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Cardiovascular Research 2003 60(1):119-130; doi:10.1016/S0008-6363(03)00320-1
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Copyright © 2003, European Society of Cardiology

Impairment of glucose metabolism in hearts from rats treated with endotoxin

Jean-Philippe Tessiera, Bernhard Thurnera, Eberhard Jünglinga, Andreas Lückhoffa and Yvan Fischera,b,*

aInstitute of Physiology, Medical Faculty, RWTH Aachen, Pauwelsstrasse 30, D-52057 Aachen, Germany
bSolvay Pharmaceuticals, Hans-Böckler-Allee 20, D-30173 Hannover, Germany

*Corresponding author. Solvay Pharmaceuticals, Hans-Böckler-Allee 20, D-30173 Hannover, Germany. Tel.: +49-511-857-2738; fax: +49-511-857-3269. Email address: yvan.fischer{at}solvay.com

Received 28 October 2002; accepted 13 February 2003


   Abstract
Top
 Abstract
1. Introduction
 2. Methods
 3. Results
 4. Discussion
 5. Concluding remarks
 References
 
Objective: In patients and animals with sepsis or critical illness, the mechanical function of the heart is often impaired. Although these conditions are accompanied by dramatic metabolic and hormonal changes, little is known about alterations of cardiac metabolism. In this study, we assessed the impact of an endotoxin-induced inflammation on cardiac glucose utilization. Methods: Bacterial endotoxin (1 mg/kg lipopolysaccharide from Salmonella typhimurium, LPS) was injected intravenously to rats. Six hours after LPS application, hearts were isolated and perfused in the Langendorff mode. Results: Left ventricular pressure was reduced by 50% in hearts from LPS-treated rats, compared to those from saline-injected control animals. With glucose as the sole fuel, there was no difference in glycolysis between the groups. However, on addition of β-hydroxybutyrate (an alternative fuel which inhibits phosphofructokinase via an increased citrate level), the glycolytic rate in the LPS group was 44 and 48% lower (in basal, and insulin-stimulated conditions, respectively; P<0.01) than in control hearts. At the end of perfusions with β-hydroxybutyrate and insulin, the cardiac citrate content was 40% higher in LPS vs. controls (P<0.001). In addition to the reduced glycolysis, the insulin-dependent increase of cardiac glycogen was 77% smaller in LPS hearts. The difference between LPS and control glycolysis was abolished if the hearts were perfused with the ceramidase inhibitor N-oleyl-ethanolamine (5 µM), and also with the cyclooxygenase-2 inhibitor NS-398 (10 µM), or the thromboxane A2 receptor antagonist SQ-29548 (1 µM). Conclusion: The inflammatory reaction caused by endotoxin impairs cardiac glucose metabolism (and in particular, the action of insulin) in at least two ways: through the exacerbation of the counterregulatory effect of alternative fuels on glycolysis, and through a reduction in net glycogen synthesis. Impairment of glycolysis may be mediated by a sphingomyelin derivative, and COX-2-derived thromboxane A2.

KEYWORDS Endotoxins; Glycolysis; Infection/inflammation; Insulin resistance; Ventricular function

Abbreviations: TNF{alpha}, tumor necrosis factor {alpha} • IL-1β, interleukin-1β • L-NAME, L-N-arginine methyl ester • NOE, N-oleyl-ethanolamine • NO, nitric oxide • LPS, lipopolysaccharide • LVDP, left ventricular developed pressure • SNAP, S-nitroso-N-acetylpenicillamine


   1. Introduction
Top
 Abstract
 1. Introduction
2. Methods
 3. Results
 4. Discussion
 5. Concluding remarks
 References
 
In patients with sepsis, injury, or critical illness, profound and complex hormonal and metabolic changes are commonly observed which, on the whole, bring about an accelerated mobilization of protein, fat, and carbohydrates (for reviews see, e.g., Refs. [1–4]). A hallmark of this hypermetabolic state is the fast appearance of a pronounced insulin resistance, especially with regard to carbohydrate metabolism [1–7]. This insulin resistance affects the liver [4,7,8], as well as skeletal muscles [5,8–10], and the heart (see below).

Systemic (as well as intracardial) inflammation, as it occurs in the above conditions, can be associated with an impairment of the heart's mechanical function [11,12]. This dysfunction probably involves inflammatory cytokines such as TNF{alpha} or IL-1β, which may act via several mechanisms, including alterations in the cardiomyocytes’ calcium homeostasis [13,14], although the relevance of these mechanisms for the sepsis- or trauma-associated heart failure is still unclear.

Despite the fact that the utilization of fuels by the heart is one of the determinants of its contractile capacity – which becomes apparent especially in pathophysiological conditions, such as ischemia or diabetes [15–18] – relatively little is known about metabolic alterations occurring in this organ in the course of local or systemic inflammatory responses. Observations in rats [5,6,19], and dogs [20] have indicated that following bacterial infections or endotoxemia the heart becomes resistant to the stimulating action of insulin on glucose utilization. The mediators and mechanisms underlying the cardiac insulin resistance related to inflammatory conditions is (are) unknown. In addition, in the above studies, only the overall uptake, but not the intracellular metabolism of glucose was assessed; it is hence not known whether only the uptake of the sugar (i.e., its transport across the cardiomyocyte plasma membrane and its hexokinase-catalyzed phosphorylation), or glycolysis, and/or glycogen synthesis are affected.

Investigations in other insulin-sensitive cells and tissues like adipocytes and skeletal muscles have shown that inflammatory mediators such as TNF{alpha} can depress the insulin-dependent uptake of glucose [21–24]. Although no comparable data are so far available for the heart, our own observations suggest that at least the stimulatory effect of insulin on the transmembrane transport of the sugar is impaired by TNF{alpha} and IL-1β in isolated cardiac myocytes (Thurner et al., manuscript in preparation). Furthermore, recent studies have revealed that nitric oxide (NO) – which is produced in the heart in at least in some models of local or systemic inflammation [11,25] – and in particular its second messenger cyclic GMP, exert an inhibitory action on glucose uptake and glycolysis in hearts [26–29] and isolated cardiac myocytes [30].

We therefore examined the influence of a systemic inflammation, induced by a bacterial endotoxin in rats, on the cardiac utilization of glucose. Our results show that in the presence of an alternative substrate of energy metabolism (β-hydroxybutyric acid), cardiac glycolysis and glycogen synthesis are impaired by this treatment.


   2. Methods
Top
 Abstract
 1. Introduction
 2. Methods
3. Results
 4. Discussion
 5. Concluding remarks
 References
 
2.1. Chemicals and reagents
Lipopolysaccharides (LPS) from Salmonella typhimurium (batch 39H4110), N-[2-(cyclohexyloxy)-4-nitrophenyl] methanesulfonamide (NS-398), and β-hydroxybutyric acid were purchased from Sigma (Munich, Germany). All chemicals for perfusion media were from Merck (Darmstadt, Germany). All enzymes, NADH, and kaolin-activated reagent were from Boehringer (Mannheim, Germany). 2-[3H]D-glucose and 3-[3H]D-glucose were obtained from Amersham (Little Chalfont, UK). Indomethacin, isoprenaline hydrochloride, sodium nitroprusside dihydrate (SNP), N-oleyl ethanolamine (NOE), S-nitroso-N-acetylpenicillamine (SNAP), and NG-nitro-L-arginine methyl ester (L-NAME) were from Calbiochem (Notthingham, UK). Bovine insulin was a generous gift from Professor Joachim Grötzinger (Institute of Biochemistry, Aachen, Germany).

