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CRISPR-based genome engineering has been widely used for producing gene-modified animals such as mice and rats, to explore the function of a gene of interest and to create disease models. However, it always requires the ex vivo handling of preimplantation embryos, as exemplified by the microinjection of genome editing components into zygotes or in vitro electroporation of zygotes in the presence of genome editing components, and subsequent cultivation of the treated embryos prior to egg transfer to the recipient females. To avoid this ex vivo process, we have developed a novel method called genome-editing via oviductal nucleic acids delivery (GONAD) or improved GONAD (i-GONAD), which enables in situ genome editing of zygotes present in the oviductal lumen of a pregnant female. This technology does not require any ex vivo handling of preimplantation embryos or preparation of recipient females and vasectomized males, all of which are often laborious and time-consuming. In this chapter, recent advances in the development of GONAD/i-GONAD will be described.
Department of Genome Medicine, National Center for Child Health and Development, Japan
Masato Ohtsuka
Department of Molecular Life Science, Division of Basic Medical Science and Molecular Medicine, Tokai University School of Medicine, Japan
Center for Matrix Biology and Medicine, Graduate School of Medicine, Tokai University, Japan
The Institute of Medical Sciences, Tokai University, Japan
Emi Inada
Department of Pediatric Dentistry, Graduate School of Medical and Dental Sciences, Kagoshima University, Japan
Shingo Nakamura
Division of Biomedical Engineering, National Defense Medical College Research Institute, Japan
Issei Saitoh
Department of Pediatric Dentistry, Asahi University School of Dentistry, Japan
Shuji Takabayashi
Laboratory Animal Facilities and Services, Preeminent Medical Photonics Education and Research Center, Hamamatsu University School of Medicine, Japan
*Address all correspondence to: sato-masa@ncchd.go.jp
1. Introduction
In mammals, embryogenesis begins when the oocytes (ovulated from an ovary of a female) fertilize with spermatozoa in the uterus of the female (Figure 1). The fertilized oocytes, called zygotes or 1-cell stage embryos, exist at the “ampulla,” an area of the oviduct located near an ovary. In mice, early zygotes are surrounded by cumulus cells (also called follicular cells) and correspond to Day 0.4 of gestation (11:00 AM in the morning after mating with a male) (box in Figure 1). In this case, Day 0 of gestation is defined as the day when the copulation plug is recognized in the morning. Late zygotes corresponding to Day 0.7 of gestation (16:00 PM) exhibit dissociation of cumulus cells (Figure 1). Then, these zygotes develop into 2-cell (~Day 1.4 of gestation), 4-cell (~Day 1.8 of gestation), 8-cell (~Day 2.4 of gestation), 16-cell (also called morula; ~Day 2.7 of gestation), early blastocyst (~Day 3.4 of gestation), and late blastocysts (~Day 4.4 of gestation) (at which stage implantation into the uterine epithelium starts). Notably, zygotes and 2-cell embryos exist in the ampulla, 4-cell to 8-cell embryos in the oviductal portion between the ampulla and isthmus, 16-cell embryos (morulae) in both the oviduct and uterus, and early- to late-blastocysts in the uterus (Figure 1).
Figure 1.
Schematic of preimplantation (Days 0.5 to 4.5; Day 0 of pregnancy is defined as the day a vaginal plug is found) and postimplantation (Days 5.5) development of mice. During preimplantation, early zygote (early 1-cell embryo) (at Day 0.4; see box), late zygote (late 1-cell embryo) (Day 0.7), 2-cell embryo (day 1.5), 8- to 16-cell embryo (Day 2.5), early blastocyst (Day 3.5), and late blastocyst (Day 4.5) float in the oviductal lumen or uterine horn. Embryos at days 0.5 to 4.5 have zona pellucida (ZP), but embryos at day 4.5 begin to escape from ZP, which is called “ZP hatching,” and are ready to implant into the uterine epithelium. Note that at early zygote stage (see box), fertilized eggs are tightly surrounded by cumulus cells, but at late zygote stage, cumulus cells begin to detach from an embryo. This figure was drawn in-house and reproduced with permission from Sato et al., “Recent Advances and Future Perspectives of In Vivo Targeted Delivery of Genome-Editing Reagents to Germ cells, Embryos, and Fetuses in Mice”; published by MDPI, 2020.
For producing genetically engineered animals through pronuclear microinjection (MI) or viral infection, early zygotes are generally used [1]. To isolate early zygotes, oviducts dissected from pregnant female mice at ~Day 0.4 of pregnancy are teased using a needle at the ampulla under paraffin oil. The exposed cumulus-oocyte-complex is then transferred to a drop of hyaluronidase (HA)-containing medium. Brief incubation (~5 min at room temperature) relieves cumulus cells. The resulting “denuded” zygotes are always used for MI or viral infection.
Historically, the first attempt to obtain transgenic (Tg) mice was performed by microinjecting SV40 viral DNA into the blastocoel cavity of blastocysts [2]. The resulting offspring had various levels of SV40 genome in their organs. In 1980, Gordon et al. [3] first produced germ-line Tg mice through MI of exogenous DNA. Since then, successful production of Tg rabbits, sheep, and pigs was reported by Hammer et al. [4]. This MI technique relies on the physical injection of any type of nucleic acids (NAs) (i.e., purified DNA fragments of 3–4 kb carrying an expression unit) using an expensive manipulator and requires personnel with specific skill. Furthermore, it generally takes 3–4 h to finish MI using ~100 zygotes per session (Figure 2A). Perry et al. [5] reported a novel technique to generate Tg mice through the intracytoplasmic injection of a sperm (which has been mixed with NAs) into a zygote. Later, this technique was recognized as a useful tool to introduce large sized DNA such as bacterial artificial chromosomes (BACs) and yeast artificial chromosomes (YACs) into the mammalian genome [6, 7, 8].
Figure 2.
Schematic of genome-edited mouse production through microinjection (MI) (A), in vitro electroporation (EP) (B), or genome-editing via oviductal nucleic acids delivery (GONAD) (or improved GONAD (i-GONAD)) (C). This figure was drawn in-house, and reproduced with permission from Sato et al., “Recent Advances in the Production of Genome-Edited Rats”; published by MDPI, 2022.
Mammalian zygotes, including those of mice, are surrounded by a translucent glycoprotein layer called zona pellucida (ZP), which exists as a barrier that protects the early embryo from environmental insults, including viral infections, and injury from chemical or physical substances [9, 10]. For example, ZP-enclosed embryos could not be transduced through simple incubation with solution containing lentivirus (LV), adenovirus (AV), or retrovirus (RV) [11, 12, 13]. However, injection of those viruses into the space between ZP and zygote (called “peri-vitelline space”) results in successful transduction [14, 15]. Furthermore, transduction of mouse zygotes was possible when they were subjected to laser perforation of ZP in the presence of LV vectors [16]. ZP can be removed by incubating ZP-enclosed embryos in the presence of proteolytic enzyme such as pronase, or under acidic conditions using acidic Tyrode’s solution [1]; therefore, gene delivery can be accomplished by incubating the ZP-removed embryos in a solution containing liposome-complexed DNA [17] or the above-mentioned viruses [11, 18]. However, these treated embryos are often vulnerable, adhesive, and are easily damaged [19]. Furthermore, transferring the ZP-removed embryos (at least up to morulae) into the oviductal lumen of pseudopregnant recipient females cannot support their normal development, because the transferred embryos tend to adhere to the oviductal epithelium [20, 21]. To avoid this, the treated embryos have to be cultured at least up to blastocysts (showing reduced adhesive property), prior to uterine transfer. Notably, acquisition of Tg founders was reported using ZP-free embryos transfected liposomally [22] or when transduced with AV vectors [11, 18].
Electroporation (EP)-mediated gene delivery is also a method that efficiently introduces exogenous NAs into a cell or zygote through electric shock-induced, transient micropores in a cell membrane [23]. It requires an expensive electroporator, but does not require a skilled person, unlike in the case of MI (Figure 2B). In an early study regarding EP-based introduction of DNA into mouse preimplantation embryos, DNA was first injected into the peri-vitelline space of zygotes, and then the embryos were subjected to in vitro EP. Unfortunately, the transfection efficiency was very low [24]. In 2002, Grabarek et al. [25] first demonstrated that in vitro EP enabled incorporation of small-sized NAs (i.e., siRNA) into mouse zygotes. In this case, prior to EP, ZP has to be weakened by a brief treatment with acidic Tyrode’s solution to facilitate transfer of NAs into an embryo and to protect the embryos from EP damage. Notably, in vitro EP was also successfully used to deliver double-stranded RNA (dsRNA) and morpholinos into mouse preimplantation embryos [26, 27].
