Over the past 20 years, stable isotopes combined with isotopomer analysis have proven to be a powerful approach to probe the dynamics of metabolism in various biological systems, including the heart. The aim of this paper is to demonstrate how isotopomer analysis of metabolic fluxes can provide novel insights into the myocardial phenotype. Specifically, building on our past experience using NMR spectroscopy and GC–MS as applied to investigations of cardiac energy metabolism, we highlight specific complex metabolic networks that would not be predicted by classical biochemistry or by static measurements of metabolite, protein and mRNA levels.

Introduction

Metabolomics has emerged as one of the major new disciplines of the post-genomic era, along with proteomics and transcriptomics. While the focus of metabolomics has been primarily on high-throughput analyses of metabolite levels, the measurement of metabolic fluxes appears also crucial in predicting the cell phenotype [15]. This information is needed by metabolic bioengineers for optimizing cellular growth and secretion of targeted compounds and by metabolic physiologists who seek to link fuel metabolism to organ function and homoeostasis. Stable isotope and isotopomer analysis by NMR or GC–MS has proven to be a very powerful approach for metabolic flux analysis in various biological systems (for reviews, see [610]). In 2004, Metabolic Engineering devoted its January issue to highlight the challenges of estimating fluxes in mammalian systems, where one has to deal with compartmentation of metabolites, dilution of tracer by endogenous pools and difficulty of sampling relevant pools [3]. In this context, we [11,12] provided an overview of the theory behind metabolic flux analysis using mass isotopomer analysis of cardiac energy metabolism, with an emphasis on the study of the CAC (citric acid cycle). We showed that, although NMR and GC–MS rely on the measurements of different metabolite isotopomer data and different mathematical approaches for the determination of metabolic fluxes, there was remarkably good agreement between substrate flux ratios extrapolated by these two techniques. There were, however, some intriguing discrepancies that emphasized our lack of understanding of the role of compartmentation at the tissue and cellular levels as well as metabolite channelling due to enzyme–enzyme interactions.

The goal of this paper is to expand on these earlier reports and to provide a guide to metabolic physiologists and bioengineers who are considering including NMR- or GC–MS-determined 13C-isotopomer and metabolic flux ratio data as part of a comprehensive model of cardiac metabolism. As ‘metabolic physiologists’ who seek ultimately to link metabolism to organ function and homoeostasis, we will highlight specific metabolic processes that emphasize both the power inherent in 13C-isotopomer analysis as well as the complexity present in apparently simple metabolic networks, specifically the pyruvate branch point, the regulation of PC (pyruvate carboxylation), an anaplerotic reaction, and fatty acid β-oxidation.

Probing the role and regulation of complex metabolic networks using isotopomer analysis

The pyruvate branch point

Tracer studies of cardiac metabolism have typically focused on the contribution of only glucose and fatty acids to energy production despite the evidence that glucose is not the only carbohydrate that contributes to acetyl-CoA production. Indeed, carbohydrates such as lactate and pyruvate represent a significant source of energy for the heart; the precise contribution is very sensitive to changes in substrate availability, hormonal milieu and disease [1319]. The situation becomes even more complex when one considers the intriguing observation that has been repeatedly reported in both in vivo and ex vivo heart studies, that where there is oxidation of exogenous lactate, there is also a simultaneous efflux of lactate originating from the metabolism of exogenous glucose [13,14,18,2022].

A practical consequence of the existence of simultaneous release and uptake of lactate by the heart is illustrated in Figure 1. This Figure depicts the concentrations of unlabelled (12C) and labelled (13C) isotopomers of lactate measured in the influent (arterial) and effluent (venous) perfusates of hearts perfused with a non-recirculating buffer containing U-13C3-labelled lactate and pyruvate, and unlabelled glucose and oleate. In such a study, the lactate uptake rate cannot be simply evaluated from the arterio-venous difference in lactate concentration. Indeed, there are minimal differences between total influent and effluent lactate concentrations despite a significantly higher concentration of unlabelled lactate in the effluent samples.