2.2. Animals and treatments
The investigation was performed in accordance with the Guide for the Care and Use of Laboratory Animals published by the US National Institute of Health (NIH publication No. 85, 23, revised 1996). The animals were acquired and used in compliance with Section 6 of the German Animal Protection Law, and the study was reviewed and approved by the appropriate authority (Bezirksregierung Cologne).

Male Sprague–Dawley rats (300 to 400 g, Charles River) were acclimated at our animal facilities for 7 days at constant temperature (20±2°C), exposed to a 12 h light–12 h dark cycle (light from 07:00) and fed ad libitum on a chow diet comprising approximately 4.9% carbohydrate, 19% protein and 3.3% fat. Animals had free access to drinking water.

Animals were assigned to either the control (saline) or to the LPS (endotoxin) group. They were anesthetized by ether inhalation (2 ml ether in a 2 l chamber) for 5 min before receiving an intravenous injection (via the caudal vein) of either 0.35 ml sterile isotonic saline (control), or endotoxin (lipopolysaccharide from S. typhimurium 1 mg/kg body weight). Rectal temperature was measured with an electronic thermometer to monitor the pyrogenic effect of LPS.

2.3. Isolated heart preparation
Six hours after the LPS injection the rats were anesthetized in ether atmosphere (0.2% v/v, 5 min). After thoracotomy, the vena cava inferior was severed and the diaphragm opened. The thymus was removed; the aorta ascendens, the pulmonary arteries and venae were cut. Hearts were rapidly excised and arrested in ice-cold (4°C) Krebs–Henselheit bicarbonate buffered solution (pH 7.4) with the following composition (mM): NaCl 128.5; KCl 4.7; KH2PO4, 1.2; MgSO4, 1.2; NaHCO3, 25; glucose, 5.6; EDTA, 0.5; CaCl2, 2.5. Non-cardiac (e.g., connective) tissue was removed, and the hearts were cannulated via the ascending aorta for retrograde perfusion in the Langendorff mode [31], using the same buffer as above, kept at 37±0.5°C and continuously gassed with 95% O2–5% CO2. The total preparation time until reperfusion was kept under 2 to 3 min to avoid prolonged ischemia. Hearts were suspended in a temperature-controlled chamber maintained at 37±0.5°C. All experiments were carried out with a perfusion pressure of 62 mmHg (84 cm H2O).

2.4. Assessment of cardiac function
Cardiac function was determined as described previously [32] using a water-filled latex balloon (volume: 0.15 ml) connected via polyethylene tubing (cavafix duo 16/18G, Braun, Germany) to a disposable pressure transducer (Model I-93-II-096, PVB Medizintechnik, Kirchseeon, Germany). The left ventricular developed pressure (LVDP) was measured isovolumetrically. LVDP, heart rate, rate of contraction (+dP/dt), and rate of relaxation (–dP/dt) were continuously recorded using a multichannel recorder. The coronary outflow was measured at regular intervals (usually 4 min). All these functional parameters were recorded throughout the perfusion period.

2.5. Validation of inflammation model
In a first series of experiments, a previously described protocol of endotoxin treatment was used to induce a generalised inflammatory reaction, coupled with a cardiac depression in rats (intraperitoneal administration of 0.5 mg/kg lipopolysaccharide from S. typhimurium [33]). However, in our hands, the occurrence and degree of contractile depression observed ex vivo (i.e., in perfused hearts isolated 6 h after the LPS injection) turned out to be variable. We therefore tried out several modifications to this protocol and found that intravenous application of LPS from S. typhimurium (1 mg/kg into the caudal vein) caused a reproducible reduction in LVDP, as measured in perfused hearts isolated 6 h after the endotoxin injection.

Thus, LVPD was diminished by ~50% when compared to values measured in time-matched control hearts from animals that had received a saline injection (Table 1). In addition, the coronary flow was lower than in controls, while the heart rate remained unchanged (Table 1). The diminished coronary flow is likely to be the consequence rather than the cause of the mechanical dysfunction because perfusion with the nitric oxide donor SNAP (50 µM) increased coronary flow by 64% without improving ventricular contractile function (not shown). The mechanical dysfunction seen ex vivo shows that this cardiac effect of LPS treatment is stable and not (or no longer) dependent on extracardial factors.


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  		Table 1 Effect of LPS treatment in vivo on functional parameters of perfused hearts ex vivo

 
As documented in Table 2, this protocol also resulted in a reproducible rise in body temperature, as well as in hematological, endocrine and metabolic changes known to occur in the course of systemic inflammatory responses.


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  		Table 2 Effect of LPS treatment on hematological, metabolic and hormonal parameters

 
2.6. Perfusion protocol and measurement of glucose uptake and glycolysis
After an equilibration period of retrograde perfusion (10 min) to wash out any blood and to reach a functional steady state, the hearts were perfused with a recirculating Krebs–Henselheit bicarbonate buffer (100 ml) containing glucose (5.6 mM) as substrate. The recirculating perfusion was started at time 0 min. Inhibitors or antagonists, glucose tracer (see below), β-hydroxybutyric acid (5 mM), and insulin (17 nM) were injected into the closed system through a milliliter syringe at the following times: 8, 10, 12, and 36 min, respectively as indicated in the figure legends. The total duration of the experiments (including the initial non-recirculating phase) was 58 min.

Glucose uptake and glycolysis were measured as previously described [27,34,35]. Briefly, the rate of 3H2O formation from 2-[3H]D-glucose, or 3-[3H]D-glucose was used to calculate glucose uptake, or glycolysis, respectively. The tracer was added to the perfusion medium (final concentration 2 µM, specific activity 2 µCi/100 ml perfusate, injection volume 500 µl) at min 10 of the recirculating perfusion. Aliquots of 1.2 ml coronary effluate were collected in plastic tubes before (i.e., at 19, 24, 29, and 34 min) and after insulin (at 37, 41, 45, and 49 min). These radioactive samples were immediately mixed with 120 µl of HClO4 (70%). After centrifugation, 1 ml of supernatant was neutralized with 400 µl KOH (3 M) and 200 µl KHCO3 (1 M). The resulting precipitate was spun down, and 1 ml of supernatant was passed over columns filled with 4 ml Dowex AG1-X8 resin, borate form, 200–400 mesh (Bio-Rad, Munich, Germany), and the tritiated water present in the eluate was counted by liquid scintillation.

2.7. Quantification of cardiac metabolites
Glycogen was measured as previously reported [36,37]. At the times indicated in the figure legends, the hearts were freeze-clamped in liquid nitrogen and stored at –80°C before they were homogenized and processed as described [36,37]. Heart citrate content was assessed using the method described by Dagley [38] with minor modifications). Briefly, 500 mg of heart tissue was mixed with 3 ml ice-cold HClO4, sonicated, centrifuged and neutralized with KOH (6 M) and filtered. A 0.1-ml volume of neutralized sample was added to 1 ml ZnCl2–Tra (tri-ethanolamine) buffer (0.1 M Tra, pH 7.6; 0.2 mM ZnCl2) and 10 µl NADH solution (10 mM). After reading the optical extinction E1 at 340 nm, 20 µl of a lactate dehydrogenase (30 U)/malate dehydrogenase (50 U) mixture were added; 3 min later 20 µl citrate lyase (2.4 U) before the final extinction E2 was measured. The citrate content was calculated from E2–E1, and expressed as µmol per g of wet tissue.

