This research was supported by the National Institutes of Health (RO1 GM079351 to D.K.). We thank Dorothea Godt for her helpful advice on pupal ovary dissections based on her original protocol. We also thank Amy Reilein for her assistance and comments on the manuscript.

1. EggLaying
<ol>
	<li>Combine approximately ten male and fifteen female adult Drosophila flies of the desired genotype in a vial of normal rich fly food supplemented with yeast. To avoid overcrowding the vial, allow mated females to lay eggs for no longer than 2&#8211;4 h14.</li>
	<li>Transfer the adults from the vial into a new vial by tapping the vial opening against a different vial with fly food. Allow the eggs to develop into larvae at room temperature for 3&#8211;4 days.</li>
</ol>
2. Selecting Female Larvae
<ol>
	<li>Using a moist fine brush with soft bristles, make a rolling movement with the brush along the wall of the vial to transfer wandering third-instar larvae from the vial to a glass well filled to the brim with 1x phosphate-buffered saline (PBS). Wash the food debris off the larvae by transferring them to another well filled with 1x PBS.</li>
	<li>Separate the male and female larvae using forceps. Identify the male larvae by a pair of large, round, and translucent testes embedded in fat body on the lateral side of the body approximately two-thirds down from its anterior end15. Female larvae are on average bigger, less translucent than males, and have much smaller gonads that are more difficult to detect.</li>
	<li>Select against male larvae to collect female larvae. Collect at least ten females from the well. Place the female larvae into a separate well filled with 1x PBS.</li>
	<li>Use forceps to transfer the female larvae gently into a new vial of fresh fly food supplemented with yeast.</li>
	<li>Place the vial with the female larvae in a dark location to facilitate pupariation16. Throughout the day, monitor the larvae for pupariation. As each larva immobilizes and develops into a prepupa, circle the prepupa against the vial and record the approximate time when it first forms into a prepupa. Identify prepupae by the protrusion of anterior spiracles and timing of puparium formation17. Allow the prepupae to develop to the desired time point (measured in hours after puparium formation, APF). 
	NOTE: Any animal that undergo puparium formation within an approximately 10 h interval can be considered a prepupa. The pupal stage begins approximately 12 h after puparium formation after an internal molt has taken place.</li>
</ol>
3. Preparing Pupae for Antibody Staining
<ol>
	<li>Form a thin glass pipet to be used in later steps for clearing out the pupal fat body.
	<ol>
		<li>Melt the glass tip of a Pasteur pipet over a Bunsen burner. As the glass melts, use forceps to pull the tip horizontally away from the rest of the pipet to form a thinner tip.</li>
		<li>After cooling, break off a small portion of the pipet tip to form a neat circular opening. Attach a bulb to the other end of the pipet.</li>
		<li>Load the pipet with 1x PBS.</li>
	</ol>
	</li>
	<li>To harvest the pupae, which are glued to the wall of the vial, apply a small drop of water along the contact zone between pupa and vial. Wait 1&#8211;2 min to let the protein glue dissolve before gently lifting the pupa off the wall with a moist fine brush. Transfer the pupa to a glass well filled with 1x PBS. Dedicate a single well per pupa to avoid overcrowding the well.</li>
	<li>While grasping the posterior end of the pupa with one pair of forceps, carefully tear the anterior portion of the pupal case with another pair until the head of the pupa is visible.</li>
	<li>Grip the anterior-most tip of the pupal head with forceps and gently pull the pupa out of its pupal case. 
	NOTE: A portion of the fat body may spill out during this process.</li>
	<li>Separate and discard the anterior half of the pupa from the posterior half until only the abdominal sack remains.</li>
	<li>Extract the fat body cells from the abdominal sack.
