Name: Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes (Video) | JoVE Journal
Uploaded: 2014-01-20T15:00:00.000+00:00
Duration: 10 min 30 s
Description: Learn cytological analysis of spermatogenesis using live and fixed Drosophila testes preparations. This method enables observation of cellular structures and protein localization through phase-contrast and immunofluorescence microscopy.

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5 minutes

The authors would like to thank Michael Anderson for establishing in the Lee lab these accepted methods for studying spermatogenesis with expert advice from Karen Hales. H. Oda and Y. Akiyama-Oda generously provided the &#947;-tubulin-GFP fly stock. This work was supported by an NIH R01 grant to L.A.L. (GM074044).

1.&#160;Testes Dissection
<ol>
	<li>Anesthetize flies in a bottle or vial using a stream of CO2 and transfer to a fly pad.</li>
	<li>Sort flies under a dissecting microscope using a small paintbrush, and collect an appropriate number (depending on the experiment) of Drosophila males of the desired genotypes. Young males (0-2 days old) are ideal for examining cells throughout the earlier stages of spermatogenesis (e.g. spermatogonia, spermatocytes, and early post-meiotic spermatids), whereas slightly older males (2-5 days old) are ideal for examining cells in the final stages of spermatogenesis (in particular, mature sperm).</li>
	<li>Use forceps to remove the wings from each fly (to prevent the flies from floating in liquid during dissection).</li>
	<li>Add ~500 ml of phosphate-buffered saline (130 mM NaCl, 7 mM Na2HPO4, 3 mM NaH2PO4; PBS) in a drop to a silicone-coated dissection dish on a black background. Other aqueous solutions have been successfully used for testes dissection3.</li>
	<li>Point forceps towards the anterior of the fly, grasp it by the thorax, and immerse it in the PBS drop. While viewing through the dissecting microscope, use another pair of forceps to grasp and pull the external genitalia (dark brown structure located at the posterior end of the ventral abdomen) posteriorly until it detaches from the abdomen. In most cases, the testes, seminal vesicles, and accessory gland will be removed from the abdomen along with the external genitalia; if not, insert a single pair of forceps into the abdomen and tease out the testes.</li>
	<li>Separate testes from accessory gland and external genitalia using two pairs of forceps in the PBS drop (Figure 2). Wild-type testes are easily distinguished from neighboring white tissues by their yellow color. Proceed immediately to step 2.</li>
	<li>To isolate testes from pharate males (i.e. enclosed within the pupal case), an additional step must first be performed that involves removing the fly from the pupal case; this step has been previously described elsewhere31. Proceed with dissection as for the adult testes beginning at step 1.2.</li>
	<li>To isolate testes from larval males, perform a modification of a protocol for isolating Drosophila larval ovaries32. Briefly, male larvae can be distinguished from female larvae by the presence of a pair of large, clear, oval structures (larval testes) embedded in the posterior third of the fat body. To isolate larval testes, partially flay open the male larva to isolate the testes and the surrounding fat body from the abdomen as described for the isolation of larval ovaries. Proceed immediately to step 2 of the protocol described herein; the testes can later be removed from the fat body just prior to mounting (steps 2.3 or 3.14) as described for the ovaries32.</li>
</ol>
2.&#160;Sample Preparation and Live Imaging
<ol>
	<li>Use a pair of forceps to gently place 2-3 pairs of testes in a drop of 4-5 ml of PBS on a square glass cover slip. Note that the ratio of testes number to PBS volume may need to be adjusted: too much liquid will prevent cells from spreading properly when squashed, whereas too little liquid will cause cells to burst when squashed. Optional: Use siliconized cover slips to minimize adherence of tissue to cover slip in step 3.2.</li>
	<li>Use a pair of forceps to tear open each testis at an appropriate position so as to maximize the presence of the desired germline cell types in the preparation (note that the contents of the testis will mostly egress from the torn region onto the slide during the squashing step.) To enrich for spermatogonia and spermatocytes, tear open the testis adjacent to its apical tip (level 1, Figure 2B). To enrich for spermatocytes and spermatids, tear open the testis at a position slightly basal to level 1 (level 2, Figure 2B). To enrich for more mature germline cells, tear open the testis closer to where the curvature begins (level 3, Figure 2B).</li>
