We would like to thank Anna Hindes at Washington University School of Medicine for her initial efforts in obtaining good-quality sperm genomic DNA using the hot shot method. This work was funded by the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institutes of Health under Award (R01AR072009 to R.S.G.).

This study was carried out in line with the guidelines in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the University of Texas at Austin Animal Care and Use Committee (AUP-2021-00254).
1. Designing the sgRNA for CRISPR mutagenesis
<ol>
	<li>Obtain the exon sequence containing the targeted loci.</li>
	<li>Design a synthetic guide RNA (sgRNA) with a protospacer adjacent motif (PAM) site that is specific to the Cas endonuclease.
	<ol>
		<li>If co-injecting a donor construct, design the sgRNA to have a predicted cut site as close to the desired edits as possible. 
		NOTE: APE is a free software with a tool to find Cas9 sgRNA sites11. Alternatively, tools such as CHOPCHOP can be used, which offers an online interface for designing sgRNAs that considers the different PAM sites of Cas9 and other Cas endonucleases12.</li>
	</ol>
	</li>
	<li>Design forward and reverse primers to amplify the region surrounding the designed CRISPR/Cas cut site. The PCR product should be approximately 200-400 base pairs (bp) in length to be able to resolve small indels on a gel later in the protocol. 
	NOTE: CHOPCHOP11 automatically designs primers for every sgRNA it outputs.</li>
	<li>Optional: Design a DNA donor construct with the desired edit(s).
	<ol>
		<li>Obtain the genomic sequence of the exon containing the loci of the desired mutation. 
		NOTE: Intronic sequences can vary between individuals, so it is best to avoid the inclusion of intronic sequences in the donor construct as this could interfere with the homology-dependent repair pathway.</li>
		<li>If a specific amino acid change is desired, alter the specific codon accordingly. 
		NOTE: When making these changes, be aware of the codon frequency. If possible, choose a codon with high usage.</li>
		<li>To prevent CRISPR/Cas digestion of the donor construct, make additional synonymous mutations that (i) disrupt the PAM and/or (ii) disrupt the sgRNA sequence, with higher preference being placed on making non-synonymous changes closer to the PAM site.</li>
		<li>Use a restriction site finder (APE11 also has this feature) to find unique restriction sites (those that only occur once) in the predicted PCR product of the wild-type sequence.
		<ol>
			<li>Repeat this process for the predicted PCR product that reflects the successful integration of the donor construct.</li>
			<li>Compare the list of restriction sites between the wild-type and donor-integrated sequences to find a restriction site that is present only in the edited sequence.</li>
			<li>If no unique restriction site is found, continue making synonymous changes within the donor construct until this is achieved. 
			NOTE: If possible, continue making synonymous mutations that would disrupt the sgRNA binding to the donor construct.</li>
		</ol>
		</li>
		<li>Trim the donor sequence to a manageable size (50-100 bp) with at least 20 bp of homology surrounding the predicted CRISPR/Cas cut site and edits, if possible.</li>
	</ol>
	</li>
	<li>Obtain the designed sgRNA and/or DNA donor oligonucleotides, as well as any primers required for genotyping. 
	NOTE: To prevent the digestion of the DNA donor oligonucleotides by nucleases, it is recommended to modify the first three phosphate bonds on the 5&#39; and 3&#39; ends of the DNA oligonucleotides with phosphorothioate.</li>
	<li>Inject 1 nL of the injection mix into one-cell stage embryos. Typically, a standard injection mix contains the following: 
	1&#160;&#181;L of 25 &#181;M locus-specific sgRNA 
	1&#160;&#181;L of 25 &#181;M Cas9 endonuclease 
	&#8203;Optional: 1 &#181;L of 3 &#181;M DNA donor construct
	<ol>
		<li>Bring the total volume to 5 &#181;L with 0.1 M RNase-free KCl.</li>
	</ol>
	</li>
	<li>Allow the F0-injected individuals to develop into sexually mature adults. The remaining protocol focuses on screening the F0-injected males for germline edits.</li>
</ol>
2. Setting up the breeding tanks
<ol>
	<li>Prime F0 males for mating by setting up breeding tanks with dividers between the F0 males and females.
	<ol>
