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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#181;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 Clostridium 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&#39;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>KH2PO4</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>Na2HPO4 &#183; 7 H2O</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&#39;s instructions.</td>
		</tr>
		<tr>
			<td>PNA TelC-Cy3</td>
			<td>PNA Bio Inc</td>
			<td>F1002</td>
			<td>Prepare as per manufacturer&#39;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&#176; 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&#39;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 ...

preparation-of-meiotic-chromosome-spreads-from-zebrafish-spermatocytes

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

Preparation of rAAV9 to Overexpress or Knockdown Genes in Mouse Hearts

Controlling the expression or activity of specific genes through the myocardial delivery of genetic materials in murine models permits the investigation of gene functions. Their therapeutic potential in the heart can also be determined. There are limited approaches for in vivo molecular intervention in the mouse heart. Recombinant adeno-associated virus (rAAV)-based genome engineering has been utilized as an essential tool for in vivo cardiac gene manipulation. The specific advantages of this technology include high efficiency, high specificity, low genomic integration rate, minimal immunogenicity, and minimal pathogenicity. Here, a detailed procedure to construct, package, and purify the rAAV9 vectors is described. Subcutaneous injection of rAAV9 into neonatal pups results in robust expression or efficient knockdown of the gene(s) of interest in the mouse heart, but not in the liver and other tissues. Using the cardiac-specific TnnT2 promoter, high expression of GFP gene in the heart was obtained. Additionally, target mRNA was inhibited in the heart when a rAAV9-U6-shRNA was utilized. Working knowledge of rAAV9 technology may be useful for cardiovascular investigations.

In this manuscript, a method to prepare recombinant adeno-associated virus 9 (rAAV9) vectors to manipulate gene expression in the mouse heart is described.

Controlling the expression or activity of specific genes through the myocardial delivery of genetic materials in murine models permits the investigation ...

Sample Preparation and Analysis of RNASeq-based Gene Expression Data from Zebrafish

The analysis of global gene expression changes is a valuable tool for identifying novel pathways underlying observed phenotypes. The zebrafish is an excellent model for rapid assessment of whole transcriptome from whole animal or individual cell populations due to the ease of isolation of RNA from large numbers of animals. Here a protocol for global gene expression analysis in zebrafish embryos using RNA sequencing (RNASeq) is presented. We describe preparation of RNA from whole embryos or from cell populations obtained using cell sorting in transgenic animals. We also describe an approach for analysis of RNASeq data to identify enriched pathways and Gene Ontology (GO) terms in global gene expression data sets. Finally, we provide a protocol for validation of gene expression changes using quantitative reverse transcriptase PCR (qRT-PCR). These protocols can be used for comparative analysis of control and experimental sets of zebrafish to identify novel gene expression changes, and provide molecular insight into phenotypes of interest.

This protocol presents an approach for whole transcriptome analysis from zebrafish embryos, larvae, or sorted cells. We include isolation of RNA, pathway analysis of RNASeq data, and qRT-PCR-based validation of gene expression changes.

The analysis of global gene expression changes is a valuable tool for identifying novel pathways underlying observed phenotypes. The zebrafish is an ...

Expedited Radiation Biodosimetry by Automated Dicentric Chromosome Identification (ADCI) and Dose Estimation

Biological radiation dose can be estimated from dicentric chromosome frequencies in metaphase cells. Performing these cytogenetic dicentric chromosome assays is traditionally a manual, labor-intensive process not well suited to handle the volume of samples which may require examination in the wake of a mass casualty event. Automated Dicentric Chromosome Identifier and Dose Estimator (ADCI) software automates this process by examining sets of metaphase images using machine learning-based image processing techniques. The software selects appropriate images for analysis by removing unsuitable images, classifies each object as either a centromere-containing chromosome or non-chromosome, further distinguishes chromosomes as monocentric chromosomes (MCs) or dicentric chromosomes (DCs), determines DC frequency within a sample, and estimates biological radiation dose by comparing sample DC frequency with calibration curves computed using calibration samples. This protocol describes the usage of ADCI software. Typically, both calibration (known dose) and test (unknown dose) sets of metaphase images are imported to perform accurate dose estimation. Optimal images for analysis can be found automatically using preset image filters or can also be filtered through manual inspection. The software processes images within each sample and DC frequencies are computed at different levels of stringency for calling DCs, using a machine learning approach. Linear-quadratic calibration curves are generated based on DC frequencies in calibration samples exposed to known physical doses. Doses of test samples exposed to uncertain radiation levels are estimated from their DC frequencies using these calibration curves. Reports can be generated upon request and provide summary of results of one or more samples, of one or more calibration curves, or of dose estimation.

Expedited Radiation Biodosimetry by Automated Dicentric Chromosome Identification ADCI and Dose Estimation

The cytogenetic dicentric chromosome (DC) assay quantifies exposure to ionizing radiation. The Automated Dicentric Chromosome Identifier and Dose Estimator software accurately and rapidly estimates biological dose from DCs in metaphase cells. It distinguishes monocentric chromosomes and other objects from DCs, and estimates biological radiation dose from the frequency of DCs.

