1 Department of Physiology, New York Medical College, Valhalla, New York 10595; 2 Department of Physiology and Biophysics, Case Western Reserve University, Cleveland, Ohio 44106; and 3 Heritage Medical Research Centre, University of Alberta, Edmonton, Canada T6G 2S2
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
To test whether the
acute reduction of nitric oxide (NO) synthesis causes changes in
cardiac substrate metabolism and in the activity of key enzymes of
fatty acid and glucose oxidation, we blocked NOS by giving
N-nitro-L-arginine methyl
ester (L-NAME; 35 mg/kg iv two times) to nine
chronically instrumented dogs. [3H]oleate,
[14C]glucose, and [13C]lactate were infused
to measure the rate of cardiac substrate uptake and oxidation.
Glyceraldehyde-3-phosphate dehydrogenase, acetyl-CoA carboxylase, and
malonyl-CoA decarboxylase activities were measured in myocardial
biopsies. In eight control dogs, ANG II was infused (20-40
ng · kg
1 · min
1) to mimic
the hemodynamic effects of L-NAME. After
L-NAME, significant changes occurred for fatty acid
oxidation (from 9.8 ± 0.8 to 7.1 ± 1.2 µmol/min), glucose
uptake (from 12.9 ± 5.5 to 45.0 ± 14.2 µmol/min), and
oxidation (from 4.4 ± 1.2 to 19.9 ± 2.3 µmol/min). ANG
caused only a significantly lower increase in glucose oxidation. Lactate uptake increased by more than twofold in both groups. The
enzyme activities did not differ significantly between the two groups.
In conclusion, the acute inhibition of NO synthesis causes marked
metabolic alterations that do not involve key rate-controlling enzymes
of fatty acid oxidation nor glyceraldehyde-3-phosphate dehydrogenase.
fatty acids; glucose; lactate; oxidation; heart
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
UNDER AEROBIC CONDITIONS, the heart derives most of the energy necessary for its contractile function from fatty acid oxidation (18, 20, 26). After an overnight fast, carbohydrate oxidation accounts for only 0-20% of total cardiac oxygen consumption, but in some pathological states, such as hypertrophic cardiomyopathy and heart failure, myocardial utilization of fatty acids can diminish significantly (18, 25, 28, 47-48). Studies in humans (33), pigs (16), dogs (22), and rats (2) indicate that the type of oxidized substrate may influence the efficiency of normal hearts. Moreover, several studies have emphasized the contribution of metabolic alterations to the mechanisms of cardiac dysfunction in pathological conditions. For instance, it has been shown that the pharmacological inhibition of fatty acid utilization to favor carbohydrate oxidation can attenuate electrophysiological alterations and improve performance in ischemic or failing hearts (27, 30, 41).
Despite the abundance of studies on this topic, the complex regulation of cardiac substrate metabolism, in health and disease, is only partially understood. Among the numerous factors involved in this regulation, an important role is likely played by nitric oxide (NO) tonically synthesized in cardiac tissue by the constitutive form of NO synthase (eNOS). We found that the fall in cardiac NO production occurring during end-stage heart failure was temporally associated with a decrease in free fatty acid (FFA) uptake and a marked increase in glucose uptake and respiratory quotient (28). Similarly, acute inhibition of nitric oxide synthase (NOS) in normal hearts caused a shift toward carbohydrate utilization that was completely reversed by the administration of exogenous NO (29). Further and strong evidence indicating the involvement of NO in cardiac substrate metabolism derives from in vitro studies. Tada et al. (40) found that baseline glucose uptake is markedly elevated in isolated hearts from eNOS knockout mice. This study and another by Depre et al. (4) showed that cGMP, the second messenger of NO, inhibits glucose uptake in isolated hearts. Despite this experimental evidence, the exact role of NO in the control of myocardial substrate utilization and the underlying mechanisms remain undetermined. To date, the only enzyme of carbohydrate metabolism known to be inhibited by pharmacological or pathological concentrations of NO is the glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which catalyzes a key reaction in the glycolytic pathway (23, 49). On the other hand, the rate of FFA oxidation is limited by the activity of carnitine palmitoyltransferase I, which is necessary for the transfer of long-chain FFA into mitochondria (20). This enzyme is reversibly inhibited by cytosolic malonyl-CoA, which is synthesized and degraded, respectively, by acetyl-CoA carboxylase (ACC) and malonyl-CoA decarboxylase (MCD; see Refs. 1, 14, 31). No studies have explored possible interactions between NO and the activity of ACC and MCD.
The first aim of the present study was to determine the metabolic fate of the three main cardiac substrates, i.e., FFA, lactate, and glucose, before and after inhibition of NO synthesis in conscious dogs. In our previous studies (28-29), we measured net substrate uptake by the heart, but we could not determine the rate of FFA and carbohydrate oxidation before and after acute NOS blockade, and we did not explore the biochemical mechanisms involved in the observed metabolic changes. Moreover, the arterial concentration of FFA fell markedly after NOS inhibition, and this phenomenon itself could have stimulated cardiac carbohydrate oxidation. In the present study, we used three different isotopic tracers to track the main cardiac substrates, while preventing marked oscillations of FFA levels by infusion of a triglyceride emulsion plus heparin. Our second aim was to determine whether inhibition of NO synthesis caused changes in the concentration of some intermediate products of substrate metabolism and in the activity of myocardial GAPDH, ACC, and MCD. GAPDH was selected among the numerous enzymes of glucose metabolism to test whether physiological concentrations of endogenously produced NO can exert a tonic inhibition on this enzyme. On the other hand, ACC and MCD are the key enzymes known to regulate the rate of FFA oxidation by controlling the cytosolic concentration of malonyl-CoA. We tested the hypothesis that NO influences the activity of one or both of these enzymes.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Surgical instrumentation. Seventeen adult, male, mongrel dogs (25-27 kg) were sedated with acepromazine maleate (1 mg/kg im), anesthetized with pentobarbital sodium (25 mg/kg iv), and ventilated with room air. A thoracotomy was performed in the left fifth intercostal space. One catheter was placed in the descending thoracic aorta, and a second catheter was placed in the coronary sinus with the tip leading away from the right atrium. A solid-state pressure gauge (P6.5; Konigsberg Instruments) was inserted in the left ventricle through the apex. A Doppler flow transducer (Craig Hartley) was placed around the left circumflex coronary artery (LcxCBF), and a pair of 3-MHz piezoelectric crystals was fixed on opposing endocardial surfaces at the base of the left ventricle. A human, screw-type, unipolar myocardial pacing lead was placed on the left ventricular (LV) wall. Wires and catheters were run subcutaneously to the intrascapular region, the chest was closed in layers, and the pneumothorax was reduced. Antibiotics were given after surgery, and the dogs were allowed to recover fully. After 7-10 days, dogs were trained to lie quietly on the laboratory table. The protocols were approved by the Institutional Animal Care and Use Committee of the New York Medical College and conform to the guiding principles for the care and use of laboratory animals published by the National Institutes of Health. We have described these methods previously (28, 29).
