Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, Tennessee 37232
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ABSTRACT |
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During chronic total parenteral nutrition
(TPN), net hepatic glucose uptake (NHGU) and net hepatic lactate
release (NHLR) are markedly reduced (~45 and ~65%,
respectively) with infection. Because small quantities of fructose are
known to augment hepatic glucose uptake and lactate release in normal
fasted animals, the aim of this work was to determine whether acute
fructose infusion with TPN could correct the impairments in NHGU and
NHLR during infection. Chronically catheterized conscious dogs received
TPN for 5 days via the inferior vena cava at a rate designed to match daily basal energy requirements. On the third day of TPN
administration, a sterile (SHAM, n = 12) or
Escherichia coli-containing (INF, n = 11)
fibrin clot was implanted in the peritoneal cavity. Forty-two hours
later, somatostatin was infused with intraportal replacement of insulin
(12 ± 2 vs. 24 ± 2 µU/ml, SHAM vs. INF, respectively) and
glucagon (24 ± 4 vs. 92 ± 5 pg/ml) to match concentrations previously observed in sham and infected animals. After a 120-min basal
period, animals received either saline (Sham+S, n = 6;
Inf+S, n = 6) or intraportal fructose (0.7 mg · kg
1 · min
1; Sham+F,
n = 6; Inf+F, n = 5) infusion for 180 min. Isoglycemia of 120 mg/dl was maintained with a variable glucose
infusion. Combined tracer and arteriovenous difference techniques were
used to assess hepatic glucose metabolism. Acute fructose infusion with
TPN augmented NHGU by 2.9 ± 0.4 and 2.5 ± 0.3 mg · kg
1 · min
1 in Sham+F
and Inf+F, respectively. The majority of liver glucose uptake was
stored as glycogen, and NHLR did not increase substantially. Therefore,
despite an infection-induced impairment in NHGU and different hormonal
environments, small amounts of fructose enhanced NHGU similarly in sham
and infected animals. Glycogen storage, not lactate release, was the
preferential fate of the fructose-induced increase in hepatic glucose
disposal in animals adapted to TPN.
liver; lactate release; dog; glycogen
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INTRODUCTION |
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THE PRESENCE OF AN INFECTION substantially alters hepatic carbohydrate metabolism. In stressed states such as trauma, injury, or infection, nutritional support is often provided to patients in the form of total parenteral nutrition (TPN). During feeding or glucose infusion, the liver removes glucose from the bloodstream and thereby limits the severity of hyperglycemia. In healthy human volunteers, the liver removes 30-35% of an acute oral glucose load (23). When nutrients are continuously infused on a chronic (i.e., days) basis, glucose uptake by the liver is markedly augmented. The liver of a TPN-adapted dog consumes 45% of the infused glucose, and ~70% of liver glucose uptake undergoes glycolytic conversion to lactate, which is released into the peripheral circulation (18). In contrast, the presence of an infection reduces net hepatic glucose uptake (NHGU) to only 25% of the glucose infused in chronic TPN-infused dogs and reduces net hepatic lactate release (NHLR) to ~45% of NHGU (18). Analogously, splanchnic (gut and liver) glucose uptake in stressed patients receiving chronic TPN accounts for ~20% of the glucose infused (13).
Despite the importance of the liver as a site of glucose disposal and
the impact of infection on liver metabolism, no studies have attempted
to specifically improve hepatic glucose disposal during nutritional
support. The stress-induced impairment in NHGU with chronic nutritional
support may be caused by inadequate suppression of hepatic glucose
production (HGP) and/or by a failure to stimulate hepatic glucose
uptake (HGU). Previous studies (19, 26) indicate that both
processes contribute to the reduced NHGU. In response to a large (8 mg · kg1 · min
1) glucose
infusion, gluconeogenesis was not suppressed to the same extent in
septic patients as in normal volunteers (25). Strategies
that improve the suppression of HGP and/or enhance HGU would facilitate
NHGU and thus improve whole body glucose disposal.
