The authors would like to thank Dr. M. Shane Hutson for establishing the laser ablation system used for pupal wounding and his contributions to and feedback on developing the protocol. This work was supported by 1R01GM130130 to APM and M. Shane Hutson. JW&#160;was supported by 2T32HD007502-21.

Drosophila melanogaster (fruit flies) were maintained at 25 &#176;C on a standard cornmeal-molasses medium. The studies were conducted on EGFP-tagged histone H2A pupae (w[*]; P{w+mC=His2Av-EGFP.C}2/SM6a). The flies were obtained from a public stock center (see Table of Materials).
1. Pupae immobilization
<ol>
	<li>Apply a 2&#34; strip of double-sided tape to a microscope slide.</li>
	<li>Identify white prepupae in vials raised at 25 &#176;C, and use a marker to indicate their location outside the vial. Return the vials to 25 &#176;C. 
	NOTE: White prepupae are characterized by their immobility, white color, and everted spiracles. These form 0-1 h after puparium formation (APF), or stage P18.</li>
	<li>12-15 h later, carefully remove 3-4 of the indicated pupae (without popping them) using a dissection scope and collect them on the microscope slide next to the tape. 
	NOTE: Pupae will now be stage P5 with an everted head sac visible at the pupae&#39;s anterior end8.</li>
	<li>Place the pupae at least one pupa width apart onto the tape with their ventral sides down.</li>
	<li>Place a drop of adhesive glue on a paraffin film (see Table of Materials) or in a centrifuge tube lid. Dip the end of a 0.1-10 &#181;L pipette tip (no pipette) in the drop of adhesive glue. Tap the pipette tip twice on a 24 mm x 60 mm (1.5 thickness) coverslip, 1 cm x 1 cm away from a corner, creating a line of adhesive glue ~1/2 the length of the pupa.</li>
	<li>Preset a 0.2-2 &#181;L (P2) pipette to 2 &#181;L, and a 200 &#181;L (P200) pipette to 200 &#181;L, and fit them with tips so they are ready to be filled with 1x PBS + 0.1 mM Ca2+, which will rapidly solidify the adhesive glue on contact.</li>
	<li>Insert forceps (see Table of Materials) near the side of the head and gently remove the case from the anterior to posterior9. Remove as much of the case as possible. Grasp the pupa&#39;s developing legs with a pair of blunt forceps and carefully pull the pupa from its case. 
	NOTE: A small rupture on the ventral portion of the pupae will not be detrimental to this procedure.</li>
	<li>Lay the pupa in the corner of the coverslip.</li>
	<li>Grasp the pupa at the posterior abdomen or the developing wing with blunt forceps, lift, and place the ventral side of the pupae down into the line of adhesive glue.</li>
	<li>Quickly fill the P2 pipette with 2 &#181;L of 1x PBS + 0.1 mM of Ca2+, and holding it in the air, expel just enough to form a small bubble at the tip (0.25-0.5 &#181;L).</li>
	<li>Touch the small bubble of the solution to one side of the pupae at the base of the thorax, then repeat&#160;on the other side. 
	NOTE: This will solidify a small amount of the adhesive glue to hold the pupa in place. Generally, all the solutions will not be used.</li>
	<li>Fill the P200 pipette with 200 &#181;L of 1x PBS + 0.1 mM of Ca2+, then place the pipette&#39;s tip over the thorax and expel the contents to submerge the pupae completely. The remainder of the adhesive glue will solidify immediately.</li>
	<li>Remove ~100 &#181;L of the PBS solution, so the pupa is barely submerged before proceeding immediately to the next step. 
	&#8203;NOTE: For unwounded samples, start from step 1.1. For wounded partially dissected samples, start at step 1.5. Wounding via laser ablation has been described previously14,21. Immobilization, dissection, and mounting steps need to be performed using a dissection microscope.</li>
</ol>
2. Dissecting the notum
<ol>
	<li>Grasp a pair of microdissection scissors bracing one side of the handle against the dominant hand&#39;s index finger and middle finger, so the thumb of the dominant hand applies the cutting force (Figure 2A,B).</li>
	<li>Stabilize the neck of the scissors against the middle finger of the non-dominant hand while bracing the coverslip with the ring finger of the non-dominant hand.</li>
	<li>Snip at the middle of the dorsal abdomen to create a small hole, ~0.2-0.5 mm. Some hemolymph will usually spill out and is a good indicator of the breach.</li>
	<li>Make small 0.5-0.75 mm cuts through the integument from the posterior to anterior, encircling the dorsal tissue to isolate it. To create a tissue as flat as possible, avoid cutting too ventrally; only the dorsal &#39;dome&#39; of the thorax should be removed along with small sections of the head and abdomen.</li>
	<li>Repeat posterior to anterior cuts on the other side of the pupae.</li>
	<li>Rotate the dissection stage to allow for a clean cut through the head if necessary. 
	NOTE: At this stage, the dorsal integument, or notum, will be separated from the rest of the pupa. If it appears separated but is not easily moved, a few cuts below the notum can help dislodge it.</li>
	<li>Add ~200 &#181;L of 1x PBS to the center of the coverslip and make a channel connecting it to the original dissection droplet by gently dragging the pipette tip across the cover glass from the new droplet to the original.</li>
	<li>Using a pair of blunt forceps, gently push or drag the isolated notum to the center of the coverslip and rotate, so the interior side faces upward. Never remove the tissue from the droplet. 
	NOTE: It is essential to move the notum away from the original dissection site to avoid any remnants of the adhesive glue occluding the sample during later imaging.</li>
