Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2021 Feb 1.
Published in final edited form as: Obesity (Silver Spring). 2020 Feb;28(2):293–302. doi: 10.1002/oby.22678

Mitochondrial pyruvate carriers are not required for adipogenesis, but regulated by high-fat feeding in brown adipose tissue

Jasmine A Burrell 1,2, Allison J Richard 2, William T King 3, Jacqueline M Stephens 1,2
PMCID: PMC6986308  NIHMSID: NIHMS1053027  PMID: 31970913

Abstract

Objective:

The objectives of this study were to assess the role of mitochondrial pyruvate carriers (MPCs) in adipocyte development in vitro and determine whether MPCs are regulated in vivo by high-fat feeding in male and female C57BL/6J mice.

Methods:

This study utilized siRNA-mediated knockdowns to assess the requirement of MPC1 for adipogenesis in the 3T3-L1 model system. Treatment with UK-5099, a potent pharmacological MPC inhibitor, was also used to assess the loss of MPC activity. Western blot analysis was performed on adipose tissue samples from mice on low fat diet (LFD) or a high fat diet (HFD) for 12-weeks.

Results:

The loss of MPC expression via siRNA-mediated knockdowns or pharmacological inhibition did not affect adipogenesis of 3T3-L1 cells. In vivo studies indicate that expression of MPCs is significantly decreased in brown adipose tissue of male, but not female, mice on HFD.

Conclusions:

Although MPCs are essential for pyruvate transport, MPCs are not required for adipogenesis in vitro suggesting that other substrates can be used for energy production when the MPC complex is not functional. Also, a significant decrease in MPC 1 and 2 expression occurred in brown fat, but not white fat, of male mice fed a high fat diet.

Keywords: adipocyte, brown adipose tissue, obesity

Introduction

The mitochondrial pyruvate carriers (MPCs) are transmembrane proteins, found on the inner mitochondrial membrane, that transport pyruvate from the cytosol into the mitochondrial matrix where pyruvate is oxidized to acetyl CoA and carbon dioxide via the pyruvate dehydrogenase complex (PDC). Acetyl CoA is utilized in the citric acid cycle to form NADH for oxidative phosphorylation (OXPHOS) to create a proton gradient for the production of ATP via the ATP synthase complex. There are two MPC proteins in mammals (MPC1 and MPC2), but there are three that have been identified in Saccharomyces cerevisiae (MPC1, MPC2, and MPC3) (1). In mammals, MPC1 and 2 form a heterodimer to actively transport pyruvate into the mitochondria (2). Since MPC1 and MPC2 act in a codependent manner, the loss or deletion of one of these proteins results in the loss of the other MPC due to destabilization (2–4). The study of MPC1 mutants has confirmed that dimerization of MPC1 and MPC2 in the inner mitochondrial membrane is essential for pyruvate uptake into the mitochondria (3).

MPC1 expression may also play a role in cancer metabolism as inhibition of MPC complex formation, via UK-5099, hinders pyruvate’s entry into the mitochondria and is associated with increased glycolysis (5). Increased glycolysis is a distinct metabolic feature of cancer cells, along with mitochondrial reprogramming and decreased lipid metabolism (6). The MPC inhibitor, 2-Cyano-3-(1-phenyl-1H-inodl-3-yl)-2-propenic acid (UK-5099), binds to MPCs, which deactivates the MPC complex and inhibits pyruvate oxidation (7). Thiazolidinediones (TZDs) inhibit MPC in several cell types, and in skeletal muscle inhibition of MPC is associated with increased glucose uptake and increased insulin sensitivity. Using the CRISPR/Cas9 system, a heterozygous MPC1 knockdown model was generated and these mice had reduced lipid accumulation, increased lipolysis, enhanced fatty acid oxidation, and decreased energy expenditure (8). During adipogenesis, mitochondrial density is increased by 20- to 30- fold along with increases in mitochondrial gene expression and oxidative capacity to meet increasing energy requirements (9–13). Insufficient or reduced mitochondrial density in tissues with a high energy demand like the brain, heart, muscles, and endocrine organs, such as adipose tissue, are associated with disease states (14). Although MPCs have been studied in several cell and tissue types, the role of these proteins in adipocyte development and their regulation in conditions of obesity is not known.

In our novel studies, we examined the requirement of MPC1 during adipocyte development using two independent approaches. Both pharmacological inhibition of MPCs as well as siRNA-mediated knockdown of MPC1 demonstrated that the expression and/or activity of MPC1 was not required for adipogenesis in 3T3-L1 cells. To our knowledge, this is the first siRNA-mediated MPC1 knockdown reported in 3T3-L1 cells. However, siRNA-mediated knockdown of MPC1 and MPC2 in 832/13 β-cells resulted in impaired insulin secretion in response to glucose and reductions in glucose-stimulated oxygen consumption (15). We also examined the modulation of MPC expression in both brown and white adipose tissue depots of male and female mice in a rodent model of diet-induced obesity. Expression levels of MPC1 and MPC2 were substantially decreased in the brown adipose tissue (BAT) of male mice following high-fat feeding, but not in female mice under the same conditions. In conclusion, our results indicate that loss of MPC1 does not have any effect on adipogenesis of 3T3-L1 preadipocytes; MPC1 and 2 are highly expressed in BAT; and MPC1 and 2 are modulated by high-fat feeding in BAT of male mice.

