Name: a drosophila model to study wound-induced polyploidization
Uploaded: 2020-06-09T13:30:00.000+00:00
Duration: 7 min 27 s
Description: Watch this Scientific Journal Video about A Drosophila Model to Study Wound-induced Polyploidization at JoVE.com

At Boston College, we would like to thank Dr. Eric Folker for use of his lab&#39;s camera and stereoscope microscope setup for imaging and Bret Judson at the Boston College Imaging Core for infrastructure and support. We would also like to thank the fly community resources: Bloomington Drosophila Stock Center (NIH P40OD018537), Vienna Drosophila Resource Center, and TRiP Center at Harvard Medical School (NIH/NIGMS R01-GM084947) for providing transgenic stocks used in this study. The mouse FasIII antibody were obtained from Developmental Studies Hybridoma Bank supported by NICHD of the NIH and maintained at The University of Iowa, Department of Biology, Iowa City, IA. Research reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under Award Number R35GM124691. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

1. Staging and wounding of adult fruit flies
<ol>
	<li>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-fzrRNAi, UAS-stg), and WIP inhibitor (UAS-E2F1RNAi; UAS-RacDN).</li>
	<li>Collect two vials of 10-15 newly enclosed female fruit flies each and age on fresh food vials at 25 &#176;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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
</ol>
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.
<ol>
	<li>Obtain all required materials for dissection: Grace&#39;s solution, forceps, Vanna&#39;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 &#181;m, and gloves (see Table of Materials).</li>
	<li>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.</li>
	<li>To start dissection, fill one well of a 9 well glass dissection dish with Grace&#39;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&#39;s solution.</li>
	<li>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.</li>
	<li>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 &#181;L of Grace&#39;s solution.</li>
	<li>Repeat steps 2.3-2.5 until all fly abdomens are dissected.</li>
	<li>Reduce the volume of Grace&#39;s solution to 30 &#181;L in the well containing pooled, dissected abdomens.</li>
	<li>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&#39;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).</li>
	<li>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 &#181;L of Grace&#39;s solution on each mounting area and transfer one filleted abdomen to each droplet.</li>
	<li>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.</li>
	<li>To fix the tissue, pipette off the Grace&#39;s solution and add 30 &#181;L of the fix solution to the pinned abdomen. 
	CAUTION: Wear gloves while handling the fix solution, as paraformaldehyde is toxic.</li>
	<li>Repeat steps 2.10-2.11 until all filleted abdomens are pinned onto the dissecting plate.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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 &#176;C until ready to immunostain within 1 week of dissection.</li>
</ol>
3. Immunofluorescence
<ol>
	<li>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 &#176;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 &#176;C and reused multiple times until signal is significantly reduced.</li>
	<li>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.</li>
	<li>Remove wash buffer and stain tissue overnight with 1.5 mL of primary antibody solution, incubating on an orbital shaker (80 rpm) at 4 &#176;C. Collect the primary antibody solution and save in a tube at 4 &#176;C for future experiments.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#176;C on an orbital shaker.</li>
	<li>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.</li>
	<li>Prepare DAPI solution by diluting DAPI to 10 &#181;g/mL in wash buffer. After the final wash, stain samples with 1.5 mL DAPI solution incubating at RT for 30 min.</li>
	<li>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 &#176;C until ready to mount on a glass slide with coverslip. The mounting step should be performed within 1 week.</li>
</ol>
4. Cell cycle activity (EdU Assay)
<ol>
	<li>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 &#181;L per tube) and stored at -80 &#176;C.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Pipette 75 &#181;L 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Wash samples for 1 h, mixing at RT in 1.5 mL of wash buffer.</li>
	<li>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 &#181;L/plate.</li>
	<li>Wash samples 1x quickly with 1x PBS and then 3x quickly with 1 mL of blocking buffer.</li>
	<li>Pipette off all remaining blocking buffer and add 500 &#181;L 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.</li>
	<li>Wash samples 1x quickly with 1.5 mL of blocking buffer.</li>
	<li>Stain with 1.5 mL of DAPI solution at 1:5,000 in wash buffer for 30 min.</li>
	<li>Wash 2x with 1x PBS quickly, wrap in foil, and store in the dark at 4 &#176;C until ready to mount samples within 3 days.</li>
</ol>
5. Mount stained tissue
<ol>
	<li>Obtain all required materials for mounting: glass slides, glass coverslips, clear nail polish, mounting media, a pair of forceps, and wipes.</li>
	<li>To mount stained fly tissue, unpin abdomens from the dissecting plate using forceps under the stereomicroscope. Transfer tissue to ~30 &#181;L 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#176;C until ready to image.</li>
</ol>
6. Imaging and processing
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Take a full z-stack image in all three channels with an optimal distance of at least 0.50 &#181;m between slices. Save captured images and open the file in the image analysis program Fiji (also known as ImageJ).</li>
	<li>For each image, create a z-stack projection using the sum of slices option for all channels.</li>
	<li>Rotate the images as necessary to ensure the nuclei are lined up horizontally across images (Figure 3A and Figure 4E).</li>
	<li>Crop all the images to a rectangular selection of 300 &#181;m x 300 &#181;m 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.</li>
</ol>
7. Endoreplication (ploidy) analysis
<ol>
	<li>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).</li>
	<li>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.</li>
	<li>Generate a region of interest (ROI) map by using the Analyze Particles function: set size to 5 &#181;m-60 &#181;m in order to capture most of the nuclei without including the background.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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)</li>
	<li>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 &#62;32C [&#62;25.0C] (Figure 3F).</li>
</ol>

