Regulation of Glycogen Synthase by Glucose and Glycogen
A Possible Role for AMP-Activated Protein Kinase
- 1School of Biochemistry and Genetics, Medical School, University of Newcastle, Newcastle upon Tyne, U.K.
- 2Cellular Stress Group, MRC Clinical Sciences Centre, Imperial College School of Medicine, Hammersmith Hospital, London, U.K.
- 3Target Cell Biology, Novo Nordisk A/S, Bagsvaerd, Denmark
We report here use of human myoblasts in culture to study the relationships between cellular glycogen concentrations and the activities of glycogen synthase (GS) and AMP-activated protein kinase (AMPK). Incubation of cells for 2 h in the absence of glucose led to a 25% decrease in glycogen content and a significant decrease in the fractional activity of GS. This was accompanied by stimulation of both the α1 and α2 isoforms of AMPK, without significant alterations in the ratios of adenine nucleotides. When glucose was added to glycogen-depleted cells, a rapid and substantial increase in GS activity was accompanied by inactivation of AMPK back to basal values. Inclusion of the glycogen phosphorylase inhibitor, CP-91149, prevented the loss of glycogen during glucose deprivation but not the activation of AMPK. However, in the absence of prior glycogen breakdown, glucose treatment failed to activate GS above control values, indicating the crucial role of glycogen content. Activation of AMPK by either 5-aminoimidazole-4-carboxamide 1-β-d-ribofuranoside (AICAR) or hydrogen peroxide was also associated with a decrease in the activity ratio of GS. AICAR treatment had no effect on total cellular glycogen content but led to a modest increase in glucose uptake. These data support a role for AMPK in both stimulating glucose uptake and inhibiting GS in intact cells, thus promoting glucose flux through glycolysis.
Glycogen synthase (GS) catalyzes a crucial and rate-limiting step in muscle nonoxidative glucose disposal (1). The regulation of GS activity is complex. Enzyme activity is sensitive to allosteric regulation by a number of metabolites (2), is subject to reversible phosphorylation, which inactivates the enzyme, (3) and is regulated by feedback inhibition by glycogen (4–6) via an unknown mechanism. GS activity is modulated by reversible phosphorylation of primarily three specific serine residues, collectively termed site 3 (3). GS is maintained in a low-activity state under basal conditions principally through the continual phosphorylation of site 3 by GS kinase 3 (GSK3) (7). Insulin is believed to activate GS mainly through the inhibition of GSK3 (8,9); however, some level of regulation may control glycogen-targeted protein phosphatases (10). A number of other kinases have been identified that can phosphorylate GS in vitro (11), including AMP-activated protein kinase (AMPK), which can phosphorylate serine 7 (termed site 2) of GS (12). Phosphorylation of site 2, which can also be catalyzed by cAMP-dependent protein kinase (PKA), primes GS for further phosphorylation at site 2a by casein kinase I, which in turn leads to a decrease in GS activity (13). AMPK is a metabolite-sensing enzyme that has been implicated in the mediation of exercise-induced glucose uptake (14), although to date, little experimental evidence has attributed a role for AMPK in the regulation of GS activity in vivo.
GS becomes activated following glycogen depletion (such as occurs during exhaustive exercise) in an insulin-independent manner (15). The molecular mechanism underlying this phenomenon has not been elucidated, although a number of hypotheses have been proposed. One possibility is the involvement of an insulin-independent pathway leading to GSK3 inactivation (16). There is evidence that this is the case in rat muscle (14), although a more recent study in humans has suggested a GSK3-independent mechanism (17). Another possibility is that decreased cellular glycogen content may directly lead to GS activation. We have recently developed a model system in vitro using cultured human muscle, in which glycogen depletion is achieved by glucose deprivation (18). Following readdition of glucose, a dramatic and sustained increase in GS activity is observed, which is independent of GSK3 inactivation. Indeed, the mechanism leading to GS activation in this model is independent of that utilized by insulin (18).
