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High-throughput Genetically Modified Animal Experiments Achieved by Next-generation Mammalian Genetics.
Animal models are essential tools for modern scientists to conduct biological experiments and investigate their hypotheses in vivo. However, for the past decade, raising the throughput of such animal experiments has been a great challenge. Conventionally, in vivo high-throughput assay was achieved through large-scale mutagen-driven forward genetic screening, which took years to find causal genes. In contrast, reverse genetics accelerated the causal gene identification process, but its throughput was also limited by 2 barriers, that is, the genome modification step and the time-consuming crossing step. Defined as genetics without crossing, next-generation genetics is able to produce gene-modified animals that can be analyzed at the founder generation (F0). This method is or can be accomplished through recent technological advances in gene editing and virus-based efficient gene modifications. Notably, next-generation genetics has accelerated the process of cross-species studies, and it will be a useful technique during animal experiments as it can provide genetic perturbation at an individual level without crossing. In this review, we begin by introducing the history of animal-based high-throughput analysis, with a specific focus on chronobiology. We then describe ways that gene modification efficiency during animal experiments was enhanced and why crossing remained a barrier to reaching higher efficiency. Moreover, we mention the Triple CRISPR as a critical technique for achieving next-generation genetics. Finally, we discuss the potential applications and limitations of next-generation mammalian genetics.
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Protocol for efficient CRISPR/AAV-mediated genome editing and erythroid differentiation of human hematopoietic stem and progenitor cells.
Here, we present a protocol for genome editing in human hematopoietic stem and progenitor cells (HSPCs) using CRISPR-Cas9 ribonucleoproteins and adeno-associated virus (AAV)-mediated homology-directed repair. We describe steps for AAV production, purification, and titration; HSPC thawing and culture; genome editing; and quantification of editing frequencies. We then detail procedures for erythroid differentiation assays. This protocol ensures high editing efficiency while maintaining cell viability and engraftment potential. For complete details on the use and execution of this protocol, please refer to Chu et al.1.
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Molecular circuits for genomic recording of cellular events.
Advances in precise genome editing are enabling genomic recordings of cellular events. Since the initial demonstration of CRISPR-based genome editing, the field of genomic recording has witnessed key strides in lineage recording, where clonal lineage relationships among cells are indirectly recorded as synthetic mutations. However, methods for directly recording and reconstructing past cellular events are still limited, and their potential for revealing new insights into cell fate decisions has yet to be realized. The field needs new sensing modules and genetic circuit architectures that faithfully encode past cellular states into genomic DNA recordings to achieve such goals. Here we review recently developed strategies to construct diverse sensors and explore how emerging synthetic biology tools may help to build molecular circuits for genomic recording of diverse cellular events.
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Standardized Iterative Genome Editing Method for Escherichia coli Based on CRISPR-Cas9.
The introduction of complex biosynthetic pathways into the hosts' chromosomes is gaining attention with the development of synthetic biology. While CRISPR-Cas9 has been widely employed for gene knock-in, the process of multigene insertion remains cumbersome due to laborious and empirical gene cloning procedures. To address this, we devised a standardized iterative genome editing system for Escherichia coli, harnessing the power of CRISPR-Cas9 and MetClo assembly. This comprehensive toolkit comprises two fundamental elements based on the Golden Gate standard for modular assembly of sgRNA or CRISPR arrays and donor DNAs. We achieved a gene insertion efficiency of up to 100%, targeting a single locus. Expression of tracrRNA using a strong promoter enhances multiplex genomic insertion efficiency to 7.3%, compared with 0.76% when a native promoter is used. To demonstrate the robust capabilities of this genome editing toolbox, we successfully integrated 5-10 genes from the coenzyme B12 biosynthetic pathway ranging from 5.3 to 8 Kb in length into the chromosome of E. coli chassis cells, resulting in 14 antibiotic-free, plasmid-free producers. Following an extensive screening process involving genes from diverse sources, cistronic design modifications, and chromosome repositioning, we obtained a recombinant strain yielding 1.49 mg L-1 coenzyme B12, the highest known titer achieved by using E. coli as the producer. Illuminating its user-friendliness, this genome editing system is an exceedingly versatile tool for expediently integrating complex biosynthetic pathway genes into hosts' genomes, thus facilitating pathway optimization for chemical production.
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[Applications of the CRISPR/Cas9 system in insects].
The CRISPR/Cas9 (clustered regularly interspaced short palindromic repeat/CRISPR-associated) system guides Cas9 to specific genomic locations by a short RNA search string. This technology enables the systematic interrogation of mammalian genome editing, repairing damaged genes, silencing harmful genes and improving quality traits. In recent years, with the introduction of the CRISPR/Cas9 system for easy, fast and efficient genetic modification, it has been possible to conduct meaningful functional studies in a broad array of insect species, such as Drosophila, Bombyx mori, Aedes aegypti and butterflies et al. In this review, we summarize the application of CRISPR/Cas9 in different insect species, discuss methods for its promotion, and consider its application for future insect studies.
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Crispr/Cas9-mediated cleavages facilitate homologous recombination during genetic engineering of a large chromosomal region.