		<li>To minimize the number of breeding tanks required, set up three males across from two females in each tank.
		<ol>
			<li>Set up as many F0 males as can effectively be screened and housed as individuals after sperm collection. The genotype of the females is not important since their gametes will not be collected during this procedure.</li>
		</ol>
		</li>
		<li>Set up one breeding tank of wild-type males across from females. Wild-type sperm will serve as a control for the interpretation of the results later in the procedure.</li>
	</ol>
	</li>
	<li>Incubate the prepared breeding tanks overnight.</li>
</ol>
3. Preparing the materials for the sperm collection procedure
<ol>
	<li>Prepare enough anesthetizing solution (0.016% tricaine-HCl in fish water) to fill a small glass bowl.</li>
	<li>Dispense 40 &#181;L of 50 mM NaOH into 0.2 mL PCR tubes. Prepare one tube for each sperm sample that will be collected. Store on ice.</li>
	<li>Prepare individual 0.8 L tanks for each fish that will be sampled during the procedure. These tanks will house the sampled fish until the genotyping is complete. 
	NOTE: This is the primary factor that limits the number of fish that can be effectively screened in this protocol. Ensure adequate space for housing the sampled individuals until the genotyping is complete.</li>
	<li>Moisten a 1-inch x 1-inch&#160;sponge (cut with a shallow, oval-shaped divot to hold a male zebrafish ventral side up during the procedure) in fish water&#160;and squeeze out excess liquid.
	<ol>
		<li>Position the sponge in the field of view of a stereomicroscope at low magnification (Figure 1A).</li>
		<li>Position oblique overhead lighting for optimal viewing.</li>
	</ol>
	</li>
</ol>
4. Anesthetizing the male fish and collecting the sperm
<ol>
	<li>Prepare a clean capillary tube.
	<ol>
		<li>Gently shake the tube container to release a single capillary tube.</li>
		<li>Place the capillary tube into the provided bulb, and set it aside.</li>
	</ol>
	</li>
	<li>Transfer a male fish into the prepared anesthetizing solution (0.016% tricaine-HCl in fish water).
	<ol>
		<li>Gently stir the solution with a slotted spoon to speed up the process of anesthesia.</li>
		<li>Once the operculum (gill) movement slows down (approximately 1-2 min), proceed with the sperm collection.</li>
	</ol>
	</li>
	<li>Use the slotted spoon to transfer the fish onto a clean stack of paper towels.
	<ol>
		<li>Use the slotted spoon to gently roll the fish to remove excess water.</li>
		<li>Gently blot the fish with clean, folded tissue paper to remove any remaining water.</li>
	</ol>
	</li>
	<li>Use the slotted spoon to transfer the fish to the prepared sponge (Figure 1B).
	<ol>
		<li>Position the fish ventral side up with its head nearest the dominant hand of the person performing the procedure.</li>
		<li>Gently blot the area around the anal fins with clean, folded tissue paper.</li>
	</ol>
	</li>
	<li>Collect the sperm.
	<ol>
		<li>Using the end of the capillary tube, gently move the pelvic fins laterally away from the midline to expose the cloaca (Figure 1C).</li>
		<li>Position the end of the capillary tube near the cloaca.</li>
		<li>Using the forceps, gently squeeze the sides of the fish beginning just below the gills and ending at the cloaca (Figure 1C&#39;).</li>
		<li>Collect sperm into the capillary tube by capillary action. Approximately 1 &#181;L is sufficient for this procedure (Figure 1D). 
		NOTE: Some labs report using p10 pipet tips instead of capillary tubes for sperm collection.</li>
		<li>If sperm is not released with gentle pressure, return the fish to the system water.
		<ol>
			<li>Monitor for recovery from the anesthesia.</li>
			<li>Do not squeeze the fish hard to expel sperm, as this can injure the fish.</li>
			<li>Rest the fish for 2 weeks before attempting to collect sperm again.</li>
		</ol>
		</li>
		<li>After collection, place the fish into a clean isolation tank with system water for individual housing.
		<ol>
			<li>Monitor the fish for recovery from the anesthesia.</li>
			<li>Rest the fish for 2 weeks before attempting to collect sperm again.</li>
		</ol>
		</li>
		<li>Place the capillary tube with the collected sperm into the prepared PCR tube (with 40 &#181;L of 50 mM NaOH).</li>
		<li>Squeeze the rubber bulb to expel the collected sample.</li>
		<li>Dispose of the used capillary tube into an approved glass waste container.</li>