Recent evidence suggests that low concentrations of fructose (<0.5 mM) augment NHGU in vivo (27), presumably by activation of glucokinase (32). Because TPN-adapted animals exhibit elevated NHGU, it is unknown whether fructose can further augment NHGU and NHLR in a chronic TPN setting. Moreover, whether fructose can enhance NHGU in the presence of an infection has not been determined. Therefore, our work addressed the following questions: 1) does fructose enhance NHGU during TPN administration; 2) does infection impair the ability of fructose to stimulate NHGU; and 3) does fructose infusion improve hepatic glucose disposition?
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METHODS |
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Animal preparation.
Twenty-three female mongrel dogs were fed a standard Kal-Kan meat
(Vernon, CA) and Purina Lab Canine #5006 (Purina Mills, St. Louis, MO)
diet once daily and had free access to water. The composition of the
diet, based on dry weight, was 52% carbohydrate, 31% protein, 11%
fat, and 6% fiber. Dogs were housed in a facility that met the
guidelines of the Association for the Accreditation of Laboratory
Animal Care International. The protocols were approved by the
Vanderbilt University Medical Center Animal Care Committee. Health of
the animals was determined before surgery and before TPN administration
as having a good appetite (i.e., consumed 75% of the daily ration)
and normal stools, hematocrit >0.35, and leukocyte count
<18,000/mm3.
Surgical preparation. A laparotomy was performed under general anesthesia (15 mg/kg iv of thiopental for induction and 1.0% isoflurane as an inhalant during surgery) on healthy dogs weighing 17-24 kg. During the laparotomy, Silastic (Dow Corning, Midland, MI) infusion catheters (0.03 in. ID) were placed in the splenic and jejunal veins, and the gastroduodenal vein was ligated. Catheters (0.04 in. ID) for blood sampling were positioned in the portal and left common hepatic veins. Two infusion catheters (0.04 in. ID) for TPN were placed in the inferior vena cava (IVC), and the free ends were exteriorized and tunneled subcutaneously behind the left clavicle. Flow probes (Transonic Systems, Ithaca, NY) were positioned about the portal vein, hepatic artery, and right external iliac artery. Incisions were made in the right and left inguinal regions, and a sampling catheter (0.04 in. ID) was placed in the left common iliac vein as the tip was advanced distal to the IVC anastomosis; another catheter was placed in the right external iliac artery, and its tip was advanced to the abdominal aorta.
All catheters were filled with 0.9% NaCl (saline) containing heparin (200 U/ml; Elkins-Sinn, Cherry Hill, NJ). The free ends of the catheters and flow probes were exteriorized and placed in subcutaneous pockets. The dogs received penicillin G (600,000 U iv; Anthony Products, Irwindale, CA) to minimize the possibility of infection. Flunixin (0.1 mg/kg body wt; Fort Dodge Lab, Fort Dodge, IA) was injected intramuscularly immediately after wound closure for acute pain relief. Dogs received ampicillin (500 mg; Bristol-Myers Squibb, Princeton, NJ) orally for 3 days after surgery.Nutritional support.
After an allowance of 14 days for recovery from surgery, the IVC
catheters were exteriorized under local anesthesia (2% lidocaine, Abbott, North Chicago, IL). TPN was infused as the sole source (i.e.,
all enteral nutrients were discontinued) of calories into one or both
of these catheters continuously for 5 days by use of an ambulatory
infusion pump (Dakmed, Buffalo, NY, or Walkmed-350, McKinley, Lakewood,
CO). Dogs wore a jacket (Alice King Chatham, Los Angeles, CA) with two
large pockets for the nutrition and the pump. The TPN was designed to
be isocaloric on the basis of predicted resting energy expenditure
(30). The composition of the TPN included glucose, lipids,
amino acids, saline, potassium phosphate (90 mg · kg
1 · day
1), and a
multivitamin supplement (MVI-12, Astra, Westborough, MA). Glucose (50%
dextrose, Abbott, ~10
mg · kg
1 · min
1) made up
75% of the nonprotein calories, whereas 20% Intralipid (Baxter,
Deerfield, IL) constituted the remaining 25% of the energy requirements. Travasol (10%, Baxter) was infused to supply basal nitrogen requirements (1.5 × body wt0.67 g
protein/day). During the 5 days of TPN, the animals had free access to water.
