DEPARTMENT OF HEALTH AND HUMAN SERVICES

Centers for Disease Control and Prevention

National Institute for Occupational Safety and Health

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

Sampling and characterization of 

bioaerosols

by William G. Lindsley, Brett J. Green, Francoise M. Blachere, Stephen B. Martin, Brandon F. 

Law, Paul A. Jensen and Millie P. Schafer, NIOSH

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1 Introduction 

2 Principles of bioaerosol collection 

3 Devices used for bioaerosol sampling 

4 Considerations for bioaerosol sampling 

5 Selection of bioaerosol samplers 

6 Sample preparation for culturable bioaerosols 

7 Identification of culturable bioaerosols 

8 Enumeration of culturable bioaerosols 

9 Sample analysis methods for non-viable and non-culturable bioaerosols 

10 Limitations of bioaerosol sampling and characterization 

11 Safety considerations 

12 Resources 

13 References 

14 Appendix 1- List of manufacturers/distributors of common bioaerosol 

 samplers and related products 

15 Appendix 2 – Commonly used bioaerosol samplers BA-104
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Sampling and Characterization of Bioaerosols 

1 Introduction

Bioaerosols are airborne particles that originate from biological sources including animals, 

plants, fungi, bacteria, protozoa, and viruses. Examples of bioaerosols encountered in 

occupational environments include plant pollen, algae, fungal spores, bacteria such as 

actinomycetes, droplets produced during coughing and sneezing that may contain bacteria 

and viruses, dust containing insect excreta, animal dander, and fragments derived from each 

of these sources. Bioaerosols are ubiquitous and can be isolated from indoor, outdoor, and 

occupational environments using a variety of methods that either enumerate viable or a 

collection of viable and non-viable bioaerosols. Photomicrographs of example viral, bacterial, 

fungal, and plant bioaerosols are presented in Figure 1.

Bioaerosol monitoring is a rapidly emerging area of industrial hygiene due to the improved 

analysis methods such as polymerase chain reaction (PCR) and the impact that occupational 

exposures may have on worker respiratory health, particularly in microbial contaminated 

environments [Eduard et al. 2012; Environment Agency 2009; Haig et al. 2016; Hung et al. 

2005; Macher 1999; Morey 2007; Nazaroff 2016]. Some human diseases encountered in 

healthcare settings such as measles and tuberculosis can be spread by bioaerosols containing 

infectious microorganisms [Ijaz et al. 2016; Jones and Brosseau 2015]. Soil saprophytic fungi 

such as Coccidioides immitis can be aerosolized during occupational disturbance activities 

and, if inhaled, can result in an acute pulmonary infection [Das et al. 2012; Wilken et al. 2014; 

Wilken et al. 2015]. The measurement of these bioaerosols in industrial hygiene includes the 

measurement of viable (culturable and non-culturable) and nonviable bioaerosols in indoor 

settings (e.g., industrial, office, education, and residential buildings), industrial facilities (e.g., 

biotechnology, composting, waste disposal, manufacturing, textile, and food processing), and 

outdoor environments (e.g., farms, feed lots, and general air quality). Monitoring for 

bioaerosols in the occupational environment is one of the many tools the industrial hygienist 

uses in the assessment of indoor air quality, infectious disease outbreaks, agricultural 

exposures, and industrial health.

Bioaerosol monitoring may be appropriate during workplace health and exposure 

assessments, epidemiological investigations, research studies, or in situations deemed 

appropriate by an occupational physician or immunologist. Sampling can also be used to 

evaluate occupational environments before and after mitigation of microbial contaminants. 

When investigating bioaerosols as a possible source of workplace exposures and health issues, 

bioaerosol sampling should be part of an integrated assessment of work conditions. This 

should also include examining heating, ventilation and air conditioning (HVAC) systems; 

checking for water infiltration and moisture control; evaluating microbial contamination in 

evaporative cooling systems, metal working fluids, and waste water; evaluating possible 
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Sampling and Characterization of Bioaerosols 

 internal and external sources of bioaerosols; and other measures [Macher 1999]. In general, if 

visible growth or contamination (microbial growth on floors, walls, or ceilings, or in the 

HVAC system) is observed, this normally should be mitigated first before indoor bioaerosol 

sampling is conducted. If personnel remain symptomatic after remediation, air sampling may 

be appropriate, but the industrial hygienist should be aware that false negative results are 

possible and should be interpreted with caution.

The industrial hygienist has a variety of tools and methodologies available to conduct an 

environmental survey [ASTM 2014a; Flannigan et al. 2011; Hung et al. 2005]. However, many 

of these approaches have lacked standardization and this has made the interpretation and 

comparison between studies challenging [Flannigan et al. 2011]. In 2005, the American 

Industrial Hygiene Association (AIHA) published the second edition of the Field Guide for 

the Determination of Biological Contaminants in Environmental Samples [Hung et al. 2005]. 

This reference provides the industrial hygienist access to the most up to date methods to 

detect and quantify bioaerosols in the environment, and covers methods of how to conduct a 

survey, sample bioaerosols, and interpret the collected data [Hung et al. 2005]. Similarly, other 

reference sources have been published by Flannigan et al. [2011] and the American 

Conference of Governmental Industrial Hygienists (ACGIH) [Macher 1999] that extensively 

outline available methods to analyze collected bioaerosols as well as strategies to conduct an 

environmental survey. ASTM International has issued a wide range of standards on indoor air 

quality, including assessment of fungal growth and collection of bioaerosols and a guide to 

developing an air sampling strategy [ASTM 2009; ASTM 2014a; ASTM 2014b; ASTM 2014d]. 

The European Committee for Standardization has also published standards on sampling for 

bioaerosols and related topics [CEN 2000; CEN 2003; CEN 2004]. The sections presented 

below provide a very broad overview of the viable and non-viable methods available to detect 

bioaerosol sources that are described in the references listed above. 
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Figure 1: Photomicrographs of acellular, prokaryotic and eukaryotic microorganisms that can 

be encountered in occupational or industrial environments. (A) Transmission electron 

micrograph of Influenza/flu (H1N1) virus particles (Photo courtesy of National Institute of 

Allergy and Infectious Diseases; CDC Public Health Image Library (PHIL) ID#: 18156); (B) 

Scanning electron micrograph of bacilli derived from the Gram-negative bacteria, Legionella 

pneumophila (Photo courtesy of National Institute of Allergy and Infectious Diseases; CDC 

Public Health Image Library (PHIL) ID#: 11150); (C) Scanning electron micrograph of 

Aspergillus species reproductive structures including chains of asexual spores (Photo courtesy 

of CDC/ Robert Simmons; CDC Public Health Image Library (PHIL) ID#: 13367); and (D) 

Scanning electron micrograph of tricolpate pollen derived from the angiosperm plant species, 

Oenothera fruticosa (Photo courtesy of CDC/ Janice Carr, Betsy Crane; CDC Public Health 

Image Library (PHIL) ID#: 8729). The CDC Public Health Image Library at 

http://phil.cdc.gov/Phil/home.asp has thousands of health-related images available to the 

public free of charge.
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Sampling and Characterization of Bioaerosols 

 

2 Principles of bioaerosol collection

a. Aerodynamic diameter

The aerodynamic diameter of an airborne particle (usually written as “da” or “dae”) is the 

single most important parameter that determines how the particle will behave in the air, 

including how long it will stay airborne and where it will deposit in the respiratory system 

if inhaled. If a particle is falling in still air, it will reach an equilibrium velocity where the 

gravitational force pulling it downward is balanced by the drag force on its surface. This 

velocity is called the terminal settling velocity, and it depends upon the size, shape and 

density of the particle. The aerodynamic diameter of a particle is defined as the diameter of 

a sphere with unit density (that is, a density of 1 g/cm3) that has the same terminal settling 

velocity as the particle. Consider, for example, the irregularly-shaped fungal fragment 

shown in Figure 2. Suppose this particle has a terminal settling velocity of 0.05 cm/sec. 

This is the same settling velocity as that of a spherical particle with a unit density that has a 

diameter of 4 μm. Thus, the fungal fragment is said to have an aerodynamic diameter of 4 

μm. Similarly, a different particle with a terminal settling velocity of 1.21 cm/sec has an 

aerodynamic diameter of 20 μm, since a 20 μm unit density sphere settles at that rate. It is 

important to note that the aerodynamic diameter may be very different from the physical 

size of a particle. A very dense and compact particle may have an aerodynamic diameter 

much larger than its actual dimensions, while a very light particle or one with fibrous 

branches may have an aerodynamic diameter that is much smaller than its physical size. It 

is possible for two particles to have very different shapes and physical sizes, but have the 

same aerodynamic diameter. Conversely, two particles may have similar physical sizes, but 

have very different aerodynamic diameters. A more detailed discussion of the 

aerodynamic diameter can be found in Hinds [1999] and Vincent [2007].
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Sampling and Characterization of Bioaerosols 

 

Figure 2: Aerodynamic diameter of an aerosol particle. In this case, the fungal fragment on 

the left is said to have an aerodynamic diameter of 4 μm, since it falls at the same terminal 

settling velocity as a 4 μm sphere with a unit density.

Aerodynamic diameter is used in aerosol science because particles with the same 

aerodynamic diameter tend to move and be collected in the same ways. For example, two 

particles with the same aerodynamic diameter will have the same likelihood of being 

collected by an impaction aerosol sampler even if they have different physical and 

morphological characteristics. For this reason, the performance of aerosol collection 

devices is usually described by giving the aerodynamic diameter of the particles that will be 

collected.

b. Collection efficiency and cut-off diameter

The collection efficiency of an aerosol sampler is the fraction of the aerosol particles of a 

particular aerodynamic diameter that will be collected by the sampler. For example, if 95% 

of the airborne particles with a 2 μm aerodynamic diameter that enter the sampler are 

deposited in the collection fluid or on the collection surface, then the sampler is said to 

have a 95% collection efficiency for 2 μm particles.

Most commonly-used aerosol filters have a high collection efficiency for particles of all 

sizes [NIOSH 2016b]. However, impactors, cyclones and impingers use the inertia of 

airborne particles to separate them from the air stream, and thus they have a high 

collection efficiency for particles with larger aerodynamic diameters and a low collection 

efficiency for smaller ones (Figure 3) [Hering 2001; Hinds 1999; Marple and Olson 2011]. 

These devices are said to have a “cut-off diameter”; that is, particles with an aerodynamic 
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Sampling and Characterization of Bioaerosols 

 diameter larger than the cut-off diameter are collected while particles with an aerodynamic 

diameter less than the cut-off diameter are not collected and pass through the device. A 

perfect collection device would have a 100% collection efficiency for particles larger than 

the cut-off diameter and 0% for smaller particles. In practice, this is not the case: the 

collection efficiency curve for an inertia-based sampler looks like the example curve shown 

in Figure 3. The aerodynamic diameter at which the collection efficiency is 50% is defined 

as the cut-off diameter (usually written as d50). A device with a more abrupt transition 

from 100% to 0% collection efficiency (that is, closer to the ideal device) is said to have a 

sharp cut-off.

For a given inertial collection device, the 50% cut-off diameter depends upon the air 

flowrate through the device. Increasing the flowrate will decrease the d50 and shift the 

collection efficiency curve to the left, while decreasing the flowrate will increase the d50 and 

shift the collection efficiency curve to the right. For example, the first stage of the NIOSH 

two-stage cyclone aerosol sampler has a d50 of 4.9 μm at 2 liters/minute of air flow, 4.1 μm 

at 3.5 liters/minute, and 2.1 μm at 10 liters/minute [Blachere et al. 2009]. For this reason, it 

is important to check the air flowrate before aerosol sampling and control it during 

sampling so that the particles are correctly segregated by size.

Figure 3: Example collection efficiency curve for an inertia-based aerosol sampler. Note 

that the collection efficiency is high for particles with large aerodynamic diameters and 

low for small particles. In this example, the 50% cut-off diameter (d50) for this device is 1 

μm.
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Sampling and Characterization of Bioaerosols 

 

c. Size-selective bioaerosol sampling in industrial hygiene

Size-selective bioaerosol sampling may be done for several reasons. Since the settling

velocity of aerosol particles is determined by the aerodynamic diameter, knowing the size 

distribution of an aerosol helps in predicting how long the particles are likely to remain 

airborne and how far they can travel. In health care settings, for example, various medical 

procedures can produce a spray of droplets containing infectious microorganisms. Large 

droplets tend to fall onto surfaces fairly close to the source, while smaller droplets can 

remain airborne and carry pathogens many feet away from a patient [Davies et al. 2009; 

Jones and Brosseau 2015]. Another application of size-selection is to isolate different types 

of bioaerosol particles, such as separating fungal fragments from intact fungal spores 

[Adhikari et al. 2013; Seo et al. 2014].

Size-selective sampling is most commonly used to help understand the potential health 

effects of bioaerosol particles, which often depend upon where the particles are deposited 

in the respiratory tract. In general, larger bioaerosol particles tend to deposit higher in the 

respiratory tract (that is, in the nasal or oral cavities or larger airways), while smaller 

particles are able to travel deeper into the lungs to the smaller airways [Hinds 1999; 

Vincent 2005]. Some pathogens such as Mycobacterium spp., Bacillus spp., and Aspergillus 

spp. are thought to be more likely to cause a pulmonary infection if they reach the deeper 

airways, and the response to bioaerosols containing immunogenic material such as 

endotoxins or fungal antigens may also vary depending upon the site of deposition. For 

this reason, size-selective sampling is often used in industrial hygiene to better understand 

the potential risks that workplace bioaerosols present.

The American Conference of Governmental Industrial Hygienists (ACGIH), the 

International Organization for Standardization (ISO) and the European Standardization 

Committee (CEN) have defined three particle collection efficiency curves for aerosol 

samplers used to conduct size-selective aerosol sampling (Figure 4) [ACGIH 2001; ISO 

2012; Vincent 2005]. The idea is that an aerosol sampler that conforms to one of the three 

criteria will collect aerosol particles in a way that approximates the fraction of particles 

that will reach different parts of the respiratory tract. These criteria are not specific to 

bioaerosols, but rather are applied to all types of aerosol particles.
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Figure 4: ACGIH/ISO sampling criteria for the inhalable, thoracic and respirable fractions 

of aerosol particles. The inhalable fraction contains all of the particles that are inhalable, 

which includes the particles in the thoracic and respirable fraction. Similarly, the thoracic 

fraction includes the particles in the respirable fraction. The 50% cut-off diameters are 100 

μm for the inhalable fraction, 10 μm for the thoracic fraction, and 4 μm for the respirable 

fraction [ACGIH 2001; ISO 2012; Vincent 2005].

A sampler that collects the inhalable fraction accumulates the fraction of aerosol particles 

of each size that would be expected to be drawn into the nose or mouth during normal 

breathing. This includes larger particles that would be expected to be deposited in the 

nasal or oral cavities as well as smaller particles that can be conveyed to the lower airways. 

An aerosol sampler that conforms to the inhalable sampling criteria collects 50% of the 

100 μm particles, 77% of the 10 μm particles, and 97% of the 1 μm particles in the ambient 

aerosol. The inhalable fraction is lower for larger particles because the greater inertia of 

these particles means they are less likely to be pulled into the body during inhalation. 

The thoracic fraction includes aerosol particles that are likely to travel into the trachea and 

bronchi. An aerosol sampler that conforms to the thoracic sampling criteria will collect 

50% of the 10 μm particles and 97% of the 1 μm particles in the ambient aerosol. This 

fraction includes fewer large particles because these particles tend to be removed from the 

airstream by the head airways.
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 The respirable fraction includes aerosol particles that are able to reach the deepest airways, 

which are the respiratory bronchioles and the alveoli. An aerosol sampler that conforms to 

the respirable sampling criteria will collect 50% of the 4 μm particles, 97% of the 1 μm 

particles, and 99% of the 0.3 μm particles in the ambient aerosol. The respiratory 

bronchioles and the alveoli are of particular concern because these airways do not have 

cilia. Non-soluble particles that land in the nasopharyngeal region or upper airways tend 

to collect in the airway mucus and are removed from the respiratory tract by the cilia 

relatively quickly. However, particles that deposit in the alveoli and respiratory 

bronchioles can remain in the lungs for longer durations (in some cases, for life) unless 

they can be broken down or removed by migrating pulmonary macrophages. This fraction 

includes only the smallest particles because the larger particles are removed from the 

airstream by the head and thoracic airways.

It should be noted that, even though larger bioaerosol particles will tend to deposit in the 

upper airways and be cleared more quickly, they can still trigger an allergic/inflammatory 

response in susceptible individuals. Particles containing viable pathogens also commonly 

cause infections after being deposited in the upper airways.

When describing size-selective sampling, particles are often said to “penetrate” to a 

particular region of the respiratory tract. This does not mean penetrate in the sense of 

entering the tissue, but rather simply being present in the air stream flowing into that 

region, as compared to particles which were deposited before reaching a particular 

location. For example, an aerosol particle that is able to remain in the air stream and reach 

the lung alveoli is said to have penetrated to the alveolar region, even if it does not 

necessarily deposit there. This is the same context as with filtration, where a particle is said 

to penetrate a filter if it flows through the filter material and remains in the air stream. It 

also should be noted that the ACGIH/ISO criteria give an approximation of the fraction of 

aerosol particles that can penetrate to different regions of the respiratory tract. However, 

they do not indicate what fraction of the aerosol particles will actually deposit in the 

airways and what fraction will be exhaled. The lung deposition of aerosol particles is 

complex and depends upon many factors. More information about this topic can be found 

in Hinds [1999] and Vincent [2005; 2007].

3 Devices used for bioaerosol sampling

Most aerosol sampling devices involve techniques that separate particles from the air stream 

and collect them in or on a preselected medium. Impactors, filters, impingers and cyclones are 

four common sampling techniques used to separate and collect bioaerosols [Haig et al. 2016; 

Macher et al. 1995; Reponen et al. 2011b; Willeke and Macher 1999]. A few systems that use 

electrostatic precipitation or condensation-based collection are also available [Haig et al. 
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Sampling and Characterization of Bioaerosols 

 2016], and some real-time bioaerosol monitoring systems are available that do not require 

that the bioaerosol particles be isolated before analysis. Below are some specific types of 

bioaerosol sampling devices employed by industrial hygienists.

a. Filters

Aerosol filters are commonly used to collect bioaerosol particles because of their simplicity 

and low cost. Filter-based sampling is particularly useful for personal bioaerosol sampling 

because filter-based collectors are small and lightweight and work well with personal 

sampling pumps. Filters can be preceded by a size-selective inlet, such as a cyclone or 

impactor, to remove larger particles and provide size-classification of the bioaerosol 

particles. Most aerosol filter media can be classified as fibrous, membrane, or capillary 

pore (also called straight-through pore) [Raynor et al. 2011]. Fibrous filters are usually 

made of a deep mesh of glass fibers. Membrane filters are manufactured in a variety of 

pore sizes from polymers such as cellulose ester, polyvinyl chloride, or 

polytetrafluoroethylene (PTFE). Capillary pore filters are made of polycarbonate. The 

choice of a filter medium depends on the contaminant of interest and the requirements of 

the analytical technique. For gravimetric analysis, non-hygroscopic materials such as glass 

fibers, silver, or polyvinyl chloride membranes are selected because their masses are less 

affected by changes in humidity. For analysis by microscopy, cellulose ester or 

polycarbonate membranes are common choices because cellulose ester membranes can be 

rendered transparent for easier visualization, while polycarbonate filters have a smooth 

collection surface that works well with light or electron microscopy. Samples also can be 

eluted from cellulose ester and polycarbonate filters, but in some cases the recovery 

efficiency can be low [Eduard et al. 1990; Rule et al. 2007]. Samples to be cultured can be 

collected on gelatin filters, and the filters can then be dissolved in water and spread on 

culture plates, dissolved in growth media, or placed directly on culture plates and allowed 

to melt. Gelatin filters are fragile and can crack or melt in use. For analysis using 

immunological assays or polymerase chain reaction (PCR), PTFE filters are a common 

choice because they do not interfere with the assays and because samples can be readily 

eluted from them.

Filters are frequently described or specified using the term “pore size” or “equivalent pore 

diameter”. It is important to note that the filter pore size does NOT indicate the minimum 

particle size that will be collected by the filter; in fact, aerosol filters generally will collect 

particles much smaller than the nominal pore size. The mechanisms by which aerosol 

filters work and the role of pore size in selecting filters is described is more detail 

elsewhere [NIOSH 2016b].
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Sampling and Characterization of Bioaerosols 

 Aerosol filters are usually supplied as disks of 25, 37 or 47-mm diameter. Because the flow 

resistance (often called the pressure drop) of a filter increases with the air velocity through 

the filter, the use of a larger filter results in a lower flow resistance for a given volumetric 

flow rate. On the other hand, the use of a smaller filter concentrates the deposit of the 

contaminant onto a smaller total area, thus increasing the density of particles per unit area 

of filter. This may be helpful for direct microscopic examination of low concentrations of 

organisms, and reduces the amount of elution media needed for immunological or PCRbased assays. In areas of high concentration, the microorganisms may have to be eluted, 

diluted, and then refiltered for microscopic analysis. Breuer [2012] reported on the flow 

resistance of common aerosol filters and its relationship to sampling pump selection. Soo 

et al. [2016] measured the filtration characteristics and flow resistance of a variety of 

commonly-used aerosol filters.

In the USA, the most common method of aerosol sampling with filters is to place the 

filters in disposable two-piece or three-piece plastic filter cassettes with a support pad to 

add rigidity. The three-piece cassette may be used either in open- or closed-face modes. 

Open-face sampling is performed by removing the end plug and the plastic cover from the 

three-piece cassette and is used when the particulate matter must be uniformly deposited 

(i.e., for microscopic analysis). If a three-piece cassette is used in the open-face 

arrangement, the plastic cover is retained to protect the filter after sampling is concluded. 

It should be noted that the aspiration efficiencies of open-face and closed-face filter 

cassettes are reported to be somewhat different [Beaulieu et al. 1980; Kenny et al. 1997]. 

In addition to collecting on the filter, aerosol particles (especially large particles) may 

collect on the internal walls of the filter cassette. Depending upon the purpose of the 

collection, wall-deposited material may need to be included in the analysis. This can be 

done by using a filter with an attached capsule or by washing or wiping the internal 

surfaces of the cassette [Ashley and Harper 2013]. 

