Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, Tennessee 37232
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ABSTRACT |
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Total parenteral nutrition
(TPN) markedly augments net hepatic glucose uptake (NHGU) and
hepatic glycolysis in the presence of mild hyperglycemia and
hyperinsulinemia. This increase is impaired by an infection. We
determined whether the adaptation to TPN alters the responsiveness of
the liver to insulin and whether infection impairs that response.
Chronically catheterized dogs received TPN for 5 days. On day
3 of TPN, either a nonlethal hypermetabolic infection was induced
(INF, n = 5) or a sham surgery was performed (SHAM,
n = 5). Forty-two hours after clot implantation,
somatostatin and glucagon (34 ± 3 vs. 84 ± 11 pg/ml in
artery, SHAM vs. INF) were infused, and a three-step (120 min each)
isoglycemic (~120 mg/dl) hyperinsulinemic (~12, 25, and 50 µU/ml)
clamp was performed to simulate levels seen in normal, infected, and
exogenous insulin treatment states. In SHAM, NHGU (3.5 ± 0.2 to
4.2 ± 0.4 to 4.6 ± 0.5 mg · kg1 · min
1)
modestly increased. In INF, NHGU was consistently lower at each insulin
step (1.1 ± 0.5 to 2.6 ± 0.5 to 2.8 ± 0.7 mg · kg
1 · min
1).
Although NHGU increased from the first to the second step in INF, it
did not increase further with the highest dose of insulin. Despite
increases in NHGU, net hepatic lactate release did not increase in SHAM
and fell in INF. In summary, in the TPN-adapted state, liver glucose
uptake is unresponsive to increases in insulin above the basal level.
Although the infection-induced increase in insulin sustains NHGU,
further increments in insulin enhance neither NHGU nor glycolysis.
liver; insulin; dog; glycogen; nutritional support
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INTRODUCTION |
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THE LIVER IS AN IMPORTANT SITE of glucose disposal, especially in response to chronic nutritional support (15). When total parenteral nutrition (TPN) is administered chronically, the liver consumes ~45% of the infused glucose. Interestingly, this high rate of net hepatic glucose uptake (NHGU) is achieved with only mild hyperglycemia and hyperinsulinemia. Moreover, the majority of the NHGU is released as lactate. The enhanced ability of the liver to take up glucose and convert it to lactate allows the body to dispose of substantial quantities of glucose without marked hyperglycemia or hyperinsulinemia.
It is unknown whether the TPN-adapted liver is as responsive and
sensitive to changes in insulin as the non-TPN-adapted liver. In the
overnight-fasted setting, the liver is very responsive to insulin. In
the presence of hyperglycemia (~200 mg/dl) a sixfold increase in
insulin increased liver glucose uptake sixfold (20). It is
also sensitive to insulin; hepatic insulin sensitivity
(ED50) in a 42-h-fasted dog is 90 µU/ml (sinusoidal
insulin levels). However, given the upregulation of liver glucose
uptake by TPN, the liver's response to insulin may be altered.
Although insulin levels are relatively low in this adapted setting, the
liver's sensitivity to insulin may also have increased
(ED50). If the mild hyperinsulinemia seen during TPN
produces a near-maximal response to insulin, the liver may not be
responsive to further increases in insulin.
An infection impairs the liver's adaptation. Both NHGU and net hepatic lactate release (NHLR) are decreased with infection (14). Consequently, peripheral tissues must directly metabolize a larger fraction of the exogenous glucose. However, the underlying peripheral insulin resistance characteristic of infection exacerbates the problem. As a consequence, a combination of marked hyperinsulinemia and hyperglycemia is required to facilitate glucose disposal. The mechanism for the impairment in NHGU with infection is unknown.
