The cellular mechanism by which high-fat feeding induces skeletal muscle insulin resistance was investigated in the present study. Insulin-stimulated glucose transport was impaired (∼40–60%) in muscles of high fat–fed rats. Muscle GLUT4 expression was significantly lower in these animals (∼40%, P < 0.05) but only in type IIa–enriched muscle. Insulin stimulated the translocation of GLUT4 to both the plasma membrane and the transverse (T)-tubules in chow-fed rats. In marked contrast, GLUT4 translocation was completely abrogated in the muscle of insulin-stimulated high fat–fed rats. High-fat feeding markedly decreased insulin receptor substrate (IRS)-1–associated phosphatidylinositol (PI) 3-kinase activity but not insulin-induced tyrosine phosphorylation of the insulin receptor and IRS proteins in muscle. Impairment of PI 3-kinase function was associated with defective Akt/protein kinase B kinase activity (−40%, P < 0.01) in insulin-stimulated muscle of high fat–fed rats, despite unaltered phosphorylation (Ser473/Thr308) of the enzyme. Interestingly, basal activity of atypical protein kinase C (aPKC) was elevated in muscle of high fat–fed rats compared with chow-fed controls. Whereas insulin induced a twofold increase in aPKC kinase activity in the muscle of chow-fed rats, the hormone failed to further increase the kinase activity in high fat–fed rat muscle. In conclusion, it was found that GLUT4 translocation to both the plasma membrane and the T-tubules is impaired in the muscle of high fat–fed rats. We identified PI 3-kinase as the first step of the insulin signaling pathway to be impaired by high-fat feeding, and this was associated with alterations in both Akt and aPKC kinase activities.

Insulin resistance represents a major pathogenic impairment in the development of type 2 diabetes (1,2). In humans and rodents, skeletal muscle is the primary site of insulin-mediated glucose disposal (2). Insulin increases glucose uptake in muscle by eliciting GLUT4 translocation from an intracellular storage site to both the plasma membrane and the transverse (T)-tubules through a complex signaling cascade (3,4,5). Impaired GLUT4 translocation has been shown to be linked to reduced glucose utilization in muscle of insulin-resistant and type 2 diabetic subjects (6,7,8). However, the precise mechanism underlying the reduced stimulatory effect of insulin on glucose transport is still unclear. Both receptor and postreceptor defects have been observed in various models of insulin resistance (9).

Insulin stimulates GLUT4 translocation by binding its receptor α-subunits, leading to autophosphorylation of the transmembrane β-subunits and intrinsic activation of receptor tyrosine kinase activity. In skeletal muscle, the activated insulin receptor (IR) increases the tyrosine phosphorylation of IR substrate (IRS)-1 and IRS-2, leading to activation of phosphatidylinositol (PI) 3-kinase (4). It is believed that PI 3-kinase activation by insulin is essential for the stimulation of GLUT4 translocation. Indeed, a large number of studies have shown that inhibitors of PI 3-kinase (wortmannin and LY294002) or overexpression of a mutated p85 adapter subunit lacking the ability to bind the p110 catalytic subunit fully inhibit insulin-mediated GLUT4 translocation and glucose transport in adipocytes and skeletal muscle cells (rev. in [10]).

Downstream effector(s) of PI 3-kinase involved in the regulation of glucose transport have yet to be clearly identified. Candidate molecules of interest include the serine/threonine kinase Akt (also termed protein kinase B [PKB] or related to A and C [RAC] protein kinase) and atypical protein kinase C (aPKC) (ζ/λ). Both aPKC and Akt lie in the PI 3-kinase/3-phosphoinositide-dependent kinase (PDK)-1 signaling pathway, giving rise to phosphorylation on Thr410 and Thr308, respectively (11,12). Full activation of Akt further requires phosphorylation on Ser473 by the putative PDK-2 (13), whereas aPKC activity appeared to be dependent on autophosphorylation by an as yet unknown mechanism (14). Evidence for the implication of Akt and aPKC in the insulin-dependent regulation of glucose transport in muscle cells arises from transfection studies using either kinase-inactive or overexpression/constitutively active forms of the kinases (15,16,17,18,19). Only a few studies have examined insulin-dependent activation of Akt in muscle from animal models of insulin resistance and diabetic subjects, and these yielded contradictory results regarding its possible involvement in impaired glucose homeostasis (20,21,22,23). However, it is still unknown whether the stimulatory effect of insulin on aPKC is altered in muscles from insulin-resistant animals or humans.

The high fat–feeding model of insulin resistance displays common features of the abdominal obesity syndrome encountered in insulin-resistant subjects (24,25). Indeed, rats fed a high-fat diet develop skeletal muscle insulin resistance, increased adiposity, hyperinsulinemia, and mild hyperglycemia (24,25,26,27). It is generally believed that GLUT4 protein expression is normal in skeletal muscle of high fat–fed rats (24,27,28), but this has not been a consistent finding (29,30,31). Recruitment of GLUT4 assessedby exofacial photolabelling (2-N-4-(1-azi2,2,2-trifluoro-ethyl)-benzoyl-1,3-bis-(d-mannose-4-yloxy)-2-propylamine) led to the view that a high-fat diet reduced insulin-stimulated GLUT4 translocation to the cell surface (27,32). However, Rosholt et al. (28) did not observe such impairment in isolated plasma membrane vesicles, suggesting that the main site of defective GLUT4 translocation may be localized to the T-tubules, the principal component of the muscle cell surface, although this remains to be studied. Early impairments in the insulin signaling cascade may lead to a decrease in GLUT4 translocation in muscle of high fat–fed rats. Insulin-induced tyrosine phosphorylation of both the IR and IRS-1 was reported to be normal in rats fed a high-fat diet for 8 weeks (32). However, IRS-1 associated PI 3-kinase activity has been found to be impaired in mice after 4 weeks of high-fat feeding (27). Whether insulin-dependent activation of either Akt or aPKC pathways is altered in skeletal muscle of high fat–fed rats is still unknown.

The aim of the present study was to clarify the cellular mechanisms leading to defective insulin-stimulated glucose transport in skeletal muscle of the high fat–fed rat. More specifically, we tested the hypothesis that potential alterations of signaling elements downstream of PI 3-kinase activation may be linked with impaired GLUT4 translocation in the muscle of high fat–fed obese rats.

