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An Optimized Editing Approach for Wheat Genes by Improving sgRNA Design and Transformation Strategies.
Hexaploid wheat has a large genome, making it difficult for transgenes to produce phenotypes due to gene redundancy and tight linkage among genes. Multiple gene copies typically necessitate multiple targeting events during gene editing, followed by several generations of self-crossing to achieve homozygous genotypes. The high cost of transgenesis in wheat is another issue, which hinders the easy availability of gene-edited materials in wheat. In this study, we developed a comprehensive approach to improve wheat gene editing efficiency. First, we established a protoplast-based system to evaluate the relative efficiency of gene editing targets, which enabled the rapid and effective selection of optimal sgRNAs. We then compared two transformation strategies: biolistic bombardment and Agrobacterium-mediated transformation for generating edited wheat lines. Although biolistic bombardment showed higher initial editing efficiency, Agrobacterium-mediated transformation proved more effective for obtaining homozygous mutants. Notably, we discovered that deploying the same sgRNA through different vectors enhanced editing efficiency, whereas overlapping but distinct sgRNAs exhibited interference effects. Finally, we optimized the VITF-edit (virus-induced transgene free editing) technique using BSMV delivery to establish a relatively simple and easily applied wheat gene editing method for general laboratories.
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Features affecting Cas9-induced editing efficiency and patterns in tomato: evidence from a large CRISPR dataset.
CRISPR/Cas9 is a cornerstone of plant genome editing, yet the determinants of editing efficiency for a given single-guide RNAs (sgRNAs) and DNA double-strand break (DSB) repair outcomes remain poorly understood, particularly in plants. Here, we generated a large experimental dataset comprising 420 sgRNAs targeting promoters, exons, and introns of 137 genes in tomato protoplasts, and quantified editing efficiency and repair footprints together with chromatin accessibility and transcriptional state in the same cellular context. Editing efficiency was consistently higher at targets in accessible chromatin and modestly higher in promoters and introns than in exons, whereas transcriptional activity had no detectable effect. Editing efficiencies were more similar among sgRNAs targeting the same gene than among different genes, revealing a local genomic influence on Cas9 activity. A distinct subset of sgRNAs achieved near-complete editing and produced characteristic repair footprints dominated by long deletions with extended microhomology tracts, indicative of microhomology-mediated end joining (MMEJ), resembling patterns associated with high-efficiency guides in human cells, and suggesting conserved sequence-driven repair biases across species. In contrast, widely used human-trained prediction models failed to accurately rank sgRNA performance in plants, highlighting the limits of cross-species predictability. Together, this dataset provides a resource for improving guide design and mechanistic understanding of plant DNA repair.
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Single-step generation of homozygous knockout/knock-in individuals in an extremotolerant parthenogenetic tardigrade using DIPA-CRISPR.
Tardigrades are small aquatic invertebrates known for their remarkable tolerance to diverse extreme stresses. To elucidate the in vivo mechanisms underlying this extraordinary resilience, methods for genetically manipulating tardigrades have long been desired. Despite our prior success in somatic cell gene editing by microinjecting Cas9 ribonucleoproteins (RNPs) into the body cavity of tardigrades, the generation of gene-edited individuals remained elusive. In this study, employing an extremotolerant parthenogenetic tardigrade species, Ramazzottius varieornatus, we established conditions that led to the generation of gene-edited tardigrade individuals. Drawing inspiration from the direct parental CRISPR (DIPA-CRISPR) technique employed in several insects, we simply injected a concentrated Cas9 RNP solution into the body cavity of parental females shortly before their initial oviposition. This approach yielded gene-edited G0 progeny. Notably, only a single allele was predominantly detected at the target locus for each G0 individual, indicative of homozygous mutations. By co-injecting single-stranded oligodeoxynucleotides (ssODNs) with Cas9 RNPs, we achieved the generation of homozygously knocked-in G0 progeny, and these edited alleles were inherited by G1/G2 progeny. This is the first example of heritable gene editing in the entire phylum of Tardigrada. This establishment of a straightforward method for generating homozygous knockout/knock-in individuals not only facilitates in vivo analyses of the molecular mechanisms underpinning extreme tolerance, but also opens up avenues for exploring various topics, including Evo-Devo, in tardigrades.
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MAGIK: A rapid and efficient method to create lineage-specific reporters in human pluripotent stem cells.
