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Duchenne muscular dystrophy (DMD) affecting 1 in 3500-5000 live male newborns is the frequently fatal genetic disease resulted from various mutations in DMD gene encoding dystrophin protein. About 70% of DMD-causing mutations are exon deletion leading to frameshift of open reading frame and dystrophin deficiency. To facilitate translating human DMD-targeting CRISPR therapeutics into patients, we herein establish a genetically humanized mouse model of DMD by replacing exon 50 and 51 of mouse Dmd gene with human exon 50 sequence. This humanized mouse model recapitulats patient's DMD phenotypes of dystrophin deficiency and muscle dysfunction. Furthermore, we target splicing sites in human exon 50 with adenine base editor to induce exon skipping and robustly restored dystrophin expression in heart, tibialis anterior and diaphragm muscles. Importantly, systemic delivery of base editor via adeno-associated virus in the humanized male mouse model improves the muscle function of DMD mice to the similar level of wildtype ones, indicating the therapeutic efficacy of base editing strategy in treating most of DMD types with exon deletion or point mutations via exon-skipping induction., (© 2024. The Author(s).)

As a miniature RNA-guided endonuclease, IscB is presumed to be the ancestor of Cas9 and to share similar functions. IscB is less than half the size of Cas9 and thus more suitable for in vivo delivery. However, the poor editing efficiency of IscB in eukaryotic cells limits its in vivo applications. Here we describe the engineering of OgeuIscB and its corresponding ωRNA to develop an IscB system that is highly efficient in mammalian systems, named enIscB. By fusing enIscB with T5 exonuclease (T5E), we found enIscB-T5E exhibited comparable targeting efficiency to SpG Cas9 while showing reduced chromosome translocation effects in human cells. Furthermore, by fusing cytosine or adenosine deaminase with enIscB nickase, we generated miniature IscB-derived base editors (miBEs), exhibiting robust editing efficiency (up to 92%) to induce DNA base conversions. Overall, our work establishes enIscB-T5E and miBEs as versatile tools for genome editing., (© 2023. The Author(s), under exclusive licence to Springer Nature America, Inc.)

Catalytically inactive CRISPR-Cas13 (dCas13)-based base editors can achieve the conversion of adenine-to-inosine (A-to-I) or cytidine-to-uridine (C-to-U) at the RNA level, however, the large size of dCas13 protein limits its in vivo applications. Here, a compact and efficient RNA base editor (ceRBE) is reported with high in vivo editing efficiency. The larger dCas13 protein is replaced with a 199-amino acid EcCas6e protein, derived from the Class 1 CRISPR family involved in pre-crRNA processing, and conducted optimization for toxicity and editing efficiency. The ceRBE efficiently achieves both A-to-I and C-to-U base editing with low transcriptome off-target in HEK293T cells. The efficient repair of the DMD Q1392X mutation (68.3±10.1%) is also demonstrated in a humanized mouse model of Duchenne muscular dystrophy (DMD) after AAV delivery, achieving restoration of expression for gene products. The study supports that the compact and efficient ceRBE has great potential for treating genetic diseases., (© 2023 The Authors. Advanced Science published by Wiley-VCH GmbH.)

The type V-F CRISPR-Cas12f system is a strong candidate for therapeutic applications due to the compact size of the Cas12f proteins. In this work, we identify six uncharacterized Cas12f1 proteins with nuclease activity in mammalian cells from assembled bacterial genomes. Among them, OsCas12f1 (433 aa) from Oscillibacter sp. and RhCas12f1 (415 aa) from Ruminiclostridium herbifermentans, which respectively target 5' T-rich Protospacer Adjacent Motifs (PAMs) and 5' C-rich PAMs, show the highest editing activity. Through protein and sgRNA engineering, we generate enhanced OsCas12f1 (enOsCas12f1) and enRhCas12f1 variants, with 5'-TTN and 5'-CCD (D = not C) PAMs respectively, exhibiting much higher editing efficiency and broader PAMs, compared with the engineered variant Un1Cas12f1 (Un1Cas12f1_ge4.1). Furthermore, by fusing the destabilized domain with enOsCas12f1, we generate inducible-enOsCas12f1 and demonstate its activity in vivo by single adeno-associated virus delivery. Finally, dead enOsCas12f1-based epigenetic editing and gene activation can also be achieved in mammalian cells. This study thus provides compact gene editing tools for basic research with remarkable promise for therapeutic applications., (© 2023. The Author(s).)

The CRISPR-Cas9 system is a genomic editing tool widely used in basic research and under investigation for potential applications in gene therapies for human diseases. To accomplish genomic editing, the system requires the expression of a prokaryotic DNA endonuclease enzyme, Cas9, in host cells. Previous studies have mainly focused on the specificity of Cas9 on the host genome, and thus it is unclear whether this bacterium-derived enzyme affects the protein homeostasis of host cells. Here we applied multi-omic analyses, including transcriptome, proteome, phosphoproteome, Cas9-associated protein interactome, protein synthesis, and histone epigenetic modification, to investigate the cellular response of human cells upon the expression of Cas9. We demonstrate that Cas9 has minimal impact on host cells. Our assessment of intracellular effects of Cas9 paves a path for its broad applications in biological studies and potential clinical translations.

