Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Effect of common dietary fatty acids on viability and fibrosis of cardiac cells Beam, Julianne Cecile 2013

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
24-ubc_2014_spring_beam_julianne.pdf [ 1.28MB ]
Metadata
JSON: 24-1.0074320.json
JSON-LD: 24-1.0074320-ld.json
RDF/XML (Pretty): 24-1.0074320-rdf.xml
RDF/JSON: 24-1.0074320-rdf.json
Turtle: 24-1.0074320-turtle.txt
N-Triples: 24-1.0074320-rdf-ntriples.txt
Original Record: 24-1.0074320-source.json
Full Text
24-1.0074320-fulltext.txt
Citation
24-1.0074320.ris

Full Text

EFFECT OF COMMON DIETARY FATTY ACIDS ON VIABILITY AND FIBROSIS OF CARDIAC CELLS  by  Julianne Cecile Beam  B.Sc., The University of British Columbia (Okanagan), 2011  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE  in  THE COLLEGE OF GRADUATE STUDIES  (Biology)   THE UNIVERSITY OF BRITISH COLUMBIA  (Okanagan)    November 2013     ? Julianne Cecile Beam, 2013 ii  Abstract  Excess dietary fatty acids (FA) are considered a major contributory factor to the high prevalence of type-2 diabetes and cardiac disease. A pathological feature of cardiac disease is cardiac fibrosis. Following cardiomyocyte death, fibroblasts deposit extracellular matrix (ECM) components (fibrosis) to maintain structural integrity of the heart. However, excess fibrosis results in cardiac dysfunction. The current North American diet consists of high levels of polyunsaturated fatty acids (PUFA), mainly linoleic acid (LA; omega-6 PUFA) from vegetable oils such as corn, sunflower and safflower oils. Unlike saturated FAs, the effects of omega-6 PUFAs, like LA, alone and during oxidative stress are not widely studied. Incubation with LA and oleic acid (OA), a monounsaturated FA used as a control, decreased cell viability of cardiomyocytes and fibroblasts. High concentrations of hydrogen peroxide, used to mimic oxidative insult, also reduced cell viability and depleted glutathione, an antioxidant in fibroblasts. Furthermore, during such oxidative insult, high concentrations or longer exposure to FAs exacerbated cell death. While hydrogen peroxide decreased collagen in fibroblasts in combination with FAs, it increased soluble collagen but decreased deposited collagen. TGF?1-stimulation was used to create a fibrotic stimulus in vitro. TGF?1 increased soluble and insoluble collagen under low but not high glucose conditions. After TGF?1-stimulation, LA but not OA increased deposited collagen levels in fibroblasts. In addition, in the presence of LA, the gene expression of fibroblasts demonstrated increased expression of TIMP1, known to inhibit degradation of the ECM. Furthermore, the COL1?1/COL3 ratio, which is directly proportional to fibrotic stiffness, was increased in fibroblasts incubated with LA. In vivo, mice were fed with either corn oil (CO) or olive oil (OO) diets, followed by isoproterenol administration to initiate cardiac injury. OO feeding increased TGF?3 expression in mice hearts. In addition, similar to in iii  vitro experiments, the COL1?1/COL3 ratio in mice hearts was increased following isoproterenol-induced cardiac injury with CO. These results identified increased LA as a cause of increased collagen deposition and altered TIMP1 expression during fibrosis. The results also point towards an augmented COL1?1/COL3 ratio, which determines cardiac stiffness following fibrosis, which is an important mediator amenable to dietary FAs.                          iv  Preface  I was responsible for all experimental data, analysis and writing that are presented in this thesis, except for the animal experiments. All animal experiments were approved by UBC Animal Care Committee (Animal Care # A12-0117, Biosafety # B10-0099) and performed at the Center for Disease Modeling (CDM) in Vancouver, BC by Chuanbin Dai under the instruction of Dr. Sanjoy Ghosh. Samples were transferred and stored at UBC Okanagan prior to analysis. I conducted all RNA extractions from heart tissues, microscopy, synthesis of cDNA and qPCR of all in vivo samples. All in vitro work was characterized and conducted by myself. Part of this work was presented as an oral presentation entitled ?Linoleic acid augments oxidative damage and initiates fibrosis in cardiomyocytes and fibroblasts? at the 2012 CDA/CSEM Professional Conference and Annual Meetings in Vancouver, BC.                 v  Table of Contents  Abstract  ........................................................................................................................................  ii Preface  .........................................................................................................................................  iv Table of Contents  ......................................................................................................................... v List of Tables  .............................................................................................................................  xii List of Figures  ...........................................................................................................................  xiii List of Abbreviations  ................................................................................................................. xv Acknowledgements  ................................................................................................................  xviii Dedication  .................................................................................................................................  xix 1.0 Introduction  ............................................................................................................................ 1 1.1 Cardiometabolic Disease  ................................................................................................... 1 1.2 Oxidative Stress: The Common Link  ............................................................................... 2 1.2.1 Superoxide and Nitric Oxide Bioavailability  ............................................................ 3 1.2.2 Biological Role of Hydrogen Peroxide  ...................................................................... 3 1.2.3 The Role of Antioxidants in Oxidative Stress  ........................................................... 4 1.2.4 Importance of Glutathione in Biological Systems  .................................................... 6 1.3 The Heart: Structure and Function  ................................................................................. 8 1.3.1 Cardiac Cells  ............................................................................................................... 9 1.3.1.1 Cardiomyocytes  .................................................................................................... 9 1.3.1.2 Fibroblasts  .......................................................................................................... 10 1.4 Heart Failure  .................................................................................................................... 11 1.4.1 Cardiomyopathy  ....................................................................................................... 12 1.4.2 Key Reasons for Heart Failure: Cardiac Remodeling  ........................................... 12 vi  1.5 Oxidative Stress in Cardiac Disease, Heart Failure and Remodeling  ........................ 14 1.6 Fibrosis  .............................................................................................................................. 16 1.6.1 Role of Fibroblasts in Cardiac Remodeling  ............................................................ 16 1.6.2 Diabetes and Cardiac Fibrosis  ................................................................................. 18 1.6.3 Signaling Molecules of the Fibrotic Process within the Heart  .............................. 19 1.6.3.1 Angiotensin II Signaling  .................................................................................... 19 1.6.3.2 Transforming Growth Factor-? Signaling  ...................................................... 19 1.6.4 Soluble and Insoluble Collagen  ................................................................................ 21 1.7 Fatty Acids  ........................................................................................................................ 22 1.7.1 The Current North American Diet  .......................................................................... 23 1.7.2 Replacement of Saturated Fatty Acids with Polyunsaturated Fatty Acids  ......... 25 1.7.3 Monounsaturated Fatty Acids and their Role in Oxidative Stress  ....................... 25 1.7.4 Omega-6 Polyunsaturated Fatty Acids and their Effect on Cardiac Disease: Conflicting Evidence  .......................................................................................................... 26 1.7.4.1 Beneficial Effects of Omega-6 Polyunsaturated Fatty Acids on Cardiac Disease  ............................................................................................................................. 26 1.7.4.2 Adverse Effects of Omega-6 Polyunsaturated Fatty Acids on Cardiac  Disease  ............................................................................................................................. 27 1.7.4.3 Recent Clinical Evidence on Omega-6 Polyunsaturated Fatty Acids ............ 28 1.8 Research Overview, Hypothesis and Aims  .................................................................... 29 2.0 Materials and Methods  ........................................................................................................ 31 2.1 Chemicals and Reagents  .................................................................................................. 31 2.2 Equipment  ........................................................................................................................ 32 2.3 Cell Models  ....................................................................................................................... 33 vii  2.4 Preparation of Oleic and Linoleic Acid  ......................................................................... 34 2.5 H9c2 Cardiomyocyte Cells  .............................................................................................. 34 2.5.1 Differentiation and Plating of H9c2 Myoblasts  ...................................................... 34 2.5.2 Treatment of Cardiomyocytes  ................................................................................. 35 2.6 NIH/3T3 Fibroblasts  ........................................................................................................ 36 2.6.1 Plating of Fibroblasts  ................................................................................................ 36 2.6.2 Treatment of Fibroblasts  .......................................................................................... 36 2.7 In Vitro Assays  .................................................................................................................. 37 2.7.1 Cell Viability Assay  ................................................................................................... 37 2.7.2 Reactive Oxygen Species Assay  ............................................................................... 37 2.7.3 Glutathione Assay  ..................................................................................................... 38 2.7.4 Standardization of the Collagen Assay  ................................................................... 39 2.7.4.1 Soluble Collagen Assay  ...................................................................................... 40 2.7.4.2 Deposited Collagen Assay .................................................................................. 40 2.8 Treatment of Mice with High Fat Diets and Isoproterenol  ......................................... 41 2.9 mRNA Isolation and Complementary DNA Synthesis  ................................................. 42 2.9.1 mRNA Isolation from Fibroblasts  ........................................................................... 42 2.9.2 mRNA Extraction from Heart Tissue Samples  ...................................................... 43 2.9.3 Complementary DNA Synthesis  .............................................................................. 44 2.10 Primer Design and the Quantitative Polymerase Chain Reaction  ............................ 44 2.10.1 Primer Design  .......................................................................................................... 44 2.10.2 Quantitative Polymerase Chain Reaction  ............................................................. 44 2.10.3 Primer Efficiencies  .................................................................................................. 45 2.11 Masson?s Trichrome Stain  ............................................................................................ 49 viii  2.12 Statistical Analysis  ......................................................................................................... 49 3.0 Results  ................................................................................................................................... 51 3.1 Cardiomyocyte Viability  ................................................................................................. 51 3.1.1 Effects of Fatty Acids and Hydrogen Peroxide on Cardiomyocyte Viability  ...... 51 3.1.1.1 Impact of Oleic or Linoleic Acid for 6 and 12 h on Cardiomyocyte Viability  ........................................................................................................................... 51 3.1.1.2 Effect of Hydrogen Peroxide for 6 and 12 h on Cardiomyocyte  Viability  ........................................................................................................................... 52  3.1.2 Effects of Fatty Acids in Combination with Hydrogen Peroxide on  Cardiomyocyte Viability  ................................................................................................... 53 3.1.2.1 Effect of Hydrogen Peroxide plus Oleic or Linoleic Acid for 6 and  12 h on Cardiomyocyte Viability  .................................................................................. 53 3.2 Fibroblast Viability and Cellular Oxidative Stress  ...................................................... 54 3.2.1 Effects of Fatty Acids on Fibroblast Viability and Reactive Oxygen Species Production  .......................................................................................................................... 54 3.2.1.1 Effects of Oleic or Linoleic Acid for 6 h on Fibroblast Viability  ................... 54 3.2.1.2 Effects of Oleic or Linoleic Acid on Reactive Oxygen Species  Production in Fibroblasts over 24 h  ............................................................................. 55 3.2.2 Effects of Fatty Acids plus Hydrogen Peroxide on Fibroblast Viability and Glutathione Levels  ............................................................................................................. 57 3.2.2.1 Effect of Oleic or Linoleic Acid Combined with Hydrogen Peroxide  after 24 h on Fibroblast Viability  ................................................................................. 57 3.2.2.2 Effects of Oleic or Linoleic Acid and Hydrogen Peroxide for 24 h on Glutathione Levels in Fibroblasts ................................................................................. 58 ix  3.3 Collagen Production of Fibroblasts  ................................................................................ 60 3.3.1 Effects of Fatty Acids on Collagen Production  ...................................................... 60 3.3.1.1 Outcome of the Combination Treatment of Oleic or Linoleic Acid with Hydrogen Peroxide on Soluble Collagen Production by Fibroblasts  ........................ 60 3.3.1.2 Effects of a 24 h Combination of Oleic or Linoleic Acid with  Hydrogen Peroxide on Deposited Collagen Production by Fibroblasts  ................... 61 3.3.2 Effects of Transforming Growth Factor-?1 and Fatty Acid Treatment on  Collagen Production in Fibroblasts  .................................................................................. 63 3.3.2.1 Effects of 24 h Transforming Growth Factor-?1 with Oleic or Linoleic  Acid on Soluble Collagen Production in Fibroblasts  .................................................. 63 3.3.2.2 Effects of Transforming Growth Factor-?1 with Oleic or Linoleic Acid  on Deposited Collagen Production in Fibroblasts ....................................................... 64 3.3.2.3 Expression of Collagen Genes in Fibroblasts following Transforming  Growth Factor-?1 with Oleic or Linoleic Acid  ........................................................... 65 3.3.3 Gene Levels In Vivo ................................................................................................... 68 3.3.3.1 Gene Expression of Collagen Genes In Vivo Following High Fat  Feeding and Mild Cardiac Injury  ................................................................................ 68 3.3.3.2 Levels of Collagen from In Vivo Heart Tissues  ............................................... 71 3.4 The Effect of Hyperglycemia on the Fibrotic Response  ............................................... 73 3.4.1 Collagen Production in Fibroblasts is Different between Low and High  Glucose Conditions  ............................................................................................................ 73 4.0 Discussion .............................................................................................................................. 76 4.1 Cardiomyocyte Viability  ................................................................................................. 76 4.1.1 Effects of Fatty Acids and Hydrogen Peroxide on Cardiomyocyte Viability  ...... 76 x  4.1.1.1 Linoleic Acid Reduced the Redox Potential of Cardiomyocytes  ................... 76 4.1.1.2 High Levels of Hydrogen Peroxide Decreased Cardiomyocyte  Viability  ........................................................................................................................... 77 4.1.2 Effects of Fatty Acids in Combination with Hydrogen Peroxide on  Cardiomyocyte Viability  ................................................................................................... 78 4.1.2.1 Effect of Hydrogen Peroxide with Oleic or Linoleic Acid for 6 and  12 h on Cardiomyocyte Viability  .................................................................................. 78 4.2 Fibroblast Viability and Cellular Oxidative Stress  ...................................................... 79 4.2.1 Effects of Fatty Acids on Fibroblast Viability and Reactive Oxygen Species Production  .......................................................................................................................... 79 4.2.1.1 Linoleic Acid Reduced Cell Viability in Fibroblasts  ...................................... 79 4.2.1.2 Oleic Acid Increased Reactive Oxygen Species in Fibroblasts  ...................... 80 4.2.2 Effects of Combination of Fatty Acids with Hydrogen Peroxide on Fibroblast Viability and Glutathione Levels  ...................................................................................... 81 4.2.2.1 Combination of Fatty Acids and Hydrogen Peroxide Further Decreased  Cell Viability  ................................................................................................................... 81 4.2.2.2 Hydrogen Peroxide, but Not Fatty Acids, Decreased Glutathione  Levels  ............................................................................................................................... 82 4.3 Collagen Production of Fibroblasts  ................................................................................ 82 4.3.1 Effects of Fatty Acids on Collagen Production  ...................................................... 82 4.3.1.1 Fatty Acids and Hydrogen Peroxides Combined Increased Soluble  Collagen Production  ...................................................................................................... 82 4.3.1.2 Effects of a 24 h Combination of Oleic or Linoleic Acid with Hydrogen Peroxide on Deposited Collagen Production by Fibroblasts ...................................... 83 xi  4.3.2 Effects of Transforming Growth Factor-?1 and Fatty Acids on Collagen  Production in Fibroblasts  .............................................................................................. 84 4.3.2.1 Transforming Growth Factor-?1 Increased Soluble Collagen  ...................... 84 4.3.2.2 Linoleic Acid Increased Deposited Collagen Following Stimulation  with Transforming Growth Factor-?1  ......................................................................... 85 4.3.2.3 COL3 and TIMP1 Genes are Increased with Oleic and Linoleic Acid  plus Transforming Growth Factor-?1, Respectively  .................................................. 85 4.3.3 Levels of Collagen In Vivo  ........................................................................................ 87 4.3.3.1 TGF?3 is Increased Following a High Fat Olive Oil Diet  .............................. 87 4.3.3.2 Different Levels of Collagen Fibers Exist Between Diets and After  Cardiac Injury In Vivo  .................................................................................................. 88 4.4 The Effect of Hyperglycemia on the Fibrotic Response  ............................................... 89 4.4.1 Responses in Collagen Production are Different Between Low and  Glucose Conditions  ........................................................................................................ 89 5.0 Conclusions and Future Work  ............................................................................................ 91 5.1 Limitations of Research  ................................................................................................... 91 5.2 Future Work  ..................................................................................................................... 91 5.3 Significance of Findings  ................................................................................................... 91 References  ................................................................................................................................... 93 Appendix A: In Vivo Dietary Components  ............................................................................ 131       xii  List of Tables  Table 1. Primer sequences used for quantitative polymerase chain reaction  .............................. 47 Table 2. Components of In vivo diets  ........................................................................................ 131                           xiii  List of Figures  Figure 1. Antioxidant functions of glutathione  .............................................................................. 7 Figure 2. Determination of primer efficiencies using LinRegPCR  ............................................. 48 Figure 3. Effect of oleic or linoleic acid for 6 and 12 h on viability of cardiomyocytes  ............. 51 Figure 4. Effect of hydrogen peroxide after 6 and 12 h on cardiomyocyte viability  ................... 52 Figure 5. Effect of the combined treatment of hydrogen peroxide plus oleic or linoleic acid   for 6 and 12 h on cardiomyocyte viability  .................................................................... 54 Figure 6. Effect of linoleic or oleic acid for 6 h on fibroblast viability  ....................................... 55 Figure 7. Effects of oleic or linoleic acid over 24 h on reactive oxygen species production   in fibroblasts .................................................................................................................. 56 Figure 8. Effects of combined treatment of hydrogen peroxide plus oleic or linoleic acid for     24h on fibroblast viability  ............................................................................................. 58 Figure 9. Effect of the combined incubation with hydrogen peroxide plus oleic or linoleic          acid following 24 h on the levels of glutathione in fibroblasts  ..................................... 59 Figure 10. Effects of combined treatment of hydrogen peroxide with oleic or linoleic acid  following 24 h on soluble collagen production by fibroblasts  ................................... 61 Figure 11. Effects of combined treatment of hydrogen peroxide with oleic or linoleic acid following 24 h on deposited collagen by fibroblasts  ................................................. 62 xiv  Figure 12. Effect of transforming growth factor-?1 on soluble collagen levels in fibroblasts      after 24 h.  ................................................................................................................... 64 Figure 13. Effect of transforming growth factor-?1 on deposited collagen levels of         fibroblasts following 24 h  .......................................................................................... 65 Figure 14. Evaluation of the collagen gene expression in fibroblasts following combined treatment of transforming growth factor-?1 with oleic or linoleic acid for 24 h  ....... 66 Figure 15. Evaluation of the COL1?1/COL3 ratio in fibroblasts following treatment with transforming growth factor-?1 with oleic or linoleic acid for 24 h  ........................... 67 Figure 16. Effect of high fat feeding and mild cardiac injury using isoproterenol on collagen  gene expression in vivo  .............................................................................................. 69 Figure 17. Evaluation of the COL1?1/COL3 ratio after high fat feeding and mild cardiac    injury, using isoproterenol in vivo  ............................................................................. 70 Figure 18. Micrographs of histological heart sections following high fat feeding and mild  cardiac injury in vivo  .................................................................................................. 72 Figure 19. Effect of low and high glucose conditions on the production of soluble collagen        by fibroblasts following stimulation with transforming growth factor-?1 plus        oleic and linoleic acid treatment for 24 h ................................................................... 74 Figure 20. Effect of low and high conditions on the production of deposited collagen by fibroblasts following stimulation with transforming growth factor-?1 plus oleic        and linoleic acid treatment for 24 h  ........................................................................... 75  xv  List of Abbreviations  4-HNE  4-Hydroxy-trans-2-nonenal Acetyl-CoA Acetyl coenzyme A Ang II  Angiotensin II ANOVA Analysis of variance ATCC  American Type Culture Collection ATP  Adenosine triphosphate BLAST Basic local alignment search tool BSA  Bovine serum albumin CAT  Catalase cDNA  Complementary DNA CO  Corn oil CO2  Carbon dioxide CVD  Cardiovascular disease DCFDA 2?, 7? - Dichlorofluorescein diacetate DMEM Dulbecco?s Modified Eagle Medium ECM  Extracellular matrix EDTA  Ethylenediaminetetraacetic acid Em  Emission EtOH  Ethanol Ex  Excitation FA  Fatty acid FBS  Fetal bovine serum GPX  Glutathione peroxidase xvi  GSH  Glutathione GSSG  Glutathione disulfide H2O2  Hydrogen peroxide H9c2  Rat myoblast cell line HCl  Hydrogen chloride HF  Heart failure HG  High glucose IDT  Integrated DNA Technologies ISO  Isoproterenol KPi  Potassium phosphate buffer LA  Linoleic acid LG  Low glucose LDL  Low density lipoprotein LOX  Lysyl oxidase LV  Left ventricular MAPK  Mitogen-activated protein kinase MIQE  Minimum information for publication of quantitative real-time PCR experiments MMP  Matrix metalloproteinase MUFA  Monounsaturated fatty acid MTT  3-(4, 5-Dimethylthiazol-2-yl)-2, 5-diphenyl tetrazolium bromide NADPH Nicotinamide adenine dinucleotide phosphate NaOH  Sodium hydroxide NC  Normal chow NCBI  National centre for biotechnology information xvii  NEM  N-Ethylmaleimide NF-?B  Nuclear factor kappa-light-chain-enhancer of activated B cells NIH/3T3 Mouse fibroblast cell line O2-  Superoxide ion OA  Oleic acid OH-  Hydroxyl radical OO   Olive oil OPA  O-phthalaldehyde PA  Palmitic acid PBS  Phosphate-buffered saline PUFA  Polyunsaturated fatty acid qPCR  Quantitative polymerase chain reaction RNS  Reactive nitrogen species ROS  Reactive oxygen species RT  Room temperature SEM  Standard error of the mean SFA  Saturated fatty acid SOD  Superoxide dismutase TIMP  Tissue inhibitor of matrix metalloproteinase TNF  Tumor necrosis factor TGF?  