Rutgers Electrostatic Passive Sampler (REPS) In Microbial Exposure Assessment

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Rutgers Electrostatic Passive Sampler (REPS) In Microbial Exposure Assessment

Emma-Reetta Musakka

Master’s degree programme in Environmental Science

University of Eastern Finland Faculty of Science and Forestry Department of Environmental and Biological Sciences

May, 2021

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Abstract

University of Eastern Finland, Faculty of Science and Forestry Department of Environmental and Biological Sciences

Master’s Degree Programme in Environmental Health

Musakka, Emma-Reetta: Rutgers Electrostatic Passive Dust Sampler (REPS) in Microbial Exposure Assessment

Master’s thesis, 44 pages, 0 appendices (44 pages)

Thesis instructors: Docent Martin Täubel, Docent Kati Huttunen February 2021

Keywords: airborne dust, indoor microbes, microbial exposure, REPS, Petri dish, settled dust

Majority of our time is spent indoors and thus, exposure to airborne particles happens mostly indoors with respect to duration of exposure. It is known that the microbiome of our environment, also of indoor environment, can have beneficial and adverse impacts on our health. For example, exposure to farm like -microbiome during early years in life has been recently shown to protect from allergic diseases later on.

On the other hand, increased microbial levels in cases of moisture damages in indoor environments are known to cause harmful effects to our health, such as allergies and asthma. Due to the great interest in this connection between microbial exposure and health, development and evaluation of new sampling methods for microbial exposure assessment in large epidemiological studies is important. Here, we studied a new settled dust sampling approach developed for microbial exposure assessment, the Rutgers Electrostatic Passive Sampler (REPS). The REPS was compared to two other sampling approaches:

settled dust samples collected with sterile Petri dishes and active air sampling conducted with Button Inhalable aerosol samplers and Harvard Impactor. The field work was done in three different locations with different expected microbial exposure levels: an office location (low microbial exposure), a home location (moderate exposure) and an outdoor location (high exposure levels). Passive dust samples (collected with sterile Petri dishes and the REPS) were collected for two weeks, four weeks and eight weeks in each location. Button Inhalable aerosol samples were collected for two weeks from all three locations in consecutive 48-hour intervals. Harvard Impactor samples were collected for eight weeks from the office and the outdoor location in consecutive intervals of one week (office location), or 3-4 days (outdoor location). The microbial levels in the various samples were determined with quantitative PCR (qPCR) assays targeting Gram-positive and Gram-negative bacteria, total fungal DNA and

Penicillium/Aspergillus spp. group.

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We observed strong and significant (p<0.05) correlation (ρ=0.80) for microbial measurements between REPS and PM2.5 samples and good correlation (ρ=0.79; p<0.05) between REPS and PM10 samples. Petri dish samples correlated better with active air samples than REPS samples: correlations between Petri dish, PM2.5 and PM10 samples were significant and strong (ρ=0.82 and ρ=0.93, respectively). REPS failed to collect microbes as efficiently as Petri dish, since the microbial levels detected with Petri dish were up to 1000-fold higher compared to levels detected with REPS.

In conclusion, better correlation with active air samples, more consistent microbial levels detected with qPCR, and the high practicality and ease-of-use of the Petri dish samplers argue that Petri dish sampling is a preferable settled dust sampling approach compared to the REPS to be used in epidemiological studies placed in low or moderate exposure environments.

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Tiivistelmä

Itä-Suomen yliopisto, Luonnontieteiden ja metsätieteiden tiedekunta Ympäristötieteen laitos

Ympäristötieteen koulutusohjelma

Musakka, Emma-Reetta: Rutgers Electrostatic Passive Sampler (REPS)- laskeutuneen pölyn keräimen käyttö mikrobialtistuksen arvioinnissa

Pro gradu, 44 sivua, 0 liite/liitettä (44 sivua)

Ohjaajat: Dosentti Martin Täubel, Dosentti Kati Huttunen Helmikuu 2021

Avainsanat: laskeutunut pöly, mikrobialtistus, REPS, keräysmenetelmä

Länsimaissa suurin osa ajasta vietetään sisätiloissa, minkä vuoksi kiinnostus sisäilman mikrobialtistusta kohtaan on kasvanut merkittävästi viimeaikoina. Ympäristömme mikrobikoostumuksella tiedetään olevan merkittäviä vaikutuksia terveyteemme. Maatilamainen mikrobialtistus lapsuusiässä on nykytiedon mukaan suojaava tekijä astmaa ja allergisia sairauksia vastaan myöhemmällä aikuisiällä.

Toisaalta, lisääntyneet sisäilmaongelmat, kuten rakennusten kosteusvauriot muuttavat sisäilman mikrobijakaumaa, minkä tiedetään edelleen aiheuttavan allergisia sairauksia ja astmaa. Tietoa yhteydestä mikrobien ja niiden terveysvaikutusten välillä tarvitaan lisää, minkä vuoksi tutkimuksissa käytettävien keräysmenetelmien tulee olla luotettavia sekä käytännöllisiä. Uusia keräysmenetelmiä kehitetään jatkuvasti, mutta ennen niiden laaja-alaisempaa käyttöä tulisi kyseisten menetelmien luotettavuus arvioida huolellisesti. Tässä tutkimuksessa selvitettiin uuden laskeutuneen pölyn keräysmenetelmän, REPS (Rutgers Electrostatic Passive Sampler) -keräimen toimivuutta ja

luotettavuutta. REPS -keräintä verrattiin kahteen eri keräysmenetelmään: Petrimalja -keräimiin, sekä aktiiviseen menetelmään, jossa näytteet kerättiin Button -keräimillä. Myös Harvard Impaktorilla kerättiin näytteet PM2.5 (pienhiukkaset, joiden läpimitta on alle 2,5µm) ja PM10 (pienhiukkaset, joiden läpimitta on alle 10µm). Tutkimus toteutettiin kolmessa eri ympäristössä, joiden mikrobialtistuksen odotettiin olevan erilainen. Nämä näytteenottokohteet olivat toimistoympäristö, kerrostaloasunto, sekä ulkoilmaympäristö. Näytteenotto kesti jokaisessa kohteessa kahdeksan viikkoa. Laskeutuneen pölyn näytteet (keräys Petrimaljoilla ja REPS-keräimellä) kerättiin kahden, neljän ja kahdeksan viikon ajan.

Aktiivisella Button -keräimellä näytteet kerättiin jokaisesta kohteesta kahden viikon ajalta kahden vuorokauden sykleissä, ja Harvard Impaktorilla kahdeksan viikon ajalta toimisto- ja

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ulkoilmaympäristöistä viikon sykleissä (toimistoympäristö) tai 3-4 vuorokauden sykleissä (ulkoilmaympäristö).

