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Cardiac lipoprotein lipase regulation : metabolic basis for diabetic heart disease Chacko, Pulinilkunnil Thomas 2005

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CARDIAC LIPOPROTEIN LIPASE REGULATION: METABOLIC BASIS FOR DIABETIC HEART DISEASE b y PULINILKUNNIL THOMAS CHACKO B.Pharm., Mumbai University, India 1998 M.S., National Institute of Pharmaceutical Education and Research (NIPER), India 2000 A thesis submitted in partial fulfilment of the requirements for the degree of DOCTOR OF PHILOSOPHY in The Faculty of Graduate Studies (Pharmaceutical Sciences) THE UNIVERSITY OF BRITISH COLUMBIA September 2005 © Pulinilkunnil Thomas Chacko, 2005 A B S T R A C T During diabetes, impaired cardiac glucose transport and utilization switches energy production to exclusive R-oxidation of fatty acid (FA). We examined the contribution of cardiac lipoprotein lipase (LPL) towards providing FA to the diabetic heart. Four days of streptozotocin (STZ) or four hours of diazoxide (DZ) induced hyperglycemia enhanced LPL activity at the coronary lumen. This increased enzyme was likely unrelated to an increase in the number of endothelial LPL binding sites and suggests that binding sites for LPL in the control rat heart are partly occupied by the enzyme and diabetes rapidly initiates filling. Phloridzin treatment of STZ animals normalized plasma glucose with no effect on luminal LPL suggesting that the effects of diabetes on LPL are also largely independent of changes in blood glucose and likely involve other circulating mediators. The influence of circulating lipoprotein triglyceride (TG) hydrolysis and their lipolytic byproducts (lysophospholipids) in facilitating LPL translocation of LPL from the underlying cardiomyocyte cell surface to the coronary lumen was evaluated. Exposure of isolated control hearts to lysophosphatidylcholine (LPC) enhanced luminal LPL to levels observed following DZ. Treatment of DZ animals with either WR 1339 or N 6 -cyclopentyladenosine (which inhibit lipolysis) decreased DZ induced augmentation of cardiac LPL. Our studies suggest that increases in LPL likely involve posttranslational processing via breakdown of circulating lipoprotein- T G and a LPC dependent mechanism. LPC maintained high luminal LPL through a protein kinase C (PKC) dependent mechanism and required formation of its metabolic byproduct lysophosphatidic acid (LPA). We examined whether LPL secretion following LPA involves actin cytoskeleton reassembly. Incubation of myocytes with LPA increased basal and heparin releasable (HR-LPL). Incubation of myocytes with cytochalasin D not only blocked LPA induced augmentation of HR-LPL but also abrogated filamentous actin (F actin) formation. Exposure of myocytes to LPA facilitated significant ii membrane translocation of RhoA, and its downstream effector Rho kinase I (ROCK 1), and blocking this effect with Y-27632 appreciably reduced basal and HR-LPL activity. Overall, our data suggest that impaired intracellular glucose utilization allows rapid vectorial transfer of LPL to unoccupied binding sites to supply the diabetic heart with excess FA potentially initiating and sustaining cardiac dysfunction during diabetes. iii T A B L E O F C O N T E N T S ABSTRACT ii TABLE OF CONTENTS iv LIST OF TABLES ix LIST OF FIGURES x LIST OF SYMBOLS xii LIST OF ABBREVIATIONS xiii ACKNOWLEDGMENTS xv DEDICATION xvi LIST OF MANUSCRIPTS xvii 1.0 Introduction 1 1.1 Clinical Problem of Diabetes 1 1.2 Cardiac Energy Metabolism 2 1.2.1 Substrate Utilization 2 1.2.2 Cardiac Metabolism during Diabetes 3 1.2.3 Role ofLPL in Heart Disease 4 1.3 Lipoprotein Lipase 5 1.3.1 Synthesis and Activation 6 13.2 Secretion 7 1.3.3 Transfer of Secreted LPL from Myocyte to Endothelial Cells 7 1.4 Physiological Regulation of LPL 9 1.4.1 Degradation of Intracellular LPL 9 1.4.2 Recycling of Intraendothelial LPL 10 1.4.3 Regulation of LPL by Substrates 10 1.5 Cardiac LPL during Hypoinsulinemia and Insulin Resistance 12 1.5.1 LPL in the STZ Diabetic Heart 12 1.5.2 Cardiac LPL following Insulin Resistance 13 1.6 Research Rationale and Hypothesis 14 1.7 Research Objectives 16 1.8 Figures 17 1.9 Bibliography 19 iv 2.0 Evidence for Rapid Metabolic Switching through Lipoprotein Lipase 38 Occupation of Endothelial Binding Sites 2.1 Introduction 38 2.2 Material and Methods 40 2.2.1 Experimental Animals and Induction of Hyperglycemia 40 2.2.2 Measurement of Total Cardiac LPL Expression, Protein and Activity... 41 2.2.3 Immunolocalization of LPL 42 2.2.4 Isolated Heart Perfusion 42 2.2.5 Purification and Perfusion of Bovine Milk LPL 43 2.2.6 Treatments 43 2.2.7 Malonyl CoA and Acetyl CoA Carboxylase Estimation 44 2.2.8 Materials 45 2.2.9 Statistical Analysis 45 2.3 Results 46 2.3.1 LPL Expression, Protein, Activity, and Immunolocalization in STZ Heart.. 46 2.3.2 LPL Binding Sites in the Coronary Lumen 46 2.3.3 In Vitro LPL Translocation 47 2.3.4 Regulation of LPL by Insulin 48 2.3.5 Regulation of LPL by Cardiac Malonyl-CoA and PDH 49 2.4 Discussion 50 2.5 Tables and Figures 54 2.6 Bibliography 63 3.0 Circulating Triglyceride Lipolysis Facilitates Lipoprotein Lipase 68 Translocation from Cardiomyocyte to Myocardial Endothelial Lining 3.1 Introduction 68 3.2 Material And Methods 70 3.2.1 Experimental Animals 70 3.2.2 Diazoxide Induced Acute Hyperglycemia 70 3.2.3 Isolated Heart Perfusion 70 3.2.4 Coronary Lumen LPL Activity 71 3.2.5 Immunolocalization of LPL in DZ Heart 71 v 3.2.6 LPL Content of Cardiomyocytes 72 3.2.7 Treatments 73 3.2.8 Plasma Measurements 74 3.2.9 Materials 74 3.2.10 Statistical Analysis 74 i 3.3 Results 75 3.3.1 Effects of DZ on Plasma Parameters 75 3.3.2 LPL Activity, Immunolocalization and Effects of Insulin Treatment 75 3.3.3 Cardiac Myocyte Cell Surface and Intracellular LPL Activity 76 3.3.4 Lysophosphatidylcholine Effects on Luminal LPL 76 3.3.5 Manipulation of Circulating TG 77 3.4 Discussion 78 3.5 Tables and Figures 82 3.6 Bibliography 92 .0 Palmitoyl Lysophosphatidylcholine Mediated Mobilization of LPL to 97 the Coronary Luminal Surface Requires PKC Activation 4.1 Introduction 97 4.2 Material and Methods 99 4.2.1 Experimental Animals 99 4.2.2 Isolated Heart Perfusion 99 4.2.3 Isolated Cardiac Myocytes 99 4.2.4 Measurement of LPL Activity 100 4.2.5 Immunoblot Analysis for PKC 100 4.2.6 Immunolocalization of PKCe 101 4.2.7 Treatments 1 0 1 4.2.8 Materials 1 0 3 4.2.9 Statistical Analysis 1 0 3 4.3 Results 1 0 4 4.3.1 LPC Maintains High LPL in DZ Treated Heart Perfused In Vitro... 1 0 4 4.3.2 LPC Mediated Increases in LPL Requires Cardiac PKC Activation 1 ° 5 4.3.3 Differential Effect of LPC on Whole Heart and Cardiomyocyte LPL 105 vi 4.3.4 LPC mediated Increases in LPL Requires Formation of LPA 106 4.4 Discussion 107 4.5 Figures 110 4.6 Bibliography 119 5.0 Lysophosphatidic Acid Mediated Augmentation of Cardiomyocyte 126 Lipoprotein Lipase Involves Actin Cytoskeleton Reorganization 5.1 Introduction 126 5.2 Material and Methods 128 5.2.1 Experimental Animals 128 5.2.2 Isolated Heart Perfusion 128 5.2.3 Isolated Cardiac Myocytes 128 5.2.4 Measurement of LPL Activity 129 5.2.5 Western Blotting for Rho A and ROCK I & II 129 5.2.6 Cardiac LPL Gene Expression 130 5.2.7 Immunolocalization of Cardiomyocyte Actin 130 5.2.8 Adipose Tissue LPL Activity 131 5.2.9 Treatments 131 5.2.10 Serum Measurements 132 5.2.11 Materials 132 5.2.12 Statistical Analysis 132 5.3 Results 133 5.3.1 LPA Augments Myocyte LPL Activity 133 5.3.2 Augmentation of HR-LPL in LPA Perfused Isolated Hearts 133 5.3.3 Inhibition of LPA Mediated Augmentation of Myocyte HR-LPL 133 5.3.4 LPA Induces Cardiomyocyte Actin Polymerization 134 5.3.5 Augmentation of HR-LPL Requires RhoA and ROCK Activation 134 5.3.6 LPA Augments Adipose Tissue LPL Activity 135 5.3.7 RhoA and ROCK are Activated during Acute Hyperglycemia 135 5.4 Discussion 136 5.5 Figures 140 5.6 Bibliography 149 vii 6.0 C o n c l u s i o n s A n d Future Direct ions 156 6.1 Summary 156 6.2 Figures 158 6.3 Bibliography 159 viii L I S T O F T A B L E S Tab les Title Page Table 2-1 General characteristics of STZ and DZ treated animals 54 Table 3-1 Body weight, serum TG and FA subsequent to DZ administration 82 either in the presence or absence of WR 1339 or CPA Table 3-2 Experimental design for in vivo DZ treatment and in vitro LPC 83 perfusion ix L I S T O F F I G U R E S F igures Title Page Figure 1-1 Substrate supply to the heart 17 Figure 1-2 LPL synthesis, activation and secretion from cardiomyocytes 18 Figure 2-1 LPL gene expression, protein and activity in STZ rat heart 55 Figure 2-2 LPL immunofluroscence in STZ rat heart 56 Figure 2-3 Effect of STZ on endothelial LPL binding sites in isolated hearts 57 Figure 2-4 In vitro effect of two heparin perfusions on LPL activity in rat hearts 58 Figure 2-5 Effect of phloridzin treatment on LPL activity in perfused STZ hearts 59 Figure 2-6 Effect of insulin on plasma glucose and cardiac LPL activity in DZ rats 60 Figure 2-7 Effect of insulin on malonyl CoA and ACC in DZ treated hearts 61 Figure 2-8 Effect of DCA in moderating the effects of insulin on LPL activity 62 Figure 3-1 Serum insulin and glucose subsequent to DZ administration 84 Figure 3-2 Serum TG and FA subsequent to DZ administration 85 Figure 3-3 LPL activity following DZ administration 86 Figure 3-4 LPL immunofluroscence in DZ rat heart 87 Figure 3-5 Effect of insulin on HR-LPL activity in DZ treated rat heart 88 Figure 3-6 LPL activity in cardiac myocytes from control and DZ treated rats 89 Figure 3-7 In vitro effect of LPC on cardiac luminal LPL activity 90 Figure 3-8 Effect of WR 1339 and CPA on serum TG and LPL activity in DZ rats 91 Figure 4-1 Effect of LPC on luminal LPL in DZ treated heart perfused in vitro 110 Figure 4-2 In vitro effects of palmitoyl, oleoyl and stearoyl LPC on cardiac LPL 111 Figure 4-3 PKC e immunoblot in hearts perfused with LPC in vitro 112 Figure 4-4 PKC s immunofluroscence in hearts perfused with LPC in vitro 113 Figure 4-5 Effect of Calphostin on PKC s and LPL in LPC perfused heart 114 Figure 4-6 Effect of LPC on cardiomyocyte LPL activity and PKC s activation 115 Figure 4-7 PKC s immunoblot in HUVECs perfused with LPC 116 Figure 4-8 Effect of LPA on cardiomyocyte LPL activity and PKC s activation 117 Figure 4-9 Scheme of probable mechanism for LPC induced LPL transcytosis 118 x F igures Title P a g e Figure 5-1 Dose dependent effect of LPA on myocyte LPL mRNA and activity 140 Figure 5-2 In vitro effect of LPA on cardiac luminal LPL activity 141 Figure 5-3 Effect of CTD and SUR in moderating the outcome of LPA on LPL 142 Figure 5-4 F and G actin immunofluroscence in myocytes incubated with LPA 143 Figure 5-5 Rho A and ROCK 1 immunoblot in myocytes incubated with LPA 144 Figure 5-6 Effect of Y-27632 in moderating the role of LPA on Rho A and LPL 145 Figure 5-7 Effect of LPA on epididymal adipose tissue LPL activity 146 Figure 5-8 Effect of DZ administration on cardiac RhoA and ROCK 1 147 Figure 5-9 Scheme of probable mechanism for LPA induced LPL translocation 148 Figure 6-1 FA induced lipotoxicity in the diabetic heart 158 xi L I S T O F S Y M B O L S ° c Degrees Celsius g Gram h Hour ul Microliter igG Immunoglobulin i.p. Intraperitoneal i.v. Intravenous Kb Kilobase kg Kilogram mg Milligram ml Milliliter mm Millimeter mM Millimolar nM Nanomolar L I S T O F A B B R E V I A T I O N S ACC Acetyl coenzyme A carboxylase Apo Apolipoprotein ATP Adenosine triphosphate BSA Bovine serum albumin CAL Calphostin cAMP Cyclic adenosine monophosphate CHYL Chylomicron Cld/CId Combined lipase deficiency CPA N6- cyclopentyladenosine CPT-1 Carnitine palmitoyl transferase-1 CTD Cytochalasin D DCA Dichloroacetate DEX Dexamethasone DNA Deoxyribonucleic acid DZ Diazoxide EDG Endothelial differentiating gene Endo H Endo-beta-N-acetylglucosaminidase ER Endoplasmic reticulum F Actin Filamentous actin Fatty Acid FA G Actin Globular actin GLUT Glucose transporter HDL High density lipoproteins HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HR-LPL Heparin releasable-lipoprotein lipase HS Heparan sulphate HSPG Heparan sulphate proteoglycan HUVEC Human umblical vein endothelial cell LPA Lysophosphatidic acid xiii LPC Lysophosphatidylcholine LPL Lipoprotein lipase MCD Malonyl coenzyme A decarboxylase mRNA Messenger ribonucleic acid NEFA Non esterified fatty acid PBS Phospahte buffered saline PDH Pyruvate dehydrogenase PDK Pyruvate dehydrogenase kinase PHZ Phloridzin PKC Protein kinase C PLD Phospholipase D PPAR Perioxisome proliferator activated receptor RNA Ribonucleic acid ROCK Rho kinase RT-PCR Real time-polymerase chain reaction SDS Sodium dodecyl sulphate SEM Standard error of means STZ Streptozotocin SUR Suramin TBS-T Tris buffered saline-Tween TG Triglyceride TGRL Triglyceride rich lipoprotein VLDL Very low density lipoprotein WR 1339 Tyloxapol;Triton xiv A C K N O W L E D G E M E N T S This section in the thesis gives the student an opportunity to thank and regard the enormous support, encouragement and knowledge garnered from different minds during the course of this intensive training program. To begin with I would like to extend my deep sense of gratitude and respect to my research supervisor and friend, Professor Dr. Brian Rodrigues, Chair, Department of Pharmacology and Toxicology, University of British Columbia. It has been a great privilege and honor to be mentored by Brian especially for his meticulous thinking, constructive criticisms, perseverance and unflagging devotion towards research. He taught me the skills to pose significant research questions, achieve systematic solutions and helped me understand the philosophy of research. His painstaking efforts and untiring enthusiasm invested on me has yielded four manuscripts out of this dissertation. I thank him for all the guidance, support, encouragement and friendship. I am also grateful to the members of my supervisory committee for their valuable suggestions and encouragement they provided me during the course of my academic and research training. I would also like to thank Ashraf and Jospy for having taught me research and administrative skills. Their technical help and gracious collegiality was an added asset during the course of my program. I sincerely acknowledge the friendship, support and encouragement of my colleagues Sanjoy, Jeff, Dake and Girish. Their intellect and brilliant presence provided a professionally stimulating atmosphere in the lab. I thank Nathan, Patsy, Claudia, Jayda, Naureen and Gloria for being talented and inquisitive summer students who worked with me during my training program. I would like to express my gratefulness to the CIHR/Rx&D, and Heart and Stroke Foundation of BC and Yukon for their financial support during my graduate program. I would like to sincerely acknowledge the collaborative efforts of Dr. Roger Brownsey, Dr. Sheila Innis and Dr. Michael Allard during the completion of this dissertation. At this juncture I also take pride in thanking the Faculty of Pharmaceutical Sciences for providing an excellent graduate training program. I thank the faculty, technical and administrative staff for the help, support and encouragement during this program. My special thanks to all my friends and well-wishers within and outside the faculty for their cheerful companionship and having made my stay in UBC pleasant and memorable. Last but not the least, no words of esteem can suffice to express my love and appreciation for my loving parents and brother. Without their moral, selfless support and sacrifice accomplishing my goal to acquire a PhD would have been impossible. xv xvi List of the Manuscripts that Constitute this Thesis 1. Pulinilkunnil, T., and Rodrigues, B. Cardiac lipoprotein lipase regulation: Metabolic basis for diabetic heart disease. Review submitted to Cardiovasc. Res. (Accepted). 2. Pulinilkunnil, T., An, D., Ghosh, S., Qi, D., Kewalramani, G., Yuen, G., Virk, N., Abraham, A., and Rodrigues, B. Lysophosphatidic acid mediated augmentation of cardiomyocyte lipoprotein lipase involves actin cytoskeleton reorganization. Am. J. Physiol. 288: H2802-H2810, 2005. 3. Pulinilkunnil, T., An, D., Yip, P., Chan, N., Qi, D., Ghosh, S., Abraham, M . , Rodrigues, B. Palmitoyl lysophosphatidylcholine mediated mobilization of LPL to the coronary luminal surface requires PKC activation. J. Mol. Cell. Cardiol. 37: 931-938, 2004. 4. Pulinilkunnil, T.. Qi, D., Ghosh, D., Cheung, C , Yip, P., Varghese, J., Abrahani, A., Brownsey, R., and Rodrigues, B. Circulating triglyceride lipolysis facilitates lipoprotein lipase translocation from cardiomyocyte to myocardial endothelial lining. Cardiovasc. Res. 59: 788-797, 2003. 5. Pulinilkunnil, T., Abrahani, A., Varghese, J., Chan, N., Tang, I., Ghosh, S., Kulpa, J., Allard, M . , Brownsey, R., and Rodrigues, B. Evidence for rapid "metabolic switching" through lipoprotein lipase occupation of endothelial binding sites. J. Mol. Cell. Cardiol. 35: 1093-1103,2003. 6. Pulinilkunnil, T., Sambandam, N., and Rodrigues, B. Altered substrate utilization in the diabetic heart: Role of lipoprotein lipase. In: Frontiers in Cardiovascular Health. N.S. Dhalla, A. Chockalingam, H.I. Berkowitz, P.K. Singal (Eds.). Kluwer Academic Publishers, Boston, M A , pp 119-142, 2003. xvii Chapter 1: Invited Review Accepted in Cardiovas. Res. 1.0 Introduction 1.1 Clinical problem of diabetes The World Health Organization estimates that approximately 170 million people suffer from diabetes worldwide, with this figure likely to more than double by 2030 (http://www.who.int/diabetes/en/). In patients diagnosed with diabetes, the Type 2 form (tissue insulin resistance followed by inadequate insulin secretion and ultimately, hyperglycemia) accounts for 90% of diagnosed cases when compared to Type 1 (pancreatic P cell death with catastrophic loss of circulating insulin and profound hyperglycemia) [1, 2]. Among the numerous complications of diabetes, cardiovascular disease is the leading cause of death [3]. Although coronary vessel disease and atherosclerosis have been identified to be the primary reasons for the increased incidence of cardiovascular dysfunction [1-3], clinical data suggests the prevalence of an additional cardiac injury termed 'diabetic cardiomyopathy' [4-6]. Indeed, a number of diabetic patients suffer from left ventricular dysfunction, cardiomegaly and clinically overt congestive heart failure in the absence of any change in epicardial coronary blood flow [7-9]. In experimental models of diabetes, myocardial abnormalities also occur without any coronary defects [10-13]. Experimental diabetic cardiomyopathy consists of two major elements: an initial phase of short-term and severe modification in fuel metabolism, followed by progressive chronic myocardial damage that involves augmented stiffness of the left ventricular wall, accumulation of connective tissue and insoluble collagen, and abnormalities of various proteins that regulate ion flux (specifically intracellular calcium) [10-13]. After 4 to 6 weeks, these changes are reflected in a measurable contractile dysfunction [12-15]. In patients with diabetes, numerous studies have confirmed all of these chronic abnormalities [16, 17]. However, verification of the initial cardiac changes [18] observed in animals has not been completely established in humans given the limited diagnostic use of sensitive techniques to measure 1 Chapter 1: Invited Review Accepted in Cardiovas. Res. changes in energy metabolism. This has led to widespread underestimation of these early cardiac events. Hence, in human diabetes, a preliminary metabolic component [19] may induce cell death [20] and deplete myocardial reserve, predisposing the diabetic heart to accelerated damage when exposed to conditions of dyslipidemia, atherosclerosis, and hypertension. 1.2 Ca rd iac energy metabo l i sm 1.2.1 Substrate utilization In the heart, cellular energy in the form of ATP is obtained via the oxidation of various substrates like fatty acids (FA), glucose, lactate and ketone bodies (Fig. 1-1), with glucose and FA being the principal fuels [21-25]. Glucose utilization provides the heart with approximately 30% of its energy requirements [24, 25]. Following insulin dependent glucose uptake and glycolysis [26], pyruvate dehydrogenase (PDH) complex facilitates entry of pyruvate into the mitochondria [27] and its oxidation eventually provides the majority of energy obtained from glucose [28]. Compared to glucose, FA is the preferred substrate, and contributes approximately 70% of the ATP necessary for normal heart function [29]. Even though the heart exhibits a preference for FA, it has limited potential to synthesize this substrate. Therefore, FA delivery and utilization by the heart involves: i) release from adipose tissue and transport to the heart after complexing with albumin [29], ii) provision through the breakdown of endogenous cardiac triglyceride (TG) stores [30, 31], iii) internalization of whole lipoproteins [32-34], and iv) hydrolysis of circulating TG-rich lipoproteins to FA by lipoprotein lipase (LPL) positioned at the endothelial surface of the coronary lumen [35]. The molar concentration of FA bound to albumin is ~10 fold less than that of FA in lipoprotein-TG [36] and studies that compared the utilization of exogenous FA and lipoprotein-TG, alone or in combination, using the isolated working control rat heart, showed a cardiac preference for non-esterified F A [37]. Recently, 2 Chapter 1: Invited Review Accepted in Cardiovas. Res. LPL-mediated hydrolysis of circulating lipoproteins to FA was suggested to be the principal source of FA for cardiac utilization [38]. 1.2.2 Cardiac metabolism during diabetes Type 1 or Type 2 diabetes impairs glucose uptake in the heart [39-41]. The major restriction to glucose utilization is the slow rate of glucose transport across the sarcolemmal membrane into the myocardium, which probably results from cellular depletion of glucose transporters (GLUT-4) [42, 43]. In animal models of diabetes, the levels of GLUT4 mRNA and protein decreases by 50% [44, 45], and both basal and insulin stimulated glucose uptake is hindered [46]. Suppression of glucose utilization can also occur secondary to the inhibition of hexokinase and PDH activity that limits glucose entry, phosphorylation and its subsequent mitochondrial oxidation [24, 47-49]. Eventually, following impaired glucose utilization, the heart switches to exclusive R-oxidation of FA [12, 25]. To support this augmented utilization of FA, a multitude of events occur. Adipose tissue lipolysis is increased, resulting in elevated circulating plasma FA leading to increased hepatic very low density lipoproteins (VLDL) secretion and circulating V L D L - T G concentrations [50-52]. There is an enhanced activity of enzymes that catalyze the synthesis of TG, and a concurrent rise in coenzyme A levels; this promotes the accumulation of intracellular T G stores [52]. Subsequent hydrolysis of this T G store may also lead to high tissue FA levels [53, 54]. Under these circumstances, LPL also increases [55]; thus, all of the above processes serve to guarantee FA supply to diabetic tissues in order to compensate for the diminished contribution of glucose as an energy source. However, the dramatic increase in FA influx is not without negative consequences. In the heart, elevated FA and subsequent T G synthesis have been implicated in a number of metabolic, morphological, and mechanical changes, and more recently, in "lipotoxicity" [56, 57]. During lipotoxicity, F A accumulates and can, either by themselves or via production of second messengers such as ceramides, provoke 3 Chapter 1: Invited Review Accepted in Cardiovas. Res. cell death [57, 58]. Given the pivotal function of LPL in FA delivery, and FA's contribution in mediating cellular lipotoxicity, examining the regulation of this enzyme is crucial for understanding the metabolic basis of diabetic heart disease. 1.2.3 Role of LPL in heart disease Hypertriglyceridemia is a common feature in patients with insulin resistance [59], diabetes [60, 61] and obesity [62] and is positively correlated with the incidence of cardiovascular complications [63]. Diminished whole body LPL activity is suggested to be one mechanism by which circulating lipoproteins are not cleared leading to hypertriglyceridemia [64]. Interestingly, administration of the LPL-activating agent, NO-1886 [65], in high-fat fed animals, suppressed lipid accumulation [66], insulin resistance [66] and intestinal polyp formation secondary to hypertriglyceridemia [67]. Moreover this compound also improved fatty liver caused by high-fat feeding in streptozotocin (STZ)-induced diabetic rats [68]. In transgenic rabbits that have global overexpression of LPL, attenuation of hypertriglyceridemia was observed, an effect suggested to contribute toward amelioration of insulin resistance and obesity [69, 70]. Taken together, these studies suggested that systemic increases in LPL levels could be associated with amelioration of insulin resistance and obesity. Conversely, other studies have documented that augmented LPL can result in insulin resistance [71] and obesity [72]. Thus, the question that emerges is how much of an increase in LPL activity is required for the enzyme to be either beneficial or pathological. In an attempt to address this issue, transgenic animals were developed that overexpressed LPL at low, moderate and high levels. High tissue-specific LPL overexpression in skeletal muscle and heart was associated with insulin resistance and severe myopathy [73], characterized by both muscle fiber degeneration and extensive proliferation of mitochondria and peroxisomes [74]. In a more recent study using genetically engineered mice that specifically overexpressed cardiomyocyte surface-bound LPL, lipid oversupply and 4 Chapter 1: Invited Review Accepted in Cardiovas. Res. deposition was observed, together with excessive dilatation and impaired left ventricular systolic function (lipotoxic cardiomyopathy) [75]. These experiments clearly demonstrate that in the absence of any vascular defects, selective overexpression of LPL in the heart is sufficient to cause cardiac failure. Diabetes is associated with heart disease and it has been unequivocally demonstrated that LPL per se has the ability to accelerate abnormal FA supply and utilization leading to lipotoxic cardiac disease. Thus specifically modulating LPL in the heart would be an important therapeutic advantage preventing initiation of diabetic heart disease. 