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Efficient CRISPR/Cas12a-Based Genome-Editing Toolbox for Metabolic Engineering in Methanococcus maripaludis.
The rapid-growing and genetically tractable methanogen Methanococcus maripaludis is a promising host organism for the biotechnological conversion of carbon dioxide and renewable hydrogen to fuels and value-added products. Expansion of its product scope through metabolic engineering necessitates reliable and efficient genetic tools, particularly for genome edits that affect the primary metabolism and cell growth. Here, we have designed a genome-editing toolbox by utilizing Cas12a from Lachnospiraceae bacterium ND2006 (LbCas12a) in combination with the homology-directed repair machinery endogenously present in M. maripaludis. This toolbox can delete target genes with a success rate of up to 95%, despite the hyperpolyploidy of M. maripaludis. For the purpose of demonstrating a large deletion, the M. maripaludis flagellum operon (∼8.9 kbp) was replaced by the Escherichia coli β-glucuronidase gene. To facilitate metabolic engineering and flux balancing in M. maripaludis, the relative strength of 15 different promoters was quantified in the presence of two common growth substrates, either formate or carbon dioxide and hydrogen. This CRISPR/LbCas12a toolbox can be regarded as a reliable and quick method for genome editing in a methanogen.
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Switching the activity of Cas12a using guide RNA strand displacement circuits.
The CRISPR effector protein Cas12a has been used for a wide variety of applications such as in vivo gene editing and regulation or in vitro DNA sensing. Here, we add programmability to Cas12a-based DNA processing by combining it with strand displacement-based reaction circuits. We first establish a viable strategy for augmenting Cas12a guide RNAs (gRNAs) at their 5' end and then use such 5' extensions to construct strand displacement gRNAs (SD gRNAs) that can be activated by single-stranded RNA trigger molecules. These SD gRNAs are further engineered to exhibit a digital and orthogonal response to different trigger RNA inputs-including full length mRNAs-and to function as multi-input logic gates. We also demonstrate that SD gRNAs can be designed to work inside bacterial cells. Using such in vivo SD gRNAs and a DNase inactive version of Cas12a (dCas12a), we demonstrate logic gated transcriptional control of gene expression in E. coli.
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When Less Is More: Targeting the Myostatin Gene in Livestock for Augmenting Meat Production.
How to increase meat production is one of the main questions in animal breeding. Selection for improved body weight has been made and, due to recent genomic advances, naturally occurring variants that are responsible for controlling economically relevant phenotypes have been revealed. The myostatin (MSTN) gene, a superstar gene in animal breeding, was discovered as a negative controller of muscle mass. In some livestock species, natural mutations in the MSTN gene could generate the agriculturally desirable double-muscling phenotype. However, some other livestock species or breeds lack these desirable variants. Genetic modification, particularly gene editing, offers an unprecedented opportunity to induce or mimic naturally occurring mutations in livestock genomes. To date, various MSTN-edited livestock species have been generated using different gene modification tools. These MSTN gene-edited models have higher growth rates and increased muscle mass, suggesting the high potential of utilizing MSTN gene editing in animal breeding. Additionally, post-editing investigations in most livestock species support the favorable influence of targeting the MSTN gene on meat quantity and quality. In this Review, we provide a collective discussion on targeting the MSTN gene in livestock to further encourage its utilization opportunities. It is expected that, shortly, MSTN gene-edited livestock will be commercialized, and MSTN-edited meat will be on the tables of ordinary customers.
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Integration, abundance, and transmission of mutations and transgenes in a series of CRISPR/Cas9 soybean lines.
