C2C12 myocytes lack an insulin-responsive vesicular compartment despite dexamethasone-induced GLUT4 expression

Lori L. Tortorella and Paul F. Pilch

Department of Biochemistry, Boston University School of Medicine, Boston, Massachusetts 02118


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Insulin regulates the uptake of glucose into skeletal muscle and adipocytes by redistributing the tissue-specific glucose transporter GLUT4 from intracellular vesicles to the cell surface. To date, GLUT4 is the only protein involved in insulin-regulated vesicular traffic that has this tissue distribution, thus raising the possibility that its expression alone may allow formation of an insulin-responsive vesicular compartment. We show here that treatment of differentiating C2C12 myoblasts with dexamethasone, acting via the glucocorticoid receptor, causes a >= 10-fold increase in GLUT4 expression but results in no significant change in insulin-stimulated glucose transport. Signaling from the insulin receptor to its target, Akt2, and expression of the soluble N-ethylmaleimide-sensitive factor-attachment protein receptor, or SNARE, proteins syntaxin 4 and vesicle-associated membrane protein are normal in dexamethasone-treated C2C12 cells. However, these cells show no insulin-dependent trafficking of the insulin-responsive aminopeptidase or the transferrin receptor, respective markers for intracellular GLUT4-rich compartments and endosomes that are insulin responsive in mature muscle and adipose cells. Therefore, these data support the hypothesis that GLUT4 expression by itself is insufficient to establish an insulin-sensitive vesicular compartment.

glucocorticoid; skeletal muscle; insulin; glucose transporter; C2C12 cells; protein trafficking


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

INSULIN PROMOTES the postprandial clearance of glucose from the blood primarily into skeletal muscle in both humans and rodents. This process entails the insulin-dependent movement or recruitment of the GLUT4 glucose transporter isoform from intracellular storage vesicles to the cell surface, where they function (42). Much remains unclear about how the insulin-dependent signal transduction pathway communicates with the intracellular, GLUT4-rich vesicles, and the exact nature of the compartments through which GLUT4 traffics is also not completely understood. Several lines of independent experimentation support the notion that insulin-dependent regulation of GLUT4 movement is similar or identical in skeletal muscle and adipocytes (24, 43). However, GLUT4 translocates in response to exercise and hypoxia in the former and not the latter tissue (16, 47). Insulin-dependent GLUT4 translocation is phosphatidylinositol (PI)3-kinase dependent in both tissues, whereas the exercise-dependent process is not (9, 57). An exercise (or hypoxia)-induced rise in AMP results in activation of the AMP-activated protein kinase (AMPK), which correlates with increased glucose transport and GLUT4 translocation (20, 29). Exposure of skeletal muscle to 5-aminoimidazole-4-carboxamide-1-beta -D-ribofuranoside, or AICAR, an AMP analog that activates AMPK, can recapitulate this effect (29, 37). Recently, transgenic animals expressing a dominant negative AMPK construct were shown not to respond to hypoxia and to respond less to exercise with regard to glucose transport and GLUT4 translocation (40). Thus there may be three independent pathways to promote glucose uptake into muscle, mediated respectively by insulin, AMPK, and unknown factors.

Much of our current understanding of GLUT4 vesicle translocation comes from experiments performed with adipocytes, where, as with cardiomyocytes and skeletal muscle, GLUT4 is expressed and undergoes vesicular translocation in response to insulin. Adipocytes have been employed because the regulation of glucose transport is important in this tissue for triglyceride storage (44). In addition, glucose uptake into this tissue affects fuel homeostasis in the whole organism, as shown recently with adipose-specific GLUT4 knockout mice (1). These mice have impaired insulin-sensitive glucose uptake in adipocytes but also exhibit insulin resistance in muscle and liver and develop glucose intolerance and hyperinsulinemia. Finally, there are long-standing and well characterized methods for the subcellular fractionation of fat cells that make them a particularly useful experimental model (51).

Overall, it is the inability of insulin to promote glucose uptake into skeletal muscle that causes insulin resistance and, eventually, type 2 diabetes, and thus it is very important to characterize GLUT4 trafficking in this tissue. Subcellular fractionation methods exist for muscle (11, 59), but they yield less precise and clean fractions because of interference from the muscle fibers and the complex T-tubule network, even though recent advances have been made in this regard (59). Because the composition of intact skeletal muscle makes it less than ideal for fractionation studies, a suitable cell line with appropriate properties of this tissue could not only allow improved fractionation procedures but could offer other experimental advantages as well. Indeed, several muscle cell lines have been described (L6, C2C12, Sol8), but they express far less GLUT4 than muscle tissue, and their insulin responsiveness is minimal. This is true even when GLUT4 levels are artificially raised by transfection (28), either because they lack the machinery to mediate GLUT4 sequestration and translocation or because the expression of endogenous GLUT1 and GLUT3 masks insulin effects on GLUT4 (18, 53).

Recently, it was shown that glucocorticoids (49, 58) and insulin-like growth factor I (IGF-I) (41, 49) cause "hypertrophy" of C2C12 muscle cells, and we thought that the increased metabolism required by hypertrophy might involve enhanced GLUT4 expression. We show here that glucocorticoids do indeed cause markedly enhanced GLUT4 expression, but they cause no increase in insulin-dependent glucose transport. Our data support the likelihood that this is due to failure of the C2C12 cells to elaborate the necessary GLUT4-containing vesicular compartments.