Biological radiation dose can be estimated from dicentric chromosome frequencies in metaphase cells. Performing these cytogenetic dicentric chromosome ...

Pooled shRNA Screen for Reactivation of MeCP2 on the Inactive X Chromosome

Forward genetic screens using reporter genes inserted into the heterochromatin have been extensively used to investigate mechanisms of epigenetic control in model organisms. Technologies including short hairpin RNAs (shRNAs) and clustered regularly interspaced short palindromic repeats (CRISPR) have enabled such screens in diploid mammalian cells. Here we describe a large-scale shRNA screen for regulators of X-chromosome inactivation (XCI), using a murine cell line with firefly luciferase and hygromycin resistance genes knocked in at the C-terminus of the methyl CpG binding protein 2 (MeCP2) gene on the inactive X-chromosome (Xi). Reactivation of the construct in the reporter cell line conferred survival advantage under hygromycin B selection, enabling us to screen a large shRNA library and identify hairpins that reactivated the reporter by measuring their post-selection enrichment using next-generation sequencing. The enriched hairpins were then individually validated by testing their ability to activate the luciferase reporter on Xi.

We report a small hairpin RNA (shRNA) and next generation sequencing-based protocol for identifying regulators of X-chromosome inactivation in a murine cell line with firefly luciferase and hygromycin resistance genes fused to the methyl CpG binding protein 2 (MeCP2) gene on the inactive X chromosome.

Forward genetic screens using reporter genes inserted into the heterochromatin have been extensively used to investigate mechanisms of epigenetic control ...

Ultralow Input Genome Sequencing Library Preparation from a Single Tardigrade Specimen

Tardigrades are microscopic animals that enter an ametabolic state called anhydrobiosis when facing desiccation and can return to their original state when water is supplied. The genomic sequencing of microscopic animals such as tardigrades risks bacterial contamination that sometimes leads to erroneous interpretations, for example, regarding the extent of horizontal gene transfer in these animals. Here, we provide an ultralow input method to sequence the genome of the tardigrade, Hypsibius dujardini, from a single specimen. By employing rigorous washing and contaminant exclusion along with an efficient extraction of the 50 ~ 200 pg genomic DNA from a single individual, we constructed a library sequenced with a DNA sequencing instrument. These libraries were highly reproducible and unbiased, and an informatics analysis of the sequenced reads with other H. dujardini genomes showed a minimal amount of contamination. This method can be applied to unculturable tardigrades that could not be sequenced using previous methods.

Contamination during the genomic sequencing of microscopic organisms remains a large problem. Here, we show a method to sequence the genome of a tardigrade from a single specimen with as little as 50 pg of genomic DNA without whole genome amplification to minimize the risk of contamination.

Tardigrades are microscopic animals that enter an ametabolic state called anhydrobiosis when facing desiccation and can return to their original state ...

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

High-throughput Identification of Gene Regulatory Sequences Using Next-generation Sequencing of Circular Chromosome Conformation Capture (4C-seq)

The identification of regulatory elements for a given target gene poses a significant technical challenge owing to the variability in the positioning and effect sizes of regulatory elements to a target gene. Some progress has been made with the bioinformatic prediction of the existence and function of proximal epigenetic modifications associated with activated gene expression using conserved transcription factor binding sites. Chromatin conformation capture studies have revolutionized our ability to discover physical chromatin contacts between sequences and even within an entire genome. Circular chromatin conformation capture coupled with next-generation sequencing (4C-seq), in particular, is designed to discover all possible physical chromatin interactions for a given sequence of interest (viewpoint), such as a target gene or a regulatory enhancer. Current 4C-seq strategies directly sequence from within the viewpoint but require numerous and diverse viewpoints to be simultaneously sequenced to avoid the technical challenges of uniform base calling (imaging) with next generation sequencing platforms. This volume of experiments may not be practical for many laboratories. Here, we report a modified approach to the 4C-seq protocol that incorporates both an additional restriction enzyme digest and qPCR-based amplification steps that are designed to facilitate a greater capture of diverse sequence reads and mitigate the potential for PCR bias, respectively. Our modified 4C method is amenable to the standard molecular biology lab for assessing chromatin architecture.

High-throughput Identification of Gene Regulatory Sequences Using Next-generation Sequencing of Circular Chromosome Conformation Capture 4C-seq

The identification of physical interactions between genes and regulatory elements is challenging but has been facilitated by chromosome conformation capture methods. This modification to the 4C-seq protocol mitigates PCR bias by minimizing over-amplification of PCR templates and maximizes the mappability of reads by incorporating an addition restriction enzyme digest step.

The identification of regulatory elements for a given target gene poses a significant technical challenge owing to the variability in the positioning and ...

