EPILOGUE
Metabolic control analysis of insulin-stimulated glucose disposal in rat skeletal muscle

Beat M. Jucker1, Nicole Barucci1, and Gerald I. Shulman2

1 Department of Internal Medicine and the 2 Howard Hughes Medical Institute, Yale University School of Medicine, New Haven, Connecticut 06520-8020


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

Metabolic control analysis was used to calculate the distributed control of insulin-stimulated skeletal muscle glucose disposal in awake rats. Three separate hyperinsulinemic infusion protocols were performed: 1) protocol I was a euglycemic (~6 mM)-hyperinsulinemic (10 mU · kg-1 · min-1) clamp, 2) protocol II was a hyperglycemic (~11 mM)-hyperinsulinemic (10 mU · kg-1 · min-1) clamp, and 3) protocol III was a euglycemic (~6 mM)-hyperinsulinemic (10 mU · kg-1 · min-1)-lipid/heparin (increased plasma free fatty acid) clamp. [1-13C]glucose was administered in all three protocols for a 3-h period, during which time [1-13C]glucose label incorporation into [1-13C]glycogen, [3-13C]lactate, and [3-13C]alanine was detected in the hindlimb of awake rats via 13C-NMR. Combined steady-state and kinetic data were used to calculate rates of glycogen synthesis and glycolysis. Additionally, glucose 6-phosphate (G-6-P) was measured in the hindlimb muscles with the use of in vivo 31P-NMR during the three infusion protocols. The clamped glucose infusion rates were 31.6 ± 2.9, 49.7 ± 1.0, and 24.0 ± 1.5 mg · kg-1 · min-1 at 120 min in protocols I-III, respectively. Rates of glycolysis were 62.1 ± 10.3, 71.6 ± 11.8, and 19.5 ± 3.6 nmol · g-1 · min-1 and rates of glycogen synthesis were 125 ± 15, 224 ± 23, and 104 ± 17 nmol · g-1 · min-1 in protocols I-III, respectively. Insulin-stimulated G-6-P concentrations were 217 ± 8, 265 ± 12, and 251 ± 9 nmol/g in protocols I-III, respectively. A top-down approach to metabolic control analysis was used to calculate the distributed control among glucose transport/phosphorylation [GLUT-4/hexokinase (HK)], glycogen synthesis, and glycolysis from the metabolic flux and G-6-P data. The calculated values for the control coefficients (C) of these three metabolic steps (CJGLUT-4/HK = 0.55 ± 0.10, CJglycogen syn = 0.30 ± 0.06, and CJglycolysis = 0.15 ± 0.02; where J is glucose disposal flux, and glycogen syn is glycogen synthesis) indicate that there is shared control of glucose disposal and that glucose transport/phosphorylation is responsible for the majority of control of insulin-stimulated glucose disposal in skeletal muscle.

nuclear magnetic resonance; flux control; glycogen synthesis; glycolysis; glucose 6-phosphate


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

MANY OF THE INITIAL STUDIES addressing the mechanism of metabolic control have focused on defining a rate-limiting or rate-controlling step in metabolism (20, 25, 26). It was generally thought that a rate-limiting enzyme was near the start of a metabolic pathway catalyzing a nonequilibrium reaction. As initially described by Kacser and Burns (15), the control of metabolism is often distributed over the entire system of enzymes composing the metabolic pathway. Consequently, it has been difficult to discover a single rate-limiting enzyme for metabolism by using traditional enzyme-expression or -inhibition techniques. With the use of a top-down approach to metabolic control theory, it is possible to determine a distribution of control over defined metabolic pathways and/or systems of enzymes that will vary depending on the physiological conditions present (16, 30). The top-down approach to metabolic control analysis (MCA) and its application to in vivo metabolic studies require only that flux through the applicable metabolic pathways and concentrations of associated substrates be measured. The quantitative measurements of metabolic fluxes and intermediate concentrations in vivo have been simplified in the advent of NMR spectroscopic techniques, which enables one to make simultaneous measurements of glycogen synthesis and glycolysis noninvasively in the hindlimb muscle of awake rats (12-14) by using 13C-NMR and glucose 6-phosphate (G-6-P) with the use of 31P-NMR spectroscopy (2, 3, 13).

In an effort to determine the control of glucose disposal in skeletal muscle during conditions of hyperinsulinemia, we used 13C- and 31P-NMR spectroscopy to measure noninvasively the partitioning of glucose disposal via glycogen synthesis and glycolysis, and G-6-P concentrations in the hindlimb skeletal muscles of rats during three separate infusion protocols, which were used to differentially modulate both the fluxes and intramuscular G-6-P concentrations. With the use of a top-down approach to MCA, elasticities (a relationship between enzyme velocity and substrate concentration) for glucose transport/phosphorylation, glycogen synthesis, and glycolysis to G-6-P were measured, and corresponding control coefficients were calculated. This study illustrates the utility of combining MCA with in vivo NMR to measure the distributed control of insulin-stimulated glucose disposal through associated transport and metabolic steps in rat skeletal muscle; it may be useful in characterizing the control of these pathways in different insulin-resistant states.


    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

Animals. Sprague-Dawley rats (Charles River, Raleigh, NC) were housed in an environmentally controlled room with a 12:12-h light-dark cycle. At a weight of 300-350 g, rats were chronically catheterized as described elsewhere (33) and allowed to recuperate for another 5-10 days.

