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Cardiac cell death in early diabetes : modulation by glutathione and polyunsaturated fatty acids Ghosh, Sanjoy 2006

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CARDIAC CELL DEATH IN EARLY DIABETES: MODULATION BY GLUTATHIONE AND POLYUNSATURATED FATTY ACIDS by Sanjoy Ghosh B.Pharm., Bangalore University, India 1999 M . S c , The University of British Columbia, 2003 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 August 2006 © Sanjoy Ghosh, 2006 A B S T R A C T Diabetes is a growing problem in Canada. Cardiovascular disease (CVD) remains the major cause of morbidity and mortality among diabetic patients. Although the majority of C V D are attributed to coronary vascular disease, an increasing body of evidence supports the existence of an additional heart muscle defect, in the absence of any vascular pathology. Among other factors, this heart disease is associated with excessive cell death in both human and animal models of diabetes. It is possible that this loss of cells progressively leads to interstitial fibrosis, myocardial hypertrophy and eventual loss of contractile function. Previous studies had established that oxidative stress due to excessive reactive oxygen species (ROS) generation and/or inadequate ROS neutralization, may be a key player in causing cardiac cell death via apoptosis in early diabetes. We used streptozotocin (STZ, 55 mg/kg) to make Wistar rats acutely diabetic and after 4 days examined the different processes that regulate cardiac glutathione (GSH), an important endogenous antioxidant. Reduction in myocyte GSH in diabetic rats was accompanied by increased oxidative stress, excessive reactive oxygen species, and an elevated apoptotic cell death. In this model, loss of GSH was not associated with any change in either biosynthesis or recycling of GSH, but with increased efflux of GSH out of diabetic cardiomyocytes. Gene expression of multidrug resistance protein 1, a transporter implicated in effluxing GSH during oxidative stress, was also elevated in these hearts. In the next study, we reported a loss of GSH specifically from the cardiac mitochondria (mitochondrial GSH, mGSH), the chief source of ROS. Such loss in mGSH could explain increased mitochondrial oxidative damage, and apoptosis via increases in caspase-9 and 3 activities in these hearts. Finally, the central role for mGSH loss in cardiac apoptosis was validated as exogenous GSH supplementation induced a robust amplification in mGSH and prevented apoptosis in diabetic rat hearts. Apart from antioxidant levels, changes in cellular lipid homeostasis have also been linked to cell death in early diabetes. However, most of the existing studies on the cytotoxicity of fatty acids are on vascular cell systems and with saturated fatty acids like palmitic acid. In the last few decades, saturated fats from the human diet have been replaced by co-6 PUFA in form of vegetable oils. Thus, the effect of the more relevant co-6 PUFA on the heart, especially during diabetes remains unknown. Male Wistar rats were first fed a diet rich in co-6 PUFA [20% (wt/wt) sunflower oil] for 4 wk followed by streptozotocin to induce diabetes. After a brief period of hyperglycemia (4 days), hearts were excised for functional, morphological, and biochemical analysis. Interestingly, in diabetic rats, co-6 PUFA decreased caspase-3 activity, crucial for myocardial apoptosis. However, cardiac necrosis, an alternative mode of cell death, increased. Interestingly, depletion of cardiac ATP stores could explain this "switch" to necrotic cell death in these animals. This was supported by mitochondrial abnormalities, impaired substrate utilization, and enhanced triglyceride accumulation in co-6 PUFA fed diabetic hearts. Further analysis revealed that even in the absence of diabetes, a pro-inflammatory response is initiated in the heart following co-6 PUFA feeding via augmented phospholipase A2 activity and breakdown of cardiolipin, a mitochondrial phospholipid. co-6 PUFA fed hearts also demonstrated elevated inducible nitric oxide synthase expression, loss of Mn superoxide dismutase, and increased mitochondrial nitrotyrosine levels. In these hearts, oxidative damage to mitochondrial DNA (mDNA) was demonstrated by 8-hydroxyguanosine immunopositivity, overexpression of DNA repair enzymes, and a decrease in the mRNA expression of specific respiratory subunits encoded by the mDNA. Thus, although promoted as being beneficial, excess dietary co-6 PUFA, with its predisposition to cause mitochondrial nitrosative damage and necrosis following hyperglycemia, could accelerate myocardial abnormalities in diabetic patients. iii TABLE OF CONTENTS Abstract i i Table O f Contents iv List O f Tables ix List O f Figures x List O f Abbreviations x i i Acknowledgments xv Dedication xvi 1. I N T R O D U C T I O N 1.1 Historical V i e w On'Diabetes ' 1 1.2 A n Overview O f Diabetes Mellitus 1 . 1.3 Diabetes Is A Cardiovascular Disease 3 1.4 Basic Mechanisms O f Oxidative Stress 5 1.5 Synthesis, Metabolism A n d Action O f Glutathione (GSH) 7 1.6 Figure 10 1.7 References 11 2. C A R D I A C C E L L D E A T H I N E A R L Y D I A B E T E S 2.1 Hyperglycemia A n d Cellular Lipids: Common Pathways To Oxidative Stress ... 19 2.2 The Relevance O f Early Diabetes 20 2.3 Basic Mechanisms O f Cel l Death 22 2.4 Oxidant Induced Cel l Death In Experimental Diabetes 24 2.5 Figures 26 2.6 References 28 3 E F F E C T O F F A T T Y A C I D S O N C A R D I A C C E L L D E A T H I N D I A B E T E S 3.1 Saturated Fatty Acids 35 3.2 Monounsaturated Fatty Acids 38 3.3 The Western Diet 39 3.4 Biochemistry O f P U F A 40 iv 3.5 Probable Effects O f coB6 P U F A In The Diabetic Heart 41 3.6 Rationale A n d Objectives 43 3.7 Figure 45 3.8 References 46 4. INCREASED EFFLUX OF GLUTATHIONE CONJUGATE IN ACUTELY DIABETIC CARDIOMYOCYTES 4.1 Introduction 57 4.2 Material A n d Methods 59 4.2.1 Experimental Animals 59 4.2.2 Cardiac Apoptosis 59 4.2.3 Cardiac L ip id Peroxidation 60 4.2.4 Preparation O f Cardiac Myocytes 60 4.2.5 Estimation O f G S H 60 4.2.6 Cardiac Gene Expression 61 4.2.7 Glutathione Reductase Activity 61 4.2.8 G S H Efflux 62 4.2.9 L D H Release 63 4.2.10 Measurement O f R O S 63 4.2.11 Statistical Analysis 63 4.3 Results 64 4.3.1 Cardiac Apoptosis A n d Oxidative Stress In S T Z Diabetes 64 4.3.2 Glutathione Status In STZ-Diabetic Hearts 64 4.3.3 M R P Like Activity In Diabetic Cardiomyocytes 65 4.3.4 Decrease In Cardiomyocyte ROS Following Inhibition O f G S H Efflux 65 4.4 Discussion 66 4.5 Tables A n d Figures 69 4.6 References 76 5 CARDIOMYOCYTE APOPTOSIS INDUCED BY SHORT-TERM DIABETES REQUIRES MITOCHONDRIAL GSH DEPLETION 5.1 Introduction 81 5.2 Material A n d Methods 83 v 5.2.1 Experimental Animals 83 5.2.2 Treatments 83 5.2.3 Preparation O f Cardiac Myocytes 84 5.2.4 Estimation O f R O S 84 5.2.5 Determination O f Mitochondrial Membrane Potential 85 5.2.6 Separation O f Cytosol A n d Mitochondrial Fractions 86 5.2.7 Estimation O f Glutathione 86 5.2.8 Cardiomyocyte Apoptosis 87 5.2.9 Cardiac Caspase Activity 87 5.2.10 L ip id Peroxidation 87 5.2.11 Protein Carbonyl Content 88 5.2.12 Serum Lactate Dehydrogenase 88 5.2.13 Statistical Analysis 88 5.3 Results 89 5.3.1 Mitochondrial Dysfunction Following Acute Diabetes 89 5.3.2 Cardiac G S H A n d Apoptosis Following Acute Diabetes 90 5.3.3 G S H Modulation A n d Effects On Cardiac Apoptosis 90 5.3.4 Cardiac Oxidative Stress A n d Caspase Activi ty 91 5.4 Discussion 93 5.5 Figures 97 5.6 References 103 6. BRIEF EPISODE OF STZ-INDUCED HYPERGLYCEMIA PRODUCES CARDIAC ABNORMALITIES IN RATS FED A DIET RICH WITH co-6 PUFA 6.1 Introduction 109 6.2 Material and Methods 111 6.2.1 Experimental Animals 111 6.2.2 Separation A n d Characterization O f Cardiac Lipids 111 6.2.3 Cardiac Caspase-3 Activi ty. 112 6.2.4 Cardiac Necrosis 112 6.2.5 Estimation O f G S H 113 6.2.6 A T P Assay 113 vi 6.2.7 E l e c t r o n M i c r o s c o p y 113 6.2.8 Heart F u n c t i o n A n d Substrate O x i d a t i o n 114 6.2.9 S e r u m Measurements 114 6.2.10 Statist ical A n a l y s i s 114 6.3 Results 115 6.3.1 G e n e r a l Characterist ics O f E x p e r i m e n t a l A n i m a l s 115 6.3.2 Free P U F A Content In T h e Heart 115 6.3.3 C a r d i a c C e l l D e a t h 116 6.3.4 C a r d i a c G S H 117 6.3.5 C a r d i o l i p i n A n d M i t o c h o n d r i a l M o r p h o l o g y 117 6.3.6 G l u c o s e A n d Palmitate O x i d a t i o n 118 6.3.7 E l e c t r o n M i c r o s c o p y A n d Ultrastructural M o r p h o l o g y 118 6.3.8 E s t i m a t i o n O f Heart F u n c t i o n 118 6.4 D i s c u s s i o n 120 6.5 Tables and Figures 126 6.6 References ••••• 136 7. C H R O N I C PROVISION O F D I E T A R Y co-6 P U F A INDUCES M I T O C H O N D R I A L N I T R O S A T I V E D A M A G E AND C A R D I A C D Y S F U N C T I O N 7.1 Introduct ion 142 7.2 M a t e r i a l A n d M e t h o d s 144 7.2.1 E x p e r i m e n t a l A n i m a l s 144 7.2.2 Separation A n d Character izat ion O f C a r d i a c L i p i d s 144 7.2.3 T o t a l Phosphol ipase A 2 ( P L A 2 ) A c t i v i t y 145 7.2.4 Separation O f C y t o s o l i c A n d Particulate Fract ions 145 7.2.5 Western B l o t t i n g Procedures 146 7.2.6 Immunof luorescence 147 7.2.7 Gene E x p r e s s i o n 147 7.2.8 C a r d i a c F u n c t i o n 148 7.2.9 Statist ical A n a l y s i s 149 7.3 Results 150 7.3.1 C a r d i a c Content O f co-3 A n d co-6 P U F A 150 vii 7.3.2 Phospholipid Metabolism 1 5 0 7.3.3 Nitrosative Stress 1 5 0 7.3.4 Mitochondrial Damage 1 5 1 7.3.5 Loss Of Gene Expression Of Respiratory Enzyme Subunits 1 5 1 7.3.6 Cardiac Functional Parameters 1 5 2 7.4 Discussion 1 5 3 7.5 Tables and Figures 1 5 8 7.6 References 1 6 8 8. C O N C L U S I O N S 1 7 4 8.1 Future Directions 1 7 6 8.2 References 1 7 7 i. vii i LIST OF TABLES Table 4.1 General characteristics of the experimental animals 69 Table 4.2 Factors regulating intracellular GSH status 70 Table 6.1 Composition of experimental diets 126 Table 6.2 General characteristics of experimental animals 127 Table 7.1 Primers used for PCR amplification of genes 158 Table 7.2 Major cardiac co-6 and co-3 PUFA , 159 Table 7.3 Heart rate and cardiac output of isolated working hearts 160 ix L I S T O F F I G U R E S Figure 1.1 Pathways of intracellular glutathione (GSH) metabolism 10 Figure 2.1 Mechanisms of ROS induced injury in diabetes 26 Figure 2.2 Overview of the basic mechanisms of apoptosis 27 Figure 3.1 Modulation of cardiac cell death by saturated (palmitic acid) and monounsaturated fatty acids (MUFA) 45 Figure 4.1 Oxidative stress in cardiac myocytes isolated from control and STZ-diabetic rats 71 Figure 4.2 mClB-GSH efflux from isolated cardiomyocytes 72 Figure 4.3 MRP-1 gene expression in hearts from control and diabetic rats 73 Figure 4.4 Effect of different inhibitors on the efflux of mClB-GSH from diabetic cardiomyocytes 74 Figure 4.5 ROS status in control and diabetic cardiomyocytes 75 Figure 5.1 Reactive oxygen species in diabetic cardiomyocytes 97 Figure 5.2 Mitochondrial dysfunction following acute hyperglycemia 98 Figure 5.3 Effect of 4 days of diabetes on cardiac cell death and glutathione 99 Figure 5.4 Effect of different treatments on GSH and cardiac apoptosis 100 Figure 5.5 Mitochondrial oxidative stress and caspase activities in control and diabetic hearts. 101 Figure 5.6 Proposed scheme of myocardial apoptosis in acute diabetes 102 Figure 6.1 Effects of sunflower oil feeding and diabetes on cardiac PUFA levels 128 Figure 6.2 Quantitative estimation of cardiac caspase-3 activity 129 Figure 6.3 Effect of 4 days of diabetes on cardiac cell death in animals fed a moderately high PUFA diet for 4 weeks 130 Figure 6.4 Status of myocardial glutathione system as estimated by enzyme-recycling assay 131 Figure 6.5 Biochemical and ultrastructural changes in cardiac mitochondria following PUFA feeding and diabetes 132 Figure 6.6 Glucose and palmitate oxidation at low and high afterloads 133 Figure 6.7 Evaluation of ultrastructural cardiac morphology Figure 6.8 Cardiac function at low (80 mm Hg) and high (135 mm Hg) afterloads 1 3 5 x Figure 7.1 Effects of co-6 PUFA feeding on cardiac cardiolipin 161 Figure 7.2 Cardiac phospholipid metabolism 162 Figure 7.3 Cardiac nitrosative stress 163 Figure 7.4 Intracellular localization of peroxynitrite and 8-hydroxyguanosine by confocal microscopy 164 Figure 7.5 Intracellular superoxide dismutase (SOD) protein expression 165 Figure 7.6 Respiratory subunit gene and PGC-1 protein expression 166 Figure 7.7 Proposed pathways of mitochondrial damage and nuclear responses following consumption of a high fat (HF) diet rich in co-6 PUFA 167 xi LIST OF ABBREVIATIONS Ai | / m Mitochondrial membrane potential y-GCS Gamma glutamylcysteine synthase ul Microliter 8-OHG 8-hydroxyguanosine A G E Advanced glycation end products AIN American institutes of Nutrition A N O V A Analysis of variance BSO Buthionine sulfoximine C A D Coronary artery disease cGSH Cytosolic GSH 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, C M - H 2 D C F D A acetyl ester COX-1 cytochrome-c-oxidase subunit 1 Cu/Zn SOD Copper-zinc superoxide dismutase CYTb cytochrome b D E M Diethyl maleate DHA docosahexaenoic acid DTNB 5,5'-dithiobis-2-nitrobenzoic acid EPA Eicosapentaenoic acid ETC Electron transport chain F.U. Fluorescence units FA Fatty acids F A D H 2 Reduced flavin adenine dinucleotide GAD Glutamic acid decarboxylase G L C Gas liquid chromatography GPx Glutathione peroxidase GSH Glutathione GSSG Glutathione disulphide, oxidized glutathione xii HNE Hydroxynonenal HPLC High pressure liquid chromatography i.v. Intravenous IAA Insulin auto antibody ICA Islet cell auto antibodies IGF-1 Insulin like growth factor 1 iNOS Inducible nitric oxide synthase PO Palm oil ip Intraperitoneal Kb Kilobase K H solution Krebs-Henseleit solution L A D A Latent autoimmune diabetes in adults L D H Lactate dehydrogenase LPC Lysophosphatidylcholine LPL Lipoprotein lipase mClB Monochlorobimane M D A Malondialdehyde mDNA Mitochondrial DNA mGSH Mitochondrial GSH mM Millimolar Mn-SOD . Manganese-superoxide dismutase mRNA Messenger ribonucleic acid MRP Multi drug resistance protein M T Metallothionein M U F A Monounsaturated fatty acids N A C N-acetyl cysteine N A D H Reduced nicotinamide adenine dinucleotide NADPH Reduced nicotinamide adenine dinucleotide phosphate ND-1 N A D H dehydrogenase subunit 1 N F - K B Nuclear factor kappa B NIDDM Non-insulin-dependent diabetes mellitus xiii nM Nanomolar NO Nitric oxide OGG-1 8-oxoG-DNA glycosylase OxPhos, OXPHOS Oxidative phosphorylation PC Protein carbonyls PGC-1 Peroxisome proliferator-activated receptor gamma coactivator 1 PI3K Phosphoinositide 3-kinase PKC Protein kinase C P L A 2 Phospholipase A 2 pNA Para-nitroaniline RNA Ribonucleic acid RNS . Reactive nitrogen species ROS Reactive oxygen species RT-PCR Real time-polymerase chain reaction SDH Succinate dehydrogenase SEM Standard error of means STZ Streptozotocin T1D Type 1 diabetes T2D Type 2 diabetes TBARS Thiobarbituric acid reactive substances TBS-T Tris buffered saline-Tween T C A Trichloroacetic acid T C A cycle Tricarboxylic acid cycle, Kreb's cycle TdT Terminal deoxynucleotidyl transferase T G Triglyceride TNF-a Tumor necrosis factor alpha TUNEL TdT mediated dUTP nick end labeling ZDF Zucker diabetic fatty co-3 PUFA Omega-3 polyunsaturated fatty acids co-6 PUFA Omega-6 polyunsaturated fatty acids xiv A C K N O W L E D G E M E N T S First of all, I would like to thank my parents and my brother, without whose blessings, well wishes and sacrifice, I would not have existed. I would also like to thank God, to have given me the opportunities and strength to fulfill my dream. I would like to thank Dr. Brian Rodrigues for all the guidance, support, and independence he has given me over the entire course of my graduate program. I am also grateful to the members of my supervisory committee for the . valuable suggestions, technical support and encouragement they provided me. I also would like to express thanks to my brilliant colleagues Thomas, Jeff, Girish and Dake for being there for me whenever I needed them. I also would like to acknowledge Howard, Simon and especially, Gloria for being such wonderful summer students. I have been privileged by your company. I would like to express my gratefulness to UBC, Heart and Stroke Foundation and CIHR/Rx&D for the financial support they provided me during my Ph.D studies. Last but not not the least, I would also thank my wife for her continuing support over the years. xv DEDICATION This thesis is dedicated to my family. I miss you and thanks for being there for me. xvi 1. Introduction 1.1. Historical View On 'Diabetes' A disease of "frequent urination" was initially reported in ancient Egypt around 3,500 years ago and was named 'diabetes' by the Greek physician Aretaeus in 100 A.D. Von Mering and Minowski first proposed the role of the pancreas in development of diabetes in 1889. Owing to the seminal discovery of Banting, Best, Collip and McLeod (1921; University of Toronto), it was revealed that diabetes was a consequence of a lack of insulin (Type 1 diabetes), a discovery for which they received the Nobel Prize in 1923. With subsequent advancements in glucose management and an increased longevity, complications of the eyes, heart, blood vessels, kidneys, limbs and nerves also increased. The Diabetes Control and Complications Trial (DCCT) [1] and the U K Prospective Diabetes Study (UKPDS) [2] have demonstrated that lack of adequate control over blood glucose is the main culprit behind these complications, and aggressive therapy with either insulin or oral glucose lowering agents could bring down their incidence in diabetic patients. 1.2. An Overview Of Diabetes Mellitus Diabetes is fast taking the form of a pandemic. An estimated 30 million people had diabetes in 1985. By the year 2000, the figure jumped to 150 million, worldwide. In the next 25 years, this figure has been projected to be as high as 300 million. As a developed nation, diabetes is a major health concern in Canada. Currently, more than two million Canadians have diabetes. More alarming is the fact that in the next few years, another 1 million new cases of diabetes would be diagnosed, taking the total up to 3 million, by 2010 [3]. Even in 1998, the total economic burden of diabetes and its chronic complications in Canada was between US$4.76 1 and $5.23 billion [4]. In the same year, the direct costs for managing just the disease without any associated complications was US $573 million [4]. Diabetes mellitus has been defined by an expert committee [5] as "a group of metabolic diseases characterized by hyperglycemia resulting from defects in insulin secretion, insulin action, or both." Based on clinical etiology, diabetes is classified into two major types: a) Type 1 diabetes (T1D) (previously termed as insulin-dependent diabetes mellitus or IDDM) characterized by a total loss of circulating insulin and therefore an obligatory requirement for exogenous insulin and b) Type 2 diabetes (T2D), caused by a combination of impaired insulin action and an inadequate compensatory insulin secretion. At a very early age, in T1D patients (5-15% of all diabetes cases), the pancreatic p cells are destroyed, leading to a total loss of insulin. Other than unknown factors that cause such a destruction (idiopathic T1D), several autoantibodies may be responsible in eliciting an autoimmune response within pancreatic p cells [6]. In this regard, islet cell auto antibodies (ICAs), insulin auto antibody (IAAs), glutamic acid decarboxylase auto antibodies (GAD 65), and tyrosine phosphatase auto antibodies (IA-2 and IA-2P) have been identified in the serum of T1D patients [6]. T2D patients comprise over 85-95% of all diabetic cases and are usually diagnosed by fasting hyperglycemia and by oral glucose tolerance tests. T2D (previously known as non-insulin-dependent diabetes mellitus or NIDDM) is often asymptomatic and characterized by insulin resistance and relative insulin insufficiency. The development of hyperglycemia is gradual over years after the demonstration of insulin resistance. A positive family history of T2D increases the probability of acquiring this disease by 2.4 fold [7]. Further confirmation of the role of heredity in the development of T2D comes from studying monozygotic twins (with 100%o concordant genes) and dizygotic twins (with 50% concordant genes) where the incidence of both having T2D are 35-58% and only 17-20% respectively [8, 9]. Inactive lifestyle, 2 abdominal fat, and overeating has been also been positively correlated with both obesity and the development of T2D. With advancing disease, as oral hypoglycemic therapy fails, T2D patients may require exogenous insulin, a factor that made the older terms of IDDM and NIDDM, invalid. Unfortunately the epidemiology of diabetes is also changing worldwide. In recent years, there has been a huge upsurge in the cases of obesity, insulin resistance and T2D in both adults and the pediatric population [10, 11]. More advanced detection techniques are identifying low titers of autoantibodies in the adult population with diabetes (latent autoimmune diabetes in adults, or LADA). These cases were previously believed to have T2D, but are now being reclassified as the new 'Type 1.5' diabetes (5-10% of all diabetic cases) [12]. These patients are characterized with a low body mass index like TID patients but with high circulating triglycerides and insulin resistance comparable to classical T2D patients. 1.3. Diabetes is a Cardiovascular Disease In patients with diabetes, cardiovascular disease is the leading cause of death [13]. In fact, the key role that diabetes plays in the development and progression of heart disease has led the American Heart Association to classify diabetic patients as having 'cardiovascular disease' [14]. Diabetes in an otherwise healthy individual increases the risk of death due to cardiac causes to levels comparable to nondiabetic patients with previous myocardial infaction [15]. The increase in cardiovascular events in diabetes is mainly attributed to coronary artery disease (CAD) [16, 17], with fatal and non-fatal C A D events being two- to four-fold higher in these patients [17]. Diabetes also alters the outcome of C A D events, and up to a seven-fold increase in mortality among patients with C A D is observed with co-existing diabetes [18, 19]. Although the majority of these patients are prone to aggressive macrovascular atherosclerotic disease [20], an increased susceptibility to heart failure is also evident [21]. The Framingham Heart 3 Study concluded that heart failure in men (2-fold) and women (5-fold) with diabetes was greater than non-diabetic individuals [22]. This conclusion was verified in subsequent trials [23-25]. CAD does not entirely account for the increased morbidity and mortality from heart failure in diabetes [26]. In 1972, autopsies on four diabetic patients with congestive heart failure and nephropathy demonstrated an absence of coronary atherosclerosis [27]. In other studies with diabetic patients, scarring and interstitial collagen deposition was observed without arterial obstruction in hearts, suggesting the presence of a heart muscle specific disease or 'cardiomyopathy' [28]. In the following years, the notion of a diabetic cardiomyopathy has been widely debated [29] as a simultaneous presence of hypertension and CAD in many diabetic patients makes it difficult to isolate this diabetes specific 'heart-muscle' disease. Nevertheless, an increasing body of evidence supports the existence of metabolic, functional, and structural changes in the diabetic heart, in the absence of any vascular pathology and has been termed as 'diabetic cardiomyopathy' [16, 30, 31]. Cardiomyopathy is defined as diseases of the myocardium associated with cardiac dysfunction [32]. Among the various types of cardiomyopathies, diabetic cardiomyopathy is classified as a 'specific cardiomyopathy' characterized by impaired diastolic function [33]. Inspite of intense research, its etiology remains poorly understood [28]. Defects in the sarcoplasmic reticulum, mitochondria, and/or metabolic pathways seem to be responsible for the development of diabetic 'cardiomyopathy' [16]. Early diastolic changes and a late systolic alteration characterize diabetic cardiomyopathy. These changes can.directly arise from alterations in the regulation of intracellular.cations, which in turn regulate the contractile properties of the heart [28]. In spite of considerable research, conclusive evidence is lacking regarding the exact status of C a + + in the diabetic heart. A depressed Na + /K + ATPase increases intracellular Na + that can decrease C a + + and K + levels in 4 the heart [34]. In addition, the N a + / C a + + exchanger mechanism is impaired together with a decreased myofibrillar C a + + ATPase activity [35]. Decreased C a + + binding to the sarcolemma and decreased C a + + intake in the sarcoplasmic reticulum are characteristics of rodent models of diabetes [28, 35]. On the other hand, C a + + overload may be a defect in the diabetic heart that could be significantly reversed by the administration of verapamil, a C a + + channel blocker [36]. Diabetic hearts also demonstrate reduced sensitivity to norepinephrine and (3-adrenoceptor mediated stimulation [37]. This may result from a decreased number o f receptors or a defect in signaling [38]. Lately, oxidative stress has also been implicated in the etiology of diabetic cardiomyopathy [28, 39] along with other diabetic complications [40]. Oxidative stress arises due to an imbalance between the production of reactive oxygen species (ROS) and' its neutralization by antioxidants. Oxidative damage by R O S and reactive nitrogen species (RNS) have been found in the hearts of diabetic rats [41]. Lately, oxidative stress induced cardiac cell death in the form of both apoptosis and necrosis has also been correlated with diabetic cardiomyopathy, both in human patients and rodent models of diabetes [42-44]. 1.4. Basic Mechanisms Of Oxidative Stress In the first step o f glucose oxidation, intracellular glucose is converted to pyruvate via glycolysis. Glycolysis also generates reduced nicotinamide adenine dinucleotide ( N A D H ) . N A D H can either reduce pyruvate to lactate, or can donate reducing equivalents to the mitochondrial electron transport chain (ETC). Pyruvate derived from glycolysis can also be utilized by the tricarboxylic acid ( T C A ) cycle, to yield carbon dioxide, water, four molecules of N A D H and one molecule of reduced flavin adenine dinucleotide (FADH2). N A D H and FADH2 are utilized for the generation of A T P by the E T C . The E T C is comprised of four inner membrane-associated respiratory complexes (termed as complex I to IV) , cytochrome c and ubiquinone (also known as coenzyme Q). N A D H , generated from glycolysis and T C A cycle 5 donates electrons to complex I, that transfers the electron to ubiquinone. Ubiquinone can also accept electrons from complex II. Electrons from reduced ubiquinone are then diverted, first to complex III and subsequently to complex IV. The final acceptor of the electron is oxygen, which gets converted to water. The proton gradient generated by such a flux of electron across complexes, activates ATP synthase and generates ATP. However the system is not.perfect and even under normal conditions, 1 % of all oxygen consumed by the mitochondria is converted to free radicals [45]. With hyperglycemia, increased levels of electron donors from the T C A cycle in the form of N A D H and F A D H 2 causes an increased efflux of protons outside the mitochondria and leads to increased mitochondrial membrane potential [40]. Augmented proton electrochemical gradient generated by the ETC, leads to transfer of electrons to molecular oxygen to generate superoxide (CV") radicals [40], the precursor for all major reactive oxygen species (ROS). It has been recently shown that overexpression of manganese-superoxide dismutase, (Mn-SOD) which dismutates superoxide in the mitochondria, prevented the three main alterations in diabetes, like polyol pathway induction, protein kinase c activation and increased formation of advanced glycation end products (AGE) in endothelial cells [40, 46]. In the heart, overexpression of Mn-SOD prevents the development of diabetic cardiomyopathy [47]. As superoxide radicals can also be generated de novo via the action of cytosolic enzymes like NADPH oxidase and xanthine oxidase or may leak from the mitochondria itself, the cytoplasmic SOD (Cu + + /Zn + + -SOD) also demonstrates beneficial effects in preventing diabetes induced glomerular injury [48]. Superoxide per se is not a good oxidizing agent due to its short half-life. However, excess superoxide radicals may generate hydrogen peroxide, which in turn may generate hydroxyl radicals (OH") via Fenton chemistry with the help of Fe + + [49]. 