Tutkimuksessa REPS näytteet korreloivat vahvasti PM2.5 näytteisiin (ρ=0.80) ja hyvin PM10 näytteisiin (ρ=0.78). Petrimalja, sekä PM2.5 ja PM10 näytteiden välillä havaittiin vahva korrelaatio (ρ=0.82, ρ=0.93).

Petrimaljanäytteet korreloivat ilmanäytteiden kanssa paremmin, minkä lisäksi ne ovat myös

käytännöllisempiä laboratorio –ja kenttätyössä, sekä niillä kerättyjen näytteiden qPCR -menetelmällä havaitut mikrobipitoisuudet olivat loogisempia keräysympäristö –ja kesto huomioonottaen. Tämän vuoksi Petrimaljamenetelmä on REPS -menetelmää toimivampi laskeutuneen pölyn keräyksessä.

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Foreword

I conducted this thesis during January 2020 to October 2020. Planning of the work was done from January 2020 to February 2020, and field work was carried out in March 2020 to May 2020. I started laboratory work as soon as the field work was finished, from May 2020 to July 2020. Planning, field work and laboratory work was done in Kuopio, Finland at Environmental Health Unit of Finnish Institute for Health and Welfare (THL). Planning was done in cooperation with Martin Täubel from Finnish Institute for Health and Welfare, field- and laboratory work was carried out largely by the writer, Emma-Reetta

Musakka, with support in the qPCR analysis by the skilled laboratory personnel at THL. Writing was supervised by Kati Huttunen from University of Eastern Finland and Martin Täubel from Finnish Institute for Health and Welfare.

The purpose of this thesis was to evaluate a new settled dust sampling approach, the Rutgers

Electrostatic Passive Sampler (REPS), for use in microbial exposure assessment. This sampling approach was developed in the State University of New Jersey, aiming at more efficient settled dust sampling due to the electrostatic material used in the sampler. In this study, the REPS-sampler was compared with three other, widely used sampling approaches: settled dust sampling conducted with sterile Petri dishes and active air sampling conducted with Button Inhalable aerosol samplers and with Harvard Impaction.

I kindly thank Gediminas Mainelis from the State University of New Jersey for providing the instructions for proper use for the REPS, and also for providing the REPS for our field work. All the personnel from Environmental Health unit at Finnish Institute for Health and Welfare provided major help with planning and conducting the field- and laboratory work: Pekka Taimisto, Heli Martikainen and Mervi Ojala: thank you for all the answers for the million questions I had during this work. A special acknowledgement for Mervi Ojala for all the help with qPCR-analysis. Even though the COVID-19 situation brought some extra challenges for all the work, all the analysis ran smoothly: thank you Maria Valkonen for all the efforts with my qPCR- analytics and organizing everything with the laboratory, and Katja Saarnio for all the help with DNA-extraction. I can’t thank both my supervisors enough for all the efforts on my thesis: I could possibly not ask for better supervisors. Martin Täubel has answered so many of my questions during my work, and I really appreciate the possibility he, and all the Environmental Health Unit provided me with carrying out this thesis with them.

Also, I want to thank my family and friends for always supporting me, and my crazy dreams.

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Abbreviations

REPS Rutgers Electrostatic Passive Sampler qPCR Quantitative Polymerase Chain Reaction

PM2,5 Airborne particulate matter, diameter less than 2,5µm PM10 Airborne particulate matter, diameter less than 10µm NCD Non-communicable disease

EDC Electrostatic Dust fall Collectors

THL Terveyden ja Hyvinvoinnin laitos, Finnish Institute for Health and Welfare PTFE Polytetrafluoroethylene

PVDF Polyvinylidenefluoride OAF Open Air Factor

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1 Introduction

The environment we spend our time in has changed radically after the industrial revolution. In large parts of today’s world, it is very common that people spend more time indoors than outdoors in their everyday lives, a trend that is further stimulated by more and more people living in urban environments.

These changes in exposure to macro-environments have gone hand-in-hand with changes of the microbiota we are exposed to. The microbiota in different environments vary, and it is known that the microbiota of outdoor and indoor environments differ from each other. In addition to the surrounding outdoor environment, the indoor microbiota is impacted by multiple factors within the building, such as ventilation, house pets, occupancy or cleaning frequency.

It is known that the microbiological factors we are exposed to affect our health, beyond just infectious diseases. Non-communicable diseases (NCDs) such as allergies and asthma have been linked to indoor microbial exposures. A recent discovery states that “farm like” microbiome exposure indoors, during the early years in life, can provide protection against the development of asthma, even when growing up in an urban environment. At the same time, certain microbial exposure is believed to cause adverse health effects, such as infections and asthma. These harmful health effects can be a result from for example exposure to microbiological factors associated with moisture damage. Beyond infectious diseases, it is still to be clarified, why exposure to some microbes have harmful, and some on the other hand, positive effects on our health. In this effort to provide answers, it is important to understand the exact exposures and mechanisms underlying these health effects. Immune system response to different microbiological factors varies individually, and we lack thorough understanding of how different individuals react to different microbial exposure situations.

Since there are many aspects still to be clarified when it comes to the association of human health and the microbiomes of indoor and outdoor environments, there is great need for future research in this specific field. For the upcoming research projects, reliable exposure assessment approaches, including microbial sampling methods will be important. There are various microbial sampling methods that are being used, none of them perfect, and all with their individual strengths and limitations. New methods are developed constantly; in order to assure those provide reliable research results, they need to be tested properly before putting them to use in field studies.

Indoor air quality is determined by multiple factors, such as chemicals, allergens, particulate matter and microbes, along with temperature, relative humidity and other parameters. In my thesis, I will focus on

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the microbial component of indoor air. Since the indoor air quality is of great research interest and relevance to public health, standardized sampling methods and laboratory techniques must be available in order to provide the reliable information needed regarding this subject. Two of the most commonly used techniques for microbial exposure assessment are active sampling of airborne particulate matter and passive sampling of settled dust. In my thesis I will focus on one of the latest applications of settled dust sampling, the Rutgers Electrostatic Passive Sampler, or REPS. In this thesis, I will provide the background on sampling approaches relevant to this topic and present a study, where REPS was

compared to two other sampling approaches – passive settled dust sampling with Petri dishes and active air sampling onto filter membranes.

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2 Background

There is a large variety of sampling applications developed for microbial exposure assessment for both indoor and outdoor environments. In this chapter, I will describe some of the most commonly used sampling methods, with focus on the ones that are relevant to this study.

