BMP signaling regulates PGC numbers and motility in organ culture

Mechanisms of Development 124 (2007) 68–77 www.elsevier.com/locate/modo

BMP signaling regulates PGC numbers and motility in organ culture Brian M. Dudley a, Chris Runyan b, Yutaka Takeuchi c, Kyle Schaible b, Kathleen Molyneaux a,* b

a Department of Genetics, Case Western Reserve University, 10900 Euclid Ave., Cleveland, OH 44106, USA Division of Developmental Biology, Cincinnati Children’s Hospital Research Foundation, 3333 Burnet Ave., Cincinnati, OH 45229, USA c Department of Aquatic Biosciences, Tokyo University of Fisheries, 4-5-7 Konan, Minato-ku, Tokyo 108-8477, Japan

Received 18 July 2006; received in revised form 8 September 2006; accepted 27 September 2006 Available online 30 September 2006

Abstract Members of the bone morphogenetic protein (BMP) family play diverse roles in multiple developmental processes. However, in the mouse, mutations in many BMPs, BMP receptors and signaling components result in early embryonic lethality making it difficult to analyze the role of these factors during organogenesis or tissue homeostasis in the adult. To bypass this early lethality, we used an organ culture system to study the role of BMPs during primordial germ cell (PGC) migration. PGCs are the embryonic precursors of the sperm and eggs. BMPs induce formation of primordial germ cells within the proximal epiblast of embryonic day 7.5 (E7.5) mouse embryos. PGCs then migrate via the gut to arrive at the developing gonads by E10.5. Addition of BMP4 or the BMP-antagonist Noggin to transverse slices dissected from E9.5 embryos elevated PGC numbers or reduced PGC numbers, respectively. Noggin treatment also slowed and randomized PGC movements, resulting in a failure of PGCs to colonize the urogenital ridges (UGRs). Based on p-Smad1/5/8 staining, migratory PGCs do not respond to endogenous BMPs. Instead, the somatic cells of the urogenital ridges exhibit elevated p-Smad1/5/ 8 staining revealing active BMP signaling within the UGRs. Noggin treatment abrogated p-Smad staining within the UGRs and blocked localized expression of Kitl, a cytokine known to regulate the survival and motility of PGCs and Id1, a transcription factor expressed within the UGRs. We propose that BMP signaling regulates PGC migration by controlling gene expression within the somatic cells along the migration route and within the genital ridges.  2006 Elsevier Ireland Ltd. All rights reserved. Keywords: Bone morphogenetic proteins; Primordial germ cells; Migration; Mouse; Kitl

1. Introduction Bone morphogenetic proteins (BMPs) are members of the transforming growth factor b (TGFb) superfamily. Members of this family signal through heteromeric complexes composed of type I and type II serine–threonine kinase receptors. Binding of BMPs to their receptors induces phosphorylation of the BMP-specific smads (Smad1, Smad5, and Smad8). p-Smads1/5/8 then associate with Smad4, translocate into the nucleus, and activate the transcription of BMP-target genes (reviewed in tenDijke et al., 2003; Zwijsen et al., 2003). In the mouse, targeted or spon*

Corresponding author. Tel.: +1 216 368 1823; fax: +1 216 368 0491. E-mail address: [email protected] (K. Molyneaux).

taneous mutations in various Bmps, Bmp receptors and Smads have confirmed the roles of bone morphogenetic proteins in the formation of the skeletal system (Kingsley et al., 1992; Storm et al., 1994; Dudley et al., 1995; Luo et al., 1995) and mesoderm induction (Mishina et al., 1995, 1999; Sirard et al., 1998; Beppu et al., 2000), as well as development of the heart, nervous system, and urogenital system (reviewed in Zhao, 2003). BMPs have known roles in germ cell development and function. BMP2 (Ying and Zhao, 2001), BMP4 (Lawson et al., 1999), and BMP8b (Ying et al., 2000) control the formation and early proliferation of primordial germ cells (PGCs). These factors are thought to be released from extraembryonic tissues and create a permissive environment for PGC formation in the epiblast (de Sousa Lopes

0925-4773/$ - see front matter  2006 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.mod.2006.09.005

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et al., 2004). Additionally, members of the BMP family are required for germ cell function in adult mice. Targeted deletion of Gdf9 blocks the growth of primary follicles resulting in infertile females (Dong et al., 1996). Additionally, targeted deletions of Bmp15 and Bmpr1b/Alk6 cause female infertility due to defects in ovulation and cumulus cell expansion (reviewed in Shimasaki et al., 2004) and mutations in Bmp7, Bmp8a, Bmp8b, and Bmp4 block spermatogenesis (Zhao et al., 1996, 1998; Hu et al., 2004). The requirement for BMPs in fertility is conserved in a number of species. Two Bmp orthologues, glass-bottom boat (Gbb) and decapentaplegic (Dpp), are expressed in the cap and sheath cells of the Drosophila ovary and regulate differentiation of the germ-line stem cells (Song et al., 2004). Gbb and Dpp play similar roles in the fly testis (Kawase et al., 2004). BMPs are important in controlling PGC formation and gametogenesis, but the early lethality caused by loss of Bmp4 (Winnier et al., 1995), the BMP type I receptors (Bmpr1a and Acvr1) (Mishina et al., 1995, 1999; Gu et al., 1999), the BMP type II receptor (Bmpr2) (Beppu et al., 2000), and Smads (Smad1, Smad5, and Smad4) (Yang et al., 1998; Chang et al., 1999; Tremblay et al., 2001) precludes analysis of BMP signaling during much of germ cell development. Additionally, redundancy amongst BMP family members and receptors complicate genetic analysis in this system. To bypass these complications, we used an organ culture system to study the role of BMP signaling in germ cell migration. After formation, PGCs proliferate and migrate via the gut to colonize the developing gonads (Molyneaux et al., 2001). Migratory PGCs express BMP receptors and Smads (Molyneaux et al., 2004) and may be exposed to BMP signals emanating from the lateral mesoderm or the extraembryonic mesoderm where Bmp2, Bmp4, Bmp5, and Bmp7 are expressed (Winnier et al., 1995; Solloway and Robertson, 1999; Fujiwara et al., 2002). BMP2 and BMP4 were shown to simulate the migration of osteoblast progenitors (Fiedler et al., 2002) and it is possible that BMPs might play a role in germ cell guidance. To test this, BMPs or the BMP-inhibitor Noggin were added to tissue dissected from E9.5 embryos. BMP treatment increased PGC numbers and Noggin treatment reduced PGC numbers and slowed PGC migration. Additionally, BMPs induced and Noggin repressed expression of Kitl and Id1. We propose that BMPs regulate the number and position of migratory PGCs by controlling expression of genes along the migratory route and within the urogenital ridges. 2. Results 2.1. PGCs and somatic cells express BMP receptors and Smads during germ cell migration BMPs signal through a complex containing type I and II serine/threonine kinase receptors. Upon binding BMPs,

