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Molecular and Cellular Biology, September 2003, p. 6159-6173, Vol. 23, No. 17
0270-7306/03/$08.00+0     DOI: 10.1128/MCB.23.17.6159-6173.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Cyclin D1 Repression of Peroxisome Proliferator-Activated Receptor {gamma} Expression and Transactivation

Chenguang Wang,1 Nagarajan Pattabiraman,1 Jian Nian Zhou,2 Maofu Fu,1 Toshiyuki Sakamaki,1 Chris Albanese,1 Zhiping Li,1 Kongming Wu,1 James Hulit,2 Peter Neumeister,2 Phyllis M. Novikoff,3 Michael Brownlee,3 Philipp E. Scherer,4 Joan G. Jones,5 Kathleen D. Whitney,5 Lawrence A. Donehower,6 Emily L. Harris,7 Thomas Rohan,8 David C. Johns,9 and Richard G. Pestell1*

Department of Oncology, Lombardi Cancer Center, Georgetown University, Washington, D.C. 20007,1 Departments of Developmental and Molecular Biology,2 Medicine,3 Epidemiology and Population Health,8 Pathology,5 Cell Biology, Albert Einstein College of Medicine, Bronx, New York 10461,4 Division of Molecular Virology, Baylor College of Medicine, Houston. Texas 77030,6 Center for Health Research, Kaiser Permanente, Portland, Oregon 97227,7 Department of Neurosurgery, The Johns Hopkins Hospital, Baltimore, Maryland 212319

Received 21 November 2002/ Returned for modification 10 January 2003/ Accepted 9 May 2003


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The cyclin D1 gene is overexpressed in human breast cancers and is required for oncogene-induced tumorigenesis. Peroxisome proliferator-activated receptor {gamma} (PPAR{gamma}) is a nuclear receptor selectively activated by ligands of the thiazolidinedione class. PPAR{gamma} induces hepatic steatosis, and liganded PPAR{gamma} promotes adipocyte differentiation. Herein, cyclin D1 inhibited ligand-induced PPAR{gamma} function, transactivation, expression, and promoter activity. PPAR{gamma} transactivation induced by the ligand BRL49653 was inhibited by cyclin D1 through a pRB- and cdk-independent mechanism, requiring a region predicted to form an helix-loop-helix (HLH) structure. The cyclin D1 HLH region was also required for repression of the PPAR{gamma} ligand-binding domain linked to a heterologous DNA binding domain. Adipocyte differentiation by PPAR{gamma}-specific ligands (BRL49653, troglitazone) was enhanced in cyclin D1-/- fibroblasts and reversed by retroviral expression of cyclin D1. Homozygous deletion of the cyclin D1 gene, enhanced expression by PPAR{gamma} ligands of PPAR{gamma} and PPAR{gamma}-responsive genes, and cyclin D1-/- mice exhibit hepatic steatosis. Finally, reduction of cyclin D1 abundance in vivo using ponasterone-inducible cyclin D1 antisense transgenic mice, increased expression of PPAR{gamma} in vivo. The inhibition of PPAR{gamma} function by cyclin D1 is a new mechanism of signal transduction cross talk between PPAR{gamma} ligands and mitogenic signals that induce cyclin D1.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The cyclin-dependent kinase holoenzymes are a family of serine/threonine kinases that play a pivotal role in controlling progression through the cell cycle (38, 47). Dysregulation of the cell cycle control apparatus is an almost uniform aberration in tumorigenesis (48). The cyclins encode regulatory subunits of the kinases which phosphorylate specific proteins, including the retinoblastoma (pRB) protein, to promote transition through specific cell cycle checkpoints (47, 57). Cyclin D1 plays a pivotal role in G1/S phase cell cycle progression in fibroblasts and is rate limiting in growth factor- or estrogen-induced mammary epithelial cell proliferation (29, 67). Cyclin D1 overexpression is found in >30% of human breast cancers, correlating with poor prognosis (23). Several different oncogenic signals induce cyclin D1 expression, including mutations of the Ras and Wnt/APC/ß-catenin pathway (2, 49). Mammary-targeted expression of cyclin D1 is sufficient for the induction of mammary adenocarcinoma, and cyclin D1-/- mice are resistant to ErbB2-induced tumorigenesis (53, 64).

In addition to binding cyclin-dependent kinases 4 and 6 (cdk4 and cdk6) and pRB, cyclin D1 forms physical associations with P/CAF (p300/CBP-associated factor), Myb, MyoD, and the cyclin D1 myb-like binding protein (DMP1) (16, 20, 31, 39). Binding of cyclin D1 to the estrogen receptor alpha (ER{alpha}) enhances ligand-independent reporter gene activity, and liganded androgen receptor reporter gene activity is inhibited by cyclin D1 (33, 39, 68). The in vivo or genetic evidence indicating a requirement for cyclin D1 in nuclear receptor function remained to be determined.

The peroxisome proliferator-activator receptors, including PPAR{alpha}, PPAR{gamma}, and PPAR{delta}, are ligand-activated nuclear receptors (42). Their modular structure resembles those of other nuclear hormone receptors with N-terminal AF-1, a DNA binding domain, and a carboxyl-terminal ligand-binding domain (LBD). PPAR{gamma} was cloned as a transcription factor involved in fat cell differentiation and is required for the induction of adipocyte differentiation (41, 51). Adenoviral delivery of PPAR{gamma} to the livers of mice induces hepatic steatosis, consistent with an important role for PPAR{gamma} in hepatocellular lipid biosynthesis (65). The PPAR{gamma} ligands include eicosanoids, such as 15-deoxy-{Delta}12,14-prostaglandin J2 (15d-PGJ2), and synthetic ligands of the thiazolidinedione (TZD) class. PPAR{gamma} agonists inhibit the growth of human colorectal cancer cells (45) and promote fibroblast and breast epithelial cell differentiation (14, 32).

