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Am J Physiol Heart Circ Physiol 287: H351-H362, 2004. First published March 4, 2004; doi:10.1152/ajpheart.00983.2003
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Impact of low-flow ischemia on substrate oxidation and glycolysis in the isolated perfused rat heart

Steven G. Lloyd,1 Peipei Wang,1 Huadong Zeng,2 and John C. Chatham1,2,3

1Division of Cardiovascular Disease, Department of Medicine, 2Comprehensive Cancer Center, and 3Department of Physiology and Biophysics and Clinical Nutrition Research Center, University of Alabama at Birmingham, Birmingham, Alabama 35294-0005

Submitted 20 October 2003 ; accepted in final form 25 February 2004


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Interventions that stimulate carbohydrate oxidation appear to be beneficial in the setting of myocardial ischemia or infarction. However, the mechanisms underlying this protective effect have not been defined, in part because of our limited understanding of substrate utilization under ischemic conditions. Therefore, we used 1H and 13C NMR spectroscopy to investigate substrate oxidation and glycolytic rates in a global low-flow model of myocardial ischemia. Isolated male Sprague-Dawley rat hearts were perfused for 30 min under conditions of normal flow (control) and low-flow ischemia (LFI, 0.3 ml/min) with insulin and 13C-labeled lactate, pyruvate, palmitate, and glucose at concentrations representative of the physiological fed state. Despite a ~50-fold reduction in substrate delivery and oxygen consumption, oxidation of all exogenous substrates plus glycogen occurred during LFI. Oxidative metabolism accounted for 97% of total calculated ATP production in the control group and ~30% in the LFI group. For controls, lactate oxidation was the major source of ATP; however, in LFI, this shifted to a combination of oxidative and nonoxidative glycogen metabolism. Interestingly, in the LFI group, anaplerosis relative to citrate synthase increased sevenfold compared with controls. These results demonstrate the importance of oxidative energy metabolism for ATP production, even during very-low-flow ischemia. We believe that the approach described here will be valuable for future investigations into the underlying mechanisms related to the protective effect of increasing cardiac carbohydrate utilization and may ultimately lead to identification of new therapeutic targets for treatment of myocardial ischemia.

nuclear magnetic resonance spectroscopy; energy metabolism; lactate; pyruvate; ATP

THERE HAVE BEEN SIGNIFICANT advances in the management and treatment of individuals suffering from ischemic heart disease during the past 50 years (50), but it remains a major public health problem (1, 2). The success and increasing availability of reperfusion therapies have led to renewed interest in the development of novel cell protection strategies that could be provided immediately before and after revascularization (49). One area that has received increasing attention in recent years is the use of interventions designed to increase myocardial glucose utilization (47). This has led to renewed interest in the beneficial effects of glucose-insulin-potassium (GIK) infusions in the setting of acute myocardial infarction (15, 17). Experimental studies have also demonstrated that pharmacological interventions that stimulate glucose oxidation directly by activation of pyruvate dehydrogenase (PDH) or indirectly by inhibition of fatty acid oxidation result in improved recovery after ischemia and reperfusion (47). Furthermore, inhibitors of fatty acid oxidation (e.g., trimetazadine and ranolazine), which shift the balance from fatty acid to carbohydrate use, have recently been evaluated clinically as anti-ischemic agents (19, 38, 42).

A variety of mechanisms have been proposed to explain the beneficial effects of increasing glucose utilization, the most common of which are as follows: 1) Increasing glucose oxidation at the expense of fatty acid oxidation results in greater "efficiency" in ATP production, because glucose oxidation requires less oxygen consumption per unit ATP produced than fatty acid oxidation. 2) Decreasing fatty acid oxidation and increasing glucose oxidation improve the coupling between glycolysis and glucose oxidation, thereby preventing detrimental proton accumulation (32, 47). However, the majority of data used to support these mechanisms are based on studies where glucose and fatty acids are the only substrates considered. We recently reported that when a physiological carbohydrate mixture including lactate and pyruvate as well as glucose and palmitate was used, a shift from predominantly fatty acid to predominantly carbohydrate oxidation did not change the estimated oxygen consumption in isolated hearts, with consequently no increase in the estimated efficiency of ATP synthesis (29). In the same study, we also found that when lactate, pyruvate, and palmitate are present at physiological concentrations, glucose contributes only 10–25% of oxidative energy production and is thus only a minor contributor to oxidative ATP production (29). This suggests that when other carbohydrate sources are present, the concept of "the coupling of glucose oxidation to glycolysis" may not be relevant.

Another potential limitation of studies into the mechanism(s) underlying the benefit of increasing carbohydrate use is that they have typically focused on the use of zero-flow models of ischemia. However, in humans and animals, data clearly demonstrate that there is a range of blood flows within an infarct zone. In humans with acute myocardial infarction, ~40% of the infarct zone was supplied by collaterals, even in severely hypokinetic or akinetic segments (43). In animal models, collateral flow during acute left anterior descending coronary artery occlusion is 10–20% of baseline flow in the infarct zone (12). It is evident that low-flow ischemia (LFI) represents a metabolic state that is different from no-flow ischemia, in part because the presence of residual flow enables the transport of lactate out of the myocyte, thereby reducing the severity of acidosis compared with no-flow ischemia. This also enables continuation of glycolytic ATP production, leading to a less severe bioenergetic deficit at the time of reperfusion. However, the presence of some flow also indicates a limited supply of oxygen, which could be used for oxidative energy production. Ascuitto et al. (3) reported that when flow was reduced to ~20% of baseline, there was significant oxygen consumption and oxidation of exogenous fatty acids in the isolated perfused neonatal pig heart. In a more severe LFI model in the perfused rat heart, Finegan et al. (18) reported significant oxidation of exogenous glucose. Interestingly, Opie et al. (37) showed that, in vivo, after arterial ligation, there was an increase in glucose oxidation relative to fatty acid oxidation. This suggests that not only was there appreciable substrate oxidation in the ischemic zone, there was also a shift in regulation of oxidative metabolism.