Materials and Methods

Animals and Diets-

Four-week-old male and female C57BL/6J mice were purchased from Jackson Laboratories (Stock #000664; Bar Harbor, ME). Animals were housed in a temperature (22 ± 2°C)- and humidity-controlled (45–55%) room under a 12-h light/dark cycle. At six weeks of age, mice were placed on respective diets. Mice were allowed ad libitum access to food and water. For twelve weeks, mice had access to either LFD containing 20% kcal protein, 70% kcal carbohydrate, and 10% kcal fat (D12450J; Research Diets, Inc. New Brunswick, NJ) or HFD containing 20% kcal protein, 20% kcal carbohydrate, and 60% kcal fat (D12492; Research Diets, Inc. New Brunswick, NJ). Body weights were obtained biweekly. Mice were fasted for four hours prior to sacrifice. All animal studies were performed with approval from the Pennington Biomedical Research Center Institutional Animal Care and Use Committee.

Animal body composition and glucose tolerance measurements -

Non-fasting body composition was measured by NMR (Bruckner Minispec) before beginning LF or HF diet intervention at 6 weeks of age and at 19 weeks of age (after 12 weeks of LFD- or HFD-feeding). Adiposity was calculated as total fat mass/total body weight x 100. Following a 4-hour fast, intraperitoneal glucose tolerance tests (IPGTTs) were performed on all animals at 17 weeks of age (10 weeks on diet). A baseline blood glucose measurement was obtained via tail nick (0 minutes), and animals were then injected with 2.5g/kg glucose. Blood glucose measurements were obtained via a drop of blood collected from the tail vein at 20, 40, 60, and 120 minutes post-injection. Blood glucose measurements were performed using a Breeze 2 glucometer (Bayer, Parsippany, NJ).

Cell culture-

Murine 3T3-L1 preadipocytes were grown in Dulbecco’s Modified Eagle’s Media (DMEM) (Sigma-Aldrich, St. Louis, MO) with 10% bovine calf serum. Two days after confluence, the preadipocytes were induced to differentiate using a standard protocol and induction cocktail composed of 3-isobutyl-methylxanthine, dexamethasone, insulin (MDI), and 10% characterized fetal bovine serum (FBS) in DMEM. HyClone calf and fetal bovine serum were purchased from Thermo Scientific (Waltham, MA) or GE Healthcare Life Sciences (Marlborough, MA). The medium was changed every 48 – 72 hours during growth and differentiation.

Pharmacological Inhibitor (UK-5099) Treatments-

3T3-L1 preadipocytes were trypsinized and seeded into 6-well plates at a density of 5.8 × 105 cells/cm2 in antibiotic-free 10% bovine calf/DMEM when approximately 70% confluent in 10-cm plates. 3T3-L1 preadipocytes were treated with 10μM UK-5099 (Sigma-Aldrich, St. Louis MO; PZ0160) added to the medium upon seeding. Preadipocytes were induced to differentiate as described above. Cells were fed with antibiotic-free media and treated with UK-5099 every 48 hours. Seven days after the induction of differentiation, the cell monolayers were harvested for protein in immunoprecipitation (IP) buffer containing 10mM Tris (pH 7.4), 150mM NaCl, 1mM EGTA, 1mM EDTA, 1% Triton X-100, 0.5% IGEPAL CA-630, protease inhibitors (1mM phenylmethylsulfonyl fluoride, 1μM pepstatin, 50 trypsin inhibitory milliunits of aprotinin, 10μM leupeptin, 1 mM 1,10-phenanthroline), and phosphatase inhibitors (0.2mM sodium vanadate and 100μM sodium fluoride), and for RNA in buffer provided in the RNeasy mini kit (Qiagen, Hilden, Germany) to assess knockdown efficiency. Three biological and technical replicates and were analyzed for each dose.

Cytotoxicity and cell viability assays-

The ToxiLight BioAssay kit (Lonza, Cologne, Germany) was utilized according to the manufacturer’s protocol. ToxiLight 100% lysis reagent (Lonza, Cologne, Germany) was utilized as a positive control. Cell viability was assessed by counting trypan blue-stained cells using a hemocytometer.

Whole cell extract preparation-

Cell monolayers were rinsed once with phosphate-buffered saline (PBS) and then scraped into non-denaturing IP buffer. The whole cell extracts were stored at −80°C before being thawed and passed through a 20-gauge needle five times. The whole cell extract was clarified via centrifugation at 13,000 × g for 10 min at 4°C.

Small interfering RNA (siRNA)-mediated knockdown-

3T3-L1 preadipocytes were trypsinized and re-plated in 6-well plates at a density of 5.8 × 105 cells/cm2 in antibiotic-free 10% bovine calf/DMEM when approximately 70% confluent in 10-cm plates. Using the protocol from Dharmacon, preadipocytes were transfected with 33nM siRNA (Dharmacon, Lafayette, CO; Non-targeting siRNA Cat #: D-001810–10-50, siRNA targeting MPC1 Cat #: L-040908–01-0005) and the DharmaFECT Duo transfection reagent (Dharmacon, Lafayette, CO, Cat #: T-2010–03) in OptiMEM reduced serum medium (Thermo Fisher, Waltham, MA; Cat #: 31985088). Non-targeting siRNA was used as negative control. Cells were treated with the siRNA cocktail during initial plating and grown to confluence. Two days after confluence, cells were induced to differentiate with the MDI-induction cocktail, as described above, and transfected again with the siRNA cocktail. After 48 hours, the cells were treated with 1/4 normal dose of insulin and transfected once again with the siRNA cocktail. Cells were fed every 48 hours with antibiotic-free media throughout the entire knockdown process. Seven days after the induction of differentiation, the cell monolayers were harvested for protein in IP buffer, and for RNA in buffer provided in the RNeasy mini kit (Qiagen, Hilden, Germany) to assess knockdown efficiency. Three biological and technical replicates were analyzed for each dose.