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type II receptors phosphorylate and activate type I receptors, which in turn phosphorylate and activate Smads. Based on data from a screen of the Affymetrix MGU74av2 chip (Molyneaux et al., 2004), both migratory (E10.5) and postmigratory (E12.5) PGCs express mRNAs for Bmp4, Bmpr1a (Alk3), and Smad4. Smad1 (based on probe set 102984_g_at) was expressed in migratory and postmigratory PGCs; however an additional probe set for Smad1 (102983_at) gave weaker and less reproducible results. The other type I receptors (Acvr1/Alk2 and Bmpr1b/Alk6), the type II receptors (Bmpr2, Acvr2, and Acvr2b), Smad5 and the inhibitory smads (Smad6 and Smad7) were not reproducibly detected in PGCs using the Affymetrix chips. As a more sensitive method, we used quantitative RTPCR to examine the expression of Bmp4, BMP receptors, and Smads in PGCs and somatic tissue at E10.5 (Fig. 1). PGCs and the adjacent soma expressed Bmp4, Bmp receptors, and Smads with the majority of these transcripts being more highly expressed in the soma than in PGCs. The germ cell marker, Stella was absent from the somatic fraction (data not shown); whereas Cxcr4, a G-protein coupled receptor implicated in PGC guidance was enriched in germ cells, but present in the soma. The somatic marker Kitl was enriched in the somatic fraction. From this expression data it is likely that both PGCs and the somatic tissue are capable of responding to BMP-signals at this time point, but the majority of the BMP signaling components are more highly expressed in the soma.

Fig. 1. BMP signaling components are enriched in the soma at E10.5. PGCs and somatic cells were purified from E10.5 Oct4DPE:GFP embryos using flow cytometry. The somatic tissue in these sorts represents cells derived from the gut, gut mesentery, ventral body wall (including the urogenital ridges) and notocord. cDNA was prepared from PGCs (GFP+) and somatic (GFP ) samples and used for quantitative RT-PCR. For each marker, the somatic sample was used to generate a standard curve (expression set to 100%). Expression of each marker was normalized to the expression of Odc. Bmp4, Bmp receptors (Bmpr1A, Bmpr1B, Acvr1, Bmpr2, Acvr2, and Acvr2b), Smad1, Smad4, and Smad5 were enriched in the soma (compare expression levels with the somatic marker Kitl/Steel). Cxcr4, the receptor for the PGC guidance factor Sdf1 and Smad8 were enriched in the PGCs. Expression of Smad7 was not detected in either the somatic or PGC fractions.

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2.2. There is a BMP signaling center within the mesonephric mesenchyme at E9.5 and E10.5 Based on in situ hybridization results, Bmp4 is expressed within the urogenital ridge (Xu et al., 2003) and BMP4 signaling is thought to regulate the outgrowth of the ureteric bud (Miyazaki et al., 2000). Immunostaining using an antibody that cross-reacts with BMP2 and BMP4 revealed that BMP protein is enriched in the mesenchyme and epithelium of the urogenital ridges at E10.5 (Fig. 2A and B). To identify cells responding to BMPs, we stained slices with an antibody that recognizes the phosphorylated epitope of Smad1, Smad5, and Smad8 (pSmad1/5/8 Chang et al., 2002). At E9.5, cells in the epithelium of the urogenital ridge as well as the adjacent mesenchyme stained positive for p-Smad1/5/8 (Fig. 2C). By E10.5, p-Smad1/5/8 staining was no longer detected in the epithelium of the genital ridge. Instead, high levels of staining were observed in the condensing mesonephric mesenchyme (Fig. 2D). By E11.5, p-Smad1/5/8 staining was absent from the developing kidney, but weak staining was detected within gonadal PGCs (Fig. 2E). p-Smad1/5/ 8 staining was not detected in migratory PGCs (stages E9.5 and E10.5) (Fig. 2C and D), which appeared to accumulate in a region of the genital ridge devoid of pSmad1/5/8 staining (Fig. 2D). These data suggest that germ cells do not respond directly to BMP signals during migration, but the surrounding somatic cells do. In particular, somatic cells within the mesonephric mesenchyme exhibited robust staining for p-Smad1/5/8. 2.3. Perturbing BMP signaling affects PGC numbers in E9.5 slice culture At E9.5, PGCs emerge from the gut and actively migrate towards the genital ridges. Chemoattractants and survival factors released by the genital ridges are thought to control this process. To assess whether BMP signaling affects PGC migration recombinant human BMP4 or recombinant Noggin were added to slices dissected from E9.5 embryos (Fig. 3). PGC numbers were quantified by optically sectioning the tissue at the start point (T0) and end point (T18 hours) of the culture period (Fig. 3A and B), and expressing the number of germ cells after 18 h of culture as a percentage of the number at T0, in order to normalize results between slices containing different numbers of germ cells. After treatment, slices were fixed and stained for p-Smad1/5/8 to verify the suppression or activation of BMP signaling (Fig. 3C–H). One thousand nanograms per milliliter Noggin reduced PGC numbers in E9.5 slices (Fig. 3A). Control slices had little change in PGC numbers (average of 96.4 ± 11.4% s.e.m.) whereas slices treated with 1000 ng/ml Noggin had reduced PGC numbers (average of 66.7 ± 7.7% s.e.m.). This effect was statistically different than controls (ANOVA followed by Fisher’s LSD post test). High doses of Noggin (2000 ng/ml) appeared to increase PGC num-

