The authors acknowledge the heroic efforts of Monica Lucas, Katherine Shaw, Ryan Taylor, Jared Thompson, Nicole Robichaud, and Hope Healey for help with fish husbandry, data collection, and early protocol development. We would also like to thank the three reviewers for their suggestions to the manuscript. We believe the final product to be of better quality after addressing their comments. This work was funded by support from the National Science Foundation #1644965 and #1455405 to JRG, and the Natural Sciences and Engineering Research Council DG grant to VLS.

The authors have nothing to disclose.

The phenotypic richness of weakly electric fish, together with a recent proliferation of genomics resources, motivates a strong need for functional genomic tools in the weakly electric fish model. This system is particularly attractive because of the convergent evolution of numerous phenotypic traits in parallel lineages of fish, which are easily kept in the laboratory.
The protocol described here demonstrates the efficacy of the CRISPR/Cas9 technique in lineages of weakly electric fish that e...

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. Two lineages of teleost fish, osteoglossiformes and siluriformes, evolved electroreception in parallel, and five lineages of teleosts (gymnotiformes, mormyrids, and the genera Astroscopus, Malapterurus, and Synodontis) evolved electrogenesis in parallel. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture1,2,3.
This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades, which produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate4, a growing community of scientists has gained tremendous insights into evolution of development1,5,6, systems and circuits neuroscience7,8,9,10, cellular physiology11,12, ecology and energetics13,14,15,16,17, behavior18,19, and macroevolution3,20,21.
More recently, there has been a proliferation of genomic, transcriptomic, and proteomic resources for electric fish1,22,23,24,25,26,27,28. Use of these resources has already produced important insights regarding the connection between genotype and phenotype in these species1,2,3,28,29,30. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools31.
One such tool is the recently developed Clustered Regularly Interspaced Short Palindromic Repeats paired with Cas9 endonuclease (CRISPR/Cas9, CRISPR) technique. CRISPR/Cas9 is a genome editing tool that has entered widespread use in both model32,33,34 and non-model organisms35,36,37 alike. CRISPR/Cas9 technology has progressed to a point where a laboratory capable of basic molecular biology can easily generate gene-specific probes called short guide RNAs (sgRNAs), at a low cost using a non-cloning method38. CRISPR has advantages over other knockout/knockdown strategies, such as morpholinos39,40, transcription activator-like effector nucleases&#160;(TALENs), and zinc finger nucleases (ZFNs), which are costly and time-consuming to generate for every target gene.
The CRISPR/Cas9 system functions to create gene knockouts by targeting a specific region of the genome, directed by the sgRNA sequence, and causing a double-stranded break. The double-stranded break is detected by the cell and triggers endogenous DNA repair mechanisms preferentially using the non-homologous end joining (NHEJ) pathway. This pathway is highly error-prone: during the repair process, the DNA molecule will often incorporate insertions or deletions (indels) at the double-stranded break site. These indels can result in a loss of function due to either (1) shifts in the open reading frame, (2) insertion of a premature stop codon, or (3) shifts in the critical primary structure of the gene product. In this protocol, we utilize CRISPR/Cas9 editing to target point mutations in target genes using the NHEJ in weakly electric fish species. While simpler and more efficient than other techniques, this method of mutagenesis is expected to result in a range of phenotypic severities in F0, which is attributed to genetic mosaicism41,42,43,44.
Selection of Organisms 
For the purposes of facilitating future studies on the comparative genomics of weakly electric fish, a representative species for both gymnotiforms and mormyrids for protocol development needed to be selected. Following discussions during the 2016 Electric Fish meeting in Montevideo, Uruguay, there was community consensus to utilize species that already could be bred in the laboratory and that had genomic resources available. The gymnotiform Brachyhypopomus gauderio and the mormyrid Brienomyrus brachyistius were selected as species that fit these criteria. In both species, natural cues to induce and maintain breeding conditions are easy to mimic in captivity. B. gauderio, a gymnotiform species from South America, has the advantage of low husbandry requirements: fish can be kept at relatively high density in relatively small (4 L) tanks. B. gauderio also has fast generational turnover under captive conditions. Under laboratory conditions, B. gauderio can develop from egg to adult in about 4 months.
B. brachyistius, a species of mormyrid fish from West-Central Africa, breeds readily in captivity. B. brachyistius is readily available through the aquarium trade, has been widely used in many studies, and now has a number of genomic resources available. Their life cycle spans 1&#8722;3 years, depending on laboratory conditions. Husbandry requirements are somewhat more intensive for this species, requiring moderately sized tanks (50&#8722;100 L) due to their aggression during breeding.
Laboratories studying other species of electric fish should be able to easily adapt this protocol as long as the species can be bred, and single cell embryos can be collected and reared into adulthood. Housing, larval husbandry, and in vitro fertilization (IVF) rates will likely change with other species; however, this protocol can be used as a starting point for breeding attempts in other weakly electric fish.
An Ideal Gene Target for Proof of Concept: scn4aa 
Weakly electric mormyrid and gymnotiform fish generate electric fields (electrogenesis) by discharging a specialized organ, called the electric organ. Electric organ discharges (EODs) result from the simultaneous production of action potentials in the electric organ cells called electrocytes. EODs are detected by an array of electroreceptors in the skin to create high-resolution electrical images of the fish&#8217;s surroundings45. Weakly electric fish are also capable of detecting features of their conspecifics&#8217; EOD waveforms18 as well as their discharge rates46, allowing EODs to function additionally as a social communication signal analogous to birdsong or frog vocalizations47.
A main component of action potential generation in the electrocytes of both mormyrid and gymnotiform weakly electric fish is the voltage-gated sodium channel NaV1.42. Non-electric teleosts express two paralogous gene copies, scn4aa and scn4ab, coding for the voltage-gated sodium channel NaV1.430. In both gymnotiform and mormyrid weakly electric fish lineages, scn4aa has evolved rapidly and undergone numerous amino acid substitutions that affect its kinetic properties48. Most importantly, scn4aa has become compartmentalized in both lineages to the electric organ2,3. The relatively restricted expression of scn4aa to the electric organ, as well as its key role in the generation of EODs, makes it an ideal target for CRISPR/Cas9 knockout experiments, as it has minimal deleterious pleiotropic effects. Because weakly electric fish begin discharging their larval electric organs 6&#8722;8 days post fertilization (DPF), targeting of scn4aa is ideally suited for rapid phenotyping following embryo microinjection.

