Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, TN 37232-0615
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ABSTRACT |
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We examined the impact of infection on hepatic
and muscle glucose metabolism in dogs adapted to chronic total
parenteral nutrition (TPN). Studies were done in five conscious
chronically catheterized dogs, in which sampling (artery, portal and
hepatic vein, and iliac vein), infusion catheters (inferior vena cava),
and Transonic flow probes (hepatic artery, portal vein, and iliac
artery) were implanted. Fourteen days after surgery, dogs were placed
on TPN. After 5 days of TPN, an infection was induced, and the TPN was continued. The balance of substrates across the liver and limb was
assessed on the day before infection
(day
0) and 18 (day
1) and 42 h
(day
2) after infection. On
day
0, the liver was a marked net consumer
of glucose (4.3 ± 0.6 mg · kg1 · min
1)
despite near normoglycemia (117 ± 5 mg/dl) and only mild
hyperinsulinemia (16 ± 2 µU/ml). In addition, the majority (79 ± 13%) of the glucose taken up by the liver was released as
lactate (34 ± 6 µmol · kg
1 · min
1).
After infection, net hepatic glucose uptake decreased markedly on
day 1 (1.6 ± 0.9 mg · kg
1 · min
1)
and remained suppressed on day
2 (2.4 ± 0.5 mg · kg
1 · min
1).
Net hepatic lactate output also decreased on
days
1 and
2 (15 ± 5 and 12 ± 3 µmol · kg
1 · min
1, respectively). This
occurred despite increases in arterial plasma glucose on
days
1 and
2 (135 ± 9 and 144 ± 9 mg/dl, respectively) and insulin levels on
days
1 and
2 (57 ± 14 and 34 ± 9 µU/ml, respectively). In summary, the liver undergoes a profound adaptation to
TPN, making it a major site of glucose disposal and conversion to
lactate. Infection impairs hepatic glucose uptake, forcing TPN-derived
glucose to be removed by peripheral tissues.
liver blood flow; lactate; intestine
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INTRODUCTION |
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INFECTION leads to marked alterations in whole body glucose metabolism, characterized by an acceleration in glucose production and utilization (17). The increase in glucose production is derived from an increase in gluconeogenesis, driven by the elevated counterregulatory hormone environment (17).
In this environment of elevated glucose flux, the ability of infected individuals to dispose of exogenous glucose is impaired, as evidenced by exaggerated hyperglycemia during glucose-based parenteral nutrition (15, 19). Peripheral insulin resistance and impaired suppression of hepatic glucose production contribute to the glucose intolerance (12). The peripheral insulin resistance is marked; insulin-stimulated whole body glucose utilization is decreased by ~50% (11, 26).
An impairment in net hepatic glucose uptake also contributes to the abnormal glucose tolerance seen during an acute infusion of glucose. During a 180-min intravenous glucose infusion, net hepatic glucose uptake was 25% of normal in infected animals (19). Consistent with previous reports (28), a failure to suppress endogenous glucose production contributed ~40% of the impairment in net hepatic glucose uptake. The remaining 60% was due to a failure to increase unidirectional hepatic glucose uptake despite marked hyperinsulinemia.
The majority of stressed patients are not given glucose alone; rather glucose is given in combination with specific fats and amino acids. In addition, nutrients are administered on a chronic basis. The response of the liver to chronic nutritional support and its interaction with infection have not been examined. In the present study, we examined the balance of glucose and substrates across the liver and muscle in chronically catheterized conscious dogs receiving continuous total parenteral nutrition (TPN). In addition, we assessed the impact of infection on the TPN-adapted liver.
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MATERIALS AND METHODS |
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Animal preparation. Experiments were carried out on five conscious female mongrel dogs (20 ± 1 kg). Before being studied, they received a diet consisting of KalKan meat (Vernon, CA) and Purina dog chow (St. Louis, MO) once daily. The composition of the diet was 52% carbohydrate, 31% protein, 11% fat, and 6% fiber based on dry weight. The dogs were housed in a facility that met American Association for Accreditation of Laboratory Animal Care guidelines, and the experimental protocols were approved by the Vanderbilt University Animal Care Subcommittee.
