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School of Clinical Medical Sciences (E.A.M., M.W.) and School of Cell and Molecular Biosciences (R.H., S.J.Y.), University of Newcastle upon Tyne, Newcastle upon Tyne, United Kingdom NE2 4HH
Address all correspondence and requests for reprints to: Prof. Mark Walker, School of Clinical Medical Sciences, The Medical School, Framlington Place, University of Newcastle, Newcastle upon Tyne, United Kingdom NE2 4HH. E-mail: mark.walker{at}ncl.ac.uk.
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Abstract |
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Introduction |
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A system has been established for culturing human skeletal muscle cells, and genetic factors are believed to contribute either wholly or in part to abnormalities of metabolism retained in the cultured cells. Human muscle cell cultures are a particularly valuable tool because they display key morphological (2) and biochemical (3, 4) features of mature skeletal muscle. Previous studies have examined insulin signaling events in cultured muscle cells (passage 1 or 2) from type 2 diabetic patients (3, 4, 5, 6, 7, 8, 9). It remains possible that the abnormalities demonstrated in these primary cultured cells result from both a carryover effect from the in vivo environment of the cell, i.e. in vivo programming, as well as gene effects in the cells. If in vivo programming is important, then the effects of obesity, physical activity, and drug treatments of the diabetic and control subjects need to be considered. These environmental factors have not always been addressed in studies of glucose uptake to date (3, 5, 9) and may partly explain inconsistencies reported. For example, Gaster et al. (5) found that diabetic cells cultured at physiological insulin concentrations increased their glucose uptake rate to a similar level as control cultures upon acute insulin stimulation, but Henry et al. (3) found the diabetic cultures had a lower rate of glucose uptake compared with control cultures after acute insulin stimulation.
To remove the confounding metabolic influence of the diabetic state, but still focus on subjects that are likely to harbor inherited defects of insulin action, we previously studied cultured muscle cells from nondiabetic, insulin-resistant, first-degree relatives of type 2 diabetic families (10). Insulin-stimulated glucose uptake was decreased in the relative muscle cell cultures as a whole, but the responses were variable within the group, such that only half were profoundly impaired. The heterogeneity demonstrated in the relatives reflects the fact that despite their heightened risk, the lifetime risk of the individual developing diabetes is between 40% and 60% (11).
The design of the current study is novel because it maximizes the chance of finding altered insulin signaling events resulting from gene effects in cultured muscle cells from a homogeneous group of type 2 diabetic patients by three principal strategies: 1) studying the cells after an extended period in culture, between passages 4 and 6, maintained in medium with physiological glucose and insulin concentrations; 2) recruiting type 2 diabetic patients with a family history of type 2 diabetes; and 3) recruiting those with clinical evidence of marked insulin resistance in the absence of severe obesity.
The aims of this study were to establish whether cultured muscle cells from insulin-resistant, type 2 diabetic subjects after prolonged culture have functionally significant insulin signaling abnormalities. If so, we aimed to determine whether an alternative pathway, stimulating glucose uptake involving AMP-activated protein kinase (AMPK) (12), through which exercise stimulates glucose uptake in vivo, was also impaired.
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Subjects and Methods |
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Ham’s F-10 medium, MEM, fetal bovine serum (FBS), penicillin-streptomycin, and trypsin-EDTA were obtained from Invitrogen (Paisley, UK). Chick embryo extract was purchased from Sera Laboratories (Salisbury, UK). All cell culture plasticware was obtained from Greiner (Gloucestershire, UK). 2-[3H]Deoxy-D-glucose and D-[U-14C]glucose were purchased from NEN Life Science Products (Cambridge, UK). Human Actrapid U-100 insulin was supplied by Novo Nordisk (Crawley, West Sussex, UK). Cytochalasin-B, 2-deoxyglucose, oyster glycogen type II, protease inhibitor cocktail, sodium dodecyl sulfate, EDTA, and the creatine kinase kit assay (CK-20) were obtained from Sigma-Aldrich Corp. (Poole, UK). Mouse antihuman desmin was purchased from Dako (High Wycombe, UK). Mouse antihuman CD56 (nerve-cell adhesion molecule) antibody was obtained from BD Biosciences (Oxford, UK). The antiglycogen synthase (anti-GS) antibody (rabbit polyclonal) was a gift from Prof. L. Groop (Malmo, Sweden), and the antiphospho antibody for sites 3a+b (residues 634–650) on GS was a gift from Calum Sutherland (Ninewells Medical School, Dundee, UK). Antiphospho-serine 473 protein kinase B (PKB) and anti-PKB antibodies were purchased from New England Biolabs (Beverley, MA). Goat, sheep, and rabbit antimouse secondary antibodies were purchased from Sigma-Aldrich Corp. Separation magnet and stand, MS+ columns, and goat antimouse Mini-Mac beads were purchased from Miltenyi Biotec (Bisley, UK). The insulin ELISA was purchased from Mercodia (Uppsala, Sweden). Coomassie protein reagent was purchased from Pierce Chemical Co. (Chester, UK). Hyperfilm and reagents for enhanced chemiluminescence were purchased from Amersham Pharmacia Biotech (Little Chalfont, UK).