Respirometry-

Mitochondrial function was assessed by respirometry of intact cells using a Seahorse XFe24 Analyzer (Agilent Technologies; Santa Clara, CA). Mature 3T3-L1 adipocytes were seeded at 1.0×105 cells per well in an XF24 specialized cell culture microplate that was coated with 0.1% gelatin for 48 hours prior to seeding. Upon seeding, cells were incubated at 37°C without CO2 for one hour prior to assay in XF base media (Agilent Technologies; Santa Clara, CA) supplemented with 25mM glucose and 2mM L-glutamine at pH 7.4. After basal oxygen consumption was measured for three cycles, 1mM sodium pyruvate ± 10μM UK-5099 was injected into each well. After three basal measurement cycles, 1μM oligomycin, 600nM carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP), and 5μM rotenone/antimycin A were serially injected, and oxygen consumption was measured for three, six, and three cycles, respectively. After the assay was complete, cells were harvested in RIPA buffer, and protein concentration was quantified using the bicinchoninic acid (BCA) assay kit (Sigma-Aldrich, St. Louis, MO; Cat #: BCA1). Oxygen consumption rate was normalized to μg of protein.

Gel electrophoresis and immunoblotting-

Protein content of cell extracts was quantified via BCA assay. Samples were separated on 7.5%, 12%, or 15% sodium dodecyl sulfate (SDS) polyacrylamide (PA) gels (acrylamide; National Diagnostics, Atlanta, GA; Cat #: EC-890) and transferred to nitrocellulose membranes (BioRad, Hercules, CA; Cat #: 162–0115) in 25 mM Tris, 192 mM glycine, and 20% methanol. After the transfer, membrane strips were blocked in 4% non-fat milk for 1 hour at room temperature and washed with TBS-T before incubating with primary antibodies overnight at 4°C. Strips were washed with TBS-T and then incubated with either anti-mouse or anti-rabbit horseradish peroxidase-conjugated secondary antibodies (Jackson ImmunoResearch, West Grove, PA) for 1h. Strips were washed with TBS-T and visualized with enhanced chemiluminescence (Pierce/Thermo Scientific, Waltham, MA).

Antibodies-

Anti-STAT5A (L-20; sc-1081; rabbit polyclonal), anti-adipsin (M-120; sc-50419; rabbit polyclonal), anti-STAT3 (C-20; sc-482; rabbit polyclonal) and anti-ERK1/2 (C-16; sc-93; rabbit polyclonal) antibodies were purchased from Santa Cruz Biotechnology (Dallas, TX). Anti-adiponectin (PA1–054; rabbit polyclonal) antibody was purchased from Thermo Scientific (Waltham, MA). Anti-MPC1 (14462S; rabbit monoclonal) and anti-MPC2 (D4I7G; rabbit monoclonal) antibodies were purchased from Cell Signaling Technology (Danvers, MA).

RNA analysis-

Total RNA from tissue samples was extracted according to TRIzol manufacturer instructions. RNA from tissue or adipocyte monolayers was purified using the RNeasy mini kit (Qiagen, Hilden, Germany). Ten microliters of purified RNA were used for reverse transcription (RT) to generate cDNA according to the Applied Biosystems protocol (Applied Biosystems, Foster City, CA; Cat #: 4368813). cDNA was quantified using the real-time quantitative PCR (qPCR) method in a total volume of 10μL (2μL DNA and 8μL reaction master mix) using an Applied Biosystems 7900HT System with SDS 2.4 software. qPCR was performed using Takara SYBR premix (Takara Bio USA Inc., Madison, WI, USA) and primers from IDT (Integrated DNA Technologies, Skokie, IL, USA). Thermal cycling conditions were as follows 2 min at 50 °C, 10 min at 95 °C, 40 cycles of 15 s at 95 °C; dissociation stage 15 s at 95 °C, 15 s at 60 °C, and 15 s at 95 °C. Cyclophilin A (Ppia) and Non-POU domain containing octamer binding protein (NoNo) were used as reference genes. The following mouse genes were examined by RT-qPCR: Oxoglutarate Dehydrogenase (Ogdh), Succinate Dehydrogenase Complex Iron Sulfur Subunit B (Sdhb), Mitochondrial Pyruvate Carrier 1 (Mpc1), Mpc2, CCAAT enhancer-binding protein alpha (Cebpa), Peroxisome Proliferator-Activated Receptor Gamma (Pparg), Adiponectin (Adpn), Citrate synthase (Cs), Fatty Acid Binding Protein 4 (aP2), Fatty Acid Synthase (Fas), and Adipsin (Cfd). Primer sequences are shown in Table 1.

Table 1.

