This work was supported by NIH grants R21 HD068949-01 and RO1 GM065303.

All animal experiments were conducted according to University of Wisconsin &#8211; Madison and Institutional Animal Care and Use Committee (IACUC) guidelines (University of Wisconsin &#8211; Madison Assurance number A3368-01).
1. Selecting Females for Egg Collection via Interrupted Mating
NOTE: IVF-based protocols rely on the extrusion of mature eggs from females through manual pressure 9. Previous protocols have used females directly from tanks or in pair matings without the females undergoing egg release behavior, but only a small fraction of these females (about 20% or less, depending on the zebrafish line) yield extruded and competent eggs upon manual pressure. In an improved procedure, females are pre-sorted for egg laying by direct visual observation, followed by immediate interruption of mating. This procedure is very effective, as nearly all females pre-selected through this interrupted mating step yield extruded and competent eggs upon manual pressure.
<ol>
	<li>The afternoon before the experiment, set up mating pairs of the desired zebrafish strain in standard zebrafish mating boxes. Keep males in the same chamber as females yet physically separated from them, either through a mating box division or by placing the male under an egg-laying insert and the female inside the insert.</li>
	<li>The morning of the experiment, remove the physical partition, placing both the male and female within the same egg-laying insert, so that mating begins.</li>
	<li>Visually inspect tanks containing mating pairs to detect extrusion of eggs during natural mating. At the first signs of egg extrusions, separate male and female to interrupt breeding. After separation from males, keep the pre-selected females either separately or pooled in the same tank. Use multiple females depending on the number of embryos desired (typically 50-150 eggs/female).</li>
</ol>
2. Preparing a Sperm Solution
NOTE: IVF relies on exposure of mature eggs to a sperm solution. This solution can be untreated, to generate diploid zygotes (which upon HS2 treatment become tetraploid embryos) or UV-treated, to generate haploid zygotes (which upon HS2 treatment become homozygous diploid embryos). Previous sperm preparation protocols suggested the use of capillary tubes to collect milt from the anal region of live males, but this was an ineffective process as only a small fraction of males yielded milt 9. The protocol presented below relies instead on sperm preparation from sheared dissected testes, which yields more reliable results.
<ol>
	<li>Prepare Hank&#39;s solution ahead of time as a Hank&#39;s premix solution, comprising of all components (Solutions 1, 2, 4 and 5) except the sodium bicarbonate solution (Solution 6). Prepare Solution 6 fresh and add to the premix the morning of the experiment (Table 1). To make Hanks&#39; solution (final solution, prepare the morning of the IVF procedure), combine 990 ul of Hank&#39;s Premix and 10 &#181;l of the freshly made Solution 6.</li>
	<li>Euthanize males by overexposure to tricaine as a 0.016% solution in conditioned water.
	<ol>
		<li>Prepare tricaine as a 0.2% stock solution in water (buffer to pH 7.0 with 1M Tris pH 9.0) and keep at 4 &#176;C. Add 8 ml of tricaine stock solution per 100 ml of water in a beaker, and use a net to transfer the males to the tricaine solution.</li>
		<li>Use the equivalent of testes for one male per clutch needed to be fertilized in a volume of Hank&#39;s solution corresponding to 100 &#181;l per male (e.g. testes from 10 males collected in 1 ml Hank&#39;s solution, to fertilize 10 clutches with 100 &#181;l/clutch).</li>
		<li>Confirm euthanasia by cessation of gill movements for 15 minutes. After euthanasia, remove the males from the beaker with a spoon. Rinse the males briefly with conditioned water and dry them lightly by placing them briefly on several locations of a paper towel.</li>
	</ol>
	</li>
	<li>To remove the testes, first decapitate euthanized males using dissecting scissors or a razor blade, and make a longitudinal cut along the abdomen with dissecting scissors.Under a dissecting microscope with a reflected light source, remove internal organs with dissecting forceps. Pull out each of the testes masses with forceps and place them inside the microcentrifuge tube containing Hank&#39;s solution. 
	NOTE: Testes can be identified as each of two elongated structures of translucent appearance found alongside the body walls and which converge near the cloaca. Testes can stay 2-3 min on a petri plate surface after dissection and before placing them in the Hank&#39;s solution.</li>