Patients
Six compliant type 2 diabetic patients with clinical evidence of marked insulin resistance, taking more than 100 U insulin/d, and with a body mass index (BMI) less than 32 kg/m2, were recruited. All patients had been treated with diet and oral hypoglycemic agents for at least 3 yr after diagnosis before starting insulin treatment; metformin was continued in five of six patients after commencing insulin treatment. The diabetic subjects had at least one first-degree relative with type 2 diabetes. Six nondiabetic control subjects with no family history of type 2 diabetes were recruited. The control and diabetic subjects were matched for age and BMI. The compliance of the diabetic subjects with their insulin treatment was checked by contacting their general practitioner to ensure the patients collected sufficient insulin prescriptions to take the prescribed dose.
All subjects gave written informed consent, and the study was approved by the Newcastle and North Tyneside joint ethics committee.
Human primary muscle cultures
Aneural muscle cultures were established using the methods of Blau and Webster (13) and Halse et al. (14) as previously described. Previous studies confirmed the expression of the key insulin-mediated glucose transporter molecule 4 (GLUT4) in our system (10). Diabetic muscle was obtained from the vastus lateralis muscle by needle biopsy under local anesthetic. Control muscle was obtained from the vastus lateralis muscle at the time of hip surgery under general anesthesia.
The diabetic and control myoblast cultures were purified using a commercially available system from Miltenyi Biotec. Briefly, harvested cells were suspended in the separation buffer (PBS containing 2 mM EDTA and 5% FBS) and incubated with a primary antibody, anti-CD56 (nerve-cell adhesion molecule), recognizing a muscle-specific cell surface antigen (15). After washing, the cell suspension was incubated with secondary antibody, goat antimouse antibody attached to a Mini-Mac bead. Then the cell suspension was passed over an MS+ column housed within a magnetic field. The cells with Mini-Mac beads attached were retained on the column, and other cells were passed through the column. Subsequently, having removed the column from the magnetic field, the cells retained in the column were eluted and returned to the culture.
Myoblast cell cultures were maintained in Ham’s F-10 nutrient mixture supplemented with 20% FBS, 1% chick embryo extract, 100 U/ml penicillin, and 100 µg/ml streptomycin in an atmosphere of 5% CO2 in air. Glucose and insulin concentrations in the myoblast culture medium were 6.1 mM and 1.7 pM, respectively. Cell monolayers were harvested by trypsinization when 70% confluent and subcultured using a split ratio of 1:4. Muscle cell origin was confirmed immunohistochemically in each culture using antibodies to the muscle-specific protein desmin (2, 16).
When the cell monolayers reached approximately 90% confluence, they were differentiated into myotubes by replacing the culture medium with MEM supplemented with 2% FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin. Glucose and insulin concentrations in the differentiation medium were 5.5 mM and 0.9 pM, respectively. The medium was changed twice during differentiation. Cell fusion and differentiation into multinucleated myotubes were monitored using phase contrast microscopy by examining the fold increase in expression of differentiation-dependent proteins, including myogenin and glycogen synthase, by Western blotting (as described below) and by measuring the fold increase in creatine kinase using a kit assay (CK-20). All experiments on myotubes were performed after 7 d in differentiation medium.
