1 Copenhagen Muscle Research Centre, Human Physiology, University of Copenhagen, 2100 Copenhagen; 2 Diabetes Biology, Novo Nordisk, 2880 Bagsværd; and 3 Medical Research Laboratory, Aarhus Kommunehospital, 8000 Aarhus C, Denmark
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ABSTRACT |
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We investigated the possible regulatory role of glycogen in insulin-stimulated glucose transport and insulin signaling in skeletal muscle. Rats were preconditioned to obtain low (LG), normal, or high (HG) muscle glycogen content, and perfused isolated hindlimbs were exposed to 0, 100, or 10,000 µU/ml insulin. In the fast-twitch white gastrocnemius, insulin-stimulated glucose transport was significantly higher in LG compared with HG. This difference was less pronounced in the mixed-fiber red gastrocnemius and was absent in the slow-twitch soleus. In the white gastrocnemius, insulin activation of insulin receptor tyrosine kinase and phosphoinositide 3-kinase was unaffected by glycogen levels, whereas protein kinase B activity was significantly higher in LG compared with HG. In additional incubation experiments on fast-twitch epitrochlearis muscles, insulin-stimulated cell surface GLUT-4 content was significantly higher in LG compared with HG. The data indicate that, in fast-twitch muscle, the effect of insulin on glucose transport and cell surface GLUT-4 content is modulated by glycogen content, which does not involve initial but possibly more downstream signaling events.
GLUT-4; skeletal muscle; fiber type; phosphoinositide 3-kinase
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INTRODUCTION |
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GLUCOSE UPTAKE in skeletal muscle is the major site of whole body insulin-mediated glucose disposal (3). Recent evidence points to glucose transport, rather than glucose phosphorylation or glycogen synthase activity, as the rate-controlling step in insulin-induced muscle glucose utilization and glycogen synthesis in healthy and diabetic humans (8). The mechanism for insulin stimulation of muscle glucose transport involves a mobilization process through exocytosis of specific glucose transporters (GLUT-4) from intracellular storage sites to the cell surface, resulting in a net increase in active GLUT-4 transporters and glucose transport rate across the membrane (9, 32). In muscles of patients with peripheral insulin resistance, GLUT-4 translocation in response to insulin is impaired (50), probably due to a defect in the insulin signaling pathway involved in glucose transport stimulation (4, 14). It is, therefore, not surprising that the interest in and the search for the molecules involved in insulin signaling leading to GLUT-4 translocation in muscle have rapidly grown in recent years. Although the complete link between the initial binding of insulin with its receptor and the final mobilization of GLUT-4-containing vesicles is presently unknown, a clear picture is now emerging of the initial events in insulin signaling (reviewed in Refs. 18 and 38). In short, the binding of insulin with the extracellular part of the receptor induces insulin receptor tyrosine kinase (IRTK) activation through autophosphorylation at the intracellular part of the receptor. Subsequent activation of insulin receptor substrate (IRS) and its binding with phosphoinositide 3-kinase (PI3K) has been shown to be essential for the stimulation of glucose transport by insulin in muscle (18, 47). The main catalytic lipid product of PI3K, phosphatidylinositol 3,4,5-trisphosphate, is believed to act as a second messenger and to activate a downstream serine-threonine protein kinase pathway involving phosphoinositide-dependent kinase-1 (PDK-1), protein kinase B (PKB; also called Akt), and protein kinase C (PKC; see Ref. 1). However, the participation of the latter kinases in the stimulation of glucose transport in insulin-stimulated muscle is still debated (21, 27, 44).
The aim of the present paper is to address the potential role of muscle glycogen in the regulation of insulin-stimulated glucose transport and insulin signaling in muscle. This idea originated from the frequently observed negative correlations between muscle glycogen content and insulin sensitivity. Jensen et al. (19) have elegantly shown that, in muscles where the glycogen content was altered by applying various fasting-refeeding protocols, the maximal insulin-stimulated glucose transport rate in incubated epitrochlearis muscles was inversely related to the glycogen content of the muscle. A similar observation was recently published by Kawanaka et al. (20) using an exercise-refeeding protocol. The authors suggest that the impaired insulin-induced glucose uptake during glycogen supercompensation can be attributed to an attenuation of the amount of GLUT-4 transporters translocated to the surface membrane. In the search for the mechanism of increased insulin sensitivity after exercise, as originally observed by Richter and colleagues (34, 35), glycogen has also been proposed as a possible regulator of muscle insulin action. As illustrated with the one-leg exercise model in humans, the increased insulin sensitivity after exercise is restricted to the exercised leg, pointing to a local factor (36). Therefore, an evident candidate for this local factor could be glycogen depletion. Nevertheless, an isolated effect of glycogen content on the insulin signaling cascade in skeletal muscle has not been elucidated.
In the present study, we therefore investigated the insulin activation
of glucose transport, cell surface GLUT-4 content, and activities of
the insulin signaling intermediates IRTK, IRS-1-associated PI3K,
and PKB- in muscles with varying glycogen levels induced by
carbohydrate deprivation or supplementation after a swimming exercise
bout. The use of the isolated perfused rat hindquarter model allows us
to study these effects simultaneously in four calf muscles with
different muscle fiber type composition.
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METHODS |
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Experimental animals. All experiments were approved by the Danish Animal Experiments Inspectorate and complied with the Principles of Laboratory Animal Care (National Institutes of Health publication no. 85-23, revised 1985). Male Wistar rats (mean weight ± SE was 87 ± 2 g, n = 66) were preconditioned to obtain three different subgroups with varying muscle glycogen concentrations, as described previously (17). Rats were subjected to 2 h of swimming in water maintained at 32-35°C with weights (5% of body weight) attached to their tails. In the 24 h preceding the swim, their food intake was restricted to 4 g (~60% of normal intake). After swimming, all rats had free access to tap water, and they were fed ad libitum with either lard (low glycogen, LG), with 4 g of normal rat chow and lard ad libitum (normal glycogen, NG), or with normal rat chow and a 20% glucose drinking solution (high glycogen, HG) until 3-6 h before perfusions or incubations. Rat muscles were perfused or incubated between 18 and 24 h after the swimming bout.
Hindlimb perfusions.
