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Reproductive Technology |
Department of Aquatic Biosciences,3 Tokyo University of Fisheries, Minato-ku, Tokyo 108-8477, Japan
PREST, Japan Science and Technology Corporation4, Kawaguchi-shi, Saitama, Japan
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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assisted reproductive technology, developmental biology, early development, embryo, gametogenesis
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In mice, blastocyst-derived embryonic stem (ES) cells contribute to all cell lineages, including the germ cells [4]. Genetically modified mouse ES cells can differentiate into functional gametes in the gonads of chimeric mouse after they have been injected into the blastocyst of the recipient. This technique eventually results in the production of individual mice carrying modified genomes through fertilization. However, despite numerous attempts in zebrafish and medaka, blastomere-derived fish ES-like cells that can contribute to the germ cell lineage of recipients following long-term cultivation have not yet been established, even though they can differentiate into various somatic cell lineages [5]. It is obvious that the single most important characteristic required in donor cells for cell-mediated gene transfer is that they have the potential to become functional gametes. Therefore, to establish a cell line retaining the ability to differentiate into gametes, we focused on primordial germ cells (PGCs), which are the embryonic precursors of germline cells [6].
In the present study, we intended to establish a technique to produce live offspring from isolated PGCs by their transplantation into developing embryos. To convert donor PGCs into functional gametes in the recipient, the transplanted cells must reenter the germline of the recipient. As in many other organisms, fish PGCs emerge in extragonadal areas and then migrate to the genital ridges [7], where they proliferate and develop into gametes. We hypothesized that for the successful production of donor-derived offspring, the donor PGCs must pass through four stages: 1) migration from the site of transplantation to the genital ridges, 2) coalescence with the somatic cells of the genital ridges, 3) avoidance of long-term immune rejection in the recipient, and 4) differentiation into functional gametes. Therefore, the experiments described here were designed to allow transplanted PGCs to participate in the migratory movements of the endogenous PGCs.
Recently, we generated transgenic rainbow trout strains in which green fluorescent protein (GFP) was expressed in PGCs under the control of a rainbow trout vasa-like gene (RtVLG) promoter [8]. Because GFP expression was specific to the PGCs, these cells could be visualized by green fluorescence in the live trout embryos. We also established a system for the mass isolation of viable GFP-labeled PGCs by flow cytometry [9]. The rainbow trout has strong advantages as a subject for PGC manipulation compared to other fish species. For example, the genital ridges, containing PGCs, can be readily isolated from hatched embryos using forceps and a dissecting microscope as a result of the large embryo size (total body length at hatching, 15 mm). Also, the high fecundity of this species allows thousands of transgenic embryos containing GFP-labeled PGCs to be produced from a single insemination. Because fish PGCs are present in very small numbers when they settle in the genital ridges [10], high fecundity is important for the mass preparation of PGCs for use in transplantation or in vitro culture experiments.
In the present study, we developed a method to transplant PGCs into developing embryos to incorporate them into the germ cell lineage of the recipient using the GFP-labeled PGCs as donor cells. Donor PGCs harvested from genital ridges of hatching embryos were transplanted into various stages of recipient embryos by microinjection.
