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CRISPR-Cas-Guided Mutagenesis of Chromosome and Virulence Plasmid in Shigella flexneri by Cytosine Base Editing.
Shigella is a Gram-negative bacterium that invades the human gut epithelium. The resulting infection, shigellosis, is the deadliest bacterial diarrheal disease. Much of the information about the genes dictating the pathophysiology of Shigella, both on the chromosome and the virulence plasmid, was obtained by classical reverse genetics. However, technical limitations of the prevalent mutagenesis techniques restrict the generation of mutants in a single reaction to a small number, preventing large-scale targeted mutagenesis of Shigella and the subsequent assessment of phenotype. We adopted a CRISPR-Cas-dependent approach, where a nickase Cas9 and cytidine deaminase fusion is guided by single guide RNA (sgRNA) to introduce targeted C→T transitions, resulting in internal stop codons and premature termination of translation. In proof-of-principle experiments using an mCherry fluorescent reporter, we were able to generate loss-of-function mutants in both Escherichia coli and Shigella flexneri with up to 100% efficacy. Using a modified fluctuation assay, we determined that under optimized conditions, the frequency of untargeted mutations introduced by the Cas9-deaminase fusion was in the same range as spontaneous mutations, making our method a safe choice for bacterial mutagenesis. Furthermore, we programmed the method to mutate well-characterized chromosomal and plasmid-borne Shigella flexneri genes and found the mutant phenotype to be similar to those of the reported gene deletion mutants, with no apparent polar effects at the phenotype level. This method can be used in a 96-well-plate format to increase the throughput and generate an array of targeted loss-of-function mutants in a few days. IMPORTANCE Loss-of-function mutagenesis is critical in understanding the physiological role of genes. Therefore, high-throughput techniques to generate such mutants are important for facilitating the assessment of gene function at a pace that matches systems biology approaches. However, to our knowledge, no such method was available for generating an array of single gene mutants in an important enteropathogen-Shigella. This pathogen causes high morbidity and mortality in children, and antibiotic-resistant strains are quickly emerging. Therefore, determination of the function of unknown Shigella genes is of the utmost importance to develop effective strategies to control infections. Our present work will bridge this gap by providing a rapid method for generating loss-of-function mutants. The highly effective and specific method has the potential to be programmed to generate multiple mutants in a single, massively parallel reaction. By virtue of plasmid compatibility, this method can be extended to other members of Enterobacteriaceae.
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An Accessible Protocol for the Generation of CRISPR-Cas9 Knockouts Using INDELs in Zebrafish.
The ability to create targeted mutations in specific genes, and therefore a loss-of-function condition, provides essential information about their endogenous functions during development and homeostasis. The discovery that CRISPR-Cas9 can target specific sequences according to base-pair complementarity and readily create knockouts in a desired gene has elevated the implementation of genetic analysis in numerous organisms. As CRISPR-Cas9 has become a powerful tool in a number of species, multiple methods for designing, creating, and screening editing efficiencies have been published, each of which has unique benefits. This chapter presents a cost-efficient, accessible protocol for creating knockout mutants in zebrafish using insertions/deletions (INDELS), from target site selection to mutant propagation, using basic laboratory supplies. The presented approach can be adapted to other systems, including any vertebrate species.
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Potent and uniform fetal hemoglobin induction via base editing.
Inducing fetal hemoglobin (HbF) in red blood cells can alleviate β-thalassemia and sickle cell disease. We compared five strategies in CD34+ hematopoietic stem and progenitor cells, using either Cas9 nuclease or adenine base editors. The most potent modification was adenine base editor generation of γ-globin -175A>G. Homozygous -175A>G edited erythroid colonies expressed 81 ± 7% HbF versus 17 ± 11% in unedited controls, whereas HbF levels were lower and more variable for two Cas9 strategies targeting a BCL11A binding motif in the γ-globin promoter or a BCL11A erythroid enhancer. The -175A>G base edit also induced HbF more potently than a Cas9 approach in red blood cells generated after transplantation of CD34+ hematopoietic stem and progenitor cells into mice. Our data suggest a strategy for potent, uniform induction of HbF and provide insights into γ-globin gene regulation. More generally, we demonstrate that diverse indels generated by Cas9 can cause unexpected phenotypic variation that can be circumvented by base editing.
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Development of a highly efficient prime editor system in mice and rabbits.
