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Current genomic editing approaches in avian transgenesis
General and Comparative Endocrinology 190 (2013) 144–148
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Current genomic editing approaches in avian transgenesis Tae Sub Park, Kyung Soo Kang, Jae Yong Han ⇑ WCU Biomodulation Major, Department of Agricultural Biotechnology, Research Institute for Agriculture and Life Sciences, College of Agriculture and Life Sciences, Seoul National University, Seoul 151-921, Republic of Korea
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Article history: Available online 14 December 2012 Keywords: Transgenesis Primordial germ cell Genomic editing Model animal
a b s t r a c t The chicken was domesticated from Red Jungle Fowl over 8000 years ago and became one of the major food sources worldwide. At present, the poultry industry is one of the largest industrial animal stocks in the world, and its economic scale is expanding significantly with increasing consumption. Additionally, since Aristotle used chicken eggs as a model to provide remarkable insights into how life begins, chickens have been used as invaluable and powerful experimental materials for studying embryo development, immune systems, biomedical processes, and hormonal regulation. Combined with advancements in efficient transgenic technology, avian models have become even more important than would have been expected. Ó 2012 Elsevier Inc. All rights reserved.
1. Introduction The chicken has several advantages as a model animal. Due to its oviparity, chicken embryos can be easily accessed and chemically processed to evaluate cellular and molecular differentiation mechanisms. In experimental procedures, chicken embryos can be treated and examined without maternal or external effects; therefore, researchers can expect and detect treatment effects alone in chicken embryos. Thus, chicken embryos provide the best model for investigating hormones and hormone regulatory networks during developmental embryonic stages. For large-scale experimental treatments, the developmental stages of chicken embryos can be easily synchronized without individual variation. In 2004, the chicken genome project was completed and chicken genomic sequences were released to the public domain [15]. Thus, genomic information and the structure of genes and target sites of interest can be readily retrieved from a Web-based database. Currently, the genomic sequences of the zebra finch and the turkey are also publicly accessible on the Internet and evolutionary comparisons among target genes can be made between the different avian species [9,56]. Finally, recent technical advances in avian transgenesis allow efficient extensions of experimental protocols and research areas, which will help provide a more comprehensive understanding of developmental processes and molecular mechanisms. 2. Aves as animal models An animal model is a living animal that is used to study human diseases as well as for basic research. To date, rodents are the most ⇑ Corresponding author. Fax: +82 2 874 4811. E-mail address: [email protected] (J.Y. Han). 0016-6480/$ - see front matter Ó 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ygcen.2012.11.020
commonly used animals but they have several limitations such as the difficult access to living mouse fetus and the indispensable maternal influences to mouse fetus [48]. Thus, novel animal models should be developed for accurate and efficient assessments in various species. In particular, chicken and quail have been used in various studies as an alternative animal model. In studies on developmental endocrinology, chicken embryos have been used to examine cellular differentiation and the maturation of endocrine glands, and ontogenic alterations in embryonic glands and their target organs [10]. Gene expression patterns have been analyzed during the functional development of embryonic glands [43], and signaling factors have been treated to verify their effects on ontogenic changes in chicken embryonic glands [2]. These studies demonstrated that the gene expression patterns and ontogeny that were observed in chicken and mouse showed high similarity; therefore, the chicken can serve as an excellent model to validate the genetic and molecular mechanisms underlying gland development. In hens, the epithelial cells of the oviduct produce large quantities of egg-white protein in daily cycles. Thus, the chicken oviduct system has a considerable advantage as a model system for studying the hormonal regulation of specific gene expression. The chicken oviduct system has been extensively examined in vivo and in vitro to study hormonal responsiveness and regulation for cell growth, gene expression, and ovarian cancer [20]. To investigate harmoniously orchestrated interactions between steroid hormones and gene expression for egg-white proteins, we developed in vitro culturing procedures for chicken oviduct epithelial cells and identified a cascade of events in cultured oviductal cells after hormone induction [20]. In 2010, Sato et al. demonstrated the vascular morphogenetic process using transgenic quail embryos [45]. Transgenic quail that
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had a yellow fluorescent protein (YFP) gene that was controlled by the mouse Tie1 promoter were generated by lentiviral transduction. Transgenic (tie1:H2B-eYFP) quail embryos expressed the reporter YFP in their endothelial cells, and early embryonic vascular morphogenesis was investigated using a time-lapse dynamic imaging system. The study demonstrated that avian transgenic technology represents a powerful approach for addressing the challenge of studying the mechanical, molecular, and cellular mechanisms involved in vascular development [45]. An alternative experimental bird, the zebra finch, is considered a suitable model for studying vocal learning processes. Recently, transgenic songbirds were successfully generated using lentiviral transduction [1]; this offers a great opportunity to develop an alternative genetic model for vocal learning.
