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

The overall goal of this seminiferous tubule squash technique is to visualize the cytological features of mouse spermatogonia, spermatocytes, and spermatids while preserving their cellular integrity. For this demonstration, we focus on the analysis of spermatocytes undergoing meiosis. This method can help answer key questions in the field of male reproductive biology, such as spermatogonial stem cell differentiation, meiotic chromosome segregation, and post-meiotic differentiation of round spermatids.

The main advantage of this technique is that the cellular integrity of male germ cells is maintained, which allows examination of cellular structures not easily visualized with other techniques. This method can be used to improve our understanding of the paternal causes of infertility in any platy. And it can be applied to mammalian systems such as mouse or organ donor derived testes.

Generally, individuals new to this method will struggle with making a good squash. Applying sufficient force and properly arranging the tubules are important factors to producing an even monolayer of cells. For each mouse to be dissected, load two 35 millimeter Petri dishes, one with three milliliters of PBS and the other with two milliliters of freshly made Fixative Lysis solution.

Select mice undergoing the first wave of spermatogenesis to observe an enriched population of interest. After sacrificing the mouse, clean the abdomen with 70%alcohol and then proceed with the surgery. Make a V-shaped opening into the abdominal pelvic cavity and remove the testes by pulling on the epididymal fat pad using forceps, but without disturbing the tunica albuginea.

Transfer the testes to the dish containing PBS. There, remove the testicular tunica albuginea by puncturing the tissue with sharp forceps and collect the loose seminiferous tubules. Next transfer the tubules into the dish with Fixative Lysis solution and let the reaction proceed for five minutes at room temperature.

During the incubation, prepare a poly-L-lysine coated glass slide by outlining the edges of the slide with a liquid blocker pen and then filling the slide with 100 microliters of Fixative Lysis solution within the borders. After the five minute incubation, use sterile forceps and scissors to gently tease apart and cut the seminiferous tubules so that 20 millimeter individual segments are obtained. Next, transfer five 20 millimeter tubule segments to the prepared slide.

Then continue to break up the segments into 1.5 to three millimeter lengths using scissors. Now with the forceps, arrange the segments so that they do not overlap. Distribute them evenly across the slide.

Ideally, 20 to 40 short lengths will occupy one slide. Begin by removing excess liquid on the slide preparation using an absorbent tissue. Now, apply a cover slip and squash the tubules using pressure from the heel of the palm for 10 to 20 seconds.

The goal is to apply enough force to disperse the spermatocytes from the seminiferous tubules. Using a dissection microscope, check that the segments were disrupted by the squash. Then, pour a little liquid nitrogen into a small Dewar and flash freeze the slide for 15 seconds or until the liquid ceases to bubble.

To proceed with amino blotting, immediately remove the cover slip using a sharp point or edge. Although not ideal, frozen slides can be immediately preserved at minus 80 degrees Celsius and saved for up to two weeks. To remove the cover slip later, immerse the slides in liquid nitrogen for 15 seconds and pry off the cover slip.

To proceed with the amino labeling, wash the slides three times in PBS for five minutes per wash using a 50 milliliter Coplin jar. After the washes, apply one milliliter of antibody dilution buffer onto each slide to serve as a block. Allow the block reaction to run for one to two hours in a humidified chamber.

Never let the slides dry out during this procedure. Next, tap the slides onto a paper towel to gently remove the buffer and return them to the humidified chamber. Then, apply 100 microliters of diluted primary antibody to each slide and transfer them to 40 degrees Celsius where the binding reaction can run overnight.

If antibody solution is at a premium, half the volume can be used if it is covered with a cover slip or Parafilm. The next day, use three washes in PBS to remove the primary antibody. Next, apply 100 microliters of diluted secondary antibody and allow this reaction to go for one to one and a half hours at room temperature.

To avoid photobleaching, keep the slides in a dark humidified chamber during this incubation. Next, wash the slides twice with PBS. Now, dot the slides using a mounting medium containing DAPI.

Then seal on cover slips using nail polish and proceed with imaging. To take images, use an epifluorescence microscope with an automated stage that enables precise z-axis movement. Also use a high resolution camera to capture images.

Now, assess the slides using a 20x objective. Determine the quality of the squash preparation. A good quality squash will have a monolayer of nuclei that are evenly distributed.

Some regions of the squash may be better than others. Note the coordinates of the best regions to easily find them under higher magnification. Now, increase the magnification to up to 100x.

Then capture images from noted regions that have a consistent monolayer of nuclei. Set the upper and lower z-stacks to ensure that all of the in-focus light is captured. The optimal number of z-stacks is generally designated by the image acquisition software and is different for each objective.

From the images, compile an extended depth image using processing software. The software will combine the in-focus light from each z-stack, which is essential for optimal imaging of tubule squash preparations. Using the described tubule squash method, transitory cell populations undergoing the prophase to metaphase one transition were visualized.

Enriched populations of metaphase one spermatocytes were visualized using the antibodies against alpha tubulin. To visualize the cells in prophase one, tubule squash preparations were immunolabeled with an antibody to the Synaptonemal Complex Protein 3. Leptotene, zygotene, and pachytene diplotene staged spermatocytes were both identified.

Antibodies against the cell cycle kinase Aurora B were used to visualize later events of meiosis one. This kinase localizes to the inner centromere during metaphase, then relocalizes to the spindle mid zone during anaphase, and finally to the cleavage furrow during cytokinesis. By maintaining cellular integrity, the tubule squash technique allows for the visualization of subcellular organization in structures associated with prophase one.

Chromosome bouquet dynamics can be followed through prophase one. Centriole duplication in centrosome disjunction can also be visualized. In early diplotene stage spermatocyte undergoing centrosome disjunction following centriole duplication was visualized with antibodies against pericentrin, a protein component of the pericentriolar matrix and antibodies against Centrin-3 to mark centrioles.

After watching this video, you should have a good understanding of how to rapidly assess the cytological features of developing mouse spermatocytes while preserving cellular integrity. Once mastered, dissection through squashing can be accomplished in under one hour per mouse. While attempting this procedure, it's important to remember to apply enough force to disperse the spermatocytes from the seminiferous tubules.

In addition, the Fixative concentration and incubation time can be optimized depending on the cellular structure to be studied. Alternative visualization strategies, such as DNA and RNA fish approaches, can be easily implemented with this technique. By complementing tubule squashes with downstream biochemical assays, it is possible to assess the mechanistic determinants of successful meiotic progression.