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Division of Endocrinology, Metabolism, and Molecular Medicine (R.S., J.N.A., W.E.T., M.B., N.F.G.-C., S.B.), Charles R. Drew University of Medicine and Science, Los Angeles, California 90059; Beth Israel Deaconess Medical Center and Harvard Medical School (X.Y.), Boston, Massachusetts 02215; and Section of Endocrinology, Diabetes, and Nutrition (S.B.), Boston Medical Center, Boston, Massachusetts 02118
Address all correspondence and requests for reprints to: Rajan Singh, Ph.D., Division of Endocrinology, Metabolism, and Molecular Medicine, Charles R. Drew School of Medicine, 1731 East 120th Street, Augustus F. Hawkins Building, Los Angeles, California 90059. E-mail: rajansingh{at}mednet.ucla.edu.
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and -
and peroxisome proliferator-activated receptor-
2 protein and mRNA. These inhibitory effects of androgens were partially blocked by flutamide or bicalutamide. Androgen treatment was associated with nuclear translocation of ß-catenin and AR. Immunoprecipitation studies demonstrated association of ß-catenin with AR and T-cell factor 4 (TCF4) in the presence of androgens. Transfection of TCF4 cDNA inhibited adipogenic differentiation, whereas a dominant negative TCF4 cDNA construct induced adipogenesis and blocked testosterone’s inhibitory effects. Our gene array analysis indicates that testosterone treatment led to activation of some Wnt target genes. Expression of constitutively activated AR fused with VP-16 did not inhibit the expression of CCAAT/enhancer binding protein- in the absence of androgens. Testosterone and dihydrotestosterone inhibit adipocyte differentiation in vitro through an AR-mediated nuclear translocation of ß-catenin and activation of downstream Wnt signaling. These data provide evidence for a regulatory role for androgens in inhibiting adipogenic differentiation and a mechanistic explanation consistent with the observed reduction in fat mass in men treated with androgens. |
Introduction |
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Androgens bind to AR, which mediates most of its physiological functions through transcriptional activation of downstream genes (17, 18, 19, 20). AR has been detected in human and rat preadipocytes and adipocytes where it may be involved in regulating proliferation and differentiation of preadipocytes or pluripotent cells (2, 17, 18, 21). In addition, Ramirez et al. (12) have shown that androgen treatment of fully differentiated fat cells can inhibit expression of LPL and GAPDH to reduce fat mass. In addition, it has been suggested that androgens elicit antiadipogenic effects in the adipose precursor cells in specific regions where AR is expressed at a high level (17, 18). Adult male AR knockout mice exhibit a pseudofemale phenotype, and they have greater amounts of body fat than wild-type, male littermates (2, 22). These AR knockout mice are testosterone resistant, and they become obese and gain weight in sc and ip white adipose tissues (2). These studies suggest that AR may serve as a negative regulator of adipocyte development. Recently, it has been reported by various groups that AR interacts with ß-catenin protein in the Wnt pathway in various cell lines (23, 24, 25, 26). However, it is not known whether antiadipogenic effects of androgens are mediated through the downstream Wnt signaling pathway.
The Wnt gene family implicated in the control of adipogenesis includes over 16 Wnt proteins, soluble secreted glycoproteins that signal through their frizzled receptors and have profound effects on cellular differentiation and growth (27, 28, 29, 30, 31, 32, 33, 34, 35). Activation of Wnt signaling inhibits glycogen synthase kinase 3ß and allows cytosolic ß-catenin to accumulate and translocate to the nucleus where it binds to the T-cell factor (TCF)/lymphoid-enhancer factor (LEF) family of transcription factors and activates transcription of Wnt-regulated target genes. In the absence of Wnt signaling, glycogen synthase kinase 3ß phosphorylates ß-catenin and targets it for ubiquitin-mediated degradation. MacDougald’s group (35, 36) has demonstrated that Wnt signaling functions as an adipogenic switch that represses adipogenesis when activated and initiates adipogenesis when it is turned off. These investigators reported that Wnt signaling, possibly mediated by Wnt 10b, maintains preadipocytes in an undifferentiated state through activation of ß-catenin and TCF4, leading to subsequent inhibition of two well-characterized, key adipogenic transcription factors, C/EBP- and peroxisome proliferator-activated receptor- (PPAR-), which are necessary and sufficient for adipogenic differentiation (35, 37, 38). Overexpression of dominant negative TCF4 (Dn-TCF4), which disrupts Wnt signaling, stimulates preadipocyte differentiation into mature adipocytes, whereas activation of Wnt signaling through TCF4 by lithium chloride blocks fat cell differentiation (35).
The role of the Wnt signaling pathway in mediating androgen effects on adipogenic differentiation has not been investigated. Although AR interacts with ß-catenin protein in the Wnt pathway in some biological systems (23, 24, 25, 26), the biological relevance of AR and ß-catenin interaction is not fully understood. We considered the possibility that AR cross-talk and interaction with ß-catenin/TCF may bypass canonical Wnt signaling to modulate adipogenic differentiation by directly activating downstream Wnt effector molecules including TCF/LEF.
