Regulation of Fgf10 Gene Expression in the Prostate: Identification of ...

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Endocrinology 145(4):1988 –1995 Copyright © 2004 by The Endocrine Society doi: 10.1210/en.2003-0842

Regulation of Fgf10 Gene Expression in the Prostate: Identification of Transforming Growth Factor-␤1 and Promoter Elements DARREN C. TOMLINSON, JUSTIN C. GRINDLEY,

AND

AXEL A. THOMSON

Medical Research Council Human Reproductive Sciences Unit (D.C.T., A.A.T.), Centre for Reproductive Biology, The University of Edinburgh, Edinburgh EH16 4SB, Scotland, United Kingdom; and Division of Pediatric Cardiology (J.C.G.), Vanderbilt University School of Medicine, Nashville, Tennessee 37232 Fibroblast growth factor 10 (FGF10) is a mesenchymal paracrine-acting factor that plays a key role in the organogenesis of the prostate, and Fgf10 transcripts exhibit a highly restricted expression pattern within prostatic mesenchyme. To study the regulation of Fgf10 we have used organ rudiments grown in vitro as well as a primary stromal cell system derived from the ventral mesenchymal pad (VMP), a condensed area of mesenchyme known to induce prostatic organogenesis. Characterization of VMP cells (VMPCs) showed that they retained expression of AR as well as transcripts for FGF10 and TGF␤1, -2, and -3. We propose that VMPCs are a good model of specialized mesenchyme involved in prostatic organogenesis and are distinct from general urogenital sinus mesenchyme/ stroma. Treatment of VMPCs with TGF␤1 resulted in a rapid and transient decrease in Fgf10 transcript levels, which were

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EVELOPMENT OF THE prostate is dependent upon the presence of androgens and involves mesenchymal/epithelial interactions. Androgens bind to androgen receptor (AR) in the mesenchyme of the urogenital sinus, and this results in the induction and growth of prostatic buds from the urothelium of the urethra (UR) (reviewed in Ref. 1). These developing prostatic buds do not express AR at their inception. In rodents, a condensed and defined area of mesenchyme has been identified that appears to be involved in prostatic organogenesis and is termed the ventral mesenchymal pad (VMP) (2). In response to androgens, paracrine factors made by the mesenchyme and VMP have been proposed to stimulate prostatic development. A mesenchymal factor known to act as a paracrine regulator of epithelial growth and branching during prostate development, as well as other organs, is fibroblast growth factor 10 (FGF10) (3). FGF10 is essential for the formation of the prostate (4) and lungs and limbs (5, 6). FGF10 stimulates growth and branching of ventral prostates (VPs) cultured in vitro and increases the proliferation of prostatic epithelial cells (3, 7). Whether or Abbreviations: AR, Androgen receptor; Cphn, Cyclophilin; CPSR, control processed serum replacement; FCS, fetal calf serum; FGF, fibroblast growth factor; RPA, RNase protection assay; SM, smooth muscle; TIEG, Tgf␤-inducible early gene; UGS, urogenital sinus; UR, urethra; URSC, urethral stromal cell; UTR, untranslated region; VMP, ventral mesenchymal pad; VMPC, VMP cell; VP, ventral prostate. Endocrinology is published monthly by The Endocrine Society (http:// www.endo-society.org), the foremost professional society serving the endocrine community.

reduced 9-fold at 3 h. TGF␤1 also inhibited Fgf10 expression in VMP organ rudiments grown in vitro. To further analyze Fgf10 regulation, 6 kb of mouse genomic sequence 5ⴕ to the translation start site was characterized by promoter analysis. Deletion analysis of the Fgf10 promoter in VMPCs identified a region of the promoter that mediated a significant proportion of promoter activity as well as mediating promoter downregulation by TGF␤1. This element was located between nucleotides –182 and –172 and contained a consensus Sp1 binding site. Taken together, our data suggest that TGF␤1 is a regulator of Fgf10 expression in prostatic mesenchyme and that a proximal element within the Fgf10 promoter plays an important role in its regulation and expression. (Endocrinology 145: 1988 –1995, 2004)

