Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, Tennessee 37232
Submitted 9 May 2003 ; accepted in final form 17 November 2003
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ABSTRACT |
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hyperinsulinemia; gluconeogenesis; glycogenolysis; hepatic glucose production
In the human, hyperglycemia decreased glycogenolysis by inhibiting flux through glycogen phosphorylase (28). In the dog, inhibition of glucose output during hyperglycemia was associated with a decrease in net hepatic glycogenolysis and a reduction in net carbon flux toward glucose 6-phosphate (G-6-P) in the gluconeogenic/glycolytic pathway (4, 40, 43). Although hyperglycemia appears to inhibit hepatic glucose production primarily by decreasing net hepatic glycogenolysis and increasing glucose cycling, it was also shown to decrease the rate of alanine and lactate gluconeogenesis when glycogen reserves were depleted (24, 40). This effect may be explained by the formation of fructose 2,6-bisphosphate (F-2,6-P2), which reciprocally regulates gluconeogenic and glycolytic flux (6, 16), since hyperglycemia would increase fasting F-2,6-P2 levels, which would then inhibit gluconeogenesis (24, 40). In addition, glucose inhibits phosphoenolpyruvate carboxykinase gene expression (18, 37) and can reduce gluconeogenesis in the human by acting as a signal to divert carbon derived from gluconeogenic flux to G-6-P into glycogen rather than plasma glucose (7). The first aim of the present studies was, therefore, to investigate the ability of hyperglycemia to acutely modify hepatic gluconeogenic flux to G-6-P in vivo.
Increases in plasma insulin inhibit hepatic glucose production directly by reducing net hepatic glycogenolytic flux (via stimulation of synthase and possibly by inhibition of phosphorylase) (28, 31, 35, 44, 47) and decreasing gluconeogenesis (as a result of the diversion of carbon derived from hepatic gluconeogenic flux into glycogen instead of plasma glucose) (5, 9, 10, 15). Insulin also reduces hepatic glucose production indirectly through its inhibitory effect on lipolysis. Decreased plasma nonesterified fatty acid (NEFA) level results in the redirection of glycogenolytic carbon to lactate instead of plasma glucose within the liver (42). During conditions in which insulin levels are deficient either acutely (10) or chronically (in an absolute or relative sense, such as long-term fasting or diabetes), gluconeogenic flux to G-6-P and the gluconeogenic rate are increased (16, 29). On the other hand, gluconeogenic flux to G-6-P is resistant to physiological increases in the insulin level (7, 9, 10), despite the well-documented in vitro effects of insulin on the activity of gluconeogenic enzymes and flux through this pathway (16, 29).
Recently, we suggested that, because acute increases in the insulin level profoundly alter liver glycogenolysis, the effects of the hormone on hepatic gluconeogenic flux might be masked by the changes in G-6-P attributable to alterations in glycogenolysis (10). Despite the presence of a phosphorylase inhibitor (BAY R3401), which prevented large changes in glycogen breakdown, a fourfold increase in insulin had no observable effect on hepatic gluconeogenic flux to G-6-P, even after 5 h. One possible explanation for insulin's lack of effect on the process is that increased glycogen synthesis may have decreased the F-2,6-P2 concentration. This could have offset the hormone's direct inhibitory effects on the gluconeogenic pathway so that flux through it was maintained.
Thus we hypothesized that hyperglycemia, occurring simultaneously with a rise in insulin, would create a condition in which gluconeogenic flux to G-6-P could be acutely reduced by the hormone. Therefore, the second aim of the present studies was to investigate the effect of hyperglycemia in combination with hyperinsulinemia on the regulation of hepatic gluconeogenic flux to G-6-P in vivo.
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MATERIALS AND METHODS |
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Experiments were conducted on 24 overnight-fasted conscious mongrel dogs (20-28 kg) of either sex. Housing and diet have been described previously (9). The surgical facility met the standards published by the American Association for the Accreditation of Laboratory Animal Care, and the protocols were approved by the Vanderbilt University Medical Center Animal Care Committee. All dogs underwent a laparotomy 2 wk before the experiment to implant infusion catheters into the jejunal and splenic veins, sampling catheters into the portal and hepatic veins and the femoral artery, and Transonic flow probes around the hepatic artery and portal vein, as described elsewhere (9). Each dog was used for only one experiment. All dogs studied were healthy, as indicated by 1) leukocyte count <18,000/mm3, 2) a hematocrit >35%, 3) a good appetite, and 4) normal stools.
