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Repurposing base editors for targeted knockin and simultaneous multiplex knockouts to generate allo-CAR T cells with minimal translocations.
Multiplex genome editing of cellular therapies frequently requires multiple DNA double-strand breaks (DSBs), which can induce genotoxicity through chromosomal rearrangements and large deletions. Base editors enable targeted sequence changes with minimal DSBs and are widely used for gene disruption, but their capacity for transgene insertion has remained unexplored. Here, we have developed base editor-mediated knockin (BEKI), a non-viral platform combining transgene insertion with multiplex gene disruption using a single enzyme. BEKI repurposes the base editor's Cas9 nickase domain to generate paired nicks (inducing a localized DSB) at the knockin locus while achieving multiplex knockouts through base editing. Optimized guide RNA orientation and spacing enabled efficient transgene insertion across multiple T cell-relevant genomic loci. DNA-PK inhibition enhanced knockin efficiency but increased kilobase-scale deletions, which were mitigated by co-inhibition of Polθ. Compared with multiplex Cas9 editing, BEKI markedly reduced chromosomal translocations while preserving cell viability. BEKI supported targeted chimeric antigen receptor (CAR) knockin alongside up to 10 simultaneous gene knockouts, enabling the generation of allogeneic CAR T cells with enhanced cytokine secretion and resistance to immunosuppressants and allo-rejection. Together, BEKI provides a streamlined and scalable strategy for multiplex CAR T cell engineering with improved genomic stability, advancing safer next-generation cell therapies for cancer and autoimmune diseases.
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Genome Editing for Familial Hemophagocytic Lymphohistiocytosis: Design Principles, Challenges, and Translational Perspectives.
Hemophagocytic lymphohistiocytosis (HLH) is a life-threatening hyperinflammatory syndrome caused by genetic defects in cytotoxic lymphocyte function. Current therapies can control disease activity, but transplantation of allogeneic hematopoietic stem and progenitor cells (HSPCs) remains the only curative option and is associated with substantial risks. These limitations have accelerated development of genome editing approaches enabling precise correction of disease-causing mutations in autologous cells. Familial HLH (FHL) represents a compelling target for genome editing, but successful and safe clinical translation has remained challenging. Preclinical studies demonstrate that targeted editing of key genes, such as PRF1 and UNC13D, can restore cytotoxic function in HSPCs and T cells. Translation to the clinic, however, depends on multiple factors, including the choice of target cell population, the level of functional correction required, and gene-specific constraints such as locus complexity and regulation of gene expression. In this review, we synthesize current progress in genome editing for FHL and highlight critical biological and technical barriers to clinical implementation. We propose a conceptual framework for designing genome editing strategies tailored to FHL, emphasizing the alignment of editing platform, gene architecture, and cellular context to enable effective and clinically translatable therapies.
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High-efficient and precise base editing of C•G to T•A in the allotetraploid cotton (Gossypium hirsutum) genome using a modified CRISPR/Cas9 system.
The base-editing technique using CRISPR/nCas9 (Cas9 nickase) or dCas9 (deactivated Cas9) fused with cytidine deaminase is a powerful tool to create point mutations. In this study, a novel G. hirsutum-Base Editor 3 (GhBE3) base-editing system has been developed to create single-base mutations in the allotetraploid genome of cotton (Gossypium hirsutum). A cytidine deaminase sequence (APOBEC) fused with nCas9 and uracil glycosylase inhibitor (UGI) was inserted into our CRISPR/Cas9 plasmid (pRGEB32-GhU6.7). Three target sites were chosen for two target genes, GhCLA and GhPEBP, to test the efficiency and accuracy of GhBE3. The editing efficiency ranged from 26.67 to 57.78% at the three target sites. Targeted deep sequencing revealed that the C→T substitution efficiency within an 'editing window', approximately six-nucleotide windows of -17 to -12 bp from the PAM sequence, was up to 18.63% of the total sequences. The 27 most likely off-target sites predicted by CRISPR-P and Cas-OFFinder tools were analysed by targeted deep sequencing, and it was found that rare C→T substitutions (average < 0.1%) were detected in the editing windows of these sites. Furthermore, whole-genome sequencing analyses on two GhCLA-edited and one wild-type plants with about 100× depth showed that no bona fide off-target mutations were detectable from 1500 predicted potential off-target sites across the genome. In addition, the edited bases were inherited to T1 progeny. These results demonstrate that GhBE3 has high specificity and accuracy for the generation of targeted point mutations in allotetraploid cotton.