	<li>Release the sperm into the solution by shearing the testes with a narrow spatula and/or gently pipetting up and down 5-6 times the testes in solution with a 1,000 &#181;l-tip micropipette, while avoiding air bubble formation. Let the testes debris settle.</li>
	<li>Store the sperm solution in ice, where it can keep its potency for up to 2-3 hr. If proceeding to UV-treatment of sperm solution, transfer the solution into a clean microcentrifuge tube leaving the pieces of testes behind.</li>
</ol>
3. UV Treatment
NOTE: UV treatment is used to crosslink sperm DNA in order to render it inactive in the embryo. This step is only used when producing gynogenetic haploid or homozygous gynogenetic diploid embryos. Sperm solution for UV treatment should be separated from pieces of testes (step 2.4), as large pieces may shield sperm from the UV treatment.
<ol>
	<li>Transfer the sperm solution to a clean, dry well of a depression glass plate sitting on an ice bed (e.g. ice inside a petri plate). Use up to 1 ml of sperm solution per glass plate well.</li>
	<li>Expose the sperm solution to UV by placing the depression glass plate on the ice bed under a UV lamp. Treat the sperm solution for 90 sec with a 115 V (60 Hz, 0.68A) UV-lamp at a 19 cm (7.5 inches) distance. With the end of a pipette tip, gently mix the solution every 30 seconds during UV treatment (use a clean pipette tip every 30 sec).</li>
	<li>Using a clean pipet tip and micropipette, transfer the treated sperm solution to a new microcentrifuge tube. Store on ice until needed for IVF (no longer than 2-3 hr from extraction).</li>
</ol>
4. Manual Extrusion of Mature Eggs
NOTE: Females obtained by interruption of natural matings will readily yield eggs under anesthesia and manual pressure. During this procedure, tricaine treatment should be carefully controlled to avoid overexposure that may prevent recovery of the females.
<ol>
	<li>Anesthetize females by light exposure to tricaine solution: transfer females with a net to tricaine solution in conditioned water (made by adding 8 ml of Tricaine 0.2% stock solution to 100 ml of conditioned fish water) for about 2-5 min, until fish stop gill movement.</li>
	<li>As soon as a female stops gill movement, use a spoon to collect it and briefly rinse it in conditioned water. Still using the spoon to move the fish, dry it lightly by placing it briefly and repeatedly on several locations of a paper towel, then transfer it to a clean, dry 10 cm diameter petri plate. Approach the fish with the spoon in the anterior to posterior direction, to avoid potentially damaging the gill operculum.</li>
	<li>Use lab wipes or soft tissue to gently further dry the anal fin area, to prevent any released eggs from prematurely being activated by water. Dampening the fingers lightly with water (to avoid them sticking to fish scales), place one finger of one hand on the female&#39;s back as support, and with a finger of the other hand apply slight pressure along the female&#39;s abdomen until eggs become extruded.</li>
	<li>Use a narrow spatula to move the eggs away from the female&#39;s body. Place the female back into a tank with conditioned water for recovery. 
	NOTE: Once eggs are extruded while the female is lying on one side of her body, she can be flipped over and the process repeated on her other side to obtain additional eggs. To insure full recovery of the female, apply only gentle manual pressure on the abdomen, and carry out the procedure from gill movement stop to return to water in less than 2 min.</li>
	<li>Activate (with water exposure) and fertilize (with sperm solution addition) eggs simultaneously and within 90 sec after extrusion (see section 5).</li>
</ol>
5. In Vitro Fertilization
NOTE: Zebrafish fertilization in natural crosses is external, dependent on the simultaneous release and activation by water of eggs and sperm during mating. In vitro fertilization mimics this process by exposing the eggs to sperm solution in the presence of water. Water volume is originally small (1 ml) in order to increase the effective sperm concentration. Binding by sperm occurs within 15-20 sec 10, and water volume can then be increased. Chorion lifting further contributes to the close synchronization of the clutch by limiting the window for competence for sperm binding 11,12. The resulting embryos therefore exhibit largely simultaneous cell division cycles during the early cleavage stages.
<ol>
	<li>Add 100 &#181;l of sperm solution (corresponding to the equivalent of testes from one male &#8211; see Part 2) to the clutch of extruded eggs on a petri plate. Gently swirl the pipette tip used to add the sperm solution among the eggs to mix the sperm and eggs together, being sure to not lift the tip from the surface of the petri plate to avoid egg damage.</li>
