"Medicine, Faculty of"@en . "Pathology and Laboratory Medicine, Department of"@en . "DSpace"@en . "UBCV"@en . "Leong, Hon Sing"@en . "2009-10-10T00:00:00"@en . "2002"@en . "Master of Science - MSc"@en . "University of British Columbia"@en . "Glycolysis, measured by \u00C2\u00B3H\u00E2\u0082\u00820 production from [5-\u00C2\u00B3H]-glucose, is accelerated in\r\nisolated working hypertrophied rat hearts. However, non-glycolytic detritiation of [5- \u00C2\u00B3H]-\r\nglucose by transaldolase in the non-oxidative pentose phosphate pathway (PPP) could\r\nlead to an overestimation of true glycolytic rates. Since the PPP may be upregulated in\r\ncardiac hypertrophy, I tested the hypothesis that detritiation of [5-\u00C2\u00B3H]-glucose does not\r\noverestimate glycolysis in hypertrophied hearts and that accelerated rates of glycolysis\r\nreported in hypertrophied hearts are real and not artifactual. Glycolysis was measured by\r\nthree independent methods in isolated working hearts from halothane-anesthetized shamoperated\r\n(Control) and aortic-constricted (Hypertrophy) rats. Two of the three methods\r\ndetermined glycolytic rates by quantifying the accumulation of glycolytic end products in\r\ntimed collections of perfusate while the last method determined glycolytic rates by\r\ndetritiation of [5-\u00C2\u00B3H]]-glucose in the glycolytic pathway by production of 3H\u00E2\u0082\u00820. The first\r\nmethod involved enzymatic determination of lactate and pyruvate combined with rates of\r\nglucose oxidation. The second method involved determination of radiolabeled [\u00C2\u00B9\u00E2\u0081\u00B4C]-\r\nlactate and [\u00C2\u00B9\u00E2\u0081\u00B4C]-pyruvate accumulation combined with rates of glucose oxidation. The\r\nthird method, which measures glycolytic flux by metabolism of [5-\u00C2\u00B3H]-glucose, was in\r\nquestion and thus was used for comparison with the aforementioned alternative methods\r\nof determining glycolytic flux. Glycolysis was accelerated in hypertrophied hearts,\r\nregardless of the method used. There was also excellent concordance between the three\r\nmethods with no significant differences in glycolysis detected between corresponding\r\ngroups. Moreover, glucose-6-phosphate dehydrogenase activity and transaldolase\r\nexpression, enzymes controlling key steps in the oxidative and non-oxidative PPP,\r\n\r\nrespectively, were not different between control and hypertrophied hearts. Thus, nonglycotytic\r\nloss of 3H\u00E2\u0082\u00820 from [5-\u00C2\u00B3H]-glucose is insignificant and 3H\u00E2\u0082\u00820 production from [5-\u00C2\u00B3H]-\r\nglucose is an accurate means to measure glycolysis in isolated working normal and\r\nhypertrophied rat hearts. Furthermore, the PPP does not appear to be increased in this model of\r\ncardiac hypertrophy."@en . "https://circle.library.ubc.ca/rest/handle/2429/13883?expand=metadata"@en . "9043796 bytes"@en . "application/pdf"@en . "Accelerated Rates of Glycolysis in Cardiac Hypertrophy: Are they a Methodological Artifact? By Hon Sing Leong B.Sc , University of Alberta, 1999 A THESIS SUMBITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE In THE F A C U L T Y OF G R A D U A T E STUDIES (Department of Pathology & Laboratory Medicine) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH C O L U M B I A October 2002 \u00C2\u00A9 Hon Sing Leong, 2002 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department The University of British Columbia Vancouver, Canada Date DE-6 (2/88) ABSTRACT Glycolysis, measured by 3H.20 production from [5-3H]-glucose, is accelerated in isolated working hypertrophied rat hearts. However, non-glycolytic detritiation of [5- H]-glucose by transaldolase in the non-oxidative pentose phosphate pathway (PPP) could lead to an overestimation of true glycolytic rates. Since the PPP may be upregulated in cardiac hypertrophy, I tested the hypothesis that detritiation of [5-3H]-glucose does not overestimate glycolysis in hypertrophied hearts and that accelerated rates of glycolysis reported in hypertrophied hearts are real and not artifactual. Glycolysis was measured by three independent methods in isolated working hearts from halothane-anesthetized sham-operated (Control) and aortic-constricted (Hypertrophy) rats. Two of the three methods determined glycolytic rates by quantifying the accumulation of glycolytic end products in timed collections of perfusate while the last method determined glycolytic rates by detritiation of [5-3H]-glucose in the glycolytic pathway by production of 3H20. The first method involved enzymatic determination of lactate and pyruvate combined with rates of glucose oxidation. The second method involved determination of radiolabeled [ 1 4C]-lactate and [14C]-pyruvate accumulation combined with rates of glucose oxidation. The third method, which measures glycolytic flux by metabolism of [5-3H]-glucose, was in question and thus was used for comparison with the aforementioned alternative methods of determining glycolytic flux. Glycolysis was accelerated in hypertrophied hearts, regardless of the method used. There was also excellent concordance between the three methods with no significant differences in glycolysis detected between corresponding groups. Moreover, glucose-6-phosphate dehydrogenase activity and transaldolase expression, enzymes controlling key steps in the oxidative and non-oxidative PPP, respectively, were not different between control and hypertrophied hearts. Thus, non-glycotytic loss of 3 H20 from [5-3H]-glucose is insignificant and 3 H20 production from [5-3H]-glucose is an accurate means to measure glycolysis in isolated working normal and hypertrophied rat hearts. Furthermore, the PPP does not appear to be increased in this model of cardiac hypertrophy. TABLE OF CONTENTS i . Abstract ii ii. Table of Contents iv iii. List of Figures and Tables viii iv. List of Abbreviations x v. Acknowledgements iix I. Introduction A. Cardiac Hypertrophy 1. Overview 1 2. Clinical Significance 1 3. Myocardial (Mal)Adaptations in Cardiac Hypertrophy 2 B. Metabolism of the Mammalian Heart 1. Overview 5 2. Myocardial Fatty Acid Metabolism 5 3. Myocardial Carbohydrate Metabolism 9 C. Pentose Phosphate Pathway 1. Overview 14 2. The Oxidative Portion of the Pentose Phosphate Pathway 16 3. The Non-Oxidative Portion of the Pentose Phosphate Pathway 18 D. Metabolic Alterations in Cardiac Hypertrophy 1. Fatty Acid Oxidation 22 iv 2. Glycolysis and Glucose Oxidation 22 3. Lactate Metabolism and Lactate Oxidation 25 4. Consequences of Altered Energy Metabolism in Hypertrophied Heart 26 5. Role of the Pentose Phosphate Pathway in the Hypertrophied Heart 27 E. Measurement of Glycolysis in Isolated Working Heart 29 Preparations F. Potential Overestimation of Glycolytic Flux in Isolated Working Rat Hearts 31 II. Hypothesis and Objectives 32 III. Experimental Methods A. Animal Model 33 B. Isolated Heart Preparation and Perfusion Protocol 33 C. Measurement of Glycolysis and Glucose Oxidation 1. Glycolysis 34 2. Glucose Oxidation 36 D. Immunoblot Analysis of Transaldolase 36 E. Measurement of Glycogen 37 F. Measurement of Glucose-6-phosphate Dehydrogenase Activity 37 G. Data Analysis 37 v IV. Results A. Heart and body weight data 1. 0.4 mM palmitate perfused (low fatty acid content group) 38 2. 1.2 mM palmitate perfused (high fatty acid content group) 38 B. Mechanical heart function 38 C. Rates of Glycolysis 1. Glycolysis from total lactate plus pyruvate and glucose oxidation a) High fatty acid conditions 42 b) Low fatty acid conditions 43 2. Glycolysis from [14C]-lactate production & [ 1 4C] -pyruvate production and glucose oxidation 48 3. Glycolysis from Detritiation of [5-3H]-Glucose 48 4. Comparison of Three Methodologies of Determining Glycolytic Flux in Isolated Working Rat Heart Preparations 51 D. Myocardial Glycogen Content 51 E. Enzymes of the Pentose Phosphate Pathway 52 V. Discussion A. Summary of Findings 56 B. Glycolysis in Hypertrophied and Control Hearts 57 C. Pentose Phosphate Pathway Flux in Hypertrophied and Control Hearts 59 D. Detritiation of f5-3HI-glucose as a Measure of Glycolytic Flux 60 E. Methodological Considerations 64 F. Importance of this Study 64 Appendices I. Modified r14C\"]-Lactate + r14Cl-Pyruvate Radiolabel Assay 66 II. Calculation of Glycolytic Flux from Accumulation of f'4C1-Lactate and r14C1-Pvruvate 74 III. Calculation of Glycolytic Flux from Accumulation of Total Lactate and Pyruvate 76 IV. Summary of Metabolic Data for Isolated Working Hearts Exposed to 1.2 mM Palmitate (High Fatty Acid Conditions') 77 V. Summary of Metabolic Data for Isolated Working Hearts Exposed to 0.4 mM Palmitate (Low Fatty Acid Conditions') 78 BIBILIOGRAPHY 79-94 vii LIST OF FIGURES AND TABLES Figure 1. Myocardial Adaptations in Cardiac Hypertrophy 3 2. Myocardial Energy Production in Hypertrophied and Normal Hearts 6 3. Energy Production in the Myocardium by Fatty Acid Oxidation 8 4. Summary of Myocardial Metabolism 11 5. Energy Production in the Myocardium by Glycolysis and Glucose Oxidation 13 6. Oxidative and Non-Oxidative Branches of the Pentose Phosphate Pathway (PPP) 15 7. Oxidative Branch of the Pentose Phosphate Pathway 17 8. Non-Oxidative Branch of the Pentose Phosphate Pathway 19 9. Potential Cycling Effect by the PPP in the Glycolytic Pathway 20 10. Glycolysis and Glucose Oxidation in Hypertrophied and Normal Hearts 23 11. The Impact of the PPP on Glycolysis and Detritiation of [5-3H]-glucose 28 12. Glycolytic Pathway and the Detritiation of [5-3H]-glucose 30 13. Mechanical Function of Control and Hypertrophied hearts perfused with either high (1.2 mM palmitate) or low fatty acid (0.4 mM palmitate) 41 14. Accumulation of Lactate and Pyruvate in Hypertrophied and Control Hearts Perfused under High Fatty Acid Conditions (1.2 mM Palmitate) 44 15. Comparison of Three Methods of Determining Glycolytic Flux from Hypertrophied and Control Hearts in High Fatty Acid Conditions (1.2 mM palmitate) 45 16. Accumulation of Lactate and Pyruvate in Hypertrophied and Control Hearts Perfused under Low Fatty Acid Conditions (0.4 mM Palmitate) 46 17. Comparison of Three Methods of Determining Glycolytic Flux from Hypertrophied and Control Hearts under Low Fatty Acid Conditions (0.4 mM palmitate) 47 18. Tritiated Water Production from Glycolysis in Hypertrophied and Control Hearts Perfused under High Fatty Acid Conditions (1.2 mM Palmitate) 49 19. Tritiated Water Production from Glycolysis in Hypertrophied and Control Hearts Perfused under High Fatty Acid Conditions (0.4 mM Palmitate) 50 20. Expression of Transaldolase 53 21. G6PDH Activity in Hypertrophied and Control hearts 54 22. Expression of Transaldolase in Various Rat Tissues 55 23. Sum of Glucose Oxidation Rates and Accumulation of Lactate and Pyruvate are Equivalent to Glycolytic Flux Measured by Detritiation of [5-3H]-glucose 67 24. Schematic of the Modified [14C]-lactate and [14C]-pyruvate Assay 68 25. Calculation of Glycolytic Flux from [14C]-lactate and [14C]-pyruvate Assay 74 Table 1. Heart and Body Weight of Hypertrophied and Control Hearts 39 2. Mechanical Function of Control and Hypertrophied Hearts in High and Low Fatty Acid Conditions (1.2 mM and 0.4 mM palmitate) 40 3. Total Lactate and Pyruvate Accumulation in Isolated Working Hypertrophied and Control Hearts 71 4. Inter/Intra Test Variability of the [14C]-Lactate Radiolabel Assay 72 5. Radiolabel Assay Efficacy in Differing Perfusate Conditions 73 LIST OF ABBREVIATIONS 1,3-BPG: 1,3-Biphosphoglycerate 2-PG: 2-Phosphoglycerate 3-PG: 3-Phosphoglycerate ATP: Adenosine triphosphate CPT1: Carnitine palmitoyltransferase 1 CoA: Coenzyme A dH 2 0: Distilled water DHAP: Dihydroxyacetone phosphate ETC: Electron Transport Chain E4P: Eiythrose 4-phosphate F A D H 2 : Reduced flavin adenine nucleotide FFA: Free Fatty Acids F6P: Fructose 6-phosphate G6P: Glucose 6-phosphate G6PDH: Glucose 6-phosphate dehydrogenase GAP: Glyeraldehyde-3-phosphate GAPDH: Glyceraldehyde-3-phosphate dehydrogenase H K : Hexokinase LPL: Lipoprotein Lipase L D H : Lactate Dehydrogenase MCT: Monocarboxylic Acid Transporter X N A D + : Oxidized nicotinamide adenine nucleotide N A D H 2 : Reduced nicotinamide adenine nucleotide PDC: Pyruvate Dehydrogenase Complex PEP: Phosphenolpyruvate PGI: Phosphoglucose isomerase PGK: Phosphoglycerate kinase P G M : Phosphoglycerate mutase PK: Pyruvate kinase PPP: Pentose Phosphate Pathway PFK-1: Phosphofructokinase-1 R5P: Ribose 5-phosphate S7P: Septulose 7-phosphate TIM: Triose phosphate isomerase TCA cycle: Tricarboxylic Cycle V L D L : Very-low density Lipoprotein Xu5P: Xylulose 5-phosphate xi ACKNOWLEDGEMENTS Thank you Dr. Allard, for all your help and effort during this degree. I am very relieved and fortunate to have met someone like you. My faith in research and in people has been restored and most of all, faith has been restored in myself. Richard Wambolt, I truly thank you for your supervision over me, and the knowledge I received under your tutelage. Mark Grist, I thank you for your discipleship and willingness to teach me as much as you can about the field of metabolism. I would like to also thank Hannah Parsons for her critical work in elucidating the expression and activity of transaldolase and G6PDH; I couldn't have done it without you. As for the rest of the Allard lab, I wish the best for all and I pray that God be as merciful to you as He was to me. I want to dedicate this work to my family, a source of inspiration and support during this degree. I greatly appreciate the devotion and faith that was placed in me during those stressful times and I pray that you guys continue to be there for me as I carry on in my studies. Finally, I want to give a big thanks to all my friends who comforted me and provided me with support during this degree. Finally, I want to thank everyone in Daniel and Phileo Fellowship and my good friends at the iCAPTUR 4E Centre. This work was sponsored and funded by grants provided by the Canadian Institutes of Health Research (CIHR) and the Heart & Stroke Foundation of British Columbia and Yukon. I. INTRODUCTION A) Cardiac Hypertrophy 1. Overview Cardiac hypertrophy is a condition where the heart adapts to prolonged increases in pressure/volume overload by an enlargement of the heart (83). This enlargement or hypertrophy of the myocardium is caused by increases in wall thickness, myocyte volume and size so that wall stress is normalized (57,83). This adaptive response also extends beyond that of a structural impact; gross mechanical function and fuel metabolism are also affected, depending on the nature of cardiac hypertrophy. There are two main types of cardiac hypertrophy: exercise-induced and pathological cardiac hypertrophy (119). This study will focus on the pathologically hypertrophied heart and the fundamental changes in its metabolism compared to the normal myocardium. 2. Clinical Significance Pathological cardiac hypertrophy is common in Western populations, present in 15-20% of populations (76), with ~90% of hospitalized individuals in advanced stages of cardiac disease possessing a hypertrophied heart (41,75,76,74,49). Although cardiac hypertrophy resulting from arterial hypertension is an initial compensatory response, cardiac failure eventually occurs due to a number of mechanisms that are provoked by the heart's adaptation to work overload (24). Therefore, cardiac hypertrophy has emerged as a major risk factor for more severe forms of cardiovascular disease (75,76,74,49) leading to sudden death, myocardial infarction, and congestive heart failure (49). In some cases, sudden death can occur without a definite ischemic event (41). Individuals with hypertrophied hearts also tend to demonstrate poorer recoveries from acute ischemic insults because the heart is more susceptible to injury in the event of an ischemic insult (14,50,6) compared to individuals with normal hearts. Therefore, the co-existence of cardiac hypertrophy and coronary artery disease, both of which are highly prevalent conditions, in any individual has dramatic implications. 