Departments of 1 Anesthesia and Critical Care and 3 Surgery, Harvard Medical School, and Anesthesia and Surgical Services, Massachusetts General Hospital and Shriners Hospital for Children, Boston, Massachusetts 02114; and 2 Center for Developmental Biology, University of Texas Southwestern Medical Center, Dallas, Texas 75390
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The molecular mechanisms underlying endotoxin-induced insulin resistance remain unclear. Endotoxin or lipopolysaccharide (LPS) injection is a potent stimulator of inducible nitric oxide synthase (iNOS). This study in rats, using the specific iNOS inhibitor aminoguanidine, investigated the role of iNOS in endotoxin-induced hyperglycemia and insulin resistance. LPS injection led to hyperglycemia, insulin resistance, and increased iNOS protein expression and activity. Aminoguanidine prevented LPS-induced hyperglycemia without affecting insulin levels or iNOS expression. Aminoguanidine attenuated the LPS-induced insulin resistance, reflected by the requirement for a higher glucose infusion rate to maintain euglycemia during a hyperinsulinemic clamp study. Aminoguanidine completely blocked the LPS-elevated hepatic glucose output and also inhibited LPS-induced increases in hepatic glycogen phosphorylase activities and phosphoenolpyruvate carboxykinase (PEPCK) mRNA expression, key enzymes for glycogenolysis and gluconeogenesis, respectively. Thus, these data demonstrate an important role for iNOS in LPS-induced insulin resistance, evidenced by the attenuation of LPS-induced hyperglycemia and reversal of increased hepatic glucose output by aminoguanidine. The protective effect of aminoguanidine on insulin resistance is probably by attenuation of hepatic glucose output via its inhibition of key enzymes for glycogenolysis and gluconeogenesis, including glycogen phosphorylase and PEPCK.
endotoxin; hepatic glucose output; euglycemic hyperinsulinemic clamp; aminoguanidine; glycogen phosphorylase; phosphoenolpyruvate carboxykinase
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
PATIENTS with, and animal models of, sepsis or endotoxicosis exhibit many metabolic alterations, including attenuated responsiveness to insulin, whose actions include stimulation of peripheral glucose uptake (25, 53, 55), glycogen synthesis (55), protein synthesis (23), inhibition of glycogenolysis (26, 43), and gluconeogenesis (7). Attenuated responsiveness to the metabolic actions of insulin is termed insulin resistance. These metabolic alterations also exacerbate infection, retard recovery from disease, and lead to muscle wasting. Despite intensive investigation for a number of years, the molecular events responsible for insulin resistance in many pathological states, including stress and inflammation, still remain uncharacterized. Endotoxicosis is a pathological state associated with release of cytokines, systemic inflammation, hyperglycemia, insulin resistance, and the hypercatabolic state (43, 53), which is related to the decreased anabolic effects of insulin (8, 48).
Bacterial lipopolysaccharide (LPS) is a key mediator of many of the host responses resulting from gram-negative bacteremia and sepsis; it induces many genes involved in the immune, inflammatory, and acute phase responses. Among those genes, inducible nitric oxide synthase (iNOS) has been implicated in both protective (e.g., bactericidal) and detrimental host responses to sepsis and endotoxemia (46, 57, 59). Mice with knockout of iNOS are resistant to LPS-induced hypotension (13, 27, 51). Consistent with the data on iNOS knockout mice, an iNOS inhibitor, aminoguanidine (AG), reduced mortality after LPS administration to wild-type mice (51). Other iNOS-involved adverse host responses to endotoxin include vascular hyperpermeability (12), myocardial dysfunction (1), liver damage (24), and gut barrier failure (10, 32).
Accumulating evidence implicates a potential link between iNOS and
insulin resistance. First, iNOS is expressed in insulin-sensitive tissues such as liver (18), skeletal muscle
(17), and adipose tissue (37). Second,
pathological conditions exhibiting insulin resistance are associated
with induction of iNOS. These conditions include infection
(6), sepsis, burn (33), autoimmune deficiency syndrome (36), and elevated levels of tumor necrosis
factor- (TNF-
) (2, 19) and endotoxin (17,
18). Third, treatment of muscle cells with LPS, TNF-
, and
interferon-
induced iNOS expression and impaired insulin-stimulated
glucose uptake; the iNOS inhibitor AG reversed the altered glucose
uptake (2). Similarly, TNF-
attenuated
insulin-stimulated p70S6K activation, a key
downstream molecule of insulin signaling, by inducing iNOS expression
in pancreatic
-cells (19). Consistent with these
findings, nitric oxide (NO) donor impaired insulin-stimulated glucose
uptake in muscle cells (17), or decreased glycogen
synthase activity and glycogen synthesis, and increased glycogen
phosphorylase activity in primary hepatocytes (44).
Furthermore, NO donor administration caused hyperglycemia in dogs
(30) and insulin resistance in elderly healthy subjects
(31). Finally, the iNOS inhibitor AG inhibited the
development of hyperglycemia in genetically obese, diabetic
(fa/fa) rats (42) and lowered blood glucose levels in genetically obese, diabetic (db/db) mice
(35).
Thus these previous findings reveal a close relationship between iNOS, or NO, and insulin resistance. However, the effects of iNOS inhibitor on insulin sensitivity have not been studied in vivo as yet, and the potential involvement of iNOS in disrupted glycemic control in endotoxicosis or sepsis remains unexplored. This study, therefore, tested the hypothesis that the endotoxin-induced increased expression of iNOS plays a role in hyperglycemia and insulin resistance. This aim was achieved by use of a specific, competitive inhibitor of iNOS, AG. The effects of AG on glycemic control in vivo and on proglycemic enzymes in the liver were examined.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Animals. Adult male Sprague-Dawley rats (176-200 g), purchased from Taconic Farms (Germantown, NY), were used for the study. The Institutional Animal Care Committee approved the study protocol. The Association accredits the animal care facility for Assessment and Accreditation of Laboratory Animal Care. The rats were housed in mesh cages in a room maintained at 25°C and illuminated in 12:12-h light-dark cycles; they were provided with standard rodent chow and water ad libitum.
