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Glycogen turnover in heart-derived H9c2 cells Cheung, Kevin K.Y. 2005

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G L Y C O G E N TURNOVER IN HEART-DERIVED H9c2 C E L L S by  KEVIN K.Y. CHEUNG B.Sc. (Hons), The University of Toronto, 1998  A THESIS SUMBITTED IN PARTIAL FULFILLMENT OF T H E REQUIREMENTS FOR T H E DEGREE OF MASTER OF SCIENCE in T H E FACULTY OF GRADUATE STUDIES  (PATHOLOGY & LABORATORY MEDICINE)  T H E UNIVERSITY OF BRITISH COLUMBIA  February 2005  © Kevin K.Y. Cheung, 2005  ABSTRACT 5'-AMP activated protein kinase (AMPK) plays a central role in modulating the energy metabolism in the heart when the heart is under metabolic stress. In theory, AMPK activation should decrease glycogen synthesis and enhance its degradation. Thus, we determined the effect of AMPK activation on glycogen metabolism in H9c2 cells, a heart-derived cell line. 5-amino-4-imidazolecarboxamide riboside (AICAR, 1 mM and 2 mM), recognized activator of AMPK, causes an increase in glycogen turnover by simultaneously stimulating degradation and synthesis of glycogen, and significantly increased the rate of glycolysis and glucose uptake. Glycogen turnover also occurs in H9c2 cells under a variety of conditions, even in the setting of very high rates of glycogen degradation. However, data on the phosphorylation state of AMPK and ACC suggested that AMPK was not activated by AICAR.  In addition, the results on the  activity level of glycogen synthase (GS) and glycogen phosphorylase (GP) did not show significant changes with AICAR stimulation. The detected activity levels of GS and GP did not account for the observed increase in glycogen synthesis. I speculate that AICAR's effect on glycogen turnover could be mediated by allosteric activation. Overall, H9c2 succeeded in providing a readily accessible and established isolated cell system with cardiac myocyte properties as a tool to investigate the role of AMPK in cardiac energy metabolism. The results and methods obtained from H9c2 could be readily adapted to be used on isolated adult cardiomyocytes for better representation of heart tissue.  ii  TABLE OF CONTENTS ABSTRACT  ii  LIST OF FIGURES  vii  LIST OF TABLES  ix  LIST OF ABBREVIATIONS ACKNOWLEDGEMENTS  x xi  Chapter 1 - INTRODUCTION  1  Ischemic Heart Disease  3  Myocardial Energy Metabolism  4  Fatty Acid Metabolism  7  Carbohydrate Metabolism  8  Lactate Metabolism  11  Pyruvate Metabolism  12  Myocardial Glycogen Metabolism  14  Contribution to Energy Production  14  Glycogen Turnover  15  Control of Glycogen Turnover  16  Glycogen Metabolism in Myocardial Ischemia & Reperfusion  19  5'AMP-activated Protein Kinase (AMPK)  21  Control of AMPK activity  21  AMPK Modulation of Lipid Metabolism in the Heart  23  AMPK Modulation of Glucose Uptake & Glycolysis in the Heart  23  AMPK Regulation of Glycogen Metabolism in the Heart  24  iii  Approaches to Investigate Cardiac Glycogen Metabolism  25  Hypothesis and Objectives  29  General Experimental Approach  29  CHAPTER 2 - MATERIALS AND METHODS  30  Cell Culture Conditions  30  Assessment of Cell Culture Development  32  Glycogen Metabolism  36  Total Glycogen Content  36  Glycogen Synthesis  37  Exogenous Glucose Metabolism  38  Glycolysis  38  Glucose Uptake  39  Lactate Content  39  Activity of Regulatory Enzymes and Kinases  40  Glycogen Phosphorylase  40  Glycogen Synthase  41  A M P K and A C C phosphorylation patterns - Western Blotting  43  Other Methods  44  Total Protein Content  44  Data Analysis  44  CHAPTER 3 - ESTABLISHING A CELLULAR M O D E L OF ENERGY METABOLISM USING THE L 6 MYOGENIC C E L L LINE Background  45 45  Experimental Design and Protocol  47  Results  49 L6 cell differentiation  .49  Morphology  49  Creatine Kinase Activity  49  Total Glycogen Content  52  Glycogen and glucose metabolism in L6 cells  54  Protocol 1  54  Glycogen Depletion and Re-Synthesis Protocol II  56 59  Glycogen Degradation and Synthesis  59  Glycolytic Flux  62  Lactate Production  62  Summary  65  Discussion  66  CHAPTER 4 - GLUCOSE AND GLYCOGEN METABOLISM IN HEART-DERIVED H9c2 CELLS  69 Introduction  69  Experimental Design and Protocol  71  Results  72 Morphologic Characteristics  72  Expression of cardiac-specific proteins  73  Myogenic Phenotype  76  v  Total Glycogen Content  76  Glycolysis  76  Cardiac Phenotype  80  Total Glycogen Content  80  Glycogen Turnover  82  Glycogen Degradation  82  Glycogen Synthesis  82  Simultaneous Synthesis and Degradation  83  Glycolysis  86  Glucose Uptake  88  Activity of Glycogen Phosphorylase  90  Activity of Glycogen Synthase  90  Phosphorylation of A M P K & A C C  93  Summary CHAPTER 5 - DISCUSSION  95 97  Metabolic Actions of the A M P K Activator: AICAR  97  Are the metabolic actions of AICAR mediated by AMPK?  99  Methodological considerations Underestimation of glycogen synthesis  101  Lack of insulin effect  101  H9c2 cells as a model  102  Future directions BIBLOGRAPHY  100  104 106  vi  LIST OF FIGURES Fig. 1.1 - Overview of Myocardial Energy Metabolism  5  Fig. 1.2 - Glycolysis  10  Fig. 1.3 - Overview of Myocardial Glycogen Metabolism  13  Fig. 1.4 - Regulation of Myocardial Glycogen Metabolism  18  Fig. 1.5 - Regulation of A M P K by allosteric factors and covalent modifications  22  Fig. 3.1 - L6 Morphological Properties: Differentiation from Day 0 to Day 8  50  Fig. 3.2 - L6 Biochemical-Metabolic Properties: Creatine Kinase Activity  51  Fig. 3.3 - L6 Biochemical-Metabolic Properties: Glycogen Content  53  Fig. 3.4 - L6 Cells Protocol I with [ H] glucose  55  Fig. 3.5 - L6 Cells Protocol I: Glycogen Synthesis & Degradation  58  Fig. 3.6 - L6 Cells Protocol II with [ H] glucose  60  Fig. 3.7 - L6 Cells: Lactate Production  64  3  3  Fig. 4.1 - H9c2 Morphological Properties: Cardiac Differentiation Vs. Myogenic Differentiation  74  Fig. 4.2 - Cardiac Troponin-T Protein Expression (Courtesy of Ramesh Saeedi, PhD student, UBC)  75  Fig. 4.3 - H9c2 Cells Skeletal Phenotype: Total Glycogen Content  78  Fig. 4.4 - H9c2 Cells Skeletal Phenotype: Rate of Glycolytic Flux  79  Fig. 4.5 - H9c2 Cells Cardiac Phenotype: Total Glycogen Content  81  Fig. 4.6 - H9c2 Cells Cardiac Phenotype: Rate of Glycogen Synthesis and Degradation  85  Fig. 4.7 - H9c2 Cells Cardiac Phenotype: Rate of Glycolytic Flux  87  Fig. 4.8 - H9c2 Cells Cardiac Phenotype: Calculated Glucose Uptake  89  Fig. 4.9 - H9c2 Cells Cardiac Phenotype: Activity of Active Glycogen Phosphorylase (Data obtained with the technical assistance of Hannah Parsons, UBC)  91  Fig. 4.10 - H9c2 Cells Cardiac Phenotype: Activity of Active Glycogen Synthase (Data obtained with the technical assistance of Hannah Parsons, UBC)  92  Fig. 4.11- H9c2 Cells Cardiac Phenotype: Irnmunoblot Analysis on Phosphorylation of A M P K and A C C (Data obtained with the technical assistance of Hannah Parsons, UBC)  94  viii  LIST OF TABLES Table 3.1 - L6 Cells: Glycogen Synthesis and Degradation  57  Table 3.2 - L6 Cells: Glycogen Synthesis and Degradation  61  Table 3.3 - L6 Cells: Rate of Glycolytic Flux  ,  Table 4.1 - H9c2 Cells Cardiac Phenotype: Glycogen Synthesis and Degradation  63 84  ix  LIST OF ABBREVIATIONS  6-PG  6-phosphogluconate  AICAR  5-ammoimidazole-4-carboxamideribonucleoside  AMPK  5'-AMP activated protein kinase  Ara-C  cytosine (3-D-Arabinofuranoside  CK  Creatine Kinase  DMEM  Dulbecco's Modified Eagle's Medium  FBS  Fetal Bovine Serum  G-l-P  Glucose-1 -Phosphate  G-6-P  Glucose-6-Phosphate  GF  Glycolytic Flux  GP  Glycogen Phosphorylase  GS  Glycogen Synthase  HK  Hexokinase  HS  Horse Serum  INS  Insulin  ISO  Isoproterenol  Lac  Lactate  Oligo  Oligomycin  Pen/Strep  Penicillin and Streptomycin  ZMP  5-aminoimidazole-4-carboxamide ribonucleoside monophosphate  x  ACKNOWLEDGEMENTS I would like to express my appreciation to many special individuals that were involved directly or indirectly in this project. First o f all, I would like to thank my wife, Joyce, for her faithful support, tireless encouragement, and intelligent advices throughout the whole journey. I could not have finished this project without her by my side. I also thank my supervisor D r . M i k e A l l a r d for his helpful guidance and endless patience i n training me as a scientist. I would also like to thank D r . Greg Bondy, Dr. Brian Rodrigues, and D r . Richard Hegele for serving in my research committee and giving me invaluable suggestions and advices along the way. I would also like to thank all present and past members o f the A l l a r d lab, especially H o n Leong, for their help, discussions and insights that contribute not only to the completion this project, but also to the development o f myself as a scientific researcher. I also thank other researchers i n the M c D o n a l d Research Labs that involved in tissue culture work, especially David Chau, for their expert advice and guidance that were fundamental to this project.  Finally, I would like to express gratitude to my parents who have faithfully loved and supported me. Thank you, M o m and Dad.  xi  CHAPTER 1 - INTRODUCTION  Overview Myocardial ischemia, which occurs in patients with coronary artery disease and during cardiac surgery, is very common in the developed world and is increasing in incidence in the developing world. One approach to lessen dysfunction in the setting of myocardial ischemia is metabolic intervention. Such an approach modulates the use of substrates by the heart, typically shifting use from fatty acids towards carbohydrates, to improve cardiac function after the onset of ischemia-reperfusion. Glycogen, the storage form of glucose in the heart, has a significant role in energy production.  Glycogen is a  significant contributor to energy production during aerobic and anaerobic respiration (49, 66, 183, 196), and is especially important to the heart in response to metabolic stress such as ischemia (189), increased workload (55, 157), or during recovery from ischemiareperfusion injury (3, 47-49).  Modulation of its metabolism may, therefore, be an  approach to decrease the detrimental effects of ischemia and reperfusion on the heart. Glycogen metabolism is controlled by two major enzymes, glycogen synthase and glycogen phosphorylase. Glycogen synthase incorporates individual glucose molecules into existing glycogen and, therefore, regulates the synthesis of glycogen.  The  degradation of glycogen is governed by glycogen phosphorylase, which controls the release of glucose molecules from glycogen. These two enzymes are the primary targets for signaling pathways involved in maintaining the homeostasis of glycogen metabolism in the cell. Ischemic stress and increases in cardiac work activate the 5'-AMP activated protein kinase (AMPK) signaling cascade (32, 72, 93, 118). Referred to as the "master  1  metabolic switch", A M P K modulates glucose and fatty acid metabolism in cardiac and skeletal muscle as well as lipogenesis and cholesterol synthesis in other tissues (41, 51, 118, 146).  AMPK's capability to influence cardiac energy metabolism may be an  important determinant of heart function in the setting of metabolic stress (147). Several key enzymes in the regulation of glycogen metabolism are reportedly targets of A M P K (10, 62, 135, 199, 204), but the role of A M P K in the control of cardiac glycogen metabolism is not yet fully understood. Because of the potential role of glycogen metabolism in determining heart function during metabolic stress, a full understanding of the relationship between A M P K and cardiac glycogen metabolism is important, at least in part a consequence of the potential to identify novel targets for therapy of heart dysfunction during ischemia and reperfusion.  The role of A M P K in control of glycogen metabolism has been studied  using isolated heart perfusion models (50, 51, 73, 93, 111, 112, 146).  However, a  cellular-based model is more useful to investigate signaling mechanisms that control substrate metabolism. Investigation of the signaling pathways is more readily achieved in a cellular-based model because manipulations of pathways can be performed in cell systems without concerns that such manipulations may detrimentally alter heart function. I set out to establish methods to measure glucose and glycogen metabolism in isolated cells and to investigate glycogen metabolism and its control by A M P K in cells derived from the heart using selected pharmacological agents. M y overall working hypothesis is that A M P K plays a significant role in the control of glycogen metabolism in heart muscle cells.  2  Ischemic Heart Disease Ischemic heart disease describes a condition that arises from insufficient supply of energy substrates and oxygen to the heart and troubles millions of North Americans annually (108). Ischemic heart disease (IHD) may manifest clinically as angina, sudden death, chronic heart failure or myocardial infarction (38, 100). Ischemic heart disease is closely related to cardiac energy metabolism.  During  the onset of IHD, energy substrates and oxygen become less available to the myocardial cells. As a result, oxidation of substrates in the mitochondria diminishes and glycolysis becomes a major ATP-producing pathway. A further consequence of impaired oxidation of substrates  is that, H , produced from glycolytically-derived A T P hydrolysis +  accumulates within the cell (40), causing acidosis.  In the presence of reduced oxidation,  lactate is also produced in greater amounts during ischemia.  The lactate content in  coronary circulation is, therefore, sometimes used as an indicator of myocardial ischemia (52). In 1962, the treatment for myocardial ischemia was revolutionized by a study by Sodi-Pollares and colleagues (159).  They successfully  treated acute myocardial  infarction (AMI) patients with a solution consisting of high concentrations of glucose, insulin, and potassium (GIK). This therapy has been tested over the next 30 years all over the world, and has been proven successful in lowering the mortality rate of A M I patients (44). GIK therapy operates by manipulating cardiac energy metabolism through increasing glucose uptake and lowering circulating fatty acid level and its uptake. The success of GIK therapy introduced the concept of metabolic intervention for myocardial ischemia.  3  Since then, much research has been done on the relationship between myocardial ischemia and metabolism.  Metabolic antianginal drugs like perhexiline, trimetazidine,  ranolazine, and etomoxir have been developed to treat IHD (100). Recent research found that alteration of myocardial glycogen metabolism can be beneficial to the recovery of heart function after IHD (174). Glycogen metabolism is very active during ischemia and reperfusion. For example, simultaneous synthesis and degradation of glycogen is known to occur during non-ischemic conditions and during ischemia and during reperfusion (49, 189). Re-synthesis of glycogen promotes the return of normal contractile function in the postischemic heart (57,  171).  Therefore, the prominent role of cardiac energy  metabolism, especially glycogen metabolism, in the functional outcome in IHD calls for a better understanding of mechanisms of its control.  Myocardial Energy Metabolism Overview (Fig. 1.1 - Overview of Myocardial Energy Metabolism) In the human heart, every day a significant amount of A T P must be synthesized in order to keep the heart functioning (170).  This A T P is important for initiating and  continuing actin-myosin interaction during systolic contraction, for pumping C a  2 +  into  the sarcoplasmic reticulum and allowing diastolic relaxation, and for maintaining ion gradients in myocardium. Under normal aerobic conditions, 66% of this A T P is devoted to contractile work and 33% is devoted to ion homeostasis.  4  Fatty A c i d Lactate Fatty acid  Glucose  Lactate  G-6-P  Glycogen  Glycolysis y ADpJ  on  J  ^-Pyruvate—''  Acyl CoA  I  PDH  Glucose R-Oxidation  cetyl Co  Oxidation  Mitochondria  Fig. 1.1 - Overview of Myocardial Energy Metabolism.  T r a n s o o r t  Contractile Work  Under usual circumstances, the intracellular A T P content is held constant because the rate of A T P hydrolysis closely matches with the rate of A T P resynthesis.  The  constant turnover of A T P in the myocardium is dependent on the coronary circulation to sufficiently supply oxygen and fuels to the myocardium and to match the energy demand of the heart. Depending on that and the availabilities of different substrates, the heart consumes a number of different energy substrates. These substrates include fatty acids, glucose, lactate, amino acids, glycogen, triglycerides and ketone bodies (66, 133, 136, 150, 151).  This diversity of substrates used has led some to describe the heart as a  metabolic omnivore (169). The heart derives its energy predominately from fatty acids, glucose and lactate. Fatty acids are the principal energy source, under aerobic conditions, supplying 60-90% of the total A T P (6, 150), while pyruvate, derived from glucose and lactate, accounts for the remaining 10-40%. Fatty acids are readily extracted and metabolized by the heart (113, 197). This substrate generates more acetyl-CoA per mole and, therefore, generates more A T P per mole than glucose or lactate.  However, glucose and lactate have an  advantage in a more efficient use of oxygen in pyruvate oxidation such that 11% more A T P is generated per atomic oxygen than that during fatty acid oxidation (161). Since supply of oxygen is critical to oxidative metabolism of fatty acids and carbohydrates, myocardial oxygen supply is a critical determinant of the pattern of myocardial substrate utilization under such conditions.  6  Fatty Acid Metabolism Free fatty acids are hydrophobic molecules and are carried in the circulation system bound to albumin or incorporated in lipoproteins.  The free fatty acid  concentration in the plasma and the fatty acid transporter proteins (1, 179) determine the rate of free fatty acid uptake. After entering into the cell, short-chain and medium chain fatty acids freely diffuse across the mitochondrial membrane without specific regulation mechanism. Conversely, long-chain fatty acids, such as oleate and palmitate, which are the major fatty acids used by the heart, are first converted into fatty acyl C o A by acyl CoA synthase on the outer mitochondrial membrane. The activated long-chain fatty acyl CoA is then transported into the mitochondrial matrix through a complex series of exchanges mediated by the carnitine palmitoyl transferase complex, comprised of carnitine palmitoyl transferase I (CPT-I), carnitine palmitoyl transferase II (CPT-II) and translocase.  Inside the mitochondria matrix, long-chain fatty acyl CoA undergoes P-  oxidation. The fatty acyl CoA is completely oxidized in repeated cycles, and generates one N A D H and one FADH2 and one acetyl CoA for each cycle of the 6-oxidation spiral. The acetyl CoA then enters the tricarboxylic acid (TCA) cycle where it is oxidized with production of N A D H . The F A D H and N A D H then enter the electron transport chain and are utilized to produce A T P .  A palmitate molecule with an 18-carbon backbone  generates a total of 129 ATP.  CPT-I is the key to mitochondrial uptake of long-chain fatty acids (113). The activity of CPT-I is inhibited by malonyl-CoA, which is converted from acetyl CoA by acetyl CoA carboxylase (ACC) (149). A C C activity is controlled allosterically by acetylCoA and covalently by protein kinase A (PKA) and 5'AMP-activated protein kinase  7  (AMPK) phosphorylation (19, 51, 93, 111). Such phosphorylation inhibits A C C activity. Malonyl-CoA decarboxylase (MCD), which may be activated by A M P K , is also involved in controlling the activity of A C C by influencing the level of malonyl-CoA (71, 152). Two isoforms of CPT-1 (muscle and liver) are expressed in the heart.  The muscle  isoform predominates and is 10 to 50 times more sensitive to malonyl-CoA inhibition (31, 92,120).  Carbohydrate Metabolism Overview Carbohydrates, including glucose and lactate, are also major energy contributors of the heart. As with fatty acid metabolism, glucose utilization is controlled at many steps. After entering the cell, glucose may be routed into forming glycogen, the storage form of glucose. For the glucose destined to be catabolized, glucose enters the glycolytic pathway and is converted into pyruvate.  The pyruvate is then transported inside the  mitochondria and oxidized for energy production.  Glucose Uptake The uptake of glucose into the myocardium is determined by the transmembrane glucose gradient as well as the activity and the number of glucose transporters present on the plasma membrane. Two kinds of glucose transporters, GLUT-1 and GLUT-4, are present in the heart and GLUT-4 is the predominant isoform. Both isoforms are found in the sarcolemmal membrane but the majority of both are found in intracellular microsomal vesicles (192). Under basal conditions, relatively more GLUT-1 is present in the  8  sarcolemma (20). The translocation of glucose transporters in the heart is in a dynamic equilibrium that can be fluctuated by changes in oxygen and substrate supply as well as circulating hormones, such as insulin, and myocardial workload (148, 192, 203).  Upon  stimulation, G L U T - 4 and GLUT-1 are recruited to the sarcolemma from intracellular vesicles (89, 99, 203). A M P K is considered to play an important role in the recruitment of GLUTs during metabolic stress (32, 103). After it is taken up into the cell, glucose is rapidly phosphorylated by hexokinase (HK) and converted to (G-6-P).  glucose-6-phosphate  G-6-P is a branch point in glucose metabolism where glucose can either be  incorporated into glycogen or can enter the glycolytic pathway.  Glycolysis (Fig. 1.2 - Glycolysis) Glycolysis is a major pathway in myocardial energy metabolism receiving most of the incoming G-6-P and converting one molecule of glucose into two pyruvate molecules. Along with pyruvate, glycolysis also generates 4 N A D H and 2 ATP. These A T P are believed to be utilized primarily for control of ionic homeostasis, such as for 9-*-  pumping Ca  back into the sarcoplasmic reticulum after contraction, and are important  for diastolic relaxation (77, 94, 191).  The overall rate of glycolytic flux is orchestrated  according to hormonal stimulation, substrate availability and other conditions like metabolic or cellular stress (81).  