Hemodynamic recordings.
The aortic catheter was attached to a P23ID strain-gauge transducer for
measurement of aortic pressure. LV pressure was measured using the
solid-state pressure gauge. The first derivative of LV pressure (LV
dP/dt) was obtained using an operational amplifier (National
Semiconductor LM 324). Coronary blood flow (CBF) was measured with a
pulsed Doppler flowmeter (model 100; Triton Technology). LV diameter
was measured by connecting the implanted piezoelectric crystals to an
ultrasonic dimension gauge. All signals were recorded on an
eight-channel direct-writing oscillograph (model RS 3800; Gould). The
analog signals were also stored in computer memory through an
analog-digital interface (National Instruments), at a sampling rate of
250 Hz. By using a custom-made program, we measured end-systolic and
end-diastolic diameters at the left upper corner and the right lower
corner of the pressure-diameter loop, respectively. We found that, at
the pacing rate used in our experiments, these diameters corresponded
to the minimum and maximum measured during the cardiac cycle.
Fractional shortening was calculated as (LV end-diastolic LV
end-systolic diameter)/LV end-diastolic diameter × 100 (28). Pressure-diameter loop areas, proportional to LV
work, were also calculated.
Total and labeled metabolite measurements. Blood gases were measured in a blood gas analyzer (Instruments Laboratory). Oxygen content was measured by a hemoglobin analyzer (CO-Oximeter; Instruments Laboratory). Total FFA concentration was determined spectrophotometrically in plasma after centrifugation of EDTA-treated blood samples using a colorimetric assay (NEFA C kit from Wako Pure Chemical Industries). Total glucose and lactate were measured in blood deproteinized with ice-cold 1 M perchloric acid (1:2 vol/vol) using spectrophotometric enzymatic assays (10, 11).
The following three isotopically labeled substrates were infused into the dogs: [9,10-3H]oleate, [U-14C]glucose, and L-[1-13C]lactate. Plasma [3H]oleate concentration was measured by extracting fatty acids from 1 ml of plasma in 4 ml of heptane-isopropanol (3:7) and counting the radioactivity of the organic phase with a liquid scintillation counter (model LS 6500; Beckman). 3H2O concentration was measured by distilling 2 ml of plasma in custom-made, modified Hickman stills (Kontes Glass; Custom Shop). We developed this method and tested it to exclude possible contamination of the distillate with [14C]- or [3H]oleate. Blood samples for [14C]glucose measurements were first deproteinized in ice-cold 1 M perchloric acid (1:2 vol/vol). The acidity of the extract was then neutralized with K2CO3, and the neutral solution was run through ion-exchange resin columns (Dowex 1 and 50) to separate secondary labeled [14C]lactate, -pyruvate, and -alanine. Total glucose concentration and 14C activity were then measured in the eluate to calculate [14C]glucose specific activity. 14CO2 concentration in blood was determined using the method of Gertz et al. (6) and Wisneski et al. (45). Briefly, 3-ml blood samples were placed in a sealed flask with a center well containing 800 µl of 1 M NaOH. The blood was acidified with 1 ml of concentrated lactic acid (98%), and the flask was placed on a rotating table at room temperature for 3 h. All of the CO2 so displaced from the blood combined with solution contained in a well inside the flask to form bicarbonate. The newly formed solution of bicarbonate was then collected to count the activity of 14C (6, 45). The isotopic enrichment of lactate with [13C]lactate was measured by gas chromatography-mass spectrometry in plasma samples deproteinized with sulfosalicylic acid, as previously described (24).Calculations.
Myocardial oxygen consumption
(MO2) was calculated by multiplying
the arterial-coronary sinus difference in oxygen content by CBF. The
rate of FFA uptake (µmol/min) was calculated as: CPF × [FFA]a × ([3H]oleatea
[3H]oleatecs)/[3H]oleatea,
where CPF is the coronary plasma flow [taken as CBF × (1
hematocrit)], [FFA]a is the arterial FFA
concentration (µmol/ml), and [3H]oleate is the
concentration of [3H]oleate in arterial (a) and coronary
sinus (cs) plasma expressed as dpm per milliliter (12).