To our knowledge, Peng et al. [27] first demonstrated that plasmid DNA can be effectively delivered into mouse preimplantation embryos when they were subjected to in vitro EP using optimal EP parameters (i.e., voltage, pulse duration, number of pulses, and repeats). Sato et al. [28] also demonstrated that plasmid DNA can be successfully introduced into early mouse embryos present within the oviductal lumen through in vivo EP. In vitro EP-based gene delivery is generally possible using over 100 zygotes per a trial and can be finished within 15–30 min. Thus, in terms of convenience, EP appears superior to the MI-based production of transgenics. However, this success appears largely to depend on the EP parameters and the type of electroporator used, as mentioned below.
Beside in vitro EP, MI, and viral transduction, substances capable of penetrating ZP (also called “ZP-penetrating reagents”) can be used with NAs to perform gene delivery towards ZP-enclosed embryos. For examples, Ivanova et al. [29] employed a receptor-mediated gene transfer system, with insulin as the admission ligand in the DNA-carrying construct, because early embryonic cells are known to have internalizable insulin receptors on their surface [30]. They first made an insulin-polylysine conjugated with plasmid DNA. Next, this complex was mixed with a conjugate consisting of streptavidin-polylysine-biotinylated adenovirus. Short (3 h) incubation of ZP-enclosed mouse and rabbit preimplantation embryos with the resulting complex [called “(insulin-polylysine)-DNA and (insulinpolylysine)-DNA-(streptavidin-polylysine)-(biotinylated adenovirus)”] penetrated the constructs through ZP and accumulated in the peri-nuclear space of the embryos, leading to ligand/receptor-mediated transgenesis. Joo et al. [31] developed amphiphilic chitosan-based nanocarriers, called VisuFect. When murine zygotes were incubated with a solution containing Cy5.5-labeled VisuFect conjugated with poly(A) oligonucleotides, the complex gradually penetrated the cytoplasm of the ZP-enclosed zygotes. This suggests that VisuFect could be used as a vehicle to deliver NAs to ZP-intact embryos. According to Joo et al. [31], VisuFect can deliver siRNA, but not large molecules such a plasmid DNA to embryos. Nanoparticles are also a promising tool to transfer exogenous NAs into ZP-enclosed embryos. Munk et al. [32] demonstrated that multiwall carbon nanotubes (MWNTs) can cross the ZP to help the delivery of plasmid DNA (carrying the green fluorescent protein (GFP) gene) into bovine embryos in vitro. According to Munk et al. [32], MWNTs themselves are non-harmful to embryos and do not affect their viability and gene expression. On the other hand, Jin et al. [33] first demonstrated that peptide nanoparticles can introduce siRNA into an intact mouse oocyte. When oocytes were incubated with peptide nanoparticle-complexed fluorescein isothiocyanate (FITC)-conjugated siRNA for 12–14 h, a cytoplasmic fluorescence of oocytes was observed together with a target gene knockdown. Jin et al. [33] suggested that peptide nanoparticle-mediated siRNA transfection was useful to explore the function of unknown genes in mouse oocytes. Unfortunately, except for the report by Ivanova et al. [29], these studies do not show germ-line transmission or chromosomal integration of transgenes.
As mentioned above, to obtain genetically-modified (GM) animals, isolation of zygotes from pregnant females or those obtained through in vitro fertilization (IVF), in vitro gene delivery towards the isolated embryos, transient cultivation of the treated embryos, and egg transfer (ET) of the cultivated embryos to the reproductive tracts of pseudopregnant recipient females to allow the GM embryos to develop in vivo further. These processes are called “ex vivo handling of embryos” (Figure 2A and B), and generally required for MI, in vitro EP, liposome- or viral transduction-based transgenesis. Notably, this ex vivo handling of embryos is costive, labor intensive, and laborious, because it requires preparation of sterile males called “vasectomized males” to create pseudopregnant females, timely supply of those pseudopregnant females, and ET technique, which is a more difficult task requiring people with specialized skill sets. To bypass this process, direct genetic manipulation must be performed in zygotes (or embryos at more advanced stages) existing within an oviductal lumen of a pregnant female. Relloso and Esponda [34, 35] first attempted to transfect epithelial cells lining oviductal lumen by injecting liposomally encapsulated DNA directly into the oviductal lumen of a female mouse. They found that 6% of oviductal epithelial cells were successfully transfected. Rios et al. [36] also demonstrated that intraoviductal injection of naked DNA or mRNA into the estradiol-treated female rats can help incorporate those substances into the oviductal cells. The introduced DNA or mRNA will then be translated into an active protein, possibly accelerating embryo transport.
Sato [37] employed in vivo EP to enhance transfection efficiency in the oviductal epithelium. In vivo EP is a method to transfect the tissue or organ in situ by injecting a fluid containing plasmid DNA into the target site, holding the injection site with tweezer-type electrodes, and subsequently giving an electric shock using an electroporator [38]. Using this system, several organs/tissues including kidney [39, 40], liver [41], brain [42], skin [43, 44], skeletal muscle [45], testis [46, 47], efferent duct [48], ovary [49], and fetuses [50, 51, 52], have been successfully transfected. According to Sato [37], ovary/oviduct/uterus were pulled out and exposed on the back of a female on Day 0.4 of pregnancy (~11:00 AM; corresponding to early zygotes). Then, a small amount (1–2 μL) of solution which contains an enhanced green fluorescent protein (eGFP) expression plasmid (0.5 μg/μL) and 0.2% (v/v) trypan blue (TB) (used as a marker for visualizing injected materials) was injected into the oviductal lumen using a mouthpiece-controlled glass pipette under a dissecting microscope. Immediately after injection, an entire oviduct was subjected to in vivo EP using tweezer-type electrodes in an attempt to transfect oviductal epithelium facing oviductal lumen and possibly zygotes (floating in the oviductal lumen) with the exogenous DNA. The EP condition was eight square-wave pulses with a pulse duration of 5 ms and an electric field intensity of 50 V, generated by a square-wave pulse generator (#T-820; BTX Genetronics Inc., San Diego, CA, USA). After EP, the treated oviduct was returned to the original position. One day after the surgery, oviducts were dissected from the female to isolate 2-cell embryos. When the eGFP-derived fluorescence in isolated embryos and oviducts was inspected, maximal 43% of oviductal epithelial cells facing oviductal lumen were fluorescent, while no fluorescence was discernible in the isolated embryos; only a cellular remnant probably derived from a part of zygotes was found to be fluorescent. Based on this finding, Sato [37] speculated that failure of gene delivery to zygotes may be due to the cumulus cells surrounding zygotes acting as a barrier. Because the oviductal epithelium can be efficiently transfected using in vivo EP method, Sato [37] named this technology “gene transfer via oviductal epithelium (GTOVE).”
Sato et al. [28] next attempted to transfect 2-cell embryos floating in the oviductal lumen using GTOVE, since those embryos are already free of cumulus cells. To address this issue, GTOVE was performed in pregnant females at Day 1.4 of pregnancy using the same conditions elaborated in the study by Sato [37]. One day after the GTOVE procedure, 8-cell embryos were collected from the GTOVE-treated females for checking eGFP-derived fluorescence. Of the 12 oviducts (6 females used) examined, 3 contained fluorescent 8-cell stage embryos (33%, 19/58 tested), but the intensity of fluorescence varied among the embryos. Unfortunately, gene expression was transient in this system, with no evidence for chromosomal integration of transgenes [28]. These results indicate that successful in vivo introduction of exogenous plasmid DNA into early mouse embryos is possible, as far as the T-820 electoporator is employed. However, the T-820 electoporator is currently unavailable. When we performed GTOVE to transfect 2-cell embryos using another electroporator (NEPA21; NEPA GENE Co., Chiba, Japan), the collected embryos failed to fluoresce [53]. This suggests a need to carefully examine the optimal EP condition enabling ZP penetration of larger sized molecules like plasmid DNA, as suggested by Peng et al. [27] and Hakim et al. [54].
Genome-editing technology includes zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (CRISPR/Cas9), all of which employ sequence-specific nucleases to induce modifications in a predefined region of the genome (reviewed by Harrison et al. [55]; Hsu et al. [56]). In the absence of the donor (or template) DNA [including longer genes (>1 kb), single-stranded (ss) sequences (>200 bp) or synthetic oligodeoxynucleotides (ODNs) 20–30 bp in size, all of which have sequences showing homology to the target sequence, these nucleases can induce double-strand breaks (DSBs) which are then repaired via nonhomologous end joining (NHEJ). This repair occurs through the cellular machinery, which frequently generates random insertions, deletions, or substitutions of nucleotides (called indels) at the break site. These indels often cause frameshift mutations, leading to the occasional failure in protein expression. In the presence of the donor (or template) DNA, it will be introduced into the DSB site through homology-directed repair (HDR), and this event is called “knock-in (KI).” According to Yoshimi et al. [57], KI is more difficult to complete successfully than inducing NHEJ-mediated indels. Furthermore, NHEJ occurs in nondividing and dividing cells, but HDR occurs preferentially in dividing cells [58, 59].