bers (although this effect was not significantly different than controls). Treating E9.5 slices with BMP4 also resulted in a biphasic effect on PGC numbers (Fig. 3B). Low doses (0.5 and 5 ng/ml) increased PGC numbers, whereas higher doses (50 and 500 ng/ml) had no effect or actually reduced PGC numbers. In the BMP experiments, control slices exhibited an average change in PGC number of 73.1 ± 9.1% s.e.m. and the most effective dose of BMP4 (5 ng/ml) had an average change in PGC number of 120.0 ± 13.9% s.e.m. This effect was statistically different than controls (ANOVA followed by Fisher’s LSD post test). p-Smad1/5/8 staining confirmed the effects of the BMP agonists and antagonists in the slice cultures. Noggin treatment reduced p-Smad1/5/8 staining (Fig. 3C and D), and abolished the high level of response to BMPs in the mesonephros (compare Fig. 3D and F). Conversely, BMP4 treatment elevated p-Smad1/5/8 staining in every cell of the slice including PGCs (Fig. 3G and H). 2.4. Noggin treatment suppresses PGC motility In addition to affecting PGC numbers, Noggin treatment also affected germ cell position in slice cultures (Fig. 4A), whereas BMP4-treatment did not (data not shown). In control slices, the majority of PGCs moved away from the midline forming two distinct clusters at the genital ridges. PGC migration is inefficient and a few PGCs remained stranded in the midline region (arrows in Fig. 4A). However, in slices treated with 1000 ng/ml Noggin there was a dramatic accumulation of PGCs across the midline region of the slice (boxed areas in Fig. 4A). In the experiment shown, 75% (3/4) of the Noggin-treated slices were affected. Similar results were seen in two additional experiments (data not shown). Accumulation of PGCs in midline structures indicates a defect in PGC migration (Molyneaux et al., 2003). To determine if Noggin treatment affects the direction and/ or speed of PGC migration, we performed time-lapse analysis of PGC behavior in control and Noggin-treated slices (Fig. 4B and C). Four control slices and four Noggin-treated slices were filmed and cell trajectories and velocity measurements were calculated (see Section 4). Fig. 4B shows start- and end-point pictures of a representative control and Noggin-treated slice. In the control slice, PGC trajectories were long and straight. In the Noggin-treated slice, PGC trajectories were shorter and twisted. PGCs starting in the midline were more severely affected than PGCs starting closer to the genital ridges. Six cells were traced per movie to yield average velocity, average maximum velocity, and average displacement measurements for each slice (Fig. 4C). PGCs in control slices had an average velocity of 21.3 ± 3 s.d. lm/h, an average maximum velocity of 46.3 ± 10.4 s.d. lm/h and an average displacement of 111.8 ± 27.4 s.d. lm. Noggin treatment reduced the average velocity, maximum veloci-

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Fig. 2. Cells of the mesonephric mesenchyme respond to endogenous BMPs. Tissue dissected from E10.5 Oct4DPE:GFP embryos was fixed and stained with naı¨ve goat IgG (A) or anti-BMP2/4 (B). Immune complexes were detected with donkey anti-goat Cy5 and visualized by confocal microscopy. Diffuse staining for BMP2/4 was present throughout the slices, but highly concentrated in both the epithelial and mesenchymal components of the mesonephros. a (aorta) and gr (genital ridge). Slices were dissected from E9.5 (C), E10.5 (D) and E11.5 (E) Oct4DPE:GFP embryos and stained with anti-p-Smad1/5/8. At E9.5, both epithelial and mesenchymal components of the genital ridge exhibited elevated p-Smad1/5/8. By E10.5, p-Smad1/5/8 staining was highly enriched within the condensing mesonephric mesenchyme (mm), but reduced in the ventral portion of the genital ridge (where PGCs accumulate). By E11.5, p-Smad1/5/8 staining was absent in the mesonephros, but was weakly detected in the PGCs. g (gonad). Scale bars are 40 lm (A) and (B) 30 lm (C) and (D) and 70 lm (E).

ty, and displacement of PGCs. PGCs in Noggin-treated slices had an average velocity of 15.9 ± 4.8 s.d. lm/h, which was statistically different than controls (Student’s t-test p = 0.00009). The average maximum velocity of PGCs in Noggin-treated slices was 38.7 ± 15.2 s.d. lm/h

(t-test p = 0.051). Displacement in Noggin-treated slices was severely reduced with the average displacement being 66.5 ± 32.6 s.d. lm (t-test p = 0.000004). We conclude that Noggin treatment randomizes and slows PGC migration.

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Fig. 3. BMP signaling regulates PGC numbers. The percent change in PGC numbers was determined in slices treated for 18 h with the indicated concentrations of Noggin (A) or BMP4 (B). Moderate doses of Noggin reduced PGC numbers, whereas moderate doses of BMP4 increased PGC numbers in slices dissected from E9.5 Oct4DPE:GFP embryos. n indicates the number of slices in each treatment group. Error bars are s.e.m. Data were analyzed by ANOVA followed by the Fisher’s least significant difference test to determine significant changes. * Samples differing from controls (post test p < 0.05). ** Samples differing from controls (post test p < 0.01). Slices treated with 1000 ng/ml Noggin (C) and (D), control slices (E) and (F) or slices treated with 10 ng/ml BMP4 (F) and (G) were fixed and stained with anti-p-smad1/5/8. Noggin treatment reduced p-Smad1/5/8 staining. BMP4 treatment elevated p-Smad1/5/8 staining over the entire slice (including in PGCs). In control slices, p-Smad1/5/8 staining was elevated in the mesonephric mesenchyme (mm), but was not observed in PGCs. gr (genital ridge). md (mesonephric duct). Scale bar for (C, E, and G) is 115 lm. Scale bar for (D, F, and H) is 52 lm. The large arrow points towards the gut and indicates the ventral axis of the slice.