3. Strategic approaches in avian transgenesis To create transgenic birds, various approaches have been tried, essentially starting by adapting techniques that were established in mammals. However, the production of transgenic birds through germline transmission either failed or the transgenesis efficiency was too low because of evolutionary and physiological differences between aves and mammals. Love et al. [25] microinjected plasmid DNA into the germinal disc of fertilized chick embryos. However, the direct microinjection technique is not currently used due to its low efficiency relative to the large number of hens that need to be killed to collect fertilized eggs. In ovo electroporation into chicken embryos was an alternative and useful method for introducing transgenesis in specific tissues during early developmental stages [19]. The in ovo electroporation technique allowed gene transfers into living embryos to analyze gene function and molecular mechanisms during embryo development. Nevertheless, this technique is only suitable for temporal transgene expression within small windows of developmental stages and can be used with regionally restricted expression in chicken embryos. Producing transgenic offspring via in ovo electroporation would be almost impossible. The most reliable method for generating transgenic chickens is virus-mediated transgenesis, regardless of whether it involves infection into blastoderms or germ cells [32,35,38,44,46]. Salter et al. [44] first applied retrovirus transduction to chicken blastoderms at stage X to produce transgenic chickens. Transgenic quail were produced by injecting a replication-defective pantropic retroviral vector based on Moloney murine leukemia virus (MoMLV) that was pseudotyped with vesicular stomatitis virus G protein (VSV-G) into the quail blastodermal layer [32]. However, in the this study, the reporter expression of the green fluorescent protein (GFP) gene that was controlled by the MoMLV long-terminal repeat (LTR) promoter was not detected because of a transgene silencing effect [32]. Retrovirus- and lentivirus-transduced primordial germ cells (PGCs) have been transplanted into recipient embryos to generate transgenic chickens and quail [35,38,46]. Compared with blastoderm injection protocols in which somatic cells and germ cells are infected by transgene-containing viruses, germ cell-targeted transduction is a more efficient tool because the germ cell is the only germline-transmittable cell type that affects the next generation. Virus transduction is a versatile and useful protocol but a virusindependent transgene delivery system is necessary due to safety issues related to infectious virus reconstruction in the host. Thus, nonviral transfection into germline-competent cells by lipofection or electroporation would represent the best option for avian transgenesis. PGC is a precursor cell of germ cells during embryo development. In the chicken, PGCs arise from the center area of the epiblast in the blastoderm, termed the area pellucida; they subse-
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quently move into the germinal crescent of an extraembryonic site [50]. At the beginning of blood vessel formation, PGCs migrate to the embryonic genital ridges through blood circulation. These PGCs can be retrieved from different embryonic sites at different stages: the germinal crescent, embryonic blood vessel, and embryonic gonads [18]. Since PGCs finally differentiate into sperm through the spermatogenetic process in male testis or into oocytes through oogenesis in the female ovary after sexual maturation, PGCs are considered a candidate vehicle for delivering transgenes to the next generation (Fig. 1). For germline transmission, Wentworth et al. [57] first characterized and manipulated quail PGCs to produce germline chimeric quial that have both their own (endogenous) germ cells and transferred (exogenous) germ cells. In the chicken, Tajima et al. [51] isolated circulating PGCs from blood vessels of embryos at stages 13–14 and transferred those PGCs into the blood vessels of recipient embryos at stages 14–15. The germline transmission efficiency ranged from 0.0% to 11.8%. As another source, chicken PGCs were isolated from embryonic gonads at 6 days, at which time PGCs had already completed their migration through the blood vessels. Notably, gonadal PGCs (gPGCs) retain the capacity to migrate into genital ridges after introduction into blood vessels in recipient embryos [6]. The manipulation of gPGCs extended the use of PGCs for in vitro