Since at least 1.6 million patients take TZDs as antidiabetic agents (42), it is important to understand PPAR{gamma} function, including its possible role in cancer. The possibility of inhibiting tumor cellular proliferation using PPAR{gamma} ligands as nontoxic therapeutics has provided the impetus to assess their efficacy in animal models. Mutation within the adenomatous polyposis coli (APC) pathway occurs frequently in human colon cancer and is sufficient for the induction of gastrointestinal polyposis in the Min mouse. PPAR{gamma} ligands inhibited the growth of implanted colonic tumors with APC mutations in one study (44); however, treatment of Min mice with PPAR{gamma} ligands increased polyposis in other studies (27, 43). Together these findings raise the possibility that PPAR{gamma} ligand effects in vivo may be governed by specific genetic determinants. Identifying the molecular genetic determinants governing PPAR{gamma} regulation of cellular proliferation and tumorigenesis in vivo is therefore of considerable importance. We examined the role of cyclin D1 as a genetic determinant of PPAR{gamma} function, since cyclin D1 has been implicated in the genesis of breast and colon cancer.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Reporter genes, expression vectors, DNA transfection, and luciferase assays. The acyl-coenzyme A oxidase triple PPAR{gamma} response element (PPRE) luciferase reporter gene [(AOX)3LUC], PPAR{gamma}-GAL4, pCMX-PPAR{gamma}, and UAS5E1B-TATALUC, the adenoviral vectors for ecdysone receptors (Ad-DB-Ecr-IRES-GFP and DB-Ecr), and retroviral vectors pBPSTR1 cyclin D1 sense and MSCV-IRES-GFP (9) were previously described (19, 39, 56). Cyclin D1 mutants were generated by PCR and subcloned into p3xFLAG-CMV-10 (Sigma). Cells were transfected by Superfect Transfection reagent (Qiagen, Valencia, Calif.). The HeLa, cyclin D1-/- mouse embryonic fibroblasts (MEFs,) and 3T3 cells (cyclin D1-/- and cyclin D1+/+) were previously described (1, 3a, 52). The medium was changed after 5 h, cells were treated with ligand or vehicle as indicated (see figure legends), and luciferase activity was determined after 24 h. Luciferase activity was normalized for transfection efficiency with ß-galactosidase reporters as an internal control. Luciferase assays were performed at room temperature with an Autolumat LB 953 (EG&G Berthold) (55). The fold effect was determined by comparison to the empty expression vector cassette, and statistical analyses were performed using the Mann Whitney U test.

Viral infection. The construction and preparation of the pBPSTR1-cyclin D1, ectopic, replication-defective helper virus was previously described (25). Retroviruses were prepared by transient cotransfection with helper virus into 293T cells using calcium phosphate precipitation (http://www.stanford.edu/group/nolan). cyclin D1-/- mouse embryonic fibroblasts (MEFs) were centrifuged for 90 min at 400 x g rpm at room temperature in DMEM medium with 10% fetal bovine serum, half volume of fresh retroviral supernatants, and 8 µg of polybrene/ml and then incubated overnight at 37°C in 5% CO2. The following day the media of the cyclin D1-/- MEFs was changed to DMEM with 10% fetal bovine serum and cultured for differentiation assay. A retroviral expression plasmid encoding green fluorescent protein (GFP) was included for monitoring infection efficiency (9).

Western blots and semiquantitative RT-PCR. The antibodies were polyclonal cyclin D1 antibody Ab3 for Western blot analysis, anti-PPAR{gamma} polyclonal antibody H100, and monoclonal E8 (Santa Cruz Biotechnology), anti-S3/12 (46), and anti-guanine dissociation inhibitor (GDI) (25) as a protein loading control. The membrane was incubated with horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology) and washed three times with 0.05% Tween 20-phosphate-buffered saline (PBS). Immunoreactive proteins were visualized by the enhanced chemiluminescence system (Amersham, Arlington Heights, Ill.), and their abundance was quantified by phosphorimaging (computing densitometer [Image Quant, version 1.11]; Molecular Dynamics, Sunnyvale, Calif.). Semiquantitative reverse transcription (RT)-PCR was conducted using 4 µg of total liver RNA from three age-matched pairs of wild-type (wt) and cyclin D1-/- mice which was reverse transcribed with the SuperScript II kit (Invitrogen) using Oligo(dT)12-18. The cDNA was diluted threefold with water sequentially three times; 1 µl of each of the dilutions was used in a PCR. Taq polymerase (TaKaRa) was used to amplify PPAR{gamma} and ß-actin using 35 cycles and 20 cycles of 94°C for 45 s, 58°C for 45 s, and 72°C for 45 s, respectively. The primers were the following: PPAR{gamma}, GTTGACACAGAGATGCCATTC (5' primer) and GGTTCTTCATGAGGCCTG (3' primer); ß-actin, TGTTACCAACTGGGACGACA (5' primer) and AAGGAAGGCTGGAAAAGAGC (3' primer).

Induction of differentiation. Primary MEFs were isolated from 14-day-postcoitus mouse embryos (1). 3T3-L1 and MEF cells were maintained at confluence for 1 day before being switched to basal differentiation medium (DMEM supplemented with 10% charcoal-stripped serum and 10 mg of insulin/liter). Differentiation was induced by serum supplemented with 0.2 mM methylisobutylxanthine (Sigma), 5 µM dexamethasone (Sigma), and insulin (Sigma) for 3 days. Subsequently, cells were maintained in basal differentiation medium supplemented with BRL49653 (0.2 µM) (P. G. Treagust [Smithkline Beecham, West Sussex, United Kingdom]) or troglitazone (5 µM) (Sanky Co., Ltd., Tokyo, Japan) or vehicle as indicated. Retroviral infection was conducted as previously described (25) (http://www.stanford.edu/group/nolan). For Oil Red-O staining, cells were fixed in 10% formaldehyde in PBS for 1 h and rinsed with water and ethanol. Cells were stained with Oil Red-O solution (six parts saturated Oil Red-O dye in isopropanol plus four parts water) at 37°C for 15 min, washed with 70% ethanol, and then rinsed with ddH2O. Cells were inspected by microscopy, and staining was quantified after incubation with 4% NP-40 in isopropanol and measuring of absorbance at 520 nm.

