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Single nucleotide substitutions effectively block Cas9 and allow for scarless genome editing in Caenorhabditis elegans.
In Caenorhabditis elegans, germline injection of Cas9 complexes is reliably used to achieve genome editing through homology-directed repair of Cas9-generated DNA breaks. To prevent Cas9 from targeting repaired DNA, additional blocking mutations are often incorporated into homologous repair templates. Cas9 can be blocked either by mutating the PAM sequence that is essential for Cas9 activity or by mutating the guide sequence that targets Cas9 to a specific genomic location. However, it is unclear how many nucleotides within the guide sequence should be mutated, since Cas9 can recognize "off-target" sequences that are imperfectly paired to its guide. In this study, we examined whether single-nucleotide substitutions within the guide sequence are sufficient to block Cas9 and allow for efficient genome editing. We show that a single mismatch within the guide sequence effectively blocks Cas9 and allows for recovery of edited animals. Surprisingly, we found that a low rate of edited animals can be recovered without introducing any blocking mutations, suggesting a temporal block to Cas9 activity in C. elegans. Furthermore, we show that the maternal genome of hermaphrodite animals is preferentially edited over the paternal genome. We demonstrate that maternally provided haplotypes can be selected using balancer chromosomes and propose a method of mutant isolation that greatly reduces screening efforts postinjection. Collectively, our findings expand the repertoire of genome editing strategies in C. elegans and demonstrate that extraneous blocking mutations are not required to recover edited animals when the desired mutation is located within the guide sequence.
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Comprehensive analysis and accurate quantification of unintended large gene modifications induced by CRISPR-Cas9 gene editing.
Most genome editing analyses to date are based on quantifying small insertions and deletions. Here, we show that CRISPR-Cas9 genome editing can induce large gene modifications, such as deletions, insertions, and complex local rearrangements in different primary cells and cell lines. We analyzed large deletion events in hematopoietic stem and progenitor cells (HSPCs) using different methods, including clonal genotyping, droplet digital polymerase chain reaction, single-molecule real-time sequencing with unique molecular identifier, and long-amplicon sequencing assay. Our results show that large deletions of up to several thousand bases occur with high frequencies at the Cas9 on-target cut sites on the HBB (11.7 to 35.4%), HBG (14.3%), and BCL11A (13.2%) genes in HSPCs and the PD-1 (15.2%) gene in T cells. Our findings have important implications to advancing genome editing technologies for treating human diseases, because unintended large gene modifications may persist, thus altering the biological functions and reducing the available therapeutic alleles.
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An overview of applications of CRISPR-Cas technologies in biomedical engineering.
Clustered Regulatory Interspaced Short Palindromic Repeats (CRISPR) is one of the major genome editing systems and allows changing DNA levels of an organism. Among several CRISPR categories, the CRISPR-Cas9 system has shown a remarkable progression rate over its lifetime. Recently, other tools including CRISPR-Cas12 and CRISPR-Cas13 have been introduced. CRISPR-Cas9 system has played a key role in the industrial cell factory's production and improved our understanding of genome function. Additionally, this system has been used as one of the major genome editing systems for the diagnosis and treatment of several infectious and non-infectious diseases. In this review, we discuss CRISPR biology, its versatility, and its application in biomedical engineering.
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CRISPR/Cas9-Mediated Genome Editing to Generate Clonal iPSC Lines.
The ability to reprogram somatic cells into induced pluripotent stem cells (iPSCs) was developed in 2006 and represented a major breakthrough in stem cell research. A more recent milestone in biomedical research was reached in 2013 when the CRISPR/Cas9 system was used to edit the genome of mammalian cells. The coupling of both human (h)iPSCs and CRISPR/Cas9 technology offers great promise for cell therapy and regenerative medicine. However, several limitations including time and labor consumption, efficiency and efficacy of the system, and the potential off-targets effects induced by the Cas9 nuclease still need to be addressed. Here, we describe a detailed method for easily engineering genetic changes in hiPSCs, using a nucleofection-mediated protocol to deliver the CRISPR/Cas9 components into the cells, and discuss key points to be considered when designing your experiment. The clonal, genome-edited hiPSC line generated via our method can be directly used for downstream applications.
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CRISPRa-mediated transcriptional activation of the SlPR-1 gene in edited tomato plants.
With the continuous deterioration of arable land due to an ever-growing population, improvement of crops and crop protection have a fundamental role in maintaining and increasing crop productivity. Alternatives to the use of pesticides encompass the use of biological control agents, generation of new resistant crop cultivars, the application of plant activator agrochemicals to enhance plant defenses, and the use of gene editing techniques, like the CRISPR-Cas system. Here, we test the hypothesis that epigenome editing, via CRISPR activation (CRISPRa), activate tomato plant defense genes to confer resistance against pathogen attack. We provide evidence that edited tomato plants for the PATHOGENESIS-RELATED GENE 1 gene (SlPR-1) show enhanced disease resistance to Clavibacter michiganensis subsp. michiganensis infection. Resistance was assessed by evaluating disease progression and symptom appearance, pathogen accumulation, and changes in SlPR-1 gene expression at different time points. We determined that CRISPRa-edited plants develop enhanced disease-resistant to the pathogen without altering their agronomic characteristics and, above all, preventing the advancement of disease symptoms, stem canker, and plant death.
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A versatile tool for gene editing in the diatom Thalassiosira pseudonana.
