Electroporation of mice zygotes with dual guide RNA/Cas9 complexes for simple and efficient cloning-free genome editing

Generation of a C57BL/6J mouse strain expressing the CD45.1 epitope to improve hematopoietic stem cell engraftment and adoptive cell transfer experiments

Adoptive cell transfer between genetically identical hosts relies on the use of a congenic marker to distinguish the donor cells from the host cells. CD45, a glycoprotein expressed by all hematopoietic cells, is one of the main congenic markers used because its two isoforms, CD45.1 and CD45.2, can be discriminated by flow cytometry. As a consequence, C57BL/6J (B6; CD45.2) and B6.SJL-Ptprca Pepcb/BoyJ (B6.SJL; CD45.1) mice are widely used in adoptive cell transfer experiments, under the presumption that they differ only at the CD45 (Ptprc) locus. However, recent studies have identified genetic variations between these congenic strains and have notably highlighted a differential expression of cathepsin E (CTSE). The B6.SJL mouse presents a number of functional differences in hematopoietic stem cell engraftment potential and immune cell numbers compared with the B6 mouse. In this study, we showed that B6 and B6.SJL mice also differ in their CD8+ T cell compartment and CD8+ T cell responses to viral infection. We identified Ctse as the most differentially expressed gene between CD8+ T cells of B6 and B6.SJL and demonstrated that the differences reported between these two mouse strains are not due to CTSE. Finally, using CRISPR-Cas9 genome editing, we generated a CD45.1-expressing B6 mouse by inserting one nucleotide mutation (A904G) leading to an amino acid change (K302E) in the Ptprc gene of the B6 mouse. We showed that this new B6-Ptprcem(K302E)Jmar/J mouse resolves the experimental biases reported between the B6 and B6.SJL mouse lines and should thus represent the new gold standard for adoptive cell transfer experiments in B6.

Efficient DNA knock-in using AAV-mediated delivery with 2-cell embryo CRISPR-Cas9 electroporation

Recent advances in CRISPR-Cas genome editing technology have been instrumental in improving the efficiency to produce genetically modified animal models. In this study we have combined four very promising approaches to come up with a highly effective pipeline to produce knock-in mouse and rat models. The four combined methods include: AAV-mediated DNA delivery, single-stranded DNA donor templates, 2-cell embryo modification, and CRISPR-Cas ribonucleoprotein (RNP) electroporation. Using this new combined approach, we were able to produce successfully targeted knock-in rat models containing either Cre or Flp recombinase sequences with knock-in efficiencies over 90%. Furthermore, we were able to produce a knock-in mouse model containing a Cre recombinase targeted insertion with over 50% knock-in efficiency directly comparing efficiencies to other commonly used approaches. Our modified AAV-mediated DNA delivery with 2-cell embryo CRISPR-Cas9 RNP electroporation technique has proven to be highly effective for generating both knock-in mouse and knock-in rat models.

Perspectives in Genome-Editing Techniques for Livestock

Genome editing of farm animals has undeniable practical applications. It helps to improve production traits, enhances the economic value of livestock, and increases disease resistance. Gene-modified animals are also used for biomedical research and drug production and demonstrate the potential to be used as xenograft donors for humans. The recent discovery of site-specific nucleases that allow precision genome editing of a single-cell embryo (or embryonic stem cells) and the development of new embryological delivery manipulations have revolutionized the transgenesis field. These relatively new approaches have already proven to be efficient and reliable for genome engineering and have wide potential for use in agriculture. A number of advanced methodologies have been tested in laboratory models and might be considered for application in livestock animals. At the same time, these methods must meet the requirements of safety, efficiency and availability of their application for a wide range of farm animals. This review aims at covering a brief history of livestock animal genome engineering and outlines possible future directions to design optimal and cost-effective tools for transgenesis in farm species.

Gait Abnormalities and Aberrant D2 Receptor Expression and Signaling in Mice Carrying the Human Pathogenic Mutation DRD2I212F

A dopamine D2 receptor mutation was recently identified in a family with a novel hyperkinetic movement disorder. That allelic variant D2-I212F is a constitutively active and G protein-biased receptor. We now describe mice engineered using CRISPR-Cas9-mediated gene editing technology to carry the D2-I212F variant. Drd2I212F mice exhibited gait abnormalities resembling those in other mouse models of chorea and/or dystonia and had striatal D2 receptor expression that was decreased approximately 30% per Drd2I212F allele. Electrically evoked inhibitory postsynaptic conductances in midbrain dopamine neurons and striatum from Drd2I212F mice, caused by G protein activation of potassium channels, exhibited slow kinetics (e.g., approximately four- to sixfold slower decay) compared with Drd2 +/+ mice. Current decay initiated by photolytic release of the D2 antagonist sulpiride from CyHQ-sulpiride was also ∼fourfold slower in midbrain slices from Drd2I212F mice than Drd2 +/+ mice. Furthermore, in contrast to Drd2 +/+ mice, in which dopamine is several-fold more potent at neurons in the nucleus accumbens than in the dorsal striatum, reflecting activation of Gα o versus Gα i, dopamine had similar potencies in those two brain regions of Drd2I212F mice. Repeated cocaine treatment, which decreases dopamine potency in the nucleus accumbens of Drd2 +/+ mice, had no effect on dopamine potency in Drd2 I212F mice. The results demonstrate the pathogenicity of the D2-I212F mutation and the utility of this mouse model for investigating the role of pathogenic DRD2 variants in early-onset hyperkinetic movement disorders. SIGNIFICANCE STATEMENT: The first dopamine receptor mutation to cause a movement disorder, D2-I212F, was recently identified. The mutation makes receptor activation of G protein-mediated signaling more efficient. To confirm the pathogenesis of D2-I212F, this study reports that mice carrying this mutation have gait abnormalities consistent with the clinical phenotype. The mutation also profoundly alters D2 receptor expression and function in vivo. This mouse model will be useful for further characterization of the mutant receptor and for evaluation of potential therapeutic drugs.

Pigs with an INS point mutation derived from zygotes electroporated with CRISPR/Cas9 and ssODN

Just one amino acid at the carboxy-terminus of the B chain distinguishes human insulin from porcine insulin. By introducing a precise point mutation into the porcine insulin (INS) gene, we were able to generate genetically modified pigs that secreted human insulin; these pigs may be suitable donors for islet xenotransplantation. The electroporation of the CRISPR/Cas9 gene-editing system into zygotes is frequently used to establish genetically modified rodents, as it requires less time and no micromanipulation. However, electroporation has not been used to generate point-mutated pigs yet. In the present study, we introduced a point mutation into porcine zygotes via electroporation using the CRISPR/Cas9 system to generate INS point-mutated pigs as suitable islet donors. We first optimized the efficiency of introducing point mutations by evaluating the effect of Scr7 and the homology arm length of ssODN on improving homology-directed repair-mediated gene modification. Subsequently, we prepared electroporated zygotes under optimized conditions and transferred them to recipient gilts. Two recipients became pregnant and delivered five piglets. Three of the five piglets carried only the biallelic frame-shift mutation in the INS gene, whereas the other two successfully carried the desired point mutation. One of the two pigs mated with a WT boar, and this desired point mutation was successfully inherited in the next F1 generation. In conclusion, we successfully established genetically engineered pigs with the desired point mutation via electroporation-mediated introduction of the CRISPR/Cas9 system into zygotes, thereby avoiding the time-consuming and complicated micromanipulation method.

CRISMERE Chromosome Engineering in Mouse and Rat

CRISPR/Cas9 technology is a versatile tool for engineering biology that has dramatically transformed our ability to manipulate genomes. In this protocol, we use its capacity to generate two double-strand breaks simultaneously, at precise positions in the genome, to generate mouse or rat lines with deletion, inversion, and duplication of a specific genomic segment. The technic is called CRISMERE for CRISpr-MEdiated REarrangement. This protocol describes the different steps to generate and validate the different chromosomal rearrangements that can be obtained with the technology. These new genetic configurations can be useful to model rare diseases with copy number variation, understand the genomic organization, or provide genetic tools (like balancer chromosome) to keep lethal mutations.

