Analysis of insulin-stimulated skeletal muscle glucose uptake
in conscious rat using isotopic glucose analogs
Robert M.
O'Doherty,
Amy E.
Halseth,
Daryl K.
Granner,
Deanna
P.
Bracy, and
David H.
Wasserman
Department of Molecular Physiology and Biophysics, Vanderbilt
University School of Medicine, Nashville, Tennessee 37232
 |
ABSTRACT |
An isotopic method
was used in conscious rats to determine the roles of glucose transport
and the transsarcolemmal glucose gradient (TSGG) in control of basal
and insulin-stimulated muscle glucose uptake. Rats received an
intravenous
3-O-[3H]methylglucose
(3-O-[3H]MG)
infusion from
100 to 40 min and a
2-deoxy-[3H]glucose
infusion from 0 to 40 min to calculate a glucose metabolic index
(Rg). Insulin was infused from
100 to 40 min at rates of 0.0, 0.6, 1.0, and 4.0 mU · kg
1 · min
1,
and glucose was clamped at basal concentrations. The ratios of soleus
intracellular to extracellular
3-O-[3H]MG
concentration and soleus glucose concentrations were used to estimate
the TSGG using principles of glucose countertransport. Tissue glucose
concentrations were compared in well-perfused, slow-twitch muscle
(soleus) and poorly perfused, fast-twitch muscle (vastus lateralis,
gastrocnemius). Data show that 1)
small increases in insulin increase soleus
Rg without decreasing TSGG,
suggesting that muscle glucose delivery and phosphorylation can
accommodate the increased flux; 2)
due to a limitation in soleus glucose phosphorylation and possibly
delivery, insulin at high physiological levels decreases TSGG, and at
supraphysiological insulin levels the TSGG is not significantly
different from 0; 3) maximum
Rg is maintained even though TSGG
decreases with increasing insulin levels, indicating that glucose
transport continues to increase and is not rate limiting for maximal
insulin-stimulated glucose uptake; and
4) muscle consisting of fast-twitch
fibers that are poorly perfused exhibits a 35-45% fall in tissue
glucose with insulin, suggesting that glucose delivery is a major
limitation in sustaining the TSGG. In conclusion, control of glucose
uptake is distributed between glucose transport and factors that
determine the TSGG. Insulin stimulation of glucose transport increases
the demands on the factors that maintain glucose delivery to the muscle
membrane and glucose phosphorylation inside the muscle.
countertransport; 3-O-methylglucose; 2-deoxyglucose
 |
INTRODUCTION |
SKELETAL MUSCLE glucose uptake in vivo is determined by
the glucose transport activity of the sarcolemma and the
transsarcolemmal glucose gradient (TSGG). Considerable work has been
done to study the role and regulation of glucose transport into muscle
(47). Far less attention has been given to the role of the TSGG and the
factors that maintain it. The TSGG is determined by the delivery of
glucose to the outer surface of the sarcolemma and removal of glucose
from the inner surface. Muscle glucose delivery is a function of the
muscle blood flow and diffusion distance, and removal of glucose from
the inner sarcolemma surface is determined by diffusion and glucose
phosphorylation. Clearly, measurement of the TSGG would give valuable
insight into the control of glucose uptake under various physiological
and pathophysiological conditions. The glucose concentrations on the
outer and inner surfaces of the sarcolemma, however, are impossible to
obtain directly. Although total tissue glucose can be measured
accurately in biopsied or excised muscle, it is impossible to translate
these measurements into a number representing the TSGG. This is because
it is very difficult to distinguish interstitial from intracellular
glucose. Furthermore, even if one were able to make the distinction
between interstitial and intracellular glucose directly, spatial and
physical barriers within extracellular and intracellular compartments
make the determination of glucose at the membrane surfaces impossible. Because the TSGG cannot be directly determined, important questions regarding control of glucose uptake remain unanswered.
The aim of these studies was to define the roles of glucose transport
and the TSGG in the regulation of basal and insulin-stimulated glucose
uptake in vivo. For these purposes, a novel method was used for
simultaneously assessing indexes of the TSGG and muscle glucose
metabolism. This method applies infusions of isotopic glucose analogs
[3-O-[3H]methylglucose
(3-O-[3H]MG),
[U-14C]mannitol
([U-14C]MN), and
2-deoxy-[3H]glucose
([2-3H]DG)] and
principles of glucose countertransport in a chronically catheterized,
awake rat model. Changes in the TSGG can be obtained because the ratio
of intracellular to extracellular
3-O-[3H]MG
concentration at equilibrium is a functional consequence of the TSGG
(6, 12, 41, 42). In addition, an index of muscle glucose metabolism is
calculated from muscle phosphorylated [2-3H]DG
([2-3H]DGP) and plasma
[2-3H]DG. These
measurements can distinguish the roles of glucose transport and the
TSGG in control of muscle glucose uptake.
 |
MATERIALS AND METHODS |
Animal maintenance and surgical procedures.
