Donor Plasmid Research Articles

Design and construction of a GAP-43 reporter system for potential identification of effective therapeutics for peripheral nerve regeneration

Background: Peripheral nerve injury (PNI) is a condition that can result in muscle paralysis and sensory disturbances. Electrical stimulation and/or the application of exogenous neurotrophic factors and cytokines are effective at enhancing nerve regeneration and is mediated via the expression of regeneration associated genes (RAGs) such as the growth associated protein GAP-43. Therapeutic upregulation of GAP-43 has potential use as a treatment for improving recovery from PNI. Few studies have investigated the potential of increasing GAP-43 for PNI therapeutic purposes and current methods for measuring GAP-43 expression are limited. Aims and Objectives: The broader aim of this work was to construct a motor neuron-like cell model with a GAP-43 reporter system. Such a model would have potential use in screening for novel therapeutics that upregulate GAP-43 and in the optimisation of electrical stimulation treatment in combination with these therapies. The key aim of the work was to design and construct a Cas9 expressing plasmid bearing a gRNA that targets GAP-43 cleavage and a donor plasmid bearing a reporter GFP or Neo cassette flanked with 5’ and 3’ GAP-43 homology arms (HAs) to facilitate the insertion of the reporter immediately 3’ of the GAP-43 promoter via CRISPR/Cas9 homology directed repair (HDR). Such an insertion would enable quantitative measurement of endogenous expression of the GAP-43 gene. Methods and Results: To guide the Cas9 nuclease to the target location, GAP-43 gRNA oligomers were designed and cloned downstream of the U6 promoter in the Cas9 expression plasmid px330, which also expresses the Cas9 gene and the cloned gRNA when transfected into cells. For CRISPR/Cas9 HDR, the 5’ and 3’ GAP-43 HAs were amplified from mouse genomic DNA and cloned into the donor plasmid using Gibson Assembly so that they flanked the reporter cassette. Results: This work successfully constructed the Cas9 gRNA expressing plasmid to target cleavage of the GAP-43 gene in mouse cell lines and has provided the complete design and construction foundation for generation of the reporter system for the endogenous GAP-43 gene in mice.

Precision and efficacy of RNA-guided DNA integration in high-expressing muscle loci

Gene replacement therapies primarily rely on adeno-associated virus (AAV) vectors for transgene expression. However, episomal expression can decline over time due to vector loss or epigenetic silencing. CRISPR-based integration methods offer promise for long-term transgene insertion. While the development of transgene integration methods has made substantial progress, identifying optimal insertion loci remains challenging. Skeletal muscle is a promising tissue for gene replacement owing to low invasiveness of intramuscular injections, relative proportion of body mass, the multinucleated nature of muscle, and the potential for reduced adverse effects. Leveraging endogenous promoters in skeletal muscle, we evaluated two highly expressing loci using homology-independent targeted integration (HITI) to integrate reporter or therapeutic genes in mouse myoblasts and skeletal muscle tissue. We hijacked the muscle creatine kinase (Ckm) and myoglobin (Mb) promoters by co-delivering CRISPR-Cas9 and a donor plasmid with promoterless constructs encoding green fluorescent protein (GFP) or human Factor IX (hFIX). Additionally, we deeply profiled our genome and transcriptome outcomes from targeted integration and evaluated the safety of the proposed sites.This study introduces a proof-of-concept technology for achieving high-level therapeutic gene expression in skeletal muscle, with potential applications in targeted integration-based medicine and synthetic biology.

STRAIGHT-IN: a platform for rapidly generating panels of genetically modified human pluripotent stem cell lines.

Targeted integration of large DNA cargoes (>10 kb) or genomic replacements in mammalian cells, such as human pluripotent stem cells (hPS cells), remains challenging. Here we describe a platform termed serine and tyrosine recombinase-assisted integration of genes for high-throughput investigation (STRAIGHT-IN) to circumvent this. First, a landing pad cassette is precisely inserted or used to replace specific genomic regions. The site-specific integrase Bxb1 then enables DNA constructs, including those >50 kb, to be integrated into the genome, while Cre recombinase excises auxiliary DNA sequences to prevent postintegrative silencing. Using a strategy whereby the positive selection marker is only expressed if the donor plasmid carrying the payload is correctly targeted, we can obtain 100% enrichment for cells containing the DNA payload. Procedures for expressing Cre efficiently also mean that a clonal isolation step is no longer essential to derive the required genetically modified hPS cells containing the integrated DNA, potentially reducing clonal variability. Furthermore, STRAIGHT-IN facilitates rapid and multiplexed generation of genetically matched hPS cells when multiple donor plasmids are delivered simultaneously. STRAIGHT-IN has various applications, which include integrating complex genetic circuits for synthetic biology, as well as creating panels of hPS cells lines containing, as necessary, hundreds of disease-linked variants for disease modeling and drug discovery. After establishing the hPS cell line containing the landing pad, the entire procedure, including donor plasmid synthesis, takes 1.5-3 months, depending on whether single or multiple DNA payloads are integrated. This protocol only requires the researcher to be skilled in molecular biology and standard cell culture techniques.

