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Synergistic effects among multiple gene mutations are involved in cancer development and progression. However, developing genetically modified mouse models to analyze various combinations of mutations is extremely labor-intensive and time-consuming. To address these problems, we developed a novel method for in vivo multiplexed genome editing of the murine uterus to model human endometrial carcinoma (EMC). To do this, we injected a CRISPR-Cas9 ribonucleoprotein complex into the uterine cavity of adult female mice, followed by electroporation. Evaluation of reporter mice demonstrated that genome editing occurred specifically in uterine epithelial cells, which are the origin of EMCs. Simultaneous targeting of Pten/Trp53/Lkb1, or targeting of Pten/Lkb1 along with the Ctnnb1ΔEx3 mutation, resulted in efficient generation of invasive tumors in wild-type females within 3 months. This novel method will enable rapid and easy validation of many combinations of gene mutations that lead to endometrial carcinogenesis.

Recent advances in the CRISPR/Cas9 system have demonstrated it to be an efficient gene-editing technology for various organisms. Laboratory mice and rats are widely used as common models of human diseases; however, the current standard method to create genome-engineered animals is laborious and involves three major steps: isolation of zygotes from females, ex vivo micromanipulation of zygotes, and implantation into pseudopregnant females. To circumvent this, we recently developed a novel method named Genome-editing via Oviductal Nucleic Acids Delivery (GONAD). This method does not require the ex vivo handling of embryos; instead, it can execute gene editing with just one step, via the delivery of a genome-editing mixture into embryos in the oviduct, by electroporation. Here, we present a further improvement of GONAD that is easily applicable to both mice and rats. It is a rapid, low-cost, and ethical approach fulfilling the 3R principles of animal experimentation: Reduction, Replacement, and Refinement. This method has been reconstructed and renamed as "improved GONAD (i-GONAD)" for mice, and "rat improved GONAD (rGONAD)" for rats.

Multiple sclerosis is a neurological disorder in which the myelin sheaths of axons are damaged by the immune response. We report here a three-dimensional structural analysis of brain and spinal cord tissues of a mouse model of multiple sclerosis, known as experimental autoimmune encephalomyelitis (EAE). EAE-induced mice were raised with or without administration of fingolimod, which is used in the treatment of multiple sclerosis. Brains and spinal cords dissected from the EAE mice were lyophilized so as to reconstitute the intrinsic contrast of tissue elements, such as axons, in X-ray images. Three-dimensional structures of the brain hemispheres and spinal cords of the EAE mice were visualized with synchrotron radiation microtomography. Microtomographic cross sections reconstructed from the X-ray images revealed dilation of capillary vessels and vacuolation in the spinal cord of the EAE mice. Vacuolation was also observed in the cerebellum, suggesting that the neuroinflammatory response progressed in the brain. The vessel networks and vacuolation lesions in the spinal cords were modelled by automatically tracing the three-dimensional image in order to analyze the tissue structures quantitatively. The results of the analysis indicated that the distribution of vacuolations was not uniform but three-dimensionally localized. The mean vessel diameter showed a linear correlation with the clinical score, indicating that vasodilation is relevant to paralysis severity in the disease model. We suggest that vasodilation and vacuolation are related with neurological symptoms of multiple sclerosis., 22 pages, 5 figures, 1 table

Contents The CRISPR/Cas9 system has enabled the editing of mammalian genomes; however, its applicability and efficiency in the pig genome has not been studied in depth. The α-gal epitope synthesized by α-1,3-galactosyltransferase gene (GGTA1) is known as a xenoantigen obtained upon pig-to-human xenotransplantation. We here employed the CRISPR/Cas9 system-mediated knock-in of endogenous GGTA1 via targeted homologous recombination (HR). Linearized donors with ~800-bp homology flanking the CRISPR/Cas9 target site [exon 4 (containing ATG) of GGTA1] served as a template for gene targeting by HR. Using a targeted toxin strategy to select clones lacking α-gal epitope expression, we successfully obtained several knock-in clones within 3 weeks of initial transfection. These results suggest that the use of CRISPR/Cas9-mediated HR to knock-in a mutated fragment at defined loci represents an efficient strategy to achieve the rapid modulation of genes of interest in swine cells and is a promising tool for the creation of KO piglets.

