We thank Trent Newman and Masuda Sharifi for comments on the manuscript and An Nguyen for helping to optimize methods for spreading and staining chromosomes from zebrafish meiocytes. This work was supported by NIH R01 GM079115 awarded to S.M.B.

All methods involving zebrafish were carried out using ethical standards approved by the Institutional Animal Care and Use Committee at UC Davis.
1. Chromosome spreading procedure
NOTE: The following protocol is designed to create 4-6 slides, with hundreds of spread meiotic nuclei per slide. The numbers of testes used will depend on the size of the fish. Expect to use 20 animals at ~60 days post fertilization (dpf) and 15 animals at ~6 months post fertilization (mpf). For large zebrafish (e.g., 12 mpf) 10 animals should be sufficient. Be aware that testes of some meiotic mutants (e.g., spo11-/-) will be somewhat smaller3. In this protocol, one pool of testes is considered as a single sample. It is recommended that no more than 4 samples are prepared in parallel.
<ol>
	<li>Make the following solutions before starting the spreading procedure 
	NOTE: It is convenient to prepare the following solutions in the quantities specified for use in future spreading experiments.
	<ol>
		<li>Prepare a 0.1 M sucrose solution by dissolving 3.42 g of sucrose in 100 mL of distilled water. Bring the sucrose solution to pH 8 using 1 M Tris-HCl that is also at pH 8. Then filter sterilize the sucrose solution. Store at room temperature.</li>
		<li>Make a 10x phosphate-buffered saline (PBS) stock solution to a final concentration of 1.37 M NaCl, 27 mM KCl, 73 mM Na2HPO4 and 27.6 mM KH2PO4 in 1.8 L of distilled water. Stir until dissolved, then adjust pH to 7.3 using NaOH and autoclave. Store at room temperature.</li>
		<li>Make a 400 &#181;g/mL DNase I solution in sterile distilled water. Store at -20 &#176;C. It is recommended to prepare 5 mL of this solution and then store as 100 &#181;L aliquots.</li>
	</ol>
	</li>
	<li>Make the following solutions on the day of the spreading protocol 
	NOTE: For time efficiency, it is best to make the collagenase solution immediately after dissecting all testes and then make the trypsin and trypsin inhibitor solutions during the time that the samples are being treated in collagenase.
	<ol>
		<li>Dissolve 4 mg of collagenase in 200 &#181;L of Dulbecco&#39;s modified Eagle medium (DMEM) per sample (2% w/v collagenase final concentration). Keep on ice until needed. The collagenase will dissociate the testes.</li>
		<li>Dissolve 1.4 mg of trypsin in 200 &#181;L of DMEM per sample (0.7% w/v trypsin final concentration). Keep on ice until needed. The trypsin will help dissociate the cells.</li>
		<li>Dissolve 10 mg of trypsin inhibitor in 500 &#181;L of DMEM per sample (2% w/v trypsin inhibitor final concentration). Keep on ice until needed. Trypsin inhibitor prevents trypsin from degrading the cells.</li>
		<li>Prepare a 1% formaldehyde solution (from a 16% pre-made solution) with 0.15% Triton X-100 in sterile distilled water. Keep on ice until needed. The 1% formaldehyde can be prepared while the testes are being treated with trypsin and DNase I (i.e., during step 1.4.7 of the spreading procedure). 
		CAUTION: Formaldehyde is hazardous.</li>
	</ol>
	</li>
	<li>Dissection of testes from adult male zebrafish
	<ol>
		<li>Euthanize male zebrafish (&#62; 60 dpf) by submerging them in ice water. The fish can be kept in ice water until ready to dissect, however they should be dissected as soon as possible. 
		NOTE: The wild-type strain AB is used for this procedure, but other wild-type strains should also be amenable. The longest time we have kept fish in ice water prior to dissecting is 3 h without any noticeable effect on the protocol.</li>
		<li>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.</li>
		<li>Dissect the testis using forceps (see Table of Materials) at 1.65x magnification under a microscope. Carry out the dissections in a silicone-coated Petri dish with covered by a shallow pool of 1x PBS. 
		NOTE: The testis is located in between the swim bladder and the intestine and will appear lighter than muscle tissue (Figure 2A). The testis should have 2 lobes when removed from the zebrafish (Figure 2B).</li>
		<li>Remove as much fat and surrounding tissue from the testis as possible. Then add each dissected testis directly into a 5 mL tube with 2 mL of DMEM and keep on ice. 
		NOTE: Steps 1.3.3 and 1.3.4 should be mastered prior to doing any experiments.</li>
	</ol>
	</li>
	<li>Dissociation of testes cells
	<ol>
		<li>Pre-warm 100 mL of 0.1 M sucrose solution to 37 &#176;C.</li>
		<li>Add 200 &#181;L of collagenase solution to the 5 mL tube with the testes. Mix the solution by inverting it several times.</li>
		<li>Gently shake the testes in an incubator shaker horizontally at 100 rpm at 32 &#176;C for 50 min to an hour until the DMEM is cloudy and the testes are in small chunks. Rapidly invert the tube every 10 min to facilitate dissociation.</li>
		<li>To wash out the collagenase, add DMEM to a final 5 mL volume and invert the tube a few times. Pellet the testes at ~200 x g for 3 min at room temperature. Remove 3 mL of the supernatant so that only 2 mL remains. 
		NOTE: The addition, removal, and transfer of DMEM is done with plastic transfer pipettes for the entirety of the chromosome spread procedure.</li>
		<li>Repeat step 1.4.4 twice more for a total of 3 DMEM washes. Do not resuspend the pellet between DMEM washes. After the last DMEM wash, remove 4 mL of the supernatant so that only 1 mL remains.</li>
		<li>Add 1 mL of DMEM for a total volume of 2 mL and add 200 &#181;L of the trypsin solution and 20 &#181;L of DNase I solution. Invert the tube a few times to mix the solution.</li>
		<li>Horizontally shake the tube at 32 &#176;C for 5-15 min at 100 rpm until the DMEM solution contains only a few clumps. Rapidly invert the tube every 5 min to facilitate dissociation. 
		NOTE: The clumps should be considerably smaller after shaking.</li>
		<li>Add 500 &#181;L of the trypsin inhibitor solution and 50 &#181;L of DNase I solution. Invert the tube a few times to mix, then briefly spin down at ~200 x g for 3 min at room temperature to remove liquid or clumps that may adhere to the tube cap or the side of the tube.</li>