Male Sprague-Dawley rats (Sasco, Omaha, NE) were individually housed at
23°C on a 0600-1800 light cycle and allowed free access to
water and a predetermined weight of chow (65% carbohydrate, 11% fat,
24% protein). The rats were housed under these conditions for ~1 wk,
by which time their weights had reached 250-300 g. During this
period, the food conversion index (FCI; ratio of weight gained to food
consumed) for each rat was calculated. Animals were anesthetized with a
50:5:1 mixture of ketamine (Avoco, Fort Dodge, IA), Rompun (Avoco), and
acepromazine (Haver, Shawnee, KS). The left common carotid artery and
the right jugular vein were catheterized (PE50, Clay Adams, Parsippany,
NJ). Catheters were tunneled under the skin, exteriorized, secured at
the back of the neck, filled (~60 µl) with a 3:1 mix of glycerol
and heparin, and flame sealed. Immediately postsurgery, each animal
received 40,000 U of penicillin G (Marsam, Cherry Hills, NJ) and 5 ml
of sterile saline subcutaneously. All procedures were preapproved by
the Vanderbilt University Animal Care Committee and followed the
National Research Council Guide for the Care and Use
of Laboratory Animals. In the postsurgery period,
animal weights and food intake were monitored daily and only animals in
which presurgery weight and FCI were restored were used for
experiments. On the day of the experiment the catheters were aspirated,
flushed with a saline-heparin solution, and connected to silcone rubber
tubing for sampling.
Experimental procedures.
Food was taken away from rats ~5 h before the beginning of a study.
At t =
100 min an infusion of
saline (n = 6) or insulin (Novo
Nordisk, Princeton, NJ) at a rate of 0.6 (n = 5), 1.0 (n = 7), or 4.0 (n = 7)
mU · kg
1 · min
1
was begun. Also at t =
100 min,
primed infusions of
[U-14C]MN (3.5 µCi
primer and 60 nCi/min infusion) and
3-O-[3H]MG
(25 µCi primer and 150 nCi/min infusion) were begun.
3-O-[3H]MG
is transported in the cell but is not further metabolized, whereas
[U-14C]MN is
restricted to the extracellular space and thus serves as a marker of
the extracellular space. In another protocol, rats were studied for a
longer duration (340 min) to assess muscle isotope equilibration (i.e.,
achievement of a steady state). Beginning at
t =
300 min,
3-O-[3H]MG
and [U-14C]MN were
primed and infused as described above. At
t = 0 min a constant-rate infusion of
[2-3H]DG (900 nCi/min)
was started and continued to the end of the study.
[2-3H]DG is
transported into the cell and phosphorylated.
Plasma glucose was maintained at ~8 mM during insulin infusions by
use of a variable glucose infusion (Abbott Laboratories, Chicago, IL)
based on feedback from frequent arterial samples. Arterial blood
samples (volumes ranging between 150 and 500 µl) for tracer or
insulin analyses were taken at t =
100,
70,
40,
10, 1, 2.5, 5, 7.5, 10, 15, 20, 25, 30, and 40 min. Whole blood from a donor rat and washed red
blood cells from the experimental animal were used to maintain
hematocrit, but neither blood cells nor whole blood was given back
after t =
10 min. At
t = 40 min the animal was killed by
decapitation, and the soleus, white superficial vastus, and
gastrocnemius-plantaris muscles were excised, frozen in liquid
N2, and stored at
70°C
for further analyses. Soleus and white superficial vastus are oxidative
and nonoxidative muscle, respectively. Gastrocnemius-plantaris is a mix
of both fiber types. The muscles were excised in <1 min.
Processing of blood and muscle samples.
Plasma glucose concentrations were measured by the glucose oxidase
method using an automated glucose analyzer (Beckman Instruments, Fullerton, CA), and immunoreactive insulin was measured using a double
antibody method (40). Total plasma radioactivity
([U-14C]MN,
3-O-[3H]MG,
[2-3H]DG) was
determined after deproteinization with barium hydroxide [Ba(OH)2, 0.3 N] and
zinc sulfate (ZnSO4, 0.3 N) and
centrifugation. Radioactivity was determined in 10 ml of Ecolite+
scintillation fluid (ICN, Irvine, CA) by dual-labeled liquid
scintillation counting (Beckman LS 5000TD, Beckman Instruments). To
distinguish plasma 3-O-[3H]MG
and [2-3H]DG
radioactivity, plasma samples were treated with
Ba(OH)2 and ZnSO4, incubated with a solution
containing (as final concentrations) 2.5 mg/ml yeast hexokinase (21 U/mg solid; Sigma, St. Louis, MO), 100 mM KCl, 40 mM
tris(hydroxymethyl)aminomethane-Cl, 20 mM
MgCl2, and 4 mM EDTA (pH 8.1),
incubated at room temperature for 30 min, and then retreated with
Ba(OH)2 and
ZnSO4. Tests in our laboratory have shown that the yeast hexokinase phosphorylates >99% of
[2-3H]DG to
[2-3H]DGP, and >98%
of the [2-3H]DGP is
removed by the final Ba(OH)2 and
ZnSO4 treatment. Muscle samples
were homogenized in 0.5% perchloric acid (PCA), centrifuged, and
neutralized with 10 N KOH. One aliquot of homogenate was counted without further treatment, as described for plasma samples, to yield
total muscle counts
([U-14C]MN,
3-O-[3H]MG,
[2-3H]DG,
[2-3H]DGP, and
[2-3H]DGP in
glycogen). A second aliquot of homogenate was treated with
Ba(OH)2 and
ZnSO4 to remove free
[2-3H]DGP and
[2-3H]DGP incorporated
into glycogen and then counted to yield
[U-14C]MN,
3-O-[3H]MG,
and [2-3H]DG
radioactivity. A third aliquot of homogenate was incubated with yeast
hexokinase (as described for plasma samples), treated with
Ba(OH)2 and
ZnSO4 to remove
[2-3H]DGP, and then
counted to give
[U-14C]MN and
3-O-[3H]MG
radioactivity. Because
[U-14C]MN is
unaffected by analytical methods [i.e., hexokinase treatment and
Ba(OH)2 and
ZnSO4], radioactivity in the
14C counting window in treated
samples was normalized to its radioactivity in untreated samples
to provide an internal control for each plasma and muscle sample.