Materials.

Reagents for SDS-PAGE and immunoblotting were from Bio-Rad (Mississauga, ON, Canada). Enhanced chemiluminescence reagent, 2-deoxy-d-[3H]glucose (2-[3H]DG), and D-14C-sucrose were from NEN Life Science Products (Boston, MA). [γ-32P]ATP, protein A- and G-Sepharose, and anti-mouse or anti-rabbit IgG conjugated to horseradish-peroxidase were purchased from Amersham Pharmacia Biotech (Baie d’Urfé, QC, Canada). Anti-goat IgG conjugated to horseradish-peroxidase, polyclonal antibodies against GLUT1 (raised against 20 COOH-terminal amino acids [C-20]), GLUT4 (C-20), IRS-1 (C-20), aPKCs (C-20), and Akt 1/2 (which recognizes both Akt 1 and 2 [H-136]), were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Anti–phospho-specific (Ser473 and Thr308) antibodies against Akt were from New England Biolabs (Beverly, MA). Akt/PKB substrate (Crosstide) and antibodies against phosphotyrosine (4G10 clone) and p85 were obtained from Upstate Biotechnology (Lake Placid, NY). Myelin basic protein was from Sigma (St. Louis, MO). Okadaic acid was purchased from Calbiochem (La Jolla, CA). l-α–PI was from Avanti Polar Lipids (Alabaster, AL). Oxalate-treated thin-layer chromatography silica gel H plates were obtained from Analtech (Newark, DE). All other chemicals were of the highest analytical grade.

Treatment of animals.

All experiments reported here were approved by the Laval University Animal Care and Handling Committee and comply with Canadian Council on Animal Care guidelines for the care and use of animals for research purposes. Male Wistar rats (Charles River, Montréal, QC, Canada) weighing 200–250 g at the beginning of the study were housed individually in plastic cages in animal quarters maintained at 22°C with a 12:12-h dark-light schedule. Animals were fed either low-fat Rodent Chow (Charles River Rodent Chow 5075, Purina Mills, St. Louis, MO) or a purified high-fat diet for 4 weeks. As percent of total energy, the high-fat diet consisted of 32.5% lard, 32.5% corn oil, 20% sucrose, and 15% protein, whereas the Rodent Chow diet contained 57.3% carbohydrate, 18.1% protein, and 4.5% fat. The energy contents of the diets were 14.3 kJ/g for the Rodent Chow diet and 25.5 kJ/g for the high-fat diet.

Hyperinsulinemic-euglycemic clamp and tracer injection.

The clamp procedure was essentially performed as previously described (33). Briefly, unrestrained conscious animals were allowed to rest for 40 min before the initial blood sample (300 μl) was obtained. For hyperinsulinemic-euglycemic clamp, a continuous intravenous infusion of insulin was then started at the rate of 4 mU · kg−1 · min−1 and continued for 2 h. The arterial blood glucose concentration was clamped using a variable-rate glucose infusion. Control rats were infused with saline for the same period of time, and no exogenous glucose was necessary to maintain euglycemia. Tracer injection (2-[3H]DG and 14C-sucrose) was administered 20 min before the end of the clamp to determine individual tissue glucose uptake, as previously reported (33). Immediately after the clamp, the rats were killed, and their hindlimb muscles (soleus, tibialis, gastrocnemius, and quadriceps) were rapidly excised, cleaned of extraneous tissues, and frozen in liquid nitrogen. The muscle were kept at −80°C until further processing.

Acute insulin stimulation.

Overnight-fasted rats were injected either with saline or insulin (8 units/kg) for 4 min, as previously described (3). Muscles were quickly excised and immediately frozen in liquid nitrogen. Muscles were homogenized in six volumes of lysis buffer containing 20 mmol/l Tris, pH 7.5, 140 mmol/l NaCl, 1 mmol/l CaCl2, 1 mmol/l MgCl2, 10% glycerol, 10 mmol/l sodium pyrophosphate, 10 mmol/l NaF, 2 mmol/l Na3VO4, 2 mg/ml benzamidine, 1 mmol/l PMSF, and protease inhibitor cocktail (Sigma). Okadaic acid (100 nmol/l) was added in lysis buffer for Akt and aPKC kinase activities. Muscle homogenates were solubilized in 1% Nonidet P-40 for 1 h at 4°C and centrifuged at 14,000g for 10 min. Supernatant was used for insulin signaling studies as described below.

Subcellular fractionation.

Plasma membranes, T-tubules, and GLUT4-enriched intracellular membranes were isolated from muscles (8–10 g, mixed gactrocnemius and quadriceps) using a procedure developed in our laboratory (3,34). This subcellular fractionation protocol has been extensively characterized with immunologic and enzymatic markers (3,34). In brief, this technique allows the simultaneous and separated isolation of plasma membrane, T-tubules, and intracellular membrane vesicles from the same muscle homogenate. GLUT4 content was determined in fractions obtained from saline- or insulin-infused rats by Western blotting, as described below.

Western blotting.

Membranes (10 μg) or muscle homogenates (50 μg) were subjected to SDS-PAGE (7.5% gel) and electrophoretically transferred to polyvinylidene difluoride (PVDF) filter membranes for 2 h. PVDF membranes were then blocked for 1 h at room temperature with buffer I (50 mmol/l Tris-HCl, pH 7.4, and 150 mmol/l NaCl) containing 0.04% NP-40, 0.02% Tween-20, and 5% nonfat milk. This step was followed by overnight incubation at 4°C with primary antibodies, as described in the figure legends. The PVDF membranes were then washed for 30 min, followed by a 1-h incubation with either anti-mouse or anti-rabbit IgG conjugated to horseradish-peroxidase in buffer I containing 1% bovine serum albumin. The PVDF membranes were washed for 30 min in buffer I, and the immunoreactive bands were detected by the enhanced chemiluminescence method. A muscle standard (an unrelated crude membrane fraction) was run on every gel for comparison of samples from different immunoblots.

Tyrosine phosphorylation of the IR and IRS.