Unlike ZFN and TALEN, the CRISPR/Cas9 gene editing system has several advantages, including easy design for any genomic targets, simplicity, and the ability to modify several target genes simultaneously (multiplexing). Owing to these properties, the CRISPR/Cas9 system is now widely used in various biological systems. It employs only two components: (1) a guide RNA (gRNA), comprised either of a duplex CRISPR RNA (crRNA)/trans-activating CRISPR RNA (tracrRNA) molecule or of single-guide RNA (sgRNA), a fusion between crRNA and tracrRNA, and (2) a Cas9 endonuclease (reviewed by Harrison et al. [55]; Hsu et al. [56]). The gRNA can bind to the specific DNA sequence together with Cas9. Once bound, the Cas9 cleaves dsDNA 3 bp upstream of the protospacer adjacent motif (PAM, 5′-NGG-3′), which is recognized by the Cas9 protein. The cleaved site is then repaired by various cellular machineries, such as NHEJ and HDR.
3. Development of novel technologies enabling EP-based genome-editing in vitro and in vivo
As described in Section 2, in the late 2013, CRISPR/Cas9 system was recognized as a useful tool to manipulate a target gene in mammalian cells and embryos (reviewed by Harrison et al. [55]; Hsu et al. [56]). Since then, genome-edited animals from various species including mice, rats, pigs, bovines, and primates (monkeys) have been generated through MI [60, 61, 62, 63, 64, 65, 66]. Kaneko et al. [67] first demonstrated that in vitro EP of rat zygotes in the presence of genome editing reagents is a powerful tool to produce GM animals (Figure 2B). According to Kaneko et al. [67], intact rat zygotes were electroporated using an NEPA21 electroporator in a solution containing ZFN (40 μg/mL) mRNA [targeted to the rat interleukin 2 receptor subunit gamma gene (Il2rg)] under the EP condition of a poring pulse (Pp) (voltage: 225 V; pulse interval: 50 ms; pulse width: 1.5 and 2.5 ms; number of pulses: 4). The mechanism underlying gene delivery by this system is described in our previous paper [68]. As a result, they obtained genome-edited rat offspring with an efficiency of 73%, which is roughly 2-fold higher than that obtained through MI-mediated genome editing. Notably, they confirmed germ-line transmission of the genome-edited traits beyond next generation. This technology was thus named “technique for animal knockout system by electroporation (TAKE).” Since then, various genome-edited animals including mice [69, 70, 71, 72, 73, 74, 75], rats [76, 77, 78], and pigs [79, 80, 81] have been successfully generated using this technology. Furthermore, this technology has been applied to introduce nucleases into frozen embryos [82] and embryos derived from freeze-dried and frozen sperm [83].
Takahashi et al. [84] developed a novel in vivo EP-based method enabling in situ CRISPR/Cas9-based genome editing towards early mouse embryos (2-cell embryos) using a technique similar to GTOVE (Figure 2C). In this case, a solution (1–1.5 μL) containing Cas9 mRNA (up to 1.1 μg/μL), sgRNA (0.6 μg/μL; targeted to eGFP cDNA), and 0.2% (v/v) TB was injected through the oviductal wall under observation into the oviductal lumen of pregnant non-Tg females at Day 1.4 of pregnancy (corresponding to the 2-cell stage) that had been mated with Tg males containing an eGFP expression cassette in a homozygous manner [85]. In this case, the pregnant female should have zygotes, all of which are expected to have eGFP expression cassette in a heterozygous manner, and be fluorescent (Figure 3). If CRISPR/Cas9-induced mutations (indels) occur in the genomes of those zygotes (eGFP cDNA), some of their offspring will most likely lose fluorescence from their entire body. Fluorescence inspection of mid-gestational fetuses dissected from the in vivo EP-treated females demonstrated that out of 6 fetuses obtained, two lost fluorescence completely, two exhibited weak fluorescence, and fluorescence in the remaining two fetuses remained unaltered. Molecular biological analysis revealed that the former fetuses comprised knock out (KO) cells, the middle fetuses contained a mixture of KO cells and intact cells (whose state is called “mosaic mutation”), and the latter fetuses had unedited intact cells. Based on these findings, the KO efficiency of these fetuses can be estimated to be approximately 29%. Based on these findings, Takahashi et al. [84] named this technology “genome-editing via oviductal nucleic acids delivery (GONAD).”
Figure 3.
Experimental flowchart for genome-editing via oviductal nucleic acids delivery (GONAD). Females (C57BL/6) are first mated to C57BL/6-Tg (CAG-eGFP) male mice that possess EGFP transgenes in a homozygous (Tg/Tg) state. All the fetuses are then expected to be eGFP-expressing fetuses carrying the transgenes in a heterozygous (Tg/+) state. Thus, successful genome editing targeted to eGFP at preimplantation stages are expected to reduce the levels of eGFP fluorescence in the mid-gestational fetuses, as a result of genome editing in the chromosomally integrated eGFP transgenes. When GONAD was performed using Cas9 mRNA and sgRNA towards pregnant females at Day 1.4 of pregnancy, there were three types of fetuses (showing complete loss of fluorescence, partial fluorescence, or no reduction in fluorescence) in view of fluorescence expression pattern.
In 2018, Ohtsuka et al. [86] in the same group of Takahashi et al. [84] further elaborated on the GONAD technology. As was mentioned in Section 3, GONAD enables in situ genome editing in 2-cell embryos. In this case, only one blastomere among two blastomeres can be genome-edited, generating “mosaic” fetuses comprising edited and non-edited cells. To avoid this risk, genome editing at zygote (one-cell) stage is desirable. As was mentioned previously, early zygotes (corresponding to Day 0.4 of pregnancy; 11:00 AM) are tightly surrounded by cumulus cells. In our previous experience, GONAD at this stage failed, because almost all of the genome editing reagents introduced intaoviductally was trapped by the cumulus cells [86]. On the other hand, as mentioned previously, the detachment of cumulus cells from a zygote commences at late zygote stage (corresponding to Day 0.7 of pregnancy; 16:00 PM). Based on this finding, Ohtsuka et al. [86] first attempted to disrupt the forkhead box protein E3 (Foxe3) locus using ribonucleoprotein (RNP) (comprised of 1 μg/μL of Cas9 protein and 30 μM of crRNA/tracrRNA) using pregnant females at Day 0.7 of pregnancy. The advantage of using RNP is to induce genome editing more rapidly than using Cas9 mRNA [87]. In vivo EP was performed using the NEPA21 apparatus under the following conditions: Pp: 50 V, 5-ms pulse, 50-ms pulse interval, three pulses, 10% decay (± pulse orientation); Tp: 10 V, 50 ms pulse, 50 ms pulse interval, three pulses, and 40% decay (± pulse orientation). This modification resulted in 97% of the embryos exhibiting indels in the target locus. They next attempted to perform KI of a sequence (coding for a gene of interest (GOI)) into the target locus using RNP containing 1–2 μg/μL of ssODN or 0.85–1.4 μg/μL of ssDNA (with ~925 bases in size) generated through a novel method, called Easi-CRISPR, a highly efficient KI technique using ssDNA as donor templates [88, 89]. As a result, ~50% and ~ 15% of embryos were found to have KI alleles for ssODN and longer ssDNA in their genome, respectively.
Ohtsuka et al. [86] also demonstrated that large deletion (LD) of a target sequence can be accomplished using this modified GONAD, which was re-named as “i-GONAD.” It uses the Cas9 protein instead of the Cas9 mRNA, and targets late zygotes. For example, they designed two gRNAs (16.2 kb distance apart), both of which recognize either the sites of the retrotransposon sequence in the intron 1 of Agouti locus in the C57BL/6 mouse genome. The i-GONAD-mediated deletion of the inserted sequence resulted in the generation of fetuses with agouti coat color with efficiencies of 50%. Molecular biological analysis of these rescued offspring revealed the evidence for LD in the Agouti locus containing retrotransposon of C57BL/6 mice.
Since the studies by Takahashi et al. [84] and Ohtsuka et al. [86], several reports have been provided using GONAD/i-GONAD technologies. In Table 1, past studies using those technologies are listed. Also, the detailed protocols for i-GONAD in mice have been provided by Gurumurthy et al. [116, 117] and Ohtsuka and Sato [118]. The GONAD/i-GONAD-based production of genome-edited rats is also possible using the same approach shown in mice. Notably, Namba et al. [119] demonstrated the protocols for GONAD/i-GONAD in rats.
Type of method (content of CRISPR/Cas9 reagents)
Genome editing tool (mode for gene modification) EP apparatus
Animal (species strain)
Outcome
Target gene
References
GONAD (mRNA)
CRISPR/Cas9 (indels) BTX T820
Mouse (MCH(ICR), eGFP Tg)
Performed at Day 1.5 of pregnancy (corresponding to 2-cell stage) in mice; first successful genome editing at mid-gestational fetuses with 28% efficiency; also frequently associated with mosaic mutations
Performed at Day 0.7 of pregnancy (corresponding to late zygote stage); successful genome editing in offspring with efficiencies of 50 ~ 100% for indels, ~50% for ssODN KI (single-base changes) and 15% for longer ssDNA KI; kilobase-sized deletions can also be induced
Showing successful genome editing (with efficiencies of 50.0% and 17.8% for indels and 26.9% and 11.1% for KI) using i-GONAD in rats; the i-GONAD in rats was renamed as “rGONAD.”