not androgens control Fgf10 expression is unclear, as there is conflicting evidence from in vitro (7) and in vivo studies (3), reviewed in Thomson (8). Fgf10 is expressed in discrete areas of prostatic mesenchyme surrounding distal epithelial buds in the prostate and appears to be down-regulated in the central part of the organ. This indicates that expression of the Fgf10 gene in the prostate is spatially restricted (3), in common with other organs such as the lung (9). TGF␤1 is involved in mesenchymal-epithelial interactions during development of the prostate (10) and has been shown to inhibit prostate growth and branching (11). Thus, TGF␤1 and FGF10 appear to play opposing roles during the development of the prostate. We propose that part of the growth inhibition induced by TGF␤ may be due to down-regulation of Fgf10. In other systems such as 3T3 cells and lung mesenchyme, TGF␤ has been shown to down-regulate Fgf10 (12, 13). The current study has examined the regulation of Fgf10 gene expression in mesenchyme involved in regulating prostatic growth. We have established and characterized a primary cell system derived from the VMP that retains many important features relevant to prostatic growth. TGF␤1 repressed endogenous Fgf10 transcript levels in this primary cell system as well as in organ rudiments grown in vitro. Deletion analysis of the Fgf10 promoter identified a conserved element that mediated a significant proportion of promoter activity as well as mediating promoter down-regulation by TGF␤1. Hence TGF␤1 may play an opposing role to FGF10 during prostate development

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by repressing Fgf10 gene expression through an element in the Fgf10 promoter. Materials and Methods Isolation and culture of primary cells Cells used for primary culture were derived from VMPs or urethral stromal tissue, microdissected from P0 Wistar rat female urogenital sinuses (UGSs). Animals were maintained and killed in accordance with United Kingdom Home Office guidelines and legislation. Microdissected VMP or urethral stromal tissue was dissociated in the presence of 1 mg ml–1 collagenase (Invitrogen, Paisley, UK) for 45 min at 37 C. Dissociated VMP [primary VMP cells (VMPCs)] was cultured in DMEM with glutaMAX I supplemented with 10% fetal calf serum (FCS) (Invitrogen) and 1⫻ penicillin/streptomycin (Invitrogen). Cells were grown until 90% confluent and then passaged. Primary VMPCs were cultured in serum-free conditions for 24 h and then treated with or without TGF␤1 (5 ng ml–1) (Invitrogen); higher concentrations (such as 10 ng ml–1, used in organ studies) were found to result in excessive cell death. Total RNA was prepared from primary cells or organ rudiments as described (14). Primary VMPC area was measured from imaged cells using NIH Image software. Changes in cell number over different passage times, and in response to testosterone, were measured by counting cells using a hemocytometer and represented as either fold change or percentage change in cell number relative to control. For transfection, 8 ␮l GenePORTER was mixed with a standardized amount of promoter construct plasmid, 0.02 ␮g of pSV-␤-galactosidase control vector (Promega, Madison, WI) and 10 ␮l PBS and incubated for 15 min at room temperature. The final volume was made up to 1 ml in serum-free DMEM and added to primary cells in six-well culture dishes (2.5 ⫻ 10 cells per well were plated out the day before transfection). Cells were cultured in the transfection reagent for 6 h and then washed in DMEM supplemented with 10% control processed serum replacement (CPSR) (Sigma Aldrich Ltd., Poole, UK). Cells were grown for 24 h, treated appropriately, and harvested for luciferase and ␤-galactosidase assays using Promega lysis buffer, as per manufacturer’s instructions. Assays were repeated twice per cell extract. Luciferase activity was normalized to ␤-galactosidase activity.

Organ culture Serum-free organ culture of P0 Wistar rat VMPs were performed as previously described (15). Culture medium was supplemented with 10 ng ml–1 TGF␤1 where indicated.

Analysis of AR and smooth muscle (SM) ␣-actin protein levels Western blotting was performed as per manufacturer’s instructions using Invitrogen products. 1 ␮g and 20 ␮g of protein were used to visualize SM ␣-actin and AR, respectively. Antibodies for AR and SM ␣-actin were used at 1:100 and 1:1000 dilutions. Staining was visualized with goat antirabbit IgG (Sigma, Poole, UK) for AR and goat antimouse IgG (Sigma) for SM ␣-actin both diluted at 1:20,000 in Tris-buffered saline. Membranes were subjected to enhanced chemifluorescence (ECF) (Amersham, Buckinghamshire, UK) reagent, and staining was visualized using a Storm phosphoimager with a blue chemiluminescence filter (Molecular Dynamics, Sunnyvale, CA).

Analysis of transcript levels A DNA template for Fgf10 riboprobes was used as described (3). DNA templates for Tgf␤1, Tgf␤2 and Tgf␤3 riboprobes were synthesized by RT-PCR from P0 UGS cDNA and subcloned into pBluescript KSII⫹ (Stratagene, La Jolla, CA). PCR primers were designed using the following GenBank-published cDNA sequences: Tgf␤1 (GenBank accession no. X52498; primers: forward, CGTGCTAATGGTGGACCGCAACAAC, and reverse, AAGACAGCCACTCAGGCGTATCAGT), Tgf␤2 (GenBank accession no. AF153012; primers: forward, CTGCTGTACCTTCATACCGTCTA, and reverse, CAATAGGCGGCATCCAAA), and Tgf␤3 (GenBank accession no. NM_013174; primers: forward, AGTGGCTGTTGCGGAGAGAGTCC, and reverse, GCACACAGCAGTTCTC-