The focus of many studies has been the modulation of metabolism at the time of reperfusion; however, the presence of residual flow in the infarct zone means that some of the myocardium at risk would be accessible to therapeutic intervention before revascularization. For example, in the recent clinical GIK trials, GIK infusion was commenced on hospital admission and before revascularization therapy (17, 51). The initiation of therapy during this time may provide valuable additional cell survival advantages. King et al. (26) demonstrated that, in a LFI-and-reperfusion protocol, perfusion of diabetic hearts with glucose and palmitate resulted in improved recovery of function compared with glucose alone. They speculated that this improvement could be due to palmitate oxidation, leading to increased ATP production during LFI, which is clearly counter to the concept that increasing carbohydrate use at the expense of fatty acids is beneficial in the setting of ischemia.

The results from the study by King et al. (26) and evidence of appreciable oxidation of glucose and fatty acids (3, 18, 37) during LFI demonstrate the need for a better understanding of substrate utilization and regulation under these conditions. We and others previously showed that, when present at physiological concentrations, lactate and pyruvate contribute significantly to energy production and, in combination, may contribute at least as much as, if not more than, glucose (10, 25). However, to our knowledge, there is no information regarding the contributions of these substrates to cardiac energy production during LFI. Therefore, the goals of this study were 1) to evaluate whether 13C NMR glutamate isotopomer analysis can be used to determine substrate utilization during LFI; 2) if so, to investigate the contributions to glucose, lactate, pyruvate, and palmitate to oxidative energy metabolism; and 3) to evaluate the importance of oxidative and nonoxidative energy metabolism to ATP production during LFI.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Heart perfusion. All animal experiments were approved by Institutional Animal Care and Use Committee and conformed to the National Institutes of Health Guide for the Care and Use of Laboratory Animals [DHHS Publication No. (NIH) 85-23, Revised 1996, Office of Science and Health Reports, Bethesda, MD 20892]. Fed, adult male Sprague-Dawley rats were anesthetized with ketamine (100 mg/kg ip) and decapitated as previously described (10, 11, 29). Hearts were excised and perfused in a modified Langendorff mode with Krebs-Henseleit bicarbonate buffer equilibrated with 95% O2-5% CO2 (38°C, pH 7.4). The buffer contained 3% bovine serum albumin (BSA, essentially fatty acid free; Intergen, Mansfield, MA), 50 µU/ml insulin, and the following components (in mM): 118 NaCl, 4.8 KCl, 1.2 MgSO4, 1.4 CaCl2, 1.2 KH2PO4, 25 Na2HCO3, 0.5 glutamine, 5 glucose, 1 sodium lactate, 0.1 sodium pyruvate, and 0.3 sodium palmitate. Concentrations of insulin and energy substrates were chosen to be representative of the physiological fed state (29). Cardiac function was monitored continuously via a fluid-filled balloon placed in the left ventricle and connected to a pressure transducer. End-diastolic pressure was set to 5 mmHg by adjustment of balloon volume, and coronary flow rate was adjusted to maintain 75 mmHg perfusion pressure. All hearts were equilibrated for 30 min; after the first 10 min, hearts were paced at a frequency of 320/min, and this was continued for the remainder of the experiments. The partial pressure of oxygen in the perfusate and in the pulmonary artery effluent was measured at the beginning and end of experiments using a pH/blood gas/electrolytes analyzer (model 1640, Instrumentation Laboratory, Lexington, MA). Molar oxygen contents were determined from these partial pressures by use of the known solubility of oxygen in the buffer and the density of oxygen.

Experimental protocols. After equilibration, hearts were divided into two perfusion groups, control (n = 11) and LFI (0.3 ml/min, n = 8), and the perfusion medium was changed from medium with unlabeled substrates to medium with 13C-labeled substrates at the same concentrations used during the equilibration period. In both groups, perfusion with 13C-labeled substrates continued for another 30 min. In the control group, perfusion pressure was maintained at 75 mmHg, requiring a flow rate of 8.9 ± 0.7 ml·min–1·g wet heart wt–1 (mean ± SD). For the LFI group, coronary flow was reduced to a fixed rate of 0.3 ml/min, corresponding to 0.25 ± 0.03 ml·min–1·g–1. The flow rate of 0.3 ml/min was chosen on the basis of the report by King et al. (26), in which they suggested that there was oxidation of palmitate during a similar LFI protocol. Over the final 5 min of the experiment, the effluent from the pulmonary artery was collected and extracted with 6% perchloric acid as described previously (8, 30); an aliquot was taken before extraction for chemical determination of total lactate concentration. After 30 min of perfusion with labeled substrates, hearts were freeze clamped, weighed, and extracted with perchloric acid in a similar manner (10, 11, 29). Heart and pulmonary artery effluent extracts were freeze dried and redissolved in a potassium phosphate buffer (pH 7.5, 50 mM) with 2H2O solvent (99.9%; Cambridge Isotope Laboratories, Andover, MA).

13C-labeling schemes. The final 30 min of perfusion in the control or LFI protocols were performed using one of two different 13C-labeling schemes: sodium [2-13C]pyruvate, sodium [3-13C]lactate, sodium [U-13C]palmitate, and unlabeled glucose (scheme 1) or sodium [2-13C]pyruvate, sodium [3-13C]lactate, unlabeled sodium palmitate, and [U-13C]glucose (scheme 2). In scheme 1, glucose and endogenous substrates are unlabeled; in scheme 2, palmitate and endogenous substrates are unlabeled. Thus, taken together, these two substrate-labeling schemes enabled us to determine the relative contributions of palmitate, lactate, pyruvate, glucose, and endogenous substrates to acetyl-CoA entry into the tricarboxylic acid (TCA) cycle (see below). Table 1 shows the numbers of perfusions performed in the control and LFI groups for each labeling scheme.


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  		Table 1. Labeling scheme and numbers of experiments in each group

 
NMR spectroscopy. As previously described, spectra were recorded on a Bruker AVANCE 500-MHz NMR spectrometer operating at 11.85 T with a TXI inverse probe at 300,000 (29). 13C-NMR spectra were recorded with 16,000 data points and a spectral width of 25 kHz, with a 30° flip angle and recycle time of 0.3 s with use of broad-band proton decoupling. Spectra were referenced to the lactate methyl carbon resonance at 21.1 ppm. 1H-NMR spectra were recorded with 6-kHz spectral width, 16,000 data points, 45° flip angle, and 2-s interpulse delay. The water 1H peak was minimized with a presaturating pulse, and spectra were referenced to the residual water 1H peak at 4.76 ppm. All spectra were analyzed using a commercial PC-based analysis software package (Nuts, Acorn NMR, Livermore, CA).

