Department of Biochemistry, Boston University School of Medicine, Boston, Massachusetts 02118
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ABSTRACT |
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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
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INTRODUCTION |
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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--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.
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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%
-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
-antibodies from Dr. Hidde Ploegh, Harvard Medical School; and
anti-
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 -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 [32
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.
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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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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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DISCUSSION |
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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 (Asn57Gln) resulted in a decrease in its
expression, perhaps due to increased degradation or slower rate of
synthesis; however, the mutant GLUT4 (Asn57
Gln) 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- 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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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