Departments of 1 Internal Medicine and 2 Diagnostic Radiology and the 3 Howard Hughes Medical Institute, Yale University School of Medicine, New Haven, Connecticut 06510
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ABSTRACT |
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To examine the mechanism by which muscle
glycogen limits its own synthesis, muscle glycogen and glucose
6-phosphate (G-6-P) concentrations were measured in seven
healthy volunteers during a euglycemic (~5.5 mM)-hyperinsulinemic
(~450 pM) clamp using 13C/31P nuclear
magnetic resonance spectroscopy before and after a muscle glycogen
loading protocol. Rates of glycogen synthase
(Vsyn) and phosphorylase
(Vphos) flux were estimated during a
[1-13C]glucose (pulse)-unlabeled glucose (chase)
infusion. The muscle glycogen loading protocol resulted in a 65%
increase in muscle glycogen content that was associated with a twofold
increase in fasting plasma lactate concentrations (P < 0.05
vs. basal) and an ~30% decrease in plasma free fatty acid
concentrations (P < 0.001 vs. basal). Muscle glycogen
loading resulted in an ~30% decrease in the insulin-stimulated rate
of net muscle glycogen synthesis (P < 0.05 vs. basal),
which was associated with a twofold increase in intramuscular
G-6-P concentration (P < 0.05 vs. basal). Muscle
glycogen loading also resulted in an ~30% increase in whole body
glucose oxidation rates (P < 0.05 vs. basal), whereas there was no effect on insulin-stimulated rates of whole body glucose uptake
(~10.5 mg · kg body
wt1 · min
1 for both clamps)
or glycogen turnover
(Vsyn/Vphos was ~23% for both clamps). In conclusion, these data are consistent with the hypothesis that glycogen limits its own synthesis through feedback inhibition of glycogen synthase activity, as reflected by an
accumulation of intramuscular G-6-P, which is then shunted into
aerobic and anaerobic glycolysis.
nuclear magnetic resonance spectroscopy; glycogen turnover; glycogen synthase
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INTRODUCTION |
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IT IS WELL ESTABLISHED that, despite wide variations in carbohydrate intake, muscle glycogen content remains remarkably stable between 60 and 90 mmol/l muscle (1, 12, 19, 25, 30) and rarely exceeds 130 mmol/l muscle in humans, even under conditions that should promote glycogen synthesis (1, 20, 25). However, the step or process by which glycogen limits its own synthesis is unknown. Glycogen could limit its own synthesis by feedback inhibition of 1) glucose transport, 2) hexokinase, or 3) glycogen synthase activity (3, 4, 7, 14, 25, 26, 28, 31). It is also possible that glycogen could limit its own synthesis through promotion of glycogen turnover where an increase in glycogen content induces an increase in phosphorylase activity (19), which in turn results in an increase in glycogen cycling. Muscle glycogen cycling has been observed previously both at rest (4) and during prolonged low-intensity exercise (24). To examine this question, rates of muscle glycogen synthesis and glycogen phosphorylase flux were measured simultaneously using 13C nuclear magnetic resonance (NMR) spectroscopy (4) under euglycemic-hyperinsulinemic conditions before and after a classic muscle glycogen loading protocol (1). This protocol is commonly used by endurance athletes to enhance performance and has been shown to increase muscle glycogen content by ~60% (1). In addition, 31P NMR spectroscopy was used to monitor changes in muscle glucose 6-phosphate (G-6-P) concentration to assess rate-controlling steps in insulin-stimulated muscle glycogen synthesis (28).
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METHODS |
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Subjects
Seven nonsmoking male volunteers (mean age 28 ± 1 yr, mean body wt 73 ± 3 kg, mean body mass index 23.6 ± 0.8 kg/m2) without a family history of diabetes mellitus, hypertension, or any major diseases were studied, and none of the subjects was taking any medications. Subjects were instructed to abstain from strenuous physical activity for at least 3 days before the first clamp study and during the loading protocol. Experimental procedures were all approved by the Yale University Human Investigation Committee. Each subject gave informed consent after the purpose, nature, and potential risks of the study were explained.Experimental Protocol
Muscle carbohydrate (CHO) metabolism was assessed at baseline and again the day after the loading protocol was completed by using 13C/31P NMR spectroscopy under euglycemic-hyperinsulinemic clamp conditions (Fig. 1). The loading protocol consisted of 60 min of exercise, 3 days of a low-CHO/high-fat diet, a second 60-min bout of exercise, and 4 days of a high-CHO/low-fat diet, as described previously (1). The exercise consisted of running on a treadmill for 55 min (Jogging Machine J2; Tuntur) and performing toe raises with both legs for the last 5 min. The workload for the running session was previously determined to reach ~75% of each subject's maximal oxygen consumption (heart rate ~160 beats/min). The daily caloric intake for both diets was 210 kJ/kg body wt with a ratio of protein-fat-carbohydrate equal to 33:57:10 for the low-CHO diet and 7.5:2.5:90 for the high-CHO diet.
