Department of Molecular Physiology and Biophysics, and Diabetes Research and Training Center, Vanderbilt University, Nashville, Tennessee 37232-0615
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ABSTRACT |
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The aim of these studies was to determine
whether prior exercise enhances net hepatic glucose uptake (NHGU)
during a glucose load. Sampling catheters (carotid artery, portal,
hepatic, and iliac veins), infusion catheters (portal vein and vena
cava), and Doppler flow probes (portal vein, hepatic and iliac
arteries) were implanted. Exercise (150 min;
n = 6) or rest
(n = 6) was followed by a 30-min
control period and a 100-min experimental period (3.5 mg · kg1 · min
1
of glucose in portal vein and as needed in vena cava to clamp arterial
blood glucose at ~130 mg/dl). Somatostatin was infused, and insulin
and glucagon were replaced intraportally at fourfold basal and basal
rates, respectively. During experimental period the arterial-portal
venous (a-pv) glucose gradient (mg/dl) was
18 ± 1 in
sedentary and
19 ± 1 in exercised dogs. Arterial insulin and
glucagon were similar in the two groups. Net hepatic glucose balance
(mg · kg
1 · min
1)
shifted from 1.9 ± 0.2 in control period to
1.8 ± 0.2 (negative rates represent net uptake) during experimental period in
sedentary dogs (
3.7 ± 0.5); with prior exercise it shifted from
4.1 ± 0.3 (P < 0.01 vs.
sedentary) in control period to
3.2 ± 0.4 (P < 0.05 vs. sedentary) during
experimental period (
7.3 ± 0.7, P < 0.01 vs. sedentary). Net
hindlimb glucose uptake (mg/min) was 4 ± 1 in sedentary
animals in control period and 13 ± 2 during experimental period; in
exercised animals it was 7 ± 1 in control period
(P < 0.01 vs. sedentary) and 32 ± 4 (P < 0.01 vs. sedentary) during experimental period. As the total glucose infusion rate (mg · kg
1 · min
1)
was 7 ± 1 in sedentary and 11 ± 1 in exercised dogs, ~30% of the added glucose infusion due to prior exercise could be accounted for
by the greater NHGU. In conclusion, when determinants of hepatic glucose uptake (insulin, glucagon, a-pv glucose gradient, glycemia) are
controlled, prior exercise increases NHGU during a glucose load due to
an effect that is intrinsic to the liver. Increased glucose disposal in
the postexercise state is therefore due to an improved ability of both
liver and muscle to take up glucose.
carbohydrates; glucose balance; portal vein; exertion; dog
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INTRODUCTION |
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EXERCISE markedly increases the metabolic demands of the organism, mostly due to increased needs of the contracting muscle (17, 29). Net muscle glucose uptake is greatly enhanced during exercise and remains elevated in the postexercise state to replenish muscle glycogen stores that were depleted during muscle contraction. Increased insulin sensitivity and glucose effectiveness facilitate muscle glucose uptake and glycogen synthesis in the postexercise state (27-29). This process is markedly accelerated if exogenous glucose is administered after the cessation of exercise (15, 26). In fact, the excess whole body glucose uptake measured during glucose infusion after exercise has been ascribed solely to skeletal muscle glucose disposal (17, 29). The liver, on the other hand, even if its glycogen stores are also depleted by prior exercise, remains a net producer of glucose in the immediate postexercise state in fasted animals (32). The liver can become a net consumer of glucose if exogenous glucose is administered after exercise (8). Nevertheless, neither net hepatic glucose uptake (NHGU) nor net hepatic glycogen deposition appears to be different from that measured in the absence of prior exercise (8). In another study, Matsuhisa et al. (20) observed that prior contraction of the rabbit hindlimb caused an increase in the rate of liver deposition of the glucose analog 3-fluoro-3-deoxy-D-glucose (3FDG) during a glucose load. In the aforementioned studies, however, direct hepatic effects of prior exercise were difficult to ascertain, because one or more of the determinants of hepatic glucose uptake [glucose load, arterial-portal venous (a-pv) glucose gradient, pancreatic hormone concentrations] were uncontrolled.