Measurement of 2-deoxy-D-[3H]glucose uptake
Cells were grown on 12-well culture plates. Before the assay, a modified version of the method described by Klip et al. (17), the medium was removed from the cell monolayers, and the monolayers were washed three times with medium (Ham’s F-10 with 0.2% BSA or MEM with 0.2% BSA, for myoblast and myotube cultures, respectively). The cell monolayers were then placed in 1 ml medium for 4 h and incubated in 5% CO2 at 37 C. The monolayers were washed twice with uptake buffer (136 mM NaCl, 4.7 mM KCl, 1.25 mM MgSO47H2O, 1.2 mM CaCl/2H2O, and 20 mM HEPES, pH 7.4) and then incubated in 1 ml uptake buffer with or without insulin or cytochalasin B for 20 min. Measurement of uptake was commenced with the addition of 2-deoxy-D-[2,6-3H]glucose. The final concentrations were 100 µM 2-deoxyglucose and 0.5 µCi/ml de-deoxy-D-[2,6-3H]glucose, respectively. After incubation for 10 min, glucose uptake was terminated by rapidly washing the culture plates five times with ice-cold PBS. The cells were then solubilized in 0.05% sodium dodecyl sulfate for 30 min. The wells were aspirated, and the cell-associated radioactivity in each well was determined in 200 µl of the aspirate by liquid scintillation counting. Fifty microliters of the cell aspirate were saved for determination of the protein content of the cell extracts. Noncarrier-mediated uptake was determined in parallel incubations for each individual cell line by exposing the cells to 10 µmol/liter cytochalasin B, a potent inhibitor of transport (18).
Glycogen synthesis
The rate of total glycogen synthesis was determined by measuring the incorporation of D-[U-14C]glucose into glycogen. Cells were grown in six-well plates. Before the assay, cells were washed three times in PBS and then placed in 1 ml of the appropriate medium for 2 h. This was then replaced with 1 ml experimental medium, with and without insulin, containing D-[14C]glucose (1.25 µCi/ml, final concentration) and incubated again for 1 h. At the end of this period, the reaction was stopped by washing the cell monolayer four times with ice-cold PBS, and then the cells were lysed by adding 200 µl 20% (wt/vol) potassium hydroxide. After 30 min the cell extracts were partially neutralized with 200 µl 1 M hydrochloric acid. The cell extracts were then transferred from the culture dishes to Eppendorf tubes, and the wells were washed with 200 µl dH2O. Glycogen was extracted from 400 µl combined extract/washings by ethanol precipitation after the addition of 12 mg/ml (final concentration) carrier glycogen. Radioactivity incorporated into glycogen was determined by scintillation counting.
Protein determination
The protein content of the cell extracts was determined by the method of Bradford (19) using the Pierce Coomassie protein reagent.
Western blotting
Myoblasts and myotubes were harvested from six-well plates in 50 µl extraction buffer [25 mM Tris-HCl (pH 7.5), 100 mM KCl, 1 mM EDTA, 25 mM potassium fluoride, 1 mM benzamidine, 0.5 mM Na3 vanadate, 0.1% (wt/vol) Triton X-100, 1 mM phenylmethylsulfonylfluoride, and 10 µl/ml protease inhibitor cocktail]. Samples were snap-frozen in liquid nitrogen and stored at –80 C until use. Upon thawing, the samples were briefly sonicated, and then the protein concentration of the samples was determined. Aliquots of 15 µg protein were boiled in loading solution [2% (wt/vol) sodium dodecyl sulfate, 10% (vol/vol) glycerol, 0.002% (wt/vol) bromophenol blue, and 6.25% (vol/vol) mercaptoethanol] for 5 min and then loaded onto 8% or 10% polyacrylamide gels. The gels were run at 10 mA through the stacking gel and then at 20 mA through the separating gel for approximately 60 min before being transferred onto nitrocellulose membranes. The membranes were then blocked in 5% milk-PBS at room temperature for 1 h with agitation. The membranes were incubated at 4 C overnight with the primary antibody. Then, after washing four times, the membranes were incubated with the appropriate secondary antibody at a dilution of 1:1000 for 1 h at room temperature. After further washes, the proteins were detected on the membranes using enhanced chemiluminescence reagents, and the photographic films were developed using a Kodak autodeveloper (Eastman Kodak, Rochester, NY). Films were scanned using a flatbed scanner onto a computer, and bands were quantified by densitometry.
Statistical analyses
Data are presented as the mean ± SEM. Comparisons between groups were determined by t test or Mann-Whitney U test for parametric and nonparametric data, respectively.
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Results |
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Table 1
summarizes the metabolic and anthropometric characteristics of the diabetic and control subjects recruited into the study. The groups were matched for age and BMI. The diabetic subjects had significantly higher glycated hemoglobin (P < 0.01), waist/hip ratio (P < 0.01), and triglyceride levels (P < 0.05) than the control subjects. In contrast to the clinical evidence of insulin resistance in the diabetic subjects, i.e. the mean insulin dose of 131.2 U insulin/d in this group, the control subjects had normal fasting insulin and glucose values. The lower total cholesterol values in the diabetic group (P < 0.01) are explained by the use of statin therapy in all participants to treat the dyslipidemia in this group. In Table 1, nonparametric data were log-transformed before statistics were performed.
