Hyperinsulinemia compensates for infection-induced impairment in net hepatic glucose uptake during TPN

Christine M. Donmoyer, Sheng-Song Chen, Scott A. Hande, D. Brooks Lacy, Joseph Ejiofor, and Owen P. McGuinness

Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, Tennessee 37232


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In animals receiving total parenteral nutrition (TPN), infection impairs net hepatic glucose uptake (NHGU) by 40% and induces mild hyperinsulinemia. In the normal animal, the majority of the glucose taken up by the liver is diverted to lactate, but in the infected state, lactate release is curtailed. Because of the hyperinsulinemia and reduced NHGU, more glucose is utilized by peripheral tissues. Our aims were to determine the role of infection-induced hyperinsulinemia in 1) limiting the fall in NHGU and hepatic lactate release and 2) increasing the proportion of glucose disposed of by peripheral tissues. Chronically catheterized dogs received TPN for 5 days via the inferior vena cava. On day 3, a fibrin clot with a nonlethal dose of E. coli was placed into the peritoneal cavity; sham dogs received a sterile clot. On day 5, somatostatin was infused to prevent endogenous pancreatic hormone secretion, and insulin and glucagon were replaced at rates matching incoming hormone concentrations observed previously in sham or infected dogs. The TPN-derived glucose infusion was adjusted to maintain a constant arterial plasma glucose level of ~120 mg/dl. after a basal blood sampling period, the insulin infusion rate was either maintained constant (infected time control, Hi-Ins, n = 6; sham time control, Sham, n = 6) or decreased (infected + reduced insulin, Lo-Ins; n = 6) for 180 min to levels seen in noninfected dogs (from 23 ± 2 to 12 ± 1 µU/ml). Reduction of insulin to noninfected levels decreased NHGU by 1.4 ± 0.5 mg · kg-1 · min-1 (P < 0.05) and nonhepatic glucose utilization by 4.8 ± 0.8 mg · kg-1 · min-1 (P < 0.01). The fall in NHGU was caused by a decline in HGU (Delta -0.6 ± 0.4 mg · kg-1 · min-1) and a concomitant increase in hepatic glucose production (HGP, Delta 0.8 ± 0.5 mg · kg-1 · min-1); net hepatic lactate release was not altered. Hyperinsulinemia that accompanies infection 1) primarily diverts glucose carbon to peripheral tissues, 2) limits the fall in NHGU by enhancing HGU and suppressing HGP, and 3) does not enhance hepatic lactate release, thus favoring hepatic glucose storage. Compensatory hyperinsulinemia plays a critical role in facilitating hepatic and peripheral glucose disposal during an infection.

total parenteral nutrition; liver; glucose; lactate; dog


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

TOTAL PARENTERAL NUTRITION (TPN) is an important mainstay in meeting the caloric requirements of individuals who are unable to ingest adequate quantities of nutrients. When individuals receive glucose-based TPN, the body provides mechanisms to dispose of the exogenous carbohydrate load efficiently without developing marked hyperglycemia and hyperinsulinemia. One of the sites important for nutrient disposal is the liver. In normal human subjects and animals receiving an acute oral glucose load, the liver removes ~40% of the glucose, storing most of it within the liver (1, 11, 25). Similarly, glucose uptake by the liver in normal dogs receiving chronic TPN (5 days) accounted for 45% of the exogenous glucose infused (21). However, rather than storing the glucose during the chronic infusion, the canine liver converts the majority of this glucose to lactate, which is subsequently released and disposed of by peripheral tissues. Peripheral lactate removal is more efficient than glucose removal and does not require additional insulin; thus conversion of glucose to lactate within the liver is effective in preventing marked hyperglycemia and hyperinsulinemia in normal animals.

Many patients receiving TPN chronically also experience an underlying stress (e.g., injury, trauma, infection). The metabolic response to infection with nutritional support is characterized by hyperglycemia and hyperinsulinemia. Hyperglycemia is attributed in part to a fall in hepatic glucose uptake (HGU) and/or an increase in glucose production. Several groups (18, 29) have observed an inability of exogenous glucose to completely suppress hepatic glucose production (HGP) in the presence of infection. Enhanced gluconeogenesis (GNG) can account for most of the defect in HGP suppression (20), and this impairment can be overcome by higher insulin levels (28). Along with the diminished ability to suppress glucose production, HGU is impaired (11, 23). In surgically stressed patients receiving chronic TPN, splanchnic (i.e., liver and gut) glucose uptake accounts for only 20% of the glucose infused (9). Infected dogs receiving TPN demonstrate markedly reduced net hepatic glucose uptake (NHGU) (21) as well as blunted hepatic lactate release. In contrast to what occurs in normal animals, during an infection, the predominant fate of the glucose taken up by the liver is storage in the form of glycogen and lipid, rather than release as lactate. This impaired glucose uptake and attenuated lactate diversion occurs in the presence of higher glucose and insulin levels. As a consequence of the lower NHGU, glucose uptake in peripheral tissues must increase to dispose of the TPN-derived glucose.

The specific role of insulin in enhancing glucose uptake and storage in the liver and muscle in the stressed TPN-adapted state is unclear. Skeletal muscle is exquisitely sensitive to insulin, so the compensatory hyperinsulinemia seen during an infection likely aids peripheral glucose uptake. Although skeletal muscle insulin resistance is well recognized with injury (14), assessment of the effectiveness of insulin in stimulating peripheral glucose uptake is complicated by the fact that infection increases whole body non-insulin-mediated glucose disposal as well (15, 16). Furthermore, the extent to which compensatory hyperinsulinemia during infection limits the fall in NHGU is unknown; the additional insulin may suppress HGP and/or stimulate HGU. In the acute setting, insulin is known to amplify hepatic glucose storage and not to facilitate glycolysis (6).

