The clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas) system is an adaptive immune system in bacteria and archaea that has recently been exploited for genome engineering. Mutant mice can be generated in one step through direct delivery of the CRISPR/Cas9 components into a mouse zygote. Although the technology is robust, delivery remains a bottleneck, as it involves manual injection of the components into the pronuclei or the cytoplasm of mouse zygotes, which is technically demanding and inherently low throughput. To overcome this limitation, we employed electroporation as a means to deliver the CRISPR/Cas9 components, including Cas9 messenger RNA, single-guide RNA, and donor oligonucleotide, into mouse zygotes and recovered live mice with targeted nonhomologous end joining and homology-directed repair mutations with high efficiency. Our results demonstrate that mice carrying CRISPR/Cas9-mediated targeted mutations can be obtained with high efficiency by zygote electroporation.

Previously we established Zygote Electroporation of Nucleases (ZEN) technology as an efficient and high-throughput way to generate genetically modified mouse models. However, there were significant variations of the targeting efficiency among different genomic loci using our previously published protocol. In this study, we improved the ZEN technology by delivering Cas9 protein into mouse zygotes through a series of electroporation. Using this approach, we were able to introduce precise nucleotide substitutions, large segment deletion and short segment insertion into targeted loci with high efficiency.

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Genetic modification of almost any species is now possible using approaches based on targeted nucleases. These novel tools now bypass previous limited species windows, allowing precision nucleotide modification of the genome at high efficiency, rapidly and economically. Here we focus on the modification of the mouse genome; the mouse, with its short generation time and comparatively low maintenance/production costs is the perfect mammal with which to probe the genome to understand its functions and complexities. Further, using targeted nucleases combined with homologous recombination, it is now possible to precisely tailor the genome, creating models of human diseases and conditions directly and efficiently in zygotes derived from any mouse strain. Combined these approaches make it possible to sequentially and progressively refine mouse models to better reflect human disease, test and develop therapeutics. Here, we briefly review the strategies involved in designing targeted nucleases (sgRNAs) providing solutions and outlining in detail the practical processes involved in precision targeting and modification of the mouse genome and the establishing of new precision genetically modified mouse lines.

The CRISPR-Cas9 system in bacteria and archaea has recently been exploited for genome editing in various model organisms, including mice. The CRISPR-Cas9 reagents can be delivered directly into the mouse zygote to derive a mutant animal carrying targeted genetic modifications. The major components of the system include the guide RNA which provides target specificity, the Cas9 nuclease that creates the DNA double-strand break, and the donor oligonucleotide or plasmid carrying the intended mutation flanked by sequences homologous to the target site. Here we describe the general considerations and experimental protocols for creating genetically modified mice using the CRISPR-Cas9 system.