2.8. Hematological measurements
All blood samples were taken from the vena cava inferior during the heart preparations. All hematological parameters (except insulin levels) were kindly performed by Dr. D. Kunz and Professor A. Gressner (Institute of Clinical Chemistry and Pathobiochemistry, Aachen, Germany). Blood cell numbers were determined with an electronic Celltek counter (Bayer Diagnostics, Munich, Germany). Plasma lactate was measured enzymatically (in samples collected in standard S-Monovette Ref. 05.1073 containing NaF and EDTA, and centrifuged at 600 g for 10 min) using a lactate oxidase (LOD) and peroxidase (POD) assay (Roche, Mannheim, Germany). Non-esterified fatty acids (NEFAs) concentrations in serum were evaluated with an enzymatic test-combination (Roche). The serum cortisol concentration was measured with the ADVIA Centaur test (competitive immunoassay) in blood samples collected in standard S-Monovette (Ref. 04.1934) and centrifuged at 2000 g for 5 min. To assess the partial thromboplastin time (PTT), blood was collected into disposable syringes containing 0.11 M sodium citrate, immediately mixed and centrifuged for 10 min at 2000 g. PTT was then measured following addition of PTT reagent to the supernatants in pre-warmed test tubes (37°C), according the manufacturer's instructions (Roche).

Plasma-immunoreactive insulin was measured in duplicate using an enzyme-linked immunosorbent assay (ELISA) kit for insulin (Mercodia, Uppsala, Sweden, kit 10-1137-01).

2.9. Statistical analysis
Values from control and LPS groups were compared using a two-tailed Student's t-test for unpaired data. For multiple comparisons one-way analysis of variance was used, where indicated. All values were expressed as means±S.E.M., and P values ≤0.05 were considered as statistically significant.


   3. Results
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
4. Discussion
 5. Concluding remarks
 References
 
3.1. Effect of lipopolysaccharide treatment on cardiac glycolysis and glucose uptake
As detailed in the Methods section, intravenous application of LPS from S. typhimurium in rats led to a reproducible depression of the mechanical myocardial function which was detectable ex vivo, i.e., in isolated perfused hearts. Using this model, we examined the influence of prior LPS treatment in vivo on cardiac glycolysis, as measured by monitoring the time-dependent cardiac production of 3H2O from 3-[3H]D-glucose. If the isolated hearts were perfused with glucose (5.6 mM) as the sole exogenous energy substrate, the absolute rates of basal and insulin-stimulated glycolysis did not differ between the control and the LPS-treated groups (Fig. 1).


Figure 1
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  		Fig. 1 Influence of LPS treatment on glycolysis of isolated hearts. Six hours after the saline (control) or LPS injection, hearts were isolated and perfused ex vivo in the presence of glucose (5.6 mM), without or with β-hydroxybutyrate (5 mM), and without (‘Basal’) or with insulin (17 nM) as indicated, and the rate of glycolysis was measured as described in Methods. Data are expressed at µmol per g of heart (wet weight) per min, and are means±S.E.M. from 5 to 9 hearts; **: P<0.01 vs. control (unpaired t-test).

 
On the other hand, Table 2 shows that the insulinemia of LPS-injected rats was dramatically increased (eightfold), whereas glycemia was only modestly elevated. Moreover, the serum concentration of non-esterified fatty acids was almost doubled. These findings are indicative of a marked systemic insulin resistance. It is well-established that cardiac (and muscle) glycolysis and glucose utilization are inhibited in situations of increased availability of alternative fuels, such as fatty acids or ketone bodies (which is part of the so-called glucose–fatty acid cycle, or Randle cycle (for recent reviews, see Refs. [39,40]).

We therefore sought to simulate these in vivo conditions by perfusing the isolated hearts with β-hydroxybutyrate in addition to glucose. As illustrated in Fig. 1, perfusion of control hearts with β-hydroxybutyrate produced, as expected, a decrease in glycolytic rate (by 36 and 55% in basal, and insulin-stimulated conditions, respectively) if compared to values measured in the absence of the ketone body. Importantly, the inhibitory effect of β-hydroxybutyrate was much more pronounced in hearts from LPS-treated animals (Fig. 1): thus the basal, and insulin-dependent glycolysis was suppressed by 55 and 76%, respectively. In other words, pretreatment with endotoxin appears to exacerbate the glycolytic depression caused by β-hydroxybutyrate.

Because the glycolytic flux may, under certain conditions, be limited by the rate of glucose uptake (i.e., transport across the plasma membrane and/or phosphorylation), we next investigated whether the observed LPS-dependent change in glycolysis resulted from, or was accompanied by, a decrease in glucose uptake (as determined by measuring the production of tritiated water from 2-[3H]D-glucose). With glucose as the sole exogenous fuel, LPS treatment did not alter basal glucose uptake, and even increased the insulin-stimulated values (Fig. 2). Addition of β-hydroxybutyrate to control hearts diminished glucose uptake by 47 and 41% in the basal, and insulin-stimulated states, respectively (Fig. 2). However, the degree of β-hydroxybutyrate-induced inhibition was not enhanced in the LPS group: basal glucose uptake was inhibited by 44%, and the insulin-stimulated uptake by 45% (Fig. 2). Hence, changes in the rate of glucose uptake cannot explain the LPS-dependent intensification of the inhibitory action of β-hydroxybutyrate at the level of glycolysis.


Figure 2
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  		Fig. 2 Influence of LPS treatment on glucose uptake. Treatment in vivo, and heart perfusions were performed as outlined in the legend of Fig. 1, and the rate of glucose uptake was measured as described in Methods. Data are means±S.E.M. from four hearts; **: P<0.01 vs. control (unpaired t-test).

 
3.2. Influence of LPS on cardiac citrate and glycogen content
In muscle and heart, the reduction of metabolic flow through the phosphofructokinase (PFK) reaction – a major rate-limiting step of glycolysis – by fatty acids and ketone bodies has been explained by an increase in the cellular content of citrate, which allosterically inhibits both PFK-1 and PFK-2 [41]. Perfusion of isolated hearts from saline-treated control rats with β-hydroxybutyrate caused a 2.3-fold increase in their citrate content, when compared to the value obtained upon perfusion with glucose as the only substrate (0.63 vs. 0.28 µmol/g wet weight) (Fig. 3).


Figure 3
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  		Fig. 3 Effect of LPS treatment on cardiac citrate content. Treatment in vivo, and heart perfusions were performed as outlined in the legend of Fig. 1, and the citrate content was determined as described in Methods. Data are means±S.E.M. from 3 to 4 hearts; ***: P<0.001 vs. control (unpaired t-test).

 
In hearts from the LPS group, the citrate content measured at the end of a perfusion with glucose alone was the same as in controls (Fig. 3). However, in the presence of β-hydroxybutyrate, there was a threefold higher level of cardiac citrate, i.e., 40% higher than in controls under the same conditions (Fig. 3). Therefore, it is conceivable that a larger gain in cellular citrate is responsible for the stronger glycolytic inhibition by β-hydroxybutyrate in hearts from the LPS group.