1.3 L ipoprote in l ipase (LPL) The human LPL gene is located on chromosome 8 [76, 77], and is approximately 30 kb pairs in length [78, 79] when compared to murine (28 kb) [80] and chicken genomic sequences (17 kb) [81]. In human and rodent models, cardiac LPL gene expression and protein synthesis begins post natal, and reaches the highest level within 3 weeks of age [82, 83]. Although the functional location of LPL-mediated lipoprotein hydrolysis is at the capillary endothelial cell surface, a number of approaches including in situ hybridization have failed to demonstrate LPL mRNA localization in endothelial cells [84]. Experimental data from the adult mouse heart suggest that LPL is synthesized and processed in cardiac myocytes, with distribution of immunoreactive LPL protein being observed in different myocardial compartments [35, 85]. Thus, electron microscopic studies of immunogold-labeled sections of mouse myocardium demonstrated that 78% of total LPL was present in cardiac myocytes, 3-6% in the interstitial space, and 18% at the capillary endothelium [86]. Within the myocytes, LPL was found localized within the sarcoplasmic reticulum, Golgi complex and secretory vesicles [35, 86]. In the human myocardium, LPL protein was also detected in interstitial cells [85]. 5 Chapter 1: Invited Review Accepted in Cardiovas. Res. 1.3.1 Synthesis and activation Synthesis, processing and translocation of LPL in the heart occur by mechanisms similar to those of adipose tissue (Fig. 1-2). LPL is synthesized as an inactive, monomeric proenzyme in the rough endoplasmic reticulum (ER), and is postulated to be activated somewhere between the ER and Golgi apparatus [87]. The catalytic activity of the dimeric enzyme is dependent on asparagine-linked glycosylation of the N-terminal domain of LPL [88-90]. During translational modifications, glycosyl moieties are added to asparagine residues on the nascent LPL polypeptide by a dolichol (a lipid)-linked oligosaccharide [87, 91]. Mannose rich glycosyl residues within this glycoprotein not only allow its retention within the ER but also make it susceptible to endo-p-N-acetylglucosaminidase H (endo-H) mediated degradation [87, 92]. Subsequently, this N-linked oligosaccharide loses its terminal glucose residues when acted upon by ER specific glycosidases I and II for its ensuing transfer to a cis-golgi compartment [87, 93]. In this compartment, the peptide undergoes a-mannosidase dependent cleavage of mannose residues giving rise to a high mannose peptide [87]. As the glycoprotein moves through the medial-golgi, mannosidase II mediated cleavage of two additional mannose residues ensues, resulting in the formation of a endo H resistant glycoprotein, that then undergoes a series of supplementary modifications in the trans-golgi, and proceeds for secretion [87, 92, 94]. Based on in vitro experiments, two different groups have proposed contrasting views on how activation of LPL occurs within the cell. One group proposes that LPL exists as an inactive monomer within the ER, and that activation of LPL occurs only after translocation to the cis/medial-Golgi [92]. Findings from other labs have suggested that translocation of LPL from ER to cis-Golgi is not required for full expression of catalytic activity [95, 96]. Interestingly, in transgenic mouse models of combined lipase deficiency (cld/cld), although LPL was glycosylated and dimerized, 6 Chapter 1: Invited Review Accepted in Cardiovas. Res. active LPL could not be produced until it was secreted out of the ER [97]. This defect was later attributed to the lack of ER specific chaperones like calnexin, calreticulin, and ERp57 that allow proteins to be folded in a manner optimal for secretion to golgi. In sf21 insect cells, calreticulin facilitated folding and dimerization of expressed human lipoprotein lipase [98]. 1.3.2 Secretion Following completion of enzyme synthesis, LPL is secreted as an active homodimer by either constitutive and/or regulated mechanisms [87]. Constitutive mechanisms involve spontaneous release of LPL, whereas regulated release occurs in response to a secretogogue (e.g., heparin) [99]. Majority of the cell types expressing LPL exhibit constitutive release. For regulated release to occur, the enzyme requires packaging and assembly in secretory vesicles [87]. In cardiomyocytes, dimerized LPL binds to cell surface heparan sulphate proteoglycans (HSPG) [87]. Subsequently, LPL is translocated across the interstitial space to the endothelial cell surface in the vascular lumen where it executes its function of hydrolyzing circulating lipoproteins [100, 101]. A number of observations support the idea that luminal HSPG bound LPL is not an outcome of uptake from the plasma pool but rather, requires transfer from the myocytes. For example perfusion of isolated rat [55] or guinea pig hearts [102-104] with heparin causes release of LPL to occur in two phases: a) a rapid phase, occurring within seconds after heparin perfusion, probably representing detachment of LPL from the endothelial luminal (apical) surface [55], and b) a sustained release, occurring likely as a result of LPL mobilization from within endothelial cells, interstitial space or cardiomyocytes [55]. 1.3.3 Transfer of secreted LPL from myocyte to endothelial cells Myocyte cell surface LPL is suggested to be transported towards the vascular endothelial cells by mechanisms that are currently unclear. In co-culture experiments using adipocytes and endothelial cells, heparanase like compounds secreted from endothelial cells have been 7 Chapter 1: Invited Review Accepted in Cardiovas. Res. suggested to cleave adipocyte bound HSPGs to release heparan sulphate oligosaccharides [105]. The oligosaccharides bound LPL non-covalently, likely preventing LPL degradation in interstitial fluid, and serve as extracellular chaperones [105], enabling transport of the enzyme in its active form across the endothelial cells to the apical surface of endothelial cells [106]. Subendothelial basement membranes likely sequester and stabilize this LPL secreted by myocytes [107]. The extracellular matrix in subendothelial basement membranes are composed of collagens, fibronectin, laminin and glycosaminoglycans like heparan, dermatan and chondroitin sulfate-proteoglycans, of which LPL binds mainly to HSPG. Binding of LPL to HSPG at the basolateral surface of the endothelium was reported to be obligatory for efficient transport to the apical surface, an effect that was susceptible to heparinases (proteases that cleave HSPGs) [108]. In addition to HSPG, movement of LPL across the endothelial cell also requires V L D L receptors [109-111]. As endothelial cells lining the microvessels of the heart are continuous, transport of macromolecules across the coronary endothelial barrier likely involves specific transcytotic mechanisms [112]. In this regard, caveolae, that are endothelium specific vesicular invaginations, are proposed to act as ferries for macromolecules, likely facilitating receptor-mediated endocytosis or transcytosis [113-115]. Given the role of actin in maintaining caveolar integrity at the plasma membrane, disruption of actin filaments by cytochalasin B inhibited vesicular transport [116]. In fibroblasts, as binding of LPL to HSPG aligns these proteoglycans along the actin cytoskeleton [117], disruption of actin filaments resulted in irregular distribution of HSPG on the fibroblast cell surface, with ensuing impairment of LPL secretion [118, 119]. Whether this organization of HSPG along the actin cytoskeleton is involved in the transcytosis of LPL from the abluminal to the luminal surface in the coronary endothelial cell is presently unknown. 8 Chapter I: Invited Review Accepted in Cardiovas. Res. 1.4 Phys io log i ca l regulat ion of L P L Potential sites for regulation of cardiac LPL include: a) nucleus (transcriptional control) [100, 120], b) rough endoplasmic reticulum (maturation of LPL by glycosylation) [96], c) golgi network (vesicular transport and secretion of LPL) [95, 121], d) plasma membrane (LPL binding to the cell surface) [84], and e) vascular endothelial cell surface (vectorial transfer and recycling of the enzyme) [122, 123]. Of these sites, the majority of studies have focused on enzyme regulation at the transcriptional and translational levels. The unique characteristic of LPL is that in some tissues like adipose and heart, changes in activity can occur independent of alterations in LPL mRNA [124]. Such changes would be desirable given that during conditions like fasting or diabetes, when FA utilization is augmented, rapid increase in LPL activity may not match the slow turnover of LPL mRNA [125]. Posttranslational regulation of this enzyme includes; 1.4.1 Degradation of intracellular LPL The process of LPL turnover by intracellular degradation represents an important post-translational mechanism by which enzyme activity is rapidly regulated [126]. In adipose tissue, this process is rapid, with a im of about 40 minutes resulting in .almost 80% of newly synthesized LPL being degraded prior to being secreted [127, 128]. Degradation probably occurs either in a leupeptin inhibitable, lysosomal compartment or a tunicamycin sensitive, ER compartment [92]. Interestingly, similar to the intracellular pool, adipocyte surface bound LPL also undergoes degradation after being internalized [87, 129]. With surface bound LPL, only a fraction (<28%) undergoes lysosomal degradation following internalization of the enzyme [129]. Moreover, this degradation is dependent on both the number of HSPG binding sites in addition to the extent of HSPG sulfation [130, 131]. Whether the cardiomyocyte internalizes surface bound LPL for degradation and if so, do alterations in this process regulate cardiac LPL during physiological or pathological stress merits further investigation; 9 Chapter 1: Invited Review Accepted in Cardiovas. Res. 1.4.2 Recycling of intraendothelial LPL Another mechanism by which heart LPL could be regulated involves recycling of endothelial surface bound LPL. LPL turnover at the endothelial lumen can occur either by detachment from HSPG and subsequent hepatic degradation, or internalization of the HSPG-LPL complex into an endothelial endocytotic compartment [132]. Within the endothelial cell, an acidic pH enables LPL to remain bound to HSPG, promoting recycling of the internalized LPL complex back onto the luminal surface, thereby allowing endothelial cells to maintain an auxiliary pool of the enzyme [132]. The acidic pH in the endocytotic vesicles favors HSPG-LPL binding, thereby enabling the complex to be released back into the medium or inserted onto the cell surface. Whether this mechanism of endothelial recycling takes place in the intact heart is yet to be ascertained. 1.4.3 Regulation of LPL by substrates LPL synthesis and activity are regulated in a tissue specific manner by conditions such as exposure to cold [133, 134], lactation [135, 136], feeding [94] and fasting [137]. During fasting, LPL activity decreases in adipose tissue, an effect opposite to that seen during feeding [138]. The fasting induced decline in adipose LPL was secondary to degradation of newly synthesized LPL [126], suggesting posttranslational regulation of the enzyme. More recently, fasting did not alter intradipocyte LPL even though the extracellular pool of enzyme was changed to the inactive form [137]. As the latter effect was sensitive to inhibition by transcription blockers, the authors suggested the presence of a gene whose product prevents the enzyme from becoming active even though synthesis of LPL protein continues unabated [137]. In the heart, fasting with ensuing hypoinsulinemia, augments heparin releasable LPL (HR-LPL) activity [139-141]. This was supported by immunocytochemical studies in the mouse myocardium, which revealed a 5-fold increase in LPL at the luminal projections of endothelium subsequent to fasting [35]. 10 Chapter 1: Invited Review Accepted in Cardiovas. Res. Changes in luminal LPL activity following fasting were independent of shifts in LPL mRNA or alterations in LPL protein and activity in cardiomyocytes, suggesting a posttranslational mechanism for this increase [139]. Some studies have suggested that this increase in HR-LPL could be attributed to increased uptake of LPL from the blood since uptake of exogenous 1 2 5I-LPL was augmented in the heart following fasting [141, 142]. However, in perfused guinea pig hearts, newly synthesized LPL can move from myocytes to the vascular lumen within 30 min and increased rate of LPL transfer could also explain this effect of fasting on HR-LPL [102, 103]. During nutritional changes, TG-lipoproteins from both exogenous (chylomicrons) and endogenous (VLDL) sources and products of lipolysis like FA likely regulate LPL activity. LPL-derived FA from T G hydrolysis can bind and competitively inhibit LPL activity [143], an effect that is abolished by incubation with albumin due to its higher affinity for FA [143]. Displacement of LPL from its binding sites is yet another potential mechanism by which LPL activity can be regulated [144]. Adipose tissue LPL bound to endothelial cells can be displaced from its binding sites by triglyceride rich lipoproteins (TGRL) and oleic acid [145, 146]. It was proposed that following LPL mediated T G hydrolysis, when supply of FA exceeds tissue demand, FA would probably bind to LPL and displace it from its binding sites, thereby regulating the enzyme activity via a negative feedback mechanism. This effect was not observed in isolated cardiac myocytes or whole hearts [147]. Alternatively, incubation of cardiac myocytes with oleic acid resulted in a decreased LPL activity [148]. This was not secondary to release of the enzyme from its binding sites but due to alterations in intracellular processing and secretion of the enzyme [148]. In addition to FA, apolipoproteins present on TGRLs and those associated with HDL also influence LPL [101]. Binding of apoCII present on the TGRL surface to LPL enhances lipolysis while binding of apoCIII inhibits LPL activity [149]. Given that 11 Chapter 1: Invited Review Accepted in Cardiovas. Res. chylomicrons obtained from nascent lymph has negligible amount of surface apoC-II, procatalytic apolipoproteins must associate with chylomicrons after they enter the blood stream via replacement of apoCs and apoE on HDL for apoA-I and apoA-IV on chylomicrons [101]. On the contrary, anticatalytic apolipoproteins inhibit LPL-mediated lipolysis [150]. Lipid metabolism studies utilizing apoC-III transgenic mice [151, 152] and apoCIII knockout mice [153, 154] have confirmed the regulatory effect of apoC-III on LPL-mediated lipolysis. ApoCIII either displaces apoCII, exhibits negative effect on the enzyme per se [155] or reduces uptake of remnant lipoproteins by not allowing them to interact with the L D L receptor-related protein [101]. ApoE, another major apolipoprotein of TGRLs, has also been shown to inhibit LPL activity [156]. HDL-associated apo A V is the most recent candidate shown to affect LPL [157]; its overexpression accelerates plasma hydrolysis of T G rich lipoproteins [158]. 1.5 Card iac L ipoprote in l ipase dur ing hypo insu l inemia a n d insul in res is tance 1.5.1 LPL in the STZ diabetic heart Although extensively studied, the relative contribution of cardiac LPL activity to the delivery of FA to the diabetic heart was inconclusive [55]. In part, this variability among different studies was due to the dosage of STZ used to induce diabetes, and the duration of diabetic state [55]. For example, induction of severe diabetes with 100 mg/kg STZ reduced heparin-releasable coronary and myocyte-associated LPL within 4 days of STZ [55]. This decline in functional activity resulted in impaired lipolysis of perfused V L D L - T G , effects that were normalized by insulin treatment [159, 160]. Other reasons for this variability in LPL activity measurements during diabetes occurred because LPL activity and protein level measurements were largely obtained using whole heart homogenates, which do not distinguish LPL localized on capillary endothelial cells from the myocyte pool [55]. Thus, moderate diabetes induced with 55 mg/kg STZ did not alter LPL when activity measurements were carried out in heart homogenates [55]. 12 Chapter 1: Invited Review Accepted in Cardiovas. Res. However, using retrograde perfusion of these hearts with heparin to displace coronary LPL, we were the first to demonstrate significantly elevated HR-LPL activity following 2 and 12 weeks of diabetes [55]. The elevated HR-LPL peak could not be explained by an enhanced LPL synthesis, as both cellular and surface-bound LPL activities in myocytes from STZ 55 mg/kg administered rats were low relative to control. Next, we confirmed that: a) the increase in LPL protein originates mainly in capillary blood vessels, presumably within or at the luminal and abluminal surface of the endothelial cell [161], b) could be regulated by short-term changes in insulin [161], and c) was capable of hydrolyzing V L D L - T G [162]. At present, the mechanisms responsible for the amplification of the heparin-releasable pool at the endothelial surface following diabetes is not fully understood, but could involve the increased vectorial transfer of LPL, an enhanced intermediate interstitial pool of LPL, or an increased number of HSPG sites on the endothelial surface in the moderately diabetic rat heart. 7.5.2 Cardiac LPL following insulin resistance Insulin resistance is a condition when normal levels of insulin are incapable of rendering its physiological effects [163, 164]. Following this condition, there is compromised glucose uptake into the peripheral tissues leading to excessive circulating glucose levels [52]. To maintain normoglycemia, a greater concentration of insulin is required, and at least in the initial stages, the pancreas hypersecretes insulin (hyperinsulinemia) [165]. Eventually, pancreatic P-cell dysfunction may occur, leading to a loss of insulin release [165]. At this stage, due to a mismatch between circulating glucose and insulin levels, diabetes develops [52, 163, 165]. Glucocorticoids impair insulin sensitivity [166]. Although injection of D E X for 4h was not associated with hyperinsulinemia, the euglycemic-hyperinsulinemic clamp showed a decrease in glucose infusion rate [167]. Similar to hypoinsulinemia induced by STZ, hearts from insulin resistant DEX animals also demonstrated enlargement of the coronary LPL pool [167]. 13 Chapter I: Invited Review Accepted in Cardiovas. Res. 1.6 R e s e a r c h Rat ionale and Hypothes i s Clinical and experimental studies have established that a cardiomyopathy occurs during diabetes [4, 11]. Although a number of factors have been implicated in the development of this disease state, increasing evidence suggests that a metabolic derangement in fuel supply and utilization by the heart could be a primary reason for the pathogenesis of diabetic cardiomyopathy [19, 168-173]. Under normal aerobic conditions, cellular energy in the form of ATP is obtained via the oxidation of various substrates, with free FA being the preferred energy substrate utilized by the heart muscle. The heart has a limited potential to synthesize FA. Hence, F A is delivered to the cardiac tissue from several sources, of which LPL derived FA is the predominant source of FA to the heart [38, 55]. In diabetes, energy production is almost entirely via R-oxidation of FA, a process that may have deleterious effects on myocardial function. In an insulin-deficient state, adipose tissue lipolysis is enhanced, resulting in an elevated circulating FA. In addition, an increased activity of myocardial enzymes that catalyze the synthesis of TG, together with a rise in CoA levels, promotes the production of T G during diabetes. Subsequent hydrolysis of this augmented T G store could also lead to high tissue FA levels. In an insulin-deficient state, augmentation of luminal LPL also serves as a process to guarantee FA supply to the diabetic heart to compensate for the diminished contribution of glucose as an energy source. Together, these mechanisms serve to guarantee F A supply to the diabetic heart to compensate for the diminished contribution of glucose as an energy source. A consequence of these effects is an aberrant intracellular handling of C a 2 + that leads to C a 2 + overload, cell death, and eventual cardiac dysfunction. In the adult heart, LPL is synthesized and processed in myocytes and is translocated onto HSPGs binding sites on the luminal surface of endothelial cells where it actively metabolizes 14 Chapter 1: Invited Review Accepted in Cardiovas. Res. lipoproteins [36, 84, 99, 100]. Since endothelial cells cannot synthesize LPL, the enzyme is synthesized by the parenchymal cells of a variety of extrahepatic tissues, including adipose, heart, skeletal muscle, brain, and ovary. In the adult heart, LPL is synthesized and processed in myocytes and is translocated onto heparan sulfate proteoglycan (HSPGs) binding sites on the luminal surface of endothelial cells where it actively metabolizes lipoproteins. The mechanisms that operate to enhance luminal LPL are largely unknown in diabetic heart. Overall we hypothesize that "following diabetes cardiac specific mechanisms are activated to facilitate an increase in LPL at the myocardial endothelial lining. This metabolic switching likely initiates the development of heart disease ". 15 Chapter 1: Invited Review Accepted in Cardiovas. Res. 1.7 R e s e a r c h object ives The objectives of the proposed research were to investigate the following postulated mechanisms responsible for changes in coronary LPL following diabetes. 1. Vascular endothelial cells synthesize HSPGs, and the negatively charged sulphate residues of these HSPG interact with the positively charged amino acid residues on the C-terminus of LPL via electrostatic interaction. Hence, it is conceivable that diabetes, either by affecting the synthesis or catabolism of these HSPGs, increases the number of capillary endothelial LPL binding sites, and hence the amount of enzyme at this location. Therefore, we will evaluate whether the augmented enzyme pool at the coronary endothelial lining is a consequence of enhanced luminal HSPG binding sites. 2. During fasting enhancement of cardiac LPL was suggested to be an outcome of plasma LPL uptake by the heart. Thus, we will study whether the enhanced coronary LPL pool following diabetes is a result of increased LPL supply from non-cardiac tissues or is a process intrinsic to the heart. 3. In perfused guinea pig hearts, LPL can move from parenchymal cells to the endothelial surface within 30 min. The effect of diabetes to augment heparin-releasable LPL activity, even in the presence of a reduced myocyte enzyme pool suggests a possible enhancement in the vectorial transfer of LPL to the luminal surface. Therefore, we will evaluate whether the increased enzyme pool at the vascular lumen is secondary to accelerated translocation of the enzyme from the myocyte to the vascular lumen. 4. Following diabetes, in addition to lack of insulin, metabolic outcomes of insulin deficiency like hyperglycemia and lipolysis could also influence cardiac LPL. Hence we will determine the contribution of circulating plasma mediators on coronary LPL. 16 1.8 Figures Chapter 1: Invited Review Accepted in Cardiovas. Res. Fig. 1-1 Fatty acid provision to the heart. FA are provided to the heart from three major sources. Adipose tissue lipolysis with release of FA into the plasma, LPL mediated breakdown of T G - rich lipoproteins from liver (VLDL) and gut (Chylomicron), and endogenous T G breakdown within the heart. The other major substrate that is utilized by the heart is glucose. HSL-hormone sensitive lipase; FAO-fatty acid oxidation. 17 Chapter 1: Invited Review Accepted in Cardiovas. Res. Lumen <gy> tf\ ffi , Translocation Interstitium P • L P L • HSPG ***•• D N A R N A V • E R J Myocyte * C D O G O L G I Fig . 1-2 LPL synthesis, activation and secretion from cardiac myocytes. Following synthesis in the ER, LPL is exported to the golgi for either secretion or lysosomal degradation. After secretion and binding to myocyte cell surface HSPG, LPL is translocated to the abluminal side of the endothelial cell. Subsequently, the enzyme is transcytosed to the apical surface where it facilitates lipoprotein hydrolysis. 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Energy metabolism of the heart: from basic concepts to clinical applications. Curr Probl Cardiol 1994;19:59-113. [171] Taegtmeyer H, McNulty P, Young M E . Adaptation and maladaptation of the heart in diabetes: Part I: general concepts. Circulation 2002;105:1727-33. [172] Young M E , McNulty P, Taegtmeyer H. Adaptation and maladaptation of the heart in diabetes: Part II: potential mechanisms. Circulation 2002;105:1861-70. [173] Lopaschuk GD, Rebeyka IM, Allard MF. Metabolic modulation: a means to mend a broken heart. Circulation 2002;105:140-2. 