Tissue glucose was measured after deproteinization with PCA by an
enzymatic method (33).
Calculations.
The distribution of mannitol between tissue and extracellular spaces
was used to calculate the fraction of extracellular to total water
space in biopsies by the equation
|
(1)
|
where
Fe is the fraction of the tissue
water that is extracellular and the subscripts t and e refer to tissue
and extracellular compartments.
[U-14C]MNe
concentrations were assumed to equal plasma
[U-14C]MN, since
mannitol is not extracted by muscle. Intracellular and extracellular
water spaces were used to calculate intracellular substrate
concentrations.
An index of skeletal muscle glucose metabolism
(Rg) was calculated during the
40-min [2-3H]DG
infusion (0-40 min) from the concentration of
[2-3H]DGP in
intracellular water and the integral of the plasma
[2-3H]DG concentration
for the infusion period. The relationship is defined as
|
(2)
|
where
[[2-3H]DGP]
is the concentration of
[2-3H]DG
that is phosphorylated (either remaining as
[2-3H]DGP or
incorporated into glycogen), [G] is glucose concentration, the subscript e refers to extracellular (as assessed in arterial plasma), and t = 40 min. The
application of 2-DG to measurement of muscle glucose metabolism has
been described in detail previously (14, 30). It should be noted that
most previous studies that used this technique underestimated muscle
glucose metabolism because the
[2-3H]DG that was
incorporated into glycogen was not considered (5, 60). As indicated
above, the analytical technique used in the present studies measures
both free [2-3H]DGP
and [2-3H]DGP in
glycogen.
Morgan et al. (42) defined countertransport as a difference in the
steady-state distribution of one sugar between intracellular and
extracellular water induced by a transmembrane gradient of a second
sugar. With this technique, the distribution of trace 3-O-[3H]MG
between the intracellular and extracellular water space is determined
at steady state to assess the transmembrane gradient of glucose. The
ratio of
3-O-[3H]MG
inside to outside the cell, defined as
Si/So,
is calculated by the following equation
|
(3)
|
The distribution of
3-O-[3H]MG
inside and outside the cell is determined by the rate constants for
entry and exit from the cell. Because, at equilibrium,
3-O-[3H]MG
movement into the cell is equal to
3-O-[3H]MG
out of the cell, the following relationships exist
|
(4)
|
and
|
(5)
|
where
kin and
kout are the rate
constants for
3-O-[3H]MG
influx and efflux, respectively, and
Si and
So refer to the concentrations of
3-O-[3H]MG
inside and outside the cell. This method relies not on direct measurements of glucose but on the functional consequence of local increases in glucose at the outer and inner cell membrane surfaces. The
distribution of
3-O-[3H]MG
across the plasma membrane will be determined by the availability of
membrane glucose transporters. Competition between glucose and
3-O-[3H]MG
for the transport system they share will determine the ratio of
apparent rate constants for inward and outward transport of 3-O-[3H]MG.
This ratio can be calculated from the ratio of
3-O-[3H]MG
inside to outside the cell at a steady state. When intracellular glucose concentrations approach those of extracellular glucose, competition for the inside face of the glucose transporter is increased, so the ratio of
3-O-[3H]MG
inside to outside the cell approaches one. The advantage of measuring
3-O-[3H]MG
is that, because
3-O-[3H]MG
is not metabolized, extracellular gradients and intracellular gradients
of this analog will not exist at steady state when this sugar is
infused at a constant rate. As a result, interstitial 3-O-[3H]MG
concentrations equal plasma measurements, and intracellular 3-O-[3H]MG
concentrations are the same throughout the contiguous intracellular water space.
Si/So
is then related to the glucose concentrations on the inner
([G]im) and outer
([G]om) surfaces of
the sarcolemma by the following equation
|
(6)
|
where
Km is the
Michaelis-Menten constant for glucose transport across the sarcolemma.