The rate of FFA oxidation (µmol/min) was calculated as: CPF × ([3H2O]cs
[3H2O]a)/([3H]oleatea/[FFA]cs),
where [3H2O] was the concentration of
[3H]water in the plasma expressed as dpm per milliliter
(12). The rate of glucose uptake was calculated as the
arterial-coronary sinus concentration difference times the CBF, and the
rate of glucose oxidation was calculated from
[14C]glucose as: CBF × ([14CO2]cs
[14CO2]a)/([14C]glucosea/[glucose]a)
as previously described (7, 45). The rate of lactate
uptake (µmol/min) was calculated as: [a] × CBF × ([13C]lactate Ea [a]
[13C]lactate Ecs
[cs])/([13C]lactate Ea [a]) where
[13C]lactate E is the fraction of the lactate in the
blood that is enriched with [1-13C]lactate, as determined
by gas chromatography-mass spectrometry analysis and corrected for
background enrichment; [a] and [cs] are the concentrations of
lactate (µmol/ml) in arterial and coronary sinus blood, respectively
(7, 24). It is known that the tracer-measured lactate uptake
matches total lactate oxidation by the heart, as previously
demonstrated in studies where ~100% of [1-14C]lactate
tracer taken up by the heart is immediately decarboxylated and released
in the venous circulation as 14CO2
(7). The lactate uptakes measured with
[U-13C]lactate and [1-14C]lactate are
equivalent (24). [13C]lactate
tracer-measured lactate output was calculated as the difference between
tracer-measured lactate uptake and the net lactate uptake
arterial-coronary sinus difference of total lactate times CBF
(46). The output quantifies the rate of nonoxidative glycolysis of endogenous and exogenous glucose. In all of the calculations described above, CBF was assumed as double the mean flow
measured in the LcxCBF (28, 29). The percentage of the M
O2 resulting from the oxidation of
glucose, lactate, and FFA was calculated by multiplying the respective
rates of oxidation (measured with [14C]glucose,
[13C]lactate, and [3H]oleate) times 6.0, 3.0, and 24.5 µmol O2 /µmol substrate, respectively, and dividing it by the M
O2
(expressed in µmol/min). For lactate it was assumed that 100% of the
[13C]lactate taken up by the heart is oxidized to
CO2 (7).
Enzyme activities and metabolic intermediates. To measure the activities of each of the enzymes GAPDH, ACC, and MCD, ~200 mg of frozen cardiac tissue were used. GAPDH activity was assessed in the reverse direction using the procedure described by Molina y Vedia et al. (23) provided by Boehringer Mannheim (assay instruction 5178) with a homogenization procedure modified from Knight et al. (15).
ACC activity was determined in the presence and absence of maximally stimulating citrate (10 mM) as described by Dyck et al. (5). Briefly, 200 mg of tissue were homogenized and dialyzed. Dialysate (25 µl) was reacted in 60 mM Tris acetate, pH 7.5, 1 mg/ml BSA, 1.32 µMProtocol.
Experiments were conducted in conscious dogs placed on the laboratory
table after overnight fasting. In all of the 17 dogs, hemodynamics were
recorded, and the isotopic tracers [9,10-3H]oleate,
[U-14C]glucose, and
L-[1-13C]lactate were infused continuously
for the entire duration of the experiment through a peripheral vein.
[3H]oleate bound to albumin was infused at a rate of 0.7 µCi/min. The bolus-continuous infusion method (7, 46)
was used for [14C]glucose (20 µCi + 0.3 µCi/min)
and for [13C]lactate (600 µmol + 700 µmol/h).
Heart rate was maintained constant throughout the experiment by pacing
the heart at 130 ± 2 beats/min. After 40 min of tracer infusion,
control paired blood samples were withdrawn from the aorta and coronary
sinus. At this point, dogs were divided into two groups. Nine dogs
received a bolus of 35 mg/kg of
N-nitro-L-arginine methyl ester
(L-NAME) intravenously. Immediately after
L-NAME administration, a bolus of 5,000 IU of heparin was given intravenously, and an infusion of 10% Intralipid plus 125 IU/ml
heparin was started at the rate of 0.006 ml · kg
1 · min
1 and
continued until the end of the experiment. This low infusion rate was
chosen since we found that, at higher rates, the cocktail of Intralipid
plus heparin caused an increase in arterial FFA concentration that
sometimes exceeded the control values. Paired arterial and coronary
sinus blood samples were withdrawn at 30, 60, 90, and 120 min after
L-NAME. A second bolus of 35 mg/kg of L-NAME
was given immediately after collection of blood samples at 90 min. A
second group of eight dogs (control group) underwent a continuous
intravenous infusion of synthetic (Sigma) human ANG II at the rate of
20-40 ng · kg
1 · min
1,
started immediately after withdrawal of control blood samples. The rate
of ANG II infusion was adjusted initially to obtain a mean arterial
pressure (MAP) of ~150 mmHg, matching the hypertension caused by NOS
inhibition, and then was maintained constant during the remaining part
of the experiment. Heparin and Intralipid were infused, and blood
samples were collected following the same protocol described for the
group receiving L-NAME. After withdrawal of the last blood
samples, all of the dogs were anesthetized with 30 mg/kg pentobarbital
sodium intravenously, intubated, and ventilated. The fifth intercostal
space was opened rapidly to harvest a tissue sample from the heart. A
large (~10 g) biopsy was obtained by punching a hole through the LV
anterior free wall with a stainless steel "cork borer" (2 cm ID)
while the heart was beating. The harvested tissue was immediately
freeze-clamped with tongs precooled in liquid nitrogen. The total time
from anesthesia to tissue harvesting was <3 min, and we found that,
during this time, the dog was not hypotensive. The time from tissue
harvesting to clamping between tongs was ~3-4 s. This approach
was established by previous studies that combined measurements of
metabolic changes in vivo with enzyme assays in vitro (15, 32,
35).
Statistical analysis. Data are presented as means ± SE. Statistical analysis was performed by employing commercially available software (Sigma Stat 2.01). Changes in hemodynamics and in rates of substrate uptake and oxidation over time were tested by one-way ANOVA for repeated measurements followed by Dunnett's test. Tukey's test was also used to obtain multiple comparisons. Changes resulting from L-NAME and ANG II infusion were compared and tested for differences by using two-way ANOVA followed by Tukey's test.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Hemodynamics.
As shown in Fig. 1, comparable changes
occurred in the main hemodynamic parameters after L-NAME
and during ANG II infusion. Additional hemodynamic data are reported in
Table 1. As described in previous
studies, mean blood flow in the LcxCBF, MAP, and LV systolic pressure
(LVSP) increased significantly after NOS inhibition (Fig. 1). By
infusing ANG II, it was possible to mimic with a good approximation
changes in LcxCBF and MAP caused by L-NAME. However, LVSP
gradually declined after 30 min of NOS blockade, and, at 90 and 120 min, it was significantly lower than LVSP measured in dogs with ANG II
infusion. Despite the fact that LV end-diastolic diameter did not
change significantly after L-NAME or during ANG II infusion
(Table 1), dp/dtmax (Fig. 1), fractional
shortening of LV diameter, and pressure-diameter area (Table 1) were
significantly lower in dogs with NOS blockade compared with the control
group.
|
|
Cardiac metabolism.
