The authors would like to thank Nolan Davis for assistance with videography and video editing.

All experimental procedures were approved by the University of Missouri&#39;s Institutional Animal Care and Use Committee (ACUC protocol #25580) and were performed according to the guidelines set forth in the Guide for the Use and Care of Laboratory Animals.
1. AAV-mediated DNA repair template delivery
<ol>
	<li>Design the DNA construct to contain the desired knock-in sequence in between two homology arms. Typical homology arm lengths are 300-500 bp in length. Ensure that the entire DNA repair template (homology arms + desired knock-in sequence) is flanked by inverted terminal repeats (ITRs) by cloning into an appropriate plasmid backbone and packaging into recombinant ssAAV1 or ssAAV6 (serotypes 1 or 6). 
	NOTE: If the sgRNA target sequence is present in the DNA repair template, it is important to engineer silent mutations to disrupt this sequence thus preventing CRISPR-Cas9 mediated editing of the correctly targeted allele. 
	CAUTION: Recombinant AAV viral vectors are non-integrating and classified as BSL-1. Standard aseptic techniques are used for pipetting and handling embryos that have been exposed to virus.</li>
	<li>Add engineered ssAAV1 or ssAAV6 (packaged with DNA repair template) to 30 &#181;L of KSOM-R media27,28 to a final concentration of 3 &#215; 107 genome copies (GC)/&#181;L under mineral oil in 35 mm Petri dish.</li>
	<li>Place zygotes with visible pronuclei into the KSOM-R media containing engineered ssAAV. 
	NOTE: Using this protocol there is no need to thin the zona pellucida as AAV serotypes 1 and 6 are able to readily pass through for efficient DNA repair template delivery.</li>
	<li>Incubate at 37 &#176;C with 5% CO2 and maximal humidity for 18-20 h to allow for sufficient viral transduction.</li>
</ol>
2. Electroporation preparation
<ol>
	<li>On the following day, make an electroporation mixture containing 100 ng/&#181;L of sgRNA and 100 ng/&#181;L of Cas9 protein diluted in Opti-MEM media in a sterile RNase-free tube. Gently mix.</li>
	<li>Incubate at room temperature for 10 min to allow for RNP formation.</li>
	<li>During the incubation time, turn on the electroporator and connect leads to a 1.5 mm gap glass slide electrode.</li>
	<li>Set electroporation parameters as described in Table 1.</li>
</ol>
3. 2-cell embryo electroporation
<ol>
	<li>Gently pipette 5 &#181;L of the CRISPR electroporation mixture between the two electrodes on the glass slide taking care not to introduce air bubbles to the solution.</li>
	<li>Remove the 2-cell stage embryo from the AAV solution. Line up 2-cell stage embryos in the mixture without allowing them to touch the sides of the electrodes. This is done to avoid lysis.</li>
	<li>Press the Ohm button to measure the impedance. Check to ensure impedance is between 0.100 and 0.300 k&#8486;. The reaction volume will alter the impedance (add volume to decrease impedance and subtract volume to increase impedance).</li>
	<li>Press Start. The process takes few seconds to complete.</li>
	<li>Remove embryos immediately following the electroporation and wash 3 times in fresh 30 &#181;L drops of KSOM-R media.</li>
	<li>Place embryos back in 37 &#176;C incubator with 5% CO2 and maximal humidity for in vitro culture in 500 &#181;L drops of KSOM-R media under mineral oil. Alternatively, surgically transfer embryos to a 1.5-day post coitum (dpc) pseudopregnant female rat to produce live offspring.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66069/66069fig1.jpg" /> 
Figure 1: Schematic of AAV-mediated DNA delivery and 2-cell embryo electroporation pipeline for CRISPR-Cas9 genome editing. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66069/66069fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have no conflicts of interest to declare.

The CRISPR-Cas genome editing system has revolutionized the field of genetic engineering by allowing the efficient production of both straightforward and complex, customized genetic modifications in a variety of animal species. Frequent refinements and improvements in techniques associated with genome editing further its versatility. Here we have described a new approach using ssAAV-mediated DNA delivery along with 2-cell embryo electroporation of CRISPR-Cas9 reagents into 2-cell rodent embryos. We have demonstrated that...

