A Drosophila Model to Study Wound-induced Polyploidization

1. Staging and wounding of adult fruit flies

1. Select D. melanogaster strain of choice (i.e., epi-Gal4/ UAS strain, see Table of Materials).
   NOTE: Here, the Gal4/UAS system is used to enable epithelial specific gene expression (epi-Gal4) of a gene or RNAi encoded downstream of UAS. This study uses fluorescent membrane protein (UAS-Cd8.mRFP), mitotic inducer (UAS-fzr^RNAi, UAS-stg), and WIP inhibitor (UAS-E2F1^RNAi; UAS-Rac^DN).
2. Collect two vials of 10-15 newly enclosed female fruit flies each and age on fresh food vials at 25 °C until 3-5 days old. One vial will serve as the uninjured control and the other vial will be wounded as described below. The female flies should be maintained with males (~5/vial).
3. To wound the flies, assemble several pin holders each with a single 0.10 mm stainless steel pin. Ensure the sharp end of the pin is facing out. Pins can easily bend or chip after puncturing the fly and hooked or damaged pins should be discarded.
4. Anesthetize aged female fruit flies on a CO₂-fly pad under a stereomicroscope and align them into a row using a paint brush. Wearing safety glasses, holding the pin holder in one hand and forceps in the other, use forceps to position a fly with its ventral abdomen facing up.
5. Puncture the adult female flies within the epithelial pleurite region of tergite A4 on either side of the ventral midline sternites (Figure 1A). Puncturing this ventral region provides optimal space away from the dissection sites where tissue edges will be torn by mechanical processing.
6. Return wounded flies to the food vial and age to desired day post injury (dpi). Epithelial wound healing starts at 1 dpi and ends by 3 dpi. Endoreplication peaks at 2 dpi, which is ideal for the EdU assay (section 4, Figure 2).

2. Fly abdominal dissection

NOTE: During this step, it is important to avoid touching the ventral abdominal tissue with the dissection tools because it will compromise the integrity of the epithelium.

1. Obtain all required materials for dissection: Grace's solution, forceps, Vanna's Spring Scissors, 0.10 mm pins, dissecting plates, 9 well glass dissection dish, fixative solution (4% paraformaldehyde in 1x PBS), 1x PBS, wipes, pipettes and tips for 30 µm, and gloves (see Table of Materials).
2. Confirm that flies were successfully injured by anesthetizing wounded flies on a CO₂-fly pad under a stereomicroscope and checking for the presence of the wound scar (i.e., a melanin spot on abdomen, see Figure 1B). Discard any flies from the experimental group that were not successfully wounded.
3. To start dissection, fill one well of a 9 well glass dissection dish with Grace's solution. Use a pair of forceps to grasp a wounded female fly by the dorsal side of the thorax and submerge the fly in the well containing Grace's solution.
4. Using forceps in the opposite hand without releasing the thorax, puncture the dorsal cuticle below tergite A6 and pull the cuticle off the rear end of the fruit fly. The internal organs (ovary and intestine) will usually come out at this step. If not, gently push on the dorsal side of the abdomen with the forceps to squeeze out the remaining organs and discard into an empty well.
5. Snap off the full abdomen at the thorax junction above tergite A2 using the forceps and transfer the abdomen to an empty well containing ~100 µL of Grace's solution.
6. Repeat steps 2.3-2.5 until all fly abdomens are dissected.
7. Reduce the volume of Grace's solution to 30 µL in the well containing pooled, dissected abdomens.
8. Fillet the abdomens open by positioning the abdomen on the dorsal side with the forceps in one hand and then inserting the bottom blade of Vanna's spring scissors into the abdominal cavity with the other hand. Cut along the dorsal midline until the abdomen is fully opened, which can require up to three cuts (Figure 1C, 1D).
9. Set up a dry dissecting plate with four 0.10 mm pins per abdominal mounting area. Each 35 mm dissecting plate can fit up to seven mounting areas. Pipette 30 µL of Grace's solution on each mounting area and transfer one filleted abdomen to each droplet.
10. Pin the filleted abdomens to the dish on the four dorsal corners (Figure 1E). Ensure that the tissue lies flat without tearing or overstretching the abdominal tissue.
11. To fix the tissue, pipette off the Grace's solution and add 30 µL of the fix solution to the pinned abdomen.
    CAUTION: Wear gloves while handling the fix solution, as paraformaldehyde is toxic.
12. Repeat steps 2.10-2.11 until all filleted abdomens are pinned onto the dissecting plate.
13. Place a tape label on the bottom of each dish to mark each control and experimental group. Fix samples for 30-60 min at room temperature (RT).
14. Wash off fix solution by pipetting on 1.5 mL of 1x PBS to each plate. Dispose of fix solution and plastics in appropriate liquid or dry chemical waste containers according to institutional guidelines.
15. Wash plates 2x with 1.5 mL of 1x PBS and store fixed tissue covered in 1.5 mL of 1x PBS in a plastic container with a lid. Add a layer of damp paper towel to the bottom of the container and store samples at 4 °C until ready to immunostain within 1 week of dissection.

