EPA 600/R-13/137 | August 2013 | www.epa.gov/ord

Evaluation of Vacuum-based 

Sampling Devices for Collection of 

Bacillus Spores from Environmental 

Surfaces

Assessment and Evaluation Report

Offi ce of Research and Development

National Homeland Security Research Center
ii

EPA 600-R-13-137

Evaluation of Vacuum-based Sampling Devices for Collection of 

Bacillus spores from Environmental Surfaces

Assessment and Evaluation Report

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711
iii

Disclaimer

The United States Environmental Protection Agency (EPA), through its Office of Research and 

Development’s National Homeland Security Research Center, managed this investigation through EP-C09-027 WA 2-10 and 3-10 with ARCADIS U.S., Inc. The effort was funded through an interagency 

agreement (RW-75-92345701) with the US Centers for Disease Control and Prevention (CDC). This 

report has been peer and administratively reviewed and has been approved for publication as an EPA

document. It does not necessarily reflect the views of the EPA. No official endorsement should be 

inferred. This report includes photographs of commercially available products. The photographs are 

included for purposes of illustration only and are not intended to imply that EPA approves or endorses the 

product or its manufacturer. The EPA does not endorse the purchase or sale of any commercial products 

or services.

Questions concerning this document or its application should be addressed to: 

M. Worth Calfee, Ph.D.

Decontamination and Consequence Management Division

National Homeland Security Research Center

U.S. Environmental Protection Agency (MD-E343-06)

Office of Research and Development

109 T.W. Alexander Drive

Research Triangle Park, NC 27711

Phone: 919-541-7600

Fax: 919-541-0496

E-mail: calfee.worth@ epa.gov
iv

Acknowledgments 

This effort was managed by the principal investigator from the National Homeland Security Research 

Center (NHSRC) within EPA’s Office of Research and Development. Laura Rose (CDC’s Clinical and 

Environmental Laboratory Branch, Division of Healthcare Quality Promotion (DHQP) within the National 

Center for Emerging and Zoonotic Infectious Diseases (NCEZID) and Dino Mattorano (EPA’s Chemical, 

Biological, Radiological, and Nuclear Consequence Management Advisory Team (CBRN CMAT) within 

the Office of Emergency Management (OEM) provided critical inputs into this investigation. Additionally, 

the efforts of Stephen Morse and Angela Weber (CDC’s Office of Environmental Microbiology, Division of 

Foodborne, Waterborne, and Environmental Diseases (DFWED), within the NCEZID are gratefully 

recognized.

The authors would like to thank the peer reviewers for their significant contributions. Specifically, the 

efforts of Lisa Delaney (National Institute for Occupational Safety and Health (NIOSH)), Matt Arduino 

(CDC), and Erin Silvestri (EPA) are recognized. 
v 

Table of Contents 

Disclaimer.............................................................................................................................................iii

Acknowledgments................................................................................................................................iv

Table of Contents ..................................................................................................................................v

List of Appendices...............................................................................................................................vii

List of Tables......................................................................................................................................viii

List of Acronyms and Abbreviations....................................................................................................x

Executive Summary.............................................................................................................................xii

1 Introduction ....................................................................................................................................1

1.1 Process .....................................................................................................................................1

1.2 Project Objectives......................................................................................................................1

1.3 Experimental Approach .............................................................................................................2

1.3.1 Vacuum-based Sampling Device Evaluation Tests – Phase 1............................................2

1.3.2 HVAC Inoculation Tests – Phase 2 ....................................................................................2

1.3.3 HVAC Sampling Tests – Phase 2.......................................................................................2

1.3.4 Inoculum Variability and Level Tests – Phase 3..................................................................3

1.3.5 Linearity Recovery Tests and Laboratory Variability Tests – Phase 3................................. 3

1.4 Definition of Sampling Efficiency................................................................................................3

2 Materials and Methods...................................................................................................................4

2.1 Test Materials and Deposition....................................................................................................4

2.1.1 Coupon Preparation...........................................................................................................4

2.1.2 Bacillus Spore Preparation................................................................................................. 6

2.1.3 Coupon Inoculation............................................................................................................6

2.2 Test Matrix ................................................................................................................................9

2.3 Sampling and Analytical Procedures........................................................................................11

2.3.1 Swab Sampling................................................................................................................11

2.3.2 Polyester-Rayon Blend (PRB) Wipe Sampling ................................................................. 11

2.3.3 Sponge-wipe Sampling....................................................................................................12

2.3.4 Vacuum Sock Sampling and Analysis ..............................................................................12

2.3.5 37 mm MCE and 37 mm PTFE Sampling.........................................................................13

2.3.6 Trace Evidence Filter Sampling .......................................................................................14

2.3.7 Operational Assessment..................................................................................................14

2.3.8 HVAC filter extraction.......................................................................................................15
vi

2.3.9 Quartz extraction .............................................................................................................15

2.3.10 Direct Extraction of Stainless Steel ..................................................................................15

2.4 Sampling Strategy ...................................................................................................................16

2.4.1 Sampling/Monitoring Points .............................................................................................16

2.5 Sampling Handling and Custody..............................................................................................18

2.5.1 Preventing Cross-contamination during Coupon Preparation............................................18

2.5.2 Preventing Cross-contamination during Sampling ............................................................18

2.5.3 Preventing Cross-contamination during Analysis..............................................................19

2.5.4 Sample Containers ..........................................................................................................19

2.5.5 Sample Identification .......................................................................................................19

2.5.6 Information Recorded by Field Personnel.........................................................................20

2.5.7 Sample Preservation .......................................................................................................20

2.5.8 Sample Holding Times.....................................................................................................20

2.5.9 Sample Custody ..............................................................................................................20

2.5.10 Sample Archiving.............................................................................................................21

2.6 Statistical Analysis Methods.....................................................................................................21

3 Results and Discussion ...............................................................................................................22

3.1 Phase 1...................................................................................................................................22

3.1.1 Results of Statistical Analyses .........................................................................................24

3.1.2 Operational Parameters...................................................................................................24

3.1.2.1 Sampling Time and Ease .........................................................................................24

3.1.2.2 Analysis Time and Ease...........................................................................................25

3.1.2.3 Cost.........................................................................................................................25

3.2 Phase 2...................................................................................................................................26

3.3 Phase 3...................................................................................................................................30

4 Quality Assurance........................................................................................................................35

4.1 Sampling, Monitoring, and Analysis Equipment Calibration ......................................................35

4.2 Data Quality ............................................................................................................................36

4.3 QA/QC Checks........................................................................................................................36

4.4 Acceptance Criteria for Critical Measurements.........................................................................38

4.5 Data Quality Audits..................................................................................................................39

4.6 QA/QC Reporting ....................................................................................................................39
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5 Summary.......................................................................................................................................40

5.1 Phase 1...................................................................................................................................40

5.2 Phase 2...................................................................................................................................40

5.3 Phase 3...................................................................................................................................40

5.4 Lessons Learned and Application of Vacuum-based Methods to Field Use .............................. 40

5.5 Future Research......................................................................................................................42

References...........................................................................................................................................43

 List of Appendices

Appendix A Miscellaneous Operating Procedures (MOPs) included as a separate document
viii

List of Figures

Figure 2-1. Carpet Coupon..................................................................................................................4

Figure 2-2. Upholstery Coupon............................................................................................................5

Figure 2-3. Concrete Coupon ..............................................................................................................5

Figure 2-4. Schematic of Deposition Apparatus for Inoculation under Flow Conditions ......................... 8

Figure 2-5. Vacuum Sock Kit and Individual Sock..............................................................................13

Figure 2-6. Filter (37 mm) Cassette with Nozzle and Tubing ..............................................................14

Figure 2-7. 3M® Trace Evidence Filter (TEF) Vacuum-based Sampling Device .................................. 15

Figure 3-1. Recovery from Materials for each Vacuum Method. . ......................................................22

Figure 3-2. Relative Recovery – Data presented as mean relative recovery (RR)............................... 23

Figure 3-3. Recovery from HVAC Filters Using 5 Sampling Methods – Test HI2................................. 27

Figure 3-4. Recovery from Mechanical MERV 8 Filter – Test HS .......................................................28

Figure 3-5. Recovery from Electrostatic HVAC Filter – Test HS..........................................................29

Figure 3-6. Recovery from Stainless Coupons during INOC Series Tests........................................... 31

Figure 3-7. Mean CFU/Sample from Linear Recovery (LR) Test.........................................................32

Figure 3-8. Laboratory Variability for Two Vacuum Methods - Test LV ...............................................34

List of Tables

Table 2-1. Phase 1Test Matrix............................................................................................................9

Table 2-2. Phase 2 Test Matrix........................................................................................................10

Table 2-3. Phase 3 Test Matrix.........................................................................................................10

Table 2-4. Frequency of Sampling Monitoring Events .......................................................................17

Table 2-5. Critical and Non-Critical Measurements ...........................................................................17

Table 3-1. Recovery from Materials for each Vacuum Method ..........................................................22

Table 3-2. Relative Recoveries from all Devices and Material Surface Types................................... 23

Table 3-3. Average Recovery from HVAC Filter Extraction – Test HI1 ..............................................26

Table 3-4. Recovery from HVAC filters – Test HI2 ............................................................................26

Table 3-5. HVAC Vacuuming Recovery – Test HS............................................................................27

Table 3-6. Student’s t-test values from Electrostatic filters ................................................................28

Table 3-7. Mean Recovery from Stainless Steel Coupons (n = 3), Phase 2 Tests ............................. 30

Table 3-8. ANOVA – Comparison of the Three Stainless Steel Recovery Methods within each 

Phase 2 Test Run (n = 3)................................................................................................. 30
ix

Table 3-9. Mean Recoveries from Stainless Steel using the Sponge-wipe Method - Test INOC 

(n = 10)............................................................................................................................30

Table 3-10. Mean Recoveries (CFU/sample) from Test LR (n=5)........................................................31

Table 3-11. Percent Relative Recovery of Vacuum Devices (Test LR) (n = 5)..................................... 32

Table 3-12. Relative Recovery of Vacuum Methods to Sponge-Wipe – Test LR (n=5)......................... 33

Table 3-13. Laboratory Variability for Two Vacuum Methods (Test LV)...............................................34

Table 4-1. Sampling and Monitoring Equipment Calibration Frequency.............................................35

Table 4-2. QA/QC Sample Acceptance Criteria ................................................................................37

Table 4-3. Critical Measurement Acceptance Criteria........................................................................38

Table 5-1. Summary of Advantages and Disadvantages of Each Vacuum-Based Sampling 

Method............................................................................................................................41
x 

List of Acronyms and Abbreviations

%Cv Percent Coefficient of Variation

ADA aerosol deposition apparatus

ANOVA Analysis of Variance

APPCD Air Pollution Prevention and Control Division

ATCC American Type Culture Collection

CBRN Chemical, Biological, Radiological, and Nuclear

CDC Centers for Disease Control and Prevention

CFM cubic feet per minute

CFU colony forming unit(s) 

CM critical measurement

cm2 square centimeter

CMAT Consequence Management Advisory Team

COC chain of custody

DCMD Decontamination and Consequence Management Division

DFWED Division of Foodborne, Waterborne, and Environmental Diseases

DHQP Division of Healthcare Quality Promotion 

DPG Dugway Proving Ground

DQI Data Quality Indicator

DQO Data Quality Objective

ECBC Edgewood Chemical Biological Center

EPA U. S. Environmental Protection Agency

EtO ethylene oxide

ft2 square feet

HAZMAT Hazardous Materials

H2O2 hydrogen peroxide

HVAC heating, ventilation, and air conditioning

ID Internal Diameter

in. inch(es) 

INL Idaho National Laboratory

INOC Inoculation

ISO International Organization for Standardization 

L liter(s) 

L min-1 liters per minute

LR Linear Range 

LV Laboratory Variability

MCE Mixed Cellulose Ester

MDI metered dose inhaler

MERV Minimum Efficiency Reporting Value

MOP Miscellaneous Operating Procedure

NCEZID National Center for Emerging and Zoonotic Infectious Diseases

NHSRC National Homeland Security Research Center

NIST National Institute of Standards and Technology

OEM Office of Emergency Management

PBS Phosphate Buffered Saline
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PBST Phosphate Buffered Saline with 0.05% TWEEN® 20

PRB Polyester-Rayon Blend

PTFE Polytetrafluoroethylene

PVC Polyvinylchloride 

rpm rotations per minute

RR% Relative Recovery Percentage

RSD Relative Standard Deviation 

QA Quality Assurance

QAPP Quality Assurance Project Plan

QC Quality Control

SCFM standard cubic feet per minute

SOP Standard Operating Procedure

STS Sodium Thiosulfate

TEF Trace Evidence Filter

VHP Vaporous Hydrogen Peroxide

WAM Work Assignment Manager
xii

Executive Summary

The existing surface sampling strategy for a biological incident involving B. anthracis spores necessitates

the use of various methods depending on the surface type. Currently-recommended surface sample 

collection methods include pre-moistened wipes (for smooth nonporous surfaces), vacuuming (for rough 

and porous surfaces), and wet swabs (for small and/or hard-to-sample areas such as keyboards). The 

currently-used vacuum-based method utilizes woven collection socks attached to a cardboard nozzle. 

Some criticisms of the current method are that the vacuum socks often come from the manufacturer with 

visible holes in the sock seams, the method is vulnerable to cross-contamination between samples, the 

socks are constructed of materials with large pore-sizes (> 1μm), and the filters are cumbersome for 

laboratory handling and extraction during analysis. This project comparatively evaluated the vacuum sock 

and two additional vacuum-based collection devices (37 mm filter cassette and 3MTM Trace Evidence 

Filter) for their sampling efficacy. The 37 mm filter cassette was evaluated with mixed cellulose ester 

(MCE) filters or polytetrafluoroethylene (PTFE) filters installed, each was considered a unique device. 

These data were generated so appropriate sampling devices could be selected following a B. anthracis

incident. 

A known quantity of Bacillus atrophaeus (B. anthracis surrogate) spores was aerosolized and deposited 

onto large coupons (1 square foot (ft2

)) of various materials common to the built environment, including

carpet, upholstery, unpainted (smooth finish) concrete, and two types (electrostatic and mechanical) of 

heating, ventilation and air conditioning (HVAC) filters. Coupons were then subjected to vacuum-based 

sampling and sample analysis according to protocols developed jointly by the US Centers for Disease 

Control and Prevention (CDC) and US Environmental Protection Agency (EPA). Recovery was 

determined for each sampling method according to culture-based microbiological assays following 

physical extraction methods developed by the CDC. 

Phase 1 of this study included the evaluation of vacuum-based sampling devices. These tests were 

conducted with four different devices and one sampling variation (fast or slow sampling rate at which the 

device passed over the sampled surface) for the vacuum sock. Samples were collected from three porous

surface types for each device, each with ten replicate samples. Vacuum and wipe samples were also 

collected from stainless steel coupons as a method to standardize results collected on different days and 

with different inoculation devices.

For carpet, the slow vacuum sock method had the highest spore recovery. For concrete, the 37 mm MCE 

had the highest spore recovery. For upholstery, the 37 mm MCE had the highest spore recovery. The 

vacuum sock method afforded more ease of use and may therefore be more desirable for larger sample 

areas. However, on concrete and upholstery, the 37 mm MCE filter method demonstrated higher 

recoveries per unit area than did the vacuum sock device. The significance of these differences was 

dependent upon the statistical test performed. There are advantages and disadvantages of each device. 

Changing the speed at which the vacuum sock sampling device passed over the surface did not 

necessarily improve the recovery of spores from the porous surfaces tested. 

During Phase 2, collection of B. atrophaeus spores from contaminated HVAC filters with two vacuumbased devices (devices with the highest recoveries from Phase 1, vacuum sock (slow rate) and 37 mm 

MCE cassette) was compared to sampling by extractive methods (removing a portion of the filter and 

extracting spores from the filter matrix). Contamination of HVAC filters during a biological incident, unlike 
xiii

most surfaces, is expected to occur under flow conditions, with HVAC blowers pulling spores to the 

interior of the filter. A method was developed which successfully deposited spores on (and within) HVAC 

filters under flow conditions. 

Two types of clean (new) HVAC filters (electrostatic and mechanical) were inoculated similarly to Phase 

1, yet under flow conditions. The contaminated surfaces were then evaluated using three methods: direct 

extraction from excised sections of the HVAC filter, vacuum sock sampling, and 37 mm MCE filter 

vacuum sampling. In addition, the magnitude and variability of the inoculating metered dose inhaler (MDI) 

was evaluated with three methods: sponge-wipe sampling of stainless steel coupons, pre-moistened

(polyester-rayon blend, PRB) wipe sampling of stainless steel coupons, and direct extraction of stainless 

steel coupons.

Three methods (two vacuum methods and one extractive) of sampling HVAC filters performed reliably on 

mechanical and electrostatic filters. The data suggest that extractive methods are more efficient than 

vacuum-based recovery methods for electrostatic filters, but not for mechanical filters. When comparing 

the two vacuum-based methods, the vacuum sock method provided higher recoveries than the 37 mm 

MCE method from electrostatic filters, but there was no statistical difference in recoveries from the

electrostatic filters. Vacuum-based methods may be more applicable to HVAC filter media that are not 

easily sectioned. 

Phase 2 data also suggested there was no statistical difference between direct extraction, sponge-wipes, 

or pre-moistened PRB wipes for recovery (inoculum magnitude) from stainless steel coupons. Mean 

variability (%Cv) was also similar across all methods, at 28%, 22%, and 20% for the extraction-based 

method, sponge-wipe method, and PRB wipe method. 

During Phase 3, Inoculum Variability and Level Tests (INOCs) were conducted to determine the 

repeatability and magnitude of inoculation from the MDI/ aerosol deposition apparatus (ADA) dosing 

method. Three inoculum levels (corresponding to three concentrations of spores within MDIs - 104

, 106, 

107 spores per coupon) were used to dose up to ten replicate stainless steel coupons. The MDI 

inoculation method was demonstrated over a broad range of surface contamination concentrations, from 

1 to 1 × 104 CFU/square centimeter (cm2

). 

Also during Phase 3, Linearity Recovery (LR) Tests and Laboratory Variability (LV) Tests were conducted 

to determine: (1) if vacuum-based method recoveries were linear over a range of concentrations; and (2) 

if recoveries varied significantly between two different technicians processing (extracting and plating) the 

samples. During these tests, carpet coupons were sampled using two chosen vacuum sampling 

methods: vacuum sock (slow) and 37 mm MCE. Following collection, all samples were extracted and 

plated by two technicians. All samples for both devices and all three inoculum levels were collected on a 

single day so that comparisons could be made across devices, inoculum levels, and technicians. One 

technician operated the vacuum-based collection device for all samples. Recoveries were compared 

between the two technicians to determine if technician-induced variability was significant. The linearity of 

recoveries for each device was determined graphically by plotting recovery versus the targeted inoculum.

The two vacuum methods were demonstrated effective for all inoculum concentrations. Compared to 

sponge-wipes (from stainless steel), the vacuum sock method recovered between 3% and 22% CFU, 

while the 37 mm MCE method recovered between 4% and 140% CFU from carpet samples. Relative

recoveries greater than 100% were possible because these values were determined by dividing vacuum 

method recovery by wipe-based recovery from stainless steel. Greater than 100% relative recovery 
xiv

indicated that the vacuum method out-performed the wipe method. Finally, the extraction procedures for 

both vacuum methods were evaluated by two independent laboratory technicians with no statistical 

difference in the two recoveries. 

These data were collected to aid in sampling device and strategy selection following a biological 

contamination incident.
1 

1 Introduction

The U.S. Environmental Protection Agency (EPA) conducted a study, in collaboration with the Centers for 

Disease Control and Prevention (CDC), to evaluate several vacuum-based sampling devices for 

collection of biological agent from environmental surfaces. Methods for detection and characterization of 

biological agent on surfaces following a bioterrorism incident include the use of swabs, wipes, and 

vacuum. Vacuum-based methods are preferred when sampling porous surfaces. Currently, there are no 

vacuum-based methods validated for collection of Bacillus anthracis spores. Further, multiple vacuumbased devices, sample collection methods, and sample extraction methods were utilized to characterize 

the extent of contamination following the 2001 anthrax incidents. Recently, some work has been 

conducted to characterize the performance of vacuum-based surface sampling devices when used to 

collect Bacillus spores [1-3]. However, significant gaps remain in our understanding of vacuum-based 

sampler performance, efficiencies, ease of use, and applicability to various material surface types. The 

current study was conducted to generate data that could be used to inform selection of appropriate 

sampling methodologies following a B. anthracis incident. Scientifically tested sampling methods will 

provide increased confidence in the ability to characterize contamination following such an incident. 

1.1 Process

Consistent with previous sampling studies [1], spores of Bacillus atrophaeus (formerly known as Bacillus 

globigii or Bacillus subtilis var. niger) served as surrogates for Bacillus anthracis spores. Collection of B.

atrophaeus spores from multiple surface types was evaluated with four vacuum-based sampling devices. 

In addition, collection of biological agent from contaminated heating, ventilation, and air conditioning 

(HVAC) filters with the two top-performing vacuum-based devices was compared to sampling by 

destructive methods (removing a portion of the filter and extracting spores directly from the filter matrix). 

A known quantity of aerosolized dry B.atrophaeus spores was gravitationally deposited onto large 

coupons (35.6 cm x 35.6 cm) of various materials common to the built environment, which included 

carpet, upholstery, unpainted (smooth finish) concrete, and two types of HVAC filters. The coupons were 

then subjected to vacuum-based sampling according to protocols developed jointly by the CDC and EPA. 

Recovery was determined for each sampling method according to culture-based microbiological assays 

following physical extraction methods developed by the CDC. All test parameters, such as coupon 

materials and sizes, sampling methods, and methods of extraction and analysis were determined by 

agreement among participating experts from EPA and the CDC. 

1.2 Project Objectives

The objective of this project was to evaluate four currently-available vacuum-based devices for their 

ability to recover Bacillus spores from environmental surfaces. Performance (relative recovery) of the

devices was compared to recoveries from pre-moistened gauze wipes, used to sample reference 

stainless steel coupons that were inoculated at the same time as the test coupons. Evaluation of 

operational parameters included time required for sample collection, the physical impact on the sampling 

team during collection, time required for sample analysis, and the cost of media and analysis equipment 

and supplies. Another objective of this study was to evaluate vacuum-based sampling of contaminated 

HVAC filters to that of extraction-based methods. 
2 

1.3 Experimental Approach

The experimental approaches that were used to meet the objectives of this project are:

• Use of controlled chambers, standardized sections and spore inoculums.

• Inoculation of material coupons via aerosol deposition of bacterial spores.

• Quantitative assessment of spore recoveries with each device, by sampling representative 

sections of materials and then extraction of sampling media to recover collected spores

(enumerated and reported as colony forming units (CFU)). 

• Use of reference stainless steel coupons to quantify and standardize results for cross-comparison 

of tests.

• Documentation of operational considerations (e.g., cross-contamination, procedural time, impacts 

on materials and personnel).

All testing was conducted at EPA’s Research Triangle Park, NC, campus. 

1.3.1 Vacuum-based Sampling Device Evaluation Tests – Phase 1 

The Vacuum-based Sampling Device Evaluation Tests were conducted with four different device types 

and one sampling variation (sampling speed) for one device type (Table 2-1). Samples were collected 

from three surface types (carpet, upholstery, and concrete) for each device with ten replicate samples. 

Pre-moistened polyester rayon blend (PRB) wipe samples were also collected from reference stainless 

steel coupons as a method to standardize results collected on different days and with different inoculation 

devices. The foundation for this test matrix was described in the Quality Assurance Project Plan (QAPP) 

entitled, “Evaluation of Vacuum-based Sampling Devices for Collection of Biological Agent 

(Decontamination and Consequence Management Division (DCMD) 3.60)” (available upon request). 

1.3.2 HVAC Inoculation Tests – Phase 2 

HVAC Inoculation (HI) Tests were conducted to determine the best inoculation method for HVAC filters. 

Contamination of HVAC filters during a biological incident, unlike most surfaces, is expected to occur 

under flow conditions, with HVAC blowers pulling spores to the interior of the filter. Also, the porous 

nature of the substrate presents challenges to even the settling-based aerosol inoculation. For this 

reason, a modified version of the method developed by Calfee et al., 2013, for depositing aerosolized 

spores on material surfaces, was utilized for this substrate. These tests were conducted to verify that the 

inoculation target (quantity of spores) could be achieved without excessive variability.

1.3.3 HVAC Sampling Tests – Phase 2 

HVAC Sampling (HS) Tests were conducted to determine spore recovery from HVAC filters. Two types of 

HVAC filters (electrostatic and mechanical) were inoculated under flow conditions. The contaminated 

surfaces were then evaluated using three methods: direct extraction, vacuum sock sampling, and 37 mm 

MCE filter vacuum sampling. In addition, the variability of the inoculating method was evaluated with two 
3 

sampling methods: sponge-wipe sampling of stainless steel coupons and direct extraction of stainless 

steel coupons.

1.3.4 Inoculum Variability and Level Tests – Phase 3 

Inoculum Variability and Level (INOC) Tests were conducted to determine the repeatability and 

magnitude of inoculation from metered dose inhaler (MDI) dosing method. Three inoculum levels (103

, 

106

, 107 spores per coupon) were deposited onto as many as ten replicate stainless steel coupons. 

Coupons were sampled by sponge-wipe following the required 18-h deposition time. 

1.3.5 Linearity Recovery Tests and Laboratory Variability Tests – Phase 3 

Linearity Recovery (LR) Tests and Laboratory Variability (LV) Tests were conducted to determine: (1) if 

vacuum-based recoveries were linear over the three inoculation levels demonstrated in the INOC tests; 

and (2) if recoveries varied significantly between two different technicians processing (extracting and 

plating) the samples. During these tests, carpet coupons were sampled using two chosen vacuum 

sampling methods: vacuum sock and 37 mm MCE filters. Following collection, all samples were extracted 

and plated by two laboratory technicians. All samples for both devices and all three inoculum levels were 

collected on a single day so that comparisons could be made across devices, inoculum levels, and 

laboratory technicians. Recoveries were compared between the two laboratory technicians to determine 

if technician-induced variability was significant. The linearity of recoveries for each device was determined 

graphically by plotting recovery versus the targeted inoculum. 

1.4 Definition of Sampling Efficiency

The recovery from vacuum sampling methods was compared to recovery from wipe-based methods (PRB 

wipes or sponge-wipes, or both, dependent upon the test) used to sample stainless steel surfaces with 

identical inoculums. Surface samples from stainless steel were considered the best estimate of the 

number of spores inoculated by the aerosol method due to the high repeatability and high recovery 

efficiencies when wipe sampling from this surface type. When side-by-side comparisons could not be 

made (from separate test days due to the large number of vacuum samples prescribed by the test matrix), 

results were normalized to the stainless steel surface samples.
4 

2 Materials and Methods 

2.1 Test Materials and Deposition

2.1.1 Coupon Preparation

Carpet coupons (Figure 2-1) were 30.5 cm x 30.5 cm (12” x 12”) carpet tiles (Shaw Living Berber sand 

loop, Home Depot, Model # 3W05300100) attached to a 35.6 cm x 35.6 cm (14” x 14”) square of 1.9 cm 

(3⁄4”)-thick plywood. The tiles were attached to the plywood using their self-adhesive backs plus three 

staples on each side. 

Figure 2-1. Carpet Coupon

Upholstery coupons (Figure 2-2) were constructed from a 61.0 cm x 61.0 cm (24” x 24”) piece of fabric 

(Indoor/Outdoor Modern Houndstooth Red Fabric, www.fabric.com, Part# UJ-849) covering a 30.5 cm x 

30.5 cm (12” x 12”) piece of upholstery padding (432 Poly Foam, 2.5 cm by 61.0 cm (1” x 24”), 

OnlineFabricStore, Item# 1243310), placed in the center of a 35.6 cm x 35.6 cm (14” by 14”) square of 

1.9 cm (3⁄4”)-thick plywood. The fabric square was stretched over the foam and wood, and excess fabric 

was folded underneath and stapled to the back side of the plywood backing.
5 

Figure 2-2. Upholstery Coupon

Concrete coupons (Figure 2-3) were formed from Sakrete Sand Mix poured into custom 35.6 cm x 35.6 

cm (14” by 14”) forms. The sand mix was prepared according to indications on the package using a 

trough and garden hose for the water supply. The coupon was smoothed with a trowel and allowed to set

and dry overnight. The coupons were allowed to cure under humid conditions for at least five days. 

Figure 2-3. Concrete Coupon

Mechanical MERV 8 HVAC filters were 35.6 cm x 35.6 cm x 2.5 cm (14” x 14” x 1”) Purafilter 2000 Blue 

series (Purafilter 2000, Las Vegas, NV; http://purafilter2000.com/products.php). Electrostatic MERV 8 

filters were 35.6 cm x 35.6 cm x 2.5 cm (14” x 14” x 1”) Eco-Aire MERV 8 High Cap (Con-Air Industried, 

Inc., Orlando, FL). Stainless steel coupons were Grade 316, cut to 35.6 cm x 35.6 cm (14” by 14”) or 10.2 

cm x 15.2 cm (4” x 6”). Only the centermost 30.5 cm x 30.5 cm (12” x 12”) of each 35.6 cm x 35.6 cm

(14” by 14”) coupon was sampled. 
6 

All coupons were placed in sterilization bags and sterilized prior to use. Carpet and upholstery coupons 

were sterilized by a minimum 250 ppmv Vaporous Hydrogen Peroxide (VHP) cycle for four hours using a 

STERIS ED1000. Concrete and stainless steel coupons were sterilized prior to use by steam autoclave 

utilizing a gravity cycle program consistent with the EPA NHSRC Microbiology Laboratory Miscellaneous 

Operating Procedure (MOP) 6570 (see Appendix A for all associated MOPs). HVAC filters were sterilized 

with ethylene oxide. Sterility was evaluated by swab sampling one coupon from each sterilization batch 

according to MOP 3135. Prior to use, the coupons treated with VHP were incubated at 30-35 °C for a 

minimum of two days to force off-gassing of residual hydrogen peroxide (H2O2) from material coupons as 

suggested by Baron et al. [4], so that biocidal activity was prevented. 

2.1.2 Bacillus Spore Preparation

The test organism for this work was a powdered spore preparation of B. atrophaeus (American Type 

Culture Collection (ATCC) 9372) and silicon dioxide particles. The preparation was obtained from the 

U.S. Army Dugway Proving Ground (DPG) Life Science Division, in Dugway Utah. The preparation 

procedure is reported in Brown et al. [5]. Briefly, after 80 – 90 percent sporulation, the suspension was 

centrifuged to generate a preparation of approximately 20 percent solids. A preparation resulting in a 

powdered matrix containing approximately 1 x 1011 viable spores per gram was prepared by dry blending 

and jet milling the dried spores with fumed silica particles (Deguss, Frankfurt am Main, Germany). The 

powdered preparation was loaded into MDIs by the U.S. Army Edgewood Chemical Biological Center 

(ECBC) or by ARCADIS according to a proprietary protocol. Control checks for each MDI were included in 

the batches of coupons contaminated with a single MDI.

2.1.3 Coupon Inoculation

Coupons were inoculated with spores of B. atrophaeus from an MDI using the procedure described by 

Calfee et al. [6] and detailed in MOPs 3161-LD and 3161-HD for low dose (≤2 x 104

) and high dose (>2 x 

104

) concentrations. Briefly, each coupon was inoculated independently by being placed into a separate 

aerosol dosing apparatus (ADA) designed to fit one 35.6 cm x 35.6 cm (14” by 14”) coupon of any 

thickness. In accordance with MOP 3161-LD or -HD, the MDI was discharged into the ADA a single time 

for most concentrations, but three discharges were administered when the 2 x 102 colony forming units 

(CFU) per dose MDI was used, for a total dose of ~6 x 102 CFU per coupon for these inoculations. The 

spores were allowed to settle onto the coupon surfaces for a minimum period of 18 h. When porous 

coupons were used (HVAC filters), coupons were placed on double-coated carpet tape (Polyken Model 

105C) to prevent re-entrainment of spores during handling and sampling. After the minimum 18-h period, 

the coupons were then removed from the ADA and sampled. The ADAs were removed from only those 

coupons required for a single sample at a time (some samples were comprised of three coupons, others 

a single coupon). After use, the coupons were placed in a bin of soapy water before disposal. The 

handling of the contaminated coupons was done in a way to minimize or control spore dispersal. One 

person was tasked with removing the clamps holding the ADA to the coupon and the removal of the ADA 

and gasket from the coupon. A second person, wearing new gloves for each coupon, was then tasked 

with placing the sampling template atop the coupon. A third person executed the actual sample collection 

procedure (e.g., vacuum-device usage, wipe-based sample collection, etc). The first person then 

removed the coupon for neutralization. All personnel conducted the same job throughout the entirety of a 

test operation. 
7 

The MDIs are claimed to provide 200 discharges per MDI. The number of discharges per MDI was 

tracked so that use did not exceed this value. For any inoculation event for all three materials, a new MDI

had to be used to avoid exceeding 200 discharges per MDI. Additionally, in accordance with MOP 3161-

LD and -HD, the weight of each MDI was determined after completion of the inoculation of each coupon. 

For quality control of the MDIs, on each day of testing at least three reference stainless steel inoculation 

control coupons were inoculated simultaneously and interspersed with test coupons. The reference 

coupons were inoculated as the first, middle, and last coupons within a single group of coupons, all

inoculated by the same MDI within a single test. These inoculation control (reference) coupons were a 

stainless steel coupon (35.6 cm by 35.6 cm) inoculated in accordance with MOP 3161-LD or -HD,

sampled and analyzed in accordance with Section 2.3.2 or Section 2.3.3. If the results from the 

inoculation controls were outside the acceptance criteria (Table 4-2), the results were discussed 

immediately to determine the corrective action.

A log was maintained for each set of coupons that was dosed via the method of MOP 3161-LD or -HD. 

Each record contained the unique coupon identifier, the MDI unique identifier, the date, the operator, the 

weight of the MDI before dissemination into the coupon dosing device, the weight of the MDI after 

dissemination, and the difference between these two weights. The coupon codes were pre-printed on the 

log sheet prior to the start of coupon inoculation (dosing).

A second method was used to inoculate HVAC filters under air flow (~1000 cubic feet per minute, CFM). 

To provide the flow, the back of the filter was connected to a blower from a high-volume air sampler

(Thermo Scientific High Volume Air Sampler VFC-PM10.for FRM RFPS-1287-063 

(https://www.thermo.com/eThermo/CMA/PDFs/Various/File_52267.pdf)). The outlet of the blower was 

directed to the intake of the H130 enclosure exhaust system to control contamination from breakthrough. 