BACKGROUND: As with many plant species, current genome editing strategies in soybean are initiated by stably transforming a gene that encodes an engineered nuclease into the genome. Expression of the transgene results in a double-stranded break and repair at the targeted locus, oftentimes resulting in mutation(s) at the intended site. As soybean is a self-pollinating species with 20 chromosome pairs, the transgene(s) in the T0 plant are generally expected to be unlinked to the targeted mutation(s), and the transgene(s)/mutation(s) should independently assort into the T1 generation, resulting in Mendellian combinations of transgene presence/absence and allelic states within the segregating family. This prediction, however, is not always consistent with observed results. RESULTS: In this study, we investigated inheritance patterns among three different CRISPR/Cas9 transgenes and their respective induced mutations in segregating soybean families. Next-generation resequencing of four T0 plants and four T1 progeny plants, followed by broader assessments of the segregating families, revealed both expected and unexpected patterns of inheritance among the different lineages. These unexpected patterns included: (1) A family in which T0 transgenes and mutations were not transmitted to progeny; (2) A family with four unlinked transgene insertions, including two respectively located at paralogous CRISPR target break sites; (3) A family in which mutations were observed and transmitted, but without evidence of transgene integration nor transmission. CONCLUSIONS: Genome resequencing provides high-resolution of transgene integration structures and gene editing events. Segregation patterns of these events can be complicated by several potential mechanisms. This includes, but is not limited to, plant chimeras, multiple unlinked transgene integrations, editing of intended and paralogous targets, linkage between the transgene integration and target site, and transient expression of the editing reagents without transgene integration into the host genome.
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DGAT2 stability is increased in response to DGAT1 inhibition in gene edited HepG2 cells.
In eukaryotic organisms, two unrelated acyl-CoA:diacylglycerol acyltransferase (DGAT) enzymes, DGAT1 and DGAT2, catalyze the final step of the triacylglycerol biosynthetic pathway. Both enzymes are highly expressed in lipogenic tissues, such as adipose tissue, small intestine and the liver. DGAT2 has a prominent role in hepatocyte lipid metabolism synthesizing triacylglycerols that are utilized for very low-density lipoprotein assembly. However, due to the lack of useful antibodies to detect endogenous DGAT2 protein, it has been difficult to determine how this enzyme functions at the cellular level. We have unsuccessfully tested many commercial antibodies as well as our own "in-house" antibodies. There is currently no evidence that DGAT2 undergoes processing such that antigenic epitopes to these antibodies are removed. As an alternative, many studies have utilized epitope tagged versions of DGAT2 overexpressed in cells. These approaches can provide valuable information about a protein, but can be subject to artifacts, such as mislocalization, misregulation, protein aggregation and abnormal protein-protein interactions. In this study, we used gene editing with CRISPR/Cas9 to add three consecutive FLAG epitopes to the C-terminus of endogenous DGAT2 in HepG2 cells. HepG2 cells, derived from a human hepatocellular carcinoma, have been routinely used as a cell model to study human hepatocyte lipid and lipoprotein metabolism. Using this system allowed us to successfully detect DGAT2 expressed from its endogenous locus in HepG2 cells by immunoblotting with anti-FLAG antibodies.
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CRISPR-Cas: a game-changer in vaccine development and the fight against viral infections.
Understanding the mechanisms of virus-host interactions is crucial for advancing our knowledge of viral pathogenesis and developing effective antiviral strategies. As novel, highly virulent pathogens continue to emerge, overcoming the challenges in virus-host biology and infection control is more important than ever. The CRISPR-Cas system has revolutionized genetic engineering, offering a precise and efficient application in synthetic biology. It is also invaluable in virology research, diagnostics, and antiviral therapeutics. In this review, we explore the latest advances in CRISPR-Cas-based approaches to elucidate virus-host interactions, focusing on the identification of key receptors and host factors essential for virus infection. Furthermore, we discuss the potential of the CRISPR-Cas system as an antiviral strategy for tackling newly emerging and re-emerging viruses. Finally, we discuss the potential of CRISPR-Cas-based gene editing for the development of innovative viral vaccines.
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Recent advances in stimuli-responsive polymeric carriers for controllable CRISPR/Cas9 gene editing system delivery.
Non-viral polymeric vectors with good biocompatibility have been recently explored as delivery systems for clustered regularly interspaced short palindromic repeat (CRISPR)-associated (Cas) nucleases. In this review, based on current limitations and critical barriers, we summarize the advantages of stimulus-responsive polymeric delivery vectors (i.e., pH, redox, or enzymes) towards controllable CRISPR/Cas9 genome editing system delivery as well as the advances in using stimulus-responsive CRISPR/Cas9 polymeric carriers towards cancer treatment. Last but not least, the key challenges and promising development strategies of stimulus-responsive polymeric vector designs for CRISPR/Cas9 systems will also be discussed.