Experimental Design
Intraportal catheters (splenic and jejunal) were used for the infusion of insulin (Lilly, Indianapolis, IN) and glucagon (Lilly). Angiocaths (Deseret Medical, Becton Dickinson, Sandy, UT) were inserted percutaneously into leg veins for [3-3H]glucose (DuPont NEN, Boston, MA), indocyanine green (Sigma, St. Louis, MO), and peripheral glucose (50% Dextrose; Baxter Healthcare, Deerfield, IL) infusion. Animals were allowed to rest quietly in a Pavlov harness for 30 min before the experiments were started. Each of the four protocols consisted of an equilibration period (-120 to -40 min), a basal period (-40 to 0 min), and an experimental period (0-300 min). At -120 min a priming dose of [3-3H]glucose (50 µCi) was given, and constant infusions of [3-3H]glucose (0.4 µCi/min) and indocyanine green (0.07 mg/min) were started. At the same time, a constant infusion of somatostatin (0.8 µg·kg-1·min-1) was started in a peripheral vein to inhibit endogenous pancreatic hormone secretion, and constant intraportal glucagon (0.5 ng·kg-1·min-1) and insulin (300 µU·kg-1·min-1) infusions were started to replace basal secretion of these hormones. Also, at -120 min a glycogen phosphorylase inhibitor (BAY R3401; 10 mg/kg in a 0.5% methyl cellulose-water solution) was given orally or intragastrically, depending on whether a stomach catheter had been implanted during surgery (there was no difference in the effectiveness of the drug with one route or the other). Shortly thereafter, a variable glucose infusion was begun to maintain euglycemia in the presence of the glycogen phosphorylase inhibitor and to raise the glucose level during the experimental period in the hyperglycemic protocols. [3-3H]glucose infusion rates were modified to agree with the cold glucose infusion rates to minimize changes in the plasma glucose specific activity. During the experimental period, two insulin infusion rates were used, and two glucose levels were maintained.
Protocol 1: basal glucose/basal insulin (Control; n = 6). No change was made in the insulin infusion rate over the course of the experiment.
Protocol 2: high glucose/basal insulin (HG; n = 6). Beginning at 0 min, the arterial glucose level was increased twofold, resulting in 5 h of hyperglycemia, while basal insulin infusion was continued.
Protocol 3: basal glucose/high insulin (HI; n = 6). Beginning at 0 min, the intraportal insulin infusion rate was increased to 1.2 mU·kg-1·min-1, resulting in 5 h of hyperinsulinemia. Euglycemia was maintained by glucose infusion.
Protocol 4: high glucose/high insulin (HGI; n = 6). Beginning at 0 min, the intraportal insulin infusion rate was increased to 1.2 mU·kg-1·min-1, and the arterial glucose level was increased twofold, resulting in 5 h of hyperinsulinemia/hyperglycemia.
Data related to the effects of BAY R3401 in the control and hyperinsulinemic states (protocols 1 and 2) were previously published (10) and are included here for ease of comparison. Although there was overlap in the time frame during which protocols 1, 2, 3, and 4 were performed, they were not completely randomized.
Immediately after the final sampling time, each animal was anesthetized, and sections of three liver lobes were freeze-clamped in situ and stored at -70°C, as previously described (9). All animals were then euthanized, and the correct positions of the catheter tips were confirmed.
Analytical Procedures
Hematocrit; plasma glucose, [3H]glucose, glucagon, insulin, cortisol, and NEFA; and blood alanine, glycine, serine, threonine, lactate, glutamine, glutamate, glycerol, -hydroxybutyrate (
-OHB), and hepatic glycogen concentrations were determined as previously described (9). Hepatic glycogen content was also measured as previously described (23). Hepatic tissue glucose, G-6-P, and fructose 6-phosphate (F-6-P) were measured fluorometrically by standard enzymatic methods (20, 22) after deproteinization with 3% perchloric acid.
Calculations
Net hepatic balances (NHB) were calculated with the arteriovenous difference method using the formula NHB = loadout - loadin, where loadout = [H] x HF; loadin = [A] x AF + [P] x PF; [H], [A], and [P] are the substrate concentrations in hepatic vein, femoral artery, and portal vein blood or plasma, respectively; and HF, AF, and PF are blood flow in the hepatic vein, hepatic artery, and portal vein, as determined by the ultrasonic flow probes. With this calculation a positive value represents net output by the liver, and a negative value represents net hepatic uptake. Plasma glucose and [3H]glucose values were multiplied by 0.73 to convert them to blood glucose values, as validated elsewhere (23). Net fractional substrate extraction by the liver was calculated as net hepatic balance ÷ loadin. The approximate insulin and glucagon levels in plasma entering the liver sinusoids were calculated using the formula [A] x %AF + [P] x %PF, where [A] and [P] are arterial and portal vein hormone concentrations, respectively, and %AF and %PF are the respective percent contributions of arterial and portal flow to total hepatic blood flow.