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Enhancement of Gene Editing and Base Editing with Therapeutic Ribonucleoproteins through In Vivo Delivery Based on Absorptive Silica Nanoconstruct.
Key to the widespread and secure application of genome editing tools is the safe and effective delivery of multiple components of ribonucleoproteins (RNPs) into single cells, which remains a biological barrier to their clinical application. To overcome this issue, a robust RNP delivery platform based on a biocompatible sponge-like silica nanoconstruct (SN) for storing and directly delivering therapeutic RNPs, including Cas9 nuclease RNP (Cas9-RNP) and base editor RNP (BE-RNP) is designed. Compared with commercialized material such as lipid-based methods, up to 50-fold gene deletion and 10-fold base substitution efficiency is obtained with a low off-target efficiency by targeting various cells and genes. In particular, gene correction is successfully induced by SN-based delivery through intravenous injection in an in vivo solid-tumor model and through subretinal injection in mouse eye. Moreover, because of its low toxicity and high biodegradability, SN has negligible effect on cellular function of organs. As the engineered SN can overcome practical challenges associated with therapeutic RNP application, it is strongly expected this platform to be a modular RNPs delivery system, facilitating in vivo gene deletion and editing.
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Off-Target Editing by CRISPR-Guided DNA Base Editors.
Base editing is a genome editing strategy that induces specific single-nucleotide changes within genomic DNA. Two major DNA base editors, cytosine base editors and adenine base editors, that consist of a Cas9 protein linked to a deaminase enzyme that catalyzes targeted base conversion directed by a single-guide RNA have been developed. This strategy has been used widely for precise genome editing because, unlike CRISPR-Cas nuclease-based genome editing systems, this strategy does not create double-strand DNA breaks that often result in high levels of undesirable indels. However, recent papers have reported that DNA base editors can cause substantial off-target editing in both genomic DNA and RNA. The off-target editing described in these studies is primarily independent of guide RNA and arises from the promiscuous reactivity of the deaminase enzymes used in DNA base editors. In this Perspective, we discuss the development of DNA base editors, the guide RNA-independent off-target activity reported in recent studies, and strategies that improve the selectivity of DNA base editors.
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CRISPR base editing lowers cholesterol in monkeys.
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Development of a universal antibiotic resistance screening system for efficient enrichment of C-to-G and A-to-G base editing.
To expand the scope of genomic editing, a C-to-G transversion-based editor called CGBE has been developed for precise single-nucleotide genomic editing. However, limited editing efficiency and product purity have hindered the development and application of CGBE. In this study, we introduced the Puromycin-Resistance Screening System, referred to as CGBE/ABE-PRSS, to select genetically modified cells via the CGBE or ABE editors. The CGBE/ABE-PRSS system significantly improves the enrichment efficiency of CGBE- or ABE-modified cells, showing enhancements of up to 59.6 % compared with the controls. Our findings indicate that the CGBE/ABE-PRSS, when driven by the CMV promoter, results in a higher enrichment of edited cells compared to the CAG and EF1α promoters. Furthermore, we demonstrate that this system is compatible with different versions of both CGBE and ABE, enabling various cell species and simultaneous multiplexed genome editing without any detectable random off-targets. In conclusion, our developed CGBE/ABE-PRSS system facilitates the selection of edited cells and holds promise in both basic engineering and gene therapy applications.
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An Agrobacterium-mediated base editing approach generates transgene-free edited banana.
