We are grateful to Ruth Lehman and Dorthea Godt who kindly supplied Vasa and Traffic Jam antibodies. We would like to acknowledge the Bloomington Stock Center at Indiana University for maintaining the provided stocks and the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by The University of Iowa. We thank all members of the Wawersik lab for their advice and support. This work was funded by the Monroe Scholars Program Grant (to AF and LB) and NSF grant IOS0823151 (to MW).

1. Preparation of a Collection Cage
<ol>
	<li>Anesthetize young, fertile flies with CO2. Transfer anesthetized flies to a cage. To obtain optimal yield, use 100-120 adult flies ranging from 2-7 days of age at a 4:1 ratio of females to males. Allow flies an appropriate acclimation period, ~24 hr prior to obtaining sample for fixation. If the cage was set up with virgin females mated to males, a use a 36 - 48 hr&#160;acclimation period.</li>
	<li>On the open end of the cage, place a pre-prepared apple juice agar plate with a dime-sized drop of yeast paste in the center over the opening of the cage. Then, secure with tape. Seal the junction between the apple juice agar plate and the cage with Parafilm.</li>
	<li>Place the cage in a secondary container with apple juice agar plate as the base and allow the flies to reproduce at a defined temperature for an appropriate period of time as determined by the experiment. 
	Note: Sample collection times will vary. For example, when collecting late stage embryos/early L1 larvae (17 - 24 hr), allow flies to lay eggs on apple juice agar plates for 7 hr at 25 &#176;C prior to aging of sample for 17 hr at 25 &#176;C. Similarly, for a collection of mid-late first instar larvae (36 - 48 hr) allow flies to lay eggs for 12 hr at 25 &#176;C prior to aging of sample for 36 hr at 25 &#176;C.</li>
	<li>Once the time period is complete, tap the cage on a table so that flies drop away from the apple juice agar plate without anesthesia. Quickly replace the used plate with a fresh one containing a drop of yeast paste. Secure the fresh plate with tape and parafilm and store the cage with apple juice agar plate as base.</li>
	<li>Carefully remove the yeast drop from the used plate with a metal spatula. Place lid on used apple juice plate and allow laid embryos to age if experimentally necessary. 
	Note: While aging samples, plates may be stored at 25 &#176;C unless higher temperatures are experimentally required. Examples of how to age embryos for collection are described above (see note for step 1.3). Removal of yeast paste prior to sample aging is performed to prevent larvae from crawling into the yeast and breaking up the agar underneath. The remaining apple juice agar and yeast paste residue provide nutrients for larval growth. While embryos laid directly in the yeast paste are lost through yeast paste removal, this loss is preferable to yeast and agar residue in the sample that can reduce staining efficiency. Should one choose not to remove the yeast paste, yeast may be liquefied with Phosphate Buffer Triton X-100 (PBTx), mixed with a paint brush, and then removed from the sample with a cell strainer prior to fixation.</li>
</ol>
2. Fixation
<ol>
	<li>Once embryos laid onto the apple juice agar plate have aged to the desired time point, use a small paintbrush wetted with Phosphate Buffer Triton X-100 (PBTx) solution to carefully remove sample from the plate. 
	Note: Older larvae are mobile and may crawl onto the inside of the lid. This sample may be collected similarly by removal with a paintbrush.</li>
	<li>Wipe the sample-laden paintbrush against the interior-facing ridge of a cell strainer. Then, rinse the walls of the strainer and the paintbrush with a squirt bottle containing PBTx solution. Place a petri dish cover underneath the strainer to capture the PBTx solution.</li>
	<li>After all the sample has been transferred to the strainer, pour enough PBTx into the strainer to raise the sample off the mesh. Then, lift the strainer and pour the contents of the petri dish into a liquid waste container.</li>
	<li>Repeat Step 2.3 three times to remove yeast and fly waste that may have been transferred along with the sample.</li>
	<li>Pour enough 50% bleach (NaOCl)/water (ddH2O) solution into the strainer to raise the larvae off the mesh. Allow larvae to sit in solution for 5 min. Then, lift the strainer and pour out the contents of the petri dish in a liquid waste container. 
	Note: Bleach is required only in embryonic samples aged 0 - 22 hr in order to remove the chorionic membrane. The application of bleach to older samples can be omitted, but its inclusion is not detrimental. If omission is desired, replace the bleach in this step with PBTx. Diluted bleach should be used within 24 hr of solution preparation.</li>