		<li>Place the tube on ice and resuspend the cell suspension by repeatedly pipetting up and down with a plastic transfer pipette for 2 min to facilitate dissociation of any remaining clumps. Do not let the cell suspension go into the bulb of the transfer pipette. After the 2 min, pipette the suspension back into the 5 mL tube.</li>
		<li>Pre-wet a 100 &#181;m cell strainer with DMEM and place it on top of a 50 mL tube on ice.</li>
		<li>Transfer the cell suspension through the strainer one drop at a time using a plastic transfer pipette. Ensure that the cell suspension does not go into the bulb of the transfer pipette.</li>
		<li>Transfer the filtrate using a plastic transfer pipette to a new 5 mL tube and add DMEM to a final volume of 5 mL. Be sure to collect the pooled cells attached to the underside of the filter with a clean plastic transfer pipette. Pellet the cells at ~200 x g for 5 min at room temperature.</li>
		<li>Remove as much supernatant as possible without disturbing the pellet.</li>
		<li>Add 5 &#181;L 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 4-5 times along an empty 1.5 mL microcentrifuge tube rack. Scraping too harshly can result in a reduction of recovered spreads.</li>
		<li>Add DMEM to a final volume of 5 mL and mix the solution by inverting the tube several times. It is common for clumps to still be present after the first DNase I treatment.</li>
		<li>Pellet the cell suspension at ~200 x g for 2 min at room temperature.</li>
		<li>Repeat steps 1.4.13-1.4.16 an additional 1-3 times until the resuspended pellet does not clump upon addition of DMEM. After the last spin, remove as much supernatant as possible without disturbing the pellet. 
		NOTE: Expect a reduction in pellet size with every DNase I treatment; exceeding 4 total DNase I washes may result in significant loss of cells. If no pellet is visible after the DNase I treatments steps, do not proceed with the procedure. Discard any unused DNase I.</li>
		<li>Add 1 mL of 1x PBS and resuspend the pellet by gently scraping the outside bottom of the tube along an empty 1.5 mL tube rack.</li>
		<li>Pellet the cell suspension at ~200 x g for 5 min then remove as much supernatant as possible without disturbing the pellet.</li>
		<li>Cut ~3 mm from the end of a 200 &#181;L pipette tip to widen the aperture and resuspend the pellet in ~80 &#181;L of 0.1 M sucrose solution by pipetting up and down with the cut pipette tip. Let the cell suspension sit at room temperature for 3 min.</li>
	</ol>
	</li>
	<li>Spreading chromosomes on glass slides
	<ol>
		<li>Coat one slide with 100 &#181;L of 1% formaldehyde with 0.15% Triton X-100 with the side of a pipette tip. Then add 18 &#181;L 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 (~60&#176;) to facilitate spreading of the cell suspension to all corners.</li>
		<li>Place the slides flat down in a slightly cracked open humidity chamber (Figure 3) to prevent the formaldehyde solution from drying. Place the humidity chamber in a dark drawer overnight.</li>
		<li>Remove the lid from the humidity chamber and allow the slides to fully dry.</li>
		<li>Place the slides in a Coplin jar then fill the Coplin jar with distilled water and incubate for 5 min with gentle shaking at room temperature. 
		NOTE: The slides can be placed in the Coplin jar in a zigzag pattern to maximize the number of slides per Coplin jar. Make sure there is space for liquid to pass between all the slides.</li>
		<li>Pour out the water and fill the jar with 1:250 wetting agent (see Table of Materials). Wash 2 times for 5 min each with gentle shaking at room temperature.</li>
		<li>Allow the slides to fully dry and store at -20 &#176;C until they are stained. 
		NOTE: The longest we have stored slides without any noticeable degradation of the chromosomes is ~2 months.</li>
	</ol>
	</li>
</ol>
2. Telomere PNA probe staining
NOTE: Telomere repeats can be stained using fluorophore-conjugated telomere PNA probes that hybridize to leading strand telomere repeats (CCCTAA). PNA probes have a neutral backbone, which increases hybridization affinity to negatively charged DNA, resulting in little to no background. The telomere probing step is optional. To proceed to antibody staining, rehydrate slides in 1x PBS as indicated in step 2.2.2., then proceed directly to &#34;Primary antibody staining&#34;.
<ol>
	<li>Make the PNA probe reagents before starting the telomere staining 
	NOTE: It is convenient to prepare the following solutions in the quantities specified for use in future experiments. Storage conditions are noted below.
	<ol>
		<li>Prepare 2 L of 20x saline-sodium citrate (SSC) solution with a final concentration of 3 M NaCl and 0.3 M sodium citrate. Autoclave and store at room temperature.</li>
		<li>Make 50 mL of pre-hybridization solution containing transfer RNA to a final concentration of 50% formamide, 5x SSC, 50 &#181;g/mL heparin, 500 &#181;g/mL transfer RNA, 0.1% Tween 20 and add 460 &#181;L of 1 M citric acid to bring the solution to pH ~6. Store at -20 &#176;C. 
		CAUTION: Formamide is hazardous. Prepare the solution in a fume hood.</li>
		<li>Make 50 mL of pre-hybridization solution prepared in the same way as in step 2.1.2 without adding transfer RNA. Store at -20 &#176;C.</li>
		<li>PNA telomere probes TelC-Cy3 and TelC-Alexa647 are prepared as 50 &#181;M stocks in formamide as per manufacturer&#39;s instructions. Store the PNA probes as 8 &#181;L aliquots at -80 &#176;C.</li>
		<li>Make at least 1 mL of 100 mg/mL stock solution of bovine serum albumin (BSA) in sterile distilled water. Store at -20 &#176;C. If preparing more than 1 mL of stock BSA, store the solution as 1 mL aliquots.</li>
		<li>Using a 2 mL tube, prepare 2 mL of hybridization solution by adding 27 &#181;L of 100 mg/mL BSA and 8 &#181;L of 50 &#181;M PNA probe to 1.965 mL of pre-hybridization solution containing transfer RNA. Store in the dark at -20 &#176;C.</li>
	</ol>
	</li>
	<li>PNA telomere probe staining
	<ol>
		<li>Preheat the hybridization oven to 82 &#176;C.</li>