[13C]glutamate isotopomer analysis. The distribution of glutamate 13C isotopomers, as determined from the labeling pattern observed in the NMR spectrum, can be used to determine the relative entry rates of acetyl-CoA derived from various 13C-labeled sources (34). Figure 1 illustrates possible isotopomers and corresponding 13C-NMR spectra of C-4 of glutamate arising from labeling scheme 1. Entry of acetyl-CoA derived from [3-13C]lactate results in glutamate isotopomers labeled at C-4 but not C-5, whereas entry of acetyl-CoA derived from [U-13C]palmitate results in isotopomers labeled at C-4 and C-5. Acetyl-CoA formed from [2-13C]pyruvate labels only C-5 and, therefore, results in no resonance at C-4. Similarly, unlabeled acetyl-CoA from glucose or glycogen also yields no C-4 resonance. The ratio of the glutamate C-4 isotopomer resonance intensities arising from lactate metabolism to those arising from palmitate metabolism provides a direct measure of the relative contributions of these two substrates to the TCA cycle without use of complex models (14). A similar analysis of the C-5 resonance would yield the ratio of palmitate (a doublet at C-5) to pyruvate (a singlet at C-5).



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  		Fig. 1. 13C-labeling pattern of acetyl-CoA and glutamate from 13C-labeled substrates into the tricarboxylic acid (TCA) cycle as used in scheme 1. Each substrate forms a uniquely labeled acetyl-CoA and glutamate. Also shown are 13C NMR spectra of individual glutamate C-4 resonances and the spectrum resulting from their superposition. Open circles, 12C; solid circles, 13C; shaded circles, 12C or 13C. Glucose, glycogen, and [2-13C]pyruvate do not label C-4; hence, they result in no resonance. PDH, pyruvate dehydrogenase; {alpha}-KG, {alpha}-ketoglutarate; LDH, lactate dehydrogenase; OAA, oxaloacetate. Peaks are labeled as lactate derived (L) or palmitate derived (P).

 
More complex models take into account all five 13C NMR resonances of glutamate and provide the fraction of total acetyl-CoA entering the TCA cycle that originates from unlabeled acetyl-CoA and [1,2-13C]-, [2-13C]-, and [1-13C]acetyl-CoA, thus enabling the relative contributions of up to four different substrates (34). We performed such an analysis here using tcaCALC, a computerized fitting routine developed and provided by Dr. Mark Jeffrey (University of Texas Southwestern Medical Center; http://www2.swmed.edu/rogersmr/). Calculation of relative substrate utilization is not dependent on reaching isotopic steady state (35); however, measurement of relative anaplerotic flux is dependent on isotopic steady state. Therefore, to determine whether the experiments were at isotopic steady state at the end of the 30-min perfusion with 13C-labeled substrates, we compared the estimates of unlabeled acetyl-CoA and [1,2-13C]-, [2-13C]-, and [1-13C]acetyl-CoA with TCA cycle flux determined by "steady-state" and "non-steady-state" calculations.

We were interested in determining the contributions of five distinct sources of acetyl-CoA: glucose, lactate, pyruvate, palmitate, and endogenous substrate. However, this cannot be achieved in a single experiment, inasmuch as only four labeling combinations are possible for acetyl-CoA. Therefore, as described above, we utilized two different labeling schemes. To quantify oxidation rates of all five sources, the mean palmitate-derived fraction determined from scheme 1 was subtracted from the unlabeled fraction in the scheme 2 experiments to determine the fraction of acetyl-CoA entering the TCA cycle from endogenous sources. Conversely, the corresponding fraction of glucose-derived acetyl-CoA was determined from the scheme 2 experiments, and this was subtracted from the unlabeled fraction in the scheme 1 experiments. This approach allowed determination of relative oxidation of all four exogenous substrates plus endogenous substrate utilization. Triglycerides and glycogen are potential sources of unlabeled acetyl-CoA and, therefore, cannot be differentiated in this study, but, using similar perfusion conditions, we previously showed that there is no appreciable contribution from endogenous triglycerides in controls (10). Thus we have assumed that glycogen is the primary endogenous source of acetyl-CoA in both experimental groups.

In addition to carbon entry into the TCA cycle metabolism via acetyl-CoA, carbon entry can also occur via so-called anaplerotic pathways, as previously described in detail elsewhere (22, 29). Such pathways include pyruvate carboxylase and malic enzyme, which catalyze the formation of oxaloacetate and malate, respectively, from pyruvate, synthesis of succinyl-CoA from propionyl-CoA, or synthesis of {propto}-ketoglutarate from glutamate. Although in principle the 13C-labeling pattern in glutamate will be different if [13C]pyruvate is metabolized via pyruvate carboxylase rather than PDH, we previously found that there is insufficient information in the [13C]glutamate isotopomer distribution to reliably determine the relative flux through pyruvate carboxylase under these experimental conditions (29). Therefore, only unlabeled anaplerosis was included in the analyses presented here.

Measurement of absolute substrate oxidation rates. The absolute oxidation rates of each labeled substrate can be calculated by combining the relative rates of TCA cycle acetyl-CoA entry determined from the glutamate 13C isotopomer analysis with measurements of tissue oxygen consumption. As originally described by Malloy et al. (33), the overall relation between the total citrate synthase rate (t) and oxygen consumption rate (t) is given by

(1)
where Fglu, Fgly, Fpyr, Flac, and Fpalm are the relative entry rates of acetyl-CoA derived from glucose, glycogen, pyruvate, lactate, and palmitate; Rglu, Rgly, Rpyr, Rlac, and Rpalm are the ratios of the number of moles of oxygen required to oxidize 1 mol of the respective substrate divided by the number of moles of acetyl-CoA generated from 1 mol of each substrate; ypro is anaplerosis (relative to t) due to propionate; and yCHO is anaplerosis (relative to t) due to exogenous lactate or glucose or endogenous glycogen. The stoichiometric proportionalities R are known from first principles and are constant; t is measured from the arteriovenous difference in perfused oxygen content, and all the other values in Eq. 1 are determined from glutamate isotopomer analysis. As described above, our data do not allow determination of the individual sources of anaplerosis. In this study, the perfusion mixtures did not contain a source of exogenous propionate; therefore, we have assumed that ypro is negligible. Thus we need only to account for yCHO; in our case, this will be given by ytotal – ypyr, where ypyr is the anaplerosis due to exogenous pyruvate. As a first approximation, we have assumed that ypyr is proportional to Fpyr so that ypyr/ytotal = Fpyr. Anaplerosis due to exogenous pyruvate needs to be accounted for separately, because, in contrast to pyruvate resulting from metabolism of glucose or exogenous lactate, no additional reducing equivalents need to be accounted for.

