composite samples to the established action levels. The same consideration that is 

given above for the LOD applies to results that are greater than the LOD. To avoid confusion 

in reporting concentrations for composite samples, it is recommended that the sample 

concentration (in μg/sample, whatever the sample size) and the total area wiped (in cm2

) be 

reported separately. For example, a result of 0.4 μg/sample for a sample consisting of four 

separate wipes of 100 cm2

 each (for a total area wiped of 400 cm2), is to be reported as 0.4 

μg/400 cm2

 and not averaged to 0.1 μg/100 cm2. This manner of reporting may be required 

by some regulatory agencies.

9. For quality assurance purposes, regulatory agencies may require duplicate samples to be taken in 

the field. If such is the case, an area contiguous with and adjacent to the first area, if possible, should 

be wiped as described under SAMPLING. Do not re-wipe the previously wiped area. This sample 

is a blind sample and should not be identifiable by the analytical laboratory as a duplicate of any 

other sample. These are distinct from the laboratory duplicates of a single sample described in step 

14 of the method. Field duplicates are useful for evaluating the consistency of sampling technique, 

assuming uniformity of contamination on adjacent sampling sites. Laboratory duplicates are useful 

for evaluating consistency of sample preparation and instrumental analysis.
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D. DESORPTION FROM MEDIA:

1. An internal standard spiking solution volume of 60 μL was selected for ease in scaling from 60 μL 

per 30 mL to 80 μL per 40 mL of desorption solution. In either case the rate of 2 μL internal standard 

spiking solution per mL desorption solution was used. However, any convenient volume of internal 

standard spiking solution (e.g. 50 μL) that can be delivered reproducibly is acceptable. Whatever 

volume is chosen, there must be no variation in the volume of the internal standard spiking solution 

used in preparing each of the calibration standards. If spiking Strategy A is used (see APPENDIX D3), 

it is critical to know the exact volume of internal standard spiking solution that is applied to each 

sample (V1

), the media blanks (V5), and the calibration standards (V2), since these volumes are used 

for internal standard spiking solution volume corrections in step 19.

2. It is not necessary to know the exact volume of desorption solution added to each sample or the 

volume of residual wetting alcohol because differences in the volumes are normalized through the 

use of internal standards added prior to desorption.

3. Alternate strategy for spiking internal standards (spiking strategy B below): By using the exact same 

volume of internal standard spiking solution in all samples, blanks, QC samples, and calibration 

standards, regardless of the volume of desorption solution added or residual wetting alcohol, the 

volume corrections in step 19, (V1

/V2 and V5/V2) drop out of the equation. However, the internal 

standard GC peak areas must still be measurable in samples where larger volumes of desorption 

solution are used (such as for composite samples). Because of the increased dilution of the internal 

standard in larger samples, this approach should be limited to desorption solution volumes of about 

120 mL or less.

NOTE: There are two separate strategies for handling larger samples requiring larger volumes of 

desorption solvent. These are outlined below as strategies A and B.

Number of 

Wipes

Size of 

Shipping 

Container 

(mL)

Volume of Internal Standard 

Spiking Solution (μL)

Strategy A Strategy B

Volume of 

Desorption Solution 

(mL) 

 (Strategies A and B)

1 40-50 60 60 30

2 50 80 60 40

4 (e.g., 100-120 160 60 80

Composite)

Apply volume Do not apply 

correction factors volume correction 

at step 19. factors at step 19.

With either strategy, if two gauze wipes were included in the samples, then use 40 mL of 

desorption solution. If four gauze wipes were included in the samples, then use 80 mL of 

desorption solution.

a. In strategy A, the volume of internal standard spiking solution is kept at a constant ratio of 2 

μL per mL of desorption solution added. This enables larger samples to be desorbed without 

diminishing the area of the GC peak for the internal standard. However, a volume correction 

factor (V1

/V2) is needed in the final calculations in step 19. Therefore, the exact volume of internal 

standard added to each of the samples relative to that added to the calibration standards must 

be known.

b. In strategy B, the volume of internal standard spiking solution is kept constant for all samples 

and calibration standards, but need not be exactly 60 μL. This enables the final calculations to 

be made in step 19 without a volume correction factor. However, the area of the GC peak for the 
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internal standard will vary with sample desorption volume and the internal standard must be 

concentrated enough to be measurable where larger volumes of desorption solution are used.

E. DRYING COLUMN PREPARATION

Using 1 cm i.d. × 12-15 cm long polypropylene columns having a fritted polyethylene disc or 

equivalent (see EQUIPMENT), add 1 gram (~0.8 cc) of anhydrous potassium carbonate (the bed 

dimension will be about 1.0 cm dia. × 1 cm long). Add 1 gram (~0.8 cc) anhydrous sodium sulfate 

on top of the potassium carbonate. Remove any particles clinging electrostatically to the outside 

surfaces.

NOTE: Particles of the drying salts must not get into the collection tube, either through the frits or 

glass wool plugs, or from particles clinging electrostatically to the outside of the columns. 

Salts appear to inhibit derivatization efficiency.

F. DERIVATIZATION:

If isopropanol was used as the wetting solvent for the wipes, some of it will be co-extracted into the 

methylene chloride. In the presence of trace isopropanol, the crystal violet will go through a series 

of color changes as the extracts are evaporated to dryness. However, if methanol was used as the 

wetting solvent, the color of the crystal violet will remain blue to blue-violet at all stages of drying. Yet 

even with methanol, the same color changes can be afforded by adding 0.1 mL of isopropanol to the 

extracts prior to evaporation. Recoveries of analyte will not be affected in the absence of isopropanol, 

however, as long as the residues are dry before proceeding to step 11b.

 With the presence of a small amount of added or co-extracted isopropanol, as each sample 

concentrates, the color of the solution will go from a blue or violet color rapidly through green to 

a yellow color as the residue approaches dryness, which is indicative of increasing hydrogen ion 

concentration in the residual alcohol. Upon continued blowing with nitrogen, the color of the residue 

turns back to a green or blue hue just at the point of dryness, which is indicative of the loss of excess 

hydrogen chloride and/or alcohol. At this stage the samples are dry and may be removed. Continued 

blowing beyond this point may turn the dried residue to a deep blue-violet or violet color. Losses of 

analyte have not been experienced even after blowing for five additional minutes beyond the violet 

stage as long as the hydrochloric acid had been added. Color changes will not be as dramatic or will 

not develop if too much crystal violet is used. 

 As the samples become concentrated, the tubes may be raised up in the water bath so that only the 

very bottoms of the tubes touch the surface of the water. This makes it easier to observe the color 

changes. The tubes may be raised out of the water bath, but blow-down times are lengthened.

Prolonged heating at high temperatures during derivatization with the acidic conditions of the acid 

anhydride derivatizing agents promotes mutual isomerization between the ephedrine diastereomers 

(ephedrine and pseudoephedrine). Dehydration of the ephedrine compounds (ephedrine, 

norephedrine, and pseudoephedrine) also occurs to some extent to yield β-amino-β-methyl styrenes. 

Heating during derivatization for longer than one hour is especially not recommended. Thirty 

minutes is sufficient.

NOTE: The color of the solution will gradually fade from purple to deep blue within about 20-30 

minutes. This is due to the known tendency of phenolphthalein to fade at high pH. It has also 

been observed that in certain bulk samples, unknown constituents will cause the color of 

phenolphthalein to fade rapidly so that a purple color cannot be obtained at a pH >9, leaving 

only the blue color of the bromothymol blue. A quick check with pH paper can confirm that 

the pH is 9 or greater.

G. MEASUREMENT:

Recoveries for the laboratory control matrix spike samples (QC and QD) must meet the guidelines of 

the specific regulatory agency involved, if applicable (80-120% is a reasonable target in the absence 

of specific guidance). 
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NOTE: The QC samples (QC and QD) in this method may be referred to in some guidance documents 

as matrix spike and matrix spike duplicate samples (MS/MSD), but serve the same purpose. 

Analyze and report field-equipment blanks as samples. Do not subtract their values from any 

other sample.

Recoveries of Continuing Calibration Verification (CCV) standards must meet guidelines of 

regulatory agency (80-120% is a reasonable target in the absence of specific guidance). The CCV 

standards may be referred to in some guidance documents as “QC samples,” but such “QCs” are 

equivalent to liquid standards (not matrix spiked samples) and serve the same purpose of the CCVs 

in this method.

With the GC/MS it is possible to achieve the lower limit of 0.05 μg or less per sample for 

methamphetamine in either the scan mode or SIM mode. The scan mode is essential where the 

identification of unknowns is an analytical objective. If lower limits of detection are desired or 

difficult to obtain in the scan mode, or for routine target compound only analyses, the instrument 

may be operated in the SIM mode.

H. MAKING DILUTIONS: 

 If the samples exceed the upper calibration range for the analysis, one of the following procedures 

may be used to estimate the high level concentrations.

1. Dilution Procedure A (dilution of the derivatized sample by reconstitution solvent): 

This option may be used only if the analytes in the sample were completely derivatized (see NOTE 

below). If derivatization was complete, transfer an aliquot of the sample from the GC vial (e.g. 0.2 

mL for a 1:5 dilution) to a clean GC vial and dilute with reconstitution solvent (e.g. 0.8 mL for a 1:5 

dilution), cap vial, mix, and reanalyze. However, dilution also dilutes the internal standard, and this 

procedure is useful only if the GC peak area for the internal standard is sufficiently measurable 

and the calibration curve is reasonably linear. Dilutions probably should not exceed a factor of 10. 

If this approach is used it is not necessary to enter a dilution factor in step 19 (V3

/ V4) since both 

internal standard and analyte are diluted equally. The accuracy of this dilution procedure depends 

upon the linearity of the calibration curve in the extrapolated region beyond the upper end of the 

calibration curve.

NOTE: Determination of Incomplete Derivatization: Incomplete derivatization can be caused 

by water, glycols, a large excess of analyte, or other contaminants that interfere with or 

compete for the derivatization reagent. If any one of the following symptoms appears, use 

Dilution Procedure B described below.

a. An “oily” film (i.e., apparently viscous liquid) or unusual residue (e.g. grit) remains after being 

blown-down under nitrogen after derivatization (step 11d). This may be due to the presence 

of water, glycols, detergents, salts, or other contaminants. Incomplete derivatization has been 

observed with such residues.

b. A very large (off scale) GC peak for any one of the derivatives (e.g. pseudoephedrine, a precursor 

for methamphetamine) indicates the possibility of incomplete derivatization for other analytes 

(e.g. methamphetamine) due to competition for the derivatization reagent.

c. A smaller than usual GC peak area for the internal standard (<50% of the average) in undiluted 

samples suggests that something was competing for or inhibiting the derivatizing reagent. Such 

inhibition or competition for the internal standard will be experienced by the target analyte as 

well.

d. Incomplete derivatization can be confirmed by the obvious presence of a GC peak for an 

underivatized target analyte. Underivatized analytes are not always detectable. Ephedrines 

usually do not show up on DB-5 capillary columns in this method, but GC peaks for underivatized 

secondary amines (e.g. methamphetamine) and for high levels of underivatized primary amines 
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(e.g. amphetamine) can be detected, usually as irregularly shaped GC peaks, depending upon GC 

column conditions.

e. The problem of incomplete derivatization can be minimized by the use of an isotopic (e.g. 

deuterated) analog for each of the target analytes as the internal standard for that compound. 

This allows quantification in spite of incomplete derivatization.

2. Dilution Procedure B (dilution of a smaller aliquot of the original desorbate): If the sample was not 

completely derivatized or if large dilutions are needed (e.g. greater than about 1:5), the following 

procedure can be used. See NOTE in part 1 above. The procedure may also be used if derivatization 

was determined to be complete.

a. Dilute an aliquot of the original aqueous acid desorbate of the wipe sample to 10 mL with 

desorption solution from a simulated sample blank, and re-extract. Add both the aliquot to 

be diluted and the diluting solution from the simulated blank directly to a clean 25-mL glass 

centrifuge tube (step 7f) and proceed to step 8. For example, to make a 1:10 dilution, transfer 1 

mL of original desorbate to the 25-mL tube and dilute with 9 mL from a simulated sample blank.

b. The simulated sample blank should be prepared identically to the sample being diluted, using 

the same volumes of internal standard spiking solution and desorption solution that were used 

with the sample in the original desorption. For example, if the original sample was desorbed with 

40 mL desorption solution with 80 μL of added internal standard spiking solution, then prepare 

the simulated blank in the same way. The volume of wetting alcohol is estimated (e.g. about 3 

mL per 3”×3” (7.5 cm x 7.5 cm) 12-ply cotton gauze wipe). Include a dilution factor (V3

/ V4) in the 

calculations in step 19 (e.g. V3

/ V4 = 10 mL divided by the volume in mL of original desorbate 

diluted to 10 mL with solution from the simulated blank). The dilution factor in the above 

example is 10 mL/ 1 mL or 10.

c. Correct for differences in internal standard spiking solution volumes in step 19 (if applicable) 

using for V1

 the volume of internal standard spiking solution which was added to the original 

undiluted sample.

NOTE: This dilution procedure gives quantitative results only if the residual volume of 

methanol (or isopropanol) used for wetting the sample wipes was exactly the same as 

the volume used in preparing the calibration standards (normally about 3 mL, see Table 

6). Deviations of a few milliliters in residual wetting alcohol will not affect the results for 

undiluted samples, but will amount to an error of a few percent in the final results of 

samples that are diluted.

d. The potential error due to differences in residual wetting solvent can be estimated for specific 

volumes of desorption solution and wetting alcohol. Assume the sample wipes and calibration 

standards are both desorbed in 30 mL of desorption solution and 3 mL of alcohol is added 

to the calibration standards. The potential error in volume (and final results) in the samples is 

approximately ±3 % (inversely proportional) per mL difference in the residual alcohol in the 

samples (i.e., ±1 mL difference in 33 mL). For 40 mL of desorption solution and 4 mL of alcohol 

added to the calibration standards, the error is ±2 % for every mL difference (i.e., ±1 mL difference 

in 44 mL). However, since the volume of residual wetting alcohol is not known and cannot be 

determined once the sample wipe has been desorbed, the actual error cannot be determined. 

However, the maximum possible error can be calculated. Since the maximum amount of alcohol 

that a recommended wipe can hold is about 6 mL when saturated (dripping wet), there can only 

be a deviation of plus or minus 3 mL from the 3 mL alcohol added to the calibration standards. 

Therefore, the maximum error in a result due to differences in the volume of residual alcohol in 

a cotton gauze sample compared to the standards can only be three times the error for a 1 mL 

difference in volume. Since the error for ±1 mL is ±3.03%, the maximum error for ±3 mL is three 

times larger, or ±9.1%. In practice, the error will be less than this because it is unlikely that the 

gauze samples will be completely dry or completely saturated after squeezing out the excess 

alcohol and wiping a surface. The practical amount of alcohol that remains in the wipes when the 

excess is squeezed out is between 1 and 2 mL. This translates into an error that is between +3% 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE . . .on Wipes by Liquid-Liquid Extraction: METHOD 9106, Issue 1, dated 17 October 2011 - Page 30 of 30

and +6% in the final results for diluted samples. Undiluted samples will not be affected. This error 

is within the overall accuracy for the method for methamphetamine.

3. Dilution Procedure C (dilution of desorbates from dried samples):

Dilution errors for over-range samples may be corrected by knowing the exact amount of residual 

alcohol in the samples. The volume (or weight) of residual solvent in each gauze wipe might be 

determined by the difference between a wet weight and dry weight. Better yet, the error might be 

eliminated for diluted samples by adding, after the samples are dried (without taking any weight), 

the same known volume of wetting alcohol that is added to the calibration standards (i.e., 3 mL). 

Thereafter, if any samples need dilution, there will be no dilution errors due to differences in residual 

alcohol, because all samples and standards will have the same volume of alcohol and total volume 

of desorption solution.

However, air drying of the samples is not recommended because of the possible loss of 

methamphetamine due to its volatility when it is not in the salt form, which form cannot be 

assured in field samples. Also, manipulating the samples for weighing and drying might introduce 

contamination. Drying is not recommended as a procedure for analytes having a vapor pressure 

high enough to be lost in the process, or that tend to form azeotropes with alcohols; this is especially 

important when the critical action levels for remedial cleanup are at the lower end of the method 

calibration range. Drying is not an option if the samples have already been desorbed.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHAMPHETAMINE and Illicit Drugs, Precursors, 9109 and Adulterants on Wipes by Solid Phase Extraction

FORMULA: Table 1 MW: Table 1 CAS: Table 1 RTECS: Table 1

METHOD: 9109, Issue 1 EVALUATION: Partial Issue 1: 17 October 2011

OSHA: none for surfaces

NIOSH: none for surfaces

Other OELs and guidelines: [1, 2, 3] 

PROPERTIES: Table 2

SYNONYMS: Table 3

SAMPLING

SAMPLER: Wipe

SAMPLE AREA: 100 cm2 or 1000 cm2

SHIPMENT: Ship refrigerated preferably, <6 °C 

SAMPLE 

STABILITY: At least 30 days at <6 °C (See Table 4)

FIELD 

BLANKS: 2 to 10 blanks per sample set 

MEASUREMENT ACCURACY

LEVEL 

STUDIED: 3.0 μg/sample

BIAS: Table 9 [4]

OVERALL

PRECISION ( ): Surface recovery not performed

ACCURACY: Table 7a and 7b [4]

MEASUREMENT

TECHNIQUE: GAS CHROMATOGRAPHY/MASS 

SPECTROMETRY

ANALYTES: Table 1

DESORPTION: 0.1 M sulfuric acid

CLEANUP/

EXTRACTION: Solid phase extraction

DERIVATIZATION: MSTFA and MBHFBA

INJECTION 

VOLUME: 2 μL Splitless

TEMPERATURE 

 Injection: 255 °C.

 Detector: 285 °C

 Column: 90 °C (2 min), to 310 °C (10 °C/ min), hold 6 

min

MASS 

SPECTROMETER: Scan mode (29 – 470 AMU), 2 scan/sec 

or selected ion monitoring (SIM) mode 

(Table 5)

CARRIER GAS: Helium, 1.5 mL/min

COLUMN: Capillary, fused silica, 30 m x 0.32 mm i.d., 

0.5 μm U.S. Pharmacopeia (USP) G27 film

CALIBRATION: Standards from spiked wipes with internal 

standard, See Table 6

RANGE: Table 7a and 7b [4].

ESTIMATED LOD: Table 4 

PRECISION ( ): Table 7a and 7b [4]

APPLICABILITY: For methamphetamine the range is 0.05 to 60 μg/sample (sample = 100 cm2 or 1000 cm2). This method was 

developed for the analysis of selected drugs and precursors on surfaces in clandestine drug labs [5]. Sampling methodology 

was tested using wipes on smooth, non-porous surfaces. The APPENDIX contains sampling information for other types of 

surfaces. 

INTERFERENCES: No chromatographic interferences detected. Water, surfactants and polyols inhibit derivatization.

OTHER WIPE METHODS: NIOSH 9106 uses liquid-liquid extraction and gas chromatography/mass spectrometry (GC/MS) 

to measure multiple drugs [6]. NIOSH 9111 uses liquid chromatography/mass spectrometry (LC/MS) to measure 

methamphet-amine [7].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 2 of 32

REAGENTS:

NOTE: See APPENDIX A for special instructions on reagents.

1. Analytes listed in Table 1*

2. Internal standards from those listed in Table 8a

3. Solvents, residue free analytical grades

a. Isopropanol (IPA)*

b. Methanol*

c. Methylene chloride (CH2

Cl2)*

d. Acetonitrile*

4. Concentrated sulfuric and hydrochloric acids (Analytical Reagent [AR] 

or trace metals analysis grades)*

5. Ammonium hydroxide (NH4

OH), 28-30%, A.C.S. grade*

6. Bromothymol blue, ≥95%, A.C.S.; crystal violet (Gentian Violet), 

≥95%, A.C.S.

7. Purified gases: helium for carrier gas, nitrogen for drying

8. MSTFA (N-methyl-N-trimethylsilyl-trifluoro-acetamide) derivatizing 

agent*

9. MBHFBA (N-methyl-N,N-bisheptafluorobutyramide) derivatizing 

agent*

10. 4,4’-Dibromooctafluorobiphenyl, 99%

11. Deionized water (ASTM type II)

SOLUTIONS:

NOTE: See APPENDIX A for special instructions on solutions.

1. Prepare solutions of analytes of interest (Table 1). Calculate 

concentrations as the free base. Keep solutions refrigerated (<6 °C). 

Protect solutions from light.

a. Stock solutions are prepared at about 1-2 mg/mL in methanol.

b. Analyte spiking solutions are prepared by diluting the stock 

solutions to about 200 μg/mL each in methanol.

2. Prepare internal standard spiking solution in methanol at about 200 

μg/mL. 

NOTE: Add about 2 mg of crystal violet per 20 mL of internal 

standard spiking solution to help indicate which 

samples have been spiked.

3. Desorption solution: 0.1 M sulfuric acid. Add 22 mL conc. sulfuric acid 

to 4 liters deionized water.

4. Bromothymol blue pH indicator solution: 1 mg/mL in 4:1 

isopropanol:deionized water.

5. Crystal violet indicator: 2-3 mg/mL in isopropanol.

6. Solid phase extraction (SPE) wash solution: Aqueous 0.1 M 

hydrochloric acid: Dilute 8.3 mL concentrated hydrochloric acid in 

about 800 mL water, dilute to 1 liter with ASTM Type II water.

7. SPE elution solution: 80:20:2 CH2

Cl2:IPA:NH4OH v/v. Prepare fresh 

daily.

8. 0.3 M hydrochloric acid in methanol: Dilute 2.5 mL conc. hydrochloric 

acid in about 80 mL methanol; dilute to 100 mL with methanol.

9. Derivatization diluent solvent: acetonitrile containing 4 μg/mL of 

4,4’-dibromo-octafluorobiphenyl (optional).

* See SPECIAL PRECAUTIONS

EQUIPMENT:

NOTE: See APPENDIX B for special instructions on equipment.

1. Wipe, cotton gauze, (7.6 cm × 7.6 cm) 12-ply or equivalent.

2. Sample storage and shipping container: 50-mL polypropylene 

centrifuge tubes with polytetrafluoroethylene (PTFE)-lined caps or 

equivalent.

3. Gas chromatograph/mass spectrometer detector, with column and 

integrator, See p. 9109-1.

4. Solid phase extraction (SPE) mixed phase cation exchange 

hydrophilic extraction columns (See Appendix E)

5. Collection tubes and GC vials:

a. Glass test tubes (13 mm x 100 mm) with PTFE-lined caps

b. GC autosampler vials, 2-mL Limited-volume, 300-500 μL (amber 

vials recommended), and caps.

