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 chemical-grade methanol. After rehydration, a properly implemented sonication process disrupts the integrity of the larval cuticle. This is critical after cuticle formation to allow for sufficient permeation of antibodies required for immunostaining. Before applying antibodies, rocking the sample in a 5% normal serum/BBTw solution prevents excessive nonspecific antibody binding. Primary antibodies are then added to detect specific proteins expressed in the target tissue. After washing to remove unbound primary antibodies, secondary antibodies bound to fluorophores are used to mark the location of bound primary antibodies. Analysis can then be performed with epifluorescence or confocal microscopy.
Some modifications may be required to optimize results in individual circumstances. In particular, sonication power and the number of sonications performed may require adjustment depending on the sonicator used. Adaptations to the height of the sonicator probe above the sample as well as to solution volume may also be required for different sonicators or if sufficient sample cannot be obtained. Increased solution turbidity can be used as an indicator of tissue lysis and, therefore, provides a gauge for when sonication is too severe. If solution turbidity becomes high, the experimenter should consider reducing the number of sonications performed, increasing solution volume, raising the height of the sonicator probe above the sample, and/or reducing sonicator power.
Alterations may also be required in the immunostaining protocol to enhance image quality. As with most immunostaining procedures the signal to noise ratio can be altered by modifying antibody dilution, time and temperature of antibody incubation with sample, and/or blocking reagents used. While each antibody must be considered independently, especially with respect to antibody dilution, we find that incubation at 4 &#176;C for longer time periods can enhance staining quality after sonication, especially for antibodies with low signal to noise ratios and when older larvae are examined. In these instances, staining quality may also benefit from a slight reduction in secondary antibody dilution. In this protocol, we include both BSA and serum as blocking reagents, and the sample is re-blocked prior to addition of secondary antibodies in order to increase staining quality. Depending on the antibodies used, re-blocking and inclusion of both blocking reagents may or may not be required to enhance staining. If the re-blocking step is omitted, longer washes and more rinses may produce cleaner images. Finally, pre-absorbing polyclonal antibodies may increase the signal to noise ratio. Pre-absorption is performed by incubating the desired antibody with rehydrated embryos or larvae prior to use in an immunostain1.
As demonstrated in our representative results, sonication allows for clear visualization of target tissues in situ and thus provides a reasonable alternative to tissue dissection in immunostaining protocols. Dissection can be cumbersome in Drosophila embryos and larvae due to difficulties in locating, isolating, and extracting undamaged target tissues. Alternately sonication allows tissues to remain in the context of the entire organism. Because sonication avoids extracting and mounting tissues between a slide and a cover slip, it more accurately preserves in situ morphology. Practically, large amounts of sample can be processed more quickly for future analysis, since many organisms can be sonicated simultaneously.
Although sonication provides a great alternative to dissection, it has limitations that must be considered. Sonication beneficially disrupts the integrity of the larval cuticle but destroys some sample in the process (see Representative Results). Rinsing to remove biological debris is required following the sonication process, which can further reduce sample size. In addition, immunostaining whole embryos and larvae leaves the tissue of interest embedded in surrounding tissues. These surrounding tissues may stain positively or show non-specific antibody binding, thus reducing final image quality. Finally, staining efficiency can be variable. This variability may depend on age of sample, sonication efficacy, and antibody specificity. As a result, sonication-based immunostaining of whole-mount tissue may not always be preferable. In particular, dissection-based immunostaining may be more efficient when studying large tissues in late-L3 larvae. Furthermore, sonication can shear early- and mid-stage embryos. Despite these limitations, sonication-based immunostaining is a highly efficient technique that is well suited for analyzing development of a variety of tissues in late-stage embryos through early/mid-L3 larvae. In our lab, we regularly use this protocol to study gonad morphogenesis in Drosophila embryos and larvae9,10. Furthermore, we anticipate that implementation of this protocol will prove equally fruitful in studying development of other&#160;tissues in Drosophila and in other organisms with a protective cuticle.

