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p38 MAPK inhibition safeguards genome integrity during gene editing of hematopoietic stem and progenitor cells.
Ex vivo genome editing of hematopoietic stem and progenitor cells (HSPCs) holds significant therapeutic potential but remains constrained by genotoxic risks associated with nuclease-induced DNA double-strand breaks, DNA donor template delivery and sensing, and proliferation-induced stress during ex vivo manipulation. These processes can lead to chromosomal instability, large on-target deletions, donor mis-integration, off-target events, and impaired long-term stem cell function, raising safety concerns for clinical translation. Here, we evaluate the impact of transient p38 MAPK inhibition on genomic integrity during clinically relevant CRISPR-Cas9 editing and show that this intervention attenuates ex vivo culture-associated stress without increasing detectable genotoxic outcomes. Comprehensive genotoxicity analyses, including quantification of large on-target deletions, adeno-associated viral vector mis-integration, and CAST-seq mapping of translocations, reveal no measurable differences in gene editing-associated structural alterations upon p38 MAPK inhibition, while micronuclei were significantly reduced. Importantly, long-term xenotransplantation followed by whole-exome sequencing shows that p38i-treated HSPCs display a reduced mutational burden without evidence of increased genomic alterations. Collectively, these findings identify transient p38 MAPK inhibition as a strategy to improve the quality and long-term fitness of gene-edited HSPCs without detectable adverse effects on the genome editing outcomes, supporting its further evaluation as a refinement to clinically relevant CRISPR-Cas9 editing workflows.
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Comparative analysis of genome editing systems, Cas9 and BE3, in silkworms.
The clustered regularly interspaced short palindromic repeats (CRISPR) system and the Cas9-derived proteins have been applied to genome editing in various organisms. Traditional Cas9 is typically used to knockout genes or specific DNA fragments based on the generation of double-stranded breaks, whereas nCas9 and dCas9 are fused with effectors to perform base pair transitions or epigenetic modification and regulation. However, this system has off-target effects and can cause genomic structure variations. Here, we comparatively analyzed Cas9 and BE3, an initial base editor based on the nCas9 fusion protein, in silkworms. Our results showed that base editing was superior to Cas9 in silkworm cultured cells. BE3 introduced accurate termination codons, whereas Cas9 did not. Moreover, Cas9 induced chromosome translocation, chromosome fragment repetition, and chromosome fragment deletion, with the deletion frequency reaching up to 4.29%. BE3 was not able to induce these changes in our study. Furthermore, Cas9-derived proteins blocked ribosome advance and mRNA transcription for 9 days, with a 9.40% repression effect by combining with double-stranded DNA when single guide RNAs were targeted in the coding region in silkworms. Overall, our findings established a strategy for choosing suitable editing tools for various applications in different organisms.
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Occurrence and applications of CRISPR-Cas systems in bifidobacteria.
Bifidobacterium is a key member of the human gut microbiota, and many strains are widely used as probiotics due to their health-promoting properties. Despite growing interest, genetic studies in Bifidobacterium have been relatively limited, primarily due to the lack of available genome editing tools. Recent advances in genomics and CRISPR-Cas systems provide opportunities for targeted genome modification in this genus. In this review, we provide an overview of the occurrence, diversity, and distribution of CRISPR-Cas systems across Bifidobacterium species and examine the editing tools developed and implemented to date. We also highlight practical challenges such as strain variability and low transformation efficiency and introduce future avenues of research such as large-payload insertion and in situ editing. Expanding the genetic toolbox for Bifidobacterium will broaden our understanding of this important genus and enable the development of next-generation probiotics.
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Genome editing in the fall armyworm, Spodoptera frugiperda: Multiple sgRNA/Cas9 method for identification of knockouts in one generation.
