Nonhepatic glucose production in humans
Alberto Battezzati,1,2
Andrea Caumo,2
Francesca Martino,2
Lucia Piceni Sereni,2
Jorgelina Coppa,3
Raffaele Romito,3
Mario Ammatuna,3
Enrico Regalia,3
Dwight E. Matthews,4
Vincenzo Mazzaferro,3 and
Livio Luzi2
1International Center for the Assessment of Nutritional Status, DiSTAM, Università degli Studi di Milano, 20133 Milano; 2Department of Medicine, Istituto Scientifico H San Raffaele, 20132 Milano; and 3Liver Transplantation Unit, Istituto Nazionale dei Tumori, 20133 Milano, Italy; and 4University of Vermont, Burlington, Vermont 05405
Submitted 7 November 2002
; accepted in final form 30 May 2003
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ABSTRACT
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Extrahepatic glucose release was evaluated during the anhepatic phase of liver transplantation in 14 recipients for localized hepatocarcinoma with mild or absent cirrhosis, who received a bolus of [6,6-2H2]glucose and L-[3-13C]alanine or L-[1,2-13C2]glutamine to measure glucose kinetics and to prove whether gluconeogenesis occurred from alanine and glutamine. Twelve were studied again 7 mo thereafter along with seven healthy subjects. At the beginning of the anhepatic phase, plasma glucose was increased and then declined by 15%/h. The right kidney released glucose, with an arteriovenous gradient of -3.7 mg/dl. Arterial and portal glucose concentrations were similar. The glucose clearance was 25% reduced, but glucose uptake was similar to that of the control groups. Glucose production was 9.5 ± 0.9 µmol·kg-1· min-1, 30% less than in controls. Glucose became enriched with 13C from alanine and especially glutamine, proving the extrahepatic gluconeogenesis. The gluconeogenic precursors alanine, glutamine, lactate, pyruvate, and glycerol, insulin, and the counterregulatory hormones epinephrine, cortisol, growth hormone, and glucagon were increased severalfold. Extrahepatic organs synthesize glucose at a rate similar to that of postabsorptive healthy subjects when hepatic production is absent, and gluconeogenic precursors and counterregulatory hormones are markedly increased. The kidney is the main, but possibly not the unique, source of extrahepatic glucose production.
kidney; intestines; amino acids
IN PHYSIOLOGICAL CONDITIONS, glucose production is assumed to be mainly hepatic in humans, although it was shown that the kidney and other organs can release newly formed glucose for the use of other tissues (22). The quantification of the extent of extrahepatic glucose production and the definition of its regulation are presently subject to debate. The kidney was studied as a major nonhepatic gluconeogenic site, but uncertainities related to methodological problems led to divergent measures of renal glucose production, which accounted for 5% up to 30% of the whole body glucose production during the postabsorptive state (8, 11, 19, 25). Most of the experimental problems are related to the accuracy in measurement of the renal blood flow (which is large), of the glycemic gradient across the renal bed (which is small or even neutral, because the kidney can simultaneously produce and take up glucose), and of the gradients in glucose tracer enrichments or specific activities across the same districts (19). In the studies that found a consistent contribution of the kidney to the postabsorptive glucose production, lactate and glutamine were preferred to alanine as gluconeogenic substrates (27). The kidney was also sensitive to the insulin suppression of gluconeogenesis (18) and to the gluconeogenic stimulation from epinephrine but not to that of glucagon (26, 27).
