Contraction-stimulated muscle glucose transport and GLUT-4
surface content are dependent on glycogen content
Wim
Derave1,2,
Sten
Lund3,
Geoffrey D.
Holman4,
Jørgen
Wojtaszewski1,
Oluf
Pedersen5, and
Erik A.
Richter1
1 Copenhagen Muscle Research
Centre, August Krogh Institute, University of Copenhagen, 2100 Copenhagen; 5 Steno Diabetes
Center and Hagedorn Research Institute, 2820 Gentofte, Copenhagen;
3 Medical Research Laboratory,
Aarhus Kommunehospital, 8000 Aarhus, Denmark;
2 Department of Movement and
Sports Sciences, University of Gent, 9000 Gent, Belgium; and
4 Department of Biology and
Biochemistry, University of Bath, Bath BA2 7AY, United
Kingdom
 |
ABSTRACT |
The influence of
muscle glycogen content on basal and contraction-induced glucose
transport and cell surface GLUT-4 content was studied in rat skeletal
muscle. Wistar rats were preconditioned by a combination of swimming
exercise and diet, resulting in 40% lower (LG) or threefold higher
(HG) muscle glycogen content compared with nonexercised controls (NG).
At rest and during contractions, 2-deoxy-D-glucose uptake in
perfused fast-twitch muscle, but not slow-twitch muscle, was
significantly lower in HG compared with LG. Cell surface GLUT-4 content
in the fast-twitch plantaris was 994 ± 180, 1,173 ± 311, and
2,155 ± 243 dpm/g in the basal condition and increased
(P < 0.05) to 2,285 ± 239, 3,230 ± 464, and 4,847 ± 654 dpm/g during contractions with HG, NG,
and LG, respectively, the increase being significantly smaller in HG
compared with LG. The contraction-induced increments in glucose
transport and in cell surface GLUT-4 content were negatively correlated
with the initial glycogen content (P <0.01). In conclusion, glucose
transport and cell surface GLUT-4 content in resting and contracting
fast-twitch muscle are dependent on the muscle glycogen content.
glucose transporters; exercise; glycogen metabolism; perfused
hindlimb; 2-deoxy-D-glucose
 |
INTRODUCTION |
THE MOLECULAR MECHANISM leading to contraction-induced
glucose uptake in skeletal muscle is still incompletely understood, but
it is known to involve translocation of GLUT-4 glucose transporters from the intracellular storage sites to the surface membrane. It has
previously also been demonstrated that the muscle glycogen content
exerts some influence on contraction-induced glucose utilization (reviewed in Ref. 23). Thus during two-legged ergometer cycling, glucose uptake was higher in the glycogen-depleted leg than in the leg
with normal muscle glycogen level (5). In the perfused rat hindlimb,
the muscle glucose uptake rate during contractions was negatively
correlated with precontraction muscle glycogen content (8). One
explanation for this inhibitory effect of glycogen on the
contraction-induced muscle glucose uptake may be that high glycogen
levels promote contraction-induced glycogen breakdown, which gives rise
to high concentrations of glucose 6-phosphate, which in turn inhibit
hexokinase. Thus this mechanism suggests that the glucose uptake rate
is limited by the inhibition of the glucose phosphorylation step rather
than the sarcolemmal glucose transport step (5, 8, 11). However,
another possibility exists. As was first shown by Hespel and Richter
(8), muscle glucose transport capacity, as measured by the uptake of
radiolabeled 3-O-methyl-D-glucose,
is increased by contractions more in glycogen-depleted than in
glycogen-supercompensated muscle (8). Recently this was indirectly
confirmed by Kawanaka et al. (12), who found a negative correlation
between contraction-induced uptake of
2-deoxy-D-glucose (2DG) and
postcontraction muscle glycogen concentrations. These findings suggest
that contraction-induced GLUT-4 translocation to the surface membrane
may be affected by glycogen levels. Likewise, hypoxia-induced surface
labeling of GLUT-4 was shown to be reduced in trained compared with
sedentary rat muscle because of a training-induced increase in glycogen
level (21). Although hypoxia and contractions do not seem to share the
same signaling pathway for increasing muscle glucose transport (3, 29),
the latter finding lends credence to the possibility that
contraction-induced muscle GLUT-4 translocation may be dependent on
muscle glycogen content. Therefore, one aim of the present study was to
study directly whether cell surface GLUT-4 content in contracting
muscle is dependent on precontraction muscle glycogen content.
After an exercise bout, glucose transport remains elevated for some
time, and the reversal is dependent on carbohydrate intake. Thus, if a
fat-rich diet is fed after exercise, muscle glucose transport is still
somewhat elevated 18 h after exercise, whereas if carbohydrates are
fed, baseline levels of transport are reached much sooner (4, 8, 31).
The mechanism underlying this effect of carbohydrate feeding and the
associated glycogen storage after exercise is not known, and
furthermore it is not known whether the increased basal glucose
transport is due to persistent GLUT-4 transporters in the surface
membrane. Therefore, a second aim of the present study was to elucidate
whether the persistent increased glucose transport rate in muscle after
exercise, when animals are fed a fat-rich diet, is due to persistent
GLUT-4 transporters at the muscle surface membrane.
 |
METHODS |
Animals.