On top of the filter was an ADA assembled exactly as described in MOP 3161-LD or -HD. Positioned 

between the HVAC filter and the blower, a sterile quartz filter was used to collect any spores that escaped 

capture by the HVAC filter (see Figure 2-4). The quartz filter was installed as outlined in MOP 3168. The 

blower was operated for 15 sec, then the MDI was activated, and the blower operated for another 15 sec

before deactivation. Once the blower ceased, the filter and ADA were lifted from the blower assembly and 

placed on carpet tape as the filters inoculated using the first method. The quartz filter was collected as 

outlined in MOP 3168. The blower adapter was cleaned and decontaminated between coupons with 

DispatchTM disinfectant wipes (Clorox Corp, CA), followed by 3% sodium thiosulfate (STS) and ethanol 

wipes. This method is described in more detail in MOP 3161-F.
8 

Figure 2-4. Schematic of Deposition Apparatus for Inoculation under Flow Conditions

Additionally, after a coupon was dosed via the above procedure, the coupon was labeled with the unique 

identifier described in Section 2.5.5. The identification (ID) was written on the ADA with a Sharpie or 

equivalent permanent marker. 

35.6 cm x 35.6 

cm HVAC filter

20.3 cm x 25.4 cm 

Quartz filter

ADAdapter

35.6 cm x 35.6 cm 

ADA

Hi-Vol Blower 

assemblyFlow
9 

2.2 Test Matrix

Testing was conducted in three phases. Data from the first phase of testing were used to compare 

recoveries from the various vacuum devices. The second phase was designed to perform side-by-side 

comparisons of the two devices with the highest recoveries (from Phase 1) when used to collect spores from

HVAC filters. The third phase was designed to better understand the variability between different 

laboratories in performing the inoculation, recovery, and extraction activities. The Phase 1 test matrix is 

shown in Table 2-1.

Table 2-1. Phase 1Test Matrix

Vacuum Device Material

Target 

Inoculum 

(CFU/coupon)

Replicate Test 

Samples (Total 

number of 

coupons)

Negative Controls

Samples

Total 

Coupons

Method 1

(Vacuum sock – 

fast sampling)

Carpet 107 10 (30 coupons) 1 (3 coupons) 33

Upholstery 107 10 (30 coupons) 1 (3 coupons) 33

Concrete 107 10 (30 coupons) 1 (3 coupons) 33

Method 1

(Vacuum sock – 

slow sampling)

Carpet 107 10 (30 coupons) 1 (3 coupons) 33

Upholstery 107 10 (30 coupons) 1 (3 coupons) 33

Concrete 107 10 (30 coupons) 1 (3 coupons) 33

Method 2

(37 mm MCE)

Carpet 107 10 coupons 1 coupon 11

Upholstery 107 10 coupons 1 coupon 11

Concrete 107 10 coupons 1 coupon 11

Method 3

(Trace Evidence 

Filter)

Carpet 107 10 (30 coupons) 1 (3 coupons) 33

Upholstery 107 10 (30 coupons) 1 (3 coupons) 33

Concrete 107 10 (30 coupons) 1 (3 coupons) 33

Method 4

(37 mm PTFE)

Carpet 107 10 coupons 1 coupon 11

Upholstery 107 10 coupons 1 coupon 11

Concrete 107 10 coupons 1 coupon 11

MCE = Mixed Cellulose Ester, PTFE = Polytetrafluoroethylene

Vacuum methods applicable to large areas were challenged with three coupons per sample (Methods 1 

and 3), while methods requiring a longer sample time per area were challenged with only one coupon per 

sample (Methods 2 and 4). 

The Phase 2 test matrix is shown in Table 2-2. 
10

Table 2-2. Phase 2 Test Matrix

Test Series Surface Type Sampling 

Methods

Deposition 

Method(s) Purpose

HI1 Clean electrostatic 

HVAC filters 

Extraction Settling vs. under flow, 

108 spores/actuation 

MDI used for 

inoculation

Measure the recovery 

based on extraction 

and test the validity of 

the inoculation 

procedure

HI2 Clean electrostatic 

HVAC filters

Vacuum sock, 37 

mm MCE, and 37 

mm PTFE vs. 

extraction

Under flow, 108

spores/actuation MDI 

used for inoculation

Compare recovery of 

vacuum methods to 

extraction method

HS Clean electrostatic 

or mechanical 

HVAC filters

Vacuum sock, 37 

mm MCE, and 37 

mm PTFE vs. 

extraction

Under flow, 108

spores/actuation MDI 

used for inoculation

Compare recovery of 

vacuum methods to 

extraction method

One test from the HS series was performed twice (Test HS1 and Test HS1b). Test HS1b included PRB wipe 

sampling of stainless steel coupons in addition to sponge-wipe sampling.

The Phase 3 test matrix is shown in Table 2-3. 

Table 2-3. Phase 3 Test Matrix

Test Series

(Test 

Name)

Target 

Inoculum 

Level

(CFU/coupon)

Material 

Type

Sample 

Method Sample Type Reps Purpose

Inoculation

(INOC) 

103 Stainless 

steel

Sponge-wipe Experimental 10 To determine the 

repeatability and 

magnitude of 

inoculation from 

the MDI/ADA 

dosing method

106 Stainless 

steel

Sponge-wipe Experimental 10

107 Stainless 

steel

Sponge-wipe Experimental 10

Linearity 

Recovery

(LR) 

103 Carpet Vacuum sock 

and 37 mm 

Experimental 5 To determine 

linearity of 

recovery as a 

function of 

inoculation dose

106 Carpet Vacuum sock 

and 37 mm 

Experimental 5 

107 Carpet Vacuum sock 

and 37 mm 

Experimental 5 

Laboratory 

Variability

(LV) 

107 Carpet Vacuum sock 

and 37 mm 

Experimental 

to Laboratory 

10 

each

To determine 

variability 

between 

laboratory 

technicians 

(laboratories)

107 Carpet Vacuum sock 

and 37 mm 

MCE

Experimental 

to Laboratory 

B 

10 

each
11

Each sample analysis day required laboratory blank samples analyzed in parallel with test samples.

2.3 Sampling and Analytical Procedures

Within a single test, surface sampling was first completed for all negative control coupons before 

sampling of any inoculated coupon was performed. Surface sampling was conducted either by PRB wipe 

sampling, sponge-wipe sampling, or vacuum sampling in accordance with the protocols documented 

below. These methods encompass those currently used by the EPA and CDC during biological 

contamination sampling events.

Laboratory surfaces were covered with new bench liner each day immediately prior to testing. 

Paper sampling templates were sterilized with ethylene oxide (EtO) before each use.

Prior to the sampling event, all materials needed for sampling were prepared using aseptic techniques. 

The materials specific to each protocol are indicated in the relevant sections below. In addition, general 

sampling supplies were also needed. A sampling material bin was stocked for each sampling event. The 

bin contained enough wipe sampling and vacuum sampling kits to accommodate all required samples for 

the specific test. Additional kits of each type were also included for backup. A sample collection bin was 

used to transport samples to the NHSRC Microbiology Laboratory after collection. The exterior of the 

transport container was decontaminated by wiping all surfaces with a bleach wipe or towelette moistened 

with a 5000 ppm hypochlorite solution prior to transport from the sampling location to the NHSRC 

Microbiology Laboratory. 

2.3.1 Swab Sampling

Swab sampling was used to verify sterility of coupons prior to inoculation. One coupon per sterilization 

batch, or one coupon per 10 coupons in large sterilization batches, was swab-sampled (approx. 25 cm2

) 

according to MOP 3135 and analyzed according to MOP 6563. 

2.3.2 Polyester-Rayon Blend (PRB) Wipe Sampling

The centermost 30.5 cm x 30.5 cm area of a coupon was delineated by a sterile paper template and 

sampled with a pre-moistened PRB wipe. Wipe sampling is typically used for small sample areas (i.e., 1 

ft2

) and is effective on nonporous, smooth surfaces such as ceramics, vinyl, metals, painted surfaces, and 

plastics [7]. Wipe sampling was used for stainless steel control samples only and conducted according to 

MOP 3144. The general approach is that a moistened sterile nonwoven PRB pad is used to wipe a 

specified area to recover bacteria, viruses, and biological toxins. The protocol that was used in this 

project is described in MOP 3144 and has been adapted from that provided by Busher et al. [7], Brown et 

al. [5], and documented in the Idaho National Laboratory (INL) 2008 Evaluation Protocols . Wipe samples 

were extracted in 20 mL Phosphate Buffered Saline (PBS) with 0.05% TWEEN®

20 (PBST) according to 

MOP 6567 and subjected to serial 10-fold dilution and spread-plating onto Tryptic Soy Agar (TSA, BD, 

Franklin Lakes, NJ) according to MOP 6535a.
12

2.3.3 Sponge-wipe Sampling

The centermost 30.5 cm x 30.5 cm area of the coupon was delineated by a sterile paper template and 

sampled with a pre-moistened sponge. Sponge-wipe samples, described in MOP 3165, were collected 

using the following five patterns: (1) using the flat side of the sponge-wipe, the surface was sampled 

using horizontal S-strokes, covering the entire template area; (2) the sponge-wipe was then flipped over 

to the opposite side to sample the surface in a vertical pattern, covering the entire template area; (3) 

using the narrow edges of the sponge-wipe, the surface was sampled using the same S-strokes but 

applied diagonally across the template, (4) rotating the sponge to use the opposite side starting at the 

midway point of the coupon; and (5) the tip of the sponge-wipe was then used to sample the perimeter of 

the sampling area. The sampling method is described in detail in the study Rose et al. [8] and are 

consistent with the CDC-developed field collection procedures (www.cdc.gov/niosh/topics/emres/surfacesampling-bacillus-anthracis.html). Sponge-wipe samples were extracted in 90 mL PBST as described in 

MOP 6580 and subjected to 10-fold serial dilution and spread-plating according to MOP 6535a.

2.3.4 Vacuum Sock Sampling and Analysis

Vacuum sock sampling was conducted in accordance with MOP 3145 with the following modification: 

three coupons were vacuumed per sample for some Phase 1 tests. A single coupon was vacuumed for 

Phase 2 and 3 tests. The centermost 30.5 cm x 30.5 cm (12” x 12”) square area was vacuumed on each 

coupon. For Phase 1, vacuum sock nozzles were moved across the coupon at two speeds, approximately 

1 sec per 30.5 cm pass (fast) and approximately 3 sec per 30.5 cm pass (slow). For example, on one 

square foot of surface area (30.5 cm x 30.5 cm), sampling with the vacuum sock (slow) or vacuum sock 

(fast) required approximately 60 passes each (30 passes in one direction, then 30 additional passes in a 

direction oriented 90 degrees to the first), and approximately 240 or 90 sec, respectively. For the three 

coupon sample, the collection time required was 720 or 270 sec for the slow or fast method, respectively. 

Each speed was considered a different vacuum method. All vacuum samples were collected using an 

OmegaVac (Atrix, Int.; Burnsville, MN), which supplied airflow of approximately 2000 L min-1. This 

equipment was powered by alternating current (120 V) supplied by a wall receptacle. Phases 2 and 3 

used only the slow speed technique. Vacuum socks were extracted in 20 mL PBST according to MOP 

6572 and subjected to 10-fold serial dilution and spread-plating according to MOP 6535a. Figure 2-5 

shows an assembled vacuum sock kit with cardboard nozzle and connection tubes, and the vacuum sock 

removed from the tubes.
13

Figure 2-5. Vacuum Sock Kit and Individual Sock.

2.3.5 37 mm MCE and 37 mm PTFE Sampling

Filter samples (37 mm MCE and PTFE) were collected according to Section A of MOP 3164, based on 

methods developed jointly by EPA and CDC. In short, a vacuum pump (Vac-U-Go, SKC, Inc., Eighty 

Four, PA) at the back end of the filter pulled 20 L min-1 of air through the filter. This pump was powered by 

120 V alternating current supplied by a wall receptacle. A section of Tygon (~3 cm ) tubing was cut to an 

angle of 45° on one end, the non-angled terminus was attached to the cassette via a polyvinyl chloride 

(PVC) adapter (SKC, Inc., P/N 225-132A), and the angled end was used as a nozzle. The nozzle and 

filter were moved along the coupon at roughly ten cm sec-1 (three seconds per 30.5 cm pass) in both 

directions (i.e., horizontally and vertically). For example, on one square foot of surface area (30.5 cm x 

30.5 cm), sampling with the 37 mm cassette (MCE or PTFE) required approximately 100 total passes and 

approximately 400 sec. A single coupon was vacuumed when sampling with either of these two devices. 

The nozzle was extracted separately (described in MOP 6579), the nozzle extract was then combined

with the filter extraction vessel, and filter extraction commenced. The combined resulting extract was 

subjected to 10-fold serial dilution and spread-plating according to MOP 6535a.

Figure 2-6 shows the 37 mm cassette with nozzle and tubing.
14

Figure 2-6. Filter (37 mm) Cassette with Nozzle and Tubing

2.3.6 Trace Evidence Filter Sampling

Trace Evidence Filter (TEF) samples were collected according to Section B of MOP 3164, based on 

methods developed jointly by EPA and CDC. In short, a vacuum (Atrix OmegaVac, operating at 50% 

voltage to prevent overcoming filter housing) pulled 790 L min-1 (28 standard cubic feet per minute 

(SCFM)) of air through the filter. The nozzle and filter were moved along the coupon at roughly ten cm 

sec-1 (three seconds per 30.5 cm pass) in both directions (i.e., horizontally and vertically). On one square 

foot of surface area (30.5 cm x 30.5 cm), sampling with the vacuum TEF required approximately 30 total 

passes and approximately 120 sec for sample collection. Three coupons per sample were vacuumed 

when sampling with this device. TEF filters were extracted in 90 mL PBST as described in MOP 6582 and 

subjected to 10-fold serial dilution and spread-plating according to MOP 6535a. Figure 2-7 shows the 

capped TEF cassette.

2.3.7 Operational Assessment 

Operational data, such as time required for sample collection, ease of sample collection, ease of device 

use, malfunction of device, and ease of laboratory analytical procedures, were collected for each 

sampling method. These data were used to qualitatively compare methods based on their ease of use.
15

Figure 2-7. 3M® Trace Evidence Filter (TEF) Vacuum-based Sampling Device

2.3.8 HVAC filter extraction

Sections (930 cm2, 1 ft2) of HVAC filters were excised and further cut into half or quarter sections (15.3 

cm x 30.5 cm (6” x 12”) or 15.3 cm x 15.3 cm (6” x 6”)) and using sterile scissors, folded, and placed in a 

sterile 1 L container (one section per container). The size chosen for filter sub-sections was based upon

optimization experiments conducted with the same 1 L containers at CDC (data not shown). Excision and 

sectioning of the filter typically required 90 sec (each 30.5 cm x 30.5 cm (12” x 12”) section) once cutting 

was initiated. HVAC filters were then extracted in PBST for 30 min using an orbital shaker (300 rpm). 

Quarter-sections of the filters were extracted in 500 mL PBST, while half-sections of the filters were 

extracted in 700 mL PBST. Details on the method are provided in MOP 6593.

2.3.9 Quartz extraction

Quartz filters were extracted in 100 mL of PBST as described in MOP 6586.

2.3.10 Direct Extraction of Stainless Steel

Six 15.3 cm x 30.5 cm (4” x 6”) stainless steel coupon parts were aseptically transferred straight from the 

ADA to a sterile 10L beaker. Two sterile glass rods, bent symmetrically at a 90 degree angle, were placed 

in the bottom of the beaker, followed by one sextant of the inoculated coupon, followed by another two 

sterile glass rods. The process of stacking the glass rods and coupons continued until there were no 

more inoculated coupon pieces. Each coupon was placed in the beaker so the inoculated side was 

upright. The glass beaker was sealed (sterile aluminum foil; the same foil with which the beaker was 

autoclaved) and transported to the NHSRC Microbiology Laboratory for extraction.
16

Sterile PBST (1.5 liters) was aseptically added to the 10L beaker containing the sample and then resealed with the aluminum foil. The entire beaker was placed into an ultrasonic cleaner (Branson model 

8510, Danbury, CT) and the sample was sonicated (40 kHz) for 15 min. Immediately following sonication, 

1L of extraction liquid was removed and transferred to a 1L specimen container. The sample was then 

homogenized by manual agitation/swirling before being 10-fold serially diluted and plated according to 

MOP 6535a. 

2.4 Sampling Strategy

2.4.1 Sampling/Monitoring Points

The experimental samples are listed in Section 2.2. For each inoculation event, additional samples 

collected from stainless steel surfaces were used as control samples. These control samples included 

wipe samples (used as inoculation controls or reference coupons) and vacuum samples (to compare 

collection efficacy among vacuum sampling methods). Wipe and vacuum samples were collected by 

sampling within a 30.5 cm x 30.5cm (12” x 12”) sampling template (SKC, Inc., P/N 225-2416) centered on 

the coupons. Direct extraction techniques were also used to quantify inoculation levels on stainless steel 

and HVAC filters. Each coupon was sampled only once. 

The time required to collect and analyze vacuum samples, both singly and in the aggregate, was logged 

in the laboratory notebook. 

Table 2-4 lists the samples collected for each test.
17

Table 2-4. Frequency of Sampling Monitoring Events 

Sample Type Sample Number Purpose

Vacuum sample 3 to 10 per test condition per 

coupon material

To determine the number of viable 

spores recovered via the vacuum method

Negative coupon 

sample (vacuum 

sample of sterile 

coupon)

1 per vacuum method per test To determine extent of crosscontamination from coupon handling and 

sampling

Field blank sample (1 

minute sample of 

laboratory air)

1 per vacuum method per test To determine extent of crosscontamination from laboratory air

Positive control 

coupons sampled 

either by extraction, 

wipe, or sponge

1 set of 3 stainless steel coupons 

per sampling method inoculated 

at the beginning, middle, and end 

of test coupon inoculations

To provide the best estimate of the 

number of viable spores deposited onto 

the material test coupons

Swab samples 1 per material or equipment per 

sterilization batch

To demonstrate sterility of coupons and 

inoculation materials

Microbiology

Laboratory material 

blanks

3 per material To demonstrate sterility of extraction and 

plating materials

Table 2-5 lists the critical and noncritical measurements for each sample. 

Table 2-5. Critical and Non-Critical Measurements

Sample Type Critical Measurements Non-critical Measurement

Vacuum sample Plated volume, incubation 

temperature, extracted volume, CFU, 

collection area

Storage time, storage temperature, 

collection time, extraction time

Negative coupon sample 

(vacuum sample of sterile 

coupon)

Plated volume, incubation 

temperature, extracted volume, CFU, 

collection area

Storage time, storage temperature, 

collection time, extraction time

Field blank sample (1 minute 

sample of laboratory air)

Plated volume, incubation 

temperature, extracted volume, CFU

Storage time, storage temperature, 

collection time, extraction time

Positive control coupons 

sampled either by extraction, 

wipe, or sponge

Plated volume, incubation 

temperature, extracted volume, CFU

Storage time, storage temperature, 

collection time, extraction time

Swab samples Plated volume, incubation 

temperature, extracted volume, CFU

Storage time, storage temperature

Microbiology Laboratory 

material blanks

Plated volume, incubation 

temperature, extracted volume, CFU

Storage time, storage temperature
18

2.5 Sampling Handling and Custody

2.5.1 Preventing Cross-contamination during Coupon Preparation

Coupon preparation included the activities performed on each pre-fabricated material coupon and 

procedural blank coupon prior to the inoculation procedure. Sterilization methods depended on the 

sample type: concrete coupons were sterilized using the gravity cycle of the autoclave; carpet and 

upholstery coupons were sterilized using VHP. Swab sampling of coupons from sterilization batches 

were used to confirm sterility of the materials after sterilization. The sterilization procedure was repeated 

if results were positive for the target organism. Swabs showing foreign contamination could be cause for 

repeating the sterilization procedure or taking other corrective action. The blank coupon sampling 

occurred before sampling of any inoculated coupons.

2.5.2 Preventing Cross-contamination during Sampling

Sampling poses a significant opportunity for cross-contamination of samples. In an effort to minimize the 

potential for cross-contamination, several management controls were followed. 

• In accordance with aseptic technique, a sampling team made up of a “sampler,” a “support 

person,” and a “sample handler” was utilized. 

• The sample handler was the only person to handle deposition pyramids (ADAs) or material 

coupons during the sampling event. The support person had the responsibility of handing sterile 

templates to the sampler. 

• The sampler handled only the sampling media and the support person handled all other supplies. 

The sampler sampled the surface according to the appropriate procedure as described in Section 

2.3.

• The collection medium was then placed into a sample container that was opened, held and 

closed by the support person.

• The sealed sample was handled only by the support person.

• All of the following actions were performed only by the support person, using aseptic technique: 

o The sealed bag with the sample was placed into another sterile plastic bag that was then 

sealed; that bag was then decontaminated using a bleach wipe.

o The double-bagged sample was then placed into a third sterile bag that was sealed and 

then placed into a sterile sample container for transport.

o The exterior of the transport container was decontaminated by wiping all surfaces with a 

bleach wipe or towelette moistened with a solution of 5000 ppm hypochlorite prior to 

transport from the sampling location to the NHSRC Microbiology Laboratory. 

• After the sample was placed into the container for transport, the sample handling team placed the 

sampled coupon in soapy water for eventual disposal.

The sampling crew then changed their gloves in preparation for working with the next sample.

Additionally, and equally important, the order of sampling was as follows: (1) first field blank; (2) all blank 

coupons; (3) second field blank (when required by test plan); (4) inoculated coupons; and (5) last field 
19

blank (when required by test plan). This order ensured that test coupons were handled in an order from 

least level of contamination to the most, and field blank control samples provided evidence that samples 

were handled properly, without cross-contamination. 

2.5.3 Preventing Cross-contamination during Analysis

General aseptic laboratory technique was followed and was embedded in the standard operating 

procedures (SOPs) and MOPs used by the NHSRC Microbiology Laboratory to recover and plate 

samples. The SOPs and MOPs document the aseptic technique employed to prevent crosscontamination. Additionally, the order of analysis was (1) all blank coupons, then (2) all inoculated 

coupons. 

2.5.4 Sample Containers

For each PRB wipe sample, the primary containment was an individual sterile 50 mL conical tube. 

Conical tubes (15mL) were primary containment for swab samples. Secondary containment for swab and 

PRB wipe samples was sterile sampling bags. The sponge-wipe primary containment was the stomacher 

bag used for extraction, and secondary containment was sterile sampling bags. The primary containment 

of the vacuum sock was a sterile sampling bag. A four inch cable tie was also used to close the open end 

of the newly-collected vacuum sock sample. The secondary containment of each vacuum method

sample was separate sterile sampling bags. All biological samples from a single test were then placed in 

a sterilized container. After samples were placed in the container for storage and transport to the NHSRC 

Microbiology Laboratory, the container was wiped with a towelette saturated with a ≥5000 ppm 

hypochlorite solution. A single container was used for storage of materials during sampling and for 

transport of samples to the NHSRC Microbiology Laboratory. 

2.5.5 Sample Identification

Each coupon or sample was identified by a unique sample ID. The sampling team maintained an explicit 

laboratory log which included records of each unique sample ID and its associated test number, inoculum 

level, sampling method, and the date sampled. Each coupon was marked with only the material 

descriptor and unique code number. Sample IDs included descriptors, where necessary, for project 

number (WA 10), test ID, coupon material type, vacuum or other sample type, inoculation type, sample 

purpose (test, control, field blank, etc.) and replicate number. The sample codes eased written 

identification. A typical sample ID was 10-HS1-F-V-4, which identified the sample as from WA 10, Test 

HS1, inoculated under Flow, sampled by Vacuum sock, replicate 4. Once samples were transferred to the 

NHSRC Microbiology Laboratory for microbiological analysis, each sample (plate) was additionally 

identified by replicate number and dilution. The NHSRC Microbiology Laboratory also included on each 

plate the date it was placed in the incubator.

The samples from blank coupons had a two-letter material code, with the first letter being “X”.

The sequence number was added to the test number to distinguish control samples in the case where all 

materials were not inoculated at the same time for a single vacuum method. For instance, a stainless 

steel PRB wipe sample control coupon for Test 1 for carpet and upholstery may be labeled 10-1.1-S-W-1, 

while for the concrete, inoculated on a different day, the sample would be labeled 10-1.2-S-W-1.
20

2.5.6 Information Recorded by Field Personnel

The sampling team members’ names, date, run number, and all sample codes with corresponding 

coupon codes were recorded in the laboratory notebook, along with sample times and durations. Any 

deviations from sampling protocols were documented in the laboratory notebook, along with any 

observations. 

Digital video was collected during sampling of each material using each vacuum method in order to 

document the process. 

2.5.7 Sample Preservation

Following transfer to the NHSRC Microbiology Laboratory, all samples were stored at 4 ± 2 °C until 

analyzed. All samples were allowed to equilibrate at room temperature for one hour prior to analysis. 

2.5.8 Sample Holding Times

After sample collection for a single test was complete, all biological samples were transported to the 

NHSRC Microbiology Laboratory immediately, with appropriate chain of custody (COC) form(s). Liquid 

samples were stored no longer than 24 h prior to analysis. Samples of other matrices were stored no 

longer than five days before the primary analysis. Typical hold times, prior to analyses, for most 

biological samples was ≤ two days.

2.5.9 Sample Custody

Careful coordination with the NHSRC Microbiology Laboratory was required to achieve successful 

transfer of uncompromised samples in a timely manner for analysis. Test schedules were confirmed with 

the Microbiology Laboratory prior to the start of each test. To ensure the integrity of samples and to 

maintain a timely and traceable transfer of samples, an established and proven chain of custody or 

possession is mandatory. Accurate records were maintained whenever samples were created, 

transferred, stored, analyzed, or destroyed. The primary objective of these procedures was to create an 

accurate written record that could be used to trace the possession of the sample from the moment of its 

creation through the reporting of the results. A sample was in custody in any one of the following states:

• In actual physical possession 

• In view, after being in physical possession

• In physical possession and locked up so that no one could tamper with it

• In a secured area, restricted except to authorized personnel

• In transit. 

Laboratory test team members received copies of the test plans prior to each test. Pre-study briefings 

were held to apprise all participants of the objectives, test protocols, and COC procedures to be followed. 

These protocols were required to be consistent with any protocols established by EPA. In the transfer of 

custody, each custodian signed, recorded, and dated the transfer on the COC. Sample transfer could be 

on a sample-by-sample basis or on a bulk basis. The following protocol was followed for all samples as 

they were collected and prepared for distribution:

• A COC record accompanied the samples. When turning over possession of samples, the transferor 

and recipient signed, dated, and noted the time on the record sheet. This record sheet allowed 
21

transfer of custody of a group of samples from the sample collection laboratory to the NHSRC 

Microbiology Laboratory. 

• If the custodian had not been assigned, the laboratory operator had the responsibility of packaging 

the samples for transport. Samples were carefully packed and hand-carried between on-site 

laboratories. The COC record showing the identity of the contents accompanied all packages. 

2.5.10 Sample Archiving

All samples and diluted samples were archived for a minimum of two weeks following completion of 

analysis. This time allowed for review of the data to determine if any re-plating of selected samples was 

required. Samples were archived by maintaining the primary extract at 4 ± 2 °C in a sealed extraction 

vessel. 

2.6 Statistical Analysis Methods

Relative recovery data were analyzed using a one-way analysis of variance (ANOVA). A three-way 

ANOVA model with full interactions between Device, Material, and Experiment/Control was used to 

analyze raw recovery data. For the three-way ANOVA, mean log10 reduction was then computed as a 

linear contrast of model coefficients to assess recovery rate. Mean differences in log10 reduction between 

devices were also computed as linear contrasts. Statistical significance (p ≤ 0.05) of differences between 

devices was then assessed via t-tests on the contrasts, and two sets of p-values were computed. First, pvalues were computed without adjusting for multiple comparisons. Subsequently, p-values which 

accounted for multiple comparisons were computed, based on the multivariate t-distribution of the test 

statistics. 

Both statistical methods were deemed valid by a statistical contractor (Neptune, Inc.). The advantage of 

the one-way ANOVA is that it compared relative recovery data that were normalized across experiments 

(normalized by PRB wipe recoveries). The disadvantage is that one-way ANOVA is not robust with 

regards to multiple sources of variation (i.e., variation in treatment groups and variation in control 

recoveries). The three-way ANOVA compared recoveries of the sampling methods using raw recovery 

values, and included recovery data from PRB wipe samples as a treatment group. While this method 

accounted for multiple sources of variation and interaction, the method, as used, was less robust for 

direct comparisons of sampling methods across tests. 

The results of both statistical approaches are presented and discussed.
22

3 Results and Discussion 

3.1 Phase 1

The results of the vacuum method scoping tests comparing recovery of five techniques from three 

materials are summarized in Table 3-1 and Figure 3-1.

Table 3-1. Recovery from Materials for each Vacuum Method

Mean CFU/cm2 (n = 10) (Standard Deviation in parentheses)

Material Vacuum Sock 

– Fast Speed

Vacuum Sock 

– Slow Speed 37 mm MCE TEF 37 mm PTFE

Carpet

2.01 x 104

(9.33 x 103

) 

2.66 x 104

(8.49 x 103

) 

2.83 x 104

(1.19 x 104

) 

1.28 x 104

(2.21 x 103

) 

2.18 x 104

(1.05 x 104

) 

Concrete

1.44 x 104

(3.55 x 103

) 

3.18 x 104

(5.55 x 103

) 

7.41 x 104

(3.88 x 104

) 

3.47 x 104

(1.06 x 104

) 

2.57 x 104

(1.49 x 104

) 

Upholstery

1.86 x 104

(4.16 x 103

) 

7.29 x 103

(2.73 x 103

) 

2.08 x 104

(5.64 x 103

) 

3.76 x 103

(3.78 x 103

) 

1.39 x 104

(8.60 x 103

) 

Figure 3-1. Recovery from Materials for each Vacuum Method. Data are plotted on a log-scale, as 

mean ± standard deviation.

These data are standardized for number of coupons sampled, with three coupons per sample for all 

methods except the 37 mm MCE and PTFE methods, which used only one coupon. Even without this 

standardization, the 37 mm MCE vacuum method shows higher recoveries from concrete (i.e., more 

1.00E+03

1.00E+04

1.00E+05

1.00E+06

Carpet Concrete Upholstery

Recovery (CFU cm-2)

vacuum sock - fast

vacuum sock - slow

37 mm MCE

TEF

37 mm PTFE
23

spores recovered by this method even though surface area sampled was one-third that of the vacuum 

sock and TEF methods).

To account for differing inoculation levels achieved across numerous test days and MDIs, recovery was 

further standardized by normalizing vacuum recoveries to PRB wipe samples collected from stainless 

steel coupons inoculated by the same MDI and collected on the same test day as the test samples. 

Figure 3-2 and Table 3-2 summarize these results. Interestingly, the 37 mm MCE method demonstrated 

higher recovery on concrete than the wipe-based method on stainless steel, resulting in a relative

recovery ≥100%. One explanation is that the 37 mm MCE device efficiently collected fine dust or debris 

particles from the concrete surface. Spores bound to these debris particles may have been collected 

more efficiently as a consequence, because the ease of particle resuspension from surfaces increases 

proportionately with cross-sectional area (i.e., larger particles are more easily resuspended than smaller 

particles). 

Figure 3-2. Relative Recovery – Data presented as mean relative recovery (RR). RR calculated as 

percent of wipe recovery from stainless steel surface. 

Table 3-2. Relative Recoveries from all Devices and Material Surface Types

Average CFU/cm2 vacuum method / Average CFU/cm2 stainless steel wipe 

Vacuum Sock 

– Fast Speed

Vacuum Sock 

– Slow Speed 37 mm MCE TEF 37 mm PTFE

Concrete 30% 26% 124% 41% 24% 

Upholstery 23% 11% 35% 3% 13%

Carpet 39% 64% 47% 12% 20% 

0%

20%

40%

60%

80%

100%

120%

140%

Concrete Upholstery Carpet

Relative Recovery (%)

Vacuum sock - fast

Vacuum sock -slow

37 mm MCE

TEF

37 mm PTFE
24

The null hypothesis that recoveries obtained during slow operation of the vacuum sock nozzle were not 

significantly different (p > 0.05) from the recoveries obtained during fast operation was not disproven by 

these tests. Recovery using the slow vacuum sock method was higher from carpet coupons but lower 

(than the fast vacuum sock method) from upholstery and concrete coupons, possibly due to the rapid 

motion of the nozzle agitating the coupon surface. Operation of the vacuum sock nozzle is expected to be 

highly variable between individual operators. Based on these data, operational variability may not greatly 

affect recoveries. 

Preliminary research (based upon direct spiking, conducted at the CDC) suggested that recovery from 

PTFE was higher than recovery from MCE for the 37 mm filter media. The current data suggest that 

overall recovery (collection from the surface, retention on the filter, then extraction from the filter) with 

MCE filters was higher. The TEF vacuum method performed poorly for upholstery and carpet coupons, 

compared to the other methods. The sock (slow) method performed poorly on upholstery.

3.1.1 Results of Statistical Analyses

For carpet, the slow vacuum sock method has the highest recovery rate (one-way ANOVA, p ≤ 0.001). 

When analyzed by three-way ANOVA without correcting for multiple comparisons, the vacuum sock had 

significantly higher recoveries than the 37mm PTFE (p = 0.004) and TEF (p = 0.003) methods. On 

carpet, none of the method comparisons were statistically significant after correcting for multiple 

comparisons (three-way ANOVA, p > 0.05). 

For concrete, the 37mm MCE method demonstrated the highest recovery (one-way ANOVA, p ≤ 0.001). 

Contrasts of the methods were also significant when analyzed by three-way ANOVA, before adjusting for 

multiple comparisons (all p ≤ 0.02). After adjusting for multiple comparisons, the 37mm MCE recoveries 

were statistically significant only when contrasted with recoveries of the 37 mm PTFE method (p = 0.03). 

For upholstery, the 37 mm MCE method demonstrated the highest recovery (one-way ANOVA, p ≤ 

0.001). When analyzed by three-way ANOVA, the recoveries using the 37 mm MCE method were

significantly higher than those of the 37 mm PTFE, TEF, and slow vacuum sock methods. After adjusting 

for multiple comparisons, was the 37 mm MCE method recoveries were only statistically higher than 

those using the TEF method. The TEF method demonstrated the lowest recovery for upholstery 

compared to all other devices. This result was statistically significant even after adjusting for multiple 

comparisons (three-way ANOVA, all p ≤ 0.001). 

3.1.2 Operational Parameters

3.1.2.1 Sampling Time and Ease

As discussed above, two sampling durations were used for vacuum sock sampling. Based on those data, 

there was conflicting effects of sampling time on vacuum sock samples. 

For the 37 mm devices, sampling is very time consuming due to the small surface area of the vacuum 

nozzle. A sample time of five min ft-2 limits the number of samples that can be collected per person per 

day, especially within the confines of Hazardous Materials (HAZMAT) operations. The TEF method 

sampling time was comparable to that of the vacuum sock.
25

The vacuum sock and 37 mm vacuums are both commercially-available items. The recommended 

vacuum for the TEF filters caused the filters to warp within the housing, which caused loss of sample. For 

these tests, the recommended vacuum had to be operated at 50% power to prevent filter collapse.

