We would like to thank Chuck Kimmel for guidance, John Dowd for help in developing this breeding strategy, Macie Walker for her work in perfecting the skeletal stain, and Charline Walker and Bonnie Ullmann for helpful zebrafish advice. This work was supported by K99/R00 DE024190 to JTN.

All experiments described in this protocol were completed in accordance and compliance with the University of Colorado and the University of Oregon Institutional Animal Care and Use Committees (IACUC).
1. Preparing the Unselected Starting Stock
<ol>
	<li>Identify heterozygous carriers of the mutant allele of interest by fin clip11 and genotyping a stock of full-sibling animals by a method of choice, such as KASP12,13,14. This protocol was performed with the zebrafish mef2cab1086 strain.</li>
	<li>KASP genotyping 
	NOTE: Allele specific primers for KASP are designed by the company that provides the reagents (see Table of Materials). To obtain the appropriate 6-carboxyl-X-rhodamine (ROX) dye concentration for the specific real-time PCR machine visit the company website.
	<ol>
		<li>Place the zebrafish tail tissue in 50 &#181;L of lysis buffer (10 mM tris (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.3% non-ionic surfactant, 0.3% Octylphenyl-polyethylene glycol). Add 10 &#181;L of 10 mg/mL Proteinase-K per well of a 96-well PCR plate and digest in a thermal cycler at 55 &#176;C for 2 - 5 h followed by a 20 min 94 &#176;C inactivation step. Refrigerate the lysis product after digestion.&#160; 
		NOTE: Lysis products can be successfully used for several months after preparation. When tissue was lysed using 50 mM NaOH, the KASP reactions were unsuccessful. These findings suggest that the NaOH method of genomic DNA preparation is not compatible with KASP.</li>
		<li>Quantify the genomic DNA using a spectrophotometer. Dilute the genomic DNA template using molecular biology grade water so that each reaction contains approximately 20-30 ng. 
		NOTE: Quantify 4 or 5 samples, pool them, then prepare dilutions for all samples based on the average. When using this protocol for adult tail tissue, a 1:100 dilution is often sufficient. The precise quantification of DNA concentration is not required.</li>
		<li>Add 0.14 &#181;L of the KASP primer mix and 5 &#181;L of the KASP master mix per sample on ice into a 1.5 mL microcentrifuge tube and mix well before briefly centrifuging to collect contents at the bottom of the tube. 
		NOTE: KASP master mix is heat and light sensitive, keep stock on ice and avoid direct light.</li>
		<li>Pipet 5 &#181;L of primer/master mix solution into the wells of a 96-well optical PCR plate suitable for the real-time PCR machine being used.</li>
		<li>Pipet 5 &#181;L of diluted (approximately 5 - 7 ng/&#181;L) DNA template samples into each well and aspirate with a pipette to mix.</li>
		<li>Add 5 &#181;L of nuclease-free water to 3 wells to serve as no-template controls (NTC) and add 5 &#181;L of diluted confirmed homozygous wild type, heterozygous and homozygous mutant DNA template for positive controls.</li>
		<li>Place an optically clear film seal over the plate ensuring that the film is fully sealed on each well.</li>
		<li>Briefly centrifuge the plate at 600 x g to collect the contents at the bottom and to remove bubbles from the mix. 
		NOTE: Keep the plate on ice and shield from light until ready to proceed.</li>
		<li>Use the cycling program shown in Table 1 to perform the main KASP reaction.</li>
		<li>View the resulting scatter plot produced by the analysis computer. A successful reaction will show 4 distinct groupings of points on the plot (Figure 3): three sample groupings corresponding to genotype, and one grouping of the NTCs near the origin. The positive controls should segregate with the appropriate sample grouping.</li>
		<li>If the computer program does not recognize the plot groupings as corresponding to specific genotypes, additional cycling sets may be required (Table 2). Repeat the additional program until the computer recognizes tight groupings by genotype.</li>
		<li>Export the genotyping results to a spreadsheet file for easier use and analysis.</li>
	</ol>
	</li>
	<li>House the identified heterozygous animals together on the main water system.</li>
</ol>
2. The First Round of Progeny Scoring
<ol>
	<li>Pairwise cross sibling heterozygotes
	<ol>
		<li>After the animals recover from the fin clip, set up pairwise intercrosses. Use dividers to ensure embryos are synchronous16.</li>
		<li>Collect the embryos and keep the parental pairs isolated in breeding cages.</li>
		<li>Monitor isolated adults so that aggressive fish can be separated or provided with cover (shredded fish nets work nicely). Fish should be fed and their water changed following IACUC guidelines while in static water.</li>
	</ol>
	</li>
	<li>Embryo rearing
	<ol>
		<li>Stage and raise the zebrafish embryos to larvae under standard conditions17.</li>
		<li>Collect phenotypically wild-type animals using a glass pipette on day 5 or 6 when the swim bladder is inflated in 75% of the clutch. Place phenotypic wild types in the nursery recording of which parents yielded them. 
