Troglitazone Induces GLUT4 Translocation in L6 Myotubes

TGZ and BRL49653 were obtained from Sankyo (Tokyo). Darglitazone was obtained from Pfizer (Groton, CT). α-Minimal essential medium (α-MEM) was purchased from Nikken (Kyoto, Japan). Fetal bovine serum (FBS) was obtained from Sanko Junyaku (Tokyo). Radioactive reagents were acquired from NEN Life Science Products (Boston, MA). Protein A/G agarose beads were from Pierce (Rockford, IL). Male Sprague-Dawley rats weighing 120–140 g were purchased from Shimizu Breeding Laboratories (Kyoto, Japan). All other chemicals were purchased from Sigma (St. Louis, MO), unless otherwise noted.

The rat L6 skeletal muscle cell line (a gift from Dr. Amira Klip) was maintained in α-MEM containing 10% FBS and 1% antibiotic solution (penicillin-streptomycin; Gibco, Gaithersburg, MD) in 80-cm² flasks in an atmosphere of 5% CO₂ at 37°C, as reported by Mitsumoto and Klip (16). L6 cells were rendered quiescent in α-MEM containing 2% FBS for 5–6 days to promote fusion into myotubes. The percentage of myotube formation was determined as the percentage of nuclei present in multinucleated myotubes by phase-contrast microscopy. In the present experiment, ∼80–90% of the myoblasts fused into myotubes.

The cells were grown in six-well plates (Corning, Corning, NY). After the indicated period of incubation with or without TGZ in α-MEM, the cells were incubated without FBS for 5 h again in the presence or absence of TGZ. The cells were then rinsed with KRPH (HEPES-buffered Krebs-Ringer phosphate), consisting of 118 mmol/l NaCl, 5 mmol/l KCl, 1.3 mmol/l CaCl₂, 1.2 mmol/l MgSO₄, 1.2 mmol/l KH₂PO₄, and 30 mmol/l HEPES (pH 7.4). The 10 μmol/l 2-deoxy-[³H]d-glucose (2-DG) (1 μCi/ml) uptake was measured over a 10-min period under conditions in which the uptake was linear. The uptake measurement was made in triplicate. Nonspecific uptake was determined in the presence of 10 μmol/l cytochalasin-B and was subtracted from the total uptake. The uptake of 2-DG was terminated after 10 min by rapidly aspirating off the radioactive incubation medium and washing the cells three times in ice-cold phosphate-buffered saline. The radioactivity associated with the cells was determined by cell lysis in 0.5 N NaOH with neutralization by the addition of 0.5 N HCl, followed by liquid scintillation (Scintisol EX-H; Dojin Chemicals, Kumamoto, Japan). Aliquots from each well were used to determine the protein concentration using the BCA Protein assay kit (Pierce). In experiments in which the effect of insulin was examined, 10⁻⁷ mol/l insulin was added to the incubation mixture in KRPH for 20 min before transport studies. Nonspecific uptake and absorption were always <15% of the total uptake.

The subcellular fractionation of myotubes was carried out according to the method of Mitsumoto and Klip (16) with slight modification. The cells from 10-cm dishes were gently scraped, centrifuged (700g for 10 min), and placed on ice. All subsequent steps were carried out at 4°C. The cells was resuspended in buffer I (250 mmol/l sucrose, 5 mmol/l NaN₃, 2 mmol/l EGTA, 200 μmol/l phenylmethylsulfonyl fluoride [PMSF], 1 μmol/l pepstatin A, 1 μmol/l aprotinin, and 20 mmol/l HEPES [pH 7.4]) and then homogenized using 20 strokes of a Dounce homogenizer. This homogenization was sufficient for the almost-complete cell breakage of myotubes, as judged by phase-contrast microscopy. The homogenate was centrifuged at 760g for 5 min to remove nuclei and unbroken cells. The supernatant was centrifuged at 31,000g for 60 min to pellet the crude plasma membrane (CPM). The light microsomes (LMs) were collected from the 31,000g supernatant by centrifugation at 190,000g for 60 min. Both the CPM and LM pellets were suspended in buffer I and frozen at −80°C.

