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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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose powder&#160;</td>
			<td>Fisher BioReagents</td>
			<td>BP1356-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Breeding tanks</td>
			<td>Carolina Biological&#160;</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&#160;</td>
			<td>Thermo Scientific&#160;</td>
			<td>FERSM0241</td>
			<td></td>
		</tr>
		<tr>
			<td>dnah10 donor construct&#160;&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry. 
			Phosphorothioate bond on the donor at the first three phosphate bonds on both the 5&#8217; and 3&#8217; ends (5&#39;-CCTCTCTCCCTTTCAGAAGCTTC 
			TGCTCATCCGCTGCTTCTGCCT 
			GGACCGAGTGTACCGTGCCGTC 
			AGTGATTACGTCACGC-3&#39;)</td>
			<td></td>
		</tr>
		<tr>
			<td>dnah10 forward primer&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5&#39;-CATGGAACTCTTTCCTAATGAGT 
			TTGGC-3&#39;)</td>
			<td></td>
		</tr>
		<tr>
			<td>dnah10 reverse primer&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (&#39;5-AGTAGAGATCACACATCAACAGA 
			ATACAGC-3&#39;)&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>dnah10 synthetic sgRNA&#160;</td>
			<td>Synthego&#160;</td>
			<td>Synthetic sgRNA, target sequence: 5&#39;-GCTCATCCGCTGCTTCAGGC-3&#39;</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrophoresis power supply</td>
			<td>Thermo Scientific&#160;</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)&#160;</td>
			<td>Thermo Scientific&#160;</td>
			<td>B2</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel imaging light box&#160;</td>
			<td>Azure Biosystems</td>
			<td>AZI200-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel stain, 10000X&#160;</td>
			<td>Invitrogen</td>
			<td>S33102</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass bowl, 250 mL&#160;</td>
			<td>Generic</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Isolation tanks, 0.8 L&#160;</td>
			<td>Aquaneering</td>
			<td>ZT080</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcap capillary tube with bulb, 20 &#181;L&#160;</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&#160;</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Microwave&#160;</td>
			<td>Generic&#160;</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&#160;</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&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5&#39;-CCCAAATGTAATCCTGACCAGT 
			-3&#39;)</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&#39;-CCAGGAACACATTAACCTGGAT 
			-3&#39;)&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>p2ry12 synthetic sgRNA&#160;</td>
			<td>Synthego&#160;</td>
			<td>Synthetic sgRNA, target sequence: 5&#39;-GGCCGCACGAGGTCTCCGCG-3&#39;</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&#160;</td>
			<td>Thermo Scientific&#160;</td>
			<td>S318; 424330010</td>
			<td></td>
		</tr>
		<tr>
			<td>Sponge, 1-inch x 1-inch cut with small oval divot</td>
			<td>Generic&#160;</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&#177;0.2)&#160;</td>
			<td>Syndel</td>
			<td>200-266</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris Base, 1M (Buffered with HCl to ph 8.0)&#160;</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

1.1K 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

Novel RNA-Binding Proteins Isolation by the RaPID Methodology

RNA-binding proteins (RBPs) play important roles in every aspect of RNA metabolism and regulation. Their identification is a major challenge in modern biology. Only a few in vitro and in vivo methods enable the identification of RBPs associated with a particular target mRNA. However, their main limitations are the identification of RBPs in a non-cellular environment (in vitro) or the low efficiency isolation of RNA of interest (in vivo). An RNA-binding protein purification and identification (RaPID) methodology was designed to overcome these limitations in yeast and enable efficient isolation of proteins that are associated in vivo. To achieve this, the RNA of interest is tagged with MS2 loops, and co-expressed with a fusion protein of an MS2-binding protein and a streptavidin-binding protein (SBP). Cells are then subjected to crosslinking and lysed, and complexes are isolated through streptavidin beads. The proteins that co-purify with the tagged RNA can then be determined by mass spectrometry. We recently used this protocol to identify novel proteins associated with the ER-associated PMP1 mRNA. Here, we provide a detailed protocol of RaPID, and discuss some of its limitations and advantages.

RNA-protein interactions lie at the heart of many cellular processes. Here, we describe an in vivo method to isolate specific RNA and identify novel proteins that are associated with it. This could shed new light on how RNAs are regulated in the cell.