The continuous infusion of Intralipid plus heparin could not prevent a
transient decrease in arterial FFA concentration after L-NAME (Fig. 2). However, FFA
concentration did not fall below 0.45 ± 0.05 mmol/l, and the
values at 120 min did not differ significantly from control. A
significant decrease in circulating FFA was observed also at 90 and 120 min of ANG II infusion. Arterial glucose concentration did not change
significantly in either of the two groups, whereas lactate increased
significantly in dogs receiving ANG II and L-NAME, with a
significant difference between the two groups at 90 min (Fig. 2). The
arterial specific radioactivity of [3H]oleate and
[14C]glucose (radioactive/total substrate in dpm/mmol),
and the lactate enrichment with [13C]lactate (fraction of
lactate in the blood that is enriched with [1-13C]lactate) are shown in Fig.
3 as a function of time. These plots demonstrate that there was isotopic steady state and no significant differences between the two groups for [14C]glucose
specific activity and for [13C]lactate enrichment from 30 to 120 min of the protocol. There was an increase in the specific
activity of [3H]oleate over time, which was more rapid in
the L-NAME group. As expected, this increase corresponded
to the time points at which the plasma concentration of total FFA fell
significantly (see Fig. 2). As shown in Fig.
4, NOS inhibition and ANG II infusion caused almost superimposable increases in
MO2. Despite matched levels of
oxygen consumption, changes in cardiac oxidation of FFA and glucose
evolved differently in the two groups of dogs (Fig. 4). Between 30 and
60 min after L-NAME, FFA oxidation fell significantly by
~28% compared with control, reaching the minimum value at 60 min.
This decrease was only transient, since FFA oxidation returned to
control values at 90 min, but it fell again at 120 min after
administration of the second dose of L-NAME. Cardiac oxidation of glucose showed a marked and stable augmentation, and a
significant difference was found at 30 and 60 min between the 400%
increase and the 270% increase occurring after L-NAME and
during ANG II infusion, respectively. Total cardiac lactate uptake
increased significantly by ~130% in both groups (Fig. 4). Changes in
FFA and glucose oxidation were not paralleled by significant changes in
uptake, except for a significant increase in glucose uptake at 60 min
after L-NAME (Fig. 5). Also,
cardiac production of lactate (as measured with
[13C]lactate tracer) did not change significantly in
either of the two groups (Fig. 5). The sum of the oxidation of glucose,
lactate, and FFA varied between 104 and 121% of the measured
M
O2 and did not change significantly
over the time course of the protocol, and no significant differences
were found between the two groups (Table
2).
|
|
|
|
|
Enzyme activities and metabolic products in cardiac tissue.
The activities of GAPDH, ACC, and MCD and the concentrations of
malonyl-CoA, free CoA, and acetyl-CoA are reported in Table 3. No significant differences were found
between the two groups, except for the levels of free CoA and
acetyl-CoA, which resulted in higher levels in
L-NAME-treated hearts.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The first aim of the present study was to measure changes in the
metabolism of cardiac substrates after inhibition of NO synthesis. Our
results show that an acute reduction of NO production causes a
reduction in FFA oxidation and an increase in glucose oxidation by
cardiac muscle. The function of NO as a metabolic modulator has been
the object of intense investigations. Endogenous NO, likely generated
by the microvascular endothelium in which eNOS is preponderant,
inhibits oxygen consumption in heart and other organs (17, 19,
39). We now provide evidence that NO plays an important role in
the complex control that directs cardiac metabolism toward preferential
oxidation of FFA. A notorious effect of acute NOS blockade is the
marked elevation of arterial blood pressure with a consequent increase
in LV afterload. This effect was present also in our study and could
have been itself a cause of cardiac metabolic alterations. To control
for this, we used for comparison a control group of dogs in which
arterial blood pressure was elevated by a continuous infusion of ANG II
to match the increase in cardiac workload and
MO2. This comparison proved that the
reduction in FFA oxidation in the L-NAME group was
specifically associated with NOS inhibition. On the other hand,
myocardial glucose oxidation increased during ANG II infusion, but to a
significantly lesser extent compared with the increase observed after
L-NAME. Finally, values of lactate uptake/oxidation in the
groups were almost overlapping, indicating that this phenomenon was not
specifically related to NO availability, but, more in general, to
elevations in cardiac workload.
The second aim of our study was to test whether alterations in myocardial substrate utilization, after NOS inhibition, were the result of changes in the activity of some key enzymes involved in FFA and carbohydrate metabolism. ACC, MCD, and GAPDH activities in L-NAME- and ANG II-treated hearts were not significantly different. And no significant difference was found between groups in the myocardial concentrations of malonyl-CoA, the physiological inhibitor of FFA transport in mitochondria. Therefore, the metabolic alterations caused by NOS inhibition likely did not involve these key enzymes regulating FFA and glucose utilization. On the other hand, tissue acetyl-CoA content was about twofold higher in the L-NAME group, which is consistent with studies in swine where the rate of myocardial carbohydrate oxidation was increased with either dichloroacetate (35) or dobutamine (11). Finally, the increased content of free CoA by ~150% was consistent with a reduced rate of FFA oxidation and a consequent fall in the synthesis of long-chain fatty acyl-CoA and the release of free CoA after L-NAME.