9  Glucose-6-phosphate ti Fructose-6-phosphate  ATP —Hfi^l^—• ADP Fructose-1,6-bisphosphate  n Dihydroxyacetone phosphate «  • Glyceraldehyde 3-phosphate NAD + P,  J-f  • NADH  1,3-Bisphosphoglycerate ADP—J-f  >ATP  3-Phosphoglycerate It 2-Phosphoglycerate H0 2  «  11 Phosphoenolpyruvate ADP—|  Pyruvate Fig. 1.2  - Glycolysis.  • ATP  Glycolysis is controlled at several enzymatic steps. Among the enzymes in the glycolytic pathway, phosphofructo-1-kinase (PFK-1) has a critical role in controlling the flux of glycolysis. PFK-1 is responsible for phosphorylating fructose-6-phosphate (F-6P) into fructose-1,6-biphosphate.  Products and by-products of glycolysis like A T P ,  citrate and H inhibit the activity of this enzyme. Its activity is stimulated by increases in +  metabolites  like A M P , A D P , C a  2 +  as well as fructose-2,6-biphosphate,  the most  important allosteric activator produced by phosphofructo-2-kinase (PFK-2) from F-6-P. PFK-2 is activated by phosphorylation by A M P K (118). The end product of glycolysis is pyruvate, which is another branch point in myocardial energy metabolism. A portion of the pyruvate produced is converted into lactate by lactate dehydrogenase to be exported out of the cell with the remainder being oxidized in the mitochondria.  Lactate Metabolism In  addition to being produced, lactate is simultaneously utilized by the  myocardium (196).  Lactate enters the myocardium through a transporter, called  monocarboxylate transporter-1 (MCT-1).  Once in the cell, intracellular lactate is  converted to pyruvate by lactate dehydrogenase (LDH), a reaction that is freely reversible. Lactate is suggested to be a major source of pyruvate and reportedly accounts for up to 50% of the pyruvate oxidized in the heart (53, 160). During vigorous exercise when the blood lactate level elevates, contribution of lactate towards pyruvate may be even higher than that of glycolysis (160). Yet, lactate is less important as a substrate when its concentration is low or when free fatty acid levels are high.  11  Pyruvate Metabolism Pyruvate decarboxylation occurs  after transport across  the mitochondrial  membrane and is the critical regulatory step in carbohydrate oxidation.  Pyruvate is  converted into acetyl-CoA, with production of N A D H and CO2, by the pyruvate dehydrogenase (PDH) enzyme complex on the inner mitochondrial membrane. P D H is controlled both allosterically and covalently: the activity of P D H is inhibited by phosphorylation mediated by P D H kinase and is stimulated by dephosphorylation by PDH phosphatase (83, 142, 193, 194).  The relative concentrations of the reaction  substrates and also products are a key determinant to the activity of PDH.  Excess  products like acetyl-CoA and N A D H competitively inhibit P D H and simultaneously activate PDH kinase (63, 142), whereas M g and stimulate P D H activity.  2 +  and C a  2 +  activate P D H phosphatase (119)  Pyruvate oxidation is also controlled by fatty acid  metabolism such that increased fatty acid oxidation inhibits the activity of PDH. In the same token, PDH activity is stimulated by depressed fatty acid oxidation (68, 84, 142). After pyruvate is decarboxylated, acetyl-CoA enters the T C A cycle.  Ultimately,  glycolysis and glucose oxidation as a whole yield 38 ATP, 3 CO2 and 3 H2O for each glucose molecule.  12  Glycogen Glycogen Phosphorylase  ATP  UDP-Glucose  ADP<-i  Glucose  •G-6-P  Fig. 1.3 - Overview of Myocardial Glycogen Metabolism.  Glycolysis  * Pyruvate  Myocardial Glycogen Metabolism Overview (Fig. 1.3 - Overview of Myocardial Glycogen Metabolism) Glycogen acts as a depot of energy in the form of stored carbohydrate. According to the model proposed by Melendez et al. (123, 124), glycogen is spherical in shape and the glucose chains are organized in a concentric tiers fashion linked by cc(l-^4) glycosidic bonds.  Each glycogen granule contains not only carbohydrate, but also  enzymes and proteins involved in its metabolism. These proteins are dynamic in a sense that they associate and disassociate depending on the energy status of the cell.  In  addition, the structure of glycogen is optimized for: storing the most amount of glucose in the least amount of space, releasing the most glucose in the shortest amount of time, and maximizing the efficiencies of enzymes involved in the release of glucose. Therefore, each glycogen granule can be recognized as a functional unit in energy metabolism.  Contribution to Energy Production Besides being a source of energy during times of metabolic stress, glycogen is now recognized as a significant contributor to energy production in aerobic condition (66, 196).  In addition, glycogen is preferentially oxidized compared to exogenous glucose  and its utilization is especially important when heart work is acutely increased (58, 66). This property of glycogen is important in protecting the heart from acute lack of energy substrate in situations of sudden increase in energy demand. During the transition to a higher workload, glycogen contributes the majority of energy needed within milliseconds (58, 157). However, it is important to recognize that tissue-specific patterns of glycogen  14  metabolism exist. Therefore, it is not possible to extrapolate findings in one tissue to another (e.g., skeletal muscle to heart muscle).  Glycogen Turnover The glycogen metabolic pathway branches off the glycolytic pathway at the level of G-6-P. Starting at the conversion of G-6-P to G-l-P, glycogen synthesis continues as the G-l-P molecule is then activated with uridine triphosphate (UTP) and becomes UTPglucose, a process that effectively uses 1 A T P per molecule of glucose activated. The molecule is then incorporated into glycogen by glycogen synthase (GS). GS attaches the activated glucose residues to the distal end of the glycogen chain with a-1,4 glycosidic bonds. Another enzyme, branching enzyme, creates new branches on existing chains via a-1,6 glycosidic bonds and transfers 7 glucosyl units onto the new branch. Each chain has 10 to 18 glucosyl units and has two branches.  The process continues until the  t  glycogen molecule matures with 12 tiers of chains (123, 124). The glycogen degradation process is controlled by glycogen phosphorylase (GP) and debranching enzyme, which are located in the outer tiers of glycogen molecule. GP cleaves up to four a-1,4 glycosidic bonds from the branch point in the glycogen chain and releases G - l - P molecules. Debranching enzyme follows by transferring the three of the last four glucosyl units to the end of another chain.  The enzyme then returns to  cleave the remaining glucosyl unit with the a-1,6 glucosyl bond and release that residue as glucose (156). The G - l - P molecule is then converted into G-6-P and subsequently enters the glycolytic pathway (123, 124).  15  Traditionally, the dogma for the control of glycogen synthesis and degradation was that the two processes were reciprocally regulated, meaning that the two processes could not take place at the same time.  However, this dogma is challenged by the  discovery that myocardial glycogen constantly undergoes synthesis and degradation despite presence of a steady supply of substrates (5, 49, 56, 66). turnover" is used to describe this phenomenon.  The term "glycogen  Since then, various studies have  demonstrated that the turnover of glycogen takes place during aerobic non-stressful conditions (49, 66), and during ischemic conditions (48,49).  Control of Glycogen Turnover (Fig. 1.4 - Regulation of Myocardial Glycogen Metabolism) GS is the key enzyme responsible for control of the synthesis of glycogen and its activity is under complex regulation. modification.  GS is controlled allosterically and by covalent  Through multiple-site (up to 10) and hierarchical phosphorylation, the  activity of GS can be intricately modulated. GS is most active in its dephosphorylated state and does not depend on its substrate, G-6-P, for activity. GS becomes least active when phosphorylated and is more dependent on G-6-P (156).  A number of kinases  phosphorylate GS including glycogen synthase kinases like glycogen synthase kinase-3 (GSK-3), protein kinase A (PKA), calcium/calmodulin kinase II, protein kinase C (PKC), and phosphorylase kinase (PhK). A M P K is also believed to phosphorylate and reduce GS activity (199). Dephosphorylation is carried out mainly by phosphoprotein phosphatase1  (PP1),  which when  activated  by  insulin, stimulates  glycogen  synthesis by  dephosphorylating many key enzymes in glycogen metabolism: GS, PhK and glycogen  16  phosphorylase (GP).  PP1, in turn, is controlled and inhibited by phosphoprotein  phosphatase inhibitor 1 (inhibitor-1). In addition to enzymes, the glycogen content itself also regulates glycogen synthesis by affecting the localization and the activity of glycogen synthase. (134) Similar to GS, GP is also controlled by substrate and by allosteric and covalent modification.  Allosterically, glycogen phosphorylase is readily activated by A M P and  inhibited by A T P and G-6-P. Phosphorylation by phosphorylase kinase (PhK), instead of inactivating in the case of GS, activates GP. Although GP could initiate glycogenolysis in both forms, the phosphorylated form has much higher activity compared to the nonphosphorylated form.  Phosphorylase kinase is activated by the binding of C a  2 +  to the  calmodulin subunit that regulates the enzyme activity and by phosphorylation through phosphorylase kinase A (PKA). Some investigators propose that A M P K phosphorylates PhK (126, 162), although this is not supported by others (17). A complex balance among all these factors modulates the extent of the synthesis and degradation of glycogen,  p-adrenergic receptor stimulation triggers adenylate  cyclase and causes cAMP levels to increase.  cAMP then activates protein kinase A  (PKA) that phosphorylates and activates PhK.  Activated PhK stimulates glycogen  phosphorylase and, subsequently, the degradation of glycogen.  At the same time,  glycogen synthase is inhibited by PKA. The combination of decreased glycogen content and increased glucose uptake may favour glycogen synthesis. Increase in metabolites including A M P , Pi and C a  2 +  also stimulates the degradation of cardiac glycogen.  17  Glycogen  lnhibitor-1  lnhibitor-1  i  PP1  GSd  ^[GPb GSK 3P CaMPK PKA PKC AMPK  ^  G-1-P<  ^  PKA AMPK  lnhibitor-1  Glucose  • G-6-P  Glycolysis  Pyruvate  Fig. 1.4 - Regulation of Myocardial Glycogen Metabolism. GSi - activated glycogen synthase, G S d - inactivated glycogen synthase; G P a - activated glycogen phosphorylase, GPb - inactivated glycogen phosphorylase; PhKa activated phosphorylase kinase, PhKb - inactivated phosphorylase kinase, G S K - glycogen syntase kinase, PKA protein kinase A, P K C - protein kinase C, C a M P K - Ca /calmodulin dependent protein kinase, PP1 - protein phosphatase-1 2+  Insulin impacts upon glycogen metabolism at the level of glycogen synthase as well as at the level of glucose metabolism.  As glycogen content decreases, glycogen  synthesis increases. Insulin initiates its signal through insulin receptor substrate 1 (IRS1), and activates the phosphatidylinositol 3-kinase (PI3K) pathway and several different signal systems.  The PI3K signal is responsible for the translocation of glucose  transporters GLUT-1 and GLUT-4 from intracellular vesicles onto plasma membrane and stimulation of the transcription of the transporter proteins.  Increased G-6-P resulting  from elevated levels of G L U T in the sarcolemma stimulates GS allosterically. Insulin also stimulates PP1, which in turn activates glycogen synthase and inhibits glycogen phosphorylase and phosphorylase kinase.  As a whole, insulin stimulates glycogen  synthesis. A M P K may also influence glycogen turnover with phosphorylation induced effects that favour reduced synthesis of glycogen and increased degradation, but increase glucose uptake favouring synthesis. Its exact role remains to be determined.  Glycogen Metabolism in Myocardial Ischemia & Reperfusion The duration and severity of ischemia have a decisive role in the changes in myocardial energy metabolism (61, 105, 136). In the case of mild ischemia with only a modest reduction of 40-60% in blood flow, myocardial oxygen consumption decreases by 10-50%. Glycolysis is stimulated along with accelerated glycogen degradation (121) and lactate production (104). Fatty acid oxidation is reduced (104, 106, 107) and contractile function could be severely impaired.  Severe ischemia with very low coronary flow  inhibits glycolysis and rapidly depletes A T P and creatine phosphate.  Compounded by  19  glycogen depletion and lactate accumulation, the heart slowly develops tissue necrosis and myocardial infarction. As mentioned above, the heart oxidizes fatty acid in preference to glucose. Fatty acid oxidation's higher oxygen consumption rate and its competitive inhibition of glucose oxidation (143) become problematic during ischemia. Ischemia induces systemic release of catecholamines and stimulates lipolysis (165).  The increase in plasma fatty acid  concentration stimulates fatty acid uptake and fatty acid oxidation. As a result, reciprocally regulated glucose metabolism is suppressed: uptake of glucose and lactate (18, 52, 96) as well as pyruvate oxidation are inhibited (115, 179).  At the same time,  glycolysis becomes a major source of ATP. The excess H ", produced by hydrolysis of 1  glycolytic A T P , cannot be consumed adequately because oxidative metabolism is reduced. The accumulation of lactate and H* gives rise to acidosis and further disrupts the function of the heart (114). The severity of ischemic heart disease is elevated by abnormally high rates of fatty acid oxidation and low rates of glucose oxidation (79, 132). However, many studies have also shown that the increased glycolysis and abundant intracellular glycogen are protective to ischemic-reperfusion injury (25, 39, 147, 183). Similar situations happened on the investigation of the role of myocardial glycogen on the recovery of cardiac function post-ischemia: while some studies indicates the beneficial role of glycogen (49, 57, 97), some studies suggest that insulin, rather than glycogen, may have a significant role in improving the post-ischemic recovery (11, 36, 37, 158, 200). Although their contributions were not completely understood, glycolysis, glucose metabolism and glycogen definitely play a decisive role in cardiac ischemia.  20  5'AMP-activated Protein Kinase ( A M P K ) Overview 5'-AMP activated protein kinase (AMPK), discovered in 1973 (13, 26), is described as "a 'fuel gauge' or 'low fuel warning system', being switched on by depletion of ATP. Once activated, it initiates energy-saving measures, and switches on reserve ATP-generating systems" (64). A M P K is a heterotrimeric complex comprised of three subunits designated a, p, and y.  The a subunit is 63 kDa in mass and is the  catalytic subunit containing a kinase domain in the N-terminal and a regulatory domain in the center (33). The 38 kDa P subunit is a scaffolding unit connecting a and y subunits (173). The function of the y subunit has not been determined, but it is speculated that the subunit involves in the binding of A M P and the activation of the enzyme (30).  In  mammals, the three subunits exist in multiple isoforms ( a l , a2, p i , P 2 , y l , y2, y3) and have multiple heterotrimeric combinations with varying tissue distributions (30, 163, 173). These combinations appear to influence the extent of allosteric activation by A M P (30).  Control of A M P K activity The ratio of AMP:ATP is an excellent indicator of the internal metabolic energy state.  From a short-term perspective, the adenosine moiety available for adenine  nucleotides, A M P , ADP and A T P is fixed and the total quantity of A M P , A D P and ATP is relatively constant.  Equilibrium of the three derivatives of the energy phosphates is  maintained by adenylate kinase, and a high level of A T P at over 80% of the total adenine nucleotide pool is maintained. The concentration of ATP drops only slightly even in  21  situations of increased heart work. The slight fall in A T P level is accompanied by a sharp rise in A M P level through an amplifying effect of adenylate kinase (45). A M P modulates the activity of A M P K in several different ways: A M P binding to A M P K allosterically activates the enzyme and exposes the Thr-172 site on the catalytic subunit for phosphorylation by A M P K kinase (AMPKK) the upstream kinase, which is also activated by A M P (35). The activation of A M P K by phosphorylating the threonine172 residue of the a subunit induces a much stronger activation than allosteric activation (164).  In addition to directly stimulating A M P K activity, A M P also inhibits the  dephosphorylation and inactivation of A M P K by protein phosphatase 2A and 2C (35). Coupled with the multiple effects of A M P on the A M P K cascade and a very low K of m  A M P K K for A M P K , the A M P K signaling system is extremely sensitive and is capable of eliciting a large response to changes in A T P concentration (65). Contrasting the stimulating effect of A M P and phosphorylation, A T P inhibits the activation of A M P K (50).  Besides intracellular metabolites or energy substrate levels,  stress responses like ischemia (93, 118) activate A M P K by increasing the AMP:ATP ratio and decreasing the PCnCr ratio.  Endocrine signals like insulin (51, 198) also  control A M P K activity by kinase signaling pathways.  Interestingly, the two signal  pathways interact and regulate each other: ischemia induces intracellular acidosis and inhibits insulin signaling, while insulin pretreatment inhibits A M P K activation by ischemia (12, 51, 72).  A M P K can also be activated pharmacologically with an agent  called 5-Aminoimidazole-4-carboxamide 1-8-D-ribofuranoside (AICAR).  The agent is  cell-permeable and is phosphorylated to 5-aminoimidazole-4-carboxamide ribonucleotide (ZMP), a 5'-AMP mimicking compound, after being taken up by the cell.  The  22  accumulation of Z M P activates the A M P K pathway through allosteric mechanisms and by phosphorylation.  A M P K Modulation of Lipid Metabolism in the Heart Lipid metabolism, particularly long-chain fatty acid metabolism, in the heart is controlled by a central factor, malonyl-CoA, that inhibits the mitochondrial oxidation of fatty acid by controlling entry of long chain fatty acids into mitochondria. Synthesis of malonyl-CoA is regulated by acetyl-CoA carboxylase (ACC) and its degradation is controlled  by  malonyl-CoA  decarboxylase  (MCD).  When  activated,  AMPK  phosphorylates and inactivates acetyl-CoA carboxylase (ACC) (51, 71, 93). At the same time, A M P K is also suggested to activate M C D (71). malonyl-CoA is dramatically decreased.  As a result, the concentration of  Thus, the rate of fatty acid oxidation is  accelerated upon the activation of A M P K because lower levels of malonyl C o A relieve inhibition of CPT-1.  A M P K Modulation of Glucose Uptake & Glycolysis in the Heart A M P K activation in the heart stimulates GLUT-4 translocation and increases glucose uptake. The stimulation involves a pathway independent from the insulin-PI3K signaling pathway and is important for ischemia-stimulated glucose transport (146). Muscle contraction also stimulates the translocation of GLUT-4 by increasing A M P K activity (205).  A M P K may stimulate glycolysis in the heart by activation of 6-  phosphofructo-2-kinase  (PFK-2) via phosphorylation of Ser-466  (118).  During  myocardial ischemia, the surge in A M P concentration stimulates PFK-1 and activates  23  AMPK.  The increase in A M P K activity activates glucose transporters and PFK-2.  Therefore, the rate of glycolysis is stimulated in metabolically-stressed hearts by multiple mechanisms.  AMPK Regulation of Glycogen Metabolism in the Heart Myocardial glycogen metabolism is closely glycolysis.  Interestingly, AICAR  connected to the myocardial  can also activate glycogenolysis via allosteric  mechanisms (112). A M P K stimulation of GLUT-4 translocation (146) directly increases the concentration of intracellular G-6-P.  The elevated  concentration of G-6-P  allosterically activates glycogen synthase and stimulates synthesis of glycogen. may  AMPK  also activate phosphorylase kinase (126), thereby activating GP. Glycogen  phosphorylase is also activated by ZMP via an allosteric mechanism (112). The gamma subunit of A M P K is responsible for the binding of A M P (30). Genetic mutations in the gamma-2 regulatory subunit (PRKAG2)  of A M P K  lead to abnormal glycogen  accumulation in the heart (54). A similar study in skeletal muscle found that gamma-3 (PRKAG3) is also important in the control of glycogen metabolism (127). Therefore, the evidence suggests that A M P K activity influences myocardial glycogen metabolism. Although the A M P K regulation of glycogen metabolism has been extensively studied in the skeletal muscle and other tissues (10, 54, 62, 70, 127, 135, 199, 204), the direct relationship between A M P K and glycogen metabolism in the heart has yet to be determined.  Since the activity and the regulation of A M P K is not the same across  different tissue types that express the enzyme (76), further understanding of the A M P K regulation on myocardial glycogen metabolism is important for the development of  24  metabolic therapies towards myocardial ischemia and reperfusion. A variety of approaches, ranging from in vitro and in vivo, and from intact organ to isolated cell are available, each of which has strengths and weaknesses. The particular usefulness of these approaches depends largely on the nature of the question asked.  Approaches to Investigate Cardiac Glycogen Metabolism Whole Organism or in vivo Studies In studies of glycogen in skeletal muscles, obtaining tissue samples with a biopsy needle is the most common approach. While this method is also used in some cases during open-heart surgery (183), obtaining biopsy sample from the human or animal heart in the research of cardiac glycogen is not commonly performed. In most cases, an animal model is used.  Use of inhibitors may be detrimental to heart function, and  systemic changes in hormones and metabolites are not easily controlled.  Isolated Heart Model Since 1970, isolated heart perfusion has had a critical role in studying the metabolism of heart. This model produces, with relative ease, highly reproducible data that has significant clinical relevance.  The isolation of the heart also eliminates  interactions of the heart with other organs, and confounding effects of the systemic circulation and hormones or neuronal factors. In addition, the system allows complete control over the concentration of nutrients and hormones or pharmacological agents to which the tissue is exposed.  