Ponasterone-inducible cyclin D1 antisense-IRES-GFP transgenic mice. The cDNA for the mouse cyclin D1 gene (a gift from V. Fantl) was subjected to internal deletion of bases 294 to 640 by PstI/StuI digestion followed by blunt-end ligation. An EcoRI fragment containing the modified murine cyclin D1 cDNA was inserted in the antisense orientation into the transgene shuttle vector, EGRE3{Delta}MMTV-BGH PolyA (3), modified to contain the IRES-GFP component from IRES2-EGFP (Clontech). The purified DNA was microinjected at the AECOM transgenic facility.

Immunostaining of human breast tumor samples. The studies were approved by the Albert Einstein College of Medicine Institutional Review Board. Material from 36 benign breast disease specimens was selected randomly from the Kaiser Permanent Northwest's Benign Breast Disease Registry using breast tissue obtained over 24 years. The tissues were evaluated by two pathologists using a standard template to identify histological features of benign breast disease, including proliferative and nonproliferative fibrocystic change, epithelial hyperplasia with and without atypia, and papillomas. Materials from 33 consecutive patients with infiltrating ductal or lobular carcinoma were obtained from the files of the Einstein Division of the Montefiore Medical Center. The benign and malignant material was formalin fixed and paraffin embedded. The primary antibodies were to cyclin D1 (monoclonal antibody [MAb] DCS-6, and polyclonal (Ab-3,), and to PPAR{gamma} (MAb E-8, Santa Cruz). The secondary antibodies were goat anti-mouse immunoglobulin G conjugated to horseradish peroxidase. For immunoperoxidase staining of paraffin-embedded tissue sections, the Santa Cruz Biotechnology ABC Staining Systems were used. Briefly, specimens were incubated for 1 h in 2% normal blocking serum derived from the same species in which the secondary antibody was raised in PBS. Sections were incubated with the primary antibody for 30 min at room temperature, washed with three changes of PBS for 5 min each, incubated for 30 min at 1 µg/ml diluted in PBS with 2% normal blocking serum, and then washed with three changes of PBS for 5 min each. Incubation was for 30 min with avidin biotin enzyme reagent. More than 300 cells were counted and scored for percent immunopositivity as previously described (26).

Morphological analysis of liver tissues. The genotyping of cyclin D1-/- mice was conducted as previously described (1). Morphological analysis of hepatic cells was determined using the methyl-green pyronin method for overall histologic study, which detects DNA and RNA in the nucleus and cytoplasm (28, 36); Oil Red-O method for staining of neutral lipids normally seen in Ito cells (28, 35); an acid phosphatase lead enzyme method using CMP as the substrate for Kupffer cells and lysosomes in hepatic cells (e.g., hepatocytes) (36); and a diaminobenzidine method at pH 9.7 for the presence of catalase in peroxisomes and microperoxisomes (34, 35). Livers from wt and knockout mice were fixed in a fixative containing cold 4% paraformaldehyde, 2% glutaraldehyde, cacodylate buffer (pH 7.4) for 3 to 5 h (22). Sections of livers were prepared on a freezing microtome after cryoprotection in increasing concentrations of sucrose. The sections were then processed by the above-described methods, after which they were viewed with the light microscope.

Structural modeling of cyclin D1/PPAR{gamma}. The coordinates for the three-dimension model of cyclin D1 were generated based on the sequence alignment between cyclin D1 and cyclin A and the published crystal structures of cyclin A (6). Modeling was performed using the homology-modeling program PROMOD (37). In the homology model of cyclin D1, the region required for PPAR{gamma} repression (from 142 to 178) formed a helix-loop-helix. Within this region a cluster of hydrophobic residues (amino acids [aa] 137 to 148, LLXXXLLLVXXL) was identified. Since this sequence forms a helix, some of the hydrophobic residues were exposed to solvent. We therefore modeled this helix binding to the same region (residues 280 to 318 in PPAR{alpha}) that corepressors and coactivators bind to. By using the published X-ray crystal structures of PPAR{alpha}-corepressor (62) and PPAR{gamma}-coactivator (61) complexes and the homology model of cyclin D1, a three-dimensional model of the complex between PPAR{gamma} and cyclin D1 was generated using the molecular modeling program INSIGHT (Accelrys Inc., San Diego, Calif.). The coordinates for the whole complex were subjected to energy minimization using the AMBER force field (8).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Cyclin D1 repression of PPAR{gamma} transactivation. We examined the role of cyclin D1 in regulating PPAR{gamma}-dependent gene activity. Cyclin D1-deficient (cyclin D1-/-) 3T3 cells, generated from cyclin D1-/- mice (3a), were transfected with the wt PPAR{gamma} and a reporter gene encoding multimeric sequences of a PPAR{gamma}-response element from the acyl coenzyme A oxidase gene promoter, (AOX)3LUC. In the presence of a PPAR{gamma} expression plasmid, the specific ligands, BRL49653 and Troglitazone, induced (AOX)3LUC three- to fivefold. Coexpression of cyclin D1 wt in cyclin D1-/- 3T3 cells repressed ligand-induced (AOX)3LUC reporter activity by 40% (Fig. 1B). In order to identify the mechanisms by which cyclin D1 repressed liganded PPAR{gamma}-dependent gene transcription, point mutations of cyclin D1 were assessed (Fig. 1B to D). Each of the mutants was expressed as well as the wt in cultured cells (Fig. 1D). The cyclin D1 KE mutant is defective in binding cdks, the GH mutant is defective in pRB binding, and the 254/255 mutant is defective in binding the p160 coactivators (66). Each of these cyclin D1 mutants inhibited liganded PPAR{gamma} reporter activity to the same level as cyclin D1 wt (Fig. 1B). The T286A mutant, which evades GSK3-ß phosphorylation in vitro and remains nuclear throughout the cell cycle in cultured cells (4), repressed PPAR{gamma} activity to a level similar to that with cyclin D1 wt. Sequential C-terminal deletion of cyclin D1 to N178 or amino-terminal deletion of the cdk- and pRB-binding regions maintained repression. N-terminal deletion from 143 to 179 (C4) abolished repression, indicating a requirement for residues between 143 and 179 for full repression (Fig. 1C). Together these studies suggest that cyclin D1 inhibits liganded PPAR{gamma} activity through a cdk-independent mechanism that requires the region of cyclin D1 between residues 143 and 179. To assess the effect of cyclin D1 on expression levels of PPAR{gamma} from the viral expression plasmid, Western blotting was conducted with 293 cells either transfected with pCMV-PPAR{gamma} alone or in the presence of cotransfected cyclin D1 expression vector. The relative abundance of PPAR{gamma} from the expression plasmid, assessed by the Flag epitope of PPAR{gamma}, was not affected by cyclin D1 (Fig. 1E).



