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Protocol for CRISPR-Cas9-mediated genome editing to study spermatogenesis in Caenorhabditis elegans.
Gene silencing by P-element-induced wimpy testis-interacting RNAs is a mechanism to maintain genome integrity in germ cells. Here, we present a protocol for knockin or knockout editing of male germline genome mediated by CRISPR-Cas9 technology in Caenorhabditis elegans. We describe steps for constructing edited plasmids, microinjecting worms with these plasmids, and screening edited worms. We then detail procedures for dissecting released sperm and their observation with fluorescence microscopy. Engineered worms provide a model for studying hermaphrodite/male fertility or protein localization in vivo. For complete details on the use and execution of this protocol, please refer to Wang et al. (2021).1.
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Lipid Nanoparticle-Mediated Delivery of CRISPR-Cas9 Components for Genome Editing.
Lipid nanoparticles (LNPs) are a clinically validated nonviral platform for the delivery of CRISPR-associated components. Composed of ionizable lipids, phospholipids, cholesterol, and PEG-lipids, LNPs enable the efficient encapsulation, protection, and cytosolic delivery of therapeutic cargo such as DNA, RNA, or proteins. The clinical relevance of LNPs has already been shown by multiple FDA-approved therapies, including siRNA-based treatments and mRNA vaccines. Compared with viral vectors, LNPs offer several advantages, including reduced immunogenicity, absence of genomic integration, scalable manufacturing, and flexibility in cargo size, while supporting transient expression, which is desirable for genome editing applications. However, challenges remain, including limited tissue specificity and inefficient endosomal escape. Recent advances in lipid chemistry optimization and surface modification have improved delivery performance. Among available formulation techniques, microfluidic mixing has emerged as a preferred method due to its reproducibility, scalability, and precise control over particle properties. This protocol describes a standardized microfluidic workflow for reproducible LNP formulation, providing practical guidance on lipid preparation, nanoparticle assembly, and quality control.
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Genome Editing in a Wide Area of the Brain Using Dendrimer-Based Ternary Polyplexes of Cas9 Ribonucleoprotein.
A preassembled Cas9/single-guide RNA complex (Cas9 ribonucleoprotein; Cas9 RNP) induces genome editing efficiently, with small off-target effects compared with the conventional techniques, such as plasmid DNA and mRNA systems. However, penetration of Cas9 RNP through the cell membrane is low. In particular, the incorporation of Cas9 RNP into neurons and the brain is challenging. In the present study, we have reported the use of a dendrimer (generation 3; G3)/glucuronylglucosyl-β-cyclodextrin conjugate (GUG-β-CDE (G3)) as a carrier of Cas9 RNP and evaluated genome editing activity in the neuron and the brain. A Cas9 RNP ternary complex with GUG-β-CDE (G3) was prepared by only mixing the components. The resulting complex exhibited higher genome editing activity than the complex with the dendrimer (G3), Lipofectamine 3000 or Lipofectamine CRISPRMAX in SH-SY5Y cells, a human neuroblastoma cell line. In addition, GUG-β-CDE (G3) enhanced the genome editing activity of Cas9 RNP in the whole mouse brain after a single intraventricular administration. Thus, GUG-β-CDE (G3) is a useful Cas9 RNP carrier that can induce genome editing in the neuron and brain.
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Protocol for establishing CRISPR-Cas12a for efficient genome editing of Pseudomonas aeruginosa phages.
We developed an efficient type V CRISPR-Cas12a system tailored specifically for Pseudomonas aeruginosa phages, showcasing its remarkable cleavage activity and the ability to precisely introduce genetic modifications, including point mutations, deletions, and insertions, into phage genomes. Here, we present a protocol for establishing CRISPR-Cas12a for genome editing of Pseudomonas aeruginosa phages. We describe steps for the construction of pCRISPR-12a plasmid and guide RNA and the utilization of the type V CRISPR-Cas12a system for precise genetic editing of phages. For complete details on the use and execution of this protocol, please refer to Chen et al.1.
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CRISPR-Based Gene Editing in Acinetobacter baumannii to Combat Antimicrobial Resistance.
Antimicrobial resistance (AMR) poses a significant threat to the health, social, environment, and economic sectors on a global scale and requires serious attention to addressing this issue. Acinetobacter baumannii was given top priority among infectious bacteria because of its extensive resistance to nearly all antibiotic classes and treatment options. Carbapenem-resistant A. baumannii is classified as one of the critical-priority pathogens on the World Health Organization (WHO) priority list of antibiotic-resistant bacteria for effective drug development. Although available genetic manipulation approaches are successful in A. baumannii laboratory strains, they are limited when employed on newly acquired clinical strains since such strains have higher levels of AMR than those used to select them for genetic manipulation. Recently, the CRISPR-Cas (Clustered regularly interspaced short palindromic repeats/CRISPR-associated protein) system has emerged as one of the most effective, efficient, and precise methods of genome editing and offers target-specific gene editing of AMR genes in a specific bacterial strain. CRISPR-based genome editing has been successfully applied in various bacterial strains to combat AMR; however, this strategy has not yet been extensively explored in A. baumannii. This review provides detailed insight into the progress, current scenario, and future potential of CRISPR-Cas usage for AMR-related gene manipulation in A. baumannii.