Once t is determined, the absolute oxidation rates of acetyl-CoA derived from each substrate are given by t multiplied by the corresponding F value for each substrate divided by the number of acetyl-CoA units generated from one unit of substrate. The absolute oxidation rate of each substrate is then given by the following equations



(2)


where the numbers in the denominators account for the number of moles of acetyl-CoA derived from 1 mol of substrate.

Lactate 1H NMR analysis. In hearts perfused with glucose and lactate as substrates, there is simultaneous uptake of exogenous lactate and efflux of glycolytically derived lactate (8). Therefore, to assess glycolytic lactate production, it is necessary to distinguish between exogenous lactate and lactate originating from exogenous glucose and glycogen. When 13C-labeling scheme 2 is utilized, exogenous lactate is labeled at C-3 only; lactate derived from [U-13C]glucose will be uniformly labeled with 13C and that originating from glycogen will be unlabeled. As previously described (30), the {beta}-proton region of the pulmonary artery effluent 1H NMR spectrum can be used to distinguish between these different lactate isotopomers to determine the amount of exogenous lactate and lactate derived from glycogen and exogenous glucose.

Therefore, with the use of 1H NMR spectra of standard samples, the 1H NMR spectra of effluent samples were deconvoluted into contributions from [U-13C]-, [3-13C]-, and unlabeled lactate (30). The rate of exogenous lactate uptake was calculated by the difference in the [3-13C]lactate concentrations in the perfusate and effluent multiplied by the coronary flow rate. If it is assumed that there is no change in the tissue concentration of lactate at the end of the experiment, the glucose and glycogen glycolytic rates are given by the concentration of [U-13C]- or [U-12C]lactate, respectively, in the effluent divided by 2 (inasmuch as 2 lactate molecules are generated from glycolysis of 1 mol of substrate) multiplied by the coronary flow rates. All values are normalized to wet heart weights.

Calculation of theoretical ATP yield. ATP may be generated within the cardiomyocyte by nonoxidative processes (substrate-level phosphorylation in glycolysis) or by oxidative phosphorylation utilizing the reducing equivalents NADH and FADH2 generated from substrate oxidation. Glycolysis of glucose or glycogen-derived glucose phosphate results in a net of 2 mol of ATP via substrate-level phosphorylation for each mole of substrate, so nonoxidative ATP production is found directly from the glycolytic rates of glucose and glycogen-derived glucose phosphate. Theoretical oxidative ATP production in each experiment was determined from the oxidation rates of each substrate multiplied by the oxidative part of the calculated ATP yield for each substrate as described by Opie (36) using the corrections for loss of energy during oxidative phosphorylation as reported by Hinkle et al. (24).

Statistical analysis. Unless otherwise stated, values are means ± SE. Unpaired t-tests were used to compare data where appropriate (Statistica, Statsoft, Tulsa, OK).


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Cardiac function. At the end of the equilibration period, baseline cardiac function as indexed by left ventricular developed pressure, rate-pressure product, and maximal positive and negative first derivatives of developed pressure (dP/dt) was not different between the control and LFI groups (Table 2). In the control group, cardiac function was stable throughout the subsequent 30 min of perfusion with 13C-labeled substrates. In the LFI group, there was rapid cessation of contractile function and gradual increase of left ventricular end-diastolic pressure after the reduction in flow rate to 75 ± 7 mmHg at the end of the ischemic period (Fig. 2).


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  		Table 2. Functional data for control and LFI groups at baseline and at the end of experiment, immediately before freeze clamping

 


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  		Fig. 2. Left ventricular end-diastolic pressure (LVEDP) after ischemia in low-flow ischemia (LFI). With onset of reduced flow (time 0), contracture develops within 10–15 min.

 
Oxidative substrate metabolism. Representative 13C NMR spectra from control and LFI extracts in hearts perfused with 13C-labeled lactate, pyruvate, and palmitate (scheme 1) are shown in Fig. 3. As anticipated on the basis of previous studies (29), there was significant enrichment of all the glutamate resonances in the control group. Resonances from [3-13C]lactate and [3-13C]alanine are also clearly evident. Interestingly, in the LFI group, there was also appreciable enrichment of glutamate, demonstrating significant oxidation of exogenous 13C-labeled substrates, despite ~97% reduction in coronary flow and oxygen delivery. As illustrated in Fig. 1, simple examination of the C-4 glutamate resonance can be used to evaluate the relative contributions of lactate and palmitate to acetyl-CoA entry into the TCA cycle. In Fig. 3 the glutamate C-4 resonances for the two protocols have been expanded and the resonances resulting from oxidation of lactate and palmitate have been identified, demonstrating that they are of sufficient quality for isotopomer analysis.



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  		Fig. 3. Representative 13C spectra of heart extracts from labeling scheme 1. Spectra are expanded to show region from 10 to 56 ppm encompassing C-2, C-3, and C-4 glutamate resonances and C-3 resonances of lactate and alanine. Insets: expansions of region of glutamate C-4 resonance. Peaks are labeled as lactate derived or palmitate derived.