	<li>Immediately activate the eggs by adding 1 ml of embryonic medium (E3) solution (conditioned water is also acceptable as a substitute for E3 in Parts 5 through 7) and again gently mix eggs and sperm by gently swirling with the pipette tip. Start a timer to initiate timing relative to fertilization.</li>
	<li>At 1 mpf in the 1 ml volume, flood the plate with E3. Before continuing, leave undisturbed until 10-12 mpf to allow egg activation, including full chorion expansion.</li>
</ol>
6. Heat Shock Treatment
NOTE: A heat shock applied in the early embryo inhibits centriole duplication, resulting in an incomplete complement of centrioles to drive spindle formation during the subsequent cell cycle 2. The absence of spindle in turn results in the lack of furrow formation 13.
<ol>
	<li>After expansion of the chorions (10-12 mpf), pour the embryos from the petri plate into a tea strainer. Rinse the petri plate using a wash bottle with E3 in order to collect any remaining embryos in the tea strainer.</li>
	<li>Place the tea strainer with the embryos inside a beaker in a pre-heated bath with E3 at 28.5 &#176;C. Pre-equilibrate the beaker and E3 to the water bath temperature, and make sure there is enough E3 so that all embryos in the tea strainer are exposed to the medium.</li>
	<li>At 22 mpf, remove the tea strainer from the 28.5 &#176;C water bath, briefly blot it onto a paper towel to remove excess moisture, and place it inside a similarly preheated E3 beaker in a heat bath at 41.4 &#176;C.</li>
	<li>At 24 mpf, transfer the tea strainer back to the E3 in the water bath at 28.5 &#176;C after brief blotting. At 29 mpf, use a wash bottle with E3 to transfer the embryos from the tea strainer to a 10 cm Petri plate.</li>
</ol>
7. Selection for Embryos with a One-cycle Cytokinesis Stall
<ol>
	<li>During the time period 35-45 mpf, under a dissecting microscope with a transmitted light source, select those embryos that are undergoing symmetrical cleavage into the 2-cell stage, and which are therefore fertilized. Remove embryos that are not undergoing cell cleavage.</li>
	<li>Continue observing the fertilized embryos, selected for normal cell division during the first cell cycle, and during the second cell cycle (50-65 mpf).</li>
	<li>Sort embryos according to the following categories (Figure 2C): 4 cells (&#34;no stall&#34;, in the 2:2 arrangement standard for a 4-cell embryo); 3 cells (&#34;partial stall&#34;, in an aberrant 2 smaller cells:1 large cell arrangement); and 2 cells (&#34;stalled&#34;, embryos exhibiting a one-cell cycle delay in a 1:1 arrangement identical to that of a 2-cell embryo). Sort the stalled embryos into a fresh petri plate. 
	NOTE: At this stage, embryos in the 2:2 arrangement correspond to those in which during the second cell cycle neither blastomere underwent a cell division stall; embryos in an aberrant 2 smaller cells:1 large cell arrangement correspond to those where heat shock caused a cell cycle stall during the second cell cycle in one blastomere but not the other; and embryos &#34;stalled&#34; in a 1:1 arrangement correspond to those where during the second cell cycle both blastomeres underwent the desired cell division stall. Stalled embryos should resume cell cleavage during the following cell cycle period (65-80 mpf). The arrangement of blastomeres may be variable, due to the incomplete cues from the previous cell cycle to stabilize the spindle 2,14, but most embryos will form a relatively normal blastula that can undergo normal development.</li>
	<li>Allow embryos to develop in the Petri plate, with a limit of 80 embryos per 10 cm plate. At 24 hpf, observe the embryos to determine whether they have a normal morphology characteristic of diploid or homozygous diploid embryos 6 (normal extent of axis extension (Figure 3A, 3C)), in contrast to reduced axis extension and increased body thickness characteristic of haploids (Figure 3B)), and/or assess diploidization using genetic pigment markers at 36 mpf, such as golden or albino and other such assays (Figure 3 and see below) 2,6,9.</li>
	<li>Remove any embryos that appear to have a haploid morphology, or that have lysed or exhibit other grossly abnormal defects. 
	NOTE: If desired, allow embryos to develop until 5 days post fertilization, while continuing to remove lysed or grossly abnormal embryos on a daily basis and adding fresh E3 to refresh the medium. NOTE: Surviving embryos can also be grown after swimbladder inflation on day 5 by transferring to a hatchery system and feeding under standard conditions.</li>
</ol>