Muscle lysates (1 mg of protein) were immunoprecipitated with 2 μg of anti-phosphotyrosine (4G10) coupled to protein A-Sepharose overnight at 4°C. The immune complex was washed three times in phosphate-buffered saline (PBS) (pH 7.4) containing 1% NP-40 and 2 mmol/l Na3VO4, resuspended in Laemmli buffer, and boiled for 5 min. Proteins were resolved on SDS-PAGE (6% gel) and processed for Western blot analysis as described above.

PI 3-kinase activity.

Muscle lysates (1 mg of protein) were immunoprecipitated with 2 μg of anti–IRS-1 coupled to protein A-Sepharose overnight at 4°C. PI 3-kinase activity was determined as described by Kristiansen et al. (35), with minor modifications. Immune complexes were washed twice with wash buffer I (PBS, pH 7.4, 1% NP-40, and 2 mmol/l Na3VO4), twice with wash buffer II (100 mmol/l Tris, pH 7.5, 500 mmol/l LiCl, and 2 mmol/l Na3VO4), and twice with wash buffer III (10 mmol/l Tris, pH 7.5, 100 mmol/l NaCl, 1 mmol/l EDTA, and 2 mmol/l Na3VO4). Beads were resuspended in 70 μl of kinase buffer (8 mmol/l Tris, pH 7.5, 80 mmol/l NaCl, 0.8 mmol/l EDTA, 15 mmol/l MgCl2, 180 μmol/l ATP, and 5 μCi [γ-32P]ATP) and 10 μl of sonicated PI mixture (20 μg l-α-PI, 10 mmol/l Tris, pH 7.5, and 1 mmol/l EGTA) for 15 min at 30°C. Reaction was stopped by the addition of 20 μl 8 mol/l HCl, mixed with 160 μl CHCl3:CH3OH (1:1), and centrifuged. Lower organic phase was spotted on oxalate-treated silica gel TLC plates and developed in CHCl3:CH3OH:H2O:NH4OH (60:47:11.6:2). The plate was dried and visualized by autoradiography with intensifying screen at −80°C.

Akt/PKB activity.

Muscle lysates (1 mg of protein) were immunoprecipitated with 4 μg of anti–Akt 1/2 coupled to protein G-Sepharose for 4 h at 4°C. Akt activity was measured essentially as described previously (20,21). Immune complexes were washed twice with wash buffer I (PBS, pH 7.4, 1% NP-40, and 100 μmol/l Na3VO4) and twice with wash buffer II (50 mmol/l Tris, pH 7.5, 10 mmol/l MgCl2, and 1 mmol/l dithiothreitol [DTT]). Beads were resuspended in 30 μl of kinase buffer (50 mmol/l Tris, pH 7.5, 10 mmol/l MgCl2, 1 mmol/l DTT, 8 μmol/l ATP, 2 μCi [γ-32P]ATP, and 50 μmol/l Crosstide) for 30 min at 30°C. Reaction product was resolved on a 40% acrylamide-urea gel and visualized by autoradiography with intensifying screen at −80°C. In preliminary experiments in which insulin was injected in intact rats for 4, 10, 20, or 30 min, it was determined that Akt/PKB kinase activation by insulin was maximal after 4 min of insulin injection (data not shown).

aPKC (ζ/λ) activity.

Muscle lysates (1 mg of protein) were immunoprecipitated with 2 μg of anti-PKC (ζ/λ) overnight at 4°C, then immune complexes were collected on protein A/G-Sepharose for 2 h. aPKC activity was determined according to the method of Chou et al. (12). Beads were washed twice with wash buffer I (PBS, pH 7.4, 1% NP-40, and 2 mmol/l Na3VO4), twice with wash buffer II (100 mmol/l Tris, pH 7.5, 500 mmol/l LiCl, and 100 μmol/l Na3VO4), and twice with wash buffer III (50 mmol/l Tris, pH 7.5, 10 mmol/l MgCl2, and 100 μmol/l Na3VO4). Beads were resuspended in 30 μl of kinase buffer (50 mmol/l Tris, pH 7.5, 10 mmol/l MgCl2, 40 μmol/l ATP, 5 μCi [γ-32P]ATP, and 5 μg myelin basic protein) for 12 min at 30°C. Reaction was stopped by the addition of Laemmli buffer and heated for 30 min at 37°C. Reaction product was resolved on 13% SDS-PAGE. Gel was dried and visualized by autoradiography with intensifying screen at −80°C. In preliminary experiments, it was determined that aPKC activation by insulin was maximal after 4 min of insulin injection (data not shown). Furthermore, we have verified that neither the M-kinase form of aPKC (36) nor Akt 1/2 (37) or PI 3-kinase (38) were recovered in the aPKC immunoprecipitates of both Rodent Chow– and high fat–fed rats (data not shown).

Data analysis.

Autoradiographs were analyzed by laser scanning densitometry using a tabletop Agfa scanner (Arcus II; Agfa-Gavaert, Morstel, Belgium) and quantified with the National Institutes of Health Image program (accessible online at rsb.info.nih.gov/nih-image/). All data are presented as means ± SE. The effects of diets and insulin were compared by a two-way analysis of variance. Differences were considered to be statistically significant at P < 0.05.

Physiological parameters of high fat–fed rats.

As expected, feeding rats a high-fat diet for 4 weeks resulted in increased adiposity, hyperinsulinemia, and moderate hyperglycemia (Table 1). High-fat diet–mediated loss of insulin sensitivity was evidenced by the decrease in glucose infusion rate (∼30%) during the euglycemic-hyperinsulinemic clamp (Table 1). These results are in close agreement with previous studies showing the diabetogenic effect of a high-fat diet (24,25,26,27).

Glucose transport and GLUT4 expression in individual muscles.

The effect of fat feeding on GLUT4 expression and glucose transport in muscles enriched in different fiber types is presented in Figs. 1A and B. Total GLUT4 content was greater (∼2.5-fold) in type I- and IIa-enriched muscles (soleus and red tibialis, respectively) compared with type IIb-enriched muscles (white gastrocnemius) in chow-fed rats. These results are consistent with our previous observations that GLUT4 expression is higher in oxidative than glycolytic fibers (39). In rats fed a high-fat diet, GLUT4 content in type I- and IIb-enriched muscle was found to be similar to chow-fed rats. However, GLUT4 abundance was reduced in type IIa-enriched muscles (∼40%) (Fig. 1A), and this observation was confirmed in the muscles used for the GLUT4 translocation assay (mixed gastrocnemius and quadriceps), which are enriched (∼50%) with type IIa fibers (Fig. 2A). Despite the latter finding, we observed similar reductions (∼40–60%) in insulin-stimulated glucose transport in all skeletal muscles tested in high fat–fed rats (Fig. 1B), whereas basal glucose transport was unaltered by fat feeding (data not shown). These results indicate that the reduced expression of GLUT4 is not the principal cause of impaired insulin-stimulated glucose transport in skeletal muscle of high fat–fed rats.