Therefore, the isolated heart perfusion model is an  25  outstanding model to study metabolic properties and the effect of pharmacologic modulators. Many excellent studies have helped characterize the energy metabolic properties of the heart in areas like glucose transport, glycolysis, glycogen metabolism, glucose oxidation, and fatty acid oxidation. Moreover, the isolated heart preparation really excels in modeling cardiac ischemia-reperfusion injury.  Many key studies of ischemia-  reperfusion and its pharmacological interventions utilize the isolated perfused heart model (3, 4, 29, 57, 80, 87, 92, 97, 114, 132, 151, 153, 168, 177, 188). The strength of the isolated heart perfusion is in the retention of the functional integrity of the tissue. However, this strength is also the major downfall of the model. Since the viability and function of the heart has to be maintained, limitations exist on the type and dose of pharmacological agents as well as the severity of physiologic or pathologic conditions that can be applied to the heart.  Isolated Cell Models Another popular model used in the investigation of energy metabolism is the use of isolated cell culture models. Rather than an intact heart, cardiomyocytes are isolated and cultured as monolayers.  Heart cells are acquired by perfusing the heart with an  oxygenated solution containing proteolytic enzymes (15). These enzymes attack the cell junctions, adhesion factors and the extracellular matrix. Gradually, tissue integrity is compromised and free cells are released and, subsequently, harvested.  Once isolated,  these cardiomyocytes, known as primary cultures, can be used immediately. Another option is to culture these cells for generations in order to establish a lineage of cells. The  26  established line of cells is usually differentiated from the primary cultures and may have a different morphology. Most differentiated cells have a finite life span that is suitable for experiments. In general, the life span of differentiated animal cells is about twenty to forty passages or generations. The isolated cell model system has a number of advantages, and the most significant one is that cells are homogeneous.  Primary cultures of cardiomyocytes  consist of only the myocytes and none of the connective tissue or endothelial cells found in the heart. Thus, metabolic data collected using primary cultures will be more specific and accurate in capturing the response of cardiac myocytes in the myocardium. Compared to the isolated heart model, the isolated cell model allows greater flexibility on the experimental conditions or parameters since the factor of functional viability in an intact heart is not an issue.  Since the cells are more accessible in the cultured form,  cellular-molecular data like phosphorylation patterns, signaling pathway mechanisms as well as enzyme activities can be studied with greater ease. metabolism  have  been  conducted  using  isolated  cells.  Many studies of energy Primary  cultures  of  cardiomyocytes (21-24, 74, 75, 110, 117, 128, 154, 207) and established cell lines like H9c2 (2, 131, 139, 166, 195, 205, 206) have played a prominent role in furthering the understanding of glycogen metabolism in heart muscle cells. A n established cell line, if available, has an advantage over primary culture in that it is more consistent in its metabolic properties and in the quality of cells at a batch-to batch level. Several cell lines derived from heart tissue are available for use. To my knowledge, none has been used to study energy metabolism, specifically glycogen and glucose metabolism and its control.  27  Besides all the benefits of the isolated cell model, there are certain aspects of the model that need to be considered. The cells are grown and differentiated in culture, and many of their properties are changed relative to those in the intact organ.  Since the  isolated cells are not doing any work like those of a beating heart, the metabolic rate of the isolated cardiomyocytes is much lower than a normal cardiac myocyte in a beating intact heart. Additionally, the amount of cellular material is small; so efficient separating techniques and sensitive analytical techniques are needed.  On balance, however, I felt  isolated cells would provide a more readily and easily useful model system to begin to characterize signaling pathways that control glycogen and glucose metabolism.  28  Hypothesis and Objectives Hypothesis: Activation of A M P K influences glycogen metabolism in cardiomyocytes by decreasing glycogen synthesis and increasing glycogen degradation.  Objectives: 1. To develop and to establish methods to measure glucose  and glycogen  metabolism in isolated muscle cells. 2. To characterize glucose metabolism and glycogen turnover in H9c2 cells, a heartderived cell line. 3. To measure glycogen turnover and its control by A M P K in H9c2 using selected pharmacologic agents.  General Experimental Approach L6, compared to H9c2, is a more established cell line in terms of metabolic properties. Therefore, protocols were first established in L6 cells.  Experimental data  collected was compared against published data on L6 to verify the correctness and effectiveness of the protocols. Subsequent studies characterizing glycogen metabolism and its control were then performed in H9c2 cells using these methods.  29  CHAPTER 2 - MATERIALS AND METHODS Cell Culture Conditions General L6 (L6.G8.C5) cells, item number CRL-1458, lot number 1334985, and H9c2 (H9c2[2-1]) cells, item number CRL-1446, lot number 2056335, were acquired from American Type Culture Collection (ATCC) (Manassas, V A ) . These cell culture stocks were cultured and were designated as passage number 1.  Cell cultures of different  passage were routinely set aside and kept frozen in liquid nitrogen until used. L6 and H9c2 cell cultures used in experiments were routinely cultured for at least 5-9 passages and no passages of 13 or beyond were used. Cell cultures were maintained in a waterjacketed cell culture incubator (Model 3326, Forma Scientific, Marietta, OH) under a humidity-controlled atmosphere comprised of 95% air and 5% CO2 at 37°C. A l l culture vessels used, including 25cm and 75cm flasks, were obtained from Sarstedt (Quebec, Canada). The media used in cell cultures were based on Dulbecco's Modified Eagle's Medium (DMEM) from Invitrogen (Carlsbad, CA) containing 5.5 m M (1000 mg/L) Dglucose, 4 m M L-glutamine, 4 mg/L pyridoxine hydrochloride, and 110 mg/L sodium pyruvate. Sodium bicarbonate (2.2 g/L; Sigma, St. Louis, MO) was added to the D M E M for pH balancing as indicated by the supplier.  Antibiotics, including 100 IU/mL  penicillin and 100 IU/mL streptomycin (Invitrogen), were present in the media at all times.  30  L6 Cell Culture Reconstituted frozen stocks of L6 were cultured in 75 cm culture flasks and the culture media was supplemented with 10% fetal bovine serum (FBS) (Invitrogen) (vol/vol). Cell cultures were subcultured by diluting the original in 1:2 to 1:4 ratios once they reached 80% confluency. When the cells reached passage 9, they were subcultured into 25 cm culture flasks with a seeding density of -7500 cells/cm . D M E M media with 2  2  10% FBS were used and the media were changed approximately every 48 hours. When the cultures reached 100% confluency, the FBS content in the media was reduced from 10% to 2% to initiate myogenic differentiation.  On Day 1 (the day the media is  switched) and Day 2, 10 u M cytosine (3-D-Arabinofuranoside (Ara-C) was added to the culture media to prevent proliferation of non-differentiated mononucleated myoblasts. Starting from Day 3, Ara-C was excluded and new media was provided every 2 days. The L6 cells were further cultured for 6 days in the presence of 2% FBS before being used for experiments. The presence of FBS in the media was specific to the experimental sample groups.  H9c2 Cell Culture Frozen stocks of H9c2 were reconstituted in 75 cm culture flasks and the D M E M 2  media was supplemented with 10% FBS and O.lmM L-Carnitine (Sigma). H9c2 cultures were subcultured by diluting the original 2 to 4 times once they reached 80% confluency. When the cells reached passage 9, they were subcultured into 25 cm culture flasks. Using the media with 10% FBS and O.lmM L-Carnitine, H9c2 cells were plated with a density of -6500 cells/cm and were cultured until they reached 80% confluency. At this  31  point, H9c2 was differentiated into cardiac-phenotype by switching to D M E M media with 1% horse serum (HS) (Invitrogen) in place of FBS. Media supplemented with retinoic acid (100 nM; Sigma) was used and was changed every day for the first 4 days. Retinoic acid was used in order to preserve the cardiac phenotype of H9c2 cells (43, 125). On the 5  th  day, retinoic acid was omitted from the culture. On the 6  th  day and 24  hours before experiment, the differentiated H9c2 cultures were provided with media comprised of only D M E M without any serum. Experiments with cardiogenic H9c2 were conducted in the same serum-free D M E M media. To induce myogenic differentiation in H9c2 cells, retinoic acid was omitted at the cellular differentiation step. Culture media with 1% (vol/vol) HS were added to the cells every 2 days for another 6 days. On the 6  th  day and 24 hours before experiment, the  differentiated H9c2 cultures were provided with media comprised of only D M E M without any serum.  Experiments with myogenic H9c2 were conducted in the same  serum-free D M E M media.  Assessment of Cell Culture Development Morphological and phenotypic properties of the cells were monitored at the early stage of the project during protocol development. Since metabolic properties of both L6 and H9c2 cells needed to be tested after the cells had undergone differentiation, tests were done to confirm the state of differentiation of the sample cells.  The parameters  chosen were cell viability, cell morphology and creatine kinase activity, the latter being recognized as changing according to stage of differentiation (186).  32  Cell Viability Cell culture was washed 2 times with ice-cold phosphate-buffered saline (PBS) (Invitrogen) and then trypsinized with 1 mL of 0.25% trypsin solution containing 2.5 g/L of trypsin (1:250) in Hanks' Balanced Salt Solution without C a C l , M g C l • 6 H 0 , and 2  2  2  MgSC>4 • 7H2O. (Invitrogen). The majority of this solution was aspirated after a brief incubation of 30 seconds, leaving only a thin film of trypsin solution over the cells. Cultures were further incubated at room temperature for 10 - 15 minutes until they were dissociated from the growing surface. To collect the detached cells, 3 mL of PBS were added to the flask and the cell suspension was mixed thoroughly by pipeting. Mobilized cells were collected into a 1.5 mL centrifuge tubes and were kept on ice.  The  concentration of cell samples was then determined using a hemacytometer and adjusted to about 20-100 cells/mm with PBS, and 200 uL of 0.4% trypan blue stain (Sigma) was 2  added and mixed. After 5 minutes of incubation at room temperature, a 20 u L aliquot of the mixture was counted with a hemacytometer. The number of stained and un-stained cells was recorded. Only the center square and four comer squares of the hemacytometer grid were counted.  A duplicate sample was counted by the same procedure for  verification purpose. Cell viability was expressed as the percentage of non-stained cells divided by the total number of cells including both stained and non-stained cells. This procedure was originally intended to measure the viability of the cells in the characterization of H9c2's metabolic properties. However, this procedure was no longer used in the experiments on AMPK's influence on glycogen metabolism in H9c2 cells.  33  Cell Morphology Images of the cell cultures were obtained to compare morphological changes that occurred during the course of growth and differentiation.  Images of the cell cultures  were acquired with a Coolpix 995 digital camera (Nikon) mounted on an inverted light microscope (model TMS-F, Nikon).  The original images were then processed using  Photoshop 5.0 L E (Adobe, San Jose, CA). Proper color settings and color contrast adjustments were made to enhance the quality of images. The shape, arrangement pattern and fusion of the cells were assessed to allow comparisons of cultures.  Creatine Kinase Activity Creatine kinase (CK) activity was measured as a benchmark of the level of differentiation in the cells: as myoblasts cells differentiate into myocytes cells, the activity of creatine kinase increases (46, 184). A commercial kit (model 47-UV, Sigma Diagnostics, St. Louis, MO) was used to measure C K activity. The method used was based on the procedures suggested by the supplier and according to the following principles.  C K catalyzes the reaction between creatine phosphate and glucose to form  creatine and ATP. Hexokinase (HK) in the assay mixture phosphorylates glucose to glucose-6-phosphate  (G-6-P) utilizing the A T P produced.  Glucose-6-phosphate  dehydrogenase (G6PDH) then oxidizes the G-6-P with nicotinamide adenine dinucleotide (NAD), forming 6-phosphogluconate (6-PG) and N A D H .  The increase in the level of  N A D H is detected by the increase in absorbance at 340 nm. The rate of change in absorbance at 340 nm is directly proportional to C K activity.  34  Cell cultures were first washed in ice-cold PBS and harvested by scraping into 100 u L homogenization buffer which consisted of: 50 m M HEPES, pH 7.4; 1 m M EDTA;  1  mM  EGTA;  protease  inhibitor  mix  containing  4-(2-aminoethyl)  benzenesulfonyl fluoride (AEBSF), pepstatinA, E-64, bestatin, leupeptin, and aprotinin (Sigma) and deionized and distilled water (ddHaO).  The harvested cells were then  transferred into 1.5 mL microcentrifuge tubes. The collected cells were homogenized by sonication for 2 intervals of 5 seconds on ice and were kept on ice until analysis. Assessment of the C K activity was performed at room temperature (20-23°C). Baseline absorbance at 340 nm ( A  340  ) was established by measuring 450 u L of C K reagent in a  cuvet for 2 minutes with a spectrophotometer (Perkin Elmer, Shelton, CT). After that, 50 uL of cell homogenate sample was added to the cuvet and mixed by rapid pipeting. The A34owas  continually monitored at 30-second intervals for a period of 180 seconds. After  the 3-minute interval, another 50 u L of cell homogenate was added to the cuvet. The rate of change in A340 was monitored for another 180 seconds to verify the detection capacity of the diagnostic reagent. The rate of change in A340 is proportional to the activity of C K in the sample, and was used to calculate the C K activity in the cells. The C K activity was normalized to protein content and reported as Units/min/mg protein where one unit 91  equals 1x10"  mole of substrate consumed.  35  Glycogen Metabolism Total Glycogen Content  To measure glycogen content in cell samples, glycogen was degraded into glucose and the resultant concentration of glucose produced was measured. Cell cultures were first washed twice with 3 mL of ice-chilled PBS (Invitrogen). After washing, cells were homogenized by adding 500 uL of 0.4 M K O H and incubated in room temperature for 15 minutes. Cell samples were then transferred to microcentrifuge tubes and were kept at 20°C until analysis. For sample analysis, frozen samples were thawed and were exposed to 95°C for 10 minutes to remove existing exogenous glucose in the samples (82). Then, 200 u L of heat-treated homogenates were spotted onto 2 cm x 2 cm 31ET filter paper (Whatman, Kent, UK). The filter papers were washed in ice-cold 66% ethanol for 3 intervals of 15 minutes and air-dried. After it was completely dried, each filter paper was cut into 8 pieces and placed into 2.0mL-centrifuge tube.  Reaction buffer (500 uL)  comprised of 0.2 M acetate buffer, pH 4.7, and 0.625 mg/mL amyloglucosidase (Sigma) was added to the sample, which was then incubated at 37°C with gentle shaking for 2 hours. A n aliquot of reaction buffer (200 uL) containing glucose released from glycogen by amyloglucosidase was then aliquoted into a standard 1.5 mL cuvet (Sarstedt). Glucose is detected by adding glucose-detecting  reagent (500 uL; Sigma  Diagnostics) to the reaction product above. This reaction took place for 30 minutes at 37°C with gentle shaking and a color reagent product detectable at 475 nm was formed. The absorbance at 475 nm was measured with a spectrophotometer and the data was compared to a glucose standard curve prepared with glucose stock solution (Sigma) to determine the glycogen content of the samples. Glycogen content was expressed as  36  nanomoles of glucose per milligram protein. Glycogen content at the beginning of the 30-minute experimental period was measured. This measurement served as a reference to determine the extent of glycogen degradation at the end of the experimental period.  Glycogen Synthesis Glycogen synthesis was determined by measuring the amount of labeled glucose incorporated into glycogen over the course of experiment (66).  In the experimental  protocol, [5'- H]-glucose (1.0 uCi/mL) was added to the cell cultures to detect synthesis 3  of glycogen.  Cell cultures were homogenized and collected as mentioned in "Total  Glycogen Content" section. To detect the [5'- H]-labeled glycogen, homogenates of cell cultures were prepared, spotted onto filter paper and washed in cold ethanol as described above. The dried filter papers were then placed into 7 mL-scintillation vials and 4 m L of scintillation fluid were added to each vial and vortexed.  Vials were left overnight in  room temperature before being counted in a scintillation counter in order to ensure the scintillation fluids were stabilized after mixing. Average specific activity of glucose, determined by measuring [ H] activity in aliquots of media, was used to determine the 3  quantity of labeled glycogen. Specifically, labeled glycogen was calculated as the quotient of [ H] activity in glycogen and specific activity of glucose in the media. Glycogen synthesized was expressed as nanomoles of glucose incorporated per milligram protein.  The rate of glycogen synthesis was calculated by dividing the measured  glycogen synthesized by thirty to get nanomoles of glucose incorporated per milligram protein per minute.  37  The  rate of glycogen synthesis was determined by measuring the rate of  accumulation of 5'-[ H]-labeled glycogen.  Comparison of different effectors and the  magnitude of change were compared to the rate of glycogen synthesis in the control condition.  Exogenous Glucose Metabolism Glycolysis The rate of glycolysis was determined by measuring the rate of production of 3  H 0 from [5'- H]-glucose (1.0 uCiyrnL) (66). 3  2  3  H 0 was separated from [ H]-glucose by 3  2  use of separation columns composed of potassium tetraborate and Dowex 1-X4 anion exchange resin (200-400 mesh.) Borate (0.4 mM; 122.196 g/L ddH 0) was dissolved in 2  appropriate amount of ddH 0 before adding 90 g/L Dowex anion exchanger and being 2  stirred overnight in room temperature.  On the day of experiment, 2 mL of the  Borate/Dowex mixture was added to each column and rinsed with ddH 0 for 3-4 times. 2  After all ddH 0 was drained, the columns were placed over 7 mL-scintillation vials. 2  Duplicate samples (200 uL) of media were added to each column followed by 2 aliquots of 400 uL ddH 0 for washing purposes. After all sample filtrates were collected into the 2  scintillation vial, scintillation fluid (4 mL) was added to each vial. Vials were vortexed and left overnight in room temperature before being counted in a scintillation counter. Glycolytic rates were calculated from glucose specific activity standard curve and [ H 0] 3  2  in the media with time taken into account. Rates of glycolysis were expressed as nanomoles of glucose catabolized per hour per milligram protein.  38  Glucose Uptake After glucose is taken up into the cell, it is either be incorporated into glycogen or catabolized via glycolysis (196). Therefore, the rate of glucose uptake can be calculated as the sum of the rate of glycogen synthesis and the rate of glycolysis. The values for the two processes, determined individually as indicated in the sections above, were thus added to yield glucose uptake, expressed as nmol glucose taken up / hour / mg protein.  Lactate Content Production of lactate from exogenous glucose and glycogen was detected by measuring the lactate present in culture media sampled when cell cultures were exposed to various experimental conditions. Lactate was measured using a diagnostic kit (No. 735, Sigma Diagnostics). A standard curve of known lactate concentrations ranging from 0 to 0.44 nmol per mg protein was generated using lactate standard solution provided with the kit.  One sigma unit of activity will transfer 1.0 umole of phosphate from  creatine phosphate to A D P per minute at 37 °C. D M E M media samples were diluted 10 times with ddJrkO. Twenty u L of diluted duplicate samples were aliquoted into 96-well plate. Lactate reagent (200 uL) was then pipetted into each well and the mixture was incubated at 37°C for 10 minutes.  The absorbance at 540 nm was measured in a  spectrophotometer (Perkin Elmer, Shelton, Connecticut) after the assay samples within 10 minutes after incubation.  The change in As^was directly proportional to the lactate  content in the sample. Lactate production was expressed as nmol per milligram protein (U/mg protein).  