	<li>Wash sample in the strainer with PBTx by pouring enough PBTx into the strainer to raise the sample off the mesh. Allow sample to sit in solution for 3 min. Then, lift the strainer and pour the contents of the petri dish into a liquid waste container.</li>
	<li>Repeat Step 2.6 five times.</li>
	<li>Dab the bottom of cell strainer dry, then transfer sample into a scintillation vial containing 1.75 ml of PEMS solution using a paintbrush.</li>
	<li>Working in a ventilated fume hood, add 250 &#956;l of 37% formaldehyde and 8 ml of reagent grade heptane to the scintillation vial. 
	Note: Formaldehyde and heptane are hazardous. For safety purposes, continue to work in a ventilated fume hood and wear appropriate gloves for steps 2.9 &#8211; 3.3.</li>
	<li>Allow the vial to shake at 200 rpm or a similar moderate speed for 20 min.</li>
	<li>Add 10 ml of methanol to scintillation vial without removing the previous aqueous phase and allow the vial to shake vigorously at 500 rpm for 1 min. 
	Note: Methanol is hazardous. Continue to work in a ventilated fume hood and wear appropriate gloves.</li>
	<li>Remove vial from shaker and immediately pour contents into a cell strainer over a liquid waste container in the hood. Add extra methanol to the vial and run through the cell strainer as necessary to ensure all sample has been removed. 
	Note: Cell strainers may drain slowly. To remedy this, touch a laboratory tissue to the bottom of the cell strainer in order to draw the solution through the strainer more quickly. Methanol, formaldehyde, and heptane should be stored in appropriate waste containers until proper disposal as per institutional guidelines.</li>
	<li>Dry bottom of cell strainer using a laboratory tissue, then use a paintbrush to transfer sample captured in the strainer to a 1.5 ml microcentrifuge tube containing 0.5 ml of methanol. 
	Note: If multiple scintillation vials are in use, complete steps 2.12 and 2.13 one vial at a time to prevent sample from drying on cell strainers.</li>
	<li>Once the sample has been added to the microcentrifuge tube, remove methanol with a pipette. Ensure sample has settled to the bottom of the tube.</li>
	<li>Rinse sample in the microcentrifuge tube by adding approximately 0.5 ml of methanol to tube. Wait for sample to settle then remove methanol with a pipette.</li>
	<li>Repeat Step 15 three times.</li>
	<li>Add 0.5 ml of methanol to the tube, and then store in a -20 &#176;C freezer for later use.</li>
</ol>
3. Rehydration and Preparation of Sample for Immunostaining
<ol>
	<li>Remove methanol from the microcentrifuge tube with a pipette, leaving the sample in the tube. Store methanol waste to the appropriate waste container until time of proper disposal. 
	Note: To enhance sonication efficiency, use no more than 3 mm of sample settled at the bottom of the 1.5 ml microcentrifuge tube. Excess sample may be transferred to a separate tube with a pipette before beginning the rehydration step above. Cutting the pipette tip with a clean razor blade can be used to prevent clogging of the pipette tip.</li>
	<li>Add&#160;1.0 ml of a 50% methanol/Phosphate Buffer Tween (PBTw) solution to the tube and rock for 3 min.</li>
	<li>Remove methanol solution to the proper waste container and rinse with 1.0 &#956;l of PBTw twice. After each rinse, allow the sample to settle before drawing off the PBTw with a pipette.</li>
	<li>Add&#160;1.0 ml of Bovine Serum Albumin/Phosphate Buffered Tween (BBTw) mix to the vial and rock. After 3 min on a rocker, allow the sample to settle and remove the BBTw with pipette.</li>
	<li>Repeat Step 3.4 two times taking care to remove as much BBTw as possible without losing sample after the last wash.</li>
	<li>Add&#160;0.5 ml of BBTw to the tube and place the tube on ice.</li>
</ol>
4. Sonication of Sample
<ol>
	<li>Set the sonicator to 10% maximum amplitude and designate a run time of 2 sec constant sonication. 
	Note: These settings are specific to the sonicator indicated in the Materials and Equipment Table and may require optimization for different sonicators.</li>
	<li>Prior to sonication of sample, clean the sonicator probe by placing the probe tip into a 50 ml conical tube filled with deionized water and run sonicator to clean the probe. Dry sonicator probe with laboratory tissue after run.</li>
	<li>Submerge the probe into the microcentrifuge tube containing sample so that it is positioned approximately 3 to 4 mm above the sample and start sonication.</li>