 
Computational analysis using tcaCALC on all available glutamate isotopomer data was performed using non-steady-state and steady-state assumptions. The results obtained from labeling scheme 1 are summarized in Table 3. There were no significant differences between the non-steady-state and steady-state analyses, indicating that in both protocols the hearts were at or near isotopic steady state after 30 min of perfusion with 13C-labeled substrates. As described in METHODS, it is possible to directly determine the ratio of lactate to palmitate contributions to acetyl-CoA entry into the TCA cycle from the glutamate C-4 resonance alone. The method requires no computational model and no assumptions regarding isotopic or metabolic steady state. The results from this direct analysis, along with the ratios obtained from the steady-state and non-steady-state tcaCALC analysis using all available data, are given in Table 4. No significant differences in the ratio were found for any of the analysis methods. Inasmuch as the directly calculated ratio is model independent and reflects the contribution of the last condensation reaction of citrate synthase (14), this provides additional evidence that the hearts were at or near isotopic steady state at the end of the experiment. Finally, to further confirm that our conditions are at or near steady state, pilot studies were performed with hearts freeze clamped and extracted after 20 and 40 min of LFI (n = 2 each). Analysis of these studies revealed no significant difference in substrate contribution to TCA cycle flux over the time points. However, anaplerosis relative to citrate synthase flux, which is a more sensitive measure of the proximity to steady state, was higher after 20 min of LFI (1.0 ± 0.3) but unchanged from 30 (0.4 ± 0.1) to 40 min (0.39 ± 0.07) of LFI.


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  		Table 3. Fractional enrichments of acetyl-CoA determined by non-steady-state and steady-state [13C]glutamate isotopomer analysis for control and LFI groups (scheme 1)

 

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  		Table 4. Ratio of lactate to palmitate contributions to acetyl-CoA formation determined by non-steady-state, steady-state, and direct analysis of [13C]glutamate isotopomer distribution for control and LFI groups

 
Any deviation from isotopic steady state will result primarily in an increase in the tcaCALC-derived estimate for relative anaplerosis, with little change in the relative contributions of substrates to TCA cycle flux. Consequently, if our assumption of isotopic steady state is incorrect, our calculated anaplerosis would be artifactually high. Therefore, to estimate the potential error in the substrate oxidation rates, we repeated the calculations with the assumption that the relative anaplerotic flux was only one-half of that obtained from the tcaCALC fit. The resulting oxidation rates were only ~5–8% higher than those obtained from the steady-state assumption, which is consistent with the fact that anaplerotic metabolism contributes minimally to total cardiac oxygen consumption (33). Consequently, all subsequent analyses were performed using the steady-state method.

The relative contributions of all exogenous substrates and glycogen to acetyl-CoA entry into the TCA cycle are summarized in Fig. 4. Consistent with the spectra in Fig. 3, there was oxidation of all exogenous substrates in control and LFI protocols. The most appreciable difference between the control and LFI groups with regard to relative contributions of these substrates was a decrease in the contribution from lactate oxidation accompanied by an increase in oxidation of glycogen in the LFI group. In the control group, there was no detectable entry of glycogen-derived acetyl-CoA into the TCA cycle. Consistent with our earlier reports (10), lactate-derived acetyl-CoA provides a large proportion of TCA cycle flux in the control group, with palmitate contributing one-third of acetyl-CoA entry and the remainder originating from glucose and pyruvate.



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  		Fig. 4. Relative contributions of all exogenous substrates and glycogen to myocardial acetyl-CoA entry into TCA cycle in control and LFI groups. In control group, oxidation of glycogen-derived pyruvate was not detected. *P < 0.05 vs. control.

 
As described in METHODS, by combining the data in Fig. 4 with measurements of oxygen consumption, we calculated the absolute rates of substrate oxidation in control and LFI groups. These data are summarized in Fig. 5. Not surprisingly, the absolute oxidation rates are markedly reduced in LFI, consistent with the reduction in oxygen consumption. In the control and LFI groups, the pattern of substrate use is somewhat different from the relative contributions (Fig. 4), primarily because the oxidation of lactate or pyruvate forms only one acetyl-CoA, whereas the oxidation of glucose forms two acetyl-CoA units and palmitate oxidation contributes eight acetyl-CoA units. Nevertheless, it is clear that, in control and LFI groups, oxidation of lactate is substantially greater than that of all other substrates.



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  		Fig. 5. Absolute oxidation rates of each substrate in control and LFI groups. Note difference in scales between A and B. In LFI group, oxidation rates for all substrates were significantly different (P < 0.05) from those in control group.

 
Anaplerosis. Relative and absolute anaplerotic fluxes for control and LFI are shown in Fig. 6. Surprisingly, in LFI, there was a large increase in relative anaplerosis compared with the control group (Fig. 6A). However, this is potentially misleading, because these data are relative to total TCA cycle flux, which is substantially lower in the LFI group. Therefore, combining this analysis with oxygen consumption data, we calculated the absolute anaplerosis fluxes (Fig. 6B). It is clear that total anaplerosis is significantly lower in the LFI than in the control group; however, it is reduced by only ~75% compared with ~100-fold reduction in substrate oxidation rates.



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  		Fig. 6. Relative and absolute anaplerosis in control and LFI groups. *P < 0.05 vs. control.

 
Nonoxidative substrate metabolism. The 1H NMR spectrum of the lactate {beta}-proton region of the extracted pulmonary artery effluent from a control heart is shown in Fig. 7. The top spectrum in Fig. 7A is the raw spectrum of the effluent. The three spectra below the raw spectrum of the effluent are those of [3-13C]-, [U-12C]-, and [U-13C]lactate isotopomer standards, scaled to the appropriate relative areas to reflect their presence in the raw spectrum (91%, 1%, and 5.5%, respectively). Figure 7B shows the remainder after subtraction of these contributions from the raw spectrum. The apparent peaks at 1.43 and 1.16 ppm represent subtraction errors due to digital offset and line width differences between the raw spectrum and those of the standards; integration of these regions reveals that their areas are zero. However, in the central region at 1.30 ppm, a quartet (doublet of doublets) representing 2.5% of the total lactate spectral area is seen with coupling constants of 7.0 and 4.4 Hz (Fig. 7B, inset). These resonances are consistent with the presence of [2-13C]lactate in the effluent sample (30). In these experiments, the only possible source of this lactate isotopomer is [2-13C]pyruvate, demonstrating that an appreciable amount of exogenous pyruvate was taken up by cardiomyocytes, reduced to lactate, and transported out of the cells in the control group. This isotopomer of lactate was not detected in the effluents from the LFI experiments.