Given the hyperinsulinemia that accompanies infection, hepatic insulin resistance may be a contributing factor to the impairment in liver glucose uptake. Insulin resistance in peripheral tissues is widely recognized during infection (8, 10, 18), but the extent of involvement of the liver is controversial. Although hepatic glucose production (HGP) is elevated, it is not resolved whether or not infection impairs the ability of insulin to suppress HGP (9, 16). Liver glucose uptake is clearly impaired by infection; whether this is due to an impairment in insulin action has not been examined. Moreover, tight metabolic control with aggressive use of insulin profoundly improves the clinical outcome (25). Yet it is unclear whether the aggressive use of insulin corrects the impairment in NHGU or glycolysis, or diverts the glucose carbon to peripheral tissues.
The aims of the present study were to determine 1) whether NHGU in the TPN-adapted setting is responsive to increases in insulin, 2) whether the presence of an infection alters this response, and 3) whether additional insulin can reverse the infection-induced derangements in hepatic metabolism. Therefore, we established an insulin dose-response curve to evaluate the response of NHGU to changes in insulin in sham and infected animals receiving chronic TPN. Tracer and arteriovenous difference methods were used to assess unidirectional liver glucose uptake and production, NHLR, and net hindlimb substrate balance.
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METHODS |
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Animal preparation.
Ten female mongrel dogs (18-24 kg) were fed a standard meat
(Pedigree; Kalkan, Vernon, CA) and chow (Purina Lab Canine no. 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. Good health of the animals was determined before surgery and
before TPN administration as having a good appetite, i.e., consumed at
least three-quarters of the daily ration, normal stools, hematocrit
>0.35, and leukocyte count <18,000 mm3.
Experimental preparation. As described previously (3), a laparotomy was performed under general anesthesia. Infusion catheters (Dow Corning, Midland, MI) were placed in the splenic and jejunal veins for intraportal hormone infusion and into the inferior vena cava (IVC) for TPN infusion. Sampling catheters were positioned in the portal vein, left common hepatic vein, left common iliac vein, and right external iliac artery. Flow probes (Transonic Systems, Ithaca, NY) were positioned about the portal vein, hepatic artery, and right external iliac artery.
All catheters were filled with 0.9% NaCl (saline) containing heparin (200 U/ml). The free ends of the catheters and flow probes were placed in subcutaneous pockets. Dogs received penicillin G (600,000 U) in saline intravenously during surgery. In addition, they received 1,000 mg of ampicillin (Principen; Bristol-Myers Squibb, Princeton, NJ) orally per day for 3 days. Flunixamine (1 mg/kg body wt; Fort Dodge Laboratory, Fort Dodge, IA) was injected intramuscularly immediately after wound closure for acute pain relief.Nutritional support.
After 14 days were allowed for recovery from surgery, the IVC
catheter was exteriorized under local anesthesia (2% Lidocaine; Abbott, North Chicago, IL). TPN was infused into one catheter continuously for 5 days using an ambulatory infusion pump (Dakmed, Buffalo, NY). The dog wore a jacket (Alice King Chatham, Los Angeles, CA) with two large pockets for the nutrition and pump. The TPN was
designed to be isocaloric on the basis of predicted resting energy
expenditure (kcal/day), calculated with the equation 144 + 62.2 × body weight (24). The TPN included glucose
[75% of the nonprotein calories (NPC)], Intralipid (25% of NPC),
Travasol, saline, potassium phosphate, and a multivitamin supplement,
as described previously (14).
Induction of infection. On the 3rd day of TPN administration, a second laparotomy was performed under anesthesia. An abdominal midline incision was made at a point below that made during the previous surgery. Dogs were randomly assigned to two groups undergoing implantation of either a sterile (SHAM, n = 5) or bacterial (INF, n = 5) fibrin clot in the peritoneal cavity. TPN was continued, and animals received saline during the laparotomy (500 ml) as well as on the next day.
The fibrin clot was prepared by filtering (0.45 µm) a 1% fibrinogen (Sigma, St. Louis, MO) solution under sterile conditions. The bacterial clot 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, washed, and reconstituted in sterile saline before addition to the filtrate. To initiate clot formation, thrombin (1,000 U; Gentrac, Middleton, WI) was added to the filtrate.Experimental protocol. A study was performed on the 5th day of TPN and 42 h after clot implantation. 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 an Ultrasonic 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 and heart rate (Micro-Med, Louisville, KY) and rectal temperature (Yellow Springs Instruments, Yellow Springs, OH) were assessed during the study.