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The authors do not have competing financial interests.

Critical steps
It is critical to work under conditions of effective in vitro fertilization. To insure a good supply of mature eggs (step 1), females set up for mating should not have been set up in mating crosses for at least 5 days and should appear gravid. During interruption of breeding, an observer can monitor 15-30 tanks adequately for the first appearance of natural egg extrusion. Interruption of mating should occur as soon as possible when the first eggs are released through natural mating...

This protocol allows the manipulation of ploidy in zebrafish embryos, such as in the generation of homozygous gynogenetic diploids from gynogenetic haploids (Figure 1) or the production of tetraploids. This is achieved by inducing a delay in cytokinesis corresponding to precisely one cell cycle (Figure 2A, 2B). The key one-cycle delay in cytokinesis is achieved by treatment with heat shock. The standard protocol of Heat Shock (HS) as originally described by Streisinger and colleagues involved a temperature pulse during the period 13-15 mpf, resulting in a one-cycle cell division stall during the first cell cycle 1. The efficiency of this protocol has been recently improved by scanning the early cell cycles with a sliding temporal window of heat shock treatment. This scan identified a later time point for a heat shock, still within the first cell cycle (22-24 mpf), that results in a higher rate of embryos with a one-cycle cell division stall, which in this case affects the second cell cycle 2. The observation that experimental manipulations during the first cell cycle interfere with cell division during the second cell cycle and cause DNA content duplication has also been reported in other fish species 3,4. We refer to this modified protocol as Heat Shock 2 (HS2 &#8211; the term &#34;2&#34; reflects that the heat pulse occurs at a later time point than the standard HS method, and that the cell cycle delay caused by HS2 occurs during the second cell cycle). These studies showed that the basis for cytokinesis arrest after heat shock is the inhibition of centriole duplication during the heat pulse, which affects spindle formation and furrow induction in the following cell cycle. HS2 results in yields of cell cycle arrest nearing 100%, and rates of ploidy duplication up to 4 times higher than standard HS 2.
Embryos treated with a heat shock during blastomere cell cycling exhibit many deleterious effects, suggesting that heat shock affects multiple processes required for cell division 2. On the other hand, if the heat shock is applied prior to the initiation of cell cycling (time period 0-30 mpf), it appears to have effects consistent with specific interference with centriole duplication and does not seem to affect other essential cell processes 2. These studies showed that the time prior to the initiation of blastomere division appears to be a developmental period amenable to using Heat Shock as a tool to specifically manipulate ploidy through centriole inhibition. The underlying cause of the apparent selectivity for heat shock on centriole duplication is unknown, but may be related to a selective degradation of centrosome substructures observed under heat stress in certain cell types, such as leukocytes 5.
Temporal synchronization of embryonic development is achieved by in vitro fertilization (IVF). Use of untreated sperm during fertilization results in diploid embryos that upon HS2-induced one-cycle cytokinesis stall become tetraploid. Use of UV-treated sperm, which carries crosslinks that inactivate its DNA, results in gynogenetic haploid embryos 6, which upon HSII-induced one-cycle cytokinesis stall become gynogenetic diploids 2. Because of the resulting whole genome duplication, the latter gynogenetic diploids are homozygous at every single locus across the genome. For conciseness, we refer to gynogenetic haploid embryos as &#34;haploids&#34;, and homozygous gynogenetic diploid embryos as &#34;homozygous diploids&#34;. If viable and fertile, homozygous diploids can be used to initiate sterile and lethal-free lines. Direct homozygosity induced by HS2 should also be readily incorporated into genetic analysis or genetic screens, since homozygous diploids from females that are heterozygous carriers of mutations exhibit rates of homozygosity at high and fixed (50%) ratios 2.
The following protocol describes steps to perform HS2 and induce ploidy duplication with full homozygosity. For tetraploid production, sperm solution should be untreated. For homozygous diploid production, sperm should be inactivated by UV treatment. In addition, as described in the Discussion, visible pigment markers can also be used to facilitate identification of homozygous diploids. Zebrafish mate primarily during the first 3 hr of the initiation of their light cycle 7, and both adults and eggs are sensitive to circadian rhythms 8, so for best results the IVF procedure should occur within this time period.