	<li>Hold the notum down with the blunt forceps by pressing into the abdominal or head sections. Using a pair of sharp forceps and/or gentle expulsions of 1x PBS from a 200 &#181;L pipette, remove any remaining fat body, muscle bands, or hemolymph (if present) to fully expose the monolayer epithelium and make the eventual staining more even. The dissected notum should appear as in Figure 3A.</li>
	<li>Once the tissue is clean, use a 200 &#181;L pipette to remove as much of the PBS solution as possible (along with debris and the ventral part of the pupae), monitoring with a dissection scope to avoid aspirating the notum.</li>
	<li>Once most of the liquid has been removed, use an absorbent tissue to carefully wipe away the rest of the adhesive glue and pupa, as well as any other debris that lingers on the coverslip. 
	NOTE: If some adhesive glue remains on the coverslip, it will not cause an issue so long as it is thinner than the dorsal tissue itself.</li>
	<li>Add 150-200 &#181;L of 4% PFA (in 1x PBS) and fix for 20 min at room temp. Depending on the dissection speed, 1 or more&#160;more pupae can be dissected during the first pupa&#39;s fixation.</li>
	<li>Remove the PFA and replace it with 1x PBS to wash the notum once for 30 s.</li>
	<li>If proceeding with antibody staining, perform 5 min washes (3 times) in 1x PBS or 1x PBST (Supplementary File 1) to permeabilize the tissue if the antigen is intracellular.</li>
	<li>Store the sample in 1x PBS + 0.02% NaN3 overnight in a humidified chamber, or if planning to stain the tissue, incubate overnight in Blocking solution (Supplementary File 1).</li>
</ol>
3. Staining the notum
NOTE: For staining using antibodies or cellular stains follow the steps below. -The notum must not be removed from the solution, as this will likely cause the tissue to curl. Thus, adapt staining protocols to be conducted entirely on the coverslip and keep in a humidified chamber for any steps longer than 5 min.&#160;Monitoring the samples under a dissection microscope can help to prevent accidental aspiration of the tissue during washes.
<ol>
	<li>To visualize the cell borders, incubate in 200 &#181;L of anti-FasIII primary Mouse IgG2a antibody (see Table of Materials) at 1:8 concentration diluted in Blocking Buffer&#160;+ 0.02% NaN3 overnight at 4 &#176;C.</li>
	<li>Wash out excess primary antibody (3 times) with 200 &#181;L of 1x PBS + 0.02% NaN3 for 1 h per wash at room temperature.</li>
	<li>Perform secondary antibody incubation with 200 &#181;L of 1:200 concentration anti-mouse IgGa2 in Cy3 in Blocking Buffer&#160;+ 0.02% NaN3 for 2 h at room temp.</li>
	<li>Wash out excess secondary antibody (3 times) with 200 &#181;L of 1x PBS + 0.02% NaN3 for 1 h per wash at room temperature.</li>
	<li>To visualize the nuclei, incubate samples in 1 &#181;g/mL of DAPI for 45 min to allow sufficient time for the stain to penetrate through muscle bands; fixing in a DAPI-containing mounting medium is not effective with this tissue.</li>
	<li>Wash out excess DAPI (3 times) with 200 &#181;L of 1x PBS + 0.02% NaN3 for 5 min per wash at room temperature, then leave in 200 &#181;L of 1x PBS + 0.02% NaN3 overnight at 4 &#176;C, or mount immediately.</li>
</ol>
4. Mounting and visualizing the notum
<ol>
	<li>Following staining, prepare a new 24&#160;&#215; 60 coverslip (topper) with supports. 
	NOTE: Because the notum is dome-shaped, flattening it completely results in distorted wrinkled tissue. Creating a gap between the two coverslips allows the notum to retain its normal shape.</li>
	<li>Create a gap of ~200 &#181;m by using spacers made from 22 x 22 coverslips (No. 0 thickness, ~100 &#181;m thick), adhered ~1 cm apart with nail polish in the middle of the topper.</li>
	<li>To adhere, place the spacers on the topper and paint the distal edges of the spacers&#160;with a thin layer of nail polish. Let dry. 
	NOTE: Only use a nail polish that is thin and runny; thick nail polish will add unnecessary additional space between the coverslip and topper.</li>
	<li>Remove as much of the aqueous solution as possible from the sample.</li>
	<li>Immediately apply two drops (~100 &#181;L) of anti-fade mounting medium (see Table of Materials) to the sample.</li>
	<li>If necessary, use clean, sharp forceps to position the notum in the center of the anti-fade mounting medium droplet.</li>
	<li>Place the coverslip with the notum onto a ~10 x 40 mm support, such as a piece of thin foam (cut from the packing material inside coverslip boxes), to elevate the sample so it will not adhere to the work surface.</li>
	<li>Under a dissection scope, slowly lower the topper onto the sample. Once the anti-fade mounting medium meets the topper, gently release and allow capillary action to pull the topper down. 
	NOTE: Within the first few seconds, minor adjustments can be made to the coverslip position without damaging the notum.</li>
	<li>Place another piece of foam onto the topper and use a standard microscope slide as a weight to gently coax the anti-fade mounting medium between the sample coverslip, topper, and spacers.</li>
	<li>After 5-10 min, use an absorbent tissue to wick away any excess anti-fade mounting medium by gently touching the edges of the coverslip.</li>
	<li>Gently apply nail polish to each corner of the coverslips to adhere them together. Once dry, paint all edges of the coverslips to seal. Avoid coating all edges first, as this can often shift the coverslip and damage the dorsal tissue.</li>
	<li>Visualize the notum under fluorescence microscopy (see Table of Materials) through the dorsal and/or the ventral side.</li>
</ol>

