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Efficient Gene Disruption via Base Editing Induced Stop in Newt Pleurodeles waltl.
Loss-of-function approaches provide strong evidence for determining the role of particular genes. The prevalent CRISPR/Cas9 technique is widely used to disrupt target gene with uncontrolled non-homologous end joining after the double strand breaks, which results in mosaicism and multiple genotypes in the founders. In animal models with long generation time such as the salamanders, producing homozygous offspring mutants would be rather labor intensive and time consuming. Here we utilized the base editing technique to create the loss-of-function F0 mutants without the random indels. As a proof of principle, we successfully introduced premature stop codons into the tyrosinase locus and produced the albino phenotype in the newts (Pleurodeles waltl). We further demonstrated that the knockout efficiency could be greatly improved by using multiplex sgRNAs target the same gene. The F0 mutated animals showed fully loss-of-function by both genotyping and phenotyping analysis, which could enable direct functional analysis in the founders and avoid sophisticated breeding. This study not only presented the high efficiency of single base editing in a gigantic animal genome (>20 G), but also provided new tools for interrogating gene function in other salamander species.
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Histone gene editing probes functions of H3K27 modifications in mammals.
A new study employs CRISPR/Cas9-based base editing for simultaneous mutagenesis of all copies of histone H3 genes in mammals, highlighting the functional importance of H3K27me3 for Polycomb-mediated gene silencing and the dispensability for H3K27ac in transcriptional activation.
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CRISPR-based engineering of phages for in situ bacterial base editing.
Investigation of microbial gene function is essential to the elucidation of ecological roles and complex genetic interactions that take place in microbial communities. While microbiome studies have increased in prevalence, the lack of viable in situ editing strategies impedes experimental progress, rendering genetic knowledge and manipulation of microbial communities largely inaccessible. Here, we demonstrate the utility of phage-delivered CRISPR-Cas payloads to perform targeted genetic manipulation within a community context, deploying a fabricated ecosystem (EcoFAB) as an analog for the soil microbiome. First, we detail the engineering of two classical phages for community editing using recombination to replace nonessential genes through Cas9-based selection. We show efficient engineering of T7, then demonstrate the expression of antibiotic resistance and fluorescent genes from an engineered λ prophage within an Escherichia coli host. Next, we modify λ to express an APOBEC-1-based cytosine base editor (CBE), which we leverage to perform C-to-T point mutations guided by a modified Cas9 containing only a single active nucleolytic domain (nCas9). We strategically introduce these base substitutions to create premature stop codons in-frame, inactivating both chromosomal (lacZ) and plasmid-encoded genes (mCherry and ampicillin resistance) without perturbation of the surrounding genomic regions. Furthermore, using a multigenera synthetic soil community, we employ phage-assisted base editing to induce host-specific phenotypic alterations in a community context both in vitro and within the EcoFAB, observing editing efficiencies from 10 to 28% across the bacterial population. The concurrent use of a synthetic microbial community, soil matrix, and EcoFAB device provides a controlled and reproducible model to more closely approximate in situ editing of the soil microbiome.
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Tools for editing the mammalian mitochondrial genome.
The manipulation of animal mitochondrial genomes has long been a challenge due to the lack of an effective transformation method. With the discovery of specific gene editing enzymes, designed to target pathogenic mitochondrial DNA mutations (often heteroplasmic), the selective removal or modification of mutant variants has become a reality. Because mitochondria cannot efficiently import RNAs, CRISPR has not been the first choice for editing mitochondrial genes. However, the last few years witnessed an explosion in novel and optimized non-CRISPR approaches to promote double-strand breaks or base-edit of mtDNA in vivo. Engineered forms of specific nucleases and cytidine/adenine deaminases form the basis for these techniques. I will review the newest developments that constitute the current toolbox for animal mtDNA gene editing in vivo, bringing these approaches not only to the exploration of mitochondrial function, but also closer to clinical use.
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Plant genome editing: ever more precise and wide reaching.