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Euglycemic-hyperinsulinemia was achieved with the glucose-insulin clamp
technique (8). On the evening before each clamp study, participants
were admitted to the General Clinical Research Center of the Yale/New
Haven Hospital and were fasted overnight (10-12 h). A Teflon
catheter was inserted in an antecubital vein in each arm for blood
drawing and for infusions. At time 0, insulin was administered
as a prime (100 pmol/kg)-continuous (6 pmol · kg1 · min
1)
infusion to raise plasma insulin concentration to ~450 pM and to
maintain that concentration for the duration of the study. At the same
time, a variable infusion of [1-13C]glucose (~20%
enriched) was begun to maintain plasma glucose levels at ~5.5 mmol/l.
After 120 min, the infusion was switched to unlabeled glucose and was
continued for another 100-120 min (chase period). Blood samples
for measurements of plasma glucose, insulin, glucagon, lactate, and
free fatty acid concentrations and plasma [3C]glucose
enrichment were obtained at 5- to 30-min intervals until completion of
the study.
Indirect Calorimetry
Continuous indirect calorimetry was performed to determine rates of total body glucose and lipid oxidation at baseline and at 100-120 min and 220-240 min into each clamp study, as previously described (29).In Vivo NMR Spectroscopy
Natural abundance 13C/31P NMR spectroscopy was performed in an interleaved fashion at 4.7 T on a Bruker Biospec (Billerica, MA) spectrometer with a 30-cm-diameter magnet bore, as previously described (23). During the measurements, subjects remained supine with the right leg positioned within the homogenous volume of the magnet and with the lower portion of that leg resting on the stage of a radiofrequency (RF) probe. The spectrometer was equipped with a modified RF relay that allowed the hardware to switch the RF power between the 13C (50.4 MHz) and 31P (81.1 MHz) channels with a 10-µs switching time. A 5.1-cm-diameter circular 13C-31P double-tuned surface coil RF probe was used for interleaved acquisitions. The double-tuned circuit was optimized for the 31P channel so that the NMR sensitivity would be enhanced to detect G-6-P. Shimming, imaging, and 1H decoupling at 200.4 MHz were performed with a 9 × 9-cm series butterfly coil. Proton water line widths were shimmed to <50 Hz. A microsphere containing 13C and 31P reference standards was fixed at the center of the double-tuned RF coil for calibration of RF pulse widths. Subjects were positioned by an image-guided localization routine that used a T1-weighted gradient-echo image (repetition time = 82 ms, echo time = 21 ms). The subject's lower leg was positioned so that the isocenter of the magnetic field was ~1 cm in the medial head of the gastrocnemius muscle. By determining the 180° flip angles at the center of the observation coil from the microsphere standard, RF pulse widths were set so that the 90° pulse was sent to the center of the muscle. This maximized suppression of the lipid signal that arises from the subcutaneous fat layer and optimized signal from the muscle.On interleaved 1H decoupled 13C-31P RF pulse sequence was designed so that 72 31P transients were acquired during the same period that 2,736 13C transients were obtained (38 13C scans/31P relaxation period), and free induction decays were saved separately in two blocks. The repetition time for 31P acquisition was 4.6 s to allow for the long T1 of 31P resonance. Power deposition, assessed by magnetic vector potential specific absorption rate calculation, was <4 W/kg. The total scan time for each interleaved spectrum was 5.5 min.
Intramuscular glycogen concentrations were determined by comparison with an external standard solution (150 mM glycogen + 50 mM KCl) in a cast of a leg that electrically loaded the RF coil to the same extent as the subject's leg (23, 27). 13C spectra were processed by methods that have been described previously (23, 27). Briefly, Gaussian-broadened spectra (30 Hz) were baseline corrected ±500 Hz on either side of the [1-13C]glycogen resonance of both subject spectra and standard spectra. Peak areas were then assessed ±200 Hz about the resonance. The 13C NMR technique for assessing intramuscular glycogen concentrations has been validated in situ in frozen rabbit muscle (11) and by comparison with human gastrocnemius muscle biopsies (30).