The rats were anesthetized with an intraperitoneal injection of
pentobarbital sodium (5 mg/100 g body wt). Surgery was performed as
described by Ruderman et al. (37) for isolated hindquarter perfusion. Rats were killed by intracardiac pentobarbital sodium injection. All perfusions lasted 25 min using a constant flow of 6 ml/min, perfusing both legs. The perfusion medium (100 ml) was
constantly gassed with a mixture of 95% oxygen-5% carbon dioxide. At
the onset of the perfusion, the first 10 ml of the perfusion medium
that passed through the hindquarter were discarded, and thereafter the
remaining 90 ml were recirculated. During the initial 20 min of
perfusions, a cell-free and glucose-free perfusate was used, consisting
of Krebs-Ringer bicarbonate buffer solution (KRBB), 4% BSA (fraction
V; Sigma Chemical), 0.15 mM pyruvate, and 4.2 IU/ml heparin, as
described previously (49). To measure the muscle glucose
transport during the last 5 min of perfusion, the hindquarters were
exposed to a new medium, similar to the previous medium but with the
following additions: 8 mM 2-deoxy-D-glucose and 1 mM
mannitol together with radioactive labeled tracers
2-deoxy-D-[2,6-3H]glucose (specific activity
51 Ci/mmol; Amersham International) and
D-[1-14C]mannitol (specific activity 57 mCi/mmol; Amersham International), yielding an activity of 0.075 and
0.05 µCi/ml, respectively, as previously described (49).
Throughout the whole perfusion period, the perfusion media contained
either 0, 100, or 10,000 µU/ml insulin (Actrapid; Novo Nordisk), so
there were nine experimental groups: basal condition and submaximal and
maximal insulin stimulation in rats with HG, NG, and LG. The
perfusion pressure ranged between 40 and 60 mmHg. At the end of the
experiment, circulation was stopped, and, from both legs, muscle
samples were taken from four different parts of the calf musculature,
representing a whole range of fiber type distributions. The respective
proportions of slow-twitch oxidative, fast-twitch oxidative glycolytic,
and fast-twitch glycolytic fibers of young rats (60-100 g) are
taken from Maltin et al. (31) and are given in
parentheses. The white, most superficial part of the gastrocnemius
(0:20:80), the plantaris (10:50:40), the red, deep proximal and medial
portion of gastrocnemius (10:55:35), and the soleus (55:40:5) were
trimmed of connective tissue, blotted, and freeze-clamped with aluminum
clamps cooled in liquid nitrogen. The biopsies were stored at 80°C
until analyzed.
Glycogen and glucose transport measurements.
Muscle glycogen content was measured as glucose residues by a
hexokinase method after acid hydrolysis (28). The
2-deoxyglucose uptake
(µmol · g1 · h
1) by the
different muscles was determined in perchloric acid extracts as
described previously (49).
Photoaffinity labeling of cell surface GLUT-4.
On a subgroup of animals of the HG and LG groups, epitrochlearis
muscles were stimulated with insulin during in vitro incubations and
subsequently exposed to Bio-LC-ATB-BMPA, synthesized as described previously (24), for surface labeling of GLUT-4. Muscle
incubation was chosen for this purpose because the high production
costs of this compound do not allow surface labeling experiments in the
perfused hindlimb. The epitrochlearis muscle was chosen because it is a
fast-twitch muscle containing ~75% fast-twitch glycolytic fibers
(43) and because it is more suitable for incubation than the fast-twitch muscles of the hindlimb. Rats were anesthetized as
described above, and the whole body was perfused (flow ~20 ml/min)
for 1 min through the left ventricle with KRBB containing 8 mM glucose,
1 mM pyruvate, and 0.2% BSA. The epitrochlearis muscles were carefully
dissected out and incubated in KRBB containing 8 mM glucose, 1 mM
pyruvate, and 0.2% albumin for at least 30 min at 29°C with constant
shaking. The incubation flasks were constantly gassed with a mixture of
95% oxygen-5% carbon dioxide. Subsequently, the muscles were
transferred to a new oxygenated KRBB solution containing 2 mM pyruvate,
0.2% albumin, and 0, 100, or 10,000 µU/ml insulin. After 30 min of
insulin stimulation, the muscles were transferred to a dark room and
incubated in 1 ml of KRBB buffer containing 400 µM Bio-LC-ATB-BMPA in
the presence of 0, 100, or 10,000 µU/ml insulin. After 8 min of
incubation at 18°C, muscles were irradiated for 6 min in a Rayonet
photochemical reactor (Southern New England Ultraviolet, Branford, CT)
using 300-nm lamps. After irradiation, the two muscles from the same rat were pooled, immediately blotted, trimmed, frozen in liquid nitrogen, and stored at 80°C. Solubilized crude membranes were prepared as described previously, with slight modification
(30). Briefly, muscles were homogenized in ice-cold 20 mM
HEPES, 5 mM NaEDTA, and 255 mM sucrose (HES buffer; pH 7.2) and later
centrifuged at 320,000 g for 60 min. The pellet was
resuspended and solubilized in the HES-buffer containing 2% (wt/vol)
Thesit. The homogenate was rotated for 1 h at 4°C and then
centrifuged at 120,000 g for 30 min. Solubilized crude
membranes (150 µg) in 500 µl HES buffer containing 1% (wt/vol)
Thesit were then mixed with 100 µl of a 50% slurry of immunopure
immobilized streptavidin on 6% beaded agarose (Pierce Chemical,
Rockford, IL). The samples were incubated overnight at 4°C, and the
precipitates were washed four times in 1% Thesit-PBS buffer, four
times in 0.1% Thesit-PBS buffer, and finally in PBS. The biotinylated
protein was separated by SDS-PAGE, transferred to nitrocellulose
membranes, and blocked with 5% nonfat milk incubated with a polyclonal
anti-COOH-terminal peptide GLUT-4 antibody (29) diluted
1:4,000 (vol/vol) in Tris-buffered saline and 0.1% (vol/vol) Tween 20. Labeled proteins were visualized by the enhanced chemiluminescence
method (Pierce Chemical) and were compared with a standard to correct
for intergel variations. Results were expressed relative to the value
of nonstimulated HG muscle (set at 1).
