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TNF-α reduces g0s2 expression and stimulates lipolysis through PPAR-γ inhibition in 3T3-L1 adipocytes
Cytokine 69 (2014) 196–205
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Cytokine journal homepage: www.journals.elsevier.com/cytokine
TNF-a reduces g0s2 expression and stimulates lipolysis through PPAR-c inhibition in 3T3-L1 adipocytes Dan Jin, Jun Sun, Jing Huang, Yiduo He, An Yu, Xiaoling Yu, Zaiqing Yang ⇑ Key Laboratory of Agricultural Animal Genetics, Breeding and Reproduction of Ministry of Education, College of Life Science and Technology, Huazhong Agricultural University, Wuhan 430070, PR China
a r t i c l e
i n f o
Article history: Received 12 August 2013 Received in revised form 22 May 2014 Accepted 4 June 2014 Available online 1 July 2014 Keywords: TNF-a G0S2 PPAR-c Lipolysis Adipocyte
a b s t r a c t Tumor necrosis factor-a (TNF-a) is a multifunctional cytokine that acts as a mediator of obesity-linked insulin resistance (IR). It is commonly accepted that macrophage-derived TNF-a acts in a paracrine manner on adjacent adipocytes, induces lipolysis, which contributes to obesity-linked hyperglycemia. Several studies suggested that G0/G1 switch gene 2 (g0s2) was up-regulated during adipogenesis, and its protein could be degraded in response to TNF-a stimulation. The aim of the present work was to investigate the transcriptional regulation of g0s2 by TNF-a stimulation. In this study, 3T3-L1 pre-adipocytes were differentiated, and treated with TNF-a for 24 h. The effects of TNF-a on lipolysis and lipase expression were then examined. Our results revealed that TNF-a exerted dose- and time-dependent lipolytic effects, which could be partially reversed by overexpression of g0s2 and peroxisome proliferator-activated receptor-c (ppar-c). In addition, TNF-a treatment significantly reduced the expression of adiponectin, ppar-c, hormone-sensitive Lipase (hsl), adipose triglyceride lipase (atgl) as well as ATGL co-factors. Interestingly, TNF-a significantly decreased adiponectin and PPAR-c protein levels, while treatment with the proteasomal inhibitor MG-132 maintained PPAR-c levels. Degradation of PPAR-c almost completely abolished the binding of PPAR-c to the g0s2 promoter in adipocytes treated with TNF-a. We propose that proteasomal degradation of PPAR-c and the reduction of g0s2 content are permissive for prolonged TNF-a induced lipolysis. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction Lipolysis is a tightly regulated process that involves the coordinate participation of several lipases and lipid droplet (LD) anchored proteins. During conditions of nutrient scarcity, triglycerides (TGs) stored in LDs of adipocytes are mobilized to provide free fatty acids (FFAs) and glycerol into the circulation as energy fuel for other organs [1]. The mobilization of TGs is mediated by three lipases: adipose triglyceride lipase (ATGL), hormone sensitive lipase (HSL), and monoglyceride lipase (MGL), which catalyze three consecutive reactions [2–6]. In a well-established model, HSL and Abbreviations: G0S2, the G0/G1 switch gene 2; ATGL, adipose triglyceride lipase; HSL, hormone sensitive lipase; MGL, monoglyceride lipase; CGI-58, comparative gene identification-58; TNF-a, tumor necrosis factor-a; PPAR-c, peroxisome proliferator-activated receptor-c; IR, insulin resistance; LD, lipid droplet; TGs, triglycerides; qPCR, quantitative polymerase chain reaction; ChIP, chromatin immunoprecipitation. ⇑ Corresponding author. Tel.: +86 27 87282669; fax: +86 27 87287376. E-mail addresses: [email protected] (D. Jin), [email protected] (Z. Yang). http://dx.doi.org/10.1016/j.cyto.2014.06.005 1043-4666/Ó 2014 Elsevier Ltd. All rights reserved.
the LD coat protein Perilipin-A can be phosphorylated by cAMPdependent protein kinase (PKA). Consequently, HSL translocates from the cytoplasm to LD, where it mediates hydrolysis of TGs [7]. Additional experiments using specific HSL inhibitors and/or si-RNA mediated knockdown of ATGL suggested that HSL is an important lipase in mediating hormone stimulated lipolysis in human fat, and also that the action of ATGL precedes HSL in the sequential hydrolysis of TGs [8,9]. ATGL is the rate-limiting enzyme for TG hydrolysis in adipocytes and is highly expressed in adipose tissue. Its TG hydrolase activity is due to a patatin domain located in its N-terminus. The catalytic site of ATGL is within the patatin domain, and is an unconventional catalytic dyad similar to that of human cytosolic phospholipase A2 (cPLA2) [10]. Deletion of the C-terminal region of ATGL retained its TG lipase activity but abrogated its LD localization [11]. In additional studies, adipose-specific overexpression of ATGL promoted lipolysis and attenuated diet-induced obesity in mice [12]. Two proteins tightly regulate ATGL activity: comparative gene identification-58 (CGI-58 or Abhd5) and G0/G1 switch gene 2 (G0S2). It has also been reported that ATGL is phosphorylated by
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AMPK at Ser406, leading to increased TG hydrolase activity [13]. CGI-58 is essential for ATGL activation, and is expressed in various tissues. Mice lacking CGI-58 exhibit increased fat mass and die a few hours after birth, most likely due to a severe skin defect [14]. A hydrophobic stretch of 45 amino acids in the C-terminal region of CGI-58 mediates the targeting of ATGL to the surface of LDs [11]. Interestingly, selective overexpression of CGI-58 in adipose tissue does not increase lipolysis nor protect against diet-induced obesity [15]. G0S2, another regulatory protein for ATGL, is abundantly expressed in metabolically active tissues such as fat and liver [16,17]. G0S2 was originally identified as a cell cycle regulator that commits a cell to enter G1 phase from G0 in blood mononuclear cells [18]. Combined with its growth arrest-associated expression in pre-adipocytes and high levels of expression in human and animal adult adipose tissue, G0S2 acts as a molecular brake for TG catabolism in adipocytes [6,19,20]. G0S2 directly interacts with ATGL and inhibits its TGs hydrolase activity, even with concurrent activation of CGI-58 on the surface of the LD [6]. Mutagenesis analysis revealed that the hydrophobic domain of G0S2 and the patatin-like domain of ATGL are the primary interacting regions of these two proteins [10]. Moreover, the TG hydrolase activity of ATGL can be selectively inhibited by G0S2 in response to fasting and caloric restriction [6,10]. TNF-a, a cytokine secreted by mature adipocytes and macrophages, plays an important role in the regulation of glucose and lipid metabolism. It is also one of the major cytokine responsible for inflammation-induced lipolysis in adipose tissue. The lipolysis induced by TNF-a leads to elevation of serum glycerol and FFAs, which may cause hyperglycemia, hyperlipemia and IR. Under conditions of nutrient overload such as in obesity, elevated TNF-a secretion induces basal adipose lipolysis, contributing to an increase in systemic FFAs levels, and thereby exacerbating peripheral lipotoxicity and IR. Yang et al. reported that silencing ATGL expression in adipocytes almost completely abolished basal and TNF-a-induced FFA and glycerol release. Conversely, coexpression of G0S2 significantly decreased TNF-a-stimulated lipolysis, even when ATGL and CGI-58 were overexpressed. It is interesting to note that treatment with TNF-a drastically decreased the level of G0S2 mRNA and protein, suggesting that early reduction of G0S2 content is permissive for TNF-a induced lipolysis [19]. While it is clear that multiple factors are involved in TNF-a induced lipolysis, very few reports have directly addressed the transcriptional regulation of G0S2 in response to TNF-a treatment. In 3T3L1 adipocytes, TNF-a treatment drastically decreased many genes’ expression and the half-life of protein. Taken together, these results suggest that degradation of proteins may contribute to the fasting lipolysis inducing. In the present study, we show that treatment with TNF-a induces lipolysis in a dose- and time-dependent manner, while incubating adipocytes with recombinant TNF-a significantly enhances glycerol release from the cells. In addition, we demonstrate that treatment with TNF-a is sufficient to down-regulate expression of ppar-c and its target genes including adiponectin, hsl, atgl, cgi-58 and g0s2. Furthermore, the proteasomal inhibitor MG-132 could block the decreased expression of PPAR-c after treatment with TNF-a. The reduction in g0s2 mRNA was demonstrated by a PPAR-c chromatin immunoprecipitation (ChIP) experiment targeted to the G0S2 promoter. Overexpression of g0s2 was able to protect against the actions of TNF-a on lipolysis, and decrease glycerol release. Additionally, PPAR-c overexpression increased the mRNA levels of g0s2 and other genes involved in fatty acid metabolism. Our results provide important information on the molecular mechanisms by which TNF-a induces lipolysis through the down-regulation of ppar-c, the degradation of its protein and the down-regulation of the expression of the ATGL co-inhibitor, g0s2.
