Role of Fibroblast Growth Factors and Their Receptors in Mouse Primordial Germ Cell Growth¹

^a Cell Biology of Development and Differentiation Group, ABL-Basic Research Program and ^b Intramural Research and Support Program, SAIC-Frederick, National Cancer Institute-Frederick Cancer Research and Development Center, Frederick, Maryland 21702–1201

	ABSTRACT

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Primordial germ cells (PGCs) are the embryonic progenitors of mature germ cells. During their proliferative stage, murine PGCs may be transiently cultured on mitotically inactive feeder layers. This culture system has permitted identification of several growth factors active toward PGCs. We and others have previously identified basic fibroblast growth factor (bFGF) as a powerful mitogen in this system. Here we characterize some of the functions of bFGF in PGC culture. Our data demonstrate that fibroblast growth factor (FGF) receptors I and II are present in the developing gonad and are consistent with expression of these receptors by PGCs. Moreover, PGCs can bind radiolabeled bFGF in vitro, demonstrating that the factor can act directly on these cells. While mitotic PGCs of either sex are shown to bind radiolabeled bFGF, oogonia that are undergoing meiotic arrest exhibit reduced bFGF binding, indicating potential developmental regulation of an FGF receptor.

	INTRODUCTION

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Mature mouse gametes can be traced back to a small population of cells in the embryo known as primordial germ cells (PGCs). The PGC-specific expression of the histochemical marker alkaline phosphatase has led to a description of their embryonic development [1]. At 7 days postcoitus (dpc), PGCs are located caudal to the primitive streak, near the base of the allantois [2]. These cells migrate through the dorsal mesentery and enter the developing fetal gonad, the genital ridge, between 10.5 and 12.5 dpc. This period of migration is accompanied by rapid proliferation, with an estimated doubling time of 16 h [3–5]. By 13.5 dpc, the final population of approximately 25 000 germ cells is present in the genital ridges. In the male, germ cells enter a mitotic arrest, while in females, germ cells arrest at meiotic prophase I [6].

PGCs can be transiently cultured on feeder layers expressing the transmembrane-bound isoform of steel factor (SLF) [7, 8], the product of the murine Sl locus. SLF, together with leukemia inhibitory factor (LIF), promotes PGC survival by suppressing apoptosis [9]. Basic fibroblast growth factor (bFGF) is also active in PGC cultures, promoting germ cell proliferation. If PGCs are cultured with SLF, LIF, and bFGF, they can form permanent cell lines, termed embryonic germ (EG) cells. EG cells share many properties with embryonic stem cells, including the ability to colonize the germ line.

We have undertaken a more thorough examination of the effects of bFGF in PGC culture, both to better understand the role of fibroblast growth factors (FGFs) in PGC development and to expedite the isolation of EG cell types from other species. Our results strongly suggest that bFGF can work directly on PGCs, as opposed to an indirect effect exerted through the feeder layer or through somatic cells plated along with PGCs. Moreover, mitotic PGCs express an FGF receptor activity. FGF-binding activity appears to be regulated in a developmental- and sex-specific manner.

	MATERIALS AND METHODS

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PGC Isolation and Culture

PGCs were isolated from B6C3F1(C57Bl/6 x C3H/HeJ) mice and cultured as previously described [8, 10]. Briefly, single-cell suspensions were prepared by trypsinization and trituration of either caudal regions of 8.5 dpc embryos or the genital ridges of 11.5–13.5 dpc embryos. The sex of 12.5 dpc and 13.5 dpc embryos was determined by microscopic inspection for the presence of testis cords. Immunomagnetic PGC purification, using TG-1 antisera recognizing a PGC cell surface antigen, was performed as described previously [11], except that 20 µl of TG-1 supernatant was used per preparation.