	<ol>
		<li>Grasp the abdominal sack against the bottom of the well with forceps in one hand while holding the thin glass pipet filled with 1x PBS in another hand.</li>
		<li>Aim the thin pipet tip towards the opening of the sack and slowly pipet 1x PBS into the abdomen to wash away the fat body cells surrounding the pupal ovaries. 
		NOTE: The ovaries are a pair of small, translucent, striated, and oblong structures that should remain inside the abdominal sack.</li>
		<li>Wash away the fat body cells until at least two-thirds of the fat body is gone or until the ovaries are visible near the opening of the abdominal sack. It is very important to execute this step slowly so as not to wash the ovaries out of the abdomen by accident.</li>
	</ol>
	</li>
	<li>Examine the well to check if the ovaries have spilled out during the fat body cell wash. If the ovaries are not visible inside the well, they should have remained in the abdomen. If the ovaries have come out, place a new pupa in the well and repeat this step.</li>
	<li>Transfer the abdomen into a clear coverglass chamber filled with fixation buffer (1x PBS, 4% paraformaldehyde) and place the lid on top. To ensure a sufficient number of ovaries withstand the staining and mounting process, dissect more ovaries than needed. 
	Caution: The fixation buffer contains paraformaldehyde which is toxic. Please wear appropriate protections such as gloves, safety glasses, etc.</li>
</ol>
4. Immunohistochemistry
<ol>
	<li>Throughout the staining process, use forceps to gently push down any floating ovaries to the bottom of the cover glass well to ensure that the ovaries are completely immersed in antibody solution.</li>
	<li>Place strips of double-sided tape onto the flat surface of a Nutator and then place the chambered coverglass on the adhesive surface.</li>
	<li>Incubate the ovaries in fixation buffer from step 3.7 for 15 min at room temperature.</li>
	<li>Rinse the ovaries three times in 1x PBS with 1% Triton X-100 for 5, 10, and 45 min (1 h in total), respectively, to allow thorough permeabilization.</li>
	<li>Block the ovaries in 10% normal goat serum in 1x PBS with 0.5% Triton X-100 for 30 min.</li>
	<li>Incubate the ovaries overnight in 600 &#181;L of primary antibody of choice diluted to the appropriate concentration in 1x PBS with 0.5% Triton X-100 at 4 &#176;C.</li>
	<li>Rinse the ovaries three times for 5 min in 1x PBS with 0.5% Triton X-100.</li>
	<li>Incubate the ovaries for 2 h in 600 &#181;L of secondary antibody of choice diluted to the appropriate concentration in 1x PBS with 0.5% Triton X-100 at room temperature.</li>
	<li>Rinse the ovaries twice for 5 min in 1x PBS with 0.5% Triton X-100 and once for 5 min in 1x PBS.</li>
</ol>
5. Dissecting and Mounting Pupal Ovaries
<ol>
	<li>Place a small drop of 1x PBS onto a microscope slide. Use forceps to transfer the abdominal sack from the chambered coverglass to a glass well filled with 1x PBS. Dedicate a single well per pupa to avoid overcrowding the well.</li>
	<li>Tear apart the abdominal sack and any remaining fat body with forceps. 
	NOTE: The ovaries, which are a pair of small, translucent, striated, and oblong structures, should be visible inside the well. The ovaries are often tightly surrounded by the fat body, so it is important to make sure all fat body is thoroughly torn apart.</li>
	<li>Transfer the ovaries from the well into the drop of 1x PBS on the microscope slide by grasping the center of the ovaries between the tips of the forceps. It is very important to use a firm grip without squeezing the ovaries so as not to lose or destroy them during the transfer. Add a few drops of 1x PBS to the slide when the solution dries out over time.</li>
	<li>Once all ovaries have been dissected and transferred to the microscope slide, pipet 40 &#181;L of mounting medium onto a 22 mm x 22 mm coverslip and place the coverslip gently on top of the ovaries. Let the slide dry overnight prior to imaging.</li>
</ol>