Transforming growth factor beta TGF?R Transforming growth factor-beta receptor    xviii  Acknowledgements  Above all, I would like to thank my family for their incredible support throughout this degree. Thanks to my fianc? for his love, support and understanding while I spent many hours and days with research, data and papers (eyes glued to a computer screen). Thanks to my grandparents for their genuine interest in my research. Great thanks to my mother, who was always there to lend a helping hand, even proofreading pages and pages of thesis drafts. I would like to express my appreciation to my supervisor, Dr. Sanjoy Ghosh, for his guidance throughout my degree. Thank you for your support throughout award application deadlines, conference presentations and research questions. I would also like to thank my committee members, Drs. Philip Ainslie, Mary Forrest and Deanna Gibson for their time, support and advice over the last couple of years. I would like to thank my lab colleagues, Amy Botta, Kirsty Brown and Daniella Decoffe, for all of their help both in and out of the lab.              xix  Dedication  To my family. I will be forever grateful for all of your love, support and encouragement.                              1  1.0 Introduction 1.1 Cardiometabolic Disease Approximately 23% of Canadian adults are obese and an additional 36% are overweight [1] and similarly, 37% of American adults are obese and 30% are overweight [2]. Such high incidences of weight gains in the North American population have given rise to cardiometabolic disease or metabolic syndrome, with an increasing prevalence parallel to the increasing incidence of obesity [3]. Approximately 19% of adults in Canada [4] and 34% of adults in the United States [5] are recognized to have cardiometabolic disease. Cardiometabolic disease is a term that includes a broad cluster of metabolic risk factors including abdominal obesity, hypertension, insulin resistance with/without glucose intolerance, proinflammatory and pro-thrombotic states, and dyslipidemia [6], all of which may result in the development of diabetes and/or cardiac disease. Several diagnoses of cardiometabolic disease are described according to individual organizations. For example, the World Health Organization?s diagnosis requires impaired glucose tolerance plus two other factors [7], the International Diabetes Federation lists abdominal obesity plus two other factors for their diagnosis [8], whereas the National Cholesterol Education Program/Adult Treatment Panel III?s diagnosis is more general in that it requires three or more of the five criteria [9]. The two primary outcomes of cardiometabolic disease, diabetes and cardiovascular disease (CVD) are public health concerns. The first outcome is diabetes, which consists of type 1 and type 2, where type 2 comprises 90% of all diabetic individuals [10]. Currently 2.4 million (6.8%) Canadian citizens [11] and 22.3 million (7%) United States [12] citizens suffer from diabetes. Additionally, more than 350 million individuals worldwide have diabetes [13], which accounts for 4.6 million deaths [14]. Diabetes and CVD are intimately linked. In fact, an 2  estimated 80% of type 2 diabetic-related mortalities are due to CVD or cardiac disease [15]. Cardiac disease not only costs $20.9 billion in Canadian health care costs [16] but is a major contributor to Canadian deaths, being responsible for 29% of all mortalities [17]. In short, cardiometabolic disease and more specifically diabetes and cardiac disease are major public health issues. 1.2 Oxidative Stress: A Common Link One common link between diabetes and cardiac disease is oxidative stress. Oxidative stress is as an imbalance between prooxidant and antioxidant species [18]. The prooxidant species refer to highly reactive oxygen and nitrogen species or ROS and RNS respectively, whereas antioxidants are molecules that exist to combat ROS/RNS. Under normal conditions, the most common source of ROS is aerobic respiration from the mitochondria [18]; however, other stressors such as radiation exposure, heat stress and nutrient availability contribute to ROS production [19]. Attractive targets for ROS are the three major macromolecules, which are proteins, DNA and lipids. For example, the interaction between ROS and these macromolecules results in the inactivation of critical enzymes and denaturation of essential proteins [20], breaks in DNA-strands and changes in gene expression [21], and irreversible damage to lipid membranes [22]. Oxidative stress is typically defined as a negative process but is also unavoidable [23] as it is required in normal cell function. For example, in immune cells like macrophages, ROS are required for the respiratory burst, which functions in the destruction of invading pathogenic [24], altered self [25] and apoptotic [26] cells. Two main mechanisms of ROS exist in the respiratory burst. The first mechanism functions through nicotinamide adenine dinucleotide phosphate (NADPH), which produces superoxide ions (O2-) and eventually hydrogen peroxide (H2O2), 3  which destroy the cells [27]. The second mechanism works through the induction of inducible nitric oxide synthase [28] to generate nitric oxide (NO), which further reacts with O2- to form peroxynitrite. The highly reactive ROS that are formed function as the mediators of cell death during the respiratory burst. In addition, in non-immune cells such as muscle and adipose, ROS are involved in the insulin-signaling pathway. For example, low levels of ROS initiate the phosphorylation of surface tyrosyl residues in order to activate insulin receptors [23]. In addition to receptor activation, this phosphorylation inhibits protein tyrosine phosphatases, which then enhances the insulin-signaling cascade [23]. Lastly, ROS signaling is required in the apoptosis pathway. Apoptosis is required within all biological systems in order to eliminate cancer, self-altered, and foreign cells. Apoptosis occurs through oxidative damage to cellular DNA [29] and intracellular proteins [30]. 1.2.1 Superoxide and Nitric Oxide Bioavailability If O2- is not converted into H2O2, the O2- can react with NO to form peroxynitrite. The interaction between NO and O2- occurs 6 times faster than the enzymatic removal of O2-. This interaction depletes the bioavailability of NO and therefore affects the vascular functions that are dependent on NO, e.g. blood pressure and vascular cell signaling [31]. Less availability of NO is believed to increase endothelial dysfunction in the presence of hypertension [32] and atherosclerosis [33]. 1.2.2 Biological Role of Hydrogen Peroxide H2O2 is an endogenous ROS produced during the dismutation of O2- radicals [34]. The O2- radicals are produced during aerobic respiration due to the leakage of electrons from the mitochondrial electron transport chain. It has been observed that the rate of mitochondrial H2O2 emission reflects the rate of electron leakage and O2- formation in the cell [35]. 4  H2O2 is categorized as a ROS but is not technically a radical molecule due to its lack of unpaired electrons. Although not radical in nature, H2O2 causes damage at concentrations as low as 15 uM [36]. Due to its non-radical nature, it is more stable with a longer life span than other ROS [37] allowing for interaction with biological macromolecules near and far from its site of origin [38]. Within biological systems, H2O2 causes damage through both direct and indirect mechanisms. Directly, H2O2 causes protein degradation, enzyme inactivation [39], DNA [40] and lipid [41] oxidation via oxidative modification. Indirectly, if H2O2 is not converted into water, it acts as an intermediate source for the production of other ROS such as the hydroxyl radical (OH-) [42]. OH- has a short life span and is the most reactive ROS in biological systems [43]. Unlike H2O2, OH- has a short life span, which prevents it from causing widespread damage like H2O2. Functionally, the OH- radical interacts with DNA, proteins and lipids resulting in cytotoxicity and cell death [43]. 1.2.3 The Role of Antioxidants in Oxidative Stress Antioxidants are chemical species that exist to prevent ROS-mediated oxidative damage by decreasing the concentration of ROS or by neutralizing ROS. Antioxidants remove or neutralize ROS by accepting or transferring electrons from ROS, which results in the deactivation of ROS or reactions occurring due to ROS [44]. The two types of antioxidants that exist are exogenous and endogenous, both of which neutralize prooxidants, and thus decrease cellular oxidative stress. Exogenous antioxidants are provided in the diet and some examples include polyphenols, lipoic acid and vitamins C and E 5  [45]. However, although exogenous antioxidants are required within the body, endogenous antioxidants are considered more vital because they possess more potent activity [46]. Endogenous antioxidants are synthesized de novo and include enzymatic antioxidants such as superoxide dismutase (SOD), catalase (CAT) and glutathione peroxidase (GPX) as well as non-enzymatic antioxidants like glutathione (GSH). These antioxidants are present in all areas of the cell including the nucleus, cytosol, mitochondria and cell membranes [47]. The enzymatic antioxidants are the initial defense system against oxidative stress and therefore their regulation is extremely important [48]. SOD is known to protect the majority of tissues from oxidative stress [49]. SODs defend against oxidative stress by catalyzing the dismutation of the O2- ions to H2O2 and molecular oxygen. The majority of H2O2 production is the result of this dismutation reaction [49]. The three isoforms of SOD are SOD1, SOD2 and SOD3 [50]. SOD1 is the first isoform and is present in the intermembrane spaces of the cytoplasm and mitochondria. The second isoform, SOD2, is found within the matrix of the mitochondria and is most concentrated in cardiomyocytes. In fact, SOD2 mutations are associated with cardiomyopathy [51]. The final isoform, SOD3, is found in a secreted form in the extracellular space [52]. The second important line of antioxidant defense is CAT. CAT catalyzes the reaction of H2O2 to oxygen and water [53]. Most cells contain CAT but at various concentrations. For instance, CAT is observed at high levels in the liver and erythrocytes [54] and at very low levels within the heart [53]. CAT is completely absent in human endothelial and vascular smooth muscle cells, which are important within the heart [55]. 6  GPX is the last major line of defense against oxidative stress. GPX is found within the cytoplasm and can inactivate both H2O2 and lipid peroxides [56]. Similar to CAT, GPX catalyzes the reaction of H2O2 to oxygen and water, but unlike CAT, this reaction requires reducing equivalents from GSH. In fact, levels of GSH correlate closely to the levels of GPX [57] in the body. Although both CAT and GPX catalyze the same reaction, GPX is the main antioxidant defense against H2O2 [58]. This difference in function is due to an overall higher concentration of GPX in cells than CAT, except for in peroxisomes [24] where CAT is dominant [25]. Indeed, the antioxidant function of CAT functions primarily to neutralize H2O2 within the peroxisomes [53]. The antioxidant function of GPX, and not CAT, is critical in the myocardium. In fact, CAT has been observed to have a lesser role in H2O2 detoxification within the heart [26]. For example, a mouse model deficient in GPX-1 results in abnormalities of the cardiac structure and also leads to cardiac endothelial dysfunction [59]. In addition, this trend is seen in humans, as patients who have low GPX-1 activity are observed to have a higher risk for cardiac disease [60]. 1.2.4 Importance of Glutathione in Biological Systems GSH (?-gluatmylcysteinylglycine tripeptide) is a ubiquitous thiol that exhibits various non-enzymatic antioxidant functions. GSH exists in two forms, which are ?reduced? called GSH and ?oxidized? called GSH disulfide (GSSG) [61]. However, only the reduced form (GSH) acts as an antioxidant. The main function of GSH is to act as the cofactor for GPX. For example, through electron donation, GSH facilitates the detoxification of H2O2 and lipid hydroperoxides [57] by GPX. The active form (GSH) is converted to the inactive form (GSSG) after it donates reducing equivalents. At this point, the inactive GSSG must be converted back into the active GSH by 7  GSH reductase [22] in the GSH-recycling pathway. GSH also functions in the regeneration of other antioxidants such as lipoic acid [57], ascorbic acid (vitamin C) [62] and ?-tocopherol (vitamin E) [57]. GSH also demonstrates the ability to neutralize or remove ROS. For example, GSH directly scavenges singlet oxygen [63] and OH- [64] radicals resulting in their neutralization. In addition, GSH neutralizes peroxynitrite and at the same time forms S-nitrosoglutathione, a substance that protects against oxidative stress [65]. GSH also functions in the removal of harmful ROS and other species from the cell. More specifically, GSH conjugates with harmful electrophiles and xenobiotics via GSH-S-transferase [57] in order for their removal from the cell. An example of a harmful species that GSH removes is 4-hydroxy-trans-2-nonenal (4-HNE), a highly toxic lipid peroxidation product of linoleic acid (LA) [66]. Figure 1 demonstrates the functions of GSH explained above.  Figure 1. Antioxidant functions of GSH. Abbreviations: DHAR, dehydroascorbate; GPX, glutathione peroxidase; GR, glutathione reductase; GSH, glutathione; GSNO, s-nitrosoglutathione; GSSG, glutathione disulfide;  GST, glutathione-s-transferase; H2O2, hydrogen peroxide; HNE, 4-hydroxynonenal; NADH, nicotinamide adenine dinucleotide; NADPH, nicotinamide adenine dinucleotide phosphate; NO, nitric oxide; ONOO-, peroxynitrite; TOH, alpha-tocopherol; VC, vitamin C. 8  1.3 The Heart: Structure and Function One major part of the cardiovascular system is the heart. The heart is an obligate aerobic organ [67] that consists of three layers of tissue: the epicardium, the myocardium and the endocardium [68]. The outermost layer or the epicardium, also known as the visceral pericardium, consists of connective tissue and functions as a protective layer for the heart. The middle layer is the myocardium and it is the thickest layer comprised of cardiac muscle cells termed cardiomyocytes. The final and innermost layer, the endocardium, comprises of a single layer of endothelial cells lining the chambers of the heart. The heart or cardiac muscle utilizes various metabolic fuels such as glucose, lactate, ketones , amino acids and fatty acids (FA) [69]. Under normal physiology, the cardiac muscle prefers to utilize FAs as its main metabolic fuel. However, when FA availability is low, the heart will switch to glucose as its primary fuel source [67]. The heart derives 60 ? 70% of its total acetyl-CoA from FA oxidation [70], whereas the remainder is derived from glucose oxidation and glycolysis. FA oxidation occurs under aerobic conditions and therefore the heart requires a constant supply of oxygen. Due to the high oxygen consumption and mitochondrial density of the heart, the susceptibility to oxidative stress is high. During cardiometabolic conditions such as diabetes, the total amount of energy derived from FAs increases within the heart [71]. Large amounts of FA oxidation occur within the heart; therefore, the impact of various dietary FAs may be vital in regards to both beneficial and detrimental effects within the heart. For instance, high levels of lipid peroxidation have been closely correlated to oxidative damage in both ageing and metabolically challenged hearts [72]. 9  1.3.1 Cardiac Cells The heart is a heterogeneous biological system. Different cell types make up the heart and are involved in its proper function. Other than endothelial cells, which are considered a part of the vasculature, the two important cardiac cells are cardiomyocytes and fibroblasts. 1.3.1.1 Cardiomyocytes Cardiomyocytes are the muscle cells of the heart and their main function is in cardiac contraction via electrical signaling [73]. In order for proper signal conduction to occur, large amounts of adenosine triphosphate (ATP) are required. This ATP requirement is fulfilled by the large stores of calcium [74] and the large amount of mitochondria (about 40% of the cardiac volume) present within the heart [75]. Calcium regulates mitochondrial function and ATP synthesis. Mitochondrial calcium activates mitochondrial metabolic machinery to produce ATP [76]. In addition to ATP production, the mitochondria, by taking up excess calcium, inhibit calcium overload and improper contraction [77]. Calcium overload also results in the production of ROS, induction of the permeability transition pore, release of cytochrome c and apoptosis [78]. Cardiomyocytes connect to form myofibers. Individually, cardiomyocytes consist of myofibrils, which are made up of contractile units termed sarcomeres. These sarcomeres consist of contractile proteins such as actin and myosin [79]. The thick myosin and thin actin filaments interact with one another in a repetitive cycle in order to produce cardiac contraction [67]. Other proteins that comprise the sarcomere structure are the troponins and tropomyosin [80]. These proteins play an important role in cardiac regulation. For example, during relaxation, troponin I and tropomyosin inhibit actin-myosin interaction [81, 82]. 10  1.3.1.2 Fibroblasts Although 75% of the myocardial volume is comprised of cardiomyocytes [83], fibroblasts account for 70% of the total myocardial cells by number [84]. Cardiac fibroblasts are found within the connective tissue of the myocardium [85] and are characterized based on their morphological and phenotypical attributes. Morphologically, fibroblasts are observed as flat spindle-shaped cells with multiple cytoplasmic extensions [86]. Fibroblasts also play a role in cardiomyocyte function. For example, through the production of mitogens, fibroblasts function in the development of cardiomyocytes and the remodeling of the adult cardiomyocyte phenotype. More specifically, cardiac fibroblasts induce hypertrophy and reduce contractility of cardiomyocytes [87]. Cardiac fibroblasts are important in the maintenance and homeostasis of the extracellular matrix (ECM). The ECM is an organizational network that surrounds, connects and provides scaffolding [88] for cardiac cells. Its main components are fibrillar collagens, glycoproteins, proteoglycans, growth factors and cytokines [89, 90]. Functionally, the ECM distributes mechanical force throughout the myocardium, transmits signals [91] and ensures proper cardiac contraction [92]. The homeostasis of the ECM is achieved by synthesis and degradation of the structure and its components. Cardiac fibroblasts are the main cell type involved in production of the cardiac ECM [93] and in addition are involved in the production of biochemical mediators such as growth factors [94]. For example, fibroblasts maintain the ECM through the synthesis/secretion and degradation of collagen types I and III [95]. Growth factors and cytokines can further stimulate fibroblasts to produce type I and III collagens, which comprise about 90% of all collagen within the heart [96]. 11  1.4 Heart Failure Heart failure (HF) occurs when the heart can no longer pump blood at a rate that is required for metabolic needs or can only fulfill metabolic requirements with an abnormally high filling pressure. HF may be a final manifestation of cardiac diseases, such as myocardial infarction, hypertension and cardiomyopathies, and is accompanied by cardiac remodeling [68]. Chronic HF is the result of various cardiovascular insults that produce systolic and/or diastolic dysfunction. When systolic dysfunction is present, the ventricle has impaired contractility or pressure overload, resulting in a decreased capacity to eject blood. On the other hand, diastolic dysfunction of the ventricle exhibits impaired early diastolic relaxation and/or increased stiffness of the ventricular wall [68]. HF most often occurs due to left ventricular (LV) dysfunction. Commonly, right-sided HF occurs due to the presence of left-sided HF yet the presence of right-sided HF without left-sided HF is uncommon. Several compensatory mechanisms against HF exist to preserve blood flow to all organs however prolonged compensation results in eventual remodeling and ventricular dysfunction. Compensatory mechanisms include changes in the Frank-Starling mechanism, alterations of the neurohormonal system and hypertrophy or remodeling of the left ventricle [68]. The first mechanism includes cardiomyocyte stretching via the Frank-Starling mechanism to create a higher stroke volume in order to preserve cardiac output. The second mechanism occurs through alterations in neurohormonal systems such as the renin-angiotensin-aldosterone system resulting in increased levels of angiotensin II (Ang II) and aldosterone. Increased secretion of hormones such as vasopressin and peptides such as endothelin-1 are also observed. The last compensatory mechanism is hypertrophy or remodeling of the left ventricle. Increased cardiac mass initially decreases wall stress and maintains contractile force. However, 12  these compensatory effects are accompanied by increased ECM deposition and myocardial stiffness [68]. All compensatory effects start as beneficial but overtime lead to increased cardiac dysfunction and failure. 1.4.1 Cardiomyopathy Cardiomyopathies arise primarily due to disorders of the cardiac muscle or myocardium. This group of heart disorders exhibits similar symptoms of HF but the etiologies for most are unknown. Cardiomyopathies do not include diseases characterized by cardiac muscle impairment from defined vascular conditions, such as hypertension or coronary disease. Cardiomyopathies are divided into three groups: (1) dilated cardiomyopathy, characterized by ventricular chamber enlargement and impaired systolic contraction, (2) hypertrophic cardiomyopathy, characterized by an abnormally thickened ventricular wall with abnormal diastolic function and (3) restrictive cardiomyopathy, characterized by an abnormally stiffened myocardium and impaired diastolic relaxation with normal systolic contractile function maintained [68]. Obesity cardiomyopathy has also been established [97, 98] and results in LV remodeling and ventricular dilation [99]. The pathology of obesity cardiomyopathy includes ventricular remodeling, myocardial abnormalities [97], which may be due to metabolic disturbances such as insulin resistance [100] and lipotoxicity [101]. Animal models of obesity cardiomyopathy have demonstrated contractile dysfunction [100]. 1.4.2 Key Reasons for Heart Failure: Cardiac Remodeling Most, if not all, cardiac diseases result in HF and cardiac remodeling. Cardiac remodeling is a physiologic condition that results from cardiac injuries/disease such as pressure or volume overload, myocardial infarction, inflammatory heart disease and cardiomyopathy. These 13  conditions are influenced by either hemodynamic or hormonal factors. Although these conditions have different etiologies, they share similar molecular, biochemical and mechanical responses. Cardiac remodeling has been defined as the cellular, molecular and interstitial changes following cardiac injury [102]. More specifically, changes in shape, size and function of the heart exemplify these broad terms. At first, these changes are compensatory or adaptive responses but when overwhelmed will lead to increased size (hypertrophy) of the heart [103] as well as cardiac dysfunction [104]. For example, the heart as well as individual cardiomyocyte cells become hypertrophied because of increased workload within the heart [105]. The structure of the heart changes during cardiac remodeling. The shape becomes less elliptical and moves toward a more spherical shape [106]. In addition, changes in ventricular mass, composition and volume of the heart occur, which leads to negative effects on cardiac function [103]. The remodeling process is focused primarily on the cardiomyocyte. During remodeling, hypertrophy of cardiomyocytes is often accompanied with cardiomyocyte cell death (apoptosis or necrosis) [102]. Apoptosis or necrosis of cardiomyocytes is an important part of HF and the remodeling process. B-cell lymphoma 2, an important regulator of cell apoptosis, promotes increased apoptosis during hypertrophy [107]. Cardiomyocyte death is associated with changes in the ECM and collagen matrix [108] around individual cardiomyocytes. Following cardiomyocyte cell death, the remodeling process leads to replacement of the surrounding necrotic area and may lead to scar formation (fibrosis), which at first may be adaptive and beneficial [109]. However, once this response is overwhelmed, excess fibrosis and remodeling of the ECM will occur. For example, pressure overload induces cardiomyocyte cell death and has demonstrated increased levels of deposited collagen between cardiomyocytes [110], which results in myocardial stiffness and cardiac 14  dysfunction. On the other hand, hypertrophy demonstrates decreased ECM components due to increased degradation of the ECM, which results in less cardiac cell support and ventricular dilation [111]. This degradation occurs through matrix metalloproteinases (MMPs), which are involved in the remodeling process [112]. Although different responses in the ECM remodeling occur, they ultimately lead to cardiac dysfunction. Research on the remodeling process has focused on LV remodeling because it is a main pathological feature of cardiac disease. A mechanism of LV remodeling at the molecular level has been proposed. When cardiomyocytes stretch, they cause increased norepinephrine activity [113] and increased release of Ang II and endothelin [114, 115]. These molecular changes stimulate cardiomyocyte hypertrophy and protein alteration, resulting in decreased cardiac function. Other molecular changes such as increased activation of aldosterone and various cytokines have demonstrated increased collagen synthesis and ECM remodeling with increased fibrosis [116, 117]. 1.5 Oxidative Stress in Cardiac Disease, Heart Failure and Remodeling A causative role for HF has been identified as myocardial oxidative stress. For example, markers of oxidative stress are highly elevated in patients with myocardial dysfunction and HF [118]. Furthermore, oxidative damage may initiate the myocardial remodeling in HF and more specifically alterations to the cardiomyocyte and myocardial ECM structure and function. Different oxidative stress markers have been implicated in the pathology of cardiac remodeling. For instance, levels of 8-iso-prostaglandin F2?, a marker of lipid peroxidation, have been shown to be elevated in hypertrophic cardiomyopathy [119]. In addition, the lipid peroxidation product, 4-HNE, is elevated in the failing myocardium of hypertrophic cardiomyopathy patients and is implicated in the exacerbation of HF. These harmful peroxidation byproducts and ROS result in 15  contractile dysfunction within cardiomyocytes. For example, 4-HNE increases the generation of H2O2 and O2- ions and in doing so initiates a calcium overload, resulting in hypercontracture of the cells and failure of the myocardium [120]. In addition to 4-HNE, OH- radicals have demonstrated the ability to decrease the contractile systolic force through increased diastolic tension, reduced response to extracellular calcium and impaired force-frequency relationship [121]. Cardiac dysfunction and disease ultimately results in cardiac remodeling and HF. Redox-sensitive changes have been implicated in the progression of cardiac hypertrophy. For example, prooxidants affect the excitation-contraction coupling by producing redox-sensitive changes [122] in cardiac cells and their key proteins. Two redox-sensitive signaling factors are mitogen-activated protein kinase (MAPK) and nuclear factor kappa-light-chain-enhancer of activated B cells (NF-?B) [122, 123]. These factors can also be activated in a ROS-dependent manner by tumor necrosis factor-alpha (TNF?) and Ang II, both of which induce a hypertrophic response [124]. For example, an increase in H2O2 by Ang II, results in the phosphorylation and subsequent activation of p38MAPK, which activates a growth signaling cascade and causes vascular hypertrophy [125]. Similarly, NF-?B, which is activated by TNF? in a ROS-dependent manner, results in increased cardiomyocyte growth. This response is thought to be due to an interaction with cardiotrophic transcription factors [126]. As mentioned earlier, changes to the ECM typically result in fibrosis, which has been linked to oxidative stress. One mechanism of oxidative stress occurs via the MMPs that are responsible for ECM degradation. Oxidative stress, measured by 8-iso-prostaglandin F2? and induced by H2O2, has demonstrated augmented MMP-2 and MMP-9 activity in cardiac fibroblasts [127]. H2O2 has been shown to decrease collagen synthesis [128]; however, a more recent study has shown the opposite trend and has demonstrated increased collagen synthesis 16  instead [129]. In agreement with the latter, concurrent administration of iron, known to increase levels of the OH- radical, with Ang II resulted in an increased fibrotic area [130]. 