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. Author manuscript; available in PMC: 2015 Apr 1.
Published in final edited form as: J Endocrinol. 2014 Nov 10;224(2):127–137. doi: 10.1530/JOE-14-0548

RGC-32 deficiency protects against high fat diet-induced obesity and insulin resistance in mice

Xiao-Bing Cui 1, Jun-Na Luan 1, Jianping Ye 3, Shi-You Chen 1,2
PMCID: PMC4293277  NIHMSID: NIHMS642227  PMID: 25385871

Abstract

Obesity is an important independent risk factor for type 2 diabetes, cardiovascular diseases and many other chronic diseases. Adipose tissue inflammation is a critical link between obesity and insulin resistance and type 2 diabetes and a contributor to disease susceptibility and progression. The objective of this study was to determine the role of response gene to complement 32 (RGC-32) in the development of obesity and insulin resistance. Wild-type (WT) and RGC-32 knockout (RGC32–/–) mice were fed normal chow or high-fat diet (HFD) for 12 weeks. Metabolic, biochemical and histologic analyses were performed. 3T3-L1 preadipocytes were used to study the role of RGC-32 in adipocytes in vitro. RGC32–/– mice fed with HFD exhibited a lean phenotype with reduced epididymal fat weight compared to WT controls. Blood biochemical analysis and insulin tolerance test showed that RGC-32 deficiency improved HFD-induced dyslipidemia and insulin resistance. Although it had no effect on adipocyte differentiation, RGC-32 deficiency ameliorated adipose tissue and systemic inflammation. Moreover, RGC32–/– induced browning of adipose tissues and increased energy expenditure. Our data indicated that RGC-32 plays an important role in diet-induced obesity and insulin resistance, and thus it may serve as a potential novel drug target for developing therapeutics to treat obesity and metabolic disorders.

Keywords: Response gene to complement 32, obesity, insulin resistance, adipose tissue

Introduction

Obesity is an important independent risk factor for type 2 diabetes and cardiovascular diseases, and it is associated with high morbidity and mortality (Friedman 2003). Although a range of therapies have been developed, these pharmaceutical therapies often have adverse side effects or limited efficacy (Bandyopadhyay 2009; Distefano and Watanabe 2010). Therefore, a continued effort is required to identify novel drug targets and develop more effective therapeutics for the treatment of obesity.

Adipose tissue plays a critical role in energy homeostasis, not only in storing triglycerides, but also in secreting hormones and biologically active molecules that regulate cellular lipid storage capacity, tissue and systemic insulin sensitivity, and metabolic energy balance (Chatterjee, et al. 2014). Although the underlying mechanism is not fully understood, adipose tissue inflammation has been acknowledged as a critical link between obesity and insulin resistance and type 2 diabetes, and as a contributor to disease susceptibility and progression (Mori, et al. 2010). Adipose tissue inflammation, characterized by increased secretion of proinflammatory cytokines and chemokines including interleukin (IL)-6, tumor necrosis factor (TNF)-α, and IL-12, contributes to low-grade systemic inflammation, insulin resistance, and metabolic disorders (Hotamisligil 2006; Xu, et al. 2003). Blocking the function of proinflammatory cytokines or chemokines results in improved insulin sensitivity and glucose homeostasis (Kanda, et al. 2006; Uysal, et al. 1997). Therefore, genes or molecules that control obesity or adipose tissue inflammation are promising therapeutic targets for insulin resistance, type 2 diabetes, and cardiovascular complications.