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Immunohistochemical staining of cultures established from each study subject, using a desmin antibody, confirmed that more than 95% of cells were of muscle origin (data not shown). The differentiation of myoblasts to myotubes was monitored using phase contrast microscopy. After 7 d of differentiation, more than 90% of cells were multinucleated (data not shown). Further experiments to determine that the diabetic and control cultures differentiated into myotubes to a similar extent are summarized in Fig. 1. In these experiments there were six diabetic and six control subjects. Experiments were performed on at least two separate occasions, and data points for individual subjects are the mean of duplicate observations. There was no significant difference in the creatine kinase activity of undifferentiated control and diabetic cultures (5.5 ± 0.6 and 4.8 ± 0.8 U/min·mg protein, respectively; P = not significant). Figure 1A shows similar and significant fold increases in the muscle creatine kinase activity of control and diabetic cultures (4-fold higher after 7 d of differentiation; P < 0.001). The expression of GS in cell extracts was examined by Western blotting. Figure 1B demonstrates a representative Western blot for GS and densitometric analysis of Western blot bands. The diabetic cell extracts had lower expression of GS, and this difference was significant after 7 d of differentiation (P < 0.01). This finding is in agreement with previous reports (8), which found lower expression of GS in diabetic myoblast and myotube cultures. Figure 1C shows that there was no difference in the fold increase in GS expression with differentiation between control and diabetic cultures; both showed a significant increase over their respective myoblast values (both P < 0.001).
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Mean basal glucose uptake rates were comparable between the control and diabetic d 7 myotube cultures (812 ± 97 and 747 ± 59 pmol/mg protein·min, respectively). After exposure to 100 nM (P < 0.05) insulin, absolute rates of glucose uptake (Fig. 2A) were decreased in the diabetic compared with the control cultures. Figure 2B shows the mean and individual values presented as the fold increase in glucose uptake after acute insulin exposure. The mean fold increases were 1.49 ± 0.07 and 1.10 ± 0.06 (P < 0.01) at 100 nM insulin in control and diabetic d 7 myotube cultures, respectively.
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). The mean fold increases in glucose uptake in response to 2 mM AICAR stimulation were 1.33 ± 0.05- and 1.30 ± 0.05-fold (P = not significant) in control and diabetic cultures, respectively. The insulin-stimulated glucose uptake was repeated at the same time as the AICAR experiments as an internal control. The diabetic cultures had a significantly lower insulin-stimulated fold increase in glucose uptake (1.17 ± 0.04; control cultures, 1.69 ± 0.04; P < 0.001). For the experiments shown in Figs. 2
and 3, there were six diabetic and six control subjects. Experiments were performed on at least two separate occasions, and data points for individual subjects are the mean of triplicate observations.
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The mean basal glycogen synthesis rates in d 7 myotube cultures were comparable in the control and diabetic cultures (109 ± 18 and 95 ± 22 pmol/mg protein·min, respectively). In this muscle culture system, stimulation of glycogen synthesis was detectable in response to lower concentrations of insulin than was glucose uptake, so the glycogen synthesis rates were measured after acute exposure to 1 and 100 nM insulin. Absolute rates of glycogen synthesis after acute insulin exposure are shown in Fig. 4A. Glycogen synthesis was significantly decreased in the diabetic compared with the control cultures at 1 nM insulin (P < 0.05). The same pattern was seen at 100 nM, but the difference was not statistically significant. Figure 4B shows the same data (mean and individual values) expressed as the fold increase in glycogen synthesis above the basal level. The mean fold increase was 1.57 ± 0.1 vs. 1.06 ± 0.01 (P < 0.01) in response to 1 nM insulin in control and diabetic d 7 myotube cultures, respectively. In contrast, there was no significant difference in the fold increase between control and diabetic cultures in response to 100 nM insulin. This indicates that the diabetic cultures have altered sensitivity to insulin, but the maximum stimulated response is normal. For the experiments shown in Fig. 4, there were six diabetic and six control subjects. Experiments were performed on at least two separate occasions, and data points for individual subjects are the mean of triplicate observations.
