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Suppression of unwanted CRISPR-Cas9 editing by co-administration of catalytically inactivating truncated guide RNAs.
CRISPR-Cas9 nucleases are powerful genome engineering tools, but unwanted cleavage at off-target and previously edited sites remains a major concern. Numerous strategies to reduce unwanted cleavage have been devised, but all are imperfect. Here, we report that off-target sites can be shielded from the active Cas9•single guide RNA (sgRNA) complex through the co-administration of dead-RNAs (dRNAs), truncated guide RNAs that direct Cas9 binding but not cleavage. dRNAs can effectively suppress a wide-range of off-targets with minimal optimization while preserving on-target editing, and they can be multiplexed to suppress several off-targets simultaneously. dRNAs can be combined with high-specificity Cas9 variants, which often do not eliminate all unwanted editing. Moreover, dRNAs can prevent cleavage of homology-directed repair (HDR)-corrected sites, facilitating scarless editing by eliminating the need for blocking mutations. Thus, we enable precise genome editing by establishing a flexible approach for suppressing unwanted editing of both off-targets and HDR-corrected sites.
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A tool for CRISPR-Cas9 sgRNA evaluation based on computational models of gene expression.
BACKGROUND: CRISPR is widely used to silence genes by inducing mutations expected to nullify their expression. While numerous computational tools have been developed to design single-guide RNAs (sgRNAs) with high cutting efficiency and minimal off-target effects, only a few tools focus specifically on predicting gene knockouts following CRISPR. These tools consider factors like conservation, amino acid composition, and frameshift likelihood. However, they neglect the impact of CRISPR on gene expression, which can dramatically affect the success of CRISPR-induced gene silencing attempts. Furthermore, information regarding gene expression can be useful even when the objective is not to silence a gene. Therefore, a tool that considers gene expression when predicting CRISPR outcomes is lacking. RESULTS: We developed EXPosition, the first computational tool that combines models predicting gene knockouts after CRISPR with models that forecast gene expression, offering more accurate predictions of gene knockout outcomes. EXPosition leverages deep-learning models to predict key steps in gene expression: transcription, splicing, and translation initiation. We showed our tool performs better at predicting gene knockout than existing tools across 6 datasets, 4 cell types and ~207k sgRNAs. We also validated our gene expression models using the ClinVar dataset by showing enrichment of pathogenic mutations in high-scoring mutations according to our models. CONCLUSIONS: We believe EXPosition will enhance both the efficiency and accuracy of genome editing projects, by directly predicting CRISPR's effect on various aspects of gene expression. EXPosition is available at http://www.cs.tau.ac.il/~tamirtul/EXPosition . The source code is available at https://github.com/shaicoh3n/EXPosition .
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Generation of genetically modified mice using CRISPR/Cas9 and haploid embryonic stem cell systems.
With the development of high-throughput sequencing technology in the post-genomic era, researchers have concentrated their efforts on elucidating the relationships between genes and their corresponding functions. Recently, important progress has been achieved in the generation of genetically modified mice based on CRISPR/Cas9 and haploid embryonic stem cell (haESC) approaches, which provide new platforms for gene function analysis, human disease modeling, and gene therapy. Here, we review the CRISPR/Cas9 and haESC technology for the generation of genetically modified mice and discuss the key challenges in the application of these approaches.
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APOBEC Reporter Systems for Evaluating diNucleotide Editing Levels.
Precision genome editing has become a reality with the discovery of base editors. Cytosine base editor (CBE) technologies are improving rapidly but are mostly optimized for TC dinucleotide targets. Here, we report the development and implementation of APOBEC Reporter Systems for Evaluating diNucleotide Editing Levels (ARSENEL) in living cells. The ARSENEL panel is comprised of four constructs that quantitatively report editing of each of the four dinucleotide motifs (AC/CC/GC/TC) through real-time accumulation of eGFP fluorescence. Editing rates of APOBEC3Bctd and AIDΔC CBEs reflect established mechanistic preferences with intrinsic biases to TC and GC, respectively. Twelve different (new and established) base editors are tested here using this system with a full-length APOBEC3B CBE showing the greatest on-target TC specificity and an APOBEC3A construct showing the highest editing efficiency. In addition, ARSENEL enables real-time assessment of natural and synthetic APOBEC inhibitors with the most potent to-date being the large subunit of the Epstein-Barr virus ribonucleotide reductase. These reporters have the potential to play important roles in research and development as precision genome engineering technologies progress toward achieving maximal specificity and efficiency.
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An unbiased method for evaluating the genome-wide specificity of base editors in rice.
Base editors can achieve targeted genomic base conversion. However, the off-target issue is one of the major concerns in their application. Whole-genome sequencing (WGS) at the individual level can provide direct information on genome-wide specificity, but it is difficult to distinguish true off-target single-nucleotide variants (SNVs) induced by base editors from background variation. Here we describe an unbiased WGS method for evaluating the specificity of base editors in rice. In this protocol, we describe the experimental design and provide details of vector construction, rice transformation and tissue culture, as well as a comprehensive WGS data analysis pipeline for overcoming two related core problems in various plant species: high background mutation rates and the heterogeneity of examined populations. Using this protocol, researchers can straightforwardly and accurately assess the genome-wide specificity of base editors and other genome editing tools in 12-15 weeks.
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QTL editing confers opposing yield performance in different rice varieties.
Grain yield is one of the most important and complex trait for genetic improvement in crops; it is known to be controlled by a number of genes known as quantitative trait loci (QTLs). In the past decade, many yield-contributing QTLs have been identified in crops. However, it remains unclear whether those QTLs confer the same yield performance in different genetic backgrounds. Here, we performed CRISPR/Cas9-mediated QTL editing in five widely-cultivated rice varieties and revealed that the same QTL can have diverse, even opposing, effects on grain yield in different genetic backgrounds.
