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CRISPR/Cas9-editing of PRNP in Alpine goats.
Misfolding of the cellular PrP (PrPc) protein causes prion disease, leading to neurodegenerative disorders in numerous mammalian species, including goats. A lack of PrPc induces complete resistance to prion disease. The aim of this work was to engineer Alpine goats carrying knockout (KO) alleles of PRNP, the PrPc-encoding gene, using CRISPR/Cas9-ribonucleoproteins and single-stranded donor oligonucleotides. The targeted region preceded the PRNPTer mutation previously described in Norwegian goats. Genome editors were injected under the zona pellucida prior to the electroporation of 565 Alpine goat embryos/oocytes. A total of 122 two-cell-stage embryos were transferred to 46 hormonally synchronized recipient goats. Six of the goats remained pregnant and naturally gave birth to 10 offspring. Among the 10 newborns, eight founder animals carrying PRNP genome-edited alleles were obtained. Eight different mutated alleles were observed, including five inducing KO mutations. Three founders carried only genome-edited alleles and were phenotypically indistinguishable from their wild-type counterparts. Among them, one male carrying a one base pair insertion leading to a KO allele is currently used to rapidly extend a PRNP-KO line of Alpine goats for future characterization. In addition to KO alleles, a PRNPdel6 genetic variant has been identified in one-third of founder animals. This new variant will be tested for its potential properties with respect to prion disease. Future studies will also evaluate the effects of genetic background on other characters associated with PRNP KO, as previously described in the Norwegian breed or other species.
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Discovery of rice essential genes by characterizing a CRISPR-edited mutation of closely related rice MAP kinase genes.
The clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein 9 nuclease (Cas9) system depends on a guide RNA (gRNA) to specify its target. By efficiently co-expressing multiple gRNAs that target different genomic sites, the polycistronic tRNA-gRNA gene (PTG) strategy enables multiplex gene editing in the family of closely related mitogen-activated protein kinase (MPK) genes in Oryza sativa (rice). In this study, we identified MPK1 and MPK6 (Arabidopsis AtMPK6 and AtMPK4 orthologs, respectively) as essential genes for rice development by finding the preservation of MPK functional alleles and normal phenotypes in CRISPR-edited mutants. The true knock-out mutants of MPK1 were severely dwarfed and sterile, and homozygous mpk1 seeds from heterozygous parents were defective in embryo development. By contrast, heterozygous mpk6 mutant plants completely failed to produce homozygous mpk6 seeds. In addition, the functional importance of specific MPK features could be evaluated by characterizing CRISPR-induced allelic variation in the conserved kinase domain of MPK6. By simultaneously targeting between two and eight genomic sites in the closely related MPK genes, we demonstrated 45-86% frequency of biallelic mutations and the successful creation of single, double and quadruple gene mutants. Indels and fragment deletion were both stably inherited to the next generations, and transgene-free mutants of rice MPK genes were readily obtained via genetic segregation, thereby eliminating any positional effects of transgene insertions. Taken together, our study reveals the essentiality of MPK1 and MPK6 in rice development, and enables the functional discovery of previously inaccessible genes or domains with phenotypes masked by lethality or redundancy.
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High fidelity CRISPR/Cas9 increases precise monoallelic and biallelic editing events in primordial germ cells.