39  Activity of Regulatory Enzymes and Kinases Glycogen Phosphorylase Cell cultures were first washed in ice-cold PBS and harvested by scraping into 1 mL homogenization buffer which consisted of: 0.5 M Tris-HCl, pH 7.8; 0.5 M E D T A , pH 7.0; 0.5 M DTT, 0.5 M NaF; 2.5 g/L glycogen (Type III) and deionized and distilled water (ddH20). The harvested cells were then transferred into 1.5 m L microcentrifuge tubes. The collected cells were homogenized by sonication for 2 intervals of 5 seconds on ice and were kept on ice until analysis. Fifty u L of sample in duplicates were put into 96-well plates and 50 u L of reaction buffer were added to the sample wells. The percentage of active GP was detected by " - A M P reaction buffer", which is comprised of: MES (31 mM), K F (200 mM), B-mercaptoethanol (0.45%), G-l-P (4.6 mg/mL), [ C]-G-1-P (18.6 uL/mL), 14  glycogen (13 mg/mL), and ddHaO. Total GP activity was detected by "+AMP reaction buffer" comprised of: MES (31 mM), K F (200 mM), B-mercaptoethanol (0.45%), G - l - P (60 g/L), [ C]-G-1-P (15 uL/mL), glycogen (53 mg/mL), A M P (3.33 mg/mL) and 14  ddHaO. All chemicals and reagents mentioned above were purchased from Sigma. The mixture of sample and reaction buffer was incubated for 1 hour at 30°C in a shaking incubator (Perkin Elmer, Boston, MA). After the incubation, 80 uL of mixture was spotted on 31ETCHR filter paper and the filter papers were washed in ice-cold 66% ethanol for 3 intervals of 15 minutes and air-dried. The dried filter paper was then put into 7 mL scintillation vials along with 5 mL of scintillation fluid and counted for radioactivity. The counts of vials with blank filter paper were subtracted from the experiment vials. The data was referenced to standards of 20 u L of +/- A M P reaction  40  buffer for specific activity of [ C]-labeled glycogen. The following formula summarized 14  the reaction: Glycogen (n) + [ C]-glucoses-l-phosphate 14  [ C]-glycogen (n+1). The 14  unit for GP activity was ^mol/min/mg protein. The assay for glycogen phosphorylase activity was based on measuring of [ C] 14  incorporation into glycogen  from  [ C]-labeled glucose-1-phosphate. 14  phosphorylase activity was expressed as a percentage  (%a)  of active  Glycogen glycogen  phosphorylase, glycogen phosphorylase a, of total glycogen phosphorylase, glycogen phosphorylase b. A M P stimulates glycogen phosphorylase activity. Therefore, glycogen phosphorylase a was assayed in the absence of A M P while glycogen phosphorylase b was assayed in the presence of 5 m M A M P . The activity of glycogen phosphorylase b did not change among all the experimental groups and was used as a common baseline for comparison.  Glycogen Synthase Cell cultures were first washed in ice-cold PBS and harvested by scraping into 1 mL homogenization buffer which consisted of: MOPS (50 mM); sucrose (250 mM); E D T A (5 mM), NaF (25 mM), and deionized and distilled water (ddF^O). The harvested cells were then transferred into 1.5 mL microcentrifuge tubes. The collected cells were homogenized by sonication for 2 intervals of 5 seconds on ice and were kept on ice until analysis. 25 u L of sample in duplicates were put into 96-well plates and 50 u L of reaction buffer were added to the sample wells.  The percentage of active GS was detected by  "0.25 m M G-6-P reaction buffer", comprised of: MOPS (75 mM), NaF (75 mM),  41  UDP-glucose (1.831 mg/mL), glycogen (10 mg/mL), [ C]-UDP-glucose (22.5 [lUmL), 14  G-6-P (0.25 mM) and ddH 0. Total GS activity was detected by "15 m M G-6-P reaction 2  buffer" comprised of the same content as the reaction buffer mentioned above, except with 15 m M G-6-P. A l l chemicals and reagents mentioned above were purchased from Sigma. The mixture of sample and reaction buffer was incubated for 30 minutes at 30°C in a shaking incubator (Perkin Elmer).  After the incubation, 50 u L of mixture was  spotted on 3 M M filter paper and the filter papers were washed in ice-cold 66% ethanol for 3 intervals of 15 minutes and air-dried. The dried filter paper was then put into 7 mL scintillation vials along with 5 mL of scintillation fluid and counted for radioactivity. Like above, the counts of vials with blank filter paper were subtracted from the experiment vials. The data was referenced to standards of 20 u L of reaction buffer for specific activity of [ C]-labeled glycogen. The reaction that took place was: UDP-[U14  l4  C]-glucose + glucose -> UDP-[ C]- glycogen. 14  The unit for GS activity was  nmol/minute/mg protein. The assay for glycogen synthase activity was based on measuring of [ C] 14  incorporation into glycogen from UDP-[U- C]glucose. Similar to that for glycogen 14  phosphorylase activity, glycogen synthase activity was expressed as a percentage (%I) of active glycogen synthase, I-form, of total glycogen synthase, D-form. As described by Guinovart et al. in 1977 (60), the active I-form of synthase was measured in the presence of 0.25 m M glucose-6-phosphate (G-6-P) while 15 m M G-6-P was present when measuring D-form synthase activity.  42  A M P K and A C C phosphorylation patterns - Western Blotting Since A M P K is controlled by phosphorylation state, its activity was determined by measuring its phosphorylation state. As a downstream target of A M P K , A C C phosphorylation can be used as an indicator of A M P K activity, both phosphorylationdependent  and  phosphorylation-independent  (e.g.,  allosteric)  (112).  Thus,  phosphorylation of both A M P K and A C C was measured to assess A M P K activity in the cultured cells. Cells in each treatment group were scraped on ice in 1 mL of homogenization buffer (pH 7.5) made up of: MOPS (20 mM), sucrose (250 mM), KC1 (150 mM), E D T A (2 mM), benzamidine (2.5 mM), pepstatin A (3 uM), leupeptin (5 uM), glutathione (2.5 mM), PMSF (0.5 mM) and microcystin-LR (0.14 ug/mL).  The cell suspension was  transferred into a 1.5 mL centrifuge tube and was centrifuged for 2 minutes at lOOOg and 5 minutes at 15000g (112). Samples of cell homogenate separated by electrophoresis on 10% SDS-polyacrylamide gels, and transferred by electroblotting to a nitrocellulose membrane. After nonspecific blocking, the blots were probed overnight with primary rabbit  antibodies  against rat phospho-AMPK  (1:1,000 dilution, Cell  Signaling  Technology, Mississauga, Ontario, Canada) or phospho-ACC (1:100 dilution, Upstate Biochemicals, Lake Placid, NY). After incubation with goat 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 total A M P K and A C C .  43  Other Methods Total Protein Content A l l enzyme activities and metabolite concentration are expressed relative to the protein content of the sample. entire series of experiments.  Therefore, protein content detection is essential to the Protein content of cell samples was determined by a  commercial kit (BCA-1, Sigma Diagnostics). A standard curve of bovine serum albumin (BSA; Sigma) ranging from 0 to 20 mg/uL was made with lOx dilution of the buffer used to homogenize the sample. Aliquots of 25 u L of a suitable dilution of the cell samples (5x - lOx) were added to a 96-well plate (BD Biosciences, Lexington, K Y ) in duplicate. This was followed by addition of 150 u L of assay reagent comprised of 98% B C A : 2%  CUSO4. The assay plate was incubated at 37°C for 30 minutes with gentle shaking. The absorbance at 562 nm was measured in a spectrophotometer (Perkin Elmer) after the assay samples were cooled to room temperature.  The change in  A562  is directly  proportional the protein content in the sample and the protein content is expressed in milligrams of protein (mg protein).  Data Analysis All data are expressed as Mean ± standard error of the Mean (SEM) and were analyzed using one way A N O V A , paired t-test, assuming unequal variances.  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.  44  CHAPTER 3 - ESTABLISHING A CELLULAR MODEL OF ENERGY METABOLISM USING THE L 6 MYOGENIC C E L L LINE Background  H9c2 cells have been recognized as a cell line that show features of the cardiomyocyte and is useful for setting up a cellular model of cardiomyocytes (27, 78, 138). However, the metabolic properties of H9c2 cells have not been fully established. Therefore, I felt it is important to use another cell model that has well determined metabolic characteristics to develop the experimental techniques for use with H9c2 cells. The cell model I have chosen is the L6 cell line. In 1968, Dr. D. Yaffe set up an investigation targeted to test the nature of the fusion and differentiation capabilities of myogenic cells. In the process, he developed a new line of cultured cells, called L6 cells, from skeletal muscles of fetal rats by selective plating and serial passage (201). Since that time, L6 cells have been widely used in skeletal muscle research and have become one of the best-characterized myogenic cell lines in terms of its morphological, biochemical, and metabolic properties. Having a spindle shape phenotype, L6 is a myoblast cell type that does not phenotypically express its differentiation potentiality beyond the myoblast level. L6 cells have shown exponential growth that "passes on virtually all their progeny the capacity to fuse and to differentiate into muscle fibers." (201)  In the myoblast state, L6 cells  replicate very rapidly with a generation time of 18-20 hours. As cell monolayers become confluent, L6 cells cease DNA synthesis and enter a stage of differentiation. differentiation,  Upon  a series of morphological changes take place: most evidently,  mononucleated myoblasts fuse into multinucleated myotubes with cross-striations.  45  Along with the fusion of myoblasts, re-organization of the maturing myotubes also takes place.  In addition, accumulation of characteristic muscle proteins, such as actin and  myosin, (184) and increasing in levels of creatine kinase and myokinase (155) occur as L6 cells differentiate.  Additionally, the development of insulin receptors and insulin  responsiveness (14) as well as the increase in the specific activities of the two major glycogen regulating enzymes, glycogen synthase and glycogen phosphorylase, can be used as markers of differentiation (185). In addition to the morphological properties, another aspect of L6 cells that has been characterized is its metabolic properties.  Since skeletal muscle has an important  role in glucose utilization, glucose transport and its regulation by insulin is one of the most well described areas in L6 cell biology. The two isoforms of glucose transporters expressed in L6 cells, namely GLUT-1 and GLUT-4, are also found in the heart (130). These two transporters are regulated by insulin (89) and the regulation by insulin depends on the differentiation state of L6 (89, 130). In addition to glucose use, the study of the insulin response in L6 cells relates closely with that of glycogen metabolism. Because they are well characterized, L6 cells serve as a good reference cell for method development. Being a cell line, L6 cells also have an advantage of consistency between batches of culture. In addition, conventional and standardized protocols for cell line cultures are also applicable to L6 cells.  Therefore, L6 cells were used to develop  and to validate various techniques used to assess glucose and glycogen metabolism in a cell culture setting. Since the L6 cell type was used as a pioneering sample for experimental method development, some of the properties and characteristics tested were not applied in the  46  finalized protocols for H9c2 cells. A n example is the determination of the creatine kinase activity in L6 cells.  Also, some of the conditions and methods used in these L6 cell  experiments differ from those used on H9c2 cells because the methods were modified when applied to H9c2 cells.  Experimental Design and Protocol The design of experimental protocols for L6 cells went through many changes and was developed on a trial-and-error basis.  The preliminary protocol was based on the  protocol used in the whole-heart perfusion model, which was a "pulse-chase" model that used two radioisotopes, [ H] and [ C], as tracers. However, the use of [ C] was never 3  14  14  implemented because of problems accounting for metabolized [ CC>2]: As a result, only 14  [ H] was used in the future experiments described below. The protocol development with [ H] also went through series of changes. The 3  original concept was to have a glycogen depletion-and-resynthesis process with the intention to label a portion of the intracellular glycogen pool with radioactive isotope (Fig. 3.5 - L6 Cells: Protocol I with [ H] radioactive isotope). The idea was that by monitoring the changes in the labeled and non-labeled glycogen content, the rate and extent of glycogen synthesis and degradation can be deduced.  However, it was later  suspected that such processes could have induced metabolic stress on the L6 cells. Therefore, a modified version of the protocol that involved no exchange of media was chosen as the final protocol (Fig. 3.7 - L6 Cells: Protocol II with [ H] radioactive 3  isotope).  The radioactively labeled glucose was introduced to the cells along with  metabolic modulators like insulin and isoproterenol.  47  L6 cells were deprived of fetal bovine serum (FBS) for 24 hours prior to being based on experiments.  FBS is commonly used in cell culture media as a nutritional  supplement and contains various kinds of hormone and neuronal factors in unknown concentrations.  These hormones and factors are known to induce differentiation in  muscle cells and change the metabolic properties of the cells (46, 129). In order to isolate the effects of differentiation, a 24 hour serum depletion procedure was used (91) in which identical cell culture media, with the absence of FBS, was substituted for 24 hours prior to the experiment. L6 cells used in the experiment were exposed to a series of agents that have specific effects on glycogen metabolism to test the basic metabolic response of L6. Agents stimulating glycogen synthesis and glycogen degradation were required for comparison purposes.  Insulin activates the phosphatidylinositol 3-kinase (PI3-K)  pathway, which subsequently stimulates glycogen synthase.  Therefore, insulin was  chosen as a standard reference for agents stimulating glycogen synthesis. For glycogen degradation,  isoproterenol  activates  the  P K A signaling  cascade  that  phosphorylase kinase, which in turn stimulates glycogen phosphorylase.  activates  As a result,  isoproterenol was used as the reference for glycogen degradation.  48  Results L6 cell differentiation Morphology (Fig. 3.1 - L6 Morphological Properties: Differentiation from Day 0 to Day 8) At the myoblast stage, L6 cells multiplied rapidly with a generation time of about 24 hours.  This rate is comparable to the generation of 18-20 hours reported by  Dr. Yaffe (201).  When L6 cell cultures become confluent (Day 4 of Fig. 3.2), the  myoblasts are induced to differentiate into myotubes by reducing serum concentration in the media. The cell monolayer shows no further signs of growth: no overgrowing or overlapping of cells in the monolayer is observed. However, the phenotype of the cells in culture changes over the next 4 to 5 days. The image in Day 8 of Fig. 3.2 shows that the myotubes are larger in size and are more prominent.  Creatine Kinase Activity (Fig. 3.2 - L6 Biochemical-Metabolic Properties: Creatine Kinase Activity) Activity of creatine kinase, which was used as a biochemical marker of differentiation, was monitored from the first day when the L6 cells were induced to differentiate (Day 0).  On Day 0, the creatine kinase activity was at 1.29 ± 0.03  units/min/mg protein. Over the course of the next few days, the C K activity increased and reached a level of 120.5 ± 8.3 % on Day 4 over the activity on Day 0 and 225.5 ± 6.8 % on Day 8 above the activity on Day 0. The creatine kinase activity had showed no further increase after Day 6.  49  ,. TH'.'.  3 1  ~ P °9' Properties: Differentiation from Day 0 to Day 8 L6 cells fused to form myotubes from Day 0 to Day 8 as cells differentiation g r a d u ^ took L  6  M o ,  h o l  c a l  place.  4.00 -,  Fig. 3.2 - L6 Biochemical-Metabolic Properties: Creatine Kinase Activity. The symbol * indicated that data point was significantly different than Day 0. All groups have p<0.001 in one-way A N O V A test and p<0.05 in Bonferroni t-test.  Total Glycogen Content (Fig. 3.3 - L6 Biochemical-Metabolic Properties: Glycogen Content) In addition to creatine kinase activity, total glycogen content was also monitored over the course of 8 days after the initiation of induced differentiation of the L 6 cells. O n Day 0, the detected glycogen was 36.9 ± 2.3 nmol/mg protein. Similar to the pattern exhibited by the change in creatine kinase activity, glycogen content also showed a dualstage increase in 8 days. Glycogen content was increased by 25.7 ± 6.6 % on D a y 2 and stayed relatively the same on D a y 4. O n D a y 6, the glycogen content further increased and was 54.2 ± 5.8 % above D a y 0. The glycogen content did not differ between D a y 6 and D a y 8.  52  Fig. 3.3 - L6 Biochemical-Metabolic Properties: Glycogen Content. The symbol * indicated that data point was significantly different than Day 0 and the symbol t indicated that data point was significantly different than Day 2. All groups have p<0.05 in one-way A N O V A test and p<0.05 in Bonferroni t-test.  Glycogen and glucose metabolism in L 6 cells Different approaches were used while attempting to establish reliable methods to measure glycogen and glucose metabolism in L6 cells. Each of these is summarized separately below.  Protocol I (Fig. 3.4 - L 6 Cells: Protocol I with [ H]-glucose radioactive isotope) 3  In this series of experiments, the original FBS-containing D M E M culture media was replaced with FBS-free D M E M media for 24 hours (91) in the FBS DEPRIVATION stage. After that, HBSS salt solution, which contained no glucose, replaced the FBS-free D M E M media, which contained glucose, in order to initiate depletion of glycogen. After 6 hours, glycogen is replenished by incubating the cells in HBSS salt solution (pH 7.4) supplemented with 5.5 m M glucose for 2 hours. Then, the HBSS salt solution was removed after 2 hours and D M E M media labeled with [5'- H]-glucose (1.0 uCi/mL) was 3  added to L6 cultures for 30 minutes to study the rate of glycogen re-synthesis and the rate of glycolysis.  54  5.5 mM Glue 0 mM Glue 10/2/0% FBS 0 % F B S  t —  K  —y— —v—*—\  A  — Y  A  V Days/24 Hrs"  6 Hours  Basal / FBS Deprivation  Depletion  I  "  5.5 mM Glue 5.5 mM Glue 0%FBS 0% FBS  1  1  1  2 Hours  Synthesis  1  1  0.5 Hour  Fig. 3.4 - L6 Cells: Protocol I with [ H]-glucose radioactive isotope. 3  Control  1  1  Glycogen Depletion and Re-Synthesis (Table 3.1 - L 6 Cells: Glycogen Synthesis and Degradation; Fig. 3.5 - L 6 Cells Protocol I: Glycogen Synthesis & Degradation)  The results summarized in Table 3.1 compared the glycogen content of the L6 cells during the protocol described in Fig. 3.5 and a modified Protocol I without a change of media or depletion step. The glycogen content of L6 cells during the various stages of Protocol I showed expected increase and decrease of glycogen content, with a surprised finding of elevated glycogen content at the end of FBS DEPRIVATION  stage as  compared to expectation that FBS provided extra nutrient factor for glycogen synthesis. Glycogen content at the end of RE-SYNTHESIS was similar to the BASELINE level, which is the basal rate of glycogen content. The measured non-labeled glycogen was 19.6 ±2.1 nmol /mg protein and the [5'H]-labeled glycogen was 0.7 ± 0 . 1 nmol /mg protein. The total glycogen content of the CONTROL group was far lower than BASELINE level. This result suggested that the L6 cells could have been shocked by the DEPLETION stage and glycogen degradation was triggered. In order to test this theory, a modified Protocol I with the identical media exchange but no glycogen depletion was used. The glycogen content at different stages of Modified Protocol I demonstrated a different pattern than the original Protocol I (Table 3.1).  The glycogen  content  remained relatively unchanged after the FBS  DEPRIVATION stage. From this result, it was concluded that the DEPLETION stage had an effect on the glycogen metabolism and should be omitted from the protocol.  56  Table 3.1 - L6 Cells Protocol I: Glycogen Synthesis and Degradation.  Protocol 1  Modified Protocol 1  Baseline (nmol / mg protein)  SerumDepletion (nmol / mg protein)  Depletion (nmol / mg protein)  Re-synthesis (nmol / mg protein)  63.4 ± 4.9  * 87.9 ± 4.4  * 14.3 ± 2 . 3  52.6 ± 4.5  65.8 ± 5.7  * 108.3 ± 5 . 4  * 110.3 ± 9 . 6  * 112.8 ± 8 . 3  F S  * indicated that data point was significantly different than BASELINE, the basal level of glycogen content.  70.0  60.0  £  50.0  40.0  *  30.0  T  20.0  10.0  0.0 BASELINE  DEPLETION  RE-SYNTHESIS  CONTROL  Fig. 3.5 - L6 Cells Protocol I: Glycogen Synthesis & Degradation. The white bar represented unlabeled glycogen. The black bar represented [ H]-labeled glycogen. The symbol * indicated that data point was significantly different than BASELINE. 3  Protocol II (Fig. 3.6 - L6 Cells: Protocol II with [ H] glucose) 3  Compared to Protocol I, Protocol II did not have a DEPLETION stage. Immediately after the 24 hour FBS DEPRIVATION stage, D M E M media labeled with [5'- H]-glucose (1.0 uCi/mL) was added to L6 cultures for 30 minutes to study the rate of 3  glycogen re-synthesis and the rate of glycolysis.  Insulin (lOOnM) and isoproterenol  (InM) was also added to the labeled D M E M media at this stage to determine their effects on glycogen metabolism.  Glycogen Degradation and Synthesis (Table 3.2 - L6 Cells: Glycogen Synthesis and Degradation in Protocol II) After the 30-minute study/experimental period, the magnitude of glycogen degradation in the C O N T R O L group of cells was almost identical to the INSULIN group. For glycogen synthesis, the insulin-treated group had significantly higher rate than the C O N T R O L group. Isoproterenol had a strong effect on L6 glycogen metabolism within the 30 minute of exposure and significantly increased glycogen degradation. Despite the strong stimulation for degradation, glycogen  synthesis was  still detected in the  ISOPROTERENOL group, although it was lower than control and insulin groups.  59  5.5 mM Glue 10/2% FBS  s '  A  7 Days  5.5 mM Glue 0% FBS  v—*  24 Hours  1  Basal  5.5 mM Glue 0% FBS  v—* 1  FBS Deprivation  Fig. 3.6 - L6 Cells: Protocol II with [ H] -glucose radioactive isotope. 3  0.5 Hour  Control  \ 1  Table 3.2 - L6 Cells: Protocol II with [ H] -glucose radioactive isotope. Glycogen Synthesis and Degradation. 3  [a] Glycogen Degradation (nmol / mg protein)  [b] Glycogen Synthesis (nmol / mg protein)  Control Protocol II  31.7 ± 7.5  2.2 ± 0.2  Insulin Protocol II  31.3 ±6.7  * 3.5 ± 0.3  Isoproterenol Protocol II  * 89.6 ± 4.3  r 1.2 ±0.1  * indicated that data point was significantly different than C O N T R O L condition, t- indicated that data point was significantly different than INSULIN condition  Glycolytic Flux (Table 3.3 - L 6 Cells: Rate of Glycolytic Flux in Protocol II) The rate of glycolysis was also measured in the above-mentioned series of experiments. The measured rates of glycolysis for both insulin-treated and isoproterenoltreated groups were significantly different from the C O N T R O L group.  Both groups  showed accelerated rate of glycolysis.  