qPCR Primer Sequences

Gene Primer 1, 5’−3’ Primer 2, 5’−3’
Cyclophilin A (Ppia) CCACTGTCGCTTTTCGCCGC TGCAAACAGCTCGAAGGAGACGC
Non-POU domain containing octamer binding protein (Nono) CATCATCAGCATCACCACCA TCTTCAGGTCAATAGTCAAGCC
Oxoglutarate Dehydrogenase (Ogdh) ATGGGAAAGACCAAAGCTGA CCATGCAGCAGGATAGACAT
Succinate Dehydrogenase Complex Iron Sulfur Subunit B (Sdhb) GGAGGGCAAGCAACAGTATC GCACACAGGATGCACTCGTA
Mitochondrial Pyruvate Carrier 1 (Mpc1) CAAGGACTTCCGGGACTATC CATCCGCCCACTGATAATCTC
Mitochondrial Pyruvate Carrier 1 (Mpc2) CCGACTCATGGATAAAGTGGAG CTAGTCCAGCACACACCAAT
Adiponectin (Adpn) AAAAGGGCTCAGGATGCTACTG TGGGCAGGATTAAGAGGAACA
Citrate synthase (Cs) AGAACTCATCCTGCCTCGT CCTGCTCCTTAGGTATCAGATTG
Fatty acid binding protein 4 (aP2) CCCTCCTGTGCTGCAGCCTTT GTGGCAAAGCCCACTCCCACTT
Fatty Acid Synthase (Fas) GGCATCATTGGGCACTCCTT ACCACCAGCTGCCATGGATC
Adipsin (Cfd) CGAGGCCGGATTCTGGG GAGTCGTCATCCGTCACTCC
CCAAT enhancer-binding protein alpha (Cebpa) ACAAGAACAGCAACGAGTACC TCATTGTCACTGGTCAACTCC
Peroxisome proliferator-activated receptor gamma (Pparg) CGAGTGGTCTTCCATCACGG TCACAAGAAATTACCATGGTTGAC
Open in a new tab

Lipid Accumulation Measurement-

Seven days after the induction of differentiation, cells were fixed and stained with Oil Red O (ORO; Sigma-Aldrich, St. Louis, MO, USA) as described previously (16).

Statistical analysis-

Statistical analyses were performed using GraphPad Prism software (version 8; La Jolla, CA, USA). Differences between groups were calculated using Student’s t-tests and two-way ANOVA. Area under the curve (AUC) calculations and standard linear regression analyses were performed to determine correlations between MPC protein expression and adiposity or GTT AUC. Results from studies of cultured adipocytes are shown as mean ± standard error of the mean (SEM). Results were considered statistically significant when p < 0.05.

Results

1.1. Loss of MPC1 has no effect on in vitro adipogenesis

Prior to determining the potential requirement for MPC1 expression for adipocyte differentiation, we examined the protein expression of MPC1 and MPC2 over a seven-day adipogenesis timecourse in 3T3-L1 cells. As shown in Figure 1, both MPC1 and MPC2 protein expressions were highly induced during differentiation. The increased expression was apparent four days after the induction of adipogenesis (Figure 1A and 1B). To assess the requirement of MPCs on adipogenesis, we used a potent pharmacological inhibitor of MPCs, UK-5099, which acts by binding to MPCs and modifying a thiol group to prevent the formation of the MPC1 and MPC2 heterodimer, thus reducing pyruvate transport into the mitochondrial matrix (17, 18). Preadipocytes were induced to differentiate in the presence of UK-5099 at various doses. As shown in Figure 2, a range of inhibitor doses did not inhibit adipocyte differentiation as judged by lipid accumulation with Oil Red O staining (Figure 2A) or adipocyte marker gene expression (Figure 2B). Three adipogenic markers (adiponectin, aP2, and FAS) were examined to demonstrate adipocyte development.

Figure 1. MPC1 and MPC2 expression is induced during adipogenesis in 3T3-L1 cells.

Figure 1.

Open in a new tab

3T3-L1 adipocytes were induced to differentiate using the MDI cocktail, and whole cell extracts were harvested at the indicated time points to assess MPC1 protein expression over a timecourse of adipocyte differentiation. A) Whole cell extracts (75 μg of protein per lane) were subjected Western blot analysis. Adiponectin was utilized as a positive control for adipogenesis. B) Densitometry quantification of MPC1 and MPC2 band intensities at the indicated time points. MPC1 and 2 band intensities were normalized to ERK 1/2. (n = 3 pooled 10-cm plates per time point). Experiments were repeated three times on independent batches of cells.

Figure 2. UK-5099, a pharmacological inhibitor of MPC1, does not inhibit adipogenesis of 3T3-L1 cells.

Figure 2.

Open in a new tab

A and B) 3T3-L1 preadipocytes were induced to differentiate either untreated or treated with DMSO vehicle or indicated dose of UK-5099 at the time of induction and every 2 days during differentiation. A) Seven days post-induction, cells were fixed and stained with Oil Red O. Photographs are from the same experiment, and wells (n = 3 wells per treatment) were treated at the same time. B) Seven days post-induction, RNA was isolated, purified and subjected to RT-qPCR to measure gene expression of Mpc1, adipogenic markers [adiponectin (Adpn), adipocyte lipid binding protein (Ap2), and fatty acid synthase (Fas)], and the mitochondrial marker, citrate synthase (Cs) (n = 3 wells per treatment). Target gene expression was normalized to the reference gene, Nono, and data are plotted as fold change over the DMSO control. C and D) Respirometry experiments were performed to demonstrate the efficacy of the MPC1 inhibitor in mature 3T3-L1 adipocytes. After introducing 1mM pyruvate ± 10μM UK-5099, the oxygen consumption rate (OCR) was measured using a Seahorse XF analyzer for 3 cycles prior to performing the Mitochondrial Stress Test using 1 μM Oligomycin (Oligo), 600 nM FCCP (carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone) and 5 μM Rotenone (Rot)/ Antimycin A (AA). C) Full respiration profile showing the mean OCR normalized by the protein content ± SEM for each well (n = 10 wells per treatment). D) Maximal respiration after the addition of 600 nM FCCP is shown (n = 10 wells per treatment). Data shown are mean oxygen consumption rates of cells treated with pyruvate (P) or pyruvate and UK-5099 (P+I). Panels C and D were analyzed by unpaired t-test. **** indicates p < 0.0001 vs. pyruvate control. Experiments were repeated at least twice on independent batches of cells. E and F) Seven days post-induction, UK-5099 treated 3T3-L1 cells were either subjected to cytotoxicity assay (E) via ToxiLight BioAssay kit or stained with trypan blue and counted to assess cell viability (F) at indicated doses (n = 3 wells per treatment for a single batch of cells).