The aim of the present study was to determine whether prior exercise enhances NHGU in response to controlled changes in arterial glucose, insulin, glucagon, hepatic glucose load, and a-pv gradient. To address this aim, isotopic and arteriovenous-difference techniques were used in the chronically catheterized, conscious dog model after either a prolonged treadmill exercise period or an equivalent period of rest.
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MATERIALS AND METHODS |
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Animals and surgical procedures. Twelve mongrel dogs of either gender (mean weight, 24 ± 2 kg) were studied. Animals were housed in a facility that met American Association for the Accreditation of Laboratory Animals Care guidelines and were fed a standard diet of meat and chow (34% protein, 14.5% fat, 46% carbohydrate, and 5.5% fiber based on dry weight). Experimental protocols were approved by the Vanderbilt University School of Medicine Animal Care and Use Committee. At least 16 days before each experiment, a laparotomy was performed under general anesthesia. Two Silastic catheters (0.03 mm ID) were inserted in the inferior vena cava for tracer and indocyanine green (ICG) infusion. Two more Silastic catheters (0.03 mm ID) were inserted in a jejunal and in a splenic vein (advanced so that the tips were in the portal vein) for intraportal infusion of glucose. Silastic catheters (0.04 mm ID) were also inserted in the portal vein (~2 cm downstream of the tips of the portal infusion catheters) and left common hepatic vein for blood sampling. Incisions were made in the neck region and the inguinal region for the insertion of Silastic sampling catheters (0.04 mm ID) in the carotid artery (advanced so that its tip rested in the aortic arch) and in a lateral circumflex vein (advanced so that its tip was in the common iliac vein). After insertion, the catheters were filled with saline containing heparin and their free ends were knotted.
Doppler flow probes (Transonic Systems, Ithaca, NY) were used to measure portal vein, hepatic artery, and external iliac artery blood flows. A small section of the portal vein, upstream from its junction with the gastroduodenal vein, was cleared of tissue, and a 6.0-mm-ID flow cuff was placed around the vessel and secured. The gastroduodenal vein was isolated and ligated proximal to its coalescence with the portal vein. A section of the main hepatic artery proximal to the portal vein was isolated, and a 3.0-mm-ID flow cuff was placed around the vessel and secured. The external iliac artery was accessed from the abdominal incision, dissected free of surrounding tissue, and fitted with a 4.0-mm-ID flow probe cuff, which was then secured around the vessel. The flow probe leads and the knotted free catheter ends, with the exception of the carotid artery and the common iliac vein catheters, were stored in subcutaneous pockets in the abdominal region so that complete closure of the skin incision was possible. The carotid artery and common iliac vein catheters were stored in subcutaneous pockets in the neck and inguinal regions, respectively.
Starting 1 wk after surgery, dogs were exercised on a motorized treadmill, so that they would be familiar with treadmill running, regardless of whether they were used for a sedentary or exercise experiment. Dogs were not exercised during the 48 h preceding an experiment. Only animals that had 1) a leukocyte count <18,000/mm3, 2) a hematocrit >36%, 3) normal stools, and 4) a good appetite (consuming all of the daily ration) were used.
Studies were conducted after a 42-h fast, because this induces a stable minimum in hepatic glycogen content in the dog (11), preventing any effect due to different liver glycogen concentrations between sedentary and exercised animals. On the day of the experiment, the subcutaneous ends of the catheters were freed through small skin incisions made under local anesthesia (2% lidocaine; Astra Pharmaceutical, Worcester, MA) in the abdominal, inguinal, and neck regions. The contents of each catheter were aspirated, and they were flushed with saline. Silastic tubing was connected to the exposed catheters and brought to the back of the dog, where they were secured with quick-drying glue. Saline was infused in the arterial catheters throughout experiments (0.1 ml/min).
Experimental procedures. Animals were
either exercised at a moderate intensity (100 m/min, 12% grade) on a
motorized treadmill (n = 6) or
remained sedentary (n = 6) from
t = 180 to
30 min (Fig.
