CRISPR Journal最新文献

Advances in CRISPR-Cas9-based genome editing have simplified the generation of isogenic induced pluripotent stem cell (iPSC) pairs differing by a single genetic perturbation, minimizing interindividual variability in disease modeling. However, imprecise editing can cause unintended mutations that distort genotype-phenotype interpretation. Here, we investigated the molecular consequences of CRISPR-Cas9 editing in human iPSCs carrying a tandem duplication of the STAG2 locus. Using hybridization capture-based targeted next-generation sequencing of genomic regions surrounding Cas9-induced double-strand breaks, we identified inversions, plasmid DNA integrations, and complex structural rearrangements that had been missed by conventional PCR-based genotyping. The main advance of our study is the application of a practical hybridization capture-based workflow for resolving such hidden editing outcomes during the generation of isogenic iPSC models. In the selected clones subjected to detailed analysis, targeted sequencing enabled us to revise the initial interpretation of editing outcomes and resolve discrepancies between genotypic and transcriptomic data. Our results demonstrate that complex on-target alterations can remain undetected by conventional PCR-based genotyping and show that hybridization capture-based targeted sequencing can serve as a practical quality-control approach for the characterization of edited iPSC clones.

The emergence of CRISPR base editors signifies a pivotal shift in genome editing, moving beyond the "cut-and-paste" approach into an era of precise "chemical rewriting." While early editors including CBEs and ABEs enabled efficient base transitions by combining deaminases with Cas9 nickase, their core mechanism limited their application to transition mutations, leaving nearly half of disease-causing transversions unresolved. This review outlines the transformation of DNA glycosylases from unwanted side-effect generators into core drivers of base editor evolution. Through strategic protein engineering, glycosylases have been harnessed to create novel editors that mediate crucial transversion edits, substantially expanding the targeting landscape of precision genome editing. We will examine how these engineered glycosylases achieve programmable DNA rewriting through redirected DNA repair pathways, compare emerging and conventional editors, and explore applications in genetic therapy and functional genomics while addressing ongoing challenges in specificity and delivery. Finally, we envision next-generation editors equipped with artificial intelligence-designed glycosylases capable of spatiotemporally controlled genome manipulation.

Prime editing (PE) enables precise insertions, deletions, and substitutions without double-strand breaks, but editing efficiency remains highly variable. To identify modulators of PE outcomes, we screened 569 compounds targeting DNA damage and repair pathways in K562 cells using targeted next-generation sequencing. PARP inhibitors selectively enhanced PE-mediated insertions at the HEK3 locus by more than 2-fold, without improving substitution or deletion editing. Genetic loss of PARP1 reproduced this phenotype, identifying PARP1 as the principal mediator. PARP1 inhibition enhanced insertion efficiency across multiple PE architectures, insertion sizes, and cellular backgrounds, including primary human T cells, and promoted more complete incorporation of pegRNA-encoded sequence changes without increasing indels or scaffold incorporation. However, this effect was dependent on genomic locus and reverse transcription template configuration. Together, these findings identify PARP1 as a context-dependent constraint on insertional PE.

Ribonucleic acid (RNA)-targeting clustered regularly interspaced short palindromic repeats-CRISPR-associated (CRISPR-Cas) systems enable modulation of gene expression without permanent genome modification, making them useful for sensitive cell types such as neurons. While CRISPR-Cas technologies have been most extensively applied and validated in primary hippocampal and cortical neurons, their use in sensory neurons remains largely unexplored. Sensory neurons are an established cellular model for studying axon growth and regeneration, pain mechanisms, sensory transduction, and neuron-environment interactions. Here, we evaluated the performance of compact RNA-targeting CRISPR-Cas effectors Cas7-11S, hfCas13X, and hfCas13d in primary rat sensory neurons in culture. Using an endogenous mRNA as the target, we compared knockdown efficiency and assessed the effects of CRISPR-Cas expression on neuronal health. The systems showed distinct differences in performance, with Cas7-11S inducing toxicity, hfCas13X showing minimal knockdown, and hfCas13d providing robust gene silencing with minimal adverse effects on neuronal health. These findings identify hfCas13d as the most effective and well-tolerated RNA-targeting CRISPR-Cas tool for sensory neurons and provide important insight into its suitability for neuroscience research and potential therapeutic applications.