37 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.0 Evidence for Rapid "Metabolic Switching" Through Lipoprotein Lipase Occupation of Endothelial Binding Sites 2.1 Introduction The "glucose-fatty acid cycle" which describes a negative relationship between FA concentration and carbohydrate oxidation suggests that the balance between glucose and F A oxidation is in effect determined by the intracellular availability of FA [1]. More recently, intracellular availability of glucose was also found to influence the nature of substrate utilization. Increase in intracellular glucose augments cytosolic citrate, leading to elevated acetyl-CoA production [2]. Malonyl-CoA that is eventually produced through the action of acetyl CoA carboxylase (ACC) inhibits carnitine palmitoyl transferase-1 (CPT-1) activity, prevents F A oxidation, and promotes glucose utilization [1]. During diabetes, impaired glucose transport and utilization by the heart switches energy production to exclusive P-oxidation of FA [3]. This occurs in part as an outcome of diminished malonyl-CoA, that relieves the inhibition on CPT-1 [3]. Whether altered cardiac malonyl-CoA also effects FA delivery to the heart by influencing LPL is unclear. Coronary endothelial-bound LPL actively metabolizes the T G core of lipoproteins to FA for myocardial energy production [4, 5]. As endothelial cells cannot synthesize LPL, it is manufactured and processed in myocytes and then translocated across the interstitial space onto HSPG binding sites on the luminal surface of endothelial cells [6, 7]. LPL protein or activity has been reported to be unchanged, increased or decreased in the diabetic rat heart [8, 9]. In part, this variability could be attributed to an inability to distinguish between enzyme localized on capillary endothelial cells from the cellular pools of cardiac LPL. Using an isolated heart, we have demonstrated significantly elevated HR-LPL activity in STZ induced diabetic rats after 2 or 12 weeks of hypoinsulinemia [9]. Interestingly, the higher luminal LPL activity in diabetic rats 38 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 could not be explained by an increase in myocyte cell surface or intracellular enzyme LPL pool that were dramatically reduced. In perfused guinea pig hearts, newly synthesized LPL can move from myocytes to the vascular lumen within 30 min [10]. The effect of diabetes to augment luminal LPL, in the presence of a reduced myocyte enzyme pool suggests an accelerated translocation of LPL from the myocyte cell surface to the vascular lumen. An alternative mechanism to explain this enlarged LPL observed during diabetes could be changes in luminal HSPG binding sites, as observed during hypoinsulinemia subsequent to 24 h fasting [11]. We hypothesize that following hypoinsulinemia, the heart switches to utilizing FA exclusively, and lipoprotein lipase is recruited to the myocardial endothelial lining via posttranslational mechanisms. In the present study we examined the regulation of luminal LPL in the diabetic heart by asking: a) Does cardiac malonyl-CoA have a role in FA delivery by way of regulating luminal LPL? b) Will the diabetic heart in vitro maintain its ability to translocate LPL from the myocyte cell surface to the vascular lumen? and c) Are changes in LPL related to an enhancement in the number of vascular LPL binding site? 39 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.2 Materials and Methods 2.2.1 Experimental animals and induction of hyperglycemia Male Wistar rats were cared for in accordance with the principles promulgated by the Canadian Council on Animal Care and the University of British Columbia. R-cell death with consequent hyperglycemia can be produced with a single iv dose of streptozotocin (STZ). Halothane-anesthetized rats were injected either with STZ (55 mg/kg iv) or an equivalent volume of saline. Following STZ, a triphasic pattern of blood glucose ensues [12]. This pattern comprises of an initial brief hyperglycemia followed by a period of hypoglycemia, before noticeable hyperglycemia is attained within 12-16 h [12]. Blood glucose then remains 2-3 times higher than normal. Although these animals are insulin deficient, they do not require insulin supplementation for survival, and do not develop ketoacidosis. Diabetic rats were kept for 4 days after STZ injection to subject them to chronic hyperglycemia, at which time they were euthanized, and hearts removed. Due to the prolonged time required to attain stable hyperglycemia, and the fact that metabolic switching in the heart from glucose to predominantly FA via LPL action likely occurs rapidly following hypoinsulinemia, we also pursued an alternate model of acute hyperglycemia to study LPL regulation. Diazoxide (DZ), a selective K + A T p channel opener inhibits secretion and causes hyperglycemia [13, 14] within one hour (chapter 3). DZ (100 mg/kg) was administered i.p., animals were euthanized after 4 h, and hearts removed. 2.2.2 Measurement of total cardiac LPL expression, protein and activity Rats were anesthetized with 65 mg/kg sodium pentobarbital i.p., the thoracic cavity opened, and the heart carefully excised. LPL gene expression was measured in the indicated groups using RT-PCR. Briefly, total RNA from hearts (100 mg) was extracted using Trizol (Invitrogen). After spectrophotometric quantification and resolving of RNA integrity using a formaldehyde agarose gel, reverse transcription was carried out using an oligo (dT) primer and superscript II 40 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 RT (Invitrogen). First strand cDNA was amplified using LPL specific primers [15]; 5'-A T C C A G C T G G G C C T A A C T T T - 3 ' (left) and 5 ' -AATGGCTTCTCCAATGTTGC-3 ' (right). The p-actin gene was amplified as an internal control using 5'-T G G T G G G T A T G G G T C AGAAGG-3 ' (left) and 5 ' - A T C C T G T C A G C G A T G C C T G G G - 3 ' (right). The linear range was found to be between 15-30 cycles. The amplification parameters were set at: 94°C for 1 min, 58°C for 1 min and 72°C for 1 min, for a total of 24 cycles. The PCR products were electrophoresed on a 1.7% agarose gel containing ethidium bromide. Expression levels were represented as the ratio of signal intensity for LPL mRNA relative to P-actin mRNA. LPL protein and activity measurement was done using 100 mg of homogenized ventricular tissue incubated with heparin (10 U/ml) for 30 min [16, 17]. The supernatant collected following centrifugation was then used either for protein (Western blot analysis) or activity measurements. Briefly, for Western blot analysis, total protein concentration was measured and 25 pg was size fractionated in a SDS-polyacrylamide gel, and blotted onto a nitrocellulose membrane. After blocking overnight at 4°C, the membrane was transferred to a solution of 1:1000 diluted primary antibody (5D2, a monoclonal mouse anti bovine LPL generously provide by Dr. J. Brunzell, University of Washington, Seattle, WA), and kept for 2 h at room temperature with gentle shaking. After washing with TBS-T, the membrane was treated with 1:3000 diluted secondary antibody (sheep anti-mouse IgG peroxidase-linked) for 1 h at room temperature. Following washing with TBS-T, the membrane was then incubated in enhanced chemiluminescent western blotting reagents for 1 min before exposing to a photographic film. Measurement of LPL activity in the supernatant was carried out using 100 pi of sample to hydrolyze a sonicated [3H] triolein substrate emulsion. 41 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.2.3 Immunolocalization of LPL Upon excision, control and STZ treated hearts were retrogradely perfused with non-circulating buffer for 3 mins to clear the heart of blood. Perfusion buffer was then changed to fixative (neutral phosphate-buffered 10% formalin) for 2 mins. Hearts were stored in 10% formalin for 24 hrs followed by paraffin processing through graded ethanol and xylene. Blocks were embedded in paraplast, sectioned at 3 um and mounted on positively charged glass slides. For immunostaining, sections were deparaffinized, rehydrated, and treated with 5% (vol/vol) heat inactivated rabbit plasma in Tris-buffered saline (TBS) to block nonspecific background. Sections were incubated with affinity-purified chicken antibovine LPL polyclonal antibody (1:400 dilution in TBS containing 1% (w/v) BSA) overnight at room temperature in a humid chamber. Samples were then washed with TBS and incubated for 1 hr at room temperature with the secondary biotinylated rabbit anti-chicken IgG (Chemicon Corp., 1:150 dilution), followed by incubation for 1 hr with streptavidin-conjugated Cy3 fluorescent probe (1:1000 dilution). The unbound fluorescent probe was rinsed with TBS buffer and sections mounted with DABCO. Slides were visualized using a Biorad 600 Confocal Microscope at 630x magnification. 2.2.4 Isolated heart perfusion Rats were not injected with heparin prior to killing, as it displaces LPL bound to HSPGs on the capillary endothelium. Consequently, it was necessary to remove and cannulate the heart as quickly as possible to avoid clotting of blood in the coronary arteries. Isolated hearts were immersed in cold (4°C) Krebs-Ringer HEPES buffer containing 10 mM glucose (pH 7.4). After the aorta was cannulated and tied below the innominate artery, hearts were perfused retrogradely by the nohrecirculating Langendorff technique as described previously [9]. Perfusion fluid was continuously gassed with 95% 02/5% C O 2 in a double-walled, water-heated chamber maintained at 37°C with a temperature-controlled circulating water bath. The flow rate was controlled at 7-8 42 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 ml/min. To measure endothelium-bound LPL, Langendorff perfusion solution was changed to Kreb's buffer containing 1% fatty acid free BSA and heparin (5 U/ml). The coronary effluent was collected in timed fractions over 5 min and assayed for LPL activity. In a separate experiment, hearts from different groups were perfused with heparin (to deplete the LPL pool at the coronary lumen), allowed to recover for 1 h during which the heart was perfused with heparin-free Krebs buffer, and a second 10 min perfusion with heparin was then performed to determine the extent of LPL recovery. 2.2.5 Purification and perfusion of bovine milk LPL Quick and reversible attachment of bovine LPL (bLPL) to endothelial cells of perfused rat hearts has previously been established [18]. To investigate if endothelial binding sites for LPL in the coronary vasculature of hearts from STZ-diabetic rats are altered, LPL was purified from fresh bovine milk by affinity chromatography [18, 19]. Briefly, skimmed milk was mixed overnight with heparin-agarose, the gel transferred to a column, and LPL enzyme eluted with 1.5 M NaCl. Peak activity fractions were pooled, and the purity of milk LPL established by 10% SDS-PAGE. Control and diabetic hearts were subjected to a 5-minute perfusion with Krebs buffer to clear the heart of blood. bLPL (0.9 ug/ml for 10 min; in preliminary experiments using control hearts, we determined this to be the optimal concentration and time required to saturate all the LPL binding sites) was then perfused through these hearts, chased with a 10 min perfusion of Krebs buffer to clear out excess bLPL, and a 4 min heparin perfusion was carried out to displace both exogenous (bovine) and endogenous (rat) LPL into the perfusate. 2.2.6 Treatments Phloridzin. In STZ-diabetic animals, the increase in cardiac LPL can be prevented or reversed by elimination of hyperglycemia with insulin. However, the effect of hyperglycemia per se on cardiac LPL is not known as insulin may alter several factors other than glucose. Phloridzin, an 43 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 inhibitor of the Na+-glucose co-transporter in proximal kidney tubules, normalizes blood glucose (without the assistance of insulin) by reducing tubular reabsorption of glucose, consequently increasing glycosuria [20-22]. Phloridzin (0.8 g/kg) dissolved in propylene glycol was injected intraperitoneally to STZ diabetic rats. In preliminary experiments, the maximal glucose lowering effect was observed after 12 h, with euglycemia persisting up to 24 h. Thus, animals were killed 12 hours after phloridzin, and cardiac LPL activity determined. Insulin. One caveat associated with DZ is that in addition to inhibiting insulin secretion, it also lowers blood pressure [13,14]. To determine whether the control of LPL by DZ is dependent on its lowering of insulin rather than its effects on blood pressure, rats were injected i.v. into the tail vein with a rapid acting insulin (Humulin R, Eli Lilly Canada Inc.), 1 h after DZ administration (following verification of hyperglycemia). Although different doses of insulin were evaluated, only the 2U and 8U insulin results are described. The animals were killed after 90 mins (time required for establishment of sustained euglycemia), and cardiac HR-LPL activity determined. Dichloroacetate (PCA). Dichloroacetate, by inhibiting pyruvate dehyrogenase kinase (PDK), a pyruvate dehydrogenase (PDH) inactivator, stimulates PDH [23-25]. Where indicated, DZ treated rats were administered D C A (100 mg/kg) in the presence or absence of 2U i.v. insulin through the right jugular vein. Animals were killed after 90 min and cardiac LPL activity determined. 2.2.7 Malonyl Co A and ACC estimation Malonyl CoA was assessed by a previously described method [26]. Briefly, hearts were removed, quickly washed in ice-cold Krebs buffer, frozen in liquid nitrogen, and stored at -70°C until analyzed. Samples of frozen tissue (-20 mg) were extracted in 200 ul (6%, w/v) perchloric acid and kept on ice for 10-15 min. After centrifugation (15 min, 10,000 g), protein-free acid supernatants were analyzed by HPLC. Elution of free CoASH and CoA esters was detected with 44 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 a flow-through monitor set at 254 nm. The column was calibrated with a range of CoA ester standards, assuming molar extinction coefficients of 14.9 and 14.4 mM'.cm"1 for acetyl-CoA and malonyl-CoA respectively. A C C content of frozen hearts was estimated as described previously [27]. Cytosolic fractions were prepared and subjected to SDS/PAGE and Western blot analysis using peroxidase labeled streptavadin. Blots were quantified by densitometry. Protein content was estimated using bicichoninic acid reagent. 2.2.8 Materials [ H] triolein was purchased from Amersham Canada. Heparin sodium injection (Hapalean; 1,000 USP U/ml) was obtained from Oraganon Teknika. All other chemicals were obtained from Sigma Canada. 2.2.9 Statistical analysis All data are reported as mean + SEM. One-way analysis of variance followed by the Newman-Keul's test or the unpaired Student's t-test was used to determine differences between group mean values. Changes in LPL activity in response to heparin perfusion, over time, were analyzed by multivariate analysis of variance followed by the Newman-Keul's test using the NCSS software. The level of statistical significance was set at P<0.05. 45 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.3 Resu l ts 2.3.1 LPL expression, protein, activity, and immunolocalization in whole hearts from STZ rats Four days of diabetes following STZ administration caused a decline in plasma insulin with ensuing hyperglycemia (Table 2-1). Other characteristics normally associated with hyperglycemia, such as polydipsia, were also observed in the STZ treated animals. Retrograde perfusion of control and STZ hearts with heparin resulted in rapid LPL discharge, and peak activity was observed within 1.5 mins. In hearts from STZ rats, peak LPL activity was almost 3-fold as much as control (Fig. 2-3). To determine whether this change in LPL is global, we measured LPL activity and protein in heart homogenates. Of note was the observation that LPL protein (Fig. 2-IB) and activity (Fig. 2-1C) in homogenates was not different between control and STZ diabetic hearts suggesting that LPL measurement using this approach may underestimate changes that occur at the coronary lumen. Immunofluorescence microscopy of myocardial sections was able to confirm a more intense LPL immunofluorescence in the diabetic myocardial blood vessels compared with control (Fig. 2-2). Interestingly, changes in luminal LPL activity were independent of shifts in mRNA levels suggesting a posttranslational increase in LPL at the luminal surface (Fig. 2-1 A). 2.3.2 LPL binding sites in the coronary lumen Irrespective of the mechanism involved, STZ diabetes (4 day), fasting (14 hours) (data not shown) or DZ (4 hours) all independently increase peak HR-LPL activity uniformly (-1200 nmol/ml/h). These data suggest that there are a finite number of LPL binding sites in the coronary lumen of the control rat heart that are only partly filled with the enzyme. To test this, both control and STZ hearts were perfused with bLPL; as rat and bLPL demonstrate significant sequence homology, exogenous milk LPL is expected to attach to the same binding sites on endothelial cells in the heart that bind endogenous rat enzyme [18]. Perfusion of control hearts 46 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 with bLPL increased peak LPL activity released (following heparin) by 3-fold (Fig. 2-3; left panel). As perfusion of diabetic hearts with bLPL was unable to further increase the amount of peak HR-LPL activity (Fig. 2-3; right panel), it is likely that luminal LPL binding sites remain unchanged following diabetes. Of note was the observation that subsequent to bLPL perfusion, although control and STZ hearts demonstrated similar peak LPL activity, the heparin release profile was distinct. Whether this altered profile is an outcome of some non-specific bLPL binding, or more likely a combination of an accumulation of myocyte rat LPL at the endothelial basolateral surface or transfer and buildup of bLPL at the abluminal surface is presently being studied. 2.3.3 In vitro LPL translocation Following STZ there is a triphasic pattern of blood glucose comprising of an initial brief hyperglycemia followed by a period of hypoglycemia before noticeable hyperglycemia is attained within 12-16 hrs [12]. As metabolic switching in the heart from glucose to predominantly FA likely occurs rapidly following hypoinsulinemia, we pursued a model of acute hyperglycemia to study LPL regulation. DZ, a selective K + A T p channel opener lowers blood pressure and inhibits insulin secretion (Table 2-1). Lowering insulin augmented plasma glucose and resulted in stable hyperglycemia within 1-h (data not shown), persisted for 4 h and blood glucose returned to normal within 7 h. In some anesthetized rats, DZ produced a significant fall in mean carotid artery blood pressure (~20 mmHg). This effect was transient, and blood pressure returned to normal within 3 h (data not shown). Given that DZ is a K +ATP channel opener with potential direct effects on the heart, we also examined whether DZ could directly influence cardiac HR-LPL activity. Isolated hearts when perfused in the absence or presence of DZ (upto 100 pM, lh), followed by heparin, did not significantly change peak LPL activity (data not shown) and rate and pattern of contraction. Even 4-h of DZ induced hyperglycemia was 47 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 sufficient to cause a substantial increase in coronary LPL activity, albeit lower when compared to 4-day STZ diabetes (Fig. 2-4, left panel). When the coronary LPL pool from control, STZ or DZ treated rats was depleted with heparin and allowed to refill for 1 h, only STZ hearts continued to demonstrate higher LPL activity on second heparin perfusion (Fig. 2-4, right panel). It should be noted that subsequent to in vitro LPL depletion after first heparin perfusion, complete recovery of the luminal LPL is not possible within 1 h. As this effect of accelerated replenishment was not observed following DZ, it implies that long-term hyperglycemia induces an intrinsic metabolic transformation in the heart allowing it to maintain augmented LPL under in vitro conditions. Whether this metabolic transformation is in part due to an accelerated transfer of enzyme from the myocyte to the vascular lumen, chronic hyperglycemia mediated LPL assembly at the abluminal surface or an ability of heparin to not only release but also recruit enzyme towards the luminal surface in the chronically diabetic rat, possibly due to endothelial dysfunction, is currently under investigation. 2.3.4 Regulation of LPL by insulin In an attempt to evaluate the contribution of hyperglycemia per se on this metabolic transformation, STZ rats were treated with phloridzin. Treatment with phloridzin restored normoglycemia within 12 h, and this effect persisted for 24 h (Fig. 2-5, inset). As expected, plasma insulin concentrations in STZ rats were unaffected by phloridzin treatment (data not shown). Correction of hyperglycemia with phloridzin was unable to reduce cardiac LPL activity in hearts from STZ rats (Fig. 2-5). We also examined the effect of insulin induced normoglycemia on LPL activity in rats made hyperglycemic with DZ and STZ. Following DZ treatment for 60 mins to induce hyperglycemia, rats were injected with 8U of rapid-acting insulin intravenously. Insulin induced normoglycemia in these hyperglycemic rats within 30 min (~5.0 mM), that lasted for 90 min (Fig. 2-6, right panel). Injection of this dose of insulin also 48 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 significantly attenuated HR-LPL activity to control levels (Fig. 2-6, left panel). Administration of 2U of insulin to DZ-hyperglycemic rats did not significantly reduce hyperglycemia for the initial 30 min (~18.0 mM). However insulin did normalize plasma glucose within 90 min; (Fig. 2-6, right panel) surprisingly, HR-LPL activity in these rats remained elevated (Fig. 2-6, left panel). Whether this inability of 2U insulin to abolish high LPL activity is related to the gradual decline in hyperglycemia is unclear. We were interested in reproducing this differential effect of insulin on LPL in STZ diabetic rats. Analogous to results observed after DZ, administration of 2U of insulin to STZ-diabetic rats normalized plasma glucose but did not abrogate the elevated HR-LPL activity (data not shown). 2.3.5 Regulation of LPL by cardiac malonyl-CoA and PDH In liver, fat and skeletal muscle, malonyl-CoA typically increases during exposure to insulin and glucose, following activation of the reaction catalyzed by A C C [28]. We determined malonyl-CoA levels in the heart following the disparate doses of insulin used to lower plasma glucose in DZ treated rats. Although marginally lower compared to control, we could not detect any significant change in malonyl-CoA levels following DZ (Fig. 2-7). Interestingly, 8U but not 2U of insulin, injected into DZ treated rats, markedly increased cardiac malonyl CoA levels (Fig. 2-7). Insulin had no significant effect on A C C protein expression (as shown by densitometry; con, 12398±985; DZ, 13737±3810; DZ+I2, 19236±5115; DZ+I8, 16312±4030, units) (Fig. 2-7, inset). Previous reports suggest that a fall in cardiac malonyl CoA is related to increased malonyl CoA decarboxylase (MCD) activity and not to changes in A C C [29, 30]. As insulin treatment did not alter A C C expression, changes in M C D might explain the high malonyl CoA. PDH contributes importantly to control of the rate of glucose oxidation [24,25]. Co-administration of D C A and 2U insulin normalized LPL activity in DZ treated animals (Fig. 2-8). DC A alone in DZ treated rats did not change cardiac LPL (data not shown). 