The Km for GLUT-4
ranges from 2 to 5 mM (21, 47). A value for
Km of 4 mM was used in these
studies, since it more closely reflects estimates obtained from muscle venous drainage in vivo (10, 63, 67). From a qualitative standpoint,
the calculation of changes in TSGG is independent of the absolute value
of Km. Moreover,
it has been shown repeatedly that
Km is unchanged
by insulin stimulation in a variety of experimental settings
(20-22, 38, 43, 45, 47, 56, 61). Although [G]im and
[G]om cannot be
measured directly, the measurements of
Si/So
and tissue glucose allow the calculation of limits for these variables.
The highest possible concentration of
[G]om is the value
obtained if one assumes that the glucose mass is confined to the
extracellular space. Although
[G]im has a finite
concentration in this scenario, it occupies a fraction of the
intracellular space that is so small that it does not contribute
significantly to the muscle glucose mass. The
[G]om calculated
assuming that [G]im
contributes negligibly to total muscle glucose is determined by the
relationship
|
(7)
|
where
the superscript
reflects local membrane concentrations when
intracellular glucose concentration is negligible, and [G]m is
[G]/µl muscle H2O.
[G]om is then
calculated assuming the opposite extreme;
[G]im is present
uniformly in the entire intracellular H2O space. This variable is
calculated as
|
(8)
|
where
the superscript
reflects local membrane concentrations when
intracellular glucose concentration is uniformly equal to
[G]im in the entire
intracellular volume. The solution to
Eq. 8
can be substituted into Eq.
6, which can then be solved for [G]im
.
TSGG
and
TSGG
can then be solved using
either
[G]im
and
[G]om
or
[G]im
and
[G]om
,
respectively
|
(9)
|
Glucose countertransport has been used to assess effects of glucose
and insulin on intracellular glucose in a variety of tissue types, and
the use of 3-O-MG to measure glucose
countertransport has been described in detail previously (6, 8, 12, 16, 38, 41, 42, 52, 61).
Statistical analyses.
Statistical comparisons were made between the basal group and each
insulin-infused group with Student's
t-test using Statview (Abacus,
Berkeley, CA) and a Macintosh computer. Differences were considered
statistically significant at P < 0.05. Data are expressed as means ± SE. Statistical differences are
presented in the legends to Figs. 1-6 and Tables 1 and 2.
 |
RESULTS |
Arterial plasma insulin, arterial plasma glucose, glucose infusion
rates, and arterial plasma
3-O-[3H]MG.
Plasma insulin concentrations (Table 1)
~1.2-, 1.8-, and 4.9-fold above values obtained during saline
infusion were achieved with 0.6, 1.0, and 4.0 mU · kg
1 · min
1
insulin infusion rates, respectively. Arterial insulin concentrations at the two higher insulin doses were significantly greater than values
obtained during saline infusion alone. There was no significant difference in plasma glucose among any of the groups (Table 1). The
glucose infusion rate required to maintain euglycemia was increased in
an insulin dose-dependent manner (Table 1).
Infusion of
3-O-[3H]MG
resulted in steady-state plasma concentrations
(disintegration · min
1 · µl
plasma
1) by
t = 0 with all insulin infusions (Fig.
1). In the absence of an insulin infusion,
arterial
3-O-[3H]MG
had not yet plateaued by t = 0 and was
still rising during the
[2-3H]DG infusion
period. Even so,
3-O-[3H]MG
concentrations during the last 20 min deviated by <10% from the mean
of that same interval.

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Fig. 1.
Arterial plasma
3-O-[3H]methylglucose
(3-O-[3H]MG)
concentrations were determined at insulin infusion rates of 0.0 (n = 6), 0.6 (n = 5), 1.0 (n = 7), and 4.0 (n = 7)
mU · kg 1 · min 1.
Data represent means ± SE.
3-O-[3H]MG
concentrations were significantly decreased by all insulin infusion
rates in relation to saline (P < 0.05); dpm, disintegrations/min.
|
|
Skeletal muscle glucose concentrations.
Skeletal muscle glucose concentration, which is the sum of tissue
glucose inside and outside the cell, fell insignificantly compared with
basal at the low insulin dose (0.6 mU · kg
1 · min
1)
in the soleus. Soleus glucose concentration was equal to basal at the
two higher insulin doses (Table 2). In
contrast, glucose concentration fell by 35-45% in muscles that
have a high composition of fast-twitch fibers (vastus lateralis and
gastrocnemius). Because the change in muscle glucose will reflect the
balance between glucose utilization and glucose delivery, these data
suggest that the muscles comprised of fast-twitch fibers have a greater
deficit in muscle glucose delivery compared with their ability to use glucose.
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Table 2.
Glucose concentrations (mmol/kg wet wt) in gastrocnemius, vastus
lateralis, and soleus during insulin infusion
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|
Skeletal muscle extracellular water space.
The fraction of H2O that was extracellular in the soleus
was 0.33 ± 0.02, 0.35 ± 0.07, 0.41 ± 0.02, and 0.38 ± 0.02 at insulin infusions of 0.0, 0.6, 1.0, and 4.0 mU · kg
1 · min
1,
respectively. The fraction of tissue
H2O that is extracellular was
increased significantly above basal at the 1.0 mU · kg
1 · min
1
insulin infusion rate (P < 0.05).
Skeletal muscle
Si/So.