In vivo experiments. All rats were fasted 24 h before the infusion experiment. Rats were placed in a customized restraining tube that allowed their left hindlimb to be secured to the outside of the tube in a manner to limit free movement of the leg for NMR measurements. The rats were transiently anesthetized (<30 s) with a low dose (2.5-5.0 mg) of thiopental (Sigma) to place them in the restraining tube. One of three infusion protocols was started: protocol I was a euglycemic (~6 mM)-hyperinsulinemic (10 mU · kg-1 · min-1; Humulin Regular, Eli Lilly) clamp, with 20% dextrose administered at 2.5 min after the commencement of the primed-continuous insulin infusion (n = 10 rats for 13C-NMR and n = 6 for 31P-NMR); protocol II was a hyperglycemic (~11 mM)-hyperinsulinemic (10 mU · kg-1 · min-1) clamp with somatostatin (1 µg · kg-1 · min-1; n = 10 for 13C-NMR and n = 6 for 31P-NMR); and protocol III was a euglycemic (~6 mM)-hyperinsulinemic (10 mU · kg-1 · min-1)-lipid/heparin (Liposyn II, Abbott; 1:3 vol/vol saline at 39 µl/min with heparin at 0.0975 IU/min) clamp (n = 9 for 13C-NMR and n = 5 for 31P-NMR). [1-13C]glucose (99% enriched, 20% wt/vol; Cambridge Isotope Laboratories, Cambridge, MA) was used during the 13C-NMR experiments. The glycolytic and glycogen synthesis flux measurements in protocols I-III originated from our previously published work (12-14). All clamps lasted for 180-240 min. Blood samples were drawn during the baseline NMR measurement, at 7.5 min, at 15 min, and every 15 min thereafter for immediate assessment of plasma glucose and lactate concentrations. At the end of the in vivo NMR experiment, rats were anesthetized with thiopental (50 mg/kg). Superficial skin was rapidly removed from the left hindquarter, followed by in situ freeze clamping of the gastrocnemius and biceps femoris muscles. Rats were euthanized with a lethal dose of thiopental. The protocol was reviewed and approved by the Yale University Animal Care Committee.

In vivo NMR spectroscopy. All in vivo 13C- and 31P-NMR experiments were performed on a Bruker Biospec 7.0-T system (horizontal/22-cm-diameter bore magnet) as previously described (11). 13C-NMR spectra were processed with the use of a Gaussian filter, followed by Fourier transformation and peak integration with the use of Bruker DISNMR processing software; 31P-NMR spectra were processed with the use of a Gaussian resolution enhancement and 100% Gaussian weighted peak fitting algorithm (Nuts NMR processing software; Acorn NMR, Fremont, CA). Concentrations of G-6-P, Pi, and phosphocreatine (PCr) were extrapolated from the in vivo spectra after correction for differential saturation with respect to the beta -ATP peak and measured ATP concentrations. The concentration of G-6-P was calculated as the average of three spectra taken between 105 and 135 min to increase the sensitivity of the measurement. Free intracellular Mg2+ was calculated with the use of the chemical shift difference between alpha - and beta -ATP resonances, with a dissociation constant of 50 µM (at pH 7.2, 37°C) for Mg-ATP (1, 8). The ADP concentration was calculated with the use of the creatine kinase equilibrium equation, and the creatine concentration was calculated from the measured total creatine and PCr concentrations (with an equilibrium constant of 1.66 × 109) (22). The intracellular pH was calculated with the use of the chemical shift difference of intracellular Pi and PCr as previously described (35).

Tissue extract analysis. Muscle tissue extracts (13C-NMR experiments) were prepared for high-field NMR analysis by homogenization of ~1 g of combined gastrocnemius and biceps femoris muscle with a variable high-speed electric homogenizer after the sample was placed in a vortex tube filled with 0.9% perchloric acid (3 vol/wt). After homogenization over ice, the sample was centrifuged at 4°C for 10 min (4,000 rpm). The supernatant was extracted, and the pellet was saved for glycogen enrichment measurements. KOH (4 N, 0.675 vol/wt) was added to the supernatant to precipitate excess perchlorate ions. The sample was centrifuged once more at 4°C for 15 min (4,000 rpm). The supernatant was extracted, and 0.5 M phosphate buffer (pH 7) was added to neutralize the sample. The sample was dried in a speed-vac (Savant, Farmingdale, NY) overnight, and 0.5 ml 2H2O was added to the dried powder before it was placed in a 5-mm NMR tube for NMR analysis at 8.4 T (Bruker WB-360 NMR spectrometer). Proton-observed carbon-enhanced spectroscopy was performed on tissue extract samples for fractional enrichment calculations. The broadband 13C inversion pulse was turned on during alternate transients, with raw data separated into two data sets, providing spectra with heteronuclear-coupled spins, inverted (spectrum B) and noninverted (spectrum A). The fractional enrichments [atom percent excess (APE)] of lactate and alanine were calculated from their respective resonances in spectrum A (A) and spectrum B (B) as follows
APE = 0.5 <FENCE><FR><NU><IT>A</IT> − <IT>B</IT></NU><DE><IT>A</IT></DE></FR></FENCE> × 100 − 1.1
Spectra were acquired with a repetition time of 6 s, number of scans of 512, 16 kilobytes of data, and broadband carbon decoupling. A repetition time value of 19 s was used to measure saturation correction factors for calculating metabolite concentrations with respect to a known internal concentration standard (lactate), which was measured in tissue extracts with the use of a 2300 Stat Plus lactate analyzer (Yellow Springs Instruments, Yellow Springs, OH).