PGC cultures were largely performed as previously described. Briefly, cell suspensions were plated on irradiated (5000 rads) STO feeder layers in 96-well microtiter dishes that had been previously treated with 0.1% gelatin. For growth assays, approximately 0.20–0.25 equivalents of 11.5 dpc embryos were plated per well in 200 µl of high-glucose Dulbecco's Modified Eagle's medium (DMEM) supplemented with 15% fetal bovine serum (FBS), sodium pyruvate, and glutamine. Cultures were re-fed daily. PGCs were identified by alkaline phosphatase histochemistry, as previously described [7, 8]. Each determination represents the average and SD of at least six wells.

FGF-Binding Assays

PGC cell suspensions were obtained as described above, but trypsinization was terminated after 5 min at 37°C by addition of one drop of FBS. The cell suspensions were quickly washed twice in 1 ml binding buffer (a 1:1 mixture of DMEM and Ham's F-12 supplemented with insulin and transferrin [Sigma Chemical Co., St. Louis, MO] and 0.1% BSA). Cell suspensions were diluted to 6 x 10⁶ total cells/ml. Cell suspension (100 µl) was incubated with 750 000 cpm of ¹²⁵I-bFGF (800–1200 Ci/mmol; Amersham, Arlington Heights, IL) with agitation for 45 min at 4°C. Where indicated, some samples had been previously supplemented with 0.5 µg unlabeled recombinant human bFGF (Life Technologies, Bethesda, MD). Suspensions were pelleted through 200 µl of cold FBS, resuspended in 0.2 ml of a 1:1 mixture of binding buffer and FBS, and pelleted in a cytospin for 10 min at 700 rpm onto 3-aminopropylethoxysilane-treated microscope slides (Digene Corp., Silver Springs, MD). The slides were fixed for 10 min in cold 70% ethanol, followed by 10 min in 4% paraformaldehyde in PBS. PGCs were histochemically stained for alkaline phosphatase as previously described [7, 8]. Slides were immersed twice in a 1:1 mixture of Kodak NTB-2 (Eastman Kodak, Rochester, NY) autoradiographic emulsion:water at room temperature and stored in the dark at 4°C for 26 days prior to development. Factor binding to the cell surface was visualized by the appearance of grains caused by ¹²⁵I exposure of the photographic emulsion.

Differences between Experimental and Plus Competitor means, as well as trend hypotheses, were determined with the use of t-tests and Wilcoxon rank sum tests. To control the type I family error rate, we used the Bonferroni criterion and set the alpha level at 0.01 to be required for significance. As outcomes from t-tests and Wilcoxon tests gave identical interpretations, probability values reported are from t-tests.

RNA Analysis

RNAs were prepared with RNAzol-B as recommended by the manufacturer (Tel-Test Inc., Friendswood, TX). NIH-3T3 cells and BM-Wt cells were grown as previously described [8, 10] and washed briefly in PBS prior to RNA extraction. Brain and liver were isolated from adult B6C3F1 animals and used immediately for RNA isolation. Genital ridges were collected from embryos derived from B6C3F1 intercrosses as described previously [8, 10]. Purified populations of PGCs were isolated as described above. RNA probes labeled with ³²P were prepared and RNase protection assays performed as described previously [12] and analyzed on 6% acrylamide-urea sequencing gels. A ß-actin internal control (Ambion, Austin, TX) was included in each assay. The protected fragments used were the 277-base pair (bp) PvuII to SphI fragment of the extracellular domain of FGF receptor (FGFR)-1 [13], the 326-bp EcoRV to EcoRI fragment in the extracellular domain of FGFR-2 [14], the DdeI fragment in the extracellular domain of FGFR-3 [15], and the 387-bp EcoRV to BamHI fragment of FGFR-4 [16]. All fragments were subcloned in pBluescript SK or KS and transcribed with T7 or T3 polymerase (Stratagene, La Jolla, CA) as appropriate.