We hypothesized that testosterone and DHT, the two potent androgens, inhibit adipogenic differentiation of preadipocytes by molecular mechanisms that involve AR and ß-catenin interaction and modulation of downstream Wnt signaling through activation of TCF4. We tested this hypothesis in a 3T3-L1 cell line that has been used widely to investigate the mechanisms of preadipocyte differentiation; these cells, when grown in appropriate culture conditions, differentiate into adipocytes and recapitulate many of the endocrine and metabolic functions of mature adipocytes in vivo (39). We evaluated the expression of AR protein and mRNA in 3T3-L1 cells and determined the effects of testosterone and DHT on preadipocyte differentiation. We investigated the possible involvement of downstream effecter molecules in the Wnt signaling pathway by studying AR-ß-catenin interaction and ß-catenin nuclear translocation, a hallmark of Wnt activation, and determined the effects of TCF4 and Dn-TCF4 overexpression on inhibition of adipogenic differentiation by testosterone. We constitutively expressed AR-VP16 and investigated its effect on the adipogenic differentiation in this cell line, in the absence of androgens or nuclear ß-catenin, to determine whether androgen-induced inhibition of adipogenesis is mediated through AR binding to androgen response elements (AREs) in the promoter DNA or through TCF4/LEF DNA binding sites. We observed that constitutively expressed AR induced the protein expression of p21, which contains ARE binding sites in the promoter region (40); however, C/EBP- level remained unchanged and ß-catenin expression was not detected in the nucleus in 3T3-L1 cells.
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Oil Red O staining
3T3-L1 cells were fixed in 2% paraformaldehyde after various treatments and stained with 0.5% Oil Red O (Sigma Chemical Co., Saint Louis, MO) for 15 min as described previously (36). For quantitative analysis of Oil Red O retention in these cells, stained adipocytes were extracted with 1 ml of 4% Igepal CA-630 (Sigma) in isopropanol, and absorbance was measured by spectrophotometry at 520 nm.
Western blot analysis
Cell lysates (50–100 µg) in lysis buffer (20 mM Tris, 0.5% SDS containing protease inhibitors) were subjected to Western blot analyses by 7.5–12% SDS-PAGE, using 1:500 dilutions of anti-C/EBP-ß, anti-C/EBP-, and anti-AR antibodies; 1:1000 anti-PPAR-; 1:300 anti-C/EBP-; 1: 500 anti-p21; or 1:5000 anti-GAPDH antibody (Santa Cruz Biotechnology, Santa Cruz, CA). The washed filters were incubated with 1:2000 dilution of secondary antibodies linked to horseradish peroxidase. Immunoreactive bands were visualized by using the ECL detection system (Amersham Biosciences, Piscataway, NJ).
Quantitating adipogenic and Wnt target gene mRNA expression by RT-PCR and real-time quantitative PCR
Total RNA was extracted by using Trizol reagent, and equal amounts (2 µg) of RNA was reverse transcribed using RNA PCR kit (Applied Biosystems, Foster City, CA). The locations of forward/reverse PCR primers for real-time RT-PCR are as follows: AR (187 bp), 1937–1958/2124–2102 on S56585; GAPDH (152 bp), 606–626/758–738 on BC023196; PPAR-2 (241 bp), 79–99/320–299 on BC021798; C/EBP- (225 bp), 843–864/1067–1047 on NM_007678; LEF1 (132 bp), 1539–1558 /1661–1643 on NM_010703.2; and fatty acid binding protein 2 (AP2) (178 bp), 221–237/399–383 on K02109. The primers for PPAR-2 are specific for PPAR-2 and do not detect PPAR-1. Mouse gene PCR primer sets (RT2) were purchased from SuperArray Bioscience (Frederick, MD) for CD44 (183 bp), PPM03628A on NM_009851; and follistatin (Fst) (150 bp), PPM04451A on NM_008046. The QIAGEN Sybr Green PCR kit with HotStar Taq DNA polymerase was used (QIAGEN, Valencia, CA) with i-Cycler PCR thermocycler and fluorescent detector lid (Bio-Rad, Hercules, CA). The protocol includes melting for 15 min at 95 C, 40 cycles of three-step PCR including melting for 15 sec at 95 C, annealing for 30 sec at 58 C, elongation for 30 sec at 72 C with an additional detection step of 15 sec at 81 C, followed by a melting curve from 55–95 C at the rate of 0.5 C per 10 sec; except that for primers Fst, CD44, and LEF1, annealing was at 55 C and detection was at 76 C. We confirmed that inverse derivatives of melting curves show sharp peaks for PPAR-2 at 83.5 C, C/EBP- at 88 C, AP2 at 84 C, CD44 at 84.5 C, LEF1 at 85.5 C, Fst at 84.5 C, and GAPDH at 87 C, indicating the correct products. Samples of 25 ng cDNA were analyzed in quadruplicate in parallel with GAPDH controls; standard curves (threshold cycle vs. log pg cDNA) were generated by log dilutions of from 0.1 pg to 100 ng standard cDNA (reverse-transcribed mRNA from 3T3-L1 in AM), and then experimental mRNA starting quantities were calculated from the standard curves and averaged using i-Cycler, iQ software as described previously (21). The ratios of marker experimental gene (e.g. PPAR-2 mRNA) to GAPDH mRNA were computed and normalized to control (untreated) samples as 100%.