It is important to verify that the filter cassette and fittings are air-tight and have no bypass 

leakage around the filter. Cassettes should not be hand-assembled; they should be pressed 

together with a mechanical or hydraulic press. All plastic cassettes should be securely 

assembled and sealed with a cellulose shrink band or tape around the seams of the cassette 

to prevent external air leakage. The cassettes should be made of conductive or static 

dissipative materials to avoid losses due to electrostatic effects. More information on using 

filter cassettes for aerosol sampling can be found elsewhere [NIOSH 2003a; NIOSH

2016a].
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Sampling and Characterization of Bioaerosols 

 

b. Impactors 

An impactor consists of a series of nozzles (circular- or slot-shaped) and an impaction 

surface [Hering 2001; Marple and Olson 2011; Marple and Willeke 1976]. Air is drawn 

into the impactor using a vacuum pump, and the air stream flows through the nozzles and 

toward the impaction surface, where particles are separated from the air stream by their 

inertia (Figure 5). Larger particles collect on the impaction surface, while small particles 

that do not impact follow the air stream. The impaction surface typically consists of a 

greased plate or tape, filter material, or growth media (agar) contained in Petri dishes. In 

some applications, impactors are not used as collection devices themselves, but rather to 

remove particles above a certain size before collection or characterization of the 

downstream aerosol.

Figure 5: Impaction. As the air stream exits the impactor nozzle, it quickly changes 

direction as shown by the arrows. Smaller particles such as those on the left flow with the 

air stream and are not collected. Larger particles cannot change direction as quickly due to 

their higher inertia and collide with the collection surface, where they accumulate.

A cascade impactor consists of a stack of impaction stages: each stage consists of one or 

more nozzles and a target or substrate. The nozzles may take the form of holes or slots. 

Each succeeding stage has smaller nozzles and thus collects smaller particles (that is, each 

succeeding stage has a smaller cut-off diameter). A filter may be used after the final 

impaction stage to collect any particles smaller than the final cut-off diameter. If the 

substrate is a greased plate or filter media, it may be weighed to determine the collected 

mass, or it may be washed and the wash solution analyzed. If the substrate is growth media 

in culture plates, they may be incubated and examined for microbial growth.
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 The most commonly used impactor for sampling airborne culturable bacteria and fungi is 

the Andersen impactor, which uses from one to six impactor stages containing Petri plates 

as seen in Figure 6 [Andersen 1958]. Since the bioaerosol particles impact directly onto the 

growth media, the samplers can be directly transferred to an incubator and observed for 

microbial growth. However, this method depends upon collecting viable microorganisms 

that are capable of growth on the specific nutrient media. 

Glass Petri plates are recommended for use with the Andersen impactor; plastic culture 

plates are often used, but this can result in loss of aerosol material due to electrostatic 

surface charges in the plastic [Andersen 1958; Kuo 2015]. NIOSH Method 0800 describes 

how to collect culturable airborne fungi and bacteria in buildings using an Andersen 

cascade impactor [NIOSH 2003b].

Figure 6: Schematic of a 6-stage Andersen cascade impactor [Andersen 1958]. Each stage 

contains a Petri plate (green) filled with nutrient agar (brown). The stages have 

progressively smaller nozzles, which create higher particle impaction velocities onto the 

agar. The aerosol particles (red) flow from the top into the first stage, where particles with 

aerodynamic diameters larger than 7 μm impact the agar. The remaining particles flow to 

the second stage, where particles with aerodynamic diameters between 7 μm and 4.7 μm 

are collected, and so on for the rest of the stages.
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One significant advantage of the Andersen impactor is that samples can be collected

directly onto culture plates and transferred to an incubator, which simplifies handling and

eliminates some losses that can occur in processing. However, there are also several

limitations. In low concentration environments, sampling time is limited to approximately

20 minutes to avoid drying the agar. The high flow rate (28.3 liters/minute) makes the 

sampler unsuitable for high concentration environments such as some agricultural sites 

(i.e. animal facilities) where a 1 minute sample may overwhelm the plates.

When using the Andersen impactor, it is also necessary to correct for “coincidence error”

using a positive-hole correction factor. This occurs because it is possible for multiple

particles, each containing one or more organisms, to pass through a particular hole during

sampling and impact onto the growth medium, with one or more bacterial or fungal

colonies forming at the same impaction sites. The colonies formed by the multiple

particles can then be inaccurately counted as a single colony. As the number of organismcontaining particles deposited onto the growth medium increases, the probability that the 

next organism-containing particle will impact an "occupied" hole increases. For example, if

75% of the holes have received at least one particle, the chance that the next particle will

impact a "clean" hole is one in four (25%). To account for this, a probability-based 

coincidence correction factor needs to be applied to the results for each impactor stage.

The basic formula for the coincidence correction is as follows [Andersen 1958; Macher

1989]: 

Where:

N = the total number of holes in the impactor stage

r = the number of colonies observed on the culture plate

Pr = the estimated culturable particle count

Andersen impactors have from one to 400 holes per stage. Macher [1989], Willeke and

Macher [1999] and Andersen [1958] provide tables of positive-hole correction factors.

Investigators often employ stationary cascade impactors either as the primary collection

mechanism, or as a preclassifier (for example, to remove nonrespirable particles from the

sampled air stream). Marple and Willeke [1976] have reported that high velocity, inlet 

losses, interstage losses, and particle reentrainment affect the performance characteristics

of an impactor. Particles larger than the cut-off diameter may bounce after impacting the

collection surface and travel to subsequent impaction stages. This is particularly a problem

with dry solid collection surfaces; for this reason, solid collection surfaces are usually

greased or oiled [Hering 2001]. Fungal spores have been shown to be prone to deaggregation and bounce when collected with an impactor, which can cause the spores to be

collected on stages with smaller cut-off diameters. This can make the spore aggregates

appear to have smaller aerodynamic diameters than is actually the case [Trunov et al.
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Sampling and Characterization of Bioaerosols 

 2001]. Although personal cascade impactors are available, these devices are not as widely 

used in personal sampling for bioaerosols as are filters [Macher and Hansson 1987].

The slit-to-agar impactor is a type of impactor in which the aerosol particles are deposited 

on a Petri plate that slowly rotates. The rotation of the plate means that particles which are 

collected at different times deposit in different locations, and thus provides an indication 

of changes in the bioaerosol concentration over time [Ho et al. 2005; Jensen et al. 1992; 

Smid et al. 1989; USP 1997]. Examples of slit-to-agar samplers include the Dycor Slit 

Sampler from Dycor and the Air Trace Environmental Slit-to-Agar Sampler from Particle 

Measuring Systems.

The Hirst/Burkard spore trap has been widely used to collect outdoor aerospora. It was 

first described by Hirst [1952] and consists of a unit that houses a vacuum pump and 

rotating drum that is lined with polyester tape. The drum rotates at 2 mm per hour and is 

continuously run for seven days. Bioaerosols pass through an orifice on the sampler and 

particles impact on the tape. Following the seven-day sampling interval, the tape is 

removed and cut in 48 mm intervals that correspond to individual sampling days. 

Bioaerosols deposited on the tape are stained and then resolved, identified, and quantified 

using bright field microscopy.

 

[Tovey et al. 2016] developed a personal aerosol sampler with a rotating surface that allows 

time-resolved collection of aerosol particles onto an electret strip or an adhesive film. They 

used the sampler to study personal exposures to dust mite allergens over time.

A novel example of an impaction-based personal bioaerosol sampler is the intranasal air 

sampler fabricated by Graham et al. [2000], which fits within the intranasal cavity of the 

subject. Bioaerosols enter the nasal cavity following inhalation and pass through slits 

where particles are deposited by impaction on either an adhesive backed tape or collection 

cup lined with silicon grease. This impaction sampler has been utilized in a number of 

studies that have evaluated exposure to indoor and occupationally relevant aeroallergen 

sources [Gore et al. 2002; Mitakakis et al. 2000; Renstrom et al. 2002]. 

Other impaction-based approaches have also been used in the assessment of outdoor 

bioaerosols, including the Rotorod, Air-o-cell and Allergenco samplers [Frenz 1999; Lee et 

al. 2004a; Pityn and Anderson 2013; Portnoy et al. 2000]. ASTM Standards D7391 and 

D7788 discuss the collection and analysis of airborne fungal structures by inertial 

impaction [ASTM 2009; ASTM 2014d].
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c. Cyclones 

A cyclone sampler consists of a circular chamber with the aerosol stream entering through 

one or more tangential nozzles as shown in Figure 7 [Hering 2001]. Like an impactor, a 

cyclone sampler depends upon the inertia of the particle to cause it to deposit on the 

sampler wall as the air stream curves around inside the chamber. Also like an impactor, a 

cyclone sampler has a collection efficiency curve like the one shown in Figure 3, and the 

collection efficiency curve depends upon the flow rate. Cyclones are less prone to particle 

bounce than impactors and can collect larger quantities of material. They also may provide 

a more gentle collection than impactors, which can improve the recovery of viable 

microorganisms. However, cyclones tend to have collection efficiency curves that are less 

sharp than impactors, and it is simpler to design a compact cascade impactor compared to 

a cascade of cyclone samplers.

In industrial hygiene, cyclone aerosol samplers are frequently used in conjunction with a 

filter to conduct size-selective aerosol sampling [Hering 2001]. For example, in NIOSH 

Method 0600, a cyclone is used to remove the non-respirable fraction from the aerosol 

(following the ACGIH/ISO criteria described earlier), and a filter is then used to collect the 

respirable fraction [NIOSH 2003c]. A sampler developed at NIOSH uses two cyclones 

followed by a filter; the first cyclone collects the non-respirable fraction of the particles, the 

second cyclone collects the respirable particles > 1 μm, and the filter collects particles < 1 

μm [Blachere et al. 2009]. The NIOSH cyclone aerosol samplers have been used in 

applications including measurements of airborne viruses in healthcare settings; airborne 

fungi and fungal fragments in residences; airborne dimorphic fungal pathogens such as 

Paracoccidioides brasiliensis in Brazil, and bioaerosols in agricultural operations [Arantes 

et al. 2013; Blachere et al. 2009; Blais Lecours et al. 2012; Kettleson et al. 2013; Lee and Liao 

2014; Lindsley et al. 2010a; Lindsley et al. 2010b; Martin et al. 2015; Seo et al. 2014; Singh 

et al. 2011a; Singh et al. 2011b].
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Figure 7: Cyclone aerosol collection. When the aerosol stream enters the body of the

cyclone through the inlet, the air flow follows the curved interior wall and flows in a spiral 

pattern. If aerosol particles are larger than the cut-off diameter, then the inertia of the 

particles causes them to collide with the wall of the cyclone and accumulate. After 

spiraling downward, the air flow comes up through the center of the cyclone and exits 

through the outlet (called a vortex finder) at the top. The illustration shows a tangential 

inlet reversed-flow cyclone, which is the most common type of cyclone sampler.

d. Impingers 

Many microorganisms can lose their viability if they are collected onto dry solid surfaces 

or filters because of impact damage and desiccation [Cox 1987; Jensen et al. 1992; Macher 

and First 1984; Verreault et al. 2008; Wang et al. 2001]. One way to avoid this is to collect 

culturable bioaerosols in liquids using an impinger [Henningson and Ahlberg 1994; 

Henningson et al. 1988; Lembke et al. 1981; Reponen et al. 2011b; Verreault et al. 2008]. A 

typical impinger is shown in Figure 8. The body of the impinger is filled with a collection 

liquid, and the aerosol stream flows down through a nozzle and enters the liquid at a high 

velocity. The aerosol particles are collected when they collide with the bottom of the 

collection vessel or disperse into the liquid. Impingers often have curved inlets to remove 

larger particles from the air stream before collection. Because impingers are essentially 

another type of inertial collection device, they have a collection efficiency curve and a cutoff diameter like impactors and cyclones. However, the collection efficiency curves tend to 

be less sharp. The high velocity air stream directed into the liquid also creates considerable 

agitation and can produce foaming if the collection liquid contains surfactants. Additives 

to the collection medium such as proteins, antifoam, or antifreeze aid in resuscitation of 

bacterial cells, prevent foaming and loss of the collection fluid, and minimize injury to the 

cells [Chang and Chou 2011; Cown et al. 1957; Dungan and Leytem 2015]. The presence 
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 of proteins and other additives can also greatly influence the survival of airborne viruses 

during collection by impingers [Ijaz et al. 1985b; Schaffer et al. 1976; Verreault et al. 2008]. 

Water loss over time reduces the liquid level in the impinger and increases the 

concentration of the non-volatile components, which limits the available collection time 

[Lin et al. 1997]. Sample losses due to re-aerosolization and particle deposition inside the 

impinger can be significant [Grinshpun et al. 1997; Han and Mainelis 2012].

Figure 8: Impingement. Bioaerosol particles exit the nozzle of the impinger at high velocity 

and impact the liquid or the bottom surface of the collection vessel. Some types of 

impingers produce air bubbles in the collection media, which can enhance particle 

collection, but can damage some types of microorganisms.

Two common impingers used for bioaerosol sampling are the Greenburg-Smith impinger 

[Greenburg 1932] and the All-Glass Impinger with the nozzle 30 mm above the base of the 

collection vessel, called the AGI-30 [May and Harper 1957]. The Greenberg-Smith and 

AGI-30 samplers operate by drawing aerosols at nominal flow rates of 28.3 and 12.5 

L/min, respectively, through an inlet tube [Macher et al. 1995]. The AGI-30 inlet tube is 

curved to simulate particle collection in the nasal passage [Cox 1987]. Investigators have 

reported problems with low sampling efficiencies and high losses due to particles in the 

collection being re-aerosolized and lost [Grinshpun et al. 1997; Kesavan et al. 2010; Lin et 

al. 1997].

When the AGI-30 is used to recover total airborne organisms from the environment, the 

curved inlet tube is washed with a known amount of collecting fluid after sampling 

because larger particles (i.e., over 15 μm) are collected on the tube wall by inertial force. 

After sampling for the appropriate amount of time, 10 mL of the full-strength collection 
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 fluid is filtered through a 0.45-μm pore size membrane filter. Serial dilutions of the 

remaining collection fluid are handled similarly [Greenberg et al. 1992]. The membrane 

filters are placed in sterile plastic petri plates filled with the appropriate medium and 

incubated for later identification and enumeration.

e. Wetted-surface bioaerosol samplers

Several types of bioaerosol sampling devices have been developed in which the aerosol 

stream impacts onto a wetted surface or onto the wall of a cyclone wetted with collection 

media [Kesavan and Sagripanti 2015; Kesavan et al. 2011]. These systems largely avoid the 

bubbling and agitation associated with conventional impingers, which may be detrimental 

to some microorganisms [Lin et al. 2000], and can provide sharper collection efficiency 

curves. One of the simplest examples of a wetted-surface sampler is the SKC BioSampler 

[Lin et al. 2000; Willeke et al. 1998]. It is similar to an AGI-30, except that it has three 

nozzles that curve so that the aerosol stream is tangential to the wall of the collection 

vessel. This causes the collection liquid to swirl and greatly reduces the agitation, bubbling 

and consequent reentrainment seen with the AGI-30. The BioSampler collects particles 

with aerodynamic diameters of approximately 0.3 μm to 8 μm into the collection media, 

although the upper cut-off diameter is not sharp [Hogan et al. 2005; Kesavan et al. 2010; 

Willeke et al. 1998]. The BioSampler reportedly can be used with non-evaporating fluids 

such as mineral oil to eliminate the collection time limits imposed by water evaporation, 

provided that the microorganism can survive collection and processing [Lin et al. 2000]. 

Alternatively, fluid can be exchanged or added to the sampler as needed [Rule et al. 2005; 

Rule et al. 2007].

The CIP10-M, a modified version of the CIP10 aerosol sampler, collects airborne 

microorganisms in a liquid layer on the interior surface of a rapidly-rotating cup. As with 

the BioSampler, the CIP10-M can be used with mineral oil as the collection fluid to avoid 

fluid evaporation. It is reported to have collection efficiencies of >80% for particles >2.8 

μm, 50% for 2.1 μm particles, and <10% for particles of <1 μm [Görner et al. 2006; Simon 

et al. 2016].

May [1966] designed a three-stage sampler in which aerosol particles are collected by 

impaction onto a wetted fritted surface in the first two stages and the third stage is a 

swirling aerosol collector similar to the BioSampler. Both glass and stainless steel versions 

are available. In his original report, May [1966] used particles with a density of 1.5 g/cm3

and reported cut-off sizes of 6 μm, 3.3 μm and 0.7 μm, which correspond to aerodynamic 

diameters of about 7.3 μm, 4 μm, and 0.86 μm. The May sampler is reported to give 

comparable results to the Andersen impactor [Zimmerman et al. 1987].
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 Several wetted-surface bioaerosol samplers recirculate the collection fluid and add 

additional fluid as needed to replace evaporative losses. This extends the collection time 

available and allows the concentration of the aerosol from a large volume of air at a high 

flow rate into a relatively small volume of liquid, which is of great advantage when 

searching for pathogens that may be present in very low concentrations. For this reason, 

such systems are often used for bioterrorism and homeland security applications. The 

Coriolis sampler [Carvalho et al. 2008], the OMNI-3000 [Zhao et al. 2014], the SASS 2000 

[Ravva et al. 2012], and the SpinCon [Yooseph et al. 2013] use a wetted wall cyclone for 

bioaerosol collection, while the BioCapture 650 [Ryan et al. 2009] collects particles onto a 

wetted rotating impactor. Kesavan and Sagripanti [2015] reported the results of 

performance tests for several of these types of bioaerosol samplers.

When conducting long-term bioaerosol collection into liquid media, it is important to 

note that if the collected bioaerosol particles remain in the collection media for an 

extended time and if steps are not taken to inhibit growth, spore germination and cell 

amplification of some fungi and bacteria can occur. This can result in the appearance of 

much higher bioaerosol concentrations than are actually present in the environment.

f. Condensation-based bioaerosol samplers

Some bioaerosol particles are too small to be readily collected by impactors or impingers. 

These particles can be collected using filters, but filter collection can reduce the viability of 

microorganisms. One solution is to humidify the aerosol stream and then cool it, which 

causes water vapor to condense on the aerosol particles and create a droplet surrounding 

the particle. This larger particle can then be collected by impaction or impingement, as 

shown in Figure 9. This is similar in principle to condensation-based particle counters, 

which are used to measure the concentration of small airborne particles. Some researchers 

showed that adding water vapor to an aerosol stream enhanced the recovery of airborne 

viruses and bacteriophages, which may work by this method (although this is unclear) 

[Hatch and Warren 1969; Trouwborst and Kuyper 1974; Warren et al. 1969]. More 

recently, Milton developed a condensation-based system to collect fine particles 

containing influenza virus from the exhaled breath of human subjects [McDevitt et al. 

2013; Milton et al. 2013]. A condensation-based bioaerosol sampler called a growth-tube 

collector has been used to collect MS2 bacteriophage and influenza virus in the laboratory, 

and is reported to be especially effective at recovering viable virus in sub-micrometer 

particles [Lednicky et al. 2016; Pan et al. 2016; Walls et al. 2016]. A version of this system 

called the Spot Sampler (Aerosol Devices, Inc.) is commercially available.
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Figure 9: Condensation-based aerosol particle collector.

g. Electrostatic samplers

Electrostatic precipitation works by using a strong electric field to create a high 

concentration of unipolar ions. The rapid motion of these ions causes them to collide with 

and charge airborne particles, and the resulting charge on the particles causes them to be 

attracted to the collection surface [Hinds 1999]. Electrostatic precipitation systems have 

been used to collect bioaerosol particles such as allergens, bacteria and viruses [Artenstein 

et al. 1968; Artenstein et al. 1967; Custis et al. 2003; Donaldson et al. 1982; Heitkamp et al. 

2006; Lee et al. 2004b; Parvaneh et al. 2000; Roux et al. 2013]. Such devices offer simplicity 

of design with few moving parts, and are generally effective at collecting small particles. 

One electrostatic bioaerosol sampling device is available commercially from Inspirotec 

[Gordon et al. 2015].

Some electrostatic bioaerosol samplers collect particles into liquid to concentrate the 

particles and help preserve the viability of microorganisms. The Large Volume Air 

Sampler (LVS) developed by Litton in the 1960’s washed the collection surface with 

recirculating fluid; this sampler was successfully used to collect pathogenic respiratory 

bacteria and viruses in a variety of settings [Artenstein et al. 1968; Artenstein et al. 1967; 

Donaldson et al. 1982]. Pardon et al. [2015] developed a system that collects particles 

directly on a microfluidic chip. The electrostatic aerosol collector devised by Han et al. 

[2015] collects the deposited aerosol into rolling water droplets, which greatly 
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concentrates the particles. The Aerosol-to-Liquid Particle Extraction System (ALPES) uses 

an electrostatic system to collect aerosol particles into recirculating liquid, which helps 

preserve the viability of microorganisms [Heitkamp et al. 2006]. 

Electrostatically-charged cloths are used to collect airborne particles that settle onto them, 

and also to wipe settled dust from surfaces. These are discussed in the next two sections.

h. Passive bioaerosol samplers

Passive bioaerosol sampling refers to the collection of bioaerosols by allowing them to

gravitationally settle onto a collection device, such as a culture plate, foil sheet, electretbased filter or electrostatically-charged cloth. Compared to active sampling, passive

bioaerosol sampling has several advantages, including simplicity, low cost, lack of

disturbance of the surrounding air, and the ability to collect for extended time periods

[Haig et al. 2016; Pasquarella et al. 2000; Vincent 2007].

Passive bioaerosol sampling can be limited by several variables including the air currents 

around the device and airborne particle size. As discussed earlier, large particles settle 

much more quickly than small particles. Thus, large particles are much more likely to be 

collected by passive samplers [Haig et al. 2016; Reponen et al. 2011b]. As a result of these 

limiting variables, results from passive bioaerosol sampling cannot be directly related to 

the concentration of airborne particles and may not correlate well with results from active 

sampling [Reponen et al. 2011b]. However, some authors have proposed that passive 

sampling may be useful in evaluating the likelihood that bioaerosol particles will 

contaminate surfaces such as open wounds in operating rooms, since they mimic the 

contamination event more closely than does an active sampler [Friberg et al. 1999; Haig et 

al. 2016; Pasquarella et al. 2000].