49 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.4 D i s c u s s i o n Under normal aerobic conditions, FA and glucose are the major providers of energy [1, 31]. However, during diabetes, metabolism of FA becomes the preferred means of energy supply [3]. To make available this increased requirement of the heart for FA, augmentation of adipose tissue lipolysis and accelerated hydrolysis of intracellular cardiac T G stores have been described in the STZ diabetic rat. Until recently, the contribution of cardiac LPL in providing FA to the diabetic heart was unclear as the majority of studies used heart homogenates that were unable to determine distribution of the enzyme. Using diabetic heart homogenates, we were also unable demonstrate any significant changes in LPL mRNA, protein or activity when compared to control. However, as reported previously, 4-day STZ hearts exhibited elevated LPL activity and immunofluroscence at the coronary luminal surface. More interestingly, DZ induced amplification of HR-LPL activity occurred within 4 h of acute hyperglycemia suggesting that posttranslational processing must predominantly control this luminal increase in enzyme. As LPL mediated hydrolysis of circulating TG-rich lipoproteins was suggested to be the principal source of FA for cardiac utilization [32], and given our data of a rapid increase of LPL following DZ, luminal LPL may be playing a predominant role in FA delivery in a setting of hypoinsulinemia. HSPGs are the most numerous and extensively expressed binding sites that attach LPL with high association and dissociation rate constants [33, 34]. Using 125I-labeled bLPL, fasting with attendant hypoinsulinemia was shown to increase the ability of the coronary lumen to bind LPL [11, 33, 34]. However, our present and previous data [9,12] were indicative of a model whereby coronary lumen of the rat heart had a finite number of LPL binding sites that are normally only partly filled. To determine whether the enhanced HR-LPL activity subsequent to STZ diabetes was due to rapid filling of these binding sites, control and STZ hearts were perfused with bLPL. 50 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 In control hearts, perfusion with bLPL increased peak HR-LPL activity to a level identical to that observed with STZ diabetes. As a similar perfusion of bLPL through the STZ heart was ineffective in producing any further increase of enzyme, our data suggest that in the control heart, only a fraction of these binding sites are occupied by LPL, and hyperglycemia is able to fill all of these sites with the enzyme. In the heart, LPL is synthesized in myocytes and subsequently transported onto HSPG binding sites on the myocyte cell surface [35]. When myocardial distribution of immunoreactive LPL protein was studied using electron microscopy of immunogold-labeled sections of mouse heart, 78% of total LPL was present in the cardiac myocytes, 3-6% in the interstitial space, and 18% in the capillary endothelium [36, 37]. At least in perfused guinea pig hearts, newly synthesized LPL can move from myocytes to the vascular lumen within 30 min [10]. During excessive luminal requirement for LPL, an anticipated translocation of the enzyme from the myocyte cell surface to the vascular lumen would be predicted. Indeed subsequent to 2 week STZ [9] or 4 h of DZ (chapter 3) administration, a 50-70% drop in surface bound myocyte LPL was observed at a time when luminal LPL activity was elevated. Given the observation that LPL in heart is predominantly located at the myocyte, we tested the significance of this decreased myocyte cell surface LPL pool in vitro using isolated hearts. Interestingly, when the luminal HR-LPL pool was allowed to recover for 1 h after removal of the enzyme, isolated hearts from STZ but not DZ continued to demonstrate a higher peak LPL activity. This suggests that a mechanism to increase coronary luminal LPL following diabetes persists after isolation of the heart, probably involves accelerated transfer of the enzyme and is only evident with chronic hyperglycemia. Both experimental and clinical hyperglycemia have been shown to induce "diabetic memory" [38]. However, as smooth muscle derived LPL can translocate to the endothelial surface of vascular tissue, the contribution of this LPL pool in explaining the effect 51 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 of DZ and diabetes cannot be completely disregarded. Similarly, at least in human myocardium, LPL protein was detected on interstitial cells and could be equally contributive in explaining the effects of DZ and diabetes [35]. The heart has a unique ability to utilize diverse substrates, to preferentially utilize a particular substrate during pathophysiological conditions, and to perform this metabolic transformation rapidly. Thus lactate is predominantly utilized following exercise [39, 40], glucose is metabolized in greater quantities by the hypertrophic heart [41] and impaired glucose utilization following diabetes allows a rapid increase in endothelial LPL derived FA supply to the heart [3, 42]. As metabolic switching in the heart from glucose to predominantly FA likely occurs rapidly following DZ induced hypoinsulinemia, it is a superior model to study cardiac LPL regulation. In the current study, both 2 and 8 units of insulin normalized plasma glucose in DZ treated animals but only 8U reversed DZ induced augmentation of cardiac luminal LPL. During hyperglycemia, and following the injection of insulin, the central tissue increasing its glucose uptake is skeletal muscle [43-45]. Increasing glucose utilization can lead to elevated concentrations of malonyl-CoA [42] that inhibits CPT-1 activity thereby preventing FA oxidation [28]. As a robust increase in cardiac malonyl CoA was observed only with 8U insulin, supraphysiological doses of insulin might have caused extensive glucose uptake into the skeletal muscle and the heart, thereby abolishing luminal LPL activity. Inability of 2U insulin to restrain the high cardiac LPL despite normoglycemia could be an outcome of preferential glucose utilization in skeletal muscle. Another caveat associated with 2U insulin treatment is that glucose entering the heart may not be metabolized due to persistent PDH inhibition while the heart is exclusively utilizing FA. Interestingly, DZ treated animals given 2U insulin and simultaneously treated with DCA (PDH activator) demonstrated decreased LPL activity. Our data suggests that enhanced glucose metabolism subsequent to increase in malonyl CoA or 52 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 activation of PDH may well turn off the machinery operating to supply the heart with excess exogenous FA. In summary, following hypoinsulinemia/hyperglycemia, the heart switches energy production to exclusive p-oxidation of FA. This metabolic transformation in part is facilitated by amplification of LPL at a "functionally" relevant location, the coronary luminal surface. As these alterations in luminal LPL occurred rapidly, posttranslational mechanisms probably regulate this enzyme at the vascular endothelial surface. A rapid vectorial transfer of LPL to unoccupied binding sites is a likely mechanism for LPL augmentation at the luminal surface. Given the persistence of increased coronary luminal LPL even in a setting of normoglycemia, our data suggest that this setting may well provide excessive FA to the heart with deleterious consequences over the long term. High LPL in skeletal and cardiac muscle have been reported to induce insulin resistance and myopathy [46-48]. In a more recent study using genetically engineered mice that specifically overexpressed cardiomyocyte surface bound LPL, lipid oversupply and impaired contractile function was observed [49]. 53 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.5 T a b l e s a n d F igures TABLE 2-1 General characteristics of the animals Control STZ DZ Body Weight 295±8 265±7 274±8 (g) Plasma Insulin 2.3±0.4 0.9±0.1* 1.3+0.2* (ng/ml) Plasma Glucose 8.7+0.2 24.2+1.7* 25.0+1.8* (mM) Values are means ± SE for 6-10 animals. Diabetes was induced by injection of STZ (55 mg/kg, i.v.) and the animals killed 4 days later. To induce acute hypoinsulinemia, animals were treated with DZ (100 mg/kg, i.p.) and the rats killed after 4 h. *Significantly different from control, P < 0.05. 54 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 A 0.20 CON STZ Fig. 2-1 Hearts from control and STZ diabetic animals were isolated and L P L gene expression, protein and activity measurement were done as described in methods using 100 mg of homogenized ventricular tissue. Results are the means ± SE of 4animals in each group. 55 Chapter 2: J. Mol. Cell. CardioL 35 (2003) 1093-1103 CON r 4 *** t F i g . 2-2 Representative photographs showing the effect of STZ (4 days) on LPL immunofluorescence as visualized by fluorescent microscopy. Heart sections were fixed, incubated with the polyclonal chicken antibody against bovine LPL followed by incubations with biotinylated rabbit anti-chicken IgG and streptavidin-conjugated Cy3 fluorescent probe respectively. Majority of LPL in the STZ heart was exclusively present in the coronary lumen (arrows); CON, Control, STZ, Streptozotocin. 56 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 CON STZ 2000 1 g 1500 c CD CD | j 1000 CD CL .~ 500 > '-4—» O < o H + bLPL -bLPL 4 A + bLPL A -bLPL 2000 1500 o 0 •*—• 1000 ^ t: Q. 500 ^ > "-4—' O < Perfusion Time (rnin) Fig. 2-3 Effect of STZ on LPL endothelial binding sites in the coronary vasculature of isolated hearts. LPL was purified from fresh bovine milk by affinity chromatography. Control and STZ hearts were then retrogradely perfused with bLPL (0.9 ug/ml for 10 min). Following a 10 min wash perfusion with Krebs buffer, perfusion with heparin was carried out to displace both endogenous (rat) and exogenous (bovine) LPL into the perfusate. Results are the mean ± SE of 5 rats in each group. 57 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 1st Heparin 2nd Heparin 0 1 2 3 4 5 0 2 4 6 8 10 12 Perfusion Time (min) Perfusion Time (min) Fig. 2-4 Graph showing the in vitro effect of two heparin perfusions on LPL activity in isolated rat hearts. Hearts from different groups were removed, perfused with heparin (5 U/ml) for 5 min (left panel) and fractions of perfusate analyzed for LPL activity. This was followed by a 1 h wash with heparin free Krebs buffer. A second perfusion for 10 min with heparin was then carried out (right panel). Results are the mean ± SE of 6 rats in each group. CON, control; STZ, 4 day diabetic; DZ, diazoxide given for 4 h. 58 Chanter2:.J. Mol. Cell. Cardiol. 35(2003) 1093-1103 16 - , 0 2 4 6 8 10 12 Perfusion Time (min) Fig . 2-5 Effect of phloridzin treatment on LPL activity in perfused hearts from STZ rats. Phloridzin (PHZ) was injected i.p. and animals killed 12 hours later for determination of LPL activity. The inset indicates plasma glucose levels of diabetic rats measured from tail vein blood samples obtained at various times after phloridzin injection. CON, control; STZ, 4 day diabetic; PHZ, STZ rats injected ip with phloridzin (0.8 g/kg). 59 1200 i o E 0 -t—» CD C/5 t CD Q_ o < 800 400 o^ o C/ia/rte/- 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 r 100 2 4 6 Time (min) 8 10 80 60 40 r 20 O Z ) < cu co o o _5 O CD E w _CD Q_ Fig . 2-6 L o CON DZ DZ DZ + + I 2 I 8 Effect of different doses of insulin (2 and 8U) on plasma glucose and cardiac LPL activity from DZ treated rats. Insulin was administered 1 h after DZ-induced hyperglycemia, and rats killed after 90 min. The right panel illustrates the integrated area under the curve for plasma glucose measured over 150 minutes. The left panel describes LPL activity measured 90 min after insulin administration. Results are the means ± SE of 5 rats in each group. * Significantly different from all other groups, P < 0.05. 60 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 15 i o 110 < o o o co CO O 0 ACC C O N DZ+I2 D Z J L CON DZ DZ+I8 DZ + I 2 DZ + I 8 Fig . 2-7 Effect of different doses of insulin (2 and 8U) on malonyl CoA in hearts from DZ treated rats. Insulin was administered 1 h after DZ-induced hyperglycemia, and rats killed after 90 min. Malonyl CoA was measured in cytosolic fractions of cardiac tissue homogenates using HPLC. Results are the means ± SE of 5 rats in each group. *Significantly different from all other groups, P < 0.05. The inset shows a Western blot for A C C using peroxidase labeled streptavadin. 61 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 1200 -i ^ 1000 E o E c •+-» CD it) CD Q_ _c >» -4—* < 800 A 600 4 400 \ 200 4 0^ heparin 0 2 DZ + 12 DZ + 12 + DCA 4 6 8 10 i 12 Time (min) Fig . 2-8 Effect of D C A in moderating the effects of 2U insulin on LPL activity. After DZ, 2U insulin and DCA (100 mg/kg) were simultaneously administered, and animals killed after 90 min for measurement of HR-LPL activity. Results are the means ± SE of 5 rats in each group. 62 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 2.6 B ib l iography [1] Randle PJ, Hales CN, Garland PB, Newsholme EA. The glucose fatty-acid cycle; its role in insulin sensitivity and the metabolic disturbances of diabetes mellitus. Lancet 1963;1:785-89. [2] Sidossis LS, Stuart CA, Shulman Gl, Lopaschuk GD, Wolfe RR. Glucose plus insulin regulate fat oxidation by controlling the rate of fatty acid entry into the mitochondria. J Clin Invest 1996;98:2244-50. [3] Rodrigues B, Cam M C , McNeill JH. Myocardial substrate metabolism: implications for diabetic cardiomyopathy. J Mol Cell Cardiol 1995;27:169-79. [4] Pillarisetti S, Saxena U. Lipoprotein lipase as a therapeutic target for dyslipidemia. Front Biosci 2003;8:D238-41. [5] Preiss-Landl K, Zimmermann R, Hammerle G, Zechner R. Lipoprotein lipase: the regulation of tissue specific expression and its role in lipid and energy metabolism. Curr Opin Lipidol 2002;13:471-81. [6] Saxena U, Klein M G , Goldberg IJ. Transport of lipoprotein lipase across endothelial cells. Proc Natl Acad Sci U S A 1991;88:2254-8. [7] Stins MF, Maxfield FR, Goldberg IJ. Polarized binding of lipoprotein lipase to endothelial cells. Implications for its physiological actions. Arterioscler Thromb 1992;12:1437-46. [8] Braun JE, Severson DL. Diabetes reduces heparin- and phospholipase C-releasable lipoprotein lipase from cardiomyocytes. Am J Physiol 1991;260:E477-85. [9] Rodrigues B, Cam M C , Jian K, Lim F, Sambandam N, Shepherd G. Differential effects of streptozotocin-induced diabetes on cardiac lipoprotein lipase activity. Diabetes 1997;46:1346-53. [10] Liu GQ, Olivecrona T. Pulse-chase study on lipoprotein lipase in perfused guinea pig heart. Am J Physiol 1991;261:H2044-50. 63 Chapter 2: J. Mol. Cell. 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[15] Brault D, Noe L, Etienne J, Hamelin J, Raisonnier A, Souli A, Chuat JC, Dugail I, Quignard-Boulange A, Lavau M , et al. Sequence of rat lipoprotein lipase-encoding cDNA. Gene 1992;121:237-46. [16] Klingenspor M , Ebbinghaus C, Hulshorst G, Stohr S, Spiegelhalter F, Haas K, Heldmaier G. Multiple regulatory steps are involved in the control of lipoprotein lipase activity in brown adipose tissue. J Lipid Res 1996;37:1685-95. [17] Vaziri ND, Liang K, Barton CH. Effect of increased afterload on cardiac lipoprotein lipase and V L D L receptor expression. Biochim Biophys Acta 1999;1436:577-84. [18] Liu L, Severson DL. Endothelial binding sites for lipoprotein lipase are not diminished in perfused hearts from diabetic rats. Can J Physiol Pharmacol 1996;74:1204-9. [19] Ramirez I, Kryski AJ, Ben-Zeev O, Schotz M C , Severson DL. Characterization of triacylglycerol hydrolase activities in isolated myocardial cells from rat heart. Biochem J 1985;232:229-36. 64 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 [20] Brichard SM, Henquin JC, Girard J. Phlorizin treatment of diabetic rats partially reverses the abnormal expression of genes involved in hepatic glucose metabolism. Diabetologia 1993;36:292-8. [21] Rossetti L, Giaccari A, DeFronzo RA. Glucose toxicity. Diabetes Care 1990;13:610-30. [22] Wen SF. Effect of phlorhizin on renal glucose and phosphate transport in the dog. Clin Sci (Lond) 1979;57:367-74. [23] Whitehouse S, Cooper RH, Randle PJ. Mechanism of activation of pyruvate dehydrogenase by dichloroacetate and other halogenated carboxylic acids. Biochem J 1974;141:761-74. [24] Wambolt RB, Lopaschuk GD, Brownsey RW, Allard MF. Dichloroacetate improves postischemic function of hypertrophied rat hearts. J Am Coll Cardiol 2000;36:1378-85. [25] Stacpoole PW. The pharmacology of dichloroacetate. Metabolism 1989;38:1124-44. [26] Boone A N , Rodrigues B, 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 1999;341:347-54. [27] Saddik M , Gamble J, Witters L A , Lopaschuk GD. Acetyl-CoA carboxylase regulation of fatty acid oxidation in the heart. J Biol Chem 1993;268:25836-45. [28] Ruderman NB, Saha AK, Vavvas D, Witters LA. Malonyl-CoA, fuel sensing, and insulin resistance. Am J Physiol 1999;276:E1-E18. [29] Longnus SL, Wambolt RB, Barr RL, Lopaschuk GD, Allard MF. Regulation of myocardial fatty acid oxidation by substrate supply. Am J Physiol Heart Circ Physiol 2001;281:H 1561-7. [30] Sakamoto J, Barr RL, Kavanagh K M , Lopaschuk GD. Contribution of malonyl-CoA decarboxylase to the high fatty acid oxidation rates seen in the diabetic heart. Am J Physiol Heart Circ Physiol 2000;278:H1196-204. 65 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 [31] Randle PJ, Priestman DA, Mistry S, Halsall A. Mechanisms modifying glucose oxidation in diabetes mellitus. Diabetologia 1994;37 Suppl 2:S155-61. [32] Augustus AS, Kako Y, Yagyu H, Goldberg IJ. Routes of FA delivery to cardiac muscle: modulation of lipoprotein lipolysis alters uptake of TG-derived FA. Am J Physiol Endocrinol Metab 2003;284:E331-9. [33] Saxena U, Klein M G , Goldberg IJ. Metabolism of endothelial cell-bound lipoprotein lipase. Evidence for heparan sulfate proteoglycan-mediated internalization and recycling. J Biol Chem 1990;265:12880-6. [34] Cisar L A , Hoogewerf AJ, Cupp M , Rapport CA, Bensadoun A. Secretion and degradation of lipoprotein lipase in cultured adipocytes. Binding of lipoprotein lipase to membrane heparan sulfate proteoglycans is necessary for degradation. J Biol Chem 1989;264:1767-74. [35] Ailhaud G. Cellular and secreted lipoprotein lipase revisited. Clin Biochem 1990;23:343-47. [36] Blanchette-Mackie EJ, Dwyer NK, Amende LA. Cytochemical studies of lipid metabolism: immunogold probes for lipoprotein lipase and cholesterol. Am J Anat 1989;185:255-63. [37] Blanchette-Mackie EJ, Masuno H, Dwyer NK, Olivecrona T, Scow RO. Lipoprotein lipase in myocytes and capillary endothelium of heart: immunocytochemical study. Am J Physiol 1989;256:E818-28. [38] Brownlee M . Biochemistry and molecular cell biology of diabetic complications. Nature 2001;414:813-20. [39] Gibala MJ, Young M E , Taegtmeyer H. Anaplerosis of the citric acid cycle: role in energy metabolism of heart and skeletal muscle. Acta Physiol Scand 2000;168:657-65. [40] Hultman E, Greenhaff PL. Skeletal muscle energy metabolism and fatigue during intense exercise in man. Sci Prog 1991;75:361-70. [41] Sambandam N, Lopaschuk GD, Brownsey RW, Allard MF. Energy metabolism in the hypertrophied heart. Heart Fail Rev 2002;7:161-73. 66 Chapter 2: J. Mol. Cell. Cardiol. 35 (2003) 1093-1103 [42] Lopaschuk GD, Rebeyka IM, Allard MF. Metabolic modulation: a means to mend a broken heart. Circulation 2002;105:140-2. [43] Gerich JE. Physiology of glucose homeostasis. Diabetes Obes Metab 2000;2:345-50. [44] Corssmit EP, Romijn JA, Sauerwein HP. Review article: Regulation of glucose production with special attention to nonclassical regulatory mechanisms: a review. Metabolism 2001;50:742-55. [45] Yki-Jarvinen H. Action of insulin on glucose metabolism in vivo. Baillieres Clin Endocrinol Metab 1993;7:903-27. [46] Kim JK, Fillmore JJ, Chen Y, Yu C, Moore IK, Pypaert M , Lutz EP, Kako Y , Velez-Carrasco W, Goldberg IJ, Breslow JL, Shulman GI. Tissue-specific overexpression of lipoprotein lipase causes tissue-specific insulin resistance. Proc Natl Acad Sci U S A 2001;98:7522-7. [47] Voshol PJ, Jong M C , Dahlmans V E , Kratky D, Levak-Frank S, Zechner R, Romijn JA, Havekes L M . In muscle-specific lipoprotein lipase-overexpressing mice, muscle triglyceride content is increased without inhibition of insulin-stimulated whole-body and muscle-specific glucose uptake. Diabetes 2001;50:2585-90. [48] Levak-Frank S, Radner H, Walsh A, Stollberger R, Knipping G, Hoefler G, Sattler W, Weinstock PH, Breslow JL, Zechner R. Muscle-specific overexpression of lipoprotein lipase causes a severe myopathy characterized by proliferation of mitochondria and peroxisomes in transgenic mice. J Clin Invest 1995;96:976-86. [49] Yagyu H, Chen G, Yokoyama M , Hirata K, Augustus A, Kako Y, Seo T, Hu Y, Lutz EP, Merkel M , Bensadoun A, Homma S, Goldberg IJ. Lipoprotein lipase (LpL) on the surface of cardiomyocytes increases lipid uptake and produces a cardiomyopathy. J Clin Invest 2003;111:419-26. 67 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.0 Circulating triglyceride lipolysis facilitates lipoprotein lipase translocation from cardiomyocyte to myocardial endothelial lining 3.1 Introduction Under physiological conditions, cardiac tissue acquires most of its energy from metabolism of two major substrates, glucose and FA, the latter being the preferred substrate consumed [1,2]. Because the heart has no potential to synthesize FA, it is dependent upon supply, with hydrolysis of TG-rich lipoproteins by LPL positioned at the endothelial surface of the coronary lumen [3] being suggested to be the principal source of F A for cardiac utilization [4]. Endothelial cells do not synthesize LPL and hence the enzyme is synthesized in cardiomyocytes [5,6]. LPL secreted as an active enzyme binds to myocyte cell surface HSPG, before it is translocated onto comparable HSPG binding sites on the luminal side of the vessel wall [7-9]. In perfused guinea pig hearts, LPL can move from myocytes to the vascular lumen within 30 min [10] by mechanisms that are not clearly understood. At least in co-culture experiments using adipocytes and endothelial cells, heparanase like compounds secreted from endothelial cells have been suggested to release subendothelial HS bound proteins and specific HS oligosaccharides that serve as extracellular chaperones allowing LPL to be transported across the interstitial space [11]. TG-rich lipoproteins and oleic acid release adipocyte cell surface LPL [12,13]. The authors concluded (but were unable to confirm) that this released LPL from adipocyte cell surface would then be transported to the luminal endothelial cell surface. With adipose tissue (as compared to heart) it is difficult to study LPL movement from underlying parenchymal cells to the vascular lumen. In addition to FA, lysophosphatidylcholine (LPC), a component of lipoprotein phospholipids, is also released during LPL mediated lipolysis of T G rich lipoproteins and would be expected to be augmented under conditions of extensive lipoprotein T G breakdown [14-16]. 68 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 Endothelial cells exposed to LPC release heparanase like compounds that cleave cell surface HSPG-bound LPL, at least in adipocytes [11]. The present study was designed to investigate the influence of circulating T G and their lipolysis in facilitating translocation of LPL from the underlying cardiomyocyte cell surface to the coronary lumen. 69 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.2 Methods 3.2.1 Experimental animals The investigation conforms with the guide for the care and use of laboratory animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996). Animals were cared for in accordance with the principles promulgated by the Canadian Council on Animal Care and the University of British Columbia. Adult male Wistar rats (270-290 g) were obtained from the UBC Animal Care Unit and maintained under a 12-h light (0700-1900)/dark cycle and supplied with a standard laboratory diet (PMI Feeds, Richmond, VA) and water ad libitum. 3.2.2 Diazoxide induced acute hyperglycemia D Z , a selective K +ATP channel opener decreases insulin secretion and causes hyperglycemia within one hour [17-19]. More importantly, changes in glucose are also associated with rapid elevation of serum FA and TG. D Z (25-100 mg/kg) was administered i.p. at 10 A M and tail vein blood samples obtained at various times after D Z injection. Animals were euthanized 4 h after D Z , and hearts removed. 3.2.3 Isolated heart perfusion Rats were anesthetized with 65 mg/kg sodium pentobarbital i.p., the thoracic cavity opened, and the heart carefully excised. The heart was immersed in cold (4°C) Krebs-Ringer HEPES buffer containing 10 mM glucose (pH 7.4). After the aorta was cannulated and tied below the innominate artery, hearts were perfused retrogradely by the nonrecirculating Langendorff technique as described previously [20]. Perfusion fluid was continuously gassed with 95% O2 /5% C O 2 in a double-walled, water-heated chamber maintained at 37°C with a temperature-controlled circulating water bath. The flow rate was controlled at 7-8 ml/min. 70 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.2.4 Coronary lumen LPL activity To measure endothelium-bourid LPL, Langendorff perfusion solution was changed to Krebs-Ringer HEPES buffer containing 1% fatty acid free BSA and heparin (5 U/ml). This concentration of heparin can maximally release cardiac LPL from its binding sites. The coronary effluent was collected in timed fractions over 10 min and assayed for LPL activity by measuring the hydrolysis of a sonicated [3H] triolein substrate emulsion [21]. LPL activity is expressed as nanomoles oleate released per hour per milliliter. Subsequent to LPL displacement with heparin, hearts were blotted dry and wet weights recorded (0.56±0.03 g). 3.2.5 Immunolocalization of LPL Upon excision, control and DZ treated hearts were retrogradely perfused with non-circulating buffer for 3 mins to clear the heart of blood. Perfusion buffer was then changed to fixative (neutral phosphate-buffered 10% formalin) for 2 mins. Hearts were stored in 10% formalin for 24 hrs followed by paraffin processing through graded ethanol and xylene. Blocks were embedded in Paraplast, sectioned at 3 um and mounted on positively charged glass slides. For immunostaining, sections were deparaffinized, rehydrated, and treated with 5% (vol/vol) heat inactivated rabbit plasma in Tris-buffered saline (TBS) to block nonspecific background. Sections were then incubated with affinity-purified chicken antibovine LPL polyclonal antibody (1:400 dilution in TBS containing 1% (w/v) BSA) overnight at room temperature in a humid chamber. Samples were then washed with TBS and incubated for 1 hr at room temperature with the secondary biotinylated rabbit anti-chicken IgG (Chemicon Corp., 1:150 dilution), followed by incubation for 1 hr with streptavidin-conjugated Cy3 fluorescent probe (1:1000 dilution). The unbound fluorescent probe was rinsed with TBS buffer and sections were mounted with DABCO. Slides were visualized using a Biorad 600 Confocal Microscope at 630x magnification. An absence of staining was observed when the primary antibody was omitted or replaced by preimmune chicken serum. 71 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.2.6 LPL content of cardiomyocytes In addition to endothelial-bound LPL, a large pool of enzyme activity is still measurable within the heart, located predominantly within myocytes. Ventricular calcium-tolerant myocytes were prepared by a previously described procedure [20]. Briefly, myocytes were made calcium-tolerant by successive exposure to increasing concentrations of calcium. Our method of isolation yields a highly enriched population of calcium-tolerant myocardial cells that are rod-shaped in the presence of 1 mM C a 2 + with clear cross striations. Intolerant cells are intact but hypercontract into vesiculated spheres. Yield of myocytes (cell number was approximately similar in both control and DZ hearts, -4.8 x 106) was determined microscopically using an improved Neubauer haemocytometer. Myocyte viability (generally between 75-85% in control and DZ hearts) was assessed as the percentage of elongated cells with clear cross striations that excluded 0.2% trypan blue. Cardiac myocytes were suspended at a final cell density of 0.4 x 106 cells/ml, incubated at 37°C, and basal LPL activity in the medium and cell pellet (after centrifugation) measured. To release surface-bound LPL activity, heparin (5 U/ml) was added to the myocyte suspension. After incubation for 10 min, an aliquot of cell suspension was removed, medium separated by centrifugation in an eppendorf microcentrifuge (1 min, 10,000g), and assayed for LPL activity. The total basal cellular LPL activity was measured by sonicating (Vibra Cel l T m Sonicator at a frequency of 40 Hz for 2 x 30 sees) the cell pellets after resuspending them in 0.2 ml of 50 mmol/1 N H 4 C I buffer (pH 8.0) containing 0.125% (vol/vol) Triton X-100. After sonication, the volume was adjusted to 1 ml using a sucrose buffer (0.25 mol/1 sucrose, 1 mmol/1 EDTA, 1 mmol/1 dithiothreitol, 10 mmol/1 HEPES, pH 7.4). Assay for cell sonicate LPL activity was done using 20 pi of the cell sonicate and heparin (2 U/ml) was included in the assay. 