Induction of infection. A 1% fibrinogen (Sigma, St. Louis, MO) solution was filtered (0.45 µm) under sterile conditions. To initiate clot formation, thrombin (1,000 U, Gentrac, Middleton, WI) was added to the filtrate. The bacterial clot also contained a nonlethal dose (2 × 109 organisms/kg body wt) of Escherichia coli determined by serial dilution followed by plating. Bacteria (American Type Tissue Culture no. 25922) were prepared by inoculation of 1 liter of Trypticase soy broth (Becton-Dickinson, Cockeysville, MD) and incubation overnight at 37°C. Bacteria were pelleted by centrifugation on the next day and were washed with and reconstituted in sterile saline before addition to the filtrate.
On the third day of the 5-day TPN infusion, a second laparotomy was performed under anesthesia, and the TPN infusion was continued. An abdominal midline incision was made at a point below the incision made during the previous surgery. Either a sterile (SHAM) or a bacterial (INF) fibrin clot was implanted in the peritoneal cavity. In addition to the TPN, animals received saline intravenously during the laparotomy and on the next day.Experimental protocol. On the 5th day of TPN and 42 h after clot implantation, a study was performed. Free ends of all catheters were exteriorized under local anesthesia, and their contents were aspirated and flushed with saline. Leads from the flow probes were also exteriorized and connected to a flowmeter. The dog was placed in a Pavlov harness for the duration of the study. Angiocaths (18 gauge, Abbott) were inserted into both cephalic veins for infusion of radioactive tracers, glucose, and somatostatin (SRIF; Bachem, Torrance, CA). Blood pressure, heart rate (Micro-Med, Louisville, KY), and rectal temperature (Yellow Springs Instruments, Yellow Springs, OH) were assessed during the basal period.
The study consisted of a 120-min basal period followed by a 180-min experimental period. Primed (44 and 27 µCi), constant infusions of [3-3H]- and [U-14C]glucose (New England Nuclear, Wilmington, DE) were begun at rates of 0.4 and 0.3 µCi/min, respectively, with syringe pumps (Harvard Apparatus, Holliston, MA) and continued for the duration of the study. A fresh TPN solution containing the nonglucose nutrients was prepared separate from the glucose. In this way, the glucose infusion rate was adjusted to maintain isoglycemia (120 mg/dl), whereas the other TPN components were infused at a constant rate. Small blood samples (0.4 ml) were taken every 10 min and centrifuged immediately to measure arterial plasma glucose concentration with a Beckman glucose analyzer II (Beckman Instruments, Fullerton, CA). The exogenous glucose infusion rate was adjusted to maintain isoglycemia. Saline was infused to replace blood volume withdrawn by sampling. During tracer equilibration, SRIF (0.8 µg · kgSample processing.
Blood samples were placed in chilled tubes containing EDTA and were
processed as described in Ref. 16. For analysis of
catecholamines, whole blood (1 ml) was treated with 20 µl of a
solution containing 90 mg/ml of EGTA and 50 mg/ml of glutathione.
