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A differentiated and durable allogeneic strategy applicable to cell therapies.
BACKGROUND AIMS: Autologous T-cell therapies have shown profound clinical responses; however, their widespread use has been limited primarily as the result of their individualized manufacturing requirements. METHODS: To develop a persistent "off-the-shelf" allogeneic (Allo) approach, a multiplex Nme2Cas9-based cytosine base editor was deployed to knockout select human leukocyte antigens (HLA) class I and II alleles (HLA-A, HLA-B and the class II transactivator [CIITA]) while retaining HLA-C to protect from natural killer (NK) cell rejection. RESULTS AND CONCLUSION: Matching the residual HLA-C allele from homozygous donors to the host prevented rejection of the donor T cells by allogeneic host T and NK cells. Site-specific integration of a tumor-specific CAR or TCR into the TRAC locus using SpyCas9 nuclease and an adeno-associated virus template allowed for a high localized insertion rate while simultaneously removing the endogenous TCR and preventing graft-versus-host disease. Using an optimized T-cell engineering process involving orthogonal CRISPR/Cas9 cleavage and base editors coupled with lipid nanoparticle delivery, we achieved efficient production of Allo-CAR T cells with high editing rates and cell expansion in a scalable manner. These allogeneic T cells demonstrated comparable functional activity to their autologous counterparts in preclinical assays. Moreover, this gene-editing approach significantly minimized the occurrence of chromosomal aberrations. This promising allogeneic approach also has been applied to induced pluripotent stem cells (iPSCs) with triple edits targeting HLA-A, HLA-B and CIITA (TKO). Pancreatic progenitor cells or cardiomyocytes derived from TKO iPSCs were protected from host peripheral blood mononuclear cell-mediated rejection when matched for HLA-C, suggesting potential applications in regenerative medicine applications.
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CRISPRpred(SEQ): a sequence-based method for sgRNA on target activity prediction using traditional machine learning.
BACKGROUND: The latest works on CRISPR genome editing tools mainly employs deep learning techniques. However, deep learning models lack explainability and they are harder to reproduce. We were motivated to build an accurate genome editing tool using sequence-based features and traditional machine learning that can compete with deep learning models. RESULTS: In this paper, we present CRISPRpred(SEQ), a method for sgRNA on-target activity prediction that leverages only traditional machine learning techniques and hand-crafted features extracted from sgRNA sequences. We compare the results of CRISPRpred(SEQ) with that of DeepCRISPR, the current state-of-the-art, which uses a deep learning pipeline. Despite using only traditional machine learning methods, we have been able to beat DeepCRISPR for the three out of four cell lines in the benchmark dataset convincingly (2.174%, 6.905% and 8.119% improvement for the three cell lines). CONCLUSION: CRISPRpred(SEQ) has been able to convincingly beat DeepCRISPR in 3 out of 4 cell lines. We believe that by exploring further, one can design better features only using the sgRNA sequences and can come up with a better method leveraging only traditional machine learning algorithms that can fully beat the deep learning models.
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Spellchecking for the Story of Life With CRISPR-Cas9 and Base, Prime Editors.
Just four letters-A, G, T, and C-make up the alphabet of the genome. It may seem simple, but a small difference in spelling can create mutations that result in life-threatening diseases. Gene variants that cause genetic diseases come in many varieties. Transition point mutations cause conditions such as progeria, the rapid aging disease. Transversion point mutations cause sickle-cell disease and other major disorders. Small insertions can cause Tay-Sachs, which stops nerves working properly and is usually fatal, and deletions can result in cystic fibrosis.
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Programmable C:G to G:C genome editing with CRISPR-Cas9-directed base excision repair proteins.
Many genetic diseases are caused by single-nucleotide polymorphisms. Base editors can correct these mutations at single-nucleotide resolution, but until recently, only allowed for transition edits, addressing four out of twelve possible DNA base substitutions. Here, we develop a class of C:G to G:C Base Editors to create single-base genomic transversions in human cells. Our C:G to G:C Base Editors consist of a nickase-Cas9 fused to a cytidine deaminase and base excision repair proteins. Characterization of >30 base editor candidates reveal that they predominantly perform C:G to G:C editing (up to 90% purity), with rAPOBEC-nCas9-rXRCC1 being the most efficient (mean 15.4% and up to 37% without selection). C:G to G:C Base Editors target cytidine in WCW, ACC or GCT sequence contexts and within a precise three-nucleotide window of the target protospacer. We further target genes linked to dyslipidemia, hypertrophic cardiomyopathy, and deafness, showing the therapeutic potential of these base editors in interrogating and correcting human genetic diseases.
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Efficient In Vivo Liver-Directed Gene Editing Using CRISPR/Cas9.
In vivo tissue-specific genome editing at the desired loci is still a challenge. Here, we report that AAV9-delivery of truncated guide RNAs (gRNAs) and Cas9 under the control of a computationally designed hepatocyte-specific promoter lead to liver-specific and sequence-specific targeting in the mouse factor IX (F9) gene. The efficiency of in vivo targeting was assessed by T7E1 assays, site-specific Sanger sequencing, and deep sequencing of on-target and putative off-target sites. Though AAV9 transduction was apparent in multiple tissues and organs, Cas9 expression was restricted mainly to the liver, with only minimal or no expression in other non-hepatic tissues. Consequently, the insertions and deletion (indel) frequency was robust in the liver (up to 50%) in the desired target loci of the F9 gene, with no evidence of targeting in other organs or other putative off-target sites. This resulted in a substantial loss of FIX activity and the emergence of a bleeding phenotype, consistent with hemophilia B. The in vivo efficacy of the truncated gRNA was as high as that of full-length gRNA. Cas9 expression was transient in neonates, representing an attractive "hit-and-run" paradigm. Our findings have potentially broad implications for somatic gene targeting in the liver using the CRISPR/Cas9 platform.
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CRISPR guides induce gene silencing in plants in the absence of Cas.
BACKGROUND: RNA-targeting CRISPR-Cas can provide potential advantages over DNA editing, such as avoiding pleiotropic effects of genome editing, providing precise spatiotemporal regulation, and expanded function including antiviral immunity. RESULTS: Here, we report the use of CRISPR-Cas13 in plants to reduce both viral and endogenous RNA. Unexpectedly, we observe that crRNA designed to guide Cas13 could, in the absence of the Cas13 protein, cause substantial reduction in RNA levels as well. We demonstrate Cas13-independent guide-induced gene silencing (GIGS) in three plant species, including stable transgenic Arabidopsis. Small RNA sequencing during GIGS identifies the production of small RNA that extend beyond the crRNA expressed sequence in samples expressing multi-guide crRNA. Additionally, we demonstrate that mismatches in guide sequences at position 10 and 11 abolish GIGS. Finally, we show that GIGS is elicited by guides that lack the Cas13 direct repeat and can extend to Cas9 designed crRNA of at least 28 base pairs, indicating that GIGS can be elicited through a variety of guide designs and is not dependent on Cas13 crRNA sequences or design. CONCLUSIONS: Collectively, our results suggest that GIGS utilizes endogenous RNAi machinery despite the fact that crRNA are unlike canonical triggers of RNAi such as miRNA, hairpins, or long double-stranded RNA. Given similar evidence of Cas13-independent silencing in an insect system, it is likely GIGS is active across many eukaryotes. Our results show that GIGS offers a novel and flexible approach to RNA reduction with potential benefits over existing technologies for crop improvement and functional genomics.
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CRISPR-Directed Therapeutic Correction at the NCF1 Locus Is Challenged by Frequent Incidence of Chromosomal Deletions.
Resurrection of non-processed pseudogenes may increase the efficacy of therapeutic gene editing, upon simultaneous targeting of a mutated gene and its highly homologous pseudogenes. To investigate the potency of this approach for clinical gene therapy of human diseases, we corrected a pseudogene-associated disorder, the immunodeficiency p47 phox -deficient chronic granulomatous disease (p47 phox CGD), using clustered regularly interspaced short palindromic repeats-associated nuclease Cas9 (CRISPR-Cas9) to target mutated neutrophil cytosolic factor 1 (NCF1). Being separated by less than two million base pairs, NCF1 and two pseudogenes are closely co-localized on chromosome 7. In healthy people, a two-nucleotide GT deletion (ΔGT) is present in the NCF1B and NCF1C pseudogenes only. In the majority of patients with p47 phox CGD, the NCF1 gene is inactivated due to a ΔGT transfer from one of the two non-processed pseudogenes. Here we demonstrate that concurrent targeting and correction of mutated NCF1 and its pseudogenes results in therapeutic CGD phenotype correction, but also causes potentially harmful chromosomal deletions between the targeted loci in a p47 phox -deficient CGD cell line model. Therefore, development of genome-editing-based treatment of pseudogene-related disorders mandates thorough safety examination, as well as technological advances, limiting concurrent induction of multiple double-strand breaks on a single chromosome.
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Heterochromatin delays CRISPR-Cas9 mutagenesis but does not influence the outcome of mutagenic DNA repair.
Genome editing occurs in the context of chromatin, which is heterogeneous in structure and function across the genome. Chromatin heterogeneity is thought to affect genome editing efficiency, but this has been challenging to quantify due to the presence of confounding variables. Here, we develop a method that exploits the allele-specific chromatin status of imprinted genes in order to address this problem in cycling mouse embryonic stem cells (mESCs). Because maternal and paternal alleles of imprinted genes have identical DNA sequence and are situated in the same nucleus, allele-specific differences in the frequency and spectrum of mutations induced by CRISPR-Cas9 can be unequivocally attributed to epigenetic mechanisms. We found that heterochromatin can impede mutagenesis, but to a degree that depends on other key experimental parameters. Mutagenesis was impeded by up to 7-fold when Cas9 exposure was brief and when intracellular Cas9 expression was low. In contrast, the outcome of mutagenic DNA repair was unaffected by chromatin state, with similar efficiencies of homology-directed repair (HDR) and deletion spectra on maternal and paternal chromosomes. Combined, our data show that heterochromatin imposes a permeable barrier that influences the kinetics, but not the endpoint, of CRISPR-Cas9 genome editing and suggest that therapeutic applications involving low-level Cas9 exposure will be particularly affected by chromatin status.
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CRISPR-prime editing, a versatile genetic tool to create specific mutations with a single nucleotide resolution in Leptospira.
Leptospirosis, caused by pathogenic bacteria from the genus Leptospira, is a global zoonosis responsible for more than one million human cases and 60,000 deaths annually. The disease also affects many domestic animal species. Historically, genetic manipulation of Leptospira has been difficult to perform, resulting in limited knowledge on pathogenic mechanisms of disease and the identification of virulence factors. The application of CRISPR/Cas9 and its variations have helped fill these gaps but the generation of knockout mutants remains challenging because double-strand breaks (DSBs) inflicted by Cas9 nuclease are lethal to Leptospira cells. The novel CRISPR prime editing (PE) strategy is the first precise genome-editing technology that allows deletions, insertions, and base substitutions without introducing DSBs. This revolutionary technique utilizes a nickase Cas9 that cleaves a single strand of DNA, coupled with an engineered reverse transcriptase and a modified single-guide RNA (termed prime editing guide RNA) containing an extended 3' end with the desired edits. We demonstrate the application of CRISPR-PE in both saprophytic and pathogenic Leptospira from multiple species and serovars by introducing deletions or insertions into target DNA with a remarkable precision of just one nucleotide. Additionally, we demonstrate the ability to genetically manipulate Leptospira borgpetersenii, a prevalent pathogenic species of humans, domestic cattle, and wildlife animals. Rapid plasmid loss by mutated strains in liquid culture allows for the generation of knockout strains without selective markers, which can be readily used to elucidate virulence factors and develop optimized bacterin and/or live vaccines against leptospirosis.IMPORTANCELeptospirosis is a geographically widespread bacterial zoonosis. Genetic manipulation of pathogenic Leptospira spp. has been laborious and difficult to perform, limiting our ability to understand how leptospires cause disease. The application of the CRISPR/Cas9 system to Leptospira enhanced our ability to generate knockdown and knockout mutants; however, the latter remains challenging. Here, we demonstrate the application of the CRISPR prime editing technique in Leptospira, allowing the generation of knockout mutants in several pathogenic species, with mutations comprising just a single nucleotide resolution. Notably, we generated a mutant in the Leptospira borgpetersenii background, a prevalent pathogenic species of humans and cattle. Our application of this method opens new avenues for studying pathogenic mechanisms of Leptospira and the identification of virulence factors across multiple species. These methods can also be used to facilitate the generation of marker-less knockout strains for updated and improved bacterin and/or live vaccines.
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A CRISPR-based method for testing the essentiality of a gene.
The CRISPR/Cas9 system is a powerful method of editing genes by randomly introducing errors into the target sites. Here, we describe a CRISPR-based test for gene essentiality (CRISPR-E test) that allows the identification of essential genes. Specifically, we use sgRNA-mediated CRISPR/Cas9 to target the open reading frame of a gene in the genome and analyze the in-frame (3n) and frameshift (3n + 1 and 3n + 2) mutations in the targeted region of the gene in surviving cells. If the gene is non-essential, the cells would carry both in-frame (3n) and frameshift (3n + 1 and 3n + 2) mutations. In contrast, the cells would carry only in-frame (3n) mutations if the targeted gene is essential, and this selective elimination of frameshift (3n + 1 and 3n + 2) mutations of the gene indicate its essentiality. As a proof of concept, we have used this CRISPR-E test in the model organism Dictyostelium discoideum to demonstrate that Dync1li1 is an essential gene while KIF1A and fAR1 are not. We further propose a simple method for quantifying the essentiality of a gene using the CRISPR-E test.
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