Diatoms are major contributors to marine primary production, with the centric diatom Thalassiosira pseudonana a powerful model for understanding biogeochemical processes including carbon fixation and silicification. While there are molecular tools available for fluorescent protein (FP) tagging and CRISPR/Cas9 genome editing in T. pseudonana, these require the delivery of multiple vectors or have limited versatility. Additionally, scarless endogenous tagging, that results in a FP fusion expressed from its native genomic location, has yet to be developed. Here, we describe a versatile modular Golden Gate-based toolkit for T. pseudonana that through the delivery of a single-episome via bacterial conjugation enables: (i) FP tagging, (ii) dual FP tagging, (iii) CRISPR/Cas9 genome editing, (iv) simultaneous FP tagging with gene editing, and (v) scarless endogenous FP tagging. We further expand the available parts for T. pseudonana by validating 3 additional FPs and 2 untested promoter/terminator pairs. We demonstrate the versatility of our system by knocking out diatom pyrenoid component 1 (DPC1) while simultaneously GFP tagging the rbcS (Rubisco small subunit), and by endogenously GFP tagging the bestrophin-like protein BST2. While DPC1 knockout does not result in a major pyrenoid structural defect due to unperturbed rbcS-GFP localization to the pyrenoid, we confirm that BST2 localizes to the pyrenoid and exhibits increased fluorescence under low CO2-supporting a role in diatom carbon fixation. Our developed genetic tools provide a robust framework for exploring cellular processes in diatoms, accelerating routine studies and enabling systematic, quantitative and large-scale studies.
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Efficient genome editing of two-cell mouse embryos via modified CRISPR/Cas electroporation.
Creating genetically modified (GM) animals using CRISPR/Cas mediated through the electroporation of two-cell stage embryos, rather than fertilized eggs, holds considerable potential. The full potential of genome editing using two-cell stage embryos is only beginning to be explored. We developed an improved electroporation method to prevent blastomere fusion in two-cell-stage embryos, enabling efficient genome editing. Using this method, we demonstrated that the indel mutation rates and ssODN knock-in (KI) efficiencies in two-cell-stage embryos are comparable to those in fertilized eggs, with a tendency for higher efficiency in long DNA KI. This study highlights the potential value of two-cell-stage embryos and provides enhanced animal model production opportunities. Furthermore, realizing genome editing in two-cell-stage embryos extends the editing timeframe from fertilized egg to two-cell-stage embryo, offering promising avenues for future research in embryo genome editing techniques.
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Research Techniques Made Simple: The Application of CRISPR-Cas9 and Genome Editing in Investigative Dermatology.
Designer nucleases have gained widespread attention for their ability to precisely modify genomic DNA in a programmable manner. These genome-editing nucleases make double-stranded breaks at specified loci, and desired changes can be made to modify, ablate, or excise target genes. This technology has been used widely to develop human disease models in laboratory animals and to study gene functions by silencing, activating, or modifying them. Furthermore, the recent discovery of a bacterially derived programmable nuclease termed clustered regularly interspaced palindromic repeats (CRISPR)-associated protein 9 (Cas9) has revolutionized the field because of its versatility and wide applicability. In this article, we discuss various modalities used to achieve genome editing with an emphasis on CRISPR-Cas9. We discuss genome-editing strategies to either repair or ablate target genes, with emphasis on their applications for investigating dermatological diseases. Additionally, we highlight preclinical studies showing the potential of genome editing as a therapy for congenital blistering diseases and as an antimicrobial agent, and we discuss limitations and future directions of this technology.
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Understanding genetic heterogeneity in gene-edited hematopoietic stem cell products.
CRISPR/Cas gene editing has transformed genetic research and is poised to drive the next generation of gene therapies targeting hematopoietic stem cells (HSCs). However, the installation of the "desired" edit is most often only achieved in a minor subset of alleles. The array of cellular pathways triggered by gene editing tools produces a broad spectrum of "undesired" editing outcomes, including short insertions and deletions (indels) and chromosome rearrangements, leading to considerable genetic heterogeneity in gene-edited HSC populations. This heterogeneity may undermine the effect of the genetic intervention since only a subset of cells will carry the intended modification. Also, undesired mutations represent a potential safety concern as gene editing advances toward broader clinical use. Here, we will review the different sources of "undesired" edits and will discuss strategies for their mitigation and control.
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CRISPR-Cas9 mediated genome editing of Kluyveromyces marxianus for iterative, multiplexed gene disruption and pathway integration.
Kluyveromyces marxianus, a thermotolerant, fast-growing, Crabtree-negative yeast, is a promising chassis for the manufacture of various bioproducts. Although several genome editing tools are available for this yeast, these tools still require refinement to enable more convenient and efficient genetic modification. In this study, we engineered the K. marxianus NBRC 104275 strain by impairing the nonhomologous end joining and enhancing the homologous recombination machinery, which resulted in improved homology-directed repair effective on homology arms of up to 40 bp in length. Additionally, we simplified the CRISPR-Cas9 editing system by constructing a strain for integrative expression of Cas9 nuclease and plasmids bearing different selection markers for gRNA expression, thereby facilitating iterative genome editing without the need for plasmid curing. We demonstrated that tRNA was more effective than the hammerhead ribozyme for processing gRNA primary transcripts, and readily assembled tRNA-gRNA arrays were used for multiplexed editing of at least four targets. This editing tool was further employed for simultaneous scarless in vivo assembly of a 12-kb cassette from three fragments and marker-free integration for expressing a fusion variant of fatty acid synthase, as well as the integration of genes for starch hydrolysis. Together, the genome editing tool developed in this study makes K. marxianus more amenable to genetic modification and will facilitate more extensive engineering of this nonconventional yeast for chemical production.
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