1.6 Fibrosis 1.6.1 Role of Fibroblasts in Cardiac Remodeling Fibrosis is the process of scar formation (made up of cross-linked collagen and other ECM components [109]) and functions through fibroblast accumulation, increased synthesis/deposition [131] and decreased degradation [132] of the ECM. These phenomena result in stiffness of the myocardium and pathological signaling, which can ultimately result in cardiac disease and HF [133]. Myocardial fibrosis is a double-edged sword. Following injury, fibrosis is required for wound healing, yet it also results in ventricular stiffening and progression into HF. When the heart becomes injured, either the same cell type (regeneration) or fibrotic tissue replaces the cell debris. In the case of cardiomyocyte death, replacement with fibrotic tissue must occur because cardiomyocytes are terminally differentiated and therefore they cannot regenerate. After cardiomyocyte death, the space previously occupied by the cardiomyocytes must be replaced with fibrotic tissue (scar) [134]. Some factors that lead to cardiac fibrosis are pressure overload [135] and other non-hemodynamic factors such as hormones [136] and growth factors [137]. Two types of fibrosis exist ? reactive interstitial and replacement fibrosis [138]. Reactive interstitial fibrosis occurs with no loss of cardiomyocytes and therefore functions as an adaptive mechanism to preserve cardiac function [139]. However, if this adaptive response becomes overwhelmed, replacement fibrosis, which is accompanied by cardiomyocyte hypertrophy and cell death [140], will occur. Unfortunately, the fibrotic process does not always occur through this two-step fibrotic process. For instance, a model of acute myocardial infarction demonstrated cardiomyocyte cell death with direct replacement fibrosis [141]. 17  After fibrosis occurs, the normal electrical signals between cardiomyocytes are impeded. Following fibrosis, fibroblasts initially increase cardiac signaling as an adaptive response [142]. The fibrotic tissue impedes the electrical stimuli but fibroblasts possess the ability to conduct electrical signals [143] due to their high cell membrane resistance [144]. For example, fibroblasts couple to cardiomyocytes through connexins-43 and -45 [145] in order to allow for the conduction of electrical signals to continue [146]. In addition, this coupling allows for the formation of bridges between cardiomyocyte regions over deposited ECM [147], which also allow the electrical signals to continue. In a normal healthy heart, collagen deposition is very limited; therefore, the degradation of the ECM is important in homeostasis. MMPs are proteolytic enzymes that degrade the components of the ECM including collagen [148]. MMPs are normally present within the myocardium [149] and are produced by fibroblasts, myofibroblasts and cardiomyocytes [150, 151]. Under normal physiological circumstances, MMP activity is tightly regulated by tissue inhibitors of MMPs (TIMPs) [152], both of which are regulated at the transcriptional level by cytokines and growth factors [153, 154]. When an imbalance between MMPs and TIMPs exist, typically during disease physiology, an initiation of pathologic remodeling occurs. This results in disrupted myocardial structure and function, and accumulation of collagen [155]. During pathological states, the regulatory mechanisms of fibroblasts become altered, which typically results in excessive and persistent collagen production [156]. The increase in collagen and other ECM components deposited between cardiomyocytes results in myocardial stiffness, diastolic dysfunction [156, 157] and impaired electrical signaling or cardiac contraction [158]. Remodeling of the ECM is important in HF and cardiac remodeling. The two main pathological features of cardiac disease are cardiomyocyte hypertrophy [159] and cardiac 18  fibrosis [160], both of which result in HF and remodeling [134]. The increased fibrotic tissue is due to the increased synthesis of collagen types I and III [161, 162] and the decreased degradation of collagen by MMPs [156]. For example, reduced levels of serum MMP-1 [156], MMP-2,  MMP-9 [163] and increased levels of TIMP-1 [164] are often present in cardiac diseases. In addition, levels of cardiac fibrosis correlate with the degree of cardiomyocyte hypertrophy [165]. 1.6.2 Diabetes and Cardiac Fibrosis Diabetes has been intimately linked to cardiac disease and studies have demonstrated that diabetic cardiomyopathy exists [166] and shows a higher incidence of HF [167]. The associated HF and remodeling demonstrate hypertrophy and fibrosis [168], similar to other cardiac diseases. More specifically, increased deposition of collagen types I and III [169] and increased collagen cross-linking [170] are observed. It is well known that wound healing is impaired in diabetes. Diabetes stimulated in vitro, under high glucose conditions (25 mM D-glucose), demonstrated impaired cell migration, which may be a factor in poor wound healing [171]. However, most in vitro studies focused on fibrosis demonstrate increased collagen synthesis and fibrosis in high glucose conditions. For example, skin fibroblasts demonstrate increased collagen III production [172]. Although few reports exist on the effects of high glucose on cardiac-specific fibroblasts, a similar trend is seen in regards to fibrosis. For instance, in neonatal rat cardiac fibroblasts, increased collagen synthesis plus increased fibronectin and TGF?1 gene expression is observed [173]. In addition, this same study showed that high glucose may promote fibrosis through increased expression of fibrous protein and decreased activity of MMP. In agreement, adult rat cardiac fibroblasts synthesized increased protein and collagen under high glucose conditions [174]. 19  1.6.3 Signaling Molecules of the Fibrotic Process within the Heart During normal and pathological physiology, a distinct pattern of cytokines, growth factors and ECM structure exist in the heart [175, 176]. This pattern may be responsible for the fate of all progenitor cells recruited to the heart. For example, a pro-fibrotic microenvironment promotes pathologic myofibroblast differentiation [177] and has also been linked to the pathology of inflammatory HF [178]. 1.6.3.1 Angiotensin II Signaling Ang II is a potent vasoconstrictor that is involved in the renin-angiotensin aldosterone system [68]. Ang II plays a role in sodium retention, vasoconstriction [68] and cardiomyocyte growth [179]. Not only does Ang II indirectly affect the heart, but it can also directly stimulate fibroblast proliferation [180], cytokine production [181] and ECM synthesis [182]. In cardiac fibroblasts, Ang II interacts with Ang II type I receptor to inhibit MMP-1 activity and therefore decreases collagen degradation [183]. In addition, Ang II regulates osteopontin, a cytokine that is associated with extensive fibrosis [184], TGF?1 [185] and plasminogen activator inhibitor-1 [186] in cardiac fibroblasts. It has been suggested that cross talk occurring between the factors produced by cardiomyocytes, macrophages and fibroblasts modulate the pro-fibrotic effects of Ang II [187]. 1.6.3.2 Transforming Growth Factor-? Signaling Transforming growth factor-beta (TGF?) is one of the major and most highly studied pro-fibrotic cytokines [188]. TGF? modulates various important cellular processes such as embryonic development, cell growth, differentiation and migration, proliferation and apoptosis. In addition, it regulates the production of collagen and deposition of the ECM [189]. Mammalian 20  TGF? consists of three isoforms ? TGF?1, TGF?2 and TGF?3 [190], where TGF?1 is the most prevalent isoform. Moreover, TGF?1 is highly expressed in the developing and adult heart and has been observed to localize in the cardiomyocytes and the ECM of adult murine hearts [191]. TGF?1 signals through a heteromeric receptor complex comprised of two serine-threonine kinase receptors, termed TGF? receptor types I and II [192] through downstream effectors known as the Smad proteins. Active TGF?1 binds to the constitutively expressed type II receptor (TGF?RII) located at the cell surface. The complex further interacts with the cytoplasmic domain of the type I receptor (TGF?RI) [193]. Phosphorylation of TGF?RI leads to the activation of the type I receptor kinase domain, which propagates the downstream intracellular signals through the Smad proteins [194] and translocation into the nucleus [195]. These two receptors are present within the heart and are expressed in cardiomyocytes and non-myocytes [196]. TGF?1 signaling and its downstream effects play a major role in fibrosis [197]. For instance, TGF?1 regulates fibrous tissue deposition through the conversion of fibroblasts to myofibroblasts [198], induction of collagen [199] and ECM synthesis and promotion of ECM preservation [200]. More specifically, TGF?1 induces recruited progenitor cells and fibroblasts into myofibroblasts [201] via Smad proteins [189]. It is known that myofibroblasts have a higher propensity to produce collagen [202] than fibroblasts. Furthermore, TGF?1 induces the deposition of ECM proteins (collagen, fibronectin, and proteoglycans) by cardiac fibroblasts [203]. In addition, the activation of the Smad2 or Smad3 signaling pathway results in TIMP upregulation [204] and thus decreased degradation of the ECM. In summary, TGF?1 increases collagen types I and III [205], decreases collagenase (MMP-1) expression, increases TIMP-1 synthesis [206], upregulates integrin expression and induces a myofibroblast phenotype [207]. 21  TGF?1 signaling occurs in other Smad-independent pathways [208]. Other factors known to activate TGF?1 signaling are MMP-9, MMP-2 [209] and ROS [210]. Furthermore, TGF?1 activates signaling cascades including extracellular signal-regulated kinases, c-Jun NH(2)-terminal kinase and p38 MAPK [211] and additionally, the pro-fibrotic effects of TGF?1 may be mediated partly through connective tissue growth factor upregulation [188]. The overexpression of TGF?1 results in cardiac hypertrophy and interstitial fibrosis within the heart [212]. Excess levels of TGF?1 accompanied by increased collagen synthesis and decreased collagen degradation are observed in patients of hypertensive heart disease [213]. 1.6.4 Soluble and Insoluble Collagen Collagen consists of a group of fibrous proteins. Soluble collagen is characterized by non-aggregated collagen fibers whereas insoluble collagen consists of aggregated collagen fibers. In fact, the bovine heart is observed to have the highest amount of insoluble collagen [214]. Collagen is synthesized by fibroblasts and myofibroblasts. Before synthesis, collagen exists in precursor forms of procollagen chains, with amino and carboxy-terminal propeptide extensions, found within the fibroblasts and myofibroblasts [215]. The procollagen chains are first combined into trimers [216]. During this process, the amino and carboxy-terminal propeptides are cleaved by procollagen proteinases to yield these triple helical monomers [217]. After secretion of procollagens, the terminal propeptides are removed by MMPs, triggering spontaneous self-assembly of mature collagen molecules into fibrils [218]. Fibrillar collagen types I and III must be cross-linked to form the final collagen types I and III fibers [219]. These fibers are highly resistant against proteolytic enzymes and exhibit physical properties of tensile strength. Once formed, a pre-existing network of fibronectin and integrins promotes the 22  deposition of collagen types I and III fibers into the ECM [220]. The initial cross linkage of the collagen fibers occur through lysyl oxidase (LOX) [219]. LOX, a copper-dependent amine oxidase, is an extracellular, matrix-embedded protein that plays a crucial role in the cross linking of the collagen fibrils and therefore the deposition of insoluble collagen fibers [221]. LOX catalyzes the oxidation of the amino groups in lysine or hydroxylysine residues, resulting in the formation of corresponding aldehydes. Either, two of these aldehydes can spontaneously react with each other or one aldehyde can bind to another amino group, both of which produce cross-links connecting two polypeptide chains [222]. LOX can be activated through other fibrotic signaling molecules. For instance, TGF?1 has been shown to stabilize LOX mRNA in rat [223] and human [224] fibroblasts resulting in increased synthesis of the LOX precursor. In addition, Ang II increases LOX as well as fibronectin expression via the activation of ras-related C3 botulinum toxin substrate 1 GTPase and connective tissue growth factor [225]. Furthermore, fibronectin has also been shown to facilitate LOX processing [226]. Oxidative stress has also been shown to mediate LOX levels. In fact, increased levels of mitochondrial ROS stimulate LOX-dependent collagen cross-linkage activity in human fibroblasts [227]. Levels of LOX have been used as an indicator for cardiac fibrosis. For example, increased levels of LOX accompanied by excess fibrillar collagen cross-linking and fiber deposition have been reported in LV dysfunction [228] and HF [229]. 1.7 Fatty Acids FAs are one of the four main macromolecules of the body. Structurally, FAs are carboxylic acids with a carbon chain and are categorized into either saturated or unsaturated FAs. Saturated fatty acids (SFAs) consist of a carbon chain that is saturated (i.e. no double bonds) whereas unsaturated FAs contain a carbon chain that is not saturated and contains one or more 23  double bonds [67]. Unsaturated FAs are further divided into monounsaturated fatty acids (MUFA), containing one double bond, or polyunsaturated fatty acids (PUFA), consisting of more than one double bond. The two main PUFA groups are omega-3 and omega-6 PUFA and their parent FAs are alpha-linolenic acid and LA respectively. At low dietary concentrations, PUFAs are required for cellular development and function. In fact, these two PUFAs are considered ?essential? FAs [230] and are required within the diet because they are not synthesized de novo. De novo production cannot occur because mammals lack the delta-3-desaturase, an enzyme required for insertion of double bonds within the carbon chain [231]. All fats including saturated, unsaturated and essential fats are consumed within the diet. For instance, SFAs are consumed through butter and other animal fats, containing palmitic and stearic acid respectively. However, plant sources of SFAs do exist and include coconut (lauric and myristic acid) and palm oil (palmitic acid; PA). The different groups of unsaturated fats are found in various sources of food. For example, the main MUFA is oleic acid (OA) and it is found in olive oil (OO) [232]. The two main groups of PUFA are found in very different sources. For instance, alpha-linolenic acid, the main omega-3 PUFA, is consumed from fish and fish oil [233], whereas the main omega-6 PUFA, LA, is found in animal meat and vegetable oils [234] including corn, safflower [235], soybean [236] and canola [237] oils. 1.7.1 The Current North American Diet Approximately 60% of North Americans including Canadians are overweight with a quarter of this value indicating obese individuals [238]. A Health Canada study reported that 50% of women and 70% of men have energy intakes greater than their nutritional needs [239]. Furthermore, this report indicated that over the last three decades Canadians have increased their 24  total caloric intake by 11%, which may be a contributing factor to increased national obesity rates [239]. The majority of this 11% increase was due to an increased intake of carbohydrates and fats. Interestingly, the intake of protein remained relatively stable. Further analysis indicated that increased dietary unsaturated FAs, more specifically of MUFAs (22%) and PUFAs (54%), dominated the increased consumption of fat. This specific increase has been attributed to the drastic replacement of SFAs with unsaturated fats that has occurred in the North American diet over the last few decades. The average consumption of LA, the main omega-6 PUFA, in the Canadian diet is approximately 7% energy, which exceeds nutritional requirements, whereas the dietary intake of alpha-linolenic acid and other omega-3 PUFAs is only 0.7% of energy [240]. These two PUFAs are functionally and metabolically different and therefore often lead to biologically different effects [241]. PUFAs regulate membrane formation [242], eicosanoid production [243] and  cellular signaling [244]. The regulation of membranes is essential in the maintenance of the composition, properties and function of cell membranes [245]. For instance, all membranes consist of a tightly regulated FA composition. Thus, the consumption of PUFAs can directly affect this composition, and therefore, intake of PUFAs should be regulated. The three main FAs (SFAs, MUFAs and PUFAs) are oxidized to different levels. PUFAs are highly susceptible to lipid peroxidation, whereas SFAs undergo very low levels of lipid peroxidation, due to the complete lack of double bonds within their structure [246]. Similarly, MUFAs undergo peroxidation at lower rates than PUFAs, because less double bonds are present within their structure. 25  1.7.2 Replacement of Saturated Fatty Acids with Polyunsaturated Fatty Acids Over the last three decades, the shift from SFAs to PUFAs in the diet was done in an attempt to reduce the prevalence of cardiometabolic diseases [247]. More specifically, SFAs were substituted with vegetable oils rich in LA [248]. Following suit, industrial animal farming also increased their feeding of LA-rich oils to animals [248], which led to faster weight gains. Therefore, the consumption of these animals and/or animal products resulted in increased dietary LA. 1.7.3 Monounsaturated Fatty Acids and their Role in Oxidative Stress Over the past few decades, research has demonstrated that MUFAs such as OA and palmitoleic acid are protective against lipid peroxidation and cardiac disease. Many studies have shown that saturated fats, specifically PA, cause detrimental effects to cells including cardiomyocytes, such as increased ROS production [249] and apoptosis [250]. However, the co-incubation of OA with PA decreased these detrimental effects. In addition, the Seven Countries Study demonstrated that certain Mediterranean populations, with a diet high in OO [251], had lower levels of plasma cholesterol and a lower prevalence of cardiac disease [252]. OO also showed beneficial effects on biomarkers associated with cardiac disease. For instance, a diet high in OO improves lipid profiles and reduces blood pressure [253]. Some researchers believe that the other components of OO (elaidic and stearic acid) are responsible for such beneficial effects but a study by Ter?s et al., demonstrated that OA and not the other components is responsible for the hypotensive effects of OO [254]. 26  1.7.4 Omega-6 Polyunsaturated Fatty Acids and their Effect on Cardiac Disease: Conflicting Evidence In regards to the risk of cardiac disease, there is conflicting evidence on the effects of the replacement of SFAs with PUFAs, mainly LA. Several meta-analyses have demonstrated possible detrimental effects of this switch [255, 256] from SFAs to LA [248, 257]. It was demonstrated that studies describing beneficial effects of omega-6 PUFA based their results on a diet consisting of omega-3 and omega-6 PUFA and not a specific diet of omega-6 PUFA [258]. From this conclusion, it was reported that omega-6 PUFA-specific diets demonstrated adverse effects on cardiac disease [258]. The omega-6 PUFA-specific diets exhibited an increased risk of 13% towards cardiac disease. This result was drastically different from studies showing a 24% risk reduction [259]. This observation clearly demonstrated that the lack of a consistent diet was a factor responsible for the conflicting evidence reported. 1.7.4.1 Beneficial Effects of Omega-6 Polyunsaturated Fatty Acids on Cardiac Disease Omega-6 PUFAs have demonstrated beneficial effects in regards to cardiac disease. Early research demonstrated that omega-6 PUFAs resulted in beneficial effects on atherosclerosis, arrhythmia and cardiac disease [260, 261]. For example, when 5% of SFAs were replaced with omega-6 PUFAs (mainly LA), there was decreased low density lipoprotein (LDL) levels [262], a 24% risk reduction for cardiac events [259] and a 26% risk reduction of deaths linked to cardiac disease [263]. Another study showed that a diet high in the omega-6 PUFA, LA, caused a decrease in whole plasma, LDL cholesterol by 20 ? 30% along with a protective effect against coronary artery atherosclerosis [260]. A diet high in PUFAs (mixed omega-3 and omega-6 PUFA) also improved insulin sensitivity [264] and reduced the risk of type II diabetes [265]. For example, a study replaced partially hydrogenated vegetable oil with SFA and demonstrated a 27  decrease in the risk of cardiac disease, indicating that SFA reduced the risk of cardiac disease [266]. The overall conclusion to these studies was that the replacement of SFAs with PUFAs was beneficial in regards to cardiac disease. 1.7.4.2 Adverse Effects of Omega-6 Polyunsaturated Fatty Acids on Cardiac Disease Although many beneficial effects of omega-6 PUFA were demonstrated, many adverse effects started to manifest in research as well. Several clinical studies and meta-analysis found no evidence between the intake of omega-6 PUFA and reduced risk of cardiac mortality [255, 267]. In fact, CO (high in LA) but not OO (high in OA) increased major cardiac events in patients with ischemic heart disease by 25% [268]. Furthermore, an excess intake of omega-6 PUFAs has been linked to the development of cardiomyopathy. For instance, rats that were fed a high-fat diet, enriched in LA, had thicker cranial left ventricle walls. In addition, these rats showed increased myocyte cross sectional areas. This type of early thickening (hypertrophy) of the LV wall and cardiomyocytes has been implicated in cardiac disease [269]. Another study raised concerns about high levels of dietary LA. This study looked at Israeli Jews, who have the world?s highest LA-intake (approximately 8% higher than USA and 11% higher than most other European nations at that time). Within their diet, this high intake of LA along with low levels of SFAs and total fats, show a higher incidence of cardiac disease, type 2 diabetes and cancer in Israeli Jews compared to non-Israeli Jews (termed as the ?Israeli Paradox?) [270]. The U.S. Veterans trial [271], involving 846 U.S. war veterans, were given supplemented diets with either CO (rich in LA) or animal fat for eight years. Eight years later, the total 28  mortality between diet groups was not different; however, there was a higher incidence of cancer-related mortalities in the CO diet. Excess dietary LA is linked to thrombotic and inflammatory events [272]. LA is metabolized into arachidonic acid, which produces potent pro-inflammatory and pro-arrhythmic molecules such as prostaglandins [236], thromboxanes [273] and hydroxyeicosatetraenoic acid [274]. An increase in these molecules could lead to increased inflammation and therefore increased detrimental effects on cardiac disease. A diet high in omega-6 PUFA has demonstrated increased risk of cardiac fibrosis, a pathological feature of most cardiac diseases. For instance, omega-6 PUFAs have been shown to upregulate levels of TGF?1. This upregulation resulted in a pro-fibrogenic and pro-inflammatory effect, which upregulated the activity of fibroblasts [275]. 1.7.4.3 Recent Clinical Evidence on Omega-6 Polyunsaturated Fatty Acids In 2010, a seminal meta-analysis summarized evidence from twenty-one studies related to cardiovascular risk with increased SFA consumption and concluded that such a relationship could not be established due to insufficient evidence [276]. The study also pointed out that most CVD or CHD could be linked to replacement nutrients used to replace SFAs in the last few decades, i.e. carbohydrates and LA [276]. With regards to LA, the Sydney Diet Heart Study was a clinical trial which evaluated the effectiveness of dietary LA on CVD and death risk reduction [277]. This study was conducted in the 1960s and 1970s and included 458 men with recent coronary events. The initial study suggested that SFAs raised LDL cholesterol, whereas LA lowered LDL cholesterol [278]. A decrease of 13% in total cholesterol with LA was shown; however, a higher all-cause mortality was reported in 1978 [279], without increased mortality 29  form CVD and coronary heart disease (CHD). In 2013, Ramsden et al. shed new light on this study as new analysis concluded that the replacement of SFAs with LA actually increased the rate of death among the patients and that no cardiovascular benefits could be associated with increased dietary LA [277]. Instead, results confirm the LA group had higher risk of all cause mortality as well as increased mortality from CVD and CHD. Using the newly analyzed data from the Sydney Diet Heart study, two more LA intervention trials were analyzed [268, 280] and the new results demonstrated an increased risk of death from CHD and CVD overall in human patients. 1.8 Research Overview, Hypothesis and Aims Due to the high consumption of FAs within the diet, a large portion of research has focused on the effects of SFAs, MUFAs and PUFAs on cardiac health. SFAs have been the main fat blamed for cardiac disease, so over the last few decades, SFAs were replaced with PUFAs to reduce the risk of cardiac disease. Interestingly, the risk of cardiac disease has not decreased but increased instead. Most research has focused primarily on SFAs and MUFAs; however, recently more studies focused on PUFAs have been becoming more prominent [281-283]. The North American diet contains high and even excess levels of LA, which has been associated with oxidative stress and cardiac disease. The main cells that are involved in cardiac disease are cardiomyocytes and fibroblasts, which maintain cardiac structure and function. However, excess dietary fats and oxidative stress may play a role in cardiac disease and more specifically on these two cell types. It is known that once cardiomyocytes undergo cell death, fibroblasts produce fibrosis in order to maintain cardiac structure and contractility. However, if this process becomes excessive, cardiac fibrosis results in HF and remodeling. The central 30  hypothesis of this research thesis is that excess LA (the main dietary omega-6 PUFA) in the diet initiates cell death and fibrosis within the heart.  The research aims of this hypothesis are: 1. Does LA with/without oxidative stress cause increased cardiomyocyte cell death? 2. Does LA with/without oxidative stress affect fibroblast viability and GSH levels? 3. Does LA cause cardiac fibrosis? 4. Does hyperglycemia alter the fibrotic response to FAs?           31  2.0 Materials and Methods 2.1 Chemicals and Reagents The following reagents were obtained from Sigma-Aldrich (Oakville, ON, Canada): beta-mercaptoethanol (Cat# M3148), bovine serum albumin (Cat# A7030, BSA), 2?, 7? ?dichlorofluorescein diacetate (Cat #D6883, DCFDA), hydrogen peroxide (Cat# H1009, H?O?), linoleic acid (Cat# L1012, LA), oleic acid (Cat# O1383, OA), o-phthalaldehyde (Cat# P1378, OPA), potassium phosphate monobasic (Cat# P9791), all-trans retinoic acid (Cat# R2625) and triton-X-100 (Cat# X100). Reagents acquired from EMD Millipore (Mississauga, ON, USA) included chloroform (Cat# 3150), N-Ethylmaleimide (Cat# 34115, NEM), isopropanol (Cat# PX1838-1) and potassium phosphate dibasic (Cat# PX1570-1). Cell culture reagents purchased from Fisher Scientific (Edmonton, AB, Canada) consisted of the following: cell culture water (Cat# SH30529.03), Dulbecco?s modified eagles medium ? high glucose (Cat# SH30081.01, DMEM-HG), Dulbecco?s modified eagles medium ? low glucose (Cat# SH30021.01, DMEM-LG), GlutaGRO (Cat# MT-25-015-CI), penicillin-streptomycin (Cat# MT-30.002.CI), sodium pyruvate (Cat# MT-25-000-CI), trypan blue (Cat# MT-25-900-CI) and trypsin/EDTA (Cat# MT-25-053-CI). Sodium hydroxide (Cat# S320-1, NaOH) was also obtained from Fisher Scientific. Reagents from VWR International (Edmonton, AB, Canada) included hydrogen chloride (Cat# RC37501, HCl), phosphate-buffered saline (Cat# CA95042-488, PBS) and xylene (Cat# 95057-822). Chemicals and reagents obtained from Amresco (Solon, OH, USA) consisted of ribozol (Cat# N580-CA), RNase-free water (Cat#E476) and TBS SHUR/Mount liquid mounting medium (Cat# CA27900-274). 