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The authors have nothing to disclose.

Critical steps 
Optimizing three steps will dramatically increase the success of this protocol. First, in step 1.5, be sparing with the adhesive glue applied to the coverslip. If too much adhesive glue is added, the pupae can become entombed in a thick layer of solidified adhesive glue, which will make dissection impossible, and if it covers the notum itself, the adhesive glue will occlude the light from the sample. Second, during steps 2.3-2.6, ensure to remove only the top dome of the notum, excluding as much of the lateral tissue as possible. If included, the lateral tissue will become compressed during mounting and cause the middle of the notum to buckle inward, often placing it outside of the working distance of high numerical aperture objectives. Third, during the cleaning step 2.9, extreme care must be taken not to damage the monolayer epithelium. If the tissue has large portions missing or no signal can be detected, this step is likely to blame.
Troubleshooting 
Issue 1: Following mounting, the pupa comes away from the double-sided tape during dissection. This is a common issue, especially for beginners. The best remedy is to ensure that the outside of the pupae case is completely dry/free of food debris. Removing food with a pair of blunt forceps and allowing the case to air dry for 10-15 min will help adhere to the tape. Alternatively, use the non-dominant hand and a pair of blunt forceps to hold the pupae down against the tape during dissection. If the problem persists, applying a small, 5-10 &#181;L drop of nail polish to the base of the posterior end of the case and allowing it to harden generally provides enough adhesion for even the unruliest of pupae.
Issue 2: Notum collapses during step 2.3, or the initial breach through the integument is not smooth. If the integument is difficult to breach, there may be too much adhesive glue from the immobilization steps. Placing the dissected pupae into less adhesive glue will improve this issue. Further, ensure that the microdissection scissors are sharp. Blunt scissors will not be able to cut into the integument and tend to cause it to collapse inwards. Once hemolymph spills out of the pupae, ensure one of the cutting blades can enter the pupa without causing the notum to deform. If the notum collapses and the blade does not enter the pupa, continue snipping until a blade enters the pupae.
Issue 3: During step 2.4, scissors catch or drag the integument. If the scissors begin to catch or drag the integument, it often helps to switch to the opposite side of the pupae and proceed to &#39;loosen&#39; the integument. Further blunt microdissection scissors will make it difficult to achieve clean cuts through the integument, and sharpened scissors must be used.
Issue 4: The sample is accidentally aspirated during staining (steps 3.1-3.6). The dissected notum is challenging to see because it is transparent. It can be helpful to place a dark blue or black sheet below the coverslip to provide contrast (an old pipette tip holder rack works well.) Additionally, all solution changes can be performed under a dissection microscope.
Issue 5: No signal is detected/patchy signal is detected. After ruling out stain-specific problems, if no signal is detected or it is patchy, step 2.9 (cleaning) is likely the culprit. An absent or patchy signal can originate from damage and removal of the epithelial tissue during cleaning. Conversely, a poor signal can be caused by occlusion from the muscle bands/fat body cells if they are not removed, as they can limit the diffusion of stains and antibodies into the notum relative to the surrounding tissue. If the tissue is damaged, being gentler during cleaning is the best solution. If, instead, the stain is visible but patchy, increasing the vigor/time dedicated to the cleaning step is recommended to remove as much of the muscle and fat body as possible. Further, increasing the stain duration can help resolve this problem with better cleaning.
Issue 6: The notum tissue has a warped/wrinkled appearance during imaging. Warping and wrinkling of the tissue come from two sources. First, compressing the notum during mounting will cause it to buckle and warp. The best solution is to remove as much of the lateral tissues as possible so the dome is as short as possible and can fit between the coverslip spacers. Second, if the notum is bent during dissection, this bend will not straighten out during mounting, so extra care must be taken not to warp the notum during dissection. Accidental bending of the notum is most common when cutting the integument away from the rest of the pupae. It is tempting to have the dissection scissors at an angle relative to the pupae instead of keeping them in the sample plane as the pupae. However, angled scissors cause the integument to buckle upwards when cut instead of remaining flat.
Existing methods, limitations, and future applications 
Wang et al.20 reported a comparable dissection protocol for the isolation of pupal epithelium. This technique requires that the pupa remains within its case and be rapidly bisected with a scalpel. This protocol is incompatible with previously live-imaged samples, as live imaging requires removing a large section of the pupal case. Because pupae lack rigidity, pupal bisection outside of the case mangled the tissue, inspiring the creation of this protocol. The technique detailed here allows for isolation and fixation of the notum, and it could be used as the first step for a wide array of other methods such as cryosectioning, in situ hybridization, or electron microscopy.
This technique has some limitations. First, dissecting, fixing, and staining the notum is more time-consuming than live-imaging fluorescently tagged proteins in the notum, which requires only a simple dissection to remove the pupal case9,23. Secondly, compared to dissections of other Drosophila tissues, this dissection is more difficult because of the thin, fragile tissue and hydrophobic cuticle. For simply visualizing proteins in Drosophila epithelia, immunohistochemistry on fixed embryos, larval wing discs, or ovaries is easier. However, this technique allows the power of live imaging to be paired with fixation and staining, making it a powerful tool once mastered.
A dissection/fixation technique has some advantages over live imaging. Basal (interior) structures can be better resolved with a basal view (Figure 5L,J). Most importantly, live imaging is limited to fluorophores that must be genetically supplied, often requiring lengthy genetic crossing schemes. In contrast, the present protocol allows the application of stains, immunohistochemistry, and other techniques which require dissection and fixation. This dramatically increases the number of signals probed in the tissue while potentially decreasing the time to experimental results.