		<li>Re-hydrate slides with 500 &#181;L of 1x PBS per slide for 5 min in a humidity chamber at room temperature. Remove the PBS by tapping the side of the slide on a paper towel.</li>
		<li>Heat the slides and the 2 mL tube containing the hybridization solution directly on a metal surface in the heated hybridization oven for 2 min.</li>
		<li>While keeping the slides on the metal surface, add 100 &#181;L of the hybridization solution per slide and then cover the slides with plastic coverslips (~25 mm x 75 mm) cut to shape out of an autoclave bag. Let slides sit at 82 &#176;C for 10-12 min.</li>
		<li>Place the slides in a humidity chamber in the dark at 37 &#176;C for 16-24 h. From this point forward and for the primary and secondary antibody staining, the slides must be kept in the dark to avoid photobleaching of the fluorescence signal. After the incubation period, the reagents for antibody staining can be prepared during step 2.2.9.</li>
		<li>Remove the coverslips with a pipette tip. 
		NOTE: To facilitate removal of the coverslip, ~50-100 &#181;L of the hybridization solution with no transfer RNA can be gently dispensed between the slide and the coverslip as the coverslip is being lifted.</li>
		<li>Pipette 500 &#181;L of pre-hybridization solution with no transfer RNA onto each slide and place the slides in the humidity chamber for 15 min at room temperature. Remove the solution by tapping the side of the slide on a paper towel. 
		NOTE: The pre-hybridization solution with no transfer RNA is not pre-heated prior to being added onto each slide.</li>
		<li>Pipette 500 &#181;L of 50% pre-hybridization solution with no transfer RNA in 1x PBS to each slide and place the slides in the humidity chamber for 15 min at room temperature. Remove the solution by tapping the side of the slide on a paper towel.</li>
		<li>Transfer the slides to a Coplin jar and wash 3 times in 1x PBS with gentle shaking for 15 min per wash at room temperature.</li>
		<li>Remove the slides from the Coplin jar and remove excess PBS by tapping the side of the slide on a paper towel. Then proceed to step 3.2 &#34;Primary antibody staining&#34;.</li>
	</ol>
	</li>
</ol>
3. Antibody staining
NOTE: Antibodies raised to known meiotic proteins can be used for immunofluorescence detection in spread chromosome preparations. Secondary antibodies conjugated to different fluorophores allows for multiple proteins to be stained simultaneously, if the primary antibodies were raised in different animals.
<ol>
	<li>Make the following solutions on the day of the primary and secondary antibody staining
	<ol>
		<li>Make 1 L of PBT with 1x PBS and 0.1% Triton X-100 diluted in distilled water. Store at room temperature. This can be prepared ahead of time and in large quantities for future spreading experiments.</li>
		<li>Prepare 500 &#181;L of antibody block per slide to a final concentration of 2 mg/mL of BSA and 2% goat serum in PBT.</li>
		<li>To make 100 &#181;L of primary or secondary antibody mix per slide, add the appropriate antibodies at the correct concentration (see Table of Materials) into the antibody block. 
		NOTE: Antibody concentration may need to be determined empirically. In the discussion, we include a list of antibodies we have tried with success (with dilution/concentrations) and those we have tried without success.</li>
	</ol>
	</li>
	<li>Primary antibody staining 
	NOTE: Rabbit anti-human SCP3 and chicken anti-zebrafish Sycp1 are typically used for primary antibody staining.
	<ol>
		<li>After removing excess PBS from the slides, add 500 &#181;L of antibody block per slide and place the slides in a humidity chamber at room temperature for a minimum of 20 min.</li>
		<li>Remove the antibody block by tapping the side of slide on a paper towel.</li>
		<li>Add 100 &#181;L of primary antibody mix in antibody block per slide and cover the slides with a plastic coverslip made from an autoclave bag. Place the slides in a humidity chamber overnight at 4 &#176;C.</li>
		<li>Remove the coverslips with a pipette tip and wash the slides 2 times for a minimum of 5 minutes each with 1x PBS in a Coplin jar with gentle shaking at room temperature.</li>
		<li>Remove the PBS by tapping the side of the slide on a paper towel.</li>
	</ol>
	</li>
	<li>Secondary antibody staining 
	NOTE: Conjugated goat anti-rabbit and anti-chicken antibodies to different fluorophores are used for staining at a 1:1000 concentration.
	<ol>
		<li>Add 500 &#181;L of antibody block per slide and place the slides in the humidity chamber at room temperature for a minimum of 5 min.</li>
		<li>Remove the antibody block by tapping the side of the slide on a paper towel.</li>
		<li>Add 100 &#181;L of secondary antibody mix in antibody block per slide and cover the slides with a plastic coverslip made from an autoclave bag. Place the slides in a humidity chamber for 1 h at 37 &#176;C.</li>
		<li>Remove the coverslips with a pipette tip and wash the slides 3 times for a minimum of 5 min each with 1x PBS in a Coplin jar with gentle shaking at room temperature.</li>
		<li>Rinse the slides one time for a minimum of 2 min with distilled water in a Coplin jar with gentle shaking at room temperature.</li>
		<li>Air dry the slides tilted, to facilitate drying.</li>
		<li>Place 20 &#181;L of anti-fade mountant with or without DAPI (see Table of Materials) as 3 evenly spaced dots on glass coverslips (24 mm x 60 mm).</li>
		<li>Place the slides face down on the glass coverslips then seal the coverslips with nail polish on the edge overlapping the frosted end of the slide.</li>
		<li>Store the prepared slides at 4 &#176;C until ready for imaging.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here we describe methods to probe the location of telomeres and chromosome-associated proteins in nuclear surface spreads from spermatocytes isolated from zebrafish testes. We expect that these methods will be applicable for analysis of spermatocytes in other teleost species with adjustment to the size of the testis.
While only a few antibodies have been raised to zebrafish meiotic proteins, we have had success using the following antibodies raised to human (h) or mouse (m) proteins. Our lab h...