Precise insertion of fluorescent proteins into lineage-specific genes in human pluripotent stem cells (hPSCs) presents challenges due to low knockin efficiency and difficulties in isolating targeted cells. To overcome these hurdles, we present the modified mRNA (ModRNA)-based Activation for Gene Insertion and Knockin (MAGIK) method. MAGIK operates in two steps: first, it uses a Cas9-2A-p53DD modRNA with a mini-donor plasmid (without a drug selection cassette) to significantly enhance efficiency. Second, a deactivated Cas9 activator modRNA and a 'dead' guide RNA are used to temporarily activate the targeted gene, allowing for live cell sorting of targeted cells. Consequently, MAGIK eliminates the need for drug selection cassettes or labor-intensive single-cell colony screening, expediting precise gene editing. We showed MAGIK can be utilized to insert fluorescent proteins into various genes, including SOX17, NKX6.1, NKX2.5, and PDX1, across multiple hPSC lines. This underscores its robust performance and offers a promising solution for achieving knockin in hPSCs within a significantly shortened time frame.
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Discovery and characterization of sgRNA-sequence-independent DNA cleavage from CRISPR/Cas9 in mouse embryos.
The CRISPR/Cas9 system can induce off-target effects in programmed gene editing, but there have been few reports on cleavage detection and their affection in embryo development. To study these events, sgRNAs with different off-target rates were designed and compared after micro-injected into mouse zygotes, and γH2AX was used for DNA cleavage sites analysis by immunostaining and CUT&Tag. Although the low off-target sgRNA were usually selected for production gene editing animals, γH2AX immunofluorescence indicated that there was a relative DSB peak at 15 h after Cas9 system injection, and the number of γH2AX foci at the peak was significantly higher in the low off-target sgRNA-injected group than in the control group. Further, the result of CUT&Tag sequencing analysis showed more double-strand breaks (DSBs) related sequences were detected in low off-target sgRNA-injected group than control and the distribution of DSB related sequences had no chromosome specificity. Gene Ontology (GO) annotation analysis of the DSB related sequences showed that these sequences were mainly concentrated at genes associated with some important biological processes, molecular functions, and cell components. In a conclusion, there are many sgRNA-sequence-independent DSBs in early mouse embryos when the Cas9 system is used for gene editing and the DSB related sequence could be detected and characterized in the genome. These results and method should also be considered in using or optimizing the Cas9 system.
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Activation of the cGAS-STING pathway combined with CRISPR-Cas9 gene editing triggering long-term immunotherapy.
Effective activation of cGAS-STING pathway combined with immune checkpoint blockade (ICB) within the immunosuppressive tumor microenvironment to induce stronger immune responsiveness yet remains challenging. CRISPR-Cas9 gene editing technology, which offers the benefits of permanence and irreversibility, could recognize the target genome sequence with sgRNA (Guide RNA) and guide the Cas9 protease to knock down the target gene. Herein, a nanoplatform (HMnMPH) for dual activation of cGAS-STING pathway in combination with CRISPR-Cas9 gene editing to silence programmed death ligand 1 (PD-L1) to trigger long-term immunotherapy was reported. The HMnMPH consists of hollow manganese dioxide (HMn) loaded with STING agonist (MSA-2) and CRISPR-Cas9/sg-PD-L1 plasmid with further modification of hyaluronic acid (HA). In acidic and GSH overexpressed tumor environment, HMnPMH was degraded to release large amounts of Mn ions and STING agonists, strongly and persistently activating the cGAS-STING pathway to promote the release of type I interferon and pro-inflammatory factors. Meanwhile, the released CRISPR-Cas9 plasmid could knockdown the PD-L1 immune checkpoint and restart immunosuppressive T cells to differentiate into cytotoxic T lymphocytes significantly, which reduced the activity of primary and distal tumors and demonstrated a long-term immune memory effect on distal tumors.
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A versatile CRISPR/Cas9 system off-target prediction tool using language model.