Defining the operation of the TEF vacuum was thus more complicated and may be more difficult to 

implement in the case of a large response.

3.1.2.2 Analysis Time and Ease

Vacuum Socks

Sample kit preparation, as well as sampling and extraction procedures, is straightforward. In the current 

study, the extraction was completed in 20 mL of PBST. Increasing the volume of PBST may be 

beneficial. With the 20 mL, the entire white (sample) portion of the sock cannot be wetted during the filter 

segmentation steps without allowing the nonsterile blue portion of the sock to touch the inside of the 

extraction cup. Durable, ergonomic scissors that are resilient to repeated sterilizations are strongly 

recommended. Custom-made racks for the orbital shaker were needed to accommodate the extraction 

vessels. Vacuum sock samples required many plating iterations (each iteration was conducted in 

triplicate) to meet QA goals for variability. This variability may be due to the presence of debris in the 

vacuum sock extraction fluid. Average processing time (all unpackaging and extraction procedures, not 

plating and analysis) for a typical sample batch (12 sock samples) was 90 min, or 7.5 min per sample.

Filter (37 mm) 

Assembly of the 37 mm sampling kits and extracting the samples was time-consuming. Transferring liquid 

from the filter cartridge can be difficult, especially when large amounts of debris are present. The nozzle 

was extracted separately and usually contained large fragments of carpet or other debris. The debris from 

the concrete, when mixed with the PBST, created a paste-like substance that was difficult to transfer via 

pipet. The carpet debris also was difficult to pipet, as it frequently obstructed pipet tips. Average 

processing time (all unpackaging and extraction procedures, not plating and analysis) for a typical sample 

batch (12 sock samples) was 120 min, or 10 min per sample.

TEF

Aseptically removing the TEF filter from the filter housing proved difficult as it was impossible to predict 

which half of the two-part housing the filter would align itself with upon opening. Rinsing the filter and 

cartridge was also time consuming and difficult, especially with samples collected from concrete. The 

dust and debris from the concrete samples generated a paste during sample extraction and was difficult 

to manipulate. Occasionally the filters punctured the stomacher bag, resulting in a compromised sample. 

Sample throughput was limited by the stomacher. Average processing time (all unpackaging and 

extraction procedures, not plating and analysis) for a typical sample batch (10 sock samples) was 180 

min, or 18 min per sample.

3.1.2.3 Cost

While the cost of all three methods was tracked, it is difficult to determine if any method was significantly 

more expensive than any another. The capital costs of each method would vary as well from laboratory to 

laboratory, depending on the availability of equipment. The following extraction equipment is 

recommended for each laboratory handling 50 samples daily:

Vacuum Sock method: Orbital shaker with appropriate rack for extraction vessels; 

37 mm filter samples: Ultrasonic water bath (sonicator); 
26

TEF filter samples: Stomacher (preferably two), bench-top centrifuge. 

3.2 Phase 2

The results from Test HI1 are summarized in Table 3-3 and suggest that MOP 3161-F, the proposed 

method for inoculating HVAC filters under flow, worked as expected and yielded repeatable results (the 

data quality objective was to achieve 1.0 x 106 recovered spores (CFU) and less than 100% RSD). During 

inoculation, an average 1.1 x 108 spores (CFU) were captured on the quartz filter having passed 

completely through the HVAC MERV 8 filter. However, the amounts recovered from HVAC filters 

inoculated under flow were similar to the HVAC filters inoculated with settling conditions only. Because 

similar recoveries were obtained from both methods and the flow-based method is likely to be more 

representative of HVAC filter contamination in the field, MOP 3161-F (flow-based method) was used on 

all subsequent HVAC filter inoculations.

Table 3-3. Average Recovery from HVAC Filter Extraction – Test HI1

Inoculation Method Mean Recovery (CFU) from HVAC Filter (n=3) RSD* (%)

Flow 3.96 x 106 54% 

Settling 9.47 x 106 39%

*RSD – Relative Standard Deviation, i.e., Coefficient of Variation as a percentage

Test HI2 was a scoping test to evaluate the feasibility of sampling flow-inoculated filters using vacuum 

methods. Table 3-4 and Figure 3-3 show the average recovery (CFU) from HVAC filters using four 

vacuum methods, as well as direct extraction. The method for direct extraction can be used on the small, 

thin filters chosen for this project but is not expected to be feasible for larger or thicker HVAC filters.

Table 3-4. Recovery from HVAC filters – Test HI2 

Vacuum Method Mean Recovery (CFU) (n=3) RSD* (%)

37 mm PTFE 1.83 x 108 15%

TEF 1.17 x 108 98%

37 mm MCE 5.41 x 107 33%

Vacuum sock (slow) 1.37 x 108 51%

Extraction 1.63 x 108 21%

*RSD – Relative Standard Deviation, i.e., Coefficient of Variation as a percentage
27

Figure 3-3. Recovery from HVAC Filters Using 5 Sampling Methods – Test HI2. Data are plotted 

on a log-scale, as mean ± standard deviation.

Results from Tests HS1 and HS2 are shown in Table 3-5. All tested methods performed well, with some 

higher variability in the TEF recovery. Though in Phase 2 the 37 mm PTFE filters provided better recovery 

than the 37 mm MCE filters (Figure 3-3), the MCE filters were chosen for side-by-side evaluation with 

vacuum socks based on the results from Phase 1 (Table 3-2). 

Table 3-5. HVAC Vacuuming Recovery – Test HS

Sample Type

Mechanical MERV 8 Filter Electrostatic MERV 8 filter

Mean

CFU/Sample

(n=5)

RSD 

(%)

Comparison

to Extraction

(T-test pvalue)

Mean

CFU/Sample

(n=5)

RSD

(%)

Comparison

to Extraction

(T-test pvalue)

Extraction 4.90 x 106 50% 4.60 x 106 84%

Vacuum sock a 1.34 x 106 43% 0.029 1.39 x 106 43% 0.26

37 mm MCE 1.58 x 106 40% 0.036 3.36 x 105 40% 0.16

a One electrostatic filter sample was lost during extraction. n = 4.

The single factor ANOVA p-value of 4.4 x 10-3 suggests a difference between all three recovery methods 

from Minimum Efficiency Reporting Value (MERV) 8 mechanical filters and the heteroscedastic Student’s 

t-test values between the vacuum methods and extraction recovery indicate neither vacuum method 

provided recoveries as high as extraction from the mechanical MERV 8 filter. Figure 3-4 shows recovery 

data from the mechanical MERV 8 filter with the three collection methods. Neither vacuum method 

provided a statistically significant difference in recovery (heteroscedastic Student’s t-test, p = 0.56). 

1.00E+06

1.00E+07

1.00E+08

1.00E+09

37 mm PTFE 37 mm MCE TEF Vacuum sock Extraction

Recovery (CFU)

Recovery from HVAC Filters
28

 

Figure 3-4. Recovery from Mechanical MERV 8 Filter – Test HS. Data are plotted on a log-scale, as 

mean ± standard deviation.

Unlike the case for mechanical type filters, the single factor ANOVA suggests no statistical difference 

between all three recovery methods from electrostatic MERV 8 filters (p = 0.16) (Figure 3-5). However, 

reanalysis using log10-transformed recovery, thus stabilizing variability, indicates there is a statistically 

significant difference between the three methods (ANOVA, p = 0.002). This difference is indicative in part 

to a high uncertainty in the number of spores deposited. Table 3-6 shows the heteroscedastic Student’s ttest between method pairs for both raw recovery and log-transformed recovery.

Table 3-6. Student’s t-test values from Electrostatic filters

Method Pairs

Heteroscedastic Student’s T-test p-values

Recovery Log10 Recovery

vacuum sock vs. 37 mm 0.036147 0.005965

vacuum sock vs. extraction 0.262986 0.187705

37 mm vs. extraction 0.157613 0.007319

Table 3-6 suggests a statistically significant difference between vacuum sock and 37 mm MCE vacuum 

methods, indicating higher recovery from vacuum sock. Table 3-6 also suggests a potential difference 

between 37 mm and extraction methods. 

1.00E+04

1.00E+05

1.00E+06

1.00E+07

Vacuum sock HVAC Extraction 37 mm MCE

Recovery (CFU)

Recovery from Mechanical MERV 8 HVAC Filters
29

Figure 3-5. Recovery from Electrostatic HVAC Filter – Test HS. Data are plotted on a log scale, as 

mean ± standard deviation.

The vacuum sock method was much easier to perform on HVAC filters than the operation of the 37 mm 

filters. However, the ease of performance may be true only for certain types of HVAC filters. Filter types 

with very deep crevices may require a sampling method such as the 37 mm filter method, which uses a 

more narrow nozzle. 

The Phase 2 test matrix also included comparisons of recovery methods from stainless steel deposition. 

While these depositions were in part included for quality control − as a measure of the stability of the 

inoculation MDI − they also provide information on spore recovery from nonporous surfaces. These data 

are not directly comparable to the HVAC data, due to the spores that passed completely through the filter 

as discussed above (Test HI1). Tables 3-7 and 3-8 summarize the recovery results from stainless steel 

surface samples, collected and analyzed during Phase 2 tests. The data are segregated by test because 

different MDIs were used for each. But taken collectively, the three recovery methods were equivalent, 

with no method producing statistically significant higher recovery.

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1.00E+08

Vacuum sock HVAC Extraction 37 mm MCE

Recovery (CFU)

Recovery from Electrostatic MERV 8 Filters
30

Table 3-7. Mean Recovery from Stainless Steel Coupons (n = 3), Phase 2 Tests 

Sampling Method Log10 CFU (± Standard Deviation)

HS2 HS1b HS1

extraction 2.2 x 107 (6.61 x 106) 1.75 x 107 (5.23 x 106) 1.11 x 108 (2.68 x 107) 

sponge-wipe 2.63 x 107 (8.8 x 106) 1.90 x 107 (6.61 x 106) 7.52 x 107 (2.10 x 107) 

PRB wipes 1.93 x 107 (3.02 x 106) 2.05 x 107 (5.11 x 106) NA

Note: Standard deviation is in parentheses.

Table 3-8. ANOVA – Comparison of the Three Stainless Steel Recovery Methods within each 

Phase 2 Test Run (n = 3)

Test Source of 

Variation df F P-value F crit

HS2 Between Groupsa 2 0.87 0.46 5.14

HS1b Between Groups 2 0.20 0.82 5.14

HS1 Between Groups 2 3.21 0.15 7.71

a Groups: Direct extraction, sponge-wipe, and PRB wipe of stainless steel. 

3.3 Phase 3

The results from the INOC test series are summarized in Table 3-9 and graphically displayed in Figure 3-

6. 

Table 3-9. Mean Recoveries from Stainless Steel using the Sponge-wipe Method - Test INOC (n = 

10)

MDI (Target Dose)

 103 104 107

Mean Recovery (CFU) 1.27 x 103 1.48 x 104 1.39 x 107

RSD 37% 31% 52%
31

Figure 3-6. Recovery from Stainless Coupons during INOC Series Tests.

These results demonstrate the ability of both the deposition method and the sponge-wipe recovery 

method over a range of spore concentrations. This approach was then applied to carpet coupons so that 

vacuum-based methods could be evaluated over a range of spore concentrations. The mean recoveries 

(CFU) for each vacuum-based method are shown in Table 3-10 and Figure 3-7. These data were 

gathered to evaluate the linearity of recoveries by each method when challenged with inocula spanning 

several orders of magnitude.

Table 3-10. Mean Recoveries (CFU/sample) from Test LR (n=5)

Mean Recoveries (CFU/sample)

Inoculation Level

Vacuum Sock

(Carpet)

37 mm MCE

(Carpet)

Extraction

(Stainless Steel)

Sponge-wipe

(Stainless Steel)

104 1.06 x 103 2.48 x 103 1.60 x 104 1.27 x 104

106 2.02 x 105 5.07 x 105 1.93 x 106 1.81 x 106

107 2.45 x 106 8.65 x 106 2.27 x 107 1.69 x 107

1.00E+02

1.00E+03

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1.00E+08

1 2 3 4 5 6 7 8 9 10

Recovery (CFU)

Replicate

Recovery from Stainless Coupons

E3

E4

E7
32

Figure 3-7. Mean CFU/Sample from Linear Recovery (LR) Test. Data are plotted on a log-scale as 

mean ± standard deviation.

Table 3-11 shows the relative recovery (percentage) based on stainless steel extraction method.

Table 3-11. Percent Relative Recovery of Vacuum Devices (Test LR) (n = 5)

Target 

Inoculum

Relative Recovery

(percent of stainless steel 

extraction recovery)

Mean Relative Recovery

(percent of stainless steel extraction 

recovery)

Vacuum Sock 37 mm MCE Vacuum Sock 37 mm MCE

104

4% 30%

7% 15%

14% 12%

3% 3%

7% 11%

6% 21%

106

11% 21%

10% 26%

7% 15%

16% 19%

9% 51%

10% 25%

107

12% 35%

11% 38%

11% 104%

16% 20%

9% 9%

6% 22%

1.00E+00

1.00E+01

1.00E+02

1.00E+03

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1.00E+08

E4 E6 E7

Recover (CFU)

Linear Recovery Test

Vacuum sock

37 mm MCE
33

The relative recovery (RR%) consists of the mean recovery (CFU) from each vacuum sample divided by 

the mean recovery (CFU) from sponge-wipe sampling stainless steel (Table 3-12). These data suggest a 

slight advantage of the 37 mm MCE vacuum method over the vacuum sock method, though the Student 

t-test returned p-values ≥ 0.05 for each comparison. 

Table 3-12. Relative Recovery of Vacuum Methods to Sponge-Wipe – Test LR (n=5). 

Target 

Inoculum

Relative Recovery

(percent of sponge-wipe recovery)

Mean Relative Recovery

(percent of sponge-wipe recovery)

Vacuum Sock 37 mm MCE Vacuum Sock 37 mm MCE

104

5% 38%

8.3% 19.6%

17% 15%

3% 4%

8% 14%

8% 27%

106

12% 22%

11.1% 28.0%

7% 16%

17% 21%

9% 54%

11% 27%

107

17% 47%

14.5% 51.2%

14% 140%1

22% 27%

12% 12%

8% 30%

1

 – Data point is within two standard deviations about the mean and the value was therefore not considered an outlier.

 ANOVA analysis of CR% over the three inoculums suggested that there was no difference in the 

recovery over the entire range for either vacuum method. (i.e., no statistical difference in CR% between 

devices at each inoculum)

Results from Test LV showing variation in laboratory and personnel for two vacuum methods are shown 

in Figure 3-8 and Table 3-13. 
34

Figure 3-8. Laboratory Variability for Two Vacuum Methods - Test LV

Table 3-13. Laboratory Variability for Two Vacuum Methods (Test LV)

Average CFU/Sample % Recovery of Extraction Controls

Sample Type n Lab 1 Lab 2 Lab 1 Lab 2

37 mm MCE 10 3.58 x 106 4.12 x 106 12.4 % 13.7 %

Vacuum Sock 10 2.01 x 106 2.47 x 106 6.8% 8.4%

Standard Deviation Standard Deviation

Sample Type n Lab 1 Lab 2 Lab 1 Lab 2

37 mm MCE 10 1.34 x 106 1.25 x 106 4.6 % 4.6%

Vacuum Sock 10 1.06 x 106 4.86 x 105 3.7 % 1.7%

CV% CV%

Sample Type n Lab 1 Lab 2 Lab 1 Lab 2

37 mm MCE 10 37% 34% 37% 34% 

Vacuum Sock 10 53% 20% 53% 20%

T-Test Between Laboratories T-Test Between Laboratories 

Sample Type n

37 mm MCE 10 0.522 0.522

Vacuum Sock 10 0.234 0.234

There was no statistically significant difference between the recoveries of the two laboratories. Pooling 

the results of the two laboratories, however, does show a significant difference between the results of the 

two vacuum methods, with a Student’s t-test value of 4.9 x 10-5. With this large number of replicates, the 

37 mm MCE vacuum method did provide better recovery than the vacuum sock method.

1.00E+05

1.10E+06

2.10E+06

3.10E+06

4.10E+06

5.10E+06

6.10E+06

7.10E+06

37 mm MCE Vacuum Sock

Recovery (CFU)

Laboratory Variability

Lab 1

Lab 2
35

4 Quality Assurance

This project was performed under an approved Category III QAPP titled Evaluation of Vacuum-based 

Sampling Devices for Collection of Biological Agent (DCMD 3.60) (October 2011).

4.1 Sampling, Monitoring, and Analysis Equipment Calibration 

There were SOPs for the maintenance and calibration of all laboratory and NHSRC Microbiology

Laboratory equipment. All equipment was verified as being certified calibrated or having the calibration 

validated by EPA’s APPCD on-site (RTP, NC) Metrology Laboratory at the time of use. Standard 

laboratory equipment such as balances, pH meters, biological safety cabinets and incubators were 

routinely monitored for proper performance. Calibration of instruments was done at the frequency shown 

in Table 4-1. Any deficiencies were noted. The instrument was adjusted to meet calibration tolerances 

and recalibrated within 24 h. If tolerances were not met after recalibration, additional corrective action was 

taken, possibly including recalibration or/and replacement of the equipment.

Table 4-1. Sampling and Monitoring Equipment Calibration Frequency

Equipment Calibration/Certification Expected Tolerance

Thermometer Compare to independent NIST thermometer (this is 

a thermometer that is recertified annually by either 

NIST or an International Organization for 

Standardization (ISO)-17025 facility) value once per 

quarter

±1°C

Stopwatch Compare against NIST Official U.S. time at 

http://nist.time.gov/timezone.cgi?Eastern/d/-5/java

once every 30 days.

±1 min/30 days

Clock Compare to office U.S. Time @ time.gov every 30 

days.

±1 min/30 days

Micropipets All micropipets will be certified as calibrated at time 

of use. Pipettes are recalibrated by gravimetric 

evaluation of pipette performance to manufacturer's 

specifications every year.

±5%

Scale Compare reading to Class S weights ±1%
36

4.2 Data Quality

The primary objective of this project was to evaluate up to four currently-available vacuum-based devices 

for biological sampling efficiency. Performance (recovery) of devices was compared to the currentlypreferred method, the "Vacuum Sock" (Midwest Filtration; Cincinnati, OH). Evaluation of operational 

parameters included time required for sample collection, the physical impact on the sampling team during 

collection, time required for sample analysis, and the cost of media and analysis equipment and supplies. 

This section discusses the Quality Assurance/Quality Control (QA/QC) checks (Section 4.3) and 

Acceptance Criteria for Critical Measurements (Section 4.4) considered critical to accomplishing the 

project objectives. 

4.3 QA/QC Checks 

Uniformity of the test materials was a critical attribute to ensuring reliable test results. Uniformity was 

maintained by obtaining a large enough quantity of material so that multiple material sections and 

coupons could be constructed with presumably uniform characteristics. Samples and test chemicals were 

maintained to ensure their integrity. Samples were stored away from standards or other samples which 

could cross-contaminate them.

Supplies and consumables were acquired from reputable sources and were National Institute of 

Standards and Technology (NIST)-traceable when possible. Supplies and consumables were examined 

for evidence of tampering or damage upon receipt and prior to use, as appropriate. Supplies and 

consumables showing evidence of tampering or damage were not used. All examinations were 

documented and supplies were appropriately labeled. Project personnel checked supplies and 

consumables prior to use to verify that they met specified task quality objectives and did not exceed 

expiration dates.

Quantitative standards do not exist for biological agents. Quantitative determinations of organisms in this 

investigation did not involve the use of analytical measurement devices. Rather, the CFU were 

enumerated manually and recorded. QC checks for critical measurements/parameters are shown in Table 

4-2. These checks also served as DQI goals. The acceptance criteria were set at the most stringent level 

that could be routinely achieved and are consistent with the DQOs described in Section 4.4. Positive 

controls and procedural blanks were included along with the test samples in the experiments so that wellcontrolled quantitative values were obtained. Verification of the sterility of samples prior to inoculation and 

other background checks were also included as part of the standard protocol. Replicate coupons were 

included for each set of test conditions. MOPs using qualified, trained and experienced personnel were 

used to ensure data collection consistency. The confirmation procedure, controls, blanks, and method 

validation efforts were the basis of support for biological investigation results. If necessary, training 

sessions were conducted by knowledgeable parties, and in-house practice runs were used to gain 

expertise and proficiency prior to initiating the research.
37

Table 4-2. QA/QC Sample Acceptance Criteria

Sample Type Purpose Frequency Acceptance Criteria Corrective 

Actions

Negative control 

(coupon without 

biological agent)

Controls for sterility 

of materials and 

methods used in 

the sampling 

procedure.

1 per test No observed CFU Identify and 

remove source 

of 

contamination.

Consult WAM.

Wipe Control

(wipe sample from 

stainless steel 

coupon 

inoculation with 

biological agent)

Verify inoculation

level on the 

coupons and to 

Demonstrate 

plate’s ability to 

support growth.

3 replicates per MDI 

use

For high inoculation, 

target loading of 107 CFU 

per sample with a 

standard deviation of 

< 0.5 log10. (5 x 106

 – 5 x 

107 CFU/sample); 

Grubbs outlier test (or 

equivalent). 

Outside target 

range: discuss 

potential impact 

on results with 

EPA WAM; 

correct loading 

procedure for 

next test and 

repeat, 

depending on 

decided impact.

Outlier: evaluate 

stability of 

MDIs.

Blank plating of 

microbiological 

supplies

Controls for sterility 

of supplies used in 

dilution plating, 

includes beads, 

PBST, dilution 

tubes, and could 

include other 

supplies if filter 

plates are needed. 

3 of each supply per 

plating event

No observed growth

following incubation

Sterilize or 

dispose of 

source of 

contamination. 

Re-plate 

samples.

Blank Tryptic Soy 

Agar Sterility 

Control

(plate incubated, 

but not inoculated)

Controls for sterility 

of plates.

Each plate is incubated 

at least 18 but fewer 

than 24 h 

No observed growth 

following incubation.

All plates are 

incubated prior 

to use; all 

contaminated 

plates will be 

discarded.

Field Blank 

Samples

(Sample matrices 

handled in 

sampling area 

without contact 

with surfaces)

The level of 

contamination 

present during 

sampling

3 per sampling event No observed growth 

following incubation

Clean up 

environment.

Sterilize 

sampling 

materials before 

use.

Tests with conditions falling outside these criteria were rejected and repeated. Decisions to accept or 

reject tests were based upon engineering judgment used to assess the likely impact of the parameter on 

the conclusions drawn from the data. For the current study, no tests required repeating. 

Potential confounding organisms were excluded or controlled by sterilization of the materials and use of 

aseptic technique, procedural blank controls, and a pure initial culture. Aseptic technique was used to 

ensure that the culture remained pure. Blank controls were set up and sampled in parallel with the 
38

contaminated materials. Infrequently, colonies were observed from negative control and procedural blank 

samples. The magnitude of the recoveries from these samples was always at least 2 orders of magnitude 

lower than test samples, and therefore did not significantly affect results. 

4.4 Acceptance Criteria for Critical Measurements

The DQOs are used to determine the CMs needed to address the stated objectives and specify tolerable 

levels of potential errors associated with simulating the prescribed decontamination environments. The 

following measurements were deemed to be critical to accomplish part or all of the project objectives:

• Enumeration of spores recovered from the surface of the coupons. 

• Total number of coupons vacuumed per sample. 

The DQIs listed in Table 4-3 are specific criteria used to quantify how well the collected data met the 

DQOs. Failure to provide a measurement method or device that meets these goals results in the rejection 

of results derived from the CM. For instance, if the plated volume of a sample is not known (i.e., is not 

100% complete), then that sample is invalid. 

Table 4-3. Critical Measurement Acceptance Criteria

Critical 

Measurement

Measurement 

Device Accuracy Precision Detection Limit Completeness

Plated Volume Pipet ±2% ±1% NA 100%

CFU/Plate Enumeration 

by visual 

inspection

±10% (between 

2 counters)

±5 1 CFU 100%

Number of 

coupons per 

plate

Enumeration 

by visual 

inspection

±1% ±0.5 1 100%

Plated volume critical measurement goals were met. All pipets are calibrated yearly by an outside 

contractor (Calibrate, Inc., Carborro, NC) and verified gravimetrically at the conclusion of testing. 

Plates were analyzed quantitatively (CFU/plate) using a visual inspection-based counting method. For 

each set of results (per test), a second enumeration was performed on 25 percent of the plates within the 

desirable range (30-300 CFU per plate). All second counts were found to be within 10 percent of the 

original count.

There are many QA/QC checks used to validate microbiological measurements. These checks include 

samples that demonstrate the ability of the NHSRC Microbiology Laboratory to culture the test organism, 

as well as to demonstrate that materials used in this effort do not themselves contain spores. The checks 

include:
39

• Field blank coupons: sterile coupons sampled at the same time as inoculated coupons. 

• Field blank sample: vacuum-based sampling media attached to vacuum device, and device was 

activated for one minute. Unfiltered laboratory air was collected, no surfaces were sampled.

• Laboratory material coupons: includes all materials, individually, used by the NHSRC Microbiology

Laboratory in sample analysis. 

• Inoculation control coupons: stainless steel coupons inoculated at beginning, middle, and end of each 

inoculation campaign. After 18 – 24 h, surfaces were sampled with PRB wipes and analyzed to 

assess the precision and accuracy of the MDI during the inoculation operation.

4.5 Data Quality Audits

This project was assigned QA Category III and did not require technical systems or performance 

evaluation audits.

4.6 QA/QC Reporting

QA/QC procedures were performed in accordance with the QAPP for this investigation.
40

5 Summary 

5.1 Phase 1

Four vacuum methods (vacuum sock, 37 mm MCE filter, 37 mm PTFE filter, and TEF), were evaluated for

sampling porous materials (concrete, carpet, and upholstery). The vacuum sock method was tested 

additionally at two sampling speeds. The vacuum sock and TEF method afforded more ease of use to 

those collecting the samples and may therefore be more desirable for larger sample areas. However, the 

37 MCE filter method demonstrated higher recoveries per unit area than did the vacuum sock device. The 

TEF and vacuum sock (slow) method performed poorly on upholstery. The TEF also performed poorly on 

carpet. There are advantages and disadvantages for each device. 

Some conclusions can be reached based on Phase 1 recovery: 

• the speed of vacuum sock sampling (rate of speed the vacuum device traversed the coupon 

surface) does not necessarily impact collection; and

• overall recovery (including collection from the surface, retention on the filter, then extraction from 

the filter) of spores using the 37 mm MCE vacuum method was often higher than the recovery 

achieved from the other devices. 

5.2 Phase 2

A method was developed which successfully deposited spores on (and through) HVAC filters under flow 

conditions. Three methods of sampling HVAC filters (two vacuum methods and one extractive) performed 

reliably on mechanical and electrostatic filters. The data suggest extractive methods may be more 

efficient than vacuum-based recovery methods, depending on filter type. Vacuum sock sampling provided 

higher recovery from mechanical filters, but both sock and 37 mm cassettes performed similarly when 

sampling from electrostatic filters. Vacuum-based methods may be more applicable to HVAC filter media 

that are not easily sectioned. 

Phase 2 data also suggested there was no statistical difference between extraction, sponge-wipes, or 

PRB wipes for recovery from stainless steel coupons.

5.3 Phase 3

The MDI inoculation method and the sponge-wipe recovery method were demonstrated to be effective 

over a broad range of concentrations, from 1 to 1 × 104 CFU/cm2 (1 x 103 to 1x107 CFU/sample). Two 

vacuum methods were also demonstrated effective over the same range. Compared to sponge-wipes 

(from stainless steel), the vacuum sock method demonstrated recoveries between 3% and 22%, while the 

37 mm MCE filter method demonstrated recoveries between 4% and 140% from carpet samples. The 

extraction procedures for both vacuum methods were evaluated by two independent laboratory 

technicians with no statistical difference in the number of spores recovered. 

5.4 Lessons Learned and Application of Vacuum-based Methods to Field Use

Based on the data generated during this study, the 37 mm MCE and the vacuum sock sampling methods 

were most efficient among the combination of materials and vacuum-based methods evaluated. When 
41

looking at these methods from an operational perspective, the vacuum sock and trace evidence filter 

methods offer advantages in that they allow a greater amount of surface area to be sampled in a given

amount of time. Increasing the amount of area sampled increases the representativeness of the sample. 

Collecting samples more rapidly decreases the sampler’s time down range (in the contaminated area), 

resulting in decreasing health and safety-related risks. However, each method has a unique set of 

advantages and disadvantages. Table 5-1 below summarizes these advantages and disadvantages.

Table 5-1. Summary of Advantages and Disadvantages of Each Vacuum-Based Sampling Method. 

37 mm (MCE or PTFE) Vacuum Sock Trace Evidence Filter

Advantages Disadvantages Advantages Disadvantages Advantages Disadvantages

High collection 

efficiency for 

some surfaces 

(i.e., concrete

and upholstery)

AC power required 

for the Vac-U-Go 

SKC sampling pump

(SKC, Inc.) 

Larger surface 

area collected

AC power required; 

however, newer Omega 

vacuum (Global 

Industrial, Port 

Washington, NY) comes 

with battery option

Larger surface 

area collected

AC power required; 

however, newer 

Omega vacuum 

comes with battery 

option

Sample is 

easily contained 

and packaged 

following 

collection 

Long sample 

collection time, >5 

min per 1 ft2

Short sample 

collection time: 

1.5 to 4 min per 1 

ft2

Heavy equipment, 

cumbersome in the field

Short sample 

collection time: 

2 min per 1 ft2

Volumetric flow 

above 790 L min-1

will cause filter to 

unseat from base,

allowing air to 

bypass filter

Relatively 

simple 

extraction 

procedure

Increases health and 

safety concerns due 

to longer sampling 

duration required for 

a given number of 

samples

Vacuum housing 

can be

decontaminated 

by wiping

Poor quality control in 

device construction 

(some have visible holes 

in the seams)

Vacuum 

housing can be 

decontaminate

d by wiping

Expensive (~$20 

per sample device)

Shortest time 

requirement of 

the methods 

evaluated for 

laboratory 

extraction

Sample is 

easily 

contained and 

packaged 

following 

sample 

collection

Longest time 

requirement of 

methods evaluated 

for laboratory 

extraction

In summary, all three methods are available and could be utilized during a large-scale event. The results 

of the current study suggest the 37 mm and vacuum sock methods perform well and with less variability 

than the TEF. When selecting a sampling method for nonporous surfaces, the method should be based 

on sampling efficiency, portability, sampling time, surface area sampled, and health and safety of 

sampling personnel.
42

5.5 Future Research

Future research efforts may focus on improving the 37 mm filter cassette sampling method. These 

improvements could involve increasing the sampling speed of the 37 mm MCE collection to determine the 

effect of sampling speed. Alternatively, the width of the 37 mm nozzle could be increased while 

maintaining the cross-sectional area of the nozzle. Further testing would need to be conducted to 

determine the effect of this nozzle alteration.

More testing should be conducted on a variety of HVAC filters to vet the methods more thoroughly. In 

addition, further study is needed to investigate the effect of surface grime or conditions on recovery. 
43

References

1. Brown, G.S., R.G. Betty, J.E. Brockmann, D.A. Lucero, C.A. Souza, K.S. Walsh, R.M. Boucher, M.S. Tezak, 

and M.C. Wilson, Evaluation of vacuum filter sock surface sample collection method for Bacillus spores from 

porous and non-porous surfaces. Journal of Environmental Monitoring, 2007. 9(7): p. 666-671.

2. Frawley, D.A., M.N. Samaan, R.L. Bull, J.M. Robertson, A.J. Mateczun, and P.C.B. Turnbull, Recovery 

efficiencies of anthrax spores and ricin from nonporous or nonabsorbent and porous or absorbent surfaces 

by a variety of sampling methods. Journal of Forensic Sciences, 2008. 53(5): p. 1102-1107.

3. Estill, C.F., P.A. Baron, J.K. Beard, M.J. Hein, L.D. Larsen, L. Rose, F.W. Schaefer, 3rd, J. Noble-Wang, L. 

Hodges, H.D. Lindquist, G.J. Deye, and M.J. Arduino, Recovery efficiency and limit of detection of 

aerosolized Bacillus anthracis Sterne from environmental surface samples. Applied and Environmental 

Microbiology, 2009. 75(13): p. 4297-306.

4. Baron, P.A., C.F. Estill, J.K. Beard, M.J. Hein, and L. Larsen, Bacterial endospore inactivation caused by 

outgassing of vapourous hydrogen peroxide from polymethyl methacrylate (Plexiglas®). Letters in Applied 

Microbiology, 2007. 45(5): p. 485-490.

5. Brown, G.S., R.G. Betty, J.E. Brockmann, D.A. Lucero, C.A. Souza, K.S. Walsh, R.M. Boucher, M. Tezak, 

M.C. Wilson, and T. Rudolph, Evaluation of a wipe surface sample method for collection of bacillus spores 

from nonporous surfaces. Applied and Environmental Microbiology, 2007. 73(3): p. 706-710.

6. Calfee, M.W., S.D. Lee, and S.P. Ryan, A rapid and repeatable method to deposit bioaerosols on material 

surfaces. Journal of Microbiological Methods, 2013. 92(3): p. 375-80.

7. Busher, A., J. Noble-Wang, and L. Rose, Sampling for Biological Agents in the Environment, P. Emanual, 

Roos, J.W.,Niyogi,K., Editor. 2008, ASM Press: Washington, DC,. p. pp 95-131.

