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Revolutionizing cancer treatment: enhancing CAR-T cell therapy with CRISPR/Cas9 gene editing technology.
CAR-T cell therapy, a novel immunotherapy, has made significant breakthroughs in clinical practice, particularly in treating B-cell-associated leukemia and lymphoma. However, it still faces challenges such as poor persistence, limited proliferation capacity, high manufacturing costs, and suboptimal efficacy. CRISPR/Cas system, an efficient and simple method for precise gene editing, offers new possibilities for optimizing CAR-T cells. It can increase the function of CAR-T cells and reduce manufacturing costs. The combination of CRISPR/Cas9 technology and CAR-T cell therapy may promote the development of this therapy and provide more effective and personalized treatment for cancer patients. Meanwhile, the safety issues surrounding the application of this technology in CAR-T cells require further research and evaluation. Future research should focus on improving the accuracy and safety of CRISPR/Cas9 technology to facilitate the better development and application of CAR-T cell therapy. This review focuses on the application of CRISPR/Cas9 technology in CAR-T cell therapy, including eliminating the inhibitory effect of immune checkpoints, enhancing the ability of CAR-T cells to resist exhaustion, assisting in the construction of universal CAR-T cells, reducing the manufacturing costs of CAR-T cells, and the security problems faced. The objective is to show the revolutionary role of CRISPR/Cas9 technology in CAR-T cell therapy for researchers.
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Commit to talks on patient data and public health.
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Jennifer Doudna: Tailoring the Genome.
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CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models.
The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the efficiency in developing gene-knockout mice by inducing small indel mutation would be sufficient enough, the efficiency of embryo genome editing for making large-size DNA knock-in (KI) is still low. Therefore, in contrast to the direct KI method in embryos, gene targeting using embryonic stem cells (ESCs) followed by embryo injection to develop chimera mice still has several advantages (e.g., high throughput targeting in vitro, multi-allele manipulation, and Cre and flox gene manipulation can be carried out in a short period). In addition, strains with difficult-to-handle embryos in vitro, such as BALB/c, can also be used for ESC targeting. This protocol describes the optimized method for large-size DNA (several kb) KI in ESCs by applying CRISPR/Cas9-mediated genome editing followed by chimera mice production to develop gene-manipulated mouse models.
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Efficient gene editing of BMP15, GDF9, and MSTN-but not the imprinted CLPG gene-in goat embryos via electrotransfection and handmade cloning.
CRISPR/Cas9 technology represents a powerful tool for advancing livestock breeding by enabling precise, on-target gene edits without the genomic mixing associated with traditional introgression methods. In this study, we employed a dual gRNA-based CRISPR/Cas9 strategy to induce targeted deletions and indel mutations in both reproductive and growth-related genes. These included the metacentric genes bone morphogenetic protein 15 (BMP15) and growth differentiation factor 9 (GDF9), which are associated with increased ovulation rate and litter size, as well as the telomeric genes myostatin (MSTN) and callipyge (CLPG), which are linked to muscle development and enhanced meat production. We employed an optimized electrotransfection protocol consisting of 10-20 µg of each plasmid DNA, 250 µL OptiMEM-GlutaMAX, and one million goat fibroblast cells. The electroporation was performed using a Bio-Rad system in a 4-mm cuvette, with two 10-millisecond pulses at 270 volts, separated by a 10-second interval. This protocol enabled efficient genome editing of goat embryonic fibroblast cells, which were subsequently used to generate cloned embryos via handmade somatic cell nuclear transfer (SCNT), involving manual enucleation and cell-oocyte fusion steps. Sequencing revealed high mutation rates (78-97%) and a predominance of biallelic edits in BMP15, GDF9, and MSTN. Notably, MSTN gRNAs with a 7-bp overlapping sequence at their 3' ends showed a high editing efficiency. In contrast, the imprinted CLPG gene exhibited a significantly lower mutation rate (~ 30%), likely due to epigenetic constraints. While overall mutation rates did not differ significantly between metacentric and telomeric genes, on-target deletions were more frequent in metacentric genes (43%) than in telomeric ones (20%). Embryo development rates from gene-edited cells were comparable to those from non-edited controls. These findings underscore the utility of combining electrotransfection with SCNT for efficient editing of non-imprinted genes and highlight the need for improved strategies to overcome barriers in editing imprinted loci.
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Improvement of Genome Editing Efficiency in Mammalian Cells by a Refined Uni-Vector Prime Editing System.
Prime editing is an advanced technology in the CRISPR/Cas genome editing field. The original multi-vector method utilized in prime editing has less efficiency and precision and is also inherently difficult in generating homozygous mutations in mammalian cells. To overcome these technical difficulties, a Uni-vector prime editing system containing the major components for prime editing has been constructed in two all-in-one plasmids, pPE3-pPuro and pePE5max-pPuro. The Uni-vector prime editing plasmids enhance the editing efficiency of prime editing and improve the generation of homozygous mutated mammalian cell lines. The editing efficiency depends on the transfection efficiency. Remarkably, the Uni-vector ePE5max system can achieve a high editing rate even in cell lines that are traditionally difficult to transfect. This Uni-vector approach simplifies the delivery of all the necessary editing components and greatly improves the editing efficiency. This approach represents an advanced progress in the field of prime editing.
