SCIENCE CHINA Fish germ cells

SCIENCE CHINA Life Sciences Topic • Special REVIEW •

April 2010 Vol.53 No.4: 435–446 doi: 10.1007/s11427-010-0058-8

Fish germ cells XU HongYan1, LI MingYou1, GUI JianFang2* & HONG YunHan1,2* 1

2

Department of Biological Sciences, National University of Singapore, Singapore 119260, Singapore; State Key Laboratory of Freshwater Ecology and Biotechnology, Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan 430072, China Received September 29, 2009; accepted December 2, 2009

Fish, like many other animals, have two major cell lineages, namely the germline and soma. The germ-soma separation is one of the earliest events of embryonic development. Germ cells can be specifically labeled and isolated for culture and transplantation, providing tools for reproduction of endangered species in close relatives, such as surrogate production of trout in salmon. Haploid cell cultures, such as medaka haploid embryonic stem cells have recently been obtained, which are capable of mimicking sperm to produce fertile offspring, upon nuclear being directly transferred into normal eggs. Such fish originated from a mosaic oocyte that had a haploid meiotic nucleus and a transplanted haploid mitotic cell culture nucleus. The first semi-cloned fish is Holly. Here we review the current status and future directions of understanding and manipulating fish germ cells in basic research and reproductive technology. fish, germ cell, germ plasm, reproduction, reproductive technology, semi-cloning

Citation:

Xu H Y, Li M Y, Gui J F, et al. Fish germ cells. Sci China Life Sci, 2010, 53: 435–446, doi: 10.1007/s11427-010-0058-8

Bisexually reproducing metazoans have two major cell lineages for two distinct lifestyle phases, i.e. soma and germ. Somatic cells form the body responsible for the individual life, and are “mortal”. Germ cells are “immortal”, because they are responsible for the species life via producing gametes transmitting genetic information from one generation to the next [1]. In fishes, as in many other organisms, germ cells are formed as primordial germ cells (PGCs) early in embryonic development. PGCs come from presumptive PGCs (pPGCs). A pPGC is the germline precursor and undergoes asymmetric cell division, producing two daughter cells with different fates: one cell is committed to the soma and the other to pPGC. When both daughter cells are committed to the germline, the parental cell is said to be a PGC. PGC formation takes place at the species-specific site(s), before the formation of somatic components of a gonad at a different location. PGCs migrate through various somatic tissues to the developing gonad called the gonadal anlage. Accord*Corresponding authors (email: [email protected]; [email protected]) © Science China Press and Springer-Verlag Berlin Heidelberg 2010

ingly, PGCs are said to be premigratory, migratory and postmigratory. Upon arrival at the prospective gonad, PGCs coalesce with the somatic cells to form an intact gonad. In the gonad, PGCs in certain species such as in mice may stop mitotic division and enter a period of mitotic quiescence in G0. Such quiescent germ cells are now called prospermatogonia or gonocytes. During sexual maturation, gonocytes become oogonia and spermatogonia, and enter meiosis for ultimate differentiation respectively into gametes, sperm or eggs in the female and male [2]. Gamo [3] was among the first fish germ cell researchers who studied PGCs by using morphology and a characteristic subcellular structure called nuage in various fish species under light microscopy. Hamaguchi et al. [4] examined the dynamic nuage ultrastructures of medaka germ cells using electron microscopy. These pioneer studies were reviewed 10 years ago [2]. Subsequently, an increasing number of germ cell marker genes were identified in fishes (Table 1), and powerful molecular and cellular techniques were developed, facilitating understanding and the manipulation of fish germ cells. In this paper, we will review new aplife.scichina.com

436

Xu HongYan, et al.

Sci China Life Sci

proaches and progress in fish germ cell biology and achievements in fish germ cell technology. We will also propose major challenges and future directions for germ cell studies and reproductive technology in fish. When appropriate, the implication of the findings from fish germ cell studies about reproduction engineering and the fertility of animals will be mentioned.

1 Features, tools and approaches for analyzing fish germ cells PGCs differ from somatic cells in several morphological characteristics, which were used for PGC identification in early studies. PGCs have a large size (~20 µm in diameter), large nuclei (6–10 µm) and relatively little cytoplasm (i.e. a high nuclear-cytoplasmic ratio). PGCs are characterized by the presence of “nuage” or germinal granules. Nuage is rich in mitochondria, and is thus also called a mitochondrial cloud. This is a membrane-less cytoplasmic organelle containing RNAs and proteins. Nuage appears as a high electron-dense body under an electronic microscope [4]. These features of PGCs remain in routine use. Recently, You et al. Table 1

April (2010) Vol.53 No.4

[5] investigated the formation of nuage in oocytes at various stages in the cyprinid teleost Spinibarbus caldwelli using light and electron microscopes. Studies about germinal granules in several animal models (e.g. fruit fly Drosophila, frog Xenopus and zebrafish Danio) have focused on the asymmetric partitioning of the structures into prospective PGCs during early embryogenesis. Putative pPGCs are fate-determined by maternally inherited factors. These maternal factors are usually concentrated in germinal granules in oocytes and cleavage embryos [6]. Maternal factors directly involved in the germ cell fate decision are called germ plasm (GP). During the early stages of embryonic development, GP is asymmetrically segregated to a subset of cells that are fated to become PGCs. Therefore, in these organisms, PGCs are determined cellautonomously by maternal factors. In mammals, GP becomes discernible at the later stages of well-developed PGCs but not before their formation. Here maternal inheritance and contribution to PGC specification has not been documented, leading to the notion that GP components in mice and comparable mammalian species function primarily in postnatal germ cell development, but not in early embryonic stages [7]. Mammalian PGCs are

Significant fish germ cells genes

Genea)

Activity

Species, expression & function

Ref.

b)

Germ plasm and germ cell formation vasa

DEAD box RNA helicase

germ plasm assembly, PGC formation & migration

[8,9]

ziwi

RNA-interacting proteins

germ cell maintenance

[10]

tudors

Tudor domain proteins

germ plasm component, highly conserved across phyla

[11]

germ cell-less

Nuclear envelope protein

PGC formation & transcriptional repression

[12]

bucky ball

Novel zebrafish gene

germ plasm assembly, Balbiani’s body formation & PGC formation

[13]

dazl & boule

RNA binding protein

DAZ family of human male infertility genes

[14,15,16]

nanos 1 to 3

RNA-binding zinc finger protein

Four nanos genes in medaka. nanos2 & 3 are germ cell markers

[17]

zebrafish PGC migration & survival, oocyte maturation

[18,19]

Germ cell transcription quiescence and survival nanos1

RNA-binding zinc finger proteins

Germ cell migration staufen

RNA binding protein

PGC migration by regulating vasa translation in zebrafish

[20]

dead end

RNA binding protein

PGC migration & survival in zebrafish

[21,22]

hmgcr

see right

3-hydroxy-3-methylglutaryl coA reductase, PGC attraction to mesoderm

[23]

quemao (qm)

Geranyl-geranyl diphosphate synthase

PGC attraction to mesoderm

[23]

Igf

Insulin-like growth factor

PGC migration

[24]

Pik3

Phosphatidylinositol 3-kinase

PGC migration speed (zebrafish)

[25]

sdf1a-cxcr4b

Stromal-derived factor 1a/Receptor

PGC chemotaxis

[26,27]

cyclopamine

Hedgehog signaling inhibitor

affecting PGC motility & adhesion via targeting Smo

[28]

gai

G-protein

PGC migration

[25]

ggt1

Geranyl-geranyl transferase

PGC attraction to mesoderm

[23]

puf-A

RNA-binding protein

PGC specification & migration

[29]

a) Genes in bold are known factors in fish for PGC development. b) According to known factors in fish PGCs. Some genes may serve as multiple factors, e.g. zebrafish nanos.

Xu HongYan, et al.