8. Rose, L.J., L. Hodges, H. O'Connell, and J. Noble-Wang, National validation study of a cellulose sponge 

wipe-processing method for use after sampling Bacillus anthracis spores from surfaces. Applied 

Environmental Microbiology, 2011. 77(23): p. 8355-9.
44

Appendix A: Miscellaneous Operating Procedures

MOP 3135 Procedure for Sample Collection using BactiSwabTM Collection and Transport Systems

MOP 3144 Procedure for Wipe Sampling of Coupons

MOP 3145 Procedure for Vacuum Sock Sampling of Large and Small Coupons

MOP 3161-F Aerosol Deposition of Spores on HVAC Filter Under Flow

MOP 3161-HD Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol 

Deposition Apparatus (ADA) – High Dosing

MOP 3161-LD Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol 

Deposition Apparatus (ADA) – Low Dosing

MOP 3164 Procedure for 37MM Cassette and Trace Evidence Filter Vacuum Sampling of Large 

and Small Coupons

MOP 3165 Sponge Sample Collection Protocol

MOP 3168 Aggressive Air Sampling (AAS) for WA 3-28: Phase 1 Sampling Approach

MOP 6535a Serial Dilution: Spread Plate Procedure to Quantify Viable Bacterial Spore

MOP 6555: Petri Dish Media Inoculation Using Beads

MOP 6562 Preparing Pre-Measured Tubes with Aliquotted Amounts of Phosphate Buffered Saline 

with Tween 20 (PBST)

MOP 6563 Swab Streak Sampling and Analysis

MOP 6565 Filtration and Plating of Bacteria from Liquid Extracts

MOP 6567 Recovery of Bacillus Spores from Wipe Samples

MOP 6570 Use of STERIS Amsco Century SV 120 Scientific Pre-vacuum Sterilizer

MOP 6572 Recovery of Spores from Vacuum Sock Samples

MOP 6579 Recovery of Bacillus Spores from 37MM Filter Cassettes

MOP 6580 Recovery of Bacillus Spores from 3M Sponge-StickTM Samples

MOP 6582 Recovery of Bacillus Spores from Trace Evidence Filters

MOP 6586 Recovery of Bacillus Spores from Quartz Filters

MOP 6593 Recovery of Spores from HVAC Filters
Appendix B: Miscellaneous Operating Procedures 

MOP 3135 Procedure for Sample Collection using BactiSwabTM Collection and Transport Systems 

MOP 3144 Procedure for Wipe Sampling of Coupons 

MOP 3145 Procedure for Vacuum Sock Sampling of Large and Small Coupons 

MOP 3161-F Aerosol Deposition of Spores on HVAC Filter Under Flow 

MOP 3161-HD Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol 

Deposition Apparatus (ADA) – High Dosing 

MOP 3161-LD Aerosol Deposition of Spores onto Material Coupon Surfaces Using The Aerosol 

Deposition Apparatus (ADA) – Low Dosing 

MOP 3164 Procedure for 37MM Cassette and Trace Evidence Filter Vacuum Sampling of Large 

and Small Coupons 

MOP 3165 Sponge Sample Collection Protocol 

MOP 3168 Aggressive Air Sampling (AAS) for WA 3-28: Phase I Sampling Approach

MOP 6535a Serial Dilution: Spread Plate Procedure to Quantify Viable Bacterial Spore 

MOP 6555: Petri Dish Media Inoculation Using Beads

MOP 6562 Preparing Pre-Measured Tubes with Aliquoted Amounts of Phosphate Buffered Saline 

with Tween 20 (PBST) 

MOP 6563 Swab Streak Sampling and Analysis 

MOP 6565 Filtration and Plating of Bacteria from Liquid Extracts 

MOP 6567 Recovery of Bacillus Spores from Wipe Samples 

MOP 6570 Use of STERIS Amsco Century SV 120 Scientific Prevacuum Sterilizer 

MOP 6572 Recovery of Spores from Vacuum Sock Samples 

MOP 6579 Recovery of Bacillus Spores from 37MM Filter Casettes 

MOP 6580 Recovery of Bacillus Spores from 3M Sponge-StickTM Samples 

MOP 6582 Recovery of Bacillus Spores from Trace Evidence Filters

MOP 6586 Recovery of Bacillus Spores from Quartz Filters 

MOP 6593 Recovery of Spores from HVAC Filters 
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Miscellaneous Operating Procedure (MOP) 3135:

Procedure for Sample Collection Using BactiSwabTM

Collection and Transport Systems

Prepared by: __________________________________________ Date: 11/15/2012

Stella McDonald, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP-3135

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MOP 3135

TITLE: Procedure for Sample Collection using BactiSwabTM Collection and Transport

Systems

SCOPE: This MOP describes the procedure for collecting swab samples for Low Tech

Decontamination Technique Testing

PURPOSE: The purpose if this MOP is to ensure all swab sampling is performed in a

consistent manner.

Equipment/Reagents

 Disposable lab coat

 Nitrile examination gloves

 P95 Respirator

 Shoe covers

 Bouffant cap

 Safety glasses

 BactiSwabTM Collection and Transport System

1.0 PROCEDURE

1. Before starting the swabbing procedure, make sure you are wearing the appropriate, projectspecific PPE (at a minimum gloves, lab coat, and safety glasses).

2. Through the sleeve, crush the BactiSwabTM ampule at midpoint.

3. Hold BactiSwabTM tip end up for at least five seconds to allow the medium to wet the swab.

4. Open the package and remove the BactiSwabTM.

5. Label the plastic tube appropriately using the following scheme:

X-Y-N where,

X is the test number,

Y is the material abbreviation, and

N is the material number
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6. Remove the cap-swab from the plastic tube.

7. Swab the surface while spinning the cap-swab between the thumb and index fingers.

Swabbing should be conducted by following the recommend guidelines for each material as

detailed in the project documentation (usually the QAPP).

8. Return cap-swab to tube.

9. Date and initial each sample tube. Enter this information into the lab notebook.

10. Complete the chain of custody form and relinquish the samples to the BioLab.
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Miscellaneous Operating Procedure (MOP) 3144:

Procedures for Wipe Sampling of Coupons

Prepared by: __________________________________________ Date: 11/15/2012

Stella McDonald, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP-3144

Revision 2

November 2012

Page 2 of 7

MOP-3144

TITLE: PROCEDURE FOR WIPE SAMPLING OF COUPONS

SCOPE: This MOP describes the procedure for wipe sampling both small and large

coupons.

PURPOSE: The purpose of this MOP is to ensure consistent and representative sampling of

such coupons.

EQUIPMENT (quantities are per sampling kit)

 Sterile sampling bag (10” x 14”) – outer bag

 Sterile sampling bag (5.5” x 9 “) – inner “sample collection sterile sampling bag”

 Two sterile 50 mL Falcon Blue-MaxTM Polypropylene Conical Tubes

 Sterile Kendall (ref. # 8402) 4-ply all-purpose sponge

 Sterile phosphate buffered saline with 0.005% TWEEN®-20, prepared according to

MOP -6562

 Pipette or other method for aseptic dispensing of 5 mL liquid

 Sterile Posi-grip© forceps

 P-95 Particulate Respirators – to prevent contamination and for respiratory protection.

(Specific projects may require additional respiratory protection and will be addressed in

the project Quality Assurance Project Plan (QAPP), e.g, SAR)

 Powder-free Nitrile gloves (support person) and Kimtech Pure G3 Sterile Nitrile gloves

(sampler)

 Dispatch®bleach wipes

1.0 PREPARATION

1. All materials needed for collection of each sample will be prepared in advance using aseptic

technique. A sample kit for a single wipe sample will be prepared as follows:

a. Two sterile sampling bags (10” x 14”, 5.5” x 9 “) and a 50 mL conical tube, capped,

will be uniquely labeled as specified in the project QAPP. These bags and conical

tube will have the same label. The 5.5” x 9” labeled sterile sampling bag will be

referred to as the sample collection sampling bag.

b. A sterile all-purpose sponge will be placed in an unlabeled sterile 50 mL conical tube

using sterile forceps and aseptic technique. The all-purpose sponge will be moistened
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by adding 2.5 mL of sterile phosphate buffered saline with 0.005% TWEEN®-20.

The tube will then be capped.

c. The labeled 50 mL conical tube (capped), the unlabeled conical tube containing the

pre-moistened all-purpose sponge, and the 5.5” x 9” labeled sampling bag will be

placed into the 10” x 14” labeled sampling bag. Hence, each labeled sampling bag

will contain a labeled 50 mL conical tube (capped), an unlabeled capped conical tube

containing a pre-moistened all-purpose sponge, and an empty labeled sampling bag.

d. Each prepared bag is one sampling kit.

2.0 SAMPLING PROCEDURE FOR SMALL 14”x14” COUPONS

1. A three person team will be used, employing aseptic technique throughout. The team will

consist of a sampler, sample handler, and support person.

2. Throughout the procedure, the support person will log anything they deem to be significant

into the laboratory notebook.

3. In general, the team works from the least contaminated sample set (i.e., control blanks)

towards the most contaminated sample set (i.e., positive controls).

4. The sampling team will each don a pair of sampling gloves (a new pair per sample, nonsterile, as they will only be handling non-sterile items); the sampler’s gloves shall be sterile

sampling gloves as they are the only member of the team in contact with the sample. All

members shall wear dust masks to further minimize potential contamination of the samples.

Depending on the situation, respiratory protection beyond a dust mask may be required to

protect the sampling team (e.g., SAR; this will be specified in the project QAPP). New

disposable lab coats are required for the sample handler when changing between different

types of materials or when direct contact between the coupon and lab coat occurs.

5. The sample handler will remove the coupon from the appropriate cabinet and place it on the

sampling area, being careful to handle the coupon only around the edges.

6. The support person will record the coupon code on the sampling log sheet.

7. The support person will remove a template from the bag and aseptically unwrap it such that

the sampler may grab it wearing sterile gloves.

8. The sampler will place the template onto the coupon surface and align it such that the edges

of the coupon are visible through the holes on the template.

9. The support person will remove a sample kit from the sampling bin and record the sample

tube number on the sampling log sheet next to the corresponding coupon code just recorded.

10. The sampler and support person will verify the sample code and ensure that the correct

coupon and location are being sampled.

11. The support person will:

a. Open the outer sampling bag touching the outside of the bag.
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b. Touching only the outside of the (10” x 14”) bag, remove and open the unlabeled

conical tube and pour the pre-moistened all-purpose sponge onto the sample or into

the sampler’s hands.

c. Discard the unlabeled conical tube.

d. Remove the sample collection sample bag (5.5” x 9”), being careful to not touch the

inside of the outer sampling bag, and open it touching only the outside.

e. Maneuver the labeled 50 mL conical tube to the end of the outer sterile sampling bag

and loosen the cap.

f. Remove the cap from 50 mL conical tube immediately preceding the introduction of

the sample into the tube.

12. The sampler will:

a. Wipe the surface of the sample horizontally using S-strokes to cover the entire sample

area of the coupon using a consistent amount of pressure.

b. Fold the all-purpose sponge concealing the exposed side and then wipe the same

surface vertically using the same technique.

c. Fold the all-purpose sponge over again and roll up the folded sponge to fit into the

conical tube.

d. Carefully place the all-purpose sponge into the 50 mL conical tube that the support

person is holding, being careful not to touch the surface of the 50 mL conical tube or

plastic sterile sampling bag.

13. The support person will then immediately close and tighten the cap to the 50 mL conical

tube and slide the tube back into the sample collection sampling bag and seal it.

14. The support person will then wipe the sample collection sampling bag with a Dispatch®

bleach wipe and place it into the outer sampling bag.

15. The support person will then seal the outer sample collection bag now containing the capped

50 mL conical tube (containing the all-purpose sponge) inside a sealed 5.5” x 9” sample

collection bag.

16. The support person will then decontaminate the outer sample bag by wiping it with a

Dispatch®bleach wipe.

17. The support person will then place the triply contained sample into the sample collection

bin.

18. All members of the sampling team will remove and discard their gloves.

19. Steps 2 – 18 will be repeated for each sample to be collected.
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3.0 SAMPLING METHOD FOR LARGE (4’x4’ or larger) COUPONS

3.1 Sample Layout

The sampling of large coupons is carried out using a sample grid to divide the large coupons into

representative sections. These sections are then numbered and selected to be sampled at different

times during the course of the experiment as a blank, a control, or an experimental group sample.

This selection grid is pre-determined and the Project Quality Assurance Project Plan (QAPP)

may overrule the template shown in Figure 1 if otherwise specified.

As in the example below, the first cell is sampled as a Blank before contamination. Starting in

cell 3, every third cell is sampled as a positive Control. This sample is to be taken postcontamination and before decontamination. Every cell directly following a Control cell is

sampled as Experimental and is taken following decontamination. The sample kit labeling will

be based on this grid and the sampling team must ensure to correctly sample the coupons based

on this template.

1

Blank

2 3

Control

4

Experimental

5

6

Control

7

Experimental

8

9

Control

10

Experimental 11 12Control

13

Experimental 14 15Control

16

Experimental

Figure 1. 4’ x 4’ Material Section Template and Sample Grid
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3.2 Sampling Procedure

1. A two-person team will be used, employing aseptic technique throughout. The team will

consist of a sampler and a support person.

2. Throughout the procedure, the support person will log anything they deem to be significant

into the laboratory notebook.

3. The sampling team will each don a pair of sampling gloves (a new pair per sample, nonsterile, as they will only be handling non-sterile items); the sampler’s gloves shall be sterile

sampling gloves as they are the only member of the team in contact with the sample. All

members shall wear dust masks to further minimize potential contamination of the samples.

Depending on the situation, respiratory protection beyond a dust mask may be required to

protect the sampling team (e.g., SAR; this will be specified in the project QAPP).

4. The support person will record the coupon code on the sampling log sheet.

5. The sampler will place the template onto the coupon surface (using clamps as necessary).

6. The support person will remove a sample kit from the sampling bin and record the sample

tube number on the sampling log sheet next to the corresponding coupon code just recorded.

7. The sampler and support person will verify the sample code and ensure that the correct

coupon and location (cell) is being sampled.

8. The support person will:

a. Open the outer sampling bag touching the outside of the bag.

b. Touching only the outside of the (10” x 14”) bag, remove and open the unlabeled

conical tube and pour the pre-moistened all-purpose sponge onto the sample or into

the sampler’s hands.

c. The unlabeled conical tube is retained for Step 9.

d. Remove the sample collection sample bag (5.5” x 9”) being careful to not touch the

inside of the outer sampling bag and open it touching only the outside.

e. Maneuver the labeled 50 mL conical tube to the end of the outer sterile sampling bag

and loosen the cap.

f. Remove the cap from 50 mL conical tube immediately preceding the introduction of

the sample into the tube.

9. The sampler will:

a. For a vertical coupon, the sampler will squeeze excess moisture from the sampling

sponge to prevent dripping down the sampling surface. The excess moisture is caught

in the unlabeled conical tube from Step 8c, and is then discarded.

b. Wipe the surface of the sample using S-strokes to cover the entire sample area of the

coupon (inside the grid) using a consistent amount of pressure.

c. Fold the all-purpose sponge concealing the exposed side and then wipe the same
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surface vertically using the same technique.

d. Fold the all-purpose sponge over again and roll up the folded sponge to fit into the

conical tube.

e. Carefully place the all-purpose sponge into the 50 mL conical tube that the support

person is holding being careful not to touch the surface of the 50 mL conical tube or

plastic sterile sampling bag.

10. The support person will then immediately close and tighten the cap to the 50 mL conical

tube and slide the tube into the sample collection sampling bag and seal it.

11. The support person will then wipe the sample collection sampling bag with a Dispatch®

bleach wipe and place it into the outer sampling bag.

12. The support person will then seal the outer sample collection bag now containing the capped

50 mL conical tube (containing the all-purpose sponge) inside a sealed 5.5” x 9” sample

collection bag.

13. The support person will then decontaminate the outer sample bag by wiping it with a

Dispatch®bleach wipe.

14. The support person will then place the triply contained sample into the sample collection

bin.

15. All members of the sampling team will remove and discard their gloves.

16. Steps 2 – 15 will be repeated for each sample to be collected.
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Miscellaneous Operating Procedure (MOP) 3145: 

Procedure for Vacuum Sock Sampling of Large and Small Coupons

Prepared by: __________________________________________ Date: 11/15/2012 

Stella McDonald, ARCADIS Work Assignment Leader 

Reviewed by: __________________________________________ Date: 11/15/2012 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 11/15/2012 

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713 
MOP-3145 

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November 2012 

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MOP-3145 

TITLE: PROCEDURE FOR VACUUM SOCK SAMPLING OF LARGE AND SMALL 

COUPONS

SCOPE: This MOP describes the procedure for vacuum sampling of porous areas. 

PURPOSE: The purpose of this MOP is to ensure consistent and representative sampling of 

such areas.

EQUIPMENT (quantities are per sampling kit)

• 1 – Gamma irradiated vacuum sock filtration kit 

• 2 - Fisherbrand bags with round wire enclosure, 5.5” x 15” (Fisher Scientific, p/n 14-955-

181) 

• 1 - Fisherbrand bag with round wire enclosure, 10” x 14” (Fisher Scientific, p/n 01-002-

53) 

• 1 – 7.5” “cable-tie” (McMaster Car, Item# 7130K59) 

• Dispatch wipes

• Fisher Scientific CLEAN_WIPES (Catalog No. 06-664-14) prepared with 400 mL of 3% 

Sodium Thiosulfate solution (90mL of 1N STS in 310mL DI water) 

• Fisher Scientific CLEAN_WIPES (Catalog No. 06-664-24) premoistened with 70% 

isopropyl alcohol 

• Permanent marker

• Nitrile gloves

• OMEGA HEPA vacuum

1.0 PREPARATION

All materials needed for each sample to be collected will be prepared in advance. A sample kit 

for a single vacuum sock sample will be prepared using the procedure in MOP-3141A, 

Procedure for Assembling Irradiated Vacuum Sock Sampling Kits. 

2.0 VACUUM SAMPLING OF SMALL (14” by 14”) COUPONS
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The following procedure will be used in this study for vacuum sock sampling of each coupon 

surface:

1. A two person team will be used, employing aseptic technique. The team will consist of a 

sampler and a support person. 

2. Both members of the sampling team will each don a pair of sampling gloves (a new pair per 

sample); the sampler’s gloves shall be sterile sampling gloves if he/she is placing a template 

onto the sample. Both members shall wear dust masks to further minimize potential 

contamination of the samples. Further respiratory protection beyond a dust mask may be 

required to protect the sampling team (e.g., SAR; this will be specified in the project QAPP).

3. The sampler will plug in the vacuum power cord and then don his/her sterile gloves. 

4. The vacuum will be powered through a foot switch to allow the sampler to turn the vacuum 

on and off. In general, care should be taken to direct the vacuum exhaust away from samples. 

5. The support person will aseptically unwrap a template (if used) from the bag and present it to 

the sampler, taking care to not touch the template. 

6. The sampler will place the template onto the coupon surface. 

7. The sampler will wipe the hose connection end (that receives the vacuum sock) first with a 

fresh dispatch wipe, followed with a fresh CLEAN-WIPE containing 3% STS, followed by a 

fresh CLEAN-WIPE containing the 70% isopropyl alcohol. 

8. The sampler will hold the vacuum nozzle for the support person to place the vacuum sock 

assembly onto the nozzle. 

9. The support person will open the sampling supply bin and remove the vacuum sock sample 

kit from the bin. 

10. The support person will record the sample collection bag ID number on the sampling log 

sheet or in the laboratory notebook. 

11. The sampler and support person will ensure that the correct sample coupon has been selected, 

referencing the coupon code on the sampling bag. 

12. The support person will record the coupon code on the sampling log sheet next to the 

corresponding vacuum sock collection bag number that was just recorded. 

13. The support person will: 

a. Open the vacuum sock sample kit outer bag and remove the unlabelled vacuum sock 

assembly bag.

b. Tear open the bag containing the vacuum sock assembly and, working from the 

outside of the bag, maneuver the assembly from the bottom to expose the cardboard 

applicator tube opening. 

c. Firmly place the vacuum sock assembly onto the nozzle of the vacuum tube, using the 

bag to handle the vacuum sock assembly, while the sampler holds the vacuum nozzle. 
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14. The sampler will:

a. Ensure that the sock is correctly placed on the nozzle and adjust, if necessary.

Care must be taken to not puncture or tear the sock. 

b. Turn on the vacuum. 

c. Vacuum “horizontally” using S-strokes to cover the entire area of the material 

surface not covered by the template, while keeping the vacuum nozzle angled so 

that the tapered opening of the vacuum sock is flush with the sample surface. 

d. Vacuum the same area “vertically” using the same technique.

e. Turn off the vacuum when sampling is completed. 

15. The sampler will remove the vacuum sock assembly from the nozzle, taking care to only 

touch the cardboard and blue sections of the sock. 

a. Remove the sock assembly from the vacuum hose. Take care not to touch the 

sock inside the tube (Figure 1a shows the sock assembly). 

b. Loosen the nozzle by pulling it free of the longer tube, while holding onto the 

blue sock filter, and lightly replacing it in the longer tube. 

c. Unfold the blue portion of the sock filter so that it is folded over the nozzle tubing 

(Figure 1b).

d. While holding both section of cardboard tubing (one in each hand), partially 

remove the angled section of tubing. 

e. Present the sock to the support person, who will cinch the sock closed using a 

plastic “cable tie” at the blue portion between the cardboard tubes (Figures 1c and 

1d). 

f. Reconnect the two cardboard tubes (Figure 1e).

g. Place nozzle end up in a labeled 5.5” x 15” bag held by the support person. 

a. b.
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c. d.

e. 

Figure 1. (a) Vacuum sock assembly, (b) blue portion of sock over nozzle tubing, (c,d) 

cinching the sock with the “cable tie”, and (e) the two cardboard tubes 

reconnected. 

16. The support person will then seal the outer sterile sampling bag and wipe it with a 

Dispatch® wipe.

17. The support person will then place this into the labeled 10” x 14” sample bag now 

containing the outer and inner bags, the inner containing the vacuum sock assembly. The 

outermost bag will then be wiped with a Dispatch® wipe.

18. The sampler will wipe down the nozzle (inside and out) and end of the tubing first with a 

Dispatch® wipe, next with a wipe pre-moistened with 3% STS, next with a wipe premoistened with 70 % ethanol. 

19. The support person will then place the triply contained sample into the sample collection 

bin. 
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20. All members of the sampling team will remove and discard their gloves.

21. Steps 2 – 20 will be repeated for each sample to be collected. 
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Miscellaneous Operating Procedure (MOP) 3161-F: 

Aerosol Deposition of Spores on HVAC Filter Under Flow 

Prepared by: _________________________________________ Date: 4/29/2013 

Matt Clayton, ARCADIS Work Assignment Leader 

Reviewed by: _________________________________________ Date: 4/29/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 4/29/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP-3161-F 

Revision 1 

April 2013 

Page 2 of 7

MOP 3161-F 

TITLE: AEROSOL DEPOSITION OF SPORES ON HVAC FILTER UNDER 

FLOW 

SCOPE: This MOP outlines the procedure for the deposition of a spore loaded aerosol onto 

an HVAC filter under high flow conditions. 

PURPOSE: To provide a procedure for the consistent deposition of spores onto HVAC filters 

under flow. 

MATERIALS 

 One 14” x 14” Aerosol Deposition Apparatus (ADA) (per inoculation) 

 One 14” x 14” gasket (per inoculation) 

 ADAdapter (see Figure 2) 

 Dispatch® wipes 

 STS Wipes 

 Isopropyl Alcohol Wipes 

 High Volume Sampler (Thermo Andersen 100042-00)

 MOP 3161 ( and associated materials) 

 MOP 3168 

 Sterile forceps – Posi-Grip® Ref # 7190 

 Quartz filter sampling kits (prepared by BioLab) 

 Sterile filter sampling bag (VHP bag sterilized with EtO) 

 Enclosure aeration duct 

 Fisherbrand ® labeling tape – Catalog # - 15-951

 Foot power switch – McMaster Item #96618 

 Sterilized Quartz filter (1 per inoculation) - WHATMAN® G653 Cat#-18209932

 Sterilized 14” x 14” HVAC filter (1 per inoculation) - Purafilter 2000® 

 Medium binder clips – Staples ® Model#-10668-CC

 Large binder clips - Staples ® Model#-10669

 Blue spring clamps – Workforce 573-682

 12” Double-sided sticky tape – Polyken 105C Carpet Tape 
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1.0 PREPARATION AND ASSEMBLY 

1. Sterilize one 14” x 14” ADA, one 14” x 14” gasket, and an ADAdapter at least 3 days prior

to allow for off-gassing. 

2. Sterilize one VHP bag per coupon using ethylene oxide (EtO) at least 3 days prior to 

testing. 

3. Set up the High Volume Sampler in the enclosure vented to the enclosure aeration system. 

NOTE: Refer to Figure 1 to ensure assembly is correct during Steps 3-8. Figure 2 shows 

the completed blower and ADA assembly.

4. The filter base on the blower assembly must be lifted above the top face of the sampler using 

angle iron (See Figure 1a) to allow the seal between the base and ADAdapter to be clamped. 

5. A two-person team is employed to aseptically install the quartz filter on top of the blower 

assembly base as in MOP 3168, except there is no need for a third person to hold the bracket 

as it is not required with this assembly (see Figure 1b). 

6. Install the ADAdapter on top of the filter ensuring contact with the gasket. It is important that 

the filter is installed such that all edges are covered by the gasket on the bottom of the 

ADAdapter to ensure all airflow passes through it. Clamp the ADAdapter to the assembly 

using small binder clips (see Figure 1c). 

7. Once the ADAdapter is securely in place, aseptically install the 14” x14” HVAC filter. 

Remove the filter from the bag using sterile gloves and place on top of the ADAdapter. 

Ensure the filter is installed in the orientation pictured in Figure 1d. 

8. Finally, Install the 14” x 14” gasket and ADA on top of the filter and secure the seal with the 

spring clamps. Ensure the ADA is equipped with 4 filters and the top port is sealed with the 

included mechanism (see Figure 1e). 

9. Before proceeding to Section 2.0, prepare adequate space for the number of filters to be 

puffed on cleaned tables. Use double-sided tape to create 16” x 16” settling areas, one for 

each puffed filter. It is necessary to clean the table surfaces with Dispatch wipes, water and 

alcohol. Do not use paper coverings due to using double-sided tape. 
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a. b.

c. d.

e.

Figure 1. (a) Blower assembly base, (b) Quartz filter installed, (c) ADAdapter installed and clamped, 

(d) HVAC filter installed, and (e) Completed assembly. 
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Figure 2. Blower and ADA assembly

 

ADAdapter

14x14 ADA

Blower Assembly 

base
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2.0 PUFFING PROCEDURE 

1. Determine the weight of the MDI canister using a balance. Record the MDI ID number and 

the weight (to the nearest 0.01g) in lab notebook. In addition, keep a record of the total 

number of ‘puffs’ dispensed for each MDI canister.

NOTE: The MDI canister full is approximately 15 grams, an empty canister is approx 9.5 

grams. To ensure the canister contains adequate spore suspension for dosing, 

canisters should be retired from use when their weight falls below 10.5 grams.

2. Next, assemble the MDI and actuator by inserting the MDI into the actuator, taking care not to 

activate the MDI. 

3. Vortex the MDI/actuator assembly for 30 seconds (the MDI canister should be in direct contact 

with the vortex mixer). 

4. Holding the MDI/actuator assembly upright, in a swift, firm motion, dispense three test ‘puffs’ 

into the aerosol trap to prime the MDI. It is important to vortex the assembly 10 seconds 

before every puff (except 30 seconds prior to the initial puff of the experiment). 

5. Vortex the assembly 10 seconds, and attach it to the ADA lid by mating the ADA adaptor to 

the hole in the ADA lid. Loosen the lid screws enough to allow the lid to be slid into the 

‘open’ position. Secure the lid in the open position by tightening the lid screws. 

NOTE: The ‘open’ position is achieved when the hole in the lid aligns with the hole in the top 

of the ADA. 

6. Using the foot switch, start the blower and allow the unit to flow for 15 seconds. 

7. At 15 seconds, dispense the spores by activating the MDI with a swift, firm motion. Hold the 

MDI in the activated position for 3 seconds before releasing. Activation is best achieved by 

grasping the MDI/actuator with two hands, and using a thumb to press the bottom of the MDI 

canister. 

8. After 30 seconds total, follow the reverse order of the lid opening procedure to close the ADA 

lid, and turn off the blower using the foot switch. 

9. Determine the weight of the actuator-MDI using a balance, and record the weight in lab 

notebook. 

NOTE: If the dosing puff is faulty, return to Step 5 and attempt a second puff on the current 

filter. Do not proceed to the next filter until a ‘successful’ puff has been delivered. A 

‘successful’ puff is achieved when the weight of the actuator- MDI assembly has a 
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0.04gram-0.07gram loss. Familiarity and professional judgment will be needed to 

determine the success of a puff. 

10. Once the filter is successfully puffed, remove the ADA , gasket, and filter assembly. Clamp the 

filter to the assembly using large binder clips, and place on the double sided tape. 

11. Remove the ADAdapter and place on a clean surface. 

12. Aseptically recover the quartz filter as in MOP 3168. 

13. Clean the ADAdapter with Dispatch® wipes, STS wipes, and isopropyl alcohol wipes before 

proceeding to Step 14. 

14. For each additional filter, proceed to apparatus assembly, using a new sterile ADA and gasket 

for each inoculation (Step 5, Section 1). 

15. Repeat Steps 1.5 to 2.14 until all filters have been puffed (see test plan for number of samples). 

16. Once all coupons have been puffed, remove the MDI from the actuator and weigh. Record the 

final weight and total number of puffs. 

17. Allow spores to settle onto the coupon surface for at least 18 hours. Settling time should not 

exceed 26 hours. 

3.0 RECOVERY 

1. After 18 hours settling time, carefully remove binder clips and remove ADA and gasket from 

coupon surface, taking care not to disturb the surface of the coupon. 

2. Don sterile gloves and transfer the filter to sterilized VHP bag. Carefully tape the bag closed, 

ensuring an air tight seal. 

3. The VHP bag should be placed in tertiary containment before transport to the APPCD 

Biocontaminant Laboratory for analysis. 
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Miscellaneous Operating Procedure (MOP) 3161-HD:

Aerosol Deposition of Spores onto Material Coupon Surfaces using

the Aerosol Deposition Apparatus (ADA) – High Dosing

Prepared by: __________________________________________ Date: 11/15/2012

Stella McDonald, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 3161-HD

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November 2012

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MOP 3161-HD

TITLE: AEROSOL DEPOSITION OF SPORES ONTO MATERIAL COUPON

SURFACES USING THE AEROSOL DEPOSITION APPARATUS (ADA) –

HIGH DOSING

SCOPE: This MOP outlines the procedure for assembly and usage of the Aerosol

Deposition Apparatus (ADA).

PURPOSE: Precise and highly repeatable aerosol deposition of bacterial spores onto material

surfaces for detection, sampling, and/or decontamination studies.

Materials:

 Aerosol Deposition Apparatus (ADA) (shown in Figure 1)

 Metered Dose Inhaler (MDI) preloaded with a bacterial spore suspension of known

concentration (i.e., 1 x 109spores per puff)

 Vertical MDI Actuator (shown in Figure 2)

 Material coupon (with dimensions at least that of the ADA)

 ADA-coupon gasket (1 per ADA) (see Figure 1)

 Clamping devices (i.e., medium-size steel binder clips, C-clamps (8 per ADA))

 Vortex mixer (shown in Figure 4)

 Aerosol trap (described in Appendix A and shown in Figure 4)

 Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles)

 pH-adjusted bleach (pAB) (MOP 3128-A)

 0.22μm pore-size syringe filters (shown in Figure 1)

 PVC tubing (3/8” OD, 1/4” ID)

 Mass balance (with 0.01 gram accuracy)

 Bench liner

1.0 STERILIZATION OF MATERIALS

Prior to the start of any experiment, all components must be sterilized and stored in a sterile

environment until usage. Sterilization is not necessary for binder clips, MDI, vortex, or the

aerosol trap.
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ADAs can be sterilized by autoclave, VHP, or by wiping with pH-adjusted bleach (pAB) with

subsequent deionized (DI) water and ethanol rinse/wipes. The ADA lid should be attached and

in the closed position during the sterilization.

Figure 1. ADA apparatus

The MDI actuator, with attached MDI adaptor, can be wiped with pAB then rinsed with DI

water.

Figure 2. MDI and vertical actuator

ADA

ADA with lid in

closed position

Gasket

2.2um

syringe filter

MDI

Vertical

Actuator
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Sterilization requirements for coupons vary by material. Regardless of the sterilization method,

quality control (QC) checks (typically by collecting a swab sample per MOP 3135) should be

administered to ensure the effectiveness of the sterilization method.

Gasket sterilization may also vary by material. Care should be taken to thoroughly degas gaskets

if sterilized via fumigation.

2.0 PROCEDURE

1. Begin by donning PPE (gloves, lab coat, and protective eyewear).

2. Clean the workspace by wiping with pAB, next with DI water , and lastly with a 70-90%

solution of denatured ethanol. Alternatively, new, clean bench liner may be placed on the

work surfaces. Make sure the workspace is clean and free of debris.

3. Discard gloves and replace with fresh pair.

4. Using aseptic techniques (when possible) assemble the coupon/ADA by first placing the

sterilized material coupon onto the clean lab bench or workspace, next place the sterilized

gasket on top of the coupon, and lastly seat the ADA on the coupon + gasket. Orient each

component so that it fits squarely with the previously placed item. Take care not to touch the

inside of the ADA or the coupon surface. Secure these components by attaching mediumsize binder clips, one at each corner, and one at the midpoint of each of the four sides of the

ADA. The binders should firmly secure the coupon to the ADA, and apply sufficient

pressure to the gasket to seal the union. If material coupons are too large to use binder clips

other methods may be used to secure the coupon and gasket to the ADA (i.e., larger clamps,

weight added to the ADA, etc.). Lastly, attach 0.2 um syringe filters to each vent tube on all

ADAs (4 per ADA). Syringe filters can be attached using PVC tubing (3/8” OD, 1/4” ID).

5. Determine the weight of the MDI canister using a balance. Record the MDI ID number and

the weight (to the nearest 0.01g) in lab notebook. In addition, keep a record of the total

number of ‘puffs’ dispensed for each MDI canister.

NOTE: The MDI canister full is approximately 15 g, an empty canister is approx 9.5 g. To

ensure the canister contains adequate spore suspension for dosing, canisters should

be retired from use when their weight falls below 10.5 g.

6. Next, assemble the MDI and actuator by inserting the MDI into the actuator, taking care not

to activate the MDI.

7. Vortex the MDI/actuator assembly for 30 seconds (the MDI canister should be in direct

contact with the vortex mixer).

8. Holding the MDI/actuator assembly upright (Figure 3), with a swift, firm motion, dispense

three test ‘puffs’ into the aerosol trap to prime the MDI. It is important to vortex the
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assembly 10 seconds before every puff (the exception being 30 seconds prior to the initial

puff of the experiment, as prescribed in Step 7).

Figure 3. MDI orientation while dispensing test puffs into the aerosol trap.

9. Vortex the assembly for 10 seconds and then attach to the ADA lid by mating the ADA

adaptor to the hole in the ADA lid. Loosen the lid screws enough to allow the lid to be slid

into the ‘open’ position. Secure the lid in the open position by tightening the lid screws.

NOTE: The ‘open’ position is achieved when the hole in the lid aligns with the hole in the

top of the ADA.

10. With a swift, firm motion, dispense the spores by activating the MDI. Hold the MDI in the

activated position for 3 seconds before releasing. Activation is best achieved by grasping the

MDI/actuator with two hands, and using a thumb to press the bottom of the MDI canister.

11. Follow the reverse order of the lid opening procedure to close the ADA lid.
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12. Determine the weight of the actuator-MDI using a balance, and record the weight in lab

notebook.

NOTE: If the dosing puff is faulty, return to Step 9 and attempt a second puff on the current

coupon. Do not proceed to the next coupon until a ‘successful’ puff has been

delivered. A ‘successful’ puff is achieved when the weight of the actuator-MDI

assembly has a 0.04 g to 0.07 g loss. Familiarity and professional judgment will be

needed to determine the success of a puff.

13. Vortex the assembly for 10 seconds, then proceed to dosing the next coupon (Step 9).

14. Repeat Steps 9 through 13 until all coupons have been dosed.

15. Once all coupons have been puffed, remove the MDI from the actuator and weigh. Record

the final weight and total number of puffs.

