This work was supported by the National Institutes of Health (R21CA182197 to J.O.), the Ralph W. and Grace M. Showalter Research Trust, and by the Purdue Agricultural Science and Extension for Economic Development (AgSEED) program. Sanger sequencing data were acquired by the Purdue Genomics Core facility supported by P30 CA023168. Mary Witucki was supported by the Purdue University Center for Cancer Research Summer Undergraduate Research Program supported by the Carroll County Cancer Association. The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript. We thank Benjamin Carter, Ellen Denning, and Taylor Sabato for providing critical feedback on the manuscript. We thank the Department of Biochemistry for support of this work, and the Center for Zebrafish Research at Purdue University for their dedication in the care and welfare of our zebrafish colony. Finally, we thank the Purdue Genomics Core Facility, and contributions of Phillip San Miguel regarding the NGS services.

Publication of this video article is sponsored by New England Biolabs.

This protocol describes the production of gene knockouts in the zebrafish vertebrate model system using CRISPR-Cas9 technology. A number of protocols have previously been described to undertake CRISPR-mediated genome engineering in zebrafish15,25,26,50,51,52. This protocol builds on previous efforts by combining a number of simple yet reproducibly consistent experimental techniques, in particular HMA and NGS of multiple heterozygous fish, to create a straightforward, economical, and experimentally robust protocol for CRISPR-mediated mutagenesis in zebrafish that is appropriate for labs staffed with personnel with a range of training and experience, as well as teaching labs.
Recommendations for design and synthesis of guide RNAs are included in this protocol. A major consideration in guide RNA design is the minimization of off-target effects. Several prediction algorithms have been developed to allow CRISPR-users to access computation tools with user-friendly graphical interfaces that predict both the activity of the on-target guide and the chance of off-target effects34,35,36. A specific advantage of the zebrafish system is lowered rates of off-target effects because the Cas9 is injected into the embryos and therefore expression is transient, which has been shown in mice to result in decreased off-target effects53. Nevertheless, off-target effects have been demonstrated to occur in zebrafish54. One way to control for off-target effects is to phenotype founder zebrafish that have been generated by two independent guide RNAs that target the same gene, as these guides would be very likely to affect different off-target sites. An alternative method to minimize off-target effects that is not described in this protocol is the use of a mutated Cas9 that generates single strand breaks at the target DNA, which are repaired with high efficiency. Pairing DNA nicks within proximity of one another that are complementary to the opposite strands results in effective indel formation at the desired locus and minimizes off-target effects55,56.
In addition to having different rates of off-target effects, different sgRNAs can have different rates of mutagenesis of the desired target57,58,59. This protocol uses HMA to analyze the efficiency of mutagenesis of a given sgRNA using heteroduplex band formation33,40. Heteroduplex bands are created by hybridization of PCR-generated DNA strands that contain mismatches, and can be easily resolved using gel electrophoresis. Unlike other methods commonly used to measure indel formation, such as the T7 endonuclease assay or high resolution melt analysis25,26, HMA does not require an expensive enzyme to cut mismatched DNA, and does not require complicated analysis of PCR melt curves. Importantly, using HMA to verify high rates of indel formation in the injected population also enables the investigator to minimize the number of fish needed for subsequent production of knock-out lines, which reduces the cost of identifying a mutation with the desired characteristics.
The relative ease of generating CRISPR-based indels enables creation of multiple alleles of multiple genes at once. Web-based software is available for analysis of single mutations from heterozygous fish using Sanger sequencing of PCR products49. In the case where three or more CRISPR-mutated alleles are analyzed, NGS to characterize the nature of the indel is likely to be more cost effective to characterize the nature of the indel as this approach allows a pool of up to 50 different alleles to be characterized at once (see Table of Materials)60,61,62. Such economy of scale would likely be particularly useful in an undergraduate laboratory setting.
In summary, this protocol provides step-by-step directions for reproducibly generating high quality CRISPR-reagents (in particular, sgRNA) such that fewer adult fish need to be created and analyzed to successfully identify the mutant alleles of interest, which also reduces the time and cost of generating the desired lines. Importantly, this protocol has been designed such that it can be applied by laboratories with limited resources to produce mutant zebrafish in an affordable manner. Furthermore, we have found that this approach is suitable for undergraduates and thus expands the opportunities for education and training of undergraduate students interested in hands-on experience in CRISPR-based genome editing.