Degenerate oligonucleotide analysis of FGFR expression by reverse transcription-polymerase chain reaction (RT-PCR) was performed as previously described [17] with the following exceptions. All RNA preparations were treated with DNase I (Life Technologies) to eliminate genomic DNA contamination. Random-primed cDNA was prepared using Superscript II (Life Technologies) as suggested by the manufacturer. PCR was performed with Expand Taq (Boehringer-Mannheim, Indianapolis, IN) in the manufacturer's supplied buffer supplemented with 10% glycerol.

	RESULTS

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Basic FGF Stimulated the Growth of Premigratory and Postmigratory PGCs

To more thoroughly define the functions of bFGF in the in vitro PGC culture system, we plated 11.5 dpc PGCs in the presence of bFGF under a variety of conditions. As previously shown, bFGF stimulated the accumulation of PGCs plated on mitotically inactivated STO fibroblast feeder layers (Fig. 1A). We have previously shown that the optimal stimulatory effect of bFGF in this system occurs at a bFGF concentration of 1 ng/ml [10]. Given the important role that LIF plays in PGC culture [18], and as bFGF can rapidly increase the accumulation of LIF RNA by STO feeder layers (unpublished results), we questioned whether the effect of bFGF on PGCs was indirect, acting primarily through the feeder layer. To determine the necessity of the feeder layer to mediate the bFGF effect, PGCs obtained from 11.5 dpc embryos were cultured on gelatinized tissue culture plastic in the absence of a STO feeder layer. While PGC survival under these conditions is poor, bFGF still exhibited a strong positive effect toward PGCs (Fig. 1B), indicating that the bFGF effect is independent of the STO feeder layer. In these crude cell suspensions, PGCs account for less than 2% of all cells (unpublished results). Under these conditions, the somatic cells that accompany the PGCs form a monolayer during 24–48 h of culture. As these somatic cells could also indirectly mediate the effects of bFGF, we plated immunomagnetically purified PGCs (more than 80% free of somatic cells) on STO feeder cells. Again, bFGF stimulated PGC growth under these conditions (Fig. 1C). Together, these results demonstrate that bFGF stimulates PGC growth either in the absence of the fibroblast feeder layer, or stimulates highly purified PGCs in the presence of a feeder layer.

View larger version (18K): [in this window] [in a new window]   FIG. 1. Analysis of the effect of bFGF on PGC growth in culture. PGCs obtained from 11.5 dpc embryos were cultured in 96-well microtiter plates in the absence (open bars) or presence (closed bars) of 1 ng/ml recombinant human bFGF, and the number of PGCs was determined by alkaline phosphatase staining. A) A single-cell suspension of urogenital ridges was plated on STO feeder layers. B) PGCs were cultured in gelatinized wells in the absence of feeder cells. C) PGCs at 11.5 dpc were immunomagnetically purified and cultured on STO feeder cells.

FGFR-1 and FGFR-2 Were Expressed in the Embryonic Gonad and Were Readily Detectable in Highly Purified PGC RNA

To further explore whether bFGF acts directly on PGCs, we sought to determine whether PGCs express an FGF receptor. Four murine high-affinity tyrosine kinase receptor genes for FGFs have been identified to date [13–16]. To investigate whether PGCs might express one of these receptors, 11.5 dpc genital ridge RNA was tested for expression for all four genes by RNase protection assays. FGFR-1 and FGFR-2 RNA were readily detected in the genital ridge RNA (Fig. 2).

View larger version (27K): [in this window] [in a new window]   FIG. 2. RNase protection analysis for FGF receptors. Five micrograms of 11.5 dpc genital ridge RNA was analyzed for the presence of each of the four known FGF receptor gene RNAs. A tRNA negative control was included in each assay. Each assay included an internal actin control for RNA quality and loading. The actin controls for each RNase protection assay are shown. The positive control RNAs are as follows: FGFR-1, NIH3T3 cells; FGFR-2, BM-wt cells [8]; FGFR-3, adult brain; FGFR-4, adult liver.