SuperArray analysis of Wnt target genes
Total cellular RNA isolated from 3T3-L1 cells undergoing adipogenic differentiation for 48 h with or without testosterone (100 nM) treatment was subjected to cDNA gene array (GEArray Q Series, MM-043; SuperArray BioScience) analysis. This series of mouse Wnt signaling pathway gene array is designed to study the genes involved in and downstream of Wnt signaling. Biotin-labeled cDNA probes were synthesized from total RNA, denatured, and hybridized overnight at 60 C in GEHybridization solution to membranes spotted with Wnt signaling pathway-specific genes. Membranes were washed, and chemiluminescent analysis was performed as per the manufacturer’s instructions. Raw data were analyzed using GEArray Expression Analysis Suite (SuperArray), and fold changes in relative gene expression were presented after background correction and normalization with a housekeeping gene.
Detection of AR, ß-catenin, and adenomatous polyposis coli (APC) by immunofluorescence
3T3-L1 cells treated with or without testosterone for various time points were grown in 10% FBS in DMEM on eight-well chamber slides and fixed in 2% paraformaldehyde for 20 min, and for AR and APC assay, they were blocked with normal goat serum and incubated with rabbit anti-AR antibody (N20; Santa Cruz Biotechnology) at 1:50 dilution or rabbit anti-APC (N15; Santa Cruz Biotechnology) at 1:300 dilution. For ß-catenin, the blocking step was done with normal horse serum followed by incubation with a monoclonal antibody against ß-catenin (C19220; BD Biosciences PharMingen, San Diego, CA) at 1:500 dilutions. The detection of AR and APC was done with a 1:200 dilution of antirabbit biotinylated secondary antibody (Calbiochem, La Jolla, CA) or for ß-catenin with antimouse biotinylated secondary antibody followed by streptavidin-fluorescein isothiocyanate (FITC) or Texas Red (Vector Laboratories, Burlingame, CA). After several washes, the slides were counterstained with 4',6-diamidino-2-phenyl-indole (DAPI) and mounted in prolong fade (Molecular Probes, Eugene, OR) and were examined under a fluorescence microscope equipped with the appropriate filters (42).
Double-labeling immunodetection of AR and ß-catenin
The double localization of AR and ß-catenin was carried out on 3T3-L1 cells treated with or without testosterone for various time points grown in 10% FBS in DMEM on eight-well chamber slides and fixed in 2% paraformaldehyde for 20 min. For AR, cells were blocked with normal goat serum and incubated with rabbit anti-AR antibody (N20; Santa Cruz Biotechnology) at 1:50 dilution, followed by a 1:200 dilution of antirabbit biotinylated secondary antibody (Calbiochem). The subsequent reaction was carried out by incubating the cells in a 20-µg/ml solution of streptavidin-FITC (Vector), followed by 10% normal horse serum and then a 1:500 dilution of anti-ß-catenin monoclonal antibody. Fluorescence labeling was performed with secondary antibody linked to Texas Red. The slides were counterstained with DAPI and mounted in prolong fade (Molecular Probes) and were examined under a fluorescence microscope equipped with the appropriate filters (42).
Cell fractionation and Western blot analysis to analyze ß-catenin nuclear translocation
3T3-L1 cells treated with androgens (100 nM testosterone and 10 nM DHT) for 0–24 h, harvested, and separated into cytoplasmic and nuclear fractions using the nuclear and cytoplasmic extraction reagent (Pierce, Rockford, IL). Nuclear and cytoplasmic fractions (20–40 µg) were separated on 10% SDS-PAGE, and proteins were transferred onto polyvinylidene difluoride membrane (Amersham). Membranes were blocked with 5% nonfat milk in 0.05% Tween 20 in PBS for 1 h and incubated with ß-catenin primary antibody (1:500 dilutions, C19220) for 2 h at room temperature. Detection of immunoreactive bands was achieved with antimouse-horseradish peroxidase-linked secondary antibody (Santa Cruz Biotechnology; sc-2031) and ECL detection reagents (Amersham; RPN 2106). The purity of nuclear and cytoplasmic fractions was assessed (data not shown) by using known cytoplasmic (hsp90) and nuclear (oct1) proteins.