72 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.2.7 Treatments Table 3-2 summarizes the experimental design for the indicated treatments given below. Insulin: In the indicated experiment, rats were injected into the tail vein with a rapid acting insulin (HumulinR, 8 units), 1 h after DZ administration (following verification of hyperglycemia). The animals were killed after 90 mins (time required for establishment of sustained euglycemia), and cardiac heparin-releasable LPL activity determined. Lysophosphatidylcholine palmitoyl (LPC): In the indicated experiment control rat hearts were perfused with L-ct-lysophosphatidylcholine palmitoyl (LPC) (0.1 nM-100 nM) for 60 mins. Cardiac HR-LPL activity was subsequently determined. Triton WR1339: WR 1339, a non-ionic detergent, physically alters lipoproteins making them inaccessible for LPL mediated hydrolysis [22,23]. When injected intravenously, newly synthesized TGs accumulate in the plasma. Rats were injected (i.v.) with WR1339 (25% w/v solution in normal saline to give a dose of 600 mg/kg body weight). WR 1339 was injected 1 h prior to DZ administration and blood samples were collected at 4 h after the injection. Serum was separated and the T G concentration was measured using Sigma Infinity diagnostic kit. N--Cyclopentyl adenosine (CPA): CPA by inhibiting adipose tissue lipolysis has been demonstrated to lower serum FA and circulating T G [24,25]. In the indicated experiment, CPA (8 mg/kg, i.p.) was administered 1 h after DZ administration. The animals were killed after 4 hrs and cardiac HR-LPL activity determined. 73 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.2.8 Plasma measurements Plasma samples were stored at -20°C until assayed. Diagnostic kits were used to measure glucose, T G (Sigma), non esterified fatty acid (NEFA,Wako), and insulin (Linco). 3.2.9 Materials [3H]triolein was purchased from Amersham Canada. Heparin sodium injection (Hapalean; 1000 USP U/ml) was obtained from Oraganon Teknika. All other chemicals were obtained from Sigma Chemical. 3.2.10 Statistical analysis Values are means ± SE. LPL activity in response to heparin perfusion over time was analyzed by multivariate analysis (two-way ANOVA) of variance using the NCSS. Wherever appropriate, one-way A N O V A followed by the Tukey or Bonferroni tests or the unpaired and paired Student's Mest was used to determine differences between group mean values (as indicated in the specific figure legends). The level of statistical significance was set at P < 0.05. 74 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.3 Resu l ts 3.3.1 Effects of DZ on plasma parameters To evaluate the dose dependent effect on serum glucose, varying doses of DZ (25, 50, & 100 mg/kg) were administered. Although doses of 25 and 50 mg/kg increased glucose, the. extent and duration of hyperglycemia were not as substantial as that seen with 100 mg/kg (Fig. 3-1, inset), which caused a rapid decline in serum insulin within 1 h (Fig. 3-1). Notably, at this dose, serum glucose reached a maximum level after 2 h, and was comparable to the hyperglycemia produced by administration of 55 mg/kg STZ [20]. Therefore, all subsequent experiments were carried out using DZ 100 mg/kg. Changes in plasma parameters with DZ also included significant and rapid increases in FA and T G (Fig. 3-2). Other characteristics normally associated with hyperglycemia, such as polydipsia, was also observed in DZ treated animals. 3.3.2 LPL activity, immunolocalization and effects of insulin treatment Retrograde perfusion of control hearts with heparin resulted in the release of LPL into the coronary perfusate (Fig. 3-3). This heparin-mediated LPL discharge was rapid, and peak activity, thought to represent LPL located at or near the endothelial surface, was observed within 1 min. Compared to control rat hearts, there was a substantial increase in coronary LPL activity (-300%, Fig. 3-3) and immunofluroscence (Fig. 3-4) at the vascular lumen following 4 h of DZ. To determine the kinetics of LPL up regulation at the vascular lumen, some DZ treated hearts were isolated at 1-4 h, and LPL activity measured. Interestingly, increase in LPL activity became apparent as early as 1 h subsequent to injection of DZ, and was maintained for an additional 3 h (Fig. 3-5, inset). One caveat associated with DZ is that in addition to inhibiting insulin secretion, it also lowers blood pressure [19]. In some anesthetized rats, DZ produced a significant fall in mean carotid artery blood pressure (-20 mmHg). This effect was transient, and blood pressure returned to normal within 3 h (data not shown). As this hypotensive effect was not modified by a dose of 75 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 insulin (8U) that normalizes cardiac LPL (Fig. 3-5) and serum glucose (Fig. 3-5, inset), control of LPL by DZ is probably dependent on its lowering of insulin rather than its effects on blood pressure. Given that DZ is a K + A T P channel opener with potential direct effects on the heart, we examined whether DZ could directly influence cardiac heparin releasable LPL activity. Isolated hearts when perfused in the absence or presence of DZ (100 uM, lh), followed by heparin, did not significantly change peak LPL activity (control, 390; DZ, 432 nmol/ml/h). Thus, the DZ-induced increase in LPL observed in vivo could not be duplicated in vitro, and suggests that the increased enzyme observed in vivo occurs subsequent to drop in insulin. 3.3.3 Cardiac myocyte cell surface and intracellular LPL activity To determine whether the increase in luminal LPL was a consequence of augmented synthesis from cardiac cells, myocytes were isolated from control and 4h DZ treated rats. There was no difference in myocyte viability or yield following DZ. Total cellular LPL activity (surface bound + intracellular) remained unchanged between control and DZ treated myocytes (Fig. 3-6, inset). Myocytes from the two groups were also incubated with heparin. Basal LPL activity released into the medium remained unchanged between the two groups. Interestingly, myocytes from DZ treated rats demonstrated a 50% reduction in surface bound HR-LPL activity at a time when luminal LPL activity was augmented suggesting an accelerated transfer of the enzyme from the myocyte cell surface to the coronary lumen (Fig. 3-6). It should be noted that establishment of a temporal relationship between the rise in luminal with a drop in myocyte cell surface LPL activity in STZ diabetic rats was never possible given the triphasic pattern of changes in glucose and insulin profile in the 24 h period subsequent to STZ administration [26]. 76 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.3.4 LPC effects on luminal LPL To investigate the role of lipolytic byproducts on translocation of LPL from the underlying myocyte cell surface to the coronary lumen, we examined the effects of varying doses of LPC on luminal LPL activity in control rat hearts. Graded concentrations of LPC (0.1 nM-100 nM) when perfused for 60 mins increased HR-LPL activity (Fig. 3-7, inset). Interestingly, LPL activity following 1 nM LPC (Fig. 3-7) was identical to that seen subsequent to in vivo DZ administration. LPC (0.1-100 nM) by itself was unable to release luminal LPL (data not shown). Additionally, following titration with these doses of LPC, hearts did not demonstrate any change in rate and pattern of contraction. 3.3.5 Manipulation of circulating TG As a lipolytic byproduct was able to influence LPL, and given the increase in T G in DZ treated animals, we hypothesized that manipulation of circulating T G in these animals would influence luminal LPL. Treatment of control animals with WR 1339 brought about a 30 fold increase in circulating T G compared to control within 4 h, with no change in LPL activity (data not shown). Although DZ administered to WR 1339 treated animals did not affect the elevated serum T G (Fig. 3-8, left panel) or glucose (Table 3-1), there was a considerable decline in serum FA (Table 3-1) and luminal LPL activity (Fig. 3-8, right panel). Using an alternative strategy, we attempted to prevent increases in circulating T G by inhibiting adipose tissue lipolysis. In preliminary experiments, CPA demonstrated a reduction in circulating FA and T G (data not shown). Interestingly, CPA prevented the DZ induced augmentation of circulating T G (Fig. 3-8, left panel) and FA (Table 3-1) with normalization of luminal LPL activity (Fig. 3-8, right panel). 77 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.4 D i s c u s s i o n Impaired glucose utilization following hypoinsulinemia increases the requirement of the heart for FA [27,28]. This excess demand is partly achieved by up regulation of LPL at the coronary luminal surface, as described in our previous studies using the STZ rat heart [20]. Although we concluded that accelerated translocation of LPL from the myocyte cell surface to the vascular lumen could be responsible for augmenting luminal LPL activity, verification of this mechanism was technically difficult due to the triphasic pattern of blood glucose following STZ. This pattern comprises of an initial brief hyperglycemia followed by a period of hypoglycemia before noticeable hyperglycemia is attained within 12-16 hrs [26]. As metabolic switching in the heart from glucose to predominantly FA likely occurs rapidly following hypoinsulinemia, we pursued an alternate model of acute diabetes to study LPL translocation. In the present study, injection of DZ caused a rapid drop in insulin and hyperglycemia within one hour, which was sustained for four hours. Our data for the first time demonstrate a rapid DZ induced increase in coronary luminal LPL activity and protein indicative of posttranslational processing. In support of this idea, changes in luminal LPL activity following four-days of STZ diabetes was independent of shifts in mRNA levels or rates of synthesis (chapter 2). Furthermore, in adipose tissue, following nutritional changes, short-term change in LPL is not associated with changes in LPL mRNA or rate of biosynthesis [29,30]. Agents that increase intracellular cyclic adenosine monophosphate (cAMP) enhance luminal LPL [31,32]. As DZ, a phosphodiesterase inhibitor also increases cAMP [33,34], we were concerned about the direct effects of this agent on cardiac LPL. However, as perfusion of the isolated heart with DZ did not increase LPL, the increased enzyme observed in vivo likely occurred subsequent to a drop in insulin. Confirmation of the inhibitory role of in vivo insulin on cardiac LPL was supported by its ability to reverse DZ induced augmentation of luminal LPL. 78 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 In the present study, the contribution of this augmented LPL towards FA utilization is unclear, given that serum FA are also elevated subsequent to DZ. In a previous study that compared the utilization of exogenous FA [non-esterified fatty acid (NEFA)] and lipoprotein-TG, alone or in combination, using the isolated working control rat heart, a cardiac preference for NEFA was suggested [35]. More recently, studies examining the in vivo mechanism of FA uptake showed that LPL mediated T G hydrolysis was the major supplier of FA for cardiac utilization [4]. Whether the latter circumstance occurs in a setting of compromised glucose utilization by the heart is uncertain. Our data suggest that following acute hypoinsulinemia and impaired glucose utilization, manifold changes in lipoprotein-TG, serum FA and luminal LPL occur to guarantee FA supply to the heart. LPL is a distinctive protein that migrates towards the lumen from underlying parenchymal cells [36,37]. During excessive luminal requirement for LPL, an anticipated translocation of the enzyme from the myocyte cell surface to the vascular lumen would be predicted. Indeed subsequent to DZ administration, a 50% drop in myocyte surface bound HR-LPL was observed. Our results suggest that this enzyme pool is a key participant for supplementing LPL at the luminal surface following an increased cardiac demand for FA. However, as smooth muscle derived LPL can translocate to the endothelial surface of vascular tissue, the contribution of this LPL pool in explaining the effect of DZ cannot be completely disregarded [38]. Similarly, at least in human myocardium, LPL protein was detected on interstitial cells and could be equally contributive in explaining the effects of DZ [39]. Mediators responsible for the cleavage and transfer of LPL from myocyte cell surface to the lumen have not been identified. At least in adipocytes, LPC (50-100 uM) formed from lipoprotein breakdown has been shown to either directly [12] or indirectly [11], through the release of heparanase like compounds, displace surface bound LPL from adipocytes. In the current study, 100 uM LPC induced a severe cardiodepressant action in the isolated heart. 79 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 However, exposure of control hearts to an extremely low concentration of LPC (1 nM) was able to enhance luminal HR-LPL to levels observed following DZ. Whether this effect of LPC in the heart is related to its ability to release heparanases or directly cleave myocyte LPL merits further investigation. At least at the luminal surface, LPC was unable to release LPL directly. The contribution of T G lipolysis in vivo towards generation of equivalent amounts of LPC is currently unknown. Although it would be useful to measure plasma LPC following DZ, it is unlikely that if DZ increases LPC in vivo, we would be able to detect this trivial amount in the plasma. More importantly, LPC release in vivo may not be retained in the plasma due to rapid transport into the adjacent tissues. LPL possesses both T G hydrolase and phosholipase activity [40]. Thus following DZ, an increase in serum T G could augment formation of lysophophospholipids like LPC releasing endothelial heparanase like compounds that enable LPL translocation from the cardiomycoyte cell surface to the luminal surface. Given the potential importance of circulating T G breakdown in influencing cardiac substrate switching, we attempted to manipulate circulating T G lipolysis using WR 1339. This agent, by virtue of its inhibitory effect on LPL mediated T G hydrolysis brought about a 30-fold increase in serum TG. Cardiac LPL activity remained unaffected in these animals. Interestingly, although serum T G remained elevated following DZ, its ability to enhance luminal LPL was dramatically restricted. These data emphasize the importance of luminal lipolysis with its potential release of LPC, and not absolute TG, in regulating cardiac LPL. Under conditions of impaired glucose utilization, inhibiting breakdown of circulating T G would compel the heart to utilize FA. Indeed in DZ treated animals pretreated with WR 1339, a three-fold decrease in serum FA was observed. Prevention of T G accumulation by CPA mediated inhibition of adipose tissue lipolysis was equally effective in lowering DZ induced hypertriglyceredemia, and the increase in cardiac LPL. Taken together our data suggest that T G hydrolysis is a key trigger for LPL transfer to the luminal surface. 80 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 In summary, following hypoinsulinemia, there is a rapid amplification of LPL at the coronary luminal surface. As lipolytic byproducts like LPC could allow transfer of LPL from the myocyte surface to the vascular lumen, luminal T G hydrolysis could be regarded as an important regulator of "functional" LPL. Transgenic mouse lines overexpressing human LPL in skeletal and cardiac muscle demonstrated cardiomyopathy characterized by muscle fiber degeneration, extensive lipid deposition and proliferation of mitochondria and peroxisomes [41]. In a more recent study, mice that specifically overexpressed cardiomyocyte surface bound functional LPL, exhibited lipid oversupply and impaired contractile function [42]. Whether the increase in luminal LPL during diabetes is a major cause of metabolic, morphological, and mechanical changes observed in the heart requires further investigation. 81 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.5 Tab les and F igures Table 3-1 C O N T R O L D Z D Z + W R 1339 D Z + C P A Body Weight (g) 265 ± 9 278±9 256 ± 7 274 ± 8 Serum FFA (mM) 0.64 ± 0.04 1.56 ± 0 . 0 3 * 0.56 ± 0.09 0.57 ± 0.06 Serum Glucose (mM) 5.47 ± 0 . 2 7 24.9+1.2* 17.52 ± 4 . 4 6 * 25.1 ± 2 . 6 9 * Values are means ± SE for 6 animals in each group. In separate group of experiments animals administered DZ were either pre-treated with WR 1339 (1 h before) or post-treated (1 h after) with CPA. Samples were obtained after 4 h of DZ administration. * Significantly different from control, P< 0.05. 82 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 Table 3-2 Groups Agents administered with DZ Total duration of study In Vivo DZ Insulin (1 h after) 2.5 h DZ Triton WR 1339 (1 h before) 4h DZ CPA (1 h after) 4h In Vitro LPC n/a 1 h DZ (100 mg/kg) was administered i.p. As indicated various pharmacological agents like insulin (i.v. 8 units, 1 h after), WR-1339 (600 mg/kg, i.v., 1 h before) and CPA (8 mg/kg, i.p., 1 h after) were administered, the animals killed at indicated times points, and cardiac HR-LPL activity determined. To investigate the effects of LPC on cardiac LPL, isolated control hearts were perfused for 1 h with 1 nM LPC and luminal LPL activity measured. 83 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 i i I i i • 1 1 i r-0 1 2 3 44 3 2 1 0 T ime (hrs) Fig. 3-1 Serum insulin and glucose subsequent to DZ. In a preliminary experiment, animals were treated with DZ (25, 50 and 100 mg/kg, i.p.; inset), and blood samples from the tail vein collected at 60 min intervals for 4-h. Following determination of the optimal dose of DZ (100 mg/kg) necessary to induce stable hyperglycemia, six animals were treated with DZ and blood samples collected for determination of glucose and insulin. Results are the means ± SE of 6 rats in each group. One-way and two-way A N O V A (inset) for repeated measures were carried out to determine statistical differences between means. 84 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 2.0 e z z z a Serum TG (mM) Fig . 3-2 Serum T G and FA subsequent to DZ. DZ (100 mg/kg, i.p.) was administered to control animals and blood samples collected over a period of 4 h. After centrifugation, serum was separated for determination of NEFA and TG. Results are the means ± SE of 6 rats in each group. *' # Significantly different from basal, P < 0.05. One-way A N O V A was carried out to determine statistical differences between means. 85 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 heparin — I 1 1 1 1 1 1 0 2 4 6 8 10 12 T ime (min) Fig. 3-3 LPL activity following DZ. Hearts from control and DZ treated animals were isolated and perfused in the nonrecirculating retrograde mode. Thereafter, coronary functional LPL was released with heparin (5 U/ml). Coronary effluents were collected (for 10 s) at different time points over 10 mins. LPL activity was assayed using radiolabeled triolein. The inset indicates a single experiment measuring peak HR-LPL activity released after varying durations of DZ (1-4 h). Following determination of the optimal time required for DZ to augment cardiac LPL, six animals were treated with DZ and LPL activity measured at 4 h. Results are the means ± SE of 6 rats in each group. Two-way A N O V A for repeated measures was carried out to determine statistical differences between means. 86 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 Fig. 3-4 This is a representative photograph showing the effect of DZ (4 h) on LPL immunofluorescence as visualized by fluorescent microscopy. Heart sections were fixed, incubated with the polyclonal chicken antibody against bovine LPL followed by incubations with biotinylated rabbit anti-chicken IgG and streptavidin-conjugated Cy3 fluorescent probe respectively. Majority of LPL in the DZ heart was exclusively present in the coronary lumen (arrows); C, Control, DZ, Diazoxide. 87 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 - O CON C O N D Z D Z + I 8 Fig. 3-5 Effect of insulin treatment on HR-LPL activity in perfused hearts from DZ treated rats. Insulin treatment (8U, 18) was initiated 1 h after DZ, and the animals were killed after 90 min. The inset indicates serum glucose levels at various times after DZ and insulin injection. LPL activity is depicted as the integrated area under the curve (AUC) released over 10 mins in the different groups. Results are the means ± SE of 6 rats in each group. *Significantly different from all other groups, P < 0.05. One-way and two-way A N O V A (inset) for repeated measures were carried out to determine statistical differences between means. 88 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 4000 Fig. 3-6 LPL activity in cardiac myocytes from control and DZ treated rats. LPL activity in the cell homogenate (inset) was determined by removing a sample of cell suspension followed by centrifugation, and sonication of the cell pellet,. Heparin (5 U/ml) was then added to the cell suspension, incubated for 10 mins, and the release of surface bound LPL activity into the medium determined. Results are mean ± SE of three rats in each group. *Significantly different from basal (B, paired t-test), #Significantly different from Control (+HEP, group t-test), P < 0.05. For the inset an unpaired t-test was carried out. 89 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 heparin T 1 1 1 1 1 1 0 2 4 6 8 10 12 Time (min) Fig . 3-7 In vitro effect of LPC on cardiac luminal LPL activity. To investigate the concentration dependent effects of LPC on cardiac LPL, isolated control hearts were perfused for 1 h with graded concentrations of LPC (0.1-100 nM) and luminal LPL activity measured. Following determination of the concentration of LPC necessary to augment cardiac LPL, six control hearts were perfused with 1 nM LPC and LPL activity determined. Results are the means ± SE of 6 hearts in each group. Two-way A N O V A for repeated measures was carried out to determine statistical differences between means. 90 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 Fig. 3-8 Effect of WR 1339 and CPA on serum TG and HR-LPL activity. Animals administered DZ were either pre-treated with WR 1339 (1 h before) or post-treated with CPA (1 h after). Rats were killed 4 h after DZ and serum TG and LPL activity measured. The left panel describes coronary HR-LPL activity whereas the right panel illustrates serum TG in the respective groups. Results are the means ± SE of 6 rats in each group. *Significantly different from control, "Significantly different from all other groups, @Significantly different DZ, P < 0.05. Mean differences between LPL activities were evaluated using two-way ANOVA for repeated measures and one-way ANOVA was carried out for comparison of serum TG. 91 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 3.6 B ib l iography [1] Randle PJ, Hales CN, Garland PB, Newsholme EA. The glucose fatty-acid cycle; its role in insulin sensitivity and the metabolic disturbances of diabetes mellitus. Lancet 1963;1:785-89. [2] Rodrigues B, Cam MC, McNeill JH. Myocardial substrate metabolism: implications for diabetic cardiomyopathy. J Mol Cell Cardiol 1995;27:169-79. [3] Blanchette-Mackie EJ, Masuno H, Dwyer NK, Olivecrona T, Scow RO. Lipoprotein lipase in myocytes and capillary endothelium of heart: immunocytochemical study. Am J Physiol 1989;256:E818-28. [4] Augustus AS, Kako Y, Yagyu H, Goldberg IJ. Routes of FA delivery to cardiac muscle: modulation of lipoprotein lipolysis alters uptake of TG-derived FA. Am J Physiol Endocrinol Metab 2003;284:E331-9. [5] Pillarisetti S, Saxena U. Lipoprotein lipase as a therapeutic target for dyslipidemia. Front Biosci 2003;8:D238-41. [6] Preiss-Landl K, Zimmermann R, Hammerle G, Zechner R. Lipoprotein lipase: the regulation of tissue specific expression and its role in lipid and energy metabolism. Curr Opin Lipidol 2002;13:471-81. [7] Blanchette-Mackie EJ, Dwyer NK, Amende LA. Cytochemical studies of lipid metabolism: immunogold probes for lipoprotein lipase and cholesterol. Am J Anat 1989;185:255-63. [8] Stins MF, Maxfield FR, Goldberg IJ. Polarized binding of lipoprotein lipase to endothelial cells. Implications for its physiological actions. Arterioscler Thromb 1992;12:1437-46. [9] Saxena U, Klein M G , Goldberg IJ. Transport of lipoprotein lipase across endothelial cells. Proc Natl Acad Sci U S A 1991;88:2254-8. [10] Liu GQ, Olivecrona T. Pulse-chase study on lipoprotein lipase in perfused guinea pig heart. Am J Physiol 1991;261:H2044-50. 92 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 [11] Pillarisetti S, Paka L, Sasaki A, Vanni-Reyes T, Yin B, Parthasarathy N, Wagner WD, Goldberg IJ. Endothelial cell heparanase modulation of lipoprotein lipase activity. Evidence that heparan sulfate oligosaccharide is an extracellular chaperone. J Biol Chem 1997;272:15753-9. [12] Sasaki A, Goldberg IJ. Lipoprotein lipase release from BFC-1 beta adipocytes. Effects of triglyceride-rich lipoproteins and lipolysis products. J Biol Chem 1992;267:15198-204. [13] Saxena U, Witte LD, Goldberg IJ. Release of endothelial cell lipoprotein lipase by plasma lipoproteins and free fatty acids. J Biol Chem 1989;264:4349-55. [14] Morley NH, Kuksis A, Buchnea D. Hydrolysis of synthetic triacylglycerols by pancreatic and lipoprotein lipase. Lipids 1974;9:481-8. [15] Morley NH, Kuksis A, Buchnea D, Myher JJ. Hydrolysis of diacylglycerols by lipoprotein lipase. J Biol Chem 1975;250:3414-8. [16] Groot PH, Van Tol A. Metabolic fate of the phosphatidylcholine component of very low density lipoproteins during catabolism by the perfused rat heart. Biochim Biophys Acta 1978;530:188-96. [17] Foy JM, Furman BL. Effect of single dose administration of diuretics on the blood sugar of alloxan-diabetic mice or mice made hyperglycaemic by the acute administration of diazoxide. Br J Pharmacol 1973;47:124-32. [18] Kersten JR, Montgomery MW, Ghassemi T, Gross ER, Toller WG, Pagel PS, Warltier DC. Diabetes and hyperglycemia impair activation of mitochondrial K(ATP) channels. Am J Physiol Heart Circ Physiol 2001;280:H1744-50. [19] Pratz J, Mondot S, Montier F, Cavero I. Effects of the K+ channel activators, RP 52891, cromakalim and diazoxide, on the plasma insulin level, plasma renin activity and blood pressure in rats. J Pharmacol Exp Ther 1991;258:216-22. 