Showing successful genome editing (with efficiencies of ~62% for indels and ~ 9% for KI) using i-GONAD in various rat strains; abnormal facial morphogenesis in fetuses was induced when Pax6 locus was targeted; strain-difference regarding the optimal in vivo EP condition was noted for BN strain
Intraoviductal injection of a solution containing two rAAV serotype 6 vectors, one carrying spCas9 gene and the other carrying gRNA expression unit, targeted Tyr into pregnant female mice at Day 0.5 of pregnancy; led to production of genome-edited pups with 10% efficiency
Showing the first successful target-specific chromosomal inversions of 7.67 megabases (Mb) in length in mice; this is longer than any previously reported inversion produced using PI-based methods.
A review showing that i-GONAD is useful for production of Tyr KO hamsters; the hamsters showed albino coat color, as expected (Hirose and Ogura, unpublished).
Showing the usefulness of two steps of i-GONAD (at Day 0.7 and Day 1.4–1.5 of pregnancy) to induce two mutations which are closely located each other; can create floxed mice carrying two lox sites flanking an exon; this approach is named “sequential i-GONAD (si-GONAD)”
For obtaining genome-edited C57BL/6 mice, setting a constant current of 100 mA upon in vivo EP is recommended; in this study, optimal EP conditions allowing the generation of a 100 mA current using two electroporators, NEPA21 and GEB15, are explored; consequently, the current and resistance were set to 40 V and 350–400 W, respectively, and were found to be suitable for i-GONAD using C57BL/6 mice.
Explored for optimal condition for obtaining genome-edited BN rats through i-GONAD; under a current of 100–300 mA using NEPA21, genome-edited BN rats were obtained with efficiencies of 75–100%; under a current of 150–200 mA using CUY21EDIT II genome-edited BN rats were obtained with efficiencies of 24–55%.
Showing successful production of floxed alleles for five genes with an efficiency of 10% through a single step under relatively low costs and minimal equipment setup; constitutive KO alleles were obtained as byproducts of these experiments.
Showing successful production of Klrc2 KO mice; Qa-1b expression levels were down-regulated in infected cells but increased in some bystander immune cells to respectively promote or inhibit their killing by activated natural killer cells.
Showing successful production of hemagglutinin (HA)-tag KI mice; HA-tag sequence was successfully inserted into the C terminus of the ATF5 coding sequence.
Showing successful production of Serpina3 KO mice which failed to evoke proper resolution, indicating that Serpina3n has a physiological function in resolving inflammation.
Showing successful production of Nrsn2 KO mice with reduced AMPAR signaling; Neurensin-2 was found to have a role as a novel modulator of emotional behavior.
Showing the first Tprkb KO mouse with an embryonic lethal mutation that was stably maintained in heterozygotes as inversion balancer strains using a B6.C3H-In(6)1 J inversion identified from C3H/HeJJcl.
Showing successful production of Mov10l1 KO hamster which is sterile in both sexes; this is in contrast with the case of Mov10l1 KO mice which were known to be sterile in males, but not in females.
Using △eGFP Tg mice carrying a single copy of disrupted eGFP (△eGFP), the feasibility of i-GONAD-mediated gene correction using AsCas12a nuclease is shown.
Showing successful production of Col4α5 KO rat model for Alport syndrome with hematuria, proteinuria, and high levels of BUN/Cre; died at 18 to 28 weeks of age.
To improve the efficiency of i-GONAD-mediated KI in rats, three gRNAs (crRNA1, crRNA2, and crRNA3), all of which recognize the target sites that are located very closely each other, were tested; consequently, KI efficiency varied among those gRNAs, suggesting that the choice of gRNA is important for determining the KI efficiency; the use of KI-enhancing drugs failed to increase the KI efficiency.
Showing successful production of RDEB mouse model with COL7A1 mutations, c.5818delC and E2857X; 5818delC homozygous mice developed severe RDEB-like phenotypes and died immediately after birth, whereas E2857X homozygous mice did not have a shortened lifespan compared to WT mice; adult E2857X homozygous mice showed hair abnormalities, syndactyly, and nail dystrophy.
Effects of AIMA treatment on an increased number of littermate in C57BL/6 mice were examined; the mean litter size following i-GONAD increased from 4.8 to 7.3 after the AIMA treatment; genetic modifications were confirmed in 80/88 (91%) of the offspring.
Rats (Rattus norvegicus) and mice (Mus musculus) are both classified into the same rodent family, and have been recognized as the most widely used models for biomedical research during the past four decades. However, these animals are different on many factors. For example, the rat is larger (roughly about eight- to ten-fold) in size than the mouse. Therefore, rats have been extensively used for pharmacological and surgical research, as exemplified by easier and more rapid microsurgery, multiple sampling of blood and tissues with relatively large amounts, and precise injection of substances into blood vessels (reviewed by Kjell and Olson [120]). Additionally, rats are considered as animals suitable for toxicological, neurobehavioral, and cardiovascular studies (reviewed by Jacob [121]). Since the first report in 1997 by Guerts et al. [122] on the production of genome-edited rats using ZFN technology, a total of 113 GM rats by MI (including pronuclear MI and cytoplasmic MI) or 9 rats by in vitro EP method have been produced during the period 1997–2021 (reviewed by Sato et al. [68]). Notably, GM rats often exhibited disease phenotype similar to that observed in humans, comparable to those shown in GM mice. For example, Zhang et al. [123] produced KO rats through MI of CRISPR/Cas9 components to obtain rat models for hereditary tyrosinemia type I (HT1), a disease caused by a deficiency in fumarylacetoacetate hydrolase (FAH) enzyme. These Fah KO rats developed remarkable liver fibrosis and cirrhosis, which have not been observed in Fah mutant mice. Furthermore, dystrophin-coding gene (dystrophin) (Dmd) KO rats (but not mice) presented cardiovascular alterations close to those observed in humans, which are the main cause of death in patients [124, 125]. These findings encourage the speculation that rats may better mimic the human situation than mice.
In 2018, two groups in Japan succeeded in generating genome-edited rats using i-GONAD. Kobayashi et al. [90] examined optimal in vivo EP conditions, allowing successful i-GONAD using NEPA21 electroporator and red fluorescent dextran (RFD) (tetramethylrhodamine-labeled dextran 3 kDa) as a fluorescent dye to monitor the fate of injected materials. Wistar Kyoto (WKY) strain rats were first subjected to the intraoviductal injection of a solution containing RFD + TB into pregnant female rats at Day 0.7 of pregnancy and subsequent in vivo EP using the NEPA21 electroporator under the following EP conditions: Pp: 30, 40, or 50 V, 5-ms pulse, 50-ms pulse interval, 3 pulses, and 10% decay (± pulse orientation); Tp: 10 V, 50-ms pulse, 50-ms pulse, 6 pulses, and 40% decay (± pulse orientation). When two-cell embryos recovered one day after i-GONAD were inspected for RFD-derived fluorescence, EP with 50 V for Pp and 6 times for Tp yielded maximal fluorescence in those embryos, with 74% efficiency. Using these optimal EP conditions, they attempted to disrupt the endogenous tyrosinase (Tyr) gene, a gene coding for protein essential for eye pigmentation, in pigmented females through intraoviductal injection of RNP (1 μg/μL of Cas9 protein +30 μM of sgRNA targeted to Tyr). As a result, genome-edited pups with albino-colored coat were obtained with an efficiency of 42%, when a Pp of 50 V was employed. They also attempted to recover the coat-color mutation in WKY females using an ssODN-based KI approach. Consequently, the KI efficiency was 27% in the pups born. They named this rat-based i-GONAD as “rGONAD.”
Takabayashi et al. [91] provided data similar to that of Kobayashi et al. [90], demonstrating that i-GONAD-based mutations (indels) resulted in the generation of fetuses (derived from pigmented Brown Norway (BN) × albino SD rat crosses) with non-pigmented eyes, with an efficiency of 56%. In this case, the in vivo EP condition used was almost the same as that used for i-GONAD in mice. They also tested the possibility of KI (targeted Tyr locus) using albino Lewis (LEW) rats and demonstrated that the i-GONAD-mediated KI efficiency was as low as ~5%, when the presence of fetuses with pigmented eyes was assessed. Takabayashi et al. [91] further attempted to disrupt another endogenous gene, paired box 6 (Pax6), an essential locus required for facial development, using i-GONAD. Out of 8 mid-gestational fetuses obtained, three had completely lacked eyes and lateral nasal prominence.