32  Aniline blue solution (Cat# 26367-06), Biebrich scarlet-acid fuchsin (Cat# 26367-04), Bouin?s fixative (Cat# 26367-01), phosphomolybdic-phosphotungstic acid (Cat# 26367-05, Sirius red dye (Cat# 26357-02), Weigert?s iron hematoxylin A (Cat# 26367-02), Weigert?s iron hematoxylin B (Cat# 26367-03) were purchased from Electron Microscopy Sciences (Hatfield, PA, USA). Fetal bovine serum (Cat# 1400-500, FBS) was obtained from Seradigm (Providence, UT, USA). Integrated DNA Technologies (IDT, San Diego, CA, USA) supplied all primers. The reagents acquired from Bio-Rad (Mississauga, ON, Canada) included iScript cDNA synthesis kit (Cat# 170-8891) and Ssofast qPCR reaction mix (Cat# 172-5201). Recombinant human TGF?1 (Cat# 100-21C) was purchased from Peprotech (Rocky Hill, NJ, USA). The cell viability assay reagent, Resazurin (Cat# 30025-1), was obtained from Biotium (Hayward, CA, USA). DNase (Cat# 79254) and proteinase-K (Cat# 19131) were bought from Qiagen (Louisville, KY, USA) and anhydrous ethyl alcohol (MSDS# 1009) was obtained through Commercial Alcohols (Brampton, ON, Canada). 2.2 Equipment All cell lines were maintained in T-75 flasks (Cat# 658175) obtained from Grenier Bio-one (Mississauga, ON, Canada) in an Air-Jacketed CO2 Symphony incubator (Model#5.3A, VWR International). Cell culture experiments were conducted using sterile 48-well tissue culture plates (Cat# 677180, Grenier Bio-one). All cell maintenance and treatments were done in a Labculture Reliant Class II, Type A2 biosafety cabinet (Model#LR2-6S2, ESCO). For passaging and plating procedures, cells were centrifuged using an Accuspin (Model#400, Fisher Scientific) centrifuge and cell counts were done using a hemocytometer. A CKX41 (Olympus) inverted microscope was used for visualization of cells. Following experimental treatments, microplate 33  assay fluorescence or absorbance readings were completed using the GloMax Multi+ Detection System (Model# E9032, Promega, Madison, WI). For filtration of FA stock solutions, 0.2 um filter (Cat# 28145-477) and 1 ml syringes (Cat# CABD309628), obtained from VWR International were used. Messenger RNA (mRNA) from heart tissues was extracted using the Qiagen RNeasy Mini Kit (Cat# 74104) with the use of a Retsch homogenizer (Model# MM 400, Fisher Scientific). mRNA was further quantified using the NanoDrop (Model# 2000c, ThermoScientific) and synthesized into cDNA using a Thermal Cycler (Model# S1000, Bio-Rad). Clear 96-well plates (Cat# 82006-650, VWR International) with PCR sealing film (Cat# 60941-072, VWR International) were used for qPCR reactions in a CFX96 Real-Time System (Bio-Rad). The Heraeus Megafuge (Model# 40R, Thermo Scientific) was used in mRNA extraction and collagen protocols. To observe histological staining, an Omax microscope (#120016) was used in conjunction with the AmScope microscope digital camera (Model# MA500).  2.3 Cell Models Adherent H9c2 (Cat# CRL-1446) rat myoblast cells were purchased from American Type Culture Collection (ATCC) and acquired through Cedarlane (Burlington, ON, Canada). H9c2 is a myoblastic cell line derived from the embryonic BD1X rat tissue and it exhibits both cardiac and skeletal phenotypes [284]. The skeletal phenotype is selected for with reduced serum concentration whereas the cardiac phenotype is selected for with reduced serum in addition to all-trans retinoic acid [285]. H9c2 cells were cultured in DMEM-HG, supplemented with 2 mM sodium pyruvate, 4 mM glutaGRO, 10% FBS and 2% penicillin streptomycin, at 37?C in a 34  humidified atmosphere of 5% CO?. Cells were passaged at 70 ? 80% confluency and media was replaced every 3 ? 4 days. NIH/3T3 (Cat# CRL-1685) adherent mouse-derived cells were used as a fibroblast model and were purchased from ATCC. NIH/3T3 is a cell line derived from NIH Swiss mouse embryonic fibroblast tissue. The cell line was established using the ?3T3? protocol of ?3 day transfer and inoculum of 300 000 cells? [286] and is used as a standard fibroblast model. Cells were maintained in either DMEM-HG, (supplemented with 2 mM sodium pyruvate, 4 mM glutaGRO, 10% FBS and 2% penicillin streptomycin) or DMEM-LG (supplemented with 10% FBS and 2% penicillin streptomycin) at 37?C in a humidified atmosphere of 5% CO?. Media was replaced every 3 ? 4 days and cells were passaged around 70 ? 80% confluency. 2.4 Preparation of Oleic and Linoleic Acid FA stocks (100 mM) were prepared in 70% ethanol (EtOH; made from 100% anhydrous ethyl alcohol and sterile cell culture water). From the 100 mM stocks, 5mM FA stocks were prepared as follows. First, FA-free, low endotoxin BSA (3%) solution was prepared and preheated to 37?C. The FA stock (100 mM) was further mixed with 3% BSA in a 20-fold dilution, resulting in a 5 mM FA: 3% BSA stock solution. Solutions were incubated at 37?C for 3 ? 4 h with shaking. Finally, the solutions were sterile filtered with 1 ml syringes through a 0.2 um syringe filter. All FA stocks were stored at -80?C. 2.5 H9c2 Cardiomyocyte Cells 2.5.1 Differentiation and Plating of H9c2 Myoblasts H9c2 are a myoblastic cell line and are observed to possess both cardiac and skeletal phenotypes [287]. At 70 ? 80% confluency, non-differentiated H9c2 cells were harvested from T-75 flasks. The cells were washed with 4 ml PBS, and then detached using 3 ml 0.25% Trypsin-35  EDTA for 3 min at 37?C and 5% CO2. Trypsin was then deactivated with 9 ml fresh supplemented DMEM and further centrifuged at 1400 rpm for 2 min at room temperature (RT). The supernatant was discarded and the cell pellet was re-suspended in fresh supplemented DMEM media. Cells were counted using a hemocytometer and viable from non-viable cells determined using 0.04% trypan blue stain. Prior to differentiation, 20,000 cells per well were plated in sterile 48-well plates. Cells were maintained in DMEM-HG until they reached ~80% confluency. In order to use these cells as a cardiomyocyte model, they were first differentiated into a cardiac phenotype. Differentiation was induced at 80% confluency because once cells reach confluency they began to differentiate into both skeletal and cardiac phenotypes. H9c2 myoblasts were differentiated into a cardiomyocyte phenotype by reducing FBS from 10 to 1% with daily addition of 0.1 uM all-trans retinoic acid (via media change) for four days as previously described [287, 288]. 2.5.2 Treatment of Cardiomyocytes Following differentiation, cardiomyocytes were treated with the following: (1) OA or LA, (2) H2O2, and (3) combination of OA or LA with H2O2. Initially, cardiomyocytes were incubated for two time points, 6 and 12 h, with either 0.1 or 0.25 mM OA or LA or control, untreated cells. Furthermore, cardiomyocytes were treated with H2O2, concentrations ranging from 100 ? 1000 uM, for 6 and 12 h and were compared to control, untreated cells. Lastly, cardiomyocytes were incubated with a combination of 0.1 or 0.25 mM OA or LA with 300 and 500 uM H2O2 for 6 and 12 h. OA or LA, or H2O2 were added to the supplemented media of each well. All treatments and controls were done in duplicate for each experiment.  36  2.6 NIH/3T3 Fibroblasts 2.6.1 Plating of Fibroblasts At 70 ? 80% confluency, fibroblasts were harvested from T-75 flasks. Cells were washed with 4 ml PBS and then detached using 3 ml 0.25% trypsin-EDTA for 3 min at 37?C and 5% CO2. Cells were further suspended in 9 ml fresh supplemented media in order to deactivate the trypsin. Cells were then centrifuged at 1400 rpm for 2 min at RT, the supernatant was discarded and the cell pellet was re-suspended in fresh supplemented DMEM. Cell counts were done in a hemocytometer and viable and non-viable cells determined with 0.04% trypan blue stain. For experiments, 20,000 cells per well were plated in sterile 48-well plates. Cells were allowed to adhere overnight before treatment. 2.6.2 Treatment of Fibroblasts Fibroblasts were treated with the following: (1) OA or LA, (2) combination of OA or LA with TGF?1 and (3) combination of OA or LA with H2O2. First, fibroblasts were incubated for several time points: 2, 4 and 6h, 0 -24 h and 24 h with either 0.1 or 0.25 mM OA or LA or control, untreated cells. Second, cells were treated with 0.1 or 0.25 mM OA or LA or control (without FA) and concurrently stimulated with 10 ng/ml TGF?1. Lastly, fibroblasts were incubated with a combination of 0.1 or 0.25 mM OA or LA with 100 or 200 uM H2O2 for 24 h. OA or LA, H2O2 and TGF?1 were added to the media in each well. Following treatment, various parameters were measured. More specifically, measurements of cell viability, GSH levels, ROS and collagen (soluble and deposited) were assessed using the Resazurin, OPA, DCFDA and Sirius Red assays respectively. All treatments and controls were completed in duplicate. 37  2.7 In Vitro Assays 2.7.1 Cell Viability Assay Cell viability was assessed with the resazurin (Alamar Blue) assay. The resazurin (7-hydroxy-10-oxidophenoxazin-10-ium-3-one) dye is non-toxic to living cells [289]. Other assays that are used for the measurement of cell viability use tetrazolium salts such as (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), which are toxic to the cells [290]. The resazurin dye is a non-fluorescent permeable blue dye, which is converted to the highly fluorescent pink dye, resorufin, due to the chemical reductions occurring within living cells. Cell growth maintains a ?reduced? environment and the lack of cell growth maintains an ?oxidized? environment [289]. NADPH and NADH dehydrogenases [291] and diaphorases [292] are enzymes that are responsible for the conversion of resazurin to resorufin. Differentiated cardiomyocytes and fibroblasts were treated as previously described in Sections 2.5.2 and 2.6.2 respectively. Media was first removed from the adherent cells and discarded. The working solution containing the resazurin dye in medium in a 10:1 dilution was transferred into each well. The adherent cells in working solution were allowed to incubate for 1 h at 37?C and 5% CO?. Fluorescence intensity was measured at 525 nm Ex. and 580-640 nm Em. using the GloMax Multi+ Detection System (Promega). Treated cells were compared to control untreated cells. 2.7.2 Reactive Oxygen Species Assay The cell permeant dye, 2?.7?-dichlorofluorescein diacetate (DCFDA), is a dye that measures non-specific levels of reactive oxygen species such as hydroxyls, peroxyls and other species [293]. DCFDA diffuses into the cell where it is deacetylated by cellular esterases to form 38  the non-fluorescent compound, 2?, 7?-dichlorodihydrofluorescin, which is then easily oxidized by reactive oxygen species into the highly fluorescent marker, 2?, 7?-dichlorofluorescein [294]. Fibroblasts were treated as previously described in Section 2.6.2. Media was first removed from the adherent cells in each well and cells were washed with 200 ul PBS. A working solution consisting of 10 uM DCFDA in fresh supplemented DMEM media was added to each well. The reaction was allowed to incubate for 0-24 h at 37?C and 5% CO?. Fluorescence intensity was measured at 490 nm Ex. and 510-570 nm Em. using the GloMax Multi+ Detection System. Treated cells were compared to control untreated cells. 2.7.3 Glutathione Assay O-phthalaldehyde (OPA) is a non-fluorescent dye that interacts with amines in the presences of thiols such as GSH to form a fluorescent isoindole [295]. The following protocol for GSH measurement was adapted for a microplate assay from Senft et al. [296]. Fibroblasts were treated as previously described in Section 2.6.2. First, the media was removed from adherent cells and all cells were washed with 200 ul PBS twice. Cells were lysed using 120 ul 1x LDH lysis buffer (30% triton-X-100; 50 mM potassium phosphate buffer (KPi; pH 7.5) through incubation for 15 min at 37?C in 5% CO?. Into a new 48-well plate, 50 ul of each lysate was added to two separate wells labeled A and B. All samples were made up to 150 ul using 1 M KPi. To column A, 20 ul NEM and 250 ul 1 M KPi were added and to column B, 250 ul 1 M KPi was added. Lysates were then incubated for 5 min at RT. Following incubation, 500 ul 0.1 M KPi and 150 ul OPA was added to all wells and incubated for 30 min at RT. Fluorescence was measured at 365 nm Ex. and 410-460 nm Em. using the GloMax Multi+ Detection System. Fluorescent values from well A were subtracted from well B for each treatment. Following this calculation, treated cells were compared to control untreated cells. 39  2.7.4 Standardization of the Collagen Assay Sirius red dye stains extracellular components such as collagen type I and III in a concentration-dependent manner whereas other extracellular components such as laminin, collagen type IV and fibronectin exhibit only weak staining. A protocol was adapted from Tullberg-Reinert and Jundt [297] and the Sircol? Assay for the measurement of deposited and soluble collagen. In order to standardize this assay, levels of deposited collagen were measured following treatment of OA or LA at 12h and 1, 2, 3, 4 and 7 days in fibroblasts grown in high glucose. In all conditions, data demonstrated no change or significantly decreased deposited collagen from control, untreated cells. In the attempt to increase levels of deposited collagen, fibroblasts were treated with OA or LA and concurrently stimulated with TGF?1, a factor known to increase deposited collagen [298]. Once again, following stimulation, levels of deposited collagen showed either no change or a decrease from control untreated cells. In another attempt to obtain positive results for deposited collagen levels (i.e. increased levels from control untreated cells), fibroblasts were grown in low glucose conditions. They were first treated with OA and LA alone, data from which demonstrated no change in deposited collagen from control but the decrease in deposited collagen (seen previously in high glucose conditions) was not observed. Fibroblasts were then treated with OA and LA with concurrent TGF?1-stimulation. After only 1 day of treatment, there was an increased level of deposited collagen. From these results, we concluded that the collagen assay is a better measurement tool in fibroblasts grown in low glucose conditions.   40  2.7.4.1 Soluble Collagen Assay Fibroblasts were treated as previously described in Section 2.6.2. From the wells of treated cells, 50 ul of media was removed and combined with 50 ul sterile cell culture water in a 1.5 ml tube. This dilution step was included in order to decrease the media serum concentration from 10 to 5%, as it is known that serum can interfere with the Sirius red dye [299]. To each tube, 250 ul Sirius red dye was added and was mixed for 30 min at RT with gentle shaking. Samples were then centrifuged at 10,000 xg for 5 min to pellet the collagen and the supernatant was carefully removed. To the pellet, 1 ml HCl was gently added to wash off any excess/unbound dye. Samples were centrifuged at 11,000 xg for 5 min and the HCl and excess dye was discarded. The excess liquid remaining on the side of the tubes was removed using a Kimwipe. To dissolve the stain, 250 ul 0.1N NaOH was added and tubes were vortexed vigorously. The dye was transferred into a new 48-well plate, from which the absorbance intensity was read at 560 nm using the GloMax Multi+ Detection System. Treated cells were compared to control untreated cells. 2.7.4.2 Deposited Collagen Assay Fibroblasts were treated as previously described in Section 2.6.2. Treated cells were washed twice with 200 ul PBS and fixed with 100 ul Bouin?s fixative for 1 h at RT. Bouin?s fixative was removed and the plate was washed for 5 ? 10 min under running water until no yellow stain remained. Water was removed from the plate using a pipette and further air-dried. To each well, 200 ul Sirius red dye was added and the plate was incubated for 1 h at RT under mild shaking. Any excess dye was removed from the wells and cells were then washed three times with 0.01 N HCl (or until all excess dye was removed). The Sirius red stain was dissolved in 200 ul 0.1 N NaOH under gentle shaking for 30 min at RT. All dye was then transferred into a 41  new 48-well plate to avoid any background readings. Absorbance intensity was measured at 560 nm using the GloMax Multi+ Detection System. Treated cells were compared to control untreated cells. 2.8 Treatment of Mice with High Fat Diets and Isoproterenol All protocols were completed in accordance with the University of British Columbia Animal Care Committee Guidelines. Male C57BL/6 mice were purchased from Jackson Laboratory (Bar Harbor, Maine, USA). Mice were housed in a temperature-controlled room (26?C) on a 12-h light/dark cycle with free access to food and water. Four-week-old male C57BL/6 mice were randomly assigned into three groups. Each group had 5 ? 8 mice. For four weeks, two groups of mice were fed with two high fat diets, OO (OO; high OA) or corn oil (CO; high LA), and one group was fed with a low fat diet, normal chow (NC, low LA). High fat diets consisted of 20% w/w of their respective oil and NC consisted of 5% CO. OO and CO diets were isocaloric and isonitrogenous; both were prepared through the addition of 200 grams of oil to 800 grams of basal mix, obtained from Harlan Teklad, USA (TD.88232). The final diet composition consisted of 13.81 kJ/g of energy and was composed of protein at 21.2% w/w (19% by energy), carbohydrates at 44.7% w/w (40.1% energy) and fat at 20.2% w/w (40.8% by energy). For the detailed compositions of Harlan Teklad Basal Mix, see Appendix A. Diets were administered to mice for 4 weeks where food and water were freely accessible. Following four weeks of feeding, a subset of each diet group was randomly assigned, and injected intraperitoneally with 120 mg/kg isoproterenol (ISO) for two consecutive days. The first subset of mice, termed the acute group, were sacrificed twenty-four hours following the last injection. Another subset of mice was maintained 21 days on their respective diets following the last ISO injection. Following the three weeks, the second subset termed the chronic group was 42  sacrificed. All tissue samples were collected immediately following sacrifice of the mice and stored at -80?C until processing. 2.9 mRNA Isolation and Complementary DNA Synthesis 2.9.1 mRNA Isolation from Fibroblasts mRNA from fibroblasts was isolated using the Trizol method. Procedures were similar to the manufacturer?s protocol with Ribozol used instead of Trizol. 20,000 cells were seeded in a 48-well plate and allowed to adhere overnight. Fibroblasts were treated with OA or LA at 0.1 mM and concurrently stimulated with 10 ng/ml TGF?1 at 37?C at 5% CO2 for 24 h. Following treatment, the media was removed from adherent fibroblasts and cells were washed with 200 ul PBS twice. For homogenization of cells, 100 ul of Ribozol was added to each well, incubated for 5 min at RT with gentle shaking and further incubated at -80?C for 5 min or stored until extraction of mRNA. mRNA from each sample was isolated from the Ribozol/cell solution. All samples were incubated for 5 min at RT and further transferred from the 48-well plate into 1.5 ml tubes. To each tube, 0.2 ml of chloroform (per 1 ml Ribozol used for homogenization) was added. Tubes were then shaken vigorously for 15 s and incubated for 2 ? 3 min at RT. Samples were spun at 12,000 xg for 15 min at 4?C. After centrifugation, the samples separated into three phases: lower phenol-chloroform (red), an interphase, and an upper colorless aqueous phase. mRNA was found within the aqueous phase only. The aqueous phase was carefully removed and placed into a new 1.5 ml tube. To each tube, 0.5 ml of 100% isopropanol (per 1ml ribozol used for homogenization) was added to the aqueous phase and incubated for 20 min at -80?C. Samples were centrifuged at 12,000 xg for 10 min at 4?C and the supernatant was then removed from the tube. The mRNA pellet was washed with 1 ml of 75% EtOH (per 1 ml ribozol used in 43  homogenization). The samples were briefly vortexed, centrifuged at 7500 xg for 5 min at 4?C and the wash was discarded. The pellet was air dried for 5 ? 10 min and further suspended in 20 ? 50 ul RNase-free water by pipetting up and down. The mRNA was incubated at 60?C for 10 ? 15 min and then analyzed for quantity and purity. 2.9.2 mRNA Extraction from Heart Tissue Samples All gene expression was conducted on the acute group of mice. mRNA was extracted from acute heart tissue samples using the RNeasy Mini Kit (Qiagen). Approximately 60 mg of heart tissue was added to 300 ul buffer RLT, 3ul beta-mercaptoethanol and a metal bead. Samples were homogenized at 30 Hz for 2 min then placed on ice, and repeated. The homogenized sample was added with 590 ul RNase free-water and 10 ul proteinase-k and further incubated at 55?C for 10 min. During incubation, preparation of 10 ul DNase and 70 ul buffer RD (per sample) was done and mixed with inversion. The samples were centrifuged at 11,000 xg for 3 min. The supernatant was then transferred to a new tube and 450 ul absolute EtOH was added, mixed by inversion. To a new spin column, 700 ul of sample was added, and spun at 11,000 xg for 15 s with all flow though being discarded. Directly to the membrane of the spin column 80 ul DNase was added and incubated for 15 min at RT. To the column 350 ul buffer RW1 was added, spun at 11,000 xg for 15 s with all flow through being discarded. To the column, 500 ul of buffer RPE was added and spun at 11,000 xg for 15 s with the flow through being discarded. An additional 500 ul buffer RPE was added to the column and spun at 11,000 xg for 2 min. The spin column was placed onto a new collection tube, spun at 11,000 xg for 1 min. The spin column was then placed on a new 1.5 ml collection tube. mRNA was eluted first with 50 ul RNase free water, let sit 2 ? 3 min at RT, and spun at 11,000 xg for 1 min. Another 30 ul RNase-free water was added, let sit for 2 ? 3 min at RT, and spun at 11,000 xg for 1 min. mRNA was further analyzed for quantity and purity. 44  2.9.3 Complementary DNA Synthesis mRNA was converted to complementary DNA (cDNA) using the iScript cDNA Synthesis Kit. Approximately 500 ng of the mRNA sample was converted to cDNA in the following mixture: 500 ng mRNA sample, 2 ul iScript reaction mix, 0.5 ul iScript reverse transcriptase enzyme and nuclease-free water to a total volume of 10 ul in a 0.2 ml microcentrifuge tube. The solution was mixed with inversion and briefly centrifuged. cDNA synthesis reactions were performed for 5 min at 25?C, 30 min at 42?C, 5 min at 85?C and held for 4?C. Samples were diluted in a 1:10 ratio with nuclease-free water. All cDNA samples were stored at -20?C. 2.10 Primer Design and the Quantitative Polymerase Chain Reaction 2.10.1 Primer Design Primers were obtained from Integrated DNA Technologies (IDT, Coralville, IA, USA). Primer design was accomplished using AutoPrime online software. Sequence specificity was checked through the National Centre for Biotechnology Information (NCBI) using the primer basic local alignment search tool (BLAST). Primers were checked for optimal annealing temperatures, primer dimer formation and efficiency following Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines [300]. 2.10.2 Quantitative Polymerase Chain Reaction All quantitative polymerase chain reactions (qPCR) were performed in duplicate with 3 ? 5 (in vitro) or 5 ? 8 (in vivo) independent biological samples in clear 96-well plates covered with clear PCR sealing film. Each reaction consisted of a total volume of 10 ul containing: 5 ul Ssofast Evagreen Supermix, 04 ul of each primer (10 uM) (for specific primers, see Table 1), 3.6 ul nuclease-free water and 1 ul of cDNA template. No control template (NTC), consisting of all 45  qPCR components with the cDNA template replaced with water, was used as a negative control to ensure that DNA contamination was not present. The qPCR protocol was run on the CFX96 Real-Time System (Bio-Rad) and consisted of the following steps: initial denaturation at 95?C for 30 s, followed by 50 cycles of denaturation at 95?C for 5 s and annealing at 58?C or 60?C for 5 s, followed by a dissociation step of 95?C for 10 s. Melt curve analysis was completed from 65?C to 95?C at 0.5?C/cycle (0.5?C/s), in order to verify amplification specificity. All primers had an annealing temperature of 58?C, except for MMP9, which had an annealing temperature of 60?C. The reference gene used was 18S ribosomal RNA. 2.10.3 Primer Efficiencies Primer efficiencies were determined using LinRegPCR (Version: 2012.1) and found to be within accepted values [300]. Gene expression of each treatment was normalized to 18S ribosomal RNA (reference gene) and further normalized to either unstimulated control (in vitro gene studies) or NC (in vivo gene studies), which were assigned a value of 1. The relative gene expression values were calculated using CFX Manager 3.0 (Bio-Rad). LinRegPCR is a program that is uses a linear regression model for the analysis of qPCR data. This program monitors PCR reactions that use SYBR green (or other fluorescent) dyes and measures the fluorescence measured per cycle of each sample and the baseline fluorescence. A window-of-linearity for each sample is determined and the intercept and slope further determined using linear regression. Individual PCR efficiencies for each sample are calculated from the slope of the line of linear regression. Selection of data points within each window-of-linearity requires at least four and up to six points in order to prevent unambiguous selection [301, 302]. The software calculates the average efficiency for amplicon group. More specifically, the Bio-Rad iCycler data file format was used in LinRegPCR in order to determine baselines from 46  imported data files (Quantification Amplification Results from the CFX Manager). Figure 2 illustrates the LinRegPCR program. 47  Table 1. Primer sequences used for quantitative polymerase chain reaction   Primer SequencesTarget Sequence Forward Sequence Accession Numbers18S ribosomal RNA  (Rn18s) F CGG CTA CCA CAT CCA AGG AA NR_003278.3R GCT GGA ATT ACC GCG GCTCollagen, type 1, alpha 1 (Col1a1) F ACC TGT GTG TTC CCT ACT CA NM_007742.3R GAC TGT TGC CTT CGC CTC TGCollagen, type III, alpha 1 (Col3a1) F AAT GGT GGT TTT CAG TTC AGC NM_009930.2R TGG GGT TTC AGA GAG TTT GGCMatrix metalloproteinase 2 (Mmp2) F CCT GAA TAC TTT CTA TGG CTG C NM_008610.2?R GTA TGT AGT GGA GCA CCA GAG CMatrix metalloproteinase 9 (Mmp9) F GCA ACG GAG ACG GCA AAC C NM_013599.3R GAC GAA GGG GAA GAC GCATissue inhibitor of metalloproteinase 1 (Timp1) F AAC TCG GAC CTG GTC ATA AG NM_011593.2; NM_001044384.1R TAA GGT GGT CTC GTT GAT TTCTissue inhibitor of metalloproteinase 2 (Timp2) F CAA AGC AGT GAG CGA GAA GG NM_011594.3R ACC CAG TCC ATC CAG AGG GATran formi g growth factor, beta 3 (Tgfb3) F CAGCCTACATAGGTGGCAAGAAT NM_009368.3R ACCCAAGTTGGACTCTCTCCTCAATransforming growth factor, beta receptor 1 (Tgfbr1) F GCA CCA TCT TCA AAA ACA GGG G NM_009370.2R GCC AAA CTT CTC CAA ACC GAC CTransforming growth factor, beta receptor 2 (Tgfbr2) F CTC AAC ACA CCA AAG TCC TC NM_009367.3R ATC AAA ACT CCC TCC CTC C48    Figure 2. Determination of primer efficiencies using LinRegPCR. (A) Data imported into LinRegPCR, (B) baseline calculation and (C) primer efficiency and information saved to excel.   49  2.11 Masson?s Trichrome Stain Masson?s trichrome is a differential stain used to differentiate collagen fibers in tissue from muscle and nuclei [303]. In this set of experiments, the chronic group of mice was used to analyze the collagen deposition qualitatively. Paraffin-fixed heart tissues were deparaffinized and rehydrated with the following steps: xylene for 5 min (twice), 100% EtOH for 3 min, 90% EtOH for 3min, 80% EtOH for 3 min, distilled water for 3 min and PBS for 3 min (shaking). Tissues were washed in distilled water for 1 min. To improve the staining quality, tissues were further fixed in Bouin?s solution at 56?C for 1 h. After fixation, slides were rinsed for 10 minutes to remove excess Bouin?s solution. Tissues were stained in Weigert?s iron hematoxylin (a solution of equal parts Weigert?s Iron Hematoxylin A and B) for 10 min and then rinsed with distilled water for 10 min. Tissues were then stained with Biebrich scarlet-acid fuchsin solution for 15 min and then rinsed briefly in distilled water. Tissues were differentiated with phosphomolybdic-phosphotungstic acid solution for 15 min to remove the red color from the collagen. Tissues were directly transferred into aniline blue solution, stained for 10 min, and then rinsed briefly in distilled water. Tissues were further differentiated using 1% acetic acid for 5 min followed by a wash in distilled water for 5 min. Tissues were dehydrated quickly through 90% EtOH for 3 min, 100% EtOH for 3 min and cleared in xylene for 5 min. All tissues were mounted with SHUR/Mount Liquid Mounting media. Masson?s trichrome stain was observed using an Omax bright field microscope and pictures were taken using the AmScope camera at 400X magnification. 