Analytic procedures. Plasma glucose concentrations were measured by the glucose oxidase method (Glucose Analyzer II; Beckman Instruments, Fullerton, CA). Plasma immunoreactive free insulin was measured with a double-antibody RIA technique (Linco Research, St. Charles, MO). 13C enrichment of plasma glucose was determined with the use of a Hewlett-Packard 5890 gas chromatograph (HP-1 capillary column, 12-m × 0.2-mm × 0.33-mm film thickness) interfaced to a Hewlett-Packard 5971A mass-selective detector, operating in the positive chemical ionization mode with methane as a reagent gas (6).

[13C]glycogen fractional enrichments were determined with the use of the precipitated glycogen from the initial muscle perchloric acid extraction (99% recovery) (13), and absolute glycogen concentrations were measured on a separate portion of muscle (17).

Skeletal muscle ATP concentrations (31P-NMR experiments) were determined enzymatically with the use of a diagnostic ATP assay kit (no. 366; Sigma Chemical, St. Louis, MO) modified for tissue analysis. Total creatine concentrations in muscle were measured by 13C-NMR analysis performed on a Bruker AM 500 NMR spectrometer system, for which the total [2-13C]creatine peak intensity [54.4 parts per million (ppm)] was referenced to the [2-13C]acetate peak (24.2 ppm), which was added as an internal concentration standard. Spectra were acquired with a repetition time of 1.4 s, number of scans of 10,000, 16 kilobytes of data, and Waltz-16 broadband proton decoupling. Peak intensities were corrected for saturation and nuclear Overhauser effect contributions.

Plasma free fatty acids (FFAs) were determined with the use of an acyl-CoA oxidase-based colorimetric kit (Wako NEFA-C; Wako Pure Chemical Industries, Osaka, Japan). Plasma lactate concentrations were measured with the use of a 2300 Stat Plus lactate analyzer.

Glycogen synthesis rate calculation. The incremental change in C-1 glycogen peak intensity from [1-13C]glucose incorporation was measured at 100.5 ppm. Incremental plasma [13C]glucose fractional enrichment as well as final [13C]glycogen enrichment and concentrations were used to back extrapolate the glycogen concentration (in µmol/g, which represents µmol glucosyl units/g muscle wet wt) at each measured time point to baseline, as described by Bloch et al. (3). Glycogen synthesis rates were determined with the use of a linear regression analysis over the individual-time-point glycogen concentrations.

Glycolytic flux calculations. Metabolic steady-state conditions were assumed for the calculation of carbon flux through the glycolytic pathway into the intermediate triose pool of lactate, pyruvate, and alanine. We have previously shown (12, 13) that these intermediates are at steady-state concentrations before and after a euglycemic- or hyperglycemic-hyperinsulinemic clamp. The 13C label incorporation from [1-13C]glucose into [3-13C]lactate and [3-13C]alanine in the hindlimb muscles was observed by 13C-NMR as an indirect marker of pyruvate labeling. Label incorporation into lactate and alanine is a qualitative indicator of glycolytic flux. Differential equations were developed from steady-state rate equations and solved for glycolytic flux (12).

MCA. A brief introduction to MCA theory is described in the APPENDIX. A top-down approach to MCA was used. In doing so, numerous enzymes associated with a particular metabolic step or pathway can be lumped together so that control coefficients for those groups of enzymes can be determined. In this manner, we determined the control of glucose disposal distributed between glucose transport through insulin-stimulated GLUT-4 and phosphorylation via hexokinase (GLUT-4/HK); glycogen synthesis, consisting of phosphoglucomutase, UDP-glucose pyrophosphorylase, and glycogen synthase enzymes; and glycolysis, consisting of numerous enzymes including phosphofructokinase and pyruvate kinase in skeletal muscle (Fig. 1).


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Fig. 1.   Top-down approach to metabolic control analysis (MCA) to determine flux control for glucose disposal distributed among 3 separate blocks [e.g., glucose transport and phosphorylation (GLUT-4/HK), glycogen synthesis, and glycolysis] in skeletal muscle. J, glucose disposal flux; Vglycogen syn, glycogen synthesis rate; Vglycolysis, glycolytic flux; Glcex, extracellular glucose; G-6-P, glucose 6-phosphate; Ala, alanine; Pyr, pyruvate; Lac, lactate.