3. Myocardial (MaDAdaptations in Cardiac Hypertrophy When the heart initially copes with the increase in workload demand, diastolic dysfunction first occurs (59,144) while dramatic changes in protein expression take place as the hemodynamic stimuli evokes growth signals, cell receptors, and transcription factors to induce structural cardiac hypertrophy (69,92). Despite the normalization of mechanical function according to workload, structural remodeling inadvertently affects other minor properties of the heart that ultimately contribute to its impending dysfunction (99). Alterations in cardiac receptors, endocrine function, contractile proteins, ion exchange and energy metabolism are observed in hypertrophied hearts (16,44,51,14) (Figure 1). For example, altered rates of shortening velocity and delayed relaxation of cardiac muscle of the heart can be attributed to alterations and deficiencies in ion exchange and handling as the cause for these changes in contraction and relaxation (58). Another significant feature is increased Ca2+accumulation in the myocytes of hypertrophied hearts; a result of a lack of C a 2 + handling (6) which is thought to result in a reversal of Na + /Ca 2 + exchange to regulate intracellular C a 2 + (105,125,93). Following this export of Ca 2 + , greater Na + accumulation is observed in the cell, which ultimately leads to ischemic dysfunction when consequent NaVrT1\" ion exchange results in myocardial acidosis (14,91,52). Although the effects of myocardial acidosis are not immediately detrimental in the aerobic perfusion setting, its effects become fully realized post-ischemia, and contribute to the increased susceptibility of the hypertrophied heart to injury in the event of ischemia; (14,91,52). The hypertrophied heart's response to ischemia is even more dramatic when its metabolism is considered. Increases in glycolytic enzyme levels (53), and altered properties of lactate dehydrogenase (21) have been reported in hypertrophied hearts, leading to increased glycolytic metabolism. This change becomes detrimental as the by-products of glycolytic metabolism (NADFfc, lactate, IT\") accumulate and contribute to the pathogenesis of myocardial ischemic damage in hypertrophied hearts (14,5). The hypertrophied heart is distinctively susceptible to myocardial ischemia, and is observed to have decreased mechanical function upon post-ischemia reperfusion when compared to normal hearts (127,11). An alteration in energy substrate use by the hypertrophied heart has been implicated as an important cause for the pathophysiology of cardiac hypertrophy (21,28) and decreased post-ischemic function in the hypertrophied heart (11,33). Although glycolysis is beneficial in generating energy during and after ischemia, oxidative metabolism of glucose is low relative to glycolysis. As a consequence, a more acidic intracellular environment is produced that complicates Ca 2 + handling which leads to myocardial dysfunction (94,6). Therefore, the hypertrophied heart is maladaptive in the ischemic setting because of alterations in glucose metabolism that may contribute to changes in myocardial ion exchange during and after an ischemic event. B) Energy Metabolism of the Mammalian Heart 1. Overview The omnivoric nature of the heart reflects its perpetual need to provide energy for the constant activity of the heart and its function in providing circulation of blood to the rest of the body. This required energy is derived from fatty acids, glucose, lactate, amino acids and ketone bodies (97,116,62). Fatty acids are the predominant source of energy for the normal heart (Figure 2), while glycolysis and the oxidation of glucose and lactate meet the remainder of the heart's energy requirements under normal circumstances. However, the extent to which all exogenous sources of energy contribute to ATP pools is entirely dependent on the availability and presence of competing energy substrates, with fatty acid oxidation and glucose catabolysis operating in an inverse relationship as outlined by the Randle cycle (111). 2. Myocardial Fatty Acid Metabolism Oxidation of fatty acids accounts for a large majority (-70-85%) of energy production (116). In the blood, these free fatty acids are bound to either albumin or lipoprotein and enter cells by passive diffusion (1) and/or saturable protein-facilitated transport (2). Thus, high extracellular FFA levels and/or low intracellular FFA levels favour the uptake of fatty acids. The proteins FAT (fatty acid translocase/CD36) and FABP (fatty acid binding protein) (135) are the main components involved in fatty acid protein- facilitated transport. Imported FFA's are then primed for mitochondrial (3- oxidation by fatty acyl-CoA synthetase (98,123). This enzyme activates each individual fatty acyl for oxidative degradation ((3-oxidation) by catalyzing the addition of a CoA group to the fatty acyl 9 > d 3 | B i \u00C2\u00A9 a \u00C2\u00A9 s CTQ \u00C2\u00AB* 09 > \u00C2\u00A7 3 S* \u00C2\u00AB - i \u00E2\u0080\u00A2 sr O hrj CfQ 3 q I s a ? E OS - a 4-o \u00E2\u0080\u00A2U a BS \u00C2\u00BB\u00E2\u0080\u00A2*\u00E2\u0080\u00A2 BS \u00C2\u00AB ^ 3 g 3 CfQ a *< BS 3 i s. m \u00C2\u00A9 BS 3 CfQ BS fi * \u00E2\u0080\u0094 SB J J O 9 o O ST \u00C2\u00A7> \u00E2\u0080\u00A2\u00C2\u00A9 3 \u00C2\u00BB 3\" 2> s 2 a- \u00C2\u00A9 8 3: o a a cr \u00C2\u00AB< TJ ft \"1 3 \u00C2\u00A9* S3 \u00C2\u00A9 BS \u00E2\u0080\u0094 sr re H 2, TP WS 3 re re \" i 88 P re sr c ft \u00C2\u00AB\u00C2\u00ABS BS V p p <*> O *\u00E2\u0080\u00A2 o\ > o \u00E2\u0080\u00A2\u00E2\u0080\u00A22. >2. \u00C2\u00A3 \"2. o o ...;>.... o o o o > > > i 8 2 n 2 S !> i P P \" i \ O O O ! \u00C2\u00BB > > >J \ U M > H - J \"CO 6 > > X X K) UJ > > H H > > H H \"CD 00 Acetyl--Oxidai -CoA ion X NJ ATP UJ ll ll UJ VO un as ATP > \u00C2\u00A3 H H hd m a. o 3 H o <-+ EL > H 3 o <-+ o 3 Although the products of fatty acid oxidation and glucose oxidation rally at the start of the TCA cycle in the form of acetyl-Co A, competition between fatty acids and glucose as the dominant source of fuel in the heart exists in a reciprocal fashion as described by the Randle Cycle (95). As outlined, glucose oxidation decreases in response to increases in fatty acid oxidation because the buildup of acetyl-Co A and NADtfe allosterically inhibit the pyruvate dehydrogenase complex (PDC) - the main regulator of glucose oxidation (129). Inversely, increases in carbohydrate metabolism result in a decrease in fatty acid oxidation. This is due to the increased production of malonyl-CoA from acetyl-CoA carboxylase (ACC) and subsequent inhibition of carnitine palmitoyltransferase I (CPT1) and fatty acid transport into the mitochondria. Balance between these metabolic pathways is also dictated by mitochondrial ratios of NADH/NAD + and acetyl-Co A/Co A, which have regulatory allosteric effects on the activity of PDC and ACC, respectively. Overall, metabolism of fatty acids and glucose is well regulated and primarily controlled by substrate availability and metabolite interplay, as well as by workload and hormonal control. 3. Myocardial Carbohydrate Metabolism Carbohydrates, such as glucose and glycogen, are a significant source of energy for the heart and are first metabolized by the glycolysis pathway, which splits glucose into two molecules of pyruvate. In doing so, glycolytic degradation of one glucose molecule produces 2 ATP molecules that are involved in ion homeostasis maintenance and muscle contractility (97,129,39). The pyruvate generated from this pathway enters the mitochondria where it is used to generate large amounts of ATP in a process known as glucose oxidation (97,129,39). Prior to glycolysis, glucose is transported into myocytes by way of glucose transport protein carriers (97,26,129,39), and once within the myocyte cytoplasm, glucose is phosphorylated by hexokinase to glucose-6-phosphate. Glucose-6-phosphate then has two major fates: either to be stored as glycogen, or to be degraded by glycolysis (Embden-Meyerhof pathway). Under normal circumstances, the majority of glucose 6-phosphate units undergo glycolysis, where one glucose molecule is ultimately split into two pyruvate molecules (a 3 carbon glycolytic intermediate) by the glycolytic pathway. In doing so, a net 2 ATP and 4 N A D H 2 are produced for every glucose molecule that passes through glycolysis. At the completion of the glycolytic pathway, pyruvate generated has three main fates: it can be transported out of the myocyte; be converted to lactate by lactate dehydrogenase; or undergo oxidation in the mitochondria, in a process known as glucose oxidation (Figure 4). The oxidation of pyruvate (essentially glucose) in the mitochondria is a very industrious process, and occurs via three main components, the Pyruvate Dehydrogenase Complex (PDC) (35), the Tricarboxylic Acid Cycle (TCA Cycle) (71,70), and the Electron Transport Chain (ETC) (90,60,17). First, the PDC decarboxylates pyruvate into acetyl-CoA, thus priming it for complete oxidation by the TCA (71,70). Once within the TCA cycle, reducing power is transferred from acetyl-CoA to cofactors such as F A D H 2 and N A D H 2 , and these reducing equivalents undergo oxidative phosphorylation at the site of the ETC to generate ATP (90,60,17). A l l in all, 38 ATP is generated alongside 3 C 0 2 and 3H2O for every glucose molecule that undergoes glycolysis and glucose I I P <\u00C2\u00A3*\u00E2\u0080\u00A2< o ^ ss rt. P P \u00C2\u00B0 -a sr o o ^ B 5-P O 02 CTQ a o \u00C2\u00A9 3 2 \u00C2\u00AE o rs a. re S3 ^ rs \u00C2\u00BB4 o \u00E2\u0080\u00A2\u00E2\u0080\u0094 S CTQ a c g \u00C2\u00AB re \u00E2\u0080\u00A2 \u00E2\u0080\u00A2 I f Is, s P \u00C2\u00A9 cr SJ \u00C2\u00A9, rs o S3 re O* s 0 i \u00C2\u00A9 a 5s a n H o p rs Hs 0 J cs ^ 3 S3 9 rt-O \"I re\" \u00C2\u00A9\" ft ^ rf P S cr \u00C2\u00A9 \u00C2\u00BB 2. i \u00C2\u00A7 3 I re P ~ 5? p o . \u00E2\u0080\u0094 i cT rs o \u00C2\u00A9 \u00C2\u00BB ^ 5-3 re <\u00C2\u00BB p a re ^ rs sr ^ fD rt-N J \u00C2\u00A9 fi t s p p~ l-( 05 sr p re s? s rt <\u00E2\u0080\u00A2> 05 fD sv sr w r* rs OS \u00C2\u00A9 re ss a \u00E2\u0080\u0094 cs ^ rt. re 00 2 - 8 8 \u00C2\u00A9 03 p \u00C2\u00B0 E . \u00C2\u00A3 \u00C2\u00A7 ^ \u00C2\u00A9 oxidation (Figure 5). However, the fraction of pyruvate that is actually oxidized is regulated by the Pyruvate Dehydrogenase Complex (PDC). This site is the major regulatory step of glucose oxidation, a regulation that is executed by a kinase-phosphatase system (pyruvate dehydrogenase kinase/pyruvate dehydrogenase phosphatase) and intramitochondrial N A D H 2 / N A D + and acetyl-CoA/CoA levels (35). Phosphorylation of PDC, catalyzed by PDK (107,108,23), leads to inactivation of the complex, while dephosphorylation of PDC is catalyzed by PDP and leads to activation of the PDC. PDP is also sensitive to the intramitochondrial environment, being dependent on Mg^and further activated in response to increases in concentrations of C a 2 + in the mitochondrial matrix (34,112,103,36,19,129). For example, during exercise an increase of C a 2 + can cause an activation of PDH phosphatase, thereby activating the PDC. In the isolated working rat heart, the fraction of pyruvate (derived from glycolysis) oxidized ranges from 20-40%, depending on the fatty acid condition (7,121,9,47,138,139,142). Under low fatty acid conditions (0.4 mM palmitate), the percentage of pyruvate oxidized increases upwards of ~35-40%, whereas isolated working hearts perfused with high fatty acid oxidize a much lower fraction of pyruvate (-10-20% of total pyruvate at 1.2 mM palmitate) (7,121,9,47,138,139,142). Under conditions of no FFA, pyruvate/glucose oxidation rates are near maximal because intracellular ratios of N A D H 2 / N A D + and acetyl- Co A/CoA are at their lowest, thus inducing an activation of PDC. There are other secondary pathways of glucose metabolism: a small portion of pyruvate is devoted to the maintenance of the Tricarboxylic Acid Cycle. In the event of excessive carbon loss in the cycle (cataplerosis) (136), pyruvate is converted to eT CTQ S ft (TQ o CL c \u00C2\u00A9' 0 3\" 3 O o C6 o re 3 f t *S P rt-f t a o* O <^ o P a 3 Q \u00C2\u00A3T o W5 f t o a P \u00C2\u00A9' 3 O O o o OO > H TJ ft ^ O oo\" X I\u00E2\u0080\u0094' \u00E2\u0080\u00A2 CL o 3 K) rt-> > H H TJ TJ > H to H > TJ o ts) ON O o K) rtl U \u00C2\u00B0 X ^ ^ > % TJ ^ oo CD o x TJ v5 CD 3 O H O CD to > H TJ 4> > H TJ 00 > H TJ > H TJ x J\u00C2\u00BB ts) > H H TJ TJ ts) ts) > H TJ > H TJ oxaloacetate and shunted to the TCA cycle to replenish intermediate pools in a process known as anaplerosis (102). Pyruvate carboxylase is one major anaplerotic enzyme that catalyzes this carboxylation of pyruvate to oxaloacetate (130,131,64,32). Another anaplerotic fate of pyruvate, one of minor metabolic significance, is the conversion of pyruvate to alanine (65). Also known as the \"pyruvate-alanine cycle\", this process is also anaplerotic by contributing to oxaloacetate pools in a reaction catalyzed by glutamine-pyruvate tranaminase (GPT). This reaction is as follows: pyruvate + glutamine <=> alanine + oxaloacetate As well, a small portion of glycolytic intermediates are also known to participate in nucleotide metabolism through a secondary pathway called the Pentose Phosphate Pathway (PPP). In this pathway, G6P is shunted away from glycolysis and used to form the carbon backbone of nucleic acids, as well as generating reducing power (in the form of NADPH2) for lipid biosynthesis processes. C) The Pentose Phosphate Pathway (PPP) 1. Overview Synonymously recognized as the Pentose Phosphate Shunt, or the Hexose Monophosphate Shunt, the PPP forms a link between carbohydrate metabolism (glycolysis and glucose oxidation), fatty acid metabolism and nucleotide synthesis in the cytosol (Figure 6). Normally, glucose is catabolized by glycolysis and glucose oxidation; however, the presence of the PPP yields a secondary pathway of glucose oxidation that seeks to replenish NADPH2 and ribose-5-phosphate pools (145,148,147), both of which serve functions unrelated to glucose and fatty acid metabolism. In terms of overall SI activity and expression, the PPP is prominent in the liver because NADPH2 is required for lipid biosynthesis, while PPP activity is extended across many other tissues including cardiac and skeletal muscle (149). Although these tissues and cell types have different metabolic requirements, the N A D P H 2 produced by this pathway is useful in protection against oxidative stress, an event not all that rare in muscle (145,148,147). To achieve these effects, the PPP consists of two main branches; the oxidative branch and the non-oxidative branch (149). Both branches work collectively in the cytosol, where the oxidative branch of the PPP is linked to glycolysis and nucleotide metabolism, and where the non-oxidative portion of the PPP serves as the reversible link between the oxidative branch of the PPP and glycolysis. 2. Oxidative Portion of the PPP Glucose-6-phosphate originating from either glucose or glycogen is predominantly catabolized by glycolysis. However, a portion of glucose-6-phosphate may be shunted away from glycolysis to enter the oxidative portion of the PPP so that ribose-5-phosphate and NADPH2 can be generated (145,148,149,147) (Figure 7). The ribose-5-phosphate produced serves as the carbon backbone for nucleic acid synthesis, while N A D P H 2 is primarily used to protect against oxidative stress by maintaining glutathione in a reduced state (145,148,147). The pivotal reaction that leads to these important functions is catalyzed by Glucose-6-Phosphate Dehydrogenase (G6PDH) and is irreversible (154). G 6 P D H activity is governed by N A D P H 2 inhibition, an inhibition that is in competition with NADP+(45). For instance, under normal physiological conditions of N A D P + and free NADPH2, this enzyme is almost completely inhibited. sr g \u00E2\u0080\u00A2O rs sr g' TO \u00C2\u00BBQ H * \u00C2\u00A9 era \u00E2\u0080\u00A2\u00E2\u0080\u00A2 s\u00C2\u00BB sr K TO re P \u00C2\u00A9 a o \u00C2\u00A3J> 3s\" M X \u00C2\u00ABv fe E-a \u00C2\u00B0 2 Vi \u00C2\u00A9 re rs \"O re sr \u00C2\u00B0 Si* 3 -a \u00C2\u00AB< sr \u00C2\u00A9 \u00C2\u00A3 ~ re 2 / \u00E2\u0080\u0094 s \u00C2\u00A9 \" Tj re ^ \u00C2\u00A9 s as TO SJ' P s & TO \u00C2\u00B0 e 3 3 P 3 CL ^ \u00C2\u00A9 3 V O CS 3 St rt- C L ST 8 8 TO rt-TO TO TO ss N 3 re J \u00C2\u00A9 rt \u00C2\u00A9' 3 \u00C2\u00A9 \u00C2\u00A9 TO M 2. & S\" p a \u00C2\u00A3 ^ P n \u00C2\u00A9 TJ 88 TJ TO *0 dn sr TO S5* TJ cr \u00C2\u00A7 1 \u00C2\u00AE 3 (/! TO TO 1? \u00C2\u00A9 rt Cfl ST T3 TO * g '\u00E2\u0080\u00A2d TO Jd P t * B P TJ '< p-j TJ \u00C2\u00A9 V a ^ g s 2 s CL TO 2 1 \u00C2\u00A33 TO TO \u00C2\u00A3 \u00C2\u00A9 TJ 5. rt \u00C2\u00A9 o- -a e sr 1 TJ \u00E2\u0080\u00A2 < \u00E2\u0080\u0094t C < CD rf CD CD \u00E2\u0080\u00A2 sr 9 X \u00C2\u00AB<. c_ o co X ? ai -o TJ 5 -N-' o CO T3 cr CD CT O CO CD \"36 ui on \"o T l cr w o CO TJ CT 0) CD 3 o i f 3' S3 Co (D CD co 3 o 5-3 CO CD tj 2 3; Co O CD a o TJ 33 C CO \u00E2\u0080\u0094 ^5 Q3 CD CD 9 1 O c o o 03 CD Therefore, G 6 P D H activity is only possible by de-inhibition; a simple matter of competition between N A D P + and N A D P H 2 for binding on G 6 P D H . The activity of the oxidative PPP is typically of a low capacity in the rat and human heart (126,45,15,153). In fact, the N A D P + pool is normally sufficiently reduced that G 6 P D H is substantially inhibited and flux through the oxidative portion of the PPP is extremely low in the heart (126,45,15,153). 