	<li>Remove the microcentrifuge tube and place it on ice for at least 30 sec to dissipate heat from sonication and allow sample to settle at the bottom of the microcentrifuge tube.</li>
	<li>Repeat steps 4.3 and 4.4 as needed based on age of sample being processed. 
	Note: For optimum sonication sample volume, embryos younger than 17 hr do not require sonication, but those between 17 and 24 hr (stage 17/early-L1) require two sonications. Larvae between 24 - 36 (early/mid-L1), 36 - 48 (mid/late-L1), 48 - 60 (early/mid-L2), 60 - 72 (mid/late-L2), 72 - 84 (early-L3), 84 - 96 (early/mid-L3), 96 - 108 (mid/late-L3) hr require 5, 9, 11, 13, 15, 18, and 22 sonications respectively. See discussion for further elaboration on sonication times.</li>
	<li>Rinse with&#160;1.0 ml BBTw twice. After each rinse allow the sample to settle before drawing off BBTw with a pipette.</li>
	<li>Add 1.0 ml of BBTw to the vial and rock. After 3 min on the rocker, allow the sample to settle and remove the BBTw with pipette.</li>
	<li>Repeat Step 4.7 two times.</li>
</ol>
5. Immunostaining
<ol>
	<li>Block sample by adding 5% normal serum diluted in BBTw to the microcentrifuge tube. Remove block after rocking for 1 hr. 
	Note: Use normal serum from the species in which the secondary antibodies are generated.</li>
	<li>Dilute primary antibodies to appropriate working concentration in 5% normal serum/BBTw solution.</li>
	<li>Rock sample in primary antibody solution O/N at 4 &#176;C or for at least 3 hr at RT (approximately 22 &#176;C. 
	Note: Antibody incubation times and temperatures may vary. O/N incubation at 4 &#176;C or 3 hr at RT is typically sufficient. However, a two-day incubation period may enhance staining quality, especially with older samples or when low-affinity primary antibodies are used. Longer incubations are typically performed at 4 &#176;C. Similar considerations must be made when using secondary antibodies (see 5.10).</li>
	<li>Allow sample to settle to bottom of microfuge tube. Draw off primary antibodies with pipette. Save antibodies for reuse if desired.</li>
	<li>Rinse with&#160;1.0 ml of BBTw twice. After each rinse allow sample to settle before drawing off BBTw with a pipette.</li>
	<li>Add 1.0 ml&#160;of BBTw to the vial and rock. After 3 min on the rocker, allow sample to settle and remove BBTw with pipette.</li>
	<li>Repeat Step 5.6 five times.</li>
	<li>Block sample by adding 5% normal serum/BBTw solution to the microcentrifuge tube. Remove block after rocking for 30 min to 1 hr. 
	Note: This additional blocking step is not required, but may improve staining quality for specific antibodies.</li>
	<li>Dilute secondary antibodies to appropriate working concentration in 5% normal serum/BBTw solution.</li>
	<li>Wrap microcentrifuge tube in aluminum foil to reduce fading due to light exposure. Rock sample in secondary antibody solution for 12 to 48 hr at 4 &#176;C or for at least 3 hr at RT (approximately 22 &#176;C.</li>
	<li>Allow sample to settle to bottom of microfuge tube. Draw off secondary antibodies with pipette.</li>
	<li>Rinse with&#160;1.0 ml of PBTw twice. After each rinse allow sample to settle before drawing off the PBTw with a pipette.</li>
	<li>Add 1.0 ml of PBTw to the vial and rock. After 5 min on the rocker, allow the sample to settle and remove the PBTw with pipette.</li>
	<li>Repeat Step 5.13 three times.</li>
	<li>OPTIONAL: Add DAPI diluted 1:1,000 in PBTw. Rock sample wrapped in aluminum foil for 3 min; then remove DAPI solution with a pipette. Rinse five times with 1.0 ml of PBTw, allowing sample to settle before removing PBTw with pipette.</li>
	<li>Add 80 - 100 &#956;l of 1,4-diazabicyclo[2.2.2]octane (DABCO) solution to microcentrifuge tube and store at -20 &#176;C. 
	Note: Another glycerol-based anti-fade agent may be substituted for .</li>
</ol>
6. Analysis
<ol>
	<li>Before mounting sample onto slides, add an extra 5 - 10 &#956;l of DABCO/p-phenylenediamine (PPD) antifade solution to microcentrifuge tube. 
	Note: Sample can be added to slides without dissection. Sample volume added to a slide varies with cover slip size. When pipetting sample onto slide, the pipette tip may be cut off using a razor blade to prevent sample shearing. It is often useful to line sample up in rows for easy analysis. The addition of PPD as an additional antifade agent may not be required, but can prevent photobleaching if samples are stored long-term.</li>
	<li>Gently place glass cover slip over sample. Then, secure cover slip in place using nail polish.</li>
	<li>View mounted sample using fluorescence microscopy.</li>
</ol>