		Note: With mutant alleles that are recessive lethal, which is true of many skeletal mutants, homozygous mutants fail to form swim bladders18,19. Therefore, phenotypic wild types can be easily discerned from their mutant siblings based on inflated swim bladders.</li>
	</ol>
	</li>
	<li>Alcian Blue/Alizarin Red cartilage and bone staining
	<ol>
		<li>Anesthetize larvae that do not inflate their swim bladders using 0.17 g/L Tricaine-S in embryo media. Collect animals into 1.5 mL microcentrifuge tubes with a wide-bore glass Pasteur pipet. Add no more than 100 larvae per tube.</li>
		<li>Remove embryo water from each tube and fix animals in 1 mL of 2% paraformaldehyde in 1x PBS per tube. 
		CAUTION: Aqueous PFA is combustible and will cause serious skin and eye irritation. Wear appropriate personal protective equipment such as gloves and eye protection, and wash hands thoroughly after use.</li>
		<li>Rock for 1 h; longer fixation impairs bone staining. Check the tubes regularly during this and all subsequent rocking steps to ensure no larvae have become stuck to the side or clumped in the bottom of the tube.</li>
		<li>Remove the fixative from tubes using a glass pipette, add 1 mL of 50% ethanol, and rock for 10 min.</li>
		<li>Prepare fresh staining solution by adding 20 &#181;L of 0.5% Alizarin Red per 1 mL of Alcian premix (0.04% Alcian Blue/10 mM MgCl2/80% ethanol). Remove 50% ethanol, add 1 mL of staining solution to each tube, and rock overnight. Larvae can remain in the stain for up to 5 days.</li>
		<li>Remove the stain solution from the tubes and add 1 mL of 80% ethanol/10 mM MgCl2 solution to each tube. Rock the tubes for at least 1 h; rinsing overnight can produce a clearer Alcian Blue stain.</li>
		<li>Remove the 80% ethanol/10 mM MgCl2 solution from the tubes and add 1 mL of 50% ethanol; rock for 5 min.</li>
		<li>Remove the 50% ethanol solution from the tubes and add 1 mL of 25% ethanol and rock for 5 min.</li>
		<li>Remove the 50% ethanol solution from the tubes and add 1 mL freshly prepared 3% H2O2/0.5% KOH bleaching solution. Leave the caps open and let the tubes stand in in a micro-tube rack for approximately 10 min or until a faint brown coloration can be seen in the upper part of the solution. 
		NOTE: A layer of bubbles form on top of the solution. Careful timing is required at this step, as over bleaching will result in a poor double stain.</li>
		<li>Remove the bleaching solution and add 1 mL of 25% glycerol/0.1% KOH; rock for 10 min to 1 h.</li>
		<li>Remove 25% glycerol/0.1% KOH solution from the tubes, add 1 mL of 50% glycerol/0.1% KOH solution to each tube, rock for 10 min to overnight.</li>
		<li>Remove 50% glycerol/0.1% KOH solution from the tubes and again add 50% glycerol/0.1% KOH solution to each tube. Rocking overnight will help remove air bubbles that can accumulate inside larvae. Store stained skeletal preparations at 4 &#176;C when not being used.</li>
	</ol>
	</li>
	<li>Phenotype scoring
	<ol>
		<li>Score stained mutant offspring for the penetrance of the selected phenotype. In Nichols et al.1 mutants were scored for any occurrence of ectopic bone. 
		Note: Because the zebrafish parents are in static water and the Alizarin Red stain can fade with time, it is important to score skeletons within a few days of completing the skeletal preparation.</li>
		<li>Penetrance is the proportion of a genotype that has a phenotype. Calculate the percent penetrance by the following formula: 
		% penetrance = (mutants with phenotype)/(total number of mutants) &#215; 100</li>
	</ol>
	</li>
</ol>
3. Family Selection
<ol>
	<li>Choose two high penetrance and two low penetrance families for the next generation. In addition to penetrance, it is important to consider fecundity and vigor when choosing which families to propagate; see Discussion for more details on this.</li>
	<li>Give each parental pair a sub stock identifier: family .01, .02, etc.</li>
	<li>House parental pairs on the main system with several mixed sex &#39;companion&#39; fish. Companion fish can be any zebrafish line with permanent, obvious distinguishing features, such as altered pigmentation or fin structure. 
	NOTE: While many researchers successfully keep only breeding pairs in a tank, favorable results are seen by housing pairs with several companion fish. This optional step allows breeding pairs of animals to be kept in larger schools and easily retrieved so they can be repeatedly crossed as needed. Parental pairs can be crossed repeatedly to generate large full-sibling families; wait at least one week before repeat crossing the same parental pair.</li>
	<li>Cull the larval families from step 2.2.2. that were not selected for the next generation.</li>
	<li>Label the tanks containing the wild-type sibling larvae from selected families with the penetrance from their mutant siblings and raise to adulthood as normal.</li>
	<li>The next generation will have at least four full-sibling families, two for the upward line and two for the downward line.</li>
	<li>When larvae reach sexual maturity, repeat the protocol starting at step one with the new generation.</li>
</ol>