In considering the difficulties of dissecting the hepatic from the extrahepatic contributions to glucose production (including these from the kidney and other potential gluconeogenic organs), it is imperative to find experimental models in humans to unequivocally demonstrate the potential of gluconeogenic organs to release glucose into the systemic circulation. The anhepatic phase of liver transplantation is the lag of time in which the recipient's liver has been removed but the donor's liver has not yet been replaced. Although it is nonphysiological, this is a clear-cut model to demonstrate nonhepatic gluconeogenesis, because there is no liver to confound results with its large output of glucose. A recent study used this model to suggest that glucose is released at the rate of 1.3 mg·kg-1·min-1 in the absence of the liver (14). In the present study, we decided to measure the glucose production during the anhepatic phase of liver transplantation and to test whether the alanine and the glutamine carbons could still be transferred to glucose in the absence of the liver. We studied 14 patients who, in the absence of the liver, did not receive exogenous glucose during this period. We found that during the anhepatic phase 1) the glucose concentration slowly declined by 15%/h, 2) the glucose production was 65% that of healthy postabsorptive subjects, 3) carbons from glutamine and to a lesser extent from alanine were incorporated in glucose.
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METHODS
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Materials. D-[6,6-2H2]glucose, L-[3-13C]alanine, and L-[1,2-13C2]glutamine were purchased from MassTrace (Woburn, MA). Chemical and isotopic purity of the tracers was determined by gas chromatography-mass spectrometry (GC-MS). Before every infusion study, sterile solutions of the tracers were prepared with an aseptic technique. Accurately weighed amounts of the labeled compounds were dissolved in weighed volumes of sterile, pyrogen-free saline and filtered through a 0.22-µm Millipore filter before use. An aliquot of the sterile solution was initially verified to be pyrogen free before administration to human subjects. Solutions were prepared
24 h before use and were kept at 4°C before administration.
Subjects. The characteristics of the subjects under study are reported in Tables 1 and 2. Fourteen subjects were studied during the anhepatic phase of liver transplantation (LTx). The subjects were transplanted for localized hepatocarcinoma [single (<5) or multiple (<2 cm diameter)] (17), with the exception of two subjects who received the liver for carcinoid. It should be stressed that 86% of the patients had mild cirrhosis or no cirrhosis at all. Accordingly, the albumin and bilirubin concentrations, the prothrombin time, and the number of platelets reflected only a minor impairment in liver function. All subjects were at stable weight before transplantation. For most of them, a combination of the following medications was prescribed according to clinical indications: diuretics (spironolactone and/or furosemide), beta-blockers (propranolol or atenolol), lactulose, or ursodeoxycholic acid. The patients were following an isocaloric diet with
250 g of carbohydrate and 70-90 g of protein daily, with salt restriction. The last meal before transplantation was consumed 6-10 h before the skin incision, from which time 5 h on average elapsed for infusion of the tracer bolus during the anhepatic phase. Thus the study was performed 11-15 h from the last meal. The bolus of isotopes was repeated in 12 of the recipients 7 mo after transplantation (POST). Seven healthy control subjects were also studied with the same protocols (CON). A number of the LTx subjects and the POST and the CON subjects were simultaneously studied to trace the glutamine, alanine, and leucine kinetics. The complete set of results relative to the latter protocol has been reported (3). The protocol was approved by the institutional ethics committees.
Surgical model of the anhepatic phase of LTx. For the purpose of this work, the anhepatic phase began when the recipient's liver was removed and terminated when the circulation through the graft was reestablished by unclamping the portal vein (17). The duration of this phase was
45-75 min. To use this model, the major difficulties are caused by the short duration of the anhepatic phase (
60 min) and by the changed metabolic environment abruptly induced by liver removal. For a number of reasons, we decided to measure the glucose kinetics by administering tracers as a single injection during the anhepatic phase and not by using a primed-continuous infusion commencing before the anhepatic phase. First, with continuous infusions, the time required to reach a tracer steady state is several times greater than the duration of the anhepatic phase itself; thus the experiment should have begun hours before the anhepatic phase. When we considered the possible displacement of body fluids resulting from the surgical procedure (blood losses and replacement with hemoderivatives, and a fluid compartmentation due to portal and caval veins clamping and unclamping) before the anhepatic phase, we doubted that we could have excluded any carryover from the isotopic dilution before the anhepatic phase. For these reasons, the surgical medical teams agreed to limit the duration of the experiment to anhepatic phase and to warrant in that period a stable hemodynamic condition without blood losses and administration of hemoderivatives. During the anhepatic phase, the subjects received saline and other electrolytes (Normosol R; Abbott Laboratories, Abbott Park, sevoflurane anesthesia with remifentanil and pancuronium bromide, and, occasionally, calcium chloride, furosemide, and human albumin solutions.