Electroporation-Mediated CRISPR/Cas9 Genome Editing in Rat Zygotes

Genetic engineering in the rat has been revolutionized by the development of CRISPR-based genome editing tools. Conventional methods for inserting genome editing elements such as CRISPR/Cas9 reagents into rat zygotes include cytoplasmic or pronuclear microinjections. These techniques are labor-intensive, require specialized micromanipulator equipment, and are technically challenging. Here, we describe a simple and effective method for zygote electroporation in which CRISPR/Cas9 reagents are introduced into rat zygotes via pores produced by precise electrical pulses applied to the cells. Zygote electroporation allows for high-throughput efficient genome editing in rat embryos.

JUN Regulation of Injury-Induced Enhancers in Schwann Cells

Schwann cells play a critical role after peripheral nerve injury by clearing myelin debris, forming axon-guiding bands of Büngner, and remyelinating regenerating axons. Schwann cells undergo epigenomic remodeling to differentiate into a repair state that expresses unique genes, some of which are not expressed at other stages of Schwann cell development. We previously identified a set of enhancers that are activated in Schwann cells after nerve injury, and we determined whether these enhancers are preprogrammed into the Schwann cell epigenome as poised enhancers before injury. Poised enhancers share many attributes of active enhancers, such as open chromatin, but are marked by repressive histone H3 lysine 27 (H3K27) trimethylation rather than H3K27 acetylation. We find that most injury-induced enhancers are not marked as poised enhancers before injury indicating that injury-induced enhancers are not preprogrammed in the Schwann cell epigenome. Injury-induced enhancers are enriched with AP-1 binding motifs, and the c-JUN subunit of AP-1 had been shown to be critical to drive the transcriptional response of Schwann cells after injury. Using in vivo chromatin immunoprecipitation sequencing analysis in rat, we find that c-JUN binds to a subset of injury-induced enhancers. To test the role of specific injury-induced enhancers, we focused on c-JUN-binding enhancers upstream of the Sonic hedgehog (Shh) gene, which is only upregulated in repair Schwann cells compared with other stages of Schwann cell development. We used targeted deletions in male/female mice to show that the enhancers are required for robust induction of the Shh gene after injury.SIGNIFICANCE STATEMENT The proregenerative actions of Schwann cells after nerve injury depends on profound reprogramming of the epigenome. The repair state is directed by injury-induced transcription factors, like JUN, which is uniquely required after nerve injury. In this study, we test whether the injury program is preprogrammed into the epigenome as poised enhancers and define which enhancers bind JUN. Finally, we test the roles of these enhancers by performing clustered regularly interspaced short palindromic repeat (CRISPR)-mediated deletion of JUN-bound injury enhancers in the Sonic hedgehog gene. Although many long-range enhancers drive expression of Sonic hedgehog at different developmental stages of specific tissues, these studies identify an entirely new set of enhancers that are required for Sonic hedgehog induction in Schwann cells after injury.

H3K27me3 at pericentromeric heterochromatin is a defining feature of the early mouse blastocyst

Early mouse development is characterized by structural and epigenetic changes while cells progress towards differentiation. At blastocyst stage, the segregation of the three primordial lineages is accompanied by establishment of differential patterns of DNA methylation and post-translational modifications of histones, such as H3K27me3. Here, we analysed the dynamics of H3K27me3 at pericentromeric heterochromatin (PCH) during early development. We also followed the localization of EZH2 and BEND3, previously shown in ESCs to drive PRC2 to hypomethylated PCH. We show that the location of H3K27me3 at PCH, in addition to H3K9me3, is a defining feature of embryonic cells in vivo. Moreover, it may play an important role in structuring PCH and preserving genomic integrity at a time of globally relaxed chromatin. At peri-implantation stages, while DNA methylation is still low, EZH2 and then H3K27me3, leave PCH in epiblast progenitors at the time of their spatial segregation from primitive endoderm cells, while BEND3 remains there up to implantation. The comparison with stem cells (ESCs and TSCs) reveals that the epigenetic marks (i.e. H3K9me3 and H3K27me3) of PCH are reset during in vitro derivation and only partially restored thereafter. This highlights possible divergences between in vitro and "in embryo" epigenetic regulation regarding constitutive heterochromatin.

The cell cycle stage of bovine zygotes electroporated with CRISPR/Cas9-RNP affects frequency of Loss-of-heterozygosity editing events

At the embryonic level, CRISPR technologies have been used to edit genomes reliably and efficiently in various mammalian models, with Ribonucleoprotein (RNP) electroporation potentially representing a superior delivery method into mammalian zygotes. However, detailed insights of the interactions between varying technical settings as well as the time point of electroporation in a bovine zygote's cell cycle on developmental metrics and the frequency and type of editing events are largely unknown. The present study uncovers that increasing pulse lengths result in higher Full Edit rates, with Mosaicism in Full-Edit embryos being significantly affected by adjusting RNP-electroporation relative to zygote cell cycle. A considerable proportion of Full Edit embryos demonstrated loss-of-heterozygosity after RNP-electroporation prior to S-phase. Some of these loss-of-heterozygosity events are a consequence of chromosomal disruptions along large sections of the target chromosomes making it necessary to check for their presence prior use of this technique in animal breeding. One out of 2 of these loss-of-heterozygosity events, however, was not associated with loss of an entire chromosome or chromosomal sections. Whether analysed loss-of-heterozygosity in these cases, however, was a false negative result due to loss of PCR primer sequences after INDEL formation at the target side or indeed due to interhomolog recombination needs to be clarified in follow up studies since the latter would for sure offer attractive options for future breeding schedules.

UVB-Induced Skin Autoinflammation Due to Nlrp1b Mutation and Its Inhibition by Anti-IL-1β Antibody

NLRP1 (NACHT and leucine-rich repeat-containing protein family, pyrin domain-containing protein 1) is an innate immune sensor that is involved in the formation of inflammasome complexes. NLRP1 hyperactivity has been reported to cause inherited autoinflammatory diseases including familial keratosis lichenoides chronica and NLRP1-associated autoinflammation with arthritis and dyskeratosis. We generated Nlrp1b (the mouse homologue of human NLRP1) gain-of-function knock-in (Nlrp1b KI) mice with UVB irradiation-induced autoinflammatory skin lesions. We demonstrated that UVB irradiation induces IL-1β upregulation and IL-1β-dependent inflammation via caspase-1 activation in these Nlrp1b KI mice. RNA sequencing revealed the upregulation of inflammasome pathway-related genes, keratinocyte stress marker genes, and keratinocyte differentiation marker genes in the Nlrp1b KI mice after UVB irradiation. The skin inflammation and hyperkeratosis from UVB irradiation in the Nlrp1b KI mice were inhibited by both intraperitoneal and subcutaneous administration of anti-IL-1β antibodies before UVB irradiation. UVB irradiation and the IL-1β pathway are important in the pathogenesis of NLRP1-associated autoinflammatory skin lesions.

Efficient and Fast Generation of Relevant Disease Mouse Models by In Vitro and In Vivo Gene Editing of Zygotes

Knockout mice for human disease-causing genes provide valuable models in which new therapeutic approaches can be tested. Electroporation of genome editing tools into zygotes, in vitro or within oviducts, allows for the generation of targeted mutations in a shorter time. We have generated mouse models deficient in genes involved in metabolic rare diseases (Primary Hyperoxaluria Type 1 Pyruvate Kinase Deficiency) or in a tumor suppressor gene (Rasa1). Pairs of guide RNAs were designed to generate controlled deletions that led to the absence of protein. In vitro or in vivo ribonucleoprotein (RNP) electroporation rendered more than 90% and 30% edited newborn animals, respectively. Mice lines with edited alleles were established and disease hallmarks have been verified in the three models that showed a high consistency of results and validating RNP electroporation into zygotes as an efficient technique for disease modeling without the need to outsource to external facilities.