Sampling methods for indoor microbial exposure assessment can be categorized in three general categories: active air sampling, passive sampling of settled dust and sampling of dust reservoirs. The basic idea in all these sampling methods is to collect particles that have been or are airborne. These particles have microbes attached to them, and these microbial signals can be detected with a variety of microbiological and/or molecular biological laboratory analysis, including traditional cultivation of viable fungi and bacteria; measurement of cell wall markers of microbes, such as bacterial endotoxins, muramic acid, or fungal glucans; and molecular DNA-based approaches, including quantitative PCR (qPCR) and next-generation sequencing. For this study, active air sampling and settled dust sampling are the relevant sampling methods, so my literature review will focus on these two sampling approaches.

2.1 Active air sampling

Active air sampling is often used as a reference method in microbial exposure assessment, because it is thought to represent the airborne microbial composition the best. It has been used as the standard method for microbial sampling in multiple studies, such as Frankel et al. (2012) and Leppänen et al.

(2018). In these studies, active air sampling was used as the “gold standard” for comparison to other sampling methods. In this study, we used filtration- and impaction techniques in active air sampling, but active air sampling can be done also via impingement. Because we used filtration and impaction here, I will introduce these active air sampling techniques in this literature review.

2.1.1 Filtration technique

With filtration, airborne particles are collected onto a filter utilizing a pump. The pump causes air to flow through the filter, which allows the airborne bioaerosols to adhere on the surface of the filter. Particles diffuse onto the filter, and electrostatic forces keep the particles adhered on the surface (Viegas et al.

(2017)). Active air sampling can be done with different pumps, depending on the purpose of the study.

For example, Karlsson et al. (2002), used personal air sampling for students when detecting cat allergen

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from classroom environment. In another study, (Hyvärinen et al. (2006)), a stationary pump was used to compare the capability of different sampling methods to collect endotoxin from indoor environment.

Also, different filters can be used depending on for example the pump used in the study. Ultimately, the purpose of the study - including the environment to be sampled; personal versus stationary sampling;

the duration of sampling relating to the aim being short-term or long-term exposure assessment; and the type of microbial agent that will be analyzed – will determine which kind of pump, sampling device, air flow rates and filter is to be used in the study.

2.1.2 Impaction technique

Impaction with inertial sampling mechanism is the most popular air sampling technique. In addition to inertial, impaction can be also centrifugal. In both impaction techniques, the air sample, which in the context of this study contains bioaerosols, can be collected straight onto a filter, agar plate, glass slide or a tape. Sample collection is determined by the purpose of the study and which microbial or molecular biological analysis is to be performed post sampling. With inertial impaction technique, a sample is collected onto a desired surface by pushing or pulling the air through a small nozzle with a help of a pump. Particles are then impacted because of their size: smaller particles make a turn before the impaction surface, while larger particles become impacted because of their inertia, with the impactor design and associated air flow determining the cut-off of collected particulate matter. In centrifugal impaction, centrifugal air forces are utilized: larger particles get impacted, because they are not able to follow the air streamlines in a way the smaller particles do (Viegas et al. (2017)). In this study, inertial impaction was utilized when collecting active air samples with Harvard Impactor, a standard device for collecting airborne PM2.5 and PM10..

2.1.3 Challenges in active air sampling

Even though active air sampling is considered as the reference method and gold-standard in airborne microbial exposure assessment, this sampling approach has also some problematic features. Hyvärinen et al. (2001) used impaction in their study to determine the temporal and spatial levels of indoor air concentrations of viable fungi. They noticed that indoor fungal concentrations in air determined from short term air samples using an Andersen impactor and traditional cultivation varied throughout the day, and concentrations were usually higher in the morning. This study very nicely illustrates one of the issues with active air sampling, which is the high variability of microbial concentrations in indoor air over time.

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Other studies have shown that concentrations of fungi in indoor air vary during the day, and is affected for example by the activity of occupants during the day (Miller et al. (1992), Verhoeff et al. (1990)).

Typically, the sampling periods applied with active air sampling is noticeably shorter – minutes to hours - compared to settled dust sampling periods (days to weeks). Due to temporal variability of airborne microbe concentrations, there is the risk that a short-term air sample does not represent the general microbial exposure levels in a space correctly.

If the study aims to evaluate short-term exposure, this variability is not an issue. If long-term exposure is of interest, however, this within -day variation is problematic in that single short-term samples may greatly mislead assumptions of long-term exposure. This can be improved with longer sampling periods, though this can create some issues also. It has been noticed that extended sampling periods in active air sampling with filtration technique can cause the filter to become saturated with collected dust,

interfering with collection efficiencies (Normand et al. (2009)). In their study, Normand et al. (2009) also noticed that long sampling periods can cause not only filter saturation, but also the collected microbes to become desiccated. This can be problematic, because microbial desiccation can result in lower detected microbial levels due to reduction in viability (if cultivation is the analysis method) or degradation of targeted molecular compounds, including cell wall markers and DNA. The microbial concentrations in indoor air typically also vary throughout the year due to the dependency of indoor microbial levels of outdoor levels and seasonal variation thereof. For example, in Northern countries with snow cover and subzero degrees during winter, usually indoor microbial levels are lower during winter than during the summer months (Reponen et al. (1992)). It is important to notice, that this issue is not unique to air sampling, but is also present with settled dust sampling –and reservoir sampling techniques. On a more technical, logistic and financial note, active air sampling is generally equipment and personnel intensive, requiring field personnel in studies which makes such sampling campaigns cost intensive. Because of these problematic features of active air sampling, settled dust sampling has become increasingly popular, specifically when the aim is to assess long –term microbial exposure.

2.2 Settled dust sampling

There are multiple approaches of how to collect settled dust, such as active collection of dust settled from elevated surfaces with a vacuum cleaner (eg. Jayaprakash et al. 2017), and sampling airborne dust passively onto a sampler utilizing electrostatic forces of a sampling surface (electrostatic dust fall

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collectors or EDCs) (Noss et al. 2008). In the following, I will introduce the approaches relevant to my study, i.e. Petri dish sampling and REPS-sampling for the collection of settling airborne particles.

2.2.1 Petri dish sampling

Petri dish sampling has been suggested to be a practical and reliable sampling approach in multiple studies, and it is a commonly used sampling approach for settled dust. For example, Adams et al. (2015) noticed in their study, that Petri dish approach for settled dust sampling coupled with DNA-based microbial determinations yielded high correlations between duplicate samples, and therefore has potential to be a useful sampling approach in epidemiological studies. In the same study, it was concluded that Petri dishes are an inexpensive sampling approach, and it also indicated highest quantitative determinations compared to for example EDC’s, which are also widely used samplers in settled dust sampling.