The pupal notum of Drosophila melanogaster has been increasingly used for live imaging studies in the last decade because the animal is both immobile and has a transparent cuticle at this stage1,2,3,4,5,6,7. However, the pupal notum is challenging to dissect and fix, making it difficult to complement live imaging studies with antibody and cell staining. The overall goal of this work is to create a reproducible protocol for dissecting and fixing the pupal notum for antibody and cell staining on new or previously live-imaged samples.
As larvae begin metamorphosis, the epidermis pulls away from the larval cuticle, forming a hard pupal case8. The larval body plan is broken down, and the new adult body plan is developed. During this time, pupae are immobile, making them ideal for live imaging. One commonly imaged tissue is the pupal notum, an adult monolayer epithelium that forms in the dorsal thorax. The notum is visually accessible after&#160;a simple dissection&#160;to remove the pupal case9. The whole animal can then be mounted, and the notum can be live imaged for hours or days, making it an ideal tissue to study epithelial cell behaviors during development, homeostasis, and following wounding10,11,12,13,14. However, the notum is challenging to dissect and fix because it is fragile and covered with a thin transparent adult cuticle that is hydrophobic. This hydrophobic cuticle makes it prone to curling in aqueous solutions when removed from the rest of the body. Thus, notum dissection and fixation has been reported only rarely and the dissection is often not described15,16,17,18. Without a detailed protocol in the literature, it is prohibitively difficult for a Drosophila researcher to complement live imaging with staining of pupae.
This technique aims to reproducibly dissect and fix samples that have been previously live-imaged, including those that have been laser-wounded. Because live imaging requires removal of the pupal case, this dissection technique begins by removing the anterior pupal case, unlike previous protocols that pin down or bisect pupae within the pupal case4,19,20. The notum is a fragile tissue, and wounding may exacerbate its fragility. Thus, to support this delicate tissue, the integument (the epithelium and attached transparent adult cuticle) of the notum and part of the head and abdomen are dissected away from the rest of the pupa while always submerged in an aqueous environment. This method reduces the likelihood of the tissue curling and being unusable. This technique has successfully stained wounded notum tissue as early as 30 min post wounding (Figure 1E-H) and at 3 h post wounding (Figure 1I-L). This protocol is expected to be effective for the duration of notum development or wound repair. The current technique will be helpful for researchers wishing to unite the live-imaging capabilities of the pupal notum with the abundance of available immunohistochemistry reagents.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#189;&#8221; Scotch Permanent Double-Sided tape</td>
			<td>Scotch 3M</td>
			<td>665</td>
			<td></td>
		</tr>
		<tr>
			<td>0.1-10 &#181;L uTIP pipette tip</td>
			<td>Biotix</td>
			<td>M-0011-9FC</td>
			<td></td>
		</tr>
		<tr>
			<td>22 x 22 mm No. 0 thickness coverslips</td>
			<td>Thomas Scientific</td>
			<td>1207Z28</td>
			<td></td>
		</tr>
		<tr>
			<td>24 x 60 mm coverslip</td>
			<td>Corning</td>
			<td>2980-246</td>
			<td></td>
		</tr>
		<tr>
			<td>3M VetBond</td>
			<td>3M</td>
			<td>1469SB</td>
			<td>Called &#34;adhesive glue&#34; in protocol</td>
		</tr>
		<tr>
			<td>Anti-FasIII primary Mouse IgG2a</td>
			<td>Developmental Studies Hybridoma Bank</td>
			<td>7G10</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride</td>
			<td>Fisher Chemical</td>
			<td>c79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Cy3-conjugated AffiniPure Goat Anti-Mouse IgG2a</td>
			<td>Jackson ImmunoResearch</td>
			<td>115-165-206</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma Aldrich</td>
			<td>D9542-1mg</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 Fine Tip Forceps</td>
			<td>Fine Science Tools</td>
			<td>No. 11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 Fine Tip Forceps: Blunt</td>
			<td>Fine Science Tools</td>
			<td>No. 11254-20</td>
			<td>Heavily used and unsharpened forceps, or dulled with a whetstone</td>
		</tr>
		<tr>
			<td>Fisherbrand Double frosted microscope slides</td>
			<td>Fisher Scientific</td>