An Overview of Cell Membrane Perforation and Resealing Mechanisms for Localized Drug Delivery

Localized and reversible plasma membrane disruption is a promising technique employed for the targeted deposition of exogenous therapeutic compounds for the treatment of disease. Indeed, the plasma membrane represents a significant barrier to successful delivery, and various physical methods using light, sound, and electrical energy have been developed to generate cell membrane perforations to circumvent this issue. To restore homeostasis and preserve viability, localized cellular repair mechanisms are subsequently triggered to initiate a rapid restoration of plasma membrane integrity. Here, we summarize the known emergency membrane repair responses, detailing the salient membrane sealing proteins as well as the underlying cytoskeletal remodeling that follows the physical induction of a localized plasma membrane pore, and we present an overview of potential modulation strategies that may improve targeted drug delivery approaches.

From genotype to phenotype: genetics of mammalian long non-coding RNAs in vivo

Genome-wide sequencing has led to the discovery of thousands of long non-coding RNA (lncRNA) loci in the human genome, but evidence of functional significance has remained controversial for many lncRNAs. Genetically engineered model organisms are considered the gold standard for linking genotype to phenotype. Recent advances in CRISPR-Cas genome editing have led to a rapid increase in the use of mouse models to more readily survey lncRNAs for functional significance. Here, we review strategies to investigate the physiological relevance of lncRNA loci by highlighting studies that have used genetic mouse models to reveal key in vivo roles for lncRNAs, from fertility to brain development. We illustrate how an investigative approach, starting with whole-gene deletion followed by transcription termination and/or transgene rescue strategies, can provide definitive evidence for the in vivo function of mammalian lncRNAs.

Recent Advances in the Production of Genome-Edited Rats

The rat is an important animal model for understanding gene function and developing human disease models. Knocking out a gene function in rats was difficult until recently, when a series of genome editing (GE) technologies, including zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and the type II bacterial clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated Cas9 (CRISPR/Cas9) systems were successfully applied for gene modification (as exemplified by gene-specific knockout and knock-in) in the endogenous target genes of various organisms including rats. Owing to its simple application for gene modification and its ease of use, the CRISPR/Cas9 system is now commonly used worldwide. The most important aspect of this process is the selection of the method used to deliver GE components to rat embryos. In earlier stages, the microinjection (MI) of GE components into the cytoplasm and/or nuclei of a zygote was frequently employed. However, this method is associated with the use of an expensive manipulator system, the skills required to operate it, and the egg transfer (ET) of MI-treated embryos to recipient females for further development. In vitro electroporation (EP) of zygotes is next recognized as a simple and rapid method to introduce GE components to produce GE animals. Furthermore, in vitro transduction of rat embryos with adeno-associated viruses is potentially effective for obtaining GE rats. However, these two approaches also require ET. The use of gene-engineered embryonic stem cells or spermatogonial stem cells appears to be of interest to obtain GE rats; however, the procedure itself is difficult and laborious. Genome-editing via oviductal nucleic acids delivery (GONAD) (or improved GONAD (i-GONAD)) is a novel method allowing for the in situ production of GE zygotes existing within the oviductal lumen. This can be performed by the simple intraoviductal injection of GE components and subsequent in vivo EP toward the injected oviducts and does not require ET. In this review, we describe the development of various approaches for producing GE rats together with an assessment of their technical advantages and limitations, and present new GE-related technologies and current achievements using those rats in relation to human diseases.

Autoinflammatory Keratinization Disease With Hepatitis and Autism Reveals Roles for JAK1 Kinase Hyperactivity in Autoinflammation

Heterozygous mutations in JAK1 which result in JAK-STAT hyperactivity have been implicated in an autosomal dominant disorder that features multi-organ immune dysregulation. This study identifies another previously unreported heterozygous missense JAK1 mutation, H596D, in an individual with a unique autoinflammatory keratinization disease associated with early-onset liver dysfunction and autism. Using CRISPR-Cas9 gene targeting, we generated mice with an identical Jak1 knock-in missense mutation (Jak1^{H595D/+;I596I/+;Y597Y/+} mice) that recapitulated key aspects of the human phenotype. RNA sequencing of samples isolated from the Jak1^{H595D/+;I596I/+;Y597Y/+} mice revealed the upregulation of genes associated with the hyperactivation of tyrosine kinases and NF-κB signaling. Interestingly, there was a strong correlation between genes downregulated in Jak1^{H595D/+;I596I/+;Y597Y/+} mice and those downregulated in the brain of model mice with 22q11.2 deletion syndrome that showed cognitive and behavioral deficits, such as autism spectrum disorders. Our findings expand the phenotypic spectrum of JAK1-associated disease and underscore how JAK1 dysfunction contributes to this autoinflammatory disorder.

Highly reliable creation of floxed alleles by electroporating single-cell embryos

BACKGROUND

Floxed (flanked by loxP) alleles are a crucial portion of conditional knockout mouse models. However, an efficient and reliable strategy to flox genomic regions of any desired size is still lacking.

RESULTS

Here, we demonstrate that the method combining electroporation of fertilized eggs with gRNA/Cas9 complexes and single-stranded oligodeoxynucleotides (ssODNs), assessing phasing of loxP insertions in founders using an in vitro Cre assay and an optional, highly specific and efficient second-round targeting ensures the generation of floxed F1 animals in roughly five months for a wide range of sequence lengths (448 bp to 160 kb reported here).

CONCLUSIONS

Floxed alleles can be reliably obtained in a predictable timeline using the improved method of electroporation of two gRNA/Cas9 ribonucleoprotein particles (RNPs) and two ssODNs.

ChREBPβ is dispensable for the control of glucose homeostasis and energy balance

Impaired glucose metabolism is observed in obesity and type 2 diabetes. Glucose controls gene expression through the transcription factor ChREBP in liver and adipose tissues. Mlxipl encodes 2 isoforms: ChREBPα, the full-length form (translocation into the nucleus is under the control of glucose), and ChREBPβ, a constitutively nuclear shorter form. ChREBPβ gene expression in white adipose tissue is strongly associated with insulin sensitivity. Here, we investigated the consequences of ChREBPβ deficiency on insulin action and energy balance. ChREBPβ-deficient male and female C57BL6/J and FVB/N mice were produced using CRISPR/Cas9-mediated gene editing. Unlike global ChREBP deficiency, lack of ChREBPβ showed modest effects on gene expression in adipose tissues and the liver, with variations chiefly observed in brown adipose tissue. In mice fed chow and 2 types of high-fat diets, lack of ChREBPβ had moderate effects on body composition and insulin sensitivity. At thermoneutrality, ChREBPβ deficiency did not prevent the whitening of brown adipose tissue previously reported in total ChREBP-KO mice. These findings revealed that ChREBPβ is dispensable for metabolic adaptations to nutritional and thermic challenges.

NLRP3 phosphorylation in its LRR domain critically regulates inflammasome assembly

NLRP3 controls the secretion of inflammatory cytokines IL-1β/18 and pyroptosis by assembling the inflammasome. Upon coordinated priming and activation stimuli, NLRP3 recruits NEK7 within hetero-oligomers that nucleate ASC and caspase-1 filaments, but the apical molecular mechanisms underlying inflammasome assembly remain elusive. Here we show that NEK7 recruitment to NLRP3 is controlled by the phosphorylation status of NLRP3 S803 located within the interaction surface, in which NLRP3 S803 is phosphorylated upon priming and later dephosphorylated upon activation. Phosphomimetic substitutions of S803 abolish NEK7 recruitment and inflammasome activity in macrophages in vitro and in vivo. In addition, NLRP3-NEK7 binding is also essential for NLRP3 deubiquitination by BRCC3 and subsequently inflammasome assembly, with NLRP3 phosphomimetic mutants showing enhanced ubiquitination and degradation than wildtype NLRP3. Finally, we identify CSNK1A1 as the kinase targeting NLRP3 S803. Our findings thus reveal NLRP3 S803 phosphorylation status as a druggable apical molecular mechanism controlling inflammasome assembly.