IRTK activity. IRTK activity was measured in duplicate by a modification (16) of the method described by Klein et al. (22). In short, microtiter wells were coated with anti-insulin receptor antibody, as described previously (22), and 30 µl of muscle homogenate supernatant were added to each well. IRTK activity was estimated as incorporation of phosphate in poly-Glu-Tyr (4:1) after addition of the substrate in the presence of [32P]ATP and scanning of filters using a PhosphorImager (Molecular Dynamics, Sunnyvale, CA; see Ref. 16). The binding capacity in each well was measured using 125I-labeled insulin.
IRS-1-associated PI3K activity.
PI3K was immunoprecipitated as previously described, with minor
modifications (46). Muscle samples (white gastrocnemius) were homogenized (OMNI 2000; Omni International, Warrenton, VA) in
ice-cold solubilization buffer consisting of 50 mM HEPES (pH 7.4), 137 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 10 mM NaF, 10 mM Na2P2O7, 2 mM EDTA, 1% Igepal,
10% glycerol, 2 mM Na3VO4, 1.5 mM
phenylmethylsulfonyl fluoride, and a protease inhibitor cocktail tablet
(Complete; Boehringer, Mannheim, Germany). Insoluble material was
removed by centrifugation (4°C; 16,900 g, TI 70.1; Beckman
Instruments, Fullerton, CA) for 30 min. IRS-1 was immunoprecipitated
with anti-IRS-1 antibody (3 µg/mg protein; UBI, Waltham, MA) for
2 h at 4°C, followed by protein A-Sepharose 6MB (4°C,
overnight; Pharmacia). The immunoprecipitates were successively washed,
as described previously (46), and PI3K activities were
measured immediately on immunoprecipitates by in vitro phosphorylation
of phosphatidylinositol. Sonicated phosphatidylinositol [20 µg
phosphatidylinositol/sample in 10 µl of 5 mM HEPES (pH 7.4)] was
added to each sample. The PI3K reaction (30°C, 10 min) was started by
addition of 10 µl of 50 mM MgCl2, 250 µM ATP, and 0.5 µCi/µl [-32P]ATP (specific activity 5,000 Ci/mmol;
Amersham International) in a buffer consisting of 20 mM HEPES (pH 7.4),
0.4 mM EGTA, and 0.4 mM Na2PO4 and was stopped
by addition of HCl. Lipids were harvested by chloroform-methanol (1:1)
extraction and were applied to a silica gel TLC plate. The plates were
developed (45-60 min) in chloroform-methanol-ammonium
hydroxide-water (45:35:3:7 by volume). Spots were quantified on a
phosphorscreen using a PhosphorImager (Molecular Dynamics) and were
compared relative to a standard sample.
PKB activity.
Muscle samples were homogenized in 10 vol of ice-cold buffer (pH 8)
containing 4 mM EDTA, 50 mM NaF, 1 mM Na3VO4, 1 µM microcystin, 0.1% 2-mercaptoethanol, and a protease inhibitor
cocktail tablet, and insoluble material was removed after a
13,000-g centrifugation for 10 min (4°C). Equal amounts of
protein (250 µg) were immunoprecipitated with 4 µg anti-PKB-
antibody (no. 06-608; UBI), and PKB activity was measured with the
PKB immunoprecipitation kinase assay kit from UBI according to the
protocol from the manufacturer except that the kinase reaction was
allowed to proceed for 30 min at room temperature. The radioactivity of
filter papers was quantified using a
-scintillation counter (TRICARP
1500; Packard, Meriden, CT). PKB activity is expressed as milliunits
per milligram protein, where 1 unit = 1 nmol phosphate
incorporated per 30 min.
PKB phosphorylation.
Supernatants from homogenates from the PKB activity assay were used for
evaluation of PKB phosphorylation. Equal amounts of protein (7.5 µg)
were separated on 10% bis-Tris NuPage gels (NOVEX, San Diego, CA) with
the PHOSPHOPLUS Akt (Ser473) antibody kit according to the
manufacturer's protocol. Bands were quantified by the use of the
FujiFilm CCD camera and the Image Gauge software (Fuji Photo Film,
Elmsford, NY). Results are expressed as percent of internal standard
(PKB-; UBI).
Statistics. Statistical evaluation of the data was done by unpaired t-tests to compare HG with LG (when no intermediate group was included) or one-way ANOVA to compare HG, NG, and LG, using the Student-Newman-Keul's method for post hoc multiple comparisons where appropriate. Linear correlations were calculated with the Pearson product moment test. Data are presented as means ± SE, and the level of significance was chosen at 0.05.
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RESULTS |
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Muscle glycogen content.
Feeding the rats different diets after the swimming exercise bout
resulted in markedly different muscle glycogen levels at the time the
experiments were performed, i.e., 18-24 h after swimming (Fig.
1). In the NG group that received a mixed
carbohydrate/fat diet, glycogen levels had returned to preexercise
levels (45-50 µmol/g wet wt in plantaris and white and red
gastrocnemius and 17 µmol/g in the soleus). The rats of the HG group
that received a carbohydrate-rich diet were glycogen supercompensated,
with calf muscle glycogen levels 2.0- to 3.6-fold higher
(P < 0.05 for all muscles investigated) compared with
the NG group (Fig. 1). The rats of the LG group were deprived of
carbohydrates and were still glycogen depleted at the time of
experiments, with values being 1.8- to 2.6-fold lower
(P < 0.05 for all muscles investigated) compared with
the NG group (Fig. 1). In the surface-labeling experiments,
epitrochlearis muscle glycogen content was not measured due to lack of
tissue. In previous experiments with an identical protocol, we had
observed a threefold higher epitrochlearis glycogen content in HG
compared with LG rats.
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Glucose transport.
Basal muscle glucose transport rate, estimated from the uptake of
isotopic 2-deoxyglucose, was significantly higher (P < 0.05) in LG compared with HG and NG in the white and red gastrocnemius but not in the plantaris and soleus (Fig.