Leppänen et al. (2017) suggested Petri dish approach to reflect well the microbial composition of indoor air, but they recommended other sampling approaches to be assessed in parallel for better quantitative assessments (for example dust reservoir samples). Similar conclusions have been made in other studies too, such as in Karlsson et al. (2002). In their study, these authors noticed significant correlation between Petri dish sampling and personal air sampling in the analysis of cat allergens (rho=0.66; p<0.0001). These results indicate that Petri dish sampling can be a more practical approach than active air sampling, when long term exposure to airborne particles is of interest.

Petri dish sampling has become a popular settled dust sampling approach not only due to the indications of its repeatability and reliability, but also due to some practical features. First of all, Petri dishes are inexpensive to purchase, and their small size and light weight is practical for field work. Also, post- sampling and pre-analysis processing of the settled dust collected in Petri dishes is relatively simple and quick, in that no time-consuming steps or expensive equipment are required. Extraction of particulate matter from EDCs for example involves multiple processing steps including a stomacher or shaker for particle release and centrifugation to concentrate the large amounts of buffer required for an efficient extraction (Adams et al. 2015).

In contrast, settled dust from Petri dishes can be collected quickly onto a cotton swab that then can be further extracted for analysis. Settled dust sampling with Petri dishes itself can be done easily without

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trained personnel, which is particularly important in large epidemiological studies that require study participants themselves to carry out indoor environmental sampling campaigns. These practical

advantages alongside the apparent good repeatability and correlation with active air sampling highlights Petri dish sampling of settled dust as a useful, promising indoor microbial sampling approach.

2.2.2 Challenges in settled dust sampling

Even though Petri dish sampling has been used as a sampling method to assess the long term microbial exposure, there are some challenges regarding the Petri dish, and also settled dust sampling approaches in general.

One issue regarding settled dust sampling approaches is the resuspension of settled dust particles. This is discussed in a study by Adams et al. 2015, and they hypothesize, that the resuspension of settled dust depends on the samplers’ material. With extended sampling periods the sampler can become saturated with dust, and this can prevent further sampling to be accurate. Also, settled dust sampling is rarely conducted in a space where absolutely no air flow is present. The air flow in the space where settled dust sampling is conducted is problematic, because it can cause resuspension of settled dust from smooth- surfaced samplers (Frankel et al. 2012). Therefore, settled dust sampling is not as efficient in a space where air flow is present, and this should be considered when appropriate sampling location is chosen in a sampling space.

Another problematic feature for settled dust sampling is the difference in sampling efficiencies between PM2.5 and PM10. In their study, Kilburg-Basnyat et al. 2016 noticed, that EDCs collected PM2.5 particles more effectively than PM2.5-PM10 particles. At the same time it is known, that the coarse particulate matter (PM2.5-PM10) contain approximately 10 times higher concentrations of endotoxin compared to the fine particulate matter (PM2.5) (Heinrich et al. 2003). These factors can result into unrealistic

measurement results of endotoxin, and other biological factors when EDCs, or settled dust sampling approach in general, is used. More research is needed in order to understand the sampling efficiencies of different particles for individual settled dust sampling approaches.

In several previous studies carried out in the research group at THL, it has been noticed that specifically Gram-positive bacterial levels detected from outdoor locations sampled with Petri dishes, followed up by qPCR-method, returned implausibly low results. This problem has not been studied in further detail, but

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there are some potential explanations for this issue. One of these explanations could relate to the

sampling approach that would make Gram-positive bacteria susceptible to degradation on the petri dish surface. More likely, however, are issues with the Gram-positive bacterial probe in the quantitative PCR assay used in studies (Kärkkäinen P, Valkonen M, et al. 2010).

2.2.3 REPS-sampling

The REPS is the least studied of all the sampling approaches discussed in this thesis, which motivated us to carry out a targeted study. This sampling approach has been introduced by Therkorn et al. (2017), where they compared the REPS with Button Inhalable sampler, PTFE (Polytetrafluoroethylene) settling filters and Agar settling plates. The REPS is a passive dust sampler, which has electrostatic properties due to the polyrized PVDF (poly(vinylidenefluoride)) film that is utilized in its structure. Therefore, the sampling of airborne particles is not depending solely on particles settling under gravity, but also on the electrostatic forces attracting the airborne particles to attach on the PVDF film utilized in the REPS.

Because of these electrostatic properties of the REPS, the sampling efficiency is supposedly higher than it is for example with Petri dish sampling approach (Thekorn et al. (2017)).

The REPS is designed in a way that the sampling surface area is as large as possible at a relatively small sampler size. This has been achieved with using a spiral shape for the REPS. The REPS has also been designed to be as practical as possible in laboratory analysis, since the shape of the sampler is designed in a way that it fits into a 50mL conical centrifuge tube. This eases the processing before and after sampling campaigns (Therkorn et al. (2017)). The structure of the REPS is presented in picture 1.

In their study, Therkorn et al. (2017) noted some promising results about the REPS. They discovered that the REPS provided greater passive collection of total bacteria and fungi compared to passive PTFE settling filters, and settling agar plates. This was thought to be the result of the electrostatic properties of the REPS, which increases the sampling efficiency compared to the sampling methods relying only on the gravitational settling of airborne particles. When Therkorn et al. (2017) compared the REPS sampling approach with active air sampling using Button Inhalable samplers they discovered, that the REPS had enhanced performance in culturable bacteria collection.

The most important result Therkorn et al. (2017) found from comparison of the REPS and active air sampling was the higher practicality of REPS compared to active air sampling, with the REPS collecting

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airborne particles relatively efficiently because of the electrostatic forces created by PVDF film, without the need for air flow, sampling pumps and personnel operating the equipment in the field.

Picture 1. Front and top views of the REPS film holder (Therkorn et al. (2017)). PVDF (polyvinyliudene fluoride) film is spiraled through the film holder openings, as presented in the top view of the picture.

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3 Aim of the study

In this study, we evaluated a new settled dust sampling approach using the Rutgers Electrostatic Passive Sampler (REPS) for assessment of airborne microbial levels. This study had two main purposes: first, we wanted to investigate, whether the REPS could be used as an advanced passive settled dust sampling method for microbial assessments in epidemiological studies instead of settled dust sampling with Petri dishes. To clarify this, we compared the performance of these two settled dust sampling methods side- by-side in different sampling locations. Second, we wanted to study how well microbial determinations from dust passively collected with the REPS correlates with microbial levels determined from active air filter samples. The purpose of this was to establish, whether the REPS represents the airborne microbial levels well, and to compare REPS with Petri dish approach also in this aspect. For this, we paralleled the settled dust assessment with active air sampling with Harvard Impactors in two sampling locations, and with Button Inhalable samplers in three sampling locations, in order to evaluate correlations in microbial levels determined by these different sampling methods.