Sexual reproduction proceeds through the combination of two haploid gametes, each carrying half the chromosome complement of a somatic cell. Meiosis is a specialized cell division that produces haploid gametes through one round of DNA replication and two successive rounds of chromosome segregation. In prophase I, homologous chromosomes (homologs) must undergo pairing, recombination, and synapsis, the latter of which is characterized by the formation of the synaptonemal complex that comprises two homolog axes bridged by the transverse filament, Sycp1 (Figure 1A,B). Failure to properly execute these processes can lead to the production of aneuploid gametes, which are a leading cause of miscarriages in humans1. Our knowledge of the coordination between pairing, recombination and synapsis has been facilitated by studies in a wide range of organisms, such as yeast, C. elegans, mouse, and Drosophila, among others2. While the general process of homologous chromosome pairing followed by segregation is well conserved, its dependency on recombination and synapsis and the order of these events varies.
Meiotic double-strand break (DSB) formation, which initiates homologous recombination, occurs near telomeres clustered in the bouquet during leptotene and synapsis ensues shortly after3,4. This configuration of DSB formation and synapsis initiation is also a characteristic of male meiosis in humans but not in mouse5,6,7,8, suggesting that zebrafish can serve as a model for human spermatogenesis. There are also several practical advantages of studying zebrafish meiosis. Both males and females undergo gametogenesis throughout adulthood, their gonads are easily accessible, and hundreds of offspring are generated from a single cross. Additionally, the embryos are transparent and develop externally, which facilitates the early detection of aberrations in embryonic development due to aneuploid gametes3,9. Disadvantages of using zebrafish are that they are slow to reach sexual maturity (~60 days) and the amount of material needed for nuclear surface spreads must be collected from ~10-20 adult animals, depending on their size.
Meiotic chromosome spread preparations are a vital tool for studying chromosome dynamics across all model organisms, since key signatures of meiotic chromosome dynamics can be probed. In zebrafish, key aspects of the progression of the meiotic program and nuclear organization have been dissected through probing nuclear surface spreads, referred to here as chromosome spreads, with antibodies for immunofluorescence detection of proteins and/or nucleic acids by FISH3,4,9,10,11,12. Indeed, the polarized localization of clustered telomeres in the bouquet can be preserved in the spread preparation (Figure 1C). Recently, we have used zebrafish spermatocyte chromosome spreads together with fluorescence detection methods and super-resolution microscopy to elucidate the detailed progression of zebrafish telomere dynamics, homologous chromosome pairing, double-strand break localization, and synapsis at key meiotic transitions3. Here we present methods to prepare chromosome spreads from spermatocytes of the zebrafish testes and subsequently stain them with fluorescent peptide nucleic acid (PNA) probes to repeated telomere sequences and immunofluorescence detection of chromosome associated proteins.