Measurement of blood glucose and insulin levels in LPS- and
AG-administered rats.
After overnight fasting (18 h), each of four groups of rats received
the injection of saline alone (group I), saline plus AG
(group II), LPS plus saline (group III), or LPS
plus AG (group IV). Just after the intraperitoneal injection
of LPS [20 mg/kg, derived from Escherichia coli Serotype
055:B5 (Sigma, St. Louis, MO)] or saline, AG (50 or 500 mg/kg, Sigma)
or saline was injected intraperitoneally. Blood samples were obtained
from the tail vein at 0, 60, 90, 120, 150, 180, and 240 min after LPS
injection to measure glucose and insulin levels. Blood glucose was
measured by an Elite glucometer (Bayer, Elkhart, IN). Insulin
concentrations in plasma, obtained from blood with heparin sodium, were
determined using an ELISA kit (Crystal Chem, Chicago, IL). The dose of
AG (500 mg/kg) was chosen on the basis of the previous studies. In those studies, the same or similar doses of AG were used with efficacy
to improve glycemic control in genetically obese, diabetic rats
(42) or to inhibit iNOS activities in animal models with various diseases, including endotoxicosis (39, 58),
pancreatic -cell damage (42, 47), diabetes (both types
I and II), wound healing (41), and radiation-induced lung
injury (50).
Euglycemic hyperinsulinemic clamp study.
The rats were anesthetized with an intraperitoneal injection of
pentobarbital sodium (60 mg/kg), and catheters (PE-50, PE-10; Becton-Dickinson, Sparks, MD) were implanted into the right jugular vein and the left carotid artery, as described previously
(15). The catheters were filled with saline containing
heparin sodium. At 5-6 days after the catheter placement,
euglycemic hyperinsulinemic clamp studies were performed in awake,
unrestrained rats in combination with the infusion of
[3H]glucose (Amersham Pharmacia, Piscataway, NJ) and
2-deoxy-[14C]glucose (Amersham Pharmacia), as previously
described (54). After an overnight fast, the infusion of
human regular insulin (Humulin R, Eli Lilly, Indianapolis, IN) was
started and maintained at the constant rate (20 mU · min1 · kg
1) via the
venous cannula. Blood samples were taken from the arterial catheter for
the measurement of blood glucose levels at 5- to 10-min intervals.
Euglycemia was maintained by varying the infusion rate of 50% dextrose
solution. At the inception of insulin infusion, [3H]glucose (6 µCi) was administered by the intravenous
bolus injection, followed by the constant infusion of 0.1 µCi/min. At
120 min after the initiation of insulin infusion,
2-deoxy-[14C]glucose (8 µCi) was administered by
intravenous bolus injection. At 120, 123, 130, 140, and 150 min after
the inception of insulin infusion, blood samples were collected to
measure [3H]glucose and
2-deoxy-[14C]glucose levels. LPS and/or AG was
administered at 60 min after the initiation of insulin infusion. At 150 min after the initiation of insulin infusion, the rats were
anesthetized with the intravenous injection of pentobarbital sodium (60 mg/kg), and then gastrocnemius muscle (skeletal muscle), liver, and
epididymal fat (adipose tissue) were taken and frozen in liquid
nitrogen. All of the tissue samples were stored at
80°C until
assayed. The rats were euthanized with an overdose of pentobarbital
(200 mg/kg ip).
Measurement of radiolabeled glucose concentrations in plasma and
tissues.
To determine the radioactivity of [3H]glucose and
2-deoxy-[14C]glucose in blood, aliquots of plasma (50 µl) obtained with heparin sodium were deproteinized with 100 µl of
Ba(OH)2 (0.3 N) and ZnSO4 (0.3 N) and then
centrifuged. The radioactivity of 3H and 14C in
the protein-free supernatants of Ba(OH)2 and
ZnSO4 precipitates was measured with a liquid scintillation
counter. To eliminate tritiated water, the supernatants were evaporated
to dryness for the measurement of [3H]glucose. Glucose
specific activities of plasma were calculated by dividing
3H and 14C radioactivity in the dried,
reconstituted aliquots by ambient plasma glucose concentration. To
measure the radioactivity of 2-deoxy-[14C]glucose in
tissues, the frozen tissue samples were digested with 1 ml of 1 M NaOH
at 60°C for 1 h and then neutralized with 1 ml of 1 M HCl.
Aliquots of 150 µl of digested tissue samples were mixed with 750 µl HClO4 (4% wt/vol) and then centrifuged. Thereafter,
the radioactivity of 14C in the supernatants was measured.
Tissue glucose uptake (Rg) was calculated as follows
(56)
![]() |
Calculation of hepatic glucose output.
Hepatic glucose output was determined as previously described
(38). Briefly, glucose appearance rate was calculated as
the ratio of the infusion rate of 3H (disintegration/min)
and the steady-state plasma [3H]glucose specific activity
(disintegration · min1 · mg
1).
The rate of hepatic glucose output was calculated as glucose appearance
rate minus the glucose infusion rate (hepatic glucose output = glucose appearance rate
glucose infusion rate).
Measurement of glycogen phosphorylase activity. The activities of glycogen phosphorylase in liver were measured as described previously (45). Briefly, after the frozen liver samples were ground with a porcelain mortar and pestle in the presence of liquid nitrogen, 125 mg of powdered liver samples were homogenized using a Polytron PT-MR 3000 (Kinematika, Littau, Switzerland) at maximum speed for 30 s in 5 ml of buffer containing 50 mM 2-(N-morpholino)ethanesulfonic acid (MES), 150 mM potassium fluoride, 5 mM EDTA, and 5 µg/ml leupeptin, pH 6.1. The homogenates were then centrifuged at 4°C for 10 min at 13,000 g. The supernatants were passed through the column (Micro Bio Spin Columns, Bio-Rad Laboratories, Hercules, CA) to remove AMP. Aliquots of 30 µl of the eluates were added to the assay buffer containing 50 mM MES, 50 mM sodium fluoride, 5 mM EDTA, 50 mM glucose 1-phosphate, 0.5 µCi/ml [14C]glucose 1-phosphate (Amersham Pharmacia), 27 mM 2-mercaptoethanol, and 10 mg/ml glycogen. Total glycogen phosphorylase activities were measured in the assay buffer containing 3 mM AMP. Glycogen phosphorylase-a activities were measured in the presence of 0.75 mM caffeine, an inhibitor of glycogen phosphorylase-b, in the absence of AMP (10). After the incubation at 25°C for 30 min, 25-µl aliquots were spotted on a Whatman filter (Whatman, Clifton, NJ). Then the filters were dropped into cold 66% ethanol for 30 min to precipitate glycogen and washed twice with 66% ethanol. The filters were then subjected to the measurement of 14C radioactivity with a liquid scintillation counter.