		<li>Incubate the samples on ice until all the males have been squeezed.</li>
	</ol>
	</li>
	<li>Repeat steps 4.1-4.5 for each individual male.</li>
	<li>Label the isolation tanks and sperm samples such that the final sperm genotyping results can be traced back to the corresponding individual with putative mutations.</li>
	<li>Monitor all the individuals, and make sure they are upright and exploring their tanks before putting the tanks back on the system.</li>
	<li>Put the females used for priming back into their respective tanks on the system.</li>
	<li>If any fish does not recover from the anesthesia after 10 min, follow the euthanasia procedure approved by the institution associated with the lab.
	<ol>
		<li>Example: Rapid chill the individual in an ice bath with fish water that is 2-4 &#176;C.</li>
		<li>Dispose of the euthanized fish in an appropriate animal carcass waste container according to the procedure approved by the institution associated with the lab.</li>
	</ol>
	</li>
</ol>
5. Extracting DNA from the sperm samples
<ol>
	<li>Briefly spin down all the sperm samples in a minicentrifuge, and place PCR tubes into a thermal cycler.</li>
	<li>Close the lid of the thermal cycler.</li>
	<li>Run the following settings in the thermal cycler:
	<ol>
		<li>Heat the samples for 40 min at 95 &#176;C.</li>
		<li>Cool the samples to 25 &#176;C.</li>
	</ol>
	</li>
	<li>Remove the samples from the thermal cycler.</li>
	<li>With a clean pipette tip for each sample, neutralize the pH by adding 10 &#181;L of 1 M Tris-HCl (buffered to pH 8).</li>
	<li>Mix well by pipetting up and down.</li>
	<li>Briefly spin down the samples in a minicentrifuge.</li>
	<li>Genomic DNA samples can be stored for several days at 4 &#176;C or placed at &#8722;20 &#176;C for up to 6 months.</li>
</ol>
6. PCR amplification (and/or restriction digest) of the desired locus
<ol>
	<li>Amplify the desired locus using a standard PCR protocol.
	<ol>
		<li>Preheat the thermal cycler to the initial denaturation temperature in the PCR reaction below.</li>
		<li>Prepare a 25 &#181;L reaction mixture for each sperm sample in PCR tubes: 
		12.5&#160;&#181;L of 2x Taq Polymerase Master Mix 
		1.5&#160;&#181;L of 10 &#181;M forward primer 
		1.5&#160;&#181;L of 10 &#181;M reverse primer 
		4.5&#160;&#181;L of nuclease-free water 
		5&#160;&#181;L of neutralized sperm DNA sample</li>
		<li>Mix well by pipetting up and down.</li>
		<li>Briefly spin down the samples in a minicentrifuge.</li>
		<li>Place the samples in the pre-heated thermal cycler with the following settings: 
		Initial denaturation: 95 &#176;C for 3 min 
		35&#160;cycles of denaturation (95 &#176;C for 30 s), annealing (55 &#176;C for 30 s), and extension (72 &#176;C for 30 s) 
		Final extension: 72 &#176;C for 5 min 
		NOTE: It may be necessary to adjust the extension temperature and/or time depending on the specific Taq polymerase and the expected length of the PCR amplification product.</li>
		<li>Remove the PCR samples from the thermal cycler.</li>
	</ol>
	</li>
	<li>Optional: Perform the donor-specific restriction digest on a small volume of the PCR product, leaving at least 5-10 &#181;L of undigested PCR product for gel electrophoresis.
	<ol>
		<li>Prepare a 30 &#181;L reaction in 0.2 mL PCR tubes: 
		10&#160;&#181;L of unpurified PCR product 
		15&#160;&#181;L of nuclease-free water 
		3&#160;&#181;L of 10x restriction enzyme buffer 
		2&#160;&#181;L of restriction enzyme (BstNI is used in the representative results for analyzing the dnah10 knock-in allele) 
		&#8203;NOTE: Many restriction enzymes are still active in a typical PCR reaction buffer; however, the addition of restriction digest buffer can help improve the results. Refer to specific manufacturer instructions for troubleshooting if any problems occur.</li>
		<li>Mix well by pipetting up and down.</li>
		<li>Briefly spin down the samples in a minicentrifuge.</li>
		<li>Place the samples in a thermal cycler with the following settings:
		<ol>
			<li>Cut the PCR amplicon at 60 &#176;C for 1 h. 
			NOTE: The cut temperature and incubation time may need to be adjusted for the specific restriction enzyme being used.</li>
			<li>Inactivate the restriction enzyme with the required temperature and incubation time, if indicated by the manufacturer. 