In view of the more pronounced depression of glycolysis by β-hydroxybutyrate in LPS hearts, one might expect that in these hearts a larger proportion of the sugar taken up (or of the glucose-6-phosphate formed from it) is directed towards the synthesis of glycogen than in the control group. We therefore measured the myocardial glycogen content in both control and LPS groups. In controls, the glycogen content (determined at the end of the perfusions, the last 15 min with added insulin) was larger when β-hydroxybutyrate was present than in the absence of the ketone body (Fig. 4, left vs. middle bar). This ‘re-routing’ of glucose towards glycogen synthesis in the presence of an alternative fuel was also observed in previous studies [42]. In contrast, virtually no re-routing occurred in hearts from LPS-treated rats (Fig. 4, compare middle bar with right bar in each group). This means that not only is glycolysis impaired under these conditions, but also glycogen metabolism. In addition, it is noteworthy that in native hearts (i.e., in freshly excised hearts that were immediately frozen without any perfusion), the glycogen level is not higher in the LPS hearts vs. controls, despite the fact that in vivo there is a higher blood concentration of free fatty acids and a massive hyperinsulinemia (Table 2), two factors that are known to produce a net glycogen gain in the heart.


Figure 4
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  		Fig. 4 Effect of LPS treatment on cardiac glycogen content. Six hours after saline control or LPS injection, the heart were excised, and either immediately frozen (‘before perfusion’) or perfused as indicated. After a 40 min perfusion phase with glucose alone or with glucose and β-hydroxybutyrate, which included 15 min in the presence of insulin, the hearts were rapidly frozen and their glycogen content was measured as described in the Methods section. *: P<0.05 versus control without β-hydroxybutyrate (unpaired t-test). Data are means±S.E.M. from four hearts in each group.

 
3.3. Signals mediating the LPS effects on cardiac glycolysis
We next sought to gain a first insight into possible signals underlying the effect of LPS on cardiac glycolysis. In many (though not all) models of systemic inflammation, the synthesis of nitric oxide (NO) is increased in many cell types and organs, including the heart, due to the expression of an inducible NO synthase (iNOS) [43,44]. Because our recent investigations have shown that cyclic GMP, which is produced in the myocardium response to NO, can depress both myocardial glucose transport, and glycolysis [27,30], we examined the possible role of the NO system in the observed effect of LPS on glycolysis. However, under our experimental conditions (i.e., up to 6 h after the injection of LPS), the rate of NO formation in perfused hearts was not different from that measured in the control group (as assessed by monitoring the nitrite content of the coronary effluate; not shown). Moreover, we could not detect any iNOS in freshly excised hearts, whereas a strong expression of the protein was seen in extracts from LPS-treated Kupffer cells used as a positive control (as assessed in Western blots; not shown). Furthermore, addition of the NOS inhibitor L-NAME (1 µM) to the perfusate failed to prevent the LPS-dependent exacerbation of the effect of β-hydroxybutyrate on cardiac glycolysis (not shown).

We therefore examined alternative signals known to be induced in the myocardium in response to inflammatory cytokines. Of particular interest are sphingomyelin metabolites, for two reasons: (i) because they are formed following the activation of sphingomyelinases by TNF{alpha} or IL-1β (two of the major cytokines whose expression is induced in the heart in the early course of inflammatory responses [11,14]); (ii) because sphingomyelin-derived products were previously found to impact on glucose metabolism in insulin-sensitive cells [23,45,46], including isolated rat cardiomyocytes (Thurner et al., manuscript in preparation).

We therefore studied the action of an inhibitor of ceramidase, an enzyme that degrades sphingomyelinase-derived ceramide. Thus, we perfused isolated hearts with the ceramidase inhibitor N-oleyl-ethanolamine (5 µM) prior to β-hydroxybutyrate addition. As illustrated in Fig. 5, the inhibitor had per se no effect on the rate of basal or insulin-stimulated glycolysis in control hearts. However, NOE completely prevented the LPS-dependent glycolytic depression (Fig. 5), indicating that the sphingomyelin-ceramide pathway might be involved in this LPS effect.


Figure 5
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  		Fig. 5 Influence of NOE (ceramidase inhibitor), and NS-398 (COX-2 inhibitor) on LPS-dependent modulation of cardiac glycolysis. Six hours after control or LPS injection, hearts were isolated and perfused ex vivo in the presence of glucose, and with β-hydroxybutyrate, and without (‘Basal’) or with insulin (17 nM) as indicated. Four min before addition of β-hydroxybutyrate, NOE (5 µM), or NS-398 (10 µM) were added to the perfusion, and the rate of glycolysis was measured as described in Methods. *: P<0.05; **: P<0.01 (ANOVA). Data are means±S.E.M. from five hearts in each experimental group.

 
Recently, the TNF{alpha}- and ceramidase-dependent mechanical depression was found to be associated with the cyclooxygenase-mediated production of eicosanoids in hearts treated with LPS [47] or TNF{alpha} [48]. Since the expression of cyclooxygenase-2 (COX-2) is known to be induced by LPS and inflammatory cytokines in many cell types, including cardiac myocytes [47,49,50], we examined whether COX-2-mediated prostanoid production may be involved in the effect of LPS on cardiac glycolysis. Fig. 5 shows that in hearts from LPS-treated rats, perfusion with the COX-2-selective inhibitor NS-398 (10 µM) nearly restored the glycolytic flux to control levels (measured in the presence of β-hydroxybutyrate) (in the control group, the inhibitor did not affect glycolysis; Fig. 5). Similar results were obtained with the non-isoform-selective COX-inhibitor indomethacin (10 µM; not shown). Because thromboxane A2 (TxA2) – a prostanoid synthesized downstream of the COX-dependent reaction – is known to be produced in the heart in response to inflammatory cytokines such as TNF{alpha} [48], we assessed the potential involvement of this mediator by using SQ-29548 (1 µM), a TxA2 receptor antagonist [51] in the perfusate (in the presence of β-hydroxybutyrate). In the basal state (i.e., in the absence of insulin), the antagonist did not modify the rate of glycolysis in control hearts, but it eliminated the inhibitory effect of LPS (Fig. 6, left). In the insulin-stimulated state, SQ-29548 increased glycolysis by 58% in the control group (Fig. 6, right). Such a potentiation of insulin's effect on glycolysis (or any other parameters of glucose metabolism) by TxA2 receptor blockade has not yet been described. On the other hand, in hearts from LPS-treated rats, SQ-29548 raised the glycolytic flux by 182%, i.e., to values not different from the control group (Fig. 6, right).


Figure 6
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  		Fig. 6 Influence of SQ-29548 (TXA2 antagonist) on LPS-dependent effect on cardiac glycolysis. Rats were treated and hearts were isolated and perfused as delineated in the legend of Fig. 5. Four minutes before addition of β-hydroxybutyrate, SQ-29548 (1 µM) was added to the perfusion, and the rate of glycolysis was measured as described in Methods. *: P<0.05; **: P<0.01; ***: P<0.001 (ANOVA). Data are means±S.E.M. from five hearts in each group.

 
3.4. Metabolic effects and contractile function
As described above, in the presence of glucose as the sole fuel, the contractility of perfused hearts from LPS-treated rats was largely depressed (Table 1), in spite of the fact that glycolysis was unaltered (Fig. 1), indicating that at least under these conditions the impairment of the heart's mechanical activity is not due to a change in glycolysis. However, addition of β-hydroxybutyrate to the perfusate (before insulin application) – which reduced the glycolytic rate (Fig. 1) – caused a small but significant decrease in the LVDP by ~10% in both control and LPS groups within 2 min (i.e., from 86±3 to 78±3 mmHg in controls, P<0.001, n = 13; and from 44±4 to 39±3 mmHg in LPS hearts, P<0.001, n = 18). Under these conditions, a subsequent application of insulin resulted in a rapid increase of LVDP (i.e., within 2 to 4 min) by 9% in controls, and by 27% in LPS hearts (Fig. 7) (although an increase was also seen in hearts perfused with glucose as the only fuel, it was clearly weaker than in the presence of β-hydroxybutyrate). Importantly, insulin's positive inotropic action was proportionally much larger in the LPS group (Fig. 7).


