In addition to their role in cell cycle progression, new data reveal an emerging role of D-type cyclins in transcriptional regulation and cellular differentiation processes. Using 3T3-L1 cell lines to study adipogenesis, we observed an up-regulation of cyclin D3 expression throughout the differentiation process. Surprisingly, cyclin D3 was only minimally expressed during the initial stages of adipogenesis, when mitotic division is prevalent. This seemingly paradoxical expression led us to investigate a potential cell cycle-independent role for cyclin D3 during adipogenesis. We show here a direct interaction between cyclin D3 and the nuclear receptor peroxisome proliferator-activated receptor γ (PPARγ). Our experiments reveal cyclin D3 acts as a ligand-dependent PPARγ coactivator, which, together with its cyclin-dependent kinase partner, phosphorylates the A-B domain of the nuclear receptor. Overexpression and knockdown studies with cyclin D3 had marked effects on PPARγ activity and subsequently on adipogenesis. Chromatin immunoprecipitation assays confirm the participation of cyclin D3 in the regulation of PPARγ target genes. We show that cyclin D3 mutant mice are protected from diet-induced obesity, display smaller adipocytes, have reduced adipogenic gene expression, and are insulin sensitive. Our results indicate that cyclin D3 is an important factor governing adipogenesis and obesity.
Our understanding of the molecular mechanisms that orchestrate adipocyte differentiation have been greatly advanced by the use of preadipocyte cell lines, such as 3T3-L1 cells capable of undergoing adipogenesis (14). Upon reaching confluence, proliferating preadipocytes become growth arrested by contact inhibition. These growth-arrested preadipocytes reenter the cell cycle after hormonal induction, arrest proliferation again, and finally undergo terminal adipocyte differentiation. Peroxisome proliferator-activated receptor γ (PPARγ), a ligand-inducible transcription factor, has been identified as a major regulator of terminal adipocyte differentiation (10, 37). PPARγ, upon activation by either fatty acid derivatives or antidiabetic thiazolidinediones, drives the expression of several adipocyte-specific genes, such as the fatty acid binding protein aP2, transforming the cell into the characteristic lipid-rich adipocyte (42). Subsequent studies have demonstrated that ectopic expression of PPARγ further induces adipocyte differentiation (43). This pivotal role of PPARγ in adipocyte differentiation is also highlighted by the phenotype observed in humans with mutations in the PPARγ gene and by PPARγ-deficient mice, which are essentially void of white adipose tissue (8).
D-type cyclins were first characterized for their ability to coordinate cell cycle progression through the G1 phase. Three D cyclins (cyclins D1, D2, and D3) bind and activate cyclin-dependent kinases 4 and 6 (CDKs 4 and 6), directing the phosphorylation of retinoblastoma protein, as well as retinoblastoma protein-related proteins p107 and p130 (4, 18, 28). This phosphorylation event disrupts the retinoblastoma protein repressor complexes, leading to derepression of E2F transcription factors and induction of E2F target genes, which are required for S-phase entry (6).
In addition to their defined role as part of the core cell cycle machinery, a new potential for D cyclins has emerged in other cellular processes, including transcriptional control and differentiation. Cyclin D1 can bind and repress the activity of several transcription factors, including b-Myb (15), MyoD (34, 40), and DMP1 (17). Although less well explored, a CDK-independent role for cyclin D3 has also been reported, including inhibition of granulocyte differentiation (19). More recent studies have attributed cyclin D3 with the ability to bind and activate certain transcription factors, such as human activating transcription factor 5 (25). In the case of cyclin D3 mutant mice it has been found that they fail to undergo development of immature T lymphocytes (39).
Recently, our laboratory explored a link between the molecular processes governing adipocyte differentiation and the molecular machinery involved in cell cycle progression. These studies have established key cell cycle regulators including the retinoblastoma protein and the E2F transcription factor family as fundamental regulators of adipogenesis through their modulation of PPARγ expression and activity (9, 11). Other recent studies have linked loss of cyclin-dependent kinase inhibitors with obesity in mice (30).
The notion that adipogenesis is regulated by proteins of the cell cycle is not unexpected since early stages of 3T3-L1 adipogenesis (days 1 to 2) are marked by active rounds of mitotic clonal expansion. An active cell cycle during the initial stages of adipogenesis is considered a prerequisite for terminal adipocyte differentiation (days 3 to 6) since CDK and MEK-1 (mitogen-activated protein kinase 1) inhibitors, which prevent mitotic clonal expansion, also block the differentiation process (41). Following a few rounds of mitotic division, CDK inhibitors mediate cell cycle exit, which sets the stage for PPARγ-driven terminal adipocyte differentiation (29).
Because D-type cyclins represent a link between cell cycle progression, cell differentiation, and transcriptional regulation, we wanted to explore their potential role during adipogenesis. We show here that cyclin D3 expression is up-regulated during terminal stages of adipogenesis and functions as a ligand-dependent coactivator of PPARγ capable of phosphorylating the A-B domain of the nuclear receptor. Knockdown of cyclin D3 diminished PPARγ activity and adipogenesis, whereas cyclin D3 overexpression had the opposite effect. Consistent with these findings, we show that cyclin D3 null mice are protected from diet-induced obesity, have reduced adipocyte size, and show increased sensitivity to insulin.