Samples for gluconeogenic metabolite, fructose, and glutamine content
were processed by adding 1 ml of whole blood to 3 ml of 4% perchloric
acid (PCA). Blood 14CO2 was assessed in
triplicate on arterial, portal, and hepatic vein samples, as described
by Chan and Dehaye (4). To assess 14C
incorporation into lactate and amino acids, 3 ml of blood were added to
3 ml of 8% PCA. The sample was centrifuged, neutralized with 2 N KOH,
and placed over ion exchange columns to separate into
[14C]lactate, [14C]glucose, and
14C-labeled amino acids (22). Blood samples
were centrifuged at 3,000 rpm for 10 min, and the remaining plasma was
stored at 70°C for later analyses. Plasma treated with aprotinin
(500 kallikrein inactivator units/ml plasma; Miles, Kankakee, IL) was
analyzed for glucagon content. Plasma glucose specific activity (SA)
was measured after deproteinization with Ba(OH)2 and
ZnSO4 and removal of charged intermediates by means of
anionic and cationic resins (29).
Analysis. Immunoreactive insulin and glucagon were assayed using a double antibody disequilibrium procedure (21) [coefficients of variation (CV) 9 and 8%, respectively], and cortisol was assayed using a single antibody technique (12) with a Diagnostic Products radioimmunoassay kit (Los Angeles, CA; CV 8%). HPLC techniques were used to assess plasma epinephrine and norepinephrine (CV 11 and 6%, respectively) (14). Analysis of lactate, alanine, and glycerol was performed on an automated centrifugal analyzer (Monarch 2000; Instrumentation Laboratory, Lexington, MA) by using a modification of the method of Lloyd et al. (15). The concentration of nonesterified fatty acids (NEFA) was determined spectrophotometrically (Wako Chemicals, Richmond, VA).
The methods of Bernt and Bergmeyer (1) and of Beutler (2) were adapted to measure blood glutamine and fructose (lower detection limit = 15 µM) content, respectively, with a Technicon Autoanalyzer II (Bran Luebbe, Buffalo Grove, IL). Fructose content was measured in blood deproteinized with 4% PCA. The extract was neutralized with 10% KOH and incubated for 60 min with an equal volume of 0.1 M phosphate buffer (pH 7.4) containing glucose oxidase (10 U/ml) and catalase (600 U/ml). Glycogen content was determined using the enzymatic method of Chan and Exton (5). Hepatic lipids were extracted with the Folch method (11), and the 14C radioactivity of the extract was determined with liquid scintillation counting.Calculations.
The rates of total glucose appearance (Ra) and utilization
(Rd) were determined according to the method of Wall et al.
(33), as simplified by de Bodo et al. (9).
Whole body endogenous glucose production (EGP) was the difference
between Ra and exogenous glucose infusion rate. The
substrate (glucose, lactate, alanine, glycerol, fructose, and NEFA)
load entering the liver was calculated as the sum of the loads in the
hepatic artery and portal vein, (As × HABF) + (Ps × PBF), where As and Ps
represent the substrate concentrations in the iliac artery and portal
vein, and HABF and PBF represent blood flow in the hepatic artery and
portal vein. Similarly, the substrate load leaving the liver was
Hs × THBF, in which Hs and THBF represent
the hepatic vein substrate concentration and total hepatic blood flow
(THBF = HABF + PBF). Net hepatic substrate balance was
calculated as the difference between the entering and exiting substrate
loads and was denoted as either uptake or output. Likewise, net
hindlimb substrate uptake was calculated using the formula
(As Vs) × ABF, where
Vs is the substrate concentration in the iliac vein and ABF
is the iliac artery blood flow. Net hepatic fractional extraction (HFE)
of substrate was calculated as the net hepatic substrate balance divided by the substrate load entering the liver. Plasma flow was used
for NEFA balance by multiplying blood flow and (1
hematocrit).
Statistics. Results are expressed as means ± SE in the basal period (average of 3 sampling points) and during the last 60 min of the experimental period (average of 3 sampling points). Fructose concentrations are expressed as means ± SE for the last 120 min of the experimental period. Basal data are presented as average of sham (SHAM, n = 12) and infected (INF, n = 11) animals, unless otherwise indicated. Student's unpaired t-test was used for comparisons of sham and infected animals in the basal period. To determine the effect of fructose, statistical comparisons over time were made with two-way ANOVA (SYSTAT, Evanston, IL) between fructose-infused groups and their corresponding saline-infused groups. Statistical significance was designated as a P value < 0.05. Results from infected + saline (Inf+S) and sham + saline (Sham+S) groups have been presented previously (10).