<table><tbody><tr><td>1.5 mL centrifuge tubes</td><td>Several commercial brands available</td><td></td><td></td></tr><tr><td>1.5 mL microcentrifuge tube rack</td><td>Several commercial brands available</td><td></td><td></td></tr><tr><td>16% formaldehyde, methanol-free</td><td>ThermoFisher Scientific</td><td>28908</td><td></td></tr><tr><td>2 mL</td><td>Several commercial brands available</td><td></td><td></td></tr><tr><td>24 x 50 mm glass coverslips</td><td>Corning</td><td>2980-245</td><td></td></tr><tr><td>24 x 60 mm glasscoverslips</td><td>VWR International</td><td>16004-312</td><td></td></tr><tr><td>50 mL conical centrifuge tubes</td><td>ThermoFisher Scientific</td><td>363696</td><td></td></tr><tr><td>Autoclave bag</td><td>Several commercial brands available</td><td></td><td>Used to make plastic coverslips.</td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Fisher Scientific</td><td>BP1605-100</td><td>Prepare a 100 mg/ml stock solution in sterile distilled water.</td></tr><tr><td>Cell Strainer, 100 &amp;micro;m</td><td>Fisher Scientific</td><td>08-771-19</td><td></td></tr><tr><td>CF405M goat anti-chicken IgY (H+L), highly cross-adsorbed</td><td>Biotium</td><td>203775-500uL</td><td>Use at 1:1000</td></tr><tr><td>Chicken anti-zfSycp1</td><td>Generated by Burgess lab</td><td>N/A</td><td>Use at 1:100</td></tr><tr><td>Collagenase from &lt;em&gt;Clostridium&lt;/em&gt; histolyticum</td><td>Sigma-Aldrich</td><td>C0130-500MG</td><td></td></tr><tr><td>Coplin jar</td><td>Several commercial brands available</td><td></td><td></td></tr><tr><td>DNase I, grade II from bovine pancreas</td><td>Roche Diagnostics</td><td>10104159001</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium (DMEM)</td><td>Fisher Scientific</td><td>MT10014CV</td><td></td></tr><tr><td>Dumont No. 5 Forceps</td><td>Fine Science Tools</td><td>11252-30</td><td>Two are required for dissecting the testes.</td></tr><tr><td>Eppendorf Tubes, 5 mL</td><td>VWR International</td><td>89429-308</td><td></td></tr><tr><td>Formamide</td><td>Fisher Scientific</td><td>BP228-100</td><td></td></tr><tr><td>Goat anti-chicken IgY (H+L) secondary antibody, Alexa Fluor 488</td><td>ThermoFisher Scientific</td><td>A-11039</td><td>Use at 1:1000</td></tr><tr><td>Goat anti-chicken IgY (H+L) secondary antibody, Alexa Fluor 594</td><td>ThermoFisher Scientific</td><td>A-11042</td><td>Use at 1:1000</td></tr><tr><td>Goat anti-hDMC1</td><td>Santa Cruz Biotechnology</td><td>sc-8973</td><td>Does not work in our hands</td></tr><tr><td>Goat anti-rabbit IgG (H+L) cross-adsorbed secondary antibody, Alexa Fluor 488</td><td>ThermoFisher Scientific</td><td>A-11008</td><td>Use at 1:1000</td></tr><tr><td>Goat anti-rabbit IgG (H+L) cross-adsorbed secondary antibody, Alexa Fluor 594</td><td>ThermoFisher Scientific</td><td>A-11012</td><td>Use at 1:1000</td></tr><tr><td>Goat serum</td><td>Sigma-Aldrich</td><td>G9023-10mL</td><td></td></tr><tr><td>Heparin sodium salt</td><td>Sigma-Aldrich</td><td>H3393-100KU</td><td></td></tr><tr><td>Humidity chamber</td><td>Fisher Scientific</td><td>50-112-3683</td><td></td></tr><tr><td>Hybridization Oven</td><td>VWR International</td><td>230401V (Model 5420)</td><td></td></tr><tr><td>Incubator Shaker</td><td>New Brunswick Scientific</td><td>Model Classic C25</td><td></td></tr><tr><td>KCl</td><td>Fisher Scientific</td><td>P217-500</td><td></td></tr><tr><td>Kimwipes</td><td>Kimerbly-Clark Professional</td><td>34155</td><td>Used for the humidity chamber</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher Scientific</td><td>P285-500</td><td></td></tr><tr><td>Microscope</td><td>Several commercial brands available</td><td></td><td>Any standard microscope capable of at least ~1.65X magnification is sufficient.</td></tr><tr><td>Microscope slides</td><td>Fisher Scientific</td><td>12-544-7</td><td></td></tr><tr><td>Mouse anti-hamsterSCP3</td><td>Abcam</td><td>ab97672</td><td>Does not work in our hands</td></tr><tr><td>Mouse anti-hMLH1</td><td>BD Biosciences</td><td>550838</td><td>Does not work in our hands</td></tr><tr><td>Mouse anti-hRPA</td><td>Sigma-Alrich</td><td>MABE285</td><td>Does not work in our hands</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; &amp;middot; 7 H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Fisher Scientific</td><td>S373-500</td><td></td></tr><tr><td>NaCl</td><td>Fisher Scientific</td><td>S271-3</td><td></td></tr><tr><td>Photo-Flo 200 solution</td><td>Electron Microscopy Sciences</td><td>74257</td><td></td></tr><tr><td>Plastic transfer pipettes</td><td>Several commercial brands available</td><td></td><td></td></tr><tr><td>PNA TelC-Alexa647</td><td>PNA Bio Inc</td><td>F1013</td><td>Prepare as per manufacturer&#039;s instructions.</td></tr><tr><td>PNA TelC-Cy3</td><td>PNA Bio Inc</td><td>F1002</td><td>Prepare as per manufacturer&#039;s instructions.</td></tr><tr><td>ProLong Diamond Antifade Mountant</td><td>ThermoFisher Scientific</td><td>P36970</td><td></td></tr><tr><td>ProLong Diamond Antifade Mountant with DAPI</td><td>ThermoFisher Scientific</td><td>P36971</td><td></td></tr><tr><td>Rabbit anti-hRPA</td><td>Bethyl</td><td>A300-244A</td><td>Use at 1:300</td></tr><tr><td>Rabbit anti-hSCP3</td><td>Abcam</td><td>ab150292</td><td>Use at 1:200</td></tr><tr><td>Rabbit anti-hRad51</td><td>GeneTex</td><td>GTX100469</td><td>Use at 1:300</td></tr><tr><td>Sodium citrate</td><td>Fisher Scientific</td><td>S279-500</td><td></td></tr><tr><td>Sucrose</td><td>Fisher Scientific</td><td>S5-500</td><td></td></tr><tr><td>Supercut Scissors, 30&amp;deg; angle, 10 cm</td><td>Fisher Scientific</td><td>50-822-353</td><td>Can also use any pair of small scissors.</td></tr><tr><td>Sylgard kit</td><td>Fisher Scientific</td><td>NC9897184</td><td>Prepare as per manufacturer&#039;s instructions.</td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>BP151-100</td><td>Dilute in sterile distilled water to make a 20% working solution. Store at room temperature. Triton X-100 forms a precipitate when diluted in water; precipitate dissolves overnight.</td></tr><tr><td>Trypsin</td><td>Worthington Biochemical</td><td>LS003708</td><td></td></tr><tr><td>Trypsin inhibitor from chicken egg white</td><td>Sigma-Aldrich</td><td>T9253-500MG</td><td></td></tr><tr><td>Tween 20</td><td>Bio-Rad</td><td>170-6531</td><td>Dilute in sterile distilled water to make a 20% working solution. Store at room temperature.</td></tr><tr><td>Vannas Spring Scissors - 4 mm (micro scissors)</td><td>Fine Science Tools</td><td>15018-10</td><td></td></tr></tbody></table>