The purpose of the present study was to further define the role of cellular glycogen content in determining GS activity and determine what role, if any, AMPK plays in controlling glycogen metabolism. Using a specific inhibitor of glycogenolysis, we have established a direct requirement for prior glycogen breakdown in the subsequent activation of GS by glucose. We have also provided evidence that AMPK is involved in the regulation of GS activity and the rate of glucose uptake in cultured human muscle cells.
RESEARCH DESIGN AND METHODS
All tissue culture trays were from Costar (Cambridge, MA). Culture media, penicillin/streptomycin, and trypsin-EDTA were from Gibco-BRL (Paisley, U.K.). Chick embryo extract and [γ-32P]ATP (148 TBq/mmol) was obtained from ICN (Costa Mesa, CA). Uridine diphospho-d-[6-3H]glucose (814 GBq/mmol) and 2-deoxy-d-[1-3H]glucose (362 GBq/mmol) were from Amersham Pharmacia Biotech (Buckinghamshire, U.K.). Glucose-6-phosphate dehydrogenase, hexokinase, and amyloglucosidase were from Boehringer Mannheim (Lewes, U.K.). 5-Aminoimidazole-4-carboxamide 1-β-d-ribofuranoside (AICAR) was from Sigma (Poole, U.K.). CP-91149 was provided by Pfizer (Pfizer Global Research & Development, Groton Laboratories, Groton, CT). Antibodies used for immunoprecipitation of α1 and α2 isoforms of AMPK were as described previously (19).
Human myoblasts were grown from needle biopsy samples taken from the vastus lateralis muscle of healthy subjects with no family history of type 2 diabetes and with normal glucose tolerance and normal insulin sensitivity, as assessed using the short insulin tolerance test (20). Myoblasts were maintained in growth medium consisting of HAMS F-10 nutrient mixture containing 20% FCS, 1% chick embryo extract, 100 units/ml penicillin, and 100 μg/ml streptomycin. All experiments were performed using cells between the 5th and 15th passage at greater than 80% confluence. Myoblasts and myotubes respond equally well to insulin and other agonists at the level of glucose metabolism, in many respects mirroring the situation in whole muscle. Differentiation of myoblasts to myotubes induces significant increases in GS expression and some lowering of the activity ratio of the enzyme (21). To avoid variations in basal GS activity, as a result of innate differences in the differentiation capacity of myoblasts from different subjects, myoblasts were used throughout this study.
Cellular glycogen content determination.
Total cellular glycogen content was assessed by modification of a previous method (22). Following treatment, cells were washed rapidly in ice-cold PBS and scraped into 100 μl of 0.2 mol/l sodium acetate, pH 4.8. Extracts were briefly sonicated, using a Soniprep 150, before addition of 250 mU amyloglucosidase per sample. Samples were incubated for 2 h at 40°C and vortexed regularly to avoid sedimentation. Sample was incubated with assay cocktail (0.1 mol/l Tris-HCl, pH 8.0, 0.3 mmol/l ATP, 6 mmol/l MgCl2, 5 mmol/l diothiothreitol [DTT], 60 μmol/l NADP+, 2.5 units/ml hexokinase, and 1 μg/ml G6P-dehydrogenase) for 30 min at room temperature. Changes in fluorescence, as a result of NADPH production, were determined using an excitation wavelength of 355 nm and an emission wavelength at 460 nm. Reaction blanks were determined as the fluorescence of samples before enzymatic treatment with amyloglucosidase.
Estimation of glucose uptake.
Glucose uptake was determined as the rate of 2-deoxy-d-[6-3H]glucose uptake, using modification of a previous method (23). Cells were maintained in the absence of serum for 5 h before the replacement of media with glucose-free Dulbecco’s Modified Eagle’s medium (DME Glu−) for 15 min at 37°C. For AICAR treatments, cells were incubated in serum free for 2 h before the addition of AICAR for the times indicated. The rate of 2-deoxyglucose uptake was determined during 5 min of incubation with 50 μmol/l 2-deoxy-d-[6-3H]glucose (specific activity 0.4 kBq/pmol). Reaction blanks were determined as the rate of 2-deoxy-d-[6-3H]glucose uptake in the presence of 0.1 mmol/l cytochalasin B.