CILF: CRISPR/Cas9 based integration of large DNA fragments in Saccharomyces cerevisiae.

Genome integration technology has markedly expedited the construction of cell factories. However, its application is currently limited by the inefficient integration of large DNA fragments. Here, we report a CRISPR/Cas9 based integration of large DNA fragments (CILF) method to efficiently integrate large DNA fragments in Saccharomyces cerevisiae. In this approach, a fusion protein, Cas9-Brex27-FadR, was employed for the targeted delivery of donor plasmid to double-strand breaks (DSBs), while simultaneously recruiting Rad51 to enhance the efficiency of homologous recombination (HR). Our findings demonstrate that this method can achieve an integration efficiency of 98% for 10 kb DNA fragments and nearly 80% for 40 kb DNA fragments at a single site, using donor plasmids with 1000 bp homology arms (HAs) and 12 FadR binding sites (BSs). The CILF technique significantly enriches the synthetic biology toolbox of S. cerevisiae, offering significant potential to propel advancements in both synthetic biology and metabolic engineering.

Genome editing using type I-E CRISPR-Cas3 in mice and rat zygotes

The type I CRISPR system has recently emerged as a promising tool, especially for large-scale genomic modification, but its application to generate model animals by editing zygotes had not been established. In this study, we demonstrate genome editing in zygotes using the type I-E CRISPR-Cas3 system, which efficiently generates deletions of several thousand base pairs at targeted loci in mice with 40%-70% editing efficiency without off-target mutations. To overcome the difficulties associated with detecting the variable deletions, we used a newly long-read sequencing-based multiplex genotyping approach. Demonstrating remarkable versatility, our Cas3-based technique was successfully extended to rats as well as mice, even by zygote electroporation methods. Knockin for SNP exchange and genomic replacement with a donor plasmid were also achieved in mice. This pioneering work with the type I CRISPR zygote editing system offers increased flexibility and broader applications in genetic engineering across different species.

In vitro conjugation of IncB/O-plasmid: Minimum inhibitory concentration of β-lactams increases 16-fold in Salmonella enterica compared with Escherichia coli.

The use of antimicrobials in poultry leaves residues in the litter, favoring the emergence of antimicrobial-resistant pathogens and making it a source of contamination. An in vitro 4 × 4 factorial trial was performed to investigate the influence of four treatments, consisting of antimicrobial sub-concentrations, on the transference of IncB/O-plasmid through conjugation in four groups. Each group was composed of one plasmid donor bacterium (Escherichia coli H2332) and a recipient bacterium (Escherichia coli J62 or Salmonella enterica serovars, Enteritidis, Typhimurium, or Heidelberg). Our results showed a little decrease in the conjugation frequency in almost all treatments between the two bacterial species, which varied according to each strain. The MIC test revealed an increase of up to 4096-fold in resistance to beta-lactams in Salmonella serovars after plasmid acquisition. This finding suggests that some genetic apparatus may be involved in increased antimicrobial resistance in Salmonella serovars after the acquisition of primary resistance determinants.

Efficient and markerless gene integration with SlugCas9-HF in Kluyveromyces marxianus

The nonconventional yeast Kluyveromyces marxianus has potential for industrial production, but the lack of advanced synthetic biology tools for precise engineering hinders its rapid development. Here, we introduce a CRISPR-Cas9-mediated multilocus integration method for assembling multiple exogenous genes. Using SlugCas9-HF, a high-fidelity Cas9 nuclease, we enhance gene editing precision. Specific genomic loci predisposed to efficient integration and expression of heterologous genes are identified and combined with a set of paired CRISPR-Cas9 expression plasmids and donor plasmids to establish a CRISPR-based biosynthesis toolkit. This toolkit enables genome integration of large gene modules over 12 kb and achieves simultaneous quadruple-locus integration in a single step with 20% efficiency. As a proof-of-concept, we apply the toolkit to screen for gene combinations that promote heme production, revealing the importance of HEM4Km and HEM12Sc. This CRISPR-based toolkit simplifies the reconstruction of complex pathways in K. marxianus, broadening its application in synthetic biology.