Chemical reagents and antibodies.Pioglitazone was provided by Takeda Pharmaceutical Company (Osaka, Japan) and rosiglitazone was purchased from Interchim (Montlucon, France). All chemicals were purchased from Sigma (St. Louis, MO). Cyclin D3 (sc-6283), PPARγ (sc-7273), PPARγ (sc-7196), and actin (sc-1615) antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The anti-bromodeoxyuridine antibody was bought from Dako A/S (Glostrup, Denmark).
Cell culture, transfections, and protein extracts.Cos and 3T3-L1 cells were grown in Dulbecco's modified Eagle's medium with 10% fetal bovine serum. In differentiation studies, 0.5 mM 3 isobutyl-1-methylxanthine, 10 μg/ml insulin, and 1 μM dexamethasone were added for 2 days. From day 3 on, 10 μg/ml insulin and in certain cases 10−6 M pioglitazone were added. Nuclear extracts and Oil Red O staining were prepared as described (35) with the exception that cells were incubated with Oil-Red-O solution for only 90 seconds. For reporter assays, cells were transfected with 10 ng of PPARγ and 300 ng of cyclin D3 expression vectors using Lipofectamine (Life Technologies, Rockville, MD). Luciferase and β-galactosidase activity was measured as described (35). Stable 3T3-L1 cell lines were carried out by transfection of the pcDNA3-cycD3 vector and the control empty vector followed by selection with neomycin (500 μg/ml) for 15 days.
Pull-down, coimmunoprecipitation, and chromatin immunoprecipitation.In vitro translation of pSG5-PPARγ and pcDNA3-cycD3 was done using [35S]methionine (Amersham, Orsay, France) in a TNT coupled reticulocyte lysate (Promega, Madison, WI). Pull-down immunoprecipitation assays were performed as described (9). Chromatin immunoprecipitation assays were performed using three 10 cm plates per point according to the Upstate chromatin immunoprecipitation assay kit (Lake Placid, NY). Oligonucleotides used to amplify the mouse aP2 PPRE were 5′-CCCAGCAGGAATCAGGTAGC-3′ and 5′-AGAGGGCGGAGCAGTTCATC.
RNA isolation, quantitative real-time PCR, and Northern blot.RNA isolation was carried out using the Rneasy minikit (QIAGEN Sciences, Maryland) according to the manufacturer's instructions. Reverse transcription of total RNA was performed at 42°C using Moloney murine leukemia virus reverse transcriptase enzyme and random hexanucleotide primers (Invitrogen, Carlsbad, CA), followed by 15 min inactivation at 70°C. Quantitative PCR was carried out by real-time PCR using a LightCycler and the DNA double-strand-specific SYBR green I dye for detection (Roche, Basel, Switzerland). Results were normalized to glyceraldehyde-3-phosphate dehydrogenase levels. Oligonucleotide sequences used for quantitative real-time PCR are available upon request. A melting temperature of 60°C was used for all of the primers used above. Northern blot analysis was performed as described previously (23).
SiRNA against cyclin D3.To target cyclin D3 expression, short interfering RNA (siRNA) sequences were designed against the 5′-CAGCGGGAGATCAAGCCGCACAT-3′ sequence of the mouse cyclin D3 transcript. Oligonucleotides coding for a hairpin siRNA were cloned into pRNAT-U6.1/Neo vector (GenScript, Piscataway, NJ) as described by the manufacturer. Stable 3T3-L1cell lines expressing the pRNAT-U6.1/Neo αD3 and the pRNAT-U6.1/Neo-control vector were created by Lipofectamine transfection of the vectors followed by selection with neomycin (500 μg/ml) for 14 days.
Western blot analysis and immunofluorescence.Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) and electrotransfer were performed as described (38). The membranes were blocked at room temperature for 45 min in phosphate-buffered saline, 0.5% Tween 20, 5% milk and incubated overnight at 4°C with the indicated antibodies followed by 1 h with a peroxidase-conjugated secondary antibody at room temperature. The complex was visualized with a 4-chloro-1-naphthol reagent. For all immunofluorescence experiments, cells were grown on coverslips and fixed with methanol at 4°C for 10 min. For bromodeoxyuridine incorporation, cells were additionally treated with 1.5 N HCl for 10 min at RT. After incubation with the indicated antibodies, cells were incubated with a combination of Texas red-conjugated anti-mouse immunoglobulin G and fluorescein isothiocyanate-conjugated anti-rabbit immunoglobulin G.
Plasmids and mutant constructs.The pcDNA3 vector was purchased from Stratagene (La Jolla, CA). The pcDNA3-cycD3 expression vector was created by excising the cyclin D3 insert from the pBABE-cycD3 vector (gift from B. Amati) at BamHI and EcoRI restriction sites followed by ligation into the pcDNA3 vector. The thymidine kinase-luciferase (TK-Luc), PPARγ response element (PPRE)-TK-Luc, upstream activation sequence (UAS)-TK-Luc, glutathione S-transferase (GST)-PPARγ DE, GST-PPARγb-AB, GST-PPARγ-AB, Gal4-PPARγ, and PPARγ2 expression vectors have been previously described (13, 35).
