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Regulation of Muscle Glucose Uptake In Vivo

Lawrence J. Mandarino, Riccardo C. Bonadonna, Owen P. Mcguinness, Amy E. Halseth, David H. Wasserman

10.1002/cphy.cp070227

Source: Supplement 21: Handbook of Physiology, The Endocrine System, The Endocrine Pancreas and Regulation of Metabolism

Originally published: 2001

Published online: January 2011

Full Article on Wiley Online Library



Abstract

The sections in this article are:

1 Methodologies for Measurement of Skeletal Muscle Glucose Uptake
1.1 Theories of Metabolic Control
1.2 Kinetic Theory of Intact Organs
1.3 Arteriovenous Differences
1.4 Muscle Biopsies
1.5 Positron Emission Tomography
1.6 Nuclear Magnetic Resonance Spectroscopy
1.7 Multiple Tracer Dilution Technique
1.8 Measurement of Whole Body Glucose Metabolism
2 Mechanism of Insulin Action at Skeletal Muscle
2.1 Insulin Signaling In Vivo
2.2 Glucose Transport and Phosphorylation in Skeletal Muscle
2.3 Glycogen Synthesis, Glycolysis, and Glucose Oxidation in Skeletal Muscle
2.4 Effect of Insulin on Blood Flow
2.5 Transport of Insulin Across the Endothelial Barrier Time‐Limiting Steps in Insulin Action
2.6 Modulation of Insulin‐Stimulated Skeletal Muscle Glucose Uptake by Nonesterified Fatty Acid Availability
3 Exercise and Muscle Glucose Uptake
3.1 Mechanism of the Exercise‐Induced Increase in Muscle Glucose Uptake
3.2 Modulation of Glucose Uptake by the Working Muscle by the Internal Milieu
3.3 Muscle Glucose Uptake in the Postexercise State
4 Conclusion
Figure 1. Figure 1.

Simplified diagram of muscle glucose metabolism. Note that while glucose transport, being a process of facilitated diffusion, is a bidirectional step, hexokinase (HK) II catalyzes an irreversible conversion of glucose to glucose‐6‐phosphate (G6P). Downstream of G6P, glucose metabolism divides into two pathways: glycogen synthesis and glycolysis. Formation of glycogen is not an irreversible phenomenon, although, for the sake of simplicity, this is not shown in the figure. Glycogen can be broken down by the enzyme glycogen phosphorylase to glucose‐l‐phosphate and, hence, G6P, which can be again reutilized to form glycogen or channeled to glycolysis. In contrast, the entry into the glycolytic pathway is an irreversible step catalyzed by phosphofructokinase.
Figure 2. Figure 2.

Plot showing the relationship between plasma insulin levels and forearm glucose uptake in healthy individuals, analyzed according to eq. 2. This empirical relationship is best fitted by a second‐order polynomial (R2 = 0.783, p < .001). According to the principles of sensitivity analysis, net sensitivity of forearm (muscle) glucose uptake to insulin is given by the first derivative of this function.

[Data Bonadonna et al. 42.]
Figure 3. Figure 3.

Net sensitivity function of forearm (muscle) glucose uptake to plasma insulin levels derived from the data shown in Figure 2. Note that net sensitivity is not a constant, but changes monotonically with the changes in plasma insulin levels. Note also the logarithmic scale of the x‐axis. If the x‐axis were in a linear scale, the net sensitivity to insulin would be described by an exponential function.
Figure 4. Figure 4.

Control strength values of glucose transporters and HK (shaded bars) in vivo on glucose uptake of human skeletal muscle in healthy individuals (n = 7) in the basal state (left panel) and during systemic physiological (˜ 75 μU/ml) hyperinsulinemia (right panel). In the postabsorptive state, HK plays the predominant role in determining muscle glucose uptake. Insulin reverses this situation and gives a prominent role to glucose transporters. These control strength values were calculated utilizing the rates of transmembrane inward and outward glucose transport, the rate of glucose phosphorylation, and extracellular and intracellular glucose concentrations.

[Data from Bonadonna et al. 41 and Saccomani et al. 276.]
Figure 5. Figure 5.