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Genome editing‑based strategies to combat geminiviruses: CRISPR/Cas9 and emerging high‑fidelity tools.
Plant virus diseases constitute a major constraint to agriculture and have adversely affected crop productivity worldwide. Over the past few decades, geminiviruses of the family Geminiviridae have emerged as some of the most destructive plant pathogens. Conventional. approaches for virus management have demonstrated varying degrees of success, their deployment is often hindered by prolonged timelines, limited availability of resistant sources, and reduced durability against rapidly evolving geminiviral pathogens. With the rapid evolution of viruses and the extensive damage they cause, it is imperative to develop rapid, cost-effective, and efficient antivirus resistance strategies. In recent years, CRISPR/Cas-based genome editing has made it possible to precisely target viral genomes, offering new avenues for developing antiviral resistance, although long-term durability remains under investigation. Derived from prokaryotic adaptive immune systems, the CRISPR/Cas platform confers resistance either by expressing genome-editing constructs that directly cleave viral genomes or by modifying host susceptibility (S) genes, representing mechanistically distinct strategies with different implications for durability of resistance. Recent advances have further expanded this toolkit with Cas-CLOVER, a dual guide RNA-dependent nuclease system designed to mitigate off-target effects associated with conventional Cas9. This review summarizes the principles of CRISPR/Cas-mediated antiviral resistance, critically evaluates viral escape, resistance durability, compares Cas9 with emerging platforms, and outlines conceptual frameworks for developing durable and field-ready genome-editing strategies.
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CRISPR-Cas9 mediated proteinase 3 autoantigen deletion as a treatment strategy for anti-neutrophil cytoplasmic autoantibody-associated vasculitis.
INTRODUCTION: Proteinase 3 (PR3) is a major autoantigen in patients with anti-neutrophil cytoplasmic autoantibody (ANCA)-associated vasculitis (AAV). Here, we performed a proof-of-principle study using ex vivo CRISPR-Cas9 guided gene editing to eliminate the PR3 autoantigen as an alternative to suppressing the autoimmune response to PR3. METHODS: A ribonucleoprotein (RNP) complex of Cas9 protein and a PR3-specific single guide-RNA was transfected into human CD34+ hematopoietic stem and progenitor cells (HSPC) by electroporation. Effects on PR3 protein abundance, neutrophil differentiation, and ANCA-dependent and -independent neutrophil responses were assessed. RESULTS: Gene editing introduced a frame shift in exon 2 of PRTN3. Consequently, PR3 protein was efficiently reduced in neutrophil-differentiated HSPCs as demonstrated by immunoblotting, ELISA, microscopy, and the complete absence of PR3-specific proteolytic activity. Human neutrophil elastase served as control and was not affected. PR3-deleted (PR3KO)- and PR3 wild-type (PR3WT)-HSPCs showed similar neutrophil differentiation. Importantly, general neutrophil defense functions to non-ANCA receptor-independent and -dependent stimuli were similar in PR3KO- and PR3WT-neutrophils as was constitutive apoptosis. Flow cytometry showed that cell membrane-PR3 was significantly reduced on PR3KO-neutrophils and consequent neutrophil activation to either monoclonal antibodies to PR3 or human PR3-ANCA was attenuated. In contrast, myeloperoxidase-ANCA stimulation was not affected. CONCLUSIONS: We show the feasibility and efficacy of depleting the PR3 autoantigen in human CD34+ HSPCs using CRISPR-Cas9. Depleting the PR3 autoantigen instead of suppressing the autoimmune response to PR3 could potentially lead to drug-free remission, particularly in patients with refractory or relapsing disease.
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Delivery of genome editing tools by bacterial extracellular vesicles.
Here we propose to use bacterial cells as factories that generate EVs harboring both the RNP complex (or any other CRISPR-mediated tool) and a specific ligand molecule. The ligand would allow specific binding of EVs to cells with the matching receptors, adding a level of specificity to the delivery, hence stimulating precise genome editing.
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CRISPR-Cas9: a world first?
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A CRISPR-Cas9 System for Genome Editing of Fusarium proliferatum.
Fusarium proliferatum causes diverse diseases of many economically important plants. The fungus produces several mycotoxins of which the fumonisins are the most toxic. Currently, deletion of key genes for mycotoxin biosynthesis is a laborious and time-consuming procedure. We developed a novel CRISPR/Cas9-based genome-editing tool for the direct delivery of preassembled Cas9 ribonucleoproteins into protoplasts of F. proliferatum. Our CRISPR-Cas9 system couples a site-specific double-strand DNA break mediated by two Cas9 ribonucleoproteins with microhomology recombination requiring only 50-bp regions flanking the target gene. This system reduces the risk of off-target mutations and minimizes the risk of altering any gene adjacent to the target region. We used this tool to delete a polyketide synthase gene (FUM1) required for fumonisin biosynthesis. The mutants generated are no longer able to produce fumonisins, confirming the key role of FUM1 in fumonisin biosynthesis. Our CRISPR-Cas9 system is an important new tool for genetic studies of Fusarium.
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