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Topology-Engineered Guide RNAs for Programmable Control of CRISPR/Cas Activity.
CRISPR/Cas systems have transformed genome editing, yet achieving precise temporal and conditional control remains challenging. Traditional strategies involving linear guide RNAs (gRNAs) modified with multiple chemical groups throughout their strands often face limitations such as heterogeneous reaction outcomes, irreversibility, and variable editing efficiencies. To overcome these issues, topology-engineered guide RNAs (TE-gRNAs) have emerged, featuring defined structural architectures including polymeric, circular, and dendrimer-like topologies that enable precise spatial control, reversibility, and programmable activation of CRISPR activity. By selectively incorporating physical or chemically responsive linkers and stimuli-sensitive groups at specific sites, TE-gRNAs facilitate dynamic and conditional genome editing that can be activated or deactivated with external triggers such as light or chemical signals. These engineered RNA structures significantly improve synthesis feasibility, stability, reduce off-target effects, and provide unprecedented control over gene editing processes. Recent advancements in TE-gRNAs demonstrate their broad applicability in synthetic biology, functional genomics, and therapeutic interventions, highlighting their potential to achieve precise spatiotemporal modulation of CRISPR systems. This review summarizes the current strategies, benefits, and challenges associated with TE-gRNAs, and discusses future directions for enhancing their performance and utility in complex genome editing applications.
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Genome Editing in Ferret Airway Epithelia Mediated by CRISPR/Nucleases Delivered with Amphiphilic Shuttle Peptides.
Gene editing strategies are attractive for treating genetic pulmonary diseases such as cystic fibrosis (CF). However, challenges have included the development of safe and effective vector systems for gene editing of airway epithelia and model systems to report their efficiency and durability. The domestic ferret (Mustela putorius furo) has a high degree of conservation in lung cellular anatomy with humans, and has served as an excellent model for many types of lung diseases, including CF. In this study, we evaluated the efficiency of amphiphilic shuttle peptide S10 for protein delivery and gene editing using SpCas9, and AsCas12a (Cpf1) ribonucleoproteins (RNPs). These approaches were evaluated in proliferating ferret airway basal cells, polarized airway epithelia in vitro, and lungs in vivo, by accessing the editing efficiency using reporter ferrets and measuring indels at the ferret CFTR locus. Our results demonstrate that shuttle peptides efficiently enable delivery of reporter proteins/peptides and gene editing SpCas9 or Cpf1 RNP complexes to ferret airway epithelial cells in vitro and in vivo. We measured S10 delivery efficiency of green fluorescent protein (GFP)-nuclear localization signal (NLS) protein or SpCas9 RNP into ferret airway basal cells and fully differentiated ciliated and nonciliated epithelial cells in vitro. In vitro and in vivo gene editing efficiencies were determined by Cas/LoxP-gRNA RNP-mediated conversion of a ROSA-TG Cre recombinase reporter using transgenic primary cells and ferrets. S10/Cas9 RNP was more effective, relative to S10/Cpf1 RNP at gene editing of the ROSA-TG locus. Intratracheal lung delivery of the S10 shuttle combined with GFP-NLS protein or D-Retro-Inverso (DRI)-NLS peptide demonstrated efficiencies of protein delivery that were ∼3-fold or 14-fold greater, respectively, than the efficiency of gene editing at the ROSA-TG locus using S10/Cas9/LoxP-gRNA. Cpf1 RNPs was less effective than SpCas9 at gene editing of LoxP locus. These data demonstrate the feasibility of shuttle peptide delivery of Cas RNPs to the ferret airways and the potential utility for developing ex vivo stem cell-based and in vivo gene editing therapies for genetic pulmonary diseases such as CF.
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RNA lipid nanoparticles as efficient in vivo CRISPR-Cas9 gene editing tool for therapeutic target validation in glioblastoma cancer stem cells.