The aim of the present study was to determine the extent to which NHGU would be impaired if the infection-induced hyperinsulinemia were not present. We hypothesized that the primary roles of the compensatory hyperinsulinemia in the fed state were to enhance peripheral glucose uptake and to suppress HGP. To assess the role of hyperinsulinemia in glucose metabolism during infection with nutritional support, it is necessary to control other factors that are known to regulate glucose metabolism (e.g., hyperglycemia and hyperglucagonemia). Experiments were performed in a controlled hormonal and glycemic environment in which the higher insulin levels observed with infection were acutely reduced to noninfected levels. The impact of the restoration to normal insulinemia on hepatic and peripheral glucose utilization was measured with arteriovenous and tracer methods.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
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Animal preparation. Eighteen female nonpregnant mongrel dogs were fed standard Kal-Kan meat (Vernon, CA) and Purina Lab Canine Diet #5006 (Purina Mills, St. Louis, MO) 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 American Association for the Accreditation of Laboratory Animal Care International guidelines. The protocols were approved by the Vanderbilt University Medical Center Animal Care Committee. The health of the animals was determined before surgery and before TPN administration as having: a good appetite (i.e., consumed at least 3/4 of the daily ration), normal stools, hematocrit >0.35, and leukocyte count <18,000 mm-3.

Experimental overview. Chronically catheterized dogs received TPN continuously for 5 days, and on day 3 of TPN administration, a bacterial or a sham clot was implanted in the peritoneal cavity. The study was performed 42 h after induction of infection and consisted of three periods: tracer equilibration, basal sampling, and experimental sampling where the insulin infusion rate was continued or reduced.

Experimental preparation. A laparotomy was performed using sterile techniques with general anesthesia (15 mg/kg thiopental sodium iv for induction and 1.0% isoflurane as an inhalant during surgery) on healthy dogs. During the laparotomy, Silastic catheters (0.03 in. ID) were placed in the splenic and jejunal veins, and the gastroduodenal vein was ligated. Blood sampling catheters (0.04 in. ID) 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. After an incision in the left and right inguinal regions, a sampling catheter (0.04 in. ID) was placed in the left common iliac vein, and the tip was positioned distal to the anastomosis with the IVC; another catheter was advanced from the right external iliac artery to the abdominal aorta.

All catheters were filled with 0.9% NaCl (saline) containing heparin (200 U/ml). The free ends of the catheters and flow probes were exteriorized and placed in subcutaneous pockets. The dogs received penicillin G (500,000 U iv) in 1 liter of saline to minimize the possibility of infection. Flunixamine (0.1 mg/kg; Fort Dodge Laboratory, Fort Dodge, IA) was injected intramuscularly immediately after wound closure for acute pain relief. Dogs also received penicillin G (600,000 U im) for 3 days after surgery.

Nutritional support. After allowing >= 14 days for recovery from surgery, each dog's IVC catheters were exteriorized under local anesthesia (2% Lidocaine, Abbott, North Chicago, IL). TPN was infused into one or both of these catheters with 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 TPN bag and pump.

The dogs received TPN as the sole exogenous caloric source for 5 days. The TPN was designed to be isocaloric, based on predicted resting energy expenditure (31). The composition of the TPN included glucose, lipids, amino acids, saline (2.9 ml · kg-1 · min-1), potassium phosphates (90 mg · kg-1 · day-1), and a multivitamin supplement (MVI-12, Astra USA, Westborough, MA). Glucose (50% dextrose, Abbott) made up 75% of the nonprotein calories, and a fat emulsion (20% Intralipid, Baxter Healthcare, Deerfield, IL) constituted the remaining 25% of the energy requirements. Travasol (Baxter) was infused to supply basal nitrogen requirements (~12 g protein/day), calculated with the formula 1.5 × body wt0.67 (in kg).

Induction of infection. A fibrin clot containing a nonlethal dose (2 × 109 organisms/kg body wt) of Escherichia coli was prepared. The dose of bacteria was determined by serial dilution followed by plating. Bacteria (ATTC #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 and then washed with and reconstituted in sterile saline. A 1% fibrinogen (10 ml/kg body wt; Sigma, St. Louis, MO) solution was filtered (0.45 µm diameter) under sterile conditions. Bacteria were mixed with the filtrate, and thrombin (1,000 U, Gentrac, Middleton, WI) was added to initiate clot formation.

On day 3 of TPN administration, a second laparotomy was performed under anesthesia. An abdominal midline incision was made at a point caudal to that made during the first surgery; the clot was then implanted in the peritoneal cavity. Animals received 0.5 liter of saline during the laparotomy and 1 liter the day after. During the course of the infection, animals were monitored closely. The same procedure was followed for Sham (n = 6) dogs, except they received a sterile clot and 0.5 liter of saline on the subsequent day. Infected animals (n = 12) were typically normotensive, hyperthermic, and tachycardic, with a characteristic increase in hepatic arterial blood flow (HABF) (21).