The following equations define the system, where C stands for control coefficient, epsilon  is elasticity, Vglycogen syn is glycogen synthesis rate, Vglycolysis is glycolytic flux, and J is glucose disposal flux in skeletal muscle
<IT>C</IT><SUP><IT>J</IT></SUP><SUB>GLUT-4/HK</SUB> + <IT>C</IT><SUP><IT>J</IT></SUP><SUB>glycogen syn</SUB> + <IT>C</IT><SUP><IT>J</IT></SUP><SUB>glycolysis</SUB> = 1
<IT>C</IT><SUP><IT>J</IT></SUP><SUB>GLUT-4/HK</SUB> &egr;<SUP>GLUT-4/HK</SUP><SUB>G-6-<IT>P</IT></SUB> + <IT>C</IT><SUP><IT>J</IT></SUP><SUB>glycogen syn</SUB> &egr;<SUP>glycogen syn</SUP><SUB>G-6-<IT>P</IT></SUB> 
+ <IT>C</IT><SUP><IT>J</IT></SUP><SUB>glycolysis</SUB> &egr;<SUP>glycolysis</SUP><SUB>G-6-<IT>P</IT></SUB> = 0
<FR><NU><IT>C</IT><SUP><IT>J</IT></SUP><SUB>glycogen syn</SUB></NU><DE><IT>V</IT><SUB>glycogen syn</SUB></DE></FR> − <FR><NU><IT>C</IT><SUP><IT>J</IT></SUP><SUB>glycolysis</SUB></NU><DE><IT>V</IT><SUB>glycolysis</SUB></DE></FR> = 0
We measured the elasticities epsilon glycogen synG-6-P and epsilon glycolysisG-6-P by varying G-6-P concentration ([G-6-P]) indirectly via a hyperinsulinemic clamp at different glycemia levels (i.e., euglycemic and hyperglycemic). We thereby maintained control of insulin, which is a potent regulator of HK-II expression and a covalent modulator of glycogen synthase in all three protocols. Concentrations of known allosteric effectors such as ADP, ATP, Mg2+, pH, and Pi were measured with the use of in vivo 31P-NMR. To measure the elasticity of GLUT-4/HK to G-6-P, we maintained external glucose homeostasis as substrate for GLUT-4/HK. It is possible to measure epsilon GLUT-4/HKG-6-P if we can modulate [G-6-P] indirectly by inhibition of glycolysis downstream, for example by increasing lipid oxidation while maintaining external glucose concentrations. This was accomplished with the use of the lipid/heparin infusion. Therefore, we maintained euglycemic-hyperinsulinemic conditions and indirectly modulated [G-6-P] simultaneously. A similar scheme with the use of ketones was used to obtain control coefficients for glucose disposal in heart (16). The control coefficients were calculated by solving the above equations simultaneously.

Statistical analysis. All data are reported as means ± SE. Single-factor ANOVA was performed on data to determine significance at a minimum P < 0.05 threshold among the three protocols. A multiple-comparison Fisher's protected least significant difference post hoc test was used when necessary to determine significance among protocols. Error analysis of control coefficients was calculated with the use of the SE in the flux and [G-6-P] measurements.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

Basal measurements. Basal plasma concentrations of glucose, insulin, and FFA were similar in all three protocols (Table 1).

                              
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Table 1.   Plasma glucose, insulin, and FFA concentrations in 3 protocols

Protocol I. During the euglycemic-hyperinsulinemic clamp experiment, the plasma glucose concentration was maintained at ~6 mM throughout the study, and the plasma insulin concentration increased to 206 ± 43 µU/ml, whereas plasma FFA decreased to 0.4 ± 0.1 mM (Table 1). The glucose infusion rate was stable throughout the clamp period (31.6 ± 2.9 mg · kg-1 · min-1 at 120 min; Table 2). The ATP and total creatine concentrations were 5.1 ± 0.3 and 31.1 ± 7.1 µmol/g, respectively.

                              
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Table 2.   Metabolic flux

Protocol II. During the hyperglycemic-hyperinsulinemic clamp experiment, the plasma glucose concentration increased and was maintained at ~11 mM throughout the study, and the plasma insulin concentration increased to 279 ± 88 µU/ml, whereas plasma FFA decreased to 0.2 ± 0.1 mM (Table 1). The glucose infusion rate was stable throughout the clamp period (49.7 ± 1.0 mg · kg-1 · min-1 at 120 min; Table 2). The ATP and total creatine concentrations were 4.9 ± 0.1 and 27.6 ± 2.2 µmol/g, respectively.

Protocol III. During the euglycemic-hyperinsulinemic-lipid/heparin clamp experiment, the plasma glucose concentration was maintained at ~6 mM throughout the study, and the plasma insulin concentration increased to 229 ± 12 µU/ml, whereas the plasma FFA increased to 2.3 ± 0.4 mM as a result of the lipid/heparin infusion (Table 1). The glucose infusion rate decreased throughout the duration of the clamp period and was 24.0 ± 1.5 mg · kg-1 · min-1 at 120 min (Table 2). The ATP and total creatine concentrations were 5.2 ± 0.1 and 27.8 ± 2.2 µmol/g, respectively.

In vivo 13C-NMR. Figure 2A illustrates a 15-min acquired baseline spectrum (bottom) and a spectrum taken at 120 min (top), when significant 13C label incorporation into metabolite intermediates was achieved. The beta - and alpha -anomer peaks of [1-13C]glucose appear at 96.8 and 93.0 ppm, respectively, and the large peak slightly downfield at 100.5 ppm corresponds to the C-1 glucosyl unit of the glycogen polymer. [3-13C]lactate and [3-13C]alanine can also be observed at 21.0 and 16.9 ppm, respectively.


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Fig. 2.   A: 15-min acquired baseline in vivo 13C-NMR spectrum of rat hindlimb skeletal muscles (bottom). Spectrum at top is indicative of [1-13C]glucose label incorporation into [1-13C]glycogen, [3-13C]lactate, and [3-13C]alanine taken at 120 min of a euglycemic-hyperinsulinemic clamp. [1-13C]glucose [beta -anomer 96.8 parts per million (ppm) and alpha -anomer 93.0 ppm], [1-13C]glycogen (100.5 ppm), [3-13C]lactate (21.0 ppm), and [3-13C]alanine (16.9 ppm) are visible as indicated. B: 15-min acquired baseline in vivo 31P-NMR spectrum of rat hindlimb skeletal muscles. Spectrum at inset illustrates increase in G-6-P (7.16 ppm) at 120 min into a hyperglycemic-hyperinsulinemic clamp. Intracellular Pi (Piin) at 4.9 ppm, phosphocreatine (PCr) at 0 ppm, and gamma -, alpha -, and beta -ATP can be observed at -2.4, -7.5, and -16.0 ppm, respectively. Piex, extracellular Pi.