93 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 [20] Rodrigues B, Cam M C , Jian K, Lim F, Sambandam N, Shepherd G. Differential effects of streptozotocin-induced diabetes on cardiac lipoprotein lipase activity. Diabetes 1997;46:1346-53. [21] Rodrigues B, Spooner M , Severson DL. Free fatty acids do not release lipoprotein lipase from isolated cardiac myocytes or perfused hearts. Am J Physiol 1992;262:E216-23. [22] Sheorain VS, Nageswara Rao T, Subrahmanyam D. On the inhibition of lipoprotein lipase by Triton WR 1339. Enzyme 1980;25:81-6. [23] Borensztajn J, Rone MS, Kotlar TJ. The inhibition in vivo of lipoprotein lipase (clearing-factor lipase) activity by triton WR-1339. Biochem J 1976;156:539-43. [24] Cheng JT, Chi TC, Liu IM. Activation of adenosine A l receptors by drugs to lower plasma glucose in streptozotocin-induced diabetic rats. Auton Neurosci 2000;83:127-33. [25] Wong EH, Smith JA, Jarett L. Adenosine effects on glucose oxidation of adipocytes isolated from streptozotocin-diabetic rats. Biochem J 1985;232:301-4. [26] Sambandam N, Abraham M A , St Pierre E, Al-Atar O, Cam M C , Rodrigues B. Localization of lipoprotein lipase in the diabetic heart: regulation by acute changes in insulin. Arterioscler Thromb Vase Biol 1999;19:1526-34. [27] Brownsey RW, Boone A N , Allard MF. Actions of insulin on the mammalian heart: metabolism, pathology and biochemical mechanisms. Cardiovasc Res 1997;34:3-24. [28] Gerich JE. Physiology of glucose homeostasis. Diabetes Obes Metab 2000;2:345-50. [29] Doolittle M H , Ben-Zeev O, Elovson J, Martin D, Kirchgessner TG. The response of lipoprotein lipase to feeding and fasting. Evidence for posttranslational regulation. J Biol Chem 1990;265:4570-7. [30] Wu G, Olivecrona G, Olivecrona T. The Distribution of Lipoprotein Lipase in Rat Adipose Tissue. Changes with nutritional state engage the extracellular enzyme. J Biol Chem 2003;278:11925-30. Chapter 3: Cardiovas. Res. 59 (2003) 788-797 [31] Eckel RH. Lipoprotein lipase. A multifunctional enzyme relevant to common metabolic diseases. N Engl J Med 1989;320:1060-8. [32] Ailhaud G. Cellular and secreted lipoprotein lipase revisited. Clin Biochem 1990;23:343-47. [33] Sen AK, Sunahara FA, Talesnik J. Prostaglandin E2 and cyclic AMP in the coronary vasodilatation due to cardiac hyperactivity. Can J Physiol Pharmacol 1976;54:128-39. [34] Sen A K , Sunahara FA, Talesnik J. Coronary reactions to cardiac hyperactivity and to hypoxia in isolated perfused heart of rat. Br J Pharmacol 1977;61:381-93. [35] Hauton D, Bennett MJ, Evans RD. Utilisation of triacylglycerol and non-esterified fatty acid by the working rat heart: myocardial lipid substrate preference. Biochim Biophys Acta 2001;1533:99-109. [36] Merkel M , Eckel RH, Goldberg IJ. Lipoprotein lipase: genetics, lipid uptake, and regulation. J Lipid Res 2002;43:1997-2006. [37] Mead JR, Irvine SA, Ramji DP. Lipoprotein lipase: structure, function, regulation, and role in disease. J Mol Med 2002;80:753-69. [38] Esenabhalu V E , Cerimagic M , Malli R, Osibow K, Levak-Frank S, Frieden M , Sattler W, Kostner G M , Zechner R, Graier WF. Tissue-specific expression of human lipoprotein lipase in the vascular system affects vascular reactivity in transgenic mice. Br J Pharmacol 2002;135:143-54. [39] O'Brien KD, Ferguson M , Gordon D, Deeb SS, Chait A. Lipoprotein lipase is produced by cardiac myocytes rather than interstitial cells in human myocardium. Arterioscler Thromb 1994;14:1445-51. [40] Bengtsson-Olivecrona G, Olivecrona T. Phospholipase activity of milk lipoprotein lipase. Methods Enzymol 1991;197:345-56. [41] Levak-Frank S, Radner H, Walsh A, Stollberger R, Knipping G, Hoefler G, Sattler W, Weinstock PH, Breslow JL, Zechner R. Muscle-specific overexpression of lipoprotein 95 Chapter 3: Cardiovas. Res. 59 (2003) 788-797 lipase causes a severe myopathy characterized by proliferation of mitochondria and peroxisomes in transgenic mice. J Clin Invest 1995;96:976-86. [42] Yagyu H, Chen G, Yokoyama M , Hirata K, Augustus A, Kako Y, Seo T, Hu Y, Lutz EP, Merkel M , Bensadoun A, Homma S, Goldberg IJ. Lipoprotein lipase (LpL) on the surface of cardiomyocytes increases lipid uptake and produces a cardiomyopathy. J Clin Invest 2003;111:419-26. 96 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.0 Palmitoyl lysophosphatidylcholine mediated mobilization of LPL to the coronary luminal surface requires PKC activation 4.1 Introduction Heart tissue acquires most of its energy from metabolism of two major substrates, glucose and FA, the latter being the preferred substrate consumed [1,2]. Hydrolysis of TG-rich lipoproteins by LPL positioned at the endothelial surface of the coronary lumen has been suggested to be the principal source of FA for cardiac utilization [3, 4]. As endothelial cells do not synthesize LPL, the enzyme is primarily synthesized in the cardiomyocytes [5, 6]. LPL secreted as an active dimer binds to myocyte cell surface HSPG, before it is translocated onto comparable HSPG binding sites on the luminal side of the vessel wall [7-9]. At this location, HR-LPL, exhibits T G hydrolase activity [10]. Impaired cardiac glucose transport and utilization switches energy production to exclusive [3-oxidation of FA [11, 12]. Under these circumstances LPL is expected to increase to supply additional FA. Indeed, following hypoinsulinemia induced by DZ or STZ, there is augmentation of LPL at the coronary luminal surface [13, 14]. We established that this increased LPL in the hyperglycemic heart is likely an outcome of empty luminal HSPG binding sites being occupied by enzyme that is recruited from the cardiac myocyte (the cell type within the heart that contains maximum LPL) [14]. Mediators responsible for LPL transfer from the underlying parenchymal cells to the vascular lumen have not been extensively characterized. At least in co-culture experiments, LPC has been suggested to be one such mediator, that by secreting heparanase like compounds from endothelial cells, cleaves and transports adipocyte surface bound LPL to the endothelial apical surface [15, 16]. 97 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 LPC, a major constituent of cellular phospholipids, is also present within T G rich lipoproteins [17]. Following phospholipase A 2 mediated phospholipid hydrolysis or LPL mediated lipolysis of T G rich lipoproteins, LPC is generated [17, 18]. We demonstrated that in isolated control rat hearts, LPC augments luminal LPL [13]. At least in endothelial cells, LPC is reported to activate protein kinase C (PKC) [19]. Additionally, activation of PKC by leptin is known to augment macrophage LPL [20]. In the present study, we hypothesized that the LPC induced increase in luminal LPL requires PKC activation. Our data for the first time demonstrates a cardiospecific regulation of LPL by lysophospholipids, a mechanism that is PKC dependent, and likely requires metabolic degradation of LPC. 98 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.2 Materials and Methods 4.2.1 Experimental animals Animals were cared for in accordance with the principles promulgated by the Canadian Council on Animal Care and the University of British Columbia. Adult male Wistar rats (270-290 g) were maintained under a 12-h light (0700-1900)/dark cycle and supplied with a standard laboratory diet and water ad libitum. 4.2.2 Isolated heart perfusion Rats were anesthetized with 65 mg/kg sodium pentobarbital i.p., the thoracic cavity opened, and hearts isolated. The heart was immersed in cold (4°C) Krebs-Henseleit HEPES buffer containing 10 mM glucose (pH 7.4). After aortic cannulation, hearts were perfused retrogradely by the non-recirculating Langendorff technique [13, 14]. To measure vascular endothelium bound LPL, perfusion solution was changed to buffer containing 1% fatty acid-free BSA and heparin (5 U/mL) to release cardiac LPL from its HSPG binding sites. Coronary effluent was collected at different time points over 10 min and assayed for LPL activity. 4.2.3 Isolated cardiac myocytes In addition to luminal LPL, considerable amount of LPL is also located on the surface and within myocytes. Ventricular calcium-tolerant myocytes were prepared by a previously described procedure [13, 14]. Cardiac myocytes were suspended at a final cell density of 0.4 x 106 cells/mL, incubated at 37°C and basal LPL activity in the medium measured. To release surface-bound LPL activity, heparin (5 U/mL) was added to the myocyte suspension and aliquots of cell suspension were removed at different time points, medium separated by centrifugation in an Eppendorf microcentrifuge, and assayed for LPL activity. 99 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.2.4 Measurement of LPL activity LPL catalytic activity in coronary perfusates and myocyte medium was determined by measuring the in vitro hydrolysis of a sonicated [3H] triolein substrate emulsion [21]. 100 pL of either myocyte medium or coronary perfusate was used to measure LPL activity. Results are expressed as nanomoles of oleate released per hour per milliliter (coronary perfusate) or per 106 cells (myocyte medium). 4.2.5 Immunoblot analysis for PKC To determine PKC activation, migration of PKC from cytosol to the membrane was determined using immunoblot analysis [22-24]. Briefly, ventricles or myocytes were homogenized in ice-cold buffer A (containing 20 mM Tris-HCl, 2 mM EDTA, 5 mM EGTA, 50 mM 2-mercaptoethanol DTT, 25 pg leupeptin, and 4 pg aprotinin, pH 7.5) and centrifuged for 1 h at 35,000 rpm; the supernatant was used as the cytosolic fraction. The pellet was resuspended in buffer B (containing 1 % NP-40, 0.1% SDS, 0.5% deoxycholic acid and 5 mM EGTA, pH 7.5), sonicated for 30 sec, and centrifuged at 35,000 rpm for 1 h; the supernatant was used as the membrane fraction. Protein content of both cytosolic and membrane fractions were determined using a Bradford protein assay kit (Bio-Rad). The protein (50 ug) for both cytosol and membrane were fractionated by 11 % SDS-PAGE and transferred onto nitrocellulose membrane. The membranes were blocked with Tris buffered saline containing 5% skim milk for 2 h at room temperature. To identify PKC protein, either polyclonal rabbit anti PKC s, PKC a or PKC Pi/o primary antibody was used at a dilution of 1:500 followed by incubation with goat anti-rabbit horseradish peroxidase conjugated secondary antibody at a dilution of 1:5000. Membranes were washed and the reaction products visualized using chemiluminescence. Blots were quantified by densitometry. PKC activation was also assessed in human umbilical vein endothelial cells 100 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 (HUVECs) by a previously described procedure [25]. HUVECs (~ 2 million) were exposed to the LPC treatment for 20 min and immunoblotting for PKC s was performed. 4.2.6 Immunolocalization of PKC s Of the many different PKC isoforms, PKC e, an abundant PKC isoform in the heart was selected to validate PKC translocation using immunolocalization. PKC £ immunolocalization was assessed in the myocardial sections by a previously described procedure [26]. Briefly, upon excision, control and LPC treated hearts were retrogradely perfused with non-circulating buffer for 3 mins to clear the heart of blood. Perfusion buffer was then changed to fixative (neutral phosphate-buffered 10% formalin) for 2 mins. Hearts were stored in 10% formalin for 24 hrs. Blocks were embedded in paraplast, sectioned at 3 urn and mounted on positively charged glass slides. For immunostaining, sections were deparaffinized, rehydrated, and treated with 5% (vol/vol) heat inactivated goat serum in Tris-buffered saline (TBS) to block nonspecific background. Sections were incubated with affinity-purified rabbit polyclonal primary antibody against phosphorylated (Ser-729) PKC 8 (1:200 dilution in TBS containing 3% (w/v) BSA and 3% goat serum) overnight at room temperature in a humid chamber. Samples were then washed with TBS and incubated for 1 hr at room temperature with the secondary antibody Alexa Fluor * 488 F(ab')2 fragment of goat anti rabbit IgG (Molecular Probes, 1:5000 dilution). The unbound fluorescent probe was rinsed with TBS buffer and sections mounted with DABCO. Slides were visualized using a Biorad 600 Confocal Microscope at 63 Ox magnification. 4.2.7 Treatments Palmitoyl Lysophosphatidylcholine (LPC): LPC (1 nM) when perfused for 60 min through isolated control hearts, augmented HR-LPL activity. To assess the contribution of the type of FA conjugated to LPC that facilitates increase in luminal LPL, hearts were also perfused with oleoyl (OL) or stearoyl (ST) L-cx-LPC, (1 and 100 nM) for 60 min, and LPL activity measured. 101 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 All of the above agents were freely soluble in Krebs-Henseleit HEPES buffer. To assess whether LPC has a similar effect in cardiomyocytes, control myocytes were incubated with LPC for 60 min and LPL activity measured. Diazoxide (DZ): DZ, a selective K +ATP channel opener decreases insulin secretion and causes hyperglycemia [13, 27, 28]. DZ (100 mg/kg) was administered i.p., the animals euthanized after 4 h, and hearts removed for determination of HR-LPL activity (0 min). To determine whether this high LPL activity can be maintained in vitro, some DZ hearts were perfused only with Krebs buffer and heparin releasable LPL activities determined at 15, 30, 45 and 60 min respectively. In a different experiment, hearts from DZ treated animals were perfused for 1 h, either in the absence or presence of LPC (1 nM), glucose (25 mM), T G (1.4 mM) or palmitic acid (1.5 mM):BSA, and LPL activity then determined. Lysophosphatidic acid (LPA): LPA, a pluripotent lipid mediator [29, 30], is generated from hydrolysis of LPC in the outer leaflet of the cell membrane, a pathway requiring phospholipase D (PLD) or lysophospholipase D activation [31-33]. To investigate the influence of LPA on LPL, isolated heart and myocytes were treated with 1 nM LPA for 1 h and LPL activity determined. LPA was solubilized in chloroform:methanol (1:2), and subsequently in Krebs-Henseleit HEPES buffer. Phospholipase D (PLD): Control myocytes were incubated simultaneously with PLD, (from Streptomyces chromofuscus, 5 U/mL) and LPC (1 nM) for 60 min and LPL activity measured. PKC Inhibition: Isolated control hearts were perfused with calphostin (100 nM, a non-selective inhibitor of PKC) [34-37] and LPC (1 nM) for 60 min, and HR-LPL activity subsequently determined. An alternate PKC inhibitor, bisindolylmaleimide HC1 (1 pM), an agent that exhibits relatively selective inhibition of conventional PKCs and PKCs [38, 39] was also tested in a group of control hearts perfused with LPC, and HR-LPL activity determined. 102 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.2.8 Materials [ H]-triolein was purchased from Amersham Canada. Heparin sodium injection (Hapalean; 1000 USP U/mL) was obtained from Oraganon Teknika. All other chemicals were obtained from Sigma Chemical. 4.2.9 Statistical analysis Values are means ± SE. LPL activity in response to heparin perfusion over time was analyzed by multivariate analysis (two-way ANOVA) of variance using the NCSS. Wherever appropriate, one-way A N O V A followed by the Tukey or Bonferroni tests or the unpaired and paired Student's /-test was used to determine differences between group mean values (as indicated in the specific figure legends). The level of statistical significance was set at P < 0.05. 103 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.3 Resu l ts 4.3.1 LPC maintains high luminal LPL in DZ treated heart perfused in vitro Injection of DZ caused a rapid increase in serum glucose within 1 h, which was sustained for 4 h (CON: 5.2±0.2, DZ: 23±2.3, P< 0.05). We have previously demonstrated that following DZ, there was a rapid and substantial increase in coronary HR-LPL activity at the vascular lumen [13], and this observation was reproduced in the present study (Fig. 4-1, 0 min). To determine whether this high luminal LPL activity can be maintained in vitro, hearts from DZ animals were perfused with Krebs buffer and at the indicated time points, heparin was added to determine the enzyme remaining at this location. Interestingly, there was a time dependent decline of luminal LPL activity over 1 h to control values, presumably through dissociation of the enzyme from endothelial HSPG binding sites into the perfusate (Fig. 4-1). To further investigate this process, in a separate experiment, basal LPL activity (in the absence of heparin) was determined in the buffer reservoir over time (Fig. 4-1, inset). Interestingly, DZ hearts showed enhanced release of basal LPL only at the early time points. In an effort to identify potential circulating mediators responsible for maintaining high LPL in vivo, some DZ treated hearts were perfused for 1 h, either in the absence or presence of glucose (25 mM), T G (1.4 mM), palmitic acid (1.5 mM) or LPC (1 nM). LPC was able to maintain the high LPL activity in DZ treated hearts (Fig. 4-1), an effect that was not observed following glucose, T G or FA perfusion (data not shown). To verify if FA other than palmitic acid conjugated to LPC can also increase luminal LPL, isolated hearts were perfused with 1 nM oleoyl and stearoyl LPC. We were unable to detect any increase in luminal LPL with these (Fig. 4-2) or higher concentrations (data not shown). 104 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.3.2 LPC mediated increases in luminal LPL requires whole heart PKC activation LPC has been reported to activate PKC in coronary endothelial and renal cells and platelets [18, 25, 40, 41]. Furthermore, activation of PKC in different cell types is known to regulate LPL [20, 42, 43]. To assess whether PLPC can also activate cardiac PKC, Western blot analysis for PKC s (an abundant PKC isoform) [22], PKC a and PKC Pi/n was carried out. In isolated LPC perfused hearts, PKC s activation peaked at 20 min (Fig. 4-3). LPC perfusion also caused membrane translocation of PKC a and PKC Pi/n (data not shown). To validate translocation, PKC s immunolocalization was carried out. Immunofluorescence microscopy of myocardial sections was able to confirm a more intense PKC £ immunofluorescence in the LPC treated myocardial blood vessels compared with control (Fig. 4-4). Given the role of PKC in regulating LPL, we hypothesized that PKC blockade would diminish the effects of LPC on luminal LPL. When perfused for 60 min, calphostin (CAL) per se did not influence luminal LPL (data not shown). Interestingly, following pretreatment with C A L for 20 min and subsequent 1 h perfusion with LPC, PKC activation (Fig. 4-5, inset), immunofluorescence (Fig. 4-4) and LPL activity (Fig. 4-5) declined significantly. Bisindolylmaleimide HC1, a more selective PKC inhibitor, also demonstrated similar effects on LPL activity (Con: 424 ± 47, LPC: 1018 ± 67, LPC+BIS-HC1: 562 ± 117 nmol/hr/mL, P< 0.05). 4.3.3 Differential effect of LPC on whole heart and cardiomyocyte LPL In the heart, as 78% of total LPL is present in cardiac myocytes, which subsequently transfers onto luminal HSPG binding sites, we evaluated if LPC can augment LPL in mycoytes [3, 8]. There were no changes in the basal and HR-LPL activity from control and LPC treated myocytes (Fig. 4-6). Interestingly, unlike the whole heart, LPC was also unable to activate myocyte PKC (Fig. 4-6, inset). To determine if other cells could contribute towards the activation of PKC, endothelial cells were cultured and subjected to LPC. Interestingly, incubation for 20 min 105 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 activated PKC in HUVECs [as measured by densitometry; CON: Cytosol:1.6, Membrane:2.6; LPC:Cytosol:1.4, Membrane:4.4; units] (Fig. 4-7) suggesting that LPC induced PKC activation in whole hearts likely occurs predominantly in endothelial cells. 4.3.4 LPC mediated increases in luminal LPL requires formation of LP A Activation of PKC is known to stimulate PLD [44], which facilitates the conversion of LPC to LPA [31-33] . Additionally, a recent report also documented sphingomyelinase D mediated degradation of LPC to LPA with consequent activation of LPA receptors [45]. Even though LPC was unable to affect myocyte LPL, the contribution of LPA towards affecting this enzyme was unclear. Interestingly, incubation of myocytes with 1 nM LPA enhanced basal and HR-LPL activity appreciably (Fig. 4-8) without influencing intracellular LPL (Con: 3578 ± 474, LPA: 3264 ± 483 nmol/hr/106 cells). This effect of LPA on myocyte LPL was independent of PKC activation [as measured by densitometry; CON: Cytosol:16.5, Membrane:5.9; LPA: Cytosol:11.7, Membrane:2.8; units] (Fig. 4-8, inset). To establish that it is LPC derived LPA that enhances LPL, we incubated control myocytes with exogenous PLD and LPC and observed a significant enhancement of basal and HR-LPL activity (Fig. 4-8) suggesting that under such circumstances, LPC may be converted to LPA. 106 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 4.4 Discussion Following hypoinsulinemia induced with DZ, HR-LPL activity and protein increased at the coronary lumen [13]. Given that this increase in luminal enzyme occurred rapidly (within 1 h) and that endothelial cells do not synthesize LPL, we proposed that this process likely involved enzyme transfer from the cardiomyocyte to the vascular lumen. Additionally, as plasma glucose, T G and lipolytic byproducts like FA increase following DZ [13], we hypothesized that these circulating factors could facilitate LPL recruitment to the lumen. In the present study, the observation that DZ treated hearts perfused with Krebs buffer for 1 h was unable to maintain high LPL activity suggests that plasma mediators are indeed mandatory for LPL augmentation. Moreover, in their absence, a gradual decline in luminal LPL was presumably due to an imbalance between enzyme dissociation from the endothelial HSPG binding sites and recruitment from the cardiomyocyte. Indeed, DZ hearts released more basal LPL compared to control at early time points supporting the above paradigm. As only LPC was able to sustain the high LPL activity in vitro, our data confirms the importance of LPC as a key metabolic trigger that facilitates enzyme transfer. Interestingly, unlike palmitoyl, the stearoyl and oleoyl species of LPC were unable to influence luminal LPL. At present, the mechanism governing this differential effect of L P C s on luminal LPL is unclear. At least in HL-60 leukemia cells, LPC mediated stimulation or inhibition of phospholipase C/Ca 2 + system also depended on the species of fatty acid residue conjugated to LPC, a phenomenon that was attributed to differences in degradation or cellular permeability [46]. LPC is known to activate PKC in platelets and renal and endothelial cells [18, 24, 40, 41]. In turn, PKC has been implicated in the regulation of LPL in adipocytes and macrophages [20, 42, 43]. In the present study, we tested if the in vitro effects of LPC on cardiac LPL occur through activation of PKC. Perfusion of isolated hearts with LPC caused membrane translocation of 107 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 PKC a, PKC pi/n and PKC e, an effect that was transient, and peaked at 20 min. Only one other study has reported LPC mediated PKC s activation in rat renal mesangial cells [47]. As pretreatment of the isolated hearts with calphostin or bisindolylmaleimide HC1 blocked PKC activation and increases in luminal LPL activity by LPC, our data suggest that whole heart PKC activation may be necessary for LPC mediated increases in LPL. Myocardial distribution of LPL protein in mouse heart demonstrates that 78% of total LPL is present in myocytes, 3-6% in the interstitial space, and 18% in the capillary endothelium [3, 8]. Since leptin augmented LPL mRNA expression in macrophages, within 1 h and in a PKC dependent manner [20], we examined whether LPC mediated augmentation of luminal LPL involves increased enzyme turnover in cardiomyocytes. Unlike its effects on the whole heart, incubation of myocytes for 1 h with LPC had no effect on either surface or intracellular LPL activity suggesting that LPL synthesis likely does not govern increases in luminal LPL. Interestingly, STZ diabetic rat hearts also demonstrate increases in coronary luminal LPL activity and protein in the absence of a shift in myocyte LPL mRNA [14]. Given the differential effects of LPC on whole heart and myocyte LPL, we hypothesized that LPC would likely be unable to activate PKC in myocytes. Indeed, unlike the whole heart, LPC was incapable of activating myocyte PKC suggesting that in the whole heart, PKC activation occurs in cells other than the myocyte or that metabolism of LPC is required for its downstream effects. Exposure of HUVECs to LPC for 20 min activated PKC s, suggesting that in the whole heart, PKC activation likely occurs in endothelial cells. To verify if metabolism of LPC is required for influencing luminal LPL, myocytes were incubated with LPA, a known breakdown product of LPC. Interestingly, incubation of myocytes with LPA significantly enhanced surface bound basal and HR-LPL activity, without influencing the intracellular pool and was independent of changes in PKC. In endothelial and vascular smooth muscle cells, activation of 108 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 PKC stimulates secretory phospholipases [48], plasma lysophospholipases [49], and phospholipase D [44, 50] that in turn promotes formation of LPA from LPC [31-33]. We thus hypothesized that at least in the whole heart and not the myocyte, it is the coronary endothelium dependent conversion of LPC to LPA that augments myocyte LPL. In an attempt to convert LPC to LPA in vitro, myocytes were co-incubated PLD and LPC and LPL measured. Our results demonstrate an increase in myocyte surface bound and not intracellular LPL, an effect similar to that observed with LPA. These data for the first time suggest that LPC, likely through conversion to LPA, is able to increase LPL at the myocyte cell surface, that is eventually recruited to the coronary lumen. Substantiation of this mechanism would require measurement of trivial concentrations of lysophospholipids within the heart. In summary, LPC causes augmentation of luminal LPL, a process that likely requires enzyme mobilization from cardiomyocytes. The differential effects of LPC on the whole heart and myocytes suggest that PKC activation and metabolism of LPC to LPA in the endothelial cells is required for this LPL augmenting property (Fig. 4-9). As lipoproteins isolated from diabetic animals and patients have a higher LPC content [51-53], it is possible that LPC by increasing luminal LPL causes lipid oversupply with resulting muscle fiber degeneration, extensive lipid deposition, proliferation of mitochondria and peroxisomes, and eventual impairment in contractile function [54-57]. At present the mechanisms by which LPA facilitates LPL release from cardiomyocyte is unclear, but could involve actin cytoskeleton reorganization. Interestingly, incubation of cultured cardiomyocytes with insulin and dexamethasone stimulated basal and HR-LPL activities, a process involving actin cytoskeleton reorganization [58], and alterations of the actin cytoskeleton during neurite retraction is one of the best-characterized effects of LPA [59]. 