Programmable large-fragment genome integration using prime-editing-coupled serine integrases, such as PASTE and PASSIGE, remains constrained by the limited activity of wild-type Bxb1 (WT Bxb1) in mammalian cells. Recently, the AI-guided protein engineering framework EVOLVEpro enabled efficient identification of functional protein variants from limited experimental sampling and nominated epBxb1(T166R) as a highly active Bxb1 variant in episomal plasmid-to-plasmid recombination screens. Here, we systematically benchmarked epBxb1 (T166R) against WT Bxb1 and the previously validated high-activity eeBxb1 (V74A/E229K/V375I) variant in genome-integrated reporter and endogenous-locus PASSIGE assays at three well-characterized benchmarking loci (AAVS1, CCR5, and ACTB). epBxb1 showed reduced and variable transferability, with no detectable advantage over WT Bxb1 in the genome-integrated reporter system or at the AAVS1 safe-harbor locus, but produced modest, locus-dependent improvements at CCR5 and ACTB, with 1.89-fold and 1.58-fold increases, respectively. By contrast, eeBxb1 consistently showed superior activity, achieving up to ∼12-fold improvement over WT Bxb1. Four additional rounds of EVOLVEpro-guided optimization on the eeBxb1-BPNLS scaffold identified no reproducibly improved variant among 40 tested single-amino-acid substitutions. These results indicate reduced and genomic context-dependent transferability of EVOLVEpro-nominated Bxb1 variants and highlight the importance of application-matched genomic benchmarking when implementing AI-guided protein optimization for therapeutic genome writing.

Dog allergy is a prevalent IgE-mediated condition, with the salivary lipocalin Can f 1 accounting for the majority of dog-specific IgE reactivity in sensitized individuals. Existing strategies for managing dog allergy primarily rely on modulating host immune responses and do not address allergen production at its source. Here, we report the generation of genetically engineered dogs lacking Can f 1. Using CRISPR-Cas9 editing and somatic cell nuclear transfer, we produced two healthy beagle puppies carrying a frameshift mutation in exon 1 of the Can f 1 gene. Salivary and hair/dander analysis demonstrated the absence of Can f 1 protein in the edited dogs. Skin prick testing in a sensitized individual demonstrated robust IgE-mediated reactivity to control dog extracts but no detectable response to extracts from the edited dogs. Together, these findings demonstrate that targeted genetic knockout of the major dog allergen is compatible with canine development and can abolish the IgE-mediated allergic response, supporting the feasibility of a gene-based approach to reducing canine allergenicity.

Base editing enables precise genome modifications without introducing DNA double-strand breaks. Using Streptococcus pyogenes Cas9 as a prototype, we previously developed a modular base editing platform in which the deaminase is recruited by an RNA aptamer engineered into the gRNA, thereby separating sequence recognition from base modification. Here, we expanded this modular base editor toolbox by engineering Staphylococcus aureus Cas9 (SaCas9) in combination with various vertebrate effectors derived from activation induced cytidine deaminase (AID) and apolipoprotein B mRNA editing enzyme, catalytic subunit 1 (APOBEC1) orthologs, from bat, lizard, human, and rat. Moreover, we adopted the SaCas9 variants with different protospacer adjacent motif requirements. These base editors generally showed high editing efficiency with low on-target indel formation and low-to-undetectable off-target activities. Quantitative and qualitative differences in editing occur among the base editors when applied to diverse loci, allowing sequence-specific optimization. Together, our study demonstrates the effectiveness of the SaCas9 modular base editors, the robustness of the platform's modularity, and its feasibility for convenient screening of target-specific base editors.

Since the advent of Cas9-based CRISPR technologies in 2012, there has been a remarkable growth in genome editing research, literature, applications, and translational impact. Much of this research has been fueled by the global dissemination of CRISPR plasmids through nonprofit distribution by Addgene, as both a repository and distributor of enabling biological material. Recently, key milestones have been reached, with over 20,000 plasmids deposited by over 1,000 labs, being distributed over 300,000 times globally. The driving trends reflect multidimensional diversification in terms of effectors (Cas9, Cas12 and beyond), editing modalities (base editing, prime editing, epigenetic modification, CRISPRi/a), and deployment across phylogenetic groups (mammalian, bacterial, plant, yeast, insects, and more). Noteworthy, guide RNA and HDR templates account for the bulk of deposits, while cloning backbones are the most requested, and lentiviral plasmids comprise the majority of expression material. The data reflect a continued diversification of the CRISPR-based toolbox, robust interest in genome editing applications across the tree of life, maturation in terms of adoption, and rising relative distribution beyond the USA and China, with Addgene continuing to play a critical role in access to equitable and disruptive technologies.

Craniosynostosis is a rare congenital bone condition where skull sutures fuse prematurely and is linked to mutations in over 60 genes. Generating mutation-specific in vitro models allows investigation of craniosynostosis-associated mutations without the need for patient-derived material or transgenic gene expression. Here, we developed a human in vitro disease model with the CRISPR-Cas9 prime editing variant, using an immortalized TERT-immortalized mesenchymal bone marrow-derived stem (MSC-TERT) cell line with osteogenic potential. MSC-TERT cells showed a higher resistance to prime editing, compared with HEK293FT cells. Addition of dnMLH1 and epegRNAs resulted in higher editing efficiencies in HEK293FT cells, but not in MSC-TERT cells. Prime editing efficiency varied between targeted loci and was found to be more efficient in nonadherent cells compared with adherent cells. Prime editing continued over 4 days in an isolated nonadherent HEK293FT culture. Our results present a foundation on the use of prime editing to establish FGFR2 mutation-specific in vitro models and their application in MSC-TERT cells.