Experimental preparation. Fourteen to seventeen days before a study, a laparotomy was performed under general anesthesia (isoflurane). Infusion catheters were placed into the inferior vena cava for infusion of TPN. Sampling catheters (0.04 inch ID) were inserted into the portal vein and the left common hepatic vein for blood sampling. Additional catheters (0.04 inch ID) for blood sampling were inserted into the femoral artery after an incision was made in the left inguinal area, and into the common iliac vein after an incision was made in the right inguinal region. The catheters were then filled with saline containing heparin (200 U/ml). Doppler flow probes were placed about the external iliac artery, portal vein, and hepatic artery after the gastroduodenal vein was ligated. The portal and hepatic vein sampling catheters and the Doppler flow probe leads were exteriorized and placed in a subcutaneous pocket in the abdominal area. The free ends of the inferior vena cava infusion catheters were exteriorized, tunneled subcutaneously, and placed under the skin between the clavicles. The femoral artery and iliac vein sampling catheters were placed under the skin in the inguinal region (20).
Two weeks after catheter implantation all animals had
1) a good appetite (consuming the
entire daily ration), 2) a normal stool, 3) a hematocrit above 35%,
and 4) a leukocyte count <18,000 mm3 before the study.
Nutritional support. Fourteen days
after catheter implantation, dogs were switched to TPN as the sole
exogenous calorie source for a 7-day period. The nutritional support
was designed to match their calculated basal energy requirements
[144 + (62.2 × BW) kcal/day (30)], where BW is body
weight in kilograms. Nitrogen (grams of protein/day)
requirements were calculated according to the formula (1.5 × BW0.67). This was met by the
infusion of Travasol (10%; Clintec Nutrition, Deerfield, IL).
Twenty-five percent of the nonprotein energy requirements were derived
from fat (20% Intralipid, Clintec Nutrition) and 75% from dextrose
(50%; Baxter, Toronto, ON, Canada). The TPN also included potassium
phosphate (45 mM; 90 mg
potassium · kg1 · day
1)
and sodium chloride (0.9%; 2.9 ml · kg
1 · day
1),
as well as a multivitamin infusion (MVI-12; Astra USA, Westborough, MA). The dogs had free access to water.
Parenteral nutrients were given via two catheters placed into the inferior vena cava at the time of surgery. The dog was placed in a jacket (Alice King Chatham, Los Angeles, CA) containing a pocket into which two portable infusion pumps were placed (INFUMED; Medfusion Systems, Norcross, GA). The nutritional support was given continuously over the 7-day period. TPN was prepared under sterile conditions and changed once daily.
Experimental protocol. After 5 days of TPN, the sampling catheters and free ends of the Transonic flow probes were removed from subcutaneous pockets under local anesthesia (2% lidocaine). The dog was then placed into a Pavlov harness. After a 60-min equilibration period, blood samples were taken at 60, 75, 90, 105, and 120 min from the four sampling catheters (femoral artery, portal vein, hepatic vein, and iliac vein; day 0). Liver and limb blood flow were assessed with Transonic flow probes. The free ends of the flow probes were then placed in their subcutaneous pockets, and an infection was induced. Eighteen (day 1) and forty-two (day 2) hours after the induction of infection, the balance of substrates was reassessed as on day 0. At the end of the study, each dog was euthanized with an overdose of pentobarbital sodium.
Induction of infection. On the day of induction of an infection, a fibrin clot was prepared from a 1% bovine fibrinogen solution (10 ml/kg; Sigma, St. Louis, MO), which was then filtered through a sterile 0.45 µM filter. Bacteria were prepared by inoculating 1 liter of trypticase soy broth (Becton-Dickinson, Cockeysville, MD) and were incubated overnight at 37°C. The next day the bacteria were pelleted by centrifugation, washed with sterile saline, and reconstituted in 20 ml of sterile saline. The dose of bacteria (Escherichia coli; ATTC no. 25922) was 2 × 109 organisms/kg. The dose of bacteria was determined by serial dilution of the bacteria followed by plating. The bacteria were mixed with the fibrinogen, and thrombin (1,000 units) was added to initiate clot formation. The model used is nonlethal (17) and is similar to the model of Fink et al. (7).
After the fibrin clot was prepared, the dog was placed under general anesthesia. An abdominal midline laparotomy incision was made at a point below that made 2 wk earlier. A clot containing viable bacteria was placed into the peritoneal cavity. Dogs received 500 ml of saline immediately after clot implantation. An additional 1,000 ml was given on day 1.
