IM is supported by the Marie Curie re-integration grant. This work was supported by the Israel Science Fund Grant no. 1146/08, by the Helen and Martin Kimmel Institute for Stem Cell Research at the Weizmann Institute of Science and the Leir Charitable Foundation.

1. Egg laying
<ol>
	<li>Five days before dissection: allow mated females to lay eggs for 2-4 hours on fresh food supplemented with yeast. To get synchronized and well-developed larvae, it is important not to have overcrowded cultured (about 30 eggs/ 25mm vial). Typically, 7-16 females are used per vial, depending on how well they lay.</li>
</ol>
2. Selecting larvae
<ol>
 <li>Prepare a 9-well glass dissecting dish filled with Ringer's medium (128mM NaCl, 2mM Kcl, 1.8mM CaCl2, 4mM MgCl2, 35.5mM Sucrose, 5mM Hepes pH 6.9).</li>
	<li>Prepare cell strainers in a six-well plate containing Ringer's medium and place it on ice. Alternatively, we use specially made molds fitted with a nylon mesh.</li>
	<li>Pick timed larvae from vial walls using fine biological tweezers (forceps) and place them in the ringer's containing dissecting dish.</li>
	<li>Transfer a female larva to a clean well. We distinguish females from males by their gonads. Male testes are easily identified as big clear ovals embedded in the posterior third of the fat body. Female ovaries, located at the same part of the fat body, can be identified as a much smaller, clear, round spheres.</li>
</ol>
3. Dissection of larva
 <ol>
 <li>Hold the larva down just posterior to the brain with forceps and remove the head with a second pair of forceps.</li>
	<li>Place the remaining posterior part on the dorsal side, with the trachea facing down.</li>
	<li>Hold the larva by the posterior spiracles with one pair of forceps and push it inward slowly while the other pair is sliding the cuticle posteriorly until about half the larval fat body emerges.</li>
	<li>Firmly hold the posterior end and use the other pair of forceps to loosely hold the cuticle. Gently and slowly pull away the posterior end so that the cuticle and the attached intestine slide through the gap. At the end of this process, the fat body should be completely separated from the intestine and the cuticle. Disconnect the intestine from the anterior part of the fat body. To make staining and mounting easier, it is important that the fat body will remain intact.</li>
	<li>Wet a pasture pipette thoroughly with Ringer's medium, preferably from a well containing dissected larvae. This helps coat the pipette and prevents the fat body from sticking to it. Use the pasture pipette to transfer the fat body into the ice cold Ringer's medium in the cell strainer.</li>
</ol>
4. Fixation and Staining
All steps are performed at room temperature, except for the incubation with first antibody, at 4 &deg;C.
<ol>
 <li>Incubate the fat body in 5% formaldehyde in Ringer's medium. for 20 minutes with gentle agitation.</li>
	<li>Wash for 5 minutes with 1% PBT (1% Triton x-100 in PBS). Repeat this step for 10 minutes and a third time for 45 minutes. 
1% Triton X-100 is required to perforate the larval ovary and allow antibodies to penetrate it.</li>
	<li>Block with 0.3% PBTB (0.3% Triton x-100 and 1% BSA in PBS) for 1 hour with gentle agitation</li>
	<li>Incubate with the desired 1st antibody diluted in 0.3% PBTB over night at 4&deg;C with gentle agitation. This step is usually performed in 0.2 ml tubes on a roller.</li>
	<li>Transfer fat body back to cell strainers.</li>
	<li>Wash 3 times, 30 minutes each, with 0.3% PBTB with gentle agitation.</li>
	<li>Block with 0.3% PBTB supplemented with 5% normal donkey serum for 1 hour with gentle agitation.</li>
	<li>Incubate with an appropriate secondary antibody diluted (according to manufacturer's specification) in the blocking solution (0.3% PBTB supplemented with 5% normal donkey serum in 0.3% PBT) for 2 hours with gentle agitation. If the antibody is fluorescent, incubate in the dark from this point onward. This step is usually performed in 0.2 ml tubes on a roller.</li>
	<li>Wash 3 times for 30 minutes each with 0.3% PBT with gentle agitation.</li>
</ol>
5. Mounting 
<ol>
 <li>Transfer the fat body to a clean eppendorf tube. Very carefully remove all liquids away and immediately cover with Vectashield mounting media. We use vectashield routinely since it does not harden while the ovaries are being separated from the fat body.
 Samples can be stored in mounting media at 4&deg;C for up to a week.
</li>
	<li>Cut the end of a pipette tip and use it to carefully transfer the fat body with minimal volume of mounting media (about 30 &mu;l for a 30mm cover-slip) onto a microscope slide.</li>
	<li>Use two nickel plated pin holders, holding 0.1 mm diameter pins to spread out the fat body. The ovaries are located at the posterior third of the fat body. The fat body that surrounds them is usually shaped like a 'flower'. This helps identify the location of the ovaries. Dissect out the 'flowers' from the rest of the fat body and discard the latter.</li>
	<li>Remove the fat body surrounding the gonad by crossing the two pins carefully around it. Place the isolated gonad on the slide away from the fat body residue.</li>
	<li>Cover with a cover slip and seal with nail polish.</li>
	<li>Visualize directly using a confocal microscope. Samples can be stored at 4&deg;C for up to three weeks.</li>
</ol>
6. Representative Results
We have used the above protocol to follow the establishment of several cell lineages within the somatic ovary, including GSCs and their somatic niches. For this purpose we use specific antibodies and markers to distinguish between the different cell types in the developing gonad. Here we show an example of two LL3 ovaries stained with different combinations of antibodies. Figure 1A highlights terminal filament and cap cells (green), which together form the somatic cells of the niche. Figure 1B, shows the Intermingled cells (ICs, magenta), which directly contact germ cells (blue).
<img src="/files/ftp_upload/2537/2537fig1.jpg" alt="Figure 1" /> Figure 1. LL3 ovaries. (A) monoclonal antibody 1B1 (magenta) outlines somatic cells and stains the fusome, an intracellular organelle within PGCs (arrows). The enhancer trap hh-lacZ (anti &beta;-galactosidase, green) is expressed in terminal filaments. At the base of the filament somatic cap cells (arrowheads) can be observed. (B) 1B1 antibody (green) outlines all somatic cells in the ovary. Anti-Vasa (blue) labels all PGCs. PGCs directly contact the Intermingled cells (ICs, anti-traffic jam, magenta). Bar (for A and B) is 20 &mu;m.

