Comet Assay for DNA Damage Detection in Cells

1. Preparation of materials for the comet assay

1. Preparation of the microscope slides
   1. Pour 1% (w/v) normal melting point agarose [dissolved in double-distilled water (ddH₂O)] in a 50 mL tube and microwave to dissolve the agarose in the ddH₂O. Store at 37 °C to prevent solidification prior to coating slides. Should solidification occur, discard and prepare fresh.
   2. Pre-coat microscope slides by dipping the slides into the 50 mL tube containing 1% (w/v) normal melting point agarose.
   3. Wipe the back of the slides quickly after dipping the slides.
      NOTE: Failure to wipe the back of the slides properly will increase the background noise of the slides during the analysis step by microscope.
   4. Label the coated slide with a permanent marker on the bottom-right corner of the frosted section (Figure 1A). This shows which side of the slide is pre-coated.
   5. Allow the agarose to set and dry overnight at room temperature.
   6. Wrap the dried slides in tissue paper and store them in a box.

2. Preparation of samples

1. Cultured cells
   NOTE: Firstly, treat cells with the damaging agent(s) prior to starting the comet assay. Then, perform the following.
   1. If the cells are adherent, trypsinize the cells to release them from the cell culture flask or cell culture Petri dishes, at the appropriate confluency of cells. Neutralize trypsin by adding serum-containing media.
   2. Transfer the cells to a 50 mL tube, centrifuge (e.g., for HaCaTs, centrifuge at 300 x g for 5 min at room temperature), gently remove the supernatant, and add 1 mL of PBS to the cell pellet.
   3. Perform cell counting.
   4. Transfer 30,000 cells to a 1.5 mL microcentrifuge tube and centrifuge at 7,607 x g for 5 min at 4 °C.
   5. Gently remove the supernatant and store the cell pellet on ice in the dark prior to performing the comet assay.
      NOTE: Cells should regularly be tested for Mycoplasma contamination prior to performing the comet assay to prevent, among other effects, the formation of artefactual DNA damage and altered DNA damage response, as reported. Centrifugation conditions may be changed, as needed, depending on the cell type used.
2. Preparation of cultured cells for repair assay
   1. Culture cells in the cell culture flask or Petri dishes.
   2. Wash the cells with 1 mL of PBS twice prior to treating the cells with damaging agents (e.g., for BE-M17 cells, treat with 50 µM of H₂O₂ for 20 min) on ice to prevent the repair from occurring during the treatment.
   3. Wash the cells gently with 1 mL of PBS twice to remove any residual damaging agents.
   4. Re-introduce the cell culture medium and allow the cells to repair for varying durations (e.g., 0 min, 30 min, 2 h, 6 h, 24 h, and 30 h) in a humidified incubator (37 °C, 5% CO₂).
   5. At each time point, collect 30,000 cells in 10% dimethyl sulfoxide (DMSO)-containing cell culture medium and store them at -80 °C.
   6. Before performing the comet assay, thaw the cells quickly at 37 °C in a water bath and centrifuge them at 7,607 x g for 5 min at 4 °C.
   7. Remove the supernatant and store the cell pellet on ice prior to performing the assay (i.e., from step 3).

3. Cell lysis

NOTE: Carry out all the procedures on ice.

1. Use 12,000 cells or 2.5 µL of whole blood per gel.
2. Prepare 0.6% (w/v) low melting point agarose dissolved in PBS using a microwave, and place in a water bath at 37 °C to prevent it from solidifying.
3. Label the frosted end of the pre-coated slides with the investigator's name, date, and treatment information using a permanent marker or pencil.
4. Place a chilling plate on a flat bench and insert the two frozen cooling packs into the sliding drawer below the metal surface (as shown in Figure 2).
5. Place the slides on the chilling plate and allow the slides to pre-chill for 1-2 min before adding the 0.6% (w/v) low melting point agarose-containing cells.
   NOTE: Leaving the slides on the chilling plate for more than 1-2 min may cause condensation to form on the slide surface due to ambient humidity. This may make the low melting point agarose gels less stable on the slides.
6. Disperse the pellet (step 2.2.7) by vortexing. Ensure that all the supernatant has been removed from the pellet. Place the sample tubes (containing the pelleted cells) immediately back on ice.
   NOTE: When placing the sample-containing tubes in the centrifuge, put them with the hinge facing outward so that the pellet will be collected on this side of the tube. Sometimes it is difficult to see the pellet, and it is easy to dislodge it while removing the supernatant. Centrifuging with the tube lid in this orientation will enable one to know where the cell pellet will be.
7. Resuspend the cell pellet with 200 µL of 0.6% low melting point agarose (LMP agarose) and mix by pipetting up and down without creating bubbles. Next, quickly transfer 80 µL of LMP agarose-containing cells onto a chilled slide and quickly place a coverslip onto the gel.
8. Allow the gel to set on the chilling plate for 1-2 min.
9. Meanwhile, prepare a 500 mL working solution of lysis buffer (Table 1) and pour it into the lysis dish (Figure 3).
10. Once the gels have been set, remove the coverslips quickly by gently holding the slide between thumb and forefinger and sliding the coverslip off the gel.
11. Place the slides containing samples inside the slide carrier (all the black "dot" marks on slides should be facing in the same direction when they are placed in a carrier) (Figure 1B), and then place the slide carrier inside the lysis dish (Figure 3).
12. Close the lid of the lysis dish and keep the lysis dish in the fridge overnight at 4 °C or 30 min at room temperature, whichever best suits the operator's time schedule.