Sci China Life Sci

induced in a subpopulation of pluripotent epiblast cells by signaling molecules (i.e. bone morphogenetic proteins) from surrounding extraembryonic ectodermal cells [30]. Therefore, PGC formation occurs in the absence of maternal factors but depends on intercellular interactions in a cell-nonautonomous manner. In many organisms, oocytes at certain early stages have a specialized structure called Balbiani’s body (BB) [14,15,31]. There are similarities and differences between BB and GP. BB is also a subcellular structure that is often associated with the Golgi apparatus, endoplasmic reticulum and mitochondria. Many GP components are also found in BB. In Xenopus, GP overlaps BB. In several mammalian species, BB is seen in early oocytes at the cyst and primordial follicle stages, but becomes disassembled in growing oocytes [32]. Convincing evidence demonstrating equivalence between BB and GP has thus far not been observed in vertebrates [33]. We have demonstrated that a component, i.e. boule RNA, is absent in the medaka BB, in which another GP component, dazl RNA, concentrates [14]. Future work will elucidate whether BB and GP are different structures or different forms of one and the same structure at different stages of germline development. GP is a ribonucleoprotein (RNP) complex comprising dozens of different RNA and protein components. These GP components assemble into a cytoplasmic architecture in a cytoskeleton-dependent manner. Differences in temporal requirements for actin and microtubules have been described in Drosophila, Xenopus and zebrafish. In Drosophila, GP components simultaneously require an intact actin and microtubule network, whereas the actin network is dispensable for GP assembly in Xenopus. In zebrafish, the initial GP assembly may be actin-dependent, whereas the subsequent segregation requires the function of the so-called furrow microtubule array [34,35]. Although the actin-based cytoskeleton is a factor in the early segregation of GP components, whether or not the cytoskeleton is directly involved in GP recruitment to the cell division furrow of early cleavage zebrafish embryos remains to be examined, as furrow formation per se cannot take place when the cytoskeleton is damaged [34]. Bontems et al. [13] have found that a gene called bucky ball (buc) is essential for GP assembly in zebrafish, as buc mutation disrupts oocyte polarity and BB formation, and buc overexpression induces ectopic germ cell formation. Buc is a divergent protein, and whether or not its homolog is present in other vertebrate species and performs a conserved function in GP assembly remains to be determined. Although the function(s) and assembly of GP in diverse species and at different stages of the germline development have not been fully understood, the GP of diverse species does share considerable similarities in morphology and molecular composition (Table 1) [30], suggesting a conserved role in germline development. There is evidence that the characteristic RNPs in germinal granules may also be asso-


437

ciated with various general processes, including RNA metabolism, retro-transposon regulation and interplay with mitochondria [7]. The identification of the zebrafish vasa gene as the first fish germ cell marker has affected germ cell studies at the molecular level [9]. Vasa has been cloned in several other fish species [36,37]. Vasa proteins across phyla show sequence conservation, allowing for profiling Vasa protein expression via immunostaining by using an anti-Vasa antibody in diverse species. For example, an anti-Vasa antibody (αVasa) we produced by using the recombinant gibel carp Vasa protein as the antigen is used for immunohistochemistry to detect Vasa protein distribution in male and female germ cells of not only the gibel carp but also several other fish species of different phylogenetic relatedness [38], revealing a conserved expression pattern, which is comparable to that described in medaka [39], and is similar to that in mice [40]. Germ cell markers have increasingly been characterized in fish. These include nanos, dnd, dazl and sdf1/cxcr4 (Table 1). The use of antisense RNAs of these genes as probes facilitates their unambiguous detection in PGCs by in situ hybridization (Figure 1). Importantly, as mentioned in the review [2], the grafting experiments in Fundulus suggest that during the mid-shield stage, the posterior shield may be the source of PGCs whereas the ventral and lateral germ ring may not give rise to PGCs. In situ hybridization at this stage revealed that vasa-positive cells are present at all these locations. Accordingly, not all vasa-positive cells will develop into PGCs. In other words, expression of vasa may be one of the criteria for identifying PGCs. It is useful to label PGCs by using multiple gene markers to improve PGC detection [21] (Figures 1A–C). To do this, we have developed a dual fluorescence in situ hybridization (FISH) procedure in medaka for utilizing several germ cell markers to label PGCs. This dual color procedure allows for simultaneously labeling PGCs with at least two germ cell marker genes (Figures 1D–F). This method clearly improves the visualization and documentation of dynamically developing germ cells. This dual color FISH procedure is expandable to multiple color FISH（Xu and Hong, unpublished）to provide more precise information for further analysis of dynamic PGC behaviors at the molecular or cellular levels. FISH detects dead samples in fixed whole mount samples and sections. Techniques have been developed to label living PGCs by producing transgenic animals. In mice, MacGregor et al. produced PGC-specific LacZ transgenic mice. Yoshimizu et al. [41] produced a transgenic mouse carrying GFP driven by the Oct-4 gene promoter, which is specifically active in PGCs at particular stages. In fishes, a gene construct upon embryo microinjection is information integrated at a low efficiency into the host genome. This uncontrollable integration often results in a low efficiency of transgenic production and an unpredictable pattern of transgene expression, demanding massive screening by a

438

Xu HongYan, et al.

Sci China Life Sci

progeny test. Consequently, although transgenic lines have been developed in fish, PGC-labeling by transgenic production has been limited to the fish models medaka and zebrafish. One exception is the trouts. In these organisms, the vasa promoter (VAS) and nanos promoter (NOS) have produced transgenics showing germline-specific transgene expression. Combinations between NOS or VAS and GFP or RFP respectively generated transgene constructs VASgfp, VASrfp and NOSgfp (a promoter linked to GFP or RFP). These constructs were used to produce transgenic trout, medaka and zebrafish [8]. Also, these constructs may be used for transgenic production in other fish species. These transgenic fishes provide a powerful tool for detection, isolation and manipulation of live germ cells [42]. Compared with mammals, producing transgenic fish is relatively easy but still represents a time-consuming task in such aquaculture species as grass carp and sturgeon which require several years to grow into sexual maturation. Moreover, a novel technique to label living PGCs has been developed for the first time in zebrafish on the basis of localized expression of a reporter (GFP or RFP) mRNA that is injected into early developing embryos. The reporter RNA is synthesized as a fusion to the 3′ untranslated region (3′ UTR) of a germ cell marker gene such as nanos [18,43]. RNA localization is determined by the cis-acting elements within the RNA. Such localization elements are located usually, but not exclusively, within the 3′UTR mediating interaction between trans-acting protein factors and RNAs.


In zebrafish, an evolutionary conserved region exists in the vasa 3′UTR targets RNA translation to the germ cells and this vasa 3′UTR works also in the Xenopus. The zebrafish nos1 3′UTR controls PGC-specific expression in several other fish species including medaka [44,45]. A 3′UTR affects RNA stability, a feature for localized RNA concentration and/or translation. In Drosophila [46], Hsp83 is a factor in the localized degradation of such RNAs as nanos. In zebrafish, mRNAs of vasa [47] and nanos1 (nos1) [48] are selectively degraded in somatic cells but stabilized and translated in PGCs. The conservation in sequence and mechanism facilitates identification of putative GP-localizing RNA sequences [49] towards tracing live PGCs in fish. This technique makes use of localized expression from injected synthetic RNA but not of transgenic expression from a stable transgenic line. This approach is called localized RNA expression (LRE). In addition to speed and simplicity, LRE provides one additional advantage, i.e. versatility: a PGClabeling RNA derived from, and tested in, one species is usable in other fish species, as has been demonstrated for the PGC-labeling ability of the zebrafish nanos 3′-UTRcontaining GFP RNA in medaka embryos [44,45]. Consequently, LRE finds an increasing use for labeling PGCs in such dynamic processes as migration, proliferation and survival during fish embryonic development [45,50]. The establishment and use of these approaches in fish have advanced fish germ cell biology and reproductive technology. It is noteworthy that transgenic production and LRE are

Figure 1 Visualization of Medaka PGCs by chemical and fluorescent WISH. A–C, Chemical in situ hybridization, showing stage-18 embryos after hybridization with anti-sense RNA probes of dazl, boule and vasa. The signal is in red to purple. PGCs are seen in two clusters bilateral to the body axis. Bars, 100 μm. D–F, Dual color fluorescent in situ hybridization, showing stage-27 embryos after hybridization with anti-sense RNA probes and fluorescence visualization. D, boule; E, dazl; F, merged of boule, dazl and nuclear staining by DAPI (blue). so, somites; no, notochord. For experimental procedures see ref. [14]. Bars, 50 μm.