Tracer-determined whole body glucose appearance, utilization (Rd), and clearance (Cl) were measured using a primed, constant infusion of [3-3H]glucose. Data calculation was carried out using the two-compartment model described by Mari (21) and by canine parameters reported by Dobbins et al. (8). Endogenous glucose appearance (Ra) was calculated by subtracting the glucose infusion rate (GIR) from whole body glucose appearance.
For calculation of unidirectional hepatic glucose uptake (HGU), the net [3H]glucose uptake was divided by the arterial [3H]glucose specific activity. This calculation is based on the assumption that intrahepatic uptake of glucose occurs before glucose production, so that the plasma glucose specific activity is not diluted. Even if this assumption is not correct, the drop in specific activity across the liver is very small; thus the assumption is of little consequence. Unidirectional hepatic glucose release (HGR) was determined by adding HGU to net hepatic glucose output (NHGO).
Gluconeogenesis, as classically defined, is the synthesis and subsequent release of glucose from noncarbohydrate precursors. Carbon produced from flux through the gluconeogenic pathway does not necessarily have to be released as glucose; it can also be stored in glycogen. Therefore, we make a distinction between gluconeogenic flux to G-6-P (conversion of precursors to G-6-P: G-6-P neogenesis) and gluconeogenesis (release of glucose derived from gluconeogenic flux). In the present studies, we estimated hepatic gluconeogenic flux to G-6-P, net hepatic gluconeogenic flux, and net hepatic glycogenolytic flux (NHGLY).
Gluconeogenic flux to G-6-P was determined by summing the net hepatic uptake rates of the gluconeogenic precursors (alanine, glycine, serine, threonine, glutamine, glutamate, glycerol, lactate, and pyruvate), converting the sums to glucose equivalents, and dividing by two to account for the incorporation of three carbon precursors into the six-carbon glucose molecule. This method assumes that there is 100% conversion of gluconeogenic precursors taken up by the liver into G-6-P and thus provides a maximal estimate of gluconeogenic flux from circulating precursors. It also assumes that intrahepatic gluconeogenic precursors do not contribute significantly to gluconeogenic flux. The errors from these assumptions are difficult to assess but appear to be small and, to the extent that they occur, are offsetting. Recently, results obtained with the arteriovenous difference technique were compared with those obtained with an independent technique that is not subject to these assumptions (9, 13). The two methods yielded similar estimates, suggesting that the assumptions are reasonable.
Net hepatic gluconeogenic flux was determined by subtracting the summed net hepatic output rates (when such occurred) of the substrates noted above (in glucose equivalents) and hepatic glucose oxidation (GO) from gluconeogenic flux to G-6-P. GO was assumed to be 0.2 mg·kg-1·min-1 in the basal period of each experiment, as in our earlier studies in the overnight-fasted dog (14, 25). This parameter was not directly measured, because it is difficult to differentiate between the small signal and the high inherent noise in the measurement. In previous studies, hepatic GO did not change appreciably under hyperglycemic euinsulinemic (Satake S, Moore MC, Neal DW, Hastings J, Cherrington AD, unpublished observations) or euglycemic hyperinsulinemic (25) states; thus GO was assumed to remain unchanged in these protocols. Previously, under conditions comparable to those seen in the HGI group (i.e., hyperglycemia-hyperinsulinemia), hepatic GO increased progressively to a rate of 0.8 mg·kg-1·min-1 (14, 38). This change in GO could have resulted from activation of pyruvate dehydrogenase by insulin in the presence of substrate (provided by hyperglycemic stimulation of glycolytic flux), which would lead to increased pyruvate oxidation and decreased availability of substrate for the lactate dehydrogenase reaction (12, 33, 46). Therefore, in the HGI group, GO was assumed to rise linearly from 0.2 to 0.8 mg·kg-1·min-1 over the test period. When net hepatic gluconeogenic flux is positive, there is net flux to G-6-P, whereas a negative number indicates net flux to CO2 or lactate.
NHGLY was estimated by subtracting net hepatic gluconeogenic flux from net hepatic glucose balance. A positive number therefore represents net glycogen breakdown, whereas a negative number indicates net glycogen synthesis.