Immunoprecipitation and immunoblot analysis to detect AR, ß-catenin, and TCF4 interactions
The interactions between AR, ß-catenin, and TCF4 were studied by immunoprecipitation. Cells were lysed using lysis buffer containing 20 mM Tris-HCl (pH 7.8), 0.5% Nonidet P-40, 137 mM NaCl, 50 µM EDTA, and protease inhibitors (Roche Diagnostics, Mannheim, Germany). Cell lysates were passed through a 30.5-gauge needle to disrupt nuclei. Protein extracts (500 µg) were incubated with either anti-AR (N20; Santa Cruz Biotechnology) or anti-TCF4 (Santa Cruz Biotechnology; Upstate, Charlottesville, VA) antibodies overnight at 4 C, followed by 1 h incubation with protein A/G-Sepharose (Calbiochem). Control immunoprecipitations were performed with rabbit preimmune serum. After three washes with 0.5 ml lysis buffer, the pellets were suspended in SDS sample buffer, boiled for 5 min, and analyzed on 10% SDS-PAGE. Proteins were transferred to a polyvinylidene difluoride membrane and blotted with anti-ß-catenin, anti-AR, or anti-TCF4 antibodies.
Transient transfection and overexpression of full-length TCF4 and dominant-negative TCF4 (Dn-TCF4)
Full-length TCF4 cDNA and Dn-TCF4 cDNA constructs were kindly provided by Dr. M. Wierman (VA Medical, Denver, CO) and Dr. Eric Fearon (University of Michigan, Ann Arbor, MI), respectively. The Dn-TCF4 clone lacks 31 amino acids from the N terminus (TCF4 
N31) and has been used successfully in previous studies to inhibit TCF function using a retroviral expression system (35, 36). However, we employed a transient transfection system using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) with approximately 35–40% efficiency.
Transfection of constitutively active pAct-AR and dual luciferase assay
3T3-L1 cells were transfected with plasmids encoding full-length AR fused to VP16 activation domain (pAct-AR), ARE-promoter-luciferase (pARE4-Luc), and empty pGL3 vectors either transfected alone or in combination using Lipofectamine 2000 with standard protocols. Cells were allowed to differentiate in adipogenic conditions for 9 d, and C/EBP- mRNA and protein levels were analyzed. As a control for endogenous AR-responsive gene, we measured the expression of p21 after transfection of 3T3-L1 cells with pARE4-Luc and pAct-AR vectors. The transfection efficiency in these experiments was measured both by immunostaining of the transfected cells with hemaglutinin (HA) antibody as well as by parallel transfection using GFP vector. Our transfection efficiency under these conditions was 40–50%. For luciferase assay, cells were simultaneously cotransfected with Renilla luciferase plasmid pRL-TK (50:1 mix of luciferase constructs and pRL-TK), harvested after 2 d using passive lysis buffer as described by the manufacturer (Promega, Madison, WI), and data are represented after normalization with Renilla luciferase. For immunofluorescence assay, cells after transfection and growth for 24 h were fixed in 2% paraformaldehyde and stained with anti-ß catenin antibodies or anti-AR (data not shown). Nuclear staining in these cells was detected by counterstaining with DAPI.
Image analysis
Oil Red O staining was quantified by image analysis using the Image Pro 4.01 software (Media Cybernetics, Silver Spring, MD), coupled to a Leica DMLB microscope/VCC video camera. After images were calibrated for background lighting, integrated OD (IOD = area x average intensity) was calculated using at least 20 pictures per treatment group (21). Results are proportional to the unweighted average OD, which was used to determine the Oil Red O staining retained in the fat cells. The experiment was repeated three times.
Statistics
Data are presented as mean ± SEM. Differences between the groups were analyzed by ANOVA. If overall ANOVA revealed significant differences, then pairwise comparisons between groups were performed using Tukey’s posttest procedure (see Figs. 2
and 6B) or Student’s t test (see Figs. 3F, 6C, and 7B). All comparisons were two-tailed, and P values < 0.05 were considered statistically significant. The experiments were repeated three times, and data from representative experiments are shown.
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). Western blot analysis confirmed that AR protein is expressed in untreated cells, and the AR protein expression levels remained unchanged after testosterone treatment (Fig. 1B). The AR mRNA expression was also detected by real-time RT-PCR in this cell line (data not shown). Thus, our data demonstrate the expression of AR mRNA and protein in this preadipocyte cell line.
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, top). We observed a dose-dependent reduction in the number of adipocytes after androgen treatment in a dose-dependent manner (Fig. 2, top); however, small changes in the overall sizes of adipocytes after androgen treatment cannot be ruled out. We further performed quantitative spectrophotometric analysis (OD, 520 nm) of Oil Red O-stained cells eluted with isopropanol, which revealed that testosterone and DHT both significantly inhibited adipogenic differentiation of these cells in a dose-dependent manner compared with the cells allowed to differentiate in control wells (Fig. 2, bottom). We observed that 100 nM testosterone caused 52 ± 6% inhibition, whereas 30 nM DHT was able to inhibit the adipogenesis by 40 ± 5%. Flutamide (200 nM), an AR antagonist, significantly blocked the inhibitory effects of both testosterone and DHT on the adipogenic differentiation of 3T3-L1 cells. Theses data suggest that testosterone and DHT both inhibit adipogenic differentiation of 3T3-L1 cells via an AR-dependent pathway.