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CTCC). DNA templates for Cyclophilin (Cphn) were from Ambion, Inc. (Cambridgeshire, UK). 32P-labeled antisense riboprobes were transcribed, and RNase protection assay (RPA) was performed as previously described (3). RNase protected products for Fgf10, Tgf␤1, Tgf␤2, Tgf␤3, and Cphn were 323, 261, 245, 324, and 103 nucleotides, respectively. Gels were imaged using a Storm phosphoimager (Molecular Dynamics), and transcript abundance was determined from the intensity of bands, calculated using ImageQuant version1.2 (Molecular Dynamics). Transcript abundance was normalized to Cphn internal standards.

Cloning and sequencing of mouse Fgf10 genomic sequence A mouse Fgf10 cDNA clone (encompassing the complete coding region) (9) was used to screen a 129 mouse genomic library in Lambda Fix II (Stratagene catalog item 946308). The clone Fgf10 luc no. 28, containing 6.6 kb upstream of the Fgf10 ATG was cloned into the luciferase reporter vector pGL3 Basic (Promega). The Fgf10 ATG site was fused to the luciferase ATG by site-directed mutagenesis.

Cloning of deletion constructs Because pGL3 was found to have a high luciferase background in our transfection studies, the 5⬘ deletion promoter constructs were amplified by PCR from the Fgf10 luc no. 28 construct and then were cloned in pGEM-T Easy (Promega). Inserts were released by restriction digestion using BamHI [which cuts in the 5⬘ untranslated region (UTR)] and SpeI (vector restriction site) and cloned into BglII and NheI sites in pA3Lucm [a modified pA3Luc vector with additional polylinker restriction sites, originally designed to prevent cryptic plasmid transcription (16)]. Region U1 was incorporated in this construct as this region was thought to be part of the core promoter. For 3⬘ deletions, PCR products were ligated into pGEM-T Easy vector and inserts were released by restriction digestion using BglII and HindIII and ligated into pA3Lucm using the same enzymes. These were amplified by PCR using a forward primer incorporating a SpeI restriction site and a reverse primer incorporating a BamHI restriction site. The inserts were released by restriction digestion using SpeI and BamHI restriction enzymes and ligated into pA3Lucm (with start site sequence), which had been digested with NheI and BglII. Vectors were confirmed correct by restriction digest and sequencing. Numbering of the Fgf10 promoter constructs is based upon identification of the Fgf10 transcriptional start site by RNase protection (Tomlinson, D. C., and A. A. Thomson, unpublished data). The region containing the Sp1 site (–214 to ⫹502) was amplified by PCR and cloned into pGEM-T Easy vector. Inserts were released by restriction digest with SpeI and BamHI and cloned into pA3lucm digested with NheI and HindIII. ⌬U7-U1-SP1 was cloned in two steps by PCR. The region 3⬘ of the Sp1 site was amplified by PCR using a primer that incorporated a BglII restriction site and extra base pairs that replaced the Sp1 sequence (CCCCCGCCCCC replaced with GGAAGATCTTC) and cloned into pGEM-T Easy vector. Inserts were released by restriction digestion using BglII and HindIII and ligated into similar sites in pA3Lucm. The 5⬘ sequence to the Sp1 was site amplified by PCR and cloned into pA3lucm containing the 3⬘ sequence via pGEM-T Easy using SpeI and the BglII restriction sites. Hence, the final construct eliminated the Sp1 site and replaced it with a nonspecific sequence.

Results Characterization of primary VMPCs

Whereas other studies have used stromal cells or cell lines derived from the UGS (17, 18), we sought to use a defined subset of mesenchyme that appears to contain key components of prostatic regulatory activity. Thus it was important to characterize these cells and establish that they were derived from the VMP and had not arisen from contaminating UGS mesenchyme that might have overtaken our VMPC primary cultures. To establish the identity and characteristics of VMPCs, they were compared with urethral stromal cells (URSCs) that consisted of the remaining mesenchyme/ stroma after removal of the VMP. The cellular size and