109 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 300 4.5 Figures 1400 -t 1200 4 O E c < D 1000 H C D C O t C D Q_ > o < C D C D 800 4 600 4 400 4 200 4 0 D Z CON 0 < 100 H 10 20 30 Perfusion Time (min) Minutes Fig. 4-1 Control (CON) and diazoxide (DZ) treated hearts were perfused in the non-recirculating retrograde mode. Following cannulation at varying time points (0-60 min), HR-LPL activity determined. Following DZ treatment and cannulation, one group of hearts were perfused with LPC (1 nM) for 60 min (DZ60) in the non-recirculating retrograde mode, HR-LPL activity subsequently determined. The inset represents basal enzyme activity in control and DZ hearts perfused with Krebs-Henseleit in the recirculating mode. Results are expressed as the means ± SE of n=6 animals, except for the 15, 30 and 45 min time points (n=l). *Significantly different from all other groups, P < 0.05. 110 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 c 5000 -1 1 o • o O 4000 \ CON P ST OL LPC F i g . 4-2 Control hearts were isolated and perfused in the recirculating retrograde mode in the presence of palmitoyl (P), stearoyl (ST) and oleoyl (OL) L-a-lysophosphatidylcholine, (1 nM) for 60 min. Thereafter, coronary LPL was released with heparin (5 U/mL), and activity measured using radiolabelled [3H]-triolein. LPL activity is depicted as the integrated area under the curve (AUC) for LPL released over 10 mins in the different groups. Results are the means ± SE of 5 rats in each group. *Significantly different from all other groups, P < 0.05. 111 Chapter 4: J. Mol. Cell. Cardiol 37 (2004) 931-938 C M C M C M o w c o .2 % ~ o S o •4—> ~*—' r; o 200 n 160 H 120 H 80 H 40 H CON Membrane Cytosol CON 10 20 LPC 1 nM 10 2 0 L P C (min) (1 nM) Fig. 4-3 Isolated hearts were perfused with LPC and after 10 and 20 min, cardiac tissue homogenates were prepared. Homogenates were subjected to cytosolic and membrane separation, and respective protein contents determined using a Bradford protein assay kit. Identification of PKC protein was carried out using polyclonal rabbit PKC £ as the primary and goat anti-rabbit horseradish peroxidase as the secondary antibody. Densitometry values are expressed as mean ± SE of 4 rats in each group. Significantly different from control, P<0.05. The inset depicts a single representative Western blot of cardiac PKC £ in cytosolic (C) and membrane (M) fractions. 112 Chapter 4: J. Mol Cell Cardiol 37 (2004) 931-938 C O N + LPC L P C + C A L Fig. 4-4 Representative photograph showing the effect of LPC (20 min) on PKC e immunofluorescence as visualized by fluorescent microscopy. Heart sections were fixed, incubated with affinity-purified rabbit polyclonal primary antibody against phosphorylated (Ser-729) PKC 8 followed by incubations with secondary antibody Alexa Fluor 488 F(ab')2 fragment of goat anti rabbit IgG fluorescent probe respectively. Majority of PKC 8 in the LPC treated heart was exclusively present along the myocardial endothelial lining (white arrows), an effect that was inhibited by calphostin; CON-Control, LPC- Palmitoyl lysophosphatidylcholine, C A L - Calphostin. 113 Chapter 4: J. Mol. Cell. Cardiol. 37(2004) 931-938 C M C M C M CON C A L L P C 1 n M 1200 i | 1000 o E c 800 CD •4—' | 600 t CD c 400 1 200 < 0 -#• LPC - O Cal+LPC 0 4 6 Time (min) 8 i 10 12 Fig. 4-5 Isolated hearts were co-perfused with calphostin (CAL, 100 nM) and LPC for 1 h, HR-LPL activity determined, and hearts frozen for subsequent PKC analysis. Results are means ± SE of 6 hearts in each group. The inset depicts a single representative Western blot of cardiac PKC e in cytosolic (C) and membrane (M) fractions. 114 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 C M C M B + H E P B + H E P Contro l L P C Fig. 4-6 Myocytes (0.4 x 106 cells/ml) were incubated either in the absence or presence of LPC (1 nM) for 1 h. Heparin (5 U/mL) was then added to the cell suspension, and after another 60 min, a 1 ml aliquot was aspirated, centrifuged and the supernatant separated. The released surface-bound LPL activity into the medium was determined. Results are means ± SE of three rats in each group. * Significantly different from respective basal. The inset represents a single representative Western blot of PKC s of myocytes isolated from the entire ventricles. 115 Chapter 4: J. Mol. Cell. Cardiol. 37(2004) 931-938 CON LPC Fig. 4-7 This graph depicts a single representative Western blot of PKC 8 of HUVECs (~ 2 million cells) incubated in the presence or absence of LPC(1 nM)for20 min. 116 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 C M C M §09 *• 1 IVfll CON LPA (1 nM) Fig. 4-8 Myocytes (0.4 x 106 cells/ml) were incubated either in the absence or presence of LPA (1 nM) for 1 h. Heparin was then added to the cell suspension, and a 1 ml aliquot was aspirated, centrifuged and the supernatant separated at the indicated time points. The released surface-bound LPL activity into the medium was determined. In a different group, myocytes were also incubated simultaneously with PLD and LPC for 1 h and HR-LPL activity subsequently determined. Results are means ± SE of three rats in each group. The inset represents a single representative Western blot of PKC 8 of myocytes isolated from the entire ventricles that were exposed to 1 nM LPA. 117 Chapter 4: J. Mol. Cell. Cardiol. 37 (2004) 931-938 LPC 0*K(^ Endothelial Cell f \ \ I A -* T / 1 I I PLD ^ I T „ . CAL/BIS LPA i Receptor I ? WI -|- Secretion and Translocation Actin Cytoskeleton^ Reassembly Cardiomyocyte Fig. 4-9 Scheme of probable mechanism for LPC induced LPL transcytosis. Activation of PKC by LPC exclusively in endothelial cells likely facilitates PLD activation with subsequent formation of LPA. LPA, possibly via its effect on actin cytoskeleton organization, promotes LPL translocation from the cardiomyocyte to the endothelial lumen where lipoprotein hydrolysis is facilitated. 118 Chapter 4: J. Mol. Cell. 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Physiol. 288 (2005) H2802-H2810 5.0 Lysophosphatidic acid mediated augmentation of cardiomyocyte lipoprotein lipase involves actin cytoskeleton reorganization 5.1 Introduction Heart disease is a leading cause of mortality in diabetic patients, with coronary vessel disease being the primary reason for the increased incidence of cardiovascular dysfunction [1,2]. More recent evidence suggests that heart failure during diabetes can also occur secondary to altered cardiac energy metabolism where impaired glucose transport and utilization switches ATP production exclusively to oxidation of FA [3]. This adaptive mechanism could eventually become counterproductive leading to "lipotoxicity" where FA accumulate and can, either by themselves or via production of second messengers such as ceramides, provoke cell death [4]. Heart tissue acquires energy from metabolism of two major substrates, glucose and FA, the latter being the preferred substrate consumed [3, 5]. FA delivery to the heart involves: i) release from adipose tissue and transport to the heart after complexing with albumin [6], ii) provision through the breakdown of endogenous cardiac T G stores [7], iii) internalization of whole lipoproteins [8], and iv) hydrolysis of circulating TG-rich lipoproteins to FA by coronary lumen LPL [9]. The molar concentration of FA bound to albumin is -10 fold less than that of FA in lipoprotein-TG [10], and recently, LPL-mediated hydrolysis of lipoproteins was suggested to be the principal source of F A for cardiac utilization [11]. As coronary endothelial cells do not synthesize LPL, this enzyme is primarily synthesized in cardiomyocytes [12]. LPL, secreted as an active dimmer, binds to myocyte cell surface HSPG before it is translocated onto comparable HSPG binding sites on the luminal side of the vessel wall [13]. At this location, LPL is HR-LPL, exhibiting T G hydrolase activity [12-14]. Given the dependence of the diabetic heart on FA, luminal LPL is expected to increase in order to supply additional FA to meet the metabolic demand [15, 16]. Indeed, following hypoinsulinemia 126 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 induced by DZ [17] or STZ [18], we reported augmentation of LPL at the coronary luminal surface. Additionally, we established that this increased LPL in the hyperglycemic heart is likely an outcome of empty luminal HSPG binding sites being occupied by enzyme that is recruited from the cardiac myocyte [18]. Mediators responsible for LPL transfer from the underlying parenchymal cells to the vascular lumen have not been extensively characterized. In co-culture experiments, LPC has been suggested to be one such mediator, that by secreting heparanase-like compounds from endothelial cells, cleaves and transports adipocyte surface LPL to the endothelial apical surface [19]. We have demonstrated that exposure of isolated control hearts to LPC also enhances luminal LPL [17, 20]. This LPL augmenting property of LPC likely required obligatory formation of LPA, leading to mobilization of enzyme from the myocyte to the coronary lumen [20]. Indeed, when cardiomyocytes were incubated with LPA, myocyte surface LPL activity increased [20]. LPA is a pluripotent lipid mediator with multiple biological actions, including promotion of cell proliferation and migration [21, 22]. LPA binds to specific cell surface G-protein-coupled receptors, initiating a variety of signal transduction pathways involving RhoA/Rho kinase (ROCK), tyrosine kinase and mitogen activated protein kinases [21, 23, 24]. The precise mechanism by which LPA induces LPL secretion from the cardiomyocyte is not clearly understood. Given that the actin cytoskeleton has been implicated in regulating cardiomyocyte LPL [25], and that actin cytoskeleton rearrangement during neurite retraction is one of the best-characterized effects of LPA [26], we hypothesized that the increase in LPL secretion following LPA involves actin cytoskeleton reassembly. We demonstrate that following exposure to LPA, the increase in cardiomyocyte LPL activity is likely secondary to activation of RhoA and its downstream effector ROCK, which in turn modulates actin cytoskeleton reorganization. 127 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.2 Materials a n d Methods 5.2.1 Experimental animals The investigation conforms to the guide for the care and use of laboratory animals published by the US National Institutes of Health and the University of British Columbia. Adult male Wistar rats (220-240 g) were obtained from the UBC Animal Care Unit and supplied with a standard laboratory diet (PMI Feeds, Richmond, VA), and water ad libitum. 5.2.2 Isolated heart perfusion Hearts were isolated and perfused as described previously [17, 18, 20]. Briefly, rats were anesthetized with 65 mg/kg sodium pentobarbital, and the hearts carefully excised. Following cannulation of the aorta, hearts were secured by tying below the innominate artery, and perfused retrogradely with Krebs-Henseleit-HEPES buffer containing 10 mM glucose (pH 7.4). Perfusion fluid was continuously gassed with 95% 02/5% C 0 2 . The rate of coronary flow (7-8 ml/min) was controlled by a peristaltic pump. 5.2.3 Isolated cardiac myocytes In addition to luminal LPL, considerable amount of enzyme is also located on the surface of and within myocytes. Ventricular calcium-tolerant myocytes were prepared by a previously described procedure [17, 18, 20]. Cardiac myocytes were suspended at a final cell density of 0.4 x 106 cells/mL, incubated at 37°C and basal LPL activity in the medium measured. To release surface-bound LPL activity, heparin (5 U/mL) was added to the myocyte suspension and aliquots of cell suspension were removed at different time points, medium separated by centrifugation, and assayed for LPL activity. 128 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.2.4 Measurement of LPL activity LPL catalytic activity in coronary perfusates and myocyte medium was determined by measuring the in vitro hydrolysis of a sonicated [3H] triolein substrate emulsion [27]. 100 pL of either myocyte medium or coronary perfusate was used to measure LPL activity. Results are expressed as nanomoles of oleate released per hour per milliliter (coronary perfusate) or per 106 cells (myocyte medium). 5.2.5 Western blotting for Rho A and Rho kinase (ROCK) I & II To determine RhoA and ROCK activation, migration of these proteins from cytosol to the membrane was determined using immunoblot analysis [28, 29]. Briefly, ventricles (200 mg) or myocytes (2 x 106) that were subjected to various agents were homogenized in ice-cold buffer A (containing 20 mM Tris-HCl, 2 mM EDTA, 5 mM EGTA, 50 mM 2-mercaptoethanol, DTT, 25 pg leupeptin, and 4 pg aprotinin, pH 7.5), and centrifuged for lh at 35,000 rpm; the supernatant was used as the cytosolic fraction. The pellet was resuspended in buffer B (containing 1 % NP-40, 0.1% SDS, 0.5% deoxycholic acid and 5 mM EGTA, pH 7.5), sonicated for 30 sec, and centrifuged at 35,000 rpm for 1 h; the supernatant was used as the membrane fraction. Protein content of both cytosolic and membrane fractions were determined using a Bradford protein assay kit (Bio-Rad). Cytosolic and membrane proteins (50 pg) were fractionated using 11 % SDS-PAGE, and transferred onto nitrocellulose membranes (PVDF membranes were used when estimating RhoA). The membranes were blocked with Tris buffered saline containing 5% skim milk for 2 h at room temperature. To identify RhoA and ROCK I & II protein, mouse monoclonal anti RhoA, or rabbit polyclonal anti ROCK I & II primary antibodies were used at dilutions of 1:250 and 1:180 respectively. This was followed by incubation with goat anti-mouse (RhoA) or goat anti-rabbit (ROCK) horseradish peroxidase conjugated secondary antibodies at dilutions of 1:3000 and 1:5000 respectively. Membranes were washed and the reaction products visualized using chemiluminescence. Blots were quantified by densitometry. 129 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.2.6 Cardiac LPL gene expression LPL gene expression was measured in the indicated groups using RT-PCR [16, 18]. Briefly, total RNA from myocytes (100 mg) was extracted using Trizol (Invitrogen). After spectrophotometric quantification and resolving of RNA integrity using a formaldehyde agarose gel, reverse transcription was carried out using an oligo (dT) primer and superscript II RT (Invitrogen). cDNA was amplified using LPL specific primers; 5'-A T C C A G C T G G G C C T A A C T T T - 3 ' (left) and 5 ' - A A T G G C T T C T C C A A T G T T G C - 3 ' (right). The p-actin gene was amplified as an internal control using 5'-T G G T G G G T A T G G G T C A G A A G G - 3 ' (left) and 5 ' - A T C C T G T C A G C G A T G C C T G GG-3' (right). The linear range was found to be between 15-30 cycles. The amplification parameters were set at: 94°C for 1 min, 58°C for 1 min and 72°C for 1 min, for a total of 30 cycles. The PCR products were electrophoresed on a 1.7% agarose gel containing ethidium bromide. Expression levels were represented as the ratio of signal intensity for LPL mRNA relative to P-actin mRNA. 5.2.7 Immunolocalization of filamentous (F) and globular (G) cardiomyocyte actin Briefly, myocytes were plated on laminin coated cover glass slides and rinsed with PBS. Myocytes were fixed for 10 min with 4 % paraformaldehyde in PBS, permeabilized with 0.1% Triton X-100 in PBS for 3 min, treated with PBS containing 1% BSA for 20 minutes, and finally rinsed with PBS. Cells were double stained with DNAasel AlexaFluor®594 and Pvhodamine®488 Phalloidin to colocalize monomelic globular actin (red, G actin), and polymerized filamentous actin (green, F actin) [30, 31]. The unbound fluorescent probe was rinsed with PBS buffer and slides were visualized using a Biorad 600 confocal microscope at 1260X magnification. For quantitative estimation of F & G actin, cells were plated on a laminin coated 24-well plate at a density of 75,000 myocytes/ per well. After incubation, cells were rinsed, fixed and permeabilized as described above. The increase in green and red fluorescence 130 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 was monitored in a spectroflurometer (green; excitation-485 nm, emission-530 nm; red; excitation-530 nm, emission-590 nm). Background was identified in wells without cells. Data is expressed as fluorescence units (F.U.), after subtracting the background. An increase in the F actin-to-G actin ratio was assumed to represent polymerization of actin filaments [32]. 5.2.8 Adipose tissue LPL activity Epididymal adipose tissue was isolated from fed rats and weighed [33]. Following incubation of adipose tissue with or without LPA, basal and HR-LPL activity was measured in the incubation medium. Results are expressed as mU of LPL per g of tissue. 5.2.9 Treatments To investigate the influence of LPA on LPL, isolated hearts and myocytes were treated with LPA for 1 h, and LPL activity determined. LPA was solubilized in chloroform:methanol, and subsequently in Krebs-Henseleit HEPES buffer. In the indicated experiment, control rat hearts were perfused with LPA (1 nM) for 60 mins, and HR-LPL activity subsequently determined. Control myocytes were also incubated with increasing concentrations of LPA (1-100 nM) for 60 min, and LPL activity measured. The effects of LPA in myocytes were also determined in the presence or absence of 1 pM cytochalasin D (CTD, an actin polymerization inhibitor) [34], 30 pM Suramin (SUR, an LPA receptor antagonist) [35], or 10 pM Y-27632 (a ROCK blocker) [36]. Diazoxide (DZ) a selective K + A T P channel opener decreases insulin secretion and causes hyperglycemia [17]. DZ (100 mg/kg) was administered i.p., the animals euthanized after 4 h, and hearts isolated for Western blot analysis. In the indicated experiment, rats were injected into the tail vein with a rapid acting insulin (Humulin R, 8 units), 1 h after DZ administration (following verification of hyperglycemia). The animals were killed after 90 mins (time required for establishment of sustained euglycemia), and hearts isolated and frozen for Western blot analysis. 131 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.2.10 Serum measurements Blood samples were removed from animals and centrifuged immediately to collect serum that was stored at -20°C until assayed. Diagnostic kits were used to measure glucose, T G (Sigma), and non-esterified fatty acid (Wako). 5.2.11 Materials [3H]-triolein was purchased from Amersham Canada. Heparin sodium injection (Hapalean; 1000 USP U/mL) was obtained from Oraganon Teknika. All other chemicals were obtained from Calbiochem and Sigma Chemicals. ECL® detection kit was obtained from Amersham. Rho A and ROCK I & II antibodies were obtained from Santa Cruz Biotechnology (CA, USA). 5.2.12 Statistical analysis Values are means + SE. LPL activity in response to heparin perfusion over time was analyzed by multivariate analysis (two-way ANOVA) of variance using the NCSS. Wherever appropriate, one-way A N O V A followed by the Tukey or Bonferroni tests or the unpaired and paired Student's Mest was used to determine differences between group mean values (as indicated in the specific figure legends). The level of statistical significance was set at PO.05. 132 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.3 Resu l ts 5.3.1 LPA augments myocyte LPL activity Myocardial distribution of LPL protein in mouse heart demonstrates that 78% of the total LPL is present in myocytes, 3-6% in the interstitial space, and 18% in the capillary endothelium [9, 37]. We evaluated whether LPA can augment LPL in myocytes. Incubation of myocytes with graded concentrations of LPA (1-100 nM) appreciably enhanced basal and HR-LPL activity in the medium (Fig. 5-1), without influencing intracellular LPL in the myocyte pellet (unpublished observation). Interestingly, changes in LPL activity were independent of shifts in mRNA, suggesting a posttranscriptional increase in myocyte LPL (Fig. 5-1, inset). 5.3.2 Augmentation of HR-LPL in LPA perfused isolated hearts LPL at the coronary lumen is an outcome of translocation of the enzyme from the myocyte cell surface [15, 18]. To determine whether the influence of LPA on myocyte LPL is subsequently reflected in an increase in enzyme at the vascular lumen, isolated control hearts were perfused retrogradely with LPA for 1 h, and subsequently, heparin was added to release luminal LPL into the coronary perfusate (Fig. 5-2). LPL discharge was rapid, and peak activity, likely representing LPL located at or near the endothelial surface, was observed within 1 min. Compared to control rat hearts, there was a substantial increase in coronary LPL activity (-200%) at the vascular lumen following 1 h of LPA perfusion (Fig. 5-2). 5.3.3 Inhibition of LPA mediated augmentation of myocyte HR-LPL Functional effects of LPA require activation of an endothelial gene differentiating (EDG) family of receptors [21, 23]. We attempted to block these receptors using SUR [35]. The ability of LPA to augment myocyte HR-LPL was inhibited by this putative LPA receptor antagonist (Fig. 5-3), suggesting that the effect of this lysophospholipid is receptor mediated. When incubated for 30 min, SUR per se did not influence myocyte LPL (Con: 195 ± 12, SUR: 221 ± 42; nmol/hr/106 cells). To investigate the involvement of the actin cytoskeleton in LPA 133 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 mediated augmentation of myocyte LPL, myocytes were pretreated with CTD, an actin polymerization inhibitor [34], prior to incubation with LPA. CTD abolished the effect of LPA to increase myocyte HR-LPL (Fig. 5-3), without any effect on basal activity (Con: 195 ± 12, CTD: 127 ± 16; nmol/hr/106 cells). 5.3.4 LPA induces cardiomyocyte actin polymerization Actin cytoskeleton has been implicated in managing myocyte LPL activity [25]. To determine whether LPA mediated augmentation of myocyte LPL involves F actin polymerization, cells were double stained with DNAsel AlexaFluor®594 and Rhodamine®488 Phalloidin to colocalize monomelic G actin (red), and polymerized F actin (green). In the resting cardiomyocyte, the proportion of G actin is consistently higher than polymerized F actin, and is predominantly localized along the cell periphery. LPA induced formation of a dense parallel F actin fibres, with complete absence of G actin (Fig. 5-4). Pretreatment with CTD markedly reduced F actin whereas G actin increased, consistent with F actin depolymerization. The ratio of fluroscence of Rhodamine and DNAse 1 (F-to-G actin ratio) was used to quantify actin cytoskeleton organization; an increase in F-to-G actin ratio indicates actin polymerization. CTD pretreatment abolished LPA's ability to increase this ratio (>60%) (Fig. 5-4). 5.3.5 LPA induced augmentation of HR-LPL requires RhoA and ROCK activation Actin cytoskeleton organization is regulated by small Rho GTPases [38, 39], a subgroup of the Ras superfamily, with RhoA being its most extensively characterized member . Following stimulation, cytosolic RhoA translocates to the cell membrane [39]. Effectors of RhoA include serine/threonine kinases like Rho kinases (ROCK I and ROCK II) [40], which phosphorylate downstream substrates [40]. Similar to RhoA, ROCKs also migrate to the cell membrane upon activation [40]. Growth factors and LPA are characterized as upstream regulators of RhoA [21, 22, 39]. Indeed, incubation of myocytes with LPA for 1 h caused significant membrane translocation of RhoA (Fig. 5-5, left panel) and ROCK I (Fig. 5-5, right panel) when compared 134 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 to control myocytes. ROCK II remained unaffected by LPA (unpublished observation). Incubation of myocytes with Y-27632 not only blocked membrane translocation of Rho kinase [as measured by densitometry; CON: 1.3; LPA:2.2; LPA+Y27632:1.6; units, PO.05], but also appreciably reduced basal and HR-LPL activity obtained with LPA (Fig. 5-6). When incubated for 30 min, Y27-632 per se did not influence myocyte basal LPL activity (Con: 195 ± 12, Y27-632: 143 ± 12; nmol/hr/106 cells). 5.3.6 LPA augments adipose tissue LPL activity Given the difficulty in differentiating sarcomeric from non-sarcomeric actin in determining the LPL secretory function of LPA, we evaluated if this lysophospholipid can augment LPL in adipose tissue. Incubation of adipose tissue with 1 nM LPA significantly enhanced basal and HR-LPL activity in the medium (Fig. 5-7). 5.3.7 Rho A and ROCK are activated during acute hyperglycemia We have previously reported that DZ, a selective K + A T P channel opener, decreases insulin secretion and causes hyperglycemia within 1 h [17]. In addition, there was a substantial increase in coronary LPL activity and immunofluorescence in DZ treated hearts. To evaluate if this change in LPL during hyperglycemia is related to RhoA and ROCK activation, we measured membrane translocation of these mediators in DZ hearts. DZ hearts demonstrated significant membrane translocation of RhoA (Fig. 5-8, left panel) and ROCK I (Fig. 5-8, right panel) when compared to control. To verify whether DZ mediated RhoA and ROCK I activation was secondary to hyperglycemia and not DZ per se, insulin was administered to DZ treated animals. Insulin not only normalized blood glucose and cardiac LPL (42), but also abrogated the DZ mediated membrane translocation of ROCK I [as measured by densitometry; CON: 0.36; DZ: 2.33; DZ+Ins: 0.39; units, PO.05] and RhoA [as measured by densitometry; CON: 0.58; DZ :3.97; DZ+Ins :1.12; units, PO.05] . 