Time course of C/EBP-ß and C/EBP- expression in differentiating 3T3-L1 cells and effect of testosterone on their expression
To determine the effects of testosterone on the expression of early markers of adipogenic differentiation, we performed time course experiments with 3T3-L1 cells undergoing differentiation either in presence or absence of testosterone (100 nM) treatment. The expression of C/EBP-ß was detectable as early as 2 h and significantly decreased after 96 h of adipogenic differentiation. Testosterone inhibited the C/EBP-ß expression at early times (2 h); however, at later time points, this effect was not significant (Fig. 3A). On the other hand, we observed that C/EBP- was expressed at a relatively later time point compared with C/EBP-ß (8 h compared with 2 h) and peaked at about 24 h, and its levels were barely detectable after 96 h. Testosterone significantly inhibited C/EBP- expression in these cells at 2–24 h.
Testosterone and DHT inhibit key adipogenic transcription factors C/EBP- and PPAR-2 in 3T3-L1 cells
We determined the effects of graded concentrations of testosterone and DHT on the protein and mRNA expression pattern of key adipogenic transcription factors C/EBP- and PPAR- by Western blot analysis and quantitative real-time RT-PCR. Incubation of 3T3-L1 cells with testosterone (0–100 nM) and DHT (0–10 nM) for 12 d significantly inhibited the expression of 54-kDa PPAR- (Fig. 3B) and 42- and 30-kDa C/EBP- (Fig. 3D) proteins in a dose-dependent manner. Densitometric analysis (Fig. 3C) of the respective bands showed 51 and 42% inhibition of PPAR- with 100 nM testosterone and 10 nM DHT, respectively. Similarly, we observed 77 and 49% inhibition of C/EBP- (sum of total intensity of 42- and 30-kDa bands) after testosterone and DHT treatment (Fig. 3E). Treatment of 3T3-L1 cells for 12 d with 0–100 nM testosterone also down-regulated PPAR-2 and C/EBP- mRNA expression (Fig. 3F), as measured by real-time RT-PCR. Figure 3E demonstrates that PPAR-2 mRNA was significantly down-regulated by testosterone treatment; maximal inhibition was observed at 100 nM testosterone (
60% inhibition). Testosterone also down-regulated C/EBP- mRNA in a dose-dependent manner; maximal inhibition was observed at 100 nM testosterone (74%).
Time course of nuclear translocation of ß-catenin in 3T3-L1 cells by testosterone and DHT treatment
To elucidate the possible mechanisms involved during testosterone- and DHT-induced inhibition of adipogenic differentiation, we performed immunofluorescence experiments to study the localization of ß-catenin in these cells after treating the cells with 10 nM DHT and 100 nM testosterone for 24 h. We analyzed the localization of ß-catenin, a key protein that translocated to the nucleus during activation of Wnt signaling after these treatments. In cells treated with medium alone, ß-catenin was localized mainly in the cytoplasm and the cytoskeleton. A fraction of ß-catenin protein translocated to the nucleus after testosterone (100 nM) and DHT (10 nM) treatment for 24 h (Fig. 4A). Simultaneous treatment of the 3T3-L1 cells with 10-fold molar excess of AR antagonist bicalutamide blocked testosterone- and DHT-induced nuclear translocation of ß-catenin (Fig. 4A), suggesting that AR plays an important role during this process of ß-catenin nuclear translocation. Lithium chloride treatment of these cells for 24 h also resulted in the nuclear translocation of ß-catenin as expected.
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). ß-Catenin immunoreactivity was detectable in the nuclear fraction as early as 1 h after testosterone and DHT treatment, and there was a concomitant decrease in ß-catenin protein in the cytoplasmic fractions, consistent with our data from the immunofluorescence experiments described in Fig. 4A. Simultaneous treatment of these testosterone-treated cells with bicalutamide resulted in decreased immunoreactivity for ß-catenin protein in the nuclear fraction (Fig. 4B, bottom) after 24 h. These data indicate that DHT and testosterone activate Wnt signaling by translocating ß-catenin to the nucleus in this preadipocyte cell line.
The shuttling of APC protein between different subcellular components and its control vary between different cell types and states (43, 44). To investigate the possible role of APC during testosterone- and DHT-induced inhibition of adipogenesis, we performed immunofluorescence experiments to study the localization of APC before and after treatment of the cells with the androgens. APC was expressed mostly in the nucleus under basal conditions (Fig. 4C), and there was no colocalization of APC with ß-catenin in the nucleus after testosterone treatment (data not shown). Nuclear localization of APC has also been demonstrated in certain cell lines under basal conditions (43, 45, 46).