The cyclin D3 LXXLL point mutations were performed by PCR of pcDNA3-cycD3 with primers GATCCCTGCCAGGAATTCTGTGAGCTCATC (for the Ct mutant), and GATGAGCTCACAGAATTCCTGGCAAGGGATC (for the Nt mutant). The Δ1-129 deletion mutant was created by PCR amplification of pcDNA3-cycD3 with the following primers: forward, CGGGATCCCGCAAGTGGGACCTGGCTGCTGTGAT, and reverse, CGGAATTCCGGCGGCCCCTCCTCTGCTTAGTGG, containing BamHI and EcoRI restriction sites, respectively. Wild-type cyclin D3, LXXLL point mutants, and Δ1-129 deletion mutants were cloned into the pGex4T1 vector at BamHI and EcoRI restriction sites. The Δ148-293 deletion mutant was created by digesting WTcycD3-pGex4T1 with PpuMI followed by ligation.
Kinase assays.Kinase assays were performed using 100 ng of an active cyclin D3/CDK6 kinase (Upstate, Charlottesville, Virginia) and 250 ng of recombinant PPARγ protein as the substrate (Active Motif, Carlsbad, CA). Reactions were performed in kinase buffer (25 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM MgCl2, 1 mM dithiothreitol) in the presence of 40 mM ATP and 8 mCi [γ-33P]ATP for 30 min at 37°C. The reaction was stopped by boiling the samples for 5 min in the presence of denaturing sample buffer. Samples were then subjected to SDS-PAGE, and the gels were then dried in a gel dryer for 1 h at 80°C and exposed to an X-ray film overnight.
Animal experiments.The cyclin D3 knockout mice were a generous gift from P. Sicinski, by whom their generation has been previously described (39). Animals were maintained according to European Union guidelines for use of laboratory animals. Sections from white adipose tissue were fixed in 4% formaldehyde and stained with hematoxylin and eosin. The intraperitoneal glucose tolerance test and insulin sensitivity tests were performed as described (33). Cyclin D3 mutant and age-matched wild-type mice were fed a lipid-rich diet (58% fat, 25% carbohydrates, and 16% protein) for 8 weeks. All experiments were performed with six age- and gender-matched mice for each group.
Cyclin D3 expression is up-regulated during adipogenesis.When hormonally stimulated, confluent 3T3-L1 preadipocytes reenter the cell cycle before they undergo differentiation (14). We correlated the expression of cyclin E, cyclin D3, and PPARγ during differentiation of 3T3-L1 cells. Protein levels of cyclin E were observed to increase after 1 day of differentiation coincident with cell cycle entry (Fig. 1A). After 2 days of differentiation cyclin E expression drops as differentiating cells exit the cell cycle and the expression of adipogenic markers such as PPARγ is switched on (Fig. 1A). Surprisingly, cyclin D3 protein levels were undetectable during the early stages of differentiation and were strongly induced during later stages after the cells had already exited from the cell cycle and began to express PPARγ. A similar expression pattern was observed by immunofluorescence microscopy (Fig. 1B). Interestingly, we observed that cyclin D3 and PPARγ coexpressed in the same cells.
Expression pattern of cyclin D3 during adipogenesis. (A) Western blot analysis of whole-cell extracts prepared at the indicated days of 3T3-L1 adipocyte differentiation. The proteins detected by cyclin E, cyclin D3, and PPARγ antibodies are indicated. (B) Analysis of cyclin D3 and PPARγ protein expression by immunofluorescence during the first 4 days of 3T3-L1 differentiation. Cells expressing PPARγ are labeled in red (Texas red), whereas cells expressing cyclin D3 are labeled in green (fluorescein isothiocyanate). Nuclei were stained with the Hoechst reagent (blue staining). (C) Northern blot analysis of cyclin D3 mRNA expression during adipocyte differentiation, 28s is used as a loading control. (D) Relative cyclin E, cyclin D3, and PPARγ gene expression measured by quantitative real-time PCR during 3T3-L1 differentiation. (E) Relative expression of cyclin D3 mRNA as measured by quantitative real-time PCR in different mouse tissues (WAT, white adipose tissue; BAT, brown adipose tissue; MUS, muscle; LIV, liver). Results were normalized to the expression of glyceraldehyde-3-phosphate dehydrogenase. (F) Analysis of cyclin D3 protein expression and bromodeoxyuridine by immunofluorescence at day 5 of differentiation. After a 24-h incubation with bromodeoxyuridine, cells, which have incorporated bromodeoxyuridine are labeled in red, while cells expressing cyclin D3 are labeled green, and nuclei are visualized with Hoechst staining.