2.12 Statistical Analysis GraphPad Prism (Version 5) software was used to conduct statistical analyses. Analysis of variance (ANOVA) was utilized to determine significance within data. Two-way ANOVA with Bonferroni post-tests and one-way ANOVA with Tukey post-tests were used to find 50  significance and determine differences between treatment groups. ANOVA was performed for all experiments and consisted of the dependent variable, the experimental data, and the independent variable, the different concentrations of compounds or times. The random factor was blocked by data collected on separate days and/or independent biological replicates. All data were presented as mean ? standard error of the mean (SEM) with a probability (p) value less than 0.05 considered statistically significant, indicated on graphs as ap<0.05 or bp<0.05. All in vitro cell assays are presented as three separate experiments. In vitro qPCR data had 3 ? 5 individual replicates and the in vivo qPCR data had 5 ? 8 individual biological replicates.                    51  3.0 Results 3.1 Cardiomyocyte Viability 3.1.1 Effects of Fatty Acids with Hydrogen Peroxide on Cardiomyocyte Viability 3.1.1.1 Impact of Oleic or Linoleic Acid for 6 and 12 h on Cardiomyocyte Viability OA and LA were tested for viability using the Resazurin assay at 0.1 and 0.25 mM concentrations for 6 and 12 h and compared to control without FA. After either 6 (Fig. 3A) or 12 h (Fig. 3B). Although following 6 h, OA demonstrated a loss of viability, this loss was recovered by 12 h. However, 0.25mM LA decreased cardiomyocyte viability at 12 h (Fig. 3B).  OALA-2000-1000010002000ControlbCell ViabilityRFU - Relative to ControlOALA-6000-4000-200002000ControlbA B6h 12h0.1 mM0.25 mM  Figure 3. Effect of oleic or linoleic acid for 6 and 12 h on viability of cardiomyocytes. Cardiomyocytes were treated with 0.1 or 0.25 mM OA or LA or control (without FA) for 6 and 12 h. Following 6 (A) and 12 h (B), cardiomyocytes were evaluated for viability using the Resazurin viability assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test. bp<0.05, significantly different between groups treated with different concentrations of the same FA.   52  3.1.1.2 Effect of Hydrogen Peroxide for 6 and 12 h on Cardiomyocyte Viability  Cardiomyocytes were treated with H2O2 in order to simulate oxidative insult in vitro and to study its effects on viability. H2O2 was tested at concentrations ranging from 100 to 1000 uM for 6 and 12 h, and all results were compared to untreated control cells. Following 6 (Fig. 4A) and 12 h (Fig. 4B), cardiomyocyte viability was assessed using the Resazurin viability assay. Cardiomyocyte viability was decreased with 500 uM H2O2 at 12 h (Fig. 4B) and 1000 uM H2O2 at both 6 (Fig. 4A) and 12 h (Fig. 4B). Lower concentrations of H2O2, 100, 200 and 300 uM, did not cause significant loss of cardiomyocyte viability following either 6 or 12 h.  1002003005001000-3 0-2000-1000010002000aControlCell ViabilityRFU - Relative to Control1002003005001000-4000-3000-2000-10000aaControlH2O2Dose (uM)A B6h 12h  Figure 4. Effect of hydrogen peroxide after 6 and 12 h on cardiomyocyte viability. Differentiated cardiomyocytes were treated with H2O2, concentrations ranging from 100 to 1000 uM, and 0 uM as a control for 6 and 12 h. Following 6 (A) and 12 h (B), Cardiomyocytes were tested for viability using the Resazurin assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of H2O2 were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control.      53  3.1.2 Effects of Fatty Acids in Combination with Hydrogen Peroxide on Cardiomyocyte Viability 3.1.2.1 Effect of Hydrogen Peroxide plus Oleic or Linoleic Acid for 6 and 12 h on Cardiomyocyte Viability Most dietary FAs induce oxidative stress in vivo given at higher concentrations. Cardiomyocytes were co-incubated with H2O2 plus either OA or LA in order to investigate their combined effects on viability. An effect of combined treatment of 300 or 500 uM H2O2 plus 0.1 or 0.25 mM OA or LA was investigated for 6 (Fig. 5A) and 12 h (Fig. 5B) on cardiomyocyte viability using resazurin assay. Cardiomyocyte viability was increased with 300 uM H2O2 for 6 h with both 0.1 mM OA and LA. However, with 0.25 mM OA or LA and 300 uM H2O2, cardiomyocytes showed a trend of decreased viability at 12 h. As treatment increased to 12 h, cardiomyocytes demonstrated decreased viability with 0.1 mM LA but not with OA. With a higher concentration of H2O2 (500 uM), 0.25 mM OA caused greater loss in viability compared to 0.1 mM OA at 6 (Fig. 5C) and 12h (Fig. 5D). In contrast, both 0.1 and 0.25 mM LA, showed similar loss in viability with 500 uM H2O2 at 6 (Fig. 5C) and 12 h (Fig. 5D).        54  OALA-4000-2000020004000Controla,b a,bCell ViabilityRFU - Relative to ControlOALA-4000-3000-2000-10000aControlA B6h 12h0.1 mM (+300 uM H2O2)0.25 mM (+300 uM H2O2) OALA-6000-4000-20000aaa,baControlCell ViabilityRFU - Relative to ControlOALA-8000-6000-4000-20000aa,baControlC D6h 12h0.1 mM (+500 uM H2O2)0.25 mM (+500 uM H2O2)  Figure 5. Effect of the combined treatment of hydrogen peroxide plus oleic or linoleic acid for 6 and 12 h on cardiomyocyte viability. Cardiomyocytes were treated with either 300 or 500 uM H2O2 plus 0.1 or 0.25 mM OA or LA for 6 and 12 h. Following 6 (A) and 12 h (B) with 300 uM H2O2 or 6 (C) and 12 h (D) with 500 uM H2O2, cardiomyocytes were evaluated for viability using the Resazurin assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of H2O2 with OA or LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control.  3.2 Fibroblast Viability and Cellular Oxidative Stress 3.2.1 Effects of Fatty Acids on Fibroblast Viability and Reactive Oxygen Species Production 3.2.1.1 Effects of Oleic or Linoleic Acid for 6 h on Fibroblast Viability OA and LA were tested for their effects on fibroblast viability. OA and LA were tested at 0.1 and 0.25 mM concentrations for 6 h. The treatment results were compared to the control 55  group, without FA. Following 6 h (Fig. 6), viability of fibroblasts was measured using the Resazurin viability assay. At 0.25 mM LA, there was decreased fibroblast viability following 6 h. However, there was no significant decrease in viability with 0.1 or 0.25 mM OA or 0.1 mM LA after 6 h.  OALA-1500-1000-50005000.1 mM0.25 mMControla6hCell ViabilityRFU - Relative to Control  Figure 6. Effect of linoleic or oleic acid for 6 h on fibroblast viability. Fibroblasts were treated with 0.1 or 0.25 mM OA or LA or control, treated without FA, for 6 h. Following 6 h, the fibroblasts were evaluated for viability using the Resazurin assay. Values are mean ? SEM. Data from 4 independent experiments are presented. The concentration-dependent effects of OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control.  3.2.1.2 Effects of Oleic or Linoleic Acid on Reactive Oxygen Species Production in Fibroblasts over 24 h OA and LA were investigated for their effects on reactive oxygen species production in fibroblasts. Fibroblasts were preloaded with the DCFDA dye in their supplemented growth media and a fluorescence reading was taken for each well, which was labeled as time 0. Following the first reading (?time 0?), fibroblasts were treated with 0.1 mM OA or LA for 1 ? 8 h and 24 h. Replicates of each treatment group were compared to their respective ?time 0? 56  control. Fibroblasts were treated with OA or LA and compared to control, without FA, for 0 ? 8 h, and 24h. For every hour up until 8 h and at 24 h (Fig. 7), reactive oxygen species levels were detected using the DCFDA assay. At 0.1 mM OA from 3 ? 8 h, there were increased levels of reactive oxygen species produced compared to control. At 24 h, 0.1 mM OA decreased levels of reactive oxygen species levels. However, 0.1 mM LA from 5 ? 8 h, and at 24 h, decreased levels of reactive oxygen species.    Figure 7. Effects of oleic or linoleic acid over 24 h on reactive oxygen species production in fibroblasts. Fibroblasts were preloaded with DCFDA in supplemented media (?time 0?) and further treated with 0.1 mM OA or LA, or control, treated without FA, for 0 ? 8h and 24h. Every hour, fibroblasts were assessed for levels of reactive oxygen species using the DCFDA assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The time-dependent effects of OA and LA were assessed using ANOVA, followed by Bonferroni post-hoc test ap<0.05, significantly different from control, treated without FA.  0 1 2 3 4 5 6 7 8240500010000ControlOALAaaaaaaaaaaaaHoursROS ProductionRFU - Relative to 0h57  3.2.2 Effects of Fatty Acids plus Hydrogen Peroxide on Fibroblast Viability and Glutathione Levels 3.2.2.1 Effect of Oleic or Linoleic Acid Combined with Hydrogen Peroxide after 24 h on Fibroblast Viability The treatment of OA or LA with/without H2O2 was investigated on fibroblast viability. Fibroblasts were treated with 100 and 200 uM H2O2 or treated with either 0.1 mM OA or LA for 24 h. In addition, cells were treated with a combination of 100 or 200 uM H2O2 and 0.1 mM OA or LA for 24 h. All results were compared to untreated control group. Combination treatments were compared to their respective FA only control. Following 24 h (Fig. 8), fibroblast viability was assessed using the Resazurin viability assay. After 24 h, 100 and 200 uM H2O2 with 0.1 mM OA and LA decreased viability. Combination of 0.1 mM OA or LA with 200 uM H2O2 demonstrated a further decrease in viability compared to OA or LA treatment alone. However, the treatment of 0.1 mM OA or LA with 100 uM H2O2 did not show any difference from OA or LA treatment alone.   58  ControlOALA-3000-2000-100000 uM H2O2100 uM H2O2200 uM H2O2aabbaa24hCell ViabilityRFU - Relative to Control/0uM  Figure 8. Effects of combined treatment of hydrogen peroxide plus oleic or linoleic acid for 24 h on fibroblast viability. Fibroblasts were treated with 100 and 200 uM H2O2 or 0.1 mM OA or LA, or in combination of 100 or 200 uM H2O2 and 0.1 mM OA or LA for 24 h. All treatment groups were compared to control, treated without FA and 0 uM H2O2, and their respective FA treated control. Following 24 h, the viability of fibroblasts was quantified for viability using the Resazurin assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of H2O2, OA and LA, and combination of H2O2 with OA or LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control, bp<0.05, significantly different between groups treated with different concentrations of H2O2 and the same FA.  3.2.2.2 Effects of Oleic or Linoleic Acid and Hydrogen Peroxide for 24 h on Glutathione Levels in Fibroblasts GSH levels were measured following treatment of OA or LA with/without H2O2 in fibroblasts. Initially, fibroblasts were treated with either 100 or 200 uM H2O2 or 0.1 mM OA or LA for 24 h. Furthermore, cells were treated with a combination of 100 or 200 uM H2O2 and 0.1 mM OA or LA for 24 h. All results were compared to untreated control group and combined treatments were compared to their FA only control. Following 24 h (Fig. 9), GSH levels of fibroblasts were measured using the o-phthalaldehyde assay. 59  With 200 uM H2O2 only, decreased GSH levels were observed. With 0.1 mM OA or LA, there was no significant change in GSH levels. Treatment of 0.1 mM OA or LA with 100 and 200 uM H2O2 did not demonstrate a decrease in GSH levels.  ControlOALA-15-10-500 uM H2O2100 uM H2O2200 uM H2O2a24hGSH LevelsRFU - Relative to Control/0uM Figure 9. Effect of the combined incubation with hydrogen peroxide plus oleic or linoleic acid following 24 h on the levels of glutathione in fibroblasts. Fibroblasts were treated with 100 or 200 uM H2O2 or 0.1 mM OA or LA for 24 h. Cells were further treated with a combination of 100 or 200 uM H2O2 and 0.1 mM OA or LA for 24 h. All treatment groups were compared to control, treated without FA and 0 uM H2O2, and their respective FA treated control. Following 24 h, GSH levels of fibroblasts were measured using the o-phthalaldehyde assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of H2O2, OA and LA, and combination of H2O2 with OA or LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control.    60  3.3 Collagen Production of Fibroblasts 3.3.1 Effects of Fatty Acids on Collagen Production 3.3.1.1 Outcome of the Combination Treatment of Oleic or Linoleic Acid with Hydrogen Peroxide on Soluble Collagen Production by Fibroblasts Soluble collagen produced by fibroblasts was assessed following treatment with OA or LA with/without H2O2. Fibroblasts were treated with 100 and 200 uM H2O2, or treated with 0.1 mM OA or LA in combination with 100 or 200 uM H2O2 for 24 h. All results were compared to control group, treated without FA and 0 uM H2O2, and co-incubated treatments were also compared to their FA treatment control. Following 24 h (Fig. 10), soluble collagen production by fibroblasts was measured using the Sirius red assay. After 24 h, incubation with 100 and 200 uM H2O2 led to lower soluble collagen in control cells. Following combination of 100 or 200 uM H2O2 with 0.1 mM OA and 200 uM H2O2 with 0.1 mM LA, there was increased levels of soluble collagen present.    61  ControlOALA-0.10-0.050.000.050.100.150 uM H2O2100 uM H2O2200 uM H2O2a aa24hSoluble CollagenRAU - Relative to Control/0uM  Figure 10. Effects of combined treatment of hydrogen peroxide with oleic or linoleic acid following 24 h on soluble collagen production by fibroblasts. Fibroblasts were treated with 100 or 200 uM H2O2 or 0.1 mM OA or LA. In addition, cells were co-incubated with 100 or 200 uM H2O2 and 0.1 mM OA or LA for 24 h. All treatment groups were compared to control, treated with no FA and 0 uM H2O2, and their respective FA treated control. Following 24 h, soluble collagen levels produced by fibroblasts were detected using the Sirius red assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of H2O2, OA and LA, and combination of H2O2 with OA or LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control.  3.3.1.2 Effects of a 24 h Combination of Oleic or Linoleic Acid with Hydrogen Peroxide on Deposited Collagen Production by Fibroblasts Deposited collagen in fibroblasts was evaluated after the treatment of OA or LA with H2O2. Fibroblasts were treated with 100 and 200 uM H2O2 or treated with 0.1 mM OA or LA for 24 h. Furthermore, cells were treated with a combination of 100 or 200 uM H2O2 for 24 h. All results were compared to untreated control group and combined treatments were compared to their FA only control. Following 24 h (Fig. 11), deposited collagen production by fibroblasts was measured using the Sirius red assay. 62  After 24 h, 100 and 200 uM H2O2 decreased deposited collagen levels. At 0.1 mM OA and LA, there was no change in deposited collagen present. However, combination of 100 or 200 uM H2O2 with 0.1 mM OA or LA decreased the deposited collagen produced. ControlOALA-0.020-0.015-0.010-0.0050.0000.0050 uM H2O2100 uM H2O2200 uM H2O2aaa,ba,ba,ba,b24hDeposited CollagenRAU - Relative to Control/0uM  Figure 11. Effects of combined treatment of hydrogen peroxide with oleic or linoleic acid following 24 h on deposited collagen by fibroblasts. Fibroblasts were treated with 100 and 200 uM H2O2 and 0.1 mM OA or LA, or co-incubated with 100 or 200 uM H2O2 with 0.1 mM OA or LA for 24 h. All treatment groups were compared to control, treated without FA and 0 uM H2O2, and their respective FA treated control. Following 24 h, deposited collagen levels produced by fibroblasts were detected using the Sirius red assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The concentration-dependent effects of H2O2, OA and LA, and combination of H2O2 with OA or LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from control, bp<0.05, significantly different between groups treated with different concentrations of H2O2 and the same FA.     63  3.3.2 Effects of Transforming Growth Factor-?1 and Fatty Acid Treatment on Collagen Production in Fibroblasts 3.3.2.1 Effects of 24 h Transforming Growth Factor-?1 with Oleic or Linoleic Acid on Soluble Collagen Production in Fibroblasts Soluble collagen in fibroblasts was assessed following OA or LA treatment following TGF?1-stimulation. Fibroblasts were stimulated with 10 ng/ml TGF?1 and concurrently treated with 0.1 mM OA or LA for 24 h. All results were compared to their control group, unstimulated or stimulated. Following 24 h (Fig. 12), soluble collagen levels in fibroblasts was measured using the Sirius red assay. Stimulated fibroblasts treated with 0.1 mM OA or LA increased soluble collagen levels, whereas unstimulated fibroblasts treated with 0.1 mM OA and LA showed a trend of increased soluble collagen. Although not significant, there is increased soluble collagen between stimulated and unstimulated fibroblasts during OA or LA treatment.    64  ControlOALA0.000.050.100.150.20UnstimulatedStimulatedaa24hSoluble CollagenRAU -  Relative to Control Figure 12. Effect of transforming growth factor-?1 on soluble collagen levels in fibroblasts after 24 h. Fibroblasts were treated with 10 ng/ml TGF?1 with concurrent treatment with 0.1 mM OA or LA for 24 h. Treatment groups were compared to their respective control and stimulated treatment groups were also compared to their unstimulated counterpart. After 24 h, soluble collagen levels in fibroblasts were measured using the Sirius red assay. Values are mean ? SEM. Data from 4 independent experiments are presented. The effects of TGF?1-stimulation on control, OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from respective control.  3.3.2.2 Effects of Transforming Growth Factor-?1 with Oleic or Linoleic Acid on Deposited Collagen Production in Fibroblasts Levels of deposited collagen following OA or LA treatment were assessed in fibroblasts. Fibroblasts were stimulated with 10 ng/ml TGF?1 and concurrently treated with 0.1 mM OA or LA for 24 h. Treatment groups were compared to their respective control and stimulated treatment groups were also compared to their unstimulated counterpart. Following 24 h (Fig. 13), deposited collagen production of fibroblasts was measured using the Sirius red assay. TGF?1-stimulation (10 ng/ml) with 0.1 mM LA increased deposited collagen levels from unstimulated fibroblasts treated with 0.1 mM LA. Although not significant, increased collagen was observed following stimulation with control and OA treatment. 65   ControlOALA-0.010.000.010.020.03UnstimulatedStimulateda,b24hDeposited CollagenRAU -  Relative to Control  Figure 13. Effect of transforming growth factor-?1 on deposited collagen levels of fibroblasts following 24 h. Fibroblasts were treated with 10 ng/ml TGF?1 with concurrent treatment with 0.1 mM OA or LA for 24 h. Treatment groups were compared to their respective control and stimulated treatment groups were also compared to their unstimulated counterpart. After 24 h, deposited collagen levels produced by fibroblasts were quantified using the Sirius red assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The effects of TGF?1-stimulation on control, OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from respective control, bp<0.05, significantly different between unstimulated and stimulated fibroblasts treated with the same FA.  3.3.2.3 Expression of Collagen Genes in Fibroblasts following Transforming Growth Factor-?1 with Oleic or Linoleic Acid Gene expression of collagen genes was measured following TGF?1-stimulation and OA or LA treatment. Fibroblasts were stimulated with 10 ng/ml TGF?1 and concurrently treated with 0.1 mM OA or LA for 24 h. Following 24 h (Fig. 14), mRNA was extracted using the Trizol extraction method and gene expression was analyzed using qPCR. All treatment groups were compared to their respective control and stimulated treatment groups were compared to their unstimulated counterpart. 66  Gene expression of COL3 (Fig. 14A) was increased after exposure to 0.1 mM OA for 24 h. In addition, TIMP1 was increased (Fig. 14B) with 0.1 mM LA following TGF?1-stimulation (10 ng/ml). No significant changes were seen in the gene expression of: COL1?1, TGF?R1 and TGF?R2 (Fig. 14A) or MMP2 and TIMP2 (Fig. 14B). ControlOALAControlOALAControlOALAControlOALA0.01.02.03.0UnstimulatedStimulatedCOL1?1  COL3     TGF?R1   TGF?R2abRelative Fold Change(Normalized to Control)ControlOALAControlOALAControlOALA0.00.51.01.52.0     MMP2        TIMP1            TIMP2aRelative Fold Change(Normalized to Control)AB Figure 14. Evaluation of the collagen gene expression in fibroblasts following combined treatment of transforming growth factor-?1 with oleic or linoleic acid for 24 h. Fibroblasts were treated with 10 ng/ml TGF?1 with concurrent treatment of 0.1 mM OA or LA for 24 h. Following 24 h, mRNA was extracted using the Trizol method, converted to cDNA and further analyzed using qPCR. Treatment groups were compared to their respective control and stimulated treatment groups were compared to their unstimulated counterpart. The following genes were evaluated: (A) COL1?1, TGF?R1 and TGF?R2 and (B) MMP2, TIMP2 and TIMP1. Values are mean ? SEM. Data from 3 ? 5 individual biological replicates are presented. The effects of TGF?1-stimulation on control, OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from unstimulated control, bp<0.05, significantly different between unstimulated and stimulated fibroblasts treated with the same FA. 67   To investigate the expression of collagen genes further, the ratio of COL1?1/COL3 was calculated. The relative amounts of these two collagens are important because they exhibit specific properties. For example, collagen type I is characterized as strong and stiff, whereas collagen type III is characterized more as elastic [304]. Therefore, a high COL1?1/COL3 ratio is indicative of increased cardiac dysfunction due to the high tensile strength of cola1a. Normalized values from each gene were determined as a ratio by dividing the value of COL1?1 by COL3. This ratio is shown in Figure 15. The COL1?1/COL3 ratio was increased following TGF?1-stimulation (10 ng/ml) when fibroblasts were treated with 0.1 mM OA for 24 h. However, OA treatment alone had a downward trend in comparison to unstimulated control cells. Interestingly, LA-treated cells showed an upward trend; however, stimulation did not increase the ratio further.  ControlOleic AcidLinoleic Acid0.00.5.01.52.02.5UnstimulatedStimulatedb24hRelative Fold Change(Normalized to Control)Col1?1/Col3 Ratio Figure 15. Evaluation of the COL1?1/COL3 ratio in fibroblasts following treatment with transforming growth factor-?1 with oleic or linoleic acid for 24 h. Fibroblasts were treated with 10 ng/ml TGF?1 with concurrent treatment of 0.1 mM OA or LA for 24 h. Following 24 h, mRNA was extracted using the Trizol method, converted to cDNA and further analyzed using qPCR Normalized values for each gene were used to calculate the ratio. Treatment groups were compared to their respective control and stimulated treatment groups were compared to their unstimulated counterpart. Values are mean ? SEM. Data from 3 ? 5 individual biological replicates are presented. The effects of TGF?1-stimulation on control, OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from unstimulated control, bp<0.05, significantly different between unstimulated and stimulated fibroblasts treated with the same FA. 68  3.3.3 Collagen Levels In Vivo 3.3.3.1 Gene Expression of Collagen Genes In Vivo Following High Fat Feeding and Mild Cardiac Injury Gene expression of collagen genes were measured in the acute group of mice following four weeks of high fat feeding and mild cardiac injury with ISO. Male C57BL/6 mice were fed for four weeks with NC, OO or CO and following feeding were injected with 120 mg/kg ISO for two consecutive days. Twenty-four hours following the last injection, mice were sacrificed and tissues were collected. mRNA was extracted from tissues, converted to cDNA and analyzed using qPCR. All results were compared to their respective control, and ISO treatment with high fat feeding was compared to high fat feeding alone. Gene expression of TGF?3 was increased (Fig. 16A) in OO fed mice. No significant changes were seen in gene expression of: TGF?R1 and TGF?R2 (Fig. 16A) or COL1?1, COL3 and FN1 (Fig. 16B) or MMP2, TIMP2, MMP9 and TIMP1 (Fig. 16C).  Figure 16. Effect of high fat feeding and mild cardiac injury using isoproterenol on collagen gene expression in vivo. Expression of collagen genes following four weeks of high fat feeding followed by ISO injections was analyzed. Male C57BL/6 mice were fed for four weeks with NC, OO or CO followed by two daily consecutive injections of 120 mg/kg ISO. Treatments were compared to their respective control, and ISO treatment with high fat feeding was compared to high fat feeding alone. From heart tissues, mRNA was extracted, converted to cDNA and analyzed using qPCR. The following genes were evaluated: (A) TGF?3, TGF?R1, TGF?R2 and (B) COL1?1, COL3, FN1 and (C) MMP2, TIMP2, MMP9 and TIMP1. Values are mean ? SEM. Data from 5 - 8 individual biological replicates are presented. The effects of high fat feeding and ISO injections were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from NC control.  69  TGF?3      TGF?R1     TGF?R2NCOOCONCOOCONCOOCO0123ControlIsoproterenolaRelative Fold Change(Normalized to Normal Chow)COL1?1       COL3         FN1NCOOCONCOOCONCOOCO01234Relative Fold Change(Normalized to Normal Chow)NCOOCONCOOCONCOOCONCOOCO02468MMP2  TIMP2   MMP9   TIMP1Relative Fold Change(Normalized to Normal Chow)ABC 70  To investigate the expression of collagen subtypes in vivo, the ratio of COL1?1/COL3 was assessed. As stated previously, COL1?1 contributes tensile strength and stiffness whereas COL3 contributes more of an elastic property [304] thus a high ratio of COL1?1/COL3 is associated with cardiac disease and dysfunction. Normalized values from each gene were determined as a ratio by dividing the value of COL1?1 by COL3. This ratio is shown in Figure 17. The COL1?1/COL3 ratio was increased following cardiac injury with ISO in the CO fed mice. There was no difference between diet groups; however, an upward trend of the ratio was demonstrated in all treatment groups following ISO injection.  Normal ChowOlive OilCorn Oil01234ControlIsoproterenolbRelative Fold Change(Normalized to Normal Chow)Col1?1/Col3 Ratio Figure 17. Evaluation of the COL1?1/COL3 ratio after high fat feeding and mild cardiac injury, using isoproterenol in vivo. Male C57BL/6 mice were fed for four weeks with NC, OO or CO followed by two daily consecutive injections of 120 mg/kg ISO. Normalized gene values were used to calculate the ratio. From heart tissues, mRNA was extracted, converted to cDNA and analyzed using qPCR. Treatments were compared to their respective control, and ISO treatment with high fat feeding was compared to high fat feeding alone. Values are mean ? SEM. Data from 5 - 8 individual biological replicates are presented. The effects of high fat feeding and ISO injections were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from NC control, bp<0.05, significantly different between unstimulated and stimulated fibroblasts treated with the same FA.  71  3.3.3.2 Levels of Collagen from In Vivo Heart Tissues To evaluate long-term effects of cardiac injury on mice fed various high fat diets, a subset of mice termed ?chronic? were sacrificed at 21 days after ISO-induced cardiac injury, to evaluate the effects on cardiac fibrosis. Levels of collagen were qualitatively assessed using Masson?s trichrome stain on heart tissue sections. Male C57BL/6 mice were fed for four weeks with NC, OO or CO and following feeding were injected with 120 mg/kg ISO for two consecutive days. The chronic group of mice was sacrificed at 21 days following the last injection of ISO. Histological sections were stained using Masson?s trichrome stain to visualize collagen fibers (Fig. 18). Interstitial and perivascular fibrosis were imaged for each treatment group. The micrographs demonstrate high levels of interstitial fibrosis in OO and the highest in OO+ISO compared to CO and CO+ISO treatments. After ISO treatment with OO or CO, there was increased fibrous collagen area as well as thicker fibers. The most perivascular fibrosis is found in CO+ISO. Perivascular fibrosis tends to be more diffuse in OO+ISO and CO groups; however, the collagen fibers seem to be more compact in the OO and CO+ISO groups.   