By histological observations, trout PGCs are firstly recognized in the lateral mesoderm at the neurula stage (7 days postfertilization ["f]) [11]. Then, PGCs start to migrate to the dorsal side of the embryo and lie near the mesonephric ducts or above the gut. During the eyed stage (20–30 dpf), the PGCs located under the mesonephric ducts or in the dorsal mesentery migrate toward the areas of future genital ridges along the peritoneal wall and coalesce with somatic gonadal support cells. At the hatching stage (36 dpf), most of the PGCs are settled within the genital ridges; however, some PGCs are occasionally observed on the peritoneal wall or dorsal mesentery [12]. According to these observations, we injected donor PGCs into the sites where endogenous PGCs were located. After the PGC transplantation, the colonization efficiency of donor PGCs in the genital ridges of the recipients was evaluated by the appearance of GFP-labeled cells. To obtain functional gametes derived from donor PGCs, we investigated whether donor-derived cells could resume gametogenesis in the gonads of the recipients. Furthermore, the production of donor-derived fry was examined through both the inheritance of the transgene and the inheritance of body color in the F1 offspring.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To enable the PGCs to be visualized in the live trout embryos, the transgene (pvasa-GFP) was constructed using a 4.7-kilobase (kb) 5' fragment of RtVLG, an enhanced GFP gene (Clontech, Palo Alto, CA); a 0.6-kb 3' untranslated region; and a 1.5-kb 3' flanking region [8]. The pvasa-GFP transgenic rainbow trout were established by a method previously described [9] and were maintained in the Oizumi Research Station, Tokyo University of Fisheries (Yamanashi, Japan). The F2 transgenic embryos were generated by crossing pvasa-GFP hemizygous, wild type-colored males (pvasa-GFP/-; wt/wt) with nontransgenic, dominant orange-colored mutant females (-/-; OR/OR). On hatching of the F2 generation, transgenic orange-colored embryos (pvasa-GFP/-; OR/wt) expressing GFP in the PGCs were screened under the fluorescent microscope (BX-50 with a BX-FLA attachment and GFP filter set; Olympus, Tokyo, Japan) and used for the preparation of donor PGCs. All embryos and fish were reared at 10°C.
Donor Cell Preparation
Donor cells were prepared from genital ridges containing GFP-labeled PGCs excised from hatched embryos at 35, 40, and 45 dpf according the procedure described by Takeuchi et al. [9]. Briefly, for each experiment, 20 pairs of excised genital ridges were incubated in 0.5% trypsin solution (pH 8.2; Worthington Biochemical Corp., Lakewood, NJ) for 2 h at 20°C. After the trypsin treatment, the medium was substituted for Dulbecco modified Eagle medium (Life Technologies, Rockville, MD) with 10% fetal calf serum. Pipetting was used to physically disperse any remaining intact portions of the genital ridges, and the cells were stored on ice until the time of injection.
Recipient Embryos and Cell Transplantation Procedure
Glass micropipettes, originally devised for the transplantation of blastomeres into the blastodiscs of trout embryos [13], were used to transplant the PGCs. To retard the adhesion of cells, the tips of the micropipettes were coated with Sigmacote (Sigma, St. Louis, MO), rinsed with water, and dried. The PGCs were distinguished from gonadal somatic cells by their green fluorescence under the fluorescent dissecting microscope (SZX-12 with an SZX-RFL attachment and a GFP filter set; Olympus); therefore, they were easily identified and isolated (Fig. 1A). Between 5 and 10 PGCs were injected into each recipient embryo. A few dozen somatic cells were also included in the injected cell suspensions, because unsorted genital ridge cells were used for the transplantation. Transplantations to early stage embryos were performed as previously reported [13]. Donor PGCs were transplanted into the lower part of the blastoderm of the blastula and the embryonic shield of the gastrula. When hatched embryos were used as recipients, individuals were anesthetized and the donor PGCs injected into the peritoneal cavity (Fig. 1B).
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Analysis of the Developmental Fate of Transplanted PGCs
Recipient embryos were observed under the fluorescent microscope between Day 1 and Day 30 to evaluate the incorporation rate of donor PGCs into the genital ridges. All transplantation experiments were performed using at least 30 embryos each and repeated at least three times. The data are represented as the mean ± SEM. Data were analyzed by one-way ANOVA followed by Duncan multiple-range test. The proliferation and differentiation of the donor-derived germ cells in the gonads of the recipients were observed at Day 180 posttransplantation. Furthermore, to confirm the distribution of donor-derived cells possessing the GFP gene in the recipient fish, genomic DNA was extracted from the gill, kidney, spleen, liver, muscle, intestine, and gonad and analyzed by PCR with GFP-specific primers as previously described [14].