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G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+essential+role+of+variant+histone+H3.3+for+ectomesenchyme+potential+of+the+cranial+neural+crest.">An essential role of variant histone H3.3 for ectomesenchyme potential of the cranial neural crest.</a> PLoS Genet. 8 (9), e1002938 (2012).</li><li>DeLaurier, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+mef2ca+in+developmental+buffering+of+the+zebrafish+larval+hyoid+dermal+skeleton.">Role of mef2ca in developmental buffering of the zebrafish larval hyoid dermal skeleton.</a> Dev Biol. 385 (2), 189-199 (2014).</li><li>McGurk, P. D., Ben Lovely, C., Eberhart, J. 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The authors have nothing to disclose.

Selective Breeding Unveils Subtleties of Gene Function
Shifting mutant phenotypes to be either more or less severe by selective breeding is a straightforward way to gain new insights into gene function. When compared with standard methods of unselected breeding, the protocol presented here can yield a much more complete understanding of mutant phenotypes. Specifically, by generating strains that are severe, the full breadth of mutant phenotypes may be revealed, including some ...

The zebrafish is a powerful model system for understanding skeletal development. With mutant zebrafish strains, biologists can decipher gene function during skeletogenesis. However, zebrafish skeletal mutant phenotypes can present with variable penetrance1,2,3,4 which can hinder developmental and genetic analyses. The purpose of this method is threefold. First, generating zebrafish mutant lines which consistently produce severe phenotypes enables downstream developmental studies like time-lapse recording5 and transplantation6. These sorts of studies can be crippled by attempting to study phenotypes that only manifest inconsistently. Second, inbreeding zebrafish strains can decrease genetic background variation, thus promoting experimental consistency and reproducibility. For example, performing all in situ hybridization analyses on one selectively inbred strain can reduce confounding variability and strengthen conclusions. Third, generating severe and mild strains will reveal the entire phenotypic series that can result from a particular mutation.
At first glance, selective breeding of lethal mutants seems impossible. How can one breed for penetrance when the animals that are scored for selection are dead? Fortunately, methods for selective breeding by family selection, specifically progeny testing, have demonstrated effectiveness in livestock breeding programs for many years7,8. These programs are mainly used for selective breeding for traits that are only present in one sex, like milk production in cows or egg production in hens. The males of these species cannot be scored directly, but their progeny are scored and a value is then assigned to the parents. Borrowing from this strategy, the protocol presented here involves scoring the fixed and stained mutant offspring from a pair of zebrafish that are heterozygous for a mutant gene of interest. The penetrance of a phenotype in the homozygous lethal mutant offspring is assigned to the parents when deciding which individuals will produce the next generation in the line. We find that this method is an effective means of shifting penetrance in zebrafish lethal skeletal mutants1.