<table><tbody><tr><td>Zebrafish mating boxes</td><td>Aqua Schwarz</td><td>SpawningBox1</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S5886</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P5405</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S3264</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>P9791</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>C7902</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;-7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>63138</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S5761</td><td></td></tr><tr><td>Tricaine</td><td>Western Chemical</td><td>Tricaine-D (MS 222)</td><td>FDA approved (ANADA 200-226)</td></tr><tr><td>Tris base</td><td>Sigma</td><td>77-86-1</td><td>to prepare 1 M Tris pH 9.0</td></tr><tr><td>HCl</td><td>Sigma</td><td>920-1</td><td>to prepare 1 M Tris pH 9.0</td></tr><tr><td>Fish net (fine mesh) (4-5 in)</td><td>PennPlax</td><td>(ThatFishThatPlace # 212370)</td><td>available in ThatFishThatPlace</td></tr><tr><td>Plastic spoon</td><td></td><td></td><td>available in most standard stores</td></tr><tr><td>Dissecting scissors</td><td>Fine Science Tools</td><td>14091-09</td><td></td></tr><tr><td>Dissecting forceps</td><td>Dumont</td><td>SS</td><td>available from Fine Science Tools</td></tr><tr><td>Dissecting stereoscope (with transmitted light source)</td><td>Nikon</td><td>SMZ645</td><td>or equivalent</td></tr><tr><td>Reflective light source (LED arms)</td><td>Fostec</td><td>KL1600 LED</td><td>or equivalent</td></tr><tr><td>Petri plates 10 cm diameter</td><td></td><td></td><td>any maker</td></tr><tr><td>Eppendorf tubes 1.5 ml</td><td></td><td></td><td>any maker</td></tr><tr><td>Ice bucket</td><td></td><td></td><td>any maker</td></tr><tr><td>Pipetteman P-1000</td><td></td><td></td><td>any maker</td></tr><tr><td>Pipette tips 1,000 &amp;micro;l</td><td></td><td></td><td>any maker</td></tr><tr><td>Narrow spatula</td><td>Fisher</td><td>14-374</td><td></td></tr><tr><td>Depression glass plate</td><td>Corning Inc</td><td>722085 (Fisher cat. No 13-748B)</td><td>available from Fisher Scientific</td></tr><tr><td>UV lamp</td><td>UVP</td><td>Model XX-15 (cat No. UVP18006201)</td><td>available from Fisher Scientific. Although not observed by us with this model, some UV sources have been observed to experience a decrease of intensity over time (if this is the case, see Modifications and Troubleshooting)</td></tr><tr><td>UV glasses</td><td></td><td></td><td>any maker</td></tr><tr><td>Paper towels</td><td></td><td></td><td>any maker</td></tr><tr><td>Kimwipes</td><td>Kimberly-Clark</td><td>06-666-11</td><td>available from Fisher Scientific</td></tr><tr><td>Timer stop watch</td><td></td><td></td><td>any maker</td></tr><tr><td>Wash bottle</td><td>Thermo Scientific</td><td>24020500</td><td>available from Fisher Scientific</td></tr><tr><td>Tea strainer</td><td></td><td></td><td>available in kitchen stores</td></tr><tr><td>beakers, 250 ml (2)</td><td>Corning Inc.</td><td>1000250</td><td>available from Fisher Scientific</td></tr><tr><td>water bath (2)</td><td></td><td></td><td>any maker, with accurary to 0.1 C (&lt;em&gt;e.g.&lt;/em&gt; Shel Lab H&lt;sub&gt;2&lt;/sub&gt;O Bath Series)</td></tr><tr><td>Hanks&amp;rsquo; Solution 1</td><td>see above</td><td>see above</td><td>8.0 g NaCl, 0.4 g KCl in 100 ml ddH&lt;sub&gt;2&lt;/sub&gt;O. Store at 4&amp;deg;C.</td></tr><tr><td>Hanks&amp;rsquo; Solution 2</td><td>see above</td><td>see above</td><td>0.358 g Anhydrous Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 0.6 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; in 100 ml ddH&lt;sub&gt;2&lt;/sub&gt;O. Store at 4 &amp;deg;C.</td></tr><tr><td>Hanks&amp;rsquo; Solution 4</td><td>see above</td><td>see above</td><td>0.72 g CaCl&lt;sub&gt;2&lt;/sub&gt; in 50 ml ddH&lt;sub&gt;2&lt;/sub&gt;O. Store at 4 &amp;deg;C.</td></tr><tr><td>Hank&#039;s Premix</td><td>see above</td><td>see above</td><td>add, in the following order: 10.0 ml Solution 1;&amp;nbsp; 1.0 ml Solution 2;&amp;nbsp; 1.0 ml Solution 4;&amp;nbsp; 86.0 ml ddH&lt;sub&gt;2&lt;/sub&gt;O;&amp;nbsp; 1.0 ml Solution 5. Store at 4 &amp;deg;C</td></tr><tr><td>Hanks&amp;rsquo; Solution 6</td><td>see above</td><td>see above</td><td>0.33 g NaHCO&lt;sub&gt;3&lt;/sub&gt; in 10 ml ddH&lt;sub&gt;2&lt;/sub&gt;O. Prepare fresh the morning of the IVF procedure.</td></tr><tr><td>Hank&#039;s Solution (final solution)</td><td>see above</td><td>see above</td><td>Combine 990 &amp;mu;l of Hank&amp;rsquo;s Premix and 10 &amp;mu;l of freshly made Solution 6 (NaHCO&lt;sub&gt;3&lt;/sub&gt; solution)</td></tr></tbody></table>