The glycolytic flux and glycogen synthesis rate measurements shown in Table 2 originated from our previously published work (12-14). From these results, it is evident that the decreased glucose disposal observed during the euglycemic-hyperinsulinemic-lipid/heparin clamp was primarily the result of an ~69% inhibition of glycolysis vs. that of the euglycemic-hyperinsulinemic clamp. The increased glucose disposal during the hyperglycemic-hyperinsulinemic clamp, however, was primarily due to a large increase in glycogen synthesis rate with little increase in glycolytic flux vs. that of the euglycemic-hyperinsulinemic clamp.

In vivo 31P-NMR. Figure 2B illustrates a 31P-NMR baseline spectrum with G-6-P, Pi (both extracellular and intracellular), PCr (assigned to 0 ppm), and alpha -, beta -, and gamma -ATP indicated. The basal [G-6-P] values were the same in all groups before the three clamp protocols, as indicated in Table 3. The incremental change in G-6-P during the hyperglycemic-hyperinsulinemic clamp can be seen in Fig. 2B (inset). [G-6-P] in all three protocol groups increased at 105-135 min, as presented in Table 3, with [G-6-P] higher in the hyperglycemic-hyperinsulinemic and euglycemic-hyperinsulinemic-lipid/heparin clamps vs. euglycemic-hyperinsulinemic clamp (265 ± 12 and 251 ± 9 vs. 217 ± 8 nmol/g; P < 0.005 and P < 0.05, respectively). Concentrations of Pi, PCr, ADP, Mg2+, and pH were not significantly different at baseline or at 120 min in all three clamp protocols (Table 3).

                              
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Table 3.   31P-NMR data

Metabolic flux control analysis. With the use of the top-down approach to calculate the elasticities associated with glucose disposal, the resulting elasticities for glucose transport/phosphorylation, glycogen synthesis, and glycolysis with respect to G-6-P were as follows: epsilon GLUT-4/HKG-6-P -2.18, epsilon glycogen synG-6-P = 3.56, and epsilon glycolysisG-6-P = 0.69, respectively (see APPENDIX). From these elasticity measurements, we were able to calculate the control coefficients by solving the three equations described above simultaneously. The majority of control of glucose disposal during hyperinsulinemia was determined to be at the glucose transport/phosphorylation step (CJGLUT-4/HK = 0.55 ± 0.10), with approximately one-half of the control distributed between glycogen synthesis (CJglycogensyn = 0.30 ± 0.06) and glycolysis (CJglycolysis = 0.15 ± 0.02).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

MCA theory was developed as an amalgamation of independent work in the early 1970s by Kacser and Burns (15) and Heinrich and Rapoport (10). This theory was formulated to determine how the intermediate steps of a pathway react to a variable that might affect it (i.e., a change in substrate and/or external effector amount or change in enzyme expression). In this manner, a system of intermediary steps can be analyzed as a whole to determine fractional contribution of any one step to the overall control of flux through a metabolic pathway. Because we cannot obtain relative information with regard to the way in which each individual enzyme in a metabolic pathway reacts to variables affecting it (bottom-up approach) from NMR metabolic flux measurements, we used a top-down approach to MCA, in which we can define control over only a few metabolic components, each consisting of a number of enzymes (e.g., glucose transport/phosphorylation, glycogen synthesis, and glycolysis). In our studies, three separate infusion protocols were necessary to indirectly modulate [G-6-P] to measure the elasticities of these three metabolic components to G-6-P (i.e., epsilon GLUT-4/HKG-6-P, epsilon glycogen synG-6-P, and epsilon glycolysisG-6-P).

It was necessary to maintain concentrations of known allosteric effectors and covalent modulation of enzymes that make up these pathways. Because the 13C-NMR measurement of glycolytic flux at euinsulinemia might not be possible (12), we maintained hyperinsulinemic conditions in all three groups, and no significant differences among groups with respect to concentrations of ADP, Mg2+, Pi, or pH were measured (Table 3). Although it has been shown that an acute elevation of plasma FFA can directly regulate insulin signaling or glucose transporter function, these events generally occur only after 3-5 h (9, 23). Additionally, we (13) and others (9) have shown that [G-6-P] values remained elevated during the initial 3-4 h of clamp before elevated plasma FFA could affect insulin signaling or glucose transporter function and result in a decrease in [G-6-P]. Therefore, with proper control of the physiological effectors of these metabolic pathways, one can couple MCA to in vivo metabolic studies in skeletal muscle.

MCA is a logical extension of traditional studies that attempted to define a rate-limiting enzyme in the pathway but were unsuccessful. For example, in heart, when intracellular glucose was not detected as extracellular concentrations varied from 2 to 16 mM, this led to the belief that glucose transport was rate limiting for glucose disposal (24). In recent heart work, MCA was used to illustrate that the control of glucose uptake is distributed among glucose transport and phosphorylation, glycogen synthesis, and glycolysis (16). Additionally, the distribution of control was shown to change, depending on substrate availability or hormonal environment.

In skeletal muscle, it has been suggested that the glycogen synthase enzyme controls the rate of glycogen synthesis (4, 21). On the other hand, insulin stimulation has been shown to increase glucose transport via an increase in GLUT-4 translocation to the cell membrane (19). The glycogen synthase enzyme is additionally regulated by insulin by reduction of its degree of phosphorylation. With the unique capability to measure net glycogen synthesis flux and intermediate substrate (G-6-P) by NMR and with the use of a top-down approach, it was shown that glucose transport/phosphorylation could indeed account for almost the entire control of insulin-stimulated glycogen synthesis in normal or diabetic humans (27, 31). In these analyses, control of glycogen synthesis was distributed over glucose transport/phosphorylation and glycogen synthesis only while glycolysis was neglected. This was thought reasonable in light of whole body extrapolation of skeletal muscle glycogen synthesis rates determined by NMR that suggested that glycogen synthesis in skeletal muscle accounts for ~90% of the whole body glucose metabolism under hyperglycemic-hyperinsulinemic conditions (32).