135 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.4 Discussion Hydrolysis of TG-rich lipoproteins by LPL positioned at the endothelial surface of the coronary lumen is suggested to be the principal source of FA for myocardial energy utilization [11]. Endothelial cells do not produce LPL and hence the enzyme is synthesized in myocytes [12]. Following enzyme activation and transport by mechanisms that are largely unknown, LPL is transported onto myocyte cell surface HSPG, before it is translocated onto comparable HSPG binding sites on the luminal side of the coronary vessel wall [13]. Heparan sulfate oligosaccharides, fragments of myocyte HSPG, have then been shown to act as extracellular chaperones for LPL, transporting the enzyme in its active form towards the apical surface of endothelial cells [14, 19]. Mediators responsible for the cleavage and transfer of LPL from the parenchymal cell surface to the lumen include the lipolytic byproduct LPC [20]. At least in the whole heart, the LPL augmenting property of LPC likely required endothelial PKC activation, and formation of LPA [20]. One of the best-characterized effects of LPA is actin cytoskeleton reassembly [21-24]. Given that incubation of cultured cardiomyocytes with insulin and dexamethasone stimulated basal and HR-LPL activities, a process involving actin cytoskeleton reorganization [25], we hypothesized that LPA augments cardiomyocyte LPL secretion through its effects on actin cytoskeleton. In this study, for the first time, we demonstrate that the LPA mediated augmentation of cardiomyocyte LPL involves the actin cytoskeleton. ATP dependent reversible polymerization of G to F actin provides the cell structural framework, in addition to mechanical force that allows for changes in contraction [41], locomotion, chemotaxis, and more importantly vesicular transport [42]. This polymerized F actin network beneath the cell membrane has been proposed to be a negative modulator of vesicular secretion, acting as a physical barrier as it depolymerizes during exocytosis [43, 44]. Other studies have suggested an obligatory role for F actin formation in facilitating ligand induced protein secretion [45]. Thus, in adrenal chromaffin cells, mobility of secretory granules 136 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 is mediated by actin filament formation, a process that is blocked by lantrunculins that disassemble cortical actin [46]. In addition, insulin mediated GLUT-4 translocation in skeletal muscle [47] and adipocytes require an intact F actin [48]. LPA is an established upstream mediator of actin polymerization, and in adrenal tumor PC 12 cells, LPA stimulated release of acetylcholine was inhibited when actin reorganization was disrupted using botulinum toxin [49]. In the current study, incubation of myocytes with LPA induced formation of a dense and organized network of thick and parallel F actin fibers, an effect that was abrogated by pretreatment with CTD. Since this effect of LPA on F actin formation occurred at a time when HR-LPL activity was augmented, our data suggests that actin polymerization likely facilitates secretion of this enzyme in myocytes. LPA induced polymerization of actin requires activation of E D G family of G-protein-coupled receptors [21-24]. In this study, we confirmed that the effects of LPA on actin cytoskeleton and myocyte LPL are receptor mediated as pretreatment with suramin, a putative LPA receptor antagonist, blocked these effects. G-proteins that participate in LPA receptor mediated effects include Gj/0, G q , and G12/13 [50], with activation of the G12/13 pathway being most closely linked to actin polymerization [51]. Stimulation of G12/13 activates small Rho GTPases [50, 52], specifically RhoA, a subgroup of the Ras superfamily that is identified as the essential upstream regulators of actin polymerization. Following receptor activation, cytosolic RhoA translocates to the cell membrane [39]. Effectors of RhoA include serine/threonine kinases like Rho kinases (ROCK I and ROCK II) [40], which also migrate to the cell membrane upon RhoA induced activation, with ensuing phosphorylation of downstream substrates [40]. Exposure of myocytes to LPA for 60 min facilitated significant membrane translocation of RhoA and ROCK 1. Interestingly, Y-27632 pretreatment not only blocked membrane translocation of ROCK 1, but also appreciably reduced basal and HR-LPL activity obtained with LPA. Overall, 137 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 these results indicate that LPA likely facilitates secretion of preformed LPL, a process that is dependent on polymerized actin. In adipose tissue, FA for T G storage are largely obtained through LPL-catalyzed hydrolysis of circulating lipoproteins [53]. Like in the heart, dietary and hormonal factors also influence the enzyme in this tissue. For example, with fasting, LPL activity decreases [33], whereas insulin is known to augment adipose tissue LPL [54]. To evaluate whether the effects of LPA on myocyte LPL can be duplicated in adipose tissue, we measured HR-LPL following treatment with LPA. As incubation of adipose tissue with LPA significantly enhanced basal and HR-LPL activity, our data suggests that sarcomeric actin likely has a limited role in influencing the LPL secretory function of LPA in the myocyte. Interestingly, peroxisome proliferator-activated receptor gamma (PPARY) was recently characterized as an intracellular receptor for LPA [55], and PPARy activation is known to enhance adipose tissue LPL activity [56]. During diabetes, LPL increases to guarantee F A supply to the diabetic heart in order to compensate for the diminished contribution of glucose as an energy source, allowing the heart to maintain its function [3, 15, 16]. Thus, following acute hypoinsulinemia induced with DZ, HR-LPL activity and protein increases at the coronary lumen [17]. As these effects of DZ were inhibited following suppression of circulating T G hydrolysis, lipolytic byproducts like LPC were implicated in explaining this enzyme increase [17]. LPA is a known metabolite of LPC [57]. We hypothesized that the increase in HR-LPL activity following DZ could be related to an LPA induced activation of RhoGTPases. Given that the effects of LPA on myocyte LPL were observed at 1 nM, and the technical difficulty associated with measuring trivial concentrations of this lysophospholipid, we measured the activation of downstream mediators facilitating actin polymerization in hearts from DZ treated animals. Similar to the results obtained when myocytes were incubated with LPA, hyperglycemia also caused significant membrane translocation of RhoA and ROCK 1 in whole hearts. Interestingly, injection of insulin 138 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 normalized blood glucose and luminal LPL (42), in addition to preventing DZ induced membrane translocation of RhoA and ROCK I. Hence, a mechanism for the acute effect of hyperglycemia in augmenting cardiac HR-LPL could involve RhoGTPase activation and actin reorganization. In summary, LPA induced increases in myocyte and adipose tissue LPL occurred via posttranslational mechanisms and processes that likely required RhoA activation and actin polymerization (Fig. 5-9). In congenital generalized lipodystrophy, where LPA conversion to phosphatidic acid is impaired, levels of LPA increase with augmented T G deposition in skeletal muscle [58] and associated insulin resistance [59]. Whether this process occurs via increases in LPL merits further investigation. In the heart, LPA, by increasing LPL, may initiate lipid oversupply with resulting muscle fiber degeneration [60], extensive lipid deposition [60], proliferation of mitochondria and peroxisomes [60], and eventual impairment of contractile function [61]. 139 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.5 Figures C O N LPA CON 1 10 50 100 (nM) LPA Fig. 5-1 Myocytes (0.4 x 106 cells/ml) were incubated either in the absence or presence of LPA for 1 h. Heparin (5 U/mL) was then added and after another 60 min, a 1 ml aliquot was aspirated, centrifuged, supernatant separated and LPL activity determined. Results are means ± SE of three rats in each group. *Significantly different from its corresponding basal, "Significantly different from control heparin releasable, +Significantly different from control basal, PO.05. Myocytes were also used to measure LPL gene expression as described in methods. The inset is a single representative gel. Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 800 i o 13 CL > O Ii < 600 -I 400 i 200 i 0 CON LPA 2 4 6 Time (min) 8 10 F i g . 5-2 Control hearts were isolated and perfused with 1 nM LPA for 1 h in the recirculating retrograde mode. Thereafter, coronary LPL was released with heparin (5 U/ml). Coronary effluents were collected (for 10 s) at different time points over 10 mins. LPL activity was assayed using radiolabeled triolein. Results are the means ± SE of four rats in each group. Two-way A N O V A for repeated measures was carried out to determine statistical differences between means. 141 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 1500 i Fig. 5-3 Myocytes were pretreated with suramin or cytochlasin D for 30 min prior to incubation with 1 nM LPA for 1 h. Heparin was then added to the cell suspension, and after another 60 min, a 1 ml aliquot was aspirated, centrifuged and the supernatant separated. The released LPL activity in the medium was determined. Results are means ± SE of four rats in each group. * Significantly different all other groups, P<0.05. 142 Chapter 5: Am. J. Heart and Circ. PhysioL 288 (2005) H2802-H2810 C O N L P A CON LPA LPA+CTD Fig. 5-4 Myocytes were double stained with DNAasel AlexaFluor®594 and Rhodamine®488 Phalloidin to colocalize red, G actin and green, F actin and visualized at 1260x magnification. Cells were also plated, at a density of 75,000 myocytes/ per well and the increase in red and green fluorescence was monitored in a spectroflurometer. Data is expressed as fluorescence units (F.U.), after subtracting the background. * Significantly different from other groups PO.05. 143 C M M Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 C M C M C O N L P A C O N L P A RHOA 100 o (A C O i o £ o 50 </> C 5cS Cytosol Membrane ROCK-1 100 n o i/) c o o +J >. o o 50 • Ep 00 C b Co 0 s CON LPA CON LPA Fig. 5-5 Myocytes were incubated with LPA for 60 min, and homogenates prepared were subjected to cytosolic and membrane separation. Identification of RhoA and ROCK I protein was carried out using mouse monoclonal anti RhoA, or rabbit polyclonal anti ROCK I primary antibodies at dilutions of 1:250 and 1:180 respectively followed by incubation with goat anti-mouse (RhoA) or goat anti-rabbit (ROCK) horseradish peroxidase conjugated secondary antibodies at dilutions of 1:3000 and 1:5000 respectively. Results are means ± SE of five rats in each group. 'Significantly different from control membrane fraction P<0.05. 144 c Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 M C M C M CON LPA LPA +Y27632 CON LPA LPA+Y27632 Fig. 5-6 Myocytes were pretreated with 10 pM Y-27632, an inhibitor of ROCK, for 30 min prior to incubation with 1 nM LPA for 1 h. Heparin was then added to the cell suspension, and after 1 h, a 1 ml aliquot was aspirated, centrifuged and the supernatant LPL activity determined. Data are means ± SE of four different hearts in each group. * Significantly different from its corresponding basal, "Significantly different from control heparin releasable, +Significantly different from control basal, PO.05. The inset depicts a single representative Western blot of cardiac ROCK I in cytosolic (C) and membrane (M) fractions of myocytes incubated with or without 1 nM LPA either in the presence or absence of 10 pM Y-27632. 145 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 CON LPA Fig. 5-7 Epididymal adipose tissue fat pads (0.5 g) were incubated either in the absence or presence of 1 nM LPA for 1 h. Heparin (5 U/mL) was then added to the cell suspension, and after another 60 min, a 1 ml aliquot was aspirated, centrifuged and the supernatant separated for measurement of LPL activity. LPL activity is expressed as milliunits (mU) of activity per g of tissue. Results are means ± SE of three rats in each group. * Significantly different from its corresponding basal, "Significantly different from control heparin releasable, """Significantly different from control basal, PO.05. 146 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 C M C M C M C M CON DZ CON DZ CON DZ CON DZ Fig. 5-8 Diazoxide (100 mg/kg) was administered to control animals and hearts isolated after 4 h. Heart homogenates were subjected to cytosolic and membrane separation, and respective protein contents determined. Identification of RhoA and ROCK I protein was carried out as described above. Results are means ± SE of five rats in each group. 'Significantly different from control membrane fraction, P<0.05. 147 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 L P A Cardiomyocyte Fig. 5-9 LPA, likely through its effects on G12/13 protein coupled receptors, stimulates RhoGTPases, specifically RhoA and its downstream effector ROCK I to promote F actin polymerization. Actin rearrangement promotes vesicular transport of the enzyme from the golgi complex to the myocyte plasma membrane. From this location, LPL translocates to the endothelial lumen where lipoprotein hydrolysis is facilitated. 148 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 5.6 B ib l iography [1] Lteif A A , Mather KJ, Clark C M . Diabetes and heart disease an evidence-driven guide to risk factors management in diabetes. Cardiol Rev 2003; 11:262-74. [2] Sowers JR, Epstein M , Frohlich ED. Diabetes, hypertension, and cardiovascular disease: an update. Hypertension 2001;37:1053-9. [3] Rodrigues B, Cam M C , McNeill JH. Myocardial substrate metabolism: implications for diabetic cardiomyopathy. J Mol Cell Cardiol 1995;27:169-79. 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Insulin and dexamethasone stimulation of cardiac lipoprotein lipase activity involves the actin-based cytoskeleton. Biochem J 1999;340 ( Pt 2):485-90. [26] Sayas CL, Avila J, Wandosell F. Regulation of neuronal cytoskeleton by lysophosphatidic acid: role of GSK-3. Biochim Biophys Acta 2002;1582:144-53. [27] Rodrigues B, Spooner M , Severson DL. Free fatty acids do not release lipoprotein lipase from isolated cardiac myocytes or perfused hearts. Am J Physiol 1992;262:E216-23. [28] Porter K E , Turner NA, O'Regan DJ, Balmforth AJ, Ball SG. Simvastatin reduces human atrial myofibroblast proliferation independently of cholesterol lowering via inhibition of RhoA. Cardiovasc Res 2004;61:745-55. [29] Levent A, Buyukafsar K. Expression of Rho-kinase (ROCK-1 and ROCK-2) and its substantial role in the contractile activity of the sheep ureter. Br J Pharmacol 2004;143:431-7. [30] Haugland RP, You W, Paragas VB, Wells KS, DuBose DA. Simultaneous visualization of G- and F-actin in endothelial cells. J Histochem Cytochem 1994;42:345-50. 151 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 [31] Aoki H, Izumo S, Sadoshima J. Angiotensin II activates RhoA in cardiac myocytes: a critical role of RhoA in angiotensin II-induced premyofibril formation. Circ Res 1998;82:666-76. [32] Begum N, Sandu OA, Duddy N. Negative regulation of rho signaling by insulin and its impact on actin cytoskeleton organization in vascular smooth muscle cells: role of nitric oxide and cyclic guanosine monophosphate signaling pathways. Diabetes 2002;51:2256-63. [33] Wu G, Olivecrona G, Olivecrona T. The distribution of lipoprotein lipase in rat adipose tissue. Changes with nutritional state engage the extracellular enzyme. J Biol Chem 2003;278:11925-30. [34] Mortensen K, Larsson LI. Effects of cytochalasin D on the actin cytoskeleton: association of neoformed actin aggregates with proteins involved in signaling and endocytosis. 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Physiol. 288 (2005) H2802-H2810 [41] Davani EY, Dorscheid DR, Lee CH, van Breemen C, Walley KR. Novel regulatory mechanism of cardiomyocyte contractility involving ICAM-1 and the cytoskeleton. Am J Physiol Heart Circ Physiol 2004;287:H1013-22. [42] Stamnes M . Regulating the actin cytoskeleton during vesicular transport. Curr Opin Cell Biol 2002;14:428-33. [43] Vitale M L , Rodriguez Del Castillo A, Tchakarov L, Trifaro JM. Cortical filamentous actin disassembly and scinderin redistribution during chromaffin cell stimulation precede exocytosis, a phenomenon not exhibited by gelsolin. J Cell Biol 1991; 113:1057-67. [44] Morita K, Oka M , Hamano S. Effects of cytoskeleton-disrupting drugs on ouabain-stimulated catecholamine secretion from cultured adrenal chromaffin cells. Biochem Pharmacol 1988;37:3357-9. [45] O'Konski MS, Pandol SJ. Cholecystokinin JMV-180 and caerulein effects on the pancreatic acinar cell cytoskeleton. Pancreas 1993;8:638-46. [46] Steyer JA, Aimers W. Tracking single secretory granules in live chromaffin cells by evanescent-field fluorescence microscopy. Biophys J 1999;76:2262-71. [47] Brozinick JT, Jr., Hawkins ED, Strawbridge AB, Elmendorf JS. Disruption of cortical actin in skeletal muscle demonstrates an essential role of the cytoskeleton in glucose transporter 4 translocation in insulin-sensitive tissues. J Biol Chem 2004;279:40699-706. [48] Kanzaki M , Pessin JE. Insulin-stimulated GLUT4 translocation in adipocytes is dependent upon cortical actin remodeling. J Biol Chem 2001;276:42436-44. [49] Ishida H, Zhang X, Erickson K, Ray P. Botulinum toxin type A targets RhoB to inhibit lysophosphatidic acid-stimulated actin reorganization and acetylcholine release in nerve growth factor-treated PC12 cells. J Pharmacol Exp Ther 2004;310:881-9. [50] Dhanasekaran N, Dermott JM. Signaling by the G12 class of G proteins. Cell Signal 1996;8:235-45. Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 [51] Kranenburg O, Poland M , van Horck FP, Drechsel D, Hall A, Moolenaar WH. Activation of RhoA by lysophosphatidic acid and Galphal2/13 subunits in neuronal cells: induction of neurite retraction. Mol Biol Cell 1999;10:1851-7. [52] Buhl A M , Johnson NL, Dhanasekaran N, Johnson GL. G alpha 12 and G alpha 13 stimulate Rho-dependent stress fiber formation and focal adhesion assembly. J Biol Chem 1995;270:24631-4. [53] Zechner R, Strauss J, Frank S, Wagner E, Hofmann W, Kratky D, Hiden M , Levak-Frank S. The role of lipoprotein lipase in adipose tissue development and metabolism. Int J Obes Relat Metab Disord 2000;24 Suppl 4:S53-6. [54] Farese RV, Jr., Yost TJ, Eckel RH. Tissue-specific regulation of lipoprotein lipase activity by insulin/glucose in normal-weight humans. Metabolism 1991;40:214-6. [55] Mclntyre T M , Pontsler A V , Silva AR, St Hilaire A, Xu Y, Hinshaw JC, Zimmerman GA, Hama K, Aoki J, Arai H, Prestwich GD. Identification of an intracellular receptor for lysophosphatidic acid (LPA): LPA is a transcellular PPARgamma agonist. Proc Natl Acad Sci U S A 2003;100:131-6. [56] Laplante M , Sell H, MacNaul K L , Richard D, Berger JP, Deshaies Y. PPAR-gamma activation mediates adipose depot-specific effects on gene expression and lipoprotein lipase activity: mechanisms for modulation of postprandial lipemia and differential adipose accretion. Diabetes 2003;52:291-9. [57] Aoki J. Mechanisms of lysophosphatidic acid production. Semin Cell Dev Biol 2004;15:477-89. [58] Agarwal A K , Garg A. Congenital generalized lipodystrophy: significance of triglyceride biosynthetic pathways. Trends Endocrinol Metab 2003;14:214-21. [59] Mingrone G, Rosa G, Di Rocco P, Manco M , Capristo E, Castagneto M , Vettor R, Gasbarrini G, Greco A V . Skeletal muscle triglycerides lowering is associated with net 154 Chapter 5: Am. J. Heart and Circ. Physiol. 288 (2005) H2802-H2810 improvement of insulin sensitivity, TNF-alpha reduction and GLUT4 expression enhancement. Int J Obes Relat Metab Disord 2002;26:1165-72. [60] Levak-Frank S, Radner H, Walsh A, Stollberger R, Knipping G, Hoefler G, Sattler W, Weinstock PH, Breslow JL, Zechner R. Muscle-specific overexpression of lipoprotein lipase causes a severe myopathy characterized by proliferation of mitochondria and peroxisomes in transgenic mice. J Clin Invest 1995;96:976-86. [61] Yagyu H, Chen G, Yokoyama M , Hirata K, Augustus A, Kako Y, Seo T, Hu Y, Lutz EP, Merkel M , Bensadoun A, Homma S, Goldberg IJ. Lipoprotein lipase (LpL) on the surface of cardiomyocytes increases lipid uptake and produces a cardiomyopathy. J Clin Invest 2003;111:419-26. 155 Chapter 6: Invited Review Accepted in Cardiovas. Res. 6.0 Conclusions and future directions 6.1 S u m m a r y In conclusion, following hypoinsulinemia/hyperglycemia, the heart undergoes metabolic transformation wherein it switches energy production to exclusive P-oxidation of FA. This process is made possible by amplification of coronary LPL, thereby allowing uninterrupted FA supply to the diabetic heart. Since this amplification of coronary LPL occurred in the absence of any change in LPL mRNA, posttranslational mechanisms likely contributed to this increase [1]. Of these posttranslational events, an increase in vascular LPL binding sites was unlikely to be the mechanism responsible for this effect [1]. Overall, our data suggest that subsequent to a decline in glucose utilization by the diabetic heart, LPL is rapidly recruited to the luminal surface in search of T G substrates [2]. Since administration of WR 1339 in DZ treated animals restricted its ability to enhance luminal LPL, circulating T G hydrolysis was proposed to be a key metabolic trigger in augmenting vectorial transfer of the enzyme [2]. In this regard, the T G breakdown products LPC and LPA were identified as prospective circulating mediators responsible for enhancing luminal and myocyte LPL [3, 4]. Finally, we established that these lysophospholipids facilitate enzyme translocation through their effect on RhoGTPases and actin cytoskeleton rearrangement [3]. We have focused on this area of research given the correlation between LPL and its ability to accelerate heart disease. Despite insulin therapy, the risk of heart disease and hypertension is increased 2-4 fold in diabetic patients and altered FA metabolism is suggested to be strong causative factor for the progression of this heart failure. Since changes in cardiac LPL can result in abnormal FA supply and utilization by the heart tissue, it could potentially initiate and sustain cardiac dysfunction during diabetes (Fig. 6-1). Since effective blood glucose control is often compromised in patients with insulin resistance and diabetes, and as pharmaceutical management 156 Chapter 6: Invited Review Accepted in Cardiovas. Res. can never duplicate the exquisite control of glucose observed in healthy humans, it is likely that increased luminal LPL, and abnormal F A supply and utilization by the heart could result in lipotoxic events that will initiate a number of metabolic, morphological, and mechanical changes, and eventually cardiac disease. Given the proposed role for L P L in initiating diabetic cardiomyopathy through its ability to enhance FA supply, restricting cardiac L P L translocation could be a therapeutic advantage as this would lead to metabolic switching to glucose utilization [5]. Interestingly, a recent study demonstrated that cardiac specific L P L knockout mice displayed enhanced expression of GLUT-1, GLUT-4 and insulin receptor substrate 2 with a parallel decline in the expression of PDK-4 and insulin receptor substrate-1 [6]. This resulted in markedly enhanced basal glucose uptake in these hearts [6]. Future studies are required to establish conclusively that increased L P L in the diabetic heart is causally related to the development of cardiomyopathy [5]. In the STZ diabetic rat, this awaits the discovery of a pharmacological agent that specifically inhibits cardiac L P L activity in vivo. In terms of drug discovery and application of this research, development of cardiac specific lysophospholipid receptor or RhoGTPase inhibitors would help in slowing L P L translocation to the coronary lumen thus interrupting F A supply to the heart. 157 Chapter 6: Invited Review Accepted in Cardiovas. Res. 6.2 Figure LIPOTOXICITY Fig . 6-1 FA induced lipotoxicity in the diabetic heart. The earliest change that occurs in the diabetic heart is altered energy metabolism, where the heart switches to exclusively using FA for energy supply. This abnormal FA utilization by cardiac tissue may have lethal consequences including a number of metabolic, morphological, and mechanical changes, and more recently, in "lipotoxicity. In lipotoxicity, FA accumulate and can, either by themselves or via production of second messengers such as ceramides, provoke cell death. 158 Chapter 6: Invited Review Accepted in Cardiovas. Res. 6.3 B ib l iography [1] Pulinilkunnil T, Abrahani A, Varghese J, Chan N, Tang I, Ghosh S, Kulpa J, Allard M , Brownsey R, Rodrigues B. Evidence for rapid "metabolic switching" through lipoprotein lipase occupation of endothelial-binding sites. J Mol Cell Cardiol 2003;35:1093-103. [2] Pulinilkunnil T, Qi D, Ghosh S, Cheung C, Yip P, Varghese J, Abrahani A, Brownsey R, Rodrigues B. Circulating triglyceride lipolysis facilitates lipoprotein lipase translocation from cardiomyocyte to myocardial endothelial lining. Cardiovasc Res 2003;59:788-97. [3] Pulinilkunnil T, An D, Ghosh S, Qi D, Kewalramani G, Yuen G, Virk N, Abrahan A, Rodrigues B. Lysophosphatidic acid mediated augmentation of cardiomyocyte lipoprotein lipase involves actin cytoskeleton reorganization. Am J Physiol Heart Circ Physiol 2005; [4] Pulinilkunnil T, An D, Yip P, Chan N, Qi D, Ghosh S, Abrahani A, Rodrigues B. Palmitoyl lysophosphatidylcholine mediated mobilization of LPL to the coronary luminal surface requires PKC activation. J Mol Cell Cardiol 2004;37:931-38. [5] Pulinilkunnil T, Sambandam N, Rodrigues B. Altered Substrate Utilization in the Diabetic Heart: Role of Lipoprotein Lipase. In: Dhalla NS, Frontiers in Cardiovascular Health, 9. Boston USA: Kluwer, 2003: 119-44 [6] Augustus A, Yagyu H, Haemmerle G, Bensadoun A, Vikramadithyan RK, Park SY, Kim JK, Zechner R, Goldberg IJ. Cardiac-specific knock-out of lipoprotein lipase alters plasma lipoprotein triglyceride metabolism and cardiac gene expression. J Biol Chem 2004;279:25050-7. 159 

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