Nlrp3 inflammasome activation requires Nek7 recruitment to drive ASC speck formation. Here the authors show how Nlrp3 phosphorylation events control this Nek7 recruitment.

Modeling a human CLP1 mutation in mouse identifies an accumulation of tyrosine pre-tRNA fragments causing pontocerebellar hypoplasia type 10

Cleavage factor polyribonucleotide kinase subunit 1 (CLP1), an RNA kinase, plays essential roles in protein complexes involved in the 3'-end formation and polyadenylation of mRNA and the tRNA splicing endonuclease complex, which is involved in precursor tRNA splicing. The mutation R140H in human CLP1 causes pontocerebellar hypoplasia type 10 (PCH10), which is characterized by microcephaly and axonal peripheral neuropathy. Previously, we reported that RNA fragments derived from isoleucine pre-tRNA introns (Ile-introns) accumulate in fibroblasts of patients with PCH10. Therefore, it has been suggested that this intronic RNA fragment accumulation may trigger PCH10 onset. However, the molecular mechanism underlying PCH10 pathogenesis remains elusive. Thus, we generated knock-in mutant mice that harbored a CLP1 mutation consistent with R140H. As expected, these mice showed progressive loss of the upper motor neurons, resulting in impaired locomotor activity, although the phenotype was milder than that of the human variant. Mechanistically, we found that the R140H mutation causes intracellular accumulation of Ile-introns derived from isoleucine pre-tRNAs and 5' tRNA fragments derived from tyrosine pre-tRNAs, suggesting that these two types of RNA fragments were cooperatively or independently involved in the onset and progression of the disease. Taken together, the CLP1-R140H mouse model provided new insights into the pathogenesis of neurodegenerative diseases, such as PCH10, caused by genetic mutations in tRNA metabolism-related molecules.

Advances in Genome Editing and Application to the Generation of Genetically Modified Rat Models

The rat has been extensively used as a small animal model. Many genetically engineered rat models have emerged in the last two decades, and the advent of gene-specific nucleases has accelerated their generation in recent years. This review covers the techniques and advances used to generate genetically engineered rat lines and their application to the development of rat models more broadly, such as conditional knockouts and reporter gene strains. In addition, genome-editing techniques that remain to be explored in the rat are discussed. The review also focuses more particularly on two areas in which extensive work has been done: human genetic diseases and immune system analysis. Models are thoroughly described in these two areas and highlight the competitive advantages of rat models over available corresponding mouse versions. The objective of this review is to provide a comprehensive description of the advantages and potential of rat models for addressing specific scientific questions and to characterize the best genome-engineering tools for developing new projects.

Electroporation-Mediated Genome Editing of Livestock Zygotes

The introduction of genome editing reagents into mammalian zygotes has traditionally been accomplished by cytoplasmic or pronuclear microinjection. This time-consuming procedure requires expensive equipment and a high level of skill. Electroporation of zygotes offers a simplified and more streamlined approach to transfect mammalian zygotes. There are a number of studies examining the parameters used in electroporation of mouse and rat zygotes. Here, we review the electroporation conditions, timing, and success rates that have been reported for mice and rats, in addition to the few reports about livestock zygotes, specifically pigs and cattle. The introduction of editing reagents at, or soon after, fertilization can help reduce the rate of mosaicism, the presence of two of more genotypes in the cells of an individual; as can the introduction of nuclease proteins rather than mRNA encoding nucleases. Mosaicism is particularly problematic in large livestock species with long generation intervals as it can take years to obtain non-mosaic, homozygous offspring through breeding. Gene knockouts accomplished via the non-homologous end joining pathway have been more widely reported and successfully accomplished using electroporation than have gene knock-ins. Delivering large DNA plasmids into the zygote is hindered by the zona pellucida (ZP), and the majority of gene knock-ins accomplished by electroporation have been using short single stranded DNA (ssDNA) repair templates, typically less than 1 kb. The most promising approach to deliver larger donor repair templates of up to 4.9 kb along with genome editing reagents into zygotes, without using cytoplasmic injection, is to use recombinant adeno-associated viruses (rAAVs) in combination with electroporation. However, similar to other methods used to deliver clustered regularly interspaced palindromic repeat (CRISPR) genome-editing reagents, this approach is also associated with high levels of mosaicism. Recent developments complementing germline ablated individuals with edited germline-competent cells offer an approach to avoid mosaicism in the germline of genome edited founder lines. Even with electroporation-mediated delivery of genome editing reagents to mammalian zygotes, there remain additional chokepoints in the genome editing pipeline that currently hinder the scalable production of non-mosaic genome edited livestock.

Targeted Transgenic Mice Using CRISPR /Cas9 Technology

CRISPR /Cas9 is a powerful technology that has transformed gene editing of mammalian genomes, being faster and more cost-effective than standard gene targeting techniques. In this chapter, we provide a step-by-step protocol to obtain Knock-Out (KO ) or Knock-In (KI ) mouse models using CRISPR /Cas9 technology. Detailed instructions for the design of single guide RNAs (sgRNA ) for KO approaches and single-strand oligonucleotide (ssODN ) matrix for generation of KI animals are included. We also describe two independent CRISPR /Cas9 delivery methods to produce gene-edited animals starting from zygote-stage embryos, based either on cytoplasmic injection or electroporation.

Harnessing CRISPR-Cas system diversity for gene editing technologies

The discovery and utilization of RNA-guided surveillance complexes, such as CRISPR-Cas9, for sequence-specific DNA or RNA cleavage, has revolutionised the process of gene modification or knockdown. To optimise the use of this technology, an exploratory race has ensued to discover or develop new RNA-guided endonucleases with the most flexible sequence targeting requirements, coupled with high cleavage efficacy and specificity. Here we review the constraints of existing gene editing and assess the merits of exploiting the diversity of CRISPR-Cas effectors as a methodology for surmounting these limitations.

Generation of mouse conditional knockout alleles in one step using the i-GONAD method

The Cre/loxP system is a powerful tool for gene function study in vivo. Regulated expression of Cre recombinase mediates precise deletion of genetic elements in a spatially– and temporally–controlled manner. Despite the robustness of this system, it requires a great amount of effort to create a conditional knockout model for each individual gene of interest where two loxP sites must be simultaneously inserted in cis. The current undertaking involves labor-intensive embryonic stem (ES) cell–based gene targeting and tedious micromanipulations of mouse embryos. The complexity of this workflow poses formidable technical challenges, thus limiting wider applications of conditional genetics. Here, we report an alternative approach to generate mouse loxP alleles by integrating a unique design of CRISPR donor with the new oviduct electroporation technique i-GONAD. Showing the potential and simplicity of this method, we created floxed alleles for five genes in one attempt with relatively low costs and a minimal equipment setup. In addition to the conditional alleles, constitutive knockout alleles were also obtained as byproducts of these experiments. Therefore, the wider applications of i-GONAD may promote gene function studies using novel murine models.