Passive bioaerosol collectors are often placed 1.5 to 2 meters above the ground to avoid 

collection of large dust particles from sources other than airborne particles, such shoes, 

clothing, skin and animals [Frankel et al. 2012; Lioy et al. 2002; Noss et al. 2008; Rintala et 

al. 2012]. Grills, screens or shields may also be placed around or over the collection device 

to screen out large debris [Brown et al. 1996; Wagner and Macher 2003; Whitehead and 

Leith 2008; Wurtz et al. 2005].

Settle plates

Settle plates (also called settling plates or sedimentation plates) are culture plates 

containing nutrient agar that are opened and placed collection-side up in a location of 

interest. Airborne particles are allowed to settle onto the plates for a specified time, 

and the plates are then closed, incubated and inspected for growth. Settle plates are 
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commonly used to assess airborne microbial contamination and are listed in methods 

and standards from the ISO, the American Public Health Association (APHA) and the 

United States Pharmacopeia (USP) [Dyer et al. 2004; ISO 2003; USP 1997]. However, 

because the results from settle plates cannot be directly compared to the amount of 

airborne microbes, they should only be used for qualitative, not quantitative, 

evaluations. The CDC recommends the use of high-volume air samplers rather than 

settle plates when investigating airborne fungal spore contamination in health care 

facilities [CDC 2003].

Settle plate methods suffer from a lack of standardization of methodology, which 

makes results difficult to compare. Pasquarella et al. [2000] reviewed the use of settle 

plates and proposed an Index of Microbial Contamination (IMA) to standardize the 

use of settling plates. To measure the IMA, 90 mm culture plates are placed 1 meter 

above the floor and 1 meter from any walls, and collect settled particles for 1 hour 

(called the 1/1/1 scheme). The number of colony-forming units (CFUs) detected on 

each plate is then used to calculate the IMA in CFUs/dm2/hour [Pasquarella et al. 

2000]. 

Electrostatic dust collectors

Noss et al. [2008] developed a method called the electrostatic dustfall collector (EDC) 

that collects settling airborne particles onto four electrostatically-charged cloths. 

EDC’s have been used in studies of culturable bacteria and fungi, endotoxin, glucan 

and inflammatory mediators in airborne particles [Adams et al. 2015; Frankel et al. 

2012; Huttunen et al. 2016; Kilburg-Basnyat et al. 2016; Kilburg-Basnyat et al. 2015; 

Noss et al. 2010; Noss et al. 2008]. Noss et al. [2008] and Frankel et al. [2012] reported 

good correlations between the EDC and active aerosol samplers. Adams et al. [2015] 

compared EDC’s to Petri dishes and other passive collection materials and found that 

the results correlated reasonably well, but that a rigorous extraction protocol was 

required to get consistent results from the EDC’s. Brown et al. [1996] developed a 

passive electrostatic-based personal aerosol sampler and reported that it gave a 

reasonable correlation with inhalable dust measurements at farms and a rubber plant.

Other passive bioaerosol samplers

The UNC Passive Aerosol Sampler consists of a 6.8 mm diameter collection substrate 

mounted on a scanning-electron microscope stub and shielded by a protective screen 

[Wagner and Macher 2003; Whitehead and Leith 2008]. Airborne particles settle or 

diffuse onto the substrate and can be analyzed by optical or electron microscopy. 

Other investigators have used aluminum sheets in boxes, Petri dishes, and sheets of 

various plastic materials as passive bioaerosol collectors [Adams et al. 2015; Meadow et 

al. 2015; Wurtz et al. 2005].
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i. Settled dust collection devices

The collection and analysis of dust that has settled onto floors, carpets, and other surfaces

is widely used as a means of identifying bioaerosols in buildings, especially allergens,

endotoxin and molds [Hung et al. 2005; Lioy et al. 2002; Martyny et al. 1999; Morey 2007;

Rintala et al. 2012]. Settled dust sampling allows for the collection of large quantities of

material, provides a long-term sample, and does not require a dedicated sampling device

for each location. Dust assays allow quantitative data to be generated per weight and

surface area of dust. Some investigators find it useful to compare different sites in a

building or to sample before and after remediation efforts to see if the source of a

bioaerosol has been eliminated.

Settled dust will vary within a building depending upon the location and collection surface 

[Lioy et al. 2002; Rintala et al. 2012]. In addition to settling from the air, dust can be 

produced by a variety of other mechanisms, making it difficult to distinguish the source. 

Floor and carpet dust, for example, will include outside material brought in by shoes, skin 

flakes, clothing fibers and animal dander. Sampling locations well above floor level are 

often chosen to minimize the amount of dust that is not from settled airborne particles 

[Frankel et al. 2012; Rintala et al. 2012].

Vacuums

The US Department of Housing and Urban Development has developed a protocol for 

the vacuum collection of home dust samples to test for allergens [HUD 2008]. 

Vacuum collection of settled dust from floors and carpets has been used to determine 

the Environmental Relative Moldiness Index (ERMI), which is a measure of mold 

contamination in homes [Kettleson et al. 2015; Reponen et al. 2012; Reponen et al. 

2011a; Taubel et al. 2016; Vesper et al. 2013; Vesper et al. 2007]. ERMI is discussed in 

more detail later in this chapter. Note that vacuuming can increase the levels of 

bioaerosols in a location. Thus, air sampling should be completed before collecting 

surface samples by vacuuming [Hung et al. 2005; Hunter et al. 1988].

Swabs

Swabs are widely used to collect airborne material that has settled onto surfaces. Swabs 

are also used to identify microbial contaminants that may be colonizing building 

materials within the indoor environment. However, obtaining consistent and reliable 

results from swab sampling is far more difficult than is often appreciated, and careful 

attention is needed to the choice of swab material, elution media, and method of 

swabbing. If swab samples are to be cultured, aseptic technique is needed to avoid 

contamination. ASTM International has a standard for collecting fungal material by 

swab [ASTM 2012]. The APHA has published a standard method for swab sampling of 
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 food-contact surfaces [Dyer et al. 2004], while the USP and ISO have standards that 

include swab sampling for microbiological contamination in clean rooms [ISO 2003; 

USP 1997]. 

An example of a validated protocol for swab sampling is that provided by NIOSH for 

surface sampling for Bacillus anthracis spores [Hodges et al. 2010; Hodges et al. 2006; 

NIOSH 2012b]. In this procedure, a defined area is first outlined using a template or a 

ruler and masking tape. A sterile macrofoam swab is then moistened using a buffer 

solution that neutralizes disinfectants. The surface is swabbed using horizontal strokes, 

followed by vertical strokes, and finally diagonal strokes, and the swab is then placed 

in a sterile tube for transport and analysis. Aseptic technique is used throughout the 

procedure.

The choice of swab material can have a significant impact on the collection of 

microorganisms from a surface. Moore and Griffith [2007] studied the recovery of 

Escherichia coli and Staphylococcus aureus from stainless steel squares using nylonflocked swabs and spatulas, cotton swabs and rayon swabs. They reported that nylonflocked and cotton swabs were equally effective at removing bacteria from dry 

surfaces, but that cotton swabs removed bacteria more effectively from wet surfaces 

than rayon or nylon-flocked swabs. However, nylon-flocked swabs and spatulas 

released the bacteria into the elution media more readily than rayon swabs, which in 

turn released more bacteria than cotton swabs. For viruses, polyester-tipped swabs 

were found to be more effective than cotton swabs or antistatic wipes at recovering 

MS2 bacteriophage from stainless steel and plastic [Julian et al. 2011], while 

macrofoam swabs performed best when recovering wet or dried norovirus from 

stainless steel surfaces, followed by cotton, rayon and polyester swabs [Park et al. 

2015]. 

The elution media used to wet the swabs and recover the bacteria from the swabs also 

can have a substantial effect on sampling. Moore and Griffith [2007] tested eleven 

different swab wetting solutions containing various combinations of salts, surfactants 

and nutrients. They found that the recovery efficiency varied widely depending upon 

the species of bacteria, type of swab, and whether the surface was wet or dry. For MS2 

bacteriophage, saline or Ringer’s solution (an isotonic salt solution) worked better 

than viral transport media or pure water [Julian et al. 2011]. It is important to note 

that the elution media must both remove the biological material from the surface and 

subsequently elute it from the swab in order to be effective.

Although they may be overlooked, storage conditions play an important role in swab 

sampling. After sample collection, room temperature storage of moist swabs may lead 
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to microbial growth if the elution media or swab contain nutrients, while the presence 

of chemicals such as Tween 80 may reduce viability over time. These problems can be 

alleviated by placing the swabs in cold storage as quickly as possible [Moore and 

Griffith 2007].

Wipes

All of the considerations and limitations of swab sampling also apply to wipe 

sampling. Swabs are typically more useful for small surfaces and hard-to-reach 

locations, while wipes are more effective at collecting dust from large non-porous 

surfaces [NIOSH 2012b]. Electrostatic wipes have been used to collect settled dust for 

studies of mold and endotoxin [Bolaños-Rosero et al. 2013; Thorne et al. 2005]. 

However, Thorne et al. [2005] found that wipes and gloves themselves were frequently 

contaminated with endotoxin and needed to be tested before use.

Adhesive tape

Adhesive tape can be used to collect dust samples from surfaces for microscopic 

examination (this is called tape lift or cellotape sampling) [ASTM 2014c; Martyny et al. 

1999; Morey 2007]. Typically, a section of adhesive tape is gently pressed onto a 

surface of interest, removed with a slow steady force, and then attached to a glass slide 

or placed in a vial. The samples are relatively simple to collect, but the results depend 

upon the ability of the examiner to identify microorganisms and their fragments, and 

do not provide a quantitative assessment of exposure.

Contact plates

Contact plates are typically round culture plates in which the agar is poured so that the 

top of the agar forms a meniscus slightly above the top rim of the plate. A surface 

sample is collected by inverting the plate and pressing the agar directly onto a flat 

surface of interest. The plate is then removed, incubated and inspected for microbial 

growth. This sampling method is often called the replicate organism direct agar 

contact (RODAC) procedure, and it is commonly used for biocontamination 

monitoring in the pharmaceutical and food industries [Dyer et al. 2004; ISO 2003; USP 

1997]. Because many of the surfaces of interest in these industries are routinely 

disinfected, contact plates are available with agars that contain neutralizers for 

disinfectants. One report indicated that nitrocellulose membranes were slightly more 

effective than RODAC plates at surface sampling, and are easier to use on curved 

surfaces [Poletti et al. 1999].
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j. Heating, ventilation and air conditioning (HVAC) filters

Building HVAC systems filter large quantities of outside and recirculated inside air as they 

maintain environmental conditions inside buildings. Researchers have taken advantage of 

these existing filtration systems as a way to study bioaerosols in a variety of structures 

[Goyal et al. 2011; Haaland and Siegel 2016; Noris et al. 2011]. Testing the collected 

particulate material on HVAC filters provides an inexpensive way of studying bioaerosols 

collected from large volumes of air over long time periods. However, some limitations 

must be kept in mind. Extracting bioaerosols from these filters can be difficult and the 

methods require validation [Farnsworth et al. 2006]. Many microorganisms lose viability 

after collection, so although PCR-based methods may be effective, culture-based methods 

likely will not work except for very hardy microbes [Farnsworth et al. 2006]. Finally, 

commonly-used HVAC filters can have relatively low collection efficiencies, especially for 

small particles [ASHRAE 2009]. Haaland and Siegel [2016] reviewed 60 studies in which 

HVAC filter analyses were used to study bioaerosols in buildings.

k. Real-time bioaerosol monitoring

Many biological molecules have an intrinsic autofluorescence, and this phenomenon has 

been used as the basis for continuous real-time bioaerosol detection systems [Pöhlker et al. 

2012]. This technique is most commonly employed for studies of atmospheric bioaerosol 

particles and for biodefense and biosecurity applications. These systems can distinguish 

biological from non-biological particles, and can usually provide information about the 

particle size and some characteristics of the bioaerosols. One device, the TSI Ultraviolet 

Aerodynamic Particle Sizer (UV-APS), was used in several studies [Bhangar et al. 2016; 

Hairston et al. 1997; Kanaani et al. 2008]; it has been replaced by an updated version called 

the Fluorescence Aerosol Particle Sensor (FLAPS) III. Other real-time bioaerosol detectors 

include the BioScout [Saari et al. 2014], the Wideband Integrated Bioaerosol Sensor 

(WIBS-4) [Toprak and Schnaiter 2013], and the Fido B2 (formerly called the 

Instantaneous Bioaerosol Analysis and Collection, IBAC) [Santarpia et al. 2013].

4 Considerations for bioaerosol sampling

a. Development of a bioaerosol sampling strategy

The first step in designing a sampling strategy for bioaerosol sampling is to determine the 

purpose of the sampling [ASTM 2014a]. For example, bioaerosol sampling may be 

conducted to estimate worker exposure to bioaerosols, or to select or evaluate engineering 

controls to reduce exposures, or to identify the source of a bioaerosol. A sampling strategy 

then should begin with an overview of the site of interest and development of initial 

hypotheses regarding the types, sources and distributions of bioaerosols. After this, the 

sampling methods, times, durations, and the analytical methods can be selected. Note that 
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Sampling and Characterization of Bioaerosols 

 bioaerosol sampling is almost always done in conjunction with the collection of other 

types of data, such as worker health information, visual observations, air flow 

measurements, surface sampling, and information about possible sources.

b. Sampling locations

The sampling locations should be selected to assist in evaluation of the working 

hypotheses about possible exposures [ASTM 2014a]. If worker exposures are being 

evaluated, then the samplers should be placed in areas occupied by the workers. If 

contamination of a ventilation system is being examined, then sampling in the system and 

at the ventilation louvers would be appropriate. Care must be exercised to ensure that 

people do not tamper with the samplers and that microorganisms on surfaces or in duct 

work are not inadvertently aerosolized.

Bioaerosol samples should be drawn directly into the sampler rather than being 

transported to the sampler by tubing. If transport tubing must be used, it should be as 

short and straight as possible. Abrupt flow constrictions and bends in the tubing should be 

especially avoided, as considerable sample deposition can occur at these locations. The 

tubing diameter should be large enough that the flow is not turbulent and that the d50 of 

any bends is well above the size of the bioaerosol particles [Pui et al. 1987; Tsai and Pui 

1990]. The tubing should be made of a material that does not lead to losses through 

electrostatic deposition [Liu et al. 1985]. A review of the many issues surrounding the 

transporting of aerosols through sampling lines is provided by Brockmann [2011].

Personal aerosol sampling provides a much better representation of worker and resident 

exposure to aerosol particles than area (static) sampling [Cherrie et al. 2011; Kissell and 

Sacks 2002; Rodes and Thornburg 2005]. However, most samplers for viable bioaerosols 

do not lend themselves to personal sampling. Thus, a combination of personal and area 

sampling may be necessary to fully characterize the exposure [Toivola et al. 2002].

c. Concentrations of indoor and outdoor bioaerosols

Indoor bioaerosol sampling is conducted in occupational (industrial, education, and office 

environments) and non-occupational (residential and buildings) settings. Outdoor 

bioaerosol sampling is often performed to provide comparative data for indoor sampling 

and to help determine possible sources of contaminants. Outdoor bioaerosol sampling also 

is conducted in occupational environments such as agricultural settings, composting sites 

and sewage treatment plants [Environment Agency 2009; Lee and Liao 2014; Masclaux et 

al. 2014]. In addition, outdoor sampling may be performed for pollen and fungi to assist 

allergists in their treatment of patients by identifying taxa distribution and concentrations 

in air over time.
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 The concentrations of bioaerosol particles vary widely depending upon the meteorological

parameters, the location of sources, the time of year and the amount of ventilation. 

Shelton et al. [2002] studied 1,717 buildings in the United States. They found that outdoor 

levels of airborne fungi are usually higher than indoor levels, and that fungal levels were 

highest in the fall and summer and lowest in the winter and spring. Outdoor levels varied 

from 1 to more than 8,200 colony-forming units (CFU)/m3 of air, with a median of 540 

CFU/m3

. Indoor levels ranged from 1 to over 10,000 CFU/m3, with a median of 82 

CFU/m3. An examination of fungi in flood-damaged homes found fungal concentrations 

of 1,100 to 8,400 spores/m3 outside and 500 to 101,100 spores/m3 inside [Reponen et al. 

2007]. An investigation of 100 large office buildings by Tsai and Macher [2005] found that 

airborne bacterial concentrations tend to be higher outdoors than indoor (except for 

Gram-positive cocci). Outdoor concentrations tended to be higher in the winter (194 vs. 

165 CFU/m3), while indoor concentrations were higher in the summer (116 vs. 87 

CFU/m3). Forty-one percent of the bioaerosol samples were below the detection limit, and 

>95% of the culturable bacteria were mesophilic (grow at moderate temperatures). In a 

report on agricultural workers working in animal confinements, Lee et al. [2006] found 

breathing zone culturable bioaerosol exposures of 300 to 36,000 CFU/m3 for fungi, 3000 to 

3.3 x 108 CFU/m3 for bacteria, and up to 2,800 CFU/m3 for actinomycetes. During grain 

harvesting, workers were exposed to culturable bioaerosol levels of 82,000 to 7.4 × 106 

CFU/m3 for fungal spores, 40,000 to 1.4 × 106 CFU/m3 for bacteria, and up to 2.6 × 104 

CFU/m3 for actinomycetes.

If one or more genera of fungi or bacteria are found indoors in concentrations greater than 

outdoor concentrations, then the source of amplification may need to be found and 

remediated. When conducting indoor bioaerosol sampling, it is advisable to sample 

before, during, and after the sampling area is occupied, including times when the heating, 

ventilating, and air conditioning system is activated and inactivated.

d. Viable and nonviable bioaerosols

Viable microorganisms are metabolically active (living) organisms with the potential to 

reproduce, grow and colonize. Viruses are not metabolically active but are considered 

viable if they are capable of reproducing in an appropriate cellular host. Viable 

microorganisms may be culturable or non-culturable. Culturable organisms reproduce 

under controlled laboratory conditions. Non-culturable organisms do not reproduce in 

the laboratory because of intracellular stress or because the conditions (e.g., culture 

medium or incubation temperature) are not conducive to growth. Some bacteria can be 

very difficult or impossible to culture from bioaerosols. For example, although human 

Mycobacterium tuberculosis is readily transmitted among people and from people to 

Guinea pigs, it has never been successfully cultured from an environmental aerosol 
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Sampling and Characterization of Bioaerosols 

 sample, probably because of its extremely low airborne concentrations and slow growth 

rate [Nardell 2016]. Other bioaerosols such as Histoplasma capsulatum or Pneumocystis 

carinii may take weeks to grow or may not even grow in culture at all [Dennis 1990; Ibach 

et al. 1954]. As the name implies, viable bioaerosol sampling involves collecting a 

bioaerosol and culturing the collected particles. Only culturable microorganisms are 

enumerated and identified, thus leading to an underestimation of bioaerosol 

concentration. Non-viable and viable but non-culturable microorganisms are often 

studied by collecting them with a dry aerosol sampler or a membrane filter. The 

microorganisms are then enumerated and identified using microscopy, classical 

microbiology, molecular biology, or immunochemical techniques [Hung et al. 2005; 

Macher 1999; Reponen et al. 2011b; Tortora et al. 2013].

Assessment of viable bacteria is also dependent on a number of variables including 

nutrient media, temperature and culture conditions. In indoor environments the 

collection of viable bacteria may be confounded by endogenous bacterial microflora such 

as Staphylococcus epidermis that sheds with skin flakes [Hung et al. 2005]. Concentrations 

of viable bacteria have been reported to be as high as 105 CFU/m3 in indoor environments; 

however, like fungi, the proportion of the total bacterial burden may be higher if nonviable bacteria are also included [Hung et al. 2005]. In addition, viable assessment of 

several bacterial species of clinical significance may not be the best approach as these 

bacteria do not remain viable in the air. Alternative methods such as immunoassays or 

molecular-based methods may provide suitable approaches for quantifying bacterial 

pathogens.

e. Bioaerosol particle sizes

As noted earlier, the aerodynamic diameter (dae) of an airborne particle is the most 

important factor determining how long it will remain in the air, how likely it is to be 

inhaled, and where it will deposit in the respiratory tract. The sizes of bioaerosol particles 

can range from tens of nanometers for small fragments to hundreds of micrometers for 

pollen, fungi or large agglomerations. However, most of the bioaerosol particles of interest 

in the indoor environment fall between about 100 nm and 10 μm [Nazaroff 2016]. For 

bacteria, vegetative cells typically have physical diameters of about 0.2 to 2 μm and are 2 to 

8 μm in length, while bacterial spores are somewhat smaller [Tortora et al. 2013]. Airborne 

particles containing bacteria were found to have aerodynamic diameters of about 1 to 3 

μm in indoor environments [Gorny et al. 1999; Kujundzic et al. 2006; Meklin et al. 2002]. 

Mycobacterium tuberculosis is a rod-shaped bacteria with a length of about 6.6 μm [Schafer 

et al. 1999]. When aerosolized from a liquid culture, M. tuberculosis DNA was found in 

particles with aerodynamic diameters of 0.6 to 1.8 μm [Schafer et al. 1999]. Aerosolized 

Mycobacterium bovis BCG (a commonly-used surrogate for M. tuberculosis) was found in 
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Sampling and Characterization of Bioaerosols 

 particles with aerodynamic diameters of 0.5 to 9.9 μm [Schafer et al. 1998]. Air sampling 

around indoor whirlpools in a public facility found airborne mycobacteria DNA in 

particles with aerodynamic diameters of 0.5 to 9.9 μm [Schafer et al. 2003]. Actinomycete

spores tend to be smaller, with aerodynamic diameters of cultured spores ranging from 0.6 

to 1.5 μm [Madelin and Johnson 1992; Reponen et al. 1998]. Fungal spores have physical 

diameters of about 0.5 to 30 μm or larger, while the aerodynamic diameters of airborne 

fungal spores and spore clusters are reported to be from 0.9 to 5 μm [Eduard 2009; 

Hussein et al. 2013; Reponen et al. 2011b].