Concentrations of lactate in the influent (arterial) and effluent (venous) perfusates of working hearts perfused with U-13C3-labelled lactate (1 mM) and pyruvate (0.2 mM), and unlabelled glucose (5.5 mM) and oleate (0.4 mM)

Figure 1
Concentrations of lactate in the influent (arterial) and effluent (venous) perfusates of working hearts perfused with U-13C3-labelled lactate (1 mM) and pyruvate (0.2 mM), and unlabelled glucose (5.5 mM) and oleate (0.4 mM)

Stacked bars depict the concentrations of unlabelled (12C; lower part) and labelled (13C; upper part) metabolites determined by GC–MS. Data are from Vincent et al. [38]. *P<0.05 for effluent versus influent perfusates.

Figure 1
Concentrations of lactate in the influent (arterial) and effluent (venous) perfusates of working hearts perfused with U-13C3-labelled lactate (1 mM) and pyruvate (0.2 mM), and unlabelled glucose (5.5 mM) and oleate (0.4 mM)

Stacked bars depict the concentrations of unlabelled (12C; lower part) and labelled (13C; upper part) metabolites determined by GC–MS. Data are from Vincent et al. [38]. *P<0.05 for effluent versus influent perfusates.

It has been suggested that the release of unlabelled lactate is a result of simple isotope exchange at the level of LDH (lactate dehydrogenase) [23,24], which would reflect the establishment of an isotopic equilibrium between lactate and pyruvate. Although rapid lactate and pyruvate interconversion by LDH clearly takes place [14], data argue against this being the only explanation for the observed lactate uptake and release by the heart. For example, there is evidence to suggest that, in the intact heart, lactate and pyruvate are not at isotopic equilibrium [21], consistent with the existence of separate non-exchanging pools of metabolites. Furthermore, heart perfusion experiments using GC–MS- or NMR-determined 13C-isotopomer analysis revealed that lactate uptake and release rates are differentially modulated under different conditions, such as ischaemia, diabetes and hypertrophy [16,18,21,25].

Altogether, the aforementioned results support a compartmentation of lactate and pyruvate metabolism in the heart, whereby glycolytically derived lactate efflux and exogenous lactate oxidation are functionally separate pathways that can be independently regulated. As discussed previously [21], this could be a result of intracellular compartmentation, as proposed by Brooks [26], or different cell populations in the intact heart or possibly channelling. Clearly, additional investigations are warranted to understand better the importance of this compartmentation of lactate and pyruvate metabolism, as well as its significance for the heart, which may be linked to the regulation of cellular redox potential [21].

Anaplerosis

The term ‘anaplerosis’ was originally coined by Kornberg in 1966 [27] to designate pathways or reactions that replenish the pool of intermediates of a metabolic cycle, such as the CAC (1). The levels of intermediates are crucial for the functioning of the CAC, whose principal role is the oxidation of acetyl-CoA to CO2, a central process in energy metabolism. The entry of catalytic carbons into the CAC by anaplerotic reactions balances the removal of intermediates due to their participation in ancillary reactions. A number of studies underlined the importance of anaplerosis for (i) biosynthetic processes, such as gluconeogenesis in liver [28] or neurotransmitter formation in brain [29], and (ii) glucose-stimulated insulin secretion in pancreatic β-cells [30,31]. However, while the crucial role of anaplerosis for hepatic gluconeogenesis is self-evident, in the heart, as in skeletal muscle, much remains to be learned about the role, regulation and sites of anaplerosis [32].

Anaplerotic pathways in the heart and their interrelationship with reactions that catalyse the removal of intermediates from the CAC

Scheme 1
Anaplerotic pathways in the heart and their interrelationship with reactions that catalyse the removal of intermediates from the CAC

OAA, oxaloacetate; BCAA, branched chain amino acids.

Scheme 1
Anaplerotic pathways in the heart and their interrelationship with reactions that catalyse the removal of intermediates from the CAC

OAA, oxaloacetate; BCAA, branched chain amino acids.