Tracer injection and sampling protocol. At the beginning of anhepatic phase, two basal samples of arterial blood were drawn spacing 5 min apart. Immediately thereafter, a bolus of 2H2]glucose (69 mg/kg) was administered to quantitate the glucose kinetics. A bolus of L-[3-13C]alanine (90 µmol/kg, n = 7) or L-[1,2-13C2]glutamine (30 µmol/kg, n = 7) was simultaneously administered to evaluate whether the labeled carbons of these tracers were incorporated into gluconeogenic glucose during the anhepatic phase. bolus was delivered in a central vein in 20 s, immediately followed a flush with 15 ml of saline. Arterial blood was drawn at the following times after the bolus: 2, 3, 4, 6, 8, 10, 12, 15, 20, 25, 30, and every min thereafter until the end of the anhepatic phase, and 15 min the portal unclamping, when the circulation through the grafted was reestablished. In some subjects, 35-45 min after the beginning the anhepatic phase, blood samples were simultaneously drawn the arterial line and from the right renal vein (n = 6) or the portal (n = 5).
The same protocol was followed in the POST (n = 6 with alanine bolus, n = 4 with the glutamine bolus) and CON (n = 4 the alanine bolus, n = 3 with the glutamine bolus) subjects, with only difference being that CON received one-half of the alanine given the other groups (45 µmol/kg), and "arterialized" venous blood was drawn from a dorsal vein of the subject's hand that was cannulated in a retrograde way and heated in a warming box, as previously described (1). In both CON and POST subjects, blood was drawn to 150 min after the bolus.
Aliquots of blood were placed in tubes containing EDTA stored on ice until the plasma was prepared by centrifugation at 4° A 0.5-ml aliquot was withdrawn, defined amounts of [2H4]alanine [2H5]glutamine were added as an internal standard for quantitation alanine and glutamine plasma concentrations, and the plasma frozen at -60°C. One-milliliter blood aliquots for the measurements of glucagon and the catecholamines were placed in tubes containing EDTA plus aprotinin and in tubes containing glutathione, respectively. Blood aliquots for insulin, cortisol, and growth hormone collected in tubes without additives for serum separation. Blood aliquots for the determination of lactate, pyruvate, glycerol,
-hydroxybutyrate were placed in tubes containing perchloric All blood samples were placed on ice until the plasma or serum prepared by centrifugation at 4°C (within 1.5 h from drawing). plasma and serum aliquots were frozen at -60°C until later analysis.
Analytical methods. Plasma glucose concentration was measured with the hexokinase method (2) (Boehringer Mannheim, Mannheim, Germany).
The 2H2-glucose enrichments were measured by GC-MS after preparation of the butyl-boronate derivative (1). Injections of the samples were made into a GC-MS instrument (model 5970; Hewlett-Packard, Palo Alto, CA) operated by electron impact ionization as previously described. Glucose 13C enrichment was measured by GC-combustion associated to IRMS as described by us (13).
Plasma amino acid concentrations and enrichments were measured by electron impact GC-MS. Before derivatization, amino acids were isolated from plasma by use of cation-exchange columns as previously described (1). Amino acids eluted from the columns were evaporated to dryness and derivatized to form the tert-butyldimethylsilyl (TBDMS) derivative. The [M-57]+ ions at m/z = 260, 261, and 264 were monitored for unlabeled alanine, [3-13C]alanine, and [2H4]alanine, respectively. The [M-57]+ ions at m/z = 431, 433, and 436 were monitored for unlabeled glutamine, [1,2-13C2]glutamine, and [2H5]glutamine, respectively. TBDMS-glutamine was chromatographically resolved from TBDMS-glutamate. For all measurements, the background-corrected tracer enrichments in mole % excess were calculated as previously defined.