To obtain further evidence of the expression pattern of cyclin D3 during adipogenesis, we performed Northern blot analysis on differentiating 3T3-L1 cells. Such analysis revealed a potent increase in cyclin D3 mRNA throughout adipogenesis (Fig. 1C). This result was confirmed by quantitative real-time PCR analysis, which revealed a threefold increase in cyclin D3 mRNA throughout adipogenesis (Fig. 1D). Consistent with protein expression (Fig. 1A), mRNA levels for cyclin E increased during days 1 to 2 and decreased thereafter, whereas expression of PPARγ increased strongly from day 1 through the end of differentiation.
To determine the relative expression of cyclin D3 mRNA in vivo, we performed quantitative real-time PCR, comparing cyclin D3 mRNA expression in mouse white adipose tissue, brown adipose tissue, muscle, and liver. Interestingly, we observed strongly elevated cyclin D3 expression in white adipose tissue compared to other tissues (Fig. 1E), suggesting a possible role of this protein in adipose tissue biology. To determine whether cyclin D3 expression during adipogenesis was associated with an active cell cycle, we incubated differentiating 3T3-L1 cells with bromodeoxyuridine to mark proliferating cells. After a 24-hour incubation at day 5 of differentiation, cells were colabeled with bromodeoxyuridine and cyclin D3 antibodies and visualized by fluorescence microscopy. We observed that only a limited percentage of cyclin D3-positive cells had incorporated bromodeoxyuridine (less than 12%) (Fig. 1F). These results suggest a cell cycle-independent role for cyclin D3 during adipogenesis.
Cyclin D3 inhibition impairs adipogenesis.To elucidate the role of cyclin D3 during adipogenesis, we silenced cyclin D3 expression using siRNA techniques. 3T3-L1 cell lines stably expressing a vector coding for a hairpin siRNA sequence against the mouse cyclin D3 transcript or an irrelevant siRNA were compared for their ability to differentiate into adipocytes. After 6 days in differentiation medium, normal lipid accumulation was observed in control cells, whereas a dramatic decrease in lipid accumulation was observed in cyclin D3 knockdown cells as assessed by Oil Red O staining (Fig. 2A and B). Differentiated cyclin D3 knockdown cells expressed significantly reduced levels of cyclin D3 and slightly reduced PPARγ protein compared to control cells (Fig. 2B). Quantitative real-time PCR performed on differentiated 3T3-L1 cells revealed a dramatic reduction of adipogenic gene markers, including adiponectin, aP2, and lipoprotein lipase, and a modest reduction of PPARγ expression when cyclin D3 was knocked down (Fig. 2C), further demonstrating the importance of cyclin D3 in adipogenesis.
Cyclin D3 knockdown inhibits 3T3-L1 adipogenesis. (A) Micrographs of Oil Red O staining of cyclin D3-silenced 3T3-L1 cells (siRNA-cycD3) and control (siRNA-mock) after 6 days of differentiation (top panel). The middle panel confirms knockdown of cyclin D3 by immunofluorescence assay. Nuclei are stained in blue (Hoechst). (B) Western blot showing knockdown of cyclin D3 expression in 3T3-L1 cells expressing a stable vector coding for a siRNA sequence directed against mouse cyclin D3 and corresponding PPARγ protein expression. (C) Quantitative real-time PCR showing gene expression of the adipogenic markers adiponectin, aP2, LPL, and PPARγ in knockdown versus control differentiation experiments. Results are expressed as fold repression compared to the control.
Cyclin D3 overexpression stimulates adipogenesis.Since knockdown of cyclin D3 was inhibiting adipogenesis, we next tested whether overexpressing cyclin D3 could accelerate adipocyte differentiation. For this we created stable cell lines expressing the pcDNA3-cyclin D3 or control vector. After just 4 days in differentiation medium, we observed increased lipid accumulation, increased PPARγ protein expression, and up-regulation of adipogenic mRNA markers as assessed by quantitative real-time PCR in cells overexpressing cyclin D3 compared to the control (Fig. 3A to 3C). Immunofluorescence performed on differentiated 3T3-L1 cells (day 5) overexpressing cyclin D3 revealed a positive correlation between the level of cyclin D3 expression and that of PPARγ in individual cells (Fig. 3D), further suggesting that cyclin D3 is an adipogenic factor.
Cyclin D3 overexpression stimulates 3T3-L1 adipogenesis. (A) Western blot showing cyclin D3 and PPARγ expression in 3T3-L1 cells overexpressing cyclin D3 and control cells. (B) Oil Red O micrographs taken from stable 3T3-L1 cells overexpressing cyclin D3 (pcDNA3-cycD3) and control (pcDNA3-) after 4 days of differentiation. (C) Gene induction change of adipogenic markers as measured by quantitative real-time PCR resulting from cyclin D3 overexpression after 4 days of differentiation. (D) Immunofluorescence showing correlation between PPARγ expression intensity and cyclin D3 in differentiated (day 5) 3T3-L1 cells overexpressing cyclin D3.
Cyclin D3 stimulates PPARγ transcriptional activity.In light of our above observation showing coexpression of cyclin D3 and PPARγ in the same cells (Fig. 3D), we decided to explore a potential functional relationship between the two proteins. To test whether cyclin D3's stimulatory role during adipogenesis may be mediated though activation of PPARγ, we performed cotransfection experiments using a PPARγ-responsive, luciferase-based reporter construct (PPRE-TK-Luc) and expression vectors for PPARγ2 and cyclin D3.
