Poultry, such as the chicken (Gallus domesticus), turkey (Meleagris gallopavo), and Japanese quail (Coturnix japonica), are commercially important species because they serve as major food sources worldwide. The poultry industry is indispensable for supporting sustainable development worldwide and is expected to significantly contribute to the United Nations’ Sustainable Development Goals. Studies on the embryos of these birds have contributed to a more detailed understanding of organogenesis ^[1][2][1,2] and human diseases [1], due to their oviparity. Chicken and quail embryos are easy to access and, thus, are amenable to introducing transgenes, normal or transgenic primordial germ cells (PGCs), and viruses ^[3][4][3,4]. The draft genome sequences of chicken, quail, turkey, and zebra finch genomes have been generated, with the information obtained supporting the generation of transgenic birds and the mass production of recombinant proteins ^[5][6][7][8][5,6,7,8]. The generation of gene knockout animals is a powerful approach for identifying essential proteins and the functions of uncharacterized genes in many species, as is the case in birds. The combination of recent technical advances is expected to provide novel insights into future aspects of avian biotechnology as well as human society.

The generation of gene knockout chickens by homologous recombination using PGC-mediated methods was initially reported in 2013 [9] following its establishment in mice [10]. Immunoglobulin light chain knockout chickens were subsequently generated using the same method [11]. Although these achievements contributed to the progression of avian gene targeting, the efficiency of this recombination was very low. In this regard, the emergence of site-specific nuclease has provided new avenues for modifications to avian genomes ^[12][13][14][12,13,14]. The earliest programmable genome editing tools for the generation of gene knockout animals were zinc finger nuclease (ZFN) and transcription activator-like effector nuclease (TALEN) [15]. These enzymes were artificially created by the fusion of FoKI endonucleases with the capacity to recognize long-chain DNA. FoKI endonucleases recognize target DNA and induce double-strand breaks (DSB) and small indels by error-prone non-homologous end joining (NHEJ).

2. CRISPR/Cas9-Mediated Genome Editing Technology

The CRISPR/Cas system, originally found in bacteria and archaea, is an RNA-based adaptive immune system that destroys invading plasmids, phages, and viruses ^[16][17][18][19,20,21]. The nucleoprotein complex, consisting of three crucial components: CRISPR-coding RNA (crRNA), trans-activating crRNA (tracrRNA), and Cas protein, recognize exogenous DNA and degrade it by endonuclease activity ^[19][22], causing error-prone NHEJ. The binding site of Cas9 is located upstream of the protospacer adjacent motif (PAM) containing the 5′-NGG base sequence. Humanized Cas9 protein or Cas9 derived from Streptococcus pyogenes Cas9 (SpCas9), combined with a synthetic single guide RNA (sgRNA) produced by fusing crRNA with tracrRNA, has been shown to trigger DSB in mammalian cells ^[20][21][23,24]. This CRISPR/Cas9 system may be easily designed and prepared for a plasmid vector by changing the sgRNA sequence to a specific region of the genome sequence instead of ZFN and TALEN described above, and its use has rapidly expanded. The recent re-engineering of Cas9 has resulted in the establishment of dead Cas9 (dCas9) with mutations in two nuclease domains, which has extended the application of the CRISPR/Cas9 system with the potential transcriptional inhibitor/activator, and point mutations ^{[22][23][24][25]}[25,26,27,28] (Figure 1A). Three viral vectors have so far been adopted in clinical trials for the delivery of the CRISPR/Cas9 system: adeno-associated viruses (AAV), adenoviruses, and lentiviruses ^[26][29]. AAV are preferred due to their low immunogenicity and stable expression. However, the construction of long Cas9 sequences in plasmids is difficult due to the packaging limitation of AAV. This issue may be partially resolved using truncated SpCas9 ^[27][30]. On the other hand, the packaging capacities of adenoviruses and lentiviruses are high. In addition, lentiviruses and adenoviruses show high infection efficiencies in non-dividing cells and non-/dividing cells, respectively. However, lentivirus vectors generally induce insertion mutations through the sustained expression of Cas9 and sgRNA, which may result in off-target effects, while other adenoviruses may induce immune toxicities ^[28][31]. In a non-viral method, three forms of the CRISPR/Cas9 system are now available for the delivery of a nucleoprotein complex into the nucleus: plasmid DNA, the RNA system of Cas9 mRNA and sgRNA, and the RNA-protein complex of sgRNA and Cas9 ribonucleoprotein (Cas9 RNP) (Figure 1B). In the plasmid delivery system, the CRISPR/Cas9 plasmid, termed pX330, was originally constructed ^[20][23]. The pX330 plasmid contains two expression cassettes, with the expression of sgRNA being driven under the U6 promoter in one and that of Cas9 being driven under the chicken β-actin promotor in the other. In addition, Cas9 is engineered to carry the nuclear localization signal, similar to SV40, which is required to transport the CRISPR/Cas9 system into the nucleus ^[29][32]. The RNA system is capable of controllable release into the cytoplasm. The Cas9 RNP system may skip the expression of the Cas9 protein and sgRNA in cellular events, but it also reduces the risk of off-target effects because it avoids overexpression. Although a gene delivery system, such as electroporation and nanoparticle transfer, may be applied to almost any cell type at any stage of the cell cycle, even large-molecule particles, low delivery efficiency remains a challenge due to the large molecular size, instability, and the low efficiency of the endosomal escape ^[30][31][33,34].

3. Current Approaches for Avian Genome Editing Based on the CRISPR/Cas9 System

Avian genome editing technology is based on a procedure for establishing transgenic chicken lines. Various approaches have been attempted to create transgenic birds, including viral infection, the chimeric method, and sperm-mediated gene transfer (Figure 2).