Experimental protocol. A study was performed 42 h after clot implantation, which was day 5 of TPN administration. The free ends of all catheters were exteriorized under local anesthesia, and their contents were aspirated and flushed with saline. The free ends of the flow probes were also exteriorized and connected to a flow meter (Transonic Systems, Ithaca, NY). 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, dextrose, and somatostatin (SRIF, Bachem, Torrance, CA). Primed (44 and 27 µCi) constant infusions (0.4 and 0.3 µCi/min) of [3-3H]- and [U-14C]glucose (New England Nuclear, Wilmington, DE), respectively, were begun at least 120 min before sampling.

A pancreatic clamp was performed in which SRIF was infused (0.8 µg · kg-1 · min-1) to suppress endogenous insulin and glucagon secretion. Insulin (porcine regular Iletin II; Eli Lilly, Indianapolis, IN) and glucagon (Eli Lilly) were replaced intraportally by infusion into the splenic and jejunal veins at rates of 1,000 µU · kg-1 · min-1 and 2.5 ng · kg-1 · min-1, respectively, to simulate the levels seen during infection. For the sham group, SRIF (0.8 µg · kg-1 · min-1), insulin (400 µU · kg-1 · min-1), and glucagon (0.1 ng · kg-1 · min-1) were infused to match the levels observed previously in sham animals. All solutions were infused by means of calibrated syringe pumps (Harvard Apparatus, Holliston, MA). The glucose infusion rate (GIR) in the TPN was adjusted regularly to maintain isoglycemia at 120 mg/dl, whereas the other components of the TPN were infused at the constant rate for the duration of the experiment.

Body temperature (Yellow Springs Instruments, Yellow Springs, OH), blood pressure, and heart rate (Micro-Med, Louisville, KY) were assessed during the study. Small blood samples (0.4 ml) were taken every 5-10 min and centrifuged immediately to measure arterial plasma glucose concentration with a Beckman Glucose Analyzer II (Beckman Instruments, Fullerton, CA). Saline was infused to replace the blood volume withdrawn by sampling. Blood samples in the artery, portal vein, hepatic vein, and iliac vein were taken every 20 min during the 40-min basal period and every 30 min during the 180-min experimental period; at least 80 min elapsed between initiation of the pancreatic clamp and blood sampling. Blood flows and hematocrits were recorded at each sampling period.

After the basal sampling period, infected animals were divided into two groups: infected time control (Hi-Ins, n = 6) and infected + reduced insulin (Lo-Ins, n = 6); the latter group's insulin infusion rate was decreased to 400 µU · kg-1 · min-1 to achieve arterial plasma insulin levels representative of the sham group. At the end of the study, the animals were killed with an overdose of pentobarbital sodium (Veterinary Lab, Lenexa, KS).

Sample processing. Blood samples were placed in chilled tubes containing potassium EDTA (15 mg). The collection and immediate processing of blood samples have been described previously (20). Blood 14CO2 was assessed in triplicate on arterial, portal vein, and hepatic vein samples, as described by Chan and Dehaye (4). An equal volume of blood was mixed with 10% sulfosalicylic acid for gluconeogenic amino acid analysis. Blood samples were centrifuged at 3,000 rpm for 10 min. For the glucagon assay, 1 ml of plasma was added to 50 µl of Trasylol (500 kallikrein inhibitor units, Miles, Kankakee, IL). Plasma (0.5 ml) samples were deproteinized with Ba(OH)2 and ZnSO4 (30) and resined to remove charged intermediates (22) and to assess plasma [3H]- and [14C]glucose specific activity (SA). The remaining plasma was stored at -70°C for later analyses.

Analysis. Immunoreactive insulin and glucagon were assayed using a double antibody technique [intra-assay coefficient of variation (CV) 11% and 10%, respectively (26)], and cortisol was assayed with Diagnostic Products (Los Angeles, CA) RIA kit (CV 12%) (8). HPLC methods were used to assess plasma epinephrine and norepinephrine (CV 15% and 12%, respectively) (10) and blood gluconeogenic amino acid (glycine, serine, threonine) concentrations (3).

Analysis of gluconeogenic metabolites (lactate, alanine, and glycerol) in blood was performed on an automated centrifugal analyzer (Monarch 2000; Instrumentation Laboratory, Lexington, MA) using a modification of the method of Lloyd et al. (17). The concentration of nonesterified fatty acids (NEFA) was determined spectrophotometrically (Wako Chemicals, Richmond, VA). Glutamine content was assayed by the method of Bernt and Bergmeyer (2), adapted for the Technicon Autoanalyzer (Tarrytown, NY).

Calculations. The hepatic substrate load (Load In) was calculated as AS × HABF + PVS × PVBF, where AS and PVS represent the blood or plasma substrate concentrations in the iliac artery and portal vein, and HABF and PVBF represent blood flow in the hepatic artery and portal vein, respectively. Similarly, the substrate load leaving the liver (Load Out) was the product of HVS × HBF, in which HVS and HBF represent the hepatic vein substrate concentration and total hepatic blood or plasma flow (HABF + PVBF). Net hepatic substrate uptake was the difference between Load In and Load Out. Net hepatic substrate fractional extraction was calculated as the ratio of net hepatic substrate uptake and Load In.

