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Found: 26 149
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Application of CRISPR-Cas9 gene editing for congenital heart disease.
Clustered regularly interspaced short palindromic repeats and CRISPR-associated protein 9 (CRISPR-Cas9) is an ancient prokaryotic defense system that precisely cuts foreign genomic DNA under the control of a small number of guide RNAs. The CRISPR-Cas9 system facilitates efficient double-stranded DNA cleavage that has been recently adopted for genome editing to create or correct inherited genetic mutations causing disease. Congenital heart disease (CHD) is generally caused by genetic mutations such as base substitutions, deletions, and insertions, which result in diverse developmental defects and remains a leading cause of birth defects. Pediatric CHD patients exhibit a spectrum of cardiac abnormalities such as septal defects, valvular defects, and abnormal chamber development. CHD onset occurs during the prenatal period and often results in early lethality during childhood. Because CRISPR-Cas9-based genome editing technology has gained considerable attention for its potential to prevent and treat diseases, we will review the CRISPR-Cas9 system as a genome editing tool and focus on its therapeutic application for CHD.
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Revolutionizing Tuberculosis Management With Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas Technology: A Comprehensive Literature Review.
Clustered regularly interspaced short palindromic repeats (CRISPR)-Cas systems have gained attention for their revolutionary potential in tuberculosis (TB) management, providing a novel approach to both diagnostics and treatment. This technology, renowned for its ability to accurately target and modify genetic material, offers a promising solution to the limitations of current TB diagnostic methods, which often rely on time-consuming culture techniques or polymerase chain reaction (PCR)-based assays. One of the key advantages of CRISPR-Cas systems is their high specificity and sensitivity, making them well-suited for detecting Mycobacterium tuberculosis, even in low-bacterial-load samples. Techniques such as CRISPR-Cas12 and Cas13 have been employed for rapid detection, utilizing their trans-cleavage activity to produce a fluorescent signal upon recognition of the TB genome. Furthermore, these methods often use isothermal amplification techniques like recombinase polymerase amplification (RPA) or loop-mediated isothermal amplification (LAMP), which require less equipment compared to traditional PCR. Beyond diagnostics, CRISPR-Cas technologies show promise in studying TB resistance mechanisms and potentially treating drug-resistant strains. Genome-editing capabilities enable researchers to manipulate the M. tuberculosis genome, investigating genes linked to virulence or antibiotic resistance. Although challenges such as the development of multiplexed CRISPR assays for detecting multiple mutations simultaneously remain, advancements continue to improve the technology's practicality for clinical use. Incorporating CRISPR into TB management could enhance early detection, inform personalized treatment, and potentially contribute to developing more effective therapies, especially in regions where TB remains a significant public health threat.
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Prime editing outperforms homology-directed repair as a tool for CRISPR-mediated variant knock-in in zebrafish.
Zebrafish serve as a valuable model organism for studying human genetic diseases. While generating knockout lines is relatively straightforward, introducing precise disease-specific genetic variants by knock-in (KI) remains challenging. KI lines, however, enable more accurate studies of molecular and physiological consequences of genetic diseases. Their generation is often hampered by low editing efficiency (EE) and potential off-target effects. Here, we optimized conventional CRISPR-Cas9-mediated homology-directed repair (HDR) strategies for precise KI of genetic variants in zebrafish and compared their efficacy with prime editing, a recently developed technique that is not yet commonly used. Using next-generation sequencing, we determined KI EE by HDR for six unique base-pair substitutions in three different zebrafish genes. We assessed the effect of (1) varying Cas9 amounts, (2) HDR templates with chemical modifications to improve integration efficiency, (3) different microinjection procedures and (4) introduction of additional synonymous guide-blocking variants in the HDR template. Increasing Cas9 amounts augmented KI EE, with optimal injected amounts of Cas9 between 200 pg and 800 pg. The use of Alt-R HDR templates further increased KI EE, while guide-blocking modifications did not. Injecting components directly into the cell was not superior to injections into the yolk. Prime editing, however, increased EE up to fourfold and expanded the F0 founder pool for four targets compared with conventional HDR editing, with fewer off-target effects. Therefore, prime editing is a very promising methodology for improving the creation of precise genomic edits in zebrafish, facilitating the modeling of human diseases.
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Rapid customization of base editors via machine learning-powered combinatorial mutagenesis.
Being able to control the complementarity and hindrance between target DNA and base editor proteins enables precise, bystander-free editing. Here, we combined combinatorial mutagenesis with machine learning to analyze and engineer these interactions at scale. By profiling DNA motif preferences across 160,000 evoAPOBEC1 and 64 million TadA variants in human cells, we used as little as 0.004% of the mutational landscape to make predictions. This identified variants with motif-specific activity and eliminated residual adenine editing in cytosine base editors. In correcting >800 disease-associated mutations, our variants outperformed previous versions in precluding unintended edits at purine motifs, achieving undetectable bystander edits in 50% of cases. Additionally, a pre-trained, structure-based deep learning model predicted functional TadA variants with 63% success across 2026 variants spanning 26 amino acid sites, without experimental data and in a single prediction round. These approaches streamline the re-engineering of base editors for enhanced precision tailored to specific targets.
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A modified glycosylase base editor without predictable DNA off-target effects.
Glycosylase base editor (GBE) can induce C-to-G transversion in mammalian cells, showing great promise for the treatment of human genetic disorders. However, the limited efficiency of transversion and the possibility of off-target effects caused by Cas9 restrict its potential clinical applications. In our recent study, we have successfully developed TaC9-CBE and TaC9-ABE by separating nCas9 and deaminase, which eliminates the Cas9-dependent DNA off-target effects without compromising editing efficiency. We developed a novel GBE called TaC9-GBEYE1, which utilizes the deaminase and UNG-nCas9 guided by TALE and sgRNA, respectively. TaC9-GBEYE1 showed comparable levels of on-target editing efficiency to traditional GBE at 19 target sites, without any off-target effects caused by Cas9 or TALE. The TaC9-GBEYE1 is a safe tool for gene therapy.