preparation of meiotic chromosome spreads from zebrafish spermatocytes

Meiosis is the key cellular process required to create haploid gametes for sexual reproduction. Model organisms have been instrumental in understanding the chromosome events that take place during meiotic prophase, including the pairing, synapsis, and recombination events that ensure proper chromosome segregation. While the mouse has been an important model for understanding the molecular mechanisms underlying these processes, not all meiotic events in this system are analogous to human meiosis. We recently demonstrated the exciting potential of the zebrafish as a model of human spermatogenesis. Here we describe, in detail, our methods to visualize meiotic chromosomes and associated proteins in chromosome spread preparations. These preparations have the advantage of allowing high resolution analysis of chromosome structures. First, we describe the procedure for dissecting testes from adult zebrafish, followed by cell dissociation, lysis, and spreading of the chromosomes. Next, we describe the procedure for detecting the localization of meiotic chromosome proteins, by immunofluorescence detection, and nucleic acid sequences, by fluorescence in situ hybridization (FISH). These techniques comprise a useful set of tools for the cytological analysis of meiotic chromatin architecture in the zebrafish system. Researchers in the zebrafish community should be able to quickly master these techniques and incorporate them into their standard analyses of reproductive function.

We have outlined a method to prepare and visualize zebrafish spermatocyte spread preparations. When performed correctly, our procedure yields well spread, non-overlapping nuclei. To recover such nuclei, it is important to have the appropriate amount of starting material (i.e., testes), treat testes for a sufficient length of time in trypsin and an adequate number of DNase I treatments. These spreads can then be stained for telomeres and meiotic proteins to study meiotic progression during...