Following incubation, cells were washed with ice-cold PBS several times and solubilized in 0.05% SDS for 30 min at room temperature. Protein content of samples was assayed using Coomassie Protein Assay Reagent, and uptake of 2-deoxy-d-[6-3H]glucose was determined by liquid scintillation counting.
Assay of GS.
Following the indicated treatments, cells were rapidly washed three times with ice-cold PBS and collected, by scraping, into GS extraction buffer (10 mmol/l Tris-HCl, pH 7.8, 150 mmol/l KF, 15 mmol/l EDTA, 60 mmol/l sucrose, 1 mmol/l 2-mercaptoethanol, 10 μg/ml leupeptin, 1 mmol/l benzamidine, and 1 mmol/l phenylmethylsulfonyl fluoride [PMSF]). Cells were then disrupted by briefly sonicating using a Soniprep 150. GS activity was determined in whole lysates as the incorporation of 3H-glucose from uridine-5′-diphosphate-[U-3H]glucose into glycogen, as described by Guinovart et al. (24). Samples were incubated with reaction cocktail (50 mmol/l Tris-HCl, pH 7.8, 20 mmol/l EDTA, 25 mmol/l KF, 1% glycogen, 0.4 mmol/l UDP-[3H]glucose [specific activity 3 kBq/nmol]), containing either 0.1 mmol/l (active) or 10 mmol/l glucose-6-phosphate (total), for 30 min at 30°C. Results were expressed as fractional activities (active/total). This assay has been optimized to detect the activity changes resulting from dephosphorylation of GS (24).
AMPK activity determinations.
Following the indicated treatments, cells were rapidly washed three times with ice-cold PBS and collected, by scraping, into buffer A (50 mmol/l Tris-HCl, pH 7.5, 1 mmol/l EDTA, 50 mmol/l NaF, 5 mmol/l NaPPI, 1 mmol/l benzamidine, 10% glycerol, 1% Triton X-100, 1 mmol/l DTT, and 0.1 mmol/l PMSF). Samples were briefly sonicated before centrifugation at 13,000g for 5 min (4°C). Immunoprecipitations of α1, α2, and total AMPK were performed on aliquots of supernatants containing 30 μg protein. In each case, immunoprecipitations were carried out over 2 h at 4°C, using appropriate antibodies and protein A or protein G immobilized on Sepharose. The immune complexes were recovered by brief centrifugation and washed twice with buffer A and twice with buffer B (50 mmol/l Hepes, pH 7.4, 1 mmol/l EDTA, 10% glycerol, and 1 mmol/l DTT).
AMPK activity in immunoprecipitates was assayed in a final volume of 25 μl containing 50 mmol/l Hepes, pH 7.4, 1 mmol/l EDTA, 10% glycerol, 1 mmol/l DTT, 0.2 mmol/l SAMS peptide substrate (25), 200 μmol/l [γ-32P]ATP (specific activity ∼1.1 kBq/nmol), 5 mmol/l MgCl2, and 0.2 mmol/l AMP. After incubation for 30 min at 30°C, samples were centrifuged briefly, and 20 μl of the supernatant containing the radiolabelled peptide product was spotted onto 1 cm2 Whatman P81 phosphocellulose paper squares. After washing in 1% phosphoric acid with four changes, the papers were dried and phosphate incorporation was determined by liquid scintillation counting. Enzyme activity (U) was defined as that which catalyzes the incorporation of 1 pmol of phosphate into peptide substrate in 1 min.
All results are expressed as means ± SE. Statistical analysis was made using a two-tailed unpaired Student’s t test, following one-way ANOVA.