Proteins (20 μg) were separated in 10% SDS-PAGE and transferred to a polyvinylidene fluoride transfer membrane (NEN Life Science Products) in 25 mmol/l Tris, 192 mmol/l glycine, and 20% methanol as described (17). After transfer, the membrane was blocked in 4% nonfat milk. The first antibodies used for the detection of GLUT1, GLUT3, and GLUT4 were generated against the COOH-terminus deduced from each transporter (gifts from Drs. S.W. Cushman and I.A. Simpson). For the detection of protein kinase B (PKB) or phosphorylated PKB, we used an anti-PKB or anti–phosphorylated (Ser473) PKB antibody, respectively (New England BioLabs, Beverly, MA). For the detection of protein kinase C (PKC)-λ, PKC-β2, and PKC-ζ, we used an anti–PKC-λ (Transduction Laboratories, Lexington, KY), anti–PKC-β2 (Gibco BRL), and anti–PKC-ζ (Gibco BRL) antibody, respectively. The results were then visualized with horseradish peroxidase–conjugated secondary antibodies and an enhanced chemiluminescence kit (Amersham, Amersham, U.K.).

To analyze the relative contribution of the three kinds of glucose transporters (GLUT4, GLUT3, and GLUT1) to 2-DG uptake in L6 cells, we performed Western blotting analysis using the plasma membranes from rat adipocytes for GLUT4 and GLUT1 and the rat brain for GLUT3 as standards. The total amount of transporter in the standard membranes was measured in a cytochalasin-B binding assay (18). The relative amounts of GLUT4 and GLUT1 in adipocytes and that of GLUT3 in the brain were estimated using the exofacial label[³H]2-N4-(1-azi-2,2,2-trifluoroethyl) benzoyl-1,3-bis-(d-mannos-4-yloxy)-2-pro-pylamine ([³H]ATB-BMPA) method and immunoprecipitation as described (18,19). The amount of each transporter in the myotubes was then calculated from the results of the Western blotting.

Total RNA was isolated from the cells by extraction with Trizol (Gibco BRL) and prepared according to the manufacturer’s instructions. Typically, 50 μg total RNA was isolated from cells cultured in a 10-cm dish. For Northern blot analysis, 40 μg total RNA was separated by electrophoresis in 1.2% agarose and 2.0 mol/l formaldehyde gels and transferred to Biodyne nylon membranes (Pall BioSupport, East Hills, NY). After ultraviolet crosslinking, the filters were prehybridized, hybridized, and subjected to analysis as described elsewhere. The Northern blot analysis was performed using ³²P-labeled rat GLUT4 and GLUT3 and mouse GLUT1 cDNA fragments as probes (20). The mRNA levels were normalized to the 28S ribosomal RNA levels in the cells to correct for differences in the amount of RNA applied. The mRNA levels (arbitrary units) are expressed in relation to those of the control cells.

L6 myotubes were incubated in the absence or presence of TGZ (10⁻⁵ mol/l) for 24 h. Rat epitrochlearis muscles were dissected and incubated in Krebs-Ringer bicarbonate buffer as described (21). L6 myotubes and muscles were then homogenized in ice-cold lysis buffer (1:100 wt/vol) containing 20 mmol/l Tris-HCl (pH 7.4), 1% Triton X-100, 50 mmol/l NaCl, 250 mmol/l sucrose, 50 mmol/l NaF, 5 mmol/l sodium pyrophosphate, 2 mmol/l dithiothreitol (DTT), 4 mg/l leupeptin, 50 mg/l trypsin inhibitor, 0.1 mmol/l benzamidine, and 0.5 mmol/l PMSF and then centrifuged at 14,000g for 20 min at 4°C. Supernatants (200 μg protein) were immunoprecipitated with an antibody raised using a synthetic peptide of the rat 5′AMP-activated protein kinase (AMPK) α2 subunit (residues 490–514) (22) and protein A/G beads. Immunoprecipitates were washed two times in lysis buffer and two times in buffer containing 240 mmol/l HEPES (pH 7.4) and 480 mmol/l NaCl. The kinase reaction was carried out in 40 mmol/l HEPES (pH 7.0), 0.2 mmol/l AMP, 80 mmol/l NaCl, 0.8 mmol/l DTT, 5 mmol/l MgCl₂, 0.2 mmol/l ATP (2 mCi [γ-³²P]ATP), and 0.1 mmol/l SAMS peptide (23) for 20 min at 30°C. Reaction products were spotted on Whatman P81 filter paper, and then the papers were extensively washed in 1% phosphoric acid. The radioactivity in the papers was measured with a scintillation counter. Kinase activity was expressed as incorporated ATP (picomoles) per immunoprecipitated protein (milligrams) per minute.