Xu HongYan, et al.

Sci China Life Sci

complementary approaches for PGC-labeling in fish. LRE features speed and versatility for PGC labeling in fishes where a transgenic line is not available. Because the rationale behind LRE is differential stability and translation of injected RNA, and this preferential expression does not occur until PGC formation, the earliest stage of PGC formation, i.e. the onset of PGC specification, is not revealed by this approach. The amount of RNA to be injected per embryo requires elaboration to obtain a reasonable contrast for clear observation and documentation. In addition, the number of embryos injected in each experiment will be limited, and different embryos will also show variation in relative signal intensity (PGC vs soma) and absolute signal intensity (e.g. PGC signal between embryos). LRE has been limited to labeling PGCs during the early stages of fish development. Transgenic production provides numerous embryos of relatively uniform labeling signal and the possibility of observing germ cells throughout development including adult gonads. From our experience, a promoter that drives a lower level of expression in germ cells (e.g. medaka nanos3) is better than a stronger promoter (medaka vasa) if PGC observation will focus on the early stages, or vice versa. For example, in VASgfp transgenic medaka embryos, the maternal supply provides a strong signal in a wide variety of cells until gastrulation, obscuring PGC detection at earlier stages. With both transgenic lines in hand, we show in medaka that embryos from crossing between NOSgfp females and VASgfp males offer an excellent opportunity for detecting PGCs at early (NOSgfp-derived signal; maternal) and late stages (VASgfp-derived signal; paternal and zygotic) [8].


439

are phosphorothioate-linked DNA (S-DNA), short interfering RNA (siRNA) and Morpholino. Owing to sequence specificity, stability and highly predictable design, Morpholinos are widely used in gene knockdown experiments, especially in those aimed at analyzing the early phenotypes of developing embryos. In fish, eggs and embryos are manipulable, with fertilization and embryonic development occurring outside the body. Together with live PGC labeling and imaging, Morpholino gene knockdown is widely used in fish to study germ cell biology in embryogenesis [8,26,43,52].

3 Germ cell development Embryonic germline development in diverse animal species proceeds in two major phases, i.e. germ cell formation and migration, as illustrated in Figure 2. Many genes involved in this process are highly conserved during evolution.

2 Tools for functional analyses of germline genes Once detection tools are available in fish, a germline strategy was established for transmitting a zygotic lethal mutation through a host animal by cell transplantation into a zebrafish embryo, resulting in the complete replacement of the host germ line with donor PGCs, allowing for maintenance of the lethal mutant strain and the analysis of the maternal effects of a zygotic lethal mutation [43]. In this elegant experiment, an approach called gene knockdown – translation inhibition by an injected antisense morpholino oligonucleotide – was adopted to reduce the translation of dead end (dnd), a gene encoding one GP component. Knockdown of dnd severely affects PGC survival and migration into the gonad, making the host fully open for colonization by donor PGCs transplanted in a pool of blastula cells [43,51]. Generally, gene knockdown achieves high sequence specificity and avoids off-target (non-specific) effects. There are three types of molecules for knockdown. These

Figure 2 PGC specification and migration in Drosophila, fish and mice. A, PGC specification. B, PGC migration initiation. C, PGC migration through somatic tissues. D, PGCs reaching the genital ridge. Germ plasm and PGCs are green in color (Modified from refs. [30,54,59]).

440

3.1

Xu HongYan, et al.

Sci China Life Sci

Fish PGC formation

The segregation between germline and soma occurs early during embryonic development. As illustrated in Figure 2, two different modes of PGC formation have been well established in diverse organisms [53]. In many organisms such as Drosophila [54], nematodes and anuran amphibians [1], the germline is segregated from the soma during the early cleavages through localization of specific cytoplasmic determinants, or “pole plasm” (GP). This mode of PGC specification is called preformation. In urodelian amphibians [55], chickens [56] and mice [30], PGCs are induced de novo by other cells in early developing embryos, in the absence of maternally supplied GP components. In these organisms, GP components are also exclusively or preferentially expressed in PGCs, but anew after PGCs have been well established PGCs. This mode of PGC formation is called epigenesis [30] The timing of PGCs specification exhibits a difference in diverse species. All mouse embryo cells up to the 8-cell stage are totipotent. A 16-cell stage embryo has 4 insider and 12 outsider cells, with insider cells making up the inner cell mass (ICM), and outsider cells producing the trophectoderm (or trophoblasts) that will become the future placenta. After implantation, the ICM develops into the pluripotent epiblast. PGCs are first visible at 6.5 d post coitum (dpc) of early gastrulation in a region of the epiblast that is close to the extraembryonic ectoderm [57]. In Drosophila and Caenorhabditis, the germline is already preformed at the onset of embryogenesis. In Xenopus, the germline occurs shortly before gastrulation [58]. In fish, the first attention focuses on the origin and timing of germ cell formation in different species. In zebrafish, by in situ hybridization using vasa as a molecular marker, pPGCs are distinctive quite early, already at the 2-cell stage. As a key GP component, vasa RNA is uniquely localized to the cleavage planes at the 2- and 4-cell stages and restricted asymmetrically to, and exclusively maintained in PGCs [9]. By analogy, zebrafish follow the preformation mode and make use of maternal determinants for PGC formation. In the killifish Fundulus, by shield grafting experiments, PGCs were first seen at the mid-gastrulation in the posterior but not the ventral or lateral shield [2]. In this fish, the mode of PGC specification remains to be determined. Medaka shows salient differences from zebrafish in the distribution of vasa transcripts prior to PGC formation. In medaka, vasa RNA does not segregate asymmetrically but is widely distributed in many cells before its restriction to visible PGCs until gastrulation [36], a feature inconsistent with preformation. In this regard, medaka resembles Fundulus. If differential vasa expression would tell when and where PGC formation occurs, then the distribution pattern of medaka vasa would imply a different timing of PGC formation. Alternatively, this discrepancy in early embryonic vasa expression and distribution perhaps reflects divergent expression of se-


lected GP component genes during teleost evolution. As described previously, medaka boule and dazl also exhibit a vasa-like expression pattern, i.e. the absence of asymmetric segregation. Medaka PGCs appear to be a specified cellautonomously, a feature supporting preformation [8,14]. This raises a question as to whether or not the zebrafish mode of PGC preformation applies also to medaka and other fish species. A question also rises as to whether or not a previously unidentified mode would operate in such fish species as medaka. Examination of the expression patterns of more GP genes and experimental analysis in medaka may provide clarification. 3.2