<table><tbody><tr><td>20 mg/mL RNA grade Glycogen</td><td>Thermo Scientific</td><td>R0551</td><td></td></tr><tr><td>50 bp DNA ladder</td><td>NEB</td><td>N3236L</td><td></td></tr><tr><td>Borosilicate glass capillary with filament</td><td>Sutter Instrument</td><td>BF100-58-10</td><td>(O.D. 1.0 mm, I.D. 0.58 mm, 10 cm length)</td></tr><tr><td>Cas9 protein with NLS; 1 mg/mL</td><td>PNA Biology</td><td>CP01</td><td></td></tr><tr><td>Dneasy Blood &amp;amp; Tissue Kit</td><td>Qiagen</td><td>69506</td><td></td></tr><tr><td>Eppendorf FemptoJet 4i Microinjector</td><td>Fisher Scientific</td><td>E5252000021</td><td></td></tr><tr><td>Eppendorf Microloader Pipette Tips</td><td>Fisher Scientific</td><td>10289651</td><td></td></tr><tr><td>Hamilton syringe</td><td>Fisher Scientific</td><td>14-824-654</td><td>referred to as &quot;precision glass syringe&quot; in the protocol</td></tr><tr><td>Kimwipe</td><td>Fisher Scientific</td><td>06-666</td><td>referred to as &quot;delicate task wipe&quot; in the protocol</td></tr><tr><td>MEGAscript T7 Transcription Kit</td><td>Invitrogen</td><td>AM1334</td><td></td></tr><tr><td>NEBuffer 3</td><td>NEB</td><td>B7003S</td><td>used for in vitro cleavage assay</td></tr><tr><td>OneTaq DNA kit</td><td>NEB</td><td>M0480L</td><td></td></tr><tr><td>Ovaprim</td><td>Syndel USA</td><td>https://www.syndel.com/ovaprim-ovammmlu010.html</td><td>referred to as &quot;spawning agent&quot; in the protocol</td></tr><tr><td>Parafilm</td><td>Fisher Scientific</td><td>S37440</td><td>referred to as &quot;thermoplastic&quot; in the protocol</td></tr><tr><td>Pipette puller</td><td>WPI</td><td>SU-P97</td><td>sutter brand</td></tr><tr><td>QIAquick PCR Purification Kit</td><td>Qiagen</td><td>28106</td><td></td></tr><tr><td>Reusable needle- requires customization</td><td>Fisher Scientific</td><td>7803-02</td><td>Customize to 0.7 inches long; point style 4 and angle 25</td></tr><tr><td>T4 DNA polymerase</td><td>NEB</td><td>M0203L</td><td>Use with the 10x NEB buffer that is included</td></tr><tr><td>Teflon coated tools</td><td>bonefolder.com</td><td>T-SPATULA4PIECE</td><td>referred to as &quot;polytetrafluoroethene&quot; in the protocol</td></tr></tbody></table>

silencing the spark: crispr/cas9 genome editing in weakly electric fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