Response gene to complement 32 (RGC-32) is expressed in numerous human organs and tissues including placenta, kidney, liver, heart and brain (Badea, et al. 1998). Functionally, RGC-32 plays an important role in cell proliferation, differentiation (Fosbrink, et al. 2009; Li, et al. 2007; Wang, et al. 2011), fibrosis (Li, et al. 2011b) and cancer (Fosbrink, et al. 2005; Kim, et al. 2011; Vlaicu, et al. 2010). Recently, we generated RGC-32 knockout (RGC32–/–) mice and found that RGC32–/– mice were born smaller than their wild-type (WT) littermates because of the impaired placental angiogenesis (Cui, et al. 2013). However, it is unknown if RGC-32 plays a role in postnatal metabolism. In the present study, we demonstrate that high-fat diet (HFD) dramatically induces RGC-32 expression in the adipose tissue. RGC-32 deficiency attenuates HFD-induced obesity and insulin resistance in mice. The beneficial effect of RGC-32 deficiency is due to the decreased adipose tissue content and systemic inflammation and increased energy expenditure of adipose tissue. This is the first report showing that RGC-32 deficiency prevents diet-induced obesity and insulin resistance in mice. Therefore, RGC-32 may serve as a potential novel drug target for preventing obesity and type 2 diabetes.

Methods

Animals and diets

RGC32–/– mice on the C57BL/6 background were generated and genotyped as described previously (Cui et al. 2013). The parallel line WT C57BL/6 mice were purchased from the Jackson Laboratory. The age-matched WT and RGC32–/– male mice were maintained on normal chow for 8 weeks, after which they were fed with either normal chow (25% protein, 62% carbohydrate, and 13% fat; 3.07 kcal/g; 5053, LabDiet) or HFD (20% protein, 40% carbohydrate, and 40% fat; 4.5 kcal/g; D12108C, Research Diets) for an additional 12 weeks. Mice were fasted overnight and anaesthetized (2.0% isoflurane), and blood was collected by direct cardiac puncture. Epididymal fat was carefully removed and weighed. A portion of the epididymal fat was fixed in 4% paraformaldehyde for histological analysis, whereas the other portion was stored at –80°C for RNA and protein preparation. All animals were housed under conventional conditions in the animal care facilities and received humane care in compliance with the Principles of Laboratory Animal Care formulated by the National Society for Medical Research and the Guide for the Care and Use of Laboratory Animals. All experimental procedures were approved by Institutional Animal Care and Use Committee (IACUC) of University of Georgia.

Body weight and metabolic studies

Mice were weighed every 4 weeks. For metabolic studies, the mice were housed individually in metabolic cages (model 3600M021, Techniplast) with free access to food and water. After a 5-day period of acclimatization, daily food and water intakes were recorded, and fecal pellets and urine were weighed. The mice were studied for 3 days in the metabolic cage and then were put back to their original cages. To assess the energy expenditure, the mice were fasted overnight and weighed. This body weight was used as a baseline. The mice were fasted for an additional 8 hours and weighed, and then the percentage change of the body weight from the baseline was calculated.

Glucose and insulin tolerance tests

For the glucose tolerance test (GTT), mice were fasted overnight followed by an intraperitoneal glucose injection (1 g/kg body weight). Blood glucose was measured by tail bleeding using the One-Touch AccuChek Glucometer (Roche) at indicated times. For the insulin tolerance test (ITT), mice without fast were injected intraperitoneally with insulin (Sigma-Aldrich) at 1.5 IU/kg body weight, and blood glucose was measured at indicated times.

Blood biochemical analysis

Serum samples were analyzed for adiponectin, leptin, insulin, triglyceride and cholesterol using the Adiponectin Mouse ELISA kit (Abcam), Leptin Mouse ELISA kit (Abcam), Rat/Mouse Insulin ELISA Kit (Millipore), Triglyceride Quantification Kit (Abcam), and HDL and LDL/VLDL Cholesterol Assay Kit (Abcam).

Cytometric bead array immunoassay

Serum from individual mouse was assayed for inflammatory markers using cytometric bead array mouse inflammation kit (BD Biosciences). Data were collected by FACSCalibur flow cytometer (Becton Dickinson).