6. Volumetric flasks: 10-, 100-, and 250-mL.

7. Reagent bottle, 4-L .

8. Liquid Transfer:

a. Syringes: 10-, 25-, and 100-μL.

b. Mechanical pipette with disposable tips, 5-mL.

c. Repeating dispensers: 1 to 5-mL.

d. Syringe or repeating dispenser: 100-μL.

e. Syringes: 250-μL.

9. Forceps.

10. Gloves: latex or nitrile. Avoid vinyl gloves (see 9109-3, Sampling, 

step 1, NOTE 2).

11. Rotating mixer capable of 10-30 rpm.

12. Vacuum manifold box with 12 to 36 vacuum ports, and adjustable 

flow rates.

13. Nitrogen blow-down apparatus with water bath capable of 

maintaining 35 oC.

14. Vortex mixer.

15. Pasteur pipettes.

16. pH paper.

17. Template: 10 cm x 10 cm (or 1 foot x 1 foot) opening made of 

relatively rigid disposable cardstock or sheet of PTFE.

18. Ice or other cold media for shipping.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 3 of 32

SPECIAL PRECAUTIONS: The solvents are flammable and have associated adverse health effects. 

Phenethylamines target the nervous system at very low concentrations and are easily absorbed 

through the skin. Avoid breathing vapors. Avoid skin contact. Work should be performed in a hood with 

adequate ventilation. Analysts must wear proper eye and hand protection (e.g., latex gloves) to prevent 

absorption of even small amounts of amines through the skin as well as for protection from the solvents 

and other reagents. Dissolving concentrated hydrochloric or sulfuric acid in water is highly exothermic. 

Goggles must be worn. The derivatization reagents react violently with water.

Caution must also be exercised in the handling and analysis of samples. Clandestine drug labs may 

produce unknown and seriously toxic by-products. For example, in the manufacture of designer 

drugs (e.g., MPPP, a homolog of Alphaprodine), at least one very neurotoxic by-product, 1-methyl-4-

phenyl-1,2,5,6-tetrahydropyridine (MPTP), has been identified that specifically and irreversibly causes 

Parkinson’s disease [8,9].

SAMPLING:

See APPENDIX C for special instructions on sampling.

1. Using a new pair of gloves, remove a gauze wipe from its protective package. Moisten the wipe with

approximately 3 to 4 mL of methanol (or isopropanol).

NOTE 1: Apply no more solvent than that needed to moisten approximately the central 80% of the 

area of the gauze wipe. Excess solvent may cause sample loss due to dripping from the 

wipe.

NOTE 2: Do not use vinyl gloves due to the potential for leaching of phthalate plasticizers and 

contamination of the samples.

2. Place the template over the area to be sampled (may tape in place along outside edge of template).

Wipe the surface to be sampled with firm pressure, using vertical S-strokes. Fold the exposed side of 

the pad in and wipe the area with horizontal S-strokes. Fold the pad once more and wipe the area 

again with vertical S-strokes.

3. Fold the pad, exposed side in, and place in shipping container and seal with cap.

NOTE: Keep samples refrigerated (<6 oC). While methamphetamine and several related amines 

are stable on the recommended wipe media for at least 7 days at room temperature, 

refrigeration is recommended as soon as possible (see Table 4).

4. Either clean the template before use for the next sample or use a new disposable template.

5. Label each sample clearly with a unique sample identifier.

6. Prepare a minimum of two field blanks with one field blank for every ten samples.

NOTE: In addition, include at least 3 media blanks for the analytical laboratory to use for their

purposes. The wipes used for the media blanks should be from the same lot as the field 

samples.

SAMPLE PREPARATION:

 See APPENDIX D for special instructions on sample preparation.

7. Desorption from media:

a. Remove cap from shipping container.

NOTE: Sample wipe should fit loosely in the container. If not, transfer sample to a larger

container.

b. Spike 60 μL of internal standard spiking solution onto each wipe sample.

c. Add 30 mL desorption solution (0.1 M sulfuric acid).

NOTE: If the samples were transferred to a larger container, rinse the original shipping container

with the desorption solution, shake, and decant the rinsate into the larger container.

Cap securely and mix contents by inverting the tubes end over end on a rotary mixer or 

equivalent at 10-30 rpm for at least one hour.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 4 of 32

d. Check the pH which should be about ≤ 4. If needed, adjust the pH with diluted (2.5 to 3 M) sulfuric

acid drop-wise, mixing the contents by shaking or inversion a few times after each addition of acid

before checking the pH.

e. After mixing, transfer 10 mL of supernatant to a 25-mL glass centrifuge tube.

NOTE: If extraction is to be performed on a subsequent day, store samples in a refrigerator. Analytes

are stable in the desorption solution for at least one week refrigerated.

8. Solid phase extraction procedure:

a. Column selection: Select one of the SPE columns listed in Appendix E. Each brand of column has a

slightly different conditioning procedure and resistance to flow. Other brands of SPE columns may

also work. Elution profiles of drugs to be analyzed need to be determined before use of columns

other than those specified.

b. Setting up columns: Attach SPE columns to vacuum ports on the manifold. Attach vacuum line to

vacuum pump capable of 25-30 psi vacuum.

c. Conditioning: Condition each column with 1 column volume (3 mL) of methanol followed by 1

column volume of Type II deionized water. For some brands the conditioning volume is 1/3 column

volume. Check product literature.

d. Loading: Load each SPE column with 5 mL of the sample acid desorbate solution. Adjust vacuum so

that the flow rate is about 1-2 mL/minute. The vacuum required to obtain that flow rate varies with

brand of SPE column.

e. First wash: Wash each column with 1 column volume (3 mL) of 0.1 M aqueous hydrochloric acid. For

some brands this volume may be decreased to 1 or 2 mL. Check product literature.

f. Second wash: Wash each column with 1 column volume of methanol. Add the methanol in 2 or 3

separate aliquots to ensure that the aqueous acid is flushed through. Discard all effluents.

g. Drying: Remove last traces of water in the SPE columns by pulling air through the columns under

increased vacuum (e.g., 25 psi) for 5 minutes. Silica-based SPE columns or columns with high

resistance to flow may require a longer time to reach dryness.

h. Elution: Position 13 x 100 mm collection tubes under each column. Elute analytes with 3 mL of

elution solution (80:20:2 methylene chloride:isopropanol:concentrated ammonium hydroxide v/v,

freshly prepared). Adjust vacuum so that the flow rate is 1 mL/minute or less. For some brands this

flow rate may occur without applied vacuum. Most of the analytes (e.g., amphetamine, ephedrine,

methamphetamine, etc.) are eluted in the first milliliter.

9. Evaporation: To each collection tube containing eluate, add about 5 μL crystal violet solution and

100 μL of 0.3 M hydrochloric acid in methanol. The samples are evaporated to dryness under gently

blowing nitrogen at 25-35 °C. The samples should be removed from the evaporation bath within a few

minutes after dryness. A mixed whitish and purple residue will remain. The purple color of the crystal

violet helps to make the residue more visible when dried. The color of the crystal violet remains a

constant blue to blue-violet during concentration and drying.

10. Derivatization: (Perform under the hood.) Add 100 μL of acetonitrile containing the optional

dibromooctafluorobiphenyl secondary internal standard. Add 25 μL MSTFA and 25 μL MBHFBA in that

order. Cap tubes between additions to prevent atmospheric humidity from affecting the reagents. (See

note below. Have no more than 5 or 6 tubes uncapped at a time.) Vortex each tube about 4-5 seconds.

Using Pasteur pipettes, transfer each mixture to low-volume (300-500 μL) amber autosampler vials and

cap vials.

NOTE 1: Some derivatization takes place at room temperature, especially trimethylsilylation.

Derivatization is completed on-column after injection. No prior heating is required or 

recommended. 

NOTE 2: The color of the reconstituted solution should be deep blue to violet. If the color turns light 

blue or turquoise upon standing, moisture may be present (the vials may not have been 

capped tightly enough). Such samples need to be reprocessed beginning at step 8 since the 

derivatives are not stable in the presence of moisture. If the vials are securely capped, the 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 5 of 32

solutions will be stable for several days at room temperature and at least a week refrigerated. 

Protect vials from light (amber vials recommended.)

11. Analyze samples, standards, blanks, and QCs by GC-MS. (See MEASUREMENT, steps 15-17 and

p. 9109-1.)

CALIBRATION AND QUALITY CONTROL:

12. Determine retention times for the derivatives of the analytes of interest using the column and

chromatographic conditions specified on page 9109-1. Table 10 gives typical retention times for

various drugs, precursors, and adulterants. Figure 1 shows a typical chromatogram.

13. Calibrate daily with at least six calibration standards and a blank selected from Table 6 to cover the

analytical range.

a. Prepare the analyte spiking solution as follows: Add known amounts of individual drug stock

solutions to a volumetric flask and dilute to volume with methanol. A recommended final

concentration for this solution is approximately 200 μg each per mL.

b. Prepare calibration standards and media blanks in clean shipping containers (e.g., 50-mL

polypropylene centrifuge tubes or equivalent).

NOTE: Liquid standards (standards without added blank wipe media) may be prepared in lieu of

media standards if cotton gauze was used for the samples.

c. Add 3 mL methanol (or isopropanol if isopropanol was used with the samples in the field) to each

calibration standard and media blank. 

NOTE: If two gauze wipes were routinely used for every sample, increase methanol (or isopropanol) 

to 4 mL. See Table 6, footnote 2.

d. Spike a known volume of analyte spiking solution into each calibration standard by spiking directly

onto the media or into solution. Use the spiking volumes suggested in Table 6 to cover the desired 

range.

e. Process each of these through the desorption, solid phase extraction (SPE), drying, and

derivatization steps (steps 7 through 11) along with the field samples.

f. Analyze these along with the field samples. (See MEASUREMENT, steps 15-17.)

14. Prepare matrix-spiked and matrix-spiked duplicate (QC and QD) quality control samples [10].

a. Cotton gauze from the same lot used for taking samples in the field should be provided to the

analytical laboratory to prepare these matrix-spiked quality control samples.

b. The quality control samples (QC and QD) must be prepared independently at concentrations within

the analytical range. (See Table 6 for applicable concentration ranges.)

c. One quality control media blank (QB) must be included with each QC and QD pair.

d. Spike QC and QD with a known amount of target analyte as suggested in Table 6.

i. Transfer clean wipes to new shipping containers.

ii. Add 3 mL of methanol (or isopropanol if isopropanol was used in wiping) to each wipe.

iii. Spike QC and QD with a known amount of analyte as suggested in Table 6.

NOTE: If two gauze wipes were used for the majority of samples in an analytical set, use two clean 

gauze wipes for each QB, QC, and QD, and increase isopropanol (or methanol) to 4 mL. See 

Table 6, footnote 2.

e. Process quality control samples through the desorption, SPE, drying, and derivatization steps (steps

7 through 11) along with the calibration standards, blanks, and field samples.

f. Analyze these along with the calibration standards, blanks, and field samples. (See MEASUREMENT,

steps 15-17.)

MEASUREMENT:

See APPENDIX G for special instructions on measurement.

15. Analyze the calibration standards, quality control samples, blanks, a continuing calibration verification

(CCV) standard consisting of one of the initial calibration standards, and samples by GC/MS. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 6 of 32

Set gas chromatograph according to manufacturer’s recommendations and to conditions listed on page 

9109-1.

a. Set mass spectrometer conditions to manufacturer’s specifications and those given on page 9109-1

for the scan mode or those in Table 5 for the SIM mode.

b. Inject sample volume with autosampler or manually.

NOTE: After the derivatives are prepared and just before analyzing any samples or standards, inject 

the highest concentrated standard several times in order to prime or deactivate the GC 

column and injection port. This will help minimize any drift in the instrument’s response to 

target analytes relative to their internal standards.

c. After analysis, the vials should be recapped promptly and refrigerated if further analysis is anticipated.

16. Using extracted ion current profiles for the primary (quantification) ions specific to each analyte,

measure GC peak areas of analyte(s) and internal standard(s) and compute relative peak areas by

dividing the peak area of the analyte by the area of the appropriate internal standard. Recommended

primary (quantification) ions and internal standards are given in Tables 5, 7 and 8. Prepare calibration

graph (relative peak area vs. μg analyte per sample).

17. Samples from initial investigations of clandestine laboratories are likely to include highly contaminated

samples. If sample results exceed the upper range of the calibration curve, the sample in the GC vial

may be diluted and reanalyzed or a smaller aliquot of the initial acid desorbate diluted, re-extracted,

derivatized, and analyzed. Refer to APPENDIX H for instructions and limitations on making dilutions.

CALCULATIONS:

18. Determine the mass in μg/sample of respective analyte found in the wipe samples, and in the media

blank from the calibration graph.

19. Calculate final concentration, C, of analyte in μg/sample:

c = concentration in sample (in μg/sample determined from the calibration curve).

  volume correction factor (needed only when the volume of internal standard spikin

solution used for spiking the samples - such as for composite samples requiring larger 

desorption solution volumes - is different from that used for spiking the calibration 

standards). (See Table 6, footnote 4).

V1

 = volume in μL of internal standard spiking solution used to spike samples.

V2

 = volume in μL of internal standard spiking solution used to spike the standards.

  dilution factor, if applicable

V3

 = 5 mL (volume of desorbate normally taken for extraction in step 8d).

V4 = volume in mL of desorbate actually taken for extraction and diluted to 5 mL with blank 

desorbing solution containing internal standard.

b = concentration in media blank (in μg/sample determined from the calibration curve).

  volume correction factor for the media blank (needed only if the volume of internal

standard spiking solution used for spiking the media blank is different from that used for 

spiking the calibration standards).
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METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 7 of 32

V5 = volume in μL of internal standard spiking solution used to spike media blank.

20. Report concentration, C’, in μg per total area wiped (in cm2

) as follows:

C = μg/sample (step 19).

A = Total area wiped in cm2 per sample.

NOTE: In general, if the area wiped was greater than or less than 100 cm2, do not convert value to 

μg/100 cm2 unless specifically required or allowed by agency having legal jurisdiction. For 

example, if the sample was a composite sample and the area was 400 cm2

, report results 

as μg/400 cm2

 and not averaged to μg/100 cm2 since regulatory agencies might not allow 

averaging of composite results to 100 cm2

. To avoid confusion, report separately both μg/

sample (C) and the total area wiped in cm2

 per sample (A) for both discrete and composite 

samples.

EVALUATION OF METHOD:

This method was developed according to the NIOSH sampling and analytical method development 

guidelines [11]. This method was evaluated for those analytes listed in Tables 7a and 7b over a range of 

approximately 0.1 μg/sample to 30 μg/sample for several types of sampling media. These concentration 

levels represent approximately the 1 through 300 times the limit of quantitation (LOQ) level for most of 

the analytes. Results are reported in the Backup Data Report [4].

The limits of detection (LOD and LOQ) were determined by preparing a series of liquid standards in 

desorption solution, processing by the SPE of NIOSH 9109, and analyzing in the scan mode. The LODs 

were estimated using the procedure of Burkart [12]. An LOD of 0.1 μg/sample for methamphetamine 

on wipes was achieved in the scan mode. The LOD was set at 0.1 μg/sample because that was the level 

of the lowest calibration standard in the LOD/LOQ study. Lower LODs (e.g., 0.02 μg/sample) have been 

achieved in practice by including calibration standards at lower concentration levels. The cleanliness and 

performance of the mass spectrometer must be maintained such that at 0.1 μg/sample a signal of at least 

5 to 10 times the baseline noise is achievable. This is more easily accomplished in the SIM mode.

Six different wipe media were evaluated. These were 3”x3” (7.5 cm x 7.5 cm) 12-ply cotton gauze, 4”x4” (10 

cm x 10 cm) AlphaWipe® (TX 1004), 4”x4” (10 cm x 10 cm) 4-ply NU GAUZE®, 4”x4” (10 cm x 10 cm) 4-ply 

MIRASORB®, 4”x4” (10 cm x 10 cm) 6-ply SOF-WICK®, and 4”x4” (10 cm x 10 cm) 4-ply TOPPER® sponges. 

Results are given in the Backup Data Report [4]. No synthetic media performed better than cotton gauze. 

Some media (NU GAUZE® and SOF-WICK®) gave inconsistent results.

Precision and accuracy were determined by analyzing 6 replicates at each of 6 concentration levels 

(nominally 0.1, 0.3, 1, 3, 10, and 30 μg/sample). Results are presented in Table 7a for cotton gauze and 

7b for AlphaWipe®. The best precision and accuracies were dependent upon the use of carefully chosen 

internal standards, especially with steric hindrance of the amine (e.g., having N-ethyl and N-propyl 

groups). Long term sample storage stability was determined for periods up to 30 days under refrigeration 

(<6 °C) and for up to 7 days at room temperature (22-24 °C). Results are given in Table 4.

Chlorodifluoroacetic anhydride (CDFAA) and pentafluoro propionic anhydride (PFPA) were evaluated 

as derivatizing agents for the SPE eluates. These were not effective, probably due to the high level of 

ammonium chloride residues in the SPE column eluates. They were most effective with the liquidliquid extraction procedure of NIOSH 9106 [6].

For SPE, the mixed silanization-acylation reagent, MSTFA and MBHFBA [13-15], proved very effective. 

The derivatization mixture is transferred directly to amber mini-GC vials and direct-injected without 

prior heating.
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METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 8 of 32

Recovery of amphetamines from six different types of surfaces using cotton gauze was evaluated (see 

Table 9a and 9b). The practice of serial wiping (wiping the same surface area a second time with a second 

gauze wipe and combining both wipes as a single sample) was evaluated. Four solvents for wetting the 

gauze were tested (distilled water, 5% distilled white vinegar, isopropanol, and methanol). Six replicates 

samples were taken on a latex painted wall. Recoveries and precisions are given in Table 9a and 9b. The 

recoveries with 5% distilled white vinegar were better than for distilled water, but not as good as for 

isopropanol. Methanol is superior to isopropanol. Recoveries with isopropanol are greatly improved with 

a repeat (serial) wipe (11% improvement compared to only about 6% improvement with methanol). 

The study and results are reported in the Backup Data Report for NIOSH 9109 [4]. Additional research on 

surface sample recovery and solvent effectiveness has been reported by Van Dyke et al [13] and Serrano 

et al. [16].

REFERENCES:

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American

Conference of Governmental Industrial Hygienists. [www.acgih.org]. Data accessed April 2014.

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database on

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ifa/GESTIS ]. Data accessed April 2014.

[3] EPA [2013]. Voluntary guidelines for methamphetamine laboratory cleanup. [https://www.epa.gov/

sites/production/files/documents/meth_lab_guidelines.pdf ]. Date accessed March 2016.

[4] Reynolds JM, Siso MC, Perkins JB [2004]. Backup Data Report for NIOSH 9109, Methamphetamine and

illicit drugs, precursors, and adulterants on wipes by solid phase extraction, Abridged. Unpublished.

Prepared under NIOSH Contract 200-2001-0800.

[5] Martyny JW, Arbuckle SL, McCammon CS Jr., Esswein EJ, Erb N [2003]. Chemical exposures associated

with clandestine methamphetamine laboratories [https://www.nationaljewish.org/NJH/media/pdf/

pdf-EnvOccChemical_Exposures.pdf]. Date accessed April 2014.

[6] NIOSH [2011]. Methamphetamine and illicit drugs, precursors, and adulterants on wipes by liquidliquid extraction: Method 9106. In: Ashley KA, O’Connor PF, eds. NIOSH manual of analytical methods.

5th ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and 

Prevention, National Institute for Occupational Safety and Health, [www.cdc.gov/niosh/nmam/].

[7] NIOSH [2011]. Methamphetamine on wipes by liquid chromatography/mass spectrometry: Method

9111. In: Ashley KA, O’Connor PF, eds. NIOSH manual of analytical methods. 5th ed. Cincinnati, OH:

U.S. Department of Health and Human Services, Centers for Disease Control and Prevention, National

Institute for Occupational Safety and Health, [www.cdc.gov/niosh/nmam/].

[8] Baum RM [1985]. New variety of street drugs poses growing problem. Chemical and Engineering

News, 63(63):7-16.

[9] Buchanan JF, Brown CR [1988]. Designer drugs, a problem in clinical toxicology. Medical Toxicology

3:1-17.

[10] NIOSH [1994]. Chapter C: Quality Assurance. In: Eller PM, Cassinelli ME, eds. NIOSH manual of analytical 

methods, 4th ed. Cincinnati, OH: Department of Health and Human Services, Centers for Disease 

Control and Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH)

Publication No. 94-113. [www.cdc.gov/niosh/nmam/].

[11] NIOSH [1995]. NIOSH Technical Report: Guidelines for air sampling and analytical method

development and evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA.

Cincinnati, OH: U.S. Department of Health and Human Services, Public Health Service,

Centers for Disease Control and Prevention, National Institute for Occupational Safety and

Health, DHHS (NIOSH) Publication No. 95-117.

[12] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene

chemistry laboratory. Applied Industrial Hygiene 1(3):153-155. 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

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[13] Van Dyke MV, Serrano KA, Kofford S, Contreras J, Martyny JW [2011]. Variability and specificity

associated with environmental methamphetamine sampling and analysis. J Occup Environ Hyg

8(11):636-4. Date accessed May, 2011.

[14] Little JL [1999]. Artifacts in trimethylsilyl derivatization reactions and ways to avoid them. Journal of

Chromatography. A, 844:1-22.

[15] Crockett DK, Frank EL, and Roberts WL [2002]. Rapid analysis of metanephrine and normetanephrine

in urine by gas chromatography-mass spectrometry. Clinical Chemistry 48: 332-337.

[16] Serrano KA, Martyny JW, Kofford S, Contreras JR, Van Dyke MV [2012]. Decontamination of clothing

and building materials associated with the clandestine production of methamphetamine. J Occup

Environ Hyg 9(3):185-197.

[17] Budavari S, ed. [1989]. Merck Index, 11th ed. Rahway, New Jersey: Merck and Co.

[18] Sweet DV, ed. [1997]. Registry of toxic effects of chemical substances. DHHS (NIOSH) Publication No.

97-119. [https://www.cdc.gov/niosh/].

[19] Sigma-Aldrich MSDS sheets [http://www.sigma-aldrich.com]. Date accessed: May 10, 2004.

[20] Cerilliant Analytical Reference Materials. 811 Paloma Drive, Suite A, Round Rock, Texas 78664 [www.

cerilliant.com].

[21] Howard PH, Meylan WM [1997]. Handbook of physical properties of organic chemicals. Boca Raton,

Florida: CRC/Lewis Publishers. See also Syracuse Research Corporation’s website [http://www.srcinc.

com for demos and updates. Date accessed May 10, 2004.

[22] Zhang JS, Tian Z, Lou ZC [1989]. Quality evaluation of twelve species of chinese ephedra (Ma Huang).

Acta Pharmaceutic Sinica 24(11):865-871.

