AJP - Heart Watch the video to learn how APS reaches out to developing nations.
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 286: H1461-H1470, 2004. First published December 11, 2003; doi:10.1152/ajpheart.00942.2003
0363-6135/04 $5.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
286/4/H1461    most recent
00942.2003v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (16)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Khairallah, M.
Right arrow Articles by Des Rosiers, C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Khairallah, M.
Right arrow Articles by Des Rosiers, C.

Profiling substrate fluxes in the isolated working mouse heart using 13C-labeled substrates: focusing on the origin and fate of pyruvate and citrate carbons

Maya Khairallah,1 François Labarthe,2 Bertrand Bouchard,2 Gawiyou Danialou,1 Basil J. Petrof,1 and Christine Des Rosiers1,2

1Department of Experimental Medicine, McGill University; and 2Department of Nutrition, University of Montreal, Notre-Dame Hospital Research Center, Montreal, Quebec, Canada H2L 4M1

Submitted 2 October 2003 ; accepted in final form 9 December 2003


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The availability of genetically modified mice requires the development of methods to assess heart function and metabolism in the intact beating organ. With the use of radioactive substrates and ex vivo perfusion of the mouse heart in the working mode, previous studies have documented glucose and fatty acid oxidation pathways. This study was aimed at characterizing the metabolism of other potentially important exogenous carbohydrate sources, namely, lactate and pyruvate. This was achieved by using 13C-labeling methods. The mouse heart perfusion setup and buffer composition were optimized to reproduce conditions close to the in vivo milieu in terms of workload, cardiac functions, and substrate-hormone supply to the heart (11 mM glucose, 0.8 nM insulin, 50 µM carnitine, 1.5 mM lactate, 0.2 mM pyruvate, 5 nM epinephrine, 0.7 mM oleate, and 3% albumin). The use of three differentially 13C-labeled carbohydrates and a 13C-labeled long-chain fatty acid allowed the quantitative assessment of the metabolic origin and fate of tissue pyruvate as well as the relative contribution of substrates feeding acetyl-CoA (pyruvate and fatty acids) and oxaloacetate (pyruvate) for mitochondrial citrate synthesis. Beyond concurring with the notion that the mouse heart preferentially uses fatty acids for energy production (63.5 ± 3.9%) and regulates its fuel selection according to the Randle cycle, our study reports for the first time in the mouse heart the following findings. First, exogenous lactate is the major carbohydrate contributing to pyruvate formation (42.0 ± 2.3%). Second, lactate and pyruvate are constantly being taken up and released by the heart, supporting the concept of compartmentation of lactate and glucose metabolism. Finally, mitochondrial anaplerotic pyruvate carboxylation and citrate efflux represent 4.9 ± 1.8 and 0.8 ± 0.1%, respectively, of the citric acid cycle flux and are modulated by substrate supply. The described 13C-labeling strategy combined with an experimental setup that enables continuous monitoring of physiological parameters offers a unique model to clarify the link between metabolic alterations, cardiac dysfunction, and disease development.

energy metabolism; citric acid cycle; 13C mass isotopomer analysis; gas chromatography-mass spectrometry; isolated working perfused heart

THERE IS A CURRENT RESURGENCE of interest in cardiac metabolism, driven in part by the availability of drugs such as ranolazine and trimetazidine, which can modulate the heart's substrate preference for energy production (38, 59, 61, 71). Substantial evidence indicates that a defect in energy substrate metabolism is an independent determining factor of cardiac dysfunction and disease development (5, 14, 33, 57, 60, 62, 66). However, much remains to be learned about the impact of manipulating substrate metabolism on the heart's contractile function and pathophysiological status. Genetically modified mice, especially if they carry inducible cardiomyocyte-specific changes in the expression of a single metabolic gene, have proven to be extremely valuable models for addressing this issue (10, 32, 37). The dynamic nature of cardiac metabolism and contractile functions requires, however, that investigations be conducted in the intact beating organ. Hence, methodologies developed for larger animals have to be applied to the mouse. Despite technical limitations associated with the small size of the organ, Grupp and colleagues (32) successfully characterized the mechanical function of the working mouse heart perfused ex vivo. The great advantage of this model is the ability to precisely manipulate and control experimental conditions such as the afterload, the preload, and the buffer composition. Subsequently, this ex vivo study model was refined by Belke et al. (8) for cardiac energy metabolism investigations. By using radioactive substrates and by providing the heart with a source of fatty acids, in addition to glucose, Belke et al. reported rates of glycolysis and glucose and fatty acid oxidation and showed a regulation of substrate selection according to the Randle cycle.

Although the prevailing view in the field of cardiac metabolism remains that long-chain fatty acids and glucose are the primary energy source, an increasing number of studies conducted both in vivo and ex vivo emphasize the importance of 1) the contribution of other energy substrates, such as lactate and pyruvate (13, 16, 17, 39, 45, 67); and 2) pyruvate metabolism through anaplerosis (19, 20, 30, 46, 52, 58, 67–69) for optimal cardiac energy metabolism. Lactate and pyruvate metabolism have been shown to be altered under pathophysiological conditions (13, 16, 30, 40, 51, 56, 58, 63, 69). Hence, it appears essential that the isotopic method used for the metabolic profiling of the mouse heart includes measurements of substrate fluxes through the pyruvate branch point. Compared with radioisotope methods, stable isotope techniques can provide, with less sample treatment, more information on the isotopic enrichment of intermediary precursor metabolites (e.g., acetyl-CoA or citrate for CO2 production) and on the metabolic fate of labeled molecules. In fact, the use of stable isotopes combined with mass isotopomer analysis has proven to be a very powerful approach for metabolic investigations in the ex vivo and in vivo perfused heart (15–20, 43, 45, 47, 51, 52, 67–69).

Thus this study was aimed at characterizing the metabolism of exogenous pyruvate and lactate in the working mouse heart perfused ex vivo. In addition, we considered it important to assess the metabolism of exogenous glucose and fatty acids to integrate our findings with those of others (1, 8). For this purpose, we expanded on a previously described 13C-labeling strategy using GCMS that was developed in the ex vivo working perfused rat heart (67, 69). Specifically, three differentially 13C-labeled carbohydrates and a 13C-labeled long-chain fatty acid were used to assess the metabolic origin and fate of tissue pyruvate as well as the relative contribution of substrates feeding acetyl-CoA (pyruvate and fatty acids) and oxaloacetate (OAA; pyruvate via anaplerosis), which are necessary for mitochondrial citrate synthesis. Perfusion conditions and buffer composition were optimized to mimic in vivo conditions in terms of cardiac functions and substrate-hormone concentrations, respectively. In addition, our experimental setup includes continuous monitoring of indexes of cardiac performance, myocardial oxygen consumption (MO2), and tissue integrity. Beyond providing data on substrate selection and oxidation rates, this study reports for the first time in the working mouse heart quantitative flux data on several metabolic processes that could potentially be altered by disease, such as lactate and citrate release (69), and highlights the substantial contribution of carbohydrates, other than glucose, in myocardial energy production.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Materials

Sources of chemicals, biological products, and 13C-labeled substrates, as well as the procedure for the dialysis of fatty acid-free BSA (BSA fraction V, Intergen), have been reported previously (19, 20, 67–69).