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Efficient CRISPR/Cas9-mediated editing of trinucleotide repeat expansion in myotonic dystrophy patient-derived iPS and myogenic cells.
CRISPR/Cas9 is an attractive platform to potentially correct dominant genetic diseases by gene editing with unprecedented precision. In the current proof-of-principle study, we explored the use of CRISPR/Cas9 for gene-editing in myotonic dystrophy type-1 (DM1), an autosomal-dominant muscle disorder, by excising the CTG-repeat expansion in the 3'-untranslated-region (UTR) of the human myotonic dystrophy protein kinase (DMPK) gene in DM1 patient-specific induced pluripotent stem cells (DM1-iPSC), DM1-iPSC-derived myogenic cells and DM1 patient-specific myoblasts. To eliminate the pathogenic gain-of-function mutant DMPK transcript, we designed a dual guide RNA based strategy that excises the CTG-repeat expansion with high efficiency, as confirmed by Southern blot and single molecule real-time (SMRT) sequencing. Correction efficiencies up to 90% could be attained in DM1-iPSC as confirmed at the clonal level, following ribonucleoprotein (RNP) transfection of CRISPR/Cas9 components without the need for selective enrichment. Expanded CTG repeat excision resulted in the disappearance of ribonuclear foci, a quintessential cellular phenotype of DM1, in the corrected DM1-iPSC, DM1-iPSC-derived myogenic cells and DM1 myoblasts. Consequently, the normal intracellular localization of the muscleblind-like splicing regulator 1 (MBNL1) was restored, resulting in the normalization of splicing pattern of SERCA1. This study validates the use of CRISPR/Cas9 for gene editing of repeat expansions.
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CRISPR-Cas9 genome engineering: Treating inherited retinal degeneration.
Gene correction is a valuable strategy for treating inherited retinal degenerative diseases, a major cause of irreversible blindness worldwide. Single gene defects cause the majority of these retinal dystrophies. Gene augmentation holds great promise if delivered early in the course of the disease, however, many patients carry mutations in genes too large to be packaged into adeno-associated viral vectors and some, when overexpressed via heterologous promoters, induce retinal toxicity. In addition to the aforementioned challenges, some patients have sustained significant photoreceptor cell loss at the time of diagnosis, rendering gene replacement therapy insufficient to treat the disease. These patients will require cell replacement to restore useful vision. Fortunately, the advent of induced pluripotent stem cell and CRISPR-Cas9 gene editing technologies affords researchers and clinicians a powerful means by which to develop strategies to treat patients with inherited retinal dystrophies. In this review we will discuss the current developments in CRISPR-Cas9 gene editing in vivo in animal models and in vitro in patient-derived cells to study and treat inherited retinal degenerative diseases.
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CRISPR/Cas9 Gene Editing: A Novel Approach Towards Alzheimer's Disease Treatment.
In defiance of the vast amount of information regarding Alzheimer's disease (AD) that has been learned over the past thirty years, progress toward developing an effective therapy has been difficult. A neurological ailment that progresses and cannot be reversed is Alzheimer's disease, which shows neurofibrillary tangles, beta-amyloid plaque, and a lack of cognitive processes that is created by tau protein clumps with hyperphosphorylation that finally advances to neuronal damage without a recognized treatment, which has stimulated research into new therapeutic strategies. The protein CAS9 is linked to CRISPR, which is a clustered Regularly Interspaced Short Palindromic Repeat that inactivates or corrects a gene by recognizing a gene sequence that produces a doublestranded break has enchanted a whole amount of interest towards its potency to cure gene sequences in AD. The novel CRISPR-Cas9 applications for developing in vitro and in vivo models to the benefit of AD investigation and therapies are thoroughly analyzed in this work. The discussion will also touch on the creation of delivery methods, which is a significant obstacle to the therapeutic use of CRISPR/Cas9 technology. By concentrating on specific genes, such as those that are significant early- onset AD risk factors and late-onset AD risk factors, like the apolipoprotein E4 (APOE4) gene, this study aims to evaluate the potential application of CRISPR/Cas9 as a possible treatment for AD.
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