Simplified diagram of muscle glucose metabolism showing the potential role of blood flow and capillary exchange in affecting muscle glucose uptake in a capillary‐tissue unit. Blood flow plays a dual role: its acceleration increases the glucose supply but also decreases the time during which the glucose molecules can be exchanged with the interstitium. According to the theory of transcapillary exchanges, the former phenomenon is always quantitatively more relevant than the latter, which only at the asymptotically highest blood flows can completely offset the increase in glucose supply.
Figure 6. Figure 6.

Plot showing the dependence of transcapillary glucose exchange on blood flow in human muscle in vivo at low (8 ml/min/kg) and average (20 ml/min/kg) values of capillary PS for glucose. It can be seen that within the range of insulin‐mediated increases in muscle blood flow and in the absence of capillary recruitment, blood flow per se has a minor impact on the amount of glucose exchanged with the interstitium. The average value of 20 ml/min/kg of capillary PS in human muscle was extrapolated from reference 44.
Figure 7. Figure 7.

The model utilized for the analysis of FDG‐PET experimental data. The time course of arterial FDG concentration is independently measured. The time course of 18F radioactivity is assumed to be the sum of three components: arterial FDG activity, cellular FDG, and FDG‐6‐phosphate irreversibly trapped inside the cell. The rate constant k1 represents transport of FDG inside the cell; the rate constant k2 quantifies the transport of FDG outside the cell, and the rate constant k3 quantifies the phosphorylation of FDG. See text for more details.
Figure 8. Figure 8.

Washout curves of an intravascular indicator and 3H2O in the deep forearm vein of a representative healthy human subject after a pulse injection of the two indicators at time = 0 into the brachial artery. Although the fractional extraction of water in human muscle is quite high (˜0.70), it is much lower than the values (>0.96) reported in the brain and in the heart.
Figure 9. Figure 9.

Washout curves of [1‐3H]‐D‐mannitol, 3‐O‐[14C]‐methyl‐D‐glucose, and [3‐3H]‐D‐glucose in the deep forearm vein of a representative healthy human subject after a pulse injection of three substances at time = 0 into the brachial artery. The difference in concentration between D‐mannitol (a nontransportable, nonmetabolizable glucose analogue) and 3‐O‐methyl‐D‐glucose (a transportable, nonmetabolizable glucose analogue) reflects the activity of transmembrane glucose transport, whereas the difference between 3‐O‐methyl‐D‐glucose and D‐glucose, which can also be metabolized, reflects the activity of phosphorylation.
Figure 10. Figure 10.

The compartmental model developed to analyze multiple tracer dilution data such as those presented in Figure 9. Three paths (compartments 2 to 7) with different speeds (fast, intermediate, and slow) are necessary and sufficient to describe D‐mannitol kinetics. There is also an irreversible exit from the injection pool (artery) because, as the forearm is drained by several veins and blood is sampled in only one of them, a variable amount of the three indicators is not recovered in the sampling pool. To these three pathways, intracellular pools 9 to 14 in bidirectional exchange (transmembrane transport of 3‐O‐methyl‐D‐glucose and D‐glucose) with the extracellular volume are appended. From these intracellular pools, D‐glucose (but not 3‐O‐methyl‐D‐glucose) can be irreversibly lost (unidirectional arrows) through the process of phosphorylation. Application of this model to experimental data such as those presented in Figure 9 allows one to compute the fluxes of glucose transport into and out of the cell, the flux of intracellular phosphorylation, and extracellular (tissue), and intracellular glucose concentrations 41, 276.
Figure 11. Figure 11.

This figure shows a typical euglycemic clamp experiment. The upper panel shows that the plasma glucose concentration (closed circles) is tightly maintained at a plasma glucose concentration of about 90 mg/dl. The lower panel shows the plasma insulin concentrations that result from a primed, continuous infusion of insulin at a rate of 1 mU/kg · min. The upper panel also shows the rate of infusion of glucose required to keep the plasma glucose constant during hyperinsulinemia.

[From DeFronzo et al. 87.]
Figure 12. Figure 12.

This figure demonstrates the ability of insulin to increase HK II, but not HK I or GS mRNA abundance, in human skeletal muscle. In this experiment 203, a euglycemic hyperinsulinemica clamp was performed in nine subjects. Needle biopsies of the vastus lateralis muscle were performed before and after 4 h of physiological hyperinsulinemia (approximately 65 μU/ml). RNA isolated from the biopsy samples was used in RNase protection assays to quantify the mRNA level of GS, HK I, and HK II. Hexokinase II mRNA was increased by threefold over basal values, but HK I and GS mRNA were not increased.
Figure 13. Figure 13.