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2. Materials and methods 2.1. Antibodies and reagents The rabbit polyclonal antibody against PPAR-c2 (#2430) was from Cell Signaling Technologies. Monoclonal b-actin antibody (#1978) was purchased from Sigma–Aldrich. The affinity purified rabbit polyclonal antibody for Adiponectin was generated against a recombinant GST fusion protein containing the C-terminal region of mouse Adiponectin (residues 111–247) by our laboratory. HRPconjugated secondary antibodies were from Pierce Chemical Co. Ltd. Protease inhibitor cocktail was obtained from Roche Diagnostics. Lipolysis assay kit (#E1012) was purchased from Applygen. Reagents for PCR were obtained from Bio-Rad. Rosiglitazone, GW9662, MG132, Cycloheximide, insulin, dexamethasone, and 3isobutyl-1-methylxanthine (IBMX) were purchased from Sigma– Aldrich. Recombinant TNF-a protein was obtained from Cusabio. Lipofectamine 2000 and reagents for cell culture were purchased from Invitrogen. 2.2. Plasmids The complete open reading frames of mouse atgl, ppar-c2 and g0s2 were amplified by PCR using Pfu DNA polymerase. The resulting PCR products were cleaved as follows: ppar-c2 using BamHI/HindIII; g0s2 with BamHI/XhoI; and atgl by HindIII/XhoI. The digested products were purified and ligated to the eukaryotic expression vector pcDNA3.1 stocks that had been pre-digested by the appropriate restriction enzymes. The pSilencer 4.1-shPPARc2 vector expressing short-hairpin RNA (shRNA) against PPARc2 was constructed by inserting the sequence 50 -GATCCCCTGGCAA AGCATTTGTATTTCAAGAGAATACAAATGCTTTGCCAGGGCA-30 into the empty vector using BamHI and HindIII. The scrambled sequence was 50 -GATCCACAA GATGAAGAGCACCAATTCAAGAGATTGGTGCTCTTCATCTTGTTGA-30 . The integrity and fidelity of all constructs was verified by DNA sequencing. 2.3. Cell culture and transient transfection Pre-adipocyte 3T3-L1 cells were obtained from American Type Culture Collection. The 3T3-L1 cells were maintained in DMEM (25 mM Glucose) supplemented with 10% fetal bovine serum (FBS) and antibiotics at 37 °C on 10 cm dishes. Cells were grown to confluence in 10% FBS/DMEM for 48 h, before adipocyte differentiation was initiated by culturing the cells in maintenance medium supplemented with 167 nM insulin, 1.0 lM dexamethasone, and 0.5 mM IBMX (Day 0). Forty-eight hours later, cells were switched to maintenance medium supplemented with 167 nM insulin. Cells were maintained in this media until day 10. Adipocytes were considered to be differentiated and treated on day 8 with 50 lM rosiglitazone, 5 lM GW9662 and recombinant TNF-a protein. For transient transfection, differentiated 3T3-L1 adipocytes were transfected with plasmids using Lipofectamine 2000 following the manufacturer’s instructions. 2.4. Oil Red O staining and lipolysis measurement The differentiated 3T3-L1 cells were washed with PBS and incubated with the Oil Red O working solutions according A. Mehlem’s method [21]. The photos were collected by the Motic inverted microscope AE2000 (Motic) and managed with ImageJ (National Institutes of Health) and Adobe Photoshop. Lipolysis was evaluated by measuring the amount of glycerol released into the cell and medium. Aliquots of culture media were centrifuged to remove debris, and directly subjected to glycerol
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detection. Cells were washed twice in ice cold PBS, harvested, and lysed on ice by sonication in 0.1 ml lysis buffer. Protein concentrations in the lysates were used to normalize the lipolysis signals. The amounts of glycerol released were determined by using an auto-analyzer to detect the absorbance at 550 nm following the enzymatic assay kit manufacturer’s instructions (Applygen Technologies Inc.). 2.5. Quantitative PCR (qPCR) Total RNA was extracted from cells using Trizol (Invitrogen), and was reverse transcribed to cDNA using M-MLV reverse transcriptase. Real time qPCR was carried out using the CFX96 real time PCR system (Bio-Rad) and SYBR Green PCR Mixture (Bio-Rad) following the manufacturer’s instructions. Gene expression levels were normalized to 36B4, and the relative expression level of each gene was calculated using the 244Ct method. The primer sets were designed by Primer 5.0 and synthesized by Sangon Biotech, Ltd. All primers are listed in Table 1. 2.6. Protein isolation and western blot Total protein was isolated from cells that had been harvested and centrifuged in PBS (pH 7.4). The pellets were re-suspended and sonicated in buffer containing 20 mM Tris, pH 7.5, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, and a proteinase inhibitor cocktail. The lysates were centrifuged at 12,000g for 5 min, mixed with an equal volume of 2 loading buffer, denatured at 100 °C for 5 min, and separated by SDS–PAGE. Proteins were transferred to a polyvinylidene fluoride (PVDF) membrane in transfer buffer containing 25 mM Tris, pH 8.3, 192 mM glycine, 0.01% SDS, and 15% methanol using a DYCZ-40D (LiuYi) electrophoresis transfer cell to which 200 mA were applied for 120 min. Membranes were blocked in 5% non-fat dry milk, 20 mM Tris, pH 7.5, 500 mM sodium chloride, and 0.5% Tween-20 (TBS-T) for 1 h. Individual proteins were identified by blotting with primary antibodies at appropriate dilutions. HRP-conjugated secondary antibodies were incubated with the membrane at a dilution of 1:5000. Signals were detected with a chemiluminescence detection system (MF-ChemBIS Bio-Imaging Systems; DNR Bio-Imaging Systems Ltd.) Equivalent protein loading between the samples was verified by blotting membranes for b-actin.
2.7. Chromatin immunoprecipitation (ChIP) ChIP assays were carried out using the ChIP assay kit (Millipore). Differentiated 3T3-L1 cells were treated with 10 ng/ml TNFa for 24 h, harvested and cross linked with 1% formaldehyde for 10 min at room temperature, lysed in SDS buffer, and sonicated to generate 500–1000 bp DNA fragments. Proteins were immunoprecipitated in ChIP dilution buffer using anti-PPAR-c2 antibody or nonspecific rabbit IgG control, followed by incubation with protein-G magnetic beads. The beads were then washed, and the protein-G bead/antibody/chromatin complexes eluted. Cross linking was reversed overnight at 65 °C, and DNA was purified by phenol/chloroform. The primers were designed to amplify the region 1535 bp to 1359 bp (forward primer 50 -CTAAGGCTCTGGACTCTAAC-30 ; reverse primer 50 -CTGTGGCTATCGGTGTAC-30 ). ChIP-qPCR data was normalized by the percentage input method (Invitrogen). 2.8. Statistical analysis The data were reported as the mean ± S.E. of at least three independent experiments. Differences between two groups were compared by Student’s t test, and P values < 0.05 were considered statistically significant. Comparisons between the values for different variables were analyzed by one-way ANOVA, followed by Student’s t test. GraphPad Prism 5.0 was used for the statistical analysis. 3. Results 3.1. Effects of TNF-a on lipolysis in 3T3-L1 adipocytes To determine the lipolytic action of TNF-a, we treated fully differentiated 3T3-L1 adipocytes with varying concentrations of TNF-a over a broad time range. As shown in Fig. 1A, TNF-a treatment resulted in a significant decrease in LD size and number. A significant dose-dependent increase in the amount of glycerol released into the medium was observed in the 3T3-L1 adipocytes treated with TNF-a (5–10 ng/ml, p < 0.05 and 50–100 ng/ml, p < 0.01) for 24 h (Fig. 1B). Moreover, the lipolytic effect of TNF-a was timedependent and, unlike catecholamine, which stimulates lipolysis in minutes, the TNF-a stimulation increase lipolysis in cultured adipocytes was observed after 3 h, became more prominent after
Table 1 Characteristics of the specific used for the qPCR analysis. Gene symbol
Product size (bp)
Forward primer (50 –30 ) and reverse primer (50 –30 )
Accession number
Mus 36B4
184
NM_007475
Mus hsl
138
Mus atgl
155
Mus mgl
144
Mus cgi-58
180
Mus g0s2
130
Mus adiponectin
169
Mus ppar-c2
179
Mus g0s2 promoter
176
F: AGATTCGGGATATGCTGTTG R: ACATCACTCAGAATTTCAATGG F: TGAGATTGAGGTGCTGTC R: TGAGATTGAGGTGCTGTC F: TTCACCATCCGCTTGTTG R: AGTTCCACCTGCTCAGAC F: AGACTGTGTGGAAATATC R: AAGAGACCTCTGATTATTG F: CCCTTTCCTTCCAGTATTC R: CGTAACAGCACCACATAG F: GACAGAGAAGGGAGACAC R: CACAGCAGCAAATCAGTC F: GCTTATGTGTATCGCTCAG R: TGTGGTAAGAGAAGTAGTAGAG F: ACTATGGAGTTCATGCTTGTG R: CCTGATGGCATTGTGAGAC F: CTAAGGCTCTGGACTCTAAC R: CTGTGGCTATCGGTGTAC
Italic values indicates the direction of the nucleic acid sequence.
NM_001039507 NM_025802 NM_001166249 NM_026179 NM_008059 NM_009605 NM_011146 NC_000067.6
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Fig. 1. Effects of TNF-a on cellular morphology and glycerol release in 3T3-L1 adipocytes. (A) Cells were treated with serum free DMEM/0.1% BSA for 24 h or with 10 ng/ml of TNF-a in serum free DMEM/0.1% BSA for 24 h, Oil Red O staining illustrate the lipid droplets. (B–E) Dose–response and time-course analysis of TNF-a treatment were performed by using differentiated 3T3-L1 adipocytes. (B and D) cells were treated with varying concentrations of TNF-a in serum-free DMEM/0.1% BSA for 24 h; (C and E) cells were treated with 10 ng/ml of TNF-a for different periods of time. Afterwards, the medium was collected and the cells were lysed, and glycerol release was measured using an autoanalyser to detect the absorbance at 550 nm following an enzymatic assay. The results are presented as the means ± S.E. of at least three independent experiments. p < 0.05; p < 0.01.