Processing of blood samples. Blood
samples were drawn into heparinized syringes and transferred to chilled
tubes containing potassium EDTA (15 mg). The collection and immediate
processing of blood samples have been previously described (3). Blood lactate, glycerol, and alanine were analyzed with the method of Lloyd
et al. (14) on a Monarch 2000 centrifugal analyzer (Instrumentation Laboratories, Lexington, MA). Plasma glucose was assayed immediately with a Beckman Glucose Analyzer II (Beckman Instruments, Fullerton, CA). Plasma nonesterified fatty acids were determined
spectrophotometrically (Wako Chemicals, Richmond, VA). Immunoreactive
insulin (31) was assayed with a double antibody technique
[Pharmacia Diagnostics, Piscataway, NJ; intra-assay coefficient
of variation (CV) of 11%]. Plasma treated with 500 kallikrein-inhibitor units of Trasylol (Miles, Kankakee, IL) was
assayed for immunoreactive glucagon (1) with a procedure similar to
that for insulin (intra-assay CV of 8%). Plasma cortisol (6) was
assayed with Clinical Assays Gamma Coat radioimmunoassay kit
(intra-assay CV of 6%). Plasma collected from blood samples that were
immediately treated with EGTA and glutathione was assayed for
epinephrine and norepinephrine with HPLC techniques (CV of 14%) (16),
as modified by Davis et al. (5). Hepatic artery, iliac artery, and
portal vein blood flow were assessed with Transonic flow probes
(Transonic Systems, Ithaca, NY). Blood flow was converted to plasma
flow by multiplying by 1 hematocrit ratio.
Calculations. Net hepatic glucose
uptake was calculated with the formula
[(Fa × A + Fp × P) H] × HBF, where H, A, and P are the blood glucose concentrations in
the hepatic vein, femoral artery, and portal vein, respectively, and
Fa and
Fp represent the fractional
contributions of the hepatic artery and portal vein, respectively, to
total hepatic blood flow (HBF). Net fractional hepatic glucose
extraction was calculated as the ratio of net hepatic glucose uptake
and hepatic glucose load. Hepatic glucose load was calculated with the
formula (Fa × A + Fp × P) × HBF. Plasma
glucose concentrations were converted to whole blood concentrations with a correction factor of 0.73 (23). The above equations were used to
calculate net hepatic lactate, alanine, nonesterified free fatty acid
(NEFA), and glycerol uptake and fractional extraction as well. However,
because the liver was a net producer (i.e., negative uptake) of
lactate, the lactate data are presented as positive values and denoted
as net output. Plasma flow was used instead of blood flow to calculate
net hepatic NEFA uptake. In an analogous way, net limb substrate uptake
was calculated with the formula [A
V] × LBF,
where V is the blood substrate concentrations in the iliac vein and LBF
is blood flow in the iliac artery.
Statistics. Hepatic blood flow and substrate flux are expressed on a per kilogram body weight basis. Statistical comparisons were made with ANOVA (Systat for Windows; Systat, Evanston, IL). A univariate post hoc F-test was used when a significant F ratio was found. Statistical significance was accepted at P < 0.05.
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RESULTS |
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Liver and limb blood flow. On
day 0 before induction of infection, hepatic artery blood flow and portal
vein blood flow were 5.9 ± 1.3 and 26.5 ± 2.7 ml · kg1 · min
1,
respectively (Table 1). After the induction
of infection, hepatic artery blood flow increased to 20.3 ± 4.6 and
11.6 ± 3.0 ml · kg
1 · min
1
on days
1 and
2 (P < 0.05). Portal vein blood flow was unaltered by infection.
Consequently, total hepatic blood flow increased from 32 ± 2 ml · kg
1 · min
1
to 51 ± 8.8 ml · kg
1 · min
1
on day
1 (P < 0.05) and 37.3 ± 3.9 ml · kg
1 · min
1
on day
2. Thus the contribution of the
hepatic artery to total hepatic blood flow increased from 19 ± 5%
on day
0 to 40 ± 6 and 31 ± 8% on
days
1 and
2, respectively
(P < 0.05). Iliac vein blood flow
was not significantly increased by infection (Table 1).
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Hormone levels. Arterial insulin levels were 16 ± 2 µU/ml on day 0 and increased to 57 ± 14 µU/ml on day 1 and 34 ± 9 µU/ml on day 2 (Table 2; P < 0.05). Arterial plasma glucagon levels increased from 30 ± 6 pg/ml to 132 ± 22 (P < 0.05) and 80 ± 28 pg/ml, on days 1 and 2, respectively. Arterial cortisol levels were 3 ± 1, 5 ± 2, and 4 ± 1 µg/dl on days 0, 1, and 2, respectively. Respective arterial plasma epinephrine levels were 199 ± 46, 271 ± 96, and 203 ± 103 pg/ml, and those for arterial plasma norepinephrine were 276 ± 81, 423 ± 121, and 477 ± 123 pg/ml.