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The authors have nothing to disclose.

This protocol provides a method to successfully immunostain target Drosophila embryonic and larval tissues in situ, thus eliminating the need for dissection. As per prior protocols for staining early embryos1,2,3, the chorionic membrane is removed using 50% bleach (NaOCl). Samples are fixed in formaldehyde and methanol. Because the larval cuticle causes older sample to float, the entire sample is then passed through a cell-strainer to ensure larval retention. Sample is stored, if desired, in ...

Drosophila embryos and larvae provide an excellent model to study developmental processes in many organs and tissues. Imaging of individual cells is often necessary in these studies in order to ascertain the complex environments in which cells develop. Visualization of cells in tissues can be accomplished through immunostaining. Well-described immunostaining protocols exist for embryonic Drosophila tissues &#60;17 hr after egg laying (AEL)1-3. However, a protective cuticle forms toward the end of embryogenesis, preventing effective antibody permeation. Thus, these immunostaining protocols are inefficient in the analysis of tissues in late-stage embryos and in subsequent stages of larval development (1st instar (L1), 2nd instar (L2), and 3rd instar (L3)). This inefficiency imposes a barrier to our understanding of dynamic processes that occur during this extended developmental period 4. Tissue dissection is a widely employed technique to circumvent this barrier 5-7. However, dissection can prove inefficient. Extraction may be encumbered by difficulty in locating or isolating embryonic and larval tissues. Furthermore, the physical removal of target tissues may cause damage by rupturing them or by failing to extract them in their entirety.
Sonication is a method that employs sound waves to disturb intermolecular interactions. It has been used to disrupt the integrity of the Drosophila larval cuticle in order to immunostain developing neural cell types 6. This protocol has been adapted to immunostain late-stage embryonic and larval gonads, which can be as small as 50&#956;m in diameter 8-10. Through such studies, the process of male germline stem cell (GSC) niche formation has been characterized in late stage Drosophila embryos8-10 and mechanisms regulating stem cell development and differentiation in late stage embryonic gonads and larvae have been elucidated 9-12. Thus, sonication provides an efficient alternative to tissue dissection that may be difficult because of tissue size. Furthermore, it enables immunostaining of Drosophila tissues in situ, leaving cells within the context of the entire organism and maintaining in situ morphology. Here, we describe a step-by-step protocol for fluorescence immunostaining of late-stage embryonic through early/mid-L3 tissues in situ. Analysis of Drosophila gonadal and neural tissue is shown in the Representative Results to demonstrate the efficacy of this protocol. Furthermore, this immunostaining protocol may be adapted to analyze other Drosophila tissues as well as tissues in other organisms with an outer cuticle.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Table 1: Reagents and Buffers</td>
		</tr>
		<tr>
			<td>Phosphate Buffer Triton X-100 (PBTx)</td>
			<td>For 5 L: 500 mL PBS 10X 4.45 L ddH2O 50 mL Triton 10%</td>
			<td>Store at room temperature.</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffer Saline 10x (PBS 10X)</td>
			<td>For 1 L in dH2O: 80 g NaCl 2 g KCl 14.4g Na2HPO4 2.4 g KH2PO4</td>