6 h of treatment (p < 0.05), and peaked by 12–24 h (p < 0.01, Fig. 1C). Furthermore, TNF-a induced a dose- and time-dependent increase in the amount of glycerol in the cells. As shown in Fig. 1D and E, treatment with low doses of TNF-a had little effect on the intracellular glycerol content, especially treated for 1–3 h or treated with 1–5 ng/ml TNF-a. However, treatment with high concentrations of TNF-a, or prolonged incubation, significantly increased the intracellular glycerol content and cell death.
3.2. Effect of TNF-a on the expression of lipolytic genes To evaluate the dose- and time-dependent effects of TNF-a on the expression of lipolytic genes, we next measured the mRNA
levels of relevant genes under basal or TNF-a-stimulated conditions (Fig. 2A and B). Quantitative-PCR analysis revealed that levels of atgl and hsl mRNA were significantly decreased by 5 ng/ml TNF-a compared with control. Similarly, treatment with 10 ng/ml TNF-a for 6 h significantly decreased the expression of atgl and hsl.
3.3. TNF-a down-regulates expression of g0s2 and cgi-58 CGI-58 and G0S2 are two selective cofactors of ATGL, and previous studies demonstrated that TNF-a down-regulated the protein levels of G0S2. We therefore focused on the expression of the g0s2 and cgi-58 genes that may play key roles in lipolysis by regulating ATGL. 3T3-L1 adipocytes were incubated for 24 h in the pres-
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Fig. 2. Effect of TNF-a on lipases and co-factor genes expression in 3T3-L1 adipocytes. Dose–response (A, C, E, G) and time-course (B, D, F, H) analysis of TNF-a incubation were performed by using differentiated 3T3-L1 cells as described in Fig.1. Total RNA was extracted from the cells and subjected to qPCR to determine the hsl, atgl, cgi -58 and g0s2 mRNA levels. The mRNA levels were normalized to the content of 36B4 and illustrated as the means ± S.E. of at least three independent experiments.
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ence (1–100 ng/ml) or absence of TNF-a, or treated with 10 ng/ml TNF-a for 0–24 h. As shown in Fig. 2C and D, TNF-a treatment resulted in a significant reduction of cgi-58 and g0s2 mRNA levels compared with control. 3.4. TNF-a down-regulates ppar-c and adiponectin expression levels Our previous studies showed that chronic TNF-a treatment down-regulates many lipolysis genes. As shown in Fig. 2, it is difficult to accurately assess the contribution of individual regulators. However, it is interesting that all of these genes are up-regulated by the transcription factor PPAR-c, which also regulates fatty acid storage and glucose metabolism in adipocytes. The mRNA levels of ppar-c and its target adiponectin were significantly decreased after TNF-a treatment compared with control in 3T3-L1 adipocytes (Fig. 3A and B). The differences in mRNA content were also reflected at the protein level in TNF-a treated cells, which had significantly lower PPAR-c and adiponectin protein levels (Fig. 3C). To test the hypothesis that the action of PPAR-c in TNF-a induced lipolysis is enhanced by the down-regulation of lipase genes or ATGL co-factors, we transfected 3T3-L1 adipocytes with PPAR-c in the presence or absence TNF-a. Fig. 3D demonstrates that overexpression of ppar-c significantly reduced the amount of glycerol released into the medium in both the presence and absence of TNF-a. In addition, treatment with the PPAR-c agonist rosiglitazone led to activation of PPAR-c and reduced glycerol release. In contrast, the PPAR-c inhibitor GW9662 significantly increased the amount of glycerol released into the medium in the absence of TNF-a, but had no effect in the presence of TNF-a (Fig. 3E). Real time-PCR analysis of the expression of atgl, g0s2, and cgi-58 showed that ppar-c overexpression significantly increased their mRNA levels upon treatment with TNF-a (Fig. 3F). 3.5. Overexpression of g0s2 protects against TNF-a induced lipolysis Previous studies demonstrated that overexpression of g0s2 increases lipogenesis, and g0s2 depletion enhances lipolysis in adipocytes. To test the hypothesis that G0S2 is a prominent regulator of TNF-a-induced lipolysis, acting by regulating ATGL activity, we assessed the effect of g0s2 overexpression on the TNF-a-induced lipolytic response. We observed that g0s2 overexpression significantly decreased glycerol release under both basal and TNF-a stimulated conditions (Fig. 3D). Cells transfected with atgl had glycerol release that was increased by 1.57-fold and 2.45-fold at basal state and in response to TNF-a incubation, respectively. 3.6. TNF-a reduced the activity of PPAR-c binding to g0s2 promoter Previous studies revealed that PPAR-c binds to the g0s2 promoter and up-regulates its expression. Here we used chromatin immunoprecipitation and qPCR to assess the differences in PPARc binding activity alteration after 10 ng/ml TNF-a treated for 24 h. As shown in Fig. 4A, TNF-a caused a 70% decrease in PPAR-c binding to g0s2 promoter. Knockdown of PPAR-c expression by shRNA decreased the expression of g0s2 (Fig. 4B). Similarly, treatment with the PPAR-c agonist rosiglitazone was able to rescue g0s2 expression in the presence of TNF-a, whereas treatment with PPAR-c inhibitor GW9662 inhibit the expression of g0s2 in the absence of TNF-a, but had no role in the presence of TNF-a (Fig. 4C). 3.7. TNF-a depredated PPAR-c through a proteasomal pathway Next, we determined whether the protein levels of PPAR-c could be maintained by preventing protein degradation. We therefore treated cells with a translational inhibitor (cycloheximide,
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CHX) to block new protein synthesis, and measured the amount of intact PPAR-c over 24 h in the presence of TNF-a. We found that PPAR-c is unusually short lived and has a half-life less than 6 h in response to TNF-a (Fig. 4D). A specific proteasomal inhibitor, MG132, was effective in preventing the PPAR-c degradation in the presence of cycloheximide and TNF-a (Fig. 4D). Compared with control, treatment with MG132 significantly increased the mRNA levels of g0s2 and other PPAR-c target genes in the presence of TNF-a, but not in the absence of TNF-a (Fig. 4E).
4. Discussion In addition to its proliferative, apoptotic and pro-inflammatory actions, TNF-a also modulates metabolic processes and cellular differentiation. These metabolic functions include blocking the actions of insulin, inducing lipolysis and suppressing lipogenesis [22–24]. Unlike catecholamine, TNF-a acts to increase the basal lipolysis rate, which contributes to elevated plasma glycerol and FFA content [19,25]. Irrespective of the type of external stimuli and the intracellular signaling pathways involved, lipolysis mainly depends on the action of three lipases and their regulators [2,5,20]. ATGL, the rate limiting lipase in adipocytes, plays a key role in both basal and hormone-stimulated lipolysis. Its hydrolysis activity is regulated by CGI-58 and G0S2 [11]. Although substantial amounts of data demonstrate that the actions of TNF-a on lipolysis occur via activation of the extracellular signal related kinase signaling pathway, the action of this cytokine on lipases and their regulators is less well established. In the present study, we showed that TNF-a induced the lipolytic response through its modulatory role on PPAR-c, lipases, and lipid droplet proteins. We provided evidence that TNF-a treatment enhanced lipolysis through transcriptional regulation of the PPAR-c-ATGL/G0S2 axis in adipocytes. Under TNF-a-stimulated conditions, downregulation of g0s2 enhanced the lipolytic rate, whereas g0s2 overexpression was able to partially repress the lipolytic response. TNF-a treatment drastically reduced PPAR-c protein levels through proteasomal degradation, reduced the efficiency of PPAR-c binding to g0s2 promoter, and decreased g0s2 expression in adipocytes. Taken together, our results indicate that there is an important PPAR-c-dependent down-regulation of the lipolytic response in adipocytes, even though PPAR-c is a master regulator of adipogenesis. Both CGI-58 and G0S2 are critically involved in TNF-a induced lipolysis. The maximal lipolytic effect of TNF-a depends on the ATGL co-activator CGI-58, whereas G0S2 down-regulation is important for TNF-a-induced TG hydrolysis in adipocytes. Interestingly, TNF-a stimulation also down-regulates atgl and hsl mRNA levels but does not alter their protein levels [19,26]. Data presented here show a significant decrease in atgl, hsl, cgi-58 and g0s2 expression in response to TNF-a stimulation in a dose- and time-dependent manner. TNF-a-induced lipolysis in adipocytes is a progressive process, and prolonged TNF-a stimulation (over 24 h) is known to suppress the expression of numerous proteins that inhibit basal lipolysis [27,28]. During the early stages of TNF-a stimulation, a rapid deactivation or degradation of protective proteins facilitates TNF-a induced lipolysis [19,29,30]. In addition, adipocytes preferentially down-regulate the transcription of these proteins that play a protective role for LD against chronic TNF-a treatment. For example, treatment of 3T3-L1 adipocytes with TNF-a for 6 h causes a rapid reduction in G0S2 protein levels, and our data showed that prolonged treatment with TNF-a causes a dramatic reduction in the mRNA expression levels of cgi-58 and g0s2. These data suggest that the maximum stimulatory effect of TNF-a on lipolysis depends on downregulation of g0s2. Based on all available data, we propose that multiple processes tightly control TNF-a-induced lipolysis. During the early stages of lipolysis,
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Fig. 3. TNF-a induced lipolysis is paralleled by a decrease in PPAR-c protein and its target genes expression. (A and B) Dose–response (up) and time-course (down) analysis of TNF-a incubation were performed by using the differentiated 3T3-L1 cells. Total RNA was extracted from the cells and subjected to qPCR to determine the ppar-c and adiponectin mRNA levels. (C) Dose–response and time-course analysis of TNF-a incubation were performed by using differentiated 3T3-L1 cells. The cells were lysed and subjected to immunoblotting to determine the PPAR-c, adiponectin and b-actin protein content in adipocytes as described in methods. (D) Differentiated 3T3-L1 cells were transfected with pcDNA3.1, atgl, g0s2 and ppar-c expression vectors for 24 h, following by incubation without or with 10 ng/ml TNF-a for another 24 h, respectively. The glycerol content in the medium was measured, and expressed as fold change from the cells receiving pcDNA3.1. (E) Serum-deprived 3T3-L1 adipocytes were incubated without or with 10 ng/ml TNF-a combinations of 50 lM rosiglitazone or 5 lM GW9662 for 24 h. The medium glycerol concentrations were measured, and illustrated as fold change from control cells. (F) Differentiated 3T3-L1 cells were transfected with pcDNA3.1 or ppar-c expression vector for 24 h, following treated without or with 10 ng/ml TNF-a for another 24 h. The total RNA was extracted and subjected to qPCR to determine the adiponectin, atgl, cgi-58 and g0s2 mRNA levels. The results are presented as the means ± S.E. of three independent experiments. p < 0.05; p < 0.01.