6 20 2" + 2H + ^ H 2 0 2 +0 2 Fe + + + H 2 0 2 • • F e + + + + OH " + OH* Hydroxyl radical is a very potent oxidant and causes changes in membrane fluidity and permeability, mitochondrial respiration and various other biochemical functions, leading to cellular dysfunction and death [49]. Thus, to prevent the deleterious effects of hydroxyl radicals, two endogenous antioxidant systems are present to neutralize its precursor, hydrogen peroxide [50]. Firstly, catalase present in the peroxisomes, can neutralize hydrogen peroxide to water and oxygen. Catalase 2H 2 0 2 ^ H 2 0 + 0 2 Secondly, glutathione peroxidase (GPx) can also reduce hydrogen peroxide to water with the help of reduced glutathione (GSH) which gets oxidized to glutathione disulfide (GSSG) in the process [51]. GPx 2GSH + H 2 0 2 ^ G S S G + 2 H 2 0 1.5. Synthesis, Metabolism And Action Of Glutathione (GSH) Due to small amounts of catalase in the heart, GSH, a tripeptide thiol (L-y-glutamyl-L-cysteinyl glycine) remains the most effective antioxidant against hydrogen peroxide [52, 53]. Its primary role is in the protection of cells and tissues against oxidative and nitrosative stress and against reactive electrophiles [54]. It has a low molecular weight and is synthesized de novo from three amino acids- cysteine, glycine and glutamate, by the action of two ATP-consuming enzymes- y-glutamylcysteine synthase (y-GCS) (Reaction a) and GSH synthase (Reaction b) [51]. The y-GCS step is rate limiting for the synthesis of GSH. 7 Reaction a: L-glutamate + L-cysteine + ATP => L-y-:glutamyl-L-cysteine + ADP + Pi Reaction b: L-y-glutamyl-L-cysteine + glycine + ATP => GSH + ADP + Pi As mentioned previously, GSH is oxidized to GSH disulfide (GSSG) by GPx, which catalyzes the reduction of H2O2. Normally, GSSG can be reduced back to GSH in an enzymatic recycling reaction by GSH reductase and NADPH. Adequate NADPH levels and GSH reductase activity maintains a GSH/GSSG ratio of > 1 0 0 : 1 (Figure 1 .1 ) . Thus, under normal levels of ROS generation, there is practically no net loss of GSH. However, if stress levels increase along with an increase in the production of ROS, and/or a fall in NADPH levels, then GSSG may accumulate. Subsequently, the redox status of the cell shifts activating certain redox-sensitive transcription factors like N F - K B [ 5 5 , 5 6 ] . Both GSH and GSSG can also be effluxed out of the cell by transporters and multidrug resistance proteins (MRPs) [ 5 7 , 5 8 ] . GSH extrusion from the cell is irreversibly related to the apoptotic process [ 5 9 ] . In this regard, inhibition of GSH efflux has also been shown to rescue cells from apoptosis and cause them to undergo regeneration [ 6 0 ] . In the heart, during reperfusion after ischemia, the heart effluxes high levels of GSSG, which persists long after the generation of ROS has ceased [ 6 1 ] . Such a loss in the GSH pool may be prevented by preadministration of N-acetyl cysteine (NAC), a source of cysteine, which increases the cardiac GSH, and is beneficial in preventing changes in the ischemic-reperfused hearts [ 6 2 , 6 3 ] . In recent years, the importance of mitochondrial GSH in preventing oxidative stress has come to light [ 6 4 - 6 6 ] . Being the major site of oxygen consumption, mitochondria are also the primary sites for the generation of ROS. Thus, although most of the cellular GSH is in the cytoplasm, a distinctly regulated pool is present in mitochondria. As mitochondria lack y-GCS mRNA, it depends exclusively on the transport of cytosolic GSH into the matrix, and accounts for 1 0 - 1 5 % of the total GSH pool [ 6 4 ] . Being negatively charged, GSH is actively transported 8 and conserved into the negatively charged mitochondria, and resists depletion even under oxidative stress [67] (Figure 1.1). It has been observed that antimycin A, a drug that increases ROS is more cytotoxic towards cells with GSH depleted in the mitochondria compared to cytosolic GSH depletion alone [68]. Further, treatment with buthionine sulfoximine (BSO), an inhibitor of y-GCS, decreases 90% of the intracellular GSH but is non-toxic in most cells. This may be due to the inability of BSO to deplete mitochondrial GSH, as under cytosolic GSH depletion, the mitochondria actively accumulates the remaining GSH within the matrix as a protective mechanism [69]. However, diethyl maleate (DEM), which can permeate into the mitochondria and bind irreversibly to all preformed GSH is more cytotoxic [69]. Interestingly, treatment with D E M can in turn trigger y-GCS activity [70]. Thus, GSH levels return to normal levels within 8-12 hours after D E M administration in vivo [71]. However, co administration of BSO and GSH in vivo has been shown to achieve chronic GSH depletion in the liver [72]. Other than its redox functions, GSH can form thioester or thiolester conjugates of different compounds including xenobiotics with the help of glutathione-S-transferases to promote their detoxification and elimination in Phase II metabolism [73]. This pathway promotes biliary excretion, and interorgan transport for final renal excretion via the mercapturate pathway. GSH has also a prominent role in signal transduction and gene expression. Protein glutathionylation of enzymes such as phosphorylase, creatine kinase, RAS, carbonic anhydrase control their intracellular activity [73]. The formation of disulfide bonds with the help of GSH has also been suggested to dictate protein-folding, and enzyme action. S-nitrosoglutathione (GS-NO) has been suggested as a storage and transfer form of nitric oxide[74]. Presence of GSH also helps to maintain the cysteine groups in the promoter regions of genes like N F - K P and AP-1, and allow their expression and biological activity [75]. 9 1.6 Figures Mitochondria Cytosol Fig.1.1 Pathways of intracellular glutathione (GSH) metabolism Extracellular cystine gets reduced intracellularly to cysteine and is utilized by y-glutamylcysteine synthase to produce y-glutamylcysteine. This step is the rate-limiting pathway for de novo GSH biosynthesis and can be blocked by buthionine sulfoximine (BSO). In the next step, y-glutamylcysteine is utilized by glutathione synthase to produce GSH (y-glutamyl-L-cysteinyl glycine). GSH can then either be transported to the mitochondria or remain in the cytosol, where they are utilized by GSH peroxidase to neutralize hydrogen peroxide (H2O2), and gets converted to oxidized glutathione (GSSG). GSH reductase present within both mitochondria and cytosol can recycle GSSG to 'active' GSH with the help of NADPH. 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Biochem Pharmacol 1992;43:2209-15 73. Sies H. Glutathione and its role in cellular functions. Free Radic Biol Med 1999; 27:916-21 74. Ji Y, Akerboom TP, Sies H, Thomas JA. S-nitrosylation and S-glutathiolation of protein sulfhydryls by S-nitroso glutathione. Arch Biochem Biophys 1999; 362:67-78 75. Arrigo AP. Gene expression and the thiol redox state. Free Radic Biol Med 1999; 27:936-44 18 2. Cardiac Cell Death In Early Diabetes 2.1 Hyperglycemia And Cellular Lipids: Common Pathways To Oxidative Stress ROS generation in the diabetic heart is a combined effect of both hyperglycemia and hyperlipidemia. Glucose oxidation generates. N A D H and F A D H 2 in the electron transport chain (ETC) of the mitochondria, to produce energy [1]. Increased proton electrochemical gradient generated by the ETC leads to transfer of electrons to molecular oxygen to generate superoxide radicals and hydrogen peroxide [2]. Hence, with onset of hyperglycemia, glucose auto-oxidation is increased [3]. Glucose auto oxidation involves the process of formation of glyoxal via the process of degradation by retroaldol condensation reactions [4]. Concurrent oxidative modifications by hydrogen peroxide also can convert glucose to glucosone and glycoaldehyde to glyoxal [4, 5]. Glyoxal may be later converted to 3-deoxyglucone and methyl glyoxal and contribute to the formation of early glycation products and oxidative stress. Free radical damage of the mitochondrial membrane can precipitate a further rise in ROS generation in the heart (ROS induced ROS release) [6]. Augmented superoxides can also increase diacylglycerol synthesis or phosphatidylcholine hydrolysis, thereby activating PKC [7]. Hyperglycemia also provokes excessive production of A G E [2]. In this process, glucose reacts non-enzymatically with amino acids in proteins to form early glycation products (Schiff bases) and intermediate glycation products (Amadori bases). These compounds are then degraded to AGEs like glyoxal, methylglyoxal, and carboxymethylysine [8]. Binding of AGEs to their receptors leads to alterations in cell signaling, and further production of free radicals [9]. Finally, hyperglycemia activates the polyol pathway, which converts excess glucose to sorbitol via aldose reductase. Increased activation of aldose reductase leads to rapid depletion of NADPH, its cofactor [10]. As NADPH is also required for the activity of 19 glutathione reductase for the regeneration of active glutathione (GSH) from its 'oxidized form (GSSG), intracellular levels of this critical antioxidant decreases rapidly in diabetes [2]. Depletion of GSH allows oxidants to go unchecked, causing irreversible cellular damage. In addition to hyperglycemia, diabetes is also characterized by dyslipidemia and 'lipid overload' in non-adipose tissues like the heart [11]. Independent of hyperglycemia, excess lipids can also precipitate oxidative stress, as observed in the Zucker diabetic fatty rat (ZDF), a model of Type 2 diabetes [12, 13]. Fatty acids (FA) can directly impair mitochondrial function via uncoupling of oxidative phosphorylation [14]. During uncoupling, electron transfer through E T C is blocked, resulting in buildup of a voltage gradient across the mitochondrial membrane. As a result, excess electrons are transferred to molecular oxygen to form ROS like-superoxide [14, 15]. Due to its short half-life, superoxide per se is not a good oxidizing agent. However, excess superoxide radicals may generate hydrogen peroxide, that in turn may generate cytotoxic hydroxyl radicals (OH") with the help of iron (Fe+ +) [16]. Recently, FA were also reported to transport iron to the mitochondria, thus further strengthening oxidative stress [17]. Although F A can also deplete intracellular GSH levels [18, 19], and activate PKC [20], the de novo production of ceramide, a proapoptotic sphingolipid appears to be the most damaging of all non-oxidative consequences of excess FA in the heart. Ceramides interact with cardiac mitochondria to induce the release of free radicals, that extensively damage cellular membranes and proteins [21, 22]. Overall, hyperglycemia and lipid overload are currently believed to be responsible for oxidative stress (Figure 2.1) and the development of heart muscle disease in diabetes. 2.2 The Relevance Of Early Diabetes Till recent years, a number of clinical studies have demonstrated the incidence of heart disease only after chronic diabetes. In patients with Type 1 diabetes, heart disease is observed at least 7 years following the diagnosis of diabetes [23]. In Type 2 patients, the time course for the 2 0 development of heart disease has been difficult to predict. This is largely a consequence of under diagnosis of about 50% of the diabetic population [24]. Additionally, as obesity, hypertension and insulin resistance are beset with their own cardiovascular problems, the causal effect of hyperglycemia on heart disease in Type 2 diabetes is unresolved. In experimental studies, most of the existing literature on diabetic cardiomyopathy suggest the development of cardiac pathology only after chronic diabetes [25-27]. In recent years, with the help of advanced imaging techniques, an increasing prevalence of heart disease is being recognized in otherwise healthy diabetic patients. While older studies reported around 30% of Type 2 diabetic patients as having diastolic dysfunction based on conventional electrocardiography, more rigorous techniques like Doppler imaging revealed uptc 60% of these subjects as having diastolic failure [28]. This early detection of heart disease points to fundamental biochemical changes during acute diabetes as the likely initiator of cardiovascular disease. In animal models, there is a growing recognition that cardiac dysfunction can occur early following induction of diabetes [29], and slower myocyte shortening and relengthening were observed within 4-6 days of diabetes [30]. In fact, high glucose perfusion for 2 hours increased peroxynitrite and superoxide levels, leading to cell death in working rat hearts, effects that were prevented by the addition of GSH to the perfusion buffer [31]. In patients, Dr. Ceriello and his group have hypothesized that acute hyperglycemia, as observed following a meal or oral glucose loading, in an otherwise 'well-controlled' diabetic individual, leads to 'postprandial hyperglycemic spikes'. These acute surges in plasma glucose precipitated a variety of cardiovascular stress signals including impaired ventricular function, arrhythmias, and accumulation of pro-thrombotic factors [32, 33]. The role of oxidative stress in these disorders was supported by direct measurement of oxidative markers or the ameliorative effect of antioxidants [33, 34]. 21 Early myocardial functional changes in diabetes are also related to acute changes in cardiac lipids. In our studies, following streptozotocin (STZ, a pancreatic P-cell toxin) injection, hypoinsulinemia and hyperglycemia ensues within 24 hours [35]. At this time point, cardiac glucose oxidation decreases, and cardiac energy provision is almost exclusively via P -oxidation of fatty acids [36]. To guarantee FA supply to the diabetic heart, lipoprotein lipase (LPL) activity at the coronary lumen is rapidly elevated within hours [37], and by hydrolyzing circulating lipoproteins, provides an augmented supply of FA to the heart [36, 38]. However, such adaptive mechanisms following acute hyperglycemia could eventually become counterproductive, and lead to 'lipotoxicity'. Increased F A oxidation can promote acidosis and generation of free radicals, making the diabetic heart more prone to injury [39]. As described, non-oxidative metabolism of FA can rapidly induce ceramide and nitric oxide formation, damage the mitochondrial membrane, and induce oxidative stress [40, 41]. Such diverse cytotoxic effects of FA on the mitochondria could also lead to respiratory chain malfunction and a lower cardiac efficiency within 2 weeks of diabetes [42]. In vitro studies have also demonstrated that addition of excess FA to cardiomyocytes can precipitate cell death via apoptosis within 16-24 hours [40, 41]. 2.3 Basic Mechanisms Of Cell Death Characteristics of the diabetic heart include myocardial hypertrophy and interstitial fibrosis [43]. It has been proposed that this hypertrophy may be an insufficient compensatory response to increased cell death in the diabetic heart. Indeed, hearts from patients with Type 2 diabetes are characterized by a 8 5-fold increase in apoptosis, and a 4-fold increase in necrosis [43]. Apoptosis is an energy dependent process of cell death characterized by nuclear condensation and formation of small apoptotic bodies that are engulfed by phagocytic cells (macrophages) without any rupture of the cell membrane [44, 45]. The molecular mechanisms 22 behind apoptotic cell death in the heart have only recently become apparent, and caspases have been proposed to be the 'central executioners' of this cellular demise (Figure 2.2). Caspases are a family of cysteine containing proteases that get activated following an apoptotic insult. Subsequently, caspases disintegrate intracellular proteins, cause nuclear DNA fragmentation and cell death [44]. Two major pathways exist to activate caspases from inactive procaspases in the cytosol. The first involves activation of downstream effector caspases like caspase-3 via 'death' receptor mediated activation of upstream caspases like caspase-8 ('extrinsic' pathway) [45]. However this pathway is likely not important in causing cell death in the acutely diabetic heart [46]. The second pathway known as the 'intrinsic' or the mitochondrial pathway of apoptosis [45], initially involves a loss of the mitochondrial membrane potential. This signals the translocation of pro-apoptotic Bcl-2 family members like Bax to the mitochondria, and the release of cytochrome c from the mitochondrial intramembrane space into the cytosol. As a protective mechanism, antiapoptotic Bcl-2 family members like Bcl-2 and B C 1 - X L interact with the proapoptotic members to prevent this release of cytochrome c. Cytochrome c once released, forms an 'apoptosome', and activates caspase-9, Caspase-9 initiates the caspase cascade, terminating in the activation of caspase-3, nuclear DNA damage and apoptotic demise [45]. Lately, it has been proposed that cardiac apoptosis may not proceed towards DNA degradation and. cell death ('apoptosis interruptus') but may lead to the disintegration of myofibrils such that the contractile properties are lost ('zombie' myocytes) [47]. As caspase activation and subsequent proteolysis in apoptosis requires energy, adequate ATP levels are required. Thus, when ATP stores are depleted, the cells may be directed towards necrosis, by the same initiating steps [48, 49]. Necrosis or 'accidental' cell death is an energy independent process characterized by cell membrane rupture [48]. Interestingly, at the earliest stage of diabetes, necrotic cell death is absent in rodent hearts [50, 51], and only appears after 15 days of hyperglycemia [52]. At this time, necrosis is temporally related to mitochondrial 23 dysfunction and ATP loss [53]. In chronic diabetes, cardiac necrosis is a common feature and is observed both in humans and animal models of the disease [26, 43]. Both apoptosis and necrosis in the diabetic heart is believed to be a result of oxidative stress. In this regard, ROS such as hydrogen peroxide (H2O2) and reactive nitrogen species (RNS) like peroxynitrite (ONOO") can induce membrane lipid peroxidation, alteration of cellular proteins, and initiation of diverse stress-signaling pathways, including cell death [54]. 2.4 Oxidant Induced Cell Death In Experimental Diabetes Development of Type 2 diabetes is a gradual process, with no clear demarcation between the 'pre-diabetic' and the hyperglycemic phase. Thus, most of the existing work on early diabetes and cell death is performed in insulin deficient STZ-diabetic rats. In these models, we and others have demonstrated that cardiomyocyte death proceeds via an apoptotic pathway, with an increase in caspase-3 activity and oxidative stress [35, 50, 51]. In vitro, exposing myoblast H9c2 cells to high glucose induced cytochrome c release into the cytosol followed by caspase-3 activation and apoptosis [55]. Increased prevalence of ROS can oxidatively damage DNA and increase p53. This transcription is factor responsible for increased production of angiotensinogen and ultimately angiotensin II [50, 56]. Binding of angiotensin II to its receptors promotes generation of free radicals like superoxide and peroxynitrite in the heart [50, 51]. Interestingly, both the angiotensin II blocker like losartan and IGF-1 diminishes oxidative stress and prevents cardiac apoptosis in diabetes [57]. Recently, taurine, an inhibitor of the angiotensin receptor also demonstrated a similar effect [58]. With hyperglycemia, upregulation of PKC5 can also precipitate cardiac apoptosis. PKC5 may exert its cytotoxicity either through enhanced generation of ROS or reduction of nuclear factor-sB, a survival signal against apoptosis [59]. Endothelin, a potent vasoconstrictor, can also activate PKC and cause cardiac apoptosis during diabetes [60]. The cellular content of cardiac GSH, a crucial 24 endogenous antioxidant is also reduced following hyperglycemia in vitro [61] and exogenous supplementation with GSH can reverse cardiac apoptosis.following diabetes [31]. In addition to GSH, metallothionein (MT), a potent antioxidant [62] has also been shown to decrease cardiac apoptosis. Additionally, administration of M T prevents hyperglycemia induced myocardial cell death and preserves cardiac function [63] whereas increasing biosynthesis of M T through exogenous administration of zinc also prevents the development of diabetic cardiomyopathy in rats [64]. Being a terminally differentiated organ with low regeneration capacity, even a low rate of cardiac cell death (ranging from 0.08-0.25% of total myocytes) over the long term could deplete a significant number cardiomyocytes, the units of cardiac contractility [65]. Thus, for the prevention of long-term cardiac problems in diabetes, early recognition and intervention of the deleterious pathways of cell death are required. 25 2.5 Figures Reactive Oxygen Species (ROS) t t LIPIDS PROTEINS DNA SUGARS Lipid Peroxidation Protein carbonyl Nitration of proteins I DNA oxidation 1 Schiff bases I C I I I I Membrane Damage \ Cell Membrane Mitochondria ^ 1) Loss of ion channel function 2 ) Increased 'leakiness' 1) Loss of enzyme function 2) Activation of deleterious pathways Nucleus/Mitochondria 1) Loss of membrane potential 2 ) Release of cytochrome c 3) more ROS \ 1) Altered gene expression 2) PolyADP ribosylation 3) Depletion of ATP Amadori products ^ r Advanced glycation end products Fig. 2.1 Mechanisms of ROS induced injury in diabetes Both hyperglycemia and hyperlipidemia can increase ROS production in the diabetic heart. Once ROS is produced, it can have multiple effects on different biomolecules like lipids, proteins, DNA and sugars. The major pathways of injury are depicted in this schematic. 26 Fig. 2.2 Overview of the basic mechanisms of apoptosis Caspases, the most important enzymes in the apoptotic pathway, can be initiated by either of two mechanisms. The first involves activation of downstream effector caspases like caspase-3 via 'death' receptor mediated activation of upstream caspases like caspase-8 ('extrinsic' pathway). However this pathway probably plays a minor role in causing cell death in the acutely diabetic heart. The second pathway known as the 'intrinsic' or the mitochondrial pathway of apoptosis initially involves a loss of the mitochondrial membrane potential, followed by the release of mitochondrial cytochrome c. The release of cytochrome c depends on the relative levels of pro-apoptotic and anti-apoptotic Bcl-2 family members on the mitochondrial membrane. Cytochrome c once released, forms an 'apoptosome', with Apaf-1 and pro-caspase 9, and activates caspase-9. Both caspases 8 and 9 can lead to the caspase cascade, terminating in the activation of caspase-3, nuclear DNA damage and apoptosis. 27 2.6 References 1. Bonnefont-Rousselot D. Glucose and reactive oxygen species. Curr Opin Clin Nutr Metab Care 2002; 5:561-8 2. Brownlee M . Biochemistry and molecular cell biology of diabetic complications. Nature 2001; 414:813-20 3. Wells-Knecht KJ, Zyzak DV, Litchfield JE, Thorpe SR, Baynes JW. Mechanism of autoxidative glycosylation: identification of glyoxal and arabinose as intermediates in the autoxidative modification of proteins by glucose. Biochemistry 1995; 34:3702-9 4. Thornalley PJ, Langborg A, Minhas HS. Formation of glyoxal, methylglyoxal and 3-deoxyglucosone in the glycation of proteins by glucose. Biochem J 1999; 344 Pt 1:109-16 5. Thornalley P, Wolff S, Crabbe J, Stern A. The autoxidation of glyceraldehyde and other simple monosaccharides under physiological conditions catalysed by buffer ions. Biochim Biophys Acta 1984; 797:276-87 6. 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Hennig B, Meerarani P, Ramadass P, Watkins BA, Toborek M . Fatty acid-mediated activation of vascular endothelial cells. Metabolism 2000; 49:1006-13 19. Toborek M , Hennig B. Fatty acid-mediated effects on the glutathione redox cycle in cultured endothelial cells. Am J ClinNutr 1994; 59:60-5 20. Griffin M E , Marcucci MJ, Cline GW, Bell K, Barucci N, Lee D, et al. Free fatty acid-induced insulin resistance is associated with activation of protein kinase C theta and alterations in the insulin signaling cascade. Diabetes 1999; 48:1270-4 29 Garcia-Ruiz C, Colell A, Mari M , Morales A, Fernandez-Checa JC. Direct effect of ceramide on the mitochondrial electron transport chain leads to generation of reactive oxygen species. Role of mitochondrial glutathione. J Biol Chem 1997; 272:11369-77 ' Di Paola M , Cocco T, Lorusso M . Ceramide interaction with the respiratory chain of heart mitochondria. Biochemistry 2000; 39:6660-8 Lehto S, Ronnemaa T, Pyorala K, Laakso M . 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Ceriello A. Postprandial hyperglycemia and diabetes complications: is it time to treat? Diabetes 2005; 54:1-7 34. Ceriello A, Hanefeld M , Leiter L, Monnier L, Moses A, Owens D, et al. Postprandial glucose regulation and diabetic complications. Arch Intern Med 2004; 164:2090-5 35. Ghosh S, An D, Pulinilkunnil T, Qi D, Lau HC, Abraham A, et al. Role of dietary fatty acids and acute hyperglycemia in modulating cardiac cell death. Nutrition 2004; 20:916-23 36. 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 37. 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 38. Pulinilkunnil T, Rodrigues B. Cardiac lipoprotein lipase: Metabolic basis for diabetic heart disease. Cardiovasc Res 2005; 31 39. Liu Q, Docherty JC, Rendell JC, Clanachan AS, Lopaschuk GD. High levels of fatty acids delay the recovery of intracellular pH and cardiac efficiency in post-ischemic hearts by inhibiting glucose oxidation. J Am Coll Cardiol 2002; 39:718-25 40. Sparagna GC, Hickson-Bick DL, Buja L M , McMillin JB. A metabolic role for mitochondria in palmitate-induced cardiac myocyte apoptosis. Am J Physiol Heart Circ Physiol 2000; 279:H2124-32 41. Rabkin SW, Huber M , Krystal G. Modulation of palmitate-induced cardiomyocyte cell death by interventions that alter intracellular calcium. Prostaglandins Leukot Essent Fatty Acids 1999; 61:195-201 42. Wang P, Lloyd SG, Zeng H, Bonen A, Chatham JC. Impact of altered substrate utilization on cardiac function in isolated hearts,from Zucker diabetic fatty rats. Am J Physiol Heart Circ Physiol 2005; 288:H2102-10 43. Frustaci A, Kajstura J, Chimenti C, Jakoniuk I, Leri A, Maseri A, et al. Myocardial cell death in human diabetes. Circ Res 2000; 87:1123-32 44. Bennett MR. Apoptosis in the cardiovascular system. Heart 2002; 87:480-7. 