2.5. Exogenous BMP4 and BMP5 elevate expression of Kitl and Id1 in organ culture Based on the p-Smad staining and organ culture data, BMP4 is unlikely to act as a chemoattractant for PGCs however BMP signaling to the soma (e.g., the mesonephric mesenchyme) may control expression of chemoattractants and survival factors that regulate PGC behavior. To test this, the expression of potential BMP-target genes was examined in organ culture using real time RT-PCR. In initial experiments, results were normalized using Ornithine Decarboxylase (Odc) as a loading control; however, Odc levels were found to drop in culture (relative to input RNA) (data not shown). Additionally, Odc levels were found to decline in vivo between E9.5 and E10.5 (data not shown). This makes Odc an unsuitable loading control for establishing a time course of gene expression at these stages. Instead, RT-PCR results were normalized using

the expression level of TATA-binding protein and b-actin, two housekeeping genes whose expression levels remain steady between E9.5 and E10.5 (data not shown). Both BMP4 and BMP5 induced expression of Kitl (Fig. 5) in organ culture. Kitl is a known survival factor for PGCs. Kitl expression was induced after 3 h in culture, but peaked between 6 and 18 h. This relatively slow induction suggested to us that BMP-regulation of Kitl might go through an intermediate step. Several transcription factors are expressed in the early genital ridge and have known roles in kidney development and/or development of the somatic components of the gonad. Quantitative RT-PCR was used to examine the effect of exogenous BMPs on transcription factors expressed within the urogenital ridge (Pax2, SF1, Lhx1, and Id1). Both BMP4 and BMP5 treatment induced rapid induction of Id1 (Fig. 5), a helix-loop helix factor and known BMP target (Valdimarsdottir et al., 2002). BMP treatment slightly reduced expression

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Fig. 4. Noggin treatment slows and randomizes PGC movements. Slices were dissected from E9.5 Oct4DPE:GFP embryos and cultured for 18 h. In control slices (A), the majority of PGCs cleared the midline and formed clusters at the genital ridges. A few PGCs remained near the midline (arrows). In Noggin-treated (1000 ng/ml) slices a large number of PGCs remained scattered across the midline (boxed areas). This effect was observed in 9 out of 12 slices from three experiments. g (gut). Four control and four Noggin-treated slices were filmed for 144 min (7.4 h) and PGC movements were traced (6 cells per movie). The start point and endpoint of a representative control slice and a representative Noggin-treated slice are shown in (B). Germ cell trajectories are overlaid in red. The average velocity, average maximum velocity and average displacement of the six traced cells is shown in (C). Movies are available as supplemental material. Each bar represents an individual slice. Error bars are standard deviation in order to display the fact that PGCs, even in control slices, exhibited a wide range of behaviors. Noggin treatment produced a statistically significant decrease in avg. velocity (t-test p < 0.001) and displacement (t-test p < 0.001). Again, note that three out of four slices were affected.

of Pax2, SF1, and Lhx1 (data not shown). Id1 was identified in an in situ screen for factors expressed within the urogenital ridge (Wertz and Herrmann, 2000). In addition, Id1 is implicated in controlling cell migration (Valdimarsdottir et al., 2002) and pluripotency (Ying et al., 2003). 2.6. Noggin treatment reduces Id1 and Kitl expression within the urogenital ridge BMP signaling is necessary for PGC formation and early patterning of the embryo. To examine the role of BMP signaling in PGC migration in vivo requires a genetic system with very fine control over the timing at which the BMP signals are ablated in order to bypass this early requirement and yet efficiently and specifically inactivate

all BMP signaling by E9.5. Instead, we have again taken an organ culture approach to address whether endogenous BMP signals regulate gene expression within the urogenital ridges. Slices were treated with Noggin for 18 h. Following treatment, the UGRs were dissected and expression levels of Kitl and Id1 were quantified in cDNA prepared from UGR and non-UGR tissue samples (see Fig. 6). Kitl expression increased an average of 3.8· in the UGRs during culture (Fig. 6C). Kitl expression also increased within the UGRs in vivo (6· comparing E10.5 genital ridges to E9.5 genital ridges). This increase was specific to the genital ridges. Adding Noggin to slices inhibited the increase in Kitl within the UGRs. Id1 expression increased in culture within both the genital ridges and in the rest of the slice albeit this increase (an average of 1.5·) was less dramatic

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Fig. 5. BMP-treatment induces expression of Kitl and Id1. E9.5 slices were treated with the indicated concentrations of BMP4 or BMP5. Slices were harvested at 0, 3, 6 or 18 h after treatment and the expression of Kitl and Id1 were compared using quantitative RT-PCR. The T0 samples were used to generate the standard curve (expression set to 100). A normalization factor was generated by averaging the expression of TATA-binding protein and bACT. Data is an average of five experiments using BMP4 and five experiments using BMP5. Data were analyzed using ANOVA followed by Fisher’s least significant difference test. * Samples that differ from T0 values (100) (p < 0.1). ** Samples that differ from T0 values (p < 0.01). Error bars are s.e.m.