Figure 7
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  		Fig. 7 Inotropic effect of insulin. Rats were treated and hearts were isolated and perfused as delineated in the legend of Fig. 1. The left ventricular developed pressure of each experimental group was compared before, and 4 min after insulin addition. **: P<0.01 versus corresponding control with β-hydroxybutyrate; §: P<0.05 vs. LPS group without β-hydroxybutyrate (ANOVA). Data are means±S.E.M. from 8 to 19 hearts per group.

 
These observations suggest that if glycolysis is depressed by β-hydroxybutyrate – and especially so in the LPS group in which this effect is particularly strong – the glycolytic flux may become one of the factors that limit the mechanical function, a limitation that is partially relieved by the stimulatory effect of insulin.


   4. Discussion
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
5. Concluding remarks
 References
 
This study provides evidence that a systemic inflammatory reaction which affects the heart's mechanical function can also impact on the myocardial utilization of glucose by at least two mechanisms: potentiation of the counterregulatory effect of an alternative fuel (β-hydroxybutyrate) on glycolysis, and impairment of insulin-dependent glycogen synthesis.

4.1. Effect of LPS treatment on glycolysis
A salient feature of endotoxemic and septic states in the present and previous studies is an exceedingly high availability of fatty acids and ketone bodies in animals and humans [7,52]. These fuels are known to be able to depress cardiac glycolysis, a phenomenon which is part of the so-called glucose fatty acid cycle (or Randle cycle) [39,40]. When isolated hearts were perfused with β-hydroxybutyrate to mimic such conditions, a clear difference between the saline-treated and the LPS-treated groups became apparent, in terms of a stronger inhibition of glycolysis in the latter. To our knowledge, this is the first demonstration of an inflammatory effect on cardiac glycolysis.

We have not investigated the concentration-dependence of β-hydroxybutyrate's effect on glycolysis, nor determined the plasma level of ketone bodies in the present study, so that we cannot make a precise prediction of the relative impact of ketone bodies on cardiac glycolysis in vivo. However, a hyperketonemia is observed in similar rat models of endotoxemia (e.g., Ref. [52]), as well as in human sepsis [7]. Moreover, we have measured an increase in serum fatty acids to 0.75 mM following LPS treatment (Table 2), a concentration which was previously shown to affect glycolysis in the same way as ketone bodies [53]. In view of these findings, one may speculate that endotoxemia could restrain cardiac glycolysis in vivo by two concurring mechanisms: (i) the increased supply of fatty acids and ketone bodies (through lipolysis, and hepatic ketogenesis), and (ii) the exacerbation of their counterregulatory action on glycolysis.

A reduction of glycolysis in heart is not necessarily in contradiction with the fact that the overall rate of body glycolysis is not decreased [9], or is even increased in critical inflammatory states [19]; first, being a relatively small organ, the heart makes only a minor contribution to the total clearance and oxidation of blood glucose, and second, it is the insulin-independent glucose utilization that is increased in sepsis, mainly in macrophage-rich tissues such as, e.g., the spleen, lung and liver [1,2].

Along with the stronger glycolytic inhibition by β-hydroxybutyrate in the LPS-group, there was a 40% higher cardiac content of citrate compared with saline-treated controls under the same conditions. Because citrate is a known allosteric inhibitor of the phosphofructokinase, which catalyzes a major rate-limiting step of glycolysis, it seems likely that the higher level of this metabolite is responsible for the glycolytic inhibition seen in the LPS group.

In the present work we have not differentiated between basal and insulin-stimulated conditions with regard to citrate accumulation (citrate was measured at the end of a perfusion consisting of a basal and an insulin-stimulated phase). However, ketone bodies have been previously shown to increase myocardial citrate content in the absence of insulin, and insulin addition on top of a ketone body does not cause any further elevation of citrate (e.g., Ref. [54]). This fact suggests, although it does not prove, that the LPS-dependent potentiation of citrate accumulation is not specific of insulin-stimulated conditions, but may involve a mechanism which is also functional in non-stimulated hearts. This mechanism remains to be elucidated, though. In principle, the rate of β-hydroxybutyrate oxidation (to acetoacetate and then to acetyl-CoA which enters the citrate synthase reaction) could be increased, thus leading to an accelerated formation of citrate; alternatively, the turnover of citrate in the tricarboxylic acid cycle could be diminished, for instance by an exacerbation of a previously described mechanism, namely the blockade of the {alpha}-ketoglutarate dehydrogenase reaction by ketone bodies [55]. However, a citrate-independent inhibition of glycolysis is also conceivable, e.g., a covalent modification of PFK.

We have not directly measured the rate of glucose oxidation in this study. However, in preliminary experiments, we had found that the rate of lactate production (in hearts perfused with glucose alone) was not modified by the treatment with LPS (not shown). This suggests that about the same proportion of the glycolytically derived pyruvate was oxidized in both control and LPS groups. Moreover, measurements of energy-rich phosphates in extracts from freshly excised hearts showed no LPS-dependent changes in the levels of ATP, ADP, AMP or creatine phosphate (not shown). Therefore the LPS treatment per se is unlikely to cause a substantial reduction of glucose oxidation. Furthermore the fact that, in hearts from LPS-treated rats, addition of β-hydroxybutyrate causes a large increase in citrate levels shows that the total oxidative capacity of mitochondria is not impaired either.

4.2. Glycogen metabolism
Another new finding of this study is the profound disturbance of cardiac glycogen metabolism following the LPS administration – as evidenced by the inability of the β-hydroxybutyrate perfusion to increase the net glycogen content, which contrasts with the massive re-routing of glucose towards glycogen synthesis seen in hearts from control rats. Of further note, the native glycogen content was not higher in hearts from the LPS-treated animals than in controls, despite the pronounced hyperinsulinemia and elevated plasma concentration of fatty acids measured in vivo, two factors that are known to produce a net glycogen gain in the heart [39,56]. This points to the existence of a marked insulin resistance of the heart's glycogen metabolism in vivo, too. Interestingly, a nearly total unresponsiveness of glycogen synthesis to insulin has also been found in skeletal muscles of endotoxemic or septic rats in vivo [9,19]. In line with this, in patients with acute infections, the insulin-dependent non-oxidative disposal of glucose (mainly by muscles) is dramatically decreased compared to healthy subjects [8].

The absence of a re-routing towards a net glycogen synthesis raises the question of the eventual fate of glucose-6-phosphate stemming from the glucose taken up, because we found that cardiac glucose uptake (as measured with 2-[3H]D-glucose) was not decreased by the endotoxemia. A possible explanation is that glucose-6-phosphate still enters glycogen synthesis, but that glycogen breakdown occurs at the same time; in other words LPS treatment may lead to a futile cycling. Interestingly, simultaneous incorporation of exogenous glucose into glycogen and degradation of glycogen has previously been reported in hearts from healthy rats [54]; it is therefore conceivable that LPS treatment reinforces such a turnover to an extent that even in the presence of insulin, no net glycogen accumulation is observed. In such a case, one should expect only a limited net consumption of hexose-phosphates, and an increase in their intracellular levels. An alternative use of glucose-6-phosphate may be that it is fed into the pentose-phosphate, or into the hexosamine pathway, although under normal conditions these pathways represent rather minor metabolic routes in the heart. Further investigations will be needed to resolve the underlying mechanism of the insulin resistance of glycogen metabolism, as well as the question of the cellular fate of the excess glucose-6-phosphate.