We used a degenerate oligonucleotide RT-PCR approach [17] to determine whether FGF receptors were encoded in highly purified (more than 90% free of somatic cells) 11.5 dpc PGC RNA. In this method, degenerate primers recognizing conserved motifs in the kinase domains of all four FGF receptors are used to amplify a 341-bp band from all FGFR transcripts. The resulting PCR fragment is then digested with restriction enzymes diagnostic for each FGFR. FGFR sequences were readily detected in 11.5 dpc PGC RNA as well as in 11.5 dpc whole embryo, which served as a positive control. No amplification was detectable in control reactions lacking reverse transcriptase (Fig. 3A). PvuII digestion of the PGC band readily revealed 195/146-bp and 233/108-bp bands indicative of FGFR-1 and FGFR-2, respectively. The 175/166-bp bands indicative of FGFR-3 were not readily apparent following PstI digestion; 216/125-bp EcoRI bands, indicative of FGFR-4, were faintly detectable (Fig. 3B). Thus, FGFR-1, FGFR-2, or potentially FGFR-4 may be expressed by proliferative PGCs.

View larger version (41K): [in this window] [in a new window]   FIG. 3. Degenerate oligonucleotide RT-PCR analysis of FGF receptor expression. A) FGF receptor sequences were amplified from RNA obtained from highly purified (> 90%) 11.5 dpc PGCs, or from whole 11.5 dpc embryos (WE) using degenerate primers that recognize conserved sequences in the kinase domains of all four FGF receptors. A 341-bp band is expected for all four receptors. The 396- and 344-bp molecular size markers are indicated. M, size marker lane; +/- indicate presence or absence of reverse transcriptase in the cDNA preparation step. B) The PGC band was purified and digested with PvuII (Pv), PstI (Ps), EcoRI (Ec), or was left uncut (Un). The sizes of expected fragments for each FGFR are FGFR-1, 195/146 in the PvuII digest; FGFR-2, 233/108 in the PvuII digest; FGFR-3, 175/166 in the PstI digest; FGFR4, 216/125 in the EcoRI digest.

PGCs Bound Iodinated bFGF

To more completely distinguish FGF receptor expression in germ cells from that in somatic cells, and to confirm that PGCs express a surface FGF-binding activity, we developed a method in which ex vivo PGCs are incubated with ¹²⁵I-bFGF, fixed on microscope slides, stained for alkaline phosphatase, and exposed to autoradiographic emulsion. Figure 4 presents a sample photomicrograph from such an experiment performed on 11.5 dpc genital ridges. The presence of autoradiographic grains over alkaline phosphatase-expressing PGCs is clearly seen. It is also notable that somatic cells (not expressing alkaline phosphatase) in this preparation bound bFGF, as might be expected for mesodermally derived genital ridge tissue. The level of ¹²⁵I-bFGF binding by alkaline phosphatase-positive and by alkaline phosphatase-negative cell types appeared comparable. The specificity of this binding was demonstrated by counting grains over stained PGCs and was compared to that of samples in which an excess of unlabeled bFGF had been added (Fig. 5). Binding of ¹²⁵I-bFGF by most somatic cells was also sensitive to the presence of unlabeled bFGF (not shown).

View larger version (144K): [in this window] [in a new window]   FIG. 4. Basic FGF binding by PGCs. Single-cell suspensions of genital ridges from 11.5 dpc embryos were incubated in the presence of 125I-bFGF as described in Materials and Methods. PGCs were identified by alkaline phosphatase staining, which is seen as a red precipitate surrounding the cell (large arrow). Cells not staining for alkaline phosphatase are considered to be somatic cells (open arrowheads). Grains (small arrow) observed in areas lacking cells are considered background.