Electroporation and genetic supply of Cas9 increase the generation efficiency of CRISPR/Cas9 knock-in alleles in C57BL/6J mouse zygotes

CRISPR/Cas9 machinery delivered as ribonucleoprotein (RNP) to the zygote has become a standard tool for the development of genetically modified mouse models. In recent years, a number of reports have demonstrated the effective delivery of CRISPR/Cas9 machinery via zygote electroporation as an alternative to the conventional delivery method of microinjection. In this study, we have performed side-by-side comparisons of the two RNP delivery methods across multiple gene loci and conclude that electroporation compares very favourably with conventional pronuclear microinjection, and report an improvement in mutagenesis efficiency when delivering CRISPR via electroporation for the generation of simple knock-in alleles using single-stranded oligodeoxynucleotide (ssODN) repair templates. In addition, we show that the efficiency of knock-in mutagenesis can be further increased by electroporation of embryos derived from Cas9-expressing donor females. The maternal supply of Cas9 to the zygote avoids the necessity to deliver the relatively large Cas9 protein, and high efficiency generation of both indel and knock-in allele can be achieved by electroporation of small single-guide RNAs and ssODN repair templates alone. Furthermore, electroporation, compared to microinjection, results in a higher rate of embryo survival and development. The method thus has the potential to reduce the number of animals used in the production of genetically modified mouse models.

Production of genome-edited mice by visualization of nucleases introduced into the embryos using electroporation

Genome editing technology contributes to the quick and highly efficient production of genetically engineered animals. These animals are helpful in clarifying the mechanism of human disease. Recently, a new electroporation technique (TAKE: Technique for animal knockout system by electroporation) was developed to produce genome-edited animals by introducing nucleases into intact embryos using electroporation instead of the microinjection method. The aim of this study was to increase the efficiency of production of genome-edited animals using the TAKE method. In the conventional protocol, it was difficult to confirm the introduction of nucleases into embryos and energization during operation. Using only embryos that introduced nucleases for embryo transfer, it will lead to increased efficiency in the production of genome-edited animals. This study examined the visualization in the introduction of nucleases into the embryos by using nucleases fluorescent labeled with ATTO-550. The embryos were transfected with Cas9 protein and fluorescent labeled dual guide RNA (mixture with crRNA and tracrRNA with ATTO-550) targeted tyrosinase gene by the TAKE method. All embryos that survived after electroporation showed fluorescence. Of these embryos with fluorescence, 43.7% developed to morphologically normal offspring. In addition, 91.7% of offspring were edited by the tyrosinase gene. This study is the first to demonstrate that the introduction of nucleases into embryos by the TAKE method could be visualized using fluorescent-labeled nucleases. This improved TAKE method can be used to produce genome-edited animals and confirm energization during operation.

Transgenic Expression and Genome Editing by Electroporation of Zebrafish Embryos

Microinjection is predominantly used to produce genetically modified fish. However, there are certain difficulties involved in some fish species. In this study, we tested an alternative method to produce genetically modified zebrafish by delivering DNA and other materials into embryos by electroporation. We optimized the electroporation conditions of a square wave electroporation system that work efficiently for the introduction of plasmid DNA, recombinant Cas9 nuclease and synthetic dual guide RNAs. Transgenic expression was observed in a wide range of tissues, which is comparable with those obtained by microinjection. We further determined that efficient gene delivery can be achieved during the cleavage stage. This study describes detailed electroporation parameters for gene delivery with high efficiency and low toxicity, providing a novel method to generate transgenic lines and genome editing.

SDR9C7 catalyzes critical dehydrogenation of acylceramides for skin barrier formation

The corneocyte lipid envelope, composed of covalently bound ceramides and fatty acids, is important to the integrity of the permeability barrier in the stratum corneum, and its absence is a prime structural defect in various skin diseases associated with defective skin barrier function. SDR9C7 encodes a short-chain dehydrogenase/reductase family 9C member 7 (SDR9C7) recently found mutated in ichthyosis. In a patient with SDR9C7 mutation and a mouse Sdr9c7-KO model, we show loss of covalent binding of epidermal ceramides to protein, a structural fault in the barrier. For reasons unresolved, protein binding requires lipoxygenase-catalyzed transformations of linoleic acid (18:2) esterified in ω-O-acylceramides. In Sdr9c7-/- epidermis, quantitative liquid chromatography-mass spectometry (LC-MS) assays revealed almost complete loss of a species of ω-O-acylceramide esterified with linoleate-9,10-trans-epoxy-11E-13-ketone; other acylceramides related to the lipoxygenase pathway were in higher abundance. Recombinant SDR9C7 catalyzed NAD+-dependent dehydrogenation of linoleate 9,10-trans-epoxy-11E-13-alcohol to the corresponding 13-ketone, while ichthyosis mutants were inactive. We propose, therefore, that the critical requirement for lipoxygenases and SDR9C7 is in producing acylceramide containing the 9,10-epoxy-11E-13-ketone, a reactive moiety known for its nonenzymatic coupling to protein. This suggests a mechanism for coupling of ceramide to protein and provides important insights into skin barrier formation and pathogenesis.

Efficient One-Step Knockout by Electroporation of Ribonucleoproteins Into Zona-Intact Bovine Embryos

Somatic cell nuclear transfer or cytoplasm microinjection have been used to generate genome-edited farm animals; however, these methods have several drawbacks that reduce their efficiency. This study aimed to develop electroporation conditions that allow delivery of CRISPR/Cas9 system to bovine zygotes for efficient gene knock-out. We optimized electroporation conditions to deliver Cas9:sgRNA ribonucleoproteins to bovine zygotes without compromising embryo development. Higher electroporation pulse voltage resulted in increased membrane permeability; however, voltages above 15 V/mm decreased embryo developmental potential. The zona pellucida of bovine embryos was not a barrier to efficient RNP electroporation. Using parameters optimized for maximal membrane permeability while maintaining developmental competence we achieved high rates of gene editing when targeting bovine OCT4, which resulted in absence of OCT4 protein in 100% of the evaluated embryos and the expected arrest of embryonic development at the morula stage. In conclusion, Cas9:sgRNA ribonucleoproteins can be delivered efficiently by electroporation to zona-intact bovine zygotes, resulting in efficient gene knockouts.

Simplified pipelines for genetic engineering of mammalian embryos by CRISPR-Cas9 electroporation†

Gene editing technologies, such as CRISPR-Cas9, have important applications in mammalian embryos for generating novel animal models in biomedical research and lines of livestock with enhanced production traits. However, the lack of methods for efficient introduction of gene editing reagents into zygotes of various species and the need for surgical embryo transfer in mice have been technical barriers of widespread use. Here, we described methodologies that overcome these limitations for embryos of mice, cattle, and pigs. Using mutation of the Nanos2 gene as a readout, we refined electroporation parameters with preassembled sgRNA-Cas9 RNPs for zygotes of all three species without the need for zona pellucida dissolution that led to high-efficiency INDEL edits. In addition, we optimized culture conditions to support maturation from zygote to the multicellular stage for all three species that generates embryos ready for transfer to produce gene-edited animals. Moreover, for mice, we devised a nonsurgical embryo transfer method that yields offspring at an efficiency comparable to conventional surgical approaches. Collectively, outcomes of these studies provide simplified pipelines for CRISPR-Cas9-based gene editing that are applicable in a variety of mammalian species.

Orthotopic T-Cell Receptor Replacement-An "Enabler" for TCR-Based Therapies

Natural adaptive immunity co-evolved with pathogens over millions of years, and adoptive transfer of non-engineered T cells to fight infections or cancer so far exhibits an exceptionally safe and functional therapeutic profile in clinical trials. However, the personalized nature of therapies using virus-specific T cells, donor lymphocyte infusion, or tumor-infiltrating lymphocytes makes implementation in routine clinical care difficult. In principle, genetic engineering can be used to make T-cell therapies more broadly applicable, but so far it significantly alters the physiology of cells. We recently demonstrated that orthotopic T-cell receptor (TCR) replacement (OTR) by clustered regularly interspaced short palindromic repeats (CRISPR)/ CRISPR-associated protein 9 (Cas9) can be used to generate engineered T cells with preservation of near-physiological function. In this review, we present the current status of OTR technology development and discuss its potential for TCR-based therapies. By providing the means to combine the therapeutic efficacy and safety profile of physiological T cells with the versatility of cell engineering, OTR can serve as an "enabler" for TCR-based therapies.