The golden hamster (Mesocricetus auratus) is a small rodent that has been extensively used in biomedical research in fields including oncology, immunology, metabolic disease, cardiovascular disease, infectious disease, physiology, and behavioral and reproductive biology [126]. In 1976, hamster oocytes were first used for IVF assay to test the fertilizing ability of human spermatozoa [127]. However, hamster embryos are highly vulnerable to in vitro conditions, hindering the generation of GM hamsters [128].
Hirose et al. [96] attempted to produce GM hamsters using i-GONAD, which allows embryo manipulation under the environment where the effects of handling embryos in vitro can be avoided as possible. Through intraoviductal injection of six sgRNAs (targeted acrosin gene) and the Cas9 protein, they produced KO hamsters lacking expression of acrosin, a protein thought to be essential for sperm penetration through ZP, to investigate how acrosin-KO hamster spermatozoa behaved both in vivo and in vitro. A total of 15 pups obtained, eight of which were weaned. Of these, five were found to have mutant alleles. Homozygous mutant males were completely sterile, as the mutant spermatozoa attached to ZP, but failed to penetrate it. This finding indicates that in hamsters, acrosin plays an indispensable role in allowing fertilizing spermatozoa to penetrate ZP.
7.1 Two-step i-GONAD can introduce mutations at two sites located close to each other
CRISPR/Cas9-based introduction of indels into two sites (which are closely located each other) through a single shot of transfection has been recognized difficult, because frequent deletion between these two sites occurs. However, sequential transfection of genome editing reagents can avoid such deletion between the two sites. To test the possibility, Sato et al. [97] tried to generate two types of indels at two target sites (that are located very close to each other; 44 bp apart) by performing i-GONAD sequentially (Figure 4). The two gRNAs were first designed to recognize the upper and lower portion of exon 4 of α-1,3-galactosyltransferase gene (GGTA1), coding for the protein essential for synthesizing the cell-surface α-Gal epitope [129]. For the 1st i-GONAD, a solution containing Cas9 protein, sgRNA (termed “A”) recognizing the upper portion of exon 4, and dye (Fast Green FCF; for monitoring injection process) was injected intraoviductally at Day 0.7 of pregnancy (corresponding to late zygote stage). Next day, a solution containing Cas9 protein, sgRNA recognizing the lower portion of exon 4 (termed “B”), and dye was injected intraoviductally at Day 1.7–1.8 of pregnancy (corresponding to 2-cell stage). One day after the final surgery, morulae were isolated for single embryo-based analysis for possible indels at the target sites. As a result, the efficiency of successful generation of morulae with indels at both two sites was 18%. In contrast, i-GONAD using two sgRNAs (A and B) + Cas9 protein at Day 0.7 of pregnancy failed to generate morulae with mutations in both sites at exon 4 of GGTA1. Based on these findings, Sato et al. [97] named this approach “sequential i-GONAD (si-GONAD).”
Figure 4.
Schematic of the detailed procedure of sequential improved genome-editing via oviductal nucleic acid delivery (si-GONAD). After first i-GONAD on Day 0.7 of pregnancy, second i-GONAD is performed as shown in the panel on Day 1.7 of pregnancy. This figure was drawn in-house, and reproduced with permission from Sato et al., “Sequential i-GONAD: An Improved In Vivo Technique for CRISPR/Cas9-Based Genetic Manipulations in Mice”; published by MDPI, 2020.
7.2 Preparation of floxed mice using i-GONAD
The Cre/loxP system is a useful tool for assessing in vivo gene function. Spatially and temporally-controlled expression of Cre recombinase enables precise deletion of loxP-floxed chromosomally integrated GOI. To realize it, two loxP sites must be simultaneously inserted in cis into the target locus. The resulting mice are called “conditional KO mice.” Previously, this process for conditional KO mouse production was achieved by embryonic stem (ES) cell-based gene targeting and subsequent chimeric mouse production, which is time-consuming and labor intensive [130]. To bypass this tedious process, attempts to produce mice carrying loxP-floxed GOI (which are generally called “floxed mice”) have been made using genome editing technology. In the initial experiments, simultaneous injection of Cas9, two pairs of gRNAs, and two ssODNs containing lox sequences into mouse zygotes generates mice containing floxed alleles [61, 131, 132, 133, 134]. This approach can generate floxed mice without using ES cells, since it does not require the construction of a KI vector, and production of floxed mice is finished in a short period of time (e.g., in a month). However, according to Horii et al. [130], the simultaneous introduction of two mutated lox sites (to which Cre recombinase bind) at a target locus is difficult as it often causes LD of a sequence. To increase the possibility that the two lox sites are knocked-in, they used the sequential introduction approach to perform KI of mutated lox sites into the introns interposing exon 3 of methyl-CpG binding protein 2 gene (Mecp2) through in vitro EP. When the resulting embryos (blastocysts) were subjected to molecular analysis, 21% (33/155 tested) of the embryos had two floxed sites in the target Mecp2 locus. Furthermore, the efficiency of generating LD was 36% (56/155 tested). Sato et al. [97] employed all the genome-editing components (gRNAs and ssODNs containing mutated lox sites as donor DNA) described by Horii et al. [130] for si-GONAD. Unfortunately, the generation of morulae with KI alleles (in which floxed sites had been knocked-in in both sides of the introns interposing exon 3 of Mecp2) failed; only a morula with one floxed site in the 5′ site of Mecp2 was successfully generated.
Recently, Shang et al. [101] developed a new approach by integrating a unique design of asymmetric loxP-ssODN to create mouse conditional KO alleles in one step using the i-GONAD method. They injected a cocktail containing Cas9 protein, two gRNAs targeting the intron 2 and 3′ region of Fos-like antigen 1 gene (Fosl1), and two short ssODNs as HDR donors for loxP insertions. Each ssODN is 161 nucleotide (nt) long, composed of 91 nt of the 5′ homology arm from the PAM-proximal side, 34 nt of loxP sequence, and 36 nt of the 3′ homology arm from the PAM-distal side. Molecular biological analysis of the resulting pups demonstrated that out of 20 F0 mice obtained one mouse had the simultaneous 5′- and 3′-loxP insertions and 6 had either 5′- or 3′-loxP integrations. Similar experiments were also conducted to obtain floxed mice for genes coding for pleomorphic adenoma gene-like 1 (Plagl1), Ak040954, cardiotrophin-like cytokine factor 1 (Clcf1), and Gm4438. The overall targeting efficiency of producing floxed alleles by i-GONAD was 10% (8/76 tested).
7.3 RFD is a useful reagent to master GONAD/i-GONAD in 2 days
RFD has been recognized as a useful reagent to judge the success of gene delivery to early mammalian embryos after in vitro EP [67]. It has also proved useful for judging the success of GONAD/i-GONAD [53, 86, 90, 91, 97, 119]. For example, when we performed i-GONAD using a solution containing RFD and TB (or Fast Green FC) at Day 0.7 of pregnancy (corresponding to late zygote stage; 16:00 PM), the success of the approach can be judged by examining the RFD-derived red fluorescence in the isolated 2-cell embryos under fluorescence microscopic observation one day after i-GONAD. The presence of fluorescent embryos means successful i-GONAD, while the absence of fluorescent embryos indicates failure of i-GONAD. This short-term experience is especially beneficial for the beginners who want to master the technique, and is thus called “2-day protocol for mastering GONAD” [117]. Notably, Sato et al. [97] showed that FITC-labeled fluorescent dextran is also effective for reporting the success of gene delivery in mouse early embryos.
7.4 Chromosomal engineering using i-GONAD
As shown in Section 7–1, CRISPR/Cas9-mediated genome editing using 2 types of gRNAs results in frequent generation of LD in the sequence flanked by the two sites recognized by these gRNAs. Iwata et al. [93] applied this technique to introduce chromosomal inversions of several megabases (Mb) in mice. When mouse zygotes were subjected to in vitro EP, a 7.67 Mb inversion was successfully introduced, which is longer than any previously reported inversion produced using MI-based methods. They confirmed that a similar event can be induced using i-GONAD. These findings suggest that CRISPR/Cas9 system via in vitro and in vivo EP is useful for examining genetic diseases with large-scale chromosomal rearrangements.
Notably, the same group [107] recently demonstrated that i-GONAD can be useful to maintain lethal mutant mice using an inversion balancer identified from the C3H/HeJJcl strain. As a proof-of-principle, they created the Tp53rk binding protein gene (Tprkb) KO strain with an embryonic lethal mutation through i-GONAD in the presence of a non-targeted B6.C3H-In(6)1 J inversion. Iwata et al. [107] demonstrated that the edited lethal genes were stably maintained in heterozygotes, as recombination is strongly suppressed within this inversion interval. This strategy may facilitate further analysis of lethal mutants.