Lactate Production (Fig 3.7 - L 6 Cells: Lactate Production) In the same series of experiments, the rate of lactate production by L6 cells was also measured.  The control group showed a detected rate of 0.92 ± 0.03 nmol  lactate/hour/ug protein. Both insulin and isoproterenol had a significantly positive effect on the rate of lactate production. The measured rate of insulin-treated L6 was 1.16 ± 0.06 nmol lactate/hour/ug protein and that of the isoproterenol-treated L6 was 1.28 ± 0.03 nmol lactate/hour/ug protein.  62  Table 3.3 - L6 Cells: Protocol II with [ H] radioactive isotope. Rate of Glycolytic Flux. 3  Glycolytic Rate (nmol/hr/mg protein) Control Protocol II  21.7 ± 0 . 9  Insulin Protocol II  * 25.2 ± 0 . 8  Isoproterenol Protocol II  * 24.8 ± 0.6  * indicated that data point was significantly different than C O N T R O L condition.  1.40  *  Fig 3.7 - L6 Cells Protocol II: Lactate Production. The symbol * indicated that data point was significantly different than C O N T R O L condition. C O N represented the control condition, INS represented L6 cells treated with 100 nM insulin, ISO represented L6 cells treated with 1nM isoproterenol. The symbol * indicated that data point was significantly different than C O N T R O L . ISO group have p<0.001 in one-way A N O V A test and p<0.05 in Bonferroni t-test.  Summary The measured specific activity of creatine kinase (CK) on Day 8 was 3.28 ± 0.07 U/mg protein, and the C K activity in the L6 culture showed a pattern of staggered increase over the 8 days. The reported C K activity values (43) for L6 cells 7 days and 10 days in culture were 1.40 ± 0.27 U/mg protein and 5.32 ± 0.59 U/mg protein respectively. Although the detected C K activity was higher than the reported value on Day 7, the likelihood of the C K activity of L6 cells to reach ~5 U/mg protein was high based on the trend observed. The detected glycogen level at basal conditions was -57 nmol/mg protein in the preliminary characterization of L6 cells and -65 nmol/mg protein with Protocol I. These results are comparable to the reported basal glycogen level of -50 nmol/mg protein (43). A comparison of glycogen content at the end of RE-SYNTHESIS stage in Table 3.1 and that of the C O N T R O L group in Fig. 3.6 showed a difference of -30 nmol/mg protein. This finding suggested that the DEPLETION stage of Protocol I might have induced a metabolic stress to L6 cells. This hypothesis was supported by the results of Modified Protocol I without a depletion stage, which the glycogen level at the end of the RE-SYNTHESIS stage remained at a much higher level. As a result, Protocol II without a glycogen depletion stage was used. The insulintreated L6 cells showed similar level of glycogen degradation as the C O N T R O L group. However, the INSULIN group showed increased glycogen synthesis. On the other hand, glycogen degradation was dramatically increased in the ISOPROTERENOL group and a small rate of glycogen synthesis was detected in this group of cells. The detected rates of glycolysis were higher in the insulin-treated and isoproterenol-treated groups.  The  65  measured rate of lactate metabolism under Control condition was 0.92 ± 0.03 nmol/ug/hr, which was higher than the reported range from 0.50 ± 0.02 to 0.73 ± 0.02 nmol/ug/hr (16).  Discussion In 1978, many of the significant metabolic properties of L6 are reported (137). Key intracellular metabolite levels involved in the regulation of glycogen metabolism were described:  intracellular glucose was detectable throughout the incubation, even  when medium glucose was as low as 0.91 mM; glucose-6-phosphate/glucose ratio is low at 5-15% in the first 73 hours and gradually increase to 37% at the end; maximal glycogen deposition takes place during the 6- to 28-hour incubation period and then net glycogenolysis  was observed; the average rate of glycogen consumption was 0.84  nmol/hr/mg protein, less than 1% of the rate of glucose utilization; a small decrease in cell glycogen between the 1- and 6- hour incubation periods is consistent among different subcultures. The extensive profile of its metabolic properties, therefore, made L6 a good reference for establishing the protocols to examine various aspects in the regulation of glycogen metabolism in H9c2 cells, cells not yet characterized metabolically. Yet, there were also limitations on the use of L6 as a reference in protocol development. Many of the recent studies on L6 cells used different culturing conditions that were not entirely comparable to each other. Therefore, making conclusions based on the metabolic data from these different studies and using these results as references for the development of experimental protocols was more challenging than originally anticipated.  66  First of all, the increase of glycogen content after serum deprivation was not expected. Most published studies on the metabolic properties of L6 cells were performed with the presence of serum in the media. The consideration of isolating the effect of serum took place only when the studies focused on the differentiation aspect of L6 (9,46, 91, 95, 98, 129, 130, 140, 141, 175).  In some of these studies, the results clearly  indicated that the presence of serum induced changes to the morphological properties of L6 cells. Fetal bovine serum (FBS) is a supplement that has numerous types of proteins, hormones, neuronal factors, etc. The type and concentration of each of these components are not generally available and it was highly likely that some of these factors or hormones in the serum affects energy metabolism of the L6 cells. Therefore, despite the lack of data support, FBS deprivation of L6 cells before experimentation was considered necessary. As summarized in Table 3.2, simultaneous synthesis and degradation of glycogen was detected in all experimental conditions. These results suggested that despite strong stimulation of glycogen degradation from isoproterenol, synthesis of glycogen continued to take place. In contrast to isoproterenol, insulin had no effect on the glycogen content of L6 cells.  Studies on insulin's effect on glucose uptake and differentiation were  conducted when the L6 cells were at the myoblast stage (46, 88).  The poor insulin  sensitivity of glycogen synthesis is a feature in differentiated L6 cells. The rate of glycolysis could not be directly referenced to any studies, because the methods or metabolic products used to measure the glycolytic flux were different (16, 202). The elevated rate of glycolysis for the ISOPROTERENOL group corresponded to the stimulation of glycogen degradation by isoproterenol. The insulin-treated group also  67  showed increased glycolytic flux. Insulin is known to stimulate glucose transport and enzymes of glycolysis (144,176,187), thereby, up-regulating glycolysis. The rates of lactate production showed identical pattern as the rates of glycolysis. The basal rate of lactate was higher than the published rate of 0.50 ± 0.02 to 0.73 ± 0.02 nmol/ug/hr (16). Although rates of lactate production are slightly higher than values reported by others, I felt that my results matched them well enough when the different protocols used were considered. Since lactate was a product of glycolysis, this result reconfirmed the authenticity of results of the detected glycolytic flux. Overall, the detected metabolic profile of L6 matched the published data reasonably well on L6 cells. Therefore, the L6 cells in this series of experiment served the purpose of verifying experimental protocols on cell culture, glycogen detection and glycolysis.  68  CHAPTER 4 - GLUCOSE AND GLYCOGEN METABOLISM IN HEART-DERIVED H 9 C 2 CELLS  Introduction In 1976, Kimes and Brandt established the H9c2 cell line and characterized its properties (86).  H9c2 cells, or H9c2(2-1) as its full designation, were derived from  embryonic BDLX rat heart ventricular tissue using Yaffe's "selective serial passage" technique (201), the same method that established L6 cells, with slight modifications. Today, H9c2 cells are recognized as one of the few cell lines that exhibit cardiac myocyte characteristics (28, 34, 67, 139, 180). Hence, H9c2 is widely considered to be a cellular model for cardiac myocytes. Despite their recognition as a model of cardiomyocytes, H9c2 cells were initially selected for their resemblance to skeletal muscle cells. Similar to skeletal muscle cells, H9c2 cells undergo morphological changes upon reaching confluency in cell culture. The fusion and re-orientation of cells and the formation of multi-nucleated cells that is common in skeletal muscle cells is also observed in H9c2 cells (43, 145, 185, 201). In addition to morphological differentiation, markers for muscle development, including an increase in key enzyme activities, like myokinase (MK) and creatine kinase (CK), were also detected in differentiated H9c2 cells (8, 69, 101, 178). Another marker of skeletal muscle phenotype is the isoform of C K expressed.  In adult skeletal muscle, the C K  isoform expressed is predominantly the M M isoform, while that in dividing myoblasts is the BB isoform. As H9c2 cells differentiate, primary C K expression changes from the BB isoform to the M M isoform.  69  Besides myokinase and creatine kinase activity and isoform differences, another marker for skeletal muscle cell phenotype is tissue-specific L-type voltage-dependent calcium channel (VDCC).  Both the cardiac and the skeletal isoform of these C a  2 +  ion  channels are simultaneously found in H9c2 cells (122) and a transition in the expression of V D C C , from a skeletal-phenotype dominant to a cardiac-phenotype dominant, occurs during the differentiation of H9c2 cells (125). The differentiated H9c2 cells also show positive immunolabeling for myogenin, troponin T and MyoD, all of which are markers for skeletal muscle myotubes and are not expressed in cardiac cells (125). Menard et al suggested that the myogenic differentiation of H9c2 was a result of a transdifferentiation process resulting in a phenotypic change from cardiac muscle to skeletal muscle. Thus, differentiated H9c2 cells show many phenotypic features that are characteristic of skeletal muscle myotubes, and not cardiac myocytes.  These results indicate that H9c2  cells, if not interfered with, will gradually lose their cardiac phenotype and transform into a more skeletal muscle-like phenotype. Despite the propensity to spontaneously develop skeletal muscle-like phenotype, an intervention to preserve the cardiac phenotype of H9c2 cells was also discovered (125).  Rather than growing the culture until it became confluent and then inducing  differentiation, H9c2 cell culture was chronically treated with all-trans-retinoic acid analogs (10 nM, R A analogs) for 5-7 days before reaching confluence. After treatment, these RA-treated H9c2 cells showed noticeably reduced growth along with distinct morphological changes.  RA-treated cells were multi-nucleated and appeared large and  rounded instead of elongated, which is the typical shape of non-RA-treated differentiated H9c2 cells. Biochemically, RA-treated H9c2 cells were negative for the skeletal muscle  70  markers, skeletal troponin T and myogenin.  W h e n checked for existence of cardiac  troponin T, a well-characterized marker for cardiac differentiation (59, 181, 182), R A treated H9c2 cells showed increased expression compared to n o n - R A treated cells. (Fig. 4.2). Together with results of functional expression of cardiac L-type C a  channels, this  finding confirmed that RA-treated H 9 c 2 maintained a cardiac-lilce phenotype.  Thus,  H9c2 cell culture can be considered a cellular model of cardiac myocytes and can be used to investigate metabolism of glucose and glycogen and its control in heart muscle cells.  Experimental Design and Protocol In the light o f the above findings, all H9c2 cell cultures used in the experiments (except where indicated) were subjected to retinoic acid treatment for 5 - 7 days in order to preserve the cardiac phenotype. In addition, the techniques and protocols derived from L 6 cell experiments in Chapter 3 were applied to H9c2 cells in investigating the control of glucose and glycogen metabolism i n these cells. H9c2 cells i n the experiment were exposed to a series of agents that have specific effects on glycogen metabolism. Agents stimulating glycogen synthesis and glycogen degradation  were  required  for  comparison  purposes.  Insulin  activates  the  phosphatidylinositol 3-kinase (PI3-K) pathway, which subsequently stimulates glycogen synthase.  Therefore, insulin served as a standard reference for agents stimulating  glycogen synthesis.  A s for the standard for glycogen degradation, isoproterenol, it  activates the P K A signaling cascade. The P K A cascade activates phosphorylase kinase, which i n turn stimulates glycogen phosphorylase.  71  Although a major overall goal of this study is to understand control of glycogen metabolism under ischemic conditions, H9c2 cell cultures cannot undergo true ischemic shock like samples in heart-perfusion protocols. Therefore, an alternative method was used.  Oligomycin is a chemical agent that abolishes the A T P synthesis function of  mitochondria (116).  Therefore, when applied to H9c2 cells, oligomycin mimics a  hypoxic conditions and produces a metabolic stress. In order to study the role of A M P K in glycogen turnover as stated in the hypothesis, a stimulating agent of A M P K , AICAR, was used. AICAR is the most commonly used pharmacological agent to stimulate the activity of A M P K .  Once within the cell, AICAR was metabolized into Z M P , an  analogue of 5'-AMP that activates A M P K . AICAR, l.OrnM and 2.0mM, were used.  In the experiment, two concentrations of  These concentrations were chosen to allow  comparison between H9c2 cells and isolated heart samples exposed to 0.8 m M and 1.2 mM AICAR (112).  Results Morphologic Characteristics (Fig. 4.1 - H9c2 Cells Morphological Properties: Cardiac Differentiation Vs. Myogenic Differentiation) H9c2 cells multiplied as a mononucleated myoblast that was large, flat and spindled-shaped (Day 1) and had a generation time of approximately 30 hours. Upon reaching confluency on the surface of tissue culture vessels, a fusion process began and the cells culture formed multinucleated tubular structures (Day 3). The orientation of the cells also changes from random in the myoblast stage to more linear and parallel arrays in the myotube stage (Day 5). In the RA-treated group, the H9c2 cells remained relatively  72  less fused and retained more of the original morphology before the fusion process began (Day 1 to 5).  In comparison, non-treated H9c2 cells exhibited L6-like morphological  changes (see Chapter 3) and extensive fusion of cells that resemble a "net-like" appearance were observed.  H9c2 cells in Day 7 exhibited identical morphological  features as in Day 5.  Expression of cardiac-specific proteins (Fig. 4.2 - H9c2 Cells Cardiac Troponin-T Protein Expression) Cardiac troponin T, a 38kDa protein, was measured in H9c2 cells after 5 days of R A treatment. As shown in Fig. 4.2, cardiac troponin-t is detectable in non-RA treated H9c2 cells.  In comparison, the RA-treated group showed bands with much higher  intensity, indicative of preservation of a cardiac phenotype.  73  Cardiac  Myogenic  Fig. 4.1 - H9c2 Morphological Properties: Cardiac Differentiation Vs. Myogenic Differentiation Myogenic cells fused to form myotubes as cells differentiation gradually took place. Cardiac cells did not fuse like myogenic cells.  (-) Retinoic acid  (+) Retinoic acid  38kDa  P<0.001  Fig. 4.2 - Cardiac Troponin-T Protein Expression. The molecular weight of the target protein was 38 kDa and the P-value from one way A N O V A test comparing the mean of two groups was P<0.001. Courtesy of Ramesh Saeedi, PhD student, U B C .  Total Glycogen Content (Fig. 4.3 - Glycogen Content in Myogenic Phenotype) Glycogen content in H9c2 cells with a myogenic phenotype was investigated in order to compare the glycogen metabolic properties of the two different H9c2 phenotypes. The Control group, served as a baseline reference, had glycogen content of 212.2 ± 11.8 nmol/mg protein. Glycogen content in the 0.1 u M insulin-treated group (180.8 ± 9.6 nmol/mg protein) did not differ significantly from Control.  The group  treated with 10 u M isoproterenol had a decrease in glycogen by about 56% with 119.1 ± 10.8 nmol/mg protein.  Comparatively, 0.5 u M oligomycin had a much stronger  stimulation on glycogen degradation and the detected glycogen was 56.8 ± 10.1 nmol/mg protein. The 1.0 m M AICAR group had 181.9 ± 9.8 nmol/mg protein glycogen content, which was very similar to the group treated with 0.1 u M insulin. As for 2.0 m M AICAR, the detected glycogen was 148.0 ± 13.4 nmol/mg protein, a value significantly lower than Control.  Glycolysis (Fig. 4.4 - Rate of Glycolysis in Myogenic Phenotype) Myogenic H9c2 cells, under control condition, had a glycolytic rate of 37.4 ± 2.9 nmol/hr/mg protein.  Insulin had minimal effect on these myogenic H9c2 cells.  The  detected glycolytic rate for the Insulin group was 38.7 ± 1 . 8 nmol/hr/mg protein and was not significantly different from the Control group.  In contrast, isoproterenol showed  significant stimulation of glycolysis and the detected rate for the isoproterenol-treated group was 55.2 ± 4.4 nmol/hr/mg protein. The two AICAR-treated groups showed no significant changes in the rate of glycolysis. The rate for 1.0 mM AICAR group was 48.5 ± 3 . 3 nmol/hr/mg protein and the rate for the 2.0 m M group was 46.2 ± 2.5 nmol/hr/mg  76  protein. However, the group with the most significant stimulation in glycolysis was the oligomycin-treated group.  The detected rate of glycolysis was twice the rate of the  Isoproterenol group at 101.4 ± 3.9 nmol/hr/mg protein.  77  8Z.  H  CO  Glycogen Content (nmol/mg protein) o b  3  st  01 o  o o b  o b  o o  2. » 2" »  >  o >  33  >  9-  O >  si  3J  CD ro  r=-  O.O  O CD  B- 52-  3 &  2.  Q  ro b  CD  5' <0 T3 >  CD O  § O  CO  CO  O  £ - > -* s TJ"8 t4 3 2  sa  ro o o  ro 01 o b  6L  Glycolytic Rate (nmol/hr/mg protein)  2 ~ CD N> 03 ©  3  O O  3  v  CD  Q. A  3  ro o  o b  3 i l > fl o /—\ rv» P ro  o b  O) o  oo o  Q  Q. >  O  O CD  u  33 CD Q  3  o _ J2 O W l  CO  i  O O  3 O £ 3 - — 9L 3  &  CD  a-.  3  sf 2  O 3  x> O 3 a. CD < w -o CD  O 2. " <P_ CD 33 w a. go  >^ O >  > 33  r» O O CD £ O  left O g CTI  O Tj  c  tz  5 ?< o_ S O to" «' § o o z  >  O  > 33  to  b  3 -g =  T-TT  5" ©, CD _ _ • m w CD *  CD  3 >2 3-  O  33 O  < -2 CD ^ "D CD  o 3  O  v co o 3 O CD a.  ?. 2 - s 3  o §  O  CD -  CD "  z 05  Z  £ s«? ^  W CD  -a < =• 2  ^ .  CD  p ©o  6 3 2.  CO  O  1:1:1:1:1:1  o o  Total Glycogen Content (Fig. 4.5 - Glycogen Content in Cardiac Phenotype) Without the presence of any effectors of metabolism or H-glucose label in the media, glycogen content for the Control group with only H-glucose label in the media was 190.1 ± 12.6 nmol/mg protein. In the presence of a high physiological concentration of 0.1 u M insulin, glycogen content remained relatively constant with a slight but insignificant increase of only 6.7 ± 6.4 %. In contrast, 10 n M isoproterenol had a much more significant effect on glycogen content, lowering glycogen content by 61.9 ± 5.7 % to 122.7 ± 11.3 nmol/mg protein. With l.OmM AICAR, total glycogen content detected decreased slightly but not significantly to 188.9 ± 14.3 nmol/mg protein, which was 95.2 ± 7.2 % of the control level.  A higher concentration of AICAR at 2.0 m M further  reduced the glycogen content slightly to 89.9 ± 7.3 % of control level, which was 178.3 ± 14.6 nmol/mg protein.  However, this difference was also not statistically significant  from the Control group. H9c2 cells treated with oligomycin led to the lowest level of glycogen amongst all the groups: the detected glycogen content after 30-minutes of treatment was 72.6 ± 1 1 . 6 nmol/mg protein, which was decreased to 36.6 ± 5.8 % of the Control group level.  80  250.0 -,  CON  AICAR 1.0  AICAR 2.0  OLIGO  INS  ISO  Fig 4.5 - H9c2 Cells Cardiac Phenotype: Total Glycogen Content. CON represented the control condition, INS represented cells treated with 100 nM insulin, ISO represented cells treated with 1nM isoproterenol, AICAR represented cells treated with 1.0 mM or 2.0 mM AICAR, and ISO represented cells treated with 0.5 uM oligomycin. The symbol * indicated that data point was significantly different than control condition in one-way ANOVA with p<0.001 and in t-test with p<0.001. The symbol t indicated that data point was significantly different than the isoproterenol-treated cells, in one-way ANOVA with p<0.001 and in t-test with p<0.001.  Glycogen Turnover Glycogen Degradation (Table 4.1 - H9c2 Cells Cardiac Phenotype: Glycogen Synthesis and Degradation) Table 4.1 clearly shows the differences in glycogen synthesis and degradation of H9c2 cells in response to the various reagents. consistently higher than those of synthesis. conditions  produced significant  change.  The magnitude of degradation was  For the degradation of glycogen, most The two  degradation-inducing agents,  isoproterenol and oligomycin, produced expected results and the detected glycogen degradation were tripled or more for the two groups as compared to Control.  The  AICAR-treated groups produced different results and only the higher concentration (2.0 mM) group had a significant effect on glycogen degradation. A n interesting finding with the Insulin group was that despite the glycogen synthesis stimulation, a moderate level of glycogen degradation was still detectable.  Glycogen Synthesis (Table 4.1 - H9c2 Cells Cardiac Phenotype: Glycogen Synthesis and Degradation) Another interesting finding is that even when H9c2 cells were exposed to agents that strongly stimulated glycogen degradation, like oligomycin and isoproterenol, significant levels of glycogen synthesis were still detected.  When focusing on the  synthesis of glycogen, insulin showed only minimal stimulation of glycogen synthesis. At  the  same time,  the  glycogen-degradation  stimulating  isoproterenol  significantly inhibit glycogen synthesis while boosting degradation.  did not  Surprisingly,  82  AICAR dramatically stimulated glycogen synthesis and the AICAR-treated groups showed the highest synthesis of glycogen.  Simultaneous Synthesis and Degradation (Turnover) (Table 4.1 - H9c2 Cells Cardiac Phenotype: Glycogen Synthesis and Degradation & Fig. 4.6 - H9c2 Cells Cardiac Phenotype: Rate of Glycogen Synthesis and Degradation) The  result above from all experiment conditions strongly supported that  simultaneous synthesis and degradation of glycogen exists in H9c2 cells. When the rate of glycogen synthesis and degradation were compared simultaneously, more intriguing findings  arose.  