16. Allow spores to settle onto the coupon surface for at least 18 hours. Settling time should not

exceed 26 hours.

17. Carefully remove binder clips (or other attachment device), and remove ADA and gasket

from coupon surface, taking care not to disturb the surface of the coupon.

18. Test coupon is now ready for use.

19. Decontaminate the ADA and associated components with the same procedures utilized

during the initial sterilization.
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APPENDIX A - Aerosol Trap

Purpose: This device allows test puffs of the MDI to be deployed without contamination of the

surrounding area. Spores are pulled into the trap, contained, and inactivated.

This device consists of a suction source, a trap (containing pAB), and an inlet funnel.

Aerosolized spores are pulled into the funnel, and forced into the trap. The spores are collected

and inactivated as the aerosol flows through the pAB solution. The effluent air traveling toward

the suction device is spore-free downstream of the trap. See Figure 4.

The aerosol trap should be assembled inside a biological safety cabinet (BSC) or chemical fume

hood.

Figure 4. Aerosol trap

Aerosol trap

Vortexer
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Miscellaneous Operating Procedure (MOP) 3161-LD: 

Aerosol Deposition of Spores onto Material Coupon Surfaces using 

the Aerosol Deposition Apparatus (ADA) – Low Dosing 

Prepared by: _________________________________________ Date: 4/29/2013 

Matt Clayton, ARCADIS Work Assignment Leader 

Reviewed by: _________________________________________ Date: 4/29/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 4/29/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP-3161-LD

Revision 1 

April 2013

Page 2 of 7 

MOP 3161-LD 

 

TITLE: AEROSOL DEPOSITION OF SPORES ONTO MATERIAL COUPON 

SURFACES USING THE AEROSOL DEPOSITION APPARATUS (ADA) –

LOW DOSING 

SCOPE: This MOP outlines the procedure for assembly and usage of the Aerosol 

Deposition Apparatus (ADA) for low dosing. 

PURPOSE: To provide a procedure for repeatable aerosol deposition of bacterial spores onto 

material surfaces for detection, sampling, and/or decontamination studies. 

MATERIALS

 Aerosol Deposition Apparatus (ADA) 

 Metered Dose Inhaler (MDI) preloaded with a bacterial spore suspension of known 

concentration (i.e., 5 x 106spores per puff or less, also designated as 5E6) 

 MDI Actuator with ADA adaptor (adaptor required for proper attachment of actuator to 

ADA lid). The MDI actuator label must match the concentration and organism of the 

MDI. 

 Material coupon (with dimensions at least that of the ADA) 

 ADA-coupon gasket (1 per ADA) 

 Clamping devices (i.e., medium-size steel binder clips, C-clamps (8 per ADA)) 

 Vortex mixer 

 Aerosol trap (described in Appendix A) 

 PPE (gloves, lab coat, P95 dust mask, cap or hair bonnet, safety goggles) 

 pH-adjusted bleach (pAB; prepared per MOP 3128) 

 0.22μm pore-size syringe filters 

 PVC tubing (3/8” OD, 1/4” ID)

 mass balance (with 1 mg accuracy) 

 canned air duster 
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1.0 DISINFECTION OF MATERIALS

Prior to the start of any experiments, all components must be disinfected, and stored in as close 

to a sterile environment as possible until usage. (Disinfection is not necessary for binder clips, 

MDI, vortex, and the aerosol trap). 

ADAs can be disinfected by autoclave, VHP, or pAB wiping with subsequent deionized water 

(diH2O) and ethanol rinse/wipes. The ADA lid should be attached and in the closed position 

during the disinfection. 

MDI actuator with attached MDI adaptor can be wiped with pAB then rinsed with diH2O. 

NOTE: DO NOT expose to ethanol, as ethanol will shatter the ADA adaptor. 

Disinfection requirements for coupons vary by material. Regardless of disinfection method, QC 

checks should be administered to ensure the effectiveness of the disinfection method. 

Gasket disinfection may also vary by material. 

2.0 PROCEDURE 

1. Begin by donning PPE (gloves, lab coat, and protective eyewear). 

2. Cover the tables with fresh bench liner. 

3. Discard gloves and replace with fresh pair. 

4. Using aseptic techniques (when possible) assemble the coupon/ADA by first placing the 

disinfected material coupon onto the clean lab bench or workspace. Next place the 

disinfected gasket on top of the coupon, and then seat the ADA on the coupon + gasket. 

Orient each component so that it fits squarely with the previously placed item. Take care not 

to touch the inside of the ADA or the coupon surface. Secure these components by attaching 

medium-size binder clips, one at each corner, and one at the midpoint of each of the four 

sides of the ADA. The binders should firmly secure the coupon to the ADA and apply 

sufficient pressure to the gasket to seal the union. If material coupons are too large to use 

binder clips, other methods may be used to secure the coupon and gasket to the ADA (i.e., 

larger clamps, weight added to the ADA, etc.). 

Lastly, attach 0.2 um syringe filters to each vent tube on all ADAs (4 per ADA). Syringe 

filters can be attached using PVC tubing (3/8” OD, 1/4” ID).

5. Determine the weight of the MDI canister using a balance. Record the weight (to the nearest 

mg) in lab notebook. In addition, keep a record of the total number of ‘puffs’ dispensed for 

each MDI canister. 
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NOTE: A full MDI canister is usually approximately 20 grams (although they are 

sometimes 15 grams); an empty canister is approximately 9.5 grams. To ensure 

the canister contains adequate spore suspension for dosing, canisters should be 

retired from use when their weight falls below 10.5 grams. 

6. Next, assemble the MDI and actuator (with ADA adaptor) by inserting the MDI into the 

actuator, taking care not to activate the MDI. Weigh the MDI and actuator assembly and 

record the weight (to the nearest mg) in lab notebook. 

7. Vortex the MDI/actuator assembly for 30 seconds (the MDI canister should be in direct 

contact with the vortex mixer). 

8. Holding the MDI/actuator assembly upright (MDI canister on top, MDI nozzle pointing 

down, and actuator + adaptor outlet pointing horizontal, in contact with the aerosol trap inlet 

adapter − see Figure 1), dispense three test ‘puffs’ into the aerosol trap to prime the MDI. It 

is important to vortex the assembly 10 seconds before every puff (except 30 seconds prior to 

the initial puff of the experiment, as prescribed in Step 7). Weigh the MDI/actuator assembly 

after each puff and record in the Deposition Log. A weight reduction of 0.04-0.07g will 

indicate a successful puff. 

Figure 1. MDI orientation while dispensing test puffs into the aerosol trap. 
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9. Vortex the assembly 10 seconds, and attach it to the ADA lid by mating the ADA adaptor to 

the hole in the ADA lid. Loosen the lid screws enough to allow the lid to be slid into the 

‘open’ position. (The ‘open’ position is achieved when the hole in the lid aligns with the hole 

in the top of the ADA.) Secure the lid in the open position by tightening the lid screws. 

10. With a swift firm motion, dispense the spores by activating the MDI. Hold the MDI in the 

activated position for 3 seconds before releasing. Activation is best achieved by grasping the 

MDI/actuator with two hands, and using a thumb to press the bottom of the MDI canister. 

Weigh the MDI and record in the Deposition Log assembly. 

NOTE: If the dosing puff is faulty, return to Step 8 and attempt a second puff on the 

current coupon. Do not proceed to the next coupon until a ‘successful’ puff has 

been delivered. A ‘successful’ puff is achieved when the plume can be seen 

through the clear ADA adaptor immediately following the puff. The puff results 

in audible characteristics typical of a successful puff, and the proper weight 

reduction is achieved. Familiarity and professional judgment will be needed to 

determine the success of a puff. 

11. Follow the reverse order of the lid opening procedure to close the ADA lid. Detach the 

MDI/actuator assembly from the lid. 

12. Remove the MDI from the actuator and determine the weight of the MDI using a balance, 

record the weight in lab notebook. 

13. Place the empty actuator into the aerosol trap and remove dosing residue from the inner 

surface of the actuator using the canned air duster. Take care to blow the compressed air into 

the bottom of the actuator so that the residue is blown directly into the aerosol trap. 

14. Vortex the assembly for 10 seconds, holding the assembly upright (MDI canister on top, 

MDI nozzle pointing down, and actuator + adaptor outlet pointing horizontal, towards the 

aerosol trap inlet − see Figure 1), dispense a test puff into the aerosol trap (expect a faulty 

puff, as the MDI requires recharging after a puff in the non-upright orientation). Repeat once 

more (vortex 10 seconds, puff). Expect the subsequent puff to be characteristic (visually,

audibly, and with expected weight loss) of a successful puff. If the second test puff is 

successful, proceed to dosing the next coupon (Step 9). If the second puff is faulty, repeat 

Step 13. 

15. Repeat Steps 9 through 13 until all coupons have been dosed. 

16. Allow spores to settle onto the coupon surface for at least 18 hours. Settling time should not 

exceed 26 hours. 

17. Wearing the proper PPE, carefully remove binder clips (or other attachment device), and 

remove ADA and gasket from coupon surface, taking care not to disturb the surface of the 

coupon. 
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18. Test coupon is now ready for use. 

19. Decontaminate the ADA and associated components with the same procedures utilized 

during the initial disinfection. 
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APPENDIX A – AEROSOL TRAP 

Purpose: This device allows test puffs of the MDI to be deployed without contamination of the 

surrounding area. Spores are pulled into the trap, contained, and inactivated. 

This device consists of a suction source, a trap (containing pAB), and an inlet funnel. 

Aerosolized spores are pulled into the funnel and forced into the trap. The spores are collected 

and inactivated as the aerosol flows through the pAB solution. The effluent air traveling toward 

the suction device is spore-free downstream of the trap. See Figure 2. 

The aerosol trap should be assembled inside a biological safety cabinet (BSC) or chemical fume 

hood. 

 

Figure 2. Schematic drawing of Aerosol Trap. 

Suction

Device

pH-adjusted bleach
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Miscellaneous Operating Procedure (MOP) 3164: 

Procedure for 37MM Cassette and Trace Evidence Filter Vacuum 

Sampling of Large and Small Coupons 

Prepared by: _________________________________________ Date: 4/29/2013 

Matt Clayton, ARCADIS Work Assignment Leader 

Reviewed by: _________________________________________ Date: 4/29/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 4/29/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
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TITLE: PROCEDURE FOR 37MM CASSETTE AND TRACE EVIDENCE FILTER 

VACUUM SAMPLING OF LARGE AND SMALL COUPONS

SCOPE: This MOP describes the procedure for vacuum sampling of porous areas. 

PURPOSE: The purpose of this MOP is to ensure consistent and representative sampling of 

such areas.

A: 37 mm filter

EQUIPMENT (starred* quantities are per sampling kit)

• *1 – 37 mm Filter cassette loaded with desired filter 

- 0.3 μm pore size PTFE membrane (SKC 225-1723) 

- 0.8 μm pore size MCE membrane (SKC 225-3-01) 

• Vac-U-Go pump (SKC 228-9605)

• Rotameter (SKC 320-100) 

• Tygon tubing, 1/4 in ID, 7/16 in OD, 50ft (SKC 225-1345) 

• *Two PVC adapters, 250/pkg (SKC 225-132A) 

• *One BD Falcon Sterile 15 mL Polyprolene Conical Tubes (fishersci.com, P/N 225-

352196) 

• *Two 5.5” x 9” 3.5 mil sterile sampling bags with flat-wire closures (fishersci.com, 

Item# 14-955-187) 

• *10” x 14” overpack sample bag with round wire enclosure (fishersci.com, Item# 01-002-

53) 

• 12” x 12” Template

• Permanent marker

• Nitrile gloves

• Timer

1.0 PREPARATION 

All materials needed for each sample to be collected will be prepared in advance. A sample kit 

for a single cassette vacuum sample will be prepared using the following procedure. 

1) Don nitrile gloves. 

2) In laboratory, assemble nozzles:
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a) Cut the sampling nozzle from tygon tubing as follows: Using scissors, cut a section of 

tygon tubing 1-inch long, then cut one end of the tubing at approximately a 45-degree 

angle. 

b) Place the nozzle onto a PVC adapter. 

c) Prepare 25% more nozzles than required by the testing protocol. 

d) Sterilize the nozzles with a 15 minute gravity cycle of the autoclave. 

3) In laboratory, assemble cassette kit: 

a) Label the 37 mm cassette with a unique sample ID. 

b) Label the 15 mL conical tube with the same unique sample ID. 

c) Aseptically remove the cassette plugs and place a PVC adaptor onto each end of the 

cassette. Save the removed plugs for step f) below. 

d) Cut a 20 cm long piece of tubing with the scissors. 

e) Place the 20 cm tubing onto the downstream end of the cassette.. 

f) Place the sampling nozzle (1-inch section of tubing) onto the upstream end of the 

cassette with the angled side furthest from the cassette. 

g) Place the cassette with PVC adaptors, 20 cm tubing, and nozzle into a 5.5” x 9” 3.5

mil bag, making sure the end of the cassette that attaches to the vacuum tubing is 

closest to the bag opening. (see Figure 1) 

h) Place the red plugs removed from the cassette in step a) above into the same 5.5” x 9” 

bag. 

i) Place the 15 mL conical tube and the 5.5” x 9” bag with 37mm cassette assembly and 

plugs into the 10” x 14” overpack bag. 

j) Place an additional sterile small bag into the overpack bag. 

4) Label the 10” x 14” bag with the sample ID. 

5) Store sample kits in a clean dry location. 
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Figure 1. 37mm Cassette Vacuum Inner Bag 

2.0 VACUUM SAMPLING OF SMALL (12” by 12”) COUPONS 

The following procedure will be used in this study for 37mm vacuum sampling of each coupon 

surface: 

1. A two-person team will be used, employing aseptic technique. The team will consist of a 

sampler and a support person. In times when a third person (sample handler) is present, he or 

she will act as an assistant to the support person for data recording. 

2. The sampler will plug in the Vac-U-Go pump power cord, and attach the calibrated 

rotameter to the pump. Turn on the pump and adjust the pump valve until the flow rate is 20 

± 2 LPM. Record the flow rate in the project notebook. The sampler will then don his/her 

sterile gloves. 

3. Both members of the sampling team will each don a pair of sampling gloves (a new pair per 

sample); the sampler’s gloves shall be sterile sampling gloves as they are the only member of 

the team in contact with the sample. Both members shall wear dust masks to further 

minimize potential contamination of the samples. Further respiratory protection beyond a 

dust mask may be required to protect the sampling team (e.g., SAR; this will be specified in 

the project QAPP). 

4. The Vac-U-Go pump will be maintained on a rolling cart for easy movement into place. 

5. The support person will aseptically unwrap a template from the bag and present it to the 

sampler, taking care to not touch the template. 

6. The sampler will place the template onto the coupon surface. 

7. The support person will open the sampling supply bin and remove the 37mm cassette sample 

kit from the bin. 
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8. The support person or assistant will record the sample collection bag ID number on the 

sampling log sheet or in the laboratory notebook. 

9. The sampler and support person will ensure that the correct sample coupon has been selected, 

referencing the coupon code on the sampling bag. 

10. The support person or assistant will record the coupon code on the sampling log sheet next to 

the corresponding 37mm cassette collection bag number that was just recorded. 

11. The support person will: 

a. Open the 37mm cassette sample kit outer bag and remove the unlabeled 37 mm 

cassette assembly bag. 

b. Open the small unlabeled bag containing the 37 mm cassette assembly. 

c. Hold the bag so that the sampler can remove the kit. 

d. The support person will hold the tubing for the sampler to place the 37mm cassette 

assembly onto the tubing. 

12. The sample handler will remove the 37 mm cassette assembly from the bag and attach to the 

tygon vacuum tube held by the support person. 

13. The support person or assistant will be prepared to record the duration of sampling. Prompts 

should be given to the sampler so that the sample duration is close to the values in the Table 

1.

Table 1: Suggested Sample Duration and Speed for 12” x 12” surface area

Material Total Sampling 

duration

Single Pass 

duration

Number of passes 

per direction

Concrete 300 seconds 3 seconds 50

Upholstery 300 seconds 3 seconds 50

Carpet 300 seconds 3 seconds 50

14. The sampler will: 

a. Ensure that the filter is correctly placed on the tygon vacuum tube and adjust, if 

necessary. 

b. Turn on the vacuum. 

c. Vacuum “horizontally” using S-strokes to cover the entire area of the material surface 

not covered by the template, while keeping the tygon nozzle angled so that the 

tapered opening of the tygon nozzle is flush with the sample surface. 

Note: a target duration of time to vacuum each coupon should be determined prior to 

testing for each material type. The sampler should pace the speed of the nozzle such 

that the target sampling time is achieved for each coupon. 
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d. Vacuum the same area “vertically” using the same technique. 

e. Turn off the vacuum when sampling is completed. 

f. Hold the nozzle and remove it from the cassette. 

15. The support person will remove the 17 mm tube from the sample kit and open the tube. 

16. The sampler will place the nozzle into the tube with the adapter end down while holding the 

cassette in the opposite hand. 

17. The support person will seal the tube and place in the small unlabeled bag. 

18. The sampler will use the “nozzle” hand to remove the tubing from the outlet side of the 

cassette and hold the cassette out to the support person. 

19. The support person will 

a. Don a fresh pair of gloves. 

b. Seal the cassette with the two red plugs found in the small unlabeled sample 

collection bag. 

20. The support person will open the small unlabeled sample collection bag, and the sampler will 

place the secured 37 mm cassette inside with the 15 mL conical tube. 

21. The support person will then seal the small unlabeled sample collection bag and wipe it with 

a Dispatch® wipe. 

22. The support person will open the labeled 10” x 14” overpack bag and place the smaller 

unlabeled collection bag containing the cassette inside. 

23. The support person will then seal the labeled 10” x 14”overpack bag and wipe it with a 

Dispatch® wipe. 

24. The sampler will remove the used 20 cm length of tubing and discard. 

25. The support person will then place the double contained sample into the sample collection bin. 

26. All members of the sampling team will remove and discard their gloves. 

27. Steps 3 – 22 will be repeated for each sample to be collected. 

28. At the completion of testing, determine the final flow rate of the vacuum using the rotameter 

and record in the project notebook. 

B: 3M Trace Evidence Filter 

Materials: 

 PPE (gloves, lab coat, safety goggles) 
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 Sterile, sealed 10”x15” Twirl-em bags 

 Alcohol wipes 

 Dry wipes moistened with 3% sodium thiosulfate 

 Sharpie and/or pre-printed labels 

 Secondary containment such as a large Tupperware bin 

 Lab notebook 

 QAPP for project that is utilizing the Trace Evidence Filter samples 

 3M Forensic Vacuum Filter (Trace Evidence Filter) (Precision Data Products, Grand Rapids, 

MI catalog #FF-1), referred to as TEF 

1.0 PROCEDURE 

1.1 Assembly of Trace Evidence Filter Kits 

TEF kits are assembled in the following manner: 

1. TEF kits can be assembled outside of the biological safety cabinet, in a dry, clean area. 

Make certain to use proper PPE, including gloves, while handling all TEF kit materials. 

Gather all materials to assemble the kits before assembly. These materials include: 

 - 3M Forensic Vacuum Filters 

- 10”x15” Twirl-em bags 

 - Sharpie or pre-printed labels 

 

2. Obtain a copy of the labeling scheme for the samples. This may be detailed in the QAPP. 

For each TEF kit, use a Sharpie and label a large 10” x 15” Twirl-em bag with the correct 

sample ID. Using 2 pre-printed labels, label the bag containing the TEF with a permanent 

label and a label that can be easily removed. The removable label will be placed on the TEF 

after sampling. 
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Figure 1: Example of a permanent and removable label on a TEF 

3. Open the labeled, 10” x 15” Twirl-em bags one at a time. Place the labeled, TEF in the 10” x 

15” Twirl-em bags that have the corresponding label. Add a non-labeled, 10” x 15” Twirlem bag in the labeled 10” x 15” Twirl-em bag containing the TEF. This completes the TEF 

kit assembly. In the lab notebook, record the date and project for which the TEF kits were 

assembled. 

4. Place the assembled TEF kits into a secondary containment, such as a large Tupperware bin. 

When moving the kits to a sampling location, always have them in secondary containment. 

Figure 2: Example of an assembled TEF kit 
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2.0 VACUUM SAMPLING OF SMALL (12” by 12”) COUPONS

The following procedure will be used in this study for TEF vacuum sampling of each coupon 

surface: 

A two-person team will be used, employing aseptic technique. The team will consist of a 

sampler and a support person. In times when a third person (sample handler) is present, he or she 

will act as an assistant to the support person for data recording. 

1) The Omega vacuum will be maintained on a rolling cart for easy movement into place. 

2) The sampler will plug in the Omega Vacuum into a variable transformer (Staco Energy 

Products Co., Model 3PN1010B) set on 50%. The sampler will wipe down the nozzle 

(inside and out) and end of the tubing first with a Dispatch® wipe, next with a wipe premoistened with 3% Sodium thiosulphate, next with a wipe pre-moistened with 70 % 

ethanol, then don his/her sterile gloves. 

3) Both members of the sampling team will each don a pair of sampling gloves (a new pair 

per sample); the sampler’s gloves shall be sterile sampling gloves as they are the only 

member of the team in contact with the sample. Both members shall wear dust masks to 

further minimize potential contamination of the samples. Further respiratory protection 

beyond a dust mask may be required to protect the sampling team (e.g., SAR; this will be 

specified in the project QAPP). 

4) The support person will aseptically unwrap a template from the bag and present it to the 

sampler, taking care to not touch the template. 

5) The sampler will place the template onto the coupon surface. 

6) The support person will open the sampling supply bin and remove the TEF sample kit from 

the bin. 

7) The support person or assistant will record the sample collection bag ID number on the 

sampling log sheet or in the laboratory notebook. 

8) The sampler and support person will ensure that the correct sample coupon has been 

selected, referencing the coupon code on the sampling bag. 

9) The support person or assistant will record the coupon code on the sampling log sheet next 

to the corresponding TEF collection bag number that was just recorded. 

10) The support person will: 

a. Open the TEF sample kit overpack bag and remove the labeled TEF assembly 

bag. 

b. Open the labeled assembly bag containing the TEF cassette assembly. 

c. Hold the bag so that the sampler can remove the TEF cassette without touching 

the outside of the bag. 

d. The sampler will remove the red round plug from the TEF cassette, and return it 

to the inside of the bag. 
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e. The sampler will attach the TEF to the vacuum hose. 

f. The sampler will remove the black end cap from the TEF cassette, and return it to 

the inside of the bag. 

11) The support person or assistant will be prepared to record the duration of sampling. 

Prompts should be given to the sampler so that the sample duration is close to the values in 

the Table 2. 

Table 2: Suggested TEF Sample Duration and Speed for 12” x 12” surface area

Material Total Sampling 

duration per 

coupon

Single Pass 

duration

Number of passes 

per direction

Concrete 90 seconds 3 seconds 15

Upholstery 90 seconds 3 seconds 15

Carpet 90 seconds 3 seconds 15

12) The sampler will: 

a. Ensure that the filter is correctly placed on the vacuum hose and adjust, if necessary. 

b. Turn on the vacuum. 

c. Vacuum “horizontally” using S-strokes to cover the entire area of the material surface 

not covered by the template, while keeping the nozzle flush with the sample surface. 

d. Vacuum the same area “vertically” using the same technique. 

e. Turn off the vacuum when sampling is completed. 

f. Reseal the inlet with the black end cap. This will be provided to him by the support 

person working the cap to the lip of the bag, touching only the outside of the bag, 

until the sampler can grab it.

g. Remove the filter from the vacuum hose. 

h. Reseal the outlet with the red plug. This will be provided to him by the support 

person working the cap to the lip of the bag, touching only the outside of the bag, 

until the sampler can grab it. 

13) The support person will remove the second removable label from the assembly bag and 

place on the filter housing. 

14) The support person will hold open the evidence bag while the sampler places the filter 

inside. 

15) The support person will seal the evidence bag with the self-adhering strip and wipe the 

outside with a Dispatch wipe. 

16) The support person will open the labeled 10” x 14” overpack bag and place the evidence 
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collection bag containing the cassette inside. 

17) The support person will then seal the labeled 10” x 14”overpack bag and wipe it with a 

Dispatch® wipe. 

18) The support person will then place the double contained sample into the sample collection bin. 

19) All members of the sampling team will remove and discard their gloves. 

20) The sampler will wipe down the nozzle (inside and out) and end of the tubing first with a 

Dispatch® wipe, next with a wipe pre-moistened with 3% STS, next with a wipe premoistened with 70 % ethanol. 

21) Steps 4 – 24 will be repeated for each sample to be collected. 
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Miscellaneous Operating Procedure (MOP) 3165:

Sponge Sample Collection Protocol

Prepared by: __________________________________________ Date: 11/15/2012 

Stella McDonald, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012 

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012 

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
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MOP 3165 

Title: SPONGE SAMPLE COLLECTION PROTOCOL 

Scope: This MOP outlines the procedure for collecting spores using a 3M Sponge-StickTM.

Purpose: To provide a procedure for the collection of spore samples using a Sponge-StickTM 

in a consistent and repeatable manner.

MATERIALS

• 3M Sponge-SticksTM (P/N SSL10NB), hereafter referred to as ‘sponge’

• One Seward stomacher bag (P/N BL6041/CLR) per kit

• Disposable gloves

• Sterilized sampling templates

• One Fisher Sterile sampling bag with flat wire enclosure (7” x 12”, P/N 14-955-194) per kit

• One Fisher Sterile sampling bag with flat wire enclosure (10” x 14”, P/N 01-002-53) per kit 

for overpack

• Dispatch wipes

1.0 PREPARATION

All materials needed for collection of each sample will be prepared in advance using aseptic 

technique. A sample kit for a single sponge sample will be prepared as follows:

1.1 One stomacher bag will be uniquely labeled as specified in the project QAPP.

1.2 A 10” x 14” bag will be labeled with the same ID as the stomacher bag.

1.3 One stomacher bag, and one 9.5” x 12” unlabeled bag will be placed in the overpack bag.

1.4 A sterile Sponge-Stick will be added to the overpack bag.

1.5 Each prepared bag is one sampling kit.

2.0 PROCEDURE 

A two person team will be used, employing aseptic technique throughout. The team will consist of 

a sampler and a sample handler. In some cases, a third person may be needed to move samples. 

Throughout the procedure, the support person will log anything they deem to be significant into the 

laboratory notebook.
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In general, the team works from the least contaminated sample set (i.e., control blanks) towards the 

most contaminated sample set (i.e.,positive controls).

All members shall wear dust masks to minimize potential contamination of the samples. 

Depending on the situation, respiratory protection beyond a dust mask may be required to protect 

the sampling team (e.g., SAR; this will be specified in the project QAPP). New disposable lab 

coats are required for the sample handler when changing between different types of materials or 

when direct contact between the coupon and lab coat occurs.

2.1 Wearing a clean pair of gloves over existing gloves, the sampler will place the disposable 

template over the area to be sampled.

2.2 The support person will remove a sample kit from the sampling bin and record the sample 

tube number on the sampling log sheet next to the corresponding coupon code just 

recorded.

2.3 The sampler and support person will verify the sample code and ensure that the correct 

coupon and location are being sampled.

2.4 The support person will:

a) Open the outer sampling bag touching the outside of the bag.

b) Touching only the outside of the (10” x 14”) bag, remove the sponge, and hand it to

the sampler.

c) Remove the stomacher bag, being careful to not touch the inside of the outer sampling 

bag, and open it touching only the outside.

2.5 The sampler will remove the sterile sponge from its package. Grasp the sponge near the top 

of the handle. Do not handle below the thumb stop.

2.6 The sampler will wipe the surface to be sampled using the moistened sterile sponge by 

laying the widest part of the sponge on the surface, leaving the leading edge slightly lifted. 

Apply gentle but firm pressure and use an overlapping ‘S’ pattern to cover the entire 

surface with horizontal strokes (Figure 1). Use the other hand to hold the template during 

sampling, being careful not to touch the surface.
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Figure 1. First pass with sponge – horizontal strokes using one side of the sponge

2.7 The sampler will turn the sponge over and wipe the same area again using vertical ‘S’-

strokes (Figure 2). 

Figure 2. Second pass with sponge – vertical strokes using the other side of the sponge

2.8 The sampler will the use the edges of the sponge (narrow sides) to wipe the same area using 

diagonal ‘S’-strokes (Figure 3). The sponge will be flipped to use the opposite side 

immediately after the longest stroke at opposite corners.
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Figure 3. Third pass with sponge – diagonal strokes using the edges of the sponge

2.9 The sampler will use the tip of the sponge to wipe the perimeter of the sampling area 

(Figure 4). 

Figure 4. Final (fourth) pass with sponge – perimeter wipe using the tip of the sponge

2.10 The sample handler will open the stomacher bag, careful not to touch the inside of the bag. 

2.11 The sampler will place the end of the sponge in the bag, holding the handle outside the 

opening of the bag.

2.12 The sample handler will grasp the sponge from outside of the bag, and help the sample break 

off the handle of the sponge. The handle below the thumbstop should not touch the inside of 

the stomacher bag.

2.13 The sample handler will securely seal the stomacher bag and wipe the outside with a 

disinfecting wipe.

2.14 The sample handler will then place the stomacher bag inside the unlabeled sterile bag.
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2.15 The sample handler will place this in the overpack bag and wipe the overpack bag with 

disinfecting wipes.

2.16 The sample handler will place the overpack bag in the sample bin.

NOTE: Remove excessive air from the re-sealable plastic bags to increase the number of 

samples that can be shipped in one container.

2.17 The sampler will dispose of the template.

2.18 Both members will remove outer gloves and discard. Clean gloves should be worn for each 

new sample.

3.0 REFERENCES

Sponge sample collection protocol adapted from:

National Validation Study of a Cellulose Sponge Wipe-Processing Method for Use after Sampling 

Bacillus anthracis Spores from Surfaces. Rose, Laura J.; Hodges, Lisa; O’Connell, Heather; 

Noble-Wang, Judith. Appl. Environ. Microbiol. 2011, 77(23):8355.

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MOP 3168

TITLE: AGGRESSIVE AIR SAMPLING (AAS) FOR WA 3-28: PHASE I SAMPLING

APPROACH

SCOPE: This MOP outlines the setup, operation, and timeline schedule for conducting

AAS testing in COMMANDER.

PURPOSE: To provide a standardized and repeatable procedure for all AAS tests to be

conducted under WA 3-28’s Phase I sampling matrix using high volume (HiVol)

samplers.

1.0 INTRODUCTION

Preparations for each test will be conducted according to the schedule listed in this procedure.

Any deviations will be noted in the laboratory notebook, along with the reason for the deviation.

Section 2.0 lists the preparation steps that need to be taken the Thursday before testing is to

occur (Day 1). Sections 3.0 and 4.0 detail the pre- and post-decon procedures to be followed for

AAS testing.

2.0 PREPARATION

NOTE: Materials needed before Day 1 include material coupons and gaskets, placed in

VHP bags, and exposed to a VHP sterilization cycle. These should be left to

degas for a minimum of 3 days before use.

Day 1 – Thursday

1. Ensure the following materials for inoculation are in airlock:

 (2) 3 x 3 Grids

 (6) ADAs and clamps for single coupons

 (18) ADAs and clamps for 3 x 3 coupons

 Oscillating fan

 Tables for coupon staging come Monday
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2. Ensure all equipment needed for testing is in COMMANDER and COMMANDER is setup

(see Figure 1):

 SAR inlet filter box, hoses and masks

 (2) Blowers and needed power cords

 (3) HiVol Samplers in open orientation (no air-tight seals)

 HiVol samplers should have been calibrated the day prior

 Hydrogen peroxide (H2O2) air monitors are calibrated and functional

Figure 1. COMMANDER layout showing HiVol sample numbers

3. Close and seal COMMANDER door.

4. Close airlock door.

5. VHP airlock per MOP-3120 (VHP Operation).

To airlock

Fan

HighVol – R1

HighVol – R2

HighVol – R3
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Day 2 – Friday

1. Aerate the airlock upon arrival in the morning.

2. VHP COMMANDER per MOP-3120 in the morning, so that external aeration can begin in

mid afternoon.

3. Clear the airlock for entry as early as possible using a Draeger tube.

4. Set up coupons in H130, outside of the enclosure, using equipment from the airlock and

previously sterilized coupons and gaskets allowed to degas in VHP bags.

5. Inoculate coupons according to MOP 6561 (Aerosol Deposition of Spores onto Material

Coupon Surfaces using the Aerosol Deposition Apparatus (ADAs)). Fill out Attachment B,

WA 3-28 Coupon Deposition Log. The list of coupon IDs follows, where M is the material

ID; L(Laminate), D(Drywall), or C(Carpet):

42” x 42” not inoculated 28-[test ID]-XM,

42” x 42” 28-[test ID]-M

14” x 14” material 28-[test ID]-M1-R[1,2,3]

14” x 14” stainless steel 28-[test ID]-S-R[1,2,3]

14” x 14” material not inoculated 28-[test ID]-XM1-F1

6. Calibrate the COMMANDER ATI H2O2 sensor outside of COMMANDER using MOP 3136

(General Procedure for Calibration of ATI Hydrogen Peroxide Gas Transmitters using

Solution Wells).

7. Place the ATI sensor back into COMMANDER.

8. Mop the enclosure with Clorox Clean-up.

9. Close enclosure doors.
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3.0 PRE-DECON AAS PROCEDURES

Day 3 - Monday

 Conduct a pre-job safety briefing prior to initiating work.

 A “buddy” equipped with appropriate respiratory protection is required to be present

outside of COMMANDER when personnel are working inside.

1. Set up the ELPI system from outside of COMMANDER and zero according to MOP 3133

(Basic Operation and Maintenance of the Dekati Electrical Low Pressure Impactor (ELPI)).

2. Wipe down the outside of the 42” x 42” coupons, the 14” x 14” test coupons, and ADA’s

with dispatch wipes.

3. Transfer the coupons from H130 to the airlock. This operation will be done with a clean team

inside the airlock, and a team in H130. The H130 team (in clean garb) will complete the wipe

down, and then change garb. The airlock team will enter the airlock in clean garb and

booties. During the moving operation, the H130 team will pass the coupon to the airlock

team with no direct contact between alternate teams. Once both coupons have been passed

into the airlock, all personnel will leave the airlock, and it will be purged for 30 minutes. An

oscillating fan will be used to prevent settling.

4. Positive control 14” x 14” coupons should be moved at this same time, but put back in their

original location. This is to simulate the movement undergone by the test coupons.

