搜索结果：CRISPR gene editing

The recently developed CRISPR-Combo technology enables simultaneous targeted mutagenesis and transcriptional activation in plants. However, its reliance on SpCas9 limits its use at AT-rich genomic loci, such as promoter regions commonly targeted for transcription activation. To overcome this limitation, we explored the usage of Cas12b and iSpyMacCas9 in the CRISPR-Combo architecture for simultaneous genome editing and gene activation. We tested these expanded CRISPR-Combo systems for hormone-free regeneration of rice plants by transcriptional activation of a morphogenic gene, OsBBM1, while knocking out the genes of interest. The Cas12b-Combo system induced mild OsBBM1 upregulation (~3-fold), which did not affect the genome editing efficiency. By contrast, iSpyMacCas9-Combo achieved approximately 12-fold OsBBM1 transcriptional activation, supporting hormone-free regeneration at a high rate (42%). As a result, iSpyMacCas9-Combo conferred higher genome editing efficiency, including improved multiplex editing, than the standard iSpyMacCas9 system, either with or without hormones during rice regeneration. Hence, our data prove iSpyMacCas9-Combo to be a more efficient system for genome editing in rice, especially at low-efficiency target sites, when coupled with OsBBM1 transcriptional activation. These findings establish iSpyMacCas9-Combo as a useful addition to the CRISPR-Combo toolkit, expanding its genomic targeting scope and enabling more efficient genome editing by activation of an appropriate endogenous gene such as OsBBM1 in rice.

The CRISPR-Cas system has evolved into a highly efficient platform for genome editing and programmable gene regulation, demonstrating broad application potential in microbiology, biotechnology, and medicine. However, traditional CRISPR tools typically rely on constitutively active nuclease activity; this constant activation state is prone to off-target effects and cytotoxicity, and lacks precise spatiotemporal regulation in complex biological environments. Therefore, developing strategies to achieve fine-tuned and context-dependent regulation of CRISPR activity has become a critical issue in this field that urgently needs to be addressed. Recent studies have demonstrated that various endogenous and exogenous regulatory modules can modulate the activity of the CRISPR-Cas system at different biological levels. Among these, anti-CRISPR proteins (Acr), which are natural inhibitory factors derived from bacteriophages, can suppress the nuclease activity of Cas proteins at the protein level by directly interfering with their function; The CRISPR interference/activation (CRISPRi/a) system, on the other hand, relies on catalytically inactivated Cas proteins to achieve sequence-specific regulation of target gene transcription; furthermore, quorum sensing (QS) networks dynamically regulate the expression of relevant genes by sensing cell density and environmental signals, thereby influencing the functional state of the CRISPR system at the population level. Based on the aforementioned regulatory mechanisms, this paper provides a comprehensive, literature-based overview of the molecular basis and recent advances in the applications of Acr proteins, the CRISPRi/a system, and QS networks in CRISPR-Cas regulation. Building on this, we propose a hierarchical regulatory framework: QS networks serve as upstream environmental sensing modules that drive CRISPRi/a-mediated programmable transcriptional regulation, while Acr proteins act as downstream rapid-response elements that finely tune CRISPR activity. This multi-tiered regulatory system holds promise for the dynamic optimization and precise control of CRISPR systems, offering new design concepts for constructing adaptive, programmable genetic regulatory networks, and demonstrating significant application potential in fields such as microbial engineering, anti-infective strategies, and precision gene regulation. The regulatory mechanism of Acr proteins on CRISPR activity has been experimentally validated in several studies. Nevertheless, the integration of Acr proteins with other regulatory modules such as CRISPRi/a systems or QS networks remains in the exploratory stage and requires further empirical research to confirm their functionality in complex biological systems.