The chronic TPN solution was replaced with a TPN solution not containing glucose. The glucose was infused using a separate pump, allowing the infusion rate of glucose to be adjusted to maintain isoglycemia while not altering the infusion rate of the other TPN components. AtSample processing and analysis. Blood samples were collected and processed on the day of the study as previously described (12). Whole blood perchloric acid extracts were analyzed for lactate, alanine, and glycerol on an automated centrifugal analyzer (Monarch 2000; Instrumentation Laboratory, Lexington, MA) (11). Whole blood amino acids were assayed using HPLC techniques (26). Blood glutamine content was measured with a Technicon Autoanalyzer II (Bran Luebbe, Buffalo Grove, IL), adapted from Bernt and Bergman (1). Blood 14CO2 and tracer incorporation into [14C]lactate, [14C]glucose, and 14C-labeled amino acids were assessed as described (21, 22). Plasma was analyzed for measurements of glucose SA (3), insulin (19), cortisol (6), glucagon (19), and nonesterified fatty acid (NEFA; Wako Chemicals, Richmond, VA) concentrations. Plasma collected from whole blood treated with EGTA and glutathione was analyzed for epinephrine and norepinephrine with HPLC techniques [coefficients of variation 11 and 6%, respectively] (7). Hepatic glycogen content was determined using the enzymatic method of Chan and Exton (2).
Calculations.
The substrate (glucose, lactate, amino acids, glycerol, and NEFA) and
hormone (insulin) 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 substrate concentrations in the artery and
portal vein and HABF and PBF represent blood flows in the hepatic
artery and portal vein. Similarly, the substrate load leaving the liver
equaled Hs × THBF, in which Hs and THBF
represent the hepatic vein substrate concentration and total hepatic
blood flow (HABF + PBF). The net hepatic substrate uptake rate was
calculated as the difference between the entering and exiting substrate
loads. In cases where the liver was a net substrate producer, output
was denoted as a positive value. Similarly, net hindlimb and gut
substrate uptake rates were calculated using the equations
(As Vs) × ABF and
(As
Ps) × PBF, where
Vs is the concentration in the iliac vein and ABF is the
iliac arterial blood flow. Net hepatic fractional extraction (HFE) of
substrate was the net hepatic substrate uptake divided by the substrate load entering the liver; a similar equation was used for the hindlimb. Plasma flow instead of blood flow was used for NEFA calculations by
multiplying blood flow by (1
hematocrit ratio).
Statistics. For the majority of data, results are expressed as means ± SE of three sampling points in the Lo-Ins, Mid-Ins, and Hi-Ins infusion periods. There were five animals in both the SHAM and the INF groups unless otherwise indicated. Student's t-test was used to compare SHAM and INF animals. Statistical comparisons between periods were made with two-way ANOVA and an F-test (SYSTAT, Evanston, IL). A P value <0.05 was considered statistically significant.
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RESULTS |
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Baseline variables.
Body temperature and heart rate were elevated in INF, but mean arterial
pressure, body weight, and liver weight were comparable to those of
SHAM (Table 1). Plasma glucose levels
were 108 ± 6 vs. 118 ± 5 mg/dl before initiation of the
somatostatin infusion. Hepatic arterial blood flows in Lo-Ins, Mid-Ins,
and Hi-Ins periods (SHAM: 8 ± 1, 9 ± 1, and 9 ± 2 ml · kg1 · min
1,
respectively) were significantly greater with infection (INF: 21 ± 4, 21 ± 3, and 23 ± 4 ml · kg
1 · min
1,
respectively). Portal venous blood flows were comparable and were not
altered with insulin infusion (18 ± 2, 18 ± 2, and 19 ± 2 vs. 18 ± 4, 19 ± 4, and 18 ± 4 ml · kg
1 · min
1
in SHAM vs. INF, respectively).