The conservation of molecular machinery across eukaryotes underlies the power of using model organisms for research. Many of these model systems facilitate the use of reverse-genetic approaches such as targeted gene knockouts to characterize the contribution of a gene product to a biological or disease process of interest. Gene disruption techniques in organisms such as zebrafish have historically relied on targeted introduction of frameshift mutations that result from imprecise repair of DSBs1,2. When a DSB is introduced into the genome, the DNA lesion is repaired through one of two pathways that are universally present in nearly all cell types and organisms: non-homologous end joining (NHEJ) and homology-directed repair (HDR)3,4. The imprecise nature of the NHEJ machinery frequently produces indels of various lengths5,6,7,8,9. Introduction of frameshift mutations in the coding sequence of a gene can produce a premature stop codon, which often renders the gene nonfunctional.
Early genome engineering strategies in zebrafish to promote indels included meganucleases, zinc-finger nucleases, and transcription activator-like effector nucleases, all of which utilized DNA-protein interactions to target a nuclease to a specific genomic target where it introduced a DSB10,11,12,13,14,15. However, these technologies are often difficult to apply due to the laborious and complex engineering needed to generate a nuclease that targets the DNA sequence of interest. Unlike previous strategies, CRISPR-based gene editing does not rely on protein-DNA interactions for targeting. Instead, the CRISPR-associated (Cas) endonuclease is directed via an RNA guide that uses nucleotide base pairing interactions to target a genomic site of interest16,17,18,19,20,21. Due to the simplicity of designing a RNA guide with the desired base pairing interactions for targeting it is relatively easy to target the Cas endonuclease to the desired locus. The type II CRISPR system in particular has been widely developed for genome editing applications due to several advantageous features including use of a single multidomain Cas nuclease (Cas9) that requires interaction with DNA to stimulate endonuclease activity and use of a single guide RNA (sgRNA) to target it to the cognate DNA sequence18. The sequence requirements necessary for targeting of the cognate sgRNA are well understood19, and the desired sgRNA is easily generated by in vitro transcription. The simplicity and robustness of the CRISPR/Cas9 approach greatly facilitates targeted genetic modification in zebrafish and a wide variety of other organisms.
The enhanced ability to undertake targeted genome editing in zebrafish as a result of developing CRISPR-based reagents has significantly increased the opportunity to study processes emblematic of vertebrate organisms such as development of the central nervous system. The zebrafish genome contains orthologs of 70% of the protein-coding genes found in the human genome as well as 84% of genes associated with diseases in humans22. Zebrafish development exhibits several key qualities that enhance its use in reverse genetic studies: the embryos are laid in large clutches, develop externally from the mother making them amenable to genetic manipulation by microinjection, and adult zebrafish sexually mature by 3 months of age, allowing for rapid propagation of desired lines23.
Numerous protocols are available that describe a variety of approaches to generate and identify CRISPR-derived indels in zebrafish24,25,26,27,28,29,30,31. However, many of these procedures are time intensive, require access to expensive equipment, and can be challenging for labs with limited expertise. The steps described herein provide a simple, robust, and economical CRISPR/Cas9-strategy to engineer zebrafish knockout lines. This protocol describes the use of a highly efficient kit to synthesize sgRNAs using DNA oligonucleotides (oligos), similar to other approaches that have been previously described32. The described protocol includes two steps in particular that greatly simplify analysis of CRISPR-mutated lines: step-by-step use of the PCR-based HMA33 to easily determine the presence of genome modifications, and sequencing analysis of heterozygous zebrafish to rapidly and easily determine the nature of multiple indels in an economical fashion. In addition, step-by-step instructions are included for robust selection, reliable production, and injection of guide RNAs. The steps provided here exemplify a robust, relatively inexpensive protocol that enables laboratory personnel with a range of expertise to contribute to the identification of gene knockouts in zebrafish.