45. Clerk A, Cole SM, Cullingford TE, Harrison JG, Jormakka M , Valks D M . Regulation of cardiac myocyte cell death. Pharmacol Ther 2003; 97:223-61 46. Cai L, Kang YJ. Cell death and diabetic cardiomyopathy. Cardiovasc Toxicol 2003; 3:219-28 47. Communal C, Sumandea M , de Tombe P, Narula J, Solaro RJ, Hajjar RJ. Functional consequences of caspase activation in cardiac myocytes. Proc Natl Acad Sci U S A 2002; 99:6252-6 48. Proskuryakov SY, Konoplyannikov A G , Gabai V L . Necrosis: a specific form of programmed cell death? Exp Cell Res 2003; 283:1-16 32 Tatsumi T, Shiraishi J, Keira N, Akashi K, Mano A, Yamanaka S, et al. Intracellular ATP is required for mitochondrial apoptotic pathways in isolated hypoxic rat cardiac myocytes. Cardiovasc Res 2003; 59:428-40 Fiordaliso F, Leri A, Cesselli D, Limana F, Safai B, Nadal-Ginard B, et al. Hyperglycemia activates p53 and p53-regulated genes leading to myocyte cell death. Diabetes 2001; 50:2363-75 Fiordaliso F, Li B, Latini R, Sonnenblick EH, Anversa P, Leri A, et al. Myocyte death in streptozotocin-induced diabetes in rats in angiotensin II- dependent. Lab Invest 2000; 80:513-27 Cai L, Wang J, Li Y, Sun X, Wang L, Zhou Z, et al. Inhibition of superoxide generation and associated nitrosative damage is involved in metallothionein prevention of diabetic cardiomyopathy. Diabetes 2005; 54:1829-37 How OJ, Aasum E, Severson DL, Chan WY, Essop MF, Larsen TS. Increased myocardial oxygen consumption reduces cardiac efficiency in diabetic mice. Diabetes 2006;55:466-73 Cai L , Kang YJ. Oxidative stress and diabetic cardiomyopathy: a brief review. Cardiovasc Toxicol 2001; 1:181 -93 Cai L , Li W, Wang G, Guo L, Jiang Y, Kang YJ. Hyperglycemia-induced apoptosis in mouse myocardium: mitochondrial cytochrome C-mediated caspase-3 activation pathway. Diabetes 2002; 51:1938-48 Monkemann H, De Vriese AS, Blom HJ, Kluijtmans L A , Heil SG, Schild HH, et al. Early molecular events in the development of the diabetic cardiomyopathy. Amino Acids 2002; 23:331-6 .33 Kajstura J, Fiordaliso F, Andreoli A M , Li B, Chimenti S, Medow MS, et al. IGF-1 overexpression inhibits the development of diabetic cardiomyopathy and angiotensin II-mediated oxidative stress. Diabetes 2001; 50:1414-24 Li C, Cao L, Zeng Q, Liu X, Zhang Y, Dai T, et al. Taurine may prevent diabetic rats from developing cardiomyopathy also by downregulating angiotensin II type2 receptor expression. Cardiovasc Drugs Ther 2005; 19:105-12 Shizukuda Y, Reyland M E , Buttrick PM. Protein kinase C-delta modulates apoptosis induced by hyperglycemia in adult ventricular myocytes. Am J Physiol Heart Circ Physiol 2002; 282:H1625-34 Chen S, Evans T, Mukherjee K, Karmazyn M , Chakrabarti S. Diabetes-induced myocardial structural changes: role of endothelin-1 and its receptors. J Mol Cell Cardiol 2000; 32:1621-9 Fiordaliso F, Bianchi R, Staszewsky L, Cuccovillo I, Doni M , Laragione T, et al. Antioxidant treatment attenuates hyperglycemia-induced cardiomyocyte death in rats. J Mol Cell Cardiol 2004; 37:959-68 Cai L, Klein JB, Kang YJ. Metallothionein inhibits peroxynitrite-induced DNA and. lipoprotein damage. J Biol Chem 2000; 275:38957-60 Song Y, Wang J, Li Y, Du Y, Arteel GE, Saari JT, et al. Cardiac metallothionein synthesis in streptozotocin-induced diabetic mice, and its protection against diabetes-induced cardiac injury. Am J Pathol 2005; 167:17-26 Wang J, Song Y, Elsherif L, Song Z, Zhou G, Prabhu SD, et al. Cardiac metallothionein induction plays the major role in the prevention of diabetic cardiomyopathy by zinc supplementation. Circulation 2006; 113:544-54 Kajstura J, Bolli R, Sonnenblick EH, Anversa P, Leri A. Cause of death: suicide. J Mol Cell Cardiol 2006; 40:425-437 34 3. Effect Of Fatty Acids On Cardiac Cell Death In Diabetes Although lipids have been acknowledged as a significant contributor to diabetic cardiac complications, most of the existing studies on the cytotoxicity of fatty acids are on cell systems like endothelial and smooth muscle cells [1-5]. It has been proposed that as these vascular cells are directly exposed to plasma fatty acids, deleterious effects on the heart are primarily mediated through this vascular component. However, in diabetes, with an increased influx of fatty acids for cardiac energy needs, the same fatty acids that injure endothelial and smooth muscle cells, also enter the cardiomyocytes and cause damage. One weakness with the traditional approach of examining fatty acid cytotoxicity has been the tendency to group ' fatty acids' as a single entity, with little attempt made to isolate the effects of individual fatty acid classes. As the cellular fate of each class of fatty acids is different with regard to their metabolism, special attention is required when assessing the effects of fatty acids on cell death. 3.1 Saturated fatty acids Both humans and chronic models of diabetes demonstrate cardiac lipid overload, suggested to cause 'lipotoxicity' [6]. . The causal effect of lipotoxicity in diabetic heart disease was strengthened in studies using Zucker diabetic fatty rats (ZDF) rats, a model of Type 2 diabetes. In these studies, amelioration of cardiac lipids by troglitazone decreased formation of ceramide, a proapoptotic sphingolipid, and reversed cardiac dysfunction [7, 8]. The first step in de novo ceramide biosynthesis is catalyzed by serine palmitoyl transferase, and involves the condensation of palmitoyl CoA and serine [9]. As the saturated fatty acid, palmitic acid (CI6:0) is the precursor of palmitoyl CoA, most studies demonstrating detrimental effects of F A on myocytes have hence focused on using saturated fatty acids. 35 Like glucose and FA, ceramides also release cytotoxic free radicals and cytochrome c from the mitochondria, thus provoking apoptotic cell death [10]. Increased ceramide levels also upregulates the expression of inducible nitric oxide synthase (iNOS) by activating the transcription factor N F K B [9]. Increase in iNOS expression leads to excess production of nitric oxide (NO), which can combine with superoxide to produce the highly damaging RNS, peroxynitrite (ONOO~) [11]. As hyperglycemia can independently augment cardiac NO production [12], a rise in ceramide levels can cause additive damage to cardiomyocytes, and a further increase in cardiac apoptosis [6]. One drawback with the conventional approach of examining lipotoxic effects on the heart is that most studies have used in vitro incubations with single fatty acids. Although this setting partly resembles fatty acid derived from plasma, it does not account for fatty acid obtained from other physiologically relevant sources like hydrolysis of lipoproteins by cardiac lipoprotein lipase (LPL). This is particularly important as: i) LPL increases in the diabetic heart [13, 14], ii) the molar concentrations of fatty acids in lipoproteins is approximately 10-fold higher than that of fatty acids bound to albumin, iii) LPL has a selective affinity towards palmitic acid containing lipoproteins (47.5% of total fatty acids released) [15] and iv) LPL derived palmitic acid causes a 8-fold greater cell death, than when it is used individually [1]. Thus, a true measure of the effects of palmitic acid on apoptosis should include an examination of its in vivo effects, and few studies have focused on the role of dietary fat on in vivo cardiac apoptosis. In a recent study, we demonstrated that feeding 20% w/w palm oil (40% energy, rich in palmitic acid, PO) increased cardiac apoptosis and oxidative stress under normoglycemic conditions [16]. Superimposition of diabetes by STZ induced a further increase in the 36 cardiac apoptotic index, likely due to the additive effects of hyperglycemia. However, only a fraction (0.02-0.03%) of total cardiomyocytes underwent apoptosis in PO fed diabetic hearts. In a recent study by Okere et al, a high fat diet rich in saturated fatty acids (60% by calorie) promoted ceramide levels and caused apoptosis, again in only 0.03%) of total cardiomyocytes [17]. This is in marked contrast to in vitro studies with palmitic acid, where up to 30%o of the total cell population perished [18, 19]. It is likely that under in vitro conditions, energy requirement is reduced in isolated, non-beating; and quiescent cardiomyocytes in culture. Thus, exogenous fatty acids may undergo a greater flux towards non-oxidative pathways, provoking an exaggerated apoptotic response. It is also possible that in vivo, phagocytes rapidly clear apoptotic cells (within 24 h), and thus determination of apoptosis in the intact heart is underestimated [20]. As palmitic acid generates RNS like peroxynitrite, this highly reactive anion can induce nitration of tyrosine residues of proteins, leading to the formation of nitrotyrosine [11]. Protein expression of nitrotyrosine increased in PO fed diabetic hearts [16]. The apoptotic pathway in the heart is controlled by a number of proteins of the Bcl-2 gene family that can either promote (e.g., Bax) or prevent (e.g., Bcl-2) apoptosis [21, 22]. Upon apoptotic stimuli, oxidative stress increases Bax activity. Other members of the same family like Bcl-2 can heterodimerize with Bax, and prevent apoptosis. Thus, estimation of Bax/Bcl-2 ratio has been used to detect pro- or anti-apoptotic environments [21]. PO feeding augmented Bax mRNA in the heart. Superimposition of diabetes in PO fed hearts further increased Bax expression, but decreased mRNA levels of the protective Bcl-2, leading to an increased Bax/Bcl-2 ratio. An augmented Bax/Bcl-2 mRNA ratio has been previously linked to an increased lipoapoptosis in pancreatic islets of Zucker Diabetic Fatty (ZDF) rats [21]. 37 3.2 Monounsaturated fatty acids Monounsaturated fatty acids (MUFA) like palmitoleic acid (CI6:1) and oleic acid (CI8:1) are biosynthesized from saturated fatty acids like palmitic and stearic acid respectively [23]. Traditionally, M U F A has been used as a control to demonstrate the higher cytotoxicity of palmitic acid. In a study on rat pancreatic P cells, palmitoleic acid prevented palmitate induced cytochrome c release, DNA fragmentation and apoptosis [24]. In human pancreatic P cells, palmitic acid decreased the expression of antiapoptotic factor Bcl-2, both under normo and hyperglycemia [25]. In the same study, co incubation with palmitoleic and oleic acids conferred protection by increasing the expression of Bcl-2 [25]. In cardiomyocytes, oleic acid also inhibits the deleterious effects of palmitic acid and prevents or delays palmitate-induced apoptosis [26, 27]. The molecular mechanism behind the antiapoptotic potential of monounsaturated fatty acids comes from studies in the Chinese hamster ovary (CHO) cells [28]. Co-supplementation with 200pM oleate, prevents palmitate-induced apoptosis in CHO cells, by either preventing the formation or scavenging of excess ceramide and free radical generation. As intracellular fatty acids in the free form can promote the formation of both these detrimental molecules, monounsaturated fatty acids like palmitoleic acid or oleic acid have the unique ability of promoting palmitate incorporation into inert triglycerides and rescuing palmitate-induced apoptosis [25]. Interestingly, oleic acid, as well as palmitic acid, is toxic in cells with impaired triglyceride synthesis [29]. Thus, in the setting of a mismatch between lipid entry and utilization, oleic acid may facilitate triglyceride accumulation in non-adipocytes as the first line of defense against lipotoxicity [29]. In addition to its effects on triglyceride storage, feeding diets rich in oleic acid (42.7-43.9% of all fatty acids) for 30 days increases the activity of critical antioxidants like 38 catalase, superoxide dismutase and glutathione peroxidase [30]. This diet was also shown to be protective against loss of myocardial fibers following ischemia-reperfusion (IR) injury. As loss of myofibers may be a result of extensive cell death, increasing endogenous cardiac antioxidants may be a beneficial feature of a monounsaturated diet [30]. Alternative mechanisms explaining the beneficial effect of oleic acid against palmitate-induced apoptosis may be its involvement in the upregulation of phosphoinositide 3-kinase (PI3K) [31], an antiapoptotic factor for most cells. PI3K activation enhances cell survival and antagonizes apoptosis via protein kinase PKB in many cell types including cardiomyocytes [32]. As hypoinsulinemia in diabetes leads to a downregulation of PI3K activation, accumulation of oleic acid may indeed play an important role in the prevention of diabetic cardiac cell death [33,34]. 3.3 The Western Diet Although majority of the work in the area of lipotoxicity, oxidative stress and diabetes has been attempted with, saturated and monounsaturated fatty acids, the current western diet is richer in polyunsaturated fatty acids than either of the two previous classes of fatty acids [35-37]. As obesity and its attendant complications like diabetes and hypertension continues to grow, increased awareness among the population has led to an indiscriminate substitution of atherogenic saturated cooking fats with vegetable oils containing co-6 PUFA [35]. Unfortunately, most of the coronary vascular benefits have been attributed to co-3 PUFA like eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), and not co-6 PUFA. As the Western diet is scarce in fish oils, the richest source of the beneficial co-3 PUFA, dependence on the other kind of PUFA (i.e. co-6 PUFA) has increased. In addition to the intentional modification of dietary fats, farmed animals are fed grains and seeds containing high amount 39 of co-6 PUFA [35]. Consumption of these animals (and their products) has led to an even greater increase in the dietary co-6 PUFA. Thus, although the total amount of dietary fat has not changed significantly, co-6 PUFA has increased up to 5-10% of the total energy intake [38]. In the United Kingdom, the daily consumption of linoleic acid has gone up from 11 g/day in 1970 to 14 g/day in 1990 [39]. In summary, a high dietary intake of co-6 PUFA has augmented our co-3: co-6 ratio to between 15:1 to 20:1, whereas during our evolution, the ratio was 1:1 or even less [40]. In our current society, the traditional diet of Greece (Crete) closely approximates the Paleolithic diet and is linked with long life expectancy and low cardiovascular disease. The traditional Greek diet contains a sizeable consumption of fish (approximately 30 times that of US population), a rich source of co-3 PUFA, as well as a large amount of vegetables (including wild plants), fruits, nuts and legumes, all rich sources of folate, calcium, GSH, antioxidants, vitamins E and C, and minerals [41]. Olive oil (rich in oleic acid, a MUFA) is the chief cooking medium. In addition, because the meat comes from animals that grazed rather than being fed grain, it contains (n-3) polyunsaturated fatty acids (PUFA) as do milk and milk products [41]. Clinical studies like the Lyon Heart study examined the effects of a modified Greek diet with a ratio of 4:1 for linolenic acid:linoleic acid [42, 43]. After 2 years, the study reported a 70% decrease in mortality. The Singh and the GISSI study also corroborated the cardiovascular benefits of the Greek diet [44-46]. 3.4 Biochemistry of P U F A Naturally occurring dietary PUFAs can be divided into two major classes. Based on the position of the terminal double bond, the PUFAs are divided into co-3 (the double bond 3 carbons away) and co-6 (the double bond 6 carbons away) classes [39]. The co-6 and co-3 40 double bonds are entered into fatty acids by plant enzymes A 12 desaturase and A 15 desaturase. As these enzymes are absent in animals, dietary co-6 and co-3 PUFAs are termed as 'essential fatty acids'. The minimal essential requirement of these fatty acids was 0.2 to 1% of daily energy [47]. The parent fatty acids of these two groups are linoleic (18:2n-6) and linolenic (18:2n-3) acids. Plant oils are typically rich in linoleic acid, rather than linolenic acid. Interconversion between co-3 and co-6 PUFA is not possible in mammalian cells [48]. However, dietary linolenic and linoleic acids can be converted to different co-3 and co-6 PUFAs respectively within the cell via the process of desaturation (which adds double bonds) and elongation (which adds carbon atoms). Linoleic acid is converted via the combined action of desaturases and elongases to yield arachidonic acid (C20:4n-6). Using the same pathways, linolenic acid is converted to eicosapentaenoic (EPA, C20:5n-3) and docosahexaenoic acid (DHA, C22:6n-3) [39]. 3.5 Probable effects of co-6 PUFA in the diabetic heart Multiple pathways have been attributed to co-6 PUFA that can adversely affect cardiac health. Both diabetes and high co-6 PUFA feeding has been shown to activate phospholipase A2 (PLA 2) in the heart [49, 50]. This class of enzymes specifically cleaves the acyl group at the sn-2 position of most phospholipids, releasing constituent co-6 PUFA and lysophosphatidylcholine (LPC) [51]. The released co-6 PUFA can undergo lipid peroxidation by reacting with ROS to produce various toxic metabolites. The highly reactive hydroxyalkenal, 4-hydroxy-2-nonenal (HNE), is formed by the reaction of superoxide with membrane co-6 PUFA [52]. HNE can induce apoptosis in various cell types including neuronal cells [53], colon carcinoma cells [54] and endothelial cells [55]. Recently it has been proposed that overproduction of HNE can directly modify mitochondrial respiratory 41 chain and affect mitochondrial respiration in the diabetic rat heart [56]. Further, a major route of FINE metabolism in the heart includes its conjugation with glutathione (GSH) and activation of the polyol pathway in the normoglycemic heart [57]. As diabetes causes a depletion of cardiac GSH, with an activation of the aldose reductase pathway, such clearance mechanism could further lower cardiac GSH, thus precipitating greater toxicity. Cardiolipin, a mitochondrial membrane phospholipid, is absolutely essential for the activity of key OxPhos enzymes [58, 59]. We and others have shown that chronic co-6 PUFA feeding causes a loss of cardiolipin in the rat heart [16, 60, 61]. High co-6 PUFA feeding increases cardiac phospholipase A 2 (PLA 2) activity [50]. P L A 2 can hydrolyse cardiolipin to induce a loss in total cardiolipin mass, as seen in rat brain and liver [62, 63]. A drop in cardiolipin may damage the mitochondria and impair energy production by the heart. Indeed, co-6 PUFA feeding can increase myocardial oxygen consumption and decrease cardiac efficiency [64], oxidize mitochondrial DNA, with structural alterations [65]. The mitochondrial damage is prevented by coenzyme Q 1 0 , a mitochondrial antioxidant [66, 67]. Incubation of human endothelial cells with linoleic acid, a co-6 PUFA selectively upregulates transcriptional activity of nuclear factor kappaB (NF-kappaB) and activator protein 1 (AP-1) [4]. Increase in activity of these factors stimulated increased mRNA levels of several inflammatory proteins including tumor necrosis factor alpha (TNF-a) in these endothelial cells [4]. TNF can stimulate neutrophils, monocytes, T cells and endothelial cells to generate free radicals, which in turn produce more damage to the myocardium [68]. As diabetes per se increases cardiac TNF-a levels within 2 weeks, additive effect of linoleic acid induced TNF-a release could precipitate increased cardiotoxicity [69]. 42 3.6 Rationale And Objectives In recent years, with the help of advanced imaging techniques, an increasing prevalence of heart disease is being recognized in early clinical diabetes [70]. In animal models, cardiac dysfunction can occur within hours of inducing hyperglycemia in vitro [12], or within 4-6 days of diabetes in vivo [71]. In most cases, oxidative stress is blamed for inducing such an acute dysfunction [12, 71-73]. Oxidative stress occurs as a result of an imbalance between the production of reactive oxygen species (ROS), and their neutralization by antioxidants [74]. In the diabetic heart, augmented ROS production can arise from multiple sources like defective mitochondria, glucose auto-oxidation, protein glycation, and increased activity of cytosolic xanthine oxidase [75]. Irrespective of its origin, ROS can induce cell death via mechanisms including lipid, protein and DNA oxidation and initiation of diverse stress-signaling pathways. In both human and experimental diabetes, loss of GSH, a crucial cardiac antioxidant has been correlated to increased cardiac oxidative stress [76-80]. Although unclear in diabetic hearts, recent studies propose a role of specific sub cellular GSH pools like that of mitochondria, in modulating oxidative stress and cell death [81-84]. Although lipids have been acknowledged as a significant contributor to diabetic cardiac complications, most of the existing studies on the cytotoxicity of fatty acids have been performed in vitro, with single saturated fatty acids like palmitic acid on quiescent cultured cardiomyocytes [5, 18, 19, 85, 86]. However, in the last few decades, saturated fats from the human diet have been replaced by co-6 PUFA in form of vegetable oils [35-37, 48]. Thus, the effect of the more relevant co-6 PUFA on the heart, especially during diabetes remains unknown. 43 in My research proposal focused on identifying the role of GSH and co-6 PUFA modulating cardiac cell death during early diabetes. We had 3 objectives: 1. To determine the source of ROS and cardiac GSH status in acute diabetes. 2 . To evaluate the role of sub cellular pools of GSH in modulating cardiac cell death in diabetes. 3. To examine the effect of co-6 PUFA feeding on the myocardium, both during normoglycemic and hyperglycemic conditions. 44 3 . 7 Figure Norma l Pa lm i t i c a c i d E x c e s s Pa lm i t i c ac id APOPTOSIS Fig. 3.1 Modulation of cardiac cell death by saturated (palmitic acid) and monounsaturated fatty acids (MUFA) Under normal conditions, moderate levels of palmitic acid is oxidized by the heart (1). However, under conditions of fatty acid excess (as during diabetes), cardiac oxidative capacity is superceded, leading to the storage of excess palmitic acid as triglycerides (TG) (2). As a non-adipose tissue, cardiac capacity of T G accumulation is low, and excess, unoxidized palmitoyl CoA is converted to ceramide, a pro-apoptotic sphingolipid (3). Ceramide, via upregulation of NF-Kp, increases the expression of inducible nitric oxide synthase (iNOS), and generates nitric oxide (NO). Excess palmitic acid can also directly damage mitochondrial membrane ( 4 ) . Palmitic acid can upregulate the expression of pro-apoptotic factors like Bax and downregulate the expression of anti-apoptotic members of the BC1-2 family, thus predisposing the heart to apoptosis (5). Increase in pro-apoptotic factors, damaged mitochondrial membrane and increased ceramide precipitates an increase in reactive oxygen species (ROS) (6). Increased mitochondrial ROS leads to the release of cytochrome c (cyto c) in the cytosol, which ultimately precipitates a caspase cascade, terminating in the activation of caspase 3, DNA damage and apoptosis ( 7 ) . MUFA (mainly oleic and palmitoleic acid) augments cardiac T G storage and removes free palmitic acid (8). Additionally, M U F A can also augment cardiac antioxidant levels and expression of PI3 kinase (PI3K), an important survival factor within the cardiomyocytes, and prevent or delay apoptosis (9). 45 3.8 References 1. Gouni-Berthold I, Berthold HK,'Seul C, Ko Y, Vetter H, Sachinidis A. 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S-Adenosyl-L-methionine and mitochondrial reduced glutathione depletion in alcoholic liver disease. Alcohol 2002; 27:179-83 84. Colell A, Garcia-Ruiz C, Miranda M , Ardite E, Mari M , Morales A, et al. Selective glutathione depletion of mitochondria by ethanol sensitizes hepatocytes to tumor necrosis factor. Gastroenterology 1998; 115:1541-51 85. Hickson-Bick DL, Sparagna GC, Buja L M , McMillin JB. Palmitate-induced apoptosis in neonatal cardiomyocytes is not dependent on the generation of ROS. Am J Physiol Heart Circ Physiol 2002; 282:H656-64. 86. Listenberger L L , Ory DS, Schaffer JE. Palmitate-induced apoptosis can occur through a ceramide-independent pathway. J Biol Chem 2001; 276:14890-5 56 4. Increased Efflux Of Glutathione Conjugate In Acutely Diabetic Cardiomyocytes 4.1 Introduction Diabetes is a significant risk factor for cardiovascular diseases with the majority of these complications being attributed to coronary vascular pathology. However, both in humans and animal models of diabetes, an additional heart muscle specific disease in the absence of any vascular pathology has also been described, and termed "diabetic cardiomyopathy"!!]. Probable candidates to explain this heart disease include autonomic abnormalities, metabolic disorders, abnormal contractile protein and enzyme function, interstitial fibrosis [1], and more recently apoptosis, a regulated, energy dependent, cell suicide mechanism.[2, 3]. Reactive oxygen species (ROS) are highly reactive oxidants that cause tissue injury and cell death. Hyperglycemia induced increase in ROS, either due to an increased production or a decrease in the antioxidant capacity, has been implicated in inducing cardiomyocyte apoptosis [4]. Once increased, ROS like hydrogen peroxide (H2O2) initiates oxidative modification of structural and functional proteins and lipids leading to apoptotic cell death. With respect to antioxidants, glutathione is a tripeptide (L-y-glutamyl-L-cysteinyl glycine, GSH) involved in the protection of cells and tissues against H2O2 induced damage [5]. y-glutamylcysteine synthetase (y-GCS) is the rate-limiting enzyme for the biosynthesis of GSH. Once formed, reduced GSH neutralizes H2O2 and is oxidized to GSH disulfide (GSSG). GSSG can be reduced back to GSH in an enzymatic recycling reaction catalyzed by GSH reductase (GR) and NADPH [5]. Another pathway contributing to GSH homeostasis is that of cellular GSH extrusion [6]. Under conditions of oxidative stress, glutathione binds to lipid peroxidation products that are then effluxed from the cell by transporters like P-glycoproteins and multidrug resistance proteins (MRPs) [7]. 57 Although the regulation of y -GCS and G S H reductase has been extensively studied in the chronically diabetic heart [8, 9], their contribution in modulating cardiac oxidative stress in the acutely diabetic heart is largely ignored. Additionally, the role of glutathione efflux in modulating cardiac oxidative stress during diabetes is unknown. Using acutely diabetic animals, the objective of the present study was to examine the glutathione status and to determine i f myocyte G S H efflux contributes towards cardiac oxidative stress. 5 8 4.2 Materials And Methods 4.2.1 Experimental Animals Animals were cared for in accordance with the principles promulgated by the Canadian Council of Animal Care and the University of British Columbia. Male Wistar rats (350-360 g) were obtained from the University of British Columbia Animal Care Unit. Animals were made diabetic with a single intravenous injection of streptozotocin (STZ, 55 mg/kg). STZ causes selective p-cell death with profound hyperglycemia developing within 24h. A commercially available kit (Sigma) was used to measure serum glucose obtained from tail vein blood samples. Diabetic rats were kept for 4 days, anesthetized with 65 mg/kg sodium pentobarbital i.p, the thoracic cavity opened, and hearts removed. 4.2.2 Cardiac Apoptosis Part of the left ventricle was fixed in 10% formalin, embedded in paraffin, and 5 pm sections prepared. TdT mediated dUTP nick end labeling (TUNEL) assay was carried out on these sections using a T U N E L kit (Oncogene) as described previously [10]. At least six random sections of each heart were quantified using the software 'Northern Eclipse'. Values are expressed as number of TdT labeled nuclei per 106 nuclei. To substantiate the incidence of apoptosis, caspase-3 (whose activity increases preceding apoptosis) was estimated using a kit (Clontech). Briefly, heart tissue (200 mg) was placed in ice-cold cell-lysis buffer (10 mM HEPES, 2 mM EDTA, 0.1 % CHAPS / NP40, 5 mM DTT, 1 mM PMSF, 10 ug/ml pepstatin A, 20 ug/ml leupeptin, 10 ug/ml aprotinin) and homogenized. Subsequently, the suspension was centrifuged at 12,000 x g for 3 min, the supernatant separated, and caspase-3 activity measured. 