1). The exercise duration and intensity
used in these experiments have been shown previously to result in a
twofold increase in heart rate and an increase in
O2 uptake to 50% of maximum (22). A period of exercise recovery or continued rest followed (
30 to
100 min). At time =
70 min, a primer of
[3-3H]glucose (30 µCi) was given, followed by venous infusions of [3-3H]glucose (0.3 µCi/min) and ICG (0.1 mg/min), which were continued for the duration
of the study. ICG was used as a backup for the Doppler method of flow
measurement. After a 10-min transition period
(t =
30 to
20 min), from
t =
20 to 0 min (basal period), three blood samples were drawn for assessment of basal levels of
metabolic variables. From t = 0-100 min (experimental period), glucose was given via a constant
intraportal infusion (3.5 mg · kg
1 · min
1)
and by a variable infusion into the inferior vena cava to clamp the
arterial blood glucose at 130 mg/dl. From
t = 0-100 min, endogenous pancreatic hormone secretion was also suppressed via a continuous somatostatin infusion into the inferior vena cava (0.8 µg · kg
1 · min
1).
Insulin and glucagon were replaced via intraportal infusions of 1.2 mU · kg
1 · min
1
(4-fold basal) and 0.5 ng · kg
1 · min
1
(basal), respectively. Arterial samples were drawn at 5-min intervals from t =
20 to 100 min. Portal,
hepatic, and common iliac venous samples were drawn at
t =
20,
10, 0, 60, 70, 80, 90, and 100 min. Portal vein, hepatic artery, and external iliac
artery blood flows were recorded continuously from the frequency shifts
of the pulse sound signal emitted from the Doppler flow probes (9, 10).
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Processing of blood samples. Plasma
and deproteinized blood samples that were not analyzed the day of the
study were stored at 70°C after the completion of the
experiment. Plasma glucose levels were determined during experiments by
the glucose oxidase method with a glucose analyzer (Beckman
Instruments, Fullerton, CA). For the determination of plasma
[3-3H]glucose
radioactivity, samples were deproteinized with barium hydroxide and
zinc sulfate, the supernatant was evaporated, and the residue was
dissolved in 1 ml of water and 10 ml
Ecolite+ (ICN Biomedicals, Irvine,
CA). Radioactivity was then determined by liquid scintillation counting
with a Beckman LS 5000TD counter. Whole blood (samples deproteinized by
1:3 dilution in 4% percloric acid) lactate, glycerol, alanine,
glucose, and plasma free fatty acids were measured by enzymatic methods
(18) on a Technicon autoanalyzer (Tarrytown, NY) or on a Monarch 2000 centrifugal analyzer (Instrumentation Laboratories, Lexington, MA).
Immunoreactive insulin was measured with a double antibody system [interassay coefficient of variation (CV) of 10%; Ref. 21]. Immunoreactive glucagon was measured in plasma samples containing 50 µl of 500 Kallikrein-inhibitor units/ml Trasylol (FBA Pharmaceuticals, NY) with a double antibody system (CV of 7%) modified from the method developed by Morgan and Lazarow (21) for insulin. Insulin and glucagon antisera, standard glucagon and insulin, and the 125I-glucagon and 125I-insulin were obtained from Linco Research (St. Charles, MO).
Calculations. The tracer-determined total rate of glucose appearance (Ra) was determined by steady-state equations for isotope ([3-3H]glucose) dilution (3). Endogenous glucose Ra was calculated by subtracting the glucose infusion rate (portal vein + vena cava) from the total glucose Ra.
Net hepatic balances of lactate, glucose, alanine, FFA, and glycerol
were determined by the following formula: HAF × ([H] [A]) + PVF × ([H]
[P]), where [A], [P], and
[H] are the arterial, portal vein, and hepatic vein blood
or plasma substrate concentrations, respectively, and HAF and PVF are
the hepatic artery and portal vein blood or plasma flows, respectively,
determined with Doppler flow probes. The load of a substrate reaching
the liver was calculated as follows: [P] × PVF + [A] × HAF. Net hepatic substrate fractional extraction was calculated as the ratio of net hepatic balance to
hepatic load.