Airborne microorganisms are often present as parts of aggregations, droplets or 

agglomerations that can be much larger than the size of the native microorganism. In 

indoor environments with large amounts of other aerosol particles like cigarette smoke, 

bacteria have been found on particles with aerodynamic diameters up to 10 μm, which was 

larger than airborne bacterial particles in cleaner environments. This was thought to occur 

because the aerosol particles were forming agglomerates [Gorny et al. 1999]. In a farm 

study, airborne Actinomycetes and fungal spores were more likely to be found in 

aggregates in environments with higher spore concentrations [Karlsson and Malmberg 

1989]. In two studies of airborne influenza virus in health care facilities, about half of the 

airborne virus was found in particles with aerodynamic diameters of 4 μm or greater, even 

though the virus itself is only about 100 nm in diameter, because the virus was contained 

in aerosolized droplets of respiratory fluids [Blachere et al. 2009; Lindsley et al. 2010a]. 

Agglomerates of fungal spores can break apart upon impaction inside an impactor and be 

collected on subsequent stages with smaller cut-off diameters [Trunov et al. 2001].

Bioaerosols may also be present as cellular fragments that are much smaller than the 

source microorganisms. Endotoxins are fragments of the cellular walls of Gram-negative 

bacteria that have been implicated in a variety of illnesses [Eduard et al. 2012; Jacobs 1989; 

Olenchock 2002]. Fragments of fungal cell walls also are thought to be associated with 

several types of adverse respiratory health effects [Green et al. 2011; Green et al. 2006b; 

Olenchock 2002]. Very high levels of fungal fragments have been measured in flooddamaged homes contaminated with mold [Reponen et al. 2007]. Fungal fragments also 

contain a variety of secondary metabolites, mycotoxins, beta-glucan, antigens and 

allergens [Green et al. 2011; Green et al. 2006b]. In one study of indoor air in homes, the 

majority of the endotoxin and fungal wall material was found in particles with 

aerodynamic diameters of less than 1 μm [Adhikari et al. 2013]. Another study found 

considerable amounts of endotoxin in aerosol particles from metalworking fluids that 

were between 0.16 and 0.39 μm [Wang et al. 2007]. Indoor and outdoor measurements of 

endotoxin levels found that the largest proportion was detected in particles with 

aerodynamic diameters of less than 1 μm [Kujundzic et al. 2006].
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 It is common to use an aerosol spectrometer in conjunction with bioaerosol sampling to 

better understand the size distribution of the airborne particles. One consideration when 

interpreting the data is, of course, that the large majority of these devices do not 

distinguish between biological and non-biological aerosols. Another less-obvious factor is 

that while a few aerosol spectrometers such as the TSI Aerodynamic Particle Sizer measure 

the aerodynamic diameter of the airborne particles, many aerosol spectrometers measure 

particles using light scattering and thus provide an approximate physical diameter instead 

[Hinds 1999; Sorensen et al. 2011]. The difference between the aerodynamic and optical 

diameters may be significant depending upon the shape and density of the particles.

f. Temperature and humidity

The temperature and humidity of the environment can affect the size of bioaerosol 

particles, the viability of airborne microorganisms, the growth of microorganisms on 

surfaces, and the amount of electrostatic charges on aerosols and surfaces. Because of these 

effects, the environmental temperature and humidity should be recorded during 

bioaerosol sampling.

Water evaporates rapidly from wet aerosol particles [Hinds 1999]. If an airborne particle is 

initially an aqueous solution containing non-volatile substances such as salts and organic 

material, and if the relative humidity is above the crystallization relative humidity (CRH, 

also called the efflorescence relative humidity), then some of the water will evaporate and 

the solution will become more concentrated, but the particle will remain liquid. If the 

relative humidity is below the CRH, then all of the water will evaporate (that is, the particle 

will desiccate) [Nicas et al. 2005]. Similarly, if an airborne particle is initially a dry 

combination of salts and organic material, and if the relative humidity is below the 

deliquescence relative humidity (DRH), then the particle will remain desiccated. However, 

if the relative humidity is above the DRH, then the particle will absorb water until it 

liquefies and becomes an aqueous solution. The DRH is always greater than the CRH 

[Nicas et al. 2005]. A particle in an environment above its CRH (or DRH if it was initially 

dry) will be larger and heavier and will settle faster than the same particle when the 

humidity is below the CRH, which can affect the size and amount of bioaerosol particles 

that are collected during sampling [Mikhailov et al. 2004]. This phenomenon was seen in a 

study of particles in human exhaled breath, where the particles detected in low humidity 

air were substantially smaller than those detected when the air was more humid 

[Holmgren et al. 2011].

Bioaerosol particles may also undergo an increase in size when the humidity increases due 

to water absorption and swelling of hygroscopic components. An increase in relative 

humidity has been shown to increase the aerodynamic diameter of fungal spores [Madelin 
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Sampling and Characterization of Bioaerosols 

 and Johnson 1992; Reponen et al. 1996]. Similar results have been reported for 

Actinomycetes spores [Madelin and Johnson 1992].

For airborne viruses, survival decreases as air temperature increases [Ijaz et al. 2016; Tang 

2009]. Exposing most viruses to temperatures of 60°C or higher for 60 minutes will 

inactivate them, although the viruses can be somewhat protected if they are encased in 

organic material [Tang 2009]. For example, in one set of experiments, airborne particles 

containing vaccinia virus, influenza virus, and Venezuelan equine encephalomyelitis virus 

all showed higher survival rates at 7-12°C than at 21-24°C, and still lower survival at 32-

34°C [Harper 1961]. Aerosol transmission of influenza virus among Guinea pigs is 

blocked at air temperatures of 30°C [Lowen et al. 2008]. The effect of humidity on virus 

survival depends upon the virus; in general, viruses with lipid envelopes tend to survive 

better at low humidity, while non-enveloped viruses survive better at high humidity [Ijaz 

et al. 2016; Tang 2009]. For example, influenza viruses and coronaviruses have enveloped 

capsids, and both survive better at low humidities compared to high [Ijaz et al. 1985a; Ijaz 

et al. 2016; Noti et al. 2013; Schaffer et al. 1976]. On the other hand, rotaviruses and 

rhinoviruses have non-enveloped capsids and survive better at high humidities compared 

to low [Ijaz et al. 1985b; Ijaz et al. 2016; Karim et al. 1985].

The survival of airborne bacteria also decreases as air temperature increases; the survival of 

virtually all airborne bacteria declines when temperatures are above 24°C [Ijaz et al. 2016; 

Tang 2009]. However, as with viruses, the effects of humidity on bacterial survival are 

much more complex, and depend not only upon species but also upon the methods of 

culture and aerosolization [Cox 1989; Tang 2009]. In field experiments in a greenhouse, 

survival of certain bacteria was 35- to 65-fold higher at 80% RH than at 40% [Walter et al. 

1990]. In laboratory experiments, survival of certain bacteria was virtually complete at low 

RH but was reduced at RH values above 80% [Cox 1968]. Higher humidities can also 

significantly decrease the efficacy of ultraviolet germicidal irradiation (UVGI) for reducing 

levels of viable airborne bacteria [Peccia et al. 2001]. Cox [1987] believes the potential for 

the movement of the solvent water is an important environmental criterion in assessing 

survivability of bacteria, viruses, and phages.

Fungi and fungal spores generally are better able to withstand environmental stresses 

compared to vegetative bacteria and viruses [Ijaz et al. 2016; Tang 2009]. Warm 

temperatures, wet substrates and humid air conditions favor the growth of fungi on 

surfaces [Eduard 2009; Tang et al. 2015]. Temperature can induce morphological changes 

in dimorphic fungi such as the pathogen Histoplasma capsulatum [Salvin 1949]. It is not 

clear, however, how air temperature and humidity affect the viability of airborne fungi and 

fungal spores [Tang 2009].
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g. Electrostatic effects

Aerosol particles in the workplace can be highly charged, and the electrostatic charge can 

vary considerably depending upon the aerosol generation mechanism and the particle 

characteristics [Johnston et al. 1985]. Aerosol particles are especially prone to develop 

electrostatic charges in low humidity environments [Baron and Deye 1990]. Like most 

particles, freshly generated microbial aerosols are nearly always electrostatically charged 

unless steps are taken to neutralize them. Lee et al. [2004b] found that airborne fungi and 

bacteria carried a net negative charge in most of the laboratory and field environments 

that they studied. Mainelis et al. [2002] found that a strong positive electrostatic charge 

reduced the viability of Pseudomonas fluorescens bacteria but did not affect Bacillus subtilis

spores.

The effect of electrostatic charge on aerosol collection is often overlooked, resulting in the 

possible bias of sampling results [NIOSH 2016a; Vincent 2007]. Aerosol samplers made of 

non-conductive plastics can develop substantial electrostatic charges, which can degrade 

their performance significantly [NIOSH 2016a; Baron and Deye 1990]. The use of 

polyethylene or polytetrafluoroethylene (PTFE) tubing to transport air streams to a 

sampler can remove a sizeable amount of aerosol particles by electrostatic deposition [Liu 

et al. 1985]. As noted above, the use of plastic Petri dishes in an Andersen impactor can 

result in bioaerosol particle losses [Andersen 1958; Kuo 2015]. Whenever possible, it is 

better to use aerosol samplers made of conductive materials such as metals or speciallytreated plastics [NIOSH 2016a].

h. Flow calibration

Accurate airflow rates are very important in calculating the concentration of 

microorganisms in the air. All samplers should be calibrated before and after sampling to 

ensure that the flow rate is within the manufacturer's specifications and does not change 

from the initial calibration. Calibration may be performed using a primary standard such 

as a spirometer or bubble calibrator. Where it is not possible to calibrate using a primary 

standard, a calibrated secondary standard such as a dry gas meter may be used. The 

calibration of such a secondary standard should be traceable to a primary standard. A 

detailed explanation of the calibration of airflow rates is given by McCammon Jr. and 

Woebkenberg [NIOSH 2016c].

i. Blanks 

Laboratory media blanks are unexposed, fresh samples of media, such as agar plates, filters 

and impinger fluids. These samples are generally not taken into the field. Before using any 

batch of media, incubate at least three culture plates under the same conditions as planned 
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Sampling and Characterization of Bioaerosols 

 for the field samples, in order to check for sterility of the media. Approximately five media 

blanks should be included with each sample set. If the samples are to be analyzed by an 

outside laboratory, consult the specific laboratory procedure for the number of blanks to 

be submitted. Similarly, blank filters should be processed in the same manner as planned 

for field samples in order to check for contamination. 

Field blanks are simply unopened, fresh media samples that are handled in the same way 

as field samples, including labeling, except that no air is drawn through the sampler. The 

generally recommended practice for the number of field blanks is to provide at least two 

field blanks for every 10 samples with a maximum of 10 field blanks for each sample set.

5 Selection of bioaerosol samplers

The first step in selecting a bioaerosol sampling device is to establish the purpose of the 

sampling. Once the goal of the bioaerosol sampling is determined, the appropriate sampling 

methods may be chosen. The selected bioaerosol sampler must be capable of high efficiency 

particle collection within the physical and biological conditions required by the 

microorganisms to be sampled. The most appropriate sampling methods will be dictated in 

part by the techniques that will be used to analyze the sample. Methods for bioaerosol sample 

analysis are discussed in the next section. A list of some manufacturers and suppliers of 

bioaerosol sampling equipment and supplies is shown in Appendix I. The characteristics of 

several commonly used bioaerosol samplers are shown in Appendix II.

a. Sampling for airborne bacteria and fungi 

Choosing a bioaerosol sampler for bacteria and fungi begins by deciding how the 

bioaerosol will be analyzed, and in particular whether the viability of the bacteria or fungi 

will be evaluated. Culturable bioaerosol sampling instruments must minimize injury 

during the collection process and maintain the culturability of the collected 

microorganisms. If the sample will not be cultured, then the samples usually can be 

collected dry using a membrane filter, cyclone, impactor, or a combination of these. Dry 

collection is typically simpler and less expensive to perform, and filters and cyclones can 

handle a wide range of particle concentrations. Organisms that are difficult or impossible 

to grow in culture are often collected using dry techniques and assessed using polymerase 

chain reaction (PCR) based methods, which have the advantage of speed and specificity. 

PCR has been used for rapid detection of Histoplasma capsulatum and mycobacteria [Reid 

and Schafer 1999; Schafer et al. 1999; Schafer et al. 2003]. A DNA-based mold specific 

quantitative PCR (msQPCR) method is widely used to evaluate indoor fungal bioaerosols 

in the academic, government and commercial sectors, and is the basis for the 

Environmental Relative Moldiness Index (ERMI) used to quantify mold contamination in 
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 homes [Kettleson et al. 2015; Vesper et al. 2013]. The ERMI and other PCR-based assays 

are discussed in greater detail later in this chapter.

If viability is to be studied, then the samples usually will need to be collected with an 

impinger or in an Andersen impactor loaded with agar plates, because many 

microorganisms will lose viability due to damage or desiccation if collected dry [Cox 1987; 

Hung et al. 2005]. For example, a membrane filter sampler is not appropriate for sampling 

culturable Escherichia coli because the cells desiccate and become either nonviable or 

viable but not culturable under these conditions [Jensen et al. 1992]. Similar results have 

been reported for other bacteria and fungi [Macher and First 1984; Wang et al. 2001]. 

Depending upon the target microorganism, impingers may be filled with distilled water or 

a buffered isotonic solution, sometimes with antifoaming agents to reduce foaming and 

proteins to enhance survival. Mineral oil has also been used in impingers instead of 

aqueous solutions to avoid evaporation [Lin et al. 2000]. Impactors are loaded with agar 

plates; the choice of agar depends upon the microorganisms of interest and the desired 

selectivity (discussed in the next section).

As noted previously, depending upon the investigation that is being conducted, the 

particle size distribution of the bioaerosol may be very important in the evaluation of the 

data obtained. If particle size information is needed to, for example, determine how much 

of the bioaerosol is in the respirable size fraction, then a size-selective sampler should be 

used for at least some of the collections if possible. For example, if an SAS-Compact 

sampler was the selected sampler for collection of culturable Escherichia coli, an Andersen 

6-Stage sampler could be used to determine the particle size distribution at each location 

sampled. The expected size of the bioaerosol particles is also an important factor in 

choosing a sampler. For example, an impactor with a d50 of 4 μm should not be used to 

collect Aspergillus niger spores (dae 1-3 μm) because most spores would remain entrained 

in the air and pass through the instrument.

NIOSH Method 0800 discusses sampling for culturable airborne bacteria and fungi with 

an Andersen cascade impactor [NIOSH 2003b]. Standard methods for the collection of 

airborne fungi by inertial impaction are presented in ASTM Standards D7788 [ASTM 

2009; ASTM 2014d]. ASTM Standard D7391 also discusses the aspects related to the 

laboratory analysis.

b. Sampling for airborne viruses

Airborne viruses are more difficult to study in bioaerosols than bacteria and fungi for a 

variety of reasons [Prussin et al. 2014; Verreault et al. 2008]. Viruses are more difficult to 

culture because they are obligatory intracellular parasites that require a host cell for 
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 reproduction [Tortora et al. 2013]. Bioaerosols of pathogenic viruses have been found in 

many settings to be present in low concentrations that can be difficult to detect [Blachere 

et al. 2009; Bonifait et al. 2015; Lindsley et al. 2010a; Tseng et al. 2010; Yang et al. 2011]. 

Viruses also are generally more susceptible to damage during aerosol collection than are 

bacteria or fungi, although their sensitivity varies widely with the collection method and 

species [Appert et al. 2012; Turgeon et al. 2014; Zuo et al. 2013]. Aerosol sampling 

methods for viruses have been reviewed by Verreault et al. [2008].

Bacteriophages are viruses that infect bacteria rather than multicellular organisms. They 

are used in laboratory aerosol studies as tracers for aerosol particles and as surrogates for 

airborne viruses that infect humans [Fisher et al. 2012; Tseng and Li 2005; Turgeon et al. 

2014]. Bacteriophages are not known to be hazardous to humans but are of interest to 

industries that rely on bacteria such as cheese manufacturers [Verreault et al. 2011].

Polymerase chain reaction (PCR) based methods are often used to study viral bioaerosols. 

PCR has the advantages of being very sensitive and very specific, and considerably easier 

to perform than viral cultural assays. For this reason, most recent field studies of airborne 

viruses have used PCR as the detection method. Examples include studies of viruses in 

healthcare facilities [Blachere et al. 2009; Booth et al. 2005; Lindsley et al. 2010a; 

Thompson et al. 2013; Tseng et al. 2010], influenza at poultry and pig farms [Corzo et al. 

2013; Jonges et al. 2015], airborne viruses in a sewage treatment plant [Masclaux et al. 

2014], and respiratory viruses in human coughs and exhaled breath [Gralton et al. 2013; 

Lindsley et al. 2010b; Milton et al. 2013].

PCR has both the advantage and disadvantage of not requiring that the virus be viable in 

order to be detected. This eliminates the need to preserve viability during and after 

collection and allows the use of dry collection methods such as cyclone samplers, dry 

impactors and filters, which are simpler and easier to carry out. On the other hand, this 

also means that it is unclear whether the airborne virus is infectious or not, which makes 

interpretation of data more difficult. This is a common criticism of PCR-based bioaerosol 

studies.

If the virus in a bioaerosol sample is to be cultured, in most cases the sample will need to 

be collected into an aqueous media using an impinger or wetted surface aerosol collector. 

Fabian et al. [2009] showed that collecting airborne influenza virus in aqueous media 

using an SKC BioSampler preserved infectivity much better than dry collection using 

filters or an impactor. A less-common method is to collect viable viruses using an 

Andersen impactor. Gustin et al. [2011] collected airborne influenza virus using an 

Andersen impactor by placing a filter and a thin layer of gelatin on top of the agar in the 

culture plates. After collection, the gelatin was removed and melted at 37°C to allow 
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Sampling and Characterization of Bioaerosols 

 subsequent culture of the virus. Note that the collection media must be compatible with 

the cell culture system used to host the virus.

6 Sample preparation for culturable bioaerosols

Collecting and culturing viable airborne microorganisms is the most common technique used 

by industrial hygienists to assess bioaerosols [Macher 1999]. However, the appropriate sample 

preparation method is highly dependent upon the microorganism(s) of interest, sample 

source, and down-stream analysis. These sampling approaches are further confounded as 

viable bioaerosols have been estimated to account for approximately 1% of the total bioaerosol 

load, and non-viable bioaerosols are often overlooked [Hung et al. 2005]. In contrast, nonculturable bacteria and fungi cannot be grown in conventional lab-based conditions, but their 

presence is still important from a health perspective [Green et al. 2011; Mitakakis et al. 2003]. 

Non-viable bioaerosols can be determined through other detection methodologies such as 

microscopy, proteomic, immunological, and molecular analysis methods, and some of these 

approaches are discussed in section 9. A list of the common bioaerosols encountered in 

indoor and outdoor environments, as well as the fungi that are common contaminants of 

indoor building materials, can be found in Flannigan et al. [2011].

a. Sample preparation for bacteria and fungi

Viable bacterial and fungal bioaerosol identification is made through the collection, 

deposition, and growth of a viable propagule or intact cell on a selected nutrient agar 

medium contained in a sterile petri dish or liquid culture suspension [Macher 1999]. 

These methods are similar for both fungal and bacterial bioaerosols [Flannigan et al. 2011; 

Hung et al. 2005]. Selection of the nutrient media, incubation conditions (time and 

temperature), and potential damage to the culturable bioaerosol during sampling are 

among several critical variables to review before the collection, growth and proliferation of 

a viable propagule [Eduard et al. 2012; Hung et al. 2005; Macher 1999]. These parameters 

have been reviewed elsewhere, but should be taken into consideration when planning an 

environmental survey [Hung et al. 2005; Macher 1999].

Growth media can be defined as either broad or selective [Macher 1999]. As the term 

implies, broad nutrient media supports the growth of a diverse number of 

microorganisms. In contrast, a selective growth medium, with appropriate energy sources, 

nutrients, and pH, is used to enrich growth of the specific microorganism in question and 

inhibit the growth of competitive organisms [Macher 1999]. A variety of broad and 

selective nutrient media for bacteria and fungi are available to the industrial hygienist and 

can be found in Hung et al. [2005] and Macher [1999].
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 Following sample collection, liquid or agar cultures are incubated at a suitable temperature 

and atmosphere (facultative versus aerobic) for an appropriate time. Fast-growing bacteria 

may develop microcolonies in hours, while fungi may take days to develop into a visible 

colony and perhaps sporulate. Organisms such as Mycobacterium tuberculosis or the 

dimorphic fungal pathogens, Histoplasma capsulatum or Blastomyces dermatitidis may 

require weeks of incubation to produce visible colonies [ATS 1990; Babady et al. 2011]. 

For fungi, plates are typically incubated at room temperature (18°C-25°C) or, if it is a 

clinically relevant isolate, at 35°C [ACGIH 1989; Baron and Finegold 1990; Hung et al. 

2005; Macher 1999]. In contrast, environmental bacteria are grown between 18°C and 

28°C, while thermophilic bacteria are grown between 50°C and 58°C [Hung et al. 2005; 

Macher 1999]. 

After allowing for vegetative growth of all viable propagules on the selected nutrient 

medium, the number of colonies is identified, quantified and presented as colony forming 

units (CFUs) [Eduard et al. 2012]. Media blanks (laboratory and field) should be processed 

using the same methods as samples to control for environmental or laboratory 

contaminants. A collection of bioaerosol identification manuals is presented in both the 

AIHA and ACGIH manuals [Hung et al. 2005; Macher 1999]. Color micrographs of 

common fungal contaminants are also presented in Flannigan et al. [2011]. Along with the 

quantification of viable microorganisms, taxonomic data and an interpretation of the 

datasets are generally reported [Hung et al. 2005].