[23] Cui JF, Zhou TH, Zhang JS, and Lou ZC [1991]. Analysis of alkaloids in chinese ephedra species by gas

chromatographic methods. Phytochemical Analysis 2:116-119.

[24] Andrews KM [1995]. Ephedra’s role as a precursor in the clandestine manufacture of

methamphetamine. Journal of Forensic Sciences 40(4):551-560.

[25] OSHA [2001]. Evaluation guidelines for surface sampling methods. Industrial Hygiene Chemistry

Division, OSHA Salt Lake Technical Center, Salt Lake City, Utah. [http://www.osha.gov/dts/sltc/

methods/surfacesampling/surfacesampling.pdf ]. Date accessed May 2011.

METHOD DEVELOPMENT BY:

John M. Reynolds, Maria Carolina Siso, and James B. Perkins, DataChem Laboratories, Inc., Salt Lake City, 

Utah under NIOSH Contract CDC 200-2001-0800.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for

the content of these websites. All web addresses referenced in this document were accessible as of

the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 10 of 32

Table 1. Formula and registry numbers of analytes

Compound

(alphabetically)

MW(1) (Daltons)

Free 

base HCl salt

Hemisulfate 

salt

Structural Formula

As free base CAS #(2) RTECS(6)

(DL)-Amphetamine 135.21 171.67 184.25 C6H5·CH2·CH(CH3)·NH2 300-62-9(3)

60-13-9(5)

SH9450000

SI1750000

(D)-Amphetamine(7) 135.21 171.67 184.25 C6

H5·CH2·CH(CH3)·NH2 51-64-9(3)

51-63-8(5)

SI1400000

(L)-Amphetamine 135.21 171.67 184.25 C6

H5·CH2·CH(CH3)·NH2 156-34-3(3) SH9050000

Caffeine 194.19 (CH3)

3

·[C5HN4O2] 58-08-2(3) EV6475000

(DL)-Ephedrine 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)·NH·CH3 90-81-3(3)

134-71-4(4)

(L)-Ephedrine(8) 165.24 201.70 214.28 C6H5·CH(OH)·CH(CH3)NH·CH3 299-42-3(3)

50-98-6(4)

134-72-5(5)

KB0700000

KB1750000

KB2625000

(D)-Ephedrine 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)NH·CH3 321-98-2(3)

24221-86-1(4)

KB0600000

KB1925000

(±)-MDEA 207.27 243.73 CH2

O2C6H3·CH2·CH(CH3)NH·C2H5 82801-81-8(3)

116261-63-2(4)

(±)-MDMA 193.24 229.71 CH2O2C6H3·CH2·CH(CH3)·NH·CH3 42542-10-9(3)

92279-84-0(4)

SH5700000

(+)-MDMA(7) 193.24 229.71 CH2

O2C6H3·CH2·CH(CH3)·NH·CH3 64057-70-1(4) SH5700000

(DL)-Methamphetamine 149.24 185.70 198.28 C6

H5·CH2·CH(CH3)·NH·CH3 4846-07-5(3)

(D)-Methamphetamine(7) 149.24 185.70 198.28 C6H5·CH2·CH(CH3)·NH·CH3 537-46-2(3)

51-57-0(4)

SH4910000

SH5455000

(L)-Methamphetamine 149.24 185.70 198.28 C6

H5·CH2·CH(CH3)·NH·CH3 33817-09-3(3) SH4905000

Phencyclidine 243.39 279.85 C6

H5·C[C5H10]·N[C5H10] 77-10-1(3)

956-90-1(4)

TN2272600

TN2272600

Phentermine 149.24 185.70 C6

H5·CH2·C(CH3)

2

·NH2 122-09-8(3)

1197-21-3(4)

SH4950000

(DL)-Norephedrine 151.21 187.67 200.25 C6

H5·CH(OH)·CH(CH3)·NH2 14838-15-4(3)

154-41-6(4)

RC2625000

DN4200000

1R,2S (-)-Norephedrine 151.21 187.67 200.25 C6

H5·CH(OH)·CH(CH3)·NH2 492-41-1(3) RC2275000

1S,2R (+)-Norephedrine 151.21 187.67 200.25 C6

H5·CH(OH)·CH(CH3)·NH2 37577-28-9(3)

1S,2S (+)-Norephedrine 151.21 187.67 200.25 C6H5·CH(OH)·CH(CH3)·NH2 36393-56-3

2153-98-2(4)

492-39-7(4)

RC9275000

(D)-Pseudoephedrine(8,9) 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)·NHCH3 90-82-4(3)

345-78-8(4)

UL5800000

UL5950000

(L)-Pseudoephedrine(10) 165.24 201.70 214.28 C6

H5·CH(OH)·CH(CH3)·NH·CH3 321-97-1(3)

(1) Molecular weights are calculated from the empirical formula using the 1987 IUPAC Atomic Weights of the Elements, Merck Index [17]. The molecular 

weight of the hemisulfate is 1⁄2 the weight of the 2:1 sulfate salt (2 moles amine + 1 mole H2

SO4).

(2) CAS from various sources: Merck Index [17], NIOSH RTECS [18], MSDS sheets from Sigma/Aldrich [19], Cerilliant [20], and other sources [21].

(3) Free base form.

(4) Hydrochloride salt.

(5) 2:1 Sulfate salt (2 moles amine + 1 mole H2SO4.

(6) RTECS = NIOSH Registry of Toxic Effects of Chemical Substances [18].

(7) More active isomer.

(8) Naturally occurring isomer.

(9) The D form of pseudoephedrine is a decongestant.

(10) The L form of pseudoephedrine is a bronchodilator. Dehydroxylation forms the less active L-methamphetamine.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 11 of 32

Table 2. Physical properties of analytes(1)

Compound (alphabetically) CAS m.p. (°C)

Vapor Pressure 

(mm Hg) pKa

(4) Log P(5)

Solubility in 

Water, g/100mL

(DL)-Amphetamine 300-62-9 — — 10.1 @ 20°C 1.76 2.8 @ 25°C

(D)-Amphetamine 51-64-9 <25 — 9.9(6) 1.76 —

(D)-Amphetamine sulfate 51-63-8 >300 — — 6.81 —

(L)-Amphetamine 156-34-3 — 0.201 @ 25 °C 10.1 @ 20°C 1.76 2.8 @ 25°C

Caffeine 58-08-2 238 15 @ 89°C 10.4 @ 40°C -0.07 2.16 @ 25°C

(DL)-Ephedrine 90-81-3 76.5 — — 0.68 —

(L)-Ephedrine 299-42-3 34 0.00083 @ 25°C 10.3 @ 0°C 1.13 63.6 @ 30°C

(L)-Ephedrine HCl 50-98-6 218 2.04E-10 @ 25°C pH 5.9 @ 

1/200 dil.(3) -2.45 25(6)

MDEA 82801-81-8 — — — — —

MDMA HCl 42542-10-9 148-149(2) — — — —

(D)-Methamphetamine 537-46-2 — 0.163 @ 25 °C 9.87 @ 25°C 2.07 1.33 @ 25°C

(D)-Methamphetamine HCl 51-57-0 170-175(2) — — — —

Phencyclidine 77-10-1 46.5 — 8.29(6) 4.69 —

Phencyclidine HCl 956-90-1 233-235(2) — — — —

Phentermine 122-09-8 — 0.0961 @ 25°C — 1.90 1.86 @ 25 °C

Phentermine HCl 1197-21-3 198(2) — — — —

(±) Phenylpropanolamine 14838-15-4 — 0.000867 @ 25°C 9.44 @ 20°C 0.67 14.9 @ 25°C

(±) Phenylpropanolamine HCl 154-41-6 194 — — -2.75 —

(L)-Norephedrine 492-41-1 51-53(3) — — — —

1S,2S (+)-Norephedrine 36393-56-3 77.5-78 0.000867 @ 25°C 9.44 @ 20 °C 0.83 14.9 @ 25 °C

1S,2S (+)-Norephedrine HCl 492-39-7 — — pH 5.9-6.1 in 

aq. soln.(3) 0.22 2 @ 25 °C

(D)-Pseudoephedrine 90-82-4 119 0.00083 @ 25°C 10.3 @ 0°C 0.89 10.6 @ 25°C

(D)-Pseudoephedrine HCl 345-78-8 181-182(2) — pH 5.9 @ 

1/200 dil.(3) — —

(1) Handbook of Physical Properties of Organic Chemicals unless otherwise noted [21].

(2) Merck Index [17].

(3) Sigma-Aldrich MSDS [19].

(4) Negative log of the acid dissociation constant for the amine in aqueous solution.

(5) Log P = octanol-water partition coefficient.

(6) Temperature not given in source.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 12 of 32

Table 3. Synonyms of analytes

Generic names(1) Trade and street names(2) Additional names(3)

(DL)-Amphetamine; 

(±)-Amphetamine

Benzedrine; Phenedrine; bennies (±)-α-Methylbenzeneethanamine(4); dl-α-Methylphenethylamine(4); dl-1-Phenyl-2-

aminopropane; (±)-Desoxynorephedrine

(D)-Amphetamine; 

(+)-Amphetamine

Dextroamphetamine; Dexedrine; dexies (S)-α-Methylbenzeneethanamine(4); d-α-Methylphenethylamine(4); d-1-phenyl-2-aminopropane; 

d-β-Phenylisopropylamine

(L)-Amphetamine; 

(-)-Amphetamine

Levoamphetamine; component of 

Adderall

(R)-α-Methylbenzeneethanamine(4); l-α-Methylphenethylamine(4); l-1-phenyl-2-aminopropane; 

(-)-1-phenyl-2-aminopropane

Caffeine Component (with ephedrine) of cloud 9 

and herbal XTC 

3,7-Dihydro-1,3,7-trimethyl-1H-purine-2,6-dione(4); 1,3,7-Trimethylxanthine

(DL)-Ephedrine; 

(±)-Ephedrine

Ephedral; Racephedrine; Sanedrine (R*,S*)-(±)-alpha-[2-(Methylamino)ethyl]benzenemethanol; DL-alpha-[1-(Methylamino)ethyl]

benzyl alcohol; dl-Ephedrine

(L)-Ephedrine; 

(-)-Ephedrine; (1R,2S)-(-)-

Ephedrine; l-Ephedrine

Primatene; Xenadrine; Ma Huang 

(Ephedra sinica and other species(5)); 

(with caffeine) cloud 9 and herbal 

ecstasy

(R-(R*,S*))-α-(1-Methylaminoethyl)benzenemethanol; L-erythro-2-(Methylamino)-

1-phenylpropan-1-ol; (1R,2S)-(-)-2-Methylamino-1-phenyl-1-propanol; (-)-alpha-(1-

Methylamino-ethyl)-benzyl alcohol; (-)-1-hydroxy-2-methylamino-1-phenylpropane; 

L-(-)-Ephedrine

(D)-Ephedrine (1S,2R)-(+)-2-Methylamino-1-phenyl-1-propanol; (+)-Ephedrine

MDEA MDE; Eve (±)-3,4-Methylenedioxy-N-ethylamphetamine; 

N-ethyl-alpha-methyl-1,3-benzodioxole-5-ethanamine

MDMA Adam, ecstasy N,α-Dimethyl-3,4-1,3-benzodioxole-5-ethanamine; 3,4-Methylenedioxymethamphetamine

(DL)-Methamphetamine; 

(±)-Methamphetamine

N,α-Dimethylbenzeneethanamine(4); N,α-Dimethylphenethylamine; dl-Desoxyephedrine; 

N-methyl-β-phenylisopropylamine

(D)-Methamphetamine; 

(+)-Methamphetamine; 

d-Methamphetamine

Methedrine; Desoxyn; chalk; crank; 

crystal; glass; ice; meth; speed; upper

(S)-N,α-Dimethylbenzeneethanamine; (S)-(+)-N,α-Dimethyl-phenethylamine(4); d-1-Phenyl-2-

methylaminopropane; d-Desoxyephedrine; d-N-methyl-β-phenyl-isopropylamine

(L)-Methamphetamine; 

(-)-Methamphetamine

Component in decongestant vapor 

inhaler (Vick’s brand)

(R)-(-)-N,α-Dimethylphenethylamine; (-)-Deoxyephedrine; 

(-)-2-(Methylamino)-1-phenylpropane 

Phencyclidine Sernylan; Sernyl; angel dust; PCP; 

peace pill

1-(1-Phenylcyclohexyl) piperidine(4)

Phentermine Fastin; Normephentermine α,α-Dimethylbenzeneethanamine(4); α,α-Dimethylphenethylamine(4); 1,1-Dimethyl-2-

phenylethylamine; α-Benzylisopropylamine

(DL)-Norephedrine; 

(±)-Norephedrine

(±)-Phenylpropanolamine; Obestat; 

Phenedrine;

(R*,S*)-(±)-α-(1-Aminoethyl)benzenemethanol(4); -(±)-α-(1-Amino-ethyl)benzyl alcohol(4); 

(±)-2-Amino-1-phenyl-1-propanol

(L)-Norephedrine; 

(-)-Norephedrine

Natural form found in Ephedra sinica 

and other species(5)

(1R,2S)- 2-Amino-1-phenyl-1-propanol; (1R,2S)-Norephedrine; 

l-erythro-2-Amino-1-phenylpropan-1-ol

(D)-Norephedrine; 

(+)-Norephedrine

Metabolite of cathinone in urine of 

Khat users.

(1S,2R)- 2-Amino-1-phenyl-1-propanol; (1S,2R)-Norephedrine; 

d-erythro-2-Amino-1-phenylpropan-1-ol

(+)-Norpseudoephedrine; 

Cathine

Amorphan; Adiposettin; Reduform; 

found naturally in Khat plant

(R*,R*)-α-(1-Aminoethyl)benzenemethanol(4); d-threo-α-2-Amino-1-hydroxy-1-phenylpropane; 

1S,2S-(+)-Norpseudoephedrine

L-(+)-Pseudoephedrine; 

(+)-Pseudoephedrine; 

d-Pseudoephedrine

Afrinol; Novafed; Sinufed; Sudafed; 

natural form found in Ephedra sinica 

and other species(5)

(S-(R*,R*))-α-[1-(Methylamino)ethyl]benzenemethanol; (1S,2S)-(+)-2-Methylamino-1-

phenylpropanol; d-(alpha-(1-Methylamino)-ethyl)benzyl alcohol; (1S,2S)-(+)-Pseudoephedrine; 

d-threo-2-Methylamino-1-phenylpropan-1-ol; (+)-ψ-Ephedrine

D-(-)-Pseudoephedrine; 

(-)-Pseudoephedrine

(1R,2R)-(-)-Pseudoephedrine; (-)-ψ-Ephedrine; l-threo-2-Methylamino-1-phenylpropan-1-ol; 

(+)-ψ-Ephedrine

(1) Common or generic names. Salts forms are not given for simplicity.

(2) Trade and street names are exemplary, not exhaustive. Street names change over time and by locality. Salts and free base forms are not distinguished.

(3) Other names from Merck Index [17], NIOSH Registry of Toxic Effects of Chemical Substances [18], and MSDS sheets [19,20]. NOTE: For amphetamine and 

methamphetamine the prefixes R-, D-, d-, and (+)- , although they mean different things, are essentially synonymous for the dextrorotatory stereoisomer and S-, L-, l-, 

and (-)- are essentially synonymous for the levorotary stereoisomer. Many other synonyms exist.

(4) Uninverted CAS name as given in Merck Index [17].

(5) Extracts of Ephedra species contain various amounts of (+)-Norephedrine, (-)-N-methylephedrine, and (+)-N-methylpseudoephedrine. (+)-Norephedrine is reduced to 

amphetamine and N-methylephedrine and N-methylpseudoephedrine reduce to N,N-dimethylamphetamine [22, 23]. The presence of these latter two compounds in 

methamphetamine samples indicate that Ephedra spp. extracts may have been used in the synthesis [24].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 13 of 32

Table 4. Limit of detection (LOD), method detection limit (MDL), and sample storage stability(1)

Compound Int. std.(2)

Estimated LOD(3)

μg/sample 

liq. stds(6)

μg/sample 

liq. stds(7)

Estimated MDL(4)

μg/sample 

cotton gauze

μg/sample 

Synthetic Wipe

Storage Stability(5)

30 days 4 oC 7 days 22 oC

(D)-Amphetamine D11-Amp 0.1 0.1 0.02 100.5 94.5

D14-Met 0.1 0.05 0.02 0.02 99.7 87.9

NMPhen 0.1 0.04 - -

Cocaine D11-Amp 0.6 0.2 (9) 99.3 98.8

D14-Met 0.4 0.1 (9) 0.1 (9) 98.5 91.9

NMPhen 0.4 0.1 (9) - -

(L)-Ephedrine D11-Amp 0.2 0.2 0.02 95.6 97.2

D14-Met 0.1 0.1 0.02 0.02 94.8 90.5

NMPhen 0.1 0.02 - -

MDEA N-PAmp 0.1 0.06 0.1 98.9 102.1

MDMA D11-Amp 0.1 0.02 99.7 111.1

D14-Met 0.1 0.02 0.04 98.9 103.2

NMPhen 0.1 0.03 - -

(D)-Methamphetamine D11-Amp 0.2 0.07 0.02 98.7 100.6

D14-Met 0.1 0.05 0.02 0.02 98.0 93.5

NMPhen 0.1 0.02 - -

Phencyclidine D11-Amp 0.6 0.1 (9) 103.7 105.2

D14-Met 0.4 0.1 (9) 0.5(9) 102.9 97.7

NMPhen 0.4 0.1 (9) - -

Phentermine D11-Amp 0.2 0.03 102.0 101.5

D14-Met 0.1 0.03 0.03 101.1 94.3

NMPhen 0.1 0.04 - -

(±)-Norephedrine(8) D11-Amp 0.1 0.05 0.03 94.3 92.7

D14-Met 0.1 0.05 0.03 0.03 93.6 86.2

NMPhen 0.1 0.03 - -

Pseudoephedrine D11-Amp 0.2 0.2 0.02 100.4 97.9

D14-Met 0.1 0.1 0.02 0.02 99.6 91.1

NMPhen 0.1 0.02 - -

(1) Backup Data Report [4].

(2) Internal standards: D11-Amp = Amphetamine-D11, D14-Met = Methamphetamine-D14, NMPhen = N-Methyl phenethylamine, N-PAmp = N-Propyl amphetamine.

(3) LODs vary according to individual GC columns, instrument conditions and cleanliness, media interferences, and internal standards used. LODs were calculated on liquid standards using 

the procedure of Burkart (LODs for linear calibration curves are calculated as 3 times the standard error of the lowest three standards analyzed in replicate divided by the slope of the 

calibration curve) [12].

(4) MDLs are provided to satisfy regulatory agencies requiring this expression of sensitivity. These MDLs are calculated as the standard deviation of six replicates on spiked media analyzed at 

the 0.1 μg/sample level (except as noted) times the Student’s t value for 6 replicates (3.365). (Normally 7 replicates are required.)

(5) Cotton gauze samples were spiked at 3 μg/sample per analyte. Six samples were analyzed immediately after preparation. Six samples were stored at room temperature (about 22 oC) for 

7 days and then analyzed. Eighteen samples were stored at >6 oC. Of the 18 samples stored at >6 oC, six each were analyzed at 7 and 21 days and three each were analyzed at 14 and 30 

days (Backup Data Report [4].) Apparent recoveries vary according to internal standard used.

(6) These LODs are conservative since the lowest calibration standard for these determinations was 0.1 μg/sample. Lower LODs are achievable with lower concentration calibration standards 

and operation of the mass spectrometer in the SIM mode.

(7) Typical LODs for a five point calibration curve with single standards at each concentration level. The lowest calibration standard for these determinations was 0.05 μg/sample.

(8) (±)-Norephedrine = (±)-phenylpropanolamine.

(9) MDLs for cocaine and phencyclidine were determined from the 0.3 μg/sample level because the GC peaks for the 0.1 μg/sample level were un-measurable. Precisions at the 0.3 μg/

sample level were such that the MDLs calculated to 0.1 μg/sample anyway. This value may be realistic since the 0.1 μg/sample level samples had been stored for one month prior to 

analysis which may have affected stability.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 14 of 32

Table 5. Example of mass spectrometer operation parameters for selected ion monitoring mode(1)

Heptafluorobutyryltrimethyl-silyl derivatives Scan window(2) Acquisition ions (m/z) per group(3)

Acquisition Group 1 8.20 to 10.20 104 118 128 132 210 213 240 244 254 261

Acquisition Group 2 10.20 to 13.20 179 240 254 282 296 456

Acquisition Group 3 13.20 to 19.00 82 162 182 200 242 254 268

GC 

Peak 

No.(4)

Target Analytes and Internal 

Standards

Retention 

Time(6)(min)

Primary Ion (m/z)

(7)

(Quantification Ion)

Secondary ion and approximate relative 

abundance(8)(relative to the Primary 

Ion)

Acquisition Group 1:

13 Amphetamine-D11 (I$)(9) 8.46 244 128 70%

5 Amphetamine 8.54 240 118 70%

92 Phentermine 8.72 254 132 12%

81 N-Methyl phenethylamine (I$)(9) 8.54 240 104 100%

68 Methamphetamine-D14 (I$)(9) 9.86 261 213 30%

64 Methamphetamine 9.94 254 210 35%

Acquisition Group 2:

95 Phenylpropanolamine 10.49 179 240 18%

97 N-Propylamphetamine (I$)(9) 11.05 282 240 85%

36 Ephedrine 11.40 179 254 17%

98 Pseudoephedrine 11.68 179 254 15%

32 Dibromooctafluorobiphenyl(10) 12.82 296 456 100%

Acquisition Group 3:

59 MDMA 13.81 254 162 80%

57 MDEA 14.19 268 162 60%

86 Phencyclidine 15.62 200 242 35%

27 Cocaine 18.65 182 82 110%

(1) In this example, 10 analytes and 5 internal standards are grouped into 3 acquisition groups having no more than 10 primary and 

secondary ions per acquisition group. For 6 analytes and internal standards or less, one acquisition group may be sufficient.

(2) Scan window is in minutes. Actual times are dependent upon GC column and instrument conditions.

(3) Ions (m/z) in bold numbers are suggested primary (quantification) ions. For best signal to noise ratio, do not exceed 10 ions per 

acquisition group. Dwell time per ion (m/z) is 50 milliseconds.

(4) GC peak numbers are those in Figures 1 and 2 and Table 10.

(5) The list of analytes and internal standards shown is an example. Analyte(s) and internal standard(s) must be selected according to 

analytical objectives.

(6) Retention times are dependent upon GC column and instrument conditions.

(7) The better ions for quantification are usually the base peak or those with masses >100 m/z and relative abundances >50% of the 

base peak. These minimize interference from co-eluting hydrocarbons. The suggested primary ions are not necessarily the base peaks 

in the mass spectra of the analytes, especially if the base peaks are ions common to aromatics (e.g., m/z 91) and paraffinic or olefinic 

hydrocarbons (e.g., m/z 42, 57, and 58). Suggested ions for other analytes and internal standards are given in Table10.