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The authors have no conflicts of interest to declare.

The most critical and difficult step of this protocol involves the preparation of pupal ovaries prior to fixation. To ensure that the ovaries, small and buried by fat body cells inside the pupal abdomen, are stained sufficiently with antibodies, it is important to not only tear a large opening in the abdominal sack with forceps, but also extract the fat body cells that obstruct the ovaries from the antibodies. Successful execution of this step requires application of subtle pressure on the Pasteur pipet bulb while washing the fat body cells out of the abdominal sack (e.g. 1&#8211;2 fat body cells should leave the abdomen per second during this step). Failure to use gentle force will likely cause the ovaries to spill out of the abdomen and into the glass well. Because the ovaries by themselves are small, translucent, and difficult to grasp with forceps, they must remain within the abdominal sack throughout the entire staining procedure. Thus, if the ovaries come out of the abdomen during the preparation step prior to fixation, it would be extremely difficult to retrieve, stain, and mount them in their isolated form during the protocol.
Fat body cells, which primarily consist of lipid droplets, are triglyceride storage organelles commonly found in insects, including Drosophila melanogaster19,20. Since some fat body cells remain in the pupal abdomen at fixation, this protocol utilizes relatively high Triton X-100 concentrations in the first three PBS rinses to extract as many lipids as possible from both the cell membranes in the ovary and any leftover fatty tissue surrounding the ovaries. As demonstrated by the bright Fasciclin III cell membrane protein antibody staining in Figure 3, using 1x PBS with 1% Triton X-100 during the first three rinses works well to maximally extract lipids while still maintaining the integrity of the cell membrane.
Once the fat body has been extracted, the second most critical step is to make sure the ovaries remain within the abdominal sack throughout the fixation and antibody staining. It is helpful to place them inside a clear, chambered coverglass rather than a microcentrifuge tube for clearer visibility of the pupal abdomens during the staining. Use forceps to gently push the abdominal sacks to the bottom of the chamber so that the abdominal tissues in the sack are fully engulfed in antibody solution.
Finally, great care should be taken to transfer the ovaries from the abdominal sack to the microscope slide during the mounting process. Though the ovaries are small and difficult to grasp with forceps without pinching them, the best way to transfer them to the slide is by firmly grasping the center of the ovaries. They will remain structurally intact once placed on the slide. It is possible to dissect the ovaries from the abdomen directly on top of the microscope slide to avoid losing the ovary, but this may lead to an excessive amount of mounting medium on the slide and could complicate the mounting process.
The limitations of this protocol involve the amount of time the entire dissection process may take, the potentially low yield of well-stained ovaries, and the dexterity needed to successfully complete each step. Because the pupal ovaries require extensive fat body extraction, preparing a single pupa for fixation may take anywhere from 10 to 20 min. Moreover, to ensure a sufficient number of ovaries withstand the entire staining procedure, it is best to dissect more pupae than needed for the experiment. This means that a large amount of time may be spent simply on preparing the pupae even before the fixation step.
Pupal ovary dissections differ largely from protocols for larval14 and adult21 ovary dissections. The encasement of the ovary within a pupal case presents unique challenges that are met by the tools used in this protocol. These methods may be applied to lineage tracing experiments to determine when FSCs and Escort cells are specified in the pupal ovary over time. A modified version of this protocol involving insect cell culture media could also potentially be used to dissect pupal ovaries for ex vivo live imaging analysis.

Stem cell research using Drosophila ovaries has widely expanded since the first documentation of a stem cell niche1,2,3,4. Following the development of lineage tracing genetic tools, Drosophila ovary dissections have been commonly used to study stem cell lineages and signaling pathways that regulate stem cell maintenance, proliferation, and fate in the stem cell niche. Knowledge of these signaling pathways may yield insights into potential causes of cancers that originate from aberrant stem cell activity5,6,7. It has also recently been shown that somatic stem cells in the Drosophila ovary, known as follicle stem cells (FSCs), strongly resemble mammalian intestinal stem cells in many aspects of their organization8. For this reason, Drosophila ovaries are a highly useful model system for studying stem cell behavior.
While larval and adult ovaries offer clues to early stem cell development and final stem cell organization in the niche, respectively, the pupal ovary is an intermediate structure in which the germline and somatic cells reorganize and establish their identities9,10. Though several studies have examined aspects of tissue development in the pupal ovary10,11,12,13, questions remain regarding the differentiation and spatial organization of ovarian cell types during pupal development. In particular, the specification of FSCs occurs during this period. This protocol outlines a method for dissecting and staining pupal ovaries at desired time points&#8212;a technique that can be used in time course experiments that analyze pupal ovary development in detail from the larval to the adult stage.
To account for the small size, translucence, and inaccessibility of the pupal ovary within the pupal abdomen, this protocol utilizes tools such as a custom-made thin-tipped Pasteur pipet to remove abdominal fat body tissue obstructing antibody access to the ovaries. A clear, chambered coverglass used during the antibody staining offers greater visibility of the pupae and a gentler platform for rocking the ovaries on a &#34;Nutator.&#34; Based on a protocol for larval ovary dissections by Maimon and Gilboa14, a relatively high concentration of Triton X-100 has been employed in the initial steps of the staining procedure to maximize cell membrane permeabilization and antibody access to the ovarian cells.