Purpose The purposes of this study are to characterize magneto-endosymbiont (ME) labeling of mammalian cells and to discern the subcellular fate of these living contrast agents. MEs are novel magnetic resonance imaging (MRI) contrast agents that are being used for cell tracking studies. Understanding the fate of MEs in host cells is valuable for designing in vivo cell tracking experiments. Procedures The ME's surface epitopes, contrast-producing paramagnetic magnetosomal iron, and genome were studied using immunocytochemistry (ICC), Fe and MRI contrast measurements, and quantitative polymerase chain reaction (qPCR), respectively. These assays, coupled with other common assays, enabled validation of ME cell labeling and dissection of ME subcellular processing. Results The assays mentioned above provide qualitative and quantitative assessments of cell labeling, the subcellular localization and the fate of MEs. ICC results, with an ME-specific antibody, qualitatively shows homogenous labeling with MEs. The ferrozine assay shows that MEs have an average of 7 fg Fe/ME, ∼30 % of which contributes to MRI contrast and ME-labeled MDA-MB-231 (MDA-231) cells generally have 2.4 pg Fe/cell, implying ∼350 MEs/cell. Adjusting the concentration of Fe in the ME growth media reduces the concentration of non-MRI contrast-producing Fe. Results from the qPCR assay, which quantifies ME genomes in labeled cells, shows that processing of MEs begins within 24 h in MDA-231 cells. ICC results suggest this intracellular digestion of MEs occurs by the lysosomal degradation pathway. MEs coated with listeriolysin O (LLO) are able to escape the primary phagosome, but subsequently co-localize with LC3, an autophagy-associated molecule, and are processed for digestion. In embryos, where autophagy is transiently suppressed, MEs show an increased capacity for survival and even replication. Finally, transmission electron microscopy (TEM) of ME-labeled MDA-231 cells confirms that the magnetosomes (the MRI contrast-producing particles) remain intact and enable in vivo cell tracking. Conclusions MEs are used to label mammalian cells for the purpose of cell tracking in vivo, with MRI. Various assays described herein (ICC, ferrozine, and qPCR) allow qualitative and quantitative assessments of labeling efficiency and provide a detailed understanding of subcellular processing of MEs. In some cell types, MEs are digested, but the MRI-producing particles remain. Coating with LLO allows MEs to escape the primary phagosome, enhances retention slightly, and confirms that MEs are ultimately processed by autophagy. Numerous intracellular bacteria and all endosymbiotically derived organelles have evolved molecular mechanisms to avoid intracellular clearance, and identification of the specific processes involved in ME clearance provides a framework on which to develop MEs with enhanced retention in mammalian cells.

There is a continual need to improve efficiency in creating precise genetic modifications in mice using embryonic stem cells (ESCs). We describe a novel approach resulting in 100% germline transmission from competent injected ESCs. We developed an F1 mouse host embryo (Perfect Host, PH) that selectively ablates its own germ cells via tissue-specific induction of diphtheria toxin. This approach allows competent microinjected ESCs to fully dominate the germline, eliminating competition for this critical niche in the developing and adult animal. This is in contrast to conventional methods, where competition from host germ cells results in offspring derived from host cells and ESCs, necessitating extensive breeding of chimeras and genotyping to identify germline. The germline transmission process is also complicated by variability in the actual number of ESCs that colonize the germline niche and the proportion that are germline competent. To validate the PH approach we used ESC lines derived from 129 F1, BALB/cByJ, and BTBR backgrounds as well as an iPS line. Resulting chimeric males produced 194 offspring, all paternally derived from the introduced stem cells, with no offspring being derived from the host genome. We further tested this approach using eleven genetically modified C57BL/6N ESC lines (International Knockout Mouse Consortium). ESC germline transmission was observed in 9/11 (82%) lines using PH blastocysts, compared to 6/11 (55%) when conventional host blastocysts were used. Furthermore, less than 35% (83/240) of mice born in the first litters from conventional chimeras were confirmed to be of ESC-origin. By comparison, 100% (137/137) of the first litter offspring of PH chimeras were confirmed as ESC-derived. Together, these data demonstrate that the PH approach increases the probability of germline transmission and speeds the generation of ESC derived animals from chimeras. Collectively, this approach reduces the time and costs inherent in the production of genetically modified animals.

Genome editing using the CRISPR/Cas9 RNA guided endonuclease system has rapidly become a driving force for discovery in modern biomedical research. This simple yet elegant system has been widely used to generate both loss-of-function alleles and precision knock-in mutations using single stranded donor oligonucleotides. Our CRISPRtools platform supports both of these applications in order to facilitate the use of CRISPR/Cas9. While there are several tools that facilitate CRISPR/Cas9 design and screen for potential off-target sites, the process is typically performed sequentially on single genes, limiting scalability for large-scale programs. Here, the design principle underlying gene ablation is based upon using paired guides flanking a critical region/exon of interest to create deletions. Guide pairs are rank ordered based upon published efficiency scores and off-target analyses, and reported in a concise format for downstream implementation. The exon deletion strategy simplifies characterization of founder animals and is the strategy employed for the majority of knockouts in the mouse. In proof-of-principle experiments, the effectiveness of this approach is demonstrated using microinjection and electroporation to introduce CRISPR/Cas9 components into mouse zygotes to delete critical exons.