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Fig. 4. Transcriptional regulation of g0s2 expression in 3T3-L1 adipocytes. (A) Serum-deprived 3T3-L1 cells were incubated without or with 10 ng/ml TNF-a for 24 h. The cells were subjected to ChIP to detect the PPAR-c binding to g0s2 promoter. The results are presented as gradation difference of target band after a PCR amplification or fold change from IgG after a qPCR analysis (NTC, no template control). (B) Differentiated 3T3-L1 cells transfected with pSilencer-shPPAR-c or control vector for twice, the total RNA was extracted and sujected to qPCR to determine g0s2 mRNA levels. (C) Serum-deprived 3T3-L1 adipocytes were incubated without or with 10 ng/ml TNF-a combinations of 50 lM rosiglitazone or 5 lM GW9662 for 24 h. Total RNA was subjected to qPCR to determine g0s2 mRNA levels. (D) 3T3L1 adipocytes were incubated with 10 ng/ml TNF-a, 5 lg/ml cycloheximide (CHX) alone or in the presence of 10 lM MG132 for varying length of time. At each time points the cells were lysed and western blotting was performed as described in methods. (E) Serum-deprived 3T3-L1 adipocytes were incubated with 10 lM MG132 in the presence or absence of 10 ng/ml TNF-a for 24 h. Total RNA was extracted and subjected to qPCR to determine adiponectin, atgl, cgi-58 and g0s2 mRNA levels. The results are presented as the means ± S.E. of at least three independent experiments. p < 0.05; p < 0.01.
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TNF-a treatment rapidly abrogated G0S2 protein levels. The elimination of G0S2 allowed full activation of ATGL. During prolonged TNF-a treatment, further decreased expression of g0s2 allows additional and sustained activation of lipolysis. It is widely accepted that PPAR-c is an important regulator of adipogenesis and lipogenesis, and accumulating evidence suggests that PPAR-c may also coordinate the balance between fat deposition and mobilization through its action on lipolysis [31,32]. Nevertheless, the relationship between PPAR-c regulated lipid metabolism and hyperglycemia and type-2 diabetes remain unclear. PPAR-c is a therapeutic target in obesity related disease, specifically through marketed drugs such as Rosiglitazone, a PPAR-c agonist that functions as an insulin sensitizer, and reduces serum glucose, fatty acid and insulin concentrations [33,34]. Tan et al. showed that functional defects in PPAR-c (P467L) lead to excessive release of free fatty acids from triglyceride-rich lipoproteins, with an impaired fatty acid trapping in peripheral tissues. Interestingly, lipolysis was also inhibited, suggesting that the capacity of adipocytes to mobilize lipids was severely compromised [35]. Specifically, defective PPAR-c is associated with decreased basal expression of atgl, and hsl, as well as lipid droplet protein genes fsp27 and adrp both in vivo and in vitro [31]. Here, we showed that PPAR-c positively regulates the expression of atgl and hsl, which contribute to the two consecutive lipolytic reactions in adipocytes. TNF-a treatment caused proteasome-mediated degradation of PPAR-c and reduced the mRNA levels of atgl, cgi-58 and g0s2. G0S2 is an inhibitor of fat mobilization, and is upregulated by PPAR-c [6,7]. The degradation of PPAR-c also led to a down-regulation of g0s2 mRNA levels. G0S2 has a half-life of less than 1 h, and could be degraded through the proteasomal pathway in response to stimulation with TNF-a [19]. This suggests that G0S2 acts as a molecular brake on lipolysis in response to TNF-a. Upon down-regulation of g0s2 transcription caused by degradation of PPAR-c, the existing G0S2 protein is rapidly degraded by the proteasome, allowing the ATGL/CGI-58 complex to stimulate lipolysis without inhibition. The lower levels of PPAR-c also caused impaired fatty acid trapping, which combined with enhanced intracellular lipolysis, means that glycerol and fatty acid levels in circulation will dramatically increase. It is therefore possible that elevated TNF-a secretion in adipose tissue may directly contribute to low-levels of PPAR-c and G0S2, which in turn enhanced lipolysis and lead to increased glycerol and free fatty acid levels in obesity. Proteasome pathway is the major proteolytic pathway in mammalian cells. It degrades most short lived cellular proteins and abnormal proteins during the cell cycle control, apoptosis and antigen presentation [36]. MG-132 reduces the degradation of ubiquitin-conjugated proteins in mammalian cells, while cyclohexmide inhibits protein biosynthesis in eukaryotic. Data present here show a pronounced decrease in PPAR-c protein upon TNF-a and CHX treatment. However, add MG-132 treatment significantly ease the PPAR protein decreasing (Fig. 4D). Before the discovery of the ubiquitin proteasome system, protein degradation in cells was thought to rely mainly on lysosomes. The mechanisms by which TNF-a cause PPAR-c degradation are unknown. It needs more work to explain the role of PPAR-c in the TNF-a induced lipolysis and the proteasome pathway. TNF-a induces oxidative stress, inflammation and apoptosis, all these signal pathway can lead to update cell components through protein degradation [37]. Four distinct pathway for protein degradation have been identified: the lysosome, Ca2+ dependent, ATP-dependent, ATP and Ca2+ independent [36,38]. Ubiquitin conjugation and proteasome degradation composed the proteasome degradation system. Here we failed to detect the ubiquitin-conjugated PPAR-c. However, it is also reported that ubiquitination is not obligatory for substrate targeting to the proteasome [39]. The next need much work to explore the PPAR-c degradation pathway.
Obesity is a major cause of type-2 diabetes, clinically represented as hyperglycemia and hyperlipemia. During adipogenesis, PPAR-c and its targets get activated, leading to the synthesis and storage of TGs. Both mature adipocytes and macrophages in adipose tissue can secrete large amounts of TNF-a, which acts in a paracrine manner on adjacent cells to regulate glucose and lipid metabolism. TNF-a induces lipolysis in mouse adipocytes by down-regulation and degradation of PPAR-c and G0S2. Decreased levels of PPAR-c lead to a down-regulation of hsl, atgl, cgi-58 and g0s2. The mechanisms by which TNF-a down-regulates the transcription of three lipases, yet still enhances lipolysis, are currently unknown. TNF-a treatment does not alter the protein levels of ATGL, HSL and CGI-58 [19,26]. We therefore speculate that the effect of TNF-a on lipolysis is predominantly due to regulation of G0S2. While overexpression of g0s2 restored the lipolytic actions of TNF-a, activation or overexpression of ppar-c increased the expression of g0s2, while inhibiting the release of glycerol into the medium. Hence the proteasomal degradation of PPAR-c and its effects on TNF-a induced lipolysis could be of etiological importance for the development of insulin resistance in obesity. Moreover, previous experiments indicate that intracellular kinases such as ERK1/2, JNK and NF-jB also regulate TNF-a-induced lipolysis [30,40–44]. The pathways activated by these kinases are involved in the regulation of PPAR-c ATGL/G0S2 expression and degradation, making them important topics for future investigation. We therefore propose that disturbances in the PPAR-cATGL/G0S2 axis may contribute to dysregulated lipolysis that is associated with common forms of human obesity. Increased TNFa secretion from adipose tissue leads to the hyperglycemia and hyperlipidemia may be an underlying mechanism for IR in obesity.
Acknowledgements This work was supported by the grants from National key Basic Research Program of China (2012CB124702), 948 Program (2013S15), Specialized Research Fund for the Doctoral Program of Higher Education (20110146130002), Program of National Natural Science Foundation of China (31172093, 30970356), and the National Science Foundation for Fostering Talents in Basic Research (J1103510). Address all correspondence and requests for reprints to: Zaiqing Yang, Ph.D., Professor, College of Life Science & Technology, Huazhong Agricultural University, Wuhan, 430070, People’s Republic of China. Email: [email protected].