However, through MCA, it has been theoretically shown that glycolytic enzymes do indeed contribute to the control of [G-6-P] (30). Additionally, in rat, glycolysis accounts for a significant portion of the glucose disposal in muscle under euglycemic-hyperinsulinemic conditions (18, 28). Therefore, the potential for glycogen synthesis and glycolysis to account for a significant fraction of control in glucose disposal is evident.

The control of glucose transport/phosphorylation when partitioned into individual flux components of glucose transport and HK activity can be determined if elasticities of these flux components to intracellular glucose are known. We have previously determined that intramuscular glucose is negligible in rat during a hyperglycemic-hyperinsulinemic clamp by using an NMR spectroscopic assay (5), and, therefore, it would most likely be negligible during a euglycemic-hyperinsulinemic clamp. Consequently, we would not have the NMR sensitivity to detect the required modulation of intracellular glucose necessary for calculating elasticities in our experiments. Hypothetically, if intracellular glucose were negligible during both euglycemic- and hyperglycemic-hyperinsulinemic clamps, then, with the use of a bottom-up approach to MCA in which known enzyme kinetic parameters are used, the control of HK would be negligible as well (29).

Insulin sensitivity varies with skeletal muscle fiber composition (11); thus there is the potential for differences in the distribution of control in slow- vs. fast-twitch muscle fibers. For example, it has been shown that the regulation of glycogen synthase is different in fast- vs. slow-twitch muscle after a glucose load (34). Because of the intrinsic sensitivity of the in vivo NMR measurement, it is not possible to make glycolytic and glycogen synthesis flux measurements in small muscles such as soleus and extensor digitorum longus. Therefore, the hindlimb placement over the NMR-sensitive volume of the radio frequency (RF) coil was such that the NMR signal obtained was predominantly from the larger mixed fiber tissue composing the gastrocnemius and biceps femoris muscles. Additionally, it must be noted that the distribution of control might vary depending on the physiological conditions (16, 30).

In conclusion, we have presented a study that takes unique advantage of the in vivo NMR measurements of glycogen synthesis and glycolytic flux and [G-6-P] to apply a top-down analysis of MCA to determine the control of insulin-stimulated glucose disposal in skeletal muscle distributed among glucose transport/phosphorylation, glycogen synthesis, and glycolysis. It was determined that during insulin stimulation, the majority of control resides at glucose transport/phosphorylation, although glycogen synthesis and glycolysis do share in the control of glucose disposal as well. This approach might be useful in characterizing the control of these pathways under conditions of insulin resistance and/or diabetes.


    APPENDIX
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

MCA Theory

The following is a brief description of MCA theory (15). The flux control coefficient is defined by the ratio of fractional change of the flux through an enzyme in the pathway to fractional change in enzyme concentration (E)
<IT>C</IT><SUP><IT>J</IT></SUP><SUB><IT>a</IT></SUB> = <FR><NU>∂<IT>J</IT></NU><DE><IT>J</IT></DE></FR> <FENCE> </FENCE> <FR><NU>∂<IT>E</IT><SUB><IT>a</IT></SUB></NU><DE><IT>E</IT><SUB><IT>a</IT></SUB></DE></FR>
where J is the flux through enzyme a, and CJa is the control coefficient for that enzyme. All of the control coefficients of a pathway are expressible in terms of elasticities. Elasticities are properties of individual enzymes that relate velocities of enzymes with their substrate concentrations in a relationship similar to control coefficients
&egr;<SUP><IT>a</IT></SUP><SUB><IT>S</IT></SUB> = <FR><NU>∂<IT>v</IT><SUB><IT>a</IT></SUB></NU><DE><IT>v<SUB>a</SUB></IT></DE></FR> <FENCE> </FENCE> <FR><NU>∂<IT>S</IT></NU><DE><IT>S</IT></DE></FR>
where va is the velocity of enzyme a, which is equal to the flux J through the enzyme in a steady-state system. S is the substrate for enzyme a, and epsilon aS is the elasticity coefficient for that enzyme.

All of the enzymes or metabolic pathways that can affect the flux through a metabolic system share control of that flux; thus, via the summation theorem, the sum of all of the coefficients equals one, i.e., <LIM><OP>∑</OP><LL><IT>i</IT>=1</LL><UL><IT>n</IT></UL></LIM>CJi = 1. If a metabolite is both a substrate and product to separate enzymes in a two-enzyme system, then the use of the connectivity theorem to relate the flux control coefficients to the kinetic properties of the enzyme gives us
<IT>C</IT><SUP><IT>J</IT></SUP><SUB>1</SUB> &egr;<SUP>1</SUP><SUB><IT>S</IT></SUB> + <IT>C</IT><SUP><IT>J</IT></SUP><SUB>2</SUB> &egr;<SUP>2</SUP><SUB><IT>S</IT></SUB> = 0
For this relationship to hold, all other allosteric effectors of the enzymes must remain constant. Additionally, the branch point theorem (7) states that the ratio of the sum of the flux control coefficients of enzymes in the branches is equal to the ratio of flux through the branches
<FR><NU><IT>C</IT><SUP><IT>J</IT><SUB>1</SUB></SUP><SUB>1</SUB></NU><DE><IT>C</IT><SUP><IT>J</IT><SUB>2</SUB></SUP><SUB>2</SUB></DE></FR> = <FR><NU><IT>v</IT><SUB>1</SUB></NU><DE><IT>v</IT><SUB><IT>2</IT></SUB></DE></FR>
Therefore, because the control coefficients of a pathway are expressible in terms of elasticities, relative fluxes, and substrate concentrations, we can use all of the above theorems together to indirectly calculate control coefficients. This can be accomplished by calculating simultaneous equations for the entire system of enzymes representing a metabolic pathway. Because we cannot define the entire system of enzymes consisting of the metabolic pathway (because of limitations of our NMR flux measurements), we must use a top-down approach whereby we assign control coefficients to blocks of enzymes contributing to the flux of a pathway measured with the use of NMR (Fig. 1). The differential flux and G-6-P data used to calculate the enzyme elasticities in the present experiment are tabulated in Table 4.