(8) Secondary ions may be used for quantification if the primary ion encounters interference. Secondary ions improve qualitative 

identification for SIM analyses. The relative abundances given are approximate (±10 to 20%) and depend upon specific instrument 

tuning and conditions. They are relative to the primary ion and not necessarily to the base peak in the mass spectrum of each analyte. The 

relative abundance of secondary ions for each analyte needs to be determined from a mass spectrum acquired on the instrument to be 

used.

(9) (I$) = internal standard. Internal standards must be paired with the appropriate analytes. Tables 7a and 7b give precision and accuracy 

data for various pairings. Other potentially useful internal standards are given in Tables 8a, 8b, and 10. Highly deuterated analogs of the 

target analytes are preferred, where available.

(10) Dibromooctafluorobiphenyl is an optional secondary internal standard useful for monitoring autosampler performance and instrument 

tuning. A shift in the mass axes or the relative abundance of m/z 296 to that of m/z 456 throughout an analytical sequence will help signal 

degraded tuning.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 15 of 32

Table 6. Suggested spiking schedule for calibration standards and quality control samples

Name

Add the following to clean shipping containers (e.g., 50-mL polypropylene centrifuge 

tubes) in the following order.

Number 

of 

Wipes(1,2)

Volume(2) of 

Isopropanol 

or 

Methanol(3)

Volume(2)

of Internal 

Standard 

Spiking 

Solution(4,5)

Volume 

of Target 

Analyte 

Spiking 

Solution(5,6)

Volume 

of Spiking 

Solution 

diluted 

1/20(5,7)

Volume(2) of 

Desorption 

Solution(8)

Resulting 

μg/sample 

as Free 

Base(9)

Calibration Standards(10)

CS0 0 3 mL 60 μL 0.0 μL 30 mL 0.00

CS1 0 3 mL 60 μL 2 μL 30 mL 0.02

CS2 0 3 mL 60 μL 5 μL 30 mL 0.05

CS3 0 3 mL 60 μL 10 μL 30 mL 0.1

CS4 0 3 mL 60 μL 20 μL 30 mL 0.2

CS5 0 3 mL 60 μL 60 μL 30 mL 0.6

CS6 0 3 mL 60 μL 10 μL 30 mL 2.0

CS7 0 3 mL 60 μL 30 μL 30 mL 6.0

CS8 0 3 mL 60 μL 100 μL 30 mL 20

CS9 0 3 mL 60 μL 300 μL 30 mL 60

CS10 0 3 mL 60 μL 1000 μL 30 mL 200

Quality Control Samples(11)

QB (media blank) 1 3 mL 60 μL 0.0 μL 30 mL 0.0

QC (matrix spike) 1 3 mL 60 μL 3-300 μL or 20-60 μL 30 mL 0.2-60

QD (matrix spike 

duplicate) 1 3 mL 60 μL 3-300 μL or 20-60 μL 30 mL 0.2-60

(1) Gauze wipes may be added to the calibration standards but are not necessary if cotton gauze is used. Blank gauze wipes must always be added to the quality 

control samples, QB, QC, and QD.

(2) a. If a sample consists of 2 gauze wipes, the volume of desorption solution must be increased to 40 mL to accommodate the second wipe. The shipping 

container should be a 50-mL polypropylene centrifuge tube or equivalent to accommodate the extra volume of desorption solution for 2 wipes. It is not critical 

to know the exact volume of desorption solution and wetting alcohol used per sample. It only needs to be enough to cover the samples and to permit free 

percolation through the samples. See step 7.

b. If a set of samples consists predominantly of 2 gauze wipes, the QB, QC, and QD should also consist of 2 wipes and treated as per the samples. The volume of 

isopropanol (or methanol) added to the QC samples should be increased to 4 mL for 2 gauze wipes to simulate samples containing 2 gauze wipes.

(3) If methanol was used for wipe sampling, it should also be used in the calibration standards, blanks, and QCs instead of isopropanol.

(4) Concentration of internal standards in the internal standard spiking solution is approximately 200 μg/mL as the free base. It is critical to know the exact 

volume of internal standard spiking solution that is added to the calibration standards, samples, blanks, and quality control samples. The volume spiked into 

the samples may vary with sample size but the volume spiked into each of the calibration standards must not vary. See step 7b. 

(5) For quality control samples, spike onto wipe media within the shipping container. For liquid calibration standards (in lieu of media calibration standards), spike 

into the isopropanol (or methanol).

(6) Concentration of analytes in the target analyte spiking solution is approximately 200 μg/mL as the free base.

(7) Concentration of analytes in the diluted spiking solution for this table is approximately 10.0 μg/mL as the free base and can be prepared by diluting 100 μL of 

the target analyte spiking solution to 2 mL in methanol.

(8) Desorption solution is 0.1 M sulfuric acid in deionized water.

(9) This is μg per total sample irrespective of the total desorption solution volume or the area wiped.

(10) Select 6 calibration standards from the list to cover the analytical range plus the blank.

(11) Prepare one set of quality control samples for every 20 samples or less.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 16 of 32

Table 7a. Precision and accuracy in scan mode for cotton gauze(1)

Compound

Internal 

Standard(2)

Range(3)

μg/sample Accuracy

Overall 

Precision 

Bias

Average Range

(D)-Amphetamine D11-Amp 0.1-30 8.1 0.0412 -0.0054 -0.0386 to +0.0428

D14-Met 0.1-30 10.3 0.0472 -0.0227 -0.0844 to +0.0199

NMPhen 0.1-30 13.2 0.0662 -0.0120 -0.0931 to +0.0290

Cocaine D11-Amp 1.0-30 15.8 0.0469 +0.0810 +0.0416 to +0.1375

D14-Met 3-30 13.3 0.0422 +0.0631 +0.0003 to +0.1294

NMPhen 0.3-30 20.2 0.0729 +0.0823 -0.0092 to +0.1359

(L)-Ephedrine D11-Amp 0.1-30 9.8 0.0499 -0.0052 -0.0608 to +0.0262

D14-Met 0.1-30 9.2 0.0397 -0.0266 -0.0463 to +0.0221

NMPhen 0.1-30 11.2 0.0493 -0.0284 -0.0775 to +0.0302

MDEA N-PAmp 0.3-29 12.4 0.0618 +0.0127 -0.0475 to +0.0869

MDMA D11-Amp 0.1-27 14.3 0.0568 +0.0497 +0.0104 to +0.1197

D14-Met 0.1-27 13.1 0.0558 +0.0389 -0.0189 to +0.0978

NMPhen 0.3-27 11.9 0.0605 +0.0007 -0.0570 to +0.0360

(D)-Methamphetamine D11-Amp 0.1-10 9.2 0.0395 +0.0270 -0.0289 to +0.0923

D14-Met 0.1-30 5.9 0.0302 +0.0015 -0.0440 to +0.0592

NMPhen 0.3-30 6.9 0.0334 +0.0113 -0.0534 to +0.0448

Phencyclidine D11-Amp 0.3-30 17.2 0.0639 +0.0670 +0.0059 to +0.1222

D14-Met 0.3-30 15.9 0.0648 +0.0521 -0.0386 to +0.1039

NMPhen 0.3-30 16.0 0.0638 +0.0547 -0.0474 to +0.0886

Phentermine D11-Amp 0.1-30 10.1 0.0444 +0.0261 -0.0067 to +0.0912

D14-Met 0.1-30 10.4 0.0527 +0.0041 -0.0600 to +0.0674

NMPhen 1.0-30 8.2 0.0400 +0.0121 -0.0378 to +0.0407

(±)-Norephedrine(4) D11-Amp 0.1-30 12.2 0.0571 +0.0241 +0.0500 to +0.0610

D14-Met 0.1-30 12.5 0.0638 -0.0005 -0.0674 to +0.0708

NMPhen 0.1-30 13.3 0.0675 +0.0036 -0.0533 to +0.0476

Pseudoephedrine D11-Amp 0.1-30 10.0 0.0507 -0.0059 -0.0530 to +0.0441

D14-Met 0.1-30 12.3 0.0507 -0.0392 -0.0737 to +0.0301

NMPhen 1.0-30 15.6 0.0716 -0.0350 -0.0813 to +0.0617

(1) Backup Data Report [4]. Values are for the heptafluorobutyryl and mixed heptafluorobutyryl-trimethylsilyl derivatives and analysis by GC-MS in scan mode. 

Each sample consisted of a pair of 3” x 3” 12-ply cotton gauze pads. There were 6 replicate samples per concentration level and six concentration levels 

evaluated from approximately 0.1 to 30 μg/sample.

(2) Internal Standards Deuterated: Non-deuterated:

D11-Amp = Amphetamine-D11 NMPhen = N-Methyl phenethylamine

 D14-Met = Methamphetamine-D14 N-PAmp = N-Propyl amphetamine

(3) Range used for calculation of precision, accuracy, and bias. The entire range studied for all analytes was approximately 0.1 to 30 μg/sample (1xLOQ to 

300xLOQ).

(4) (±)-Norephedrine = (±)-phenylpropanolamine.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 17 of 32

Table 7b. Precision and accuracy in scan mode for synthetic wipe(1)

Compound

Internal 

Standard(2)

Range(3) μg/

sample Accuracy Overall Precision 

Bias

Average Range

(D)-Amphetamine D14-Met 0.1-30 17.2 0.0611 -0.0712 -0.1066 to -0.0468

Cocaine D14-Met 0.3-30 17.7 0.0901 -0.0014 -0.0246 to +0.0252

(L)-Ephedrine D14-Met 0.1-30 10.7 0.0432 -0.0362 -0.0638 to -0.0039

MDEA N-PAmp 0.3-29 9.6 0.0425 -0.0240 -0.0453 to +0.0416

MDMA D14-Met 0.3-27 11.4 0.0498 -0.0297 -0.0612 to +0.0095

(D)-Methamphetamine D14-Met 0.1-30 8.7 0.0430 -0.0114 -0.0483 to +0.0625

Phencyclidine D14-Met 0.3-30 13.0 0.0391 +0.0658 +0.0216 to +0.1418

Phentermine D14-Met 0.3-30 10.4 0.0295 -0.0560 -0.0917 to -0.0266

(±)-Norephedrine(4) D14-Met 0.1-30 12.6 0.0577 +0.0282 -0.0220 to +0.0937

Pseudoephedrine D14-Met 0.1-30 13.5 0.0592 -0.0352 -0.1001 to -0.0020

(1) Backup Data Report [4]. Values are for the heptafluorobutyryl and mixed heptafluorobutyryl-trimethylsilyl and analysis by GC-MS in scan mode (see p 9109-1 

for GC and MS conditions). Each sample consisted of a pair of 3” x 3” 12-ply cotton gauze pads. There were 6 replicate samples per concentration level and six 

concentration levels evaluated from approximately 0.1 to 30 μg/sample.

(2) Internal Standards: D14-Met = Methamphetamine-D14, N-PAmp = N-Propyl amphetamine.

(3) Range used for calculation of precision, accuracy, and bias. The entire range studied for all analytes was approximately 0.1 to 30 μg/sample (1xLOQ to 300xLOQ).

(4) (±)-Norephedrine = (±)-phenylpropanolamine.

Table 8a. Recommended internal standards(1) and best application

COMPOUND NAME CAS

MW as

free base

Quant.

Ion

Secondary

Ion COMMENTS

(±)-Amphetamine-D11 not available 146.12 244 128 Preferred analog for amphetamine

(±)-Amphetamine-D8 145225-00-9 143.15 243 126 Alternate for amphetamine-D11

(±)-Amphetamine-D6 not available 141.16 244 123 Alternate for amphetamine-D11

(±)-Methamphetamine-D14 not available 163.12 261 213 Preferred methamphetamine analog

(±)-Methamphetamine-D11 152477-88-8 160.15 260 213 Alternate for methamphetamine-D14

(±)-Methamphetamine-D9 not available 158.16 261 213 Alternate for methamphetamine-D14

N-Methylphenethylamine 589-08-2 135.23 240 104 Alternate for methamphetamine-D14

Phencyclidine-D5 60124-86-9 248.35 205 96 Use only for phencyclidine

MDEA-D6

(2) 160227-44-1 213.22 268 162 Use only for MDEA

N-Propylamphetamine(2) not available 177.29 282 240 Alternate for MDEA-D6

(1) Care must be exercised in the selection of internal standards for each analyte because of differences in derivatization efficiencies due to structural differences.

a. Deuterated analogs of each target analyte may be acceptable as internal standards if they are isotopically pure enough and their ions do not interfere with the 

quantification ions (usually base peaks) of the target analyte, especially at the limit of detection for the target analyte. Conversely it is also important that ions 

in the target analyte, especially at high concentrations, do not interfere with the quantification ion (usually base peaks) of any deuterated analog used as the 

internal standard.

b. The more highly deuterated an analog, the more it will chromatographically separate from the target analyte, reducing interference from common ions.

c. Phentermine and mephentermine have been used as internal standards. Such use is not advised in this method because of their reported occasional use as 

adulterants in certain illicit drugs such as MDMA.

(2) N-Propylamphetamine and MDEA-D6

 are only applicable to MDEA and other hindered amines (e.g., fenfluramine and MBDB) due to similar steric hindrance at the 

nitrogen (N-ethyl or N-propyl substitution) which affects derivatization efficiency.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 18 of 32

Table 8b. Recommended best application of internal standards(1)

Target Analyte

Recommended Deuterated Internal Standards

Amphetamine-D11

(2)

Methamphetamine-D14

(2) MDEA-D6(1) Phencyclidine-D5

Recommended Alternate Non-deuterated 

Internal Standards(3)

N-Methylphenethylamine N-Propylamphetamine(1)

Amphetamine X X X

Cocaine X X X

Ephedrine X X X

MDEA X X

MDMA X X X

Methamphetamine X X X

Phencyclidine X X X X

Phentermine X X X

(±)-Norephedrine(4) X X X

Pseudoephedrine X X X

(1) N-Propylamphetamine and MDEA-D6 are only applicable to MDEA and other hindered amines (e.g., fenfluramine and MBDB) due to similar steric hindrance at 

the nitrogen (N-ethyl or N-propyl substitution) which affects derivatization efficiency.

(2) The alternate deuterated compounds listed in part A above may be used. Avoid ring-labeled amphetamine-D5

 (CAS 65538-33-2) since the primary 

(quantification) ion is the same as for amphetamine and GC peaks overlap significantly. Also avoid methamphetamine-D5

 (CAS 60124-88-1) since GC peaks 

significantly overlap.

(3) The listed non-deuterated compounds are acceptable as internal standards for the listed target analytes for the applicable ranges and limits of detection listed 

in Tables 7a and 7b respectively. Non-deuterated internal standards might not be permissible. Consult regulations of agency having legal jurisdiction.

(4) (±)-Norephedrine is the same as (±)-phenylpropanolamine.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 19 of 32

Table 9a. Recovery from wall (latex painted) with various solvents; one wipe compared with the sum of 

two wipes(1,2)

Test Compounds(5)

Water(3)

First Wipe

Plus 

Second 

Wipe(4)

Isopropanol

First Wipe

Plus 

Second 

Wipe(4)

Methanol

First Wipe

Plus 

Second 

Wipe(4)

Percent %RSD Percent Percent %RSD Percent Percent %RSD Percent

Amphetamine 51 14 56 67 6.0 78 90 4.0 96

Cocaine 36 22 36 69 22 80 89 9.1 94

Ephedrine 48 23 52 76 7.4 85 91 4.4 96

MDMA 40 20 44 61 9.0 70 88 5.3 94

MDEA 45 22 50 69 12 80 90 11 97

Methamphetamine 46 16 50 64 7.4 75 87 3.5 94

Phencyclidine 27 26 30 64 9.6 73 86 5.2 91

Phentermine 53 9.2 58 78 6.6 91 95 2.9 101

Phenylpropanolamine 58 21 62 80 9.3 95 85 5.0 94

Pseudoephedrine 49 20 53 73 7.0 85 95 3.3 101

(1) Backup Data Report for NIOSH 9109 [4]. Area of each sample was 100 cm2.

(2) Wall was an existing standard gypsum board wall painted with a latex based paint. Painted surface was at least one year old. There were six replicates for each 

solvent tested. 

(3) Water was deionized water (ASTM type II). Note low recovery and high %RSD.

(4) For the serial wipe study, each 100-cm2 area was wiped again with a fresh pre-wetted gauze wipe and the amount recovered was determined separately. In 

practice, a second (serial) wipe is included with the first gauze wipe; both gauze wipes constitute a single sample. The percent recoveries shown in the column 

represent the sum of the amounts recovered in both the first and second wipes.

(5) Each pre-measured area was spiked with 3 μg of each analyte in methanol and the methanol allowed to dry for several minutes prior to wipe sampling.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 20 of 32

Table 9b. Recovery of methamphetamine from various surfaces with various solvents; one wipe 

compared with the sum of two wipes(1)

Surface Material(3) Replicates

Isopropanol

First Wipe

Percent %RSD

Plus Second 

Wipe(2)

Percent

Methanol

First Wipe

Percent %RSD

Plus Second 

Wipe(2)

Percent

Enamel (lid of 

washing machine) 

4(4) 58 5.7 68 81 2.4 87

Vinyl veneer on 

particle board 

4 (5) 60 5.2 68 81 4.8 89

Latex painted wall 6(4) 64 7.4 75 87 3.5 94

Refrigerator door 2(5) 65 2.9 76 91 4.0 92

Varnished 

hardwood panel 

2(6) 72 5.4 76 82 3.7 86

Formica® 

countertop 

4(5) 75 4.9 82 87 3.8 91

(1) Backup Data Report for NIOSH 9109 [4]. Area of each sample was 100 cm2.

(2) For the serial wipe study, each 100-cm2 area was wiped again with a fresh pre-wetted gauze wipe and the amount recovered was determined separately. In 

practice, a second (serial) wipe is included with the first gauze wipe; both gauze wipes constitute a single sample. The percent recoveries shown in the column 

represent the sum of the amounts recovered in both the first and second wipes.

(3) The refrigerator door and the washing machine lid were from used appliances. The vinyl-veneered particle board (a book shelf), the Formica® countertop, and 

the varnished hardwood paneling were all purchased new. All surfaces of used and new materials were pre-cleaned with multiple rinses of methanol prior to 

spiking. Each pre-measured 100-cm2 square was spiked with 3 μg methamphetamine.

(4) Samples were taken using the side-to-side and then top-to-bottom wiping technique.

(5) Half of the samples were wiped using the side-to-side wiping technique and half were wiped using the concentric squares wiping technique. There were no 

significant differences in recoveries. Percent recoveries and %RSDs are for both techniques combined.

(6) Samples were taken each time using only top-to-bottom wiping with the grain of the wood in an “N” pattern.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 21 of 32

TABLE 10. Gas chromatographic retention times for heptafluorobutyryl and trimethylsilyl derivatives of 

selected drugs of abuse, precursors, and potential adulterants(1)

GC 

Peak 

No. Compound Derivative Form(2) Notes(3)

Retention 

Time 

Minutes(4)

Relative Retention Time Ions (Significant m/z)

(7)

(5) (6) 1’ 2’(7) 3’(7)

1 Acetaminophen(8) N,N’- bis-TMS- Pri.deriv. 12.30 0.9594 1.2374 206 280 [90] 295 [70]

2 Acetaminophen(8) N-HFB-N’-TMS- Minor peak 10.37 0.8089 1.0433 330 404 [80] 419 [30]

3 Aminorex N,N’- bis-HFB- Major peak 14.12 1.1014 1.4205 385 342 [30] 169 [40]

4 Aminorex N-HFB-N’-TMS- Major peak 16.59 1.2941 1.6690 261 146 [48] 128 [45]

5 Amphetamine N-HFB- Pri.deriv. 8.54 0.6661 0.8592 240 118 [70] 169 [20]

6 Amphetamine N-HFB-N-TMS- OS artifact 9.21 0.7184 0.9266 312 91 [50] 313 [10]

7 Amphetamine-D5

, ring labeled (I$)(9) N-HFB- Pri.deriv. 8.47 0.6607 0.8521 240 123 [85] 96 [55]

8 Amphetamine-D5

,ring labeled (I$)(9) N-HFB-N-TMS- OS artifact 9.17 0.7153 0.9225 312 96 [45] 73 [95]

9 Amphetamine-D6

 (I$)(9) N-HFB- Pri.deriv. 8.45 0.6591 0.8501 244 123 [70] 93 [45]

10 Amphetamine-D6

 (I$)(9) N-HFB-N-TMS- OS artifact 9.14 0.7129 0.9195 316 93 [40] 73 [75]

11 Amphetamine-D8

 (I$)(9) N-HFB- Pri.deriv. 8.46 0.6599 0.8511 243 126 [75] 96 [40]

12 Amphetamine-D8

 (I$)(9) N-HFB-N-TMS- OS artifact 9.16 0.7145 0.9215 315 96 [25] 73 [55]

13 Amphetamine-D11 (I$)(9) N-HFB- Pri.deriv. 8.46 0.6599 0.8511 244 128 [70] 98 [45]

14 Amphetamine-D11 (I$)(9) N-HFB-N-TMS- OS artifact 9.14 0.7129 0.9195 316 98 [60] 73 [70]

15 Atropine(8) O-TMS- Pri.deriv. 18.86 1.4711 1.8974 124 361 [9] 82 [17]

16 BDB N-HFB- Pri.deriv. 13.35 1.0413 1.3431 135 176 [50] 254 [12]

17 BDB N-HFB-N-TMS- OS artifact 13.65 1.0647 1.3732 326 135 [60] 73 [90]

18 Benzoyl ecgonine O-TMS- 19.18 1.4961 1.9296 82 240 [45] 361 [25]

19 Benzyl piperazine(10) (“Legal XTC”) N-HFB- Pri.deriv. 13.73 1.0710 1.3813 91 372 [30] 281 [30]

20 4-Bromo-2,5-DMPEA(11) (Nexus) N-HFB- Pri.deriv. 15.79 1.2317 1.5885 242 244 [98] 229 [75]

21 4-Bromo-2,5-DMPEA(11) (Nexus) N-HFB-N-TMS- OS artifact 16.22 1.2652 1.6318 229 231 [98] 298 [85]