RNA-binding proteins (RBPs) play important roles in every aspect of RNA metabolism and regulation. Their identification is a major challenge in modern ...

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Systematic identification and characterization of genetic perturbations have proven useful to decipher gene function and cellular pathways. However, the conventional approaches of permanent gene deletion cannot be applied to essential genes. We have pioneered a unique collection of ~70 temperature-sensitive (ts) lethal mutants for studying cell cycle regulation in the unicellular green algae Chlamydomonas reinhardtii1. These mutations identify essential genes, and the ts alleles can be conditionally inactivated by temperature shift, providing valuable tools to identify and analyze essential functions. Mutant collections are much more valuable if they are close to comprehensive, since scattershot collections can miss important components. However, this requires the efficient collection of a large number of mutants, especially in a wide-target screen. Here, we describe a robotics-based pipeline for generating ts lethal mutants and analyzing their phenotype in Chlamydomonas. This technique can be applied to any microorganism that grows on agar. We have collected over 3000 ts mutants, probably including mutations in most or all cell-essential pathways, including about 200 new candidate cell cycle mutations. Subsequent molecular and cellular characterization of these mutants should provide new insights in plant cell biology; a comprehensive mutant collection is an essential prerequisite to ensure coverage of a broad range of biological pathways. These methods are integrated with downstream genetics and bioinformatics procedures for efficient mapping and identification of the causative mutations that are beyond the scope of this manuscript.

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Temperature-sensitive (ts) lethal mutants are valuable tools to identify and analyze essential functions. Here we describe methods to generate and classify ts lethal mutants in high throughput.

Systematic identification and characterization of genetic perturbations have proven useful to decipher gene function and cellular pathways. However, the ...

Robust DNA Isolation and High-throughput Sequencing Library Construction for Herbarium Specimens

Herbaria are an invaluable source of plant material that can be used in a variety of biological studies. The use of herbarium specimens is associated with a number of challenges including sample preservation quality, degraded DNA, and destructive sampling of rare specimens. In order to more effectively use herbarium material in large sequencing projects, a dependable and scalable method of DNA isolation and library preparation is needed. This paper demonstrates a robust, beginning-to-end protocol for DNA isolation and high-throughput library construction from herbarium specimens that does not require modification for individual samples. This protocol is tailored for low quality dried plant material and takes advantage of existing methods by optimizing tissue grinding, modifying library size selection, and introducing an optional reamplification step for low yield libraries. Reamplification of low yield DNA libraries can rescue samples derived from irreplaceable and potentially valuable herbarium specimens, negating the need for additional destructive sampling and without introducing discernible sequencing bias for common phylogenetic applications. The protocol has been tested on hundreds of grass species, but is expected to be adaptable for use in other plant lineages after verification. This protocol can be limited by extremely degraded DNA, where fragments do not exist in the desired size range, and by secondary metabolites present in some plant material that inhibit clean DNA isolation. Overall, this protocol introduces a fast and comprehensive method that allows for DNA isolation and library preparation of 24 samples in less than 13 h, with only 8 h of active hands-on time with minimal modifications.

This article demonstrates a detailed protocol for DNA isolation and high-throughput sequencing library construction from herbarium material including rescue of exceptionally poor-quality DNA.

Herbaria are an invaluable source of plant material that can be used in a variety of biological studies. The use of herbarium specimens is associated with ...

Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly defined states of chromatin organization. The dynamic architecture of the nucleus is controlled by multiple mechanisms and shapes the transcriptional output programs. It is, therefore, important to determine locus-specific chromatin accessibility in a reliable fashion that is preferably independent from antibodies, which can be a potentially confounding source of experimental variability. Chromatin accessibility can be measured by various methods, including the Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) assay, that allow the determination of general chromatin accessibility in a relatively low number of cells. Here we describe a FAIRE protocol that allows simple, reliable, and fast identification of genomic regions with a low protein occupancy. In this method, the DNA is covalently bound to the chromatin proteins using formaldehyde as a crosslinking agent and sheared to small pieces. The free DNA is afterwards enriched using phenol:chloroform extraction. The ratio of free DNA is determined by quantitative polymerase chain reaction (qPCR) or DNA sequencing (DNA-seq) compared to a control sample representing total DNA. The regions with a looser chromatin structure are enriched in the free DNA sample, thus allowing the identification of genomic regions with lower chromatin compaction.