2). Stimulation with a submaximal insulin
concentration (100 µU/ml) in muscles with NG caused an approximately
half-maximal stimulation of 2-deoxyglucose uptake in the four muscle
types. At a maximally stimulating (35) insulin
concentration (10,000 µU/ml) 2-deoxyglucose uptake in NG rats was
higher in the oxidative than in the glycolytic muscles (soleus > red gastrocnemius > plantaris > white gastrocnemius). With
regard to the different muscle glycogen levels, there appear to be
major differences both in insulin sensitivity and in responsiveness. In
white and red gastrocnemius, plantaris, and soleus, respectively, 2-deoxyglucose uptake at submaximal insulin stimulation was 4.6-, 3.2-, 2.6-, and 1.6-fold higher (P < 0.05, except for the
soleus) in LG compared with HG rats (Fig. 2). When correcting for the different basal glucose transport values, the respective increases in
2-deoxyglucose uptake from basal to submaximal insulin stimulation were
2.4, 9.4, 6.5, and 10.2 µmol · g1 · h
1 in
glycogen-supercompensated muscles (HG) compared with 15.2, 30.5, 17.3, and 17.3 µmol · g
1 · h
1
in glycogen-depleted muscles (LG; P < 0.05 for all
muscles except soleus). Maximally insulin-stimulated (10,000 µU/ml)
glucose transport rates were only affected (P < 0.05)
by glycogen levels in the white gastrocnemius, with the increase from
basal to maximal insulin stimulation being 7.0, 12.6, and 15.0 µmol · g
1 · h
1 in HG, NG,
and LG, respectively (Fig. 2A). Whenever a significant difference in 2-deoxyglucose uptake was observed between HG and LG, the
value of the NG group was intermediate to those extremes. Thus we found
significant negative correlations between muscle glycogen content and
glucose transport rate at submaximal (white and red gastrocnemius) and
maximal insulin stimulation (white gastrocnemius), as shown in Table
1. It is noteworthy that, in glycogen-depleted rats (LG), a high physiological insulin concentration of 100 µU/ml caused a maximally effective increase in glucose transport, as no further increase was noticed at 10,000 µU/ml insulin
in white and red gastrocnemius and plantaris.
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GLUT-4 cell surface content.
To measure cell surface GLUT-4 content in predominantly fast-twitch
glycolytic muscle fibers, where the largest effect of glycogen content
on glucose transport was observed, it was necessary to use the intact
epitrochlearis muscles, since the white part of the gastrocnemius
muscle could not be incubated. As described in METHODS, the
epitrochlearis, like the white gastrocnemius, contains 75-80%
fast-twitch glycolytic fibers. The GLUT-4 cell surface content in
incubated epitrochlearis muscles in the presence of 0, 100, and 10,000 µU/ml insulin, respectively, was 1.8-fold, 3.5-fold
(P < 0.05), and 2.3-fold (P < 0.05)
higher in LG compared with HG (Fig. 3).
The insulin-induced increase in GLUT-4 cell surface content in muscles
with HG vs. LG, respectively, was 2.2 ± 0.4 vs. 8.6 ± 1.7 (arbitrary units) at submaximal insulin (P < 0.05) and
5.8 ± 0.8 vs. 13.4 ± 1.9 (P < 0.05) at
maximal insulin concentration (Fig. 3).
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Early insulin signaling activity in white gastrocnemius. Because the largest effect of glycogen content on glucose transport was observed in the white gastrocnemius, we initially decided that possible glycogen effects on insulin signaling would be largest in this muscle. Thus initial signaling assays were performed only in white gastrocnemius. Furthermore, we assayed muscles with LG and HG, and only if differences between these groups were found was NG assayed also.
Basal IRTK and PI3K activities were similar in LG and HG (Fig. 4). The IRTK activity increased (P < 0.05) ~2-fold with submaximal and ~16-fold with maximal insulin stimulation in both groups. IRTK activity was not affected by the muscle glycogen content at any insulin concentration. It should be noted that the increase in IRTK activity in response to 100 µU/ml insulin was only 5 and 3% of the increase induced by 10,000 µU/ml in HG and LG, respectively (Fig. 4A), whereas the glucose transport rate at submaximal insulin levels was 34 and 94% of the maximal response, pointing to a large spare IRTK activity. By contrast, PI3K activity (Fig. 4B) increased (P < 0.05) approximately twofold with submaximal and approximately fivefold with maximal insulin stimulation and was not affected by the muscle glycogen content at any insulin concentration.
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PKB- phosphorylation and activity.
In the present study, we initially evaluated the degree of Akt1/PKB-
activity by the degree of Ser473 phosphorylation. Akt/PKB
phosphorylation was increased more by insulin in LG than in HG, but the
difference only reached statistical significance at maximal insulin
levels (Fig. 5). Because differences in
Akt/PKB phosphorylation were found between LG and HG, we next studied
Akt/PKB activity at all three glycogen levels and in both the white and
red gastrocnemius (Fig. 6). The basal PKB
activity was not different between the glycogen groups. In the white
gastrocnemius, PKB activity increased significantly with submaximal and
maximal insulin stimulation in all groups, but the increases were
significantly affected by the muscle glycogen content. In HG, NG, and
LG, respectively, submaximal insulin stimulation caused 1.5-, 2.2-, and
3.8-fold increases (P < 0.05 for LG vs. NG and HG),
and maximal insulin stimulation caused 5.0-, 11.0-, and 16.7-fold
increases (P < 0.05 for LG vs. HG) in PKB activity.
Thus PKB activity was negatively correlated with muscle glycogen
content at both insulin concentrations, and PKB activity also
correlated positively (r = 0.68; P < 0.01) with glucose transport rates at submaximal insulin stimulation (Table 1). In the red gastrocnemius, insulin stimulation of PKB activity was not dependent on glycogen (Fig. 6B).
Interestingly, although PKB activity was not dependent on muscle
glycogen in this fiber type, there still was a significant positive
correlation between PKB activity and glucose transport at submaximal
insulin concentrations (Table 1).
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DISCUSSION |
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In the present study, we have shown that the effect of glycogen on insulin-stimulated glucose transport and GLUT-4 translocation is restricted to fast-twitch muscle fibers. In addition, we have shown that this glycogen effect on insulin action may involve downstream (Akt/PKB) but not upstream (IRTK- and IRS-1-associated PI3K) insulin signaling events. This fiber-type specificity could not be attributed to fiber type-dependent differences in effectiveness in glycogen depletion/supercompensation protocols, since the degree of difference in glycogen content between HG and LG was not smaller in slow-twitch (6.4-fold in soleus) compared with fast-twitch (4.5-fold in white gastrocnemius) muscle. We have recently reported a similar fiber type-specific regulatory role of glycogen in contraction stimulation of glucose transport (10).