A 3.5-fold induction of luciferase activity was observed upon transfection of PPARγ2 in the presence of the PPARγ agonist pioglitazone (Fig. 4A). This induction was further enhanced up to 5.5-fold by cotransfection of cyclin D3. Transfection of cyclin D3 alone stimulated the PPARγ response element over twofold. No effects of either PPARγ or cyclin D3 were observed on the parental reporter vector TK-Luc. which does not contain a PPRE (Fig. 1A, right panel). No induction of PPRE-TK-Luc was observed after transfecting expression vectors coding for cyclin D1 and D2 (data not shown).
Cyclin D3 stimulates PPARγ transcriptional activity. (A) Activity of the PPRE-TK-Luc reporter carrying the PPARγ specific response elements measured in Cos cells upon transfecting expression vectors for cyclin D3, PPARγ or both plasmids together. The experiments were performed in triplicate in the presence or absence of the PPARγ agonist pioglitazone (10−6M) and were normalized for β-galactosidase activity. The same transfections were performed using a TK-luc vector which does not contain the PPRE. (B) Knockdown of cyclin D3 by transfection of expression vectors coding for siRNA against cyclin D3 abrogates PPARγ transcriptional activity.
We next tested whether the cotransfection of expression vectors coding for siRNA against cyclin D3 could attenuate the PPARγ-mediated luciferase response. Cotransfection of siRNA vector against cyclin D3 reduced PPARγ-mediated luciferase activation by approximately twofold (Fig. 4B). Together, these results suggest that cyclin D3's stimulatory role during adipogenesis is likely the result of its ability to modulate PPARγ activity.
Cyclin D3 physically interacts with PPARγ.To test whether the induction of PPARγ activity in the presence of cyclin D3 is the consequence of an interaction between PPARγ and cyclin D3, nuclear extracts from Cos cells transfected with cyclin D3 and PPARγ expression vectors were immunoprecipitated with an anti-PPARγ antibody. A 33-kDa protein was recognized by a cyclin D3 antibody, indicating that cyclin D3 is associated with PPARγ (Fig. 5A, top panel). We performed the same immunoprecipitation on endogenous PPARγ from differentiated 3T3 L1 cells (day 5) and also revealed an association between the two proteins (Fig. 5A, bottom panel).
Cyclin D3 interacts with PPARγ. (A) Coimmunoprecipitation of PPARγ and cyclin D3 from Cos cells transfected with the expression vectors for both proteins (top panel) and in 5-day differentiated 3T3-L1 cells (bottom panel). Whole-cell extracts were immunoprecipitated with either PPARγ or mock immunoprecipitated (no antibody) and blots were revealed by an anti-cyclin D3 antibody. (B) GST pull-down assay showing details of the PPARγ-cyclin D3 interaction. In vitro-translated 35S-radiolabeled cyclin D3 protein was incubated with GST-PPARγ AB (residues 1 to 146), b-AB (contains an additional 30 amino acids specific for the PPARγ 2 transcript), GST-PPARγ DEF (residues 203 to 477), and GST alone. (C) The same experiment was performed using the DEF domain in the presence and absence of rosiglitazone, 10−6 M. (D) GST pull-down between GST-cyclin D3 and full-length, in vitro-translated 35S-radiolabeled PPARγ protein in the presence and absence of rosiglitazone 10−6M. (E) GST-wild-type cyclin D3 (WTD3), deletion, and point mutants were constructed and tested for their ability to bind full-length recombinant PPARγ. After washing the beads, PPARγ was revealed by Western blot. (F) Chromatin immunoprecipitation assays demonstrating binding of cyclin D3 to the region of the aP2 promoter carrying the PPARγ-responsive element. Cross-linked chromatin from either confluent (top panel) or 5-day-differentiated 3T3-L1 cells (middle panel) was incubated with antibodies against PPARγ, cyclin D3, or nonspecific antibody (mock). Immunoprecipitates were analyzed by PCR using primers specific for the mouse aP2 promoter containing the PPRE and by primers amplifying a region outside the PPRE (negative control). A sample representing 0.1% of total chromatin was included in the PCR (Input). (G) The same experiment was performed on NIH 3T3 cells transfected or not with PPARγ.
To identify the PPARγ domain responsible for the interaction with cyclin D3, GST-PPARγ DEF, AB, and b-AB (where "b" contains an additional 30-amino-acid subunit specific to the PPARγ2 transcript) fusion proteins were incubated with in vitro-translated 35S-radiolabeled cyclin D3. We found that cyclin D3 binds to all three constructs of PPARγ (Fig. 5B).
Next, to see if the association between cyclin D3 and the DEF construct of PPARγ, which contains the ligand binding region of the receptor could depend on ligand, we performed the same pull-down assay in the presence and absence of the PPARγ ligand rosiglitazone. Interestingly, we observed a strong enhancement of the interaction between cyclin D3 and the DEF-PPARγ construct in the presence of rosiglitazone (Fig. 5C). To see if the ligand-dependent interaction between cyclin D3 and PPARγ DEF could also be observed using full-length PPARγ, we incubated GST-cyclin D3 with full-length in vitro-translated 35S radiolabeled PPARγ. We observed no interaction enhancement between cyclin D3 and PPARγ in the presence of ligand, (Fig. 5D), possibly due to the masking of the ligand-dependent effect by the additional contribution of the AB domain.