22 Bupropion (Wellbutrin®, Zyban®) parent 12.15 0.9477 1.2223 44 100 [45] 111 [20]

23 Caffeine(8) parent 14.89 1.1615 1.4980 194 109 [45] 67 [45]

24 S-(-)-Cathinone (from Khat plant) N-HFB- Pri.deriv. 10.21 0.7964 1.0272 105 77 [45] 240 [15]

25 S-(-)-Cathinone (from Khat plant) N-HFB-N-TMS- OS artifact 10.89 0.8495 1.0956 105 312 [68] 77 [55]

26 Chlorpheniramine(8) parent 16.74 1.3058 1.6841 203 205 [32] 167 [22]

27 Cocaine parent 18.65 1.4548 1.8763 82 182 [90] 303 [20]

28 Codeine O-HFB- Minor peak 19.59 1.5281 1.9708 282 283 [20]

29 Codeine O-TMS- Pri.deriv. 20.72 1.6162 2.0845 371 343 [25] 234 [55]

30 Dextromethorphan(8) parent 18.10 1.4119 1.8209 271 270 [62] 214 [40]

31 Diazepam (Valium® etc.) parent 20.80 1.6225 2.0926 256 283 [90] 284 [75]

32 Dibromooctafluorobiphenyl (I$)(9) parent 12.82 1.0000 1.2897 296 456 [100] 454 [50]

33 N,N-Dimethyltryptamine (DMT) N-HFB- Pri.deriv. 13.00 1.0140 1.3078 58 129 [15] 42 [15]

34 N,N-Dimethyltryptamine (DMT) N-TMS- Minor peak 15.02 1.1716 1.5111 58 73 [12] 202 [10]

35 Ecgonine, methyl ester O-TMS- 11.72 0.9142 1.1791 82 96 [75] 83 [75]

36 Ephedrine N-HFB-O-TMS- Pri.deriv. 11.40 0.8892 1.1469 179 254 [17] 327 [10]

37 1S,2R(+)-Ephedrine-D3

 (I$)(9) N-HFB-O-TMS- Pri.deriv. 11.36 0.8861 1.1429 179 257 [20] 330 [10]

38 N-Ethyl amphetamine N-HFB- Pri.deriv. 10.33 0.8058 1.0392 268 240 [35] 118 [15]

39 Fenfluramine(8) N-HFB- Pri.deriv. 10.12 0.7894 1.0181 268 240 [35] 159 [22]

40 Fenfluramine-D10 (I$)(9) N-HFB- Pri.deriv. 10.01 0.7808 1.0070 277 245 [35] 160 [15]

Table 10 continued
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 22 of 32

TABLE 10 (continued). Gas chromatographic retention times for heptafluorobutyryl and trimethylsilyl 

derivatives of selected drugs of abuse, precursors, and potential adulterants(1)

GC 

Peak 

No. Compound

Derivative 

Form(2) Notes (3)

Retention 

Time 

Minutes (4)

Relative Retention Time Ions (Significant m/z) (7)

(5) (6) 1’ 2’ (7) 3’ (7)

41 Fentanyl (Sublimaze® etc.) parent 22.97 1.7917 2.3109 245 146 [60] 189 [33]

42 Flunitrazepam (Rohypnol®, roofies)(10) parent 22.20 1.7317 2.2334 312 285 [95] 286 [90]

43 Hydrocodone (Lortab® etc.) HFB- Minor peak 19.47 1.5187 1.9588 495 438 [50] 298 [40]

44 Hydrocodone (Lortab® etc.) TMS- Minor peak 20.82 1.6240 2.0946 371 356 [50] 234 [55]

45 Hydrocodone (Lortab® etc.) parent Pri.deriv. 20.93 1.6326 2.1056 299 242 [50] 243 [35]

46 Hydromorphone (Dilaudid®) O-HFB-O’-TMS- Minor peak 19.85 1.5484 1.9970 308 267 [92] 358 [75]

47 Hydromorphone (Dilaudid®) O,O’-bis-TMS- Minor peak 20.98 1.6365 2.1107 414 429 [100] 234 [75]

48 Hydromorphone (Dilaudid®) O-TMS- Pri.deriv. 21.21 1.6544 2.1338 357 300 [55] 342 [28]

49 Ketamine (“special K”)(8)(10) parent Major peak 15.24 1.1888 1.5332 180 182 [32] 209 [22]

50 Lidocaine(8) N-TMS- Major peak 13.69 1.0679 1.3773 86 220 [75] 73 [45]

51 Lidocaine(8) parent Major peak 15.28 1.1919 1.5372 86 58 [10] 91 [5]

52 LSD (MW-519, scanned only to 470) HFB- Pri.deriv. 24.61 1.9197 2.4759 417 221 [95] 418 [45]

53 MBDB N-TMS- Minor peak 14.30 1.1154 1.4386 144 73 [50] 135 [15]

54 MBDB N-HFB- Pri.deriv. 14.44 1.1264 1.4527 268 176 [75] 210 [50]

55 MDA N-HFB- Pri.deriv. 12.54 0.9782 1.2616 135 162 [55] 240 [12]

56 MDA N-HFB-N-TMS- OS artifact 12.88 1.0047 1.2958 312 73 [58] 135 [48]

57 MDEA(10) N-HFB- Pri.deriv. 14.19 1.1069 1.4276 268 162 [60] 240 [50]

58 MDEA-D6 (I$)(9) N-HFB- Pri.deriv. 14.13 1.1022 1.4215 274 165 [46] 244 [35]

59 MDMA(10) N-HFB- Pri.deriv. 13.81 1.0772 1.3893 254 162 [80] 135 [45]

60 Meperidine (Demerol® etc.) parent 13.97 1.0897 1.4054 247 246 [55] 218 [50]

61 Mephentermine N-HFB- Pri.deriv. 10.38 0.8097 1.0443 268 210 [95]

62 Mescaline N-HFB- Pri.deriv. 14.68 1.1451 1.4769 181 194 [45] 179 [30]

63 Mescaline N-HFB-N-TMS- OS artifact 15.26 1.1903 1.5352 181 73 [35]

64 Methamphetamine N-HFB- Pri.deriv. 9.94 0.7754 1.0000 254 210 [35] 118 [22]

65 Methamphetamine-D5 (I$)(9) N-HFB- Pri.deriv. 9.86 0.7691 0.9920 258 213 [30] 92 [20]

66 Methamphetamine-D9 (I$)(9) N-HFB- Pri.deriv. 9.84 0.7676 0.9899 261 213 [30] 123 [18]

67 Methamphetamine-D11 (I$)(9) N-HFB- Pri.deriv. 9.84 0.7676 0.9899 260 213 [25] 126 [20]

68 Methamphetamine-D14 (I$)(9) N-HFB- Pri.deriv. 9.86 0.7691 0.9920 261 213 [30] 128 [20]

69 Methaqualone parent 18.31 1.4282 1.8421 235 250 [30] 233 [28]

70 S-(-)-Methcathinone (“Cat”) N-HFB- Pri.deriv. 10.55 0.8229 1.0614 254 210 [35] 105 [100]

71 4-Methoxyamphetamine N-HFB- Pri.deriv. 11.40 0.8892 1.1469 121 148 [40] 240 [10]

72 4-Methoxyamphetamine N-HFB-N-TMS- OS artifact 11.87 0.9259 1.1942 312 121 [100] 73 [100]

73 cis-(±)-4-Methylaminorex (“U4Euh”) N,N’-bis-HFB- Minor peak 13.78 1.0749 1.3863 399 169 [70] 160 [75]

74 cis-(±)-4-Methylaminorex (“U4Euh”) N-HFB-N’-TMS- Pri.deriv. 16.78 1.3089 1.6881 275 160 [60] 117 [30]

75 (-)-N-Methylephedrine(11) O-TMS- Pri.deriv. 9.66 0.7535 0.9718 72 73 [13] 163 [5]

76 (+)-N-Methylephedrine(11) O-TMS- Pri.deriv. 9.71 0.7574 0.9769 72 73 [13] 163 [5]

77 N-Methylphenethylamine (I$) (9) N-HFB- Pri.deriv. 9.54 0.7441 0.9598 240 104 [100] 169 [40]

78 Methylphenidate (Ritalin®) N-HFB- Pri.deriv. 15.38 1.1997 1.5473 280 281 [10]

Table 10 continued
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 23 of 32

TABLE 10 (continued). Gas chromatographic retention times for heptafluorobutyryl and trimethylsilyl 

derivatives of selected drugs of abuse, precursors, and potential adulterants(1)

GC 

Peak 

No. Compound Derivative Form(2) Notes(3)

Retention 

Time 

Minutes(4)

Relative Retention 

Time Ions (Significant m/z)

(7)

(5) (6) 1’ 2’ (7) 3’ (7)

79 N-Methyl pseudoephedrine(12) O-TMS- Pri.deriv. 9.66 0.7535 0.9718 72 73 [13] 163 [5]

80 Morphine O-HFB-O’-TMS- Minor peak 19.97 1.5577 2.0091 340 324 [28] 341 [25]

81 Morphine O,O’-bis-TMS- Pri.deriv. 21.08 1.6443 2.1207 429 414 [50] 401 [35]

82 Nicotine parent 8.86 0.6911 0.8913 84 133 [35] 162 [18]

83 Norpseudoephedrine (Cathine) N-HFB-O-TMS- Pri.deriv. 10.39 0.8105 1.0453 179 180 [18] 240 [18]

84 Norpseudoephedrine (Cathine) N-HFB-N,O-bis-TMS- OS artifact 11.26 0.8783 1.1328 179 180 [18] 312 [10]

85 Oxycodone (OxyContin®) TMS- Pri.deriv. 21.66 1.6895 2.1791 387 388 [30] 372 [30]

86 Phencyclidine (PCP) parent Major peak 15.62 1.2184 1.5714 200 242 [35] 243 [25]

87 Phencyclidine (PCP) N-HFB-dehydro- Artifact 19.85 1.5484 1.9970 91 159 [60] 280 [10]

88 Phencyclidine-D5 (I$)(9) parent Major peak 15.59 1.2161 1.5684 205 96 [42] 246 [25]

89 Phencyclidine-D5 (I$)(9) N-HFB-dehydro- Artifact 19.83 1.5468 1.9950 96 164 [65] 280 [10]

90 Phenethylamine(8) N-HFB- Pri.deriv. 8.58 0.6693 0.8632 104 91 [60] 169 [15]

91 Phenethylamine(8) N-HFB-N-TMS- Pri.deriv. 9.51 0.7418 0.9567 298 105 [40] 220 [10]

92 Phentermine(8) N-HFB- Pri.deriv. 8.72 0.6802 0.8773 254 132 [12] 214 [8]

93 4-Phenyl-1-butylamine (I$)(9) N-HFB- Pri.deriv. 11.47 0.8947 1.1539 91 104 [25] 176 [22]

94 Phenylephrine(8) N-HFB-O,O’-bis-TMS- Pri.deriv. 13.94 1.0874 1.4024 267 268 [25] 240 [12]

95 Phenylpropanolamine N-HFB-O-TMS- Pri.deriv. 10.49 0.8183 1.0553 179 180 [18] 240 [18]

96 Phenylpropanolamine N-HFB-N,O-bis-TMS- OS artifact 11.01 0.8588 1.1076 179 180 [18] 312 [10]

97 N-Propyl amphetamine (I$)(9) N-HFB- Pri.deriv. 11.05 0.8619 1.1117 282 240 [85] 118 [20]

98 Pseudoephedrine N-HFB-O-TMS- Pri.deriv. 11.68 0.9111 1.1751 179 254 [15] 73 [75]

99 Theophylline (8) parent Major peak 15.50 1.2090 1.5594 237 252 [57] 223 [14]

100 Trifluoromethylphenyl piperazine (10) N-HFB- Pri.deriv. 13.76 1.0733 1.3843 200 229 [70] 172 [73]

(1) Actual retention times may vary depending on individual GC column and GC conditions. Gas chromatographic and mass spectrometer conditions used as on p. 9109-1. 

(2) Derivative form. HFB = heptafluorobutyryl derivative. TMS = trimethylsilyl derivative. N- = attachment to nitrogen atom. O- = attachment to oxygen atom. Not all forms are presented. 

Trifluoroacetyl derivatives are not presented. Underivatized compounds are identified as a “parent” compound. Parent compounds that have poor chromatographic peak shapes under the 

conditions used are not presented. Spectra for the derivatives are given in the Backup Data Report (Appendix-II) [4].

(3) Major and minor peaks are identified where two or more forms are possible. In some cases two major peaks may exist. Pri.deriv. = Primary derivative, a major peak. The major peak 

or the primary derivative should be used for quantitation. OS artifact = Oversilylation artifact [14]. Oversilylation artifacts occur where a primary amine is substituted with both a 

heptafluorobutyryl and a trimethylsilyl group. Under the specified conditions of extraction and derivatization these remain as minor components and are of little concern. 

(4) Retention times are not the same as in Table 10 or Figures 1 and 2 in this method since these data were obtained on a different instrument. Relative retention times should be 

approximately the same. 

(5) Retention time relative to 4,4’-dibromooctafluorobiphenyl.

(6) Retention time relative to the heptafluorobutyryl derivative of methamphetamine.

(7) Significant ions that can be used for quantification and qualitative identification are given. The base peaks are not necessarily included, especially if they are low mass (<100 AMU). 

Numbers in brackets indicate the approximate relative abundance of the secondary (2’) and tertiary (3’) ions relative to the primary (1’) ion and not necessarily to the base peak of each 

mass spectrum. Relative abundance varies with different tuning criteria and cleanliness of the mass spectrometer source. The 1’ or 2’ ions are recommended for quantification. All ions are 

selected as much as possible above m/z 100 to avoid interference from low mass co-eluting interferences. The 2’ and 3’ ions are selected as much as possible for nearness to the primary 

ion to minimize false negatives from skewing of spectra as the mass spectrometer source becomes contaminated with use. Ubiquitous ions (e.g., m/z 73, 91, and 169) are avoided as 

much as possible.

(8) Intentional or unintentional adulterants. For example, phentermine may be added to MDMA and caffeine added to methamphetamine intentionally. Chlorpheniramine is an 

unintentional adulterant when pseudoephedrine containing chlorpheniramine is used as a methamphetamine precursor.

(9) (I$) = Internal standard. The best results are obtained using internal standards that are deuterated analogs of the target analyte, or those that are chemically and structurally similar to 

the target analytes.

(10) Typical “club drugs” (piperazine analogs as ecstasy substitutes, ketamine and flunitrazepan as predatory drugs).

(11) 4-Bromo-2,5-DMPEA = 4-Bromo-2,5-dimethoxyphenethylamine (Nexus).

(12) Presence of (+)-norephedrine, N-methylpseudoephedrine and/or N-methylephedrine in pseudoephedrine or ephedrine indicates extracts of Ephedra spp. as source of methamphetamine 

precursor. Presence of amphetamine and N,N-dimethylamphetamine in methamphetamine final product also suggests the same source [22, 23, 24].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 24 of 32

Figure 1. Typical chromatogram of mixed heptafluorobutyryl and trimethylsilyl derivatives 

by GC-MS in scan mode (time in minutes) 

GC Peak Identification: See Table 10 for identification of numbered GC peaks. (But note that 

retention times in Table 10 do not correspond to those in Figure 1 because a different U.S. 

Pharmacopeia (USP) G27 column and instrument was used.)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 25 of 32

Figure 2. Typical extracted ion chromatograms (EIC) of mixed heptafluorobutyryl and trimethylsilyl 

derivatives by GC-MS in scan mode (time in minutes)

GC Peak Identification: See Table 10 for identification of numbered GC peaks. (But note that retention times in

Table 10 do not correspond to those in Figure 1 because a different U.S. Pharmacopeia (USP) G27 column and 

instrument was used.) 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 26 of 32

APPENDIX:

A. REAGENTS and SOLUTIONS: 

1. 4,4’-Dibromooctafluorobiphenyl is optional. It is useful for monitoring instrument tuning and

autosampler performance.

2. Primary amines form Schiff bases and enamines with ketones and aldehydes. These may in turn

form derivatives with the acylating reagents. The use of acetone must strictly be avoided prior to 

the analytes being derivatized. Glassware and equipment rinsed with acetone must be thoroughly 

dried. Toluene should be avoided for making up standard solutions because it usually contains 

benzaldehyde, an oxidation product of toluene. Condensation products have been observed 

between primary amines and benzaldehyde. The only solvents recommended for the preparation of 

stock solutions and dilutions thereof are methanol (preferably) and isopropanol.

B. EQUIPMENT:

1. Wipe media: Besides cotton gauze, 4”X4” (10 cm x 10 cm) 4-ply MIRASORB® (Johnson and Johnson),

and 4”X4” (10 cm x 10 cm) AlphaWipe® (TX1004, Texwipe Corp) were acceptable wipe media and 

can be used in the absence of cotton gauze. MIRASORB®, a non-woven cotton/polyester blend, is 

discontinued but counterparts exist that claim to be of identical construction and fiber composition. 

AlphaWipe® is a hydrophilic, highly adsorbent, tightly knitted continuous filament polyester wipe. 

Precision and accuracy data for MIRASORB® are given in the Backup Data Report [4].

2. Shipping containers: The 50-mL polypropylene centrifuge tubes with caps are preferred for one or

two gauze wipes and are not as breakable as the 40-mL VOA vials. The 40-mL VOA vials are acceptable 

for single gauze wipes. Larger containers (glass with a PTFE lined cap) should be used for combining 

more than two gauze wipes into a single sample. The size of the container for two or more wipes 

should be approximately 25 mL per gauze wipe (e.g., a minimum size of 100-mL for up to four gauze 

wipe samples). There needs to be enough extra headspace in the shipping container to allow the 

desorption solution to cover the gauze wipes and to percolate freely through the wipe sample(s) 

during mixing.

3. Each regulatory agency having legal jurisdiction over the contaminated site may require different

but specific off-site preparation and on-site sampling procedures. It is important to consult local 

regulatory agencies or departments of health having legal jurisdiction over contaminated sites to 

determine specific sampling, quality control, analyses, and reporting requirements.

C. SAMPLING:

1. Follow specific requirements of surface area to be wiped (usually 100 cm2or 1000 cm2) and action

threshold (or maximum allowable residual level) set by the state or specified by the client. Uptake 

rates depend upon the wipe sampling method used, so the specific wipe technique used must be 

specified, and any deviations from the required wipe sampling requirements noted.

Note: To ensure that samples have not been tampered with, the use of custody seals and a chain-ofcustody form is strongly recommended.

2. Prepare a rigid template from disposable cardstock or a sheet of PTFE having either a 10 cm × 10 cm

or 32 cm × 32 cm square-cut hole. The template must be able to retain its shape during wiping to 

ensure that the areas wiped were either 100 cm2 or 1000 cm2. Secure the template(s) to the area(s) 

to be wiped (e.g., with tape along outside edge of template). If a single-use disposable template 

is not used, clean the template between samples to avoid cross-contamination, and provide the 

laboratory with a blank wipe of the cleaned template between samples to determine that no crosscontamination has occurred. 

3. A template might not always be applicable, as in curved or odd-shaped areas such as around burners

on stove tops or a fan blade. In such cases sample an area as close to either 100 cm2or 1000 cm2 as 

feasible and provide the measurement to the regulatory agency and to the analytical laboratory for 

proper reporting. Tape can be used to delineate the sampling area.

4. It is recommended to provide extra wipe media from the same lot for required media blanks, fieldequipment blanks, samples, and quality-control samples.
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METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 27 of 32

5. Gauze in sterile packaging is recommended to minimize the chance for cross-contamination, which

can more easily occur with open bulk packaged cotton gauze.

6. To prevent contamination in the field, another alternative is to pre-wet and insert the gauze

wipes into the sample containers off-site. This avoids any possibility of the bottle of methanol or

isopropanol becoming contaminated on-site with methamphetamine (or other analytes). If the wipes

were prepared off-site, then remove pre-wetted gauze wipe from sample container, opening only one

sample container at a time. In either case, squeeze out and discard any excess solvent from the gauze

wipe. Use fresh latex or nitrile gloves for each separate sample and blank. Do not use vinyl gloves due

to the potential for leaching of phthalate plasticizers and contamination of the samples.

7. Wipe techniques

a. Concentric Squares Wiping Technique (particularly suitable for smooth and non-porous surfaces

and described by OSHA [25]): Fold the pre-wetted gauze in half and then fold in half again. Using

firm pressure wipe the area within the template. Start at one of the inside corners of the template

and wipe in concentric squares, progressing toward the center. End with a scooping motion.

Without allowing the gauze to touch any other surface, reverse the last fold so that the exposed

side of the gauze is facing inward and using a fresh surface of the gauze, wipe the same area in the

same manner as before. Roll or fold the gauze again and insert into the shipping container.

b. Side-to-side Wiping (or Blotting) Technique (particularly suitable for rough, porous, and/or soiled

surfaces): Fold the pre-wetted gauze in half and then fold in half again. Using firm pressure wipe or

blot the area within the template with at least five overlapping side-to-side horizontal passes (see

NOTE) beginning at the top and progressing to the bottom in a “Z” pattern. End with a scooping

motion. If blotting, blot at least five times on each horizontal pass (see NOTE). Without allowing

the gauze to touch any other surface, reverse the last fold so that the exposed side of the gauze

is facing inward. Using a fresh surface of the gauze, wipe or blot the area again with at least five

overlapping top-to-bottom vertical passes beginning at the left side and progressing to the right

in an “N” pattern. If blotting, blot at least five times on each vertical pass. Roll or fold the gauze

again and insert into the shipping container. Blotting is suggested in areas so soiled or rough that

the threads of the gauze media are continually snagged.

NOTE: On areas larger than 100 cm2, more than five passes and blots will be needed.

c. Repeat or Serial Wiping: If isopropanol is used for wiping, a serial or repeat wipe sample of the

same area with a fresh gauze wipe will improve sampling efficiency. (See recoveries for second

wipe in Tables 9a and 9b.) For serial wiping, repeat the wiping procedure described above

(APPENDIX 6a or 6b) with a fresh gauze wipe. Place the second gauze wipe into the same shipping

container as the first gauze. The 50-mL polypropylene centrifuge tubes are large enough to

contain up to two gauze wipes.

NOTE: If the area to be wiped remains substantially wet from the first gauze, the second gauze

wipe might be used in the dry state to soak up the residual solvent from the first gauze 

wipe.

8. Composite sampling: Composite samples are allowed by some regulatory agencies. Their use for

quantitative purposes may be subject to the permission and guidance of regulatory agencies. Refer 

to guidelines of regulatory agency for directions on composite sampling. A basic default guideline for 

composite sampling is as follows: Do not mix inconsistent samples, that is, areas wiped must be equal 

in area, sampled areas must have the same high or low probability of contamination, and sampled 

areas must relate to a specific target appliance or site and not to several appliances or incongruous 

sites combined. 

NOTE: Composite samples cannot meet specific action-threshold requirements for discrete sampling 

locations. Nor do composite samples consisting of four wipes, for example, improve the 

sensitivity by decreasing the LOD four fold; instead it raises the LOD by a factor related to 

the extra volume of desorption solution that is required to desorb a larger number of wipes. 