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><tbody><tr><td colspan="4">Table 1: Reagents and Buffers</td></tr><tr><td>Phosphate buffer Triton X-100 (PBTx)</td><td></td><td></td><td>For 5 L: 500 ml PBS 10X, 4.45 L ddH&lt;sub&gt;2&lt;/sub&gt;O, 50 ml Triton 10%. Store at RT.</td></tr><tr><td>Phosphate buffer saline 10X (PBS 10X)</td><td></td><td></td><td>For 1 L in dH&lt;sub&gt;2&lt;/sub&gt;O: 80 g NaCl, 2 g KCl,14.4g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 2.4 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;. Add components and fill to appropriate volume. Store at 4 &amp;deg;C.</td></tr><tr><td>Triton 10%</td><td></td><td></td><td>For 50 ml: 5 ml of Triton, 5 ml of PBS 10X, 45 ml of ddH&lt;sub&gt;2&lt;/sub&gt;O. Rock to mix. Store at RT.</td></tr><tr><td>PEMS</td><td></td><td></td><td>0.1 M Pipes (pH 6.9), 2.0 mM MgSO&lt;sub&gt;4&lt;/sub&gt;, 1.0 mM EGTA. Store at RT.</td></tr><tr><td>Pipes</td><td></td><td></td><td>For a 400 ml of a 0.25 M solution (pH 6.9): 30.24 g Pipes dH&lt;sub&gt;2&lt;/sub&gt;0 NaOH. Dissolve Pipes in 300 ml dH&lt;sub&gt;2&lt;/sub&gt;O and then adjust to pH 6.9 with NaOH. Bring the total volume to 400 ml with dH&lt;sub&gt;2&lt;/sub&gt;O and autoclave. Store at RT.</td></tr><tr><td>Formaldehyde</td><td></td><td></td><td>37% formaldehyde by weight in methanol. Store at RT. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.</td></tr><tr><td>Heptane</td><td></td><td>CAS 142-82-5</td><td>n-Heptane. Store at RT. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.</td></tr><tr><td>Methanol</td><td></td><td>CAS 67-56-1</td><td>Store at RT. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.</td></tr><tr><td>Phosphate buffer Tween (PBTw)</td><td></td><td></td><td>To make 1 L: 100 ml PBS 10X, 890 ml ddH&lt;sub&gt;2&lt;/sub&gt;O, 10 ml Tween 10%. Filter sterilize after adding all components. Store at 4 &amp;deg;C.</td></tr><tr><td>Tween 10%</td><td></td><td></td><td>For 50 ml: 5 ml Tween, 5 ml of PBS 10X, 40 ml of ddH&lt;sub&gt;2&lt;/sub&gt;O. Rock to mix. Store at RT.</td></tr><tr><td>Bovine serum albumin/phosphate buffer Tween (BBTw)</td><td></td><td></td><td>To make 1 L: 100 ml PBS 10X, 890 ml ddH&lt;sub&gt;2&lt;/sub&gt;O, 10 ml Tween 10%, 1 g Bovine Serum Albumin (BSA). Add BSA then sterilize using a 0.2 &amp;mu;m vacuum filter unit. Store at 4 &amp;deg;C.</td></tr><tr><td>Normal goat serum (NGS)</td><td>ackson ImmunoResearch Laboratories</td><td>005-000-121</td><td>To make 10 ml: Normal goat serum 10 ml ddH&lt;sub&gt;2&lt;/sub&gt;O. Add ddH&lt;sub&gt;2&lt;/sub&gt;O to vial of NGS and sterilize using a 0.2 &amp;mu;m syrninge filter. Store aliquots at -20 &amp;deg;C.</td></tr><tr><td>1,4-diazabicyclo[2.2.2]octane (DABCO)</td><td></td><td>CAS: 281-57-9</td><td>To make 100 ml: 25 ml ddH&lt;sub&gt;2&lt;/sub&gt;O, 1 ml Tris HCl (1M, pH 7.5), 2.5 g of DABCO solid, 3.5 ml 6N HCl, 250 &amp;mu;l 10N NaOH, 70 ml glycerol. In 250 ml beaker with stir bar, add ddH&lt;sub&gt;2&lt;/sub&gt;O, Tris HCl and DABCO. Stir and then add 6N HCl, 10 N NaOH, and glycerol. Then add 10N NaOH dropwise until solution reaches pH 7.5. Aliquot. Store aliquots at -20 &amp;deg;C.</td></tr><tr><td>DABCO + p-phenylenediamine (PPD) solution</td><td></td><td></td><td>1.765 ml NaHCO&lt;sub&gt;3&lt;/sub&gt;, 0.353 Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;, 0.02 g PPD (CAS: 106-50-3). Dissolve PPD in NaHCO&lt;sub&gt;3&lt;/sub&gt; and NaCO&lt;sub&gt;3&lt;/sub&gt; solution. Add 60 &amp;mu;l of PPD solution to 500 &amp;mu;l of DABCO. Store aliquots at -20 &amp;deg;C.</td></tr><tr><td>Apple juice plates</td><td></td><td></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 ddH&lt;sub&gt;2&lt;/sub&gt;O. Add agar to ddH&lt;sub&gt;2&lt;/sub&gt;O in 4 L flask then autoclave for 30 min. Mix apple juice and sugar on heated stir plate. Gradually add apple juice mixture to autoclaved agar. Mix on heated stir plate then aliquot 10 ml volumes into 35 mm petri dishes and let stand at RT to solidify. Store at 4 &amp;deg;C.