We next set out to identify the region of cyclin D3 responsible for the interaction with PPARγ. Upon amino acid sequence screening of cyclin D3, we identified two LXXLL nuclear receptor interaction motifs located at the N- and C-terminal regions of the transcript (Fig. 5E). To test the contribution of these LXXLL motifs on the interaction with PPARγ, we performed site-specific mutagenesis, converting the second L to I, and performed GST pull-down with purified full-length PPARγ. Despite mutations of both LXXLL sites, the interaction with PPARγ was not disrupted, indicating that these sites do not contribute to the interaction with PPARγ (Fig. 5E, lane 4). Next, we created two deletion mutants of cyclin D3, Δ1-149, which lacks the cyclin box (CDK binding unit), and deletion mutant Δ148-293. GST pull-down assays reveal that amino acids 1 to 149 of cyclin D3 are required for the interaction with PPARγ (Fig. 5E).
To demonstrate that cyclin D3 could regulate the expression of PPARγ target genes in vivo, we performed chromatin immunoprecipitation experiments on differentiating 3T3-L1 cells. As expected, when chromatin of cells after 5 days of differentiation was immunoprecipitated with an anti-PPARγ antibody, we observed amplification of the region of the aP2 promoter containing the PPARγ response element (PPRE). Immunoprecipitation of cyclin D3 in the same conditions also resulted in amplification of the aP2 promoter (Fig. 5F, center panel). No amplification of the aP2 promoter was observed when either PPARγ or cyclin D3 was immunoprecipitated from confluent, undifferentiated 3T3-Ll cells, which do not express PPARγ or cyclin D3 (Fig. 5F, top panel). Binding of cyclin D3 and PPARγ was specific to the PPARγ binding site of the aP2 enhancer, since no amplification of a promoter region located outside the PPRE was observed (Fig. 5F, bottom panel).
To determine if the recruitment of cyclin D3 to PPARγ target genes is dependent on the presence of PPARγ, we performed additional chromatin immunoprecipitation experiments on NIH 3T3 cells, which do not express PPARγ, transfected or not with PPARγ2. We show that cyclin D3 is targeted to the PPRE of the aP2 promoter only when PPARγ is introduced into the cells by transfection (Fig. 5G). The recruitment of cyclin D3 to the aP2 promoter was, however, not found to depend on ligand (data not shown). The results of these chromatin immunoprecipitation assays demonstrate that cyclin D3 is recruited to the promoter of PPARγ target genes during adipogenesis and that this recruitment is dependent on the presence of PPARγ.
Cyclin D3/CDK6 complex phosphorylates PPARγ.Because cyclin D3, together with its cyclin-dependent kinase partners CDK 4 and 6, constitutes an active kinase that phosphorylates retinoblastoma protein during the cell cycle, we wanted to investigate whether cyclin D3 could also participate in PPARγ phosphorylation. To test this hypothesis, we performed in vitro kinase assays using an active cyclin D3/CDK6 kinase complex and purified PPARγ protein as the substrate. Kinase assays resulted in the appearance of a 60-kDa band corresponding to the size of recombinant PPARγt (Fig. 6A, lane 3). As a control, the kinase assay was performed in the absence of active kinase and in the absence of PPARγ (Fig. 6A, lanes 1 and 2, respectively).
Cyclin D3/CDK6 phosphorylates PPARγ. (A) In vitro kinase assay showing phosphorylation of purified full-length PPARγ by cyclin D3/CDK6 complex (lane 3). As a control, assays were performed in the absence of PPARγ (lane 1) or cyclin D3/CDK6 (lane 2). (B) The same kinase assay was performed using PPARγ-AB and DEF domain fusion proteins. (C) Activity of the UAS-TK-Luc reporter carrying the Gal4-specific response element measured in Cos cells upon transfecting expression vectors for cyclin D3, and chimeric Gal4-PPARγ containing a Gal4 DNA-binding domain and lacking the AB domain. The experiments were performed in triplicate in the presence or absence of the PPARγ agonist pioglitazone and were normalized for β-galactosidase activity. (D) Chromatin immunoprecipitation assays showing association of CDK6 with the PPRE of the aP2 promoter in differentiated 3T3-L1 cells (day 5). Cross-linked chromatin was incubated with antibodies against PPARγ, cyclin D3, CDK6, or nonspecific antibody (mock). Immunoprecipitates were analyzed by PCR using primers specific for the mouse aP2 promoter containing the PPRE and by primers amplifying a region outside the PPRE (negative control). A sample representing 0.1% of total chromatin was included in the PCR (Input).