Activation of GS by glucose following time-dependent decrease in cellular glycogen in response to glucose deprivation has been reported in human muscle cells in culture (18), suggesting that induced changes in cellular glycogen content might be responsible for alterations in the activity of GS. However, it was unclear whether changes in the activity state of GS were a direct result of alterations in the concentration of cellular glycogen or of other metabolic consequences of glucose deprivation. Therefore, a well-characterized inhibitor of liver glycogen phosphorylase, CP-91149 (26), was used to dissociate changes in intracellular glycogen levels from other experimental variables. The ability of CP-91149 to affect intracellular glycogen levels was assessed in human myoblasts in culture (Table 1). Incubation of myoblasts in the absence of glucose for 2 h caused an ∼25% decrease in intracellular glycogen concentrations, consistent with an earlier work (18). This decrease was essentially blocked by 10 and 100 μmol/l CP-91149, indicating that this compound can also inhibit the human muscle isoform of glycogen phosphorylase. At an inhibitor concentration of 10 μmol/l, no change in glycogen content was observed in cells maintained in normal glucose (6.1 mmol/l); however, 100 μmol/l CP-91149 caused a small but significant increase in glycogen accumulation in these cells. For further studies, 10 μmol/l CP-91149 was used to inhibit glycogenolysis in order to avoid alterations in basal glycogen levels. Glycogen levels were not depleted in the presence of 10 μmol/l CP-91149 following up to 7 h of glucose deprivation, whereas in the absence of inhibitor, glycogen levels fell by >50% in 5 h (Table 1).
The effect of CP-91149 on starvation-induced changes in 2-deoxyglucose uptake was then examined (Fig. 1). Glucose withdrawal from myoblasts for 5 h caused a 1.6-fold increase in the rate of 2-deoxyglucose uptake, as compared with cells maintained in glucose-containing media. This is consistent with an earlier work (18). The basal rate of 2-deoxyglucose uptake was unaffected by the presence of CP-91149 during glucose deprivation; however, a slight decrease was observed in the rate of uptake following glucose-deprivation (644.7 ± 24.1 pmol · min−1 · mg−1 in untreated cells vs. 574.1 ± 15.2 pmol · min−1 · mg−1 in CP-91149 treated cells; P < 0.05). In the presence of CP-91149, and therefore in the absence of glycogen breakdown (Table 1), a 1.5-fold increase in the rate of glucose uptake following glucose deprivation persisted, indicating that in this system the stimulation of glucose uptake during glucose deprivation is not dependent on cellular glycogen content.
A direct relationship between intracellular glycogen concentration and GS activity has been suggested (5,6,27). We have also previously reported a dramatic and sustained increase in the fractional activity of GS following glucose treatment of glucose-deprived, glycogen-depleted, human myoblasts (18). CP-91149 was used to dissociate changes in the activity of GS resulting from changes in cellular glycogen concentration from other variables (Fig. 2). Glucose-deprivation of cells for 2 h caused a significant decrease in the fractional activity of GS before re-admission of glucose (0.020 in untreated cells to 0.004 in glucose-starved cells) without altering the activity of GS in the presence of saturating G6P (10 mmol/l) concentrations (46.14 ± 5.3 nmol · min−1 · mg−1 in untreated cells to 54.14 ± 6.4 nmol · min−1 · mg−1 in glucose-starved cells). Glucose treatment (5.5 mmol/l) of previously glucose-starved cultures increased GS activity approximately fivefold over that observed in control cultures. The inclusion of 10 μmol/l CP-91149 had no effect on GS activity in either control or glucose-starved cells; however, the fivefold increase in GS activity over control cultures observed during glucose re-admission was completely inhibited by prior treatment with CP-91149, indicating that this effect is totally dependent on prior depletion of glycogen. It is noteworthy that the fractional activity of GS in glucose-deprived cells was restored to levels observed in control cells following glucose re-admission of cultures maintained in the presence of CP-91149 (0.007 before glucose re-admission vs. 0.03 ± 0.02 following glucose re-admission). Therefore, in the absence of glycogen breakdown, glucose deprivation induces a decrease in GS activity that can be reversed by re-admission of glucose. However, glycogen depletion is required for full activation of GS observed following the treatment of previously glucose-starved cells with glucose.