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RESULTS |
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General characteristics.
Body temperature, heart rate, mean arterial pressure, and blood flow in
the hepatic artery and portal vein are shown in Table 1. Infected animals were typically
hyperthermic, normotensive, and tachycardic, with significantly
elevated hepatic arterial blood flow. There were no changes in blood
flow over time in any group (data not shown).
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Hormones.
As expected with the infusion rates, arterial plasma insulin
concentrations were higher (P < 0.05) in the basal and
experimental periods of the INF group, 24 ± 2 and 25 ± 2 µU/ml, than in the SHAM group, 12 ± 2 and 13 ± 2 µU/ml
(Fig. 1). Basal arterial plasma glucagon
concentrations were also higher in INF (24 ± 4 vs. 92 ± 5 pg/ml, SHAM vs. INF, respectively; P < 0.05) and did
not change in the experimental period (22 ± 3 and 89 ± 7 pg/ml). The calculated insulin (40 ± 2 and 100 ± 14 µU/ml) and glucagon (32 ± 3 and 285 ± 26 pg/ml)
concentrations in the portal vein for the SHAM and INF groups were also
higher. Basal concentrations of cortisol, epinephrine, and
norepinephrine were similar in SHAM and INF groups (Table 1) and did
not change in the experimental period (data not shown).
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Fructose.
Arterial fructose concentrations in the basal period were 41 ± 11 and 23 ± 10 µM in Sham+F (n = 6) and Inf+F
(n = 3). During the final 120 min of the intraportal
fructose infusion, arterial fructose concentrations increased modestly
to 101 ± 10 and 116 ± 24 µM, Sham+F and Inf+F,
respectively; Fig. 2. Fructose
concentrations reached 321 ± 10 and 367 ± 36 µM in the
portal vein and 91 ± 8 µM and 127 ± 29 µM in the
hepatic vein. Hepatic fructose loads were similar (7.8 ± 0.6 vs.
9.2 ± 2.0 µmol · kg1 · min
1, Sham+F
vs. Inf+F). Net hepatic fructose uptake (4.9 ± 0.3 vs. 4.6 ± 1.7 µmol · kg
1 · min
1)
and HFE of fructose (0.63 ± 0.03 vs. 0.46 ± 0.11) were also similar.
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Whole body glucose kinetics.
Arterial plasma glucose concentrations were 120 ± 2 mg/dl in both
SHAM and INF groups and were maintained for the duration of the study
(Fig. 3). Exogenous glucose infusion rate
(GIR) was 8.6 ± 0.6 and 11.5 ± 0.6 mg · kg1 · min
1 in SHAM
(n = 11) and INF (n = 11, P < 0.05) and did not increase with saline infusion.
In the fructose-infused groups, GIR increased by 3.7 ± 1.0 and
2.1 ± 0.6 mg · kg
1 · min
1, Sham+F and
Inf+F [not significant (NS)]. Similar to GIR, basal glucose
Rd was 10.3 ± 0.6 and 13.0 ± 0.9 mg · kg
1 · min
1 in SHAM and
INF (P < 0.05) and did not change with saline
infusion. During fructose infusion, Rd increased by
3.8 ± 0.6 and 2.9 ± 0.6 mg · kg
1 · min
1, Sham+F and
Inf+F, respectively. Basal endogenous glucose production rates were
similar (1.5 ± 0.3 and 1.6 ± 0.7 mg · kg
1 · min
1, SHAM and
INF) and did not change significantly with fructose (
0.7 ± 0.5 vs.
0.9 ± 0.4 mg · kg
1 · min
1, Sham+F vs.
Inf+F) or saline (
0.1 ± 0.4 vs.