Reverse transcription (RT) was performed on 2 μg RNA using the Superscript Preamplication System (Gibco BRL) according to the manufacturer’s instructions. The cDNA was amplified by polymerase chain reaction (PCR) using a pair of primers to amplify the PPAR-γ fragment as follows (nucleotide positions according to GenBank): 5′-GGATTCATGACCAGGGAGTTCCTC-3′ (residues 1120–1143, sense strand) and 5′-GCGGTCTCCACTGAGAATAATGAC-3′ (residues 1252–1275, antisense strand).

Results are expressed as means ± SE. Statistical significance was tested with a one-way analysis of variance followed by Duncan’s multiple range test and Wilcoxon’s signed-rank test, as appropriate, and differences were accepted as significant at the P < 0.05 level.

We examined the effect of TGZ (10⁻⁵ mol/l) on the 2-DG uptake in L6 myotubes over a 30-h time course. As shown in Fig. 1A, TGZ increased the 2-DG uptake in a time-dependent fashion, and the uptake was maximal after 15 h (0 h, 0.25 ± 0.13 vs. 30 h, 2.95 ± 0.26 nmol · min⁻¹ · well⁻¹, P < 0.02). The dose-response curve (Fig. 1B) showed a maximum response at concentrations >7.5 × 10⁻⁶ mol/l. This TGZ-stimulated glucose uptake was completely blocked by adding cytochalasin-B (10 μmol/l) to the incubation buffer just before the 2-DG assay (Fig. 1B). TGZ treatment had no significant effect on cell number or protein content in each well (data not shown). There were no obvious differences in cell morphology after the treatment.

We next investigated whether the TGZ-induced 2-DG uptake was accompanied by an increase in the total amount of glucose transporters in the myotubes. It has been reported that L6 myotubes contain three isoforms of glucose transporters: GLUT4, GLUT1, and GLUT3 (24). After incubating the cells with TGZ (10⁻⁵ mol/l) for 24 h, we assessed the total amount of GLUT4, GLUT1, and GLUT3 protein by Western blotting using the specific antibody against each transporter (Figs. 2A and B). TGZ did not affect the amount of each transporter in the myotubes. Comparable results were obtained by Northern blotting (Figs. 2C and D).

To analyze the relative contribution of the three isoforms of glucose transporters to 2-DG uptake in L6 cells, we performed Western blot analysis using the plasma membranes from rat adipocytes for GLUT4 and GLUT1 and the rat brain for GLUT3 as standards (the left lanes in Fig. 2A). The amount of each glucose transporter in the standard membrane was estimated by a cytochalasin-B binding assay, photo-affinity labeling of the exofacial label [³H]ATB-BMPA, and immunoprecipitation as described (18,19). In the L6 myotubes, the amounts of GLUT4, GLUT1, and GLUT3 were estimated to be ∼8, 1, and 1 pmol · mg⁻¹ · protein⁻¹, respectively (Fig. 2B). Thus, TGZ did not change the total amount of glucose transporters, and GLUT4 appeared to be the most abundant transporter in the L6 myotubes.

The above-mentioned results suggested that TGZ exposure causes the translocation or an increase in the intrinsic activity of glucose transporters, especially GLUT4. To further test this concept, we performed a GLUT4 translocation assay on membrane fractions from L6 myotubes (Fig. 3). The cells were exposed or not exposed to TGZ (10⁻⁵ mol/l) for 24 h and then fractionated into membrane compartments, i.e., CPMs and LMs. In the TGZ-treated myotubes, there was an increase in GLUT4 in the CPMs and a corresponding decrease of GLUT4 in the LMs, a result indicative of transporter translocation to the CPMs. The level of GLUT4 protein present in the CPMs of the TGZ-treated cells was much higher than that of the insulin-treated cells (10⁻⁷ mol/l for 20 min) (P < 0.05). In addition, significant changes in GLUT1 or GLUT3 translocation were not produced by TGZ treatment (data not shown). GLUT4 translocation to the plasma membrane thus appears to be a primary cause of the enhancement of glucose uptake by TGZ in L6 myotubes.