The interest in detecting and quantifying fungi has increased following consensus 

documents that reported associations between fungi in damp indoor environments and 

adverse respiratory health effects [IOM 2004; Mendell et al. 2011; WHO 2009]. Compared 

to bacteria, additional variables need to be taken into consideration by the industrial 

hygienist when evaluating viable fungal bioaerosols including water activity, colony 

competition, and carbohydrate nutrient sources [Hung et al. 2005; Macher 1999]. Broad 

viable culture approaches favor species belonging to the phylum Ascomycota, as well as 

species that outcompete slower-growing species. Several different types of media and 

physiological conditions (e.g. temperature) may also need to be employed to assess 

complete fungal diversity using this approach. For fungi, selection of the nutrient media 

may potentially bias the growth of specific viable fungal bioaerosols. Common nutrient 

media include malt extract agar (MEA) supplemented with chloramphenicol or rose 

bengal agar to suppress bacterial growth [Hung et al. 2005; Macher 1999]. Cellulose agar 

can also be used for the selection of indoor fungal contaminants such as Stachybotrys 

chartarum [Hung et al. 2005]. Dichloran glycerol (DG18) can be used to select for those 

fungi that are xerotolerant [Flannigan et al. 2011; Hocking and Pitt 1980; Macher 1999]. 

Temperature and incubation time can also be used to select for specific fungal bioaerosols 

such as Aspergillus fumigatus which are capable of growth within human hosts [Flannigan 
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 et al. 2011; Hung et al. 2005]. A selection of nutrient media and growth conditions that can 

be used for viable fungal culture can be found in Hung et al. [2005], Flannigan et al. [2011] 

and Macher [1999].

b. Sample preparation for viruses

Because viruses are obligate intracellular parasites, special precautions must be taken in an 

effort to minimize damage to the collected virus-laden aerosol. Environmental factors 

such as humidity, temperature and gas composition of the air can significantly impact the 

infectiousness of a virion and should be monitored closely [Ijaz et al. 2016]. Several studies 

have shown that the inactivation of an airborne virus is directly related to the relative 

humidity and temperature [Weber and Stilianakis 2008]. In one study, high humidity 

levels caused a loss of infectious influenza virus from simulated coughs [Noti et al. 2013]. 

Similarly, using a ferret animal model, Lowen et al. [2007] demonstrated a correlation 

between airborne transmission of influenza and the relative humidity and temperature. 

Through the use of an ozone-oxygen delivery system, researchers were able to show that 

ozone-mediated reactive oxygen species (ROS) caused lipid peroxidation and subsequent 

damage to the lipid envelope and viral capsid [Murray et al. 2008]. Such studies highlight 

the importance of collecting viral aerosols under optimal environmental conditions and, 

when possible, minimizing the detrimental effects of environmental factors on collected 

samples.

Air sampling techniques also may cause damage to the virus and compromise analysis. 

Before collecting viral aerosols, the hardiness of the target virus must be taken into 

account. Currently, there are over 200 known respiratory viruses that fall under one family 

of DNA viruses (Adenoviridae) and four families of RNA viruses (Orthomyxoviridae, 

Paramyxoviridae, Picornaviridae and Coronaviridae) [Abed and Boivin 2006]. While all 

viruses package their genome in a protective protein coat known as the capsid, some 

viruses also possess a lipid bilayer envelope that, as the name implies, surrounds the viral 

capsid. Once outside the host, the viral envelope is highly sensitive to desiccation, 

temperature fluctuations and readily undergoes degradation. Variations in temperature 

can greatly affect viral enzymatic activity and nucleic acid stability [Tang 2009]. As noted 

earlier, viruses with lipid envelopes tend to survive better at low humidity, while nonenveloped viruses survive better at high humidity [Tang 2009]. Also, RNA viruses are 

inherently more unstable than DNA viruses due to the presence of the 2’-hydroxyl group 

on the ribose sugar molecule of RNA that is susceptible to base-catalyzed hydrolysis and 

degradation. Therefore it is critical that collection methods do not disrupt the lipid 

membrane and/or compromise the integrity of nucleic acids. Impairment of either the 

lipid envelope or nucleic acids can significantly impact detection of the viral aerosol and 

lead to false negatives.
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 To optimize collection efficiency while maintaining infectiousness of the viral aerosol, 

researchers must be discriminating when deciding on what type of aerosol sampler to use 

and how long the sampling collection period should be. As discussed in Section 3, some 

commercially available bioaerosol sampling devices exist, each of which possesses unique 

collection properties. In a review by Verreault et al. [2008], liquid impingers were found to 

be the most effective sampling devices for capturing small viral particles while maintaining 

virion integrity and infectiousness [Verreault et al. 2008].

While liquid impingement preserves and maintains viral integrity (in comparison to dry 

impaction), the type of collection medium must also be considered. With culture-based 

identification methods, it is of utmost importance to maintain the stability of the collected 

viruses while using cell-culture compatible media. The type of liquid medium, as well as 

the volume used, are important variables to consider. Virus collection and transport media 

are typically isotonic solutions with a buffer to control the pH, protein to protect the virus, 

and antibiotics to prevent microbial growth. If used in an impinger, the viral collection 

media may also include an antifoaming agent. Specimen handling, storage time and 

storage temperature can significantly impact the integrity of sample analysis. Collection 

and storage under suboptimal conditions can result in viral inactivation and degradation 

of nucleic acids. Bioaerosol samples containing viruses should be processed as soon as 

possible after collection. They should be refrigerated or frozen and transported as quickly 

as possible. The stability and retention of viability depend upon the virus [Johnson 1990].

7 Identification of culturable bioaerosols

Identification of the microbial taxa is a critical element in the determination of the viable 

bioaerosol load in an industrial or occupational environment. The science of classification, 

especially the classification of living forms, is called taxonomy. The objective of taxonomy is 

to classify living organisms to establish the relationship between one group of organisms and 

another, and to differentiate between them based on phenotypic and genotypic characteristics. 

The identification of viable fungal bioaerosols has been challenging due to the confusion of 

current nomenclature [Flannigan et al. 2011]. As a result, investigators often use synonymous 

names that over time have been placed in another group. Familiarity with taxonomy and 

nomenclature is critical when undertaking assessments of the viable microbial burden in an 

environment [Flannigan et al. 2011]. Several criteria and methods for the classification of 

culturable microorganisms are briefly discussed in the following subsections. Besides using 

these methods, the nonviable and non-culturable methods of identification discussed in 

Section 9 may also be used in combination with these viable methods.

Classical microbiology includes general methods for classifying or identifying 

microorganisms. The least specific of these is the observation of growth characteristics. 
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 Growth characteristics include the appearance of the microorganisms in a liquid medium, 

colony morphology on solid medium, pigmentation, and arrangement of reproductive 

structures such as fungal sexual or asexual spores.

a. Bacteria 

Bacteria are prokaryotes with distinguishing morphological characteristics that include the 

cell shape, cell size, arrangement of cells, and the presence or absence of flagella, capsules, 

or endospores. Simple and differential staining may be performed on bacteria to enhance 

visualization and to aid in grouping and identification [Tortora et al. 2013]. In simple 

staining, a single basic dye is used that highlights the cellular morphology. Stains such as 

methylene blue, carbolfuchsin, crystal violet, or safranin may be used for bacteria. 

A differential stain distinguishes among structures or microorganisms based on varying 

reactions to the staining procedure. Two examples of differential stains are the Gram stain 

and the acid-fast stain. In Gram staining, bacteria are stained and then washed with 

alcohol. Gram-positive bacteria possess a cell wall composed of a relatively thick 

peptidoglycan layer and teichoic acids, which retains the dye complex. Gram-negative 

bacteria possess a cell wall composed of a thin peptidoglycan layer and an outer membrane 

which consists of lipoproteins, lipopolysaccharides, and phospholipids, and do not retain 

the dye complex when washed [Tortora et al. 2013]. A few of the commercially available 

identification kits require a Gram-stain prescreening to assure that the correct reagents are 

used. Acid-fast stains are used for some species of bacteria, particularly those of the genus 

Mycobacterium, which do not stain readily. In the acid-fast staining process, the 

application of heat facilitates the staining of the microorganism [Tortora et al. 2013].

b. Legionella 

Bacteria that are placed in the genus Legionella, are the etiological agents of pulmonary 

infections called Legionellosis [Fields 2002]. Legionella pneumophila (Figure 1B) is the 

most widely known species that has been implicated in Legionnaires’ disease, which can 

result in pneumonia. Milder illness with fever and body aches is referred to as Pontiac 

fever. Bacteria placed in this genus consist of Gram negative rods and are associated with 

freshwater in the environment [ASTM 2015; Macher 1999]. Exposure to warm 

temperatures (25-42°C) can result in the growth and proliferation of the bacteria, a 

problem that has emerged in water and air handling systems within the indoor built 

environment [Hung et al. 2005]. Growth and persistence within Protista have also been 

reported [Hung et al. 2005]. For health care facilities, ASHRAE Guideline 12–2000 

recommends storing and distributing cold water at <20°C (68°F), whereas hot water 

should be stored at >60°C (140°F) and circulated with a minimum return temperature of 

51°C (124°F). In other settings, hot water should be stored at ≥40°C (≥120°F) [ASHRAE 
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 2000]. Building air conditioning cooling towers, humidifiers, or structures used for 

bathing such as hot tubs are particularly susceptible to Legionella amplification. Legionella 

can be aerosolized within water droplets via abiotic disturbance mechanisms and 

disseminated into the breathing zone of the subject. Airborne levels lower than 10 

CFU/mL have been associated with Legionnaires’ disease [ASTM 2015; Demirjian et al. 

2015; Hung et al. 2005].

Since the identification of L. pneumophila and association with Legionnaires’ disease in 

1976, there have been many studies that have focused on a variety of approaches to detect 

and mitigate this bacterial species from the built environment. These approaches are 

broadly reviewed in Hung et al. [2005]. Along with industrial hygiene practices and 

building maintenance, environmental monitoring and surveillance programs are critical to 

ensure effectiveness of employed engineering controls and maintenance/disinfection 

programs [ASTM 2015]. ASTM International has published a standard for inspecting 

water systems and investigating outbreaks [ASTM 2015]. The CDC has also published a 

sampling procedure [CDC 2015].

In 2015, ASHRAE published a consensus standard for the primary prevention of 

Legionnaires’ disease in building water systems [ASHRAE 2015]. Similar environmental 

assessment methods are utilized in maintenance programs and outbreak cases. In this 

approach, bulk water samples are typically collected (250 mL to 1 L for non-potable and 

1000 mL for potable water), concentrated via filtration, resuspended, and then plated on a 

growth medium (such as buffered charcoal yeast extract media) to enable the propagation 

and identification of Legionella spp. [CDC 2015]. Samples may also be direct plated, acid 

treated, or heat treated to enhance the recovery of the bacterium. Colonies represent the 

viable fraction of Legionella in a water sample and these colonies are quantified and 

reported as CFU/mL

 

In addition to viable culture-based approaches, molecular-based methods such as PCR as 

well as antibody-based methods have been developed to enable the detection of Legionella. 

Air sampling is not considered a reliable method for Legionella surveillance in the built 

environment [Hung et al. 2005]. 
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 Using an environmental microbiology laboratory with expertise in propagating Legionella

spp. is important when evaluating Legionella contamination of water systems within the 

built environment. A number of laboratories participate in the CDC’s Environmental 

Legionella Isolation Techniques Evaluation (ELITE) Program. Participation in the 

program is voluntary and enables laboratories to test their proficiency in Legionella

isolation and identification techniques against standardized samples. A list of ELITE 

member laboratories can be accessed at

https://wwwn.cdc.gov/elite/Public/MemberList.aspx. 

c. Fungi 

In general, culturable fungi are classified by colony features including the septation of 

hyphae and colony morphological phenotypes, including pigmentation and the 

presentation of asexual and sexual spores on hyphae. Stains such as Calberla’s solution, 

lactophenol cotton blue, periodic acid-Schiff stain, Grocott’s methenamine silver stain, 

and calcofluor white may be used in combination with potassium hydroxide (10% KOH) 

to resolve these colony structures using microscopic-based approaches [Hung et al. 2005]. 

The identification and classification of fungal colonies should be performed by an 

examiner that is skilled in microbiology and mycology. A number of commercial labs that 

employ examiners skilled in the identification of microorganisms encountered in indoor 

and occupational environments are accredited by the AIHA Environmental Microbiology 

Laboratory Accreditation Program (EMLAP). 

In addition to viable culture-based approaches, biochemical, physiological, and nutritional 

tests for bacteria and fungi can be used [Flannigan et al. 2011]. These testing strategies 

offer identification based on numerous variables including cell wall constituents, pigment 

biochemicals, storage inclusions, antigens, optimum temperature and temperature range, 

the effect of oxygen on growth, pH tolerance, osmotic tolerance, salt requirement and 

tolerance, antibiotic sensitivity, energy sources, carbon sources, nitrogen sources, 

fermentation products, and modes of metabolism (autotrophic, heterotrophic, 

fermentative, respiratory). As a rule, batteries of such tests, rather than any one individual 

test, are used to identify or classify microorganisms. A few commercially available test 

batteries are discussed briefly in the section on biochemical approaches.
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8 Enumeration of culturable bioaerosols

a. Enumeration of bacteria and fungi

The total concentration of culturable airborne microorganisms in a sample is determined 

by collecting the bioaerosol sample on a culture plate or plates (or inoculating culture 

plates with a bioaerosol sample), incubating the plates, and dividing the number of 

colonies observed on the culture plates by the volume of air sampled. Note that, as 

discussed in section 2, the number of colonies counted on an agar plate from a bioaerosol 

impactor must be adjusted using a positive-hole correction factor to correct for multiple 

microorganisms depositing beneath an impactor hole [Andersen 1958; Leopold 1988; 

Macher 1989]. A colony is defined as a macroscopically visible growth of microorganisms 

on a solid nutrient medium. Concentrations of culturable bioaerosols collected during air 

sampling are normally reported as colony forming units (CFU) per unit volume of air 

[Eduard et al. 2012]. CFUs also can be determined from samples collected in a swab or 

dust sample collected from the floor or area of contamination [Hung et al. 2005].

Often, it is difficult to identify multiple colonies at one location on a plate because of the 

lack of differential colony morphology [Burge et al. 1977]. In addition, some organisms 

produce large, spreading colonies while others produce microcolonies. Analysis of plates 

containing multiple types of microorganisms can be difficult because the chemicals 

secreted by one microorganism might inhibit the growth of other microorganisms at that 

same location [Burge et al. 1977]. The morphology of the colony of one microorganism 

also may completely obscure that of another, and a fast-grower might obscure a slowgrower.

b. Enumeration of viruses

Before the advent and mainstreaming of molecular-based detection methodologies, cell 

culture-based methods and serological assays were considered the gold standard for the 

detection of viral pathogens. Typically, through the use of commercially available 

immortal cell lines, researchers can screen collected bioaerosols by inoculating cells and 

looking for common cytopathic effects (CPEs) such as rounding of infected cells, fusion 

with adjacent cells and lysis of cells. Examples of well-known cell lines that are routinely 

used in viral diagnostics include primary rhesus monkey kidney (RhMK) cells, primary 

rabbit kidney cells, human lung fibroblasts (MRC-5), human epidermoid carcinoma cells 

(HEp-2), human lung carcinoma cells (A549) and Madin Darby kidney cells (MDCK) 

[Leland and Ginocchio 2007]. Selection of the appropriate cell line is based on the 

specimen source and the suspected causal viral pathogen. Certain viral pathogens may 

require several cell passages before CPEs can be observed. More information on 

enumeration assays for viable viruses can be found in standard virology reference texts, 

such Principles of Virology [Flint et al. 2009b] and Fields Virology [Fields et al. 2007].
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Viral plaque assay

A widely used approach for detecting viral pathogens and quantifying viral titers is the 

viral plaque assay (VPA) [Condit 2007]. Under biological safety controls, the 

appropriate cell line is propagated and plated, usually in a 6-well format, at a 

concentration at which cells form a monolayer. The cell monolayer is next treated for a 

defined period (30-60 minutes) with a specific volume (0.1 mL to 1.5 mL) of the 

collected bioaerosol and incubated at the specified temperature and CO2 levels for 24 

to 72 hours. Throughout the incubation period, cells are routinely inspected and CPEs 

are documented. Upon completion of the incubation period, cells are chemically fixed, 

stained and plaques (zones of cellular clearing) are enumerated. The final 

concentration of the collected viral aerosols is calculated based on the number of 

plaques, dilution of the inoculum and volume plated, and is expressed in plaque 

forming units per mL (PFU/mL]). While the VPA is a cost-effective method of 

assessing sample viral loads, results can take anywhere from 3-5 days. Other 

limitations such as the inability to detect low viral titers and inactivated 

(noninfectious) virus and the failure of some viruses to form plaques, may 

compromise detection and underestimate the viral loads in an aerosol sample. 

Likewise, common indoor and outdoor contaminants (such as fungi and bacteria) can 

impair the VPA by disrupting the cellular monolayer or outcompeting for nutrients in 

the cell culture medium.

Tissue culture infectious dose assay

Another cell culture-based approach to identifying viral aerosols is the Tissue Culture 

Infectious Dose assay (TCID50), also known as an endpoint dilution assay [Condit 

2007]. As with the VPA, select cells are plated at a desired concentration in a 96-well 

format and inoculated with serial dilutions of the collected sample. Following a 

specified incubation period, cells are examined for CPEs. The TCID50 is defined as the 

dilution of virus required to infect 50% of the cell culture wells [Reed and Muench 

1938]. Based on the number of cells that are infected at the designated virus dilution, 

viral titers are mathematically calculated. Limitations of the TCID50 are similar to 

those observed with the VPA.

Immunofluorescence antibody (IFA) assays

To enhance viral detection and quantify viral loads, immunofluorescence antibody 

(IFA) assays (direct or indirect) are frequently used in combination with cell culturebased methods [Flint et al. 2009a; Tortora et al. 2013]. By combining infected cells 

with a fluorescently-labeled, antigen-specific antibody, it is possible to increase 

detection levels without the lengthy incubation periods that are typically necessary 

with VPAs and TCID50 assays.
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c. Interpretation of data

In industrial hygiene surveys that evaluate bioaerosols, indoor bioaerosol levels are usually 

compared to outdoor levels or to a control area. In general, indoor bioaerosol levels are 

lower than outdoor levels, and the taxa are similar [ACGIH 1989; Burge et al. 1977; Hung 

et al. 2005; Macher 1999; Solomon et al. 1980]. However, elevated indoor bioaerosol levels 

may be a sign of dampness, water infiltration, or microbial contamination [Hung et al. 

2005]. In 2010, The ACGIH published a variety of occupational exposure limits for 

aerosols that are derived from biological material in specific industries and include 

subtilisins derived from Bacillus subtilis, as well as cotton, grain, flour, wood, and organic 

dusts [ACGIH 2015; Eduard et al. 2012]. To date, no occupational exposure limits for 

specific fungal bioaerosols exist and these typically fall under particulate matter not 

otherwise regulated (10 mg/m3 for inhalable dust; [ACGIH 2015; Eduard et al. 2012]. 

Proposed limits developed in other regions of the world are provided in Eduard et al. 

[2012]. 

Although the quantification of viable fungal propagules can provide helpful datasets to 

evaluate differences between indoor and outdoor fungal diversity, the interpretation of 

results should be evaluated closely. Total fungal exposure will be underestimated as nonviable fungal bioaerosols are not captured in the analysis [Eduard et al. 2012]. Fungal 

genera, including Cladosporium, Alternaria, and Epicoccum, and Basidiomycetes are 

predominantly localized in outdoor environments and the presence of elevated 

concentrations may be an indicator of indoor fungal contamination [Hung et al. 2005]. 

Similarly, the presence of certain hydrophilic species including Stachybotrys chartarum

and Aspergillus versicolor, and Chaetomium globosum may be signs of indoor fungal 

contamination and may require immediate inspection [Flannigan et al. 2011; Hung et al. 

2005]. 

Where local amplification and dissemination of bacteria have not occurred in an occupied, 

indoor environment, Gram-positive cocci (e.g., Micrococcus and Staphylococcus) are 

normally dominant [Morey et al. 1986]. Airborne human skin scales and respiratory 

secretions may contain Gram-positive cocci [ACGIH 1989; Hung et al. 2005]. Detection of 

high levels of these microorganisms may be an indication of over-crowding and 

inadequate ventilation. Indoor air that tests high for Gram-negative bacteria indicates a 

need to identify and eliminate the source of contamination. Concentrations ranging from 

4,500-10,000 CFU/m3 have been suggested as the upper limit for ubiquitous bacterial 

aerosols [ACGIH 1989; Nevalainen 1989]. These exposure limits, however, do not apply to 

pathogenic microorganisms. Actinomycetes (mesophilic and thermophilic) are commonly 

found in agricultural areas. Their presence in indoor environments is an indicator of 

contamination [ACGIH 1989; Banaszak et al. 1970; Lacey and Crook 1988]. Thermophilic 
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 Actinomycetes at concentrations above 70 CFU/m3 in an affected person's work area have 

been regarded as the threshold for triggering remedial action [Otten et al. 1986].

9 Sample analysis methods for non-viable and 

non-culturable bioaerosols

The collection and classification of nonviable and non-culturable microorganisms cannot be 

performed by using viable culture methods. A large proportion of fungal bioaerosols are nonviable and would not grow and proliferate on nutrient media [Eduard et al. 2012]. This 

fraction of the bioaerosol load is equally important to assess in industrial hygiene surveys that 

investigate the role of personal bioaerosol exposure on respiratory health [Brasel et al. 2005; 

Green et al. 2011; Mitakakis et al. 2003]. These bioaerosols can also contain antigens, 

allergens, microbial volatile organic compounds and even mycotoxins [Brasel et al. 2005; 

Eduard et al. 2012; Green et al. 2011; Green et al. 2006b]. Identification of nonviable or nonculturable microorganisms or components of microorganisms (such as cell wall fragments) 

can be performed using a variety of other available assessment strategies such as microscopy, 

immunoassays and, more recently, molecular biology techniques [Afanou et al. 2015; Brasel et 

al. 2005; Eduard et al. 2012; Flannigan et al. 2011; Green et al. 2011; Hung et al. 2005; Macher 

1999; Rittenour et al. 2012]. 

