NMAM Chapter BA

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 D

evices 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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Sampling and Characterization of Bioaerosols

Figure 1: Photomicrographs of acellular, prokaryotic and eukaryotic microorganisms that can

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

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

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

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

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

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

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

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

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

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

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

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

public free of charge.

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Sampling and Characterization of Bioaerosols

2 Principles of bioaerosol collection

a. Aerodynamic diameter

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

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

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

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

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

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

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

a sphere with unit density (that is, a density of 1 g/cm

) 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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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 d

). 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 d

and shift the

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

and

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

two-stage cyclone aerosol sampler has a d

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 (d

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

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

bioaerosol sampling devices employed by industrial hygienists.

a. Filters

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

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

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

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

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

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

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

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

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

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

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

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

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

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

polycarbonate membranes are common choices because cellulose ester membranes can be

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

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

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

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

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

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

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

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

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

eluted from them.

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

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

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

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

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

elsewhere [NIOSH 2016b].

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

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

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

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

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

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

organisms, and reduces the amount of elution media needed for immunological or PCR-

based assays. In areas of high concentration, the microorganisms may have to be eluted,

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

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

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

commonly-used aerosol filters.

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

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

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

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

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

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

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

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

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

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

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

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

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

surfaces of the cassette [Ashley and Harper 2013].

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

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

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

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

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

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

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

2016a].

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b. Impactors

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

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

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

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

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

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

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

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

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

downstream aerosol.

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

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

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

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

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

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

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

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

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

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

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

in culture plates, they may be incubated and examined for microbial growth.

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The most commonly used impactor for sampling airborne culturable bacteria and fungi is

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

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

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

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

that are capable of growth on the specific nutrient media.

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

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

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

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

cascade impactor [NIOSH 2003b].

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

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

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

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

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

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

are collected, and so on for the rest of the stages.

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One significant advantage of the Andersen impactor is that samples can be collected

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

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

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

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

sampler unsuitable for high concentration environments such as some agricultural sites

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

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

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

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

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

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

particles can then be inaccurately counted as a single colony. As the number of organism-

containing 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

= 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 de-

aggregation 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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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 cut-

off 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/cm

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, electret-

based 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 nylon-

flocked swabs and spatulas, cotton swabs and rayon swabs. They reported that nylon-

flocked and cotton swabs were equally effective at removing bacteria from dry

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

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

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

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

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

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

macrofoam swabs performed best when recovering wet or dried norovirus from

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

2015].

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

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

different swab wetting solutions containing various combinations of salts, surfactants

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

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

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

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

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

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

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

sampling. After sample collection, room temperature storage of moist swabs may lead

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to microbial growth if the elution media or swab contain nutrients, while the presence

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

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

Griffith 2007].

Wipes

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

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

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

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

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

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

contaminated with endotoxin and needed to be tested before use.

Adhesive tape

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

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

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

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

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

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

do not provide a quantitative assessment of exposure.

Contact plates

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

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

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

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

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

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

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

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

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

disinfectants. One report indicated that nitrocellulose membranes were slightly more

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

surfaces [Poletti et al. 1999].

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j. Heating, ventilation and air conditioning (HVAC) filters

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

maintain environmental conditions inside buildings. Researchers have taken advantage of

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

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

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

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

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

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

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

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

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

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

HVAC filter analyses were used to study bioaerosols in buildings.

k. Real-time bioaerosol monitoring

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

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

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

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

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

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

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

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

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

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

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

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

4 Considerations for bioaerosol sampling

a. Development of a bioaerosol sampling strategy

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

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

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

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

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

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

sampling methods, times, durations, and the analytical methods can be selected. Note that

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

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

measurements, surface sampling, and information about possible sources.

b. Sampling locations

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

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

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

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

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

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

work are not inadvertently aerosolized.

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

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

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

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

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

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

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

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

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

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

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

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

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

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

c. Concentrations of indoor and outdoor bioaerosols

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

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

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

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

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

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

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

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

in air over time.

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The concentrations of bioaerosol particles vary widely depending upon the meteorological

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

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

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

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

from 1 to more than 8,200 colony-forming units (CFU)/m

of air, with a median of 540

CFU/m

. Indoor levels ranged from 1 to over 10,000 CFU/m

, with a median of 82

CFU/m

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

of 1,100 to 8,400 spores/m

outside and 500 to 101,100 spores/m

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/m

), while indoor concentrations were higher in the summer (116 vs. 87

CFU/m

). 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/m

for fungi, 3000 to

3.3 x 108 CFU/m

for bacteria, and up to 2,800 CFU/m

for actinomycetes. During grain

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

CFU/m

for fungal spores, 40,000 to 1.4 × 106 CFU/m

for bacteria, and up to 2.6 × 104

CFU/m

for actinomycetes.