    MATERIALS AND METHODS
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Cell culture. C2C12 (56) and L6 (55) myoblast cells were maintained at subconfluent conditions in growth media containing DMEM with 4.5 g/l glucose, 100 U/ml penicillin, 100 µg/ml streptomycin, and 20% fetal bovine serum (Sigma Chemical, St Louis, MO). Near-confluent cells (~80% confluence) were differentiated by lowering the serum concentration to 2% calf serum (Sigma). Cells were maintained for 3-7 days to obtain myotubes. The 3T3-L1 cells (17) were grown to confluence in 10% calf serum. Two days postconfluence, the cells were switched to induction medium (10% fetal bovine serum, 1 µM dexamethasone, 100 nM insulin, and 50 µM IBMX in DMEM) for 2 days, after which they were maintained in 10% fetal bovine serum until use on days 6-8 of differentiation. All cell lines were grown in a humidified, 37°C incubator with ambient oxygen and 5% CO2.

Subcellular fractionation of C2C12 cells. Cells were washed three times on ice with PBS, pH 7.4 (in mM: 137 NaCl, 2.7 KCl, 10 Na2HPO4, and 1.8 KH2PO4) and then homogenized with 30-40 strokes of a Kontes glass tissue grinder (no. 885451-0021) in HES buffer [255 mM sucrose, 4 mM disodium EDTA, 20 mM HEPES pH 7.4, 10 µM leupeptin, 1 µM pepstatin, 1 µM aprotinin, 1 mM phenylmethylsulfonyl fluoride (PMSF), and 5 mM benzamidine]. The homogenate was centrifuged at 19,000 g (16 K in Ti70 rotor) for 20 min. The pellet was saved and fractionated further to crude "plasma membrane" (P1) and crude nuclear (N)/endoplasmic reticulum (ER) fractions. The supernatant was centrifuged at 40,000 g (23 K in Ti70 rotor) for 20 min. The pellet (P2) was resuspended in PBS plus protease inhibitors. The supernatant was centrifuged at 180,000 g (48 K in Ti70 rotor) for 1.5 h. The pellet (P3) was resuspended in PBS plus protease inhibitors. The supernatant of the 180,000-g spin contains the cytosol. The P2 and P3 fractions most likely correspond to high- and low-density microsomes. To obtain the plasma membrane (PM) fraction, the pellet from the first spin was resuspended in HES, layered onto a 1.12 M sucrose cushion in 20 mM HEPES and 1 mM disodium EDTA, and centrifuged at 100,000 g (27 K in an AH627 rotor) for 1 h. The pellet (N/ER-containing fraction) was resuspended in a buffer containing 20 mM Tris (pH 7.4), 50 mM NaCl, 2% Nonidet P-40, 0.5% deoxycholate, 0.2% SDS, and the protease inhibitor cocktail. The interphase of the sucrose cushion was collected and pelleted at 40,000 g (23 K in a Ti70 rotor) for 20 min. This PM-containing pellet (P1) was resuspended in PBS plus protease inhibitors. All centrifugations were performed at 4°C with a Sorvall Ultraspeed centrifuge (no. OTD70B).

Labeling with sulfo-NHS-SS-biotin. Cells were washed three times in Krebs-Ringer phosphate (KRP) buffer (in mM: 128 NaCl, 4.7 KCl, 1.25 CaCl2, 1.25 MgSO4, 5 Na2HPO4, and 20 HEPES, pH 7.4) and then incubated with 1 mM EZ Link sulfo-NHS-SS-biotin (Pierce, Rockford, IL) in KRP buffer at 37°C for the indicated time. The reaction was quenched by washing the cells twice with 20 mM Tris (pH 7.4), 25 mM ethanolamine, and 150 mM NaCl. After the reaction was quenched, the cells were incubated with 2 mM N-ethylmaleimide for 8 min to prevent reduction of the disulfides in the biotinylated proteins. Cells were lysed in 1% Triton X-100, 150 mM NaCl, 20 mM Tris (pH 7.4), 10 µM leupeptin, 1 µM pepstatin, 1 µM aprotinin, and 1 mM PMSF, and equal amounts were incubated with Agarose Immobilized Streptavidin (Pierce) at 4°C for 16 h. Streptavidin-agarose complexes were washed four times in lysis buffer, eluted in Laemmli SDS sample buffer containing 5% beta -mercaptoethanol, and then analyzed by Western blotting.

Gel electrophoresis and Western blotting. Proteins were separated by SDS-PAGE (30) and then transferred to a 0.2-µm polyvinylidene difluoride membrane by use of a buffer containing 25 mM Tris and 192 mM glycine at 4°C for 1,600 mAmp/h. Membranes were blocked in 10% nonfat milk in PBS-T (PBS plus 0.1% Tween 20) before incubation for 1 h at room temperature with primary antibody. Membranes were then washed in PBS-T and incubated with horseradish peroxidase-conjugated secondary antibodies (Sigma). Protein bands were detected using enhanced chemiluminescent reagents (NEN Life Sciences, Boston, MA). The antibodies used in the present study were obtained from the following sources: anti-myosin heavy chain (MHC, MF20) and anti-myogenin (F5D) antibodies from the Developmental Studies Hybridoma Bank, University of Iowa; anti-transferrin receptor (TfR) antibody from Zymed Laboratory, San Francisco, CA; anti-glucocorticoid receptor antibody from Affinity Bioreagents, Golden, CO; anti-insulin receptor antibody from Transduction Laboratories, Lexington KY; anti-insulin receptor substrate-1 (IRS-1) and p85 PI 3-kinase antibodies from Upstate Biotechnology, Lake Placid, NY; anti-phospho-Akt and insulin-responsive aminopeptidase (IRAP) antibodies from Quality Control Biochemicals/Biosource; and anti-GLUT4 antibody, 1F8 (23). The following antibodies were obtained as gifts: anti-GLUT 1 antibody from Dr. Christin Carter-Su, University of Michigan; anti-Akt2 antibody from Dr. Morris Birnbaum, University of Pennsylvania; anti-protein disulfide isomerase (PDI) and Sec61 alpha -antibodies from Dr. Hidde Ploegh, Harvard Medical School; and anti-beta 1-integrin antibody from Dr. Carlos Enrich, Universitat de Barcelona.