The development of CRISPR-based genome editing tools has accelerated our ability to efficiently generate new and more sophisticated genetically engineered rat models. Single-guide RNA, along with Cas9 nuclease, is combined to form Ribonucleoprotein (RNP) complexes that target DNA sequences of interest within the genome and result in double-stranded DNA breaks. Because cellular DNA repair mechanisms are error-prone, insertions and deletions (INDELs) are introduced during the repair process that can disrupt a target gene&#39;s function. When there is co-delivery of a desired engineered DNA sequence (repair template) along with genome editing reagents, insertion of the repair template in the region containing the double-stranded DNA break occurs through a process called homology-directed repair (HDR). This is an effective strategy for generating animal models with targeted DNA insertions/substitutions (knock-ins). One limitation is that knock-in sequences are often large in size, which has been shown to reduce gene editing efficiency, thus making generating the desired model more difficult1. Strategies to increase knock-in efficiencies have included linearization of both double-stranded DNA (dsDNA) and single-stranded DNA (ssDNA) repair templates and chemical modification of DNA repair templates2,3,4. In addition, pronuclear microinjection along with HDR stimulating compounds, application of electrical pulses in conjunction with microinjection, and timed microinjection into 2-cell embryos have all been attempted5,6,7. Despite the success of some of these approaches, the incorporation of DNA sequences larger than 1.0 kb remains technically challenging.
Electroporation, which is a common method for introducing reagents into cultured cell lines, offers an alternative to microinjection for delivering CRISPR-Cas9 components into embryos. Embryo electroporation, first demonstrated in rat embryos8, has since been successfully used as a delivery method in mice9,10,11,12,13, pigs14,15, and other animal model organisms16,17,18. Embryos, suspended in the medium containing CRISPR-Cas9 reagents, are placed into a cuvette or onto a glass slide between two electrodes and subjected to direct pulses of electrical currents. This creates transient openings in the zona pellucida and embryo plasma membrane through which the CRISPR-Cas9 components enter the embryos. Typically, mid-level electrical &#34;poring&#34; pulses are used to create the temporary openings followed by lower level electrical &#34;transfer pulses&#34; that facilitate movement of the negatively charged genome editing components. Embryo electroporation is efficient, has a high throughput, and is easy to perform. However, while embryo electroporation has been shown to be highly successful for the introduction of small (&#60;200 bp) ssDNA repair templates, there are few reports of successful electroporation of larger (&#62;1.0 kb) repair templates13,19. This size restriction represents a major limitation of embryo electroporation for generating knock-in animal models requiring large insertions.
In the context of gene therapy, adeno-associated viruses (AAVs) have long been used as vehicles to deliver genetic material due to their efficient in vivo infectivity of both dividing and non-dividing cells, lack of pathogenicity, and rare genomic integration20,21. Recently, more studies have combined AAVs with CRISPR-Cas9 technology to introduce DNA repair templates and CRISPR reagents22,23,24. This approach allows delivery of larger DNA repair templates without the need for microinjection techniques.
The HDR pathway is more active in the late S and G2 phases of the cell cycle25,26. In studies performed in vitro, significant increases in knock-in efficiency were achieved by delivery of CRISPR-Cas9 RNPs and DNA repair templates into G2-synchronized cells or by restriction of the presence of Cas9 protein to late S and G2 phases using a Cas9-Geminin fusion protein2. Moreover, there is a major zygotic genome activation (ZGA) event which occurs during the extended G2 phase of the 2-cell stage embryo, and this is associated with an open chromatin state. It is speculated that this provides the CRISPR-Cas9 RNPs and repair templates with greater accessibility to the genomic DNA.
Our goal was to build on all these observations, by combining the AAV approach with embryo electroporation to introduce CRISPR-Cas9 RNPs at the 2-cell stage of embryo development. This strategy takes advantage of the larger DNA repair template delivery capacity of AAV, the technical ease of electroporation and the more optimal 2-cell time point for genomic accessibility during embryo development to create an efficient method for targeted genetic engineering of DNA insertions. As highlighted in this protocol, our optimized method allows for the production of targeted knock-in rat models carrying insertions of DNA sequences from 1.2 to 3.0 kb in size without the need for microinjection techniques.