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Hormones.
Infusion of insulin achieved comparable circulating insulin
concentrations in SHAM and INF at each insulin dose (Fig.
1). Arterial plasma insulin
concentrations in the Lo-Ins, Mid-Ins, and Hi-Ins periods were 10 ± 1, 24 ± 2, and 54 ± 4 µU/ml in SHAM and 11 ± 2, 27 ± 4, and 55 ± 6 µU/ml in INF, respectively. Sinusoidal insulin concentrations (35 ± 5, 83 ± 6, and 163 ± 14 µU/ml vs. 29 ± 3, 66 ± 6, and 130 ± 7 µU/ml,
respectively) were higher in SHAM (P < 0.05 at
Mid-Ins) due to lower THBF. The arterial plasma glucagon concentration
was significantly elevated in INF (34 ± 3 vs. 84 ± 11 pg/ml), which was similar to previously reported values
(14). Infection did not alter cortisol, epinephrine, or
norepinephrine concentrations (Table 1), and these hormones did not
change with insulin infusion in either group (data not shown).
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Hepatic glucose metabolism.
The exogenous glucose infusion rate (GIR; Table
2) was varied to maintain the arterial
plasma glucose concentration near 120 mg/dl (Fig.
2). As expected, GIR increased as the
insulin infusion rate was increased, and infection impaired this
increase.
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Glucose disposition.
Arterial blood lactate concentrations were similar in the Lo-Ins period
(978 ± 92 vs. 813 ± 210 µM, SHAM vs. INF) and did not
change with higher insulin doses (Fig.
4). NHLR rates were also comparable in
the basal period and did not increase as insulin was raised in SHAM
(1.6 ± 0.2 to 1.4 ± 0.1 to 1.2 ± 0.2 mg · kg1 · min
1)
and in INF (1.3 ± 0.4 to 1.0 ± 0.3 to 0.6 ± 0.5 mg · kg
1 · min
1).
Thus the proportion of NHGU released as lactate declined as insulin was
increased (46 ± 5, 33 ± 1, and 28 ± 4% in SHAM and 119 ± 25, 37 ± 7, and 14 ± 14% in INF). The
proportion of hepatic [14C]glucose uptake released as
[14C]lactate in SHAM was similar (51 ± 9%) to the
percentage of NHGU released as lactate in the Lo-Ins period
(n = 3); data for the other insulin periods, however,
were not available. The proportion of hepatic
[14C]glucose uptake released as
[14C]lactate in INF decreased as the insulin
concentration was elevated (70 ± 7, 30 ± 19, and 20 ± 24%; n = 4). Thus increases in insulin did not enhance
lactate release even though they enhanced NHGU.
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Metabolites.
Arterial plasma NEFA concentrations were similar in SHAM and INF at
each insulin dose and declined as insulin was increased (369 ± 64 to 225 ± 37 to 184 ± 30 µM in SHAM and 316 ± 48 to
210 ± 30 to 172 ± 23 µM in INF). Net hepatic NEFA uptake
rates were also comparable and fell in response to increases in insulin
(0.6 ± 0.3 to 0.3 ± 0.4 to 0.1 ± 0.2 µmol · kg
1 · min
1
and 0.9 ± 0.2 to 0.5 ± 0.2 to 0.1 ± 0.1 µmol · kg
1 · min
1).
Neither arterial plasma glycerol concentrations (54 ± 10, 53 ± 8, and 49 ± 6 µM vs. 63 ± 7, 59 ± 8, and 56 ± 10 µM, SHAM vs. INF) nor net hepatic glycerol uptake rates
(1.0 ± 0.2, 0.9 ± 0.2, and 0.9 ± 0.2 µmol · kg
1 · min
1
vs. 1.4 ± 0.2, 1.5 ± 0.1, and 1.4 ± 0.2 µmol · kg
1 · min
1)
changed significantly with increasing insulin concentrations.
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Gut substrate balance.