Northern blotting. RNA was obtained as described by Chomczynski and Sacchi (6). Cells were rinsed three times with PBS, lysed in solution D [4 M guanidinium isothiocyanate, 25 mM sodium citrate (pH 7.0), 0.5% sarkosyl, and 0.1 M beta -mercaptoethanol], followed by acid phenol (pH 4.2)-chloroform extraction. Total RNA was precipitated in isopropanol at -20°C, followed by centrifugation at 10,000 g at 4°C for 20 min. Equal amounts (20 µg) of RNA were separated on a 1% agarose-6% formaldehyde gel and transferred to Genescreen nylon membrane (NEN Life Sciences) by capillary action with 10× standard sodium citrate (SSC: 1.5 M NaCl and 0.15 M sodium citrate pH 7.0) as the liquid phase. The RNA was cross-linked to the membrane using the ultraviolet (UV) Stratalinker (Stratagene, La Jolla, CA). RNA was checked for equal loading and transfer by UV visualization of ethidium bromide rRNA staining. Prehybridization of nylon membranes was for 4-6 h at 42°C in 50% formamide, 4× SSC, 5× Denhardt's solution (0.1% Ficoll, 0.1% polyvinyl pyrrolidone, and 0.1% BSA), 0.05 M sodium phosphate (pH 7.0), 0.5 mg/ml sodium pyrophosphate, 1% SDS, and 0.1 mg/ml tRNA carrier. Hybridization of the blot was for 16-20 h at 42°C in 50% formamide, 4× SSC, 1× Denhardt's solution, 0.05 M sodium phosphate (pH 7.0), 0.5 mg/ml sodium pyrophosphate, 1% SDS, 0.1 mg/ml tRNA carrier, and the labeled probe. The cDNA inserts used for probes were as follows: GLUT4, GenBank accession no. M23383; GLUT1, from G. Bell; IRAP, GenBank accession no. U32990; hexokinase (HK) II, GenBank accession no. M68971; myogenin, (see Ref. 54), MHC (see Ref. 33), and myocyte-enhancer factor 2C (MEF2C) (see Ref. 34). The cDNA probes were labeled with [32alpha P]dATP by random priming (13) and purified with a NucTrap column (Stratagene). The specific activity of the probes was >= 108 cpm. Blots were washed in a stepwise gradient at moderate stringency (0.2× SSC-0.1% SDS at 42°C) and exposed to film at -80°C.

2-Deoxy-[3H]glucose uptake assay. Cells were serum starved in DMEM for 2 h at 37°C and then stimulated with or without 100 nM insulin for 15 min at 37°C. Cells were washed twice with Krebs-Ringer-Henseleit (KRH) buffer [in mM: 121 NaCl, 4.9 KCl, 1.2 MgSO4, 0.33 CaCl2, and 12 HEPES (pH 7.4)] before addition of 2-deoxy-[3H]glucose at 1 µCi/ml in KRH buffer. 2-Deoxy-[3H]glucose was added to the cells in the presence or absence of 5 µM cytochalasin B to determine nonspecific uptake. Cells were incubated for 5 min at room temperature with tritiated glucose before chasing and washing with excess unlabeled 2-deoxy-D-glucose in KRH buffer. Cells were lysed in 0.25 M mannitol, 17 mM MOPS, 2.5 mM EDTA, and 8 mg/ml digitonin. The sample was then counted using a scintillation counter, or protein concentration was determined using the Bradford assay. Data are presented as picomoles of 2-deoxy-[3H]glucose per milligram protein per minute.


    RESULTS
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INTRODUCTION
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RESULTS
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Dexamethasone dramatically enhances GLUT4 expression in C2C12 myotubes. Previous studies in C2C12 myocytes showed that dexamethasone exposure results in their hypertrophy (49, 58) and thus may require more metabolic substrates, i.e., glucose. To determine whether this glucocorticoid could affect expression of GLUT4 in these cells, Northern blot analysis was performed. Total mRNA was isolated from C2C12 cells at the indicated time during the differentiation process, and the expression of GLUT4, GLUT1, IRAP, a protein that colocalizes with GLUT4 in insulin-sensitive vesicles (26), and various differentiation markers were examined (Fig. 1). GLUT4 mRNA is minimal in cultured myoblasts and, consistent with previous reports (28, 39), its expression is induced as differentiation progresses. Furthermore, dexamethasone-treated cells exhibit markedly enhanced GLUT4 mRNA levels beginning on day 4 of differentiation. Levels of IRAP rise during the differentiation of untreated cells, and the timing of its expression is advanced upon addition of dexamethasone. The expression of mRNA for the muscle-specific transcription factors, myogenin and MEF2C, and for the ubiquitously expressed glucose transporter GLUT1 is not altered with dexamethasone treatment. Interestingly, hexokinase II, a muscle/fat-specific enzyme involved in the glycolytic pathway (31), is not affected by dexamethasone, suggesting that dexamethasone may not affect all the molecules involved in glucose metabolism but perhaps may regulate the proteins involved in insulin-sensitive glucose uptake. Moreover, these results indicate that dexamethasone acts at the transcriptional (RNA) level in C2C12 cells and is specific for the regulation of a set of genes rather than inducing a general increase in transcription.