Physical interaction between AR, ß-catenin, and TCF4 in DHT-treated 3T3-L1 cells
Using double immunofluorescence, we observed that AR and ß-catenin were colocalized in the nucleus in both testosterone-treated (100 nM) and DHT-treated (10 nM) cells after 24 h (Fig. 5A) as observed by merging the AR- and ß-catenin-stained pictures (yellow). The respective positions of the nuclei were detected by counterstaining the cells with DAPI (blue). We also performed immunoprecipitation experiments to evaluate the interaction between AR, ß-catenin, and TCF4 using cell lysates obtained from 3T3-L1 cells after DHT (0–30 nM) treatment (Fig. 5B). Protein extracts were immunoprecipitated using anti-AR (Fig. 5B, top) or anti-TCF4 antibody (Fig. 5B, bottom), and immunoprecipitates were analyzed by immunoblotting with anti-ß-catenin, anti-AR, or anti-TCF4 antibodies (Fig. 5B, top) or with anti-ß-catenin and anti-TCF4 or anti-AR antibodies. Except for a minimal AR-TCF4 interaction, these interactions were not detected in untreated cells, suggesting that DHT treatment was necessary for these interactions and complex formation. These interactions were significantly inhibited in the presence of 10-fold molar excess of bicalutamide (100 nM), an AR antagonist. A higher concentration of DHT (30 nM) led to an enhancement of this complex formation (Fig. 5B). These immunoprecipitation studies are consistent with our findings from immunofluorescence studies and demonstrate the translocation of ß-catenin to the nucleus in a complex with AR and TCF4. Bicalutamide (100 nM), an AR antagonist, blocked the DHT induction of ß-catenin nuclear translocation. These data thus provide evidence of both ß-catenin-AR and ß-catenin-TCF4 interactions.
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As expected, in 3T3-L1 cells, overexpressing TCF4 cDNA expression construct was associated with a significant inhibition of fat cell formation, as analyzed by Oil Red O staining of the fat cells (Fig. 6A). Testosterone treatment of 3T3-L1 cells that had been transfected with the TCF4 cDNA construct had no additional inhibitory effect on fat cell formation or adipogenic differentiation of 3T3-L1 cells, as measured by quantitative image analysis of the Oil Red O-positive cells (Fig. 6B). In contrast, 3T3-L1 cells transfected with Dn-TCF4 cDNA construct had a significantly greater number of adipocytes than control wells. Testosterone’s inhibitory effects on preadipocyte differentiation were blocked by overexpression of Dn-TCF4 cDNA construct (Fig. 6B), suggesting that TCF4 signaling pathway was necessary for mediating testosterone-induced inhibition of adipogenesis. Overexpression of TCF4 protein in these transfected cells was verified by Western blotting (data not shown).
To examine adipose gene regulation downstream from TCF4, we measured expression of mRNA in 3T3-L1 cells transfected with TCF4 or Dn-TCF4, by using real-time quantitative RT-PCR for the key adipogenic early marker C/EBP- and late marker AP2 normalized to GAPDH (Fig. 6C). Overexpression of TCF4 in these preadipocytes significantly inhibited expression of C/EBP- by over 50% and blocked AP2 by about 80% (Fig. 6C). However, Dn-TCF4 induced expression of both C/EBP- by 50% and AP2 by 100% and prevented inhibition by testosterone (Fig. 6C). As a standard control, treatment of 3T3-L1 cells grown in AM with 20 mM LiCl, which activates Wnt signaling, resulted in almost complete inhibition of adipogenesis and also blocked expression of C/EBP- and AP2 mRNA, as measured by quantitative RT-PCR (data not shown). Taken together, these data suggest that activation of downstream Wnt signaling pathway via AR/ß-catenin/TCF4 plays an important role in mediating testosterone’s inhibitory effects on adipogenic differentiation.
Testosterone up-regulation of Wnt target genes
Because androgen treatment induced AR interaction with ß-catenin and androgen-induced nuclear translocation of ß-catenin bypassed the canonical Wnt signaling for activation of TCF/LEF, we tested the ability of androgens to activate expression of some well-known downstream Wnt target genes including Lef1 and Fst. We performed SuperArray analysis of 3T3-L1 cells undergoing adipogenic differentiation either in the presence or absence of testosterone (100 nM), using a Wnt gene-specific array. We observed that testosterone treatment led to significant activation of CD44 antigen, Fst, and Lef1 genes (Fig. 7A and Table 1).
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). These data confirm that although testosterone did not induce canonical Wnt signaling, it activated AR/ß-catenin interaction, their nuclear translocation, and activation of TCF4/LEF to activate known downstream Wnt-pathway target genes.