Net limb balances were calculated as follows: LF × ([A] [I]). LF is limb blood flow
through the external iliac artery, and [I] is the substrate
level in the common iliac vein. Limb fractional substrate extraction
was calculated as the limb substrate uptake divided by the limb
substrate load (LF × [A]). Blood levels and flows
were used for the calculation of all hepatic and limb balances, with
the exception of FFA balances for which plasma concentrations and flows
were used. The ratio of blood to plasma glucose was calculated for the
basal period and the glucose infusion period for each dog at each of
the four sampling sites. Plasma glucose values were then multiplied by
their corresponding ratio (i.e., blood glucose to plasma glucose) to
convert to blood glucose concentrations. The advantage of plasma
glucose measurements is that a large number of replicates can be
obtained quickly and at little added cost. The ability to measure
replicate samples reduces the measurement CV. The conversion to blood
values alleviates the need for assumptions regarding the equilibration
of substrates between red cell and plasma water.
When glucose is infused in the slow, laminar flow of the portal vein, mixing of the glucose in the blood can be problematic. To assess whether at a given time point t during intraportal glucose infusion good mixing of glucose was present, we used the following equation
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Data are expressed as mean ± SE. Data in some instances are
expressed as the mean of three measurements for the basal period (20 to 0 min) and of five measurements for the glucose
steady-state portion of the experimental period (60-100 min).
Statistics were performed with SuperAnova (Abacus Concepts, Berkeley,
CA) on a Macintosh PowerPC. Statistical comparison between groups and
over time was made with ANOVA designed to account for repeated
measures. Specific time points were examined for significance with
contrasts solved by univariate repeated measures. Pooled data from
basal and glucose infusion periods were compared with unpaired
t-tests. Statistics are reported in
the corresponding table or figure legend for each variable. Differences
were considered significant when P
values were <0.05.
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RESULTS |
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Arterial blood glucose, a-pv glucose gradient, and
pancreatic hormone levels. Arterial blood glucose was
similar between the two groups at baseline and rose by ~80% during
the experimental period in both groups (Fig.
2). The a-pv glucose gradient was positive
in both groups at baseline (3.1 ± 0.4 in sedentary and 4.3 ± 0.4 mg/dl in exercised dogs) and became markedly negative during the
experimental period (17.9 ± 1.8 in sedentary and
19.3 ± 3.4 mg/dl in exercised dogs). There were no differences in the a-pv glucose gradient between the two groups.
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As expected, arterial plasma insulin at baseline was slightly lower in the exercised than in the sedentary dogs. Insulin levels rose by about threefold during the experimental period and were similar in the two groups. Arterial plasma glucagon was higher at baseline in exercised compared with sedentary dogs. During the experimental period, the pancreatic clamp equalized glucagon levels in the two groups. The presence of a metabolic steady state was reflected by the stability of the measurements described previously during the last 40 min of the experimental period.
Tracer-determined total and endogenous glucose
Ra.
Ra was not significantly higher in
the exercised compared with the sedentary animals at baseline (4.0 ± 0.5 vs. 2.8 ± 0.3 mg · kg1 · min
1).
Total Ra (endogenous + exogenous)
became significantly greater in the exercised animals during the
experimental period (11.1 ± 0.8 vs. 7.0 ± 0.5 mg · kg
1 · min
1).
Endogenous glucose Ra was
completely suppressed during the experimental period in both groups.
Hepatic glucose metabolism. The basal
hepatic glucose load was 23 ± 1 mg · kg1 · min
1
in the sedentary dogs and 25 ± 1 mg · kg
1 · min
1
in the exercised dogs (Fig. 3). It
increased to 36 ± 2 mg · kg
1 · min
1
in sedentary and to 40 ± 1 mg · kg
1 · min
1
in exercised dogs during the experimental period. Exercised dogs had a
significantly higher basal net hepatic glucose output compared with
sedentary animals (4.2 ± 0.3 vs. 1.9 + 0.2 mg · kg
1 · min
1,
P < 0.01). Animals in
both groups shifted to NHGU during the experimental period. There was
significantly more net glucose uptake in exercised than in sedentary
dogs (
3.2 ± 0.4 vs.