In addition to these two main aspects of this study, we also wanted to evaluate the practicality of the REPS in field work and laboratory processing. In epidemiological studies that assess indoor microbial exposures it is often the case that high numbers of samples are to be collected from a large study population. It is therefore crucial to have a feasible and simple sampling approach, which can be

implemented by study participants themselves. Because of this, we were interested in the handling and practicality of the REPS compared to Petri dish sampling approach, which is currently a widely used sampling approach for settled dust.

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4 Materials and methods

4.1 Sampling locations

Sampling was done in three different locations with different expected microbial exposure levels.

Description of the locations, numbers of individual samples, and sampling durations for each sampling approach applied in this study are presented in table 1. EXP 1 was an office indoor location (about 10 m²) with low expected microbial levels in air. EXP 2 was a residential flat in an apartment building (48 m², 8th floor) in an urban area in Eastern Finland, where moderate microbial levels were expected. EXP 3 was an outdoor location, where high microbial levels in air were expected. The outdoor location (EXP 3) was located in a green, park-like area in vicinity of a heavily trafficked (about 450 meters) and a less heavily trafficked (about 100 meters) road. Some office buildings were located near EXP 3 sampling location. All three sampling locations were located within 4 km distance. In each sampling location, samples were collected for two months, between February 2020 and April 2020.

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Table 1. Description of sampling locations (EXP1-3), sample numbers and sampling durations for the various sampling approaches

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4.2 Sampling approaches

4.2.1 Active air sampling Impaction

Active air samples using PM2.5 and PM10 Harvard impactors were collected from two locations: EXP 1 office location and the EXP 3 outdoor location. Sampling in EXP 2 location was not feasible due to the noise levels caused by active sampling pumps in the small apartment. In the indoor location, PM samples were collected in eight consequent one-week periods, covering the entire eight-week sampling period. In the outdoor location, samples were collected in consequent 3-4 day-sampling intervals, covering the entire eight week-period as well. The shorter sampling intervals for impaction sampling in the outdoor setting was necessary in order to avoid filter saturation and overload due to the higher expected particle levels in the outdoor environment. Two field blanks were included for each sampling location.

PTFE filters (3 µm pore size, 37 mm diameter [Merck Millipore]) were equilibrated for 24 hours in

standardized conditions (temperature: 22,7 C°-23,7 C°; RH: 29%-38%; pressure: 745 mmHg-767 mmHg).

After equilibration, pre-sampling weight of filters was determined on a Mettler Toledo scale, (model XP 6), filters were transferred into sterile filter holders and placed in a plastic cassette, and each filter cassette was coded with a sample code. Filters were kept at +5 °C prior to sampling, including transportation between laboratory and sampling locations. In the outdoor sampling environment, Harvard Impactors were set at a height of 160 cm, and sampling was performed with a Wageningen UR pump (model ASP-100R). Flow rates during the sampling periods were between 9.1066 l/min and 9.3622 l/min for the PM2.5 impactor. For PM10 samples, the flow rates during sampling were between 9.5449 l/min and 9.9834 l/min. In the office sampling location, Harvard Impactors were set at 135 cm height, and flow rate during the sampling periods were between 9.241 l/min and 9.9433 l/min for PM2.5 and between 9.1198 l/min and 10.018 l/min for PM10 impactor.

In the office location, impactors were attached to an Air Diagnostics and Engineering Inc. pump (model SP-280E). The flow rates were measured with a calibrator (Bios International Corporation: Defender 510- H) before and after each sampling period and marked into a sample log sheet. After sampling, filters were stored at -20 °C until post-sampling weighing (Mettled Toledo, model XP 6). Before post-sampling

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weighing, the filters were equilibrated in the same conditions as before pre-sampling weighing.

Immediately after post-weighing, filters were cut in half using sterile scissors and tweezers. One half was transferred into a coded glassbead tube and stored at -20 °C until DNA extraction. The other half was transferred into petri slide for possible later analysis and stored at -20 °C.

Button Inhalable sampling

For Button sampling, we performed sampling for a total of two weeks for each location, and button samples were collected in consecutive 48 hours intervals. The Button samples were collected in duplicate, summing up to a sample number of 14 for each sampling location. Two field blanks were included in each of the sampling locations.

For Button sampling we used PTFE filters with a pore-size of 0.45 µm (Millipore: Fluoropore™, 25 mm Membrane filters). Button samplers (pre-cleaned by 30 min sonicating in water and subsequently washed with Etax A) were assembled in the laboratory with these filters using sterile tweezers, coded with a sample code, and attached to a pump (Air Diagnostics and Engineering Inc., model SP-280E). The details of Button sampler assembly is presented in picture 2. Flow rates on the samplers were between 3.4645 l/min and 4.5087 l/min during sampling. The flow rates were measured (Bios International Corporation:

Defender 510-H) before and after each sampling, and marked into a sample log. After each sampling, Button samplers were dissembled in the laboratory and filters were transferred into a sterile petri slide coded with the sample code, and stored in -20 C° until further analysis.

Picture 2. The assembly of Button Inhalable sampler.

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4.2.2 Settled dust sampling

Settled dust samples were collected with two different methods: sterile Petri dishes and the REPS.

Settled dust samples were collected in triplicate with both methods in parallel, over three different time periods: two weeks, four weeks and eight weeks.

Petri dish sampling

Two sterile Petri dishes were combined into one sample, thus six Petri dishes (resulting in triplicate samples) were set up in each sampling location for each sampling period. Petri dishes were coded with an individual sample code and taken to the sampling location unopened in a sterile plastic bag. Opening of the Petri dish samplers to initiate sampling was done with laboratory gloves to avoid any

contamination. After sampling, Petri dishes were closed – again using laboratory gloves - sealed with parafilm and transported to the laboratory for further processing. Pre-processing of Petri dishes was done right after collecting the samples in the field. Pre-processing was done by swabbing the Petri dishes (both the dish and the lid) with sterile cotton swabs. Swabs were wetted with a buffer (sterilized

deionized water+0.05% tween 20) before swabbing and as mentioned, always two Petri dishes were combined onto one swab, representing one sample. After swabbing, the tip of the cotton swab was transferred into a glassbead tube suitable for DNA extraction coded with sample code, by cutting it with sterile scissors. The samples were transferred to -20 °C immediately after pre-processing, until DNA extraction. In the office location (EXP 1), sampling height was 210 cm. In the home location (EXP 2), sampling height was 150 cm, and for outdoor location (EXP 3), 130 cm.