Although AMPK has been shown to phosphorylate site 2 of GS in vitro (12), to date no evidence has been offered to suggest that GS is a physiological substrate of AMPK. However, a reduction in the intracellular ATP-to-AMP ratio following glucose-deprivation has been demonstrated in a pancreatic cell line (28). In addition, in view of the fact that the intracellular ratio of ATP to AMP plays a major role in the regulation of AMPK activity (29), we wished to explore whether AMPK mediated the effects of glucose withdrawal on GS activity. The activity of α1 and α2 AMPK isoforms was therefore examined in human myoblasts following glucose-withdrawal and glucose re-admission (Fig. 3). Significant levels of both the α1 and α2 AMPK isoforms were detected in control cells. An approximate fourfold increase in the activity of both the α1 and α2 AMPK isoforms from 7.04 ± 1.4 and 1.8 ± 0.7 units/mg to 26.8 ± 6.1 and 7.5 ± 2.0 units/mg, respectively, was observed in myoblasts following 2 h of glucose withdrawal, during which time the fractional activity of GS fell significantly (Fig. 2). Somewhat surprisingly, no significant alteration in the ratio of ATP to ADP was apparent following glucose starvation (glucose-fed control ATP/ADP 3.6 ± 0.3 compared with glucose-starved ATP/ADP 3.6 ± 0.2; n = 6). AMPK activity returned to control values following treatment of cells with 5.5 mmol/l glucose for 10 min, conditions associated with dramatic activation of GS but again without significant change in the ATP/ADP ratio (4.7 ± 0.5). It is noteworthy that AMPK activity returned to control values after 10 min of glucose re-administration, a time at which glycogen levels are still significantly depleted (18). In the presence of CP-91149, a greater increase in AMPK activity was observed in glucose-starved cells, with a dramatic decrease in the ATP-to-ADP ratio (0.53). Again, glucose re-admission caused AMPK levels to return to control values, which was associated with a restoration of the fractional activity of GS to control value. However, the dramatic reactivation of GS was not observed, indicating that an additional glycogen-dependent mechanism is involved.
AICAR, the cell-permeable precursor of ZMP, has been shown to selectively activate AMPK in a number of model systems (30,31). To assess further the involvement of AMPK in the regulation of GS and glucose uptake, human myoblasts were treated with AICAR (2 mmol/l) (Fig. 4). Incubation of cells with AICAR caused a time-dependent increase in AMPK activity, with stimulation being observed within 30 min and reaching a maximum of approximately twofold after 90 min. AICAR treatment also caused a time-dependent decrease in the activity of GS (0.063 ± 0.01 in control cells vs. 0.029 following 2 h of AICAR treatment) and an increase in the rate of 2-deoxyglucose uptake (1.3-fold over basal levels following 2 h of AICAR treatment), again consistent with a role for AMPK in controlling both parameters. Total cellular glycogen content was unaffected following AICAR treatment, suggesting that observed decreases in GS activity were not a result of glycogen accumulation. In comparison to maximal treatments of AICAR (2 h at 2 mmol/l), glucose-deprivation of myoblasts (2 h) caused a greater activation of AMPK (19.2 ± 3 nmol · min−1 · mg−1 in AICAR-treated cells vs. 27.9 ± 4 nmol · min−1 · mg−1 in glucose-starved cells) and inhibition of GS (0.029 ± 0.0 in AICAR-treated cells vs. 0.017 ± 0.01 in glucose-starved cells). Cells in normal glucose conditions were also treated with hydrogen peroxide, another known activator of AMPK (32), to further examine the role of this enzyme in modulating GS activity (Fig. 5). H2O2 rapidly activated AMPK in a transient manner, and this was mirrored by a transient decrease in GS activity.