Homologous recombination over large genomic regions is difficult to achieve due to low efficiencies. Here, we report the successful engineering of a humanized mTert allele, hmTert, in the mouse genome by replacing an 18.1-kb genomic region around the mTert gene with a recombinant fragment of over 45.5 kb, using homologous recombination facilitated by the Crispr/Cas9 technology, in mouse embryonic stem cells (mESCs). In our experiments, with DNA double-strand breaks (DSBs) generated by Crispr/Cas9 system, the homologous recombination efficiency was up to 11% and 16% in two mESC lines TC1 and v6.5, respectively. Overall, we obtained a total of 27 mESC clones with heterozygous hmTert/mTert alleles and three clones with homozygous hmTert alleles. DSBs induced by Crispr/Cas9 cleavages also caused high rates of genomic DNA deletions and mutations at single-guide RNA target sites. Our results indicated that the Crispr/Cas9 system significantly increased the efficiency of homologous recombination-mediated gene editing over a large genomic region in mammalian cells, and also caused frequent mutations at unedited target sites. Overall, this strategy provides an efficient and feasible way for manipulating large chromosomal regions.
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Proximity-activated guide RNA of CRISPR-Cas12a for programmable diagnostic detection and gene regulation.
The flexibility and programmability of CRISPR-Cas technology have made it one of the most popular tools for biomarker diagnostics and gene regulation. Especially, the CRISPR-Cas12 system has shown exceptional clinical diagnosis and gene editing capabilities. Here, we discovered that although the top loop of the 5' handle of guide RNA can undergo central splitting, deactivating CRISPR-Cas12a, the segments can dramatically restore CRISPR function through nucleic acid self-assembly or interactions with small molecules and aptamers. This discovery forms the basis of an engineered Cas12a system with a programmable proximity-activated guide RNA (PARC-Cas12a) that links targets of interest to dsDNA. Leveraging the efficient trans- and cis-cleavage of Cas12, our findings further inspired a detection platform design for RNAs or non-nucleic acid biomarkers, enabling highly sensitive and multiplexed analysis. We further demonstrated the feasibility of RNA-controllable gene knockout/knockdown in Escherichia coli. Notably, we successfully validated the gene regulatory capabilities of the PARC-Cas12a system within mammalian cell systems by utilizing the classical theophylline molecule-aptamer system. Our results introduce a programmable toolbox for precise diagnostics and cell regulation, allowing the development of versatile diagnostic tools, complex synthetic biological circuits, and cellular biosensors.
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Protocol for the design, conduct, and evaluation of prime editing in human pluripotent stem cells.
Prime editing introduces single-nucleotide polymorphism changes, small deletions, or insertions at a specific genome site without double-stranded DNA breaks or the need for the donor template. Here, we present a protocol to design, conduct, and evaluate prime editing in human pluripotent stem cells. We describe steps for pegRNA and nicking sgRNA design and cloning, the prime editing tool electroporation, and the efficiency evaluation using Miseq. We elaborate the process of GBA (N370S) mutation induction and correction as an example. For complete details on the use and execution of this protocol, please refer to Li et al. (2022).1.
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A mathematical model of HIV dynamics treated with a population of gene-edited haematopoietic progenitor cells exhibiting threshold phenomenon.
The use of gene-editing technology has the potential to excise the CCR5 gene from haematopoietic progenitor cells, rendering their differentiated CD4-positive (CD4+) T cell descendants HIV resistant. In this manuscript, we describe the development of a mathematical model to mimic the therapeutic potential of gene editing of haematopoietic progenitor cells to produce a class of HIV-resistant CD4+ T cells. We define the requirements for the permanent suppression of viral infection using gene editing as a novel therapeutic approach. We develop non-linear ordinary differential equation models to replicate HIV production in an infected host, incorporating the most appropriate aspects found in the many existing clinical models of HIV infection, and extend this model to include compartments representing HIV-resistant immune cells. Through an analysis of model equilibria and stability and computation of $R_0$ for both treated and untreated infections, we show that the proposed therapy has the potential to suppress HIV infection indefinitely and return CD4+ T cell counts to normal levels. A computational study for this treatment shows the potential for a successful 'functional cure' of HIV. A sensitivity analysis illustrates the consistency of numerical results with theoretical results and highlights the parameters requiring better biological justification. Simulations of varying level production of HIV-resistant CD4+ T cells and varying immune enhancements as the result of these indicate a clear threshold response of the model and a range of treatment parameters resulting in a return to normal CD4+ T cell counts.
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Development of a CRISPR/Cas9-degron system that enables in vivo specific gene depletion in leukemia models.
The CRISPR/Cas9 system has transformed genome editing, yet precise temporal control of Cas9 activity remains challenging. We developed a Cas9-degron platform that couples degron-tagged Cas9 with a dTAG-based chemical degradation strategy. In the presence of dTAG, Cas9 is rapidly and near-completely degraded, preventing editing; upon dTAG withdrawal, Cas9 activity is restored, enabling precise temporal control. Using this system, we achieved selective in vivo gene depletion in acute myeloid leukemia (AML) models and confirmed that SETDB1, a histone H3K9 methyltransferase, is essential for the in vivo growth of both human (MOLM13) and murine (cSAM) AML cells. By maintaining SETDB1 intact prior to transplantation and depleting it afterward, we avoided culture-induced pre-selection bias inherent to sgRNA transduction and validated its critical role in AML progression within the in vivo context. The Cas9-degron retains activity and delivery efficiency comparable to conventional Cas9 in the absence of dTAG. Thus, this versatile system provides a superior alternative to conventional Cas9 and a powerful platform for in vivo CRISPR screening, gene function studies, and potentially temporally controlled gene therapy.
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