In the early 1980s, radioactive tracer experiments in isolated hearts demonstrated that anaplerotic flux of carbon into the CAC is a normal part of metabolism [33]. In the 1990s, ex vivo perfusion studies revealed a decline in contractile function following depletion of CAC intermediates, a phenomenon that is reversed by the addition of anaplerotic substrates [34]. In addition, investigations suggested that stimulating anaplerosis from exogenous substrates, such as pyruvate and propionyl-CoA precursors, could become part of the treatment of myocardial reperfusion injury and other cardiomyopathies [32,35,36]. More recently, research on cardiac anaplerosis has benefited from 13C-isotopomer analysis of CAC intermediates by GC–MS [14,19,35,37,38] and of glutamate by NMR [15,39]. Note that, with GC–MS, the anaplerotic flux is determined with a good precision and is specific for PC. In contrast, with NMR, the site of entry of carbon into the CAC is not defined; however, it includes all possible sources of anaplerosis.

Numerous studies of anaplerosis have been conducted using 13C-isotopomer analysis by GC–MS, in various models, including ex vivo Langendorff and work-performing perfused rat and mice hearts and in situ pig hearts [14,19,25,37,38]. Taken together, these studies demonstrated that, in the heart, anaplerotic PC represents between 3 and 12% of citrate synthase flux and is subject to regulation by substrate and oxygen supply, as well as disease. 13C-NMR studies also revealed that anaplerosis was enhanced by insulin [15]. GC–MS analysis of 13C-labelling of tissue CAC intermediates provided additional information regarding the sites and extent of anaplerosis other than PC [18,19,38]. Data from GC–MS studies can also be used to test the relationship between measured anaplerotic flux ratios and tissue levels of CAC intermediates under various conditions of substrate supply and energy demand. On the basis of our current understanding of the role of anaplerosis in the heart, one would expect parallel variations between these two parameters, but as illustrated by the following data taken from recent publications [14,19], this is not always the case.

Figure 2 depicts the effects of increasing exogenous fatty acid concentrations on the tissue levels of CAC intermediates and the anaplerotic PC flux ratio measured from 13C-isotopomer analysis by GC–MS in work-performing mice heart (Figures 2A and 2B) and rat heart (Figures 2C and 2D), perfused with physiological concentrations of substrates and hormones (glucose, lactate, pyruvate, oleate and insulin). Note that, in both animal models, raising the fatty acid concentration, as expected, inhibited pyruvate decarboxylation, enhanced oleate β-oxidation, and increased tissue levels of CAC intermediates, principally citrate. However, while the anaplerotic PC flux ratio was slightly increased in mouse hearts as might be expected, it was lowered 2.4-fold in the rat hearts. This finding of an inhibition of the PC flux ratio by high fatty acid concentration was unexpected. Indeed, according to the prevailing view, acetyl-CoA, which would be expected to increase with fatty acid concentrations, stimulates pyruvate carboxylase activity. However, there is also one report suggesting that the activity of this enzyme may be inhibited at acetyl-CoA levels above 200 μM [40]. Overall, these results emphasize the complexity of inter-regulatory mechanisms governing flux through anaplerotic PC and maintaining CAC pool size in the heart as well as the need to combine isotopomer and concentration data to obtain a comprehensive view of metabolic regulation. Clearly, additional investigations appear warranted to increase our understanding of factors involved in regulation of anaplerosis given that it has been linked to cardioprotection [36].