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growth rate of VMPCs and URSCs were compared as well as expression of markers normally present in the organ rudiments from which VMPCs were derived (AR, SM ␣-actin, Tgf␤1, Tgf␤2, Tgf␤3, and Fgf10). The size and growth rate of primary VMPCs and URSCs at different passage numbers were measured to determine the passage number on which the cells were least stressed or senescent, as numerous reports have indicated that an increase in cell size and a decline in growth rate may be a result of stress or senescence (19). On the second passage, the growth rate of VMPCs rapidly decreased from a 10-fold to a 3-fold increase in cell number (Fig. 1A, gray bars), and the cell size increased (from 3604 to 8210 ␮m2) (Fig. 1B, filled circles). Hence it appeared that primary VMPCs were least stressed on passage one of culture and were used for experiments on the first passage. Comparison of the growth rate and cellular size of VMPCs with URSCs showed that VMPC growth slowed more rapidly than URSCs and that VMPCs were larger at early passage numbers (Fig. 1, A and B). These data suggest that VMPCs and URSCs are composed of different cell populations. The effect of testosterone on the growth rate of primary

Tomlinson et al. • Regulation of Fgf10 Expression in the Prostate

VMPCs was investigated to determine whether the cells were growth responsive to testosterone. Testosterone did not significantly affect the growth rate of primary VMPCs in the presence or absence of FCS (Fig 1C). Primary VMPCs expressed AR at all passage numbers (Fig. 1D), and immunohistochemistry (Fig. 1E) demonstrated that AR was expressed in almost all cells of the culture (⬎95%) similar to VMP in vivo. In VMPCs, AR showed a cytoplasmic and nuclear distribution in the absence of added androgens, although cultures included 10% FCS, consistent with AR distribution the VMP in vivo (20). Western blotting demonstrated a higher level of AR expression in the VMP rudiment than VMPCs, suggesting AR expression was somewhat reduced upon cellular disaggregation and growth in vitro. Next the expression of SM ␣-actin was examined. Cells within the VMP in vivo do not express SM actin but are located in close proximity to a layer of SM (20). Primary VMPCs expressed SM ␣-actin at all passages (Fig. 1D) as well as in almost all (⬎99%) cells in the culture (Fig. 1E). The microdissected VMP rudiment used in our studies appeared to contain SM actin by Western blot (Fig. 1D); this may have arisen from contaminating cells of SM layer included during

FIG. 1. Characterization of primary VMPCs: cell growth, cell size, and expression of markers. A, Comparison of growth rate of VMPCs with URSCs. Cell number was counted after passages at 4-d intervals to examine growth rate of VMPCs (gray bars) and URSCs (black bars). VMPC growth slowed more rapidly than URSC growth. B, Cell size of VMPCs and URSCs. Cellular area of VMPCs (F) and URSCs (f) was measured at different passage number. VMPC size increased more rapidly than URSCs. C, VMPCs were grown in the absence (F) or presence (f) of FCS with or without testosterone. Testosterone had no effect on primary VMPC growth in the presence or absence of FCS. D, Western blot for SM ␣-actin (42 kDa) and AR (119 kDa) on VMP organ rudiments and primary VMPCs. For SM ␣-actin blotting, 1 ␮g of protein was loaded, and 15 ␮g of protein was loaded per sample for AR protein blotting. The top image of each panel shows a Western blot and the bottom image of each panel shows a Coomassie stain of each blot to demonstrate consistent protein loading. Expression of SM ␣-actin was detected in the VMP dissected rudiment, and primary VMPCs expressed SM ␣-actin at similar levels at each passage. AR expression was highest in the VMP organ rudiment, and primary VMPCs expressed AR at each passage at similar levels (n ⫽ 3). E, Immunohistochemistry showing expression of SM actin (blue) and AR (green) in VMPCs; top, without primary antibodies. Nuclei are stained red with propidium iodide.

Tomlinson et al. • Regulation of Fgf10 Expression in the Prostate

microdissection. The expression of SM ␣-actin in VMPCs may have been due to contamination with, or overgrowth of, cells from the SM layer or due to the response of the VMPCs to conditions in cell culture. We believe the latter is more likely than the former. To ensure that primary VMPCs were derived from the VMP and not the SM layer, other markers of VMP tissue were examined. Tgf␤ transcript levels in the VMP, primary VMPCs, UR, and URSCs were compared to determine whether primary VMPCs were derived from the VMP and not contaminating urethral stroma or SM. Figure 2A shows quantification of Tgf␤ mRNA levels by RPA and phosphoimager analysis (numbers at the bottom of the panel are averages of two to four independent experiments). Figure 2A shows a comparison of TGF␤ mRNA levels between VMP and UR organ