Microscopy includes a variety of approaches that utilize bright-field, light, phase contrast, 

fluorescence or even electron-based approaches [Eduard et al. 2012; Macher 1999]. These 

methods enable the enumeration of both viable and nonviable microorganisms [Macher 1999] 

as well as other non-culturable bioaerosols including cell wall fragments, plant pollen and 

pteridophyte and bryophyte spores [Green et al. 2011; Rittenour et al. 2012]. These 

approaches provide a platform to visualize particle morphology and to identify reproductive 

fungal structures of individual genera that are based on a combination of propagule 

phenotypes [Flannigan et al. 2011; Macher 1999]. However, these approaches can be 

confounded by observer bias, especially when it comes to differentiating bioaerosols that 

contain similar morphological phenotypes such as amerospores (e.g. Aspergillus conidia) 

[Flannigan et al. 2011; Hung et al. 2005]. The ASTM has published a standard method for the 

use of optical microscopy to categorize and quantify fungal structures in samples collected by 

inertial impaction [ASTM 2009].

To overcome these methodological challenges, alternative methods based on the 

quantification of bioaerosol biomarkers (proteins or DNA) have been developed that enable 

the quantification of these bioaerosol sources [Eduard et al. 2012]. These include a variety of 

assessment methods that can be used to qualitatively and quantitatively assess bioaerosol 

exposure by detecting cell wall components, proteins, carbohydrates, or oligonucleotides (e.g. 
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endotoxin or β-glucan) [Eduard et al. 2012]. Other chemical and proteomic methods 

including HPLC, flow cytometry, and mass spectrometry-based approaches can also be used 

to detect and quantify cell wall components such as microbial volatile organic compounds and 

mycotoxins. These approaches have been reviewed more extensively by Flannigan et al. 

[2011]. 

Several modifications of classical biochemical procedures have been used in recent years to 

facilitate inoculation of media, decrease the incubation time, automate the procedure, and 

systematize the determination of species based on reaction patterns. Historically, clinical 

microbiological techniques have been used for analysis of environmental samples. However, 

clinical strains and environmental isolates may differ, requiring modification of clinicallybased techniques.

a. Microscopy

Bright-field or light

In bright-field or light microscopy, an ordinary microscope is used for the

morphological observation and sizing of sampled bioaerosols. Visible light from an

incandescent source is used for illumination and the specimen appears against a bright

backfield. Objects smaller than 0.2 μm cannot be resolved. The image contrast

(visibility) decreases as the refractive index of the substance/microorganism under

observation and the mounting medium become similar. To maximize the contrast, the

mounting medium should have the same refractive index as glass or the immersion oil.

Membrane filters are often "cleared" by using the appropriate immersion oil or acetone

vapor/triacetin combination. This method is commonly used to observe various

stained specimens and to identify and count viable and non-viable bioaerosols. In

addition, pollen grains are often identified and enumerated in this manner [Eduard et

al. 1990].

Collection of fungal bioaerosols onto an adhesive surface followed by microscopic

identification based on the morphological characteristics of the spores (size, shape,

septation etc.) is another common method of assessment. This non-viable method

overcomes limitations introduced in viable analyses and many genera can be

differentiated based on differences in spore morphology [Eduard et al. 2012; Macher

1999]. Microscopic examination of fungi captured on filters or an adhesive tape are

divided into seven spore morphological characteristics and include amero-, didymo-,

helico-, stauro-, dictyo-, phragmo-, and scoleco- spores [Kendrick 2000]. Amerospores

are the most common spore morphology encountered in air samples and are the most

challenging to differentiate taxonomically [Kendrick 2000]. Amerospores are usually

placed in a group represented as Aspergillus/Penicillium group [Hung et al. 2005].
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Many other common environmental fungal bioaerosols share similar morphologies 

which can make taxonomic placement challenging for the untrained or inexperienced 

observer [Eduard et al. 2012]. Typical magnification used in the assessment of fungal 

propagules ranges from 400-1000X. A standard operating procedure for the 

assessment of microscopic non-viable samples is presented in Hung et al. [2005] and 

by ASTM [2009]. 

The confounding factors associated with traditional fungal exposure assessment 

methods have limited our understanding of the spectrum of fungal bioaerosols in 

industrial and occupational environments. Measures using these approaches also 

cannot be acquired in real time. Furthermore, identifying and quantifying the 

complete diversity of fungal bioaerosols using a standardized methodology is critical 

in the determination of fungal bioaerosols within occupational environments [ASTM 

2009; Hung et al. 2005].

Phase contrast

Phase-contrast microscopy is used when the microorganism under observation (e.g., 

Escherichia coli) is hyaline and an alternative mounting medium is not possible. As 

light passes through the specimen, variations in the index of refraction of the 

components cause phase shifts in the light. A phase-contrast microscope uses a special 

condenser and diffraction plate that cause these phase shifts to appear as differences in 

brightness and contrast. One cannot see an object exactly matching the refractive 

index of the mounting liquid; however, very slight differences produce visible images. 

This type of microscope is commonly used to provide detailed examination of the 

internal structures of living specimens; no staining is required.

Fluorescence

Fluorescence microscopy uses an ultraviolet or near-ultraviolet source of illumination 

that causes fluorescent compounds in a specimen to emit light. Fluorescence 

microscopy for the direct count of microorganisms has been described in a number of 

studies [Eduard et al. 2012]. Direct-count methods to enumerate microorganisms 

(especially bacteria) have been developed using fluorescence microscopy and some 

stains such as acridine orange, fluorescein isothiocyanate (FITC) and 4’,6-diamidino2-phenyl-indole (DAPI) [Macher 1999; Thermo Fisher Scientific 2014]. Utilization of

stains such as calcofluor white can resolve fungal spores and hyphal structures

[Haghani et al. 2013]. This stain binds to chitin; however, other plant-derived and

insect bioaerosol sources (e.g. dust mite, plant pollen) may also be resolved using this

stain. Viability stains also have been developed and are available commercially for the

detection of viable fungi and bacteria bioaerosols in collected air samples [Thermo

Fisher Scientific 2014].
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Electron

Electron microscopy consists of a beam of electrons that enable structures smaller than 

0.2 μm, such as viruses, to be resolved. Scanning electron microscopy (SEM) and, 

more recently, field emission scanning electron microscopy (FESEM), are approaches 

used to study the surface features of prokaryote and eukaryote cells as well as viruses 

(usually magnified 1,000-10,000X). These bioaerosols are immobilized onto a semisolid filter submicron membrane or in the form of a liquid suspension and a threedimensional image of the area is generated. Images from SEM can provide vital 

information about the size, morphology and concentration of the collected bioaerosol 

[Afanou et al. 2014; Eduard et al. 2012]. However, SEM does not provide information 

on viability of the collected bioaerosol. FESEM has been recently used to resolve fungal 

fragments that are produced from fungal colonies following abiotic disturbance 

[Afanou et al. 2015; Afanou et al. 2014]. Transmission electron microscopy can also be 

used to examine viruses or the internal ultrastructure in thin sections of cells (usually 

magnified 10,000-100,000X), although the image produced is not three- dimensional. 

Compared to other methods of assessment described in this chapter, SEM and 

FESEM-based approaches require a highly trained technician to obtain images from 

bioaerosols captured on filter membranes.

b. Endotoxin assays

The lipopolysaccharide endotoxin is a virulence factor possessed by all Enterobacteriaceae

(as well as other Gram-negative bacteria) that is found in the outer membrane of the cell

wall. Airborne endotoxin has been found in high concentrations in agricultural, industrial,

and office environments [Eduard et al. 2012; Milton et al. 1990; Rylander and Vesterlund

1982; Singh et al. 2011b]. Individuals may experience disseminated intravascular

coagulopathy, respiratory tract problems, cellular and tissue injury, fever, and other

debilitating problems. Endotoxin can be detected in air samples collected on glass filters

using the Limulus amebocyte lysate (LAL) assay [Eduard et al. 2012]. This assay uses

amebocytes from the blood-like circulating fluid of the Limulus polyphemus (horseshoe

crab). After exposure to the lysed amebocyte cells, the chromogenic version of the LAL

enables endotoxins to be quantified [Eduard et al. 2012]. Laboratories use this assay to test

for contamination by Gram-negative bacteria [Baron and Finegold 1990].

Although widely used, endotoxin aerosol measurement techniques lack comparability 

between results obtained in different laboratories because of differing sampling, 

extraction, and analytical methods [Jacobs 1989; Milton et al. 1990; Olenchock et al. 1983; 

Rylander and Vesterlund 1982] and water-insoluble endotoxins are not detected [Eduard 

et al. 2012]. A monoclonal antibody assay has also been developed but it is less sensitive 
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 than the LAL method. Similarly, chemical-based approaches are available to detect 

endotoxins including gas chromatography-mass spectrometry [Eduard et al. 2012].

c. Biochemical analysis methods

Because of the high frequency of isolation of Gram-negative rods in clinical settings, 

several commercial multi-test systems have been developed for identification of members 

of the family Enterobacteriaceae and other pathogenic microorganisms. These 

microorganisms are indistinguishable except for characteristics determined by detailed 

biochemical testing. These systems require that a pure culture be examined and 

characterized. A list of some commercially available identification kits is provided in Table 

III. All of these multitest systems have documented accuracies greater than 90% in clinical 

settings [Baron and Finegold 1990; Koneman 1988]. For fungi, API (Analytab Products, 

Plainview, NY) and BIOLOG can also be used to differentiate yeasts based on the 

respective biochemical and physiological profiles [Flannigan et al. 2011].

d. Chemotaxonomic approaches 

Cellular fatty acids (CFA) of bacteria are structural in nature, occurring in the cell 

membrane or cell wall of all bacteria. When the bacteria are grown under standardized 

growth conditions, the CFA profiles are reproducible within a genus, down to the 

subspecies or strain level in some microorganisms. The Sherlock Microbial Identification 

System (MIS), developed by MIDI (Newark, DE), provides a chromatographic technique 

and software libraries capable of identifying various microorganisms based on their CFA 

composition [Sasser 1990a; Sasser 1990b]. The chromatographic technique is also known 

as gas chromatography fatty acid methyl ester analysis (GC-FAME). MIS has a database 

containing the analysis libraries for culturable Gram-negative and Gram-positive bacteria, 

and yeasts. In a comparison study [Amy et al. 1992], only 8 of 18 isolates, identified by 

either API multitest or MIDI MIS, were identified accurately using BIOLOG multitest. A 

prototype method for extracting and analyzing fungi is currently being distributed by 

MIDI.

e. Chemical-based approaches

A variety of chemical-based approaches are available for the detection and quantification 

of bacteria and fungi in environmental samples [Flannigan et al. 2011; Hung et al. 2005]. 

Common approaches include thermal desorption - gas chromatography - mass 

spectrometry (TD-GC-MS), high performance liquid chromatography, gas 

chromatography-tandem mass spectrometry, and matrix-assisted laser 

desorption/ionization time of flight (MALDI-TOF) mass spectrometry [Flannigan et al. 

2011]. These methods require the formation of a library of markers or spectral signatures 

that are used to discriminate between various prokaryote and eukaryote species. Examples 
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of spectral signatures that have been used for the detection of fungi include microbial 

volatile organic compounds (mVOCs), mycotoxins, ergosterol, 3-hydroxy fatty acids, 

muramic acid as well as intracellular and extracellular proteins. NIOSH Method 2549 is a 

TD-GC-MS based approach that allows for the detection of mVOCs in environmental 

samples [NIOSH 2003d].

f. High performance liquid chromatography

High performance liquid chromatography (HPLC) is commonly used for bioaerosol

“fingerprinting” and biomass determination. Techniques such as proteomics and

identifying variations in chromatographic patterns can be used to determine the source of

airborne material. For example, ergosterol has been used for decades to detect fungal

contamination [Seitz et al. 1979] and even determine taxonomy [Axelsson et al. 1995;

Pasanen et al. 1999; Schnurer 1993]. Keratin analysis can be used to identify bioaerosols

derived from vertebrates and possibly the habitat within an environment [Staton et al.

2013]. Detection can be as straightforward as using UV absorbance or as complex and

specific as employing an ion trap mass spectrometer.

HPLC can be adventitious compared to other methods of analysis; it is an established 

technology, fairly inexpensive after initial equipment costs, fast and accurate. Its 

disadvantages include a lack of specificity inherent with detectors using UV absorbance 

(especially at lower wavelengths), and that complex matrices associated with bioaerosols 

can prove to be troublesome and may require multi-step enhancement procedures such as 

solid-phase extraction. Buffered solvent systems are sometimes required, which can be 

technically difficult to use.

Mycotoxins

Fungal contamination is a concern in food production because it can modify the 

nutritional content of feed and cereal grains and introduce potentially adverse

mycotoxins. Before the use of HPLC, methods of identifying fungal contamination 

were time consuming or missed non-viable organisms which still contributed to the 

biomass [Seitz et al. 1979]. HPLC can be used to detect ergosterol, which is a structural 

sterol nearly universally present in fungi but not naturally present in grains [Pasanen 

et al. 1999]. HPLC has also been used to detect ergosterol, mannitol and arabitol in 

bioaerosols [Buiarelli et al. 2013].

Ergosterol is extracted using a liquid-liquid extraction and concentrated. The clean 

samples are then analyzed by HPLC with UV detection. The ergosterol UV spectrum 

varies significantly from the UV spectrum of higher plant sterols, making it specific to 

fungal contaminants. Though the method is still fairly time consuming, it eliminates 
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the need for fungal culturing and can detect the presence of non-viable fungi. When 

combined with other sterols this information can be used to help determine fungal 

species [Schnurer 1993]. 

In addition to using sterols to identify fungal contamination, mycotoxins can also be 

analyzed by liquid chromatography with mass spectrometry (LC-MS). Identification of 

specific mycotoxins can help identify the species of fungi present [Bennett and Klich 

2003; Castillo et al. 2016]. 

Mycotoxins can play a role in indoor air quality (IAQ), food safety and possibly 

bioterrorism. The use of LC-MS as a screening tool to identify mycotoxins can reduce 

the use of more intensive molecular techniques for identification and quantitation. 

When using mass spectral analysis it is important to have a reliable and accurate 

database for identification and a qualified analyst to correctly interpret data.

Other biomolecules

More recently, HPLC has been proposed as a forensic tool to identify and track 

vertebrate species using keratin profiles. Like ergosterol in fungi, keratin is found 

mainly in dander left by vertebrates [Plowman 2007]. If patterns can be established, it 

may be possible to identify what species had been present in a specific dwelling and 

possibly even track movements [Staton et al. 2013]. Bacterial contamination can also 

be tracked using endotoxin analysis and bacterial peptidoglycan fingerprinting [Staton 

et al. 2013].

Sample preparation and enhancement

Bioaerosols can have complex matrices with many interfering constituents. Ultraviolet 

absorption detectors are fairly inexpensive and straightforward to use, but they can 

suffer from a lack of specificity and sensitivity. Although mass spectrometry detectors 

do not suffer from a lack of specificity or sensitivity, a dirty sample can still present 

challenges. 

Another potential pitfall is the presence of large particles in bioaerosol samples. In 

general, analytical HPLC systems and detectors use small diameter tubing and small 

orifice injectors that are easily clogged or contaminated by particles. Most analytical 

systems have some sort of less-expensive trapping or pre-column that can be sacrificed 

in order to spare the more expensive analytical columns. However, trap columns are 

still very much an expense and should only be considered if other options are not 

available. A multitude of cleanup procedures are available to lessen or eliminate 

problems due to particles, the most common of which are simple centrifugation and 
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filtration. Both methods will lessen the likelihood of clogging or damaging the system, 

but neither offers target analyte enrichment.

Liquid-liquid extraction

In order to enrich analytes, samples can be concentrated, chemical interferences can be 

removed, or a combination of both methods can be used. One of the oldest methods of 

enrichment is a liquid-liquid extraction, which separates analytes based on relative 

solubility in a given solvent [Koncsag and Barbulescu 2011]. In general, two 

immiscible solvents are mixed, one solvent containing the whole extract (called the 

“feed”) and one that ideally solubilizes the analyte of interest preferentially. Based on 

solubility, the chemical constituents will either stay in the feed solvent or partition into 

the other solvent. Once the solvents are allowed to phase-separate, the solvent 

containing the enriched analyte (called the raffinate) is removed. 

Liquid-liquid extraction is usually done using an aqueous solvent and an organic 

solvent. Solvent selection is critical and can be difficult. The goal of the extraction is to 

choose a solvent that leaves behind as much of the interfering matrix as possible in the 

feed solvent while also being able to preferentially solvate the analyte of interest. The 

solvent also must be compatible with the analytical instrumentation. Liquid-liquid 

extractions tend to use large amounts of solvent, which can result in the analyte of 

interest being below analytical levels of detection and/or levels of quantitation. Many 

organic solvents can be easily concentrated by evaporation, but this can pose problems 

with labile or volatile analytes.

Solid-phase extraction

Another option that can help avoid these pitfalls is the use of solid-phase extraction 

(SPE) [Sigma-Aldrich 1998]. SPE exploits the analyte affinity (or lack thereof) for a 

solid material packed inside a column. Sample enrichment can occur by the column 

retaining interfering compounds and the analyte of interest passing through, or by the 

column retaining the analyte and the interfering chemicals passing through. 

Concentration of the analyte of interest can be achieved by eluting the analyte in a 

smaller volume of solvent, and filtration can be achieved simultaneously. SPE may 

make it possible to achieve analyte concentration and avoid potential losses that could 

arise from concentrating a large volume of solvent. Appropriate solvent selection is 

also critical to successful SPE.
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g. Immunoassays

The immunoassay is an analytical technique for measuring a targeted antigen, which is

also referred to as an analyte. A critical component of the immunoassay is the antibody or

ligand, which binds a specific antigen or binding site. The binding of the antibody or

ligand forms the basis for the immunoassay, and numerous formats have been devised

which permit visual or instrumental measurements of this reaction. Antibodies include

either monoclonal or polyclonal antibodies and these are commonly employed to detect

organisms by binding to antigens, usually proteins, polysaccharides or other cell wall

components [Hung et al. 2005; Macher 1999]. The analysis is usually performed following

extraction of the analytes to form a heterogeneous matrix. In most immunoassays, there is

little need for extensive sample cleanup. Following the development of

radioimmunoassays, many immunoassays that use monoclonal antibodies are now readily

available from commercial sources, permitting laboratories to use standardized

immunoassays or rapidly develop in-house immunochemical assays. In addition,

commercially available immunoassays or multiplex platforms are available to quantify a

variety of indoor or occupational bioaerosol sources [King et al. 2013]. Some of the more

widely used immunoassay formats are as follows:

Enzyme immunoassays (EIA)

Enzyme Immunoassays (EIA) are composed of a variety of assay formats that can be 

used to quantify bioaerosols in an air or dust sample. The binding of an antibody or 

antigen to an enzyme, such as horseradish peroxidase (HRP) or alkaline phosphatase 

(AP), is the basis of EIA techniques. Enzymatic activity, in the presence of a 

chromogen, results in a colored end-product that is quantified using a 

spectrophotometer. Many, if not most, commercially available EIAs are enzyme-linked 

immunosorbent assays (ELISAs). There are four types of ELISAs which include direct 

ELISA, indirect ELISA, sandwich ELISA and competitive ELISA. 

The sandwich ELISA method is typically used for the detection for airborne viruses 

and aeroallergens [Hung et al. 2005]. In this assay, a capture antibody is bound to a 

solid surface, usually a 96-well plate. The bioaerosol extract is added to the plate 

containing the capture antibody and incubated for a specified length of time, washed 

with a phosphate buffer solution and probed with an enzyme-labeled antibody that 

enables detection and quantification, either through colorimetric changes or 

fluorescence emissions. The advantages to using a sandwich-based ELISA method are 

that it is highly specific and can be used on complex samples such as aerosols. Some 

disadvantages to using a sandwich-based ELISA are poor antibody recognition and/or 

minimal detection due to low sample concentration. Lastly, it should be noted that 

ELISA, like any protein-based assay, does not distinguish between viable and non-
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viable viral aerosols. Multiplexed technologies that enable the detection and 

quantification of multiple allergen sources in one sample are also available from a 

variety of commercial sources [King et al. 2013].

EIA methods to assess fungal bioaerosols based on the detection of fungal cell wall 

components, enzymes, antigens and allergens have become commercially available. β1,3-D glucan, extracellular polysaccharides, mycotoxins and a variety of fungal 

antigens can now be quantified in air and dust samples using immunochemical, 

enzymatic and chemical detection platforms [Chew et al. 2001; Douwes et al. 1997; 

Eduard et al. 2012]. Although many of these biomarkers and methods serve as a proxy 

measure for total fungal biomass, these approaches can be confounded by limitations 

associated with component extraction biases. In addition, complex extraction, 

washing, amplification or immunochemistry steps are required that may add hours or 

even days before a dataset is finalized for analysis and interpretation.

To date, there are a number of commercial companies that have developed ready to 

use EIA kits for the detection and quantification of a variety of indoor and 

occupational biomarkers including dog, cat, dust mite, fungal and rodent allergens 

[Filep et al. 2012]. These EIA approaches enable the collection and quantification of 

these biomarkers in the work environment and provide a ready to use platform for the 

industrial hygienist.

Fluorescent immunoassays (FIA)

Utilization of fluorescent-labeled antibodies to detect bacterial antigens was 

introduced by Coons et al. [1941; 1942]. Various FIA techniques have now evolved 

and are commonly utilized in laboratories. These include: (1) direct FIA, to detect cellbound antigens using a fluorescent antibody; (2) indirect FIA to detect cell-bound 

antigens using a primary antibody and a fluorescent secondary antibody; and (3) 

indirect FIA to detect serum antibodies using an antigen, serum, and a fluorescent 

antibody. Various fluorescent dyes, such as fluorescein, fluorescein isothiocyanate, and 

rhodamine isothiocyanate, may be employed [Thermo Fisher Scientific 2014]. A 

fluorescent or confocal microscope is used to evaluate the samples and to count the 

number of fluorescently stained organisms [Garvey et al. 1977; Popp et al. 1988]. 

Similarly, flow cytometry-based approaches using fluorescent-labelled antibodies have 

also been employed to evaluate a variety of bioaerosol sources including bacteria, 

pollen and fungi [Rittenour et al. 2012; Rule et al. 2007; Rydjord et al. 2007]. 

Multiplexed approaches have been developed for the detection of multiple allergens in 

the same extracted sample [King et al. 2013].

