Analysis of the Ambient Particulate Matter-induced Chromosomal Aberrations Using an In Vitro System

1. Particle Collection and Preparation

1. Calibration of the high volume total suspended particles sampler
   1. Disconnect the sampler motor from the mass flow controller and connect the motor to a stable AC power source (Figure 1).
      NOTE: A filter is not used during this procedure.
   2. Mount the calibrator orifice and top loading adapter plate to the sampler. Tighten the top loading adapter hold-down nuts securely to ensure that no air leaks are present. Allow the sampler motor to warm up to its normal operating temperature (approximately 10-15 min).
   3. Conduct a leak test by covering the holes on top of the orifice and pressure tap on the orifice with hands. Listen for a high-pitched squealing sound made by escaping air. If this sound is heard, re-tighten the top loading adapter hold-down nuts as a leak is present. If the sound is lower, the leak is near one of the other gaskets in the system.
   4. Connect one side of a water manometer to the pressure tap on the side of the orifice with a rubber vacuum tube. Leave the opposite side of the manometer open to the atmosphere. Hold a manometer vertically to ensure accurate readings. Tapping the backside of the continuous flow recorder will help to center the pen and provide accurate readings.
   5. Repeat step 1.1.2 using five flow rates representing operating flow rates from 30 to 60 ft/min (approximately every 5-6 ft³/min). Adjust the knob on the variable orifice to five different positions and take five different readings.
   6. Record the ambient air temperature, ambient barometric pressure, sampler serial number, orifice serial number, orifice slope and intercept with date last certified, the date, site location, and operator's initials on the calibration sheet.
2. Total suspended particles sample collection and gravimetric analysis
   1. Wrap an 8" x 10" quartz fiber filter in two layers of aluminum foil. Place the wrapped filter in the furnace and set the temperature at 550 °C. Bake the filter for at least 4 hr. Let the furnace cool and retrieve the filter. Place the filter in the desiccator with CaCO₃.
   2. Weigh the filter in a microbalance with precision of 10 µg. Repeat the measurement three times during a day. Individual measurements should be within 25 µg. If this is not achieved, return the wrapped filter to the desiccator and repeat the process the following day.
   3. Open the shelter lid (Figure 2). Remove filter holder frame by loosening the four wing nuts, allowing the brass bolts and washers to swing down out of the way. Unfold the filter from the aluminum foil, and use cleaned forceps to carefully center it, rougher side up, on the supporting screen. Properly align the filter on the screen.
   4. Place the frame on the screen and secure it with the brass bolts and washers while applying sufficient pressure to avoid air leakage at the edges. Wipe any dust accumulation from around the filter holder with a clean cloth. Close shelter lid carefully and secure it.
   5. Make sure all cords are plugged into their appropriate receptacle sockets and the tubing between the blower motor pressure tap and the continuous flow recorder is connected.
   6. Prepare the flow recorder. Depress pen arm lifter to raise pen point and insert a new chart. Align the tab of the chart to the drive hub of the recorder and press with thumb to lower chart center onto hub. Set time by rotating the drive hub clock-wise until the correct time on chart is aligned with time index pointer.
   7. Manually switch on the sampler and start the collection. Monitor the recorder to be sure it is inking correctly and the elapsed time indicator to be sure it is working properly.
   8. At the end of the sampling period, switch off the sampler, record the elapsed time and retrieve the chart. Remove the frame to expose the filter. Remove the exposed filter from the supporting screen by holding it gently at the ends with the forceps. Fold the filter lengthwise so that sample touches sample twice and wrap it in the original aluminum foil.
   9. Store the filter in a desiccator with CaCO₃ for at least 24 hr prior to weighing. Repeat step 1.2.2 to measure the weight of the filters after sampling. Place the filter in a zip plastic bag and store it in -80 °C until analysis.
3. Analysis of collected total suspended particles sample
   1. Retrieve the stored filter, unfold the aluminum foil and transfer the filter on a clean Teflon surface. Using forceps and scalpel, cut outer (white) part of the filter and dispose.
      1. Next, cut the filter in 0.5" x 0.5" pieces and place pieces in a 100 ml flask. Add 50 ml of ultrapure water. Place the flask in an ultrasound bath and sonicate for 60 min at 33±3 kHz.
   2. Filter extract through a 0.45 µm polypropylene syringe filter to a 100 ml round bottom flask. Evaporate the water using a rotavap instrument under pressure at 50 °C until about 5 ml of extract remains in the flask.
      1. Evaporate the remaining water in a 15 ml pear-shaped flask of known weight. Weigh the flask with the dry extract and record the mass of the dry extract.
   3. Dissolve the dry sample with 400 µl of D₂O by sonicating for 15 min and transfer into a 5 mm standard NMR tube. Centrifuge for 5 min at 12,000 x g at room temperature to remove any undissolved matter.
   4. Repeat with 300 µl of D₂O. Add 10 µl of NaN₃ 1.4% (final conc.: 0.02% w/v) in the NMR tube. Add 10 µl (32 µg) of total suspended particles (TSP)-d4 (3.2 mg/ml) to D₂O (final concentration 0.258 mM) in the NMR tube.
   5. Acquire ¹H NMR spectra using a NMR spectrometer operating at 600.17 MHz.
      1. Insert NMR tube into the triple resonance cryoprobe. Calibrate the instrument in the following order: lock, tune, shim, 90° pulse length calibration and receiver gain adjustment.
      2. Measure spectra at 25 °C with a 1 D excitation sculpting sequence using 180 water-selective pulses in Digital Quad detection mode, 8,192 scans, 4 dummy scans, sweep width of 8012.57 Hz, frequency offset for water signal suppression set at 4.70 ppm, 1.70 sec acquisition time, 32,768 time domain data points (TD), 1 msec duration gradient (p16), 100 µsec recovery delay after gradient (d16).
   6. Process ¹H NMR spectra by applying a line broadening of 1 Hz and zero filling by a factor of 2 (65,536 data points). Correct the phase and baseline manually and integrate. Set TSP-d4 peak at 0.0 ppm.