Si/So
in the soleus was 0.45 ± 0.10 after a 140-min infusion of isotopes
alone. Extending the infusion of
[U-14C]MN and
3-O-[3H]MG
by 200 min did not lead to a further increase in
Si/So
(0.53 ± 0.03), indicating that
3-O-[3H]MG
had reached equilibrium in the soleus. A clear steady-state Si/So
was not apparent in other muscles, and ratios are not presented. The
Si/So
response to increasing insulin in the soleus is shown in Fig.
2.
Si/So
was increased above values seen with saline infusion at the highest
insulin dose (P < 0.005).

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Fig. 2.
Ratio of intracellular to extracellular
3-O-[3H]MG
concentration
(Si/So)
was calculated in soleus at insulin infusion rates of 0.0 (n = 6), 0.6 (n = 5), 1.0 (n = 7), and 4.0 (n = 7)
mU · kg 1 · min 1.
Data represent means ± SE.
Si/So
was increased at highest insulin dose in soleus
(P < 0.005).
|
|
Skeletal muscle [G]om,
[G]im, and TSGG.
[G]om,
[G]im, and TSGG were
calculated under two extreme theoretical conditions, which are defined
in Calculations. In the first
condition, [G]im is
contained in such a small fraction of the intracellular
H2O that, regardless of whether
[G]im is relatively high or low, it contributes negligibly to the tissue glucose mass (the
symbol
represents terms calculated in this condition). In the
second condition, the opposite is assumed. Glucose was assumed to be
distributed evenly throughout intracellular
H2O (designated with the symbol
). Within the bounds of these extremes lay the true values of
[G]om,
[G]im, and TSGG.
[G]om
was virtually unchanged with increasing insulin.
[G]om
decreased gradually with increasing insulin concentration (Fig. 3).
[G]im
was not significantly different from zero under basal conditions and at
insulin doses of 0.6 and 1.0 mU · kg
1 · min
1
but rose to 2.4 ± 0.6 at an insulin dose of 4.0 mU · kg
1 · min
1
(P < 0.05; Fig. 3).
[G]im
was also not significantly different from zero in the basal state and
at insulin infusion rates of 0.6 and 1.0 mU · kg
1 · min
1.
At the higher insulin dose, however, it rose significantly, reaching
0.9 ± 0.2 mM at an insulin infusion of 4.0 mU · kg
1 · min
1
(P < 0.005).

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Fig. 3.
Outer sarcolemmal glucose concentrations
([G]om and
[G]om )
(top) and inner sarcolemmal glucose
concentrations ([G]im and
[G]im )
(bottom) were calculated in soleus
at insulin infusion rates of 0.0 (n = 6), 0.6 (n = 5), 1.0 (n = 7), and 4.0 (n = 7)
mU · kg 1 · min 1.
Data represent range bounded by means ± SE for
[G]om
and
[G]om
and
[G]im
and
[G]im .
[G]om
decreased gradually with increasing insulin concentration.
[G]im
and
[G]im
were not significantly different from a glucose concentration of 0 at
basal insulin and at insulin doses of 0.6 and 1.0 mU · kg 1 · min 1
but rose significantly at the insulin dose of 4.0 mU · kg 1 · min 1
(P < 0.05-0.005).
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|
TSGG fell gradually with increasing plasma insulin (Fig.
4). This response was independent of
whether [G]im and
[G]om were calculated
assuming that glucose was distributed in a negligible volume or the
entire intracellular H2O space.
TSGG
fell from 4.9 ± 0.9 mM
in the basal state to 4.7 ± 0.3, 2.9 ± 0.9 and 0.7 ± 0.7 mM
at insulin doses of 0.6, 1.0, and 4.0 mU · kg
1 · min
1,
respectively. TSGG
was
significantly reduced compared with basal at the 4.0 mU · kg
1 · min
1
insulin infusion (P < 0.02) and was
not significantly different from zero.
[G]om
was 5.4 ± 1.2 mM in the basal state and fell to 4.8 ± 0.6, 2.6 ± 0.8, and 0.6 ± 0.5 mM at insulin doses of 0.6, 1.0, and 4.0 mU · kg
1 · min
1.
TSGG
was significantly reduced
compared with basal at insulin infusions of 1.0 and 4.0 mU · kg
1 · min
1
(P < 0.05-0.01). At the highest
insulin dose, TSGG
was not
significantly different from zero.

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Fig. 4.
Transsarcolemmal glucose gradients
(TSGG and
TSGG ) were calculated in soleus
at insulin infusion rates of 0.0 (n = 6), 0.6 (n = 5), 1.0 (n = 7), and 4.0 (n = 7)
mU · kg 1 · min 1.
Data represent range bounded by means ± SE for
TSGG and
TSGG .
TSGG was significantly reduced
compared with basal at 4.0 mU · kg 1 · min 1
insulin infusion (P < 0.02).
TSGG was reduced compared with
basal at insulin infusions of 1.0 and 4.0 mU · kg 1 · min 1
(P < 0.05-0.01).
TSGG and
TSGG were not significantly
different from 0 at highest insulin dose.