The plasticity of nuclear architecture is controlled by dynamic epigenetic mechanisms including non-coding RNAs, DNA methylation, nucleosome repositioning as well as histone composition and modification. Here we describe a Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) protocol which allows the determination of chromatin accessibility in a reproducible, inexpensive, and straightforward manner.

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly ...

Mating-based Overexpression Library Screening in Yeast

Budding yeast has been widely used as a model in studying proteins associated with human diseases. Genome-wide genetic screening is a powerful tool commonly used in yeast studies. The expression of a number of neurodegenerative disease-associated proteins in yeast causes cytotoxicity and aggregate formation, recapitulating findings seen in patients with these disorders. Here, we describe a method for screening a yeast model of the Amyotrophic Lateral Sclerosis-associated protein FUS for modifiers of its toxicity. Instead of using transformation, this new screening platform relies on the mating of yeast to introduce an arrayed library of plasmids into the yeast model. The mating method has two clear advantages: first, it is highly efficient; second, the pre-transformed arrayed library of plasmids can be stored for long-term as a glycerol stock, and quickly applied to other screens without the labor-intensive step of transformation into the yeast model each time. We demonstrate how this method can successfully be used to screen for genes that modify the toxicity of FUS.

This article presents a mating-based method to facilitate overexpression screening in budding yeast using an arrayed plasmid library.

Budding yeast has been widely used as a model in studying proteins associated with human diseases. Genome-wide genetic screening is a powerful tool ...

Screening Cotton Genotypes for Reniform Nematode Resistance

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium hirsutum) varieties to improve nematode management. Most protocols involve extracting vermiform nematodes or eggs from the cotton root system or potting soil to determine population density or reproduction rate. These approaches are generally time-consuming with a small number of genotypes evaluated. An alternative approach is described here in which the root system is visually examined for nematode infection. The protocol involves inoculating cotton seedling 7 days after planting with vermiform nematodes and determining the number of females attached to the root system 28 days after inoculation. Data are expressed as the number of females per gram of fresh root weight to adjust for variation in root growth. The protocol provides an excellent method for evaluating host-plant resistance associated with the ability of the nematode to establish an infection site; however, resistance that hinders nematode reproduction is not assessed. As with other screening protocols, variation is commonly observed in nematode infection among individual genotypes within and between experiments. Data are presented to illustrate the range of variation observed using the protocol. To adjust for this variation, control genotypes are included in experiments. Nonetheless, the protocol provides a simple and rapid method to evaluate host-plant resistance. The protocol has been successfully used to identify resistant accessions from the G. arboreum germplasm collection and evaluate segregating populations of more than 300 individuals to determine the genetics of resistance. A vegetative propagation method for recovering plants for resistance breeding was also developed. After removal of the root system for nematode evaluation, the vegetative shoot is replanted to allow the development of a new root system. More than 95% of the shoots typically develop a new root system with plants reaching maturity.

Here, a protocol is presented for the rapid non-destructive screening of cotton genotypes for reniform nematode resistance. The protocol involves visually examining the roots of nematode-infected cotton seedlings to determine infection response. The vegetative shoot from each plant is then propagated to recover plants for seed production.

A rapid non-destructive reniform nematode (Rotylenchulus reniformis) screening protocol is needed for the development of resistant cotton (Gossypium ...

Characterizing Histone Post-translational Modification Alterations in Yeast Neurodegenerative Proteinopathy Models

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives each year. Effective treatment options able to halt disease progression are lacking. Despite the extensive sequencing efforts in large patient populations, the majority of ALS and PD cases remain unexplained by genetic mutations alone. Epigenetics mechanisms, such as the post-translational modification of histone proteins, may be involved in neurodegenerative disease etiology and progression and lead to new targets for pharmaceutical intervention. Mammalian in vivo and in vitro models of ALS and PD are costly and often require prolonged and laborious experimental protocols. Here, we outline a practical, fast, and cost-effective approach to determining genome-wide alterations in histone modification levels using Saccharomyces cerevisiae as a model system. This protocol allows for comprehensive investigations into epigenetic changes connected to neurodegenerative proteinopathies that corroborate previous findings in different model systems while significantly expanding our knowledge of the neurodegenerative disease epigenome.