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  		FIG. 1. The region of cyclin D1 required for repression of PPAR{gamma} activity. (A) Cyclin D1-/- 3T3 cells were transfected with reporter plasmid (AOX)3TK LUC carrying response elements specific for PPAR{gamma} (1 µg). prl-TK LUC was cotransfected as an internal control. Cells were cotransfected with the expression vector encoding human PPAR{gamma} in either the presence or absence of cyclin D1 (pCMV-cyclin D1) or (B and C) expression vectors for cyclin D1 mutants. The effect of the coexpressed cyclin D1 construction on ligand-induced luciferase reporter activity is shown as mean ± standard error of the mean. Activity of the parental prl-TK Luc reporter, which lacks the (AOX)3 element, was not affected by cyclin D1. (D) Western blotting for the Flag epitope of the cyclin D1 constructs. (E). Western blot of cells transfected with the pCMV-cyclin D1 and PPAR{gamma} expression vector.

 
We examined further the role of cyclin D1 in regulating PPAR{gamma}-dependent gene activity using HeLa cells as previously described (54), transfecting cells with the wt PPAR{gamma} and a reporter gene (AOX)3LUC (Fig. 2A). Addition of 15d-PGJ2, which is also capable of activating PPAR{gamma}, induced reporter activity 3.5-fold. Coexpression of cyclin D1 inhibited 15d-PGJ2-induced PPAR{gamma}-dependent reporter activity by 60% (Fig. 2A). Cyclin D1 repressed ligand-induced (AOX)3LUC reporter activity at several different concentrations of vector (Fig. 2A) and ligand (1, 2, 5, 5, and 10 µm) (data not shown). Transfection of cells with a tetracycline-regulated expression vector for cyclin D1 repressed 15d-PGJ2-induced (AOX)3LUC reporter activity by 60% (Fig. 2B). 15d-PGJ2-induced (AOX)3LUC reporter activity was unaffected by either cyclin D2 or cyclin D3. The activities of the viral promoters (CMV-LUC and RSV-LUC) and the cyclin E, c-Jun, and JunB luciferase reporter vectors were not regulated by cyclin D1 (39; also data not shown). We next assessed the PPAR{gamma}1 promoter, since it is a PPAR{gamma}-responsive reporter gene (15). PPAR{gamma}1 promoter activity was twofold more active in cyclin D1-/- 3T3 compared with wt 3T3 cells, normalized for transfection efficiency. The PPAR{gamma}1 promoter was repressed two- to threefold by cyclin D1 overexpression in cyclin D1-/- 3T3 cells (Fig. 2C). The relative abundance of PPAR{gamma}1 mRNA was assessed by semiquantitative RT-PCR using liver cell extracts from either cyclin D1 wt or cyclin D1-/- mice. The relative abundance of PPAR{gamma}1 mRNA was increased twofold in the cyclin D1-/- mice (Fig. 2D). To determine whether cyclin D1 was capable of inhibiting the transactivation domain of PPAR{gamma}, a heterologous reporter system was used. To assess DNA-binding independent PPAR{gamma} activity, the PPAR{gamma} LBD linked to the Gal4 DNA binding domain was used with a reporter construct consisting of a multimeric DNA binding site for the Gal4 DNA binding domain linked to the luciferase reporter gene. The heterologous reporter system was introduced into cyclin D1-deficient (cyclin D1-/-) 3T3 cells, and luciferase activity was normalized to that of a cotransfected Renilla luciferase reporter gene. Cotransfection of the expression vector for cyclin D1 into cyclin D1-/- 3T3 cells inhibited PPAR{gamma}-Gal4 activity 50% (Fig. 2E). As with repression of liganded (AOX)3LUC reporter activity, the region from 143 to 179 was required for repression of PPAR{gamma}-Gal4 activity.



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  		FIG.2. Cyclin D1 inhibits liganded PPAR{gamma} transactivation function. (A) The (AOX)3 luciferase reporter (1 µg) was transfected into HeLa cells with the expression vector encoding the human PPAR{gamma} in either the presence or absence of cyclin D1 (pCMV-cyclin D1). Comparison was made with the effect of the expression of equal amounts of empty expression vector cassette (pRC/CMV). 15d-PGJ2 (10 µM) was added as indicated. The results are shown as mean ± standard error of the mean throughout. (B) (AOX)3LUC reporter activity in HeLa cells transfected with tetracycline-inducible vector pcz-cyclin D1. (C). PPAR{gamma}1 promoter activity in cyclin D1-/- or cyclin D1+/+ 3T3 cells cotransfected with pCMV-cyclin D1 or control vector as indicated. (E) Semiquantitative RT-PCR for PPAR{gamma}1 from mRNA of livers of cyclin D1-/- or cyclin D1+/+ mice. The PPAR{gamma} LBD construct linked to the Gal4 DNA binding domain was assessed for activity using the heterologous reporter (UAS)5E1BTATA LUC in the presence or absence of the expression vectors for cyclin D1 in cyclin D1-/- 3T3 cells. The relative transactivation level was shown as luciferase activity represented by light units measured in cells cotransfected with a specific receptor expression plasmid. Reporter gene activity was normalized to prl-TK LUC activity.