culture as well as germline chimera production because of the ease of isolation and the relatively large number of PGCs that can be harvested from embryonic gonads (Fig. 1). In the first report of germline chimera production using gPGCs, the efficiency of germline transmission was between 1.3% and 3.1% [6]. Park et al. [39] advanced the short-term culture technique for chicken gPGCs and improved the germline transmission efficiency to 10.7–49.7%. More recently, a chicken PGC in vitro culture system has been established [7,26,55]. Using cultured chicken PGCs that were isolated from embryonic blood vessels, van de Lavoir et al. [55] successfully produced transgenic chickens as well as germline chimeras after genetic modification and transplantation. In the last year, two separate groups reported the combination of chicken PGC culture with transfection of transposon elements, piggyBac and Tol2, to enhance stable genomic integration and the expression of transgenes for transgenic production [27,40]. Transposon elements that are mediated by the transposase catalytic process can be used in strategic applications to achieve efficient avian genomic modification. Using piggyBac-mediated gene transfer into chicken PGCs, we demonstrated that the efficiency of germline transmission of donor PGCs after piggyBac transposition was 95.2% on average, and as was expected, half of the donor-derived offspring (52.2%) were transgenic due to heterozygous transgenes in the donor PGCs [40]. The transposon-mediated approach can be widely adapted to transgenesis in other avian species without running risks such as using viral vectors. Other germline-competent cells, specifically blastodermal cells, embryonic stem (ES) cells, embryonic germ (EG) cells, and spermatogonial cells, have also been used to transmit manipulated cells into the next generation, but the efficiency of germline transmission is generally too low compared with PGC techniques [21,37,41,42].
4. Specific genetic modifications in the chicken genome 4.1. Overexpression and knockdown To overexpress specific genes, constitutive and strong promoters have been used in various species, including aves. One strong promoter was isolated from the Rous sarcoma virus (RSV), which was the first oncovirus and one that causes sarcoma in chickens. Since Gorman et al. [13] verified the strong transcriptional activity
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Fig. 1. Summary of germline chimera production and transgenesis strategies in aves.
of RSV LTR in different cell lines, many researchers have used this strong promoter to achieve overexpression of specific genes in different species. The immediate early cytomegalovirus (CMV) promoter is the most frequently used promoter. Chicken PGCs that were transfected with a CMV-GFP or CMV-DsRed expression vector showed high and stable expression of exogenous transgenes without gene silencing (Fig. 2). In addition, we generated transgenic chickens with CMV-GFP transgene flanked with piggyBac transposons; various tissues constantly expressed GFP in the transgenic offspring [40]. The promoter of the CMV early enhancer element and the chicken b-actin promoter are frequently used in combination to achieve high levels of specific gene expression in transgenic animals [36]. RNA interference (RNAi) is a valuable research tool that can be used to knock down specific transcripts, both in cell culture and in living organisms [49]. RNAi is an RNA-dependent silencing process that is modulated by the RNA-induced silencing complex (RISC) [18,22]; the RNAi pathway is found in many eukaryotes, including avian species. Basically, two types of small RNA, microRNA (miRNA) and small interfering RNA (siRNA), are the major RNAi molecules used to attenuate gene expression [22,49]. We applied RNAi knockdown systems to chicken PGCs as well as chicken cell lines and efficiently downregulated the target transcripts [23]. Overexpression and RNAi knockdown techniques can also be combined with tissue- or stage-specific promoters and inducible expression promoters to achieve spatiotemporal expression. Using combinations of regulatory elements, the initiation or termination of exogenous transgene expression and the silencing of endogenous genes can be controlled in a tissue-specific and time-dependent manner. 4.2. Gene targeting Gene targeting is a genetic modification technique that can be used to knock out or knock in a specific gene by homologous recombination. In 1987, Thomas and Capecchi [53] first succeeded in generating a knockout mouse for the endogenous hypoxanthine phosphoribosyl transferase (HPRT) gene using mouse embryonic stem cells (ES cells) [53]. Currently, site-specific mutagenesis by gene targeting has become one of the major tools used to study functional genomics in various species. In particular, somatic cell