5. Verify the camera is recording, and that the DAS is operational and recording data.

6. Assemble the supplies:

a. Sterilized PM10 filters (8)

b. Sterilized collection shims (8)

c. Sterile gloves (2 packs) and laboratory gloves

d. Sterile garb x 8(coat, p95, hair net, 12 boot covers)

e. EtO’d notebook, AAS event log and pen

f. Digital timer (2)

g. Duct tape

h. Blank sample kits (Filters Sample IDs : 28-05a-XAF-1-R(1-3), and 28-05a-XAF1-F1),( Shims Sample IDs : 28-05a-XAS-1-R(1-3) and 28-05a-XAS-1-F1)
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(Sponge sticks: 28-05a-M-1-R(1-3), 28-05a-XM-1-F1 28-05a-S-1-R(1-3)

and 28-05a-S-1-F1)

i. Cooling vests

j. Dispatch wipes

k. Molykote® Grease Spray

l. Anemometer

7. Three personnel (referred to as “Clean Man”, “Dirty Man 1”, and Dirty Man 2”, or CM,

DM1 and DM2) will enter the enclosure and wipe in bins of supplies, then close the

enclosure doors.

- Plug in and record position of power cords for blowers and fans.

- Install the relative humidity (RH) and vaprous hydrogen peroxide (VHP) sensors to

the COMMANDER chamber.

- Verify supply quantities against the WA 28 packing list (found on DTRL/WA 3-28).

- ELPI should be monitoring enclosure PM, and should be changed to COMMANDER

immediately before entry.

- Personnel should spend 10 minutes minimum in enclosure before opening airlock

door.

8. Three personnel (CM, DM1, and DM2) will enter the airlock wearing cooling vests, C-suits

and carrying the supply bins:

9. Once the airlock door is closed, open the door to COMMANDER. Put on clean garb over the

C-suit. Install the sterile filters on top of the vents of the three HiVol samplers (vent seen in

Figure 2a). To do this, the support person (DM1) will grab the filter frame with sterile gloves

and hold it out of the way. The sample handler (DM2) will open the bag containing the filter

and the sampler (CM) will use sterile thumb forceps to put the filter in place. The DM1 will

then return the frame and secure it in place. Unlatch the hinge of the head unit and pull down.

Secure latches around the base of the unit (Figure 2b). CM can help with this last step before

changing into new sterile gloves.

10. The DM1 will unwrap the aluminum foil covering the collection shim, leaving the shim

inside and exposed. The DM2 will spray the shim with Molykote® Grease Spray, ensuring

full coverage of the shim. Wearing sterile gloves, the CM will aseptically pull the shim out of

the foil and place in each of the three HiVol inlet heads, minimizing contact with the greased

surface. Align the shim and secure in place with the Teflon tabs (Figure 2c).

11. CM will unlatch the top covering of the head unit and pull down. Secure in place with the

latches (Figure 2d).
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12. DM1 and DM2 will return to the airlock.

13. Wearing sterile gloves, the DM1 will remove the apparatus brace, ADAs, and gaskets from

the sterile coupon and put aside in the airlock. DM1 and DM2 will pick up the coupon, being

careful not to touch the surface. Move the coupon into COMMANDER and place in the

designated sampling position. To do this, the coupon will need to be rotated vertically so that

it can fit through the door.

14. DM1 and DM2 remove their gowns and don SAR. The CM returns to the airlock and closes

the airlock door.

 Inspect SAR masks and hoses and the filter box prior to donning.

a. b.
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c. d.

Figure 2. a) HiVol sampler with head unit and top open, and filter vent secured;

b) head unit closed and latched; c) placement of shim; and d) top closed and

latched.

15. Turn on the fan to the 70% speed setting. Measure the wind speed at the HiVol inlets. These

should be less than 30 mph.

16. Test the force of the leaf blower, pointed away from the coupon, using the anemometer.

17. The DM1 will agitate the surface of the coupon using a leaf blower according to MOP 3166

(Aerosolization of Contaminated Coupons Using the Toro Power Sweep Electric Blower for

Aggressive Air Sampling (AAS)). At the very moment the blower is turned on, DM2 will turn

on all three HiVol samplers. DM2 will monitor the blowing time as per MOP 3166 and

document the intervals by filling out the Aggressive Air Sampling (AAS) Event Log

(Attachment A).

18. At the completion of the agitation, the DM1 and DM2 remain in COMMANDER. After 20

minutes total (10 minutes after finishing aggressive agitation), DM2 turns off the HiVol

samplers.

19. Turn off (unplug) the fan.

20. CM will re-enter COMMANDER.
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21. Collect blank samples:

Filters Sample IDs : XAF-1-R(1-3) + XAF-1-F1 (Field blank)

Shims Sample IDs : XAS-1-R(1-3) + XAS-1-F1 (Field blank)

a. DM1 and DM2 don outer layer of clean garb.

b. The DM1 will open the first HiVol sampler and latch in it place.

c. The DM1 will don a pair of sterile gloves and remove the collection shim, being careful

not to touch the sides of the flow ports.

d. The DM2 and CM will then use a sponge kit to sample the top surface of the shim

according to MOP 3169 (Sponge Sample Collection Protocol for AAS Shims).

e. The DM1 will then discard the shim to the airlock for sterilization. The DM1 will unlatch

the base of the HiVol head unit and tilt the unit up to reveal the vent (see Figure 2a).

f. The DM1 will don a new pair of sterile gloves and remove the vent bracket and top grate,

being careful not to touch the filter.

g. Donning new gloves, the CM will remove the filter using sterile thumb forceps. Fold the

filter in half and then in half again, keeping the top side of the filter inside of the fold.

Using the forceps, transfer the filter directly to a stomacher bag. The DM2 will hold the

stomacher bag open for this transfer.

h. Sample the shim plates and remove the filters for the two remaining HiVol samplers

using the same protocol.

22. Remove the blank coupon and place against the COMMANDER wall.

23. All personnel change into new sterile garb.

24. CM will install sterile filters on top of the vents of the three HiVol samplers and secure in

place. Unlatch the hinge of the head unit and pull down. Secure latches around the base of

the unit.

25. The DM1 will unwrap the aluminum foil covering the collection shim, leaving the shim

inside and exposed. The DM2 will spray the shim with Molykote® Grease Spray, ensuring

full coverage of the shim. Wearing sterile gloves, the DM2 will aseptically pull the shim out
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of the foil and place in each of the three HiVol inlet heads, minimizing contact with the

greased surface. Align the shim and secure in place with the Teflon tabs (see Figure 2c).

26. DM2 will unlatch the top covering of the head unit and pull down. Secure in place with the

latches.

27. Personnel will return to the airlock and DM1 and DM2 will don 3 pairs of boot covers.

28. Wearing gloves, the DM1 will remove the apparatus brace, ADAs, and gaskets from the

inoculated coupon and put aside in the airlock. DM1 and DM2 will pick up the coupon, being

careful not to touch the inoculated surface, and move the coupon into COMMANDER and

place in the designated sampling position. DM1 and DM2 will remove one pair of boot

covers while crossing the threshold into COMMANDER with the help of CM.

29. In the same aseptic manner, remove ADAs from material coupons and place inside on

COMMANDER floor just above the test coupon, removing a second pair of boot covers

while entering COMMANDER. This process will make one large 42” x 56” coupon.

30. The DM1 and DM2 remove their gowns and don SAR. CM closes the airlock door.

31. CM communicates to the buddy outside to unplug blowers 2 and 3. Confirm this is done to

DM2. Alternatively, the CM can unplug as he leaves the airlock.

32. DM2 then plugs in blowers 2 and 3 (now, or after the 12 minute AAS operation).

33. DM2 turns on the fan to the 70% speed setting.

34. Agitate the surface of the inoculated coupon according to MOP 3166. At the very moment

the blower is turned on, a DM2 will turn on the first HiVol sampler as specified in the QAPP.

The DM2 will monitor the blowing time as per MOP 3166 and document the intervals by

filling out the AAS Event Log.

35. When agitation is complete, remove SAR and leave in COMMANDER.

36. CM to exit airlock, leaving clean garb in the airlock upon exit.

37. DM 1 and DM2 will open the airlock door, move to the airlock and close the airlock door.

38. DM1 and DM2 will shower out and exit the airlock.
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39. After 20 minutes of air sampling, turn off the first HiVol sampler. Immediately turn on the

second sampler.

40. After 20 minutes of sampling with the second HiVol sampler, turn off the second sampler

and immediately turn on the third. Again, sample for 20 minutes and then turn off.

41. Wearing clean garb, collect samples from the stainless steel and laminate control coupons

according to MOP 3165 (Sponge Sample Collection Protocol). A blank laminate coupon (L1-

F1) and a field blank for sponge sticks (S1-F1) should also be sampled at this time.

Reference Positives IDs : S-1-R(1-3), L1-R(1,2,3), L1-F1, and S1-F1

42. VHP airlock and start aeration as soon as possible (MOP-3120). Allow 18 hours for the

agitated spores to settle in COMMANDER.

Day 4 – Tuesday

1. Verify the camera is recording, and that the DAS is operational and recording data.

2. After waiting the specified spore-settling time, a three-person sampling team (CM, DM1,

and DM2) will enter COMMANDER in C-suits carrying :

a. Sterile gloves and laboratory gloves

b. Sterile garb (coat, p95, hair net, 8 boot covers)

c. EtO’d notebook, AAS event log and pen

d. Digital timer

e. Duct tape

f. Sample kits: Kits: 28-05a-MAF-1-R{1-3) and 28-05a-MAF-1-F1

28-05a-MAS-1-R{1-3) and 28-05a-MAS-1-F1

g. Dispatch wipes

3. Each will don sterile garb and boot covers.

4. Enter COMMANDER, removing one pair of boot covers while crossing the threshold.

5. Remove the air filters from the HiVol samplers. The team will consist of a CM, DM2 and a

DM1.
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Filter Sample IDs: MAF-1-R(1-3), where M is material ID – C-carpet, L-laminate, and

W-drywall

Shim Sample IDs: MAS-1-R(1-3), where M is material ID – C-carpet, L-laminate, and

W-drywall

6. The DM1 will open the first HiVol sampler and latch in it place.

7. The DM1 will don a pair of sterile gloves and remove the collection shim, being careful not

to touch the sides of the flow ports.

8. The DM2 and CM will then use a sponge kit to sample the top surface of the shim according

to MOP 3169 (Sponge Sample Collection Protocol for AAS Shims). CM will manipulate the

sponge stick.

9. The DM1 will then discard the shim into the airlock for disinfection. The DM1 will unlatch

the base of the head unit and tilt the unit up to reveal the vent.

10. The DM1 will don a new pair of gloves and remove the vent bracket and top grate, being

careful not to touch the filter.

11. Donning new gloves, the CM will remove the filter using sterile thumb forceps. Fold the

filter in half and then in half again, keeping the top side of the filter inside of the fold. Using

the forceps, transfer the filter directly to a stomacher bag. The DM2 will hold the stomacher

bag open for this transfer.

12. Sample the shim plates and remove the filters for the two remaining HiVol samplers using

the same protocol.

13. All HiVol samplers should be in the fully open position.

14. All personnel shower out of the airlock.

15. VHP COMMANDER as per MOP 3120. Conditions will be determined by the WAM.

16. Start aeration of COMMANDER.

17. VHP the airlock as per MOP 3120.
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4.0 POST-DECON AAS PROCEDURES

Day 5 - Wednesday

1. Start airlock aeration as soon as possible and ensure conditions are safe for entry.

2. Verify the camera is recording, and that the DAS is operational and recording data.

3. Three personnel will enter the airlock wearing C-suits and carrying:

a. Sterilized PM10 filters (3)

b. Sterilized collection shims (3)

c. Sterile gloves and laboratory gloves

d. Sterile garb (coat, p95, hair net, boot covers)

e. EtO’d laboratory notebook, AAS event log and pen

f. Digital timer

g. Duct tape

h. Dispatch wipe

i. Sample kits Sponge stick kits 28-05a-L-2-R(1-3) and 28-05a-L-2-F1

4. Once the airlock door is closed, open the door to COMMANDER. Don boot covers (2 pairs),

being careful not to contaminate the first pair with the airlock floor.

5. Move into COMMANDER, removing one pair of boot covers when crossing the threshold.

6. Collect post-decon sponge stick surface samples (or vacuum sock samples of carpet coupons)

of the three positive coupons per MOP 3165. CM will serve as the sampler, DM1 will serve

as the support person.

Sample IDs: M-2-R(1-3), where M is material type as listed above

7. DM2 will place coupons in soapy water in airlock after sampling.

8. All personnel to change into new clean garb.

9. CM will install the sterile filters on top of the vents of the HiVol samplers and secure in

place. DM2 will unlatch the hinge of the head unit and pull down. Secure latches around the

base of the unit.
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10. The DM1 will unwrap the aluminum foil covering the collection shim, leaving the shim

inside and exposed. The DM2 will spray the shim with Molykote® Grease Spray, ensuring

full coverage of the shim. Wearing sterile gloves, the DM2 will aseptically pull the shim out

of the foil and place in each of the three HiVol inlet heads, minimizing contact with the

greased surface. Align the shim and secure in place with the Teflon tabs (see Figure 2c).

11. The DM1 and DM2 will remove their gowns and don SAR, while CM retires to airlock.

12. CM to communicate to buddy outside to verify Blower 1 power cord is plugged in, and

Blowers 2 and 3 are off.

 Inspect the SAR masks and hoses and the filter box prior to donning.

13. Turn on the fan to the 70% speed setting. Measure the wind speed at the HiVol inlets. These

should be less than 30 mph.

14. Test the force of the leaf blower, pointed away from the coupon, using the anemometer.

15. Agitate the surface of the coupon according to MOP 3166. At the very moment the blower is

turned on, DM2 will plug in the first HiVol sampler. The DM2 will monitor the blowing time

as per MOP 3166 and document the intervals by filling out the AAS event log.

16. When agitation is complete, DM1 and DM2 move to airlock, remove SAR and drop their

hose and mask in COMMANDER, close the airlock door and shower out of the

COMMANDER airlock.

17. After 20 minutes of air sampling, turn off the first HiVol sampler. Immediately turn on the

second sampler.

18. After 20 minutes of sampling with the second HiVol sampler, turn off the second sampler

and immediately turn on the third. Again, sample for 20 minutes and then turn off.

19. Allow chamber to settle for 2 hours.

20. After the settling time, a three-person sampling team will enter COMMANDER carrying

post-decon sample kits: filter kits 28-05a-LAF-2-R{1-3) and 28-05a-LAF-2-F1 and shim

kits 28-05a-LAS-2-R{1-3) and 28-05a-LAS-2-F1

21. Sample the collection shims and collect the filters.

Filter Sample IDs: MAF-2-R(1-3), where M is material code , listed above

Shim Sample IDs: MAS-2-R(1-3), where M is material code , listed above
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a. The DM1 will open the first HiVol sampler and latch in it place.

b. The DM1 will don a pair of sterile gloves and remove the collection shim, being

careful not to touch the sides of the flow ports.

c. The DM2 and CM will then use a sponge kit to sample the top surface of the shim

according to MOP 3169.

d. The DM1 will then discard the shim to the airlock. The DM1 will unlatch the base of

the head unit and tilt the unit up to reveal the vent.

e. The DM1 will don a new pair of gloves and remove the vent bracket and top grate,

being careful not to touch the filter.

f. Donning new gloves, the CM will remove the filter using sterile thumb forceps. Fold

the filter in half and then in half again, keeping the top side of the filter inside of the

fold. Using the forceps, transfer the filter directly to a stomacher bag. The DM2 will

hold the stomacher bag open for this transfer.

g. Sample the shim plates and remove the filters for the two remaining HiVol samplers

using the same protocol.

22. Conduct post-test flow calibration as outlined in MOP 3170 (HiVol Calibration Check).

23. Verify that two blowers are placed in COMMANDER and that the samplers are opened.

Ensure all SAR related equipment is located in COMMANDER.

24. Exit COMMANDER and shower out of the airlock.

25. VHP COMMANDER per MOP-3120.
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Attachment A

Aggressive Air (AA) Sampling Event Log

Test ID ______________

Agitator

DM2

Buddy 2

Reference:

Dimensions

(in2)

Agitation

Time (s)

Horizontal

Sweeps

Vertical

Sweeps

Sweep

rate

(s/sweep)

14 x 14 60 10 10 3

28 x 28 240 20 20 6

42 x 42 540 30 30 9

Test Day/

date

Coupon ID High Vol 1

start

High Vol

2 start

High Vol 3

start

2/

Sterile

Blank

2/ Test

4/ Test

Test Day/

date

Coupon ID Vertical

start

Vertical

end

# sweeps Horizontal

start

Horizontal

end

#

sweeps

2/

Sterile

Blank 30 30

2/ Test 30 30

3/ Positive 1 10 10

3/ Positive 2 10 10

3/ Positive 3 10 10

4/ Test 30 30
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Attachment B

WA 3-28 Coupon Deposition Log Page ____ of ___

Test ID MDI ID:

Date Initial Weight (g):

Personnel / Title Final Weight (g):

Time

Coupon

ID Vortex Interval (s) Puff Number

MDI

Weight (g) Comments
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Miscellaneous Operating Procedure (MOP) 6535a: 

Serial Dilution: Spread Plate Procedure to Quantify Viable 

Bacterial Spores

Prepared by: __________________________________________ Date: 2/11/2013 

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 2/11/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 2/11/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP 6535a 

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MOP 6535a 

TITLE: SERIAL DILUTION: SPREAD PLATE PROCEDURE TO QUANTIFY 

VIABLE BACTERIAL SPORES

SCOPE: Determine the abundance of bacterial spores in a liquid extract 

PURPOSE: Determine quantitatively the number of viable bacterial spores in a liquid 

suspension using the spread plate procedure to count colony-forming units (CFU) 

Materials: 

• Liquid suspension of bacterial spores 

• Sterile centrifuge tubes

• Diluent as specified in QAPP or Test Plan (e.g., sterile water, Phosphate Buffered Saline with 

Tween 20 (PBST)) 

• Media plates as specified in QAPP or Test Plan (e.g., Trypticase Soy Agar (TSA) plates)

• Microliter pipettes with sterile tips

• Sterile beads placed inside a test tube (used for spreading samples on the media surface

according to MOP 6555 (Petri Dish Media Inoculation Using Beads) or cell spreaders

• Vortex mixer

1.0 PROCEDURE (This protocol is designed for 10-fold dilutions.) 

1. For each bacterial spore suspension to be tested label microcentrifuge tubes as follows: 10-1, 

10-2, 10-3, 10-4, 10-5, 10-6... (The number of dilution tubes will vary depending on the 

concentration of spores in the suspension). Aseptically, add 900 uL of sterile diluent to each 

of the tubes. 

2. Label three media plates for each dilution that will be plated. These dilutions will be plated 

in triplicate. 

3. Mix original spore suspension by vortexing thoroughly for 30 seconds. Immediately after the 

cessation of vortexing, transfer 100 uL of the stock suspension to the 10-1 tube. Mix the 10-1

tube by vortexing for 10 seconds, and immediately pipette 100 uL to the 10-2 tube. Repeat 

this process until the final dilution is made. It is imperative that used pipette tips be 

exchanged for a sterile tip each time a new dilution is started.

4. To plate the dilutions, vortex the dilution to be plated 10 seconds, immediately pipette 100 

uL of the dilution onto the surface of a media plate, taking care to dispense all of the liquid
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from the pipette tip. If less than 10 seconds elapses between inoculation of all replicate 

plates, then the initial vortex mixing before the first replicate is sufficient for all replicates of 

the sample. Use a new pipette tip for each set of replicate dilutions. 

5. Carefully and aseptically spread the aliquotted dilution on the surface of the media either by 

use of glass beads (MOP 6555) or cell spreader (the method used may be directed in the 

QAPP or Test Plan) until the entire sample is distributed on the surface of the agar plate. 

Repeat for all plates. 

6. Incubate the plates for the optimum time period at the optimum growth temperature for the 

target organism (incubation conditions will vary depending on the organism’s optimum 

growth temperature and generation time. This information can be found in Bergey’s Manual 

of Determinative Bacteriology or it will be provided with the ATCC certification.

7. Manually enumerate the colony forming units (CFU) on the media plates by manually 

counting with the aid of a plate counting lamp and a marker (place a mark on the surface of 

the Petri dish over each CFU when counting, so that no CFU is counted twice). A hand held 

tally counter or an electronic counting pen may be used to assist the person counting, but 

may not be used as the primary source for the count. 

Quality control (QC) requirements for bacterial enumeration will be addressed per QAPP or 

test plan. However, in general, the following QC practices should always be adhered to: 

a. The arrangement of plates and tubes, and the procedure for preparing dilutions and 

enumerating CFU should be done the exact same way each time. This helps prevent 

systematic errors and often helps determine the cause of problems when a discrepancy is 

found. 

b. A visual check of the graduated pipette tip should be made during each use to ensure the 

pipette is pulling properly. 

c. Samples should acclimate to room temperature for 1 hour prior to plating. 

d. Samples should be processed (extracted and plated) from the least contaminated to the 

most contaminated.

e. When a target range of CFU is known, three dilution factors are plated to bracket the 

expected results (0, -1, and -2, if the -1 dilution factor was the target).

f. Enumerated colonies and results should be verified that the results are the target 

organism, and that second counts have been performed. Second counts must be 

completed on 25% of significant data, and must be within 10% of the first count. If 

CFUs are found to have more than a 10% difference between first and second counts, 

then a third count is to be completed. 
MOP 6535a 

Revision 4 

January 2013 

Page 4 of 8 

g. Pictures should be taken of any plates that are contaminated or have results out of the 

normal 

8. Record all quantitative data in the “Serial Dilution/Plating Results Sheet”. Target range for 

statistically significant counts is 30-300 CFU. Data that fall out of the 30-300 CFU range are 

addressed in MOP 6584 (Procedure for Replating Bacteria Spore Extract Samples) and MOP 

6565 (Filtration and Plating of Bacteria from Liquid Extracts). 

 

2.0 CALCULATIONS

Total abundance of spores (CFU) within extract: 

(Avg CFU / volume (mL) plated) X (1 / tube dilution factor) X extract volume

For example: 

 

Tube Dilution Volume plated Replicate CFU

 10-3 100 μL (0.1 mL) 1 150 

 10-3 100 μL (0.1 mL) 2 250 

10-3 100 μL (0.1 mL) 3 200 

 

 

Extract total volume = 20 mL 

 (200 CFU / 0.1 mL) X (1/10-3) X 20 mL = 

 (2000) X (1000) X 20 = 4.0 X 107 CFU

Note: The volume plated (mL) and tube dilution can be multiplied to yield a ‘decimal factor’ 

(DF). DF can be used in the following manner to simplify the abundance calculation.

Spore Abundance per mL = (Avg CFU) X (1 / DF) X extract volume 
MOP 6535a 

Revision 4 

January 2013 

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Serial Dilution/Plating Results Sheet Page 1 of ______

TEST INFORMATION

EPA Project No. PI

Technician Name Test Date

Technician Signature Test No.

RESULTS

Date: Volume Plated:

Tube Dilution

Sample ID Plate

Repl. 100 10-1 10-2 10-3 10-4 10-5 10-6

A 

B 

C 

A 

B 

C 

A 

B 

C 

A 

B 

C 

A 

B 

C 

A 

B 

C 

NOTES: 

 
MOP 6535a 

Revision 4 

January 2013 

Page 6 of 8 

Page 2 of ______

Sample ID Plate

Repl. 100 10-1 10-2 10-3 10-4 10-5 10-6

A

B 

C 

A 

B 

C

A 

B 

C 

A 

B 

C 

A 

B 

C 

A

B 

C 

A 

B 

C

A 

B

C 

A 

B 

C 

A 

B

C 

NOTES: 

 

 
MOP 6535a 

Revision 4 

January 2013 

Page 7 of 8 

Page _____ of ______

Sample ID Plate

Repl. 100 10-1 10-2 10-3 10-4 10-5 10-6

A

B 

C 

A 

B 

C

A 

B 

C 

A 

B 

C 

A 

B 

C 

A

B 

C 

A 

B 

C

A 

B

C 

A 

B 

C 

A 

B

C 

NOTES: 

 

 
MOP 6535a 

Revision 4 

January 2013 

Page 8 of 8 

Page _____ of ______

Sample ID Plate

Repl. 100 10-1 10-2 10-3 10-4 10-5 10-6

A

B 

C 

A 

B 

C

A 

B 

C 

A 

B 

C 

A 

B 

C 

A

B 

C 

A 

B 

C

A 

B

C 

A 

B 

C 

A 

B

C 

NOTES: 

 

 
MOP 6555

Revision 2

November 2012

Page 1 of 4

Miscellaneous Operating Procedure (MOP) 6555:

Petri Dish Media Inoculation Using Beads

Prepared by: __________________________________________ Date: 11/15/2012

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6555

Revision 2

November 2012

Page 2 of 4

MOP 6555

TITLE: PETRI DISH MEDIA INOCULATION USING BEADS

SCOPE: This MOP outlines the procedure for cleaning, assembling, and using beads to

inoculate agar plates.

PURPOSE: To provide an easily repeatable method for spreading liquid inoculation onto

agar plates.

Equipment:

 #13 test tubes

 6 x 12 test tube racks (which hold 72 tubes)

 Beads of various sizes (glass)

 Glass autoclavable trays (stainless steel is eventually corroded by the bleach and

autoclaving processes)

 Bleach

 DI water

 Hot gloves

 Amber bottle for collecting hazardous waste (with hazardous waste label)

 Funnel

 Aluminum foil

 Label tape

 Chemical hood

 Autoclave

 Oven

 Labeled bead container or cup (All mold and bacteria beads must be kept separately)

1.0 CLEANING BEADS

1. When a sufficient number of beads have been collected, or at least once a day when

beads are being used to spread colonies, place the used beads into a tray with a solution

containing a 1:5 ratio of bleach to deionized water.

Add the bleach to the beads first, under the protection of a chemical safety cabinet. Then
MOP 6555

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November 2012

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add the deionized water. Cover the pan with aluminum foil and label it with the

contents (for example: “bacteria beads in 1:5 bleach to DI water solution”). Soak the

beads 12-24 hours (usually overnight) in a chemical fume hood.

2. After soaking, take a bottle brush and thoroughly scrub the beads.

3. Decant the bleach solution (collect the bleach for proper disposal) and rinse with

deionized water 6 to 8 times, collecting the rinsate after the first rinse for disposal

(subsequent rinses can be discarded in the lab sink). Rinse until the decanted liquid is

clear. Use a funnel to add the bleach waste to a labeled amber waste bottle. These

liquids must be labeled “hazardous waste” and can then be stored, collected or disposed

of properly.

4. Cover the beads with deionized water and autoclave for 1 hour on the liquid cycle.

5. Decant the deionized water and place the tray of beads in the Thelco lab oven at 121 C

until dry (a minimum of 3 hours).

6. Remove the beads from the oven using proper safety equipment (heat gloves) and cover

with clean aluminum foil to prevent contamination. Label each tray with the following

information:

“Clean bacteria (or mold) beads,” the date beads were cleaned, initials of the person

who cleaned them.

7. These beads are then ready for use as described in “PLACING BEADS IN TUBES”.

3.0 PLACING BEADS IN TUBES

1. Fill a 6 x 12 rack with tubes

2. Place clean beads into a shallow pan, and then manually fill each tube with 7-15

beads/tube.

Note: Beads vary in size and will therefore fill the tubes to different heights.

3. Tightly attach a cap to each tube

4. Autoclave for 1 hour using a gravity sterilization cycle (see MOP 6570). Autoclave tape

must be placed on the top of each rack to provide evidence that the beads have been

sterilized
MOP 6555

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November 2012

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4.0 SPREADING BEADS

1. To spread inoculum on the agar surface, one tube of beads should be used for each

individual plate.

2. After the beads have been added, the plates can be stacked up to six plates high. The

plates are then shaken 10 times from side-to-side. Turn the stack of plates 1⁄4 turn, and

again shake 10 times from side-to-side. Repeat this procedure (1⁄4 turn and 10 shakes) two

more times, so that the beads are shaken a total of forty times.

3. Turn the plates over (upside down), and tap the beads into the lid.

4. Aseptically dump the beads into a labeled bead container (mold and bacteria beads must

be labeled and collected separately), which should be considered contaminated one plate

at a time, replacing the lid as quickly as possible to prevent contamination.

References

1. http://serc.carleton.edu/microbelife/k12/LIMW/dilution.html

2. http://www.qbiogene.com/businessdivisions/platformnews/news0005-b.shtml

3. http://www.genlantis.com/objects/catalog/Product/extras/C400050.pdf

4. EPA - SHEM Chemical Hygiene Plan
MOP 6562

Revision 1

February 2013

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Miscellaneous Operating Procedure (MOP) 6562:

Preparing Pre-Measured Tubes with Aliquoted Amounts of

Phosphate Buffered Saline with Tween 20 (PBST)

Prepared by: __________________________________________ Date: 2/12/2013

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 2/12/2013

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 2/12/2013

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6562

Revision 1

February 2013

Page 2 of 6

MOP 6562

TITLE: PREPARING PRE-MEASURED TUBES WITH ALIQUOTED AMOUNTS

OF PHOSPHATE BUFFERED SALINE WITH TWEEN 20 (PBST)

SCOPE: This MOP provides the procedure for preparing PBST.

PURPOSE: This procedure will ensure that that the PBST is prepared correctly and that all

measured tubes are filled aseptically.

1.0 PREPARING STERILE PHOSPHATE BUFFERED SALINE WITH TWEEN 20

(PBST)

Phosphate Buffered Saline with Tween 20 (PBST) is prepared 1 L at a time in a 1 L flask.

1. Add 1 packet of SIGMA Phosphate Buffered Saline with Tween 20 (P-3563) to 1 L of

deionized (DI) water.

2. Shake vigorously to mix until dissolved.

3. Label bottle as “non-sterile PBST” and include date and initials of person who made

PBST.

4. Filter sterilize into two 500 mL reagent bottles using 150 ml bottle top filter (w/ 33mm

neck and .22 μm cellulose acetate filter) for sterilization. Complete this by pouring the

liquid into the non-sterile PBST into the top portion of the filtration unit 150 ml at a time,

while using the vacuum to suck the liquid through the filter. Continue to do this until 500

ml have been sterilized into a 500 ml bottle. Change bottle top filter units between each

and every 500 ml bottle.

5. Change label to reflect that the PBST is now sterile. Include initials and date of

sterilization. The label should now include information on when the PBST was initially

made and when it was sterilized and by whom.

6. Each batch of PBST should be used within 90 days.

2.0 PREPARING 20 ML/5 ML PBST TUBES FOR USE DURING

EXPERIMENTATION

Twenty (20) ml or five (5) ml of the prepared PBST will be added to each sterile 50-ml

conical tube as detailed below. Each flat of conical tubes contains 25 tubes, so one 500 ml

sterile bottle of PBST should fill approximately one flat when 20 ml tubes are needed and

four flats when 5 ml tubes are needed.
MOP 6562

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February 2013

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1. Prepare the hood by wiping down with ethanol, followed by bleach, followed by DI water

and a clean Kimwipe or Techwipe. Then stock the hood with the following items if they

are not already there:

- The flats of sterile conical tubes you need to fill with PBST.

- Sufficient bottles of sterile PBST to fill these tubes.

- Ample 25 ml serological pipettes (at least 3 per flat) for 20 ml transfers and 10

ml serological pipettes for the 5 ml transfers.

- Serological pipetter (automatic, hand-held pipette).

- Burner and striker.

2. Light the burner and adjust the flame for a width adequate to flame the lips of the PBST

bottles.

3. Take one flat of sterile conical tubes and loosen each cap on the outside edges (about 1⁄2

turn).

4. Open a serological pipette and insert into the serological pipetter, taking care to not touch

the tip to any surface.

5. Hold the pipetter with the first three fingers of your right (or dominant) hand. With your

left hand (or non-dominant hand), pick up a bottle of the PBST and use the bottom of

your right hand to unscrew the lid. Place the lid upside down on the benchtop and

quickly flame the lip of the bottle. Turn the bottle and repeat, taking care to thoroughly

flame the lip without getting the glass so hot that it shatters.

6. Inset the tip of the pipette into the bottle and fill to the 20 ml line. Flame the bottle lip

and place the bottle on the benchtop.

NOTE: If the tip of the pipette touches the outside of the bottle or any other

surface in the hood, consider it contaminated. Discard the pipette

and reload a new one.

7. Quickly pick up one of the tubes that you have loosened the cap on, and use the bottom

of your right hand to remove the cap. Completely discharge the entire pipette into the

tube, taking care to not touch anything with the tip of the pipette. Recap the tube and

place back into the flat (the lid does not have to be tight – you will tighten the lids after

you have completed filling the 10 outside tubes).

NOTE: If the tip touches the outside or rim of the tube (or any other surface

in the hood), consider the tube and pipette contaminated. Discard

both the tube and the pipette.
MOP 6562

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February 2013

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8. Pick up the PBST bottle and flame the lip. Repeat Steps 6 and 7 until all 10 of the tubes

on the outside of the flat have been filled. Flame the lip of the PBST bottle and replace

the cap. Slide the used pipette back into the plastic sleeve and put to the side of the hood

for disposal. Then tighten the lid of each tube you just filled. But rather than placing it

back into its original spot in the flat, switch it for the empty tube from the next row.

When this has been completed, go around the outside of the flat again and loosen the lids

of these 10 tubes. Repeat steps 4 through 7 to fill and cap these tubes.

9. This same procedure is used to fill the middle row of tubes from the flat, and if more

than one flat of tubes is being filled, can be done at the same time as the outside rows of a

second flat.

10. When all tubes have been filled, label each flat as follows, and place on the shelf in room

E390B:

“PBST Tubes (20 ml or 5 ml)”

Date prepared

Your initials

11. These tubes should be made at least 14 days before they need to be used so that they can

be verified as sterile. Any tubes that are cloudy or that have any floating matter/turbidity

should be discarded. The tubes are stable for and should be used within 90 days.

3.0 CLEANUP FOR 20 ML/5 ML PBST TUBES

1. Dispose of the used pipettes in the nonregulated waste.

2. Plug in the serological pipetter so that it can recharge.

3. Replace any unused PBST in the liquid containment on the shelf. Make sure that the

bottle is labeled as having been opened (date opened and initials of whomever used it).

4. Turn off the burner.

5. Wipe down the hood benchtop with ethanol, followed by bleach, followed by DI water

and a clean Kimwipe or TechWipe.

4.0 PREPARING 900μL PBST TUBES FOR USE DURING EXPERIMENTATION

1. Prepare the hood by wiping down with ethanol, followed by bleach, followed by DI water

and a clean Kimwipe or Techwipe. Then stock the hood with the following items if they

are not already there:
MOP 6562

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- A sterile beaker of microcentrifuge tubes.

- Sufficient tubes of sterile PBST to fill these tubes (PBST may be aseptically

transferred to 50 ml conical tubes for an easier aseptic transfer to the

microcentrifuge tubes- it is easier than working from a 500 ml reagent bottle.

Make certain that these 50 ml conical tubes are labeled to when the PBST was

made, sterilized, etc.).

- 1000 μL micropipette.

- 1000 μL sterile pipette tips

- Microcentrifuge tube racks.

- Labeled beaker or waste container used to hold non-regulated waste, such as

tips, under the hood.