Plasma hormone concentrations were measured by radioimmuno-assay with commercial kits, as previously described (23). The catecholamine concentrations were measured by an HPLC method (12). The concentrations of whole body lactate, pyruvate, glycerol, and
-hydroxybutyrate were measured as previously described (15).
Data analysis. Because the liver utilizes and produces glucose, it stands to reason that its removal provokes a brisk change in both the plasma clearance rate (PCR) and the rate of appearance (Ra). The sudden change in PCR and Ra forces the glucose plasma concentration to change during the anhepatic phase toward a new level (dictated by the new ratio Ra/PCR); the rate of disappearance (Rd) changes accordingly. To quantify PCR, Ra, and Rd after liver removal, we analyzed the tracer disappearance curve, which, normalized to the administered tracer dose, is the unitary impulse response of the system. Characterization of the impulse response was performed using standard noncompartmental techniques (7). Briefly, a two-exponential function was found to be necessary and sufficient to describe satisfactorily the impulse response. The measurement error was assumed white, gaussian, of zero mean, and with experimentally determined standard deviation. Parameter estimation was performed by weighted nonlinear least squares. Weights were chosen optimally, i.e., equal to the inverse of the variance of the experimental error (6). The kinetic parameters that were derived from the analysis of the impulse response were the initial distribution volume [V1 (ml/kg)], the plasma clearance rate [PCR (ml·kg-1·min-1)], the total distribution volume [VT (ml·kg-1·min-1)], and the total body mass [QT (mmol/kg)]. Ra (mmol·kg-1·h-1) was estimated from the impulse response and tracee data by deconvolution (6), and Rd was then derived from the equation of the accessible pool: Rd(t) = Ra - V1·dC/dt, where C is the tracee concentration, and dC/dt is its rate of change during the anhepatic phase. The kinetic analysis of the impulse response measured in normal subjects and posttransplant patients followed the same approach as that just outlined. Because in such groups the tracee was in steady state, calculation of Ra and Rd from the impulse response was straightforward: Ra = PCR·C, and Rd = Ra. The kinetic analysis in the control groups (i.e., normal and posttransplant patients) was performed by relying on the tracer data collected in the first 75 min of the tracer decay curve. In this way, we ruled out the possibility that the different observation periods for tracer decay (150 min in the control groups and 60-75 min in patients during the anhepatic phase) could differently affect the estimation of the kinetic parameters in the three groups. It is worth pointing out that 75 min is a relatively short period for the analysis of glucose kinetics, and thus we were exposed to the risk of overestimating the slower component of glucose kinetics and, in turn, overestimating PCR. To assess the magnitude of such error, in the control groups we compared the clearances obtained by analyzing the first 75 min of the tracer decay curve with those obtained by analyzing the full set of data (150 min). We found that the clearances were never overestimated >10% when only the first 75-min data were used. We reasoned that, for proving our hypothesis, a <10% systematic error in glucose clearance (applied to both the study and control groups) was not relevant compared with the advantage of performing the study of the anhepatic metabolism with the tracer bolus technique.
Statistical analysis. We used t-tests for paired data to compare patients during LTx (ANHEP) and POST, and t-tests for independent data to compare ANHEP and CON, and we applied to both tests the Bonferroni correction. During the anhepatic phase, the changes over time in plasma substrate concentrations were defined as a significant correlation (P > 0.05) with time by standard linear regression.