 
PPAR{gamma} expression is reduced in human benign breast disease and cancers correlating with increased cyclin D1 abundance. We determined PPAR{gamma} abundance in murine mammary tumors, reasoning that expression of cyclin D1 and PPAR{gamma} may be reciprocal. In mammary tumors induced by mammary tissue-targeted oncogenes (ErbB2, Ras, and Src), the relative abundance of cyclin D1 was increased and that of PPAR{gamma} was decreased compared with results for normal mammary epithelium (Fig. 3A and B). Cyclin D1 abundance was also increased in the mammary tumor compared with that in the adjacent mammary epithelium in the same animal, with reciprocal changes in PPAR{gamma} expression (Fig. 3A and B).



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  		FIG. 3. PPAR{gamma} and cyclin D1 abundance in mammary tumors induced by distinct oncogenes and human benign and malignant breast disease. (A) Expression of PPAR{gamma} and cyclin D1 in murine mammary tumors. Western blotting of normal mammary epithelium or mammary tumors of MMTV-directed oncogene mice (MMTV-Ras [Ras], MMTV-Src [Src], or MTV-ErbB2 [ErbB2]) is shown. Abundance of PPAR{gamma}, cyclin D1, or the loading control GDI is shown. MMTV-ErbB2 mammary tumors (T) and adjacent normal mammary epithelium (N) are represented below. (B). The mean levels of cyclin D1 and PPAR{gamma} protein are shown for n > 7 tumors induced by each oncogene, compared with normal mammary epithelium. (C). Representative immunostaining of normal mammary epithelium, benign breast disease, and ER{alpha}+ infiltrating breast cancers (n = 40) (magnification, x388). (D). The percent immunopositive staining for PPAR{gamma} and cyclin D1 is shown for premalignant lesions (n = 36) with comparison to normal mammary epithelium.

 
Immunohistochemical assessment was made of cyclin D1 and PPAR{gamma} in normal breast epithelium, benign breast lesions, and infiltrating ER{alpha}+ breast cancers (Fig. 3C). PPAR{gamma} was detectable in normal mammary epithelium as previously reported (32). Preimmune sera showed no nuclear staining of breast epithelial cells (not shown). The majority of human breast cancers were cyclin D1 immunopositive and PPAR{gamma} negative. The samples of human benign disease included a spectrum of histopathological subtypes (nonproliferative, proliferative [epithelial cell hyperplasia with or without atypia, sclerosing adenosis, papilloma, and fibroadenoma]). The benign breast lesions displayed increased cyclin D1 immunopositivity and reduced PPAR{gamma} staining compared with normal mammary epithelium (Fig. 3D). Thus, in human ER{alpha}-positive breast tumors, reduced PPAR{gamma} abundance is found in conjunction with increased cyclin D1 levels.

Inhibition of PPAR{gamma} ligand-induced differentiation by cyclin D1. Although cyclin D1 inhibited PPAR{gamma}-responsive reporter gene activity, it was important to determine whether cyclin D1 inhibited the in vivo function of liganded PPAR{gamma}. Liganded PPAR{gamma} is both necessary and sufficient for the induction of Oil Red-O positive staining as a marker of adipocyte phenotype in immortalized 3T3 cells (40). We compared PPAR{gamma} ligand-regulated adipogenesis in cyclin D1+/+ versus cyclin D1-/- MEFs prior to immortalization. 3T3-L1 adipogenesis medium was not capable of inducing MEF differentiation into adipocytes. Addition of the PPAR{gamma} agonists rosiglitazone (0.2 µM) or troglitazone (5 µM) induced a modest (8% positive) but significant lipid droplet accumulation in cyclin D1+/+ wt MEFs. In contrast, more than 35% of the cyclin D1-deficient MEFs showed Oil Red-O positive staining (Fig. 4A and C).



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  		FIG. 4. Cyclin D1 deficiency enhances PPAR{gamma} function. (A) MEFs derived from either cyclin D1+/+ or cyclin D1-/- mice were treated with vehicle, BRL-49653 (1 µM), and troglitazone (5 µM), and lipid accumulation was assessed. cyclin D1-/- MEFs exhibit lipid accumulation by OilRed-O staining in the presence of BRL-49653 or troglitazone, shown as mean ± standard error of the mean for three experiments in panel C. (B) cyclin D1-/- MEFs were infected with either a parental retrovirus or retroviruses encoding cyclin D1. Two days postconfluence, cells were treated with differentiation media and troglitazone (5 µM). Seven days later, cells were stained with Oil Red-O to visualize the degree of lipid accumulation. The results were quantitated for two separate experiments (D). Representative infection of cyclin D1-/- MEFs with viral GFP expression construct is shown. Tro, troglitazone; DMSO, dimethyl sulfoxide.

 
These findings suggest that the deficiency of cyclin D1 or factors regulated by cyclin D1 contributes to the adipocyte phenotype induced by the specific PPAR{gamma} ligands. To determine whether cyclin D1 deficiency of the cyclin D1-/- MEFs was the key regulator of the PPAR{gamma} ligand-induced adipocyte phenotype, cyclin D1-/- MEFs were infected with a retroviruses encoding cyclin D1 prior to the differentiation protocol. Expression of cyclin D1 inhibited rosiglitazone (0.2 µM)-induced or troglitazone (5 µM)-induced lipid accumulation in the cyclin D1-/- MEFs (Fig. 4B and D). The transduction efficiency of the MEFs was >90%, assessed by transduction with the viral vector encoding GFP (Fig. 4E). These studies provide genetic evidence that cyclin D1 is necessary and sufficient to regulate the adipogenic differentiation function of PPAR{gamma}.