Genome editing for the development of improved varieties is supported by the possibility of segregating out the editor T-DNA cassette after genome editing in many crop species. Removal of the T-DNA cassette prevents potential continuous editing activity in the transformed plant and furthermore facilitates regulatory approval. While transgene outcrossing of exogenous sequences is possible for many crops, this is not the case for vegetatively propagated and sterile crops, such as Cavendish bananas. Therefore, gene editing techniques leading to transgene-free edited plants are essential to untap the potential of genome editing for those crops. Here, we present a method for transgene-free gene editing in sterile banana (Musa spp.) through a co-editing strategy. A novel Agrobacterium tumefaciens-mediated transgene-free gene editing approach combining embryogenesis and chlorsulfuron selection was established in sterile banana and validated through whole genome sequencing. Editing of the acetolactate synthase (MaALS) genes in banana using a plant base editor allows effective selection of edited plants. Moreover, transgene-free plantlets were regenerated with mutations at two target sites, indicating that the strategy can be used to target multiple genomic sites. The presented method allows for efficient transgene-free gene editing and represents the first report of a co-editing strategy in sterile crop species.
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Multiple gene substitution by Target-AID base-editing technology in tomato.
The use of Target activation-induced cytidine deaminase (Target-AID) base-editing technology with the CRISPR-Cas 9 system fused with activation-induced cytidine deaminase (AID) resulted in the substitution of a cytidine with a thymine. In previous experiments focusing on a single target gene, this system has been reported to work in several plant species, including tomato (Solanum lycopersicum L.). In this research, we used Target-AID technology to target multiple genes related to carotenoid accumulation in tomato. We selected 3 genes, SlDDB1, SlDET1 and SlCYC-B, for their roles in carotenoid accumulation. Among 12 edited T0 lines, we obtained 10 independent T0 lines carrying nucleotide substitutions in the three targeted genes, with several allelic versions for each targeted gene. The two edited lines showed significant differences in carotenoid accumulation. These results demonstrate that Target-AID technology is a highly efficient tool for targeting multiple genes with several allelic versions.
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Beyond Precision: A Multidimensional Framework for Selecting Genetic Medicine Platforms.
Gene therapy is undergoing continued clinical translation and technological development. This progress has been marked by regulatory approvals and broadened therapeutic indications across genetic, metabolic, and oncologic diseases and disorders. The field has evolved over decades from early viral-mediated gene addition to approaches capable of targeted editing, regulation, or replacement of genetic information. These systems include base and prime editors, epigenetic modulators, CRISPR-Cas, RNA therapeutics and programmable integration platforms. When paired with increasingly sophisticated viral and nonviral delivery strategies, these technologies enable greater control over tissue targeting, duration of activity, and therapeutic exposure. Recent clinical successes, including approved ex vivo CRISPR-based therapies for hemoglobinopathies, in vivo CRISPR editing for transthyretin amyloidosis, and emerging clinical applications of base and prime editing, provide growing clinical evidence for the feasibility of genetic medicines. However, technological advancement has also made platform selection increasingly complex. Therapeutic performance is determined not by editing efficiency alone, but by the interaction among genetic precision, temporal control, dosage tunability, delivery efficiency, durability, and disease-specific safety requirements. A molecularly efficient platform may still have limited therapeutic value if it cannot reach the disease-relevant cell population at sufficient and safe exposure. In this review, we examine recent technological and clinical advances in genetic medicine with particular emphasis on developments during the past approximately five years. We propose a multidimensional framework in which gene therapy platforms are evaluated according to three intrinsic properties-genetic precision, temporal control, and dosage tunability-while delivery, clinical maturity, and disease context act as major translational constraints. This framework highlights that no single platform is universally optimal; rather, successful therapeutic design depends on matching the biological characteristics of the intervention to the requirements of the disease and target tissue. Remaining challenges in extrahepatic delivery, genomic safety, immunogenicity, manufacturing, and long-term monitoring remain important determinants of broader clinical implementation.
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