This protocol outlines experimental procedures to characterize genome-wide changes in the levels of histone post-translational modifications (PTM) occurring in connection with the overexpression of proteins associated with ALS and Parkinson&#39;s disease in Saccharomyces cerevisiae models. After SDS-PAGE separation, individual histone PTM levels are detected with modification-specific antibodies via Western blotting.

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives ...

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding sites for transcriptional regulators to modulate gene expression. Alterations of CREs have been implicated in various diseases including cancer. While promoters and enhancers have been the primary CREs for studying gene regulation, very little is known about the role of silencer, which is another type of CRE that mediates gene repression. Originally identified as an adaptive immunity system in prokaryotes, CRISPR/Cas9 has been exploited to be a powerful tool for eukaryotic genome editing. Here, we present the use of this technique to delete an intronic silencer in the human RUNX1 gene and investigate the impacts on gene expression in OCI-AML3 leukemic cells. Our approach relies on electroporation-mediated delivery of two preassembled Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complexes to create two double-strand breaks (DSBs) that flank the silencer. Deletions can be readily screened by fragment analysis. Expression analyses of different mRNAs transcribed from alternative promoters help evaluate promoter-dependent effects. This strategy can be used to study other CREs and is particularly suitable for hematopoietic cells, which are often difficult to transfect with plasmid-based methods. The use of a plasmid- and virus-free strategy allows simple and fast assessments of gene regulatory functions.

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

Direct delivery of preassembled Cas9/guide RNA ribonucleoprotein complexes is a fast and efficient means for genome editing in hematopoietic cells. Here, we utilize this approach to delete a RUNX1 intronic silencer and examine the transcriptional responses in OCI-AML3 leukemic cells.

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding ...

A Screening Method for Identification of Heterochromatin-Promoting Drugs Using Drosophila

Drosophila is an excellent model organism that can be used to screen compounds that might be useful for cancer therapy. The method described here is a cost-effective in vivo method to identify heterochromatin-promoting compounds by using Drosophila. The Drosophila&#39;s DX1 strain, having a variegated eye color phenotype that reflects the extents of heterochromatin formation, thereby providing a tool for a heterochromatin-promoting drug screen. In this screening method, eye variegation is quantified based on the surface area of red pigmentation occupying parts of the eye and is scored on a scale from 1 to 5. The screening method is straightforward and sensitive and allows for testing compounds in vivo. Drug screening using this method provides a fast and inexpensive way for identifying heterochromatin-promoting drugs that could have beneficial effects in cancer therapeutics. Identifying compounds that promote the formation of heterochromatin could also lead to the discovery of epigenetic mechanisms of cancer development.

A Screening Method for Identification of Heterochromatin-Promoting Drugs Using Drosophila

Drosophila is a widely used experimental model suitable for screening drugs with potential applications for cancer therapy. Here, we describe the use of Drosophila variegated eye color phenotypes as a method for screening small-molecule compounds that promote heterochromatin formation.

Drosophila is an excellent model organism that can be used to screen compounds that might be useful for cancer therapy. The method described here is a ...

In Vivo CRISPR/Cas9 Screening to Simultaneously Evaluate Gene Function in Mouse Skin and Oral Cavity

Genetically modified mouse models (GEMM) have been instrumental in assessing gene function, modeling human diseases, and serving as preclinical model to assess therapeutic avenues. However, their time-, labor- and cost-intensive nature limits their utility for systematic analysis of gene function. Recent advances in genome-editing technologies overcome those limitations and allow for the rapid generation of specific gene perturbations directly within specific mouse organs in a multiplexed and rapid manner. Here, we describe a CRISPR/Cas9-based method (Clustered Regularly Interspaced Short Palindromic Repeats) to generate thousands of gene knock-out clones within the epithelium of the skin and oral cavity of mice, and provide a protocol detailing the steps necessary to perform a direct&#160;in vivo&#160;CRISPR&#160;screen for tumor suppressor genes. This approach can be applied to other organs or other CRISPR/Cas9 technologies such as CRISPR-activation or CRISPR-inactivation to study the biological function of genes during tissue homeostasis or in various disease settings.

Here we describe a rapid and direct in vivo CRISPR/Cas9 screening methodology using ultrasound-guided in utero embryonic lentiviral injections to simultaneously assess functions of several genes in the skin and oral cavity of immunocompetent mice.

Genetically modified mouse models (GEMM) have been instrumental in assessing gene function, modeling human diseases, and serving as preclinical model to ...