</td></tr><tr><td>Tegosept 10%</td><td></td><td></td><td>To make 100 ml: 10 g Tegosept, 100 ml ethanol. Store aliquots at -20 &amp;deg;C.</td></tr><tr><td>Yeast paste</td><td></td><td></td><td>~50 g dry active yeast. Gradually add ddH&lt;sub&gt;2&lt;/sub&gt;O to beaker containing yeast while stirring until paste-like consistency reached. Store at 4 &amp;deg;C.</td></tr><tr><td colspan="4">Table 2: Staining Materials</td></tr><tr><td>DAPI</td><td>Invitrogen</td><td>D3571</td><td>1:1000, stock at 5 mg/ml.</td></tr><tr><td>Rabbit anti-Vasa</td><td></td><td></td><td>1:250, a gift from Ruth Lehmann.</td></tr><tr><td>Mouse anti-Fasciclin III</td><td>Developmental Studies Hybridoma Bank (DSHB)</td><td>7G10</td><td>1:10</td></tr><tr><td>Mouse anti-1B1</td><td>Developmental Studies Hybridoma Bank (DSHB)</td><td>1B1</td><td>1:4</td></tr><tr><td>Guniea pig anti-Traffic Jam</td><td></td><td></td><td>1:2500, a gift from Dorthea Godt (Li et al, 2003).</td></tr><tr><td>Mouse anti-Prospero</td><td>Developmental Studies Hybridoma Bank (DSHB)</td><td>Prospero MR1A</td><td>1:10</td></tr><tr><td>Rat anti-Elav</td><td>Developmental Studies Hybridoma Bank (DSHB)</td><td>7EBA10</td><td>1:30</td></tr><tr><td>mouse anti-Repo</td><td>Developmental Studies Hybridoma Bank (DSHB)</td><td>8D12</td><td>1:10</td></tr><tr><td>Goat anti-rabbit Alexa546</td><td>Invitrogen</td><td>A11010</td><td>1:500</td></tr><tr><td>Goat anti-mouse Alexa488</td><td>Invitrogen</td><td>A11029</td><td>1:500</td></tr><tr><td>Goat anti-guniea pig Alexa633</td><td>Invitrogen</td><td>A21105</td><td>1:500</td></tr><tr><td>Goat anti-rat Alexa488</td><td>Invitrogen</td><td>A11006</td><td>1:500</td></tr><tr><td colspan="4">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 &lt;em&gt;Drosophila&lt;/em&gt; 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 &amp;mu;m 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></td><td>Microscope: BX51 equipped with DSU spinning disc, Light source: X-Cite 120Q, Camera: RETIGA-SRV, Imaging Software: Slidebook 5.0</td></tr><tr><td>Vacuum filter unit</td><td>Nalgene</td><td>450-0020</td><td>0.2 &amp;mu;m 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 late-embryonic through early/mid-L3 stages of Drosophila development.
Results from Testis Immunostain: 
Testis development is a particularly good system for illustrating protocol efficacy since testes maturation is dynamic throughout larval development. Adult Drosophila testes form a coiled tube with one blind end, where the germline stem cell (GSC) niche is situated (see 13,14 for reviews). In this niche, GSCs are arrayed around a tight cluster of non-mitotic somatic cells called the hub. GSCs undergo asymmetric division to produce one GSC that remains anchored to the hub and a daughter gonialblast that is displaced away from the stem cell niche. As the gonialblast divides incompletely, cytoplasmic extensions called fusomes form, connecting the cells within the spermatogonium. After 4 successive divisions, the spermatogonium initiates meiosis to form sperm.
Testis formation begins with the association of primordial germ cells (PGCs) and somatic gonadal precursor cells (SGPs) during embryogenesis (see 15,16 for reviews). This association results in formation of a functional GSC niche by the end of embryogenesis 8-10. By mid-L1, asymmetric GSC divisions within the GSC niche give rise to differentiating spermatogonia with branched fusomes 9. Asymmetric GSC division continues throughout larval development, resulting in production of additional spermatogonia and a progressive increase in gonad size. Representative images of late embryonic and early/mid-L1, L2, and L3 testes, immunostained for germ cells, hub cells, and fusomes, are shown (Figure 1A-D). These images illustrate the dynamic changes in gonad size and germ cell differentiation observed in testes over time.