nuclear transfer (SCNT) techniques allow scientists to efficiently create specific site-mutated lines in various mammals, including mice. However, due to the lack of germline-competent ES cells and the difficulty of adapting SCNT techniques to avian species, successful gene targeting by homologous recombination has not been reported. Currently, chicken PGCs can be successfully maintained in vitro without the loss of germ cell capacity [7,26,55]; therefore, it is possible to generate gene-targeted chickens using cultured chicken PGCs. Recently, novel approaches have been developed to achieve efficient and specific genomic modifications: zinc-finger nuclease (ZFN) and transcription activator-like effector nuclease (TALEN). State-of-the-art technical platforms for ZFN and TALEN represent next-generation tools for customized genomic editing in transgenic animals as well as cultured cells in vitro. ZFN consists of a sequence-specific zinc-finger DNA-binding domain and engineered Fok I endonucleases [5,30]. Similar to ZFN, TALENs are generated by fusing the TAL effector of a DNA-binding domain and a DNA cleavage nuclease domain; it is an indispensable genomic editing tool that can be used to modify complex genomes for applied biotechnology as well as functional studies. Transcription activatorlike effector (TALE) is a transcription factor that was first discovered in the plant pathogen bacteria Xanthomonas. In plant cells, it is localized in the nucleus and binds specific sequences of gene promoters to modulate host resistance mechanisms [17,52]. Thus, synthetic transcription factors that use TALE domains can be used for gene expression regulation by binding to the promoter region of a specific gene [4,31,34,59]. After DNA sequence-specific binding, the fused catalytic nuclease Fok I domain cleaves and disrupts the targeted genome site. In both ZFN and TALEN units, Fok I endonuclease generates a DNA double-strand break (DSB); subsequently, the damaged DSB is repaired by homologous recombination (HR) or nonhomologous end-joining (NHEJ) processes [8,24]. During these repair processes, deletion or addition of several nucleotides occurs to create mutagenesis or to generate a frameshift in the open reading frame of the specific target gene [8,24]. The advantages of ZFN and TALEN technologies for specific genomic editing are their high precision and high efficiency, their versatile application to various animal species, and the lack of genomic integration of exogenous transgenes, even in genetically modified animals.
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Fig. 2. G418-selected chicken PGCs after transfection with a piggyBac CMV-GFP (A) or a piggyBac CMV-DsRed expression vector (B). Left panels in (A) and (B) are phasecontrast images; right images are fluorescent images. (Scale bar = 50 lm).
ZFNs have been used to generate genome modifications in plants [47,54], Drosophila [3], and Caenorhabditis elegans [33]. More recently, these targeted genome editing approaches were adapted to vertebrates such as zebrafish [11,29] and rats [12,28,52]. In addition, Hockemeyer et al. [16] demonstrated that the ZFN-mediated approach can be used to efficiently modify the genome of human embryonic stem cells (hES cells) and human induced pluripotent stem cells (hiPS cells) [16]. Additionally, Hockemeyer et al. [17] accomplished genomic locus-specific insertions of transgenic cassettes in hES cells and hiPS cells using TALENs and generated distinct architectures in both stem cell lines [17]. Thus, in human stem cells, both ZFN and TALEN are useful for generating genetic tools to study cell fate decisions and cell type-specific reporter systems to improve hESC differentiation protocols [16,17]. In domestic animals, Hauschild et al. [14] reported the generation of biallelic knockout pigs using ZFN. For xenotransplantation, they disrupted the porcine a1,3-galactosyltransferase (GGTA1) gene in porcine fibroblasts and subsequently produced knockout pigs using the SCNT technique. In cattle, Yu et al. [58] modified the blactoglobulin gene using specific zinc-finger nucleases. In the future, combined with chicken PGC culturing techniques, it may be possible to generate specific genomic locus-targeted chickens using ZFN and TALEN.