Genome editing with the CRISPR/Cas9 system has revolutionized life and medical sciences, particularly in treating monogenic genetic diseases by enabling long-term therapeutic effects from a single intervention. However, the CRISPR/Cas9 system can tolerate mismatches and DNA/RNA bulges at target sites, leading to unintended off-target effects that pose challenges for gene-editing therapy development. Existing high-throughput detection and in silico prediction methods are often limited to specifically designed single guide RNAs (sgRNAs) and perform poorly on unseen sequences. To address these limitations, we introduce CCLMoff, a deep learning framework for off-target prediction that incorporates a pretrained RNA language model from RNAcentral. CCLMoff captures mutual sequence information between sgRNAs and target sites and is trained on a comprehensive, updated dataset. This approach enables accurate off-target identification and strong generalization across diverse NGS-based detection datasets. Model interpretation reveals the biological importance of the seed region, underscoring CCLMoff's analytical capabilities. The development of CCLMoff lays the foundation for a comprehensive, end-to-end sgRNA design platform, enhancing both the precision and efficiency of CRISPR/Cas9-based therapeutics. CCLMoff is a versatile tool and is publicly available at github.com/duwa2/CCLMoff .
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Exosome-mediated delivery of CRISPR/Cas9 for targeting of oncogenic KrasG12D in pancreatic cancer.
CRISPR/Cas9 is a promising technology for gene editing. To date, intracellular delivery vehicles for CRISPR/Cas9 are limited by issues of immunogenicity, restricted packaging capacity, and low tolerance. Here, we report an alternative, nonviral delivery system for CRISPR/Cas9 based on engineered exosomes. We show that non-autologous exosomes can encapsulate CRISPR/Cas9 plasmid DNA via commonly available transfection reagents and can be delivered to recipient cancer cells to induce targeted gene deletion. As a proof-of-principle, we demonstrate that exosomes loaded with CRISPR/Cas9 can target the mutant Kras G12D oncogenic allele in pancreatic cancer cells to suppress proliferation and inhibit tumor growth in syngeneic subcutaneous and orthotopic models of pancreatic cancer. Exosomes may thus be a promising delivery platform for CRISPR/Cas9 gene editing for targeted therapies.
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An engineered hypercompact CRISPR-Cas12f system with boosted gene-editing activity.
Compact CRISPR-Cas systems offer versatile treatment options for genetic disorders, but their application is often limited by modest gene-editing activity. Here we present enAsCas12f, an engineered RNA-guided DNA endonuclease up to 11.3-fold more potent than its parent protein, AsCas12f, and one-third of the size of SpCas9. enAsCas12f shows higher DNA cleavage activity than wild-type AsCas12f in vitro and functions broadly in human cells, delivering up to 69.8% insertions and deletions at user-specified genomic loci. Minimal off-target editing is observed with enAsCas12f, suggesting that boosted on-target activity does not impair genome-wide specificity. We determine the cryo-electron microscopy (cryo-EM) structure of the AsCas12f-sgRNA-DNA complex at a resolution of 2.9 Å, which reveals dimerization-mediated substrate recognition and cleavage. Structure-guided single guide RNA (sgRNA) engineering leads to sgRNA-v2, which is 33% shorter than the full-length sgRNA, but with on par activity. Together, the engineered hypercompact AsCas12f system enables robust and faithful gene editing in mammalian cells.
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An Introduced RNA-Only Approach for Plasmid Curing via the CRISPR-Cpf1 System in Saccharomyces cerevisiae.
The CRISPR-Cas system has been widely used for genome editing due to its convenience, simplicity and flexibility. Using a plasmid-carrying Cas protein and crRNA or sgRNA expression cassettes is an efficient strategy in the CRISPR-Cas genome editing system. However, the plasmid remains in the cells after genome editing. Development of general plasmid-curing strategies is necessary. Based on our previous CRISPR-Cpf1 genome-editing system in Saccharomyces cerevisiae, the crRNA, designed for the replication origin of the CRISPR-Cpf1 plasmid, and the ssDNA, as a template for homologous recombination, were introduced for plasmid curing. The efficiency of the plasmid curing was 96 ± 4%. In addition, we further simplified the plasmid curing system by transforming only one crRNA into S. cerevisiae, and the curing efficiency was about 70%. In summary, we have developed a CRISPR-mediated plasmid-curing system. The RNA-only plasmid curing system is fast and easy. This plasmid curing strategy can be applied in broad hosts by designing crRNA specific for the replication origin of the plasmid. The plasmid curing system via CRISPR-Cas editing technology can be applied to produce traceless products without foreign genes and to perform iterative processes in multiple rounds of genome editing.
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