4.3. Signal(s) mediating the LPS-dependent glycolytic inhibition
The cardiac production of NO (and of its second messenger cGMP) has been shown to be increased in some [43,44], though not all [47,57] models of LPS-induced inflammation. However, in the present model, we could not detect any sign of a cardial activation of the NO system, as evidenced by a lack of iNOS expression, and of an increase in NO production in hearts from LPS-treated animals. Therefore, it can be concluded that in our model NO and/or cGMP do not participate in the effect of LPS on glycolysis seen in this study ex vivo. On the other hand, it cannot be ruled out that an extracardial increase in NO production may affect the utilization of glucose in the heart in vivo.

Endotoxins are known to induce the release of inflammatory cytokines such as TNF{alpha} and IL-1β in the heart [11,14]. We therefore took the sphingomyelin-ceramide cascade into consideration, which is a rapidly activated in response to these cytokines. The action of the ceramidase inhibitor NOE to suppress the LPS-dependent potentiation of β-hydroxybutyrate on glycolysis indicates that this signaling pathway is involved here. This is to our knowledge the first demonstration that the regulation of glycolysis may be modulated by sphingomyelin derivatives. Previous studies performed in adipose and muscle cells have shown that hydrolysis of endogenous sphingomyelin or direct application of ceramide (a product of this hydrolysis) leads to an inhibition of the insulin signal transduction chain and depresses insulin's stimulating action on the glucose transport system of these cells [23,45,46]. The fact that under our experimental conditions cardiac glycolysis is impaired in the absence of a change in glucose uptake suggests that the effect observed here is independent of a modulation at the level of the glucose transport.

We next examined possible signals downstream of sphingosine (the product of the NOE-sensitive ceramide reaction). TNF{alpha} [58], as well as IL-1β [49] can activate a phospholipase A2, so having the potential to release arachidonic acid and to increase the cyclooxygenase-mediated synthesis of prostanoids. Sphingosine was shown to be at least in part responsible for, or to promote, this activation of the phospholipase A2 [58]. The recent demonstration that an interaction between sphingomyelinase and phospholipase A2/cyclooxygenase signaling also occurs in hearts perfused with LPS [47] or TNF{alpha} [48] prompted us to assess the involvement of the latter (i.e., cyclooxygenase) pathway in LPS-dependent glycolysis modulation.

Our finding that blockade of either the sphingomyelin pathway (with NOE) or the prostanoid pathway (with the COX-2 inhibitor NS-398, or the thromboxane receptor antagonist SQ-29548) results in the complete prevention of the LPS effect on glycolysis may be interpreted in terms of a sequential activation of these signaling systems. A possible sequence of events also consistent with the studies quoted in the previous paragraph would be that sphingosine (or a metabolite thereof) causes the release of arachidonic acid and – via the cyclooxygenase reaction – the production of prostanoids such as TxA2 which, in turn, would act as effector(s) of the observed glycolytic modulation. The inflammation-dependent induction of COX-2 expression in heart which was detected following LPS-treatment [47], but also in patients with congestive heart failure [59], may be an essential element of this cascade, as suggested by the action of NS-398. Thus, in this scenario, the signaling cascade may be as follows: LPS -> cytokine (e.g., TNF{alpha}) expression -> activation of sphingomyelinase pathway (blocked by NOE) -> phospholipase A2 activation -> arachidonic acid release -> COX-2-mediated prostanoid (TxA2) production (blocked by NS-398) -> TxA2 receptor activation (blocked by SQ-29548).

However, a different sequence of events may also exist, in which for instance the increased availability of arachidonic acid and TxA2 would stimulate the formation of sphingomyelin derivatives [60–62]. Whatever scenario may be envisaged, it is unlikely that PGE2, another prostanoid reported to be formed in response to TNF{alpha} stimulation [58], or to COX-2-mediated arachidonic metabolism [49], plays a role in the present setting, because no effect of PGE2 on insulin-stimulated glycolysis was detected in hearts [63]; moreover, insulin sensitivity correlated positively with PGE2 levels in muscle [64]. Of interest, the present study provides the first known evidence of a TxA2-mediated regulation of glycolysis.

A major implication of the observation that the effect on glycolysis of LPS administered in vivo could be totally reversed by the ex vivo application of inhibitors of the sphingomyelin and eicosanoid pathways is that factors such as the TNF{alpha}, COX-2, and signaling metabolites are induced or produced locally in the myocardium and have become independent of the primary stimulus in vivo (e.g., LPS, and possibly non-cardiac factors induced by it).

Regardless of the mediators/signals involved in the observed metabolic changes, a further of interest is whether the insulin signaling is affected by LPS treatment. The stimulation of glucose uptake by insulin was not blunted by LPS treatment, suggesting that there is at least no proximal impairment (i.e., at an early step) of the insulin signaling in our model. Moreover, the LPS-dependent change in glycolysis was also seen in the basal state, which indicates that it cannot be entirely related to an altered insulin signaling. On the other hand, because in perfused hearts, glycogen synthesis is largely insulin-dependent, the massive reduction in glycogen accumulation in LPS hearts may be indicative of a more specific inhibition of insulin's signaling and/or action, although this issue remains to be investigated.

4.4. Glucose metabolism and contractility
The large depression of the left ventricular contractility observed ex vivo following the in vivo treatment with endotoxin is unlikely to be due to an impairment of glycolysis because this contractile dysfunction was also observed under conditions where glycolysis was not altered in the LPS group, namely if glucose was used as the sole substrate. However, under certain conditions, glycolysis may become more essential for the maintenance of contractility. This may for instance be the case when the glycolytic flux falls below a critical level. It is noteworthy that insulin had its most pronounced inotropic effect in the experimental group with the lowest basal glycolytic rate, i.e., in LPS hearts perfused with β-hydroxybutyrate. Interestingly, earlier studies performed in canine heart preparations following endotoxin treatment have shown a partial recovery of mechanical parameters with insulin infusions [65]. Alternatively, a limitation of glycolysis may become critical when energy demand is higher, or during ischemic episodes, where oxidative phosphorylation of alternative fuels is limited and glycolysis’ contribution to ATP synthesis becomes more important.

Thus, although a reduction in glycolysis is certainly not a major cause of the LPS-induced mechanical failure, the metabolic effect of LPS may, under in vivo conditions, contribute to, or worsen the cardiodepression. The main cause(s) of this cardiodepression may lie in previously suggested mechanisms such as e.g., changes in myocardial calcium homeostasis (for review see, e.g., Ref. [13]).