Detection of iNOS protein expression. iNOS expression was examined by immunoblotting. Sample preparation and immunoblotting were performed as previously described with minor modifications (16). Briefly, the frozen liver samples were ground with a porcelain mortar and pestle in the presence of liquid nitrogen and thereafter were homogenized using a Polytron PT-MR 3000 at maximum speed for 30 s in lysis buffer (50 mM HEPES/NaOH, pH 7.5, 150 mM NaCl, 2 mM EDTA, 1% Nonidet P-40, 10% glycerol, 10 mM sodium fluoride, 2 mM sodium vanadate, 1 mM phenylmethylsulfonyl fluoride, 10 mM sodium pyrophosphate, 5 µg/ml aprotinin, and 5 µg/ml leupeptin). After the incubation on ice for 30 min, the homogenized samples were centrifuged at 13,000 g for 30 min. Aliquots of the supernatant containing equal amounts of protein, as determined by the Bradford protein assay, were subjected to 10% SDS-PAGE after the addition of Laemmli sample buffer and boiling for 5 min. After transferring electrophoretically to nitrocellulose membrane (Bio-Rad), the membranes were blocked in 5% nonfat dry milk for 2 h at room temperature and incubated with anti-iNOS antibody (Upstate Biotechnology, Lake Placid, NY) for 2 h at room temperature. This was followed by the incubation with anti-rabbit IgG antibody conjugated with horseradish peroxidase (Amersham Pharmacia) for 30 min. Western blotting chemiluminescence luminol reagent (Santa Cruz Biotechnology, Santa Cruz, CA) visualized the blots. Bands of interest were scanned by MD-4000 (Alps Electric, San Jose, CA) and were quantified by use of National Institutes of Health Image 1.61 software (NTIS, Springfield, VA).
Measurement of iNOS activity. NOS activity was assayed by the conversion of L-[14C]arginine to L-[14C]citrulline, as described previously (4). Twenty-five microliters of tissue homogenate were added to 75 µl of reaction buffer containing 50 mM HEPES (pH 7.4), 1 mM NADPH, 0.1 mM tetrahydrobiopterin, 600 nM calmodulin, 0.24 mM CaCl2, 10 µM FAD, 1 mM EDTA, 1 mM dithiothreitol, and 20 µM L-arginine plus L-[14C]arginine (Amersham Pharmacia). After incubation for 10 min at 37°C, the reaction was terminated with 500 µl of stop solution containing 20 mM HEPES (pH 5.5) and 2 mM EDTA; it was applied to a 1-ml column of Dowex AG50Wx8 (Na+form) (Bio Rad Laboratories), which was eluted with 500 µl of water. L-[14C]citrulline was quantified by a liquid scintillation counter. iNOS activity was measured as Ca2+-independent NOS activities in the presence of 1 mM EGTA, being determined from the difference between samples with and without NOS inhibitor NG-monomethyl-L-arginine (L-NMMA, 10 mM; Calbiochem, San Diego, CA).
RNase protection assay.
Total RNA was isolated from liver tissue using the QuickPrep Total RNA
Extraction Kit (Amersham Pharmacia) according to the manufacturer's
instructions. The fragment of cDNA for phosphoenolpyruvate carboxykinase (PEPCK) was obtained from rat total liver RNA (Ambion, Austin, TX) by RT-PCR by use of the primers of
5'-ggaggtcactcaggaatccagttcttc-3' and
5'-gcaacacgactctctagttataactac-3', which was then subcloned into the
pCR2.1 vector (Invitrogen, Carlsbad, CA). An antisense riboprobe was
transcribed with T7 RNA polymerase (Ambion) by using [32P]UTP (Amersham Pharmacia). RNase protection assays
were carried out by using the RPA III kit (Ambion). Briefly, the
riboprobe for PEPCK was hybridized with 20 µg of total RNA extracted
from rat liver at 42°C for 18 h followed by RNase A/T1 digestion
at 37°C for 30 min. Protected fragments were heat denatured and
separated on 5% denaturing polyacrylamide gels, dried, and exposed to
film (Kodak BioMax MR, Rochester, NY). A rat -actin
antisense probe was used as a control. A cDNA fragment for
-actin
was purchased from Ambion. Radioactive signals were scanned and
quantitated by use of National Institutes of Health Image 1.61.
Statistical analysis. The data were compared using one-way ANOVA followed by Fisher's protected least significant difference test. The null hypothesis was rejected when P < 0.05. All values are expressed as means ± SE.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Effects of AG in LPS-induced hyperglycemia and iNOS
induction.
LPS administration led to hyperglycemia within 60 min. The maximal
hyperglycemic response was observed at 90 min after LPS injection. The
50 mg/kg dose of AG did not change blood glucose levels (data not
shown). However, 500 mg/kg of AG treatment abrogated the LPS-induced
hyperglycemia but did not affect blood glucose levels in LPS-naive
(saline-injected) rats (Fig.
1A). LPS increased the insulin
level. AG failed to modulate insulin levels in both saline- and
LPS-injected rats (Fig. 1B), suggesting that AG improved glycemic control in the LPS-injected rats through an effect on peripheral insulin sensitivity, and not by increasing insulin secretion
from pancreatic -cells.
|
|
Effects of LPS and AG on whole body insulin sensitivity.