Modification of i-GONAD Suitable for Production of Genome-Edited C57BL/6 Inbred Mouse Strain

Improved genome editing via oviductal nucleic acid delivery (i-GONAD) is a novel method for producing genome-edited mice in the absence of ex vivo handling of zygotes. i-GONAD involves the intraoviductal injection of clustered regularly interspaced short palindromic repeats (CRISPR) ribonucleoproteins via the oviductal wall of pregnant females at 0.7 days post-coitum, followed by in vivo electroporation (EP). Unlike outbred Institute of Cancer Research (ICR) and hybrid mouse strains, genome editing of the most widely used C57BL/6J (B6) strain with i-GONAD has been considered difficult but, recently, setting a constant current of 100 mA upon EP enabled successful i-GONAD in this strain. Unfortunately, the most widely used electroporators employ a constant voltage, and thus we explored conditions allowing the generation of a 100 mA current using two electroporators: NEPA21 (Nepa Gene Co., Ltd.) and GEB15 (BEX Co., Ltd.). When the current and resistance were set to 40 V and 350–400 Ω, respectively, the current was fixed to 100 mA. Another problem in using B6 mice for i-GONAD is the difficulty in obtaining pregnant B6 females consistently because estrous females often fail to be found. A single intraperitoneal injection of low-dose pregnant mare’s serum gonadotrophin (PMSG) led to synchronization of the estrous cycle of these mice. Consequently, approximately 51% of B6 females had plugs upon mating with males 2 days after PMSG administration, which contrasts with the case (≈26%) when B6 females were subjected to natural mating. i-GONAD performed on PMSG-treated pregnant B6 females under conditions of average resistance of 367 Ω and average voltage of 116 mA resulted in the production of pregnant females at a rate of 56% (5/9 mice), from which 23 fetuses were successfully delivered. Nine (39%) of these fetuses exhibited successful genome editing at the target locus.

Recent Advances and Future Perspectives of In Vivo Targeted Delivery of Genome-Editing Reagents to Germ Cells, Embryos, and Fetuses in Mice

The recently discovered clustered regularly interspaced short palindromic repeats (CRISPR)-associated protein 9 (Cas9) systems that occur in nature as microbial adaptive immune systems are considered an important tool in assessing the function of genes of interest in various biological systems. Thus, development of efficient and simple methods to produce genome-edited (GE) animals would accelerate research in this field. The CRISPR/Cas9 system was initially employed in early embryos, utilizing classical gene delivery methods such as microinjection or electroporation, which required ex vivo handling of zygotes before transfer to recipients. Recently, novel in vivo methods such as genome editing via oviductal nucleic acid delivery (GONAD), improved GONAD (i-GONAD), or transplacental gene delivery for acquiring genome-edited fetuses (TPGD-GEF), which facilitate easy embryo manipulation, have been established. Studies utilizing these techniques employed pregnant female mice for direct introduction of the genome-editing components into the oviduct or were dependent on delivery via tail-vein injection. In mice, embryogenesis occurs within the oviducts and the uterus, which often hampers the genetic manipulation of embryos, especially those at early postimplantation stages (days 6 to 8), owing to a thick surrounding layer of tissue called decidua. In this review, we have surveyed the recent achievements in the production of GE mice and have outlined the advantages and disadvantages of the process. We have also referred to the past achievements in gene delivery to early postimplantation stage embryos and germ cells such as primordial germ cells and spermatogonial stem cells, which will benefit relevant research.

Sequential i-GONAD: An Improved In Vivo Technique for CRISPR/Cas9-Based Genetic Manipulations in Mice

Improved genome-editing via oviductal nucleic acid delivery (i-GONAD) is a technique capable of inducing genomic changes in preimplantation embryos (zygotes) present within the oviduct of a pregnant female. i-GONAD involves intraoviductal injection of a solution containing genome-editing components via a glass micropipette under a dissecting microscope, followed by in vivo electroporation using tweezer-type electrodes. i-GONAD does not involve ex vivo handling of embryos (isolation of zygotes, microinjection or electroporation of zygotes, and egg transfer of the treated embryos to the oviducts of a recipient female), which is required for in vitro genome-editing of zygotes. i-GONAD enables the generation of indels, knock-in (KI) of ~ 1 kb sequence of interest, and large deletion at a target locus. i-GONAD is usually performed on Day 0.7 of pregnancy, which corresponds to the late zygote stage. During the initial development of this technique, we performed i-GONAD on Days 1.4–1.5 (corresponding to the 2-cell stage). Theoretically, this means that at least two GONAD steps (on Day 0.7 and Day 1.4–1.5) must be performed. If this is practically demonstrated, it provides additional options for various clustered regularly interspaced palindrome repeats (CRISPR)/Caspase 9 (Cas9)-based genetic manipulations. For example, it is usually difficult to induce two independent indels at the target sites, which are located very close to each other, by simultaneous transfection of two guide RNAs and Cas9 protein. However, the sequential induction of indels at a target site may be possible when repeated i-GONAD is performed on different days. Furthermore, simultaneous introduction of two mutated lox sites (to which Cre recombinase bind) for making a floxed allele is reported to be difficult, as it often causes deletion of a sequence between the two gRNA target sites. However, differential KI of lox sites may be possible when repeated i-GONAD is performed on different days. In this study, we performed proof-of-principle experiments to demonstrate the feasibility of the proposed approach called “sequential i-GONAD (si-GONAD).”

Suppression of mosaic mutation by co-delivery of CRISPR associated protein 9 and three-prime repair exonuclease 2 into porcine zygotes via electroporation

Gene-modified animals, including pigs, can be generated efficiently by introducing CRISPR associated protein 9 (CRISPR/Cas9) into zygotes. However, in many cases, these zygotes tend to become mosaic mutants with various different mutant cell types, making it difficult to analyze the phenotype of gene-modified founder animals. To reduce the mosaic mutations, we introduced three-prime repair exonuclease 2 (Trex2), an exonuclease that improves gene editing efficiency, into porcine zygotes along with CRISPR/Cas9 via electroporation. Although the rate of porcine blastocyst formation decreased due to electroporation (25.9 ± 4.6% vs. 41.2 ± 2.0%), co-delivery of murine Trex2 (mTrex2) mRNA with CRISPR/Cas9 did not affect it any further (25.9 ± 4.6% vs. 31.0 ± 4.6%). In addition, there was no significant difference in the diameter of blastocysts carrying CRISPR/Cas9 (164.7 ± 10.2 μm), and those with CRISPR/Cas9 + mTrex2 (151.9 ± 5.1 μm) as compared to those from the control group (178.9 ± 9.0 μm). These results revealed that mTrex2 did not affect the development of pre-implantation embryo. We also found bi-allelic, as well as mono-allelic, non-mosaic homozygous mutations in the blastocysts. Most importantly, co-delivery of mTrex2 mRNA with CRISPR/Cas9 increased non-mosaic mutant blastocysts (29.3 ± 4.5%) and reduced mosaic mutant blastocysts (70.7 ± 4.5%) as compared to CRISPR/Cas9 alone (5.6 ± 6.4% and 92.6 ± 8.6%, respectively). These data suggest that the co-delivery of CRISPR/Cas9 and mTrex2 is a useful method to suppress mosaic mutation.

ElectroPen: An ultra-low-cost, electricity-free, portable electroporator

Electroporation is a basic yet powerful method for delivering small molecules (RNA, DNA, drugs) across cell membranes by application of an electrical field. It is used for many diverse applications, from genetically engineering cells to drug- and DNA-based vaccine delivery. Despite this broad utility, the high cost of electroporators can keep this approach out of reach for many budget-conscious laboratories. To address this need, we develop a simple, inexpensive, and handheld electroporator inspired by and derived from a common household piezoelectric stove lighter. The proposed "ElectroPen" device can cost as little as 23 cents (US dollars) to manufacture, is portable (weighs 13 g and requires no electricity), can be easily fabricated using 3D printing, and delivers repeatable exponentially decaying pulses of about 2,000 V in 5 ms. We provide a proof-of-concept demonstration by genetically transforming plasmids into Escherichia coli cells, showing transformation efficiency comparable to commercial devices, but at a fraction of the cost. We also demonstrate the potential for rapid dissemination of this approach, with multiple research groups across the globe validating the ease of construction and functionality of our device, supporting the potential for democratization of science through frugal tools. Thus, the simplicity, accessibility, and affordability of our device holds potential for making modern synthetic biology accessible in high school, community, and resource-poor laboratories.