3T3-L1 cell culture and differentiation

3T3-L1 preadipocytes (ATCC) were maintained in Dulbecco's modified Eagle's medium (DMEM, Invitrogen) supplemented with 10% FBS (Sigma). At confluence, adipocyte differentiation was induced by treatment with 1 μmol/L dexamethasone (Sigma), 500 μmol/L isobutylmethylxanthine (Sigma), 10 μmol/L pioglitazone, and 100 nmol/L insulin (Sigma) for 2 days. Cells were then incubated in 100 nmol/L insulin-containing medium for 6 days to induce lipid accumulation. Medium was replaced every other day.

Oil red O staining

Culture plates were washed by PBS, and cells were fixed in 4% formaldehyde for 1 h, followed by staining with Oil Red O (Sigma) for 1 h. Oil Red O was prepared by diluting a stock solution (0.5% in isopropanol) with water (60:40 vol/vol) followed by filtration. After staining, plates were washed twice in water and photographed. The dye was then extracted with 100% isopropanol, and the absorbance was determined at 520 nm.

RNA extraction and real-time quantitative RT-PCR (qPCR)

qPCR was performed in a Mx3005P qPCR machine using SYBR Green master mix (Agilent Technologies) as previously described (Li, et al. 2011a). Each sample was amplified in triplicate. The expression of each gene was normalized with cyclophilin. Primer sequences are summarized in Table 1.

Table 1.

Primer sequences utilized for qPCR evaluation of gene expression

Gene Primer sequence
Cyclophilin 5’-GTG GTC TTT GGG AAG GTG AA-3’ (forward)
5’-TTA CAG GAC ATT GCG AGC AG-3’ (reverse)
RGC-32: 5’-CCG ATC TGG ACA GGA CCT TA-3’ (forward)
5’-AGC TTC ACT CTC CGA ACT GC-3’ (reverse)
PPAR-γ: 5’-ATCCCTGGTTTCATTAACCT-3’ (forward)
5’-GCTCCATAAAGTCACCAAAG-3’ (reverse)
PPAR-α: 5’-GGATGTCACACAATGCAATTCG-3’ (forward)
5’-TCACAGAACGGCTTCCTCAGGT-3’ (reverse)
C/EBPα: 5’-CAAGAACAGCAACGAGTACCG-3’ (forward)
5’-GTCACTGGTCAACTCCAGCAC-3’ (reverse)
Adiponectin: 5’-TGTTCCTCTTAATCCTGCCCA-3’ (forward)
5’-CCAACCTGCACAAGTTCCCTT-3’ (reverse)
Leptin: 5’-GTGGCTTTGGTCCTATCTGTC-3’ (forward)
5’-CGTGTGTGAAATGTCATTGATCC-3’ (reverse)
HSL: 5’-AAGGACTTGAGCAACTCAGA-3’ (forward)
5’-TTGACTATGGGTGACGTGTA-3’ (reverse)
PGC1α: 5’-ATACCGCAAAGAGCACGAGAA-3’ (forward)
5’-CTCAAGAGCAGCGAAAGCGTCACA-3’ (reverse)
IL-6 5’-CTGCAAGAGACTTCCATCCAG-3’ (forward)
5’-AGTGGTATAGACAGGTCTGTTGG-3’ (reverse)
IL-12 5’-TGGTTTGCCATCGTTTTGCTG-3’ (forward)
5’-ACAGGTGAGGTTCACTGTTTCT-3’ (reverse)
TNF-α 5’-CAGGCGGTGCCTATGTCTC-3’ (forward)
5’-CGATCACCCCGAAGTTCAGTAG-3’ (reverse)
UCP1 5’-TATCATCACCTTCCCGCTG-3’ (forward)
5’-GTCATATGTTACCAGCTCTG-3’ (reverse)
PRDM16 5’-AGCCCTCGCCCACAACTTGC-3’ (forward)
5’-TGACCCCCGGCTTCCGTTCA-3’ (reverse)
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Western blot analysis

Western blotting was performed as described previously (Cui, et al. 2012). Antibodies against hormone-sensitive lipase (HSL, Cell Signaling Technology, 4107S), peroxisome proliferator-activated receptor (PPAR)-α, (Abcam, ab8934), PPAR gamma coactivator (PGC)1α (Abcam, ab54481), RGC-32, and α-tubulin (Cell Signaling Technology, 9099S) were used. All the antibodies were applied at 1:1000 dilution. Protein expressions were detected using an enhanced chemiluminescence kit (Millipore).