Test for Germline Transmission of Donor-Derived Phenotypes
Transplantation of PGCs was performed to produce germline chimeras using a combination of donor PGCs prepared from 35-dpf embryos and 35-dpf recipients. The PGC-transplanted recipients were reared until they reached maturity. Semen was collected from 1-yr-old PGC-transplanted fish. The DNA was extracted from 1 µl of semen and subjected to PCR with GFP-specific primers according to the method of Takeuchi et al. [9]. To determine the production of offspring from donor PGC-derived spermatozoa, semen obtained from PCR-positive fish was fertilized with nontransgenic eggs from wild type-colored females (-/-; wt/wt). Progeny tests were performed and repeated at three times, and the data are represented as the mean ± SEM. Eggs obtained from 2-yr-old, PGC-transplanted fish were fertilized with sperm from wild-type fish.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To select the most suitable developmental stage for transplantation, recipient embryos were compared at three different developmental stages: blastula (2.5 dpf), gastrula (6 dpf), and hatched embryo (35 dpf) (Table 1). When blastulae and gastrulae were used as recipients, the survival rates of the manipulated embryos (26% and 62%, respectively) were much lower than those of the control embryos (99%) at Day 30 posttransplantation. In contrast, when hatched embryos were used as recipients, the survival rate (94%) was similar to that of the untreated control group at Day 30.
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The distribution of donor PGCs in the recipient embryos was confirmed by observations of cell fluorescence at Day 30 posttransplantation (Fig. 1, C–G). When blastulae and gastrulae were used as the recipients, 60% (9 of 15) and 5.6% (1 of 18) of the recipients contained GFP-labeled cells, respectively (Table 1). However, these GFP-labeled cells were ectopically distributed in the epidermis of the head (Fig. 1C), the epidermis of the trunk (Fig. 1D), and the fin membrane and yolk sac (data not shown). In addition, the size and the morphology of these GFP-labeled cells were not altered in these areas: The cells retained PGC-like morphological characteristics, such as large size and round shape. The number of GFP-labeled cells found in the ectopic areas ranged from one to three, and signs of cell proliferation, such as the clustering of GFP-labeled cells, were not observed. In contrast, when hatched embryos were used as the recipients, colonization was observed in 21.6% (16 of 74) of the recipients (Table 1). The GFP-labeled cells were incorporated into the genital ridges, which are located on the dorsal wall of the peritoneal cavity (Fig. 1, E and F). The number of GFP-labeled cells observed in the gonads at Day 30 posttransplantation ranged from 1 to 35. Because both the dividing GFP-labeled cells and the resultant cell clusters were found in the recipient gonads (Fig. 1E), they likely had undergone proliferation. The incorporation of GFP-labeled cells was further confirmed by examination of the excised genital ridges (Fig. 1G).
Distribution of Transplanted PGCs in the Peritoneal Cavity of Recipients
To determine the process involved in the incorporation of donor PGCs into the genital ridges, the distribution of transplanted cells in the peritoneal cavities of the recipients was monitored from Day 1 to Day 30 posttransplantation (Fig. 2A). Donor PGCs were not attached on the peritoneal wall or incorporated into the genital ridges at Day 1 (data not shown). At Day 5, donor PGCs were found attached to the peritoneal wall or the dorsal mesentery of the recipients, but they were not observed in the genital ridges. The first incorporation of donor PGCs into the genital ridges was observed at Day 10. The frequency of colonization increased at Day 30, whereas the number of recipients with donor PGCs attached to the peritoneal wall decreased. Interestingly, GFP-labeled PGCs with extended lobopodia were often observed during this period (Fig. 2B).
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Influence of the Age of Donor PGCs and Recipient Embryos on the Frequency of Colonization
Genital ridges were difficult to excise in embryos before 35 dpf, because they were not sufficiently developed. These embryos were also unsuitable as recipients, because their peritoneal cavities were too small to allow insertion of the transplant needle. Therefore, the transplantation procedure established in the present study was not applicable to rainbow trout embryos before 35 dpf.