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rudiments, and average transcript levels (⫾ sem) of VMP compared with UR were as follows: Tgf␤1, 128 ⫾ 13.6; Tgf␤2, 182 ⫾ 26.2; and Tgf␤3, 583 ⫾ 55.5. In vivo, VMP Tgf␤ transcript levels were approximately 1.3-fold (Tgf␤1), 1.8-fold (Tgf␤2), and 5.8-fold (Tgf␤3) higher than in the UR (Fig. 2A). In vitro, primary VMPC Tgf␤ transcript levels were approximately 1.5-fold (Tgf␤1), 2.8-fold (Tgf␤2) and 4.8-fold (Tgf␤3) higher than in primary URSCs (Fig. 2A). Average levels of Tgf␤ transcripts in VMPCs compared with URSCs were as follows: Tgf␤1, 162 ⫾ 22.9; Tgf␤2, 277 ⫾ 18.9; Tgf␤3, 478 ⫾ 13.8. The difference in Tgf␤ levels between VMPCs and URSCs was statistically significant when examined by t test: Tgf␤1, P ⫽ 0.01; Tgf␤2, P ⫽ 1.8e– 05; and Tgf␤3, P ⫽ 3.3e– 08. In addition, expression of the Tgf␤ receptor 1 in VMPCs was determined by RT-PCR (data not shown). The expression of Tgf␤ isoforms in organ rudiments was highly similar to those observed in the primary cells demonstrating that VMPCs retain VMP features and are different from URSCs and general mesenchyme. However, it remains possible that VMPCs are contaminated by a small proportion of URSCs included during microdissection. Fgf10 transcript levels in VP, VMP, and UR organ rudiments were compared with primary VMPCs at different passage numbers. The same P0 VP sample was included in each RPA so that different experiments could be compared. Fgf10 mRNA levels in the urethra were 31 ⫾ 8.0, and in the VMP 410 ⫾ 22.8, when compared with the VP (t test P ⫽ 0.0005 and P ⫽ 8.5e– 05, respectively, and P ⫽ 4.8e– 05 when compared with each other). Fgf10 transcript levels in the VMP were 14 times higher than those expressed in the UR and four times higher than levels in VPs (Fig. 2B). Fgf10 transcript levels in the VMP were four times higher than those expressed in primary VMPCs. The levels of Fgf10 transcripts in VMPCs showed little change over three passages: passage 1, 115 ⫾19.0; passage 2, 107 ⫾ 10.7; and passage 3, 92 ⫾ 11.4 (t test P ⫽ 0.23, P ⫽ 0.29, and P ⫽ 0.27, respectively; statistically, not significant differences). Using RPA, we have demonstrated the presence of Fgf10 transcripts in the UR that were not detected by in situ hybridization (3). This is most likely due to in situ hybridization not being sensitive enough to detect very low levels of transcripts and is not due to contaminating VMP, because Fgf10 transcript levels were regulated by TGF␤1 in the VMP but not in the UR (see below). Regulation of Fgf10 transcripts in VMPC and VMP by TGF␤

FIG. 2. Expression of Tgf␤ isoforms and Fgf10 transcript levels in organ rudiments and primary cells. Total cellular RNA (10 ␮g) from primary cells and organ rudiments was hybridized with 32P-labeled antisense riboprobes for Tgf␤1, Tg␤2, Tgf␤3, and Cphn, or Fgf10 and Cphn transcript levels by RPA. Transcript levels were normalized to Cphn. Numbers shown below autoradiographs show percent transcript abundance relative to UR and primary URSCs (A) or VP (B) and were calculated using a phosphoimager based upon an average of two to four independent experiments. A, The expression pattern of Tgf␤ isoform transcript levels was maintained from organ rudiment (UR or VMP; n ⫽ 2) to their respective primary cells (URSCs or VMPCs; n ⫽ 4). B, Fgf10 transcripts were expressed in the VP, UR, and VMP and were maintained in primary VMPCs at each passage (n ⫽ 3).

The regulation of Fgf10 transcript levels was investigated in primary VMPCs by treating them with TGF␤1 for different lengths of time in serum-free conditions (Fig. 3). Levels of Fgf10 mRNA in VMPCs compared with VP were as follows: with serum, 118 ⫾ 6.4; without serum, 171 ⫾ 33.2; 1-h Tgf␤ treatment, 138 ⫾ 30.9; 3-h Tgf␤ treatment, 20 ⫾ 12.4; 7-h Tgf␤ treatment, 39 ⫾ 20.0; 48-h Tgf␤ treatment, 139 ⫾ 40.8; and 48-h without-serum treatment, 127 ⫾ 27.3. Statistical analysis by t test showed a significant difference (P ⫽ 0.006) after 3 h of Tgf␤1 treatment. After a 1-h TGF␤1 treatment, Fgf10 transcript levels decreased slightly, but by 4 h, transcript levels had fallen by 9-fold. At 7 h, Fgf10 transcript levels were

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Tomlinson et al. • Regulation of Fgf10 Expression in the Prostate