Fish PGC migration

3.2.1 Migration pathway Early in developing embryos, PGCs arise in one place that is different or far away from their final destination, the genital ridge or embryonic gonad. Therefore, PGCs migrate a long way through the tissues to reach the genital ridges. The processes of PGCs migration have been extensively studied in Drosophila, chickens, mice and zebrafish [54,59]. As illustrated in Figure 2, there are two major pathways of PGC migration. One is the blood circulation, which is used in birds. The other makes use of the gut, which takes place in all examined organisms except birds. PGC migration in several fish species, examined by using histological and morphological criteria, was found to follow the gut pathway (Figure 2), similar to Drosophila and mice [2]. Studies using molecular markers and live imaging following PGC labeling by transgenic expression and LRE have confirmed this. The best example is zebrafish. A zebrafish embryo at the 1000-cell stage (3 hpf) contains four PGCs in a square-like configuration, which are randomly oriented with respect to the dorsal–ventral axis. Upon initial proliferation, PGCs form four clusters in the marginal regions of embryos at the sphere stage (~4000-cell stage). When embryogenesis proceeds, PGCs undergo six sequential steps of migration, leading to two bilateral groups that ultimately reach the gonad [2,59]. A similar picture has been drawn for medaka PGC migration [36]. 3.2.2 Molecules/genes controlling PGC migration Recent work using PGCs labeling and functional analysis of germ cell regulators has led to substantial progress in elucidating genes and signal pathways in fish PGC migration. It has been shown that fish PGC migration depends on several GP components and on the GP integrity. For example, PGCs cannot properly migrate upon knockdown of such GP components as nanos [18] and dnd [22], resulting in mislocalized PGCs that ultimately die [60]. In medaka, vasa is cell-autonomously required for PGCs migration [8]. Zebrafish migratory PGCs take passive and active movements. Passive movements depend on gastrulation rearrangements (e.g. epiboly). Active migration depends on

Xu HongYan, et al.

Sci China Life Sci

motility, as evidenced by the formation and dynamic changes of pseudopodia that were identified in zebrafish PGCs during gastrulation. Soon after formation, zebrafish PGCs undergo transition from round, immotile cells (passive movement) into polarized migratory population for active movements [60]. Although the molecular or cytoskeletal basis for this transition is not clear, two genes have been implicated in PGC maturation before migration. One encodes phosphatidylinositol 3-kinase (PI3K). PGCs expressing a dominant negative form of PI3K show reduced cell polarity and stability of filopodia, as well as a slower migration speed [25]. The other gene is dnd, a novel vertebrate GP component encoding an RNA-binding protein. As mentioned above in the germline replacement experiment, dnd is essential for pseudopodia formation in PGCs, and dnd knockdown thus affects PGC motility and the onset of migration [22]. The establishment and/or maintenance of cellular motility in fish PGCs appears to be a cell-autonomous process, as PGCs exhibit motility when they are observed in culture [8,60]. Identifying more genes or dnd target genes and determining their roles might provide new insights into the understanding of molecular mechanisms underlying motility establishment and migration initiation, a critical step for normal germline development and fertility, and offer tools for manipulating these steps in germline engineering. After the onset of migration, PGCs undergo directed migration towards the gonad. PGC migration depends on cell-autonomous and non-autonomous mechanisms. Zebrafish PGCs are sequentially translocated to the anterior and lateral boundaries of the mesoderm during gastrulation, move toward an intermediate region bordering the mesoderm during somitogenesis, become bilaterally aligned at 24 hpf to the anterior yolk extension (between the eighth and tenth somite) and finally reach the gonad where PGCs coalesce with somatic cells into an intact gonad [61]. As in Drosophila, fish PGC migration is also guided by certain repulsing and attracting chemokines acting on receptor molecules in PGCs. For example, the G-protein-coupled receptor CXCR4b [26,27] and its ligand, stromal cellderived factor 1a (SDF1a) [26], provide directional cues to PGCs and guide them towards the gonadal ridge in zebrafish. Besides the SDF-1 and CXCR4b, other chemokine– receptor pairs may also exist which guide PGC migration, because zebrafish PGCs travel through many SDF-1 expressing tissues but ignore other SDF-1 expressing tissues [62]. In zebrafish, the SDF-1 and CXCR7 have been reported to make a second pair of ligands and receptors. CXCR7 is crucial for PGC migration, but its function is distinct from that of CXCR4: CXCR4b translates the polarized distribution of SDF-1 into directed PGC migration, whereas CXCR7 acts as a high-affinity decoy receptor and facilitates PGC migration by shaping the distribution of the chemokine in the environment [63]. Similar mechanisms of


441

PGC migration have been documented also in medaka. Medaka PGC movements are controlled by two sdf1 genes (arisen from whole genome duplication) encoding SDF1a and SDF1b. The two genes show only partially overlapping expression patterns and their products cooperate in the correct positioning of PGCs [45]. Perhaps multiple pairs of chemokines and receptors are adopted in the regulation of PGC migration. In zebrafish, a new mechanism controlling PGC migration has been described [23], in which hydroxymethylglutaryl coenzyme A reductase (HMGCoAR) is a key factor. This gene encodes a key enzyme in cholesterol biosynthesis whose activity is needed to provide the geranylgeranyl transferase (GGT1) substrates, and GGT1 triggers the specific prenylation necessary for PGC migration. There are many examples illustrating that the mechanisms underlying PGC migration appear to be highly conserved in diverse animal phyla. HMGCoAR was found to also function in Drosophila [64]. In mice, directional PGC migration also requires the repellant activity of IFITM1, expressed in the mesoderm [65]. In mutant mice lacking functional SDF-1 or CXCR4, PGCs cannot coalesce with gonadal somatic cells into gonads [66,67]. In both mice and chickens, SDF-1 functions during the second phase of PGC migration, but not at earlier phases. SDF-1 is required for PGCs to execute the final migration step as they transmigrate through the blood vessel endothelium (chicken) or the gut epithelium (mouse) towards the prospective gonad [56], in contrast to its guidance role for PGC migration in fish [26]. Although the SDF-1a functions in different steps of germ cell migrations in diverse species, its role in PGCs migration is evolutionarily conserved across phyla. Therefore, the findings regarding fish PGCs migration may provide clues for investigating PGC migration in other vertebrates including mammals. The formation of a functional gonad requires directed PGC migration towards the somatic gonad-precursor cells. Although male and female animals result from embryos with PGC migration failure, they will be partially or completely infertile [51,68]. Data from fish PGC migration may potentially shed insights into animal (in) fertility. 3.2.3 PGCs as a model for mechanistic understanding of cell migration Cell migration is essential for many biological processes, including embryonic morphogenesis, immune response, wound repair and tumor metastasis. Molecular mechanisms underlying cell migration in different organisms and different cell systems share many features in common. It is widely accepted that data from model organisms or cell systems provide insights into the understanding of these processes in humans. In this regard, fish PGCs offer an excellent system for analyzing a highly dynamic process of cell migration in externally developing embryos. As men-

442

Xu HongYan, et al.

Sci China Life Sci

tioned above, the chemokine SDF-1 is required for PGC migration in fish [26,52]. SDF1 is well known for its attractant role in the migration of human lymphocytes and monocytes [69]. In addition, SDF-1 and CXCR4 have been shown to be involved in tumor pathogenesis and metastasis [70], and infectious and inflammatory processes as well [71]. It is anticipated that ongoing work regarding fish PGC migration will improve understanding of cell migration in normal and abnormal situations.

4 Germ cells and reproduction modes Lower vertebrates display a wide variety of reproduction modes, ranging from fully unisexual to typical bisexual reproduction. Certain fish species, such as a population of the crucian goldfish called “gibel carp” (Carassius auratus gibelio), undergo a unique variant of gynogenetic reproduction called “allogynogenesis”. This fish consists predominantly of females and variably occasional males capable of sperm production. However, unlike “purely” bisexually or gynogenetically reproducing animals, it is unusual in that it has the ability to switch between asexual and sexual reproduction. It often undergoes gynogenesis in nature. Gibel carp reproduce also gynogenetically following artificial insemination of their eggs with heterologous sperm from a closely related species such as the common carp (Cyprinus carpio) [72]. Gibel carp undergo bisexual reproduction, with biparental genetic contribution to both the egg and sperm genomes to the offspring. The choice between reproduction modes depends ultimately on the role/behavior of germ cells, which provokes interest in studying germ cells in this unique organism. For example, we have cloned several germ cell marker genes and analyzed their RNA and protein expression pattern, such as vasa [38] and dazl [15], as well as a novel oocyte-specific variant of H2A [73], implying a possible association between chromatin structure and reproduction mode.