2. Particle Exposure

1. Culture set up
   1. Warm complete growth media containing 10% fetal bovine serum (FBS) and penicillin-streptomycin solution (100 U/ml), 0.25% trypsin solution and calcium magnesium free phosphate buffer saline (PBS) in a water bath (at 37 °C) for at least 30 min before culture initiation.
   2. Take out a T75 flask from 5% CO₂ incubator containing cultured RAW264.7 cells and wash the cells with 2 ml of pre-warmed PBS two times.
   3. Dislodge adherent RAW264.7 cells from the bottom of tissue culture vessel using 1 ml of pre-warmed trypsin solution followed by scraping.
   4. Collect the cells from tissue culture vessel and make a single cell suspension by repeated pipetting. Count the number of cells with trypan blue and plate at a density of 8,000/cm² (500,000 cells total) in a 100 mm tissue culture dish with pre-warmed complete media. Adjust media volume to 10 ml for a final concentration of 50,000 cells per ml.
      NOTE: Double the number of plates if DNA methylation analysis is also performed.
   5. Place the newly plated RAW264.7 cells back in 5% CO₂ incubator at 37 °C and allow to attach for 24 hr.
   6. Also in a biosafety cabinet, resuspend filtered particles. For example, dilute particles at a concentration of 0.5 mg/ml in sterile PBS for a 5 µg/ml treatment dose and at 5 mg/ml for a 50 µg/ml treatment dose. Dilutions can be prepared up to month in advance and stored at -80 °C.
2. Treatment with particulate matter
   1. Take out the cells from 5% CO₂ incubator and check under 10x magnification using an inverted microscope for cell growth, cell morphology, culture condition, and contamination. One should expect to have at least 30% confluent cells at this stage.
   2. Aseptically, aspirate the media using sterilized pipette and wash the culture vessel with 2 ml 37 °C PBS two times to completely remove the media and floating or dead cells.
   3. Add pre-determined dose of particulate matter with fresh media in the culture vessels and incubate for desired time at 37 °C in 5% CO₂ incubator. In this study, treat RAW264.7 cells with PM for 72 hr to parallel the exposure used for concurrent genotoxic and epigenotoxic analysis.
      NOTE: Cytotoxicity may negatively affect the quality of the cytogenetic analysis. Therefore, additional assays such as cytotoxicity, DNA methylation, Western blotting, or gene expression analysis are typically performed on treated cells. Expose cells as described and proceed with the specific assay desired. Inexperienced users may also want to include a positive control such as methyl methanesulphonate to test their ability to detect CAs.