These equations were used to calculate net hepatic glucose, lactate, alanine, glycerol, amino acid, and NEFA balances. Plasma glucose was converted to blood glucose by a correction factor of 0.73. Unidirectional HGU and hepatic glucose oxidation were calculated as the ratios of hepatic [3H]glucose uptake and hepatic 14CO2 production divided by the corresponding (3H or 14C) inflowing glucose specific activity, respectively. In cases where the liver was a producer of substrate (i.e., negative uptake), these data were presented as positive values and denoted as net output. Because the liver produced and consumed glucose simultaneously, hepatic glucose production (HGP) was the difference between unidirectional uptake and net uptake (NHGU = HGU - HGP). Similarly, net hindlimb glucose uptake was calculated with the formula (Ag - Vg) × ABF, where Ag and Vg represent glucose concentrations in the iliac artery and iliac vein and ABF represents blood flow in the iliac artery. Plasma flow was calculated by multiplying blood flow by (1 - hematocrit). Net nonhepatic glucose uptake was the difference between exogenous GIR and NHGU. The rate of whole body glucose appearance (Ra) was calculated with a two-compartmental model, as described by Mari et al. (19). Endogenous glucose production (EGP) was the difference between Ra and GIR.

Statistics. All mean and SE values reported for the basal period are the means of the 40-min period, using three time points. For the experimental period, the mean of 120-180 min (i.e., the final 60 min of the study) was used. Statistical comparisons were made with two-way ANOVA vs. change from basal for infected groups followed by an F test (SYSTAT, Evanston, IL), and one-way ANOVA when comparing Sham and infected groups. P < 0.05 was regarded as significant.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Hemodynamics. Basal blood flow (hepatic arterial, portal vein, and iliac arterial), mean arterial blood pressure, heart rate, body temperature, and noncontrolled hormone concentrations were measured in Sham, infected control (Hi-Ins), and infected + reduced insulin (Lo-Ins) groups (Table 1). Infected groups showed characteristic increases in body temperature, heart rate, and hepatic arterial blood flow.

                              
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Table 1.   Hemodynamic and hormone concentrations in the basal period of Sham, Hi-Ins, and Lo-Ins groups receiving TPN for 5 days

Hormone levels. Arterial plasma insulin levels were 10 ± 2 µU/ml (n = 6) in the Sham group. The basal insulin level was elevated in the infected groups (23 ± 1 µU/ml, n = 12; Fig. 1). The insulin levels in Lo-Ins (23 ± 2 µU/ml) were reduced to achieve levels seen in sham animals (12 ± 1 µU/ml). The glucagon concentration was higher in the infected groups (91 ± 5 pg/ml, n = 12) compared with Sham (22 ± 3 pg/ml, n = 6) and remained stable throughout the experimental period.


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Fig. 1.   Arterial plasma glucose (mg/dl), insulin (µU/ml), and glucagon (pg/ml) concentrations in Sham, infected control (Hi-Ins), and infected + reduced insulin (Lo-Ins) groups receiving total parenteral nutrition (TPN) for 5 days. SRIF, somatostatin. Data are expressed as means ± SE.

Whole body glucose metabolism. The arterial plasma glucose concentration (Fig. 1) was maintained at similar levels (~120 mg/dl) by varying the exogenous GIR, which is equivalent to whole body glucose utilization. GIR was 8.0 ± 1.0 and 11.1 ± 0.8 mg · kg-1 · min-1 in Sham and infected groups, respectively. In the Lo-Ins group, GIR decreased by 6.3 ± 0.7 mg · kg-1 · min-1 between the basal and experimental periods (P < 0.01); GIR in Hi-Ins did not change (Delta -0.2 ± 0.4 mg · kg-1 · min-1; Fig. 2). Whole body Ra was 9.3 ± 0.9 and 12.9 ± 0.9 mg · kg-1 · min-1 in Sham and infected groups, respectively. Upon lowering of insulin, Ra decreased by 5.0 ± 0.4 mg · kg-1 · min-1 (P < 0.01), whereas Ra in Hi-Ins did not change (Delta 0.2 ± 0.4 mg · kg-1 · min-1; Table 2). EGP increased by 1.2 ± 0.5 mg · kg-1 · min-1 (P < 0.05) when insulin was reduced; EGP did not change in Hi-Ins (Delta 0.4 ± 0.2 mg · kg-1 · min-1).


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Fig. 2.   Exogenous glucose infusion rate (basal, 11.1 ± 0.7 vs. 11.1 ± 0.9 mg · kg-1 · min-1, Hi-Ins vs. Lo-Ins, respectively), net nonhepatic glucose uptake rate (9.3 ± 0.9 vs. 8.3 ± 1.0 mg · kg-1 · min-1), and net hindlimb glucose uptake rate (13.8 ± 1.9 vs. 15.8 ± 2.9 mg/min), expressed as change from basal period, in infected animals receiving SRIF with glucagon replacement in which insulin levels were held constant (Hi-Ins) or lowered to levels seen in the absence of an infection (Lo-Ins). Data are expressed as means ± SE.


                              
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Table 2.   GIR, Ra, and EGP in Sham, Hi-Ins, and Lo-Ins groups during the basal period and last 60 min of the experimental period

Hepatic glucose metabolism. Basal NHGU in Sham was 3.4 ± 0.5 mg · kg-1 · min-1 (n = 6), whereas NHGU in the infected groups was lower (2.3 ± 0.3 mg · kg-1 · min-1, n = 12). Similarly, net hepatic fractional extraction (HFE) of glucose was lower in the infected groups (0.06 ± 0.01) compared with Sham (0.13 ± 0.01). Basal NHGU rates were not statistically different (P = 0.06) in the two infected groups. Both NHGU and HFE fell when insulin was lowered; NHGU decreased by 1.4 ± 0.5 mg · kg-1 · min-1 (P < 0.05), and HFE fell by 0.04 ± 0.01 (P < 0.05) (Fig. 3). In the Hi-Ins group, neither NHGU nor HFE fell (Delta 0.3 ± 0.5 mg · kg-1 · min-1 and Delta 0.01 ± 0.01).