5. Conclusions Chicken embryos are an excellent model for studying developmental endocrinology, and the application of transgenic techniques to chicken model development will help provide a comprehensive understanding of the regulatory processes and the complicated networks in avian endocrinology. Furthermore, state-of-the-art approaches for specific genomic editing, such as
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General and Comparative Endocrinology journal homepage: www.elsevier.com/locate/ygcen
Current genomic editing approaches in avian transgenesis Tae Sub Park, Kyung Soo Kang, Jae Yong Han ⇑ WCU Biomodulation Major, Department of Agricultural Biotechnology, Research Institute for Agriculture and Life Sciences, College of Agriculture and Life Sciences, Seoul National University, Seoul 151-921, Republic of Korea
a r t i c l e
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Article history: Available online 14 December 2012 Keywords: Transgenesis Primordial germ cell Genomic editing Model animal
a b s t r a c t The chicken was domesticated from Red Jungle Fowl over 8000 years ago and became one of the major food sources worldwide. At present, the poultry industry is one of the largest industrial animal stocks in the world, and its economic scale is expanding significantly with increasing consumption. Additionally, since Aristotle used chicken eggs as a model to provide remarkable insights into how life begins, chickens have been used as invaluable and powerful experimental materials for studying embryo development, immune systems, biomedical processes, and hormonal regulation. Combined with advancements in efficient transgenic technology, avian models have become even more important than would have been expected. Ó 2012 Elsevier Inc. All rights reserved.
1. Introduction The chicken has several advantages as a model animal. Due to its oviparity, chicken embryos can be easily accessed and chemically processed to evaluate cellular and molecular differentiation mechanisms. In experimental procedures, chicken embryos can be treated and examined without maternal or external effects; therefore, researchers can expect and detect treatment effects alone in chicken embryos. Thus, chicken embryos provide the best model for investigating hormones and hormone regulatory networks during developmental embryonic stages. For large-scale experimental treatments, the developmental stages of chicken embryos can be easily synchronized without individual variation. In 2004, the chicken genome project was completed and chicken genomic sequences were released to the public domain [15]. Thus, genomic information and the structure of genes and target sites of interest can be readily retrieved from a Web-based database. Currently, the genomic sequences of the zebra finch and the turkey are also publicly accessible on the Internet and evolutionary comparisons among target genes can be made between the different avian species [9,56]. Finally, recent technical advances in avian transgenesis allow efficient extensions of experimental protocols and research areas, which will help provide a more comprehensive understanding of developmental processes and molecular mechanisms. 2. Aves as animal models An animal model is a living animal that is used to study human diseases as well as for basic research. To date, rodents are the most ⇑ Corresponding author. Fax: +82 2 874 4811. E-mail address: [email protected] (J.Y. Han). 0016-6480/$ - see front matter Ó 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ygcen.2012.11.020
commonly used animals but they have several limitations such as the difficult access to living mouse fetus and the indispensable maternal influences to mouse fetus [48]. Thus, novel animal models should be developed for accurate and efficient assessments in various species. In particular, chicken and quail have been used in various studies as an alternative animal model. In studies on developmental endocrinology, chicken embryos have been used to examine cellular differentiation and the maturation of endocrine glands, and ontogenic alterations in embryonic glands and their target organs [10]. Gene expression patterns have been analyzed during the functional development of embryonic glands [43], and signaling factors have been treated to verify their effects on ontogenic changes in chicken embryonic glands [2]. These studies demonstrated that the gene expression patterns and ontogeny that were observed in chicken and mouse showed high similarity; therefore, the chicken can serve as an excellent model to validate the genetic and molecular mechanisms underlying gland development. In hens, the epithelial cells of the oviduct produce large quantities of egg-white protein in daily cycles. Thus, the chicken oviduct system has a considerable advantage as a model system for studying the hormonal regulation of specific gene expression. The chicken oviduct system has been extensively examined in vivo and in vitro to study hormonal responsiveness and regulation for cell growth, gene expression, and ovarian cancer [20]. To investigate harmoniously orchestrated interactions between steroid hormones and gene expression for egg-white proteins, we developed in vitro culturing procedures for chicken oviduct epithelial cells and identified a cascade of events in cultured oviductal cells after hormone induction [20]. In 2010, Sato et al. demonstrated the vascular morphogenetic process using transgenic quail embryos [45]. Transgenic quail that
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had a yellow fluorescent protein (YFP) gene that was controlled by the mouse Tie1 promoter were generated by lentiviral transduction. Transgenic (tie1:H2B-eYFP) quail embryos expressed the reporter YFP in their endothelial cells, and early embryonic vascular morphogenesis was investigated using a time-lapse dynamic imaging system. The study demonstrated that avian transgenic technology represents a powerful approach for addressing the challenge of studying the mechanical, molecular, and cellular mechanisms involved in vascular development [45]. An alternative experimental bird, the zebra finch, is considered a suitable model for studying vocal learning processes. Recently, transgenic songbirds were successfully generated using lentiviral transduction [1]; this offers a great opportunity to develop an alternative genetic model for vocal learning.
