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Switchable genome editing via genetic code expansion.
Multiple applications of genome editing by CRISPR-Cas9 necessitate stringent regulation and Cas9 variants have accordingly been generated whose activity responds to small ligands, temperature or light. However, these approaches are often impracticable, for example in clinical therapeutic genome editing in situ or gene drives in which environmentally-compatible control is paramount. With this in mind, we have developed heritable Cas9-mediated mammalian genome editing that is acutely controlled by the cheap lysine derivative, Lys(Boc) (BOC). Genetic code expansion permitted non-physiological BOC incorporation such that Cas9 (Cas9BOC) was expressed in a full-length, active form in cultured somatic cells only after BOC exposure. Stringently BOC-dependent, heritable editing of transgenic and native genomic loci occurred when Cas9BOC was expressed at the onset of mouse embryonic development from cRNA or Cas9BOC transgenic females. The tightly controlled Cas9 editing system reported here promises to have broad applications and is a first step towards purposed, spatiotemporal gene drive regulation over large geographical ranges.
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Kinetics and Fidelity of the Repair of Cas9-Induced Double-Strand DNA Breaks.
The RNA-guided DNA endonuclease Cas9 is a powerful tool for genome editing. Little is known about the kinetics and fidelity of the double-strand break (DSB) repair process that follows a Cas9 cutting event in living cells. Here, we developed a strategy to measure the kinetics of DSB repair for single loci in human cells. Quantitative modeling of repaired DNA in time series after Cas9 activation reveals variable and often slow repair rates, with half-life times up to ∼10 hr. Furthermore, repair of the DSBs tends to be error prone. Both classical and microhomology-mediated end joining pathways contribute to the erroneous repair. Estimation of their individual rate constants indicates that the balance between these two pathways changes over time and can be altered by additional ionizing radiation. Our approach provides quantitative insights into DSB repair kinetics and fidelity in single loci and indicates that Cas9-induced DSBs are repaired in an unusual manner.
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Dual sgRNA-directed gene deletion in basidiomycete Ganoderma lucidum using the CRISPR/Cas9 system.
Ganoderma lucidum is an important medicinal mushroom in traditional Chinese medicine. However, the lack of adequate genetic tools has hindered molecular genetic research in and the genetic modification of this species. Here, we report that the presence of an intron is necessary for the efficient expression of the heterologous phosphinothricin-resistance and green fluorescent protein genes in G. lucidum. Moreover, we improved the CRISPR/Cas9-mediated gene disruption frequency in G. lucidum by adding an intron upstream of the Cas9 gene. Our results showed that the disruption frequency of the orotidine 5'-monophosphate decarboxylase gene (ura3) in transformants containing the glyceraldehyde-3-phosphate dehydrogenase gene intron in the Cas9 plasmid is 14-18 in 107 protoplasts, which is 10.6 times higher than that in transformants without any intron sequence. Furthermore, genomic fragment deletions in the ura3 and GL17624 genes were achieved via a dual sgRNA-directed CRISPR/Cas9 system in G. lucidum. We achieved a ura3 deletion frequency of 36.7% in G. lucidum. The developed method provides a powerful platform to generate gene deletion mutants and will facilitate functional genomic studies in G. lucidum.
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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.
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Comprehensive deletion scan of anti-CRISPR AcrIIA4 reveals essential and dispensable domains for Cas9 inhibition.
Delineating a protein's essential and dispensable domains provides critical insight into how it carries out its function. Here, we developed a high-throughput method to synthesize and test the functionality of all possible in-frame and continuous deletions in a gene of interest, enabling rapid and unbiased determination of protein domain importance. Our approach generates precise deletions using a CRISPR library framework that is free from constraints of gRNA target site availability and efficacy. We applied our method to AcrIIA4, a phage-encoded anti-CRISPR protein that robustly inhibits SpCas9. Extensive structural characterization has shown that AcrIIA4 physically occupies the DNA-binding interfaces of several SpCas9 domains; nonetheless, the importance of each AcrIIA4 interaction for SpCas9 inhibition is unknown. We used our approach to determine the essential and dispensable regions of AcrIIA4. Surprisingly, not all contacts with SpCas9 were required, and in particular, we found that the AcrIIA4 loop that inserts into SpCas9's RuvC catalytic domain can be deleted. Our results show that AcrIIA4 inhibits SpCas9 primarily by blocking PAM binding and that its interaction with the SpCas9 catalytic domain is inessential.
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No off-target mutations in functional genome regions of a CRISPR/Cas9-generated monkey model of muscular dystrophy.
CRISPR/Cas9 is now widely used in biomedical research and has great potential for clinical applications. However, the safety and efficacy of this gene-editing technique are significant issues. Recent reports on mouse models and human cells have raised concerns that off-target mutations could hamper applying the CRISPR technology in patients. The high similarities of nonhuman primates to humans in genome content and organization, genetic diversity, physiology, and cognitive abilities have made these animals ideal experimental models for understanding human diseases and developing therapeutics. Off-target mutations of CRISPR/Cas9 have been analyzed in previous studies of nonhuman primates, but no report has investigated genome-wide off-target effects in living monkeys. Here, we used rhesus monkeys in which a genetic disorder mimicking Duchenne muscular dystrophy had previously been produced with CRISPR/Cas9. Using whole-genome sequencing to comprehensively assess on- and off-target mutations in these animals, we found that CRISPR/Cas9-based gene editing is active on the expected genomic sites without producing off-target modifications in other functional regions of the genome. These findings suggest that the CRISPR/Cas9 technique could be relatively safe and effective in modeling genetic disease in nonhuman primates and in future therapeutic research of human diseases.