7.5 Viral transduction using i-GONAD
Virus-based gene delivery approaches have been widely used in the biomedical sciences, especially for gene therapeutic purposes (reviewed by Sung and Kim [135]). The viral vectors widely used are RV, AV, LV, and adeno-associated viral vector (AAV). Each of these vectors have specific properties. For example, RV and LV can infect both dividing and senescent cells and enable chromosomal integration. AV can infect mainly dividing cells efficiently, but cannot integrate into host chromosomes. These viral vectors (but not AAV) can infect ZP-removed early embryos, but not ZP-enclosed (or intact) embryos [13]. Notably, the simple incubation of ZP-enclosed embryos with recombinant AAV (rAAV)-containing medium was recently shown to lead to the transduction of those embryos [13, 92]. Notably, there are over 10 different serotypes of AAVs, each of which exhibits different infectious ability depending on the type of cells [136]. Mizuno et al. [13] examined which serotype of AAV could effectively transduce ZP-intact mouse 2-cell embryos. The embryos were co-incubated for 16 h with several types of rAAVs carrying an eGFP expression unit and then transferred to normal medium; the morulae developing after co-incubation with AAV serotype 6 (which is hereinafter called rAAV-6) exhibited strong fluorescence [13]. The next vector showing relatively strong infectivity was rAAV-1. A similar observation was also made by Yoon et al. [92]. Importantly, rAAV-6 can transduce rat and bovine embryos [13], suggesting the multi-species infectivity of this vector.
However, genome editing could not be induced in early embryos through transduction with rAAV-6. The rAAV carrying Cas9 gene (~9 kb in size) and that carrying gRNA could not be co-delivered, because rAAV was unable to incorporate over 4.5-kb of an insert [137]. To perform successful CRISPR-based KI in mice, Mizuno et al. [13] employed a two-step gene delivery approach. Zygotes were first subjected to in vitro EP in the presence of RNP and then transduced with rAAV-6 carrying a 1.8-kb GFP expression cassette flanked by two 100-bp Rosa26 homology arms. Molecular biological analysis of the newborn pups demonstrated that the KI efficiency in the Rosa26 locus was 6%. Yoon et al. [92] performed intraoviductal injection of a solution containing rAAV6-Cas9 (carrying spCas9 gene derived from Streptococcus pyogenes) and rAAV6-gTyr (carrying gRNA expression unit targeted Tyr) into the pregnant female mice at Day 0.5 of pregnancy, similar to GONAD/i-GONAD. Molecular biological analysis of the newborn pups demonstrated that the indel efficiency was 6%. All mutated founder (F0) mice generated albino offspring, indicating germ-line transmission; this suggested that AAV is a powerful tool for inducing genome editing in the ZP-enclosed early embryos in vivo. According to Sato et al. [138], this in vivo approach is referred to as “AAV-based GONAD.”
As was mentioned earlier, AAV-based GONAD appears to be more convenient than GONAD/i-GONAD, since the former does not require any of the apparatus required for EP. Unfortunately, it still requires more detailed information concerning (1) which rAAV serotype is effective for in vivo transduction towards early mouse embryos, (2) the stage allowing maximal expression of GOI (included in rAAV) after infection at late zygote stage, and (3) whether the oviductal epithelial cells are infected through AAV-based GONAD. Sato et al. [139] first examined the above possibility using 4 types (1, 2, 5, and 6) of rAAVs carrying a unit for expression of eGFP (as GOI). When AAV-based GONAD was performed at Day 0.7 of pregnancy, and 2 days later the morulae were isolated to inspect eGFP fluorescence, rAAV-6 gave strongest fluorescence, though the fluorescence intensity varied among embryos. The fluorescence intensity provided by rAAV-1 was the next highest, but transduction with the other remaining serotypes (2 and 5) resulted in negative or faint fluorescence in the embryos. These results are consistent with the in vitro data from Mizuno et al. [13] and Yoon et al. [92]. A similar mode of transduction was also seen in the oviductal epithelial cells, suggesting the use of rAAV-6 (or possibly rAAV-1) for genome manipulation of those cells.
AAV-based GONAD using rAAV-6 was performed at Day 0.7 of pregnancy, and one day later, 2-cell embryos were isolated and cultured until the late blastocyst stage to monitor the eGFP fluorescence expression. Under the fluorescence microscope, fluorescence was first discernible at the 2-cell stage, attained at a maximal level at the morula stage, and declined towards late blastocyst stage [139]. These results suggest that one-day infection with rAAV-6 is enough to transduce ZP-enclosed zygotes floating in the oviductal lumen. Furthermore, the GOI expression was transient, peaking at the morula stage. These findings suggest a possibility that early mouse embryos from zygote to morula stages can be effectively transduced in a sequential manner, like si-GONAD.
7.6 Effect of HA treatment on the efficiency of i-GONAD-mediated genome editing
As was mentioned in Section 4, i-GONAD at Day 0.4 of pregnancy (corresponding to early zygote) often failed to obtain genome-edited embryos/fetuses. This appears to be solely due to the presence of cumulus cells that tightly surround a zygote. One idea to overcome this problem may be that early zygotes are pretreated with HA, an enzyme capable of dispersing cumulus cells from a zygote [1], prior to i-GONAD. Kaneko and Tanaka [100] examined the possibility by injecting 1 μL of 0.1% HA into the ampulla of a female (ICR) at Day 0.4 of pregnancy (10:00–11:00) using a thin glass needle. As a control, the solvent (PBS) was similarly injected. Several minutes after the injection, a solution (1 μL) containing genome editing reagents (2 μg/μL Cas9 protein +60 mM dual gRNA (targeted fibroblast growth factor 10 gene (Fgf10) + 0.08% TB) was intraoviductally introduced and subsequently the entire oviducts were subjected to in vivo EP using tweezer-type electrodes. After that, the developing fetal offspring were isolated for examining the presence of possible genome editing in those samples. Consequently, the samples isolated from HA-treated group exhibited 2.5-fold higher genome editing (indels) efficiency than those isolated from the control group (68% vs. 27%). The i-GONAD on Day 0.7 of pregnancy (16:00–17:00; in which case no HA is used) yielded genome edited pups with an efficiency of 54%. These findings indicate that HA-mediated removal of cumulus cells at Day 0.4 of pregnancy is effective when in situ genome editing towards early zygotes are intended. According to Kaneko and Tanaka [100], the operation time for introducing genome editing reagents into embryos in the oviducts can be adjusted by treatment with HA before EP. This improved protocol can also be used for efficient production of genome-edited mice and rats.
7.7 Strain-difference can affect the efficiency of i-GONAD-mediated genome editing
According to Ohtsuka et al. [86], successful i-GONAD relies on the mouse strain used. For example, it worked successful under relatively stringent electrical conditions (40 V/100–200 Ω/~300 mA) when random-bred mice (such as MCH(ICR) and B6C3F1, a hybrid between C3H/He and C57BL/6), but not C57BL/6 strain, were used. Under less stringent conditions (40 V/350–400 Ω/~100 mA), i-GONAD was successful in the inbred C57BL/6 strain [86, 98, 117]. These findings suggest the importance of selecting the appropriate EP conditions, particularly when different mouse strains are used for i-GONAD experiment.
This is also true for i-GONAD using rats. For example, when a current of >500 mA was employed using the NEPA21 electroporator, albino SD and albino LEW rats were successfully genome-edited; however, no offspring were derived from pigmented BN rats (fetuses/newborns) [91]. In contrast, i-GONAD was performed under a current of 100–300 mA using the NEPA21 electroporator, leading to the production of genome-edited BN rats at efficiencies of 75–100% [99]. Similar success in producing GM BN rats was achieved with efficiencies of 24–55% when another electroporator CUY21EDIT II (BEX Co., Ltd., Tokyo, Japan) was employed under a current of 150–200 mA [99].
Notably, the most widely used electroporators (as exemplified by NEPA21) employ a constant voltage. Also, other electroporators (as exemplified by GEB15 (BEX Co., Ltd.)) employ a constant current. Kobayashi et al. [98] explored the conditions allowing the generation of a 100 mA current in C57BL/6 mice using two electroporators, NEPA21 and GEB15. As a result, i-GONAD performed under conditions of average resistance of 367 Ω and average voltage of 116 mA resulted in the production of genome-edited fetuses with efficiency of 39%.
7.8 Attempt to increase the efficiency of KI using i-GONAD
In our previous study using i-GONAD to produce GM rats and KO/KI rats, the success rate of producing KI rats was lower than that of KI mice (5% vs. 60%, respectively) when ssODNs were used as KI donors [91, 117]. To improve the efficiency of i-GONAD in rats, Aoshima et al. [112] examined the effects of commercially available KI-enhancing drugs (including SCR7, L755,507, RAD51-stimulatory compound 1 (RS-1) and Alt-RⓇ HDR Enhancer (HDR enhancer)), some of which have been known to increase KI efficiency in culture cells and early embryos [140, 141, 142, 143]. For example, i-GONAD was applied to SD female rats (albino) using a solution containing RNP complex (consisting of Cas9 protein and gRNA targeted Tyr locus), ssODN (used as a KI donor oligodeoxynucleotide), and various amounts (5 or 15 μM) of L755,507 on Day 0.7 of pregnancy. Inspection of mid-gestational fetuses revealed that 12% of fetuses obtained showed pigmented eyes when 5 μM L755,507 was used for i-GONAD, suggesting successful KI [112]. In addition to L755,507, some drugs (e.g., SCR7 and HDR enhancer) were found to be effective in i-GONAD in rats, but their effects were limited.