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  		Fig. 7. A: lactate {beta}-proton region of 1H NMR spectrum of a representative pulmonary artery effluent from control group in labeling scheme 2. A: spectrum showing raw data. Spectra of standard samples of [3-13C]lactate, [U-12C]lactate, and [U-13C]lactate scaled to fit raw spectrum with their corresponding relative areas are shown below raw data. Decomposition in this manner allows determination of efflux rates of lactate derived from exogenous glucose and endogenous glycogen. B: remainder when standard spectra are subtracted from raw spectrum. Apparent signals at 1.43 and 1.16 ppm are digital subtraction errors and have zero area. Inset, quartet remaining in the center at 1.30 ppm (vertical scale increased 16 times), representative of spectrum of [2-13C]lactate formed from metabolism of exogenous [2-13C]pyruvate.

 
As described in METHODS, isotopomer analysis of the lactate {beta}-proton resonance distinguishes the various sources of lactate and allows determination of the rates of uptake of exogenous lactate and efflux of lactate derived from other sources (glucose, glycogen, and pyruvate). With the use of this analysis, the total efflux rates of lactate derived from these sources ([U-13C]-, [U-12C]-, and [2-13C]lactate) and uptake rates of [3-13C]lactate were determined and are shown in Fig. 8A for the control and LFI groups. The rate of lactate uptake during LFI is ~4.5% of that during control, whereas flow is ~3% of the control value, suggesting that the decrease in the rate of lactate uptake is primarily due to decreased rate of delivery. The rate of lactate efflux resulting from glycolytic metabolism of exogenous glucose and glycogen is not significantly different between the two groups. However, the contributions of each lactate source to total lactate efflux is clearly different between the groups (Fig. 8B); in the LFI group, the predominant source of glycolytic lactate production shifts from glucose to glycogen. Lactate derived from exogenous pyruvate makes up ~20% of the total lactate efflux in the controls but was not detectable in the LFI group.



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  		Fig. 8. A: total lactate uptake and efflux rates in control and LFI groups as measured in pulmonary artery effluents using labeling scheme 2. B: contribution of each source of lactate to lactate efflux, measured as described in RESULTS. *P < 0.05 vs. control.

 
Estimated ATP synthesis rates. Using the absolute oxidative and nonoxidative flux data, we estimated the total theoretical ATP yield under control and LFI protocols (Table 5). Not surprisingly, the oxidative ATP yield was much smaller in the LFI group than in controls. Nevertheless, substrate oxidation remains a relatively important source of cellular energy production, responsible for nearly one-third of ATP produced in LFI. There was no difference in nonoxidative ATP production rates between control and LFI groups.


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  		Table 5. Rates of oxidative, nonoxidative, and total ATP production in control and LFI groups

 
The relative contributions of each possible energy source to total ATP production in the control and LFI groups are compared in Fig. 9. Similar to the data shown in Fig. 4, the major differences between control and LFI in the relative contributions of the substrates are due to a decreased contribution from lactate and the increased contribution from glycogen.



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  		Fig. 9. Relative contribution of each substrate to total ATP production in control and LFI groups. *P < 0.05 vs. control; {dagger}P = 0.06.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
We have shown for the first time that, using 13C NMR glutamate isotopomer analysis, we can obtain detailed information regarding the contributions of multiple exogenous substrates to oxidative energy production under LFI in a manner similar to that previously reported by us (29) and others (25, 34) under normoxic conditions. We found that, despite a 97% reduction in coronary flow and a corresponding decrease in oxygen consumption, there was significant oxidation of all exogenous substrates plus glycogen during LFI. By combining glutamate isotopomer analysis of tissue extracts with lactate isotopomer analysis of perfusate effluent samples, we were also able to quantify total oxidative and nonoxidative ATP production in the control and LFI groups (Table 5), demonstrating that, even during this very-low-flow ischemia protocol, oxidative metabolism still accounted for approximately one-third of the total ATP production. These results clearly demonstrate the utility of 13C NMR glutamate isotopomer analysis in evaluating cardiac substrate utilization under pathophysiological conditions. Furthermore, the observation that oxidation of all exogenous substrates occurs even in LFI may have important implications for the underlying mechanisms related to protective effect of increasing cardiac carbohydrate utilization.

One of the constraints of 13C NMR glutamate isotopomer analysis is the relatively low sensitivity of the method compared with traditional radioisotope techniques. Consequently, we were uncertain as to whether there would be sufficient enrichment of the glutamate pool in LFI to permit analysis. The spectra in Fig. 3 clearly demonstrate that, despite the very low coronary flow and the resulting reduction in 13C-labeled substrate and oxygen delivery coupled with the decrease in TCA cycle flux, there was appreciable enrichment of the glutamate pool in the LFI group. One advantage of this method over the use of radioisotope techniques is the ability to determine the contributions of up to four different substrates to energy production, as well as to determine anaplerotic flux, all within a single experiment. However, for all of this information to be obtained from a single experiment, it is necessary for the enrichment of the glutamate pool to have reached isotopic steady state. As shown in Table 4, there was no significant difference between the non-steady-state and steady-state analyses for the control or the LFI group. Furthermore, the ratio of lactate to palmitate entry into the TCA cycle determined from direct analysis of the C-4 glutamate resonance, which is independent of any assumptions regarding isotopic or metabolic steady state, was also similar to the ratios determined by the other analyses in the control and LFI groups. We also found that relative anaplerotic flux was unchanged between 30 and 40 min of LFI, further supporting the notion that isotopic steady state had been reached after 30 min of LFI. Thus we concluded that in these experiments the reactions were at or near isotopic equilibrium, and it was appropriate to use the steady-state analysis. However, it should be emphasized that, although the 13C data suggest that we have reached an isotopic steady state, this does not necessarily reflect a biological steady state. Indeed, although function was stable throughout the control experiments, there was a marked increase in left ventricular end-diastolic pressure in the LFI group during the 13C perfusion (Fig. 2) that reached a new steady state at 20 min.