REPS-sampling

One REPS was used always as one sample, thus three REPS (for triplicate analysis) were set up in each sampling location for each sampling period (2, 4 and 8 weeks, in parallel with Petri dish samplers). The REPS is presented in picture 1. Samplers are composed of sterile 3D-printed housing containing a sterile PVDF film (provided assembled as such by Rutgers University), fixed with autoclaved paper clips to REPS holders that are placed on the sampling surface (located at the same height and in immediate proximity to Petri dishes). The REPS were transported between laboratory and the sampling locations in sterile conical 50 mL centrifuge tubes coded with sample code.

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The REPS-samples were pre-processed immediately after each sampling. For pre-processing, 35 mL of sterile, deionized water was pipetted into the 50 mL centrifuge tube with the REPS-sampler in it. After this, two minutes of vortexing with maximum speed, and ten minutes of sonicating in an ultrasonic water bath (Branson Bransonic-3510) were applied. After vortexing and sonicating, the sample tube was

shaken vigorously, and the liquid was poured into a sterile beaker coded with the sample code. The liquid was divided for further analysis: 200 µL was transferred into a glass bead tube for DNA extraction, 30 mL was transferred into a new sterile conical centrifuge tube, and the rest into a sterile 10 mL conical centrifuge tube. All the vials were stored into -20 C° until further analysis.

Because only a weak microbial signal was detected during the first qPCR–analysis of the REPS-samples using the approach above with a volume of 200 µL REPS buffer solution for DNA extraction, we further concentrated the REPS-samples from 30 mL aliquots in order to yield a stronger microbial signal. For this, we centrifuged the 30 mL aliquots of the REPS-samples for 15 minutes with 6000 x G (Thermo Fisher Scientific, Sorvall Lynx 4000). After centrifugation, we removed the supernatant down to a volume of 500 µl carefully without disturbing the pellet at the bottom of the centrifugation tube. This was followed by resuspending the pellet in the remaining buffer (sterile deionized water). The 500 µL suspension was then transferred into a sterile Eppendorf tube (Eppendorf® Safe-Lock microcentrifuge tube, 1.5 mL). The original 30 mL centrifugation tube was rinsed with 250 µl of sterile deionized water by vortexing, and this buffer (sterile deionized water) was added to the same Eppendorf tube as the initial sample, thus ending up with approximately 750 µl of REPS suspension. After vortexing, 200 µl of this suspension was pipetted into a glass bead tube for final DNA extraction, and rest of the suspension was kept in the 1.5 mL

Eppendorf tube for future analysis. Both of these aliquots were kept in -20 °C until further processing.

4.3 DNA-extraction

All samples were kept in -20 °C before DNA extraction. DNA was extracted from all sample types as follows: a 200 µl aliquot from the REPS -samples, a cotton swab from Petri dish sampling, a 25 mm filter from button sampling, and a half 37 mm filter from Harvard impaction. All the samples were transferred into a glass bead tube for DNA extraction as described previously.

For DNA extraction, we used 1 minute beat beating step in lysis buffer for mechanical cell disruption (MiniBeadbeater, Biospec Products Inc., USA), followed by DNA purification with Chemagic DNA plant kit (PerkinElmer chemagen Technologie GmbG, Germany). We performed the extractions on a KingFisher

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DNA extraction robot (Thermo Scientific, Finland). As an internal standard, we added salmon testis DNA (Sigma Aldrich Co., USA) to all samples and controls, in order to control for qPCR inhibition and

differences in DNA extraction efficiency (Haugland et al. (2005)). The DNA extraction method we used is described previously by Jayaprakash et al. (2017). Negative and positive controls were included in DNA extraction, alongside real samples and blanks. Negative controls were added in order to control for possible contamination of the used reagents. Positive controls (known bacterial and fungal mock communities) were added to confirm the success of DNA extraction and qPCR reactions.

4.4 Quantitative PCR analysis

The qPCR methods used in this thesis have previously been described in detail, for example by Hyytiäinen et al. (2018). Previously published qPCR assays were utilized to determine these following bacterial and fungal groups from all the samples: Gram-positive and Gram-negative bacteria (Kärkkäinen et al. (2010)); group of Penicillium spp., Aspergillus spp. and Paecilomyces variotii (referred to as PenAsp group in the results section); total fungal DNA (Haugland et al. (2003)); and internal standard salmon testis DNA (Haugland et al. (2005)). QPCR reactions were performed in this study with minor

modifications to the original publications, as detailed in Hyytiäinen et al. (2018). QPCR reactions were performed with the Stratagene Mx3005P qPCR System (Agilent Technologies Inc., CA, USA) equipment in a 96-well plate (volume 0,2 mL, Agilent Technologies Inc., CA, USA). Positive and negative controls, as well as no template controls were included in the qPCR runs. Microbial cell equivalents (CE) were calculated as describer earlier (Haugland et al. (2004)). Number of microbial cell equivalents was normalized for each sample type: cells per sample for the REPS, cells per m³ for the Button Inhalable samples, cells per sample for the Impaction samples and cells per two Petri dishes for the Petri dish samples. Results were blank corrected in all cases, using the average of the background CEs of blanks per each sample type and qPCR assay. For Button Inhalable samples, five field blanks were included, for Impaction samples four, for REPS three, and for Petri dish three, respectively.

4.5 Statistical analysis

Statistical analysis was conducted using Microsoft Excel and IBM SPSS Statistics, version 26. Correlations between microbial levels in different sample types were analyzed using Spearman’s rank correlation.

Correlation between REPS, Petri dish, PM2.5 and PM10 samples was analyzed, from EXP 1 and EXP 3 locations. For the outdoor EXP 3 location, qPCR results for Gram-positive bacteria were discarded from

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the analysis, because the levels for Gram-positive bacteria detected from outdoor locations have been found to be unreliable. Button Inhalable samples were not included in correlation analysis, because only a two-week sampling period was applied for this sampling method. The Button samples were used for later studies of microbial communities in different sample types via sequencing analysis, but this module is not included in this thesis.