0.4 ± 0.2 mg · kg
1 · min
1, Sham+S vs.
Inf+S) infusion.
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Hepatic glucose balance.
Basal NHGU was higher in SHAM (3.7 ± 0.3 mg · kg1 · min
1) than in
INF (1.7 ± 0.3 mg · kg
1 · min
1;
P < 0.05; Fig. 3). This was paralleled by higher basal
unidirectional HGU, 3.9 ± 0.3 vs. 2.2 ± 0.2 mg · kg
1 · min
1, SHAM vs.
INF (P < 0.05). There were no changes in NHGU over time in the groups that received saline. Within 30 min of fructose infusion, NHGU markedly increased (P < 0.05) in both
Sham+F and Inf+F groups and remained elevated for the duration
of the study. NHGU increased to 6.8 ± 0.7 mg · kg
1 · min
1 in Sham+F
and 4.3 ± 0.4 mg · kg
1 · min
1 in Inf+F
(
2.9 ± 0.4 and
2.5 ± 0.3 mg · kg
1 · min
1,
respectively). Net HFE of glucose in the basal period was 0.14 ± 0.01 and 0.05 ± 0.01 in SHAM and INF (P < 0.05).
Analogous to NHGU, HFE increased during fructose infusion (
0.10 ± 0.01 and
0.08 ± 0.02, Sham+F and Inf+F, respectively).
Basal HGP was similar in SHAM and INF (0.2 ± 0.2 and 0.5 ± 0.2 mg · kg
1 · min
1) and
did not change significantly with fructose or saline (data not shown).
Hepatic glucose disposition.
Arterial lactate concentrations in the basal period were higher in SHAM
relative to INF (892 ± 62 vs. 669 ± 64 µM;
P < 0.05; Fig. 4). When
fructose was infused, arterial lactate concentrations rose
(P < 0.05) by 340 ± 107 and 324 ± 31 µM
in Sham+F and Inf+F. Basal net hepatic lactate release (NHLR) was
23 ± 2 µmol · kg1 · min
1 in SHAM
and did not change significantly in the experimental period
(
1.3 ± 1.0 and
1.3 ± 1.7 µmol · kg
1 · min
1, Sham+F
and Sham+S). As expected, basal NHLR was lower during infection
(10 ± 3 µmol · kg
1 · min
1,
P < 0.05). NHLR tended to increase with fructose
infusion (
7 ± 1 µmol · kg
1 · min
1,
Inf+F), although this was not significant with respect to Inf+S (
1 ± 2 µmol · kg
1 · min
1).
Hepatic glucose oxidation rates in the basal period were 0.4 ± 0.1 and 0.5 ± 0.1 mg · kg
1 · min
1 in SHAM
(n = 12) and INF (n = 10) and did not
increase with saline or fructose infusion (data not shown).
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Hepatic metabolites.
Arterial alanine concentrations in blood were lower with infection
(471 ± 48 vs. 205 ± 15 µM, SHAM vs. INF;
P < 0.05) and did not change during the experimental
period. Basal net hepatic alanine uptake rates (1.4 ± 0.2 and
2.7 ± 0.3 µmol · kg1 · min
1;
P < 0.05) and net HFE of alanine (0.20 ± 0.02 vs. 0.29 ± 0.02, P < 0.05; Table
2) were higher during infection and did
not increase with fructose infusion.
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Hindlimb.
Net hindlimb glucose uptake rates during the basal period in SHAM and
INF were 14 ± 2 and 16 ± 2 mg/min, and net hindlimb fractional extraction of glucose was 0.09 ± 0.02 and 0.12 ± 0.02. Basal net hindlimb lactate uptake was 2.0 ± 0.3 and
0.5 ± 0.3 µmol · kg1 · min
1 in SHAM
(n = 11) and INF (n = 11); net hindlimb
fractional extraction of lactate was 0.37 ± 0.07 and 0.09 ± 0.05. Fructose infusion did not alter these variables (data not shown).