'Active' caspase-3 can remove the chromophore pNA (para-nitroaniline) from the labeled 59 substrate DEVD-pNA; measuring free pNA in the reaction mixture is diagnostic for caspase activity [11]. Values are expressed as O.D.U. (optical density units) per 100 mg wet weight. 4.2.3 Cardiac Lip id Peroxidation Oxidative stress was determined by the appearance of thiobarbituric acid-reactive substances like malondialdehyde (MDA) [12]. Heart tissue was ground under liquid nitrogen, and homogenized. After acidification of the homogenate with 1% phosphoric acid and treatment with 0.6 % T B A solution, the mixture was heated in the presence of 0.4% butylated hydroxytoluene to prevent further tissue oxidation. After cooling, the adduct was extracted into n-butanol, and absorbance measured at 540 nm against 1,1,3,3-tetramethoxypropane as the standard. 4.2.4 Preparation of Cardiac Myocytes Hearts were digested by perfusing modified Joklik minimal essential medium containing collagenase (228 U/ml), 0.5% BSA, and 50 mM CaCl 2 retrogradely through the heart [13]. Myocytes were made calcium-tolerant by successive exposure to increasing concentrations of calcium upto 1 mM Ca 2 + . Yield of cardiomyocytes (90-95%) of total cell number) was determined microscopically using a Neubauer haemocytometer. Myocyte viability (generally between 75-85%) was assessed as the percentage of elongated cells with clear cross striations that excluded 0.2% trypan blue. Cardiac myocytes, were either used fresh for estimation of GSH or GSH efflux or were plated in laminin coated 6 well culture plates using colorless Media-199 (Sigma) to a cell density of 75,000 cells/well, and incubated at 37°C under an atmosphere of 95% 0 2 - 5 % C 0 2 for fiuorimetric estimation of ROS. 4.2.5 Estimation of G S H GSH content of freshly isolated cardiomyocytes was measured using a commercially available kit (Cayman Chemicals). GSH was reacted with 5,5'-dithiobis-2-nitrobenzoic acid (DTNB) to 60 produce a yellow colored 5-thio-2-nitrobenzoic acid (TNB). GS-TNB, produced in parallel, is recycled back to GSH by glutathione reductase to produce more TNB. Rate of TNB production is directly proportional to this recycling reaction, which is in turn directly proportional to the total concentration of GSH in the sample. Oxidized glutathione (GSSG) was estimated using 2-Vinylpyridine[14]. The difference between total GSH and GSSG gave the value of the reduced GSH. Protein assay was performed according to the Bradford Method. Values are expressed as nM GSH per mg protein. 4.2.6 Cardiac Gene Expression Cardiac gene expression was measured using semiquantitative reverse transcriptase polymerase chain reaction (RT-PCR) as described previously [10]. PCR primers were designed from rat sequences available in GenBank, and are as follows: gamma glutamylcysteine synthase heavy chain (j-GCSHc); 5' -C A A G A G A A G G G G G A G A G G AC-3' (forward) and 5'-T T A T C G T G C A A A G A G C C T G A - 3 ' (reverse) (accession # NM012815); multidrug resistance protein 1 (MRP-1); 5' - C C T T T T C C T G T G C A A T C A T G T A - 3 ' (forward) and 5'-A G A A C C T C T G C A C A A A G A A G T A -3' (reverse) (accession # NM022281); p-actin; 5'-C G T A A A G A C C T C T A T G C C A A - 3 ' (forward) and 5' - A G C C A T G C C A A A T G T G T C A T - 3 ' (reverse) (accession # J00691). Levels of expression were indicated as the ratio of signal intensity for the mRNA of interest relative to that for P-actin mRNA. 4.2.7 Glutathione Reductase Activity Ventricular tissue was powdered under liquid nitrogen, homogenized in cold buffer (50 nM potassium phosphate, pH 7.4, ImM EDTA), and centrifuged at 7000g for 15 minutes at 4°C. 100 ul of supernatant was removed for protein analysis (Bradford Assay). For the assay, GSSG and 20 ul of samples were added to a cuvette. The reaction was initiated by the addition of 2 mM NADPH and absorbance recorded at 340 nm. Glutathione reductase (GR) activity was 61 measured by the decrease in absorbance, which accompanies oxidation of NADPH to NADP+[15], Values were expressed as nmol of NADPH/min/ml. 4.2.8 G S H Efflux The bimanes are heterocyclic compounds that form fluorescent conjugates with thiols [16, 17]. Within the cell, monochlorobimane [syn-(QCH2,CH3)-l,5-diazabicyclo-[3,3,0]-octa-3,6-dione-2,8-dione; mClB), a freely permeable non-fluorescent dye, that conjugates preferably with reduced GSH [18]. Once complexed, mClB emits a strong fluorescence (excitation, 380 nm; emission 485 nm). Briefly, freshly isolated cardiomyocytes (0.5 x 106) were transferred in duplicate to pre-warmed 10 ml test tubes. The cells were spun lightly to remove all media (1000 x g x 1 min) and 2 ml of loading media (50pM mClB, lOpM GSH, lg/ml BSA, prepared in M-199) added. After 15 mins incubation, cells were spun again to remove loading media and 2 ml of the efflux media was added (loading media lacking mClB). 50ul of the media was taken out immediately for estimation of the initial value. Cells were then incubated at 37°C under an atmosphere of 95% 0 2-5%C0 2 either in the presence or absence of MK-571 (lOpM) or verapamil (lOpM). The cells were periodically shaken and 50pl of media aspirated at 15, 30 and 45 minutes for estimation of mCIB-GSH complex (fluorimetry) and lactate dehydrogenase (LDH; spectrophotometry). Finally, the cells were separated from the media by centrifugation for final estimation of cellular and media mCIB-GSH complex (at 45 minutes). Cellular content of the complex was estimated by sonication and precipitation of proteins with lOpl of 60% perchloric acid as described previously [16]. The precipitate was separated by centrifugation and 50pl of the resultant supernatant was estimated by fluorimetry. Total (media+cell) GSH was estimated by adding the fluorescence of the media at 45 minutes to the value obtained from the cellular mCIB-GSH complex (from 0.5x106 cells) at this time. Negative controls were generated using diethyl maleate (DEM) that complexes with GSH and 6 2 prevents formation of the mClB-GSH complex. D E M (0.02%) was added 15 minutes before the start of the experiment. 4.2.9 LDH Release To ascertain that the release of mClB-GSH is not due to cell membrane leakage, lactate dehydrogenase (LDH) release into the media was estimated using an in vitro toxicology assay kit (Sigma). In brief, L D H reduces nicotinamide adenine dinucleotide (NAD +) that then converts a tetrazolium dye to a soluble, colored formazan derivative that was measured using a spectrophotometer (490 nm). 4.2.10 Measure ment of ROS A redox sensitive dye, 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H 2 DCFDA) was used to assess the presence of ROS [19]. C M - H 2 D C F D A is a cell-permeant indicator for ROS like H 2 0 2 that reacts with the probe, oxidizes it and emits green fluorescence. For the quantative estimation of ROS, plated cells (65-75,000 myocytes/ per well) were either left untreated or incubated with MK-571 (lOuM), verapamil (10uM), or N-acetylcysteine (NAC, 5mM) for 2 hours. Subsequently, 0.5 ml colourless M-199 and C M -H 2 DCFDA (final concentration, 5 uM) were added per well. The increase in green fluorescence was monitored in a fluorimeter over the next 90 minutes (excitation: 480nm; emission: 530nm). Background was identified in wells without cells. Data was expressed per 75,000 cells as fluorescence units (F.U.) after subtracting the background at 90 minutes. 4.2.11 Statistical Analysis Values are means ± SE. One-way A N O V A followed by the Bonferroni test or the unpaired Student's Mest was used to determine differences between group mean values. The level of statistical significance was set at P < 0.05. 63 4.3 Results 4.3.1 Cardiac apoptosis and oxidative stress in STZ diabetes Table 4.1 depicts the general characteristics of control and 4-day diabetic animals. STZ injection induced hyperglycemia and a loss of body weight. Based on previous reports of increased apoptosis in hearts from STZ diabetic rats and mice, we attempted to corroborate these results in our model of diabetes. TUNEL staining showed that diabetic hearts had an approximately 5-fold increase in apoptotic cells compared to controls with a corresponding 2-fold increase in active caspase-3 activity. As hyperglycemia-induced cardiac apoptosis has been intimately linked to an increase in ROS production [19], cardiac oxidative stress (TBARS like MDA) was also determined and found to be elevated in 4 day STZ-diabetic rats (Fig. 4.1, top panel). 4.3.2 Glutathione status in STZ-diabetic hearts Acute diabetes was associated with a significant decrease in cardiomyocyte GSH levels (Fig. 4.1, lower panel). Given that GSH can decrease, either due to changes in glutathione biosynthesis or a decrease in GSH regeneration [20], we measured gene expression of the heavy subunit of y-glutamylcysteine synthetase ( Y - G C S H ) and the activity of glutathione reductase, and determined that both remained unchanged following diabetes (Table 4.2). Interestingly, evaluation of a third pathway by which GSH can fall revealed that diabetic cardiomyocytes demonstrate a higher rate of GSH efflux, as measured by extrusion of mGB-GSH into the incubation media (Fig. 4.2). This increase in GSH efflux was not a result of a difference in cell loading of the mCIB-GSH complex, as both control and diabetic cardiomyocytes had nearly identical total GSH-mClB fluorescence (Fig. 4.2; right inset). Isolation of cardiomyocytes with collagenase leads to membrane damage and associated L D H release. As L D H release between control and diabetic cardiomyocytes were similar, it is 64 unlikely that the increase in GSH efflux in diabetic cardiomyocytes is a result of non-specific changes in membrane permeability (Fig. 4.2; left inset). 4.3.3 M R P like activity in diabetic cardiomyocytes As MRP-1 has been associated with glutathione efflux under conditions of oxidative stress, its contribution towards efflux of mClB-GSH in diabetic cardiomyocytes was investigated [7, 21]. 4 days of hyperglycemia led to an increase in cardiac MRP-1 gene expression (Fig. 4.3). Additionally, incubation of these myocytes with MK-571, a specific blocker of MRP-1 [22] reduced mClB-GSH efflux to control levels [i.e., similar to cells in which mClB-GSH efflux is completely inhibited using D E M , (Fig.4.4)]. To investigate the contribution of transporters other than MRP-1, diabetic cardiomyocytes were also incubated with verapamil, an inhibitor of P-glycoproteins [16]. Although verapamil lowered mClB-GSH release into the incubation media, efflux was still higher than that observed with MK-571 (Fig.4.4). 4.3.4 Decrease in cardiomyocyte R O S following inhibition of G S H efflux Following isolation of cardiomyocytes and plating for 2 hours, measurement of ROS using H 2 D C F D A fluorescence revealed higher ROS levels in diabetic cardiomyocytes (Fig. 4.5). Given the association between GSH and ROS, cardiomyocytes were treated with either verapamil or MK-571 for 2 hours, and ROS levels measured. Unlike verapamil, MK-571, an efficient blocker of GSH extrusion in diabetic cardiomyocytes, was more effective in reducing ROS levels. Additionally, incubation of diabetic cardiomyocytes with N A C (that increases intracellular GSH levels) [23, 24] also decreased ROS. 65 4.4 Discussion Heart disease in patients with diabetes can occur due to intrinsic defects in the heart muscle [1]. Of the many proposed mechanisms contributing towards this cardiac dysfunction, regulated cell death is a more recent addition to the list. In this regard, an increase in apoptosis was observed in hearts from diabetic patients and STZ-diabetic animals [2, 4]. As a number of issues like strain, dosage of STZ and duration of the diabetic state may influence apoptosis; we were concerned about the prevalence of this mode of cell death in our acute model of diabetes. This is important as in strains like Wistar-Kyoto (WKY) rats, 55mg/kg STZ does not induce diabetic cardiomyopathy [25]. This could be related either to the lower incidence of hypertriglyceridemia, or an inherent resistance to the diabetogenic action of STZ, resulting in greater pancreatic insulin content in these rats [25]. However, with Wistar rats, we detected a 5-fold increase in apoptosis in the diabetic heart using TUNEL assay, and further validated this observation by identifying an increase in caspase-3 activity, the proteolytic enzyme involved in the execution of apoptosis [26]. Given that the STZ-diabetic rat demonstrates cardiomyopathy 4-6 weeks after STZ injection, it is possible that this initial loss of cells progressively leads to interstitial fibrosis, myocardial hypertrophy, and eventual contractile impairment [27]. Oxidative stress due to excessive ROS generation and/or inadequate ROS neutralization is believed to be a key player in inducing apoptosis. Once generated, ROS can cause cell death via lipid peroxidation reactions, alterations of cellular proteins and initiating diverse signaling pathways [28, 29]. Both ROS and lipid peroxidation were elevated in the diabetic heart and could be implicated in explaining cardiac apoptosis in these animals. In an attempt to elucidate the mechanisms behind the increase in ROS, we measured GSH, the main antioxidant responsible for detoxification of ROS like H2O2 in the heart [30]. At least in chronic human and experimental diabetes, GSH levels decrease in various cells and organs like platelets [31], polymorphonuclear cells [32], liver, kidney, neurons and heart [33-35]. Our 66 results suggest that cardiac GSH levels are also reduced following acute hyperglycemia. In chronically diabetic rats (2-8 weeks), fall in GSH is often linked to decrease in biosynthesis and/or altered enzymatic recycling as a consequence of reduced gene expression [36] or activity [8] of y-GCS, and decreased activity of glutathione reductase [8]. Interestingly, in our acute model of diabetes, fall in cardiomyocyte GSH could not be explained by either of the above mechanisms and indicated participation of GSH extrusion behind this drop. Indeed diabetic cardiomyocytes demonstrated higher GSH conjugate efflux compared to controls. Similar to our results, ischemia-reperfusion leads to loss of GSH, not by changes in glutathione reductase activity but by active GSH efflux [37]. Overall, our data suggests that at least at this early time point, GSH extrusion may be a major contributor in the fall of cardiac GSH. MRP-1, a 190kDa membrane glycoprotein and member of the ATP-binding cassette (ABC) transporter family is well recognized for conferring multidrug resistance by effluxing anti-cancer drugs [16]. However, its affinity is highest for endogenous GSH conjugates [38], and can transport GSH out of astrocytes and cancer cell lines in response to oxidative stress [7, 21]. MRP-1 has ubiquitous distribution, including expression in the heart [39, 40]. In our study, cardiac MRP-1 gene expression increased following diabetes, potentially increasing GSH efflux with subsequent decrease in cardiomyocyte GSH. MK-571, a leukotriene D 4 receptor antagonist and an MRP-1 selective inhibitor [22] blocked mClB-GSH efflux in diabetic cardiomyocytes. As both MK-571 and N A C (which increases the intracellular concentration of glutathione) decreased ROS, our data suggests that MRP-1 has an important role in GSH and ROS homeostasis in the acutely diabetic heart. In the heart, other A B C transporters like P-glycoproteins could potentially efflux GSH [41-43]. Verapamil, a P-glycoprotein blocker, partially decreased mClB-GSH efflux, and suggested a role for this transporter in diabetic cardiomyocytes. However, as verapamil can also influence MRP-1 when used along with GSH, the influence of P-glycoproteins in our 67 efflux experiments is likely minimal. Additionallyfwhether MRPs other than MRP-1 could explain this partial drop of mClB-GSH efflux with verapamil is yet to be determined [17]. At present, the purpose for this enhanced MRP-1 expression in the diabetic heart is unknown. It is possible that MRP-1 through its efforts to reduce 4-hydroxynonenal (4-HNE; a toxic aldehyde responsible for the modification of proteins), inadvertently reduces GSH to which 4-HNE is conjugated [7, 44, 45]. Alternatively, for successful induction and maintenance of the apoptotic pathway in diabetic cardiomyocytes, it may be necessary for dying cells to lose their reducing capacity in favor of a pro-oxidizing environment, a goal that can be achieved easily through export of GSH, the most important cytosolic antioxidant [6]. It should be noted that MRP-1 has also been implicated in exporting the oxidized form of glutathione (GSSG). [46]. However, loss of GSSG also translates into net loss of GSH. In conclusion, our findings demonstrate that the acutely diabetic heart is characterized by increased expression of MRP-1, reduction in GSH, increased oxidative stress and an elevated apoptotic index. Mechanisms responsible for the increase in MRP-1 expression in the diabetic heart are unknown and are currently being investigated. 68 4.5 Tables And Figures Table 4.1 General characteristics of the experimental animals C O N T R O L D I A B E T I C r> A wi ' u+1 \ 384 ± 5 366 ± 3 * Body Weight (g) c n . . 1 / T , 8.3 ±0 .2 22.9 ± 0 . 3 * Serum Glucose (mmol/L) TdT Labeled Cardiomyocytes (per 106 nuclei) 101 ± 16 495 ± 20 n i •• - w n r » T T / inn ^ 0.16 ±0.01 0.39 ±0 .01* Caspase-3 activity (O.D.U./per 100 mg w.w) Values are means ± S E for 4-6 animals in each group. Animals were made diabetic by S T Z administration (55 mg/kg; i.v.). Diabetic animals were kept for 4 days before being killed. Values are those obtained before death and were from fed animals. * Significantly different from controls. P < 0.05. TdT, terminal deoxynucleotidyl transferase; O . D . U , optical density unit; w.w.,wet weight. 69 Table 4.2 Factors Regulating Intracellular GSH Status CONTROL DIABETIC y-GCS heavy chain/p-actin mRNA expression GR activity (nmol NADPH/min/ml/mg protein) Values are means ± SE for 4-6 animals in each group. Animals were made diabetic by STZ administration (55 mg/kg; i.v.). Diabetic animals were kept for 4 days before being killed. Values are those obtained before death and were from fed animals. * Significantly different from controls. P < 0.05. y-GCS, gamma glutamylcysteine synthase; GR, glutathione reductase. 0.66 ± 0 . 0 8 0.70 ± 0 . 0 3 594 ± 29 492 ± 99 7 0 Fig. 4.1 Oxidative stress in cardiac myocytes isolated from control and STZ-diabetic rats Lipid peroxidation was estimated with a thiobarbituric acid-reactive substance (TBARS) assay, with malondialdehyde (MDA) levels depicted in the upper panel. The lower panel shows cardiomyocyte GSH levels that were determined using an enzyme-recycling assay. *Significantly different from control, P < 0.05. 71 - I — 1 1 1 0 15 30 45 T i m e Fig. 4.2 mClB-GSH efflux from isolated cardiomyocytes Control and diabetic cardiomyocytes were isolated, and incubated with 50 uM mClB and 10 uM GSH. At time 0, the medium was removed and replaced with medium lacking mClB, and the amount of mClB-GSH release measured in the media over 45 minutes. The results are expressed per 0.5 x 106 cells in fluorescence units (F.U.) as a percentage of the initial value (time 0). Inset (left): Total L D H released into the media at termination of the experiment (45 minutes). Inset (right): Total mClB-GSH fluorescence calculated by adding the fluorescence from cell lysates and the media at termination.. Significantly different from control, P < 0.05. CON, Control; DIA, Diabetic; A.U. , arbitrary units; F.U.; fluorescence units 72 CONTROL DIABETIC CONTROL DIABETIC Fig. 4.3 MRP-1 gene expression in hearts from control and diabetic rats Bars represent the mean ±SEM (n=3) of MRP-1 /p-actin ratio. The inset is a representative blot of MRP-1 RT-PCR products. *Significantly different from control, P < 0.05. 73 Fig. 4.4 Effect of different inhibitors on the efflux of mCIB-GSH from diabetic cardiomyocytes Diabetic cardiomyocytes were first incubated with media containing 50 uM mClB and 10 uM GSH. After 15 minutes, it was replaced with medium lacking mClB and media fluorescence calculated as percentage of the initial value (time 0). mCIB-GSH release in the media in the presence or absence of either MK-571 (10 pM) or verapamil (10 pM) was measured over the next 45 minutes. D E M (0.02%) was added 15 minutes before the start of the experiment and used as the negative control. Significantly different from untreated diabetic myocytes. "Significantly different from verapamil treated myocytes, P < 0.05. UTX, Untreated 74 F i g . 4.5 ROS status in control and diabetic cardiomyocytes For the qualitative estimation of ROS, plated cells were either left untreated or incubated with MK-571 (10 pM), verapamil (10 pM), or N-acetylcysteine (NAC, 5 mM) for 2 hours. Subsequently, media with C M - H 2 D C F D A (final concentration, 5 pM) was added to each well. The increase in fluorescence was monitored in a fluorometer over the next 90 minutes (excitation: 480nm; emission: 530nm). The bars represent H 2 D C F D A fluorescence (i.e. ROS content) in cardiomyocytes expressed in fluorescence units (F.U.) per 7.5 x 104 cells at termination of the experiment (45 minutes). Significantly different from untreated diabetic myocytes; "Significantly different from verapamil treated myocytes. P < 0.05. U T X , Untreated 75 4.6 References 1. Rodrigues B, Cam M C , McNeill JH. 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However, both in humans and animal models of diabetes, an additional heart muscle specific disease in the absence of any vascular pathology has also been described [1]. Probable candidates to explain this heart disease include autonomic abnormalities, metabolic disorders, abnormal contractile protein and enzyme function, interstitial fibrosis [1], and more recently apoptosis, a regulated, energy dependent, cell suicide mechanism [2-4]. In both human and animal models of diabetes, heart disease is associated with excessive apoptotic cell death [2, 3]. It is possible that this loss of cells progressively leads to interstitial fibrosis, myocardial hypertrophy, contractile impairment and eventual heart disease [5, 6]. Various factors including protein kinase C activation [7], upregulation of the renin-angiotensin system [2], p53 mediated gene transcription [8], and pro-apoptotic free fatty acids [6] have been implicated in inducing myocardial apoptosis during diabetes. Interestingly, all of these different elements appear to trigger cell death by a common 'oxidative stress' mediated pathway [9]. Oxidative stress occurs as a result of an imbalance between the production of reactive oxygen species (ROS), and their neutralization by antioxidants. In the diabetic heart, augmented ROS production can arise from defective mitochondria, glucose auto-oxidation, protein glycation, and increased activity of cytosolic xanthine oxidase [10]. Irrespective of its 81 origin, ROS such as hydrogen peroxide (H2O2) can induce cell death via mechanisms including lipid peroxidation, alteration of cellular proteins, and initiation of diverse stress-signaling pathways [11]. Among antioxidants, glutathione (L-y-glutamyl-L-cysteinyl glycine, GSH) is a non-protein thiol crucial for protecting the heart against H2O2 induced damage [11]. In both human and experimental diabetes, loss of GSH has been demonstrated and correlated to increased cardiac oxidative stress [12-14]. Reduced GSH, present in the cytosol (cGSH, 80% of total cellular GSH) and mitochondria (mGSH, 10-15%) of total GSH), acts as a hydrogen donor in the neutralization of H2O2, and is itself converted to the inactive glutathione disulfide (oxidized glutathione; GSSG) [15]. Despite the importance of mGSH in preventing mitochondrial oxidative stress, mitochondria are incapable of de novo synthesis of GSH and rely on the import of GSH from the cytosol where it is manufactured from constituent amino acids. Recently, we have reported that short-term diabetes induces cardiac GSH depletion, increased ROS levels, and cardiac apoptosis as early as 4 days after streptozotocin (STZ) administration [16]. Interestingly, similar observations were reported when cardiomyocytes were exposed to 24 hours of high glucose in vitro [17]. In both these studies, neither the source of ROS nor the site of GSH loss was explored. This is important because cytosolic and mitochondrial sources of ROS may affect specific pools of GSH [i.e. cGSH or mGSH], and differentially regulate apoptosis [15, 18]. Using acutely diabetic rats, the objective of the present study was to: (i) determine the source of cardiac ROS, and (ii) investigate the contribution of the respective intracellular GSH pools to cardiac apoptosis. We report that in the diabetic heart, mitochondria may be the chief source of ROS, with mGSH depletion being' crucial in the activation of the cell suicide program in vivo. 82 5.2 Materials And 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 (NIH Publication No. 85-23, revised 1985), and the University of British Columbia. Male Wistar rats (220-240 g) were obtained from the University of British Columbia Animal Care Unit, and maintained under a 12-h light (0700-1900)/dark cycle. All animals were fed standard laboratory chow diet (PMI Feeds, Richmond, VA). Water was provided ad libitum. In some rats, diabetes was induced with a moderate dose of streptozotocin (STZ; 55 mg/kg, i.v.), a selective beta cell toxin, and hyperglycemia verified after 24 hours with a glucometer (Advantage Comfort, Roche Diagnostics). All diabetic rats were kept for 4 days following STZ administration, anesthetized with sodium pentobarbital (65 mg/kg, i.p.), the thoracic cavity opened, and hearts removed [5, 6, 16]. Following termination, plasma insulin was measured using rat insulin standards by radioimmunoassay (Novo, Copenhagen, Denmark). 5.2.2 Treatments To examine the influence of acute diabetes on specific pools of GSH, in addition to determining how manipulation of these pools of GSH could affect myocardial apoptosis, some rats were left untreated or injected with L-buthionine sulfoximine (BSO; 10 mmol/kg, i.p.; dissolved in saline) and/or diethyl maleate (DEM; 4 mmol/kg, i.p.; dissolved in corn oil) for 4 days following STZ-administration. A control group with both BSO and D E M treatments was utilized to negate the possibility of non-specific toxic effects of these agents. BSO inhibits the activity of y-glutamyl cysteine synthetase (y-GCS), and blocks the de novo synthesis of GSH [11], whereas D E M conjugates and inactivates all GSH within the cell [18-20]. In contrast, to 83 augment cardiac GSH, some rats were treated with daily injections of GSH (400 mg/kg; pH 7.4; i.p.) for 10 days, prior to induction of diabetes. Administration of GSH was continued for 4 days following STZ-administration. 5.2.3 Preparation Of Cardiac Myocytes Hearts were digested by perfusing modified Joklik minimal essential medium containing collagenase (228 U/ml), 0.5% BSA, and 50 mM CaCl 2 retrogradely through the heart. Myocytes were made calcium-tolerant by successive exposure to increasing concentrations of calcium (up to 1 mM). Yield of cardiomyocytes (90-95%) of total cell number) was determined microscopically using a Neubauer haemocytometer. Myocyte viability (generally between 75-85%>) was assessed as the percentage of elongated cells with clear cross striations that excluded 0.2%) trypan blue. There was no difference in myocyte viability (percent of live cells) or yield (total number of cells x 106) between control and diabetic rats at 4 days after the induction of diabetes. Immediately after isolation, fractions of cardiomyocytes were snap frozen in liquid nitrogen for estimation of various caspases. The remaining cardiomyocytes were either fractionated for determination of sub cellular pools of GSH, or plated on laminin coated culture plates as described previously [16]. 