Histological analysis

After fixing in 4% paraformaldehyde overnight, the epididymal fat were dehydrated and embedded in paraffin. Sections (5 μm) were cut with a microtome. Sections were deparaffinized and stained with hematoxylin and eosin (H&E). Images were captured by a Nikon microscope. For quantitative analysis of adipocyte area, eight images of H&E-stained sections were acquired from each animal, and cross-sectional area of each adipocyte was measured using ImageJ software.

Statistical analysis

Data are presented as means ± SD, and the numbers of independent experiments are indicated for each data set. For statistical analysis, the two groups were compared using two-tailed student's t-tests, and the four groups were evaluated by two-way ANOVA followed by Bonferroni post-hoc tests for multiple comparisons, whereas the three and five groups were evaluated by one-way ANOVA followed by Tukey's multiple comparisons using GraphPad Prism 5.0 software. P < 0.05 was considered statistically significant.

Results

RGC-32 deficiency prevented HFD-induced obesity

Our previous study has shown that the body size of RGC32–/– mice is smaller compared with their WT littermates when they are born (Cui et al. 2013). Interestingly, the difference is not as dramatic as they age, suggesting that RGC-32 has little effect on the post-natal growth on the regular chow diet. It is unknown, however, if RGC-32 affects HFD-induced obesity. To test this, we fed WT mice with HFD for 12 weeks, and then detected RGC-32 expression in adipose tissue. We found that RGC-32 expression was dramatically up-regulated by the HFD (Figure 1A). To investigate the potential role of RGC-32 in obesity, the WT and RGC32–/– mice were fed with HFD for 12 weeks. The HFD-fed WT mice gained significantly more weight than the normal chow controls. However, RGC32–/– appeared to diminish the weight gain (Figure 1B). The weight of epididymal fat pads was also markedly lower in HFD-fed RGC32–/– mice as compared to HFD-fed WT mice although it was increased compared to the normal chow controls (Figure 1C). Histological analysis of epididymal fat showed that HFD induced a significant adipocyte hypertrophy (more than 5 folds) in WT mice. However, this effect was significantly reduced in RGC32–/– mice (Figure 1D and 1E). To determine if the lean phenotype of RGC32–/– mice was due to a reduced energy intake, we housed the mice individually in metabolic cages and monitored the food intake. As shown in Figure 1F, the energy intake of WT and RGC32–/– mice fed with HFD was increased compared to the normal chow controls, while there was no difference between WT and RGC32–/– mice fed on either normal chow or HFD. There were also no significant differences in the water intake, urine, and feces (data not shown). To assess the energy expenditure, we measured the body weight before and after an 8-hour fast. In the absence of energy intake, greater loss of body weight indicates increased energy expenditure. After fasting, although there was no significant difference between WT and RGC32–/– mice under chow conditions, HFD-fed RGC32–/– mice lost more body weight than HFD-fed WT mice (Figure 1G), suggesting that the energy expenditure was increased in HFD-fed RGC32–/– mice, which may, at least partially, responsible for the lean phenotype of HFD-fed RGC32–/– mice.

Figure 1.

Figure 1

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RGC-32 deficiency prevented HFD-induced obesity. (A) RGC-32 expression in adipose tissue of wild-type (WT) mice fed with normal chow or a 12-week high-fat diet (HFD) were detected by western blot and normalized to α-tubulin (n = 3). (B) Body weight of WT and RGC32–/– mice fed with normal chow (n = 6) or HFD (n = 10). (C) Weights of epididymal fat from WT and RGC32–/– mice fed on normal chow and HFD. (D) Representative H&E-stained images of epididymal fat from WT and RGC32–/– mice fed on normal chow and HFD. (E) Quantitative analysis of the mean adipocyte area. The areas were normalized to the mean adipocyte area of WT mice fed on normal chow. (F) Energy intake of WT and RGC32–/– mice fed on normal chow and HFD. (G) Body weight change of WT and RGC32–/– mice after an 8-hour fast. **P<0.01 compared with WT chow group, ##P<0.01 compared with WT HFD group.