GLUT4 translocation to the plasma membrane and the T-tubules.

We next investigated the effect of high-fat feeding on GLUT4 translocation to both cell surface compartments of muscle cells (i.e., the plasma membrane and the T-tubules) to precisely determine the locus of insulin resistance in this animal model (Figs. 2B–D). The characteristics of the subcellular membrane fractions are presented in Table 2 and are in good agreement with previous studies (3,34,40). Insulin stimulation induced translocation of GLUT4 from the intracellular membranes (−35%, from 330 ± 58 to 215 ± 28 relative densitometric units [RDU], P < 0.05) to the plasma membrane (+100%, from 98 ± 18 to 196 ± 28 RDU, P < 0.05) and the T-tubules (+35%, from 53 ± 7 to 71 ± 5 RDU, P < 0.05) in control rats fed a standard chow diet. In marked contrast, insulin failed to induce GLUT4 translocation to either the plasma membrane or the T-tubules in skeletal muscle of rats fed the high-fat diet (Figs. 2B–D). These results clearly show that the reduced insulin-stimulated glucose uptake in muscle of high fat–fed rats is linked to a defective translocation of GLUT4 glucose transporters to both cell surface compartments of skeletal muscle cells.

Tyrosine phosphorylation of IR and IRS proteins.

To investigate whether high-fat feeding causes insulin resistance via the alteration of an early insulin signaling step, we measured insulin-induced tyrosine phosphorylation of the IR and IRS proteins in anti-phosphotyrosine immune complexes. In rats fed either the control chow or the high-fat diets, insulin stimulated the tyrosine phosphorylation of IR and IRS proteins (approximately six- and threefold, respectively) (Figs. 3A and B). Thus, high-fat feeding for 4 weeks did not affect the stimulatory effect of insulin on IR/IRS tyrosine phosphorylation in skeletal muscle.

PI 3-kinase activity.

Although high-fat feeding did not alter the activation of proximal events in insulin signaling (IR/IRS), we next evaluated insulin-induced activation of PI 3-kinase, a lipid kinase that mediates most of the metabolic action of insulin (10). Protein levels of PI 3-kinase p85 subunit were similar in muscle from chow- and high fat–fed rats (0.92 ± 0.04 and 0.81 ± 0.05 RDU, respectively; NS). The kinase activity of the enzyme was measured in anti–IRS-1 precipitates because IRS-1 is the main isoform responsible for insulin-stimulated glucose transport in skeletal muscle (41,42). In the basal state, PI 3-kinase activity was not different among the dietary groups (Fig. 4). Following insulin stimulation, PI 3-kinase activity in the muscle of control rats was markedly increased (approximately eightfold). However, in muscle of rats fed the high-fat diet, insulin-dependent PI 3-kinase activation was severely attenuated (∼60% reduction versus chow-fed rats, P < 0.05).

Akt phosphorylation and activity.

Because impairment of PI 3-kinase stimulation by insulin may lead to a concomitant decrease in the activation of downstream effector(s) in rats fed a high-fat diet, we next measured insulin-induced phosphorylation of Akt using phospho-specific (Ser473 and Thr308) antibodies. Akt phosphorylation was robustly enhanced by insulin in skeletal muscle of rats fed the standard chow diet. The effect of insulin was similar in high fat–fed rats (Fig. 5A). We then assessed the kinase activity of the enzyme in anti-Akt immunoprecipitates using Crosstide (a peptide containing a glycogen synthase kinase-3 motif known to be phosphorylated by Akt) as substrate (43). We observed that basal Akt kinase activity tended to be reduced (∼50%, P = 0.16) in high fat–fed animals compared with control rats. Insulin increased Akt kinase activity by ∼2.2-fold in both dietary groups compared with their respective basal activity (Fig. 5B). However, maximal activation of Akt (insulin-treated groups) was reduced by ∼40% (P < 0.01) in rats fed the high-fat diet compared with chow-fed rats (Fig. 5B). Furthermore, the reduced Akt kinase activity could not be attributed to a decrease in Akt protein expression in the muscle of high fat–fed rats (0.93 ± 0.10 and 0.82 ± 0.07 RDU for chow- and high fat–fed rats, respectively; NS).

aPKC activity and translocation.

Another downstream effector of the PI 3-kinase pathway is the diacylglycerol- and calcium-insensitive aPKC (ζ/λ). We first examined if insulin stimulated the kinase activity of aPKC in skeletal muscle. We found that insulin stimulates aPKC activity in anti-PKC (ζ/λ) complexes by 2.2-fold (P < 0.01), as measured by 32P incorporated in myelin basic protein (Fig. 6A). We next determined whether defective PI 3-kinase activity in skeletal muscle of high fat–fed rats was linked to impaired aPKC activation by insulin. In marked contrast to the situation observed in chow-fed animals, aPKC activity was already elevated in control muscles of high fat–fed rats. Moreover, insulin could not further activate the enzyme in the muscle of these insulin-resistant animals. aPKC has been shown to be translocated to membranes when activated by insulin (14). Accordingly, we also found that insulin increases the association of aPKC with the plasma membrane (∼50%, P < 0.01) in the skeletal muscle of chow-fed rats after a hyperinsulinemic clamp (Fig. 6B). As observed for aPKC kinase activity, high-fat feeding in rats was found to increase basal aPKC association with the plasma membrane as compared with chow-fed rats (+35%, P < 0.05), and insulin infusion failed to further increase this association in high fat–fed rats. Muscle expression of aPKC protein was similar between both groups (1.11 ± 0.36 and 1.01 ± 0.15 RDU for chow- and high fat–fed rats, respectively; NS).