Next, to investigate which domain of PPARγ is phosphorylated by cyclin D3/CDK6, we performed the same kinase assays on GST-PPARγ DEF and AB domains. The AB domain but not the DEF domain was found to be phosphorylated by cyclin D3/CDK6 (Fig. 6B). To further investigate the functional significance of the PPARγ-AB domain in the activation by cyclin D3, we performed cotransfection experiments using a chimeric Gal4-PPARγ lacking the AB domain whose activity is measured via the UAS-TK-Luc reporter construct. As expected, when chimeric Gal4-PPARγ was introduced into the system in the presence of ligand, a strong induction of luciferase activity was observed. However, unlike experiments performed with full-length PPARγ, cotransfection of cyclin D3 with PPARγ failed to increase the luciferase response (Fig. 6C). These results suggest that the AB domain of PPARγ is important for its activation by cyclin D3, possibly due the presence of important phosphorylation sites.
Next, to see if CDK6 could associate on the PPRE of the aP2 promoter and, thereby, contribute to the adipogenic process, we performed chromatin immunoprecipitation assays of CDK6 on differentiated 3T3-L1 cells. PCR analysis of CDK6 immunoprecipitations confirmed its presence on the PPRE of the aP2 promoter (Fig. 6D), further suggesting a role for CDK6 during adipogenesis.
Cyclin D3 null mice display a compromised adipose tissue phenotype.We have shown that cyclin D3 and PPARγ are expressed during the same time in the differentiation process and that cyclin D3 binds PPARγ and activates its transcriptional potential. To determine whether the activating effect of cyclin D3 could apply to in vivo models, we analyzed the adipose tissue phenotype of cyclin D3 mutant mice. Cyclin D3 mutant mice showed normal weight gain and initial examination of fat tissue mass revealed no significant differences in weight between cyclin D3 mutant and wild-type mice (data not shown and Fig. 7A).
Analysis of adipose tissue phenotype in cyclin D3-deficient mice. (A) Epididymal fat pad weight relative to total body weight for cyclin D3 mutant and wild-type mice. (B and C) Histological analysis of epididymal white adipose tissue (WAT) from cyclin D3 mutant and wild-type mice and corresponding histograms of adipocyte cell size. (D) Gene expression profiles of pertinent adipogenic genes from white adipose tissue measured by quantitative real-time PCR. (E) Weight gain curves for cyclin D3 mutant and wild-type mice fed a lipid-rich diet for 8 weeks. Each group was composed of six animals weighed on a weekly basis. Weight gain is adjusted to total initial body weight. (F) Fasting plasma glucose levels in cyclin D3 mutant and wild-type mice. (G) The intraperitoneal glucose tolerance test (IPGTT) measuring glucose levels at different times after intraperitoneal injection of glucose in cyclin D3 mutant and wild-type mice. (H) Glucose clearance after intraperitoneal injection of insulin (0.75 IU/kg) as a measure of insulin sensitivity in cyclin D3 mutant and wild-type mice.
Histological analysis of epididymal fat pads showed that adipocytes from cyclin D3 mutant mice were significantly smaller than wild-type mice (Fig. 7B and 7C), suggesting that cyclin D3 is important for normal adipocyte differentiation. Interestingly, this difference was observed in mice at 5 months of age but was absent in 3-month-old mice whose adipocytes were still of premature size in both the cyclin D3 mutant and wild-type groups (data not shown). Gene expression analysis of white adipose tissue taken from 5-month-old mice demonstrated a marked decrease in adipogenic markers aP2, LPL, and PPARγ (Fig. 7D).
We next analyzed the effect of challenging cyclin D3mutant mice with a high-fat diet. After feeding the mice a lipid-rich diet for 8 weeks, we observed a 30% decrease in weight gain in cyclin D3 mutant mice compared to their wild-type littermates (Fig. 7E). Because adipocyte size is also known to affect glucose homeostasis, we measured both glucose tolerance and insulin sensitivity in cyclin D3 mutant mice. Initial glucose measurements indicated that cyclin D3 mutant mice have a 30% decrease in fasting glucose levels (Fig. 7F). The intraperitoneal glucose tolerance test revealed that cyclin D3 mutants cleared glucose more efficiently than wild-type mice (Fig. 7G). Consistent with this observation, glucose decreased over two times more efficiently in cyclin D3 mutant compared to wild-type mice after insulin injection, indicating that the absence of cyclin D3 improves insulin sensitivity (Fig. 7H). Taken together, these in vivo studies confirm our in vitro data and suggest a crucial role for cyclin D3 in adipose tissue development.
The results presented in this study establish a new role for cyclin D3 as a PPARγ cofactor. We show that cyclin D3 is preferentially expressed in adipose tissue and that its expression is strongly induced during terminal stages of 3T3-L1 adipogenesis. We have identified cyclin D3 as a PPARγ coactivator capable of phosphorylating its AB domain. The essential role that cyclin D3 plays during adipogenesis was highlighted by the observation that silencing its expression strongly inhibited adipogenesis, whereas its overexpression promoted adipogenesis. Finally, we show that cyclin D3 mutant mice have a compromised adipose tissue phenotype. Our finding that cyclin D3 mutant mice display reduced adipocyte size, stunted adipogenic gene expression, resistance to high-fat weight gain, and insulin sensitivity is reminiscent of the phenotype observed in PPARγ+/− mice (21). This observation is consistent with the hypothesis that the phenotype observed in cyclin D3 mutant mice is due to reduced PPARγ activity.