All experiments were approved by the Danish Animal Experiments
Inspectorate and complied with the "European Convention for the
Protection of Vertebrate Animals Used for Experiments and Other
Scientific Purposes" (Council of Europe no. 123, Strasbourg, France,
1985). Male Wistar rats (3-4 wk old, 60-100 g) were
preconditioned so that we could obtain three different subgroups with
varying muscle glycogen concentrations, as described previously (8). A
control group with normal muscle glycogen levels (normal glycogen, NG)
had free access to regular rat chow and tap water until 3-6 h
before perfusion. The rats of the other two groups were subjected to 2 h of swimming in water maintained at 32-35°C, with weights (6% 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, they were fed ad libitum with either lard and
tap water (low glycogen, LG) or with normal rat chow, tap water, and a
20% glucose drinking solution (high glycogen, HG) until 3-6 h
before perfusions. Rats were perfused between 18 and 24 h after the
swimming bout.
Surgical procedure.
The rats were anesthetized by an intraperitoneal injection of
pentobarbital sodium (5 mg/100 g body weight). Surgery was performed as
described by Ruderman et al. (26) for isolated hindquarter perfusion.
Perfusion medium.
All perfusions were carried out using a cell-free and glucose-free
perfusate consisting of Krebs-Ringer bicarbonate buffer solution
(KRBB), 4% bovine serum albumin (fraction V, Sigma Chemical) dialyzed
(pore size 10-15 kDa) twice for 24 h against 11 volumes of KRBB,
0.15 mM pyruvate, and 4.2 IU/ml heparin, as previously described (28).
Media having this composition were used throughout the perfusion in the
surface labeling experiments and in the initial stage during the
glucose transport experiments. For measurement of glucose transport, 8 mM of 2DG (Sigma) and 1 mM mannitol (Sigma) together with radioactive
labeled tracers
2-deoxy-D-[2,6-3H]glucose
(specific activity 51 Ci/mmol; Amersham International, UK) and
D-[1-14C]mannitol
(specific activity 57 mCi/mmol; Amersham International) yielding
activities of 0.075 and 0.05 µCi/ml, respectively, were used as
previously described (28).
Perfusion procedure.
The perfusion medium (100 ml) was gassed with a mixture of 95%
oxygen-5% carbon dioxide. The oxygen pressure and pH of the recirculated arterial perfusion medium ranged between 450 and 550 mmHg
and 7.3 and 7.4, respectively. The temperature of the perfusate was
35°C, which resulted in a muscle temperature in the calf muscles of
~32°C. With respect to the viability of the presently used muscle
preparation, we have previously shown that muscle ATP and creatine
phosphate values during 45 min of basal perfusion with a cell-free
medium do not change compared with values obtained from rested
anesthetized rat muscles (27). The perfusion pressure ranged between 30 and 50 mmHg in resting conditions and between 40 and 60 mmHg during
contractions. The initial 10 ml of perfusate were discarded, and
thereafter a 15-min equilibration period was carried out, with the
hexose-free medium recirculating at a flow of 5 ml/min perfusing both
legs (~0.4
ml · min
1 · g
muscle
1). After 10 min of
equilibration, the left common iliac artery and vein were ligated, and
muscle biopsies were taken from the left leg (for determination of
precontraction glycogen levels). After the remaining 5 min of the
equilibration perfusion, the right leg was made to contract
isometrically by electrical stimulation of the sciatic nerve for 10 min
while flow was maintained at 5 ml/min. The electrical stimulation was
performed with supramaximal trains (25 V) of 100 ms delivered at 2-s
intervals and an impulse duration and frequency within the train of 0.1 ms and 100 Hz, respectively. During the last 4 min of contractions, the
hindlimb was perfused with the hexose- and tracer-containing perfusate, without recirculation (to ensure a constant concentration of 2DG in the
arterial perfusate throughout the exposure time). Immediately after the
perfusion, the muscles of the right (stimulated) leg were biopsied.
Muscle samples were taken from four different parts of the calf muscles
representing a whole range of fiber type distributions. The approximate
frequencies of slow-twitch oxidative (SO), fast-twitch oxidative
glycolytic (FOG), and fast-twitch glycolytic (FG) fibers of rats aged
20-34 days (60-100 g) are taken from Maltin et al. (16) and
are given in parentheses (SO:FOG:FG, together with an abbreviation of
the muscle name used in this paper). The white, most superficial part
of the gastrocnemius (0:20:80; WG), the plantaris (10:50:40; Plant),
the red, deep proximal and medial portion of gastrocnemius (10:55:35;
RG), and the soleus (55:40:5; Sol) 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.
For measurement of basal glucose transport, both hindlimbs were
perfused at a flow of 5 ml/min for 10 min with recirculating
hexose-free medium and subsequently for 20 min with the hexose- and
tracer-containing medium without recirculation. Immediately after
perfusion, the right leg was biopsied. Longer periods of radioactive
exposure time were used during basal perfusions (20 min) than during
contractions (4 min) to get high enough counts in the muscle extracts
during basal perfusions. In a subsample of experiments, arterial and
venous perfusate samples were taken at rest and after 5 min of
contractions and were analyzed immediately for
O2 pressure (ABL 510 acid-base laboratory, Radiometer). Oxygen uptake was calculated as described previously (3).