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Fig. 3.   Net hepatic glucose uptake and net hepatic glucose fractional extraction, expressed as change from basal, in infected animals receiving SRIF with glucagon replacement in which insulin levels were held constant (Hi-Ins: basal 1.8 ± 0.5 mg · kg-1 · min-1, 0.05 ± 0.01) or lowered to levels seen in the absence of an infection (Lo-Ins: basal 2.8 ± 0.2 mg · kg-1 · min-1, 0.08 ± 0.01). Data are expressed as means ± SE.

Unidirectional HGU in the basal period was 4.0 ± 0.6 and 2.8 ± 0.4 mg · kg-1 · min-1 in the Sham (n = 6) and infected (n = 12) groups. HGU fell during the experimental period in both infected groups: Delta -0.6 ± 0.4 and Delta -0.5 ± 0.4 mg · kg-1 · min-1 (Lo-Ins and Hi-Ins), respectively. Basal HGP rates were similar in all groups (0.4 ± 0.3, 0.4 ± 0.3, and 0.6 ± 0.7 mg · kg-1 · min-1; Sham, Hi-Ins, and Lo-Ins, respectively). HGP tended to fall in Hi-Ins (Delta -0.7 ± 0.6 mg · kg-1 · min-1), but when insulin was lowered, HGP increased (Delta 0.8 ± 0.5 mg · kg-1 · min-1). Basal hepatic glucose oxidation rate was 0.4 ± 0.1 mg · kg-1 · min-1 in each group and did not change in the experimental period (data not shown).

Hindlimb glucose metabolism. Basal net nonhepatic glucose uptake was 8.8 ± 0.6 mg · kg-1 · min-1 (n = 12) in the infected groups and 4.5 ± 0.9 mg · kg-1 · min-1 in Sham. When the insulin level was lowered, nonhepatic glucose uptake decreased by 58 ± 6% to 3.5 ± 0.5 mg · kg-1 · min-1 (P < 0.01), whereas it did not change with time in Hi-Ins (Delta -0.5 ± 0.6 mg · kg-1 · min-1; Fig. 2). Consistent with nonhepatic glucose uptake, hindlimb glucose uptake fell from 15.8 ± 2.9 to 8.2 ± 3.0 mg/min upon reduction of insulin (Delta -7.6 ± 1.1 mg/min, P < 0.01; Hi-Ins, Delta 1.5 ± 1.6 mg/min). In comparison, basal limb glucose uptake in the Sham group was 12.0 ± 3.4 mg/min.

Metabolites. The basal arterial lactate concentration tended to be lower (720 ± 61 µM, n = 12) in the infected groups compared with Sham (930 ± 104 µM, P = 0.066), and levels in both infected groups rose in the experimental period (Lo-Ins, Delta 184 ± 64 µM, P < 0.05; Hi-Ins, Delta 82 ± 54 µM, not significant; Fig. 4). Net hepatic lactate release in the Sham group was higher than that in the infected groups (24 ± 2 vs. 10 ± 2 µmol · kg-1 · min-1, P = 0.001). Net hepatic lactate release did not increase in either infected group during the experimental period (Delta 0.1 ± 2.1 vs. Delta -0.5 ± 1.9 µmol · kg-1 · min-1, Lo-Ins vs. Hi-Ins, respectively).


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Fig. 4.   Arterial blood lactate concentrations and net hepatic lactate release, expressed as change from basal, in infected animals receiving SRIF with glucagon replacement in which insulin levels were held constant (Hi-Ins: basal 684 ± 84 µM, 7.8 ± 3.2 µmol · kg-1 · min-1) or lowered to levels seen in the absence of an infection (Lo-Ins: basal 755 ± 102 µM, 13.0 ± 3.2 µmol · kg-1 · min-1). Data are expressed as means ± SE.

The arterial alanine concentration was decreased with infection (224 ± 14 µM, n = 12, vs. 501 ± 69 µM in Sham, P < 0.001). However, net hepatic alanine uptake was elevated during infection (2.5 ± 0.3 vs. 1.5 ± 0.3 µmol · kg-1 · min-1, infected vs. Sham, respectively, P < 0.05) because of an increase in net HFE of alanine (0.24 ± 0.02 vs. 0.11 ± 0.03, P < 0.01). Circulating levels of all gluconeogenic amino acids except threonine were lower with infection (Table 3); however, net uptake rates of the gluconeogenic precursors were not altered when the insulin level was reduced.

                              
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Table 3.   Arterial plasma metabolite concentrations and metabolite uptake rates in Sham, Hi-Ins, and Lo-Ins groups during the basal period and last 60 minutes of the experimental period

Arterial glycerol concentrations were not altered by infection; however, net hepatic glycerol uptake was increased (Table 3). The acute lowering of insulin altered neither glycerol concentration nor net hepatic glycerol uptake. Arterial plasma NEFA concentrations in the basal period were 234 ± 11 µM (n = 12) with infection and 291 ± 32 µM in the Sham group (Sham vs. infected, P < 0.05). Net hepatic NEFA uptake rates were similar (0.9 ± 0.1 vs. 0.6 ± 0.2 µmol · kg-1 · min-1, infected vs. Sham). When insulin was reduced, NEFA levels increased by 107 ± 29 µM (P < 0.05), but hepatic NEFA uptake did not increase (Delta 0.4 ± 0.3 µmol · kg-1 · min-1; Fig. 5). In Hi-Ins, NEFA levels and net hepatic NEFA uptake did not change (Delta -14 ± 10 µM and Delta -0.1 ± 0.2 µmol · kg-1 · min-1).