Reduction of cyclin D1 by ponasterone-regulated cyclin D1 antisense transgenic mice induces PPAR{gamma} abundance. The correlative reciprocal expression profile of cyclin D1 and PPAR{gamma} in mammary epithelium was assessed further to determine whether cyclin D1 inhibited PPAR{gamma} expression. PPAR{gamma} levels were increased in untreated cyclin D1-/- 3T3 cells (Fig. 5A) and cyclin D1-/- MEFs (Fig. 5B), suggesting that cyclin D1 may inhibit PPAR{gamma} expression, consistent with our finding that cyclin D1 repressed the PPAR{gamma}1 promoter. To examine further whether cyclin D1 inhibited PPAR{gamma} abundance in vivo, we generated transgenic mice in which the murine cyclin D1 antisense cDNA linked to a GFP transgene was regulated under control of the ecdysone enhancer (Fig. 5Ca). We had previously used ponasterone-inducible cyclin D1 antisense to reduce cyclin D1 abundance in cultured cells (63). Transgene transmission was confirmed by genomic Southern blotting (Fig. 5Cb). To induce expression of the transgene, an ecdysone receptor (Bbyx), adenoviruses were used that expresses GFP from a second cistron (DB-Ecr-IRES-GFP, Fig. 5Cc). The entire experiment was conducted on two separate occasions, and representative results are shown. Comparison was made with equal titer of an adenovirus for the ecdysone receptor that does not express GFP.



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  		FIG. 5. Transgenic inducible cyclin D1 antisense regulates PPAR{gamma} abundance in vivo. Western blot of cyclin D1-/- 3T3 (A) and cyclin D1-/- MEFs (B) with antibodies as indicated. (Ca) An ecdysone enhancer-driven cyclin D1 antisense transgene (schematic shown), was used to generate transgenic mice (Tg) confirmed by genomic Southern blot (Cb). (Cc) Adenoviral vectors encoding receptors for the Bombyx LBD either with or without a second cistron for GFP were used to infect the cyclin D1 antisense transgenic mice (Cd). (Ce) Transgenic mice were treated with ponasterone to induce transgene expression, and Western blotting of hepatic extracts was performed at 5 days for the loading control (GDI), cyclin D1 abundance and GFP from the second cistron of the cyclin D1 antisense, and PPAR{gamma}.

 
The adenoviral receptors encoding the ecdysone receptor were introduced into transgenic cyclin D1 antisense mice or control strain-matched mice, and activity of the receptors was induced using ponasterone pellets (Fig. 5Cd). Serum samples demonstrated the induction of ecdysone enhancer-driven SEAP or luciferase activity in parallel experiments (data not shown). Analysis was performed on the livers of transgenic and antisense transgenic mice. PPAR{gamma} is expressed in hepatocytes, and the expression of PPAR{gamma} is induced by TZDs in vivo (13). It has been hypothesized that the insulin-sensitizing effects of TZDs may result in part from effects on PPAR{gamma} expression and function in muscle and liver (12, 13). The induction of the GFP from the second cistron of the cyclin D1 antisense transgene was observed by Western blotting (Fig. 5Ce). Analysis of hepatic tissue evidenced the presence of GFP by immunofluorescence and by Western blotting with mice transduced with the DB-Ecr-IRES-GFP adenovirus, used as a positive control for GFP in parallel experiments (data not shown). The abundance of cyclin D1 protein was reduced by 90% for the cyclin D1 antisense transgenic mice compared with results for control mice (n = 2). The reduction in cyclin D1 protein levels correlated with the induction in PPAR{gamma} abundance in vivo (Fig. 5Ce), providing further evidence that cyclin D1 inhibits PPAR{gamma} expression in vivo.

Cyclin D1 inhibits PPAR{gamma} ligand-induced gene expression without affecting C/EBP{alpha} or C/EBPß. The expression of genes known to be either upstream (C/EBPß) (60) or downstream (adipocyte complement related protein of 30 kDa [ACRP30], C3/12) of PPAR{gamma} in the adipocyte differentiation program was next assessed. C/EBP{alpha}-mediated adipocyte differentiation requires PPAR{gamma}; however, PPAR{gamma}-mediated differentiation is independent of CEBP{alpha}, placing PPAR{gamma} as a downstream effector of CEBP{alpha} (40). Analysis was performed during differentiation of cyclin D1+/+ and cyclin D1-/- MEFs. Normalization of protein loading was performed using GDI (25). Cells were harvested every day for 7 days and lysed, and the cell lysates were subjected to Western blotting for ACRP30 and C3/12. Prior to the induction of differentiation by the PPAR{gamma} ligands BRL49653 (Fig. 6) and troglitazone (data not shown), cyclin D1 was detectable in the cyclin D1+/+ but not the cyclin D1-/- MEFs. PPAR{gamma} levels were increased two- to threefold in the cyclin D1-/- MEFs compared with results in the cyclin D1 +/+ MEFs (Fig. 6), consistent with the finding that cyclin D1 inhibits both PPAR{gamma} expression and transactivation. In the undifferentiated state, cyclin D1+/+ and cyclin D1-/- MEFs expressed similar levels of C/EBP{alpha} and C/EBPß.



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  		FIG. 6. Enhanced PPAR{gamma}-responsive gene expression in cyclin D1-/- cells. (A) MEFs derived from either cyclin D1+/+ or cyclin D1-/- mice were treated with BRL-49653 (1 µM). Western blotting was conducted for PPAR{gamma} and the PPAR{gamma}-responsive genes, ACRP30 (11) and S3/12. Levels of cyclin D1, C/EBPß, C/EBP{alpha}, and the loading control (GDI) are shown. (B) Western blot analysis of cyclin D1-/- MEFs or cyclin D1-/- MEFs infected with a cyclin D1 retrovirus. MEFs were treated for the time points indicated.