Glycogen and glucose transport measurements.
Muscle glycogen content was measured as glucose residues by a
hexokinase method after acid hydrolysis (13). The 2DG taken up by the
different muscles was determined in perchloric acid extracts and
corrected for label in the extracellular space determined by the
14C counts for mannitol.
Radioactivity was measured in a liquid scintillation counter (model
2000 Tri-Carb, Packard Instruments, Downers Grove, IL). From the
intracellular accumulation of
2-deoxy-D-[2,6-3H]glucose,
the rate of glucose transport was calculated using a specific activity
of hexose determined by the hexose concentration and
2-deoxy-D-[2,6-3H]glucose
counts in the perfusate and was expressed as micromoles per gram of
muscle per hour.
Photoaffinity labeling of cell surface GLUT-4.
Rats were perfused as described previously, with the hexose-free
perfusate only. The rested and stimulated legs were perfused simultaneously at a flow of 5 ml/min during an equilibration period (15 min) and 8 ml/min during the 10-min contraction period (100-ms trains
at 2-s intervals). A flow of 8 ml/min was chosen in two-legged perfusions because this flow resulted in the same pressure in the
arterial tubing as 5 ml/min in one-legged perfusions during contractions. After stimulation, perfusion was stopped, and the plantaris muscles were rapidly but carefully dissected out from rested
and stimulated legs and without further stimulation were immediately
transferred to a dark room and incubated at 18°C for 8 min in KRBB
containing 1 mCi/ml
ATB-[2-3H]BMPA,
2-N-4-(1-azi-2,2,2-trifluoroethyl)benzoyl-1,3-bis(D-mannose-4-yloxy)-2-propylamine (ATB-BMPA; specific activity
10 Ci/mmol), prepared as previously described (2). The muscles were then irradiated for 6 min with manual
turning over after 3 min in a Rayonet RPR 100 photochemical reactor (RPR 3000 lamps). After irradiation, muscles were
blotted and trimmed of visible tendons and frozen in liquid nitrogen. The frozen muscles were weighed and further processed as described previously (14), except that the immunoprecipitation was carried out
overnight at 4°C. Labeled GLUT-4 protein was expressed as disintegrations per minute per gram wet muscle weight.
Total muscle GLUT-4 content.
Total crude membranes (TCM) were prepared from 30-40 mg of
individual muscles, as described previously (20). TCM samples were
separated using a 10% SDS-PAGE gel (Mini Protean II, Bio-Rad Laboratories, Hercules, CA) and then transferred to an Immobilon P
membrane (Millipore, Bedford, MA) by semi-dry blotting. Subsequently the Immobilon P membrane was blocked for 2 h in 10 mM Tris and 0.9%
NaCl, pH 7.4, or TBS buffer, containing 5% defatted milk powder,
followed by incubation with a primary GLUT-4 antibody (goat polyclonal
IgG, Santa Cruz Biotechnology, Santa Cruz, CA) overnight at 4°C.
Then the membrane was washed and incubated with a secondary alkaline
phosphatase conjugated rabbit anti-goat antibody (Pierce, Rockford,
IL). A chemifluorescence substrate for alkaline phosphatase (Attophos,
Amersham International) was added to the washed membrane, and the
antibody-antigen complexes were visualized and quantified with a
chemifluorescence scanner (Storm 840, Molecular Dynamics, Sunnyvale,
CA). TCM GLUT-4 protein content per microgram protein (measured with
Pierce Microtiter Protocol, Rockford, IL) was expressed in arbitrary
units relative to a TCM standard.
Statistics.
Statistical evaluation of the data was done by
t-tests or one-way ANOVA by use of the
Student-Newman-Keuls method for post hoc multiple comparisons, where
appropriate. 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.
 |
RESULTS |
Muscle glycogen.
Muscle glycogen content in glycogen-supercompensated rats (HG) was
four- to sixfold higher than in glycogen-depleted (LG) rats (Table
1). The effect of the
pretreatment (swimming and diet) on glycogen levels was
similar in all of the muscles studied. However, the absolute muscle
glycogen content was approximately two- to threefold lower in Sol than
in the RG, WG, and Plant. The glycogen breakdown during contractions
was two- to threefold higher in HG muscles compared with LG muscles in
WG, RG, and Plant (P < 0.05) but not
significantly different in Sol. At the end of electrical stimulation,
the glycogen content was 4- (Sol), 10- (WG), 12- (Plant), and 16-fold
(RG) higher in HG muscles than in LG muscles
(P < 0.05).
View this table:
[in this window]
[in a new window]
|
Table 1.
Initial and final muscle glycogen levels and glycogen breakdown in
contracting muscles with high and low glycogen content
|
|
Glucose transport.
Basal glucose transport, as measured by uptake of 2DG, was
approximately twofold higher in LG than in HG in WG, RG, and Plant, whereas in Sol, basal glucose transport was unaffected by glycogen levels (Fig.