Ideally, the gluconeogenic flux rate would be calculated using unidirectional hepatic uptake and output rates for each substrate, but this would be difficult, as it would require the simultaneous use of multiple stable isotopes, which could themselves induce a mild perturbation of the metabolic state. Therefore, net hepatic balance was used instead, necessitating consideration of the limits of this approach. There is little or no hepatic production of alanine (40) (quantitatively the most important gluconeogenic amino acid) or glycerol (30), so in this case the compromise is of little consequence. Such is, however, not the case for lactate. Our estimate of the rate of gluconeogenic flux to G-6-P will be quantitatively accurate only if we assume that lactate flux is unidirectional at a given moment (i.e., either in or out of the liver). In a given cell this does not seem like an unreasonable assumption in light of the reciprocal control of gluconeogenesis or glycogenolysis (29). Jungermann and Katz (17) have suggested, however, that there is spatial separation of metabolic pathways, so that gluconeogenic periportal hepatocytes primarily synthesize glucose and glycogen from lactate and other noncarbohydrate precursors, whereas glycolytic perivenous hepatocytes generally consume plasma glucose, which is predominantly oxidized or released as lactate (31). Therefore, it is possible that under normal nutritional conditions, in a net sense, hepatic gluconeogenic and glycolytic flux occur simultaneously, with lactate output or uptake occurring in different cells. To the extent that flux occurs in both directions simultaneously, the arteriovenous difference method will underestimate the absolute gluconeogenic flux to G-6-P. It should be noted, however, that net hepatic gluconeogenic flux and net hepatic glycogenolytic flux can be calculated accurately regardless of the validity of the assumptions related to whether or not simultaneous gluconeogenic and glycolytic substrate flux occur, so long as equilibrium has been attained across all compartments.
The total net contribution (TNC) of carbon to hepatic glycogen was determined by using the average net hepatic glycogen synthetic rate during the experimental period to predict the total net accumulation of glycogen over that period of time (300 min) with the following formula
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The direct net contribution (DNC) of plasma glucose to hepatic glycogen stores (mg/g) was estimated by dividing [3H]glycogen counts/g liver by the average arterial plasma [3H]glucose specific activity present during the experimental period (dpm/g ÷ dpm/mg). This is a maximal estimate, because initially, before the glycogen specific activity reaches a steady state, a disproportionate amount of tracer glucose (as opposed to unlabeled glucose) would be deposited in glycogen as a result of ongoing glycogen cycling. The indirect net contribution (INC) of gluconeogenic precursors to hepatic glycogen stores was estimated by subtracting DNC from TNC and is thus a minimal estimate.
Statistical Analysis
The data were analyzed for differences from the basal period and for differences from the control group. Statistical comparisons were carried out using two-way ANOVA (JMP IN Software, SAS Institute). One-way ANOVA comparison tests were used post hoc when significant F ratios were obtained. Significance was established when P < 0.05 (two-sided test).
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RESULTS |
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Glucose infusion was required to maintain euglycemia during the basal period in each group as a result of the inhibition of glycogen breakdown. Although the GIR did not change significantly over time in the control group (3 ± 1 mg·kg-1·min-1), it rose to an average of 7 ± 1, 12 ± 1, and 15 ± 1 mg·kg-1·min-1 in the HG, HI, and HGI groups, respectively, by the last hour of the experiment (P < 0.05; Table 1). In the control and insulin excess groups, euglycemia was maintained throughout. The plasma glucose level was increased twofold in the two hyperglycemic protocols (HG and HGI; P < 0.05; Fig. 2).
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Net hepatic glucose balance (NHGB) remained close to zero throughout the control protocol (Fig. 2). In the HG, HI, and HGI groups, net HGU occurred, averaging 4.11 ± 0.50, 1.58 ± 0.31, and 3.93 ± 0.56 mg·kg-1·min-1, respectively, during the last hour of the study (P < 0.05). HGU did not change over time in the control group (averaging 1.12 ± 0.09 mg·kg-1·min-1) but rose to 3.45 ± 0.40, 1.80 ± 0.16, and 3.37 ± 0.47 mg·kg-1·min-1 in the HG, HI, and HGI groups, respectively (P < 0.05; Table 1). HGR tended to decline over the course of the experiment in the control group (1.23 ± 0.21 to 0.84 ± 0.31) and was completely inhibited in the other three groups (P < 0.05; Table 1). Glucose Ra was reduced relative to normal in the basal period of each group as a result of phosphorylase inhibition. It changed little over time in the control group. On the other hand, endogenous Ra decreased during the experimental period in the other groups, although the magnitude of the fall did not reach significance in response to hyperglycemia alone (Table 1). Glucose Rd did not change in the control group, whereas it rose (from basal to the last hour of the experimental period) by 3.60 ± 0.30, 8.06 ± 1.82, and 13.43 ± 1.69 mg·kg-1·min-1 in the HG, HI, and HGI groups, respectively (Table 1). Glucose Cl, an index of the avidity with which tissues take up glucose, was unchanged in the control and HG groups and rose during hyperinsulinemia (HI and HGI groups; P < 0.05; Table 1).