<table><tbody><tr><td>CRISPOR</td><td></td><td></td><td>sgRNA analysis software. http://crispor.tefor.net/</td></tr><tr><td>Breaking-Cas</td><td></td><td></td><td>sgRNA analysis software. https://omictools.com/breaking-cas-tool</td></tr><tr><td>ChopChop</td><td></td><td></td><td>sgRNA analysis software. https://chopchop.rc.fas.harvard.edu/</td></tr><tr><td>Primer 3</td><td></td><td></td><td>PCR primer design. http://bioinfo.ut.ee/primer3-0.4.0/</td></tr><tr><td>Rnase free technique</td><td></td><td></td><td>http://genomics.no/oslo/uploads/PDFs/workingwithrna.pdf</td></tr><tr><td>RNase-free water</td><td>Any brand</td><td></td><td>Synthesis of guide RNAs. Thermo Fisher and Qiagen both carry suitable water.</td></tr><tr><td>Rnase-free ethanol</td><td>Any brand</td><td></td><td>Molecular biology grade ethanol is Rnase free. Fischer, VWR sell suitable ethanol.</td></tr><tr><td>Cas9 Protein</td><td>PNA Bio</td><td>CP01</td><td>Cas9 protein with nuclear localization signal.</td></tr><tr><td>Target specific oligos</td><td>Integrated DNA Technologies (IDT)</td><td></td><td>Synthesize guide RNAs.</td></tr><tr><td>PCR primers flanking guide RNA cut site</td><td>Integrated DNA Technologies (IDT)</td><td></td><td>Use for heteroduplex analysis, 100-300 bp product.</td></tr><tr><td>PCR primers flanking guide RNA cut site</td><td>Integrated DNA Technologies (IDT)</td><td></td><td>Use for sequencing CRISPR-alleles,300-600 bp product recommended.</td></tr><tr><td>EnGen sgRNA Synthesis Kit</td><td>New England BioLabs</td><td>E3322</td><td>In vitro transccribe guide RNAs.</td></tr><tr><td>10X TBE</td><td>Thermo Fisher</td><td>B52</td><td>RNase-free buffer for electrophoresis of nucleic acids.</td></tr><tr><td>6X Load Dye</td><td>New England BioLabs</td><td>B7025S</td><td>Loading DNA for electropheresis.</td></tr><tr><td>5M Ammonium acetate</td><td>Thermo Fisher</td><td>AM9071</td><td>Clean up reagent for purification of guide RNAs.</td></tr><tr><td>Ethanol (200 proof, nuclease-free)</td><td>Any brand</td><td></td><td>Clean up reagent for purification of guide RNAs. Molecular grade materials are RNase free.</td></tr><tr><td>Nuclease-free Water</td><td>Any brand</td><td></td><td>For synthesis and purification of guide RNAs.</td></tr><tr><td>RNase away</td><td>Thermo Fisher</td><td>10328011</td><td>Eliminates RNase contamination from surfaces, equipment, glassware.</td></tr><tr><td>DNA Ladder, 100 bp</td><td>New England BioLabs</td><td>N0467S</td><td>Molecular weight standards for gel electrophoresis of DNA.</td></tr><tr><td>DNA Ladder, 1 kb</td><td>New England BioLabs</td><td>N0552S</td><td>Molecular weight standards for gel electrophoresis of DNA.</td></tr><tr><td>RNA Ladder</td><td>New England BioLabs</td><td>N0364S</td><td>Single strand RNA marker to verify that full length sgRNA has been synthesized</td></tr><tr><td>TEMED</td><td>Thermo Fisher</td><td>17919</td><td>Casting polyacrylamide gel.</td></tr><tr><td>Ammonium persulfate (APS)</td><td>Sigma-Aldrich</td><td>A3678</td><td>Casting polyacrylamide gel.</td></tr><tr><td>40% Polyacrylamide (19:1)</td><td>BioRad</td><td>161-0154</td><td>Casting denaturing polyacrylamide gel.</td></tr><tr><td>Urea</td><td>Sigma-Aldrich</td><td>U5378-100G</td><td>Casting denaturing polyacrylamide gel.</td></tr><tr><td>RNA Gel Loading Dye (2x)</td><td>Thermo Fisher</td><td>R0641</td><td>Loading guide RNA onto denaturing gel for electrophoresis.</td></tr><tr><td>Ethidium bromide</td><td>Sigma-Aldrich</td><td>E1510-10ML</td><td>Visualization of nucleic acids.</td></tr><tr><td>SYBER Green</td><td>Thermo Fisher</td><td>S7563</td><td>Alternate method to ethidium bromide for detection of dsDNA in agarose or polyacrylamide gels.</td></tr><tr><td>Microcentrifuge</td><td>Eppendorf</td><td>5424</td><td>RNA clean up, PCR purification.</td></tr><tr><td>MyTaq Polymerase</td><td>Bioline</td><td>BIO-21105</td><td>For amplification of DNA using PCR.