Targeted insertion of conditional expression cassettes into the mouse genome using the modified i-PITT

BackgroundTransgenic (Tg) mice are widely used in biomedical research, and they are typically generated by injecting transgenic DNA cassettes into pronuclei of one-cell stage zygotes. Such animals often show unreliable expression of the transgenic DNA, one of the major reasons for which is random insertion of the transgenes. We previously developed a method called “pronuclear injection-based targeted transgenesis” (PITT), in which DNA constructs are directed to insert at pre-designated genomic loci. PITT was achieved by pre-installing so called landing pad sequences (such as heterotypic LoxP sites or attP sites) to create seed mice and then injecting Cre recombinase or PhiC31 integrase mRNAs along with a compatible donor plasmid into zygotes derived from the seed mice. PITT and its subsequent version, improved PITT (i-PITT), overcome disadvantages of conventional Tg mice such as lack of consistent and reliable expression of the cassettes among different Tg mouse lines, and the PITT approach is superior in terms of cost and labor. One of the limitations of PITT, particularly using Cre-mRNA, is that the approach cannot be used for insertion of conditional expression cassettes using Cre-LoxP site-specific recombination. This is because the LoxP sites in the donor plasmids intended for achieving conditional expression of the transgene will interfere with the PITT recombination reaction with LoxP sites in the landing pad.ResultsTo enable the i-PITT method to insert a conditional expression cassette, we modified the approach by simultaneously using PhiC31o and FLPo mRNAs. We demonstrate the strategy by creating a model containing a conditional expression cassette at the Rosa26 locus with an efficiency of 13.7%. We also demonstrate that inclusion of FLPo mRNA excludes the insertion of vector backbones in the founder mice.ConclusionsSimultaneous use of PhiC31 and FLP in i-PITT approach allows insertion of donor plasmids containing Cre-loxP-based conditional expression cassettes.

Large T antigen mediated target gene replication improves site-specific recombination efficiency.

With advantages of high-fidelity, monoclonality and large cargo capacity, site-specific recombination (SSR) holds great promises for precise genomic modifications. However, broad applications of SSR have been hurdled by low integration efficiency, and the amount of donor DNA available in nucleus for SSR presents as a limiting factor. Inspired by the DNA replication mechanisms observed in double-stranded DNA virus SV40, we hypothesized that expression of SV40 large T antigen (TAg) can increase the copy number of the donor plasmid bearing an SV40 origin, and in consequence promote recombination events. This hypothesis was tested with dual recombinase-mediated cassette exchange (RMCE) in suspension 293F cells. Results showed that TAg co-transfection significantly enhanced SSR in polyclonal cells. In the monoclonal cell line carrying a single landing pad at an identified genomic locus, 12% RMCE efficiency was achieved, and such improvement was indeed correlated with donor plasmid amplification. The developed TAg facilitated RMCE (T-RMCE) was exploited for the construction of large libraries of >107 diversity, from which GFP variants with enhanced fluorescence were isolated. We expect the underlying principle of target gene amplification can be applicable to other SSR processes and gene editing approaches in general for directed evolution and large-scale genomic screening in mammalian cells.

Gene targeting in adult organs using in vivo cleavable donor plasmids for CRISPR-Cas9 and CRISPR-Cas12a

The CRISPR-Cas system for in vivo genome editing is a powerful tool for gene therapy against several diseases. We have previously developed the pCriMGET_9-12a system, an in vivo cleavable donor plasmid for precise targeted knock-in of exogenous DNA by both Cas9 and Cas12a. Here, we show that the pCriMGET_9-12a system can be applied for in vivo in-frame knock-in of exogenous DNA in adult mouse liver by hydrodynamic delivery of the targeting plasmids. The in vivo cleavable pCriMGET_9-12a donor plasmids significantly increased the knock-in efficiency of both CRISPR-Cas9 and CRISPR-Cas12a in the adult mouse liver compared to uncleavable donor plasmids. This strategy also achieved in-frame reporter gene knock-in without indel mutations. Therefore, in vivo gene targeting using the pCriMGET_9-12a system may contribute to the establishment of safer, more precise, versatile and efficient gene therapy methods in adult organs.