FIG. 3. Effect of TGF␤1 on Fgf10 transcript levels in primary VMPCs. Transcript levels were measured by RPA and standardized to Cphn. Numbers shown below autoradiographs show percent transcript abundance as an average of three experiments, relative to P0 VP. Total cellular RNA(10 ␮g) was hybridized with 32P-labeled antisense riboprobes for Fgf10 and Cphn as an internal control. Primary VMPCs were cultured either in the presence of FCS, the absence of FCS (for 0 or 48 h), or in the absence of FCS and presence of TGF␤1 (5 ng ml–1) for 1, 3, 7, and 48 h. Primary VMPCs cultured in the presence of serum express similar levels of Fgf10 transcripts to the VP. Serum starvation of the cells resulted in approximately a 2-fold increase in Fgf10 transcript levels. TGF␤1 repressed Fgf10 transcript levels by 9-fold after 3 h and 4-fold after 7 h in culture. Fgf10 transcript levels returned to control values after 48 h (n ⫽ 3).

approximately 4-fold less than at the beginning of culture, but by 48 h, levels had recovered to those observed in the control. Also, Fgf10 transcript levels were increased when serum was removed from the media, suggesting that factors within the serum may be acting as inhibitors of Fgf10. Organ cultures of female UGTs were used to investigate whether the regulation of Fgf10 transcript levels in primary VMPCs by TGF␤1 could be extended to organ rudiments grown in vitro. P0 female UGTs were placed in culture and treated with TGF␤1 for 7 h, followed by dissection to separate the VMP and UR and comparison of Fgf10 mRNA levels in different parts of the UGT (Fig. 4A). A 3-fold decrease in Fgf10 transcript levels was observed in the VMP treated with TGF␤1 for 7 h (Fig. 4B). Levels of Fgf10 mRNA in VMP with TGF␤1 (compared with VMP without TGF␤1) were 33 ⫾ 10.3 (P ⫽ 0.0002 when examined by t test). No change was observed in Fgf10 transcript levels in the UR, although Fgf10 levels in the UR are considerably lower than in the VMP. Primary VMPCs showed a maximal decrease of Fgf10 transcript levels at 3 h, but the organ rudiments were treated with TGF␤1 protein for 7 h to allow penetration of the VMP tissue. A 3-h TGF␤1 treatment of VMP rudiments resulted in a 3-fold repression of Fgf10 transcript levels (n ⫽ 2, data not shown). These results demonstrate that TGF␤1 regulated Fgf10 gene expression in the VMP but did not regulate Fgf10 gene expression in the UR. This suggests that different mechanisms may control Fgf10 gene expression in cells located in different parts of the UGS. Characterization of the Fgf10 promoter

The characterization of the Fgf10 promoter was performed in primary VMPCs to identify a region of the 5⬘ genomic sequence (relative to the transcription start site) that might regulate Fgf10 expression in the prostate. Many regions of the Fgf10 promoter were highly conserved between mouse and human (marked as boxes U1–U7 in Fig. 5) (O’Rear, L. D., and J. C. Grindley, manuscript in preparation), and these conserved regions might contain regulatory elements. The re-

FIG. 4. Effect of TGF␤1 on Fgf10 transcript levels in VMP and UR rudiments in vitro. Female UGTs were cultured overnight in serumfree conditions, and then cultured for 7 h with TGF␤1. A, Regions of the UGT microdissected for analysis of Fgf10 transcript levels by RPA. RNA (5 ␮g) from microdissected VMP and UR rudiments was hybridized with 32P-labeled antisense riboprobes for Fgf10 and Cphn as an internal control. Transcript levels were measured by RPA and standardized to Cphn. Numbers shown below autoradiographs show percent transcript abundance as an average of three experiments, relative to untreated VMP. B, TGF␤1 repressed Fgf10 transcript levels by 3-fold in the VMP, but no regulation of Fgf10 transcript levels was detected in the UR (n ⫽ 3).

gions of homology are as follows: U7, –5492 to –5222; U6, – 4731 to – 4523; U5, –3590 –3140; U4, –3184 to –3140; U3, –2123 to –1915; U2, –1265 to – 667; and U1, –198 to –50. These regions were characterized by deleting them from the Fgf10 promoter in a luciferase expression vector, pA3Lucm. In primary VMPCs, 5⬘ to 3⬘ deletions up to the ⌬U7–U2 construct and 3⬘ to 5⬘ deletions showed no significant change in activity between constructs (when compared by t test). ⌬U7–U1 and ⌬U7–U1 ⌬5⬘UTR deletions resulted in a 2-fold