Concentrations of phosphorylated compounds were calculated from
31P NMR spectra as described previously (28). The area of
the -ATP resonance peak was used as an internal concentration
standard assuming a constant concentration of 5.5 mM for resting muscle (12). Chemical shifts are referenced to phosphocreatine at 0.00 ppm.
The resonance for G-6-P is in a region of the 31P
NMR spectrum in close proximity to other phosphomonoester resonances. Any potential contribution from other phosphomonoester resonances, which have chemical shifts upfield (lower ppm) to G-6-P, was
minimized by integrating over the chemical shift range of the
downfield half of the G-6-P resonance (7.43-7.13 ppm)
and multiplying by two, as described previously (27, 28). The
G-6-P measurement has been validated in an animal model
(2).
Calculations
During the euglycemic-hyperinsulinemic clamp study, the increment in muscle glycogen concentration ([
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Analytical Procedures
Plasma glucose was measured by the glucose oxidase method using a Beckman glucose analyzer (Fullerton, CA). Plasma immunoreactive insulin and glucagon were measured using commercially available double-antibody RIA kits [insulin (Diagnostic Systems Laboratories, Webster, TX); glucagon (Linco Research, St. Charles, MO)]. Plasma lactate concentrations were measured by the lactate dehydrogenase method. Plasma free fatty acids (FFA) were measured using a microfluorometric assay. 13C atom percent enrichment of plasma glucose was determined by gas chromatography-mass spectrometry, as described previously (29).Statistics
All values are expressed as means ± SE. A one-way ANOVA with repeated measurements was used to analyze time course changes in plasma substrate and hormone concentrations throughout each clamp period. A two-way ANOVA with repeated measures was used to analyze time course differences in plasma substrate/hormone concentrations between both studies. When significant changes were obtained over time, post hoc comparisons were made using a paired t-test. Pairwise comparisons for metabolic flux measurements were made using a paired t-test. All data are expressed as means ± SE. ![]() |
RESULTS |
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Muscle Glycogen Loading
Basal muscle glycogen concentrations before both clamp studies are depicted in Fig. 2. Basal muscle glycogen concentrations were similar before the first clamp study and on day 1 of the loading protocol (73 ± 2 and 72 ± 9 mmol/l muscle, respectively). Performing 1 h of running on the treadmill caused glycogen concentration to decrease by 44 ± 5 mmol/l muscle on day 1 (P < 0.001 vs. preexercise). Muscle glycogen content then returned to close to preexercise levels after 3 days of the low-CHO/high-fat diet, although muscle glycogen content still remained slightly lower than baseline (day 4: 62 ± 2 mmol/l muscle, P < 0.01 vs. baseline). Performing the second bout of exercise resulted in glycogen depletion to a similar extent as in the first exercise bout (
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Effects of CHO Loading on Basal Metabolism
Plasma glucose, insulin, and glucagon concentrations were similar before the first clamp study (glucose, 89 ± 2 mg/dl; insulin, 48 ± 6 pM; glucagon, 47 ± 2 pg/ml) and after glycogen loading (glucose, 89 ± 1 mg/dl; insulin, 48 ± 6 pM; glucagon, 45 ± 3 pg/ml). In contrast, a significant increase in 12-h fasting concentrations of plasma lactate (first clamp, 0.43 ± 0.14 mM vs. second clamp, 0.81 ± 0.14 mM; P < 0.05) and a significant decrease in plasma FFA concentration (first clamp, 551 ± 51 µM vs. second clamp, 380 ± 24 µM, P < 0.001) were found on completion of the glycogen-loading protocol. This was paralleled by an increase in the basal whole body glucose oxidation rate (first clamp, 0.41 ± 0.16 mg · kg body wtEuglycemic-Hyperinsulinemic Clamps
Plasma glucose, insulin, glucagon, lactate, and FFA concentrations.