72   Figure 18. Representative micrographs of histological heart sections demonstrating interstitial or perivascular fibrosis following high fat feeding and mild cardiac injury in vivo. Male C57BL/6 mice were fed for four weeks with NC, OO or CO followed by two daily consecutive injections of 120 mg/kg ISO. Heart tissue sections were stained with Masson?s trichrome. Collagen fibres are depicted as blue, whereas nuclei are stained black and muscle is stained red. Magnification of 400X.  73  3.4 The Effect of Hyperglycemia on the Fibrotic Response 3.4.1 Collagen Production in Fibroblasts is Different between Low and High Glucose Conditions Soluble and deposited collagen in fibroblasts was assessed after TGF?1-stimulation with OA or LA treatment in low and high glucose conditions. Fibroblasts were stimulated with 10 ng/ml TGF?1 and concurrently treated with 0.1 mM OA or LA for 24 h in high or low glucose. All results were compared to their respective control and stimulated treatments were compared to their unstimulated counterparts. Following 24 h, soluble (Fig. 19) and deposited (Fig. 20) collagen production by fibroblasts was measured using the Sirius red assay. Soluble collagen produced by stimulated control cells was greater in high (Fig. 19A) than low glucose (Fig. 19B). All other treatments demonstrated a similar trend between low and high glucose. In high glucose conditions, there was no change or decreased deposited collagen in all treatment groups (Fig, 20A); however, in low glucose conditions, there were increased levels of deposited collagen with 0.1 mM LA (Fig. 20B). There was also a trend of increased deposited collagen following stimulation with control and 0.1 mM OA in low glucose.     74  ControlOALA0.000.050.100.150.20aaLow GlucoseASoluble CollagenRAU -  Relative to ControlControlOALA0.000.050.100.150.20aHigh GlucoseB UnstimulatedStimulated  Figure 19. Effect of low and high glucose conditions on the production of soluble collagen by fibroblasts following stimulation with transforming growth factor-?1 plus oleic and linoleic acid treatment for 24 h. Fibroblasts were treated with 10 ng/ml TGF?1 with concurrent treatment of 0.1 mM OA or LA for 24 h in high or low glucose. Treatment groups were compared to their respective control and stimulated treatments were compared to their unstimulated counterpart. Following 24 h, soluble collagen levels from fibroblasts grown in low (A) and high (B) glucose conditions were quantified using the Sirius red assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The effect of high and low glucose on soluble collagen production by TGF?1-stimulated fibroblasts with control, OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from respective control.               75  ControlOALA-0.010.000.010.020.03a,bLow GlucoseADeposited CollagenRAU -  Relative to ControlControlOALA-0.015-0.010-0.0050.0000.005aaaHigh GlucoseB UnstimulatedStimulated Figure 20. Effect of low and high conditions on the production of deposited collagen by fibroblasts following stimulation with transforming growth factor-?1 plus oleic and linoleic acid treatment for 24 h. Fibroblasts were treated with 10 ng/ml TGF?1 with concurrent treatment of 0.1 mM OA or LA for 24 h in high or low glucose. Treatment groups were compared to their respective control and stimulated fibroblasts were compared to their unstimulated counterpart. Following 24 h, deposited collagen levels from fibroblasts grown in low (A) and high (B) glucose conditions were measured using the Sirius red assay. Values are mean ? SEM. Data from 3 independent experiments are presented. The effect of high and low glucose on deposited collagen production by TGF?1-stimulated fibroblasts with control, OA and LA were assessed using ANOVA, followed by Tukey?s post-hoc test ap<0.05, significantly different from respective control; bp<0.05, significantly different between unstimulated and stimulated fibroblasts treated with the same FA.                 76  4.0 Discussion 4.1 Cardiomyocyte Viability 4.1.1 Effects of Fatty Acids and Hydrogen Peroxide on Cardiomyocyte Viability 4.1.1.1 Linoleic Acid Reduced the Redox Potential of Cardiomyocytes OA and LA were tested for their effects on cardiomyocyte viability in order to investigate the relevance of fatty-acid induced cardiomyocyte death that occurs in cardiac disease. I hypothesized that excess levels of FAs could cause decreased cardiomyocyte viability. In answer to my first hypothesis, I found that 0.25 mM LA for 12 h caused significant loss of cardiomyocyte viability (Fig. 3). I also observed that 0.25 mM OA for 6 h decreased cardiomyocyte viability but that this loss in cardiomyocyte viability was recovered at 12 h.  The cardiomyocytes had not been exposed to FAs prior; therefore, the initial response from the cells towards FAs could have been to reduce the cell viability at 6 h. As resazurin measures redox potential of the cell (explained in Section 2.7.1), such effects seen with FAs at 6 h could be a compensatory response to mild oxidative stress, which is known to augment redox potential and cell survival of cells [305]. Loss in cell viability was demonstrated at higher or ?excess? levels of both LA and OA within 12 h; however, cells treated with OA recovered from the initial loss in viability. In other cell models such as mouse neural cells, 50 uM LA demonstrated loss of viability after 12 h and increasing loss up until 24 h [306]. Although there are few studies looking at the effect of LA on cardiomyocytes specifically, high levels of omega-6 PUFA (like LA) is correlated with cardiac disease and type 2 diabetes [307], both of which demonstrate cardiomyocyte death. In fact, it is well known that cardiac disease involves cardiomyocyte death, which is known to occur through both apoptosis and necrosis [308]. 77  Moreover, in hearts of type 2 diabetic patients, there is an 85-fold increase in apoptosis and a 4-fold increase in necrosis of myocytes [309]. In general, OA is thought of as beneficial in regards to cardiac disease [310]. For instance, studies have shown that 0.5 mM OA for 48 h does not alter the cell viability of neonatal ventricular myocytes [311]. Moreover, the viability of chick embryonic ventricular myocyte cells was not altered up to 0.1 mM OA for 24 h; however cell death was observed with 0.5 and 1 mM OA [312]. These results demonstrate that excess levels of FAs, including OA, or presence of FAs over long periods can become toxic. A possible mechanism for this observation is the build-up of toxic lipid byproducts [313, 314] through lipid metabolism or lipid peroxidation. 4.1.1.2 High Levels of Hydrogen Peroxide Decreased Cardiomyocyte Viability H2O2 was next investigated for its ability to decrease cardiomyocyte viability. H2O2 was used to simulate different levels of oxidative stress in vitro. In further investigation into my first hypothesis, I found that higher or ?excess? H2O2 at 500 uM (12 h) and 1000 uM (6, 12 h) but not low levels (100 ? 300 uM) caused increased loss of cardiomyocyte viability (Fig. 4). It is well established that low levels of oxidative stress are required within the cell for normal function [23, 29]. Previous studies have demonstrated similar findings of loss in cell viability at higher concentrations of H2O2. For example, 1 mM H2O2 decreased H9c2 myoblast viability by 40% at 24 h [315]. Further studies differentiated between cell necrosis and apoptosis during H2O2 treatment. For instance, H9c2 myoblasts treated with 450 to 600 uM H2O2 for 24h had increased cell necrosis, whereas cells treated with 250 uM H2O2 demonstrated apoptosis of 45% of the cells. Interestingly, cells that survived H2O2 treatment (250 uM or less) demonstrated increased size (hypertrophy) [316]. Moreover, even with short incubation periods like 6 h, H9c2 78  myoblasts showed decreased cell viability starting at 500 uM and an even greater decrease with 750 uM with accompanied cell hypertrophy [317]. 4.1.2 Effects of Fatty Acids in Combination with Hydrogen Peroxide on Cardiomyocyte Viability 4.1.2.1 Effect of Hydrogen Peroxide with Oleic or Linoleic Acid for 6 and 12 h on Cardiomyocyte Viability I have shown that both a combination of excess LA and H2O2 decreased cardiomyocyte viability (Fig. 5, 6). I wanted to investigate the combined effect of FAs with H2O2 because it is known that the prevalence of obesity, cardiac disease and diabetes is linked by oxidative stress and affected by dietary fats. In the last set of experiments for investigation of my first hypothesis, I found that 300 uM H2O2 for 6 h with either 0.1 mM OA or LA increased cardiomyocyte viability (Fig. 5A). A downward trend of viability was seen after 12 h with 0.25 mM OA or LA and 300 uM H2O2 (Fig. 5B). As treatment was increased from 6 to 12 h, 300 uM H2O2 and 0.1 mM LA, but not OA, decreased cardiomyocyte viability. As resazurin measures redox potential of the cell (explained in Section 2.7.1), such effects seen with 300 uM H2O2 and 0.1 mM FAs (6 h) could be a compensatory response to mild oxidative stress, which is known to augment redox potential and cell survival of cells [305]. Excess FAs in combination with oxidative stress decreased viability at short incubation like 6 h. In addition, a low FA concentration like 0.1 mM in the presence of H2O2 also decreased cell viability following longer treatment duration of 12 h. This demonstrates that oxidative stress exacerbates the effects of FA-induced toxicity both at low and high concentration depending on treatment duration. Through the addition of oxidative stress, it is likely that FAs are undergoing lipid peroxidation and the lipid peroxide products are involved in the decreased viability. In fact, 79  increased levels of H2O2 has been shown to increase lipid peroxidation [318]. This increased lipid peroxidation and production of lipid peroxides is associated with atherosclerosis [319], cardiac risk factors [320] and directly to cardiomyocyte toxicity [321]. At a higher concentration of H2O2 (500 uM), I found that there is loss in viability with both 0.1 and 0.25 mM OA and LA (Fig. 5C, D). When the concentration of FA is increased from 0.1 to 0.25 mM, there is an increased loss of viability with OA. I demonstrated that a higher concentration of OA (0.25 mM) in combination with high oxidative stress (500 uM H2O2) has a greater effect in regards to decreased cell viability than LA. After both 6 and 12 h, the change from 0.1 to 0.25 mM OA shows a drastic decrease in cell viability. Although most studies demonstrate the beneficial effects of OA, there is toxicity associated with high concentrations of OA. For example, OA (0.5 and 1 mM) drastically increased cell death from a lower concentration of 0.2 mM in chick embryonic ventricular myocyte cells [312]. In this study, the toxicity of OA may be occurring at a lower concentration like 0.25 mM due to the synergistic toxic effects of H2O2. As H9c2 cells are myofibroblasts with embryonic characteristics, which were induced to have a mature phenotype using retinoic acid, similar toxicities of OA can be present in this particular cell type.  4.2 Fibroblast Viability and Cellular Oxidative Stress 4.2.1 Effects of Fatty Acids on Fibroblast Viability and Reactive Oxygen Species Production 4.2.1.1 Linoleic Acid Reduced Cell Viability in Fibroblasts  LA was demonstrated to decrease fibroblast viability. To test my second hypothesis, OA and LA were first tested for their effects on fibroblast viability. I found that LA at 0.25 mM but 80  not 0.1 mM after 6 h decreased the viability (Fig. 6). In addition, both 0.1 and 0.25 mM OA did not alter viability. These same results were demonstrated at 2 and 4 h as well.  In general, cardiomyocytes are more sensitive to oxidative stress than fibroblasts [322]; however, after 6 h exposure, fibroblasts were more sensitive to 0.1 and 0.25 mM LA and 0.1 mM OA than cardiomyocytes. Previous studies have demonstrated that 0.05 mM OA or LA does not decrease cell viability and actually increases cell number of fibroblasts following 24 h incubation [323]. Our results agree with this study as no significant decrease in cell viability is seen with the lower concentration (0.1 mM) of OA or LA; however, a downward trend of viability was observed with 0.1 mM LA. However, in a model of chondrocytes, which are cells responsible for cartilage matrix formation, a lower concentration (0.05 mM) of LA decreased cell viability [324]. 4.2.1.2 Oleic Acid Increased Reactive Oxygen Species in Fibroblasts  Oxidative stress, high levels of ROS, lead to cell damage and death. In this study, I have demonstrated that LA decreased both cardiomyocyte and fibroblast viability. We tested OA at the lower concentration of 0.1 mM on ROS production in fibroblasts. A lower concentration was used in order to cause the least amount of loss in cell viability. OA increased, whereas LA decreased, ROS levels produced in fibroblasts (Fig. 7). However, at 24 h, there was observed to be less ROS with OA. It is possible that higher levels of ROS are present in OA and not LA-incubated fibroblasts because LA causes more loss in cell viability. Most FAs increase ROS production from cardiomyocytes irrespective of the type [325, 326]. Additionally, LA has been observed to increase the efflux of mitochondrial calcium, which may disrupt mitochondrial function [283] and impact viability. Another study demonstrated that there was a direct correlation of the increase of O2- ions with the concentration of OA or LA for 81  10 min exposure in fibroblasts [325]. Nicotinamide adenine dinucleotide phosphate (NADPH) oxidase is a membrane associate enzyme complex that is known to produce O2- [327]. These FAs caused increased O2- levels at 100 and 200 uM in the presence of beta-NADH or NADPH, involved in the NADPH oxidase reactions; however, OA showed greater values of O2- measurement in comparison to LA. However, in regards to the maximal O2- production, in the presence of beta-NADH or NADPH, LA showed a maximal O2- production at around 4 min whereas OA had maximal O2- production at around 8 min after FA drop [325]. This demonstrates that LA shows a quicker ?oxidative burst? than OA. 4.2.2 Effects of Combination of Fatty Acids with Hydrogen Peroxide on Fibroblast Viability and Glutathione Levels 4.2.2.1 Combination of Fatty Acids and Hydrogen Peroxide Further Decreased Cell Viability  An environment of oxidative stress in fibroblasts was simulated using H2O2. In combination to this simulated environment, FAs were also added to the environment. The combined effect of H2O2 and FAs was used to investigate my second hypothesis further. I found that after 24 h, 100 and 200 uM H2O2 as well as both 0.1 mM OA and LA decreased cell viability (Fig. 8).  A synergistic effect of FAs and 200 uM H2O2 on loss of cell viability was observed; however, this synergistic effect was not seen with combination of 100 uM H2O2. This could demonstrate that different levels of oxidative stress within the cell modulate effects of FAs. Higher FA-induced toxicity may be present in environments of oxidative stress due to increased lipid peroxidation [328]. For example, the lipid peroxidation products of LA cause toxicity in 82  human fibroblasts [329]. In addition, LA increases TNF-alpha-induced oxidative stress and apoptosis in endothelial cells [330]. 4.2.2.2 Hydrogen Peroxide, but Not Fatty Acids, Decreased Glutathione Levels GSH is an endogenous antioxidant that is used in the neutralization of H2O2 and other lipid peroxides by GSH peroxidase [57]. To test the second part of my second hypothesis regarding levels of GSH in fibroblasts, GSH levels were measured following combination of H2O2 and FAs. I found that 200 uM H2O2 decreased GSH levels (Fig. 9). FAs alone or in combination with H2O2 did not decrease levels of GSH significantly but all demonstrated a downward trend of GSH levels. The depletion of GSH has been associated with oxidative stress and cardiac disease. For example, GSH depletion resulted in increased levels of nitrotyrosine accumulation [331] and lipid peroxidation accompanied by cellular damage [332]. In addition, depletion of GSH demonstrated exacerbated cardiac dysfunction and remodeling [333] as well as increased apoptosis, as measured by caspase-3 and -9 activity [334]. 4.3 Collagen Production of Fibroblasts 4.3.1 Effects of Fatty Acids on Collagen Production 4.3.1.1 Fatty Acids and Hydrogen Peroxides Combined Increased Soluble Collagen Production For investigation of my third hypothesis, soluble collagen levels in fibroblasts were measured. I found that 100 and 200 uM H2O2 plus 0.1 mM OA or LA increased soluble collagen 83  (Fig. 10). However, H2O2 alone showed a downward trend of soluble collagen where FAs demonstrated an upward trend of soluble collagen. Studies have demonstrated that O2- [335], OH- and ozone oxidants [336] demonstrate the ability to directly degrade soluble collagen. This agrees with the results in this study that demonstrate H2O2 resulting in a downward trend of soluble collagen. When FAs are added, the soluble collagen content is drastically increased. Therefore, the addition of FA tends to result in increased soluble collagen even in the presence of oxidants, such as H2O2. 4.3.1.2 Effects of a 24 h Combination of Oleic or Linoleic Acid with Hydrogen Peroxide on Deposited Collagen Production by Fibroblasts For further investigation of my third hypothesis, deposited collagen levels of fibroblasts were measured. H2O2 in combination with FAs decreased deposited collagen; however, H2O2 alone decreased deposited collagen, whereas FAs alone caused no difference in deposited collagen levels (Fig. 11). In regards to FA treatment alone, 0.05 mM LA has demonstrated low levels, whereas 0.05 mM OA showed no alteration in collagen synthesis in avian chondrocytes [324]. Our data demonstrate similar findings that LA has a very small downward trend of deposited collagen whereas OA showed no alteration from control. As previously described, oxidants are involved in the degradation of soluble collagen [335, 336]. Furthermore, low levels of these oxidants result in collagen modification and sensitization of the collagen structure to proteolytic degradation [336]. It is reasonable to say that if soluble collagen degradation is increased, then there will be less conversion of remaining soluble collagen into deposited collagen. 84  When FAs are added in combination with H2O2, we see an even further decrease in deposited collagen. This data is in agreement with previous data in this study demonstrating increased soluble collagen with FAs plus H2O2. Therefore, FAs (even in combination with H2O2) tend to have increased soluble but not deposited collagen levels. Another possible explanation of decreased deposited collagen may be that since the structure of the deposited collagen has become more sensitive to degradation due to oxidants, that addition of FAs may further degrade these sensitized structures. 4.3.2 Effects of Transforming Growth Factor-?1 and Fatty Acids on Collagen Production in Fibroblasts 4.3.2.1 Transforming Growth Factor-?1 Increased Soluble Collagen In further investigation of my third hypothesis, I found that following 24 h stimulation with TGF?1, both OA and LA at 0.1 mM caused increased soluble collagen production from stimulated control cells (Fig. 12). Although not significant, there was an increase from unstimulated to stimulated cells incubated with OA and LA. TGF?1 is known to increase soluble collagen production in rat cardiac fibroblasts after 24 and 48 h exposure [298]. This increase in collagen production was attributed to ROS. In addition, when adult rat cardiac fibroblasts were treated with Ang II, known to stimulate TGF?1, increased soluble collagen was present following 24 h [337]. As demonstrated earlier in this study, FAs tended to favor increased soluble collagen and decreased deposited collagen content within unstimulated cells. We see the same trend here with unstimulated cells in regards to soluble collagen content. 85  4.3.2.2 Linoleic Acid Increased Deposited Collagen Following Stimulation with Transforming Growth Factor-?1 In further investigation of my third hypothesis, I found that following 24 h stimulation with TGF?1, LA at 0.1 mM caused increased deposited collagen production compared to unstimulated cells treated with LA (Fig. 13). Although not significant, there was an increase from unstimulated to stimulated cells with control and OA treatment. TGF?1 has been demonstrated to increase non-soluble collagen production [298]. One mechanism of this increase involves LOX, which is an oxidase responsible for the cross-linking of collagen and therefore synthesis of deposited collagen [221]. Studies have demonstrated that TGF?1 increases LOX activity and gene expression in rat aortic smooth muscle cells [338] and lung fibroblasts [339] and fibroblasts [340], respectively. In addition, TNF? increased LOX activity through TGF?1 signaling in adult rat cardiac fibroblasts [341]. Furthermore, it has been demonstrated that 0.01 mM OA did not alter TGF?1-induced collagen synthesis in adult cardiac fibroblasts [342]. 4.3.2.3 COL3 and TIMP1 Genes are Increased with Oleic and Linoleic Acid plus Transforming Growth Factor-?1, Respectively  To investigate my third hypothesis further, I measured expression of collagen genes. Overall, gene expression was not altered except for COL3 and TIMP1. COL3 expression was increased with exposure to 0.1 mM OA for 24 h (Fig. 14A). In addition, TIMP1 expression was increased following treatment with 0.1 mM LA and 10 ng/ml TGF?1 for 24 h (Fig. 14B).  Studies have demonstrated that TGF?1 increases TIMP1 mRNA and protein in bone [343] and cardiac fibroblasts [206] and lung fibroblasts [344], respectively. LA derived 86  peroxides have also demonstrated increases in TIMP-1 levels in rheumatoid synovial fibroblasts [345]. Collagen type III is characterized as a more elastic collagen [304]. Our results demonstrate that OA increases the gene expression of collagen type III. Studies have shown that COL3 mRNA levels are increased in MUFA-fed BALB/c mice with skin wounds. In this model, COL3 acts as a reparative response and the initial wound structure in skin wounds [346]. Collagen type III (COL3) is more elastic and has shown beneficial effects in skin wounds, thus it may provide beneficial fibrosis within the heart for continued cardiac function. However, if excess fibrosis occurs, this elasticity may become adverse to the heart where increased stiffness may be required for contractile force [110]. The ratio of collagens type I and III (COL1?1/COL3) are therefore important in regards to cardiac disease and more specifically in cardiac fibrosis. From the gene expression values, the ratio of COL1?1/COL3 was further evaluated. I found that the ratio was increased in OA treatment following TGF?1-stimulation (Fig. 15). Interestingly, an upward trend of increased ratio was present with LA treatment, which was not altered after TGF?1-stimulation. The Western diet was simulated in C57BL6/J mice and was found to be associated with increased collagen 1 expression and an increased ratio of COL1?1/COL3 [347]. We demonstrate a similar finding in our data with an upward trend of this increased ratio with LA treatment. Increased levels of collagen type I and this ratio are accompanied with increased stiffness and less compliance in the heart [348].  87  4.3.3 Levels of Collagen In Vivo 4.3.3.1 TGF?3 is Increased Following a High Fat Olive Oil Diet  I used an in vivo approach to examine my third hypothesis. I found that following four weeks of an OO high fat diet, TGF?3 was increased compared to NC fed mice (Fig. 16A). Overall, the remaining collagen genes showed no significantly altered expressions.  Although TGF?3 is related to TGF?1, it demonstrates anti-fibrotic effects in various model systems. In fact, TGF?3 down regulates the expression of TGF?1[349]. In addition, TGF?3 has been shown to decrease the deposition of fibronectin and collagen I [350] in the liver [351] and following labioplasty [352]. TGF?3 has been shown to improve wound architecture [350] and when present in higher levels than both TGF?1 and TGF?2, wounds have been shown to heal without scarring [349]. In human cardiac fibroblasts, TGF?3 increases levels of desmosomal genes [353], which when decreased impair wound healing [354]. As we investigated the ratio of COL1?1/COL3 in vitro, we also assessed it in vivo. I found that the ratio was increased after ISO-induced cardiac injury with the CO diet (Fig. 17). Diet treatments alone did not alter the ratio; however, there was an upward trend in the ratio following ISO mediated injury in all diet groups. As previously described, the COL1?1/COL3 ratio is indicative of cardiac function and fibrosis. ISO was used to induce cardiac injury. We see that CO exacerbates this ratio following injury. These results are different from the in vitro COL1?1/COL3 ratio results. To induce increased collagen content or fibrosis, TGF?1-stimulation and ISO injection was used in vitro and in vivo, respectively. We see that the greatest increase of this ratio in vitro occurs in OA treated cells after stimulation where in vivo, it occurs following injury in the CO diet group. 88  However, in vitro data demonstrates that the ratio is high in LA treated cells with or without stimulation. These data may indicate that LA, a component of CO, causes more fibrosis alone or that there may be a component of CO that is protective during diet alone. Although CO consists of very high levels of LA, it also contains OA [355]. On the other hand, the OO diet after injury seems more protective in vivo than OA following stimulation in vitro. Some protective effects in vivo may be attributed not only to OA but to other components of OO such as squalene [356]. The increase in COL1?1/COL3 ratio in vivo could be attributed to increased lipid peroxidation products affecting collagen deposition and fibrosis. ISO injection results in increased lipid peroxidation and O2- production in the heart [357] and has also been attributed to increased type I collagen gene expression [357]. 4.3.3.2 Different Levels of Collagen Fibers Exist Between Diets and After Cardiac Injury In Vivo There are higher amounts of interstitial fibrosis present in OO and OO+ISO then CO and CO+ISO. However, the collagen fibers of interstitial fibrosis tend to be thinner in the OO and CO diets, whereas they tend to have a thicker form after treatment with ISO. In general, collagen type I is thicker than collagen type III [358]. It may be probable that the thinner fibers represent collagen type III, whereas the thicker fibers represent collagen type I. From in vivo gene expressions from this study, the OO and OO+ISO diets demonstrate a lower COL1?1/COL3 ratio indicating a greater amount of COL3, whereas CO+ISO demonstrates a higher COL1?1/COL3 ratio and therefore greater COL1?1. OO also demonstrated increased TGF?3 expression, indicating increased collagen synthesis, possibly collagen type III. This is speculation only at this point, while further investigations on these aspects are currently being carried out in our lab. A study focused on the quantification of collagen discussed the possibility 89  that the thinner fibers may be immature thin collagen type I fiber or that during histological sectioning, a thick collagen type I fiber may have been smeared, thus decreasing its thickness [358]. 4.4 The Effect of Hyperglycemia on the Fibrotic Response 4.4.1 Responses in Collagen Production are Different between Low and Glucose Conditions American type culture collection (ATCC) recommends NIH-3T3 fibroblasts to be cultured in high glucose conditions, which is 25mM D-glucose. Studies in our lab have shown that differences in cell responses exist between high and low glucose conditions (unpublished data). When culturing cells in vitro, the glucose conditions can be manipulated. For example, the three main glucose concentrations are used in order to stimulate three different biological conditions: 5.5 mM (normoglycemic), 10 mM (pre-diabetic) and 25 mM (hyperglycemic or diabetic). In respect to these concentrations, low glucose media has considerably less glucose than high glucose. To evaluate my final and fourth hypothesis, I found that soluble collagen produced by stimulated control fibroblasts was greater in high than in low glucose (Fig. 19). In addition, levels of deposited collagen with LA were increased in low but not high glucose (Fig. 20). Furthermore, under high glucose conditions, there was no further increase in soluble or deposited collagen following TGF?1-stimulation. The initial lack of collagen deposition in high glucose may be directly associated with impaired wound healing that is demonstrated in diabetes (41). Furthermore, high glucose conditions have been associated with increased RhoA and Rac1 activation resulting in decreased cell adhesion and a morphological change from elongated to round cells, indicating loss of cell polarity activation indicating defects in wound healing [171] or fibrosis. 90  However, other studies have shown that high glucose synthesized increased collagen levels in adult rat cardiac fibroblasts [174]. Interestingly, the addition of Ang II [174] and aldosterone [359] do not demonstrate synergistic effects during co-incubation. High levels of D-glucose has also been shown to increase the stimulation of myofibroblasts [359].                