                              
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Table 4.   Differential flux and G-6-P data


    ACKNOWLEDGEMENTS

We are indebted to Veronika Walton, Laura Burdon, and Kevin Cadman for expert technical assistance. We are grateful to electrical engineers Terry Nixon and Scott Mcyntire for NMR technical improvements and Peter Brown for radio frequency antenna design and construction. We also thank Dr. Douglas L. Rothman for helpful discussions on MCA.


    FOOTNOTES

This study was supported by grants from the National Institute of Diabetes and Digestive and Kidney Diseases (RO1-DK-40936 and P30-DK-45735), the American Diabetes Association (Mentor-Based Postdoctoral Fellowship to B. M. Jucker), and an unrestricted grant from Bristol-Myers Squibb.

G. I. Shulman is an investigator for the Howard Hughes Medical Institute.

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: B. M. Jucker, Dept. of Internal Medicine, Yale Univ. School of Medicine, PO Box 208020, Fitkin 1, 333 Cedar St., New Haven, CT 06520-8020 (E-mail: Jucker{at}mrclin1.med.yale.edu).

Received 9 November 1998; accepted in final form 21 April 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
APPENDIX

1.   Altura, B. M., A. Gebrewold, A. Zhang, B. T. Altura, and R. K. Gupta. Short-term reduction in dietary intake of magnesium causes deficits in brain intracellular free Mg2+ and [H+]i but no high-energy phosphates as observed by in vivo 31P-NMR. Biochim. Biophys. Acta 1358: 1-5, 1997[Medline].

2.   Bloch, G., J. R. Chase, M. J. Avison, and R. G. Shulman. In vivo 31P NMR measurement of glucose-6-phosphate in the rat muscle after exercise. Magn. Reson. Med. 30: 347-350, 1993[Medline].

3.   Bloch, G., J. R. Chase, D. B. Meyer, M. J. Avison, G. I. Shulman, and R. G. Shulman. In vivo regulation of rat muscle glycogen resynthesis after intense exercise. Am. J. Physiol. 266 (Endocrinol. Metab. 29): E85-E91, 1994[Abstract/Free Full Text].

4.   Bogardus, C., S. LiDioja, K. Stone, and D. Mott. Correlation between muscle glycogen synthase activity and in vivo insulin action in man. J. Clin. Invest. 73: 1185-1190, 1984[Medline].

5.   Cline, G. W., B. M. Jucker, Z. Trajanoski, A. J. M. Rennings, and G. I. Shulman. A novel 13C NMR method to assess intracellular glucose concentration in muscle, in vivo. Am. J. Physiol. 274 (Endocrinol. Metab. 37): E381-E389, 1998[Abstract/Free Full Text].

6.   Cline, G. W., and G. I. Shulman. Quantitative analysis of the pathways of glycogen repletion in periportal and perivenous hepatocytes in vivo. J. Biol. Chem. 266: 4094-4098, 1991[Abstract/Free Full Text].

7.   Fell, D. A., and H. M. Sauro. Metabolic control and its analysis. Eur. J. Biochem. 148: 555-561, 1985[Abstract].

8.   Gupta, R. K., J. L. Benovic, and Z. B. Rose. The determination of the free magnesium level in the human red blood cell by 31P NMR. J. Biol. Chem. 253: 6172-6176, 1978[Abstract].

9.   Hawkins, M., N. Barzilai, R. Liu, M. Hu, W. Chen, and L. Rossetti. Role of the glucosamine pathway in fat-induced insulin resistance. J. Clin. Invest. 99: 2173-2182, 1997[Abstract/Free Full Text].

10.   Heinrich, R., and T. A. Rapoport. A linear steady-state treatment of enzymatic chains. General properties, control and effector strength. Eur. J. Biochem. 42: 89-105, 1974[Medline].

11.   James, D. E., A. B. Jenkins, and E. W. Kraegen. Heterogeneity of insulin action in individual muscles in vivo: euglycemic clamp studies in rats. Am. J. Physiol. 248 (Endocrinol. Metab. 11): E567-E574, 1985[Abstract/Free Full Text].

12.   Jucker, B. M., A. J. M. Rennings, G. W. Cline, K. F. Petersen, and G. I. Shulman. In vivo NMR investigation of intramuscular glucose metabolism in conscious rats. Am. J. Physiol. 273 (Endocrinol. Metab. 36): E139-E148, 1997[Abstract/Free Full Text].

13.   Jucker, B. M., A. J. M. Rennings, G. W. Cline, and G. I. Shulman. 13C and 31P NMR studies on the effects of increased plasma free fatty acids on intramuscular glucose metabolism in the awake rat. J. Biol. Chem. 272: 10464-10473, 1997[Abstract/Free Full Text].