3. Strategic approaches in avian transgenesis To create transgenic birds, various approaches have been tried, essentially starting by adapting techniques that were established in mammals. However, the production of transgenic birds through germline transmission either failed or the transgenesis efficiency was too low because of evolutionary and physiological differences between aves and mammals. Love et al. [25] microinjected plasmid DNA into the germinal disc of fertilized chick embryos. However, the direct microinjection technique is not currently used due to its low efficiency relative to the large number of hens that need to be killed to collect fertilized eggs. In ovo electroporation into chicken embryos was an alternative and useful method for introducing transgenesis in specific tissues during early developmental stages [19]. The in ovo electroporation technique allowed gene transfers into living embryos to analyze gene function and molecular mechanisms during embryo development. Nevertheless, this technique is only suitable for temporal transgene expression within small windows of developmental stages and can be used with regionally restricted expression in chicken embryos. Producing transgenic offspring via in ovo electroporation would be almost impossible. The most reliable method for generating transgenic chickens is virus-mediated transgenesis, regardless of whether it involves infection into blastoderms or germ cells [32,35,38,44,46]. Salter et al. [44] first applied retrovirus transduction to chicken blastoderms at stage X to produce transgenic chickens. Transgenic quail were produced by injecting a replication-defective pantropic retroviral vector based on Moloney murine leukemia virus (MoMLV) that was pseudotyped with vesicular stomatitis virus G protein (VSV-G) into the quail blastodermal layer [32]. However, in the this study, the reporter expression of the green fluorescent protein (GFP) gene that was controlled by the MoMLV long-terminal repeat (LTR) promoter was not detected because of a transgene silencing effect [32]. Retrovirus- and lentivirus-transduced primordial germ cells (PGCs) have been transplanted into recipient embryos to generate transgenic chickens and quail [35,38,46]. Compared with blastoderm injection protocols in which somatic cells and germ cells are infected by transgene-containing viruses, germ cell-targeted transduction is a more efficient tool because the germ cell is the only germline-transmittable cell type that affects the next generation. Virus transduction is a versatile and useful protocol but a virusindependent transgene delivery system is necessary due to safety issues related to infectious virus reconstruction in the host. Thus, nonviral transfection into germline-competent cells by lipofection or electroporation would represent the best option for avian transgenesis. PGC is a precursor cell of germ cells during embryo development. In the chicken, PGCs arise from the center area of the epiblast in the blastoderm, termed the area pellucida; they subse-
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quently move into the germinal crescent of an extraembryonic site [50]. At the beginning of blood vessel formation, PGCs migrate to the embryonic genital ridges through blood circulation. These PGCs can be retrieved from different embryonic sites at different stages: the germinal crescent, embryonic blood vessel, and embryonic gonads [18]. Since PGCs finally differentiate into sperm through the spermatogenetic process in male testis or into oocytes through oogenesis in the female ovary after sexual maturation, PGCs are considered a candidate vehicle for delivering transgenes to the next generation (Fig. 1). For germline transmission, Wentworth et al. [57] first characterized and manipulated quail PGCs to produce germline chimeric quial that have both their own (endogenous) germ cells and transferred (exogenous) germ cells. In the chicken, Tajima et al. [51] isolated circulating PGCs from blood vessels of embryos at stages 13–14 and transferred those PGCs into the blood vessels of recipient embryos at stages 14–15. The germline transmission efficiency ranged from 0.0% to 11.8%. As another source, chicken PGCs were isolated from embryonic gonads at 6 days, at which time PGCs had already completed their migration through the blood vessels. Notably, gonadal PGCs (gPGCs) retain the capacity to migrate into genital ridges after introduction into blood vessels in recipient embryos [6]. The manipulation of gPGCs extended the use of PGCs for in vitro culture as well