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To further investigate possible abnormalities demonstrated in the acute signaling effects of insulin in the diabetic cultures, we studied PKB expression and phosphorylation. Firstly we compared the expression levels of PKB by Western blotting using an antibody that recognized all isoforms of PKB. We found no difference between diabetic and control cultures, independent of day of differentiation. A representative Western blot is shown in Fig. 5A. We then studied the PKB phosphorylation in response to acute insulin stimulation with 20 nM insulin for 10 min as a measure of insulin-stimulated PKB activity. PKB is phosphorylated at two sites (Thr308 and Ser473) for full activation in response to insulin. The phospho-PKB antibody used in this study detected the Ser473phosphorylation site of PKB, detecting the 
and ß isoforms of PKB (PKB
lacks Ser473). Figure 5B shows a representative Western blot demonstrating detection of PKB phosphorylation after stimulation with 20 nM insulin and subsequent densitometric analysis of the Western blots. There was no significant difference between diabetic and control cultures in levels of insulin-stimulated PKB phosphorylation, measured as the fold increase. The signal from the antiphospho-PKB blot was also standardized for blots against total PKB. In the diabetic and control d 7 myotubes, the phosphorylated PKB densitometry means (arbitrary units), expressed as a percentage of the signal for total PKB protein expression, were 65.1 ± 1.5% and 63.2 ± 2.5% respectively, confirming that the degree of phosphorylation was similar in control and diabetic cells. Because of the different reactivities of the primary antibodies, this does not, of course, give a value for the actual percentage of PKB molecules that are phosphorylated.
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, showing a representative Western blot (A) and the subsequent densitometric analysis (B), demonstrates dephosphorylation of sites 3 in response to 20 nM insulin stimulation for 10 min in the diabetic and control cells. There was no significant difference in the degree of dephosphorylation observed between the two groups.
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Discussion |
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An important feature of our study is the recruitment of a homogeneous group of insulin-treated type 2 diabetic patients with clinical evidence of insulin resistance based upon a very high daily insulin requirement. To ensure that the high insulin requirements were not simply due to associated obesity, we excluded patients with a BMI greater than 32 kg/m2. Furthermore, all recruited patients had a family history of type 2 diabetes to optimize the chance of detecting inherited abnormalities.
This approach appears to have proven successful, as a key observation was that all cell cultures from the diabetic patients retained a marked and comparable impairment of insulin-stimulated glucose uptake. In studies by other groups, cultured muscle cells from type 2 diabetic patients have been shown to have either decreased (3, 9) or normal (5) insulin-stimulated glucose uptake. This variability may simply reflect the heterogeneity within type 2 diabetic subjects because the other groups appear not to have rigorously selected patients with both a family history of type 2 diabetes and clinical evidence of insulin resistance. They also studied myotubes at an earlier passage (passage 1, i.e. after 4–6 wk in culture) when the possibility of retained environmental influences is a greater concern. It is worth noting that although our patients and control subjects were carefully matched for BMI and age, the ratio of male to female subjects differed within the two groups. However, unpublished data from our group suggest that the gender of the muscle donor does not influence the effect of insulin in the cultured muscle cells.
Exercise is an important stimulus for glucose uptake by skeletal muscle and acts alone or in concert with insulin’s stimulatory effects (12, 24, 25). The exercise signaling pathway is independent of the proximal signaling components of the insulin signaling pathway, namely phosphatidylinositol 3-kinase (21, 26), and is known to involve the protein kinase, AMPK. AMPK is activated by depletion of ATP levels (27) and by an increase in creatine to phosphocreatine levels (28). The antidiabetic drug metformin has been shown to activate AMPK (29), principally in the liver, but also in skeletal muscle (30). Experiments in vivo demonstrate that diabetic muscle exposed chronically to significantly higher insulin concentrations than control muscle had similar levels of AMPK protein expression and AMPK activation in response to exercise compared with the control muscle (31). AICAR, which is taken up by cells and converted to the monophosphorylated form, 5-aminoimidazole-4-carboxamide ribonucleotide, an analog of AMP, has been shown to activate AMPK both by direct allosteric activation and by activating an upstream kinase (32). Our group has shown that AICAR increases AMPK activity in control cultured muscle cells, resulting in an increase in the rate of glucose uptake by the cells (33). As in the case of insulin, the increased glucose uptake induced by AICAR is associated with increased translocation of GLUT4 to the plasma membrane (34). A novel finding of this study is that after prolonged culture, thus minimizing any effect of metformin therapy from in vivo, myotubes from the type 2 diabetic patients had an increase in the rate of glucose uptake in response to AICAR similar to that in control cultures. It has been recently reported that both insulin- and AICAR-stimulated glucose uptake rates were decreased in skeletal muscle strips from diabetic patients (35). However, as mentioned above, a difficulty with this tissue model is that there will be a general down-regulation of GLUT4-mediated glucose transport due to the effects of chronic hyperglycemia mediated through the hexosamine pathway (36). We have been able to remove this confounding effect by studying cultured myotubes and have shown that AICAR-stimulated, but not insulin-stimulated, glucose uptake is normal in the cultured diabetic muscle cells. Indeed, this is supported by the observation by Koistinen et al. (35) that AICAR-activated phosphorylation of AMPK was, in fact, normal in diabetic muscle strips. From a clinical perspective, these findings highlight the potential importance of AMPK as a therapeutic target for glucose-lowering in type 2 diabetes.