The results of the first experiment demonstrated that the donor PGCs isolated from 35-dpf embryos could be incorporated into the genital ridges of 35-dpf recipient embryos. The second experiment investigated the optimal developmental stage of both the donor PGCs and the recipient embryos for use in the transplant procedure. The PGCs prepared from hatched donor embryos at 35, 40, and 45 dpf were injected into recipient embryos at the same three stages (Fig. 3). The frequency of colonization obtained by the transplantation of 40-dpf donor PGCs into the 35-dpf recipients was similar to that obtained using 35-dpf donor PGCs. However, the rate declined significantly when 45-dpf donor PGCs were transplanted into 35-dpf recipients. Both 35- and 40-dpf donor PGCs were able to colonize the genital ridges of 40-dpf recipients, whereas colonization attempts by the 45-dpf donor PGCs were unsuccessful. These results indicated that the ability of donor PGCs to migrate toward the genital ridges decreased between 40 and 45 dpf.
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The age of the recipient embryos also affected the colonization rate, and we investigated whether a critical period existed during which recipient embryos provided suitable embryonic environments for the guidance of PGC migration (Fig. 3). No colonization was observed when donor PGCs prepared from 35- and 40-dpf embryos were transplanted into 45-dpf recipients, whereas they could successfully colonize the genital ridges of 35- and 40-dpf recipients. These results indicated that conditions in the peritoneal cavities of recipients suitable for the migration of transplanted PGCs were lost between 40 and 45 dpf.
Proliferation and Differentiation of Donor-Derived Germ Cells in Recipient Gonads
To examine whether the incorporated donor PGCs could resume gametogenesis, we examined the gonads of recipient fish at Day 180 posttransplantation. Donor PGCs carried a transgene, pvasa-GFP, which expressed the GFP gene specifically in all stages of germline cells other than spermatocytes, spermatids, and spermatozoa (unpublished data); therefore, the proliferation and differentiation of donor-derived germ cells could be readily analyzed. The GFP-labeled cells in the testes of the recipients had proliferated and differentiated into at least spermatogonia at Day 180 (Fig. 4, A and B). Similarly, two types of donor-derived germ cell were distinguished in the ovaries at Day 180: oogonia and primary oocytes, with cellular diameters of 10–15 and 50–80 µm, respectively (Fig. 4, C and D). The GFP-labeled cells were not observed outside of the gonads at this stage. The frequency of germline chimerism at Day 180 is summarized in Table 2. When donor embryos and recipient embryos were obtained from the same brood, 9.8% of the recipients possessed donor-derived germ cells; a similar result was observed when embryos obtained from different parents were used as recipients. No difference was found in the frequency of colonization between male and female recipients (data not shown). The PCR analysis using primers against the GFP gene revealed that the gonads containing donor-derived germ cells displayed GFP gene-specific signals, whereas no such signals were observed in other tissue samples, including the contralateral gonads (Fig. 4E). This confirmed that donor-derived cells were only present in the gonads at this stage.
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Germline Transmission of Donor PGCs
Spermatozoa obtained from PGC-transplanted recipients were tested for the presence of the transgene by PCR analysis. Exogenous GFP genes were detected in the spermatozoa obtained from 17% (2 of 12) of the matured recipients (Fig. 5A). The PCR-positive spermatozoa were used to perform a progeny test by artificial insemination, and the spermatozoa derived from one of the two males produced 115 orange-colored fry among 5825 progenies (2.04% ± 0.21%) (Fig. 5B). No orange-colored fry was found among the offspring of nontransplanted control males. Furthermore, approximately half the orange-colored embryos (54.8%, 63 of 115) showed GFP expression specifically in PGCs, because the transferred PGCs were hemizygous for the transgene (Fig. 5C). Because the transferred PGCs were heterozygous for the dominant orange-colored phenotype, half the progeny that developed from the donor-derived spermatozoa were expected to have orange body color; therefore, the contribution rate of the donor-derived spermatozoa to the germline was estimated as 4%. Of 14 matured females obtained from PGC-transplanted recipients, 2 (14%) yielded donor-derived offspring. The contribution rates of the donor-derived eggs to the germline were estimated as 3.03% (16 of 528) and 2.08% (20 of 960).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Although colonization of GFP-labeled donor PGCs was not observed in the genital ridges of recipients at Day 5 posttransplantation, it was evident at Day 10, and a marked increase in colonization efficiency was observed over a 30-day posttransplantation period. In contrast, the percentage of recipients with donor PGCs attached to the peritoneal wall gradually declined. Furthermore, some donor PGCs that were attached near the genital ridges had extended pseudopodia, which is a typical morphological feature of cells during locomotion. These results indicated that the transplanted PGCs actively migrated, using their extended pseudopodia, to the genital ridges, where they were subsequently incorporated. This process is schematically represented in Figure 2C. It has been reported that tropic or diffusible agents emanating from gonadal somatic precursors act as guidance cues for PGC migration in Drosophila sp. [15], chicks [16], mice [17], and zebrafish [18]. Although no direct evidence was found that attractant molecules were involved in the migration of trout PGCs toward the genital ridges, the results obtained in the present study indicate that the transplanted PGCs might sense and respond to molecules released from the genital ridges of the recipient embryos.