Heart Perfusions in the Semirecirculating Mode

Animal experiments were approved by the local ethics committee in agreement with the guidelines of the Canadian Council on Animal Care. Male C57BL/10 mice (10–12 wk old; average body weight: 29.4 ± 0.2 g; Jackson Laboratories) were anesthetized (1 µl/g ip) with a mixture of ketamine (100 mg/ml) and xylazine (20 mg/ml) and were heparinized (5,000 U/kg ip) 15 min before surgery. After rapid insertion of an 18-gauge steel cannula into the aorta, hearts were excised and placed on the perfusion system, where they first underwent Langendorff perfusion (55 mmHg perfusion pressure) for ~5 min. The modified Krebs-Henseleit buffer (containing 119 mM NaCl, 4.8 mM KCl, 2.5 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4, 25 mM NaHCO3, 0.1 mM EDTA, 11 mM glucose, 0.8 nM insulin, 50 µM L-carnitine, 1.5 mM lactate, and 0.2 mM pyruvate) was gassed with 95% O2-5% CO2 (pH 7.4) and maintained at 37.5°C. During this retrograde perfusion period, a polyethylene catheter (PE-50) was inserted through the pulmonary vein into the left ventricle, anchored at the apex of the heart by a fluted end, and connected to a pressure transducer for the continuous monitoring of intraventricular functions (Digi-Med Heart Performance Analyzer, Micro-Med). Furthermore, the left atrium was connected to the preload reservoir (oxygenator) by cannulating the pulmonary vein with a 16-gauge steel cannula. Spontaneously beating hearts were then switched to the anterograde work-performing mode and perfused via the left atria cannula with a semirecirculating modified Krebs-Henseleit buffer containing oleate complexed to 3% albumin (see Perfusion Protocols for details). The perfusion setup for the working mouse heart in the semirecirculating mode, depicted in Fig. 1, was adapted from that previously described for the working rat heart (67, 69). Briefly, in contrast to a recirculating setup, the coronary perfusate, which contains various metabolites released by the heart, is not recirculated but is continuously collected. Thus only the aortic outflow is recirculated into the buffer reservoir. The preload and afterload pressures were maintained at 15 and 50 mmHg, respectively, through continuous monitoring by pressure transducers (Digi-Med Blood Pressure Analyzer and Low Pressure Analyzer, Micro-Med). Other parameters that were continuously monitored (10-s duration for each recording) are 1) the atrial and aortic flow rates (monitored with calibrated electromagnetic flow probes; Carolina Medical Electronics); 2) left ventricular functions, namely, heart rate (HR), maximum left ventricular systolic pressure, left ventricular end-diastolic pressure, and maximum value for the first derivative of maximum left ventricular systolic pressure (measured by using the intraventricular cannula linked to a pressure transducer; Digi-Med Heart Performance Analyzer, Micro-Med); and 3) heart temperature (measured with a thermocouple; Yellow Springs Instruments). In addition, biochemical parameters, among which PO2, PCO2, pH, Ca2+, and other ion concentrations, were evaluated by using a blood gas, electrolytes, and pH analyzer (ABL 77 series, Radiometer Copenhagen) in the atrial influent and the coronary effluent perfusates collected every 10 min. Finally, effluent samples were collected every 10 min to assess lactate dehydrogenase (LDH) release, an index of tissue necrosis, by using a standard enzyme activity assay on a Roche Cobas Fara apparatus.



View larger version (23K):
[in this window]
[in a new window]
  		Fig. 1. Schematic overview of the semirecirculating working mouse heart setup. See MATERIAL AND METHODS for details.

 

Perfusion Protocol

Working mouse hearts were perfused for 30 min with a semirecirculating modified Krebs-Henseleit buffer, whose composition was identical to that described above (see Heart Perfusions in the Semirecirculating Mode) except that it also contained oleate bound to 3% albumin and 5 nM epinephrine. In addition, for any given perfusion, one of the unlabeled substrates was replaced by its corresponding labeled substrate. Substrate and hormone concentrations are within the range of reported values of mouse plasma concentrations in the non-fasting state (Jackson Laboratories "Phenome Database"; and refs. 1, 3, 8, 25, 64, 72, and 73). We chose a lactate concentration of 1.5 mM, which is below values reported for the normal rested mouse (between 2.7 and 4.7 mM) (44, 54, 72). However, in blood drawn by cardiac puncture in a syringe containing sulfosalicylic acid to precipitate proteins and prevent anaerobic glycolysis, we evaluated lactate concentration to be 1.8 ± 0.3 mM (n = 4) by using an enzymatic assay (69). Perfusion protocols can be subdivided into two groups that differed with respect to their oleate concentration. In a first group of experiments, oleate concentration was 0.7 mM. Four different 13C-labeled substrates were used: [U-13C3]lactate [initial molar percent enrichment (MPE): 99%; n = 5], [U-13C3]pyruvate (initial MPE: 99%; n = 4), [U-13C6]glucose (initial MPE: 50%; n = 6), or [U-13C18]oleate (initial MPE: 25%; n = 4). In a second set of experiments, we tested whether perfused working mouse hearts responded to a change in substrate supply resulting from a lowering of oleate concentration from 0.7 to 0.4 mM. Two different 13C-labeled substrates were used: [U-13C3]pyruvate (initial MPE: 99%; n = 4) or [U-13C18]oleate (initial MPE: 25%; n = 4).

Throughout the perfusion, influent and effluent perfusates were collected at the following times: 1) 20–25 min to evaluate the citrate and succinate release rates; and 2) 25–30 min to document the lactate and pyruvate uptake and release rates, of which 1 ml was immediately treated with sodium borodeuteride. These samples were stored at –80°C until further analysis. Subsequent to each perfusion period, hearts were freeze clamped with metal tongs chilled in liquid nitrogen, weighed, and stored at –80°C.

Analytical Procedures

Perfusate and tissue processing. Procedures for the determination of 1) citrate and succinate release rates and 2) 13C enrichment and concentration of the citric acid cycle (CAC) intermediates (citrate, {alpha}-ketoglutarate, succinate, fumarate, malate, and OAA) were adapted from those previously described for the working rat heart (67–69). Changes made concerned the sample size. In addition, the previously described procedure for citrate cleavage was optimized to the smaller tissue sample (30 mg). Briefly, after tissue homogenization and centrifugation, the supernatants were reacted with sodium borohydride, acidified, and neutralized before incubation with 2.24 ml of 500 mM triethanolamine buffer (pH 7.6) containing 10 mM MgSO4, 20 mM EDTA, 20 mM methoxylamine, and 1.5 g/l of citrate lyase.

Conditions for the operation and analysis of all metabolites as their t-butyldimethylsilyl derivatives by GCMS (Hewlett-Packard 6890N gas chromatograph coupled to a 5973N mass spectrometer) were previously described (67). Areas under each fragmentogram were determined by computer integration and corrected for naturally occurring stable isotopes.

Calculations

Biochemical and functional status. MO2 (in µmol/min), intracellular pH, rate-pressure product (in mmHg·beats·min–1·10–3), cardiac power (in mW), and cardiac efficiency (in mW·µmol–1·min–1) were calculated from previously reported equations (67–69).

Flux parameters. GCMS data are expressed as MPE, as defined previously (19, 20, 68). Briefly, mass isotopomers of metabolites containing 1 to n 13C-labeled atoms were identified as Mi, with i = 1, 2,... n, and the absolute MPE of individual 13C-labeled mass isotopomers (Mi) of a given metabolite was calculated as follows

(1)
where AM and AMi represent the peak areas from ion chromatograms corrected for natural abundance, corresponding to unlabeled (M) and 13C-labeled (Mi) mass isotopomers, respectively. The equations to calculate flux ratios relevant to citrate synthesis in hearts perfused with [U-13C3]lactate, [U-13C3]pyruvate, or [U-13C18]oleate were adapted from those previously described (19, 20, 68) for the use of individual labeling. Equations used for [U-13C3] pyruvate or lactate also apply to [U-13C6]glucose. In brief, flux ratios were calculated from the measured mass isotopomer distribution (MID) of the following tissue metabolites: 1) citrate and its OAA moiety (OAACit), from which we extrapolated the acetyl moiety of citrate (ACCit); 2) pyruvate; and 3) succinate. In this study, we reported the following flux rates, expressed relative to that of citrate synthase (CS): 1) oleate oxidation (OLE): OLE/CS = M2 ACCit/M18 oleate; 2) pyruvate decarboxylation (PDC): PDC/CS = M2 ACCit/M3 pyruvate (Eq. 5 of Ref. 19); 3) pyruvate carboxylation (PC): pyruvate (Eq. 4 of Ref. 19); and 4) the contribution of other substrates (OS), such as endogenous fatty acids and/or amino acids, to the formation of acetyl-CoA: OS/CS = 1 – [(PDC/CS) + (OLE/CS)]. The measured M3 OAACit was corrected for the fraction of M3 OAA molecules coming from citrate isotopomers metabolized in the CAC, as described in Ref. 19 (Eqs. 8–10). To extrapolate the MPE M2 of ACCit from the measured MID of citrate and of its OAA moiety, we used a mathematical approach, previously described by Vincent et al. (67, 69) (Eqs. 2 and 2a)