Watch this Scientific Journal Video about Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes at JoVE.com

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.

Meiosis is the key cellular process required to create haploid gametes for sexual reproduction. Model organisms have been instrumental in understanding ...

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Research

JoVE Journal

Genetics

7.8K Views.  University of California, Davis. 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.

Video: Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

Chromatin Spread Preparations for the Analysis of Mouse Oocyte Progression from Prophase to Metaphase II

Chromatin spread techniques have been widely used to assess the dynamic localization of various proteins during gametogenesis, particularly for spermatogenesis. These techniques allow for visualization of protein and DNA localization patterns during meiotic events such as homologous chromosome pairing, synapsis and DNA repair. While a few protocols have been described in the literature, general chromatin spread techniques using mammalian prophase oocytes are limited and difficult due to the timing of meiosis initiation in fetal ovaries. In comparison, prophase spermatocytes can be collected from juvenile male mice with higher yields without the need for microdissection. However, it is difficult to obtain a pure synchronized population of cells at specific stages due to the heterogeneity of meiotic and post-meiotic germ cell populations in the juvenile and adult testis. For later stages of meiosis, it is advantageous to assess oocytes undergoing meiosis I (MI) or meiosis II (MII), because groups of mature oocytes can be collected from adult female mice and stimulated to resume meiosis in culture. Here, methods for meiotic chromatin spread preparations using oocytes dissected from fetal, neonatal and adult ovaries are described with accompanying video demonstrations. Chromosome missegregation events in mammalian oocytes are frequent, particularly during MI. These techniques can be used to assess and characterize the effects of different mutations or environmental exposures during various stages of oogenesis. As there are distinct differences between oogenesis and spermatogenesis, the techniques described within are invaluable to increase our understanding of mammalian oogenesis and the sexually dimorphic features of chromosome and protein dynamics during meiosis.

Oogenesis in mammals is known to be error-prone, particularly due to chromosome missegregation. This manuscript describes chromatin spread preparation methods for mouse prophase, metaphase I and II-staged oocytes. These fundamental techniques allow for the study of chromatin-bound proteins and chromosome morphology throughout mammalian oogenesis.

Chromatin spread techniques have been widely used to assess the dynamic localization of various proteins during gametogenesis, particularly for ...

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Developmental Biology

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, specifically the generation of homozygous gynogenetic diploids is useful in genetic analysis because it allows the direct production of homozygotes from a single heterozygous mother. This article describes a modified protocol for ploidy duplication based on a heat pulse during the first cell cycle, Heat Shock 2 (HS2). Through inhibition of centriole duplication, this method results in a precise cell division stall during the second cell cycle. The precise one-cycle division stall, coupled to unaffected DNA duplication, results in whole genome duplication. Protocols associated with this method include egg and sperm collection, UV treatment of sperm, in vitro fertilization and heat pulse to cause a one-cell cycle division delay and ploidy duplication. A modified version of this protocol could be applied to induce ploidy changes in other animal species.

A modified protocol for ploidy manipulation uses a heat shock to induce a one-cycle stall in cytokinesis in the early embryo. This protocol is demonstrated in the zebrafish but may be applicable to other species.

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, ...

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

The advent of targeted CRISPR-Cas nuclease technologies has revolutionized the ability to perform precise genome editing in both established and emerging model systems. CRISPR-Cas genome editing systems use a synthetic guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to specific genomic DNA loci, where the Cas endonuclease generates a double-strand break. The repair of double-strand breaks by intrinsic error-prone mechanisms leads to insertions and/or deletions, disrupting the locus. Alternatively, the inclusion of double-stranded DNA donors or single-stranded DNA oligonucleotides in this process can elicit the inclusion of precise genome edits ranging from single nucleotide polymorphisms to small immunological tags or even large fluorescent protein constructs. However, a major bottleneck in this procedure can be finding and isolating the desired edit in the germline. This protocol outlines a robust method for screening and isolating germline mutations at specific loci in Danio rerio (zebrafish); however, these principles may be adaptable in any model where in vivo sperm collection is possible.

CRISPR-Cas technologies have revolutionized the field of genome editing. However, finding and isolating the desired germline edit remains a major bottleneck. Therefore, this protocol describes a robust method for quickly screening F0 CRISPR-injected zebrafish sperm for germline edits using standard PCR, restriction digest, and gel electrophoresis techniques.

The advent of targeted CRISPR-Cas nuclease technologies has revolutionized the ability to perform precise genome editing in both established and emerging ...

Making Gynogenetic Diploid Zebrafish by Early Pressure

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from heterozygous females allow F1 mutagenized females to be screened directly for deleterious mutations in an accelerated forward genetic screen. Streisinger et al.1,2 described methods for making gynogenetic (homozygous) diploid zebrafish by activating zebrafish eggs with ultraviolet light-inactivated sperm and preventing either the second meiotic or the first zygotic cell division using physical treatments (heat or pressure) that deploymerize microtubules. The "early pressure" (EP) method blocks the meiosis II, which occurs shortly after fertilization. The EP method produces a high percentage of viable embryos that can develop to fertile adults of either sex. The method generates embryos that are homozygous at all loci except those that were separated from their centromere by recombination during meiosis I. Homozygous mutations are detected in EP clutches at between 50% for centromeric loci and less than 1% for telomeric loci. This method is reproduced verbatim from the Zebrafish Book3.