<table><tbody><tr><td>Dumont #5 Forceps, biology</td><td>Fine Scientific Tools</td><td>11252-20</td><td></td></tr><tr><td>Nunc Lab-Tek Chambered Coverglass</td><td>Thermo Fisher Scientific</td><td>155383</td><td></td></tr><tr><td>Dissection microscope</td><td>Nikon</td><td>SMZ-10A</td><td></td></tr><tr><td>Confocal Microscope</td><td>Carl Zeiss</td><td>LSM 700</td><td></td></tr><tr><td>Analysis software</td><td>Carl Zeiss</td><td>Zen</td><td></td></tr><tr><td>9 Depression Glass Spot Plates</td><td>Pyrex</td><td>7220-85</td><td></td></tr><tr><td>Pasteur pipet</td><td>Fisher Scientific</td><td>13-678-6B</td><td></td></tr><tr><td>Pasteur pipet bulb</td><td>Various vendors</td><td></td><td></td></tr><tr><td>Bunsen burner</td><td>Various vendors</td><td></td><td></td></tr><tr><td>Fisherfinest Premium Frosted Microscope Slides</td><td>Thermo Fisher Scientific</td><td>12-544-2</td><td></td></tr><tr><td>22 x 22 mm glass coverslips No 1</td><td>VWR</td><td>48366-067</td><td></td></tr><tr><td>Dapi Fluoromount-G</td><td>SouthernBiotech</td><td>0100-20</td><td></td></tr><tr><td>Double-sided tape</td><td>Scotch</td><td></td><td></td></tr><tr><td>Nutator</td><td>Clay Adams</td><td></td><td></td></tr><tr><td>Fine brush #0, #3-#5</td><td>Various vendors</td><td></td><td></td></tr><tr><td>Gilson Pipetman Starter Kit</td><td>Thomas Scientific</td><td>F167300</td><td>Contains p20, p200, p1000 pipettors</td></tr><tr><td>16% Paraformaldehyde</td><td>Electron Microscopy Sciences</td><td>15710</td><td>Dilute to 4% paraformaldehyde in 1x PBS</td></tr><tr><td>Triton</td><td>Sigma-Aldrich</td><td>9002-93-1</td><td></td></tr><tr><td>10x PBS</td><td>Ambion</td><td>AM9624</td><td>Dilute to 1x PBS</td></tr><tr><td>Normal Goat Serum</td><td>Jackson ImmunoResearch</td><td>5000121</td><td>Dilute to 10% normal goat serum in PBST with 0.5% Triton concentration</td></tr><tr><td>Primary antibodies (in protocol: 7G10 anti-Fasciclin III diluted 1:250, rabbit anti-phosphohistone H3 diluted 1:1000)</td><td>Various vendors (in protocol: Developmental Studies Hybridoma Bank, Millipore)</td><td></td><td>Dilute in PBST with 0.5% Triton concentration</td></tr><tr><td>Secondary antibodies (in protocol: Alexa-546, FITC-conjugated anti-rabbit serum)</td><td>Various vendors (in protocol: Molecular Probes, Jackson ImmunoResearch Laboratories, Inc.)</td><td></td><td>Dilute in PBST with 0.5% Triton concentration</td></tr><tr><td>Fly vials</td><td>Denville Scientific</td><td>V9406</td><td></td></tr><tr><td>Cotton Balls, For Wide Vials</td><td>Genesee Scientific</td><td>51-102W</td><td></td></tr><tr><td>Yeast, Bakers Dried Active</td><td>MP Biomedicals</td><td>101400</td><td></td></tr><tr><td>Fly food</td><td>Produced in laboratory</td><td></td><td>Mixture of water, brewer&#039;s yeast, cornmeal, molasses, agar, EtOH, penicillin, methyl 4-hydrobenzoate, and propionic acid</td></tr><tr><td>Male and female &lt;em&gt;Drosophila&lt;/em&gt; flies (genotype used in protocol: yw; P[Fz3-RFP, w+]/TM2)</td><td>Bloomington Drosophila Stock Center</td><td></td><td></td></tr></tbody></table>

dissection and staining of drosophila pupal ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Successful execution of this procedure should result in clear antibody staining that reveals the structure and cellular organization of a Drosophila pupal ovary. Immunohistochemistry outlined in this protocol can be used to identify cell types commonly stained in larval and adult ovaries. Cells of the pupal stalk derived from swarm cells18 (outlined by Fasciclin III in white) are shown in Figure 3. In addition to highlighting the cellular organization of pupal ovaries, antibody staining specific to cell proliferation (such as phospho-histone H3 staining in Figure 3, shown in green) can be used to study cell division patterns of stem cells and other mitotically active cell types. If fluorescent antibody signals are weak when examined under a confocal microscope, it is likely that the ovaries were not sufficiently exposed to the antibodies due to insufficient fat body extraction in the pupal abdomen. Another possibility may be that the pupal sack opening collapsed during the staining.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56779/56779fig1.jpg" /> 
Figure 1: Side-by-side comparison of male vs. female larvae. (A) Male larva in PBS solution identified by a pair of translucent, oblong testes located approximately two-thirds down from its anterior end. (B) Female larva in PBS solution identified by the absence of large, translucent testes. Scale bars = 2mm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56779/56779fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56779/56779fig2.jpg" /> 
Figure 2: Dissection of pupal ovaries after immunohistochemistry. (A) Image of a pair of striated, translucent pupal ovaries (red circles) that have been removed from the abdominal sack upon immunohistochemistry staining. Pupal ovaries were dissected approximately 48 h APF. The remaining fat body cells (black arrow) that were not extracted in step 3.6 are dispersed in PBS solution. (B) Image of a single pupal ovary in PBS solution dissected approximately 48 h APF. Scale bars = 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56779/56779fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56779/56779fig3.jpg" /> 
Figure 3: Stained wild-type pupal ovary dissected 48 hours APF. Image was acquired using a confocal microscope. The Wnt pathway reporter, Fz3 RFP8 (red), is expressed in anterior somatic cells. Antibodies directed to Fasciclin III (white) outlines cells of the pupal stalk. Nuclei were counterstained with DAPI (blue). Phospho-histone H3 staining (green) highlights cells undergoing mitosis. Diagonal white lines indicate the edges of the original image. Scale bar = 20 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56779/56779fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