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.

In spite of the one-cell cycle cytokinesis stall, DNA replication occurs normally in such embryos, resulting in the duplication of the DNA content of the embryo (Figure 1). The Streisinger Heat Shock protocol (standard HS) involves a heat pulse during the period 13-15 minutes post fertilization (mpf) and induces primarily cytokinesis arrest during the first embryonic cell division at 35 mpf 1,2, whereas the derived method described here, referred to as Heat Sho...

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Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

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, ...

ploidy-manipulation-of-zebrafish-embryos-with-heat-shock-2-treatment

Research

JoVE Journal

Developmental Biology

10.0K Views.  University of Wisconsin.  Een gewijzigde protocol voor ploïdie manipulatie maakt gebruik van een heat shock naar een één-cyclus kraam in cytokinese induceren in de vroege embryo. Dit protocol wordt gedemonstreerd in de zebravis, maar kan voor andere soorten. 

Video: Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

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 ...

Genetics

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

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

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 ...

Rapid and Efficient Spatiotemporal Monitoring of Normal and Aberrant Cytosine Methylation within Intact Zebrafish Embryos

Cytosine methylation is highly conserved across vertebrate species and, as a key driver of epigenetic programming and chromatin state, plays a critical role in early embryonic development. Enzymatic modifications drive active methylation and demethylation of cytosine into 5-methylcytosine (5-mC) and subsequent oxidation of 5-mC into 5-hydroxymethylcytosine, 5-formylcytosine, and 5-carboxylcytosine. Epigenetic reprogramming is a critical period during in utero development, and maternal exposure to chemicals has the potential to reprogram the epigenome within offspring. This can potentially cause adverse outcomes such as immediate phenotypic consequences, long-term effects on adult disease susceptibility, and transgenerational effects of inherited epigenetic marks. Although bisulfite-based sequencing enables investigators to interrogate cytosine methylation at base-pair resolution, sequencing-based approaches are cost-prohibitive and, as such, preclude the ability to monitor cytosine methylation across developmental stages, multiple concentrations per chemical, and replicate embryos per treatment. Due to the ease of automated in vivo imaging, genetic manipulations, rapid ex utero development time, and husbandry during embryogenesis, zebrafish embryos continue to be used as a physiologically intact model for uncovering xenobiotic-mediated pathways that contribute to adverse outcomes during early embryonic development. Therefore, using commercially available 5-mC-specific antibodies, we describe a cost-effective strategy for rapid and efficient spatiotemporal monitoring of cytosine methylation within individual, intact zebrafish embryos by leveraging whole-mount immunohistochemistry, automated high-content imaging, and efficient data processing using programming language prior to statistical analysis. To current knowledge, this method is the first to successfully detect and quantify 5-mC levels in situ within zebrafish embryos during early development. The method enables the detection of DNA methylation within the cell mass and also has the ability to detect cytosine methylation of yolk-localized maternal mRNAs during the maternal-to-zygotic transition. Overall, this method will be useful for the rapid identification of chemicals that have the potential to disrupt cytosine methylation in situ during epigenetic reprogramming.