			<td>Add components and fill to appropriate volume. Store at 4 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton 10%</td>
			<td>For 50 mL: 5 mL of Triton 5 mL of PBS 10X 45 mL of ddH2O</td>
			<td>Rock to mix. Store at room temperature.</td>
			<td></td>
		</tr>
		<tr>
			<td>PEMS</td>
			<td>0.1 M Pipes (pH 6.9) 2.0 mM MgSO4 1.0 mM EGTA</td>
			<td>Store at room temperature.</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipes</td>
			<td>For a 400ml of a 0.25 M solution (pH 6.9): 30.24 g Pipes dH20 NaOH</td>
			<td>Dissolve Pipes in 300 mL dH2Oand then adjust to pH 6.9 with NaOH. Bring the total volume to 400 mL with dH2O and autoclave. Store at room temperature.</td>
			<td></td>
		</tr>
		<tr>
			<td>Formaldehyde</td>
			<td>37% formaldehyde by weight in methanol</td>
			<td>Store at room temperature. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.</td>
			<td></td>
		</tr>
		<tr>
			<td>Heptane</td>
			<td>n-Heptane</td>
			<td>CAS 142-82-5. Store at room temperature. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Methanol</td>
			<td>CAS 67-56-1. Store at room temperature. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffer Tween (PBTw)</td>
			<td>To make 1 L: 100 mL PBS 10X 890 mL ddH2O 10 mL Tween 10%</td>
			<td>Filter sterilize after adding all components. Store at 4 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 10%</td>
			<td>For 50 mL: 5 mL Tween 5 mL of PBS 10X 40 mL of ddH2O</td>
			<td>Rock to mix. Store at room temperature.</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin/Phosphate Buffer Tween (BBTw)</td>
			<td>To make 1 L: 100 mL PBS 10X 890 mL ddH2O 10 mL Tween 10% 1 g Bovine Serum Albumin (BSA)</td>
			<td>Add BSA then sterilize using a 0.2 micrometer vacuum filter unit. Store at 4 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal Goat Serum (NGS)</td>
			<td>To make 10 mL: Normal Goat Serum (Jackson ImmunoResearch Laboratories Code: 005-000-121) 10 mL ddH2O</td>
			<td>Add ddH2O to vial of NGS and sterilize using a 0.2 micrometer syrninge filter. Store aliquots at -20 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>1,4-diazabicyclo[2.2.2]octane (DABCO)</td>
			<td>To make 100 mL: 25 mL ddH2O 1 mL Tris HCl (1M, pH 7.5) 2.5 g of DABCO solid (CAS: 281-57-9) 3.5 mL 6N HCl 250 uL 10N NaOH 70 mL glycerol</td>
			<td>In 250 mL beaker with stir bar, add ddH2O, Tris HCl and DABCO. Stir and then add 6N HCl, 10 N NaOH, and glycerol. Then add 10NaOH dropwise until solution reaches pH 7.5. Aliquot. Store aliquots at -20 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>DABCO + p-phenylenediamine (PPD) Solution</td>
			<td>1.765 mL NaHCO3 0.353 Na2CO3 0.02 g PPD (CAS: 106-50-3)</td>
			<td>Dissolve PPD in NaHCO3 and NaCO3 solution. Add 60 uL of PPD solution to 500 uL of DABCO. Store aliquots at -20 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>Apple juice plates</td>
			<td>To make ~200 plates: 45 g agar (CAS#9002-18-0) 45 g granulated sugar (store bought) 500 mL Apple juice (store bought) 15 mL Tegosept 10% 1.5 mL ddH2O</td>
			<td>Add agar to ddH2O in 4L flask then autoclave for 30 minuntes. Mix apple juice and sugar on heated stir plate. Gradually add apple juice mixture to autoclaved agar. Mix on heated stir plate then aliquot 10mL volumes into 35 mm petri dishes and let stand at room temperature to solidfy. Store at 4 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td>Tegosept 10%</td>
			<td>To make 100mL: 10g Tegosept 100 mL Ethanol</td>