Effects of increasing exogenous oleate concentrations on tissue concentrations of individual CAC intermediates (A, C) and the anaplerotic PC/citrate synthase flux ratio (B, D) in working mice (A, B) and rat (C, D) heart perfusion experiments

Figure 2
Effects of increasing exogenous oleate concentrations on tissue concentrations of individual CAC intermediates (A, C) and the anaplerotic PC/citrate synthase flux ratio (B, D) in working mice (A, B) and rat (C, D) heart perfusion experiments

Results are means±S.E.M. Data from mice hearts perfused with U-13C3-labelled pyruvate (0.2 mM), and unlabelled lactate (1.5 mM), glucose (11 mM) and oleate (0.4 or 0.7 mM) are from Khairallah et al. [14]. Data from rat hearts perfused with U-13C3-labelled pyruvate (0.2 mM) and lactate (1 mM), and unlabelled glucose (5.5 mM) and oleate (0.4 or 1 mM) are from Vincent et al. [38]. Magnifications of the scale (insets) present the concentrations of isocitrate. *P<0.05, 1 or 0.7 mM oleate versus 0.4 mM oleate. CS, citrate synthase; gww, g wet weight; αKG, α-ketoglutarate (2-oxoglutarate).

Figure 2
Effects of increasing exogenous oleate concentrations on tissue concentrations of individual CAC intermediates (A, C) and the anaplerotic PC/citrate synthase flux ratio (B, D) in working mice (A, B) and rat (C, D) heart perfusion experiments

Results are means±S.E.M. Data from mice hearts perfused with U-13C3-labelled pyruvate (0.2 mM), and unlabelled lactate (1.5 mM), glucose (11 mM) and oleate (0.4 or 0.7 mM) are from Khairallah et al. [14]. Data from rat hearts perfused with U-13C3-labelled pyruvate (0.2 mM) and lactate (1 mM), and unlabelled glucose (5.5 mM) and oleate (0.4 or 1 mM) are from Vincent et al. [38]. Magnifications of the scale (insets) present the concentrations of isocitrate. *P<0.05, 1 or 0.7 mM oleate versus 0.4 mM oleate. CS, citrate synthase; gww, g wet weight; αKG, α-ketoglutarate (2-oxoglutarate).

Fatty acid oxidation

Recently, heart perfusion experiments using differently 13C-labelled fatty acid and isotopomer analysis by GC–MS have revealed the partitioning of fatty acid between mitochondrial and peroxisomal β-oxidation [41]. Specifically, results demonstrated partial peroxisomal β-oxidation of long-chain fatty acids forming C12 and C14 acyl-CoAs and contributing >50% of the fatty acid-derived acetyl groups that end up in malonyl-CoA, a key regulator of mitochondrial fatty acid β-oxidation. Surprisingly, these data suggest that, by supplying acetyl-CoA to malonyl-CoA synthesis, peroxisomal fatty acid β-oxidation might participate in the control of mitochondrial fatty acid β-oxidation. It is noteworthy that peroxisomal fatty acid β-oxidation consumes O2 and produces H2O2; it is not linked to the formation of reducing equivalents and ATP. Hence, the extent of peroxisomal β-oxidation of fatty acids should be considered when ATP production rates are extrapolated from the stoichiometric relationships between oxygen consumption and citrate formation from fats, as well as theoretical yields of ATP per mole of fatty acid oxidized.

Practical considerations, future challenges and conclusion

The aforementioned examples illustrate complex metabolic networks involved in cardiac energy substrate metabolism, which can be probed by isotopomer analysis, but are not predicted by classical biochemistry or measurements of metabolite concentrations. Other complex metabolic networks prevailing in the heart include glycogen [22] and triacylglycerol cycling [42], NADH transfer from the cytosol to mitochondria by the α-ketoglutarate/malate shuttle [43], glutamate/glutamine cycling [44], the ICDH (isocitrate dehydrogenase) substrate cycle [45] and the fumarate reduction pathway [46]. These pathways may consume energy, for example glutamine synthesis or the citrate cleavage pathway, or affect redox balance such as the ICDH cycle, or proton production such as triacylglycerol cycling, and are thus critical to the overall energy balance and metabolic integrity of the intact heart. Inasmuch as all these processes are affected by substrate supply and disease, as well as potentially affecting cardiac function and pathogenesis, we believe that they need to be part of any comprehensive model of cardiac metabolism. Their exclusion will lead to an incomplete description of the system, resulting in potentially erroneous conclusions.