To examine whether the change in the TGZ-treated cells was caused by a protein synthesis–dependent process, we studied the effect of cycloheximide (3.5 × 10⁻⁶ mol/l) on TGZ-induced 2-DG uptake. In the presence of cycloheximide, the effect of TGZ was completely abolished (Fig. 4A: control, 0.9 ± 0.2; TGZ, 2.7 ± 0.2; TGZ and cycloheximide, 1.0 ± 0.2 nmol · min⁻¹ · well⁻). The 2-DG uptake in the cells treated with cycloheximide alone (0.5 ± 0.1 nmol · min⁻¹ · well⁻) was significantly lower than that in the control cells (P < 0.05).

The removal of TGZ by the washing procedure completely reversed the 2-DG uptake to the basal level after further incubation without TGZ for 24 h (Fig. 4A). Moreover, the reversed 2-DG uptake of the cells was restimulated by insulin (Fig. 4A). Thus, the TGZ-stimulated GLUT4 translocation was reversible, suggesting that GLUT4, which was translocated to the plasma membranes by TGZ, could enter the recycling compartment again.

To clarify the mechanism of the enhancement of GLUT4 translocation to the plasma membrane by TGZ, we first studied whether insulin enhances the TGZ-stimulated 2-DG uptake additively. As shown in Fig. 4B, the 2-DG uptake in TGZ-treated cells (10⁻⁵ mol/l, 24 h) was not further enhanced by insulin (10⁻⁷ mol/l, 20 min), thus indicating the likelihood that a complete insulin-dependent transporter translocation has already occurred.

We next examined whether the TGZ-induced 2-DG uptake was reversed by wortmannin, which is a specific inhibitor for phosphatidylinositol (PI) 3-kinase that blocks the insulin-signaling pathway. After treatment of the cells with TGZ (10⁻⁵ mol/l) for 24 h, wortmannin (5 × 10⁻⁸ mol/l) was added to the culture medium for 1 h, and the 2-DG uptake was then measured. In the TGZ-treated cells, 2-DG uptake was only partially reversed by wortmannin to ∼80% (Fig. 4C). In a parallel experiment, the 2-DG uptake in insulin-treated cells (10⁻⁷ mol/l, 20 min) was completely reversed to the basal level by wortmanin within 1 h.

To examine whether TGZ (10⁻⁵ mol/l) affects the post–PI 3-kinase step in the insulin-signaling system for glucose transport, we examined the levels of PKB, phosphorylated (Ser473) PKB, and PKCs (PKC-λ, PKC-β2, and PKC-ζ) in the total cell lysate using Western blots. As shown in Fig. 5A, the protein level of PKB was not changed by the TGZ treatment. The levels of phosphorylated PKB in the basal and insulin-stimulated (10⁻⁷ mol/l) states were not changed by TGZ, although insulin alone significantly increased the phosphorylation of PKB. As shown in Fig. 5A (upper panel), the mobility of PKB was shifted in parallel with its phosphorylation. Figure 5B shows that the protein levels of PKC-λ, PKC-β2, and PKC-ζ were not changed by the TGZ treatment (10⁻⁵ mol/l, 24 h).

Recent studies (21,25) have provided evidence that AMPK is involved in enhancing glucose transport by an insulin-independent signaling mechanism in skeletal muscle. AMPK is a heterotrimeric protein consisting of one catalytic subunit (α) and two noncatalytic subunits (β and γ) (26). Two isoforms of the α subunit have been identified (α1 and α2) in mammalian skeletal muscle. The α2 isoform is highly expressed in skeletal muscle, heart, and liver, whereas the α1 isoform is widely distributed in many tissues (27). A recent report has shown that the activity of AMPK α2 is closely correlated with glucose transport activity in rat skeletal muscle (28). To examine whether AMPK α2 is involved in the TGZ-induced increase in glucose transport, we determined AMPK α2 activity in L6 myotubes. Figure 6 shows that TGZ (10⁻⁵ mol/l, 24 h) did not change AMPK α2 activity in L6 myotubes. Furthermore, AMPK activity in L6 myotubes was 1/70 of that in nonstimulated rat skeletal muscle.