The following example illustrates these two points. Assume that the action level was 0.1 

μg/100 cm2. If the analysis gave an LOD of 0.06 μg/sample for a single wipe or discrete sample 

covering an area of 100 cm2, then the LOD for the analysis could be expressed as 0.06 μg/100 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 28 of 32

cm2, which is low enough to be able to determine whether any discrete sample is at or exceeds 

the action level. Now if a composite of four wipes was taken, each with an area of 100 cm2 

for a total area wiped of 400 cm2, the LOD for that composite sample is not 0.06 μg/400 cm2 

nor is it 0.015 μg/100 cm2; it is actually several times larger than 0.06 μg/400 cm2. First of all it 

increases relative to the ratio of the volume of desorption solution used to desorb the sample 

compared to that used for the calibration standards. Secondly it has nothing to do with the 

AREA that was wiped, because the LOD for the calibration curve is determined in terms of μg 

per sample, independent of the area. To explain the first point, assume approximately 90 mL 

was used (for ease in calculation) to desorb the four wipes and 30 mL (the normal amount for 

a single wipe) was used to desorb each calibration standard. The calculation of the LOD for the 

four composited samples would be μg/sample x (desorption volume for 4 wipes) ∕ (desorption 

volume for the calibration standards), or 0.06 μg/sample x (90 mL/30 mL), or 0.18 μg/sample for 

the composited sample. Since the area wiped for the composite sample was 400 cm2, the LOD 

for that sample could be expressed as 0.18 μg/400 cm2. Regarding the second point, this value, 

0.18 μg/400 cm2, cannot be construed or mathematically reduced to 0.045 μg/100 cm2 because 

it cannot be known whether three of the four wipes were blank and the fourth wipe just under 

the value of 0.18 μg. Hence, the effective LOD per individual wipe has to be regarded not only 

as 0.18 μg/400 cm2 but also as 0.18 μg/100 cm2 because any value determined for entire 400 

cm2 might have come from just one of those 100 cm2 areas. Thus, for composite samples, the 

LOD must be expressed in terms of the entire area wiped and not extrapolated to some portion 

thereof. In this example, an LOD of 0.18 μg/100 cm2 is above the action threshold of 0.1 μg/100 

cm2, meaning that this composite sample cannot satisfy the requirement that residual levels be 

below 0.1 μg/100 cm2. It remains for the regulatory agency and not the laboratory to determine 

how to apply results for composite samples to the established action levels. The same 

consideration that is given above for the LOD applies to results that are greater than the LOD. 

To avoid confusion in reporting concentrations for composite samples, it is recommended that 

the sample concentration (in μg/sample, whatever the sample size) and the total area wiped (in 

cm2) be reported separately. For example, a result of 0.4 μg/sample for a sample consisting of 

four separate wipes of 100 cm2 each (for a total area wiped of 400 cm2), is to be reported as 0.4 

μg/400 cm2 and not averaged to 0.1 μg/100 cm2. This manner of reporting may be required by 

some regulatory agencies.

9. For quality assurance purposes, regulatory agencies may require duplicate samples to be taken in

the field. If such is the case, an area contiguous with and adjacent to the first area, if possible, should 

be wiped as described under SAMPLING. Do not re-wipe the previously wiped area. This sample 

is a blind sample and should not be identifiable by the analytical laboratory as a duplicate of any 

other sample. These are distinct from the laboratory duplicates of a single sample described in step 

14 of the method. Field duplicates are useful for evaluating the consistency of sampling technique, 

assuming uniformity of contamination on adjacent sampling sites. Laboratory duplicates are useful for 

evaluating consistency of sample preparation and instrumental analysis.

D. DESORPTION FROM MEDIA:

1. An internal standard spiking solution volume of 60 μL was selected for ease in scaling from 60 μL

per 30 mL to 80 μL per 40 mL of desorption solution. In either case the rate of 2 μL internal standard 

spiking solution per mL desorption solution was used. However, any convenient volume of internal 

standard spiking solution (e.g., 50 μL) that can be delivered reproducibly is acceptable. Whatever 

volume is chosen, there must be no variation in the volume of the internal standard spiking solution 

used in preparing each of the calibration standards. If spiking strategy A is used (see step D3 of 

APPENDIX), it is critical to know the exact volume of internal standard spiking solution that is applied 

to each sample (V1

), the media blanks (V5), and the calibration standards (V2), since these volumes are 

used for internal standard spiking solution volume corrections in step 19.

2. It is not necessary to know the exact volume of desorption solution added to each sample or the

volume of residual wetting alcohol because differences in the volumes are normalized through the use 

of internal standards added prior to desorption.
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3. Alternate strategy for spiking internal standards (spiking strategy B below): By using the exact same

volume of internal standard spiking solution in all samples, blanks, QC samples, and calibration

standards, regardless of the volume of desorption solution added or residual wetting alcohol, the

volume corrections in step 19 (V1

/V2 and V5/V2) drop out of the equation. However, the internal

standard GC peak areas must still be measurable in samples where larger volumes of desorption

solution are used (such as for composite samples). Because of the increased dilution of the internal

standard in larger samples, this approach should be limited to desorption solution volumes of about

120 mL or less.

NOTE: There are two separate strategies for handling larger samples requiring larger volumes of

desorption solvent. These are outlined below as strategies A and B.

Number of 

Wipes

Size of 

Shipping 

Container 

(mL)

Volume of Internal Standard 

Spiking Solution (μL)

Strategy A Strategy B

Volume of Desorption 

Solution (mL) 

 (Strategies A and B)

1 40 to 50 60 60 30

2 50 80 60 40

4 (e.g., 100 to 120 160 60 80

composite)

Apply volume Do not apply 

correction volume correction 

factors at step factors at step 19.

19.

With either strategy, if two gauze wipes were included in the samples, then use 40 mL of 

desorption solution. If four gauze wipes were included in the samples, then use 80 mL of 

desorption solution.

a. In strategy A, the volume of internal standard spiking solution is kept at a constant ratio of 2 μL per

mL of desorption solution added. This enables larger samples to be desorbed without diminishing

the area of the GC peak for the internal standard. However, a volume correction factor (V1

/V2 ) is

needed in the final calculations in step 19. Therefore, the exact volume of internal standard added

to each of the samples relative to that added to the calibration standards must be known.

b. In strategy B, the volume of internal standard spiking solution is kept constant for all samples and

calibration standards, but need not be exactly 60 μL. This enables the final calculations to be made

in step 19 without a volume correction factor. However, the area of the GC peak for the internal

standard will vary with sample desorption volume and the internal standard must be concentrated

enough to be measurable where larger volumes of desorption solution are used.

E. SOLID PHASE EXTRACTION PROCEDURE:

 The following SPE columns were investigated and found to work adequately for this method: BOND 

ELUT-CERTIFY® (200 mg/3mL); Clean Screen® (300 mg/3 mL) ; Oasis® MCX (60 mg/3 cc); or Speedisk® 

H2O-Philic SC-DVB. Two columns (Clean Screen® and BOND ELUT-CERTIFY®) are based upon a silica 

support. The other two (Oasis® and Speedisk®) are based upon an organic polymer support. The 

precision and accuracy data in Tables 7a and 7b were generated using the Waters Oasis® MCX 3cc/60 

mg column.

F. DERIVATIZATION:

There are unique advantages and disadvantages in using the mixed MSTFA + MBHFBA reagent. The 

disadvantages with some possible remedies are listed as follows.

1. A few percent of trifluoroacetyl derivatives of secondary amines are formed (presumably from MSTFA)

in competition to the intended heptafluorobutyryl derivatives.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 30 of 32

a. Remedy #1: This artifact is eliminated by replacing MSTFA with MSHFBA (N-methyl-N-trimethylsilyl

heptafluorobutyramide, Alltech Associates, Deerfield, IL). However, precision and accuracy were not

evaluated for NIOSH 9109 using MSHFBA instead of MSTFA.

b. Remedy #2: If ephedrine compounds or compounds containing free hydroxyl groups are not to be

analyzed, MSTFA might be omitted and MBHFBA used alone.

2. Use of the mixed reagent often results in over-silylation, the production of unintended silylation

artifacts [14], particularly of amides. The primary over-silylation artifact with primary amines is the Ntrimethylsilyl derivative of the N-acyl derivative. The GC peak area for this artifact can be significant;

under certain circumstances it is nearly equal to that of the intended N-acyl derivative.

a. Remedy #1: The presence of ammonium chloride from the SPE eluates seems to prevent or greatly

reduce over-silylation of amides. These artifacts can be ignored when using the SPE columns with

the 80:20:2 methylene chloride:isopropanol:ammonium hydroxide eluent.

b. Remedy #2: If ephedrine compounds or compounds containing free hydroxyl groups are not to be

analyzed, silylating reagents (MSTFA or its alternate, MSHFBA) might be omitted and MBHFBA used

alone.

3. The mass spectrometer may need more frequent cleaning to maintain sensitivity. This is offset by the

shorter sample preparation time, especially for large numbers of samples.

4. When the fused silica capillary columns become exposed to the mixed silanization-acylation reagents,

the column may become unsuitable for other types of samples.

5. The chromatograms are cluttered with silylation by-products making it difficult to detect low levels of

unknown (non-target) compounds if a drug screen for unknown compounds is an objective. For this

objective, the liquid-liquid extraction procedure of NIOSH 9106 [6] provides cleaner chromatograms

with less interference from reagent by-products.

6. The advantages of the mixed MSTFA+ MBHFBA reagent, when used with SPE, are as follows:

a. Faster preparation time (no heating in an oven, no cool-down time, no evaporation or neutralization

of the reagents, and no reconstitution with solvent thereafter).

b. No heat or acid induced isomerization or dehydroxylation of the ephedrine or other hydroxyl

containing compounds (e.g., ephedrine, norephedrine, pseudoephedrine, phenylephrine, etc.).

c. The method can be extended to easily hydrolyzed phenolic and polyhydroxy compounds of

aryl-alkyl-amines (e.g., Albuterol, epinephrine and metabolites [15], metabolites of MDMA, and

phenylephrine) because of the thermal stability of the trimethylsilyl ether groups on phenols and

trimethylsilyl ester groups.

d. Hindered amines such as MDEA are derivatized more completely but still require an internal

standard with structural similarity.

G. MEASUREMENT:

Recoveries for the laboratory control matrix spike samples (QC and QD) must meet the guidelines 

of the specific regulatory agency involved (80-120% is a reasonable target in the absence of specific 

guidance). 

NOTE 1: The QC samples (QC and QD) in this method may be referred to in some guidance documents 

as matrix spike and matrix spike duplicate samples (MS/MSD) but serve the same purpose. 

Analyze and report field-equipment blanks as samples. Do not subtract their values from any 

other sample. 

Recoveries of CCV standards must meet guidelines of regulatory agency (80-120% is a reasonable 

target in the absence of specific guidance). The CCV standards may be referred to in some guidance 

documents as ‘QC samples’ but such QCs are equivalent to liquid standards (not matrix spiked samples) 

and serve the same purpose of the CCVs in this method.

NOTE 2: With the GC/MS it is possible to achieve the lower limit of 0.05 μg or less per sample for 

methamphetamine in either the scan mode or SIM mode. The scan mode is essential where 

the identification of unknowns is an analytical objective. If lower limits of detection are 

desired or difficult to obtain in the scan mode, or for routine target compound only analyses, 

the instrument may be operated in the SIM mode.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 31 of 32

H. MAKING DILUTIONS:

If the samples exceed the upper calibration range for the analysis, one of the following procedures 

may be used to estimate the high level concentrations.

1. Dilution procedure A (dilution of the derivatization mixture within a GC vial): Transfer an aliquot of 

the derivatization sample mixture from the GC vial to a clean low-volume GC vial and add acetonitrile, 

MSTFA, and MBHFBA. For example, for a 10:1 dilution transfer 20 μL of sample to a clean vial and 

add 120 μL of acetonitrile and 30 μL each of MSTFA and MBHFBA, for a total volume of 200 μL. For a 

4:1 dilution, transfer 50 μL of sample to a clean vial and add 100 μL of acetonitrile and 25 μL each of 

MSTFA and MBHFBA, for a total volume of 200 μL. Cap the GC vial, mix by inversion a few times, and 

analyze diluted sample. Do not include the dilution factor in step 19 since the internal standard will be 

diluted along with the target analyte. 

NOTE: For dilutions greater than 10, the internal standard may become too diluted to quantify. In such 

a case, use the following procedure B.

2. Dilution procedure B (dilution of the original sample desorbate): In this procedure, an aliquot of the 

original sample desorbate is diluted with a simulated blank solution and then transferred to a SPE 

column in step 8d. For example, for a 10:1 dilution, dilute 0.5 mL of sample desorbate solution from 

step 7f in a clean test tube containing 4.5 mL of a simulated blank solution, mix, and then transfer the 

entire contents to a pre-conditioned SPE column. For a 50:1 dilution, dilute 0.1 mL of sample desorbate 

solution from step 7f in a clean test tube containing 4.9 mL of a simulated blank solution, mix, and 

then transfer the entire contents to a pre-conditioned SPE column. Proceed thereafter to step 8d as 

normal. The simulated sample blank should be prepared identically to the sample needing dilution, 

using the same volumes of internal standard spiking solution and desorption solution that were used 

with the sample in the original desorption. For example, if the original sample was desorbed with 

40 mL desorption solution with 80 μL of added internal standard spiking solution, then prepare the 

simulated blank in the same way. The volume of wetting alcohol is estimated (e.g., about 3 mL per 

3”x3” 12-ply cotton gauze wipe). Include a dilution factor (V3

/V4) in the calculations in step 19 

(e.g., V3

/V4 = 5 mL divided by the volume in mL of original desorbate diluted to 5 mL with solution 

from the simulated blank). The dilution factor in the above examples are 5 mL/0.5 mL or 10 for a 10:1 

dilution and 5 mL/0.1 mL or 50 for a 50:1 dilution. Correct for differences in internal standard spiking 

solution volumes in step 19 (if applicable) using for V1

 the volume of internal standard spiking solution 

which was added to the original undiluted sample.

Caution: This dilution procedure gives quantitative results only if the residual volume of methanol (or 

isopropanol) used for wetting the sample wipes was exactly the same as the volume used in 

preparing the calibration standards (normally about 3 mL, see Table 6). Deviations of a few 

milliliters in residual wetting alcohol will not affect the results for undiluted samples but will 

amount to an error of a few percent in the final results of samples that are diluted.

The potential error due to differences in residual wetting solvent can be estimated for 

specific volumes of desorption solution and wetting alcohol. Assume the sample wipes and 

calibration standards are both desorbed in 30 mL of desorption solution and 3 mL of alcohol 

is added to the calibration standards. The potential error in volume (and final results) in the 

samples is approximately ±3.03% (inversely proportional) per mL difference in the residual 

alcohol in the samples (i.e. ±1 mL difference in 33 mL). For 40 mL of desorption solution 

and 4 mL of alcohol added to the calibration standards, the error is ±2.27% for every mL 

difference (i.e. ±1 mL difference in 44 mL). However, since the volume of residual wetting 

alcohol is not known and cannot be determined once the sample wipe has been desorbed, 

the actual error cannot be determined.

 However, the maximum possible error can be calculated. Since the maximum amount 

of alcohol that a 3”x3” 12-ply (or 4”x4” 8-ply) cotton gauze can hold is about 6 mL when 

saturated (dripping wet), there can only be a deviation of plus or minus 3 mL from the 3 

mL alcohol added to the calibration standards. Therefore, the maximum error in a result 

due to differences in the volume of residual alcohol in a cotton gauze sample compared to 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE . . . on Wipes by SPE: METHOD 9109, Issue 1, dated 17 October 2011 - Page 32 of 32

the standards can only be three times the error for a 1 mL difference in volume. Since the 

error for ±1 mL is ±3.03%, the maximum error for ±3 mL is three times larger, or ±9.1%. In 

practice, the error will be less than this because it is unlikely that the gauze samples will be 

completely dry or completely saturated after squeezing out the excess alcohol and wiping 

a surface. The practical amount of alcohol that remains in the 3”x3” 12-ply (or 4”x4” 8-ply) 

cotton gauze wipes when the excess is squeezed out is between 1 and 2 mL. This translates 

into an error that is between +3% and +6% in the final results for diluted samples. Undiluted 

samples will not be affected. This error is within the overall accuracy for the method for 

methamphetamine.

3. Dilution procedure C (dilution of desorbates from dried samples): Dilution errors for over-range 

samples may be corrected by knowing the exact amount of residual alcohol in the samples. The 

volume (or weight) of residual solvent in each gauze wipe might be determined by the difference 

between a wet weight and dry weight. Better yet, the error might be eliminated for diluted samples 

by adding, after the samples are dried (without taking any weight), the same known volume of 

wetting alcohol that is added to the calibration standards (i.e. 3 mL). Thereafter, if any samples need 

dilution, there will be no dilution errors due to differences in residual alcohol, because all samples and 

standards will have the same volume of alcohol and total volume of desorption solution.

However, air drying of the samples is not recommended because of the possible loss of 

methamphetamine due to its volatility when it is not in the salt form, which form cannot be 

assured in field samples. Also, manipulating the samples for weighing and drying might introduce 

contamination. Drying is not recommended as a procedure for analytes having a vapor pressure high 

enough to be lost in the process, or that tend to form azeotropes with alcohols, especially when the 

critical action levels for remedial cleanup are at the lower end of the method calibration range. Drying 

is not an option if the samples have already been desorbed.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

BERYLLIUM in Surface Wipes

by Fluorometry 9110

Be MW: 9.0121 CAS: 7440-41-7 RTECS: DS1750000 

METHOD: 9110, Issue 2 EVALUATION: FULL Issue 1: 6 April 2007

Issue 2: 12 December 2015

OSHA: none for surfaces

MSHA: none for surfaces

DOE: 3 μg per 100 cm2 (housekeeping), 0.2 μg per 

 100 cm2 (equipment release) [1]

OTHER OELs: [2]

PROPERTIES: solid, d 1.85 g/mL, MP 1,278 °C, VP 0 kPa

 (0 mm Hg) @ 25 °C 

SYNONYMS: beryllium metal, beryllia (BeO)

SAMPLING

SAMPLER: WIPE (cellulosic or polyvinyl alcohol)

WIPE AREA: 100 cm2 minimum

SHIPMENT: routine

SAMPLE

STABILITY: stable

BLANKS: 3 field blanks min.

ACCURACY

RANGE 

STUDIED: 0.0001 to 6 μg per wipe [3,4]

BIAS: negligible [3,4]

OVERALL

PRECISION (SS�rrrr): 0.094

ACCURACY: 18.9%

MEASUREMENT

TECHNIQUE: UV/VIS FLUOROMETRY

ANALYTE: complex of hydroxybenzoquinoline 

sulfonate (HBQS) with beryllium

DISSOLUTION: ammonium bifluoride (aqueous), 10 g/L

DETECTION 

SOLUTION: contains 63.4 μmol/L HBQS, 2.5 mmol/L 

EDTA, and 50.8 mmol/L lysine 

monohydrochloride (optional); pH 

adjusted to 12.85 with 10 mol/L NaOH,

as necessary

DETECTOR: excitation, 360 nm to 390 nm; emission, 

integrated between 470 and 480 nm 

(λmax ≈ 475 nm)

CALIBRATION: beryllium standard solutions

RANGE: (0.005 to 6) μg per wipe [3,4]

ESTIMATED LOD: 0.0001 μg per wipe

PRECISION (SS�rr): 0.021 at ≈0.2 μg per wipe, 0.076 at ≈1.5 

μg per wipe, 0.052 at ≈3 μg per wipe

APPLICABILITY: The working range of the method is 0.0005 μg to 6 μg for surface wipe samples. The analysis is for total beryllium 

and is not compound specific.

INTERFERENCES: Minor interference from iron can result if iron concentrations are high. Samples high in iron demonstrate a 

yellow or gold coloration. This interference can be minimized by allowing the solution to sit for at least two hours, during which 

time the solution clears, and then filtering the sample extract before use. An alternative method is to filter the solution (after 30 

minutes of standing) through a hydrophilic filter of pore size of 0.2 μm or smaller.

OTHER METHODS: Method 7300 (hot plate digestion and inductively coupled plasma atomic emission spectrometry) is an 

alternative procedure for the determination of elemental beryllium [5], but with higher detection limits. ASTM method D7202 is a 

similar procedure to detect elemental beryllium by fluorescence [6].
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 2 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

REAGENTS: 

1. Ammonium bifluoride.*

2. Ethylenediaminetetraacetic acid (EDTA), 

disodium salt, dihydrate. 

3. 10-Hydroxybenzo[h]quinoline-7-sulfonate 

(HBQS) [5]. 

4. L-Lysine monohydrochloride 

5. Sodium hydroxide.* 

6. Water, deionized.

7. Dissolution solution:* aqueous ammonium 

bifluoride, 10 g/L (prepared by dissolving 

ammonium bifluoride in deionized water)

8. Detection solution:* 63.4 μmol/L HBQS, 2.5 

mmol/L EDTA, and 50.8 mmol/L lysine 

monohydrochloride; pH adjusted to 12.85 

with 10 mol/L NaOH). An alternative 

preparation of dye solution without lysine 

(lysine-free) may be made by adding 1.104 g 

of EDTA and 64 μmoles of the 10-HBQS dye 

in 900 ml of water. After a clear solution is 

obtained, 114.5 ml of 2.5 N NaOH is added 

and mixed to obtain the final dye solution. 

The pH of the dye solution is 13.2. The lysinefree dye solution (commercially available) 

may be used for all analytical purposes and 

also provides superior detection limits.

9. Beryllium standard solution,* 1,000 mg/L 

(commercially available).

10. Beryllium-spiked media* (commercially 

available).

*See SPECIAL PRECAUTIONS.

EQUIPMENT:

1. Sampler: wipe, cellulosic, 47 mm diameter 

minimum.

NOTE: Polyvinylalcohol (PVA) media are also 

suitable for this method [7]

2. Template, disposable/reusable, 100 cm2

minimum area.

3. Tape, masking

4. Ultraviolet/visible (UV/Vis) fluorometer, with 

excitation lamp (λ = 380 nm) and timeintegrating visible detector (400 nm to 700 

nm, λmax ≈ 475 nm) or optical filters for 

appropriate wavelengths (excitation of 360 

nm to 390 nm; emission of ≈475 nm, with full 

width at half maximum of ±5 nm).

5. Mechanical agitator, shaker, or rotator.

6. Hot block (for beryllium oxide extraction).

7. Fluorescence cuvettes, disposable, 10 mm 

diameter, transparent to UV/Vis radiation.

8. Centrifuge tubes, plastic, 15 mL

9. Syringe filters, hydrophilic polypropylene, 0.2 

μm pore size, 25-mm diameter, in plastic 

housings. 

NOTE: Polytetrafluoroethylene (PTFE) filters 

are unsuitable for this method

10. Pipettors, mechanical, of assorted sizes. Pipet 

tips, plastic, disposable, of assorted sizes.

11. Labware, plastic (e.g., beakers, flasks, 

graduated cylinders), of assorted sizes.