			<td>22-034-486</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent Light Source</td>
			<td>Lumencor</td>
			<td>Celesta 90-10512</td>
			<td></td>
		</tr>
		<tr>
			<td>Fly Stock: EGFP-tagged Histone H2A</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>24163</td>
			<td></td>
		</tr>
		<tr>
			<td>Fly Vial Foam Plugs &#34;Flugs&#34;</td>
			<td>Genesee Scientific</td>
			<td>49-101</td>
			<td></td>
		</tr>
		<tr>
			<td>Humidified Chamber: Foil-wrapped small container lined with filter paper saturated with water</td>
			<td>Cole-Parmer</td>
			<td>759075D</td>
			<td>A great humidified chamber can be made from the styrofoam box containing Cole-Parmer cuvettes, 759075D, filled with 50 ml water.</td>
		</tr>
		<tr>
			<td>KimWipe</td>
			<td>KimTech</td>
			<td>34155</td>
			<td>Called &#34;Absorbent Tissue&#34; in protocol</td>
		</tr>
		<tr>
			<td>Nikon Ti2 Eclipse</td>
			<td>Nikon</td>
			<td>Eclipse Ti2-E</td>
			<td></td>
		</tr>
		<tr>
			<td>NIS Elements Software</td>
			<td>Nikon</td>
			<td>AR</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Pechiney Plastic Packaging</td>
			<td>PM-996</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde (PFA) 16%</td>
			<td>Ted Pella, Inc</td>
			<td>18505</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic Vials</td>
			<td>Genesee Scientific</td>
			<td>32-114</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Azide (NaN3)</td>
			<td>Fisher Scientific</td>
			<td>19038-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo Microdissection Scope</td>
			<td>Carl Zeiss</td>
			<td>STEMI 2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-00</td>
			<td>New or freshly sharpened scissors</td>
		</tr>
		<tr>
			<td>Vecta Shield</td>
			<td>Vector Laboratories</td>
			<td>H-1000</td>
			<td>Called &#34; antifade mounting medium&#34; in protocol</td>
		</tr>
		<tr>
			<td>Vecta Shield with DAPI</td>
			<td>Vector Laboratories</td>
			<td>H-1200</td>
			<td>Not ideal for pupal notum.</td>
		</tr>
		<tr>
			<td>X-Light V2 Spinning Disc</td>
			<td>Crest Optics</td>
			<td>V2 L-FOV</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The presented technique works well on the unwounded notum (Figure 1A-D), allowing for investigation of the development and homeostasis of the tissue, e.g., the formation of the polyploid mechanosensory bristle cells18, or the anterior to the posterior flow of epithelial cells10. This protocol is also applicable to a laser-ablated notum (Figure 1E-L), where the cellular response to injury can be analyzed live with endogenous fluorophores such as Histone2-EGFP (Figure 1B,F,J). Post-staining with immunohistochemistry (step 3) can reveal many features, such as Fasciclin III, which labels cell borders (Figure 1C,G,K). Additionally, quantitative stains such as DAPI (Figure 1A,E,I) can be used to assess DNA content changes, including wound-induced polyploidy22.
The current protocol is particularly beneficial as it can be used following long-term live imaging experiments. As pupae are immobile, they can be imaged for hours (Figure 4A,B). Importantly, this protocol does not cause considerable changes to the wound epithelium&#39;s overall architecture or morphology following dissection (Figure 4B,C). Thus, features within the tissue could be imaged long-term and then further investigated with immunohistochemistry or cellular stains.
Imaging deep within tissues is complex due to their opacity, and Drosophila are coated in a waxy cuticle8 making deep imaging even more difficult. However, with this technique, a dissected notum can be placed between two coverslips allowing for imaging of the epithelial monolayer from either side: the apical side through the cuticle (Figure 5A-D) and/or the basal side of the epithelium which faces the body cavity (Figure 5E-H). These different views are ideally suited to visualize different structures within the tissue. For instance, the apical view is ideal for visualizing epithelial cell borders and nuclei that lie just below the cuticle (Figure 5A-D). With the basal view, these apical signals are less visible (Figure 5E-H). However, basal structures are observed at the wound margin (Figure 5J,L yellow, white arrows). These basal structures are much brighter as there is less occlusion in the basal view than in the apical view.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63682/63682fig01.jpg" /> 