Glycogen levels are depleted during exhaustive exercise (15). Restoration of these levels is in part independent of insulin action (15) and involves increases in both glucose uptake (33,34) and GS activity (35). However, to date the molecular mechanisms underpinning these events have been poorly understood. A role for glycogen in influencing the metabolic steps involved in glycogen repletion has been described (4). We have previously reported the use of human myoblasts as an experimental system to study the relationship between glycogen content and subsequent glucose metabolism (18).
Glucose deprivation of human muscle cells in culture induces a decrease in both cellular glycogen content and GS fractional activity. Subsequent glucose treatment of cells causes a dramatic increase in the fractional activity of GS, reflecting a decrease in the phosphorylation state of the enzyme. A significant increase in the rate of glucose uptake was also observed in glucose-starved cells. It could be argued that the observed changes in glucose uptake and GS activity in this model are a result of the glucose withdrawal/re-admission protocol per se. To address this concern, a specific inhibitor of glycogen phosphorylase (CP-91149) was used to discriminate effects of glycogen depletion from glucose deprivation. CP-91149 treatment of human myoblasts completely inhibited glycogen breakdown during glucose deprivation and, at maximal concentrations, caused some slight glycogen accumulation. The increased rate of glucose uptake in glucose-starved cells was largely unaffected by inhibition of glycogenolysis, implying that cellular glycogen content was not controlling metabolite entry into the cell, following glucose-deprivation. In contrast, the large increase in GS activity observed following re-admission of glucose to cells was severely blunted by CP-91149, suggesting that glycogen depletion is necessary for the superactivation of GS in this model. It is noteworthy, however, that neither the decrease in GS fractional activity during glucose deprivation nor the recovery of fractional activity to basal values following glucose re-admission was affected by inhibition of glycogen breakdown. This suggests that two separate mechanisms are responsible for modulating GS activity in this system, one that is dependent on glycogen depletion and one that is not. The dramatic activation of GS by glucose is clearly dependent on the preexisting glycogen content of the cell. This provides direct evidence that intracellular glycogen concentration can affect GS activity in the presence of physiological concentrations of glucose. Moreover, the low fractional activity of GS in human myoblasts, as compared wth human muscle in vivo, may be explained by the elevated intracellular glycogen concentration reported here and observed by others in human myotubes (27).
A potential candidate for mediating the glycogen-independent effects on glucose uptake and GS fractional activity is AMPK. In rat epitrochlearis muscles, AMPK activity was strongly correlated with the rate of glucose uptake following challenge with a variety of fuel-depleting stimuli (14). In addition, AMPK has been implicated in mediating contraction-induced increases in glucose uptake (31,35). It is worth noting that in a recent study of transgenic mice where a dominant inhibitory AMPK mutant was overexpressed in muscle, exercise-stimulated glucose uptake and translocation of Glut 4 to the cell surface was only partly inhibited, implicating other AMPK-independent pathways in this process (36). Furthermore, reported effects of AICAR on GS activity in rats are confusing, apparently being dependent on muscle type and experimental design (37,38).
AMPK is acutely sensitive to changes in cellular energy balance and is therefore a candidate for mediating the inhibitory effects on GS. Indeed, inhibition of GS as a result of fuel depletion in glucose-starved cells would be a sensible energy-preserving response in order to meet changing energy demands. AMPK has already been implicated in inhibition of other energy-consuming processes, including fatty acid and cholesterol synthesis (29). In human myoblasts, both AMPK α1 and α2 isoforms were stimulated in response to glucose withdrawal in both the absence and presence of the glycogen phosphorylase inhibitor. AMPK activity rapidly returned to control values in both instances following glucose re-administration. The reverse correlation between GS fractional activity and AMPK activity are consistent with a causal relationship between the two.