Glucose uptake by the gut was 0.8 ± 0.1, 0.6 ± 0.1, and
0.9 ± 0.2 mg · kg1 · min
1
in SHAM and 0.8 ± 0.2, 0.9 ± 0.2, and 1.2 ± 0.3 mg · kg
1 · min
1
in INF. Gut glucose oxidation (0.2 ± 0.1 vs. 0.2 ± 0.1 mg · kg
1 · min
1,
SHAM vs. INF, Lo-Ins) did not change with insulin (data not shown). The
gut was a net consumer of glutamine (1.6 ± 0.6, 1.5 ± 0.5, and 1.1 ± 0.3 µmol · kg
1 · min
1
vs. 1.4 ± 0.5, 1.0 ± 0.4, and 0.7 ± 0.2 µmol · kg
1 · min
1,
SHAM vs. INF).
Hindlimb substrate balance.
As expected in response to elevations in insulin, net hindlimb glucose
uptake (LGU) increased substantially in both groups, from 11 ± 4 to 21 ± 3 to 39 ± 7 mg/min in SHAM and from 6 ± 1 to
15 ± 4 to 27 ± 5 mg/min in INF (Fig.
5). LGU was not significantly different
between groups at comparable insulin doses. Net hindlimb fractional
extraction of glucose also increased in response to insulin (0.07 ± 0.02 to 0.19 ± 0.05 to 0.34 ± 0.07 in SHAM and 0.05 ± 0.01 to 0.13 ± 0.02 to 0.23 ± 0.03 in INF). In the
Hi-Ins period, net hindlimb fractional extraction of glucose was
significantly decreased with infection. Net hindlimb lactate uptake was
lower in INF relative to SHAM in the Lo-Ins period (4.5 ± 1.3 vs.
0.9 ± 0.4 mg/min, P < 0.05); it tended to fall
when insulin was increased (to 2.6 ± 1.2 and 0.1 ± 1.4 mg/min in SHAM and to 0.8 ± 0.6 and 0.3 ± 0.6 mg/min in
INF), although the changes were not significantly different.
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DISCUSSION |
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The liver consumes ~45% of the glucose infused in chronic TPN, and peripheral tissues consume the remaining 55%. The presence of an infection sharply decreases NHGU; consequently, peripheral glucose uptake increases to sustain whole body glucose uptake. We previously demonstrated that the compensatory rise in insulin limits the fall in NHGU and increases peripheral glucose uptake during infection (3). Our study demonstrates that both hepatic insulin sensitivity (ED50) and responsiveness are enhanced in the TPN-adapted state (i.e., SHAM). We also demonstrate that infection impairs both the responsiveness and the sensitivity of the liver to insulin. Although the liver is reliant on the compensatory hyperinsulinemia to sustain liver glucose uptake during an infection, NHGU does not improve further with even higher doses of insulin. Moreover, the modest benefit of improving NHGU gained by the additional insulin predisposes the liver to accumulate glycogen rather than to improve hepatic glycolysis.
Chronic TPN enhanced the responsiveness and sensitivity (i.e.,
ED50) of NHGU to insulin. As we have reported previously
(22), NHGU in sham TPN-adapted dogs was substantially
higher than rates achieved in normal 42-h-fasted dogs (3.5 vs. 0.55 mg · kg
1 · min
1)
despite lower insulin and glucose levels (sinusoidal insulin, 35 vs. 51 µU/ml; arterial plasma glucose, 120 vs. 240 mg/dl). Moreover, NHGU
with chronic TPN increased to 4.6 mg · kg
1 · min
1
at the higher insulin level (163 µU/ml), which corresponded to an HFE
of 0.20. In contrast, in the 42-h-fasted dog, NHGU and HFE were only
3.0 mg · kg
1 · min
1
and 0.04, respectively, after ~4 h of combined hyperglycemia (200 mg/dl) and hyperinsulinemia (191 µU/ml) (20). TPN also enhances the sensitivity of NHGU to insulin. In the TPN-adapted state,
when the sinusoidal insulin concentration was halved (19 µU/ml) in a
SHAM dog, NHGU fell by ~50%. We estimate hepatic insulin sensitivity
(ED50) to be below a sinusoidal insulin concentration of 25 µU/ml in TPN-adapted animals, which is substantially lower than that
seen in the 42-h-fasted dog (~90 µU/ml) (20). Thus chronic TPN administration enhances both the sensitivity and the responsiveness of the liver to insulin.