Plasma glucose concentrations were maintained within a range of
90-110 mg/dl throughout both clamp studies. Plasma insulin concentrations increased rapidly and reached steady-state values within
15 min [time = 30-210 min; first clamp, 440 ± 13 pM; second clamp, 447 ± 13 pM), whereas plasma glucagon concentrations
remained at baseline levels during both clamp studies. During the
measurement of glycogen synthase and phosphorylase fluxes (60-120
and 140-210 min, respectively), there was no difference in the
mean glucose infusion rate between the two studies (Table
1). During the glucose infusion, plasma
lactate concentrations rose significantly (P < 0.0001 vs.
baseline) and were significantly higher throughout the glycogen-loaded
clamp study (90-210 min, 1.13 ± 0.19 mM, P < 0.05
vs. baseline clamp) compared with the first clamp study (90-210
min, 0.73 ± 0.16 mM). Plasma FFA concentrations decreased to
similar levels during each clamp study (first clamp, 124 ± 4 µM;
second clamp, 140 ± 13 µM).
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Flux measurements.
Vsyn were linear in both studies (first clamp,
r = 0.99 ± 0.01; second clamp, r =
0.98 ± 0.01). Vsyn was suppressed by 23% in
the glycogen-loaded study compared with the first clamp
(0.127 ± 0.031 vs. 0.165 ± 0.029 mmol · l
muscle1 · min
1,
P < 0.01; Fig. 3).
Vphos was slightly but not significantly lower
in the second clamp study (0.029 ± 0.005 vs. 0.040 ± 0.010 mmol · l
muscle
1 · min
1, not
significant). Because muscle glycogen turnover
(Vphos/Vsyn) was similar in
the two studies (first clamp, 23 ± 3%; second clamp, 28 ± 3%), the lower rate of net glycogen synthesis (
29%,
P < 0.05 vs. baseline clamp study) observed in the
glycogen-loaded studies could be attributed to a marked reduction in
Vsyn. Despite lower rates of insulin-stimulated
muscle glycogen synthesis, rates of insulin-stimulated whole body
glucose metabolism were unchanged in the muscle glycogen-loaded
studies. Assuming that muscle mass is equivalent to ~26% body wt
(5), the contribution of muscle glycogen net synthesis accounted for
less of nonoxidative glucose metabolism (first clamp, 75 ± 4% vs.
second clamp, 51 ± 11%, P < 0.05) in the
presence of supercompensated muscle glycogen levels.
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Phosphorylated metabolites.
Basal intramuscular concentrations of G-6-P were 2.2-fold
higher after the glycogen loading protocol (first clamp, 178 ± 21 µM vs. second clamp, 393 ± 66 µM, P < 0.05). Under
euglycemic-hyperinsulinemic conditions, intramuscular G-6-P
concentrations rose by a similar increment in both clamp studies (first
clamp, 85 ± 12 µM; second clamp,
101 ± 46 µM) but
remained higher (P < 0.001 vs. baseline study) in absolute
concentrations throughout the glycogen-loaded clamp study (Fig.
4). Finally, although no change in muscle
pH was observed during the first clamp study, a slight but significant decrease in pH was noted during the second clamp study (
0.04 pH
units vs. first clamp study, P < 0.05). Glycogen loading
had no detectable effect on intramuscular concentrations in
phosphocreatine (~20 mM) or inorganic phosphate (~3.3 mM).
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DISCUSSION |
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In this study, we found that a 1.6-fold increase in muscle glycogen content to ~130 mmol/l muscle resulted in an ~30% reduction in the rate of insulin-stimulated net muscle glycogen synthesis. This reduction in insulin-stimulated muscle glycogen synthesis could occur through a reduction in glucose transport, hexokinase, or glycogen synthase activity. To distinguish between these possibilities, we used 31P NMR to noninvasively measure intramuscular G-6-P concentrations, which, unlike enzyme activity measurements, provides unique information regarding rate-controlling steps in muscle glycogen synthesis. Because G-6-P is an intermediate between glucose transport/hexokinase and glycogen synthase, its concentration will reflect the relative activities of these two steps. After glycogen loading, we observed an approximately twofold increase in intracellular G-6-P concentration, suggesting that a decrease in glycogen synthase activity was responsible for this lower rate of muscle glycogen synthesis. These data are consistent with the hypothesis that glycogen limits its own synthesis through inhibition of glycogen synthase activity and are concordant with the in vitro observations that glycogen inhibits the stimulatory effect of insulin on glycogen synthase activity (19) through inactivation of glycogen phosphatase activity (18, 31). It is noteworthy that this rate-controlling step is different from that found in other conditions of decreased insulin-stimulated muscle glycogen synthesis, such as obesity (22), type II diabetes (28), insulin-resistant offspring of type II diabetic parents (27), and poorly controlled type I diabetes (4), where reduced glucose transport/phosphorylation activity has been shown to be responsible for the lower rates of muscle glycogen synthesis. Although it is possible that the preceding high-CHO/exercise protocol and the lower plasma FFA concentrations in the glycogen-loading studies might have contributed to these findings, it is unlikely since all of these factors might be expected to promote muscle glycogen synthesis (1, 2, 21, 23, 26, 32).