The present findings are in agreement with the conclusions that we drew from our previous studies in conscious dogs in which we blocked NOS synthesis (28, 29) and reversed metabolic changes by infusing an NO donor (29). An important difference, however, consists in the separate measurements of oxidation and uptake allowed, in the present case, by the use of isotopes. To our knowledge, this is the first study in which cardiac metabolism was explored by employing, simultaneously, three different isotopes labeling the three main substrates consumed by the heart. To rule out possible artifacts resulting from isotope infusion and blood sampling, we performed pilot experiments in conscious dogs not subjected to any pharmacological intervention and did not observe any significant changes in substrate oxidation over a period of 3 h (data not shown). The differentiation between uptake and oxidation revealed that these two events did not necessarily vary in parallel after NOS inhibition. FFA uptake, for instance, did not change significantly after L-NAME, despite a decrease in oxidation by almost 30% that occurred at 60 and 120 min. In our previous study, we hypothesized a similar dissociation between FFA uptake and oxidation based on the calculation of cardiac respiratory quotient. However, in that study, FFA uptake fell markedly at 60 min after administration of NOS inhibitor. Such a phenomenon was likely because of a significant drop in the arterial concentration of FFA that we observed after nitro-L-arginine (NLA) administration. To avoid the confounding effects of fluctuations in FFA availability that could have specifically altered myocardial metabolism, in the present set of experiments we tried to clamp the level of circulating FFA by continuously infusing Intralipid plus heparin. Although we did not obtain a complete clamp, FFA concentration remained well above the low value of 0.25 mmol/l that we observed in our earlier studies with NOS inhibition (28, 29). The decrease in FFA oxidation found in the present investigation cannot be explained simply by a fall in plasma fatty acid levels and uptake. In fact, arterial FFA concentration was constantly lower than control from 30 to 90 min after the first bolus of L-NAME, even though FFA oxidation was significantly reduced at 60 min only. Furthermore, at 120 min, after the second bolus of L-NAME, FFA concentration increased and was not different from control. Nonetheless, oxidation surprisingly fell again by ~30% despite the fact that hemodynamics did not change further. We have no data to explain this phenomenon. It seems that the depression of FFA oxidation was reversible and occurred only at high concentrations of NOS inhibitor. Unfortunately, we could collect a large cardiac biopsy only at the end of the experiment, and our measurement of enzyme activities could not follow the entire dynamics of metabolic change.
A key physiological modulator of FFA oxidation is cytosolic malonyl-CoA, the concentration of which is largely determined by the activities of ACC and MCD. None of these factors changed significantly in L-NAME-treated dogs compared with the control group. Based on these data, it is only possible to speculate that NO is one of the regulatory factors of the FFA oxidation pathway, but its action is exerted at a site of regulation that remains unknown. Alternatively, it is possible that NO reversibly modulates the activities of ACC and/or MCD and/or other enzymes of the FFA metabolic pathway in vivo, but this function is lost in tissue homogenates.
After L-NAME (60 min), a significant increase in glucose oxidation was paralleled by a significant increase in uptake. This finding is consistent with our previous studies documenting a marked increase in glucose uptake associated with diminished NO synthesis in failing hearts (28), in normal hearts treated with NLA (28-29), and in hearts isolated from eNOS knockout mice (40). It is also consistent with a decrease in glucose uptake found in isolated hearts after infusion of 8-bromo-cGMP (8-BrcGMP; see Refs. 4 and 40), a putative second messenger of NO. Endogenous NO likely inhibits glucose uptake via a mechanism that remains undetermined. Glucose entry in cells is strongly dependent on the incorporation of specific transporters in the cell membrane (34). Depre et al. (4) hypothesized a direct inhibition by NO, via the second messenger cGMP, on glucose transporters in myocytes. In support of this hypothetical mechanism, Tada et al. (40) found that 1-H-(1,2,4)oxadiazolo[4,3-a]quinoxaline-1-one, an inhibitor of cGMP synthesis, markedly stimulates glucose uptake in isolated mouse hearts. Specific experiments at the cellular level will be necessary to elucidate the possible modulatory action of cGMP or NO on the function of glucose transporters, specifically on the translocation and kinetics of GLUT-1 and GLUT-4. Interestingly, in our experiments, the effect of NOS inhibition on cardiac glucose uptake was transient, limited to one time point and not so pronounced as observed in isolated hearts. This could be because of the incomplete inhibition of NO synthesis by pharmacological blockade of NOS compared with the complete shut off in eNOS knockouts. It also could be due in part to the competition by other substrates available in intact animal preparations. In the present study, however, glucose oxidation was constantly higher than control after NOS inhibition, although it tended to decrease over time so that no significant differences vs. the ANG II group were found at 90 and 120 min. It is noteworthy that, even if less pronounced, there was a significant increase in glucose oxidation also during ANG II infusion. This could have been simply because of the increased metabolic demand, but we cannot exclude additional mechanisms, such as an ANG II-mediated elevation of intracellular pH (9) with consequent phosphofructokinase activation.
Glucose oxidation includes the following two phases: the first one, cytosolic, consists of the glycolytic pathway with pyruvate as the end product; the second one leads to complete oxidation of pyruvate in mitochondria. We tested the hypothesis that physiological concentrations of NO exert a tonic inhibition on GAPDH, a key enzyme of the glycolytic pathway. This hypothesis was based on the previous findings that pharmacological or pathological concentrations of NO inhibit GAPDH in vivo and in vitro (23, 49). We did not find significant differences between the two groups. Therefore, it is likely that GAPDH function is unaffected by changes in the physiological concentration of NO. As for the enzymes regulating FFA oxidation, an alternative hypothesis is that the action of physiological amounts of NO on GAPDH is rapidly reversible in vivo; thus, it was lost in tissue homogenates, in which spontaneous production of NO did not occur, exogenous NO was not added, and tissutal scavengers could have rapidly neutralized NO bound to thiol groups. Another important limiting step shared by glucose and lactate oxidation is the irreversible dehydrogenation of pyruvate by the pyruvate dehydrogenase enzyme complex in mitochondria (38). We did not test this enzyme, but similar increases in lactate oxidation in both groups suggested that the activity of pyruvate dehydrogenase was not stimulated exclusively by reduced availability of NO but rather by generic elevations in ventricular afterload. Other investigators have found reduced glycolysis, but no changes in pyruvate dehydrogenase activity, in isolated hearts during infusion of 8-BrcGMP (4). It is likely, therefore, that the inhibitory action exerted by NO on myocardial glucose metabolism is limited to the transmembrane transport and, perhaps, to one or more enzymes of glycolysis other than GAPDH. For instance, it has been shown that NO can inhibit phosphofructokinase in pancreatic islets (42). The release of tonic inhibition of myocardial phosphofructokinase after NOS blockade could thus be a candidate for the mechanism of stimulated glucose metabolism. Future studies specifically exploring the role of NO in the control of the glycolytic pathway, in well-controlled conditions in vitro, will be necessary to answer these open questions. It is also important to note that the [13C]lactate tracer-measured output of lactate, an index of anaerobic glycolysis, did not change significantly during the experiments, indicating a preserved match between oxygen demand and supply.