dissection-and-staining-of-drosophila-pupal-ovaries

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

9.3K Views.  Columbia University. De Drosophila eierstok is een uitstekende modelsysteem voor het bestuderen van de stamcel niche ontwikkeling. Hoewel de methoden voor het ontleden van larvale en volwassen eierstokken zijn gepubliceerd, vereisen pop eierstok dissecties verschillende technieken die niet zijn gepubliceerd in detail. Hier duidelijk naar voren komt een protocol voor ontleden, kleuring en montage van de pop eierstokken.

Video: Dissection and Staining of Drosophila Pupal Ovaries

Dissection of Drosophila Ovaries

Biology

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Visualizing the Effects of Oxidative Damage on Drosophila Egg Chambers using Live Imaging

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ribonucleoprotein particle movement, mRNA localization, organelle movement, and cytoskeletal dynamics. There are several methods for live imaging that have been developed. Due to the fact that each method involves dissecting individual ovarioles placed in media or halocarbon oil, cellular damage due to hypoxia and/or physical manipulation will inevitably occur over time. One downstream effect of hypoxia is to increase oxidative damage in the cells. The purpose of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries after induction of controlled cellular damage. Here, we use hydrogen peroxide to induce cellular oxidative damage and give examples of the effects of such damage on two subcellular structures, mitochondria and Clu bliss particles. However, this method is applicable to any subcellular structure. The limitations are that hydrogen peroxide can only be added to aqueous media and would not work for imaging that uses halocarbon oil. The advantages are that hydrogen peroxide is readily available and inexpensive, acts quickly, its concentrations can be modulated, and oxidative damage is a good approximation of damage caused by hypoxia as well as general tissue damage due to manipulation.

The objective of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries.

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ...

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Dissection and Staining of Drosophila Larval Ovaries

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

Dissection and Staining of Drosophila Larval Ovaries

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

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

Drosophila Pupal Abdomen Immunohistochemistry

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

Drosophila Pupal Abdomen Immunohistochemistry

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

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

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 ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

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

Studying Mitochondrial Structure and Function in Drosophila Ovaries

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

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

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and diseases, such as obesity and fatty liver. Usually, lipids are stored in lipid droplets, which are the multifunctional lipid storage organelles in cells. Lipid droplets vary in size and number in different tissues and under different conditions. It has been reported that lipid droplets are tightly controlled through regulation of its biogenesis and degradation. In Drosophila melanogaster, the oenocyte is an important tissue for lipid metabolism and has been recently identified as a human liver analogue regarding lipid mobilization in response to stress. However, the mechanisms underlying the regulation of lipid droplet metabolism in oenocytes remain elusive. To solve this problem, it is of utmost importance to develop a reliable and sensitive method to directly visualize lipid droplet dynamic changes in oenocytes during development and under stressful conditions. Taking advantage of the lipophilic BODIPY 493/503, a lipid droplet-specific fluorescent dye, described here is a detailed protocol for the dissection and subsequent lipid droplet staining in the oenocytes of Drosophila larvae in response to starvation. This allows for qualitative analysis of lipid droplet dynamics under various conditions by confocal microscopy. Furthermore, this rapid and highly reproducible method can also be used in genetic screens for indentifying novel genetic factors involving lipid droplet metabolism in oenocytes and other tissues.

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Presented here are detailed methods for the dissection and lipid droplet staining of oenocytes in Drosophila larvae using BODIPY 493/503, a lipid droplet-specific fluorescent dye.

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and ...

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

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

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

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

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

Dissection and Staining of Drosophila Pupal Ovaries

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