			&#8203;NOTE: The BstNI enzyme does not require heat inactivation.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
7. Performing gel electrophoresis to separate PCR amplicons of varying sizes
<ol>
	<li>Prepare 500 mL of 0.5x TBE running buffer: add 50 mL of 10x TBE buffer concentrate to 450 mL of deionized water.</li>
	<li>Prepare 100 mL of 4% agarose gel in 0.5x TBE running buffer: add 4 g of agarose with 10 &#181;L of 10,000x gel stain to 100 mL of 0.5x TBE running buffer. Microwave until the agarose powder is fully dissolved.
	<ol>
		<li>Pour the gel solution into an appropriately sized gel casting frame, and insert the comb on one side of the gel. 
		NOTE: A larger gel size (recommended: 15 cm x 15 cm) is advantageous for this procedure, as it provides the amplicons with more space to resolve on the gel.
		<ol>
			<li>If a restriction digest product is to be analyzed, it is best to run it on the same gel as the PCR product so that the results can be compared. Adjust the comb well size as necessary to fit all the samples on the gel.</li>
		</ol>
		</li>
		<li>Leave the gel at room temperature until solidified. Carefully remove the comb. Carefully remove the gel from the casting frame.</li>
	</ol>
	</li>
	<li>Set up the gel electrophoresis rig.
	<ol>
		<li>Position the gel in the electrophoresis rig such that the wells are nearest the negative electrode. Pour 0.5x TBE running buffer into the electrophoresis rig until the wells are fully submerged.</li>
	</ol>
	</li>
	<li>Load the gel.
	<ol>
		<li>Load 5 &#181;L of DNA ladder into the first well (choose a DNA ladder that contains DNA fragments that are similar in size to the PCR amplicon).</li>
		<li>Using a new pipette tip for each sample (including the wild-type control sample), load 5-10 &#181;L of the PCR product into the remaining wells.</li>
		<li>Optional: If a restriction digest product is to be analyzed, load these samples onto the gel as well. 
		NOTE: It is best to load equal amounts of DNA between the PCR and the restriction digest product. For example, in the 30 &#181;L restriction digest reaction mixture, 10 &#181;L of PCR product was added. Therefore, the DNA concentration of the 15 &#181;L of digest product was comparable to the 5 &#181;L of unpurified PCR product.</li>
	</ol>
	</li>
	<li>Run the gel.
	<ol>
		<li>Connect the electrodes to the gel electrophoresis rig, and apply 150 V for at least 1 h to ensure adequate separation of the amplicons.</li>
		<li>Image the gel to view the DNA bands. 
		&#8203;NOTE: Strongly mutagenized CRISPR-targeted loci will run as multiple band sizes representing a combination of insertion/deletion alleles with the wild-type amplicons.
		<ol>
			<li>If the gel bands are not resolved after 1 h, apply 150 V to the gel for 15 min intervals until the bands are sufficiently separated on the gel.</li>
			<li>Optional: If screening for the insertion of a precise edit such as an SNP, epitope tag, or fluorescent tag, screen the sperm DNA with PCR methods that are specific to the desired edit. For example, when identifying a donor-specific restriction site, an additional band will be present in the restriction digest product compared to the PCR product of the sperm sample.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
8. Isolating germline stable alleles
<ol>
	<li>After 2 weeks of rest from the sperm squeezing procedure, outcross the F0 individuals whose sperm genomic DNA contained the desired edits. 
	NOTE: Sperm generation from individual spermatogonia clones may be cyclical in zebrafish; thus, the desired allele might not be producing sperm at the time of the outcross. For this reason, additional crossing may have to be performed more than once to isolate the allele implicated by the initial screening.</li>
	<li>Once the F1 outcrossed fish are sexually mature, proceed with standard genotyping methods (for example, fin-clip DNA) using the same PCR and gel electrophoresis protocol performed during the screening of the sperm genomic DNA.
	<ol>
		<li>Sanger sequence the PCR products to identify the F1 individuals with the same desired edit.</li>
		<li>Optional: If the desired edit introduces or disrupts a restriction site, screen with an appropriate restriction digest protocol.</li>
	</ol>
	</li>
	<li>In-cross F1 heterozygotes with the same desired edit.</li>
	<li>Screen the F2 progeny for the expected phenotype and/or genomic edits.</li>
</ol>