In conditions such as failure or pathological hypertrophy, the utilization of glucose by cardiac muscle increases (20, 25, 28, 47, 48). NO production is impaired in both of these pathological states (21, 28). In the light of the present findings, it is possible to hypothesize that NO plays a role in the metabolic changes typical of diseased myocardium. Understanding this role assumes a particular importance if, as speculated by some authors on the basis of recent data, the performance of the failing heart may be affected by the type of substrate oxidized (30). Furthermore, solid evidence supports the concept that, during cardiac ischemia, the stimulation of glucose, rather than FFA oxidation, is advantageous in preserving function and facilitating recovery of the ischemic myocardium (20, 27, 41). On the other hand, evidence also exists that the recovery of stunned myocardium is potentiated by oxidation of FFA (43). Like failure and hypertrophy, cardiac ischemia and stunning are pathological conditions in which the synthesis of NO is altered (8). In these cases, the changed availability of NO might be in part responsible for metabolic alterations and functional consequences.
Limitations of the study.
A number of limitations should be pointed out. Our study was aimed at
measuring changes in cardiac metabolism in conscious animals. The heart
was therefore studied under optimal conditions of perfusion,
oxygenation, and supply of the entire range of physiological substrates. This represented a remarkable advantage relative to isolated heart preparations. A disadvantage consisted of the
impossibility to control simultaneously numerous variables, including
substrate concentration, cardiac load, and neurohumoral factors that
might have been responsible for part of the myocardial metabolic
alteration. For instance, the switch in substrate oxidation could have
been the result of the release of insulin triggered in pancreatic
-cells by the arginine analog L-NAME (44).
This was not likely the case, however, since in our previous study
(29) we found that arterial insulin levels were rather
decreased after NOS inhibition. The hemodynamic changes occurring
after L-NAME were in part mimicked, in the control group,
by infusion of ANG II, since we found this vasopressor well tolerated
by conscious dogs, easy to titrate, and rapidly reversible. We could
indeed reproduce the increase in MAP resulting from NOS blockade,
although ANG II enhanced or preserved cardiac contractility, as
indicated by the differences in dP/dtmax, LV
diameter fractional shortening, and pressure-diameter areas between the
two groups. We did not measure ventricular volume and aortic flow, but
cardiac output was likely unchanged (13). However, despite
these differences in hemodynamics, the groups presented a similar
increase in oxygen consumption, i.e., in metabolic demand. Coronary
flow increased in both groups in response to the augmented metabolic
demand. Another limitation regards the calculated
M
O2 that systematically exceeded the
measured M
O2 (Table 2). This
suggests that there was an overestimation in the rate of oxidation of
one or more of the substrates for both groups. This could be because of
a greater fractional extraction and oxidation of oleate compared with
other fatty acids in plasma or because of equilibration between
intracellular lactate and pyruvate and overestimation of exogenous
lactate oxidation with the [13C]lactate tracer (37,
38). Even with this limitation, the calculated ratio did not
change significantly over time, despite marked alterations in the rate
of substrate oxidation. A limitation regarding the calculation of
glucose uptake was based on measurements of the arteriovenous
difference in the total concentration. The low sensitivity of this
method could explain the large SEs reported for glucose uptake. The
best alternative would have been the use of two different isotopic
tracers to label glucose, one to measure uptake and the other to
measure oxidation. This method would also have allowed precise
estimates of glycolytic flux. Unfortunately, we already infused all of
the three isotopes universally used to label cardiac substrates.
Finally, the negative findings relative to enzyme activities could have
been caused simply by the techniques used and may not necessarily
reflect regulatory processes that occurred in vivo. Unfortunately, we
could adopt only the method presently available; metabolic changes
observed in vivo were correlated with enzyme activities tested in
biopsied tissue ex vivo, under nonphysiological conditions. We measured
maximal activities of the enzymes in the presence of saturating
concentrations of substrate, and we cannot exclude that the affinity
for the substrate, indicated by the Michaelis-Menter constant
(Km), was significantly different between the
two groups. Further studies in vitro will be necessary to test the
effects of NO/cGMP on the Km of purified enzymes.
![]() |
ACKNOWLEDGEMENTS |
---|
This study was supported by the National Heart, Lung, and Blood Institute Grant RO1 HL-62573 (F. A. Recchia), in part by PO1 HL-43023 (T. H. Hintze), RO1 HL-58653, and HL-64848 (W. C. Stanley), and by a grant from the Canadian Institutes of Health (G. D. Lopaschuk).
![]() |
FOOTNOTES |
---|
Address for reprint requests and other correspondence: F. A. Recchia, Dept of Physiology, BSB 622, New York Medical College, Valhalla, NY 10595 (E-mail: fabio_recchia{at}nymc.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 29 June 2001; accepted in final form 30 August 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Awan, AA,
and
Saggerson ED.
Malonyl-CoA metabolism in cardiac myocytes and its relevance to the control of fatty acid oxidation.
Biochem J
295:
61-66,
1993[ISI][Medline].
2.
Burkhoff, D,
Weiss RG,
Schulman SP,
Kalil-Filho R,
Wannenburg T,
and
Gerstenblith G.
Influence of metabolic substrate on rat heart function and metabolism at different coronary flows.
Am J Physiol Heart Circ Physiol
261:
H741-H750,
1991
3.
Corkey, BE.
Analysis of acyl-coenzyme A esters in biological samples.
Methods Enzymol
166:
55-70,
1988[ISI][Medline].
4.
Depre, C,
Gaussin V,
Ponchaut S,
Fischer Y,
Vanoverschelde JL,
and
Hue L.
Inhibition of myocardial glucose uptake by cGMP.
Am J Physiol Heart Circ Physiol
274:
H1443-H1449,
1998
5.
Dyck, JRB,
Barr AJ,
Barr RL,
Kolattukudy PE,
and
Lopaschuk GD.