Primordial germ cells (PGCs), the embryonic precursors of the sperm and egg, are used for the introduction of genetic modifications into avian genome. Introduction of small defined sequences using genome editing has not been demonstrated in bird species. Here, we compared oligonucleotide-mediated HDR using wild type SpCas9 (SpCas9-WT) and high fidelity SpCas9-HF1 in PGCs and show that many loci in chicken PGCs can be precise edited using donors containing CRISPR/Cas9-blocking mutations positioned in the protospacer adjacent motif (PAM). However, targeting was more efficient using SpCas9-HF1 when mutations were introduced only into the gRNA target sequence. We subsequently employed an eGFP-to-BFP conversion assay, to directly compare HDR mediated by SpCas9-WT and SpCas9-HF1 and discovered that SpCas9-HF1 increases HDR while reducing INDEL formation. Furthermore, SpCas9-HF1 increases the frequency of single allele editing in comparison to SpCas9-WT. We used SpCas9-HF1 to demonstrate the introduction of monoallelic and biallelic point mutations into the FGF20 gene and generate clonal populations of edited PGCs with defined homozygous and heterozygous genotypes. Our results demonstrate the use of oligonucleotide donors and high fidelity CRISPR/Cas9 variants to perform precise genome editing with high efficiency in PGCs.
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Protocol for genetic dissection of class switch recombination using genome-editing tools.
Antibody class switch recombination is achieved through programmed DNA damage, and the processing of programmed DNA lesions requires the coordinated action of many DNA metabolic factors. Here, we present a protocol for inducing class switch recombination using base editors or CRISPR-Cas9. We provide optimized guide RNA (gRNA) sequences and describe steps for cytokine activation, electroporation, surface immunoglobulin detection, and data analysis. This method allows researchers to investigate the involvement of specific factors in antibody diversification and elucidate their functional roles. For complete details on the use and execution of this protocol, please refer to Dai et al.1.
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Editing of DNA methylation using CRISPR/Cas9 and a ssDNA template in human cells.
Programmable DNA methylation is required for understanding of transcriptional regulation and elucidating gene functions. We previously reported that MMEJ-based promoter replacement enabled targeted DNA methylation in human cells. ssDNA-mediated knock-in has recently been reported to completely reduce random integrations. We speculated that by changing MMEJ-to ssDNA-based knock-in, targeted DNA methylation may be achieved through a hemimethylation-symmetric methylation pathway. We herein successfully developed a new system that enables the replacement of an unmethylated promoter with a methylated ssDNA promoter through ssDNA-based knock-in. A DNA methylation ratio of approximately 100% was achieved at the cancer-associated gene SP3 in HEK293 cells. The present results provide a promising framework for artificial epigenetic modifications.
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Improved Genome Editing Efficiency and Flexibility Using Modified Oligonucleotides with TALEN and CRISPR-Cas9 Nucleases.
Genome editing has now been reported in many systems using TALEN and CRISPR-Cas9 nucleases. Precise mutations can be introduced during homology-directed repair with donor DNA carrying the wanted sequence edit, but efficiency is usually lower than for gene knockout and optimal strategies have not been extensively investigated. Here, we show that using phosphorothioate-modified oligonucleotides strongly enhances genome editing efficiency of single-stranded oligonucleotide donors in cultured cells. In addition, it provides better design flexibility, allowing insertions more than 100 bp long. Despite previous reports of phosphorothioate-modified oligonucleotide toxicity, clones of edited cells are readily isolated and targeted sequence insertions are achieved in rats and mice with very high frequency, allowing for homozygous loxP site insertion at the mouse ROSA locus in particular. Finally, when detected, imprecise knockin events exhibit indels that are asymmetrically positioned, consistent with genome editing taking place by two steps of single-strand annealing.
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CRISPR/Cas9-Based Efficient Genome Editing in Clostridium ljungdahlii, an Autotrophic Gas-Fermenting Bacterium.
Acetogenic bacteria have the potential to convert single carbon gases (CO and CO2) into a range of bulk chemicals and fuels. Realization of their full potential is being impeded by the absence of effective genetic tools for high throughput genome modification. Here we report the development of a highly efficient CRISPR/Cas9 system for rapid genome editing of Clostridium ljungdahlii, a paradigm for the commercial production of ethanol from synthesis gas. Following the experimental selection of two promoters (Pthl and ParaE) for expression of cas9 and the requisite single guide RNA (sgRNA), the efficiency of system was tested by making precise deletions of four genes, pta, adhE1, ctf and pyrE. Deletion efficiencies were 100%, >75%, 100% and >50%, respectively. The system overcomes the deficiencies of currently available tools (more rapid, no added antibiotic resistance gene, scarless and minimal polar effects) and will find utility in other acetogens, including the pathogen Clostridium difficile.