To examine the efficacy of UK-5099, respirometry experiments were performed on 3T3-L1 cells to measure oxygen consumption in the presence of pyruvate with or without the addition of the inhibitor. There were no differences in oxygen consumption observed during baseline readings after the addition of pyruvate ± UK-5099, or oligomycin. After the addition of FCCP, the maximal oxygen consumption rates in 3T3-L1 cells treated with the inhibitor were significantly reduced in comparison to the control (1.167 vs. 3.411pmol/min/μg protein) (Figure 2C and 2D). Oxygen consumption rates were reduced by UK-5099 in the presence of the mitochondrial uncoupler, FCCP, likely due to a reduction of pyruvate substrate entry into the mitochondria. Cell viability and cytotoxicity assays show that UK-5099-treated 3T3-L1 cells had no changes in cytotoxicity (Figure 2E) or cell viability (Figure 2F) in comparison to DMSO, which indicates that UK-5099 had no toxic effects on the cells.

An independent approach using siRNA-mediated knockdowns of MPC1 was also performed to examine the role of MPCs on adipogenesis in 3T3-L1 cells. Despite substantial loss of MPC1 protein expression (Figure 3A and 3B), there were no observed changes in adiponectin protein expression or lipid accumulation (Figure 3A and 3C). Though Mpc1 gene expression was significantly reduced, mRNA expression of Mpc2 and adipogenic markers, such as Adpn, Ap2, and Pparg were not affected (Figure 3D). Although Cebpa gene expression increased in the absence of MPC1, this increase did not affect adipogenesis. These data confirm that loss of MPC1 expression does not have any significant effects on adipogenesis of 3T3-L1 cells.

Figure 3. Knockdown of MPC1 gene expression with siRNA does not inhibit adipogenesis of 3T3-L1 cells.

Figure 3.

Open in a new tab

3T3-L1 preadipocytes were transfected with non-targeting siRNA (NT) or MPC1 siRNA upon plating and every 48 hours following induction of differentiation with MDI cocktail until endpoint assessments protein and gene expression as well as Oil Red O staining) were conducted on the cells at seven days post-MDI. A) Whole cell extracts were isolated and 50 μg protein per lane were analyzed by Western blot analysis (n = 3 replicate wells per treatment). Adiponectin was utilized as a differentiated adipocyte marker. B) MPC1 band intensities were quantified by densitometry and normalized to ERK 1/2 for each treatment. C) The cells were fixed and stained with Oil Red O to examine lipid accumulation. D) RNA was isolated, purified and subjected to RT-qPCR to show gene expression of Mpc1, Mpc2, adipogenic markers [adiponectin (Adpn), adipocyte lipid binding protein (Ap2), CCAAT enhancer-binding protein alpha (Cebpa) and peroxisome proliferator-activated receptor gamma (Pparg)], and mitochondrial markers [citrate synthase (Cs), oxoglutarate dehydrogenase (Ogdh) and succinate dehydrogenase (Sdhb)]. Data was analyzed by two-way ANOVA. * indicates p < 0.05, ** indicates p < 0.01, and *** indicates p < 0.001 vs. NT control (n = 3 wells per treatment).

1.2. MPCs are regulated in the adipose tissue of mice during high-fat feeding

To further study MPCs, we examined MPC1 and MPC2 protein expression in several adipose tissue depots and compared its expression with other tissues in mice. As shown in Figure 4, MPC1 and MPC2 were highly expressed in brown-adipose tissue (BAT), heart, liver and kidneys in C57BL/6J mice. When directly compared, it was apparent that there are differences in protein expression of MPC1 and MPC2 in some tissues. MPC1 had a much higher protein expression than MPC2 in the skeletal muscle, both the extensor digitorum longus (EDL) and the gastrocnemius (Gastroc), and in the brain (Figure 4). Also, the levels of MPC1 and MPC2 expression were low in all of the white adipose tissue (WAT) depots (Figure 4).

Figure 4. MPC1 and MPC2 proteins are highly expressed in brown adipose tissue, liver, and heart.

Figure 4.

Open in a new tab

Select tissues were harvested from a 27-week old C57BL/6J mouse to assess the tissue distribution of MPC1 and MPC2 proteins. A) Tissue samples were homogenized and 100 μg of total protein per lane were analyzed by Western blotting. B) MPC1 and MPC2 band intensities were quantified by densitometry and normalized to respective ERK 1/2 intensities for each tissue depot. gWAT, gonadal WAT; rpWAT, retroperitoneal WAT; iWAT, inguinal WAT; mWAT, mesenteric WAT; BAT, brown AT; EDL, extensor digitorum longus; gastroc, gastrocnemius.

To determine if MPC expression was regulated by diet-induced obesity, mice were placed on either low-fat (LF) or high-fat (HF) diets for 12 weeks. BAT, inguinal WAT (iWAT), and gonadal/epididymal WAT (gWAT) were harvested and analyzed in both male and female mice on LF or HF diets. In both male and female mice, Mpc1 mRNA expression was significantly decreased in the BAT, iWAT, and gWAT during high-fat feeding of male and female mice while Mpc2 mRNA expression was decreased in the gWAT of males and the BAT and gWAT of females during high-fat feeding (Figure 5). We also observed an expected decrease in fatty acid synthase (Fas) gene expression during high-fat feeding (19). Protein expression of both MPC1 and MPC2 in BAT was significantly reduced in male mice on HF diets (Figure 6A and 6B). This significant decline in MPC1 and MPC2 levels was not observed in the female mice fed a HF diets (Figure 6C and 6D). In iWAT, MPC1 protein expression was extremely low, near the detection limit, and no changes in protein levels were observed as a result of HFD-feeding (data not shown). These data indicate that MPC gene expression is downregulated in a depot and sex specific manner by high-fat feeding.