Figure 1: Dissected, fixed, and stained Drosophila pupal nota. (A-D) Unwounded notum. (E-H), Wounded notum 30 min after laser ablation. (I-L) Wounded notum 3 h after laser ablation. (A,E,I) DAPI stain shows nuclei. (B,F,J) Transgenic Histone2-EGFP, used in live imaging, is visible after fixing and staining. (C,G,K) The anti-FasIII antibody shows that antibody stains work well on the fixed notum. (D,H,L) Merged image. The images are captured with 40x objective using&#160;spinning disc microscopy, maximum intensity projections of Z-stacks are shown with 0.3 &#181;m Z-slices. A-D represents 263 Z-slices. E-H represents 195 slices. I-L represents 53 Z-slices. The orange dashed line denotes wound margin. Scale bar in L is 25 &#181;m, applicable to A-L. <a href="https://www.jove.com/files/ftp_upload/63682/63682fig01large.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/63682/63682fig02.jpg" /> 
Figure 2: Hand placement for notum dissection. (A-B) Left and right views of hand placement for holding the microdissection scissors. To prevent hand shaking, the neck of the scissors is placed against the middle finger of the non-dominant hand. Scissors need to be parallel to the microscope slide to avoid the warping of the notum tissue. <a href="https://www.jove.com/files/ftp_upload/63682/63682fig02large.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/63682/63682fig03.jpg" /> 
Figure 3: Correctly dissected notum vs. curled notum. (A) Pupal notum post dissection and cleaning uncurled and ready for fixation. (B) Pupal notum curled on itself after being removed from 1x PBS, thus rendering it unusable. <a href="https://www.jove.com/files/ftp_upload/63682/63682fig03large.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/63682/63682fig04.jpg" /> 
Figure 4: Pre and post-fixed images are largely similar. A pupal notum expressing Histone2-EGFP in the nuclei was imaged (A) live before ablation and (B) 3 h after laser-ablation. (C) The notum was retrieved, dissected, and fixed as mentioned in the protocol and re-imaged after fixing. The wound site is labeled with a red star. The images were captured with 40x objective using spinning disc microscopy, Z-slices were taken every 0.3 &#181;m. Maximum intensity projections of 34 Z-slices pre-wounding (A), 48 Z-slices 3 h after wounding (B), and 103 Z-slices post dissection, fixation, and staining (C) are shown. Scale bar in C is 25 &#181;m, applicable to A-C. <a href="https://www.jove.com/files/ftp_upload/63682/63682fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63682/63682fig05.jpg" /> 
Figure 5: Imaging apical and basal sides of a wounded notum. The same wounded pupal notum is shown in all panels. The wound is marked with a red star. The orange dotted line indicates the wound margin. (A-D) The notum is imaged from the apical side, through the cuticle. (E-H) The notum is imaged from the basal interior side. (I-L) The top panels show an X-Z image of the notum, apical side up, with the epithelial sheet designated by a white triangle. The orange line denotes the apical-basal plane of the X-Y slice, shown in the lower panels. Within the lower panels, the orange line shows the plane of the above X-Z image. (I) Apically imaged notum - apical slice. The top X-Z view shows precise imaging of the apical epithelial sheet (white triangle). This view is equivalent to a live imaging view. (J) Apically imaged notum - basal slice, partially obstructed by apical tissue. Other tissues denoted by yellow arrow (possible muscle band) and white arrows (possible blood cells) are visible on the basal side. (K) Basally imaged notum - apical slice, partially obstructed by basal tissue and wound scab (dark central area). (L) Basally imaged notum - basal slice. This view best shows the basal tissues denoted with yellow and white arrows. A,E: DAPI stain, B,F: Histone-EGFP, C,G: FasIII staining. The images were captured with 20x objective with spinning disc microscopy. Z-slices were taken every 0.9 &#181;m. A-D shows the maximum intensity projection of 19 slices. E-H represents the maximum intensity projection of 17 slices. 25 &#181;m scale bars in D,H,L apply to A-D, E-H, I-L, respectively. <a href="https://www.jove.com/files/ftp_upload/63682/63682fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: Preparation of solutions and buffers. Recipes for the following solutions are detailed: 1xPBS, 1x PBST, blocking solution, and paraformaldehyde fixative. <a href="https://www.jove.com/files/ftp_upload/63682/Supplementary File 1.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum at JoVE.com