(2)

The fractional contribution (FC) of individual carbohydrate precursors to pyruvate formation was calculated from the MPE M3 of tissue pyruvate measured in hearts perfused with [U-13C6]glucose, [U-13C]lactate, or [U-13C3]pyruvate by using the following general equation

(3)
where the term Mi refers to the 13C enrichment of the exogenous carbohydrate precursor. Finally, the FC of other carbohydrates (FCother carbohydrates -> pyruvate), such as glycogen and amino acids, was calculated as follows

Lactate and pyruvate uptake and efflux. Lactate and pyruvate uptake and efflux rates were determined by a modification of the procedure described in detail by Vincent et al. (69) for the working rat heart. In brief, rates of uptake and efflux of unlabeled (M) and [U-13C3]labeled (M3) lactate and pyruvate are quantified from 1) the difference between their influent and effluent perfusate concentrations, determined by GCMS and enzymatic assays; and 2) the coronary flow rate. However, the use of three different 13C-labeled carbohydrate substrates in this study enables the determination of additional flux parameters. First, in hearts perfused with unlabeled lactate and pyruvate and [U-13C6]glucose, the efflux rate of [U-13C3]-labeled lactate and pyruvate (M3) reflects that of glycolysis from exogenous glucose. Second, in hearts perfused with unlabeled glucose and lactate and [U-13C3]pyruvate, the rate of efflux of [U-13C3]lactate reflects the rate of conversion of pyruvate into lactate by the LDH reaction. Finally, in hearts perfused with unlabeled glucose and pyruvate and [U-13C3]lactate, the rate of efflux of [U-13C3]pyruvate reflects the rate of conversion of lactate into pyruvate by the LDH reaction. It is noteworthy that with all three 13C-labeled carbohydrates, the rate of efflux of unlabeled lactate and pyruvate reflects more than one metabolic process, namely glycolysis, lactate-pyruvate interconversion by LDH and/or alanine transamination.

Absolute CAC flux rate. The CAC flux rate was calculated from MO2, and the stoichiometric relationships between oxygen consumption and citrate formation, from carbohydrates and fats. The equation of Vincent et al. (Eq. 3 of Ref. 67) was modified to take into account 1) the specific contribution of exogenous oleate to citrate formation, as assessed from the flux ratios OLE/CS; 2) the contribution of other sources, as assessed from the flux ratio OS/CS, which we assumed to be endogenous triglyceride stores consisting of equal proportions of oleate and palmitate; and 3) the contribution of carbohydrates to citrate synthesis, as assessed from the flux ratio PDC/CS and the FC of each carbohydrate to pyruvate synthesis, assuming that other carbohydrate sources are mainly glycogen. That is, we used Eq. 4 below, which considers that 1 µmol of consumed O2 results in the formation of 0.333 µmol of citrate from glucose and lactate and 0.4, 0.353, and 0.348 µmol of citrate from pyruvate, oleate, and palmitate, respectively

(4)

ATP production. Total rates of ATP production were calculated from 1) the various absolute flux rates by using theoretical yields of ATP per mole of substrate oxidized (50), and 2) rates of anaerobic glycolysis determined from the efflux rate of [U-13C3]lactate measured in hearts perfused with [U-13C6]glucose.

Statistical Analysis

Data are expressed as means ± SE of n = 4–15 heart perfusions. Statistical significance was reached at P < 0.05 by using an unpaired t-test or a one-way analysis of variance followed by a Bonferroni selected comparison test.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Functional and Physiological Parameters

During the 30-min perfusion period, isolated working hearts perfused with 11 mM glucose, 0.8 nM insulin, 50 µM carnitine, 1.5 mM lactate, 0.2 mM pyruvate, 5 nM epinephrine, 0.7 mM oleate, and 3% albumin maintained stable values for the various functional and physiological parameters reported in Table 1. HR was stable without the need for external pacing. The integrity of our preparation is indicated by the low values for LDH and succinate release rates, which are used as indexes of necrosis and ischemia, respectively (41, 49, 70). Reducing the oleate concentrations from 0.7 to 0.4 mM did not affect any parameter (data not reported).


View this table:
[in this window]
[in a new window]
  		Table 1. Functional and physiological parameters of ex vivo perfused working mouse hearts

 

Pyruvate Branch Point

The use of individual uniformly labeled carbohydrate substrates allowed the direct assessment of several metabolic flux parameters relevant to pyruvate metabolism in hearts perfused in the presence of 0.7 mM oleate. From the 13C-labeling of tissue pyruvate (reported in Table 2), we can conclude the following. For all three 13C-labeled substrates, tissue pyruvate was only enriched in M3 isotopomers. The absence of M1 or M2 isotopomers (data not reported) indicates negligible decarboxylation of 1) malate to pyruvate through the malic enzyme reaction or 2) glucose in the oxidative pentose phosphate pathway. From the values of MPE M3 of tissue pyruvate, we conclude that, among all carbohydrates, exogenous lactate contributes the most to pyruvate synthesis (42.0 ± 2.3%; Table 2). However, when the contribution of exogenous carbohydrate substrates to pyruvate synthesis is normalized to the pyruvate equivalents produced by these substrates at their respective concentration, it becomes apparent that the contribution of exogenous pyruvate exceeds that of both exogenous lactate and glucose (53.5 ± 2.0 vs. 28.0 ± 1.5 and 1.0 ± 0.1, respectively; Table 2). It is noteworthy that 26.0 ± 0.5% of tissue pyruvate remained unaccounted for and could arise from unlabeled endogenous sources, such as glycogen and alanine.


View this table:
[in this window]
[in a new window]
  		Table 2. Fractional contribution of exogenous carbohydrates to tissue pyruvate and its normalization to pyruvate equivalents

 

Second, from the measurements of 13C enrichment of lactate and pyruvate in the influent and effluent perfusates and of coronary flow rates, we assessed the rates of pyruvate and lactate efflux, uptake, and interconversion by the LDH (Fig. 2). Perfused working mouse hearts took up exogenous pyruvate and lactate as determined by using [U-13C3]pyruvate or [U-13C3]lactate, respectively, as the 13C-labeled substrate (0.11 ± 0.02 and 0.09 ± 0.03 µmol/min, respectively; Fig. 2, B and C). Although lactate and pyruvate were supplied at the physiological concentration ratio of 7.5:1, their uptake rates were similar, indicating a preferential pyruvate uptake. Consequently, the fractional extraction of lactate was significantly less than that of pyruvate (2.9 ± 1.0 and 34.6 ± 3.6%, respectively; P < 0.001), consistent with the results of Table 2. Working mouse hearts perfused with [U-13C3]pyruvate also constantly released unlabeled lactate, represented in Fig. 2B (0.40 ± 0.18 µmol/min). This release of unlabeled lactate reflected glycolysis from exogenous glucose as indicated from the rate of release of [U-13C3]lactate in hearts perfused with [U-13C6]glucose (0.42 ± 0.05 µmol/min; Fig. 2C). Similarly, mouse hearts perfused with [U-13C3]lactate released unlabeled pyruvate at a rate that was similar to the release rate of [U-13C3]pyruvate in hearts perfused with [U-13C6]glucose ([U-13C3]lactate: 0.05 ± 0.05 µmol/min; [U-13C6]glucose: 0.09 ± 0.03 µmol/min). The ratio of lactate to pyruvate produced from glycolysis, an index of the cytosolic redox state, was calculated to be 6.59 ± 0.36, which is within the physiological range.



View larger version (23K):
[in this window]
[in a new window]
  		Fig. 2. Lactate (Lac) and pyruvate (Pyr) uptake and efflux rates assessed in isolated working mouse hearts perfused with 3 different 13C-labeled carbohydrates. Data are means ± SE of 4 heart perfusion experiments, as described in Table 1. Hatched bars are unlabeled substrates, and solid bars are 13C-labeled substrates. The uptake (negative values) and release (positive values) of lactate and pyruvate by the hearts were quantified in hearts perfused with one of the following three different 13C-labeled carbohydrates: [U-13C6]glucose (A), [U-13C3]pyruvate (B), or [U-13C3]lactate (C). Lactate and pyruvate uptake and efflux rates were calculated from the product of values of coronary flow rates and concentration differences in the influent and effluent perfusates, determined by GCMS and enzymatic assays.