5.2.4 Estimation of ROS A redox sensitive dye, 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H 2 DCFDA) was used to assess the presence of ROS [4, 16]. C M - H 2 D C F D A is a cell-permeant indicator that is oxidized in the presence of ROS such as H 2 0 2 , emitting green fluorescence. Myocytes were isolated and plated for 16 hours. Thereafter, the media was discarded and wells loaded with C M - H 2 D C F D A (5 pM, 30 min, 37°C). Subsequently, cells were washed and incubated for a further 45 mins, after which green fluorescence was monitored in a fluorimeter at 485/530 nm. To identify the sources of ROS production, some 84 cardiomyocytes were incubated with rotenone (5pg/ml) or antimycin A (100 ng/ml), for 1 hour [21]. Rotenone blocks Complex I of the mitochondrial electron transport chain (ETC) and decreases mitochondrial ROS generation [21]. Conversely, incubation with antimycin A blocks Complex III and increases mitochondrial ROS [22]. Finally, N-acetylcysteine (NAC, 5 mM) which can decrease ROS production by increasing the intracellular GSH concentration [23], was used to monitor the effect of GSH supplementation in vitro. The emission was calculated per 75,000 cells in fluorescence units. For confocal microscopic images, chambered coverslips were utilized as described above. Myocytes were visualized using a confocal scanning microscope at 485/530 nm wavelength at 300x magnification for visualization of green fluorescence [16]. 5.2.5 Determination Of Mitochondrial Membrane Potential JC-1, a carbocyanine dye was utilized to assess changes in mitochondrial membrane potential (Avpm) [24]. At low concentrations, this dye exists as a monomer in the cytosol, yielding green fluorescence at 530 nm [25]. Being cationic in nature, it accumulates in mitochondria and forms dimers, exhibiting red fluorescence at 590 nm. Formation of these dimers is directly proportional to A\|/ m [24]. Cells were isolated, plated and cultured for 16 hours. Thereafter, the media was discarded and the wells loaded with 10 pg/ml JC-1 for 10 minutes, washed in ice-cold phosphate buffered saline, and scanned immediately with a CYTOFLUOR™ fluorimeter at 530/580 nm wavelength for dimers, and 485/530 nm for the monomeric JC-1. The emission was estimated per 75,000 cells in fluorescence units, and the red to green fluorescence ratio calculated. For confocal microscopic images, chambered coverslips were utilized instead of culture plates. Following procedures described above, myocytes were visualized using a confocal scanning microscope (Biorad MRC 600) at 530/580 nm wavelength, and 600x magnification. A single scan was performed to prevent mitochondrial damage during the 85 experiment. 100 uM of the mitochondrial uncoupler, m G C C P was utilized as a negative control [25]. 5.2.6 Separation Of Cytosol And Mitochondrial Fractions Separation of cytosol and mitochondria was achieved by digitonin fractionation [26]. In brief, digitonin (0.24 mg/ml) was added to the cardiomyocyte pellet (2 x 106 cells) to disrupt the plasma membrane. After 2 mins, the mixture was transferred to eppendorf tubes containing 40% glycerol-silicone-oil mixture, and buffer containing 19.8 mM EDTA, 250 mM mannitol and 17 mM HEPES, and centrifuged at 13,000g for 3 mins. The top oil layer was separated and discarded without perturbing the middle buffer layer (cytosolic fraction). Triton X was added to the bottom glycerol layer (mitochondrial fraction), and sonicated for 15-20 sees. To determine fractionation efficiency, the glycerol and buffer samples were each assayed spectrophotometrically for succinate dehydrogenase (SDH; mitochondrial marker), and lactate dehydrogenase (LDH, cytosolic marker) activities. Since only ~5% of the total SDH was released into the buffer layer, mitochondria were recovered intact in the glycerol layer [26] before sonication. Additionally, L D H activity within the glycerol layer was also minimal (-5%). 5.2.7 Estimation Of Glutathione Intracellular fractions were immediately subjected to deproteination with 5% metaphosphoric acid to remove protein thiol groups, snap frozen in liquid nitrogen and stored at -80°C. Determination of glutathione was performed within the next 48 hours. Cardiac GSH content was measured using a commercially available kit (Trevigen). GSH was reacted with 5,5'-dithiobis-2-nitrobenzoic acid (DTNB) to produce a yellow colored 5-thio-2-nitrobenzoic acid (TNB). GS-TNB, produced in parallel, is recycled back to GSH by glutathione reductase to produce more TNB. Rate of TNB production is directly proportional to this recycling reaction, 86 which is in turn directly proportional to the total concentration of GSH in the sample. Oxidized glutathione (GSSG) was estimated using 2-Vinylpyridine [27]. The difference between total GSH and GSSG gave the value of the reduced GSH. Protein assay was performed according to the Bradford method using a non-deproteinated portion of the samples. Values are expressed as nM GSH per mg protein. 5.2.8 Cardiomyocyte Apoptosis Where indicated, part of the left ventricle was fixed in 10% formalin, embedded in paraffin, and 5 urn sections prepared. To estimate cardiomyocyte apoptosis, TdT mediated dUTP nick end labeling (TUNEL) assay with cardiomyocytes specific labeling was carried out as described previously [5, 16]. At least six random sections of each heart were quantified using the software 'Northern Eclipse'. Values are expressed as number of TdT labeled nuclei per 106 nuclei. 5.2.9 Cardiac Caspase Activity Activities of cardiac caspase-3, 8 and 9 were determined using the corresponding fluorescent caspase specific substrates D E V D - A M C (Molecular Probes), IETD-AMC (Biovision) and L E H D - A M C (Biosource), respectively. Briefly, 50-75 pg of total cell protein was added to reaction buffer containing 50 uM of the respective substrate, and incubated at 37°C for lh. The enzyme-catalyzed release of A M C (aminomethylcoumarin) was quantified in a fiuorimeter at 380/450 nm. 5.2.10 Lipid Peroxidation ^ As previously described, oxidative damage on lipids was determined by the appearance of thiobarbituric acid-reactive substances (TBARS) like malondialdehyde (MDA) in each of the subcellular fractions [6]. 87 5.2.11 Pro tein Carbonyl Content Protein carbonyls (PC) were assayed as an index of oxidative modification of proteins [28]. Briefly, 50 pi of the fractions were added to an equal volume of 10% TCA, centrifuged at 6000g at 4°C for 5 min, and the supernatant discarded. The precipitated proteins were resuspended in 0.2% 2,4-dinitrophenyl hydrazine, and incubated for 1 h at 37°C. Subsequently, proteins were precipitated again with T C A , centrifuged, washed with ethanol:ethyl acetate, dissolved in 6 mM guanidine hydrochloride, and the absorbance measured spectrophotometrically at 370 nm. 5.2.12 Serum Lac tate Dehydrogenase As described previously, serum samples were isolated, kept at 4°C and processed within 24 hours for lactate dehydrogenase (LDH) using a commercially available kit (Sigma) [5]. 5.2.13 Statistical Analysis Values are means ± SEM. One-way A N O V A followed by the Tukey or Holm-Sidak test was used to determine differences between group mean values. The level of statistical significance was setatP<0.05. 88 5.3 Results 5.3.1 Mitochondrial Dysfunction Following Acute Diabetes STZ administration increased blood glucose within 24 hours, which was sustained for the next 3 days (CON 7.1±0.3 vs. DIA 23.4±0.7 mM; PO.05), with a reduction in plasma insulin levels (CON 3.3±0.4 vs. DIA 0.7±0.1, ng/ml; PO.05). One of the earliest diabetes-induced alterations is generation of ROS [4, 16]. To evaluate ROS levels, we utilized CM-H2DCFDA, a dye that fluoresces green when oxidized by ROS (Fig. 5.1, upper panel). Using confocal microscopy (upper panel) and fluorimetry (Fig. 5.1, lower panel), diabetic myocytes revealed an increase in green fluorescence, suggesting increased prevalence of ROS within these cells. Treatment with rotenone reduced ROS production in diabetic myocytes to control levels. Diabetic cells were more sensitive towards antimycin A and demonstrated greater fold increases in ROS compared to control myocytes. Additionally, incubation with N A C decreased the fluorescence to less than control levels. ROS generation in the diabetic heart may be regulated by changes in A\ym [29, 30]. Using confocal microscopy with JC-1, a dual fluorescent dye that dimerizes in the presence of intact Avj/m (with emission of red fluorescence), diabetic myocytes consistently demonstrated less red fluorescence than control cells (Fig. 5.2, top panel), suggestive of inadequate dimer formation, and reduction in Aipm [24]. Using a fluorimeter, diabetic myocytes demonstrated a lower red to green fluorescence ratio for JC-1 as compared to controls (Fig. 5.2, lower panel), implying mitochondrial damage. Incubation of some cardiomyocytes with mClCCP demonstrated almost no red fluorescence (data not shown). 89 5.3.2 Cardiac GSH And Apoptosis Following Acute Diabetes As reported previously, estimation of cardiac apoptosis revealed a nearly 4-fold rise in TUNEL positive nuclei in diabetic hearts (Fig 5.3, upper panel). As the most important cardiac antioxidant, in addition to being a key regulator of apoptosis [16, 17], reduced GSH was estimated in both the cytosolic and mitochondrial fractions. Interestingly, unlike the cytoplasmic pool, a depletion of mGSH (Fig 5.3, lower panel) was observed in 3-day diabetic rat hearts, implicating the importance of this specific pool of GSH in the cell suicide process. 5.3.3 GSH Modulation And Effects On Cardiac Apoptosis To further investigate the relationship between GSH and cardiac apoptosis, we used different GSH-depleting agents and quantified TUNEL positive nuclei in diabetic hearts. BSO, known to inhibit de novo synthesis of GSH, decreased cytoplasmic (Fig 5.4, upper left panel) but not mitochondrial GSH (Fig 5.4, upper right panel) in diabetic hearts, likely due to the fact that mitochondria lack gamma glutamyl cysteine synthase, the target enzyme for BSO [31]. In an attempt to also influence mGSH, D E M , an agent that conjugates GSH and depletes both mGSH and cGSH in vitro, was used [18, 19]. Like BSO, D E M induced a substantial drop in cGSH (Fig 5.4, upper left panel), but was ineffective in reducing mGSH in vivo (Fig. 5.4, upper right panel). Interestingly, although both BSO and D E M treatments brought about substantial reductions in cGSH, they were ineffective in augmenting cardiomyocyte apoptosis in the diabetic heart (Fig 5.4, lower panel). Although co-administration of BSO and D E M did not further decrease cGSH levels compared to D E M only, mGSH was almost undetectable with this treatment (Fig 5.4, upper right panel). In this setting, cardiac apoptosis was further aggravated compared to the untreated diabetic group (Fig 5.4, lower panel). In control animals similar treatment resulted in the depletion of only cytosolic (UTX CON 18.7±3.9 vs CON+BSO+DEM 9.7+3.1, pmol./mg protein; PO.05) but not mitochondrial GSH (UTX CON 9 0 7.2±1.9 vs CON+BSO+DEM 6.3±0.9, pmol./mg protein). Apoptotic rates remained unchanged compared to controls (data not shown). In an attempt to prevent cardiac GSH loss, animals were pre-treated with exogenous GSH prior to the induction of diabetes, with the treatment continuing until termination. Unexpectedly, GSH supplementation decreased cGSH levels (Fig 5.4, upper left panel). However, there was a robust amplification in mGSH content in diabetic hearts (Fig 5.4, upper right panel), together with a substantial drop in the apoptotic index to control levels (Fig 5.4, lower panel). 5.3.4 Cardiac Oxidative Stress And Caspase Activity Having implicated mitochondrial glutathione loss as a key participant in cardiac apoptosis, we investigated the outcome of GSH depletion on the initiation pathways of apoptosis. Within 3 days of diabetes, mitochondrial lipid (mTBARS) and protein (protein carbonyls, mPC) oxidation increased (Fig 5.5, upper panels), with no change in these parameters in the cytosol (data not shown). Estimation of caspase-9 (initiator caspase of the mitochondrial apoptotic pathway), caspase-8 (initiator caspase of the 'death receptor' mediated pathway) and caspase 3 (effector caspase of the above pathways) revealed an increase in caspase-9 and 3 activities only (Fig 5.5, lower panels), implicating predominance of the mitochondrial pathway in cardiac apoptosis at this early time point [32]. Treatment of diabetic rats with BSO or D E M increased cytosolic oxidative stress [TBARS (UTX DIA 23.8±2.5 vs DIA+BSO 36.9±4.9, DIA+DEM 36.2±3.6 nM MDA/mg protein; PO.05), protein carbonyl content (UTX DIA 98.5+21.1 vs DIA+BSO 212.6+16.3, DIA+DEM 208+28.8 A.U./mg protein; PO.05)] and caspase-8 activity (UTX DIA 19.7+2.5 vs DIA+BSO 33.1+2.7, DIA+DEM 36+3.2 F.U./mg protein; PO.05) compared to the untreated diabetic group. These treatments were ineffective in further aggravating either mitochondrial oxidative 91 stress (Fig 5.5, upper panels) or caspase-9 and 3 activities (Fig 5.5, lower panels). Co-administration of both BSO and D E M led to a depletion of mGSH, specific increases in mitochondrial oxidative stress (Fig 5.5, upper panels) and caspase-9 and 3 activities (Fig 5.5, lower panels), and no change in caspase-8 (data not shown) compared to untreated diabetic rats. Co administration of BSO and D E M in control animals was unable to influence either oxidative stress or cardiac caspase activities (Fig.5.5), suggesting that it is not simply depletion of GSH, but the added effect of the prevailing diabetic environment that promotes cell death in diabetic hearts. As severe glutathione depletion may precipitate necrotic death [5, 33], serum L D H was measured and found to be unchanged in all groups, suggesting that cell death was predominantly apoptotic in nature (data not shown). As GSH supplementation prevented cardiac apoptosis, reduced mitochondrial oxidative stress (Fig 5.5, upper panels) and activities of caspase 9 and 3 (Fig 5.5, lower panels), these data further confirm mitochondrial participation in cardiac apoptosis during acute diabetes [32]. 92 5.4 Discussion ROS are unstable oxygen containing molecules capable of modifying various cellular structures. Should the generation of ROS exceed their neutralization by antioxidants, oxidative 'stress' and premature cellular demise may occur [6, 16, 17]. In both clinical and experimental models of diabetes, oxidative stress is believed to be a key participant in causing cardiac injury [9]. In this study, using C M - H 2 D C F D A , we confirmed the presence of ROS such as H2O2 and OH" in cardiomyocytes isolated from acutely diabetic rats [34]. Given that under normal conditions, 1-2% of all oxygen consumed is converted to ROS by electron transfer in the mitochondrial ETC [35], we assessed the contribution of this pathway towards ROS generation in acutely diabetic rat hearts. In this regard, ubisemiquinone at complex III appears to be a major site for cardiac mitochondrial ROS generation [21]. Using rotenone, an inhibitor of the upstream complex I, we were able to decrease the excess ROS production in diabetic cardiomyocytes to control levels, likely as a consequence of inadequate electron transfer to the downstream complex III [21]. These results with rotenone precluded examination of the role of extra-mitochondrial-ROS-generating pathways in the acutely diabetic heart. As an alternate strategy, antimycin A, which inhibits electron transfer from complex III thereby prolonging the half-life of ubisemiquinone and increasing ROS, was used [21]. Antimycin A augmented ROS generation in both control and diabetic cardiomyocytes. However, the fold increase in ROS was greater in diabetic compared to control myocytes, supporting our conclusion that in diabetes, cardiac ROS is predominantly mitochondrial in origin. Amplification of mitochondrial ROS may result from changes in Ai)/m (crucial in maintaining electron flux in the ETC thereby minimizing ROS generation). Using confocal microscopy and measurement of the red (A\\im < -100 mV) to green (A\\im > -100 mV) fluorescence ratio of the dye JC-1, a loss of negative A\\im was observed in diabetic 93 cardiomyocytes, suggesting that ROS generation may indeed be related to changes in Au/m [25]. Other than its role in ROS generation, the cell's commitment towards apoptosis may become irreversible following collapse of Au/m [29]. In this regard, changes in Ai | / m induces the opening of 'channels' in the mitochondrial membrane [permeability transition (PT) pore] leading to release of low-molecular-weight proapoptotic factors from the mitochondria like cytochrome c and apoptosis-inducing factor. These in turn can specifically activate initiator caspases like caspase-9 and effector caspases like caspase-3, leading to apoptosis [29]. As caspase-9 and 3 were both increased along with a substantial rise in cardiac apoptotic death following only 3 days of diabetes, our data suggests that the 'mitochondrial' pathway of apoptosis may predominate at this early time point. In addition to changes in Ai[/m, PT pore formation has also been linked to insufficiency in antioxidants like glutathione, and oxidative stress [30] In this study, although cGSH remained unaltered, diabetic hearts were characterized by a decrease in mGSH, with a parallel increase in mitochondrial oxidative stress. Decrease in mGSH induces extensive ROS induced damage to mitochondrial membranes, which in turn causes a collapse of Aij/m and activation of the mitochondrial death pathway [30, 36] (Fig. 5.6). As similar results have also been reported in chronically diabetic rodent hearts [12, 13], these data imply that loss of mGSH and the consequent ROS induced damage, may be an essential cause for the diabetes-induced early apoptosis in diabetic rat hearts. To further examine the relationship between the different subcellular pools of GSH and apoptosis, we used BSO and D E M , agents that are known to deplete cellular GSH through different mechanisms. Administration of BSO, an inhibitor of y-GCS activity, resulted in a 30% decline in cGSH, with no change in mGSH or apoptosis. To overcome the resistance of mGSH to change, D E M , an agent that irreversibly inactivates all pre-formed subcellular pools 94 of GSH, was used [18, 19]. Although D E M decreased cGSH (almost 70%), it was also incapable of either reducing mGSH or increasing cardiac apoptosis compared to untreated diabetic hearts. In other studies, D E M treatment prior to ischemia/reperfusion was also incapable of worsening damage in rat hearts [37]. It has been suggested that although D E M administration decreases mGSH in vitro, conjugation of cellular GSH with D E M in vivo triggers a compensatory increase in y-GCS activity [38, 39], with newly formed GSH being transferred preferentially to the mitochondria. In this regard, during extensive cytosolic GSH depletion, the carrier system responsible for the transport of GSH may act to preserve mGSH levels [20, 31]. We speculated that abrogation of mGSH in vivo would only be possible if the compensatory rise in y-GCS activity is prevented following D E M administration. To achieve this goal, BSO and D E M were co-administered over 3 days to STZ-diabetic rats. Using this novel approach, we were successful in depleting mGSH in diabetic rat hearts. Under these conditions, mitochondrial lipid peroxidation, protein oxidation and apoptosis were exaggerated, suggesting an obligatory role for mGSH in minimizing oxidative stress and apoptosis in acutely diabetic rat hearts. As cGSH levels was similar to that seen with D E M alone, it is unlikely that the increased cytotoxicity was due to cGSH depletion. Exogenous GSH supplementation in pigs significantly increases myocardial GSH content and resistance to ischemia/reperfusion injury [40, 41]. Additionally, apoptosis and oxidative stress in isolated hearts perfused with high glucose were attenuated by addition of GSH to the perfusion buffer [42]. To prevent depletion of cardiac GSH following diabetes, rats were administered exogenous GSH. In acutely diabetic rat hearts, apoptosis and mitochondrial oxidative stress were averted by GSH supplementation. Measurement of GSH in the different pools revealed a selective increase in mGSH only, likely reflective of the need to protect the 95 diabetic mitochondria from oxidative damage. Unexpectedly, cGSH levels decreased, and the mechanism responsible for such an effect is presently unknown. Two independent pathways can cause cardiac apoptosis, with cellular GSH levels influencing both the death-receptor-mediated 'extrinsic' and the mitochondrial 'intrinsic' pathways [43, 44]. Caspase-8 and 9 are initiator caspases for the extrinsic and intrinsic pathways respectively [32]. In our study, induction of diabetes was only associated with caspase-9 activation, an effect that was amplified using BSO and D E M in combination. As similar results have been reported in diabetic kidney and neuronal cells [45, 46], and as BSO and D E M increased caspase-8 without intensifying apoptosis, our data points towards a major role for the mitochondrial apoptotic pathway in the acutely diabetic heart. In summary, at least following short-term diabetes, cardiac oxidative stress and apoptosis appear to have mitochondrial origins, with mitochondrial but not cytoplasmic GSH playing an essential role. As large-scale trials have failed to demonstrate any cardiovascular benefits using antioxidants like vitamin E and P-carotene in diabetic patients [47, 48], it is possible that: (i) the patients were in an advanced stage of the disease where damage reversal is difficult, (ii) agents like vitamin E only prevent lipid peroxidation, whereas ROS can affect a wide range of biomolecules, and (iii) the antioxidants were unable to reach therapeutic levels in the mitochondria where most of the ROS is generated. Thus, when using antioxidants, therapy should be initiated early, with scavenging of mitochondrial ROS being an essential property for preventing diabetic heart disease [35]. 96 5.5 Figures + A A + A A + R O T + N A C Fig. 5.1 Reactive oxygen species in diabetic cardiomyocytes To estimate ROS content, cardiomyocytes were incubated with C M - H 2 D C F D A for 30 minutes, and visualized under confocal microscopy at 400x magnification (upper panel). To examine the origin of ROS, cardiomyocytes were either left untreated or pre-treated with rotenone (ROT, 5 pg/ml) or antimycin A (AA, 100 ng/ml) for 1 hour and subsequently, D C F D A fluorescence was monitored using a fluorimeter (excitation-480 nm; emission-530 nm) (lower panel). At termination (45 mins), DCFDA fluorescence was expressed in fluorescence units (F.U.) per 7.5 x 10 cells. Pre-incubation of diabetic myocytes with 5 mM N A C (1 h) was used to evaluate the relationship between GSH and ROS. Results are means±SEM of 4 rats per group. *Significantly different from untreated control myocytes; "Significantly different from untreated diabetic, P < 0.05. 97 Control Diabetic 120 -| Fig.5.2 Mitochondrial dysfunction following acute hyperglycemia. Isolated control and diabetic myocytes were incubated for 10 mins with a mitochondrial membrane potential sensitive dye, JC-1. Red fluorescence due to JC-1 dimers was visualized under confocal microscopy at 600x magnification (upper panel). JC-1 fluorescence ratio (red to green) was quantitated using a fluorimeter (lower panel). Results are means±SEM of 4 rats in each group. * Significantly different from controls; P < 0.05. 98 25 ! 20 1 Cytosolic Mitochondrial Fig. 5.3 Effect of 4 days of diabetes on cardiac cell death and glutathione TUNEL assay was performed on formalin-fixed rat heart sections • and quantification of cardiac apoptosis was done using the imaging software 'Northern Eclipse.' The number of cardiomyocytes undergoing apoptosis is expressed per 106 nuclei (upper panel). The lower panel describes the subcellular glutathione status in control and diabetic hearts. Cytosolic and mitochondrial fractions were separated using 'digitonin fractionation,' and assayed for GSH sing an enzyme-recycling assay. Results are the means±SE of 6 rats in each group. Significantly different from control; P < 0.05. 99 Fig. 5.4 Effect of different treatments on GSH and cardiac apoptosis In an effort to deplete cardiac GSH, daily doses of BSO (10 mmol/kg) and/or D E M (4 mmol/kg) were administered for 3 days. For exogenous GSH supplementation, control rats were administered GSH (400 mg/kg) for 10 days prior to STZ administration, and the treatment continued for a further 4 days. Following the above treatments, rats were killed, hearts isolated, and used either intact, or after isolation of cardiomyocytes. The upper panel depicts myocardial GSH in sub cellular fractions from diabetic hearts, whereas the lower panel represents cardiac apoptosis as determined by T U N E L staining. Results are the means±SE of 6 rats in each group. *Significantly different from untreated diabetic; "Significantly different from BSO; P < 0.05. 100 i 1 UTX C O N Hi UTX DIA vma DIA+BSO Fig. 5.5 Mitochondrial oxidative stress and caspase activities in control and diabetic hearts Control hearts were either left untreated or treated with BSO+DEM. Diabetic rats were either left untreated, or treated with BSO, D E M , BSO+DEM or GSH. Mitochondrial oxidative stress was estimated by lipid peroxidation (mTBARS) and protein carbonyl content (mPC) (upper panels). The lower panels represent the quantitative estimation of cardiac caspase activities as determined using the corresponding fluorescent caspase specific substrates, DEVD-A M C (for caspase 3) and L E H D - A M C (caspase-9). 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High glucose evokes an intrinsic proapoptotic signaling pathway in mesangial cells. Kidney Int 2005; 67:82-93 47. Lonn E, Yusuf S, Hoogwerf B, Pogue J, Yi Q, Zinman B, et al. Effects of vitamin E on cardiovascular and microvascular outcomes in high-risk patients with diabetes: results of the HOPE study and MICRO-HOPE substudy. Diabetes Care 2002; 25:1919-27 48. MRC/BHF Heart Protection Study of antioxidant vitamin supplementation in 20,536 high-risk individuals: a randomised placebo-controlled trial. Lancet 2002; 360:23-33 108 6. Brief Episode Of STZ-Induced Hyperglycemia Produces Cardiac Abnormalities In Rats Fed A Diet Rich With co-6 PUFA 6.1 Introduction Obesity is a common nutritional condition in the Western world with attendant disorders like hypertension, insulin resistance and Type 2 diabetes. Over time, patients who develop diabetes are susceptible to heart disease, a leading cause of death in these patients [1]. Factors that largely account for this increased incidence of cardiovascular dysfunction include large and small coronary vessel disease [1]. However, a significant number of diabetic patients continue to suffer from left ventricular dysfunction and clinically overt congestive heart failure in the absence of vascular defects [1, 2]. Probable candidates to explain this heart disease include autonomic abnormalities, metabolic disorders, abnormal contractile protein and enzyme function, [1], and more recently apoptosis, a regulated, energy dependent, cell suicide mechanism [3, 4]. Given that animal models of diabetes [like the Zucker Diabetic Fatty rat [5] or the S T Z diabetic rat [6] also display progressive myocardial abnormalities in the absence of vascular changes but in the presence of accelerated cell death, it is possible that this loss of cells progressively leads to interstitial fibrosis, myocardial hypertrophy, and eventual contractile impairment. Increase in plasma glucose during diabetes has been suggested to predispose cardiomyocytes to death by apoptosis [3]. However, in diabetes, cardiac energy production is almost exclusively from fatty acids (FA), that are supplied in excess to the heart and have been implicated in causing cardiac l ipid overload and cell death [7, 8]. Thus in vitro, palmitic acid, a saturated fatty acid causes intracellular accumulation of ceramide and reactive oxygen species 109 (ROS) in cardiomyocytes and can precipitate apoptotic cell death, either in the presence or absence of hyperglycemia [8, 9]. Recently, we have demonstrated that feeding a 20% w/w palm oil diet (rich in palmitic acid) to diabetic rats enhances cardiac apoptosis in vivo [10]. The majority of studies that have looked at the relationship between lipotoxicity and cardiovascular complications of diabetes have utilized lard or other sources of saturated dietary fat rich in palmitic acid [10-12]. However, in humans, increased awareness of obesity and its cardiovascular complications have led to an indiscriminate substitution of atherogenic saturated cooking fats with "heart-friendly" refined vegetable oils like sunflower o i l , rich in co-6 polyunsaturated fatty acids (co-6 P U F A ) [13]. In several studies, co-6 P U F A conferred protection against arrythmias [14] and coronary artery disease [15], and at least in human primary fibroblasts and Ley dig cells, prevented apoptosis [16, 17]. Substantial data exists on the role of saturated F A in the development of cardiac apoptosis and ultimately cardiomyopathy in animal models of diabetes. However, in the light of current dietary recommendations, the impact of the more relevant co-6 P U F A on cardiac function and morphology, especially during diabetes, is not clearly defined. We hypothesized that given the role of saturated fatty acids in accelerating cardiac apoptosis following diabetes, switching to a co-6 P U F A rich diet may well be protective against cell death. Instead of preventing cardiac cell death, our data for the first time suggests that during diabetes, along with other morphological and functional abnormalities, co-6 P U F A provision converts the mode of cellular demise to necrosis, an alternate form of cell death. 