We also examined whether increased glycogen cycling might have contributed to decreased insulin-stimulated muscle glycogen synthesis in the presence of elevated muscle glycogen concentration. We found that glycogen turnover (Vsyn/Vphos) was similar under both normal and glycogen-loaded conditions, suggesting that increased muscle glycogen turnover does not play a major role in limiting net synthesis of muscle glycogen under glycogen-loaded conditions. Although muscle (14, 19) and liver (13) phosphorylase activity have both been shown to be stimulated by an increase in glycogen content, it is possible that the increased intramuscular G-6-P concentration observed in the glycogen-loaded subjects inhibited phosphorylase activity under these conditions (15).
Despite a lower rate of muscle glycogen synthesis in the
glycogen-loaded state, rates of insulin-stimulated whole body glucose disposal were similar in the two protocols. This was somewhat surprising, because muscle glycogen synthesis typically accounts for
the majority of insulin-stimulated glucose disposal (29). These data
imply that, under glycogen-loaded conditions, a significant portion of
the infused glucose was being shunted into alternative pathways.
Consistent with this hypothesis were the higher rates of whole body
glucose oxidation (+30%) and plasma lactate concentrations (+90%)
observed in the subjects after the glycogen-loading protocol. These
data suggest that, under glycogen-loaded conditions, glucose is
diverted from muscle glycogen synthesis into aerobic and anaerobic glycolysis, with some of the glycolytically derived lactate taken up by
the liver for gluconeogenesis and/or lipogenesis and other tissues for
oxidation (Fig. 5). Although it is also
possible that decreased lactate clearance contributed to this increase
in plasma lactate concentration, the decrease in intramuscular pH
observed during the glycogen-loaded clamp studies would support the
former possibility. These observations are consistent with previous in vitro rat studies that have demonstrated that muscles with high glycogen content converted less of the glucose entering the cell into
glycogen and more into lactate (32). However, they are in contrast to
the results of other in vitro rat studies that have demonstrated a
negative correlation between muscle glycogen content and
insulin-stimulated glucose uptake (9). One possible explanation for
this latter finding is that these studies examined glucose uptake when
muscle glycogen was kept low by CHO restriction, resulting in a
relatively high proportion of fat in the diet that is well known to
cause insulin resistance (16).
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In summary, these are the first in vivo studies in humans to demonstrate that an elevated concentration of muscle glycogen is associated with a reduction in the rate of insulin-stimulated net muscle glycogen synthesis. This reduced rate of muscle glycogen synthesis could be attributed to a reduction in glycogen synthase activity as opposed to a reduction in glucose transport/hexokinase activity or an increase in the rate of muscle glycogen cycling. This mechanism likely plays an important role in limiting muscle glycogen synthesis in humans under fed conditions.
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ACKNOWLEDGEMENTS |
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We acknowledge the staff of the Yale/New Haven Hospital General Clinical Research Center for assistance with the studies and Veronika Walton for assistance with the GC-MS measurements. In addition, we thank Dr. Douglas L. Rothman and Terry Nixon for help with the NMR spectroscopy.
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FOOTNOTES |
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This research was supported by National Institutes of Health (NIH) Grants RO1 DK-49230, P30 DK-45735, and MO1 RR-00125 and United States Army Grant DAMD17-96-C-6097 (T. B. Price). K. F. Petersen is the recipient of a Clinical Associate Physician award from the NIH. G. I. Shulman is an investigator of 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: G. I. Shulman, Howard Hughes Medical Institute, Yale Univ. School of Medicine, Dept. of Internal Medicine, 295 Congress Ave., Box 9812, New Haven, CT 06510 (E-mail: gerald.shulman{at}yale.edu).
Received 22 July 1999; accepted in final form 5 November 1999.
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