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TNF-a reduces g0s2 expression and stimulates lipolysis through PPAR-c inhibition in 3T3-L1 adipocytes Dan Jin, Jun Sun, Jing Huang, Yiduo He, An Yu, Xiaoling Yu, Zaiqing Yang ⇑ Key Laboratory of Agricultural Animal Genetics, Breeding and Reproduction of Ministry of Education, College of Life Science and Technology, Huazhong Agricultural University, Wuhan 430070, PR China
a r t i c l e
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Article history: Received 12 August 2013 Received in revised form 22 May 2014 Accepted 4 June 2014 Available online 1 July 2014 Keywords: TNF-a G0S2 PPAR-c Lipolysis Adipocyte
a b s t r a c t Tumor necrosis factor-a (TNF-a) is a multifunctional cytokine that acts as a mediator of obesity-linked insulin resistance (IR). It is commonly accepted that macrophage-derived TNF-a acts in a paracrine manner on adjacent adipocytes, induces lipolysis, which contributes to obesity-linked hyperglycemia. Several studies suggested that G0/G1 switch gene 2 (g0s2) was up-regulated during adipogenesis, and its protein could be degraded in response to TNF-a stimulation. The aim of the present work was to investigate the transcriptional regulation of g0s2 by TNF-a stimulation. In this study, 3T3-L1 pre-adipocytes were differentiated, and treated with TNF-a for 24 h. The effects of TNF-a on lipolysis and lipase expression were then examined. Our results revealed that TNF-a exerted dose- and time-dependent lipolytic effects, which could be partially reversed by overexpression of g0s2 and peroxisome proliferator-activated receptor-c (ppar-c). In addition, TNF-a treatment significantly reduced the expression of adiponectin, ppar-c, hormone-sensitive Lipase (hsl), adipose triglyceride lipase (atgl) as well as ATGL co-factors. Interestingly, TNF-a significantly decreased adiponectin and PPAR-c protein levels, while treatment with the proteasomal inhibitor MG-132 maintained PPAR-c levels. Degradation of PPAR-c almost completely abolished the binding of PPAR-c to the g0s2 promoter in adipocytes treated with TNF-a. We propose that proteasomal degradation of PPAR-c and the reduction of g0s2 content are permissive for prolonged TNF-a induced lipolysis. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction Lipolysis is a tightly regulated process that involves the coordinate participation of several lipases and lipid droplet (LD) anchored proteins. During conditions of nutrient scarcity, triglycerides (TGs) stored in LDs of adipocytes are mobilized to provide free fatty acids (FFAs) and glycerol into the circulation as energy fuel for other organs [1]. The mobilization of TGs is mediated by three lipases: adipose triglyceride lipase (ATGL), hormone sensitive lipase (HSL), and monoglyceride lipase (MGL), which catalyze three consecutive reactions [2–6]. In a well-established model, HSL and Abbreviations: G0S2, the G0/G1 switch gene 2; ATGL, adipose triglyceride lipase; HSL, hormone sensitive lipase; MGL, monoglyceride lipase; CGI-58, comparative gene identification-58; TNF-a, tumor necrosis factor-a; PPAR-c, peroxisome proliferator-activated receptor-c; IR, insulin resistance; LD, lipid droplet; TGs, triglycerides; qPCR, quantitative polymerase chain reaction; ChIP, chromatin immunoprecipitation. ⇑ Corresponding author. Tel.: +86 27 87282669; fax: +86 27 87287376. E-mail addresses: [email protected] (D. Jin), [email protected] (Z. Yang). http://dx.doi.org/10.1016/j.cyto.2014.06.005 1043-4666/Ó 2014 Elsevier Ltd. All rights reserved.
the LD coat protein Perilipin-A can be phosphorylated by cAMPdependent protein kinase (PKA). Consequently, HSL translocates from the cytoplasm to LD, where it mediates hydrolysis of TGs [7]. Additional experiments using specific HSL inhibitors and/or si-RNA mediated knockdown of ATGL suggested that HSL is an important lipase in mediating hormone stimulated lipolysis in human fat, and also that the action of ATGL precedes HSL in the sequential hydrolysis of TGs [8,9]. ATGL is the rate-limiting enzyme for TG hydrolysis in adipocytes and is highly expressed in adipose tissue. Its TG hydrolase activity is due to a patatin domain located in its N-terminus. The catalytic site of ATGL is within the patatin domain, and is an unconventional catalytic dyad similar to that of human cytosolic phospholipase A2 (cPLA2) [10]. Deletion of the C-terminal region of ATGL retained its TG lipase activity but abrogated its LD localization [11]. In additional studies, adipose-specific overexpression of ATGL promoted lipolysis and attenuated diet-induced obesity in mice [12]. Two proteins tightly regulate ATGL activity: comparative gene identification-58 (CGI-58 or Abhd5) and G0/G1 switch gene 2 (G0S2). It has also been reported that ATGL is phosphorylated by
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AMPK at Ser406, leading to increased TG hydrolase activity [13]. CGI-58 is essential for ATGL activation, and is expressed in various tissues. Mice lacking CGI-58 exhibit increased fat mass and die a few hours after birth, most likely due to a severe skin defect [14]. A hydrophobic stretch of 45 amino acids in the C-terminal region of CGI-58 mediates the targeting of ATGL to the surface of LDs [11]. Interestingly, selective overexpression of CGI-58 in adipose tissue does not increase lipolysis nor protect against diet-induced obesity [15]. G0S2, another regulatory protein for ATGL, is abundantly expressed in metabolically active tissues such as fat and liver [16,17]. G0S2 was originally identified as a cell cycle regulator that commits a cell to enter G1 phase from G0 in blood mononuclear cells [18]. Combined with its growth arrest-associated expression in pre-adipocytes and high levels of expression in human and animal adult adipose tissue, G0S2 acts as a molecular brake for TG catabolism in adipocytes [6,19,20]. G0S2 directly interacts with ATGL and inhibits its TGs hydrolase activity, even with concurrent activation of CGI-58 on the surface of the LD [6]. Mutagenesis analysis revealed that the hydrophobic domain of G0S2 and the patatin-like domain of ATGL are the primary interacting regions of these two proteins [10]. Moreover, the TG hydrolase activity of ATGL can be selectively inhibited by G0S2 in response to fasting and caloric restriction [6,10]. TNF-a, a cytokine secreted by mature adipocytes and macrophages, plays an important role in the regulation of glucose and lipid metabolism. It is also one of the major cytokine responsible for inflammation-induced lipolysis in adipose tissue. The lipolysis induced by TNF-a leads to elevation of serum glycerol and FFAs, which may cause hyperglycemia, hyperlipemia and IR. Under conditions of nutrient overload such as in obesity, elevated TNF-a secretion induces basal adipose lipolysis, contributing to an increase in systemic FFAs levels, and thereby exacerbating peripheral lipotoxicity and IR. Yang et al. reported that silencing ATGL expression in adipocytes almost completely abolished basal and TNF-a-induced FFA and glycerol release. Conversely, coexpression of G0S2 significantly decreased TNF-a-stimulated lipolysis, even when ATGL and CGI-58 were overexpressed. It is interesting to note that treatment with TNF-a drastically decreased the level of G0S2 mRNA and protein, suggesting that early reduction of G0S2 content is permissive for TNF-a induced lipolysis [19]. While it is clear that multiple factors are involved in TNF-a induced lipolysis, very few reports have directly addressed the transcriptional regulation of G0S2 in response to TNF-a treatment. In 3T3L1 adipocytes, TNF-a treatment drastically decreased many genes’ expression and the half-life of protein. Taken together, these results suggest that degradation of proteins may contribute to the fasting lipolysis inducing. In the present study, we show that treatment with TNF-a induces lipolysis in a dose- and time-dependent manner, while incubating adipocytes with recombinant TNF-a significantly enhances glycerol release from the cells. In addition, we demonstrate that treatment with TNF-a is sufficient to down-regulate expression of ppar-c and its target genes including adiponectin, hsl, atgl, cgi-58 and g0s2. Furthermore, the proteasomal inhibitor MG-132 could block the decreased expression of PPAR-c after treatment with TNF-a. The reduction in g0s2 mRNA was demonstrated by a PPAR-c chromatin immunoprecipitation (ChIP) experiment targeted to the G0S2 promoter. Overexpression of g0s2 was able to protect against the actions of TNF-a on lipolysis, and decrease glycerol release. Additionally, PPAR-c overexpression increased the mRNA levels of g0s2 and other genes involved in fatty acid metabolism. Our results provide important information on the molecular mechanisms by which TNF-a induces lipolysis through the down-regulation of ppar-c, the degradation of its protein and the down-regulation of the expression of the ATGL co-inhibitor, g0s2.