It was apparent that the 35- and 40-dpf recipient embryos were able to guide the donor PGCs toward their genital ridges but that 45-dpf recipients could not. Therefore, we suggest that the genital ridges of 35- and 40-dpf recipient embryos produced PGC attractants, but that this production had ceased in the 45-dpf recipients. Colonization efficiency was also affected by the age of the donor PGCs: The PGCs prepared from the genital ridges of both 35- and 40-dpf embryos had a greater ability to colonize the genital ridges of recipients than did the PGCs prepared from 45-dpf embryos. In vitro emigration assays that had been done with mouse PGCs showed that the PGCs only exhibited motile behavior when they were isolated during their migratory period [19]. Our results also suggested that the migratory competency of trout PGCs had a distinct and narrow window during embryonic development. Note that PGCs isolated from genital ridges retained their ability to migrate, their capacity to sense and respond to the attractant molecules produced by the genital ridges in this experiment. However, this competency gradually decreases during gonadal development. Therefore, we conclude that both the presence of suitable conditions in the peritoneal cavity of recipients and the migratory ability of donor PGCs themselves are essential for colonization of donor PGCs in the recipient gonad. To date, little is known about the genes, expressed in PGCs, that are involved with PGC migration in any organism. It has been shown that the somatic cells surrounding PGCs produce several kinds of proteins that might have a role in PGC migration. However, to our knowledge, only one gene involved in migration, which is expressed in the PGCs themselves, has been identified [18]. Therefore, detailed analysis of the gene expression profile of trout PGCs isolated at different developmental stages (e.g., 40 and 45 dpf) might facilitate the identification of the key molecules governing PGC migration.
The GFP-labeled cells were observed in both the head and the trunk of recipient embryos when donor PGCs were transplanted into blastulae and gastrulae. These results demonstrated that donor PGCs collected from the genital ridges did not follow the correct migratory path in the early stages of embryonic development. It has been reported that the migration of zebrafish PGCs can be summarized in two steps [20]. First, zebrafish PGCs migrate from their site of origin to the intermediate target, which comprises the precursor of the pronephros, during gastrulation and early somite genesis. Second, the PGCs migrate toward the region where the genital ridges are formed. Therefore, we suggest that the donor PGCs injected into recipient blastulae and gastrulae strayed from the migratory pathway, because they could not respond to guidance cues from the intermediate target.
In the present study, transplanted PGCs were converted into functional gametes and, subsequently, produced live offspring through fertilization, to our knowledge for the first time in nonavian vertebrates. Although donor PGCs colonized in the genital ridges were present in small numbers at Day 10 posttransplantation, large numbers were observed in the gonads of recipients at Day 180. Furthermore, donor-derived germ cells appeared as one cluster in the gonads of recipients at both Day 30 and Day 180. Therefore, each GFP-labeled cell cluster likely originated from a single donor PGC. These results demonstrated that even when the numbers of PGCs colonizing the genital ridges were low, a high degree of proliferation could be obtained. Surprisingly, donor-derived germ cells avoided immune rejection in the gonads of the recipients. One possible explanation for this phenomenon is that the donor PGCs acquired immune tolerance because they were transplanted before the onset of cellular immunity. In fact, the immune system is relatively immature at the time of hatching in a number of fish species [21]. These findings indicated that the PGC-transplantation technique established in the present study would be suitable for both allogenic and xenogenic transplantation in a wide range of fish species. Therefore, PGCs are a highly suitable cell type for efficient germ cell transplantation in fish. In the present study, the germline transmission of PGCs was 2–4%. However, the use of sterile recipients, such as triploid [22] or 
-ray irradiated [23] embryos, might increase the contribution efficiency of donor-derived gametes.