110 6.2 Materials And Methods 6.2.1 Experimental Animals The investigation conforms to the guide for the care and use of laboratory animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1985) and the University of British Columbia. Previous studies have established that feeding of 20% w/w sunflower oil for 4 weeks increases the co-6 PUFA content of circulating lipoproteins, [18] a major source of FA to the heart. To investigate potential beneficial effects of co-6 PUFA, male Wistar rats (220-240 g) were first fed a co-6 PUFA diet [(AIN)-76A supplemented with 20% w/w sunflower oil, Table 6.1] (Research Diets Inc.) for 4 weeks to increase cardiac PUFA content (PUFA Control; PC). Energy contribution from fats in this diet was 40% of the total energy intake, similar to human consumption [19]. Some PUFA fed animals were then given a moderate dose of streptozotocin (STZ; 55 mg/kg, i.v.), a selective beta cell toxin [20]. With this dose of STZ, there is only a partial destruction of fi-cells with ~50%> reduction in plasma insulin levels and stable hyperglycemia (PUFA Diabetic; PD) within 24 hours [21]. An acute time period (4 days) was chosen based on previous studies that demonstrate peak incidence of cardiac apoptosis after 3 days of STZ-diabetes in normal chow fed rats [4]. PUFA feeding was continued subsequent to STZ administration and after 4 days of diabetes, rats were anesthetized with sodium pentobarbital (65 mg/kg, i.p.), blood collected, and hearts excised for histological, morphological and biochemical analysis. Additional control (Normal Control, NC) and STZ (Normal Diabetic, ND) groups fed normal laboratory chow [5% corn oil; PMI Feeds, Table 1] were utilized to compare the effects of co-6 PUFA feeding and diabetes per se. 6.2.2 Separation And Characterization Of Cardiac Lipids Total cardiac lipids were extracted and solubilized in chloroform:methanol:acetone:hexane (4:6:1 :lv/v/v/v). Separation of cardiolipin, triglycerides and FA was achieved using HPLC 111 (Waters 2690 Alliance H P L C , Milford, M A ) equipped with an auto-sampler and column heater. F A were quantified as their respective methyl esters using heptadecaenoic acid (17:0) as the internal standard with a Varian 3400 G L C equipped with a flame ionization detector, a Varian Star data system, and a SP-2330 capillary column (30 m x 0.25 mm internal diameter) (Supelco, Bellefonte, P A ) [22]. Value of cardiac F A , triglycerides and cardiolipin were expressed as p,g/mg protein. 6.2.3 Cardiac Caspase-3 Activity To investigate the contribution of the apoptotic pathway, caspase-3 activity, the prime effector of cardiomyocyte apoptosis, was estimated (Molecular Probes). Caspase-3 is a cysteine containing protease crucial in the 'execution' phase of apoptosis and breaks down the contractile proteins in the heart [23]. Briefly, ventricular tissue (200 mg) was placed in cell-lysis buffer and homogenized (4°C). Subsequently, the suspension was centrifuged at 12,000g for 3 min, and the supernatant separated and reacted with aminomethylcoumarin (AMC)-derived substrate Z - D E V D - A M C . Caspase-3 activity is proportional to A M C released and is expressed as u M A M C released per mg protein. 6.2.4 Cardiac Necrosis Necrosis leads to a loss of cell membrane integrity with release of lactate dehydrogenase (LDH) into the serum [24]. To verify necrosis, serum L D H was estimated using an appropriate kit (Sigma). However release of necrosis does not necessarily imply cardiac necrosis. Thus to verify cardiac necrosis, sections were evaluated histologically using Masson's Trichrome stain [25]. Briefly, ventricular tissue was fixed in 10% formalin, and sectioned at 5 u M . Sections were deparaffinized in xylene, rehydrated using various grades of ethanol and refixed in Bouins fluid for l h at 56°C. Subsequently, sections were stained, dehydrated with acetone, cleared 112 with xylene, and mounted. Positive controls were generated by injection of isoproterenol (50 mg/kg; sc) to a normal chow fed rat, and the heart removed after 48 h [26]. 6.2.5 Estimation Of G S H Cardiac GSH content was measured from frozen, ground tissue using a commercially available kit (Cayman Chemicals). GSH was reacted with 5,5'-dithiobis-2-nitrobenzoic acid (DTNB) to produce a yellow colored 5-thio-2-nitrobenzoic acid (TNB). GS-TNB, produced in parallel, is recycled back to GSH by glutathione reductase to produce more TNB. Rate of TNB production is directly proportional to this recycling reaction, which is in turn directly proportional to the total concentration of GSH in the sample. Oxidized glutathione (GSSG) was estimated using 2-Vinylpyridine [27]. The difference between total GSH and GSSG gave the value of the reduced GSH. Protein assay was performed according to the Bradford method. Values are expressed as nM GSH per mg protein. 6.2.6 A T P Assay ATP from heart tissue was extracted using 2.5% T C A , pH adjusted to 7.8 by IM K H C O 3 and ATP (nmol/mg protein) measured by a luciferase-luciferin assay kit (Sigma). 6.2.7 Electron Microscopy. Morphological evaluation of hearts was carried out using transmission electron microscopy. Briefly, left ventricular tissue was fixed in 1.5% glutaraldehyde and paraformaldehyde, cut into small blocks (~1 mm3) and fixed for 8 h at 4°C. After washing, tissue was post fixed with 1% osmium tetraoxide and further treated with 1% uranyl acetate and dehydrated using increasing concentrations of ethanol (50-100%). Blocks were embedded in molds, polymerized, and sectioned at ~100 nm. Sections were stained with 1% uranyl acetate and Reynolds lead citrate. Images of the longitudinal-sections were obtained with a Hitachi H7600 electron microscope. 113 6.2.8 H e a r t F u n c t i o n A n d Subs t ra te O x i d a t i o n Hearts from halothane (2-3%) anesthetized rats were perfused in the working mode with modified Krebs-Henseleit buffer (including 1.0 mM [9,10-3H] palmitate pre-bound to 3% BSA, 5.5 mM [U- 1 4C] glucose, 2.0 mM calcium, and 100 U/L insulin.) at a preload of 11.5 mmHg as described previously [28]. An afterload of 80 mmHg was kept for the first 30 min. During this time, samples of perfusate and hyamine hydroxide were taken every 10 min for measurement of glucose and FA oxidation. Subsequently, the afterload is clamped off causing the peak systolic pressure to increase to -135 mmHg, and the perfusion continued for another 20 minutes. Heart function [rate pressure product (RPP)-heart rate, [mmHg. min" 10"3] was measured using a Direcwin physiograph (Raytech). 6.2.9 S e r u m M e a s u r e m e n t s Serum samples were stored at -20°C until assayed. Diagnostic kits were used to measure glucose (Sigma), insulin and leptin (Linco) and non-esterified fatty acids (Wako). 6.2.10 S ta t i s t i ca l A n a l y s i s . Values are means ± SEM. One-way A N O V A followed by the Tukey or Holm-Sidak test was used to determine differences between group mean values. The level of statistical significance was set at .PO.05. Statistical analysis 114 6.3 Results 6.3.1 General Characteristics Of Experimental Animals Table 6.2 describes the general characteristics of the experimental groups. There was no difference in food intake (data not shown) between rats fed co-6 PUFA diet or normal chow for 4 weeks. However, co-6 PUFA feeding increased body weight compared to normal chow. When compared to fed NC rats, PC animals had higher circulating T G and increased serum insulin (likely as a consequence of insulin resistance), but no overt hyperglycemia. STZ precipitated loss of circulating insulin and caused overt hyperglycemia in both ND and PD groups. Interestingly, unlike ND (pre-STZ, 361±9; post-STZ, 358±9, g), diabetes in PUFA animals was associated with a profound loss of body weight (pre-STZ, 450±11; post-STZ, 393±11, g). This loss in body weight could not be attributed to any change in food or fluid intake but could be a result of excessive lipolysis and loss of adipose tissue mass with subsequent increases in both serum FFA and T G in PD animals. Additionally, dietary co-6 PUFA for 4 weeks increased serum leptin levels compared to the NC group, whereas diabetes reduced leptin in both ND and PD groups, co-6 PUFA feeding induced a 6-7 fold increase in total cardiac FFA and a 40-80 fold increase in total cardiac T G in both PC and PD groups, and indicated a cardiac lipid overload in these animals.. 6.3.2 Free PUFA Content In The Heart Although several fatty acid classes (e.g., saturated and monounsaturated FA) were measured, only the cardiac PUFA content in the free form are described in Fig. 6.1 (as per the focus of this study). Feeding 20% w/w sunflower oil successfully magnified both co-6 (mainly linoleic and arachidonic acid) and co-3 (mainly eicosapentaenoic acid and docosahexaenoic acid) PUFA content in the heart. However, the increase in co-6 PUFA was substantially greater than the 115 increase in co-3 PUFA, and was reflected in an approximately 2 fold higher co-6/co-3 PUFA ratio in the PC group. Diabetes had no further effect on either the co-6 or co-3 PUFA content. 6.3.3 Cardiac Cell Death Previous studies have associated cardiac lipid overload to cell death via apoptosis. To determine cardiac apoptosis following co-6 PUFA feeding, caspase-3 activity, crucial for myocardial apoptosis, was estimated and determined to be unchanged in both PC and PD groups compared to NC (Fig. 6.2). In ND, the caspase-3 like activity was almost 2.5 fold higher than control and indicates acceleration of apoptotic cell death, as has been previously described [4]. As apoptosis can proceed in a caspase-3-independent manner, TdT mediated dUTP nick end labeling (TUNEL) on myocytes was also performed and found to be increased in ND but insignificant in PD hearts (data not shown). Absence of apoptosis may not signify lack of cell death as cell demise may also progress via a necrotic pathway. Incidence of necrosis by measurement of serum L D H revealed that only PD had higher circulating levels of this enzyme marker (Fig. 6.3, bottom right). However, as serum L D H alone does not substantiate the incidence of cardiac necrosis, Masson's Trichrome staining was performed on cardiac ventricles [25]. In hearts treated with isoproterenol, a P-agonist, severe disruption of the contractile apparatus (solid arrow; Fig. 6.3E) together with focal necrosis (indicated by grayish discoloration instead of the more usual deep red coloration demonstrated by intact muscles) (hollow arrow; Fig. 6.3E) was observed. No such morphology was evident either in NC, ND or PC hearts (Fig. 6.3, A, B or C; magnification, 400X). Following diabetes, PD hearts showed necrotic changes, which on higher magnification (600X) demonstrated extensive grayish patches and focal necrosis (hollow arrow; Fig. 6.3D) with hypercontracted muscle fibers (solid arrow; Fig. 6.3D) characteristic of contraction band necrosis [29]. 116 6.3.4 Cardiac GSH Factors like GSH levels, which dictate the redox environment of the cell and also the mode of cell death [30], were also measured. PUFA feeding was associated with a significant decrease in GSH. Interestingly, diabetes for 4 days in normal chow fed rat hearts could not decrease cardiac GSH, whereas superimposition of diabetes in PUFA fed animals led to a further decrease in cardiac GSH levels (Fig. 6.4, top panel). Although cardiac GSSG was marginally increased in ND compared to NC hearts, this difference did not reach significance. PUFA feeding had no effect on GSSG. However, induction of diabetes in PD animals increased cardiac GSSG almost 3-fold (Fig. 6.4, middle panel). Evaluation of the GSH/GSSG ratio revealed that PD hearts had the lowest values, indicating substantial oxidative stress in this group (Fig. 6.4, lower panel) [31]. 6.3.5 Cardiolipin And Mitochondrial Morphology Cardiolipin is a mitochondrial membrane phospholipid that is essential for optimal functioning of oxidative phosphorylation enzyme complexes, and ATP generation [32]. Diabetes in both normal chow and high fat groups did not alter cardiolipin levels compared to the respective controls (Fig. 6.5, top panel), co-6 PUFA feeding induced a profound loss of cardiac cardiolipin levels (Fig. 6.5, top panel) together with a drop in total ATP (NC 8.9±0.6, ND 8.1±0.8 and PC 6.9±0.6 vs. PD 5.1 ±0.2; nmol ATP/mg protein; P<0.05). As changes in cardiolipin have also been associated with mitochondrial structural alterations, we scrutinized mitochondrial morphology in all of the above groups. Figure 6.5 (lower panel, A) depicts a mitochondrion with double membrane and lamellar cristae that are typical in NC, ND or PC hearts. A novel observation in this study was that abnormal condensed mitochondria were observed, but only in the PD group (Fig. 6.5, lower panel, B). 117 6 .3 .6 Glucose And Palmitate Oxidation As condensed mitochondria suggests lack of energy production, glucose and palmitate oxidation (Fig. 6.6) in the isolated heart were studied in parallel with heart function. At low afterload (80 mm Hg), rates of oxidation of exogenous glucose decreased in ND and PC with a parallel increase in palmitate oxidation in these groups. Interestingly, in PD hearts, although glucose oxidation dropped further compared to both ND and PC hearts, palmitate oxidation remained unchanged. To meet the energy demand at high afterload (135 mm Hg), all groups except PD increased their glucose oxidation. Regarding F A oxidation, only NC was able to increase palmitate oxidation suggesting that in the ND, PC and PD groups, FA oxidation was already operating at its maximum at low afterloads. 6 .3 .7 Electron Microscopy And Ultrastructural Morphology In the milieu of increased intracardiac fatty acid levels (Table 6.2) and in the absence of any further increase in F A oxidation, we hypothesized that intracardiac T G will build up in PD hearts. Figure 6.7 (a-d) demonstrates ultrastructural morphology of hearts isolated from the different groups. Expectedly, although lipid droplets were visible in both co-6 PUFA fed groups, only PD hearts demonstrated abundant lipid-like vacuoles (L). Additionally, NC, ND and PC hearts showed highly packed myofibrils separated by rows of mitochondria. Myofibrillar sarcomeres in these two groups exhibited clear Z-lines (indicated by hollow arrows) whereas PD hearts demonstrated more diffuse Z-lines indicating myofibrillar damage (indicated by hollow arrows). These findings are consistent with necrosis present in the PD hearts (Fig. 6.3), and could precipitate an early contractile defect. 6.3 .8 Estimation Of Heart Function Cardiac mechanical function measured as rate pressure product (RPP) of the different groups is summarized in Fig. 6.8. There were no differences in RPP between the groups at normal workload (80 mmHg). Subsequent to an increase in workload to 135 mmHg, RPP increased in 1 1 8 a l l groups. C o m p a r e d to N C , neither fat feeding nor STZ-diabetes affected heart funct ion i n response to this h i g h w o r k l o a d . Interestingly, super imposi t ion o f diabetes for 4 days i n P D rats induced the least increase i n cardiac funct ion. 119 6.4 Discussion Numerous clinical and epidemiological reports have confirmed that human diabetics are particularly susceptible to cardiomyopathy, independent of vascular disease and recent evidence points towards cell death as a potential contributing factor [1]. Indeed, in a recent study, we detected a 5-fold increase in apoptosis in the diabetic heart using TUNEL assay, and further validated this observation by identifying an increase in caspase-3 activity, the proteolytic enzyme involved in the execution of apoptosis [10]. In this study, an increase in cardiac caspase-3 activity was also observed in ND rats. Despite this augmented apoptosis in the diabetic heart, the number of cells affected in this and other studies [3, 4] is relatively minor and may explain the lack of functional changes at this early time point. However, given that apoptotic cells are cleared rapidly by phagocytes [23], and in the absence of any significant regeneration of the adult myocardium, low cell death rates over chronic periods of hyperglycemia may still result in progressive loss of myocytes and contractile failure [23]. Interestingly, compared to ND, superimposition of diabetes in 20% w/w PUFA fed rats did not change caspase-3 activity or TUNEL positive myocytes, and questions the prevalence of apoptosis in this group. At least in cell lines, co-6 PUFA is also known to block apoptosis [16, 17]. Overall, our data suggest that rather than the amount; it is the type of FA that controls lipotoxic apoptosis [33]. Indeed, feeding a 20% w/w palm oil diet [rich in palmitic acid that is converted to pro-apoptotic ceramide [33]] increases apoptosis in control hearts, a process that was further aggravated following diabetes [10]. Although apoptosis is the predominant form of cellular demise, the diabetic heart is also characterized by necrosis, an 'accidental' form of cell death [3]. Assessment of serum markers or histochemical detection of necrosis did not reveal any necrotic change in NC, ND or PC hearts. However, PD animals demonstrated a rise in serum L D H , and severe cardiac necrosis as evidenced by light microscopy and appropriate staining. Furthermore, ultrastructural 120 evidence demonstrated diffuse Z-lines suggesting myofibrillar damage in PD hearts, indicative of necrotic changes. In an effort to determine the elements that induce necrosis, we measured the cardiac FFA species and determined that sunflower oil feeding increased cardiac co-6 PUFA, predominantly linoleic acid. In clinical studies, accumulation of linoleic acid has been strongly associated with an increased inflammatory response in endothelial cells, in addition to an augmented risk of myocardial infarction and necrosis [34]. Additionally in vitro, linoleic acid precipitates necrosis via pro-inflammatory pathways in various cell types [35, 36]. Oxidative stress has also been implicated in the etiology of several lipotoxic and diabetic complications [2, 8], and can arise as a result of an imbalance between the production of free radicals and its neutralization by antioxidants like GSH [2, 31]. In ventricular myocytes, under conditions of oxidative stress, GSH is converted to GSH disulfide (GSSG) and the GSH/GSSG ratio decreases (i.e., a high GSH/GSSG ratio maintains a normal reducing intracellular environment). Interestingly, co-6 PUFA feeding induced a loss of reduced GSH that decreased further with diabetes, an effect that could be directly linked to an increased conversion of reduced GSH to GSSG under conditions of augmented oxidative stress [31]. A major functional impact of a decreased GSH/GSSG ratio is oxidation of regulatory proteins at cysteine residues [30, 31]. For caspase activation and propagation of the apoptotic pathway, an intact cysteine moiety is necessary, that can easily be oxidized under increased oxidative stress and GSH depleted conditions [37]. Overall, our data suggests that increased cardiac co-6 PUFA content and depleted cardiac GSH operate in concert to allow a 'switch' to necrosis in PD hearts. Compromised myocardial energy status may also dictate necrotic death. Cardiolipin, a mitochondrial membrane phospholipid essential for optimal functioning of oxidative phosphorylation enzyme complexes and ATP generation [32], was estimated along with total 121 myocardial ATP levels. In myocardial ischemia, disappearance of total cardiolipin resulted in loss of OXPHOS activity [38] and could explain the drop in mitochondrial ATP synthesis. Total cardiolipin decreased almost 6-fold following co-6 PUFA feeding with a significant drop of ATP only in PD hearts, that could have contributed towards cardiac necrosis [39]. Changes in cardiolipin have been associated with mitochondrial morphological alterations. In Barth Syndrome, a genetic disorder characterized by dilated cardiomyopathy, a lack of cardiolipin in mitochondria has been described together with abnormal mitochondria with concentric cristae [40]. A novel observation in this study was that analogous to Barth Syndrome, co-6 PUFA feeding for 4 weeks together with 4 days of hyperglycemia led to similar changes in some mitochondria. Interestingly, abnormal condensed mitochondria have been recently recognized in skeletal muscle and cardiac atrial neurons from diabetic patients [41]. Given that both control and diabetic PUFA groups showed similar changes in cardiolipin, it could be assumed that the altered mitochondrial morphology in PD animals is a consequence of an additional effect of hyperglycemia i.e., when hyperglycemia sets in, there is augmented oxidative stress and extensive damage to the mitochondria, already compromised by depleted cardiolipin levels [42]. Structural damage to the mitochondria is associated with derangements in fuel utilization, and could lead to cardiac contractile impairment. Compared to glucose, FA's are the preferred substrate consumed by the heart, providing approximately 70% of its energy requirements. With hyppinsulinemia or insulin resistance that characterizes ND and PC respectively, glucose utilization is compromised and the heart is forced to switch to using higher levels of FA [7]. Interestingly, PD animals, characterized by a combination of insulin resistance initially followed by eventual hypoinsulinemia after STZ, demonstrated a precipitous drop in cardiac glucose utilization with no further increase in F A oxidation. Despite these changes in substrate preference, heart function at low afterloads remained unchanged. At higher afterloads, both 122 glucose and FA oxidation increased in the NC group. In ND and PC hearts, the only increase observed was that of glucose oxidation as presumably, F A oxidation was already operating at its maximum. PD was the only group that failed to increase its glucose oxidation in response to a higher energy demand, an aspect that could have contributed to the drop in cardiac function. Interestingly, when diabetic hearts are perfused with both glucose and FA, addition of dichloroacetate, a stimulator of glucose oxidation to the perfusion buffer improved heart function [43]. More recently, shifting substrate utilization from FA to glucose using trimetazidine improved left ventricular function in patients with diabetes and ischemic cardiomyopathy [44], and in transgenic db/db mice that overexpress GLUT-4 glucose transporters, glucose metabolism and contractile function were normalized [45]. Our study provides novel evidence that unlike other models of diabetic cardiomyopathy that exhibit cardiac dysfunction only after chronic hyperglycemia, co-6 PUFA feeding coupled with only four days of diabetes can precipitate metabolic and contractile abnormalities in the heart. The heart and other non-adipose tissues have inadequate ability to handle excess lipids and accumulation of T G has been proposed to impair cardiac function. [5, 46]. In our study, co-6 PUFA feeding substantially increased cardiac FA in both PC and PD groups, but only PD hearts demonstrated a considerable increase in lipid droplets. Given that fatty acid oxidation is likely operating at maximum in PC hearts, the excess circulating fatty acids in PD is likely channeled towards T G synthesis and could explain the increase in lipid droplets in these hearts. Interestingly, in Zucker fatty rats, when glucose utilization is compromised (as during fasting), increased lipid availability is associated with cardiac lipid accumulation [46]. At present it is unclear why PD hearts were unable to increase its fatty acid oxidation even in the presence of excess fatty acids. In this regard, leptin, a liporegulatory hormone increases FA entry and T G storage in adipose tissue, but activates fatty acid oxidation and decreases T G accumulation in 123 the heart [47]. Although leptin was high in PC animals, PD animals demonstrated low insulin and leptin levels, factors that could have contributed towards FA being channeled from oxidation to increased cardiac T G assembly. It should be noted that although co-6 PUFA rich diet was utilized as a beneficial regimen in this study, sunflower oil per se induced obesity and insulin resistance like other saturated fat diets. However, .insulin resistance, hyperinsulinemia and impaired glucose tolerance that characterize obese humans, is usually associated with transition to frank diabetes occurring when secretion of insulin is no longer able to overcome insulin resistance and impaired glucose disposal. In rodents fed high fat diets, insulin resistance does not progress to hyperglycemia in the absence of genetic defects [20]. Thus, our current model in which a low dose of STZ was used to precipitate overt hyperglycemia following insulin resistance is a close approximation of the human condition where there is a gradual evolution of insulin resistance to hyperglycemia. In summary, chronic caloric excess of co-6 PUFA when coupled with acute diabetes of only 4 days precipitated mitochondrial abnormalities, a steep drop in GSH, altered substrate utilization and myocardial T G deposition. Given that these hearts also demonstrated necrosis and extensive myocardial cell loss, a feature that is predominant only in chronic diabetes [3, 4, 6, 48], our data suggests that this mode of cell death in PUFA fed diabetic hearts is an important factor in accelerating diabetic cardiomyopathy. Although these effects of co-6 PUFA in the diabetic animal would seem contrary to accepted belief as being beneficial, in countries like Israel, with high dietary co-6 PUFA consumption, there is an excessive incidence of obesity, insulin resistance, hypertension, and Type 2 diabetes [49]. Thus, the dilemma with recommending vegetable oils like sunflower oil as a source of PUFA is that it contains high levels of co-6 PUFA but negligible amounts of co-3 PUFA, the primary candidate for conferring cardiovascular benefit. Given the concurrent scarcity of fish oil in the Western diet, the main source of co-3 PUFA, the co-6 /co-3 PUFA ratio has amplified from a traditional balance of 1:1 124 to an astounding 15:1 to 20:1 in the modern diet [50] and thus advocating diets rich in co-6 PUFA to diabetic patients could accelerate the impairment of myocardial contractility. 