The focus of the current article is the role of the AMPK in controlling glucose metabolism, not its mechanism of activation. However, several aspects of the nucleotide ratio measurements merit discussion. Firstly, the ATP-to-ADP ratio is lower than in most other culture cell models, although it is consistent with the value in neonatal cardiomyocytes (D.C., unpublished data). The absolute values of the nucleotides is also low, making measurements difficult, particularly that of AMP. As ATP, ADP, and AMP are maintained in equilibrium, where AMP concentrations are difficult to detect, the ratio of ATP to ADP can be used as an indicator of AMP levels (39). In the present study, the main observation is that the ATP-to-ADP ratio does not change during glucose deprivation—presumably, glycogen breakdown provides the necessary energy. Consistent with this is the dramatic drop in ratio when glycogen breakdown is inhibited by CP-9149. However, there are now several reports where the activation of AMPK is apparently independent of changes in the nucleotide ratios. These include activation of AMPK in hepatocytes (40) and skeletal muscle (41) in response to the glucose-lowering drug metformin, activation of AMPK in skeletal muscle by leptin (42), and the effects of a number of metabolites in perfused rat hearts (43). It is apparent from these studies that other mechanisms regulate AMPK in addition to the nucleotide ratio.
AICAR, a relatively selective activator of AMPK (30,31), was used to further substantiate the role of AMPK in the regulation of glucose uptake and GS activity. AICAR is an intermediate in de novo purine biosynthesis and, once metabolized (to ZMP), is a potent activator of AMPK (30). In cultured human myoblasts, AICAR was a less potent stimulator of AMPK activity than has been reported in some other cell systems (19). This may be due to a lower rate of AICAR metabolism and, thus, production of ZMP. Despite relatively low levels of AMPK activation, AICAR treatment of cultured human muscle cells stimulated glucose uptake and inhibited GS activity in a time-dependent manner; also, there was a strong correlation between GS and AMPK activity during both AICAR treatment and glucose starvation. Furthermore, H2O2 treatment of cells potently activated AMPK and inhibited GS activity. Therefore, several lines of evidence indicate that AMPK can regulate GS in vivo, although whether AMPK is directly phosphorylating GS remains to be established.
We thank Dr. Dennis J. Hoover and Dr. Judith L. Treadway for generously providing the glycogen phosphorylase inhibitor CP-91149 (Pfizer Inc., Groton Laboratories). This work was supported in part by Diabetes U.K., Xcellsyz Ltd., and Medical Research Council, U.K. R.H. held a CASE studentship from the Biotechnology and Biological Sciences Research Council, U.K., partly funded by Novo Nordisk A/S. We thank Dr. Mark Walker for his continued assistance in obtaining muscle biopsies.
Address correspondence and reprint requests to Stephen J. Yeaman, School of Biochemistry and Genetics, The Medical School, University of Newcastle, Newcastle upon Tyne NE2 4HH, U.K. E-mail:.
Received for publication 28 March 2001 and accepted in revised form 24 September 2002.
J.G.M. is currently affiliated with OSI Pharmaceuticals Ltd., Oxford, U.K.
R.H. receives consulting fees from Xcellsyz, Ltd., a start-up company engaged in deriving immortalized cell lines for study of diabetes. J.G.M. is employed by and holds stock in Novo Nordisk A/S. D.C. is on the Scientific Advisory Board for Xcellsyz. S.J.Y. holds stock in Xcellsyz and has received honoraria from Novo Nordisk and Glaxo Wellcome.
AICAR, 5-aminoimidazole-4-carboxamide 1-β-d-ribofuranoside; AMPK, AMP-activated protein kinase; DME Glu−, glucose-free Dulbecco’s Modified Eagle’s medium; DTT, diothiothreitol; GS, glycogen synthase; GSK3, GS kinase 3; PKA, cAMP-dependent protein kinase; PMSF, phenylmethylsulfonyl fluoride.