The predominant substrates for cardiac energy production are usually considered to be glucose and fatty acids, despite the fact that a number of in vitro and in vivo studies have demonstrated that when lactate is present, it is readily oxidized and is utilized in preference to glucose (4, 10, 16, 21, 25). Our results reinforce our earlier observation that, when present at physiologically relevant concentrations, lactate and pyruvate contribute appreciably to cardiac energy production (29). Indeed, in this study in the control group, lactate is clearly utilized to a greater extent than glucose or palmitate, whether it is evaluated in relative terms (Fig. 4), absolute oxidation rates (Fig. 5A), or contribution to total ATP production (Fig. 9). Despite obvious differences between the isolated heart and the heart in vivo, our results are consistent with reports of cardiac metabolism in vivo. For example, Liedtke et al. (28) reported that under control conditions the lactate extraction rate by the heart was three to four times higher than the rate of glucose utilization, which is consistent with our observation that the contribution of lactate to myocardial energy production is greater than that of glucose. More recently, Hart et al. (23) reported fatty acid, lactate, and glucose uptake rates across the heart in vivo in dogs that are in fairly good agreement with the rates of palmitate, lactate, and glucose oxidation in the control group (Fig. 5).

Under the conditions of our experiments, which were designed to mimic the fed state, we found that, in the control group, palmitate contributed ~30% of total ATP production, which is substantially lower than the ~70% usually attributed to fatty acids. However, the majority of in vivo studies of cardiac metabolism are necessarily performed on fasted animals, which increases serum fatty acid concentrations and lowers insulin levels compared with the fed state, and both of these factors will tend to increase fatty acid utilization compared with carbohydrate oxidation. Clearly, the contributions of glucose, lactate, pyruvate, and palmitate to oxidative metabolism are very sensitive to circulating insulin levels and substrate concentrations; consequently, the results presented here may not be applicable to other hormonal and/or metabolic conditions.

Of particular interest is that, during LFI, there was appreciable oxidation of all exogenous substrates as well as oxidation of glycogen, which was not detected in the control group. The increase in glycogen oxidation during LFI is consistent with the well-established effect of hypoxia on glycogenolysis and phosphofructokinase in the heart (36). Although there have been other reports of glucose and fatty acid oxidation during LFI (3, 18), we believe that this is the first report of exogenous lactate and pyruvate oxidation under conditions of severely reduced perfusion. However, Opie et al. (37) found an increase in the rate of glucose oxidation relative to fatty acid oxidation during flow reduction, which is consistent with the decrease in the contribution of palmitate to ATP production in the LFI group (Fig. 9).

Although the decrease in absolute substrate oxidation rates was similar to the reduction in oxygen consumption during LFI, surprisingly anaplerotic flux was not reduced to the same extent. Indeed, although oxidation rates decreased ~50-fold, anaplerosis decreased by only 5-fold. Although we are unable to identify the specific anaplerotic pathways in these experiments, we previously showed that total anaplerosis determined by 13C NMR methods is similar to flux through pyruvate carboxylase (14). Therefore, if we assume that anaplerotic flux in these experiments primarily reflects flux through pyruvate carboxylase, it is clear that the partitioning of pyruvate between PDH and pyruvate carboxylase is shifted toward pyruvate carboxylase in LFI. Several studies have shown that anaplerosis may be modulated by changes in the metabolic or physiological state (52); however, the significance of the apparent increase in flux through pyruvate carboxylase relative to PDH is unclear. It has been proposed that an increase in anaplerotic flux via oxaloacetate during LFI could lead to NADH-dependent reduction of fumarate to succinate, resulting in increased ATP production via complex I of the respiratory chain (40). It is possible, therefore, that the shift in pyruvate flux from PDH to pyruvate carboxylase in LFI may represent an adaptive response that could be important for mitochondrial ATP synthesis during low-oxygen conditions. We have shown that anaplerosis is insulin sensitive under normoxic conditions (29); consequently, this could represent a possible mechanism by which GIK therapy, as well as other interventions designed to increase carbohydrate use, may exert their anti-ischemic effect. Clearly, more work is needed to investigate the role of anaplerosis during LFI.

We previously showed that with the use of 13C-labeled lactate and 1H NMR spectroscopic lactate isotopomer analysis of effluent samples, it is possible to quantify the efflux of glycolytic lactate and the uptake of exogenous lactate (8). Using this approach, we found that the lactate uptake rate was slightly greater than the efflux rate in the control group; however, both rates were similar to those reported previously (8). Although lactate uptake was measurable in the LFI group, it was markedly diminished compared with the control group, reflecting the reduction in flow and decrease in lactate oxidation, as well as reduced transport into the cells due to an accumulation of intracellular lactate and, consequently, a transmembrane concentration gradient less favorable to uptake. Lactate efflux in the LFI group was similar to that in the controls, which is somewhat surprising, because glycolysis is usually thought to increase in response to reduced oxygen. However, this may reflect limitations in glucose delivery and lactate efflux as a result of the reduction in flow.

In addition to determining total lactate efflux, the choice of 13C-labeled substrates in scheme 2 enabled us to determine the rates of lactate efflux from different sources. In the control group, we found that lactate efflux primarily resulted from metabolism of exogenous glucose; however, there were also contributions from glycogen, and, perhaps most surprisingly, lactate derived from exogenous pyruvate also represented a significant proportion of total efflux. In the controls, we were unable to detect any oxidation of glycogen, suggesting that glycogen-derived pyruvate is preferentially metabolized to lactate, rather than oxidized to acetyl-CoA. However, other studies have reported that glycogen is preferentially oxidized, rather than metabolized to lactate (20). This discrepancy could be due to the fact that, in the study by Goodwin et al. (20), glucose was the only substrate provided, and glycogen metabolism was evaluated after stimulation with epinephrine. This is in contrast to the present study, where other carbohydrates were provided as well as palmitate, and glycogenolysis was not actively stimulated.