3. The Non-Oxidative Portion of the PPP The non-oxidative branch of the PPP is involved in the interconversion of 3- , 4- , 5- , 6-, and 7-carbon sugars catalyzed by a series of non-oxidative reactions that occur in cytoplasm (145,148,149,147) (Figure 8). Given that the synthesis of NADPH 2 by the oxidative PPP is essential for reductive biosynthetic processes, the non-oxidative portion of the PPP fulfills this demand by cycling and converting excess ribose-5-phosphate generated by the oxidative PPP back into glycolysis. In doing so, the back conversion of the ribose-5-phosphate to triose and hexose glycolytic intermediates allows continual cycling of both portions of the PPP with the aid of the glycolytic pathway, for continual N A D P H 2 production (Figure 9). Therefore, the non-oxidative branch of the PPP serves to prevent the depletion of the intermediate pools of the oxidative portion of the PPP (145,148,149,147). As a result, a cycling effect is produced (Figure 9) where NADPH2 may be produced without significant loss of carbon. Overall: 2 xylulose 5-phosphate (5C) + 1 ribose 5-phosphate (5C) \u00E2\u0080\u00A2 2 fructose 6-phosphate (6C) + 1 glyeraldehyde 3-phosphate (3C) 2 K S3 ft ~ S 3 \u00C2\u00B0 CM ^ a *a 2 \u00C2\u00BB c 2\" s* ^ \u00E2\u0080\u00A2 \u00E2\u0080\u00A2 o rt crQ 3. 3 3 oo P \u00E2\u0080\u00A2\u00E2\u0080\u00A2 ^ sr 5* \u00C2\u00AB rt \u00E2\u0080\u00A2 s r ^ i\u00E2\u0080\u0094\u00C2\u00BB as \u00E2\u0080\u00A2 -\u00C2\u00B0 St -a a oro ^ 2 ~ p s\u00C2\u00BB i t , 5? a.? \u00C2\u00A7 o I 3* a 2. \u00C2\u00A3 t a r t * SS rt e 2? P ss sr rt ON 3 \u00C2\u00AB 5\" T J g. 5' o ST rt rt g 3 \u00C2\u00A9 C 8. ? I rt CC rt-. I\u00E2\u0080\u0094I On' 3 -< o 3 ^ 5 ^ E a t ? \u00E2\u0080\u00A2 ' * ss 4*. T3 13 P f 1 w as! \u00C2\u00A9 B p s 5 2 .. a rt X \u00E2\u0080\u00A2<_ o 1/1 fD X 1 I O i/> T3 3 \" n> Q -< n ro E i Q . fD n Q-\"> fD 1 o T3 3 \" Glucose Pyruvate Figure 9: The Cycling of Glycolytic Intermediates by the Pentose Phosphate Pathway as Proposed by Goodwin et al (56). 20 Transketolase and transaldolase are the key enzymes of this pathway (Figure 7), where transketolase catalyzes the conversion of xylulose-5-phosphate (5C) and ribulose-5-phosphate (5C) into glyceraldehyde-3-phosphate (3C) and sedoheptulose-7-phosphate (7C) (145,148,149,147). Transaldolase then catalyzes a conversion between these two end products to synthesize a 4-carbon sugar (E4P) and a 6-carbon sugar (F6P). The glycolytic intermediates fructose-6-phosphate and glyceraldehyde-3-phosphate are also formed at the end of the PPP when transketolase catalyzes a conversion between xylulose-5-phosphate and erythrose 4-phosphate. This branch of the PPP is controlled by the availability of substrates and because all interconversion reactions are reversible, there is great metabolic flexibility and versatility (45). For example, when ribose 5-phosphate is in higher demand than N A D P H 2 , glucose 6-phosphate will be catabolized by glycolysis to fructose 6-phosphate and glyceraldehyde 3-phosphate. Transaldolase and transketolase will then convert these substrates in a reversible fashion to synthesize ribose 5-phosphate which will be available for purine and pyrimidine nucleotide synthesis (146). Conversely, if the demand for N A D P H 2 is greater than the need for ribose-5-phosphate, then glucose-6-phosphate is completely catabolized by glycolysis and its intermediates will be shunted to G 6 P D H and the oxidative PPP. Both the oxidative and non-oxidative branches of the PPP will generate N A D P H 2 while excess ribose-5-phosphate will be recycled back to its glycolytic intermediate forms by the non-oxidative PPP (146,148,147). Thus, the non-oxidative PPP provides much needed versatility by adjustments according to specific metabolic situations and demands. D) Metabolic alterations in Cardiac Hypertrophy 1. Fatty Acid Oxidation Oxidation of fatty acids has been observed to be lower in hypertrophied hearts compared to normal hearts (46,7,120,47), an observation dependent on the degree of hypertrophy, fatty acid levels and workload (8,118). Reduced expression of fatty acid uptake/transport proteins (104,3,135) and reduced carnitine levels (46,7,120,47) is thought to be responsible for low fatty acid oxidation in hypertrophied hearts; carnitine being a structural component of the CPT system for long chain fatty acid translocation, as well as binding to individual fatty acyls-CoA throughout this transport (7,8,29). At the site of the CPT, impaired cooperation between the acyl-CoA dehydrogenase and CPT has also been implicated as being responsible for low fatty acid oxidation rates (29). Finally, it has been shown that the mRNA expression of enzymes necessary for the oxidation of fatty acids is depressed in the hypertrophied heart (115,18). However, the extent to which the down-regulation expression of such enzymes plays a role in reducing fatty acid oxidation has not yet been determined (115,18). 2. Glycolysis and Glucose Oxidation In the hypertrophied heart, rates of glycolysis have been shown to be accelerated when compared to control hearts in the isolated working mode (5,121,47). In hypertrophied hearts, expression and activities of a number of glycolytic enzymes are increased (53,132), and may even shift towards more fetal, anaerobic isoforms (68) as part of the adaptive response to workload. However, oxidation of pyruvate is not correspondingly increased (Figure 10) (7,121,9,47,138,137,143). In fact, the fraction of nmol glucose / min / g dry wt > H B P 2 re 2. S3 S 5? 3 O ^ \u00C2\u00A3 wi SS \u00C2\u00A9' 6 8 O CL re p * *J cr Q\ \u00E2\u0080\u00A2 \u00E2\u0080\u00A2 cn O \u00C2\u00A9 sr ^ S * p QTQ P rt- i-s a \u00C2\u00AB> re O 'Q M si \u00C2\u00A9 o ^, Wi . . V\u00C2\u00A9 p rt p re TJ re a a* t s re \u00C2\u00BBi 3 55\" Ml \u00C2\u00A9* s< \u00C2\u00A9 re i . W *t g * re f* re B r f 3 re 3 wa P ~ B \" ^ \"i i\u00E2\u0080\u0094i re ^ QTQ a ts y C re 2 v <*> 2-C H rtj wi 55\" n g > ST 52 SI B s* CL \u00E2\u0080\u009E ws L*-\u00C2\u00A9 ^| CfQ Wi ^ re 2 c 3 rs r - v re o * rt 5 \u00C2\u00AB * 3 re re \u00C2\u00A9 5 \u00C2\u00AB P S * W3 C. 2 B a 2 2 \u00C2\u00BB Si re rt rt- Q , \u00C2\u00AB \u00E2\u0080\u00A2 rt. \u00C2\u00A9 o e B ~ 2. / - s \u00E2\u0080\u00A2o 3 W A <\u00C2\u00BB ^ o ^ ^ \u00C2\u00A9 V R \"T 3' <& rs B \u00C2\u00A9 2 B wa *S re \u00C2\u00A9, ^ p 5\" B 2 f re 2 Wi rt, _ rt-rtj Q tjq TJ B 3*. o re 9 CL wi re nmol glucose / min / g dry wt CD *< TJ CD O TJ pyruvate oxidized (from glycolysis) in hypertrophied hearts is lower than that found in control hearts (5,120,121,9,47,138,10,140,141) because rates of pyruvate/glucose oxidation are comparable or even lower than those in control hearts (46,9,141). These findings are particularly unusual because decreases in fatty acid oxidation in control hearts are typically accompanied by increases in glucose oxidation that result in a higher fraction of pyruvate being completely oxidized (80). This disparity in glucose oxidation between control and hypertrophied hearts has significant functional relevance since low fractional oxidation of glucose has been linked to contractile dysfunction, especially after an ischemic episode. Normally, decreases in fatty acid oxidation in the normal heart result in low intracellular levels of N A D H 2 and acetyl-Co A, conditions that induce an activation of PDC and glucose oxidation (112,29). Moreover, accelerated rates of glycolysis in hypertrophied heart produce large quantities of pyruvate that normally maintain a high PDC activation state that also lead to high glucose oxidation rates (112,103,113,109,110). Despite such circumstances, glucose oxidation is still limited in the hypertrophied heart. Hence, the proportion of glycolytically-derived pyruvate that is oxidized is lower in hypertrophied hearts than in normal hearts. It is thought that these alterations in energy metabolism are important factors contributing to the functional changes in the heart and its increased susceptibility to injury during ischemia (14,50,6) (see section Consequences of Altered Energy Metabolism in Hypertrophied Heart for details). The rate of glucose oxidation is mostly dependent on the activation state of the PDC (129), and it is possible that an alteration in its expression or activation is responsible for the low glucose oxidation rates in hypertrophied hearts. Since intracellular pyruvate and acetyl-Co A levels in the hypertrophied heart are theoretically capable of inducing high glucose oxidation rates, any alteration in PDC regulatory machinery could be responsible for depressed glucose oxidation rates. However, a study by Lydell et al. (84) determined that subunit expression, activity, and covalent regulation of PDC machinery were not different between control and hypertrophied hearts. Therefore, the limitation of glucose oxidation in hypertrophied hearts is not due to a reduction or alteration in PDC expression, activity or covalent regulation (84) and that other mechanisms are responsible for this limitation (84). 3. Lactate Metabolism and Lactate Oxidation Upon the degradation/catabolysis of glucose through glycolysis, the major fate of pyruvate is the conversion to lactate by lactate dehydrogenase in the isolated working heart setting (99). This is especially true for pyruvate produced by the hypertrophied myocardium where glycolytic rates are accelerated and rates of glucose oxidation remain relatively stagnant and depressed (21,132). Under such circumstances, lactate production is significantly higher in the hypertrophied heart as proportionally less pyruvate undergoes glucose oxidation compared to normal hearts. In doing so, this increased production of lactate regenerates and stabilizes the cytosolic NADH2/NAD+ redox state, the stability of which the continuation of glycolysis is dependent on (21). The formation of N A D + for every lactate molecule synthesized serves to restore this glycolytic capability and is a major source of N A D + in the cytosol. Enhanced lactate production and release in hypertrophied hearts is believed to be caused by the emergence of a fetal isoform of lactate dehydrogenase in hypertrophied heart that possesses a higher affinity for pyruvate (132). The emergence of this fetal isoform is associated with the overall alteration in gene expression observed in pressure-overload hypertrophied hearts (21,42). Although this fetal isoform of L D H is characterized as such, the extent to which pyruvate is intentionally converted to lactate is unknown. However, both hypertrophied and normal hearts oxidize lactate (97,7,120), and lactate oxidation in hypertrophied hearts is equivalent or less than that found in normal hearts (7,120). Therefore, although lactate production is enhanced in hypertrophied heart, its oxidation and contribution to ATP pools is not accordingly increased. 4. Consequences of Altered Energy Metabolism in Hypertrophied Heart Accelerated rates of glycolysis and high physiological levels of lactate are of major concern in the hypertrophied heart due to high \u00C2\u00A5t load in the cytosol (14,5). Collectively, high lactate production may compound the problem of proton overload in hypertrophied heart. Surprisingly, it is the dramatic reduction in the oxidative metabolism of glucose that causes a net buildup of H4\", and not the dissociation of lactate which produces ^(63). When all glycolytically-derived pyruvate undergoes complete oxidation, net IT production is zero. However, when there is low fractional pyruvate oxidation, a net 2 it is produced for every glucose molecule passing through glycolysis and not through glucose oxidation (63). Consequent accumulation of protons results when proton release from ATP hydrolysis exceeds proton export capacity of the myocyte. During ischemia, many Jrt transporters are also believed to be inactive and therefore a large intracellular concentration of fT is created, often resulting in ion imbalances, membrane damage and contractile dysfunction. The accumulation of H1\" during ischemia primes the myocardium for C a 2 + overload during reperfusion by means of NaVrT\" and Na + /Ca 2 + exchange (72). Upon reperfusion, a massive export of proton erupts, causing damage to the cell membranes as well as creating an influx of C a 2 + into the cell (133,80). Low glucose oxidation rates seen in the hypertrophied heart often compound this predicament by not fully oxidizing pyruvate produced by glycolysis (80), a process which would otherwise cancel out proton release by ATP hydrolysis. Instead, excessive amounts of glycolytic product result in an acidic intracellular environment which increases calcium influx and decreases contractility (48). This enhancement of calcium transport often results in poor contractile activity as the myocyte attempts to deal with the imbalance of ions in the myocyte. However, the amount of energy expended to reach ion homeostasis decreases cardiac efficiency and increases dysfunction. 5. The Role of the PPP in the Hypertrophied Heart There are claims that non-glycolytic detritiation of [5-3H]-glucose may occur at the transaldolase reaction of the Non-Oxidative portion of the PPP (Figure 11). It was observed that glycolytic flux as measured by three independent methods in control hearts was inconsistent and that glycolytic flux as measured by the [5-3H]-glucose method was neither constant nor linear (44). Therefore, a very important corollary was the possibility that rates of glycolysis are overestimated by the [5-3H]-glucose method in the isolated working rat heart. This overestimation may be even more pronounced in the pathologically hypertrophied heart setting because it is possible that the PPP may be elevated or altered due to the heart's increasing requirements for nucleic acids for structural enhancement as the heart enlarges, or to protect the myocardium from reactive oxygen species as a consequence of accelerated cellular processes. Therefore, if there is significant PPP activity in the rat heart, the well-documented metabolic profile of the pathologically hypertrophied heart will require further review. Other studies have also established the presence of a long-term control mechanism regarding G6PDH in which increased activity and expression of G6PDH occurs in skeletal muscle and in rat heart during the development of cardiac hypertrophy and/or by the influence of catecholamines (155,156). Long-term responses to cardiac hypertrophy (by constriction of the abdominal aorta) have also demonstrated varying increases in G6PDH expression and activity in the heart (155,156). Overall, both branches of the PPP may be significant in the hypertrophied isolated working rat heart. E) Measurement of Glycolysis in the Isolated Working Heart Preparation In the isolated working heart preparation, tritium-labeled glucose ([5-3H]-glucose or [2-3H]-glucose) is added to the perfusion buffer solution to measure glycolysis (82,37). This method has proven to be a popular method of determining glycolytic rates in the isolated working rat heart because of its commercial availability and specificity. In particular, [5-3H]-glucose is preferentially used because non-glycolytic detritiation of [2-3H]-glucose has been observed during glucose transport (31). Therefore, rates of glycolytic flux are determined by the release of 3H20 from [5-3H]-glucose during its passage through glycolysis. It is assumed that [5-3H]-glucose is completely detritriated by the triose phosphate isomerase and enolase reactions of glycolysis (82,37) (Figure 12). The standard measurement of glycolysis by timed collection of perfusate samples and [5-3H] Glucose I2\"3\"] D H A P P - 0 - C H , - C - C H , O H II O 5H P - 0 - C H , - C H - C OH OP 5H I / P - 0 - C H , - C H - C ^ O I OH o-I / CH2\u00E2\u0080\u0094CH\u00E2\u0080\u0094C v l l x GAPDH [2-3H]13_BpG PGK [2-3H] 3_pG PGM 3 H 2 0 o [2-3H]2.pG OH OP C H r C H - C ' OP o-V o I Enolase P E P CHo\u00E2\u0080\u0094CH\u00E2\u0080\u0094C v I \ o o-PK Pyruvate Figure 12: The degradation of glucose via the glycolytic pathway. The tritium label at the fifth carbon of glucose will be released as water at either: 1) the triose isomerase reaction (TIM) or 2) at the enolase reaction. Adapted with permission from: Figure 16-3; Biochemistry 2nd Edition, Voet and Voet, Copyright \u00C2\u00A9 (1993). Reprinted by permission of John Wiley & Sons. separation of 3 H 2 0 from [5-3H]-glucose, is a well-established means of measuring glycolytic rates in isolated heart preparations (95,82,37). F) Potential Overestimation of Glycolytic Flux in Isolated Working Rat Hearts Recently, Goodwin et al have suggested that rates of glycolysis obtained by quantitation of 3 H 2 0 production from [5-3H]-glucose overestimate true rates of glycolysis in isolated working normal rat hearts (56). They propose that non-glycolytic detritiation of [5-3H]-glucose occurs in the non-oxidative portion of the pentose phosphate pathway (PPP) by means of the transaldolase reaction in the heart. In other studies, key enzymes of the oxidative pentose phosphate pathway, such as glucose-6-phosphate dehydrogenase, were found to be elevated in hypertrophied hearts (155,61). If enzymes of the non-oxidative portion of the pentose phosphate pathway are also increased