Insulin, with lowered but not statistically  significant  glycogen  degradation, showed a tendency of decreased turnover of glycogen than the Control group. In a paired t-test detecting a change of 0.5, with expected standard deviation of change equals to 1, desired power equal to 0.8 and alpha equals 0.05, a sample size of 34. Therefore, the insulin group could be exhibiting a statistical insignificance rather than a biological insignificance.  In contrast, AICAR-treated groups had stimulated glycogen  turnover due to increased synthesis and degradation.  Oligomycin and Isoproterenol  groups also showed increase glycogen turnover, but were strongly skewed towards degradation.  Overall, only AICAR groups had an even increase in both glycogen  synthesis and degradation.  From the perspective of glycogen metabolism, only  conditions that stimulate B O T H synthesis and degradation should be considered to stimulated turnover. Therefore, AICAR-treated group was the only group that stimulated the turnover of glycogen in H9c2 cells.  83  Table 4.1 - H9c2 Cells Cardiac Phenotype: Glycogen Synthesis and Degradation.  Glycogen Synthesis  Glycogen Degradation  (nmol / mg protein)  (nmol / mg protein)  Control  8.3 ± 0.7  33.8 ± 6.4  AICAR (1.0 mM)  * 14.4 ± 1 . 0  45.7 ± 7.0  AICAR (2.0 mM)  * 15.1 ± 1 . 0  * 54.4 ± 4.9  *t 4.2 ± 0 . 4  *t 153.2 ± 6 . 1  Insulin (0.1 uM)  10.3 ± 0 . 9  17.5 ± 6 . 7  Isoproterenol (10 uM)  6.6 ± 0.5  * 103.1 ± 6 . 2  Oligomycin (0.5 uM)  * indicated that data point was significantly different than control condition in one-way ANOVA with p<0.05 and in t-test with p<0.05. The symbol t indicated that data point was significantly different than the isoproterenol-treated cells, in one-way ANOVA with p<0.05 and in t-test with p<0.05.  nmol/min/mg protein Fig. 4.6 - H9c2 Cells Cardiac Phenotype - Rate of Glycogen Synthesis and Degradation. CON represented the control condition, INS represented cells treated with 100 nM insulin, ISO represented cells treated with 1nM isoproterenol, AICAR represented cells treated with 1.0 mM or 2.0 mM AICAR, and ISO represented cells treated with 0.5 uM oligomycin. The symbol * indicated that data point was significantly different than control condition in one-way ANOVA with p<0.05 and in t-test with p<0.05. The symbol t indicated that data point was significantly different than the isoproterenol-treated cells, in one-way ANOVA with p<0.05 and in t-test with p<0.05.  Glycolysis (Fig. 4.7 - H9c2 Cells Cardiac Phenotype: Rate of Glycolytic Flux) Generally, the results of glycolytic flux were very similar respectively to the results of glycogen degradation for each group in the experiment.  Isoproterenol  stimulated the rate of glycolysis by 38.5 ± 5.9 % over the Control condition to 56.9 ± 2.4 nmol/hr/mg protein. In comparison, insulin induced less stimulation in glycolytic rate of only 14.8 ± 5.8 % and was not statistically significant.  As with the case of glycogen  degradation, the AICAR-treated and the oligomycin-treated groups showed significant changes in glycolytic rates. Both concentrations of AICAR yielded an almost identical stimulation in glycolytic rate with 58.2 ± 3.2 nmol/hr/mg protein for ImM and 57.9 ± 2.9 nmol/hr/mg protein for 2 m M AICAR, resulting in 41.7 ± 7.8 % increase for ImM and 41.1 ± 7.0 % increase for 2 m M AICAR. The response of glycolysis in the presence of 0.5 u M oligomycin was even more dramatic at 104.1 ± 2.6 nmol/hr/mg protein, which was an increase in glycolytic flux of 153.5 ± 6.2 % over the control.  86  140.0 120.0 c  &  o 100.0 a.  D)  E  o  80.0  E c  a>  & a 60.0  OC o  !  o o  >. O  40.0  I I  20.0  0.0 CON  AICAR 1.0  AICAR 2.0  0LIG0  INS  ISO  Fig 4.7 - H9c2 Cells Cardiac Phenotype: Rate of Glycolytic Flux. CON represented the control condition, INS represented cells treated with 100 nM insulin, ISO represented cells treated with 1nM isoproterenol, AICAR represented cells treated with 1.0 mM or 2.0 mM AICAR, and ISO represented cells treated with 0.5 uM oligomycin. The symbol * indicated that data point was significantly different than control condition in one-way ANOVA with p<0.05 and in t-test with p<0.05. The symbol f indicated that data point was significantly different than the isoproterenol-treated cells, in one-way ANOVA with p<0.05 and in t-test with p<0 05  Glucose Uptake (Fig. 4.8 - H9c2 Cells Cardiac Phenotype: Calculated Glucose Uptake) Insulin, which had a mild stimulating effect on glycolysis and glycogen synthesis in the H9c2 cells, induced a small increase in the rate of exogenous glucose uptake as compared to the Control. However, this increase was not significant. Isoproterenol had a modest stimulation on glycolysis, but also displayed a low level of glycogen synthesis. As a result, glucose uptake, calculated as the sum of glycolysis and glycogen synthesis, was insignificantly increased above the Control by 21.9 %, a value that was similar to that of the insulin-treated group. With a greater stimulation in glycogen synthesis and an equally strong stimulation in glycolytic flux, AICAR induced a sizable increase of 51 to 53 % in glucose uptake in H9c2 cells above Control values. Oligomycin showed a very dramatic stimulation in glycolytic flux that was highest amount all the groups tested. Although the rate of glycogen synthesis was low, the oligomycin-treated group had a significant increase in the rate of glucose uptake, nearly doubling the rate above Control value.  88  140.0  £  -i  120.0  a> ** o k.  a. E  100.0  o  1  U H  80.0  (0 (0  60.0  o o °  40.0  «(0  20.0  liiil  i i i l i t  mm  u 0.0 CON  iLVr? "th  1nn  C a  [  AICAR 1.0  d i a  "!TS^' H fion^n 'T ° nr ? n 1 M  A  c i  o  S  r ? A o  I  S  r  e  D  r  P  h e n 0 t  °  r e  e  s  e  yP  e :  n  e  d  c e l l s  t  n  n  Way  INS  ISO  c e l l s t r e a t e d w i  h  ^  t r e a t e d  £ S ^ T d ^ thTn In" ' ° ?" f ° significantly different than the isoproterenol-treated 0  OLIGO  Calculated Glucose Uptake. CON represented the control condition, INS represented cells * isoproterenol, AICAR represented cells treated with 1.0 mM with 0.5 uM oligomycin. The symbol * indicated that data point was significantly  P  r e s e n t e d  t  AICAR 2.0  N  V  A  °- « y m t indicated that data point was cells, in one-way ANOVA with p<0.05 and in t-test with p<0.05.  W i t h  P <  0 5  a  n  d  i n  W  e  S  t  W i t h  P  <  0  0  5  T  h  e  w  7  Activity of Glycogen Phosphorylase (Fig. 4.9 - H9c2 Cells Cardiac Phenotype: Activity of Active Glycogen Phosphorylase) The results on glycogen phosphorylase were surprising: besides insulin that stimulated glycogen synthesis, the glycogen degradation-inducing agents oligomycin and isoproterenol did not significantly alter the activity of glycogen phosphorylase. The only group the showed altered GP activity was the 2 m M AICAR group, which had a significantly lowered activity level.  Activity of Glycogen Synthase (Fig. 4.10 - H9c2 Cells Cardiac Phenotype: Activity of Active Glycogen Synthase) Comparing to the results for GP, the results for GS were even more surprising: insulin did not significantly increase GS activity, but the isoproterenol-treated group did. Also, neither 2 mM AICAR nor oligomycin significantly changed GP activity.  90  16  Rate of Activity (nmol/min/mg protein) OS S 0 3 o P ^-  i,  O" O l 3 S» CD O C L 9. T J  5TT0  CD  ^  £ co rr 3 CD  b o  CO  O  CJl  b 0  b 0  ro  ro  0 b 0  cn b 0  u  0 b 0  CO  01 b 0  I  I  »  5'  8 > 24- = ^ rr —  3  o  o 3  ^  ro O CD  w" O 3  o 0)  s'  o  SL8  J 0> TCO CO CO  OP  3  3"  -I  CL „  CD 3  CO  > o  CO  0) ICD 3 o TO CD V  O  I  b 0 3 a en w <  3 3 CD 3"  O 3 CD  0)  TJ S 0) 0) 3 •<  » -g  s =r co  CD CD 0) 3 CD CD  o  3 CO  o  N> O  c?  CD O CD ( Q CD CL  CD > CD  33  T"T  &? P  c <  P  a  >  p >  3> 3  >  O  CL  3  CO  o  '  (0 CD  O  TJ CO CD CD CL TJ  •™i  CD CO CD  CO  CO  O  £ 8-1 s  ~  3  o  2  o  o  3 CL  CD CD CD CL  CD CO CD 3 CD  CL o CD CO  3  Si CD CL  I.I.I.I.I.I.I.I.I.  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Interestingly,  the  isoproterenol-treated  group showed a  decreased phosphorylation level and was the only group that had a negative effect on A M P K phosphorylation.  Surprisingly, AICAR, a commonly used activator of A M P K ,  had no effect on the phosphorylation level of A M P K .  The only reagent that strongly  induced phosphorylation of A M P K was oligomycin. The phosphorylation of the substrate of A M P K , A C C , was also assessed in order to evaluate the possibility of allosteric activation of A M P K , which was not detectable from the western blot analysis on A M P K  phosphorylation.  The results of the  phosphorylation of A C C (pACC) showed identical trends as the pAMPK: Isoproterenol had negative effect on the phosphorylation of A C C and oligomycin considerably stimulated the phosphorylation of A C C .  93  Fig. 4.11 - H9c2 Cells Cardiac Phenotype: Immunoblot Analysis on Phosphorylation of A M P K and A C C . Data obtained with the technical assistance of Hannah Parsons, UBC.  Summary The metabolic properties of AICAR-treated H9c2 cells were very interesting, being dependent upon the cellular phenotype. While AICAR decreased total glycogen content with increasing concentration in H9c2 cells with a skeletal muscle phenotype, cardiac phenotype H9c2 cells showed no change in total glycogen content. Detailed analysis, however, showed that simultaneous glycogen synthesis and degradation (or glycogen turnover) in the cardiac H9c2 cells were enhanced.  AICAR also caused  different effects on glycolysis in the two phenotypes of H9c2 cells. AICAR had no effect on skeletal H9c2 cells but significantly stimulated glycolysis and calculated-glucoseuptake in cardiac H9c2 cells. intriguing:  although  AICAR  The detected results for activity of GS and GP were simultaneously  increased  glycogen  synthesis  and  degradation, neither the activity of active GP nor GS was increased. Oligomycin-treated cells yielded expected results.  The chemical-hypoxia  condition induced a massive decrease in glycogen content and stimulation in glycogen degradation in both skeletal and cardiac phenotypes of H9c2 cells.  Despite that  oligomycin drastically stimulating degradation, glycogen turnover was still present. The glycolytic rate was also significantly increased for both phenotypes. The activities of active GS and GP were not significantly different from the C O N T R O L conditions. In general, out of the two glycogen-metabolism effectors that served as references for glycogen synthesis and glycogen degradation, insulin and isoproterenol, only isoproterenol showed expected results in the experiments. However, the impacts of insulin on the H9c2 cells from the various aspects of glycogen metabolism tested were minimal.  The INSULIN group showed results similar to the C O N T R O L group in  95  glycogen content,  glycogen synthesis and glycolytic  isoproterenol, displayed expected  flux.  On the other hand,  properties of beta-adrenergic  agonists,  causing  significantly increased glycogen degradation and glycolytic flux, as well as activity of active glycogen phosphorylase.  Although isoproterenol was known to inhibit glycogen  synthesis and to stimulate degradation, glycogen  turnover was  detected in the  ISOPROTERENOL group. The assay on the activities of glycogen synthase and glycogen phosphorylase, as well as immunoblot analysis of A M P K and A C C phosphorylation produced controversial results, because they did not account for the changes in metabolism observed. A possible explanation for that was the small sample size used (n=3). Given the volatile nature of energy pathway enzymes, such a small sample size could not accurately determine the true activities of the enzymes.  Nevertheless, the data provided a general trend of  responses for H9c2 cells on the various reagents.  However, the sample size for the  experiment is lower than the required sample size for paired t-test. Therefore, a larger sample size is needed for more accurate comparison of the effects of the various reagents.  96  CHAPTER 5 - DISCUSSION With reference to the three objectives of this research project, glucose and glycogen metabolism in H9c2 cells were successfully measured and the control of A M P K on the glycogen turnover of H9c2 cells was investigated. The pharmacological activator of A M P K , AICAR, demonstrated unexpected results in both glycogen metabolism and A M P K activation.  This project remained novel from the perspective that the glycogen  metabolism was, for the first time, characterized in the established cardiac-like H9c2 cell line. In addition, the characteristics of cardiac phenotype of the H9c2 cells were ensured to have maintained for better modeling of cardiomyocytes, as demonstrated by phenotype dependent metabolic responses to AICAR. Overall, this project succeeded in developing a protocol for studying glycogen metabolism from a cellular perspective through the use of H9c2 cells.  Metabolic Actions of AMPK Activator, AICAR Research on the effects of AICAR on cardiac cells, especially on glucose and glycogen metabolism, has been relatively little and results reported controversial. Conventionally, published data suggested " A M P K activation, via AICAR, increases cardiac muscle glucose uptake through translocation of GLUT-4 via a pathway that is independent of PI3K" (146).  Yet more recent findings on cardiomyocytes reported  contradictory results on the role of AICAR on glucose uptake (103, 147, 190).  These  results suggested more data were needed to draw a definitive conclusion of AICAR and A M P K on the energy metabolism in cardiac cells.  97  The 2.0 m M AICAR-treated group in the experiment showed increased glycogen degradation and both AICAR groups showed significantly increased glycogen synthesis. I would add a comment here about glycogen synthesis and degradation both being present in H9c2 cells, even when degradation is very substantial, such as with oligomycin. This finding strongly supports the concept that the turnover of glycogen, independent of net glycogen synthesis or degradation, exists in H9c2 under all conditions tested. Although it had no significant effect on the total glycogen content, AICAR had a significant effect on the turnover of glycogen. These effects could not be accounted for by changes in activity of relevant kinases or enzymes.  Z M P , the metabolized  intracellular form of AICAR, could allosterically activate glycogen phosphorylase and increase glycogen degradation (112).  The activation of glycogen synthesis could be  caused by AMPK-induced elevated glucose uptake (103, 146). In addition, low glycogen content could lead to increased glycogen synthesis by decreasing inhibition of GS (109). PFK-1 is also reported to be allosterically activated by AICAR (112) which could lead to increased glycolysis.  Therefore, increase in glycogen turnover could be caused by the  allosteric activation of GS and PFK-1 by AICAR. The rate of glycolysis was also significantly increased in both AICAR-treated groups. As a result, the calculated glucose uptake, based on the rate of glycolysis and the rate of glycogen synthesis, was significantly higher in the AICAR-treated group. Although the A M P K and the A C C enzymes were not phosphorylated, the metabolized from of AICAR, ZMP, could, as mentioned above, cause the acceleration of glycolysis via allosteric effects on PFK-1 (112).  98  As discussed in the Introduction, the key metabolic property of A M P K in the cardiac cells is the modulation of the energy substrate utilization.  In this series of  experiments, the cell culturing media did not include fatty acids as constituents.  The  lack of fatty acids could be a factor determining the activation of A M P K in response to AICAR in the regulation of energy metabolism of H9c2 cells.  Are the metabolic actions of AICAR mediated by AMPK? Utilization of AICAR as an activator of 5'-AMP-activated protein kinase was first reported in 1994 (167). However, in 2003, our lab discovered that AICAR stimulated glycogen degradation via an allosteric mechanism without the activation of A M P K (112). As a result, more data from different enzymes involved in glycogen metabolism were required to interpret the effect of A M P K on glycogen metabolism.  A n advantage of  AICAR is that it does not disturb the intracellular energy state. However, the property that AICAR closely resembles the activation mechanism of A M P toward A M P K is also its major weakness. AICAR cannot specifically activate A M P K without having effects on other pathways via ZMP. The results in Fig. 4.11 summarize the phosphorylation state of A M P K and A C C when exposed to the selected metabolic modulators. AICAR did not significantly change the phosphorylation state of the A M P K and its downstream substrate A C C . This result showed that there was no measured activation of A M P K under these conditions.  In  addition, AICAR did not have significant effect on the detected activity of glycogen synthase (GS) and glycogen phosphorylase (GP). However, a very important point in interpreting the data from GS and GP was that the results were obtained from a small  99  sample pool (n=3) and is much lower than the required sample size of 10 required to detect a difference of 1.0.  Therefore, results were affected by high variances.  Summarizing the above data, the observed change in glycogen turnover cannot be explained by the measured activation of A M P K or alteration of GS and GP activity. The observation is entirely consistent with data obtained in the intact heart (112). Although the results suggested that the covalent modification of A M P K by phosphorylation did not take place, other mechanisms such as the allosteric mechanism by Z M P (112) could have led to activation of PFK-1 and GP and induced glycogen degradation and stimulated glycolysis, respectively. Other signaling pathways like NOguanylate cyclase pathway (103) and p38 mitogen-activated protein kinase (MAPK) (102) were also reported to be activated in the presence of AICAR.  Therefore, these  pathways could also be contributing to the metabolic effects observed.  Methodological considerations Underestimation of glycogen synthesis When compared to glycogen degradation, glycogen synthesis showed both a lower rate and a lower absolute amount.  Although an underestimation of glycogen  synthesis is a possible explanation, the experimental procedure was already optimized to allow the maximum amount of glycogen to be captured for measurement.  The  concentration of [5'- H]-glucose used in this series of experiments (1.0 uCi/mL) was ten times the concentration used in the whole-heart perfusion protocols (0.1 uCi/mL). Also, the largest cell culture flasks possible for the designed protocol was used to maximize the absolute content of cells and to minimize the risk of measurement and extrapolation. The  100  retrieval and measurement protocol for [5'- H]-glycogen was already developed in previous studies (66, 111, 112) and optimized. In some conditions, glycogen degradation was very high. A possibility of underestimating glycogen synthesis could exist because some newly synthesized glycogen could be lost and not be accounted for. Therefore, a solution to the possible underestimation of glycogen synthesis would require further studies.  Non-specificity of AICAR as an activator of AMPK AICAR  was a commonly recognized pharmacological activator of A M P K .  However, there was an inherent problem of non-specificity with AICAR because the ZMP produced could theoretically activate other pathways and enzymes besides A M P K . In addition, AICAR-derived Z M P was shown to activate glycogen degradation through a allosteric activation mechanism (112).  Therefore, AICAR  could also potentially  influence other pathways indirectly through the allosteric mechanism. Therefore, a more specific activator of A M P K would be required for a more accurate study of A M P K .  Lack of insulin effect Essential to glycogen metabolism in humans and many animals, insulin has a central role in the regulation of glycogen. Insulin impacts upon the cellular response to energy substrates like glucose, lactate, fatty acids, and amino acids. However, a lack of insulin effect was observed in these experiments.  A close examination on the cell  culturing protocol suggested that the difference in culturing protocol could be the cause  101  of detected lowered insulin response.  For example, in one protocol that showed  significant effect of insulin (205), H9c2 was cultured in D M E M medium supplemented with 10% FBS. In contrast, H9c2 cells were switched to D M E M medium with 1% horse serum (HS) after 7 days of D M E M with 10 % FBS in the current experiments. The H9c2 cells were also FBS-deprived for 24 hours prior to experiment. As a result of serum deprivation, hormonal factors and components could be depleted and enzymes could be inactivated in the various pathways activated by insulin. Activities and subcellular distribution of components in glycolysis including glucose transporters, 6-phosphofructo-l-kinase (PFK-1), 6-phosphofructo-2-kinase (PFK-2) and pyruvate  dehydrogenase  (PDH),  as  well  as  glycogen  synthase  and  glycogen  phosphorylase could be affected and rendered a low response to insulin stimulation. A larger sample size would be used in future experiments to determine the effect of insulin on glycogen content in H9c2 cells.  H9c2 cells as a good model to study metabolism H9c2 cells had been widely used in cardiac research and were recognized as cardiomyocytes (2, 28, 42, 85, 131, 172). experiment  did not  stimulate  cardiomyocytes (116, 146).  the  However, AICAR used in the H9c2 cells  activity  of  AMPK  as  reported in  isolated  Although the detected allosteric activation of A M P K was  consistent with the finding in intact heart perfusion models (111, 112), the challenge to cross-reference metabolic data obtained from studies with different sample types arose.  102  Besides the issue of comparability among other types of heart tissue preparation, and the methodological details in each H9c2 cells study must also be considered. A n example mentioned above was the difference in response to insulin with the difference in serum content in the cell culture media (205). The culturing conditions for H9c2 played a role in metabolic properties. However the lack of standard culturing protocol for H9c2 made comparing studies with H9c2 cells more difficult.  