The cellular mechanism(s) responsible for impaired insulin-stimulated glucose uptake in the peripheral tissues of insulin-resistant subjects is still unclear. In the present study, we used the high fat–fed rat model to clarify the cellular defects behind the occurrence of skeletal muscle insulin resistance. We first looked at possible alterations in GLUT4 expression and/or translocation because both are important determinants of glucose uptake in adipocytes and muscle cells (1). Whereas impaired glucose transport in response to insulin in the adipose tissue of high fat–fed rats has been attributed to decreased GLUT4 content (44,45), discrepancies still exist regarding whether fat feeding alters GLUT4 expression in skeletal muscle (24,27,28,29,30,31). These discrepant findings may be partly explained by the fact that different types of muscles were used in these studies.

In this study, we found that feeding a high-fat diet caused a selective downregulation of GLUT4 in type IIa-enriched muscle. Despite this, total GLUT4 content does not appear to predict the extent of insulin resistance in the muscle of high fat–fed rats. As shown in this study and in the results from other studies (27,32), the cellular localization of GLUT4, rather than its amount, is the principal determinant of impaired insulin-stimulated glucose transport in the muscle fibers of high fat–fed rats. We previously observed that GLUT4 translocation was selectively impaired to the T-tubules but normal to the plasma membrane of muscle from insulin-resistant type 1 diabetic rats (40). In this study, we found that impaired GLUT4 translocation in rats fed a high-fat diet was generalized to both cell surface compartments. There are currently no data available concerning insulin-induced GLUT4 translocation to the T-tubules in insulin-resistant subjects. Regulation of GLUT4 translocation to this surface compartment is particularly important because the T-tubules cover most of the cell surface area in muscle cells. Surprisingly, the extent of insulin resistance on GLUT4 translocation (not detectable) in high fat–fed animals was more pronounced than that observed for glucose transport (40–60% inhibition). This discrepancy cannot be explained by a greater contribution from the GLUT1 transporter in the muscle of high fat–fed rats because its levels were also found to be reduced (by 49%, P < 0.05) in the plasma membrane of high fat–fed animals (GLUT1 was not detectable in the T-tubules). Recently, Ryder et al. (8) reported similar results in the muscle of type 2 diabetic subjects, showing that the inhibition of GLUT4 translocation (∼90%) did not match the reduction of insulin-stimulated glucose transport (∼50%). These authors suggested that another insulin-sensitive glucose transporter (e.g., the newly described GLUTX1 [46]) may have partly rescued the lack of GLUT4 translocation to the cell surface. Alternatively, one might speculate that in high fat–fed rats, insulin is still able to promote glucose transport by increasing the intrinsic activity of cell surface GLUT4 via the p38 mitogen-activated protein (MAP) kinase pathway (47). Indeed, it has been recently reported that SB 203580, an inhibitor of the p38 MAP kinase pathway, decreased glucose transport activity in L6 myotubes and in rat skeletal muscle (47,48) without interfering with GLUT4 translocation and insertion at the cell surface, implying that it inhibited GLUT4 activation (47).

There are currently two models that have been proposed to explain the defective insulin-induced activation of glucose transport in the high fat–feeding model of insulin resistance. On one hand, it has been reported that an alteration of insulin signaling (i.e., activation of PI 3-kinase) was an early occurrence in the pathogenesis of impaired glucose transport in the skeletal muscle of mice fed a high-fat diet (27). On the other hand, it has been suggested that defects in insulin signaling is a late event and is not the primary defect causing muscle insulin resistance in high fat–fed animals (32). The latter model was based on the observation that IR function and IRS-1 phosphorylation were not affected in rats fed a high-fat diet for 8 weeks, despite significant reductions in insulin-mediated GLUT4 translocation. However, PI 3-kinase activity was not assessed in the latter study.

In the present study, we found that impaired insulin-stimulated glucose transport and GLUT4 translocation in skeletal muscle of high fat–fed rats was associated with defective PI 3-kinase activation. Furthermore, we confirmed that this defect occurred without alteration of IR/IRS-1 tyrosine phosphorylation in the high-fat feeding model of insulin resistance (32). The latter finding is consistent with the fact that impairments of IR/IRS-1 signaling occurred later and are not the primary cause of decreased insulin-stimulated glucose transport in this model. A putative mechanism for the impairment of PI 3-kinase is an increased serine phosphorylation of IRS-1, which in turn would act as a inhibitor of PI 3-kinase function. For instance, 14–3-3 protein has been shown to bind to phosphoserine residues of IRS-1 and subsequently inhibit insulin-stimulated PI 3-kinase despite normal tyrosine phosphorylation of IRS-1 and binding of p85 regulatory subunit of PI 3-kinase (49). Another possibility is that serine phosphorylation of the p85 subunit of PI 3-kinase is increased, which has been reported to decrease the lipid kinase activity of the enzyme (50). In rats, injection of angiotensin II impaired insulin-mediated PI 3-kinase activation via increased serine phosphorylation of p85 without any change in the level of IR/IRS-1 tyrosine phosphorylation (51,52). Furthermore, increased serine kinase activity has been observed in insulin-resistant states (53).

Intense interest has been focused on the identification of those signaling steps downstream of PI 3-kinase that may be implicated in glucose transport activation by insulin. One of these is the serine-threonine kinase Akt, which has been shown to be involved in the control of glucose transport at a step downstream of PI 3-kinase (10). However, it is still unclear whether insulin-stimulated Akt activity is impaired in insulin-resistant skeletal muscle. Insulin activation of Akt in the muscle of glucosamine-infused rats (21), as well as in human diabetic subjects (20), was found to be normal. However, insulin-stimulated Akt in the muscle of type 2 diabetes subjects was reported to be impaired at a maximal dose of insulin in a separate study (23). In the present study, we found that insulin-dependent Akt kinase activity is reduced in skeletal muscle of high fat–fed rats, despite normal phosphorylation of the enzyme on both Ser473 and Thr308. Whether this alteration represents a primary defect leading to impaired GLUT4 translocation in this animal model is still uncertain. In L6 myoblasts, it has been shown that a >50% decrease in Akt activity does not significantly affect GLUT4 translocation (18). If such data could be extrapolated to rat skeletal muscle, this would suggest that the reduced Akt activity is unlikely to be responsible for the lack of insulin action in muscle of high fat–fed animals. Nevertheless, Akt kinase activity in insulin-stimulated muscle of high fat–fed rats was reduced by as much as 40% compared with insulin-stimulated chow-fed controls. Thus, although Akt activation by insulin appears normal based on Akt phosphorylation status, the actual ability of the kinase to phosphorylate an exogenous substrate is significantly altered. The mechanism behind this defect in Akt kinase activity remains to be determined.