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Presented is a detailed protocol on how to dissect and use the adult D. melanogaster abdominal epithelium to study how genes regulate WIP by altering re-epithelialization and endoreplication during wound repair16. Using this method, the proto-oncogene Myc was recently identified as a key regulator of WIP. Myc is required for epithelial cells to endoreplicate post injury and is sufficient for quiescent epithelial cells to endocycle both in adult fly epithelium and accessory glands16,22. It was also found that switching epithelial cells to a mitotic cell cycle by expression of stg, fzrRNAi is detrimental to wound repair. Continued studies using this method will identify other genes required to regulate re-epithelialization and endoreplication during WIP, revealing both similarities and differences to how polyploidy is regulated and functions in a variety of tissues.
This model and method offer unique advantages, including the easy induction of polyploidy with a mechanical puncture and the fact that polyploid cells are generated within days4. The tissue dissection and preparation protocols are based on larval dissection techniques23, but the adult fly abdomen is more rigid and therefore easily perturbed. As a result, this protocol requires practice and precision to isolate an intact tissue to study WIP. Once dissected, however, the epithelium is clearly visible and easily imaged, yielding a snapshot of the wound healing process. This method provides a wealth of information on the adult fly&#39;s epithelial organization, cell and syncytium size, and the ploidy of cells and individual nuclei. While live imaging is not yet possible within the intact fruit fly due to its opaque cuticle, this protocol could be adapted to include currently available ex vivo culture conditions used in D. melanogaster to perform short-term live imaging studies24.
In the future, this model will be ideal to study cell-to-cell crosstalk and the contribution of other cell types to WIP by regulating gene expression with the Gal4/UAS system in other cell types of interest. Similar questions can also be answered using a variety of genetic and mutant backgrounds. The dissected adult fly abdomen contains a variety of cell types that can be easily visualized using this method, including fat body and oenocytes, lateral muscle fibers, sensory neurons, trachea, and macrophage-like hemocytes. In addition, this model will allow researchers to investigate how physiological variables influence WIP, including sex, diet, infection, age, and environmental stressors. While the protocol uses the adult female fly due to its larger size, WIP also occurs in the male fruit fly (Gjelsvik and Losick, unpublished). Polyploid cells have been found to arise during aging and age-associated disease in the mammalian liver, brain, eye, and heart12. The fruit fly model will enable researchers to study polyploidization in physiological and disease contexts&#160;because human disease-related genes are highly conserved.

Drosophila melanogaster is an attractive model system to study the cellular and molecular mechanisms of epithelial wound repair. As in mammals, the tissue repair mechanisms used depend&#160;on both the tissue and its developmental stage. Scarless wound healing occurs in the fruit fly embryo where an actomyosin &#34;purse string&#34; forms at the epithelial leading edge enabling the wound to seamlessly close1,2. Post-embryonic wound healing in larvae, pupae, and adult fruit flies results in extracellular matrix remodeling, melanin scar formation, and epithelial cell growth3,4,5,6. The epithelial cells increase in size by cell fusion and the endocycle, an incomplete cell cycle that bypasses mitosis3,4,7,8. As result, cell loss is compensated by polyploid cell growth instead of cell division. The adult fly hindgut, midgut, and follicular epithelium also rely on polyploid cell growth to compensate for cell loss after tissue damage9,10,11.
Polyploidy is a well-known aspect of organismal development in plants and insects, but in the last few years it has become more apparent that polyploidy is a conserved tissue repair strategy in vertebrates12. The zebrafish, which has the capacity to regenerate its heart, relies on polyploid cell growth to heal damaged epicardium13. Polyploidy also contributes to mammalian liver regeneration and kidney tubule epithelium repair after acute injury14,15. In these examples, polyploid cells are generated by endoreplication via either endocycle or endomitosis, which results in a binucleated cell due to a block in cytokinesis12. The enigma is why polyploid cells arise during wound repair and how polyploidy affects tissue function. Recent studies have provided new insight into the question of whether polyploidy offers a healing advantage or disadvantage. In zebrafish epicardium, polyploidy enhanced the speed of wound healing13. In D. melanogaster hindgut and mammalian liver, polyploidy was found to be protective against oncogenic growth11,14. In adult fly epithelium, it was recently found that polyploidy enables wound repair in the presence of genotoxic stress16. Endoreplication is resistant to DNA damage, allowing wound healing when cell proliferation would otherwise be compromised17. For cardiomyocytes in mouse and zebrafish hearts, however, polyploidy slows healing, resulting in enhanced scar formation18,19. Therefore, depending on the organ and/or cell type, polyploidy can be a beneficial or detrimental tissue repair strategy. The accessibility of D. melanogaster genetics coupled with analysis of the wound-induced polyploidization (WIP) response make it an ideal model system for elucidating the molecular and cellular mechanisms that guide this wound healing strategy.
Here, we present a protocol for analyzing WIP in the adult D. melanogaster epithelium. Included are instructions for fruit fly injury, dissection, immunostaining, mounting, imaging, and analysis of re-epithelialization, cell fusion, and endoreplication (ploidy). The imaging and ploidy analysis can also be adapted to other models to test whether WIP occurs. It should be noted that with an increase in nuclear DNA content there is often a corresponding increase in nuclear size. However, there are many examples in biology where nuclear size does not reflect a corresponding change in ploidy20. Even more caution should be taken when interpreting nuclear size in the context of a wound environment where cells will often spread or stretch to cover the wound site. Therefore, the only definitive proof of change in ploidy is to measure DNA content by this method (or others, such as whole genome sequencing)21. This method increases the suitability of the adult D. melanogaster abdominal epithelium as a model to study the role and regulation of polyploidy in wound repair.