4. Electrophoresis

1. Carefully remove the slide carrier from the lysis dish. Take care not to disturb the gels.
2. Gently place the slide carrier in a washing dish pre-loaded with ice-cold ddH₂O and leave it for 30 min ensuring that slides are completely covered with ddH₂O.
3. Insert a frozen cooling pack inside the sliding drawer under the electrophoresis tank to maintain optimal buffer temperature.
4. Carefully add ice-cold electrophoresis working solution (Table 1) to the electrophoresis tank and transfer the slide carrier into the electrophoresis tank. Orientate the slides such that their clear parts with the cell-containing gels (i.e., NOT the frosted/labeling ends) point toward the cathode (red electrode).
5. Allow the slides to sit in the electrophoresis tank for 20 min so that the DNA relaxes and unwinds. Keep the power supply switched off during this step.
6. If needed, insert a new frozen cooling pack to maximize chilling.
7. Perform electrophoresis for 20 min at 1.19 V/cm, or whatever conditions have been optimized.
   NOTE: Optimization of electrophoresis running conditions and volume of buffer is recommended for every laboratory. Using only a single slide carrier during electrophoresis does not cause any effect of slides on the resistance of the electrophoresis buffer, and the authors did not see a significant effect in the voltage or current when the number of the slides changed.
8. Switch off the power supply, carefully remove the slide carrier from the electrophoresis tank, and allow it to drain on tissue paper for 30 s.
9. Place the slide carrier into the dish containing neutralization buffer (Table 1). Leave it for 20 min.
10. Remove the slide carrier from the neutralization dish, place it in the washing dish containing ice-cold ddH2O, and leave it for 20 min.
11. Remove the slide carrier from the water and allow the slides to dry in an incubator at 37 °C for 1 h, or at room temperature overnight, or do not dry, depending upon the operator's schedule.
    NOTE: If there is no drying in step 4.11, perform the staining step from 5.2.

5. Propidium iodide (PI) staining

1. Transfer the slide carrier to a washing dish containing ice-cold ddH₂O to rehydrate the slides and leave for 30 min.
2. Place the slide carrier into a staining dish containing 2.5 µg/mL propidium iodide solution.
   NOTE: Propidium iodide is light-sensitive, so handle it in a darkened area. It is also toxic.
3. Close the lid of the staining dish and incubate it for 20 min in the dark at room temperature.
4. Transfer the slide carrier to a separate dish, and wash it with ice-cold ddH₂O for 20 min.
5. Remove the slide carrier from the dish and dry it completely in the dark, either in a 37 °C incubator or at room temperature, depending upon the operator's schedule or preference.
6. Once the slides are fully dried, remove them from the slide carrier and store them in a slide box in the dark until ready for image analysis.
   NOTE: The slides will remain readable indefinitely and can be re-stained if necessary.

6. Comet scoring and data analysis

NOTE: The term "comet" derives from the images of damaged cells when viewed under a microscope after the assay has been performed (Figure 4). Under electrophoresis conditions, DNA in the undamaged cells largely does not migrate but remains in a spheroid termed as comet "head." However, the presence of strand breaks allows the cell's DNA to migrate out of the head and form a "tail," thus leading to an appearance like a comet (Figure 4). The more DNA in the tail, the more damage is present.

1. Turn on the fluorescence microscope with the PI (red) filter (λ = 536/617 nm) and the comet assay scoring software.
2. Add a drop of water using a Pasteur pipette to the gel and cover with a coverslip.
3. Place the slides into the fluorescence microscope and "score" the comets.
   NOTE: Scoring is a means by which the comets are assessed, to determine the amount of damage present in each comet. Broadly, this can be achieved by using two approaches, according to the user's chosen preference, either by eye (gauging the size of the comets on a scale of zero to four) or by using freely, or commercially available software30. Generally, both approaches assess the size of the comet tail, although a variety of comet-related endpoints can be determined. If using the software, click on the middle of the comet head and wait until the software detects the comet automatically, and then assesses the chosen endpoint (Figure 4).
4. Score 50 comets per gel and 100 comets per sample (i.e., each sample corresponding to different DNA damaging treatments, or their replicates).
5. Replicate the experiments (n = 2) or triplicate the experiments (n = 3).
   NOTE: If only n=2 replicate experiments are undertaken, statistical analysis cannot be performed, but if n=3, perform the D'agostino normality test. Most comet assay data does not pass a normality test. In this case, use a nonparametric test (Kruskal-Wallis test with Dunn's multiple comparisons test, and Mann-Whitney tests significance set at p < 0.05).

Table 1: Composition of reagents used in high throughput alkaline comet assay (HTP ACA). The stock and working concentrations of lysis, electrophoresis, neutralization, and staining buffers are shown.

Figure 1: Representative images of a comet assay slide and HTP rack (microscope slide carrier). (A) For correct orientation, the pre-coated face of the microscope slide is recognized by a black dot in the right-hand corner of a microscope slide. (B) The image of the HTP rack illustrates how the slides are kept in a tight vertical orientation, with tabs on the carrier to fix its orientation within the electrophoresis tank. Each carrier can accommodate up to 25 slides.

Figure 2: Representation of the chilling plate with sample slides and freezer packs in place.

Figure 3: The materials comprising the high-throughput comet electrophoresis system. HTP electrophoresis tank, HTP racks, and the dishes for lysis, wash, neutralization, and staining are shown.

Figure 4: Screenshot of representative comets taken during scoring. HaCaTs (A) without treatment and (B) treated with 1 J/cm2 UVB prior to performing HTP ACA. Most software packages can calculate a variety of comet endpoints, but the most common ones are the % tail DNA (preferred) or tail moment based upon these images (blue: start of the head, green: middle of the head, and purple: end of tail). The scale bar is 10 µm