Alternatively, use of donors and recipients of the same sex (e.g., transplantation of male donor PGCs to male recipient embryos) may increase the transmission rate, because the retardation of donor-PGC development in recipient gonads of the opposite sex was reported in the chicken [24]. This technique, combined with the cryopreservation of PGCs and interspecies transplantation, has several potential applications in fish breeding, including the preservation of endangered species and the seed production of commercially valuable species, whose bloodstocks are difficult to maintain in captivity.
Because isolated PGCs can be converted into individuals via the maturation and fertilization processes using the PGC-transplantation technique, PGC-derived cell lines will be a suitable material for cell-mediated gene transfer. Furthermore, gene targeting could be performed using in vitro-cultured PGCs in combination with homologous recombination techniques. To reintroduce in vitro-cultured PGCs with genetic modifications into the germ cell lineage of recipient embryos, the cells must retain both the ability to migrate toward, and to colonize, the genital ridges and the ability to form functional gametes during the cultivation period. Establishment of a cell line possessing these characteristics, which would be suitable for cell-mediated gene transfer, is currently in progress.
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FOOTNOTES |
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2 Correspondence: Goro Yoshizaki, Department of Aquatic Biosciences, Tokyo University of Fisheries, 4-5-7 Konan, Minato-ku, Tokyo 108-8477, Japan. FAX: 81 3 5463 0558; goro{at}tokyo-u-fish.ac.jp
Received: 28 March 2003.
First decision: 16 April 2003.
Accepted: 8 May 2003.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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(1,3)galactosyl transferase (GGTA1) gene and the prion protein (PrP) gene in sheep. Nat Biotechnol 2001 19:559-562[CrossRef][Medline]This article has been cited by other articles:
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  T. Saito, R. Goto-Kazeto, K. Arai, and E. Yamaha Xenogenesis in Teleost Fish Through Generation of Germ-Line Chimeras by Single Primordial Germ Cell Transplantation Biol Reprod, January 1, 2008; 78(1): 159 - 166. [Abstract] [Full Text] [PDF]
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 T. Okutsu, S. Shikina, M. Kanno, Y. Takeuchi, and G. Yoshizaki Production of Trout Offspring from Triploid Salmon Parents Science, September 14, 2007; 317(5844): 1517 - 1517. [Abstract] [Full Text] [PDF]
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 T. Okutsu, K. Suzuki, Y. Takeuchi, T. Takeuchi, and G. Yoshizaki Testicular germ cells can colonize sexually undifferentiated embryonic gonad and produce functional eggs in fish PNAS, February 21, 2006; 103(8): 2725 - 2729. [Abstract] [Full Text] [PDF]
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G. Yoshizaki, Y. Tago, Y. Takeuchi, E. Sawatari, T. Kobayashi, and T. Takeuchi Green Fluorescent Protein Labeling of Primordial Germ Cells Using a Nontransgenic Method and Its Application for Germ Cell Transplantation in Salmonidae Biol Reprod, July 1, 2005; 73(1): 88 - 93. [Abstract] [Full Text] [PDF]
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) or inside (
) the genital ridges. B) A GFP-labeled PGC (arrow) attached to the peritoneal wall, near a genital ridge (GR), at Day 10 posttransplantation. The inset shows a high-magnification image of a PGC with an irregular shape and elongated lobopodium (arrowhead). C) Schematic representation of the migration of donor PGCs on the peritoneal wall. The inset shows a donor PGC (green) sensing and responding to chemoattractants (purple) produced by the genital ridges (orange). Red arrows indicate the direction of PGC movement. Bar = 20 µm