The arterial blood lactate level did not change significantly in the control or HI groups, although there was a tendency for it to fall slightly and then rise modestly (Fig. 3). In the HG and HGI groups, the lactate level rose twofold (P < 0.05). Net hepatic lactate balance did not change significantly over time in the control group but decreased slightly during hyperinsulinemia ( = -2.98 ± 2.18 µmol·kg-1·min-1 from basal to last hour of the experimental period; P < 0.05; Fig. 3). On the other hand, in the HG and HGI groups, net hepatic lactate balance increased (
= 11.20 ± 3.74 and 4.73 ± 2.05 µmol·kg-1·min-1, respectively, from basal to the last hour of the experimental period).
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The net hepatic fractional extraction of gluconeogenic amino acids did not change significantly in any group but tended to rise, particularly when insulin was increased (Fig. 4). Therefore, although the arterial gluconeogenic amino acid levels decreased in the HI and HGI groups (Fig. 4 and Table 2), the net hepatic gluconeogenic amino acid uptake rates did not change significantly over time in any group (Fig. 4 and Table 3).
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Despite an initial drift down, there was no significant change in arterial blood glycerol level or net hepatic glycerol uptake in the control group (Fig. 5). During hyperglycemia, the change in glycerol levels and uptake followed similar patterns to those evident in the control group. On the other hand, when insulin was increased, with or without hyperglycemia, the arterial glycerol levels and net hepatic glycerol uptakes decreased by 60% (P < 0.05).
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The arterial plasma NEFA levels decreased modestly in the control and HG groups (P < 0.05; Fig. 6). On the other hand, the NEFA level decreased by 85% in the hyperinsulinemic groups (HI and HGI; P < 0.05). Net hepatic NEFA uptake showed a similar pattern, tending to fall modestly in the control and hyperglycemic groups but decreasing 90% during insulin excess with and without hyperglycemia (P < 0.05; Fig. 6). In the control group, there was no significant change in the arterial blood -OHB level or in net hepatic
-OHB output (Table 4). In the other three groups, the arterial
-OHB levels and outputs declined (P < 0.05; Table 4).
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Net hepatic glycogenolytic flux decreased over time in the control group (-0.30 ± 0.32 to -1.20 ± 0.18 mg·kg-1·min-1; P < 0.05), the HG group (-0.76 ± 0.36 to -3.37 ± 0.51 mg·kg-1·min-1; P < 0.05), the HI group (-0.81 ± 0.28 to -2.44 ± 0.35 mg·kg-1·min-1; P < 0.05), and the HGI group (-0.98 ± 0.14 to -3.77 ± 0.60 mg·kg-1·min-1; P < 0.05). In a net sense, glycogen synthesis increased (from basal to the last hour of the experimental period) by 0.90 ± 0.43, 2.62 ± 0.48, 1.63 ± 0.36, and 3.19 ± 0.60 mg·kg-1·min-1 in the control, HG, HI, and HGI groups, respectively (P < 0.05; Fig. 7). The maximal DNC values of plasma glucose to glycogen stores were 7 ± 1, 14 ± 1, 10 ± 1, and 13 ± 2 mg/g in the control, HG, HI, and HGI groups, respectively (Table 5). The minimal INC values of gluconeogenic substrates to liver glycogen were 5 ± 1, 25 ± 7, 11 ± 3, and 21 ± 8 mg/g in the four groups, respectively (Table 5). The TNC of carbon to hepatic glycogen over the 5-h experimental period was 11 ± 2, 39 ± 7, 21 ± 3, and 34 ± 8 mg/g liver in the control, HG, HI, and HGI groups, respectively (Table 5).
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The hepatic tissue glucose level was not altered by insulin excess (0.23 ± 0.02 vs. 0.21 ± 0.02 mg/g in the control and HI groups, respectively); however, it was increased in response to hyperglycemia (0.46 ± 0.02 and 0.45 ± 0.01 mg/g in the HG & HGI groups, respectively; P < 0.05; Table 6). The terminal hepatic G-6-P and F-6-P levels were not significantly altered by treatment with insulin or glucose (Table 6).
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Hepatic gluconeogenic flux to G-6-P did not change significantly over time in the control group (1.05 ± 0.17 and 1.15 ± 0.17 mg·kg-1·min-1 in the two periods) or the HI group (0.86 ± 0.07 and 1.06 ± 0.01 mg·kg-1·min-1), whereas in the HG and HGI groups it decreased 33% (P < 0.05). The changes in hepatic gluconeogenic flux to G-6-P between basal and the last hour of the experimental period were +0.10 ± 0.10, -0.24 ± 0.12, +0.21 ± 0.13, and -0.40 ± 0.14 mg·kg-1·min-1 in the control, HG, HI, and HGI groups, respectively (Figs. 7 and 8).