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Development of deaminase-free T-to-S base editor and C-to-G base editor by engineered human uracil DNA glycosylase.
DNA base editors enable direct editing of adenine (A), cytosine (C), or guanine (G), but there is no base editor for direct thymine (T) editing currently. Here we develop two deaminase-free glycosylase-based base editors for direct T editing (gTBE) and C editing (gCBE) by fusing Cas9 nickase (nCas9) with engineered human uracil DNA glycosylase (UNG) variants. By several rounds of structure-informed rational mutagenesis on UNG in cultured human cells, we obtain gTBE and gCBE with high activity of T-to-S (i.e., T-to-C or T-to-G) and C-to-G conversions, respectively. Furthermore, we conduct parallel comparison of gTBE/gCBE with those recently developed using other protein engineering strategies, and find gTBE/gCBE show the outperformance. Thus, we provide several base editors, gTBEs and gCBEs, with corresponding engineered UNG variants, broadening the targeting scope of base editors.
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Application of Base Editor-Mediated Genome Editing in Mouse Retina.
Base editor is a newly developed genome editing technology that enables conversion of single nucleotides without DNA double-strand breaks (DSB) and maintains a low rate of insertion-deletion (INDEL) errors. With these flexibility and safety, base editor has been widely used in many fields, including inherited retinal disease. The majority of retinal genome editing requires intravitreal and subretinal injection delivery of the therapeutic vector in order to transduce the target cells. Here, we provide an application guide of base editor as performed in the mouse retina.
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β Cell Replacement after Gene Editing of a Neonatal Diabetes-Causing Mutation at the Insulin Locus.
Permanent neonatal diabetes mellitus (PNDM) can be caused by insulin mutations. We generated induced pluripotent stem cells from fibroblasts of a patient with PNDM and undetectable insulin at birth due to a homozygous mutation in the translation start site of the insulin gene. Differentiation of mutant cells resulted in insulin-negative endocrine stem cells expressing MAFA, NKX6.1, and chromogranin A. Correction of the mutation in stem cells and differentiation to pancreatic endocrine cells restored insulin production and insulin secretion to levels comparable to those of wild-type cells. Grafting of corrected cells into mice, followed by ablating mouse β cells using streptozotocin, resulted in normal glucose homeostasis, including at night, and the stem cell-derived grafts adapted insulin secretion to metabolic changes. Our study provides proof of principle for the generation of genetically corrected cells autologous to a patient with non-autoimmune insulin-dependent diabetes. These cases should be readily amenable to autologous cell therapy.
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Efficient Method for Generating Point Mutations in the Vaccinia Virus Genome Using CRISPR/Cas9.
The vaccinia virus (VACV) was previously used as a vaccine for smallpox eradication. Nowadays, recombinant VACVs are developed as vaccine platforms for infectious disease prevention and cancer treatment. The conventional method for genome editing of the VACV is based on homologous recombination, which is poorly efficient. Recently, the use of CRISPR/Cas9 technology was shown to greatly improve the speed and efficiency of the production of recombinant VACV expressing a heterologous gene. However, the ability to rapidly recover viruses bearing single nucleotide substitutions is still challenging. Notwithstanding, ongoing studies on the VACV and its interaction with the host cell could benefit from viral gene targeted mutagenesis. Here, we present a modified version of the CRISPR/Cas9 system for the rapid selection of mutant VACV carrying point mutations. For this purpose, we introduced a silent mutation into the donor gene (which will replace the wildtype gene) that serves a double function: it is located in the PAM (NGG) sequence, which is essential for Cas9 cleavage, and it alters a restriction site. This silent mutation, once introduced into the VACV genome, allows for rapid selection and screening of mutant viruses carrying a mutation of interest in the targeted gene. As a proof of concept, we produced several recombinant VACVs, with mutations in the E9L gene, upon which, phenotypic analysis was performed.
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Versatile and efficient in vivo genome editing with compact Streptococcus pasteurianus Cas9.
Compact CRISPR-Cas9 systems that can be packaged into an adeno-associated virus (AAV) show promise for gene therapy. However, the requirement of protospacer adjacent motifs (PAMs) restricts the target scope. To expand this repertoire, we revisited and optimized a small Cas9 ortholog derived from Streptococcus pasteurianus (SpaCas9) for efficient genome editing in vivo. We found that SpaCas9 enables potent targeting of 5'-NNGYRA-3' PAMs, which are distinct from those recognized by currently used small Cas9s; the Spa-cytosine base editor (CBE) and Spa-adenine base editor (ABE) systems efficiently generated robust C-to-T and A-to-G conversions both in vitro and in vivo. In addition, by exploiting natural variation in the PAM-interacting domain, we engineered three SpaCas9 variants to further expand the targeting scope of compact Cas9 systems. Moreover, mutant mice with efficient disruption of the Tyr gene were successfully generated by microinjection of SpaCas9 mRNA and the corresponding single guide RNA (sgRNA) into zygotes. Notably, all-in-one AAV delivery of SpaCas9 targeting the Pcsk9 gene in adult mouse liver produced efficient genome-editing events and reduced its serum cholesterol. Thus, with distinct PAMs and a small size, SpaCas9 will broaden the CRISPR-Cas9 toolsets for efficient gene modifications and therapeutic applications.
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