The recently developed prime-editing (PE) technique is more precise than previously available techniques and permits base-to-base conversion, replacement, and insertions and deletions in the genome. However, previous reports show that the efficiency of prime editing is insufficient to produce genome-edited animals. In fact, prime-guide RNA (pegRNA) designs have posed a challenge in achieving favorable editing efficiency. Here, we designed prime binding sites (PBS) with a melting temperature (Tm) of 42 °C, leading to optimal performance in cells, and we found that the optimal Tm was affected by the culture temperature. In addition, the ePE3max system was developed by updating the PE architecture to PEmax and expressing engineered pegRNA (epegRNA) based on the original PE3 system. The updated ePE3max system can efficiently induce gene editing in mouse and rabbit embryos. Furthermore, we successfully generated a Hoxd13 (c. 671 G > T) mutation in mice and a Tyr (c. 572 del) mutation in rabbits by ePE3max. Overall, the editing efficiency of modified ePE3max systems is superior to that of the original PE3 system in producing genome-edited animals, which can serve as an effective and versatile genome-editing tool for precise genome modification in animal models.
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CRISPys: Optimal sgRNA Design for Editing Multiple Members of a Gene Family Using the CRISPR System.
The development of the CRISPR-Cas9 system in recent years has made eukaryotic genome editing, and specifically gene knockout for reverse genetics, a simple and effective task. The system is directed to a genomic target site by a programmed single-guide RNA (sgRNA) that base-pairs with it, subsequently leading to site-specific modifications. However, many gene families in eukaryotic genomes exhibit partially overlapping functions, and thus, the knockout of one gene might be concealed by the function of the other. In such cases, the reduced specificity of the CRISPR-Cas9 system, which may lead to the modification of genomic sites that are not identical to the sgRNA, can be harnessed for the simultaneous knockout of multiple homologous genes. We introduce CRISPys, an algorithm for the optimal design of sgRNAs that would potentially target multiple members of a given gene family. CRISPys first clusters all the potential targets in the input sequences into a hierarchical tree structure that specifies the similarity among them. Then, sgRNAs are proposed in the internal nodes of the tree by embedding mismatches where needed, such that the efficiency to edit the induced targets is maximized. We suggest several approaches for designing the optimal individual sgRNA and an approach to compute the optimal set of sgRNAs for cases when the experimental platform allows for more than one. The latter may optionally account for the homologous relationships among gene-family members. We further show that CRISPys outperforms simpler alignment-based techniques by in silico examination over all gene families in the Solanum lycopersicum genome.
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Research progress of gene correction for genetic diseases via in vivo prime editing.
Curative therapies for genetic diseases caused by pathogenic mutations demand precise and safe genome editing technologies. Prime editing (PE), a CRISPR-Cas9-derived technology that enables precise base conversions without double-strand breaks (DSBs), has emerged as a transformative tool with the ability to correct 89% of human pathogenic mutations. However, transitioning PE from preclinical models to clinical applications faces critical challenges, particularly in the development of in vivo delivery systems and scalable manufacturing. This review systematically summarizes advances of in vivo PE therapeutics, focusing on the therapeutic outcomes across some retinal, metabolic, and hematologic genetic diseases. We further dissect barriers to clinical translation, including in the immunogenicity of viral vectors, off-target risks in nondividing cells, and large-scale production bottlenecks. By highlighting the unique advantages of in vivo PE and proposing actionable solutions for scalable manufacturing, the work provides a roadmap for accelerating in vivo PE-based therapies toward clinical reality.
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Glycosylase-based base editors for efficient T-to-G and C-to-G editing in mammalian cells.
Base editors show promise for treating human genetic diseases, but most current systems use deaminases, which cause off-target effects and are limited in editing type. In this study, we constructed deaminase-free base editors for cytosine (DAF-CBE) and thymine (DAF-TBE), which contain only a cytosine-DNA or a thymine-DNA glycosylase (CDG/TDG) variant, respectively, tethered to a Cas9 nickase. Multiple rounds of mutagenesis by directed evolution in Escherichia coli generated two variants with enhanced base-converting activity-CDG-nCas9 and TDG-nCas9-with efficiencies of up to 58.7% for C-to-A and 54.3% for T-to-A. DAF-BEs achieve C-to-G/T-to-G editing in mammalian cells with minimal Cas9-dependent and Cas9-independent off-target effects as well as minimal RNA off-target effects. Additional engineering resulted in DAF-CBE2/DAF-TBE2, which exhibit altered editing windows from the 5' end to the middle of the protospacer and increased C-to-G/T-to-G editing efficiency of 3.5-fold and 1.2-fold, respectively. Compared to prime editing or CGBEs, DAF-BEs expand conversion types of base editors with similar efficiencies, smaller sizes and lower off-target effects.