2. Carefully remove the microcentrifuge tubes one at a time from the beaker and close the

top on each one before placing it in the tube rack. Place the tubes in the rack skipping

every other row. Fill up two racks doing this.

3. Add 900 μL of PBST to the microcentrifuge tubes by aseptically transferring the PBST

from the sterile 50 ml conical tube containing the PBST. Do this by using the 1000 μL

micropitte and tips. Change tips whenever after two rows of tubes are completed or

whenever a contamination event (such as touching the outside of the 50 ml tube or the

microcentrifuge tube) occurs. Put the dirty tips in the beaker or container used to contain

waste (tips, tubes) in the hood. If any 900 μL tubes are contaminated during the transfer,

dispose of them in the waste container used to hold tips under the hood. If a new box of

tips has to be opened, make certain the date it was opened and initials of the person who

opened it are clearly labeled on the box.

4. After both racks are full, carefully move all the tubes from one rack to fill in the empty

rows on the other rack. In this manner, one rack should be completely filled with tubes at

this point.

5. Label the rack of tubes as “Sterile 900 μL PBST Tubes”, along with the name of the

person who completed the transfer, along with the date. Also, include the date that the

original stock of PBST was made and the date it was sterilized, along with the initials of

the person who completed those steps.

5.0 CLEANUP FOR 900μL PBST TUBES

1. Dispose of the waste that was put in the labeled beaker or waste container (micropipette

tips and tubes) in the nonregulated waste. Then, place this beaker in the “To be

decontaminated via sterilization- contaminated glassware” bin or if it is a disposable

container, then it can be put in the non-regulated waste container.

2. Put the unused sterile tips and the micropipetter back in its original location.
MOP 6562

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3. Replace any unused 50 ml conicals of PBST in the liquid containment on the shelf.

Make sure that the tube is labeled as having been opened (date opened and initials of

whomever used it). If the tube could possibly be contaminated in any way, dispose of it

in non-regulated waste.

4. Wipe down the hood benchtop with ethanol, followed by bleach, followed by DI water

and a clean Kimwipe or TechWipe.
MOP 6563

Revision 2

November 2012

Page 1 of 6

Miscellaneous Operating Procedure (MOP) 6563:

Swab Streak Sampling and Analysis

Prepared by: __________________________________________ Date: 11/15/2012

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6563

Revision 2

November 2012

Page 2 of 6

MOP 6563

TITLE: SWAB STREAK SAMPLING AND ANALYSIS

SCOPE: This MOP provides the procedure for the process of completing a swab streak

plate and subsequent qualitative analysis.

PURPOSE: This procedure will ensure that the swab streak plate sampling and analysis

methods are standardized and that the collection and plating of samples are

free from contamination.

1.0 PREPARING THE MATERIALS

There are two types of prepared swabs that can be used in this procedure:

Environmental Transport Swabs – purchased swabs that are individually packaged

and pre-sterilized.

In-house Sterilized Swabs – swabs placed into autoclave pouches and sterilized

using a 1-hour gravity cycle.

This procedure requires the following materials and equipment:

 Tryptic soy agar (TSA) media plates

 32 C incubator

 Nitrile (non-sterile) gloves

 Sharpie for writing on plates

2.0 COLLECTING AND PLATING SAMPLES

The procedure for collecting and plating samples is dependent on the type of swab being

used. Appropriate PPE should be worn in both cases and includes a lab coat, nitrile gloves,

and safety glasses.
MOP 6563

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November 2012

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2.1 Environmental Transport Swabs

2.1.1 Collection of Environmental Transport Swab Sample

1. Break the seal on the individually packaged and sterile swab. Collect the specimen with

the swab applicator as detailed in the specific test protocol, then replace the swab in the

tube.

2. Label the tube with what is being swabbed (sample ID), the date, time, and initials of the

person performing the procedure.

3. Place the swab into a secondary container, such as a sterile bag, and label the bag with

the same information placed on the tube label.

4. Transport the sample(s) to the Microbiology Laboratory for processing.

2.1.2 Plating of Environmental Transport Swab Sample

1. When the sample is received in the Microbiology Laboratory, label one TSA plate using

a Sharpie with the information from the swab packaging. Verify that the sample ID and

date match.

2. Place labeled plates and swab samples under the biological safety cabinet. Remove the

sample swab from the secondary container and the tube. Press onto the plate in an Sstroke motion, turning the swab as it is plated to ensure that all of the surface area of the

swab touches the plate. Press firmly, but not so hard that the surface of the media is

broken.

3. Replace the swab into its tube and discard in the non-regulated waste container.

4. Repeat steps #1 through #4 for each sample.

5. Label three TSA plates as Swab Blank A, Swab Blank B, and Swab Blank C. These

plates will serve as negative controls for the swabs.

6. Open a new/unused Environmental Transport Swab and use it to plate the three blank

plates as detailed in Step #2.

7. Stack the triplicate plates media side up and place in a 35 C ± 2 C incubator for at least

18 hours. Note the time the plates were placed in the incubator.
MOP 6563

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2.2 In-house Sterilized Swabs

When In-house Sterilized Swabs are being used to collect samples, they need to be plated

immediately (unlike the Environmental Transport Swabs which are transported back to the

Microbiology Laboratory for plating). Therefore, prior to travelling to the sample site, collect

the following materials and supplies which will be needed:

 One TSA media plate (in a media bag) per sample to be collected plus three additional

plates to be used as negative controls for swab blanks.

 One In-house Sterilized Swab (in their autoclave pouches) per sample to be collected, one

swab for the control plates, plus a few extras.

 Sharpie for labeling plates.

Use the following procedure to collect and plate samples.

1. Once at the sample collection site, take the TSA plates out of the media bag and label one

plate for each sample with what is being swabbed (sample ID), date, time, and initials of

the person performing the procedure.

2. As carefully and as aseptically as possible, remove the swab from the autoclave pouch by

the stick end. Be sure and not touch the swab end to anything but the sample. If the

swab’s sterility is compromised, dispose of the swab and use one of the extras.

3. Collect the specimen with the swab applicator as detailed in the specific test protocol.

4. Press onto the plate in an S-stroke motion, turning the swab as it is plated to ensure that

all of the surface area of the swab touches the plate. Press firmly, but not so hard that the

surface of the media is broken. Because these samples are being plated in the open air

and not in a biological safety cabinet, be certain to limit the time that the lid is removed

from the TSA plate.

5. Replace the swab into the autoclave pouch it came in and discard in the non-regulated

waste container.

6. Repeat steps #1 through #6 for each sample.

7. Label three TSA plates as Swab Blank A, Swab Blank B, and Swab Blank C. These

plates will serve as controls for both the swabs and the TSA.

8. Open another in-house sterilized swab from the autoclave pouch and use it to plate the

three blank plates as detailed in Step #2.
MOP 6563

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9. Put the TSA plates back into the media bag and transport to the Microbiology

Laboratory.

10. When received by the laboratory, the plates should be stacked media side up and placed

in a 35 C ± 2 C incubator for at least 18 hours. Note the time the plates were placed in

the incubator.

3.0 ANALYZING THE SAMPLES

The Swab Results Template, which follows this section, is used to record the results of the

sampling. Some quantities of samples may require more than one form. Make certain that

the data is filled in completely on each page. The analyst will use the information on the

TSA plates to fill in the following blanks at the top of the form:

 Swab samples taken on: (date)

 Swabbed by: (person)

 Plating completed on: (date)

 Plated by: (person)

The following procedure is used to analyze the samples and complete the remainder of the

Swab Results Template form.

1. Fill in the final two sections at the top of the form: Plate results read on and Results

read by.

2. Observe the agar surface on the plates and note the sample IDs on the first three lines in

the Sample column.

3. For each plate, check whether there was growth (G) or no growth (NG). Growth is

indicative of an organism(s) being present, and should be described on the form. Be as

detailed as possible, noting colony morphology (size, shape, color and any other

distinctive things that can be seen concerning the growth).

4. The Swab Results Template form serves as the sample report and should be provided to

the Project Manager.
MOP 6563

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Swab Results Template

Swab samples taken

on:

Swabbed by:

Plating completed on: Plated by:

Plate results read on: Results read

by:

Sample Name Result If growth, describe

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

G  NG 

Controls Result If growth, describe

Swab blank A G  NG 

Swab blank B G  NG 

Swab blank C G  NG 

Key

G = Growth.

NG = No Growth.

All plates are plated in triplicate resulting in sample identification of “A”, “B”, and “C”.
MOP 6565

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November 2012

Page 1 of 4

Miscellaneous Operating Procedure (MOP) 6565:

Filtration and Plating of Bacteria from Liquid Extracts

Prepared by: __________________________________________ Date: 11/15/2012

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6565

Revision 4

November 2012

Page 2 of 4

MOP 6565

TITLE: FILTRATION AND PLATING OF BACTERIA FROM LIQUID

EXTRACTS

SCOPE: This MOP outlines the procedure for filtration and subsequent cultivation

of bacterial spores from a liquid extract.

PRUPOSE: This method is deployed when results from spread-plate methods yield

less than 30 colony-forming units (CFU) per plate. This method allows a

lower limit of detection for bacterial recovery/survivorship assays. This

method can also be used to analyze liquid samples such as decon rinsates.

Materials:

 Petri dishes with appropriate agar

 0.2 μm pore-size disposable analytical filter units (2-3 per sample)

 P1000 pipette and sterile tips

 Sterile forceps

 Pipettman and sterile serological pipettes

1.0 PROCEDURE

1. For each liquid sample to be analyzed, gather the required number of disposable

analytical filter units and Petri dishes containing the desired sterilized/QC’d media.

NOTE #1: For analysis of 5 to 30 ml extracts, 1 ml and remainder should be

filtered; for 31 to 200 ml samples, 1 ml, 10 ml, and remainder should

be filtered; for samples over 200 ml, more filter samples may be

needed.

NOTE #2: For previously plated samples where 10 – 19 CFU were observed,

replating using a 400 μl inoculum, and plates where 20 – 29 CFU

were observed, replating using a 200 μl inoculum can be executed

rather than filter plating. For inoculua greater than 200 μl, a sterile

spreader should be used rather than the bead method).
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2. Label plates.

3. Vortex liquid extract vigorously for 2 minutes, using 10 second bursts. (for larger

volume samples, a vigorous mixing by shaking of the sample container can be

substituted for vortex mixing)

4. Using a P1000, sterile tip, and aseptic techniques, immediately following vortexing,

pipette 1 ml of the extract into one of the filter units.

5. Apply vacuum to the filter unit, to pull the liquid through the filter and collect the

spores on the surface of the filter.

6. Using a sterile serological pipette, rinse the filter unit by pipetting 10 ml of sterile

deionized water along the inner sides of the unit while it is under vacuum.

7. Aseptically remove the filter from the filter apparatus using sterile forceps, and lay

the filter onto the agar surface within the Petri dish (spore side up).

8. Vortex the liquid extract vigorously for 10 seconds.

9. Use the appropriate volume serological pipette to transfer the remaining aliquots into

their respective filtration units (one at a time).

10. Repeat steps 5 through 7 taking time to vortex or mix the sample 10 seconds

immediately before removing an aliquot.

Important: Be sure to note and record the volume of the “remainder” sample.

11. Incubate all plates at the optimal growth temperature for the organism used for 16 –

28 hours.

12. Enumerate and record the number of CFU on each plate.

2.0 DATA CALCULATIONS

Utilize the following equation to determine the total abundance of recovered spores:

filtered

Extract

V

V

CFUN 
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Where, N is the total number of spores recovered in the extract, CFU is the abundance of

colonies on the agar plate, VExtract is the total volume of the extract (before any aliquots

were removed), VFiltered is the volume of the extract filtered.
MOP-6567

Revision 1

November 2012

Page 1 of 4

Miscellaneous Operating Procedure (MOP) 6567:

Recovery of Bacillus Spores from Wipe Samples

Prepared by: __________________________________________ Date: 11/15/2012

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP-6567

Revision 1

November 2012

Page 2 of 4

MOP 6567

TITLE: RECOVERY OF BACILLUS SPORES FROM WIPE SAMPLES

SCOPE: This MOP outlines the procedure for recovering Bacillus spores from wipe

samples.

PURPOSE: To aseptically extract and quantify Bacillus spores from wipe samples in order to

determine viability and obtain quantifiable data.

Materials:

 PPE (gloves, lab coat, safety goggles)

 Biological Safety Cabinet (Class II)

 pH-amended bleach

 Deionized water

 70% solution of denatured ethanol

 Kimwipes

 Disbatch®bleach wipes

 Non-regulated waste container

 50 mL sterile conical tubes containing 20 mL of sterile phosphate buffered saline with Tween

20 solution (PBST) (MOP 6562)

 Vortex mixer

 Cart

 Wire or foam rack for 50mL conical tubes

 Tryptic soy agar plates

 900 μL tubes of sterile PBST

 Pipettor and pipette tips for dilutions

 Incubator set to appropriate growth temperature for target organism (35°C or 55°C)

 Light box for counting colonies

 Lab notebook

 QAPP for project that is utilizing the wipe samples
MOP-6567

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1.0 PROCEDURE

1. Begin by donning PPE (gloves, lab coat, and protective eyewear).

2. Obtain wipe samples that may contain Bacillus spores. Wipe samples should be received as

one wipe/sponge in a sterile 50 mL conical tube delivered in secondary containment. Make

certain that all of the samples are labeled. Review any chain of custody forms that may

accompany the samples to ensure that all of the labels are consistent and that there is no notable

variation in the samples. If variation has occurred, make a note of it in the notebook.

3. Clean the workspace (biological safety cabinet) by wiping surfaces with pH-amended bleach,

next with deionized water, and lastly with a 70-90 % solution of denatured ethanol. Wipe with

a Kimwipe to remove any excess liquid. Make sure the workspace is clean and free of debris.

Gather all necessary items to perform the task, place these items on a clean cart beside the

biological safety cabinet, within arm’s reach so that, once the procedure has begun, the task

may be performed without interruptions.

4. Discard gloves and replace with fresh pair.

5. One at a time, under the biological safety cabinet, remove the sample tube containing the wipe

sample from the secondary containment bag in which it arrived. Using the Dispatch®bleach

wipes, wipe each sample tube with one wipe, and then wipe it with a clean Kimwipe. Discard

the used bleach wipe and the used Kimwipe in the secondary containment bag and place them

in the non-regulated waste container. Remove gloves and don a fresh pair of gloves. Repeat

this procedure for every sample. After each sample has been cleaned, place the tubes

containing the wipe samples in an appropriate sized wire or foam rack to hold the tubes in an

upright, vertical position.

6. Leaving the tubes in the rack underneath the biological safety cabinet, aseptically add 20 mL of

PBST solution (this should be in a pre-measured, sterile conical tube, per MOP 6562) to each

sample tube containing a wipe, one a time. Remove the rack containing wipe samples from

hood when all samples have had the PBST added. Place the rack with the samples on the cart.

7. Using the procedure to clean the biological safety cabinet, as found in Step 3, clean the

biological safety cabinet again. Afterwards don a fresh pair of gloves.

8. Using a vortex mixer, agitate the wipe samples, four at a time, in a biological safety cabinet, for

ten second bursts for two minutes total. Make certain to clean the biological safety cabinet

after each set of four samples and change gloves between each set of samples.

NOTE: The reason that four samples are done at one time is to limit the time between

agitation and plating. The samples need to be processed immediately after
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agitation, and agitation of more than four samples at a time leaves too much

time between agitation and spread plating.

9. Using tryptic soy agar media plates that are appropriately labeled with the sample number,

dilution set and date, complete dilution plating for the wipe samples immediately after the two

minute agitation step is completed. The samples should also be agitated again for ten seconds

directly prior to removing an aliquot from the sample tube. Each dilution tube should also be

agitated for ten seconds prior to removal of aliquots. Dilutions should be completed using the

techniques and methodology as described in MOP 6535a, and the 900 μL tubes should be

made with sterile PBST to stay consistent with materials/solutions. Plating in this manner

should be repeated for all samples, with any changes in protocol noted in the lab notebook.

10. Once the dilution plating has been completed, the plates are to be placed in an incubator. For

non-thermophilic Bacillus species, the plates should be placed at 35°C +/- 2° C for 12-24

hours. For thermophilc Bacillus species, such as Geobacillus stearothermophilis, the plates

should be incubated at 55°C ±2 °C for 12-24 hours. The target Bacillus organism that will be

used for the wipe samples will be specific to the project and noted in the QAPP.

11. After the plates have incubated for a sufficient amount of time (12-24 hours) and the growth

from any Bacillus colonies are quantifiable, the colonies should be manually counted using the

light box and the data should be properly recorded as dictated per project by the QAPP. All

results will be checked for quality assurance and all data will be reported to the proper

personnel as listed in the QAPP.
MOP 6570

Revision 3 

March 2013 

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Miscellaneous Operating Procedure (MOP) 6570: 

Use of Steris Amsco Century SV 120 Scientific Prevacuum 

Sterilizer 

Prepared by: __________________________________________ Date: 3/21/2013 

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader 

Reviewed by: __________________________________________ Date: 3/21/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 3/21/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP 6570

Revision 3 

March 2013 

Page 2 of 4 

MOP 6570 

TITLE: USE OF STERIS AMSCO CENTURY SV 120 SCIENTIFIC 

PREVACUUM STERILIZER 

SCOPE: Basic instructions for use of the large Steris autoclave. 

PURPOSE: To outline proper procedural use of the autoclave, using preprogrammed 

cycles, to effectively sterilize items, while complying with quality control 

standards. 

Materials: 

 Amsco Century SV 120 Scientific Prevacuum Sterilizer 

 Items to be sterilized (liquids, solids, waste, etc) 

 Pouches to contain materials during sterilization and maintain sterility until use 

 Aluminum foil 

 Autoclave indicator tape 

 Sterilization verification ampoules (such as Raven ProSpore Ampoules) 

 Thermally resistant gloves 

 De-Ionized (DI) water 

1.0 PROCEDURE

1.1 Start Up 

1. Turn on the autoclave. The power switch is located behind the door in the top right 

corner. The digital touch screen on the front of the unit will power up and indicate 

that a memory test is in progress. 

2. After the memory test is complete, the device will request that it be flushed. This 

should be conducted daily to minimize scaling inside the boiler. The flush valve is 

located behind the door on the bottom, left of the device (yellow handle). Move the 

valve to the open position and then press the “Start Timer” button on the touch 

screen. The flush will run for 5 minutes and will alert at completion with a single 

chime. 

3. Once the flush is complete, close the flush valve and press the “Continue” button on 

the touch screen. The screen should then return to its default menu which has 2 

choices “Cycle Menu” and “Options”
MOP 6570

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March 2013 

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1.2 Basic Operation 

1. Prepare any items that need to be sterilized. The items must be carefully wrapped or 

sealed in sterilization pouches in order to maintain sterility when removed from the 

autoclave. Examples of this include: wrapping any orifices with aluminum foil, 

placing whole items in autoclave pouches, loosely applying a cap on a bottle (to allow 

for the pressure changes inside). 

2. Once prepared, each item should be outfitted with a sterility indicator such as a small 

piece of autoclave indicator tape; or by utilizing an autoclave pouch with a built-in 

sterility indicator strip. These indicators provide a visual verification that the 

sterilizing temperature (121°C) was reached. 

3. To add items to the autoclave, open the autoclave door by pressing down on the foot 

pedal on the bottom right corner on the front of the device. 

4. Place items that need to be sterilized into the autoclave, adding or moving racks to 

accommodate the load. If liquids are being autoclaved, then they must have secondary 

containment (usually a large plastic autoclave-safe tray) to contain any fluids in the 

event of a leak, spill or boil-over. Add an indicator ampoule to the first autoclave 

cycle of the day, regardless of the type of cycle. 

5. Once the autoclave is loaded, press the foot pedal to close the autoclave door. 

6. Once the door is sealed, a menu of the cycles can be seen by pressing the button on 

the touch screen labeled “Cycle Menu”. Then choose the appropriate cycle by 

touching the corresponding button. If the cycle chosen is the one desired for the 

sterilization process, press the “Start Cycle” button. Otherwise, press “Back” to 

return to the prior menu screen. 

7. After the cycle has started, the type of cycle, the number of the cycle, the items 

placed in the autoclave during the cycle, the time, whether or not an indicator 

ampuole was included in the load, and the initials of the person starting the cycle 

must be recorded in the autoclave log book, located in the drawer across form the unit 

labeled “Autoclave Supplies.”

8. Quality control (QC) indicator ampoules, usually Raven ProSpore Ampoules with 

Geobacillus stearothermophilus (at a concentration 10E6), are added to one cycle 

each day to ensure that the autoclave is functioning properly. These ampoules are 

used according to manufacturer’s instructions. These ampoules must be properly 

labeled with the date in which they were autoclaved and the initials of the individual 

that completed the cycle. At the beginning of each week, a positive control ampoule 

must be processed, where the ampoule is placed directly into the 55°C water bath, 

without being autoclaved. The positive control indicator ampoule should change 

from purple to yellow in color, indicating growth. All test ampoules should be placed 

in a water bath following the end of the cycle in which they are run. These ampoules 

should not change color (from purple to yellow, but instead should remain a purple to 
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purple-brown color). Ampoules should be checked at both 24 and 48 hour intervals 

for growth and then finally recorded and disposed of after 48 hours. All QC 

information concerning ampoules should be recorded in the autoclave notebook. 

9. Upon completion of any cycle, the autoclave will alarm with a repeating beep for 

approximately one minute. Any time after this alarm starts, it is safe to open the main 

door (take caution because the steam escaping the chamber will be very hot when the 

door is opened). The contents from the autoclave will be very hot; use protection to 

remove items from the autoclave (thermally resistant gloves).

10. Place the contents of the autoclave in an appropriate place to cool, and close the 

autoclave door using the foot pedal. 

1.3 Cycles 

1.3.1 Gravity Cycles 

Gravity cycles are used to sterilize glassware and other utensils, which are not submerged 

in nor contain any volume of liquid. These cycles are typically used for “dry” materials.

Currently there are two different gravity cycles programmed for daily operations: a 1-

hour cycle and a 30-minute cycle. The time that the chamber is held at the sterilization 

temperature (121 °C) is the only difference between these two cycles. The different 

sterilization times allow for the compensation of the various sizes of materials and more 

resilient organisms. The 30-minute cycle is primarily used for a small quantity of 

material. The 1 hour cycle is used for large loads or items containing a large amount of 

contamination. The 1 hour cycle is recommended for inactivation of gram positive sporeforming bacteria. 

1.3.2 Liquid Cycles 

Liquid cycles are used to sterilize a variety of liquids and solutions. The solutions are 

typically mixed prior to sterilization. It is important to have secondary containment to 

contain any fluids in the event of a leak, spill or boil-over. The 30-minute liquid cycle is 

used to sterilize small volumes of liquid (usually less than 2L total). When attempting to 

sterilize any volume larger than 2L, the 1-hour liquid cycle should be used to ensure 

complete sterilization. The 1-hour liquid cycle is the preferential cycle used as the 

destruction cycle for waste. In the event of materials (liquid or otherwise) being 

contaminated/exposed microorganisms, the 1-hour liquid cycle will be used as the initial 

means of decontamination. When completing a decontamination cycle, if there is no 

liquid inside of a container, then deionized water must be added to the container or the 

item must be submerge prior to the start of the cycle. Only items that are being 

decontaminated can go in destruction cycles. Decontamination cycles cannot be mixed 

with sterilization cycles. 
MOP 6572

Revision 2

November 2012

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Miscellaneous Operating Procedure (MOP) 6572:

Recovery of Spores from Vacuum Sock Samples

Prepared by: __________________________________________ Date: 11/15/2012

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 11/15/2012

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 11/15/2012

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6572

Revision 2

November 2012

Page 2 of 4

MOP 6572

TITLE: RECOVERY OF SPORES FROM VACUUM SOCK SAMPLES

SCOPE: This MOP outlines the procedure for recovering spores from vacuum sock samples

PURPOSE: To aseptically extract and quantify spores from vacuum sock samples in order to

determine viability and obtain quantifiable data

MATERIALS

 PPE (gloves, lab coat, safety goggles)

 Biological Safety Cabinet (Class II)

 pH-amended bleach

 Deionized water

 70% solution of denatured ethanol

 Kimwipes

 Disbatch®bleach wipes

 Non-regulated waste container

 3 oz. sterile specimen cup containing 20 mL of sterile phosphate buffered saline with Tween 20

solution (PBST) (MOP 6562)

 Sterile scissors

 Vortex mixer

 Cart

 Tryptic soy agar plates

 900 μL tubes of sterile PBST

 Pipettor and pipette tips for dilutions

 Incubator set to appropriate growth temperature for target organism (35 °C or 55 °C)

 Light box for counting colonies

 Lab notebook

 QAPP for project that is utilizing the vacuum sock samples
MOP 6572

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November 2012

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1.0 PROCEDURE

1. Begin by donning PPE (gloves, lab coat, and protective eyewear).

2. Obtain vacuum sock samples that may contain Bacillus spores. Vacuum sock samples should

be received as one vacuum sock in a sterile 5.5’ x 9 bag secondarily contained in a 10’ x 15’

bag. Make certain that all of the samples are labeled. Review any chain of custody forms that

may accompany the samples to ensure that all of the labels are consistent and that there is no

notable variation in the samples. If variation has occurred, make a note of it in the notebook.

3. Clean the workspace (biological safety cabinet) by wiping surfaces with pH-amended bleach,

next with deionized water, and lastly with a 70-90 % solution of denatured ethanol. Wipe with

a Kimwipe to remove any excess liquid. Make sure the workspace is clean and free of debris.

Gather all necessary items to perform the task, place these items on a clean cart beside the

biological safety cabinet, within arm’s reach so that once the procedure has begun the task may

be performed without interruptions.

4. Discard gloves and replace with fresh pair.

5. Label a 3 oz. specimen cup to match the vacuum sock sample ID. The specimen cup contains

20 mL of sterile PBST.

6. When extracting samples, handle one sample at a time from start to finish. Begin by removing

the inner bag from the outer bag. Discard the outer bag in the non-regulated waste container.

Place the inner bag containing the vacuum sock under the hood. Loosen the cap on the 3 oz.

specimen cup and open a pack of sterile scissors. Open the bag and remove the sock, careful

not to touch the white part. Roll the non-sterile blue portion of the vacuum sock onto the

smaller cardboard ring. Dispose of the larger cardboard ring. Wet the vacuum sock by holding

the upper blue portion of the vacuum sock (around the smaller cardboard ring) and dipping the

lower 1-inch of the vacuum sock into the PBST. The vacuum sock will be allowed to absorb

the PBST for a few seconds. After wetting, the vacuum sock will be lifted up just above the

opening of the specimen bottle, and a 1-inch vertical slit will be cut up the center from the

bottom of the sock using sterile scissors (a new pair of scissors should be used for each

sample). The vacuum sock is then cut horizontally from side to side, about 1 inch from the

bottom allowing the two pieces to fall into the specimen bottle. The vacuum sock should be cut

only where the sock has been wetted. Repeat the dip/cutting procedure until the entire

collection portion of the sock has been excised. The upper top blue portion of the vacuum sock

will then be discarded. Place used scissors in a discard pan. After samples are all extracted,

scissors will be immediately autoclaved using a one hour gravity destruction cycle in

preparation for use with the next sample batch. Remove gloves and don a fresh pair of gloves.

Repeat the extraction procedure for every sample, while maintaining aseptic technique.

7. After cutting all vacuum sock samples, all specimen cups (up to sixty samples at a time) should

be loaded into the sample cup holder of the orbital shaker-incubator. The samples are then
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agitated in the shaker incubator at 300 rpm for 30 minutes at room temperature. The samples

are then removed from the shaker incubator and brought to the Biological Safety Cabinet for

dilution plating.

8. Using the procedure to clean the biological safety cabinet, as found in Step 3, clean the

biological safety cabinet again. Afterwards don a fresh pair of gloves.

9. Using tryptic soy agar media plates that are appropriately labeled with the sample number,

dilution set and date, complete dilution plating for the vacuum sock samples immediately after

the thirty minute agitation step is completed. The samples should also be agitated again for ten

seconds directly prior to removing an aliquot from the specimen cup. Each specimen cup

should also be agitated for ten seconds prior to removal of aliquots. Dilution-plating should be

carried out according to MOP 6535a. Dilution tubes used in MOP 6535a should contain

PBST to stay consistent with materials/solutions. Repeat procedure for all samples.

10. Once the dilution plating has been completed, the plates should be incubated. For nonthermophilic Bacillus species, the plates should be placed at 35°C ±2 °C for 18-24 hours. For

thermophilc Bacillus species such as Geobacillus stearothermophilis, the plates should be

incubated at 55°C ±2 °C for 18-24 hours. The target Bacillus organism that will be used for

the vacuum sock samples will be specific to the project and noted in the QAPP.

11. After the plates have incubated for a sufficient amount of time (18-24 hours) and the growth is

quantifiable, the colonies should be manually counted with the assistance of a light box. The

data should be properly recorded as dictated per project by the QAPP. All results will be

checked for quality assurance and all data will be reported to the proper personnel as listed in

the QAPP.
MOP 6579 

Revision 4

April 2013

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Miscellaneous Operating Procedure (MOP) 6579: 

Recovery of Bacillus Spores from 37MM Filter Cassettes

Prepared by: __________________________________________ Date: 4/29/2013 

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader 

Reviewed by: __________________________________________ Date: 4/29/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 4/29/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP 6579 

Revision 4

April 2013

Page 2 of 6 

MOP: 6579 

Title: RECOVERY OF BACILLUS SPORES FROM 37MM FILTER CASSETTES 

Objective: To aseptically extract and quantify recovery of Bacillus spores from 37mm filters used to 

sample porous surfaces. 

MATERIALS AND EQUIPMENT 

Equipment: 

 Biological Safety Cabinet (BSC, Class II) 

 Sonicating cleaner (such as Branson 8510, 40KHz) 

 Incubator set to appropriate growth temperature for target organism (35 °C) 

 Light box for counting colonies 

 Vortex mixer 

 Cassette opening tool (SKC cat# 225-8372, www.SKCshopping.com) 

 Specimen cup rack (cat # 5534M1, http://www.medicus-health.com) 

Materials: 

 Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles) 

 1:10 Bleach, pH-amended to 7.0 with acetic acid 

 Dispatch bleach wipes 

 Disposable 500 ml beaker (such as Fisher Scientific cat #14-955-111D) (for discarding used tips 

and beads/spreaders) 

 70% Solution of denatured ethanol 

 Kimwipes 

 Waste container 

 Sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562) 

 Tryptic Soy Agar (TSA) plates 

 Sterile scissors or scalpels, preferably one for each sample 

 Sterile 1500 μL tubes filled with 900μL of sterile PBST 

 15 ml centrifuge tube containing 6 ml of sterile PBST (one for each sample) 

 Autoclaved 2 oz polypropylene jars with lids (cat # J037, cat # L208 

www.containerandpackaging.com)
MOP 6579 

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 Pipettor and pipette tips for dilutions (100 μl, 1000 μl, and 5000 μl.)

 Pall Filter units 0.45 um MCE (VWR; catalog #28143-544) 

 Vacuum Pressure Gauge (McMaster; catalogue # 38545K16) 

 Disposable cell spreaders or glass beads for cell spreading 

 Disposable sterile thumb forceps 

1.0 REMOVING FILTER AND DUST FROM CASSETTE 

1. Begin by donning PPE (gloves, lab coat, and protective eyewear). 

2. Each cassette should be received separately in a small sealable bag (primary containment), then 

placed within secondary containment. Make certain that all of the samples are labeled. Review any 

chain of custody forms that may accompany the samples to ensure that all of the labels are consistent 

and that there is no notable variation in the samples. If discrepancies are found, make a note of each 

one in the notebook. 

3. Clean the workspace (BSC) by wiping surfaces with pH-amended bleach, next with deionized water, 

and lastly with a 70-90 % solution of denatured ethanol. Wipe with a kimwipe to remove any excess 

liquid. Make sure the workspace is clean and free of debris. Gather all necessary items to perform 

the task. 

4. Discard gloves and replace with fresh pair. 

5. For each cassette sample, label one 2 oz polypropylene jar with the sample identifier. 

6. One at a time, under the BSC, remove the conical tube and cassette from the primary containment 

bag in which it arrived. 

7. First wipe down the outside of the conical tube containing the vacuum tubing and adapter with a 

disinfecting wipe, and place it into a rack. Then, aseptically add 5ml of premeasured PBST. Set the 

rack to the side. 

8. Remove the band from around the cassette using a sterile scalpel or sterile pair of scissors. Using 

the Dispatch bleach wipes, wipe each cassette with a wipe, and then a clean kimwipe. Discard the 

used bleach wipe and the used kimwipe in the secondary containment bag and place them in the nonregulated waste container. Remove gloves and don a fresh pair of gloves before proceeding. 

9. With the plug in place on the back side of the filter, remove the red plug from the front filter side of 

the cassette, use a pipette and aseptically dispense about 1mL μl of PBST (taken from the tube 

containing 6ml PBST) into the cassette. Roll the cassette around to allow the liquid to touch all 

surfaces of the inside of the cassette. If there is a large quantity of dust inside, more PBST may be 

needed to dampen the dust before opening the cassette. 

10. Once a visual check has been completed to assure that the dust is dampened enough to prevent its 

aerosolization, use the cassette tool to pry open the top section of the cassette by carefully rotating 

the cassette while using the tool to pry the edges up. Be careful to hold the cassette right side up 

while prying open, so as not to spill the liquid inside. 
MOP 6579 

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11. When the cassette is open, use a pipette to aseptically rinse the inside walls of the cassette with 1 to

2 ml of PBST (additional aliquots from the same 5ml tube used previously), then remove the rinse 

eluent (using the same transfer pipette) and place it in the 2 oz sterile cup labeled with the sample 

identifier. 

12. Use the cassette opening tool to then remove the middle section of the cassette in the same manner 

as the top section, thereby allowing the filter to be removed. Using sterile forceps, aseptically 

remove the filter, being careful not to pick up the support filter underneath, and place it in the 2 oz 

polypropylene jar with the rinse eluent. 

13. Use the remainder of the 6 ml PBST to rinse the walls of the middle and top sections of the cassette 

while holding them over the open 2 oz cup. The entire 6 ml PBST, the filter and as much dust as can 

be rinsed from the cassette should now be inside the 2 oz polypropylene jar. 

14. Discard the cassette sections, support filter, cassette plugs, disposable pipettes and disposable thumb 

forceps into the proper waste. Close the 2 oz jar tightly. Wipe outside of the jar with disinfecting 

wipe and place in specimen cup rack or sample retention bin. 

15. Wipe down the cassette opening tool thouroughly with a disinfecting wipe, and wipe down the 

surface of the BSC as described in Step 3 above. 

16. Change gloves, and repeat from Step 5 for each remaining sample set. 

2.0 PROCESSING 37 MM FILTERS 

1. Place the rack of conical tubes containing the 5mL of PBST with the vacuum tubing and adapters 

into the sonicating bath with a weighted, waterproof, rectangular, flat surface on top of the tubes to 

keep them from floating when submerged. Sonicate the conical tubes for 1 minute. 