This figure illustrates the variability in the pathways of insulin‐stimulated leg muscle glucose metabolism in humans. The full height of each bar represents insulin‐stimulated leg glucose uptake (mmoles of glucose per minute · 100 ml leg tissue) for each of seven healthy male subjects (a‐g). The components of leg glucose metabolism are shown within each bar. Glucose oxidation is shown as the hatched bar, nonoxidized glycolysis (net release of lactate and alanine) is the stippled area, and glucose storage (glycogen formation) is the open area. There is more than a twofold difference among these normal volunteers in the magnitude of total leg glucose metabolism. It can be appreciated that variability in leg muscle glucose storage as glycogen is the primary determinant of the variability in overall glucose uptake.

[From Kelley et al. 171.]
Figure 14. Figure 14.

This figure illustrates how insulin and G6P increase GS activity. Glucose‐6‐phosphate binds allosterically to GS and increases its activity. The physiological range of G6P is shown as a shaded area, and it can be seen that small changes in the G6P concentration can have a large impact on GS activity. Insulin results in dephosphorylation of the enzyme and a resultant shift in the G6P dose‐response curve to the left. As a result, GS activity at a given G6P concentration is increased by insulin. This effect is especially pronounced within the physiological range of G6P concentrations.
Figure 15. Figure 15.

Hindlimb glucose uptake (left), glucose + lactate oxidation (middle) and nonoxidative glucose + lactate metabolism (right) versus plasma glucose concentration during rest and exercise in chronically catheterized dogs. The glucose clamp technique was used to maintain glucose at one of four glucose levels. Exercise samples were obtained during the last 40 min of a 90 min work period. Insulin levels were fixed at basal using somatostatin and intraportal insulin replacement. Oxidative and nonoxidative metabolism were calculated including 14C‐lactate specific activity when net limb lactate uptake is present. Five samples show a glucose level of 5.0 mM, and four samples represent all other glucose levels. Data are mean ± SE.

[Modified from Zinker et al. 347.]
Figure 16. Figure 16.

Effect of exercise alone (n = 19) and exercise plus somatostatin with (n = 8) and without (n = 6) insulin replacement on whole body glucose disappearance in dogs. Data are mean ± SE.

[Modified from Wasserman et al. 321.]
Figure 17. Figure 17.

Comparison of the effect of 1 μM wortmannin on basal and insulin‐ and contraction‐stimulated skeletal muscle glucose transport. Intact soleus muscles were preincubated with or without wortmannin for 10 min and then studied during a second incubation using the same medium under basal, insulin‐stimulated, or contraction‐stimulated conditions. p < .01 vs. a corresponding experiment performed in the absence of wortmannin.

[Modified from Lund et al. 198.]


Figure 1.

Simplified diagram of muscle glucose metabolism. Note that while glucose transport, being a process of facilitated diffusion, is a bidirectional step, hexokinase (HK) II catalyzes an irreversible conversion of glucose to glucose‐6‐phosphate (G6P). Downstream of G6P, glucose metabolism divides into two pathways: glycogen synthesis and glycolysis. Formation of glycogen is not an irreversible phenomenon, although, for the sake of simplicity, this is not shown in the figure. Glycogen can be broken down by the enzyme glycogen phosphorylase to glucose‐l‐phosphate and, hence, G6P, which can be again reutilized to form glycogen or channeled to glycolysis. In contrast, the entry into the glycolytic pathway is an irreversible step catalyzed by phosphofructokinase.



Figure 2.

Plot showing the relationship between plasma insulin levels and forearm glucose uptake in healthy individuals, analyzed according to eq. 2. This empirical relationship is best fitted by a second‐order polynomial (R2 = 0.783, p < .001). According to the principles of sensitivity analysis, net sensitivity of forearm (muscle) glucose uptake to insulin is given by the first derivative of this function.

[Data Bonadonna et al. 42.]


Figure 3.