	<li>Gently place a glass microscope slide over the cover slip to squash the testes; do not apply pressure manually as the weight of the cover slip alone is sufficient to obtain a properly squashed sample. Try to avoid trapping air bubbles. Optional: Use poly-L-lysine coated microscope slides to promote adherence of tissue to slide in step 3.2.</li>
	<li>Use preparation immediately (ideally within 15 min of preparation) to observe live cells by phase-contrast microscopy; for transgenic flies with expression of fluorescently tagged proteins in the testes, live cells can be examined by fluorescence microscopy at this step. Alternatively, proceed with fixation and antibody staining (Protocol 3).</li>
	<li>Gently wick any excess liquid from under the coverslip using a cleaning wipe to allow flattening of the preparation until the germ cells are clearly in focus.</li>
</ol>
3.&#160;Formaldehyde Fixation and Antibody Staining
<ol>
	<li>Snap freeze each slide containing squashed testes (from Protocol 2) using a pair of metal tongs to immerse it briefly in liquid nitrogen (until liquid nitrogen stops bubbling).</li>
	<li>Remove the cover slip immediately using a razor blade.</li>
	<li>Use metal tongs to transfer slides to a prechilled glass slide rack filled with ice-cold 95% ethanol (spectrophotometric grade, methanol-free). Store at -20 &#176;C for 10 min.</li>
	<li>Use metal tongs to transfer slides to a glass slide rack filled with 4% formaldehyde in PBS plus 0.1% Triton X-100 (PBS-T). Store at room temperature for 7 min.</li>
	<li>Use metal tongs to transfer slides to a glass slide rack filled with PBS. Wash slides in PBS for 5 min at room temperature. Repeat 1x. Perform all washes by discarding solution in the glass slide rack (i.e. by pouring it out) and replacing with fresh solution.</li>
	<li>Discard the PBS and immerse slides in PBS-T for 30 min at room temperature to permeabilize cell membranes.</li>
	<li>Wash slides in PBS for 5 min at room temperature. Repeat 2x.</li>
	<li>Blocking step (optional): Immerse slides in PBS plus 1% BSA for 45 min at room temperature.</li>
	<li>Use a hydrophobic barrier pen to draw a circle on the slide around squashed tissue (easily visible by eye) in order to confine the antibody solutions (added in steps 3.9 and 3.11). The tissue should be kept moist at all times while performing immunostaining.</li>
	<li>Add 30-40 ml of primary antibody (diluted in PBS-T, 1:400 to 1:50, depending on antibody) to tissue within the circle. If blocking was performed, dilute primary antibody in PBS-T plus 1% BSA. Anti-gamma-tubulin antibody (for staining of centrosomes in Figure 4) was diluted 1:100. Incubate in a moist, dark chamber (e.g. closed plastic box with damp paper towels) for 2 hr at room temperature or overnight at 4 &#176;C.</li>
	<li>Wash slides in PBS for 5 min at room temperature 3x. If blocking was performed, wash twice in PBS-T and once in PBS (5 min at room temperature each).</li>
	<li>Add 30-40 ml of fluorophore-conjugated secondary antibody (diluted 1:400 in PBS) to the tissue and incubate in the dark for 1-2 hr at room temperature.</li>
	<li>Wash slides in PBS for 5 min at room temperature. Repeat 2x.</li>
	<li>Add 30-40 ml of DAPI solution (0.2 mg/ml in PBS) to the tissue within the circle.</li>
	<li>Gently place a glass cover slip over the tissue, taking care to avoid trapping air bubbles. If air bubbles should appear, carefully move around the cover slip without destroying the squash until the bubbles escape from the sides of the cover slip.</li>
	<li>Use a cleaning wipe to blot excess DAPI from the edges of the slide.</li>
	<li>Seal the cover slip to the slide using clear nail polish.</li>
	<li>Use this preparation within the next 3-4 hr to view immunostained cells by fluorescent microscopy. For longer-term storage of slides (up to at least several weeks), use a glycerol-based hard mount media with DAPI, and store slides at -20 &#730;C.</li>
	<li>Alternatively, fix samples in methanol instead of formaldehyde (depending on the antigen). After completing the entire protocol through step 3.2, immerse slides of squashed testes in methanol for 10 min at -20 &#176;C and proceed with step 3.5 onward.</li>
</ol>