1A).
Basal glucose transport correlated negatively with the glycogen content
in Plant (r =
0.50;
P < 0.05) and RG (r =
0.40;
P < 0.05) but not in the WG
(r =
0.32) and Sol
(r =
0.12). After 10 min of
electrical stimulation, glucose transport in WG, RG, and Plant was
increased four- to eightfold over basal (Fig.
1B). Contraction-induced 2DG uptake
in LG was approximately twofold higher than in HG in these muscles
(P < 0.05). In the Sol, the
contraction-induced increase in glucose transport was very limited
(~2-fold), and no effect of glycogen content could be observed (Fig.
1B). To ensure that the lack of
effect of glycogen content in the Sol was not due to this limited
increase in glucose transport, we in additional experiments observed
that stimulation with a more intense stimulation protocol (200-ms
trains at 1-s intervals, instead of 100-ms trains at 2-s intervals)
could increase glucose transport in the Sol nearly fivefold over basal,
with no effect of glycogen content (see Table
2). It is our experience that applying this
stimulation protocol to muscles of larger rats results in larger
increases in glucose transport. Thus, in another set of experiments, we
applied the same submaximal and maximal stimulation protocols during
perfusion of rats weighing 200-250 g. In these larger rats, the
respective increases in Sol 2DG uptake were 6- and 15-fold over basal.
Still, there were no differences between HG and LG (Table 2).

View larger version (16K):
[in this window]
[in a new window]
|
Fig. 1.
Basal (A) and 10-min
contraction-stimulated (B)
2-deoxy-D-glucose (2DG) uptake
in individual muscles with high (filled bars, HG) and low (open bars,
LG) muscle glycogen content. WG and RG, white and red gastrocnemius,
respectively; Plant, plantaris; Sol, soleus. Measurements of basal and
contraction-stimulated 2DG uptake were done on separate groups of rats.
Data are presented as means ± SE
(n = 5-11). * Significant
difference between HG and LG (P < 0.05).
|
|
The glucose transport rate during contractions in the small rats was
significantly negatively correlated with the precontraction glycogen
levels in WG (r =
0.53;
P < 0.05), RG (r =
0.59;
P < 0.05), and Plant (r =
0.74;
P < 0.05), but not in the Sol
(r =
0.31). The glucose
transport rate during contractions was also correlated with
postcontraction glycogen levels in WG
(r =
0.49; P < 0.05), RG
(r =
0.61;
P < 0.05), and Plant
(r =
0.73;
P < 0.05), but not in the Sol
(r =
0.18). Finally, in WG
(r =
0.54; P < 0.05) and in Plant
(r =
0.69;
P < 0.05), there was also a significant correlation between glycogen breakdown (calculated as the
difference between initial and final glycogen content) and glucose transport.
Cell surface GLUT-4 content.
To study whether differences in glucose transport were due to different
recruitment of GLUT-4, surface labeling of the Plant was performed.
This muscle was chosen because, of the three muscles (WG, RG, and
Plant) in which glycogen was found to have an effect on glucose
transport, the Plant was the only one that could be dissected out as an
intact muscle and incubated. In addition to the HG and LG groups, we
also studied a third group of muscles from rested rats with normal
glycogen levels (NG). The muscle glycogen content in this NG group (40 ± 3 µmol/g) was intermediate and ~40% higher than in LG
(P < 0.05) and threefold lower
compared with HG (P < 0.05; Fig.
2C). In the basal state, the glucose
transport and the GLUT-4 content at the muscle cell surface were
significantly higher in the LG muscles compared with the HG and NG
groups (Fig. 2,
A and
B). Although there was a significant
negative linear correlation between glycogen content on the one hand
and basal glucose transport (r =
0.50; P < 0.05) or basal
GLUT-4 surface content (r =
0.53; P < 0.05) on the other,
the relation appears to be curvilinear (Fig.
3, A and
B). Thus the basal glucose transport and cell surface GLUT-4 content are only stimulated in the muscles where glycogen content is below normal, and very high glycogen levels
do not affect basal glucose transport or surface membrane GLUT-4
content, compared with normal glycogen levels. Contractions increased
the amount of GLUT-4 on the muscle cell surface two- to threefold
(P < 0.05; Fig.
2B). The contraction-induced GLUT-4 cell surface content in LG was over twofold higher compared with HG and
50% higher compared with NG (P < 0.05). Cell surface GLUT-4 content in NG tended to be higher than in HG
(P = 0.06; Fig.
2B). The contraction-induced
increments in both glucose transport rate (as calculated by the
contraction-induced glucose transport rate minus the mean glucose
transport rate of the basal group) and GLUT-4 cell surface content (as
calculated by the difference between cell surface GLUT-4 contents in
contracted and rested legs) were significantly negatively correlated
with the initial glycogen content (correlation coefficients of
0.70 and
0.59, respectively; P < 0.05; Fig. 4,
A and
B) and the postcontraction glycogen
level (correlation coefficients of
0.69 and
0.53,
respectively; P < 0.05). Although
glucose transport and cell surface GLUT-4 were not measured in the same
muscle samples, a positive correlation (r = 0.95, P < 0.01) was found between the six mean values
of both parameters in Plant muscles with HG, NG, and LG at rest and during contractions (Fig. 5).