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CRISPR-Cas9 in leukemia immunotherapy: precision engineering of CAR-T cells and tumor-microenvironment modulation.
Together with base editors, prime editors, and alternative nuclease platforms, CRISPR-Cas9 technology has transformed the field of genetic engineering by providing unprecedented precision in genome editing and creating new opportunities for therapeutic interventions. This technology comprises a versatile genome-editing toolkit for achieving a variety of therapeutic objectives. In the context of leukemia, a group of life-threatening hematologic malignancies, CRISPR-Cas9 has emerged as a transformative tool for immunotherapy. By enabling targeted modifications of immune cells, such as T cells, this technology enhances their ability to recognize and eradicate leukemic cells. CRISPR-Cas9 facilitates the disruption of immune checkpoint inhibitors, the insertion of chimeric antigen receptors (CARs), and the correction of genetic mutations that drive leukemia progression. These advancements have led to the development of more potent and personalized immunotherapies, such as CAR-T cell therapies, with improved efficacy and reduced off-target effects. Moreover, CRISPR-Cas9 allows researchers to model leukemia in vitro, providing deeper insights into disease mechanisms and accelerating the discovery of novel therapeutic targets. Despite challenges such as delivery efficiency and potential immunogenicity, the integration of CRISPR-Cas9 into leukemia immunotherapy represents a paradigm shift, offering hope for durable remissions and potentially curative outcomes. As clinical trials progress, this technology promises to redefine the standard of care for leukemia patients, ushering in a new era of precision medicine. This narrative review explores the revolutionary applications of CRISPR-Cas9 in redefining therapeutic strategies for leukemia.
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Efficient genotype-independent cotton genetic transformation and genome editing.
Cotton (Gossypium spp.) is one of the most important fiber crops worldwide. In the last two decades, transgenesis and genome editing have played important roles in cotton improvement. However, genotype dependence is one of the key bottlenecks in generating transgenic and gene-edited cotton plants through either particle bombardment or Agrobacterium-mediated transformation. Here, we developed a shoot apical meristem (SAM) cell-mediated transformation system (SAMT) that allowed the transformation of recalcitrant cotton genotypes including widely grown upland cotton (Gossypium hirsutum), Sea island cotton (Gossypium barbadense), and Asiatic cotton (Gossypium arboreum). Through SAMT, we successfully introduced two foreign genes, GFP and RUBY, into SAM cells of some recalcitrant cotton genotypes. Within 2-3 months, transgenic adventitious shoots generated from the axillary meristem zone could be recovered and grown into whole cotton plants. The GFP fluorescent signal and betalain accumulation could be observed in various tissues in GFP- and RUBY-positive plants, as well as in their progenies, indicating that the transgenes were stably integrated into the genome and transmitted to the next generation. Furthermore, using SAMT, we successfully generated edited cotton plants with inheritable targeted mutagenesis in the GhPGF and GhRCD1 genes through CRISPR/Cas9-mediated genome editing. In summary, the established SAMT transformation system here in this study bypasses the embryogenesis process during tissue culture in a conventional transformation procedure and significantly accelerates the generation of transgenic and gene-edited plants for genetic improvement of recalcitrant cotton varieties.
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Introducing a Spectrum of Long-Range Genomic Deletions in Human Embryonic Stem Cells Using Type I CRISPR-Cas.
CRISPR-Cas systems enable microbial adaptive immunity and provide eukaryotic genome editing tools. These tools employ a single effector enzyme of type II or V CRISPR to generate RNA-guided, precise genome breaks. Here we demonstrate the feasibility of using type I CRISPR-Cas to effectively introduce a spectrum of long-range chromosomal deletions with a single RNA guide in human embryonic stem cells and HAP1 cells. Type I CRISPR systems rely on the multi-subunit ribonucleoprotein (RNP) complex Cascade to identify DNA targets and on the helicase-nuclease enzyme Cas3 to degrade DNA processively. With RNP delivery of T. fusca Cascade and Cas3, we obtained 13%-60% editing efficiency. Long-range PCR-based and high-throughput-sequencing-based lesion analyses reveal that a variety of deletions, ranging from a few hundred base pairs to 100 kilobases, are created upstream of the target site. These results highlight the potential utility of type I CRISPR-Cas for long-range genome manipulations and deletion screens in eukaryotes.
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CRISPR-Cas9-based mutagenesis frequently provokes on-target mRNA misregulation.
The introduction of insertion-deletions (INDELs) by non-homologous end-joining (NHEJ) pathway underlies the mechanistic basis of CRISPR-Cas9-directed genome editing. Selective gene ablation using CRISPR-Cas9 is achieved by installation of a premature termination codon (PTC) from a frameshift-inducing INDEL that elicits nonsense-mediated decay (NMD) of the mutant mRNA. Here, by examining the mRNA and protein products of CRISPR targeted genes in a cell line panel with presumed gene knockouts, we detect the production of foreign mRNAs or proteins in ~50% of the cell lines. We demonstrate that these aberrant protein products stem from the introduction of INDELs that promote internal ribosomal entry, convert pseudo-mRNAs (alternatively spliced mRNAs with a PTC) into protein encoding molecules, or induce exon skipping by disruption of exon splicing enhancers (ESEs). Our results reveal challenges to manipulating gene expression outcomes using INDEL-based mutagenesis and strategies useful in mitigating their impact on intended genome-editing outcomes.
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