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 faster 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.

Microinjection is considered the gold standard technique for delivery of nucleic acids (NAs; transgenes or genome editing tools such as CRISPR/Cas9 systems) into embryos, for creating genetically modified organisms. It requires sophisticated equipment as wel as well-trained and highly skilled personnel to perform the micro-injection technique. Here, we describe a novel and simple microinjection-independent technique, called Genome-editing via Oviductal Nucleic Acids Delivery (GONAD). Using GONAD, we show that NAs (e.g., eGFP mRNA or Cas9 mRNA/sgRNAs) can be effectively delivered to pre-implantation embryos within the intact mouse oviduct by a simple electroporation method, and result in the desired genetic modification in the embryos. Thus GONAD can bypass many complex steps in transgenic technology such as isolation of zygotes, microinjection of NAs into them, and their subsequent transfer to pseudo-pregnant animals. Furthermore, this method can potentially be used for genome editing in species other than mice.

Genome editing using clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) predominantly induces non-homologous end joining (NHEJ), which generates random insertions or deletions, whereas homology-directed repair (HDR), which generates precise recombination products, is useful for wider applications. However, the factors that determine the ratio of HDR to NHEJ products after CRISPR/Cas9 editing remain unclear, and methods by which the proportion of HDR products can be increased have not yet been fully established. We systematically analyzed the HDR and NHEJ products after genome editing using various modified guide RNAs (gRNAs) and Cas9 variants with an enhanced conformational checkpoint to improve the fidelity at endogenous gene loci in HEK293T cells and HeLa cells. We found that these modified gRNAs and Cas9 variants were able to enhance HDR in both single-nucleotide substitutions and a multi-kb DNA fragment insertion. Our results suggest that the original CRISPR/Cas9 system from the bacterial immune system is not necessarily the best option for the induction of HDR in genome editing and indicate that the modulation of the kinetics of conformational checkpoints of Cas9 can optimize the HDR/NHEJ ratio.

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.

Microphthalmia is a severe ocular disorder, and this condition is typically caused by mutations in transcription factors that are involved in eye development. Mice carrying mutations in these transcription factors would be useful tools for defining the mechanisms underlying developmental eye disorders. We discovered a new spontaneous recessive microphthalmos mouse mutant in the Japanese wild-derived inbred strain KOR1/Stm. The homozygous mutant mice were histologically characterized as microphthalmic by the absence of crystallin in the lens, a condition referred to as aphakia. By positional cloning, we identified the nonsense mutation c.444C>A outside the genomic region that encodes the homeodomain of the paired-like homeodomain transcription factor 3 gene (Pitx3) as the mutation responsible for the microphthalmia and aphakia. We examined Pitx3 mRNA expression of mutant mice during embryonic stages using RT-PCR and found that the expression levels are higher than in wild-type mice. Pitx3 over-expression in the lens during developmental stages was also confirmed at the protein level in the microphthalmos mutants via immunohistochemical analyses. Although lens fiber differentiation was not observed in the mutants, strong PITX3 protein signals were observed in the lens vesicles of the mutant lens. Thus, we speculated that abnormal PITX3, which lacks the C-terminus (including the OAR domain) as a result of the nonsense mutation, is expressed in mutant lenses. We showed that the expression of the downstream genes Foxe3, Prox1, and Mip was altered because of the Pitx3 mutation, with large reductions in the lens vesicles in the mutants. Similar profiles were observed by immunohistochemical analysis of these proteins. The expression profiles of crystallins were also altered in the mutants. Therefore, we speculated that the microphthalmos/aphakia in this mutant is caused by the expression of truncated PITX3, resulting in the abnormal expression of downstream targets and lens fiber proteins.

The Rinshoken cataract (rct) mutation, which causes congenital cataracts, is a recessive mutation found in SJL/J mice. All mutants present with opacity in the lens by 2 months of age. The rct locus was mapped to a 1.6-Mb region in Chr 4 that contains the Foxe3 gene. This gene is responsible for cataracts in humans and mice, and it plays a crucial role in the development of the lens. Furthermore, mutation of Foxe3 causes various ocular defects. We sequenced the genomic region of Foxe3, including the coding exons and UTRs; however, no mutations were discovered in these regions. Because there were no differences in Foxe3 sequences between the rct/rct and wild-type mice, we inferred that a mutation was located in the regulatory regions of the Foxe3 gene. To test this possibility, we sequenced a 5' noncoding region that is highly conserved among vertebrates and is predicted to be the major enhancer of Foxe3. This analysis revealed a deletion of 22-bp located approximately 3.2-kb upstream of the start codon of Foxe3 in rct mice. Moreover, we demonstrated by RT-PCR and in situ hybridization that the rct mutant has reduced expression of Foxe3 in the lens during development. We therefore suggest that cataracts in rct mice are caused by reduced Foxe3 expression in the lens and that this decreased expression is a result of a deletion in a cis-acting regulatory element.

The excision of genomic sequences using paired CRISPR-Cas nucleases is a powerful tool to study gene function, create disease models and holds promise for therapeutic gene editing. However, our understanding of the factors that favor efficient excision is limited by the lack of a rapid, accurate measurement of DNA excision outcomes that is free of amplification bias. Here, we introduce ddXR (droplet digital PCR eXcision Reporter), a method that enables the accurate and sensitive detection of excisions and inversions independent of length. The method can be completed in a few hours without the need for next-generation sequencing. The ddXR method uncovered unexpectedly high rates of large (> 20 kb) excisions and inversions, while also revealing a surprisingly low dependence on linear distance, up to 170 kb. We further modified the method to measure precise repair of excision junctions and allele-specific excision, with important implications for disease modeling and therapeutic gene editing.

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