FIA can be used to detect viruses. An example of a direct FIA assay for the detection of 

virus-laden aerosols is the Focus Forming Assay (FFA) [Flint et al. 2009a]. With the 
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FFA, fluorescent microscopy is used to visualize immunostained cells and viral titers 

are quantified as focus forming units per milliliter, or FFU/mL. While the FFA is more 

sensitive than culture-based methods alone, samples with low viral titers may weakly 

fluoresce and possibly be considered undetectable. To overcome weak detection levels, 

an indirect IFA assay may be more appropriate [Leland and Emanuel 1995; Madeley 

and Peiris 2002]. With indirect IFA assays, a primary, unconjugated antibody is used 

in combination with a fluorophore-conjugated, secondary antibody directed against 

the primary antibody. Because the secondary antibody is able to bind to multiple 

epitopes on the primary antibody, it increases fluorescence and enhances overall 

detection. While IFA is a trusted method of viral detection and quantification, it is 

fraught with limitations including excessive cost and the necessity of a skilled 

technician experienced in the reading of immunofluorescence. Likewise, because 

viruses are constantly undergoing antigenic drift and occasionally antigenic shift, 

changes in viral antigens can affect the binding affinity of the primary antibody and 

may result in false negatives.

Ligand-based assays

As an alternative to cell culture and immunofluorescent-based assays, there are several 

protein-based methods of detection that can be used to quantify viral loads in an 

aerosol sample. The hemagglutination (HA) assay is a non-fluorescence quantitative 

assay that is based upon the ability of certain viral pathogens to agglutinate speciesspecific erythrocytes [Condit 2007]. In a serial twofold dilution, viral samples are 

mixed with a 1% solution of erythrocytes and incubated at room temperature for 30-

60 minutes. Viral samples which form an agglutinated lattice are able to prevent red 

blood cells from precipitating out of solution by the binding of the hemagglutinin 

protein (present on the surface of the viral pathogen) to the sialic acid receptors 

(present on the surface of red blood cells). The titer of the sample is based on the well 

with the last agglutinated appearance, immediately before the well in which the red 

blood cells have settled out of solution. Hemagglutination units (HAUs) are typically 

used to quantify the viral concentration. 

One of the major limitations of the HA assay is that it does not distinguish between 

infectious and non-infectious viral particles. Likewise, certain bacteria and fungi 

possess hemolytic activity and when present in a collected bioaerosol, can alter the HA 

assay and result in false positive readings. To circumvent this issue, a variation of the 

HA assay, known as the Hemagglutination-Inhibition test, can be performed [Stewart 

et al. 1967]. The HA inhibition test measures serum antibodies that are directed 

against the viral pathogen. When present in sufficient concentration, the serum 

antibodies are able to prevent agglutination of red blood cells thereby providing an 

alternative means of quantifying viral loads.
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Direct and indirect immunostaining

Direct and indirect immunostaining methods have been previously described for the 

detection of bioaerosol sources. Popp et al. [1988] developed a staining technique to 

enumerate bioaerosol samples directly captured on microscope slides. These 

approaches have enabled the identification of specific bioaerosol sources, especially 

those that do not contain morphological phenotypes that would be used by a trained 

microbiologist to resolve and identify a specific microorganism [Popp et al. 1988]. 

Alternatives to this approach have been developed and utilized in a variety of indoor 

and occupational environments. A press blotting approach that included immobilizing 

proteins from collected bioaerosols captured on an adhesive tape provided insight into 

the bioaerosols that contain allergen in the outdoor environment [Takahashi et al. 

1993; Takahashi and Nilsson 1995]. An alternative method, called the Halogen 

Immunoassay, enables the immunostaining of allergen and antigen around bioaerosols 

captured on a protein binding membrane such as PVDF or mixed cellulose ester 

[Green et al. 2006c; Tovey et al. 2000]. This immunoassay approach has been used in a 

variety of indoor and occupational settings to evaluate allergen sources including, cat, 

dog, latex, rodent, plant, and fungi [Green et al. 2006a; Green et al. 2003; Green et al. 

2005a; Green et al. 2011; Green et al. 2005b; Green et al. 2005c; Green et al. 2006b; 

Green et al. 2006c; Mitakakis et al. 2001; Poulos et al. 2002; Poulos et al. 1999; 

Razmovski et al. 2000; Renstrom 2002; Tovey and Green 2004]. Recently these 

approaches have been adapted to FESEM applications and have been used to detect 

morphologically indiscernible fungal fragments [Afanou et al. 2015].

Biosensors

To overcome some of the technical challenges associated with traditional methods to 

assess bioaerosol exposure, real-time sensor technologies are being developed for the 

detection of bioaerosols [Fronczek and Yoon 2015; Hook-Barnard et al. 2014]. The 

sensor technologies are based on a variety of signal detection strategies that include 

optical, mechanical, electrical, or magnetic sensing approaches. These methods have 

resulted in platforms that have enabled the rapid detection of microbial pathogens and 

in an automated format. Using this technology requires little technological skill and 

can be developed into an automated handheld device. These developments have 

resulted in the fabrication of remote monitoring units in the agricultural sector that 

can process data remotely and apply it to geographical information systems or 

forecasting models.

Biosensors have been developed for a variety of applications including the detection of 

microbial pathogens [Fronczek and Yoon 2015; Hook-Barnard et al. 2014]. Typically, 

a biologically derived analyte (such as fungal spores, hyphae, antigens, peptides or 

nucleotides) is collected and interacts with a selected bioreceptor immobilized on a 
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 sensor surface. Examples of bioreceptors include oligonucleotides, 

monoclonal/polyclonal antibodies, enzymes, cells, and even phages. The bioreceptor 

system consists of a physiochemical transducer that produces a measurable signal. 

Transducers can be optical and include surface plasmon resonance (SPR), which is a 

method that measures changes in the refractive index during molecular binding events 

[Unser et al. 2015; Usachev et al. 2014]. In contrast, electrochemical transducers 

measure changes in current, potential, impedance, and conductance across an 

electrode surface for detection events [Patolsky et al. 2006]. Examples of 

electrochemical detection include the measurement of electrical conductance 

produced by antibody-antigen binding events [Patolsky et al. 2004]. Both optical and 

electrochemical transducers have been developed for the detection of a variety of 

pathogens in the biosecurity, medical and agricultural sectors.

h. Gene-based assays

Cell culture, protein-based, and immunological-based assays are invaluable diagnostic 

tools. However, the evolution of nucleic acid-based molecular diagnostics are rapidly 

becoming the preferred method for detecting and quantifying bioaerosols [West et al. 

2008]. Nucleic acid-based molecular diagnostics can be divided into two categories, (1) 

Direct sample analysis and (2) Indirect sample analysis, such as the Viral Replication 

Assay (VRA), which requires cultivation of the target microorganism prior to molecular 

analysis [Blachere et al. 2011]. Regardless of which approach is taken, there are three steps 

involved: extraction and purification of nucleic acids; amplification of the gene target; and 

detection of the amplicon. For viral, bacterial, plant and fungal nucleic acid extraction and 

purification, a number of kits are commercially available. Such kits generally rely upon 

either silica adsorption (spin-column) or affinity purification (magnetic separation) 

methodologies. Because bioaerosols are typically dilute in nature, investigators should 

determine which method yields the greatest amount of nucleic acids while minimizing 

sample handling, contamination and degradation. These are important variables to 

consider and can vary depending on the selected approach.

With the advent of molecular assays such as the Polymerase Chain Reaction (PCR) and 

Real-Time Quantitative PCR (RT-qPCR), which detect specific genetic sequences in the 

sample DNA or RNA, it has been possible to provide standardized assays, reduce 

turnaround time, and enhance assay sensitivity and detection specificity [Cella et al. 2013; 

Life Technologies 2014; Mahony 2008]. Using gene-specific oligonucleotides coupled with 

either an intercalating fluorescent dye (e.g. SYBR green) or a fluorogenically labeled gene 

probe (e.g. VIC, 6FAM), industrial hygienists are able to monitor indoor and outdoor 

bioaerosols for the presence of microorganisms. Likewise, multiplexing PCR and beadbased multiplexing PCR, which couples PCR and flow-cytometry, can be used for high-
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 throughput screening of multiple respiratory viruses in a single reaction mixture. It should 

be noted that poor assay design, primer and probe base-pair mismatches and degraded 

template nucleic acids can lead to false negatives or reduced detection sensitivity. 

Investigators should ensure optimal assay design, validate limits of detection, and run 

valid PCR controls in parallel.

Within the last two decades, a variety of molecular technologies have been used to 

quantify eukaryotic biomass including fungi and plant pollen in occupational, health, 

residential, and industrial samples [Rittenour et al. 2012; Scott et al. 2011; Summerbell et 

al. 2011]. Examples of these technologies include molecular based methods to evaluate 

specific or conserved gene loci (internal transcribed spacer region of ribosomal RNA) such 

as Sanger [Rittenour et al. 2014] or next generation sequencing [Kettleson et al. 2015], 

denaturing gradient gel electrophoresis [Johansson et al. 2014] and quantitative PCR 

[Eduard et al. 2012; Vesper et al. 2007]. The latter approach includes examples such as a 

DNA-based mold specific quantitative PCR (msQPCR) method that enables the detection 

and quantification of 36 indicator fungal species [Vesper et al. 2007]. This msQPCR 

method has been used to develop an Environmental Relative Moldiness Index (ERMI) to 

quantify the mold burden in homes. Originally developed by the U.S. Environmental 

Protection Agency (EPA) [Vesper et al. 2007], this approach has been licensed to a variety 

of companies in the commercial sector and is widely used to evaluate indoor fungal 

bioaerosol particles in settled dust during investigations of indoor air quality [BolañosRosero et al. 2013; Kettleson et al. 2015; Reponen et al. 2012; Reponen et al. 2011a; Taubel 

et al. 2016; Vesper et al. 2013; Vesper et al. 2007]. The development of this methodology 

has provided the first step towards a standardized approach to quantify fungal bioaerosol 

sources within the indoor environment. Other metagenomic molecular methods including 

Sanger, 454, and Illumina miSeq sequencing platforms have also provided new insights 

into the complete diversity of bacterial and fungal bioaerosols in indoor, outdoor and 

occupational environments.

10 Limitations of bioaerosol sampling and 

characterization

Bioaerosol sampling can be a useful tool to study occupational exposures, potential health 

hazards, and the transmission of infectious diseases. However, bioaerosol sampling has 

significant limitations, and these need to be kept in mind when deciding whether or not to 

collect bioaerosol samples, preparing a sampling plan, and interpreting the results.

The first and most important limitation is the lack of standards and guidelines for acceptable 

bioaerosol exposure limits. NIOSH and other organizations have set recommended exposure 
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 limits for several organic materials which may contain microorganisms and their fragments, 

such as cotton dust, grain dust, starch and wood dust [NIOSH 2010]. Although numerous 

studies have suggested a connection between exposure to various bioaerosols and respiratory 

illnesses, exposure limits do not currently exist in the US for airborne pollen, fungi, protozoa, 

bacteria, viruses, or their fragments. These limits have not been established largely because it 

is not possible to definitively state that a particular bioaerosol concentration will or will not 

lead to adverse health outcomes [Eduard et al. 2012; Heederik 2013; Morey 2007; Nevalainen 

et al. 2015; NIOSH 2012a]. This is true for several reasons: bioaerosols are often a complex 

mixture of microorganisms and organic materials; thousands of species of microorganisms 

exist, and most have not been studied; microorganisms and their fragments can cause illnesses 

in a variety of ways, including allergic reactions, infections and toxicity; the health effects of 

biological materials can vary substantially from person to person; and sampling and analytical 

procedures are not standardized, which makes it difficult to compare results [Eduard et al. 

2012; Heederik 2013; Morey 2007; Nevalainen et al. 2015; Taubel et al. 2016]. Although 

research is ongoing, no standards for acceptable levels of bioaerosols in the environment have 

been established by the US government or organizations such as the ACGIH or the AIHA.

In addition to the lack of occupational exposure limits for bioaerosols, measuring and 

interpreting bioaerosol concentrations are more complex than is often appreciated. Bioaerosol 

concentrations can vary significantly from location to location within a building, especially if 

the bioaerosol has one or a few localized sources. A study of bioaerosol exposure in a large 

engine plant found that levels of airborne fungi, bacteria and endotoxin varied from location 

to location within the plant [Thorne et al. 1996]. A study of airborne fungi in two residences 

found significant differences between two rooms sampled at the same time [Hyvarinen et al. 

2001]. In healthcare settings, patients with certain respiratory infections expel bioaerosol 

particles containing infectious pathogens. Because of the dispersion of the aerosol and the 

settling of larger droplets, the bioaerosol concentration decreases rapidly as the distance from 

the patient increases [Jones and Brosseau 2015]. A study of airborne influenza in a healthcare 

clinic found that the concentrations were much higher in examination rooms containing 

patients with influenza than other locations, and that the airborne influenza concentration 

also varied from location to location within the waiting room [Lindsley et al. 2010a].

Most bioaerosol collection methods provide a snapshot of the environmental bioaerosols at a 

specific time. Temporal variations in bioaerosol concentrations are commonly observed, 

especially if the bioaerosol generation occurs during episodic events rather than continuously. 

One study of indoor airborne mold in a residence found that day-to-day concentrations of 

airborne fungi varied considerably, and that levels were 26 times higher in the summer than in 

the winter [LeBouf et al. 2008]. Another residential study found that more airborne fungi were 

present in the morning than the afternoon and earlier in the winter compared to later 

[Hyvarinen et al. 2001]. In the influenza study mentioned above, day-to-day levels of airborne 
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Sampling and Characterization of Bioaerosols 

 influenza virus varied considerably depending upon the number of influenza patients present 

[Lindsley et al. 2010a].

Other factors also influence bioaerosol concentrations. Building airflow patterns and the 

operation of the HVAC system can affect bioaerosol levels, particularly if the HVAC system is 

a source of bioaerosol particles [Macher 1999]. Areas occupied by people show increased 

levels of bioaerosols compared to empty spaces, both because people themselves shed 

bioaerosol particles and because human activities such as walking and sitting can re-suspend 

dust from floors and furniture [Buttner and Stetzenbach 1993; Ferro et al. 2004; Hung et al. 

2005; Qian et al. 2012]. Outdoor air is an important source of airborne fungi in many indoor 

environments due to fresh air being drawn in by HVAC systems and infiltration through 

cracks and openings [Eduard 2009].

Because of these issues, if bioaerosol sampling is to be conducted, it needs to be a part of a 

well-planned and comprehensive sampling strategy. The development of a sampling plan 

should begin with a thorough inspection and understanding of the workplace, including the 

building, HVAC system, and possible sources of bioaerosols. The sampling plan should 

integrate other types of data collection with the bioaerosol sampling, such as bulk sampling of 

possible source materials, surface sampling of settled dust, and health surveys of workers. 

Collections will need to be carried out at multiple locations and multiple time points, and 

even then it must be kept in mind that such samples may not fully characterize the exposure 

and that false negative results are quite possible [ASTM 2014a; Hung et al. 2005; Macher 1999; 

Morey 2007].

Bioaerosol sampling can be beneficial when done in the appropriate context [Hung et al. 2005; 

Macher 1999; Morey 2007]. It can be helpful to compare indoor and outdoor levels of 

bioaerosols to identify possible indoor problem microorganisms. Sampling for specific 

microorganisms of concern can be useful, especially if there is a known source, such as a 

composting operation or an aeration tank in a sewage treatment facility [Environment Agency 

2009; Masclaux et al. 2014]. Bioaerosol sampling is also a valuable research tool for better 

understanding sources and exposures. On the other hand, sampling for bioaerosols will not be 

helpful if there is not some basis for interpreting the resulting data. For example, because of 

the lack of dose-response information and the variability associated with bioaerosol sampling, 

NIOSH does not recommend routine air sampling when investigating possible respiratory 

illness due to exposures in damp buildings. Instead, NIOSH recommends inspections of the 

building and its HVAC systems to locate moisture and microbial growth problems, followed 

by remediation [NIOSH 2012a].
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Sampling and Characterization of Bioaerosols 

 

11 Safety considerations

Investigators should use appropriate personal protective equipment (PPE) and practice good 

personal hygiene when conducting indoor environmental quality, disease outbreaks, and 

agricultural health investigations that have resulted in medically diagnosed symptoms. PPE 

may include respiratory protection to prevent inhalation of microbes and microorganismresistant clothing to prevent transmission to investigators by bodily contact with 

microorganisms. Good personal hygiene practices include washing exposed skin and clothing 

thoroughly and refraining from eating, drinking, or smoking in a contaminated area. These 

simple steps will help minimize the ingestion, inhalation, or uptake of microorganisms.

All samplers, culture plates, and other equipment should be handled aseptically to prevent 

contamination of the samples and, more importantly, to prevent the spread of potential 

human pathogens to the worker or the work environment. All surfaces, including washed 

hands, may harbor microorganisms or spores unless they are specifically sterilized. Practically 

speaking, however, not all objects can be sterilized. While disinfection with an oxidizing 

chemical or alcohol destroys most vegetative cells, these agents do not destroy all spores. 

Samplers should be disinfected or, if possible, sterilized after each sample collection. Special 

care should be given to samplers with convoluted inlets or air pathways where 

microorganisms may accumulate.

Information on the safe handling of biological specimens can be found in in the free online 

manual “Biosafety in Microbiological and Biomedical Laboratories” from the Centers for 

Disease Control and Prevention at 

(http://www.cdc.gov/biosafety/publications/bmbl5/index.htm) [CDC 2009]. Information on 

the handling of some specific pathogens can also be found at the CDC website (www.cdc.gov).

12 Resources

The NIOSH Manual of Analytical Methods has several chapters discussing other aspects of 

aerosol sampling, including general considerations and factors affecting aerosol sampling, an 

explanation of filter pore size, sampling airborne fibers, sampler wall losses, and avoiding 

bypass leakage in filter cassettes [NIOSH 2003e]. NIOSH also maintains a web page on indoor 

environmental quality with more information and links to additional resources at 

http://www.cdc.gov/niosh/topics/indoorenv/. The American Industrial Hygiene Association 

(AIHA; https://www.aiha.org) has reference materials and resources on indoor air quality and 

bioaerosols, as does the American Conference of Governmental Industrial Hygienists 

(ACGIH; http://www.acgih.org). ASTM International (http://www.astm.org) publishes 

numerous standards and guides on the evaluation of indoor air quality, including developing 
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Sampling and Characterization of Bioaerosols 

 an air sampling strategy and the collection and evaluation of bioaerosols [ASTM 2009; ASTM 

2014a; ASTM 2014b; ASTM 2014d; ASTM 2014e]. The Centers for Disease Control and 

Prevention has guidelines on environmental infection control in healthcare facilities that 

include recommendations on environmental sampling [CDC 2003].

Disclaimer

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

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

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

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

were accessible as of the publication date.

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Vincent JH [2005]. Health-related aerosol measurement: a review of existing sampling criteria 

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Vincent JH [2007]. Aerosol sampling: science, standards, instrumentation and applications. 

Hoboken, NJ: John Wiley & Sons.

Wagner J, Macher JM [2003]. Comparison of a passive aerosol sampler to size-selective pump 

samplers in indoor environments. Am Ind Hyg Assoc J 64(5):630-639.

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[2016]. Generation and sampling of nanoscale infectious viral aerosols. Aerosol Sci Technol 

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modes of transmission: a critical review. J Infect 57(5):361-373.

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Trends Microbiol 16(8):380-387.

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Sampling and Characterization of Bioaerosols 

 Yooseph S, Andrews-Pfannkoch C, Tenney A, McQuaid J, Williamson S, Thiagarajan M, 

Brami D, Zeigler-Allen L, Hoffman J, Goll JB, Fadrosh D, Glass J, Adams MD, Friedman R, 

Venter JC [2013]. A metagenomic framework for the study of airborne microbial 

communities. PLoS ONE 8(12):e81862.

Zhao Y, Aarnink AJ, Wang W, Fabri T, Groot Koerkamp PW, de Jong MC [2014]. Airborne 

virus sampling: efficiencies of samplers and their detection limits for infectious bursal disease 

virus (IBDV). Ann Agric Environ Med 21(3):464-471.

Zimmerman NJ, Reist PC, Turner AG [1987]. Comparison of two biological aerosol sampling 

methods. Appl Environ Microbiol 53(1):99-104.

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[2013]. Association of airborne virus infectivity and survivability with its carrier particle size. 

Aerosol Sci Technol 47(4):373-382.
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Sampling and Characterization of Bioaerosols 

 

14 Appendix 1- List of manufacturers/distributors of 

common bioaerosol samplers and related 

products

This list is not inclusive, and the inclusion of a specific product or company does not 

constitute endorsement by the National Institute for Occupational Safety and Health 

(NIOSH). If your bioaerosol equipment manufacturing or supply company is not listed or the 

information is incorrect or out-of-date, please contact us and we will review your information 

for inclusion as the list is updated.

Manufacturers/Distributors of Common Bioaerosol Samplers and Related Products

A.P. Buck Inc.

7101 Presidents Drive, Suite 110

Orlando, FL 32809 USA

Phone: (800) 330-BUCK [2825]

Phone: (407) 851-8602

Fax: (407) 851-8910

http://www.apbuck.com

Ace Glass Incorporated

1430 North West Boulevard

P.O. Box 688

Vineland, NJ 08362 USA

Phone: (800) 223-4524

Phone: (856) 692-3333

Fax: (800) 543-6752

Fax: (856) 692-8919

http://www.aceglass.com

Aerosol Devices Inc.

2614 S. Timberline Road, #109-125

Fort Collins, CO 80525 USA

Phone: (970) 744-3244

http://aerosoldevices.com

Aquaria srl

Via della Levata, 14

Lacchiarella (Milan) 20084 Italy

Phone: +39 02-90091399

Fax: +39 02-9054861

http://www.aquariasrl.com

Barramundi Corporation

6449 South Tex Point

PO Drawer 4259

Homosassa, FL 34448 USA

Phone: (800) 382-1817

Phone: (352) 628-0200

Fax: (352) 628-0203

http://barramundicorp.com

BD Biosciences 

2350 Qume Drive 

San Jose, CA 95131 

Phone: 877-232-8995

http://www.bdbiosciences.com

Beijing SENNON Technology

Development Company, Ltd.