Streptomycetes are prolific producers of bioactive natural products, but many of the biosynthetic gene clusters (BGCs) are silent in the laboratory. Genetic manipulation is important to unlock their full potential. CRISPR-Cas-based genome editing has greatly advanced genetic engineering in Streptomyces. However, several challenges remain, including Cas nuclease toxicity, unintended genomic rearrangements, and elimination of the delivery plasmid. Here, we present a novel genome editing strategy that harnesses cumate-inducible CRISPR interference (CRISPRi) to transiently knockdown essential genes such as divIVA or dnaA as counterselectable marker. This enforces loss of the vector backbone, promotes homologous recombination, and yields markerless mutants by loss of the antibiotic resistance cassette during the final recombination step. We demonstrate the versatility of the ICE system (Inducible CRISPRi targeting an Essential gene) by (i) deleting four BGCs in Streptomyces coelicolor M145, (ii) inserting both a promoter and a large BGC, and (iii) introducing precise single-nucleotide substitutions. Furthermore, deletion of the prodigiosin BGC elicited expression of a poorly expressed BGC for prolinolexin lipopeptides in Streptomyces roseifaciens DSM 106196T. Considering that different essential genes may be targeted, we anticipate that inducible CRISPRi-based counterselection may be adaptable to genome editing strategies in a broad range of microbial systems.

Plant diseases remain a major constraint to crop productivity and global food security. Conventional breeding has long been used to develop resistant cultivars through the introgression of resistance traits from wild relatives and the selection of favorable phenotypes. However, this process is often slow and limited by linkage drag, known genetic diversity, intrinsic genetic limitations, and the rapid evolution of pathogen populations. Molecular breeding strategies, including marker-assisted selection and genomic selection, have improved the precision of resistance breeding but still rely on existing genetic variation. Recent advances in genome editing technologies are transforming plant breeding by enabling precise modification of gene targets. CRISPR-based systems allow targeted gene knockouts, promoter editing, allelic replacement, and multiplex editing to rapidly generate resistance traits. Many studies have demonstrated that editing susceptibility genes or regulatory regions can enhance resistance to diverse pathogens. Recent research shows that resistance can also be improved by targeting non-classical genes involved in plant immunity, including transcription factors, membrane transporters, heat shock proteins, cell wall-related genes, metabolic enzymes, and epigenetic regulators. Emerging tools such as base editing, prime editing, regulatory tools, and transposon-associated genome engineering systems are further expanding the precision and versatility of plant genome editing. Despite these advances, challenges related to delivery systems, editing efficiency, regulatory frameworks, and field validation remain. Continued technological progress and improved knowledge of plant immune networks will be essential to fully integrate genome editing into crop improvement programs.

Endothelial dysfunction is a key characteristic of many diseases, including atherosclerosis, hypertension, heart failure, stroke, cancer, acute respiratory distress syndrome (ARDS), peripheral vascular disease, coronavirus 2019 (COVID-19), and pulmonary arterial hypertension (PAH). To improve understanding of the roles of endothelial cells (ECs) in health and disease, EC-specific genome editing technologies have been developed in recent years. Therapeutic strategies that aim to restore a healthy endothelial monolayer include the inhibition of endothelial genes that cause EC injury and dysfunction and the induction or activation of endothelial genes that drive EC repair and regeneration. In this review, we describe established recombinase-mediated genetic modification technologies and emerging EC-specific genome editing technologies including viral and non-viral delivery of the CRISPR/Cas9 genome editing system, and we summarize the strengths and limitations of each technology. We then discuss possible avenues for future research, including the development of organ-specific EC genome editing technologies. In short, EC-specific genome editing technologies can be used to modulate gene expression selectively in ECs and even within a specific vascular bed and/or distinctive EC subtype, and, in doing so, greatly improve the understanding of vascular biology and help develop precision genetic medicine targeting the disease-causing vascular bed(s) to effectively treat diseases caused by vascular endothelial dysfunction.