12. Tweezers, plastic or plastic-coated.

13. Laboratory wipes. 

14. Personal protective wear (e.g., respirators, 

gloves, lab coats, safety eyewear), as needed

SPECIAL PRECAUTIONS: Wear appropriate personal protection during sampling activities and analysis. It 

is essential that suitable gloves, eye protection, laboratory coat, etc., be used when working with the 

chemicals. Perform sample preparation and analysis in a clean, well-ventilated area that is well removed 

from any possible beryllium contamination. Any area of skin affected by the dissolution or detection 

solutions must be immediately washed with plenty of water. Ammonium bifluoride will etch glass, so it is 

essential that all ammonium bifluoride solutions be contained in plastic labware. Avoid exposure by 

contact with skin or eyes, or by inhalation of vapor. 

SAMPLING [8,9]: 

1. Don a clean pair of gloves. 

2. Demarcate the sampling area (100 cm2 minimum) using a clean template or tape. If a template is used, 

tape the outside edges of the template to the surface to prevent its moving during sampling. 

3. Wet a clean wipe with 0.2 mL of deionized water and wipe the surface to be sampled with firm pressure, using 3 to 4 vertical S-strokes. Fold the exposed side of the wipe in and wipe the area with 3 to 4 

horizontal S-strokes. Fold the wipe once more and wipe the perimeter of the area. 

4. Fold the wipe sample, exposed side in, and place into a labeled 15 mL plastic centrifuge tube.
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 3 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

SAMPLE PREPARATION:

5. Add 5 mL of the dissolution solution (ammonium bifluoride, 10 g/L) to each 15 mL centrifuge tube 

containing a wipe sample, and cap each tube.

6. Place each tube into a mechanical rotator, and rotate for at least 30 min.

NOTE: Rotator may also be substituted by a shaker or an agitator as long as the dissolution solution 

wets the wipe well. Sonication has also been shown to be effective. For dissolution of refractory 

materials such as high-fired beryllium oxide, agitation of the dissolution solution with the 

media must be replaced by heating to 85 °C for 60 minutes or more. Any standard dissolution 

process is particle-size dependent [10]. The two sources of BeO used to validate the method are 

described in the backup data report [11].

7. Filter each solution with a hydrophilic polypropylene syringe filter into a clean tube.

NOTE: This tube should be able to accept a cap so that the solution may be saved and used later for 

reanalysis if required.

8. Pipet 0.1 mL of each sample filtrate into cuvettes containing 1.9 mL of the detection solution. Cap and 

mix briefly.

NOTE: The above procedure is typically used to analyze a range of 0.05 μg to 6 μg of beryllium on the 

sampling media. Alternative ratios of dissolution solution and detection solution may be used 

for analyzing alternative ranges of beryllium concentration. To test a range of 0.005 μg to 0.4 μg 

of beryllium on the sampling media, 0.4 mL of the sample filtrate is added to 1.6 mL of the 

detection solution in the cuvettes. The lysine-free dye solution may also be used for obtaining 

even lower detection limits at a dilution ratio of 3x, where 1.33 mL of the dye solution is mixed 

with 0.67 mL of the filtered solution extracts (Table 3), and beryllium in the range of 0.0005 μg 

to 0.4 μg may be determined.

NOTE: If high iron or titanium concentration is suspected or is evident (owing to the appearance of 

suspended precipitate), allow the solution to settle and filter the solution using a hygroscopic 

syringe filter (e.g., polyethersulfone, or hydrophilic polypropylene).

NOTE: The stability of the detection and the dissolution solution is more than six months and of the 

mixed measurement solution comprising both is greater than 30 days. The solutions must be 

kept in sealed containers, and the detection and mixed solutions must be stored away from 

light. 

NOTE: If the samples are suspected of having a contaminant that fluoresces and has excitation and 

emission spectra that overlap with that of the signal produced by the fluorescent dye bound to 

beryllium, then this contaminant needs to be removed. The presence of such a contaminant 

can be verified by subjecting the filtered sample to fluorescence excitation after the extraction 

step (without adding the fluorescent dye). If a fluorescence signal is detected, then that signal is 

ascribed to the presence of a fluorescent contaminant. To remove the contaminant, high-purity 

activated charcoal is added to the beryllium extraction solution (~10mg/mL) and the extraction 

procedure is carried out at elevated temperature (80 to 90 °C for at least 45 minutes). If the 

beryllium extraction procedure has already been performed, then after the addition of 

activated charcoal, the extraction process is repeated at the elevated temperature. The solution 

is filtered to remove the activated charcoal before adding this to the detection solution to make 

the measurement solution. Details of this process have been published [12,13].

CALIBRATION AND QUALITY CONTROL:

9. Calibrate the fluorometer with beryllium stock standard solutions. Prepare a calibration graph of fluorescence intensity vs. beryllium concentration (ng/mL) in the stock standard.

NOTE: To test a range of 0.05 μg to 6 μg of beryllium on the sampling media, beryllium stock standard 

solutions are made up using beryllium spectrometric standards diluted with the ammonium

bifluoride dissolution solution. A recommended series of stock standard solutions is (800, 200, 

40, 10, and 0) ng/mL. As with the samples, the stock standards are prepared for analysis by 
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 4 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

adding 0.1 mL of beryllium stock standard into 1.9 mL of detection solution (20-fold dilution). 

Please see Table 1. Either of the two detection solutions may be used. 

NOTE: To test a range of 0.005 μg to 0.4 μg of beryllium on the sampling media, a recommended series 

of stock standard solutions is (80, 20, 4, 1, and 0) ng/mL. These standards with lower beryllium 

concentration can be prepared by 10-fold dilution of the stock standards mentioned in the note 

above. As with the samples, these stock standards are prepared for analysis by adding 0.4 mL of 

beryllium stock standard into 1.6 mL of detection solution (5-fold dilution). Please see Table 2. 

Either of the two detection solutions may be used.

NOTE: When using the lysine-free dye solution ONLY, a range can be tested of 0.0005 to 0.4 μg of 

beryllium on the media using a recommended series of stock solutions is (0, 0.15, 0.3, 0.6 and 

2.4) ng/mL. These standards with lower beryllium concentration can be prepared by dilution of 

the stock standards mentioned in the note above. The standards are prepared for analysis by 

adding 0.67 mL of beryllium stock standard into 1.33 mL of detection solution (3-fold dilution). 

This dilution will result in 0, 0.05, 0.1, 0.2 and 0.8 ppb of beryllium in these standards; see 

Table 3. Cellulosic filters (47 mm in diameter) were spiked with a solution of beryllium acetate 

and analyzed in triplicate after extracting beryllium in 5 ml of 1% ABF solution at 85o

C for 60 

minutes and then mixed with lysine-free dye solution in a 3-fold dilution; these results are 

shown in Table 4. The difference in the average fluorescent signals from blanks and the 0 ppb 

standard were subtracted from the fluorescent readings of the spiked filters.

NOTE: If alternative ratios of dissolution solution and detection solution are used for sample preparation, then a similar ratio for calibration solutions is required.

10. Analyze a stock standard, a reagent blank, and a media blank at least once every 20 samples. Ensure 

that the concentration range of the stock standards spans the beryllium levels found in the samples. 

11. Analyze one media spike and one quality control blind spike per 20 samples (minimum of three each 

per sample set) to insure that percent recovery is in control (e.g., 100 ± 15). Correct sample results for 

the average recovery if it differs significantly from 100%.

NOTE: If it is suspected that beryllium oxide may be present, then it is recommended to use beryllium 

oxide for media and blind spikes.

MEASUREMENT:

12. For each sample, obtain the fluorescence intensity at λmax or with optical filter for appropriate wavelength.

13. If the fluorescence response for any of the samples is above the range of responses for the stock 

standards, dilute the sample filtrate with dissolution solution, reanalyze, and apply the appropriate 

dilution factor (D) in subsequent calculations.

CALCULATIONS:

14. Obtain the solution concentration for each sample filtrate, Cs (ng/mL), and the average media blank, Cb

(ng/mL) from the calibration graph.

15. Using the dissolution volumes (normally 5 mL) of sample, Vs (mL), and media blank, Vb (mL), calculate 

the concentration, C (μg/m2

), of Be in the surface area sampled, A (cm2), while accounting for the 

dilution factor (D). 

C=D x (CsVs-CbVb)/(10 x A) μg/100 cm2

NOTE: Tables 1, 2 and 3 can be used for correlating the amount of beryllium in the sampling media 

with the concentrations of beryllium in solution. Table 1 is for testing media with 0.05 μg to 6 μg 

of beryllium at 20-fold dilution; Table 2 is for testing media with 0.005 μg to 0.4 μg of beryllium 

at 5-fold dilution; and Table 3 is for testing media with 0.0005 μg to 0.012 μg of beryllium at 3-

fold dilution. Lysine-free dye solution may be used for any of these dilutions, but for 3x dilution, 

lysine-free dye solution must be used.
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 5 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 1. Correlation of amount of Be in sampling media with Be concentration in stock standard and 

Be concentration as analyzed, assuming 0.1 mL of sample or stock standard is added to 1.9 mL of 

detection solution (20-fold dilution). 

Be concentration in stock 

standard (ng/mL)

Be concentration as analyzed 

(ng/mL)

Amount of Be in the media* 

(ng)

0 0 0 

10 0.5 50

40 2 200

200 10 1000

800 40 4000

*Equals stock standard Be concentration (ng/mL) × volume (5 mL) of dissolution solution used to extract 

media.

Table 2. Correlation of amount of Be in sampling media with Be concentration in stock standard and 

Be concentration as analyzed, assuming 0.4 mL of sample or stock standard is added to 1.6 mL of 

detection solution (5-fold dilution).

Be concentration in stock 

standard (ng/mL)

Be concentration as 

analyzed (ng/mL)

Amount of Be in the media* 

(ng)

0 0 0 

1 0.2 5 

4 1 25

20 4 100

80 16 400

*Equals stock standard Be concentration (ng/mL) × volume (5 mL) of dissolution solution used to extract 

media.

Table 3. Correlation of amount of Be in sampling media with Be concentration in stock standard and 

Be concentration as analyzed, assuming 0.67 mL of sample or stock standard is added to 1.33 mL of 

lysine-free dye solution (3-fold dilution). 

Be concentration in stock 

standard (ng/mL)

Be concentration as 

analyzed (ng/mL)

Amount of Be in the media* 

(ng)

0 0 0 

0.15 0.05 0. 75

0.3 0.1 1.5

0.6 0.2 3 

2.4 0.8 12

*Equals stock standard Be concentration (ng/mL) × volume (5 mL) of dissolution solution used to extract 

media.
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 6 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

Table 4: Analysis of beryllium spiked cellulosic filters (47mm in diameter) using 3-fold dilution of the 

extraction solution with lysine-free dye solution. Beryllium concentration of calibration solutions 

after mixing with the dye solutions were 0, 0.05, 0.1, 0.2 and 0.8 ng/ml. Samples analyzed in 

triplicate, Averages and standard deviations are shown both in ppb in the solution and as μg on the 

filter.

Nominal Be concentration 

on the spiked filter, μg

Be concentration measured in 

the mixture of dye and sample 

solution, ppb ± Std Dev

Be concentration measured 

on the wipes, μg ± Std Dev

0 0.0003±0.0057 0.0000±8.5E-5 

0.0005 0.0375±0.0073 0.00056±11.0E-5 

0.001 0.0735±0.0058 0.0011±8.7E-4 

0.002 0.1262±0.0047 0.0019±7.11E-4 

0.005 0.3396±0.0093 0.0051±1.39E-4 

0.05 3.275 ±0.051 0.049±7.58E-4 

0.48 32.29±0.293 0.484±4.40E-3 

EVALUATION OF METHOD:

The method was evaluated [3,4,11,14] in accordance with published guidelines [15]. Experiments were 

conducted [11] using an Ocean Optics® portable fluorescence device with the following components:

USB 200 spectrometer with spectral grating #2 (UV/Vis 600), LS-1 lamp (380 nm) in LS-450 housing, 

UV-2 casting, OFLV linear filter 200-850, L2 collection lens and slit-200.

Tests were carried out in relative irradiance mode using 2- or 5-second integration times.

The method was evaluated using beryllium oxide spiked onto mixed cellulose ester (MCE) filters at levels of 

(0, 0.02, 0.1, 0.2, 0.3, 0.4, 1.5, 3.0, and 6.0) μg (five samples at each level) [12]. The procedure was also 

evaluated on polyvinyl alcohol (PVA) wipes in an interlaboratory trial with media spiked with BeO at 0.030, 

0.16, 0.32, 1.8, 2.8 and 5.6 μg [7].

Long-term stability of samples was verified from spikes (number [n] = 30) of 0.1 μg Be on MCE filters [9]. 

Samples were analyzed at day one (n = 12) and then one week (n = 6), ten days (n = 3), two weeks (n = 3), 

three weeks (n = 3), and one month (n = 3) after spiking. No diminution of fluorescence signal was 

observed from samples prepared and analyzed after having been stored for up to thirty days.

Interference tests were carried out using solutions of 0 nmol/L, 100 nmol/L, and 1.0 μmol/L Be in the presence of 0.4 mmol/L Al, Ca, Co, Cu, Fe, Ti, Li, Ni, Pb, Sn, U, V, W, or Zn (separate experiments were carried out 

for each potential interferant) [13]. Interlaboratory evaluations of the method were also performed [4,7,14].

The method using the lysine-free detection solution was compared and tested (with the detection solution 

with lysine for comparison) (Table 4) and this was carried out on a GlomaxTM spectrometer (Turner 

Biosystems, Sunnyvale, CA) with an emission filter of 475 ± 5 nm and the excitation was at 360 nm

REFERENCES:

[1] US Department of Energy [1999]. 10 CFR 850, Chronic Beryllium Disease Prevention Program. 

Washington: Code of Federal Regulations.

[2] Dion C, Perrault G [2005]. Summary of good cleanup and decontamination practices for workplaces 

with beryllium-containing dust (Publ. no. R-409). Montreal, Canada: IRSST.
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 7 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

[3] Agrawal A, Cronin J, Tonazzi J, McCleskey TM, Ehler DS, Minogue EM, Whitney G, Brink C, Burrell AK, 

Warner B, Goldcamp MJ, Schlecht PC, Sonthalia P, Ashley K [2006]. Validation of a standardized 

portable fluoresence method for determining trace beryllium in workplace air and wipe samples. J 

Environ Monit 8: 619- 624.

[4] Ashley K, Agrawal A, Cronin J, Tonazzi J, McCleskey TM, Burrell AK, Ehler DS [2007]. Ultra-trace determination pf beryllium in occupational hygiene samples by ammonium bifluoride extraction and 

fluorescence detection using hydroxybenzoquinoline sulfonate. Anal Chim Acta 584: 281-286.

[5] NIOSH [1994]. Elements by ICP (nitric/perchloric acid ashing): Method 7300 (supplement issued 

3/15/03). In: NIOSH Manual of Analytical Methods. 4th ed. Cincinnati, OH: National Institute for 

Occupational Safety and Health, DHHS (NIOSH) Publication No. 94–113.

[6] ASTM International [2015]. D7202 – Standard test method for determination of beryllium in the 

workplace using field-based extraction and fluorescence detection. West Conshohocken, PA: ASTM 

International. 

[7] Ashley K, Wise TJ, Marlow D, Agrawal A, Cronin JP, Adams L, Ashley E, Lee PA [2011]. Trace beryllium 

determination in polyvinyl alcohol wipes by extraction and fluorescence detection – Interlaboratory 

analysis. Anal Methods 3: 1906-1901.

[8] ASTM International [2013]. D6966 - Standard practice for wipe sampling of surfaces for subsequent 

determination of metals. West Conshohocken, PA: ASTM International.

[9] ASTM International [2011]. D7707 - Standard Specification for Wipe Sampling Materials for Beryllium 

in Surface Dust. West Conshohocken, PA: ASTM International.

[10] Goldcamp MJ, Goldcamp DM, Ashley K, Fernback JE, Agrawal A, Millson M, Marlow D,Harrison K 

[2009]. Extraction of beryllium from refractory beryllium oxide with dilute ammonium bifluoride and 

determination by fluorescence – A multi-parameter performance evaluation. J Occup Environ Hyg 6: 

735-744.

[11] McCleskey TM, Agrawal A, Ashley K [2007]. Backup data—method nos. 7704 and 9110 / beryllium, 

Issue 1. NIOSH Docket Office, Cincinnati, Ohio 45226; e-mail: NIOSHDocket@cdc.gov. 

[12] Youmans-McDonald L, Brisson MJ, Bernard M, Agrawal A, Cronin J, Adams, L [2011]. Determining 

beryllium by optical fluorescence using pre-wetted wipes. J ASTM Int 8(3) [JAI 103489].

[13] Agrawal A, Adams LL, Agrawal A, Cronin JP, Tonazzi JCL, Duran BL [2011]. Beryllium measurement by 

optical fluorescence in samples contaminated by fluorescent impurities, J ASTM Int 8(8) [JAI 103411].

[14] Ashley K, McCleskey TM, Brisson MJ, Goodyear G, Cronin J, Agrawal A [2005]. Interlaboratory evaluation of a portable fluorescence method for the measurement of trace beryllium in the workplace. J 

ASTM Int 2(9) [JAI13156].

[15] Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA [1995]. Guidelines for air sampling and analytical method development and evaluation. Cincinnati, OH: National Institute for Occupational 

Safety and Health; DHHS (NIOSH) Publication No. 95–117 
BERYLLIUM in Surface Wipes by Fluorometry: METHOD 9110, Issue 2, 12 December 2015- Page 8 of 8 

NIOSH Manual of Analytical Methods (NMAM), Fifth Edition

METHOD REVISED BY:

T. Mark McCleskey, Los Alamos National Laboratory, Los Alamos, NM; Kevin Ashley, CDC/NIOSH, Division of 

Applied Research and Technology; Anoop Agrawal, Berylliant Inc., Tucson, AZ.

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In addition, 

citations to websites external to NIOSH do not constitute NIOSH endorsement of the sponsoring 

organizations or their programs or products. Furthermore, NIOSH is not responsible for the content of 

these websites. All web addresses referenced in this document were accessible as of the publication date.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

 METHAMPHETAMINE on Wipes by Liquid 9111

 Chromatography/Mass Spectrometry

C6H5·CH2·CH(CH3)·NH·CH3 MW: 149.2 CAS: 537-46-2 RTECS: SH4910000

METHOD: 9111, Issue 1 EVALUATION: Partial Issue 1: 17 October 2011

OSHA: None for surfaces

NIOSH: None for surfaces

Other OELs and guidelines: [1, 2, 3]

PROPERTIES: White solid, MP 171 - 175 °C; VP 0.163 mm Hg 

@ 25°C; pKa = 9.87 @ 25°C; Water Sol. (1.33 

g/100 mL @ 25°C). Log P = 2.07 (octanolwater partition coefficient).

SYNONYMS: (S)-N,α-Dimethylbenzeneethanamine; (S)-(+)-N,α-Dimethylphenethylamine: d-1-Phenyl-2-

methylaminopropane. Methedrine; Desoxyn; chalk; crank; crystal; glass; ice; meth, speed; upper.

SAMPLING

SAMPLER: Wipe

SAMPLE AREA: 100 cm2 or 1000 cm2

SHIPMENT: Ship refrigerated preferably

SAMPLE

STABILITY: At least 7 days at 22 °C

At least 30 days at <6 °C 

FIELD BLANKS: 2 to 10 blanks per set

MEASUREMENT ACCURACY

RANGE STUDIED: Not Determined

BIAS: Not Determined

OVERALL 

PRECISION ( ): Not Determined

ACCURACY: Not Determined

MEASUREMENT

TECHNIQUE: LIQUID CHROMATOGRAPHY/MASS 

SPECTROMETRY-SIM mode. See Table 1.

ANALYTE: Methamphetamine

DESORPTION: 0.1 M sulfuric acid

INJECTION 

VOLUME: 50 μL

MOBILE

PHASE: A: 5/95 acetonitrile/water, 0.1% acetic acid 

B: 95/5 acetonitrile/water, 0.1% acetic acid

COLUMN: 150 mm x 4.6 mm, 5 μm film of dimethyln-octadecylsilane (C18) stationary phase 

to silica gel. Column temperature 40 °C. 

GRADIENT: 100% A for one min, gradient 

to 100% B (9 min), hold 5 min, 

gradient to 100% A (2 min), hold 8 

min. Flow rate is 0.50 mL/min

CALIBRATION: Media spiked standards to cover 

the range (See Table 1)

RANGE: 0.1 – 100 μg/sample

ESTIMATED

LOD: 0.1 μg/sample

PRECISION ( ): 0.067 [4] 

APPLICABILITY: For methamphetamine the range is approximately 0.1 to 100 μg/sample (sample = 100 cm2).

INTERFERENCES: No chromatographic interferences detected.

OTHER WIPE METHODS: NIOSH 9106 uses liquid-liquid extraction and gas chromatography/mass spectrometry (GC/MS) to 

measure multiple drugs [5]. NIOSH 9109 uses solid-phase extraction and GC/MS to measure multiple drugs [6].
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 2 of 10

REAGENTS:

1. Methamphetamine.* 1 mg/mL in methanol.

2. Methamphetamine-D14 1 mg/mL in methanol.

3. Solvents, residue free analytical grades:

a. Isopropanol (IPA) *

b. Acetic Acid *

c. Acetonitrile *

d. Methanol *

4. Concentrated sulfuric acid (AR or trace metals

analysis grade).*

5. Purified gas: nitrogen for drying.

6. Deionized water (ASTM type II).

SOLUTIONS:

1. Prepare spiking solutions of target analyte and

internal standard. Keep solutions refrigerated.

Protect solutions from light.

a. Target analyte spiking solutions are

prepared by diluting the 1000 μg/mL

methamphetamine stock solution to 200 μg/

mL and 20 μg/mL each in methanol.

b. Dilute 1 mL of 1000 μg/mL

methamphetamine-D14 stock solution to 10

mL for a 100 μg/mL (0.1 μg/μL) solution.

2. Desorption solution: 0.1 M sulfuric acid. Add 22

mL conc. sulfuric acid to 4 L deionized water.

3. Prepare Mobile Phase A: 0.1% acetic acid, 5%

acetonitrile in water.

4. Prepare Mobile Phase B: 0.1% acetic acid, 95%

acetonitrile in water.

* See SPECIAL PRECAUTIONS

EQUIPMENT:

1. Wipe, cotton gauze, 3” x 3” (7.6 cm x 7.6 cm)

12-ply or 4” x 4” (10.2 cm x 10.2 cm) 8-ply or

equivalent.

2. Sample storage and shipping container: 50-mL

polypropylene centrifuge tubes with caps or

equivalent.

3. Liquid chromatograph/mass spectrometer,

with integrator or computerized data collection

system, and column.

