The emergence of CRISPR/Cas9 gene editing technology with its simplicity, versatility, and efficiency has considerably improved the time frame and the process of creating these modified alleles. The technology has enabled rapid production of knockout, conditional alleles or mice carrying single point mutations, which mimic those in human patients, in only several weeks. For the generation of knockout alleles, the microinjection of a single guide RNA (sgRNA) in zygotes is sufficient to create indels in a critical exon, inducing a frameshift mutation and therefore functionally abolishing the gene of interest [4]. However, the generation of more complex alleles, including conditionals, is more challenging and requires at least two sgRNAs and a repair template in the form of two short single-stranded DNA (ssDNA) repair templates each containing a loxP site [5]. The repair template replaces the endogenous locus of interest by homology-directed repair after Cas9 cleavage and the double-strand break (DSB) of the region of interest. Unfortunately, the process of generating these conditional alleles using programmable nucleases remains lengthy and relatively inefficient, as it requires microinjection of the DNA and the correct insertion of multiple repair templates into the same allele of the mouse genome, without recombination or mutation within the repair template or the exon of interest.

A first research article from Quadros and colleagues aimed to improve the generation of conditional alleles in mice using programmable nucleases [2]. The authors made the simple observation that since the efficiency to repair DNA after DSB is higher for the homology-directed repair pathway than homologous recombination, the delivery of a longer repair template would result in a higher efficiency for generating mutant alleles. This technique, called efficient addition with ssDNA inserts-CRISPR (Easi-CRISPR), involves targeting by two sgRNAs which flank the endogenous exon and are complexed with Cas9 to form a ribonucleoprotein complex for cellular delivery. The exon is replaced after the DSB of the DNA and repaired with a long-stranded oligonucleotide template containing two loxP sites and spanning the entire exon. The authors demonstrated the power of this approach by showing a high efficiency in editing and allelic replacement, averaging a 50% success rate, and up to 100% editing for certain alleles, which is a marked improvement compared with conventional methods. Future work and replication studies from various research groups and transgenesis core facilities will confirm or disprove these observations. However, while efficient, this technique does not solve the limiting issues of CRISPR/Cas9, such as the requirement for highly trained staff and the use of an expensive microinjection apparatus only available in transgenesis core facilities.

A second research paper considerably simplified the procedure of generating complex alleles using CRISPR/Cas9 technology [3]. Ohtsuka and colleagues reasoned that direct delivery of the CRISPR/Cas9 reagents into the mouse oviduct would be as efficient, and more effective, than microinjection and surgical transfer of zygotes [3]. This technique would bypass all lengthy and complex procedures, from the isolation of zygotes from the oviduct, to mouse embryo transfer into recipient females. The other advantage of this method would be the reduction of the number of animals required to generate a knockout or knockin allele, since it is no longer necessary to sacrifice females for zygote collection, and the authors show that the recipient females are able to be impregnated again after their first transgenic litter.

Ohtsuka and colleagues [3] postulated that in situ delivery of the CRISPR/Cas9 reagents to the mouse oviduct by electroporation would allow Cas9 protein access to the zygote DNA to edit the genome. To ascertain this hypothesis, Ohtsuka and colleagues optimized the delivery protocol of CRISPR/Cas9 reagents to the oviduct and determined the optimum editing efficiency was at 0.7 dpc. The authors then hypothesized that an in vivo delivery of the CRISPR/Cas9 reagents combined with an Easi-CRISPR approach would display a similar, if not better, efficiency of editing simple or complex alleles when compared with microinjection into mouse zygotes. They tested this approach, called improved-genome editing via oviductal nucleic acid delivery (i-GONAD) (Fig. 1b), and after optimization the success of editing observed was up to 97% for straight knockout alleles and 50% for gene tags, which gives similar results to microinjected zygotes [3]. Importantly, Ohtsuka and colleagues demonstrated the feasibility of this approach targeting various genes in multiple mouse strains. Interestingly, the recipient females could be used for multiple experiments, suggesting that it is possible to considerably reduce the number of mice needed to generate these modified alleles. Whereas the efficiency of the i-GONAD approach to generate complex alleles using CRISPR/Cas9 genome editing technologies seems remarkable and promising, the frequency of mosaicism appears to remain up to 30% of edited alleles. Future optimization of this technique, combined with replication studies from various research groups, will enable improvements to the technology, address technical hurdles, and hopefully enable the successful and efficient generation of conditional alleles using the i-GONAD technique.

CRISPR/Cas9 gene editing technology has considerably changed transgenesis technologies. In the last 3–5 years, at a rapid pace, remarkable achievements have been observed. Easi-CRISPR and i-GONAD techniques have the potential to entirely reshape the traditional route of generating modified alleles in mice if the techniques are widely adopted by many research groups and transgenesis core facilities. It is predictable that soon all conventional steps to efficiently generate knockout or knockin alleles in mice will be bypassed and the CRISPR/Cas9 reagents will be delivered in situ into the oviduct. It will require less highly skilled personnel or specialized equipment since a stereomicroscope and an electroporation device would suffice to generate editing for simple or complex alleles. Importantly, it will enable the reduction of animal usage inline with the 3Rs rule for animal work. I would predict the recent technological development in gene editing and assisted reproduction will redefine decades of transgenesis. The future will tell what the pace of these changes will be.