Infection impairs both the sensitivity and the responsiveness of NHGU
to insulin in the TPN-adapted setting. At the low sinusoidal insulin
concentration (29 µU/ml), NHGU was 70% lower in INF than in SHAM.
When insulin was raised to the levels normally seen during infection,
NHGU increased to 2.6 mg · kg1 · min
1.
However, when the sinusoidal insulin concentration was raised above 66 µU/ml, NHGU was not significantly elevated, suggesting that a
near-maximal response was attained. Compared with SHAM, NHGU (2.8 mg · kg
1 · min
1)
and HFE (0.08) were 38 and 60% lower, respectively, in INF at the
highest insulin dose studied. Moreover, we observed that NHGU was
completely suppressed at a sinusoidal insulin level of 14 µU/ml.
Hepatic insulin sensitivity (ED50) was estimated to be at a
sinusoidal insulin level of ~40 µU/ml for infected animals receiving TPN. There are no other reports of the impact of infection on
the sensitivity of liver glucose uptake to insulin. There are, however,
conflicting results as to infection's impact on the sensitivity of HGP
to insulin. Lang et al. (9) found no impairment in the suppression of tracer-determined HGP by insulin in conscious septic rats. McLane et al. (16) observed that insulin's
suppression of HGP was impaired in anesthetized septic dogs. In the
latter study, in contrast to our study, glucagon, a potent regulator of
HGP and HGU, was not controlled. Thus our studies demonstrate for the
first time that infection reduces both the sensitivity and the
responsiveness of NHGU to insulin.
One factor contributing to the infection-induced decrease in NHGU at
low, but not at high, insulin levels was an elevated rate of HGP. The
magnitude of NHGU is determined by the balance between HGU and HGP,
which was determined by combining the tracer and arteriovenous
difference methods. At the lowest insulin dose, HGP was greater in INF
than in SHAM. As insulin was increased, HGP in both groups declined to
a rate not different from zero. Consistent with our previous finding
(3), the insulin-induced improvement in NHGU in INF
results from a fall in HGP (0.8 ± 0.3 mg · kg
1 · min
1)
as well as a stimulation of HGU (
0.8 ± 0.3 mg · kg
1 · min
1).
In contrast, HGP in SHAM was already suppressed at the lowest insulin
dose; therefore, the modest increase in NHGU in response to insulin was
due solely to an increase in HGU.
The persistent HGP in the INF group in the presence of relatively low insulin levels was derived from both glycogenolysis and gluconeogenesis. Consistent with activation of glycogenolysis during infection, the percentage of glucose released as lactate was greater than 100%, suggesting an intrahepatic source of glucose carbon to support lactate release. This percentage rapidly decreased as insulin was increased, which is consistent with a suppression of glycogenolysis by insulin. As was seen in studies in the fasted setting, gluconeogenic precursor uptake is increased by infection (12), and this increase persisted in the TPN-adapted setting. The increase in gluconeogenic precursor uptake was supported by the elevated HFE of the gluconeogenic amino acids, which is likely driven by the characteristic hyperglucagonemia seen during infection (15). Total gluconeogenic precursor uptake was not altered by the stepwise increase in insulin. Because glucose production was suppressed by insulin, presumably, the gluconeogenic carbon was either oxidized or diverted to glycogen. The lack of an acute effect of insulin on gluconeogenesis is consistent with recent work (5) that suggests that the primary acute action of insulin on the liver is on glycogenolysis. Thus administration of additional insulin does restrain the infection-induced increase in glycogenolysis but does not suppress the infection-induced increase in gluconeogenic precursor uptake.