4. LC autosampler vials, 2-mL, and caps.

5. Volumetric flasks: various sized flasks for making

standards and spiking solutions. A 4-L bottle for

making the desorption solution.

6. Liquid Transfer: 

a. Various microliter syringes for making and

spiking standard solutions.

b. Adjustable 10 to 50-mL desorption solution

dispenser to fit 4-L bottle.

7. Forceps.

8. Latex or nitrile gloves. Avoid vinyl gloves. (See

9111-3, Sampling, Step 1, NOTE 2.)

9. Rotating mixer capable of 10 to 30 rpm.

10. Pasteur pipettes.

11. Template: 10 cm x 10 cm hole. Template made of

relatively rigid disposable cardstock or sheet of

PTFE.

12. Filters: Ion Chromatography filter media, 25

mm syringe filter with 0.45 μm film hydrophilic

polyethersulfone (PES) membrane.

13. Ice or other cold media for shipping.

SPECIAL PRECAUTIONS: The solvents are flammable and have associated adverse health effects. Avoid 

breathing vapors. Avoid skin contact. Work should be performed in an adequate hood. Analysts must wear 

proper eye and hand protection (e.g. latex gloves) to prevent absorption of even small amounts of amines 

through the skin as well as for protection from the solvents and other reagents. Dissolving concentrated 

sulfuric acid in water is highly exothermic. Goggles must be worn. 

Caution must also be exercised in the handling and analysis of samples. Clandestine drug labs may 

produce unknown and seriously toxic by-products.

SAMPLING:

See APPENDIX for special instructions on sampling.

1. Using a new pair of gloves, remove a gauze wipe from its protective package. Moisten the wipe with

approximately 3 to 4 mL of methanol (or isopropanol).

NOTE 1: Apply no more solvent than that needed to moisten approximately the central 80% of the area 

of the gauze wipe. Excess solvent may cause sample loss due to dripping from the wipe.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 3 of 10

NOTE 2: Do not use vinyl gloves due to the potential for leaching of phthalate plasticizers and 

contamination of the samples.

2. Place the template over the area to be sampled (may tape in place along outside edge of template). Wipe

the surface to be sampled with firm pressure, using vertical S-strokes. Fold the exposed side of the pad in 

and wipe the area with horizontal S-strokes. Fold the pad once more and wipe the area again with vertical 

S-strokes.

3. Fold the pad, exposed side in, and place in shipping container and seal with cap.

NOTE: Keep samples refrigerated (<6 oC). While methamphetamine and several related amines are 

stable on the recommended wipe media for at least 7 days at room temperature, refrigeration is 

recommended as soon as possible.

4. Clean the template before use for the next sample or use a new disposable template.

5. Label each sample clearly with a unique sample identifier.

6. Prepare a minimum of two field blanks with one field blank for every ten samples.

NOTE: In addition, include at least 3 media blanks for the analytical laboratory to use for their purposes.

The wipes used for the media blanks should be from the same lot as the field samples.

SAMPLE PREPARATION: 

7. Desorption from media:

a. Remove cap from shipping container. Sample media should fit loosely in the container. If not,

rearrange media carefully with rinsed forceps or transfer to a larger container. If the sample media are

transferred to a larger container, do not discard the original container. Samples may consist of more

than one wipe. If this is the case, internal standard and desorption solution volumes may be adjusted

accordingly.

b. Spike exactly 50 μL of internal standard spiking solution onto each wipe sample.

c. Add 30 mL desorption solution (0.1 M sulfuric acid). If the samples were transferred to a larger

container, the original shipping container must be rinsed with the desorption solution first, shaken,

and the rinsate decanted into the larger container.

d. Cap securely and mix contents by inverting the tubes end over end on a rotary mixer at 10-30 rpm for

at least one hour.

NOTE 1: The desorption solution must percolate freely through the gauze wipes.

NOTE 2: If there is reason to believe that the samples may be alkaline enough to overcome the acidity

of the desorption solution (e.g. wipes of unpainted concrete or stucco surfaces), then the pH 

must be adjusted to about ≤ 4. See APPENDIX for instructions.

e. Filter an aliquot of the sample through a 0.45 μm pore coated with hydrophilic polyether sulfone on

the ion chromatography filter media (25-mm diameter) for analysis.

8. Transfer the filtered sample into a vial and cap.

9. Analyze samples, standards, blanks, and Quality Control samples (QCs) by LC-MS. (See MEASUREMENT,

steps 13-15.)

CALIBRATION AND QUALITY CONTROL:

10. Determine retention time using the column and chromatographic conditions specified on page 9111-1.

11. Calibrate daily with at least six media spiked calibration standards and a blank.

a. Prepare the target analyte spiking solutions. (See SOLUTIONS, 9111-2)

b. Prepare calibration standards and media blanks in clean shipping containers (e.g. 50-mL

polypropylene centrifuge tubes.)

c. Spike a known volume of target analyte spiking solution into each calibration standard by spiking

directly onto the media. Use the spiking volumes suggested in Table 2 to cover the desired range.

d. Analyze these along with the field samples. (See MEASUREMENT, steps 13-15.)
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 4 of 10

12. Prepare matrix-spiked and matrix-spiked duplicate quality control samples (QC and QD).

a. Cotton gauze from the same lot used for taking samples in the field should be provided to the

analytical laboratory to prepare these matrix-spiked QC samples.

b. The quality control samples (QC and QD) must be prepared independently at concentrations within the

analytical range. (See Table 2 for applicable concentration ranges.)

c. One quality control media blank (QB) must be included with each QC and QD pair.

d. The quality control samples must be prepared at the rate of one set (QB, QC, and QD) per 20 samples or

less.

e. Transfer clean gauze wipes to new shipping containers.

NOTE: If two gauze wipes were used for the majority of samples in an analytical set, use two clean

gauze wipes for each QB, QC, and QD. 

f. Spike QC and QD with a known amount of target analyte as suggested in Table 2.

g. Process quality control samples along with the calibration standards, blanks, and field samples through

steps 7 and 8.

h. Analyze these along with the calibration standards, blanks, and field samples. (See MEASUREMENT,

steps 13-15.)

MEASUREMENT:

13. Analyze the calibration standards, quality control samples, blanks, and samples by LC-MS.

a. Use the following suggested analytical sequence.

i. Calibration standards.

ii. Matrix spiked quality control samples (QC and QD), one set for every 20 samples or less.

iii. A media blank (QB), one for every 20 samples or less.

iv. Samples (up to 10) including one sample duplicate.

v. A continuing calibration verification (CCV) standard consisting of one of the initial calibration

standards.

vi. A media blank.

b. Set liquid chromatograph according to manufacturer’s recommendations and to conditions listed

previously.

c. Set mass spectrometer to scan for ions 119, 150, and 164 in SIM mode. Further suggestions for MS

conditions are listed in Table 1 but will vary for particular instruments and conditions. See Note 2 in

Table 1.

d. Inject 50 μL of the sample aliquot into liquid chromatograph.

e. After analysis, the vials should be promptly recapped and refrigerated if further analysis is anticipated.

Samples are stable refrigerated for at least seven days.

14. Using extracted ion current profiles for the primary (quantification) ions specific to methamphetamine

and the internal standard, measure the LC peak area of each respective peak and compute relative peak

areas by dividing the peak area of the analyte by the area of the internal standard. Recommended primary

(quantification) ions and internal standard ions are given in Table 1. Prepare a calibration graph (relative

peak area vs. μg analyte per sample).

15. Samples from initial investigations of clandestine laboratories are likely to include highly contaminated

samples. If sample results exceed the upper range of the calibration curve, the sample in the LC vial may

be diluted with the sulfuric acid desorption solution and reanalyzed.

CALCULATIONS:

16. Determine the mass in μg/sample of methamphetamine found in the wipe samples and in the media

blank from the calibration graph.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 5 of 10

17. Calculate final concentration, C, of methamphetamine in μg/sample:

Where:

c = concentration in sample (in μg/sample determined from the calibration curve)

dilution factor, if applicable 

V1 = volume in μL of internal standard spiking solution used to spike samples.

V2

 = volume in μL of internal standard spiking solution used to spike the standards.

b = concentration in media blank (in μg/sample determined from the calibration curve).

18. Report concentration, C’, in μg per total area wiped (in cm2) as follows:

Where:

C = μg/sample (step 17).

A = Total area wiped in cm2 per sample.

NOTE: For example, if the sample was a composite sample and the area was 400 cm2, report results as 

μg/400 cm2

. In general, if the area wiped was greater than or less than 100 cm2, do not convert 

value to μg/100 cm2. To avoid confusion, report separately both μg/sample (C) and the total area 

wiped in cm2

 per sample (A) for both discrete and composite samples.

EVALUATION OF METHOD:

This method was evaluated for methamphetamine over a range of approximately 0.4 μg/sample to 17.8 μg/

sample on cotton gauze. These concentration levels represent approximately 3 through 100 times the limit of 

quantitation (LOQ) level. Results are reported in the Backup Data Report for 9111s[4].

The limit of detection (LOD) and LOQ were determined by preparing a series of media spiked standards, 

desorbing in the sulfuric acid desorption solution and analyzing in the SIM mode. The LODs were estimated 

using the procedure of Burkart [7]. An LOD of less than 0.02 μg/sample for methamphetamine on wipes was 

achieved in the SIM mode. The LOD was set at 0.05 μg/sample and the LOQ at 0.15 μg/sample for method 

development purposes. Lower LODs can be achieved in practice by including calibration standards at lower 

concentration levels and with proper instrument maintenance. The cleanliness and performance of the mass 

spectrometer must be maintained such that at a minimum of 0.1 μg/sample a signal of at least 5 to 10 times 

the baseline noise is achievable.

Precision and accuracy were determined by analyzing 6 replicates at each of 4 concentration levels 

(nominally 0.44, 1.8, 4.4, and 18 μg/sample). Accuracy was calculated using equations and methodology 

found in the NIOSH Technical Report “Guidelines for Air Sampling and Analytical Method Development and 

Evaluation” [8]. Using all data, method precision ( ) was 0.06663. Accuracy was 20.7% and mean bias was 

-0.09753.

Long term sample storage stability was determined for periods up to 30 days under refrigeration (4 °C ± 2 °C) 

and for up to 7 days at room temperature (22-24 °C). Since long term storage measures only the viability of 
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 6 of 10

analytes on a particular media over time, this determination was not repeated for this particular method; the 

reader is directed to NIOSH 9106 [5] for more detail. All recoveries were found to be 93.5% or better.

Recovery of amphetamines from six different types of surfaces using cotton gauze was evaluated. The study 

and results are reported in NIOSH 9109 [6]. The practice of serial wiping (wiping the same surface area a 

second time with a second gauze wipe and combining both wipes as a single sample) was evaluated. Four 

solvents for wetting the gauze were tested (distilled water, 5% distilled white vinegar, isopropanol, and 

methanol). Six replicate samples were taken on a latex painted wall. Recovery and precision results are 

presented in the previously mentioned Backup Data Report. In summary, the effectiveness of the various 

solvents using a single wipe on a latex painted wall were as follows: water, 46% recovery; 5% distilled white 

vinegar, 55% recovery; isopropanol, 64% recovery and methanol, 87% recovery. Average recoveries with 

isopropanol from all the surfaces tested were greatly improved with a repeat (serial) wipe (11% improvement 

compared to only about 6% improvement with methanol). The serial wipe is added to the first wipe and 

constitutes a single sample.

REFERENCES:

[1] ACGIH [2013]. TLVs and BEIs based on the documentation of the Threshold Limit Values for chemical

substances and physical agents and Biological Exposure Indices. Cincinnati, Ohio: American Conference

of Governmental Industrial Hygienists. [www.acgih.org]. Data accessed April 2014.

[2] Institut fur Arbeitsschutz der Deutschen Gesetzlichen Unfallversicherung [2013]. GESTIS database on

hazardous substances (German Social Accident Insurance). Sankt Augustin, FRG: [http://www.dguv.de/

ifa/GESTIS]. Data accessed April 2014.

[3] EPA [2013]. Voluntary guidelines for methamphetamine laboratory cleanup. [https://www.epa.gov/sites/

production/files/documents/meth_lab_guidelines.pdf ]. Date accessed March 2016.

[4] Siso MC, McKay TT, Reynolds JM, Wickman DC [2005]. Backup data report for NIOSH 9111,

Methamphetamine on wipes by liquid chromatography-mass spectrometry. Unpublished. Prepared

under NIOSH Contract 200-2001-0800.

[5] NIOSH [2011]. Methamphetamine and illicit drugs, precursors, and adulterants on wipes by liquid-liquid

extraction: Method 9106. In: Ashley KA, O’Connor PF, eds. NIOSH manual of analytical methods. 5th

ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and

Prevention, National Institute for Occupational Safety and Health, [www.cdc.gov/niosh/nmam/].

[6] NIOSH [2011]. Methamphetamine and illicit drugs, precursors, and adulterants on wipes by solid phase

extraction: Method 9109. In: Ashley KA, O’Connor PF, eds. NIOSH manual of analytical methods. 5th

ed. Cincinnati, OH: U.S. Department of Health and Human Services, Centers for Disease Control and

Prevention, National Institute for Occupational Safety and Health, [www.cdc.gov/niosh/nmam/].

[7] Burkart JA [1986]. General procedures for limit of detection calculations in the industrial hygiene

chemistry laboratory. Appl Ind Hyg 1(3):153-155.

[8] NIOSH [1995]. NIOSH Technical Report: Guidelines for air sampling and analytical method development

and evaluation. By Kennedy ER, Fischbach TJ, Song R, Eller PM, Shulman SA. Cincinnati, OH: U.S.

Department of Health and Human Services, Public Health Services, Centers for Disease Control and

Prevention, National Institute for Occupational Safety and Health, DHHS (NIOSH) Publication No. 95-117.

[9] Serrano KA, Martyny JW, Kofford S, Contreras JR, Van Dyke MV [2012]. Decontamination of clothing and

building materials associated with the clandestine production of methamphetamine. J Occup Environ

Hyg 9(3):185-197.

[10] Van Dyke MV, Serrano KA, Kofford S, Contreras J, Martyny JW [2011]. Variability and specificity associated

with environemental methamphetamine sampling and analysis. J Occup Environ Hyg 8(11):636-41. 

METHOD DEVELOPMENT BY:

Maria Carolina Siso, Thomas T. McKay, John M. Reynolds, and Don C. Wickman, DataChem Laboratories, Inc., 

Salt Lake City, Utah under NIOSH Contract CDC 200-2001-0800.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 7 of 10

Disclaimer: Mention of any company or product does not constitute endorsement by NIOSH. In

addition, citations to websites external to NIOSH do not constitute NIOSH endorsement of the

sponsoring organizations or their programs or products. Furthermore, NIOSH is not responsible for

the content of these websites. All web addresses referenced in this document were accessible as of

the publication date.

Table 1. Suggested Mass Spectrometer SIM Conditions

 Ionization Mode: API-ES (Atmospheric Pressure Ionization –Electrospray)

 Polarity: Positive

Fragmentator: 100

Gain: 3.0 EMV

Actual Dwell: 294

SIM ions: 119 Methamphetamine confirmation ion

150 Quantitation ion for methamphetamine

164 Ion for methamphetamine-D14

Spray Chamber: (Optimize for the particular instrument in use.)

Gas Temperature: 200 °C

Drying Gas: 12.0 L/min

Nebulizer Pressure 50 psig

Note 1: Methamphetamine and the internal standard essentially co-elute. Monitor 

m/z ions 119 and 150 for methamphetamine quantitation and 164 for 

methamphetamine-D14.

Note 2: These instrumental conditions are suggestions and should be optimized by the 

analyst. Each mass spectrometer will be different and the acid concentration and 

composition will alter the conditions. Furthermore there is the possibility of using 

MS/MS for these analyses if the laboratory is equipped with that instrumentation. 

This tandem MS could add specificity and sensitivity to the method but was not 

part of this method development.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 8 of 10

Table 2. Suggested spiking schedule for calibration standards

Standard

Volume (μL) of 

Methamphetamine Spiking 

Solution Spiked on Media

200 μg/mL 

solution

20 μg/mL 

solution

Internal 

Standard 

Spike (μL)

Desorption 

Solution 

(mL)

Final Concentration (μg/

sample)

1 500 50 30 100

2 100 50 30 20

3 25 50 30 5

4 5 50 30 1

5 25 50 30 0.5

6 5 50 30 0.1

7 2.5 50 30 0.05

8 1.2 50 30 0.024

Figure 1. LC-MS total ion chromatogram of methamphetamine (m/z 119, 150, 164).
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 9 of 10

APPENDIX: 

SAMPLING

NOTE: For further information and data on the effects of surface composition and porosity as well as the 

use of different solvents on the efficacy of wipe sampling, please see reports by Serrano et al [9] 

and Van Dyke et al [10].

1. Follow specific requirements of surface area to be wiped (usually 100 cm2

) and action threshold (or

maximum allowable residual level) set by the agency having legal jurisdiction or specified by the client. 

Uptake rates depend upon the wipe sampling method used so the specific wipe technique used must be 

specified and any deviations from the required wipe sampling requirements noted.

2. The following steps only summarize the overall sampling procedure and are not intended to be used as a

shortcut or wipe sampling procedures that may be specified by the legal jurisdiction or the client.

3. Prepare a rigid template from disposable cardstock or a sheet of PTFE having either a 10 cm x 10 cm or

1 ft x 1 ft (1000 cm2) square hole cut according to the dimensions required by the regulatory agency. 

The template must be able to retain its shape during wiping to ensure that the areas wiped were 100 

cm2 or 1 ft2. Single-use disposable cardstock is preferred because it eliminates the possibility for crosscontamination and the necessity to take a blank wipe between samples in step 3.

4. Provide enough wipe media from the same lot to cover all required laboratory media blanks, fieldequipment blanks, samples and sample duplicates, and quality control samples. Use gauze in sterile 

packaging to minimize the chance for cross-contamination which might more easily occur with open 

bulk packaged cotton gauze. The gauze wipes needed for the laboratory media blanks and QC samples 

are to be sent to the laboratory in their unopened sterile packages.

5. Secure the template(s) to the area(s) to be wiped (e.g. with tape along outside edge of template). If

a single-use disposable template is not used, clean the template between samples to avoid crosscontamination and provide laboratory with a blank wipe of the cleaned template between samples to 

ensure that no cross-contamination has occurred.

6. With freshly gloved hand, take one gauze and wet it with isopropanol or methanol (about 3-4 mL

for either the 3” x 3” (7.5 cm x 7.5 cm) 12-ply or the 4” x 4” (10 cm x 10 cm) 8-ply cotton gauze wipes). 

Alternatively, pre-wet and insert the gauze wipes into the sample containers off-site. This avoids 

any possibility of the bottle of methanol or isopropanol becoming contaminated on-site with 

methamphetamine. If the wipes were prepared off-site, then remove pre-wetted gauze wipe from sample 

container, opening only one sample container at a time. In either case, squeeze out and discard any 

excess solvent from the gauze wipe. Use fresh latex or nitrile gloves for each separate sample and blank. 

Do not use vinyl gloves due to the potential for leaching of phthalate plasticizers and contamination of 

the samples.

7. Wipe Techniques

a. Concentric Squares Wiping Technique (particularly suitable for smooth and non-porous surfaces):

Fold the pre-wetted gauze in half and then fold in half again. Using firm pressure wipe the area

within the template. Start at one of the inside corners of the template and wipe in concentric squares,

progressing toward the center. End with a scooping motion. Without allowing the gauze to touch any

other surface, reverse the last fold so that the exposed side of the gauze is facing inward and using a

fresh surface of the gauze, wipe the same area in the same manner as before. Roll or fold the gauze

again and insert into the shipping container.

b. Side-to-side Wiping (or Blotting) Technique (particularly suitable for rough, porous, and/or soiled

surfaces): Fold the pre-wetted gauze in half and then fold in half again. Using firm pressure wipe or

blot the area within the template with at least five overlapping side-to-side horizontal passes (see

NOTE) beginning at the top and progressing to the bottom in a “Z” pattern. End with a scooping

motion. If blotting, blot at least five times on each horizontal pass (see NOTE). Without allowing the

gauze to touch any other surface, reverse the last fold so that the exposed side of the gauze is facing

inward. Using a fresh surface of the gauze, wipe or blot the area again with at least five overlapping

top-to-bottom vertical passes beginning at the left side and progressing to the right in an “N” pattern.
NIOSH Manual of Analytical Methods (NMAM), Fifth Edition 

METHAMPHETAMINE on Wipes by LC/MS : METHOD 9111, Issue 1, dated 17 October 2011 - Page 10 of 10

If blotting, blot at least five times on each vertical pass. Roll or fold the gauze again and insert into 

the shipping container. Blotting is suggested in areas so soiled or rough that the threads of the gauze 

media are continually snagged. 

NOTE: On areas larger than 100 cm2, more than five passes and blots will be needed.

c. Repeat or Serial Wiping: If isopropanol is used for wiping, a serial or repeat wipe sample of the same

area with a fresh gauze wipe will improve sampling efficiency. For serial wiping, repeat the wiping 

procedure described above (APPENDIX 7a or 7b) with a fresh gauze wipe. Place the second gauze wipe 

into the same shipping container as the first gauze. The 50-mL polypropylene centrifuge tubes are 

large enough to contain up to two gauze wipes of either the 3” x 3” 12-ply or 4” x 4” 8-ply sizes. 

NOTE: If the area to be wiped remains substantially wet from the first gauze, the second gauze wipe 

might be used in the dry state to soak up the residual solvent from the first gauze wipe.

8. Cap shipping containers securely and keep refrigerated (<6 °C). Make sure caps are not cross-threaded.

Containers must have no chips, fractures, or other irregularities on the sealing edge. Do not use 

polyethylene plastic bags. While methamphetamine and several related amines are stable on the 

recommended wipe media for at least 7 days at room temperature, refrigeration is recommended as soon 

as possible.

9. Label each sample clearly with a unique sample identification number or name, and the date, time,

location, and initials or identification number of the individual taking the sample. The above information 

and a description of the sample and the area wiped should also be recorded in a logbook for later 

correlation with the analytical results.

SAMPLE PREPARATION

Samples requiring pH adjustment: If there is reason to believe that the samples may be alkaline enough to 

overcome the acidity of the desorption solution (e.g. wipes of unpainted concrete or stucco surfaces), then 

the pH must be adjusted to about ≤ 4. The pH may be checked with pH paper or monitored with the addition 

of about 2 drops of the mixed pH indicator solution of bromothymol blue and phenolphthalein. (The color 

should be yellow and not green or blue.) The preparation of this indicator solution can be found in NIOSH 

9106 or NIOSH 9109 [5, 6]. If the pH needs to be adjusted, a solution of dilute (2.5 to 3 M) sulfuric acid is used 

and added dropwise. Mix the contents by shaking or inversion a few times by hand after each addition of acid 

before checking the pH.