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Cell Synchronization Enhances Nuclear Transformation and Genome Editing via Cas9 Enabling Homologous Recombination in Chlamydomonas reinhardtii.
In Chlamydomonas reinhardtii, the model organism for eukaryotic green algae and plants, the processes of nuclear transformation and genome editing in particular are still marked by a low level of efficiency, and so intensive work is required in order to create and identify mutants for the investigation of basic physiological processes, as well as the implementation of biotechnological applications. In this work, we show that cell synchronization during the stages of the cell cycle, obtained from long-term cultivation under specific growth conditions, greatly enhances the efficiency of transformation and allows the identification of DNA repair mechanisms that occur preferentially at different stages of the cell cycle. We demonstrate that the transformation of synchronized cells at different times was differentially associated with nonhomologous end joining (NHEJ) and/or homologous recombination (HR), and makes it possible to knock-in specific foreign DNA at the genomic nuclear location desired by exploiting HR. This optimization greatly reduces the overall complexity of the genome editing procedure and creates new opportunities for altering genes and their products.
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Multiplex gene editing of the Yarrowia lipolytica genome using the CRISPR-Cas9 system.
Yarrowia lipolytica is categorized as a generally recognized as safe (GRAS) organism and is a heavily documented, unconventional yeast that has been widely incorporated into multiple industrial fields to produce valuable biochemicals. This study describes the construction of a CRISPR-Cas9 system for genome editing in Y. lipolytica using a single plasmid (pCAS1yl or pCAS2yl) to transport Cas9 and relevant guide RNA expression cassettes, with or without donor DNA, to target genes. Two Cas9 target genes, TRP1 and PEX10, were repaired by non-homologous end-joining (NHEJ) or homologous recombination, with maximal efficiencies in Y. lipolytica of 85.6 % for the wild-type strain and 94.1 % for the ku70/ku80 double-deficient strain, within 4 days. Simultaneous double and triple multigene editing was achieved with pCAS1yl by NHEJ, with efficiencies of 36.7 or 19.3 %, respectively, and the pCASyl system was successfully expanded to different Y. lipolytica breeding strains. This timesaving method will enable and improve synthetic biology, metabolic engineering and functional genomic studies of Y. lipolytica.
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Use of designer nucleases for targeted gene and genome editing in plants.
The ability to efficiently inactivate or replace genes in model organisms allowed a rapid expansion of our understanding of many of the genetic, biochemical, molecular and cellular mechanisms that support life. With the advent of new techniques for manipulating genes and genomes that are applicable not only to single-celled organisms, but also to more complex organisms such as animals and plants, the speed with which scientists and biotechnologists can expand fundamental knowledge and apply that knowledge to improvements in medicine, industry and agriculture is set to expand in an exponential fashion. At the heart of these advancements will be the use of gene editing tools such as zinc finger nucleases, modified meganucleases, hybrid DNA/RNA oligonucleotides, TAL effector nucleases and modified CRISPR/Cas9. Each of these tools has the ability to precisely target one specific DNA sequence within a genome and (except for DNA/RNA oligonucleotides) to create a double-stranded DNA break. DNA repair to such breaks sometimes leads to gene knockouts or gene replacement by homologous recombination if exogenously supplied homologous DNA fragments are made available. Genome rearrangements are also possible to engineer. Creation and use of such genome rearrangements, gene knockouts and gene replacements by the plant science community is gaining significant momentum. To document some of this progress and to explore the technology's longer term potential, this review highlights present and future uses of designer nucleases to greatly expedite research with model plant systems and to engineer genes and genomes in major and minor crop species for enhanced food production.
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SNIPSNP: precision design of CRISPR/Cas9 knock-in reagents for variant correction and disease modeling.
We present SNIPSNP (crisprtools.org/snipsnp), a comprehensive bioinformatics pipeline for designing experiments for CRISPR-induced homology-directed repair (HDR). The tool addresses the critical challenge of Cas9 re-cleavage by simplifying the selection of "blocking" silent variants that are effective at inhibiting RNP binding upon donor-templated editing. SNIPSNP handles complex edits, including indels, and uses multi-objective optimization to balance editing efficiency with biological safety. From user-defined wild-type and desired HDR alleles, the pipeline identifies candidate guides, annotating them with integrated efficiency scores and genome-wide off-target assessments. Uniquely, SNIPSNP evaluates guide binding against the post-edit genome to determine whether the therapeutic variant alone disrupts repeated Cas9 recognition. When necessary, it introduces synonymous blocking variants, prioritizing PAM and seed regions to minimize re-cleavage probability and editing of the wild-type (WT) allele when editing heterozygous variants. All candidate modifications undergo safety profiling and prioritization of known benign variants from dbSNP. We experimentally validated SNIPSNP and benchmarked it on pathogenic inborn error of immunity variants in primary patient T-cells. Across loci, SNIPSNP-designed templates outperform standard "correction-only" strategies, demonstrating enhanced precision editing, and reduced re-cleavage, establishing SNIPSNP as a robust platform for genome editing and disease modeling.
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