Another downstream target of PI 3-kinase that has been suggested to be involved in the regulation of glucose transport is the atypical member of the PKC family. It has been shown to be activated by insulin in rat adipocytes (54), 3T3-L1 adipocytes, (19,55) and L6 myocytes (15). The use of inhibitor (pseudosubstrate), transfection of kinase-inactive or overexpression/constitutively active forms of the kinase, as well as microinjection of antibodies against aPKC, argued for its role in insulin-regulated glucose transport (15,16,19,54,55,56). However, to the best of our knowledge, there is as yet no experimental evidence for insulin-dependent activation of aPKC in skeletal muscle, the main target for insulin-stimulated glucose disposal. In this study, we provide evidence that insulin stimulates aPKC activity in skeletal muscle. This effect was demonstrated by measurements of direct kinase activity as well as by increased membrane association of aPKC. These data are consistent with the proposition that aPKC is involved in the stimulatory effect of insulin on glucose transport in skeletal muscle. More importantly, we found that aPKC was unresponsive to the action of insulin in skeletal muscle of high fat–fed rats. The fact that insulin-induced aPKC membrane association was also impaired in muscle from insulin-infused (euglycemic-clamped) high fat–fed animals further suggests that the lack of insulin action on aPKC is sustained for at least 2 h of insulin exposure. Impaired insulin-dependent activation of PI 3-kinase is likely to explain part of the failure of insulin to activate aPKC in rats fed the high-fat diet. A recent report by Kanoh et al. (57) showed that activation of aPKC was resistant to the action of insulin in adipocytes isolated from diabetic animals, which was restored by thiazolidinedione treatment with a concomitant increase in insulin-stimulated glucose transport. The latter finding, together with our results, supports a role for impaired aPKC signaling in mediating insulin-resistant glucose transport in both adipose tissue and skeletal muscle. Interestingly, we also found that basal (non–insulin-stimulated) aPKC activity and membrane localization are increased in high fat–fed animals. It is not clear at the present time why aPKC activity in the absence of insulin is abnormally elevated in high fat–fed rats. This increase in aPKC activity was observed even though basal glucose uptake was not affected by high-fat feeding, although this may be explained by the observation of decreased levels of both GLUT1 and GLUT4 in muscle from these animals. Whereas the higher insulin levels in high fat–fed rats may partly explain the enhanced basal aPKC activity, the fact that neither PI 3-kinase nor Akt basal activities were increased in the same muscles suggest that hyperinsulinemia is unlikely to be the main factor involved. Another possibility is that other factors in high fat–fed rats could have elevated muscle aPKC activity. Indeed, aPKCs are involved in many cellular processes, and it is therefore possible that only a small fraction of the total pool of aPKCs participates in the stimulation of glucose transport. Among other roles of aPKC in cellular signaling, it has been shown to participate in tumor necrosis factor-α (TNF-α)-induced formation of ceramides by sphingomyelinase (58,59). Although muscle TNF-α levels were found to be similar between both dietary groups (0.018 ± 0.02 vs. 0.020 ± 0.02 pg/μg protein for chow- and high fat–fed rats, respectively), we confirmed previous reports of overexpression of the cytokine in white adipose tissue of obese animals (0.91 ± 0.35 vs. 1.76 ± 0.36 pg/μg DNA for chow- and high fat–fed rats, respectively; P < 0.05) (60). It may therefore be speculated that local production of TNF-α by surrounding adipose tissue increases basal aPKC activity in muscle and subsequently makes it unresponsive to the action of insulin in high fat–fed rats.

In summary, the present study provides an extensive characterization of the insulin signal transduction pathway in skeletal muscle of the high fat–fed rat model of insulin resistance. These animals showed a complete absence of GLUT4 translocation in response to insulin not only to the plasma membrane but also to the T-tubules, the major component the muscle cell surface. We identified PI 3-kinase as the first step of the insulin signaling pathway to be altered by fat feeding. This was associated with reduced Akt kinase activity in insulin-stimulated muscle, despite normal Akt phosphorylation. Moreover, we characterized for the first time the insulin-dependent activation and translocation of aPKC in normal and insulin-resistant skeletal muscle. We found a complete failure of insulin to activate aPKC in high fat–fed animals. Thus, alterations in both Akt and aPKC signaling may be involved in the PI 3-kinase–dependent impairment in GLUT4 translocation in the skeletal muscle of high fat–fed rats.

FIG. 1.

Comparison between GLUT4 protein expression and insulin-stimulated glucose uptake in muscles enriched in different fiber types. A: GLUT4 protein expression was measured in muscle homogenates from type I-, type IIa-, and type IIb- (soleus, red tibialis, and white gastrocnemius, respectively) enriched muscles of chow- and high fat–fed rats. Proteins (50 μg) were resolved on 7.5% SDS-PAGE and immunoblotted with polyclonal anti-GLUT4, as described in research design and methods. The means ± SE from 4–10 determinations for each muscle are shown. B: In vivo 2-[3H]DG uptake in muscle fibers was determined after bolus injection of 2-[3H]DG and 14C-sucrose in insulin-infused rats (four mU · kg−1 · min−1) at the end of the euglycemic-hyperinsulinemic clamp as described under research design and methods. The means ± SE from 7–9 determinations are shown. □, chow-fed; ▪, high fat–fed. #P < 0.05 vs. chow-fed rats.

FIG. 1.

Comparison between GLUT4 protein expression and insulin-stimulated glucose uptake in muscles enriched in different fiber types. A: GLUT4 protein expression was measured in muscle homogenates from type I-, type IIa-, and type IIb- (soleus, red tibialis, and white gastrocnemius, respectively) enriched muscles of chow- and high fat–fed rats. Proteins (50 μg) were resolved on 7.5% SDS-PAGE and immunoblotted with polyclonal anti-GLUT4, as described in research design and methods. The means ± SE from 4–10 determinations for each muscle are shown. B: In vivo 2-[3H]DG uptake in muscle fibers was determined after bolus injection of 2-[3H]DG and 14C-sucrose in insulin-infused rats (four mU · kg−1 · min−1) at the end of the euglycemic-hyperinsulinemic clamp as described under research design and methods. The means ± SE from 7–9 determinations are shown. □, chow-fed; ▪, high fat–fed. #P < 0.05 vs. chow-fed rats.

Close modal
FIG. 2.

Effect of high fat–feeding on GLUT4 translocation in skeletal muscle. Rats were clamped with either saline (basal) or insulin (4 mU · kg−1 · min−1) for 2 h. Immediately after the clamp, muscles (mixed gastrocnemius and quadriceps) were quickly excised, cleaned of extraneous tissues, and frozen in liquid nitrogen. GLUT4 content was assessed by Western blotting in total homogenate (A), plasma membrane (B), T-tubules (C), and intracellular membranes (D) isolated from saline- or insulin-infused rats as described in research design and methods. The means ± SE from 4–5 individual membrane preparations are shown. □, Basal; ▪, insulin. IM, intracellular membranes; PM, plasma membrane; TT, T-tubules. #P < 0.05 vs. chow-fed rats. *P < 0.05 vs. corresponding basal value.

FIG. 2.

Effect of high fat–feeding on GLUT4 translocation in skeletal muscle. Rats were clamped with either saline (basal) or insulin (4 mU · kg−1 · min−1) for 2 h. Immediately after the clamp, muscles (mixed gastrocnemius and quadriceps) were quickly excised, cleaned of extraneous tissues, and frozen in liquid nitrogen. GLUT4 content was assessed by Western blotting in total homogenate (A), plasma membrane (B), T-tubules (C), and intracellular membranes (D) isolated from saline- or insulin-infused rats as described in research design and methods. The means ± SE from 4–5 individual membrane preparations are shown. □, Basal; ▪, insulin. IM, intracellular membranes; PM, plasma membrane; TT, T-tubules. #P < 0.05 vs. chow-fed rats. *P < 0.05 vs. corresponding basal value.

Close modal
FIG. 3.

Effect of high-fat feeding on insulin-induced tyrosine phosphorylation of IR/IRS proteins. Overnight-fasted rats were injected with either saline or insulin (8 units/kg) for 4 min. Phosphoproteins were immunoprecipitated from muscle homogenates, resolved on 6% SDS-PAGE, and immunoblotted using anti-phosphotyrosine (4G10) as described in research design and methods. Quantification of tyrosine phosphorylation of the IR (A) and IRS (B) proteins was expressed relative to chow-fed basal values. Representative immunoblots are shown at the top of each panel. The location of molecular weight markers is shown on the right. The means ± SE from 4–6 determinations from different animals are shown. □, Basal; ▪, insulin.

FIG. 3.

Effect of high-fat feeding on insulin-induced tyrosine phosphorylation of IR/IRS proteins. Overnight-fasted rats were injected with either saline or insulin (8 units/kg) for 4 min. Phosphoproteins were immunoprecipitated from muscle homogenates, resolved on 6% SDS-PAGE, and immunoblotted using anti-phosphotyrosine (4G10) as described in research design and methods. Quantification of tyrosine phosphorylation of the IR (A) and IRS (B) proteins was expressed relative to chow-fed basal values. Representative immunoblots are shown at the top of each panel. The location of molecular weight markers is shown on the right. The means ± SE from 4–6 determinations from different animals are shown. □, Basal; ▪, insulin.

Close modal
FIG. 4.

Effect of high-fat feeding on PI 3-kinase activity. PI 3-kinase was measured in anti–IRS-1 immunoprecipitates of muscle homogenate as described in research design and methods. Quantification of 32P incorporated into PI 3-phosphate was expressed relative to chow-fed basal values. A representative autoradiograph is shown at the top of the figure. The means ± SE of 4–6 determinations from different animals are shown. □, Basal; ▪, insulin. *P < 0.05 vs. corresponding basal value; #P < 0.05 vs. chow-fed insulin values. PI(3)P, PI 3-phosphate.

FIG. 4.

Effect of high-fat feeding on PI 3-kinase activity. PI 3-kinase was measured in anti–IRS-1 immunoprecipitates of muscle homogenate as described in research design and methods. Quantification of 32P incorporated into PI 3-phosphate was expressed relative to chow-fed basal values. A representative autoradiograph is shown at the top of the figure. The means ± SE of 4–6 determinations from different animals are shown. □, Basal; ▪, insulin. *P < 0.05 vs. corresponding basal value; #P < 0.05 vs. chow-fed insulin values. PI(3)P, PI 3-phosphate.

Close modal
FIG. 5.

Effect of high-fat feeding on Akt/PKB phosphorylation and activity. Overnight fasted rats were injected with either saline or insulin (8 units/kg) for 4 min. A: Phosphorylation state of Akt (Ser473 and Thr308) was measured in muscle homogenates. Protein (50 μg) was separated on 7.5% SDS-PAGE and immunoblotted with anti–phospho-specific antibody against Akt as described in research design and methods. B: Akt kinase activity was measured in anti–Akt-1/2 immunoprecipitates as described in research design and methods. Quantification of 32P incorporated into Crosstide was expressed relative to chow-fed basal values. Representative immunoblot (A) and autoradiograph (B) are shown at the top of each panel. The means ± SE of 4–6 determinations from different animals are shown. □, Basal; ▪, insulin. *P < 0.05 vs. corresponding basal value; #P < 0.01 vs. chow-fed insulin values.

FIG. 5.

Effect of high-fat feeding on Akt/PKB phosphorylation and activity. Overnight fasted rats were injected with either saline or insulin (8 units/kg) for 4 min. A: Phosphorylation state of Akt (Ser473 and Thr308) was measured in muscle homogenates. Protein (50 μg) was separated on 7.5% SDS-PAGE and immunoblotted with anti–phospho-specific antibody against Akt as described in research design and methods. B: Akt kinase activity was measured in anti–Akt-1/2 immunoprecipitates as described in research design and methods. Quantification of 32P incorporated into Crosstide was expressed relative to chow-fed basal values. Representative immunoblot (A) and autoradiograph (B) are shown at the top of each panel. The means ± SE of 4–6 determinations from different animals are shown. □, Basal; ▪, insulin. *P < 0.05 vs. corresponding basal value; #P < 0.01 vs. chow-fed insulin values.

Close modal
FIG. 6.

Effect of high-fat feeding on aPKC activity and translocation. Overnight-fasted rats were injected either with saline or insulin (8 units/kg) for 4 min. A: aPKC kinase activity was measured in anti-PKC (ζλ) immunoprecipitates as described in research design and methods. Quantification of 32P incorporated into myelin basic protein was expressed relative to chow-fed basal values. B: Membrane recovery of aPKC in control and insulin-infused rats was assessed by Western blotting as described in research design and methods. Representative autoradiograph (A) and immunoblot (B) are shown at the top of each panel. The location of molecular weight markers is shown on the right. The means ± SE of 4–6 determinations from different animals are shown. □, Basal; ▪, insulin. *P < 0.01 vs. corresponding basal value; #P < 0.05 vs. chow-fed basal values.

FIG. 6.

Effect of high-fat feeding on aPKC activity and translocation. Overnight-fasted rats were injected either with saline or insulin (8 units/kg) for 4 min. A: aPKC kinase activity was measured in anti-PKC (ζλ) immunoprecipitates as described in research design and methods. Quantification of 32P incorporated into myelin basic protein was expressed relative to chow-fed basal values. B: Membrane recovery of aPKC in control and insulin-infused rats was assessed by Western blotting as described in research design and methods. Representative autoradiograph (A) and immunoblot (B) are shown at the top of each panel. The location of molecular weight markers is shown on the right. The means ± SE of 4–6 determinations from different animals are shown. □, Basal; ▪, insulin. *P < 0.01 vs. corresponding basal value; #P < 0.05 vs. chow-fed basal values.

Close modal
TABLE 1

Physiological parameters of rats fed a Rodent Chow– or a high fat–diet

Rodent ChowHigh-Fat
Body weight (g) 351 ± 20 380 ± 20 
Epididymal fat pad (g) 1.92 ± 0.10 3.33 ± 0.30* 
Retroperitoneal fat pad (g) 1.17 ± 0.15 2.67 ± 0.35* 
Fasting glucose (mmol/l) 7.2 ± 0.3 8.9 ± 0.2* 
Fasting insulin (nmol/l) 0.13 ± 0.02 0.25 ± 0.03* 
GIR (mg · kg–1 · min–116.9 ± 2.0 12.2 ± 1.1* 
Rodent ChowHigh-Fat
Body weight (g) 351 ± 20 380 ± 20 
Epididymal fat pad (g) 1.92 ± 0.10 3.33 ± 0.30* 
Retroperitoneal fat pad (g) 1.17 ± 0.15 2.67 ± 0.35* 
Fasting glucose (mmol/l) 7.2 ± 0.3 8.9 ± 0.2* 
Fasting insulin (nmol/l) 0.13 ± 0.02 0.25 ± 0.03* 
GIR (mg · kg–1 · min–116.9 ± 2.0 12.2 ± 1.1* 

Values are means ± SE. GIR, glucose infusion rate.

*

P < 0.05 vs. chow value.

TABLE 2

Characterization of membrane fractions from skeletal muscle

FractionsDietInsulinProtein recoveries (μg/g muscle)5′-nucleotidase (nmol · mg–1 · min–1)
PM Rodent Chow − 30 ± 5 557 ± 57 
 Rodent Chow 30 ± 5 546 ± 74 
 High-Fat − 38 ± 5 417 ± 138 
 High-Fat 31 ± 4 355 ± 77 
T-tubules Rodent Chow − 234 ± 24 90 ± 7 
 Rodent Chow 279 ± 12 72 ± 8 
 High-Fat − 258 ± 10 59 ± 14 
 High-Fat 265 ± 24 52 ± 14 
 Rodent Chow − 95 ± 21 ND 
IM Rodent Chow 99 ± 27 ND 
 High-Fat − 136 ± 19 ND 
 High-Fat 219 ± 45 ND 
FractionsDietInsulinProtein recoveries (μg/g muscle)5′-nucleotidase (nmol · mg–1 · min–1)
PM Rodent Chow − 30 ± 5 557 ± 57 
 Rodent Chow 30 ± 5 546 ± 74 
 High-Fat − 38 ± 5 417 ± 138 
 High-Fat 31 ± 4 355 ± 77 
T-tubules Rodent Chow − 234 ± 24 90 ± 7 
 Rodent Chow 279 ± 12 72 ± 8 
 High-Fat − 258 ± 10 59 ± 14 
 High-Fat 265 ± 24 52 ± 14 
 Rodent Chow − 95 ± 21 ND 
IM Rodent Chow 99 ± 27 ND 
 High-Fat − 136 ± 19 ND 
 High-Fat 219 ± 45 ND 

Values are means ± SE. PM, plasma membranes; IM, Intracellular membranes; ND, nondetectable.

This work was supported by grants from the Canadian Diabetes Association (H.J. and A.M.). A.M. was supported by scholarships from the Medical Research Council of Canada and the Fonds de la Recherche en Santé du Québec. F.T. was supported by a studentship from the Québec Hypertension Society.

We thank Luce Dombrowski for expert technical assistance. We are grateful to Romel Somwar, Dr. Philip Bilan, and Dr. Amira Klip for their helpful advice on the PI 3-kinase assay and for critical reading of the manuscript.

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Address correspondence and reprint requests to André Marette, Department of Physiology and Lipid Research Unit, Laval University Hospital Research Center, 2705, Laurier Blvd., Ste-Foy, Québec, Canada, G1V 4G2. Email: andre.marette@crchul.ulaval.ca.

Received for publication 30 June 2000 and accepted in revised form 7 May 2001.

aPKC, atypical protein kinase C; DTT, dithiothreitol; 2-[3H]DG, 2-deoxy-d-[3H]glucose; IR, insulin receptor; IRS, insulin receptor substrate; MAP, mitogen-activated protein; PBS, phosphate-buffered saline; PDK, 3-phosphoinositide-dependent kinase; PI, phosphatidylinositol; PKB, protein kinase B; PKC, protein kinase C; PVDF, polyvinylidene difluoride; RAC, related to A and C; RDU, relative densitometric units; T, transverse; TNF-α, tumor necrosis factor-α.