 
ACRP30 (adiponectin/apM1/AdipoQ/GBP28) is an adipocyte-secreted protein induced during fat cell differentiation (5) induced by the PPAR{gamma} ligand TZD (13). At day 6, the levels of the adipocyte differentiation markers downstream of PPAR{gamma} (ACRP30 and S3/12) were substantially higher in the cyclin D1-/- MEFs than in the cyclin D1+/+ MEFs. The levels of ACRP30 was >100-fold higher in the cyclin D1-/- MEFs than in the wt MEFs. In both wt and cyclin D1-/- MEFs, the levels of ACRP30 were increased 10-fold by BRL49653 or troglitazone. Upon induction of differentiation, both cyclin D1+/+ and cyclin D1-/- MEFs showed a similar pattern of C/EBP{alpha} and C/EBPß expression. These studies suggest that inhibition of adipocyte differentiation by cyclin D1 occurs distal to C/EBPs and involves inhibition of PPAR{gamma} function. Transduction of cyclin D1-/- MEFs with a cyclin D1 retrovirus inhibited the induction of differentiation by BRL49653 and reduced the time-dependent induction of ACRP30 (Fig. 6B).

Adipogenic steatosis in cyclin D1-/- mouse liver. In previous studies, adenoviral delivery of PPAR{gamma}1 overexpression in the murine liver induced hepatic steatosis (65). If repression of PPAR{gamma}1 activity by cyclin D1 was functionally significant in vivo, it would be predicted that the cyclin D1-/- mice would demonstrate features of hepatic steatosis. Detailed morphometric and histological analysis was therefore conducted of the livers from cyclin D1-/- and wt littermate controls. Acid phosphatase cytochemistry revealed that the Kupffer cells in the cyclin D1-/- mice were enlarged, appeared increased in numbers (Fig. 7), and developed an extensive lysosomal system filled with lipid deposits (Fig. 7). In addition, acid phosphatase cytochemistry revealed that the lysosomes of hepatocytes appeared larger and appeared to be distributed more widely in the cytoplasm in contrast to the wt hepatocytes lysosomes, which were more concentrated near the bile canaliculus. Neutral lipid staining using Oil Red-O showed a substantial increase in neutral lipid droplets in the cyclin D1-/-. Ito cells from cyclin D1-/- mice were increased in numbers and enlarged, with more numerous and larger lipid spheres, compared with Ito cells of wt littermate controls. Changes were also found in hepatocytes and included increased numbers of microperoxisomes, and cytoplasmic lipid spheres were noted (Fig. 7). Thus, the livers of cyclin D1-/- mice display the features of hepatic steatosis consistent with increased PPAR{gamma} activity.



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  		FIG. 7. Morphological analysis of hepatic cells from cyclin D1 wt and cyclin D1-/- mice. Hepatic tissues were stained for catalase, CMP, methyl-green pyronin (MPG), and Oil Red-O for staining of neutral lipids as described in Materials and Methods.

 
Three-dimensional modeling of the cyclin D1 143-179 region. In view of the importance of the region of cyclin D1 from 143 to 179 in repression of PPAR{gamma} transactivation, we determined the predicted structure of this domain using the homology modeling program PROMOD (37) (see Materials and Methods). The coordinates for the three-dimensional model of cyclin D1 were generated based on the sequence alignment between cyclin D1 and cyclin A and the published crystal structures of cyclin A (6). We generated models from the published crystal structure of PPAR{alpha} corepressor (62) (Fig. 8A, left) and PPAR{gamma}-coactivator (61) (Fig. 8A, right). The coactivator is shown in yellow. PPAR{gamma} is shown in grey with the region of PPAR{gamma} contacting the coactivator shown in green. In the homology model of cyclin D1, the region from 142 to 179 forms a helix-loop-helix, shown in Fig. 8B in yellow and red. The cluster of hydrophobic residues (aa 137 to 148, LLXXXLLLVXXL) was then modeled binding to the same region where corepressors and coactivators bind to PPAR{gamma} (61). Using these two PPAR structures and the homology model of cyclin D1, a three-dimensional model of the complex between PPAR{gamma} and cyclin D1 was generated (Fig. 8C) (INSIGHT; Accelrys Inc., San Diego, Calif.). The hydrophobic cluster region of the cyclin D1 HLH is shown in red in Fig. 8C.



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  		FIG. 8. Prediction model of cyclin D1 helix-loop-helix region binding to PPAR{gamma}. (A) Ribbon model of left panel: nuclear corepressor (red) binding to PPAR{alpha} (grey, with helix 12 in magenta and coactivator binding region of PPAR{alpha} in green); right panel, nuclear receptor coactivator (yellow) binding to PPAR{gamma} (grey, with helix 12 in magenta and coactivator binding region of PPAR{gamma} in green). (B). Ribbon model of PPAR{gamma} and cyclin D1 helix-loop-helix region. Predicted cyclin D1 helix-loop-helix structure is in yellow with component of hydrophobic cluster in red. (C). PPAR{gamma} (grey, with helix 12 in magenta and coactivator binding region of PPAR{gamma} in green) and cyclin D1 hydrophobic cluster of helix-loop-helix motif colored red. (The position alignment, determined by energy minimization using the AMBER force field, displays the cyclin D1 helix-loop-helix motif in an alignment similar to that of a nuclear receptor corepressor).

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present studies demonstrate for the first time in vivo antagonism between a collaborative oncogene, cyclin D1, and a nuclear receptor, PPAR{gamma}. Cyclin D1 inhibited PPAR{gamma}-dependent reporter activity and repressed the trans activity of PPAR{gamma} when linked to a heterologous DNA binding domain. Consistent with previous findings that PPAR{gamma} expression is induced by PPAR{gamma} (12), cyclin D1 also inhibited PPAR{gamma} expression and promoter activity. The reduction in cyclin D1 abundance in cyclin D1-/- 3T3 cells, cyclin D1-/- MEFs, and cyclin D1 antisense transgenic mice correlated with the induction of PPAR{gamma} expression (Fig. 5). The induction of adipocyte differentiation by PPAR{gamma}-specific ligands was substantially enhanced in cyclin D1-deficient cells. The reintroduction of cyclin D1 into cyclin D1-deficient cells abolished the adipogenic phenotype, consistent with a key role for cyclin D1 as an inhibitor of PPAR{gamma}-specific functional activity. Since cyclin D1 abundance is regulated by diverse oncogenic and mitogenic stimuli, the inhibition of PPAR{gamma} transactivation by cyclin D1 may have important implications for signal transduction and tumorigenesis.