Technologies for editing epigenetic modifications and controlling transcription in mammalian cells have revolutionized targeted gene perturbation, functional genomics, and basic research. By avoiding the generation of DNA breaks, epigenome editing serves as a safe and precise approach for altering gene expression and has emerged as a promising platform for therapeutic applications. The advent of CRISPR has contributed significantly to the expansion of the existing toolkit for programmable modulation of epigenetic and transcriptional states. This review highlights recent discoveries in engineering novel tools for epigenome editing and transcriptional modulation through rational design, high throughput screening methods, and mutational scans, which leverage the endogenous reservoir of chromatin and transcriptional effectors for targeted gene repression and activation. We also discuss the therapeutic potential of epigenome modulators and highlight the key challenges that need to be addressed to improve their safety and efficacy. Advancing our understanding of the complex mechanisms driving gene expression and overcoming current limitations will pave the way for the development of novel technologies that advance fundamental research and translational applications.

Over the past decade, CRISPR-based technologies have revolutionized our capacity to manipulate genomes, thereby reshaping the landscape of functional genomics research. Among the CRISPR toolkit, CRISPR/Cas9-mediated homology-directed repair (HDR) enables precise genome editing with predefined mutations, rendering it an indispensable tool for gene functional analysis, disease model construction, and the development of gene therapy strategies. Nevertheless, despite the robust efficiency of CRISPR/Cas9 in mediating gene knockouts, HDR-dependent gene knock-in (KI) remains a major bottleneck due to its inherently low efficiency. Herein, we report that the co-expression of PALB2 with the CRISPR/Cas9 nuclease could trigger an enhanced HDR effect. Specifically, the fusion of Cas9 with PALB2 elevated KI efficiency by approximately 1.7-fold in human HEK293T cells. Furthermore, this Cas9-PALB2 fusion strategy exhibited cross-cell-type efficacy, demonstrating its broad applicability beyond a single cell line. Notably, the combined application of the Cas9-PALB2 fusion system and Nocodazole further boosted KI efficiency to a remarkable 25.5%. Collectively, these findings establish the Cas9-PALB2 fusion as a highly potent and versatile strategy to augment HDR-mediated KI efficiency, highlighting its substantial potential for widespread utilization in applications that demand high-fidelity genome editing.

The U6 promoter is a critical component of the CRISPR/Cas9 system, as it drives the transcription of single-guide RNAs (sgRNAs) to enable precise genome editing. Endogenous promoters typically exhibit higher transcriptional activity than their exogenous counterparts, which can significantly enhance editing efficiency. However, the endogenous U6 promoter in kenaf (Hibiscus cannabinus L.), an important fiber crop, has not yet been characterized. Using the Arabidopsis U6-26 (AtU6-26) promoter as a reference, we performed a homologous sequence search and identified two candidate U6 promoters in kenaf, designated HcU6-1 and HcU6-14. Promoter fragments were amplified from the kenaf cultivar 'Fuhong 952' and cloned into a β-glucuronidase (GUS) reporter vector. Histochemical GUS staining assays revealed that both HcU6 promoters were transcriptionally active, with HcU6-14 showing significantly stronger expression levels compared to HcU6-1. To further evaluate the utility of these promoters for genome editing, we constructed CRISPR/Cas9 vectors targeting the kenaf acetolactate synthase (ALS) gene, driven by either HcU6-14P or the exogenous cotton GbU6-9P promoter. Agrobacterium rhizogenes K599-mediated transformation was used to induce hairy roots, and mutation analysis of the ALS gene was performed via Sanger sequencing. Notably, targeted mutations in the ALS gene were detected in hairy roots transformed with the HcU6-14P-driven CRISPR/Cas9 vector, whereas no mutations were observed in roots transformed with the exogenous GbU6-9P promoter. These results demonstrate that the endogenous HcU6-14 promoter confers superior genome editing efficiency compared to the heterologous promoter, which facilitates the development of improved varieties with enhanced agronomic traits.