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Fig. 5.   Nonesterified fatty acid (NEFA) concentrations and net hepatic NEFA uptake, expressed as change from basal, in infected animals receiving SRIF with glucagon replacement in which insulin levels were held constant (Hi-Ins: basal 232 ± 17 µM, 0.7 ± 0.2 µmol · kg-1 · min-1) or lowered to levels seen in the absence of an infection (Lo-Ins: basal 236 ± 17 µM, 1.1 ± 0.2 µmol · kg-1 · min-1). Data are expressed as means ± SE.


    DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Previous work in dogs adapted to TPN demonstrated that the presence of an infection led to a 40% fall in NHGU and a corresponding increase in nonhepatic glucose uptake with a concomitant doubling of insulin concentration (21). In the present studies, when the insulin concentration in infected dogs was reduced to levels observed in sham dogs, glucose uptake rates in both liver and hindlimb (i.e., skeletal muscle) fell by ~50%. Thus the infection-induced increase in insulin facilitates liver and skeletal muscle glucose disposal. The decline in NHGU was caused by a combined increase in HGP and decrease in unidirectional HGU. Because the infection-induced rise in insulin enhanced NHGU but did not increase hepatic lactate release, it preferentially favors carbon storage within the liver.

The liver is responsible for limiting hyperglycemia in response to nutritional support during an infection by increasing HGU and decreasing HGP. Infection impairs the increase in NHGU observed in response to an acute glucose infusion (23), and TPN-adapted animals show a similar pattern of impaired NHGU (21). The mechanism of the impairment in NHGU with infection is unclear, but endocrine changes such as hyperglucagonemia (7, 13) or elevated cortisol and catecholamines, hepatic insulin resistance, and/or enhanced cytokine synthesis (e.g., interleukin-1 and -6, tumor necrosis factor-alpha ) may be involved. Basal NHGU tended to be lower in Hi-Ins than in Lo-Ins, which may indicate greater stress, but both infected groups had lower NHGU than Sham did. Some basal parameters were not equivalent in the two infected groups (heart rate and epinephrine); however, all other counterregulatory hormone (glucagon, norepinephrine, cortisol) concentrations and hemodynamic variables were similar. When insulin was reduced, NHGU declined in every animal (Lo-Ins), and the magnitude of the fall was independent of the basal NHGU; NHGU in Hi-Ins animals did not change over time. The decrease in unidirectional HGU contributed 42% to the fall in NHGU; however, this could be an overestimate. Tracer accumulated in hepatic glycogen during the equilibration and basal periods may have been released upon reduction of the insulin infusion rate. Thus the absolute HGU during the last hour may have been an underestimate, leading to an overestimate of the fall in HGU.

The mechanism for the increase in HGP when insulin was reduced was an increase in the rate of glycogenolysis. The rise in HGP was confirmed by a parallel increase in tracer-determined EGP. The elevated insulin level observed during infection, therefore, attenuates the rise in HGP. Endogenous sources of glucose carbon include glycogenolysis and gluconeogenesis (GNG), and the latter process is frequently enhanced during infection (20). Because net hepatic gluconeogenic precursor uptake (of alanine, glycerol, serine, glycine, threonine, and glutamine) did not increase (Delta -0.2 ± 0.1 mg · kg-1 · min-1 of glucose equivalents) when insulin was reduced, the increase in HGP was caused by an increase in glycogenolysis. Insulin at moderate doses has a very potent inhibitory effect on glycogenolysis (5). It is possible that an increase in GNG would have been observed as well if the lower insulin level had been maintained beyond 3 h. In addition, intrahepatic proteolysis may have increased with the change in insulin but could not be detected by the arteriovenous difference method.

In normal TPN-adapted animals, the conversion of glucose to lactate in the liver is effective in preventing marked hepatic glucose storage, but hepatic lactate release is blunted in the presence of an infection. Although the baseline for hepatic lactate release was higher for Lo-Ins relative to Hi-Ins (not statistically different), reflecting the higher basal NHGU in Lo-Ins, the proportion of NHGU released as lactate was similar in the infected groups (32 ± 3% and 23 ± 7%, Lo-Ins and Hi-Ins, respectively). Because NHGU fell when insulin was reduced, hepatic lactate release would be expected to decrease. This fall was not observed, however, presumably because glycogenolytically derived glucose carbon was shunted to lactate. Thus, although the infection-induced increase in insulin limits the fall in NHGU and facilitates glucose entry, it does not enhance hepatic lactate release. Instead, the hormone minimizes glucose production by restraining glycogen breakdown, which sustains glucose storage within the liver. Although glycogen formation with chronic nutritional support is favored by the accompanying hyperinsulinemia (1), lipid synthesis may become a major storage fate (32) when liver glycogen capacity is saturated.