When trying to understand how glycogen, directly or indirectly, affects insulin action on glucose uptake, it is critical to know the precise site of regulation of the glucose utilization process. By using the sensitive biotinylated bis-mannose photolabel (Bio-LC-ATB-BMPA; see Ref. 24), we presently show that most, if not all, of the variation in glucose transport with varying glycogen levels can be explained by the number of GLUT-4 proteins on the surface membrane. This was also recently shown by Kawanaka et al. (20) in maximally insulin-stimulated muscles using the radioactive bis-mannose photolabel.
It thus appears that a high glycogen content in muscle is associated with a state of insulin resistance to translocate GLUT-4-containing vesicles. It was previously hypothesized that glycogen inhibits GLUT-4 translocation by an association between GLUT-4-containing vesicles and glycogen particles, making the former "unavailable" for translocation. However, we presently believe that regulation of glucose transport by glycogen is not situated at the translocation level itself but is more upstream at the signaling level. Kawanaka et al. (20) have recently shown that glycogen supercompensation inhibits not only insulin-stimulated glucose transport but also insulin-stimulated amino acid transport. It is therefore tempting to speculate that glycogen levels affect insulin signaling in muscle. Tyrosine phosphorylation of IRS-1 by IRTK activity and subsequent binding to and activation of PI3K have been identified as two important initial events in the insulin transduction pathway leading to increased glucose transport. As shown in Fig. 4, neither IRTK activity nor IRS-1-associated PI3K activity is different between muscles with HG or LG during stimulation with 100 or 10,000 µU/ml insulin. By analogy, in vivo studies in humans (46) and rats (13) have failed to detect increased insulin-stimulated IRTK and PI3K after a glycogen-depleting exercise bout. From a teleological point of view, however, it is not completely surprising that a metabolic feedback signal (e.g., a signal indicating "intracellular energy/substrate abundance" during glycogen supercompensation) will not interfere with the initial step of insulin signaling (IRTK activity), since this step is shared by other signaling pathways of insulin, leading to nonmetabolic events (e.g., the mitogen-activated protein kinase signaling cascade leading to cell growth and mitosis). In other words, it could be expected that the "product inhibition" exerted by glycogen is restricted to processes that lead to its synthesis.
PKB (also known as Akt or Rac) has been proposed to be a mediator of
the insulin signaling pathway downstream of PI3K (reviewed in Refs. 1
and 38). PKB is activated by phosphorylation on Thr308 and
Ser473 by PDK-1 and phosphoinositide-dependent protein
kinase-2 (PDK-2) and binding of phosphatidylinositol
3,4,5-trisphosphate, the major catalytic product of activated PI3K
(38). A new finding of the present study is that
PKB-/Akt1 activation (assessed by an in vitro activity assay and by
the phosphorylation status on Ser473) in response to a
standardized submaximal or maximal insulin stimulus is dependent on the
muscle glycogen content. It is speculative but tempting to attribute
causality to the relationship between the changes in PKB activation and
the parallel changes in glucose transport and cell surface GLUT-4
content in insulin-stimulated muscles with varying glycogen content.
The majority of recent papers support the notion of a functional link
between PKB activation and glucose transport stimulation in muscle
(15, 23, 25, 26, 42), whereas others do not (27,
39). In the present study, glycogen content in the red
gastrocnemius muscle affected insulin-stimulated glucose transport but
not PKB activity. The data on this muscle type with mixed fiber-type
composition (red gastrocnemius) therefore indicate that a causal link
between the PKB activity and the glucose transport rate should be taken
with precaution. Furthermore, it should be noted that not only PKB but
also some isoforms of PKC have recently been suggested to be downstream
effectors of PI3K and to be involved in insulin-stimulated glucose
transport regulation. However, the present literature in this area is
still inconclusive, because most work so far has been limited to
cultured cell lines, and controversy still exists about which isoforms
of PKC are actually involved. Most reports indicate the atypical
isoforms
and
(40) as the ones that participate in
insulin stimulation of glucose transport, whereas others also report
the involvement of the conventional isoform
2
(6) and the novel isoform
(5). In
relation to the present study, it will be most interesting to
investigate in mammalian skeletal muscle whether the insulin activation
of various PKC isoforms is dependent on the muscle glycogen content.
In a subset of experiments, the in vivo venous plasma glucose concentrations were measured before perfusion in rats from the HG and LG groups. As expected from their food intake, the results show significantly (P < 0.05; n = 10-12) higher circulating glucose levels in HG (12.4 ± 1.1 mM) compared with LG (6.3 ± 0.8 mM). Thus it might be suggested that the downregulation of insulin activation of PKB and glucose transport might in fact not be directly related to glycogen levels but rather to the hyperinsulinemia and hyperglycemia that will prevail in the carbohydrate-fed HG rats compared with the fat-fed LG rats. Such a notion is difficult to evaluate, because the different muscle glycogen levels in the present study cannot be separated from hyperinsulinemia and hyperglycemia. However, the following arguments support the notion that it is the glycogen level per se rather than hyperinsulinemia and hyperglycemia that is responsible. First, there was no effect of glycogen levels on the proximal insulin signaling steps, which might be expected if we are dealing with a downregulation of insulin signaling by chronic insulin stimulation, as recently shown by Pryor et al. (33). Second, muscles were preperfused for 20 min with a standardized glucose-free medium before measurements were made, so that, at least for the last 20 min, insulin and glucose were identical in the groups. Finally, in a previous study in which insulin resistance of glucose uptake in muscle was found in muscles with high glycogen levels (19), the glycogen manipulation protocol in some of the studies included a period of 12-h fasting before the experiment in groups, ending up with different glycogen levels and different insulin effect on glucose uptake. Although plasma insulin and glucose levels in these rats were not reported, the 12-h fasting in both groups is believed to result in largely similar values. Taken together, although final proof is lacking, we believe that the evidence favors a direct role of glycogen in insulin action rather than an indirect role related to high insulin and glucose levels in the blood.
