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

Podsumowanie

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

Streszczenie

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 and testes^3-12. 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.

Wprowadzenie

Drosophila testes are an ideal model system for the study of many biological processes including the regulation of stem cells, meiosis, and sperm development^13-18. 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 spermatogenesis^21-23.

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 literature^3-12. 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.

Protokół

1. Testes Dissection

1. Anesthetize flies in a bottle or vial using a stream of CO₂ and transfer to a fly pad.
2. 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).
3. Use forceps to remove the wings from each fly (to prevent the flies from floating in liquid during dissection).
4. 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.
5. 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.
6. 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.
7. 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.
8. 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.

2. Sample Preparation and Live Imaging

1. 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.
2. 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).
3. 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.
4. 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).
5. 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.

3. Formaldehyde Fixation and Antibody Staining

1. 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).
2. Remove the cover slip immediately using a razor blade.
3. 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 °C for 10 min.
4. 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.
5. 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.
6. Discard the PBS and immerse slides in PBS-T for 30 min at room temperature to permeabilize cell membranes.
7. Wash slides in PBS for 5 min at room temperature. Repeat 2x.
8. Blocking step (optional): Immerse slides in PBS plus 1% BSA for 45 min at room temperature.
9. 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.
10. 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 °C.
11. 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).
12. 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.
13. Wash slides in PBS for 5 min at room temperature. Repeat 2x.
14. Add 30-40 ml of DAPI solution (0.2 mg/ml in PBS) to the tissue within the circle.
15. 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.
16. Use a cleaning wipe to blot excess DAPI from the edges of the slide.
17. Seal the cover slip to the slide using clear nail polish.
18. 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 ˚C.
19. 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 °C and proceed with step 3.5 onward.

Wyniki

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...

Dyskusje

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...

Materiały

Name	Company	Catalog Number	Comments
Sylgard	World Precision Instruments	SYLG184	Two-part silicon elastomer for making silicone-coated dissection dish from Kimax Petri dish
PAP pen	Fisher Scientific	NC9888126	Ted Pella #22309
Clear nail protector	Wet n Wild	7780235001
ProLong Gold Antifade Reagent with DAPI	Life Technologies	P36931
Mouse anti-gamma-tubulin antibody (clone GTU-88)	Sigma-Aldrich	T6557
Cy3-AffiniPure Goat Anti-Mouse IgG	Jackson ImmunoResearch	115-165-003
Triton X-100	Fisher Scientific	BP151-100
Ethanol	Fisher Scientific	AC61511-0040
Methanol	Fisher Scientific	A412-4
16% Formaldehyde	Thermo Fisher Scientific	28908
Sigmacote	Sigma-Aldrich	SL2	Use according to manufacturer's directions to siliconize cover slips
DAPI	Sigma-Aldrich	D-9542	0.5 mg/ml in 75% ethanol; store at -20 °C
NaCl	Research Products International Corp.	S23020
Na2HPO4	Sigma-Aldrich	S9763
NaH2PO4	Sigma-Aldrich	S0751
Kimwipes delicate task wipers	Fisher Scientific	S47299
BSA	Research Products International Corp.	A30075	Molecular biology grade
Glass Coplin staining jar, screw cap	Electron Microscopy Sciences	70315
Single frosted microscope slides	Corning	2948-75X25
Poly-L-lysine coated microscope slides	Polysciences, Inc.	22247-1	Optional (to replace untreated microscope slides )
Square cover glass	Corning	2865-22
Razor blades	Fisher Scientific	12-640
Kimax Petri dish	Fisher Scientific	S31473	Kimble #23060 10015 EMD
Forceps	Dumont	52100-51S	Pattern 5 INOX
Stemi 2000-CS stereoscope	Carl Zeiss
Eclipse 80i	Nikon
Plan-Fluor 40X objective	Nikon
Axiophot	Carl Zeiss
Plan-Neofluar Ph2 40X objective	Carl Zeiss