Preparation and Gene Modification of Nonhuman Primate Hematopoietic Stem and Progenitor Cells

Hematopoietic stem and progenitor cell (HSPC) transplantation has been a cornerstone therapy for leukemia and other cancers for nearly half a century, underlies the only known cure of human immunodeficiency virus (HIV-1) infection, and shows immense promise in the treatment of genetic diseases such as beta thalassemia. Our group has developed a protocol to model HSPC gene therapy in nonhuman primates (NHPs), allowing scientists to optimize many of the same reagents and techniques that are applied in the clinic. Here, we describe methods for purifying CD34+ HSPCs and long-term persisting hematopoietic stem cell (HSC) subsets from primed bone marrow (BM). Identical techniques can be employed for the purification of other HSPC sources (e.g., mobilized peripheral blood stem cells [PBSCs]). Outlined is a 2 day protocol in which cells are purified, cultured, modified with lentivirus (LV), and prepared for infusion back into the autologous host. Key readouts of success include the purity of the CD34+ HSPC population, the ability of purified HSPCs to form morphologically distinct colonies in semisolid media, and, most importantly, gene modification efficiency. The key advantage to HSPC gene therapy is the ability to provide a source of long-lived cells that give rise to all hematopoietic cell types. As such, these methods have been used to model therapies for cancer, genetic diseases, and infectious diseases. In each case, therapeutic efficacy is established by enhancing the function of distinct HSPC progeny, including red blood cells, T cells, B cells, and/or myeloid subsets. The methods to isolate, modify, and prepare HSPC products are directly applicable and translatable to multiple diseases in human patients.

The goal of this protocol is to isolate nonhuman primate CD34+ cells from primed bone marrow, to gene-modify these cells with lentiviral vectors, and to prepare a product for infusion into the autologous host. The total protocol length is approximately 48 h.

Hematopoietic stem and progenitor cell (HSPC) transplantation has been a cornerstone therapy for leukemia and other cancers for nearly half a century, ...

A Non-random Mouse Model for Pharmacological Reactivation of Mecp2 on the Inactive X Chromosome

X chromosome inactivation (XCI) is the random silencing of one X chromosome in females to achieve gene dosage balance between the sexes. As a result, all females are heterozygous for X-linked gene expression. One of the key regulators of XCI is Xist, which is essential for the initiation and maintenance of XCI. Previous studies have identified 13 trans acting X chromosome inactivation factors (XCIFs) using a large-scale, loss-of-function genetic screen. Inhibition of XCIFs, such as ACVR1 and PDPK1, using short-hairpin RNA or small molecule inhibitors, reactivates X chromosome-linked genes in cultured cells. But the feasibility and tolerability of reactivating the inactive X chromosome in vivo remains to be determined. Towards this goal, a Xist&#916;:Mecp2/Xist:Mecp2-Gfp mouse model has been generated with non-random XCI due to deletion of Xist on one X chromosome. Using this model, the extent of inactive X reactivation was quantitated in the mouse brain following treatment with XCIF inhibitors. Recently published results show, for the first time, that pharmacological inhibition of XCIFs reactivates Mecp2 from the inactive X chromosome in cortical neurons of the living mouse brain.

Here, we describe a protocol to generate a viable female murine model with non-random X chromosome inactivation, i.e., the maternally-inherited X chromosome is inactive in 100% of the cells. We also describe a protocol to test feasibility, tolerability, and safety of pharmacological reactivation of the inactive X chromosome in vivo.

X chromosome inactivation (XCI) is the random silencing of one X chromosome in females to achieve gene dosage balance between the sexes. As a result, all ...

Identification of Enhancer-Promoter Contacts in Embryoid Bodies by Quantitative Chromosome Conformation Capture (4C)

During mammalian development, cell fates are determined through the establishment of regulatory networks that define the specificity, timing, and spatial patterns of gene expression. Embryoid bodies (EBs) derived from pluripotent stem cells have been a popular model to study the differentiation of the main three germ layers and to define regulatory circuits during cell fate specification. Although it is well-known that tissue-specific enhancers play an important role in these networks by interacting with promoters, assigning them to their relevant target genes still remains challenging. To make this possible, quantitative approaches are needed to study enhancer-promoter contacts and their dynamics during development. Here, we adapted a 4C method to define enhancers and their contacts with cognate promoters in the EB differentiation model. The method uses frequently cutting restriction enzymes, sonication, and a nested-ligation-mediated PCR protocol compatible with commercial DNA library preparation kits. Subsequently, the 4C libraries are subjected to high-throughput sequencing and analyzed bioinformatically, allowing detection and quantification of all sequences that have contacts with a chosen promoter. The resulting sequencing data can also be used to gain information about the dynamics of enhancer-promoter contacts during differentiation. The technique described for the EB differentiation model is easy to implement.

Identification of Enhancer-Promoter Contacts in Embryoid Bodies by Quantitative Chromosome Conformation Capture 4C

We report the application of quantitative chromosome conformation capture followed by high-throughput sequencing in embryoid bodies generated from embryonic stem cells. This technique allows to identify and quantitate the contacts between putative enhancers and promoter regions of a given gene during embryonic stem cell differentiation.

During mammalian development, cell fates are determined through the establishment of regulatory networks that define the specificity, timing, and spatial ...