			<td>Store aliquots at -20 degrees C</td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast paste</td>
			<td>~50 g Dry Active Yeast</td>
			<td>Gradually add ddH2O to beaker containing yeast while stirring until paste-like consistency reached. Store at 4 degrees C.</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Table 2: Staining Materials</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>1:1000</td>
			<td>Invitrogen</td>
			<td>D3571 //// Stock at 5mg/mL</td>
		</tr>
		<tr>
			<td>rabbit anti-Vasa</td>
			<td>1:250</td>
			<td></td>
			<td>A gift from Ruth Lehmann</td>
		</tr>
		<tr>
			<td>mouse anti-Fasciclin III</td>
			<td>1:10</td>
			<td>Developmental Studies Hybridoma Bank (DSHB)</td>
			<td>7G10</td>
		</tr>
		<tr>
			<td>mouse anti-1B1</td>
			<td>1:4</td>
			<td>Developmental Studies Hybridoma Bank (DSHB)</td>
			<td>1B1</td>
		</tr>
		<tr>
			<td>guniea pig anti-Traffic Jam</td>
			<td>1:2500</td>
			<td></td>
			<td>A gift from Dorthea Godt (Li et al, 2003)</td>
		</tr>
		<tr>
			<td>mouse anti-Prospero</td>
			<td>1:10</td>
			<td>Developmental Studies Hybridoma Bank (DSHB)</td>
			<td>Prospero MR1A</td>
		</tr>
		<tr>
			<td>rat anti-Elav</td>
			<td>1:30</td>
			<td>Developmental Studies Hybridoma Bank (DSHB)</td>
			<td>Rat-elav 7EBA10 anti-elav</td>
		</tr>
		<tr>
			<td>mouse anti-Repo</td>
			<td>1:10</td>
			<td>Developmental Studies Hybridoma Bank (DSHB)</td>
			<td>8D12 anti-Repo</td>
		</tr>
		<tr>
			<td>goat anti-rabbit Alexa546</td>
			<td>1:500</td>
			<td>Invitrogen</td>
			<td>A11010</td>
		</tr>
		<tr>
			<td>goat anti-mouse Alexa488</td>
			<td>1:500</td>
			<td>Invitrogen</td>
			<td>A11029</td>
		</tr>
		<tr>
			<td>goat anti-guniea pig Alexa633</td>
			<td>1:500</td>
			<td>Invitrogen</td>
			<td>A21105</td>
		</tr>
		<tr>
			<td>goat anti-rat Alexa488</td>
			<td>1:500</td>
			<td>Invitrogen</td>
			<td>A11006</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Table 3: Materials and Equipment</td>
		</tr>
		<tr>
			<td>Fly Cages</td>
			<td>Hand-made; Genesee Scientific Corporation</td>
			<td>Not applicable; Bottles: 32-130; Pre-made cage: 59-101</td>
			<td>Made by cutting clear cast acrylic tubing (1 3/4 inch in diameter) into 4 inch tall segments with a compound miter saw at 400 rpm. Ultrafine stainless steel screening (was attached to one end of the tub with acrylic compund glue. An alternate method using an empty fly food bottle can be found in Drosophila Protocols ISBN 0-87969-584-4. Cages may also be purchased from the Genesee Scientific Corporation.</td>
		</tr>
		<tr>
			<td>Sonicator: Branson 250 Digital Sonifier</td>
			<td>Branson</td>
			<td>Model: Branson Digital Sonifier 250</td>
			<td></td>
		</tr>
		<tr>
			<td>Sonicator Probe</td>
			<td>Branson</td>
			<td>Model #: 102C (CE)</td>
			<td>EDP: 101-135-066; S/N: OBU06064658</td>
		</tr>
		<tr>
			<td>Syringe filter</td>
			<td>Nalgene</td>
			<td>190-25-20</td>
			<td>0.2 micrometer cellulose, acetate membrane filter</td>
		</tr>
		<tr>
			<td>Imaging system: Spinning disc confocal microscope with multichromatic light source, digital CCD camera, and imaging software</td>
			<td>Microscope: Olympus Light source: Lumen Dynamics Camera: Q-Imaging Imaging Software: Intelligent Imaging Inc.</td>
			<td>Microscope: BX51 equipped with DSU spinning disc Light source: X-Cite 120Q Camera: RETIGA-SRV Imaging Software: Slidebook 5.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum filter unit</td>
			<td>Nalgene</td>
			<td>450-0020</td>
			<td>0.2 micrometer cellulose nitrate membrane filter</td>
		</tr>
	</tbody>
</table>