125 6.5 Tables and Figures Table 6.1 Composition of experimental diets Caloric Composition PUFA diet Normal diet g% g% Protein 24.1 23.4 Carbohydrate 44.9 49.9 Fat 20.7 4.5 Fibre 6.0 8.3 kcal/g 4.6 4.0 Fatty Acids (g%) Saturated 1.9 1.5 Monounsaturated 3.8 1.6 co-6 PUFA 10.6 1.2 co-3 PUFA 0.1 0.3 Ingredients PUFA Diet: Casein, 80 mesh, DL-Methionine, LoDex 10, Corn Starch, Sucrose, Cellulose, BW200, Corn Oil, Sunflower Oil, Mineral Mix,1" Vitamin Mix,* Choline Bitartrate Normal Diet: Ground corn, soybean meal, beet pulp, fish meal, wheat, cane molasses, ground oats, Mineral Mix,T- Vitamin Mix* Mineral Mix- Di-Calcium Phosphate, Magnesium Oxide, Potassium citrate, Potassium Sulfate, Sodium Chloride, Chromium KSO4.I2H2O, Cupric Carbonate, Potassium Iodate, Ferric Citrate, Manganous Carbonate, Sodium Selenite, Zinc Carbonate * Vitamin Mix- Vitamin A, Vitamin D 3 , Vitamin E, Menadione Sodium bisulfite, Biotinl%, Cyancocobalamin 0.1%, Folic Acid, Nicotinic Acid, Calcium Pantothenate, Pyridoxine-HCl, Riboflavin, Thiamin HC1 126 Table 6.2 General characteristics of experimental animals NC ND PC PD Body weight (g) 381 ± 9 358 ± 8.7 485 ± 14f 393 ± 1 1 * Cardiac FFA (pg/mg protein) 1.9±0.3 1.7±0.2 13 .9±1 ! 1 5 . 5 ± l t Cardiac T G (pg/mg protein) 0 . 0 1 ± 0 . 0 1 0.02 ± 0.0 0.5 ± 0.21" 0.8 ± 0.2* Serum parameters Glucose (mmol/1) 8.6 ± 0 . 2 2 4 . 2 ± 0 . 8 T 11.1 ± 0 . 5 2 3 . 8 ± 0 . 5 * t FFA(mmol/l) 0.5 ± 0 . 1 0.6 ± 0 . 1 0.4 ± 0 . 1 1.8 ± 0 . 8 * Triglyceride (mmol/1) 0.8 ± 0 . 1 1.2 ± 0 . 2 4 . 5 ± 1 . 2 t 11.8 ±2 .9* Insulin (ng/ml) 2.8 ± 0.4 0.8 ± 0.31" 5 . 5 ± 1 . 1 T 1.5 ± 0 . 6 * Leptin (ng/ml) 8.0± 0.8 1 . 9 ± 0 . 4 t 12.7± 1.6T 1 .2±0 .2* t Values are means ± SE for 6 animals in each group. Animals were maintained on normal chow (NC) or a co-6 PUFA (PC) rich diet for 4 weeks prior to diabetes induction (55 mg/kg STZ). Diabetic animals (ND, PD) were kept for 4 days before being killed. Values are those obtained before death and were from fed animals. * Significantly different from PC, P. < 0.05. ^Significantly different from NC, P < 0.05. ^Significantly different from all groups, P < 0.05; FFA, free fatty acids; T G , triglycerides. 127 Fig. 6.1 Effects of sunflower oil feeding and diabetes on cardiac PUFA levels Cardiac FFA's were extracted with chloroform:methanol:acetone:hexane solvent and were converted to their respective methyl esters, and separated by gas chromatography. Results are the means ± SE of 4 rats in each group. Top panel: Total cardiac co-6 PUFA levels (mainly linoleic and arachidonic acid). Middle Panel: Total cardiac co-3 PUFA levels (mainly eicosapentaenoic acid and docosahexaenoic acid). Lower panel: Ratio of co-6 and co-3 PUFA in the heart. * Significantly different from normal chow fed groups; P < 0.05. 128 2.0 0) o Q . O I E •B <u in C D JD <u i — O < 1.5 1.0 o C D C L in C D O 0.5 Control Diabetic 0.0 Normal Diet co-6 P U F A Diet Fig.6.2 Quantitative estimation of cardiac caspase-3 activity Heart tissue was placed in cell-lysis buffer and homogenized. Subsequently, the suspension was centrifuged, supernatant separated and reacted with Z - D E V D - A M C substrate. Results are the means + SE of 4 rats in each group. * Significantly different from all other groups; P < 0.05. 1 2 9 Momwl Otex «> * P U f A Diet Fig. 6.3 Effect of 4 days of diabetes on cardiac cell death in animals fed a moderately high PUFA diet for 4 weeks Heart sections were treated with Masson's Trichrome stain. Solid arrows depict severe disruption of the contractile apparatus (hypercontracted muscle and thickened fibres) whereas the hollow arrows indicate focal necrosis (indicated by grayish discoloration) in positive control ^Isoproterenol; E , 400x) and P D hearts (D, 600x). Cardiac necrosis was absent in N C (A, 400x), N D (B, 400x) and P C (C, 400x) hearts. Bottom right panel: Serum was collected at death and L D H activity measured with an appropriate kit. Results were expressed as a percentage of normal chow fed controls and are the means ± SE of 4 rats in each group. * Significantly different from all other groups; P < 0.05. 130 Fig. 6.4 Status of myocardial glutathione system as estimated by enzyme-recycling assay Upper panel: Reduced GSH Middle panel: Oxidized GSSG. Lower panel: Cardiac GSH/GSSG ratio. Results are the means ± SE of 4 rats in each group. ^Significantly different from normal chow fed control, * Significantly different from all groups. P < 0.05. 131 12 ' i Control Normal Diet co-6 P U F A diet Fig . 6.5. Biochemical and ultrastructural changes in cardiac mitochondria following PUFA feeding and diabetes Top Panel: Separation of total cardiolipin was achieved by HPLC. Results are the means ± SE of 4 rats in each group. * Significantly different from normal chow fed groups, P < 0.05. Lower Panel: Representative electron micrographs of myocytes showing normal (A) and condensed (B) mitochondrion. Some condensed mitochondria were only observed in PD hearts. The scale bar for the micrograph in A (100 nm) also apply to B. 1 3 2 2 0 0 0 1 5 0 0 H. 1 0 0 0 H Af te r load - 8 0 m m H g 'cu s "D .O) C E o E c c o ns 5 0 0 H * 2 0 0 0 O 0) co 1 5 0 0 A f te r l oad - 1 3 5 m m H g 1 0 0 0 H 5 0 0 Fig. 6 . 6 . Glucose and palmitate oxidation at low (top panel, 80 mm Hg) and high flower panel, 135 mm Hg) afterloads During measurement of heart function, samples of perfusate and hyamine hydroxide were taken every 10 min for measurement of substrate oxidation. Results are the means ± SE of 4 rats in each group. Top panel: "Significantly different from NC; ^Significantly different from all groups. Lower panel: Comparison was only made between results obtained at high and low afterloads, *Significantly different from low afterload; P < 0.05. 133 Fig. 6.7 Evaluation of ultrastructural cardiac morphology A representative electron micrograph of hearts from N C , N D , P C and PD animals are depicted. The scale bar on PD represents 1pm. M , mitochondria; Nu, nucleus; L , lipid like vacuoles. Hollow arrows indicate Z-lines. 134 42 35 X E E CL (/) £ Q. 0 28 H 21 Control Diabetic Afterload - 135mm Hg 1 Afterload - 80mm Hg Normal Diet co-6 P U F A Diet Normal Diet co-6 P U F A Diet Fig.6.8 Cardiac function at low (80 mm Hg) and high (135 mm Hg) afterloads. Isolated hearts were perfused in the working mode at an afterload of 80 mmHg for the first 30 min. Subsequently, the afterload is clamped off causing the peak systolic pressure to increase to -135 mmHg, and the perfusion continued for another 20 minutes. Results are the means ± SE of 4 rats in each group. RPP, rate pressure product. * Significantly different from all other groups at afterload ~135 mmHg, P < 0.05 135 6.6 References 1. Rodrigues B, Cam M C , McNeill JH. Myocardial substrate metabolism: implications for diabetic cardiomyopathy. J Mol Cell Cardiol 1995; 27:169-79 2. Cai L, Kang YJ. Oxidative stress and diabetic cardiomyopathy: a brief review. Cardiovasc Toxicol 2001; 1:181-93 3. 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Biochemical, histological and echocardiographic changes during experimental cardiomyopathy in STZ-induced diabetic rats. Pharmacol Res 2003; 48:429-35 49. Berry E M . Who's afraid of n-6 polyunsaturated fatty acids? Methodological considerations for assessing whether they are harmful. Nutr Metab Cardiovasc Dis 2001;11:181-8 50. Simopoulos AP. Essential fatty acids in health and chronic disease. Am J Clin Nutr 1999; 70:560S-569S 141 7. Chronic Provision Of Dietary co-6 PUFA Induces Mitochondrial Nitrosative Damage And Cardiac Dysfunction 7.1 Introduction Obesity, a common disorder in the Western world, increases the incidence of diseases like hypertension and diabetes. In the last few decades, heightened awareness of obesity and its complications has led to an indiscriminate substitution of atherogenic saturated cooking fats with vegetable oils containing co-6 polyunsaturated fatty acids (PUFA) [1]. In addition to this intentional modification of human dietary intake, farmed animals are fed grains and seeds containing high amount of co-6 PUFA like linoleic and arachidonic acid [2]. Consumption of these animals (and their products) has led to an even greater increase in dietary co-6 PUFA. Thus, intake of linoleic acid in the US and Western Europe has risen from <3% of energy in the 1950s to the current 6-7%, with a parallel decrease in saturated fats [3]. . A few populations (Northern Belgium and Israel) consume even higher proportions of linoleic acid (8-12% of their total energy) [3]. Additionally, following increased co-6 PUFA consumption, competition for desaturase enzymes reduces the biosynthesis of the beneficial co-3 PUFA like eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) [1]. Finally, a reduction in dietary fish oils, a rich source of co-3 PUFA, has also been documented. Taken together, the present human dietary co-6: co-3 PUFA ratio has increased from a balanced 1:1 to an unexpected 15:1 to 20:1 in the current Western diet [1]. Given that the anti-arrhythmic and hypocholesterolemic effects of PUFA have mainly been attributed to co-3, excess co-6 PUFA may not provide the desired cardiovascular benefits [1,4]. In fact, a number of recent studies have identified co-6 PUFA to be pro-inflammatory, with effects ranging from smooth muscle 142 proliferation to oxidative endothelial injury, mitochondrial damage and activation of transcription factors in the vasculature [ 5 - 8 ] The majority of studies that have evaluated the effects of co-6 PUFA on cell systems used high levels of individual fatty acids like linoleic or arachidonic acid [ 5 - 7 ] . Little attempt was made to evaluate these effects in the presence of low co-3 PUFA, which would mimic the current Western diet. Additionally, although widely studied with respect to coronary vascular disease, the consequences of increased co-6 PUFA on the heart muscle itself, are largely unknown. In a recent study, we reported that in diabetic rats fed 2 0 % w/w sunflower oil (which increases the cardiac co-6: co-3 PUFA ratio), abnormal mitochondria and decreased ATP levels were evident [ 9 ] . In these hearts, cardiac damage was characterized by necrosis, a process of cell death that requires extensive mitochondrial damage and loss of ATP [ 1 0 , 1 1 ] . Interestingly, in experimental models of diabetes fed normal chow (with a balanced co-6: co-3 PUFA ratio), or saturated fats, cardiac damage is usually characterized by an increase in apoptosis, a process of programmed cell death characterized by transient mitochondrial permeability changes [ 1 0 , 1 1 ] . Thus, one of the key 'switches' that converts apoptotic to necrotic cell death is persistent mitochondrial damage [ 1 0 , 1 1 ] . Interestingly, simply feeding 2 0 % w/w sunflower oil to control rats led to an extensive loss of cardiolipin, a mitochondrial phospholipid [ 9 ] . Depletion of cardiolipin is not only associated with loss of oxidative phosphorylation, but also extensive generation of free radicals [ 1 2 ] . We hypothesized that chronic dietary co-6 PUFA can induce free radical generation, predisposing the cardiac mitochondria to oxidative damage. In this study, we demonstrate that co-6 PUFA feeding for 4 weeks, in addition to inducing loss of cardiolipin, can precipitate mitochondrial nitrosative stress, DNA damage and lead to cardiac dysfunction at high workloads. 143 7.2 Materials And Methods 7.2.1 Experimental Animals The investigation conforms to the guide for the care and use of laboratory animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996), and the University of British Columbia. Previous studies have established that feeding 20% w/w sunflower oil for 4 weeks increases the co-6 PUFA content of the heart [9, 13]. To investigate the cardiac effects of this increased co-6 PUFA, male Wistar rats (220-240 g) were fed a diet [(AIN)-76A; Research Diets Inc.] supplemented with 20% w/w sunflower oil (High PUFA; HP) for 4 weeks. The composition of the HP diet in g%> was as follows: protein, 24.1; carbohydrate, 44.9; fats, 20.7; fiber, 6.0; kcal/g, 4.6; total co-6 PUFA, 10.6; total co-3 PUFA, 0.3. In this diet, the energy contribution from co-6 PUFA was 10.6 g%, and is reflective of the human diets in North America and Europe (around 5-10% of the total energy intake) [14, 15]. A control group fed normal laboratory chow [5%> corn oil; Low PUFA, LP] (PMI Feeds) was utilized to compare the effects of co-6 PUFA feeding. The composition of the LP diet in g%> was as follows: protein, 23.4; carbohydrate, 49.9; fats, 4.5; fiber, 8.3; kcal/g, 4.0; total co-6 PUFA, 1.2; total co-3 PUFA, 0.1. Water was provided ad libitum. Following 4 weeks of feeding the two diets, glucose was measured using a glucometer from blood obtained from the tail vein (Advantage Comfort, Roche Diagnostics). Subsequently, the rats were anesthetized with sodium pentobarbital (65 mg/kg, i.p.), the thoracic cavity opened, and hearts removed. 7.2.2 Separation And Characterization O f Cardiac Lipids Total cardiac lipids were extracted and solubilized in chloroform:methanol:acetone:hexane (4:6:1 :lv/v/v/v). Separation of cardiolipin, lysophosphatidylcholine and free fatty acids was achieved using HPLC (Waters 2690 Alliance HPLC, Milford, MA) equipped with an auto-sampler and column heater, as previously described [9, 13]. Individual fatty acids were 144 quantified as their respective methyl esters using heptadecaenoic acid (17:0) as the internal standard with a Varian 3400 G L C equipped with a flame ionization detector, a Varian Star data system, and a SP-2330 capillary column (30 m x 0.25 mm internal diameter) (Supelco, Bellefonte, PA). Value of cardiac FA, cardiolipin and lysophosphatidylcholine were expressed as p.g/mg protein. 7.2.3 Total Phospholipase A 2 (PLA2) Activity After removal of the heart, the apex was cut, washed in cold PBS buffer, and homogenized in cold buffer containing 50mM HEPES and ImM EDTA (pH 7.4). The sample was centrifuged at 10,000g for 15 mins, and supernatant separated. Supernatant corresponding to 50 pg of protein from each heart was then subjected to P L A 2 assay. Briefly, P L A 2 cleaves off fatty acids at the sn-2 position of glycerophospholipids to yield free fatty acid and lysophospholipid as by-products. Arachidonyl thio-phosphatidylcholine is a synthetic substrate that contains an arachidonyl thioester bond at the sn-2 position. Hydrolysis of this bond releases free thiol that is detected by DTNB [5,5'-dithiobis(2-dinitrobenzoic acid)] to give yellow-colored 5-thio-2-nitrobenzoic acid (Xmax 412nm) [16]. The sample was reacted with assay buffer (160 mM HEPES, 300mM NaCl, 20mM CaCl 2 , 8mM Triton X, 60%glycerol and 2mg/ml BSA) and 1.5mM Arachidonyl thio-PC for 60 mins at room temperature. Bee venom P L A 2 was used a positive control. Finally, lOul of 25mM DTNB/EDTA was added to develop color, which was measured spectrophotometrically at 405nm. 7.2.4 Separation Of Cytosolic And Particulate Fractions Heart sections were immediately snap frozen in liquid nitrogen and stored at - 80°C. 50 mg of tissue was ground under liquid nitrogen, and homogenized in cytosolic buffer (20 mM Tris HC1, 250mM E G T A , 200mM EDTA, lOOmM PMSF, lOOmM NaF, 2-mercaptoethanol, leupeptin and aprotinin). After centrifugation at 35,000g for 1 hour at 4°C, the 'cytosolic 145 fraction' (supernatant) was isolated. The pellet containing all membrane bound fractions including nucleus and mitochondria were washed, recentrifuged and supernatant discarded. The pellet was resuspended in particulate buffer containing detergents (cytosolic buffer + NP-40, 10%SDS and 5%DCA) to rupture and dissolve all membranes. The mixture was centrifuged at 35,000g for 1 hour at 4°C, and the supernatant collected as the 'particulate fraction'. The purity of the fractions was estimated by performing immunoblotting for manganese superoxide dismutase (MnSOD; a mitochondrial protein) and copper/zinc SOD (Cu/Zn SOD; a cytosolic protein). No detectable signals for MnSOD or Cu/Zn SOD were present in the cytosolic or particulate fractions respectively. Protein content of the fractions was quantified using a Bradford protein assay. 7.2.5 Western Blotting Procedures Either heart fractions or whole heart homogenates were diluted, boiled with sample loading dye, and samples corresponding to 50 pg protein were used in SDS-polyacrylamide gel electrophoresis. After transfer, nitrocellulose membranes were blocked in 5% skim milk overnight in Tris-buffered saline containing 0.1% Tween-20 (TBS-T). Membranes were incubated with antibodies raised in rabbit [inducible nitric oxide synthase (iNOS), peroxisome proliferator-activated receptor gamma coactivator 1 (PGC-1)], sheep [Cu/Zn superoxide dismutase (Cu/ZnSOD), Mn superoxide dismutase (MnSOD)], mouse (nitrotyrosine) or goat (p22 p h o x , p47 p h o x) for 2 hours at room temperature. Following 3x wash in TBS-T, the membranes were incubated for the next 2 hours at room temperature with secondary goat anti-rabbit, goat anti-mouse, donkey anti-sheep or donkey anti-goat HRP-conjugated antibodies, and visualized using E C L detection kit. 146 7.2.6 Immunofluorescence Part of the left ventricle was fixed in 10% neutral buffered formalin, embedded in paraffin, and 5 urn sections prepared. Double immunostaining was performed for both nitrotyrosine and 8-hydroxyguanosine (8-OHG) [17]. For the first labeling, sections were deparaffinized in xylene, rehydrated using various grades of ethanol, and pretreated with 10pg/ml proteinase K (to permeabilize the nucleus) for 30 min at 37°C. Non-specific binding of immunoglobulins was blocked by incubating the sections in 10% BSA for 20 minutes. Nitrotyrosine immunolabeling was performed using a rabbit polyclonal anti-nitrotyrosine antibody (1:50, 2 hours at 37°C) (Molecular Probes, Oregon, USA) with Alexa Fluor 594® conjugated goat anti-rabbit IgG (Molecular Probes; 90 min at 37°C). For the second labeling, the same sections were incubated with primary goat anti-8-OHG antibody (Alpha Diagnostics) (1:200, overnight at room temperature), and secondary donkey anti-goat antibody conjugated with Alexa Fluor 488®. Sections were finally counterstained with DAPI (to visualize nuclei), and examined using confocal fluorescent microscopy. Slides were visualized using FITC, DAPI and Texas Red filters under a mutiphoton confocal microscope. 7.2.7 Gene Expression Gene expressions of mitochondrial respiratory subunits encoded by mitochondrial DNA (mfDNA) and 8-oxoG-DNA glycosylase (OGG-1), a mitochondrial oxidative DNA repair enzyme were measured using reverse transcriptase-polymerase chain reaction (RT-PCR). Briefly, total RNA from snap frozen hearts (lOOmg) was extracted using Tri-Zol (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 specific primers for N A D H dehydrogenase subunit 1 (ND-1), cytochrome b (CYTb), cytochrome-c-oxidase subunit 1 (COX 1), ATPase 6 147 and OGG-1 (primers are listed in Table 1). 18S rRNA or P-actin was amplified as an internal control. The linear range was found to be between 15 and 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 and identified according to bands corresponding to specific base pairs (as per Table-1). Expression levels were represented as the ratio of signal intensity for target mRNA relative to either 18S or p-actin mRNA. 7.2.8 Cardiac Function Hearts from halothane (2-3%)-anesthetized rats were perfused for 30 min with Krebs-Henseleit (KH) solution in the working heart mode at a preload of 11.5 mmHg, as previously described [18]. The K H solution contained 1.2mM palmitate prebound to 3% fatty acid-free albumin, 5.5mM glucose, 0.5mM lactate, 2.5mM calcium, and 100mU/l of insulin, and was continuously circulated through the closed perfusion system. A high concentration, of palmitate was used to minimize differences in fatty acid oxidation, between HP and LP hearts. To ensure that glucose uptake was not limiting, a high physiological concentration of insulin was utilized. The solution was oxygenated with 95% 02-5% C 0 2 and maintained at 37°C throughout the perfusion. A pressure transducer (Viggo-Spectramed, Oxnard, CA) inserted in the afterload line was used to measure heart rate and peak systolic pressure. Cardiac output and aortic flow was measured via external flow probes (Transonic Systems, Ithaca, NY) on the preload and afterload lines, respectively. An afterload of 80mmHg was kept for the first 30 min. Subsequently, the afterload was clamped off causing the peak systolic pressure to increase to ~135 mmHg, and the perfusion continued for another 20 minutes [9]. 148 7.2.9 Statistical Analysis. Values are means ± SE. Paired student's /-test was used to determine differences between group mean values. The level of statistical significance was set at P < 0.05. 149 7.3 Results 7.3.1 Cardiac Content Of co-3 And co-6 PUFA Compared to LP fed animals, HP feeding for 4 weeks did not affect food or water intake (data not shown), body weight (LP, 431 ± 7 ; HP, 454 ± 23; g) or blood glucose (LP, 5.8 ± 0.4; HP, 6.4 ± 0 . 1 ; mmol). The cardiac PUFA content in the free form is described in Table 7.2. Feeding 20% w/w sunflower oil successfully magnified both the co-6 (mainly linoleic and arachidonic acid) and co-3 (mainly linolenic, EPA and DHA) PUFA content in HP fed hearts. However, the increase in co-6 was substantially greater than the increase in co-3 PUFA, and resulted in an approximate 3-fold increase in the co-6: co-3 ratio in HP fed rats. 7.3.2 Phospholipid Metabolism Cardiolipin is a mitochondrial membrane phospholipid that is essential for oxidative phosphorylation (OXPHOS). HP feeding brought about a severe drop in total cardiolipin levels. Evaluation of individual fatty acids within this phospholipid revealed a substantial reduction in the linoleic acid fraction of cardiolipin, with no significant change in the saturated and monounsaturated fatty acid fraction (Fig 7.1). Hearts from HP rats also demonstrated a substantial increase in P L A 2 activity (Fig 7.2A). Given the non-selective nature of P L A 2 , we also measured total phospholipids. A 50%> decrease in total cardiac phospholipids with a corresponding increase in lysophosphatidylcholine (LPC), a lysophospholipid by-product of increased P L A 2 activity, was noted (Fig 7.2B, C). 7.3.3 Nitrosative Stress Under pathological conditions, excess nitric oxide (NO) is predominantly generated by nitric oxide synthases (NOS) like inducible nitric oxide synthase (iNOS). HP hearts demonstrated increased iNOS protein expression (Fig 7 . 3 A ) . NO when combined with superoxide, can form the most damaging free radical, peroxynitrite (ONOO"). ONOO" nitrates tyrosine residues of 150 proteins to produce nitrotyrosine. HP hearts were characterized by a significant rise in the particulate, but not cytosolic nitrotyrosine levels (Fig 7.3B). 7.3.4 Mitochondrial Damage In addition to proteins, peroxynitrite can also oxidize macromolecules like DNA, present in both mitochondria and nucleus, to produce 8-hydroxyguanosine (8-OHG) [19]. As evidence of peroxynitrite activity in the particulate fraction could indicate mitochondrial and/or nuclear damage, immunofluorescence for nitrotyrosine and 8-OHG was carried out on rat heart sections (Fig 7.4). LP fed hearts demonstrated negligible 8-OHG and nitrotyrosine staining (Fig 7.4A, B, C). In contrast, the HP group demonstrated widespread 8-OHG staining (Fig 7.4D, G) that co localized with increased nitrotyrosine (Fig 4E, H) staining in the extra nuclear area (Fig 4 I), suggesting a predominantly mitochondrial location for this DNA damage. The mRNA expression of OGG-1, the DNA repair enzyme specific for 8-OHG, also increased in HP hearts (Fig. 7.4J). Nuclear staining for nitrotyrosine and 8-OHG was absent in all groups. The mitochondrial origin of peroxynitrite was also supported by decreased levels of Mn superoxide dismutase (SOD), the antioxidant responsible for neutralizing superoxide in the mitochondria (Fig 7.5A). Interestingly, Cu/Zn SOD, the cytosolic antioxidant for superoxide demonstrated no change in protein levels (Fig 7.5B) between LP and HP fed hearts. To evaluate the role of cytosol in the generation of free radicals, we measured p22 p h o x and p47 p h o x subunit expression of the NADPH oxidase system. Both p22 p h o x (LP, 2.5±0.3; HP, 3.0±0.4, arbitrary units) and p 4 7 p h o x 2.8±0.2; HP, 3.0±0.4, arbitrary units) demonstrated no change in their protein expression. 7.3.5 Loss Of Gene Expression Of Respiratory Enzyme Subunits A mammalian mitochondrion encodes 13 essential components of oxidative phosphorylation enzyme complexes. In lieu of investigating the gene expression of all 13 subunits, we 151 identified 4 genes, each representative of a different enzyme complex, (viz. N A D H dehydrogenase subunit 1 (ND-1) from Complex I, cytochrome b (CYTb) from Complex III, cytochrome-c-oxidase subunit 1 (COX 1) from Complex IV and ATPase 6 from Complex V) which is affected by oxidative stress in the heart. It should be noted that subunits from Complex II are not encoded by mitochondrial DNA but by nuclear DNA, and were thus excluded from gene expression studies. Interestingly, RT-PCR studies revealed a drop in gene expression of only ND-1 (Fig 7.6B) and ATPase 6 (Fig 6 C ) , but not Cytb (LP, 2.0±0.57; HP, 1.7+0.1, arbitrary units; Fig 6A) or COX-1 (LP,-1.0110.07; HP, 1.02±0.07, arbitrary units; Fig 6A) in the HP group. Protein expression of PGC-1 (a nuclear co activator for biogenesis of mitochondrial OXPHOS enzymes) was negligible in LP hearts but increased almost 5-fold in the HP hearts (Fig7.6D). 7 . 3 . 6 Cardiac Functional Parameters Cardiac output (CO) is the total volume of blood pumped by the ventricle per minute. There were no differences in CO or heart rate (HR) between the LP and HP groups at normal workload (80 mmHg). However, subsequent to an increase in workload to 135 mmHg, CO and HR decreased in all groups. Interestingly, compared to LP, HP hearts demonstrated a more robust decline in both CO and HR (Table 7.3). 