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</ol>

This protocol describes a method for rapidly characterizing putative genome edits or targeted mutations using CRISPR-Cas technology by focused analysis on F0 male sperm genomes. This protocol should be amenable to other animal models where sperm is readily available for sampling without euthanasia. This method will increase the throughput of screening for desired edits and is especially useful for identifying rare HDR-mediated knock-in events. This approach also serves to reduce the number of experimental animals used to...

The CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas system is a powerful tool to perform loci-specific mutagenesis and precise genome editing in the Danio rerio (zebrafish) model system1,2,3,4. The Cas-ribonucleoprotein (RNP) is comprised of two main components: a Cas endonuclease (commonly Cas9 or Cas12a) and a locus-specific synthetic guide RNA (sgRNA)5. Together, the Cas-RNP generates a double-stranded break (DSB) in the desired locus that can be repaired by one of two intrinsic repair mechanisms. The non-homologous end joining (NHEJ) repair mechanism is error-prone and often results in a variety of insertions or deletions (indels) around the DSB. These indels can be deleterious if they introduce a frameshift mutation or premature stop in the resultant protein sequence. Alternatively, the homology-directed repair (HDR) mechanism uses a donor template with regions of homology surrounding the DSB site to repair the damage. Researchers can take advantage of the HDR system to generate precise genomic edits. Specifically, they can co-inject a double-stranded DNA donor construct that contains the desired edits as well as regions of homology flanking the DSB site in the genome. The increased economy of scale for these commercially produced CRISPR components has greatly reduced the barriers to screening multiple loci and to setting up larger-scale efforts for precise genome editing. However, in sexually reproducing animal models, a major bottleneck is the identification and isolation of germline-stable mutant animals.
The zebrafish model system exhibits several key qualities that enhance its use in reverse genetic studies. They are easy to rear in large numbers with basic aquatic housing equipment, and females exhibit high fecundity all year round6. Moreover, their external egg-laying and fertilization make them amenable to the microinjection of CRISPR/Cas components. The Cas-RNP is commonly injected into one-cell stage zebrafish embryos to generate DSBs/repair that is, in theory, inherited by all the daughter cells. However, diploid genomes require two DSB/repair events to mutagenize both homologous chromosomes. Furthermore, although Cas-RNP is injected at the one-cell stage, the DSB/repair may not occur until later points in development. Together, these factors contribute to the mosaic nature of F0-injected fish. A common practice is to outcross F0-injected fish and screen the F1 progeny for indels/specific edits. However, since not all F0-injected fish possess germline mutations, this practice results in many unproductive crosses that do not generate the desired edit. Screening the F0 germline rather than F1 somatic tissue increases the probability of isolating the desired germline edit and reduces the number of animals required in this process.
Sperm can easily be collected from F0-injected zebrafish without the need for euthanasia. This feature allows for the cryopreservation and rederivation of frozen sperm stocks7 but can also be exploited to rapidly screen, identify, and isolate the germline carriers of desired genomic mutations8,9. Brocal et al. (2016) previously described a sequencing-based method for screening germline edits in F0-injected male zebrafish10. Although useful for identifying the mutated alleles present in the germline, this approach can become costly in high throughput and may not be accessible for all labs. In contrast, the current protocol offers an approachable and cost-effective electrophoresis-based strategy for identifying germline edits. Specifically, this protocol outlines a robust method for screening and isolating germline mutations at specific loci using high-resolution agarose gel electrophoresis. In addition, this protocol describes a similar strategy for identifying the successful integration of a donor construct containing specific edits. As always, if specific edits are desired, sequencing-based strategies can be performed in tandem with the protocol described below. Although this protocol is specific to the zebrafish model system, these principles should be adaptable to any model in which the collection of sperm is a routine procedure. Together, these strategies will allow for the identification of F0-injected males with germline indels/edits&#160;that can be resolved on a gel after standard polymerase chain reaction (PCR) and/or restriction digest.