5 Germline manipulation The understanding of germ cell development provides a basis for germline engineering. One such approach is germ cell transplantation (GCT). GCT was established initially in chickens [74] and mice [75]. GCT has been applied in a number of animal species including domestic mammals [76], birds [77] and fish [69]. Owing to the availability of transgenic fish [78] and its ease for in vitro fertilization and embryogenesis, GCT has been used in fish [79,80]. Fish GCT was reported first in the rainbow trout by grafting a testicular cell mixture into an isogeneic immature gonad with a low efficiency of donor-derived spermatogenesis [81]. The efficiency was improved by transplantation of FACS-sorted PGCs into the migration route of em-


bryos or fry [82,83], and the transplanted syngeneic or xenogeneic spermatogonia also colonized the recipient gonads [83,84]. Recent studies in salmonids revealed two prerequisites for the migration of intraperitoneally transplanted donor spermatogonia to the recipient genital ridges. One is that the age of recipient larvae affects the colonization efficacy by donor PGCs or spermatogonia, and the other is that only type-A spermatogonia possess the ability to colonize the recipient genital ridge [82,84]. There is increasing interest in exploring surface markers to enrich them for spermatononia in various animals e.g. mammals [85]. In the rainbow trout, Notch1 appears to be such a marker [86].

6 Germ cell culture For basic studies and germline engineering, germ cell cultures hold great potential. The cell culture of PGCs and adult germ cells in mammals, chickens and fish has been attempted. Matsui et al. [87] succeeded in the cultivation of mouse PGCs, leading to the formation of embryonic germ cells. PGC cultures also have been reported in chickens [88]. In fish, Yoshizaki’s laboratory isolated trout PGCs by FACS sorting and examined growth responses in short-term culture (Table2). Fan et al. [89] have produced transgenic zebrafish expressing RFP under the vasa promoter, and cultivated FACS-sorted PGCs for up to 4 months. These studies demonstrate the ability to isolate and cultivate PGCs from various vertebrate species including fish. Throughout the adult life of most animals, e.g. fish, male germ cells in the testis produce sperm that transmit genetic information between generations. At the onset of spermatogenesis, testicular gonocytes or prospermatogonia resume proliferation and become undifferentiated type-A spermatogonia, the male germ stem cells that self-renew themselves to maintain the stem cell pool or differentiate through meiosis into fertile sperm [90]. As the adult testis has germ stem cells, efforts have been made towards isolating and cultivating testis-derived cell cultures including germ cells. Consequently, cell cultures were extensively attempted in the mouse model and several cell lines were established from the adult testis [91]. Feng et al. [92] reported spermatogonial cell culture from a virus-transformed mouse testis and demonstrated that transformation is necessary for serial culture. In medaka, we succeeded in a SG3, a normal spermatogonial cell line from an adult testis without the need for transformation (Table 2). Later, Guan et al. [93] have established normal mouse spermatogonial cell cultures. Germ cell culture provides an in vitro system for germ cell biology and a tool for germ cell transplantation. In this regard, embryonic stem (ES) cells provide a routine for germline transmission of highly demanding transgenic mice, leading to the production of knockout models by germline chimera formation.

Xu HongYan, et al.

Table 2 Year

Sci China Life Sci

443


Major events of fish germ cell researcha) Laboratory

Event

Publication

1982

Hamaguchi

Microscopic study on PGC migration in medaka

Cell Tissue Res 227: 139–151

1996

ES cell lines from medaka as the 2nd vertebrate

1998

Hong Olsen Yoon Hong

ES cell-derived medaka chimeras

Mech Dev 60: 33–44 Mech Dev 66: 95–105 Development 124: 3157–3165 Proc Natl Acad Sci USA 95: 3679–3684

2000

Yoshizaki

Transgenic rainbow trout with labeled PGCs

Int J Dev Biol 44: 323–326

2001

Raz

zebrafish nanos, knockdown & PGC labeling by RNA

Genes Dev 15: 2877–2885

1997

zebrafish vasa as a germ cell marker

2002

Schier

Zebrafish germline replacement by blastomere transfer

Proc Natl Acad Sci USA 99: 14919–14924

2002

Raz

Guidance of PGC migration by SDF-1

Cell 111: 647–659

2002

Knaut

vasa 3'UTR and RNA translation to zebrafish PGCs

Curr Biol 12: 454–466

2002

Yoshizaki

Mass isolation of transgenic rainbow trout PGCs

Biol Reprod 67: 1087–1092

2003

Yoshizaki

Rainbow trout fry from PGC transplantation

Biol Reprod 69: 1142–1149

2003

Raz

zebrafish dead end in initial PGC migration and survival

Curr Biol 13: 1429–1434

2004

Hong

Normal spermatogonial cell line in medaka


2004

Yoshizaki

Trout production in salmon by PGC transplantation

Nature 430: 629–630

2005

Raz

Female-to-male sex reversal by PGC ablation


2006

Yoshizaki

Trout production in salmon by transplanted testicular cells


2007

Yoshizaki

“Pure” trout production in salmon from PGCs

Science 317: 1517

2009

Hong

Haploid ES cells & semi-cloned animal Holly in medaka

Science 326(5951): 430–433

a) Milestones are highlighted in bold. The emails of major laboratories are below: Erez Raz, [email protected]; Hong YunHan, [email protected]; Goro Yoshizaki, [email protected]

Fish ES culture has been extensively attempted in model species. Medaka represents a unique lower vertebrate that has given rise to chimera-competent ES lines [94,95]. Ma et al. [96] have reported that short-term ES cell culture in zebrafish colonizes the germ line. The promise from the model fish has provoked similar attempts in aquaculture species. Chen’s laboratory reported ES-like cells from several marine fish species [97,98]. Ongoing work aims to achieve a high efficiency of germline chimera formation. Data from the experimental analyses of fish germ cell biology may provide direct information for ES cell-mediated germline transmission. Generally, the approach, transplanting the early embryonic cells or ES cells to host embryos for germline chimera production, in itself is one type of GCT, as early embryonic cells are undetermined cells and contain a few PGCs, and ES cells are considered to have potential for giving rise to germ cells [99–101].

7 Novel reproductive technology The first haploid ES cells have been obtained from medaka and used to test a novel reproductive technology [102]. In normal reproduction, fertilization leads to the fusion of two haploid meiotic nuclei, one from sperm and the other from an egg. To mimic fertilization, intracytoplasmic sperm injection has been developed and increasingly used for treating infertile men who have germ cells but defects in post

meiotic progression [103]. Embryos from diploid somatic cell nuclear transfer to enucleated oocytes grew into viable offspring in frogs, fish and mammals [104–106], providing a powerful tool for animal cloning, human ES cell derivation and analyzing nuclear reprogramming [104]. However, the application of this cloning strategy to human assisted reproduction has widely been debated because of low efficiency and ethical concerns about identical progeny from the nuclear donor. An assisted reproductive technology on the basis of creating mosaic oocytes has been proposed for treating infertile patients lacking any germ cells [107]. This approach, called semi-cloning (SC), uses nuclear transfer to combine a haploid somatic nucleus and a haploid gamete nucleus in the oocyte. SC not only ensures the biparental contribution to the progeny but also creates a new and unpredictable combination of genetic traits from both parents similar to normal fertilization. SC has remained hypothetical because viable SC offspring have not yet been obtained [108]. The availability of haploid ES cells allowed the testing of this SC in medaka as a first model. Transplantation of these haploid ES cell nuclei into normal, non-enucleated eggs has led to reconstituted eggs capable of normal development. One of those developed even to a viable and fertile female capable of germline transmission. This semi-cloned fish was Holly [102]. The birth of Holly provided a direct proof-of-principle that haploid ES cells are capable of mimicking sperm to produce fertile offspring. This fish

444

Xu HongYan, et al.