3.1. Viral Infection

Viral infection is the most reliable method because of its high efficiency for incorporation of the transgene. The viral gene delivery system was primarily applied to blastoderm stage-X embryos for the purpose of inducing the transgene into the genome of germline cells in the blastoderm ^{[32][33][34][35][36][37][38][39]}[36,37,38,39,40,41,42,43]. Salter et al. ^[32][36] was the first to attempt retrovirus transfection into a blastoderm. The infection of a replication-defective pantropic retrovirus vector based on Moloney murine leukemia virus (MoMLV) pseudotyped with the vesicular stomatitis virus G protein was subsequently attempted in Japanese quail stage-X embryos ^[33][37]. In that study, the viral vector sequence was detected in the tissue of all the hatched quails, and the efficiency of the germline transmission from G0 to G1 was very high (80%). In addition, the injection of the lentiviral vector derived from the lentivirus equine infectious anemia virus into stage-X embryos was successfully adopted to produce transgenic chickens ^[34][38]. Although the expression of the anti-prion single-chain Fv protein was successfully achieved in the eggs of transgenic chickens transfected with the MoMLV-based mouse stem cell virus, the expression level of this protein in the G2 generation decreased to less than that of the G0 founder due to a transgene silencing effect ^[35][39]. Therefore, the appropriate selection of key regulator regions for gene silencing or increases in transcriptional activity may be required in future studies that focus on the expression of reporter genes and analyze their functions. The generation of genome-edited quails was initially achieved using an adenovirus injection containing the CRISPR/Cas9 system targeting the melanophilin (MLPH) gene ^[40][44] (Figure 2A). Since MLPH functions in feather pigmentation, the feathers of the genome-edited quails were gray. In this study, germline chimeric G0 quails produced genome-edited progeny, and the efficiency of the germline transmission ranged between 2.4 and 10%, suggesting the efficacy of adenovirus infection as a gene delivery system to transduce the CRISPR/Cas9 system for proliferating blastodermal cells. In addition, myostatin gene-edited quails with a higher body weight and muscle mass were successfully produced using the same adenovirus injection ^[41][45]. The effects of the adenoviral transduction of the CRISPR/Cas9 system into local tissue were also demonstrated in chick leg muscle ^[42][46]. More recently, genome-edited ducklings were successfully produced by an adenoviral infection into blastoderms, indicating the efficacy of genome editing technology with adenoviruses in poultry and water birds other than chickens and quails ^[43][47].

3.2. Chimeric Method Using PGCs

Since the viral method may introduce a potentially hazardous infection, non-viral methods for the integration of exogenous DNA into the host genome are preferred. Germ cells are the sole source of the transmission of genetic information to the next generation; therefore, the use of PGCs is currently the most common method for cell-mediated gene transfer because of the higher competency of PGCs than other stem cells, such as embryonic stem cells and embryonic germ cells ^[44][45][46][50,51,52]. Chicken PGCs appear from the early blastoderm stage and are present as colonies in the central region of the blastoderm at stage X ^[47][48][49][53,54,55]. After migrating to the germinal crescent region, they circulate in embryonic blood vessels and settle in the embryonic gonads ^[50][56]. Due to the unique migratory features of avian PGCs, they may be isolated from various embryonic stages and subjected to long-term cultures without the loss of germ cell potency ^[51][57]. Wentworth et al. ^[52][58] reported that cultured PGCs settled in recipient gonads after their injection into embryonic blood vessels, resulting in the production of a germ-line chimera in chickens. Isolated gonadal PGCs retained the capacity to migrate and differentiate into mature gametes in the recipient embryo transplanted into their blood vessels ^[53][54][55][59,60,61]. Using the PGC transplantation method, van de Lavoir et al. ^[51][57] reported the successful expression of green fluorescent protein, the reporter gene, in germline chimeras. The transposon system has since been identified as the most efficient for transgenesis ^[56][57][62,63]. Transposon elements have the ability to change their positions within a genome, which mediates the integration of the intended foreign gene. Park and Han reported about 50% germline chimerisms using piggyBac and the Tol2 transposon ^[57][63]. Another method involved the direct injection of transposon plasmids into embryonic blood vessels with lipofectamine, with the transformation of the circulating PGCs and the subsequent production of transgenic chickens ^[58][64] (Figure 2B). Zhu et al. successfully integrated human immunoglobulin-coded genes in embryonic stem cells derived from stage X ^[59][65], indicating the utility of transgenic poultry in industries such as agriculture and biomedicine. Genome-editing technologies using the PGC method have been developed in the last decade for basic research on embryogenesis as well as for agriculture and biomedicine. In 2016, two egg-white genes, ovalbumin and ovomucoid, which are well-known allergenic proteins, were efficiently mutagenized in cultured chicken PGCs by the lipofection of the px330 plasmid and an ovomucoid gene-edited germline chimeric rooster was established ^[60][66] (Figure 2C). In the same year, transgenic chickens carrying a loxP site in the immunoglobulin heavy chain gene were successfully generated by PGCs using the CRISPR/Cas9-mediated homology-directed repair system (HDR) ^[61][67]. HDR is a cellular repair mechanism performed by the homologous recombination pathway that modifies the target genome sequence when an exogenous donor sequence is present at the sgRNA-targeted site in the CRISPR/Cas9 system. A germline chimera carrying a human interferon-β (hIFN-β) gene recombined in the ovalbumin locus was then generated by the CRISPR/Cas9-mediated HDR method, and high hIFN-β protein levels were produced in the egg white ^[62][68]. Even in basic research fields, a recent study successfully generated a genome-edited chicken targeting doublesex and mab-3-related transcription factor 1 (DMRT1) as the testis-determining gene ^[63][69].