View larger version (16K):
[in this window]
[in a new window]
|
Fig. 2.
2DG uptake (A), GLUT-4 cell surface
content (B), and glycogen content
(C) in plantaris muscles at rest
(filled bars) and after 10 min of contractions (open bars; 100-ms
trains with 2-s intervals). w.w., Wet weight. Rats were pretreated to
obtain muscles with high (HG), normal (NG), or low (LG) muscle glycogen
content. After hindlimb perfusion, plantaris muscles were dissected out
of the rested and electrically stimulated legs and were incubated in
ATB-BMPA to label cell surface GLUT-4. Glycogen and 2DG uptake were
determined in separate experiments, as in Fig. 1. Data are presented as
means ± SE (n = 5-8). w.w,
Wet wt. * Significantly different from HG
(P < 0.05);
significantly different from NG
(P < 0.05).
|
|

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 3.
Basal 2DG uptake (A) and basal
GLUT-4 cell surface content (B) as a
function of muscle glycogen content in plantaris muscles with HG
(triangles), NG (squares), and LG (circles). Individual data points are
given as filled symbols, and group means (± SE) are given as open
symbols.
|
|

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 4.
Effect of initial glycogen content on increments in 2DG uptake
(A) and cell surface GLUT-4 content
(B) from rest to contractions,
measured in plantaris muscles with HG ( ), NG ( ), and LG ( ).
Data are means ± SE (n = 5-8).
|
|

View larger version (16K):
[in this window]
[in a new window]
|
Fig. 5.
Correlation (r = 0.95, P < 0.01) between GLUT-4 cell
surface content and 2DG uptake in basal (closed symbols) and
contraction-stimulated (open symbols) plantaris muscles with HG
(triangles), NG (squares), and LG (circles). Each point is the mean ± SE of 6-8 observations.
|
|
Total muscle GLUT-4 content.
The total GLUT-4 protein content in the muscle of HG, NG, and LG,
respectively, was 1.4 ± 0.4, 1.1 ± 0.2, and 1.5 ± 0.4 (arbitrary units) in the Plant (n = 6). There were no significant differences between groups.
Body weight, contraction force, and oxygen uptake.
Body weight was 86 ± 3, 77 ± 2, and 80 ± 2 g immediately
before perfusion in HG, NG, and LG, respectively. The maximal force development during electrical stimulation in the perfused hindlimb was
1.27 ± 0.06, 1.26 ± 0.05, and 1.28 ± 0.04 N/g, and the mean force development was 0.50 ± 0.03, 0.46 ± 0.03, and 0.42 ± 0.02 N/g during 10 min of electrical stimulation in HG, NG, and LG, respectively (n = 12-14). There
were no significant differences between groups in maximal or mean
contraction force. Muscle oxygen uptake was measured immediately before
and at the end of the 10-min contraction period in a subsample of HG
(n = 8) and LG
(n = 7) rats. Oxygen uptakes were 12 ± 1 and 15 ± 1 µmol · g
1 · h
1
in resting hindquarters and increased to 30 ± 7 and 29 ± 7 µmol · g
1 · h
1
during contractions in HG and LG groups, respectively.
 |
DISCUSSION |
During the last two decades, a number of papers have suggested that
glycogen plays a role in regulating glucose entry in unstimulated skeletal muscle cells and in muscle cells stimulated with either insulin, hypoxia, or contractions (1, 4, 5, 8-10, 12, 18, 22, 31).
The present study is the first to demonstrate that in fast-twitch
muscle, the increment in cell surface GLUT-4 content in response to a
standardized contraction stimulus is inversely correlated to
precontraction muscle glycogen content. These findings thus suggest
that glycogen in muscle modifies GLUT-4 translocation in response to
muscle contractions. In addition, we have shown that when muscle
glycogen is maintained low by fat feeding after exercise, the surface
membrane content of GLUT-4 is increased in the basal state.
The effect of muscle glycogen content on basal and contraction-induced
glucose transport was consistently observed in hindlimb muscles with a
mixed (RG and Plant) or primarily glycolytic (WG) fiber-type
composition. In contrast, glycogen levels did not at all affect glucose
transport in the Sol, containing primarily slow-twitch fibers (Table 2
and Fig. 1). This could be interpreted in several ways. First, the Sol
is known to have a limited glycogen storage capacity (24, 25), causing
threefold lower glycogen levels in Sol compared with the other hindlimb
muscles (Table 1). Thus the inhibition of contraction-induced glucose
transport during glycogen supercompensation may only be detectable when glycogen levels exceed 100 µmol/g wet weight, a value that can hardly
be reached in rat Sol muscles (25). However, this interpretation does
not explain why there is no enhanced glucose transport in the highly
glycogen-depleted (precontraction glycogen levels below 15 µmol/g in
the LG group) Sol muscles. Second, the regulatory role of glycogen on
glucose transport may be truly fiber type specific and may therefore be
restricted to glycolytic fibers. In this context, it should be realized
that the effect of glycogen in fast-twitch muscle may not be due to
glycogen itself but rather to another metabolite or reporter of
intracellular energy status that changes concomitantly with glycogen.
