Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

Nuclear surface spreads are an indispensable tool for studying chromosome events during meiosis. Here we demonstrate a method to prepare and visualize meiotic chromosomes during prophase I from zebrafish spermatocytes.

Transcript

Zebrafish is emerging as an excellent vertebrate model for studying the chromosome events of meiosis including homologous chromosome pairing, synapsis and recombination. This nuclear surface spreading protocol has enabled us to visualize key features of meiotic chromosome architecture using super resolution microscopy. Researchers can expect hundreds of well-spread nuclei per slide with less debris than other zebrafish spread protocols.

The most technically challenging part of this procedure is the dissection of the zebrafish testes as it is in close proximity to the skin and is also attached to other organs. Visualizing the dissection procedure will allow researchers to know what to expect when they see the gonad embedded in the tissue of the animal. After euthanizing male zebrafish, decapitate one fish at a time with small scissors and then use micro scissors to cut along the ventral midline to expose the body cavity.

Place the fish in a silicone-coated Petri dish and cover by a shallow pool of 1X PBS. Under a microscope at 1.65X magnification, dissect the testes using forceps. Remove as much as fat and surrounding tissue from the testes as possible.

The gonad will appear translucent under the dissection scope light while the surrounding fat may have a slightly darker appearance. Some amount of fat may be left on the gonad without impeding the procedure. Add each dissected testes directly into a five milliliter tube with two milliliters of DMEM and keep on ice.

To dissociate the testes cells, add 200 microliters of collagenase solution to the five milliliter tube with the testes. Mix the solution by inverting it several times. Gently shake the testes in an incubator shaker horizontally at 100 RPM at 32 degrees Celsius.

Rapidly invert the tube every 10 minutes to facilitate dissociation. After an hour, the DMEM is cloudy and the testes are in small chunks. While the testes are incubated in collagenase, prewarm 100 microliters of 0.1 molar sucrose solution to 37 degrees Celsius.

To wash out the collagenase, add DMEM to a final volume of five milliliters and invert the tube a few times. Pellet the testes at 200 g for three minutes at room temperature. Remove three milliliters of the supernatant so that only two milliliters remain.

Repeat the DMEM wash two additional times. After the last DMEM wash, remove four milliliters of the supernatant and add one milliliter of DMEM, 200 microliters of trypsin solution, and 20 microliters of DNase I solution. Invert the tube a few times to mix the solution.

Horizontally shake the tube at 32 degrees Celsius at 100 RPM. Rapidly invert the tube every five minutes to facilitate dissociation. After five to 15 minutes, the DMEM solution contains only a few clumps.

Add 500 microliters of the trypsin inhibitor solution and 50 microliters of DNase I solution. Invert the tube a few times to mix then briefly spin down at 200 g for three minutes at room temperature to remove liquid or clumps that may adhere to the tube cap or the side of the tube. Place the tube on ice and resuspend the cell suspension by repeatedly pipetting up and down with a plastic transfer pipette for two minutes to facilitate dissociation of any remaining clumps.

Ensure that the cell suspension does not go into the bulb of the transfer pipette. Then pre-wet a 100 micron cell strainer with DMEM and place it on top of a 50 milliliter tube on ice. Transfer the cell suspension through the strainer one drop at a time using a plastic transfer pipette.

Transfer the filtrate using a plastic transfer pipette to a new five milliliter tube. Be sure to collect the pooled cells attached to the underside of the filter. Add DMEM into the tube to a final volume of five milliliters.

Pellet the cells at 200 g for five minutes at room temperature. Remove as much supernatant as possible without disturbing the pellet and add five microliters of DNase I solution directly into the pellet. Then mix the pellet with the DNase I solution by gently scraping the outside bottom of the tube four to five times along an empty microcentrifuge tube rack.

Add DMEM to a final volume of five milliliters and mix the solution by inverting the tube several times. Pellet the cell suspension at 200 g for two minutes at room temperature. Repeat the DNase treatment an additional one to three times until the resuspended pellet does not clump upon addition of DMEM.

After the last spin, remove as much as supernatant as possible without disturbing the pellet. Next, add one milliliter of 1X PBS and resuspend the pellet by gently scraping the outside bottom of the tube along an empty tube rack. Pellet the cell suspension at 200 g for five minutes and then remove as much supernatant as possible without disturbing the pellet.

Cut three millimeters from the end of a 200 microliter pipette tip to widen the aperture. With the cut pipette tip, resuspend the pellet in 80 microliters of 0.1 molar sucrose solution by pipetting up and down. Let the cell suspension sit at room temperature for three minutes.

Next, with a pipette tip, coat one slide with 100 microliters of 1%formaldehyde with 0.15%Triton X-100. Then add 18 microliters of the sucrose cell suspension onto the center of the slide in a straight line perpendicular to the long edge. Tilt the slide back and forth to facilitate spreading of the cell suspension to all corners.

Place the slides flat down in a slightly cracked open humidity chamber to prevent the formaldehyde solution from drying. Place the humidity chamber in a dark drawer overnight. In the morning, remove the lid from the humidity chamber and allow the slides to fully dry.

Place the slides in a coplin jar and then fill the coplin jar with distilled water. Incubate for five minutes with gentle shaking at room temperature. Pour out the water and fill the jar with one to 250 wetting agent.

Wash two times for five minutes each with gentle shaking at room temperature. Allow the slides to fully dry and store at minus 20 degrees Celsius until they are stained. This protocol outlined a method to prepare and visualize zebrafish spermatocyte spread which yields well-spread non-overlapping nuclei.

The panels on the left show three stages of meiotic prophase, leptotene, early zygotene, and pachytene. Telomeres were stained magenta for each stage. An example of poor quality spreads due to insufficient DNase I treatments are shown here.

Meiotic chromosomes stained for SYCP3 indicate overlapping nuclei due to a viscous sucrose cell suspension that prevents cells from properly spreading on the slide. It is very important to start with the correct amount of starting material, treat zebrafish testes for the appropriate amount of time in trypsin, and carry out the necessary number of DNase I treatments as failure to do so can lead to very few nuclei or clumped nuclei. Proper PPE should always be worn when working with formaldehyde.

Following the spreading procedure, chromosomes can be stained to visualize various chromosomal structures as well as double-strand breaks to analyze the timing dependency and localization of meiotic events. Our technique has paved the way for showing that zebrafish may be a better model of human spermatogenesis than mouse based on the chromosome features and the telomere bouquet.

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Journal (Genetics)

Journal (Genetics)

Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

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