The CRISPR/Cas9 system is an efficient genome editing method that can be used in functional genomics research. The fall armyworm, Spodoptera frugiperda, is a serious agricultural pest that has spread over most of the world. However, very little information is available on functional genomics for this insect. We performed CRISPR/Cas9-mediated site-specific mutagenesis of three target genes: two marker genes [Biogenesis of lysosome-related organelles complex 1 subunit 2 (BLOS2) and tryptophan 2, 3-dioxygenase (TO)], and a developmental gene, E93 (a key ecdysone-induced transcription factor that promotes adult development). The knockouts (KO) of BLOS2, TO and E93 induced translucent mosaic integument, olive eye color, and larval-pupal intermediate phenotypes, respectively. Sequencing RNA isolated from wild-type and E93 KO insects showed that E93 promotes adult development by influencing the expression of the genes coding for transcription factor, Krüppel homolog 1, the pupal specifier, Broad-Complex, serine proteases, and heat shock proteins. Often, gene-edited insects display mosaicism in which only a fraction of the cells are edited as intended, and establishing a homozygous line is both costly and time-consuming. To overcome these limitations, a method to completely KO the target gene in S. frugiperda by injecting the Cas9 protein and multiple sgRNAs targeting one exon of the E93 gene into embryos was developed. Ten percent of the G0 larvae exhibited larval-pupal intermediates. The mutations were confirmed by T7E1 assay, and the mutation frequency was determined as >80%. Complete KO of the E93 gene was achieved in one generation using the multiple sgRNA method, demonstrating a powerful approach to improve genome editing in lepidopteran and other non-model insects.
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Efficient generation of targeted large insertions by microinjection into two-cell-stage mouse embryos.
Rapid, efficient generation of knock-in mice with targeted large insertions remains a major hurdle in mouse genetics. Here, we describe two-cell homologous recombination (2C-HR)-CRISPR, a highly efficient gene-editing method based on introducing CRISPR reagents into embryos at the two-cell stage, which takes advantage of the open chromatin structure and the likely increase in homologous-recombination efficiency during the long G2 phase. Combining 2C-HR-CRISPR with a modified biotin-streptavidin approach to localize repair templates to target sites, we achieved a more-than-tenfold increase (up to 95%) in knock-in efficiency over standard methods. We targeted 20 endogenous genes expressed in blastocysts with fluorescent reporters and generated reporter mouse lines. We also generated triple-color blastocysts with all three lineages differentially labeled, as well as embryos carrying the two-component auxin-inducible degradation system for probing protein function. We suggest that 2C-HR-CRISPR is superior to random transgenesis or standard genome-editing protocols, because it ensures highly efficient insertions at endogenous loci and defined 'safe harbor' sites.
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The versatile electric condition in mouse embryos for genome editing using a three-step square-wave pulse electroporator.
Technique for Animal Knockout system by Electroporation (TAKE) is a simple and efficient method to generate genetically modified (GM) mice using the clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein 9 (Cas9) systems. To reinforce the versatility of electroporation used for gene editing in mice, the electric condition was optimized for vitrified-warmed mouse embryos, and applied to the fresh embryos from widely used inbred strains (C57BL/6NCr, BALB/cCrSlc, FVB/NJcl, and C3H/HeJJcl). The electric pulse settings (poring pulse: voltage, 150 V; pulse width, 1.0 ms; pulse interval, 50 ms; number of pulses, +4; transfer pulse: voltage, 20 V; pulse width, 50 ms; pulse interval, 50 ms; number of pulses, ±5) were optimal for vitrified-warmed mouse embryos, which could efficiently deliver the gRNA/Cas9 complex into the zygotes without zona pellucida thinning process and edit the target locus. These electric condition efficiently generated GM mice in widely used inbred mouse strains. In addition, electroporation using the electrode with a 5 mm gap could introduce more than 100 embryos within 5 min without specific pretreatment and sophisticated technical skills, such as microinjection, and exhibited a high developmental rate of embryos and genome-editing efficiency in the generated offspring, leading to the rapid and efficient generation of genome editing mice. The electric condition used in this study is highly versatile and can contribute to understanding human diseases and gene functions by generating GM mice more easily and efficiently.
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Gene editing and CRISPR-dependent homology-mediated end joining.