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REEP6 deficiency leads to retinal degeneration through disruption of ER homeostasis and protein trafficking.
Retinitis pigmentosa (RP) is the most common form of inherited retinal dystrophy. We recently identified mutations in REEP6, which encodes the receptor expression enhancing protein 6, in several families with autosomal recessive RP. REEP6 is related to the REEP and Yop1p family of ER shaping proteins and potential receptor accessory proteins, but the role of REEP6 in the retina is unknown. Here we characterize the disease mechanisms associated with loss of REEP6 function using a Reep6 knockout mouse generated by CRISPR/Cas9 gene editing. In control mice REEP6 was localized to the inner segment and outer plexiform layer of rod photoreceptors. The Reep6-/- mice exhibited progressive photoreceptor degeneration from P20 onwards. Ultrastructural analyses at P20 by transmission electron microscopy and 3View serial block face scanning EM revealed an expansion of the distal ER in the Reep6-/- rods and an increase in their number of mitochondria. Electroretinograms revealed photoreceptor dysfunction preceded degeneration, suggesting potential defects in phototransduction. There was no effect on the traffic of rhodopsin, Rom1 or peripherin/rds; however, the retinal guanylate cyclases GC1 and GC2 were severely affected in the Reep6 knockout animals, with almost undetectable expression. These changes correlated with an increase in C/EBP homologous protein (CHOP) expression and the activation of caspase 12, suggesting that ER stress contributes to cell death. Collectively, these data suggest that REEP6 plays an essential role in maintaining cGMP homeostasis though facilitating the stability and/or trafficking of guanylate cyclases and maintaining ER and mitochondrial homeostasis.
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A phage DUF669 protein mediates post-cleavage DNA repair against host CRISPR immunity.
Phages have evolved various anti-CRISPR (Acr) proteins or RNA-based anti-CRISPR (Racr) to evade host CRISPR immunity by direct interference. However, whether other counter-CRISPR mechanisms exist remains unexplored. Here, we report a phage-encoded two-protein system, termed Healer, which neutralizes CRISPR immunity via a post-cleavage DNA repair mechanism. This phage replication-associated system comprises two proteins: Gp63 (a DUF669 domain-containing protein), and Gp64 (an AAA domain-containing protein). Mechanistic investigations elucidate that Gp63 acts as a rapid-response effector of CRISPR-induced DNA breaks. After Gp63 binds ssDNA, it promotes Gp64-mediated homologous recombination, which repairs CRISPR-generated phage DNA breaks and supports phage viability. Notably, co-expression of the Healer system with Cas9/Cas12 in E. coli, P. aeruginosa, and A. baumannii demonstrated high phage genome-editing efficiency. Collectively, our findings reveal an important anti-CRISPR complementary strategy, providing a promising tool for genome engineering.
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A general genome editing strategy using CRISPR lipid nanoparticle spherical nucleic acids.
Genome editing with CRISPR-Cas systems hold promise for treating a wide range of genetic disorders and cancers. However, efficient delivery of genome editors remains challenging due to the requirement for the simultaneous delivery or intracellular generation of Cas proteins, guide RNAs, and, in some applications, donor DNAs. Furthermore, the immunogenicity and toxicity of delivery vehicles can limit the safety and efficacy of genetic medicines. Here, we combine two nucleic acid delivery approaches to create CRISPR lipid nanoparticle-spherical nucleic acids (LNP-SNAs) that are both efficient and biocompatible. Compared to lipid nanoparticles (LNPs) lacking a surface-bound DNA shell, CRISPR LNP-SNAs exhibit two- to three-fold higher cellular uptake, reduced cytotoxicity, and improved gene transfection efficiency. Across multiple cell lines and genomic loci, CRISPR LNP-SNAs induce insertion-deletion mutations at average frequencies two- to three-fold higher than those observed with LNPs. When codelivered with donor templates, CRISPR LNP-SNAs enable homology-directed repair at an average efficiency of 21 ± 7%, a 2.5-fold improvement over LNPs (8 ± 4%). The ease of synthesis and biocompatibility of CRISPR LNP-SNAs highlight their potential as a versatile delivery platform for CRISPR-Cas and other gene therapies.
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Abrogation of HLA surface expression using CRISPR/Cas9 genome editing: a step toward universal T cell therapy.
As recent advancements in the chimeric antigen receptor-T cells have revolutionized the way blood cancers are handled, potential benefits from producing off-the-shelf, standardized immune cells entail the need for development of allogeneic immune cell therapy. However, host rejection driven by HLA disparity in adoptively transferred allogeneic T cells remains a key obstacle to the universal donor T cell therapy. To evade donor HLA-mediated immune rejection, we attempted to eliminate T cell's HLA through the CRISPR/Cas9 gene editing system. First, we screened 60 gRNAs targeting B2M and multiple sets of gRNA each targeting α chains of HLA-II (DPA, DQA and DRA, respectively) using web-based design tools, and identified specific gRNA sequences highly efficient for target deletion without carrying off-target effects. Multiplex genome editing of primary human T cells achieved by the newly discovered gRNAs yielded HLA-I- or HLA-I/II-deficient T cells that were phenotypically unaltered and functionally intact. The overnight mixed lymphocyte reactions demonstrated the HLA-I-negative cells induced decreased production of IFN-γ and TNF-α in alloreactive T cells, and deficiency of HLA-I/II in T cells further dampened the inflammatory responses. Taken together, our approach will provide an efficacious pathway toward the universal donor cell generation by manipulating HLA expression in therapeutic T cells.
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