   5. Concluding remarks
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 5. Concluding remarks
References
 
Our investigations have revealed new mechanisms that can potentially impair the myocardial metabolism of glucose in an inflammatory situation: the exacerbation of the glycolytic inhibition by alternative fuels (present study), the impairment of glycogen synthesis (present study), as well as an inhibitory action of the NO/cGMP system on glucose transport (in models and situations in which this system is activated). Under which conditions and to what extent one or several of these processes may cause an impairment of cardiac glucose utilization in vivo, and possibly affect the heart's mechanical function remains to be established. In principle, all these effects could contribute to the cardial insulin resistance that is seen in sepsis or injury. In view of the recent demonstration that an intensive insulin therapy – used to overcome the insulin resistance in intensive care patients – substantially decreased the overall mortality and morbidity [66], a further elucidation of mechanisms affecting the cardiac action of inflammatory mediators in critical conditions may be of interest.


   Acknowledgements
 
We acknowledge the kind help provided by Dr. D. Kunz, Ms. Sauer-Lehnen, and Professor A. Gressner (Institute of Clinical Chemistry, Aachen, Germany) who performed the hematological measurements. This work was funded by the Deutsche Forschungsgemeinschaft (SFB 542). We are also grateful to Professor P.C. Heinrich for his logistic, and moral support as head of the SFB 542.


   Notes
 
Time for primary review 28 days.


   References
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 5. Concluding remarks
 References
 