If so, this mechanism would have to be more active in fast-twitch than
in slow-twitch fibers, either because of inherent differences between
the muscle fiber types or simply because of the different glycogen
levels in the different fiber types. Of interest in this regard is the recent work by W. W. Winder and colleagues (see Hayashi et al., Ref. 7,
and Merrill et al., Ref. 17), suggesting a role for 5'-AMP-activated protein kinase (AMPK) in contraction-stimulated glucose transport. Differential activation of AMPK, which is dependent on the fuel status of the cell, may help to explain the differential regulation of glucose transport in contracting glycogen-depleted and
supercompensated muscles.
It is noteworthy that the results obtained in the nonexercised and
non-dietary-manipulated control (NG) group were well within the data
obtained in the two groups with extreme muscle glycogen levels (Figs. 2
and 4). This is an important observation, because it signifies that the
differences in contraction-induced glucose transport between the groups
are not caused by some nonspecific effect of the combination of
exercise and diet but rather are genuinely related to differences in
muscle glycogen content. Neither were differences in
contraction-induced muscle glucose transport and GLUT-4 cell surface
content related to differences in total muscle GLUT-4 content, which
was similar in the three groups.
It was previously shown in perfused rat hindlimbs that the
contraction-induced increase in glucose uptake and transport is dependent on precontraction muscle glycogen concentration (8). In that
study, it was concluded that the decrease in contraction-induced glucose uptake in HG compared with LG muscles was due both to decreased
glucose phosphorylation and decreased glucose transport capacity in HG.
In the present study, we have provided further strong evidence that a
major effect of high muscle glycogen levels is a blunted
contraction-induced increase in muscle glucose transport capacity.
Furthermore, we have for the first time shown that this blunted
increase in glucose transport in HG muscle is due to a reduced amount
of functional GLUT-4 transporters at the muscle surface membrane.
In the present study, qualitative changes in glucose transport with
varying muscle glycogen levels are fully reflected by changes in cell
surface GLUT-4, supporting the opinion that the main mechanism for
increased glucose transport in contracting muscles is GLUT-4
translocation. However, the absolute increases of contractions over
basal are smaller in cell surface GLUT-4 data than in glucose transport
data. This is in apparent contrast to a previous study, in which the
contraction-mediated increase in GLUT-4 cell surface content fully
accounted for the increment in glucose transport after contractions in
the soleus (15). This discrepancy may have several explanations. First,
in contrast to the in situ measurement of basal glucose transport in
the perfused hindlimb, the handling and possibly also stretching of the
plantaris (while taking it out for incubation) may have led to
increased basal cell surface GLUT-4 content. Second, in the present
study, the glucose transport was measured during contractions in the perfused hindlimb, whereas the photolabeling experiments took place a
few minutes after cessation of contractions, the time necessary to take
out the plantaris and transfer it to the incubation medium. Reversal of
the contraction effect may have led to an underestimation of cell
surface GLUT-4 content during contractions. Third, the relatively thick
plantaris muscle may have represented a diffusion barrier to the
photolabel, which might contribute to the lower degree of increase in
surface labeling compared with glucose transport.
After a bout of glycogen-depleting exercise, carbohydrate feeding
speeds and carbohydrate restriction slows the reversal of exercise-induced glucose transport (8, 30, 31). The present results
indicate that, in muscles where normal glycogen stores have not yet
been completely restored 18-24 h after exercise (as in our LG
group), the increased glucose transport is due to a higher number of
GLUT-4 molecules at the surface of the muscle cell (Figs. 2 and 3). It
seems that, as soon as normal glycogen levels are restored (~40
µmol/g wet weight), the glycogen stores exert a negative feedback
signal to stop the GLUT-4 recruitment to the surface membrane. However,
muscle glycogen content is not the only regulator of postexercise
glucose uptake. Several in vitro studies have shown that the reversal
of glucose transport after exercise is not always dependent on glycogen
synthesis (6, 19, 30). In these studies, glucose transport returned to
baseline or near-baseline levels despite glycogen concentrations being maintained at a low level. Therefore, one may question whether glycogen
depletion itself, or another metabolite or signal, is the promoting
factor of increased cell surface GLUT-4 content postexercise.
Additionally, it is not clear whether GLUT-4 is trapped at the cell
surface and unable to return to the intracellular storage sites
(decreased endocytosis), or whether GLUT-4 endocytosis is unaffected,
but the GLUT-4 translocation to the membrane is continuously stimulated
(increased exocytosis).