CRISPR-Cas technology has evolved rapidly from a bacterial adaptive immune system to transformative use in molecular diagnostic and genomic engineering. Beyond traditional genome-editing capabilities, newly engineered versions of CRISPR/Cas can be used for programmable transcriptional regulation, epigenetic modification, molecular imaging, and ultrasensitive nucleic acid detection. Specifically, catalytic-inactive Cas proteins like dCas9 and dCas12 retain their ability to bind specific sequences on DNA but do not cleave it. Therefore, these proteins can be reversibly regulated by either CRISPRi or CRISPRa to alter gene expression. Thus, they represent powerful tools for both functional genomic studies and synthetic biological applications. Advances in CRISPR engineering have recently greatly increased the diagnostic potential of Cas12 and Cas13 enzymes. For example, collateral cleavage activity allowed the creation of CRISPR-based diagnostic platforms (SHERLOCK, DETECTR and FELUDA), which can detect target DNA/RNA sequences at high sensitivity and specificity. Moreover, they were demonstrated to work in detecting several infectious pathogens (SARS-CoV-2, Zika virus, and M. tuberculosis) and thus have significant value in point-of-care testing, especially when there is limited availability of resources. CRISPR systems are also being combined with increasing frequency with epigenetic regulators, fluorescence microscopy methods, biosensors, and lab-on-a-chip platforms that incorporate microfluidics to provide improved molecular analysis and automated diagnosis. The purpose of this review is to describe how engineered CRISPR-Cas systems have been developed from primarily genome editing tools into multi-functional platforms for transcriptional regulation, epigenetic engineering, diagnostics, imaging, and emerging microfluidic integrations. Additionally, this review will address some of the current challenges that exist with using CRISPR-based technologies, including off-target effects, delivery efficiency, diagnostic standardization, scaling up production, and translating these technologies clinically.

CRISPR/Cas9 gene editing is a transformative tool for genetic studies in non-model organisms like the southern green stinkbug, Nezara viridula. However, current protocols depend on embryonic microinjection of CRISPR/Cas9, which remains technically difficult. An alternative method of delivering Cas9 ribonucleoprotein directly into female ovaries has been tested in only a few insect species, such as mosquitoes and whiteflies. Here, we developed a simple technique for gene editing by injecting Cas9 ribonucleoprotein into adult Nezara viridula females using ReMOT control or BAPC delivery methods previously described. We observed gene editing of the eye color marker white using the ReMOT method and HhKV ligand. These results demonstrate proof of concept for creating germline mutations in N. viridula. The protocol presented in this study could help advance genetic research in hemipteran pest species.

Bacillus cereus GW-01, an efficient degrader of β-cypermethrin (β-CY), has a high safety profile and probiotic potential for regulating intestinal flora and fermented foods, which is difficult to genetically engineer for modification due to its restrictive modification system. This study successfully developed a CRISPR/enCas12f-based genome editing system, first selecting the plcR gene for proof-of-concept validation with 100% knockout efficiency. Subsequently, this system was utilized to delete the virulence gene nheABC in GW-01, yielding a safer probiotic strain. Compared with the wild-type strain GW-01, the probiotic-related indicators of the ΔnheABC mutant, including cell surface hydrophobicity, auto-aggregation ability and biofilm formation ability, were 80%, 90% and 2.9 (OD₅₉₅), respectively. There were no significant differences in these indicators between the mutant and the wild type. Meanwhile, the ΔnheABC mutant still maintained a high β-cypermethrin degradation efficiency of 80% at the concentration of 30 μg/mL. This work facilitates functional genomic research and genetic modification of Bacillus cereus GW-01. The established CRISPR/enCas12f system enables targeted gene deletion to explore gene functions and phenotypic mechanisms, and paves the way for its development into safe probiotics and excellent microbial chassis.

Multiplex editing is crucial for analyzing complex multiple-gene traits in woody plants, yet its application remains limited by low transformation efficiency and lengthy regeneration cycles. To overcome these barriers, this study establishes an efficient protoplast isolation protocol for Pyrus, employing 1.0% cellulase R10 and 0.4% macerozyme R10 with an 8.5 h digestion, and demonstrates its broad applicability across seven economically important woody plants. Coupling a 40% PEG-4000-mediated transfection regimen with DNA-free CRISPR/Cas9 ribonucleoprotein (RNP) delivery enabled multiplex genome editing in isolated protoplasts. Using this platform, simultaneous disruption of PbrARC3, PbrPARC6, and PbrFtsZ2-1a, key components of the chloroplast division apparatus, consistently reproduced macro-chloroplast abnormalities, confirming effective multigene perturbation within a single cellular context. Notably, chloroplast division failure activated chloroplast-to-nucleus retrograde signaling, evidenced by induction of nuclear stress-response genes PbrRBOHD and PbrZAT12, a concomitant surge in reactive oxygen species, and progression to severe cellular deformation. These results establish a rapid, cross-genus protoplast-RNP workflow that enables DNA-free multiplex editing and accelerates genotype-to-phenotype analyses in woody perennials. The approach provides a practical foundation for functional genomics and supports advances in non-transgenic precision breeding of tree crops.