  1. Kinney J.M. Metabolic responses of the critically ill patient. Crit Care Clin (1995) 11:569–585.[ISI][Medline]
  2. Mizock B.A. Alterations in carbohydrate metabolism during stress: a review of the literature. Am J Med (1995) 98:75–84.[CrossRef][ISI][Medline]
  3. Chiolero R., Revelly J.P., Tappy L. Energy metabolism in sepsis and injury. Nutrition (1997) 13:45S–51S.[CrossRef][Medline]
  4. Wolfe R.R. Substrate utilization/insulin resistance in sepsis/trauma. Ballieres Clin Endocrinol Metab (1997) 11:645–657.[CrossRef][ISI][Medline]
  5. Lang C.H., Dobrescu C., Meszaros K. Insulin-mediated glucose uptake by individual tissues during sepsis. Metabolism (1990) 39:1096–1107.[CrossRef][ISI][Medline]
  6. Shangraw R.E., Jahoor F., Wolfe R.R., Lang C.H. Pyruvate dehydrogenase inactivity is not responsible for sepsis-induced insulin resistance. Crit Care Med (1996) 24:566–574.[CrossRef][ISI][Medline]
  7. Chambrier C., Laville M., Rhzioual Berrada K., et al. Insulin sensitivity of glucose and fat metabolism in severe sepsis. Clin Sci (Lond) (2000) 99:321–338.[Medline]
  8. Virkamaki A., Puhakainen I., Koivisto V.A., Vuorinen-Markkola H., Yki-Jarvinen H. Mechanisms of hepatic and peripheral insulin resistance during acute infections in humans. J Clin Endocrinol Metab (1992) 74:673–679.[Abstract]
  9. Virkamaki A., Yki-Jarvinen H. Mechanisms of insulin resistance during acute endotoxemia. Endocrinology (1994) 134:2072–2080.[Abstract]
  10. Raymond R.M., Klein D.M., Gibbons D.A., Jacobs H.K., Emerson T.E. Jr. Skeletal muscle insulin unresponsiveness during chronic hyperdynamic sepsis in the dog. J Trauma (1985) 25:845–855.[ISI][Medline]
  11. Kelly R.A., Smith T.W. Cytokines and cardiac contractile function. Circulation (1997) 95:778–781.[Free Full Text]
  12. Parker M.M. Pathophysiology of cardiovascular dysfunction in septic shock. New Horiz (1998) 6:130–138.[ISI][Medline]
  13. Werdan K., Müller-Werdan U. Elucidating molecular mechanisms of septic cardiomyopathy—the cardiomyocyte model. Mol Cell Biochem (1996) 163–164:291–303.
  14. Meldrum D.R. Tumor necrosis factor in the heart. Am J Physiol (1998) 274:R577–R595.[ISI][Medline]
  15. Opie L.H. Myocardial ischemia—metabolic pathways and implications of increased glycolysis. Cardiovasc Drugs Ther (1990) 4:777–790.[CrossRef][ISI][Medline]
  16. Vanoverschelde J.L., Janier M.F., Bakke J.E., Marshall D.R., Bergmann S.R. Rate of glycolysis during ischemia determines extent of ischemic injury and functional recovery after reperfusion. Am J Physiol (1994) 267:H1785–H1794.[ISI][Medline]
  17. McVeigh J.J., Lopaschuk G.D. Dichloroacetate stimulation of glucose oxidation improves recovery of ischemic rat hearts. Am J Physiol (1990) 259:H1079–H1085.[ISI][Medline]
  18. Nicholl T.A., Lopaschuk G.D., McNeill J.H. Effects of free fatty acids and dichloroacetate on isolated working diabetic rat heart. Am J Physiol (1991) 261:H1053–H1059.[ISI][Medline]
  19. Lang C.H. IGF-I stimulates muscle glucose uptake during sepsis. Shock (1996) 5:22–27.[ISI][Medline]
  20. Raymond R.M., McLane M.P., Law W.R., King N.F., Leutz D.W. Myocardial insulin resistance during acute endotoxin shock in dogs. Diabetes (1988) 37:1684–1688.[CrossRef][ISI][Medline]
  21. Lang C.H., Dobrescu C., Bagby G.J. Tumor necrosis factor impairs insulin action on peripheral glucose disposal and hepatic glucose output. Endocrinology (1992) 130:43–52.[Abstract]
  22. Hotamisligil G.S., Murray D.L., Choy L.N., Spiegelman B.M. Tumor necrosis factor alpha inhibits signaling from the insulin receptor. Proc Natl Acad Sci USA (1994) 91:4854–4858.[Abstract/Free Full Text]
  23. Begum N., Ragolia L. Effect of tumor necrosis factor-alpha on insulin action in cultured rat skeletal muscle cells. Endocrinology (1996) 137:2441–2446.[Abstract]
  24. Bedard S., Marcotte B., Marette A. Cytokines modulate glucose transport in skeletal muscle by inducing the expression of inducible nitric oxide synthase. Biochem J (1997) 325:487–493.[ISI][Medline]
  25. Schulz R., Nava E., Moncada S. Induction and potential biological relevance of a Ca(2+)-independent nitric oxide synthase in the myocardium. Br J Pharmacol (1992) 105:575–580.[ISI][Medline]
  26. Depre C., Vanoverschelde J.L., Goudemant J.F., Mottet I., Hue L. Protection against ischemic injury by nonvasoactive concentrations of nitric oxide synthase inhibitors in the perfused rabbit heart. Circulation (1995) 92:1911–1918.[Abstract/Free Full Text]
  27. Depre C., Gaussin V., Ponchaut S., et al. Inhibition of myocardial glucose uptake by cyclic GMP. Am J Physiol (1998) 274:H1443–H1449.[ISI][Medline]
  28. Tada H., Thompson C.I., Recchia F.A., et al. Myocardial glucose uptake is regulated by nitric oxide via endothelial nitric oxide synthase in Langendorff mouse heart. Circ Res (2000) 86:270–274.[Abstract/Free Full Text]
  29. Recchia F.A., McConnell P.I., Bernstein R.D., et al. Reduced nitric oxide production and altered myocardial metabolism during the decompensation of pacing-induced heart failure in the conscious dog [see comments]. Circ Res (1998) 83:969–979.[Abstract/Free Full Text]
  30. Bergemann C., Löken C., Becker C., et al. Inhibition of glucose transport by cyclic GMP in cardiomyocytes. Life Sci (2001) 69:1391–1406.[CrossRef][ISI][Medline]
  31. Langendorff O. Untersuchungen am überlebenden Säugetierherzen. Pflügers Arch (1895) 61:291–332.[CrossRef]
  32. Wieüngling E., Kammermeier H. Enzyme release into the interstitial space of the isolated rat heart induced by changes in contractile performance. Cardiovasc Res (1994) 28:1292–1298.[Abstract/Free Full Text]
  33. Meng X., Ao L., Meldrum D.R., et al. TNF-alpha and myocardial depression in endotoxemic rats: temporal discordance of an obligatory relationship. Am J Physiol (1998) 275:R502–R508.[ISI][Medline]
  34. Bontemps F., Hue L., Hers H.-G. Phosphorylation of glucose in isolated rat hepatocytes. Sigmoidal kinetics explained by activity of glucokinase alone. Biochem J (1978) 174:603–611.[ISI][Medline]
  35. Hue L., Hers H.G. On the use of (3 H,14C)glucose in the study of the so-called ‘futile cycles’ in the liver and muscle. Biochem Biophys Res Commun (1974) 58:532–539.[ISI][Medline]
  36. Giesen J., Sondermann M., Juengling E., Kammermeier H. Time dependent partial loss of the effects of isoproterenol on function and energy metabolism of isolated rat hearts. Basic Res Cardiol (1980) 75:515–525.[CrossRef][ISI][Medline]
  37. Hue L., Bontemps F., Hers H.G. The effect of glucose and potassium ions on the interconversion of the two forms of glycogen phosphorylase and of glycogen synthase in rat liver preparations. Biochem J (1975) 152:105–114.[ISI][Medline]
  38. Dagley J. Methods of enzymatic analysis, 2nd ed. Bergmeyer H.U., ed. (1974) vol. 3. New York, London: Academic Press. 1562–1565.
  39. Randle P.J. Regulatory interactions between lipids and carbohydrates: the glucose fatty acid cycle after 35 years. Diabetes Metab Rev (1998) 14:263–283.[CrossRef][ISI][Medline]
  40. Kraegen E.W., Cooney G.J., Ye J.M., Thompson A.L., Furler S.M. The role of lipids in the pathogenesis of muscle insulin resistance and beta cell failure in type II diabetes and obesity. Exp Clin Endocrinol Diabetes (2001) 109(suppl. 2):S189–S201.[CrossRef][ISI][Medline]
  41. Depre C., Ponchaut S., Deprez J., Maisin L., Hue L. Cyclic AMP suppresses the inhibition of glycolysis by alternative oxidizable substrates in the heart. J Clin Invest (1998) 101:390–397.[ISI][Medline]
  42. Depre C., Rider M.H., Hue L. Mechanisms of control of heart glycolysis. Eur J Biochem (1998) 258:277–290.[ISI][Medline]
  43. Kelly R.A., Balligand J.L., Smith T.W. Nitric oxide and cardiac function. Circ Res (1996) 79:363–380.[Free Full Text]
  44. Nishigaki R., Aramaki T., Hirakawa K., Asano G. Nitric oxide production and energy state in the heart after endotoxin administration. Jpn Circ J (1998) 62:527–531.[CrossRef][Medline]
  45. Kanety H., Hemi R., Papa M.Z., Karasik A. Sphingomyelinase and ceramide suppress insulin-induced tyrosine phosphorylation of the insulin receptor substrate-1. J Biol Chem (1996) 271:9895–9897.[Abstract/Free Full Text]
  46. Summers S.A., Garza L.A., Zhou H., Birnbaum M.J. Regulation of insulin-stimulated glucose transporter GLUT4 translocation and Akt kinase activity by ceramide. Mol Cell Biol (1998) 18:5457–5464.[Abstract/Free Full Text]
  47. Grandel U., Fink L., Blum A., et al. Endotoxin-induced myocardial tumor necrosis factor alpha synthesis depresses contractility of isolated rat hearts. Circulation (2000) 102:2758–2764.[Abstract/Free Full Text]
  48. Edmunds N.J., Woodward B. Effects of tumour necrosis factor-alpha on the coronary circulation of the rat isolated perfused heart: a potential role for thromboxane A2 and sphingosine. Br J Pharmacol (1998) 124:493–498.[CrossRef][ISI][Medline]
  49. LaPointe M.C., Isenovic E. Interleukin-1beta regulation of inducible nitric oxide synthase and cyclooxygenase-2 involves the p42/44 and p38 MAPK signaling pathways in cardiac myocytes. Hypertension (1999) 33:276–282.[Abstract/Free Full Text]
  50. Schuette R., LaPointe M.C. Phorbol ester stimulates cyclooxygenase-2 expression and prostanoid production in cardiac myocytes. Am J Physiol (2000) 279:H719–H725.[ISI]
  51. Monshizadegan H., Hedberg A., Webb M.L. Characterization of binding of a specific antagonist, [3H]-SQ 29,548, to soluble thromboxane A2/prostaglandin H2 (TP) receptors in human platelet membranes. Life Sci (1992) 51:431–447.[CrossRef][ISI][Medline]
  52. Malaisse W.J., Nadi A.B., Ladriere L., Zhang T.M. Protective effects of succinic acid dimethyl ester infusion in experimental endotoxemia. Nutrition (1997) 13:330–341.[ISI][Medline]
  53. Randle P.J., Newsholme E.A., Garland P.B. Regulation of glucose uptake by muscle. Effects of fatty acids, ketone bodies and pyruvate, and of alloxan-diabetes and starvation, on the uptake and metabolic fate of glucose in rat heart and diaphragm muscles. Biochem J (1964) 93:652–665.[ISI][Medline]
  54. Russell R.R., Cline G.W., Guthrie P.H., et al. Regulation of exogenous and endogenous glucose metabolism by insulin and acetoacetate in the isolated working rat heart. J Clin Invest (1997) 100:2892–2899.[ISI][Medline]
  55. Russell R.R., Taegtmeyer H. Coenzyme A sequestration in hearts oxidizing ketone bodies. J Clin Invest (1992) 89:968–973.[ISI][Medline]
  56. Stanley W.C., Lopaschuk G.D., Hall J.L., McCormack J.G. Regulation of myocardial carbohydrate metabolism under normal and ischemic conditions. Potential for pharmacological interventions. Cardiovasc Res (1997) 33:243–257.[Free Full Text]
  57. Roberts A.B., Vodovotz Y., Roche N.S., Sporn M.B., Nathan C.F. Role of nitric oxide in antagonistic effects of transforming growth factor-beta and interleukin-1 beta on the beating rate of cultured cardiac myocytes. Mol Endocrinol (1992) 6:1921–1930.[Abstract]
  58. Candela M., Barker S.C., Ballou L.R. Sphingosine synergistically stimulates tumor necrosis factor alpha induced protaglandin E2 production in human fibroblasts. J Exp Med (1991) 174:1363–1369.[Abstract/Free Full Text]
  59. Wong S.C.Y., Mitsumasa F., Melnyk P., Rodger I., Giaid A. Induction of cyclooxygenase-2 and activation of nuclear factor-kB in myocardium of patients with congestive heart failure. Circulation (1998) 98:100–103.[Abstract/Free Full Text]
  60. Amadou A., Nawrocki A., Best-Belpomme M., Pavoine C., Pecker F. Arachidonic acid mediates dual effect of TNF-alpha on Ca2+ transients and contraction of adult ra