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|
Skeletal muscle Rg.
The Rg response in the soleus is
shown in Fig. 5.
Rg was 13.3 ± 3.0, 36.3 ± 9.2, 31.8 ± 4.2, and 35.1 ± 2.1 µmol · 100 g
1 · min
1
at insulin infusions of 0.0, 0.6, 1.0, and 4.0 mU · kg
1 · min
1,
respectively. The increment in soleus
Rg at the 0.6 mU · kg
1 · min
1
insulin infusion occurred without a decrease in the TSGG. Comparison of
Figs. 4 and 5 shows that the TSGG narrowed at the higher insulin infusion rates even after Rg had
reached a maximum. These data indicate that glucose transport is not
maximal and that those factors responsible for sustaining the glucose
gradient (glucose delivery to muscle and/or glucose
phosphorylation in muscle) have become rate limiting. The increase in
[G]im at the highest
insulin dose suggests that glucose phosphorylation is one of those
factors.

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Fig. 5.
Glucose metabolic index (Rg) in
soleus was determined at insulin infusion rates of 0.0 (n = 6), 0.6 (n = 5), 1.0 (n = 7), and 4.0 (n = 7)
mU · kg 1 · min 1.
Data represent means ± SE. An insulin infusion increased
Rg significantly above values seen
with saline infusion alone (P < 0.05-0.001).
|
|
 |
DISCUSSION |
A great number of studies have assessed the means by which insulin
stimulates glucose transport, and much has been learned. Far fewer
studies have assessed the means by which the downhill glucose gradient
from outside to inside the muscle cell is maintained in the face of the
marked increase in glucose transport that occurs during insulin
stimulation. The results of this study show that a small physiological
increase in insulin can stimulate muscle Rg by approximately threefold
without significantly affecting the TSGG in vivo. This suggests that
mechanisms of glucose delivery to soleus and glucose phosphorylation
within the soleus keep pace with the increase in glucose transport at
this concentration of insulin. If the increase in insulin-stimulated
glucose transport is uncompensated for by parallel increases in muscle
glucose delivery and/or intracellular glucose phosphorylation,
the TSGG will be reduced. This is, in fact, what occurred at higher
insulin concentrations. Stimulation of glucose transport exceeded the
stimulation of mechanisms that maintain the TSGG, and the TSGG fell
from 5 mM in the basal state to <1 mM at an insulin infusion rate of
4.0 mU · kg
1 · min
1.
The fact that Rg is sustained
despite a progressive fall in TSGG indicates that glucose transport
continues to increase even after
Rg has achieved a maximum. The
TSGG at the high insulin infusion rate was not significantly different
from zero (i.e., no gradient existed), suggesting that TSGG, and not
glucose transport, had become rate limiting for muscle glucose uptake.
The decrease in TSGG is due, at least in part, to a limitation in
glucose phosphorylation. Even our most conservative estimate showed an
increase in [G]im at
the highest insulin concentration, suggesting that glucose
phosphorylation is insufficient to maintain TSGG. This finding is
consistent with the conclusions of several other studies that used
different approaches (11, 15, 31, 65). It is less clear whether glucose
availability to the outer surface of the muscle is also a limitation.
[G]om does not fall when it is calculated assuming
[G]im occupies a
negligible volume, but falls to one-third of the basal value when it is
calculated assuming that
[G]im exists
throughout the entire intracellular space. It is probable that a
deficit in glucose availability is more important in fast-twitch
muscles, since the vastus lateralis and gastrocnemius both exhibit
significant decreases in muscle glucose concentration in response to
insulin infusions. The question of whether insulin may play a role in
increasing the delivery of glucose to skeletal muscle is controversial.
Although some investigators have reported an insulin-induced increase
in limb blood flow (for review, see Ref. 1) and capillary perfusion (50), others have not observed these effects (e.g., 27, 44). This issue
is further complicated by the results of one study in which insulin
increased muscle blood flow but the areas of the muscle that received
the increased flow were distinct from the areas in which the largest
increases in glucose uptake occurred (49). Whether or not insulin has
hemodynamic effects, the fall in total muscle glucose in type II
muscles is evidence that the rate of glucose entry into the muscle as a
whole must be less than the rate of glucose metabolism.
Three markedly different modeling approaches have been conducted using
data from rats (15, 64) or humans (4). Each of these shows a greater
insulin stimulation of 3-O-MG
transport into the cell compared with transport out of the cell. This
is exactly what is predicted from the increase in
Si/So
in the present study. These models all give results consistent with the
present study in that they predict a greater antagonism of
3-O-MG transport out of the cell in
the presence of hyperinsulinemia in vivo. The principle of glucose
countertransport has been used to estimate intracellular glucose in
studies conducted in diverse model systems (6, 8, 12, 16, 38, 41, 42,
52). The linking of the transmembrane glucose distribution to
3-O-MG countertransport assumes that
1) glucose and
3-O-MG share the same transport
system, 2) the reaction between
carrier and sugar is rapid compared with carrier mobility,
3) the relative affinity of each
sugar for the transport proteins is the same on the extracellular and
intracellular sides of the plasma membrane, and
4) carrier mobility is independent of whether or not the transporter is bound to either sugar. These assumptions have been discussed in detail by Foley et al. (12). The
validity of the first two assumptions has been repeatedly demonstrated
(6, 7, 17, 18, 52, 59, 62). The third assumption is supported by the
demonstration that the kinetic parameters for
3-O-MG are equal for entry and exit
into adipocytes (56, 61), which use the same glucose transport proteins
as muscle. Morever, this symmetry is independent of the method used to
assess transport kinetics and whether insulin is present or absent
(56). The fourth assumption is consistent with the lack of any marked
asymmetry of the transport system in adipocytes, even when
intracellular glucose and consequently the bound state of the inner
aspect of the glucose transporter are less than the extracellular (18,
56).