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2. Materials and methods 2.1. Antibodies and reagents The rabbit polyclonal antibody against PPAR-c2 (#2430) was from Cell Signaling Technologies. Monoclonal b-actin antibody (#1978) was purchased from Sigma–Aldrich. The affinity purified rabbit polyclonal antibody for Adiponectin was generated against a recombinant GST fusion protein containing the C-terminal region of mouse Adiponectin (residues 111–247) by our laboratory. HRPconjugated secondary antibodies were from Pierce Chemical Co. Ltd. Protease inhibitor cocktail was obtained from Roche Diagnostics. Lipolysis assay kit (#E1012) was purchased from Applygen. Reagents for PCR were obtained from Bio-Rad. Rosiglitazone, GW9662, MG132, Cycloheximide, insulin, dexamethasone, and 3isobutyl-1-methylxanthine (IBMX) were purchased from Sigma– Aldrich. Recombinant TNF-a protein was obtained from Cusabio. Lipofectamine 2000 and reagents for cell culture were purchased from Invitrogen. 2.2. Plasmids The complete open reading frames of mouse atgl, ppar-c2 and g0s2 were amplified by PCR using Pfu DNA polymerase. The resulting PCR products were cleaved as follows: ppar-c2 using BamHI/HindIII; g0s2 with BamHI/XhoI; and atgl by HindIII/XhoI. The digested products were purified and ligated to the eukaryotic expression vector pcDNA3.1 stocks that had been pre-digested by the appropriate restriction enzymes. The pSilencer 4.1-shPPARc2 vector expressing short-hairpin RNA (shRNA) against PPARc2 was constructed by inserting the sequence 50 -GATCCCCTGGCAA AGCATTTGTATTTCAAGAGAATACAAATGCTTTGCCAGGGCA-30 into the empty vector using BamHI and HindIII. The scrambled sequence was 50 -GATCCACAA GATGAAGAGCACCAATTCAAGAGATTGGTGCTCTTCATCTTGTTGA-30 . The integrity and fidelity of all constructs was verified by DNA sequencing. 2.3. Cell culture and transient transfection Pre-adipocyte 3T3-L1 cells were obtained from American Type Culture Collection. The 3T3-L1 cells were maintained in DMEM (25 mM Glucose) supplemented with 10% fetal bovine serum (FBS) and antibiotics at 37 °C on 10 cm dishes. Cells were grown to confluence in 10% FBS/DMEM for 48 h, before adipocyte differentiation was initiated by culturing the cells in maintenance medium supplemented with 167 nM insulin, 1.0 lM dexamethasone, and 0.5 mM IBMX (Day 0). Forty-eight hours later, cells were switched to maintenance medium supplemented with 167 nM insulin. Cells were maintained in this media until day 10. Adipocytes were considered to be differentiated and treated on day 8 with 50 lM rosiglitazone, 5 lM GW9662 and recombinant TNF-a protein. For transient transfection, differentiated 3T3-L1 adipocytes were transfected with plasmids using Lipofectamine 2000 following the manufacturer’s instructions. 2.4. Oil Red O staining and lipolysis measurement The differentiated 3T3-L1 cells were washed with PBS and incubated with the Oil Red O working solutions according A. Mehlem’s method [21]. The photos were collected by the Motic inverted microscope AE2000 (Motic) and managed with ImageJ (National Institutes of Health) and Adobe Photoshop. Lipolysis was evaluated by measuring the amount of glycerol released into the cell and medium. Aliquots of culture media were centrifuged to remove debris, and directly subjected to glycerol
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detection. Cells were washed twice in ice cold PBS, harvested, and lysed on ice by sonication in 0.1 ml lysis buffer. Protein concentrations in the lysates were used to normalize the lipolysis signals. The amounts of glycerol released were determined by using an auto-analyzer to detect the absorbance at 550 nm following the enzymatic assay kit manufacturer’s instructions (Applygen Technologies Inc.). 2.5. Quantitative PCR (qPCR) Total RNA was extracted from cells using Trizol (Invitrogen), and was reverse transcribed to cDNA using M-MLV reverse transcriptase. Real time qPCR was carried out using the CFX96 real time PCR system (Bio-Rad) and SYBR Green PCR Mixture (Bio-Rad) following the manufacturer’s instructions. Gene expression levels were normalized to 36B4, and the relative expression level of each gene was calculated using the 244Ct method. The primer sets were designed by Primer 5.0 and synthesized by Sangon Biotech, Ltd. All primers are listed in Table 1. 2.6. Protein isolation and western blot Total protein was isolated from cells that had been harvested and centrifuged in PBS (pH 7.4). The pellets were re-suspended and sonicated in buffer containing 20 mM Tris, pH 7.5, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, and a proteinase inhibitor cocktail. The lysates were centrifuged at 12,000g for 5 min, mixed with an equal volume of 2 loading buffer, denatured at 100 °C for 5 min, and separated by SDS–PAGE. Proteins were transferred to a polyvinylidene fluoride (PVDF) membrane in transfer buffer containing 25 mM Tris, pH 8.3, 192 mM glycine, 0.01% SDS, and 15% methanol using a DYCZ-40D (LiuYi) electrophoresis transfer cell to which 200 mA were applied for 120 min. Membranes were blocked in 5% non-fat dry milk, 20 mM Tris, pH 7.5, 500 mM sodium chloride, and 0.5% Tween-20 (TBS-T) for 1 h. Individual proteins were identified by blotting with primary antibodies at appropriate dilutions. HRP-conjugated secondary antibodies were incubated with the membrane at a dilution of 1:5000. Signals were detected with a chemiluminescence detection system (MF-ChemBIS Bio-Imaging Systems; DNR Bio-Imaging Systems Ltd.) Equivalent protein loading between the samples was verified by blotting membranes for b-actin.
2.7. Chromatin immunoprecipitation (ChIP) ChIP assays were carried out using the ChIP assay kit (Millipore). Differentiated 3T3-L1 cells were treated with 10 ng/ml TNFa for 24 h, harvested and cross linked with 1% formaldehyde for 10 min at room temperature, lysed in SDS buffer, and sonicated to generate 500–1000 bp DNA fragments. Proteins were immunoprecipitated in ChIP dilution buffer using anti-PPAR-c2 antibody or nonspecific rabbit IgG control, followed by incubation with protein-G magnetic beads. The beads were then washed, and the protein-G bead/antibody/chromatin complexes eluted. Cross linking was reversed overnight at 65 °C, and DNA was purified by phenol/chloroform. The primers were designed to amplify the region 1535 bp to 1359 bp (forward primer 50 -CTAAGGCTCTGGACTCTAAC-30 ; reverse primer 50 -CTGTGGCTATCGGTGTAC-30 ). ChIP-qPCR data was normalized by the percentage input method (Invitrogen). 2.8. Statistical analysis The data were reported as the mean ± S.E. of at least three independent experiments. Differences between two groups were compared by Student’s t test, and P values < 0.05 were considered statistically significant. Comparisons between the values for different variables were analyzed by one-way ANOVA, followed by Student’s t test. GraphPad Prism 5.0 was used for the statistical analysis. 3. Results 3.1. Effects of TNF-a on lipolysis in 3T3-L1 adipocytes To determine the lipolytic action of TNF-a, we treated fully differentiated 3T3-L1 adipocytes with varying concentrations of TNF-a over a broad time range. As shown in Fig. 1A, TNF-a treatment resulted in a significant decrease in LD size and number. A significant dose-dependent increase in the amount of glycerol released into the medium was observed in the 3T3-L1 adipocytes treated with TNF-a (5–10 ng/ml, p < 0.05 and 50–100 ng/ml, p < 0.01) for 24 h (Fig. 1B). Moreover, the lipolytic effect of TNF-a was timedependent and, unlike catecholamine, which stimulates lipolysis in minutes, the TNF-a stimulation increase lipolysis in cultured adipocytes was observed after 3 h, became more prominent after
Table 1 Characteristics of the specific used for the qPCR analysis. Gene symbol
Product size (bp)
Forward primer (50 –30 ) and reverse primer (50 –30 )
Accession number
Mus 36B4
184
NM_007475
Mus hsl
138
Mus atgl
155
Mus mgl
144
Mus cgi-58
180
Mus g0s2
130
Mus adiponectin
169
Mus ppar-c2
179
Mus g0s2 promoter
176
F: AGATTCGGGATATGCTGTTG R: ACATCACTCAGAATTTCAATGG F: TGAGATTGAGGTGCTGTC R: TGAGATTGAGGTGCTGTC F: TTCACCATCCGCTTGTTG R: AGTTCCACCTGCTCAGAC F: AGACTGTGTGGAAATATC R: AAGAGACCTCTGATTATTG F: CCCTTTCCTTCCAGTATTC R: CGTAACAGCACCACATAG F: GACAGAGAAGGGAGACAC R: CACAGCAGCAAATCAGTC F: GCTTATGTGTATCGCTCAG R: TGTGGTAAGAGAAGTAGTAGAG F: ACTATGGAGTTCATGCTTGTG R: CCTGATGGCATTGTGAGAC F: CTAAGGCTCTGGACTCTAAC R: CTGTGGCTATCGGTGTAC
Italic values indicates the direction of the nucleic acid sequence.
NM_001039507 NM_025802 NM_001166249 NM_026179 NM_008059 NM_009605 NM_011146 NC_000067.6
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Fig. 1. Effects of TNF-a on cellular morphology and glycerol release in 3T3-L1 adipocytes. (A) Cells were treated with serum free DMEM/0.1% BSA for 24 h or with 10 ng/ml of TNF-a in serum free DMEM/0.1% BSA for 24 h, Oil Red O staining illustrate the lipid droplets. (B–E) Dose–response and time-course analysis of TNF-a treatment were performed by using differentiated 3T3-L1 adipocytes. (B and D) cells were treated with varying concentrations of TNF-a in serum-free DMEM/0.1% BSA for 24 h; (C and E) cells were treated with 10 ng/ml of TNF-a for different periods of time. Afterwards, the medium was collected and the cells were lysed, and glycerol release was measured using an autoanalyser to detect the absorbance at 550 nm following an enzymatic assay. The results are presented as the means ± S.E. of at least three independent experiments. p < 0.05; p < 0.01.