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FIG. 5. Fgf10 promoter analysis. Promoter constructs were cloned into pa3Lucm, a luciferase reporter vector, and used for transfection studies in VMPCs. Constructs containing the following regions of Fgf10 promoter sequence are shown on the left side of the figure: Full, 5.8 kb; ⌬U7, 5.2 kb; ⌬U7–U5, 3.1 kb; ⌬U7–U4, 2.4 kb; ⌬U7–U2, 307 bp; and ⌬U7– U1, 63 bp; ⌬U7–U1 ⌬5⬘UTR, 63 bp of DNA 5⬘ and 72 bp of DNA 3⬘ to the transcription start site. ⌬U2, ⌬U2–U5, ⌬U2–U6, and ⌬U2–U7 represent internal 5⬘ DNA deletions from ⫺63 of 2.1, 3.5, 4.7, and 5.4 kb, respectively. Promoter constructs were cotransfected with a ␤-gal-expressing vector into primary VMPCs and cultured in CPSR-containing media for 36 h. Results represent luciferase values normalized for transfection efficiency (by ␤-gal assay) and standardized to the Full promoter luciferase activity. The 5⬘ to 3⬘ deletions up to ⌬U7–U2 and internal deletions showed no significant change in activity between constructs. ⌬U7–U1 and ⌬U7–U1 ⌬5⬘UTR deletions resulted in a down-regulation of activity.

FIG. 6. Effects of TGF␤1 on Fgf10 promoter activity. Promoter constructs were cotransfected with a ␤-gal-expressing vector into primary VMPCs. Cells were cultured for 36 h in CPSR-containing media and treated with (gray bars) or without (black bars) TGF␤1 (5 ng ml–1) for 8 h. Results represent luciferase values normalized for transfection efficiency (by ␤-gal assay) and standardized to the Full promoter luciferase value. The average core promoter activity is represented as a dotted line running down the graph. TGF␤1 treatment resulted in an approximately 2-fold down-regulation of activity in the Full, ⌬U7–U4, and ⫹Sp1 promoter constructs. No down-regulation was observed in the ⌬U7–U4, ⌬Sp1, and core promoter constructs. Deletion of the Sp1 site from ⌬U7–U4 resulted in a down-regulation of activity, similar to levels in the core promoter.

down-regulation of activity relative to the longest promoter construct (Fig. 5). Region U1 appeared to mediate a significant proportion of Fgf10 promoter activity and was further investigated. TGF␤ represses Fgf10 promoter activity

A consensus Sp1 binding site (–182 to –172) was identified in region U1 by DNA sequence analysis (Gene Jockey Software, Biosoft, Cambridge, UK). Sp1 sites may regulate gene

transcription and also may mediate the effect of TGF␤1 on gene transcription (21–24). Fgf10 promoter constructs were made to investigate both the effect of the Sp1 site on the activity of Fgf10 transcription and to determine whether the effect of TGF␤1 was mediated via this region (Fig. 6). Constructs were transfected into primary VMPCs, and TGF␤1 treatment resulted in approximately 2-fold down-regulation of activity in the Full, ⌬U7–U4, and ⫹Sp1 promoter constructs to similar levels as those observed in the core pro-

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moter. No down-regulation was observed in the ⌬U7–U4 ⌬Sp1 and core promoter constructs after TGF␤1 treatment. Deletion of the Sp1 site from ⌬U7–U4 resulted in a decrease of activity, similar to levels of the core promoter. Discussion

We suggest that primary VMPCs are a good model to examine the regulation of Fgf10 gene expression in the prostate as they maintained characteristics and markers specific to the VMP and not urethral stroma or general mesenchyme. VMPCs showed differences in growth rate and cell size as well as elevated levels of expression of Tgf␤3 mRNA, when compared with URSCs. TGF␤1 repressed Fgf10 transcript levels in primary VMPCs and in VMP organ rudiments. The repression observed in primary VMPCs was rapid and reversible. This result is consistent with data published in 3T3 fibroblast cells (12) and also with data in lung mesenchyme (13). Hence, TGF␤1 repression of Fgf10 may be a regulatory mechanism conserved between the lung and the prostate. TGF␤1 did not regulate Fgf10 transcript levels in the UR, although levels of Fgf10 mRNA were very low in this tissue. The lack of regulation of Fgf10 mRNA in UR suggests that regulation in the VMP and VMPC is specific. Lack of Fgf10 regulation by TGF␤1 in UR may be due to the close proximity of UR stroma with epithelia of the urethra. In the lung, epithelial cells have been proposed to produce factors that regulate Fgf10 transcript levels in the mesenchyme (9). We observed that TGF␤s were expressed in both organ rudiments as well as primary cells (Fig 2A), which raises the question of why the addition of exogenous TGF␤1 downregulated Fgf10 mRNA, as TGF␤s are likely to be made by VMP and VMPCs. TGF␤1 is secreted in a latent form and its activity is regulated by extracellular processing. Endogenous TGF␤ protein made in our cells and organs may not act as a regulator of Fgf10 gene expression until extracellular processing has occurred. In prostate stromal cells grown in vitro, it has been demonstrated that less than 0.2% of secreted TGF␤ was biologically active (25), supporting the idea that in our primary cells and rudiments the endogenous TGF␤1 most probably would have no effect on Fgf10 transcript levels unless processed. It is possible that some of the TGF␤ made by VMP or VMPCs is active, but it is unlikely that there is full activity as the addition of exogenous TGF␤ down-regulated Fgf10. It is possible that proteases that activate TGF␤ are produced by cells outside the VMP, such as cells in the SM layer or in prostatic epithelia. Deletion analysis of the Fgf10 promoter revealed a region of the promoter (U1) in close proximity to the transcription start site that was involved in promoter activity. Sequences close to the Fgf10 transcriptional start site have also been identified as containing elements regulating expression of Fgf10 during limb development using a transgenic approach (26). This study identified a region of 700 nucleotides 5⬘ to the translational initiation site as being important for expression in vivo but did not identify a specific element within the sequence required for expression. Given that our studies and those of Sasak et al. (26) have independently identified the proximal promoter region of the Fgf10 promoter, it suggests that sequences at the transcriptional start site are important