Another concern was the  morphological properties of the differentiated H9c2 cell line versus the cardiomyocytes in the intact heart. The two cell types differ not only in morphological properties, but also in the intrinsic level of activity because there were no contractile activities in cultured H9c2 cells. In summary, H9c2 cells provided the convenience and accessibility of established cell lines with certain properties of cardiomyocytes.  However, the fundamental  difference between H9c2 and isolated cardiomyocytes and the possible differences in metabolic properties and signaling pathways could not be ignored. Nonetheless, H9c2 could qualify as a valuable tool to develop protocols that could be applied to isolated cardiomyocytes and intact hearts.  103  Future directions As discussed above, H9c2 cells do not qualify as a perfect isolated cell model of the heart that completely recapitulates the whole-heart perfusion model.  However,  techniques and methods acquired from this project could be readily translated to apply on isolated cardiomyocytes. The biggest constraints of using adult cardiac myocytes are the short life span of the cells and the high level of technical knowledge and experience to successfully harvest adult cardiac myocytes. Since H9c2 cells had been profiled in this project, protocols could be optimized on H9c2 cells before implemented on adult cardiac myocytes, which would be a better model for the heart. Besides choosing another cell type for the investigation, using a modulated form of A M P K could be another effective approach to the characterization of the role of A M P K on energy metabolism. A n isolated cell model has the advantage of accepting genetic modification via viral gene transfer or transfection. For example, constitutively active forms of A M P K could be introduced into the cells to assess effects of increased A M P K activity. In the contrary, dominant negative A M P K mutants or siRNA could also be used to reduce its activity. Enzymes upstream or downstream of the A M P K signaling pathways were also potential targets to study the signaling mechanics of A M P K activation. A M P K plays an essential role in regulating metabolism and could involve many signaling pathways.  Studies indicated that A M P K signaling stimulated glucose uptake  and G L U T translocation via the nitric oxide (NO) pathway (103).  Since the level of  glucose uptake had a direct relationship to intracellular G-6-P and glycogen synthesis, the contribution of NO signaling pathway could be significant. Other studies also indicated  104  the relative contribution of phosphorylation by A M P K kinase (AMPKK), and positive allosterism by the ratios of A M P : A T P and Cr:PCr in A M P K activation (7). If fatty acid oxidation would be considered in the study, F A translocase (FAT)/CD36 could be another target. The ability of insulin to inhibit A M P K may be controlled via an Aktmediated mechanism (90) make Akt another target for investigation of AMPK's regulation of cardiac metabolism in the future.  105  Bibliography  1.  Abumrad N, Harmon C, and Ibrahimi A. Membrane transport of long-chain  fatty acids: evidence for a facilitated process. J Lipid Res 39: 2309-2318., 1998. 2.  Aki T, Mizukami Y, Oka Y, Yamaguchi K, Uemura K, Fujimiya T, and  Yoshida K. Phosphoinositide 3-kinase accelerates necrotic cell death during hypoxia. Biochem J 358: 481-487., 2001. 3.  Allard MF, Emanuel PG, Russell JA, Bishop SP, Digerness SB, and  Anderson PG. Preischemic glycogen reduction or glycolytic inhibition improves postischemic recovery of hypertrophied rat hearts. Am J Physiol 261: H66-74., 1994. 4.  Allard MF, Flint JD, English JC, Henning SL, Salamanca MC, Kamimura  CT, and English DR. Calcium overload during reperfusion is accelerated in isolated hypertrophied rat hearts. J Mol Cell Cardiol 26:1551-1563., 1994. 5.  Allard MF, Henning SL, Wambolt RB, Granleese SR, English DR, and  Lopaschuk GD. Glycogen metabolism in the aerobic hypertrophied rat heart. Circulation 96: 676-682,1997. 6.  Allard MF, Schonekess BO, Henning SL, English DR, and Lopaschuk GD.  Contribution of oxidative metabolism and glycolysis to ATP production in hypertrophied hearts. Am J Physiol 267: H742-750., 1994. 7.  Altarejos JY, Taniguchi M, Clanachan AS, and Lopaschuk GD. Myocardial  ischemia differentially regulates LKB1 and an alternate 5'AMP-activated protein kinase kinase. / Biol Chem, 2004. 106  8.  Apostolova MD, Ivanova IA, and Cherian MG. Metallothionein and  apoptosis during differentiation of myoblasts to myotubes: protection against free radical toxicity. Toxicol Appl Pharmacol 159:175-184., 1999. 9.  Apostolova MD, Ivanova IA, and Cherian MG. Signal transduction  pathways, and nuclear translocation of zinc and metallothionein during differentiation of myoblasts. Biochem Cell Biol 78: 27-37., 2000. 10.  Aschenbach WG, Hirshman MF, Fujii N, Sakamoto K, Howlett KF, and  Goodyear LJ. Effect of AICAR treatment on glycogen metabolism in skeletal muscle. Diabetes 51: 567-573., 2002. 11.  Asimakis GK. Myocardial glycogen depletion cannot explain the  cardioprotective effects of ischemic preconditioning in the rat heart. J Mol Cell Cardiol 28: 563-570., 1996. 12.  Beauloye C, Marsin AS, Bertrand L, Krause U, Hardie DG, Vanoverschelde  JL, and Hue L. Insulin antagonizes AMP-activated protein kinase activation by ischemia or anoxia in rat hearts, without affecting total adenine nucleotides. FEBS Lett 505: 348-352., 2001. 13.  Beg ZH, Allmann DW, and Gibson DM. Modulation of 3-hydroxy-3-  methylglutaryl coenzyme A reductase activity with cAMP and wth protein fractions of rat liver cytosol. Biochem Biophys Res Commun 54:1362-1369., 1973. 14.  Beguinot F, Kahn CR, Moses AC, and Smith RJ. The development of insulin  receptors and responsiveness is an early marker of differentiation in the muscle cell line L6. Endocrinology 118: 446-455., 1986.  107  15.  Berry MN and Friend DS. High-yield preparation of isolated rat liver  parenchymal cells: a biochemical and fine structural study. / Cell Biol 43: 506-520., 1969. 16.  Bevington A, Brown J, Pratt A, Messer J, and Walls J. Impaired glycolysis  and protein catabolism induced by acid in L6 rat muscle cells. Eur J Clin Invest 28: 908-917., 1998. 17.  Beyer A, Kitzerow A, Crute B, Kemp BE, Witters LA, and Heilmeyer LM,  Jr. Muscle phosphorylase kinase is not a substrate of AMP-activated protein kinase. Biol Chem 381: 457-461., 2000. 18.  Boden G, Chen X, Ruiz J, White JV, and Rossetti L. Mechanisms of fatty  acid-induced inhibition of glucose uptake. J Clin Invest 93: 2438-2446., 1994. 19.  Boone AN, Rodrigues B, and Brownsey RW. Multiple-site phosphorylation  of the 280 kDa isoform of acetyl-CoA carboxylase in rat cardiac myocytes: evidence that cAMP-dependent protein kinase mediates effects of beta-adrenergic stimulation. Biochem J 341: 347-354., 1999. 20.  Brownsey RW, Boone AN, and Allard MF. Actions of insulin on the  mammalian heart: metabolism, pathology and biochemical mechanisms. Cardiovascular Research 34: 3-24,1997.  21.  Buczek-Thomas JA, Jaspers SR, and Miller TB, Jr. Adrenergic activation of  glycogen phosphorylase in primary culture diabetic cardiomyocytes. Am J Physiol 262: H649-653., 1992.  108  22.  Buczek-Thomas JA, Jaspers SR, and Miller TB, Jr. Post-receptor defect  accounts for phosphorylase hypersensitivity in cultured diabetic cardiomyocytes. Mol Cell Biochem 117: 63-70., 1992.  23.  Buczek-Thomas JA and Miller TB, Jr. Cyclic GMP accumulation in normal  and diabetic primary culture adult rat ventricular cardiomyocytes: a minor role for nitric oxide in phosphorylase activation. Cell Signal 7: 591-598., 1995. 24.  Buczek-Thomas JA and Miller TB, Jr. Identification of the molecular basis  for phosphorylase hypersensitivity in cultured diabetic cardiomyocytes. Mol Cell Biochem 145:131-139., 1995. 25.  Burelle Y, Wambolt RB, Grist M, Parsons HL, Chow JC, Antler C, Bonen A,  Keller A, Dunaway GA, Popov KM, Hochachka PW, and Allard MF. Regular exercise is associated with a protective metabolic phenotype in the rat heart. Am J Physiol Heart Circ Physiol 287: H1055-1063,2004.  26.  Carlson CA and Kim KH. Regulation of hepatic acetyl coenzyme A  carboxylase by phosphorylation and dephosphorylation. J Biol Chem 248: 378-380., 1973. 27.  Ceccarelli F, Scavuzzo MC, Giusti L, Bigini G, Costa B, Carnicelli V, Zucchi  R, Lucacchini A, and Mazzoni MR. ETA receptor-mediated Ca2+ mobilisation in H9c2 cardiac cells. Biochem Pharmacol 65: 783-793,2003. 28.  Chen QM, Tu VC, Wu Y, and Bahl JJ. Hydrogen peroxide dose dependent  induction of cell death or hypertrophy in cardiomyocytes. Arch Biochem Biophys 373: 242-248., 2000.  109  29.  Chen TM, Goodwin GW, Guthrie PH, and Taegtmeyer H. Effects of insulin  on glucose uptake by rat hearts during and after coronary flow reduction. American Journal of Physiology 273: H2170-2177,1997.  30.  Cheung PC, Salt IP, Davies SP, Hardie DG, and Carling D. Characterization  of AMP-activated protein kinase gamma-subunit isoforms and their role in AMP binding. Biochem J 346 Pt 3: 659-669., 2000. 31.  Cook GA and Lappi MD. Carnitine palmitoyltransferase in the heart is  controlled by a different mechanism than the hepatic enzyme. Mol Cell Biochem 116: 39-45., 1992. 32.  Coven DL, Hu X, Cong L, Bergeron R, Shulman GI, Hardie DG, and Young  LH. Physiological role of AMP-activated protein kinase in the heart: graded activation during exercise. Am J Physiol Endocrinol Metab 285: E629-636, 2003. 33.  Crute BE, Seefeld K, Gamble J, Kemp BE, and Witters LA. Functional  domains of the alphal catalytic subunit of the AMP-activated protein kinase. J Biol Chem 273: 35347-35354., 1998. 34.  Dangel V, Giray J, Ratge D, and Wisser H. Regulation of beta-adrenoceptor  density and mRNA levels in the rat heart cell-line H9c2. Biochem J 317: 925-931., 1996. 35.  Davies SP, Helps NR, Cohen PT, and Hardie DG. 5'-AMP inhibits  dephosphorylation, as well as promoting phosphorylation, of the AMP-activated protein kinase. Studies using bacterially expressed human protein phosphatase-2C alpha and native bovine protein phosphatase-2AC. FEBS Lett 377:421-425., 1995.  110  36.  Doenst T, Guthrie PH, and Taegtmeyer H. Ischemic preconditioning in rat  heart: no correlation between glycogen content and return of function. Molecular & Cellular Biochemistry 180:153-161,1998.  37.  Doenst T, Richwine RT, Bray MS, Goodwin GW, Frazier OH, and  Taegtmeyer H. Insulin improves functional and metabolic recovery of reperfused working rat heart. Ann Thorac Surg 67:1682-1688., 1999. 38.  Duseja R and Feldman JA. Missed acute cardiac ischemia in the ED:  limitations of diagnostic testing. Am J Emerg Med 22: 219-225,2004. 39.  Dyck JR, Cheng JF, Stanley WC, Barr R, Chandler MP, Brown S, Wallace  D, Arrhenius T, Harmon C, Yang G, Nadzan AM, and Lopaschuk GD. Malonyl coenzyme a decarboxylase inhibition protects the ischemic heart by inhibiting fatty acid oxidation and stimulating glucose oxidation. Circ Res 94: e78-84, 2004. 40.  Dyck JR and Lopaschuk GD. Glucose metabolism, H+ production and  Na+/H+-exchanger mRNA levels in ischemic hearts from diabetic rats. Mol Cell Biochem 180: 85-93,1998. 41.  Dyck JR and Lopaschuk GD. Malonyl CoA control of fatty acid oxidation in  the ischemic heart. J Mol Cell Cardiol 34:1099-1109., 2002. 42.  El-Ani D and Zimlichman R. TNFalpha stimulated ATP-sensitive potassium  channels and attenuated deoxyglucose and Ca uptake of H9c2 cardiomyocytes. Ann N YAcad Sci 1010: 716-720,2003.  Ill  43.  Eisner P, Quistorff B, Hermann TS, Dich J, and Grunnet N. Regulation of  glycogen accumulation in L6 myotubes cultured under optimized differentiation conditions. Am J Physiol 275: E925-933., 1998. 44.  Fath-Ordoubadi F and Beatt KJ. Glucose-insulin-potassium therapy for  treatment of acute myocardial infarction: an overview of randomized placebocontrolled trials. Circulation 96:1152-1156., 1997. 45.  Fell DA and Sauro HM. Metabolic control and its analysis. Additional  relationships between elasticities and control coefficients. Eur J Biochem 148: 555561., 1985. 46.  Florini JR, Ewton DZ, Evinger-Hodges MJ, Falen SL, Lau RL, Regan JF,  and Vertel BM. Stimulation and inhibition of myoblast differentiation by hormones. In Vitro 20: 942-958,1984. 47.  Fraser H, Davidge ST, and Clanachan AS. Enhancement of post-ischemic  myocardial function by chronic 17 beta - estradiol treatment: role of alterations in glucose metabolism. / Mol Cell Cardiol 31:1539-1549., 1999. 48.  Fraser H, Lopaschuk GD, and Clanachan AS. Alteration of glycogen and  glucose metabolism in ischaemic and post-ischaemic working rat hearts by adenosine A l receptor stimulation. British Journal of Pharmacology 128:197-205, 1999. 49.  Fraser H, Lopaschuk GD, and Clanachan AS. Assessment of glycogen  turnover in aerobic, ischemic, and reperfused working rat hearts. American Journal of Physiology 275: H1533-1541,1998.  112  50.  Frederich M and Balschi JA. The relationship between AMP-activated  protein kinase activity and AMP concentration in the isolated perfused rat heart J Biol Chem 277:1928-1932., 2002. 51.  Gamble J and Lopaschuk GD. Insulin inhibition of 5' adenosine  monophosphate-activated protein kinase in the heart results in activation of acetyl coenzyme A carboxylase and inhibition of fatty acid oxidation. Metabolism: Clinical & Experimental 46:1270-1274,1997. 52.  Gertz EW, Wisneski JA, Neese R, Houser A, Korte R, and Bristow JD.  Myocardial lactate extraction: multi-determined metabolic function. Circulation 61: 256-261., 1980. 53.  Gertz EW, Wisneski JA, Stanley WC, and Neese RA. Myocardial substrate  utilization during exercise in humans. Dual carbon- labeled carbohydrate isotope experiments. J Clin Invest 82: 2017-2025., 1988. 54.  Gollob MH. Glycogen storage disease as a unifying mechanism of disease in  the PRKAG2 cardiac syndrome. Biochem Soc Trans 31: 228-231., 2003. 55.  Goodwin GW, Ahmad F, Doenst T, and Taegtmeyer H. Energy provision  from glycogen, glucose, and fatty acids on adrenergic stimulation of isolated working rat hearts. American Journal of Physiology 274: H1239-1247,1998. 56.  Goodwin GW, Arteaga JR, and Taegtmeyer H. Glycogen turnover in the  isolated working rat heart. J Biol Chem 270: 9234-9240., 1995. 57.  Goodwin GW and Taegtmeyer H. Metabolic recovery of isolated working rat  heart after brief global ischemia. Am J Physiol 267: H462-470., 1994.  113  58.  Goodwin GW, Taylor CS, and Taegtmeyer H. Regulation of energy  metabolism of the heart during acute increase in heart work. J Biol Chem 273: 29530-29539., 1998. 59.  Guan K, Furst DO, and Wobus AM. Modulation of sarcomere organization  during embryonic stem cell-derived cardiomyocyte differentiation. Eur J Cell Biol 78: 813-823,1999. 60.  Guinovart JJ, Salavert A, Massague J, Ciudad CJ, Salsas E, and Itarte E.  Glycogen synthase: a new activity ratio assay expressing a high sensitivity to the phosphorylation state. FEBS Lett 106: 284-288., 1979. 61.  Guth BD, Wisneski JA, Neese RA, White FC, Heusch G, Mazer CD, and  Gertz EW. Myocardial lactate release during ischemia in swine. Relation to regional bloodflow.Circulation 81:1948-1958., 1990. 62.  Halse R, Fryer LG, McCormack JG, Carling D, and Yeaman SJ. Regulation  of glycogen synthase by glucose and glycogen: a possible role for AMP-activated protein kinase. Diabetes 52: 9-15., 2003. 63.  Hansford RG and Cohen L. Relative importance of pyruvate dehydrogenase  interconversion and feed- back inhibition in the effect of fatty acids on pyruvate oxidation by rat heart mitochondria. Arch Biochem Biophys 191: 65-81., 1978. 64.  Hardie DG and Carling D. The AMP-activated protein kinase-fuel gauge of  the mammalian cell? Eur J Biochem 246: 259-273., 1997.  114  65.  Hardie DG, Salt IP, Hawley S A, and Davies SP. AMP-activated protein  kinase: an ultrasensitive system for monitoring cellular energy charge. Biochem J 338: 717-722., 1999. 66.  Henning SL, Wambolt RB, Schonekess BO, Lopaschuk GD, and Allard MF.  Contribution of glycogen to aerobic myocardial glucose utilization. Circulation 93: 1549-1555., 1996. 67.  Hescheler J, Meyer R, Plant S, Krautwurst D, Rosenthal W, and Schultz G.  Morphological, biochemical, and electrophysiological characterization of a clonal cell (H9c2) line from rat heart. Circ Res 69:1476-1486., 1991. 68.  Higgins AJ, Morville M, Burges RA, Gardiner DG, Page MG, and Blackburn  KJ. Oxfenicine diverts rat muscle metabolism from fatty acid to carbohydrate oxidation and protects the ischaemic rat heart. Life Sci 27: 963-970., 1980. 69.  Hoch B, Haase H, Schulze W, Hagemann D, Morano I, Krause EG, and  Karczewski P. Differentiation-dependent expression of cardiac delta-CaMKII isoforms. / Cell Biochem 68: 259-268., 1998. 70.  Holmes BF, Kurth-Kraczek EJ, and Winder WW. Chronic activation of 5'-  AMP-activated protein kinase increases GLUT-4, hexokinase, and glycogen in muscle. JAppl Physiol 87:1990-1995., 1999. 71.  Hopkins TA, Dyck JR, and Lopaschuk GD. AMP-activated protein kinase  regulation of fatty acid oxidation in the ischaemic heart. Biochem Soc Trans 31: 207212., 2003.  115  72.  Hue L, Beauloye C, Marsin AS, Bertrand L, Herman S, and Rider MH.  Insulin and ischemia stimulate glycolysis by acting on the same targets through different and opposing signaling pathways. J Mol Cell Cardiol 34:1091-1097., 2002. 73.  Ingwall JS. Is creatine kinase a target for AMP-activated protein kinase in  the heart? J Mol Cell Cardiol 34:1111-1120., 2002. 74.  Janero DR, Burghardt C, and Feldman D. Amphiphile-induced heart  muscle-cell (myocyte) injury: effects of intracellular fatty acid overload. J Cell Physiol 137:1-13., 1988. 75.  Jaspers SR, Garnache AK, and Miller TB, Jr. Factors affecting the  activation of glycogen synthase in primary culture cardiomyocytes. J Mol Cell Cardiol 25:1171-1178., 1993. 76.  Javaux F, Vincent MF, Wagner DR, and van den Berghe G. Cell-type  specificity of inhibition of glycolysis by 5-amino-4- imidazolecarboxamide riboside. Lack of effect in rabbit cardiomyocytes and human erythrocytes, and inhibition in FTO-2B rat hepatoma cells. Biochem J 305: 913-919., 1995. 77.  Jeremy RW, Koretsune Y, Marban E, and Becker LC. Relation between  glycolysis and calcium homeostasis in postischemic myocardium. Circ Res 70:11801190., 1992. 78.  Kageyama K, Ihara Y, Goto S, Urata Y, Toda G, Yano K, and Kondo T.  Overexpression of calreticulin modulates protein kinase B/Akt signaling to promote apoptosis during cardiac differentiation of cardiomyoblast H9c2 cells. / Biol Chem 277:19255-19264,2002.  116  79.  Kantor PF, Dyck JR, and Lopaschuk GD. Fatty acid oxidation in the  reperfused ischemic heart. American Journal of the Medical Sciences 318: 3-14,  1999. 80.  Kantor PF, Lucien A, Kozak R, and Lopaschuk GD. The antianginal drug  trimetazidine shifts cardiac energy metabolism from fatty acid oxidation to glucose oxidation by inhibiting mitochondrial long-chain 3-ketoacyl coenzyme A thiolase [see comments]. Circulation Research 86: 580-588,2000. 81.  Kashiwaya Y, Sato K, Tsuchiya N, Thomas S, Fell DA, Veech RL, and  Passonneau JV. Control of glucose utilization in working perfused rat heart. J Biol Chem 269: 25502-25514., 1994. 82.  Katz J, Golden S, and Wals PA. Stimulation of hepatic glycogen synthesis by  amino acids. Proc Natl Acad Sci USA73:  83.  3433-3437., 1976.  Kerbey AL, Randle PJ, Cooper RH, Whitehouse S, Pask HT, and Denton  RM. Regulation of pyruvate dehydrogenase in rat heart. Mechanism of regulation of proportions of dephosphorylated and phosphorylated enzyme by oxidation of fatty acids and ketone bodies and of effects of diabetes: role of coenzyme A, acetylcoenzyme A and reduced and oxidized nicotinamide-adenine dinucleotide. Biochem J154: 327-348., 1976. 84.  Kerner J, Zaluzec E, Gage D, and Bieber LL. Characterization of the  malonyl-CoA-sensitive carnitine palmitoyltransferase (CPTo) of a rat heart mitochondrial particle. Evidence that the catalytic unit is CPTi. J Biol Chem 269: 8209-8219., 1994.  117  85.  Kim JM, Yoon M, Kang I, Kim SS, and Ha J. Evidence that acetyl-CoA  carboxylase isoforms play different biological roles in H9c2 cardiomyocyte. Biochem Biophys Res Commun 248: 490-496., 1998.  86.  Kimes BW and Brandt BL. Properties of a clonal muscle cell line from rat  heart. Exp Cell Res 98: 367-381., 1976. 87.  King L M and Opie LH. Glucose delivery is a major determinant of glucose  utilisation in the ischemic myocardium with a residual coronary flow. Cardiovasc Res 39: 381-392., 1998. 88.  Klip A, Li G, and Logan WJ. Induction of sugar uptake response to insulin  by serum depletion in fusing L6 myoblasts. Am J Physiol 247: E291-296,1984. 89.  Koivisto UM, Martinez-Valdez H, Bilan PJ, Burdett E, Ramlal T, and Klip  A. Differential regulation of the GLUT-1 and GLUT-4 glucose transport systems by glucose and insulin in L6 muscle cells in culture. J Biol Chem 266: 2615-2621., 1991. 90.  Kovacic S, Soltys CL, Barr AJ, Shiojima I, Walsh K, and Dyck JR. Akt  activity negatively regulates phosphorylation of AMP-activated protein kinase in the heart. / Biol Chem 278: 39422-39427, 2003. 91.  Kubo Y. Comparison of initial stages of muscle differentiation in rat and  mouse myoblastic and mouse mesodermal stem cell lines. J Physiol 442: 743-759, 1991. 92.  Kudo N, Barr AJ, Barr RL, Desai S, and Lopaschuk GD. High rates of fatty  acid oxidation during reperfusion of ischemic hearts are associated with a decrease  118  in malonyl-CoA levels due to an increase in 5'-AMP-activated protein kinase inhibition of acetyl-CoA carboxylase. / Biol Chem 270:17513-17520., 1995. 93.  Kudo N, Gillespie JG, Kung L, Witters LA, Schulz R, Clanachan AS, and  Lopaschuk GD. Characterization of 5'AMP-activated protein kinase activity in the heart and its role in inhibiting acetyl-CoA carboxylase during reperfusion following ischemia. Biochim Biophys Acta 1301: 67-75., 1996.  94.  Kusuoka H and Marban E. Mechanism of the diastolic dysfunction induced  by glycolytic inhibition. Does adenosine triphosphate derived from glycolysis play a favored role in cellular Ca2+ homeostasis in ferret myocardium? / Clin Invest 93: 1216-1223., 1994. 95.  LaDu MJ and Palmer WK. Expression of lipoprotein lipase during  differentiation of cultured L6 muscle cells. Can J Physiol Pharmacol 72: 243-247., 1994. 96.  Lassers BW, Kaijser L, and Carlson LA. Myocardial lipid and carbohydrate  metabolism in healthy, fasting men at rest: studies during continuous infusion of 3 H-palmitate. Eur J Clin Invest 2: 348-358., 1972. 97.  