In our studies, the proportion of the TPN-derived glucose taken up by nonhepatic tissues was enhanced with infection relative to the Sham group. It is difficult to assess insulin-mediated nonhepatic glucose uptake between sham and infected groups (8.8 ± 0.6 vs. 4.5 ± 0.9 mg · kg-1 · min-1, infected vs. Sham, respectively), because insulin levels were different and insulin-independent glucose uptake is increased with infection (24). However, when the insulin concentration was reduced to sham insulin levels, nonhepatic glucose uptake in infected dogs decreased to 3.5 ± 0.5 mg · kg-1 · min-1, which was not significantly lower than that in the Sham group. Because insulin-independent glucose uptake (one component of nonhepatic glucose uptake) is enhanced during infection, it is likely that insulin-mediated glucose uptake (i.e., skeletal muscle) is decreased. A 48% fall in hindlimb glucose uptake was observed when the insulin level was reduced. The impairment was likely amplified by a rise in NEFA levels, which inhibit insulin-stimulated glucose uptake (27). At comparable insulin levels, hindlimb glucose uptake in the infected group (Lo-Ins 8.2 ± 3.0 mg/min) was 40% less than the Sham group (14.3 ± 4.3 mg/min), although this was not statistically different. This result is consistent with insulin resistance in the periphery and is in agreement with a previous finding (12) that insulin-mediated glucose uptake in peripheral tissues is impaired in the presence of an infection.

The presence of an infection serves to shunt glucose away from the liver toward peripheral tissues. Although the rise in insulin observed during infection maintains glucose disposal in insulin-sensitive tissues such as the liver and skeletal muscle, it may also contribute to the shift of glucose from the liver. Insulin did not alter the proportion of the glucose load taken up by hepatic and nonhepatic tissues. At the high insulin level (basal period), the liver disposed of 26 ± 3% of the whole body glucose utilization (WBGU or GIR), whereas nonhepatic tissues disposed of 74 ± 3%. By lowering the insulin level, WBGU was reduced by 6.3 ± 0.7 mg · kg-1 · min-1, but the proportion of glucose taken up by the liver and periphery remained the same (25 ± 9% and 75 ± 9%, respectively). Thus the infection-induced hyperinsulinemia maintains, but does not amplify, the infection-induced diversion of glucose carbon to nonhepatic tissues. However, in terms of the mass of glucose, the infection-induced elevation in insulin supported glucose uptake by the peripheral tissues more than it improved HGU.

During TPN administration, infection impairs WBGU, which is compensated for by the ensuing hyperinsulinemia. The rise in insulin facilitates both hepatic and peripheral glucose disposal, and if this had not occurred, greater hyperglycemia would have developed. Within the liver, moderate hyperinsulinemia serves a dual role by suppressing HGP and enhancing HGU. Thus, in a catabolic environment, hyperinsulinemia performs an important function; it preserves the body's ability to consume glucose. Compensatory hyperinsulinemia diverts the majority of glucose carbon to peripheral tissues but does not attenuate or correct the underlying impairments in hepatic glucose metabolism.


    ACKNOWLEDGEMENTS

The authors are grateful for the technical assistance of Pamela Y. Venson and Eric J. Allen for hormone analysis, Ying Yang for amino acid analysis, and Mary C. Moore, Amy E. Halseth and David T. Duong for critical reading of the manuscript.


    FOOTNOTES

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. §1734 solely to indicate this fact.

Received 25 October 1999; accepted in final form 24 February 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Askanazi, J, Carpentier YA, Elwyn DH, Nordenstrom J, Jeevanandam M, Rosenbaum SH, Gump FE, and Kinney JM. Influence of total parenteral nutrition on fuel utilization in injury and sepsis. Ann Surg 191: 40-46, 1980[ISI][Medline].

2.   Bernt, E, and Bergmeyer HU. L-Glutamate: determination with glutamic dehydrogenase. In: Methods of Enzymatic Analysis, edited by Bergmeyer HU. Weinheim: Verlag Chemie, 1963, p. 384-388.

3.   Bidlingmeyer, BA, Cohen SA, and Tarvin TL. Rapid analysis of amino acids using precolumn derivatization. J Chromatogr 336: 93-104, 1984[Medline].

4.   Chan, TM, and Dehaye JP. Hormone regulation of glucose metabolism in the genetically obese-diabetic mouse (db/db). Diabetes 30: 211-218, 1981[ISI][Medline].

5.   Cherrington, AD. The acute regulation of hepatic glucose production. In: The Role of the Liver in Maintaining Glucose Homeostasis, edited by Pagliassotti MJ, Davis SN, and Cherrington AD. Austin, TX: RG Landes, 1994, p. 19-43.

6.   Davis, SN, McGuinness OP, and Cherrington AD. Insulin action in vivo. In: Diabetes Annual IV, edited by Alberti KGMM, and Krall LP. Amsterdam: Elsevier Science, 1990, p. 585-602.

7.   Del Prato, S, Castellino P, Simonson D, and DeFronzo R. Hyperglucagonemia and insulin-mediated glucose metabolism. J Clin Invest 79: 547-556, 1987[ISI][Medline].

8.   Farmer, RW, and Pierce CE. Plasma cortisol determination: radioimmunoassay and competitive protein binding compared. Clin Chem 20: 411-414, 1974[Abstract/Free Full Text].

9.   Gil, KM, Gump FE, Starker PM, Askanazi J, Elwyn DH, and Kinney JM. Splanchnic substrate balance in malnourished patients during parenteral nutrition. Am J Physiol Endocrinol Metab 248: E409-E419, 1985[Abstract/Free Full Text].