Net sensitivity function of forearm (muscle) glucose uptake to plasma insulin levels derived from the data shown in Figure 2. Note that net sensitivity is not a constant, but changes monotonically with the changes in plasma insulin levels. Note also the logarithmic scale of the x‐axis. If the x‐axis were in a linear scale, the net sensitivity to insulin would be described by an exponential function.



Figure 4.

Control strength values of glucose transporters and HK (shaded bars) in vivo on glucose uptake of human skeletal muscle in healthy individuals (n = 7) in the basal state (left panel) and during systemic physiological (˜ 75 μU/ml) hyperinsulinemia (right panel). In the postabsorptive state, HK plays the predominant role in determining muscle glucose uptake. Insulin reverses this situation and gives a prominent role to glucose transporters. These control strength values were calculated utilizing the rates of transmembrane inward and outward glucose transport, the rate of glucose phosphorylation, and extracellular and intracellular glucose concentrations.

[Data from Bonadonna et al. 41 and Saccomani et al. 276.]


Figure 5.

Simplified diagram of muscle glucose metabolism showing the potential role of blood flow and capillary exchange in affecting muscle glucose uptake in a capillary‐tissue unit. Blood flow plays a dual role: its acceleration increases the glucose supply but also decreases the time during which the glucose molecules can be exchanged with the interstitium. According to the theory of transcapillary exchanges, the former phenomenon is always quantitatively more relevant than the latter, which only at the asymptotically highest blood flows can completely offset the increase in glucose supply.



Figure 6.

Plot showing the dependence of transcapillary glucose exchange on blood flow in human muscle in vivo at low (8 ml/min/kg) and average (20 ml/min/kg) values of capillary PS for glucose. It can be seen that within the range of insulin‐mediated increases in muscle blood flow and in the absence of capillary recruitment, blood flow per se has a minor impact on the amount of glucose exchanged with the interstitium. The average value of 20 ml/min/kg of capillary PS in human muscle was extrapolated from reference 44.



Figure 7.

The model utilized for the analysis of FDG‐PET experimental data. The time course of arterial FDG concentration is independently measured. The time course of 18F radioactivity is assumed to be the sum of three components: arterial FDG activity, cellular FDG, and FDG‐6‐phosphate irreversibly trapped inside the cell. The rate constant k1 represents transport of FDG inside the cell; the rate constant k2 quantifies the transport of FDG outside the cell, and the rate constant k3 quantifies the phosphorylation of FDG. See text for more details.



Figure 8.

Washout curves of an intravascular indicator and 3H2O in the deep forearm vein of a representative healthy human subject after a pulse injection of the two indicators at time = 0 into the brachial artery. Although the fractional extraction of water in human muscle is quite high (˜0.70), it is much lower than the values (>0.96) reported in the brain and in the heart.



Figure 9.

Washout curves of [1‐3H]‐D‐mannitol, 3‐O‐[14C]‐methyl‐D‐glucose, and [3‐3H]‐D‐glucose in the deep forearm vein of a representative healthy human subject after a pulse injection of three substances at time = 0 into the brachial artery. The difference in concentration between D‐mannitol (a nontransportable, nonmetabolizable glucose analogue) and 3‐O‐methyl‐D‐glucose (a transportable, nonmetabolizable glucose analogue) reflects the activity of transmembrane glucose transport, whereas the difference between 3‐O‐methyl‐D‐glucose and D‐glucose, which can also be metabolized, reflects the activity of phosphorylation.



Figure 10.

The compartmental model developed to analyze multiple tracer dilution data such as those presented in Figure 9. Three paths (compartments 2 to 7) with different speeds (fast, intermediate, and slow) are necessary and sufficient to describe D‐mannitol kinetics. There is also an irreversible exit from the injection pool (artery) because, as the forearm is drained by several veins and blood is sampled in only one of them, a variable amount of the three indicators is not recovered in the sampling pool. To these three pathways, intracellular pools 9 to 14 in bidirectional exchange (transmembrane transport of 3‐O‐methyl‐D‐glucose and D‐glucose) with the extracellular volume are appended. From these intracellular pools, D‐glucose (but not 3‐O‐methyl‐D‐glucose) can be irreversibly lost (unidirectional arrows) through the process of phosphorylation. Application of this model to experimental data such as those presented in Figure 9 allows one to compute the fluxes of glucose transport into and out of the cell, the flux of intracellular phosphorylation, and extracellular (tissue), and intracellular glucose concentrations 41, 276.