This Community Page article describes an ultra-low–cost (23-cent) 3D-printed electroporator, inspired by a common barbecue lighter, designed to enable broader access to synthetic biology in high-school, community, and budget-conscious laboratories.

Introducing gene deletions by mouse zygote electroporation of Cas12a/Cpf1

CRISPR-associated (Cas) nucleases are established tools for engineering of animal genomes. These programmable RNA-guided nucleases have been introduced into zygotes using expression vectors, mRNA, or directly as ribonucleoprotein (RNP) complexes by different delivery methods. Whereas microinjection techniques are well established, more recently developed electroporation methods simplify RNP delivery but can provide less consistent efficiency. Previously, we have designed Cas12a-crRNA pairs to introduce large genomic deletions in the Ubn1, Ubn2, and Rbm12 genes in mouse embryonic stem cells (ESC). Here, we have optimized the conditions for electroporation of the same Cas12a RNP pairs into mouse zygotes. Using our protocol, large genomic deletions can be generated efficiently by electroporation of zygotes with or without an intact zona pellucida. Electroporation of as few as ten zygotes is sufficient to obtain a gene deletion in mice suggesting potential applicability of this method for species with limited availability of zygotes.

Defective tubulin detyrosination causes structural brain abnormalities with cognitive deficiency in humans and mice

Reversible detyrosination of tubulin, the building block of microtubules, is crucial for neuronal physiology. Enzymes responsible for detyrosination were recently identified as complexes of vasohibins (VASHs) one or two with small VASH-binding protein (SVBP). Here we report three consanguineous families, each containing multiple individuals with biallelic inactivation of SVBP caused by truncating variants (p.Q28* and p.K13Nfs*18). Affected individuals show brain abnormalities with microcephaly, intellectual disability and delayed gross motor and speech development. Immunoblot testing in cells with pathogenic SVBP variants demonstrated that the encoded proteins were unstable and non-functional, resulting in a complete loss of VASH detyrosination activity. Svbp knockout mice exhibit drastic accumulation of tyrosinated tubulin and a reduction of detyrosinated tubulin in brain tissue. Similar alterations in tubulin tyrosination levels were observed in cultured neurons and associated with defects in axonal differentiation and architecture. Morphological analysis of the Svbp knockout mouse brains by anatomical magnetic resonance imaging showed a broad impact of SVBP loss, with a 7% brain volume decrease, numerous structural defects and a 30% reduction of some white matter tracts. Svbp knockout mice display behavioural defects, including mild hyperactivity, lower anxiety and impaired social behaviour. They do not, however, show prominent memory defects. Thus, SVBP-deficient mice recapitulate several features observed in human patients. Altogether, our data demonstrate that deleterious variants in SVBP cause this neurodevelopmental pathology, by leading to a major change in brain tubulin tyrosination and alteration of microtubule dynamics and neuron physiology.

Reproducibility of CRISPR-Cas9 methods for generation of conditional mouse alleles: a multi-center evaluation

BACKGROUND

CRISPR-Cas9 gene-editing technology has facilitated the generation of knockout mice, providing an alternative to cumbersome and time-consuming traditional embryonic stem cell-based methods. An earlier study reported up to 16% efficiency in generating conditional knockout (cKO or floxed) alleles by microinjection of 2 single guide RNAs (sgRNA) and 2 single-stranded oligonucleotides as donors (referred herein as "two-donor floxing" method).

RESULTS

We re-evaluate the two-donor method from a consortium of 20 laboratories across the world. The dataset constitutes 56 genetic loci, 17,887 zygotes, and 1718 live-born mice, of which only 15 (0.87%) mice contain cKO alleles. We subject the dataset to statistical analyses and a machine learning algorithm, which reveals that none of the factors analyzed was predictive for the success of this method. We test some of the newer methods that use one-donor DNA on 18 loci for which the two-donor approach failed to produce cKO alleles. We find that the one-donor methods are 10- to 20-fold more efficient than the two-donor approach.

CONCLUSION

We propose that the two-donor method lacks efficiency because it relies on two simultaneous recombination events in cis, an outcome that is dwarfed by pervasive accompanying undesired editing events. The methods that use one-donor DNA are fairly efficient as they rely on only one recombination event, and the probability of correct insertion of the donor cassette without unanticipated mutational events is much higher. Therefore, one-donor methods offer higher efficiencies for the routine generation of cKO animal models.

Creation of CRISPR-based germline-genome-engineered mice without ex vivo handling of zygotes by i-GONAD

Methods to create genetically engineered mice involve three major steps: harvesting embryos from one set of females, microinjection of reagents into embryos ex vivo and their surgical transfer to another set of females. Although tedious, these methods have been used for more than three decades to create mouse models. We recently developed a method named GONAD (genome editing via oviductal nucleic acids delivery), which bypasses these steps. GONAD involves injection of CRISPR components (Cas9 mRNA and guide RNA (gRNA)) into the oviducts of pregnant females 1.5 d post conception, followed by in vivo electroporation to deliver the components into the zygotes in situ. Using GONAD, we demonstrated that target genes can be disrupted and analyzed at different stages of mouse embryonic development. Subsequently, we developed improved GONAD (i-GONAD) by delivering CRISPR ribonucleoproteins (RNPs; Cas9 protein or Cpf1 protein and gRNA) into day-0.7 pregnant mice, which made it suitable for routine generation of knockout and large-deletion mouse models. i-GONAD can also generate knock-in models containing up to 1-kb inserts when single-stranded DNA (ssDNA) repair templates are supplied. i-GONAD offers other advantages: it does not require vasectomized males and pseudo-pregnant females, the females used for i-GONAD are not sacrificed and can be used for other experiments, it can be easily adopted in laboratories lacking sophisticated microinjection equipment, and can be implemented by researchers skilled in small-animal surgery but lacking embryo-handling skills. Here, we provide a step-by-step protocol for establishing the i-GONAD method. The protocol takes ∼6 weeks to generate the founder mice.

Biomaterials as vectors for the delivery of CRISPR-Cas9

The emergence of the CRISPR-Cas9 gene editing system has brought much hope and excitement to the field of gene therapy and the larger scientific community. However, in order for CRISPR-based therapies to be translated to the clinical setting, there is an urgent need to develop optimized vectors for their delivery. The delivery vector is a crucial determinant of the therapeutic efficacy of gene editing and should be designed to accommodate various factors including the form of the payload, the physiological environment, and the potential immune responses. Recently, biomaterials have become an attractive option for the delivery of Cas9 due to their tunability, biocompatibility and increasing efficacy at drug delivery. Biomaterials offer a unique solution for creating tailored vectors to meet the demands of various applications that cannot be easily met by other delivery methods. In this review, we will discuss the various biomaterial systems that have been used to deliver Cas9 in its plasmid, mRNA and protein forms. In addition, the functions of these materials will be reviewed to understand their roles in Cas9 delivery. Finally, the immune challenges associated with Cas9 and the delivery materials will be discussed as an understanding of the immune responses along with the functions of biomaterials will ultimately guide the field in designing new delivery systems for the clinical applications of CRISPR-Cas9.

i-GONAD: A method for generating genome-edited animals without ex vivo handling of embryos

The recent development of genome editing technologies has enabled the creation of genome-edited animals, with alterations at the desired target locus. The clustered regularly interspaced short palindromic repeats (CRISPR) system is widely used for this purpose because it is simpler and more efficient than other genome editing technologies. The conventional methods for creation of genome-edited animals involve ex vivo handling of embryos (zygotes) for microinjection or in vitro electroporation. However, this process is laborious and time-consuming, and relatively large numbers of animals are used. Furthermore, these methods require specialized skills for handling embryos. In 2015, we reported a novel method for the creation of genome-edited animals without ex vivo handling of embryos. The technology known as Genome-editing via Oviductal Nucleic Acids Delivery (GONAD) involved intraoviductal instillation of genome editing components into a pregnant female and subsequent in vivo electroporation of an entire oviduct. The genome editing components present in the oviductal lumen are transferred to preimplantation embryos in situ for introducing insertion or deletion (indel) mutations at the desired loci. This technology was further improved by optimizing several parameters to develop improved GONAD (i-GONAD) for the efficient generation of mutant or knock-in animals. In this review, we discuss the historical background, potential applications, advantages, and future challenges of GONAD/i-GONAD technology.