A euglycemic hyperinsulinemic clamp study was performed to examine the
effects of AG (500 mg/kg) on LPS-induced insulin resistance. Steady-state glucose level was achieved during the 60- to 90-min period
after LPS injection. During steady-state glucose infusion, blood
glucose levels did not differ among the four groups. The average blood
glucose levels during the steady state of the animals treated with
saline alone (LPS naive), AG alone, LPS alone, and LPS+AG were
115.1 ± 5.4 (SE), 109.8 ± 3.8, 117.7 ± 4.3, and
122.1 ± 4.9 mg/dl, respectively. Whole body insulin sensitivity
was judged by the glucose infusion rate required to maintain
steady-state blood glucose levels between 60 and 90 min after LPS
injection. The glucose infusion rate to maintain steady-state glucose
level was significantly decreased in the rats administered LPS. AG
reversed significantly the LPS-induced decrease in the requirement of
glucose infusion rate. AG, when administered to LPS-naive
(saline-injected) rats, also decreased glucose infusion rate
significantly (Fig. 3).
|
Effects of AG on insulin-stimulated glucose uptake in tissues.
LPS significantly reduced tissue glucose uptake by skeletal muscle
during the euglycemic hyperinsulinemic clamp. AG did not affect
skeletal muscle glucose uptake in either saline- or LPS-injected rats
(Fig. 4). Glucose uptake by liver was
unaltered by LPS; the trend for decreased uptake by liver was not
statistically significant. AG failed to alter insulin-stimulated
glucose uptake by the liver. In contrast to liver and muscle, glucose
uptake in adipose tissue was relatively low in all groups, with no
differences observed between groups with and without AG.
|
Effects of AG on hepatic glucose output.
Hepatic glucose output was calculated as glucose appearance rate minus
the infusion rate. Hepatic glucose output was almost zero in
saline-treated (LPS-naive) rats, because a large infusion dose of
insulin suppressed hepatic glucose output completely. AG did not alter
hepatic glucose output in control animals (LPS naive). LPS
significantly increased hepatic glucose output. AG effectively
decreased LPS-induced hepatic glucose output to the level of controls
(Fig. 5). This, therefore, contrasts with
the ineffectiveness of AG on glucose uptake by liver and muscle in LPS-injected rats.
|
Effects of AG on glycogen phosphorylase activity and PEPCK mRNA
expression.
The data we have described suggest that the protective effects of AG on
LPS-induced hyperglycemia and whole body insulin resistance might be
accounted for mainly by inhibition of LPS-induced elevation of hepatic
glucose output. Augmented hepatic glucose output is attributable to
increased glycogenolysis and/or gluconeogenesis. Therefore, we examined
the effects of AG on glycogen phosphorylase and PEPCK, key enzymes for
glycogenolysis and gluconeogenesis, respectively. LPS upregulated both
total glycogen phosphorylase activity and glycogen phosphorylase-a
activity. AG restored both phosphorylase activities to the basal levels
(Fig. 6, A and B). In contrast, phosphorylase activities were unaltered by AG in saline-injected rats (controls). The RNase protection assay
demonstrated that LPS significantly increased PEPCK mRNA expression
levels. AG inhibited LPS-induced increase in PEPCK expression (Fig.
7, A and B) but did
not affect PEPCK mRNA level in saline-injected rats. The expression
levels of housekeeping gene -actin mRNA were unaltered by either LPS
or AG.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The detrimental effects of LPS on glucose metabolism observed in the present study confirm the previous reports of altered glucose homeostasis in septic patients and animals administered LPS. The adverse effects on glucose homeostasis induced by LPS include hyperglycemia (Fig. 1) (14, 21, 40), whole body insulin resistance, as judged by the glucose infusion rate during euglycemic hyperinsulinemic clamp (Fig. 3) (25, 55), attenuated insulin-stimulated glucose uptake in skeletal muscle (Fig. 4) (25, 53, 55), elevated hepatic glucose output (Fig. 5) (40), increased glycogen phosphorylase activities (Fig. 6) (26, 43), and increased PEPCK expression (Fig. 7) (7).
The salient new findings of the present study are that 1) the iNOS inhibitor AG decreased LPS-induced hyperglycemia and inhibited partially but significantly LPS-induced whole body insulin resistance (Figs. 1, 3); 2) AG completely blocked LPS-induced elevation in hepatic glucose output but did not alter insulin-stimulated glucose uptake in tissues (Figs. 4 and 5); 3) the LPS-mediated increases in glycogen phosphorylase activities (Fig. 6) and PEPCK mRNA expression (Fig. 7) were reversed by AG. The beneficial effects of AG were observed in LPS-injected rats in which iNOS was induced, but not in saline-injected rats where iNOS was undetectable (Fig. 1-7). On the basis of these data, one can conclude that iNOS plays a pivotal role in hyperglycemia and hepatic insulin resistance of endotoxin and that AG inhibits LPS-induced hyperglycemia and hepatic insulin resistance mainly by reversing the detrimental effects of LPS on hepatic glucose output. The abrogation of LPS-induced increases in glycogen phosphorylase activities and PEPCK expression probably contributed to the protective effects of AG. These findings are consistent with other reports that iNOS plays a key role in the deleterious host responses to endotoxin.
The protective effects of the specific iNOS inhibitor (Fig. 5) contrasts with an earlier study demonstrating that nonselective iNOS inhibition of all NO synthase isoenzymes (iNOS, endothelial NOS, and neuronal NOS) did not cause a statistically significant decrease in elevated hepatic glucose output in LPS-administered pigs (49). The nonspecific nature of the NOS inhibition may account for the lack of significant difference or effectiveness.