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[Plant prime editing technique: a new genome editing tool for plants].
The CRISPR/Cas9 based prime editing (PE) technique enables all 12 types of base substitutions and precise small DNA deletions or insertions without generating DNA double-strand breaks. Prime editing has been successfully applied in plants and plays important roles in plant precision breeding. Although plant prime editing (PPE) can substantially expand the scope and capabilities of precise genome editing in plants, its editing efficiency still needs to be further improved. Here, we review the development of PPE technique, and introduce structural composition, advantages and limitations of PPE. Strategies to improve the PPE editing efficiency, including the Tm-directed PBS length design, the RT template length, the dual-pegRNA strategy, the PlantPegDesigner website, and the strategies for optimizing the target proteins of PPE, were highlighted. Finally, the prospects of future development and application of PPE were discussed.
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Precise Correction of Heterozygous SHOX2 Mutations in hiPSCs Derived from Patients with Atrial Fibrillation via Genome Editing and Sib Selection.
Patient-specific human induced pluripotent stem cells (hiPSCs) offer unprecedented opportunities for the investigation of multigenic disease, personalized medicine, and stem cell therapy. For heterogeneous diseases such as atrial fibrillation (AF), however, precise correction of the associated mutation is crucial. Here, we generated and corrected hiPSC lines from two AF patients carrying different heterozygous SHOX2 mutations. We developed a strategy for the scarless correction of heterozygous mutations, based on stochastic enrichment by sib selection, followed by allele quantification via digital PCR and next-generation sequencing to detect isogenic subpopulations. This allowed enriching edited cells 8- to 20-fold. The method does not require antibiotic selection or cell sorting and can be easily combined with base-and-prime editing approaches. Our strategy helps to overcome low efficiencies of homology-dependent repair in hiPSCs and facilitates the generation of isogenic control lines that represent the gold standard for modeling complex diseases in vitro.
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CRISPR-Cas9 guided RNA-based model for the silencing of spinal bulbar muscular atrophy: A functional genetic disorder.
This study explores a novel therapeutic approach for spinal bulbar muscular atrophy (SBMA), a neurodegenerative disorder caused by a mutation in the Androgen Receptor (AR) gene. The aim is to investigate the potential of CRISPR-Cas9 technology in targeting the mutant AR gene to inhibit its production. The objectives include assessing the accuracy and efficacy of CRISPR-Cas9 guided RNAs in silencing the mutant gene and evaluating the feasibility of this approach as a treatment for SBMA. Computational and in-silico approaches are used to evaluate the feasibility of using CRISPR-Cas9 technology for treating SBMA. Computational analysis is used to design CRISPR-Cas9 guided RNAs targeting the mutant AR gene, assessing their on-target and off-target scores, GC content, and structural accuracy. In-silico simulations predict the potential therapeutic outcomes of the CRISPR-Cas9 approach in an artificial environment. Three guided RNA (gRNA) sequences were designed using the CHOPCHOP tool, targeting specific regions of the AR gene with high efficiency and 100% match. These gRNAs demonstrated effective targeting with minimal off-target scores and optimal GC content. Additionally, lentiCRISPR v2 plasmids were designed for the delivery of CRISPR materials, enabling high-efficiency multiplex genome editing of the AR gene. Thermodynamic ensemble predictions indicated favorable secondary structure stability of the designed gRNAs, further supporting their suitability for gene editing. The evaluation of designed gRNAs confirmed their strong binding ability to the target sequences, validating their potential as effective tools for genome editing. The study highlights the potential of CRISPR-Cas9 technology for targeting the Androgen Receptor gene associated with spinal bulbar muscular atrophy (SBMA). The findings support the feasibility of this approach for gene editing and suggest further exploration in preclinical and clinical settings. Recommendations include continued research to optimize CRISPR-Cas9 delivery methods and enhance specificity for therapeutic applications in SBMA.
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