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.

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 ...

dissecting-fixing-and-visualizing-the-drosophila-pupal-notum

Research

JoVE Journal

Biology

3.0K Views.  Vanderbilt University. 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. 

Video: Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

In situ Protocol for Butterfly Pupal Wings Using Riboprobes

Here we present, in video format, a protocol for in situ hybridizations in pupal wings of the butterfly Bicyclus anynana using riboprobes.  In situ hybridizations, a mainstay of developmental biology, are useful to study the spatial and temporal patterns of gene expression in developing tissues at the level of transcription. If antibodies that target the protein products of gene transcription have not yet been developed, and/or there are multiple gene copies of a particular protein in the genome that cannot be differentiated using available antibodies, in situs can be used instead. While an in situ technique for larval wing discs has been available to the butterfly community for several years, the current protocol has been optimized for the larger and more fragile pupal wings.

In order to examine gene expression in the pupal wing tissue of Bicyclus anynana, we present an optimized protocol for in situ hybridizations using riboprobes. We also provide guidelines for the further optimization of this protocol for use in pupal wings of other Lepidopteran species.

Here we present, in video format, a protocol for in situ hybridizations in pupal wings of the butterfly Bicyclus anynana using riboprobes.  In situ ...

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Dissecting and Recording from The C. Elegans Neuromuscular Junction

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale. This complex process requires the coordinated activity of many pre- and post-synaptic proteins to ensure appropriate synaptic connectivity, conduction of electrical signals, targeting and priming of secretory vesicles, calcium sensing, vesicle fusion, localization and function of postsynaptic receptors and finally, recycling mechanisms. As neuroscientists it is our goal to elucidate which proteins function in each of these steps and understand their mechanisms of action. Electrophysiological recordings from synapses provide a quantifiable read out of the underlying electrical events that occur during synaptic transmission. By combining this technique with the powerful array of molecular and genetic tools available to manipulate synaptic proteins in C. elegans, we can analyze the resulting functional changes in synaptic transmission. 
The C. elegans NMJs formed between motor neurons and body wall muscles control locomotion, therefore, mutants with uncoordinated locomotory phenotypes (known as unc s) often perturb synaptic transmission at these synapses 1. Since unc mutants are maintained on a rich supply of a bacterial food source, they remain viable as long as they retain some pharyngeal pumping ability to ingest food. This, together with the fact that C. elegans exist as hermaphrodites, allows them to pass on mutant progeny without the need for elaborate mating behaviors. These attributes, coupled with our recent ability to record from the worms NMJs 2,3,7 make this an excellent model organism in which to address precisely how unc mutants impact neurotransmission. 
The dissection method involves immobilizing adult worms using a cyanoacrylic glue in order to make an incision in the worm cuticle exposing the NMJs. Since C. elegans adults are only 1 mm in length the dissection is performed with the use of a dissecting microscope and requires excellent hand-eye coordination. NMJ recordings are made by whole-cell voltage clamping individual body wall muscle cells and neurotransmitter release can be evoked using a variety of stimulation protocols including electrical stimulation, light-activated channel-rhodopsin-mediated depolarization 4 and hyperosmotic saline, all of which will be briefly described.

Application of electrophysiology to accessible synapses provides a quantifiable measure of synaptic activity, useful in analyzing synaptic mutants. This article describes a dissection method used to expose the neuromuscular junctions (NMJ) of Caenorhabditis elegans (C. elegans) and briefly discusses some of the uses to which this preparation can be applied.

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale.   This ...

Dissecting the Non-human Primate Brain in Stereotaxic Space

The use of non-human primates provides an excellent translational model for our understanding of developmental and aging processes in humans1-6. In addition, the use of non-human primates has recently afforded the opportunity to naturally model complex psychiatric disorders such as alcohol abuse7. Here we describe a technique for blocking the brain in the coronal plane of the vervet monkey (Chlorocebus aethiops sabeus) in the intact skull in stereotaxic space. The method described here provides a standard plane of section between blocks and subjects and minimizes partial sections between blocks. Sectioning a block of tissue in the coronal plane also facilitates the delineation of an area of interest. This method provides manageable sized blocks since a single hemisphere of the vervet monkey yields more than 1200 sections when slicing at 50&mu;m. Furthermore by blocking the brain into 1cm blocks, it facilitates penetration of sucrose for cyroprotection and allows the block to be sliced on a standard cryostat.