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RESULTS
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Glucose concentrations. The time course of the glucose concentration in the plasma from arterial blood is reported in Fig. 1. After laparotomy, glucose concentration increased to reach a plateau of 174 ± 11 mg/dl, which was maintained until the portal vein was clamped, at which time the glucose concentration began to fall. During the anhepatic phase, glucose concentration decreased at the rate of 18 ± 3 mg·dl-1·h-1. Fifteen minutes after the liver replacement, the glucose concentration rose by 97 ± 10 mg/dl. At all times, the glucose concentration was significantly higher than in POST and in CON, whereas no difference was apparent between the two control groups.

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Fig. 1. Time course of glucose concentration. ANHEP, subjects during liver transplant; POST, recipients 7 mo after transplant; CON, healthy control subjects. Surgical stress increased glucose concentration until the portal vein was clamped. After liver removal, glucose concentration slowly declined, but it remained in the hyperglycemic range. Liver replacement immediately increased the glucose concentration.
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In six subjects, the glucose concentration in the plasma from the right renal vein was higher than in the artery (+3.7 ± 0.9 mg/dl, P < 0.01, Fig. 2). This subset of subjects had a mean glucose Ra of 7.5 ± 1.4 µmol·kg-1·min-1. In contrast, no significant difference in glucose concentration was apparent across the portal system by portal blood sampling.

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Fig. 2. A: arteriovenous difference in glucose concentration across the right (R) kidney. The renal venous sampling in 6 subjects demonstrated that the kidney is a net releaser of glucose during the anhepatic phase. B: arteriovenous difference in glucose concentration across the portal vein. Glucose concentration in the portal blood was not different from that in the arterial blood.
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Glucose kinetics. The time course of the concentration of [6,6-2H2]glucose after the bolus is reported in Fig. 3. The pattern of the bolus disappearance was similar among the groups, even though the area under the curve during the anhepatic phase was increased, indicating a 25% reduction in both the glucose space and glucose clearance, as shown in Table 3. The rate of glucose uptake was similar to that of control groups. The rate of glucose production was 30% decreased compared with the control groups.

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Fig. 3. Time course of labeled glucose after the bolus. The tracer disappearance profile was similar among the groups, even though the area under the curve during the anhepatic phase was increased, indicating a 25% reduction in glucose clearance.
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The carbons from alanine and glutamine were incorporated into glucose as shown in Fig. 4. The [13C]glucose concentration, normalized for the 13C dose infused, was markedly reduced compared with the control groups when the precursor was [13C]alanine, but it was similar to that of healthy subjects when the precursor was [13C2]glutamine. The area under the curve in the first 60 min after the bolus was only 10% of the area of the control groups with alanine infused, and it was similar to that of healthy control subjects when glutamine was infused. These data qualitatively show that the liver removal impaired alanine but not glutamine gluconeogenesis.

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Fig. 4. Incorporation in glucose of 13C from alanine (A) and glutamine (B). After the bolus of labeled alanine and glutamine (solid lines), 13C was incorporated in glucose (dashed lines). The [13C]glucose concentration (normalized to the bolus 13C dose) was markedly reduced compared with that of control groups when the precursor was [13C]alanine, but it was similar to that of healthy subjects when the precursor was [13C2]glutamine. The area under the curve in the first 60 min after the bolus was only 10% that of control groups with alanine infused, and it was comparable to that of healthy control subjects when glutamine was infused. These data qualitatively show that liver removal impaired alanine but not glutamine gluconeogenesis.
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Gluconeogenic precursors. The time course of glutamine and alanine is reported in Fig. 5. In ANHEP, the alanine concentration was three times greater and the glutamine concentration was 50% greater than in the control groups. The liver replacement produced a marked reduction in circulating alanine (-20 ± 4%, P < 0.001) and glutamine (-22 ± 4%, P < 0.001) concentration within 15 min. The time course of lactate, pyruvate, and glycerol is shown in Fig. 6. The three metabolites were increased severalfold during the anhepatic phase. The liver replacement further increased the concentration of lactate and pyruvate, whereas it decreased that of glycerol.