<table><tbody><tr><td>Agarose powder&amp;nbsp;</td><td>Fisher BioReagents</td><td>BP1356-100</td><td></td></tr><tr><td>Breeding tanks</td><td>Carolina Biological&amp;nbsp;</td><td>161937</td><td></td></tr><tr><td>BstNI Restriction Enzyme</td><td>NEB</td><td>R0168S</td><td></td></tr><tr><td>Cas9 Endonuclease</td><td>IDT</td><td>1081060</td><td></td></tr><tr><td>DNA Ladder, 100 bp&amp;nbsp;</td><td>Thermo Scientific&amp;nbsp;</td><td>FERSM0241</td><td></td></tr><tr><td>dnah10 donor construct&amp;nbsp;&amp;nbsp;</td><td>Sigma-Aldrich</td><td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry.&lt;br/&gt; Phosphorothioate bond on the donor at the first three phosphate bonds on both the 5&amp;rsquo; and 3&amp;rsquo; ends (5&#039;-CCTCTCTCCCTTTCAGAAGCTTC&lt;br/&gt; TGCTCATCCGCTGCTTCTGCCT&lt;br/&gt; GGACCGAGTGTACCGTGCCGTC&lt;br/&gt; AGTGATTACGTCACGC-3&#039;)</td><td></td></tr><tr><td>dnah10 forward primer&amp;nbsp;</td><td>Sigma-Aldrich</td><td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5&#039;-CATGGAACTCTTTCCTAATGAGT&lt;br/&gt; TTGGC-3&#039;)</td><td></td></tr><tr><td>dnah10 reverse primer&amp;nbsp;</td><td>Sigma-Aldrich</td><td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (&#039;5-AGTAGAGATCACACATCAACAGA&lt;br/&gt; ATACAGC-3&#039;)&amp;nbsp;</td><td></td></tr><tr><td>dnah10 synthetic sgRNA&amp;nbsp;</td><td>Synthego&amp;nbsp;</td><td>Synthetic sgRNA, target sequence: 5&#039;-GCTCATCCGCTGCTTCAGGC-3&#039;</td><td></td></tr><tr><td>Electrophoresis power supply</td><td>Thermo Scientific&amp;nbsp;</td><td>105ECA-115</td><td></td></tr><tr><td>Filter forceps</td><td>Millipore</td><td>XX6200006P</td><td></td></tr><tr><td>Fish (system) water</td><td>Generic</td><td>n/a</td><td></td></tr><tr><td>Gel electrophoresis system (including casting frame, comb, and electrophoresis chamber)&amp;nbsp;</td><td>Thermo Scientific&amp;nbsp;</td><td>B2</td><td></td></tr><tr><td>Gel imaging light box&amp;nbsp;</td><td>Azure Biosystems</td><td>AZI200-01</td><td></td></tr><tr><td>Gel stain, 10000X&amp;nbsp;</td><td>Invitrogen</td><td>S33102</td><td></td></tr><tr><td>Glass bowl, 250 mL&amp;nbsp;</td><td>Generic</td><td>n/a</td><td></td></tr><tr><td>Isolation tanks, 0.8 L&amp;nbsp;</td><td>Aquaneering</td><td>ZT080</td><td></td></tr><tr><td>Microcap capillary tube with bulb, 20 &amp;micro;L&amp;nbsp;</td><td>Drummond</td><td>1-000-0020/CA</td><td></td></tr><tr><td>Minicentrifuge</td><td>Bio-Rad</td><td>12011919EDU</td><td></td></tr><tr><td>Micropipettes, various with appropriate tips</td><td>Generic&amp;nbsp;</td><td>n/a</td><td></td></tr><tr><td>Microwave&amp;nbsp;</td><td>Generic&amp;nbsp;</td><td>n/a</td><td></td></tr><tr><td>Nuclease free water</td><td>Promega</td><td>P119-C</td><td></td></tr><tr><td>Paper towels</td><td>Generic</td><td>n/a</td><td></td></tr><tr><td>PCR tubes, 0.2 mL</td><td>Bioexpress</td><td>T-3196-1</td><td></td></tr><tr><td>Plastic spoon, with drilled holes/slots&amp;nbsp;</td><td>Generic</td><td>n/a</td><td></td></tr><tr><td>KCl solution, 0.2 M RNAse Free</td><td>Sigma-Aldrich</td><td>P9333</td><td></td></tr><tr><td>p2ry12 forward primer&amp;nbsp;</td><td>Sigma-Aldrich</td><td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5&#039;-CCCAAATGTAATCCTGACCAGT&lt;br/&gt; -3&#039;)</td><td></td></tr><tr><td>p2ry12 reverse primer</td><td>Sigma-Aldrich</td><td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5&#039;-CCAGGAACACATTAACCTGGAT&lt;br/&gt; -3&#039;)&amp;nbsp;</td><td></td></tr><tr><td>p2ry12 synthetic sgRNA&amp;nbsp;</td><td>Synthego&amp;nbsp;</td><td>Synthetic sgRNA, target sequence: 5&#039;-GGCCGCACGAGGTCTCCGCG-3&#039;</td><td></td></tr><tr><td>Restriction Enzyme 10X Buffer</td><td>NEB</td><td>B6003SVIAL</td><td></td></tr><tr><td>NaOH solution, 50 mM&amp;nbsp;</td><td>Thermo Scientific&amp;nbsp;</td><td>S318; 424330010</td><td></td></tr><tr><td>Sponge, 1-inch x 1-inch cut with small oval divot</td><td>Generic&amp;nbsp;</td><td>n/a</td><td></td></tr><tr><td>Stereomicroscope</td><td>Zeiss</td><td>Stemi 508</td><td></td></tr><tr><td>Taq polymerase master mix, 2X</td><td>Promega</td><td>M7122</td><td></td></tr><tr><td>TBE Buffer Concentrate, 10X</td><td>VWR</td><td>E442</td><td></td></tr><tr><td>Thermal Cycler</td><td>Bio-Rad</td><td>1861096</td><td></td></tr><tr><td>Tissue paper</td><td>Fisher Scientific</td><td>06-666</td><td></td></tr><tr><td>Tricaine-methanesulfonate solution (Syncaine, MS-222), 0.016% in fish water (pH 7.0&amp;plusmn;0.2)&amp;nbsp;</td><td>Syndel</td><td>200-266</td><td></td></tr><tr><td>Tris Base, 1M (Buffered with HCl to ph 8.0)&amp;nbsp;</td><td>Promega</td><td>H5131</td><td></td></tr></tbody></table>