In a study by Aoshima et al. [112], three gRNAs (called crRNA1, crRNA2, and crRNA3) were used. As shown in Figure 5, these gRNAs recognize different portions of the target locus, but also overlap each other in the target locus. Surprisingly, the KI efficiency in rat fetuses generated after i-GONAD with crRNA2 and ssODN was significantly higher (24%) than crRNA1 (5%) or crRNA3 (0%). The KI efficiency of i-GONAD with triple gRNAs was 11%. These findings demonstrated that the choice of gRNA is important for determining KI efficiency.
Figure 5.
Schematic of knock-in (KI) experiment in rats towards the mutated tyrosinase gene (Tyr) locus performed by Aoshima et al. [112]. The target sequence (exon 2 of Tyr) recognized by crRNA1, 2, and 3 is shown in green. PAM sequences are underlined. Single-stranded oligodeoxynucleotide (ssODN) (containing wild-type nucleotide “G” that corresponds to mutated nucleotide “A”) is shown in orange below the target sequence. The nucleotide “A/T” marked in red is the mutation causing the albino phenotype. This figure was drawn in-house, and reproduced with permission from Aoshima et al., “Modification of improved-genome editing via oviductal nucleic acids delivery (i-GONAD)-mediated knock-in in rats”; published by BioMed Central Ltd, 2021.
7.9 Regulated timing of i-GONAD by administration of gonadotrophins
The i-GONAD experiment using C57BL/6 strain is always associated with the difficulty in consistently obtaining pregnant females, because estrous females are not always available. The administration of gonadotrophins has been frequently used for inducing superovulation in many mouse strains to obtain a number of early embryos [1]. This approach has an additional advantage in that it is capable of synchronizing the estrous cycle of females; thus, the estrous cycle need not be examined through smear testing or through visual inspection of the vagina.
Administration of higher dose (in this case, more than 5 international units (IU)) of gonadotrophins can induce superovulation, but often causes failure to deliver pups [144, 145, 146]. Administering low doses (less than 5 IU) of gonadotrophins facilitates ovulation of natural number of oocytes and successful delivery of pups [147, 148]. Notably, Sato et al. [53] reported that intraperitoneal (IP) administration of low-dose (2–0.5 IU) serum gonadotrophin (PMSG) from a pregnant mare on 11:00, followed by 5 IU of human chorionic gonadotrophin (hCG) 48 h later, is effective for inducing natural ovulation before i-GONAD. In case of administration of 5 IU PMSG, females having vaginal plug failed to deliver their pups. When females were inspected later, some had dead fetuses in their uterus. When 2 or 0.5 IU PMSG was administered, all females successfully delivered viable pups (average: 8 in each group). These findings suggest that i-GONAD can be performed on 11:00 at Day 0.7 of pregnancy when females were induced to ovulate by administering a low dose of PMSG. Indeed, Sato et al. [97] demonstrated that i-GONAD on 11:00 at Day 0.7 of pregnancy leads to generation of genome-edited morulae.
Unfortunately, the regime shown by Sato et al. [53] has only been used successfully on B6C3F1 hybrid mice. Kobayashi et al. [98] examined whether the administration of a single IP injection of low-dose PMSG (1.2 IU/10 g) is effective for synchronizing the estrous cycle in C57BL/6 females. Consequently, approximately 51% of C57BL/6 females had plugs upon mating with males 2 days after PMSG administration, which contrasts with the case (~26%) when C57BL/6 females were subjected to natural mating. Furthermore, 44% hormone-injected and plugged females delivered pups with an average litter size of six, which was comparable to the rate obtained from females that were not injected with hormones. These findings indicate that a single IP injection of low-dose PMSG increases the rate of acquiring plugged females before mating. This is particularly beneficial for i-GONAD which always requires desired number of plugged females obtained through a scheduled mating.
7.10 Combinational use of i-GONAD with anti-inhibin monoclonal antibodies (AIMAs) treatment to increase the number of GM mice
Many attempts have been made to increase litter sizes (which is determined by the number of oocytes naturally ovulated) using conventional superovulation regimens (e.g., using PMSG/hCG), but had limited success because of unexpected decreases in the numbers of embryos surviving to term, as mentioned in the Section 7–9. Hasegawa et al. [115] attempted to overcome this problem using rat-derived AIMAs. They administrated progesterone (P4) once a day for 2 days (days 1 and 2) to synchronize the estrous cycle of female C57BL/6 mice, and AIMAs were injected into the same animals at Day 4 followed by mating with male C57BL/6 mice. When i-GONAD targeting Tyr was applied to the AIMA-treated C57BL/6 female mice on the day of vaginal plug formation during Days 6–8, a 1.5-fold increase in litter size was observed (7.3 vs. 4.8 for the untreated control). Notably, genome editing efficiency did not differ between these two groups. Therefore, AIMA treatment can reduce the number of females used for the i-GONAD experiment, which will fulfill the 3R principles of animal experimentation (i.e., Reduction, Replacement, and Refinement), and can be applied to other mouse strains and animals.
7.11 GONAD/i-GONAD as a useful tool to check in vivo gene correction event
As was shown previously, GONAD/i-GONAD enable gene delivery to early embryos present within the oviductal lumen and to the epithelial cells facing the lumen [37, 53, 139]. The success of gene delivery to the oviductal epithelial cells can be easily judged through direct observation of fluorescence under a fluorescence microscope [37, 53, 139]. Therefore, in vivo gene delivery approach targeting oviductal epithelial cells are excellent for testing the function of the GOI.
Miura et al. [109] attempted to examine whether in vivo CRISPR/Cas9-mediated gene correction is possible using GONAD/i-GONAD technologies. They first generated ΔeGFP KO mouse strain through MI of a solution containing Cas9 mRNA and gRNA (targeted eGFP cDNA) into zygotes from Tg mice carrying eGFP cDNA [85]. The resulting ΔeGFP KO mice failed to exhibit systemic eGFP expression, due to frame-shift mutations in the coding region of eGFP cDNA that was chromosomally integrated in their genome. Next, they injected a solution containing sgRNA targeted to the mutation site, Cas9 protein, ssODN (as donor DNA sequence; in some cases), and Fast Green FCF into the oviductal lumen of female ΔeGFP KO mouse. Subsequently, the entire oviducts were electroporated, similar to that performed in GONAD/i-GONAD. Three to 13 days later, the eGFP fluorescence was inspected in the oviducts dissected from the treated females. Consequently, fluorescence was detected in a portion of an oviduct, suggesting gene editing at the mutated site in the eGFP cDNA through HDR-mediated KI of ssODN or NHEJ-mediated indels. Molecular biological analysis of the oviduct confirmed the above events. Notably, in this system, editing of mutated site can be easily monitored by visually inspecting the gene-edited oviducts under UV illumination.
7.12 GONAD/i-GONAD as a useful tool to generate mouse models for ovarian cancer
As was mentioned previously, GONAD/i-GONAD could transfect both preimplantation embryos and the oviductal epithelium facing the oviductal lumen. Ovarian cancer is the most lethal gynecologic cancer to date. High-grade serous ovarian carcinoma (HGSOC) is the most common type of ovarian cancer and has the lowest rate of survival. Teng et al. [149] recreated the mutations found in ovarian cancer to generate somatic ovarian cancer mouse models, using an in vivo oviductal EP method similar to GONAD/i-GONAD. Using the CRISPR/Cas9 genome editing approach, they mutated the tumor suppressor genes (transformation related protein 53 (Trp53), breast cancer susceptibility gene I (Brca1), neurofibromin 1 (Nf1), and phosphatase and tensin homolog (Pten)) to study how these genes contribute to tumor development. When mutations were introduced in three of the four genes, namely Trp53, Brca1, and either Nf1 or Pten, the sites transfected with the genome editing reagents displayed effects that were similar to human HGSOC and changes in chromosome number. Teng et al. [149] concluded that the in vivo oviductal EP method is highly useful for generating mouse models to advance the understanding and treatment of ovarian cancer.