above normal and if substantial non-glycolytic detritiation of [5-3H]-glucose occurs in myocardium, as suggested, it is conceivable that the acceleration of glycolysis observed in isolated working hypertrophied rat hearts is an artifact. n. HYPOTHESIS AND OBJECTIVES If there is substantial non-oxidative PPP activity in the normal and hypertrophied heart, in which non-glycolytic detritiation of [5-3H]-glucose occurs, then glycolytic flux measured by this tracer method should be exaggerated when compared to other independent measures of glycolytic flux. If this is the case, then the postulated elevation of the PPP in hypertrophied heart may be responsible for accelerated rates of glycolysis observed when using this technique. However, based upon the long-standing reputation of the method, I hypothesize that 3H20 production from [5-3H]-glucose does not overestimate true rates of glycolysis in hypertrophied rat hearts due to the PPP. To test this hypothesis, rates of glycolysis in isolated working hypertrophied and non-hypertrophied hearts were determined by three independent methods. To determine if both oxidative and non-oxidative portions of the pentose phosphate pathway are altered in the hypertrophied heart, I also measured activity or expression of two key enzymes, glucose-6-phosphate dehydrogenase and transaldolase, of the oxidative and non-oxidative portions of the pentose phosphate pathway, respectively. III. EXPERIMENTAL METHODS A. Animal Model Pressure-overload left ventricular hypertrophy was produced in 3-week old male Sprague-Dawley rats (50-75 g) by constriction of the suprarenal abdominal aorta with a metallic clip (0.4mm diameter) (7). In sham-operated control rats, the aorta was isolated but not constricted. Experiments were performed 8 weeks after surgery. Rats, housed in a temperature-controlled (22\u00C2\u00B11\u00C2\u00B0C) and light-controlled (12:12-h light-dark cycle) room, had free, unlimited access to feed and water. Care of the animals was performed in accordance with guidelines set out by the Canadian Council on Animal Care. B. Isolated heart preparation and perfusion protocol Hearts from halothane (2-3%) anesthetized sham-operated and aortic-constricted rats were perfused for 30min with Krebs-Henseleit (KH) solution in the working heart mode at a preload of 11.5 mrnHg and an afterload of 80 mmHg, as previously described (7,9,10,141,79). Anesthesia with halothane is expected to have minimal effect on cardiac integrity and metabolism (157). The K H solution contained 1.2 mM or 0.4 mM palmitate prebound to 3% fatty acid-free albumin, 5.5 mM [5-3H/U14C]-glucose, 0.5 mM lactate, 2.5 mM calcium, and 100 mU/L insulin and was continuously circulated through the closed perfusion system Two different concentrations of palmitate (high fatty acid-1.2 mM and low fatty acid-0.4 mM), that represent the physiological range, were used in these perfusions . To ensure that glucose uptake was not limiting, a high physiologic concentration of insulin was utilized. The solution was oxygenated with 95% 02/5% CO2 and maintained at 37\u00C2\u00B0 C throughout the perfusion. A pressure transducer (Viggo-Spectramed, Oxnard, CA, USA) inserted in the afterload line was used to measure heart rate and peak systolic pressure. Cardiac output and aortic flow was measured via external flow probes (Transonic Systems Inc., Ithaca, NY, USA) on the preload and afterload lines, respectively. Coronary flow was calculated as the difference between cardiac output and aortic flow. Rate-pressure product, the product of heart rate and peak systolic pressure, and hydraulic work, the product of cardiac output and peak systolic pressure, were used to measure external work performed by the heart (7,9,10,141,79). These measures of heart function were assessed every 10 min throughout the working heart perfusion. At the end of 30 min, hearts were quickly frozen using aluminum tongs cooled to the temperature of liquid nitrogen. Frozen heart tissue was powdered using a mortar and pestle and then stored in cryovials at -70\u00C2\u00B0C until use. C. Measurement of Glycolysis and Glucose Oxidation 1. Glycolysis Rates of glycolysis were determined simultaneously in the same hearts by different and independent methods. In one method, glycolysis was calculated as the sum of the rate of lactate and pyruvate accumulation in the perfusate and the rate of glucose oxidation (56). For this calculation, accumulation of total lactate and pyruvate or [ 1 4C]-lactate and [14C]-pyruvate in the perfusate was determined (See Appendix I & II for calculation). Total lactate and pyruvate were measured enzymatically using a commercially available assay (Sigma, St. Louis, MO). Accumulation of radiolabelled lactate and pyruvate was determined by means of a modified version of the two-step assay described by Lehoux et al (73). Briefly, the [14C]-lactate in the sample was enzymatically converted to [14C]-pyruvate in the first step. Al l the [14C]-pyruvate in the sample, including that originating from [14C]-lactate, was then decarboxylated enzymatically to 1 4 C 0 2 in the second step. The 1 4 C 0 2 produced as a gas and from [ 1 4C]-bicarbonate in the buffer, released after addition of H2SO4, was collected into hyamine hydroxide-soaked filter paper, suspended in a centre well in the reaction vial, and subsequently counted by standard techniques. Recovery rates using this method, which were tested by spiking K H solution with known quantities of [14C]-lactate, range from 85 to 90%. A more detailed description of this assay is provided in Appendix I. Rates of accumulation of lactate and pyruvate (total or 14C-labelled), determined by taking volume of the perfusate, perfusion time, and dry heart weight as well as specific activity of perfusate [14C]-glucose, where appropriate, into account are expressed as glucosyl units/min/g dry heart weight. In the third method, glycolysis was determined by quantitatively measuring the rate of 3 H 2 0 liberated from [5-3H]-glucose (7,9,10,141,79). 3 H 2 0 was separated from perfusate using columns containing Dowex 1X4 anion exchange resins (200-400 mesh) dissolved in 0.4 M potassium tetraborate. The Dowex resin was extensively washed with dHzO before use. Duplicate samples (0.2 mL each) were added to the column and eluted into scintillation vials with 0.8 mL dEfeO. Following the addition of scintillation fluid (4 mL) to the tubes containing the eluent, the samples were subjected to double isotope counting procedures to measure 3 H 2 0 and residual [3H/14C]-glucose. Rates are expressed as glucosyl uruts/min/g dry weight. 2. Glucose Oxidation Glucose oxidation rates were measured by quantitative collection of 14CC>2 from [U-14C]-glucose released as a gas and dissolved in the perfusate as [14C]-bicarbonate, as previously described (7,9,10,141,79). Perfusate and gaseous samples were required to measure glycolysis and glucose oxidation and taken every 10 min of perfusion. Samples for determination of glycolysis and glucose oxidation were ultimately placed in vials containing scintillation cocktail and counted by standard techniques. D. Immunoblot Analysis of Transaldolase Expression of transaldolase protein in myocardium was determined by a previously described method (10). Briefly, samples of frozen ventricular tissue homogenate (containing 40 to 50|ig total protein) were solubilized by boiling in reducing sample buffer, separated by electrophoresis on 10% SDS-polyacrylamide gels, and transferred by electroblotting to a nitrocellulose membrane. After non-specific blocking, the blots were probed overnight with a primary rabbit anti rat transaldolase antibody (kindly donated by Dr. A. Perl, SUNY). After incubation with anti-rabbit secondary antibody, the signal was detected by an ECL based detection system. Bands were quantified by densitometry. Equivalence of protein loading was confirmed by detection of glyceraldehyde-3-phosphate dehydrogenase. E. Measurement of Glycogen Myocardial glycogen content was determined following extraction of frozen, powdered ventricular tissue with 30% KOH, ethanol precipitation, and acid hydrolysis of glycogen (9,79). F. Measurement of Glucose-6-phosphate dehydrogenase activity Glucose-6-phosphate dehydrogenase (G6PDH) activity in myocardium was measured by a standard spectrophotometric technique (40) which measures the rate of increase in absorbance at 339 nm. The assay reaction is as follows: G6PDH G l u c o s e - 6 - P + N A D P + - a \u00C2\u00BB 6 -phosphog lucono lac tone + N A D P H + H + G. Data Analysis Data are expressed as mean \u00C2\u00B1 standard error of the mean (SEM). Individual group means were compared using t-tests. The Bonferroni procedure was applied to all tests to correct for multiple tests and comparisons. A corrected value of p<0.05 was considered significant. 37 IV. RESULTS A. Heart and body weight data a. 0.4 mM palmitate perfused (low fatty acid content group) The heart and body weight data are summarized in Table 1. Heart weight of aortic-constricted rats (1.85\u00C2\u00B10.05 g, n=4) was not greater than that of the control rats (1.8710.02 g, n=4, p=NS). However, the heart weight/body weight ratio was higher in hypertrophied hearts compared to normal hearts, a finding consistent with our previous studies (5,9,10,141), demonstrating that this model produces a mild cardiac hypertrophy. Body weight was not significantly different between aortic-constricted (431+7 g) and control (459\u00C2\u00B13 g) rats (p=NS). b. 1.2 mM palmitate perfused (high fatty acid content group) The heart and body weight data are summarized in Table 1. Heart weight of aortic-constricted rats (1.9910.05 g, n=6) was ~ 11% greater than that of the control rats (1.7810.04 g, n=7, p<0.05), and consistent with our previous studies (5,9,10,141). Body weight was not significantly different between aortic-constricted (464112 g) and control (472110 g) rats. B. Mechanical Heart Function Mechanical function in isolated working rat hearts from control and aortic-banded rats is shown in Table 2. Mechanical function was stable throughout the perfusion in both groups under both fatty acid concentrations (Figure 13) while heart rate and rate-pressure product were similar in control and hypertrophied hearts. In high fatty acid \u00C2\u00AB< . \"O | 3 CD ii a-\"D rr co l+ oo cn l+ o b cn k> co i+ o O Z> o 11 2. ^ 3 cn co i+ co co ->j 1+ o b b 1+ o *< T3 ZJ CD II 3-2 3 T3 -rt M 1+ CD CD CO 1+ O b \u00C2\u00BB -rt i+ o _ O r> o 11 H \u00E2\u0080\u0094 o -rt -rt 1+ 00 1+ O b O l 00 bo 1+ o CD o CL CD C Q ' 3 1 CD CD ~ CD 00 o ^ 3 CD \u00C2\u00A3. <\u00E2\u0080\u00A2 0 5 \u00C2\u00AB\u00C2\u00A7\u00E2\u0080\u00A2 c? \u00C2\u00BB-f- \u00E2\u0080\u0094 Of # < Cfl n o B 3 c Cfl CD cr A O O m p CJ H-00 W x 3. 3 ' CD o o o 3 CO \u00E2\u0080\u00A23 \u00E2\u0080\u0094h o CO CD CO 1+ o 4^ CO 1+ o bo CO 1+ 45> 1+ cn co Q . \u00E2\u0080\u0094I CO C o M O CO 1+ IO 1+ O co * 4^ CO 1+ N> 4^ CO CO 1+ CO co O 03 CO O =\u00E2\u0080\u00A2 O \"D -vl \u00E2\u0080\u00A2vl 1+ U l CO cn CO o 1+ ro * CT) Ul i+ CT) CT) U l b 1+ 00 3 S. CQ X 7] 0) f\u00E2\u0080\u0094I-CD T3 \u00E2\u0080\u0094^ CD Cfl cn c \u00E2\u0080\u00941 CD \u00E2\u0080\u00A2a \u00E2\u0080\u0094t o Q . C o CO 1+ o CD co CO 1+ o CT) tO CT) 1+ CT) ro co l+ co 3 & T l CD CO T T Cfl *< Cfl CQ O T3 B Cfl Cfl c CD CO 1+ CT> CO 1+ Ul o CT) 1+ ro CT) 1+ co * cr CD 0) I \u00E2\u0080\u0094 f -Cfl X i 3 I CD CD 3-CD CD ro CT) U l 1+ co ro co ui l+ CT> to U l ro i+ CO ro ro CO 1+ ro co O o 3 I \u00C2\u00BB< TD CD \u00E2\u0080\u0094* O T3 CT) O O 3 3 II 4i> T3 CD 3 T3 3 II 4^ ro \"D co CD O 3 TJ CO CO r\u00E2\u0080\u0094f CD 5 P 3 a o P H P e s o rt-\u00C2\u00A9 ' S \u00C2\u00A9 \u00C2\u00A9 rt-rt, \u00C2\u00A9 , p a I rt a o E Iro p rt-cr rt P rt a IP Rate Developed Pressure Product (bpmxmmHg/100) CTQ CfO ST C re 2 . g O. re s s-| B S\" s* C to \u00C2\u00A9 \u00E2\u0080\u0094 A \u00C2\u00A7 SB O ^ a e o P s 3 5-& O P O TJ S3 TO SJ-SB 3 w \u00C2\u00A9 a xs sr TO I 3 \u00E2\u0080\u00A2< SA rt, sr p p P W TO M a g TO A \u00C2\u00A9 a s i \u00C2\u00A9 rt-SA O ^ T> 8- ~ 2- a TO ^ 5 s TO SA sr \u00E2\u0080\u00A2 P TJ TO SA TO P rt-SA 3 CD K) ^ O 3. 5' CO o \u00C2\u00A9 ro o co o \u00E2\u0080\u00A2Irt o o conditions, other parameters of heart function were lower in hypertrophied hearts than in control hearts, with statistically significant differences in cardiac output, hydraulic work and coronary flow. In low fatty acid content perfusions, only peak systolic pressure was significantly higher in hypertrophied hearts compared to control hearts. A l l other parameters of heart function were not statistically significant between hypertrophied and control hearts. C. Rates of Glycolysis 1. Glycolysis from Total Lactate plus Pyruvate and Glucose Oxidation a. High Fatty Acid Conditions Over the duration of the perfusion under high fatty acid conditions, accumulation of lactate and pyruvate was linear in both hypertrophied and control hearts (Figure 14). Lactate and pyruvate were released into the perfusate at significantly higher rates by hypertrophied hearts than by control hearts which lead to a greater overall accumulation of lactate and pyruvate in hypertrophied hearts (Table 3). Under high fatty acid (1.2 mM palmitate) conditions, rates of accumulation of lactate (Hypertrophy, 2948 \u00C2\u00B1375 vs. Control, 1095 \u00C2\u00B1 288 nmol glucose equivalents/min/g dry wt, n=7, p<0.05) and pyruvate (Hypertrophy, 647 \u00C2\u00B1 99 vs. Control, 423 \u00C2\u00B1 54 nmol glucose equivalents/min/g dry wt, p<0.05) were significantly greater in hypertrophied hearts than in control hearts. In addition, glucose oxidation rates (measured directly by 1 4 C02 production from [U- 1 4 C]-glucose) were not significantly different between hypertrophied hearts (619 \u00C2\u00B1 48 nmol/min/g dry wt) and control hearts (475 \u00C2\u00B1 70 rimol/min/g dry wt, p=NS). Overall, glycolysis calculated from these rates of lactate and pyruvate accumulation and glucose oxidation were higher in hypertrophied hearts compared to control hearts (Figure 15). b. Low Fatty Acid Conditions Perfusions performed under low fatty acid conditions, resulted in accumulation of lactate and pyruvate that was also linear in both hypertrophied and control hearts (Figure 16). Although lactate was released into the perfusate at similar rates in hypertrophied and control hearts, there was a greater overall accumulation of lactate in hypertrophied hearts than in control hearts (Table 3). Pyruvate accumulation was higher in hypertrophied hearts than in control hearts over the 30 min, but this was not statistically significant. Thus, rates of accumulation of lactate (Hypertrophy, 2453 \u00C2\u00B1 273 vs. Control, 1364 \u00C2\u00B1238 nmol glucose equivalents/min/g dry wt, p<0.05) were significantly greater in hypertrophied hearts than in control hearts under these conditions. Meanwhile, rates of accumulation of pyruvate (Hypertrophy, 423 \u00C2\u00B1 128 vs. Control, 317 \u00C2\u00B1 35 nmol glucose equivalents/min/g dry wt, p=NS) were not significantly different. Glucose oxidation rates (measured directly by 1 4 C02 production from [U-14C]-glucose) did not differ significantly between hypertrophied hearts (1472 \u00C2\u00B1198 nmol/min/g dry wt) and control hearts (1336 \u00C2\u00B1 270 nmol/min/g dry wt, p=NS). Under low fatty acid conditions, rates of glycolysis in hypertrophied hearts were higher compared to control hearts (Figure 17), but this difference was not statistically significant. The low physiological level of fatty acids (0.4 mM palmitate) resulted in an enhancement of glucose oxidation in both hypertrophied and control hearts (as compared to 1.2 mM palmitate). In doing so, a greater degree of glucose passing through glycolysis was completely oxidized in hearts perfused with low fatty acid perfusate compared to glucosyl units (umol) 9v 3 rt CTQ 3\" \u00C2\u00A9 hcj 3 M -53 St 2. \u00E2\u0080\u00A2 s 2. 1 3 a \u00C2\u00A9 O ~ o 3 a o 3 H 3\" rt rt nmol/min/g dry wt io o o o o o o o o o fl) o rt> fl) CD + < rt C < fl) rt> CD * \u00C2\u00A9 rt i-t\u00C2\u00BB 2 a 5. n - 3 ^ 5 ' \u00C2\u00AE o rt \u00E2\u0080\u00A2 O rt 5\" n 9 3 rt-|-S 3 a M \u00E2\u0080\u00A2vj \u00E2\u0080\u00A2o rt *i rt-\u00E2\u0080\u00A2 t o ns sr rt\" a O fl) o rt> fl) rt> CD + o I \u00E2\u0080\u00A2o *< rt c < fl) CD \u00E2\u0080\u00A2 w X \u00E2\u0080\u00A2 o o 0) CD CO CD CO CO CO CD to LP nmol/min/g dry wt 9 3 S3 rt* O 3 O o P T j rt> CD O D) O rt> Q) r f CD + 9 ~ o I TJ *< c < Q) rt> CD c n \u00E2\u0080\u00A2 w \u00E2\u0080\u00A2 O c o o C/> CD \u00E2\u0080\u00A2r> cn hearts perfused with high fatty acid perfusate (36% vs. 16% of glucose oxidized respective to fatty acid concentration) 2. Glycolysis from f14C]-Lactate Production & f14C]-Pyruvate Production & Glucose Oxidation Similar results were obtained when glycolysis was calculated from [14C]-lactate and [14C]-pyruvate in the perfusate (Figures 15 & 17). Rates of glycolytic flux as measured by this method were higher in hypertrophied hearts compared to control hearts under high fatty acid conditions (4393 \u00C2\u00B1 369 nmol/min/g dry wt vs. 1972 \u00C2\u00B1 363 nmoVmin/g dry wt in high fatty acid perfusions and 4454 \u00C2\u00B1 487 nmoymin/g dry wt vs. 4181 \u00C2\u00B1 693 nmol/min/g dry wt in low fatty acid perfusions; hypertrophy and control hearts respectively). These values are not different from those measured by the aforementioned method of accumulation of lactate + pyruvate + glucose oxidation (Figure 23). Linearity of rates of glycolytic flux determined by this radiolabel assay was not assessed because the assay was only performed on perfusate collected at time point t=30 min. Data collection at earlier time points (10 min and 20 min) was not possible because of the low accumulation of [l4C]-lactate and [14C]-pyruvate and because sensitivity of the assay was not considered optimal at such low counts/activity. 