The targeting efficiency of knockin sequences via homologous recombination (HR) is generally low. Here we describe a method we call Tild-CRISPR (targeted integration with linearized dsDNA-CRISPR), a targeting strategy in which a PCR-amplified or precisely enzyme-cut transgene donor with 800-bp homology arms is injected with Cas9 mRNA and single guide RNA into mouse zygotes. Compared with existing targeting strategies, this method achieved much higher knockin efficiency in mouse embryos, as well as brain tissue. Importantly, the Tild-CRISPR method also yielded up to 12-fold higher knockin efficiency than HR-based methods in human embryos, making it suitable for studying gene functions in vivo and developing potential gene therapies., (Copyright © 2018 Elsevier Inc. All rights reserved.)

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

Despite rapid progresses in the genome-editing field, in vivo simultaneous overexpression of multiple genes remains challenging. We generated a transgenic mouse using an improved dCas9 system that enables simultaneous and precise in vivo transcriptional activation of multiple genes and long noncoding RNAs in the nervous system. As proof of concept, we were able to use targeted activation of endogenous neurogenic genes in these transgenic mice to directly and efficiently convert astrocytes into functional neurons in vivo. This system provides a flexible and rapid screening platform for studying complex gene networks and gain-of-function phenotypes in the mammalian brain.

Background: The CRISPR/Cas9 system has become an efficient gene editing method for generating cells carrying precise gene mutations, including the rearrangement and deletion of chromosomal segments. However, whether an entire chromosome could be eliminated by this technology is still unknown., Results: Here we demonstrate the use of the CRISPR/Cas9 system to eliminate targeted chromosomes. Using either multiple cleavages induced by a single-guide RNA (sgRNA) that targets multiple chromosome-specific sites or a cocktail of multiple sgRNAs, each targeting one specific site, we found that a sex chromosome could be selectively eliminated in cultured cells, embryos, and tissues in vivo. Furthermore, this approach was able to produce a targeted autosome loss in aneuploid mouse embryonic stem cells with an extra human chromosome and human induced pluripotent stem cells with trisomy 21, as well as cancer cells., Conclusions: CRISPR/Cas9-mediated targeted chromosome elimination offers a new approach to develop animal models with chromosome deletions, and a potential therapeutic strategy for human aneuploidy diseases involving additional chromosomes.

The CRISPR/Cas9 system is an efficient gene-editing method, but the majority of gene-edited animals showed mosaicism, with editing occurring only in a portion of cells. Here we show that single gene or multiple genes can be completely knocked out in mouse and monkey embryos by zygotic injection of Cas9 mRNA and multiple adjacent single-guide RNAs (spaced 10-200 bp apart) that target only a single key exon of each gene. Phenotypic analysis of F0 mice following targeted deletion of eight genes on the Y chromosome individually demonstrated the robustness of this approach in generating knockout mice. Importantly, this approach delivers complete gene knockout at high efficiencies (100% on Arntl and 91% on Prrt2) in monkey embryos. Finally, we could generate a complete Prrt2 knockout monkey in a single step, demonstrating the usefulness of this approach in rapidly establishing gene-edited monkey models.

Targeted integration of transgenes can be achieved by strategies based on homologous recombination (HR), microhomology-mediated end joining (MMEJ) or non-homologous end joining (NHEJ). The more generally used HR is inefficient for achieving gene integration in animal embryos and tissues, because it occurs only during cell division, although MMEJ and NHEJ can elevate the efficiency in some systems. Here we devise a homology-mediated end joining (HMEJ)-based strategy, using CRISPR/Cas9-mediated cleavage of both transgene donor vector that contains guide RNA target sites and ∼800 bp of homology arms, and the targeted genome. We found no significant improvement of the targeting efficiency by the HMEJ-based method in either mouse embryonic stem cells or the neuroblastoma cell line, N2a, compared to the HR-based method. However, the HMEJ-based method yielded a higher knock-in efficiency in HEK293T cells, primary astrocytes and neurons. More importantly, this approach achieved transgene integration in mouse and monkey embryos, as well as in hepatocytes and neurons in vivo, with an efficiency much greater than HR-, NHEJ- and MMEJ-based strategies. Thus, the HMEJ-based strategy may be useful for a variety of applications, including gene editing to generate animal models and for targeted gene therapies.

Precisely targeted genome editing is highly desired for clinical applications. However, the widely used homology-directed repair (HDR)-based genome editing strategies remain inefficient for certain in vivo applications. We here demonstrate a microhomology-mediated end-joining (MMEJ)-based strategy for precisely targeted gene integration in transfected neurons and hepatocytes in vivo with efficiencies up to 20%, much higher (up to 10 fold) than HDR-based strategy in adult mouse tissues. As a proof of concept of its therapeutic potential, we demonstrate the efficacy of MMEJ-based strategy in correction of Fah mutation and rescue of Fah -/- liver failure mice, offering an efficient approach for precisely targeted gene therapies., (Copyright © 2017 The Authors. Published by Elsevier B.V. All rights reserved.)