<table><tbody><tr><td>Leads with alligator clips for electrodes</td><td>NepaGene</td><td>C117</td><td></td></tr><tr><td>Mineral oil</td><td>MilliporeSigma</td><td>M5310</td><td></td></tr><tr><td>NepaGene21 Super Electroporator&amp;nbsp;</td><td>NepaGene</td><td>NEPA21</td><td></td></tr><tr><td>Platinum plate electrodes on slide glass</td><td>NepaGene</td><td>CUY501P1-1.5</td><td></td></tr><tr><td>PURedit Cas9 Protein</td><td>MilliporeSigma</td><td>PECAS9</td><td></td></tr><tr><td>sgRNA (chemically-modified)</td><td>Synthego</td><td>N/A</td><td></td></tr><tr><td>ssAAV1 or ssAAV6 packaged DNA repair template</td><td>Vectorbuilder</td><td>N/A</td><td></td></tr></tbody></table>

crispr-cas9 genome editing of rat embryos using adeno-associated virus (aav) and 2-cell embryo electroporation

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair templates and CRISPR-Cas reagents is the most common delivery method into embryos. However, this type of micromanipulation necessitates access to specialized equipment, is laborious, and requires a certain level of technical skill. Moreover, microinjection techniques often result in lower embryo survival due to the mechanical stress on the embryo. In this protocol, we developed an optimized method to deliver large DNA repair templates to work in conjunction with CRISPR-Cas9 genome editing without the need for microinjection. This protocol combines AAV-mediated DNA delivery of single-stranded DNA donor templates along with the delivery of CRISPR-Cas9 ribonucleoprotein (RNP) by electroporation to modify 2-cell embryos. Using this novel strategy, we have successfully produced targeted knock-in rat models carrying insertion of DNA sequences from 1.2 to 3.0 kb in size with efficiencies between 42% and 90%.

Following the protocol, there is effective AAV-mediated delivery of the DNA repair template allowing for highly efficient HDR after 2-cell embryos are electroporated with CRISPR-Cas9 RNPs. As shown in the video, a successful electroporation process results in bubbles forming on each electrode (Figure 2C) and the impedance remaining within the range of 0.100 and 0.300 k&#8486;. After electroporation it is not uncommon for embryos to swell filling the perivitel...

Watch this Scientific Journal Video about CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus AAV and 2-Cell Embryo Electroporation at JoVE.com

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus AAV and 2-Cell Embryo Electroporation

This protocol presents an optimized approach for producing genetically modified rat models. Adeno-associated virus (AAV) is used to deliver a DNA repair template, and electroporation is used to deliver CRISPR-Cas9 reagents to complete the genome editing process in 2-cell embryo.

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair ...

crispr-cas9-genome-editing-rat-embryos-using-adeno-associated-virus

Universidad San Sebastian

Research

JoVE Journal

Genetics

1.7K Views.  University of Missouri. This protocol presents an optimized approach for producing genetically modified rat models. Adeno-associated virus (AAV) is used to deliver a DNA repair template, and electroporation is used to deliver CRISPR-Cas9 reagents to complete the genome editing process in 2-cell embryo.

Video: CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus AAV and 2-Cell Embryo Electroporation

In Utero Intraventricular Injection and Electroporation of E16 Rat Embryos

In-utero in-vivo injection and electroporation of the embryonic rat neocortex provides a powerful tool for the manipulation of individual progenitors lining the walls of the lateral ventricle. This technique is now widely used to study the processes involved in corticogenesis by over-expressing or knocking down genes and observing the effects on cellular proliferation, migration, and differentiation. In comparison to traditional knockout strategies, in-utero electroporation provides a rapid means to manipulate a population of cells during a specific temporal window. In this video protocol, we outline the experimental methodology for preparing rats for surgery, exposing the uterine horns through laporatomy, injecting DNA into the lateral ventricles of the developing embryo, electroporating DNA into the progenitors lining the lateral wall, and caring for animals post-surgery. Our laboratory uses this protocol for surgeries on E15-E21 rats, however it is most commonly performed at E16 as shown in this video.

In-utero in-vivo injection and electroporation of the embryonic rat neocortex provides a powerful tool for the manipulation of individual progenitors ...

Biology

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 ...

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 ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...

Generation of Topically Transgenic Rats by In utero Electroporation and In vivo Bioluminescence Screening

