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

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.

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.

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.