10.   Goldstein, DS, Feuerstein G, Izzo JL, Jr, Kopin IJ, and Keiser HR. Validity and reliability of liquid chromatography with electrochemical detection for measuring plasma levels of norepinephrine and epinephrine in man. Life Sci 28: 467-475, 1981[ISI][Medline].

11.   Gump, FE, Long C, Killian P, and Kinney JM. Studies of glucose intolerance in septic injured patients. J Trauma 14: 378-388, 1974[ISI][Medline].

12.   Lang, CH. Sepsis-induced insulin resistance in rats is mediated by a beta -adrenergic mechanism. Am J Physiol Endocrinol Metab 263: E703-E711, 1992[Abstract/Free Full Text].

13.   Lang, CH, Bagby GJ, Blakesley HL, and Spitzer JJ. Importance of hyperglucagonemia in eliciting the sepsis-induced increase in glucose production. Circ Shock 29: 181-191, 1989[ISI][Medline].

14.   Lang, CH, and Dobrescu C. In vivo insulin resistance during nonlethal hypermetabolic sepsis. Circ Shock 28: 165-178, 1989[ISI][Medline].

15.   Lang, CH, and Dobrescu C. Gram-negative infection increases non-insulin-mediated glucose disposal. Endocrinology 128: 645-653, 1991[Abstract].

16.   Lang, CH, Obih JC, Bagby GJ, Bagwell JN, and Spitzer JJ. Increased glucose uptake by intestinal mucosa and muscularis in hypermetabolic sepsis. Am J Physiol Gastrointest Liver Physiol 261: G287-G294, 1991[Abstract/Free Full Text].

17.   Lloyd, B, Burrin J, Smythe P, and Alberti KGMM Enzymatic fluorometric continuous flow assays for blood glucose, lactate, pyruvate, alanine, glycerol and 3-hydroxybutyrate. Clin Chem 24: 1724-1729, 1978[Abstract/Free Full Text].

18.   Long, CL, Kinney JM, and Geiger JW. Nonsuppressability of gluconeogenesis by glucose in septic patients. Metabolism 25: 193-201, 1976[ISI][Medline].

19.   Mari, A, Cobelli C, Cherrington AD, and McGuinness OP. A model for the study of glucose kinetics in nonsteady state. In: Modelling and Control in Biomedical Systems, edited by Cobelli C, and Mariani L. Frankfurt: Pergamon, 1988, p. 357-362.

20.   McGuinness, OP. The impact of infection on gluconeogenesis in the conscious dog. Shock 2: 336-343, 1994[ISI][Medline].

21.   McGuinness, OP, Donmoyer CM, Ejiofor J, McElligott S, and Lacy DB. Hepatic and muscle glucose metabolism during total parenteral nutrition: impact of infection. Am J Physiol Endocrinol Metab 275: E763-E769, 1998[Abstract/Free Full Text].

22.   McGuinness, OP, Fujiwara T, Murrell S, Bracy D, Neal D, O'Connor D, and Cherrington AD. Impact of chronic stress hormone infusion on hepatic carbohydrate metabolism in the conscious dog. Am J Physiol Endocrinol Metab 265: E314-E322, 1993[Abstract/Free Full Text].

23.   McGuinness, OP, Jacobs J, Moran C, and Lacy DB. Impact of infection on hepatic disposal of a peripheral glucose infusion in the conscious dog. Am J Physiol Endocrinol Metab 269: E199-E207, 1995[Abstract/Free Full Text].

24.   Mészáros, K, Lang CH, Bagby GJ, and Spitzer JJ. In vivo glucose utilization by individual tissues during nonlethal hypermetabolic sepsis. FASEB J 2: 3083-3086, 1988[Abstract/Free Full Text].

25.   Moore, MC, Cherrington AD, Cline G, Pagliassotti MJ, Jones EM, Neal DW, Badet C, and Shulman GI. Sources of carbon for hepatic glycogen synthesis in the conscious dog. J Clin Invest 88: 578-587, 1991[ISI][Medline].

26.   Morgan, CR, and Lazarow AL. Immunoassay of insulin: two antibody system. Plasma insulin of normal, subdiabetic, and diabetic rats. Am J Med 257: 415-419, 1963.

27.   Randle, PJ, Priestman DA, Mistry SC, and Halsall A. Glucose fatty acid interactions and the regulation of glucose disposal. J Cell Biochem 55: Suppl: 1-11, 1994[ISI][Medline].

28.   Shaw, JHF, Klein S, and Wolfe RR. Assessment of alanine, urea and glucose interrelationships in normal subjects and in patients with sepsis with stable isotopes. Surgery 97: 557-567, 1985[ISI][Medline].

29.   Shaw, JH, and Wolfe RR. Response to glucose and lipid infusions in sepsis: a kinetic analysis. Metabolism 34: 442-449, 1985[ISI][Medline].

30.   Somogyi, M. Determination of blood sugar. J Biol Chem 160: 69-73, 1945[Free Full Text].

31.   Subcommittee on Dog Nutrition. Nutrient Requirements for Dogs. Washington, DC: National Academy, 1985, p. 2-50.

32.   Wolfe, RR, O'Donnell TF, Stone MD, Richmand DA, and Burke JF. Investigation of factors determining the optimal glucose infusion rate in total parenteral nutrition. Metabolism 29: 892-900, 1980[ISI][Medline].


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