Repression of PPAR{gamma} transactivation by cyclin D1 was independent of the cdk and pRb binding functions and required a C-terminal region of cyclin D1 that is predicted to form a helix-loop-helix structure. The mechanism by which cyclin D1 regulated PPAR{gamma} activity is thus distinct from that regulating the ER{alpha} through recruiting a p160 coactivator SRC-1 in vitro (66), as an SRC-1 binding point mutation of cyclin D1 maintained wt repression of liganded PPAR{gamma} activity. Herein, genetic deletion of cyclin D1 enhanced PPAR{gamma} ligand-mediated differentiation of MEFs into adipocytes. Although interactions between cyclin D1 and nuclear receptors have been previously described, the present studies provide strong genetic evidence for a functional interaction between cyclin D1 and a nuclear receptor. Ectopic expression of PPAR{gamma} induced differentiation of NIH-3T3 fibroblast cells into fat-laden adipocyte cells (51). In the present studies, the cyclin D1-/- MEF adipogenic phenotype induced by PPAR{gamma} ligands was reversed by cyclin D1 overexpression. ACRP30 was increased 50-fold in the cyclin D1-/- MEFs, consistent with the enhanced induction of the adipogenic phenotype. The relatively modest increase in PPAR{gamma} abundance, together with the dramatic enhancement of PPAR{gamma} activity in the cyclin D1-/- MEFs, is consistent with the reporter gene studies in which cyclin D1 inhibited PPAR{gamma} trans activity.

These studies raise the possibility that reduced PPAR{gamma} expression, together with increased cyclin D1, may be a genetic feature of the transition from normal breast epithelium, to benign breast disease and adenocarcinoma. PPAR{gamma} immunopositivity was decreased in benign breast disease compared with normal mammary epithelium and was reduced further in adenocarcinomas. Cyclin D1 immunopositivity increased from normal epithelium to benign disease and adenocarcinomas. The reduction in PPAR{gamma} expression in the cyclin D1-infected MEFs, together with the finding of increased levels of PPAR{gamma} mRNA and protein in cyclin D1-/- livers by microarray (not shown) and Western blotting, suggest that cyclin D1 inhibits PPAR{gamma} expression. The overexpression of cyclin D1 with ER{alpha} reflects poor prognosis in human breast cancer. Given the repression of PPAR{gamma} function and expression by cyclin D1 and the cytoinhibitory role of PPAR{gamma} in breast epithelium, these studies raise the question of whether reduced PPAR{gamma} may contribute to poor prognosis in a subset of patients. The reduction in PPAR{gamma} staining in proliferative breast disease suggests that further studies of PPAR{gamma} as a prognostic indicator and candidate target for prevention or therapy of human breast cancer warrants consideration.

The present studies are important in demonstrating a functional antagonism between a collaborative oncogene, cyclin D1, and a candidate tumor suppressor, PPAR{gamma}. Several lines of evidence suggest that PPAR{gamma} may function as a tumor suppressor (42). Consistent with a role for PPAR{gamma} as an inhibitor of tumorigenesis, heterozygous mutations of PPAR{gamma} were detected in 4 of 55 patients with colon cancer (45). In follicular thyroid cancer, a fusion oncoprotein has been described, formed by a chromosomal translocation between PPAR{gamma}1 and PAX8 with a deletion in its C-terminal activation domain. The PPAR{gamma} fusion protein functioned as a powerful dominant-negative of wt PPAR{gamma} and was not observed in benign follicular adenomas (24). The addition of PPAR{gamma} ligands (TZD or 15d-PGJ2) inhibited breast and colonic cellular proliferation (7, 14, 32, 52). In contrast, cyclin D1 abundance is induced by diverse oncogenic and mitogenic signals in breast and colonic epithelial cells and functions as a collaborative oncogene (18, 38). Cyclin D1 antisense inhibits the growth of murine mammary tumors derived from MMTV-ErbB2 mice (25), and cyclin D1-/- mice are resistant to the induction of tumor formation by ErbB2 (64). Since PPAR{gamma} inhibits the expression of several genes promoting tumor invasion (those encoding iNOS, gelatinase B, matrix metalloproteinases [10, 21, 30], and UPA) (data not shown), cyclin D1 antagonism of PPAR{gamma} function may enhance expression of tumor invasion genes.

The functional antagonism between PPAR{gamma} and cyclin D1 may also have implications for signal transduction cross talk. Cyclin D1 is induced by diverse mitogenic signaling pathways, including those of Src, Rac mutants, Dbl proteins, and ß-catenin, and the NF-{kappa}B signaling pathway (1, 2, 17, 26, 49, 55, 56, 58, 59). PPAR{gamma} activity is also induced by a large number of synthetic and natural ligands, including prostaglandins and fatty acids (42, 50). The inhibition of PPAR{gamma} function by cyclin D1 may contribute to altered metabolism and altered inflammatory responses and remains to be further explored.


   ACKNOWLEDGMENTS
 
We thank R. Evans, V. Fantl, C. Glass, P. Sicinski, and M. Lazar for reagents and helpful discussion. We thank M. Caparas for assistance with the manuscript.

This work was supported in part by awards from the Susan Komen Breast Cancer Foundation, Breast Cancer Alliance Inc., R01CA70896, R01CA75503, R01CA86072, R01CA86071 (R.G.P.), R03AG20337 (C.A.), NIH CA06576 (P.N.), and R01DK55758 (P.E.S.). R.G.P was a recipient of the Irma T. Hirschl and Weil Caulier award and was the Diane Belfer Faculty Scholar in Cancer Research. M.D. was a recipient of the New York State mentored EMPIRE award. Work conducted at the Lombardi Cancer Center was supported by the NIH Cancer Center Core grant.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Oncology, Lombardi Cancer Center, Research Building, Room E501, 3970 Reservoir Rd. NW, Box 571468, Georgetown University, Washington, DC 20007. Phone: (202) 687-2110. Fax: (202) 687-6402. E-mail: pestell{at}georgetown.edu. Back


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