Fig. 6. Noggin treatment represses Kitl and Id1 expression in the genital ridges. E9.5 slices (four slices per treatment group) were treated with 100 ng/ml (0.1 N), 1000 ng/ml (1 N), or 2000 ng/ml (2 N) of Noggin. After 18 h in culture the genital ridges were dissected away from the rest of the body wall and assayed for expression of Kitl and Id1 using real time PCR. (A) Schematic of how the dissections are performed. (B) A confocal image of a slice showing dissection of the genital ridge on one side. (C) Real time PCR expression levels for Kitl. (D) Real time PCR expression levels for Id1. Data is an average of three experiments except for the E10.5 data which represents a single comparison. In each experiment the T0 genital ridge sample was used to generate a standard curve (expression level set to 100). A normalization factor was generated by averaging the expression of TATA-binding protein and bACT. Data were analyzed using ANOVA followed by Fisher’s least significant difference test. * Samples that differ from T0 genital ridge values (100) (post test p < 0.1). ** Samples that differ from T0 genital ridge values (post test p < 0.01). Error bars are s.e.m.

than the increase in Kitl expression (Fig. 6D). Noggin treatment suppressed this induction and reduced Id1 levels to 0.4–0.6· starting levels. 3. Discussion A founding population of approximately 50 PGCs forms at the posterior end of the mouse embryo at E7.5

(Lawson et al., 1999). From there PGCs migrate and proliferate until 1000 cells colonize the genital ridges by E10.5. Chemoattractants, repellents, adhesive gradients, and/or localized survival factors have all been proposed to regulate PGC migration (reviewed in Molyneaux and Wylie, 2004), but molecular evidence for these processes is scant. In a screen to identify transcripts expressed in migratory PGCs, we found that PGCs express BMP

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receptors and hence might be able to respond to endogenous BMPs expressed in the lateral plate and developing urogenital ridges. BMP signaling is required for PGC formation and mice lacking Bmp4 (Lawson et al., 1999), Bmp2 (Ying and Zhao, 2001), Bmp8b (Ying et al., 2000), Smad1 (Tremblay et al., 2001) or Smad5 (Chang and Matzuk, 2001) have a reduced founding population of PGCs as well as defects in mesoderm formation and development of the allantois. In order to address the role of BMPs in germ cell migration, we used an organ culture system to bypass the early role of BMPs in PGC formation and embryonic patterning. Quantitative RT-PCR revealed that migratory (E10.5) PGCs and the surrounding somatic tissue express Bmp4, Bmp receptors, and Smads and that these transcripts were more abundant in the soma than in PGCs. At E9.5, pSmad staining was detected in somatic cells along the migratory route and by E10.5 intense p-Smad staining was observed within the condensing mesonephric mesenchyme within the urogenital ridges. Surprisingly, PGCs did not exhibit p-Smad staining until after colonizing the UGRs (E11.5). A similar expression pattern was observed by Chuva de Sousa Lopes et al. (2005). This suggests that migratory PGCs are not responding to BMP signals or do so using a non-Smad pathway such as ERK or p38 MAPK. Takeuchi et al. (2005) have shown that 15% of migrating PGCs exhibit pp-ERK, but this signaling is likely due to FGFs and not BMPs. Migratory PGCs did not stain for phospho-p38 MAPK (data not shown). Although migratory PGCs did not appear to be responding to endogenous BMPs, perturbing BMP signaling affected both PGC numbers and motility in an organ culture assay. Addition of 5 ng/ml of BMP4 increased PGC numbers whereas high doses decreased PGC numbers. Noggin treatment had the converse effect, 1000 ng/ ml of Noggin decreased PGC numbers whereas high doses slightly increased PGC numbers. One thousand nanograms per milliliter Noggin treatment also reduced the speed of and partially randomized PGC movements causing a large number of germ cells to remain on the midline of cultured slices. BMP4-treatment did not have a significant effect on the direction or speed of germ cell migration (data not shown). This suggests that BMPs do not act directly as chemoattractants for PGCs. If migratory PGCs are not responding to BMPs, why should Noggin treatment affect germ cell numbers and motility? Addition of BMP4 and BMP5 induced expression of Kitl and Id1 in cultured slices. Conversely, Noggin treatment inhibited Id1 expression in slices and blocked expression of Kitl specifically within the urogenital ridges. Kitl and its receptor c-Kit control PGC survival and proliferation during migration (reviewed in Wylie, 1999). Strong mutations in both the receptor and ligand result in a complete loss of PGCs early in development (Besmer et al., 1993). Additionally, changing expression patterns of Kitl may control germ cell motility by either attractive (via the secreted form of Kitl) or adhesive (via the membrane

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bound form of Kitl) mechanisms. Godin et al. (1991) have shown that soluble Kitl does not attract PGCs in culture, but certain mutations in Kitl cause mislocalization of PGCs within the gut (Buehr et al., 1993) in addition to affecting PGC numbers. Because of the role of Kitl in PGC survival and proliferation it has been difficult to draw conclusions about whether this protein has a specific effect on PGC motility. However recently, Mahakali Zama et al. (2005) used an allelic series of mutations in the Kitl gene to demonstrate that the role of this protein in PGC survival/ proliferation may be separable from its role in migration. Our data suggest a model whereby BMP signaling regulates PGC behavior by controlling expression of Kitl within the somatic tissue along the migration route. BMPs are required at multiple stages of germ cell development and in many cases mediate their effects at least in part by control of the soma. de Sousa Lopes et al. (2004) have shown that BMP signals from the extraembryonic ectoderm regulate PGC formation by controlling the release of secondary signals from the visceral endoderm. Recently, Kirilly et al. (2005) showed that BMPs in the fly ovary not only signal directly to the developing oocyte, but are also required to control proliferation of the somatic cells within the germ cell niche. Likewise in the mammalian ovary, BMP15 and GDF9 released from the oocyte control proliferation of granulosa cells (Otsuka et al., 2000). Intriguingly, BMP15 and GDF9 have opposing effects on Kitl expression. BMP15 was shown to induce Kitl expression within granulosa cells (Otsuka and Shimasaki, 2002) whereas GDF9 was shown to inhibit Kitl expression both in vivo (Elvin et al., 1999) and in ovary organ culture (Joyce et al., 2000). We have shown that both BMP4 and BMP5 can induce Kitl expression in an organ culture system. It will be intriguing to see if there are additional BMP family members (perhaps GDFs) expressed within the trunk of the E9.5 embryos that are capable of downregulating Kitl expression. If this is the case, it may explain one of the puzzling aspects of our organ culture data. We have found that both Noggin and BMP4 have bi-phasic effects on germ cell numbers. Moderate doses of BMP4 were found to increase PGC numbers whereas high doses actually reduced numbers. Noggin treatment had the converse effect. Alternatively, it may be possible that different levels of BMP signaling may have different effects on PGC behavior. For example, it has been shown that low doses of BMP4 can induce hematopoietic stem cell precursors to differentiate whereas high doses promote self renewal (Bhatia et al., 1999). In summary, cells along the migration route and within the developing urogenital ridges respond to BMPs; however signaling was not detected in migratory PGCs. Germ cells exhibited p-SMAD staining after colonizing the gonads. We propose that BMP signaling regulates PGC numbers and may control PGC position by controlling expression of genes along the migration route. Tissue specific targeting will be required to definitively address the role of BMPs in PGC migration in vivo. However, the large

