Name: preparation of drosophila larval and pupal testes for analysis of cell division in live, intact tissue
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Description: Watch this Scientific Journal Video about Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue at JoVE.com

This work was supported by Department of Defense start-up funds to J.T.S.

1. Prepare Animals, Tools, and Media for Dissection
<ol>
	<li>Cross male and female flies to obtain progeny of the desired genotype. Useful transgenic markers for identification and staging of meiotic spermatocytes include GFP-tubulin to label the meiotic spindle and RFP-histone 2A to label chromsomes8. Use five to ten females per cross and maintain crosses on standard fly food in vials in a 25 &#176;C incubator until third instar larvae begin crawling up the sides of the vial (4&#8211;5 days). Early white pupae will begin forming one day later.</li>
	<li>Obtain two pairs of straight forceps with fine tips. Ensure that the tips are sharp, even in length, and straight (Figure 1A). Use these forceps only for dissections, and cap with 10 &#956;L pipette tips when not in use.</li>
	<li>Prepare a scalpel tool.
	<ol>
		<li>Insert the blunt end of a black anodized steel insect pin into a pin holder, and twist the screw of the holder to secure the pin in place. Using a pair of forceps and under a dissecting microscope, grasp the insect pin in the middle and bend it to form an internal angle of 135&#176; (Figure 1B). The sharp end is for cutting tissue, while the edge is used for transferring testes between drops of media.</li>
		<li>Cap with a 1,000 &#956;L pipette tip when not in use.</li>
	</ol>
	</li>
	<li>Working in a tissue culture hood, prepare 5 mL aliquots of Schneider&#8217;s media and store at 4 &#176;C until the time of use. When ready to begin dissections, warm one 5 mL media aliquot to room temperature. Then transfer the warmed dissection media to a 10 mL syringe and attach a 0.22 &#181;m syringe filter.</li>
</ol>
2. Dissection of Testes from Larvae and Early Pupae
<ol>
	<li>Use the media filled filter syringe to expel three drops of media (50 &#181;L each) at the top, middle and bottom of a glass slide.</li>
	<li>Use a dissecting probe to transfer five to ten third instar larvae or early pupae (pupae that are still white or yellow) to the top drop. Gently agitate the larvae and pupae in the media with the dissecting probe to remove any food or debris.</li>
	<li>Under a dissecting microscope, turn a single larva or pupa on its side with a dissecting probe to identify the two bilateral testes if present, which appear as oval shaped translucent structures in the posterior third of the body (Figure 2A, B). Animals that lack testes are female and should be discarded.</li>
	<li>When a male larva or pupa is found and is clean, move it to the second drop of media. Repeat until 3 or 4 male larvae and/or pupae have been transferred to the second drop of media.</li>
	<li>Working in the second drop of media, grasp a single male larva or pupa with a pair of forceps at its mid-region, just anterior to the testes. Use the second pair of forceps to gently tear the animal in half.</li>
	<li>Hold the posterior end of the larva or pupa down on the glass slide with a pair of forceps. Starting just next to the forceps, push down on the cuticle using the edge of the scalpel tool and move the scalpel toward the cut end of the animal. This will expel the animal&#8217;s internal organs, which include the guts, fat bodies, and testes.</li>
	<li>Gently tease apart the testes from the rest of the organs. The testes are clear, oval shaped organs embedded in ribbons of fat body (Figure 2C).</li>
	<li>Transfer the testes one at a time to the third drop of media. To accomplish this, insert the edge of the scalpel under the fat body, lift the tissue out of the media, and quickly move it to the third drop.</li>
	<li>Alternatively, if there is not enough fat body still attached to the testes to successfully lift the tissue, gently transfer the tissue using a glass Pasteur pipette that has been pre-wetted with dissection media.</li>
	<li>Use the sharp end of the scalpel tool to gently slice away excess fat body from the testes, leaving just a small rim of fat body around the edges (Figure 2C). It is not necessary to remove all of the fat body, and attempting to do so will likely result in damaging or rupturing of the testes.</li>
	<li>Repeat the above steps for all of the male larvae on the glass slide.</li>
	<li>Proceed either to step 3 or step 5.</li>
</ol>
3. Mounting Testes for Live Microscopic Imaging
NOTE: This mounting procedure was adapted from a recently published protocol for imaging of Drosophila larval brain neuroblasts20. Additional details can be found in this reference.
<ol>
	<li>Use the filter syringe to deposit a single drop of Schneider&#8217;s media (about 30 &#181;L) onto the center of a 50 mm gas-permeable culture dish.</li>
	<li>Use a pre-wetted Pasteur pipette to transfer the prepared testes to the drop on the gas-permeable dish. The testes will generally float to the surface of the drop.</li>
	<li>Use the scalpel tool to gently push the testes down onto the gas-permeable membrane. Take care not to rupture the gas-permeable membrane with the sharp point of the scalpel. Push all of the testes to the center of the drop.</li>
	<li>Use a 1 mL syringe filled with halocarbon oil to make four drops of oil, about 30 &#956;L each, on the gas permeable membrane. Space the drops of oil around the drop of media containing the testes to correspond to the four corners of a 22 mm glass coverslip.</li>