Calculation of [G]om,
[G]im, and TSGG
requires knowledge of the interstitial [G] and the
Km of glucose
transport. Boundaries for interstitial [G] were achieved by
using two extreme theoretical conditions. The first condition is one in
which [G]im is
confined to such a small portion of the intracellular
H2O that it contributes negligibly
to the total tissue glucose measured and all the glucose is confined to
the extracellular space (Eq.
7). The opposite case is one in
which glucose is dispersed uniformly in intracellular H2O so that
[G]im is the glucose
concentration of the entire intracellular H2O. The values for
[G]om that are
obtained with these approaches give a representative interstitial
glucose concentration for these conditions. At some cells, however,
this value may be too high and in others it may be too low. Regardless,
the relationship of
[G]om to
[G]im will be the
same. It is also difficult to precisely know
Km, since a range
of 2 to 5 mM for the glucose transporter has been reported (47). In the
calculations described above a
Km of 4 mM was
used, since it is within the range measured for glucose transport by
GLUT-4 in vitro and it is comparable to the venous glucose at which the
Km occurs in vivo
(10, 63, 67). The responses of
[G]im,
[G]om, and TSGG to
insulin are similar regardless of the
Km used to
calculate them. It is also apparent that the value used for
Km has a much
greater influence on our calculated values when
Si/So
is relatively small, as in the basal state (as shown in Fig.
6). At an
Si/So
of 1 (which is not different from the value in soleus at the highest
insulin infusion rate), Km does not
influence this calculation.

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|
Fig. 6.
Influence of Michaelis-Menten constant
(Km) on values of [G]om
(top),
[G]im
(middle), and TSGG
(bottom) calculated in soleus at
insulin infusion rates of 0.0 (n = 6),
0.6 (n = 5), 1.0 (n = 7), and 4.0 (n = 7)
mU · kg 1 · min 1.
Data are mean values. These data show that the same types of responses
are evident regardless of
Km used in
calculations.
|
|
The measurement of intracellular
3-O-MG concentration could also
conceivably be increased if its intracellular volume of distribution were expanded. One could speculate that insulin could increase the
volume of 3-O-MG distribution by
either 1) accelerating exchange with
a pool that equilibrates slowly under basal conditions or 2) increasing permeability of an
intracellular physical barrier. The first postulate is unlikely to
explain the increase in intramuscular 3-O-MG, since
3-O-MG is already in equilibrium with
the contiguous cell water. It is difficult to envision the second,
since insulin has not been shown to increase the permeability of
intracellular organelles to glucose. Earlier studies have demonstrated
that insulin stimulation does not affect the intracellular volume of 3-O-MG distribution in adipocytes (18,
59) or the tissue distribution volume of
3-O-MG in skeletal muscle (42).
Studies in isolated cells (38) and tissues (6, 42) clearly show that
the insulin-induced increase in
3-O-[3H]MG
inside vs. outside the cell is secondary to effects of insulin on
transmembrane glucose distribution and are not due to a primary increase in the volume of the intracellular glucose compartment. The
reliance of
Si/So
on transmembrane glucose distribution is exemplified by the
demonstration that, if glucose is removed entirely from the incubation
medium, the rate constants for 3-O-MG
transport in and out are equivalent regardless of the insulin
concentration. Conversely, rat skeletal muscle
[3-O-MG] is reduced by
hyperglycemia even in the presence of high serum insulin concentrations
(42), as would be predicted by Eq.
6.
It is impossible to get an accurate value for intracellular glucose
from direct glucose measurements in excised and biopsied tissue for
several reasons. Estimates of intracellular glucose concentration
require that interstitial glucose in tissue samples is known. Blood
glucose concentrations have been used in place of interstitial glucose
(66). This leads to spurious numbers, since estimates of skeletal
muscle interstitial glucose concentration made using microdialysis show
that it is well below arterial or venous plasma concentrations (34).
Because 3-O-MG is not metabolized, its
plasma concentration equals its interstitial concentration and can be
used to calculate intracellular
[3-O-MG] from measurements in tissue samples. Even if the intracellular glucose concentration could be determined, it may not be the most sensitive determinant of
the glucose diffusion gradient in the cell, since physical barriers and
spatial glucose gradients may compartmentalize glucose within the cell.