Although AG is known to be capable of inhibiting advanced glycation end product (AGE) formation as well as iNOS, the doses of AG needed to inhibit AGE formation and iNOS differ substantially. Relatively low doses (50 mg/kg body wt) of AG are sufficient to inhibit AGE formation when administered to rodents. In contrast, higher doses of AG (125-500 mg/kg body wt) are needed to inhibit LPS-induced iNOS activity (39). There is no report of increased AGE formation induced by LPS. Taken together, on the basis of our data that the low dose of AG (50 mg/kg body wt) was ineffective but that the higher dose of AG (500 mg/kg body wt) was required to inhibit LPS-induced hyperglycemia, AG is likely to exert its protective effects by inhibiting iNOS activity but not by blocking AGE formation. In addition, our preliminary experiment (data not presented) showed that another iNOS inhibitor, S-methylthiourea (SMT), also significantly decreased blood glucose levels in LPS (3 mg/kg iv)-injected rats, despite the fact that SMT does not inhibit AGE formation. [Blood glucose levels in rats treated with LPS+saline, LPS+SMT (5 mg/kg), and LPS+SMT (50 mg/kg) were 106.3 ± 14.3 (SE), 95.7 ± 9.7, and 56.3 ± 0.8 mg/dl, respectively; P < 0.01, LPS+saline vs. LPS+SMT (50 mg/kg)]. These data are supportive of a role for iNOS in LPS-induced derangements in glucose metabolism.
LPS impaired insulin-stimulated glucose uptake in skeletal muscle, the
major tissue for insulin-mediated glucose uptake (Fig. 4). LPS-induced
alterations in the insulin-stimulated glucose uptake were not observed,
however, in liver and adipose tissue. This selective impairment in
skeletal muscle is in accordance with the previous reports in septic
rats (20, 22) but not with another study showing that LPS
impaired insulin-stimulated glucose uptake not only in skeletal muscle
but also in liver and adipose tissue (25). This apparent
discrepancy may be explained by the differences in the experimental
design, such as insulin infusion rates, the body weight of animals, and
the doses of endotoxin. AG was ineffective in reversing the attenuated
insulin-stimulated glucose uptake by skeletal muscle in LPS-injected
rats (Fig. 4). Therefore, LPS seems to exert its adverse effects on
insulin-stimulated glucose uptake in skeletal muscle by an
iNOS-independent mechanism. LPS-induced insulin resistance in skeletal
muscle is postulated to be mediated by a -adrenergic mechanism, a
conclusion that was based on the protective effects of the
-adrenergic antagonist propranolol (20).
AG administration resulted in a modest but significant decrease in glucose infusion rate during the hyperinsulinemic euglycemic clamp in control (LPS-naive) rats, whereas it induced a significant increase in glucose infusion rate in LPS-injected rats (Fig. 3). The biological significance and mechanism for the decreased glucose infusion rate in controls during AG therapy are unclear. However, the AG-induced decrease in glucose infusion rate in controls is unlikely to be attributable to the effects of iNOS, because iNOS was not induced (Fig. 2); neither glucose uptake in skeletal muscle, liver, and adipose tissue nor hepatic glucose output was affected by AG in control rats (Figs. 4 and 5).
Increased hepatic glucose output is responsible for fasting
hyperglycemia (48). Because AG did not alter insulin
levels, one can conclude that the normoglycemic action of AG is not via pancreatic -cells to increase insulin secretion but a peripheral action on target tissues. The attenuation of elevated hepatic glucose
output by the iNOS inhibitor accounts for its potential normoglycemic
effects after a hyperglycemic response to LPS. In contrast, the
relatively modest amelioration of glucose infusion rate by AG in
LPS-treated animals during euglycemic hyperinsulinemic clamp may be
explained by the failure to mitigate the impaired glucose uptake in
skeletal muscle. When these observations are taken together, therefore,
one can conclude that the mechanisms underlying LPS-induced insulin
resistance differ between tissues, and that iNOS-dependent and
iNOS-independent mechanisms may play major roles in the derangement in
insulin actions in liver and skeletal muscle of LPS-injected rats, respectively.
Glycogenolysis and gluconeogenesis are the two major biochemical pathways for hepatic glucose production. Glycogen phosphorylase is a key enzyme responsible for glycogenolysis. Glycogenolysis is increased by LPS (5, 28). A recent study (29) demonstrating that the inhibition of glycogen phosphorylase lowered hyperglycemia of genetically obese, diabetic (ob/ob) mice emphasizes the important role of glycogen phosphorylase in hyperglycemia in diabetes mellitus. In our study, the parallel relationship between induction of glycogen phosphorylase activity and hyperglycemia and the normalization of the enzyme activity with AG and attenuation of hyperglycemia suggest a role for glycogen phosphorylase activity in the hyperglycemic response to LPS. However, little is known about how glycogenolysis and glycogen phosphorylase activity are upregulated by endotoxin.
Glycogen phosphorylase, when phosphorylated by cAMP-dependent glycogen phosphorylase kinase, becomes activated and is called "glycogen phosphorylase-a"; the dephosphorylated form is inactive and referred to as "glycogen phosphorylase-b." Glycogen phosphorylase-b (inactive form) is subject to allosteric activation with AMP, and in the presence of AMP, glycogen phosphorylase-b is as active as glycogen phosphorylase-a. Thus the activities measured in the presence and absence of AMP represent those of total (a + b) glycogen phosphorylase and glycogen phosphorylase-a alone, respectively. The present study reveals that LPS increased the activities of both total and glycogen phosphorylase-a without affecting the ratio between the total enzyme and the phosphorylated fraction of the enzyme (Fig. 6). These results indicate that the activation of glycogen phosphorylase by LPS is not mediated by glycogen phosphorylase kinase or the phosphorylation status of the enzyme.
It is noteworthy that -adrenergic agonists activate and insulin
inactivates glycogen phosphorylase by altering phosphorylation status,
namely the ratio of phosphorylase-a to total phosphorylase (a+b)
[a/total(a+b)] (34, 52). In this study, LPS and AG
modulated the enzyme activities, with no change in a/total(a+b) (Fig.
6C). This finding is consistent with the assumption that
LPS-induced hepatic insulin resistance is
-adrenergic independent.
It is also suggested that phosphorylase activation might be one of the primary events of hepatic insulin resistance, rather than just a
consequence secondary to insulin resistance, considering that phosphorylase activation resulting from insulin resistance, namely, decreased insulin action, would be accompanied by an increase in
a/total(a+b). Combined with a previous report showing that an NO donor
increased glycogen phosphorylase activity and glycogenolysis (3,
44, 52), the protective effects of AG on LPS-induced glycogen
phosphorylase activation in the present study indicate that LPS
upregulated glycogen phosphorylase activity by inducing iNOS.