To our knowledge, the observation that exogenous pyruvate is taken up by the heart during normoxia, reduced to lactate, and released (Figs. 7 and 8B) has not been reported previously. Reduction of pyruvate to lactate by lactate dehydrogenase (LDH) requires NADH and, therefore, consumes reducing equivalents that could have been used for ATP production through oxidative phosphorylation. In fact, this may seem to be a "futile" process, particularly in light of the ongoing oxidation of lactate to pyruvate and formation of acetyl-CoA from the pyruvate produced by this oxidation. In simple biological systems such as red blood cells, there is rapid isotopic equilibrium between lactate and pyruvate (41); therefore, one possible explanation for the simultaneous reductive and oxidative reactions of LDH is that this merely reflects the establishment of an isotopic equilibrium for lactate and pyruvate. However, there is also evidence that, in the intact heart, lactate and pyruvate may not be at isotopic equilibrium (7–9, 27, 39). This could be a result of intracellular compartmentation, as proposed by Brooks et al. (5, 6), or different cell populations in the intact heart. Thus the uptake and interconversion of pyruvate to lactate and its subsequent release are consistent with the reports of cellular compartmentation of pyruvate and lactate metabolism (6–8, 27, 39) and may be a means of regulating the cellular redox potential.

The combination of 1H NMR lactate isotopomer analysis with 13C NMR glutamate isotopomer analysis enabled us to determine oxidative and nonoxidative metabolic rates of all exogenous substrates plus glycogen and to estimate the contributions of substrates to total ATP production in the control and LFI groups (Table 5, Fig. 9). In the controls, oxidative metabolism represented 97% of total ATP production. Although nonoxidative glycolysis becomes the major source of ATP production during LFI, oxidative metabolism was not negligible, resulting in one-third of the calculated total ATP produced (Table 5). Thus drugs designed to influence myocardial substrate oxidation may be useful in altering cellular energetics, even during severe ischemia. However, at least in these experiments, carbohydrates represent the predominant source of ATP in the control and LFI groups, with palmitate contributing ~20% and ~10% to total ATP production in controls and LFI, respectively; thus it is unclear whether interventions targeted at decreasing fatty acid oxidation will be especially effective. The substrate and insulin levels used here were chosen to represent a typical fed state. However, the conditions of stress, which accompany a myocardial infarction, include increased adrenergic stimulation and decreased insulin, which clearly affect myocardial metabolic rate and fuel selection. These factors were not considered here. Furthermore, in vivo the relative contributions of substrates to ATP production may also change as a result of the increased serum glucose, lactate, and fatty acids frequently seen after acute myocardial infarction (13, 31, 44–46, 48).

An important factor that should be considered is that the results from the [13C]glutamate isotopomer analysis represents a time average of the metabolism over the duration of perfusion with 13C-labeled substrates, rather than at a specific point in time. However, the good agreement between non-steady-state, steady-state, and direct analyses suggests that we have a reliable measure of exogenous substrate oxidation, despite the very low coronary flow. Any deviation from isotopic steady state would result primarily in an apparent increase in the estimate for relative anaplerosis and would cause little change in the relative contributions of substrates to TCA cycle flux. The calculation of substrate oxidation rates is determined primarily by the relative TCA cycle fluxes and the oxygen consumption rate; therefore, substrate oxidation rates are insensitive to even fairly large differences in anaplerotic flux. If our experiments were not at isotopic steady state after 30 min of LFI, analysis reveals that relative and absolute anaplerotic flux would be significantly decreased, but only a 5–8% increase in substrate oxidation rates would be expected.

In contrast to the time-averaged nature of the glutamate analysis, measurements of lactate uptake and efflux and, thus, nonoxidative metabolic rates were determined at a single time point at the end of the 30 min of LFI. Further studies are necessary to assess the time dependence of total lactate efflux and the contributions of glucose, glycogen, and pyruvate to lactate efflux during LFI. Another potential limitation is that we did not include tissue lactate levels and enrichments in our calculation of nonoxidative metabolic fluxes. Although during no-flow ischemia the increase in tissue lactate can be assumed to reflect glycolytic flux, it is unclear whether this is also the case during LFI, because lactate is being continually transported out of the cell. Although tissue lactate would be expected to increase early after the reduction in flow, only changes in tissue lactate levels over the last 5 min of the protocol would affect our lactate production rate determinations, because this is the time during which the effluents were collected for analysis. Three potential scenarios exist for the tissue lactate levels over these last 5 min. 1) If the rate of the lactate efflux is equal to the lactate production from glycolysis, then any change in tissue lactate levels presumably reflects a change in the equilibrium of LDH and would not contribute to the calculation of nonoxidative ATP production. In this situation, our calculated lactate production rates will be correct. 2) Lactate production may exceed the efflux rate during the effluent collection time. If this is the case, our calculated rates will underestimate the true lactate production (i.e., glycolysis) rates. 3) There is the possibility that lactate efflux exceeds production during these last 5 min. In this case, our calculated rates will be an overestimate of the true glycolytic rates. Although these limitations may affect the calculations of nonoxidative and oxidative flux rates, they do not negate the primary observation that there was significant oxidation of all exogenous substrate plus glycogen, despite a 97% reduction in flow and oxygen consumption.

In summary, we have shown that in the isolated perfused heart the combination of 13C NMR analysis of glutamate isotopomers in heart extracts, 1H NMR analysis of the perfusate effluent, chemical measurement of the lactate concentration in the effluent, and oximetry of the perfusate and effluent enable us to quantify all glycolytic and oxidative fluxes in a single type of experiment using just two 13C-labeling schemes. Furthermore, these measurements are sensitive enough to determine these fluxes, even during a severe reduction in coronary flow, and we demonstrated that, even with a 97% reduction in oxygen delivery, substrate oxidation still occurred, contributing an estimated 33% of total ATP produced during LFI. We believe that the approach described here will be valuable for future investigations into the underlying mechanisms related to a protective effect of increasing cardiac carbohydrate utilization and may ultimately lead to the identification of new therapeutic targets for the treatment of myocardial ischemia.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported in part by National Heart, Lung, and Blood Institute Grants HL-67464 and HL-48789 (to J. C. Chatham) and 5T32 HL-07703 (to S. G. Lloyd) and American Heart Association Grant-in-Aid 0050545N (to J. C. Chatham). All NMR studies were performed at the University of Alabama Birmingham NMR Core Facility funded by National Cancer Institute Cancer Center Support Grant CA-13148.


   ACKNOWLEDGMENTS
 
We thank Charlye Brocks and Clarence Forrest for technical support.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. C. Chatham, Univ. of Alabama at Birmingham, McCallum Bldg., Rm. 684 1530 3rd Ave. South, Birmingham, AL 35294-0005 (E-mail: jchatham{at}uab.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.


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