<table><tbody><tr><td>35 mm Petri dishes</td><td>Fisher Scientific</td><td>FB0875713</td><td>For creating plates to dissect in</td></tr><tr><td>50 mL Conical Centrifuge Tubes</td><td>Fisher Scientific</td><td>14-432-22</td><td>For preparing staining reagents in</td></tr><tr><td>AxioImager M2 with Apotome</td><td>Zeiss</td><td>NA</td><td>For imaging samples</td></tr><tr><td>Blowgun mini</td><td>Genesee Scientific</td><td>54-104M</td><td>For anethesizing D. melanogaster strains</td></tr><tr><td>Bovine Serum Albumin, 30%</td><td>Sigma</td><td>A7284-500ML</td><td>For immuostaining</td></tr><tr><td>Carbon dioxide tank</td><td>various distributors</td><td>N/A</td><td>For anethesizing D. melanogaster strains</td></tr><tr><td>Click-iT EdU 594 Kit</td><td>Thermofisher</td><td>C10339</td><td>For EdU assay</td></tr><tr><td>Coverslips</td><td>Thermofisher</td><td>3406</td><td>For mounting</td></tr><tr><td>DAPI</td><td>Sigma</td><td>D9542-10MG</td><td>For immuostaining</td></tr><tr><td>Dissecting Plates (use Sylgard 184 Sil Elastic Kit)</td><td>Ellsworth Adhesives</td><td>184SIL</td><td>For creating plates to dissect in. Mix epoxy as directed, let dry overnight</td></tr><tr><td>Donkey anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 488 conjugate</td><td>Thermofisher</td><td>A21206</td><td>For secondary immuostaining</td></tr><tr><td>Donkey anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 568 conjugate</td><td>Thermofisher</td><td>A10042</td><td>For secondary immuostaining</td></tr><tr><td>Drosophila tubing and fittings</td><td>Genesee Scientific</td><td>59-124C, 59-123, 59-140</td><td>For anethesizing D. melanogaster strains</td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11252-20</td><td>For dissecting</td></tr><tr><td>epi-Gal4</td><td>Bloomington Drosophila Stock Center (b)</td><td>b38793</td><td>Losick et al. Current Biology, 2013</td></tr><tr><td>epi-Gal4, UAS-mCD8.RFP</td><td>Bloomington Drosophila Stock Center (b)</td><td>b38793, b27392</td><td>Losick et al. Current Biology, 2013</td></tr><tr><td>Excel</td><td>Microsoft</td><td></td><td>For performing ploidy calculations</td></tr><tr><td>Fiji/ImageJ (image analysis software)</td><td>NIH</td><td>&lt;a href=&quot;https://imagej.nih.gov/ij&quot; target=&quot;_blank&quot;&gt;https://imagej.nih.gov/ij&lt;/a&gt;</td><td>For image analysis</td></tr><tr><td>Fly food</td><td>Archon Scientific</td><td>N/A</td><td>Corn Syrup/Soy food</td></tr><tr><td>Flystuff Flypad</td><td>Genesee Scientific</td><td>59-114</td><td>For anethesizing D. melanogaster strains</td></tr><tr><td>Glass dissecting dish</td><td>Fisher Scientific</td><td>13-748B</td><td>For performing dissections in</td></tr><tr><td>Glass slides</td><td>Fisher Scientific</td><td>12-518-104C</td><td>For mounting</td></tr><tr><td>Goat anti-Mouse IgG (H+L) Secondary Antibody, Alexa Fluor 488 conjugate</td><td>Thermofisher</td><td>A11001</td><td>For secondary immuostaining</td></tr><tr><td>Goat anti-Mouse IgG (H+L) Secondary Antibody, Alexa Fluor 568 conjugate</td><td>Thermofisher</td><td>A11031</td><td>For secondary immuostaining</td></tr><tr><td>Grace&#039;s Insect Medium, unsupplemented</td><td>Thermofisher</td><td>11595030</td><td>For dissecting in</td></tr><tr><td>Insect pins</td><td>Fine Science Tools</td><td>26002-10</td><td>For wounding and pinning fly abdomens flat</td></tr><tr><td>Mouse anti-Fasciclin III (Drosophila) Primary Antibody</td><td>Developmental Studies Hybridoma Bank</td><td>7G10</td><td>For immunostaining epithelial cell-cell junctions</td></tr><tr><td>Mouting media</td><td>Vector Laboratories</td><td>H-1000</td><td>Anti-fade mounting media to prevent photo bleaching during imaging</td></tr><tr><td>Nail polish</td><td>Electron Microscopy Sciences</td><td>72180</td><td>For sealing slides</td></tr><tr><td>Ortibal shaker</td><td>Fisher Scientific</td><td>02-217-988</td><td>For immuostaining</td></tr><tr><td>Phosphate Buffered Saline, PH 7.4</td><td>Sigma</td><td>P3813-10PAK</td><td>For staining</td></tr><tr><td>Pin holders</td><td>Fine Science Tools</td><td>91606-07</td><td>For wounding</td></tr><tr><td>Rabbit anti-Grainyhead Primary Antibody</td><td>N/A</td><td>N/A</td><td>For immunostaining epithelial nuclei. Protocol to make antibody can be found (Ref. #4 and 8)</td></tr><tr><td>Rabbit anti-RFP Primary Antibody</td><td>MBL</td><td>PM005</td><td>For immunostaining mCD8-RFP fly epithelium</td></tr><tr><td>Stereomicroscope</td><td>Olympus</td><td>SZ51</td><td>For dissecting and mounting fly tissue</td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>10789704001</td><td>For immuostaining</td></tr><tr><td>UAS-E2F RNAi, UAS-RacDN</td><td>VDRC (v) and Bloomington Drosophila Stock Center (b)</td><td>v108837, b6292</td><td>Losick et al. Current Biology, 2013</td></tr><tr><td>UAS-fzr RNAi, UAS-Stg</td><td>VDRC (v) and Bloomington Drosophila Stock Center (b)</td><td>v25550, b56562</td><td>Grendler et al. Development, 2019</td></tr><tr><td>UAS-Myc</td><td>Bloomington Drosophila Stock Center (b)</td><td>b9674</td><td>Grendler et al. Development, 2019</td></tr><tr><td>UAS-myc RNAi</td><td>Bloomington Drosophila Stock Center (b)</td><td>b36123</td><td>Grendler et al. Development, 2019</td></tr><tr><td>Vannas Spring Scissors</td><td>Fine Science Tools</td><td>15000-00</td><td>For dissecting</td></tr></tbody></table>