screening sperm for the rapid isolation of germline edits in zebrafish

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

The experimental approaches described in this protocol allow for the more rapid identification of genome edits or putative deleterious alleles by focusing on the analysis of thousands of genomes derived from the collection of F0-injected male sperm. Figure 2 highlights how to interpret the results obtained using this protocol.
To generate mutations in the p2ry12 locus, one-cell stage zebrafish embryos were injected with Cas9 endonuclease and a p2ry12<...

Watch this Scientific Journal Video about Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish at JoVE.com

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

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

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

screening-sperm-for-the-rapid-isolation-of-germline-edits-in-zebrafish

Research

JoVE Journal

Genetics

1.2K Views.  The University of Texas at Austin. CRISPR-Cas technologies have revolutionized the field of genome editing. However, finding and isolating the desired germline edit remains a major bottleneck. Therefore, this protocol describes a robust method for quickly screening F0 CRISPR-injected zebrafish sperm for germline edits using standard PCR, restriction digest, and gel electrophoresis techniques.

Video: Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Developmental Biology

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 ...

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be difficult to study, and can lead to confounding results. The protocol described here is a straightforward breeding paradigm to increase and decrease penetrance in lethal zebrafish skeletal mutants. Because lethal mutants cannot be selectively bred directly, the classic selective breeding strategy of progeny testing is employed. This method also includes protocols for Kompetitive Allele Specific PCR (KASP) genotyping zebrafish and staining larval zebrafish cartilage and bone. Applying the husbandry strategy described here can increase the penetrance of an interesting skeletal phenotype enabling more reproducible results in downstream applications. In addition, decreasing the mutant penetrance through this selective breeding strategy can reveal the developmental processes that most crucially require the function of the mutated gene. While the skeleton is specifically considered here, we propose that this methodology will be useful for all zebrafish mutant lines.