Results from Ovary Immunostain: 
Adult Drosophila ovaries are composed of 16-20 individual egg-producing units called ovarioles (see 17,18 for reviews). At one end of each ovariole, a structure called the germarium contains a stem cell niche. The ovariole niche is composed of undifferentiated GSCs and two populations of somatic cells: cap cells and terminal filament cells (TFs). Similar to the testis, GSCs undergo asymmetric division to produce one GSC that remains adjacent to the cap cells and a differentiating daughter cell, called a cystoblast, which is pushed away from the niche. The cystoblast subsequently undergoes 4 rounds of cell division to form a germ-line cyst, interconnected by a branched fusome. Each germline-cyst is then surrounded by follicle cells to produce an egg chamber that continues to mature as it moves down the length of the ovariole.
Like testes, ovaries are first formed during embryogenesis 16,18. Multiple ovarioles arise from a single embryonic gonad comprised of PGCs and SGPs. By mid-L3, SGPs give rise to TF precursors at the ovary anterior, while germ cells localize to the ovary posterior and associate with SGP-derived intermingled cells (ICs)19-22. By late-L3, TF cells differentiate and organize into the stacks found in the adult stem cell niche, GSCs and cap cells are established, and germ-line cyst differentiation is observed 19,20,23. Representative images of late-stage embryonic and early/mid-L3 ovaries, immunostained for germ cells, SGPs, ICs, TF precursors, and fusomes, are shown (Figure 1E,F). These images show normal morphology and developmental progression for their age.&#160;
Results from Neural Immunostain: 
The Drosophila central nervous system (CNS) is derived from neural stem cells, called neuroblasts (NBs), that arise from embryonic neuroepithelium (see 24,25 for reviews). NBs in embryos and larvae divide asymmetrically to produce ganglion mother cells (GMC) that, in turn, generate neurons and glial cells found in the adult brain and ventral nerve cord. Because larval NBs give rise to the majority of adult neurons and can be distinguished based on their position within the brain, larval brains have become an important model for studying NB behavior and neural differentiation.
The L3 brain is composed of two lobes and a centrally located ventral nerve cord 24. Representative images of mid-L3 brains immunostained for NBs, GMCs, undifferentiated neurons, immature and primary neurons and glial cells are shown (Figure 2A-C)26-30. These images show strong staining in distinct expression patterns within the brain lobes and the ventral nerve cord. Depending on mounting orientation, imaging allows for brain tissue visualization from the dorsal surface (Figure 2A), ventral surface (Figure 2B), or in sagittal cross section (Figure 2C).
Staining efficiency: 
Our representative results demonstrate that a sonication-based immunostaining procedure is versatile. Not only can it be used to identify specific cell types but also can be used to indicate the presence of specific proteins and organelles. However, ambiguous results may stem from expected variation in staining efficiency. For example, anti-vasa stained at least one gonad in 73.2% of all larvae examined (n=781) while anti-Elav stained brains in 86% of larvae (n=115). Furthermore, we observe that staining efficiency is roughly inversely proportional to developmental stage. In late-stage embryos, anti-vasa stained 89.8% of gonads (n=206), while staining was present in only 59.8% and 46.9% of larvae at L2 and mid-L3, respectively (n=132 and 49). In addition, sonication results in sample loss. When the number of embryos and larvae present in an optimum sonication sample volume was counted before and after sonication, an average of 41.0% of sample was determined unsuitable for analysis (n=1165). To maximize staining efficiency, protocols should be optimized for specific antibodies by altering published antibody concentrations or antibody incubation times.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51528/51528fig1highres.jpg" width="500" /> 