a drosophila model to study wound-induced polyploidization

Polyploidy is a frequent phenomenon whose impact on organismal health and disease is still poorly understood. A cell is defined as polyploid if it contains more than the diploid copy of its chromosomes, which is a result of endoreplication or cell fusion. In tissue repair, wound-induced polyploidization (WIP) has been found to be a conserved healing strategy from fruit flies to vertebrates. WIP has several advantages over cell proliferation, including resistance to oncogenic growth and genotoxic stress. The challenge has been to identify why polyploid cells arise and how these unique cells function. Provided is a detailed protocol to study WIP in the adult fruit fly epithelium where polyploid cells are generated within 2 days after a puncture wound. Taking advantage of D. melanogaster&#39;s extensive genetic tool kit, the genes required to initiate and regulate WIP, including Myc, have begun to be identified. Continued studies using this method can reveal how other genetic and physiological variables including sex, diet, and age regulate and influence WIP&#39;s function.

A detailed protocol is provided to use D. melanogaster as a model to study wound-induced polyploidization (WIP). This wound healing model provides many advantages over mammalian and other fly models of WIP. Polyploidy was easily induced by a mechanical puncture with an insect pin and polyploid cells were generated within a short period of time (2-3 dpi) (Figure 1A, 1B)4. The major challenge is the dissection of intact abdominal tissue without any perturbations to the epithelium. The D. melanogaster epithelium is easily accidentally bumped or scratched with the sharp dissection tools. Therefore, the steps of this protocol should be practiced prior to use and analysis.
First, the injury was restricted to the ventral female abdomen, which provides a large, flat opaque tissue area ideal for imaging. The puncture wounds were inflicted in the pleurite epithelium, which lies on either side of the ventral midline sternites and targeted between tergite (T) segments T4-T5 (Figure 1A-C). This wound placement provides a large visible area that is not disrupted by the dissection. Challenging steps include the abdominal spring scissor cut and pinning steps (Figure 1D, 1E). The spring cutting step worked best when the abdomens were cut in a reduced volume of Grace&#39;s solution (~30 &#181;L) to decrease tissue movement. A well-centered cut along the dorsal midline was necessary to provide sufficient area on the abdominal dorsal flaps to pin open on the dissection plate (Figure 1C). The abdomen must be gently pinned on the four corners without excessive force (Figure 1E). A pin push that is too hard will distort the abdominal tissue and could even push the tissue into the dissection plate. If this happens, the tissue must be discarded. Once the abdominal tissue was fixed, it remained on the dissection plate until immunofluorescence staining was complete and the abdomens were mounted on a glass coverslip for imaging (Figure 1F).
Wound healing requires a continuous epithelial sheet to form, which is dependent on endoreplication and cell fusion4,16. The septate junction protein FasIII, which labels cell-cell junctions, provided an indicator for whether any processing perturbations occurred during preparation (Figure 1G, 1H). Abdomens with large scratches (unstained area) that perturb the wound area must be discarded and were not used for further analysis (Figure 1H).
The next step was to analyze intact samples for any defects in WIP. This protocol includes distinct assays to detect different aspects of the WIP response (Figure 2). Wound repair was complete when a central, large, multinucleated cell covered the wound scab (Figure 3A). Here cell fusion was detected by staining for FasIII/Grh and quantifying the number of Grh+ epithelial nuclei encompassed in the FasIII outlined area4. Defects in wound closure or re-epithelialization were detected when gaps of &#62;10 &#181;m in the epithelial sheet were observed (Figure 3B, red arrow). This was the case, for example, when WIP was inhibited by the activation of mitotic cycle via expression of stg, fzrRNAi, as recently reported16. In this genetic condition, 52% of the wounds were not able to form a continuous epithelial sheet over the wound scab (Figure 3B, 3C).
Another method to measure wound repair in this model was by visualizing the epithelial membrane with epi-Gal4 expression of UAS-mCD8-ChRFP4 (Figure 2, Figure 3D). In the control, 91% of epithelial wounds closed completely by 3 dpi, but inhibiting WIP by blocking endoreplication (E2f1RNAi) and cell fusion (RacDN) simultaneously, as previously reported, caused 92% of epithelial wounds to remain completely open (Figure 3D, 3E)8,16. The activation of mitotic cell cycle by expression of stg, fzrRNAi also resulted in an epithelial wound closure defect. However, by visualizing the epithelial cell membrane, the extent of re-epithelialization defect could be determined. The WIP mutant (E2f1RNAi, RacDN) fly wounds were more open than the stg, fzrRNAi wounds (Figure 3D, dashed red outline)16. This membrane wound healing assay provided more information on the extent of the wound repair defect. As a result, re-epithelialization defects could be grouped as either completely open, partially closed (i.e., &#62;10 &#181;m gaps), or completely closed (Figure 3D, 3E).
In addition to cell fusion, epithelial cells grow in size by endoreplication, an incomplete cell cycle that doubles the nuclear DNA content. Endoreplication was assayed by both cell cycle activity and direct nuclear DNA ploidy measurements (Figure 2 and Figure 4). Here, cell cycle activity was detected by incorporation of the thymidine analog, EdU (Figure 4A, 4B). D. melanogaster epithelial cells were found to enter the endocycle, an incomplete cell cycle that oscillates between the S and G phases without an intervening M phase4,12. The adult D. melanogaster diet was supplemented with EdU+ food prior to injury and the flies were maintained on an EdU+ diet until dissection at 2 dpi (Figure 4A). The EdU was then detected using the manufacturer&#39;s Click-iT protocol. This EdU assay was used to determine where, when, and how many nuclei were triggered to enter S phase in response to a wound. Using the Gal4/ UAS system, it was recently found that epithelial specific expression of myc can either block (mycRNAi) or exacerbate (Myc overexpression) the competence of epithelial cells to enter S phase. As result, it has been shown that Myc is sufficient to induce endoreplication in postmitotic cells, even without injury16,22.