 

Finally, the 13C-labeling data provided insights into the dynamics of the LDH reaction. The release rate of [U-13C3]-lactate in hearts perfused with exogenous [U-13C3]pyruvate reflects the rate of conversion of pyruvate to lactate by the LDH (0.08 ± 0.01 µmol/min). Likewise, when [U-13C3]lactate is the 13C labeled substrate, the release rate of [U-13C3]-pyruvate reflects the rate of conversion of lactate to pyruvate by the LDH (0.07 ± 0.01 µmol/min). However, the latter rate is likely to represent a minimal estimate, because intracellular pyruvate has metabolic fates other than its cellular export.

Substrate Selection for Citrate Synthesis

13C-labeling data relevant to the calculation of the relative contribution of substrates to citrate synthesis are shown in Tables 3, 4, 5. Table 3 portrays the MID of the CAC intermediates isolated from mouse hearts perfused with physiological concentrations of [U-13C3]pyruvate, [U-13C3]lactate, [U-13C6]-glucose, or [U-13C18]oleate. Table 4 reports the MPE values for ACCit, the corrected OAACit, and the various flux ratios relevant to pyruvate metabolism. Table 5 reports substrate flux ratios relevant to fatty acid {beta}-oxidation for hearts perfused with the two different oleate concentrations.


View this table:
[in this window]
[in a new window]
  		Table 3. 13C-labeling of CAC intermediates from working mouse hearts perfused with [U-13C3]lactate, [U-13C3]pyruvate, [U-13C6]glucose, or [U-13C18]oleate

 

View this table:
[in this window]
[in a new window]
  		Table 4. Effects of differential labeling and oleate concentration on relative carbohydrate contribution to acetyl-CoA and OAA for citrate synthesis in ex vivo perfused working mouse hearts

 

View this table:
[in this window]
[in a new window]
  		Table 5. Effects of oleate concentration on relative contribution of fatty acids to acetyl-CoA for citrate synthesis

 

Let us first consider 13C-labeling data from hearts perfused with 0.7 mM oleate. From the 13C enrichment data of Table 3, it is apparent that the extent of 13C-labeling of the various CAC intermediates varied with the type of 13C-labeled substrate. This can be explained by differences in the initial 13C enrichment of these substrates (~50% for [U-13C6]glucose, ~25% for [U-13C18]oleate, and >99% for [U-13C3]lactate and [U-13C3]-pyruvate) and their relative concentration as well as their respective contribution to citrate formation. However, for any given 13C-labeled substrate, there were only small differences in the labeling patterns as well as the total 13C-labeling of the various CAC intermediates. In fact, the values of the 13C dilution factors for the various 13C-labeled substrates were similar and indicated little, if any, entry of unlabeled carbon through anaplerosis at sites other than OAA ([U-13C3]lactate: 1.01 ± 0.03; [U-13C3]pyruvate: 0.95 ± 0.13; [U-13C6]glucose: 1.00 ± 0.10; and [U-13C6]oleate: 1.43 ± 0.22; P = not significant; Eq. 10 of Ref. 19).

As shown in Table 4, there was a fairly good agreement in flux values obtained with the three 13C-labeled carbohydrate substrates; small differences between the flux values of PDC/CS and PC/CS did not reach significance. According to the relative flux ratio PDC/CS, pyruvate contributed between 25 and 30% to acetyl-CoA for citrate synthesis. The contribution of anaplerotic pyruvate carboxylation (PC/CS) to OAA for citrate synthesis was on average 5% and was approximately sixfold lower than its decarboxylation. As indicated in Table 5, the oxidation of exogenous oleate represented the predominant source of acetyl-CoA for citrate synthesis (OLE/CS: 63.5 ± 3.9%). The relative contribution of OS, presumably endogenous triglycerides or amino acids, was evaluated to be <10% (OS/CS: 9.8 ± 5.7%).

Lowering of oleate concentration from 0.7 to 0.4 mM resulted in the expected increase in the flux ratio PDC/CS (1.5-fold; Table 4), although it did not reach significance, at the expense of a decreased OLE/CS flux (1.3-fold; Table 5). Interestingly, at 0.4 mM oleate, the relative rate of anaplerotic pyruvate carboxylation was also decreased significantly compared with hearts perfused with 0.7 mM oleate (Table 4). To expand on our finding of a lower flux ratio PC/CS at 0.4 than 0.7 mM oleate, we documented the rates of citrate release and tissue levels of CAC intermediates in both perfusion conditions (Fig. 3). Compared with hearts perfused with 0.7 mM oleate, those perfused with 0.4 mM oleate showed a significantly lower citrate release rate (Fig. 3A) and tissue concentration (Fig. 3B). Tissue concentrations of other CAC intermediates did not vary with oleate concentration.



View larger version (17K):
[in this window]
[in a new window]
  		Fig. 3. Effects of oleate concentration on citrate release rates and citric acid cycle (CAC) intermediates tissue concentrations. Data are means ± SE of 8–13 heart perfusion experiments. Hearts were perfused with either 0.7 (solid bars; n = 13) or 0.4 mM (hatched bars; n = 8) oleate and other substrates-hormones listed in Table 1. A: citrate release rates were quantified in effluent perfusate samples collected between 25 and 30 min by isotope dilution GCMS and flow rate measurements. B: tissue levels of CAC intermediates were quantitated by GCMS in tissue homogenates spiked with standards. {alpha}-KG, {alpha}-ketoglutarate. Inset: isocitrate tissue levels with the use of a magnified scale. *P < 0.05 and ***P < 0.001 vs. 0.7 mM.

 


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
With the use of a 13C-labeling methodology, this study profiles metabolic pathway fluxes linked to energy production in the working mouse heart perfused ex vivo under conditions in which cardiac flows and work as well as the buffer substrate-hormone composition mimic those prevailing in vivo. In addition to measurements of exogenous glucose (glycolysis, oxidation) and fatty acid utilization for energy production, our 13C-labeling strategy provides for the first time quantitative data on 1) the contribution of various exogenous carbohydrate sources to pyruvate synthesis; 2) the rates of pyruvate and lactate efflux, uptake, and interconversion by the LDH; 3) the cytosolic redox state; and 4) the pyruvate anaplerosis and citrate efflux. Our data substantiate the notions of a predominant use of fatty acids over glucose and a regulation of substrate fuel selection according to the Randle cycle (27). However, more importantly, they reveal new information on the metabolism of other exogenous substrates, namely, lactate and pyruvate. As discussed sequentially in greater detail below, we show that these substrates contribute significantly to energy production and participate in anaplerosis. Furthermore, with the use of MO2 values, our relative flux values are converted into quantitative flux rates to enable comparisons with previous studies using radioactive substrates (1, 8).

Our results demonstrate a substantial role for lactate and pyruvate as energy fuels and underscore the dynamic aspects of the pyruvate metabolic branch point in the mouse heart. These concepts were supported by the measured 13C enrichments of 1) tissue pyruvate as well as 2) influent (arterial) and effluent (venous) lactate and pyruvate with the three different 13C-labeled carbohydrates. From the former data, we calculated the relative contributions of exogenous lactate, pyruvate, and glucose to tissue pyruvate formation (Table 2). Clearly, exogenous lactate is the primary contributor to tissue pyruvate formation (42.0 ± 2.3%). Interestingly, when data were normalized to pyruvate molar equivalents produced by these substrates and expressed as a percentage, exogenous pyruvate became the predominant contributor (>50%). The importance of exogenous pyruvate as an energy substrate for the heart when compared with lactate is also emphasized by their similar uptake rates despite a 7.5-fold difference in their "arterial" perfusate concentration. A higher percentage of pyruvate extraction relative to that of lactate, which was also observed by Laughlin et al. (42) in vivo in the dog heart and emphasized by Lloyd et al. (45) in perfused rat hearts, can possibly be explained by the capacity of pyruvate to inhibit both lactate transport (34) and its metabolism by LDH (21). It is noteworthy that the contribution of exogenous glucose to tissue pyruvate (21.3 ± 3.0%) is subject to some uncertainties. It may represent a minimal estimate. Consequently, the contribution of endogenous sources, possibly glycogen or amino acids, to tissue pyruvate (26%) would be overestimated. Indeed, the [13C]glucose enrichment could be diluted if glucose was incorporated into glycogen, estimated to be ~11.6% (35), before its conversion to tissue pyruvate. To further clarify the relative contributions of exogenous glucose and endogenous glycogen to tissue pyruvate formation, one would need to design a different 13C-labeling protocol. Furthermore, potential compartmentation of glycogen metabolism should be taken into account in the data interpretation (35).