Constitutive activation of AR in 3T3-L1 cells does not inhibit adipogenic differentiation
To determine whether constitutively active AR protein fused with artificial coactivator VP16 inhibits adipogenesis in the absence of androgens, we expressed pACT- full-length AR (AR-VP16) and pARE4-Luc and allowed the cells to differentiate in AM for 9 d. Cells were lysed, and C/EBP- protein and mRNA levels were analyzed by Western blot and real-time RT-PCR analysis. We were unable to detect any significant changes in C/EBP- mRNA (left) or protein (right) expression (Fig. 8A). Our densitometric scan of C/EBP- protein (sum of 42- and 30-kDa) bands did not show a substantial change after normalization with GAPDH in samples where pAct-AR was expressed (data not shown). As a positive control to verify expression of activated AR-VP16 protein, plasmids encoding either pGL3 or pARE4-Luc alone or AR-VP16 in combination with pARE4-Luc along with pRL-TK-Luc were transfected and changed to AM for another 48 h. The transfection efficiency under the conditions varied from 45–55%. The values were expressed after normalization with Renilla luciferase. We observed that luciferase activity was significantly higher (75 ± 6-fold) in cells cotransfected with both AR-VP16 and pARE4-Luc compared with individual vectors alone (Fig. 8B), providing evidence that AR is constitutively active. We also analyzed the subcellular localization of AR (data not shown) and ß-catenin in these cells in which AR was constitutively active; we were unable to detect any nuclear translocation of ß-catenin under these condition (Fig. 8C). To measure expression of an endogenous AR-responsive gene, we further analyzed the expression of p21 protein after transfection. We observed that p21 level was significantly induced in cells transfected with pAct-AR compared with pARE4-Luc vector (Fig. 8D), suggesting that constitutively activated AR is able to activate endogenous AR-responsive genes.
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, C/EBP-, and PPAR-2, in a time- and concentration-dependent manner. These antiadipogenic effects of testosterone and DHT were blocked by an AR antagonist, flutamide, indicating that these effects were transduced through an AR-mediated pathway, supported by the fact that AR mRNA and protein were expressed in these cells. Both testosterone and DHT induced AR translocation and colocalization with ß-catenin in the nucleus within 1 h of treatment. The AR protein formed a molecular complex with ß-catenin and TCF4, and this interaction was androgen dependent because it was blocked when cells were treated simultaneously with 10-fold molar excess of a specific AR antagonist, bicalutamide. Androgen inhibition of fat cell differentiation and C/EBP- and AP2 mRNA expression was mimicked by TCF4 overexpression but blocked by Dn-TCF4 in 3T3-L1 cells. These data lead us to conclude that androgens inhibit adipogenic differentiation of 3T3-L1 preadipocytes by activating noncanonical Wnt through activation of AR/ß-catenin-TCF4.
During early adipogenesis, a cascade of transcription factors is sequentially induced, including C/EBP-ß and C/EBP- at early time points, followed by C/EBP- and PPAR-2 (47). The expression of C/EBP-ß and C/EBP- was induced within 2–24 h in 3T3-L1 cells grown in AM for various times. Treatment with testosterone significantly inhibited expression levels of C/EBP- and subsequently C/EBP- and PPAR-2.
In a series of elegant studies, McDougald and colleagues (35, 36) have demonstrated that the Wnt signaling pathway plays an important role in regulation of adipogenic differentiation; however, the regulation of Wnt signaling pathway by androgens during adipogenic differentiation has not been previously studied. Wnts, a group of soluble secretory proteins, are important molecular switches that govern adipogenesis by inhibiting C/EBP- and PPAR-2 (35, 36, 38, 41). Disruption of extracellular Wnt signaling by soluble Wnt inhibitors, soluble frizzled receptor proteins-1 and –2 (36) or by overexpression of a Dn-TCF4 construct results in up-regulation of C/EBP- and PPAR-2 and spontaneous adipogenesis (35, 47). Our data provide direct evidence of the androgen-dependent interaction of AR and ß-catenin in 3T3-L1 preadipocytes. Using double immunofluorescence, we demonstrated the colocalization of AR and ß-catenin in the nucleus in a ligand-dependent manner. The immunoprecipitation studies provided additional confirmation of the physical interaction between AR and ß-catenin and between ß-catenin and TCF4; these interactions were androgen dependent and were significantly inhibited in the presence of the AR antagonist bicalutamide.
The most compelling evidence for the involvement of an AR/ß-catenin/TCF4 pathway in mediating androgen inhibition of adipogenic differentiation emerged from our experiments that used Dn-TCF4 cDNA constructs. In 3T3-L1 preadipocytes, transfection with Dn-TCF4 cDNA construct not only increased adipogenesis but also blocked testosterone’s effects on adipogenic differentiation. Thus, disruption of TCF4 and the consequent inhibition of downstream Wnt signaling resulted in an induction in overall adipogenic differentiation as expected and attenuation of testosterone’s effects on adipogenesis. Conversely, when TCF4 was constitutively overexpressed using a cDNA construct, adipogenesis was inhibited, and concomitant incubation with testosterone had no additional effect on adipogenesis. Although this does not prove that the TCF4 factor, as opposed to LEF1, is the endogenous molecule that mediates the androgen effect, it shows that this downstream Wnt signaling pathway is involved during the process. These results were confirmed in other experiments in which we used nucleofection of these constructs in 3T3-L1 cells and achieved over 70% transfection efficiency. Taken together, these data point to the pivotal role of a functional AR/ß-catenin/TCF4 pathway in mediating androgen effects on modulating adipogenesis.