1.8 ± 0.2 mg · kg
1 · min
1,
P < 0.05). The total change in net
hepatic glucose balance from baseline to the experimental period was
therefore 3.7 ± 0.5 mg · kg
1 · min
1
in sedentary dogs and 7.3 ± 0.7 mg · kg
1 · min
1
in exercised animals (P < 0.001 vs.
sedentary). The net hepatic fractional extraction of glucose during the
experimental period was greater in the exercised than in the sedentary
dogs (0.08 ± 0.01 vs. 0.05 ± 0.01, P < 0.02).
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Hindlimb glucose metabolism. The basal net hindlimb glucose uptake was 4.0 ± 0.9 mg/min in the sedentary dogs and 8.4 ± 0.9 mg/min in the exercised dogs (P < 0.05 vs. sedentary). During the experimental period, it rose to 13.2 ± 1.4 mg/min in sedentary dogs and to 32.8 ± 3.5 mg/min in exercised dogs (P < 0.01 vs. sedentary). Net hindlimb glucose fractional extraction paralleled the response of net hindlimb glucose uptake (2.6 ± 0.5% in sedentary dogs and 4.6 ± 0.4% in exercised dogs at baseline, P < 0.05; 4.7 ± 0.5% in sedentary dogs and 13.4 ± 1.0% in exercised dogs during the experimental period, P < 0.05).
Blood levels and hepatic and hindlimb balances of
gluconeogenic precursors. Arterial lactate
concentrations were similar at baseline and increased by ~80% in
both groups (Fig. 4). Exercised animals
displayed significantly more basal net hepatic lactate uptake
(12.5 ± 1.2 vs.
7.3 ± 1.7 µmol · kg
1 · min
1,
P < 0.05). During the
experimental period, all animals shifted to net hepatic lactate output,
with no significant difference between the two groups. The basal net
hindlimb lactate output was
21.8 ± 4.0 and
13.8 ± 4.7 µmol/min (P > 0.05;
nonsignificant) in exercised and sedentary dogs, respectively. During
the experimental period, net hindlimb lactate output was suppressed in
both groups.
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Net hepatic alanine and glycerol uptakes were higher at baseline in exercised dogs; similar values of hepatic uptake of both metabolites were detected during the experimental period (data not shown).
Blood flows. Although basal hepatic
artery blood flow was higher in the exercised dogs than in the
sedentary dogs (P < 0.05), total
splanchnic blood flow was similar between the two groups throughout the
study (Table 1). The external iliac artery
blood flow was similar between groups throughout the study.
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DISCUSSION |
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It is well established that prior exercise increases insulin
sensitivity and glucose effectiveness (12), leading to enhanced rates
of net muscle glucose uptake and glycogen synthesis (7, 8, 14). By
comparison, far less is known about how prior exercise affects NHGU. In
the present study, the influence of prior exercise on the net hepatic
uptake of an intraportal venous glucose load was investigated in
conscious dogs in which circulating glucose, insulin, glucagon, a-pv
glucose gradient, and hepatic glucose load were carefully controlled.
Prior exercise led to a 74% greater NHGU and a 58% greater net
hepatic glucose fractional extraction compared with sedentary controls.
The total change in net hepatic glucose balance from the basal period
to the glucose infusion period was 3.7 and 7.3 mg · kg1 · min
1
in sedentary and exercised dogs, respectively. Whole body glucose uptake was 57% greater in exercised compared with sedentary animals. It could be calculated that 26 and 28% of the whole body glucose uptake could be accounted for by NHGU in the sedentary and exercised dogs, respectively. Although the exercised dogs consumed more glucose,
the percent contribution of hepatic and nonhepatic tissues to whole
body glucose uptake was similar.