Characterization of cardiac malonyl-Coa decarboxylase and its putative role in regulating fatty acid oxidation.
Am J Physiol Heart Circ Physiol
275:
H2122-H2129,
1998
6.
Gertz, EW,
Wisneski JA,
Neese RA,
Bristow JD,
Searle GL,
and
Hanlon JT.
Myocardial lactate metabolism: evidence of lactate release during net chemical extraction in man.
Circulation
63:
1273-1279,
1981[Abstract].
7.
Gertz, EW,
Wisneski JA,
Stanley WC,
and
Neese RA.
Myocardial substrate utilization during exercise in humans: dual carbon-labeled carbohydrate isotope experiments.
J Clin Invest
82:
2017-2025,
1988[ISI][Medline].
8.
Giraldez, RR,
Panda A,
Xia Y,
Sanders SP,
and
Zweier JL.
Decreased nitric-oxide synthase activity causes impaired endothelium-dependent relaxation in the post-ischemic heart.
J Biol Chem
272:
21420-21426,
1997
9.
Gunasegaram, S,
Haworth RS,
Hearse DJ,
and
Avkiran M.
Regulation of sarcolemmal Na(+)/H(+) exchanger activity by angiotensin II in adult rat ventricular myocytes: opposing actions via AT(1) versus AT(2) receptors.
Circ Res
85:
919-930,
1999
10.
Hall, JL,
Lopaschuk GD,
Pizzurro RD,
Bringas J,
and
Stanley WC.
Increased cardiac fatty acid uptake with dobutamine infusion in swine is accompanied by a decrease in malonyl CoA levels.
Cardiovasc Res
32:
879-885,
1996[ISI][Medline].
11.
Hall, JL,
Stanley WC,
Lopaschuk GD,
Wisneski JA,
Pizzurro RD,
Hamilton CD,
and
McCormack JG.
Impaired pyruvate oxydation but normal glucose uptake in diabetic pig heart during dobutamine-induced work.
Am J Physiol Heart Circ Physiol
271:
H2320-H2329,
1996
12.
Havel, RJB,
Pernow B,
and
Jones NL.
Uptake and release of free fatty acids and other metabolites in the legs of exercising men.
J Appl Physiol
23:
90-99,
1967
13.
Heyndrickx, GR,
Boettcher DH,
and
Vatner SF.
Effects of angiotensin, vasopressin, and methoxamine on cardiac function and blood flow distribution in conscious dogs.
Am J Physiol
231:
1579-1587,
1976[ISI][Medline].
14.
Kim, YS,
and
Kolattukudy PE.
Purification and properties of malonyl-CoA decarboxylase from rat liver mitochondria and its immunological comparison with the enzymes from rat brain, heart, and mammary gland.
Arch Biochem Biophys
190:
234-246,
1978[ISI][Medline].
15.
Knight, RJ,
Kofoed KF,
Schelbert HR,
and
Buxton DB.
Inhibition of glyceraldehyde-3-phosphate dehydrogenase in post-ischemic myocardium.
Cardiovasc Res
32:
1016-1023,
1996[ISI][Medline].
16.
Korvald, C,
Elvenes OP,
and
Myrmel T.
Myocardial substrate metabolism influences left ventricular energetics in vivo.
Am J Physiol Heart Circ Physiol
278:
H1345-H1351,
2000
17.
Laycock, SK,
Vogel T,
Forfia PR,
Tuzman J,
Xu X,
Ochoa M,
Thompson CI,
Nasjletti A,
and
Hintze TH.
Role of nitric oxide in the control of renal oxygen consumption and the regulation of chemical work in the kidney.
Circ Res
82:
1263-1271,
1998
18.
Liedtke, AJ.
Alterations of carbohydrate and lipid metabolism in the acutely ischemic heart (Abstract).
Prog Cardiovasc Dis
23:
321,
1981[ISI][Medline].
19.
Loke, KE,
McConnell PI,
Tuzman JM,
Shesely EG,
Smith CJ,
Stackpole CJ,
Thompson CI,
Kaley G,
Wolin MS,
and
Hintze TH.
Endogenous endothelial nitric oxide synthase-derived nitric oxide is a physiological regulator of myocardial oxygen consumption.
Circ Res
84:
840-845,
1999
20.
Lopaschuk, GD,
Belke DD,
Gamble J,
Itoi T,
and
Schonekess BO.
Regulation of fatty acid oxidation in the mammalian heart in health and disease.
Biochim Biophys Acta
1213:
263-276,
1994[ISI][Medline].
21.
MacCarthy, PA,
and
Shah AM.
Impaired endothelium-dependent regulation of ventricular relaxation in pressure-overload cardiac hypertrophy.
Circulation
101:
1854-1860,
2000
22.
Mjøs, OD.
Effect of free fatty acids on myocardial function and oxygen consumption in intact dogs.
J Clin Invest
50:
1386-1389,
1971[ISI][Medline].
23.
Molina y Vedia, L,
McDonald B,
Reep B,
Brune B,
Di Silvio M,
and
Billiar TR.
Lapetina EG Nitric oxide-induced s-nitrosylation of glyceraldehyde-3-phosphate dehydrogenase inhibits enzymatic activity and increases endogenous ADP-rybosilation.
J Biol Chem
267:
24929-24932,
1992
24.
Neese, RE,
Gertz EW,
Wisneski JA,
Gruenke LD,
and
Craid LC.
A stable isotope technique for investigating lactate metabolism in humans.
Biomed Mass Spectrom
10:
458-462,
1983[ISI][Medline].
25.
Olson, RE.
Myocardial metabolism in congestive heart failure.
J Chron Dis
9:
442-447,
1959[Medline].
26.
Opie, LH.
The Heart: Physiology and Metabolism. New York: Raven, 1991, p. 208-276.
27.
Pepine, CJ,
and
Wolff AA.
A controlled trial with a novel anti-ischemic agent, ranolazine, in chronic stable angina pectoris that is responsive to conventional antianginal agents. Ranolazine study group.
Am J Cardiol
84:
46-50,
1999[ISI][Medline].
28.