A striking finding of the present study is that the differences in insulin activation of PKB with varying muscle glycogen levels did not appear to involve IRS-1-associated PI3K. Interestingly, two papers published last year (27, 39) reported observations that support ours, namely that hyperglycemia-induced insulin resistance in fast-twitch fibers coincides with impaired PKB activation but not PI3K activation. This supports the attractive hypothesis that the mechanisms by which hyperglycemia and glycogen supercompensation induce insulin resistance are similar and involve a downregulation of the insulin signaling at the site of PKB. Both situations indeed share the situation of abundant carbohydrate availability and the necessity to downregulate the entry of more glucose into the muscle cells to prevent overaccumulation. Furthermore, we show that the opposite situation, i.e., decreased carbohydrate availability, is associated with facilitated insulin stimulation of PKB. To further illustrate the pivotal role of PKB in insulin action, Chen et al. (7) have shown that osmotic shock inhibits insulin-stimulated glucose transport by downregulating PKB phosphorylation without affecting IRS-1 or PI3K activity. In an attempt to explain decreased PKB activation in the face of normal PI3K activity, an alternative PI3K-independent upstream signaling pathway could be hypothesized. Interestingly, Wojtaszewski et al. (48) have reported that insulin alone can modestly activate PKB, and insulin plus exercise can markedly activate PKB in mice in which the muscle-specific insulin receptor was knocked out and in which there consequently was no apparent activation of IRTK or PI3K.
One can only speculate on how high glycogen levels, directly or through another metabolite or signal, regulate insulin activation of PKB. In this context, one might speculate why the relationship is only observed in fast-twitch glycolytic muscle. Glycogen is known to form complexes with the sarcoplasmic reticulum and the enzymes that regulate glycogen metabolism. In fast-twitch glycolytic muscle, sarcoplasmic reticulum is more abundant than in oxidative muscle (2), and it might be that some of the enzymes (e.g., PKB) of the insulin signaling cascade can also bind to the glycogen-sarcoplasmic reticulum complex, which might make them less available in the cascade. This remains to be established, however.
One important health benefit of exercise is the increased insulin sensitivity in the period after an exercise bout. The mechanism for this increased insulin sensitivity is poorly understood but could possibly be ascribed to the low muscle glycogen levels caused by exercise; hence, the mechanism involves the stimulatory role of glycogen depletion on downstream insulin signaling, as illustrated by the present results in fast-twitch rat muscle. Indeed, Thorell et al. (41) and Wojtaszewski et al. (48) reported in humans and in mice, respectively, that insulin-stimulated muscle glucose uptake was enhanced immediately after glycogen-depleting exercise, which could be associated with the higher PKB phosphorylation compared with the same insulin stimulus without prior exercise. In contrast, one study reported that 4 h after exercise, when insulin sensitivity is still markedly increased, PKB phosphorylation and activity is not different between exercised and nonexercised muscles (45). Thus some but not all recent studies suggest that increased postexercise insulin sensitivity may be caused by enhanced downstream signaling through an exercise factor that might be glycogen depletion. However, glycogen depletion is clearly not the only factor, since some studies have shown that increased insulin sensitivity persists when glycogen levels have returned to baseline levels (12, 35), and another study has indicated that the exercise-induced increase in insulin sensitivity requires a serum factor (11).
In conclusion, this study indicates that the activation of muscle
glucose transport and GLUT-4 translocation in response to a submaximal
or maximal insulin stimulus is modulated by the glycogen content in
fast-twitch but not slow-twitch skeletal muscle. Furthermore, we have
shown that insulin activation of PKB-/Akt1 but not IRTK- or
IRS-1-associated PI3K activity is also modulated by muscle glycogen
content in fast-twitch muscle in a similar manner. Thus inhibition and
enhancement of insulin action by high and low muscle glycogen levels,
respectively, is proposed to involve downstream but not initial insulin
signaling events.
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ACKNOWLEDGEMENTS |
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G. D. Holman (University of Bath, UK) is acknowledged for kind donation of the surface labeling compound. We are grateful to B. Bolmgren, E. Hornemann, and P. Jensen for superior technical assistance.
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FOOTNOTES |
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This study was supported by Danish National Research Foundation Grant 504-14, by the Novo Nordisk Foundation, and by the Danish Diabetes Research Foundation.
Address for reprint requests and other correspondence: E. A. Richter, Copenhagen Muscle Research Centre, Human Physiology, Univ. of Copenhagen, 13 Universitetsparken, DK-2100 Copenhagen, Denmark (E-mail: erichter{at}aki.ku.dk).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 15 March 2000; accepted in final form 31 May 2000.
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REFERENCES |
---|
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---|
1.
Alessi, DR,
and
Downes CP.
The role of PI 3-kinase in insulin action.
Biochim Biophys Acta
1436:
151-164,
1998[ISI][Medline].
2.
Åstrand, PO,
and
Rodahl K.
Textbook of Work Physiology. New York: McGraw-Hill, 1986, p. 33.
3.
Baron, AD,
Brechtel G,
Wallace P,
and
Edelman SV.
Rates and tissue sites of non-insulin and insulin-mediated glucose uptake in humans.
Am J Physiol Endocrinol Metab
255:
E769-E774,
1988
4.
Bjornholm, M,
Kawano Y,
Lehtihet M,
and
Zierath JR.
Insulin receptor substrate-1 phosphorylation and phosphatidylinositol 3- kinase activity in skeletal muscle from NIDDM subjects after in vivo insulin stimulation.
Diabetes
46:
524-527,
1997[Abstract].
5.
Braiman, L,
Alt A,
Kuroki T,
Ohba M,
Bak A,
Tennenbaum T,
and
Sampson SR.
Protein kinase Cdelta mediates insulin-induced glucose transport in primary cultures of rat skeletal muscle.
Mol Endocrinol
13:
2002-2012,
1999
6.
Braiman, L,
Sheffi-Friedman L,
Bak A,
Tennenbaum T,
and
Sampson SR.
Tyrosine phosphorylation of specific protein kinase C isoenzymes participates in insulin stimulation of glucose transport in primary cultures of rat skeletal muscle.
Diabetes
48:
1922-1929,
1999[Abstract].
7.
Chen, D,
Fucini RV,
Olson AL,
Hemmings BA,
and
Pessin JE.