Next, epithelial ploidy was determined by directly measuring nuclear DNA content. Epithelial nuclei were identified by immunofluorescence staining for the epithelial specific marker, Grh (Figure 4D). In the Fiji imaging software, epithelial nuclei were systemically identified and then thresholded using Grh nuclear stain. Nuclei were then separated and the ROIs overlaid on to the SUM of stacks DAPI image (Figure 4D, green arrow). Any overlapping nuclei were manually deleted before the Integrated Density of the selected nuclei was measured (Figure 4D, red arrows). This semiautomated method allows one to quantify the distribution and ploidy of most nuclei throughout the uninjured and repaired fly abdominal epithelium8. As recently reported, the epithelial nuclei surrounding the wound were composed of 44% polyploid nuclei with DNA content more than 3C at 3 dpi (Figure 4E, 4F)16. As expected from the EdU results, knockdown of myc led to a significant block in endoreplication, as only 9% of epithelial nuclei were polyploid, whereas overexpression of Myc resulted in 100% polyploid epithelial nuclei around the wound site (Figure 4F)16. Epithelial nuclear size was also visibly affected by myc expression with either reduced or enlarged nuclei present. However, nuclear area is not an accurate measure of ploidy and physiological effects, because factors such as cell stretching can also influence nuclear size without affecting nuclear DNA content20.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61252/61252fig1.jpg" /> 
Figure 1: Adult fruit fly abdominal wounding, dissecting, and tissue mounting. (A) Diagram of the adult abdominal wounding assay. Flies should be injured on either side of the abdomen at tergite 4 (T4). (B) Adult female fruit fly 3 dpi with melanin scab formed from wound healing (white box). Scale bar = 50 &#181;m. (C) Dissected adult abdomen, dorsal view, with tergites labeled. Abdomens were filleted down the midline of the dorsal side (white dashed line). Scale bar = 50 &#181;m. (D) Dissected and filleted adult abdomens prior to pinning. Scale bar = 50 &#181;m. (E) Pinned adult abdomen on a dissecting plate. A pin was placed in each of the four corners of the abdomen on the dorsal side. The tissue was gently opened but not stretched, to avoid tearing. (F) Adult abdomens were mounted and placed on a glass coverslip with the inside of the abdomen facing down towards the coverslip and cuticle oriented toward the glass slide. (G) FasIII staining of intact wound area with no processing perturbation and a central syncytium (dashed yellow line). Scale bar = 50 &#181;m. (H) Image of a scratched wound area (*) with an unstained FasIII region that disrupts the syncytium. <a href="https://www.jove.com/files/ftp_upload/61252/61252fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61252/61252fig2.jpg" /> 
Figure 2: WIP analysis workflow. The flowchart depicts the three assays described in this study and overlapping and distinct steps to detect and measure the WIP response. The EdU assay measures cell cycle activity (blue boxes), ploidy and re-epithelialization are detected by Grh/FasIII immunostaining (green boxes), and expression of membrane RFP allows measurement of the extent of epithelial wound closure (pink boxes). Common steps are in gray boxes and D. melanogaster strain genotypes are listed above. <a href="https://www.jove.com/files/ftp_upload/61252/61252fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61252/61252fig3.jpg" /> 
Figure 3: Methods to detect re-epithelialization during WIP. Re-epithelialization was perturbed when WIP was genetically inhibited. Immunofluorescent images of control (A) and stg, fzrRNAi (B) at 3 dpi. Epithelial nuclei and septate junctions were stained with Grh (green) and FasIII (magenta), respectively. (A&#39; and B&#39;) FasIII staining alone showed that re-epithelialization was impaired (red arrow) in stg, fzrRNAi epithelium. Scale bar = 50 &#181;m. (C) Quantification of re-epithelialization defects (%) at 3 dpi (gray): control (n = 8), stg, fzrRNAi (n = 6). Error bars indicate standard error; statistical significance was measured via Student&#39;s T-test, **P &#60; 0.01. Re-epithelialization during wound repair could also be detected by expression of a membrane linked RFP using epi-Gal4, UAS-mCD8-RFP. (D) Immunofluorescent images of control, E2F1RNAi, RacDN, and stg, fzrRNAi at 3 dpi. Scale bar = 20 &#181;m. Wound scab (yellow outline) and open epithelial wound area (red outline). (E) Quantification of wound closure in ctrl (n = 11), E2F1RNAi, RacDN (n = 13), and stg, fzrRNAi (n = 13). Adapted from Grendler et al.16. <a href="https://www.jove.com/files/ftp_upload/61252/61252fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61252/61252fig4.jpg" /> 
Figure 4: Methods to detect endoreplication during WIP. (A) Timeline of EdU assay: adult Drosophila were fed 75 &#181;L of 5 mM yeast-EdU every day 2 days prior to injury and continued until 2 dpi. (B) Immunofluorescent images of EdU label in fly strains expressed with epi-Gal4/ UAS system at 2 dpi. Wound scab (W). Scale bar = 50 &#181;m. (C) Average number of EdU+ epithelial nuclei per fly at 2 dpi: ctrl (n = 37), mycRNAi#1 (n = 10), and Myc (n = 8). Error bars indicate standard error; statistical significance was measured via Student&#39;s T-test, *P &#60; 0.05, **P &#60; 0.01. (D) Schematic of detection and measurement of epithelial nuclear ploidy. Epithelial nuclei were identified and thresholded by the anti-Grh stain in Fiji. Overlapping epithelial nuclei were separated (green arrowheads) or removed (red arrowheads) if overlaid by nonepithelial nuclei. The integrated density and nuclear area of corresponding DAPI stained nuclei image were measured. (E) Epithelial nuclear size (Grh) was altered by myc expression at 3 dpi. (F) Epithelial nuclear ploidy (%) at 3 dpi: ctrl (n = 4), mycRNAi#1 (n = 6), and Myc (n = 3). Adapted from Grendler et al.16. <a href="https://www.jove.com/files/ftp_upload/61252/61252fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Drosophila Model to Study Wound-induced Polyploidization at JoVE.com