Figure 1.&#160;Immunostain of Drosophila gonads in situ.&#160;(A-D)&#160;Images of testes in situ in late-stage embryos (stage 17/early-L1) through early/mid-L3 larvae immunostained with anti-Vasa (A-D, red; A&#8217;-D&#8217; alone) to detect germ cells, anti-Fasicilin III (Fas III) and anti-1B1 (A-D, green; A&#8217;&#8217;-D&#8217;&#8217; alone) to detect hub cells and fusomes respectively, and DAPI (A-D, blue; A&#8217;&#8217;&#8217;-D&#8217;&#8217;&#8217; alone) to detect nuclei. (A) Stage 17/early L1 testis shows the newly coalesced hub and spherical fusomes in undifferentiated germ cells. (B) Early/mid-L1 testis shows spherical fusomes in GSCs localized near the hub and in some posterior germ cells elongated fusomes indicative of early spermatogonial differentiation. (C) Early/mid-L2 and (D) early/mid-L3 testes show maintenance of GSCs with spherical fusomes near the hub and fusome branching indicative of late spermatagonial differentiation in posterior germ cells. Early/mid-L3 testis has increased in size substantially and shows significant fusome branching. (E-F)&#160;Images of ovaries in situ in late-stage embryos and mid-L3 larvae immunostained with anti-Vasa (E-F, red; E&#8217;-F&#8217; alone) to detect germ cells, anti-1B1 (E-F, green; E&#8217;&#8217;-F&#8217;&#8217; alone) to detect fusomes and somatic cell membranes at L3, and anti-Traffic Jam (TJ)(E-F, blue; E&#8217;&#8217;&#8217;-F&#8217;&#8217;&#8217; alone) to detect somatic ovarian cells. (E) Stage 17/early L1 ovary shows that somatic cells are distributed throughout the gonad and that germ cells have spherical fusomes. (F) Early/mid-L3 ovary is slightly enlarged and membrane-associated 1B-1 expression indicative of TF progenitor development is detected in anterior somatic cells. Germ cells at mid-L3 are located in the testes posterior, show spherical fusomes, and associate with TJpositive/1B1dim somatic ICs. All images are Z-projections of 4 successive confocal slices with the gonad anterior oriented to the left. Gonads are outlined and the hub is indicated with a yellow arrow in testes. Scale bar is 10 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51528/51528fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51528/51528fig2highres.jpg" width="500" /> 
Figure 2. Immunostain of Drosophila neural tissues in situ.&#160;Early/mid-L3 larvae immunostained with anti-Elav (A-C, red; A&#8217;&#8217; alone) to detect nuclei of immature and primary neurons, and either anti-Prospero (Pros) (A, green; A&#8217; alone) to detect nuclei in GMCs and undifferentiated neurons or anti-Reversed Polarity (Repo) (B-C, green; B&#8217;-C&#8217; alone) to detect glial cells. nuclei and DAPI (A-C, blue; A&#8217;&#8217;&#8217; alone) was used to detect all cell nuclei.&#160; All images oriented with anterior up. Sagittal section oriented with ventral to the left. (A) Dorsal section shows strong staining for Pros and Elav in peripheral regions of the brain lobe (BL) and along the midline and periphery of the ventral nerve cord (VNC). Inset in panel A shows Pros and Elav staining in distinct nuclei along the VNC midline. (B) Ventral cross-section shows broad-based expression of Repopositive glial cells with stronger staining along the VNC midline and periphery. (C) Sagittal section shows enrichment of Repo staining on the ventral face of the VNC. All images are single confocal sections. Scale bar is 20 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51528/51528fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