Our studies provide additional evidence of the cross-communication between AR-mediated intracellular signaling and the downstream Wnt signal transduction cascade. Addition of androgen to the incubation medium was associated with AR translocation into the nucleus and its association with ß-catenin, as confirmed by double-immunofluorescence studies. This interaction was ligand dependent and blocked by an AR antagonist. The immunoprecipitation experiments provided additional proof of the physical interaction between AR and ß-catenin in the nuclear fraction of 3T3-L1 cells. Our data are thus consistent with previous reports that AR promotes nuclear translocation of ß-catenin in prostate cancer and other cells through specific protein-protein interaction in a ligand-dependent manner (23, 24, 25, 26). A strong interaction between the ligand binding domain of AR and the first six armadillo repeats of ß-catenin has been demonstrated (23, 24). ß-Catenin has been shown to interact with AR but not with other steroid hormone receptors.
During canonical Wnt signaling, ß-catenin is translocated to the nucleus in association with APC protein, which remains in the cytoplasm under normal conditions and is translocated to the nucleus after Wnt activation along with ß-catenin, to activate Wnt target genes through the TCF4-mediated pathway. However, in our experiments, nuclear translocation of ß-catenin was independent of APC. APC immunofluorescence was predominantly localized in the nucleus, and we were unable to detect colocalization of ß-catenin and APC either by double immunofluorescence or by immunoprecipitation either under basal conditions or after androgen treatment. Similar nuclear localization of APC has been reported in several other cells under basal conditions (44, 45, 46).
The data reported in this manuscript provide evidence that androgens inhibit differentiation of preadipocytes into mature adipocytes; these data are consistent with previous reports that testosterone inhibits fat cell differentiation of adipocyte precursor cells (10, 11, 12, 17). However, these studies do not exclude the possibility that androgens might exert additional effects at other steps in the adipogenic process. Our demonstration of androgen suppression of preadipocyte differentiation is only one possible mechanism of fat mass inhibition. In fact, previous reports suggest that testosterone may modulate body composition by exerting additional effects on lipid metabolism in fully differentiated fat cells (12). We have demonstrated previously that testosterone and DHT promote myogenic commitment and/or differentiation and inhibit adipogenic differentiation and PPAR-2 and C/EBP- expression in mesenchymal C3H 10T1/2 pluripotent cells (21). Testosterone also has been reported to suppress lipid uptake and LPL activity in adipocytes (12). Dieudonne et al. (11) reported that testosterone and DHT inhibit GAPDH activity significantly in epididymal preadipocytes. The net result of these androgen actions at multiple sites in the adipogenic and lipogenic processes is a reduction in fat mass.
Our findings have clinical relevance because they provide a rational mechanistic explanation for the observed decrease in whole-body fat mass during testosterone supplementation in human studies. We understand that our current findings using 3T3-L1 cells as a model system is not an accurate measure of what occurs in human preadipocytes. However, it still has important implications for drug discovery, especially development of selective AR modulators. Testosterone and DHT inhibit preadipocyte differentiation by activation of downstream target genes through noncanonical Wnt signaling. The intracellular events triggered by androgen binding to the AR are cross-communicated to the Wnt signaling pathway through an association of AR with ß-catenin and translocation of this complex to the cell nucleus. This results in downstream activation of TCF4/LEF and inhibition of adipogenesis, involving suppression of key adipogenic factors PPAR-2, C/EBP-, and C/EBP- and also activation of several Wnt pathway target genes including Fst, Lef1, and CD44 antigen (48, 49, 50). We also tested whether constitutively active AR fused with artificial coactivator VP16 (51, 52) is sufficient to inhibit adipogenesis in the absence of androgen and nuclear ß-catenin. We observed that constitutive AR activation alone in these cells does not inhibit C/EBP- mRNA and protein expression or ß-catenin translocation, although it significantly induced the protein expression of p21, an AR-responsive gene containing ARE in the promoter region (22). These findings might suggest that inhibition of adipogenesis in 3T3-L1 cells by androgens is mediated preferably through TCF4/LEF sites. Additional work is in progress to investigate this important aspect of androgen regulation of adipogenesis in details. The molecular events that follow TCF4 activation and result in inhibition of adipogenic differentiation are not well understood and need future investigation.
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First Published Online October 6, 2005
Abbreviations: AM, Adipogenic medium; AP2, fatty acid binding protein 2; APC, adenomatous polyposis coli; AR, androgen receptor; ARE, androgen response element; C/EBP, CCAAT/enhancer binding protein; DHT, dihydrotestosterone; Dn, dominant negative; FBS, fetal bovine serum; FITC, fluorescein isothiocyanate; Fst, follistatin; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GM, growth medium; LEF, lymphoid-enhancer factor; LPL, lipoprotein lipase; PPAR, peroxisome proliferator-activated receptor; TCF, T-cell factor.
Received December 22, 2004.
Accepted for publication September 9, 2005.
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2. Endocrinology 141:649–656