Genetic Knock-Ins of Endogenous Fluorescent Tags in RAW 264.7 Murine Macrophages Using CRISPR/Cas9 Genome Editing.

CRISPR/Cas9 genome editing is a widely used tool for creating genetic knock-ins, which allow for endogenous tagging of genes. This is in contrast with random insertion using viral vectors, where expression of the inserted transgene changes the total copy number of a gene in a cell and does not reflect the endogenous chromatin environment or any trans-acting regulation experienced at a locus. There are very few protocols for endogenous fluorescent tagging in macrophages. Here, we describe a protocol to design and test CRISPR guide RNAs and donor plasmids, to transfect them into RAW 264.7 mouse macrophage-like cells using the Neon transfection system and to grow up clonal populations of cells containing the endogenous knock-in at various loci. We have used this protocol to create endogenous fluorescent knock-ins in at least six loci, including both endogenously tagging genes and inserting transgenes in the Rosa26 and Tigre safe harbor loci. This protocol uses circular plasmid DNA as the donor template and delivers the sgRNA and Cas9 as an all-in-one expression plasmid. We designed this protocol for fluorescent protein knock-ins; it is best used when positive clones can be identified by fluorescence. However, it may be possible to adapt the protocol for non-fluorescent knock-ins. This protocol allows for the fairly straightforward creation of clonal populations of macrophages with tags at the endogenous loci of genes. We also describe how to set up imaging experiments in 24-well plates to track fluorescence in the edited cells over time. Key features • CRISPR knock-in of fluorescent proteins in RAW 264.7 mouse macrophages at diverse genomic loci. • This protocol is optimized for the use of the Neon transfection system. • Includes instructions for growing up edited clonal populations from single cells with one single-cell sorting step and efficient growth in conditioned media after cell sorting. • Designed for knocking in fluorescent proteins and screening transfected cells by FACS, but modification for non-fluorescent knock-ins may be possible.

Decoding the complexity of on-target integration: characterizing DNA insertions at the CRISPR-Cas9 targeted locus using nanopore sequencing

BackgroundCRISPR-Cas9 technology has advanced in vivo gene therapy for disorders like hemophilia A, notably through the successful targeted incorporation of the F8 gene into the Alb locus in hepatocytes, effectively curing this disorder in mice. However, thoroughly evaluating the safety and specificity of this therapy is essential. Our study introduces a novel methodology to analyze complex insertion sequences at the on-target edited locus, utilizing barcoded long-range PCR, CRISPR RNP-mediated deletion of unedited alleles, magnetic bead-based long amplicon enrichment, and nanopore sequencing.ResultsWe identified the expected F8 insertions and various fragment combinations resulting from the in vivo linearization of the double-cut plasmid donor. Notably, our research is the first to document insertions exceeding ten kbp. We also found that a small proportion of these insertions were derived from sources other than donor plasmids, including Cas9-sgRNA plasmids, genomic DNA fragments, and LINE-1 elements.ConclusionsOur study presents a robust method for analyzing the complexity of on-target editing, particularly for in vivo long insertions, where donor template integration can be challenging. This work offers a new tool for quality control in gene editing outcomes and underscores the importance of detailed characterization of edited genomic sequences. Our findings have significant implications for enhancing the safety and effectiveness of CRISPR-Cas9 gene therapy in treating various disorders, including hemophilia A.

High mechanical property silk produced by transgenic silkworms expressing the Drosophila Dumpy.

Extensive research has been conducted on utilizing transgenic silkworms and their natural spinning apparatus to produce high-performance spider silk fibers. However, research on using non-spider biological proteins to optimize the molecular structure of silk protein and improve the mechanical performance of silk fibers is still relatively scarce. Dumpy, a massive extracellular matrix polypeptide, is essential for preserving the shape and structural integrity of the insect cuticle due to its remarkable tension and elasticity. Here, we constructed two transgenic donor plasmids containing the fusion genes of FibH-Dumpy and FibL-Dumpy. The results indicated the successful integration of two exogenous gene expression cassettes, driven by endogenous promoters, into the silkworm genome using piggyBac-mediated transgenic technology. Secondary structure analysis revealed a 16.7% and 13.6% increase in the β-sheet content of transgenic silks compared to wild-type (WT) silk fibers. Mechanical testing demonstrated that, compared to the WT, HDUY and LDUY transgenic silk fibers exhibited respective increases of 39.54% and 21.45% in maximum stress, 44.43% and 45.02% in toughness, and 24.91% and 28.51% in elastic recovery rate. These findings suggest that Drosophila Dumpy significantly enhanced the mechanical properties of silk, positioning it as an excellent candidate for the development of extraordinary-performance fibers. This study provides rich inspiration for using other biological proteins to construct high-performance silk fibers and expands the possibilities for designing and applying novel biomaterials.