RGC-32 deficiency improved metabolic homeostasis in HFD-fed mice

Diet-induced obesity is typically accompanied by dyslipidemia and insulin resistance. Therefore, we measured serum triglyceride and cholesterol concentrations. No difference was observed between WT and RGC32–/– mice on normal chow (Figure 2A and 2B). However, on HFD, WT mice exhibited significantly increased serum concentrations of triglyceride, high-density lipoprotein (HDL) cholesterol, and low-density lipoprotein/very-low-density lipoprotein (LDL/VLDL) cholesterol (Figure 2A and 2B). Importantly, RGC32–/– mice appeared to be resistant to the HFD-induced increase of serum triglyceride and cholesterol. The serum triglyceride and LDL/VLDL cholesterol concentrations in RGC32–/– mice were not altered by the HFD feeding and thus were much lower compared to the HFD-fed WT control. HDL cholesterol was slightly lower in HFD-fed RGC32–/– mice than the WT control, although it was increased compared to RGC32–/– mice fed with normal chow (Figure 2A and 2B).

Figure 2.

Figure 2

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RGC-32 deficiency improved metabolic homeostasis in HFD-fed mice. (A) Serum triglyceride (TG), (B) high-density lipoprotein (HDL) cholesterol and low-density lipoprotein/very-low-density lipoprotein (LDL/VLDL) cholesterol concentrations in wild-type (WT) and RGC32–/– mice fed on normal chow and HFD (n = 6). (C) Fasting blood glucose, (D) insulin concentration, and (E) homeostasis model assessment-insulin resistance (HOMA-IR= fasting glucose × fasting insulin/22.5) in WT and RGC32–/– mice fed on normal chow and HFD (n = 6). **P<0.01 compared with WT chow group, ##P<0.01 compared with WT HFD group.

To determine if RGC-32 affects insulin sensitivity, blood glucose and serum insulin levels were detected. RGC32–/– mice showed similar fasting blood glucose and insulin levels compared with WT mice fed with normal chow (Figure 2C and 2D). Thus, the homeostasis model assessment-insulin resistance (HOMA-IR) scores had no difference (Figure 2E). HFD significantly increased fasting blood glucose level, insulin concentration, and HOMA-IR score in WT mice but not in RGC32–/– mice (Figure 2C-2E), suggesting that insulin sensitivity is improved due to RGC-32 deficiency. Improved insulin sensitivity in HFD-fed RGC32–/– mice was also confirmed using insulin tolerance test (ITT) although there was no difference between WT and RGC32–/– mice fed with normal chow (Figure 3A-3C). Furthermore, glucose tolerance test (GTT) showed that on normal chow, blood glucose levels appeared to be improved at 15 and 30 min (Figure 3D), and the area under the curve (AUC) was significantly lower in RGC32–/– mice compared with WT controls (Figure 3F), indicating that the glucose tolerance was improved in RGC32–/– mice under chow conditions. HFD-fed WT mice developed severe glucose intolerance, whereas RGC32–/– mice showed a significantly improved glucose tolerance compared to the HFD-fed WT mice (Figure 3E and 3F). These data demonstrate that RGC-32 deficiency improves glucose tolerance and protects mice from HFD-induced insulin resistance.

Figure 3.