2. Remove the tubes from the sonicator and dry them with a kimwipe and wipe them with a 

disinfecting wipe. 

3. Vortex the conical tubes for 2 continuous minutes. 

4. Aseptically transfer the 5mL of PBST from the conical tube samples to the 2oz specimen cup 

containing the filter sample that corresponds with the conical tube sample. The labels on the conical 

tube should match the label on the specimen cup. 

5. After the aseptic transfer of extraction liquid from the conical tube to the specimen cup, using a 10 

ml serological pipette, measure the amount of PBST extraction liquid that has been collected in the 

2oz specimen cup. Record this number. 

6. Place all 2 oz polypropylene cups containing the 37mm filters and 5 ml PBST in the specimen cup 

rack, place a weighted waterproof, rectangular, flat surface on top of the jars to keep them from 

floating when submerged in the sonicating bath. 

7. Place the rack with the cups and the weighted surface into the bath so that they are submerged about 

1 – 2 inches, leaving about 1 inch between the level of the bath water and the jar lid. 
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NOTE: This may require one or two wire racks (test tube racks or additional cup holders) in the 

bottom of the sonicating bath to achieve the correct level for the cups. Make sure the 

sonicating bath is filled with water to the correct level as designated by the line on the 

inside of the bath. A series of open racks is preferable to a basket for this purpose, as 

the baskets provided by the bath manufacturers tend to have more solid walls with 

small holes, which can restrict the sonic waves more than a wire rack. 

8. When the correct level of submersion is achieved, turn on the sonicating bath (no heat) and sonicate 

for 3 minutes, rotating the cup rack within the bath at the 1 min and 2 min mark (pick up, turn 90 

degrees, re-submerge). 

9. Remove the rack of cups from the sonicating bath and place on an absorbent pad. Remove each 2 oz 

cup from the rack, wipe with a kimwipe to dry and place back inside the BSC. 

3.0 FILTRATION AND PLATING OF BACTERIA FROM LIQUID EXTRACTS 

1. Follow MOP 6535a for spread-plating samples in triplicate from the 2 oz cups. 

2. Filtration – detailed in the steps below – is required only if extremely low recovery is expected. 

a. Label three TSA plates with information specific to the sample and amount to be filtered. 

b. Set up filter manifold with three filter unit cups. 

c. Set the vacuum, with proper gauge, to no more than 10 inches Hg (4.9 psi). 

d. Add 10 ml PBST to filter cup. 

e. Add 1 ml sample eluent from sample cup to each filter cup on the manifold. 

f. Open the vacuum valve and vacuum liquid through filter. 

g. Rinse sides of filter cup with 10 ml of sterile PBST, vacuum through again. 

h. Pop off sides of the filter cup by squeezing and lifting. 

i. Use sterile forceps to lift filter and place it on the TSA, making sure good contact with the 

agar surface is made and air pockets are minimized. 

3. Repeat for subsequent sample jars, completing samples in batches of no more than 4. This will help 

to avoid too much settling time between vortexing and plating. 

4. Place all TSA plates in incubator set at 35 oC and incubate 18 – 24 hrs before counting colonies. 
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4.0 CALCULATIONS

For spread plates: total CFU recovered = (mean CFU on plate / 0.1ml volume plated) x (1/dilution 

factor) x (5 ml total in jar) 

For vacuum filter plates: total CFU recovered = (mean CFU on vacuum filter / volume filtered) x 5 ml 

total in jar 
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Miscellaneous Operating Procedure (MOP) 6580:

Recovery of Bacillus Spores from 3M Sponge-StickTM Samples

Prepared by: __________________________________________ Date: 2/12/2013

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 2/12/2013

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 2/12/2013

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6580

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MOP 6580

TITLE: RECOVERY OF BACILLUS SPORES FROM 3M SPONGE-STICKTM

SAMPLES

SCOPE: This MOP provides the procedure for recovering spores from 3M SpongeStickTM samples.

PURPOSE: To extract and quantify bacterial spores from 3M Sponge-StickTM samples

using a highly repeatable procedure.

MATERIALS

 pH-amended bleach

 70-90 % Solution of denatured ethanol

 Deionized (DI) water

 Kimwipes

 3M Sponge-StickTM samples (P/N SSL10NB), hereafter referred to as ‘sponge’

 Seward Stomacher® bags (P/N BA6041/CLR)

 Phosphate buffered saline with 0.05% TWEEN®20 (PBST) (SIGMA-ALDRICH, Co,

P/N P3563-10PAK)

 MicroFunnel Disposable Filter Funnels, Pall Life Sciences (VWR P/N 55095-060) or

Nalgene Sterile Analytical Filter Unit (Fisher P/N 130-4020)

 Disposable polystyrene serological pipettes (5mL and 10mL)

 Tryptic Soy Agar (TSA) plates

 Vortex mixer

 Disposable sterile 10μl loops

 Disposable sterile forceps

 Disposable gloves

 Cell spreaders or glass beads for spreading

 Racks for 15 mL and 50 mL centrifuge tubes

 Sterile, plastic, screw-cap 50 mL centrifuge tubes (e.g. Fisher Cat# 14-959-49A)
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 Sterile, plastic, screw-cap 15 mL centrifuge tubes (e.g. Fisher Cat# 14-959-49D)

 Pipette tips with aerosol filter for 1 mL and 200 μL 

1.0 PREPARATION

Personnel must be familiar with all procedures prior to start.

1.1 Equipment Preparation

a) Begin by donning personal protective equipment (PPE) such as gloves, lab coat,

and protective eyewear.

b) Clean the workspace (Biological Safety Cabinet; BSC) by wiping surfaces with pHamended bleach, next with DI water, and lastly with a 70-90% solution of denatured

ethanol. Allow any excess liquid to dry prior to beginning procedure. Make sure

the workspace is clean and free of debris.

c) Assemble equipment in the BSC as needed: vortex mixer, filtration manifold,

automatic pipettors, tips, racks, etc.

d) Assemble extra supplies, such as stomacher and reagents, near BSC.

1.2 Supply Preparation

a) Unpack shipping containers directly into a BSC.

b) If sponges are not in Stomacher® bags, label one Stomacher® bag for each sponge

and place in a bag rack.

c) Label two sterile 50 mL centrifuge tubes for each sponge sample and place in tube

rack.

d) For each sample, label TSA plates on the agar side of the plate with the sample

number and the appropriate dilution factors, as per MOP 6535a (Serial Dilution:

Spread Plate Procedure to Quantify Viable Bacterial Spores).

e) Label two additional plates for filter-plate analysis.
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2.0 PERFORM SPORE EXTRACTION, ELUTION, AND CULTURE PROCEDURE

2.1 Dislodge Spores from the Sample Sponges

a) Begin by donning a new pair of gloves. All subsequent procedures involving

manipulation of sponges or spore suspensions must be carried out in a BSC.

(Stomaching may occur outside the BSC when samples are double-contained inside

the indicated bags.)

b) If the sponges are not in Stomacher® bags, aseptically transfer each sponge to a

Stomacher® bag (labeled during step 1.2b) using sterile disposable forceps. Change

forceps between samples.

c) Aseptically add 90 mL of PBST to each bag that contains a sponge.

d) Stomach sponges in the PBST by completing the following:

 Make certain the Stomacher® is set to MANUAL. Program the Stomacher®

speed to 260 RPM and the timer to 1 minute.

 Open the Stomacher® door by raising the lid fully upward and back. The

DOOR OPEN icon will be displayed.

 Place the stomacher bag containing the sponge sample into a second stomacher

bag to contain any leakage in the event the primary containment is

compromised. Place the combined bags such that 50 to 60 mm of the top

portions protrude above the bag clamp, while making certain that the sponge

sample rests evenly between the homogenizer paddles.

 Close the door to the Stomacher®. The DOOR OPEN icon will no longer be

illuminated.

 Stomach each sponge for 1 min by pressing the START button.

 When the cycle ends, the Stomacher® will stop. If there is an emergent reason

to stop the stomacher during the 1 minute stomaching period, press the red

button or the power button to do so prior to opening the Stomacher®. Stopping

the Stomacher® by opening the door can damage the equipment.

 Open the door of the Stomacher® and remove the bags containing the sponge.

Grab the sponge from the outside of the bag with your hands. Move the sponge
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to the top of the bag while using your hands to squeeze excess liquid from the

sponge.

 Remove and discard the sponge using sterile forceps.

e) Repeat steps (b) through (d) for all samples.

f) Allow bags to sit for 10 min to allow elution suspension foam to settle before

beginning the concentration step.

2.2 Remove Sponge Elution Suspension

a) Gently mix elution suspension up and down with a 50 mL pipette three times.

b) Split elution suspension volume equally.

 Remove half of the suspension volume (~45 mL) with a sterile 50 mL pipette

and place it in a 50 mL screw capped centrifuge tube.

 Place remaining suspension (~45 mL) into a second 50 mL tube.

c) Record suspension volumes on tubes and data sheet.

d) Repeat steps (a) through (c) for all samples.

2.3 Concentrate Sponge Elution Suspension (Optional)

a) Centrifuge 50 mL centrifuge tubes

 Prior to daily use and before placing tubes into centrifuge, follow MOP 6558

(Centrifuge Cleaning Procedure) for cleaning this equipment.

 Add centrifuge tubes to rotor, evenly distributing weight.

 Centrifuge tubes at 3500 x g for 15 min. Do not use the brake option on the

centrifuge to slow the rotor, as re-suspension of pellet may occur.

b) Carefully remove about 42mL of supernatant with a 50 mL pipette and discard to

leave approximately 3 mL in each tube. The pellet may be easily disturbed and not

visible, so place pipette tip away from the tube bottom or side.
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c) Vortex and sonicate tubes as follows:

 Set vortex mixer to level 10 and touch activation.

 Turn on sonicator water bath.

 Vortex tubes for 30 sec.

 Transfer tubes to sonicator bath and sonicate for 30 sec.

 Repeat vortex and sonication cycles two additional times.

d) Remove suspension from one tube with a sterile 5 mL pipette and place it in the

other tube of the same sample. The combined result is the final sponge elution

suspension.

e) Measure final volume of the final sponge elution suspension with 5 mL pipette

and record on tube and data sheet.

f) Repeat steps (e) through (i) for all samples.

2.4 Serially Dilute and Plate the Final Spore Elution Suspension

a) Use MOP 6535a to serially dilute and plate samples.

NOTE: If the samples are turbid, wide-orifice pipette tips may be used to prevent

clogging of pipette tips.

b) Place all plates in an incubator set at 35 ± 2 oC for a maximum of 3 days. Plates

should be examined within 18-24 hours after start of incubation. Manually

enumerate CFU of target organism and record data.

 If the CFU is <300/plate, record actual number.

 If the CFU is >300/plate, record as “too numerous to count” (TNTC)

2.5 Capture Spores on Filter Membranes and Culture on TSA

Choose one of the following to methods to filter the final spore elution suspension:

a) Complete filter plating using MOP 6565 (Filtration and Plating of Bacteria from

Liquid Extracts).
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b) Complete filter plating using the following method:

1) Place two 0.45 μm (pore-size) Microfunnels on a Pall vacuum manifold (Pall Cat#

15403).

2) Moisten Microfunnel membranes with 5 ml PBST, open vacuum, and vacuum

through the filter. All filtering should be done with a vacuum pressure <20 cm Hg.

3) Make certain that the manifold vacuum valve is closed. Turn on the vacuum.

4) With the vacuum valve closed, place 10 mL of PBST into each filter cup.

5) Add 1.0 mL of the final sponge elution suspension to each filter cup.

6) Open valves and allow the suspension to flow through the filter, close the valve.

7) Rinse the walls of each Microfunnel cup with 10 mL of PBST. Reopen the valve to

allow the suspension to flow through the filter.

8) Close the valve, turn off the vacuum pump. Slowly reopen the valve to equalize the

pressures.

9) Squeeze the walls of the Microfunnel cup gently and separate the walls from the

base holding the filter. Remove each filter membrane with sterile disposable

forceps and place grid-side up on a TSA plate. Make sure that the filter is in good

contact with the surface of the agar. If an air pocket occurs under the filter, use the

sterile forceps to lift the edge of the filter to release the air pocket for better contact

with the agar.

10) Record exact volume of the sponge elution suspension filtered on each plate. It

should be 1 mL. (Greater sample volumes may be used to lower detection limits)

11) Repeat steps (1) through (8) for all each sample.

12) Incubate TSA plates with filter membranes at 35 ± 2 oC for a maximum of 3 days.

Plates should be examined within 18-24 hours after start of incubation. Manually

enumerate CFU of target organism and record data.

 If the CFU is <300/plate, record actual number.

 If the CFU is >300/plate, record as “too numerous to count” (TNTC)
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Miscellaneous Operating Procedure (MOP) 6582: 

Recovery of Bacillus Spores from Trace Evidence Filters 

Prepared by: __________________________________________ Date: 4/29/2013 

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader 

Reviewed by: __________________________________________ Date: 4/29/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 4/29/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP 6582

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MOP: 6582

Title: RECOVERY OF BACILLUS SPORES FROM TRACE EVIDENCE FILTERS 

Objective: To aseptically extract and quantify recovery of Bacillus spores from 3M Trace Evidence 

filters (TEF) used to sample porous surfaces. 

MATERIALS AND EQUIPMENT 

Equipment: 

 Biological Safety Cabinet (BSC, Class II) 

 Incubator set to appropriate growth temperature for target organism (35 °C) 

 Light box for counting colonies 

 Vortex mixer 

 Stomacher (Seward Stomacher®400 Circulator (Seward; catalog #0400/001/AJ) 

 Closure bags (catalog # BA6141/CLR) 

 Stomacher bag rack (Seward catalog # BA6090) 

 Specimen cup rack 

 50 mL Tube rack 

 Materials: 

 Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles) 

 1:10 Bleach, pH-adjusted (pAB) to 7.0 with acetic acid 

 Dispatch bleach wipes 

 Disposable 500 mL beaker (such as Fisher Scientific cat #14-955-111D) (for discarding used tips 

and beads/spreaders) 

 70% Solution of denatured ethanol 

 Kimwipes 

 Waste container 

 Sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562) 

 Tryptic Soy Agar (TSA) plates 

 3M Forensic Vacuum Filter (Trace Evidence Filter) (Precision Data Products, Grand Rapids, MI 

catalog #FF-1) 

 Stomacher bags, closure type (Seward catalog # BA6141/CLR) 
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 Sterile scalpels, preferably one for each sample 

 Sterile 50 mL centrifuge tubes filled with 45 mL of sterile PBST, two for each sample 

 Pipettor and sterile 50 mL pipettes 

 Pipettor and pipette tips for dilutions (100 μl, 1000 μl, and 5000 μl.)

 Disposable cell spreaders or glass beads for cell spreading 

 Disposable sterile thumb forceps 

 Disposable sterile 2 mL bulb pipettors (such as Fisher Sci cat #13-711-7M) 

1.0 REMOVING FILTER AND DUST FROM TRACE EVIDENCE FILTER 

1. Begin by donning PPE (gloves, lab coat, and protective eyewear). 

2. Each TEF should be received separately in a small sealable bag (primary containment), and within 

secondary containment. Make certain that all of the samples are labeled. Review any chain of 

custody forms that may accompany the samples to ensure that all of the labels are consistent and that 

there is no notable variation in the samples. If discrepancies are found, make a note of each one in 

the notebook. 

3. Clean the workspace (BSC) by wiping surfaces with pAB, next with deionized water, and lastly with 

a 70-90% solution of denatured ethanol. Wipe with a kimwipe to remove any excess liquid. Make 

sure the workspace is clean and free of debris. Gather all necessary items to perform the task. 

4. Discard gloves and replace with fresh pair. 

5. For each TEF sample, label one closure-type stomacher bag and two 50 ml centrifuge tubes 

(containing 45 ml PBST each) with the sample identifier. 

6. One at a time, under the BSC, remove the TEF from the primary containment bag in which it 

arrived. Using the Dispatch bleach wipes, wipe each cassette with a wipe, and then a clean 

kimwipe. Discard the used bleach wipe and the used kimwipe in the secondary containment bag and 

place them in the non-regulated waste container. Remove gloves and don a fresh pair of gloves 

before proceeding. 

7. With the plug in place on the back side of the filter, remove the black nozzle cover from the front 

filter side of the cassette, use a pipette and aseptically dispense 5 mL of PBST (taken from the 

centrifuge tubes containing 45 ml PBST) into the cassette (Figure 1). 
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Figure 1. Dispensing 5 mL PBST into TEF nozzle 

8. Replace the black nozzle cover and roll the TEF around to allow the liquid to touch all surfaces of 

the inside of the cassette. If there is a large quantity of dust inside, more PBST may be needed to 

dampen the dust before opening the TEF. Be careful not to hold or squeeze the black nozzle cover, 

as this may cause the seal to be disrupted, resulting in loss of PBST. 

9. Once a visual check has been completed to assure that the dust is dampened enough to prevent its 

aerosolization, use the sterile scalpel to cut through the red tape seal around the circumference of the 

TEF. 

10. Holding the TEF nozzle side up, open the plastic case of the filter. Turn the top of the filter case

upside down (nozzle side down with the black nozzle cover in place) and set aside for a moment,

inside the BSC on a rack (specimen cup rack will work) so that it will not be touching anything. 

(Figure 2) 
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Figure 2. Place the top half of the filter case onto a rack so that the filter can be removed from 

the bottom half of the filter case. 

11. Position the labeled stomacher bag in the bag rack so that the closure wires can be bent in a manner 

so that the bag remains open while standing up in the rack (Figure 3). 

 

Figure 3. Stomacher bag in bag holder with wires in open position, ready to accept filter and 

liquid. 
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12. Remove the moistened filter with sterile forceps and place in the stomacher bag (Figure 4).

Figure 4. Placing moistened filter into open stomacher bag. 

13. Use a pipette to aseptically remove liquid that has collected under the filter during the moistening 

step (Step 7 above). Place the liquid into the stomacher bag with the filter (Figure 5). 

Figure 5. Transferring moistening liquid to stomacher bag. 
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14. Use a disposable bulb pipette to aseptically rinse the inside walls of the bottom portion of the filter 

case with 1 to 2 mL of PBST (additional aliquots from the same 45 mL tubes used previously), then 

remove the rinse eluent (using the same transfer pipette) and place it into the stomacher bag along 

with the filter (Figure 6). 

Figure 6. Rinsing inside walls of filter case. 

15. Repeat the rinsing step for the top portion of the filter case that was set aside (Step 10). 

16. Pipette or pour the remainder of the 90 mL PBST (two tubes of 45 mL each) into the stomacher bag 

that contains the filter and the rinsate of the filter case (Figure 7). 

Figure 7. Pouring remainder of 90 mL PBST into stomacher bag. 
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17. Discard the filter case, red plug nozzle cover, disposable pipettes and disposable thumb forceps into 

the proper waste containers. DO NOT discard the 50 mL centrifuge tubes, as you will use these 

again for centrifuging the eluent after processing in the stomacher. 

18. Close stomacher bags by folding multiple times and folding the wire closures into the center of the 

bag. Wipe the outside of the bag with a disinfecting wipe and place in a secondary stomacher bag 

rack or sample retention bin. 

19. Wipe down the surface of the BSC as described in Step 3 above. 

20. Change gloves, and repeat from Step 5 for each remaining TEF sample. 

2.0 PROCESSING TRACE EVIDENCE FILTERS

1. Set the stomacher to 1 min, 260 rpm (setting P3). Place one stomacher bag from Section 1.0, Step 18 

(above) into the stomacher and run the cycle. Remove the stomacher bag and place back in the bag 

rack. 

2. Repeat for each sample. 

3. Hold the samples at room temp for 10 min to allow the foam to settle in the stomacher bag (Figures 

8 and 9). 

Figure 8. Stomacher bag with sample; foam in stomacher bag immediately after 1 min 

stomacher cycle. 
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Figure 9. Stomacher bag with sample; settled foam after 10 min holding time. 

4. Using a 50 mL pipette, transfer the eluent evenly (so the tubes will be balanced in the centrifuge) 

into the same two labeled 50 mL centrifuge tubes saved from Section 1.0, Step 17 above (~43 mL in 

each tube, since ~ 4 mLs is absorbed by the filter). 

5. Weigh the tubes to guarantee that they will be balanced (+/- 0.25g). 

6. Centrifuge the eluent at 3500g for 15 min.

7. Carefully remove all but approximately 3 mL (use the 5mL gradient on the side of the tube to guide 

the amount) from each tube using a 50 mL pipette, being careful not to disturb the pellet. Vortex 

vigorously to disperse the pellet. 

8. Vortex the remaining liquid in the tubes and then aseptically combine the remaining eluent from the 

two tubes so that approximately 6 mL of eluent is now in one tube. Vortex and sonicate to break up 

aggregates before plating.

9. Measure and record the exact volume of the remaining liquid within each sample. 

3.0 PLATING AND FILTRATION OF BACTERIA FROM LIQUID EXTRACTS 

1. Follow MOP 6535a for serial dilutions and spread-plating samples in triplicate from the ~6mL 

solution containing the concentrated sample. 

2. Filtration – detailed in the steps below – is required only if extremely low recovery is expected. 
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a. Label three TSA plates with information specific to the sample and amount to be filtered. 

b. Set up filter manifold with three filter unit cups. 

c. Set the vacuum, with proper gauge, to no more than 10 inches Hg (4.9 psi). 

d. Add 10 mL PBST to filter cup. 

e. Add 1 mL sample eluent from sample cup to each filter cup on the manifold. 

f. Open the vacuum valve and vacuum liquid through filter.

g. Rinse sides of filter cup with 10 mL of sterile PBST, vacuum through again.

h. Pop off sides of the filter cup by squeezing and lifting. 

i. Use sterile forceps to lift filter and place it on the TSA, making sure good contact with the 

agar surface is made and air pockets are minimized. 

3. Repeat for subsequent sample tubes, completing samples in batches of no more than 4. This will help 

to avoid too much settling time between vortexing and plating. 

4. Place all TSA plates in incubator set at 35 oC and incubate 18 – 24 hrs before counting colonies. 

4.0 CALCULATIONS 

For spread plates: 

 total CFU recovered = (mean CFU on plate / 0.1 mL volume plated) x (1/dilution factor) x (total 

volume in centrifuge tube after decanting) 

For vacuum filter plates: 

 total CFU recovered = (mean CFU on vacuum filter /volume filtered) x total volume in 

centrifuge tube after decanting 
MOP 6586

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Miscellaneous Operating Procedure (MOP) 6586:

Recovery of Bacillus Spores from Quartz Filters

Prepared by: __________________________________________ Date: 2/12/2013

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader

Reviewed by: __________________________________________ Date: 2/12/2013

Dahman Touati, ARCADIS Project Manager

Approved by: __________________________________________ Date: 2/12/2013

Worth Calfee, EPA Work Assignment Manager

Prepared for

National Homeland Security Research Center

Office of Research and Development

U.S. Environmental Protection Agency

Research Triangle Park, NC 27711

Prepared by

ARCADIS U.S., Inc.

4915 Prospectus Drive, Suite F

Durham, NC 27713
MOP 6586

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MOP: 6586

Title: RECOVERY OF BACILLUS SPORES FROM QUARTZ FILTERS

Objective: To aseptically extract and quantify recovery of Bacillus spores from quartz filters used to

sample air.

MATERIALS AND EQUIPMENT

Equipment:

 Biological Safety Cabinet (BSC, Class II)

 Seward 400 Circulator Stomacher®

 Incubator set to appropriate growth temperature for target organism (35 °C ± 2 oC)

 Light box for counting colonies

 Vortex mixer

 Stomacher bag rack

 Serological pipette

Materials:

 Personal Protective Equipment (PPE) (gloves, lab coat, safety goggles)

 1:10 Bleach, pH-amended to 7.0 with acetic acid (pAB)

 Dispatch bleach wipes

 Disposable 500 ml beaker (such as Fisher Scientific, cat #14-955-111D) (for discarding used tips

and beads/spreaders)

 70% Solution of denatured ethanol

 Kimwipes

 Waste container

 Sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562)

 Tryptic Soy Agar (TSA) plates

 Sterile 1500 μL tubes filled with 900μL of sterile PBST

 120 mL specimen cup containing 100 mL of sterile PBST
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 Pipettor and pipette tips for dilutions (100 μl, 1000 μl, and 5000 μl.) 

 Disposable cell spreaders or glass beads for cell spreading

 100 mL serological pipette tips

 Disposable sterile thumb forceps

 10” x 15” Twirl’em bag (Fisher, cat # 01-002-53)

 Quartz filter (Whatman, cat # 18209932)

 Seward stomacher 400 bags (BA6141/CLR closure bags x 10)

1.0 PREPARATION

Personnel must be familiar with all procedures prior to start.

1.1 Equipment Preparation

a) Begin by donning personal protective equipment (PPE) such as gloves, lab coat, and protective

eyewear.

b) Clean the workspace (Biological Safety Cabinet; BSC) by wiping surfaces with pH-amended

bleach (pAB), next with deionized (DI) water, and lastly with a 70-90% solution of denatured

ethanol. Allow any excess liquid to dry prior to beginning procedure. Make sure the

workspace is clean and free of debris.

c) Assemble equipment in the BSC as needed: vortex mixer, filtration manifold, automatic

pipettors, tips, racks, etc.

d) Assemble extra supplies, such as stomacher and reagents, near BSC.

e) Wipe down workspace once again with 70-90% solution of denatured ethanol.

1.2 Supply Preparation

a) Unpack shipping containers onto a cart near a BSC.

b) For each sample, label TSA plates on the agar side of the plate with the sample number and the

appropriate dilution factors, as per MOP 6535a (Serial Dilution: Spread Plate Procedure to

Quantify Viable Bacterial Spores).
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c) Pre-label sterile, empty 120 mL specimen cups to collect the extraction liquid from each

sample.

2.0 PERFORM SPORE EXTRACTION AND CULTURE PROCEDURE

2.1 Dislodge Spores from the Sample Filters

a) Begin by donning a new pair of gloves. All subsequent procedures involving manipulation of

the filters or spore suspensions must be carried out in a BSC. (Stomaching may occur outside

the BSC when samples are double-contained inside the indicated bags.)

b) Outside of the BSC, open the secondary (10” x 15”) containment bag containing the labeled

stomacher bag containing the sample filter. Check to make sure the secondary bag and the

stomacher bag have matching labels. Wipe down the stomacher bag with a dispatch wipe and

place in the stomacher rack located in the BSC. Discard the outer 10” x15” bag in the nonregulated waste container.

c) Aseptically add 100 mL of PBST to each labeled stomacher bag containing a filter. The 100

mL of PBST should be pre-measured in a 120 mL specimen cup. Discard this specimen cup

when empty.

d) Place the stomacher bag containing the quartz filter sample and PBST into a second stomacher

bag to contain any leakage in the event the primary containment is compromised. Place the

combined bags such that 50 to 60 mm of the top portions protrude above the bag clamp, while

making certain that the quartz filter rests evenly between the homogenizer paddles.

e) Stomach filters in the PBST by completing the following:

1) Make certain the Stomacher® is set to MANUAL. Program the Stomacher® speed to 230

RPM and the timer to 2 minutes.

2) Open the Stomacher® door by raising the lid fully upward and back. The DOOR OPEN

icon will be displayed.

3) Close the door to the Stomacher®. The DOOR OPEN icon will no longer be illuminated.

4) Stomach each filter for 2 min by pressing the START button.

5) When the cycle ends, the Stomacher® will stop.

Note: If there is an emergency, stop the stomacher by pressing the red button or

the power button prior to opening the Stomacher®. Stopping the Stomacher® by

opening the door can damage the equipment.
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6) Open the door of the Stomacher® and remove the bags containing the filter. Remove the

labeled stomacher bag containing the filter sample and place it into the stomacher bag rack.

The same secondary stomacher bag may be used for each sample provided there was no

leak. If a leak was detected, leave the labeled stomacher bag in the secondary bag and

place both into the stomacher bag rack.

f) Repeat steps (b) through (d) for all samples.

g) Allow bags to sit for 10 min to allow the suspension foam to settle before beginning the

extraction step.

2.2 Recovery of Suspension

a) Obtain a new, empty, pre-labeled 120 mL specimen cup with the same sample ID as that of the

stomacher bag.

b) The suspension will be removed using a 100 mL pipette. Open the stomacher bag and remove

the liquid by first tilting the bag to one side, moving the liquid away from the filter. Remove

as much liquid as possible using the pipette.

c) When liquid can no longer be collected this way, simultaneously squeeze the filter and collect

liberated liquid via pipette. Make attempts to locate pipette tip in a location within the bag that

reduces collection of filter debris.

NOTE: The filter particles can become lodged in the pipette tip. If this happens it may be

necessary to eject some of the liquid to dislodge the particle.

d) After all the liquid has been removed from the bag, place it into the pre-labeled specimen cup.

Make sure to note the volume collected on the specimen cup as well.

e) Discard the labeled stomacher bag with the filter into a non-regulated waste container.

f) Repeat steps (a) through (e) for each sample.

2.3 Serially Dilute and Plate the Final Spore Elution Suspension

a) Use MOP 6535a to serially dilute and plate samples.

NOTE: If the samples are turbid, wide-orifice pipette tips may be used to prevent clogging of

the pipette tips.
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Miscellaneous Operating Procedure (MOP) 6593: 

Recovery of Spores from HVAC Filters

Prepared by: __________________________________________ Date: 4/30/2013 

Nicole Griffin Gatchalian, ARCADIS Work Assignment Leader 

Reviewed by: __________________________________________ Date: 4/30/2013 

Dahman Touati, ARCADIS Project Manager 

Approved by: __________________________________________ Date: 4/30/2013 

Worth Calfee, EPA Work Assignment Manager 

Prepared for 

National Homeland Security Research Center 

Office of Research and Development 

U.S. Environmental Protection Agency 

Research Triangle Park, NC 27711 

Prepared by 

ARCADIS U.S., Inc. 

4915 Prospectus Drive, Suite F 

Durham, NC 27713 
MOP 6593

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MOP: 6593 

Title: RECOVERY OF SPORES FROM HVAC FILTERS 

Scope: This MOP outlines the procedure for recovering spores from HVAC filter sections 

Purpose: To aseptically extract and quantify spores from sections of HVAC filters in order to 

determine viability and obtain quantifiable data 

1.0 MATERIALS 

 PPE (gloves, lab coat, safety goggles) 

 Class II Biological Safety Cabinet (BSC) 

 pH-amended bleach 

 Deionized (DI) water 

 70% solution of denatured ethanol 

 Kimwipes 

 Dispatch bleach wipes 

 Non-regulated waste container 

 Bottles of sterile Phosphate Buffered Saline with Tween 20 solution (PBST) (MOP 6562) 

 Vortex mixer 

 Shaker table with clamps or brackets to hold Nalgene bottles 

 Cart 

 Trypticase soy agar (TSA) plates 

 900uL dilution tubes of sterile PBST 

 Pipettor and pipette tips for dilutions 

 Incubator set to appropriate growth temperature for target organism (35°C+/-2°C) 

 Light box for counting colonies 

 Lab notebook 

 Sterile graduated cylinder 

 Parafilm 

2.0 PROCEDURE 

1. Begin by donning PPE (gloves, lab coat, and protective eyewear). 

2. Samples will be received as follows: Half portions of HVAC filters will be in 32oz white capped Nalgene 

jars (Fisher cat no. 2118-0032) and quarter portions of HVAC filters will be in 1L clear bottles (Fisher cat 

no. 02-893D). The samples should be secondarily contained by large sterile bags (10” x 15” sterile 

bags Fisher cat no. 01-002-53). Chain of custody forms that accompany the samples will need to be 
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reviewed to ensure that all of the samples IDs are consistent and that there is no notable variation in the 

samples. If variation has occurred, make a note of it in the notebook. 

3. Clean BSC workspace by wiping surfaces with pH-amended bleach, followed by DI water, and lastly 

with a 70% solution of denatured ethanol. Wipe with a kimwipe to remove any excess liquid. Make 

sure the workspace is clean and free of debris. Gather all necessary items to perform the task, place 

these items on a clean cart beside the BSC, within arm’s reach so that once the procedure has begun 

the task may be performed without interruptions. 

4. Discard gloves and replace with a fresh pair. 

5. Place the samples one at a time, under the BSC. Samples should be handled in order of least 

contaminated to most contaminated, with any negative controls or blank samples being handled first, 

test samples being handled next, and lastly the positive controls. Carefully unwrap the secondary 

containment bag from around the jar or bottle, discard in non-regulated waste container and then wipe 

down the jars or bottles with a dispatch wipe, followed by a Kimwipe. Change gloves after handling 

each sample. 

6. Once all of the samples are under the BSC and disinfected, change gloves and place the sterile 

graduated cylinder under the BSC, along with several bottles of sterile PBST. Again, in order from 

least contaminated to most contaminated, handle the samples individually, and carefully open the jars 

or bottles to aseptically add 700 mL of PBST to the sample. The PBST is first aseptically poured into 

the graduated cylinder and measured at 700 mL, and then it is aseptically poured from the graduated 

cylinder into the sample. Carefully transfer all liquid as to not spill any or disturb/touch the filter or its 

container in any way. Once the 700 mL of PBST has been added, carefully place the jar lid or bottle top 

back on the sample, and make certain it is tightly closed. 

7. Again, wipe the outside of the samples with Dispatch wipes, followed by a Kimwipe to remove any 

moisture. After all samples have 700 mL of sterile PBST added, wrap the sample lids/bottle tops with 

Parafilm to ensure a tight seal and to prevent sample leakage. 

8. Place all sample jars or bottles into an orbital shaker table, using the specially designed platform 

holders. Make certain all samples are secure, and shake at room temperature for 30 min at 300 rpm. 

9. Immediately after the thirty minute agitation step is completed, remove the samples from the shaker and 

place them, again from least contaminated to most contaminated, into the BSC for dilution plating. Don 

a fresh pair of gloves, remove and discard the Parafilm, and follow MOP 6535a to complete serial 

dilutions using TSA media plates that are appropriately labeled with the sample ID, dilution set, and 

date. Each sample should also be manually agitated for ten seconds prior to removal of aliquots. 

Dilution tubes used in MOP 6535a should contain PBST to stay consistent with materials/solutions. 

 

10. Once the dilution plating has been completed, incubate the plates at 35°C +/- 2°C for 18-24 hours, and 

archive the samples in at refrigerator at 4°C+/-2°C. 

11. Count the colonies with the assistance of a light box. Record data. 

12. Colony forming units for each 6” x 12” half section or for each 6” x 6” quarter section will be determined 

and the sum of the CFUs for the two or four sections making up one 12” x 12” sample will be 

determined. Data will be reported as total CFU for the 12” x 12” sample. 
Offi ce of Research and Development (8101R)

Washington, DC 20460

Offi cial Business

Penalty for Private Use

$300

PRESORTED STANDARD

POSTAGE & FEES PAID

EPA

PERMIT NO. G-35