Figure 11.

This figure shows a typical euglycemic clamp experiment. The upper panel shows that the plasma glucose concentration (closed circles) is tightly maintained at a plasma glucose concentration of about 90 mg/dl. The lower panel shows the plasma insulin concentrations that result from a primed, continuous infusion of insulin at a rate of 1 mU/kg · min. The upper panel also shows the rate of infusion of glucose required to keep the plasma glucose constant during hyperinsulinemia.

[From DeFronzo et al. 87.]


Figure 12.

This figure demonstrates the ability of insulin to increase HK II, but not HK I or GS mRNA abundance, in human skeletal muscle. In this experiment 203, a euglycemic hyperinsulinemica clamp was performed in nine subjects. Needle biopsies of the vastus lateralis muscle were performed before and after 4 h of physiological hyperinsulinemia (approximately 65 μU/ml). RNA isolated from the biopsy samples was used in RNase protection assays to quantify the mRNA level of GS, HK I, and HK II. Hexokinase II mRNA was increased by threefold over basal values, but HK I and GS mRNA were not increased.



Figure 13.

This figure illustrates the variability in the pathways of insulin‐stimulated leg muscle glucose metabolism in humans. The full height of each bar represents insulin‐stimulated leg glucose uptake (mmoles of glucose per minute · 100 ml leg tissue) for each of seven healthy male subjects (a‐g). The components of leg glucose metabolism are shown within each bar. Glucose oxidation is shown as the hatched bar, nonoxidized glycolysis (net release of lactate and alanine) is the stippled area, and glucose storage (glycogen formation) is the open area. There is more than a twofold difference among these normal volunteers in the magnitude of total leg glucose metabolism. It can be appreciated that variability in leg muscle glucose storage as glycogen is the primary determinant of the variability in overall glucose uptake.

[From Kelley et al. 171.]


Figure 14.

This figure illustrates how insulin and G6P increase GS activity. Glucose‐6‐phosphate binds allosterically to GS and increases its activity. The physiological range of G6P is shown as a shaded area, and it can be seen that small changes in the G6P concentration can have a large impact on GS activity. Insulin results in dephosphorylation of the enzyme and a resultant shift in the G6P dose‐response curve to the left. As a result, GS activity at a given G6P concentration is increased by insulin. This effect is especially pronounced within the physiological range of G6P concentrations.



Figure 15.

Hindlimb glucose uptake (left), glucose + lactate oxidation (middle) and nonoxidative glucose + lactate metabolism (right) versus plasma glucose concentration during rest and exercise in chronically catheterized dogs. The glucose clamp technique was used to maintain glucose at one of four glucose levels. Exercise samples were obtained during the last 40 min of a 90 min work period. Insulin levels were fixed at basal using somatostatin and intraportal insulin replacement. Oxidative and nonoxidative metabolism were calculated including 14C‐lactate specific activity when net limb lactate uptake is present. Five samples show a glucose level of 5.0 mM, and four samples represent all other glucose levels. Data are mean ± SE.

[Modified from Zinker et al. 347.]


Figure 16.

Effect of exercise alone (n = 19) and exercise plus somatostatin with (n = 8) and without (n = 6) insulin replacement on whole body glucose disappearance in dogs. Data are mean ± SE.

[Modified from Wasserman et al. 321.]


Figure 17.

Comparison of the effect of 1 μM wortmannin on basal and insulin‐ and contraction‐stimulated skeletal muscle glucose transport. Intact soleus muscles were preincubated with or without wortmannin for 10 min and then studied during a second incubation using the same medium under basal, insulin‐stimulated, or contraction‐stimulated conditions. p < .01 vs. a corresponding experiment performed in the absence of wortmannin.

[Modified from Lund et al. 198.]
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Article Title: Regulation of Muscle Glucose Uptake In Vivo
 

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Lawrence J. Mandarino, Riccardo C. Bonadonna, Owen P. Mcguinness, Amy E. Halseth, David H. Wasserman. Regulation of Muscle Glucose Uptake In Vivo. Compr Physiol 2011, Supplement 21: Handbook of Physiology, The Endocrine System, The Endocrine Pancreas and Regulation of Metabolism: 803-845. First published in print 2001. doi: 10.1002/cphy.cp070227