Recombinant CHO Cell Pool Generation Using PiggyBac Transposon System.

CHO cell pools with desirable characteristics of high titer and consistent product quality are useful for rapid production of recombinant proteins. Here, we describe the generation of CHO cell pools using the piggyBac transposon system for mediating gene integration. The method describes the co-transfection of cells with the donor plasmid (coding for the gene of interest) and the helper plasmid (coding for the transposase) using polyethyleneimine (PEI). This is followed by a genetic selection for the generation of a cell pool. The resulting cell pool can be used to start a batch or fed-batch culture. Alternatively, it can be used for generation of clonal cell lines or generation of cell banks for future use.

Multiplex Marker-Less Genome Integration in Pichia pastoris Using CRISPR/Cas9.

Pichia pastoris is known for its excellent protein expression ability. As an industrial methyl nutritional yeast, it can effectively utilize methanol as the sole carbon source, serving as a potential platform for C1 biotransformation. Unfortunately, the lack of synthetic biology tools in P. pastoris limits its broad applications, particularly when multigene pathways should be manipulated. Here, the CRISPR/Cas9 system is established to efficiently integrate multiple heterologous genes to construct P. pastoris cell factories. In this protocol, with the 2,3-butanediol (BDO) biosynthetic pathway as a representative example, the procedures to construct P. pastoris cell factories are detailed using the established CRISPR-based multiplex genome integration toolkit, including donor plasmid construction, competent cell preparation and transformation, and transformant verification. The application of the CRISPR toolkit is demonstrated by the construction of engineered P. pastoris for converting methanol to BDO. This lays the foundation for the construction of P. pastoris cell factories harboring multi-gene biosynthetic pathways for the production of high-value compounds.

Efficient Gene-Editing in Human Pluripotent Stem Cells Through Simplified Assembly of Adeno-Associated Viral (AAV) Donor Templates.

Gene-edited human pluripotent stem cells provide attractive model systems to functionally interrogate the role of specific genetic variants in relevant cell types. However, the need to isolate and screen edited clones often remains a bottleneck, in particular when recombination rates are sub-optimal. Here, we present a protocol for flexible gene editing combining Cas9 ribonucleoprotein with donor templates delivered by adeno-associated virus (AAV) vectors to yield high rates of homologous recombination. To streamline the workflow, we designed a modular system for one-step assembly of targeting vectors based on Golden Gate cloning and developed a rapid protocol for small-scale isolation of AAV virions of serotype DJ. High homology-directed repair (HDR) rates in human pluripotent stem cells (hPSCs), ~70% in ACTB and ~30% in LMNB1, were achieved using this approach, also with short (300 bp) homology arms. The modular design of donor templates is flexible and allows for the generation of conditional and/or complex alleles. This protocol thus provides a flexible and efficient strategy workflow to rapidly generate gene-edited hPSC lines. Key features • Versatile approach combining AAV-DJ donors and CRISPR ribonucleoproteins, providing an efficient method for long and short edits, insertions, and deletions in human pluripotent stem cells. • One-step cloning method for rapid generation of customized AAV donor plasmids. • Simplified AAV purification pipeline for ready-to-infect virion preparations.

Optimal tagging strategies for illuminating expression profiles of genes with different abundance in zebrafish

CRISPR-mediated knock-in (KI) technology opens a new era of fluorescent-protein labeling in zebrafish, a preferred model organism for in vivo imaging. We described here an optimized zebrafish gene-tagging strategy, which enables easy and high-efficiency KI, ensures high odds of obtaining seamless KI germlines and is suitable for wide applications. Plasmid donors for 3′-labeling were optimized by shortening the microhomologous arms and by reducing the number and reversing the sequence of the consensus Cas9/sgRNA binding sites. To allow for scar-less KI across the genome, linearized dsDNA donors with 5′-chemical modifications were generated and successfully incorporated into our method. To refine the germline screen workflow and expedite the screen process, we combined fluorescence enrichment and caudal-fin junction-PCR. Furthermore, to trace proteins expressed at a low abundance, we developed a fluorescent signal amplifier using the transcriptional activation strategy. Together, our strategies enable efficient gene-tagging and sensitive expression detection for almost every gene in zebrafish.