3. Glycolysis from Detritiation of f5-3H]-Glucose Cumulative rates of 3 H 2 0 production from [5-3H]-glucose were linear in both control and hypertrophied hearts at both fatty acid levels (Figure 18 & 19). As with the other methods of determining glycolysis, rates of glycolysis determined by this glucosyl units (pmol) methodology were greater in hypertrophied hearts than in control hearts under high fatty acid conditions but not significantly different under low fatty acid conditions (1487\u00C2\u00B1132 nmol/min/g dry wt vs. 3962\u00C2\u00B1298 nmol/min/g dry wt in high fatty acid perfusions and 4108+298 nmol/min/g dry wt vs. 36521477 nmol/min/g dry wt in low fatty acid perfusions; hypertrophy and control hearts respectively; Figures 15 & 17). Glycolytic rates obtained in this way did not differ significantly from rates calculated on the basis of accumulation of total or [14C]-labeled lactate and pyruvate in either hypertrophied or non-hypertrophied hearts. Thus, there was excellent concordance between all three methods of determining glycolytic flux in isolated working rat hearts (Figure 15 & 17). 4. Comparison of Three Methodologies of Determining Glycolytic Flux in Isolated Working Heart Preparations As evidenced by Figures 15 and 17, glycolytic flux is not different between the three techniques over differing fatty acid concentrations. The excellent concordance observed in these figures demonstrates no overestimation of glycolytic flux by the [5-3H]-glucose method. Overall, this method is equivalent to two other independent methods. D. Myocardial Glycogen Content At the end of the 30 minute aerobic perfusion under high fatty acid conditions, control hearts had a glycogen level of 113.9\u00C2\u00B1 12.6 |imol/g dry wt, whereas hypertrophied hearts had a glycogen level of 107.5+12.6 |imol/g dry. Therefore, glycogen content at the end of the perfusion was not significantly different between hypertrophied and control hearts. Under low fatty acid conditions, control hearts had a glycogen level of 50.6.2\u00C2\u00B14.4 u,mol/g dry (n=4), whereas glycogen in hypertrophied hearts was significantly higher at a level of 71.4.7\u00C2\u00B15.4 (imol/g dry (n=4, p<0.05). E. Enzymes of the Pentose Phosphate Pathway Expression of transaldolase (based on protein abundance, Figure 20), and of glucose- 6-phospate dehydrogenase (based on enzyme activity, Figure 21) did not differ between hypertrophied and control hearts. Expression of transaldolase in heart was higher than muscle but lower than liver (Figure 22). >a 3 \u00C2\u00A7 \u00E2\u0080\u00A2 < \u00C2\u00A7 \u00C2\u00B0 5 5T \u00C2\u00AB rt a is \u00C2\u00AB i sr o a 2. 3 1 o 5 ST - a a >*i fD m m * ^ s rt 3 O Ni era \u00C2\u00A9 rt \u00E2\u0080\u00A2\u00E2\u0080\u00A2 5 S es rt % *o rt \u00C2\u00BBi _. rt rt w 3 rt O rt-3 g 3 ^ \u00C2\u00A3 rt O 3 l i es \u00C2\u00A9\" g sr rt- rt es \u00E2\u0080\u00A2 g s H rt (*i 3 es > 2 sr jr -a a rt- sr \u00C2\u00A9 3 & 3 sr rt era I Sr'Sf rt. rs \u00C2\u00A9 re w \u00C2\u00BB a rt/ o 3 es 5' (ra. 8T era rt- \u00C2\u00AB 3 6 8 sT 2 rt \u00C2\u00A9 * sr \u00C2\u00AB i rt es es rt-rt (D > \"D D > I O o Z3 I i v; \"O CD 3 \u00E2\u0080\u00A2 D g re ere n ^ rs p O V3 re H ; ^ i re 9 ra = re TJ sr o .. re re rT 3 -,sr a, P ^ 2 o 3 3'\u00C2\u00A7 CTQ sr HH re re 3 p 5^ p P-J TJ ri\" re re \"1 - i - o hd fe o TJ sr re\" a P P sr s >i re CL rt re o g a \u00C2\u00A7 3 \u00C2\u00A3 -B re H r w re II 3 ff TJ TJ ^ \u00C2\u00BB \u00C2\u00BB 8L S? rc SS -S TJ ^ 3. re \u00C2\u00A9 s Q, rt-- re \u00C2\u00A3 s \u00C2\u00A9 ^ rts M CfQ \u00C2\u00A9 = TJ mU/mg protein cn cn o to cn sr \u00C2\u00AB \u00C2\u00B0 T3 *H TO ST ffi TO W5 a T J TO -I sr o a S 3 2. CTQ c/> TO O S3 TO a \u00E2\u0080\u00A2 \u00C2\u00A3\u00E2\u0080\u00A2 3 1 3 T J g: sr ~ \u00C2\u00A9 ff \u00C2\u00A9 rt -t TO crq \u00E2\u0080\u0094 CO w s TJ * \u00C2\u00A9 H ff > <' tt TO \" TO rt 2 B S3 p 2- \u00C2\u00A9 * \u00C2\u00A3\" wa TO TO N> \u00E2\u0080\u00A2 \u00E2\u0080\u00A2 TO TJ i l TO wa TO S < TO SB era ^ TO TO rt. a TO o. \u00C2\u00BB i g, C v >*j te 2. cfq' 3 ^ e 1 a \u00C2\u00A9 *i T J \u00C2\u00A9 TO crq \u00C2\u00BB i TO p s rt- wa TO p C v wa ^ \u00C2\u00BB \u00C2\u00A7 S3* 3 \u00C2\u00BB3 2. as p wa 3 \u00C2\u00AE ff E TO rt rt-' \u00C2\u00A9 =3 * S3 C 3 it B a S \u00C2\u00AE TO \u00E2\u0080\u0094 2 \u00C2\u00A9 O rt-* g TO ^ rt-2 2 ^ 3 \u00C2\u00B0 S3 TO g. \" ss sr \u00C2\u00A9 wa <^ sr 3 X wa TO 3 \u00C2\u00BB TO *t rt gr sr crq a i r ? TO TO \u00C2\u00A9 a \" i f c v P O TO 2-ff \u00C2\u00A3 \u00C2\u00BBs v sr b J rt\" &3 rt \" & ff \u00C2\u00A7 TO a TO 0) c/> 0 fi) < 0 3 ffi TO wa C o (D TJ O o *< lrt\u00C2\u00BB IV. DISCUSSION The use of [5-3H]-glucose as a tracer to determine glycolytic rates in the heart is a well-established method that quantitatively measures glycolytic rates by collection of 3 H 2 0 released from [5-3H]-glucose to the perfusate (95,82,38). [5-3H]-glucose, during its catabolism in the glycolytic pathway, is assumed to be completely detritriated by the triose phosphate isomerase and enolase reactions (95,82,38). However, Goodwin et al. claim that 3 H 2 0 production from [5-3H]-glucose overestimates \"true\" rates of glycolysis in rat hearts due to non-glycolytic detritiation in the non-oxidative PPP (56). This aberration, if it exists, may be even more severe in the pathologically hypertrophied heart, raising the possibility that accelerated rates of glycolytic flux (measured using [5-3H]-glucose) may be artifactual. A . Summary o f Findings In this study, I confirm that glycolysis is accelerated in isolated working hypertrophied rat hearts as assessed with three different methods. I also demonstrate that non-glycolytic loss of 3 H 2 0 from [5-3H]-glucose is insignificant in isolated working hypertrophied and non-hypertrophied rat hearts. Most importantly, my data conclusively show that glycolytic rates deterrnined by measuring rates of 3 H 2 0 production from [5-3H]-glucose do not overestimate true rates of glycolysis and, thereby, confirm that this method is an accurate means to measure glycolysis in this setting. Lastly, I provide evidence that the pentose phosphate pathway is not increased in rat hearts hypertrophied by constriction of the abdominal aorta. B. Glycolysis in Hypertrophied and Control Hearts To investigate if glycolytic flux is overestimated by use of [5-3H]-glucose, three independent methods of determining glycolysis were used: detritiation of [5-3H]-glucose; the accumulation of lactate and pyruvate combined with rates of glucose oxidation; and the accumulation of [I4C]-lactate and [14C]-pyruvate combined with rates of glucose oxidation. When hearts were perfused under high fatty acid conditions (1.2 mM palmitate), excellent concordance was found between these three different and independent methods. Glycolytic rates were significantly higher in hypertrophied hearts than in control hearts, and this was consistent across all three methodologies. As well, glycolytic production was linear and glycolytic rates in both hypertrophied hearts and control hearts are typical of previous results in other experiments with similar perfusion conditions (at 1.2 mM palmitate) (7,8,121,122,139,10,141) demonstrating a linear trend of glycolytic flux. Hypertrophied and control hearts perfused under low fatty acid conditions (0.4 mM palmitate) also exhibited similar values of glycolytic flux amongst the three methodologies used. In hypertrophied hearts, rates of glycolytic flux were equivalent in all three methodologies. In control hearts exposed to 0.4 mM palmitate, the glycolytic flux measured by lactate and pyruvate accumulation and glucose oxidation was lower than values of glycolytic flux determined by the other two independent methods, but this difference was not statistically significant. Moreover, the glycolytic flux determined by the accumulation of [14C]-lactate and [14C]-pyruvate and glucose oxidation remained equivalent to the [5-3H]-glucose method, thus confirming the validity of all three methodologies as being equivalent techniques of measuring glycolytic flux. At high fatty acid levels (1.2 mM), glycolytic flux is accelerated in hypertrophied hearts compared to control hearts, presumably a result of low fatty acid oxidation rates and enhanced activity and expression of glycolytic enzymes. Under low fatty acid levels (0.4 mM palmitate), both hypertrophied and control hearts exhibited similar values of glycolytic flux. The abolishment of disparity in glycolytic flux between both groups can be mainly attributed to the availability and presence of glucose over fatty acid in the perfusate. This situation results in very high rates of glycolysis and glucose oxidation in both groups of hearts. Although the three independent methods of determining glycolytic flux are equivalent and accurate, there is some concern regarding the depletion of glycogen in hearts exposed to low fatty acid conditions and its possible contribution to lactate and pyruvate production. Control and hypertrophied hearts perfused with low fatty acids typically demonstrate glycogen depletion, with the added benefit that such endogenous forms of glucose are preferentially oxidized (9,139,79). However, excessive recruitment of glycogen as a source of unlabelled glucose into glycolysis could dilute the exogenous glucose pool and overestimate lactate and pyruvate accumulation levels upon glycolysis. If so, then glycolytic flux (as measured by accumulation of lactate & pyruvate + glucose oxidation) is overestimating. However, endogenous glucose is typically preferentially oxidized over exogenous sources of glucose (54,55,9). Therefore, this endogenous glucose likely does not contribute to lactate and pyruvate production and measured glycolytic rates because glycogen-derived glucose is catabolized by glycolysis and pre-destined for oxidation in the mitochondria. A l l in all, a more suitable explanation for the low accumulation of lactate, pyruvate, and glucose oxidation in this group may simply be due to error in handling or analysis of the perfusate samples. Our finding that rates of glycolysis are accelerated in hypertrophied hearts exposed to 1.2 mM palmitate, regardless of the method used, is in keeping with previous in vivo and in vitro observations based on a variety of independent parameters. Specifically, increased accumulation of 2-deoxyglucose-6-phosphate has been observed in hypertrophied dog (150) and rat (66) hearts in vivo . Furthermore, activity of a number of glycolytic enzymes is greater in hearts exposed to a pressure overload than in normal hearts (132) and isoenzymes of lactate dehydrogenase (21) and enolase (68) shift toward more anaerobic, fetal forms in hypertrophied hearts. The acceleration of glycolysis in hypertrophied hearts has been proposed as a compensatory response to low fatty acid oxidation rates in these hearts (5,7,47). As a result, rates of glycolysis may be near maximal in hypertrophied hearts despite any changes in fatty acid content. C. Pentose Phosphate Pathway Flux in Hypertrophied and Control Hearts In the event that loss of 3 H from [5-3H]-glucose does occur in the heart by way of the non-oxidative PPP, the extent of flux through the PPP can be calculated by comparison of fluxes determined by the [5-3FTJ-glucose method and the accumulation of lactate & pyruvate. A comparison of both these methods could determine non-glycolytic detritiation of [5-3Ff]-glucose, a detritiation assumed to be mainly caused by the PPP. However, in this study, glycolytic flux as measured by the accumulation of lactate, pyruvate and glucose oxidation is consistently higher than or equal to rates of glycolytic flux as measured by the [5-3H]-glucose method, but this difference was not statistically different. Although flux through the PPP cannot be accurately assessed through this method, it appears that flux through the PPP is minimal. Therefore, in light of these crude calculations, significant or detectable production of 3 H 2 0 from [5-3 H]-glucose by detritiation in the PPP is unlikely. This view is consistent with evidence that the capacity of the oxidative pentose phosphate pathway in heart is very low compared to liver and other tissues. Maximal cardiac oxidative PPP flux is generally less than 50 nmoVmin/g dry wt, a value far lower than that of glycolysis (~2500 nmol/min/g dry wt) (126,27,124,106,147). Furthermore, non-oxidative pentose phosphate pathway flux between the hexose and ribose pools in the heart is likely even lower, probably not exceeding 2 nmol/rnin/g dry wt (151,114,43). Expression analysis of transaldolase and G6PDH in control and hypertrophied cardiac muscle provides proof that both branches of the PPP are not elevated in hypertrophied heart (Figure 20 & 22)). As well, expression of transaldolase in both control and hypertrophied cardiac muscle was significandy lower than that found in liver. These two pieces of data provide further evidence that activity of PPP is low in the heart and supports the view that the PPP does not cause any non-glycolytic detritiation of any magnitude. This conclusion is also consistent with previous studies on the PPP in cardiac hypertrophy, in which changes in PPP enzyme activities, when observed, were transient and reversed with time after aortic constriction (89,30,152). D. Detritiation of [5-3H|-Glucose as a Measure of Glycolytic Flux Recently, Goodwin et al suggested that use of [5-3H]-glucose to measure glycolysis leads to an overestimation of \"true\" rates of glycolysis in isolated working rat hearts because of non-glycolytic loss of 3 H 2 0 from [5-3H]-glucose (56). This suggestion was based upon their finding that rates of glycolysis determined by measuring 3 H 2 0 production from [5-3H]-glucose were significantly higher than rates of glycolysis determined as the sum of lactate and pyruvate released to the perfusate and glucose oxidation rates measured using [ , 4C]-glucose. They proposed that glucose utilization by the non-oxidative PPP, specifically, the enzyme transaldolase, was responsible for any non-glycolytic detritiation of [5-3H]-glucose. In contrast to Goodwin et al (56), we found an excellent concordance between rates of glycolysis determined by different and independent methods (Figure 15 &17) and confirmed that glycolysis is accelerated in hypertrophied hearts. Our data indicate the non-glycolytic loss o f 3 H20 from [5-3H]-glucose is insignificant, relative to loss in the glycolytic pathway, in both hypertrophied and non-hypertrophied hearts under both high fat and low fat conditions used in our experiments. The concordance of glycolytic rates determined by the different methods also indicates that glycogen did not contribute significantly to total lactate and pyruvate accumulation in the perfusate. This is in keeping with the finding the glycogen content did not differ in hearts exposed to high fatty acids, and that the observed loss of glycogen levels in hearts exposed to low fatty acid conditions cannot easily account for the differences observed. We attribute the discrepancy between the two studies primarily to two unexpected observations by Goodwin et al. The first is the exceedingly low glucose oxidation rates reported by Goodwin et al (~100nmol'min/g dry wt), especially when considered in relation to rates of glycolysis (56). In their study, glycolytic rates are, at least, 26 fold higher than glucose oxidation rates, which contrasts dramatically with other studies with isolated working hearts perfused with Krebs-Henseleit solution containing a similarly low (i.e., 0.4 mM) concentration of fatty acid (67). In Kantor et al.'s study with similar perfusion conditions (67), glycolytic rates are only approximately 2 fold higher than glucose oxidation rates (~1900nmol/min/g dry wt). The relatively high rates of glucose oxidation observed in these studies are to be expected in the presence of low concentrations of fatty acid because of the well-established reciprocal relationship between