Graphical abstract, Highlights • A Cas9 target that cleaves precisely at the ROSA26 original provirus integration site. • Combining CRISPR/Cas9 with the PITT system can enable insertion of larger DNAs. • Anomalous and mosaic sequence insertions can occur with the CRISPR/Cas9 system., Targeted transgenic mouse models, where an exogenous gene is inserted into a specified genomic locus to achieve its stable and reliable expression, have been widely used in biomedical research. However, the available methodologies for targeted insertion of sequences require many laborious steps that involve the use of embryonic stem (ES) cells. We recently developed Pronuclear Injection-based Targeted Transgenesis (PITT), a method that uses a recombinase-mediated cassette exchange (RMCE) to enable insertion of sequences at a predetermined genomic locus, such as ROSA26. The PITT technique uses fertilized eggs (instead of ES cells) collected from ‘seed mice’ that contain the RMCE landing pad. The PITT method can rapidly generate reliable targeted transgenic mice; it requires a seed mouse, which in our previous study was generated using ES cell targeting approaches. Here, we demonstrate that seed mice containing the RMCE landing pad can be developed rapidly by using the CRISPR/Cas9 system. One of the CRISPR targets tested in this study enabled the insertion of sequences precisely at the original ROSA26 provirus integration site. We anticipate that using a similar approach, PITT landing pad sequences can be rapidly and precisely inserted at other genomic loci to develop an array of PITT tools. This two-step strategy combines the best features of the two newer technologies—rapid creation of PITT landing pads using the CRISPR/Cas9 system and efficient and precise insertion of larger cassettes at the landing pads using PITT. This study also revealed that anomalous and mosaic sequence insertions can occur with the CRISPR/Cas9 system.

Isolation of cells harboring exogenous DNA is typically achieved by the introduction of plasmids, but its efficiency remains still low. In this study, we developed a novel strategy to obtain stable transfectants efficiently. Porcine embryonic fibroblasts were transfected with two plasmids: (i) pTransIEnd, which comprises the ubiquitous promoter, the piggyBac (PB) transposase gene, an internal ribosomal entry site, the Clostridium perfringens-derived endo-β-galactosidase C (EndoGalC) gene, and a poly(A) tail and (ii) a PB-based plasmid, termed pT-EGFP, which contains enhanced green fluorescent protein (EGFP) expression unit flanked by PB acceptor sites. The PB transposase can accelerate the chromosomal integration of transposon vectors. EndoGalC expression results in removal of a cell surface α-Gal epitope, which is specifically recognized by Bandeiraea simplicifolia isolectin-B4 (IB4). Four days after transfection, cells were treated with IB4SAP (IB4 conjugated to saporin, which eliminates any α-Gal epitope-expressing cells) for a short period, followed by standard culture for approximately 10 days. Several colonies emerged, most of which were positive for EGFP expression and lacked TransIEnd. These results indicated that the proposed approach is useful and efficient for obtaining stable transfectants without the use of drug-resistance genes, and offers a novel route for gene manipulation in cultured nonhuman mammalian cells.

Background The recent development of the type II clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 system has enabled genome editing of mammalian genomes including those of mice and human; however, its applicability and efficiency in the pig have not been studied in depth. Here, using the CRISPR/Cas9 system, we aimed to destroy the function of the porcine α-1,3-galactosyltransferase (α-GalT) gene (GGTA1) whose product is responsible for the synthesis of the α-Gal epitope, a causative agent for hyperacute rejection upon pig-to-human xenotransplantation. Methods Porcine embryonic fibroblasts were transfected with a Cas9 expression vector and guide RNA specifically designed to target GGTA1. At 4 days after transfection, the cells were incubated with IB4 conjugated with saporin (IB4SAP), which eliminates α-Gal epitope-expressing cells. Therefore, the cells surviving after IB4SAP treatment would be those negative for α-Gal epitope expression, which in turn indicates the generation of GGTA1 biallelic knockout (KO) cells. Results Of the 1.0 × 106 cells transfected, 10–33 colonies survived after IB4SAP treatment, and almost all colonies (approximately 90%) were negative for staining with red fluorescence-labeled IB4. Sequencing of the mutated portion of GGTA1 revealed a frameshift of the α-GalT protein. Porcine blastocysts derived from the somatic cell nuclear transfer of these α-Gal epitope-negative cells also lacked the α-Gal epitope on their surface. Conclusions These results demonstrated that the CRISPR/Cas9 system can efficiently induce the biallelic conversion of GGTA1 in the resulting somatic cells and is thus a promising tool for the creation of KO cloned piglets.