as germline chimera production because of the ease of isolation and the relatively large number of PGCs that can be harvested from embryonic gonads (Fig. 1). In the first report of germline chimera production using gPGCs, the efficiency of germline transmission was between 1.3% and 3.1% [6]. Park et al. [39] advanced the short-term culture technique for chicken gPGCs and improved the germline transmission efficiency to 10.7–49.7%. More recently, a chicken PGC in vitro culture system has been established [7,26,55]. Using cultured chicken PGCs that were isolated from embryonic blood vessels, van de Lavoir et al. [55] successfully produced transgenic chickens as well as germline chimeras after genetic modification and transplantation. In the last year, two separate groups reported the combination of chicken PGC culture with transfection of transposon elements, piggyBac and Tol2, to enhance stable genomic integration and the expression of transgenes for transgenic production [27,40]. Transposon elements that are mediated by the transposase catalytic process can be used in strategic applications to achieve efficient avian genomic modification. Using piggyBac-mediated gene transfer into chicken PGCs, we demonstrated that the efficiency of germline transmission of donor PGCs after piggyBac transposition was 95.2% on average, and as was expected, half of the donor-derived offspring (52.2%) were transgenic due to heterozygous transgenes in the donor PGCs [40]. The transposon-mediated approach can be widely adapted to transgenesis in other avian species without running risks such as using viral vectors. Other germline-competent cells, specifically blastodermal cells, embryonic stem (ES) cells, embryonic germ (EG) cells, and spermatogonial cells, have also been used to transmit manipulated cells into the next generation, but the efficiency of germline transmission is generally too low compared with PGC techniques [21,37,41,42].
4. Specific genetic modifications in the chicken genome 4.1. Overexpression and knockdown To overexpress specific genes, constitutive and strong promoters have been used in various species, including aves. One strong promoter was isolated from the Rous sarcoma virus (RSV), which was the first oncovirus and one that causes sarcoma in chickens. Since Gorman et al. [13] verified the strong transcriptional activity
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Fig. 1. Summary of germline chimera production and transgenesis strategies in aves.
of RSV LTR in different cell lines, many researchers have used this strong promoter to achieve overexpression of specific genes in different species. The immediate early cytomegalovirus (CMV) promoter is the most frequently used promoter. Chicken PGCs that were transfected with a CMV-GFP or CMV-DsRed expression vector showed high and stable expression of exogenous transgenes without gene silencing (Fig. 2). In addition, we generated transgenic chickens with CMV-GFP transgene flanked with piggyBac transposons; various tissues constantly expressed GFP in the transgenic offspring [40]. The promoter of the CMV early enhancer element and the chicken b-actin promoter are frequently used in combination to achieve high levels of specific gene expression in transgenic animals [36]. RNA interference (RNAi) is a valuable research tool that can be used to knock down specific transcripts, both in cell culture and in living organisms [49]. RNAi is an RNA-dependent silencing process that is modulated by the RNA-induced silencing complex (RISC) [18,22]; the RNAi pathway is found in many eukaryotes, including avian species. Basically, two types of small RNA, microRNA (miRNA) and small interfering RNA (siRNA), are the major RNAi molecules used to attenuate gene expression [22,49]. We applied RNAi knockdown systems to chicken PGCs as well as chicken cell lines and efficiently downregulated the target transcripts [23]. Overexpression and RNAi knockdown techniques can also be combined with tissue- or stage-specific promoters and inducible expression promoters to achieve spatiotemporal expression. Using combinations of regulatory elements, the initiation or termination of exogenous transgene expression and the silencing of endogenous genes can be controlled in a tissue-specific and time-dependent manner. 4.2. Gene targeting Gene targeting is a genetic modification technique that can be used to knock out or knock in a specific gene by homologous recombination. In 1987, Thomas and Capecchi [53] first succeeded in generating a knockout mouse for the endogenous hypoxanthine phosphoribosyl transferase (HPRT) gene using mouse embryonic stem cells (ES cells) [53]. Currently, site-specific mutagenesis by gene targeting has become one of the major tools used to study functional genomics in various species. In particular, somatic cell