A Drosophila Model to Study Wound-induced Polyploidization

Wound-induced polyploidization is a conserved tissue repair strategy where cells grow in size instead of dividing to compensate for cell loss. Here is a detailed protocol on how to use the fruit fly as a model to measure ploidy and its genetic regulation in epithelial wound repair.

A Drosophila Model to Study Wound-induced Polyploidization

Polyploidy is a frequent phenomenon whose impact on organismal health and disease is still poorly understood. A cell is defined as polyploid if it ...

a-drosophila-model-to-study-wound-induced-polyploidization

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

5.2K Views.  Boston College. Wound-induced polyploidization is a conserved tissue repair strategy where cells grow in size instead of dividing to compensate for cell loss. Here is a detailed protocol on how to use the fruit fly as a model to measure ploidy and its genetic regulation in epithelial wound repair.

Video: A Drosophila Model to Study Wound-induced Polyploidization

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the context of a whole organism. Sophisticated techniques to generate genetic mosaics facilitate induction of visually marked, genetically defined clones surrounded by normal tissue. The clones can be analyzed through diverse molecular, cellular and omics approaches. This study describes how to generate fluorescently labeled clonal tumors of varying malignancy in the eye/antennal imaginal discs (EAD) of Drosophila larvae using the Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. It describes procedures how to recover the mosaic EAD and brain from the larvae and how to process them for simultaneous imaging of fluorescent transgenic reporters and antibody staining. To facilitate molecular characterization of the mosaic tissue, we describe a protocol for isolation of total RNA from the EAD. The dissection procedure is suitable to recover EAD and brains from any larval stage. The fixation and staining protocol for imaginal discs works with a number of transgenic reporters and antibodies that recognize Drosophila proteins. The protocol for RNA isolation can be applied to various larval organs, whole larvae, and adult flies. Total RNA can be used for profiling of gene expression changes using candidate or genome-wide approaches. Finally, we detail a method for quantifying invasiveness of the clonal tumors. Although this method has limited use, its underlying concept is broadly applicable to other quantitative studies where cognitive bias must be avoided.