catabolism of fatty acids and glucose in the heart (95,112,129). Moreover, if Kantor's rates of glucose oxidation are substituted for Goodwin's rates of glucose oxidation, the methodological differences observed by Goodwin are eliminated, with all three methodologies being equivalent. Use of [5-3H]-glucose, is therefore, an accurate means to determine glycolytic rates in the isolated working rat heart. The explanation for the dramatically lower glucose oxidation rates obtained in the study by Goodwin et al (56) is unclear but could be methodological. They used a modified working heart preparation in which the coronary flow was not recirculated (56) (56), as it is in the more traditional preparation (82). Thus, rather than calculating rates of glucose oxidation from the accumulation of 1 4 C 0 2 over the duration of the perfusion, perfusate containing 14CC>2 released in a single pass through the myocardium is collected into open pre-weighed vials over short time periods (i.e., 5min). An aliquot (10ml) of this perfusate is subsequently used to measure the 1 4 C 0 2 produced. Given the fact that Re-labelled lactate and pyruvate accumulate in the recirculating working heart preparation and likely contribute to overall 1 4 C 0 2 production, it may be argued that the low glucose oxidation rates reported with the use of a non-recirculating perfusion by Goodwin et al are a better representation of true glucose oxidation rates. However, review of elegant studies in which flux through the pyruvate dehydrogenase reaction (i.e., equivalent to glucose oxidation) was determined by isotopomer analysis in non-recirculating isolated rat hearts perfused with 0.4mM oleate shows that this is not the case (32,136). In these isotopomer studies, glucose oxidation rates in isolated normal hearts were at least ~l.l|amol/min/g dry wt or about 10-fold higher than those reported by Goodwin et al. and comparable to values obtained in a recirculating working heart preparation under similar conditions (67). The second unexpected observation by Goodwin et al. is the lack of linearity of 3 H 2 0 production over a 30-min time period. At the outset of perfusion, glycolytic rates determined from all three methodologies were relatively similar; however, over time, 3 H 2 0 production rates increased, whereas lactate and pyruvate production rates did not. This increase, which is the primary basis for their conclusions with respect to detritiation of [5-3H]-glucose by the PPP, is inconsistent with the findings of others, including those in the present study and previous investigations originating in Goodwin's laboratory (96,7,22,82). As summarized in Figures 17 and 18, we do not observe a time-dependent increase in 3 H 2 0 production rates, a finding consistent with the published literature (95,82,10). Of additional importance is the fact that the recent conclusions by Goodwin et al. (56) contradict their own data from a previous study (22) in which they concluded that detritiation of [5-3 H]-glucose traces glycolytic flux from exogenous glucose (22) where they found that 3 H 2 0 production from [5-3H]-glucose did not differ from 3 H 2 0 production from [2-3H]-glucose. Other studies have also corifirmed that the use of either [5-3H]-glucose and [2-3H]-glucose is equivalent (80,117,85). This is important because detritiation of [2-3H]-glucose, which occurs at the hexose-6-phosphate isomerase reaction, is not affected by the reactions in the pentose phosphate pathway implicated as being responsible for non-glycolytic loss of 3 H . E. Methodological Considerations As the majority of radiolabelled by-products will be accounted for by the production of lactate, pyruvate and glucose oxidation, some radiolabelled intermediates may be lost or unaccounted for, as in the case of [14C]-alanine via the \"pyruvate-alanine cycle\". [ 1 4C]-alanine, an essential amino acid, may be reincorporated into the myocardium and thus hidden away from major metabolic processes. However, flux through this particular pathway is typically small (~50 nmol/min/g dry wt) and is generally considered negligible. However, to confirm flux through this pathway, direct measurement of [14C]-Alanine release from [14C]-glucose will be required to determine if this is the case. F. Importance of this Study The results of our study also have great general relevance to investigators in the field. Specifically, a very important corollary of the data obtained is that measurement of the rate of 3 H i O production from [5-3H]-glucose is an accurate means to determine rates of glycolysis in isolated working normal and pathologic rat hearts. Glycolysis and 3H20 production in isolated working rat hearts are also linear and constant. This method can be applied in many different experimental settings, ranging from cell culture to isolated working rat heart preparations. Moreover, the data obtained alleviate any doubts about conclusions from many studies over the years that have used this methodology to measure glycolysis in isolated hearts and is an ideal technique for measuring glycolytic flux in many experimental settings. Appendix I - Modified [14C1-Lactate+f14Cl-Pvruvate Radiolabel Assay The quantification of lactate and pyruvate concentrations in perfusate can be used as a method of determining myocardial glycolytic rates, assuming glycogen degradation is minimal. In the event of significant contribution of glycogen to total lactate and pyruvate accumulation, this assay was developed to determine only [14C]-lactate and [14C]-pyruvate accumulation, a method that only measures exogenous glycolysis. In general, the amount of lactate and pyruvate produced combined with the amount of glucose oxidized in a given amount of time equals glycolytic flux as shown in Figure 23. Moreover, the use of U-[14C]-glucose generates [14C]-lactate and [14C]-pyruvate, end products of glycolysis which can be used to calculate glycolytic flux in conjunction with glucose oxidation rates. The assay developed for this study converts all [14C]-lactate into [14C]-pyruvate, and then decarboxylates all [14C]-lactate-derived [14C]-pyruvate and [14C]-pyruvate from glycolysis, producing 1 4 C 0 2 gas, which can be collected to calculate glycolytic flux. The main advantage of using this method will confirm the idea that all lactate produced and released into the perfusate is glycolytically derived. Because the glucose is also U-[ 1 4C] labeled, concordance between both methodologies of determining glycolytic flux should confirm that all lactate produced is from glycolysis. This assay was initially described by Lehoux et al. (73), and is a convenient and accurate method that can deterrnine the specific activities of [14C]-lactate and [ 1 4C]-pyruvate in perfusate. In order to determine accumulations for both metabolite species simultaneously, a modification was made to this assay as will be described in this section (and in Figure 24). This two-point assay initially enzymatically decarboxylzes [ 1 4C]-lactate to [14C]-pyruvate catalyzed by lactate oxidase in an enclosed environment. The 66 Cytoso l Glycolytic Flux collected K ^ - N Pyruvate > Pyruvate >collected \u00E2\u0080\u0094 \u00E2\u0080\u0094 i / f 1 / Glucose ~ s i\" Oxidation 1 ' Mitochondria CO. Collected Figure 23: Accumulation of Lactate and Pyruvate Combined with Rates of Glucose Oxidation are Equivalent to Glycolytic Flux. 67 [14C]-Lactate Lactate oxidase [14C]-Pyruvate Pyruvate oxidase \ v ^ H C O , [14C]-Acetate Hyamine trap Scintillation Counter Figure 24: Modified Two Step [14C]-Lactate & [14C]-Pyruvate Assay. 68 second half of the assay enzymatically decarboxylzes [14C]-pyruvate to [14C]-acetate by pyruvate oxidase. This half of the reaction releases one 1 4 C 0 2 per [14C]-pyruvate molecule, which is then collected by the hyamine trap. The detailed protocol is described as follows: r14Cl-Lactate+r14Cl-Pvruvate Radiolabel Assay Protocol 1. Place filter paper into scintillation tube and soak with 300 \iL hyamine hydroxide. 2. Reaction Mixture (for one reaction) -125 u L d B , 0 -40 uL KH 2 P0 4 -NaOH buffer -25 uT 2% BSA soln -20 uL 10 mM M g C | soln -5 units cocarboxylase -2 uU FAD soln -2 units catalase -1 unit Lactate Oxidase -2 units Pyruvate Oxidase -50 uT Sample 3. Incubate at 37\u00C2\u00B0C for 90 minutes with constant gentle shaking. 4. With a lOcc syringe, inject 0.5 mL of 9N H2SO4 into the reaction mixture via the rubber adaptor. 5. Incubate at 37\u00C2\u00B0C for 90 minutes with constant gentle shaking. 6. Remove scintillation vial and add 4 mL of scintillation fluid. Accumulation of [14CJ-lactate and [14C]-pyruvate was determined by the use of a two-step assay system that, 1) oxidizes all [14C]-lactate in a given sample to [14C]-pyruvate and then, 2) all remaining [14C]-pyruvate is decarboxylated by pyruvate oxidase to [14C]-acetate releasing 1 4 C 0 2 gas. All 1 4 C 0 2 gas is collected by hyamine soaked filter papers in an enclosed vessel. To ensure specificity and consistency of this two-step assay, mock trials of this assay were done with three different lactate standards with varying specific activities. The three standards consisted of lactate spiked with [14C]-lactate at: lOmM (3000dpm); 5mM (lOOOdpm); and 1 mM (500dpm). Upon proper examination of the assay protocol, [14C]-lactate recovery efficiencies ranged from ~75% to ~100%, regardless of the standard used or the day of assay (Table 4). Consistently high recoveries were found regardless of the day of experiment or the standard used. Additionally, these results do not differ whether or not the substrates ([14C]-Lactate and [14C]-Pyruvate) were dissolved in distilled water, or dissolved in 3% BSA, or when dissolved in K H buffer complexed with varying palmitate concentrations (Table 5). Positive control standards were used to test for inter- (between days) and intra-test (duplicates of standards/samples) variability and assay consistency. The three standards tested were: lOmM (3000 dpm), 5mM (1000 dpm) and 1 mM (500 dpm), all dissolved in dH 20. TJ-[14C]-lactate (0.5|iCi/mL) was used for the construction of the standards. Due to the uniform 1 4 C labeling on all three carbons of the tracer, by way of enzymatic decarboxylation under optimal conditions, one third of the original specific activity of lactate should theoretically be released as 14CC>2 and collected by the hyamine sink trap. Efficiency of recovery by the assay was calculated as follows, where \"n\" denotes number of samples: (Counts recovered by Sample * 3 Counts in Standard Statistically, intra- test variability was low (coefficient of variability= 8.1,19.8, 9.3 on three separate occasions), approximating a 10.0% range of variability, while inter-test variability across all three experiment occasions was also low (coefficient of variability = 9.5) and also approximating a 10% range of variability. Therefore, quenching or complexing of [14CJ-lactate and [14C]-pyruvate in perfusate is not significant and does not affect the efficacy of this radiolabel assay. fl * 100 = Assay Efficiency \u00C2\u00A9 2 8L & eT \u00C2\u00AB 3 ^ rt 2 co ^ CD rt o- ^ P CT rt P 5' i+ ts oo o W 0 pa ^ cT 2 rt co o O* Z 0 P m | >T3 rt a P rt = CO g co rt ft 5 P 3 co p ^ P - a _j CD P CD co \u00E2\u0080\u009E\u00E2\u0080\u00A2 a CO p ' O O. o I I t3 O ' A M 0 c 3 o CQ\" D_ CO o 3 zz TJ *< c < \u00C2\u00A30 CD CO b 1+ co b 1+ cn i+ co co c 3 o CQ CO o \u00C2\u00A30 o 1\u00E2\u0080\u00941-\u00C2\u00A30 f\u00E2\u0080\u0094I-CD CT) 1+ ->1 CO CO CD \u00E2\u0080\u0094 * CO Ko l+ i+ CO M cn * Ko * 00 bo i+ co -i rt-rt O a 0* O E 0 ro p rt-0\" rt> P H p ST \u00E2\u0080\u00A2 \u00E2\u0080\u00A2 rt> rt. 0 S P rt-n ST o rt-P rt-rt> P 0 a a < rt. 0 P rt-re 0 O a c o rt \u00C2\u00A9* 0 3' o o 0 rt-rt o n o 0 rt-rt. P 0 a 0* < a n \"I rt rt o a sr rV a ZL 70 CD O O \u00E2\u0080\u00A22 TJ 3 > CD C L 3 v i j i CD| co o oo J i cn bo o o o CO CO c n o o CD CD io oo co co C D Ko oil C D ro o ho co o CO J i -4 cn| co CO c n I cn| CD Ko CO o CO 4-CD CO CO cn co o ro I oi o c n I cn| CD Ko j - co ro CO -vl -v| 00 \u00E2\u0080\u0094>\u00E2\u0080\u00A2 co o \u00E2\u0080\u0094>\u00E2\u0080\u00A2 CD CO O \u00E2\u0080\u0094^ Ko i t - \"-vj co ro ro ro ro CD cn CD -vl s cn cn *> w ^ ro C D co cn CD cn co ro CD co ro Ko j i oo co N) o CO 00 -v| v l S CO 00 co co cn co ro ro ro ro i\i -vi cn -vi ro co CD \u00E2\u0080\u0094^ co c o _\u00C2\u00BB._\u00C2\u00BB._i.ro CO CD CO J i j \u00E2\u0080\u0094 c o o c n s b J s CO CO CO cn J - cn -\u00C2\u00BB- j i - -ro CO CD CD co ro ro ro J i CO co co cn CD -_ CD j _ co j - bo o o 00 00 -v| J i J i CO CO CO CD ro ro o o c n o o C L \" O 3 CO o o o C L T 3 3 o o o C L T 3 3 i C/) _> 05 _J C L _ \u00E2\u0080\u00945 C L co ro ro io o o cn o o C L T 3 3 CO o o o C L T 3 3 o o o C L \" O 3 I CO I cu Ki _> \u00E2\u0080\u00941 C L CO ro CO io o o cn o o a. T 3 H P re* re -s p s a P re w> rt-< P 5\" a\" n P o rt-P rt-re p E3 a o = P rt-re p O CD > tc 00 > o o t CD X CD p--v l CD o CD O IO CO cn o j i o bo CD ho tn 00 CO CD \u00E2\u0080\u00A2vi J i J i v j \u00E2\u0080\u0094-bo co co CD 00 oxi CD IO - v l ro 00 00 - v l 00 CD j i 00 -CD o 1 00 o J i v j o . vg bo 00 00 00 00 o o CD 00 ro 00 Oi CO j i 00 v4 CD vg CD CT CO + CD CO > IO . 00 o J i oo b cn o o cn 00 ro i o . N> on CD to v j b CD - to o b oo o o Ul 00 cn - v l cn 00 j i b to to ro 00 00 J i ro 00 to oo o ro ro ro ro ro b j i 00 \" v l 00 - v l to \u00E2\u0080\u0094- J i j i Ul j i v j 00 o co CD 3 \u00E2\u0080\u00A2D CD O O \u00E2\u0080\u009E > 3 eg CD 73 CD T3 Appendix II - Calculation of Glycolytic Flux from Specific Activity of [14Cl-Lactate and [14Cl-Pyruvate Glycolytic flux is determined from the amount of lactate and pyruvate produced combined with the amount of glucose oxidation that occurs in the 30 minute isolated working heart perfusion period. The counts recovered from the radiolabel assay consist of 1 4 C 0 2 gas released from the enzymatic decarboxylation of [14C]-Lactate and [ 1 4C]-Pyruvate. These 1 4 C 0 2 counts are converted to glucosyl equivalents/min/gram dry weight by extrapolating the counts registered by the 50 \xL aliquot sample to the counts in 100 mL of perfusate. Then the total amount of I 4 C dpm counts is divided by the initial specific activity of glucose in the perfusate prior to the isolated working heart mode. The amount of glucose equivalents catabolized is then divided by the time (min) and dry wt of the heart (g dry wt). The formula is on Figure 25: 74 SL 8s 0 0 re c *n re \u00E2\u0080\u00A2 \u00E2\u0080\u00A2 rs\" P\" rt-\u00C2\u00A9 * s \u00C2\u00A9 rtj O rs \u00C2\u00A9 rs \u00C2\u00A3T x rtj o 3 T3 re rs 5 rs > rs rt-O n rs rt-P rt-re p s a *>. O = P rt-re o a CD CO 9 \u00E2\u0080\u00A2< O o J> i-t fD o \u00C2\u00A9 N> ? O o o l/l N> o CO 5i o o o 3 O O o o 1 o \u00E2\u0080\u00A2o o o mol o o o o Act] 6c? o o 3^ o o <_ 1 o o % 3 r o o o glue 3 ~~ r 3 eluc glue o eluc o l/i O CD 3 r CD B' CM o H 5- \" 9L o E_ rT e_ ET \u00E2\u0080\u0094 + 5' 3 cn cr cn CD C\u00E2\u0080\u0094 o CTQ t p 3 Cu 3 3 C u G> cn O a 3 C u 3' OQ C u n -t cn ' Sa I sr o-o fl> CD _> sr sr ?S ? \u00C2\u00B0 Q) fi) sr o\" c sr a-. o 3 CO 1633 i A \u00E2\u0080\u00A2 D D CD CD CD CD 3 3 _J _J CD CD Q. CL cr >< 0) CL CL CL CL 3 ' ZJ' CQ CQ ico I--+ + 14- iro + Ico l_r \u00C2\u00BB s a S* HN HH HN I n cT c_ ST \u00C2\u00A9' 3 \u00C2\u00A9 H t \u00C2\u00BB o rt O rt \u00E2\u0080\u0094 rt fe? a* > rt rt c 3 s=_ ST rt \u00C2\u00A9\" s 65 rt rt rt-rt as s a H e ti as rt rt Appendix IV: Summary of Metabolic Data for Hearts Perfused with High Fatty Acid High Fat (1.2 mm Pa m) i HH1 Method 1 Method 2 Method 3 -Lactate+Pyruvate+GO H C Lactate Measurement of ^ 0 1832.6 4214.8 SEM 1972.1 4393.5 S E M SEM Sham 264.3 n=7 363.2 n=4 \" 1487 2 132.6 n=7 \ Hypertrophy 424.0 n=7 368.6 n=4 3962.4 298.1 n=7 \u00E2\u0080\u009E _ _ \u00E2\u0080\u0094 Control Hearts L1 L4 L5 L8 L36 \u00E2\u0080\u00A2 9 L40 Average SEM GF 1068 1230 1803 1779 1198 1380 1953 1487.2 132.6 nmol/min/g dry wt GO 248 389 499 622 331 792 445 475.1 69.6 nmol/min/g dry wt Lactate (umol) 11 6 13 11 48.1 34 22 20.8 5.8 umol/g dry wt 38 20 43 36 145 111 72 66.4 17.3 umol/g dry wt/min 1.3 0.7 1.4 1.2 4.8 3.7 2.4 2.2 0.6 Lactate (glc eqvs) 631 339 724 598 2413 1847 1115 1095.4 287.6 nmol/min/g dry wt Pyruvate (umol) 4.8 5.7 7.5 6.8 4.4 5.6 5,1 5.7 0.4 umol/g dry wt 16 19 25 23 15 19 17 19.0 1.4 umol/g dry wt/min 0.5 0.6 0.8 0.8 0.5 0.6 0.6 0.6 0.0 Pyruvate (glc eqvs) 236 281 385: 378 489 622 567 422.5 54.2 nmol/min/g dry wt 1 4 - C Lactate 1385 722 2018 2005 1532.6 435.5 nmol/min/g dry wt GF (PYR+LAC) 1115 1009 1608 1599 2902 2469 2127 1832.6 264.3 nmol/min/g dry wt GF ( 1 4C) nmol/min/g dry wt 1633 1111 2517 2628 1972.1 274.6 Hypertrophied Hearts L2 L3 L6 L7 L37 L41 L18 Average S E M GF 4632 2782 3957 3763 5098 3266 4238 3962.4 298.1 nmol/min/g dry wt GO 529 608 527 636 500 659 874 619.0 48.3 nmol/min/g dry wt Lactate (umol) 30 66 78 61 92.4 74 51 64.6 7.6 umol/g dry wt 99 220 259 203 257 205 143 198.2 22.2 umol/g dry wt/min 3.3 7.3 8.6 6.8 8.6 6.8 4.8 6.6 0.7 Lactate (glc eqvs) 1290 2970 3746 2532 4290 3422 2389 2948.4 374.7 nmol/min/g dry wt Pyruvate _(umol)_ 9.7 8.2 15.0 9.7 8.4 6.0 9.14 9.5 1.0 umol/g dry wt 32 27 50 32 28 20 30 31.5 3.5 umol/g dry wt/min 1.1 0.9 1.7 1.1 0.9 0.7 1.0 1.1 0.1 Pyruvate (glc eqvs) 421 367 722 406 933 667 1016 647.4 98.9 nmol/min/g dry wt 1 4 - C Lactate 4347 2692 4086 4150 3818.6 379.7 nmol/min/g dry wt \ nmol/min/g dry wt G F ( 1 4 C ) 4876 3300 1 4613 4786 j 4393.5 368.6 13 13 i s . o 3 f--' o Si i o . CD = TJ 2. 