North Building No. 2

Dongdajie xili, Fengtai District

Beijing 100071 P.R. China

Phone: +86 10-6381 8024

Fax: +86 10-6380 6170

http://www.sennon.net/eng

Bertin Corporation

9700 Great Seneca Highway

Suite # 662

Rockville, MD 20850 USA

Phone: (240) 428-1047

http://www.coriolis-airsampler.com

Bi-Air Corporation

1349 Montevideo Ave

Placentia, CA 92870 USA

Phone: (714) 985-9659

Fax: (714) 528-5429

http://expertonmold.com

bioMérieux, Inc.

595 Anglum Road

Hazelwood, MO 63042 USA

Phone: (800) 634-7656

Fax: (800) 657-3053

http://www.biomerieux-usa.com

Bioscience International

11333 Woodglen Drive

Rockville, MD 20852 USA

Phone: (301) 231-7400

Fax: (301) 231-7277

http://www.biosci-intl.com

Burkard Manufacturing Company 

Ltd.

Woodcock Hill Industrial Estate

Rickmansworth, Hertfordshire WD3 

1PJ England

Phone: +44 (0) 1923 773134

Fax: +44 (0) 1923 774790

http://www.burkard.co.uk

Climet Instruments Company

1320 W. Colton Avenue

Redlands, CA 92374 USA

Phone: (909) 793-2788

http://www.climet.com

Droplet Measurement Technologies

2545 Central Avenue

Boulder, CO 80301 USA

Phone: (303) 440-5576Fax: (303) 440-

1965

http://www.dropletmeasurement.com
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Sampling and Characterization of Bioaerosols 

 Manufacturers/Distributors of Common Bioaerosol Samplers and Related Products -

Continued

Dycor Technologies, Ltd.

1851 94th Street

Edmonton, AB T6N 1E6 Canada

Phone: (800) 663-9267

Phone: (780) 486-0091

Fax: (780) 486-3535

http://www.dycor.com

EMD Millipore

290 Concord Road

Billerica, MA 01821 USA

Phone: (800) MILLIPORE [645-5476]

Fax: (781) 533-6000

http://www.emdmillipore.com

EMSL Analytical, Inc

200 Route 130 North

Cinnaminson, NJ 08077

Phone: (800) 220-3675

http://www.emsl.com Environics Oy

Environmental Monitoring Systems, Inc.

3864 Leeds Avenue

Charleston, SC 29405 USA

Phone: (800) 293-3003

Phone: (843) 724-5708

Fax: (866) 724-5702

Fax: (843) 724-5702

http://www.emssales.net

Evogen, Inc.

10513 W. 84th Terrace

Lenexa, KS 66214 USA

Phone: (888) 450-4321

Phone: (913) 948-5640

Fax: (913) 948-5664

http://evogen.com

F.W. Parrett Limited

7 Coppergate Close

Bromley, Kent BR1 3JG England

Phone: +44 020-8460-2116

Fax: +44 020-7504-3536

http://www.parrett.uk.com

FLIR Systems, Inc.

70 Castilian Drive

Goleta, CA 93117 USA

Phone: (888) 747-FLIR [3547]

http://www.flir.com

GE Healthcare Life Sciences 

(Whatman)

800 Centennial Avenue

P.O. Box 1327

Piscataway, NJ 08855 USA

Phone: (800) 526-3593

Fax: (877) 295-8102

http://www.gelifesciences.com

Indoor Biotechnologies Inc

700 Harris Street

Charlottesville, VA 22903 USA

https://inbio.com

InnovaPrep

132 East Main Street

Drexel, MO 65742 USA

Phone: (816) 619-3375

http://innovaprep.com

InnovaTek, Inc.

3100 G. Washington Way

Suite 108

Richland, WA 99354 USA

Phone: (509) 375-1093

Fax: (509) 375-5183

http://www.innovatek.com

Inspirotec

2319 West Wabansia Avenue #1

Chicago, IL 60647

Phone: 847 302 1839

Fax: 847 234 2089

http://www. inspirotec.com 

MSP Corporation

5910 Rice Creek Parkway

Suite 300

Shoreview, MN 55126 USA

Phone: (651) 287-8100

Fax: (651) 287-8140

http://www.mspcorp.com

Pall Corporation

25 Harbor Park Drive

Port Washington, NY 11050 USA

Phone: (800) 521-1520

Phone: (516) 484-3600

Fax: (516) 801-9754

http://www.pall.com

Particle Measuring Systems

5475 Airport Boulevard

Boulder, CO 80301 USA

Phone: (800) 238-1801

Phone: (303) 443-7100

Fax: (303) 449-6870

http://pmeasuring.com

Research International, Inc.

17161 Beaton Road SE

Monroe, WA 98272 USA

Phone: (800) 927-7831

Phone: (360) 805-4930

Fax: (360) 863-0439

http://www.resrchintl.com

RJ Lee Group, Inc.

350 Hochberg Road

Monroeville, PA 15146

Manufacturers/Distributors of 

Common Bioaerosol Samplers and 

Related Products

724) 325-1776

Fax: (724) 733-1799

http://www.rjlg.com 

Sammonkatu 12

P.O. Box 349

FI-50101 Mikkeli Finland

Phone: +358 201 430 430

Fax: +358 201430 440

http://www.environics.fi/

Sartorius Stedim Biotech GmbH

August-Spindler-Strasse 11

Goettingen 37079 Germany

Phone: (800) 368-7178

Phone: +49 551-308-0

Fax: +49 551-308-3289

https://www.sartorius.com

SKC, Inc.

863 Valley View Road

Eighty Four, PA 15330 USA

Phone: (800) 752-8472

Phone: (724) 941-9701

Fax: (724) 941-1369

http://www.skcinc.com
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Sampling and Characterization of Bioaerosols 

 Manufacturers/Distributors of Common Bioaerosol Samplers and Related Products -

Continued

Tecora 

211-215 Rue la Fontaine 

94134 Fontenay sous Bois Cedex, France 

Phone: Tel: +33 1 48 75 82 82 

Fax: +33 1 48 75 82 96 

http://www.tecora.com/en/

Thermo Scientific

27 Forge Parkway

Franklin, MA 02038 USA 

Phone: (866) 282-0430 

Phone: (508) 520-0430 

Fax: (508) 520-1460 

http://www.thermoscientific.com

TSI Incorporated

500 Cardigan Road

Shoreview, MN 55126 USA 

Phone: (800) 874-2811 

Phone: (651) 483-0900 

Fax: (651) 490-3824 

http://www.tsi.com

Veltek Associates, Inc.

15 Lee Boulevard

Malvern, PA 19355 USA 

Phone: (888) 4-STERILE [478-3745] 

Phone: (610) 644-8335 

http://sterile.com

Zefon International, Inc. 

5350 SW 1st Lane 

Ocala, FL 34474 USA 

Phone: (800) 282-0073 

Phone: (352) 854-8080 

Fax: (352) 854-7480 

http://www.zefon.com
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-104 of BA-115

Sampling and Characterization of Bioaerosols 

15 Appendix 2 – Commonly used bioaerosol samplers

This list is not inclusive, and the inclusion of a specific product or company does not constitute endorsement by the National Institute for 

Occupational Safety and Health (NIOSH). If a bioaerosol sampler is not listed or the information is incorrect or out-of-date, please contact 

us and we will review your information for inclusion as the list is updated.

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques

Table I. Common Commercially-Available Filter Samplers for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

d50

(μm) Application¥

Bi-Air Bi-Air Filter Cassette Filter 1 – 5 Varies C,M,O

Burkard High Throughput 'Jet' Spore and Particle Sampler Filter 850 Varies C,M,O

Dycor CSU-1 Low Volume Air Sampler Filter 10 Varies C,M,O

EMD Millipore Various filter and membrane media Filter Varies Varies C,M,O

GE (Whatman) Various filter and membrane media Filter Varies Varies C,M,O

InnovaPrep ACD-200 Bobcat Air Sampler Filter 100 or 200 C,M,O

Pall Corp. Various filter and membrane media Filter Varies Varies C,M,O

Research 

International SASS® 3100 Dry Air Sampler Filter 50 – 310 0.3 – 0.5 C,M,O

Sartorius AirPort MD8 Air Sampler Filter 30, 40, or 50 - C

SKC Button Aerosol Sampler Filter 4 - M,O

Zefon Various filter and membrane media Various Varies Varies C,M,O
Table II A. Common Commercially-Available Single-stage Impactor Samplers for Bioaerosol Collection

Manufacturer/ Collection Flowrate d50 Sampler Name Application¥

Distributor Media (L/min) (μm)

BioAireTM B6 Single Stage Microbial Sampler Agar 28.3 - C

A.P. Buck Bio-CultureTM Microbial Air Sampler Agar 30 – 120 - C

BioSlideTM Microbial Air Sampler Slide 10 – 20 - M

MICROFLOW 60 Microbiological Air Sampler Agar 30 – 120 - C

Aquaria MICROFLOW 60-90/C Microbiological Air Sampler Agar 30 – 120 - C

MICROFLOW 90/C Microbiological Air Sampler Agar 30 – 120 - C

Barramundi Mattson-Garvin Model 220 Air Sampler (240V is Model 270) Agar 28.3 - C

Handy Microbial Air Sampler® Agar/Filter - - C,M,O

Beijing JWL-IIA Mini® Microbial Air Sampler Agar 20 - C

SENNON JWL-IIB202 Universal® Microbial Air Sampler Agar 20 - C

JWL-IIC Professional® Microbial Air Sampler Agar 20 - C

airIDEAL® 3PTM Traceability Air Sampler Agar 100 - C bioMérieux Samp'airTM Microbial Air Sampler Agar 100 - C

SAS Duo 360 High Volume Microbial Air Sampler Agar 360 - C

Bioscience SAS Isolator Microbial Air Sampler Agar 180 - C

International SAS Super 100 Microbial Air Sampler Agar 100 - C

SAS Super 180 Microbial Air Sampler Agar 180 - C

24-Hour Recording Volumetric Spore Trap Slide 10 - M

Continuous Recording Air Sampler Slide 10 - M

Personal Volumetric Air Sampler Slide 10 - M Burkard Portable Air Sampler for Agar Plates Agar 10 or 20 - C

Recording Air Sampler Side 10 - M

Seven-Day Recording Volumetric Spore Trap Slide 10 - M

CI-90 & CI-90+ Airborne Microbial Sampler Agar 100 - C

Climet CI-95 & CI-95+ Airborne Microbial Sampler Agar 100 - C

CI-99 Microbial Air Sampler Agar 100 - C

Dycor Dycor Slit Sampler Agar 15 – 50 - C

MAS-100 Iso MH® Air Sampler Agar 100 - C

MAS-100 Iso NT® Air Sampler Agar 100 - C

MAS-100 NT® and MAS-100 NT Ex® Air Sampler Agar 100 - C EMD Millipore MAS-100 VF® Active Air Sampler Agar 100 - C

RCS® Isolator Microbial Air Sampler Agar 100 - C

RCS® Plus Ex Explosion-Proof Microbial Air Sampler Agar 100 - C

NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-105 of BA-115

Sampling and Characterization of Bioaerosols 

 
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-106 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II A. Common Commercially-Available Single-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection 

Media

Flowrate

(L/min)

d50

(μm) Application¥

EMD Millipore RCS® High Flow Touch Portable Air Sampler Agar 100 - C

Environmental 

Monitoring 

Systems

Allergenco MK-III Slit-Impaction Sampler Slide 15 - M

Allergenco-D Disposable IAQ Air Monitoring Cassettes Slide 15 - M

Allergenco-D Posi-Track Full Slide, IAQ Impactor Cassette Slide 15 1.7 M

BioSIS Slit Impaction Air Sampler Slide 5 – 50 - M

cyclex-d Cassettes Slide 20 < 1.0 M

E6 Single-Stage Bioaerosol Impaction Sampler Agar 28.3 - C

Micro5 MicroCell Cassettes Slide 5 < 1.0 M

Micro5 Posi-Track Full Slide, IAQ Impactor Cassette Slide 5 <1.0 M

F.W. Parrett MicroBio MB1 Air Sampler Agar 100 - C

MicroBio MB2 Air Sampler Agar 100 - C

Particle 

Measuring 

Systems

Air Trace® Environmental Slit-to-Agar Sampler Agar 28.3 - C

BioCaptTM Impactor Active Microbial Air Sampling Atrium Agar 25 - C

MiniCaptTM Portable Microbial Air Sampler Agar 50 or 100 - C

Sartorius AirPort MD8 Air Sampler Agar 125 - C

SKC

BioStage® Standard Single-Stage Viable Cascade Impactor Agar 28.3 - C

BioStage® 200 Single-Stage Viable Cascade Impactor Agar 14.15 - C

VersaTrap® Spore Trap Cassette Slide 15 - M

Thermo Scientific

IUL Basic Air Air Sampler Agar 60 – 100 - C

IUL Spin Air Air Sampler Agar 60 – 100 - C

IUL Spin Air Basic Air Sampler Agar 60 – 100 - C

N6 Single-Stage Viable Andersen Impactor Agar 28.3 0.65 C

Oxoid Air Sampler Agar 100 - C

Veltek SMA MicroPortable® Air Sampler Agar 1 or 5 - C

Zefon 

International

A-6 Bioaerosol Impactor Agar 28.3 0.65 C

Air-O-Cell® Sampling Cassette Slide 15 2.6 M

Via-Cell® Bioaerosol Sampling Cassette Slide 15 1.56 C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - 

C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-107 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

0.18 

0.32 

0.56 

Model 100-NR (non-rotating)/100-R 

Impactor

(rotating) MOUDITM Filter 30 8 1.0

1.8

3.2

M,O

5.6

10

0.056

0.10

0.18

0.32

MSP 

Corporation

Model 110-NR (non-rotating)/100-R 

Impactor

(rotating) MOUDITM Filter 30 10 0.56

1.0

1.8

M,O

3.2

5.6

10

1.0

Model 100-S4 MOUDITM Impactor Filter 30 3 2.5

10

M,O

0.010

Model 115 Nano-MOUDITM Impactor Filter 10 3 0.018

0.032

M,O

0.010

Model 116 Nano-MOUDITM Impactor Filter 30 3 0.018

0.032

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-108 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

MSP 

Corporation

continued

Model 120 R (rotating) Moudi-IITM Impactor Filter 30 10

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 122-NR/122-R Moudi-IITM and NanoMoudi-IITM Impactor Filter 30 13

0.010

0.018

0.032

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-109 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

0.010

0.018

0.032

0.056

0.10

0.18

Model 125-NR/125-R Moudi-IITM and NanoMoudi-IITM Impactor Filter 10 13 0.32

0.56

M,O

1.0

1.8

3.2

MSP 

Corporation

continued

5.6

10

Model 135-6 MiniMOUDITM (Marple Personal II) Impactor Filter 2 6 

0.56

1.0

1.8

3.2 M,O

5.6

10

0.18

0.32

0.56

Model 135-8 MiniMOUDITM (Marple Personal II) Impactor Filter 2 8 1.0

1.8 M,O

3.2

5.6

10
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-110 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

MSP 

Corporation 

continued

Model 135-10 MiniMOUDITM Impactor Filter 2 10

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Model 135-13 MiniMOUDITM Impactor Filter 2 13

0.010

0.018

0.032

0.056

0.10

0.18

0.32

0.56

1.0

1.8

3.2

5.6

10

M,O

Thermo 

Scientific Eight Stage Non-Viable Cascade Impactor Filter 28.3 8 

0.43

0.65

1.1

2.1

3.3

4.7

5.8

9.0

M,O
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-111 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table II B. Common Commercially-Available Multi-stage Impactor Samplers for Bioaerosol Collection - Continued

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

3.5

Marple 294 Personal Cascade Impactor Filter 2 4 9.8

14.8 M,O

21.3

0.52

0.93

Marple 296 Personal Cascade Impactor Filter 2 6 1.55

3.5 M,O

6.0

9.8

0.52

Thermo 

Scientific 

continued Marple 298 Personal Cascade Impactor Filter 2 8 

0.93

1.55

3.5

6.0 M,O

9.8

14.8

21.3

0.65

1.1

Six Stage Viable Andersen Cascade Impactor Agar 28.3 6 2.1

3.3 C 

4.7

7.0

Two-Stage Viable Andersen Cascade Impactor Agar 28.3 2 0.8

8 C 

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - 

C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-112 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table III. Common Commercially-Available Cyclones and Impinger Samplers for Bioaerosol Collection

Cyclones

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

Burkard Cyclone Sampler for Airborne Particles Dry Vial 20 1 - C,M,O

Cyclone Sampler for Field Operation Dry Vial 16.5 1 - C,M,O

Evogen Sceptor DryCloneTM Dry Vial 400 1 - C,M,O

FLIR Systems C100 Modular Tactical Collector Dry Vial 150 1 - C,M,O

Impingers

Ace Glass

AGI-30 Impinger Liquid 12.5 1 - C,M,O

Greenburg-Smith Impinger Liquid 28.3 1 - C,M,O

Midget Impinger Liquid 1 - C,M,O

4

Burkard Multistage Liquid Impinger Liquid 20 3 10

> 10

C,M,O

Dycor

XMX/102 High Volume Bioaerosol Sampling System Serum Tube 530 1 - C,O

XMX/2L-MIL Bioaerosol Sampler – Military Liquid 530 1 - C,M,O

XMX-CV Microbial Air Sampler – Civilian Liquid 530 1 - C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - 

C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-113 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table IV. Common Commercially-Available Wetted-Surface and Condensation-Based Samplers for Bioaerosol Collection

Wetted-Surface Bioaerosol Samplers

Manufacturer/

Distributor Sampler Name Collection Media

Flowrate

(L/min)

# of 

Stages

d50

(μm) Application¥

Bertin Coriolis® μ Microbial Air Sampler Liquid 100 – 300 1 < 0.5 C,M,O

Coriolis® RECON Portable Air Sampler Liquid 600 1 ≈ 0.5 C,M,O

Bioscience 

International SAS Cyclone Air Sampler Liquid 1200 1 - C,M,O

Evogen Sceptor SpinConTM Advanced Air Sampler Liquid 450 1 - C,M,O

FLIR Systems Fido® B1 (BioCapture® 650) Portable Air Sampler Liquid 

Cartridge 200 1 - C,M,O

InnovaPrep SpinCon® II Advanced Air Sampler Liquid 450 1 - C,M,O

InnovaTek BioGuardian® Air Sampler Liquid 100, 350, 

or 1000 1 - C,M,O

Research 

International

BioHawk® 8-Channel Collector/Bioidentifier Liquid 325 1 - O

SASS® 2300 Wetted-Wall Air Sampler Liquid 325 1 - C,M,O

SASS® 2400 Low-Volume Wetted-Wall Air Sampler Liquid 40 1 - C,M,O

SKC BioSampler® Liquid 12.5 1 - C,M,O

Tecora CIP10-M personal bioaerosol sampler Liquid 10 1 2.1 C,M,O

Condensation-Based Bioaerosol Samplers

Aerosol Devices LSS100 Series Liquid Spot Sampler Liquid 1.0 – 1.5 1 n/a C,M,O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol 

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-114 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table V. Common Commercially-Available Electrostatic, Passive Aerosol and Settled Dust Collection Samplers for Bioaerosol Collection

Electrostatic Samplers

Manufacturer/

Distributor

Sampler Name Collection 

Media

Flowrate

(L/min)

# of 

Stages

d50

(μm)

Application¥

Inspirotec Inspirotec sampler Cartridge 130 1 n/a O

Passive electrostatic dust collectors are listed under Passive Aerosol Samplers. Electrostatic cloths used for 

Settled Dust Collection Devices.

wipe sampling are listed under 

Passive Aerosol Samplers

BD Biosciences Ready-to-use settle plates Agar n/a n/a n/a C

EMD Millipore Ready-to-use settle plates Agar n/a n/a n/a C

Thermo-Scientific Ready-to-use settle plates Agar n/a n/a n/a C

Department of Occupational and 

Environmental Health, University of Iowa

Electrostatic Dust Collector (EDC) Electrostaticallycharged clothsn/a n/a n/a O

RJ Lee Group UNC Passive Aerosol Sampler Various 

substrates

n/a n/a n/a M, O

Settled Dust Collection Devices

Indoor Biotechnologies DUSTREAM® Collector (DU-ST-1) 40 μm nylon 

mesh filter

n/a n/a n/a O

d50 – aerodynamic diameter at which the collection efficiency is 50% and is defined as the cut off diameter

Application¥ - C = culture-based analysis for viability

M = microscopic examination of collected bioaerosol

O = other laboratory analyses, such as immunoassays, bioassays, chemical assays, or molecular detection techniques
NIOSH Manual of Analytical Methods  5th Edition  Chapter BA March 2017 Page BA-115 of BA-115

Sampling and Characterization of Bioaerosols 

 

Table VI. Common Commercially-Available Real-Time Bioaerosol Monitors for Bioaerosol Collection

Manufacturer/

Distributor Sampler Name Detection Method Flowrate(L/min) Application¥

Bioscience 

International SAS-PCR Pathogenic Microorganisms Air Sampler PCR - R 

Droplet 

Measurement 

Technologies

Wideband Integrated Bioaerosol Sensor (WIBS) Fluorescence 0.3 R 

Dycor C-FLAPS Biological Detection System Fluorescence 350 R

Environics ENVI BioScoutTM Fluorescence 2 R

FLIR Systems Fido® B2 Instantaneous Biological Aerosol Detector Fluorescence 3.8 R

Particle 

Measuring

Systems

BioLazTM Real-Time Microbial Monitor Fluorescence 3.6 R 

Research 

International

BioHawk® 8-Channel Collector/Bioidentifier Fluorometric Bioassay 325 R

TacBioTM Biological Aerosol Detector Fluorescence 1 R

TSI BIOTRAK® Real-Time Viable Particle Counter 9510-BD Fluorescence 28.3 R

Fluorescence Aerosol Particle Sensor (FLAPS) 3317 (FLAPS III)TM Fluorescence 1 R

Application¥ - 

R= Real-time or near-real time bioaerosol detection

PCR= Polymerase chain reaction