91  5.0 Conclusions and Future Work 5.1 Limitations of Research The first limitation of this study is that the in vitro experimental data was completed with the use of cell lines (e.g. cardiomyocytes and fibroblasts). The H9c2 and NIH/3T3 cells are derived from rat and mouse models respectively, but both may exhibit different responses from their primary cell counterparts. In order to overcome this first limitation, in vivo studies were completed in mice following ISO injury in order to stimulate cardiomyocyte cell death and fibroblast function.  Another limitation in this study is the ROS measurement in fibroblasts. The experimental data demonstrated increased ROS production with OA and decreased ROS production with LA. However, I also demonstrate that there is less viability of cells with LA but greater viability with OA. ROS production correlates not only to oxidative stress but also to the number of cells/viable cells present. To overcome this limitation, the ROS production of fibroblasts figure can be compared to the viability of fibroblasts at the 6 h and 24 h figures. I also added an explanation for this occurrence in the discussion chapter. 5.2 Future Work  In this study, qPCR was used to look at levels of collagen gene expressions. In future, western blot analysis will be completed to further look at the protein levels of these collagen genes. In addition, primary cardiomyocyte and fibroblast cells were studied to assess the responses similarly to H9c2 cardiomyocyte and NIH/3T3 fibroblast cells in this study. 5.3 Significance of Findings Overall, this study demonstrates that LA and CO cause pathological cardiac fibrosis, whereas OA and OO demonstrated more adaptive beneficial effects. Overall, these results 92  indicate that it is important to implement dietary guidelines in respect to the consumption of dietary fats. In addition, by reducing omega-6 PUFAs, mainly LA, the prevalence of fibrosis in cardiac disease could decrease. Not many studies have investigated the role of LA or OA in regards to cardiomyocyte viability and fibroblast function, i.e. fibrosis. However, many studies demonstrate beneficial effects with OA or OO and detrimental effects with LA or CO in regards to cardiac disease. My findings demonstrate that both OA and LA play a role in fibrosis but result in opposite responses. LA demonstrated an adverse fibrotic modeling process. For example, LA decreased viability of cardiomyocytes and fibroblasts, increased deposited collagen in vitro and inhibited ECM/fibrotic area degradation via TIMP1 in vivo. Following in vivo cardiac injury, the COL1?1/COL3 ratio was increased with the CO diet indicating a greater stiffness of the fibrotic area. In addition, LA demonstrated a high COL1?1/COL3 ratio in vitro before and after stimulation. On the other hand, OA demonstrated a more beneficial or adaptive fibrotic process. For example, although OA demonstrated loss of viability after a longer exposure time in vitro, it demonstrated an increased COL3 gene expression in vitro. In addition, a diet high in OO increased TGF?3 gene expression and demonstrated a low COL1?1/COL3 ratio in vivo. As collagen type III is more elastic, the increase in col3 and the low COL1?1/COL3 ratio indicate a more elastic fibrotic area, which is more beneficial for maintenance of cardiac function in early fibrosis. Although the COL1?1/COL3 ratio increases following TGF?1-stimulation in vitro, there is no change in this ratio following cardiac injury by ISO in vivo. However, the in vitro and in vivo results did not match completely. These differences can be attributed to the use of non-primary cell lines. These cells lines are most likely metabolically different from primary cell lines, which may be a better representative in comparison to in vivo models. However, the exact mechanisms of these discrepancies remain outside the scope of my Master?s thesis. 93  References 1. Shields, M. and M. Tjepkema, Trends in adult obesity, in Health Rep, S. Canada, Editor 2006. p. 53-9. 2. Ogden, C.L., et al., Prevalence of overweight and obesity in the United States, 1999-2004. JAMA - J Am Med Assoc, 2006. 295(13): p. 1549-55. 3. Grundy, S.M., Metabolic syndrome pandemic. Arterioscler Thromb Vac Biol, 2008. 28(4): p. 629-36. 4. Riediger, N.D. and I. Clara, Prevalence of metabolic syndrome in the Canadian adult population. Can Med Assoc J, 2011. 183(15): p. E1127-34. 5. Ervin, R.B., Prevalence of metabolic syndrome among adults 20 years of age and over, by sex, age, race and ethnicity, and body mass index: United States, 2003-2006. Natl Health Stat Report, 2009(13): p. 1-7. 6. Grundy, S.M., et al., Definition of metabolic syndrome: Report of the National Heart, Lung, and Blood Institute/American Heart Association conference on scientific issues related to definition. Circulation, 2004. 109(3): p. 433-8. 7. Alberti, K.G. and P.Z. Zimmet, Definition, diagnosis and classification of diabetes mellitus and its complications. Part 1: diagnosis and classification of diabetes mellitus provisional report of a WHO consultation. Diabet Med, 1998. 15(7): p. 539-53. 8. Alberti, K.G., P. Zimmet, and J. Shaw, Metabolic syndrome--a new world-wide definition. A Consensus Statement from the International Diabetes Federation. Diabet Med, 2006. 23(5): p. 469-80. 9. III, N.A., Third Report of the National Cholesterol Education Program (NCEP) Expert Panel on Detection, Evaluation, and Treatment of High Blood Cholesterol in Adults (Adult Treatment Panel III) Final Report. Circulation, 2002. 106(25): p. 3143. 94  10. Diabetes: Canada at the tipping point. Charting a new path, D.Q. Canadian Diabetes Association, Editor 2011. 11. Pelletier, C., et al., Report summary. Diabetes in Canada: facts and figures from a public health perspective. Chronic Dis Inj Can, 2012. 33(1): p. 53-4. 12. Economic costs of diabetes in the U.S. in 2012. Diabetes Care, 2013. 36(4): p. 1033-46. 13. Whiting, D.R., et al., IDF Diabetes Atlas: Global estimates of the prevalence of diabetes for 2011 and 2030. Diabetes Res Clin Pract, 2011. 94(3): p. 311-321. 14. Diabetes Atlas, Fifth Edition, I.D. Federation, Editor 2011: Brussels, Belgium. 15. The Prevalence and Costs of Diabetes, C.D. Association, Editor 2008. 16. Browarski, S., Stonebridge, C., Theriault, L., The Canadian Heart Health Strategy: Risk Factors and Future Cost Implications, T.C.B.o. Canada, Editor 2010. 17. Mortality, Summary List of Causes, S. Canada, Editor 2009. 18. Avery, S.V., Molecular targets of oxidative stress. Biochem J, 2011. 434(2): p. 201-10. 19. Yang, H., et al., Oxidative stress and diabetes mellitus. Clin Chem Lab Med, 2011. 49(11): p. 1773-82. 20. Fucci, L., et al., Inactivation of key metabolic enzymes by mixed-function oxidation reactions: Possible implication in protein turnover and ageing. Proc Natl A Sci, 1983. 80(6): p. 1521-1525. 21. Konat, G., H2O2-induced higher order chromatin degradation: A novel mechanism of oxidative genotoxicity. J Biosci, 2003. 28(1): p. 57-60. 22. Thollon, C., et al., Nature of the cardiomyocyte injury induced by lipid hydroperoxides. Cardiovasc Res, 1995. 30(5): p. 648-655. 95  23. Mahadev, K., et al., The NAD(P)H oxidase homolog Nox4 modulates insulin-stimulated generation of H2O2 and plays an integral role in insulin signal transduction. Mol Cell Biol, 2004. 24(5): p. 1844-54. 24. Janeway, C., Immunobiology : the immune system in health and disease. 6th ed2005, New York: Garland Science. xxiii, 823 p. 25. Kunin, S. and R. Gallily, Recognition and lysis of altered-self cells by macrophages. I. Modification of target cells by 2,4,6-trinitrobenzene sulphonic acid. Immunology, 1983. 48(2): p. 265-272. 26. Savill, J., et al., Phagocyte recognition of cells undergoing apoptosis. Immunol Today, 1993. 14(3): p. 131-136. 27. Fang, F.C., Antimicrobial reactive oxygen and nitrogen species: concepts and controversies. Nat Rev Microbiol, 2004. 2(10): p. 820-832. 28. Fang, F.C. and A. Vazquez-Torres, Nitric oxide production by human macrophages: there's NO doubt about it. Am J Physiol-Lung C, 2002. 282(5): p. L941-L943. 29. Zhivotovsky, B., et al., Ca2+ and Endonuclease Activation in Radiation-Induced Lymphoid Cell Death. Exp Cell Res, 1993. 207(1): p. 163-170. 30. Giordano, F.J., Oxygen, oxidative stress, hypoxia, and heart failure. J Clin Invest, 2005. 115(3): p. 500-8. 31. Cai, H. and D.G. Harrison, Endothelial Dysfunction in Cardiovascular Diseases: The Role of Oxidant Stress. Circ Res, 2000. 87(10): p. 840-844. 32. Zalba, G., et al., Oxidative Stress in Arterial Hypertension: Role of NAD(P)H Oxidase. Hypertension, 2001. 38(6): p. 1395-1399. 96  33. Cifuentes, M.E., et al., Upregulation of p67 phox and gp91 phox in aortas from angiotensin II-infused mice. American Journal of Physiology - Heart and Circulatory Physiology, 2000. 279(5): p. H2234-H2240. 34. Xu, Y., et al., Formation of Hydrogen Peroxide and Reduction of Peroxynitrite via Dismutation of Superoxide at Reperfusion Enhances Myocardial Blood Flow and Oxygen Consumption in Postischemic Mouse Heart. J Pharmacol Exp Ther, 2008. 327(2): p. 402-410. 35. Anderson, E.J., et al., Mitochondrial H2O2 emission and cellular redox state link excess fat intake to insulin resistance in both rodents and humans. J Clin Invest, 2009. 119(3): p. 573-581. 36. Rosignoli, P., et al., Protective activity of butyrate on hydrogen peroxide-induced DNA damage in isolated human colonocytes and HT29 tumour cells. Carcinogenesis, 2001. 22(10): p. 1675-1680. 37. Halliwell, B.G., J. M. C. , Free radicals in biology and medicine. 3rd ed1999: Clarendon Press; Oxford University Press  38. Schubert, J. and J.W. Wilmer, Does hydrogen peroxide exist ?free? in biological systems? Free Radic Biol Med, 1991. 11(6): p. 545-555. 39. Nulton-Persson, A.C. and L.I. Szweda, Modulation of Mitochondrial Function by Hydrogen Peroxide. J Biol Chem, 2001. 276(26): p. 23357-23361. 40. Ballinger, S.W., et al., Hydrogen Peroxide? and Peroxynitrite-Induced Mitochondrial DNA Damage and Dysfunction in Vascular Endothelial and Smooth Muscle Cells. Circ Res, 2000. 86(9): p. 960-966. 97  41. Minotti, G. and S.D. Aust, The requirement for iron (III) in the initiation of lipid peroxidation by iron (II) and hydrogen peroxide. J Biol Chem, 1987. 262(3): p. 1098-104. 42. Thomas, C., et al., Hydroxyl radical is produced via the Fenton reaction in submitochondrial particles under oxidative stress: implications for diseases associated with iron accumulation. Redox Rep, 2009. 14(3): p. 102-108. 43. Chen, S.-x. and P. Schopfer, Hydroxyl-radical production in physiological reactions. Eur J Biochem, 1999. 260(3): p. 726-735. 44. Ungvari, Z., et al., Free radical production, antioxidant capacity, and oxidative stress response signatures in fibroblasts from Lewis dwarf rats: effects of life span-extending peripubertal GH treatment. J Gerontol A Biol Sci Med Sci, 2011. 66(5): p. 501-10. 45. Rahman, K., Studies on free radicals, antioxidants, and co-factors. Clin Interv Aging, 2007. 2(2): p. 219-36. 46. Sies, H., Oxidative stress: oxidants and antioxidants. Exp Physiol, 1997. 82(2): p. 291-5. 47. Maritim, A.C., R.A. Sanders, and J.B. Watkins, 3rd, Diabetes, oxidative stress, and antioxidants: a review. J Biochem Mol Toxicol, 2003. 17(1): p. 24-38. 48. Kumar, D. and B.I. Jugdutt, Apoptosis and oxidants in the heart. J Lab Clin Med, 2003. 142(5): p. 288-97. 49. Mao, G.D., et al., Superoxide dismutase (SOD)-catalase conjugates. Role of hydrogen peroxide and the Fenton reaction in SOD toxicity. J Biol Chem, 1993. 268(1): p. 416-20. 50. Packer, L., Methods in Enzymology: Superoxide Dismutase2002: Academic Press. 51. Xu, Y., et al., Mutations in the SOD2 Promoter Reveal a Molecular Basis for an Activating Protein 2-Dependent Dysregulation of Manganese Superoxide Dismutase Expression in Cancer Cells. Mol Can Res, 2008. 6(12): p. 1881-1893. 98  52. Marklund, S.L., Human copper-containing superoxide dismutase of high molecular weight. P Natl A Sci, 1982. 79(24): p. 7634-7638. 53. Kang, Y.J., Y. Chen, and P.N. Epstein, Suppression of Doxorubicin Cardiotoxicity by Overexpression of Catalase in the Heart of Transgenic Mice. J Biol Chem, 1996. 271(21): p. 12610-12616. 54. Deisseroth, A. and A.L. Dounce, Catalase: Physical and chemical properties, mechanism of catalysis, and physiological role. Physiol Rev, 1970. 50(3): p. 319-75. 55. Shingu, M., et al., Human vascular smooth muscle cells and endothelial cells lack catalase activity and are susceptible to hydrogen peroxide. Inflammation, 1985. 9(3): p. 309-320. 56. Michiels, C., et al., Importance of SE-glutathione peroxidase, catalase, and CU/ZN-SOD for cell survival against oxidative stress. Free Radic Biol Med, 1994. 17(3): p. 235-248. 57. Hayes, J.D. and L.I. McLellan, Glutathione and glutathione-dependent enzymes represent a co-ordinately regulated defence against oxidative stress. Free Radic Res, 1999. 31(4): p. 273-300. 58. Jones, D.P., et al., Detoxification reactions in isolated hepatocytes. Role of glutathione peroxidase, catalase, and formaldehyde dehydrogenase in reactions relating to N-demethylation by the cytochrome P-450 system. J Biol Chem, 1978. 253(17): p. 6031-7. 59. Forgione, M.A., et al., Heterozygous Cellular Glutathione Peroxidase Deficiency in the Mouse: Abnormalities in Vascular and Cardiac Function and Structure. Circulation, 2002. 106(9): p. 1154-1158. 60. Blankenberg, S., et al., Glutathione Peroxidase 1 Activity and Cardiovascular Events in Patients with Coronary Artery Disease. N Engl J Med, 2003. 349(17): p. 1605-1613. 99  61. Meister, A. and M.E. Anderson, GLUTATHIONE. Annu Rev Biochem, 1983. 52: p. 711-760. 62. Arrigoni, O. and M.C. De Tullio, Ascorbic acid: much more than just an antioxidant. Biochim Biophys Acta, 2002. 1569(1?3): p. 1-9. 63. Devasagayam, T.P.A., et al., Activity of thiols as singlet molecular oxygen quenchers. J Photoch Photobio B, 1991. 9(1): p. 105-116. 64. Yadav, A. and P. Mishra, Modeling the activity of glutathione as a hydroxyl radical scavenger considering its neutral non-zwitterionic form. J Mol Model, 2013. 19(2): p. 767-777. 65. Jaworski, K., et al., S-nitrosothiols do not induce oxidative stress, contrary to other nitric oxide donors, in cultures of vascular endothelial or smooth muscle cells. Eur J Pharmacol, 2001. 425(1): p. 11-19. 66. Goicoechea, E., et al., Fate in digestion in vitro of several food components, including some toxic compounds coming from omega-3 and omega-6 lipids. Food Chem Toxicol, 2011. 49(1): p. 115-24. 67. Moran, L.A. and J.D. Rawn, Biochemistry. 2nd ed1994, Englewood Cliffs, NJ: Neil Patterson Publishers. 68. Lilly, L.S. and Harvard Medical School., Pathophysiology of heart disease : a collaborative project of medical students and faculty. 5th ed2011, Baltimore, MD: Wolters Kluwer/Lippincott Williams & Wilkins. xiv, 461 p. 69. Grynberg, A. and L. Demaison, Fatty Acid Oxidation in the Heart. J Cardiovasc Pharm, 1996. 28: p. 11-17. 70. Bing, R.J., et al., Metabolism of the human heart: II. Studies on fat, ketone and amino acid metabolism. Am J Med, 1954. 16(4): p. 504-515. 100  71. Herrero, P., et al., Increased Myocardial Fatty Acid Metabolism in Patients With Type 1 Diabetes Mellitus. J Am Coll Cardiol, 2006. 47(3): p. 598-604. 72. Mir?, ?., et al., Aging is associated with increased lipid peroxidation in human hearts, but not with mitochondrial respiratory chain enzyme defects. Cardiovasc Res, 2000. 47(3): p. 624-631. 73. Campbell, N.A. and J.B. Reece, Biology. 7th ed2005, San Francisco: Pearson, Benjamin Cummings. xl, 1231 p. 74. Bers, D.M., Calcium cycling and signaling in cardiac myocytes. Annu Rev Physiol, 2008. 70: p. 23-49. 75. Voet, D.V., J.G., Biochemistry. 3 ed2005, New York: J. Wiley. 1616. 76. Jouaville, L.S., et al., Regulation of mitochondrial ATP synthesis by calcium: Evidence for a long-term metabolic priming. Proc Natl A Sci, 1999. 96(24): p. 13807-13812. 77. Crompton, M., The role of calcium in the function and dysfunction of heart mitochondria. In: Langer GA, ed. Calcium and the Heart. New York: Raven Press, 1990: p. 167-198. 78. Brookes, P.S., et al., Calcium, ATP, and ROS: a mitochondrial love-hate triangle. American Journal of Physiology - Cell Physiology, 2004. 287(4): p. C817-C833. 79. Gregorio, C.C. and P.B. Antin, To the heart of myofibril assembly. Trends Cell Biol, 2000. 10(9): p. 355-362. 80. Kitsis, R.N. and J. Scheuer, Functional significance of alterations in cardiac contractile protein isoforms. Clin Cardiol, 1996. 19(1): p. 9-18. 81. Chalovich, J.M., P.B. Chock, and E. Eisenberg, Mechanism of action of troponin . tropomyosin. Inhibition of actomyosin ATPase activity without inhibition of myosin binding to actin. J Biol Chem, 1981. 256(2): p. 575-578. 101  82. Noland Jr, T.A. and J.F. Kuo, Protein Kinase C Phosphorylation of Cardiac Troponin I and Troponin T Inhibits Ca2+-Stimulated MgATPase Activity in Reconstituted Actomyosin and Isolated Myofibrils, and Decreases Actin-Myosin Interactions. J Mol Cell Cardiol, 1993. 25(1): p. 53-65. 83. Moore, G.W., et al., Constituents of the human ventricular myocardium: connective tissue hyperplasia accompanying muscular hypertrophy. Am Heart J, 1980. 100(5): p. 610-6. 84. Nag, A.C., Study of non-muscle cells of the adult mammalian heart: a fine structural analysis and distribution. Cytobios, 1980. 28(109): p. 41-61. 85. Kanekar, S., et al., Cardiac Fibroblasts: Form and Function. Cardiovasc Pathol, 1998. 7(3): p. 127-133. 86. Fernandez-Madrid, F., S. Noonan, and J. Riddle, The "spindle-shaped" body in fibroblasts: intracellular collagen fibrils. J Anat, 1981. 132(Pt 2): p. 157-66. 87. LaFramboise, W.A., et al., Cardiac fibroblasts influence cardiomyocyte phenotype in vitro. Am J Physiol-Cell Ph, 2007. 292(5): p. C1799-C1808. 88. Borg, T.K. and J.B. Caulfield, The collagen matrix of the heart. Fed Proc, 1981. 40(7): p. 2037-41. 89. Schaper, J. and B. Speiser, The extracellular matrix in the failing human heart. Basic Res Cardiol, 1992. 87 Suppl 1: p. 303-9. 90. Huang, J.X., et al., Mechanisms of cell transformation in the embryonic heart. Ann N Y Acad Sci, 1995. 752: p. 317-30. 91. Zoumi, A., A. Yeh, and B.J. Tromberg, Imaging cells and extracellular matrix in vivo by using second-harmonic generation and two-photon excited fluorescence. P Natl A Sci, 2002. 99(17): p. 11014-11019. 102  92. Parker, K.K. and D.E. Ingber, Extracellular matrix, mechanotransduction and structural hierarchies in heart tissue engineering. Philos T Roy Soc B, 2007. 362(1484): p. 1267-1279. 93. Eghbali, M. and K.T. Weber, Collagen and the myocardium: fibrillar structure, biosynthesis and degradation in relation to hypertrophy and its regression. Mol Cell Biochem, 1990. 96(1): p. 1-14. 94. Fisher, S.A. and M. Absher, Norepinephrine and ANG II stimulate secretion of TGF-beta by neonatal rat cardiac fibroblasts in vitro. Am J Physiol-Cell Ph, 1995. 268(4): p. C910-C917. 95. Siwik, D.A., D.L.-F. Chang, and W.S. Colucci, Interleukin-1? and Tumor Necrosis Factor-? Decrease Collagen Synthesis and Increase Matrix Metalloproteinase Activity in Cardiac Fibroblasts In Vitro. Circ Res, 2000. 86(12): p. 1259-1265. 96. Bosman, F.T. and I. Stamenkovic, Functional structure and composition of the extracellular matrix. J Pathol, 2003. 200(4): p. 423-8. 97. Hubert, H.B., et al., Obesity as an independent risk factor for cardiovascular disease: a 26-year follow-up of participants in the Framingham Heart Study. Circulation, 1983. 67(5): p. 968-77. 98. He, J., et al., Risk factors for congestive heart failure in us men and women: Nhanes i epidemiologic follow-up study. Arch Intern Med, 2001. 161(7): p. 996-002. 99. Karason, K., et al., Effects of obesity and weight loss on left ventricular mass and relative wall thickness: survey and intervention study. Brit Med J, 1997. 315(7113): p. 912-916. 100. Mizushige, K., et al., Alteration in Left Ventricular Diastolic Filling and Accumulation of Myocardial Collagen at Insulin-Resistant Prediabetic Stage of a Type II Diabetic Rat Model. Circulation, 2000. 101(8): p. 899-907. 103  101. McGavock, J.M., et al., Adiposity of the Heart*, Revisited. Ann Intern Med, 2006. 144(7): p. 517-524. 102. Cohn, J.N., R. Ferrari, and N. Sharpe, Cardiac remodeling?concepts and clinical implications: a consensus paper from an international forum on cardiac remodeling. J Am Coll Cardiol, 2000. 35(3): p. 569-582. 103. Rumberger, J.A., et al., Nonparallel changes in global left ventricular chamber volume and muscle mass during the first year after transmural myocardial infarction in humans. J Am Coll Cardiol, 1993. 21(3): p. 673-82. 104. Anversa, P., G. Olivetti, and J.M. Capasso, Cellular basis of ventricular remodeling after myocardial infarction. Am J Cardiol, 1991. 68(14): p. 7D-16D. 105. Watkins, S., G. Borthwick, and H. Arthur, The H9C2 cell line and primary neonatal cardiomyocyte cells show similar hypertrophic responses in vitro. In Vitro Cell Dev-An, 2011. 47(2): p. 125-131. 106. Mitchell, G.F., et al., Left ventricular remodeling in the year after first anterior myocardial infarction: a quantitative analysis of contractile segment lengths and ventricular shape. J Am Coll Cardiol, 1992. 19(6): p. 1136-44. 107. G?lvez, A.S., et al., Distinct Pathways Regulate Proapoptotic Nix and BNip3 in Cardiac Stress. J Biol Chem, 2006. 281(3): p. 1442-1448. 108. Kehat, I. and J.D. Molkentin, Molecular Pathways Underlying Cardiac Remodeling During Pathophysiological Stimulation. Circulation, 2010. 122(25): p. 2727-2735. 109. Gurtner, G.C., et al., Wound repair and regeneration. Nature, 2008. 453(7193): p. 314-321. 110. Weber, K.T., et al., Collagen remodeling of the pressure-overloaded, hypertrophied nonhuman primate myocardium. Circ Res, 1988. 62(4): p. 757-65. 104  111. Spinale, F.G., Myocardial matrix remodeling and the matrix metalloproteinases: influence on cardiac form and function. Physiol Rev, 2007. 87(4): p. 1285-342. 112. Matsusaka, H., et al., Targeted deletion of matrix metalloproteinase 2 ameliorates myocardial remodeling in mice with chronic pressure overload. Hypertension, 2006. 47(4): p. 711-7. 113. Sadoshima, J. and S. Izumo, Mechanical stretch rapidly activates multiple signal transduction pathways in cardiac myocytes: potential involvement of an autocrine/paracrine mechanism. Embo J, 1993. 12(4): p. 1681-92. 114. Kojima, M., et al., Angiotensin II receptor antagonist TCV-116 induces regression of hypertensive left ventricular hypertrophy in vivo and inhibits the intracellular signaling pathway of stretch-mediated cardiomyocyte hypertrophy in vitro. Circulation, 1994. 89(5): p. 2204-11. 115. Yamazaki, T., et al., Endothelin-1 Is Involved in Mechanical Stress-induced Cardiomyocyte Hypertrophy. J Biol Chem, 1996. 271(6): p. 3221-3228. 116. Bunda, S., et al., Aldosterone Induces Elastin Production in Cardiac Fibroblasts through Activation of Insulin-Like Growth Factor-I Receptors in a Mineralocorticoid Receptor-Independent Manner. Am J Pathol, 2007. 171(3): p. 809-819. 117. DUNCAN, M.R., et al., Connective tissue growth factor mediates transforming growth factor ?-induced collagen synthesis: down-regulation by cAMP. FASEB J, 1999. 13(13): p. 1774-1786. 118. Nain, S., et al., The role of oxidative stress in the development of congestive heart failure in a chicken genotype selected for rapid growth. Avian Pathol, 2008. 37(4): p. 367-73. 119. Dimitrow, P.P., et al., Enhanced oxidative stress in hypertrophic cardiomyopathy. Pharmacol Rep, 2009. 61(3): p. 491-5. 105  120. Nakamura, K., et al., 4-Hydroxy-2-nonenal induces calcium overload via the generation of reactive oxygen species in isolated rat cardiac myocytes. J Card Fail, 2009. 15(8): p. 709-16. 121. Flesch, M., et al., Effect of ?-Blockers on Free Radical?Induced Cardiac Contractile Dysfunction. Circulation, 1999. 100(4): p. 346-353. 122. Grieve, D.J. and A.M. Shah, Oxidative stress in heart failure - More than just damage. Eur Heart J, 2003. 24(24): p. 2161-2163. 123. Mercurio, F. and A.M. Manning, NF-kappaB as a primary regulator of the stress response. Oncogene, 1999. 18(45): p. 6163-71. 124. Nakamura, K., et al., Inhibitory Effects of Antioxidants on Neonatal Rat Cardiac Myocyte Hypertrophy Induced by Tumor Necrosis Factor-? and Angiotensin II. Circulation, 1998. 98(8): p. 794-799. 125. Ushio-Fukai, M., et al., p38 Mitogen-activated Protein Kinase Is a Critical Component of the Redox-sensitive Signaling Pathways Activated by Angiotensin II. J Biol Chem, 1998. 273(24): p. 15022-15029. 126. Higuchi, Y., et al., Involvement of Reactive Oxygen Species-mediated NF- ? B Activation in TNF- ? -induced Cardiomyocyte Hypertrophy. J Mol Cell Cardiol, 2002. 34(2): p. 233-240. 127. Kameda, K., et al., Correlation of oxidative stress with activity of matrix metalloproteinase in patients with coronary artery disease. Eur Heart J, 2003. 24(24): p. 2180-2185. 128. Siwik, D.A., P.J. Pagano, and W.S. Colucci, Oxidative stress regulates collagen synthesis and matrix metalloproteinase activity in cardiac fibroblasts. Am J Physiol-Cell Ph, 2001. 280(1): p. C53-C60. 106  129. Wang, P., et al., Hydrogen peroxide-mediated oxidative stress and collagen synthesis in cardiac fibroblasts: Blockade by tanshinone IIA. J Ethnopharmacol, 2013. 145(1): p. 152-161. 130. Ishizaka, N., et al., Iron Overload Augments Angiotensin II?Induced Cardiac Fibrosis and Promotes Neointima Formation. Circulation, 2002. 106(14): p. 1840-1846. 131. Bujak, M. and N.G. Frangogiannis, The role of TGF-? signaling in myocardial infarction and cardiac remodeling. Cardiovasc Res, 2007. 74(2): p. 184-195. 132. Meiners, S., et al., Downregulation of Matrix Metalloproteinases and Collagens and Suppression of Cardiac Fibrosis by Inhibition of the Proteasome. Hypertension, 2004. 44(4): p. 471-477. 133. Kapur, N.K., et al., Reduced Endoglin Activity Limits Cardiac Fibrosis and Improves Survival in Heart Failure. Circulation, 2012. 125(22): p. 2728-2738. 134. Beltrami, C.A., et al., Structural basis of end-stage failure in ischemic cardiomyopathy in humans. Circulation, 1994. 89(1): p. 151-63. 135. Bishop, J.E., et al., Increased collagen synthesis and decreased collagen degradation in right ventricular hypertrophy induced by pressure overload. Cardiovasc Res, 1994. 28(10): p. 1581-1585. 136. Crowley, S.D., et al., Angiotensin II causes hypertension and cardiac hypertrophy through its receptors in the kidney. P Natl A Sci, 2006. 103(47): p. 17985-17990. 137. Tomita, H., et al., Early Induction of Transforming Growth Factor-? via Angiotensin II Type 1 Receptors Contributes to Cardiac Fibrosis Induced by Long-term Blockade of Nitric Oxide Synthesis in Rats. Hypertension, 1998. 32(2): p. 273-279. 138. Anderson, K.R., M.G. Sutton, and J.T. Lie, Histopathological types of cardiac fibrosis in myocardial disease. J Pathol, 1979. 128(2): p. 79-85. 107  139. Grimm, D., et al., Differential effects of growth hormone on cardiomyocyte and extracellular matrix protein remodeling following experimental myocardial infarction. Cardiovasc Res, 1998. 40(2): p. 297-306. 140. Isoyama, S. and Y. Nitta-Komatsubara, Acute and chronic adaptation to hemodynamic overload and ischemia in the aged heart. Heart Fail Rev, 2002. 7(1): p. 63-9. 141. Hasenfuss, G., Animal models of human cardiovascular disease, heart failure and hypertrophy. Cardiovasc Res, 1998. 39(1): p. 60-76. 142. Zhou, B., et al., Genetic fate mapping demonstrates contribution of epicardium-derived cells to the annulus fibrosis of the mammalian heart. Dev Biol, 2010. 338(2): p. 251-61. 143. Kohl, P., Heterogeneous cell coupling in the heart: an electrophysiological role for fibroblasts. Circ Res, 2003. 93(5): p. 381-3. 144. Kohl, P., et al., Mechanosensitive fibroblasts in the sino-atrial node region of rat heart: interaction with cardiomyocytes and possible role. Exp Physiol, 1994. 79(6): p. 943-956. 145. Zhang, Y., et al., Connexin43 expression levels influence intercellular coupling and cell proliferation of native murine cardiac fibroblasts. Cell Commun Adhes, 2008. 15(3): p. 289-303. 146. Gaudesius, G., et al., Coupling of cardiac electrical activity over extended distances by fibroblasts of cardiac origin. Circ Res, 2003. 93(5): p. 421-8. 147. Camelliti, P., et al., Fibroblast Network in Rabbit Sinoatrial Node: Structural and Functional Identification of Homogeneous and Heterogeneous Cell Coupling. Circ Res, 2004. 94(6): p. 828-835. 148. Nagase, H. and J.F. Woessner, Matrix Metalloproteinases. J Biol Chem, 1999. 274(31): p. 21491-21494. 108  149. Montfort, I. and R. Perez-Tamayo, The distribution of collagenase in normal rat tissues. J Histochem Cytochem, 1975. 23(12): p. 910-20. 150. Awad, A.E., et al., Tumor necrosis factor induces matrix metalloproteinases in cardiomyocytes and cardiofibroblasts differentially via superoxide production in a PI3K?-dependent manner. Am J Physiol-Cell Ph, 2010. 298(3): p. C679-C692. 151. Romanic, A.M., et al., Matrix metalloproteinase expression in cardiac myocytes following myocardial infarction in the rabbit. Life Sci, 2001. 68(7): p. 799-814. 152. Aicher, W.K., et al., Transcription factor early growth response 1 activity up-regulates expression of tissue inhibitor of metalloproteinases 1 in human synovial fibroblasts. Arthritis Rheum, 2003. 