Therefore there are a number of important questions that need to be considered when developing a model of cardiac metabolism, such as: which 13C-labelled substrate(s) and measured metabolite 13C-isotopomer data are most informative? Which provide the most reliable and precise flux ratio data? Which experimental metabolite isotopomer data should be selected for modelling? Clearly, it is unlikely that the use of a single 13C-labelled substrate and the measurements of 13C-isotopomer data in a single metabolite will be sufficient to probe the complexity of the metabolic networks involved in cardiac metabolism. Nevertheless, if one chooses to restrict analyses to simple data sets, then the inherent limitations of such analyses need to be understood and clearly articulated. For example, if one uses [U-13C]glucose as the only 13C-labelled substrate and 13C-labelled isotopomer data from a single metabolite, either citrate by GC–MS or glutamate by NMR, then this will provide precise relative flux data solely on the contribution of glucose to acetyl-CoA formation (energy) and anaplerosis, and as we have shown above, this will not be relevant to conditions existent in the heart in vivo. To dissect out the contribution of other substrates to acetyl-CoA, such as lactate, pyruvate and fatty acids, these substrates also need to be 13C-labelled. In this regard, one advantage of NMR over current GC–MS methodology is the possibility to use up to four differently labelled substrates in a single heart perfusion experiment [16].

As discussed previously in detail [12], with the appropriate choice of labelled substrates both NMR- or GC–MS-determined glutamate or citrate isotopomer data provide precise flux ratios for substrate oxidation in the CAC, as well as for anaplerosis using physiologically relevant substrate mixtures. Nevertheless, additional metabolite isotopomer data are needed to dissect out the regulation of other metabolic networks. For example, as described above, 13C-isotopomer analysis of lactate, pyruvate and alanine from both heart tissue and perfusate, which can be readily determined using NMR and/or GC–MS methods, provides valuable information on the regulation of pyruvate partitioning [1416,21]. Similarly, the comparison of GC–MS-determined 13Clabelling patterns of malonyl-CoA and of mitochondrial acetyl-CoA was used to reveal partitioning of fatty acid between mitochondrial and peroxisomal β-oxidation [41]. Clearly, the significance of many of the aforementioned metabolic processes may vary under various physiological and pathophysiological conditions and, hence, need to be evaluated in separate experiments using different 13C-labelled substrates over a range of physiological conditions. For example, [U-13C]glutamate has been used to probe reverse flux through the ICDH reaction [45], [U-13C]fumarate for the fumarate reductase pathway [46] and [1-13C]glutamine for glutamate/glutamine cycling [44]. The determined metabolite flux ratios may be used subsequently as constraint parameters for metabolic models. It is noteworthy that one should be careful in extrapolating data from other organs or tissues to the heart, where the significance of these processes may be different. For example, mitochondrial citrate efflux, pyruvate recycling and flux through the reversal of the ICDH reaction are considerably less active in the heart than in liver [19,45,47].

The combination of 13C-isotopomer analysis by GC–MS and NMR will ultimately provide maximal metabolic flux information from a given experiment and consequently play an important role in developing a comprehensive understanding of cellular function. Given the complexity of physiological experiments and the mathematical tools needed, we, like others [3], believe that the successful building of a comprehensive model of cardiac metabolism will most likely result from an interactive consortium between metabolic physiologists and bioengineers. This concerted effort of investigators also needs to address the intra- and extra-cellular compartmentation of cardiac energy metabolism, as well as the potential for channelling due to protein–protein interactions, which remains a major challenge for all researchers in this field [12,4852].

Large-Scale Screening: A Focus Topic at BioScience2005, held at SECC Glasgow, U.K., 17–21 July 2005. Edited by B. Baum (Ludwig Institute, London, U.K.), K. Brindle (Cambridge, U.K.), S. Eaton (Institute of Child Health, London, U.K.) and I. Johnstone (Glasgow, U.K.).

Abbreviations

     
  • CAC

    citric acid cycle

  •  
  • ICDH

    isocitrate dehydrogenase

  •  
  • LDH

    lactate dehydrogenase

  •  
  • PC

    pyruvate carboxylation

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