This is a method for generating gynogenetic diploid zebrafish embryos (embryos whose only genetic contribution comes from the mother) by blocking the second meiotic division immediately after fertilization with ultraviolet light-inactivated sperm. EP embryos are not fully homozygous due to recombination during the first meiotic division, however they are homozygous at all loci that have not been separated from their centromere by recombination.

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from ...

Biology

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

Isolation of meiotic spermatocytes is essential to investigate molecular mechanisms underlying meiosis and spermatogenesis. Although there are established cell isolation protocols using Hoechst 33342 staining in combination with fluorescence-activated cell sorting, it requires cell sorters equipped with an ultraviolet laser. Here we describe a cell isolation protocol using the DyeCycle Violet (DCV) stain, a low cytotoxicity DNA binding dye structurally similar to Hoechst 33342. DCV can be excited by both ultraviolet and violet lasers, which improves the flexibility of equipment choice, including a cell sorter not equipped with an ultraviolet laser. Using this protocol, one can isolate three live-cell subpopulations in meiotic prophase I, including leptotene/zygotene, pachytene, and diplotene spermatocytes, as well as post-meiotic round spermatids. We also describe a protocol to prepare single-cell suspension from mouse testes. Overall, the procedure requires a short time to complete (4-5 hours depending on the number of needed cells), which facilitates many downstream applications.

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 meiotic spermatocytes is essential to investigate molecular mechanisms underlying meiosis and spermatogenesis. Although there are established ...

Separation of Follicular Cells and Oocytes in Ovarian Follicles of Zebrafish

Zebrafish has become an ideal model to study the ovarian development of vertebrates. The follicle is the basic unit of the ovary, which consists of oocytes and surrounding follicular cells. It is vital to separate both follicular cells and oocytes for various research purposes such as for primary culture of follicular cells, analysis of gene expression, oocyte maturation and in vitro fertilization, etc. The conventional method uses forceps to separate both compartments, which is laborious, time consuming and has high damage to the oocyte. Here, we have established a simple method to separate both compartments using a pulled glass capillary. Under a stereomicroscope, oocytes and follicular cells can be easily separated by pipetting in a pulled fine glass capillary (the diameter depends on the follicle diameter). Compared with the conventional method, this new method has high efficiency in separating both oocytes and follicular cells and has low damage to the oocytes. More importantly, this method can be applied to early-stage follicles including at the pre-vitellogenesis stage. Thus, this simple method can be used to separate follicular cells and oocytes of zebrafish.

Here, we present a simple method for separating follicular cells and oocytes in zebrafish ovarian follicles, which will facilitate investigations of ovarian development in zebrafish.

Zebrafish has become an ideal model to study the ovarian development of vertebrates. The follicle is the basic unit of the ovary, which consists of ...

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

Preparation of Meiotic Chromosome Spreads from Mouse Spermatocytes

Mammalian meiosis is a dynamic developmental process that occurs in germ cells and can be studied and characterized. Using a method to spread nuclei on the surface of slides (rather than dropping them from a height), we demonstrate an optimized technique on mouse spermatocytes that was first described in 1997. This method is widely used in laboratories to study mammalian meiosis because it yields a plethora of high quality nuclei undergoing substages of prophase I. Seminiferous tubules are first placed in a hypotonic solution to swell spermatocytes. Then spermatocytes are released into a sucrose solution to create a cell suspension, and nuclei are spread onto fixative-soaked glass slides. Following immunostaining, a diversity of proteins germane to meiotic processes can be examined. For example, proteins of the synaptonemal complex, a tripartite structure that connects the chromosome axes/cores of homologs together can be easily visualized. Meiotic recombination proteins, which are involved in repair of DNA double-strand breaks by homologous recombination, can also be immunostained to evaluate progression of prophase I. Here we describe and demonstrate in detail a technique widely used to study mammalian meiosis in spermatocytes from juvenile or adult male mice.

Meiosis is the developmental process by which gametes are formed through a single round of DNA replication and two successive rounds of chromosome segregation. Mammalian meiosis can be examined by utilizing a technique to prepare meiotic chromosome spreads. Here, we demonstrate a method of preparing surface-spread nuclei from mouse spermatocytes.

Mammalian meiosis is a dynamic developmental process that occurs in germ cells and can be studied and characterized. Using a method to spread nuclei on ...

Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

Summary

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Chapters in this video

Chromatin Spread Preparations for the Analysis of Mouse Oocyte Progression from Prophase to Metaphase II

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Isolation and Characterization of Single Cells from Zebrafish Embryos

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

Making Gynogenetic Diploid Zebrafish by Early Pressure

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

Separation of Follicular Cells and Oocytes in Ovarian Follicles of Zebrafish

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Preparation of Meiotic Chromosome Spreads from Mouse Spermatocytes