In utero electroporation (IUE) is a technique which allows genetic modification of cells in the brain for investigating neuronal development. So far, the use of IUE for investigating behavior or neuropathology in the adult brain has been limited by insufficient methods for monitoring of IUE transfection success by non-invasive techniques in postnatal animals. 
 For the present study, E16 rats were used for IUE. After intraventricular injection of the nucleic acids into the embryos, positioning of the tweezer electrodes was critical for targeting either the developing cortex or the hippocampus. 
 Ventricular co-injection and electroporation of a luciferase gene allowed monitoring of the transfected cells postnatally after intraperitoneal luciferin injection in the anesthetized live P7 pup by in vivo bioluminescence, using an IVIS Spectrum device with 3D quantification software. 
 Area definition by bioluminescence could clearly differentiate between cortical and hippocampal electroporations and detect a signal longitudinally over time up to 5 weeks after birth. This imaging technique allowed us to select pups with a sufficient number of transfected cells assumed necessary for triggering biological effects and, subsequently, to perform behavioral investigations at 3 month of age. As an example, this study demonstrates that IUE with the human full length DISC1 gene into the rat cortex led to amphetamine hypersensitivity. Co-transfected GFP could be detected in neurons by post mortem fluorescence microscopy in cryosections indicating gene expression present at &ge;6 months after birth.
 We conclude that postnatal bioluminescence imaging allows evaluating the success of transient transfections with IUE in rats. Investigations on the influence of topical gene manipulations during neurodevelopment on the adult brain and its connectivity are greatly facilitated. For many scientific questions, this technique can supplement or even replace the use of transgenic rats and provide a novel technology for behavioral neuroscience.

Generation of Topically Transgenic Rats by In utero Electroporation and In vivo Bioluminescence Screening

Genes can be manipulated during development of the cortex or the hippocampus of the rat via in utero electroporation (IUE) at E16, to enable fast and targeted modifications in neuronal connectivity for later studies of behavior or neuropathology in adult animals. Postnatal in vivo imaging for control of IUE success is performed by bioluminescence of activating co-transfected luciferase.

 In utero electroporation (IUE) is a technique which allows genetic modification of cells in the brain for investigating neuronal development. So far, the ...

Neuroscience

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (&#62;1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily ...

Developmental Biology

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the efficiency in developing gene-knockout mice by inducing small indel mutation would be sufficient enough, the efficiency of embryo genome editing for making large-size DNA knock-in (KI) is still low. Therefore, in contrast to the direct KI method in embryos, gene targeting using embryonic stem cells (ESCs) followed by embryo injection to develop chimera mice still has several advantages (e.g., high throughput targeting in vitro, multi-allele manipulation, and Cre and flox gene manipulation can be carried out in a short period). In addition, strains with difficult-to-handle embryos in vitro, such as BALB/c, can also be used for ESC targeting. This protocol describes the optimized method for large-size DNA (several kb) KI in ESCs by applying CRISPR/Cas9-mediated genome editing followed by chimera mice production to develop gene-manipulated mouse models.

Here we present a protocol for developing genetically modified mouse models using embryonic stem cells, especially for large DNA knock-in (KI). This protocol is tuned up using CRISPR/Cas9 genome editing, resulting in significantly improved KI efficiency compared with the conventional homologous recombination-mediated linearized DNA targeting method.

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the ...

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 ...

A Novel Strategy Combining Array-CGH, Whole-exome Sequencing and In Utero Electroporation in Rodents to Identify Causative Genes for Brain Malformations

Birth defects that involve the cerebral cortex &#8212; also known as malformations of cortical development (MCD) &#8212; are important causes of intellectual disability and account for 20-40% of drug-resistant epilepsy in childhood. High-resolution brain imaging has facilitated in vivo identification of a large group of MCD phenotypes. Despite the advances in brain imaging, genomic analysis and generation of animal models, a straightforward workflow to systematically prioritize candidate genes and to test functional effects of putative mutations is missing. To overcome this problem, an experimental strategy enabling the identification of novel causative genes for MCD was developed and validated. This strategy is based on identifying candidate genomic regions or genes via array-CGH or whole-exome sequencing and characterizing the effects of their inactivation or of overexpression of specific mutations in developing rodent brains via in utero electroporation. This approach led to the identification of the C6orf70 gene, encoding for a putative vesicular protein, to the pathogenesis of periventricular nodular heterotopia, a MCD caused by defective neuronal migration.

A Novel Strategy Combining Array-CGH, Whole-exome Sequencing and In Utero Electroporation in Rodents to Identify Causative Genes for Brain Malformations

Periventricular nodular heterotopia (PNH) is the most common form of malformation of cortical development (MCD) in adulthood but its genetic basis remains unknown in most sporadic cases. We have recently developed a strategy to identify novel candidate genes for MCDs and to directly confirm their causative role in vivo.

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation

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In Utero Intraventricular Injection and Electroporation of E16 Rat Embryos

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

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

Generation of Topically Transgenic Rats by In utero Electroporation and In vivo Bioluminescence Screening

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

A Novel Strategy Combining Array-CGH, Whole-exome Sequencing and In Utero Electroporation in Rodents to Identify Causative Genes for Brain Malformations