14.   Jucker, B. M., G. W. Cline, N. Barucci, and G. I. Shulman. Differential effects of safflower oil versus fish oil feeding on insulin stimulated glycogen synthesis, glycolysis, and pyruvate dehydrogenase flux in skeletal muscle: a 13C NMR study. Diabetes 48: 134-140, 1999[Abstract].

15.   Kacser, H., and J. A. Burns. The control of flux. Symp. Soc. Exp. Biol. 27: 65-104, 1973[Medline].

16.   Kashiwaya, Y., K. Sato, N. Tsuchiya, S. Thomas, R. L. Veech, and J. V. Passonneau. Control of glucose utilization in working perfused rat heart. J. Biol. Chem. 269: 25502-25514, 1994[Abstract/Free Full Text].

17.  Keppler, D., and K. Decker. Glycogen: Determination with Amyloglucosidase. Methods of Enzymatic Analysis, edited by H. U. Bergmeyer. New York: Verlag Chemie Weinheim, Academic1974, p. 1127-1131.

18.   Kim, J. K., J. K. Wi, and J. H. Youn. Plasma free fatty acids decrease insulin-stimulated skeletal muscle glucose uptake by suppressing glycolysis in conscious rats. Diabetes 45: 446-453, 1996[Abstract].

19.   Klip, A., and M. R. Paquet. Glucose transport and glucose transporters in muscle and their metabolic regulation. Diabetes Care 13: 228-243, 1990[Abstract].

20.   Krebs, H. A. Cyclic processes in living matter. Enzymologia 12: 88-100, 1946.

21.   Kubo, K., and J. E. Foley. Rate-limiting steps for insulin-mediated glucose uptake into perfused rat hindlimb. Am. J. Physiol. 250 (Endocrinol. Metab. 13): E100-E102, 1986[Abstract/Free Full Text].

22.   Lawson, J. W. R., and R. L. Veech. Effects of pH and free Mg2+ on the Keq of the creatine kinase reaction and other phosphate hydrolyses and phosphate transfer reactions. J. Biol. Chem. 254: 6528-6537, 1979[Abstract].

23.   Marcucci, M., M. Griffin, P. Estrada, N. Barucci, G. Cline, and G. I. Shulman. Elevations in free fatty acids induce insulin resistance via inhibition of IRS-1 associated PI-3 kinase activity in vivo (Abstract). Diabetes 47: A284, 1998.

24.   Neely, J. R., H. Liebermeister, and H. E. Morgan. Effect of pressure development on membrane transport of glucose in isolated rat heart. Am. J. Physiol. 212: 815-822, 1967[Medline].

25.   Newsholme, E. A, and C. Start. Regulation in Metabolism. London: Wiley and Sons, 1973.

26.   Rognstad, R. Rate-limiting steps in metabolic pathways. J. Biol. Chem. 254: 1875-1878, 1979[Abstract].

27.   Rothman, D. L., R. G. Shulman, and G. I. Shulman. 31P nuclear magnetic resonance measurements of muscle glucose-6-phosphate. Evidence for reduced insulin-dependent muscle glucose transport or phosphorylation activity in non-insulin-dependent diabetes mellitus. J. Clin. Invest. 89: 1069-1075, 1992[Medline].

28.   Rossetti, L., and A. Giaccari. Relative contribution of glycogen synthesis and glycolysis to insulin-mediated glucose uptake. A dose-response euglycemic clamp study in normal and diabetic rats. J. Clin. Invest. 85: 1785-1792, 1990[Medline].

29.   Roussel, R., P. G. Carlier, J.-J. Robert, G. Vehlo, and G. Bloch. 13C/31P NMR studies of glucose transport in human skeletal muscle. Proc. Natl. Acad. Sci. USA 95: 1313-1319, 1998[Abstract/Free Full Text].

30.   Schulz, A. R. Control analysis of muscle glycogen metabolism. Arch. Biochem. Biophys. 353: 172-180, 1998[Medline].

31.   Shulman, R. G., G. Bloch, and D. L. Rothman. In vivo regulation of muscle glycogen sythase and the control of glycogen synthesis. Proc. Natl. Acad. Sci. USA 92: 8535-8542, 1995[Abstract].

32.   Shulman, G. I., D. L. Rothman, T. Jue, P. Stein, R. A. DeFronzo, and R. G. Shulman. Quantitation of muscle glycogen synthesis in normal subjects and subjects with non-insulin-dependent diabetes by 13C nuclear magnetic resonance spectroscopy. N. Engl. J. Med. 322: 223-228, 1990[Abstract].

33.   Smith, D., L. Rossetti, E. Ferrannini, C. M. Johnson, L. Cobelli, G. Toffolo, L. D. Katz, and R. A. DeFronzo. In vivo glucose metabolism in the awake rat: tracer and insulin clamp studies. Metabolism 36: 1167-1174, 1987[Medline].

34.   Sugden, M. C., M. J. Holness, and L. G. D. Fryer. Differential regulation of glycogen synthase by insulin and glucose in vivo in skeletal muscles of the rat. Am. J. Physiol. 273 (Endocrinol. Metab. 36): E479-E487, 1997[Abstract/Free Full Text].

35.   Taylor, D. J., P. Styles, P. M. Matthews, D. A. Arnold, D. G. Gadian, P. Bore, and G. K. Radda. Energetics of human muscle: exercise-induced ATP depletion. Magn. Reson. Med. 3: 44-54, 1986[Medline].


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