The sgRNA target sites were identified within exon 1 of scn4aa in both B. gauderio and B. brachyistius as described in Section 1. The sgRNAs were generated as described in Section 2. Following successful sgRNA selection and synthesis (Figure 1), in vitro cleavage was tested (Figure 2). The sgRNAs demonstrating in vitro cutting were then selected for single cell microinjections.
Adult fish were conditioned fo...

Watch this Scientific Journal Video about Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish at JoVE.com

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

silencing-the-spark-crisprcas9-genome-editing-in-weakly-electric-fish

Research

JoVE Journal

Biology

9.9K Views.  Michigan State University. CRISPR/Cas9 genbewerking bij zwak elektrische vissen, met name in een vergelijkende context, zal onderzoekers in staat stellen hypothesen te testen over hoe specifieke genen en genetische variantie bijdragen aan fenotypische variatie. CRISPR/Cas9 is een efficiënte methode voor het genereren van gemuteerde F-zero embryo's. Dit protocol stelt onderzoekers in staat om CRISPR-Cas9 manipulaties uit te voeren in twee convergent ontwikkelde soorten van zwak elektrische vissen.De sleutel tot ...

Video: Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

Genetics

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 ...

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

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 ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Developmental Biology

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, ...

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 ...

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal experiments. When generating animal models, the electroporation of zygotes offers higher efficiency, simplicity, cost, and throughput as an alternative to the gold standard method of microinjection. Electroporation is also gentler, with higher viability, and reliably delivers Cas9/single-guide RNA (sgRNA) ribonucleoproteins (RNPs) into the zygotes of common laboratory mouse strains (e.g., C57BL/6J and C57BL/6N) that approaches 100% delivery efficiency. This technique enables insertion/deletion (indels) mutations, point mutations, the deletion of whole genes or exons, and small insertions in the range of 100-200 bp to insert LoxP or short tags like FLAG, HA, or V5. While constantly being improved, here we present the current state of CRISPR-EZ in a protocol that includes sgRNA production through in vitro transcription, embryo processing, RNP assembly, electroporation, and the genotyping of preimplantation embryos. A graduate-level researcher with minimal experience manipulating embryos can obtain genetically edited embryos in less than 1 week using this protocol. Here, we offer a straightforward, low-cost, efficient, high-capacity method that could be used with mouse embryos.

Here, we describe a simple technique intended for the efficient generation of genetically modified mice called CRISPR RNP Electroporation of Zygotes (CRISPR-EZ). This method delivers editing reagents by electroporation into embryos at an efficiency approaching 100%. This protocol is effective for point mutations, small genomic insertions, and deletions in mammalian embryos.

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal ...

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases (TALENs)

Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs. Towards this end, much recent work has focused on identifying candidate genes for the evolution of traits in a variety of species. However, until recently it has been challenging to functionally validate interesting candidate genes. Recently developed tools for genetic engineering make it possible to manipulate specific genes in a wide range of organisms. Application of this technology in evolutionarily relevant organisms will allow for unprecedented insight into the role of candidate genes in evolution. Astyanax mexicanus (A. mexicanus) is a species of fish with both surface-dwelling and cave-dwelling forms. Multiple independent lines of cave-dwelling forms have evolved from ancestral surface fish, which are interfertile with one another and with surface fish, allowing elucidation of the genetic basis of cave traits. A. mexicanus has been used for a number of evolutionary studies, including linkage analysis to identify candidate genes responsible for a number of traits. Thus, A. mexicanus is an ideal system for the application of genome editing to test the role of candidate genes. Here we report a method for using transcription activator-like effector nucleases (TALENs) to mutate genes in surface A. mexicanus. Genome editing using TALENs in A. mexicanus has been utilized to generate mutations in pigmentation genes. This technique can also be utilized to evaluate the role of candidate genes for a number of other traits that have evolved in cave forms of A. mexicanus.

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases TALENs

Gene-targeting mutagenesis is now possible in a wide range of organisms using genome editing techniques. Here, we demonstrate a protocol for targeted gene mutagenesis using transcription activator like effector nucleases (TALENs) in Astyanax mexicanus, a species of fish that includes surface fish and cavefish.

Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs.  Towards this end, much recent ...

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

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

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

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

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

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