3. Immunofluorescence

1. Freshly prepare reagents (see Table of Materials): wash buffer solution (0.3% Triton X 100, 0.3% BSA in 1x PBS). Leftover wash buffer can be saved at 4 °C and used for the duration of the 2 day staining protocol. Prepare enough primary antibody solution per assay (Figure 2) using Anti-FasIII (1:50 mouse anti-Fasciclin-III) in wash buffer with either anti-Grh (1:300 affinity purified rabbit anti-Grainyhead⁸) or anti-RFP (1:1,000 rabbit anti-RFP). Primary antibody solutions can be saved at 4 °C and reused multiple times until signal is significantly reduced.
2. Permeabilize tissue by pipetting off 1x PBS, adding 1.5 mL of wash buffer, and incubating for at least 30 min on an orbital shaker (80 rpm) at RT.
3. Remove wash buffer and stain tissue overnight with 1.5 mL of primary antibody solution, incubating on an orbital shaker (80 rpm) at 4 °C. Collect the primary antibody solution and save in a tube at 4 °C for future experiments.
4. First rinse the sample quickly with 1x PBS and then wash 3x with 1.5 mL of wash buffer. For each wash, incubate samples at RT on an orbital shaker for at least 30 min.
5. During the final wash, prepare the secondary antibody solution: 1:1,000 donkey anti-rabbit Alexa 488 or 568 and 1:1,000 goat anti-mouse Alexa 488 or 568 (or fluorophores of choice) in wash buffer.
6. Remove wash buffer and stain tissues with 1.5 mL of secondary antibody solution. Cover samples with aluminum foil and incubate on an orbital shaker at RT for 3 h. Alternatively, samples can be incubated overnight at 4 °C on an orbital shaker.
7. Wash samples by first discarding secondary antibody solution and then rinse sample quickly with 1x PBS followed by three washes with 1.5 mL of wash buffer. For each wash, incubate samples at RT on an orbital shaker for at least 30 min.
8. Prepare DAPI solution by diluting DAPI to 10 µg/mL in wash buffer. After the final wash, stain samples with 1.5 mL DAPI solution incubating at RT for 30 min.
9. Discard DAPI solution and rinse samples 2x in 1.5 mL of 1x PBS. Store stained tissue in 1.5 mL of 1x PBS in the dark, covered with aluminum foil, at 4 °C until ready to mount on a glass slide with coverslip. The mounting step should be performed within 1 week.

4. Cell cycle activity (EdU Assay)

1. Make up a 10 mM EdU stock solution from Click-iT kit (see Table of Materials) by dissolving EdU powder in dH₂0 and mixing for ~15 min until completely dissolved. The stock solution can be aliquoted (250 µL per tube) and stored at -80 °C.
2. Feed flies EdU by first diluting EdU stock to 5 mM in dH₂O. Add dry yeast until the solution is cloudy and briefly vortex to mix. Cut off a 0.5 mL tube cap and place it at the bottom of the fly food vial. Push the cap into the food, so it is stable.
3. Anesthetize the flies and transfer 3-5 day old flies into the vial. Tap the flies to one edge so none are stuck in the cap.
4. Pipette 75 µL of yeast-EdU solution into the cap. Flies should be fed fresh yeast-EdU solution every day and transferred to a fresh food vial with a cap every other day to ensure that the flies do not get stuck in the bottom of the vial.
5. To transfer flies, flip to a new vial with a cap, put flies to sleep, tap flies to one side, and add fresh yeast-EdU solution.
6. On the third day, injure the flies and continue to feed yeast-EdU until dissection at 2 dpi (Figure 4A). See protocol section 2 for dissection and fixation methods.
7. Prepare EdU staining reagents: wash buffer (0.3% Triton X 100, 0.3% BSA in 1x PBS), permeabilization buffer (0.5% Triton X 100 in 1x PBS), blocking buffer (3% BSA in 1x PBS), and prepare reagents from the EdU assay kit (see Table of Materials), including 1x reaction buffer and 1x reaction buffer additive as directed by the manufacturer.
8. Wash samples for 1 h, mixing at RT in 1.5 mL of wash buffer.
9. Add 1.5 mL of permeabilization buffer and incubate samples for 20 min.
   NOTE: Thaw and prepare reaction cocktail solution using a volume of 500 µL/plate.
10. Wash samples 1x quickly with 1x PBS and then 3x quickly with 1 mL of blocking buffer.
11. Pipette off all remaining blocking buffer and add 500 µL of reaction cocktail solution per plate. Swirl plates to ensure tissues are completely covered. Incubate in a drawer in the dark for 1 h at RT.
12. Wash samples 1x quickly with 1.5 mL of blocking buffer.
13. Stain with 1.5 mL of DAPI solution at 1:5,000 in wash buffer for 30 min.
14. Wash 2x with 1x PBS quickly, wrap in foil, and store in the dark at 4 °C until ready to mount samples within 3 days.