	<li>Line up the corners of a 22 mm glass coverslip with the four drops of halocarbon oil and gently lower the coverslip onto the media and oil. Allow the coverslip to settle and for the media containing the testes to spread between the drops of oil.</li>
	<li>Remove excess media to allow the coverslip to settle onto the surface of the testes. While observing the testes under a dissecting microscope, insert the corner of a delicate task wipe into the media under the coverslip to wick away a small amount of liquid. Wick away enough media until the glass coverslip just makes contact with the surface of the largest testis. It is critical not to remove too much media and lower the coverslip too far, as this will exert pressure on the testes and cause them to rupture (see Figure 5).</li>
</ol>
4. Live Imaging of Meiotic Spermatocytes
NOTE: Spermatocytes can be imaged using a laser scanning or spinning disk confocal microscope. The system must have adequate speed and sensitivity to avoid photobleaching of fluorescent proteins or photodamage of the tissue.
<ol>
	<li>Place a drop of immersion oil on the glass coverslip just above the testes. Flip over the dish and place it on the microscope stage and move the microscope objective (40x or 60x) into the oil until it is just below the testes. Use transmitted light to find a single testis and bring it into focus.</li>
	<li>While capturing fast images with the confocal, move the testis around to examine the organization of the cells. One end of the testis will have many small, densely packed cells. This end contains the germline stem cells and spermatogonia. Moving toward the other end of the organ, identify progressively larger cells organized into discrete clusters or cysts. These large cells are the spermatocytes (Figure 3A).</li>
	<li>If imaging GFP-tubulin, identify spermatocytes that will commence meiosis within 30&#8211;60 min by the presence of two bright microtubule asters (i.e., centrosomes) located at the cortex of the cell (Figure 3B, C).</li>
	<li>Once a cyst of meiotic spermatocytes has been identified, set up acquisition parameters to capture z-stacks of images over time. Typically, acquisition of 15&#8211;20 images spaced 1.5 &#956;m apart in the z-dimension is sufficient to capture the full volume of several spermatocytes within the same cyst.</li>
	<li>Acquire full z-stacks every 2 min if imaging the entire first meiotic division, which takes about 1.5 hours. It is possible to use faster acquisition rates, particularly if only a specific event of meiosis is to be analyzed. However, take care not to cause photobleaching or photodamage due to repeated exposure of the sample to laser excitation.</li>
	<li>Use a continuous, automatic focus control system to prevent focal drift over the long time-course of acquisition. Temperature control of the sample on the microscope stage generally is not necessary, assuming room temperature of approximately 22&#8211;23 &#176;C. If temperature control is available, maintain the sample at 25 &#176;C on the microscope stage.</li>
	<li>If meiotic spermatocytes are not found, store the sample for several hours or overnight in a 25 &#176;C incubator and image again.</li>
</ol>
5. Fixation and Immunostaining
<ol>
	<li>From step 2.10 above, use a glass Pasteur pipette to transfer testes to a well in a 9-well glass dissecting dish containing 0.5 mL of 8% paraformaldehyde in PBSTx (phosphate buffered saline with 0.3% Triton X-100). Ensure that the testes are laying on the bottom of the dish. 
	NOTE: Paraformaldehyde is toxic and should be handled with gloves in a fume hood.</li>
	<li>Fix the testes for 20 min at room temperature.</li>
	<li>Transfer the testes to a well containing 0.5 mL PBSTx. Wash for 5 min with gentle agitation. Repeat the wash step in new wells with fresh PBSTx two more times.</li>
	<li>Dilute primary antibody to the desired concentration in 0.5 mL PBSTx containing 5% bovine serum albumin (BSA) in a 1.5 mL microcentrifuge tube. Transfer testes from the 9-well glass dissecting dish to the tube of diluted primary antibody and incubate overnight at 4 &#176;C with gentle rocking or nutation.</li>
	<li>Wash the testes in 0.5 mL of PBSTx for 5 min in a well of a 9-well glass dissecting dish. Repeat the wash step with fresh PBSTx two more times.</li>
	<li>Dilute secondary antibody in 250 &#956;L of PBSTx containing 5% BSA and dispense into a clean well of a 9-well glass dissecting dish. If desired, add DAPI to stain DNA. Transfer testes to the well containing secondary antibody, cover with plastic wrap, and incubate in the dark with gentle agitation for 4 hours at room temperature.</li>
	<li>Transfer the testes to a clean well filled with 0.5 mL of PBSTx. Wash with fresh PBSTx twice for 5 min and a third time for 30 min.</li>
</ol>
6. Mounting Fixed Testes
<ol>
	<li>Transfer the testes from the last wash onto a glass microscopy slide using a Pasteur pipette. If necessary, use the edge of the scalpel tool to push the testes down onto the surface of the slide.</li>
	<li>Use the corner of a delicate task wipe to wick away excess liquid from the testes. Remove as much liquid as possible.</li>
	<li>Apply a 30&#8211;50 &#181;L drop of microscopy mounting medium to the testes.</li>
	<li>Place a 22 mm coverslip onto the drop of mounting medium and allow the coverslip to settle and the mounting medium to spread under the coverslip. Apply gentle pressure to the coverslip if necessary to squeeze out excess mounting medium and allow the coverslip to rest on the surface of the testes.</li>
	<li>Use nail polish to seal the edges of the coverslip.</li>
</ol>