In this regard, there is evidence that the glucose transporters (9, 13,
19, 36, 37, 39) and hexokinases (2, 26, 32, 58) are localized to
specific regions within the skeletal muscle cell. Glucose concentration
gradients may exist within the intracellular space so that
concentrations are diminished from regions of high glucose transport to
regions of high glucose phosphorylating activity. Because intracellular
3-O-[3H]MG
is not metabolized, it can be measured and, at equilibrium, is
homogenous within the contiguous intracellular space.
Rg increases in soleus without an
increase in
Si/So
and [G]im at an
insulin dose of 0.6 mU · kg
1 · min
1,
suggesting that hexokinase can accommodate the increase in
insulin-stimulated glucose transport at this insulin concentration. It
is also possible that insulin increases the hexokinase activity. This
is consistent with the increase in the rate constant for glucose
phosphorylation that was predicted by compartmental analysis (54) and
studies using 18F-2-deoxyglucose
(28). The basis for this change may relate to an increase in the
fraction of hexokinase II that is bound to mitochondria and thus in its
more active form (3). One study showed, however, that there is no
change in the fraction of bound hexokinase II in human muscle during
hyperinsulinemia (25). Although it is also possible that more
hexokinase II is synthesized during the hyperinsulinemic clamp (35,
48), the time needed for this to occur in skeletal muscle is >6 h. An
increased ability to phosphorylate glucose may also result from the
reduction in glucose 6-phosphate that has been shown to occur in rat
skeletal muscle at about the same insulin concentration as was obtained with the 0.6 mU · kg
1 · min
1
infusion (53). At higher insulin concentrations, glucose 6-phosphate begins to return to basal concentrations (53). In the present study,
this correlated to the increase in
[G]im.
Basal and insulin-stimulated Rg
are higher in oxidative than nonoxidative muscle (30). Hexokinase II
(46, 58) and GLUT-4 (23, 29) are both more abundant, and oxidative
fibers are better perfused than nonoxidative fibers of the rat. In the
present study, insulin was shown to have different effects on the
tissue glucose concentration of slow- and fast-twitch muscles. Soleus glucose concentration fell insignificantly and only transiently with
increasing insulin concentrations. In contrast, vastus lateralis and
gastrocnemius glucose concentrations both fell significantly (Table 2).
Results obtained in human skeletal muscle, which contains both fast- and slow-twitch muscles, support the findings of the present study by showing that glucose concentration either falls slightly or stays the same during a hyperinsulinemic, euglycemic clamp
(24, 51, 55, 57). The rat model is advantageous in that certain muscles
are homogeneous for fiber type, permitting fiber type-specific effects
to be assessed. Just as in the present study, the glucose concentration
in a rat muscle that is predominantly fast twitch (rectus abdominus)
was shown to decrease in the presence of hyperinsulinemia (66). The
greater decrease in muscle glucose content in fast-twitch muscle
suggests that glucose availability is a more serious limitation in
these tissues and is more likely to compromise insulin-stimulated
glucose uptake. This deficit is probably due to the lower blood flow
and greater diffusion distances in these tissues.
Skeletal muscle comprises ~50% of total body mass and exhibits the
greatest increases in glucose uptake in response to insulin and
exercise. A knowledge of the regulation of skeletal muscle glucose
uptake therefore is a prerequisite to understanding normal and
pathophysiological whole body glucose uptake. These studies provide
insight into the control of glucose uptake by showing that
1) increases in
Rg can occur in response to
insulin levels in the physiological range without decreasing TSGG,
suggesting that muscle glucose delivery and glucose phosphorylation are
adequate to accommodate the increased glucose transport flux;
2) insulin at high physiological or
supraphysiological levels leads to a decrease in TSGG, which suggests
that the increase in transport activity has exceeded glucose delivery
to the muscle or glucose phosphorylation within the muscle;
3) maximum
Rg is sustained even though TSGG
continues to fall, indicating that glucose transport still has the
capacity to increase and is not rate limiting for insulin-stimulated
glucose uptake; and 4) in contrast
to the soleus, which exhibits only a transient fall in muscle glucose,
muscle consisting of fast-twitch fibers that are poorly perfused
actually exhibits a 35-45% fall in tissue glucose, suggesting
that glucose delivery is a major limitation in sustaining the TSGG
during insulin stimulation in these tissues. In
conclusion, control of glucose uptake is distributed between glucose
transport and factors that determine the TSGG. The stimulation of
glucose transport that occurs with increasing insulin concentrations
places more importance on the factors that maintain glucose delivery to
the muscle membrane and glucose phosphorylation inside the muscle.
 |
ACKNOWLEDGEMENTS |
We are grateful to Drs. David Regen, James May, Richard Whitesell,
and Richard Printz for their insights in the preparation of the
manuscript.
 |
FOOTNOTES |
This work was supported by a grant from the American Diabetes
Association and National Institute of Diabetes and Digestive and Kidney
Diseases Grant RO1 DK-50277. A. Halseth was supported by Training Grant
5 T32 DK07563-08.
Address for reprint requests: D. H. Wasserman, Dept. of Molecular
Physiology and Biophysics, Vanderbilt Univ. School of Medicine,
Nashville, TN 37232.
Received 20 August 1997; accepted in final form 28 October 1997.
 |
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