The hypothesis that liver glucose uptake could be stimulated by prior
exercise is consistent with previous studies in humans (16) and rabbits
(20). Kawamori et al. (16) showed that the splanchnic disposal of an
oral glucose load during a euglycemic-hyperinsulinemic clamp was about
twice as high after exercise as after rest in non-insulin-dependent
diabetes mellitus patients. In this study, glucagon levels were not
controlled or even measured, and only an indirect estimate of the NHGU
was made. Matsuhisa et al. (20) measured the accumulation of
phosphorylated 3FDG in the liver of anesthetized rabbits after 30 min
of electrical stimulation of the hindlimb muscles. The accumulation of
phosphorylated 3FDG in the liver was approximately twofold greater
after muscle contraction compared with nonelectrically stimulated
control rabbits. The effect of prior exercise directly on the liver was
difficult to assess because the animals that underwent muscle
contraction had higher circulating insulin. Although the studies
previously cited provide indirect support for the hypothesis that prior
exercise stimulates NHGU, in none were the factors that are known to
influence NHGU controlled nor was NHGU measured directly. Hamilton et
al. (8) showed that rates of NHGU were similar in exercised dogs and in
sedentary controls, when glucose was infused itraduodenally at 8 mg · kg1 · min
1.
Because a greater suppression of endogenous
Ra was seen in exercised animals
at a time when rates of NHGU were similar to sedentary controls,
unidirectional hepatic glucose uptake must also have been reduced by
prior exercise. Endogenous pancreatic hormone secretion was not
suppressed in the study by Hamilton et al., and circulating glucagon
levels were greater in the exercised dogs. Because glucagon can
antagonize NHGU (13), the higher glucagon levels may have masked a NHGU
response in the exercised dogs. Maehlum et al. (19) have reported that
a greater proportion of an oral glucose load escapes hepatic retention
in exercised than in sedentary humans. Although this finding may appear
contrary to the findings from the present study, it should be noted
that in the work by Maehlum et al. the experimental setting was
significantly different from ours. The human subjects in their study
were administered glucose orally in a bolus. In this situation, the
glucose load reaching the intestine is the same in the two experimental
groups, but the load reaching the liver is greater in the exercised
subjects, as exercise enhances the ability of the gut to absorb
ingested glucose (5, 8). Because this exercise-induced increase in hepatic glucose load is greater than the increase in NHGU, a greater amount of glucose leaves the splanchnic region in response to oral
glucose after exercise.
All factors known to stimulate NHGU were controlled in the present study. It could be argued, however, that other extrahepatic determinants such as catecholamines, cortisol, and growth hormone, which were not measured, may also have influenced liver metabolism. Arterial catecholamine levels have been previously demonstrated to increase two- to threefold during an exercise bout comparable with that used in this study (2). Nevertheless, the experimental period started 30 min after the cessation of exercise, at which time catecholamine levels have been shown to return to basal levels (2). Further, catecholamines mainly affect liver glucose metabolism by increasing the availability of gluconeogenic precursors from the periphery. In this study, the net liver uptake of glycerol and alanine and the net liver output of lactate were similar in exercised and sedentary dogs. Cortisol and growth hormone levels are also increased during exercise; their influences on hepatic glucose metabolism, nevertheless, have not to date been clearly defined. It should be noted, however, that the increased secretion of the aforementioned counterregulatory hormones favors net glucose output by the liver, not NHGU. Controlling the levels of these hormones should, if anything, have enhanced the effect of prior exercise on NHGU.
A strong positive correlation exists between NHGU and both insulin
concentration and hepatic glucose load (23, 24). Increased sensitivity
to one of these factors, or both, may cause the increase in NHGU after
exercise. A positive correlation also exists at rest between NHGU and
the magnitude of a negative a-pv glucose gradient until values of
approximately 20 mg/dl, above which no further enhancement of
NHGU was observed (25). The a-pv glucose gradients measured in our
study (
18 and
19 mg/dl) were very close to the value that
maximally stimulates NHGU (25). It is possible, however, that prior
exercise increases the sensitivity or responsiveness of NHGU to the
a-pv glucose gradient. At the cellular level, glucose transport across
the hepatocyte membrane is primarily mediated by the GLUT-2 isoform of
the glucose transporter family. Once in the cytoplasm, glucose is
phosphorylated through a reaction catalyzed by the enzyme glucokinase
(GK). GLUT-2 is not believed to play a limiting role in the movement of
glucose from the extracellular to the intracellular space of the
hepatocyte. GK activity, on the other hand, is thought to be rate
limiting for glucose uptake in the liver and can be regulated in
response to different stimuli, such as glucose, fructose, and glucagon concentrations (1). An increase in GK activity occurs because of the
translocation of GK from the nucleus of the hepatocyte to the cytosol
and the loss of the inhibitory binding with its regulatory protein.