The goal of this protocol is to alter the penetrance of lethal skeletal mutant phenotypes in zebrafish by selective breeding. Lethal mutants cannot be grown to adulthood and bred themselves, therefore this protocol describes a method for tracking and selecting penetrance through multiple generations by progeny testing.

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be ...

Determining the Role of Maternally-Expressed Genes in Early Development with Maternal Crispants

Early development depends on a pool of maternal factors incorporated into the mature oocyte during oogenesis that perform all cellular functions necessary for development until zygotic genome activation. Typically, genetic targeting of these maternal factors requires an additional generation to identify maternal-effect phenotypes, hindering the ability to determine the role of maternally-expressed genes during development. The discovery of the biallelic editing capabilities of CRISPR-Cas9 has allowed screening of embryonic phenotypes in somatic tissues of injected embryos or "crispants," augmenting the understanding of the role zygotically-expressed genes play in developmental programs. This article describes a protocol that is an extension of the crispant method. In this method, the biallelic editing of germ cells allows for the isolation of a maternal-effect phenotype in a single generation, or "maternal crispants." Multiplexing guide RNAs to a single target promotes the efficient production of maternal crispants, while sequence analysis of maternal crispant haploids provides a simple method to corroborate genetic lesions that produce a maternal-effect phenotype. The use of maternal crispants supports the rapid identification of essential maternally-expressed genes, thus facilitating the understanding of early development.

Early development is dependent on maternally-inherited products, and the role of many of these products is currently unknown. Herein, we described a protocol that uses CRISPR-Cas9 to identify maternal-effect phenotypes in a single generation.

Early development depends on a pool of maternal factors incorporated into the mature oocyte during oogenesis that perform all cellular functions necessary ...

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

Fluorescent Protein Tagging in Zebrafish Larvae Using CRISPR/Cas9 Knock-In

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in vivo imaging due to its transparency during the early developmental stage. Here, we provide a detailed protocol for performing high-efficiency fluorescence gene KI, rapid screening for KI founders, and low-abundance protein tracing in zebrafish larvae, which will lay a critical foundation for subsequent physio-pathological studies in zebrafish. The current protocol includes complete steps for the sgRNA design for the gene of interest, sgRNA in vitro transcription, Cas9 mRNA in vitro transcription, in vivo sgRNA screen for the one with the highest efficiency, donor plasmid design and construction, microinjection in zebrafish larvae, KI founder screen and zebrafish live imaging. Critical steps, troubleshooting tips, quality control methods, and advantages and applications of this protocol are included and discussed. This protocol assures quick and accurate results at a low cost and has been validated by multiple trials.

Here, we present a protocol to label proteins with a fluorescence protein tag in zebrafish larvae, a newly developed and modified in vivo system especially useful for visualizing low-abundance proteins in zebrafish.

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in ...

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

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 ...

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological phenomena. Zebrafish have been used as an effective genetic model to study numerous questions related to heritable disease, development, and toxicology at the whole-organ and -organism level. Due to the well-annotated and mapped zebrafish genome, numerous tools for gene editing have been developed. However, the efficacy of generating and ease of detecting precise knock-in edits using CRISPR is a limiting factor. Described here is a CRISPR-Cas9-based knock-in approach with the simple detection of precise edits in a gene responsible for cardiac repolarization and associated with the electrical disorder, Long QT Syndrome (LQTS). This two-single-guide RNA (sgRNA) approach excises and replaces the target sequence and links a genetically encoded reporter gene. The utility of this approach is demonstrated by describing non-invasive phenotypic measurements of cardiac electrical function in wild-type and gene-edited zebrafish larvae. This approach enables the efficient study of disease-associated variants in a whole organism. Furthermore, this strategy offers possibilities for the insertion of exogenous sequences of choice, such as reporter genes, orthologs, or gene editors.

This protocol describes an approach to facilitate precise knock-in edits in zebrafish embryos using CRISPR-Cas9 technology. A phenotyping pipeline is presented to demonstrate the applicability of these techniques to model a Long QT Syndrome-associated gene variant.

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological ...

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

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Przeglądaj więcej filmów

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

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

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

Determining the Role of Maternally-Expressed Genes in Early Development with Maternal Crispants

Making Gynogenetic Diploid Zebrafish by Early Pressure

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts