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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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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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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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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-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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Gene Editing-Driven Engineering of Microbial Systems and Metabolites for Precision Medicine and Sustainable Environmental Remediation.
The engineering of microorganisms is undergoing a fundamental paradigm shift, transitioning from the construction of static cell factories to the programming of dynamically responsive living materials. However, translating molecular interventions into robust macroscopic functions requires overcoming distinct microbial-specific barriers, including delivery bottlenecks and genetic stability. In this review, we establish a unified Edit-Reprogram-Functionalize conceptual framework that systematically delineates transient genetic regulation from permanent genomic engineering. We critically examine the evolutionary trajectories of five foundational technologies: plasmid engineering, CRISPR-Cas systems, base editors, prime editors, and enzyme engineering. Rather than analyzing these toolsets in isolation, we map their convergence into an integrated engineering continuum that drives the precise synthesis of two distinct output classes: engineered living microbial materials and robust microbial metabolite-derived materials. By evaluating representative breakthroughs-from ultrasound-actuated bacterial therapeutics to ultra-tough, biosynthesized protein composites-through the strict lens of host-dependent constraints, we reveal the mechanistic principles governing successful preclinical translation. Finally, we propose an actionable roadmap centered on systemic miniaturization, closed-loop control, and multi-scale integration, providing a definitive blueprint for the next generation of precision medicine, advanced biomanufacturing, and ecological remediation.
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Genome editing in butterflies reveals that spalt promotes and Distal-less represses eyespot colour patterns.
Butterfly eyespot colour patterns are a key example of how a novel trait can appear in association with the co-option of developmental patterning genes. Little is known, however, about how, or even whether, co-opted genes function in eyespot development. Here we use CRISPR/Cas9 genome editing to determine the roles of two co-opted transcription factors that are expressed during early eyespot determination. We found that deletions in a single gene, spalt, are sufficient to reduce or completely delete eyespot colour patterns, thus demonstrating a positive regulatory role for this gene in eyespot determination. Conversely, and contrary to previous predictions, deletions in Distal-less (Dll) result in an increase in the size and number of eyespots, illustrating a repressive role for this gene in eyespot development. Altogether our results show that the presence, absence and shape of butterfly eyespots can be controlled by the activity of two co-opted transcription factors.
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Dual-sgRNA CRISPR/Cas9 knockout of PD-L1 in human U87 glioblastoma tumor cells inhibits proliferation, invasion, and tumor-associated macrophage polarization.
Programmed death ligand 1 (PD-L1) plays a key role in glioblastoma multiforme (GBM) immunosuppression, vitality, proliferation, and migration, and is therefore a promising target for treating GBM. CRISPR/Cas9-mediated genomic editing can delete both cell surface and intracellular PD-L1. This systemic deliverable genomic PD-L1 deletion system can be used as an effective anti-GBM therapy by inhibiting tumor growth and migration, and overcoming immunosuppression. To target PD-L1 for CRISPR/Cas9 gene editing, we first identified two single guide RNA (sgRNA) sequences located on PD-L1 exon 3. The first sgRNA recognizes the forward strand of human PD-L1 near the beginning of exon 3 that allows editing by Cas9 at approximately base pair 82 (g82). The second sgRNA recognizes the forward strand of exon 3 that directs cutting at base pair 165 (g165). A homology-directed repair template (HDR) combined with the dual-sgRNAs was used to improve PD-L1 knockout specificity and efficiency. sgRNAs g82 and g165 were cloned into the multiplex CRISPR/Cas9 assembly system and co-transfected with the HDR template in human U87 GBM cells (g82/165 + HDR). T7E1 analysis suggests that the dual-sgRNA CRISPR/Cas9 strategy with a repair template was capable of editing the genomic level of PD-L1. This was further confirmed by examining PD-L1 protein levels by western blot and immunofluorescence assays. Western blot analysis showed that the dual-sgRNAs with the repair template caused a 64% reduction of PD-L1 protein levels in U87 cells, while immunostaining showed a significant reduction of intracellular PD-L1. PD-L1 deletion inhibited proliferation, growth, invasion and migration of U87 cells, indicating intracellular PD-L1 is necessary for tumor progression. Importantly, U87 cells treated with g82/165 + HDR polarized tumor-associated macrophages (TAM) toward an M1 phenotype, as indicated by an increase in TNF-α and a decrease in IL-4 secretions. This was further confirmed with flow cytometry that showed an increase in the M1 markers Ly6C + and CD80 +, and a decrease in the M2 marker CD206 + both in vitro and in vivo. Utilizing dual-sgRNAs and an HDR template with the CRISPR/Cas9 gene-editing system is a promising avenue for the treatment of GBM.
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PhieCBEs: Plant High-Efficiency Cytidine Base Editors with Expanded Target Range.
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