Gluconeogenesis is upregulated in sepsis in the fasting condition (9). PEPCK, a rate-limiting enzyme for gluconeogenesis, is known to be induced by LPS and sepsis (7). The activities of PEPCK are considered to be regulated by its expression levels, and increased expression of PEPCK has been assumed to play a critical role in elevated hepatic glucose production in sepsis and endotoxicosis, as well as type 2 diabetes. Our results of the inhibitory effects of AG on PEPCK expression (Fig. 7) suggest that iNOS may also be involved in PEPCK induction by LPS.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Drs. J. Avruch and K. Ueki for their helpful discussion.
![]() |
FOOTNOTES |
---|
This work was supported by grants from the National Institute of General Medical Sciences (R01 GM-31569, GM-61411, and GM-55082) and Shriners Hospital for Children.
Address for reprint requests and other correspondence: J. A. J. Martyn (E-mail: jmartyn{at}etherdome.mgh.harvard.edu) and M. Kaneki (E-mail: mkaneki{at}helix.mgh.harvard.edu), Dept. of Anesthesia and Critical Care, Massachusetts General Hospital, 55 Fruit St., Boston, MA 02114.
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.
10.1152/ajpendo.00087.2001
Received 2 March 2001; accepted in final form 11 October 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Afulukwe, IF,
Cohen RI,
Zeballos GA,
Iqbal M,
and
Scharf SM.
Selective NOS inhibition restores myocardial contractility in endotoxemic rats; however, myocardial NO content does not correlate with myocardial dysfunction.
Am J Respir Crit Care Med
162:
21-26,
2000
2.
Bedard, S,
Marcotte B,
and
Marette A.
Cytokines modulate glucose transport in skeletal muscle by inducing the expression of inducible nitric oxide synthase.
Biochem J
325:
487-493,
1997[ISI][Medline].
3.
Borgs, M,
Bollen M,
Keppens S,
Yap SH,
Stalmans W,
and
Vanstapel F.
Modulation of basal hepatic glycogenolysis by nitric oxide.
Hepatology
23:
1564-1571,
1996[ISI][Medline].
4.
Bredt, DS,
and
Snyder SH.
Isolation of nitric oxide synthetase, a calmodulin-requiring enzyme.
Proc Natl Acad Sci USA
87:
682-685,
1990[Abstract].
5.
Buday, AZ,
Lang CH,
Bagby GJ,
and
Spitzer JJ.
Glycogen synthase and phosphorylase activities during glycogen repletion in endotoxemic rats.
Circ Shock
19:
149-163,
1986[ISI][Medline].
6.
Burgner, D,
Rockett K,
and
Kwiatkowski D.
Nitric oxide and infectious diseases.
Arch Dis Child
81:
185-188,
1999
7.
Chang, CK,
Moskal SF, II,
Srivenugopal KS,
and
Schumer W.
Altered levels of mRNA encoding enzymes of hepatic glucose metabolism in septic rats.
Circ Shock
41:
35-39,
1993[ISI][Medline].
8.
Choo, JJ,
Horan MA,
Little RA,
and
Rothwell NJ.
Muscle wasting associated with endotoxemia in the rat: modification by the beta 2-adrenoceptor agonist clenbuterol.
Biosci Rep
9:
615-621,
1989[ISI][Medline].
9.
Deaciuc, IV,
Lang CH,
Bagby GJ,
and
Spitzer JJ.
Effect of subcutaneous Escherichia coli-induced hypermetabolic sepsis on hepatic gluconeogenesis and its hormonal responsiveness in the rat.
Circ Shock
41:
82-87,
1993[ISI][Medline].
10.
Dickinson, E,
Tuncer R,
Nadler E,
Boyle P,
Alber S,
Watkins S,
and
Ford H.
NOX, a novel nitric oxide scavenger, reduces bacterial translocation in rats after endotoxin challenge.
Am J Physiol Gastrointest Liver Physiol
277:
G1281-G1287,
1999
11.
Franch, J,
Aslesen R,
and
Jensen J.
Regulation of glycogen synthesis in rat skeletal muscle after glycogen-depleting contractile activity: effects of adrenaline on glycogen synthesis and activation of glycogen synthase and glycogen phosphorylase.
Biochem J
344:
231-235,
1999[ISI][Medline].
12.
Fujii, E,
Yoshioka T,
Ishida H,
Irie K,
and
Muraki T.
Evaluation of iNOS-dependent and independent mechanisms of the microvascular permeability change induced by lipopolysaccharide.
Br J Pharmacol
130:
90-94,
2000
13.
Gunnett, CA,
Chu Y,
Heistad DD,
Loihl A,
and
Faraci FM.
Vascular effects of LPS in mice deficient in expression of the gene for inducible nitric oxide synthase.
Am J Physiol Heart Circ Physiol
275:
H416-H421,
1998
14.
Hargrove, DM,
Bagby GJ,
Lang CH,
and
Spitzer JJ.
Adrenergic blockade does not abolish elevated glucose turnover during bacterial infection.
Am J Physiol Endocrinol Metab
254:
E16-E22,
1988
15.
Hawkins, M,
Hu M,
Yu J,
Eder H,
Vuguin P,
She L,
Barzilai N,
Leiser M,
Backer JM,
and
Rossetti L.
Discordant effects of glucosamine on insulin-stimulated glucose metabolism and phosphatidylinositol 3-kinase activity.
J Biol Chem
274:
31312-31319,
1999
16.
Kaneki, M,
Kharbanda S,
Pandey P,
Yoshida K,
Takekawa M,
Liou JR,
Stone R,
and
Kufe D.
Functional role for protein kinase Cbeta as a regulator of stress-activated protein kinase activation and monocytic differentiation of myeloid leukemia cells.
Mol Cell Biol
19:
461-470,
1999
17.