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Hepatic metabolism.
Arterial plasma glucose levels increased from 117 ± 5 mg/dl on
day 0 to 135 ± 9 and 144 ± 9 mg/dl on
days 1 and
2, respectively
(P < 0.05; Fig.
1). On day
0, the liver was a net consumer of
glucose (4.3 ± 0.6 mg · kg1 · min
1).
On days
1 and
2, despite hyperglycemia and
hyperinsulinemia, net hepatic glucose uptake decreased to 1.6 ± 0.9 and 2.4 ± 0.5 mg · kg
1 · min
1
(P < 0.05). Net fractional hepatic
glucose extraction was markedly elevated with TPN and decreased
markedly after infection. The fraction of TPN-infused glucose (9.5 ± 0.1 mg · kg
1 · min
1)
removed by the liver was 45 ± 6% on
day 0 and decreased markedly to 16 ± 9 and 25 ± 5% (Fig.
2) on days
1 and
2.
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Arterial lactate levels were 1.5 ± 0.1 mM on
day
0, were not altered on
day 1 (1.5 ± 0.2 mM), and were decreased on
day 2 (P < 0.05; 1.0 ± 0.1 mM; Fig.
3). The liver was a marked producer of
lactate on day
0 (34 ± 6 µmol · kg1 · min
1).
After infection, net hepatic lactate release decreased to 15 ± 5 and 12 ± 3 µmol · kg
1 · min
1
on days
1 and
2 (P < 0.05). On day
0, 79 ± 13% of the glucose taken
up by the liver was converted to lactate. After infection, this
percentage did not decrease on day
1 (84 ± 51%). However, on
day
2, it was decreased (44 ± 14%;
P < 0.05).
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Arterial alanine levels fell from 509 ± 51 µM on
day 0 to 273 ± 23 and 342 ± 46 µM on
days
1 and
2 (P < 0.05; Fig. 4). Net hepatic alanine
uptake increased (P < 0.05) from 1.4 ± 0.3 µmol · kg1 · min
1
to 2.7 ± 0.4 on day
1 and tended to return toward normal
by day 2 (1.9 ± 0.3 µmol · kg
1 · min
1).
A parallel rise in net fractional hepatic alanine extraction also
occurred (0.08 ± 0.2 to 0.21 ± 0.03 to 0.16 ± 0.02 on
days 0, 1,
and 2, respectively.).
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Arterial glycerol levels were not altered by infection (Table 3). Net hepatic glycerol uptake was increased on days 1 and 2. Net fractional hepatic glycerol extraction was not altered.
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Arterial NEFA levels were not altered by infection (Table 3). However, both net hepatic NEFA uptake and fractional hepatic extraction were increased on day 1. On day 2, net hepatic NEFA uptake was not significantly elevated, although net fractional hepatic NEFA extraction remained elevated.
Gut metabolism. On
day
0, net gut glucose uptake was 0.6 ± 0.1 mg · kg1 · min
1.
After the induction of an infection, net gut glucose uptake increased
to 1.2 ± 0.3 mg · kg
1 · min
1
on day
1, and on
day 2 it returned to the basal rate, 0.7 ± 0.1 mg · kg
1 · min
1.
The intestine was a net consumer of lactate on
day 0 (2.5 ± 0.9 µmol · kg
1 · min
1).
On days
1 and
2, the gut lactate uptake decreased to
0.8 ± 0.4 and 0.7 ± 0.8 µmol · kg
1 · min
1,
respectively. The gut was a net producer of alanine (0.7 ± 0.1 µmol · kg
1 · min
1)
on day
0, and net gut alanine release was not
altered on days 1 and
2 (0.6 ± 0.2 and 0.3 ± 0.2 µmol · kg
1 · min
1).
Limb metabolism. On day 0, limb glucose uptake and fractional glucose extraction were 6 ± 1 mg/min and 0.04 ± 0.01, respectively (Fig. 5). On day 1, after the induction of infection, limb glucose uptake increased (18 ± 8 mg/min) and returned toward normal on day 2 (10 ± 3 mg/min). Limb glucose fractional extraction was 0.08 ± 0.04 and 0.05 ± 0.01 on days 1 and 2, respectively.