sonication-facilitated immunofluorescence staining of late-stage embryonic and larval drosophila tissues in situ

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and organogenesis. Immunostaining allows for the visualization of developing tissues and organs. However, a protective cuticle that forms at the end of embryogenesis prevents permeation of antibodies into late-stage embryos and larvae. While dissection prior to immunostaining is regularly used to analyze Drosophila larval tissues, it proves inefficient for some analyses because small tissues may be difficult to locate and isolate. Sonication provides an alternative to dissection in larval Drosophila immunostaining protocols. It allows for quick, simultaneous processing of large numbers of late-stage embryos and larvae and maintains in situ morphology. After fixation in formaldehyde, a sample is sonicated. Sample is then subjected to immunostaining with antigen-specific primary antibodies and fluorescently labeled secondary antibodies to visualize target cell types and specific proteins via fluorescence microscopy. During the process of sonication, proper placement of a sonicating probe above the sample, as well as the duration and intensity of sonication, is critical. Additonal minor modifications to standard immunostaining protocols may be required for high quality stains. For antibodies with low signal to noise ratio, longer incubation times are typically necessary. As a proof of concept for this sonication-facilitated protocol, we show immunostains of three tissue types (testes, ovaries, and neural tissues) at a range of developmental stages.

To demonstrate the efficacy of sonication-based immunostaining in analysis of late-stage embryonic and larval tissues in situ, wild-type embryos and larvae were processed for immunostaining of testes, ovaries, and neural tissue. Samples were imaged via confocal microscopy and representative results are shown (Figure 1 and Figure 2). Results reveal that the described protocol is effective for visualizing morphological features as well as individual cells in situ during l...

Watch this Scientific Journal Video about Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ at JoVE.com

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and ...

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12.6K Views.  College of William & Mary. Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae. 

Video: Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

Dissection z larw CNS na Drosophila melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Drosophila Larval NMJ Immunohistochemistry

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 31 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. Immunohistochemistry can be used to clearly image the components of the NMJ. In this article, we demonstrate how to use antibody staining to visualize the Drosophila larval NMJ.

Drosophila Larval NMJ Immunohistochemistry

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Microdissection of Zebrafish Embryonic Eye Tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...

In Situ Immunofluorescent Staining of Autophagy in Muscle Stem Cells

Increasing evidence points to autophagy as a crucial regulatory process to preserve tissue homeostasis. It is known that autophagy is involved in skeletal muscle development and regeneration, and the autophagic process has been described in several muscular pathologies and age-related muscle disorders. A recently described block of the autophagic process that correlates with the functional exhaustion of satellite cells during muscle repair supports the notion that active autophagy is coupled with productive muscle regeneration. These data uncover the crucial role of autophagy in satellite cell activation during muscle regeneration in both normal and pathological conditions, such as muscular dystrophies. Here, we provide a protocol to monitor the autophagic process in the adult Muscle Stem Cell (MuSC) compartment during muscle regenerative conditions. This protocol describes the setup methodology to perform in situ immunofluorescence imaging of LC3, an autophagy marker, and MyoD, a myogenic lineage marker, in muscle tissue sections from control and injured mice. The methodology reported allows for monitoring the autophagic process in one specific cell compartment, the MuSC compartment, which plays a central role in orchestrating muscle regeneration.

In Situ Immunofluorescent Staining of Autophagy in Muscle Stem Cells

Active autophagy is associated with productive muscle regeneration, which is essential for Muscle Stem Cell (MuSC) activation. Here, we provide a protocol for the in situ detection of LC3, an autophagy marker in MyoD-positive MuSCs of muscle tissue sections from control and injured mice.

Increasing evidence points to autophagy as a crucial regulatory process to preserve tissue homeostasis. It is known that autophagy is involved in skeletal ...