Figure 3

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RGC-32 deficiency prevents HFD-induced insulin resistance in mice. (A and B) Insulin tolerance test (ITT) in wild-type (WT) and RGC32–/– mice fed on normal chow (A) and HFD (B) (n = 6). (C) Inverse area under the curve (AUC) of ITT. (D and E) Glucose tolerance test (GTT) in WT and RGC32–/– mice fed on normal chow (D) and HFD (E) (n = 6). (F) Quantification of the AUC of GTT. *P<0.05, **P<0.01 compared with WT chow group, #P<0.05, ##P<0.01 compared with WT HFD group.

RGC-32 deficiency attenuated adipose tissue and systemic inflammation in HFD-fed mice

Since adipose tissue inflammation and low levels of systemic inflammation are important factors contributing to the development of obesity, we detected adipose tissue and serum inflammatory status of the mice. RGC-32 deficiency appeared to decrease IL-6 and TNF-α mRNA expression in the fat tissue under chow conditions while had no effect on other adipokines such as adiponectin, leptin, and IL-12 (Figure 4A and 4B). HFD dramatically decreased adiponectin, an anti-inflammatory adipokine while increased the expression of pro-inflammatory adipokines leptin, IL-6, TNF-α, and IL-12 in the fat tissue of WT mice. However, these effects were alleviated in RGC32–/– mice (Figure 4A and 4B). Similar results were observed with the circulating adipokines (Figure 4C and 4D) although the circulating IL-6, TNF-α, and IL-12 were undetectable under chow conditions (data not shown). These data demonstrate that RGC-32 deficiency attenuates adipose tissue and systemic inflammation in HFD-fed mice.

Figure 4.

Figure 4

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RGC-32 deficiency attenuated adipose tissue and systemic inflammation in HFD-fed mice.(A and B) mRNA expression of adiponectin, leptin, interleukin (IL)-6, tumor necrosis factor (TNF)-α and IL-12 in epididymal adipose tissue from wild-type (WT) and RGC32–/– mice (n = 6) was measured by qPCR. (C and D) Protein concentration of adiponectin, leptin, IL-6, TNF-α and IL-12 in the serum from WT and RGC32–/– mice (n = 6) was measured by cytometric bead array immunoassay. *P<0.05, **P<0.01 compared with WT chow group, #P<0.05, ##P<0.01 compared with WT HFD group.

RGC-32 deficiency increased the expression of metabolic genes in adipose tissues

Since HFD-fed RGC32–/– mice had reduced fat mass and improved adipose tissue inflammation, we further assessed the expression of metabolic genes related with β-oxidation, lipolysis, and thermogenesis in adipose tissue. As shown in Figure 5A, RGC-32 deficiency increased the mRNA expression of PPAR-α, HSL, and PGC1α in epididymal fat tissue as compared to WT controls both under normal chow and HFD conditions. Since the alterations in HFD groups were greater than in the normal chow groups, and the body weight difference was observed in HFD groups, we confirmed the protein expression of these genes in the epididymal adipose tissue of HFD-fed mice (Figure 5B and 5C).

Figure 5.

Figure 5

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RGC-32 deficiency increased the expression of metabolic genes in adipose tissues. (A) mRNA expression of PPAR-α, HSL and PGC1α in epididymal adipose tissues from wild-type (WT) and RGC32–/– mice (n = 6). (B and C) PPAR-α, HSL and PGC1α protein expression in epididymal adipose tissues from WT and RGC32–/– mice were detected by western blot and normalized to α-tubulin (n = 6). (D and E) mRNA expression of PGC1α, UCP1, and Prdm16 in interscapular (D) and inguinal (E) fat tissues from WT and RGC32–/– mice (n = 6). *P<0.05, **P<0.01 compared with WT chow group, #P<0.05, ##P<0.01 compared with WT HFD group.

It is known that PGC1α can induce browning of subcutaneous adipose tissue through regulating uncoupling protein 1 (UCP1) (Bostrom, et al. 2012). We further determined adipose tissue browning in interscapular and inguinal fat tissues of WT and RGC32–/– mice. As shown in Figure 5D and 5E, PGC1α, UCP1, and Prdm16 were all increased in both interscapular and inguinal fat tissues of RGC32–/– mice compared with WT controls under both normal chow and HFD conditions. These data suggest that RGC-32 deficiency induces browning of adipose tissues, leading to increased energy expenditure.