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On day
0, limb lactate uptake and limb
lactate fractional extraction were 34 ± 5 µmol/min and 0.18 ± 0.04, respectively. On day
1, limb lactate uptake and fractional
extraction were decreased (3 ± 17 µmol/min and 0.0 ± 0.04, respectively). They remained decreased on
day 2 (2 ± 8 µmol/min and 0.05 ± 0.04 .
The limb increased its release of glycerol from 0.2 ± 1.1 µmol/min on day 0 to 5.3 ± 1.6 and 4.1 ± 1.9 µmol/min on days 1 and 2, respectively. The limb release of NEFA was unaltered (0.4 ± 2.7 to 3.5 ± 6.8 to 2.9 ± 6.6 µmol/min on days 0, 1, and 2).
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DISCUSSION |
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These studies demonstrate that the liver has a remarkable ability to adapt to chronic (5 days) total parenteral nutrition. In the presence of mild hyperinsulinemia and normoglycemia, the liver became a marked consumer of glucose (~45% of exogenous glucose infusion). Surprisingly, the liver did not retain the glucose carbon. Rather, it converted ~80% of the glucose to lactate, which was consumed by peripheral tissues such as those of the limb (i.e., muscle). After the induction of infection, net hepatic glucose uptake and lactate release decreased markedly. A parallel rise in limb glucose uptake and a fall in limb lactate uptake were observed. Thus the adaptation of the liver to TPN is essential in limiting the hyperglycemia and hyperinsulinemia that would otherwise be seen. Infection attenuates this adaptation, forcing peripheral tissues to consume a greater portion of the glucose, which results in hyperglycemia and hyperinsulinemia.
The most surprising finding of the study was that the liver could take
up substantial quantities of glucose (4.3 mg · kg1 · min
1)
in the absence of hyperglycemia (117 mg/dl) and only mild
hyperinsulinemia (17 µU/ml). The substantial uptake of glucose by the
liver after TPN (45% of the exogenous glucose) is greater than that
which would have been predicted on the basis of acute studies (4). The
three factors that regulate net hepatic glucose uptake are the hepatic
glucose load (flow × level), the hormonal milieu (insulin and
glucagon levels), and the route of glucose delivery. Physiological increases in glucose alone will not induce net hepatic glucose uptake.
However, in the presence of elevated glucose levels, net hepatic
glucose uptake increases in a dose-dependent manner with a rise in
insulin. This is further enhanced if the glucose is administered into
the portal vein (which activates the "portal" signal) (23, 25).
To acutely increase net hepatic glucose uptake to ~4.3
mg · kg
1 · min
1,
the arterial plasma glucose levels would have to be greater than 200 mg/dl, and the arterial insulin levels would have to exceed 40 µU/ml
(23). In addition, the glucose would have to be infused into the portal
vein. Thus TPN has increased the capacity of the liver to consume
glucose. This response is similar to but somewhat greater than those
responses seen in malnourished humans receiving TPN in which the
splanchnic bed removed ~20% of the TPN-derived glucose (9). The
lower glucose uptake in the human may be due, in part, to the fact that
the subjects were not healthy and they had elevated glucagon levels.
One likely explanation for the enhancement in liver glucose uptake is
an increase in glucokinase activity (2, 24), possibly initiated by the
mild hyperinsulinemia and the very low glucagon levels seen in the normal dog receiving TPN. Other glycolytic enzymes must also be involved, because activation of glucokinase alone cannot explain the
marked increase in hepatic glycolysis observed (29).
The major fate of glucose taken up by the liver of TPN-adapted dogs was
conversion to lactate and subsequent release. If a liver is acutely
induced to consume ~4
mg · kg1 · min
1
of glucose by exposure to combined hyperinsulinemia and hyperglycemia, only a small fraction (~10%) of the glucose is released as lactate (4). The majority of the consumed glucose is converted to glycogen. In
response to chronic TPN, however, a large fraction (~80%) of the
glucose was converted to lactate. As is the case for the high rate of
glucose uptake, the mechanism is unclear.
The hepatic conversion of glucose to lactate allows the peripheral tissues to consume ~90% of the exogenous carbohydrate without marked hyperinsulinemia in the normal TPN-adapted dog. Insulin levels were only increased twofold over those seen in overnight-fasted dogs (4). Consistent with this, limb fractional extraction and uptake of glucose were elevated compared with normal overnight-fasted dogs (32). Whereas the limb was consuming more glucose than in a fasted state, nearly 50% of the limb carbohydrate uptake was supplied as lactate. The increased dependency on lactate allows the muscle to utilize large quantities of carbohydrate without exaggerated hyperinsulinemia.