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number of BMP family members and receptors makes this a challenging task. It may be necessary to target multiple BMP receptors and/or Smads to reveal the impact of BMP signals on PGCs at this stage.

p-Smad1/5/8 antibody. We thank Greg Matera and Joeseph Nadeau for critical reading of the manuscript and we acknowledge Patti Conrad for microscopy assistance. Appendix A. Supplementary data

4. Experimental procedures 4.1. Organ culture All animal procedures have been approved by the Case Western Institutional Animal Care and use Committee. Oct4DPE:GFP +/+ homozygous males (Anderson et al., 1999) established on the FVB background were crossed to CD1 females (Charles River Laboratories). Embryonic day 0.5 (E0.5) was assumed to be noon on the day that a vaginal plug was detected. E9.5 embryos were harvested and slices were cut and cultured in millicell Organ culture chambers pre-coated with collagen IV as previously described (Molyneaux et al., 2003). Recombinant human BMP4 (R&D systems) or mouse Noggin-FC (R&D systems) were added to the medium at the indicated concentrations. Germ cells were counted at T = 0 h and T = 18 h by optically sectioning the slices as described (Molyneaux et al., 2003). Slices were filmed using the LSM510 confocal system and germ cell movements were quantitated as described (Molyneaux et al., 2003). Briefly, one frame was captured every 7 min for 7.5 h (63 frames). Individual germ cells (six cells per movie) were manually traced using NIH Image (http://rsb.info.nih.gov/nih-image/Default.html). The position of a germ cell was determined every 35 min (5 frames) generating a trace and a velocity measurement. Velocity measurements (12 per cell) were averaged to yield the average velocity of that cell (Avg. V.). At the end of the trace, the maximum velocity of the cell was recorded (Max. V) (fastest of the 12 measurements) and the absolute distance that the cell moved from its starting point was recorded (displacement, D). For an individual slice, data from the six germ cells that were tracked were averaged to yield the Avg. V., Avg. Max. V. and Avg. D. for that slice.

4.2. Immunostaining E9.5, E10.5, and E11.5 embryos were dissected from the uterus in PBS/ 2% FCS. The heads were removed and the bodies were fixed overnight in 4% PFA/PBS. Embryos were washed in PBS/0.1% TX-100 and a scalpel was used to cut transverse slices from the trunk of the fixed embryos. Slices (200 lm thick) were permeabilized at 4 C o/n in PBS/0.1% TX-100, and then blocked overnight in 2% donkey serum/PBS (for donkey secondary antibodies) or 2% goat serum/PBS (for goat secondary antibodies). Slices were incubated (overnight at 4 C) with primary antibodies used at a 1:100 dilution in blocking buffer. Rabbit polyclonal antibodies against p-smad1/5/8 were generously supplied by Peter ten Dijke. Goat polyclonal antibodies against BMP2/4 (sc-6267) were purchased from Santa Cruz BioTech. Slices were washed 5· 1 h in PBS/0.1% TX-100 and incubated (overnight at 4 C) with 1:100 Cy5-conjugated Goat anti-rabbit (Jackson Immuno Research) or 1:100 Cy5-conjugated Donkey anti-goat secondary antibodies (Jackson Immuno Research) diluted in blocking buffer.

4.3. Semi-quantitative RT-PCR Germ cells and somatic cells were isolated by flow cytometry as previously described (Molyneaux et al., 2004). cDNAs were prepared from 24 ng of mRNA using the Superscript III kit (Invitrogen). Semi-quantitative RTPCR was performed as described (Molyneaux et al., 2004). Primers were designed using Primer 3 (http://frodo.wi.mit.edu/cgi-bin/primer3/ primer3_www.cgi) and are shown in Table S1 of the supplemental material.

Acknowledgements Funding support was provided by the March of Dimes (#5-FY05-1233). We thank Peter ten Dijke for the

Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.mod.2006. 09.005. References Anderson, R., Fassler, R., Georges-Labouesse, E., Hynes, R.O., Bader, B.L., Kreidberg, J.A., Schaible, K., Heasman, J., Wylie, C., 1999. Mouse primordial germ cells lacking beta1 integrins enter the germline but fail to migrate normally to the gonads. Development 126, 1655– 1664. Beppu, H., Kawabata, M., Hamamoto, T., Chytil, A., Minowa, O., Noda, T., Miyazono, K., 2000. BMP type II receptor is required for gastrulation and early development of mouse embryos. Dev. Biol. 221, 249–258. Besmer, P., Manova, K., Duttlinger, R., Huang, E.J., Packer, A., Gyssler, C., Bachvarova, R.F., 1993. The kit-ligand (steel factor) and its receptor c-kit/W: pleiotropic roles in gametogenesis and melanogenesis. Dev. Suppl., 125–137. Bhatia, M., Bonnet, D., Wu, D., Murdoch, B., Wrana, J., Gallacher, L., Dick, J.E., 1999. Bone morphogenetic proteins regulate the developmental program of human hematopoietic stem cells. J. Exp. Med. 189, 1139–1148. Buehr, M., McLaren, A., Bartley, A., Darling, S., 1993. Proliferation and migration of primordial germ cells in We/We mouse embryos. Dev. Dyn. 198, 182–189. Chang, H., Matzuk, M.M., 2001. Smad5 is required for mouse primordial germ cell development. Mech. Dev. 104, 61–67. Chang, H., Huylebroeck, D., Verschueren, K., Guo, Q., Matzuk, M.M., Zwijsen, A., 1999. Smad5 knockout mice die at mid-gestation due to multiple embryonic and extraembryonic defects. Development 126, 1631–1642. Chang, W., ten Dijke, P., Wu, D.K., 2002. BMP pathways are involved in otic capsule formation and epithelial–mesenchymal signaling in the developing chicken inner ear. Dev. Biol. 251, 380–394. Chuva de Sousa Lopes, S.M., van den Driesche, S., Carvalho, R.L., Larsson, J., Eggen, B., Surani, M.A., Mummery, C.L., 2005. Altered primordial germ cell migration in the absence of transforming growth factor beta signaling via ALK5. Dev. Biol. 284, 194–203. de Sousa Lopes, S.M., Roelen, B.A., Monteiro, R.M., Emmens, R., Lin, H.Y., Li, E., Lawson, K.A., Mummery, C.L., 2004. BMP signaling mediated by ALK2 in the visceral endoderm is necessary for the generation of primordial germ cells in the mouse embryo. Genes Dev. 18, 1838–1849. Dong, J., Albertini, D.F., Nishimori, K., Kumar, T.R., Lu, N., Matzuk, M.M., 1996. Growth differentiation factor-9 is required during early ovarian folliculogenesis. Nature 383, 531–535. Dudley, A.T., Lyons, K.M., Robertson, E.J., 1995. A requirement for bone morphogenetic protein-7 during development of the mammalian kidney and eye. Genes Dev. 9, 2795–2807. Elvin, J.A., Yan, C., Wang, P., Nishimori, K., Matzuk, M.M., 1999. Molecular characterization of the follicle defects in the growth differentiation factor 9-deficient ovary. Mol. Endocrinol. 13, 1018–1034. Fiedler, J., Roderer, G., Gunther, K.P., Brenner, R.E., 2002. BMP-2, BMP-4, and PDGF-bb stimulate chemotactic migration of primary human mesenchymal progenitor cells. J. Cell. Biochem. 87, 305–312. Fujiwara, T., Dehart, D.B., Sulik, K.K., Hogan, B.L., 2002. Distinct requirements for extra-embryonic and embryonic bone morphogenetic protein 4 in the formation of the node and primitive streak and

B.M. Dudley et al. / Mechanisms of Development 124 (2007) 68–77 coordination of left–right asymmetry in the mouse. Development 129, 4685–4696. Godin, I., Deed, R., Cooke, J., Zsebo, K., Dexter, M., Wylie, C.C., 1991. Effects of the steel gene product on mouse primordial germ cells in culture. Nature 352, 807–809. Gu, Z., Reynolds, E.M., Song, J., Lei, H., Feijen, A., Yu, L., He, W., MacLaughlin, D.T., van den Eijnden-van Raaij, J., Donahoe, P.K., Li, E., 1999. The type I serine/threonine kinase receptor ActRIA (ALK2) is required for gastrulation of the mouse embryo. Development 126, 2551– 2561. Hu, J., Chen, Y.X., Wang, D., Qi, X., Li, T.G., Hao, J., Mishina, Y., Garbers, D.L., Zhao, G.Q., 2004. Developmental expression and function of Bmp4 in spermatogenesis and in maintaining epididymal integrity. Dev. Biol. 276, 158–171. Joyce, I.M., Clark, A.T., Pendola, F.L., Eppig, J.J., 2000. Comparison of recombinant growth differentiation factor-9 and oocyte regulation of KIT ligand messenger ribonucleic acid expression in mouse ovarian follicles. Biol. Reprod. 63, 1669–1675. Kawase, E., Wong, M.D., Ding, B.C., Xie, T., 2004. Gbb/Bmp signaling is essential for maintaining germline stem cells and for repressing bam transcription in the Drosophila testis. Development 131, 1365–1375. Kingsley, D.M., Bland, A.E., Grubber, J.M., Marker, P.C., Russell, L.B., Copeland, N.G., Jenkins, N.A., 1992. The mouse short ear skeletal morphogenesis locus is associated with defects in a bone morphogenetic member of the TGF beta superfamily. Cell 71, 399–410. Kirilly, D., Spana, E.P., Perrimon, N., Padgett, R.W., Xie, T., 2005. BMP signaling is required for controlling somatic stem cell self-renewal in the Drosophila ovary. Dev. Cell 9, 651–662. Lawson, K.A., Dunn, N.R., Roelen, B.A., Zeinstra, L.M., Davis, A.M., Wright, C.V., Korving, J.P., Hogan, B.L., 1999. Bmp4 is required for the generation of primordial germ cells in the mouse embryo. Genes Dev. 13, 424–436. Luo, G., Hofmann, C., Bronckers, A.L., Sohocki, M., Bradley, A., Karsenty, G., 1995. BMP-7 is an inducer of nephrogenesis, and is also required for eye development and skeletal patterning. Genes Dev. 9, 2808–2820. Mahakali Zama, A., Hudson 3rd, F.P., Bedell, M.A., 2005. Analysis of hypomorphic KitlSl mutants suggests different requirements for KITL in proliferation and migration of mouse primordial germ cells. Biol. Reprod. 73, 639–647. Mishina, Y., Suzuki, A., Ueno, N., Behringer, R.R., 1995. Bmpr encodes a type I bone morphogenetic protein receptor that is essential for gastrulation during mouse embryogenesis. Genes Dev. 9, 3027–3037. Mishina, Y., Crombie, R., Bradley, A., Behringer, R.R., 1999. Multiple roles for activin-like kinase-2 signaling during mouse embryogenesis. Dev. Biol. 213, 314–326. Miyazaki, Y., Oshima, K., Fogo, A., Hogan, B.L., Ichikawa, I., 2000. Bone morphogenetic protein 4 regulates the budding site and elongation of the mouse ureter. J. Clin. Invest. 105, 863–873. Molyneaux, K., Wylie, C., 2004. Primordial germ cell migration. Int. J. Dev. Biol. 48, 537–544. Molyneaux, K.A., Stallock, J., Schaible, K., Wylie, C., 2001. Time-lapse analysis of living mouse germ cell migration. Dev. Biol. 240, 488–498. Molyneaux, K.A., Zinszner, H., Kunwar, P.S., Schaible, K., Stebler, J., Sunshine, M.J., O’Brien, W., Raz, E., Littman, D., Wylie, C., Lehmann, R., 2003. The chemokine SDF1/CXCL12 and its receptor CXCR4 regulate mouse germ cell migration and survival. Development 130, 4279–4286. Molyneaux, K.A., Wang, Y., Schaible, K., Wylie, C., 2004. Transcriptional profiling identifies genes differentially expressed during and after migration in murine primordial germ cells. Gene Expr. Patterns 4, 167–181. Otsuka, F., Shimasaki, S., 2002. A negative feedback system between oocyte bone morphogenetic protein 15 and granulosa cell kit ligand: its role in regulating granulosa cell mitosis. Proc. Natl. Acad. Sci. USA 99, 8060–8085.