The aforementioned data on the contribution of exogenous glucose, lactate, and pyruvate to tissue pyruvate provided, however, only a partial picture of the complexity of the metabolic trafficking at the pyruvate branch point. Indeed, as revealed by measurements of the 13C enrichments and concentrations of lactate and pyruvate in the influent and effluent perfusates, the ex vivo perfused mouse heart simultaneously releases and takes up lactate and pyruvate. These results in the mouse heart are consistent with those reported by others both in vivo and ex vivo in humans, lambs, and rats (2, 6, 16, 29, 67). However, by labeling three different carbohydrates in this study, we demonstrate with precision that the metabolic origin of the lactate released by the heart is predominantly exogenous glucose. In fact, when hearts were perfused with [U-13C6]-glucose, rates of [U-13C3]lactate production were similar to rates of unlabeled lactate obtained when [U-13C3]pyruvate was the labeled substrate. These findings are in agreement with the proposed compartmentation of lactate metabolism in the heart (11, 16, 35).

Let us now consider the relative contribution of exogenous 13C-labeled pyruvate, lactate, and glucose to citrate synthesis. With these three 13C-labeled substrates, the 13C-labeling of the tissue ACCit and OAACit (Table 4) showed a pattern similar to that observed for tissue pyruvate (Table 2). The contribution of carbohydrate oxidation to acetyl-CoA formation for citrate synthesis, as reflected by the flux ratio PDC/CS (on average 27%), is approximately twofold lower than that of fatty acids (63.5 ± 3.9%; Table 5). These patterns of substrate selection for energy production concur with those observed by Belke et al. (8) using radioisotopes. To enable a direct comparison of our flux data obtained with 13C-labeling methods and expressed as relative flux rates (i.e., flux ratios) with those of Belke et al. (8), which were obtained with 14C-labeling methods and expressed as absolute oxidation rates, our relative flux rates were converted to absolute values by using values of MO2, expressed in a heart weight-specific manner [6.25 ± 0.40 µmol·min–1·g wet wt–1]. The calculation is based on theoretical stoichiometric equations for complete substrate oxidation. The following values (in µmol·min–1·g dry wt–1) were calculated by assuming a conversion factor of 0.2 for dry-to-wet weight conversion (8) and an average wet weight for the mouse heart of 0.24 ± 0.01 g: 1) CAC flux rate: 9.4 ± 1.4; 2) oleate {beta}-oxidation: 0.7 ± 0.3; and 3) pyruvate decarboxylation: 2.5 ± 0.5. Converting these absolute flux rates to rates of ATP production revealed a pattern of substrate contributions, which was similar to that observed with data expressed as flux ratios (Fig. 4). It is noteworthy that the total mitochondrial ATP production rate calculated from the extrapolated absolute substrate oxidation rates was similar to that calculated from the theoretical ATP/O ratio of 2.83 (50) and MO2 (8.1 ± 1.3 vs. 8.3 ± 0.6 µmol/min, respectively).



View larger version (26K):
[in this window]
[in a new window]
  		Fig. 4. Relative contribution of various substrates to acetyl-CoA and ATP production. Data are means ± SE of 4–15 heart perfusion experiments, as described in Table 1. The contribution of carbohydrates (lactate, pyruvate, and glucose; closed bars), exogenous fatty acids (oleate; hatched bars), and other sources (possibly endogenous triglycerides or proteins; open bars) to energy production is depicted as follows. A: contribution to acetyl moiety of citrate, calculated from the flux ratios pyruvate decarboxylation to citrate synthase, oleate oxidation to citrate synthase, and other substrate oxidation to citrate synthase, respectively. B: contribution to ATP production, calculated from 1) flux ratios, myocardial oxygen consumption values, and Eq. 4 for mitochondrial oxidative phosphorylation; and 2) lactate release rates for anaerobic glycolysis (shaded bar).

 

Overall, our range of values for absolute substrate oxidation rates and for substrate contributions to ATP synthesis concurs with data previously reported for perfused working mouse hearts (1, 8). However, there are some slight differences. For example, the pattern of substrate selection for ATP production that we observed at the physiological concentration of 0.7 mM oleate (~62% fatty acid and ~34% carbohydrates) was closer to that observed by Belke et al. (8) when palmitate was added at 0.4 mM. Moreover, our glycolytic rates are also superior to those of Belke et al., except in one instance when insulin was added to their buffer. To explain this variability, the following considerations should be taken into account. In our perfusion buffer, we routinely added physiological concentrations of the following substrates and hormones: lactate, pyruvate, carnitine, insulin, and epinephrine. On the one hand, the last four factors have been reported to individually enhance cardiac function, glycolysis, and glucose oxidation (7, 22, 23, 36, 46, 55). Specifically, at a physiological concentration, carnitine mimics insulin-like effects on glycolysis and carbohydrate oxidation (55). Lactate, on the other hand, can compete effectively with fatty acids for oxidation (65), possibly through its capacity to stimulate cardiac acetyl-CoA carboxylase activity, which gives rise to elevated levels of cytoplasmic malonyl-CoA, a known inhibitor of the mitochondrial fatty acid transporter (carnitine palmitoyltransferase-I) (9). Insulin also inhibits fatty acid oxidation through a similar mechanism (26).

Taken as a whole, our data suggest that substrate selection for energy production in the mouse heart resembles that of other species such as dogs (48), pigs (52), and humans (4). However, it appears to differ somewhat from that of the rat heart (67). When perfused with a similar mixture containing 13C-labeled substrates as in our study (including lactate, pyruvate, carnitine, insulin, and 0.4 mM oleate), working rat hearts show a higher contribution of carbohydrates than fatty acids to ATP synthesis (60 ± 4 and 30 ± 4% compared with 34 ± 4 and 62 ± 10% in mice, respectively) (67). Despite this different pattern of substrate selection, ex vivo perfused rat hearts and mouse hearts have similar CAC flux rates (1.7 ± 0.2 for the rat vs. 1.9 ± 0.3 µmol·min–1·g wet wt–1) and cardiac efficiency (1.7 ± 0.2 for the rat vs. 1.8 ± 0.1 mW·µmol–1·min–1 for the mouse).

In addition to providing data on substrate selection for ATP synthesis, we also report for the first time metabolic data on pyruvate anaplerosis and citrate efflux. In fact, using 13C-labeled carbohydrates, we were able to evaluate the contribution of pyruvate to OAA formation necessary for citrate synthesis, referred to as anaplerosis (PC/CS; Table 4). We show that anaplerosis through pyruvate carboxylation is active in the perfused mouse heart, representing ~5% of CAC flux. Measured rates expressed as a percentage of the CAC flux rate or as absolute values (0.09 ± 0.03 µmol·min–1·g wet wt–1) are comparable to those found in the perfused pig and rat hearts (52, 67). Furthermore, the mouse heart perfused with a physiological substrate mixture releases small amounts of citrate at a constant rate of 17 ± 1 nmol·min–1·g wet wt–1 that is similar to that of perfused pig (12 ± 7 nmol·min–1·g wet wt–1) (52) and rat hearts (8 ± 1 nmol·min–1·g wet wt–1) (67). Also similar to pig and rat hearts is the observation that citrate efflux rate in the mouse heart represents 18% of the rate of pyruvate carboxylation, suggesting the presence of other sites for efflux of CAC intermediates. In fact, our results suggest that succinate is another site, representing ~13% of anaplerosis. Another potential site is {alpha}-ketoglutarate.