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The authors have nothing to disclose.

We have described a protocol for the preparation of larval or early pupal Drosophila testes, optimized for long-term, live imaging of spermatocyte cell division. This is a powerful method for analysis of cell division in the physiological context of intact tissue. The power of this method is further expanded when combined with Drosophila genetic tools, such as specific gene mutations, tissue-specific RNAi-mediated suppression, and fluorescently labeled protein and organelle markers. In addition, the sma...

Cell division is often studied using cell lines grown in culture1. While we have gained a wealth of invaluable insight and understanding of fundamental mechanisms from these studies2, cells grown in culture cannot fully recapitulate the physiology of cell division as it occurs in intact, living tissue. For example, in intact tissues and organs, cells must divide at the right place and at the right time so that progeny cells are properly situated within the tissue, so that they can undergo appropriate differentiation or functional programs, and so that cell proliferation is properly coordinated with tissue growth or homeostasis3. For cells grown in culture on the other hand, cell division is generally regulated by growth factors in the culture medium4, and thus we cannot learn from these cells how in vivo environmental factors such as tissue architecture or developmental signaling influence the division process. It is also important to note that many of the cell lines used to study cell division, such as HeLa and U2OS cells, were derived from metastatic tumors5. Therefore, many aspects of the basic physiology of these cancer cells, such as cell cycle regulatory mechanisms and chromosomal stability, have likely been altered compared to healthy cells. Complete understanding of cell division physiology, therefore, depends on our ability to study dividing cells in their native, in vivo environments that preserve physiological regulatory mechanisms and tissue architecture.
Advances in understanding how cell division operates within intact tissues and organs are hampered by difficulties inherent to in vivo or ex vivo analysis. First, it can be difficult or impossible to access dividing cells for microscopic analysis within large organs or thick tissues. Second, it is often difficult to predict when individual cells will divide in vivo. Third, tissue physiology may rapidly deteriorate during ex vivo culture. In this protocol, we describe readily accessible methods for live and fixed analysis of Drosophila melanogaster spermatocytes as they undergo meiotic cell divisions within fully intact testes. These cells are ideal for live, ex vivo analysis because they are readily accessible with standard optical methods such as confocal microscopy, they divide at predictable times and locations, and intact testes can be maintained in ex vivo culture for up to about 24 h. In addition, Drosophila spermatocytes are large round cells (approximately 20&#8211;30 &#181;m in diameter) that do not change shape when they divide, making them ideal for high resolution, time-lapse imaging of cellular components such as the spindle apparatus and cytoplasmic organelles. Although these cells undergo meiosis as opposed to mitosis, many of the essential cell division processes are very similar between these two cell division mechanisms6. These advantages, combined with powerful Drosophila genetic tools, have made Drosophila spermatocytes an extensively used model for ex vivo analysis of essential cell division processes including spindle formation and regulation, cytokinesis, and organelle remodeling and partitioning7,8,9,10,11.