Muscle glycogen mass was not different between groups (data not shown).
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DISCUSSION |
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Although chronic TPN administration markedly enhances NHGU and NHLR in dogs, the presence of an infection sharply reduces both processes despite elevated insulin and glucose concentrations (18). Fructose is known to augment liver glucose uptake in non-TPN adapted animals. In the present study, we demonstrated that fructose enhanced NHGU during chronic TPN administration in both sham and infected dogs. Although acute fructose infusion overcame the infection-induced impairment in NHGU, it did not correct the impairment in NHLR.
Unexpectedly, the ability of fructose to enhance glucose disposal in
normal TPN-adapted animals was comparable to the effect observed in
fasted glucose-infused animals. Acute fructose infusion during chronic
TPN increased NHGU by 2.9 mg · kg1 · min
1 in the
presence of slightly elevated glucose (120 mg/dl) and insulin (12 µU/ml) concentrations. In 42-h-fasted dogs in the presence of greater
hyperglycemia (225 mg/dl) and hyperinsulinemia (30 µU/ml), a similar
fructose infusion raised liver glucose uptake by 7 mg · kg
1 · min
1
(27). The increments in HFE of glucose, however, were
similar (
0.14 vs.
0.10; fasted vs. TPN), suggesting that
nutritional support does not substantially blunt the response to
fructose. The mechanism by which fructose activates NHGU is presumably
via an increase in fructose 1-phosphate concentration, which allows translocation and activation of glucokinase (31, 32). The ability of fructose to stimulate NHGU beyond that seen with TPN implies
that, despite the already high NHGU with TPN, the majority of GK
protein is not already maximally translocated.
Given the different hormone environments, the response to fructose in
infected animals was remarkably similar to that of sham animals.
Insulin and glucagon concentrations were clamped at the elevated levels
seen during infection, allowing us to examine the effects of fructose
in the absence of changes in these hormones. As we have observed
previously (18), infection impaired NHGU by ~50%. Given
the impaired NHGU and the known in vitro effects of glucagon to
partially reverse the effect of fructose (3, 8), we
expected infection to impair the response to fructose; in fact,
infection did not impair the response to fructose. It is possible that
the acute inhibitory effect of glucagon on fructose action does not
persist chronically. Consistent with this, the acute and chronic
effects of glucagon are markedly different. Acute changes in glucagon
primarily alter hepatic glycogen metabolism, the primary fate of
glucose in response to fructose. In contrast, glucagon chronically
modulates gluconeogenesis (17, 20). In four additional
experiments, acute reduction (40%) of the glucagon replacement rate
did not alter the ability of fructose to stimulate NHGU (data not
presented). However, the glucagon replacement rate was still
substantially higher than in sham animals receiving chronic TPN;
therefore, the inhibitory effect of glucagon may still be present.
Nevertheless, it is also possible that the underlying impairment in
NHGU seen with infection is distinct from the site of action of fructose.
The effect of fructose was mediated rapidly and specifically on the
liver, as hindlimb glucose uptake was not altered by acute fructose
infusion. This is consistent with a previous report in the dog
(27). Circulating arterial levels of fructose were
relatively low (~100 µM) because of the high first-pass extraction
by the liver. The fructose-induced increase in NHGU (2.9 ± 0.4 and
2.5 ± 0.3 mg · kg
1 · min
1 in Sham+F
and Inf+F) roughly accounted for the increase in whole body glucose
disposal (
3.8 ± 0.6 and 2.9 ± 0.6 mg · kg
1 · min
1).