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

This protocol demonstrates how to generate fluorescently marked, genetically defined clonal tumors in the Drosophila eye/antennal imaginal discs (EAD). It describes how to dissect the EAD and brain from the third instar larvae and how to process them to visualize and quantify gene expression changes and tumor invasiveness.

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the ...

Cancer Research

Manipulation of Ploidy in Caenorhabditis elegans

Mechanisms that involve whole genome polyploidy play important roles in development and evolution; also, an abnormal generation of tetraploid cells has been associated with both the progression of cancer and the development of drug resistance. Until now, it has not been feasible to easily manipulate the ploidy of a multicellular animal without generating mostly sterile progeny. Presented here is a simple and rapid protocol for generating tetraploid Caenorhabditis elegans animals from any diploid strain. This method allows the user to create a bias in chromosome segregation during meiosis, ultimately increasing ploidy in C. elegans. This strategy relies on the transient reduction of expression of the rec-8 gene to generate diploid gametes. A rec-8 mutant produces diploid gametes that can potentially produce tetraploids upon fertilization. This tractable scheme has been used to generate tetraploid strains carrying mutations and chromosome rearrangements to gain insight into chromosomal dynamics and interactions during pairing and synapsis in meiosis. This method is efficient for generating stable tetraploid strains without genetic markers, can be applied to any diploid strain, and can be used to derive triploid C. elegans. This straightforward method is useful for investigating other fundamental biological questions relevant to genome instability, gene dosage, biological scaling, extracellular signaling, adaptation to stress, development of resistance to drugs, and mechanisms of speciation.

Manipulation of Ploidy in Caenorhabditis elegans

This method allows for the generation of tetraploid and triploid Caenorhabditis nematodes from any diploid strain. Polyploid strains generated by this method have been used to study chromosome interactions in meiotic prophase, and this method is useful for examining important basic questions in cell, developmental, evolutionary, and cancer biology.

Mechanisms that involve whole genome polyploidy play important roles in development and evolution; also, an abnormal generation of tetraploid cells has ...

Genetics

Microinjection Wound Assay and In vivo Localization of Epidermal Wound Response Reporters in Drosophila Embryos.

The Drosophila embryo develops a robust epidermal layer that serves both to protect the internal cells from a harsh external environment as well as to maintain cellular homeostasis. Puncture injury with glass needles provides a direct method to trigger a rapid epidermal wound response that activates wound transcriptional reporters, which can be visualized by a localized reporter signal in living embryos or larvae. Puncture or laser injury also provides signals that promote the recruitment of hemocytes to the wound site. Surprisingly, severe (through and through) puncture injury in late stage embryos only rarely disrupts normal embryonic development, as greater than 90% of such wounded embryos survive to adulthood when embryos are injected in an oil medium that minimizes immediate leakage of hemolymph from puncture sites. The wound procedure does require micromanipulation of the Drosophila embryos, including manual alignment of the embryos on agar plates and transfer of the aligned embryos to microscope slides. The Drosophila epidermal wound response assay provides a quick system to test the genetic requirements of a variety of biological functions that promote wound healing, as well as a way to screen for potential chemical compounds that promote wound healing. The short life cycle and easy culturing routine make Drosophila a powerful model organism. Drosophila clean wound healing appears to coordinate the epidermal regenerative response, with the innate immune response, in ways that are still under investigation, which provides an excellent system to find conserved regulatory mechanisms common to Drosophila and mammalian epidermal wounding.

Microinjection Wound Assay and In vivo Localization of Epidermal Wound Response Reporters in Drosophila Embryos.

The embryonic epidermis of very late stage Drosophila embryos provides an in vivo system for rapid puncture wound response analysis and can be combined with genetic manipulations or chemical microinjection treatments to advance studies in wound healing for translation into mammalian models.

The Drosophila embryo develops a robust epidermal layer that serves both to protect the internal cells from a harsh external environment as well as to ...

Bioengineering

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

Wing development in Drosophila melanogaster is an ideal model for studying morphogenesis at tissue level. These appendages develop from a group of cells named wing imaginal discs formed during embryonic development. In the larval stages the imaginal discs grow, increasing its number of cells and forming monolayered epithelial&#160;structures. Inside the pupal case, the imaginal discs bud out and fold into bilayers along a line that becomes the future margin of the wing. During this process, the longitudinal primodia veins originate vein cells on the prospective dorsal and ventral surfaces of the wing. During the pupal stage the stripes of vein cells of each surface communicate in order to generate tight tubes; at the same time, the cross-veins begin their formation.
With the help of appropriate molecular markers, it is possible to identify the major elements composing the wing during its development. For this reason, the ability to accurately detect transcripts or proteins in this structure is critical for studying their abundance and localization related to the development process of the wing.
The procedure described here focuses on manipulating pupal wings, providing detailed instructions on how to dissect the wing during the pupal stage. The dissection of pupal tissue is more difficult to perform than their counterparts in third instar larvae. This is why this approach was developed, to obtain rapid and efficient high quality samples. Details of how to immunostain and mount these wing samples, to allow the visualization of proteins or cell components, are provided in the protocol. With little expertise it is possible to collect 8-10 high quality pupal wings in a short amount of time.