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5 Results

5.1 Comparison between REPS and Petri dish sampling

In order to compare the performance of REPS with Petri dish settled dust sampling we measured

microbial levels from settled dust sampling campaigns carried out in parallel with REPS and Petri dishes in three locations and over three sampling durations (2, 4 and 8 weeks). The results are summarized in Table 2, presenting the average and standard deviation of triplicate measurements for each timepoint and qPCR measurement. It is clearly visible that Petri dish samples return higher microbial levels compared to REPS in all cases, with microbial levels determined from settled dust collected with one Petri dish being up to 1000-fold higher compared to determinations from one REPS. The low microbial exposure environment EXP1 does not return measureable microbe levels when using REPS, while Petri dish samples do return detectable levels (note: after correcting blank levels). Datapoints for Gram- positive bacteria from Petri dish samples in the outdoor location (EXP3) are missing from Table 2 due to these determinations from Petri dish settled dust samples found to be unreliable, possibly for reasons related to qPCR assay.

For Petri dish samples (Chart 1) we observe that microbial levels generally, but not always, tend to increase from shorter to longer term sampling periods, but increases are not linear and depend to some extent on the microbial group measured. For outdoor sampling location EXP 3 levels are decreasing in all microbial groups from two to eight weeks sampling periods in petri dish samples. For REPS samples (Chart 1) only weak microbial signal was detected from the low exposure environment (EXP 1) with Penasp and Unifung qPCR assay. In EXP 2 sampling environment, microbial levels were detected, but they were not increasing simultaneously with extended sampling periods. For EXP 3 environment, the REPS seemed to collect microbial matter more consistently with increasing levels with increasing sampling duration, outcompeting Petri dish with that respect.

Table 2. Microbial levels from REPS and Petri dish settled dust samples in three sampling locations (EXP 1-3) and from three sampling periods (2,4,8 weeks) for the microbial groups determined with qPCR assays (Grampos = Gram-positive bacteria; Gramneg = Gram-negative bacteria; PenAsp = group of Penicillium spp./Aspergillus spp./Paecilomyces variotii; Unifung = total fungal DNA). Mean represents the average of triplicate samples; SD represents standard deviation in the triplicate assessment. Results are expressed as cells per sample [REPS-sampler] and cells per two Petri dishes.

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Chart 1. Microbial levels in settled dust samples in three locations and three sampling durations (2, 4, 8 weeks). Columns represent means of triplicate samples. For Petri dish, a sample is two Petri dishes and for REPS, a sample is one REPS. (Note: In EXP 2 environment the detected microbial levels for Gram- positive and Gram-negative bacteria from 2 week period by REPS sampling is noticeably high. This is likely a false result caused by contamination and thus should be considered with caution).

5.2 Active air sampling compared to sampling with REPS and Petri dishes

In order to compare active air sampling methods applied in this study (Impaction and Button Inhalable sampling) with the REPS and Petri dish samplers, we applied active air sampling campaign parallel to settled dust sampling. In table 3 an increasing trend of microbial levels can be seen when active air sampling is performed. The microbial levels increase simultaneously with extended sampling periods with all microbial levels and sample types.

When comparing the microbial levels detected with REPS compared to other sampling methods, it is important to notice the following: REPS fails to collect any microbial groups in EXP 1 sampling environment. In EXP 3 sampling environment, REPS performs better than in EXP 1 sampling

environment. In EXP 3 for Unifung assay and Gram-negative bacteria, the microbial levels increase as the sampling times are extended. Even though the microbial levels increase over extended sampling periods, the microbial levels detected with active air sampling and Petri dish are 100-1000 –fold higher. For Gram- positive bacteria, the REPS fails to collect efficiently in EXP 3 sampling environment. (Chart 2)

The settled dust sampling applications yield different microbial levels compared with Button Inhalable samples in all three sampling locations. The same trend is seen for all the microbial groups: REPS fails to collect microbes as efficiently as the Button Inhalable samples or Petri dishes do, since the microbial levels detected with Petri dishes and Button Inhalable samples are approximately 10-1000 –folds higher than the levels detected with REPS. (Chart 3)

Table 3. Microbial levels in PM2.5 and PM10 impaction and Button Inhalable Aerosol samples in different sampling locations (EXP1-3) and from three sampling periods (2,4,8 weeks), determined with qPCR assays (Grampos = Gram-positive bacteria; Gramneg = Gram-negative bacteria; PenAsp = group of

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Penicillium spp./Aspergillus spp./Paecilomyces variotii; Unifung = total fungal DNA). Results are expressed as cells per sample collected over a given sampling period.

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Chart 2. Microbial levels (Gram-positive bacteria, Gram-negative bacteria, Total fungal DNA) in PM2.5 and PM10 filter samples as well as in Petri dish and REPS settled dust in two locations (EXP1, office and EXP3, outdoor) and over three sampling durations (2, 4, 8 weeks). Cells per sample in the case of REPS refers to cells per one REPS, for Petri dish it refers to cells per two Petri dishes and for PM2.5 and PM10 samples it means cells per filter. (Note: non-detect results were replaced with value “1” to allow presentation in these log-scale graphs.)

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Chart 3. Microbial levels in each sampling location at timepoint 2 weeks. Cells per sample refers to cells per one REPS; cells per two Petri dishes; and for Button Inhalable samples cells per m³. (Note: non-detect results were replaced with value “1” to allow presentation in these log-scale graphs.)

5.3 Correlation between active and passive sampling methods

Finally, we performed correlation analysis between the microbial levels in actively collected air (PM2.5

and PM10) and passively collected settled dust samples (Petri dish and REPS). We did correlation using datapoints from two sampling locations at three timepoints (2, 4, 8 weeks) and all available different qPCR assays, totaling 21-24 datapoints. We found significant and strong correlations between microbial levels detected with qPCR between PM2.5 and PM10 samples, Petri dish and PM2.5 samples, REPS and PM2.5

samples, and Petri dish and PM10 samples (Table 4). Also, a good correlation was found between

microbial levels detected with qPCR between REPS and PM10 samples, and REPS and Petri dish samples.

All the measured correlations were significant (p-value <0.050).

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Table 4. Spearman rank -correlation matrix between the sampling methods applied in this study (n=21- 24). Datapoints are derived from two sampling locations (EXP 1, EXP 3), three timepoints (2, 4, 8 weeks) and 3-4 qPCRs (Gram-positive from EXP 3 was excluded). Significant correlation (p-value <0.050) is marked with **.

5.4 Practicality of the REPS –sampler compared with other sampling approaches

In addition to comparing microbial levels detected with different sampling approaches applied in this study, we also evaluated the practicality of these sampling approaches during field and laboratory work.

An overview of this assessment is provided in table 5.

Table 5. Comparison of three different sampling approaches applied in this study, evaluating the practicality during field and laboratory work.