Kapur, S,
Bedard S,
Marcotte B,
Cote CH,
and
Marette A.
Expression of nitric oxide synthase in skeletal muscle: a novel role for nitric oxide as a modulator of insulin action.
Diabetes
46:
1691-1700,
1997[Abstract].
18.
Kurrek, MM,
Zapol WM,
Holzmann A,
Filippov G,
Winkler M,
and
Bloch KD.
In vivo lipopolysaccharide pretreatment inhibits cGMP release from the isolated-perfused rat lung.
Am J Physiol Lung Cell Mol Physiol
269:
L618-L624,
1995
19.
Kwon, G,
Xu G,
Marshall CA,
and
McDaniel ML.
Tumor necrosis factor alpha-induced pancreatic beta-cell insulin resistance is mediated by nitric oxide and prevented by 15-deoxy-Delta12,14-prostaglandin J2 and aminoguanidine. A role for peroxisome proliferator-activated receptor gamma activation and inos expression.
J Biol Chem
274:
18702-18708,
1999
20.
Lang, CH.
Sepsis-induced insulin resistance in rats is mediated by a beta-adrenergic mechanism.
Am J Physiol Endocrinol Metab
263:
E703-E711,
1992
21.
Lang, CH,
Bagby GJ,
Blakesley HL,
and
Spitzer JJ.
Fever is not responsible for the elevated glucose kinetics in sepsis.
Proc Soc Exp Biol Med
185:
455-461,
1987[Abstract].
22.
Lang, CH,
Dobrescu C,
and
Meszaros K.
Insulin-mediated glucose uptake by individual tissues during sepsis.
Metabolism
39:
1096-1107,
1990[ISI][Medline].
23.
Lang, CH,
Frost RA,
Jefferson LS,
Kimball SR,
and
Vary TC.
Endotoxin-induced decrease in muscle protein synthesis is associated with changes in eIF2B, eIF4E, and IGF-I.
Am J Physiol Endocrinol Metab
278:
E1133-E1143,
2000
24.
Liaudet, L,
Rosselet A,
Schaller MD,
Markert M,
Perret C,
and
Feihl F.
Nonselective versus selective inhibition of inducible nitric oxide synthase in experimental endotoxic shock.
J Infect Dis
177:
127-132,
1998[ISI][Medline].
25.
Ling, PR,
Bistrian BR,
Mendez B,
and
Istfan NW.
Effects of systemic infusions of endotoxin, tumor necrosis factor, and interleukin-1 on glucose metabolism in the rat: relationship to endogenous glucose production and peripheral tissue glucose uptake.
Metabolism
43:
279-284,
1994[ISI][Medline].
26.
Liu, MS,
and
Kang GF.
Liver glycogen metabolism in endotoxin shock. II. Endotoxin administration increases glycogen phosphorylase activities in dog livers.
Biochem Med Metab Biol
37:
73-80,
1987[ISI][Medline].
27.
MacMicking, JD,
Nathan C,
Hom G,
Chartrain N,
Fletcher DS,
Trumbauer M,
Stevens K,
Xie QW,
Sokol K,
and
Hutchinson N.
Altered responses to bacterial infection and endotoxic shock in mice lacking inducible nitric oxide synthase.
Cell
81:
641-650,
1995[ISI][Medline].
28.
Mandl, J,
Wall C,
Lerant I,
Falus A,
Machovich R,
and
Thurman RG.
Endotoxin and fibrinogen degradation product-D have different actions on carbohydrate metabolism: role of Kupffer cells.
FEBS Lett
376:
65-66,
1995[ISI][Medline].
29.
Martin, WH,
Hoover DJ,
Armento SJ,
Stock IA,
McPherson RK,
Danley DE,
Stevenson RW,
Barrett EJ,
and
Treadway JL.
Discovery of a human liver glycogen phosphorylase inhibitor that lowers blood glucose in vivo.
Proc Natl Acad Sci USA
95:
1776-1781,
1998
30.
McGowder, D,
Ragoobirsingh D,
and
Dasgupta T.
The hyperglycemic effect of S-nitrosoglutathione in the dog.
Nitric Oxide
3:
481-491,
1999[ISI][Medline].
31.
Meneilly, GS,
Battistini B,
and
Floras JS.
Lack of effect of sodium nitroprusside on insulin-mediated blood flow and glucose disposal in the elderly.
Metabolism
49:
373-378,
2000[ISI][Medline].
32.
Mishima, S,
Xu D,
Lu Q,
and
Deitch EA.
Bacterial translocation is inhibited in inducible nitric oxide synthase knockout mice after endotoxin challenge but not in a model of bacterial overgrowth.
Arch Surg
132:
1190-1195,
1997[Abstract].
33.
Paulsen, SM,
Wurster SH,
and
Nanney LB.
Expression of inducible nitric oxide synthase in human burn wounds.
Wound Repair Regen
6:
142-148,
1998[Medline].
34.
Pickett-Gies, CA,
Carlsen RC,
Anderson LJ,
Angelos KL,
and
Walsh DA.
Characterization of the isolated rat flexor digitorum brevis for the study of skeletal muscle phosphorylase kinase phosphorylation.
J Biol Chem
262:
3227-3238,
1987
35.
Piercy, V,
Toseland CD,
and
Turner NC.
Potential benefit of inhibitors of advanced glycation end products in the progression of type II diabetes: a study with aminoguanidine in C57/BLKsJ diabetic mice.
Metabolism
47:
1477-1480,
1998[ISI][Medline].
36.
Rafalowska, J.
HIV-1-infection in the CNS. A pathogenesis of some neurological syndromes in the light of recent investigations.
Folia Neuropathol
36:
211-216,
1998[ISI][Medline].
37.
Ribiere, C,
Jaubert AM,
Gaudiot N,
Sabourault D,
Marcus ML,
Boucher JL,
Denis-Henriot D,
and
Giudicelli Y.
White adipose tissue nitric oxide synthase: a potential source for NO production.
Biochem Biophys Res Commun
222:
706-712,
1996[ISI][Medline].
38.
Rossetti, L,
Stenbit AE,
Chen W,
Hu M,
Barzilai N,
Katz EB,
and
Charron MJ.