RGC-32 had no effect on adipocyte differentiation

Since HFD induced a significant adipocyte hypertrophy in WT mice (Figure 1D and 1E), we sought to determine if RGC-32 promotes adipocyte differentiation. RGC-32 has been shown to induce smooth muscle cell differentiation (Li et al. 2007). In the differentiation of 3T3-L1 preadipocytes, RGC-32 expression was dramatically increased accompanied with elevated expression of adipogenic differentiation specific genes PPAR-γ and C/EBPα (Figure 6A-6C). To test RGC-32 function in this process, we knocked down or overexpressed RGC-32 in 3T3-L1 cells followed by induction of adipocyte differentiation and found that neither knockdown nor overexpression of RGC-32 had any effect on the expression of adipogenic specific genes (Figure 6D) or lipid accumulation (Figure 6E and 6F). These data indicate that the role of RGC-32 in obesity was not due to an increased adipocyte differentiation. To confirm the regulation of energy expenditure-related genes by RGC-32 as observed in the fat tissue (shown in Figure 5A-5C), we detected the protein expression of PPAR-α, HSL, and PGC1α in adipocytes differentiated from 3T3-L1 cells. Consistently, all of these genes were increased when RGC-32 was knocked down (Figure 6G and 6H). The expression of HSL and PGC1α was decreased when RGC-32 was overexpressed. PPAR-α expression appeared not to be dramatically affected by RGC-32 overexpression (Figure 6G and 6H). These data indicate that RGC-32 deficiency prevents mice from the diet-induced obesity through up-regulating energy expenditure-related genes in adipose tissues.

Figure 6.

Figure 6

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RGC-32 had no effect on adipocyte differentiation. (A) mRNA expression of RGC-32, PPAR-γ and C/EBPα during 3T3-L1 preadipocyte differentiation at the indicated times. (B and C) RGC-32 protein expression during 3T3-L1 preadipocyte differentiation was detected by western blot and normalized to α-tubulin. *P<0.05, **P<0.01 compared with vehicle-treated group (0 day). (D-H) 3T3-L1 preadipocyte was transduced with Ad-GFP, Ad-shRGC32 or Ad-RGC32 for 24 h and then was induced for adipocyte differentiation. (D) mRNA expression of RGC-32, PPAR-γ and C/EBPα and (E and F) lipid droplet accumulation were determined at the indicated times. (G and H) PPAR-α, HSL, PGC1α, and RGC-32 expression was detected by western blot and normalized to α-tubulin. *P<0.05, **P<0.01 compared with Ad-GFP group. All results are representatives of at least three independent experiments.

Discussion

Our present study demonstrates for the first time that RGC-32 plays an important role in HFD-induced obesity. RGC-32 is strongly up-regulated in adipose tissue of HFD-fed mice. RGC-32 deficiency prevents the development of HFD-induced obesity and insulin resistance because RGC32–/– mice fed a HFD gain less weight without changing energy intake and have a reduced fat mass. Interestingly, HFD increases the fat mass of RGC32–/– mice without altering their body weight, which is probably because the increase of the fat mass in these mice accounts for a relatively small portion of the whole body mass and thus does not cause a significant change in their body weight (as shown in Figure 1B). HFD-fed RGC32–/– mice also exhibit significantly improved insulin sensitivity as indicated by improved HOMA-IR and insulin tolerance. Our results are consistent with previous studies showing that HFD causes obesity and insulin resistance (Winzell and Ahren 2004), which is a characteristics of type 2 diabetes and several cardiometabolic syndromes including hypertension and dyslipidemia (Kim, et al. 2008). Although adipocyte differentiation is involved in diet-induced obesity (Berry, et al. 2012; Federico, et al. 2012), it may not be involved in RGC-32-mediated obesity because either overexpression or knockdown of RGC-32 has no effect on the expression of adipogenic differentiation specific genes PPAR-γ and C/EBPα and lipid accumulation of the adipocytes.