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Fig. 5. Time course of alanine and glutamine concentrations. In ANHEP, the alanine concentration was 3 times greater and the glutamine concentration was 50% greater than in the control groups. Liver replacement produced a rapid reduction in circulating alanine and glutamine.
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Fig. 6. Time course of lactate, pyruvate, and glycerol concentrations. The 3 metabolites were increased severalfold during the anhepatic phase. Liver replacement further increased the concentration of lactate and pyruvate, whereas it decreased that of glycerol.
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Hormones. The hormone concentrations are reported in Fig. 7: insulin and all of the counterregulatory hormones were markedly increased during the anhepatic phase. The hormone concentrations were similar in CON and in POST, with the exception of a modest increment in insulin and in C-peptide in POST.

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Fig. 7. Hormone concentrations. Insulin and all of the counterregulatory hormones were markedly increased during the anhepatic phase. Hormone concentrations were similar in CON and in POST, with the exception of a modest increment in insulin and in C-peptide in POST.
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DISCUSSION
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In this study we showed that, in certain conditions, the extrahepatic tissues can synthesize an amount of glucose that is close to the whole body glucose production in healthy postabsorptive subjects. Our conclusion is based on several observations. First, we found that, during the anhepatic phase, the glucose concentration remained in the hyperglycemic range. The glucose concentration diminished during that period of time, but it did so at a rate that was very slow compared with what we would have expected if the glucose production had been consistently reduced with the liver removal. The decrement in circulating glucose was only 15%/h, whereas we previously found that it decreased by 30-40% within 30 min during hypoglycemic clamp studies in which the glucose production was reduced by 50% and the glucose disappearance was not appreciably changed (1, 4). This slow reduction in circulating glucose during the anhepatic phase might have occurred either because of a substantially maintained glucose production or because of a markedly decreased glucose uptake. The bolus of labeled glucose allowed us to discriminate between the two possibilities on the basis of direct and model-independent considerations. The area under the tracer concentration curve was 25% higher than in the control groups, indicating an
25% reduction in glucose clearance. This finding was expected, because deep anesthesia should have reduced the brain glucose uptake. Because the circulating glucose concentration increased 30% on average, we deduced that the glucose disappearance resulting from an increased glucose concentration and an equally decreased glucose clearance had to be grossly comparable to that in the control groups. Glucose production had to be similar to the glucose disappearance minus the amount of glucose that disappeared from the glucose space during the anhepatic phase. Assuming a volume of 0.1 l/kg for the glucose-accessible pool, a glucose decrement of 20 mg·dl-1·h-1 would have indicated a glucose production
0.3 mg·kg-1·min-1 smaller than the disappearance. On the basis of these qualitative observations, we could conclude that, during the anhepatic phase, glucose is released at a rate close to that of postabsorptive subjects. The calculations based on the two-exponential model of the glucose system quantified the glucose fluxes and confirmed the qualitatively drawn conclusions. We excluded that glucose was administered from any exogenous source during the anhepatic phase. Thus all of the released glucose had to be of endogenous origin. To qualitatively prove that it was at least in part of gluconeogenic origin, we evaluated whether any carbon transfer occurred between simultaneously infused 13C-labeled alanine or glutamine tracers and glucose. Because we found detectable 13C enrichments into glucose (remarkably in the experiments in which glutamine was infused), we concluded that extrahepatic organs have the potential to produce glucose from amino acids and especially from glutamine.