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<li>Wang, Z., Mann, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Requirement+for+two+nearly+identical+TGIF-related+homeobox+genes+in+Drosophila+spermatogenesis%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Requirement for two nearly identical TGIF-related homeobox genes in Drosophila spermatogenesis.</a> Development. 130, 2853-2865 (2003).</li>
</ol>

The authors declare that they have no competing financial interests.

Although the testes of wild-type flies can be readily identified due to their yellow color (in contrast the neighboring white tissues), the testes of white mutant flies are white and thus can occasionally be confused with the gut. Most transgenic strains, which are typically in a white background, also have white testes because the mini-white gene found in P-elements does not promote pigment accumulation in the testes. When Drosophila testes cannot be distinguished by color, o...

Drosophila testes are an ideal model system for the study of many biological processes including the regulation of stem cells, meiosis, and sperm development13-18. The spermatocytes and their meiotic spindles are large and hence convenient for cytological analysis, and relaxed cell cycle checkpoints during spermatogenesis facilitate the study of mutations in cell cycle genes. Different cell types can be observed in ordered progression along the length of the testes, and any disruption in spermatogenesis can lead to changes in this overall arrangement. These features combined with Drosophila genetic tools have facilitated the mutational analysis of spermatogenesis21-23.
The stages of Drosophila spermatogenesis have been well defined. Germline cells that develop synchronously within cysts progress sequentially through the stages of spermatogenesis along the length of the testis. During both the mitotic and meiotic divisions of the male germ cells, cytokinesis occurs incompletely such that the daughter cells remain connected by cytoplasmic bridges known as ring canals (Figure 1). The apical tip of the testis contains a population of germline stem cells that gives rise to spermatogonial cells, which undergo four mitotic divisions with incomplete cytokinesis to generate 16-cell cysts of primary spermatocytes. After premeiotic S phase, primary spermatocytes enter G2, a prolonged growth period of ~90 hr during which cellular volume increases ~25-fold. Progression through meiosis I and meiosis II results in the formation of 32-cell cysts of secondary spermatocytes and 64-cell cysts of haploid spermatids, respectively. The immature, round spermatids undergo extensive cellular remodeling to form mature sperm. Post-meiotic cells, in particular the bundles of elongating and mature spermatids, occupy much of the volume of the testis.
The successful transport of functional sperm to female flies requires coordination between the different parts of the male reproductive system, which is composed of several paired structures (the testes, seminal vesicles, and accessory glands) and a single ejaculatory duct (Figure 2). Sperm are produced within the testes and stored within the seminal vesicles until copulation24. The accessory glands contain secretory cells that produce seminal fluid. The sperm migrating from the seminal vesicles are mixed with seminal fluid within the ejaculatory duct, which is connected to both the seminal vesicles and the accessory glands. This mixture of sperm and seminal fluid is ultimately pumped out of the male into the vagina of the female fly through the ejaculatory bulb located at the posterior end of the male abdomen25. Proteins within the seminal fluid are essential for prolonged storage of sperm within specialized organs known as spermathecae in the reproductive tract of Drosophila females26.
Excellent methods for the isolation of Drosophila testes and visualization of cells at various stages of spermatogenesis are available in the scientific literature3-12. We herein add to this body of knowledge by presenting examples of these protocols with an accompanying video demonstration. The protocol for preparation of live testes samples for phase-contrast microscopy is based on a previously described method27. The protocol for formaldehyde fixation and immunostaining of testes is also based on a previously described method28. The approaches described herein have been used in many studies of Drosophila spermatogenesis (for example, to assess the roles of dynein, a minus-end-directed microtubule motor, during Drosophila spermatogenesis).
In addition to the basic protocols, suggestions are provided for varying the dissection so as to enrich for spermatogonia, spermatocytes, or mature sperm. Different methods for processing the testes such that cysts either remain intact or are disrupted as needed are described. An advantage in using Drosophila testes as a model system is that, compared to Drosophila oocytes and embryos, antibodies and dyes can easily penetrate cells following their dispersal from the testes, and fewer washing steps are required; thus, protocols can be performed in a relatively short time.