Once consumed by the liver, glucose undergoes one of three possible fates: oxidation, release as lactate, or storage in the form of glycogen or lipid. Acute fructose infusion did not increase hepatic glucose oxidation. Given the marked activation of glycolysis during TPN administration, we expected fructose infusion to substantially increase lactate release; however, an increase was not observed. In fact, in Sham+F, the proportion of NHGU diverted to lactate fell in response to fructose infusion, whereas the proportion of [14C]glucose diverted to [14C]lactate was unaltered. These data suggest that the lack of a rise in NHLR, despite a rise in NHGU, was probably a result of an inhibition of hepatic glycogenolysis and subsequent conversion to lactate. In contrast, in infected animals, although basal NHLR was lower, NHLR tended to increase in Inf+F during fructose infusion. Thus glycogenolysis may not have decreased to the same extent as in Sham+F. In previous studies in fasted animals (27), fructose infusion did enhance NHLR, possibly because hepatic glycogenolysis was already suppressed by the accompanying hyperglycemia and hyperinsulinemia. Why glycogenolysis remains active in the TPN-adapted state is unknown.
Sham and infected animals demonstrated a twofold increase in hepatic
glucose storage in response to fructose. The increase reflects greater
glycogen deposition because tracer incorporation into lipid was
minimal. Glycogen was also the major glucose fate during acute fructose
infusion in fasted dogs (27). Fructose is known to enhance
both the direct and indirect pathways of glycogen synthesis
(24), and our data support increases in both pathways. Although fructose increased tracer incorporation into glycogen in
Sham+F (2.0 mg · kg1 · min
1), which
accounts for the majority of the increase in NHGU, we did not observe a
net increase in glycogen mass. It is possible that 1)
near-maximal hepatic glycogen capacity is achieved with chronic TPN or
2) we lacked the statistical power to detect an increase in
glycogen mass in Sham+F because of a large variance. In contrast, the
15 mg/g increase in hepatic glycogen mass in Inf+F approximates the
theoretical increase in mass (12 mg/g), assuming that all of the
additional carbohydrate taken up was stored as glycogen. However, the
increase (0.7 ± 0.4 mg · kg
1 · min
1) in
tracer-determined glycogen synthetic rate attributed to fructose is
lower than predicted, suggesting that some of the glycogen was formed
via the indirect pathway. Although our data cannot differentiate
between a stimulation of glycogen synthase and/or an inhibition of
phosphorylase, they are consistent with in vitro evidence suggesting
that fructose stimulates glycogen synthase (6).
Acute fructose infusion had no effect on hepatic lipid metabolism. Tracer incorporation into lipid was insignificant, and neither arterial NEFA concentration nor hepatic NEFA uptake increased significantly during fructose infusion. In contrast, some studies have found adverse effects (increased triglyceride and cholesterol levels) of fructose in at-risk patients when larger amounts of fructose (e.g., 10% of calories) were consumed for several weeks (7). In the present study, however, a 3-h infusion of small amounts of fructose did not alter hepatic lipid metabolism.
Thus, in the TPN-adapted state, intraportal fructose infusion enhanced NHGU in sham and infected dogs. Although the ability of fructose to stimulate NHGU was not impaired by infection, fructose did not correct the impairment in hepatic lactate release. Glycogen was a major fate of the additional glucose utilized by the liver. Future studies can be done to determine whether the addition of small amounts of fructose to TPN will chronically improve liver glucose uptake and limit the hyperglycemia seen during stress.
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ACKNOWLEDGEMENTS |
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The authors thank Wanda Snead, Pamela Venson, and Eric Allen of the Vanderbilt Hormone Core Laboratory for hormone analysis and Mary C. Moore, Masakazu Shiota, and Patrick Fueger for critical reading of the manuscript.
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FOOTNOTES |
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This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-43748 (Principal Investigator, O. P. McGuinness), Diabetes Research and Training Center Grant P60-DK-20593, and Clinical Nutrition Research Unit Grant P30-DK-26657.
Address for reprint requests and other correspondence: O. P. McGuinness, 702 Light Hall, Dept. of Molecular Physiology and Biophysics, Vanderbilt Univ., Nashville, TN 37232-0615 (E-mail: owen.mcguinness{at}mcmail.vanderbilt.edu).
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.
Received 30 June 2000; accepted in final form 15 January 2001.
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