Gene editing is the intentional modification of a genetic locus in a living cell and is used for two general applications of great importance and wide interest. One is the inactivation of genes ('knockouts'), a process utilized to delineate the loss-of-function phenotype(s) of a particular gene. The second application ('knock-ins') is essentially the process of gene therapy, which predominately involves correcting a pre-existing mutated allele(s) of a gene back to wild-type to ameliorate some pathological phenotype associated with the mutation. Importantly, although these applications are conceptually exact reciprocal opposites of one another, they are achieved via mechanistically different pathways. In the case of knockouts, breakage (usually in the form of double-stranded breaks) of the chromosomal DNA at the site of targeting is used to engage a repair process (nonhomologous end joining) that is error prone. The ensuing repair frequently results in insertions/deletions at the cleavage site, which, in turn, results in out-of-frame mutations and, hence, a knockout of the gene in question. In the case of knock-ins, breakage (again, usually in the form of double-stranded breaks) of the DNA is used to engage a repair process (homology-dependent repair/recombination) in which homologous sequences between an incoming donor DNA (containing new genetic information) and the chromosomal DNA are exchanged. Although homology-directed repair was known to predominate in bacteria and lower eukaryotes, the competing process of nonhomologous end joining predominates in higher eukaryotes and was presumed to prevent the use of knock-in gene editing in human somatic cells in culture. A series of molecular and technical advances disproved this notion but still resulted in a process that was cumbersome, labor intensive, highly inefficient and slow. In 2013, however, a new RNA-programmable nuclease, CRISPR-Cas9 was described that has revolutionized the field and made gene editing accessible to anyone with even a rudimentary knowledge of molecular biology. Thus, gene editing in a wide variety of model organisms, as well as human somatic cells in culture, has become not only extremely feasible but also extremely facile, and it harbingers a golden age for directed mutagenesis, directed evolution and improvements in gene therapy.
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Optimization of gene knockout approaches and sgRNA selection in hPSCs with inducible Cas9 expression.
CRISPR/Cas9 has been extensively used for gene knockout, enabling functional studies of genetic loss-of-function in human pluripotent stem cells (hPSCs). However, commonly used Cas9 systems usually exhibit limited and variable efficiencies, and identifying single-guide RNAs (sgRNAs) with high cleavage activity-while avoiding ineffective ones-remains a major challenge. To address these issues, we generated a doxycycline-inducible spCas9-expressing hPSCs (hPSCs-iCas9) line and developed it into an optimized gene knockout system through systematically refining critical parameters. Through this optimization, the system achieved stable INDELs (Insertions and Deletions) efficiencies of 82-93% for single-gene knockouts, over 80% for double-genes knockouts, and up to 37.5% homozygous knockout efficiency for large DNA fragment deletions. Moreover, using this optimized system, we precisely evaluated three widely used gRNA scoring algorithms and integrated Western blotting to rapidly identify the ineffective sgRNA. As a result, among the tested algorithms, Benchling provided the most accurate predictions. Notably, we identified an ineffective sgRNA targeting exon 2 of ACE2, where the edited cell pool exhibited 80% INDELs but retained ACE2 protein expression. Together, these findings provide a robust framework for improving gene knockout efficiency in hPSCs and offer practical guidance for reliable sgRNA selection in gene editing experiments.
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Developmental competence of porcine genome-edited zygotes.
Genome editing in pigs has tremendous practical applications for biomedicine. The advent of genome editing technology, with its use of site-specific nucleases-including ZFNs, TALENs, and the CRISPR/Cas9 system-has popularized targeted zygote genome editing via one-step microinjection in several mammalian species. Here, we review methods to optimize the developmental competence of genome-edited porcine embryos and strategies to improve the zygote genome-editing efficiency in pigs.
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Comparison of Gene-Editing Approaches for Severe Congenital Neutropenia-Causing Mutations in the ELANE Gene.
Safety considerations for gene therapies of inherited preleukemia syndromes, including severe congenital neutropenia (CN), are paramount. We compared several strategies for CRISPR/Cas9 gene editing of autosomal-dominant ELANE mutations in CD34+ cells from two CN patients head-to-head. We tested universal and allele-specific ELANE knockout, ELANE mutation correction by homology-directed repair (HDR) with AAV6, and allele-specific HDR with ssODN. All strategies were not toxic, had at least 30% editing, and rescued granulopoiesis in vitro. In contrast to published data, allele-specific indels in the last exon of ELANE also restored granulopoiesis. Moreover, by implementing patient-derived induced pluripotent stem cells for GUIDE-Seq off-target analysis, we established a clinically relevant "personalized" assessment of off-target activity of gene editing on the background of the patient's genome. We found that allele-specific approaches had the most favorable off-target profiles. Taken together, a well-defined head-to-head comparison pipeline for selecting the appropriate gene therapy is essential for diseases, with several gene editing strategies available.
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