<table><tbody><tr><td>Sylgard</td><td>World Precision Instruments</td><td>SYLG184</td><td>Two-part silicon elastomer for making silicone-coated dissection dish from Kimax Petri dish</td></tr><tr><td>PAP pen</td><td>Fisher Scientific</td><td>NC9888126</td><td>Ted Pella #22309</td></tr><tr><td>Clear nail protector</td><td>Wet n Wild</td><td>7780235001</td><td></td></tr><tr><td>ProLong Gold Antifade Reagent with DAPI</td><td>Life Technologies</td><td>P36931</td><td></td></tr><tr><td>Mouse anti-gamma-tubulin antibody (clone GTU-88)</td><td>Sigma-Aldrich</td><td>T6557</td><td></td></tr><tr><td>Cy3-AffiniPure Goat Anti-Mouse IgG&amp;nbsp;</td><td>Jackson ImmunoResearch</td><td>115-165-003</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>BP151-100</td><td></td></tr><tr><td>Ethanol</td><td>Fisher Scientific</td><td>AC61511-0040</td><td></td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A412-4</td><td></td></tr><tr><td>16% Formaldehyde</td><td>Thermo Fisher Scientific</td><td>28908</td><td></td></tr><tr><td>Sigmacote</td><td>Sigma-Aldrich</td><td>SL2</td><td>Use according to manufacturer&#039;s directions to siliconize cover slips</td></tr><tr><td>DAPI</td><td>Sigma-Aldrich</td><td>D-9542</td><td>0.5 mg/ml in 75% ethanol; store at -20 &amp;deg;C</td></tr><tr><td>NaCl</td><td>Research Products International Corp.</td><td>S23020</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S9763</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S0751</td><td></td></tr><tr><td>Kimwipes delicate task wipers</td><td>Fisher Scientific</td><td>S47299</td><td></td></tr><tr><td>BSA</td><td>Research Products International Corp.</td><td>A30075</td><td>Molecular biology grade</td></tr><tr><td>Glass Coplin staining jar, screw cap</td><td>Electron Microscopy Sciences</td><td>70315</td><td></td></tr><tr><td>Single frosted microscope slides</td><td>Corning</td><td>2948-75X25</td><td></td></tr><tr><td>Poly-L-lysine coated microscope slides</td><td>Polysciences, Inc.</td><td>22247-1</td><td>Optional (to replace untreated microscope slides )</td></tr><tr><td>Square cover glass</td><td>Corning</td><td>2865-22</td><td></td></tr><tr><td>Razor blades</td><td>Fisher Scientific</td><td>12-640</td><td></td></tr><tr><td>Kimax Petri dish</td><td>Fisher Scientific</td><td>S31473</td><td>Kimble #23060 10015 EMD</td></tr><tr><td>Forceps</td><td>Dumont</td><td>52100-51S</td><td>Pattern 5 INOX</td></tr><tr><td>Stemi 2000-CS stereoscope</td><td>Carl Zeiss</td><td></td><td></td></tr><tr><td>Eclipse 80i</td><td>Nikon</td><td></td><td></td></tr><tr><td>Plan-Fluor 40X objective</td><td>Nikon</td><td></td><td></td></tr><tr><td>Axiophot</td><td>Carl Zeiss</td><td></td><td></td></tr><tr><td>Plan-Neofluar Ph2 40X objective</td><td>Carl Zeiss</td><td></td><td></td></tr></tbody></table>

cytological analysis of spermatogenesis: live and fixed preparations of drosophila testes

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.