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Fig. 1.   Dexamethasone induces GLUT4 mRNA expression in C2C12 myocytes. A: C2C12 myoblasts were differentiated in 2% calf serum (CS) with or without 1 µM dexamethasone. Total RNA was collected by acid phenol-chloroform extraction on the indicated day. Equal amounts (20 µg) of RNA were separated and analyzed by Northern blotting with 32P-labeled probes identified at right. IRAP, insulin-responsive aminopeptidase; MEF2C, myocyte-enhancer factor 2C. Ethidium bromide staining of the rRNA shows equal loading. The experiment was performed twice with essentially identical results. B: densitometric analysis of select blots from A with a Molecular Dynamics densitometer (Amersham Pharmacia Biotech, Piscataway, NJ) with MD ImageQuant Software version 3.2.

To confirm that the increase in GLUT4 mRNA results in a concomitant increase in protein level, the expression of the GLUT1 and GLUT4 isoforms was examined in C2C12 myoblasts that were differentiated in the presence or absence of dexamethasone. On day 7, the total particulate fraction was collected and analyzed by Western blot for the indicated proteins (Fig. 2). The expression of GLUT4 is increased ~10-fold in treated vs. untreated cells, whereas the overall expression of GLUT1 remains unchanged, although its glycosylation appears altered compared with untreated cells. The expression of IRAP is enhanced with dexamethasone treatment. IRAP is also a glycoprotein, and its carbohydrate modification appears unchanged with glucocorticoid treatment. The muscle differentiation markers myogenin and MHC do not show an increase in expression, suggesting specificity with respect to the induction by dexamethasone.


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Fig. 2.   Dexamethasone-treated C2C12 cells express elevated GLUT4 and IRAP proteins. C2C12 myoblasts were differentiated in 2% CS with or without 1 µM dexamethasone. On day 7, cells were homogenized in HES buffer, pH 7.4, and a total particulate fraction was collected by centrifugation at 180,000 g. Equal amounts (100 µg) of protein were separated by SDS-PAGE and analyzed by Western blotting for the indicated proteins. MHC, myosin heavy chain. The experiment was performed 5 times with similar results.

Dexamethasone regulates gene expression through activation of its intracellular target, the glucocorticoid receptor. The expression of the glucocorticoid receptor was examined in C2C12 myotubes by Western blot analysis. The glucocorticoid receptor is expressed in C2C12 myocytes, and its level is similar in both dexamethasone-treated and untreated cells (Fig. 3A). To determine whether dexamethasone induces GLUT4 expression via the glucocorticoid receptor, C2C12 cells were differentiated in the presence or absence of dexamethasone and/or RU-486, an antagonist of the glucocorticoid receptor (3). Then, the expression of GLUT4 was determined by Northern blot (Fig. 3B). Increasing concentrations of RU-486 are able to completely inhibit dexamethasone-induced GLUT4 expression. This repression is observed at the mRNA level, suggesting that the glucocorticoid receptor acts by regulating gene transcription. These results do not indicate whether or not the glucocorticoid receptor is acting directly on the GLUT4 promoter. However, no obvious glucocorticoid-responsive binding sites were found upon analysis of published promoter sequences in rat and mouse [GenBank accession nos. L36125 (rat), M29960 (mouse)], although a potential glucocorticoid receptor-binding element was found in one of two described human GLUT4 promoter sequences [GenBank accession nos. X58489 (human), and M61126 (human)].


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Fig. 3.   The dexamethasone effect on GLUT4 expression is dose dependent and is mediated by the glucocorticoid receptor. A: C2C12 myoblasts were differentiated in 2% CS with or without 1 µM dexamethasone, and total protein was collected on day 4. Equal amounts (100 µg) were separated by SDS-PAGE and analyzed by Western blot by use of an anti-glucocorticoid receptor antibody. B: C2C12 myoblasts were differentiated in 2% CS with or without 1 µM dexamethasone and with 10 nM, or 1 or 10 µM RU-486. Total RNA was harvested on day 3, and equal amounts (20 µg) were separated and analyzed by Northern blotting with a 32P-labeled GLUT4 cDNA probe. Ethidium bromide staining of the rRNA shows equal loading. C: C2C12 myoblasts were differentiated in 2% CS in the presence or absence of the indicated concentration of dexamethasone. Total RNA was harvested on day 3, and equal amounts (20 µg) were separated and analyzed by Northern blotting with a 32P-labeled GLUT4 cDNA probe. Ethidium bromide staining of the rRNA shows equal loading. Experiments were performed twice with similar results.

To further characterize dexamethasone's effect, C2C12 cells were differentiated in the absence or presence of the indicated concentration of dexamethasone, and GLUT4 expression was analyzed by Northern blot (Fig. 3C). A dose-dependent effect on GLUT4 expression is observed with dexamethasone treatment, and the maximum GLUT4 expression is achieved at a concentration between 10 and 1,000 nM.