sonication-facilitated-immunofluorescence-staining-late-stage

Nanyang Technological University

Research

JoVE Journal

Biology

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

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of tissues, we have developed numerous modifications and improvements to our original fluorescent in situ hybridization (FISH) protocol. To facilitate throughput and cost effectiveness, all steps, from probe generation to signal detection, are performed using exon 96-well microtiter plates. Digoxygenin (DIG)-labelled antisense RNA probes are produced using either cDNA clones or genomic DNA as templates. After tissue fixation and permeabilization, probes are hybridized to transcripts of interest and then detected using a succession of anti-DIG antibody conjugated to biotin, streptavidin conjugated to horseradish peroxidase (HRP) and fluorescently conjugated tyramide, which in the presence of HRP, produces a highly reactive intermediate that binds to electron dense regions of immediately adjacent proteins. These amplification and localization steps produce a robust and highly localized signal that facilitates both cellular and subcellular transcript localization. The protocols provided have been optimized to produce highly specific signals in a variety of tissues and developmental stages. References are also provided for additional variations that allow the simultaneous detection of multiple transcripts, or transcripts and proteins, at the same time.

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

The described RNA in situ hybridization protocol allows the detection of RNA in whole Drosophila embryos or dissected tissues. Using 96-well microtiter plates and tyramide signal amplification, transcripts can be detected at high resolution, sensitivity, and throughput, and at a relatively low cost.

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of ...

Genetics

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and apoptosis, development and ageing, to neuronal synaptic plasticity and regeneration have been associated with Ryanodine receptors (RyRs). Despite the importance of calcium signaling, the exact mechanism of its function in early development is unclear. As an organism with a short gestational period, the embryos of Drosophila melanogaster are prime study subjects for investigating the distribution and localization of CICR associated proteins and their regulators during development. However, because of their lipid-rich embryos and chitin-rich chorion, their utility is limited by the difficulty of mounting embryos on glass surfaces. In this work, we introduce a practical protocol that significantly enhances the attachment of Drosophila embryo onto slides and detail methods for successful histochemistry, immunohistochemistry, and in-situ hybridization. The chrome alum gelatin slide-coating method and embryo pre-embedding method dramatically increases the yield in studying Drosophila embryo protein and RNA expression. To demonstrate this approach, we studied DmFKBP12/Calstabin, a well-known regulator of RyR during early embryonic development of Drosophila melanogaster. We identified DmFKBP12 in as early as the syncytial blastoderm stage and report the dynamic expression pattern of DmFKBP12 during development: initially as an evenly distributed protein in the syncytial blastoderm, then preliminarily localizing to the basement layer of the cortex during cellular blastoderm, before distributing in the primitive neuronal and digestion architecture during the three-gem layer phase in early gastrulation. This distribution may explain the critical role RyR plays in the vital organ systems that originate in from these layers: the suboesophageal and supraesophageal ganglion, ventral nervous system, and musculoskeletal system.

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Here, we describe a protocol for detection and localization of Drosophila embryo protein and RNA from collection to pre-embedding and embedding, immunostaining, and mRNA in situ hybridization.

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and ...

Developmental Biology

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, germ cell development, piRNA biology, and niche formation. Here, we present a dissection technique to live-image the gonad ex vivo during a period when in vivo live-imaging is highly ineffective. This protocol outlines how to transfer embryos to an imaging dish, choose appropriately-staged male embryos, and dissect the gonad from its surrounding tissue while still maintaining its structural integrity. Following dissection, gonads can be imaged using a confocal microscope to visualize dynamic cellular processes. The dissection procedure requires precise timing and dexterity, but we provide insight on how to prevent common mistakes and how to overcome these challenges. To our knowledge this is the first dissection protocol for the Drosophila embryonic gonad, and will permit live-imaging during an otherwise inaccessible window of time. This technique can be combined with pharmacological or cell-type specific transgenic manipulations to study any dynamic processes occurring within or between the&#160;cells in their natural gonadal environment.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of embryogenesis, these organelles undergo dramatic movements powered by actin-based cytoplasmic streaming and motor-driven trafficking along microtubules. The development of a multitude of small, organelle-specific fluorescent probes (FPs) makes it possible to visualize a wide range of different lipid-containing structures in any genotype, allowing live imaging without requiring a genetically encoded fluorophore. This protocol shows how to inject vital dyes and molecular probes into Drosophila embryos to monitor the trafficking of specific organelles by live imaging. This approach is demonstrated by labeling lipid droplets (LDs) and following their bulk movement by particle image velocimetry (PIV). This protocol provides a strategy amenable to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the ER, and for tracking the motion of individual LDs along microtubules. Using commercially available dyes brings the benefits of separation into the violet/blue and far-red regions of the spectrum. By multiplex co-labeling of organelles and/or cytoskeletal elements via microinjection, all the genetic resources in Drosophila are available for trafficking studies without the need to introduce fluorescently tagged proteins. Unlike genetically encoded fluorophores, which have low quantum yields and bleach easily, many of the available dyes allow for rapid and simultaneous capture of several channels with high photon yields.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of ...