</td></tr><tr><td>Thermocycler</td><td>Any brand</td><td></td><td>PCR amplification.</td></tr><tr><td>Spectrophotometer</td><td>Any brand</td><td></td><td>NanoDrop or related product to quantify the amout of DNA/RNA.</td></tr><tr><td>E3 embryo media</td><td>Made in house</td><td></td><td>Media for raising embryos and making injeciton plate. Recipe: http://cshprotocols.cshlp.org/content/2011/10/pdb.rec66449</td></tr><tr><td>Methylene Blue</td><td>Sigma-Aldrich</td><td>M9140</td><td>Added to 1x E3 media to prevent fungal growth on embryos.</td></tr><tr><td>Petri dish</td><td>Thermo Fisher</td><td>FB0875711Z</td><td>Store embryos, cast injection plate.</td></tr><tr><td>Agarose</td><td>Denville</td><td>CA3510-8</td><td>Casting injection plate, agarose gels.</td></tr><tr><td>Microinection mold</td><td>Adaptive Science Tools</td><td>TU-1</td><td>To create wells to hold embryos during injection.</td></tr><tr><td>Phenol Red</td><td>Sigma-Aldrich</td><td>P0290&amp;nbsp;</td><td>Dye for visualization of injection.</td></tr><tr><td>Mineral Oil</td><td>Sigma-Aldrich</td><td>M5904-5ML</td><td>To calibrate needle injection volume.</td></tr><tr><td>Transfer pipettes</td><td>Any brand</td><td></td><td>Moving embryos.</td></tr><tr><td>Razor blade</td><td>Thermo Fisher</td><td>11295-10</td><td>Cutting injection needle, tail clipping adult fish.</td></tr><tr><td>Incubator</td><td>Any brand</td><td></td><td>Maintaining embryos at 28.5 &lt;sup&gt;o&lt;/sup&gt;C.</td></tr><tr><td>Verticle pipette puller</td><td>David Kopf Instruments</td><td>700C</td><td>Geneate needles for injection. Additional needle pulling instructions: http://cshprotocols.cshlp.org/content/2006/7/pdb.prot4651.long</td></tr><tr><td>Capillary tubes</td><td>Sutter Instruments</td><td>BF100-58-10</td><td>Geneate needles for injection.</td></tr><tr><td>Microloader tips</td><td>Eppendorf</td><td>930001007</td><td>Load solution into injection needles.</td></tr><tr><td>Microinjector</td><td>World Precision Instruments</td><td>PV 820</td><td>Injecting embryos.</td></tr><tr><td>Disecting microscope</td><td>Leica</td><td></td><td>Injecting embryos.</td></tr><tr><td>30% Polyacylamide (29:1)</td><td>BioRad</td><td>161-0156</td><td>Heteroduplex gel casting.</td></tr><tr><td>MS-222 (Tricaine)</td><td>Sigma-Aldrich</td><td>A-5040</td><td>Anesthetize zebrafish for tail clipping and gDNA extraction from embryos. Recipe: https://wiki.zfin.org/display/prot/TRICAINE</td></tr><tr><td>Microwave</td><td>Any brand</td><td></td><td>Casting injection plate, agarose gels.</td></tr><tr><td>Scale</td><td>Any brand</td><td></td><td>Casting injection plate, agarose gels.</td></tr><tr><td>Gloves</td><td>Any brand</td><td></td><td>For all aspects of the protocol.</td></tr><tr><td>N2</td><td>Any brand</td><td></td><td>To expell liquid from the capillary for embryo injection.</td></tr><tr><td>1,000 &amp;micro;L&amp;nbsp; tips</td><td>Any brand</td><td></td><td>For all aspects of the protocol.</td></tr><tr><td>200 &amp;micro;L tips</td><td>Any brand</td><td></td><td>For all aspects of the protocol.</td></tr><tr><td>10 &amp;micro;L tips</td><td>Any brand</td><td></td><td>For all aspects of the protocol.</td></tr><tr><td>PCR strip tubes</td><td>Any brand</td><td></td><td>For all aspects of the protocol.</td></tr><tr><td>Micropipettes</td><td>Any brand</td><td></td><td>For all aspects of the protocol.</td></tr><tr><td>NGS Sequencing platform: Wideseq</td><td></td><td></td><td>If your institution does not offer this servicce visit: https://www.purdue.edu/hla/sites/genomics/wideseq-2/ (External cost: $35).</td></tr><tr><td>Software for sequence analysis</td><td></td><td></td><td>For Sanger sequencing of heterozygous fish. Visit: http://yosttools.genetics.utah.edu/PolyPeakParser/</td></tr><tr><td>Software for sequence analysis</td><td></td><td></td><td>SnapGene Viewer, Sequence Scanner for analysis of NGS sequencing.</td></tr></tbody></table>