Similar to other studies, this selective breeding protocol takes under consideration criteria like clutch size, survival of offspring, normal development of embryos, and sex ratio9. However, these factors are all considered in the context of a mutant background with the objective of shifting the mutant penetrance. Therefore, this protocol extends previous selective breeding paradigms by offering a method to strengthen developmental mutant analyses as well as increase background homogeneity.
This protocol requires extensive genotyping, so it is important to develop a reliable, rapid genotyping protocol in advance. There are many genotyping protocols available10,11, however we find the KASP genotyping12,13,14 is faster, more cost efficient, and more reliable than methods based on restriction enzyme cleavage of amplified sequences10. Therefore, we include a KASP protocol in this work. Additionally, we focus on skeletal mutant phenotypes in this protocol and include a procedure for Alcian Blue/Alizarin Red staining modified from previous protocols15.
The method described here is a straightforward strategy for shifting lethal mutant penetrance upward or downward. While this protocol focuses on skeletal mutant phenotypes, we believe it will be a useful strategy for husbandry of all mutant zebrafish lines. In fact, the utility of this breeding strategy likely extends beyond zebrafish. We predict that this protocol can be modified to shift penetrance in a broad range of organisms. Shifting lethal penetrance by progeny testing can help push forward the progress of any developmental geneticist.

<table><tbody><tr><td>Paraformaldehyde, pelleted, solid</td><td>Ted Pella Co.</td><td>18501</td><td>Pelleted PFA is a safer alternative to powdered PFA</td></tr><tr><td>Magnesium Chloride, solid</td><td>Acros Organics</td><td>223210010</td><td></td></tr><tr><td>10x PBS, Aqueous</td><td>Fisher</td><td>BP3994</td><td></td></tr><tr><td>190 proof Ethanol</td><td></td><td></td><td></td></tr><tr><td>Alcian Blue, solid</td><td>Anatech Ltd.</td><td>867</td><td>Must be from Anatech</td></tr><tr><td>Alizarin Red, solid</td><td>Sigma</td><td>A5533-25G</td><td></td></tr><tr><td>Glycerol, liquid</td><td>Fisher</td><td>BP229 1</td><td></td></tr><tr><td>Hydrogen peroxide, liquid</td><td>Fisher</td><td>BP263500</td><td></td></tr><tr><td>Potassium hydroxide,&amp;nbsp; solid</td><td>Fisher</td><td>P250 500</td><td></td></tr><tr><td>StepOnePlus Real-time PCR Machine</td><td>Applied Biosystems</td><td></td><td></td></tr><tr><td>MicroAmp Fast Optical 96-well Reaction Plate with Barcode (0.1 mL)</td><td>Applied Biosystems</td><td>4346906</td><td></td></tr><tr><td>Microseal &#039;B&#039; seal</td><td>BioRad</td><td>MSB1001</td><td></td></tr><tr><td>KASP Master Mix, High ROX</td><td>LGC</td><td>KBS-1016-022</td><td>https://www.lgcgroup.com/products/kasp-genotyping-chemistry/#.WOPX41UrKUk</td></tr><tr><td>KASP By Design Primer Mix</td><td>LGC</td><td>KBS-2100-100</td><td></td></tr><tr><td>Tris HCl, solid</td><td>Fisher</td><td>BP153 500</td><td></td></tr><tr><td>potassium chloride, solid</td><td>Fisher</td><td>BP366 500</td><td></td></tr><tr><td>Tween-20, liquid</td><td>Fisher</td><td>BP337 100</td><td></td></tr><tr><td>Nonidet P40</td><td>ThermoFisher</td><td>28324</td><td></td></tr><tr><td>Tricaine-S</td><td>Western Chemicals</td><td></td><td></td></tr><tr><td>Proteinase K</td><td>Fisher</td><td>BP1700 100</td><td></td></tr><tr><td>T100 Thermal Cycler</td><td>BioRad</td><td>1861096</td><td></td></tr><tr><td>Controlled Drop Pasteur Pipets</td><td>Fisher</td><td>13-678-30</td><td></td></tr><tr><td>Nanodrop</td><td>ThermoFisher</td><td></td><td>for DNA quantitation</td></tr></tbody></table>