3. Cytogenetic Assay

1. Treatment to arrest cells at metaphase
   1. Warm the colcemid solution (10 µg/ml) in a 37 °C water bath for 30 min. Wipe the lid with 70% alcohol under a biosafety cabinet to minimize the risk of contamination
   2. Completely remove the media aseptically and wash the culture vessels with 2 ml pre-warmed PBS two times.
   3. Add pre-warmed fresh media (2 ml media/10⁶ cells) containing colcemid solution (5 µl/ml) in the culture vessel and incubate for 2 hr.
2. Cell harvest
   1. Warm trypsin solution and PBS in a water bath maintained at 37 °C for 30 min. From this step onwards, there is no need to maintain aseptic condition.
   2. Completely remove media, wash the culture vessels with 2 ml 37 °C PBS for two times, add required volume of pre-warmed trypsin solution (usually 2 ml for 60 mm culture dish or T25 flask and 4 ml for 100 mm dish or T75 flask), and place the culture vessel back in 5% CO₂ incubator at 37 °C for 1 min.
   3. Take out the culture vessel, detach the cells using a scraper, and add 10 ml of PBS to stop trypsin action.
   4. Pipette up and down at least ten times to make single cell suspension and transfer cells in a 15 ml conical bottom centrifuge tube.
   5. Centrifuge cell suspension at 400 x g for 5 min in a swing out centrifuge at room temperature.
   6. Remove the supernatant, break the cell pellet by gentle tapping, add 10 ml of PBS, and again centrifuge for five min.
3. Hypotonic treatment and fixation
   1. Pre-warm 0.075 M potassium chloride solution in a 37 °C water bath at least for 30 min.
   2. Remove supernatant without disturbing the cell pellet and leave approximately 0.5 ml of PBS.
   3. Gently break the cell pellet and make single cell suspension using a P200 pipette.
   4. Add 4 ml of pre-warmed potassium chloride solution drop by drop with gentle shaking.
   5. Incubate the cells in hypotonic potassium chloride solution in a 37 °C water bath for 20 min.
   6. Make fresh acetomethanol solution by adding 3-parts methanol with 1-part glacial acetic acid. Keep this freshly prepared fixative at room temperature.
   7. After hypotonic treatment, add an equal volume (4 ml) of fixative in the tube and gently mix the solution by inverting the tube.
   8. Centrifuge at 400 x g for 5 min at room temperature, and remove the supernatant. At this stage cell pellet size increases, become whitish and more visible.
   9. Remove supernatant, add 4 ml fresh fixative and keep at room temperature for 30 min.
   10. Centrifuge at 400 x g for 5 min, remove supernatant, and give two more washes with 4 ml fixative.
4. Slide cleaning and metaphase spread preparation
   1. Immerse glass slides completely in 10% liquid detergent for 30 min.
   2. Rinse the slides thoroughly in cold running tap water for 30 min.
   3. Rinse slides three times with distilled water to completely remove detergent, immerse in distilled water, and store in the refrigerator for future use.
      NOTE: Cleaning slides before use facilitates uniformity of spreading and increases metaphase quality.
   4. Drop 10 µl cell suspension on chilled wet slide and allow it to air dry overnight at room temperature.
5. Staining and mounting
   1. Prepare 50 ml Giemsa staining solution by mixing one part of Giemsa stain solution with one part of PBS and pour in a Coplin jar.
   2. Immerse the slides in staining solution for 20 min.
   3. Rinse the slides in distilled water to remove excess stain and immediately dry the slides using a hair-dryer.
   4. Leave the slides at room temperature overnight.
   5. Apply a single drop of mounting medium on the slide, gently place a coverslip on the mounting medium, allow the medium to spread over the slide slowly, and remove the excess medium with a paper towel. Allow the mounted slide to dry for at least 1 hr prior to moving or viewing under a microscope to avoid movement of the coverslip.
      Caution: It is important to place the coverslip carefully without forming any bubbles during this step. The use of excessive heat to remove bubbles is not recommended.
6. Microscopic observation and aberration types
   1. Examine the structural CAs in metaphase spreads at 100X magnification using a good-quality bright-field microscope.
   2. Score at least 50 to 100 metaphase spreads for each treatment group for treatments that induce a large number of CAs. For smaller effect sizes, the analysis of up to 300 metaphase spreads is advised.