2 ~- < 3 K. 5\" ro to a. fl N J CO 4b. N J CD CO 3 3 \u00C2\u00AB 0 o g .3: 5* < ICQ CQ su O . C L CD K x 1 i 3 3 <=\u00C2\u00AB 3-CO - v l N J CO CO CO \u00E2\u0080\u0094i. CO m N J N J cn CD N J cn co CO CO CO CO 03 CD 1 a \u00E2\u0080\u0094 su 3; 3' fl - v | N J ( D o \u00E2\u0080\u00A2 t i CO o N J o cn CJ} CD CD C J N J -v j a P \u00E2\u0080\u00A2 \u00E2\u0080\u009E * \u00E2\u0080\u00A2*\u00C2\u00BB i n m io c j _\u00C2\u00BB 01 01 01' o N J P jg M CO 01 - v | \u00E2\u0080\u00A2 C P M U ) CO 01 CZ tr 1\u00E2\u0080\u0094 3 3-8 o o S-~ v T . S U to C Q CL CL CD 3 O 3 to\" CL fl CD N J N J CO w a co CO 01 01 - v l N J 01 45. eg O J - v l 01 M CO N J - v | N J 01 \u00E2\u0080\u00A2 I v b . 01 co Lo O J L0 CD ffl (JJ . L O 4a. O l O J N J cn u i cn co 0 P - co - v | -v N J r 1 4s. 4*. CD - v l O l p. ^ U l O J O l 3 Q 3 -n o to fl CD cotS O J ^ 0 1 CO co m CD CD 3. o 3\" 3-su 3 NJ II 4b. CD CO 3 ii l a su CD + T)| IK CD + O o -n su 3 3 su CO o p CO m 4a. cn CO CD - v l CO su o CO CD + T l CD 3-o Q _ I eg 1 I 45. 4a. eg O J 01 N J 00 m su cn cz o C L l N J 4b. CD - v l - v l O J - v l O J CO X N J O B I B L I O G R A P H Y 1. Abumrad, N . A., and al., e., Mechanism of long chain fatty acid permeation in the isolated working rat heart. Journal of Biological Chemistry, 1981. 256(17): pp. 9183-91. 2. Abumrad, N . A., Coburn, C , and Ibrahimi, A., Membrane proteins implicated in long-chain fatty acid uptake by mammalian cells: CD36, FATP, and FABPm. Biochim Biophys Acta, 1999. 1441: pp. 4-13. 3. Airman, T. J., Glazier, A. M . , Wallace, C. A., Cooper, L. D., Norsworthy, P. J., Wahid, F. N . , Al-Majali, K. M . , Trembling, P. M . , Mann, C. J., Shoulders, C. C , Graf, D., St. Lezin, E., Kurtz, T. W., Kren, V., Pravenec, M . , Ibrahimi, A. , Abumrad, N . A., Stanton, L. W., and Scott, J., Identification of Cd36 (Fat) as an insulin-resistance gene causing defective fatty acid and glucose metabolism in hypertensive rats [see comments]. Nat Genet, 1999. 21(1): pp. 76-83. 4. Alam, N . , and Saggerson, E. D., Malonyl-CoA and the regulation of fatty acid oxidation in soleus muscle. Biochem J, 1998. 334: pp. 233-241. 5. Allard, M . F., Emanuel, P. G., Russell, J. A., Bishop, S. P., Digerness, S. B., and Anderson, P. G., Preischemic glycogen reduction or glycolytic inhibition improves postischemic recovery of hypertrophied rat hearts. Am J Physiol, 1994. 267(1 Pt 2): pp. H66-74. 6. Allard, M . F., Flint, J. D., English, J. C , Henning, S. L., Salamanca, M . C , Kamimura, C. T., and English, D. R., Calcium overload during reperfusion is accelerated in isolated hypertrophied rat hearts. J Mol Cell Cardiol, 1994. 26(12): pp. 1551-63. 7. Allard, M . F., Schonekess, B. O., Henning, S. L., English, D. R., and Lopaschuk, G. D., Contribution of oxidative metabolism and glycolysis to ATP production in hypertrophied hearts. Am J Physiol, 1994. 267(2 Pt 2): pp. H742-H750. 8. Allard, M . F., and Lopaschuk, G. D., Ischemia and reperfusion injury in the hypertrophied heart. 1996, in: Myocardial ischemia:mechanisms, reperfusion, protection (M. Karmazyn, ed.), Birkhauser Verlag, Basel, Switzerland, pp. 423-452. 79 9. Allard, M . F., Henning, S. L., Wambolt, R. B., Granleese, S. R., English, D. R., and Lopaschuk, G. D., Glycogen metabolism in the aerobic hypertrophied rat heart. Circulation, 1997. 96(2): pp. 676-682. 10. Allard, M . F., Wambolt, R. B., Longnus, S. L., Grist, M . , Lydell, C. P., Parsons, H. L., Rogrigues, B., Hall, J. L., Stanley, W. C , and Bondy, G. P., Hypertrophied rat hearts are less responsive to the metabolic and functional effects of insulin. Am J Physiol, 2000. 273(3): pp. E487-E493. 11. Anderson, P. G., Bishop, S. P., and Digerness, S. B., Transmural progression of morphologic changes during ischemic contracture and reperfusion in the normal and hypertrophied rat heart. AmJPathol, 1987. 129(1): pp. 152-67. 12. Anderson, P. G., Bishop, S. P., and Digerness, S. B., Coronary vascular function and morphology in hydralazine treated DOCA salt rats. J Mol Cell Cardiol, 1988. 20(10): pp. 955-67. 13. Anderson, P. G., Bishop, S. P., and Digerness, S. B., Vascular remodeling and improvement of coronary reserve after hydralazine treatment in spontaneously hypertensive rats. Circ Res, 1989. 64(6): pp. 1127-36. 14. Anderson, P. G., Allard, M . F., Thomas, G. D., Bishop, S. P., and Digerness, S. B., Increased ischemic injury but decreased hypoxic injury in hypertrophied rat hearts. Circ Res, 1990. 67(4): pp. 948-59. 15. Andres, A., Satrustegui, J., and Machado, A., Development ofNADPH-producing pathways in rat heart. Biochem J, 1980. 186(3): pp. 799-803. 16. Anversa, P., Ricci, R., and Olivetti, G., Quantitative structural analysis of the myocardium during physiologic growth and induced cardiac hypertrophy: a review. J Am Coll Cardiol, 1986. 7: pp. 1140-9. 17. Babcock, G. T., and Wikstrom, M , Oxygen activation and the conservation of energy in cell respiration. Nature, 1992. 356: pp. 301-309. 18. Barger, P. M , and Kelly, D. P., Fatty acid utilization in the hypertrophied and failing heart: molecular regulatory mechanisms. Am J Med Sci, 1999. 318(1): pp. 36-42. 80 19. Behal, R. H., Buxton, D. B., Robertson, J. G., and Olson, M . S., Regulation of the pyruvate dehydrogenase multienzyme complex. Annu Rev Nutr, 1993. 13: pp. 497-520. 20. Bird, M . I., and Saggerson, E. D., Binding of malonyl-CoA to isolated mitochondria. Evidence for high- and low- affinity sites in liver and heart and relationship to inhibition of carnitine palmitoyl transferase activity. 1984. 21. Bishop, S. P., and Altschuld, R. A., Evidence for increased glycolytic metabolism in cardiac hypertrophy and congestive heart failure. 1971, in: Cardiac Hypertrophy (N. Alpert, ed.), Academic Press, Inc., New York, pp. 567-585. 22. Bolukoglu, H., Goodwin, G. W., Guthrie, P. H., Carmical, S. G., Chen, T. M . , and Taegtmeyer, H., Metabolic fate of glucose in reversible low-flow ischemia of the isolated working rat heart. Am J Physiol Heart Circ Physiol, 1996. 270: pp. H817-H826. 23. Bowker-Kinley, M . M . , Davis, W. I., Wu, P., Harris, R. A., and Popov, K. M . , Evidence for existence of tissue-specific regulation of the mammalian pyruvate dehydrogenase complex. Biochem J, 1998. 329(Pt 1): pp. 191-196. 24. Braunwald E, R. J., Sonnenblick EH, Mechanism of contraction of the normal and hypertrophied failing heart. 1976, in: (B. Little, ed.). 25. Brownsey, R., and Denton, R. M . , Acetyl-coenzyme A carboxylase. 1987, in: The Enzymes (P. D. Boyer, and E. G. Krebs, eds.), Academic Press, pp. 123-146. 26. Brownsey, R. W., Boone, A. N . , and Allard, M . F., Actions of insulin on the mammalian heart: metabolism, pathology and biochemical mechanisms. Cardiovasc Res, 1997. 34(1): pp. 3-24. 27. Burns, A. H., and Reddy, W. J., Hexose monophosphate shunt in isolated cardiac myocytes from normal rats. Am J Physiol Endocrinol Metab Gastrointest Physiol, 1977. 232: pp. E570-E573. 28. Buser, P. T., Wikman-Coffelt, J., Wu, S. T, Derugin, N . , Parmley, W. W., and Higgins, C. B., Postischemic recovery of mechanical performance and energy metabolism in the presence of left ventricular hypertrophy. A 3 IP-MRS study. Circ Res, 1990. 66(3): pp. 735-46. 81 29. Christian, B., El Alaoui-Talibi, Z., Moravec, M . , and Moravec, J., Palmitate oxidation by the mitochondria from volume-overloaded rat hearts. Mol Cell Biochem, 1998. 180(1-2): pp. 117-28. 30. Clark, M . G., Williams, J. F., Kolos, G., and Hickie, J. B., The role of the pentose phosphate pathway in myocardial hypertrophy. Int J Biochem, 1972. 3: pp. 629-636. 31. Collins-Nakai, R. L., Noseworthy, D., and Lopaschuk, G. D., Epinephrine increases ATP production in hearts by preferentially increasing glucose metabolism. Am J Physiol, 1994. 267(5 Pt 2): pp. H1862-H1871. 32. 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, 1997. 272: pp. 26125-26131. 33. Cunningham, M . J., Apstein, C. S., Weinberg, E. O., Vogel, W. M . , and Lorell, B. H., Influence of glucose and insulin on the exaggerated diastolic and systolic dysfunction of hypertrophied rat hearts during hypoxia. Circ Res, 1990. 66(2): pp. 406-15. 34. Denton, R. M . , Randle, P. J., and Martin, B. R., Stimulation by calcium ions of pyruvate dehydrogenase phosphate phosphatase. Biochem J, 1972. 128(1): pp. 161-3. 35. Denton, R. M . , and Halestrap, A. M . , Regulation of pyruvate metabolism in mammalian tissues. Essays Biochem, 1979. 15: pp. 37-77. 36. Denton, R. M . , Tavare, J. M . , Borthwick, A., Dickens, M . , Diggle, T. A., Edgell, N . J., Heesom, K. J., Isaad, T., Lynch, D. F., Moule, S. K., and et al., Insulin-activated protein kinases in fat and other cells. Biochem Soc Trans, 1992. 20(3): pp. 659-64. 37. Depre, C , Rider, M . H., and Hue, L., Mechanisms of control of heart glycolysis. Eur J Biochem, 1998. 258(2): pp. 277-90. 38. Depre, C , Rider, M . H., and Hue, L., Mechanisms of control of heart glycolysis. Eur J Bbchem, 1998. 258: pp. 277-290. 39. Depre, C , Vanoverschelde, J. L., and Taegtmeyer, H., Glucose for the heart. Circulation, 1999. 99(4): pp. 578-588. 82 40. Deutsch, J., Glucose-6-phosphate dehydrogenase. 1983, in: Methods of Enzymatic Analysis (H. Bergmeyer, ed.), Wiley, New York. 41. Devereux, R. B., Casale, P. N . , Hammond, I. W., Savage, D. D., Alderman, M . H., Campo, E., Alonso, D. R., and Laragh, J. H., Echocardiographic detection of pressure-overload left ventricular hypertrophy: effect of criteria and patient population. J Clin Hypertens, 1987. 3(1): pp. 66-78. 42. Do, E., Baudet, S., Verdys, M . , Touzeau, C , Bailly, F., Lucas-Heron, B., Sagniez, M . , Rossi, A., and Noireaud, J., Energy metabolism in normal and hypertrophied right ventricle of the ferret heart. J Mol Cell Cardiol, 1997. 29(7): pp. 1903-1913. 43. Dow, J. W., Nigdikar, S., and Bowditch, J., Adenine nucleotide synthesis de novo in mature rat cardiac myocytes. Biochim Biophys Acta, 1985. 847: pp. 425-435. 44. Dzau, V. J., and Re, R. N . , Evidence of the existence of renin in the heart. Circulation, 1987. 75(Suppl 1): pp. I-134-I-136. 45. Eggleston, L. V., and Krebs, H. A., Regulation of the pentose phosphate cycle. Biochemical Journal, 1974. 138: pp. 425-435. 46. el Alaoui-Talibi, Z., Landormy, S., Loireau, A., and Moravec, J., Fatty acid oxidation and mechanical performance of volume-overloaded rat hearts. Am J Physiol, 1992. 262(4 Pt 2): pp. H1068-H1074. 47. El Alaoui-Talibi, Z., Guendouz, A., Moravec, M . , and Moravec, J., Control of oxidative metabolism in volume-overloaded rat hearts: effect of propionyl-L-carnitine. Am J Physiol, 1997. 272(4 Pt 2): pp. H1615-H1624. 48. Fabiato, A., and Fabiato, F., Effects of pH on the myofilaments and the sarcoplasmic reticulum of skinned cells from cardiace and skeletal muscles. J Physiol (Lond), 1978. 276: pp. 233-55. 49. Frohlich, E. D., Apstein, C , Chobanian, A. V., Devereux, R. B., Dustan, H. P., Dzau, V., Fauad-Tarazi, F., Horan, M . J., Marcus, M . , Massie, B., and et al., The heart in hypertension [published erratum appears in N Engl J Med 1992 Dec 10;327(24):1768] [see comments]. N Engl J Med, 1992. 327(14): pp. 998-1008. 50. Gaasch, W. H., Zile, M . R., Hoshino, P. K., Weinberg, E. O., Rhodes, D. R., and Apstein, C. S., Tolerance of the hypertrophic heart to ischemia. Studies in 83 compensated and fading dog hearts with pressure overload hypertrophy. Circulation, 1990. 81(5): pp. 1644-53. 51. Gardner, D. G., Wu, J. P., LaPointe, M . C , Hedges, B. K., and Deschepper, C. F., Expression of the gene for the atrial natriuetic peptide in cardiac myocytes in vitro. Cardiovasc Drugs Ther, 1988. 2: pp. 479-86. 52. Golden, A. L., Bright, J. M . , Pohost, G. M , and Pike, M . M . , Ischemic dysfunction and impaired recovery in hypertensive hypertrophied hearts is associated with exaggerated intracellular sodium accumulation. American Journal of Hypertension, 1994. 7(8): pp. 745-54. 53. Goodwin, G. W., Zhang, B., Paxton, R., and Harris, R. A., Determination of activity and activity state of branched-chain alpha-keto acid dehydrogenase in rat tissues. Methods in Enzymology, 1988. 166: pp. 189-201. 54. Goodwin, G. W., Arteaga, J. R., and Taegtmeyer, H., Glycogen turnover in the isolated working rat heart. Journal of Biological Chemistry, 1995. 270(16): pp. 9234-40. 55. Goodwin, G. W., Ahmad, F., and Taegtmeyer, H., Preferential oxidation of glycogen in isolated working rat heart. Journal of Clinical Investigation, 1996. 97(6): pp. 1409-16. 56. Goodwin, G. W., Cohen, D. M . , and Taegtmeyer, H., [5-SH]-glucose overestimates glycolytic flux in isolated working rat heart: role of the pentose phosphate pathway. Am J Physiol, 2001. 280: pp. E502-E508. 57. Grossman, W., Jones, D., and McLaurin, L. P., Wall stress and patterns of hypertrophy in the human left ventricle. J Clin Invest, 1975. 56(1): pp. 56-64. 58. Gwathmey, J. K., and Morgan, J. P., Altered calcium handling in experimental pressure-overload hypertrophy in the ferret. Circ Res, 1985. 57(6): pp. 836-43. 59. Hanrath, P., Mathey, D. G., Siegert, R., and Bleifeld, W., Left ventricular relaxation and filling pattern in different forms of left ventricular hypertrophy: an echocardiographic study. Am J Cardiol, 1980. 45(1): pp. 15-23. 60. Harris, D. A., and Das, A. M . , Control of mitochondrial ATP synthesis in the heart. Biochem J, 1991. 280: pp. 561-573. 84 61. Heckmann, M . , and Zimmer, H. G., Effects of triiodothyronine in spontaneously hypertensive rats. Studies on cardiac metabolism, function, and heart weight. Basic Research in Cardiology, 1992. 87(4): pp. 333-343. 62. Henning, S. L., Wambolt, R. B., Schonekess, B. O., Lopaschuk, G. D., and Allard, M . F., Contribution of glycogen to aerobic myocardial glucose utilization. Circulation, 1996. 93(8): pp. 1549-1555. 63. Hochachka, P. W., and Mommsen, T. P., Protons and anaerobiosis. Science, 1983.219(4591): pp. 1391-7. 64. Jeffrey, F. M . H., Storey, C. J., Sherry, A. D., and Malloy, C. R., 1 3 C isotopomer model for estimation of anaplerosis substrate oxidation via acetyl-CoA. Am J Physiol Endocrinol Metab, 1996. 271: pp. E788-E799. 65. Jessen, M . E., Kovarik, T. E., Jeffrey, F. M . H., Sherry, A. D., Storey, C. J., Chao, R. Y. , Ring, W. S., and Malloy, C. R., Effects of amino acids on substrate selection, anaplerosis, and left-ventricular function in the ischemic reperfused rat hearts. J Clin Invest, 1993. 92: pp. 831-839. 66. Kagaya, Y. , Kanno, Y. , Takeyama, D., Ishide, N . , Maruyama, Y. , Takahashi, T., Ido, T., and Takishima, T., Effects of long-term pressure overload on regional myocardial glucose and free fatty acid uptake in rats. A quantitative autoradiographic study. Circulation, 1990. 81(4): pp. 1353-1361. 67. Kantor, P. F., Lucien, A., Kozak, R., and Lopaschuk, G. D., 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 [see comments]. Circulation Research, 2000. 86(5): pp. 580-588. 68. Keller, A., Rouzeau, J. D., Farhadian, F., Wisnewsky, C , Marotte, F., Lamande, N . , Samuel, J. L., Schwartz, K., Lazar, M . , and Lucas, M . , Differential expression of alpha- and beta-enolase genes during rat heart development and hypertrophy. Am J Physiol, 1995. 269(6 Pt 2): pp. H1843-H1851. 69. Komuro, I., Kaida, T., Shibazaki, Y. , and al., e., Stretching cardiac myocytes stimulatesprotooncogene expression. J Biol Chem, 1990. 265: pp. 3595-3598. 70. Kornberg, H. L., Tricarboxylic acid cycles. BioEssays, 1987. 7: pp. 236-238. 85 71. Krebs, H. A., The history of the tricarboxylic acid cycle. Perspect Biol Med, 1970. 14: pp. 154-170. 72. Lazdunski, M . , Frelin, C , and P., V., The sodium/hydrogen exchange system in cardiac cells: its biochemical and pharmacological properties and its role in regulating internal concentrations of sodium and internal pH. J Mol Cell Cardiol, 1985. 17: pp.1029-1042. 73. Lehoux, E. A., Svedruzic, Z., and Spivey, H. O., Determination of the specific radioactivity of [14C]-lactate by enzymatic decarboxylation and 14C02 collection. Anal Biochem, 1997. 253: pp. 190-195. 74. Levy, D., and Kannel, W. B., Cardiovascular risks: new insights from Framingham. Am Heart J, 1988. 116(1 Pt 2): pp. 266-72. 75. Levy, D., Left ventricular hypertrophy. Epidemiological insights from the Framingham Heart Study. Drugs, 1988. 35 Suppl 5: pp. 1-5. 76. Levy, D., Anderson, K. M . , Savage, D. D., Kannel, W. B., Christiansen, J. C , and Castelli, W. P., Echocardiographically detected left ventricular hypertrophy: prevalence and risk factors. The Framingham Heart Study. Ann Intern Med, 1988. 