In conclusion, we have demonstrated that the increased basal glucose
transport in glycogen-depleted muscle 18-24 h after exercise is
the result of increased appearance of active GLUT-4 molecules at the
muscle cell surface. We also provide evidence that contraction-induced glucose transport and GLUT-4 cell surface content are affected by the
muscle glycogen content in a concentration-dependent manner. Basal and
contraction-induced glucose transport in the soleus muscle is
unaffected by glycogen levels, possibly pointing to a fiber
type-specific mechanism. These data for the first time show the role of
glycogen in regulating GLUT-4 cell surface content in muscle at rest
and during contractions.
 |
ACKNOWLEDGEMENTS |
We thank Prof. Peter Hespel (University Leuven, Belgium) for
constructive criticism and inspiration throughout this work, and E. Hornemann and Betina Bolmgreen for excellent technical assistance.
 |
FOOTNOTES |
The present study was supported by grants from the Danish National
Research Foundation (no. 504-14 to E. A. Richter), from Novo-Nordisk Research Council (to E. A. Richter), from the Danish Sports Research Council (to E. A. Richter), and from the Medical Research Council, United Kingdom (to G. D. Holman).
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. §1734 solely to indicate this fact.
Address for correspondence and reprint requests: E. A. Richter,
Copenhagen Muscle Research Centre, August Krogh Institute, Univ. of
Copenhagen, 13, Universitetsparken, DK-2100 Copenhagen, Denmark
(E-mail: erichter{at}aki.ku.dk).
Received 9 April 1999; accepted in final form 13 July 1999.
 |
REFERENCES |
1.
Cartee, G. D.,
D. A. Young,
M. D. Sleeper,
J. Zierath,
H. Wallberg-Henriksson,
and
J. O. Holloszy.
Prolonged increase in insulin-stimulated glucose transport in muscle after exercise.
Am. J. Physiol.
256 (Endocrinol. Metab. 19):
E494-E499,
1989[Abstract/Free Full Text].
2.
Clark, A. E.,
and
G. D. Holman.
Exofacial photolabelling of the human erythrocyte glucose transporter with an azitrifluoroethylbenzoyl-substituted bismannose.
Biochem J.
269:
615-622,
1990[Medline].
3.
Derave, W.,
and
P. Hespel.
Role of adenosine in regulating muscle glucose uptake during contractions and hypoxia in rat skeletal muscle.
J. Physiol. (Lond.)
515:
255-263,
1999[Abstract/Free Full Text].
4.
Fell, R. D.,
S. E. Terblanche,
J. L. Ivy,
J. C. Young,
and
J. O. Holloszy.
Effect of muscle glycogen content on glucose uptake following exercise.
J. Appl. Physiol.
52:
434-437,
1982[Abstract/Free Full Text].
5.
Gollnick, P. D.,
B. Pernow,
B. Essén,
E. Jansson,
and
B. Saltin.
Availability of glycogen and plasma FFA for substrate utilization in leg muscle of man during exercise.
Clin. Physiol.
1:
27-42,
1981.
6.
Gulve, E. A.,
G. D. Cartee,
J. R. Zierath,
V. M. Corpus,
and
J. O. Holloszy.
Reversal of enhanced muscle glucose transport after exercise: roles of insulin and glucose.
Am. J. Physiol.
259 (Endocrinol. Metab. 22):
E685-E691,
1990[Abstract].
7.
Hayashi, T.,
M. Hirshman,
E. J. Kurth,
W. W. Winder,
and
L. Goodyear.
Evidence for 5'AMP-activated protein kinase mediation of the effect of muscle contraction on glucose transport.
Diabetes
47:
1369-1373,
1998[Abstract].
8.
Hespel, P.,
and
E. A. Richter.
Glucose uptake and transport in contracting, perfused rat muscle with different pre-contraction glycogen concentrations.
J. Physiol. (Lond.)
427:
347-359,
1990[Abstract].
9.
Host, H. H.,
P. A. Hansen,
L. A. Nolte,
M. M. Chen,
and
J. O. Holloszy.
Glycogen supercompensation masks the effect of a training-induced increase in GLUT-4 on muscle glucose transport.
J. Appl. Physiol.
85:
133-138,
1998[Abstract/Free Full Text].
10.
Jensen, J.,
R. Aslesen,
J. L. Ivy,
and
O. Brørs.
Role of glycogen concentration and epinephrine on glucose uptake in rat epitrochlearis muscle.
Am. J. Physiol.
272 (Endocrinol. Metab. 35):
E649-E655,
1997[Abstract/Free Full Text].
11.
Katz, A.,
K. Sahlin,
and
S. Broberg.
Regulation of glucose utilization in human skeletal muscle during moderate dynamic exercise.
Am. J. Physiol.
260 (Endocrinol. Metab. 23):
E411-E415,
1991[Abstract/Free Full Text].
12.
Kawanaka, K.,
I. Tabata,
A. Tanaka,
and
M. Higuchi.
Effects of high-intensity intermittent swimming on glucose transport in rat epitrochlearis muscle.
J. Appl. Physiol.
84:
1852-1857,
1998[Abstract/Free Full Text].
13.
Lowry, O. H.,
and
J. V. Passonneau.
A Flexible System of Enzymatic Analysis. London: Academic, 1972, p. 177-179.
14.
Lund, S.,
G. D. Holman,
O. Schmitz,
and
O. Pedersen.
GLUT4 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[Medline].
15.
Lund, S.,
G. D. Holman,
O. Schmitz,
and
O. Pedersen.
Contraction stimulates translocation of glucose transporter GLUT4 in skeletal muscle through a mechanism distinct from that of insulin.