An example of a properly dissected pair of Drosophila male reproductive organs is shown in Figure 2A. Testes removed from the abdomen of the adult male fly are typically attached to the ejaculatory duct (brown, Figure 2A') and a pair of accessory glands (green, Figure 2A') via a pair of seminal vesicles (blue, Figure 2A'). To separate the testes from most of the accompanying somatic tissue, the ejaculatory duct and the accessory glands should be...

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 ...

The overall goal of this procedure is to dissect and prepare oph testes for microscopy. This is accomplished by first dissecting out testes pairs from the abdomen of male flies. The second step is to tear the testes at the appropriate level of development and to squash the testes to release the cells within.Next, the squashed testes are fixed and then incubated in primary and secondary antibodies. The final step is to mount the tissue on a slide for imaging. Ultimately, phase contrast or immunofluorescence microscopy is used to observe cells at various stages of spermatogenesis for defects and morphology, and to observe the subcellular localizations of various proteins.This method can help answer key questions in the field of developmental biology, such as the identification of critical roles of proteins during the progression of spermatogenesis. Visual demonstration of the dissection of RAs testes is critical because a novice to this procedure may initially have difficulty in locating the testes within the fly abdomen. Additionally, it's easier to demonstrate rather than to describe in words the most successful way to perform the squashing of the tissue.To begin dissection, anesthetize flies in a bottle or vial using a stream of carbon dioxide. Then transfer the anesthetized flies to a fly pad. Sort flies under a dissecting microscope using a small paintbrush to collect approximately six to eight oph males of the desired genotype.Next, remove the wings from each fly with forceps. This will prevent the flies from floating in the liquid. During dissection, add 500 microliters of phosphate buffered saline or PBS in a single drop onto a silicone coated dissection dish on a black background.Then point forceps towards the anterior end of one of the de winged flies. Grasp it by the thorax and immerse it in the PBS drop. Identify the dark brown structure located at the posterior end of the ventral abdomen and use another pair of forceps to grasp and pull the external genitalia posteriorly until it detaches from the abdomen.In most cases, the testes, seminal vesicles and accessory gland will be removed from the abdomen along with the external genitalia. However, if they are not, insert a pair of forceps into the abdomen and tease out the testes. Separate testes from accessory glands and external genitalia using two pairs of forceps.Distinguish wild type testes from neighboring white tissues by their yellow color. To prepare the sample for live imaging, use a pair of forceps to gently place two to three pairs of testes in a four to five microliter drop of PBS on a square glass cover slip. Next, tear.Open each testes with a pair of forceps at an appropriate level position to maximize the presence of the desired germline cell types in the preparation. The contents of the testes will egress from the torn region onto the slide. During the squashing step to enrich for spermatogonia and spermatocytes tear open the testes adjacent to its apical tip to enrich.For spermatocytes and spermatids tear open the testes at a position slightly basal to level one to enrich for more mature germline cells. Tear open the testes closer to where the curvature begins. The next step is to squash the testes by gently placing a microscope slide over the cover slip.Do not apply pressure manually and avoid trapping air bubbles. Use the sample within 15 minutes of preparation to observe live cells. By phase contrast microscopy gently wick any excess liquid from under the cover slip using a cleaning wipe to allow flattening of the preparation until the germ cells are clearly in focus.Snap freeze prepared slides containing squash testes by immersing them with a pair of metal tongs into liquid nitrogen until the liquid stops bubbling. Remove the cover slip immediately with a razor blade. Then transfer slides with metal tongs to a pre chilled glass slide rack filled with ice cold 95%ethanol and store the sample at minus 20 degrees Celsius for 10 minutes.Transfer the equilibrated slides to a glass slide rack filled with 4%formaldehyde in PBS plus 0.1%Triton X 100 or PBST and store at room temperature for seven minutes. Then transfer the slides to a rack filled with PBS. Wash the slides in PBS for five minutes at room temperature by discarding the solution in the rack and replacing it with fresh PBS solution.Repeat the wash once. Discard the PBS and immerse slides in PBST for 30 minutes at room temperature to perme cell membranes. Then wash slides three times in PBS for five minutes At room temperature, draw a circle around the squash tissue on the slide with a hydrophobic barrier pen in order to confine the antibody solutions and to keep the tissue moist at all times while performing immuno staining.Then add 30 to 40 microliters of primary antibody diluted in PBST to the tissue within the circle. Incubate the slide in a moist dark chamber for two hours at room temperature or overnight at four degrees Celsius following incubation in primary antibody. Wash the slides three times in PBS for five minutes at room temperature.Then add 30 to 40 microliters of a floor, four conjugated secondary antibody diluted in PBS to the tissue and incubate in the dark for one to two hours at room temperature. Next, wash the slides three times in PBS for five minutes at room temperature. Then add 30 to 40 microliters of dappy solution to the tissue within the circle.Gently place a glass cover slip over the tissue, taking extra care to avoid trapping any air bubbles. If air bubbles appear carefully move the cover slip around without destroying the sample until the bubbles escape from the sides of the cover. Slip blot excess dappy from the edges of the slide with a cleaning wipe.Then seal the cover slip using nail polish view immuno stain cells using a fluorescent microscope within three to four hours. This cartoon depicts the location of various cell maturation levels within drosophila Testes. Level one contains younger cells that are in the early stages of spermatogenesis and level three contains more mature sperm cells.Level two contains a mixed population of miotic and post miotic germline cells. A corresponding phase contrast image shows the release of spermatogonia and early and late primary spermatocytes when the testes is torn at level one. The yellow arrow also points to a fusion of two or more interconnected cells from incomplete cytokinesis.An artifact of the squashing procedure shown here is a corresponding phase contrast image showing the release of late primary spermatocytes and post myotic cells, including round spermatids, elongating spermatids, and mature sperm when the testes was torn at level two. Phase contrast imaging of squash testes can also be used to readily identify defects in the abundant round. Spermatids wild type spermatids with one phase light nucleus are shown here attached to a phase dark single mitochondrial aggregate of roughly equal size.In contrast, spermatids from Aun testes contain multiple small nuclei and one large mitochondrial aggregate as a result of failed cytokinesis and errors in chromosome segregation. The gray scale images shown here are from a preparation of squash testes stained with reagents that identify the microtubules DNA and centrosomes in color. The images combined show a dividing spermatocyte, large field of round spermatids and a bundle of mature sperm Once mastered preparation of samples for phase contrast, microscopy can be done in 20 to 30 minutes.Preparation of samples for immunofluorescence microscopy can be done in six hours if performed properly. In addition To the processing of oph testes for microscopy, dissected testes can be alternatively used for the biochemical analysis of proteins.