Osmotic shock inhibits insulin signaling by maintaining Akt/protein kinase B in an inactive dephosphorylated state.
Mol Cell Biol
19:
4684-4694,
1999
8.
Cline, GW,
Petersen KF,
Krssak M,
Shen J,
Hundal RS,
Trajanoski Z,
Inzucchi S,
Dresner A,
Rothman DL,
and
Shulman GI.
Impaired glucose transport as a cause of decreased insulin-stimulated muscle glycogen synthesis in type 2 diabetes.
N Engl J Med
341:
240-246,
1999
9.
Cushman, SW,
and
Wardzala LJ.
Potential mechanism of insulin action on glucose transport in the isolated rat adipose cell. Apparent translocation of intracellular transport systems to the plasma membrane.
J Biol Chem
255:
4758-4762,
1980
10.
Derave, W,
Lund S,
Holman GD,
Wojtaszewski J,
Pedersen O,
and
Richter EA.
Contraction-stimulated muscle glucose transport and GLUT-4 surface content are dependent on glycogen content.
Am J Physiol Endocrinol Metab
277:
E1103-E1110,
1999
11.
Gao, J,
Gulve EA,
and
Holloszy JO.
Contraction-induced increase in muscle insulin sensitivity: requirement for a serum factor.
Am J Physiol Endocrinol Metab
266:
E186-E192,
1994
12.
Garetto, LP,
Richter EA,
Goodman MN,
and
Ruderman NB.
Enhanced muscle glucose metabolism after exercise in the rat: the two phases.
Am J Physiol Endocrinol Metab
246:
E471-E475,
1984
13.
Goodyear, LJ,
Giorgino F,
Balon TW,
Condorelli G,
and
Smith RJ.
Effects of contractile activity on tyrosine phosphoproteins and PI 3-kinase activity in rat skeletal muscle.
Am J Physiol Endocrinol Metab
268:
E987-E995,
1995
14.
Goodyear, LJ,
Giorgino F,
Sherman LA,
Carey J,
Smith RJ,
and
Dohm GL.
Insulin receptor phosphorylation, insulin receptor substrate-1 phosphorylation, and phosphatidylinositol 3-kinase activity are decreased in intact skeletal muscle strips from obese subjects.
J Clin Invest
95:
2195-2204,
1995[ISI][Medline].
15.
Hajduch, E,
Alessi DR,
Hemmings BA,
and
Hundal HS.
Constitutive activation of protein kinase B alpha 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,
1998[Abstract].
16.
Hansen, BF,
Danielsen GM,
Drejer K,
Sørensen AR,
Wiberg FC,
Klein HH,
and
Lundemose AG.
Sustained signaling from the insulin receptor after stimulation with insulin analogues exhibiting increased mitogenic potency.
Biochem J
315:
271-279,
1996[ISI][Medline].
17.
Hespel, P,
and
Richter EA.
Glucose uptake and transport in contracting, perfused rat muscle with different pre-contraction glycogen concentrations.
J Physiol (Lond)
427:
347-359,
1990[Abstract].
18.
Holman, GD,
and
Kasuga M.
From receptor to transporter: insulin signalling to glucose transport.
Diabetologia
40:
991-1003,
1997[ISI][Medline].
19.
Jensen, J,
Aslesen R,
Ivy JL,
and
Brors O.
Role of glycogen concentration and epinephrine on glucose uptake in rat epitrochlearis muscle.
Am J Physiol Endocrinol Metab
272:
E649-E655,
1997
20.
Kawanaka, K,
Han DH,
Nolte LA,
Hansen PA,
Nakatani A,
and
Holloszy J.
Decreased insulin-stimulated GLUT-4 translocation in glycogen-supercompensated muscles of exercised rats.
Am J Physiol Endocrinol Metab
276:
E907-E912,
1999
21.
Kitamura, T,
Ogawa W,
Sakaue H,
Hino Y,
Kuroda S,
Takata M,
Matsumoto M,
Maeda T,
Konishi H,
Kikkawa U,
and
Kasuga M.
Requirement for activation of the serine-threonine kinase Akt (protein kinase B) in insulin stimulation of protein synthesis but not of glucose transport.
Mol Cell Biol
18:
3708-3717,
1998
22.
Klein, HH,
Kowalewski B,
Drenckhan M,
Neugebauer S,
Matthaei S,
and
Kotzke G.
A microtiter well assay system to measure insulin activation of insulin receptor kinase in intact human mononuclear cells.
Diabetes
42:
883-890,
1993[Abstract].
23.
Kohn, AD,
Summers SA,
Birnbaum MJ,
and
Roth RA.
Expression of a constitutively active Akt Ser/Thr kinase in 3T3-L1 adipocytes stimulates glucose uptake and glucose transporter 4 translocation.
J Biol Chem
271:
31372-31378,
1996
24.
Koumanov, F,
Yang J,
Jones AE,
Hatanaka Y,
and
Holman GD.
Cell surface biotinylation of GLUT-4 using bis-mannose photolabels.
Biochem J
330:
1209-1215,
1998[ISI][Medline].
25.
Krook, A,
Kawano Y,
Song XM,
Efendic S,
Roth RA,
Wallberg-Henriksson H,
and
Zierath JR.
Improved glucose tolerance restores insulin-stimulated Akt kinase activity and glucose transport in skeletal muscle from diabetic Goto-Kakizaki rats.
Diabetes
46:
2110-2114,
1997[Abstract].
26.
Krook, A,
Roth RA,
Jiang XJ,
Zierath J,
and
Wallberg-Henriksson H.
Insulin-stimulated Akt kinase activity is reduced in skeletal muscle from NIDDM subjects.
Diabetes
47:
1281-1286,
1998[Abstract].
27.
Kurowski, TG,
Lin Y,
Luo S,
Tsichlis PN,
Buse MG,
Heydrick SJ,
and
Ruderman N.
Hyperglycemia inhibits insulin activation of Akt/protein Kinase B but not phosphatidylinositol 3-kinase in rat skeletal muscle.
Diabetes
48:
658-663,
1999[Abstract].
28.
Lowry, OH,
and
Passonneau JV.
A Flexible System of Enzymatic Analysis. London: Academic, 1972, p. 1-291.