nuclear transfer (SCNT) techniques allow scientists to efficiently create specific site-mutated lines in various mammals, including mice. However, due to the lack of germline-competent ES cells and the difficulty of adapting SCNT techniques to avian species, successful gene targeting by homologous recombination has not been reported. Currently, chicken PGCs can be successfully maintained in vitro without the loss of germ cell capacity [7,26,55]; therefore, it is possible to generate gene-targeted chickens using cultured chicken PGCs. Recently, novel approaches have been developed to achieve efficient and specific genomic modifications: zinc-finger nuclease (ZFN) and transcription activator-like effector nuclease (TALEN). State-of-the-art technical platforms for ZFN and TALEN represent next-generation tools for customized genomic editing in transgenic animals as well as cultured cells in vitro. ZFN consists of a sequence-specific zinc-finger DNA-binding domain and engineered Fok I endonucleases [5,30]. Similar to ZFN, TALENs are generated by fusing the TAL effector of a DNA-binding domain and a DNA cleavage nuclease domain; it is an indispensable genomic editing tool that can be used to modify complex genomes for applied biotechnology as well as functional studies. Transcription activatorlike effector (TALE) is a transcription factor that was first discovered in the plant pathogen bacteria Xanthomonas. In plant cells, it is localized in the nucleus and binds specific sequences of gene promoters to modulate host resistance mechanisms [17,52]. Thus, synthetic transcription factors that use TALE domains can be used for gene expression regulation by binding to the promoter region of a specific gene [4,31,34,59]. After DNA sequence-specific binding, the fused catalytic nuclease Fok I domain cleaves and disrupts the targeted genome site. In both ZFN and TALEN units, Fok I endonuclease generates a DNA double-strand break (DSB); subsequently, the damaged DSB is repaired by homologous recombination (HR) or nonhomologous end-joining (NHEJ) processes [8,24]. During these repair processes, deletion or addition of several nucleotides occurs to create mutagenesis or to generate a frameshift in the open reading frame of the specific target gene [8,24]. The advantages of ZFN and TALEN technologies for specific genomic editing are their high precision and high efficiency, their versatile application to various animal species, and the lack of genomic integration of exogenous transgenes, even in genetically modified animals.
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Fig. 2. G418-selected chicken PGCs after transfection with a piggyBac CMV-GFP (A) or a piggyBac CMV-DsRed expression vector (B). Left panels in (A) and (B) are phasecontrast images; right images are fluorescent images. (Scale bar = 50 lm).
ZFNs have been used to generate genome modifications in plants [47,54], Drosophila [3], and Caenorhabditis elegans [33]. More recently, these targeted genome editing approaches were adapted to vertebrates such as zebrafish [11,29] and rats [12,28,52]. In addition, Hockemeyer et al. [16] demonstrated that the ZFN-mediated approach can be used to efficiently modify the genome of human embryonic stem cells (hES cells) and human induced pluripotent stem cells (hiPS cells) [16]. Additionally, Hockemeyer et al. [17] accomplished genomic locus-specific insertions of transgenic cassettes in hES cells and hiPS cells using TALENs and generated distinct architectures in both stem cell lines [17]. Thus, in human stem cells, both ZFN and TALEN are useful for generating genetic tools to study cell fate decisions and cell type-specific reporter systems to improve hESC differentiation protocols [16,17]. In domestic animals, Hauschild et al. [14] reported the generation of biallelic knockout pigs using ZFN. For xenotransplantation, they disrupted the porcine a1,3-galactosyltransferase (GGTA1) gene in porcine fibroblasts and subsequently produced knockout pigs using the SCNT technique. In cattle, Yu et al. [58] modified the blactoglobulin gene using specific zinc-finger nucleases. In the future, combined with chicken PGC culturing techniques, it may be possible to generate specific genomic locus-targeted chickens using ZFN and TALEN.
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