Sci China Life Sci

originated from a mosaic oocyte that had a haploid meiotic oocyte nucleus and a transplanted haploid mitotic cultured cell nucleus. Therefore, a reconstituted mitotic-meiotic diploid egg supports normal development, demonstrating the feasibility of SC for reproductive medicine. Ongoing work will determine whether or not haploid ES cells may be established in other vertebrate species and used for SC.

1 2 3 4 5

8 Perspectives Fish germ cells have gained increasing interest. The driving force is threefold. First, fish species offer uniquely easy embryology for experimentation and bioimaging of dynamic processes of PGC development. Second, fish represents a large group of vertebrates, with numerous species threatened and endangered, and germ cell cultures and transplantation provide a powerful tool for cryopreservation and propagation of these species by surrogate reproduction. Third, as vertebrates, fish share many common features with mammals including humans. It is widely accepted that data from fish germ cells will shed insights into the understanding of reproduction and infertility. For example, the recent birth of Holly demonstrated the possibility of semi-cloning as a novel reproductive technology. We anticipate that the work on fish germ cells will continue to provide invaluable information for germ cell biology and biotechnology in basic and applied research fields. In fish, the experimental analysis of gene functions in germline development has mainly relied on the use of gene knockdown. All germ plasm genes analyzed in fish appear to be indispensable for PGC formation, in contrast to the Drosophila situation, where many of them, e.g. vasa, are essential for PGC formation as revealed by mutant analysis. Such variations in the steps of PGC development may be due to speciesspecific differences or to technical approaches, e.g. complete loss of function (null mutation) versus reduction of activity (gene knockdown). Development of ES cell technology for producing knockout fishes will be highly desirable in the future. Derivation of stable PGC cultures will represent another future direction. Isolation, cryopreservation and transplantation of germ cells from adult fishes (e.g. endangered species) into germ-cell-absent host species (embryos or gonads) for surrogate reproduction will be of substantial value in sustainable aquaculture and conservation biology.

6 7

8 9

10 11 12 13 14 15

16 17 18 19

20 21 22

This work was supported by the Biomedical Research Council of Singapore (Grant Nos. R-05-1-21-19-404, R-08-1-21-19-585 and SBIC-SSCC C-002-2007), the Ministry of Education of Singapore (Grant No. R-154-000-285-112), the National University of Singapore (Grant No. R-154-000-153-720), and the National Key Basic Research Program of China (Grant Nos. 2004CB117406 and 2010CB126301). The authors would like to express their appreciation to Deng JiaoRong and Zeng QingHua for fish breeding.

23 24


Ikenishi K. Germ plasm in Caenorhabditis elegans, Drosophila and Xenopus. Dev Growth Differ, 1998, 40: 1–10 Braat A K, Speksnijder J E, Zivkovic D. Germ line development in fishes. Int J Dev Biol, 1999, 43: 745–760 Gamo H. On the origin of germ cells and formation of gonad primordia in the medaka, Oryzias latipes. Japan J Zoo, 1961, 13: 9 Hamaguchi S. A light- and electron-microscopic study on the migration of primordial germ cells in the teleost, Oryzias latipes. Cell Tissue Res, 1982, 227: 139–151 You Y L, Lin D J, Su M. The formation of nuage in the oogenesis of the teleost Spiniba rbus caldwelli (Nichols). Acta Zool Sin, 2004, 50: 231–239 Strome S, Lehmann R. Germ versus soma decisions: lessons from flies and worms. Science, 2007, 316: 392–393 Chuma S, Hosokawa M, Tanaka T, et al. Ultrastructural characterization of spermatogenesis and its evolutionary conservation in the germline: Germinal granules in mammals. Mol Cell Endocrinol, 2009, 306: 17–23 Li M, Hong N, Xu H, et al. Medaka vasa is required for migration but not survival of primordial germ cells. Mech Dev, 2009, 126: 366– 381 Yoon C, Kawakami K, Hopkins N. Zebrafish vasa homologue RNA is localized to the cleavage planes of 2- and 4-cell-stage embryos and is expressed in the primordial germ cells. Development, 1997, 124: 3157–3165 Houwing S, Kamminga LM, Berezikov E, et al. A role for Piwi and piRNAs in germ cell maintenance and transposon silencing in Zebrafish. Cell, 2007, 129: 69–82. Strasser M J, Mackenzie N C, Dumstrei K, et al. Control over the morphology and segregation of Zebrafish germ cell granules during embryonic development. BMC Dev Biol, 2008, 8: 58 Li W, Deng F, Wang H, et al. Germ cell-less expression in zebrafish embryos. Dev Growth Differ, 2006, 48: 333–338 Bontems F, Stein A, Marlow F, et al. Bucky ball organizes germ plasm assembly in zebrafish. Curr Biol, 2009, 19: 414–422 Xu H, Li Z, Li M, et al. Boule is present in fish and bisexually expressed in adult and embryonic germ cells of medaka. PLoS One, 2009, 4: e6097 Peng J X, Xie J L, Zhou L, et al. Evolutionary conservation of Dazl genomic organization and its continuous and dynamic distribution throughout germline development in gynogenetic gibel carp. J Exp Zool B Mol Dev Evol, 2009, 312: 855–871 Xu H, Li M, Gui J, et al. Cloning and expression of medaka dazl during embryogenesis and gametogenesis. Gene Expr Patterns, 2007, 7: 332–338 Aoki Y, Nagao I, Saito D, et al. Temporal and spatial localization of three germline-specific proteins in medaka. Dev Dyn, 2008, 237: 800–807 Koprunner M, Thisse C, Thisse B, et al. A zebrafish nanos-related gene is essential for the development of primordial germ cells. Genes Dev, 2001, 15: 2877–2885 Draper B W, McCallum C M, Moens C B. nanos1 is required to maintain oocyte production in adult zebrafish. Dev Biol, 2007, 305: 589–598 Ramasamy S, Wang H, Quach H N, et al. Zebrafish Staufen1 and Staufen2 are required for the survival and migration of primordial germ cells. Dev Biol, 2006, 292: 393–406 Liu L, Hong N, Xu H, et al. Medaka dead end encodes a cytoplasmic protein and identifies embryonic and adult germ cells. Gene Expr Patterns, 2009, 9: 541–548 Weidinger G, Stebler J, Slanchev K, et al. Dead end, a novel vertebrate germ plasm component, is required for zebrafish primordial germ cell migration and survival. Curr Biol, 2003, 13: 1429–1434 Thorpe J L, Doitsidou M, Ho S Y, et al. Germ cell migration in zebrafish is dependent on HMGCoA reductase activity and prenylation. Dev Cell, 2004, 6: 295–302 Schlueter P J, Sang X, Duan C, et al. Insulin-like growth factor receptor 1b is required for zebrafish primordial germ cell migration and survival. Dev Biol, 2007, 305: 377–387

Xu HongYan, et al.