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.

The experimental approaches described in this protocol allow for efficient, cost-effective production of zebrafish knock-out lines using CRISPR/Cas9 technology. The following figures have been included in this article to facilitate interpretation and troubleshooting of the results obtained using this protocol. Following successful production and microinjection of CRISPR-reagents, the zebrafish embryos can be analyzed for overt phenotypes and for indel formation using HMA. A helpful control to visualize the success of the CRISPR-experiment is the use of the sgRNA described in step 1.5 to target the pigment-producing gene tyrosinase. Cas9-induced indel formation at tyrosinase results in loss of pigmentation and is easily scored by 48 hpf (Figure 1). Another helpful control to ensure that preparation of the CRISPR-reagents for injection has been successful, is to verify that full-length (120 nt) sgRNA has been synthesized using a denaturing polyacrylamide gel (Figure 2, Lane 1 and 2). If the RNA has been degraded it may appear as a smear, for example Lane 3 (Figure 2) shows degraded RNA that is not suitable for injection.
To analyze the indel formation frequency of genes targeted by CRISPR-Cas9 that do not result in overt phenotypes such as tyrosinase, HMA analysis is a simple and reliable method. sgRNA/Cas9 injected embryos analyzed using HMA results in the formation of heteroduplex bands, and reduction of the intensity of the homoduplex band (Figure 4). The presence of heteroduplex bands is further utilized in this protocol to identify potential founder fish from the microinjected embryos and as adults (Figure 4 and Figure 5), to analyze the germline transmission efficiency of a founder (Figure 6), and to verify presence of an indel in a heterozygous F1 fish (Figure 7). The heterozygous fish that contain an indel are candidates for NGS to identify the nature of the indel and to determine if a premature stop codon is present in the coding region of the target gene.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56969/56969fig1.jpg" /> 
Figure 1: Zebrafish embryos exhibit a pigment defect when injected with a sgRNA targeting tyrosinase at the one-cell stage. (A) Wild-type, uninjected embryo at 48 hpf and (B) injected embryo at 48 hpf. <a href="https://cloudflare.jove.com/files/ftp_upload/56969/56969fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56969/56969fig2.jpg" /> 
Figure 2: In vitro transcription of sgRNA using synthesis kit. Oligos were synthesized using in vitro transcription according to the sgRNA synthesis kit instructions. 500 ng of RNA was run on a urea/PAGE gel as described. sgRNA loaded in lanes 1 and 2 shows a band corresponding to the full length, intact 120 nt RNA. The sgRNA in lane 3 shows a degraded RNA sample that is not suitable for injection.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56969/56969fig3.jpg" /> 