A CRISPR-Cas9-mediated versatile method for targeted integration of a fluorescent protein gene to visualize endogenous gene expression in Xenopus laevis

Xenopus laevis is a widely used model organism in developmental and regeneration studies. Despite several reports regarding targeted integration techniques in Xenopus, there is still room for improvement of them, especially in creating reporter lines that rely on endogenous regulatory enhancers/promoters. We developed a CRISPR-Cas9-based simple method to efficiently introduce a fluorescent protein gene into 5′ untranslated regions (5′UTRs) of target genes in Xenopus laevis. A donor plasmid DNA encoding an enhanced green fluorescent protein (eGFP) flanked by a genomic fragment ranging from 66 bp to 878 bp including target 5′UTR was co-injected into fertilized eggs with a single guide RNA and Cas9 protein. Injections for krt12.2.L, myod1.S, sox2.L or brevican.S resulted in embryos expressing eGFP fluorescence in a tissue-specific manner, recapitulating endogenous expression of target genes. Integrations of the donor DNA into the target regions were examined by genotyping PCR for the eGFP-expressing embryos. The rate of embryos expressing the specific eGFP varied from 2.1% to 13.2% depending on the target locus and length of the genomic fragment in the donor plasmids. Germline transmission of an integrated DNA was observed. This simple method provides a powerful tool for exploring gene expression and function in developmental and regeneration research in X. laevis.

The strategy of knock-in with homology-directed genome editing in the model ornamental plant Petunia using CRISPR/Cas9 ribonucleoprotein complex

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated system 9 (Cas9) ribonucleoprotein (RNP) complex offers numerous advantages, including increased genome editing efficiency, reduced off-target effects, minimized risk of genomic integration, expanded applicability, and the ability to address specific ethical concerns. These benefits establish the CRISPR/Cas9 RNP complex as a valuable technique for precise genome editing in plant research and applications. Nevertheless, achieving precise knock-in using CRISPR/Cas9-mediated precision knock-in of genes by homology-directed repair (HDR) remains challenging due to the infrequent occurrence of HDR and the limited availability of DNA repair templates near double-strand breaks (DSBs). In this study, we presented the efficient utilization strategy of HDR-mediated gene knock-in through co-infection of the CRISPR/Cas9 RNP complex and donor plasmid in Petunia protoplast. We accomplish this by designing a donor plasmid targeting the chalcone synthase (CHS) in Petunia, consisting of a CRISPR RNA (crRNA)-PAM sequences flanked by a donor DNA cassette with homology arms. These findings offer insights into the application of these techniques in ornamental crops to enhance CRISPR/Cas9 RNA complex-mediated gene knock-in strategies.

Targeted genome engineering based on CRISPR/Cas9 system to enhance FVIII expression in vitro

BackgroundHemophilia A is caused by a deficiency of coagulation factor VIII in the body due to a defect in the F8 gene. The emergence of CRISPR/Cas9 gene editing technology will make it possible to alter the expression of the F8 gene in hemophiliacs, while achieving a potential cure for the disease. MethodsInitially, we identified high-activity variants of FVIII and constructed donor plasmids using enzymatic digestion and ligation techniques. Subsequently, the donor plasmids were co-transfected with sgRNA-Cas9 protein into mouse Neuro-2a cells, followed by flow cytometry-based cell sorting and puromycin selection. Finally, BDD-hF8 targeted to knock-in the mROSA26 genomic locus was identified and validated for FVIII expression. ResultsWe identified the p18T-BDD-F8-V3 variant with high FVIII activity and detected the strongest pX458-mROSA26-int1-sgRNA1 targeted cleavage ability and no cleavage events were found at potential off-target sites. Targeted knock-in of BDD-hF8 cDNA at the mROSA26 locus was achieved based on both HDR/NHEJ gene repair approaches, and high level and stable FVIII expression was obtained, successfully realizing gene editing in vitro. ConclusionsKnock-in of exogenous genes based on the CRISPR/Cas9 system targeting genomic loci is promising for the research and treatment of a variety of single-gene diseases.