View larger version (36K): [in this window] [in a new window]   FIG. 5. Analysis of 125I-bFGF binding by PGCs. PGCs from embryos of the indicated age and sex were analyzed for binding of 125I-bFGF. The average number and SD of autoradiographic grains over an alkaline phosphatase-expressing PGC was determined (solid bars). In each case, nonspecific binding was monitored by a control incubation containing 0.5 µg unlabeled bFGF competitor (gray bars). At least 10 cells were counted to determine an average number of grains. The mean levels of grains per PGC under the Experimental condition (total binding) significantly exceeded (p = 0.0001) those under the Plus Competitor condition (nonspecific binding) at every time except for females at 13.5 dpc (p = 0.1085).

PGCs Demonstrated Developmental- and Sex-Specific Changes in bFGF Binding

We also investigated potential developmental regulation of this FGF-binding activity by performing this procedure on germ cells obtained from 8.5 and 11.5 dpc embryos as well as samples from sex-segregated 12.5 and 13.5 dpc embryos, the earliest stages at which gender is morphologically distinguishable. The results from this experiment, also presented in Figure 5, illustrate that at all stages tested, male germ cells exhibited specific binding of ¹²⁵I-bFGF. Surprisingly, while germ cells obtained from 12.5 dpc female embryos also bound bFGF, binding was reduced to nonspecific levels by 13.5 dpc. At this stage most oogonia are entering meiotic prophase I [19].

	DISCUSSION

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The results presented here confirm previous observations that bFGF stimulates PGC growth in culture. This effect is apparent in the absence of a preformed feeder layer or with purified PGCs plated on a feeder layer. Basic FGF stimulates PGC growth on a variety of feeder layers including bone marrow-derived stromal cells (Sl220) [20], primary mouse fibroblasts [21], and the established mouse fibroblast cell lines STO, NIH3T3, and 10T1/2 cells (unpublished results). The ability of bFGF to stimulate PGC growth independently of the feeder cell type suggests that it may exert its action, in part, directly on the PGCs. The demonstration that ¹²⁵I-bFGF can bind to PGCs shows that PGCs have a cell surface FGF receptor. RNase protection analyses indicate that FGFR-1 and FGFR-2 are expressed in the 11.5 dpc urogenital ridge. Single primer pair RT-PCR analysis suggests that FGFR transcripts are encoded in the RNA of highly purified 11.5 dpc PGCs. While it is possible that bFGF can promote PGC growth indirectly through both the STO feeder cells and the somatic cells of the urogenital ridge, our data indicate that PGCs can bind and respond to bFGF. Together, these results are most consistent with a model in which bFGF acts directly on PGCs.

At present, there are 16 members of the FGF ligand family. The typical FGF receptor has a split intracellular tyrosine kinase domain and an extracellular domain containing up to three ligand-binding immunoglobin-like loops [22]. Alternative splicing within the extracellular domains of FGFR genes can generate great diversity of ligand-binding domains, which can result in different ligand-binding specificities [23–25]. Additionally, each FGF receptor may transduce a particular mitogenic potential [23]. However, as individual receptors can be activated by several different ligands [14, 23–25], specificity may partly depend upon spatial and temporal expression patterns. An exception is FGF7 (keratinocyte growth factor, KGF), which selectively activates a differentially spliced form of FGFR-2 [26]. Using RNase protection analysis, we have detected KGF receptor transcripts in urogenital ridge RNA (unpublished results). Neither germ cell nor gonadal defects have been observed in KGF-deficient mice [27].