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

The ability to accurately detect transcripts or proteins in Drosophila tissues is critical for studying their abundance and localization related to the development process. Here is the description of a straightforward procedure to dissect pupal wings. These wings can be used as samples in several techniques (immunohistochemistry, PCR assay, etc.).

Wing development in Drosophila melanogaster is an ideal model for studying morphogenesis at tissue level. These appendages develop from a group of cells ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

During the rapid inflammatory response to tissue damage, cells of the innate immune system are quickly recruited to the injury site. Once at the wound, innate immune cells perform a number of essential functions, such as fighting infection, clearing necrotic debris, and stimulating matrix deposition. In order to fully understand the diverse signaling events that regulate this immune response, it is crucial to observe the complex behaviors of (and interactions that occur between) multiple cell lineages in vivo, and in real-time, with the high spatio-temporal resolution. The optical translucency and the genetic tractability of Drosophila embryos have established Drosophila as an invaluable model to live-image and dissect fundamental aspects of inflammatory cell behavior, including mechanisms of developmental dispersal, clearance of apoptotic corpses&#160;and/or microbial pathogens, and recruitment to wounds. However, more recent work has now demonstrated that employing a much later stage in the Drosophila lifecycle &#8211; the Drosophila pupa &#8211; offers a number of distinct advantages, including improved RNAi efficiency, longer imaging periods, and significantly greater immune cell numbers. Here we describe a protocol for imaging wound repair and the associated inflammatory response at the high spatio-temporal resolution in live Drosophila pupae. To follow the dynamics of both re-epithelialization and inflammation, we use a number of specific in vivo fluorescent markers for both the epithelium and innate immune cells. We also demonstrate the effectiveness of photo-convertible fluorophores, such as Kaede, for following the specific immune cell subsets, to track their behavior as they migrate to, and resolve from, the injury site.

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

Here we present a protocol for live-imaging wound repair and the associated inflammatory response at high spatio-temporal resolution in vivo. This method utilizes the pupal stage of Drosophila development to enable long-term imaging and tracking of specific cell populations over time and is compatible with efficient RNAi-mediated gene inactivation.

During the rapid inflammatory response to tissue damage, cells of the innate immune system are quickly recruited to the injury site. Once at the wound, ...

Immunology and Infection

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Medicine

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

Biology

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

The pupae of Drosophila melanogaster are immobile for several days during metamorphosis, during which they develop a new body with a thin transparent adult integument. Their immobility and transparency make them ideal for in vivo live imaging experiments. Many studies have focused on the dorsal epithelial monolayer of the pupal&#160;notum because of its accessibility and relatively large size. In addition to the studies of epithelial mechanics and development, the notum has been an ideal tissue to study wound healing. After an injury, the entire epithelial repair process can be captured by live imaging over 6-12 h. Despite the popularity of the notum for live imaging, very few published studies have utilized fixed notum samples. Fixation and staining are common approaches for nearly all other Drosophila tissues, taking advantage of the large repertoire of simple cellular stains and antibodies. However, the pupal notum is fragile and prone to curling and distortion after removal from the body, making it challenging to complement live imaging. This protocol offers a straightforward method for fixing and staining the pupal notum, both intact and after laser-wounding. With this technique, the ventral side of the pupa is glued down to a coverslip to immobilize the pupa, and the notum is carefully removed, fixed, and stained. The notum epithelium is mounted on a slide or between two coverslips to facilitate imaging from the tissue&#39;s dorsal or ventral side.

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

The present protocol details the preparation and visualization of fixed tissue of the Drosophila pupal notum. It can be used for either intact or wounded tissue, and the original architecture of the tissue is preserved. The procedures for dissecting, fixing, and staining are all described in this article.

The pupae of Drosophila melanogaster are immobile for several days during metamorphosis, during which they develop a new body with a thin transparent ...

Drosophila Development and Reproduction

One of the many reasons that make Drosophila an extremely valuable organism is that the molecular, cellular, and genetic foundations of development are highly conserved between flies and higher eukaryotes such as humans. Drosophila progress through several developmental stages in a process known as the life cycle and each stage provides a unique platform for developmental research. This video introduces each stage of the Drosophila life cycle and details the physical characteristics and major developmental events that occur during each stage. Next, the video discusses the genetic regulation of pattern formation, which is important for establishing the body plan of the organism and specifying individual tissues and organs. In addition, this video gives an overview of Drosophila reproduction, and how to use the reproductive characteristics of Drosophila to set up a genetic cross. Finally, we discuss examples of how the principles of Drosophila development and reproduction can be applied to research. These applications include RNA interference, behavioral assays of mating behaviors, and live imaging techniques that allow us to visualize development as a dynamic process. Overall, this video highlights the importance of understanding development and reproduction in Drosophila, and how this knowledge can be used to understand development in other organisms.

Development and Reproduction of Drosophila melanogaster

One of the many reasons that make Drosophila an extremely valuable organism is that the molecular, cellular, and genetic foundations of development are ...

A Drosophila Model to Study Wound-induced Polyploidization

Summary

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Chapters in this video

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

Manipulation of Ploidy in Caenorhabditis elegans

Microinjection Wound Assay and In vivo Localization of Epidermal Wound Response Reporters in Drosophila Embryos.

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

Drosophila Development and Reproduction