Microinjection of DNA expression cassettes into fertilized zygotes has been a standard method for generating transgenic animal models. While efficient, the injected DNA integrates randomly into the genome, leading to potential disruption of endogenous genes or regulatory elements, variation in copy number, or integration into heterochromatic regions that inhibit transgene expression. A recently developed method addresses such pitfalls of traditional transgenesis by targeting the transgene to predetermined sites in the genome that can safely harbor exogenous DNA. This method, called Pronuclear Injection-based Targeted Transgenesis (PITT), employs an enzymatic transfer of exogenous DNA from a donor vector to a previously created landing-pad site in the mouse genome. DNA transfer is achieved using molecular tools such as the Cre-LoxP recombinase and PhiC31-attB/P integrase systems. Here, we provide protocols for performing PITT and an overview of the current PITT tools available to the research community. © 2016 by John Wiley & Sons, Inc.

Transgenic technologies used for creating a desired genomic change in animals involve three critical steps: isolation of fertilized eggs, microinjection of transgenic DNA into them and their subsequent transfer to recipient females. These ex vivo steps have been widely used for over 3 decades and they were also readily adapted for the latest genome editing technologies such as ZFNs, TALENs, and CRISPR/Cas9 systems. We recently developed a method called GONAD (Genome editing via Oviductal Nucleic Acids Delivery) that does not require all the three critical steps of transgenesis and therefore relieves the bottlenecks of widely used animal transgenic technologies. Here we provide protocols for the GONAD system.

Porcine embryonic fibroblasts (PEFs) are widely used as donor cells for somatic cell nuclear transfer (SCNT) in pigs. Transfection of PEFs with exogenous DNA is essential for producing genetically modified (GM; transgenic or knockout) pigs via SCNT. In this case, selectable markers are strictly required selecting and enriching stably transfected cells. The most frequently used selective drug for this purpose is a neomycin analogue (G418/geneticin); neo has been widely used as a selectable marker gene in the genomic manipulation of pigs. However, little is known about optimal concentrations of other selection drugs. This often hampers functional analysis of the porcine genome and development of individual GM pigs. This study explores the optimal concentrations of selective drugs, other than neomycin, that can be used for the selection of transfected PEFs. Porcine embryonic fibroblasts were incubated in media containing different concentrations of drugs for up to 10 days, to determine the optimal drug concentrations fatal for PEFs. The following concentrations were found to be optimal selective concentrations for use with PEFs: G418/geneticin, 400 μg/ml; blasticidin S, 8 μg/ml; hygromycin B, 40 μg/ml; puromycin, 2 μg/ml; and zeocin, 800 μg/ml. Repeated transfections with plasmids carrying selectable markers resulted in the generation of multidrug-resistant swine transfectants. Furthermore, these markers were found to be independent. The present information will be useful for the production of SCNT-mediated GM piglets that express multiple transgenes.

Porcine embryonic fibroblasts (PEFs) have been used extensively as donor nuclei for the production of cloned pigs via somatic cell nuclear transfer (SCNT). Somatic cell nuclear transfer of gene-targeted PEFs has been the only way to produce gene-targeted pigs, given the lack of germ-line-competent porcine embryonic stem cells. Unlike other primary-cultured cells, such as murine embryonic fibroblasts, a single porcine PEF is unable to proliferate under normal conditions in which a certain number of PEFs (likely over 100) can grow normally. This limitation greatly hampers re-cloning of gene-modified PEFs, which is required for SCNT. Herein, we demonstrate the cultivation of a single PEF transfectant carrying the pEGFP-N1 plasmid with intact normal PEFs (>100) in a Terasaki microtest plate, which resulted in stimulation of the growth of the former cell (doubling time = 2.6 days). In contrast, when a single cell was cultured, it could typically divide only once and never divided more than twice. When a single transfectant was seeded in a well of a 96-well plate together with 5 × 10(4) untransfected PEFs and was subsequently selected in the presence of G418, we obtained a pure cell population of single-cell origin. Thus, this method should be useful for the purification of target recombinant pig clones from mosaic populations that cannot be cultured as a single cell or for which suitable cell growth-promoting conditions are unclear.