Genome-editing technologies consisting of targeted mutagenesis and gene targeting enable us to modify genes of interest rapidly and precisely. The discovery in 2012 of CRISPR/Cas9 systems and their development as sequence-specific nucleases has brought about a paradigm shift in biology. Initially, CRISPR/Cas9 was applied in targeted mutagenesis to knock out a target gene. Thereafter, advances in genome-editing technologies using CRISPR/Cas9 developed rapidly, with base editing systems for transition substitution using a combination of Cas9 nickase and either cytidine or adenosine deaminase being reported in 2016 and 2017, respectively, and later in 2021 bringing reports of transversion substitution using Cas9 nickase, cytidine deaminase and uracil DNA glycosylase. Moreover, technologies for gene targeting and prime editing systems using DNA or RNA as donors have also been developed in recent years. Besides these precise genome-editing strategies, reports of successful chromosome engineering using CRISPR/Cas9 have been published recently. The application of genome editing to crop breeding has advanced in parallel with the development of these technologies. Genome-editing enzymes can be introduced into plant cells, and there are now many examples of crop breeding using genome-editing technologies. At present, it is no exaggeration to say that we are now in a position to be able to modify a gene precisely and rearrange genomes and chromosomes in a predicted way. In this review, we introduce and discuss recent highlights in the field of precise gene editing, chromosome engineering and genome engineering technology in plants.
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CRISPR-Cas9 and precision editing technologies linking functional genomics to clinical translation in genetic diseases.
BACKGROUND: CRISPR-Cas9 and derivative precision-editing platforms increasingly connect pathogenic variant interpretation with functional genomics and therapeutic development in genetic diseases. This narrative review focuses on a variant-mechanism-driven framework for matching editing strategies to mutation structure, functional consequence, disease-model evidence, delivery feasibility, safety risk, and translational readiness. MAIN BODY: The review summarizes how monogenic, polygenic, coding, non-coding, mitochondrial, and complex disease contexts influence the choice of canonical Cas9 editing, base editing, prime editing, Cas variants, CRISPR interference/activation, epigenome editing, and disease-model systems. It further compares ex vivo and in vivo delivery routes, safety assessment strategies, immunogenicity and genotoxicity concerns, and clinical implementation barriers, including CMC/manufacturing scalability, long-term follow-up, affordability, and regulatory oversight. Current evidence supports the clinical maturity of ex vivo hematopoietic editing, whereas most in vivo and precision-repair approaches remain constrained by delivery, durability, product heterogeneity, and safety uncertainties. CONCLUSION: The central conclusion is that future CRISPR-based interventions should be judged not only by editability, but by whether molecular correction can be translated into durable, safe, manufacturable, and clinically meaningful benefit.
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In Silico Analysis of Pathogenic CRB1 Single Nucleotide Variants and Their Amenability to Base Editing as a Potential Lead for Therapeutic Intervention.
Mutations in the Crumbs homolog 1 (CRB1) gene cause both autosomal recessive retinitis pigmentosa (RP) and Leber congenital amaurosis (LCA). Since three separate CRB1 isoforms are expressed at meaningful levels in the human retina, base editing shows promise as a therapeutic approach. This retrospective analysis aims to summarise the reported pathogenic CRB1 variants and investigate their amenability to treatment with currently available DNA base editors. Pathogenic single nucleotide variants (SNVs) were extracted from the Leiden open-source variation database (LOVD) and ClinVar database and coded by mutational consequence. They were then analyzed for their amenability to currently available DNA base editors and available PAM sites from a selection of different Cas proteins. Of a total of 1115 unique CRB1 variants, 69% were classified as pathogenic SNVs. Of these, 62% were amenable to currently available DNA BEs. Adenine base editors (ABEs) alone have the potential of targeting 34% of pathogenic SNVs; 19% were amenable to a CBE while GBEs could target an additional 9%. Of the pathogenic SNVs targetable with a DNA BE, 87% had a PAM site for a Cas protein. Of the 33 most frequently reported pathogenic SNVs, 70% were targetable with a base editor. The most common pathogenic variant was c.2843G>A, p.Cys948Arg, which is targetable with an ABE. Since 62% of pathogenic CRB1 SNVs are amenable to correction with a base editor and 87% of these mutations had a suitable PAM site, gene editing represents a promising therapeutic avenue for CRB1-associated retinal degenerations.