The amount of GLUT4 protein in dexamethasone-treated C2C12 cells was compared with that in other cell lines presently used to study GLUT4 trafficking, including 3T3-L1 adipocytes and L6 myocytes (Fig. 4). The levels of GLUT4 and IRAP detected in 3T3-L1 and glucocorticoid-treated C2C12 cells are comparable, with ~1.5-fold more GLUT4 in the adipocytes than in the treated myocytes. Interestingly, L6 myocytes express appreciable levels of GLUT4, but exposure to dexamethasone greatly diminishes GLUT4 expression under the condition used.


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Fig. 4.   Comparison of GLUT4 expression in C2C12, L6, and 3T3-L1 cells. C2C12 and L6 myoblasts were differentiated in the presence or absence of 1 µM dexamethasone. A total membrane fraction was collected from these or from 3T3-L1 adipocytes, and 100 µg of each were separated by SDS-PAGE. After transfer, analysis was by Western blot using anti-GLUT4 or IRAP antibodies. Two independent experiments were performed and yielded similar results.

Dexamethasone-treated C2C12 myocytes do not exhibit insulin-sensitive glucose uptake, even though they have increased levels of GLUT4 protein. We determined whether dexamethasone-treated cells gain insulin-sensitive glucose uptake as a result of increased GLUT4 expression. C2C12 myoblasts and untreated or dexamethasone-treated myocytes were analyzed for uptake of labeled 2-deoxyglucose after insulin stimulation (Fig. 5). No significant increase in labeled glucose tracer is observed in response to insulin in myoblasts or differentiated cells, regardless of dexamethasone treatment. The uptake that is observed in these cells is most likely mediated by GLUT1, GLUT3, or cell surface GLUT4, because nontransporter-mediated uptake is negligible (see bars labeled cyto B in Fig. 5). These data suggest that C2C12 cells do not gain an insulin-sensitive phenotype upon differentiation, even with the dramatic increase in GLUT4 expression caused by dexamethasone. Reasons that the C2C12 myocytes may not exhibit insulin-sensitive glucose uptake include the lack of insulin-sensitive vesicles containing GLUT4 or a deficiency in insulin signaling.


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Fig. 5.   GLUT4 expression does not lead to insulin-sensitive glucose uptake in dexamethasone-treated C2C12 cells. C2C12 myoblasts or myotubes (day 4) that were differentiated in 2% CS with or without 1 µM dexamethasone were serum starved for 2 h and then stimulated with 100 nM insulin for 15 min at 37°C. 2-Deoxy-[3H]glucose was added to a final concentration of 1 µCi/ml with or without 5 µM cytochalasin B (cyto B) and incubated for 5 min at room temperature. Uptake of labeled tracer was terminated by addition of excess unlabeled glucose. Cells were lysed in digitonin buffer, and the amount of 2-deoxy-[3H]glucose uptake was quantitated by scintillation counting. Data were normalized to protein content at each time point and are expressed as pmol 2-deoxy-[3H]glucose per mg protein per min. (-)/(+), insulin treatment. Data are representative of 3 independent experiments.

To verify whether or not insulin signaling was functional in dexamethasone-treated C2C12 myotubes, the expression of molecules involved in this signal transduction pathway was analyzed by Western blot. The expression of insulin receptor, IRS-1, PI 3-kinase, and Akt2 is detected in both untreated and dexamethasone-treated cells (Fig. 6A), although less insulin receptor and IRS-1 are observed after dexamethasone treatment. The activation of the signaling cascade was tested by Western blotting for the activated form of Akt2 by use of an antibody that recognizes the Ser473 phosphorylated form. Cells treated as described for Fig. 6A were serum starved and then stimulated with insulin for 10 min. Insulin stimulates the phosphorylation of Akt2 in both dexamethasone-treated and untreated myocytes, and little phosphorylated Akt is detected in cells not treated with insulin (Fig. 6B). These results suggest that insulin activates a signaling pathway in dexamethasone-treated C2C12 cells that would normally cause GLUT4 translocation (5), and therefore a defective insulin signal is not responsible for the lack of glucose uptake.


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Fig. 6.   Proteins involved in insulin-dependent signaling and GLUT4 vesicle targeting are expressed in dexamethasone-treated C2C12 myocytes. A and C: C2C12 myoblasts were differentiated in 2% CS with or without 1 µM dexamethasone, and total protein was collected on day 4 or 5. Equal amounts (100 µg) were separated by SDS-PAGE and analyzed by Western blot using the indicated antibodies. B: C2C12 cells treated as in A were starved for 2 h in serum-free media, and then 100 nM insulin was added for 7 min at 37°C. Total protein was collected, separated by SDS-PAGE, and analyzed by Western blot with anti-Akt2 or anti-phosho-Akt (Ser473) antibodies. Data are representative of 2 independent experiments.

The trafficking of vesicles to distinct subcellular destinations is mediated in part by the interaction of specific targeting proteins on the vesicle with their cognate target membrane receptors (V- and T- SNARE proteins) (15, 19). VAMP2 and syntaxin4 are the vesicle and target membrane proteins known to be important for directing GLUT4 to the plasma membrane (4, 19). To verify whether or not these proteins are present in dexamethasone-treated C2C12 myotubes, their expression was analyzed by Western blot. VAMP2 and syntaxin4 are detected at similar levels in untreated and dexamethasone-treated cells (Fig. 6C). These data suggest that the appropriate SNAREs are present in C2C12 myocytes, and thus a defect in vesicle targeting is likely not the reason for the lack of glucose uptake in response to insulin.