Proc. Natl. Acad. Sci. USA
92:
5817-5821,
1995[Abstract/Free Full Text].
16.
Maltin, C. A.,
M. I. Delday,
A. G. S. Baillie,
D. A. Grubb,
and
P. J. Garlick.
Fiber-type composition of nine rat muscles. I. Changes during the first year of life.
Am. J. Physiol.
257 (Endocrinol. Metab. 20):
E823-E827,
1989[Abstract/Free Full Text].
17.
Merrill, G. F.,
E. J. Kurth,
D. G. Hardie,
and
W. W. Winder.
AICA riboside increases AMP-activated protein kinase, fatty acid oxidation, and glucose uptake in rat muscle.
Am. J. Physiol.
273 (Endocrinol. Metab. 36):
E1107-E1112,
1997[Medline].
18.
Nolte, L. A.,
E. A. Gulve,
and
J. O. Holloszy.
Epinephrine-induced in vivo muscle glycogen depletion enhances insulin sensitivity of glucose transport.
J. Appl. Physiol.
76:
2054-2058,
1994[Abstract/Free Full Text].
19.
Ploug, T.,
H. Galbo,
J. Vinten,
M. Jørgensen,
and
E. A. Richter.
Kinetics of glucose transport in rat muscle: effects of insulin and contractions.
Am. J. Physiol.
253 (Endocrinol. Metab. 16):
E12-E20,
1987[Abstract/Free Full Text].
20.
Ploug, T.,
J. Wojtaszewski,
S. Kristiansen,
P. Hespel,
H. Galbo,
and
E. A. Richter.
Glucose transport and transporters in muscle giant vesicles: differential effects of insulin and contractions.
Am. J. Physiol.
264 (Endocrinol. Metab. 27):
E270-E278,
1993[Abstract/Free Full Text].
21.
Reynolds, T. H., IV,
J. T. Brozinick, Jr.,
M. A. Rogers,
and
S. W. Cushman.
Effects of exercise training on glucose transport and cell surface GLUT-4 in isolated rat epitrochlearis muscle.
Am. J. Physiol.
272 (Endocrinol. Metab. 35):
E320-E325,
1997[Abstract/Free Full Text].
22.
Reynolds, T. H., IV,
J. T. Brozinick, Jr.,
M. A. Rogers,
and
S. W. Cushman.
Mechanism of the hypoxia-stimulated glucose transport in rat skeletal muscle: potential role of glycogen.
Am. J. Physiol.
274 (Endocrinol. Metab. 37):
E773-E778,
1998[Abstract/Free Full Text].
23.
Richter, E. A.
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.
24.
Richter, E. A.,
and
H. Galbo.
High glycogen levels enhance glycogen breakdown in isolated contracting skeletal muscle.
J. Appl. Physiol.
61:
827-831,
1986[Abstract/Free Full Text].
25.
Richter, E. A.,
B. F. Hansen,
and
S. A. Hansen.
Glucose-induced insulin resistance of skeletal-muscle glucose transport and uptake.
Biochem. J.
252:
733-737,
1988[Medline].
26.
Ruderman, N. B.,
C. R. Houghton,
and
R. Hems.
Evaluation of the isolated perfused rat hindquarter for the study of muscle metabolism.
Biochem. J.
124:
639-651,
1971[Medline].
27.
Wojtaszewski, J. F. P.,
B. F. Hansen,
B. Ursø,
and
E. A. Richter.
Wortmannin inhibits both insulin- and contraction-stimulated glucose uptake and transport in rat skeletal muscle.
J. Appl. Physiol.
81:
1501-1509,
1996[Abstract/Free Full Text].
28.
Wojtaszewski, J. F. P.,
A. B. Jakobsen,
T. Ploug,
and
E. A. Richter.
Perfused rat hindlimb is suitable for skeletal muscle glucose transport measurements.
Am. J. Physiol.
274 (Endocrinol. Metab. 37):
E184-E191,
1998[Abstract/Free Full Text].
29.
Wojtaszewski, J. F. P.,
J. L. Laustsen,
W. Derave,
and
E. A. Richter.
Hypoxia and contractions do not utilize the same signalling mechanism in stimulating skeletal muscle glucose transport.
Biochim. Biophys. Acta
1380:
369-404,
1998[Medline].
30.
Young, D. A.,
H. Wallberg-Henriksson,
M. D. Sleeper,
and
J. O. Holloszy.
Reversal of the exercise-induced increase in muscle permeability to glucose.
Am. J. Physiol.
253 (Endocrinol. Metab. 16):
E331-E335,
1987[Abstract/Free Full Text].
31.
Young, J. C.,
S. M. Garthwaite,
J. E. Bryan,
L.-J. Cartier,
and
J. O. Holloszy.
Carbohydrate feeding speeds reversal of enhanced glucose uptake in muscle after exercise.
Am. J. Physiol.
245 (Regulatory Integrative Comp. Physiol. 14):
R684-R688,
1983[Abstract/Free Full Text].
Am J Physiol Endocrinol Metab 277(6):E1103-E1110
0002-9513/99 $5.00
Copyright © 1999 the American Physiological Society