108(1): pp. 7-13. 77. Lewandowski, E. D., Chari, M . V., and Roberts, R., NMR studies of beta-oxidation and short chain fatty acid metabolism during recovery of reperfused hearts. Am J Physiol, 1991. 261: pp. H354-H363. 78. Lewandowski, E. D., Kudej, R. K., White, L. T., O'Donnell, J. M . , and Vatner, S. F., Mitochondrial preference for short chain faty acid oxidation during coronary artery constriction. Circulation, 2002. 105(3): pp. 367-72. 79. Longnus, S. L., Wambolt, R. B., Barr, R. L., Lopaschuk, G. D., and Allard, M . F., Regulation of myocardial fatty acid oxidation by substrate supply. Am J Physiol, 2001. 281(4): pp. H1561-H1567. 80. Lopaschuk, G. D., Wambolt, R. B., and Barr, R. L., An imbalance between glycolysis and glucose oxidation is a possible explanation for the detrimental effects of high levels offatty acids during aerobic reperfusion of ischemic hearts. J Pharmacol Exp Ther, 1993. 264(1): pp. 135-144. 86 81. Lopaschuk, G. D., Belke, D. D., Gamble, J., Itoi, T., and Schonekess, B. O., Regulation of fatty acid oxidation in the mammalian heart in health and disease. Biochim Biophys Acta, 1994. 1213(3): pp. 263-76. 82. Lopaschuk, G. D., and Barr, R. L., Measurements of fatty acid and carbohydrate metabolism in the isolated working rat heart. Mol Cell Biochem, 1997. 172(1-2): pp. 137-147. 83. Lorell, B. H., and Grossman, W., Cardiac hypertrophy: the consequences for diastole. J Am Coll Cardiol, 1987. 9(5): pp. 1189-93. 84. Lydell, C. P., Chan, A., Wambolt, R. B., Sambandam, N. , Parsons, H., Bondy, G. P., Rodrigues, B., Popov, K. M . , Harris, R. A., Brownsey, R. W., and Allard, M . F., Pyruvate dehydrogenase and the regulation of glucose oxidation in hypertrophied rat hearts. Cardiovasc Res, 2002. 53(4): pp. 841-51. 85. McCormack, J. G., Barr, R. L., Wolff, A. A., and Lopaschuk, G., Ranolazine stimulates glucose oxidation in normoxic, ischemic, and reperfused ischemic rat hearts. Circulation, 1996. 93: pp. 135-142. 86. McGarry, J. D., Malonyl-CoA and carnitine palmitoyltransferase I: an expanding partnership. Biochemical Society Transactions, 1995. 23(3): pp. 481-5. 87. McGarry, J. D., The mitochondrial carnitine palmitoyltransferase system: its broadening role in fuel homoeostasis and new insights into its molecular features. Biochemical Society Transactions, 1995. 23(2): pp. 321-4. 88. McGarry, J. D., and Brown, N . F., The mitchondrial carnitine palmitoyl transferase system. From concept to molecular analysis. Eur J Biochem, 1997. 244: pp. 1-14. 89. Meerson, F. Z., Spiritchev, V. B., Pshennikova, M . G., and Djachkova, L. V. , The role of the pentose-phosphate pathwayin adjustment of the heart to a high load and the development of myocardial hypertrophy. Experientia, 1967. 23: pp. 530-532. 90. Mitchell, P., Vectorial chemistry and the molecular mechanics of chemiosmotic coupling: power transmission by proticity. Biochem Soc Trans, 1976. 4: pp. 398-430. 87 91. Mochizuki, T., Eberli, F. R., Ngoy, S., Apstein, C. S., and Lorell, B. H., Effects of brief repetitive ischemia on contractility, relaxation, and coronary flow. Exaggerated postischemic diastolic dysfunction in pressure-overload hypertrophy. Circ Res, 1993. 73(3): pp. 550-8. 92. Morgan, H. E., and Baker, K. M . , Cardiac hypertrophy. Mechanical, neural, and endocrine dependence. Circulation, 1991. 83: pp. 13-25. 93. Nakanishi, H., Makino, N. , and Hata, T., Sarcolemmal Ca2+ transport activities in cardiac hypertrophy caused by pressure overload. Am J Physiol, 1989. 257: pp. H349-356. 94. Nayler, W. G., Calcium and cell death. Eur Heart J, 1983. 4([Suppl.]): pp. 33-41. 95. Neely, J. R., Denton, R. M . , England, P. J., and Randle, P. J., The effects of increased work on the tricarboxylate cycle and its interactions with glycolysis in the perfused rat heart. Biochem J, 1972. 128: pp. 147-159. 96. Neely, J. R., Rovetto, M . J., and Oram, J. F., Myocardial utilization of carbohydrate and lipids. Prog Cardiovasc Dis, 1972. 15: pp. 289-329. 97. Neely, J. R., and Morgan, H. E., Relationship between carbohydrate and lipid metabolism and the energy balance of heart muscle. Annu Rev Physiol, 1974. 36: pp. 413-457. 98. Normann, P. T., Norseth, J., and Flatmark, T., Acyl-CoA synthetase activity of rat heart mitochondria. Substrate specificity with special reference to very-long-chain fatty acids. Biochimica et Biophysica Acta, 1983. 752(3): pp. 474-81. 99. Opie, L. H., 1998, The heart physiology, from cell to circulation, Lippincott_Raven Publishers, Philadelphia, USA, pp. 295-342. 100. Oram, J. F., Bennetch, S. L., and Neely, J. R., Regulation of fatty acid utilization in isolated perfused rat hearts. Journal of Biological Chemistry, 1973. 248(15): pp. 5299-309. 101. Oram, J. F., Wenger, J. I., and Neely, J. R., Regulation of long chain fatty acid activation in heart muscle. Journal of Biological Chemistry, 1975. 250(1): pp. 73-8. 102. Panchal, A. R., Comte, B., Huang, H , Kerwin, T., Darvish, A., Des Rosiers, C , Brunengraber, H., and Stanley, W. C , Pardoning of pyruvate between oxidation 88 and anaplerosis in swine hearts. Am J Physiol Heart Circ Physiol, 2000. 279: pp. H2390-H2398. 103. Patel, M . S., and Roche, T. E., Molecular biology and biochemistry of pyruvate dehydrogenase complexes. FASEB J, 1990. 4(14): pp. 3224-3233. 104. Pauly, D. F., Yoon, S. B., and McMillin, J. B., Carnitine-acylcarnitine translocase in ischemia: evidence for sulfhydryl modification. Am J Physiol (Heart Circ Physiol.), 1987. 253: pp. H1557-H1565. 105. Pernollet, M . G., Devynck, M . A., and Meyer, P., Abnormal calcium handling by isolated cardiac plasma membrane from spontaneously hypertensive rats. Clin Science, 1981. 61: pp. 45s-48s. 106. Pfeifer, R., Karl, G., and Scholz, R., Does the pentose cycle paly a major role for NADPH supply in the heart? Biol Chem Hoppe Seyler, 1986. 367: pp. 1061-1068. 107. Popov, K. M . , Kedishvili, N . Y. , Zhao, Y. , Shimomura, Y. , Crabb, D. W., and Harris, R. A., Primary structure of pyruvate dehydrogenase kinase establishes a new family of eukaryotic protein kinases. J Biol Chem, 1993. 268(35): pp. 26602-26606. 108. Popov, K. M . , Kedishvili, N . Y. , Zhao, Y. , Gudi, R., and Harris, R. A., Molecular cloning of the p45 subunit ofpyruvate dehydrogenase kinase. J Biol Chem, 1994. 269(47): pp. 29720-29724. 109. Priestman, D. A., Orfali, K. A., and Sugden, M . C , Pyruvate inhibition of pyruvate dehydrogenase kinase. Effects of progressive starvation and hyperthyroidism in vivo, and of dibutyryl cyclic AMP and fatty acids in cultured cardiac myocytes. FEBS Letters, 1996. 393(2-3): pp. 174-178. 110. Priestman, D. A. , Orfali, K. A., and Sugden, M . C , Inhibition of pyruvate dehydrogenase kinase by pyruvate in cultured cardiac myocytes. Biochemical Society Transactions, 1997. 25(1): pp. 101S. 111. Randle, P. J., and Tubbs, P. K., Carbohydrate and fatty acid metabolism. 1979, in: Handbook of Physiology. The Cardiovascular System: The Heart (R. M . Berne, and N . Sperelakis, eds.), American Physiological Society, Bethesda, Maryland, pp. 805-844. 89 112. Randle, P. J., Fuel selection in animals. Biochem Soc Trans, 1986. 14(5): pp. 799-806. 113. Randle, P. J., Priestman, D. A., Mistry, S. C , and Halsall, A., Glucose fatty acid interactions and the regulation of glucose disposal. J Cell Biochem, 1994. 55(Suppl): pp. 1-11. 114. Reimer, K. A., Hill, M . L., and Jennings, R. B., Prolonged depletion of ATP and of the adenine nucleotide pool due to delayed resynthesis of adenine nucleotides following reversible myocardial ischemic injury in dogs. J Mol Cell Cardiol, 1981. 13: pp.229-239. 115. Sack, M . N . , Rader, T. A., Park, S., Bastin, J., McCune, S. A. , and Kelly, D. P., Fatty acid oxidation enzyme gene expression is downregulated in the failing heart. Circulation, 1996. 94(11): pp. 2837-42. 116. Saddik, M . , and Lopaschuk, G. D., Myocardial triglyceride turnover and contribution to energy substrate utilization in isolated working rat hearts. J Biol Chem, 1991. 266(13): pp. 8162-70. 117. Saddik, M , and Lopaschuk, G., Triacylglycerol turnover in isolated working rat hearts of acutely diabetic rats. Can J Physiol Pharmacol, 1994. 72: pp. 1110-1119. 118. Sambandam, N. , Lopaschuk, G., Brownsey, R., and Allard, M . , Energy metabolism in the hypertrophied heart. Heart Failure Reviews, 2002. 7: pp. 161-173. 119. Scheuer, J., and Buttrick, P., The cardiac hypertrophic responses to pathologic and physiologic loads. Circulation, 1987. 75(1 Pt 2): pp. 163-8. 120. Schonekess, B. O., Brindley, P. G., and Lopaschuk, G. D., Calcium regulation of glycolysis, glucose oxidation, and fatty acid oxidation in the aerobic and ischemic heart. Can J Physiol Pharmacol, 1995. 73(11): pp. 1632-40. 121. Schonekess, B. O., Allard, M . F., and Lopaschuk, G. D., Recovery of glycolysis and oxidative metabolism during postischemic reperfusion of hypertrophied rat hearts. Am J Physiol, 1996. 271(2 Pt 2): pp. H798-H805. 122. Schonekess, B. O., Allard, M . F., Henning, S. L., Wambolt, R. B., and Lopaschuk, G. D., Contribution of glycogen and exogenous glucose to glucose 90 metabolism during ischemia in the hypertrophied rat heart. Circ Res, 1997. 81(4): pp. 540-9. 123. Schultz, H., An overview of the pathways for the beta-oxidation offatty acids. World Review of Nutrition & Dietetics, 1994. 75: pp. 18-21. 124. Severin, S. E., and Stepanova, N . G., Interrelationship between glycolysis and the anaerobic part of the pentose phosphate pathway of carbohydrate metabolism in the myocardium. Adv Enzyme Regul, 1980. 19: pp. 235-255. 125. Sharma, R. V. , Buttler, C. A., and Bhalla, R. C , Alterations in plasma membrane properties of the myocardium of spontaneously hypertensive rats. Hypertension, 1986. 8: pp. 583-591. 126. Shipp, J. C , Delcher, H. K., and Crevasse, L. E., Glucose metabolism by the hexose monophosphate pathway in the perfused rat heart. Biochim Biophys Acta, 1964. 86: pp.399-402. 127. Snoeckx, L. H., van der Vusse, G. J., Coumans, W. A., Willemsen, P. H., van der Nagel, T., and Reneman, R. S., Myocardial function in normal and spontaneously hypertensive rats during reperfusion after a period of global ischaemia. Cardiovasc Res, 1986. 20(1): pp. 67-75. 128. Stanley, W. C , Lopaschuk, G. D., and McCormack, J. G., Regulation of energy substrate metabolism in the diabetic heart. Cardiovasc Res, 1997. 34(1): pp. 25-33. 129. Stanley, W. C , Lopaschuk, G. D., Hall, J. L., and McCormack, J. G., Regulation of myocardial carbohydrate metabolism under normal and ischaemic conditions. Potential for pharmacological interventions. Cardiovasc Res, 1997. 33(2): pp. 243-257. 130. Sundqvist, K. E., Heikkila, J., Hassinen, I. E., and Hiltunen, J. K., Role of NADP+ (correct)-linked malic enzymes as regulators of the pool size of tricarboxylic acid-cycle intermediates in the perfused rat heart. Cardiovasc Res, 1987. 33: pp. 853-857. 131. Sundqvist, K. E., Hiltunen, J. K., and Hassinen, I. E., Pyruvate carboxylation in the rat heart: role ofbiotin-dependent enzymes. Biochem J, 1989. 257: pp. 913-916. 91 132. Taegtmeyer, H., and Overturf, M . L., Effects of moderate hypertension on cardiac function and metabolism in the rabbit. Hypertension, 1988. 11(5): pp. 416-426. 133. Tani, M . , and Neely, J. R., Role of intracellular Na+ in Ca2+ overload and depressed recovery of ventricular function of reperfused ischemic rat hearts. Possible involvement of IT-Na+ and Na+-Ca2+ exchange. Circ Res, 1989. 65(4): pp. 1045-56. 134. Tanjiri, H., Cardiac hypertrophy in spontaneously hypertensive rats. Jpn Heart J, 1975. 16: pp.174-188. 135. van der Vusse, G. J., van Bilsen, M . , and Glatz, J. F., Cardiac fatty acid uptake and transport in health and disease. Cardiovasc Res, 2000. 45(2): pp. 279-93. 136. Vincent, G., Comte, B., Poirier, M . , and Rosiers, C. D., Citrate release by perfused rat hearts: a window on mitochondrial cataplerosis. Am J Physiol Endocrinol Metab, 2000. 278(5): pp. E846-E856. 137. Wambolt, R. B., Henning, S. L., English, D. R., Bondy, G. P., and Allard, M . F., Regression of cardiac hypertrophy normalizes glucose metabolism and left ventricular function during reperfusion. J Mol Cell Cardiol, 1997. 29(3): pp. 939-48. 138. Wambolt, R. B., English, D. R., Henning, S. L., Bondy, G. P., and Allard, M . F., Dichloroacetate improves post-ischemic function of hypertrophied rat hearts. J Mol Cell Cardiol, 1997. 29: pp. A213. 139. Wambolt, R. B., Henning, S. L., English, D. R., Dyachkova, Y. , Lopaschuk, G. D., and Allard, M . F., Glucose utilization and glycogen turnover are accelerated in hypertrophied rat hearts during severe low-flow ischemia. J Mol Cell Cardiol, 1999.31(3): pp. 493-502. 140. Wambolt, R. B., Grist, M . , and Allard, M . , Accelerated rates of glycolysis in hypertrophied rat hearts: are they a methodological artifact? FASEB Journal, 2000. 14(4): pp. A417. 141. Wambolt, R. B , Lopaschuk, G. D., Brownsey, R. W., and Allard, M . F., Dichloroacetate improves post-ischemic function of hypertrophied rat hearts. J Am Coll Cardiol, 2000. 36(4): pp. 1378-1385. 92 142. Wambolt, R. B., Lopaschuk, G. D., Brownsey, R. W., and Allard, M . F., Dichloroacetate improves postischemic function of hypertrophied rat hearts. Journal of the American College of Cardiology, 2000. 36(4): pp. 1378-1385. 143. Wambolt, R. B., Grist, M . , Bondy, G. P., and Allard, M . F., Accelerated glycolysis and greater post-ischemic dysfunction in hypertrophied rat hearts are independent of coronary flow. Can J Cardiol, 2001. (in press). 144. Wexler, L. F., Lorell, B. H., Momomura, S., Weinberg, E. O., Ingwall, J. S., and Apstein, C. S., Enhanced sensitivity to hypoxia-induced diastolic dysfunction in pressure-overload left ventricular hypertrophy in the rat: role of high-energy phosphate depletion. Circ Res, 1988. 62(4): pp. 766-75. 145. Williams, J. F., A critical examination of the evidence for the reactions of the pentose phosphate pathway in animal tissues. Trends Biochem Sci, 1980. 5: pp. 315-320. 146. Williams, J. F., and Blackmore, P. F., Non-oxidative synthesis of pentose 5-phosphate from hexose 6-phosphate and triose phosphate by the L-type pathway. International Journal of Biochemistry, 1983. 15: pp. 797-816. 147. Williams, J. F., Arora, K. K., and Longenecker, J. P., The pentose pathway: a random harvest. Impediments which oppose acceptance of the classical (F-type) pentose cycle for liver, some neoplasms and photosynthetic tissue. The case for the L-type pentose pathway. Int J Biochem, 1987. 19(9): pp. 749-817. 148. Wood, T., 1985, The Pentose Phosphate Pathway, Academic Press, Inc., London. 149. Wood, T., Physiological functions of the pentose phosphate pathway. Cell Biochem Funct, 1986. 4: pp. 241-247. 150. Zhang, J., Duncker, D. J., Ya, X. , Zhang, Y. , Pavek, T., Wei, H., Merkle, Ft., Ugurbil, K., From, A. H., and Bache, R. J., Effect of left ventricular hypertrophy secondary to chronic pressure overload on transmural myocardial 2-deoxyglucose uptake. A 31P NMR spectroscopic study. Circulation, 1995. 92(5): pp.1274-1283. 151. Zimmer, H. G., and Gerlach, E., Stimulation of myocardial adenine nucleotide biosynthesis by pentoses andpentitols. Pflugers Arch, 1978. 376(3): pp. 223-227. 93 152. Zirnmer, H. G., Ibel, H., and Gerlach, E., Significance of the hexose monophosphate shunt in experimentally induced cardiac hypertrophy. Basic Res Cardiol, 1980. 75(1): pp. 207-213. 153. Zirnmer, H. G., Bunger, R., Koschine, H., and Steinkopff, G., Rapid stimulation on the hexose monophosphate shunt in the isolated perfused rat heart: possible involvement of oxidized glutathione. J Mol Cell Cardiol, 1981. 13(5):pp. 531-535. 154. Zirnmer, H. G., Ibel, H., Suchner, U., and Schad, H., Ribose intervention in the cardiac pentose phosphate pathway is not species-specific. Science, 1984. 223: pp. 712-714. 155. Zirnmer, H. G., Ibel, H., and Suchner, U., Beta-adrenergic agonists stimulate the oxidative pentose phosphate pathway in the rat heart. Circ Res, 1990. 67(6): pp. 1525-1534. 156. Zirnmer, H. G., Lankat-Buttgereit, B., Kolbeck-Ruhmkorff, C , Nagano, T., and Zierhut, W., Effects of norepinephrine on the oxidative pentose phosphate pathway in the rat heart. Circulation Research, 1992. 71: pp. 451-459. 157. Kashimoto, S., Tsuji, Y. , and Ktimazawa, T., Effects ofhalothane and enflurane on myocardial metabolism during postischaemic reperfusion in the rat. Acta Anaesthesiol Scand., 1987. 31(1): pp. 44-47. "@en . "Thesis/Dissertation"@en . "2003-05"@en . "10.14288/1.0090969"@en . "eng"@en . "Pathology"@en . "Vancouver : University of British Columbia Library"@en . "University of British Columbia"@en . "For non-commercial purposes only, such as research, private study and education. Additional conditions apply, see Terms of Use https://open.library.ubc.ca/terms_of_use."@en . "Graduate"@en . "Accelerated rates of glycolysis in cardiac hypertrophy : are they a methodological artifact?"@en . "Text"@en . "http://hdl.handle.net/2429/13883"@en .