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Net hepatic gluconeogenic flux did not change significantly over time in the control group (0.58 ± 0.26 and 0.87 ± 0.19 mg·kg-1·min-1 in the two periods). During insulin excess it rose from 0.47 ± 0.11 to 0.85 ± 0.11 mg·kg-1·min-1 (P < 0.05) but was not different from the control group. During hyperglycemia, net hepatic gluconeogenic flux decreased (0.35 ± 0.16 to -0.74 ± 0.16 and 0.79 ± 0.14 to -0.16 ± 0.20 in the HG and HGI groups, respectively; P < 0.05), indicating that in a net sense glycolysis was occurring. The changes in net hepatic gluconeogenic flux between basal and the last hour of the experimental period were 0.29 ± 0.21, -1.09 ± 0.26, 0.38 ± 0.14, and -0.95 ± 0.16 mg·kg-1·min-1 in the control, HG, HI, and HGI groups, respectively (P < 0.05; Figs. 7 and 8).
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DISCUSSION |
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The glycogen phosphorylase inhibitor (BAY R3401) used in these studies reduces hepatic glycogenolysis through allosteric inhibition and dephosphorylation of glycogen phosphorylase a, and it acts without producing other apparent direct metabolic alterations (3, 10, 39). Activation of glycogen synthase will occur, however, if phosphorylase is inactivated below the threshold level inhibitory to glycogen synthase phosphatase (31, 35, 44, 47). Its purpose in these studies was to allow us to examine the effects of hyperglycemia on the hepatic gluconeogenic process under conditions in which glycogen breakdown was inhibited before glucose or insulin administration, thereby preventing the effects of a sudden drop in glycogen breakdown from masking the effects of hyperglycemia on gluconeogenesis. Although net hepatic glycogen breakdown was fully inhibited by the drug, glycogen synthesis increased during hyperglycemia. Thus it should be kept in mind that these studies did not address the effects of hyperglycemia on the gluconeogenic pathway completely independently from changes in glycogen metabolism (i.e., a "pull" effect could still have occurred via activation of glycogen synthase).
Hepatic gluconeogenic flux to G-6-P is determined by the supply (load) of gluconeogenic substrates reaching the liver and the gluconeogenic capacity of the organ, whereas net hepatic gluconeogenic flux is derived from the net balance of gluconeogenic flux to G-6-P and the glycolytic flux rate (predominantly lactate production and glucose oxidation). During hyperglycemia or hyperinsulinemia alone, the supply of gluconeogenic amino acids and glycerol reaching the liver decreased by 10 and 20%, respectively, whereas there was no change in the control group. The effects of hyperglycemia and hyperinsulinemia (HGI group) were additive, and the two together resulted in a 30% decrease in the load of these precursors reaching the liver. There was an offsetting increase in the hepatic lactate load in the three experimental groups, however, so that the total gluconeogenic precursor supply to the liver was unaffected by the changes in glucose and/or insulin. Despite unchanged total precursor availability, hepatic gluconeogenic flux to G-6-P tended to decrease over time during hyperglycemia (HG and HGI groups), suggesting a mild inhibitory effect of glucose on the gluconeogenic pathway (Fig. 8). As in the control group, gluconeogenic flux to G-6-P did not decrease during hyperinsulinemia alone, suggesting that a fourfold rise in insulin has no effect on this process over a 5-h period.
Net hepatic gluconeogenic flux was decreased when hyperglycemia was present (HG and HGI groups) due to a change in lactate balance such that the net carbon flux in the glycolytic/gluconeogenic pathway was in the direction of glycolysis. The change in net lactate balance in the HG group was twice that observed in the HGI group, however (+11 vs. +5 µmol·kg-1·min-1 in the two groups, respectively). This difference may have been the result of an increase in glucose oxidation due to activation of pyruvate dehydrogenase (PDH) by insulin (12, 46). This would be consistent with our observation that, despite similar glucose uptake and glycogen synthetic rates, net hepatic lactate output did not occur during hyperglycemia in the presence of hyperinsulinemia. These findings are in agreement with those from previous studies, which showed that although gluconeogenic flux to G-6-P was not significantly inhibited by a twofold rise in glucose in the 18-h-fasted dog (43), in the 36-h-fasted dog net hepatic lactate uptake decreased from 7 µmol·kg-1min-1 to near zero, and the conversion of labeled alanine and lactate into glucose decreased by 60% after 2 h of hyperglycemia (40). In both of those studies, hyperglycemia caused a shift in net hepatic lactate balance toward lactate output, but a reduction in gluconeogenic flux to G-6-P was only visible when preexisting lactate uptake was initially present (i.e., during extended fasting). Net hepatic gluconeogenic flux, which takes into account both gluconeogenic and glycolytic fluxes, was reduced during hyperglycemia in both circumstances, however, as it was in the present study. In the control and HI groups, there was no reduction in net hepatic gluconeogenic flux (Fig. 8). These data demonstrate that hyperglycemia inhibits net hepatic gluconeogenic flux, whereas physiological hyperinsulinemia per se has minimal impact on the process.