cytological-analysis-spermatogenesis-live-fixed-preparations

Research

JoVE Journal

Biology

Imaging of centrosomal proteins during Drosophila spermatogenesis is a powerful method to identify new proteins critical for centrosome biology as well as to elucidate the particular function of known players in this process.

Imaging Centrosomes in Fly Testes

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

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

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.

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

We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

The goal of this tubule squash technique is to rapidly assess cytological features of developing mouse spermatocytes while preserving cellular integrity. This method allows for the study of all stages of spermatogenesis, and can be easily implemented alongside other biochemical and molecular biological approaches for the study of mouse meiosis.

A Seminiferous Tubule Squash Technique for the Cytological Analysis of Spermatogenesis Using the Mouse Model

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

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.

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

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

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

Here we present a simple and efficient method to isolate live meiotic and post-meiotic germ cells from adult mouse testes. Using a low-cytotoxicity, violet-excited DNA binding dye and fluorescence-activated cell sorting, one can isolate highly enriched spermatogenic cell populations for many downstream applications.

Isolation of Murine Spermatogenic Cells using a Violet-Excited Cell-Permeable DNA Binding Dye

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.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

This article reports an in vivo inhibition of CENP-E through abdominal surgery and testicular injection of GSK923295, a valuable model for male meiotic division. Using the immunofluorescence, flow cytometry and transmission electron microscopy assays, we show that CENP-E inhibition results in chromosome misalignment and genome instability in mouse spermatocytes.

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry

Learn cytological analysis of spermatogenesis using live and fixed Drosophila testes preparations. This method enables observation of cellular structures and protein localization through phase-contrast and immunofluorescence microscopy.

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

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