8. Challenges, limitations, concerns and future perspective
8.1 Challenges
GONAD/i-GONAD is performed by injecting a small amount (1–2 μL) of a solution containing genome editing components into the specific site of an oviduct (called ampulla) of a pregnant female at Day 0.7 to 1.4 of pregnancy, using oral breath-controlled glass micropipette under a dissecting microscope. The genome editing components introduced within an oviductal lumen exist around ZP-enclosed early embryos (zygotes to 2-cell embryos), but are never incorporated into those embryos in an intact state. However, in vivo EP enables delivery of substances (present outside the embryos) into the internal portion of an embryo, leading to generation of genome-edited embryos. Another method to induce genome editing in early embryos in situ is viral transduction using rAAV. Unfortunately, EP requires expensive apparatus such as an electroporator, and rAAV transduction requires labor-intensive and time-consuming preparation of viral particles. In this context, the use of ZP-penetrating agents (e.g., MWNT and VisFect) would be ideal, because it does not require an electroporator or viral preparation. To date, there is no report on successful genome editing in early embryos using these ZP-penetrating agents. We expect that these agents could be useful for facilitating NA delivery to embryos and subsequent induction of genome editing.
Kaneko and Tanaka [100] demonstrated that pretreatment of early zygotes (tightly surrounded by cumulus cells) with HA led to increased efficiency of genome editing in those embryos after i-GONAD. This is based on the concept that cumulus cells surrounding a zygote hamper rapid transfer of genome editing reagents to zygotes [86]. In the previous approach using GONAD/i-GONAD, in vivo EP is applied immediately after intraoviductal injection of NAs [84, 86, 116]. In this case, it is highly likely that the reagents injected might not have been fully infiltrated between the intercellular space connecting cumulus cells. Waiting for several minutes after intraoviductal injection may permit sufficient infiltration of the reagents before in vivo EP, leading to increased efficiency of genome editing. This line of experiment has now been undertaken by Takabayashi and his colleagues.
8.2 Limitations
GONAD/i-GONAD can be applied to larger animals, as exemplified by its use in pigs; the demand for GM pigs has been rising due to the needs of the biomedical and agricultural fields [150, 151]. The current strategy for creating GM pigs is based on “ex vivo handling of embryos,” where MI or EP is carried in vitro towards zygotes collected from individuals or produced through IVF, following which the treated embryos have to be subjected to ET to recipient females [138]. Similar to the case of MI or in vitro EP-mediated production of GM mice and rats, creation of GM pigs is highly costive, time-consuming, and labor-intensive. Additionally, it requires recipient female pigs, which are also expensive. In this context, GONAD/i-GONAD should be theoretically performed in more convenient and inexpensive manner, since it does not require “ex vivo handling of embryos.” Notably, the porcine oviduct is generally ca. 100 mm in length and their form is linear, unlike the spiral form in rodents (mice and rats). Furthermore, it does not have an enlarged site called “ampulla” where rodent zygotes always stay. As a result, porcine zygotes will exist in a broad area throughout the oviduct. To perform GONAD/i-GONAD in pigs, researchers must inject a large amount of fluid (probably over 1 mL) and electroporate towards several sites whenever tweezer-type electrodes are used. It remains uncertain whether this is feasible. Therefore, GONAD/i-GONAD in pigs remains a challenge.
8.3 Concerns
As was mentioned in Section 1, in the initial step of development of GONAD/i-GONAD, gene delivery to early murine embryos using plasmid DNA were successful when the T-820 electroporator from BTX Co. was used [28]. However, it was impossible when the other electroporators such as NEPA21 and CUY21EDIT II were employed (Sato, Ohtsuka, unpublished). Notably, Hakim et al. [54] recently checked several in vitro EP parameters to seek optimal conditions enabling gene (plasmid DNA) delivery into mouse follicles, oocytes, and early embryos. When they were electroporated in the 1-mm gap cuvettes using Gene Pulser Mxcell System (Bio-rad Laboratories, Hercules, CA, USA), EP under 3 pulses of 30 V of 1 ms each at an interval of 10 s was ideal, with no need to weaken or loosen the ZP layer. This suggests that exploration of optimal EP condition using the above apparatuses (NEPA21 and CUY21EDIT II) may enable the transfection of ZP-intact embryos with plasmid DNA. If it is realized, transgenesis via introduction of plasmid-based transposons (e.g., piggyBac (PB) transposon + PB transposase mRNA) may be possible.
8.4 Future perspective
Preimplantation embryos at zygote to morula stages, all of which exist within the oviductal lumen of a pregnant female, can be a target for gene delivery through GONAD/i-GONAD. As cleavage embryos (at 2-cell to early 8-cell stages) are comprised of blastomeres, each of which is facing the outside environment, genome editing at these stages may frequently result in the generation of mosaic offspring containing edited and unedited cells. This mosaic nature is tedious for the researchers who want to produce GM animals with high efficiency, but in turn beneficial for investigating the properties of embryonic lethal genes, because mosaic fetuses or pups produced through MI of genome editing components into one blastomere of two-cell embryos should be viable and carry heritable lethal mutations [152]. On the other hand, in the case of compacted 8-cell embryos and morulae, only blastomeres facing the external environment (but not the inner cells present inside an embryo) can be susceptible to genome editing. The outer blastomeres of morulae are thought to contribute to the formation of a trophectodermal cell, which is a cell involving implantation and placenta formation. Therefore, the GONAD/i-GONAD-mediated genome manipulation at these stages may be a novel tool to explore the molecular mechanisms underlying implantation and placenta formation.
Seven years have passed since the first report [84] on the development of GONAD using mice. During this period, successful genome editing was reported in other animals including rats and hamsters. The genes targeted by GONAD/i-GONAD technologies were eGFP cDNA (chromosomally integrated in the eGFP Tg mice) [84, 109] and endogenous genes such as Acrosin, Agouti, Ak040954, ATF5, Axdnd1, Cdkn1a, Cdkn2a, Clcf1, COL4A3, COL4A4, COL4A5, COL7A1, Dll1, eEF2, Fgf10, Fosl1, Foxe3, Gbx2, GGTA1, Gm30413, Gm44386, Hprt, Kit, Klrc2, Mecp2, Mov10l1, Nrsn2, Pafah1b1, Pax6, Pitx3, Plagl1, Serpina3n, Tis21, Tprkb, and Tyr (Table 1). All these genes were disrupted by indel mutations (KO) or modified through HDR-mediated KI using the CRISPR/Cas9 system. As for the components used for CRISPR/Cas9-mediated genome editing, Cas9 mRNA was initially employed, but later Cas9 protein was mainly used. For KI experiments, ssODN or in vitro synthesized long ssDNA was used as donor DNA. The dye used for monitoring successful injection process was also changed; while TB was initially employed, Fast Green FCF was used for later experiments. According to Ohtsuka et al. [86], TB (but not Fast Green FCF) often generates visible precipitates, when RNP is mixed with the dye. GONAD/i-GONAD technologies are very simple systems that only require the intraoviductal injection of NA-containing solution and subsequent in vivo EP. EP is now recognized as a powerful tool enabling efficient gene delivery, but often causes deleterious effects on cell/embryo survival, leading to reduction in litter size, as suggested by Kaneko and Tanaka [100]. Ohtsuka et al. [86] first demonstrated that the success of in vivo EP depends on the mouse strains used. This was also true for generating GM rats using i-GONAD [91]. Therefore, exploration of optimal EP conditions is required before GONAD/i-GONAD experiments start using new strains.
GONAD/i-GONAD requires an expensive electroporator. Notably, the intraoviductal injection of rAAV into a pregnant female mouse was also useful for the in situ transduction of zygotes [92, 139]. Furthermore, it has already been shown that AAV-mediated genome editing at zygotes through AAV-based GONAD is powerful for the convenient acquisition of GM animals [92]. In this AAV-based GONAD system, the electroporator need not be used, although it is always associated with laborious and time-consuming tasks such as viral vector preparation. For developing systems that are simpler than the present GONAD/i-GONAD, the use of ZP-penetrating reagents will be highly desirable.
Recently, new genome editing systems known as prime editing [153, 154, 155, 156] and base editing [157, 158, 159] were reported. These systems enable precise gene correction at a single nucleotide level. To date, these reagents have not been used for GONAD/i-GONAD-mediated production of GM animals. Future application of these new genome editing systems to GONAD/i-GONAD is highly expected.
We thank Kazusa Inada for her support for in-house drawing of the Figures, shown in Figures 1–4. This study was partly supported by a grant (no. 24580411 for Masahiro Sato; no. 21 K10165 for Emi Inada; no. 16H05049 for Shingo Nakamura; no. 22H03277 for Issei Saitoh; no. 16 K07087 for Shuji Takabayashi) from the Ministry of Education, Science, Sports, and Culture, Japan, and a fund for the Promotion of Joint International Research (Fostering Joint International Research) (no. 16KK0189 for Masato Ohtsuka) from JSPS, Japan.
Conflict of interest
The founding sponsors had no role in the design of the study, collection, analyses, or interpretation of data, writing of the manuscript, and decision to publish the results.
Notes
Masahiro Sato and Shuji Takabayashi designed the study and drafted the manuscript; Masato Ohtsuka, Emi Inada, Shingo Nakamura and Issei Saitoh critically revised the manuscript.
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