The plasma NEFA and glycerol levels tended to decrease in each of the four groups; however, they decreased more in the presence of elevated insulin (HI and HGI groups). Hyperglycemia alone had little effect on lipolysis (as indicated by the similarity in fall of blood glycerol and plasma NEFA in the HG and control groups), whereas insulin decreased plasma NEFA levels by both decreasing lipolysis and increasing fatty acid reesterification (the fall in NEFA levels was greater than the fall in glycerol levels). Hepatic fatty acid oxidation was also reduced by hyperglycemia and hyperinsulinemia (HG, HI, and HGI groups), as demonstrated by the fall in ketone body production relative to the control group (Table 4). During hyperinsulinemia, fatty oxidation was completely inhibited because there was no substrate to oxidize (net hepatic NEFA uptake was close to zero). In the HG group, the fall in the net hepatic output of -OHB (
60%) was greater than the decrease in net hepatic NEFA uptake (
35%), whereas in the other groups the reduction in
-OHB production was similar to the fall in NEFA uptake. Thus hyperglycemia appeared to inhibit hepatic fatty acid oxidation, at least in part by a direct, nonlipolytic mechanism. These findings are in agreement with the hypothesis of Randle (32), who suggested that glucose promotes lipid storage and inhibits fatty acid oxidation.
In the HG and HGI groups, net hepatic gluconeogenic flux was reduced from basal (1.1 mg·kg-1·min-1 in both groups) as the result of decreased gluconeogenic flux to G-6-P and increased glycolytic flux. The fall in hepatic fatty acid oxidation observed during hyperglycemia would be expected to result in a decrease in the mitochondrial ratios of [acetyl-CoA]/[CoA] and [NADH]/[NAD+] (11, 32, 48). These changes would result in the stimulation of the PDH complex and a decrease in citrate [an inhibitor of phosphofructokinase (PFK-1 and PFK-2)], and thereby the levels of F-2,6-P2 and F-1,6-P2 would increase. This in turn would result in increased glycolysis and decreased gluconeogenic flux to G-6-P (16, 29, 32). Reduction of acetyl-CoA or long-chain fatty acetyl-CoA could also decrease pyruvate carboxylase activity, and a fall in free fatty acid oxidation would reduce the supply of the reducing equivalents (NADH) and energy (ATP) necessary for gluconeogenic flux (2, 11, 32, 34, 48, 49). Thus the reduction in hepatic fatty acid oxidation could at least partially explain the changes in gluconeogenic flux to G-6-P and glycolytic flux seen during hyperglycemia (Figs. 7 and 8). On the other hand, despite complete inhibition of fatty acid oxidation during hyperinsulinemia alone, there was neither a reduction in gluconeogenic flux to G-6-P nor net gluconeogenic flux. Thus, in this in vivo setting, changes in hepatic fatty acid oxidation may have affected net hepatic gluconeogenic flux only in the presence of hyperglycemia; however, further studies where free fatty acid levels are controlled are needed to allow this conclusion.
Previous studies in the dog and human demonstrated that peripheral glucose infusion, when brought about in the presence of basal insulin, is relatively ineffective in stimulating net hepatic glucose uptake (4, 7, 27). On the other hand, the liver takes up glucose at high rates when glucose is delivered via the portal vein, particularly when portal insulin concentrations are also increased (4, 7). In the present studies, net hepatic glucose uptake increased from basal modestly in the control and insulin-alone (HI) groups (Fig. 2). During hyperglycemia (created by peripheral, not portal vein glucose infusion), however, net hepatic glucose uptake was 12-fold greater than in the control group and was thus similar in magnitude to the uptake seen after maximal activation of the portal signal in the presence of hyperglycemia (27). Glycogen synthase activity is reduced by glycogen phosphorylase's inhibitory effect on glycogen synthase phosphatase (31). Thus it is likely that activation of glycogen synthase occurred because of the inactivation of glycogen phosphorylase, which in turn was responsible for the marked increase in hepatic glucose uptake during hyperglycemia in these studies.
