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

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.