We investigated the effects of two more thiazolidinediones, darglitazone and BRL49653, on glucose uptake in L6 myotubes. Figure 7 shows that TGZ, darglitazone, and BRL49653 increased the 2-DG uptake in L6 myotubes in a dose-dependent manner. The rank order of potency for the glucose uptake in L6 myotubes was as follows: TGZ > darglitazone > BRL49653.

Figure 8 shows the result of the RT-PCR using a primer to amplify the sequences of PPAR-γ cDNA. PPAR-γ mRNA was detected in L6 myotubes only when PCR was performed for 35 cycles, whereas in adipocytes (fat), it was detected after both 25 and 35 cycles. The mRNA level of PPAR-γ was not enhanced by the TGZ treatment (10⁻⁵ mol/l, 24 h). PPAR-γ protein was not detected by Western blot analysis in contrast with the adipocytes (data not shown).

The action of TGZ may be attributable to the similarity of its molecular structure to that of α-tocopherol (29,30). Figure 9 shows that α-tocopherol (10⁻⁵ mol/l, 24 h) did not increase glucose uptake in L6 myotubes.

A number of studies have demonstrated that insulin resistance in skeletal muscles plays a role in the insulin resistance of obesity and type 2 diabetes. A decrease in GLUT4 translocation from LMs to the plasma membranes has been implicated as a possible cause of insulin resistance, as has the reduced kinase function of the insulin receptor. Investigations of the mechanisms underlying insulin resistance in type 2 diabetes have revealed tissue-specific regulation of GLUT4 with decreased gene expression in adipose cells but not in skeletal muscle (31). This has led to the hypothesis that alterations in the trafficking of the GLUT4 vesicle or in the exposure or activation of the GLUT4 transporter may cause insulin resistance in skeletal muscle in obesity and diabetes. In a human study, TGZ has been reported to increase peripheral glucose disposal, mainly in skeletal muscles, but to have no significant effect on hepatic glucose production (10). Moreover, Burant et al. (11) reported the antidiabetic action of TGZ in aP2/DTA mice, in which white fat and brown fat are virtually eliminated by the fat-specific expression of diphtheria toxin A chain. These results suggest that TGZ is one of the keys to improving insulin resistance in skeletal muscle. To investigate this hypothesis, we tested the effect of TGZ on glucose uptake and changes in glucose transporters in L6 myotubes.

In the present study, the TGZ-stimulated 2-DG uptake was completely blocked by cytochalasin-B, suggesting that this effect was caused by changes in glucose transporters in the plasma membrane and not by an increase in simple diffusion across the membrane. The total amount of glucose transporters, including GLUT1, GLUT3, and GLUT4, was not changed by the TGZ treatment. We have also shown that GLUT4 was expressed much more intensely than GLUT1 and GLUT3 in L6 myotubes. The ability of TGZ to translocate GLUT4 from LM to the CPM was more augmented than that of insulin. In another study (12), the short-term (2-h) exposure of fully differentiated L6 myotubes to TGZ had no effect on glucose transport activity, but the long-term (72-h) treatment of myotubes with TGZ resulted in a doubling of glucose transport in the absence of insulin. This is compatible with the present results. However, the authors of that study did not refer to the changes of glucose transporters in their conditions. Recently, Cooksey et al. (32) reported that transgenic mice overexpressing the rate-limiting enzyme for hexosamine synthesis, glutamine:fructose-6-phosphate amidotransferase, had hexosamine-induced insulin resistance because of a decrease in the GLUT4 translocation in the skeletal muscle and that TGZ improved the glucose disposal in those transgenic mice. These reports further support the hypothesis that TGZ directly acts on skeletal muscles and affects the translocation of GLUT4.