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

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

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

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

ventilating, and air conditioning system is activated and inactivated.

d. Viable and nonviable bioaerosols

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

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

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

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

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

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

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

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

Mycobacterium tuberculosis is readily transmitted among people and from people to

Guinea pigs, it has never been successfully cultured from an environmental aerosol

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

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

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

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

bioaerosol and culturing the collected particles. Only culturable microorganisms are

enumerated and identified, thus leading to an underestimation of bioaerosol

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

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

microorganisms are then enumerated and identified using microscopy, classical

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

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

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

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

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

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

of viable bacteria have been reported to be as high as 105 CFU/m

in indoor environments;

however, like fungi, the proportion of the total bacterial burden may be higher if non-

viable 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 (d

) 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 flood-

damaged 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 specially-

treated 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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for the field samples, in order to check for sterility of the media. Approximately five media

blanks should be included with each sample set. If the samples are to be analyzed by an

outside laboratory, consult the specific laboratory procedure for the number of blanks to

be submitted. Similarly, blank filters should be processed in the same manner as planned

for field samples in order to check for contamination.

Field blanks are simply unopened, fresh media samples that are handled in the same way

as field samples, including labeling, except that no air is drawn through the sampler. The

generally recommended practice for the number of field blanks is to provide at least two

field blanks for every 10 samples with a maximum of 10 field blanks for each sample set.

5 Selection of bioaerosol samplers

The first step in selecting a bioaerosol sampling device is to establish the purpose of the

sampling. Once the goal of the bioaerosol sampling is determined, the appropriate sampling

methods may be chosen. The selected bioaerosol sampler must be capable of high efficiency

particle collection within the physical and biological conditions required by the

microorganisms to be sampled. The most appropriate sampling methods will be dictated in

part by the techniques that will be used to analyze the sample. Methods for bioaerosol sample

analysis are discussed in the next section. A list of some manufacturers and suppliers of

bioaerosol sampling equipment and supplies is shown in Appendix I. The characteristics of

several commonly used bioaerosol samplers are shown in Appendix II.

a. Sampling for airborne bacteria and fungi

Choosing a bioaerosol sampler for bacteria and fungi begins by deciding how the

bioaerosol will be analyzed, and in particular whether the viability of the bacteria or fungi

will be evaluated. Culturable bioaerosol sampling instruments must minimize injury

during the collection process and maintain the culturability of the collected

microorganisms. If the sample will not be cultured, then the samples usually can be

collected dry using a membrane filter, cyclone, impactor, or a combination of these. Dry

collection is typically simpler and less expensive to perform, and filters and cyclones can

handle a wide range of particle concentrations. Organisms that are difficult or impossible

to grow in culture are often collected using dry techniques and assessed using polymerase

chain reaction (PCR) based methods, which have the advantage of speed and specificity.

PCR has been used for rapid detection of Histoplasma capsulatum and mycobacteria [Reid

and Schafer 1999; Schafer et al. 1999; Schafer et al. 2003]. A DNA-based mold specific

quantitative PCR (msQPCR) method is widely used to evaluate indoor fungal bioaerosols

in the academic, government and commercial sectors, and is the basis for the

Environmental Relative Moldiness Index (ERMI) used to quantify mold contamination in

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homes [Kettleson et al. 2015; Vesper et al. 2013]. The ERMI and other PCR-based assays

are discussed in greater detail later in this chapter.

If viability is to be studied, then the samples usually will need to be collected with an

impinger or in an Andersen impactor loaded with agar plates, because many

microorganisms will lose viability due to damage or desiccation if collected dry [Cox 1987;

Hung et al. 2005]. For example, a membrane filter sampler is not appropriate for sampling

culturable Escherichia coli because the cells desiccate and become either nonviable or

viable but not culturable under these conditions [Jensen et al. 1992]. Similar results have

been reported for other bacteria and fungi [Macher and First 1984; Wang et al. 2001].

Depending upon the target microorganism, impingers may be filled with distilled water or

a buffered isotonic solution, sometimes with antifoaming agents to reduce foaming and

proteins to enhance survival. Mineral oil has also been used in impingers instead of

aqueous solutions to avoid evaporation [Lin et al. 2000]. Impactors are loaded with agar

plates; the choice of agar depends upon the microorganisms of interest and the desired

selectivity (discussed in the next section).

As noted previously, depending upon the investigation that is being conducted, the

particle size distribution of the bioaerosol may be very important in the evaluation of the

data obtained. If particle size information is needed to, for example, determine how much

of the bioaerosol is in the respirable size fraction, then a size-selective sampler should be

used for at least some of the collections if possible. For example, if an SAS-Compact

sampler was the selected sampler for collection of culturable Escherichia coli, an Andersen

6-Stage sampler could be used to determine the particle size distribution at each location

sampled. The expected size of the bioaerosol particles is also an important factor in

choosing a sampler. For example, an impactor with a d50 of 4 µm should not be used to

collect Aspergillus niger spores (dae 1-3 µm) because most spores would remain entrained

in the air and pass through the instrument.