Breast cancer (BC) is the most prevalent cancer among women and the second leading cause of cancer-related deaths globally, after lung cancer. Despite advances in treatment, BC remains a major contributor to cancer mortality worldwide, underscoring the need for innovative therapeutic approaches. The TopBP1 (DNA Topoisomerase II Binding Protein 1) gene, involved in DNA damage response and cell cycle regulation, has been associated with cancer progression and resistance to chemotherapy. This study investigates the potential of using CRISPR/Cas9 technology to knockout the TopBP1 gene as a novel strategy in breast cancer research. A pair of guide RNAs (gRNAs) was specifically designed to target the TopBP1 gene, inducing the deletion of exon 4. These gRNAs were transfected into the MCF7 breast cancer cell line, and the efficacy of genomic editing was validated using PCR and Sanger sequencing. Subsequent analyses employing real-time PCR and Western blotting were conducted to investigate the downstream effects of this genetic modification on gene expression. The CRISPR/Cas9 system successfully knocked out exon 4 of the TopBP1 gene in MCF7 breast cancer cells, as validated by PCR and Sanger sequencing. Real-time PCR analysis revealed a significant increase in Bax expression and a decrease in Bcl-2 expression in the knockout cells compared to controls. These changes indicate enhanced apoptotic activity following TopBP1 knockout, suggesting that MCF7 cells may become more sensitive to apoptosis. Overall, the findings support the hypothesis that targeting TopBP1 could play a critical role in promoting cell death in breast cancer, potentially offering a new therapeutic strategy. This study successfully employed the CRISPR/Cas9 system to knockout exon 4 of the TopBP1 gene in MCF7 breast cancer cells, resulting in reduced TopBP1 expression. The subsequent increase in the pro-apoptotic Bax gene and decrease in the anti-apoptotic Bcl-2 gene suggest that targeting TopBP1 could enhance apoptosis in breast cancer cells, offering a promising alternative to conventional treatments. Further research is necessary to fully explore the therapeutic potential of this approach.

Neurological disorders are complex and often very challenging for patients. Many of these conditions result from mutations in genes that are essential for normal function. Most existing treatments only alleviate symptoms, highlighting the urgent need for more effective therapeutic strategies. In the current drug development landscape, gene therapy offers hope as a promising approach. Specifically, CRISPR-Cas9 technology enables precise gene editing across diverse cell types and organisms. An increasing number of research groups are investigating innovative therapies and the molecular mechanisms behind neurological diseases. This review highlights the use of CRISPR-based gene therapies for various brain diseases, including multiple sclerosis, Alzheimer's, Parkinson's disease, epilepsy, stroke, and brain tumors. It consistently recognizes significant challenges in clinical applications, including overcoming the blood-brain barrier (BBB), managing off-target effects, ensuring efficient delivery, and addressing immunogenicity and ethical concerns.