As shown in Fig. 4B, exposure of the cells to TGZ resulted in an increase in basal glucose uptake. More important, there was no further increase in insulin-stimulated glucose transport, thus indicating the likelihood that complete insulin-dependent transporter translocation in the newly steady state was already achieved and that the signaling pathway activated by TGZ converges with that activated by insulin. In the presence of insulin, GLUT4 is recycled and a new steady state is achieved in rat adipose cells (33), increasing exocytosis, although it is still unclear whether insulin also affects the endocytosis of GLUT4. Acting as an inhibitor of the lipid kinase PI 3-kinase (34), wortmannin blocks the insulin-stimulated translocation of GLUT4 from its basal compartment to the plasma membrane (35). In the present study, wortmannin reversed the insulin-stimulated 2-DG uptake to the basal level when added after full stimulation by insulin (Fig. 4C). The addition of wortmannin to the TGZ-stimulated cells, however, did not reverse the uptake to the basal level. In addition, the amount of PKB and its phosphorylated state (Ser473) were not changed by the TGZ treatment (Fig. 5A). TGZ did not affect the protein levels of each isoform of PKC (Fig. 5B). Although the site of action on TGZ is still not clear, one possibility is that TGZ acts at a step beyond PI 3-kinase, PKB, and PKC in which GLUT4 exocytosis is enhanced according to the present results. Another possibility is that TGZ reduces the endocytosis of GLUT4 and thus allows the accumulation of GLUT4 in the plasma membrane. The latter mechanisms have been described in several recent articles. The disassembly of clathrin lattices by potassium depletion results in the accumulation of GLUT4 at the cell surface (36), and the expression of a dominant interfering dynamin mutant in rat (37) and 3T3-L1 (38) adipocytes inhibits GLUT4 endocytosis. Further study will be necessary to elucidate the precise site of action of TGZ with regard to the exocytosis and endocytosis steps.

The antioxidant action of TGZ, which is attributable to the similarity of its molecular structure to that of α-tocopherol, is considered to be of benefit in preventing diabetic vascular complications, in addition to having hypoglycemic and hypolipidemic effects (29,30). We studied 2-DG uptake in the presence of α-tocopherol. Figure 9 shows that α-tocopherol (10⁻⁵ mol/l, 24 h) did not increase glucose uptake in L6 myotubes. Various thiazolidinediones increased glucose uptake in L6 myotubes as shown in Fig. 7. We believe that the structure of the thiazolidinedione is needed for enhancing glucose uptake in L6 myotubes, but that of α-tocopherol is not.

The onset of the augmentation of the basal glucose transport by TGZ was slow; 6–12 h passed before significant effects were observed, suggesting that the effect requires the synthesis of new proteins. To examine this possibility, we incubated L6 cells together with cycloheximide (3.5 × 10⁻⁶ mol/l) and TGZ for 24 h. Figure 4A illustrates that the addition of the protein synthesis inhibitor cycloheximide prevented the TGZ-induced 2-DG uptake. Furthermore, the washing procedure to remove TGZ from the cells completely reversed the 2-DG uptake to the level of the control cells, and the uptake could be restimulated by insulin. Thus, the increase of glucose uptake with time apparently reflects the ongoing synthesis of a new protein that affects GLUT4 translocation. Further study will be necessary to find the protein(s).

The recent discovery that thiazolidinediones are synthetic ligands for PPAR-γ reveals a potential mechanism by which TGZ could regulate gene expression. PPAR-γ mRNA levels are lower in skeletal muscle biopsies and cultured muscle than in adipose tissue (39,40), although PPAR-γ is expressed in skeletal muscles (39,40,41,42) and provides a potential target for TGZ action. TGZ treatment has also been reported to increase mRNA levels for PPAR-γ in skeletal muscles (8). In fact, PPAR-γ mRNA was detected in L6 myotubes only when RT-PCR was performed in our experiments (Fig. 8), and the mRNA level of PPAR-γ was not enhanced by the TGZ treatment (10⁻⁵ mol/l, 24 h).