Figure 5. Steady state mRNA expression of MPC1 is substantially decreased in adipose tissue after high-fat feeding in male and female C57BL/6J mice.

Figure 5.

Open in a new tab

Six-week old C57BL/6J mice were fed either a high-fat (HF) or low-fat (LF) diet for 12 weeks ad libitum before euthanizing and collecting tissue. BAT, iWAT, and gWAT samples were homogenized and RNA was isolated and purified from each tissue depot for male (A) and female (B) mice. Each sample was subjected to RT-qPCR to show gene expression of Mpc1, Mpc2, adipogenic markers (Adpn, Ap2, and Fas), adipokine (Adipsin), and mitochondrial marker (Cs). Data was analyzed by unpaired t-tests. * indicates p < 0.05, ** indicates p < 0.01, *** indicates p < 0.001, and **** indicates p < 0.0001 vs. LF diet (n = 6 per condition). Each sample was run in duplicate. Target gene expression was normalized to the reference gene, Cyclophilin A, and data are plotted as fold change over the LF control.

Figure 6. MPC1 and MPC2 expression is significantly decreased in the brown adipose tissue of male, but not female mice following high-fat feeding.

Figure 6.

Open in a new tab

Six-week old male and female C57BL/6J mice were fed either a high-fat (HF) or low-fat (LF) diet for 12 weeks ad libitum before euthanizing and collecting BAT. Tissue samples were homogenized, and 30 μg of total protein per lane were subjected to Western blot analysis (n = 6 per condition) for males (A) and females (C). MPC1 and MPC2 band intensities were quantified by densitometry and normalized to respective STAT3 intensities for each sample; males (B) and females (D). Data was analyzed by two-way ANOVA. ** indicates p < 0.01 vs. LF diet.

Because female C57BL/6 mice are typically less susceptible to metabolic dysfunction during HF diet-induced obesity than their male counterparts (20–22), we examined the relationships between MPC1 or MPC2 protein expression and adiposity or glucose tolerance. Protein expression of MPC1 and MPC2 for males significantly correlates with adiposity and glucose tolerance test (GTT) AUC for male, but not female mice (Figure 7). For the male mice, the linear fits of MPC1 or MPC2 versus adiposity or GTT AUC have an R2 value of greater than 0.5 and the slopes are significantly non-zero (p < 0.01), while female mice R2 values are less than 0.4 and none of the slopes are significantly non-zero. These data show that decreased levels MPC proteins in male mice are associated with increased adiposity and decreased glucose tolerance.

Figure 7. Decreased MPC1 and MPC2 protein expression in BAT during DIO correlates with increased adiposity and decreased metabolic health in male, but not female, mice.

Figure 7.

Open in a new tab

A and B) MPC protein expression from the immunoblot quantification in Figure 6B and 6D was correlated with percent fat mass (adiposity) for male and female mice on LFD and HFD. Protein extracts were collected and adiposity measured at 19 weeks of age (12 weeks on diet). C and D) MPC protein expression values for male and female mice were also plotted against area under the curve (AUC) of glucose tolerance tests (GTTs) performed on LFD and HFD-fed mice at 17 weeks of age (10 weeks on diet) for male and female mice. Linear regression analyses were performed and are shown for data in A-D. R2 values indicate goodness of the linear fit for each data set, and whether the slope is significantly differently from zero is denoted by p value or ns (not significant).

Discussion

Our novel data demonstrate the dispensability of MPC1 during adipogenesis in vitro as well as showing the expression and modulation of MPCs in brown adipose tissue in vivo during diet-induced obesity (DIO). The observed increase in protein expressions of both MPC1 and MPC2 during adipocyte differentiation of 3T3-L1 cells (Figure 1) has not been previously reported in 3T3-L1 cells, but MPC1 and MPC2 expression is increased upon differentiation of LGR5+ intestinal stem cells (ISCs) (23). The increase in MPC1 and 2 expressions in adipocytes is likely due to increased mitochondrial density and gene expression that occurs to generate energy needed to accommodate the enhanced metabolic requirements during adipogenesis (9, 12, 13). Because of enhanced protein expression of MPCs during differentiation, we hypothesized that MPCs were necessary for the differentiation of 3T3-L1 cells.

Contrary to our premise, two independent approaches using both pharmacological inhibition with UK-5099 (Figure 2) and siRNA-mediated knockdowns (Figure 3) of MPCs did not affect adipocyte development. Even at doses as low as 50nM, UK-5099 has been able to inhibit formation of the MPC1 and 2 heterodimer, thus inhibiting pyruvate transport via this complex (4, 17). In BAT progenitor cells, the addition of UK-5099 inhibited 13C-glucose incorporation into acetyl CoA (17) suggesting that the pyruvate produced from glycolysis of 13C-glucose was not utilized for acetyl CoA production due to impaired pyruvate uptake by the MPC heterodimer via UK-5099 inhibition. Liver-specific loss of MPC2 in C57BL/6J mice resulted in compromised, but not eliminated, pyruvate metabolism and suggested that mechanisms such as pyruvate-alanine cycling were activated to compensate for the loss of the MPC complex functionality (24). A robust decline (~80%) in MPC1 expression via siRNA knockdowns also had no effect on adipocyte development (Figure 3). Our data further supports the current literature. Our observations suggest that although pyruvate is the primary substrate for energy production in the mitochondria, other substrates, such as alanine, glutamine (2, 24, 25), or branched chain amino acids (26), might bypass the MPC1/2 complex and enter into the mitochondrial matrix to compensate for the lack of pyruvate transport under conditions where MPC1 or MPC2 is not present or functional. A clear limitation of our adipogenesis studies is that they are solely in vitro observations. Although our findings show that the loss of MPC expression in vitro has no effect on adipocyte development, other in vivo studies have found that C57BL/6 mice with an adipocyte-specific loss of MPC1 had increased fatty acid oxidation, increased levels of triglycerides in circulation, deficiencies in storage of TGs, and mitochondrial damage. It was also observed that heterozygous MPC1 knockdown mice had decreased body weight, activity, fat accumulation, and low body shell temperatures during cold exposure (8, 27).