25

26

27

28

29

30 31

32

33

34

35

36

37

38

39 40

41

42

43

44

45 46

Sci China Life Sci

Dumstrei K, Mennecke R, Raz E. Signaling pathways controlling primordial germ cell migration in zebrafish. J Cell Sci, 2004, 117: 4787–4795 Doitsidou M, Reichman-Fried M, Stebler J, et al. Guidance of primordial germ cell migration by the chemokine SDF-1. Cell, 2002, 111: 647–659 Knaut H, Werz C, Geisler R, et al. A zebrafish homologue of the chemokine receptor Cxcr4 is a germ-cell guidance receptor. Nature, 2003, 421: 279–282 Mich J K, Blaser H, Thomas N A, et al. Germ cell migration in zebrafish is cyclopamine-sensitive but Smoothened-independent. Dev Biol, 2009, 328: 342–354 Kuo M W, Wang S H, Chang J C, et al. A novel puf-A gene predicted from evolutionary analysis is involved in the development of eyes and primordial germ-cells. PLoS One, 2009, 4: e4980 Saga Y. Mouse germ cell development during embryogenesis. Curr Opin Genet Dev, 2008, 18: 337–341 Kosaka K, Kawakami K, Sakamoto H, et al. Spatiotemporal localization of germ plasm RNAs during zebrafish oogenesis. Mech Dev, 2007, 124: 279–289 Pepling M E, Wilhelm J E, O’Hara A L, et al. Mouse oocytes within germ cell cysts and primordial follicles contain a Balbiani body. Proc Natl Acad Sci USA, 2007, 104: 187–192 Kobayashi H, Iwamatsu T. Development and fine structure of the yolk nucleus of previtellogenic oocytes in the medaka Oryzias latipes. Dev Growth Differ, 2000, 42: 623–631 Knaut H, Pelegri F, Bohmann K, et al. Zebrafish vasa RNA but not its protein is a component of the germ plasm and segregates asymmetrically before germline specification. J Cell Biol, 2000, 149: 875–888 Theusch E V, Brown K J, Pelegri F. Separate pathways of RNA recruitment lead to the compartmentalization of the zebrafish germ plasm. Dev Biol, 2006, 292: 129–141 Shinomiya A, Tanaka M, Kobayashi T, et al. The vasa-like gene, olvas, identifies the migration path of primordial germ cells during embryonic body formation stage in the medaka, Oryzias latipes. Dev Growth Differ, 2000, 42: 317–326 Yoshizaki G, Sakatani S, Tominaga H, et al. Cloning and characterization of a vasa-like gene in rainbow trout and its expression in the germ cell lineage. Mol Reprod Dev, 2000, 55: 364–371 Xu H, Gui J, Hong Y. Differential expression of vasa RNA and protein during spermatogenesis and oogenesis in the gibel carp (Carassius auratus gibelio), a bisexually and gynogenetically reproducing vertebrate. Dev Dyn, 2005, 233: 872–882 Aoki Y, Nakamura S, Ishikawa Y, et al. Expression and syntenic analyses of four nanos genes in medaka. Zool Sci, 2009, 26: 112–118 Chuma S, Hiyoshi M, Yamamoto A, et al. Mouse tudor repeat-1 (MTR-1) is a novel component of chromatoid bodies/nuages in male germ cells and forms a complex with snRNPs. Mech Dev, 2003, 120: 979–990 Yoshimizu T, Sugiyama N, De Felice M, et al. Germline-specific expression of the Oct-4/green fluorescent protein (GFP) transgene in mice. Dev Growth Differ, 1999, 41: 675–684 Kobayashi T, Yoshizaki G, Takeuchi Y, et al. Isolation of highly pure and viable primordial germ cells from rainbow trout by GFP-dependent flow cytometry. Mol Reprod Dev, 2004, 67: 91– 100 Ciruna B, Weidinger G, Knaut H, et al. Production of maternalzygotic mutant zebrafish by germ-line replacement. Proc Natl Acad Sci USA, 2002, 99: 14919–14924 Saito T, Fujimoto T, Maegawa S, et al. Visualization of primordial germ cells in vivo using GFP-nos1 3'UTR mRNA. Int J Dev Biol, 2006, 50: 691–699 Herpin A, Rohr S, Riedel D, et al. Specification of primordial germ cells in medaka (Oryzias latipes). BMC Dev Biol, 2007, 7: 3 Ding D, Parkhurst S M, Halsell S R, et al. Hsp83 RNA localization during Drosophila oogenesis and embryogenesis. Mol Cell Biol, 1993, 13: 3773–3781

47

48

49

50

51

52

53 54 55

56

57 58 59 60

61

62 63

64 65

66

67

68

69

70

71


445

Wolke U, Weidinger G, Koprunner M, et al. Multiple levels of posttranscriptional control lead to germ line-specific gene expression in the zebrafish. Curr Biol, 2002, 12: 289–294 Mishima Y, Giraldez A J, Takeda Y, et al. Differential regulation of germline mRNAs in soma and germ cells by zebrafish miR-430. Curr Biol, 2006, 16: 2135–2142 Rangan P, DeGennaro M, Lehmann R. Regulating gene expression in the Drosophila germ line. Cold Spring Harb Symp Quant Biol, 2008, 73: 1–8 Hashimoto Y, Maegawa S, Nagai T, et al. Localized maternal factors are required for zebrafish germ cell formation. Dev Biol, 2004, 268: 152–161 Weidinger G, Wolke U, Koprunner M, et al. Regulation of zebrafish primordial germ cell migration by attraction towards an intermediate target. Development, 2002, 129: 25–36 Herpin A, Fischer P, Liedtke D, et al. Sequential SDF1a and b-induced mobility guides Medaka PGC migration. Dev Biol, 2008, 320: 319–327 Saffman E E, Lasko P. Germline development in vertebrates and invertebrates. Cell Mol Life Sci, 1999, 55: 1141–1163 Santos A C, Lehmann R. Germ cell specification and migration in Drosophila and beyond. Curr Biol, 2004, 14: R578–589 Maufroid J P, Capuron A P. A demonstration of cellular interactions during the formation of mesoderm and primordial germ cells in Pleurodeles waltlii. Differentiation, 1985, 29: 20–24 Stebler J, Spieler D, Slanchev K, et al. Primordial germ cell migration in the chick and mouse embryo: the role of the chemokine SDF-1/CXCL12. Dev Biol, 2004, 272: 351–361 Albert M, Peters A H. Genetic and epigenetic control of early mouse development. Curr Opin Genet Dev, 2009, 19: 113–121 Wylie C. Germ cells. Cell, 1999, 96: 165–174 Raz E. Primordial germ-cell development: the zebrafish perspective. Nat Rev Genet, 2003, 4: 690–700 Blaser H, Eisenbeiss S, Neumann M, et al. Transition from nonmotile behaviour to directed migration during early PGC development in zebrafish. J Cell Sci, 2005, 118: 4027–4038 Weidinger G, Wolke U, Koprunner M, et al. Identification of tissues and patterning events required for distinct steps in early migration of zebrafish primordial germ cells. Development, 1999, 126: 5295–5307 Kunwar P S, Lehmann R. Developmental biology: Germ-cell attraction. Nature, 2003, 421: 226–227 Mahabaleshwar H, Boldajipour B, Raz E. Killing the messenger: The role of CXCR7 in regulating primordial germ cell migration. Cell Adh Migr, 2008, 2: 69–70 Van Doren M, Broihier H T, Moore L A, et al. HMG-CoA reductase guides migrating primordial germ cells. Nature, 1998, 396: 466–469 Tanaka S S, Yamaguchi Y L, Tsoi B, et al. IFITM/Mil/fragilis family proteins IFITM1 and IFITM3 play distinct roles in mouse primordial germ cell homing and repulsion. Dev Cell, 2005, 9: 745–756 Molyneaux K A, Zinszner H, Kunwar P S, et al. The chemokine SDF1/CXCL12 and its receptor CXCR4 regulate mouse germ cell migration and survival. Development, 2003, 130: 4279–4286 Ara T, Nakamura Y, Egawa T, et al. Impaired colonization of the gonads by primordial germ cells in mice lacking a chemokine, stromal cell-derived factor-1 (SDF-1). Proc Natl Acad Sci USA, 2003, 100: 5319–5323 Slanchev K, Stebler J, de la Cueva-Mendez G, et al. Development without germ cells: The role of the germ line in zebrafish sex differentiation. Proc Natl Acad Sci USA, 2005, 102: 4074–4079 Kim C H, Broxmeyer H E. In vitro behavior of hematopoietic progenitor cells under the influence of chemoattractants: stromal cell-derived factor-1, steel factor, and the bone marrow environment. Blood,1998, 91: 100–110 Staller P, Sulitkova J, Lisztwan J, et al. Chemokine receptor CXCR4 downregulated by von Hippel-Lindau tumour suppressor pVHL. Nature, 2003, 425: 307–311 Hernandez P A, Gorlin R J, Lukens J N, et al. Mutations in the chemokine receptor gene CXCR4 are associated with WHIM syndrome, a combined immunodeficiency disease. Nat Genet, 2003,

446

72

73 74

75

76 77

78

79

80

81

82

83

84

85

86

87

88

Xu HongYan, et al.