6 h of treatment (p < 0.05), and peaked by 12–24 h (p < 0.01, Fig. 1C). Furthermore, TNF-a induced a dose- and time-dependent increase in the amount of glycerol in the cells. As shown in Fig. 1D and E, treatment with low doses of TNF-a had little effect on the intracellular glycerol content, especially treated for 1–3 h or treated with 1–5 ng/ml TNF-a. However, treatment with high concentrations of TNF-a, or prolonged incubation, significantly increased the intracellular glycerol content and cell death.
3.2. Effect of TNF-a on the expression of lipolytic genes To evaluate the dose- and time-dependent effects of TNF-a on the expression of lipolytic genes, we next measured the mRNA
levels of relevant genes under basal or TNF-a-stimulated conditions (Fig. 2A and B). Quantitative-PCR analysis revealed that levels of atgl and hsl mRNA were significantly decreased by 5 ng/ml TNF-a compared with control. Similarly, treatment with 10 ng/ml TNF-a for 6 h significantly decreased the expression of atgl and hsl.
3.3. TNF-a down-regulates expression of g0s2 and cgi-58 CGI-58 and G0S2 are two selective cofactors of ATGL, and previous studies demonstrated that TNF-a down-regulated the protein levels of G0S2. We therefore focused on the expression of the g0s2 and cgi-58 genes that may play key roles in lipolysis by regulating ATGL. 3T3-L1 adipocytes were incubated for 24 h in the pres-
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Fig. 2. Effect of TNF-a on lipases and co-factor genes expression in 3T3-L1 adipocytes. Dose–response (A, C, E, G) and time-course (B, D, F, H) analysis of TNF-a incubation were performed by using differentiated 3T3-L1 cells as described in Fig.1. Total RNA was extracted from the cells and subjected to qPCR to determine the hsl, atgl, cgi -58 and g0s2 mRNA levels. The mRNA levels were normalized to the content of 36B4 and illustrated as the means ± S.E. of at least three independent experiments.
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ence (1–100 ng/ml) or absence of TNF-a, or treated with 10 ng/ml TNF-a for 0–24 h. As shown in Fig. 2C and D, TNF-a treatment resulted in a significant reduction of cgi-58 and g0s2 mRNA levels compared with control. 3.4. TNF-a down-regulates ppar-c and adiponectin expression levels Our previous studies showed that chronic TNF-a treatment down-regulates many lipolysis genes. As shown in Fig. 2, it is difficult to accurately assess the contribution of individual regulators. However, it is interesting that all of these genes are up-regulated by the transcription factor PPAR-c, which also regulates fatty acid storage and glucose metabolism in adipocytes. The mRNA levels of ppar-c and its target adiponectin were significantly decreased after TNF-a treatment compared with control in 3T3-L1 adipocytes (Fig. 3A and B). The differences in mRNA content were also reflected at the protein level in TNF-a treated cells, which had significantly lower PPAR-c and adiponectin protein levels (Fig. 3C). To test the hypothesis that the action of PPAR-c in TNF-a induced lipolysis is enhanced by the down-regulation of lipase genes or ATGL co-factors, we transfected 3T3-L1 adipocytes with PPAR-c in the presence or absence TNF-a. Fig. 3D demonstrates that overexpression of ppar-c significantly reduced the amount of glycerol released into the medium in both the presence and absence of TNF-a. In addition, treatment with the PPAR-c agonist rosiglitazone led to activation of PPAR-c and reduced glycerol release. In contrast, the PPAR-c inhibitor GW9662 significantly increased the amount of glycerol released into the medium in the absence of TNF-a, but had no effect in the presence of TNF-a (Fig. 3E). Real time-PCR analysis of the expression of atgl, g0s2, and cgi-58 showed that ppar-c overexpression significantly increased their mRNA levels upon treatment with TNF-a (Fig. 3F). 3.5. Overexpression of g0s2 protects against TNF-a induced lipolysis Previous studies demonstrated that overexpression of g0s2 increases lipogenesis, and g0s2 depletion enhances lipolysis in adipocytes. To test the hypothesis that G0S2 is a prominent regulator of TNF-a-induced lipolysis, acting by regulating ATGL activity, we assessed the effect of g0s2 overexpression on the TNF-a-induced lipolytic response. We observed that g0s2 overexpression significantly decreased glycerol release under both basal and TNF-a stimulated conditions (Fig. 3D). Cells transfected with atgl had glycerol release that was increased by 1.57-fold and 2.45-fold at basal state and in response to TNF-a incubation, respectively. 3.6. TNF-a reduced the activity of PPAR-c binding to g0s2 promoter Previous studies revealed that PPAR-c binds to the g0s2 promoter and up-regulates its expression. Here we used chromatin immunoprecipitation and qPCR to assess the differences in PPARc binding activity alteration after 10 ng/ml TNF-a treated for 24 h. As shown in Fig. 4A, TNF-a caused a 70% decrease in PPAR-c binding to g0s2 promoter. Knockdown of PPAR-c expression by shRNA decreased the expression of g0s2 (Fig. 4B). Similarly, treatment with the PPAR-c agonist rosiglitazone was able to rescue g0s2 expression in the presence of TNF-a, whereas treatment with PPAR-c inhibitor GW9662 inhibit the expression of g0s2 in the absence of TNF-a, but had no role in the presence of TNF-a (Fig. 4C). 3.7. TNF-a depredated PPAR-c through a proteasomal pathway Next, we determined whether the protein levels of PPAR-c could be maintained by preventing protein degradation. We therefore treated cells with a translational inhibitor (cycloheximide,
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CHX) to block new protein synthesis, and measured the amount of intact PPAR-c over 24 h in the presence of TNF-a. We found that PPAR-c is unusually short lived and has a half-life less than 6 h in response to TNF-a (Fig. 4D). A specific proteasomal inhibitor, MG132, was effective in preventing the PPAR-c degradation in the presence of cycloheximide and TNF-a (Fig. 4D). Compared with control, treatment with MG132 significantly increased the mRNA levels of g0s2 and other PPAR-c target genes in the presence of TNF-a, but not in the absence of TNF-a (Fig. 4E).
4. Discussion In addition to its proliferative, apoptotic and pro-inflammatory actions, TNF-a also modulates metabolic processes and cellular differentiation. These metabolic functions include blocking the actions of insulin, inducing lipolysis and suppressing lipogenesis [22–24]. Unlike catecholamine, TNF-a acts to increase the basal lipolysis rate, which contributes to elevated plasma glycerol and FFA content [19,25]. Irrespective of the type of external stimuli and the intracellular signaling pathways involved, lipolysis mainly depends on the action of three lipases and their regulators [2,5,20]. ATGL, the rate limiting lipase in adipocytes, plays a key role in both basal and hormone-stimulated lipolysis. Its hydrolysis activity is regulated by CGI-58 and G0S2 [11]. Although substantial amounts of data demonstrate that the actions of TNF-a on lipolysis occur via activation of the extracellular signal related kinase signaling pathway, the action of this cytokine on lipases and their regulators is less well established. In the present study, we showed that TNF-a induced the lipolytic response through its modulatory role on PPAR-c, lipases, and lipid droplet proteins. We provided evidence that TNF-a treatment enhanced lipolysis through transcriptional regulation of the PPAR-c-ATGL/G0S2 axis in adipocytes. Under TNF-a-stimulated conditions, downregulation of g0s2 enhanced the lipolytic rate, whereas g0s2 overexpression was able to partially repress the lipolytic response. TNF-a treatment drastically reduced PPAR-c protein levels through proteasomal degradation, reduced the efficiency of PPAR-c binding to g0s2 promoter, and decreased g0s2 expression in adipocytes. Taken together, our results indicate that there is an important PPAR-c-dependent down-regulation of the lipolytic response in adipocytes, even though PPAR-c is a master regulator of adipogenesis. Both CGI-58 and G0S2 are critically involved in TNF-a induced lipolysis. The maximal lipolytic effect of TNF-a depends on the ATGL co-activator CGI-58, whereas G0S2 down-regulation is important for TNF-a-induced TG hydrolysis in adipocytes. Interestingly, TNF-a stimulation also down-regulates atgl and hsl mRNA levels but does not alter their protein levels [19,26]. Data presented here show a significant decrease in atgl, hsl, cgi-58 and g0s2 expression in response to TNF-a stimulation in a dose- and time-dependent manner. TNF-a-induced lipolysis in adipocytes is a progressive process, and prolonged TNF-a stimulation (over 24 h) is known to suppress the expression of numerous proteins that inhibit basal lipolysis [27,28]. During the early stages of TNF-a stimulation, a rapid deactivation or degradation of protective proteins facilitates TNF-a induced lipolysis [19,29,30]. In addition, adipocytes preferentially down-regulate the transcription of these proteins that play a protective role for LD against chronic TNF-a treatment. For example, treatment of 3T3-L1 adipocytes with TNF-a for 6 h causes a rapid reduction in G0S2 protein levels, and our data showed that prolonged treatment with TNF-a causes a dramatic reduction in the mRNA expression levels of cgi-58 and g0s2. These data suggest that the maximum stimulatory effect of TNF-a on lipolysis depends on downregulation of g0s2. Based on all available data, we propose that multiple processes tightly control TNF-a-induced lipolysis. During the early stages of lipolysis,
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Fig. 3. TNF-a induced lipolysis is paralleled by a decrease in PPAR-c protein and its target genes expression. (A and B) Dose–response (up) and time-course (down) analysis of TNF-a incubation were performed by using the differentiated 3T3-L1 cells. Total RNA was extracted from the cells and subjected to qPCR to determine the ppar-c and adiponectin mRNA levels. (C) Dose–response and time-course analysis of TNF-a incubation were performed by using differentiated 3T3-L1 cells. The cells were lysed and subjected to immunoblotting to determine the PPAR-c, adiponectin and b-actin protein content in adipocytes as described in methods. (D) Differentiated 3T3-L1 cells were transfected with pcDNA3.1, atgl, g0s2 and ppar-c expression vectors for 24 h, following by incubation without or with 10 ng/ml TNF-a for another 24 h, respectively. The glycerol content in the medium was measured, and expressed as fold change from the cells receiving pcDNA3.1. (E) Serum-deprived 3T3-L1 adipocytes were incubated without or with 10 ng/ml TNF-a combinations of 50 lM rosiglitazone or 5 lM GW9662 for 24 h. The medium glycerol concentrations were measured, and illustrated as fold change from control cells. (F) Differentiated 3T3-L1 cells were transfected with pcDNA3.1 or ppar-c expression vector for 24 h, following treated without or with 10 ng/ml TNF-a for another 24 h. The total RNA was extracted and subjected to qPCR to determine the adiponectin, atgl, cgi-58 and g0s2 mRNA levels. The results are presented as the means ± S.E. of three independent experiments. p < 0.05; p < 0.01.