Recchia, FA,
McConnell PI,
Bernstein RD,
Vogel TR,
Xu XB,
and
Hintze TH.
Reduced nitric oxide production and altered myocardial metabolism during the decompensation of pacing-induced heart failure in the conscious dog.
Circ Res
83:
969-979,
1998
29.
Recchia, FA,
McConnell PI,
Loke KE,
Xu XB,
Ochoa M,
and
Hintze TH.
Nitric oxide controls cardiac substrate utilization in the conscious dog.
Cardiovasc Res
44:
325-332,
1999[ISI][Medline].
30.
Sabbah, HN,
Mishima T,
Suzuki G,
Chaudhry P,
Nass O,
Blackburn B,
Huang BL,
and
Stanley WC.
Ranolazine, a partial fatty acid oxidation inhibitor, improves left ventricular function in dogs with heart failure but not in normals (Abstract).
Circulation
102, SupplII:
721,
2000[ISI].
31.
Saddik, M,
Gamble J,
Witters LA,
and
Lopaschuk GD.
Acetyl CoA carboxylase regulation of fatty acid oxidation in the heart.
J Biol Chem
286:
25836-25845,
1993.
32.
Schoder, H,
Knight RJ,
Kofoed KF,
Schelbert HR,
and
Buxton DB.
Regulation of pyruvate dehydrogenase activity and glucose metabolism in post-ischemic myocardium.
Biochim Biophys Acta
1406:
62-72,
1998[ISI][Medline].
33.
Simonsen, S,
and
Kjekshus JK.
The effect of free fatty acids on myocardial oxygen consumption during atrial pacing and catecholamine infusion in man.
Circulation
58:
484-941,
1978[Abstract].
34.
Slot, JW,
Geuze HJ,
Gigengack S,
James DE,
and
Lienhard GE.
Translocation of the glucose transporter GLUT4 in cardiac myocytes of the rat.
Proc Natl Acad Sci USA
88:
7815-7819,
1991[Abstract].
35.
Stanley, WC,
Hernandez LA,
Spires DA,
Bringas J,
Wallace S,
and
McCormack JG.
Pyruvate dehydrogenase activity and malonyl-CoA levels in normal and ischemic swine myocardium: effects of dichloroacetate.
J Mol Cell Cardiol
28:
905-914,
1996[ISI][Medline].
36.
Stanley, WC,
Lopaschuk GD,
Hall JL,
and
McCormack JG.
Regulation of myocardial carbohydrate metabolism under normal and ischaemic conditions.
Cardiovasc Res
33:
243-257,
1997[ISI][Medline].
37.
Stanley, WC,
and
Lehman SL.
A model for measurement of lactate turnover with isotopic tracers.
Biochem J
256:
1035-1038,
1988[ISI][Medline].
38.
Stanley, WC,
and
Lehman SL.
Calculation of lactate disappearance with isotopic tracers using tissue lactate specific radioactivity (Abstract).
Biochem J
259:
935,
1989[ISI][Medline].
39.
Suto, N,
Mikuniya A,
Okubo T,
Hanada H,
Shinozaki N,
and
Okumura K.
Nitric oxide modulates cardiac contractility and oxygen consumption without changing contractile efficiency.
Am J Physiol Heart Circ Physiol
275:
H41-H49,
1998
40.
Tada, H,
Thompson CI,
Recchia FA,
Loke KE,
Ochoa M,
Smith CJ,
Shesely EG,
Kaley G,
and
Hintze TH.
Myocardial glucose uptake is regulated by nitric oxide via endothelial nitric oxide synthase in Langendorff mouse heart.
Circ Res
86:
270-274,
2000
41.
Taniguchi, M,
Wilson C,
Hunter CA,
Pehowich DJ,
Clanachan AS,
and
Lopaschuk GD.
Dichloroacetate improves cardiac efficiency after ischemia independent of changes in mitochondrial proton leak.
Am J Physiol Heart Circ Physiol
280:
H1762-H1769,
2001
42.
Tsuura, Y,
Ishida H,
Shinomura T,
Nishimura M,
and
Seino Y.
Endogenous nitric oxide inhibits glucose-induced insulin secretion by suppression of phosphofructokinase activity in pancreatic islets.
Biochem Biophys Res Commun
252:
34-38,
1998[ISI][Medline].
43.
Van de Velde, M,
Wouters PF,
Rolf N,
Van Aken H,
Flameng W,
and
Vandermeersch E.
Long-chain triglycerides improve recovery from myocardial stunning in conscious dogs.
Cardiovasc Res
32:
1008-1015,
1996[ISI][Medline].
44.
Weinhaus, AJ,
Poronnik P,
Tuch BE,
and
Cook DI.
Mechanisms of arginine-induced increase in cytosolic calcium concentration in the beta-cell line NIT-1.
Diabetologia
40:
374-382,
1997[ISI][Medline].
45.
Wisneski, JA,
Gertz EW,
Neese RA,
and
Mayr M.
Myocardial metabolism of free fatty acids: studies with 14C-labeled substrates in humans.
J Clin Invest
79:
359-366,
1987[ISI][Medline].
46.
Wisneski, JA,
Stanley WC,
Neese RA,
and
Gertz EW.
Effects of acute hyperglycemia on myocardial glycolytic activity in humans.
J Clin Invest
85:
1648-1656,
1990[ISI][Medline].
47.
Wittels, B,
and
Spann JF, Jr.
Defective lipid metabolism in the failing heart.
J Clin Invest
47:
1787-1794,
1968[ISI][Medline].
48.
Zhang, J,
Duncker DJ,
Ya X,
Zhang Y,
Pavek T,
Wei H,
Merkle H,
Ugurbil K,
From AH,
and
Bache RJ.
Effect of left ventricular hypertrophy secondary to chronic pressure overload on transmural myocardial 2-deoxyglucose uptake. A 31P NMR spectroscopic study.
Circulation
92:
1274-1283,
1995
49.
Zhang, J,
and
Snyder SH.
Nitric oxide stimulates auto-ADP-ribosylation of glyceraldehyde-3-phosphate dehydrogenase.
Proc Natl Acad Sci USA
89:
9382-9385,
1992[Abstract].