NIOSH Method 0800 discusses sampling for culturable airborne bacteria and fungi with

an Andersen cascade impactor [NIOSH 2003b]. Standard methods for the collection of

airborne fungi by inertial impaction are presented in ASTM Standards D7788 [ASTM

2009; ASTM 2014d]. ASTM Standard D7391 also discusses the aspects related to the

laboratory analysis.

b. Sampling for airborne viruses

Airborne viruses are more difficult to study in bioaerosols than bacteria and fungi for a

variety of reasons [Prussin et al. 2014; Verreault et al. 2008]. Viruses are more difficult to

culture because they are obligatory intracellular parasites that require a host cell for

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reproduction [Tortora et al. 2013]. Bioaerosols of pathogenic viruses have been found in

many settings to be present in low concentrations that can be difficult to detect [Blachere

et al. 2009; Bonifait et al. 2015; Lindsley et al. 2010a; Tseng et al. 2010; Yang et al. 2011].

Viruses also are generally more susceptible to damage during aerosol collection than are

bacteria or fungi, although their sensitivity varies widely with the collection method and

species [Appert et al. 2012; Turgeon et al. 2014; Zuo et al. 2013]. Aerosol sampling

methods for viruses have been reviewed by Verreault et al. [2008].

Bacteriophages are viruses that infect bacteria rather than multicellular organisms. They

are used in laboratory aerosol studies as tracers for aerosol particles and as surrogates for

airborne viruses that infect humans [Fisher et al. 2012; Tseng and Li 2005; Turgeon et al.

2014]. Bacteriophages are not known to be hazardous to humans but are of interest to

industries that rely on bacteria such as cheese manufacturers [Verreault et al. 2011].

Polymerase chain reaction (PCR) based methods are often used to study viral bioaerosols.

PCR has the advantages of being very sensitive and very specific, and considerably easier

to perform than viral cultural assays. For this reason, most recent field studies of airborne

viruses have used PCR as the detection method. Examples include studies of viruses in

healthcare facilities [Blachere et al. 2009; Booth et al. 2005; Lindsley et al. 2010a;

Thompson et al. 2013; Tseng et al. 2010], influenza at poultry and pig farms [Corzo et al.

2013; Jonges et al. 2015], airborne viruses in a sewage treatment plant [Masclaux et al.

2014], and respiratory viruses in human coughs and exhaled breath [Gralton et al. 2013;

Lindsley et al. 2010b; Milton et al. 2013].

PCR has both the advantage and disadvantage of not requiring that the virus be viable in

order to be detected. This eliminates the need to preserve viability during and after

collection and allows the use of dry collection methods such as cyclone samplers, dry

impactors and filters, which are simpler and easier to carry out. On the other hand, this

also means that it is unclear whether the airborne virus is infectious or not, which makes

interpretation of data more difficult. This is a common criticism of PCR-based bioaerosol

studies.

If the virus in a bioaerosol sample is to be cultured, in most cases the sample will need to

be collected into an aqueous media using an impinger or wetted surface aerosol collector.

Fabian et al. [2009] showed that collecting airborne influenza virus in aqueous media

using an SKC BioSampler preserved infectivity much better than dry collection using

filters or an impactor. A less-common method is to collect viable viruses using an

Andersen impactor. Gustin et al. [2011] collected airborne influenza virus using an

Andersen impactor by placing a filter and a thin layer of gelatin on top of the agar in the

culture plates. After collection, the gelatin was removed and melted at 37°C to allow

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subsequent culture of the virus. Note that the collection media must be compatible with

the cell culture system used to host the virus.

6 Sample preparation for culturable bioaerosols

Collecting and culturing viable airborne microorganisms is the most common technique used

by industrial hygienists to assess bioaerosols [Macher 1999]. However, the appropriate sample

preparation method is highly dependent upon the microorganism(s) of interest, sample

source, and down-stream analysis. These sampling approaches are further confounded as

viable bioaerosols have been estimated to account for approximately 1% of the total bioaerosol

load, and non-viable bioaerosols are often overlooked [Hung et al. 2005]. In contrast, non-

culturable 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 non-

enveloped 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 slow-

grower.

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 culture-

based 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 non-

viable 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/m

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/m

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 non-

viable 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 non-

culturable 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 clinically-

based 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-diamidino-

2-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 semi-

solid filter submicron membrane or in the form of a liquid suspension and a three-

dimensional 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. β-

1,3-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 cell-

bound 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 species-

specific 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 bead-

based 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 [Bolaños-

Rosero 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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Sampling and Characterization of Bioaerosols

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 microorganism-

resistant 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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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