152 7.4 Discussion Changes that disrupt mitochondrial structure or function can compromise cardiac contractility. Cardiolipin, a mitochondrial membrane phospholipid, is absolutely essential for the activity of key OxPhos enzymes [12, 20]. Recently, we demonstrated that cardiac cardiolipin levels were reduced in rats fed 20% w/w sunflower oil (rich in linoleic acid, an co-6 PUFA) [9]. In the current study, we confirmed this observation and extended our findings to a specific loss of the linoleic acid fraction in cardiac cardiolipin, following HP feeding. As linoleic acid represents around 80% of the total cardiolipin mass [12], and given that the HP diet is rich in linoleic acid, this observation appeared paradoxical. It should be noted that in the rat heart, other studies have also demonstrated a loss of cardiolipin following chronic co-6 PUFA feeding [8, 21]. A mechanism behind such an occurrence is yet to be determined. However, high co-6 PUFA feeding has been shown to increase cardiac phospholipase A 2 (PLA 2) activity [22]. P L A 2 specifically cleaves the acyl group at the sn-2 position of most phospholipids, releasing constituent arachidonic acid and lysophosphatidylcholine (LPC). Similar effects on linoleic acid rich cardiolipin can lead to a reduction in this fatty acid fraction, and a loss in total cardiolipin mass [12]. Indeed, increased P L A 2 activity has been reported to decrease cardiolipin levels in rat brain and liver [23, 24]. In the current study, the increase in total P L A 2 activity matched the drop in total phospholipids, and the increase in LPC in HP fed hearts. Given the ability of LPC and arachidonic acid to generate superoxide (CV), this free radical can also lead to peroxidation of the highly unsaturated linoleic acid fraction of cardiolipin, leading to a further loss in this phospholipid [25-27]. In addition to generation of superoxide, LPC and other P L A 2 metabolites are known to increase inducible nitric oxide synthase (iNOS) expression, as observed following HP feeding [28]. iNOS overexpression produces large amounts of nitric oxide (NO), for a sustained period. Excess NO, when combined with superoxide, forms peroxynitrite (ONOO"), a highly 153 reactive anion [29]. Although NO was not directly measured, its increased generation is critical for the formation of peroxynitrite [30, 31]. Once formed, peroxynitrite can oxidize lipids and nitrate proteins to form nitrotyrosine. Based on its intracellular location, peroxynitrite can differentially affect cellular functions. Mitochondrial peroxynitrite can oxidize sulfhydryl groups on aconitase and other enzymes in the mitochondrial respiratory chain, thereby impairing energy production [32]. Additionally, peroxynitrite can oxidize the C-8 position of guanine in the DNA of both mitochondria and nucleus to form 8-hydroxyguanosine (8-OHG) [19]. Oxidative damage to nuclear D N A (nDNA) triggers DNA repair, increased N A D consumption and cell death [29]. Measurement of cytosolic and 'particulate' (consisting both nucleus and mitochondria) nitrotyrosine following HP feeding revealed an increase in particulate, and not cytosolic nitrotyrosine. To determine the nuclear and/or mitochondrial location of peroxynitrite, we used immunofluorescent co-localization techniques to identify 8-OHG and nitrotyrosine on intact cardiac sections. Mechanical separation of nucleus from mitochondria often leads to high levels of background oxidation and artificial DNA damage, and was avoided to prevent false positivity for nitrotyrosine and oxidized DNA [33, 34]. Our double-labeling immunofluorescence analysis revealed that both nitrotyrosine and 8-OHG were localized predominantly in the extranuclear space in HP heart sections, thus negating the oxidative effects of peroxynitrite on nuclear DNA. In spite of the presence of extensive free radical induced mitochondrial, damage, absence of nuclear DNA damage may explain the lack of cell death (both apoptosis and necrosis) in rat hearts following HP feeding [9]. Thus, in the current study, cardiac mitochondria (the only other organelle with DNA) were believed to be the source for 8-OHG in HP hearts. This preferential damage to mDNA likely occurs as a consequence of a lack of protective histones and its proximity to the electron transport 154 complexes that produce free radicals [35]. Accumulation of 8-OHG in the mitochondria can cause GC to A T mutations [36]. To prevent this event, nuclear DNA encoded OGG-1, an 8-oxoG-DNA glycosylase can be targeted towards mitochondria to remove 8-OHG. Our data suggests that similar to humans and rodents, augmented expression of OGG-1 in HP hearts could result from increased formation of 8-OHG [36-38]. HP hearts were characterized by a specific loss of MnSOD, the antioxidant responsible for neutralization of mitochondrial O2", an essential precursor for peroxynitrite formation [39]. As the cytosolic antioxidant Cu/Zn SOD, and the NADPH oxidase subunits p22 p h o x and p47 p h o x protein expression (chief source for extramitochondrial ROS generation within the cytosol), demonstrated no change in their protein expression, our data supports a predominantly mitochondrial location of peroxynitrite. It should be noted that high fat feeding with lard has been shown to induce p22 p h o x and p47 p h o x protein expression in rat brains [40]. It is likely that this differential effects of high fat on the NADPH system is due to differences in organs studied, and the type of dietary fat used (lard is a source of saturated fatty acids). The mammalian mitochondrial DNA contains 13 polypeptide genes, all of which encode essential subunits of OxPhos complexes including complex I (ND 1, 2, 3, 4, 4L, 5, and 6), III (cytochrome b), IV (COX I, II and III) and V (ATPase 6 and 8) [17, 41]. Although mutations in these genes are known to decrease the rate of energy production, the relative importance of individual genes remains controversial. Nevertheless, with 13 potential candidates, we selected 4 genes from the different complexes that are affected by cardiac oxidative stress. For example, oxidative stress induced by lipopolysaccaride or doxorubicin alters ND-1 and ATPase-6, and C O X I expression, respectively [42, 43]. Additionally, in humans, oxidative damage following myocardial infarction decreases ND-1 and cytochrome b [44]. In the present study, HP feeding reduced the gene expression of ND-1 of complex I, and ATPase 6 of complex V. Although the reason behind such losses in gene expression of specific subunits is 155 unknown, complex I inhibition has been suggested to work as a protective response to limit electron flow to Complex III, in an attempt to decrease superoxide generation [45]. Loss of mitochondrial respiratory subunits may promote a compensatory response from the nuclear genome through activation of PGC-1, as seen in HP hearts. PGC-1 interacts with nuclear respiratory factors (NRF-1 and NRF-2) to induce genes for OXPHOS and other mitochondrial functions [43]. With regard to the heart, oxidative stress induced by lipopblysaccaride exposure damages mDNA, decreases respiratory subunits and concurrently elevates gene expression for PGC-1 [43]. In spite of considerable mitochondrial damage, both LP and HP fed hearts had identical cardiac output and heart rate at basal afterloads of 80 mmHg. It is likely that during such conditions, the vast mitochondrial reserves in cardiomyocytes (around 30% of the cardiomyocytes volume is occupied by mitochondria) may be responsible for this absence of any functional alterations in HP hearts [46]. However with the existing mitochondrial nitrosative damage, we hypothesized that HP hearts may not function optimally following increased workloads. Consistent with our hypothesis, both LP and HP fed hearts had identical cardiac output and heart rates at basal afterloads. However, when these hearts were exposed to a high afterload, HP hearts demonstrated a greater decline in both these functional parameters than LP hearts. A limitation of this study is the lack of an isocaloric diet (e.g., a diet enriched with saturated fatty acid). Thus, the conclusions of mitochondrial nitrosative damage cannot be attributed solely to co-6 PUFA, but could be a result of a high fat diet per se. However, it should be noted that the pro-inflammatory properties demonstrated in this study (activation of PLA2, breakdown of membrane phospholipids, and induction of nitrosative stress) is a hallmark of co-6 PUFA and not saturated fatty acids [4-7]. 156 In summary, we conclude that a high fat diet enriched with co-6 PUFA leads to activation of pro-inflammatory pathways within the heart causing mitochondrial nitrosative damage (Fig. 7.7) and cardiac dysfunction at higher afterloads. Given the postmitotic nature of cardiomyocytes, the heart would be particularly vulnerable to this damage inflicted by co-6 PUFA [47]. Interestingly, accumulation of co-6 PUFA has been strongly associated with an augmented risk of myocardial damage in humans [48, 49]. With the current shift towards dietary co-6 PUFA, our study raises other epidemiological concerns. During aging and diseases associated with aging, loss of cardiolipin [8, 21], oxidative stress, mitochondrial membrane and DNA damage [50] are observed, and are similar to changes seen in this study. Whether the current 'epidemic' of cardiovascular disease is a function of high fat diet enriched with co-6 PUFA warrants further investigation. 157 7.5 Tables and Figures Table 7.1 Primers used for PCR amplification of genes. Name GeneBank Acc. No. Forward (5'-3') ND1 X 14848 (2729.3685) CytB X 14848 (14124..15266) C O X 1 X 14848 (5309..6853) ATPase6 X 14848 (7904..8584) 18S rRNA XO 3205 OGG1 NMO 30870 P-actin N M 031144 C A C C C C C T T A T C A A C C T C A A T C T C A T C A G T C A C C C A C A T C A G C A G G A A T A G T A G G G A C A G C C G A A C C T G A G C C C T A A T A T C A A T C T C G G G T G G C T G A A C G A T C T G T T C T T C C A A C A A C A A C T T G T C A C C A A C T G G G A C G A T A T G Reverse (5'-3') Base pair ATTTGTTTCTGCGAGGGTTG 305 CATTCTGGTTTGATGTGGGG 627 T G A G A G A A G T A G T A G G A C G G C 519 G T A G C T C C T C C G A T T A G A 337 G G A C C A G A G C G A A A C C A T T T G 495 G C C A G C A T A A G G T C C C C A C A G 500 G A T C T T G A T C T T C A T G G T G C T A G 763 Table 7.2 Major cardiac co-6 and co-3 PUFA L o w P U F A H I G H PUFA Linoleic Acid (18:2 co-6) 0.42 ± 0 . 0 1 2.1± 0.09* Arachidonic Acid (20:4 co-6) 0.12 ± 0 . 0 0 3.7 ± 0 . 0 7 * Linolenic Acid (18:3 co-3) 0.14 ± 0 . 0 0 0.71 ± 0 . 1 3 * Eicosapentaenoic Acid (20:5 co-3) 0.10 ± 0 . 0 0 0.20± 0.05* Docosahexaenoic Acid (22:6 co-3) 0.07 ± 0 . 0 2 0.17 ± 0 . 0 8 * Animals were fed either low or high PUFA for 4 weeks before termination and removal of the hearts. Cardiac lipids including phospholipids were extracted with chloroform: methanol: acetone: hexane solvent. Separation of phospholipids and fatty acids was achieved using HPLC, followed by their conversion to their respective methyl esters, and quantification by GLC. Data are expressed in pg/mg protein. The results are means ± SEM for 4-6 animals in each group. * Significantly different from low PUFA fed group, P < 0.05. 159 Table 7.3 Heart Rate and Cardiac Output of isolated working hearts P A R A M E T E R Low P U F A H I G H P U F A B A S A L A L E L E V A T E D A L B A S A L A L E L E V A T E D A L Heart Rate 257±11 305±7 243±3 283±6* (beats/min) Cardiac Output (ml/min) 7 1 ± 2 ' 2 4 1 ± 4 " 6 6 8 ± L 3 2 8 ± L 4 * Animals were fed either low or high PUFA for 4 weeks before termination and removal of the hearts. Various parameters of cardiac function were measured at either basal (80 mm Hg) or elevated (135 mm Hg) afterloads (AL). The results are means ± SEM for 4-6 animals in each group. * Significantly different from corresponding group fed low PUFA, P < 0.05. 160 15 -i Fig. 7.1 Effects of co-6 PUFA feeding on cardiac cardiolipin Separation of total cardiolipin was achieved by HPLC. Cardiolipin fatty acids were quantified as their respective methyl esters using heptadecaenoic acid (17:0) as the internal standard with a G L C equipped with a flame ionization detector. Results are the means ± SE of 4-6 rats in each group. SFA, saturated fatty acid; M U F A , monounsaturated fatty acid. * Significantly different from low PUFA fed group; P< 0.05. 161 CO Fig . 7.2 Cardiac phospholipid metabolism Total cardiac phospholipase A 2 (PLA 2) activity as measured by a spectrophotometric assay using arachidonyl thio-phosphatidylcholine as a substrate (A). Total cardiac phospholipid (B) and lysophosphatidylcholine (C) levels were measured by HPLC. Results are the means ± SE of 4-6 rats in each group. * Significantly different from low PUFA fed group; P< 0.05. 162 Low P U F A High P U F A Low P U F A High P U F A cNitrotyrosine —»-pNitrotyrosine ' * W • (B) Fig. 7.3 Cardiac nitrosative stress Inducible nitric oxide synthase (iNOS) (A) and cytosolic and particulate nitrotyrosine (B) protein expression were measured by Western blot, followed by densitometry of the respective bands. Results are the means ± SE of 4-6 rats in each group. * Significantly different from low PUFA fed group; P < 0.05. 163 Low PUFA High PUFA J . OGG-1 G U E S S E S ^ " * ^ ^ i r > n Fig. 7.4 Intracellular localization of peroxynitrite and 8-hydroxyguanosine (8-OHG) by confocal microscopy Formalin-fixed heart sections were analyzed by double immunostaining for 8-OHG (A, D, G) and nitrotyrosine (B, E, H). C, F and I represent composite images of 8-OHG, nitrotyrosine and nucleus. Images A-C represent low PUFA fed heart sections that were negative for both nitrotyrosine and 8-OHG (400x magnification). Images D-F represent high PUFA fed heart sections with both 8-OHG and nitrotyrosine at 400x magnification. Images G-I represent high PUFA fed hearts at a higher magnification (1200x) and demonstrate a predominantly extra nuclear location of 8-OHG and nitrotyrosine (arrows). OGG-1 gene expression is depicted in representative gels, followed by densitometric analysis (J). p-actin mRNA was utilized as an internal standard. Results are the means ± SE of 4-6 rats in each group. * Significantly different from low PUFA fed groups; P < 0.05. OGG-1, 8-oxo-DNA glycosylase. 164 Low PUFA High PUFA Low PUFA High PUFA (B) ro 3 i_ 5-Q o cn c s o Fig. 7.5 Intracellular superoxide dismutase (SOD) protein expression Mitochondrial MnSOD (A) and cytosolic Cu/Zn SOD (B) protein expression were measured by Western blot followed by densitometry of the representative bands. Results are the means ± SE of 4-6 rats in each group. * Significantly different from low PUFA fed group; P < 0.05. 165 (A) L O W P U F A H IGH P U F A 18s R N A -C O X - 1 -18s R N A -CYT. B -4MMPtH WWMtt 18s R N A -(B) N D - 1 -1.0-1 < or 0.5 4 Q 0.0 L O W P U F A H IGH P U F A r T l i i Low P U F A High P U F A L O W P U F A H IGH P U F A (C) 18s R N A -A T P a s e - 6 -0.6 -, oo S 0.3 0 tn co 0.0 (D) 0.6 ^ 0.4 C O O O Q-0.2 0.0 Low P U F A High P U F A Fig. 7.6 Respiratory subunit gene and PGC-1 protein expression Gene expression of mitochondrial respiratory subunits encoded by mitochondrial DNA (mtDNA) was measured using reverse transcriptase-polymerase chain reaction. Four genes, each representative of a different enzyme complex were chosen. Representative gels of COX-1 and Cyt b gene expression are depicted in (A). No change was observed between the two groups. ND-1 (B) and ATPase 6 (C) gene expression are depicted in representative gels, followed by densitometric analysis. 18s RNA was utilized as an internal standard. PGC-1 protein expression was measured by Western blot followed by densitometry of the representative bands in (D). Results are the means ± SE of 4-6 rats in each group. * Significantly different from low PUFA fed group; P < 0.05. ND1, N A D H dehydrogenase subunit 1 from Complex I; CYTb, cytochrome b from Complex III; COX 1, cytochrome-c-oxidase subunit 1 from Complex IV; ATPase 6 from Complex V; PGC-1, peroxisome proliferator-activated receptor gamma co activator 1. 166 HF diet with co-6 PUFA PLA2 Phosphol ipid Breakdown IPQ AA J. •> iNOS Cardiolipin Loss \ MnSOD +°2 T NO Respiratory <— Gene expression 4 i ONOO 8-OHG DA//A 4 oxidation _ ^ x " PGC-1 Mitochondria IGG-1 Nucleus Fig. 7.7 Proposed pathways of mitochondrial damage and nuclear responses following consumption of a high fat (HF) diet rich in co-6 PUFA Excess co-6 PUFA increases cardiac phospholipase A 2 (PLA 2) activity, breaking down cardiac phospholipids, including cardiolipin, and promoting free radical generation. Byproducts of P L A 2 activity like lysophosphatidylcholine (LPC), and A A (arachidonic acid) can stimulate inducible nitric oxide synthase (iNOS) expression, and production of nitric oxide (NO). 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Up-regulation of base excision repair activity for 8-hydroxy-2'-deoxyguanosine in the mouse brain after forebrain ischemia-reperfusion. J Neurochem 2000; 74:1098-105 37. Moller P, Loft S, Lundby C, Olsen NV. Acute hypoxia and hypoxic exercise induce DNA strand breaks and oxidative DNA damage in humans. Faseb J 2001; 15:1181-6 171 38. Tsuruya K , Furuichi M , Tominaga Y , Shinozaki M , Tokumoto M , Yoshimitsu T, et al. Accumulation of 8-oxoguanine in the cellular D N A and the alteration of the OGG1 expression during ischemia-reperfusion injury in the rat kidney. D N A Repair (Amst) 2003;2:211-29 39. L i Y , Huang TT, Carlson EJ , Melov S, Ursell P C , Olson JL , et al. Dilated cardiomyopathy and neonatal lethality in mutant mice lacking manganese superoxide dismutase. Nat Genet 1995; 11:376-81 40. Zhang X , Dong F, Ren J, Driscoll M J , Culver B . High dietary fat induces N A D P H oxidase-associated oxidative stress and inflammation in rat cerebral cortex. Exp Neurol 2005; 191:318-25 41. Wallace D C . Mitochondrial diseases in man and mouse. Science 1999; 283:1482-8 42. Lebrecht D , Setzer B , Ketelsen U P , Haberstroh J, Walker U A . Time-dependent and tissue-specific accumulation of m t D N A and respiratory chain defects in chronic doxorubicin cardiomyopathy. Circulation 2003; 108:2423-9 43. Suliman H B , Wel ty-Wolf K E , Carraway M , Tatro L , Piantadosi C A . Lipopolysaccharide induces oxidative cardiac mitochondrial damage and biogenesis. Cardiovasc Res 2004; 64:279-88 44. Ide T, Tsutsui H , Hayashidani S, Kang D , Suematsu N , Nakamura K , et al. Mitochondrial D N A damage and dysfunction associated with oxidative stress in failing hearts after myocardial infarction. Circ Res 2001; 88:529-35 45. Chen Q, Vazquez EJ , Moghaddas S, Hoppel C L , Lesnefsky EJ . Production of reactive oxygen species by mitochondria: central role of complex III. J B i o l Chem 2003; 278:36027-31 172 46. Ferrari R. The role of mitochondria in ischemic heart disease. J Cardiovasc Pharmacol 1996;28Suppl F.Sl-10 47. Larsson N G , Holme E , Kristiansson B , Oldfors A , Tulinius M . Progressive increase of the mutated mitochondrial D N A fraction in Kearns-Sayre syndrome. Pediatr Res 1990; 28:131-6 48. Kark JD, Kaufmann N A , Binka F, Goldberger N , Berry E M . Adipose tissue n-6 fatty acids and acute myocardial infarction in a population consuming a diet high in polyunsaturated fatty acids. A m J C l i n Nutr 2003; 77:796-802 49. Valek J, Hammer J, Kohout M , Grafnetter D , Vondra K , Topinka V . Serum linoleic acid and cardiovascular death in postinfarction middle-aged men. Atherosclerosis 1985; 54:111-8 50. Beckman K B , Ames B N . The free radical theory of aging matures. Physiol Rev 1998; 78:547-81 173 8. Conclusions There is strong experimental evidence supporting the involvement of apoptosis in the development of diabetic myocardial disease [1]. However, although the significance of cell death in the diabetic heart was highlighted more than 6 years ago, factors modulating such death are unclear [2]. Oxidative stress has been hypothesized to be the "unifying link" behind all diabetic complications including heart disease [3, 4]. However, large-scale trials have failed to demonstrate any cardiovascular benefits using antioxidants like vitamin E and |3-carotene in diabetic patients [5, 6]. These regimens have failed probably because as 'chain-breaking' antioxidants, they could only prevent lipid peroxidation reactions, but not oxidative protein modifications or D N A damage, which can be only prevented by antioxidants that can directly inhibit free radical generation at its source. Further, it is possible that these patients with a long-standing history of diabetes were poor candidates for damage reversal. A s this thesis and other laboratories have demonstrated mitochondria to be the main, source of free radical generation in the diabetic heart, antioxidants that neutralize free radicals within the cardiac mitochondria are warranted [7]. Glutathione (GSH) is such an endogenous antioxidant that can directly prevent free radical formation in the cardiac mitochondria. With regards to G S H , this thesis provides the following evidence, which could shed new light on antioxidant therapy during diabetes: 1 The major source for free radical generation in the diabetic heart is the mitochondria and not the cytosol. Thus, therapies increasing cytosolic antioxidant levels only, may not prevent diabetic heart disease. 2 Decrease in mitochondrial glutathione and oxidative stress can be observed as early as 3 days following the induction of diabetes. Thus, therapy against oxidative stress should be initiated early. 174 3 Replenishment of mitochondrial GSH can reduce oxidative stress and cell death in the diabetic heart by reducing the generation of free radicals from the source (i.e. the mitochondria). Although lipotoxicity is also considered a major player in the induction of diabetic heart muscle death, most of the work has been done in vitro with saturated fatty acids. Looking at the current western dietary trends, more emphasis on the role of PUFAs, specifically co-6 PUFA are warranted. This thesis demonstrates that feeding a high fat diet enriched with co-6 PUFA in rats leads to activation of pro-inflammatory pathways within the heart causing mitochondrial nitrosative damage. Given the postmitotic nature of cardiomyocytes, the heart would be particularly vulnerable to this damage inflicted by co-6 PUFA [8]. Interestingly, accumulation of co-6 PUFA has been strongly associated with an augmented risk of myocardial damage in humans [9, 10]. Upon superimposition of diabetes, with preexisting mitochondrial nitrosative damage, this thesis also demonstrates loss of ATP, increased cardiac necrosis and functional cardiac impairment at higher afterloads. The findings of this thesis also raise some important epidemiological concerns. The loss of cardiolipin [11], oxidative stress [12], DNA damage and energy stress [13] are all hall marks of aging. Indeed in human studies, high co-6 PUFA or low co-3 PUFA feeding is positively correlated with metabolic and degenerative diseases like cancer [14] atherosclerosis [15, 16], Alzheimer's and Parkinson's disease [17], insulin resistance and diabetes [18, 19]. Even in the absence of any disease, high co-6 PUFA feeding in young Wistar rats decreases cardiac inotropic responses to adrenergic agonists to that seen in senescent rats [20]. Whether the current epidemic of cancer, neurological or cardiovascular disease is a function of high co-6 PUEA diet is a question that warrants further investigation. 175 8.1 Future Directions This thesis demonstrates that cardiac mitochondrial GSH is lost following the induction of diabetes. It is possible that following generation of excessive ROS from the electron transport chain, the flux of cytosolic GSH to the mitochondria is impaired in the diabetic heart, which in turn leads to lower mitochondrial GSH levels and oxidative damage. Two major transporters, the dicarboxylate and 2-oxaloglutarate carriers, transport mitochondrial GSH in the liver [21]. However their regulation in the diabetic heart is still unknown. Apart from defective transporters, alteration of mitochondrial membrane phospholipids during diabetes [22] can also affect this transport and can precipitate a deficit in mitochondrial GSH [21]. My MSc. project also demonstrated a severe depletion of cardiac GSH following co-6 PUFA feeding [23], a conclusion supported by other studies [24-26]. Interestingly, increasing dietary co-3 PUFA like docosahexaenoic acid can reverse such a defect [27, 28]. In all df these cases, the exact sub cellular site of GSH depletion and the exact mechanisms leading to such a loss of cardiac GSH following co-6 PUFA feeding remains unknown. Thus, future experiments should target the study of mitochondrial GSH transport and the effects of both co-3 PUFA and co-6 PUFA in modulating cardiac GSH levels in the diabetic heart. 176 8.2 References 1. Cai L , Kang Y J . Ce l l death and diabetic cardiomyopathy. Cardiovasc Toxicol 2003; 3:219-28 2. Frustaci A , Kajstura J, Chimenti C , Jakoniuk I, Leri A , Maseri A , et al. Myocardial cell death in human diabetes. Circ Res 2000; 87:1123-32 3. Dhalla N S , L i u X , Panagia V , Takeda N . Subcellular remodeling and heart dysfunction in chronic diabetes. Cardiovasc Res 1998; 40:239-47 4. Brownlee M . Biochemistry and molecular cell biology of diabetic complications. Nature 2001;414:813-20 5. Lonn E , Yusuf S, Hoogwerf B , Pogue J, Y i Q, Zinman B , et al. Effects of vitamin E on cardiovascular and microvascular outcomes in high-risk patients with diabetes: results of the H O P E study and M I C R O - H O P E substudy. Diabetes Care 2002; 25:1919-27 6. M R C / B H F Heart Protection Study of antioxidant vitamin supplementation in 20,536 high-risk individuals: a randomised placebo-controlled trial. Lancet 2002; 360:23-33 7. Green K , Brand M D , Murphy M P . Prevention of mitochondrial oxidative damage as a therapeutic strategy in diabetes. Diabetes 2004; 53 Suppl 1:S110-8 8. Larsson N G , Holme E , Kristiansson B , Oldfors A , Tulinius M . Progressive increase of the mutated mitochondrial D N A fraction in Kearns-Sayre syndrome. Pediatr Res 1990; 28:131-6 9. Kark JD, Kaufmann N A , Binka F, Goldberger N , Berry E M . Adipose tissue n-6 fatty acids and acute myocardial infarction in a population consuming a diet high in polyunsaturated fatty acids. A m J C l in Nutr 2003; 77:796-802 1 7 7 10. Valek J, Hammer J, Kohout M, Grafnetter D, Vondra K, Topinka V. Serum linoleic acid and cardiovascular death in postinfarction middle-aged men. Atherosclerosis 1985; 54:111-8 11. Pepe S, Tsuchiya N, Lakatta EG, Hansford RG. PUFA and aging modulate cardiac mitochondrial membrane lipid composition and Ca2+ activation of PDH. Am J Physiol 1999; 276:H149-58 12. Beckman K B , Ames B N . The free radical theory of aging matures. Physiol Rev 1998; 78:547-81 13. Drew B, Leeuwenburgh C. Ageing and subcellular distribution of mitochondria: role of mitochondrial DNA deletions and energy production. Acta Physiol Scand 2004; 182:333-41 14. Bartsch H, Nair J, Owen RW. Dietary polyunsaturated fatty acids and cancers of the breast and colorectum: emerging evidence for their role as risk modifiers. Carcinogenesis 1999; 20:2209-18 15. Hodgson JM, Wahlqvist M L , Boxall JA, Balazs ND. Can linoleic acid contribute to coronary artery disease? Am J Clin Nutr 1993; 58:228-34 16. Leng GC, Taylor GS, Lee AJ , Fowkes FG, Horrobin D. Essential fatty acids and cardiovascular disease: the Edinburgh Artery Study. Vase Med 1999; 4:219-26 17. Young G, Conquer J. Omega-3 fatty acids and neuropsychiatric disorders. Reprod Nutr Dev 2005; 45:1-28 18. Raheja BS, Sadikot S M , Phatak RB, Rao MB. Significance of the N-6/N-3 ratio for insulin action in diabetes. Ann N Y Acad Sci 1993; 683:258-71 19. Lardinois CK , Starich GH, Mazzaferri EL, DeLett A. Polyunsaturated fatty acids augment insulin secretion. J Am Coll Nutr 1987; 6:507-15 178 20. Skuladottir GV, Johannsson M . Inotropic response of rat heart papillary muscle to alpha 1- and beta-adrenoceptor stimulation in relation to dietary n-6 and n-3 polyunsaturated fatty acids (PUFA) and age. Pharmacol Toxicol 1997; 80:85-90 21. Lash L H . Mitochondrial glutathione transport: Physiological, pathological and toxicological implications. Chem Biol Interact 2006; 22. Panagia V, Taira Y, Ganguly PK, Tung S, Dhalla NS. Alterations in phospholipid N-methylation of cardiac subcellular membranes due to experimentally induced diabetes in rats. J Clin Invest 1990; 86:777-84 23. Ghosh S, An D, Pulinilkunnil T, Qi D, Lau HC, Abraham A, et al. Role of dietary fatty acids and acute hyperglycemia in modulating cardiac cell death. Nutrition 2004; 20:916-23 24. Ovide-Bordeaux S, Grynberg A. Docosahexaenoic acid affects insulin deficiency- and insulin resistance-induced alterations in cardiac mitochondria. Am J Physiol Regul Integr Comp Physiol 2004; 286:R519-27 25. Ovide-Bordeaux S, Bescond-Jacquet A, Grynberg A. Cardiac mitochondrial alterations induced by insulin deficiency and hyperinsulinaemia in rats: targeting membrane homeostasis with trimetazidine. Clin Exp Pharmacol Physiol 2005; 32:1061-70 26. Mori TA, Woodman RJ, Burke V, Puddey IB, Croft K D , Beilin LJ. Effect of eicosapentaenoic acid and docosahexaenoic acid on oxidative stress and inflammatory markers in treated-hypertensive type 2 diabetic subjects. Free Radic Biol Med 2003; 35:772-81 27. Hunkar T, Aktan F, Ceylan A, Karasu C. Effects of cod liver oil on tissue antioxidant pathways in normal and streptozotocin-diabetic rats. Cell Biochem Funct 2002; 20:297-302 179 Yilmaz O, Ozkan Y, Yildirim M , Ozturk AI, Ersan Y. Effects of alpha lipoic acid, ascorbic acid-6-palmitate, and fish oil on the glutathione, malonaldehyde, and fatty acids levels in erythrocytes of streptozotocin induced diabetic male rats. J Cell Biochem 2002; 86:530-9 180 

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