Do FGFs contribute to the in vivo development of the germ line? The pattern of expression of several FGFs, but most notably FGF4, suggests that it may be expressed in or near PGCs. In the early postimplantation embryo, FGF4 is expressed in the embryonic ectoderm at the egg cylinder stage, then along the primitive streak and in the primitive dorsal mesoderm [28–30]. These areas of FGF4 expression in the preimplantation and postimplantation embryo closely match expected locations for PGCs. Other FGFs, especially FGF-3 and FGF-5, have also been shown to be expressed in tissues immediately adjacent to those expressing FGF-4 [28, 29, 31–34]. These observations suggest that FGFs may be continuously available to premigratory and migratory PGCs. Mice lacking either a functional FGF-3 [34] or FGF-5 [35] gene are fertile, indicating that some PGCs mature in the absence of either factor. Embryos deficient in FGF-4 cease development prior to PGC proliferation [36], as do embryos lacking functional FGFR-1 [37, 38], preventing an analysis of PGC development in these embryos. Mice lacking a functional FGFR-2 are not presently available. FGF may play important roles during gametogenesis, as bFGF has been detected both in murine ovarian follicles [39] and in rat spermatocytes [40]. Another member of this growth factor family, FGF-8, may also be present in prespermatogonia between 15 and 17 dpc [41].

The reduction in FGF binding by 13.5 dpc oogonia as they enter meiosis was an unexpected finding. The ability to generate EG cell colonies from PGCs declines beyond 11.5 dpc, with only male EG cells being obtained from 12.5 dpc embryos [42]. Reduced expression of an FGF receptor in female cells provides an explanation for this observation and suggests a mechanism. In both male and female germ cells, c-kit RNA levels decline at 13.5 dpc [43]. It is interesting to speculate that a function of FGFR activation is to maintain c-kit expression in PGCs, thereby contributing to the proliferative stage. This model would predict that in females, FGF receptor expression is reduced by 13.5 dpc, such that the down-regulation of c-kit cannot be prevented by treatment with bFGF. In male PGCs, continuing FGFR expression and subsequent activation may maintain c-kit expression.

When transplanted back into an animal, EG cells can generate teratomas and teratocarcinomas [20, 44] (unpublished results). If a combination of SLF, LIF, and FGF is sufficient to increase the proliferative potential of PGCs resulting in EG cells, what are the limits on PGC proliferation in vivo? Two explanations seem possible. First, in vivo, soluble factors may limit PGC growth. As suggested from the work of Godin and Wylie [45], members of the transforming growth factor ß family may be candidates for such an activity. Bone morphogenetic protein 2, a member of the transforming growth factor ß family, is known to reverse the positive effects of FGF-4 on limb bud development [46], suggesting that an analogous system may limit PGC growth. Alternatively, PGC growth may be limited by physical isolation from one or more factors. In this regard, it is interesting to speculate that FGFs may not be widely expressed in the genital ridge. As PGCs enter the genital ridge they may become isolated from one or more FGFs, contributing to the cessation of proliferation. Further examination of the expression patterns of FGFs and FGFRs in the developing gonad may help elucidate the role of FGFs in germline development.

	ACKNOWLEDGMENTS

 
We gratefully acknowledge Dr. Linzhao Cheng for many helpful discussions. We also thank Drs. Claudio Basilico, David Givol, Mitchell Goldfarb, Jean Hebert, Gail Martin, and Andrew McMahon for sharing cDNA probes. We thank Lynn White, Theresa Shatzer, and Mike Ziemak for their contribution to this work. We also acknowledge Dr. W. Gregory Alvord for his help with statistical analysis.

	FOOTNOTES

 
¹ J.R. was supported by the Cooperative States Research Service, U.S. Department of Agriculture under Agreement No. 93–37205–9074. Research is sponsored in part by the National Cancer Institute, DHHS, under contract with ABL and SAIC. The contents of this publication do not necessarily reflect the views or policies of the Department of Health and Human Services, nor does the mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government.

² Correspondence: Peter J. Donovan, Thomas Jefferson University, BLSB Room 706, 233 South Tenth St., Philadelphia, PA 19107. FAX: (215) 923-4153.

³ Current address: College of Medicine, Molecular Genetics and Microbiology, University of Florida, Gainesville, FL 32610–0266.

Accepted: July 8, 1998.

Received: October 29, 1997.

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