Despite marked activation of hepatic glycolysis (lactate release) in the TPN-adapted state, physiological increases in insulin did not augment NHLR in either SHAM or INF animals. Instead, elevations in insulin modulated glucose storage (e.g., glycogen synthesis). This result agrees with data from normal fasted animals, in which the major intrahepatic fate of glucose in response to insulin was glycogen synthesis, with only ~10% of NHGU being diverted to lactate release (22). Thus the capacity to enhance glycogen synthesis far exceeds the glycolytic capacity of the liver, even in the TPN-adapted state.
Glycogen deposition was the principal fate of additional glucose
consumed by the liver. Terminal glycogen content was ~60% lower in
INF (83 ± 9 vs. 36 ± 6 mg/g), which supports our previous observation that infection markedly reduces glycogen content during TPN
administration [77 ± 2 vs. 21 ± 3 mg/g (4)].
By use of these historical controls, insulin-mediated net glycogen
deposition was calculated to be 6 and 15 mg/g in SHAM and INF,
respectively. These are very similar to the theoretical increments in
glycogen mass attributed to insulin [7 and 11 mg/g; NHGU
mg · kg
1 · min
1 × time/(liver mass/body wt ratio); SHAM and INF] if all of the additional glucose consumed by the liver were deposited as glycogen. Moreover, the theoretical increase in glycogen mass in infected animals
was likely higher than the calculated value. When the insulin level in
the infected animals was reduced to the level observed in sham-infected
dogs, glycogenolysis was enhanced (3), resulting in a
further decline in glycogen content before the experimental increases
in insulin. Thus the increment in glycogen mass can account for the
majority of the insulin-mediated increase in NHGU.
The GIR required to maintain isoglycemia during each insulin period was reduced in infected animals, suggesting a state of whole body insulin resistance. Peripheral insulin resistance, predominantly in skeletal muscle, is a well-known characteristic of critical illness (10). Figure 5 shows that nonhepatic tissues were responsible for the majority of the increase in whole body glucose uptake in response to insulin. In the Lo-Ins period, net nonhepatic glucose uptake rates were equivalent in SHAM and INF animals. This is not surprising. An increase in insulin-independent glucose uptake during inflammatory stress could offset any decrease in insulin-mediated glucose disposal. This would be most evident at low insulin concentrations, where insulin-mediated glucose uptake is a small fraction of the total glucose uptake (17). A corresponding impairment in LGU was not detected at any insulin dose because of a large variability in iliac arterial blood flow. However, a variable that is less sensitive to variation in blood flow, fractional extraction of glucose across the hindlimb, was reduced in INF at the highest insulin dose.
In summary, our in vivo studies demonstrate that, following the adaptation to TPN, 1) the liver is more sensitive and responsive to insulin than in a normal animal, 2) the presence of an infection reduces both the responsiveness and the sensitivity of NHGU to insulin, and 3) despite marked augmentation of NHLR in the TPN-adapted state, increases in insulin do not correct the infection-induced impairment in glycolysis. These results suggest that, although an increase in arterial insulin commonly seen during infection will minimize the infection-induced impairment in liver glucose uptake, insulin did not reverse the inhibition of glycolysis. Additional insulin will not enhance NHGU further. The implication of this study is that, although the administration of insulin will correct the hyperglycemia induced by an infection or critical illness, insulin will not overcome the defects in hepatic metabolism. Any beneficial effects of insulin on liver glucose uptake do not lead to a corresponding rise in glycolysis and in fact favor glucose storage, initially in the form of glycogen. The long-term consequence of sustained hepatic storage of carbohydrate is unclear.
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ACKNOWLEDGEMENTS |
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We thank Ying Yang, Eric Allen, Angelina Penaloza, and Wanda Snead for assistance with amino acid and hormone analyses.
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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 University, 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.
First published November 19, 2002;10.1152/ajpendo.00035.2002
Received 30 January 2002; accepted in final form 14 November 2002.
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