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.

This protocol is a long-term husbandry technique useful for understanding zebrafish skeletal mutants (Figure 1). Selective breeding by progeny testing should yield a shift in overall penetrance both downward and upward in a few generations (Figure 2). In our previous work, two rounds of selective breeding drove the average penetrance downward from 17% to 3%1. Similarly, in our upward line, we shifted the a...

Watch this Scientific Journal Video about Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing at JoVE.com

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

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 ...

shifting-zebrafish-lethal-skeletal-mutant-penetrance-progeny

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7.5K Views.  University of Colorado Anschutz Medical Campus. 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. 

Video: Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

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 ...

Genetics

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 ...

Zebrafish Model of Neuroblastoma Metastasis

Zebrafish has emerged as an important animal model to study human diseases, especially cancer. Along with the robust transgenic and genome editing technologies applied in zebrafish modeling, the ease of maintenance, high-yield productivity, and powerful live imaging altogether make the zebrafish a valuable model system to study metastasis and cellular and molecular bases underlying this process in vivo. The first zebrafish neuroblastoma (NB) model of metastasis was developed by overexpressing two oncogenes, MYCN and LMO1, under control of the dopamine-beta-hydroxylase (d&#946;h) promoter. Co-overexpressed MYCN and LMO1 led to the reduced latency and increased penetrance of neuroblastomagenesis, as well as accelerated distant metastasis of tumor cells. This new model reliably reiterates many key features of human metastatic NB, including involvement of clinically relevant and metastasis-associated genetic alterations; natural and spontaneous development of metastasis in vivo; and conserved sites of metastases. Therefore, the zebrafish model possesses unique advantages to dissect the complex process of tumor metastasis in vivo.

This paper introduces the method of developing, characterizing, and tracking in real-time the tumor metastasis in zebrafish model of neuroblastoma, specifically in the transgenic zebrafish line with overexpression of MYCN and LMO1, which develops metastasis spontaneously.

Zebrafish has emerged as an important animal model to study human diseases, especially cancer. Along with the robust transgenic and genome editing ...

Cancer Research

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

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

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

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

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in developmental biology for decades. Traditionally, most gene expression studies relied on visual evaluation of the ISH signal, a method that is prone to bias, particularly in cases where sample identities are known a priori. We have previously reported on a method to circumvent this bias and provide a more accurate quantification of ISH signals. Here, we present a simple guide to apply this method to quantify the expression levels of genes of interest in ISH-stained embryos and correlate that with their corresponding genotypes. The method is particularly useful to quantify spatially restricted gene expression signals in samples of mixed genotypes and it provides an unbiased and accurate alternative to the traditional visual scoring methods.

Gene editing technologies have enabled researchers to generate zebrafish mutants to investigate gene function with relative ease. Here, we provide a guide to perform parallel embryo genotyping and quantification of in situ hybridization signals in zebrafish. This unbiased approach provides greater accuracy in phenotypical analyses based on in situ hybridization.

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in ...