This paper describes a protocol for the rapid and efficient spatiotemporal monitoring of normal and aberrant cytosine methylation within intact zebrafish embryos.

Cytosine methylation is highly conserved across vertebrate species and, as a key driver of epigenetic programming and chromatin state, plays a critical ...

The Zebrafish Tol2 System: A Modular and Flexible Gateway-Based Transgenesis Approach

Fetal alcohol spectrum disorders (FASD) are characterized by a highly variable set of structural defects and cognitive impairments that arise due to prenatal ethanol exposure. Due to the complex pathology of FASD, animal models have proven critical to our current understanding of ethanol-induced developmental defects. Zebrafish have proven to be a powerful model to examine ethanol-induced developmental defects due to the high degree of conservation of both genetics and development between zebrafish and humans. As a model system, zebrafish possess many attributes that make them ideal for developmental studies, including large numbers of externally fertilized embryos that are genetically tractable and translucent. This allows researchers to precisely control the timing and dosage of ethanol exposure in multiple genetic contexts. One important genetic tool available in zebrafish is transgenesis. However, generating transgenic constructs and establishing transgenic lines can be complex and difficult. To address this issue, zebrafish researchers have established the transposon-based Tol2 transgenesis system. This modular system uses a multisite Gateway cloning approach for the quick assembly of complete Tol2 transposon-based transgenic constructs. Here, we describe the flexible Tol2 system toolbox and a protocol for generating transgenic constructs ready for zebrafish transgenesis and their use in ethanol studies.

This work describes a protocol for the modular Tol2 transgenesis system, a gateway-based cloning method to create and inject transgenic constructs into zebrafish embryos.

Fetal alcohol spectrum disorders (FASD) are characterized by a highly variable set of structural defects and cognitive impairments that arise due to ...

Triggering Cell Stress and Death Using Conventional UV Laser Confocal Microscopy

Using a standard confocal setup, a UV ablation method can be utilized to selectively induce cellular injury and to visualize single-cell responses and cell-cell interactions in the CNS in real-time. Previously, studying these cell-specific responses after injury often required complicated setups or the transfer of cells or animals into different, non-physiological environments, confounding immediate and short-term analysis. For example, drug-mediated ablation approaches often lack the specificity that is required to study single-cell responses and immediate cell-cell interactions. Similarly, while high-power pulsed laser ablation approaches provide very good control and tissue penetration, they require specialized equipment that can complicate real-time visualization of cellular responses. The refined UV laser ablation approach described here allows researchers to stress or kill an individual cell in a dose- and time-dependent manner using a conventional confocal microscope equipped with a 405-nm laser. The method was applied to selectively ablate a single neuron within a dense network of surrounding cells in the zebrafish spinal cord. This approach revealed a dose-dependent response of the ablated neurons, causing the fragmentation of cellular bodies and anterograde degeneration along the axon within minutes to hours. This method allows researchers to study the fate of an individual dying cell and, importantly, the instant response of cells-such as microglia and astrocytes-surrounding the ablation site.

Targeted manipulations to cause directed stress or death in individual cells have been relatively difficult to accomplish. Here, a single-cell-resolution ablation approach to selectively stress and kill individual cells in cell culture and living animals is described based on a standard confocal UV laser.

Using a standard confocal setup, a UV ablation method can be utilized to selectively induce cellular injury and to visualize single-cell responses and ...