77

Otsuka, F., Yao, Z., Lee, T., Yamamoto, S., Erickson, G.F., Shimasaki, S., 2000. Bone morphogenetic protein-15. Identification of target cells and biological functions. J. Biol. Chem. 275, 39523–39528. Shimasaki, S., Moore, R.K., Otsuka, F., Erickson, G.F., 2004. The bone morphogenetic protein system in mammalian reproduction. Endocr. Rev. 25, 72–101. Sirard, C., de la Pompa, J.L., Elia, A., Itie, A., Mirtsos, C., Cheung, A., Hahn, S., Wakeham, A., Schwartz, L., Kern, S.E., Rossant, J., Mak, T.W., 1998. The tumor suppressor gene Smad4/Dpc4 is required for gastrulation and later for anterior development of the mouse embryo. Genes Dev. 12, 107–119. Solloway, M.J., Robertson, E.J., 1999. Early embryonic lethality in Bmp5;Bmp7 double mutant mice suggests functional redundancy within the 60A subgroup. Development 126, 1753–1768. Song, X., Wong, M.D., Kawase, E., Xi, R., Ding, B.C., McCarthy, J.J., Xie, T., 2004. Bmp signals from niche cells directly repress transcription of a differentiation-promoting gene, bag of marbles, in germline stem cells in the Drosophila ovary. Development 131, 1353–1364. Storm, E.E., Huynh, T.V., Copeland, N.G., Jenkins, N.A., Kingsley, D.M., Lee, S.J., 1994. Limb alterations in brachypodism mice due to mutations in a new member of the TGF beta-superfamily. Nature 368, 639–643. Takeuchi, Y., Molyneaux, K., Runyan, C., Schaible, K., Wylie, C., 2005. The roles of FGF signaling in germ cell migration in the mouse. Development 132, 5399–5409. ten Dijke, P., Korchynskyi, O., Valdimarsdottir, G., Goumans, M.J., 2003. Controlling cell fate by bone morphogenetic protein receptors. Mol. Cell. Endocrinol. 211, 105–113. Tremblay, K.D., Dunn, N.R., Robertson, E.J., 2001. Mouse embryos lacking Smad1 signals display defects in extra-embryonic tissues and germ cell formation. Development 128, 3609–3621. Valdimarsdottir, G., Goumans, M.J., Rosendahl, A., Brugman, M., Itoh, S., Lebrin, F., Sideras, P., ten Dijke, P., 2002. Stimulation of Id1 expression by bone morphogenetic protein is sufficient and necessary for bone morphogenetic protein-induced activation of endothelial cells. Circulation 106, 2263–2270. Wertz, K., Herrmann, B.G., 2000. Large-scale screen for genes involved in gonad development. Mech. Dev. 98, 51–70. Winnier, G., Blessing, M., Labosky, P.A., Hogan, B.L., 1995. Bone morphogenetic protein-4 is required for mesoderm formation and patterning in the mouse. Genes Dev. 9, 2105–2116. Wylie, C., 1999. Germ cells. Cell 96, 165–174. Xu, P.X., Zheng, W., Huang, L., Maire, P., Laclef, C., Silvius, D., 2003. Six1 is required for the early organogenesis of mammalian kidney. Development 130, 3085–3094. Yang, X., Li, C., Xu, X., Deng, C., 1998. The tumor suppressor SMAD4/ DPC4 is essential for epiblast proliferation and mesoderm induction in mice. Proc. Natl. Acad. Sci. USA 95, 3667–3672. Ying, Y., Zhao, G.Q., 2001. Cooperation of endoderm-derived BMP2 and extraembryonic ectoderm-derived BMP4 in primordial germ cell generation in the mouse. Dev. Biol. 232, 484–492. Ying, Y., Liu, X.M., Marble, A., Lawson, K.A., Zhao, G.Q., 2000. Requirement of Bmp8b for the generation of primordial germ cells in the mouse. Mol. Endocrinol. 14, 1053–1063. Ying, Q.L., Nichols, J., Chambers, I., Smith, A., 2003. BMP induction of Id proteins suppresses differentiation and sustains embryonic stem cell self-renewal in collaboration with STAT3. Cell 115, 281–292. Zhao, G.Q., 2003. Consequences of knocking out BMP signaling in the mouse. Genesis 35, 43–56. Zhao, G.Q., Deng, K., Labosky, P.A., Liaw, L., Hogan, B.L., 1996. The gene encoding bone morphogenetic protein 8B is required for the initiation and maintenance of spermatogenesis in the mouse. Genes Dev. 10, 1657–1669. Zhao, G.Q., Liaw, L., Hogan, B.L., 1998. Bone morphogenetic protein 8A plays a role in the maintenance of spermatogenesis and the integrity of the epididymis. Development 125, 1103–1112. Zwijsen, A., Verschueren, K., Huylebroeck, D., 2003. New intracellular components of bone morphogenetic protein/Smad signaling cascades. FEBS Lett. 546, 133–139.