An Efficient Protocol to Assess ERK Activity Modulation in Early Zebrafish Noonan Syndrome Models via Live FRET Microscopy and Immunofluorescence

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. Despite a broad genetic heterogeneity, germline gain-of-function mutations in key regulators of the RAS-MAPK pathway underlie the majority of the cases, and, thanks to advanced sequencing techniques, potentially pathogenic variants affecting the RAS-MAPK pathway continue to be identified. Functional validation of the pathogenicity of these variants, essential for accurate diagnosis, requires fast and reliable protocols, preferably in vivo. Given the scarcity of effective treatments in early childhood, such protocols, especially if scalable in cost-effective animal models, can be instrumental in offering a preclinical ground for drug repositioning/repurposing. 
Here we describe step-by-step the protocol for rapid generation of transient RASopathy models in zebrafish embryos and direct inspection of live disease-associated ERK activity changes occurring already during gastrulation through real-time multispectral F&#246;rster resonance energy transfer (FRET) imaging. The protocol uses a transgenic ERK reporter recently established and integrated with the hardware of commercial microscopes. We provide an example application for Noonan syndrome (NS) zebrafish models obtained by expression of the Shp2D61G. We describe a straightforward method that enables registration of ERK signal change in the NS fish model before and after pharmacological signal modulation by available low-dose MEK inhibitors. We detail how to generate, retrieve, and assess ratiometric FRET signals from multispectral acquisitions before and after treatment and how to cross-validate the results via classical immunofluorescence on whole embryos at early stages. We then describe how, via examining standard morphometric parameters, to query late changes in embryo shape, indicative of a resulting impairment of gastrulation, in the same embryos whose ERK activity is assessed by live FRET at 6 h post fertilization.

RASopathies are multisystem genetic syndromes caused by RAS-MAPK pathway hyperactivation. Potentially pathogenic variants awaiting validation emerge continuously while poor preclinical evidence limits therapy. Here, we describe our in vivo protocol to test and cross-validate RASopathy-associated ERK activation levels and its pharmacological modulation during embryogenesis by live FRET imaging in Teen-reporter zebrafish.

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. ...

Biochemistry

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present methods for the development of assays to query potentially clinically significant nonsynonymous changes using in vivo complementation in zebrafish. Zebrafish (Danio rerio) are a useful animal system due to their experimental tractability; embryos are transparent to enable facile viewing, undergo rapid development ex vivo, and can be genetically manipulated.1 These aspects have allowed for significant advances in the analysis of embryogenesis, molecular processes, and morphogenetic signaling. Taken together, the advantages of this vertebrate model make zebrafish highly amenable to modeling the developmental defects in pediatric disease, and in some cases, adult-onset disorders. Because the zebrafish genome is highly conserved with that of humans (~70% orthologous), it is possible to recapitulate human disease states in zebrafish. This is accomplished either through the injection of mutant human mRNA to induce dominant negative or gain of function alleles, or utilization of morpholino (MO) antisense oligonucleotides to suppress genes to mimic loss of function variants. Through complementation of MO-induced phenotypes with capped human mRNA, our approach enables the interpretation of the deleterious effect of mutations on human protein sequence based on the ability of mutant mRNA to rescue a measurable, physiologically relevant phenotype. Modeling of the human disease alleles occurs through microinjection of zebrafish embryos with MO and/or human mRNA at the 1-4 cell stage, and phenotyping up to seven days post fertilization (dpf). This general strategy can be extended to a wide range of disease phenotypes, as demonstrated in the following protocol. We present our established models for morphogenetic signaling, craniofacial, cardiac, vascular integrity, renal function, and skeletal muscle disorder phenotypes, as well as others.

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present a systematic approach for developing physiologically relevant, sensitive and specific in vivo assays for interpreting variation in human pathology. Transient genetic manipulation via microinjection of WT and mutant human mRNA and morpholino (MO) antisense oligonucleotides harness the tractability of the developing zebrafish embryo to rapidly assay pathogenic mutations, especially, but not exclusively, in the context of human developmental disorders.

Here,  we present methods for the development of assays to query potentially clinically  significant nonsynonymous changes using in  vivo complementation ...

Biology

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

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

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

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

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

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

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

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

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

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

Summary

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Chapters in this video

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

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

Zebrafish Model of Neuroblastoma Metastasis

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

An Efficient Protocol to Assess ERK Activity Modulation in Early Zebrafish Noonan Syndrome Models via Live FRET Microscopy and Immunofluorescence

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

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