Although the effect of exercise on GK activity has not been completely
elucidated, a significant increase in GK activity has been reported
after exercise in rats (4). Hamilton et al. (8) observed that GK
activity was not significantly different in exercised and sedentary
dogs after an intraduodenal glucose load. In these previous studies, GK
analysis was performed on crude liver tissue homogenates, and enzymatic
movement from the nuclear membrane into the cytosol could not be
detected. In addition, other determinants of liver glucose uptake were
not controlled.
Because liver glycogen content and synthetic rates were not measured in the present study, it is impossible to specify the intrahepatic fate of the excess glucose taken up by the liver in response to a glucose load in the postexercise state. The net hepatic balances of the gluconeogenic precursors lactate, alanine, and glycerol, on the other hand, were similar in sedentary and exercised animals. This suggests that glycogen storage was probably the preferential pathway through which the liver of exercised dogs directed the excess glucose taken up compared with sedentary animals.
Prior exercise induced greater net hindlimb glucose uptake during glucose infusion, despite similar hindlimb glucose and insulin loads in exercised and sedentary animals. This is consistent with numerous previous studies in which prior exercise led to greater net skeletal muscle glucose fractional extraction due to enhanced muscle insulin sensitivity (28, 30) as well as increased insulin-independent glucose uptake (6, 29). The greater net glucose uptake by the skeletal muscle has been previously shown to result in greater rates of intramuscular glycogen synthesis (19, 29). Although muscle glycogen was not measured in the present study, these previous reports are consistent with our observation that net muscle lactate output was similar in sedentary and exercised animals.
As stated previously, the contribution of the liver to whole body glucose uptake is quantitatively smaller than that of the skeletal muscle (in the present study the liver accounted for 26-28% of whole body glucose uptake, whereas the large majority of the remainder could be accounted for by total whole body glucose uptake). The importance of the liver as a contributor to this process, on the other hand, appears more clearly if glucose uptake rates are expressed per unit of tissue mass. In dogs of the size used (average, 24 kg), the skeletal muscle of a single hindlimb and the liver have similar masses (~650-700 g; Ref. 31). After exercise, glucose infusion resulted in a net hindlimb glucose uptake of ~33 mg/min or 0.047 mg/g of muscle tissue; NHGU was ~77 mg/min or 0.110 mg/g of liver tissue. The difference in muscle vs. liver glucose uptake is even more striking if the whole body muscle mass is considered. The whole body, nonhepatic glucose uptake was 158 mg/min in exercised dogs during glucose infusion. Assuming that all this amount was due to muscle glucose uptake, dividing this rate of 158 mg/min by 10.8 kg (~45% of our dogs average total body weight; Ref. 31) yields an average muscle glucose uptake of 0.015 mg/g of tissue, or ~15% of the liver uptake per gram tissue.
In summary, the work presented here provides for the first time clear evidence that prior exercise can directly influence liver carbohydrate metabolism by significantly increasing net glucose uptake by this organ. This result was obtained by strictly controlling the fundamental determinants of NHGU (concentrations of glucose and pancreatic hormones, a-pv glucose gradient, hepatic glucose load), thereby isolating effects of prior exercise per se. In this setting, prior exercise resulted in a 74% increase in NHGU and in a 58% increase in net hepatic glucose fractional extraction compared with sedentary controls. NHGU comprised one-third of the added whole body glucose uptake after exercise. Our data show that the liver is a major site of glucose removal when a carbohydrate load is administered in the postexercise state.
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ACKNOWLEDGEMENTS |
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We thankfully acknowledge Wanda Snead, Pam Venson, and Brittina Murphy for excellent technical assistance.
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FOOTNOTES |
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This work was supported by the National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-50277. P. Galassetti was supported by the National Institute of Health Training Grant DK-07061.
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 reprint requests and other correspondence: P. Galassetti, 754 MRB-I, Vanderbilt Univ. Medical Center, Nashville, TN 37232-0615 (E-mail: pietro.galassetti{at}mcmail.vanderbilt.edu).
Received 23 October 1998; accepted in final form 23 February 1999.
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