Over the past several years, a series of studies using 13C-labeled substrates were conducted to further understand the role and regulation of pyruvate anaplerosis and citrate cataplerosis in the regulation of the pool size of CAC intermediates and fuel selection in the heart (19, 20, 51, 52, 67–69). Anaplerotic pyruvate carboxylation, which was shown to be essential for the maintenance of cardiac function (30, 58), was found to be inhibited by high fatty acids (67) and hibernation (51), whereas citrate efflux was reduced in hibernation (51), increased in spontaneously hypertensive rat hearts (69), and modulated by substrate supply (67, 68). In this study conducted in the working mouse heart, lowering of oleate concentration from 0.7 to 0.4 mM decreased anaplerosis, citrate efflux, and tissue citrate concentration. These data substantiate the importance of substrate supply as a determining factor in the regulation of tissue citrate pool size by cardiac pyruvate anaplerosis and citrate efflux.

We recognize that several considerations must be kept in mind in our study. First, although the perfusion system was optimized to obtain cardiac functions similar to those in vivo (28), the HR (between 318 and 428 beats/min) remained slightly below the normal resting physiological values in vivo; nevertheless it was in the upper range of that reported for other ex vivo working mouse heart preparations (1, 8, 24, 31, 32). In fact, because the isolated heart is denervated, it is difficult to obtain in vivo values of HRs without external pacing. Second, due to the greater cost associated with the use of Millar catheter system in the mouse heart, as well as its fragility at such a small size, we instead chose to use a fluid-filled catheter inserted at the apex of the heart for the measurements of left ventricular function. By doing so, we are limited to a low resonant frequency, which could lead to slight underestimation of some functional values, such as the first derivative of maximum left ventricular systolic pressure. Nevertheless, we routinely obtained values of 6,280 ± 270 mmHg/s for this parameter (data not reported) (28). Finally, concerning our 13C-labeling data, we cannot exclude the possibility that the small, but not significant, differences in the flux ratios PC/PDC (Table 4) calculated from individual 13C-labeled carbohydrates may be due to compartmentation of glucose and pyruvate metabolism (12, 35, 43, 53).

In conclusion, this study, based on measurements of substrate fluxes using 13C-labeling methods, provides a detailed quantitative profile of the fate and origin of pyruvate and citrate carbons in the working mouse heart perfused ex vivo under conditions in which cardiac flows and work as well as the buffer substrate-hormone composition mimic those prevailing in vivo. We believe that the described strategy of combining 13C-labeled substrates with an experimental system enabling the continuous monitoring of cardiac mechanics offers a unique and powerful tool for clarifying the link between metabolic alterations and cardiac contractile dysfunction. This should also allow dynamic flux data to be integrated with static data of metabolite concentrations, enzyme activities, protein levels, and gene expression, and provides an integrative view of the regulation of metabolic pathways at all levels, namely, posttranslational (acute) and transcriptional (chronic) levels. Finally, this methodology should provide valuable insights when applied to genetically altered mouse models of disease.


   ACKNOWLEDGMENTS
 
The authors thank Johanne Bourdon and Duska Gvozdic for assistance with animal care.

This work was presented at the Heart Failure 2003 Congress in Strasbourg and at the Society Heart and Vascular Metabolism in Freiburg in June 2003.

Present address of F. Labarthe: EMI-U 02-11, Laboratoire Nutrition, Croissance et Cancer, Faculté de Médecine, 2 Bis Blvd. Tonnellé, 37032 Tours Cedex, France.

GRANTS

This study was supported by Canadian Institutes of Health Research Grants 9575 and 201819 (to C. Des Rosiers) and a studentship (to M. Khairallah).


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. Des Rosiers, Laboratory of Intermediary Metabolism, CHUM Research Center, Notre-Dame Hospital, 1560 Sherbrooke East, Rm. Y-3616, Montreal, Quebec, Canada H2L 4M1 (E-mail: christine.des.rosiers{at}umontreal.ca).