An analysis of several mouse tissue samples revealed that MPCs are highly expressed in BAT, heart, lung, and skeletal muscle (Figure 4). This is an expected finding, as each of these are mitochondria-rich tissues. However, it has not been reported that MPC1 and MPC2 are more highly enriched in BAT than other mitochondria-rich tissues (Figure 4). Because MPCs are highly enriched in BAT depots, but not in WAT depots, we assessed whether MPCs were regulated in AT depots during diet-induced obesity (DIO). Although MPC1 was downregulated in BAT and WAT during high-fat feeding at the mRNA level (Figure 5), MPC1 protein expression was barely detectable and not obviously changed in the WAT of either male or female mice during high-fat feeding (data not shown). The differences in gene and protein expression could be due to a variety of factors including translational rates, protein degradation rates, or a combination of both factors (28). However, it is a novel finding that MPC gene and protein expressions are downregulated in BAT of male, but not female, mice during DIO (Figures 5 and 6). It has been reported that reductions in diet-induced thermogenesis contribute to weight gain after consumption of high fat meals (29). In addition, glucose oxidation rates are significantly decreased in subjects after consuming high-fat meals (29). This literature suggests that the decreased MPC expression correlates with decreased diet-induced thermogenesis that occurs with prolonged high-fat feeding. Also, the sex-specific regulation of MPCs (Figure 6) is likely attributed to sex-specific metabolic responses to high fat feeding in C57BL/6J mice (21). Unlike male mice, previous studies have shown that female mice are more metabolically healthy, have reduced inflammation, and are more insulin sensitive despite having obesity while on HFD (20–22). Accordingly, in our study the female mice on HFD had 27% higher adiposity than the male HFD-fed mice, but improved glucose metabolism during an intraperitoneal glucose tolerance test. Data in Figure 7 demonstrate that decreased MPC protein levels correlate with increased adiposity and glucose intolerance in male, but not female, mice. We speculate that the variations in expression of MPC1 and MPC2 between male and female mice are due to differences in metabolic health and that MPCs are specifically regulated in brown, but not white, AT depots in conditions of metabolic dysfunction.

In summary, our findings demonstrate that (1) the expression of MPCs is induced during adipocyte development, but not required for adipogenesis, (2) a loss of MPC expression or activity during adipogenesis likely promotes the activation of alternative mechanisms to compensate for the loss of pyruvate transport, and (3) MPC expression is highly enriched in BAT but decreased as a result of high-fat feeding or diet-induced obesity. Future studies will be needed to determine which substrates can be utilized to compensate for the loss or dysfunction of MPCs. In addition, it is necessary to determine whether adipogenesis in BAT is dependent on MPCs. Also, future studies will be necessary to determine whether the observed in vitro effects on adipogenesis are similar to in vivo models.

Study Importance.

1. What is already known about this subject?

Mitochondrial pyruvate carriers are essential for pyruvate to be transported into the mitochondria and utilized as substrate by the pyruvate dehydrogenase complex (PDC) for energy production.

2. What does your study add?

This study is novel because it demonstrates that MPCs are not required for the adipogenesis of 3T3-L1 cells, suggesting that other substrates may compensate for energy production during in vitro adipogenesis.

This study is also novel because it shows that MPCs are regulated by diet-induced obesity in brown adipose tissue.

Acknowledgements

The authors would like to acknowledge Hardy Hang and Tamra Mendoza for help with the animal component of this study, and Timothy Allerton for assistance with correlation analyses.

Funding: This work was supported by National Institutes of Health Grant R01DK052968 to JMS. This research project utilized the Genomics Core facilities that are supported in part by COBRE (NIH8 1P30GM118430-02) and NORC (NIH 2P30DK072476) center grants from the National Institutes of Health.

Footnotes

Disclosure: The authors declare no conflict of interest.