In vitro and ex-vivo target identification strategies often fail to predict in vivo efficacy, particularly for glioblastoma (GBM), a highly heterogenous tumor rich in resistant cancer stem cells (GSCs). An in vivo screening tool can improve prediction of therapeutic efficacy by considering the complex tumor microenvironment and the dynamic plasticity of GSCs driving therapy resistance and recurrence. This study proposes lipid nanoparticles (LNPs) as an efficient in vivo CRISPR-Cas9 gene editing tool for target validation in mesenchymal GSCs. LNPs co-delivering mRNA (mCas9) and single-guide RNA (sgRNA) were successfully formulated and optimized facilitating both in vitro and in vivo gene editing. In vitro, LNPs achieved up to 67 % reduction in green fluorescent protein (GFP) expression, used as a model target, outperforming a commercial transfection reagent. Intratumoral administration of LNPs in GSCs resulted in ∼80 % GFP gene knock-out and a 2-fold reduction in GFP signal by day 14. This study showcases the applicability of CRISPR-Cas9 LNPs as a potential in vivo screening tool in GSCs, currently lacking effective treatment. By replacing GFP with a pool of potential targets, the proposed platform presents an exciting prospect for therapeutic target validation in orthotopic GSCs, bridging the gap between preclinical and clinical research.
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Reduced Susceptibility to Phytophthora in Non-Transgenic Cacao Progeny Through CRISPR-Cas9 Mediated TcNPR3 Mutagenesis.
Black pod disease, caused by a complex of Phytophthora species, poses a severe threat to global cacao production. This study explores the use of CRISPR-Cas9 genome editing to reduce disease susceptibility in Theobroma cacao L. by targeting the TcNPR3 gene, a known negative regulator of plant defence. Transgenic T0 lines carrying mutations predicted to disrupt TcNPR3 function exhibited reduced susceptibility to Phytophthora infection in in vitro foliar assays. These T0 plants were advanced to maturity and outcrossed with non-transgenic cacao to eliminate T-DNA sequences associated with the CRISPR-Cas9 transgene. Whole-genome sequencing of the T0 parents and 22 progeny revealed single T-DNA insertion sites in each T0 line; seven progeny retained the edited npr3 alleles but lacked T-DNA insertions. Transcriptome analysis of the mutant lines showed upregulation of genes associated with reactive oxygen species (ROS) generation, defence-related transcription factors and pathogenesis-related proteins. Several genes were also downregulated, suggesting that TcNPR3 may function as both a repressor and an activator in regulating basal transcriptional states. Genome-edited plants were phenotypically comparable to wild-type controls and displayed a 42% reduction in lesion size upon Phytophthora challenge. These findings demonstrate the feasibility of generating non-transgenic cacao with reduced susceptibility to Phytophthora through CRISPR-Cas9-mediated genome editing, offering a promising strategy for sustainable cacao cultivation and improved farmer livelihoods. Field trials are underway to evaluate long-term agronomic performance under natural conditions.
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Development of a marker-free mutagenesis system using CRISPR-Cas9 in the pathogenic mould Aspergillus fumigatus.
Aspergillus fumigatus is a saprophytic fungal pathogen that is the cause of more than 300,000 life-threatening infections annually. Our understanding of pathogenesis and factors contributing to disease progression are limited. Development of rapid and versatile gene editing methodologies for A. fumigatus is essential. CRISPR-Cas9 mediated transformation has been widely used as a novel genome editing tool and has been used for a variety of editing techniques, such as protein tagging, gene deletions and site-directed mutagenesis in A. fumigatus. However, successful genome editing relies on time consuming, multi-step cloning procedures paired with the use of selection markers, which can result in a metabolic burden for the host and/or unintended transcriptional modifications at the site of integration. We have used an in vitro CRISPR-Cas9 assembly methodology to perform selection-free genome editing, including epitope tagging of proteins and site-directed mutagenesis. The repair template used during this transformation use 50 bp micro-homology arms and can be generated with a single PCR reaction or by purchasing synthesised single stranded oligonucleotides, decreasing the time required for complex construct synthesis.
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Amount of Cas9 protein introduced into mouse embryos via electroporation affects the genome-editing rate.