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Tracking-seq reveals the heterogeneity of off-target effects in CRISPR-Cas9-mediated genome editing.
The continued development of novel genome editors calls for a universal method to analyze their off-target effects. Here we describe a versatile method, called Tracking-seq, for in situ identification of off-target effects that is broadly applicable to common genome-editing tools, including Cas9, base editors and prime editors. Through tracking replication protein A (RPA)-bound single-stranded DNA followed by strand-specific library construction, Tracking-seq requires a low cell input and is suitable for in vitro, ex vivo and in vivo genome editing, providing a sensitive and practical genome-wide approach for off-target detection in various scenarios. We show, using the same guide RNA, that Tracking-seq detects heterogeneity in off-target effects between different editor modalities and between different cell types, underscoring the necessity of direct measurement in the original system.
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Characterization of targeted knock-in achieved via tandem paired nicking mediated by CRISPR/Cas9 nickases.
Targeted knock-in of specific DNA sequences using CRISPR/Cas9 is an advanced technology that enables programmed genome alterations including insertions, deletions, and base substitutions exactly as designed. Despite its utility in life sciences and promise for medical and industrial applications, it remains critical to establish a methodology for highly precise and efficient targeted knock-in to facilitate the practical use of this technology. Tandem paired nicking (TPN) is a genome editing methodology leveraging nicking variants of CRISPR/Cas9 nucleases (Cas9 nickases) to create site-specific nicks within the homologous region of the genome and donor DNA. Such nicking configuration promotes precise and efficient targeted knock-in while repressing the formation of unintended insertions and deletions and p53-mediated DNA damage response. In this study, we conducted a detailed characterization of TPN-based targeted knock-in by performing genome editing assays with various nicking configurations modified from TPN. Our results demonstrated that genomic nicks remarkably contribute to TPN-based targeted knock-in, whereas donor nicks play a less critical role. The introduction of additional nicks beyond the standard two-nick configuration did not further improve the efficiency of TPN-based targeted knock-in. Comparison with other Cas9 nickase-based methodologies for targeted knock-in demonstrated largely equivalent knock-in efficiencies achieved by these methodologies. High-throughput long-read sequencing confirmed a lower incidence of undesired insertions and deletions of various lengths by TPN, in comparison with a conventional Cas9 nuclease-based approach. These findings underscore TPN as a methodology for precise and efficient targeted knock-in, and highlight the broad potential of Cas9 nickase-based targeted knock-in for clinical and industrial applications.
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A redox switch regulates the assembly and anti-CRISPR activity of AcrIIC1.
Anti-CRISPRs (Acrs) are natural inhibitors of bacteria's CRISPR-Cas systems, and have been developed as a safeguard to reduce the off-target effects of CRISPR gene-editing technology. Acrs can directly bind to CRISPR-Cas complexes and inhibit their activities. However, whether this process is under regulation in diverse eukaryotic cellular environments is poorly understood. In this work, we report the discovery of a redox switch for NmeAcrIIC1, which regulates NmeAcrIIC1's monomer-dimer interconversion and inhibitory activity on Cas9. Further structural studies reveal that a pair of conserved cysteines mediates the formation of inactive NmeAcrIIC1 dimer and directs the redox cycle. The redox switch also applies to the other two AcrIIC1 orthologs. Moreover, by replacing the redox-sensitive cysteines, we generated a robust AcrIIC1 variant that maintains potent inhibitory activity under various redox conditions. Our results reveal a redox-dependent regulation mechanism of Acr, and shed light on the design of superior Acr for CRISPR-Cas systems.
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