References

  • 1.Bender T, Pena G, Martinou J-C. Regulation of mitochondrial pyruvate uptake by alternative pyruvate carrier complexes. EMBO J 2015;34:911–24. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Gray LR, Sultana MR, Rauckhorst AJ, et al. Hepatic mitochondrial pyruvate carrier 1 is required for efficient regulation of gluconeogenesis and whole-body glucose homeostasis. Cell Metab 2015;22:669–681. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Bricker DK, Taylor EB, Schell JC, et al. A mitochondrial pyruvate carrier required for pyruvate uptake in yeast, Drosophila, and humans. Science 2012;337:96–100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.McCommis KS, Finck BN. Mitochondrial pyruvate transport: a historical perspective and future research directions. Biochem J 2015;466:443–454. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Yang C, Ko B, Hensley CT, et al. Glutamine oxidation maintains the TCA cycle and cell survival during impaired mitochondrial pyruvate transport. Mol Cell 2014;56:414–24. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Sciacovelli M, Gaude E, Hilvo M, Frezza C. The Metabolic Alterations of Cancer Cells. In: Methods in enzymology.Vol 542, 2014, pp 1–23. [DOI] [PubMed] [Google Scholar]
  • 7.Divakaruni AS, Wiley SE, Rogers GW, et al. Thiazolidinediones are acute, specific inhibitors of the mitochondrial pyruvate carrier. Proc Natl Acad Sci U S A 2013;110:5422–5427. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Zou S, Zhu L, Huang K, Luo H, Xu W, He X. Adipose tissues of MPC1± mice display altered lipid metabolism-related enzyme expression levels. PeerJ 2018;6:1–18. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Wilson-Fritch L, Burkart A, Bell G, et al. Mitochondrial biogenesis and remodeling during adipogenesis and in response to the insulin sensitizer rosiglitazone. Mol Cell Biol 2003;23:1085–1094. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Yang C, Ko B, Hensley CT, et al. Glutamine oxidation maintains the TCA cycle and cell survival during impaired mitochondrial pyruvate transport. Mol Cell 2014;56:414–424. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Moyes CD, Mathieu-Costello OA, Tsuchiya N, Filburn C, Hansford RG. Mitochondrial biogenesis during cellular differentiation. Am J Physiol 1997;272:C1345–C1351. [DOI] [PubMed] [Google Scholar]
  • 12.Kadowaki T, Kitagawa Y. Enhanced transcription of mitochondrial genes after growth stimulation and glucocorticoid treatment of Reuber hepatoma H-35. FEBS Lett 1988;233:51–56. [DOI] [PubMed] [Google Scholar]
  • 13.Leary SC, Battersby BJ, Hansford RG, Moyes CD. Interactions between bioenergetics and mitochondrial biogenesis. Biochim Biophys Acta - Bioenerg 1998;1365:522–530. [DOI] [PubMed] [Google Scholar]
  • 14.De Pauw A, Tejerina S, Raes M, Keijer J, Arnould T. Mitochondrial (dys)function in adipocyte (de)differentiation and systemic metabolic alterations. Am J Pathol 2009;175:927–939. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Patterson JN, Cousteils K, Lou JW, Manning Fox JE, MacDonald PE, Joseph JW. Mitochondrial metabolism of pyruvate is essential for regulating glucose-stimulated insulin secretion. J Biol Chem 2014;289:13335–13346. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Richard AJ, Fuller S, Fedorcenco V, et al. Artemisia scoparia enhances adipocyte development and endocrine function in vitro and enhances insulin action in vivo. PLoS One 2014;9:e98897. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Colca JR, McDonald WG, Cavey GS, et al. Identification of a mitochondrial target of thiazolidinedione insulin sensitizers (mTOT)--relationship to newly identified mitochondrial pyruvate carrier proteins. PLoS One 2013;8:1–10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Hildyard JCW, Ämmälä C, Dukes ID, Thomson SA, Halestrap AP. Identification and characterisation of a new class of highly specific and potent inhibitors of the mitochondrial pyruvate carrier. Biochim Biophys Acta - Bioenerg 2005;1707:221–230. [DOI] [PubMed] [Google Scholar]
  • 19.Jensen-Urstad APL, Semenkovich CF. Fatty acid synthase and liver triglyceride metabolism: housekeeper or messenger? Biochim Biophys Acta 2012;1821:747–753. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Macotela Y, Boucher J, Tran TT, Kahn CR. Sex and depot differences in adipocyte insulin sensitivity and glucose metabolism. Diabetes 2009;58:803–82. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Heydemann A An overview of murine high fat diet as a model for Ttype 2 diabetes mellitus. J Diabetes Res 2016;2016:1–14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Morselli E, Criollo A, Rodriguez-Navas C, Clegg DJ. Chronic high fat diet consumption impairs metabolic health of male mice. Inflamm cell Signal 2014;1:1–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Schell JC, Wisidagama DR, Bensard C, et al. Control of intestinal stem cell function and proliferation by mitochondrial pyruvate metabolism. Nat Cell Biol 2017;19:1027–1036. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.McCommis KS, Chen Z, Fu X, et al. Loss of mitochondrial pyruvate carrier 2 in the liver leads to defects in gluconeogenesis and compensation via pyruvate-alanine cycling. Cell Metab 2015;22:682–694. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Li A, Liu Q, Li Q, Liu B, Yang Y, Zhang N. Berberine reduces pyruvate-driven hepatic glucose production by limiting mitochondrial import of pyruvate through mitochondrial pyruvate carrier 1. EBioMedicine 2018;34:243–255. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Green CR, Wallace M, Divakaruni AS, et al. Branched-chain amino acid catabolism fuels adipocyte differentiation and lipogenesis. Nat Chem Biol 2016;12:15–21. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Zou S, Lang T, Zhang B, et al. Fatty acid oxidation alleviates the energy deficiency caused by the loss of MPC1 in MPC1+/−mice. Biochem Biophys Res Commun 2018;495:1008–1013. [DOI] [PubMed] [Google Scholar]
  • 28.Perl K, Ushakov K, Pozniak Y, et al. Reduced changes in protein compared to mRNA levels across non-proliferating tissues. BMC Genomics 2017;18:305. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Tentolouris N, Pavlatos S, Kokkinos A, Perrea D, Pagoni S, Katsilambros N. Diet-induced thermogenesis and substrate oxidation are not different between lean and obese women after two different isocaloric meals, one rich in protein and one rich in fat. Metabolism 2008;57:313–320. [DOI] [PubMed] [Google Scholar]

RESOURCES