Insulin's inability to stimulate glucose uptake in dexamethasone-treated C2C12 cells may result from lack of an insulin-sensitive vesicular compartment or from mistargeting of GLUT4 to this compartment. To test these possibilities, the subcellular localization of GLUT4 was investigated in dexamethasone-treated cells that were or were not stimulated with insulin. Cells were homogenized and then fractionated by differential centrifugation into five relatively crude subcellular fractions, including the P1, P2, and P3 membrane fractions (see below), cytosol, and the N/E-containing fraction. Each fraction was analyzed by Western blot for GLUT4 and IRAP content, as well as for various organelle markers (Fig. 7A). The majority of the ER resident proteins Sec61alpha (45) and PDI (32) were detected in the N/E fraction. The nuclei and ER copurify in the same low-speed fraction (N/E), probably because the continuity of the ER membrane and nuclear envelope is not disrupted by the homogenization step. The pellet P1 contains the PM, as marked by the localization of beta 1-integrin, and likely contains cytoskeletal-associated fractions. The P2 and P3 fractions contain intracellular membrane compartments and likely correspond to the heavy (Golgi) and light (vesicles) microsome fraction, respectively. GLUT1 and the TfR are found in all membrane-containing compartments, and this distribution is consistent with that of proteins that constitutively traffic through the endosomal pathway. On the other hand, GLUT4 and IRAP colocalize to the same cellular compartments, primarily the P1 and P2 fractions, with 84% of the total GLUT4 found in these fractions (Fig. 7, A and B). Interestingly, less than 3% of the GLUT4 is found in the P3 fraction. Trafficking of GLUT4 from intracellular compartments to the PM is apparently absent in dexamethasone-treated cells, since there is no significant change in the percentage of GLUT4 in any subcellular fraction after insulin stimulation (Fig. 7, A and B). In addition, GLUT4 was not detected in light microsome fractions that were separated by sucrose gradient centrifugation (data not shown). Moreover, light microsomes from C2C12 myocytes were immunoadsorbed by use of the anti-GLUT4 antibody 1F8, and, again, vesicles containing GLUT4 could not be isolated (data not shown). Thus it is likely that a vesicle population containing GLUT4 is lacking in C2C12 myocytes.


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Fig. 7.   GLUT4 is does not traffic in response to insulin in dexamethasone-treated C2C12 cells. C2C12 myoblasts were differentiated in 2% CS in the presence of 1 µM dexamethasone. Cells were serum starved for 2 h in DMEM and stimulated with 100 nM insulin for 15 min at 37°C. Then subcellular fractions were collected, as described in MATERIALS AND METHODS. Equal amounts of protein from the resulting fractions were separated by SDS-PAGE and then analyzed by Western blot with the indicated antibodies. TfR, transferrin receptor; PDI, protein disulfide isomerase; P1, crude plasma membrane; N/E, nuclei and endoplasmic reticulum; cyto, cytosol; P2, high-density microsomes; P3, low-density microsomes. The experiment was performed 4 times with similar results.

Although the majority of the GLUT4 is found in the P1 and P2 fractions in dexamethasone-treated cells as determined by fractionation protocols, it is possible that an insulin-sensitive vesicle pool is present and the GLUT4 could be mistargeted. In skeletal muscle and fat, vesicular transport in response to insulin leads not only to GLUT4 transit to the cell surface, but other proteins are targeted there as well, including IRAP and TfR (25). Therefore, to determine whether C2C12 cells exhibit insulin-sensitive protein transport, cells were stimulated or not with insulin, and then cell surface proteins were labeled with membrane-impermeable sulfo-NHS-SS-biotin reagent. Streptavidin-conjugated agarose was used to separate biotin-labeled proteins from those unlabeled, and the avidin-associated complexes were analyzed by Western blot with anti-IRAP and TfR antibodies (Fig. 8). C2C12 myoblasts exhibit a small but reproducible translocation of both IRAP and TfR upon insulin addition. On the other hand, C2C12 myocytes that were differentiated with or without dexamethasone do not have any discernible amount of IRAP or TfR trafficking to the cell surface in response to insulin. These data suggest that C2C12 myocytes do not intrinsically contain an insulin-sensitive compartment, and this is independent of dexamethasone treatment.


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Fig. 8.   Dexamethasone-treated cells do not contain an insulin-sensitive vesicle compartment. C2C12 myoblasts were differentiated in 2% CS with or without 1 µM dexamethasone. Cells were serum starved for 2 h in DMEM and stimulated or not with 100 nM insulin for 15 min at 37°C. Cell surface proteins were labeled for 2 or 10 min at 37°C with 1 mM sulfo-NHS-SS-biotin. After the biotinylation reaction was quenched with 20 mM Tris, pH 7.4, and 25 mM ethanolamine, cells were lysed in buffer containing 1% Triton X-100. Equal amounts of protein were incubated with streptavidin-conjugated agarose beads, and biotinylated proteins were eluted with SDS sample buffer containing 5% beta -mercaptoethanol and were separated by SDS-PAGE. Analysis of recovered proteins was by Western blot with anti-IRAP and anti-TfR antibodies. The experiment was performed 3 times with similar results.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The trafficking of GLUT4 between intracellular vesicular compartments and the plasma membrane is a highly regulated process. Several proteins besides GLUT4 translocate to the plasma membrane in response to insulin, including IRAP and TfR (25). These vesicle proteins were first identified as such in adipocytes, and much of the work done thus far toward understanding the mechanism of insulin-sensitive protein traffic has been performed using adipocyte models. Both adipocytes and striated muscles exhibit insulin-sensitive trafficking and therefore may utilize a similar mechanism in this regard. In addition, muscle fibers translocate GLUT4 in response to contraction, and data suggest that separate GLUT4 vesicle pools may exist, one being sensitive to insulin and another responsive to contraction (7, 27). Therefore, multiple signaling mechanisms may be important for translocation of distinct GLUT4 vesicle populations under different physiological states. Even though these physiological differences between adipose and muscle tissues exist and skeletal muscle is the major depot for glucose in the postprandial state, adipocytes (in vitro and in vivo) have been studied the most thus far. A skeletal muscle cell line that could be used to study GLUT4 translocation by insulin and other stimuli could provide new insights as to other regulatory mechanisms and/or novel proteins involved in this process.