The protection from diet-induced obesity in RGC32–/– mice is linked to the attenuated adipose tissue and systemic inflammation and increased adipose tissue energy expenditure. Recent data show that adipose tissue inflammation is a key mechanism leading to obesity and insulin resistance (Chakrabarti, et al. 2009; Lee, et al. 2010). RGC-32 is essential for C5b-9-induced cell cycle activation (Fosbrink et al. 2009), indicating an important role in regulating inflammation. Indeed, RGC-32 deficiency suppresses adipose tissue and systemic inflammation in HFD-fed mice as evidenced by the reduced production of proinflammatory adipokines including leptin, IL-6, TNF-α, and IL-12 and increased anti-inflammatory adipokine adiponectin. Dietary excess and obesity have been shown to cause lipid accumulation in adipocytes, initiating a state of cellular stress and activation of NF-κB signaling pathway (Shoelson, et al. 2006), leading to an increased adipocyte production of proinflammatory cytokines. Our previous study has shown that RGC-32 activates NF-κB in vascular endothelial cells (Cui et al. 2013). Therefore, RGC-32 may increase inflammation of adipose tissue through activating NF-κB signaling pathway, which may be studied in the future.

RGC-32 appears to regulate the obesity development by influencing the thermogenesis. Particularly, RGC-32 may suppress the expression of metabolic genes PPAR-α, HSL, and PGC1α because dramatic elevation of PPAR-α, HSL, and PGC1α is observed in adipose tissue of HFD-fed RGC32–/– mice. HSL is a rate-limiting enzyme to cleave fatty acids from the triglyceride molecule (Schweiger, et al. 2006). PPAR-α promotes uptake, utilization, and catabolism of fatty acids by upregulation of genes involved in fatty acid transport, fatty binding and activation, and peroxisomal and mitochondrial fatty acid β-oxidation (Rakhshandehroo, et al. 2010). PGC1α enhances thermogenesis and oxidative metabolism of adipose tissue (Jun, et al. 2014; Liang and Ward 2006). Indeed, RGC-32 deficiency promotes the browning of adipose tissues. Brown adipose tissue is known to dissipates chemical energy and protect against obesity through a process termed nonshivering thermogenesis (Bi, et al. 2014). Active brown adipose tissue burns lipids to produce heat, resulting in an increase in energy expenditure. PGC1α and UCP1 are highly expressed in brown adipose tissue (Fisher, et al. 2012), and Prdm16 is a brown adipose determination factor (Seale, et al. 2011). Increased expression of these genes in interscapular and inguinal fat tissues may have collectively caused the increased energy expenditure and the lean phenotype of HFD-fed RGC32–/– mice. Besides adipose tissue, defective energy expenditure of skeletal muscle also contributes to the diet-induced obesity and insulin resistance. However, since RGC-32 expression is undetectable in skeletal muscle, contribution of skeletal muscle energy expenditure to the lean phenotype of HFD-fed RGC32–/– mice is likely to be minimal.

In summary, our study demonstrates that RGC-32 mediates HFD-induced obesity through enhancing inflammation while decreasing energy expenditure in adipose tissues.

Acknowledgments

Funding agencies: This work was supported by NIH grants HL107526, HL119053, and HL123302 and National Natural Science Foundation of China grant 81328002 (to S.Y.C.). X.B.C. is supported by an American Heart Association Postdoctoral Fellowship (14POST20480015).

Footnotes

Author contributions: X.B.C. conceived and carried out the experiments including data collection, analysis and interpretation, and wrote the manuscript. J.N.L. performed the qPCR, western blot and adipocyte differentiation experiments. J.Y. offered critical logistical advice on experimental design and data interpretation and critically read the manuscript. S.Y.C. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis. All authors had final approval of the submitted and published versions.

Disclosure: The authors declared no conflict of interest.

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