Spermatocytes are cells that are at the meiotic stage of spermatogenesis. In Drosophila, spermatogenesis begins with a group of germline stem cells that are located in a small region or &#8220;hub&#8221; at one pole of the testis12,13. These cells divide by an asymmetric mitosis, giving rise to a single differentiated spermatogonium. Spermatogonia then undergo four synchronous mitoses to produce a group of 16 cells that remain closely associated within a single cyst. At this stage, the cells transition from a mitotic to a meiotic cell cycle and are referred to as spermatocytes. Spermatocytes spend about two to three days in an extended G2 stage of the cell cycle, during which they grow dramatically and undergo cytological changes in preparation for the two meiotic divisions and subsequent spermiogenesis13,14. The entire group of 16 spermatocytes within a single cyst then enter the first meiotic division at the same time. Thus, several meiotic spermatocytes can be imaged simultaneously as they proceed through cell division. The first meiotic division proceeds for approximately 1.5 h and is followed almost immediately by the second meiotic division, yielding 64 total spermatids that go on to differentiate into mature spermatozoa.
A unique advantage of using spermatocytes to study cell division in live, intact tissue is that groups or cysts of cells constantly progress through the different stages of spermatogenesis, and cells at all stages of spermatogenesis can usually be identified in any given testis (see Figure 3A). Therefore, it is relatively easy to find meiotic cells in whole testes. We generally focus our analyses on the first as opposed to second meiosis because these cells are much larger and more amenable to high resolution imaging, but the entire process encompassing both meiotic divisions can be successfully imaged. It should also be noted that the general protocol for testis preparation and culture can be used to analyze other processes of spermatogenesis as well, such as the earlier mitotic stem cell or spermatogonial divisions or the cytological changes that occur as spermatids mature into spermatozoa15. Many of these aspects of spermatogenesis are highly conserved between Drosophila and humans16.
Drosophila spermatocytes begin to reach the meiotic phase of spermatogenesis during the third instar larval stage of Drosophila development13. Therefore, testes isolated from lifecycle stages beginning with third instar larvae and including pupae and adults can be used for analysis of dividing spermatocytes. Several excellent protocols are available for extraction of testes from adult male flies for live and fixed analysis of spermatogenesis17,18,19. These protocols may be preferable for studying late stages of spermatogenesis or if genetic markers that are only visible in adults must be used. The protocol focuses instead on testis preparation from larvae and early pupae, because testes at these stages have several advantages that are specifically pertinent to cell division analysis by high resolution, time-lapse microscopy. First, testes from larvae and pupae are smaller than those from adults, and the cells within the organ are less crowded. Because of this, dividing spermatocytes can often be imaged near the outer surface of larval testes, without having to penetrate through multiple layers of light scattering tissue. Second, adult testes move rhythmically due to contractions of the attached accessory organs, and these movements make time-lapse imaging of individual cells challenging. And third, larval testes are advantageous when studying gene mutations that cause pupal or adult lethality. Our methods are optimized for long-term culture of testes on the microscope stage, allowing for imaging of multiple rounds of cell division or the progression of individual cells through multiple stages of spermatogenesis in the same preparation. We also describe a protocol for fixation and immunostaining of whole testes. Overall, our protocols are particularly useful for those interested in studying cell division in intact tissue, and the ability to combine spermatocyte analysis with highly tractable Drosophila genetic tools makes this an especially powerful approach.