Akasaka E, Watanabe S, Himaki T, Ohtsuka M, Yoshida M, Miyoshi K, Sato M. Enrichment of xenograft-competent genetically modified pig cells using a targeted toxin, isolectin BS-I-B4 conjugate. Xenotransplantation 2010; 17: 81–89. © 2010 John Wiley & Sons A/S. Abstract: Background: The recent availability of α-1,3-galatosyltransferase knockout pigs has eliminated anti-Gal antibodies to the galα1-3gal (αgal epitope) as the major barrier to xenotransplantation. These αgal epitope-negative animals can also be produced by somatic cell nuclear transfer of cells overexpressing endo-β-galactosidase (EndoGalC), an enzyme capable of digesting the αgal epitope. For this, selection of cells with highly reduced synthesis of αgal epitope is a prerequisite. In this study, we developed a novel method of selection using isolectin BS-I-B4-conjugated saporin (IB4-SAP), a targeted cytotoxin, that is specific for the terminal αgal epitope. Methods: A mixture of αgal epitope-expressing and non-expressing pig cells was obtained by transfection with an EndoGalC expression vector. These cells were incubated with a solution containing IB4-SAP for 2 h at 37 °C, and subsequently cultivated for over 2 months under general conditions. Results: Almost all (98%) of surviving cells were completely negative for expression of αgal epitope, as confirmed by cytochemical staining using fluorescence-labeled IB4. FACS analysis also confirmed that the IB4-SAP-treated cells exhibited a staining pattern similar to that of the IB4-negative human cells. Extended cultivation (more than 6 months) of these IB4-SAP-treated cells did not alter the above staining pattern. RT-PCR analysis revealed the presence of EndoGalC mRNA in these cells. Conclusions: This IB4-SAP-mediated method of selection of αgal epitope-negative cells will provide an alternative to the present method of cytotoxicity-based selection using specific antibody and complement.

Attempts to image reporter gene expression driven by weak promoters are often hampered by the poor transcriptional activity of such promoters. Most tissue-specific promoters are weak compared with stronger but constitutively expressing viral promoters. In this study, we validated methods of enhancing the transcriptional activity of weak promoters using a Cre-loxP system in vitro and in vivo. We constructed a tester vector, pCTL, which carries a strong systemic cytomegalovirus enhancer/chicken beta-actin promoter (CAG), loxP-flanked CAT, and firefly luciferase (luc) cDNAs. Herpes simplex virus-thymidine kinase (HSV-tk) promoter was used as a weak and systemic promoter and ligated to Cre for construction of pTC. Luc activity was higher (about 10-fold enhancement) in co-transfected (with pCTL and pTC) than in singly (with HSV-tk promoter-driven luc expression vector pTL) transfected NIH3T3 cells. In vivo electroporation-mediated gene delivery of both pCTL and pTC into murine oviductal epithelium yielded results (about 16-fold enhancement) similar to those obtained with in vitro-transfected NIH3T3 cells. To evaluate tissue-specific enhancement of gene expression, podocyte (glomerular visceral epithelial cell)-specific nephrin promoter was ligated to the Cre gene or luc cDNA to create pNC and pNL, respectively. We achieved 2.4-fold improvement of luc gene expression in the mouse kidney in vivo when pCTL and pNC were co-transfected via the tail vein via the lipoplex method. The combination of a weak tissue-specific promoter with the Cre-loxP system could thus be used to enhance the strength of tissue-specific promoters in vitro and in vivo.

The availability of techniques to create desired genetic mutations has enabled the laboratory mouse as an extensively used model organism in biomedical research including human genetics. A new addition to this existing technical repertoire is the CRISPR/Cas system. Specifically, this system allows editing of the mouse genome much more quickly than the previously used techniques, and, more importantly, multiple mutations can be created in a single experiment. Here we provide protocols for preparation of CRISPR/Cas reagents and microinjection into one-cell mouse embryos to create knockout or knock-in mouse models.