Over 20 years ago, it was discovered how the ability of cyclins to bind and induce their CDK partners was dependent on their fluctuating expression pattern during the cell cycle (7). Here we show that the ability of cyclin D3 to bind PPARγ and help drive adipogenesis is also dependent on its differential expression pattern during adipogenesis. Strikingly, the stimulatory function of cyclin D3 during adipogenesis seems to fall deliberately outside its cell cycle role, as evidenced by its protein expression pattern; cyclin D3 is almost undetectable during the mitotic clonal expansion phase of adipogenesis, and then its expression is strongly induced during the noncycling terminal differentiation stage. Interestingly, an up-regulation of cyclin D3 expression has also been documented in other differentiation processes, including hematopoiesis (12, 26) and colon development (3); incidentally, PPARγ is also strongly induced during the latter (23). In the present study, we not only demonstrate an up-regulation of cyclin D3 during adipocyte differentiation but also identify PPARγ as a functional partner through which cyclin D3 mediates its proadipogenic effects.
The stimulatory effects of cyclin D3 on PPARγ could be the direct result of phosphorylation of PPARγ (Fig. 4C). Regulation of PPARγ activity by phosphorylation has already been documented. While some studies have linked PPARγ phosphorylation with its activation (2), others have shown PPARγ phosphorylation inhibits its activity, as is the case with mitogen-activated protein kinase-mediated phosphorylation (16). Other cyclin/CDK complexes are able to phosphorylate nuclear receptors. This includes cyclin A/CDK2-driven phosphorylation and activation of the estrogen receptor alpha (36), the progesterone receptor (46), and the glucocorticoid receptor (20).
The notion that cyclin D3 functions as a PPARγ coactivator during adipogenesis is not completely unexpected. Several studies have emerged showing regulation of nuclear receptor biology by D-type cyclins. Cyclin D1 can activate estrogen receptor alpha transcription through a direct interaction with the ligand binding domain of the receptor (47). On the other hand, cyclin D1 was shown to repress transcriptional activity of the thyroid hormone receptor (24) and the androgen receptor (31). Moreover, cyclin D1 can interact with several cofactors, including SRC1 (47), P/CAF (27), histone deacetylase 3 (24, 31), and TAF250 (1). While less well explored, cyclin D3 has also been implicated in the regulation of nuclear receptors including activation of the retinoic acid receptor (5). In addition, cyclin D3 can bind the SRC family coactivator GRIP-1, thereby disrupting its association with transcriptional regulators (22).
Recently, Pestell and colleagues reported that cyclin D1 represses PPARγ expression (45). In addition to inhibiting PPARγ promoter activity, they show that cyclin D1 retards adipogenesis and correlate this block with reduced PPARγ expression and activity. Remarkably they show that mouse embryo fibroblasts from cyclin D1 mutant mice have elevated levels of PPARγ even prior to inducing differentiation. Our laboratory and others have observed cyclin D1 expression to rapidly decrease after the mitotic clonal expansion phase of 3T3-L1 differentiation (32) (data not shown), consistent with the finding that PPARγ inhibits cyclin D1 expression (44).
Together, this information has allowed us to develop a model for how D-type cyclins orchestrate the molecular events taking place during adipocyte differentiation. Before adipogenesis is induced and immediately after its induction, elevated levels of cyclin D1 block immature expression of PPARγ. After a couple of rounds of mitotic clonal expansion, cyclin D1 levels rapidly shoot down, releasing its inhibitory effect on PPARγ expression and poising the cell for terminal differentiation. Once PPARγ protein is induced, cyclin D3 expression is increased, allowing it to bind and activate PPARγ. Such a model is dependent on the timely expression of both cyclin D1 and D3 and highlights their distinctive cell cycle-independent roles during adipogenesis.
In the present study we have established a link between the cell cycle machinery and adipogenesis. The metabolic response needed for growth and/or calorie storage is under the direct control of extracellular nutrients, growth factors, and hormones. Growth stimuli, including glucose, insulin, and glucocorticoids, are known to have an immediate mitogenic effect, however, the pathways by which these nutrients initiate the metabolic response are poorly understood. Here, we shed new light on this question by showing how cyclin D3 can functionally bind and activate the master regulator of adipogenesis, PPARγ. Thus, we propose cyclin D3 as a type of metabolic sensor linking external nutritional stimuli with the metabolic growth response.
Jacques Teyssier is acknowledged for his technical assistance. We thank P. Sicinski for providing the cyclin D3 knockout mice and E. Sicinska and J. S. Annicotte for their helpful support and discussion.
This work was supported by grants from INSERM (Avenir), FRM, Alfediam, and ARC. Anna Abella is supported by an Inserm Poste Vert grant, and Irena Iankova is supported by La Ligue National Contre le Cancer Ph.D. fellowship. David Sarruf is supported by the Boehringer Ingelheim Fonds Ph.D. scholarship program.
Returned for modification 8 April 2005.
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