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6 Discussion

6.1 REPS compared with Petri dish settled dust sampling

When comparing the performance of microbial determinations from settled dust samples collected with the REPS and with Petri dishes, one of the key observations was that REPS was inferior to Petri dish sampler with respect to collecting detectable microbial levels in low exposure office environment.

During two and four week sampling periods, the majority of qPCR assays – targeting Gram-positive bacteria, Gram-negative bacteria, total fungal DNA and Penicillium/Aspergillus group - returned non- detectable results from REPS, while Petri dish samples did return clearly detectable values. During the eight week sampling period in the office environment, some qPCR assays still did not detect signal from REPS samples, and approximately 8 -fold higher levels of Penasp group was detected with Petri dish than with the REPS-sampler. In the residential home environments we noticed a similar trend in that Petri dish samples yielded higher levels for almost all the sampling periods and qPCR assays, compared to REPS.

One exception for this can be seen in the residential home environment in two week –sampling period.

Here, the Gram-positive (and in part also Gram-negative) bacteria seemed to be collected more

efficiently by the REPS sampler than the Petri dish sampler as the REPS sampler yielded approximately 1.5 -times higher concentrations for Gram-positive bacteria than the Petri dish sampler. This result , however, is likely an artefact. The high values in Gram-positive and Gram-negative bacterial levels in this timepoint are driven by an oddly high result from one (out of the triplicate) REPS sampler, whereas the other two REPS samplers gave a very low result. We have to assume that this one REPS sampler was actually contaminated at some point during the laboratory analysis or the field work, resulting in an artificially high value from the bacterial measurements. An observation supporting this hypothesis is also that we noted that the concentrations for Gram-positive and Gram-negative bacteria were again

decreasing with prolonged sampling period instead of increasing after the two week sample.

When it comes to the Petri dish samples, we observed that the levels of Gram-positive bacteria increased when sampling periods were prolonged from two weeks to four weeks in the residential home

environment. For eight week –sampling period, the concentrations decreased, which can be a result of resuspension of settled dust from the samplers, as explained by Frankel et al. 2012. On the other hand, for other qPCR –assays in Petri dish sampling the detected levels from each assay actually increased as

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the sampling periods were prolonged in the home sampling environments. Similarly, this was the case in Petri dish sampling in the low exposure office environment for all microbial groups, with the exception of Pen/Asp fungal group. This trend is not seen with the REPS-samples in residential home environment.

Generally, for the two indoor environments our assessment is that i) Petri dish sampling resulted in higher microbial yields compared to REPS; and ii) that we observed an increase of microbial levels with prolonged sampling durations for Petri dish sampling for most microbial groups. These increases were, however, not consistent for all microbial groups measured and in no case were they linear. The latter may be due to variable microbial levels in the air, or/and saturation effects on the Petri dishes. Based on our observations, a two week sampling duration could be considered preferable for settled dust

sampling with Petri dishes, obviously depending also on the study question. Additional Petri dishes (for example 4 petri dishes rather than two) to increase sample amounts yielded from low exposure

environments could be opted for. As for REPS, the low microbial levels yielded from indoor sampling environments represent a challenge for the applicability of this approach to indoor microbial exposure assessment, in particular in sampling environments with expected low exposure levels.

Our observations in outdoor locations were somewhat different: while REPS still collected considerably lower levels of microbes, it performed well in that increases in microbial levels were observed with increasing sampling duration. For Petri dish sampling, microbial levels decreased in all cases from two to eight week samples. This strongly points towards the susceptibility of this sampling approach to strong air currents resulting in resuspension of collected particles, as certainly observed outdoors, but maybe also in some semi-ideal indoor locations, eg. close to doors, windows or ventilation in- and outlets. Thus, when using Petri dish sampling, care needs to be taken with respect to selecting largely undisturbed sampling locations. Moreover, detection of Gram-positive bacteria from Petri-dish dust with the qPCR assay we used (Kärkkäinen et al. 2010) was problematic in our study specifically from the outdoor environment, confirming observations made by the research group in earlier studies. The leading hypothesis to explain this is that particularly high levels of bacteria in settled dust might cause issues with the qPCR assay, more specifically with the Gram-positive bacterial probe

Alternatively, a phenomenon referred to as “open air factor” (OAF) may offer an explanation for the low levels of Gram-positive bacteria detected in settled dust samples from long sampling periods in outdoor sampling environments. The “open air factor” first described by May et al. (1969) has been explained recently by Hobday et al. (2019). Both this earlier study as well as the study by Hood et al. (2009)

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suggested that the “open air factor” killed certain bacterial species and preferentially Gram-positive ones, such as Staphylococcus epidermidis. They also noticed some reductions in Gram-negative bacteria species –levels, caused by OAF. Bailey et al. (2007) compared different factors and how well they reduced a specific bacterial species levels, and noticed that the OAF reduced the levels of aerosolized Micrococcus luteus, another Gram-positive bacterial species, more effectively than ozone treatment alone.

Ultimately, based on our results REPS could be a good method when performing outdoor microbial sampling over longer time periods. If using Petri dishes, shorter sampling periods of maximum two weeks should be used, and care needs to be taken when interpreting total microbial levels from such sample material. The REPS sampler, however, failed to collect as high concentrations of microbes as the Petri dish sampler did during this trial. Therefore, in sampling environments where low microbial exposure is expected, the REPS-sampler may not perform microbial sampling as efficiently as the Petri dish sampler.

6.2 Settled dust sampling compared with active air sampling

The microbial levels detected from samples collected with active air sampling techniques – PM2.5 and PM10 Harvard impactors and Button Inhalable Aerosol Samplers - behaved as expected within different sampling periods and different sampling environments. The collected microbial levels were lowest in office environment, second highest in home residence (PM2.5 and PM10 -samples were not collected from the home residence), and highest in outdoor location. Also, all the collected active air samples from PM2.5

and PM10 impactors followed the expected microbial levels when it comes to the sampling periods:

microbial levels in each sampling location were lowest in the two week samples and highest in the eight week -samples.

Active air samples were collected also with Button Inhalable samplers, but only for two week-sampling periods in each location. Microbial levels were lower compared to the PM10 samples collected with Harvard impactors, likely at least in part relating to the higher flow rates in Harvard impaction

(~10L/min) compared to Button sampler (~4L/Min). The Button Inhalable Aerosol samples will be used in further studies applying next generation sequencing techniques for bacterial and fungal community characterization, comparing to settled dust samples, but this more time consuming and expensive analysis was not included in this thesis.

Rutgers Electrostatic Passive Sampler (REPS) In Microbial Exposure Assessment