The rank order for clinical efficacy has been reported to be BRL49653 > darglitazone > TGZ (43,44,45). It has also been reported that BRL49653 has high affinity for PPAR-γ among thiazolidinediones (6), and the potency of thiazolidinediones as PPAR-γ agonists correlates with their antidiabetic efficacy in vivo (46,47). In our experiments, the rank order of potency for the glucose uptake in L6 myotubes was as follows: TGZ > darglitazone > BRL49653 (Fig. 7), which is inconsistent with the clinical efficacies and affinities for PPAR-γ reported previously. Recently, one of the thiazolidinediones has been reported to possess strong action of lowering blood glucose in spite of its low binding affinity for PPAR-γ (48). This effect is partly explained by unique partial agonism of coactivator recruitment to PPAR-γ. Thiazolidinedione action in skeletal muscle (L6 myotubes) may include activation of PPAR-γ coactivators. Another possibility is that the bioavailability is different among such thiazolidinediones in skeletal muscle or in L6 myotubes. For example, the intracellular concentration of each drug could be different from the extracellular concentration. It is also possible that the thiazolidinediones activate an alternative signal transduction pathway and increase glucose uptake through PPAR-γ–independent mechanisms in skeletal muscle.

In our study, thiazolidinediones (TGZ, BRL49653, and darglitazon) increased glucose transport in L6 myotubes in the absence of insulin. This observation is consistent with previous in vitro results obtained using the L6 cell (12,49), 3T3-L1 adipocyte (50), and cardiomyocyte (51). However, many studies have shown that insulin is necessary for thiazolidinediones to be effective in vivo. Insulin lowers plasma glucose levels both by stimulating glucose uptake into muscle and adipose tissue and by inhibiting hepatic glycogen breakdown and gluconeogenesis. Insulin is also necessary for lipid metabolism. Therefore, it may be reasonable that increased glucose transport by thiazolidinediones is not sufficient to ameliorate the metabolic pertubation under conditions of insulin deficiency such as type 1 diabetes or in animal models of streptozocin-induced diabetes.

AMPK is involved in enhancing glucose transport by an insulin-independent signaling mechanism in skeletal muscle (21). A recent report showed that the activity of AMPKα2 is closely correlated with glucose transport activity in rat skeletal muscle (28). Figure 6 shows that TGZ did not change AMPKα2 activity in L6 myotubes and that the magnitude of the activities was 1/70 that of skeletal muscle of rat per same protein content. The result suggests that AMPK has no major role in TGZ-induced glucose uptake in L6 cells.

According to one report (52), in L6 myotubes, cytokines and lipopolysaccharide (LPS) significantly stimulated nitric oxide (NO) production and induced inducible NO synthase (iNOS) protein and mRNA expression. Cytokines and LPS markedly increased basal glucose transport, but inhibited insulin-stimulated glucose transport. It has been suggested that cytokines/LPS exposure significantly increased GLUT1 transporter protein levels but decreased GLUT4 transporter protein levels by inducing iNOS expression and NO production in L6 cells. Another study (53) found that TGZ upregulated cytokine-stimulated NO synthase in vascular smooth muscle cells, but TGZ alone did not stimulate it. In our study, L6 myotubes were not exposed to cytokines. TGZ did not change the protein levels of GLUT4 and GLUT1. TGZ increased basal glucose transport and did not inhibit insulin-stimulated glucose transport. Thus, we consider the NO pathway not to be involved in the TGZ-induced glucose uptake in L6 myotubes.

In summary, prolonged TGZ treatment stimulates glucose transport in L6 myotubes by activating GLUT4 translocation from the LM to plasma membrane without changing the total amount of GLUT4. Synthesis of protein(s) in the GLUT4 translocation machinery may play an important role in activating transport, although further investigation is necessary to identify the protein(s). The effects of TGZ on GLUT4 translocation may include a new mechanism for improving glucose transport in skeletal muscle.

This work was supported in part by research grants from the Japanese Ministry of Education, Science, and Culture; the Japanese Ministry of Health and Welfare; the Smoking Research Foundation; and the Insulin Research Foundation.

We would like to thank Drs. S.W. Cushman and I.A. Simpson for their advice and for providing collagenase and albumin, Drs. Amira Klip and Y. Mitsumoto for donating L6 cells, and Dr. G.I. Bell for donating rat GLUT4 and GLUT3 and mouse GLUT1 cDNA fragments as probes.

A part of this work was presented at the 58th Annual Meeting of the American Diabetes Association, San Diego, CA, June 1999.