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No conflicts of interest declared.

This video demonstrates an isolation and staining protocol of late third instar larval ovaries. To perform this protocol routinely and reliably, attention should be paid to the following points:
<ol>
	<li>For synchronized and well-developed larvae, over crowding must be avoided.</li>
	<li>To avoid loss of the small translucent ovaries, make sure to dissect the fat body intact. This will also help in locating the ovaries at the mounting stage, particularly when mounting younger (second or beginning of third instar)...

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>NaCl</td>
<td>&nbsp;</td>
<td>J.T.Baker</td>
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<td>&nbsp;</td>
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<td>&nbsp;</td>
<td>Merck</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>CaCl2</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>MgCl2</td>
<td>&nbsp;</td>
<td>Merck</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Sucrose</td>
<td>&nbsp;</td>
<td>J.T.Baker</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Hepes</td>
<td>&nbsp;</td>
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<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>&nbsp;</td>
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<td>Triton X-100</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
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<td>Albumin Bovine Fraction V</td>
<td>&nbsp;</td>
<td>MP Biomedicals</td>
<td>160069</td>
<td>&nbsp;</td>
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<td>Dumont biology tweezers 5 dumstar polished</td>
<td>&nbsp;</td>
<td>FST Fine Science Tools</td>
<td>11295-10</td>
<td>&nbsp;</td>
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<td>Nickel plated pin holder</td>
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<td>FST Fine Science Tools</td>
<td>26018-17</td>
<td>&nbsp;</td>
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<td>s.s minutien pins 0.1mm diam, 10mm long</td>
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<td>FST Fine Science Tools</td>
<td>26002-10</td>
<td>&nbsp;</td>
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<td>9 well plates 85X100 mm, 22mm o.d.x7mm deep</td>
<td>&nbsp;</td>
<td>CORNING</td>
<td>7220-85</td>
<td>&nbsp;</td>
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<td>Stereo Microscope MZ 16.5 with a standard transmitted light base TL ST</td>
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<td>Leica</td>
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<td>Menzel Glaser GmbH</td>
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<td>Cover slips</td>
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<td>Corning</td>
<td>2940-223</td>
<td>&nbsp;</td>
<tr>
<td>Mounting media</td>
<td>&nbsp;</td>
<td>Vectashield</td>
<td>H-1200</td>
<td>&nbsp;</td>
<tr>
<td>Cell strainer</td>
<td>&nbsp;</td>
<td>Falcon</td>
<td>FAL352350</td>
<td>&nbsp;</td>
<tr>
<td>1B1 antibody</td>
<td>&nbsp;</td>
<td>Developmental Studies Hybridoma Bank</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Anti-Traffic Jam</td>
<td>&nbsp;</td>
<td>Laboratory of Dr. Dorothea Godt</td>
<td>&nbsp;</td>
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<td>Anti-Vasa</td>
<td>&nbsp;</td>
<td>Laboratory of Dr. Ruth Lehmann</td>
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<td>&nbsp;</td>
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<td>Anti &beta;-Galactosidase</td>
<td>&nbsp;</td>
<td>Cappel</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Secondary Antibodies</td>
<td>&nbsp;</td>
<td>Jackson ImmunoResearch</td>
<td>&nbsp;</td>
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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.

Watch this Scientific Journal Video about Dissection and Staining of Drosophila Larval Ovaries at JoVE.com

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.

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 ...

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26.1K Views.  Weizmann Institute of Science. 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.

Video: Dissection and Staining of Drosophila Larval Ovaries

Dissection of Drosophila Ovaries

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.

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

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.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...

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.

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.

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

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called imaginal discs. These discs give rise to a high percentage of adult structures that are found within the adult fly. Here we describe a protocol that has been optimized to recover these discs and prepare them for analysis with antibodies, transcriptional reporters and protein traps. This procedure is best suited for thin tissues like imaginal discs, but can be easily modified for use with thicker tissues such as the larval brain and adult ovary. The written protocol and accompanying video will guide the reader/viewer through the dissection of third instar larvae, fixation of tissue, and treatment of imaginal discs with antibodies. The protocol can be used to dissect imaginal discs from younger first and second instar larvae as well. The advantage of this protocol is that it is relatively short and it has been optimized for the high quality preservation of the dissected tissue. Another advantage is that the fixation procedure that is employed works well with the overwhelming number of antibodies that recognize Drosophila proteins. In our experience, there is a very small number of sensitive antibodies that do not work well with this procedure. In these situations, the remedy appears to be to use an alternate fixation cocktail while continuing to follow the guidelines that we have set forth for the dissection steps and antibody incubations.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called ...

Dissection and Mounting of Drosophila Pupal Eye Discs

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

Dissection and Mounting of Drosophila Pupal Eye Discs

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

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