Figure 3: Comparison of the health of 24 hpf injected embryos. A living embryo (A) developed to 24 hpf, is easily distinguished from an embryo that has aborted development (B). Embryos that resemble (B) or have drastically altered features to (A), such as spinal curvature or altered head and eye development should be removed from dish. <a href="https://cloudflare.jove.com/files/ftp_upload/56969/56969fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56969/56969fig4.jpg" /> 
Figure 4: Heteroduplex mobility assay of sgRNA-Cas9 microinjected zebrafish embryos. Pools of 5 embryos per sample were collected at 72 hpf and gDNA was extracted. Heteroduplex analysis was performed as described, samples were loaded equally with 500 ng of DNA. Lanes: M = 100 bp marker; 1 = uninjected control; 2 = injection sample 1; 3 = injection sample 2. Expected band size = 98 bp.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56969/56969fig5.jpg" /> 
Figure 5: Heteroduplex mobility assay of gDNA extracted from the tail of an adult CRISPR-injected zebrafish. Embryos that were injected with an sgRNA and Cas9 protein were grown to adulthood (3 months). Fish B and C exhibit heteroduplex bands and were subsequently bred to identify germline transmitted indels; fish A was not used in subsequent analysis because it does not exhibit a positive heteroduplex band. Lanes: 1 = wild-type control; 2 = Fish A; 3 = Fish B; 4 = Fish C. Expected band size = 98 bp. <a href="https://cloudflare.jove.com/files/ftp_upload/56969/56969fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56969/56969fig6.jpg" /> 
Figure 6: Heteroduplex mobility assay of single embryos generated by breeding a F0 CRISPR-injected zebrafish to a wild-type fish to identify germline transmitted indels. Zebrafish were mated, and the F1 embryos grown for 72 h. Single embryos were collected and heteroduplex analysis performed as described. Lanes: 1 = wild-type control; 2-10 = a single F1 embryo per lane. This gel shows that 7 out of 10 embryos show a positive heteroduplex band, indicating a germline transmission rate of 70% of the indel. Expected band size = 98 bp. <a href="https://cloudflare.jove.com/files/ftp_upload/56969/56969fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56969/56969fig7.jpg" /> 
Figure 7: Heteroduplex mobility assay of adult F1 zebrafish tail clips. Adult F1 fish were scored by HMA to identify indels. Fish that exhibited a positive heteroduplex band were PCR amplified and submitted for wide sequencing analysis to determine the nature of the mutation. (A) Lanes: 1 = wild-type control; 2 = fish A (4 bp deletion, 1 bp mismatch). (B) These F1 fish were identified from the same founder, and yet show different heteroduplex patterning, indicating germline transmission of multiple modified alleles from a single founder. Lanes: 1 = wild-type control; 2 = fish B (4 bp insertion, 7 bp mismatch), 3 = fish C (4 bp deletion, 4 bp mismatch). Each of these indels created a premature stop codon in the coding sequence of the target gene, as determined by NGS. <a href="https://cloudflare.jove.com/files/ftp_upload/56969/56969fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio at JoVE.com