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Advancing CRISPR genome editing into gene therapy clinical trials: progress and future prospects.
Genome editing has recently evolved from a theoretical concept to a powerful and versatile set of tools. The discovery and implementation of CRISPR-Cas9 technology have propelled the field further into a new era. This RNA-guided system allows for specific modification of target genes, offering high accuracy and efficiency. Encouraging results are being announced in clinical trials employed in conditions like sickle cell disease (SCD) and transfusion-dependent beta-thalassaemia (TDT). The path finally led the way to the recent FDA approval of the first gene therapy drug utilising the CRISPR/Cas9 system to edit autologous CD34+ haematopoietic stem cells in SCD patients (Casgevy). Ongoing research explores the potential of CRISPR technology for cancer therapies, HIV treatment and other complex diseases. Despite its remarkable potential, CRISPR technology faces challenges such as off-target effects, suboptimal delivery systems, long-term safety concerns, scalability, ethical dilemmas and potential repercussions of genetic alterations, particularly in the case of germline editing. Here, we examine the transformative role of CRISPR technologies, including base editing and prime editing approaches, in modifying the genetic and epigenetic codes in the human genome and provide a comprehensive focus, particularly on relevant clinical applications, to unlock the full potential and challenges of gene editing.
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Cytosine and adenosine base editing in human pluripotent stem cells using transient reporters for editing enrichment.
Deaminase fused-Cas9 base editing technologies have enabled precise single-nucleotide genomic editing without the need for the introduction of damaging double-stranded breaks and inefficient homology-directed repair. However, current methods to isolate base-edited cell populations are ineffective, especially when utilized with human pluripotent stem cells, a cell type resistant to genome modification. Here, we outline a series of methods that employ transient reporters of editing enrichment (TREE) to facilitate the highly efficient single-base editing of human cells at precise genomic loci. Briefly, these transient reporters of editing enrichment based methods employ a transient episomal fluorescent reporter that allows for the real-time, flow-cytometry-based enrichment of cells that have had single nucleotide changes at precise genomic locations. This protocol details how these approaches can enable the rapid (~3-4 weeks) and efficient (clonal editing efficiencies >80%) generation of biallelic or multiplexed edited isogenic hPSC lines using adenosine and cytosine base editors.
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Precision A3G base editors for disease modeling and correction.
Cytosine base editors (CBEs) enable efficient cytosine-to-thymine substitutions at targeted genomic loci without introducing double-stranded breaks. Among CBEs, APOBEC3G BEs (A3G-BEs) preferentially edit the second cytosine within a 5'-CC-3' motif in human cells, reducing potential bystander editing. However, A3G-BEs often unintentionally edit multiple CC motifs within their editing window and are limited by protospacer adjacent motif (PAM) constraints imposed by SpCas9, which restricts their applicability. Here, we engineered A3G-BE variants through linker optimization, rational mutagenesis, and the integration of SpG and SpRY Cas9 effectors with relaxed PAM constraints. These improvements enhanced the precision of single-cytosine editing within CC motifs and broadened the targeting scope to previously inaccessible genomic sites. We then validated the engineered A3G-BE variants by precisely installing and correcting cystic fibrosis-causing mutations in HEK293T cells. When applied to 16HBE14o-human bronchial epithelial cells, precise editing modulated cystic fibrosis transmembrane conductance regulator mRNA levels, protein expression, and channel function, establishing precision A3G-BE variants as powerful tools for modeling and treating cystic fibrosis and other human diseases.
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