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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Aasum E, Belke DD, Severson DL, Riemersma RA, Cooper M, Andreassen M, and Larsen TS. Cardiac function and metabolism in Type 2 diabetic mice after treatment with BM 17.0744, a novel PPAR-alpha activator. Am J Physiol Heart Circ Physiol 283: H949–H957, 2002.[Abstract/Free Full Text]
  2. Allard MF, Schonekess BO, Henning SL, English DR, and Lopaschuk GD. Contribution of oxidative metabolism and glycolysis to ATP production in hypertrophied hearts. Am J Physiol Heart Circ Physiol 267: H742–H750, 1994.[Abstract/Free Full Text]
  3. Amrani A, Durant S, Throsby M, Coulaud J, Dardenne M, and Homo-Delarche F. Glucose homeostasis in the nonobese diabetic mouse at the prediabetic stage. Endocrinology 139: 1115–1124, 1998.[Abstract/Free Full Text]
  4. Ballard FB, Danforth WH, Naegle S, and Bing RJ. Myocardial metabolism of fatty acids. J Clin Invest 39: 717–730, 1960.[Web of Science][Medline]
  5. Barger PM and Kelly DP. Fatty acid utilization in the hypertrophied and failing heart: molecular regulatory mechanisms. Am J Med Sci 318: 36–42, 1999.[CrossRef][Web of Science][Medline]
  6. Bartelds B, Knoester H, Beaufort-Krol GC, Smid GB, Takens J, Zijlstra WG, Heymans HS, and Kuipers JR. Myocardial lactate metabolism in fetal and newborn lambs. Circulation 99: 1892–1897, 1999.[Abstract/Free Full Text]
  7. Belke DD, Betuing S, Tuttle MJ, Graveleau C, Young ME, Pham M, Zhang D, Cooksey RC, McClain DA, Litwin SE, Taegtmeyer H, Severson D, Kahn CR, and Abel ED. Insulin signaling coordinately regulates cardiac size, metabolism, and contractile protein isoform expression. J Clin Invest 109: 629–639, 2002.[CrossRef][Web of Science][Medline]
  8. Belke DD, Larsen TS, Lopaschuk GD, and Severson DL. Glucose and fatty acid metabolism in the isolated working mouse heart. Am J Physiol Regul Integr Comp Physiol 277: R1210–R1217, 1999.[Abstract/Free Full Text]
  9. Bielefeld DR, Vary TC, and Neely JR. Inhibition of carnitine palmitoyl-CoA transferase activity and fatty acid oxidation by lactate and oxfenicine in cardiac muscle. J Mol Cell Cardiol 17: 619–625, 1985.[Web of Science][Medline]
  10. Bockamp E, Maringer M, Spangenberg C, Fees S, Fraser S, Eshkind L, Oesch F, and Zabel B. Of mice and models: improved animal models for biomedical research. Physiol Genomics 11: 115–132, 2002.[Abstract/Free Full Text]
  11. Brooks GA. Intra- and extra-cellular lactate shuttles. Med Sci Sports Exerc 32: 790–799, 2000.
  12. Bunger R. Compartmented pyruvate in perfused working heart. Am J Physiol Heart Circ Physiol 249: H439–H449, 1985.[Abstract/Free Full Text]
  13. Bunger R, Mallet RT, and Hartman DA. Pyruvate-enhanced phosphorylation potential and inotropism in normoxic and postischemic isolated working heart. Near-complete prevention of reperfusion contractile failure. Eur J Biochem 180: 221–233, 1989.[Web of Science][Medline]
  14. Carvajal K and Moreno-Sanchez R. Heart metabolic disturbances in cardiovascular diseases. Arch Med Res 34: 89–99, 2003.[CrossRef][Web of Science][Medline]
  15. Chatham JC, Bouchard B, and Des Rosiers C. A comparison between NMR and GCMS 13C-isotopomer analysis in cardiac metabolism. Mol Cell Biochem 249: 105–112, 2003.[CrossRef][Web of Science][Medline]
  16. Chatham JC, Des Rosiers C, and Forder JR. Evidence of separate pathways for lactate uptake and release by the perfused rat heart. Am J Physiol Endocrinol Metab 281: E794–E802, 2001.[Abstract/Free Full Text]
  17. Chatham JC, Gao ZP, and Forder JR. Impact of 1 wk of diabetes on the regulation of myocardial carbohydrate and fatty acid oxidation. Am J Physiol Endocrinol Metab 277: E342–E351, 1999.[Abstract/Free Full Text]
  18. Coggan AR. Use of stable isotopes to study carbohydrate and fat metabolism at the whole-body level. Proc Nutr Soc 58: 953–961, 1999.[Web of Science][Medline]
  19. Comte B, Vincent G, Bouchard B, and Des Rosiers C. Probing the origin of acetyl-CoA and oxaloacetate entering the citric acid cycle from the 13C labeling of citrate released by perfused rat hearts. J Biol Chem 272: 26117–26124, 1997.[Abstract/Free Full Text]
  20. Comte B, Vincent G, Bouchard B, Jette M, Cordeau S, and Des Rosiers C. A 13C mass isotopomer study of anaplerotic pyruvate carboxylation in perfused rat hearts. J Biol Chem 272: 26125–26131, 1997.[Abstract/Free Full Text]
  21. Coulson CJ and Rabin BR. Inhibition of lactate dehydrogenase by high concentrations of pyruvate: the nature and removal of the inhibitor. FEBS Lett 3: 333–337, 1969.[CrossRef][Web of Science][Medline]
  22. Depre C, Rider MH, and Hue L. Mechanisms of control of heart glycolysis. Eur J Biochem 258: 277–290, 1998.[Web of Science][Medline]
  23. Depre C, Vanoverschelde JL, and Taegtmeyer H. Glucose for the heart. Circulation 99: 578–588, 1999.[Free Full Text]
  24. De Windt LJ, Willems J, Reneman RS, van der Vusse GJ, Arts T, and van Bilsen M. An improved isolated, left ventricular ejecting, murine heart model. Functional and metabolic evaluation. Pflügers Arch 437: 182–190, 1999.[CrossRef][Web of Science][Medline]
  25. Fan W, Dinulescu DM, Butler AA, Zhou J, Marks DL, and Cone RD. The central melanocortin system can directly regulate serum insulin levels. Endocrinology 141: 3072–3079, 2000.[Abstract/Free Full Text]
  26. Gamble J and Lopaschuk GD. Insulin inhibition of 5' adenosine monophosphate-activated protein kinase in the heart results in activation of acetyl coenzyme A carboxylase and inhibition of fatty acid oxidation. Metabolism 46: 1270–1274, 1997.[CrossRef][Web of Science][Medline]
  27. Garland PB, Newsholme EA, and Randle PJ. Regulation of glucose uptake by muscle. Biochem J 93: 665–678, 1964.[Web of Science][Medline]
  28. Georgakopoulos D, Mitzner WA, Chen CH, Byrne BJ, Millar HD, Hare JM, and Kass DA. In vivo murine left ventricular pressure-volume relations by miniaturized conductance micromanometry. Am J Physiol Heart Circ Physiol 274: H1416–H1422, 1998.[Abstract/Free Full Text]
  29. 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.[Web of Science][Medline]
  30. Gibala MJ, Young ME, and Taegtmeyer H. Anaplerosis of the citric acid cycle: role in energy metabolism of heart and skeletal muscle. Acta Physiol Scand 168: 657–665, 2000.[CrossRef][Web of Science][Medline]
  31. Grupp IL, Lorenz JN, Walsh RA, Boivin GP, and Rindt H. Overexpression of alpha1B-adrenergic receptor induces left ventricular dysfunction in the absence of hypertrophy. Am J Physiol Heart Circ Physiol 275: H1338–H1350, 1998.[Abstract/Free Full Text]
  32. Grupp IL, Subramaniam A, Hewett TE, Robbins J, and Grupp G. Comparison of normal, hypodynamic, and hyperdynamic mouse hearts using isolated work-performing heart preparations. Am J Physiol Heart Circ Physiol 265: H1401–H1410, 1993.[Abstract/Free Full Text]
  33. Hajri T, Ibrahimi A, Coburn CT, Knapp FF Jr, Kurtz T, Pravenec M, and Abumrad NA. Defective fatty acid uptake in the spontaneously hypertensive rat is a primary determinant of altered glucose metabolism, hyperinsulinemia, and myocardial hypertrophy. J Biol Chem 276: 23661–23666, 2001.[Abstract/Free Full Text]
  34. Henderson AH, Craig RJ, Gorlin R, and Sonnenblick EH. Lactate and pyruvate kinetics in isolated perfused rat hearts. Am J Physiol 217: 1752–1756, 1969.[Free Full Text]
  35. Henning SL, Wambolt RB, Schonekess BO, Lopaschuk GD, and Allard MF. Contribution of glycogen to aerobic myocardial glucose utilization. Circulation 93: 1549–1555, 1996.[Abstract/Free Full Text]
  36. Hiraoka T, DeBuysere M, and Olson MS. Studies of the effects of {beta}-adrenergic agonists on the regulation of pyruvate dehydrogenase in the perfused rat heart. J Biol Chem 255: 7604–7609, 1980.[Free Full Text]
  37. Hunter JJ, Zhu H, Lee KJ, Kubalak S, and Chien KR. Targeting gene expression to specific cardiovascular cell types in transgenic mice. Hypertension 22: 608–617, 1993.[Abstract/Free Full Text]
  38. Kantor PF, Lucien A, Kozak R, and Lopaschuk GD. The antianginal drug trimetazidine shifts cardiac energy metabolism from fatty acid oxidation to glucose oxidation by inhibiting mitochondrial long-chain 3-ketoacyl coenzyme A thiolase. Circ Res 86: 580–588, 2000.[Abstract/Free Full Text]
  39. Kemppainen J, Fujimoto T, Kalliokoski KK, Viljanen T, Nuutila P, and Knuuti J. Myocardial and skeletal muscle glucose uptake during exercise in humans. J Physiol 542: 403–412, 2002.[Abstract/Free Full Text]
  40. Khogali SE, Harper AA, Lyall JA, and Rennie MJ. Effects of L-glutamine on post-ischaemic cardiac function: protection and rescue. J Mol Cell Cardiol 30: 819–827, 1998.[CrossRef][Web of Science][Medline]
  41. Laplante A, Vincent G, Poirier M, and Des Rosiers C. Effects and metabolism of fumarate in the perfused rat heart A 13C mass isotopomer study. Am J Physiol Endocrinol Metab 272: E74–E82, 1997.[Abstract/Free Full Text]