Sci China Life Sci

34: 70–74 Gui J F, Liang S C, Zhu L F. Preliminary confirmation of gynogenetic reproductive mode in artificially multiple tetraploid allogynogenetic silver crucial carp. Chinese Sci Bull, 1993, 38: 327– 331 Wu N, Yue H M, Chen B, et al. Histone H2A has a novel variant in fish oocytes. Biol Reprod, 2009, 81: 275–283 Tajima A, Naito M, Yasuda Y, et al. Production of germ line chimera by transfer of primordial germ cells in the domestic chicken (Gallus domesticus). Theriogenology, 1993, 40: 509–519 Brinster R L, Zimmermann J W. Spermatogenesis following male germ-cell transplantation. Proc Natl Acad Sci USA, 1994, 91: 11298–11302 Honaramooz A, Megee S O, Dobrinski I. Germ cell transplantation in pigs. Biol Reprod, 2002, 66: 21–28 Kang S J, Choi J W, Kim S Y, et al. Reproduction of wild birds via interspecies germ cell transplantation. Biol Reprod, 2008, 79: 931–937 Yoshizaki G, Takeuchi Y, Sakatani S, et al. Germ cell-specific expression of green fluorescent protein in transgenic rainbow trout under control of the rainbow trout vasa-like gene promoter. Int J Dev Biol, 2000, 44: 323–326 Okutsu T, Yano A, Nagasawa K, et al. Manipulation of fish germ cell: visualization, cryopreservation and transplantation. J Reprod Dev, 2006, 52: 685–693 Takeuchi Y, Higuchi K, Yatabe T, et al. Development of spermatogonial cell transplantation in Nibe croaker, Nibea mitsukurii (Perciformes, Sciaenidae). Biol Reprod, 2009, 81: 1055–63 Nagler J J, Cloud J G, Wheeler P A, et al. Testis transplantation in male rainbow trout (Oncorhynchus mykiss). Biol Reprod, 2001, 64: 644–646 Takeuchi Y, Yoshizaki G, Takeuchi T. Generation of live fry from intraperitoneally transplanted primordial germ cells in rainbow trout. Biol Reprod, 2003, 69: 1142–1149 Saito T, Goto-Kazeto R, Arai K, et al. Xenogenesis in teleost fish through generation of germ-line chimeras by single primordial germ cell transplantation. Biol Reprod, 2008, 78: 159–166 Okutsu T, Suzuki K, Takeuchi Y, et al. Testicular germ cells can colonize sexually undifferentiated embryonic gonad and produce functional eggs in fish. Proc Natl Acad Sci USA, 2006, 103: 2725–2729 Fong C Y, Peh G S, Gauthaman K, et al. Separation of SSEA-4 and TRA-1-60 labelled undifferentiated human embryonic stem cells from a heterogeneous cell population using magnetic-activated cell sorting (MACS) and fluorescence-activated cell sorting (FACS). Stem Cell Rev Rep, 2009, 5: 72–80 Yano A, von Schalburg K, Cooper G, et al. Identification of a molecular marker for type A spermatogonia by microarray analysis using gonadal cells from pvasa-GFP transgenic rainbow trout (Oncorhynchus mykiss). Mol Reprod Dev, 2009, 76: 246–254 Matsui Y, Zsebo K, Hogan B L. Derivation of pluripotential embryonic stem cells from murine primordial germ cells in culture. Cell, 1992, 70: 841–847 van de Lavoir M C, Diamond J H, Leighton P A, et al. Germline transmission of genetically modified primordial germ cells. Nature,

89

90 91

92

93

94

95

96

97

98

99 100

101 102 103 104 105

106 107 108


2006, 441: 766–769 Fan L, Moon J, Wong T T, et al. Zebrafish primordial germ cell cultures derived from vasa::RFP transgenic embryos. Stem Cells Dev, 2008, 17: 585–597 Brinster R L. Germline stem cell transplantation and transgenesis. Science, 2002, 296: 2174–2176 Hofmann M C, Hess R A, Goldberg E, et al. Immortalized germ cells undergo meiosis in vitro. Proc Natl Acad Sci USA, 1994, 91: 5533–5537 Feng L X, Chen Y, Dettin L, et al. Generation and in vitro differentiation of a spermatogonial cell line. Science, 2002, 297: 392–395 Guan K, Nayernia K, Maier L S, et al. Pluripotency of spermatogonial stem cells from adult mouse testis. Nature, 2006, 440: 1199–1203 Hong Y, Winkler C, Schartl M. Pluripotency and differentiation of embryonic stem cell lines from the medakafish (Oryzias latipes). Mech Dev, 1996, 60: 33–44 Hong Y, Winkler C, Schartl M. Production of medakafish chimeras from a stable embryonic stem cell line. Proc Natl Acad Sci USA, 1998, 95: 3679–3684 Ma C, Fan L, Ganassin R, et al. Production of zebrafish germ-line chimeras from embryo cell cultures. Proc Natl Acad Sci USA, 2001, 98: 2461–2466 Chen S L, Sha Z X, Ye H Q, et al. Pluripotency and chimera competence of an embryonic stem cell line from the sea perch (Lateolabrax japonicus). Mar Biotechnol (NY), 2007, 9: 82–91 Chen S L, Ren G C, Sha Z X, et al. Establishment of a continuous embryonic cell line from Japanese flounder Paralichthys olivaceus for virus isolation. Dis Aquat Organ, 2004, 60: 241–246 Yi M S, HONG N, Li Z D, et al. Medaka fish stem cells and their applications. Sci China Life Sci, 2010, 53: 426–434 Hong N, Li M, Zeng Z, et al. Accessibility of host cell lineages to medaka stem cells depends on genetic background and irradiation of recipient embryos. Cell Mol Life Sci. 2010, 10. [Epub ahead of print]PMID: 20063174 Hong Y, Gui J, Chen S, et al. Embryonic stem cell in fish. Acta Hydrobiol Sin, 2003, 49: 14 Yi M, Hong N, Hong Y. Generation of medaka fish haploid embryonic stem cells. Science, 2009, 326: 4 Ramasamy R, Ricci J A, Palermo G D, et al. Successful fertility treatment for Klinefelter’s syndrome. J Urol, 2009, 182: 1108–1113 Gurdon J B, Melton D A. Nuclear reprogramming in cells. Science, 2008, 322: 1811–1815 Wakamatsu Y, Ju B, Pristyaznhyuk I, et al. Fertile and diploid nuclear transplants derived from embryonic cells of a small laboratory fish, medaka (Oryzias latipes). Proc Natl Acad Sci USA, 2001, 98: 1071–1076 Campbell K H, McWhir J, Ritchie W A, et al. Sheep cloned by nuclear transfer from a cultured cell line. Nature, 1996, 380: 64–66 Tsai M C, Takeuchi T, Bedford J M, et al. Alternative sources of gametes: reality or science fiction? Hum Reprod, 2000, 15: 988–998 Tesarik J. Reproductive semi-cloning respecting biparental embryo origin: embryos from syngamy between a gamete and a haploidized somatic cell. Hum Reprod, 2002, 17: 1933–1937