Cas9 Ribonucleoprotein Complex Delivery: Methods and Applications for Neuroinflammation

The CRISPR/Cas9 system is a revolutionary gene editing technology that combines simplicity of use and efficiency of mutagenesis. As this technology progresses toward human therapies, valid concerns including off-target mutations and immunogenicity must be addressed. One approach to address these issues is to minimize the presence of the CRISPR/Cas9 components by maintaining a tighter temporal control of Cas9 endonuclease and reducing the time period of activity. This has been achieved to some degree by delivering the CRISPR/Cas9 system via pre-formed Cas9 + gRNA ribonucleoprotein (RNP) complexes. In this review, we first discuss the molecular modifications that can be made using CRISPR/Cas9 and provide an overview of current methods for delivering Cas9 RNP complexes both in vitro and in vivo. We conclude with examples of how Cas9 RNP delivery may be used to target neuroinflammatory processes, namely in regard to viral infections of the central nervous system and neurodegenerative diseases. We propose that Cas9 RNP delivery is a viable approach when considering the CRISPR/Cas9 system for both experimentation and the treatment of disease. Graphical Abstract.

CRISPR links to long noncoding RNA function in mice: A practical approach

Next generation sequencing has uncovered a trove of short noncoding RNAs (e.g., microRNAs) and long noncoding RNAs (lncRNAs) that act as molecular rheostats in the control of diverse homeostatic processes. Meanwhile, the tsunamic emergence of clustered regularly interspaced short palindromic repeats (CRISPR) editing has transformed our influence over all DNA-carrying entities, heralding global CRISPRization. This is evident in biomedical research where the ease and low-cost of CRISPR editing has made it the preferred method of manipulating the mouse genome, facilitating rapid discovery of genome function in an in vivo context. Here, CRISPR genome editing components are updated for elucidating lncRNA function in mice. Various strategies are highlighted for understanding the function of lncRNAs residing in intergenic sequence space, as host genes that harbor microRNAs or other genes, and as natural antisense, overlapping or intronic genes. Also discussed is CRISPR editing of mice carrying human lncRNAs as well as the editing of competing endogenous RNAs. The information described herein should assist labs in the rigorous design of experiments that interrogate lncRNA function in mice where complex disease processes can be modeled thus accelerating translational discovery.

Programmed genome editing of the omega-1 ribonuclease of the blood fluke, Schistosoma mansoni

CRISPR/Cas9-based genome editing has yet to be reported in species of the Platyhelminthes. We tested this approach by targeting omega-1 (ω1) of Schistosoma mansoni as proof of principle. This secreted ribonuclease is crucial for Th2 polarization and granuloma formation. Schistosome eggs were exposed to Cas9 complexed with guide RNA complementary to ω1 by electroporation or by transduction with lentiviral particles. Some eggs were also transfected with a single stranded donor template. Sequences of amplicons from gene-edited parasites exhibited Cas9-catalyzed mutations including homology directed repaired alleles, and other analyses revealed depletion of ω1 transcripts and the ribonuclease. Gene-edited eggs failed to polarize Th2 cytokine responses in macrophage/T-cell co-cultures, while the volume of pulmonary granulomas surrounding ω1-mutated eggs following tail-vein injection into mice was vastly reduced. Knock-out of ω1 and the diminished levels of these cytokines following exposure showcase the novel application of programmed gene editing for functional genomics in schistosomes.

Schistosomiasis is a tropical disease that can cause serious health problems, including damage to the liver and kidneys, infertility and bladder cancer. Nearly a quarter billion people are currently infected, mostly in poor regions of sub-Saharan Africa, the Philippines and Brazil.

A freshwater worm known as Schistosoma mansoni causes the disease. These parasites enter the human body by burrowing into the skin; once in the bloodstream, they move to various organs where they rapidly start to reproduce. Their eggs release several molecules, including a protein known as omega-1 ribonuclease, which can damage the surrounding tissues.

A gene editing technique called CRISPR/Cas9 allows scientists to precisely target and then deactivate the genetic information a cell needs to produce a given protein. While the tool has been used in other species before, it was unknown if it could be applied to S. mansoni. Here, Ittiprasert et al. harnessed CRISPR/Cas9 to deactivate the gene that codes for omega-1 ribonuclease and create parasites that do not produce the protein, or only very little of it. The experiments showed that mice infected with the gene-edited worm eggs displayed far fewer symptoms of schistosomiasis compared to those that carry the non-edited parasites.

Alongside this work, Arunsan et al. used CRISPR/Cas9 to inactivate a gene in another species of worm that can cause liver cancer in humans. Together, these findings demonstrate for the first time that the gene editing method can be adapted for use in parasitic flatworms, which are a major public health problem in tropical climates. This tool should help scientists understand how the parasites invade and damage our bodies, and provide new ideas for treatment and disease control.

An optimized electroporation approach for efficient CRISPR/Cas9 genome editing in murine zygotes

Electroporation of zygotes represents a rapid alternative to the elaborate pronuclear injection procedure for CRISPR/Cas9-mediated genome editing in mice. However, current protocols for electroporation either require the investment in specialized electroporators or corrosive pre-treatment of zygotes which compromises embryo viability. Here, we describe an easily adaptable approach for the introduction of specific mutations in C57BL/6 mice by electroporation of intact zygotes using a common electroporator with synthetic CRISPR/Cas9 components and minimal technical requirement. Direct comparison to conventional pronuclear injection demonstrates significantly reduced physical damage and thus improved embryo development with successful genome editing in up to 100% of living offspring. Hence, our novel approach for Easy Electroporation of Zygotes (EEZy) allows highly efficient generation of CRISPR/Cas9 transgenic mice while reducing the numbers of animals required.

Reproductive technologies for the generation and maintenance of valuable animal strains

Many types of mutant and genetically engineered strains have been produced in various animal species. Their numbers have dramatically increased in recent years, with new strains being rapidly produced using genome editing techniques. In the rat, it has been difficult to produce knockout and knock-in strains because the establishment of stem cells has been insufficient. However, a large number of knockout and knock-in strains can currently be produced using genome editing techniques, including zinc-finger nuclease (ZFN), transcription activator-like effector nuclease (TALEN), and the clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated protein 9 (Cas9) system. Microinjection technique has also contributed widely to the production of various kinds of genome edited animal strains. A novel electroporation method, the “Technique for Animal Knockout system by Electroporation (TAKE)” method, is a simple and highly efficient tool that has accelerated the production of new strains. Gamete preservation is extremely useful for maintaining large numbers of these valuable strains as genetic resources in the long term. These reproductive technologies, including microinjection, TAKE method, and gamete preservation, strongly support biomedical research and the bio-resource banking of animal models. In this review, we introduce the latest reproductive technologies used for the production of genetically engineered animals, especially rats, using genome editing techniques and the efficient maintenance of valuable strains as genetic resources. These technologies can also be applied to other laboratory animals, including mice, and domestic and wild animal species.