5. Mount stained tissue

1. Obtain all required materials for mounting: glass slides, glass coverslips, clear nail polish, mounting media, a pair of forceps, and wipes.
2. To mount stained fly tissue, unpin abdomens from the dissecting plate using forceps under the stereomicroscope. Transfer tissue to ~30 µL of mounting media on a glass coverslip by gently grabbing the tissue with forceps by its dorsal flanks, taking care to avoid touching the ventral area with forceps.
3. Under the stereomicroscope orient the abdominal tissue so that the inside is facing down toward the coverslip (i.e., the external cuticle/bristles are facing up). Pull the oriented abdomens to the edge of the media droplet using the forceps. Surface tension will help keep the tissue flat (Figure 1F).
   NOTE: It is helpful for imaging to organize the abdomens in a column or row at this stage.
4. Label a glass slide (i.e., control or experimental) and pick up the coverslip by slowly bringing the slide closer to the coverslip. Flip slide over and gently blot with a wipe to remove excess mounting media.
5. Seal the edges of the coverslip with clear nail polish and repeat for all remaining experimental groups. Store slides in a slide box at 4 °C until ready to image.

6. Imaging and processing

1. Image the fly abdominal wound area by first locating the melanin scar with a confocal microscope (Figure 1B), either a point scanner or a structured illumination (ApoTome) with a 40x oil or dry objective.
2. Check the exposure on each channel, ensuring that the signal is below saturation. The imaging settings should be based on the brightest sample group. This is particularly important for ploidy analysis as the DAPI channel needs to stay in the linear range to accurately measure DNA content.
3. Take a full z-stack image in all three channels with an optimal distance of at least 0.50 µm between slices. Save captured images and open the file in the image analysis program Fiji (also known as ImageJ).
4. For each image, create a z-stack projection using the sum of slices option for all channels.
5. Rotate the images as necessary to ensure the nuclei are lined up horizontally across images (Figure 3A and Figure 4E).
6. Crop all the images to a rectangular selection of 300 µm x 300 µm centered around the wound site or center of uninjured control. Identify the area by drawing a rectangle and selecting Edit | Selection | Specify. Check that scaled units are in microns to ensure the same sized box is used for all images to be analyzed.

7. Endoreplication (ploidy) analysis

1. Using Fiji, select the Grh channel window and duplicate the image. Then use the threshold tool to create a mask. Manually adjust the threshold by sliding the top bar to minimize background without causing the nuclei to shrink drastically (Figure 4D).
2. If any nuclei in the Grh channel are touching in the threshold image, use the paintbrush tool (2 pixel width) in the same color as the background, to draw a line between the nuclei. Click to apply 1x when finished to generate the final mask.
3. Generate a region of interest (ROI) map by using the Analyze Particles function: set size to 5 µm-60 µm in order to capture most of the nuclei without including the background.
4. Manually adjust the ROI map as needed in the ROI manager. Delete any selections that are not nuclei and add any nuclei to the list that were not identified by outlining the nucleus with the freehand selection tool and adding it to the ROI manager (Figure 4D).
5. Select the DAPI channel then click Show All in the ROI manager to apply the generated ROI map in the Grh channel onto the DAPI channel.
6. Delete any selections where the epithelial nuclei outline overlaps with nonepithelial nuclei (e.g., nuclei from muscle or fat) from the ROI map. Grainyhead only stains epithelial nuclei, whereas DAPI stains all nuclei. Check that each outlined selection contains only one nucleus and delete or edit any selections with more than one nucleus. Save edited ROI list.
7. Measure the area and integrated density of each epithelial nuclei in the ROI map using the analysis tools in Fiji. Export the values into a spreadsheet program.
8. Measure the average image background using the circular selection tool. Draw three circles that do not overlap with any nuclei in different areas of the DAPI image. Add the area and integrated density of each of the circles to a spreadsheet program to establish the background image brightness.
9. Start by calculating the average background per unit area for each image by dividing each background integrated density value by its corresponding area. Then average the three integrated density per area measurements for the image in order to obtain the average background per unit area.
10. Next, calculate the total background of each DAPI nucleus by multiplying the area of the nucleus by the average background per unit area. The normalized DAPI intensity for each nucleus measured can then be calculated by subtracting the total background of each nucleus from its measured integrated density.
11. Average all the normalized DAPI intensity values from the uninjured epithelial control. The uninjured epithelial nuclei were previously calculated to have a ploidy value of 2C and can serve as a reference for calculating ploidy in the epithelial nuclei from the experimental conditions⁸.
12. Calculate the ploidy of each nucleus by dividing the normalized DAPI intensity of each nucleus by the normalized value from reference uninjured epithelial control (2C), then multiply the value by 2 to equal the normalized ploidy (C-value):
    (Nuclear Integrated Density – Background Nuclear Integrated Density)/Average Nuclear Integrated Density (uninjured epithelial nuclei = 2C) x 2 = epithelial nuclear ploidy (C)
13. Graph nuclei with ploidy values as a dot plot, histogram, or grouped into a bar graph accordingly (i.e., 2C [0.6-2.9C], 4C [3.0-5.9C], 8C [6.0-12.9C], 16C [13.0-24.9C], and >32C [>25.0C] (Figure 3F).