Cardiomyopathies constitute a heterogeneous group of myocardial disorders representing leading causes of heart failure and cardiovascular mortality worldwide. While genetic mutations have been extensively characterized across different cardiomyopathy subtypes, the mechanistic links between genotype and phenotype remain incompletely understood. This review synthesizes current knowledge regarding chromatin remodeling complexes and their roles in cardiac gene regulation under physiological and pathological conditions. Moreover, disease-specific chromatin remodeling patterns were examined across dilated, hypertrophic, arrhythmogenic, and restrictive cardiomyopathies, highlighting both conserved mechanisms and subtype-specific alterations. Chromatin remodeling alterations contribute significantly to cardiomyopathy pathogenesis across multiple subtypes. The reversibility of epigenetic modifications presents therapeutic opportunities not available with genetic interventions. Selective HDAC inhibitors and EZH2 antagonists show promise in preclinical models, though clinical translation requires development of cardiac-specific delivery systems. CRISPR-based epigenetic editing technologies offer future potential for precise genomic locus-specific interventions to reverse pathological transcriptional programs. Chromatin remodeling complexes including SWI/SNF (BAF), NuRD, Polycomb, ISWI, CHD, and INO80 families- modulate disease expression, progression, and phenotypic variability through epigenetic modifications and ATP-dependent chromatin remodeling. Emerging evidence demonstrates that chromatin remodelers interact dynamically with DNA methylation machinery, histone-modifying enzymes, and cardiac transcription factors to orchestrate pathological gene expression programs. Understanding these epigenetic mechanisms offers unprecedented opportunities for developing novel therapeutic strategies targeting the chromatin regulatory apparatus, potentially reversing maladaptive transcriptional programs that drive disease progression.

Cone-rod dystrophy (CORD) and achromatopsia (ACHM) are inherited retinal dystrophies for which conventional adeno-associated virus (AAV) gene augmentation has important limitations, particularly in autosomal-dominant gain-of-function CORD and recessive ACHM. Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) offers the potential for one-time, mutation-specific gene correction or allele ablation. This systematic review summarizes preclinical evidence on CRISPR/Cas9-based approaches for CORD and ACHM, focusing on editing efficiency, phenotypic rescue, and safety. This review followed PRISMA guidelines. PubMed, Google Scholar, and ScienceDirect were searched through June 2025 for original experimental studies using CRISPR/Cas9 in CORD or ACHM animal models or human-derived cell lines. Dual independent screening and data extraction were performed. Outcomes related to editing efficiency, structural or functional rescue, and safety were synthesized narratively. Four studies were included: three targeting CORD and one targeting ACHM. In vivo studies used AAV-delivered SaCas9 to disrupt GUCY2D (or murine orthologs) in mouse and macaque photoreceptors, achieving approximately 8-45% on-target editing in mice and approximately 13% in macaques. Although ablation alone reduced retGC1 expression, it did not improve retinal function; however, a dual-AAV "ablate-and-replace" strategy preserved outer nuclear layer thickness for up to 24 weeks in CORD6 mice. In vitro, PROM1 correction in patient-derived iPSCs restored CD133 expression, and SpCas9-HiFi-mediated PDE6C correction in ACHM iPSCs achieved approximately 80% editing efficiency while preserving pluripotency and showing no detectable off-target effects. Safety data were limited, with immune responses assessed in only one primate study. CRISPR/Cas9 shows promising preclinical efficacy for CORD and ACHM, particularly allele-specific ablate-and-replace strategies for CORD and precise HDR-based correction for ACHM. However, the available evidence remains limited, underscoring the need for expanded safety assessment, non-human primate studies, and standardized functional outcomes measures before clinical translation.

Advancements in genome editing have established a new frontier for the treatment of various genetic diseases, including sickle cell disease (SCD). SCD, the most prevalent monogenic blood disorder, causes severe pain, organ damage, and reduced life expectancy. The recent clinical approval of clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9-based gene therapies for severe sickle cell anemia marks a significant milestone in treating genetic diseases. Despite these breakthroughs, limitations in CRISPR technology persist, requiring further innovation. Alternative approaches, such as the Fanzor (Fz) system, are being developed to complement CRISPR's capabilities. Unlike CRISPR, which is typically encoded within prokaryotic organisms, Fz is encoded in the eukaryotic genome, offering a universal RNA-guided mechanism applicable across all life kingdoms. Fz's eukaryotic origin may facilitate more efficient delivery across diverse cell types and tissues, enhancing its therapeutic potential. Here, we will review the current successes and limitations of the CRISPR technology in editing mutation associated with SCD. Additionally, we will explore the potential role of Fz as a genome-editing tool for SCD, a field where its application has not yet been studied.