Lavanchy N, Grably S, Garnier A, and Rossi A. Crucial role of intracellular  effectors on glycogenolysis in the isolated rat heart: potential consequences on the myocardial tolerance to ischemia. Mol Cell Biochem 160-161: 273-282., 1996. 98.  Lawson MA and Purslow PP. Differentiation of myoblasts in serum-free  media: effects of modified media are cell line-specific. Cells Tissues Organs 167:130137,2000.  119  99.  Laybutt DR, Thompson AL, Cooney GJ, and Kraegen EW. Selective chronic  regulation of GLUT1 and GLUT4 content by insulin, glucose, and lipid in rat cardiac muscle in vivo. Am J Physiol 273: H1309-1316., 1997. 100.  Lee CD, Folsom AR, Pankow JS, and Brancati FL. Cardiovascular events in  diabetic and nondiabetic adults with or without history of myocardial infarction. Circulation 109: 855-860,2004. 101.  Lee JK and Kim KH. Roles of acetyl-CoA carboxylase beta in muscle cell  differentiation and in mitochondrial fatty acid oxidation. Biochem Biophys Res Commun 254: 657-660., 1999. 102.  Lemieux K, Konrad D, Klip A, and Marette A. The AMP-activated protein  kinase activator AICAR does not induce GLUT4 translocation to transverse tubules but stimulates glucose uptake and p38 mitogen-activated protein kinases alpha and beta in skeletal muscle. Faseb J17:1658-1665, 2003. 103.  Li J, Hu X, Selvakumar P, Russell RR, 3rd, Cushman SW, Holman GD, and  Young LH. Role of the nitric oxide pathway in AMPK-mediated glucose uptake and GLUT4 translocation in heart muscle. Am J Physiol Endocrinol Metab 287: E834841,2004. 104.  Liedtke AJ. Alterations of carbohydrate and lipid metabolism in the acutely  ischemic heart. Prog Cardiovasc Dis 23: 321-336., 1981. 105.  Liedtke AJ, DeMaison L, Eggleston AM, Cohen LM, and Nellis SH. Changes  in substrate metabolism and effects of excess fatty acids in reperfused myocardium. Circ Res 62: 535-542., 1988.  120  106.  Liedtke AJ, Nellis S, and Neely JR. Effects of excess free fatty acids on  mechanical and metabolic function in normal and ischemic myocardium in swine. Circ Res 43: 652-661., 1978. 107.  Liedtke AJ, Nellis SH, and Mjos OD. Effects of reducing fatty acid  metabolism on mechanical function in regionally ischemic hearts. Am J Physiol 247: H387-394., 1984. 108.  Lilly LS and Harvard Medical School. Pathophysiology of heart disease: a  collaborative project of medical students and faculty. Baltimore: Williams & Wilkins,  1998. 109.  Liu J and Brautigan DL. Glycogen synthase association with the striated  muscle glycogen- targeting subunit of protein phosphatase-1. Synthase activation involves scaffolding regulated by beta-adrenergic signaling. J Biol Chem 275: 2607426081., 2000. 110.  Lokuta A, Kirby MS, Gaa ST, Lederer WJ, and Rogers TB. On establishing  primary cultures of neonatal rat ventricular myocytes for analysis over long periods. / Cardiovasc Electrophysiol 5: 50-62., 1994. 111.  Longnus SL, Wambolt RB, Barr RL, Lopaschuk GD, and Allard MF.  Regulation of myocardial fatty acid oxidation by substrate supply. Am J Physiol Heart Circ Physiol 281: H1561-1567., 2001.  112.  Longnus SL, Wambolt RB, Parsons HL, Brownsey RW, and Allard MF. 5-  Aminoimidazole-4-carboxamide 1-beta -D-ribofuranoside (AICAR) stimulates  121  myocardial glycogenolysis by allosteric mechanisms. Am J Physiol Regul Integr Comp Physiol 284: R936-R944., 2003. 113.  Lopaschuk GD, Belke DD, Gamble J, Itoi T, and Schonekess BO. Regulation  of fatty acid oxidation in the mammalian heart in health and disease. Biochim BiophysActa 1213: 263-276., 1994. 114.  Lopaschuk GD, Spafford MA, Davies NJ, and Wall SR. Glucose and  palmitate oxidation in isolated working rat hearts reperfused after a period of transient global ischemia. Circ Res 66: 546-553., 1990. 115.  Lopaschuk GD, Wambolt RB, and Barr RL. An imbalance between  glycolysis and glucose oxidation is a possible explanation for the detrimental effects of high levels of fatty acids during aerobic reperfusion of ischemic hearts. J Pharmacol Exp Ther 264:135-144., 1993.  116.  Luiken JJ, Coort SL, Willems J, Coumans WA, Bonen A, van der Vusse GJ,  and Glatz JF. Contraction-induced fatty acid translocase/CD36 translocation in rat cardiac myocytes is mediated through AMP-activated protein kinase signaling. Diabetes 52:1627-1634, 2003. 117.  Luque MA, Gonzalez N, Marquez L, Acitores A, Redondo A, Morales M,  Valverde I, and Villanueva-Penacarrillo ML. Glucagon-like peptide-1 (GLP-1) and glucose metabolism in human myocytes. / Endocrinol 173: 465-473., 2002. 118.  Marsin AS, Bertrand L, Rider MH, Deprez J, Beauloye C, Vincent MF, Van  den Berghe G, Carling D, and Hue L. Phosphorylation and activation of heart PFK-  122  2 by AMPK has a role in the stimulation of glycolysis during ischaemia. Curr Biol 10:1247-1255., 2000. 119.  McCormack JG, Halestrap AP, and Denton RM. Role of calcium ions in  regulation of mammalian intramitochondrial metabolism. Physiol Rev 70: 391-425., 1990. 120.  McGarry JD, Mills SE, Long CS, and Foster DW. Observations on the  affinity for carnitine, and malonyl-CoA sensitivity, of carnitine palmitoyltransferase I in animal and human tissues. Demonstration of the presence of malonyl-CoA in non-hepatic tissues of the rat. Biochem J 214: 21-28., 1983. 121.  McNulty PH, Sinusas AJ, Shi CQ, Dione D, Young LH, Cline GC, and  Shulman GI. Glucose metabolism distal to a critical coronary stenosis in a canine model of low-flow myocardial ischemia. J Clin Invest 98: 62-69., 1996. 122.  Mejia-Alvarez R, Tomaselli GF, and Marban E. Simultaneous expression of  cardiac and skeletal muscle isoforms of the L-type Ca2+ channel in a rat heart muscle cell line. / Physiol 478: 315-329., 1994. 123.  Melendez R, Melendez-Hevia E, and Cascante M. How did glycogen  structure evolve to satisfy the requirement for rapid mobilization of glucose? A problem of physical constraints in structure building. J Mol Evol 45: 446-455., 1997. 124.  Melendez-Hevia E, Waddell TG, and Shelton ED. Optimization of molecular  design in the evolution of metabolism: the glycogen molecule. Biochem J 295:477483., 1993.  123  125.  Menard C, Pupier S, Mornet D, Kitzmann M, Nargeot J, and Lory P.  Modulation of L-type calcium channel expression during retinoic acid- induced differentiation of H9C2 cardiac cells. J Biol Chem 274: 29063-29070., 1999. 126.  Michell BJ, Stapleton D, Mitchelhill KI, House CM, Katsis F, Witters LA,  and Kemp BE. Isoform-specific purification and substrate specificity of the 5'-AMPactivated protein kinase. / Biol Chem 271: 28445-28450,1996. 127.  Milan D, Jeon JT, Looft C, Amarger V, Robic A, Thelander M, Rogel-  Gaillard C, Paul S, Iannuccelli N, Rask L, Ronne H, Lundstrom K, Reinsch N, Gellin J, Kalm E, Roy PL, Chardon P, and Andersson L. A mutation in PRKAG3 associated with excess glycogen content in pig skeletal muscle. Science 288:12481251., 2000. 128.  Millart H and Seraydarian MW. Influence of plating density on individual  cell growth, cell division and differentiation of neonatal rat heart primary cultures. Tissue Cell 18: 209-218,1986.  129.  Minotti S, Scicchitano BM, Nervi C, Scarpa S, Lucarelli M, Molinaro M, and  Adamo S. Vasopressin and insulin-like growth factors synergistically induce myogenesis in serum-free medium. Cell Growth Differ 9:155-163,1998. 130.  Mitsumoto Y, Burdett E, Grant A, and Klip A. Differential expression of the  GLUT1 and GLUT4 glucose transporters during differentiation of L6 muscle cells. Biochem Biophys Res Commun 175: 652-659., 1991.  124  131.  Mizukami Y, Kobayashi S, Uberall F, Hellbert K, Kobayashi N, and Yoshida  K. Nuclear mitogen-activated protein kinase activation by protein kinase czeta during reoxygenation after ischemic hypoxia. / Biol Chem 275:19921-19927., 2000. 132.  Neely JR and Grotyohann LW. Role of glycolytic products in damage to  ischemic myocardium. Dissociation of adenosine triphosphate levels and recovery of function of reperfused ischemic hearts. Circ Res 55: 816-824., 1984. 133.  Neely JR, Rovetto MJ, and Oram JF. Myocardial utilization of carbohydrate  and lipids. Prog Cardiovasc Dis 15: 289-329., 1972. 134.  Nielsen JN, Derave W, Kristiansen S, Ralston £ , Ploug T, and Richter EA.  Glycogen synthase localization and activity in rat skeletal muscle is strongly dependent on glycogen content. / Physiol 531: 757-769., 2001. 135.  Nielsen JN, Wojtaszewski JF, Haller RG, Hardie DG, Kemp BE, Richter EA,  and Vissing J. Role of 5'AMP-activated protein kinase in glycogen synthase activity and glucose utilization: insights from patients with McArdle's disease. J Physiol 541: 979-989., 2002. 136.  Opie LH. The heart: physiology, from cell to circulation. Philadelphia:  Lippincott-Raven, 1998. 137.  Pardridge WM, Davidson MB, and Casanello-Ertl D. Glucose and amino  acid metabolism in an established line of skeletal muscle cells. / Cell Physiol 96: 309318., 1978. 138.  Park C, So HS, Shin CH, Baek SH, Moon BS, Shin SH, Lee HS, Lee DW, and  Park R. Quercetin protects the hydrogen peroxide-induced apoptosis via inhibition  125  of mitochondrial dysfunction in H9c2 cardiomyoblast cells. Biochem Pharmacol 66: 1287-1295,2003. 139.  Persky AM, Green PS, Stubley L, Howell CO, Zaulyanov L, Brazeau GA,  and Simpkins JW. Protective effect of estrogens against oxidative damage to heart and skeletal muscle in vivo and in vitro. Proc Soc Exp Biol Med 223: 59-66., 2000. 140.  Pinset C and Whalen RG. Manipulation of medium conditions and  differentiation in the rat myogenic cell line L6. Dev Biol 102: 269-277., 1984. 141.  Portier GL, Benders AG, Oosterhof A, Veerkamp JH, and van Kuppevelt  TH. Differentiation markers of mouse C2C12 and rat L6 myogenic cell lines and the effect of the differentiation medium. In Vitro Cell Dev BiolAnim 35: 219-227., 1999. 142.  Randle PJ. Fuel selection in animals. Biochem Soc Trans 14: 799-806., 1986.  143.  Randle PJ, Garland PB, Hales CN, and Newsholme EA. The glucose fatty-  acid cycle: its role in insulin sensitivity and the metabolic disturbances of diabetes mellitus. Lancet i: 785-789,1963. 144.  Rett K, Wicklmayr M, Dietze GJ, and Haring HU. Insulin-induced glucose  transporter (GLUT1 and GLUT4) translocation in cardiac muscle tissue is mimicked by bradykinin. Diabetes 45 Suppl 1: S66-69,1996. 145.  Richler C and Yaffe D. The in vitro cultivation and differentiation capacities  of myogenic cell lines. Dev Biol 23:1-22., 1970. 146.  Russell RR, 3rd, Bergeron R, Shulman GI, and Young LH. Translocation of  myocardial GLUT-4 and increased glucose uptake through activation of AMPK by AICAR. Am J Physiol 277: H643-649., 1999.  126  147.  Russell RR, 3rd, Li J, Coven DL, Pypaert M, Zechner C, Palmeri M,  Giordano FJ, Mu J, Birnbaum MJ, and Young LH. AMP-activated protein kinase mediates ischemic glucose uptake and prevents postischemic cardiac dysfunction, apoptosis, and injury. J Clin Invest 114: 495-503,2004. 148.  Russell RR, 3rd, Yin R, Caplan MJ, Hu X, Ren J, Shulman GI, Sinusas AJ,  and Young LH. Additive effects of hyperinsulinemia and ischemia on myocardial GLUT1 and GLUT4 translocation in vivo. Circulation 98: 2180-2186., 1998. 149.  Saddik M, Gamble J, Witters LA, and Lopaschuk GD. Acetyl-CoA  carboxylase regulation of fatty acid oxidation in the heart. J Biol Chem 268: 2583625845., 1993. 150.  Saddik M and Lopaschuk GD. Myocardial triglyceride turnover and  contribution to energy substrate utilization in isolated working rat hearts. J Biol Chem 266: 8162-8170., 1991. 151.  Saddik M and Lopaschuk GD. Myocardial triglyceride turnover during  reperfusion of isolated rat hearts subjected to a transient period of global ischemia. J Biol Chem 267: 3825-3831., 1992.  152.  Saha AK and Ruderman NB. Malonyl-CoA and AMP-activated protein  kinase: an expanding partnership. Mol Cell Biochem 253: 65-70,2003. 153.  Schonekess BO, Allard MF, Henning SL, Wambolt RB, and Lopaschuk GD.  Contribution of glycogen and exogenous glucose to glucose metabolism during ischemia in the hypertrophied rat heart. Circulation Research 81: 540-549,1997.  127  154.  Schroedl NA, Hartzell CR, Ross PD, and McCarl RL. Glucose metabolism,  insulin effects, and developmental age of cultured neonatal rat heart cells. / Cell Physiol 113: 231-239., 1982. 155.  Schubert D, Tarikas H, Humphreys S, Heinemann S, and Patrick J. Protein  synthesis and secretion in a myogenic cell line. Dev Biol 33:18-37., 1973. 156.  Shearer J and Graham TE. New perspectives on the storage and  organization of muscle glycogen. Can J Appl Physiol 27:179-203., 2002. 157.  Shulman RG and Rothman DL. The "glycogen shunt" in exercising muscle:  A role for glycogen in muscle energetics and fatigue. Proc Natl Acad Sci USA 98: 457-461., 2001. 158.  Soares PR, de Albuquerque CP, Chacko VP, Gerstenblith G, and Weiss RG.  Role of preischemic glycogen depletion in the improvement of postischemic metabolic and contractile recovery of ischemia- preconditioned rat hearts. Circulation 96: 975-983., 1997. 159.  Sodi-Pollares D, Testelli MR, Fishleder BL, Bisteni A, Medrano GA, and  DeMicheli A. Effects of an intravenous infusion of a potassium-glucose-insulin solution on teh electrocardiographic signs of myocardial infarction. Am J Cardiol 9: 166-181,1962. 160.  Stanley WC. Myocardial lactate metabolism during exercise. Med Sci Sports  Exerc 23: 920-924., 1991. 161.  Stanley WC and Chandler MP. Energy metabolism in the normal and failing  heart: potential for therapeutic interventions. Heart Fail Rev 7:115-130., 2002.  128  162.  Stapleton D, Mitchelhill KI, Gao G, Widmer J, Michell BJ, Teh T, House  CM, Fernandez CS, Cox T, Witters LA, and Kemp BE. Mammalian AMP-activated protein kinase subfamily. J Biol Chem 271: 611-614., 1996. 163.  Stapleton D, Woollatt E, Mitchelhill KI, Nicholl JK, Fernandez CS, Michell  BJ, Witters LA, Power DA, Sutherland GR, and Kemp BE. AMP-activated protein kinase isoenzyme family: subunit structure and chromosomal location. FEBS Lett 409: 452-456., 1997. 164.  Stein SC, Woods A, Jones NA, Davison MD, and Carling D. The regulation  of AMP-activated protein kinase by phosphorylation. Biochem J 345 Pt 3: 437-443., 2000. 165.  Steinberg D and Khoo JC. Hormone-sensitive lipase of adipose tissue. Fed  Proc 36:1986-1990., 1977. 166.  Su CY, Chong KY, Chen J, Ryter S, Khardori R, and Lai CC. A  physiologically relevant hyperthermia selectively activates constitutive hsp70 in H9c2 cardiac myoblasts and confers oxidative protection. J Mol Cell Cardiol 31: 845-855., 1999. 167.  Sullivan JE, Brocklehurst KJ, Marley AE, Carey F, Carling D, and Beri RK.  Inhibition of lipolysis and lipogenesis in isolated rat adipocytes with AICAR, a cellpermeable activator of AMP-activated protein kinase. FEBS Lett 353: 33-36,1994. 168.  Sutherland FJ and Hearse DJ. The isolated blood and perfusion fluid  perfused heart. Pharmacol Res 41: 613-627., 2000.  129  169.  Taegtmeyer H. Carbohydrate interconversions and energy production.  Circulation 72: IV1-8., 1985. 170.  Taegtmeyer H. Energy metabolism of the heart: from basic concepts to  clinical applications. CurrProbl Cardiol 19: 59-113., 1994. 171.  Taegtmeyer H, Goodwin GW, Doenst T, and Frazier OH. Substrate  metabolism as a determinant for postischemic functional recovery of the heart. American Journal of Cardiology 80: 3A-10A, 1997.  172.  Tanaka H, Sakurai K, Takahashi K, and Fujimoto Y. Requirement of  intracellular free thiols for hydrogen peroxide-induced hypertrophy in cardiomyocytes. / Cell Biochem 89: 944-955,2003. 173.  Thornton C, Snowden MA, and Carling D. Identification of a novel AMP-  activated protein kinase beta subunit isoform that is highly expressed in skeletal muscle. / Biol Chem 273:12443-12450., 1998. 174.  Tracey WR, Treadway JL, Magee WP, Sutt JC, McPherson RK, Levy CB,  Wilder DE, Yu LJ, Chen Y, Shanker RM, Mutchler AK, Smith AH, Flynn DM, and Knight DR. Cardioprotective effects of ingliforib, a novel glycogen phosphorylase inhibitor. Am J Physiol Heart Circ Physiol 286: H1177-1184,2004.  175.  Turo KA and Florini JR. Hormonal stimulation of myoblast differentiation  in the absence of DNA synthesis. Am J Physiol 243: C278-284., 1982. 176.  Uphues I, Kolter T, Goud B, and Eckel J. Insulin-induced translocation of  the glucose transporter GLUT4 in cardiac muscle: studies on the role of smallmolecular-mass GTP-binding proteins. Biochem J 301 (Pt 1): 177-182,1994.  130  177.  van Bilsen M, van der Vusse GJ, Snoeckx LH, Arts T, Coumans WA,  Willemsen PH, and Reneman RS. Effects of pyruvate on post-ischemic myocardial recovery at various workloads. PflugersArch 413:167-173., 1988. 178.  van den Eijnde SM, van den Hoff MJ, Reutelingsperger CP, van Heerde WL,  Henfling ME, Vermerj-Keers C, Schutte B, Borgers M, and Ramaekers FC. Transient expression of phosphatidylserine at cell-cell contact areas is required for myotube formation. J Cell Sci 114: 3631-3642., 2001. 179.  van der Vusse GJ, van Bilsen M, and Glatz JF. Cardiac fatty acid uptake and  transport in health and disease. Cardiovasc Res 45: 279-293., 2000. 180.  Van Nieuwenhoven FA, Luiken JJ, De Jong YF, Grimaldi PA, Van der  Vusse GJ, and Glatz JF. Stable transfection of fatty acid translocase (CD36) in a rat heart muscle cell line (H9c2). / Lipid Res 39: 2039-2047., 1998. 181.  Velez C, Aranega AE, Fernandez JE, Melguizo C, Alvarez L, and Aranega A.  Modulation of contractile protein troponin-T in chick myocardial cells by catecholamines during development. Cell Mol Biol (Noisy-le-grand) 40:1189-1199, 1994. 182.  Velez C, Aranega AE, Melguizo C, Fernandez JE, Prados J, and Aranega A.  Modulation of contractile protein troponin-T in chick myocardial cells by basic fibroblast growth factor and platelet-derived growth factor during development. J Cardiovasc Pharmacol 24: 906-913,1994.  131  183.  Vogt AM, Elsasser A, Nef H, Bode C, Kubler W, and Schaper J. Increased  glycolysis as protective adaptation of energy depleted, degenerating human hibernating myocardium. Mol Cell Biochem 242:101-107., 2003. 184.  Wahrmann JP, Drugeon G, Delain E, and Delain D. Gene expression during  the differentiation of myogenic cells of the L6 line. Biochimie 58: 551-562., 1976. 185.  Wahrmann JP, Gros F, and Luzzati D. Phosphorylase and glycogen  synthetase during myoblast differentiation. Biochimie 55: 457-463., 1973. 186.  Wahrmann JP, Recouvreur M, and Favard-Sereno C. Development and  regulation of the phosphorylase-glycogen complex in myogenic cells of the L6 line. J Cell Sci 26: 77-91., 1977. 187.  Walker PS, Ramlal T, Sarabia V, Koivisto UM, Bilan PJ, Pessin JE, and Klip  A. Glucose transport activity in L6 muscle cells is regulated by the coordinate control of subcellular glucose transporter distribution, biosynthesis, and mRNA transcription. J Biol Chem 265:1516-1523,1990. 188.  Wambolt RB, Henning SL, English DR, Bondy GP, and Allard MF.  Regression of cardiac hypertrophy normalizes glucose metabolism and left ventricular function during reperfusion. Journal of Molecular & Cellular Cardiology 29: 939-948,1997. 189.  Wambolt RB, Henning SL, English DR, Dyachkova Y, Lopaschuk GD, and  Allard MF. Glucose utilization and glycogen turnover are accelerated in hypertrophied rat hearts during severe low-flow ischemia. Journal of Molecular & Cellular Cardiology 31: 493-502,1999.  132  190.  Webster I, Huisamen B, and Lochner A. The effect of aicar and ZMP on  myocardial glucose uptake. Cardiovasc J S Afr 15: S10,2004. 191.  Weiss J and Hiltbrand B. Functional compartmentation of glycolytic versus  oxidative metabolism in isolated rabbit heart. J Clin Invest 75: 436-447., 1985. 192.  Wheeler TJ, Fell RD, and Hauck MA. Translocation of two glucose  transporters in heart: effects of rotenone, uncouplers, workload, palmitate, insulin and anoxia. Biochim Biophys Acta 1196:191-200., 1994.  193.  Wieland O, Funcke H, and Loffler G. Interconversion of pyruvate  dehydrogenase in rat heart muscle upon perfusion with fatty acids or ketone bodies. FEBS Lett 15: 295-298., 1971.  194.  Wieland O, Siess E, Schulze-Wethmar FH, yon Funcke HG, and Winton B.  Active and inactive forms of pyruvate dehydrogenase in rat heart and kidney: effect of diabetes, fasting, and refeeding on pyruvate dehydrogenase interconversion. Arch Biochem Biophys 143: 593-601., 1971.  195.  Wiley C and Beeson C. Continuous measurement of glucose utilization in  heart myoblasts. Anal Biochem 304:139-146., 2002. 196.  Wisneski JA, Gertz EW, Neese RA, Gruenke LD, Morris DL, and Craig JC.  Metabolic fate of extracted glucose in normal human myocardium. J Clin Invest 76: 1819-1827., 1985. 197.  Wisneski JA, Gertz EW, Neese RA, and Mayr M. Myocardial metabolism of  free fatty acids. Studies with 14C-labeled substrates in humans. / Clin Invest 79: 359-366., 1987.  133  198.  Witters LA and Kemp BE. Insulin activation of acetyl-CoA carboxylase  accompanied by inhibition of the 5'-AMP-activated protein kinase. J Biol Chem 267: 2864-2867., 1992. 199.  Wojtaszewski JF, Jorgensen SB, Hellsten Y, Hardie DG, and Richter EA.  Glycogen-dependent effects of 5-aminoimidazole-4-carboxamide (AICA)- riboside on AMP-activated protein kinase and glycogen synthase activities in rat skeletal muscle. Diabetes 51: 284-292., 2002. 200.  Yabe K and Takeo S. Does glycogen depletion play an important role in  ischemic preconditioning? Heart Vessels 12:136-142,1997. 201.  Yaffe D. Retention of differentiation potentialities during prolonged  cultivation of myogenic cells. Proc Natl Acad Sci US A 61: 477-483., 1968. 202.  Yang H, Egan JM, Wang Y, Moyes CD, Roth J, Montrose MH, and  Montrose-Rafizadeh C. GLP-1 action in L6 myotubes is via a receptor different from the pancreatic GLP-1 receptor. Am J Physiol 275: C675-683., 1998. 203.  Young LH, Renfu Y, Russell R, Hu X, Caplan M, Ren J, Shulman GI, and  Sinusas AJ. Low-flow ischemia leads to translocation of canine heart GLUT-4 and GLUT-1 glucose transporters to the sarcolemma in vivo. Circulation 95: 415-422., 1997. 204.  Young ME, Radda GK, and Leighton B. Activation of glycogen  phosphorylase and glycogenolysis in rat skeletal muscle by AICAR~an activator of AMP-activated protein kinase. FEBS Lett 382: 43-47., 1996.  134  205.  Yu B, Poirier LA, and Nagy LE. Mobilization of GLUT-4 from intracellular  vesicles by insulin and K(+) depolarization in cultured H9c2 myotubes. Am J Physiol 277: E259-267., 1999. 206.  Yu B, Schroeder A, and Nagy LE. Ethanol stimulates glucose uptake and  translocation of GLUT-4 in H9c2 myotubes via a Ca(2+)-dependent mechanism. Am J Physiol Endocrinol Metab 279: E1358-1365., 2000.  207.  Ziegler B and Lippmann HG. The effect of insulin on the glycolysis and  glycogen content of beating rat heart cells in the primary culture. Experientia 27: 138-139., 1971.  135  

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