Previously, our laboratory has ectopically expressed GLUT4 in C2C12 myoblasts, but its overexpression did not result in the recapitulation of regulated GLUT4 trafficking (28). Using a denervation model of insulin resistance, our laboratory has shown that loss of GLUT4 expression, per se, in skeletal muscle does not diminish insulin-sensitive vesicle trafficking (60). These results suggest that GLUT4 itself is not responsible for vesicle formation, and other factors must be involved in both vesicle biogenesis and movement. This is in contrast to results reported for L6 cells overexpressing myc-tagged GLUT4 (L6-Glut4-myc), where its expression alone was postulated to confer insulin-sensitive trafficking of GLUT4 vesicles in both myoblasts and myotubes. In L6-Glut4-myc myoblasts, GLUT4 was present in both PM and intracellular compartments, while it was localized intracellularly in L6-Glut4-myc myotubes. In both L6-Glut4-myc myoblasts and myotubes, insulin stimulated GLUT4 translocation, which suggests that L6 cells may contain insulin-responsive vesicles regardless of whether or not they are differentiated, in contrast to skeletal muscle in vivo, where GLUT4 expression is regulated during development (48). The overexpression of myc-tagged GLUT4 in L6 cells results in increased insulin-sensitive glucose uptake in both myoblasts and myotubes, albeit only a 1.5-fold induction (52), whereas overexpression of untagged GLUT4 in L6 cells results in a 6-fold induction (46). In the latter case, only a small portion of GLUT4 was translocated, unlike in vivo, where 50% or more of the intracellular pool can translocate (59, 60). The differences in these two L6 transfection experiments may arise from clonal differences between cells from each laboratory or from variation intrinsic to the cDNAs used (tagged vs. untagged). With the exception of the studies in L6 cells, the ectopic expression of GLUT4 results in its intracellular sequestration in an incompletely characterized compartment(s) without any gain in insulin-sensitive glucose uptake, as shown previously in NIH-3T3 fibroblasts (21), Chinese hamster ovary cells (50), 3T3-L1 fibroblasts (50), and C2C12 myoblasts (28). The questions are whether GLUT4 expression is responsible for the formation of insulin-sensitive vesicles and what proteins specifically regulate their formation and trafficking.

Dexamethasone enhances GLUT4 and IRAP expression in C2C12 myocytes, and we tested whether this was a useful in vitro skeletal muscle cell line for studying insulin-sensitive trafficking. Although dexamethasone induces the expression of GLUT4 >= 10-fold in C2C12 myocytes, its expression does not convert the cells to an insulin-sensitive phenotype. Several reports have shown that dexamethasone induces insulin resistance in rat skeletal muscle by inhibiting translocation of GLUT4 to the plasma membrane (10) or by recruitment of GLUT4 to this locale in the basal state (8). There are several possible reasons why dexamethasone-treated C2C12 cells do not recruit GLUT4 to the PM in a regulated fashion. We have ruled out the possibility that insulin signaling is deficient in dexamethasone-treated C2C12 cells, since the pertinent signaling molecules involved are expressed, and Akt2, a downstream target of insulin receptor signaling that has been implicated in GLUT4 translocation in skeletal muscle (5), is activated with insulin stimulation.

The data presented suggest that GLUT4 is targeted to an insulin-insensitive compartment in dexamethasone-treated C2C12 cells. One conclusion from the fractionation data is that GLUT4 does not move to an appreciable extent from one membrane fraction to another with insulin treatment. It is difficult with subcellular fractionation alone to interpret the exact localization of GLUT4. We further analyzed GLUT4 localization in dexamethasone-treated C2C12 myocytes by immunofluorescent microscopy, and it appears that GLUT4 is localized intracellularly regardless of insulin stimulation (data not shown). Similar results were observed previously in cells ectopically expressing GLUT4 (21, 28, 50). Therefore, it is plausible that GLUT4 is targeted to an insulin-insensitive compartment in dexamethasone-treated C2C12 cells. Moreover, the amount of glucose uptake is the same in myoblasts, untreated myotubes, and dexamethasone-treated myotubes, even though the dexamethasone-treated myotubes express 10-fold more GLUT4 than the two aforementioned groups, and >50% of the GLUT4 is in the P1 fraction, which corresponds, in part, to the PM. These data suggest that, if GLUT4 is present at the cell surface, either it is not actively transporting glucose or its activity could be masked by the action of GLUT1 and/or GLUT3, which are also expressed in skeletal muscle cell lines. Moreover, dexamethasone-treated or untreated C2C12 myocytes do not appear to contain an insulin-sensitive vesicle compartment, since the cell surface labeling of IRAP and TfR does not increase with insulin stimulation. We were unable to test directly by cell-surface biotinylation whether or not GLUT4 is at the cell surface. GLUT4 contains on its exofacial loops only one lysine that lies in close proximity to the transmembrane region and therefore is not easily accessible to labeling with the biotin reagent we employed. Overall, these results provide further evidence that GLUT4 does not contain information in its primary sequence that aids in the formation of an insulin-sensitive compartment.