Like the plasma glucose level, during hyperglycemia (HG and HGI groups) the hepatic tissue glucose level doubled (Table 6). Despite the remarkable increase in net hepatic glucose uptake, however, the hepatic G-6-P concentration was not significantly increased at the end of the experimental period. One possibility is that flux through glucokinase (GK) was maximal in the control state, but because GK is not inhibited by G-6-P and is half-saturated at 10-15 mM (16, 45), and because glucose uptake increased during hyperglycemia, this cannot be the case. Therefore, the discordance between hepatic glucose and G-6-P levels suggests that flux out of G-6-P increased proportionally to the rate of entry into the pool. Although glucose cycling (through GK and G-6-Pase) was not measured in this study, in previous experiments it was increased during hyperglycemia (15, 36, 39). It is also possible that hyperglycemia, per se, caused an initial increase in the hepatic G-6-P concentration, which then activated the glycolytic and glycogen synthetic pathways, returning the G-6-P level to a new steady state near, but slightly above, the original concentration (Table 6). Indeed, net hepatic glycogen synthesis increased, as did net hepatic glycolytic flux. The increase in glycogen synthesis was rapid and occurred maximally within the first hour, whereas net hepatic glycolytic flux increased steadily over the 5-h experimental period (Fig. 7). Previous studies demonstrated that when fasted rats were infused with glucose, there was a lag of 2-3 h before hepatic F-2,6-P2 levels began to rise (19). Thus a steady rise in F-2,6-P2 levels during hyperglycemia in the present study could explain the reciprocal increase in net glycolytic flux and equivalent decrease in net flux into glycogen over the course of the experiments.
In each group, the changes in net hepatic glucose uptake were similar to the changes in net hepatic glycogen synthesis (Figs. 2 and 7). It has been suggested that glucose controls glycogen deposition by stimulating synthase, and therefore by a pull mechanism (31). Activation of glycogen synthase could have occurred via inactivation of phosphorylase a, in response to the phosphorylase inhibitor, with hyperglycemia providing the substrate (G-6-P) for rapid flux into glycogen in the HG and HGI groups (1, 3, 31, 35, 44, 47). In previous human experiments, 13C NMR spectroscopy was used to demonstrate that hyperglycemia, per se, inhibits net hepatic glycogenolysis primarily through inhibition of glycogen phosphorylase flux and not an activation of glycogen synthesis, whereas hyperinsulinemia, per se, inhibits the process primarily through stimulation of glycogen synthase flux (28). Recent studies in cultured hepatocytes demonstrated, however, that hepatic glycogen synthesis is highly sensitive to phosphorylase activity (1, 3). The latter is confirmed here in the dog, where we show that net hepatic glycogen synthesis is maximally stimulated by hyperglycemia after inhibition of glycogen phosphorylase, because there was no additional increase in glycogen synthesis when the plasma insulin level was also increased (Fig. 7).
The increase in glycogen synthesis during hyperglycemia resulted in a fourfold increase in total net contribution of carbon to hepatic glycogen (Table 5). Because net hepatic gluconeogenic flux decreased, whereas net hepatic glucose uptake rose, the direct pathway might be predicted to be the major contributor to glycogen synthesis. On the contrary, however, there was only a twofold increase in the direct contribution, whereas the increase in the indirect route was over fivefold compared with the control group. This suggests that activation of glycogen synthase was not, in fact, pulling plasma glucose directly into glycogen. Previously, Newgard et al. (26) found that, under normal refeeding conditions in the rat, the majority of liver glycogen is formed by an indirect pathway involving the conversion of glucose lactate
G-6-P
glycogen. Although in theory this loop could occur in a single cell, the gluconeogenic/glycolytic pathways are reciprocally regulated, in part by F-2,6-P2. Therefore, it is more likely that, during hyperglycemia, cycling from plasma glucose
G-6-P
pyruvate and pyruvate
G-6-P
glycogen was occurring in metabolically distinct hepatocytes. Thus labeled glucose entering glycolytic perivenous hepatocytes could have been released as unlabeled lactate, which was then subsequently converted into glycogen by gluconeogenic periportal hepatocytes (17, 31).
In summary, hyperglycemia inhibited net hepatic gluconeogenic flux in these studies, mainly by stimulation of glycolysis. Decreased hepatic fatty acid oxidation may have played a role in this response. Furthermore, hyperglycemia markedly stimulated the hepatic uptake of glucose, which appeared to be stored as glycogen via the indirect pathway (presumably via the release of lactate derived from plasma glucose from perivenous hepatocytes and its subsequent conversion into glycogen in periportal hepatocytes). Finally, the effects of hyperglycemia in combination with hyperinsulinemia were similar to the effect of hyperglycemia in the presence of basal insulin.
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ACKNOWLEDGMENTS |
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This work was presented in part at the 60th Annual Meeting of the American Diabetes Association, San Diego, CA, June 2000.
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FOOTNOTES |
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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. Section 1734 solely to indicate this fact.
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