We sampled the renal and the portal venous blood to locate the possible sources of the glucose released during the anhepatic phase. We found an appreciable glycemic gradient across the right kidney that was greater than previously found in postabsorptive, starved, or hypoglycemic humans (8, 11, 18, 19, 25, 26). With the assumption of a conversion factor of 0.85 between plasma and whole blood glucose concentration and a renal blood flow of 1.5 l/min during the anhepatic phase, a plasma glucose gradient of 3.7 mg/dl in 70-kg individuals would give a net glucose release of 3.7 µmol·kg-1·min-1. When we consider the fact that the kidney may utilize 0.5-1.5 µmol·kg-1·min-1 (8, 19), glucose production could have been as large as 5.2 µmol·kg-1·min-1, i.e., the majority (70%) of the observed (7.5 µmol·kg-1·min-1) endogenous glucose production. Given the analytical precision of the methods employed, it is possible that glucose release from the kidney could have accounted for all of the endogenous glucose release, although it is possible that minor contributions were provided from other sources. The gradient across the portal vein-drained viscera was not significantly different from zero. The absence of a significant net extraction of glucose from the portal blood does not exclude a gut glucose release, consistent with the recent demonstration that the glucose-6-phosphatase gene is expressed in the human intestine (24) and that the rat intestine is a gluconeogenic organ (10).
Regarding the gluconeogenic precursors, lactate is one important candidate, because it is very abundant during the anhepatic phase, and it was shown that it is a primary gluconeogenic precursor in the kidney of dogs (9) and humans (27). We were interested in alanine and especially in glutamine, because the latter substrate is the most important carrier of protein-derived carbons for gluconeogenesis (21). Glutamine and alanine also shuttle the amino groups derived from amino acid transamination to the organs capable of disposing nitrogen. Once deaminated, the glutamine and the alanine carbon skeletons are suitable gluconeogenic precursors, and gluconeogenesis may actually represent a means to recycle these carbons without oxidizing them. We studied the disposal of alanine and glutamine during the anhepatic phase, and we found that their fluxes were actually increased in such a condition (3). Glutamine and alanine disposal and cycling through gluconeogenic precursors occur not only within the liver with its periportal and perivenous hepatocytes, but also in the kidney, in the gut, and in the brain (20, 28). These are the candidate organs for nonhepatic glucose production, and all of them express to various degrees the enzymes needed to handle amino nitrogen and to release the newly synthesized glucose (5, 24).
Despite the marked hyperinsulinemia, the counterregulatory hormones increased the glucose production before the anhepatic phase. As expected, we found that surgery per se increased the glucose concentration. This was probably due to the combined effect of glucagon, epinephrine, and cortisol (16). When the liver was removed, part of this effect could still be maintained through a possible effect of increased epinephrine on renal gluconeogenesis (26, 27). It should be noted that, during the anhepatic phase, the glucose production was close to that of postabsorptive subjects, but this does not strictly mean that it was appropriate to the degree of surgical stress and the consequent increment in the counterregulatory hormones. The increment in glucose concentration after the skin incision until the removal of the liver was probably sustained by an increment in glucose production. Thus our experiments represented a sort of challenge for the extrahepatic organs to produce the maximum possible amount of glucose, being strongly stimulated by the counterregulatory hormones, surgical stress, and abundance of gluconeogenic precursors.
In conclusion, we demonstrated that, in the absence of the liver, extrahepatic organs can release glucose at a rate close to that of healthy postabsorptive subjects. These results support the concept of hepatorenal reciprocity, according to which the renal glucose release compensates for the failure of hepatic glucose release, and vice versa, to preserve normoglycemia (29). The kidney accounts for most and possibly all of endogenous glucose release in anhepatic individuals due to an increase in the availability of gluconeogenic precursors and catecholamines and cortisol, which would be expected to increase renal gluconeogenesis. Part of the glucose was synthesized from circulating alanine, and a larger amount was synthesized from glutamine.
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GRANTS
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This work was supported in part by grants from the Italian Ministero della Sanità (RF 97.4-2126) and from Associazione Italiana Ricerca Cancro (98-01 no. 163).
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FOOTNOTES
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Address for reprint requests and other correspondence: A. Battezzati, Amino Acids and Stable Isotopes Laboratory, San Raffaele Scientific Institute, Via Olgettina, 60, 20132 Milano, Italy (E-mail: battezzati.alberto{at}hsr.it).
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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