Enhancing Protein Visualization with Antibody-Mediated Live Imaging

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein localization, dynamics, and function. Compared to immunofluorescence, live imaging more accurately reflects protein localization without potential artifacts arising from tissue fixation. Importantly, live imaging enables quantitative and temporal characterization of protein levels and localization, crucial for understanding dynamic biological processes such as cell movement or division. However, a major limitation of fluorescent tagging approaches is the need for sufficiently high protein expression levels to achieve successful visualization. Consequently, many endogenously tagged fluorescent proteins with relatively low expression levels cannot be detected. On the other hand, ectopic expression using viral promoters can sometimes lead to protein mislocalization or functional alterations in physiological contexts. To address these limitations, an approach is presented that utilizes highly sensitive antibody-mediated protein detection in living embryos, essentially performing immunofluorescence without the need for tissue fixation. As proof of principle, endogenously GFP-tagged Notch receptor that is barely detectable&#160;in living embryos can be successfully visualized after antibody injection. Furthermore, this approach was adapted to visualize post-translational modifications (PTMs) in living embryos, allowing the detection of temporal changes in tyrosine phosphorylation patterns during early embryogenesis and revealing a novel subpopulation of phosphotyrosine (p-Tyr) underneath apical membranes. This approach can be modified to accommodate other protein-specific, tag-specific, or PTM-specific antibodies and should be compatible with other injection-amenable model organisms or cell lines. This protocol opens new possibilities for live imaging of low-abundance proteins or PTMs that were previously challenging to detect using traditional fluorescent tagging methods.

Author Spotlight: Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research 

This protocol describes the customized antibody-based fluorescence labeling and injection into early Drosophila embryos to enable live imaging of low-abundance proteins or post-translational modifications that are challenging to detect using traditional GFP/mCherry-tag approaches.

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein ...

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Assessing the expression pattern of a gene, as well as the subcellular localization properties of its transcribed RNA, are key features for understanding its biological function during development. RNA in situ hybridization (RNA-ISH) is a powerful method used for visualizing RNA distribution properties, be it at the organismal, cellular or subcellular levels 1. RNA-ISH is based on the hybridization of a labeled nucleic acid probe (e.g. antisense RNA, oligonucleotides) complementary to the sequence of an mRNA or a non-coding RNA target of interest 2. As the procedure requires primary sequence information alone to generate sequence-specific probes, it can be universally applied to a broad range of organisms and tissue specimens 3. Indeed, a number of large-scale ISH studies have been implemented to document gene expression and RNA localization dynamics in various model organisms, which has led to the establishment of important community resources 4-11. While a variety of probe labeling and detection strategies have been developed over the years, the combined usage of fluorescently-labeled detection reagents and enzymatic signal amplification steps offer significant enhancements in the sensitivity and resolution of the procedure 12. Here, we describe an optimized fluorescent in situ hybridization method (FISH) employing tyramide signal amplification (TSA) to visualize RNA expression and localization dynamics in staged Drosophila embryos. The procedure is carried out in 96-well PCR plate format, which greatly facilitates the simultaneous processing of large numbers of samples.

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Here we describe a whole-mount fluorescent in situ hybridization (FISH) protocol for determining the expression and localization properties of RNAs expressed during embryogenesis in the fruit fly, Drosophila melanogaster.

Assessing  the expression pattern of a gene, as well as the subcellular localization  properties of its transcribed RNA, are key features for ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm development. However, immunolabeling of the whole Drosophila testis is often associated with significant non-uniformity of staining due to antibody penetration. Squashed preparations only partially overcome the problem since it decreases the 3D quality of the analyses. Herein, we describe a whole-mount protocol using NP40 and heptane during fixation together with immunolabeling in liquid media. It preserves the volume suitable for confocal microscopy together with reproducible and reliable labeling. We show different examples of 3D reconstitution of spermatocyte nuclei from confocal sections. The intra- and inter-testes reproducibility allows 3D quantification and comparison of fluorescence between single cells from different genotypes. We used different components of the intranuclear MINT structure (Mad1-containing Intra Nuclear Territory) as well as two components associated with the nuclear pore complex to illustrate this protocol and its applications on the largest cells of the testis, the S4-S5 spermatocytes.

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

We present here an improved protocol for the whole-mount preparation and immunostaining of Drosophila testes, suitable for confocal microscopy and also allowing reproducible and reliable labeling. We illustrate this protocol by 3D representation and quantification of colocalization experiments on the large S4-S5 spermatocytes.

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm ...

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

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Chapters in this video

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Upright Imaging of Drosophila Embryos

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes