Isoform-Specific Regulation of 5′ AMP-Activated Protein Kinase in Skeletal Muscle From Obese Zucker (fa/fa) Rats in Response to Contraction

Obese male Zucker (fa/fa) rats (aged 11–12 weeks) and lean littermates were purchased from Charles River (Uppsala, Sweden) and housed at the animal facility at the Karolinska Institute for 4 weeks before use. Rats were maintained on a 12-h light-dark cycle and given free access to standard rodent chow and water. All studies we performed on rats in the overnight-fasted state. The regional animal ethical committee approved all experimental procedures.

Glucose (2 g/kg body wt) was administered by intraperitoneal injection. Blood samples were obtained via the tail vein before and 15, 30, 60, and 120 min after glucose injection. Blood glucose levels were measured using a One Touch Basic glucose meter (Lifescan, Milpitas, CA).

All incubation media were prepared from a stock solution of Krebs-Henseleit bicarbonate buffer (KHBB) supplemented with 5 mmol/l HEPES and 0.1% BSA (radioimmunoassay grade). Rats were anesthetized with sodium pentobarbital (60 mg/kg body wt), and epitrochlearis muscles were carefully isolated for in vitro incubation. The viability of the incubated muscle preparation from large rats has been previously established by measuring high-energy phosphate levels of epitrochlearis muscle from adult rats that are of similar weight to the animals used in our study (26). Muscles were preincubated at 30°C for 40 min in KHBB containing 5 mmol/l glucose and 15 mmol/l mannitol, without (basal) or with insulin (60 nmol/l) for either 6 min (PI 3-kinase activity) or 40 min (glucose transport activity). Unless otherwise stated, all media were continuously gassed with 95% O₂/5% CO₂. After preincubation, muscles were immediately frozen between aluminum tongs cooled to the temperature of liquid nitrogen or further incubated to assess glucose transport activity.

Epitrochlearis muscles were placed in a temperature-controlled (30°C) incubation chamber and immersed in 4 ml preincubation media without or with insulin as described above. Muscles were positioned between two platinum electrodes. The distal tendon was mounted to the bottom of the incubation chamber. The proximal tendon was connected to an isometric force transducer (Harvard Apparatus, South Natick, MA), and resting tension was adjusted to 0.5 g. Electrical stimulation (contraction group only) was applied during the final 10 min of the 40-min preincubation period. Basal and insulin-stimulated muscles were treated as described above minus the application of electrical stimulation. Muscle contraction was induced via electrical stimulation. Muscles were stimulated at 100 Hz (0.2-ms pulse duration, 20 V), at a rate of one 0.2-s contraction every 2 s for 10 min, as previously described (27). Muscles were frozen immediately for AMPK or PI 3-kinase activity measurements or further incubated for the assessment of glucose transport activity.

Epitrochlearis muscles were preincubated at 30°C for 80 min in KHBB containing 5 mmol/l glucose and 15 mmol/l mannitol, under a gas phase of 95% O₂/5% CO₂ (basal) or 95% N₂/5% CO₂ (hypoxia) (28). Muscles were frozen immediately for AMPK activity or further incubated for the assessment of glucose transport activity.

After preincubation, muscles were incubated at 30°C for 10 min in glucose-free KHBB containing 20 mmol/l mannitol. When effects of hypoxia were studied, muscles were incubated for 15 min in KHBB containing 2 mmol/l pyruvate and 18 mmol/l mannitol under a gas phase of 95% O₂/5% CO₂. Thereafter, muscles were transferred to vials containing 1 mmol/l 2-deoxy-[1,2,³H]glucose (2.5 μCi/ml) and 19 mmol/l [¹⁴C]mannitol (0.7 μCi/ml) without or with insulin and incubated for 15 min. The rate of 2-deoxyglucose uptake is linear between 5 and 120 min in rat epitrochlearis muscle (29). Glucose transport activity is expressed as micromoles per milliliter of intracellular water per hour (29).

Muscles were homogenized in ice-cold lysis buffer containing 50 mmol/l Tris-HCl (pH 7.5), 1 mmol/l EDTA, 1 mmol/l diothiothreitol (DTT), 10% (vol/vol) glycerol, 50 mmol/l NaF, 5 mmol/l Na pyrophosphate, 1 mmol/l benzamidine, 0.1 mmol/l phenylmethylsulfonyl fluoride (PMSF), and 1% (vol/vol) Triton X-100. Muscle homogenates were subjected to centrifugation at 14,000g for 10 min at 4°C. Supernatants were removed and used for determination of protein content using a commercially available kit based on the Bradford method (Bio-Rad, Hercules, CA). Aliquots (400 μg protein) were sequentially immunoprecipitated for 2 h at 4°C with sheep AMPKα1 and -α2 antibodies precoupled to protein G-Agarose (5 mg/sample). The antibodies were raised against peptides predicted from the rat AMPKα1 and -α2 amino acid sequences, as previously described (30). Immunoprecipitates were washed three times with lysis buffer and twice with 50 mmol/l HEPES (pH 7.5), 10% (vol/vol) glycerol, 1 mmol/l EDTA, and 1 mmol/l DTT. AMPK activity in the immune complex was determined by in vitro phosphorylation of the SAMS (full sequence: HMRSAMSGLHLVKRR) synthetic peptide substrate (30), as previously described (18). Kinase reactions performed in reaction buffer (40 mmol/l HEPES buffer, pH 7.0, 0.2 mmol/l SAMS peptide, 0.2 mmol/l AMP, 80 mmol/l NaCl₂, 0.8 mmol/l DTT, 5 mmol/l MgCl₂, and 0.2 mmol/l ATP (containing 2 μCi [γ-³²P]ATP). Reactions were incubated on a vibrating platform for 60 min at 30°C and terminated by centrifugation (9,000g for 30 s). Immediately thereafter, aliquots were spotted onto phosphocellulose units (Pierce, Rockford, IL). Phosphocellulose units were washed with 1% phosphoric acid according to the manufacturer specifications. Incorporation of [³²P] into peptide substrate was measured by liquid scintillation counting.

Epitrochlearis skeletal muscle was homogenized in ice-cold lysis buffer (20 mmol/l Tris, pH 8.0, 137 mmol/l NaCl, 2.7 mmol/l KCl, 1 mmol/l MgCl₂, 0.5 mmol/l Na₃VO₄, 1% Triton X-100, 10% vol/vol glycerol, 10 μg/ml leupeptin, 0.2 mmol/l PMSF, 10 mmol/l NaF, and 10 μg/ml aprotinin). Homogenates were subjected to centrifugation at 8,000g for 10 min at 4°C, and protein content was determined as described above. Supernatants (800 μg) were subjected to immunoprecipitation with a polyclonal anti-phosphotyrosine antibody (Signal Transduction Laboratories, Lexington, KY) coupled to protein A-Sepharose. PI 3-kinase activity was assayed for 20 min at room temperature as described (31). Reaction products were separated by thin-layer chromatography, and quantification of [ ³²P] incorporation into PI was determined using a phosphoimager (Bio-Rad Laboratories, Hercules, CA).

Differences between two groups were determined by unpaired t test. Differences between more than two groups were determined by one-way ANOVA followed by the Fisher’s least significant difference post hoc analysis. Significance was accepted at P < 0.05.

The body weight of 11-week-old obese Zucker rats was significantly greater than that of age-matched lean rats (393 ± 7 vs. 267 ± 3 g; P < 0.05). Epitrochlearis muscle weights were reduced in obese Zucker rats (42.2 ± 1.6 vs. 59.4 ± 2.1 mg, obese versus lean rats; P < 0.05). Obese Zucker rats displayed marked glucose intolerance, with blood glucose levels significantly elevated versus lean rats at all time points measured (P < 0.05) (Fig. 1).

Skeletal muscle glucose uptake was determined in response to insulin, contraction, or hypoxia. Insulin increased 2-deoxyglucose transport 3.4-fold in isolated skeletal muscle from lean rats (P < 0.05) (Fig. 2). Obese rats exhibited marked skeletal muscle insulin resistance. The absolute rate of insulin-stimulated glucose transport in obese rats was reduced 47% compared with lean controls (1.27 ± 0.08 vs. 0.68 ± 0.06 μmol · ml⁻¹ · h⁻¹; P < 0.05). Rates of contraction- and hypoxia-stimulated glucose transport in skeletal muscle were similar between lean and obese rats (Fig. 2). In lean rats, contraction- and hypoxia-stimulated glucose transport was increased 3.1-fold (P < 0.05) and 3.4-fold (P < 0.05), respectively. In insulin-resistant obese Zucker rats, contraction- and hypoxia-stimulated glucose transport was increased 3.9-fold (P < 0.05) and 4.1-fold (P < 0.05), respectively.

Phosphotyrosine-associated PI 3-kinase activity was determined in epitrochlearis skeletal muscle from lean and obese rats, in response to insulin or contraction. Insulin increased phosphotyrosine-associated PI 3-kinase activity 2.7-fold in lean rats (P < 0.05) (Fig. 3), with a 42% reduction noted in skeletal muscle from obese rats (P < 0.05 vs. lean). PI 3-kinase activity was not increased after muscle contraction.

Basal AMPKα1 (Fig. 4A) and AMPKα2 (Fig. 4B) activities were similar in skeletal muscle between lean and obese rats during experiments involving insulin and muscle contraction. Regardless of genetic background, insulin was without effect on AMPKα1 or -α2 activity. Contraction elicited an increase in activation of both the α1 and α2 isoforms in skeletal muscle. In lean rats, AMPKα1 and -α2 activity was increased 2.3- and 4.8-fold, respectively (P < 0.05). Interestingly, in skeletal muscle from obese rats, an isoform-specific effect of contraction on AMPK was noted: AMPKα1 activity was not altered (P = 0.10), whereas AMPKα2 activity was increased 4.3-fold (P < 0.05). Furthermore, when comparing responses between both genotypes, the contraction-mediated response of AMPKα1 tended to be reduced in obese rats (P < 0.08). In contrast, contraction-mediated AMPKα2 activity was comparable between lean and obese rats. In response to hypoxia, AMPKα1 (Fig. 5A) and -α2 (Fig. 5B) activity in lean rats was increased 6.6-fold (P < 0.05) and 5.5-fold (P < 0.05), respectively. In obese rats, hypoxia-stimulated AMPK was preserved, with increases in AMPKα1 and -α2 activity 8.4-fold (P < 0.05) and 6.5-fold (P < 0.05) over basal values, respectively. Basal AMPKα1 and -α2 activity in obese rats was slightly reduced (P < 0.05).

In skeletal muscle, separate and distinct signaling pathways can activate glucose transport. Insulin increases glucose transport via activation of PI 3-kinase (13,14) and possibly via activation of the CAP/TC10 pathway (32,33). However, the latter pathway has not been validated in skeletal muscle. Exercise (34), muscle contraction (35,36), and hypoxia (28,37) increase glucose transport via insulin-independent pathways (13–15), presumably via activation of AMPK (17,18). In type 2 diabetic patients, defects in insulin-mediated whole-body glucose uptake are coupled to impairments in glucose transport in skeletal muscle (4,10), which arise from aberrant signal transduction at the level of insulin receptor substrate-1, PI 3-kinase (6,8,9), and GLUT4 translocation (10,38). Since muscle contraction and hypoxia increase glucose transport via an alternative mechanism that bypasses defective insulin signaling (37,39,40), strategies to identify and characterize components of this insulin-independent pathway will potentially reveal novel entry points that can be targeted to increase glucose uptake in insulin-resistant muscle. Recent interest has focused on AMPK, as it is the most distal signaling molecule identified in the insulin-independent regulation of glucose transport. AMPK mediates contraction- and hypoxia-regulated glucose transport in skeletal muscle (17) and has been proposed as a promising target for treatment of altered glucose homeostasis in type 2 diabetes (41). Contraction-stimulated glucose transport in skeletal muscle is normal in severely insulin-resistant obese Zucker rats (39,40); however, effects of contraction and hypoxia on isoform-specific activation of AMPK in skeletal muscle from this animal model have not been determined.

Consistent with earlier studies (39,40), insulin resistance at the level of PI 3-kinase and glucose transport was observed in skeletal muscle from obese Zucker rats, whereas stimulation of AMPKα2 activity and glucose transport in response to muscle contraction or hypoxia was normal. These findings are also consistent with observations in humans, whereby moderate aerobic exercise elicits an appropriate increase in AMPKα2 in skeletal muscle from type 2 diabetic subjects (42). Thus, contraction and hypoxia increase AMPKα2 activity and glucose transport in insulin-resistant skeletal muscle. These studies provide evidence to suggest that exercise- and hypoxia-induced AMPK activity is not impaired in insulin-resistant skeletal muscle. Furthermore, they support the recent observation in cultured skeletal muscle, indicating direct activation of AMPK alone is sufficient for stimulation of glucose uptake via an increase in cell surface expression of GLUTs (18).

An isoform-specific reduction in AMPKα1 activity in skeletal muscle from obese Zucker rats was observed in response to contraction, but not hypoxia. Contraction increased AMPKα1 activity in epitrochlearis muscle from lean rats, whereas no effect was observed in obese rats. In contrast, hypoxia-stimulated AMPKα1 activity was similar between lean and obese rats; however, basal AMPKα1 activity was slightly reduced. Interestingly, basal AMPKα1, although not significant, tended to be lower in skeletal muscle from type 2 diabetic subjects (42). A previous study provides evidence that total AMPK activity in calf muscles in response to AICAR infusion was similar between lean and obese Zucker rats (24). However, since total rather than isoform-specific AMPK activity was measured (24), subtle defects in AMPKα1 activity may have been masked.

An explanation for the differential response of AMPKα1 in response to contraction versus hypoxia in the present study may be related to the amount of cellular stress applied to the muscle. The hypoxia stimulus is likely to have elicited a greater degree of stress on the muscle (21). Thus, the AMPKα1 defect may represent reduced sensitivity of kinase to cellular stress. Physiological evidence for this comes from human studies in which AMPKα1 activity is increased in response to anaerobic (sprint) (43), but not aerobic (endurance) (44,45), cycle ergometry. The contraction protocol used in the present study is more closely related to anaerobic (high-intensity) rather than aerobic (low-intensity) exercise. An alternative possibility is that the lack of a contraction effect on AMPKα1 in obese muscle may be due to the fact that the contraction protocol was not intense enough, although it did stimulate glucose transport to the same extent. Despite the isoform-specific defect in AMPKα1 activity in response to contraction, normal glucose transport was achieved in skeletal muscle from insulin-resistant Zucker rats.

Hypoxia increases AMPK activity to a greater degree than in vitro muscle contraction (21). Since only subtle defects in contraction-induced AMPKα1 activity were observed in obese rats in the present investigation, the observed defect is not likely to have a major impact on the rate of glucose transport in response to muscle contraction. Importantly, full activation of AMPK may not be necessary for complete activation of exercise/contraction-stimulated glucose transport. Transgenic overexpression of a dominant inhibitory mutant of AMPK in skeletal muscle completely blocks the ability of hypoxia or AICAR to activate glucose uptake while only partially (30–40%) reducing contraction-stimulated glucose uptake (17). Although these results (17) confirm a role of AMPK in contraction-induced glucose transport, they also demonstrate that AMPK is only partially responsible for this effect. Thus, AMPK-dependent and -independent pathways contribute to the regulation of glucose uptake in skeletal muscle in response to exercise (17). Consistent with this hypothesis, glucose transport in slow-twitch muscle can be markedly activated in response to contraction, without measurable changes in AMPK activity (46). Collectively, these studies illustrate the complexity in identifying the precise role of AMPK signaling in the regulation of metabolic events and they strongly suggest that additional factors contribute to the regulation of exercise-mediated glucose uptake.

In summary, obesity-related insulin resistance in skeletal muscle is associated with reduced contraction-stimulated AMPKα1 activity, whereas AMPKα2 activity is normal. However, when muscles were challenged by cellular hypoxia, activation of both AMPKα1 and -α2 were normal. In skeletal muscle from lean animals, hypoxia was more effective than contraction in stimulating AMPKα1 activity. Thus, the impairment in AMPKα1 in insulin-resistant muscle may be related to a reduced sensitivity of the enzyme in response to cellular stress. Importantly, contraction-mediated glucose transport is normal, despite impaired AMPKα1 activity in muscles from obese Zucker rats. Thus, AMPKα1 does not appear to play a major role in stimulation of glucose transport by muscle contraction. Our results are consistent with the hypothesis that activation of AMPKα2 in response to muscle contraction/exercise is associated with a parallel and normal increase in glucose transport in insulin-resistant skeletal muscle.

This study was supported by grants from the Swedish Medical Research Council, the Swedish Diabetes Association, the Foundation for Scientific Studies of Diabetology, the Swedish National Center for Research in Sports, Diabetes U.K., the Medical Research Council U.K., and the following research foundations: Karolinska Institutet, Novo-Nordisk, Thurings, Wiberg, Tore Nilson, and the AFA-Insurance Jubilee Foundation for Research in National Diseases.