Neuroscience

Analysis of Oxidative Stress in Zebrafish Embryos

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes oxidation of molecules (lipid, DNA, protein) and leads to cell death. Oxidative stress also impacts the progression of several pathological conditions such as diabetes, retinopathies, neurodegeneration, and cancer. Thus, it is important to define tools to investigate oxidative stress conditions not only at the level of single cells but also in the context of whole organisms. Here, we consider the zebrafish embryo as a useful in vivo system to perform such studies and present a protocol to measure in vivo oxidative stress. Taking advantage of fluorescent ROS probes and zebrafish transgenic fluorescent lines, we develop two different methods to measure oxidative stress in vivo: i) a &#8220;whole embryo ROS-detection method&#8221; for qualitative measurement of oxidative stress and ii) a &#8220;single-cell&#160;ROS detection method&#8221; for quantitative measurements of oxidative stress. Herein, we demonstrate the efficacy of these procedures by increasing oxidative stress in tissues by oxidant agents and physiological or genetic methods. This protocol is amenable for forward genetic screens and it will help address cause-effect relationships of ROS in animal models of oxidative stress-related pathologies such as neurological disorders and cancer.

Here we report a protocol to measure oxidative stress in living zebrafish embryos. This procedure allows reactive oxygen species (ROS) detection in both whole embryo tissues and single-cell populations. This protocol will accomplish both qualitative and quantitative analyses.

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes ...

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for forward genetic analysis of developmental processes due to their external fertilization, embryonic size, rapid ontogeny, and optical clarity &#8211; a constellation of traits that enable the direct observation of events ranging from gastrulation to organogenesis with a basic stereomicroscope. Further, zebrafish embryos can survive for several days in the haploid state. The production of haploid embryos in vitro is a powerful tool for mutational analysis, as it enables the identification of recessive mutant alleles present in first generation (F1) female carriers following mutagenesis in the parental (P) generation. This approach eliminates the necessity to raise multiple generations (F2, F3, etc.) which involves breeding of mutant families, thus saving the researcher time along with reducing the needs for zebrafish colony space, labor, and the husbandry costs. Although zebrafish have been used to conduct forward screens for the past several decades, there has been a steady expansion of transgenic and genome editing tools. These tools now offer a plethora of ways to create nuanced assays for next generation screens that can be used to further dissect the gene regulatory networks that drive vertebrate ontogeny. Here, we describe how to prepare haploid zebrafish embryos. This protocol can be implemented for novel future haploid screens, such as in enhancer and suppressor screens, to address the mechanisms of development for a broad number of processes and tissues that form during early embryonic stages.

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish is a powerful model system for developmental biology and human disease research due to their genetic similarity with higher vertebrates. This protocol describes a methodology to create haploid zebrafish embryos that can be utilized for forward screen strategies to identify recessive mutations in genes essential for early embryogenesis.

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for ...

Efficient PAM-Less Base Editing for Zebrafish Modeling of Human Genetic Disease with zSpRY-ABE8e

CRISPR/Cas9 technology has increased the value of zebrafish for modeling human genetic diseases, studying disease pathogenesis, and drug screening, but protospacer adjacent motif (PAM) limitations are a major obstacle to creating accurate animal models of human genetic disorders caused by single-nucleotide variants (SNVs). Until now, some SpCas9 variants with broad PAM compatibility have shown efficiency in zebrafish. The application of the optimized SpRY-mediated adenine base editor (ABE), zSpRY-ABE8e, and synthetically modified gRNA in zebrafish has enabled efficient adenine-guanine base conversion without PAM restriction. Described here is a protocol for efficient adenine base editing without PAM limitation in zebrafish using zSpRY-ABE8e. By injecting a mixture of zSpRY-ABE8e mRNA and synthetically modified gRNA into zebrafish embryos, a zebrafish disease model was constructed with a precise mutation that simulated a pathogenic site of the TSR2 ribosome maturation factor (tsr2). This method provides a valuable tool for the establishment of accurate disease models for studying disease mechanisms and treatments.

This protocol describes how to perform efficient adenine base editing without PAM limitation to construct a precise zebrafish disease model using zSpRY-ABE8e.

CRISPR/Cas9 technology has increased the value of zebrafish for modeling human genetic diseases, studying disease pathogenesis, and drug screening, but ...

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

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