The non-human primate is an important translational species for our understanding of the normal processing of the brain. The anatomical organization of the primate brain can provide important insights into normal and pathological conditions in humans.

The use of non-human primates provides an excellent translational model for our understanding of developmental and aging processes in humans1-6. In ...

The Gateway to the Brain: Dissecting the Primate Eye

The visual system in humans is considered the gateway to the world and plays a principal role in the plethora of sensory, perceptual and cognitive processes. It is therefore not surprising that quality of vision is tied to quality of life . Despite widespread clinical and basic research surrounding the causes of visual disorders, many forms of visual impairments, such as retinitis pigmentosa and macular degeneration, lack effective treatments. Non-human primates have the closest general features of eye development to that of humans. Not only do they have a similar vascular anatomy, but amongst other mammals, primates have the unique characteristic of having a region in the temporal retina specialized for high visual acuity, the fovea1. Here we describe a general technique for dissecting the primate retina to provide tissue for retinal histology, immunohistochemistry, laser capture microdissection, as well as light and electron microscopy. With the extended use of the non-human primate as a translational model, our hope is that improved understanding of the retina will provide insights into effective approaches towards attenuating or reversing the negative impact of visual disorders on the quality of life of affected individuals.

The non-human primate is an important translational species for our understanding of development and aging. The anatomical organization of the primate retina may provide important insights into normal and pathological conditions in humans.

The visual system in humans is considered the  gateway  to the world and plays a principal role in the plethora of sensory, perceptual and cognitive ...

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes differentiation from a stem cell population, mitotic amplification, and meiosis. In addition, post-meiotic germ cells undergo a dramatic morphological reshaping process as well as a global epigenetic reconfiguration of the germ line chromatin&#8212;the histone-to-protamine switch.
Studying the role of a protein in post-meiotic spermatogenesis using mutagenesis or other genetic tools is often impeded by essential embryonic, pre-meiotic, or meiotic functions of the protein under investigation. The post-meiotic phenotype of a mutant of such a protein could be obscured through an earlier developmental block, or the interpretation of the phenotype could be complicated. The model organism Drosophila melanogaster offers a bypass to this problem: intact testes and even cysts of germ cells dissected from early pupae are able to develop ex vivo in culture medium. Making use of such cultures allows microscopic imaging of living germ cells in testes and of germ-line cysts. Importantly, the cultivated testes and germ cells also become accessible to pharmacological inhibitors, thereby permitting manipulation of enzymatic functions during spermatogenesis, including post-meiotic stages.
The protocol presented describes how to dissect and cultivate pupal testes and germ-line cysts. Information on the development of pupal testes and culture conditions are provided alongside microscope imaging data of live testes and germ-line cysts in culture. We also describe a pharmacological assay to study post-meiotic spermatogenesis, exemplified by an assay targeting the histone-to-protamine switch using the histone acetyltransferase inhibitor anacardic acid. In principle, this cultivation method could be adapted to address many other research questions in pre- and post-meiotic spermatogenesis.

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

This protocol describes the dissection and cultivation of intact testes and germ-line cysts from Drosophila melanogaster pupae. This method allows microscopic observation of spermatogenesis ex vivo. Furthermore, we describe a pharmacological assay of the effect of inhibitors on specific stages of germ-cell development in pupal testes.

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes ...

Dissection and Mounting of Drosophila Pupal Eye Discs

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes. It contains approximately 800 separate eye units, termed ommatidia1. Each ommatidium contains eight neuronal photoreceptors that develop from undifferentiated cells following the passage of the morphogenetic furrow in the third larval instar2. Following the sequential differentiation of the photoreceptors, non-neuronal cells develop, including cone and pigment cells, along with mechanosensory bristle cells3. Final differentiation processes, including the structured arrangement of all the ommatidial cell types, programmed cell death of undifferentiated cell types and rhodopsin expression, occurs through the pupal phase4-7. This technique focuses on manipulating the pupal eye disc, providing insight and instruction on how to dissect the eye disc during the pupal phase, which is inherently more difficult to perform than the commonly dissected third instar eye disc. This technique also provides details on immunostaining to allow the visualization of various proteins and other cell components.

Dissection and Mounting of Drosophila Pupal Eye Discs

The goal of this technique is to enable researchers to perform dissection, immunostaining and mounting of pupal eye discs from Drosophila melanogaster of any age.

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes.  It contains approximately 800 ...