GLUT4 is a glycoprotein, and its expression in C2C12 cells is not detected as a single band by SDS-PAGE but rather as a group of closely migrating forms that collapse to one ~38-kDa band after endoglycosidase F digestion (data not shown). This glycosylation pattern is unlike the single GLUT4 band we observe in 3T3-L1 adipocytes, where it normally traffics in response to insulin. GLUT4 contains only one possible N-glycosylation site in its first extracellular loop; therefore, extensive variation in the carbohydrate processing is responsible for the multiple forms seen in C2C12 cells. It is known that proper processing is important for protein folding, stability, sorting, and function (14), and it is possible that the aberrant glycosylation of GLUT4 is nonpermissive regarding its correct targeting or function. Mutation of the N-glycosylation site in GLUT4 (Asn57right-arrowGln) resulted in a decrease in its expression, perhaps due to increased degradation or slower rate of synthesis; however, the mutant GLUT4 (Asn57right-arrowGln) was not detected at the cell surface after insulin stimulation (22). Likewise, examination of GLUT1 glycosylation with either N-glycosylation site mutants or inhibitors of carbohydrate processing shows that this modification is important for GLUT1 stability, targeting, and glucose transport activity (2, 35, 36). In C2C12 cells, both GLUT4 and GLUT1 appear as multiple bands on SDS-PAGE, yet glucose uptake is occurring, just not in an insulin-sensitive fashion. An interesting point is that IRAP is also a glycoprotein, but its carbohydrate moiety is apparently unaltered in C2C12 cells, and thus aberrant glycosylation is not seen for all such proteins in C2C12 cells and is unlikely to account for the lack of formation of insulin-responsive vesicles.

The genetic program that leads to insulin-sensitive vesicle formation does not appear to be activated in C2C12 myocytes, and it is not induced by dexamethasone or by GLUT4 expression. There are obvious differences in insulin-sensitive glucose transport and dexamethasone-induced GLUT expression if C2C12 and L6 cells are compared. In contrast to C2C12 cells, which exhibit little if any insulin-sensitive glucose uptake, both L6 myoblasts and myotubes appear to have this property to some degree, albeit considerably less than in intact muscle. Moreover, with respect to GLUT4 expression, these cell lines differ in their response to glucocorticoid treatment. The glucocorticoid effect on GLUT4 in C2C12 cells is seen at the mRNA level, and no change in GLUT1 expression was observed under these conditions. In L6 cells, dexamethasone caused an increase in both GLUT1 and GLUT4 protein expression (12). The effect on GLUT1 was attributed to altered protein translation via a rapamycin-sensitive pathway. However, in this study, the levels of GLUT1 and GLUT4 mRNA were not examined (12). In our hands, dexamethasone caused a marked decrease in GLUT4 expression in L6 cells, although the mechanism for this remains unknown. The timing of treatment in L6 myoblasts vs. myocytes could affect the cells' response to glucocorticoid treatment. Thus Ewart et al. (12) examined the effect of dexamethasone after a 24-h exposure to differentiated muscle cells. In the present study, both L6 and C2C12 myoblasts were exposed to dexamethasone at the start of differentiation, and exposure to dexamethasone was maintained until the time of the experiment. The time of dexamethasone exposure (24 h in Ewart et al. vs. >= 4 days in the present study) could therefore result in the different responses observed for GLUT4 expression.

The treatment of C2C12 cells with dexamethasone will be a useful system for identifying those proteins that are involved in GLUT4 vesicle formation and trafficking, for example by transfecting in candidate genes that may rescue the insulin-insensitive phenotype. It is also likely that we can use this system to investigate the mechanism that controls GLUT4 expression. Recently, it was shown that ectopic expression of the peroxisome proliferator-activated receptor-gamma coactivator-1 (PGC-1) in C2C12 cells can induce GLUT4 expression (38). To determine whether dexamethasone was inducing GLUT4 expression via the upregulation of PGC-1, we tested whether or not PGC-1 was expressed in dexamethasone-treated C2C12 myocytes, but we were unable to detect PGC-1 mRNA or protein in these cells (data not shown). Therefore, future experiments will utilize dexamethasone as a tool to determine what transcription factors directly induce GLUT4 expression and to identify molecular components that regulate vesicle formation and trafficking.


    ACKNOWLEDGEMENTS

We thank Drs. Nadia Rosenthal and Antonio Musaro, as well as members of the Pilch laboratory, for helpful discussions, suggestions, and technical assistance. We thank Dr. Bruce Spiegelman (Dana-Farber Cancer Institute and Harvard Medical School) for the anti-PGC-1 antibody.


    FOOTNOTES

This work was supported by research grants from the National Institute of Diabetes and Digestive and Kidney Diseases (DK-30425 and DK-49147 to P. F. Pilch). L. L. Tortorella was supported by National Institutes of Health Training Grant DK-27201 (to N. B. Ruderman).

Address for reprint requests and other correspondence: P. F. Pilch, Dept. of Biochemistry, Boston Univ. School of Medicine, 715 Albany St., Boston, MA 02118 (E-mail: ppilch{at}bu.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

May 28, 2002;10.1152/ajpendo.00092.2002

Received 1 March 2002; accepted in final form 1 April 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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