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Fig. 4. Transcriptional regulation of g0s2 expression in 3T3-L1 adipocytes. (A) Serum-deprived 3T3-L1 cells were incubated without or with 10 ng/ml TNF-a for 24 h. The cells were subjected to ChIP to detect the PPAR-c binding to g0s2 promoter. The results are presented as gradation difference of target band after a PCR amplification or fold change from IgG after a qPCR analysis (NTC, no template control). (B) Differentiated 3T3-L1 cells transfected with pSilencer-shPPAR-c or control vector for twice, the total RNA was extracted and sujected to qPCR to determine g0s2 mRNA levels. (C) Serum-deprived 3T3-L1 adipocytes were incubated without or with 10 ng/ml TNF-a combinations of 50 lM rosiglitazone or 5 lM GW9662 for 24 h. Total RNA was subjected to qPCR to determine g0s2 mRNA levels. (D) 3T3L1 adipocytes were incubated with 10 ng/ml TNF-a, 5 lg/ml cycloheximide (CHX) alone or in the presence of 10 lM MG132 for varying length of time. At each time points the cells were lysed and western blotting was performed as described in methods. (E) Serum-deprived 3T3-L1 adipocytes were incubated with 10 lM MG132 in the presence or absence of 10 ng/ml TNF-a for 24 h. Total RNA was extracted and subjected to qPCR to determine adiponectin, atgl, cgi-58 and g0s2 mRNA levels. The results are presented as the means ± S.E. of at least three independent experiments. p < 0.05; p < 0.01.
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TNF-a treatment rapidly abrogated G0S2 protein levels. The elimination of G0S2 allowed full activation of ATGL. During prolonged TNF-a treatment, further decreased expression of g0s2 allows additional and sustained activation of lipolysis. It is widely accepted that PPAR-c is an important regulator of adipogenesis and lipogenesis, and accumulating evidence suggests that PPAR-c may also coordinate the balance between fat deposition and mobilization through its action on lipolysis [31,32]. Nevertheless, the relationship between PPAR-c regulated lipid metabolism and hyperglycemia and type-2 diabetes remain unclear. PPAR-c is a therapeutic target in obesity related disease, specifically through marketed drugs such as Rosiglitazone, a PPAR-c agonist that functions as an insulin sensitizer, and reduces serum glucose, fatty acid and insulin concentrations [33,34]. Tan et al. showed that functional defects in PPAR-c (P467L) lead to excessive release of free fatty acids from triglyceride-rich lipoproteins, with an impaired fatty acid trapping in peripheral tissues. Interestingly, lipolysis was also inhibited, suggesting that the capacity of adipocytes to mobilize lipids was severely compromised [35]. Specifically, defective PPAR-c is associated with decreased basal expression of atgl, and hsl, as well as lipid droplet protein genes fsp27 and adrp both in vivo and in vitro [31]. Here, we showed that PPAR-c positively regulates the expression of atgl and hsl, which contribute to the two consecutive lipolytic reactions in adipocytes. TNF-a treatment caused proteasome-mediated degradation of PPAR-c and reduced the mRNA levels of atgl, cgi-58 and g0s2. G0S2 is an inhibitor of fat mobilization, and is upregulated by PPAR-c [6,7]. The degradation of PPAR-c also led to a down-regulation of g0s2 mRNA levels. G0S2 has a half-life of less than 1 h, and could be degraded through the proteasomal pathway in response to stimulation with TNF-a [19]. This suggests that G0S2 acts as a molecular brake on lipolysis in response to TNF-a. Upon down-regulation of g0s2 transcription caused by degradation of PPAR-c, the existing G0S2 protein is rapidly degraded by the proteasome, allowing the ATGL/CGI-58 complex to stimulate lipolysis without inhibition. The lower levels of PPAR-c also caused impaired fatty acid trapping, which combined with enhanced intracellular lipolysis, means that glycerol and fatty acid levels in circulation will dramatically increase. It is therefore possible that elevated TNF-a secretion in adipose tissue may directly contribute to low-levels of PPAR-c and G0S2, which in turn enhanced lipolysis and lead to increased glycerol and free fatty acid levels in obesity. Proteasome pathway is the major proteolytic pathway in mammalian cells. It degrades most short lived cellular proteins and abnormal proteins during the cell cycle control, apoptosis and antigen presentation [36]. MG-132 reduces the degradation of ubiquitin-conjugated proteins in mammalian cells, while cyclohexmide inhibits protein biosynthesis in eukaryotic. Data present here show a pronounced decrease in PPAR-c protein upon TNF-a and CHX treatment. However, add MG-132 treatment significantly ease the PPAR protein decreasing (Fig. 4D). Before the discovery of the ubiquitin proteasome system, protein degradation in cells was thought to rely mainly on lysosomes. The mechanisms by which TNF-a cause PPAR-c degradation are unknown. It needs more work to explain the role of PPAR-c in the TNF-a induced lipolysis and the proteasome pathway. TNF-a induces oxidative stress, inflammation and apoptosis, all these signal pathway can lead to update cell components through protein degradation [37]. Four distinct pathway for protein degradation have been identified: the lysosome, Ca2+ dependent, ATP-dependent, ATP and Ca2+ independent [36,38]. Ubiquitin conjugation and proteasome degradation composed the proteasome degradation system. Here we failed to detect the ubiquitin-conjugated PPAR-c. However, it is also reported that ubiquitination is not obligatory for substrate targeting to the proteasome [39]. The next need much work to explore the PPAR-c degradation pathway.
Obesity is a major cause of type-2 diabetes, clinically represented as hyperglycemia and hyperlipemia. During adipogenesis, PPAR-c and its targets get activated, leading to the synthesis and storage of TGs. Both mature adipocytes and macrophages in adipose tissue can secrete large amounts of TNF-a, which acts in a paracrine manner on adjacent cells to regulate glucose and lipid metabolism. TNF-a induces lipolysis in mouse adipocytes by down-regulation and degradation of PPAR-c and G0S2. Decreased levels of PPAR-c lead to a down-regulation of hsl, atgl, cgi-58 and g0s2. The mechanisms by which TNF-a down-regulates the transcription of three lipases, yet still enhances lipolysis, are currently unknown. TNF-a treatment does not alter the protein levels of ATGL, HSL and CGI-58 [19,26]. We therefore speculate that the effect of TNF-a on lipolysis is predominantly due to regulation of G0S2. While overexpression of g0s2 restored the lipolytic actions of TNF-a, activation or overexpression of ppar-c increased the expression of g0s2, while inhibiting the release of glycerol into the medium. Hence the proteasomal degradation of PPAR-c and its effects on TNF-a induced lipolysis could be of etiological importance for the development of insulin resistance in obesity. Moreover, previous experiments indicate that intracellular kinases such as ERK1/2, JNK and NF-jB also regulate TNF-a-induced lipolysis [30,40–44]. The pathways activated by these kinases are involved in the regulation of PPAR-c ATGL/G0S2 expression and degradation, making them important topics for future investigation. We therefore propose that disturbances in the PPAR-cATGL/G0S2 axis may contribute to dysregulated lipolysis that is associated with common forms of human obesity. Increased TNFa secretion from adipose tissue leads to the hyperglycemia and hyperlipidemia may be an underlying mechanism for IR in obesity.
Acknowledgements This work was supported by the grants from National key Basic Research Program of China (2012CB124702), 948 Program (2013S15), Specialized Research Fund for the Doctoral Program of Higher Education (20110146130002), Program of National Natural Science Foundation of China (31172093, 30970356), and the National Science Foundation for Fostering Talents in Basic Research (J1103510). Address all correspondence and requests for reprints to: Zaiqing Yang, Ph.D., Professor, College of Life Science & Technology, Huazhong Agricultural University, Wuhan, 430070, People’s Republic of China. Email: [email protected].
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