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.

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 ...

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21.9K Views.  Purdue University. Gerichte genoom bewerken in het modelsysteem Danio rerio (zebravis) is sterk bevorderd door de opkomst van de CRISPR gebaseerde benaderingen. Hierin beschrijven we een gestroomlijnde, robuust protocol voor generatie en van onzin CRISPR afkomstige allelen waarin de heteroduplex mobility assay en identificatie van mutaties met behulp van de volgende generatie sequencing.

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

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play in the functioning of cells and whole organisms. This cause has been tremendously aided by the recent introduction of the CRISPR/Cas9 system-a versatile tool that allows researchers to manipulate the genome and transcriptome in order to, among other things, knock out, knock down, or knock in genes in a targeted manner. For the purpose of knocking out a gene, CRISPR/Cas9-mediated double-strand breaks recruit the non-homologous end-joining DNA repair pathway to introduce the frameshift-causing insertion or deletion of nucleotides at the break site. However, an individual guide RNA may cause undesirable off-target effects, and to rule these out, the use of multiple guide RNAs is necessary. This multiplicity of targets also means that a high-volume screening of clones is required, which in turn begs the use of an efficient high-throughput technique to genotype the knockout clones. Current genotyping techniques either suffer from inherent limitations or incur high cost, hence rendering them unsuitable for high-throughput purposes. Here, we detail the protocol for using fluorescent PCR, which uses genomic DNA from crude cell lysate as a template, and then resolving the PCR fragments via capillary gel electrophoresis. This technique is accurate enough to differentiate one base-pair difference between fragments and hence is adequate in indicating the presence or absence of a frameshift in the coding sequence of the targeted gene. This precise knowledge effectively precludes the need for a confirmatory sequencing step and allows users to save time and cost in the process. Moreover, this technique has proven to be versatile in genotyping various mammalian cells of various tissue origins targeted by guide RNAs against numerous genes, as shown here and elsewhere.

The genotyping technique described here, which couples fluorescent polymerase chain reaction (PCR) to capillary gel electrophoresis, allows for high-throughput genotyping of nuclease-mediated knockout clones. It circumvents limitations faced by other genotyping techniques and is more cost effective than sequencing methods.

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

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

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

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

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

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 ...

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

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

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

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

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

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of energy homeostasis and obesity. The small size of zebrafish allows for assays on embryos to be conducted in a 96- or 384-well plate format, Morpholino and CRISPR based technologies promote ease of genetic manipulation, and drug treatment by bath application is viable. Moreover, zebrafish are ideal for forward genetic screens allowing for novel gene discovery. Given the relative novelty of zebrafish as a model for obesity, it is necessary to develop tools that fully exploit these benefits. Herein, we describe a method to measure energy expenditure in thousands of embryonic zebrafish simultaneously. We have developed a whole animal microplate platform in which we use 96-well plates to isolate individual fish and we assess cumulative NADH2 production using the commercially available cell culture viability reagent alamarBlue. In poikilotherms the relationship between NADH2 production and energy expenditure is tightly linked. This energy expenditure assay creates the potential to rapidly screen pharmacological or genetic manipulations that directly alter energy expenditure or alter the response to an applied drug (e.g. insulin sensitizers).

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism for the study of energy homeostasis. By utilizing a NADH2 sensitive redox indicator, alamar Blue, we have developed an assay that measures the metabolic rate of zebrafish larvae in a 96 well plate format and can be applied to drug or gene discovery.

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of ...

Biology

One-step CRISPR-based Strategy for Endogenous Gene Tagging in Drosophila melanogaster

The study of protein subcellular localization, dynamics, and regulation in live cells has been profoundly transformed by the advent of techniques that allow the tagging of endogenous genes to produce fluorescent fusion proteins. These methods enable researchers to visualize protein behavior in real time, providing valuable insights into their functions and interactions within the cellular environment. Many current gene tagging studies employ a two-step process where visible markers, such as eye color changes, are used to identify genetically modified organisms in the first step, and the visible marker is excised in the second step. Here, we present a one-step protocol to perform precise and rapid endogenous gene tagging in Drosophila melanogaster, which enables screening for engineered lines without the visible eye marker, offering a significant advantage over past methods. To screen for successful gene-tagging events, we employ a PCR-based technique to genotype individual flies by analyzing a small segment from their middle leg. Flies that pass the screening criteria are then used to produce stable stocks. Here, we detail the design and construction of CRISPR editing plasmids and methods for screening and confirmation of engineered lines. Together, this protocol improves the efficiency of endogenous gene tagging in Drosophila significantly and enables studies of cellular processes in vivo.

One-step CRISPR-based Strategy for Endogenous Gene Tagging in Drosophila melanogaster

Here, we present a simplified endogenous gene tagging protocol for Drosophila, which utilizes a PCR-based technique for marker-free identification of successful genetic modifications, facilitating the development of stable knock-in lines.

The study of protein subcellular localization, dynamics, and regulation in live cells has been profoundly transformed by the advent of techniques that ...

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

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