29.
Lund, S,
Flyvbjerg A,
Holman GD,
Larsen FS,
Pedersen O,
and
Schmitz O.
Comparative effects of IGF-I and insulin on the glucose transporter system in rat muscle.
Am J Physiol Endocrinol Metab
267:
E461-E466,
1994
30.
Lund, S,
Holman GD,
Schmitz O,
and
Pedersen O.
GLUT-4 content in the plasma membrane of rat skeletal muscle: comparative studies of the subcellular fractionation method and the exofacial photolabelling technique using ATB-BMPA.
FEBS Lett
330:
312-318,
1993[ISI][Medline].
31.
Maltin, CA,
Delday MI,
Baillie AS,
Grubb DA,
and
Garlick PJ.
Fiber-type composition of nine rat muscles. I. Changes during the first year of life.
Am J Physiol Endocrinol Metab
257:
E823-E827,
1989
32.
Ploug, T,
Van Deurs B,
Ai H,
Cushman S,
and
Ralston E.
Analysis of GLUT-4 distribution in whole skeletal muscle fibers: Identification of distinct storage compartments that are recruited by insulin and muscle contractions.
J Cell Biol
142:
1429-1446,
1998
33.
Pryor, PR,
Liu SC,
Clark AE,
Yang J,
Holman GD,
and
Tosh D.
Chronic insulin effects on insulin signalling and GLUT-4 endocytosis are reversed by metformin.
Biochem J
348:
83-91,
2000[ISI][Medline].
34.
Richter, EA.
Glucose utilization.
In: Handbook of Physiology: Exercise: Regulation and Integration of Multiple Systems. Bethesda, MD: Am Physiol Soc, 1996, sect. 12, chapt. 20, p. 912-951.
35.
Richter, EA,
Garetto LP,
Goodman MN,
and
Ruderman NB.
Muscle glucose metabolism following exercise in the rat. Increased sensitivity to insulin.
J Clin Invest
69:
785-793,
1982[ISI][Medline].
36.
Richter, EA,
Mikines KJ,
Galbo H,
and
Kiens B.
Effect of exercise on insulin action in human skeletal muscle.
J Appl Physiol
66:
876-885,
1989
37.
Ruderman, NB,
Houghton CR,
and
Hems R.
Evaluation of the isolated perfused rat hindquarter for the study of muscle metabolism.
Biochem J
124:
639-651,
1971[ISI][Medline].
38.
Shepherd, PR,
Withers DJ,
and
Siddle K.
Phosphoinositide 3-kinase: the key switch mechanism in insulin signalling.
Biochem J
333:
471-490,
1998[ISI][Medline].
39.
Song, XM,
Kawano Y,
Krook A,
Ryder JW,
Efendic S,
Roth RA,
Wallberg-Henriksson H,
and
Zierath JR.
Muscle fiber type-specific defects in insulin signal transduction to glucose transport in diabetic GK rats.
Diabetes
48:
664-670,
1999[Abstract].
40.
Standaert, ML,
Bandyopadhyay G,
Perez L,
Price D,
Galloway L,
Poklepovic A,
Sajan MP,
Cenni V,
Sirri A,
Moscat J,
Toker A,
and
Farese RV.
Insulin activates protein kinases C-zeta and C-lambda by an autophosphorylation-dependent mechanism and stimulates their translocation to GLUT-4 vesicles and other membrane fractions in rat adipocytes.
J Biol Chem
274:
25308-25316,
1999
41.
Thorell, A,
Hirshman MF,
Nygren J,
Jorfeldt L,
Wojtaszewski JF,
Dufresne SD,
Horton ES,
Ljungqvist O,
and
Goodyear LJ.
Exercise and insulin cause GLUT-4 translocation in human skeletal muscle.
Am J Physiol Endocrinol Metab
277:
E733-E741,
1999
42.
Turinsky, J,
and
Damrau-Abney A.
Akt kinases and 2-deoxyglucose uptake in rat skeletal muscles in vivo: study with insulin and exercise.
Am J Physiol Regulatory Integrative Comp Physiol
276:
R277-R282,
1999
43.
Wallberg-Henriksson, H.
Glucose transport into skeletal muscle.
Acta Physiol Scand Suppl
564:
1-80,
1987[Medline].
44.
Wang, Q,
Somwar R,
Bilan PJ,
Liu Z,
Jin J,
Woodgett JR,
and
Klip A.
Protein kinase B/Akt participates in GLUT-4 translocation by insulin in L6 myoblasts.
Mol Cell Biol
19:
4008-4018,
1999
45.
Wojtaszewski, J,
Hansen BF,
Gade J,
Kiens B,
Markuns JS,
Goodyear L,
and
Richter EA.
Insulin signaling and insulin sensitivity after exercise in human skeletal muscle.
Diabetes
49:
325-331,
2000
46.
Wojtaszewski, J,
Hansen BF,
Kiens B,
and
Richter EA.
Insulin signaling in human skeletal muscle. Time course and effect of exercise.
Diabetes
46:
1775-1781,
1997[Abstract].
47.
Wojtaszewski, JF,
Hansen BF,
Urso B,
and
Richter EA.
Wortmannin inhibits both insulin- and contraction-stimulated glucose uptake and transport in rat skeletal muscle.
J Appl Physiol
81:
1501-1509,
1996
48.
Wojtaszewski, JF,
Higaki Y,
Hirshman MF,
Michael MD,
Dufresne SD,
Kahn CR,
and
Goodyear LJ.
Exercise modulates postreceptor insulin signaling and glucose transport in muscle-specific insulin receptor knockout mice.
J Clin Invest
104:
1257-1264,
1999
49.
Wojtaszewski, JFP,
Jakobsen AB,
Ploug T,
and
Richter EA.
The perfused hindlimb is suitable for skeletal muscle glucose transport measurements.
Am J Physiol Endocrinol Metab
274:
E184-E191,
1998
50.
Zierath, JR,
He L,
Guma A,
Odegoard WE,
Klip A,
and
Wallberg-Henriksson H.
Insulin action on glucose transport and plasma membrane GLUT-4 content in skeletal muscle from patients with NIDDM.
Diabetologia
39:
1180-1189,
1996[ISI][Medline].