<table><tbody><tr><td>5&quot; Dissecting Probe</td><td>Fisher</td><td>08-965-A</td><td></td></tr><tr><td>5.75&quot; Glass Pasteur Pipet</td><td>Fisher</td><td>13-678-20A</td><td></td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Fisher</td><td>BP9703-100</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11252-20</td><td>Straight forcepts with fine tips</td></tr><tr><td>Frosted Microscope Slides</td><td>Fisher</td><td>12-544-2</td><td>Slides for mounting fixed tissue, with frosted writing surface</td></tr><tr><td>Halocarbon oil 700</td><td>Sigma</td><td>H8898-100 mL</td><td></td></tr><tr><td>Lumox Dish 50</td><td>Sarstedt</td><td>SAR946077410</td><td>Gas-permeable tissue culture dish</td></tr><tr><td>Microscope Cover Glass</td><td>Fisher</td><td>12-541-B</td><td>22 mm x 22 mm, #1.5 glass coverslip</td></tr><tr><td>Minutien Pins</td><td>Fine Science Tools</td><td>26002-15</td><td>Insect pins used to make scalpel tool</td></tr><tr><td>Nickel Plated Pin Holder</td><td>Fine Science Tools</td><td>26018-17</td><td></td></tr><tr><td>Paraformaldehyde 32% Solution, EM Grade</td><td>Electron Microsocopy Sciences</td><td>15714</td><td></td></tr><tr><td>Plan Beveled Edge Microscope Slides</td><td>Fisher</td><td>12-549-5</td><td>Slides used for dissections</td></tr><tr><td>PYREX Spot Plates</td><td>Fisher</td><td>13-748B</td><td>9-well glass dissecting dish</td></tr><tr><td>Schneider&#039;s &lt;em&gt;Drosophila&lt;/em&gt; medium</td><td>Fisher</td><td>21720-024</td><td></td></tr><tr><td>Syringe Filter, 0.22 &amp;micro;m</td><td>EMD Millipore</td><td>SLGS033SB</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher</td><td>BP151-500</td><td></td></tr><tr><td>Vectashield</td><td>Vector Labs</td><td>H-1000</td><td>Microscopy mounting medium</td></tr></tbody></table>

preparation of drosophila larval and pupal testes for analysis of cell division in live, intact tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

When this protocol is successfully executed, testes will remain fully intact for imaging by confocal microscopy or other fluorescence microscopy methods. As seen in Figure 3A, the cellular organization of the testes is preserved, and the progression of cell differentiation from one end of the testis to the other including spermatogonia, spermatocytes, and haploid spermatids is visible. GFP-tubulin is a useful marker for identifying dividing spermatocytes and for live imaging of the progressi...

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Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

preparation-drosophila-larval-pupal-testes-for-analysis-cell-division

Research

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Developmental Biology

8.0K Views.  Uniformed Services University of the Health Sciences. The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Video: Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

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.

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.

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

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

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, ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Biology

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Flow cytometry has been widely used to obtain information about DNA content in a population of cells, to infer relative percentages in different cell cycle phases. This technique has been successfully extended to the mitotic tissues of the model organism Drosophila melanogaster for genetic studies of cell cycle regulation in vivo. When coupled with cell-type specific fluorescent protein expression and genetic manipulations, one can obtain detailed information about effects on cell number, cell size and cell cycle phasing in vivo. However this live-cell method has relied on the use of the cell permeable Hoechst 33342 DNA-intercalating dye, limiting users to flow cytometers equipped with a UV laser. We have modified this protocol to use a newer live-cell DNA dye, Vybrant DyeCycle Violet, compatible with the more common violet 405nm laser. The protocol presented here allows for efficient cell cycle analysis coupled with cell type, relative cell size and cell number information, in a variety of Drosophila tissues. This protocol extends the useful cell cycle analysis technique for live Drosophila tissues to a small benchtop analyzer, the Attune Acoustic Focusing Cytometer, which can be run and maintained on a single-lab scale.

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

A protocol for cell cycle analysis of live Drosophila tissues using the Attune Acoustic Focusing Cytometer is described. This protocol simultaneously provides information about relative cell size, cell number, DNA content and cell type via lineage tracing or tissue specific expression of fluorescent proteins in vivo.

Flow cytometry has  been widely used to obtain information about DNA content in a population of  cells, to infer relative percentages in different cell ...

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 ...

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

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

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

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

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

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 ...

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 ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

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Dissection and Staining of Drosophila Pupal Ovaries

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

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

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

Studying Mitochondrial Structure and Function in Drosophila Ovaries

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

Super-Resolution Live Cell Imaging of Subcellular Structures