Infection led to a marked inhibition of liver glucose uptake. The mechanism for the infection-induced decrease in net hepatic glucose uptake is unknown. A fall in net hepatic glucose uptake could be due to either a decrease in the entry of glucose into the liver and/or a corresponding increase in hepatic glucose production. In response to an acute infusion of glucose into a peripheral vein, infection decreases net hepatic glucose uptake because of an impaired entry of glucose as well as an impaired suppression of hepatic glucose production (19). In septic patients receiving overnight glucose infusion, endogenous glucose production and alanine gluconeogenesis are not completely suppressed (27). Because the response of the liver to acute glucose infusion and chronic TPN infusion is markedly different, it is uncertain whether a similar explanation would apply to an infected animal receiving nutritional support. The factors responsible for the alterations in net hepatic glucose uptake are unclear. However, it is likely that the infection-induced hyperglucagonemia plays a central role. Hyperglucagonemia when combined with the available nutrients (amino acids and lipid supplied by the TPN) can increase the rate of gluconeogenesis (15, 18, 19).
After infection, the peripheral tissues were forced to dispose of the
TPN-derived glucose directly, because liver glucose uptake was markedly
decreased. Extrahepatic glucose uptake (exogenous glucose infusion
rate net hepatic glucose uptake) increased by 52 and
37% on days
1 and
2. Some of this increase occurred in
muscle, because limb glucose uptake increased markedly on
day 1 and remained somewhat elevated on day
2 (the hindlimb in the dog is ~65%
skeletal muscle by weight). The decrease in net hepatic lactate release
was paralleled by a near-complete cessation of net limb lactate uptake
on days
1 and
2. The prevailing hyperinsulinemia likely contributed to the increase in limb glucose uptake. However, because infection also increases insulin-independent glucose uptake (10), this may also contribute to the increase.
Arterial alanine levels were likely altered by infection because of the infection-induced rise in glucagon. On day 0, both arterial glucagon levels and net fractional hepatic alanine extraction were markedly lower than in overnight-fasted dogs (30 vs. 50 pg/ml and 0.08 vs. 0.30). After infection, arterial alanine levels fell because of the rise in net hepatic alanine fractional extraction. The rise in net fractional hepatic alanine extraction is consistent with the known potent effects of glucagon on this parameter (21).
The liver became more reliant on fatty acids after infection. Nonesterified fatty acid uptake and extraction were increased during infection. In addition, net hepatic glycerol uptake was increased. These data are consistent with the known acceleration of lipolysis during infection. Because the levels of these substrates did not increase, the primary mechanism for the increase in uptake is an increase in hepatic blood flow and fractional extraction of these precursors.
Infection leads to a marked increase in liver blood flow, predominantly by enhancing hepatic artery blood flow. This increase has been reported in the rat (8). The mechanism is unclear. However, because liver energy demands increase after infection, it may activate the hepatic artery buffer response (13) or possibly an infection-induced increase in local release of vasodilators, such as nitric oxide (22).
In summary, after adaptation to TPN, the liver becomes a major site of glucose disposal. However, the liver does not retain the glucose skeleton; rather, it converts it to lactate, which is subsequently released and consumed by peripheral tissues. This adaptation allows peripheral tissues to remove large amounts of carbohydrate while minimizing the insulin requirements. After infection, this cycle is disrupted. Net hepatic glucose uptake is markedly inhibited, and the conversion to lactate is diminished. The result is that the peripheral tissues must directly remove a greater fraction of the TPN-derived glucose. Because infection also leads to peripheral insulin resistance, the insulin requirements are further amplified, resulting in hyperglycemia.
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ACKNOWLEDGEMENTS |
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We are grateful for the technical assistance of Pamela Venson and Eric Allen from the hormone core laboratory of the Vanderbilt University Diabetes Research and Training Center.
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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) and Diabetes Research and Training Center Grant P60-DK-20593.
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.
Address for reprint requests: O. P. McGuinness, 702 Light Hall, Dept. of Molecular Physiology and Biophysics, Vanderbilt Univ., Nashville, TN 37232-0615.
Received 17 April 1998; accepted in final form 3 August 1998.
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