Genetically engineered animals can be produced quickly using genome editing technology. A new electroporation technique, technique for animal knockout system by electroporation (TAKE), aids in the production of genome-edited animals by introducing nucleases into intact embryos using electroporation instead of microinjection. It is difficult to confirm nuclease delivery into embryos after electroporation using the conventional TAKE method. We previously reported the successful visualization of fluorescently-labeled tracrRNA in embryos after electroporation Cas9 paired with the crRNA:tracrRNA-ATTO550 duplex. However, the amount of fluorescence signal from labeled tracrRNA in embryos did not correlate with the genome editing rate of the offspring. This study examined the visualization of Cas9 protein in embryos after electroporation and its correlation with the genome editing rate of the offspring using a fluorescent Cas9 fusion protein. The fluorescent Cas9 protein was observed in all embryos that survived following electroporation. We found that the efficiency of Cas9 protein delivery into embryos via electroporation depended on the pulse length. Furthermore, we demonstrated that the amount of fluorescent Cas9 protein detected in the embryos correlated with the genome editing efficiency of the embryos. These data indicate that the TAKE method using fluorescently-labeled nucleases can be used to optimize the delivery conditions and verify nuclease delivery into individual embryos prior to embryo transfer for the efficient production of genome-edited animals.
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Empowering chimeric antigen receptor T-cell therapy with CRISPR.
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Epigenetic editing of marine medaka ( Oryzias melastigma) fgf2 using CRISPR/dCas9-Tet1CD.
CRISPR/dCas9-mediated epigenetic editing offers a versatile approach for transcriptional regulation without introducing DNA strand breaks. Although this strategy has been explored in a limited number of species, its application in aquatic vertebrates remains largely uncharacterized. In this study, ten-eleven translocation methylcytosine dioxygenase 1 ( tet1) was cloned and molecularly characterized in marine medaka ( Oryzias melastigma). Decitabine treatment identified fibroblast growth factor 2 ( fgf2) as a methylation-sensitive gene, with a regulatory CpG island located within its promoter region. Subsequently, a CRISPR/dCas9-Tet1CD activation system was constructed by fusing the catalytic domain of Tet1 (Tet1CD, Ala1352-Thr2034) to dCas9, enabling locus-specific DNA demethylation. Targeting fgf2, this CRISPR/dCas9-Tet1CD system induced efficient and selective demethylation of the CpG island, resulting in a maximal 2.41-fold increase in fgf2 transcript levels. Whole-genome bisulfite sequencing and transcriptomic analysis confirmed high on-target precision with minimal off-target effects. Epigenetic activation of fgf2 further modulated downstream gene networks associated with growth, promoting durable transcriptional enhancement and increased cellular proliferation. Collectively, these results establish a robust and highly specific epigenetic editing platform in marine medaka, providing a powerful tool for functional genomics studies and regulatory element analysis in aquatic models.
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CD46 Gene Editing Confers Ex Vivo BVDV Resistance in Fibroblasts from Cloned Angus Calves.
A previous study demonstrated that a 19-nucleotide edit, encoding a six amino acid substitution in the bovine CD46 gene, dramatically reduced bovine viral diarrhea virus (BVDV) susceptibility in a cloned Gir (Bos indicus) heifer. The present study aimed to replicate this result in American Angus (Bos taurus) using genetically matched controls and larger sample sizes. CRISPR/Cas9-mediated homology-directed repair introduced the identical CD46 edit, encoding the A82LPTFS amino acid sequence, into exon 2 of CD46 in primary Angus fibroblasts. Thirty-three cloned embryos (22 CD46-edited and 11 unedited) were transferred to recipient cows. However, all pregnancies resulted in pre- and perinatal losses due to cloning-related abnormalities, preventing in vivo BVDV challenge. Consequently, ex vivo BVDV susceptibility assays were performed on primary fibroblast cell lines rescued from deceased cloned Angus calves. Infection studies revealed significantly reduced susceptibility in the edited lines, comparable to the resistance previously observed from the edited Gir heifer. These studies extend the applicability of this finding from Gir to the most common US beef breed, Angus, suggesting the potential for broad application of CD46 editing in BVDV control. Continued advancements in cloning technology will enhance the potential of gene-editing for producing disease-resistant livestock.
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Plant prime editing goes prime.
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