We have shown that insulin-stimulated glycogen synthesis was significantly decreased in the diabetic myotubes after acute exposure to 1 nM insulin. Other groups have reported decreased insulin-stimulated glycogen synthesis (7), glycogen synthase activity (expressed as fractional velocity) (4, 5, 8), and glycogen synthase protein expression (8) in cultured diabetic muscle cells. It is likely, therefore, that the decreased glycogen synthase protein expression observed in the cultured diabetic muscle cells in the present study contributes to the decreased glycogen synthesis.
In view of our findings of both decreased insulin-stimulated glucose uptake and glycogen synthesis, PKB was considered an attractive intermediary of the insulin signaling pathway for further study. Overexpression of constitutively active PKB increases glucose uptake in muscle cells (37, 38), and PKB inhibits glycogen synthase kinase-3 (GSK-3), leading to increased glycogen synthesis (39). Previous studies have examined PKB activity in whole muscle preparations from subjects with diabetes showing either reduced (40) or normal PKB activity (41). Furthermore, insulin-stimulated PKB phosphorylation (a measure of PKB activity) in whole muscle (42) and cultured muscle cells (4) from subjects with diabetes has been shown to be normal. Hence, there is a consensus from published studies to date that PKB activity is normal in diabetic muscle; however, it remains possible that the use of antibodies recognizing multiple PKB isoforms in some studies may result in subtle differences between groups being overlooked. Recently, Brozinick et al. (43) reported that insulin activated all three PKB isoforms in lean muscle, but only PKB in muscle from obese, insulin-resistant, nondiabetic subjects. Our study has shown that the expression of PKB and phosphorylation of the and ß isoforms (Ser473 is absent in the isoform) in response to acute insulin exposure was normal in our diabetic myotubes, although we cannot exclude minor changes in phosphorylation of the specific isoforms that do not affect total PKB phosphorylation.
In the current study we also demonstrated similar dephosphorylation of sites 3a+b on GS in response to insulin in both groups of cultures, presumably resulting from phosphorylation and inactivation of GSK-3 by PKB (39). This indicates that events downstream of PKB are functionally intact in the diabetic myotube cultures. Our findings support those of a recently published in vivo study examining insulin’s activation of GS in diabetic muscle, which demonstrated normal activation of PKB and inactivation of GSK-3, with then normal dephosphorylation of sites 3a+b on GS (42). Therefore, in summary, current data suggest that decreased activity of GS in diabetic muscle is not a direct effect of defective signaling through the classical insulin signaling cascade ending with PKB and GSK-3.
In conclusion, we have observed defects of insulin-stimulated glucose uptake and glycogen synthesis in cultured myotubes from type 2 diabetic patients with clinical evidence of insulin resistance. The persistence of these defects in the muscle cells after several passages in culture suggests that the retained abnormalities have a genetic basis. In contrast, AMPK-stimulated glucose uptake was normal in the diabetic myotube cultures in this cell model, and this supports observations from other studies that glucose uptake and AMPK activation are normal in response to exercise in type 2 diabetes. These observations have important clinical implications for developing strategies to improve glucose uptake and lower circulating blood glucose levels in type 2 diabetes.
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Acknowledgments |
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Footnotes |
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Abbreviations: AICAR, 5-Amino-4-imidazolecarboxamide; AMPK, AMP-activated protein kinase; BMI, body mass index; FBS, fetal bovine serum; GLUT4, glucose transporter molecule 4; GS, glycogen synthase; GSK-3, glycogen synthase kinase 3; PKB, protein kinase B.
Received November 5, 2003.
Accepted March 18, 2004.
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by membrane targeting promotes glucose and system A amino acid transport, protein synthesis, and inactivation of glycogen synthase kinase 3 in L6 muscle cells. Diabetes 47:1006–1013[Abstract]
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