Tomlinson et al. • Regulation of Fgf10 Expression in the Prostate

for Fgf10 expression. Additionally, certain genes have been shown to be Sp1 dependent where the Sp1 site participated in transcription initiation (21–23, 27, 28). Because Fgf10 transcription was reduced when the Sp1 site (–182 to –172) was deleted from the Fgf10 promoter construct, it is likely that the Sp1 site was participating in Fgf10 transcription. Also, several studies have shown that regulation of gene expression by TGF␤ involves interactions of the Sp1 transcription factor and Sp1 binding sites (21–24, 29 –32). In our system, the Sp1 site seemed to not only regulate Fgf10 gene expression but may have also mediated TGF␤1 repression of Fgf10 gene expression to levels observed in the core promoter. A key question is: what are the proteins binding to this site? It is possible that either Sp1 or TGF␤-inducible early gene (TIEG) proteins may be binding to this sequence. TIEGs are induced by the presence of TGF␤ (33, 34) and may cause a repression of gene transcription (35–37). TIEGs also have the ability to bind to GC-rich regions, like Sp1 sites (34, 37, 38); therefore, they may mediate their effect via competing for binding with Sp1 (33, 34). Comparison of Fgf10 mRNA levels between VMP and VMPCs indicated that levels were higher in the VMP. This raises that possibility that key factors that stimulate Fgf10 expression in vivo may have been lost upon cellular dissociation and growth in vitro (in the VMPC system). Our studies have shown that Fgf10 mRNA levels can be down-regulated by TGF␤ in primary cells and organs and have identified a region of the promoter involved in this regulation. The down-regulation observed was rapid and transient, and this may have important ramifications for the speed with which interactions occur in vivo. It might suggest that a modest change in FGF10 signaling for 24 – 48 h is sufficient to affect subsequent decisions regarding cell fate or cell-cell communication. However, no defects have been reported in mice heterozygous for a deletion of the Fgf10 gene, suggesting that a potential 50% reduction in expression may have no biological effect. In either of these examples it is assumed that changes in mRNA level rapidly lead to changes in protein level. We have attempted to study levels of FGF10 protein but have been unable to identify a suitable antiFGF10 antibody (either from commercial sources or raised by ourselves). It is known that factors produced in the epithelium and stroma can regulate Fgf10 expression in the mesenchyme. TGF␤1 protein is expressed in prostatic stroma, and it has been shown to be a factor that regulates prostate growth and development. We have shown that TGF␤1 can regulate Fgf10 expressed in inductive prostate cells and tissue, and we are proposing that this regulation may occur during prostate development. Also we have identified a region of the Fgf10 promoter that not only regulates the level of Fgf10 expression but also mediates the effect of TGF␤1. Acknowledgments We thank Cathal Grace, Denis Doogan, and James MacDonald for technical assistance. Received July 8, 2003. Accepted January 6, 2004. Address all correspondence and requests for reprints to: Axel A. Thomson, Medical Research Council Human Reproductive Sciences Unit, Centre for Reproductive Biology, The University of Edinburgh

Tomlinson et al. • Regulation of Fgf10 Expression in the Prostate

Chancellor’s Building, 49 Little France Crescent, Old Dalkeith Road, Edinburgh EH16 4SB, Scotland, United Kingdom. E-mail: axel.thomson@ hrsu.mrc.ac.uk. This work was supported by grants from the Medical Research Council UK and Congressionally Directed Medical Research Program for Prostate Cancer Research (DAMD17-00-1-0034 to A.A.T.).

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