Peripheral but not hepatic insulin resistance in mice with one disrupted allele of the glucose transporter type 4 (GLUT4) gene.
J Clin Invest
100:
1831-1839,
1997
39.
Roth, J,
Storr B,
Goldbach J,
Voigt K,
and
Zeisberger E.
Dose-dependent attenuation of lipopolysaccharide-fever by inhibitors of inducible nitric oxide-synthase in guinea pigs.
Eur J Pharmacol
383:
177-187,
1999[ISI][Medline].
40.
Santak, B,
Radermacher P,
Adler J,
Iber T,
Rieger KM,
Wachter U,
Vogt J,
Georgieff M,
and
Trager K.
Effect of increased cardiac output on liver blood flow, oxygen exchange and metabolic rate during longterm endotoxin-induced shock in pigs.
Br J Pharmacol
124:
1689-1697,
1998[Abstract].
41.
Schaffer, MR,
Tantry U,
Thornton FJ,
and
Barbul A.
Inhibition of nitric oxide synthesis in wounds: pharmacology and effect on accumulation of collagen in wounds in mice.
Eur J Surg
165:
262-267,
1999[ISI][Medline].
42.
Shimabukuro, M,
Ohneda M,
Lee Y,
and
Unger RH.
Role of nitric oxide in obesity-induced beta cell disease.
J Clin Invest
100:
290-295,
1997
43.
Spitzer, JJ,
Bagby GJ,
Meszaros K,
and
Lang CH.
Altered control of carbohydrate metabolism in endotoxemia.
Prog Clin Biol Res
286:
145-165,
1989[Medline].
44.
Sprangers, F,
Sauerwein HP,
Romijn JA,
van Woerkom GM,
and
Meijer AJ.
Nitric oxide inhibits glycogen synthesis in isolated rat hepatocytes.
Biochem J
330:
1045-1049,
1998[ISI][Medline].
45.
Stalmans, W,
and
Hers HG.
The stimulation of liver phosphorylase b by AMP, fluoride and sulfate. A technical note on the specific determination of the a and b forms of liver glycogen phosphorylase.
Eur J Biochem
54:
341-350,
1975[Abstract].
46.
Symeonides, S,
and
Balk RA.
Nitric oxide in the pathogenesis of sepsis.
Infect Dis Clin North Am
13:
449-463,
1999[ISI][Medline].
47.
Tanaka, Y,
Gleason CE,
Tran PO,
Harmon JS,
and
Robertson RP.
Prevention of glucose toxicity in HIT-T15 cells and Zucker diabetic fatty rats by antioxidants.
Proc Natl Acad Sci USA
96:
10857-10862,
1999
48.
Taylor, SI.
Deconstructing type 2 diabetes.
Cell
97:
9-12,
1999[ISI][Medline].
49.
Trager, K,
Radermacher P,
Rieger KM,
Vlatten A,
Vogt J,
Iber T,
Adler J,
Wachter U,
Grover R,
Georgieff M,
and
Santak B.
Norepinephrine and N-monomethyl-L-arginine in porcine septic shock: effects on hepatic O2 exchange and energy balance.
Am J Respir Crit Care Med
159:
1758-1765,
1999
50.
Tsuji, C,
Shioya S,
Hirota Y,
Fukuyama N,
Kurita D,
Tanigaki T,
Ohta Y,
and
Nakazawa H.
Increased production of nitrotyrosine in lung tissue of rats with radiation-induced acute lung injury.
Am J Physiol Lung Cell Mol Physiol
278:
L719-L725,
2000
51.
Tunctan, B,
Uludag O,
Altug S,
and
Abacioglu N.
Effects of nitric oxide synthase inhibition in lipopolysaccharide-induced sepsis in mice.
Pharmacol Res
38:
405-411,
1998[ISI][Medline].
52.
Villar-Palasi, C.
On the mechanism of inactivation of muscle glycogen phosphorylase by insulin.
Biochim Biophys Acta
1224:
384-388,
1994[ISI][Medline].
53.
Virkamaki, A,
Puhakainen I,
Koivisto VA,
Vuorinen-Markkola H,
and
Yki-Jarvinen H.
Mechanisms of hepatic and peripheral insulin resistance during acute infections in humans.
J Clin Endocrinol Metab
74:
673-679,
1992[Abstract].
54.
Virkamaki, A,
Rissanen E,
Hamalainen S,
Utriainen T,
and
Yki-Jarvinen H.
Incorporation of [3-3H]glucose and 2-[1-14C]deoxyglucose into glycogen in heart and skeletal muscle in vivo: implications for the quantitation of tissue glucose uptake.
Diabetes
46:
1106-1110,
1997[Abstract].
55.
Virkamaki, A,
and
Yki-Jarvinen H.
Role of prostaglandins in mediating alterations in glucose metabolism during acute endotoxemia in the rat.
Endocrinology
136:
1701-1706,
1995[Abstract].
56.
Wang, JL,
Chinookoswong N,
Scully S,
Qi M,
and
Shi ZQ.
Differential effects of leptin in regulation of tissue glucose utilization in vivo.
Endocrinology
140:
2117-2124,
1999
57.
Weimann, J,
Bloch KD,
Takata M,
Steudel W,
and
Zapol WM.
Congenital NOS2 deficiency protects mice from LPS-induced hyporesponsiveness to inhaled nitric oxide.
Anesthesiology
91:
1744-1753,
1999[ISI][Medline].
58.
Wildhirt, SM,
Weismueller S,
Schulze C,
Conrad N,
Kornberg A,
and
Reichart B.
Inducible nitric oxide synthase activation after ischemia/reperfusion contributes to myocardial dysfunction and extent of infarct size in rabbits: evidence for a late phase of nitric oxide-mediated reperfusion injury.
Cardiovasc Res
43:
698-711,
1999[ISI][Medline].
59.
Wong, JM,
and
Billiar TR.
Regulation and function of inducible nitric oxide synthase during sepsis and acute inflammation.
Adv Pharmacol
34:
155-170,
1995[Medline].