  42. Laughlin MR, Taylor J, Chesnick AS, DeGroot M, and Balaban RS. Pyruvate and lactate metabolism in the in vivo dog heart. Am J Physiol Heart Circ Physiol 264: H2068–H2079, 1993.[Abstract/Free Full Text]
  43. Lewandowski ED. Metabolic heterogeneity of carbon substrate utilization in mammalian heart: NMR determinations of mitochondrial versus cytosolic compartmentation. Biochemistry 31: 8916–8923, 1992.[CrossRef][Medline]
  44. Lewis W, Haase CP, Raidel SM, Russ RB, Sutliff RL, Hoit BD, and Samarel AM. Combined antiretroviral therapy causes cardiomyopathy and elevates plasma lactate in transgenic AIDS mice. Lab Invest 81: 1527–1536, 2001.[Web of Science][Medline]
  45. Lloyd S, Brocks C, and Chatham JC. Differential modulation of glucose, lactate, and pyruvate oxidation by insulin and dichloroacetate in the rat heart. Am J Physiol Heart Circ Physiol 285: H163–H172, 2003.[Abstract/Free Full Text]
  46. Mallet RT. Pyruvate: metabolic protector of cardiac performance. Proc Soc Exp Biol Med 223: 136–148, 2000.[Abstract/Free Full Text]
  47. Malloy CR, Sherry AD, and Jeffrey FM. Carbon flux through citric acid cycle pathways in perfused heart by 13C NMR spectroscopy. FEBS Lett 212: 58–62, 1987.[CrossRef][Web of Science][Medline]
  48. Myears DW, Sobel BE, and Bergmann SR. Substrate use in ischemic and reperfused canine myocardium: quantitative considerations. Am J Physiol Heart Circ Physiol 253: H107–H114, 1987.[Abstract/Free Full Text]
  49. Nishimura M, Takami H, Kaneko M, Nakano S, Matsuda H, Kurosawa K, Inoue T, and Tagawa K. Mechanism of mitochondrial enzyme leakage during reoxygenation of the rat heart. Cardiovasc Res 27: 1116–1122, 1993.[Abstract/Free Full Text]
  50. Opie LH. The Heart: Physiology, From Cell to Circulation (3rd ed.). New York: Lippincott-Raven, 1998, p. 304.
  51. Panchal AR, Comte B, Huang H, Dudar B, Roth B, Chandler M, Des Rosiers C, Brunengraber H, and Stanley WC. Acute hibernation decreases myocardial pyruvate carboxylation and citrate release. Am J Physiol Heart Circ Physiol 281: H1613–H1620, 2001.[Abstract/Free Full Text]
  52. Panchal AR, Comte B, Huang H, Kerwin T, Darvish A, Des Rosiers C, Brunengraber H, and Stanley WC. Partitioning of pyruvate between oxidation and anaplerosis in swine hearts. Am J Physiol Heart Circ Physiol 279: H2390–H2398, 2000.[Abstract/Free Full Text]
  53. Peuhkurinen KJ, Hiltunen JK, and Hassinen IE. Metabolic compartmentation of pyruvate in the isolated perfused rat heart. Biochem J 210: 193–198, 1983.[Web of Science][Medline]
  54. Ren JM, Marshall BA, Mueckler MM, McCaleb M, Amatruda JM, and Shulman GI. Overexpression of Glut4 protein in muscle increases basal and insulin-stimulated whole body glucose disposal in conscious mice. J Clin Invest 95: 429–432, 1995.[Web of Science][Medline]
  55. Rodgers RL, Christe ME, Tremblay GC, Babson JR, and Daniels T. Insulin-like effects of a physiologic concentration of carnitine on cardiac metabolism. Mol Cell Biochem 226: 97–105, 2001.[CrossRef][Web of Science][Medline]
  56. Roe CR, Sweetman L, Roe DS, David F, and Brunengraber H. Treatment of cardiomyopathy and rhabdomyolysis in long-chain fat oxidation disorders using an anaplerotic odd-chain triglyceride. J Clin Invest 110: 259–269, 2002.[CrossRef][Web of Science][Medline]
  57. Rupp H, Schulze W, and Vetter R. Dietary medium-chain triglycerides can prevent changes in myosin and SR due to CPT-1 inhibition by etomoxir. Am J Physiol Regul Integr Comp Physiol 269: R630–R640, 1995.[Abstract/Free Full Text]
  58. Russell RR, III, and Taegtmeyer H. Pyruvate carboxylation prevents the decline in contractile function of rat hearts oxidizing acetoacetate. Am J Physiol Heart Circ Physiol 261: H1756–H1762, 1991.[Abstract/Free Full Text]
  59. Sabbah HN, Chandler MP, Mishima T, Suzuki G, Chaudhry P, Nass O, Biesiadecki BJ, Blackburn B, Wolff A, and Stanley WC. Ranolazine, a partial fatty acid oxidation (pFOX) inhibitor, improves left ventricular function in dogs with chronic heart failure. J Card Fail 8: 416–422, 2002.[CrossRef][Web of Science][Medline]
  60. Sambandam N and Lopaschuk GD. AMP-activated protein kinase (AMPK) control of fatty acid and glucose metabolism in the ischemic heart. Prog Lipid Res 42: 238–256, 2003.[CrossRef][Web of Science][Medline]
  61. Stanley WC. Partial fatty acid oxidation inhibitors for stable angina. Expert Opin Investig Drugs 11: 615–629, 2002.[CrossRef][Web of Science][Medline]
  62. Taegtmeyer H, Razeghi P, and Young ME. Mitochondrial proteins in hypertrophy and atrophy: a transcript analysis in rat heart. Clin Exp Pharmacol Physiol 29: 346–350, 2002.[CrossRef][Web of Science][Medline]
  63. Tejero-Taldo MI, Caffrey JL, Sun J, and Mallet RT. Antioxidant properties of pyruvate mediate its potentiation of {beta}-adrenergic inotropism in stunned myocardium. J Mol Cell Cardiol 31: 1863–1872, 1999.[CrossRef][Web of Science][Medline]
  64. Tsunoda M, Takezawa K, Yanagisawa T, Kato M, and Imai K. Determination of catecholamines and their 3-O-methyl metabolites in mouse plasma. Biomed Chromatogr 15: 41–44, 2001.[CrossRef][Web of Science][Medline]
  65. Van der Vusse GJ and de Groot MJ. Interrelationship between lactate and cardiac fatty acid metabolism. Mol Cell Biochem 116: 11–17, 1992.[CrossRef][Web of Science][Medline]
  66. Van der Vusse GJ, Van Bilsen M, and Glatz JF. Cardiac fatty acid uptake and transport in health and disease. Cardiovasc Res 45: 279–293, 2000.[Abstract/Free Full Text]
  67. Vincent G, Bouchard B, Khairallah M, and Des Rosiers C. Differential modulation of citrate synthesis and release by fatty acids in perfused working rat hearts. Am J Physiol Heart Circ Physiol 286: H257–H267, 2004.[Abstract/Free Full Text]
  68. Vincent G, Comte B, Poirier M, and Des Rosiers C. Citrate release by perfused rat hearts: a window on mitochondrial cataplerosis. Am J Physiol Endocrinol Metab 278: E846–E856, 2000.[Abstract/Free Full Text]
  69. Vincent G, Khairallah M, Bouchard B, and Des Rosiers C. Metabolic phenotyping of the diseased rat heart using 13C-substrates and ex vivo perfusion in the working mode. Mol Cell Biochem 242: 89–99, 2003.[CrossRef][Web of Science][Medline]
  70. Vork MM, Glatz JF, Surtel DA, and van der Vusse GJ. Release of fatty acid binding protein and lactate dehydrogenase from isolated rat heart during normoxia, low-flow ischemia, and reperfusion. Can J Physiol Pharmacol 71: 952–958, 1993.[Web of Science][Medline]
  71. Wolff AA, Rotmensch HH, Stanley WC, and Ferrari R. Metabolic approaches to the treatment of ischemic heart disease: the clinicians' perspective. Heart Fail Rev 7: 187–203, 2002.[CrossRef][Medline]
  72. Wu C, Okar DA, Newgard CB, and Lange AJ. Overexpression of 6-phosphofructo-2-kinase/fructose-2, 6-bisphosphatase in mouse liver lowers blood glucose by suppressing hepatic glucose production. J Clin Invest 107: 91–98, 2001.[Web of Science][Medline]
  73. Yokogawa K, Higashi Y, Tamai I, Nomura M, Hashimoto N, Nikaido H, Hayakawa J, Miyamoto K, and Tsuji A. Decreased tissue distribution of L-carnitine in juvenile visceral steatosis mice. J Pharmacol Exp Ther 289: 224–230, 1999.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 J. Nutr.Home page
S. W. El-Kadi, R. L. Baldwin VI, K. R. McLeod, N. E. Sunny, and B. J. Bequette
Glutamate Is the Major Anaplerotic Substrate in the Tricarboxylic Acid Cycle of Isolated Rumen Epithelial and Duodenal Mucosal Cells from Beef Cattle
J. Nutr., May 1, 2009; 139(5): 869 - 875.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
R. Gelinas, F. Labarthe, B. Bouchard, J. Mc Duff, G. Charron, M. E. Young, and C. Des Rosiers
Alterations in carbohydrate metabolism and its regulation in PPAR{alpha} null mouse hearts
Am J Physiol Heart Circ Physiol, April 1, 2008; 294(4): H1571 - H1580.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
D. An and B. Rodrigues
Role of changes in cardiac metabolism in development of diabetic cardiomyopathy
Am J Physiol Heart Circ Physiol, October 1, 2006; 291(4): H1489 - H1506.
[Abstract] [Full Text] [PDF]

Home page
 Exp Biol MedHome page
R. T. Mallet, J. Sun, E. M. Knott, A. B. Sharma, and A. H. Olivencia-Yurvati
Metabolic Cardioprotection by Pyruvate: Recent Progress
Exp Biol Med, July 1, 2005; 230(7): 435 - 443.
[Abstract] [Full Text] [PDF]

Home page
 Physiol. Rev.Home page
W. C. Stanley, F. A. Recchia, and G. D. Lopaschuk
Myocardial Substrate Metabolism in the Normal and Failing Heart
Physiol Rev, July 1, 2005; 85(3): 1093 - 1129.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
286/4/H1461    most recent
00942.2003v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (16)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Khairallah, M.
Right arrow Articles by Des Rosiers, C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Khairallah, M.
Right arrow Articles by Des Rosiers, C.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2004 by the American Physiological Society.