48(2): p. 348-359. 153. Kajanne, R., et al., EGF-R regulates MMP function in fibroblasts through MAPK and AP-1 pathways. J Cell Physiol, 2007. 212(2): p. 489-497. 154. Li, Y.Y., C.F. McTiernan, and A.M. Feldman, Proinflammatory cytokines regulate tissue inhibitors of metalloproteinases and disintegrin metalloproteinase in cardiac cells. Cardiovasc Res, 1999. 42(1): p. 162-172. 155. Wang, L., et al., Dynamic expression profiles of MMPs/TIMPs and collagen deposition in mechanically unloaded rat heart: implications for left ventricular assist device support-induced cardiac alterations. J Physiol Biochem, 2013: p. 1-9. 156. Laviades, C., et al., Abnormalities of the extracellular degradation of collagen type I in essential hypertension. Circulation, 1998. 98(6): p. 535-40. 157. Bradshaw, A.D., et al., Age-dependent alterations in fibrillar collagen content and myocardial diastolic function: role of SPARC in post-synthetic procollagen processing. Am J Physiol Heart Circ Physiol, 2010. 298(2): p. H614-22. 109  158. Spach, M.S. and J.P. Boineau, Microfibrosis produces electrical load variations due to loss of side-to-side cell connections: a major mechanism of structural heart disease arrhythmias. Pacing Clin Electrophysiol, 1997. 20(2 Pt 2): p. 397-413. 159. Izzo Jr, J.L. and A.H. Gradman, Mechanisms and management of hypertensive heart disease: from left ventricular hypertrophy to heart failure. Med Clin North Am, 2004. 88(5): p. 1257-1271. 160. Campbell, S.E., et al., Myocardial Fibrosis in the Rat With Mineralocorticoid Excess Prevention of Scarring by Amiloride. Am J Hypertens, 1993. 6(6 Pt 1): p. 487-495. 161. D?ez, J., et al., Serum Markers of Collagen Type I Metabolism in Spontaneously Hypertensive Rats: Relation to Myocardial Fibrosis. Circulation, 1996. 93(5): p. 1026-1032. 162. Laviades, C., G. Mayor, and J. D?ez, Treatment With Lisinopril Normalizes Serum Concentrations of Procollagen Type III Amino-Terminal Peptide in Patients With Essential Hypertension. Am J Hypertens, 1994. 7(1): p. 52-58. 163. Zervoudaki, A., et al., Plasma levels of active extracellular matrix metalloproteinases 2 and 9 in patients with essential hypertension before and after antihypertensive treatment. J Hum Hypertens, 2003. 17(2): p. 119-24. 164. Sundstr?m, J., et al., Relations of plasma total TIMP-1 levels to cardiovascular risk factors and echocardiographic measures: the Framingham heart study. Eur Heart J, 2004. 25(17): p. 1509-1516. 165. O'Hanlon, R. and D.J. Pennell, Cardiovascular magnetic resonance in the evaluation of hypertrophic and infiltrative cardiomyopathies. Heart Fail Clin. 5(3): p. 369-87. 166. Rubler, S., et al., New type of cardiomyopathy associated with diabetic glomerulosclerosis. Am J Cardiol, 1972. 30(6): p. 595-602. 110  167. Fein, F.S. and E.H. Sonnenblick, Diabetic cardiomyopathy. Cardiovasc Drugs Ther, 1994. 8(1): p. 65-73. 168. Fischer, V.W., H.B. Barner, and L.S. Larose, Pathomorphologic aspects of muscular tissue in diabetes mellitus. Hum Pathol, 1984. 15(12): p. 1127-36. 169. Shimizu, M., et al., Collagen remodelling in myocardia of patients with diabetes. J Clin Pathol, 1993. 46(1): p. 32-6. 170. Aronson, D., Cross-linking of glycated collagen in the pathogenesis of arterial and myocardial stiffening of aging and diabetes. J Hypertens, 2003. 21(1): p. 3-12. 171. Lamers, M.L., et al., High Glucose-Mediated Oxidative Stress Impairs Cell Migration. PLoS ONE, 2011. 6(8): p. e22865. 172. Trevisan, R., et al., Enhanced collagen synthesis in cultured skin fibroblasts from insulin-dependent diabetic patients with nephropathy. J Am Soc Nephrol, 1997. 8(7): p. 1133-9. 173. Tokudome, T., et al., Direct effects of high glucose and insulin on protein synthesis in cultured cardiac myocytes and DNA and collagen synthesis in cardiac fibroblasts. Metabolism, 2004. 53(6): p. 710-5. 174. Asbun, J., A.M. Manso, and F.J. Villarreal, Profibrotic influence of high glucose concentration on cardiac fibroblast functions: effects of losartan and vitamin E. Am J Physiol Heart Circ Physiol, 2005. 288(1): p. H227-34. 175. Wang, H., et al., Cytokines Regulate the Pattern of Rejection and Susceptibility to Cyclosporine Therapy in Different Mouse Recipient Strains After Cardiac Allografting. J Immunol, 2003. 171(7): p. 3823-3836. 176. Iemitsu, M., et al., Physiological and pathological cardiac hypertrophy induce different molecular phenotypes in the rat. Am J Physiol-Reg I, 2001. 281(6): p. R2029-R2036. 111  177. Blyszczuk, P., et al., Profibrotic potential of Prominin-1+ epithelial progenitor cells in pulmonary fibrosis. Resp Res, 2011. 12(1): p. 126. 178. Fairweather, D., et al., Interferon-? Protects against Chronic Viral Myocarditis by Reducing Mast Cell Degranulation, Fibrosis, and the Profibrotic Cytokines Transforming Growth Factor-?1, Interleukin-1?, and Interleukin-4 in the Heart. Am J Pathol, 2004. 165(6): p. 1883-1894. 179. Schultz, J.E.J., et al., TGF-?1 mediates the hypertrophic cardiomyocyte growth induced by angiotensin II. J Clin Invest, 2002. 109(6): p. 787-796. 180. Sadoshima, J. and S. Izumo, Molecular characterization of angiotensin II--induced hypertrophy of cardiac myocytes and hyperplasia of cardiac fibroblasts. Critical role of the AT1 receptor subtype. Circ Res, 1993. 73(3): p. 413-23. 181. Ikeda, U., et al., Angiotensin II Augments Cytokine-Stimulated Nitric Oxide Synthesis in Rat Cardiac Myocytes. Circulation, 1995. 92(9): p. 2683-2689. 182. Kawaguchi, H. and A. Kitabatake, Altered signal transduction system in hypertrophied myocardium: angiotensin II stimulates collagen synthesis in hypertrophied hearts. J Card Fail, 1996. 2(4 Suppl): p. S13-9. 183. Funck, R.C., et al., Regulation and role of myocardial collagen matrix remodeling in hypertensive heart disease. Adv Exp Med Biol, 1997. 432: p. 35-44. 184. Szalay, G., et al., Osteopontin: a fibrosis-related marker molecule in cardiac remodeling of enterovirus myocarditis in the susceptible host. Circ Res, 2009. 104(7): p. 851-9. 185. Sun, Y., et al., Angiotensin II, Transforming Growth Factor-?1and Repair in the Infarcted Heart. J Mol Cell Cardiol, 1998. 30(8): p. 1559-1569. 186. Kawano, H., et al., Angiotensin II has multiple profibrotic effects in human cardiac fibroblasts. Circulation, 2000. 101(10): p. 1130-7. 112  187. Diez, J., Profibrotic effects of angiotensin II in the heart: a matter of mediators. Hypertension, 2004. 43(6): p. 1164-5. 188. Leask, A. and D.J. Abraham, TGF-beta signaling and the fibrotic response. FASEB J, 2004. 18(7): p. 816-27. 189. Bujak, M., et al., Essential role of Smad3 in infarct healing and in the pathogenesis of cardiac remodeling. Circulation, 2007. 116(19): p. 2127-38. 190. Gold, L.I., et al., TGF-beta isoforms are differentially expressed in increasing malignant grades of HaCaT keratinocytes, suggesting separate roles in skin carcinogenesis. J Pathol, 2000. 190(5): p. 579-88. 191. Thompson, N.L., et al., Expression of transforming growth factor-beta 1 in specific cells and tissues of adult and neonatal mice. J Cell Biol, 1989. 108(2): p. 661-669. 192. Schiller, M., D. Javelaud, and A. Mauviel, TGF-?-induced SMAD signaling and gene regulation: consequences for extracellular matrix remodeling and wound healing. J Dermatol Sci, 2004. 35(2): p. 83-92. 193. Letourneur, O., et al., Ligand-Induced Dimerization of the Extracellular Domain of the TGF-? Receptor Type II. Biochem Bioph Res Co, 1996. 224(3): p. 709-716. 194. Shi, Y. and J. Massagu?, Mechanisms of TGF-? Signaling from Cell Membrane to the Nucleus. Cell, 2003. 113(6): p. 685-700. 195. Moustakas, A., S. Souchelnytskyi, and C.-H. Heldin, Smad regulation in TGF-? signal transduction. J Cell Sci, 2001. 114(24): p. 4359-4369. 196. Brand, T. and M.D. Schneider, The TGF beta superfamily in myocardium: ligands, receptors, transduction, and function. J Mol Cell Cardiol, 1995. 27(1): p. 5-18. 197. Lijnen, P.J., V.V. Petrov, and R.H. Fagard, Induction of Cardiac Fibrosis by Transforming Growth Factor-?1. Mol Genet Metab, 2000. 71(1?2): p. 418-435. 113  198. Desmouliere, A., et al., Transforming growth factor-beta 1 induces alpha-smooth muscle actin expression in granulation tissue myofibroblasts and in quiescent and growing cultured fibroblasts. J Cell Biol, 1993. 122(1): p. 103-11. 199. Petrov, V.V., R.H. Fagard, and P.J. Lijnen, Stimulation of Collagen Production by Transforming Growth Factor-?1 During Differentiation of Cardiac Fibroblasts to Myofibroblasts. Hypertension, 2002. 39(2): p. 258-263. 200. Schiller, M., D. Javelaud, and A. Mauviel, TGF-beta-induced SMAD signaling and gene regulation: consequences for extracellular matrix remodeling and wound healing. J Dermatol Sci, 2004. 35(2): p. 83-92. 201. Phan, S.H., The myofibroblast in pulmonary fibrosis. Chest, 2002. 122(6 Suppl): p. 286S-289S. 202. Petrov, V.V., R.H. Fagard, and P.J. Lijnen, Stimulation of collagen production by transforming growth factor-beta1 during differentiation of cardiac fibroblasts to myofibroblasts. Hypertension, 2002. 39(2): p. 258-63. 203. Heimer, R., et al., TGF-beta modulates the synthesis of proteoglycans by myocardial fibroblasts in culture. J Mol Cell Cardiol, 1995. 27(10): p. 2191-8. 204. Zhou, X., et al., Mechanisms of tissue inhibitor of metalloproteinase 1 augmentation by IL-13 on TGF-?1?stimulated primary human fibroblasts. J Allergy Clin Immun, 2007. 119(6): p. 1388-1397. 205. Eghbali, M., et al., Differential effects of transforming growth factor-beta 1 and phorbol myristate acetate on cardiac fibroblasts. Regulation of fibrillar collagen mRNAs and expression of early transcription factors. Circ Res, 1991. 69(2): p. 483-90. 206. Chua, C.C., et al., Effect of growth factors on collagen metabolism in cultured human heart fibroblasts. Connect Tissue Res, 1991. 26(4): p. 271-81. 114  207. Cucoranu, I., et al., NAD(P)H Oxidase 4 Mediates Transforming Growth Factor-?1?Induced Differentiation of Cardiac Fibroblasts Into Myofibroblasts. Circ Res, 2005. 97(9): p. 900-907. 208. Moustakas, A. and C.H. Heldin, The regulation of TGFbeta signal transduction. Development, 2009. 136(22): p. 3699-714. 209. Ignotz, R.A. and J. Massagu?, Transforming growth factor-beta stimulates the expression of fibronectin and collagen and their incorporation into the extracellular matrix. J Biol Chem, 1986. 261(9): p. 4337-4345. 210. Iglesias-de la Cruz, M.C., et al., Hydrogen peroxide increases extracellular matrix mRNA through TGF-beta in human mesangial cells. Kidney Int, 2001. 59(1): p. 87-95. 211. Derynck, R. and Y.E. Zhang, Smad-dependent and Smad-independent pathways in TGF-beta family signalling. Nature, 2003. 425(6958): p. 577-84. 212. Rosenkranz, S., et al., Alterations of beta-adrenergic signaling and cardiac hypertrophy in transgenic mice overexpressing TGF-beta(1). Am J Physiol Heart Circ Physiol, 2002. 283(3): p. H1253-62. 213. Laviades, C., N. Varo, and J. Diez, Transforming growth factor beta in hypertensives with cardiorenal damage. Hypertension, 2000. 36(4): p. 517-22. 214. Siddigi, N.J., Alhomida, A.S., Investigation into the distribution of total, free, peptide-bound, protein-bound, soluble- and insoluble-collagen hydroxyproline in various bovine tissues. J Biochem Mol Biol, 2003. 36(2): p. 154-8. 215. Souders, C.A., S.L. Bowers, and T.A. Baudino, Cardiac fibroblast: the renaissance cell. Circ Res, 2009. 105(12): p. 1164-76. 115  216. Engel, J. and D.J. Prockop, The Zipper-Like Folding of Collagen Triple Helices and the Effects of Mutations that Disrupt the Zipper. Annu Rev Biophys Bio, 1991. 20(1): p. 137-152. 217. Prockop, D.J. and K.I. Kivirikko, Collagens: molecular biology, diseases, and potentials for therapy. Annu Rev Biochem, 1995. 64: p. 403-34. 218. Prockop, D.J., A.L. Sieron, and S.-W. Li, Procollagen N-proteinase and procollagen C-proteinase. Two unusual metalloproteinases that are essential for procollagen processing probably have important roles in development and cell signaling. Matrix Biol, 1998. 16(7): p. 399-408. 219. Reiser, K., R.J. McCormick, and R.B. Rucker, Enzymatic and nonenzymatic cross-linking of collagen and elastin. FASEB J, 1992. 6(7): p. 2439-49. 220. Velling, T., et al., Polymerization of Type I and III Collagens Is Dependent On Fibronectin and Enhanced By Integrins ?11?1and ?2?1. J Biol Chem, 2002. 277(40): p. 37377-37381. 221. Smith-Mungo, L.I. and H.M. Kagan, Lysyl oxidase: properties, regulation and multiple functions in biology. Matrix Biol, 1998. 16(7): p. 387-98. 222. Kagan, H.M. and P.C. Trackman, Properties and function of lysyl oxidase. Am J Respir Cell Mol Biol, 1991. 5(3): p. 206-10. 223. Boak, A.M., et al., Regulation of lysyl oxidase expression in lung fibroblasts by transforming growth factor-beta 1 and prostaglandin E2. Am J Respir Cell Mol Biol, 1994. 11(6): p. 751-5. 224. Roy, R., et al., Regulation of lysyl oxidase and cyclooxygenase expression in human lung fibroblasts: interactions among TGF-beta, IL-1 beta, and prostaglandin E. J Cell Biochem, 1996. 62(3): p. 411-7. 116  225. Adam, O., et al., Increased lysyl oxidase expression and collagen cross-linking during atrial fibrillation. J Mol Cell Cardiol, 2011. 50(4): p. 678-85. 226. Fogelgren, B., et al., Cellular Fibronectin Binds to Lysyl Oxidase with High Affinity and Is Critical for Its Proteolytic Activation. J Biol Chem, 2005. 280(26): p. 24690-24697. 227. Majora, M., et al., Functional Consequences of Mitochondrial DNA Deletions in Human Skin Fibroblasts: Increased Contractile Strength in Collagen Lattices Is Due to Oxidative Stress-Induced Lysyl Oxidase Activity. Am J Pathol, 2009. 175(3): p. 1019-1029. 228. Hermida, N., et al., A synthetic peptide from transforming growth factor-?1 type III receptor prevents myocardial fibrosis in spontaneously hypertensive rats. Cardiovasc Res, 2009. 81(3): p. 601-609. 229. L?pez, B., et al., Impact of Treatment on Myocardial Lysyl Oxidase Expression and Collagen Cross-Linking in Patients With Heart Failure. Hypertension, 2009. 53(2): p. 236-242. 230. Rustan, A.C., M.S. Nenseter, and C.A. Drevon, Omega-3 and omega-6 fatty acids in the insulin resistance syndrome. Lipid and lipoprotein metabolism and atherosclerosis. Ann N Y Acad Sci, 1997. 827: p. 310-26. 231. Ruxton, C.H.S., et al., The health benefits of omega-3 polyunsaturated fatty acids: a review of the evidence. J Hum Nutr Diet, 2004. 17(5): p. 449-459. 232. Newmark, H.L., Squalene, olive oil, and cancer risk: a review and hypothesis. Cancer Epidem Biomar, 1997. 6(12): p. 1101-1103. 233. Connor, W.E., ?-Linolenic acid in health and disease. Am J Clin Nutr, 1999. 69(5): p. 827-828. 234. Catala, A., A synopsis of the process of lipid peroxidation since the discovery of the essential fatty acids. Biochem Biophys Res Commun, 2010. 399(3): p. 318-23. 117  235. Prasad, A., M.S. Bloom, and D.O. Carpenter, Role of calcium and ROS in cell death induced by polyunsaturated fatty acids in murine thymocytes. J Cell Physiol, 2010. 225(3): p. 829-36. 236. Schmitz, G. and J. Ecker, The opposing effects of n-3 and n-6 fatty acids. Prog Lipid Res, 2008. 47(2): p. 147-55. 237. Czernichow, S., D. Thomas, and E. Bruckert, n-6 Fatty acids and cardiovascular health: a review of the evidence for dietary intake recommendations. Br J Nutr, 2010. 104(6): p. 788-96. 238. Lau, D.C., et al., 2006 Canadian clinical practice guidelines on the management and prevention of obesity in adults and children [summary]. Can Med Assoc J, 2007. 176(8): p. S1-13. 239. HealthCanada, Do Canadian Adults meet their nutrient requirements through food intake alone?, 2009, Health Canada: Ottawa. p. H164-112/3-2009E-PDF. 240. Madden, S.M., C.F. Garrioch, and B.J. Holub, Direct diet quantification indicates low intakes of (n-3) fatty acids in children 4 to 8 years old. J Nutr, 2009. 139(3): p. 528-32. 241. Simopoulos, A.P., Omega-3 Fatty Acids in Inflammation and Autoimmune Diseases. J Am Coll Nutr, 2002. 21(6): p. 495-505. 242. Shaikh, S.R. and M. Edidin, Polyunsaturated fatty acids, membrane organization, T cells, and antigen presentation. Am J Clin Nutr, 2006. 84(6): p. 1277-1289. 243. Tapiero, H., et al., Polyunsaturated fatty acids (PUFA) and eicosanoids in human health and pathologies. Biomed Pharmacother, 2002. 56(5): p. 215-222. 244. Calder, P.C., n?3 Polyunsaturated fatty acids, inflammation, and inflammatory diseases. Am J Clin Nutr, 2006. 83(6): p. S1505-1519S. 118  245. Safarinejad, M.R., et al., Relationship of omega-3 and omega-6 fatty acids with semen characteristics, and anti-oxidant status of seminal plasma: a comparison between fertile and infertile men. Clin Nutr, 2010. 29(1): p. 100-5. 246. Spiteller, G., The relation of lipid peroxidation processes with atherogenesis: a new theory on atherogenesis. Mol Nutr Food Res, 2005. 49(11): p. 999-1013. 247. Simopoulos, A.P., The importance of the ratio of omega-6/omega-3 essential fatty acids. Biomed Pharmacother, 2002. 56(8): p. 365-379. 248. Simopoulos, A.P., Essential fatty acids in health and chronic disease. Am J Clin Nutr, 1999. 70(3 Suppl): p. 560S-569S. 249. Gao, D., et al., Palmitate promotes monocyte atherogenicity via de novo ceramide synthesis. Free Radic Biol Med, 2012. Epub ahead of print. 250. Miller, T.A., et al., Oleate prevents palmitate-induced cytotoxic stress in cardiac myocytes. Biochem Bioph Res Co, 2005. 336(1): p. 309-315. 251. Trichopoulou, A., et al., Adherence to a Mediterranean diet and survival in a Greek population. N Engl J Med, 2003. 348(26): p. 2599-2608. 252. Keys, A., et al., Epidemiological studies related to coronary heart disease: characteristics of men aged 40-59 in seven countries. Acta Med Scand Suppl, 1966. 460: p. 1-392. 253. Estruch, R., et al., Effects of a Mediterranean-style diet on cardiovascular risk factors: a randomized trial. Ann Intern Med, 2006. 145(1): p. 1-11. 254. Teres, S., et al., Oleic acid content is responsible for the reduction in blood pressure induced by olive oil. Proc Natl Acad Sci U S A, 2008. 105(37): p. 13811-13816. 255. Mente, A., et al., A systematic review of the evidence supporting a causal link between dietary factors and coronary heart disease. Arch Intern Med, 2009. 169(7): p. 659-69. 119  256. Micha, R. and D. Mozaffarian, Saturated fat and cardiometabolic risk factors, coronary heart disease, stroke, and diabetes: a fresh look at the evidence. Lipids, 2010. 45(10): p. 893-905. 257. Lands, W.E., Dietary fat and health: the evidence and the politics of prevention: careful use of dietary fats can improve life and prevent disease. Ann N Y Acad Sci, 2005. 1055: p. 179-92. 258. Ramsden, C.E., et al., n-6 Fatty acid-specific and mixed polyunsaturate dietary interventions have different effects on CHD risk: a meta-analysis of randomised controlled trials. Brit J Nutr, 2010. 104(11): p. 1586-1600. 259. Gordon, D.J., Lowering cholesterol and total mortality. In: Rifkin BM, ed. Lowering Cholesterol in High-Risk Individuals and Populations, 1995, Marcel Dekker, Inc: New York, NY. p. 33-48. 260. Rudel, L.L., et al., Dietary polyunsaturated fat modifies low-density lipoproteins and reduces atherosclerosis of nonhuman primates with high and low diet responsiveness. Am J Clin Nutr, 1995. 62(2): p. 463S-470S. 261. McLennan, P.L., Relative effects of dietary saturated, monounsaturated, and polyunsaturated fatty acids on cardiac arrhythmias in rats. Am J Clin Nutr, 1993. 57(2): p. 207-12. 262. Hegsted, D.M., et al., Dietary fat and serum lipids: an evaluation of the experimental data. Am J Clin Nutr, 1993. 57(6): p. 875-83. 263. Jakobsen, M.U., et al., Major types of dietary fat and risk of coronary heart disease: a pooled analysis of 11 cohort studies. Am J Clin Nutr, 2009. 89(5): p. 1425-32. 120  264. Summers, L.K.M., et al., Substituting dietary saturated fat with polyunsaturated fat changes abdominal fat distribution and improves insulin sensitivity. Diabetologia, 2002. 45(3): p. 369-377. 265. Salmeron, J., et al., Dietary fat intake and risk of type 2 diabetes in women. Am J Clin Nutr, 2001. 73(6): p. 1019-26. 266. Mozaffarian, D. and R. Clarke, Quantitative effects on cardiovascular risk factors and coronary heart disease risk of replacing partially hydrogenated vegetable oils with other fats and oils. Eur J Clin Nutr, 2009. 63: p. S22-S33. 267. Ravnskov, U., The questionable role of saturated and polyunsaturated fatty acids in cardiovascular disease. J Clin Epidemiol, 1998. 51(6): p. 443-60. 268. Rose, G.A., W.B. Thomson, and R.T. Williams, Corn Oil in Treatment of Ischaemic Heart Disease. Br Med J, 1965. 1(5449): p. 1531-3. 269. Jeckel, K.M., et al., The role of dietary fatty acids in predicting myocardial structure in fat-fed rats. Lipids Health Dis, 2011. 10: p. 92. 270. Yam, D., A. Eliraz, and E.M. Berry, Diet and disease--the Israeli paradox: possible dangers of a high omega-6 polyunsaturated fatty acid diet. Isr J Med Sci, 1996. 32(11): p. 1134-43. 271. Pearce, M.L. and S. Dayton, Incidence of cancer in men on a diet high in polyunsaturated fat. Lancet, 1971. 1(7697): p. 464-7. 272. Harris, W., Omega-6 and omega-3 fatty acids: partners in prevention. Curr Opin Clin Nutr Metab Care, 2010. 13(2): p. 125-9. 273. Thomas, D.W., et al., Proinflammatory actions of thromboxane receptors to enhance cellular immune responses. J Immunol, 2003. 171(12): p. 6389-95. 121  274. Hoff, U., et al., Inhibition of 20-HETE synthesis and action protects the kidney from ischemia/reperfusion injury. Kidney Int, 2011. 79(1): p. 57-65. 275. Biasi, F., C. Mascia, and G. Poli, The contribution of animal fat oxidation products to colon carcinogenesis, through modulation of TGF-?1 signaling. Carcinogenesis, 2008. 29(5): p. 890-894. 276. Siri-Tarino, P.W., et al., Meta-analysis of prospective cohort studies evaluating the association of saturated fat with cardiovascular disease. The American Journal of Clinical Nutrition, 2010. 91(3): p. 535-546. 277. Ramsden, C.E., et al., Use of dietary linoleic acid for secondary prevention of coronary heart disease and death: evaluation of recovered data from the Sydney Diet Heart Study and updated meta-analysis. BMJ, 2013. 346. 278. Katan, M.B., P.L. Zock, and R.P. Mensink, Dietary oils, serum lipoproteins, and coronary heart disease. The American Journal of Clinical Nutrition, 1995. 61(6): p. 1368S-1373S. 279. Woodhill, J.M., et al., Low Fat, Low Cholesterol Diet in Secondary Prevention of Coronary Heart Disease, in Drugs, Lipid Metabolism, and Atherosclerosis, D. Kritchevsky, R. Paoletti, and W. Holmes, Editors. 1978, Springer US. p. 317-330. 280. Frantz, I.D., et al., Test of effect of lipid lowering by diet on cardiovascular risk. The Minnesota Coronary Survey. Arterioscler Thromb Vac Biol, 1989. 9(1): p. 129-35. 281. Ghosh, S., et al., Induction of mitochondrial nitrative damage and cardiac dysfunction by chronic provision of dietary omega-6 polyunsaturated fatty acids. Free Radic Biol Med, 2006. 41(9): p. 1413-24. 282. Ramsden, C.E., et al., Lowering dietary linoleic acid reduces bioactive oxidized linoleic acid metabolites in humans. Prostag Leukotr Ess, 2012. 87(4?5): p. 135-141. 122  283. Zhang, H.-M., et al., Linoleic Acid-Induced Mitochondrial Ca(2+) Efflux Causes Peroxynitrite Generation and Protein Nitrotyrosylation. PLoS ONE, 2009. 4(6): p. e6048. 284. Kimes, B.W. and B.L. Brandt, Properties of a clonal muscle cell line from rat heart. Exp Cell Res, 1976. 98(2): p. 367-381. 285. Menard, C., et al., Modulation of L-type calcium channel expression during retinoic acid-induced differentiation of H9C2 cardiac cells. The Journal of biological chemistry, 1999. 274(41): p. 29063-70. 286. Todaro, G.J. and H. Green, QUANTITATIVE STUDIES OF THE GROWTH OF MOUSE EMBRYO CELLS IN CULTURE AND THEIR DEVELOPMENT INTO ESTABLISHED LINES. The Journal of Cell Biology, 1963. 17(2): p. 299-313. 287. M?nard, C., et al., Modulation of L-type Calcium Channel Expression during Retinoic Acid-induced Differentiation of H9C2 Cardiac Cells. J Biol Chem, 1999. 274(41): p. 29063-29070. 288. Saeedi, R., et al., AMP-activated protein kinase influences metabolic remodeling in H9c2 cells hypertrophied by arginine vasopressin. Am J Physiol-Heart C, 2009. 296(6): p. H1822-H1832. 289. Zhang, H., et al., Glutathione-dependent reductive stress triggers mitochondrial oxidation and cytotoxicity. FASEB J, 2012. 26(4): p. 1442-1451. 290. Mosmann, T., Rapid colorimetric assay for cellular growth and survival: Application to proliferation and cytotoxicity assays. J Immunol Methods, 1983. 65(1?2): p. 55-63. 291. Page, B., M. Page, and C. Noel, A new fluorometric assay for cytotoxicity measurements in-vitro. Int J Oncol, 1993. 3(3): p. 473-6. 123  292. Matsumoto, K., et al., Fluorometric determination of carnitine in serum with immobilized carnitine dehydrogenase and diaphorase. Clin Chem, 1990. 36(12): p. 2072-6. 293. Cathcart, R., E. Schwiers, and B.N. Ames, Detection of picomole levels of hydroperoxides using a fluorescent dichlorofluorescein assay. Anal Biochem, 1983. 134(1): p. 111-116. 294. LeBel, C.P., H. Ischiropoulos, and S.C. Bondy, Evaluation of the probe 2',7'-dichlorofluorescin as an indicator of reactive oxygen species formation and oxidative stress. Chem Res Toxicol, 1992. 5(2): p. 227-231. 295. Sternson, L.A., J.F. Stobaugh, and A.J. Repta, Rational design and evaluation of improved o-phthalaldehyde-like fluorogenic reagents. Anal Biochem, 1985. 144(1): p. 233-246. 296. Senft, A.P., T.P. Dalton, and H.G. Shertzer, Determining Glutathione and Glutathione Disulfide Using the Fluorescence Probe o-Phthalaldehyde. Anal Biochem, 2000. 280(1): p. 80-86. 297. Tullberg-Reinert, H. and G. Jundt, In situ measurement of collagen synthesis by human bone cells with a sirius red-based colorimetric microassay: effects of transforming growth factor beta2 and ascorbic acid 2-phosphate. Histochem Cell Biol, 1999. 112(4): p. 271-6. 298. Purnomo, Y., et al., Role of reactive oxygen species in the transforming growth factor-beta1-induced collagen production and differentiation of cardiac fibroblasts into myofibroblasts. Oxid Antioxid Med Sci, 2013. 2(1): p. 5-10. 299. Lareu, R.R., et al., Essential modification of the Sircol Collagen Assay for the accurate quantification of collagen content in complex protein solutions. Acta Biomater, 2010. 6(8): p. 3146-3151. 124  300. Bustin, S.A., et al., The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments. Clin Chem, 2009. 55(4): p. 611-622. 301. Ramakers, C., et al., Assumption-free analysis of quantitative real-time polymerase chain reaction (PCR) data. Neurosci Lett, 2003. 339(1): p. 62-6. 302. Ruijter, J.M., et al., Amplification efficiency: linking baseline and bias in the analysis of quantitative PCR data. Nucleic Acids Res, 2009. 37(6): p. e45. 303. Ukang, S., S. Ampawong, and K. Kengkoom, Collagen Measurement and Staining Pattern of Wound Healing Comparison with Fixations and Stains. J Microsc Soc Thailand, 2008. 22: p. 37-41. 304. Pauschinger, M., et al., Dilated Cardiomyopathy Is Associated With Significant Changes in Collagen Type I/III ratio. Circulation, 1999. 99(21): p. 2750-2756. 305. Trachootham, D., et al., Redox regulation of cell survival. Antioxid Redox Signal, 2008. 10(8): p. 1343-74. 306. Khanna, S., et al., Characterization of the potent neuroprotective properties of the natural vitamin E alpha-tocotrienol. J Neurochem, 2006. 98(5): p. 1474-86. 307. Yam, D., A. Eliraz, and E.M. Berry, Diet and disease--the Israeli paradox: possible dangers of a high omega-6 polyunsaturated fatty acid diet. Isr J Med Sci, 1996. 32(11): p. 1134-43. 308. Kajstura, J., et al., Apoptotic and necrotic myocyte cell deaths are independent contributing variables of infarct size in rats. Lab Invest, 1996. 74(1): p. 86-107. 309. Frustaci, A., et al., Myocardial Cell Death in Human Diabetes. Circ Res, 2000. 87(12): p. 1123-1132. 310. Kris-Etherton, P.M. and f.t.N. Committee, Monounsaturated Fatty Acids and Risk of Cardiovascular Disease. Circulation, 1999. 100(11): p. 1253-1258. 125  311. de Vries, J.E., et al., Saturated but not mono-unsaturated fatty acids induce apoptotic cell death in neonatal rat ventricular myocytes. J Lipid Res, 1997. 38(7): p. 1384-94. 312. Rabkin, S.W. and P. Lodha, Stearic acid-induced cardiac lipotoxicity is independent of cellular lipid and is mitigated by the fatty acids oleic and capric acid but not by the PPAR agonist troglitazone. Exp Physiol, 2009. 94(8): p. 877-887. 313. Hill, B.G., et al., Importance of the bioenergetic reserve capacity in response to cardiomyocyte stress induced by 4-hydroxynonenal. Biochem J, 2009. 424(1): p. 99-107. 314. Spiteller, P., et al., Aldehydic lipid peroxidation products derived from linoleic acid. BBA-Mol Cell Biol L, 2001. 1531(3): p. 188-208. 315. Diestel, A., et al., Hypothermia protects H9c2 cardiomyocytes from H2O2 induced apoptosis. Cryobiology, 2011. 62(1): p. 53-61. 316. Chen, Q.M., et al., Hydrogen Peroxide Dose Dependent Induction of Cell Death or Hypertrophy in Cardiomyocytes. Arch Biochem Biophys, 2000. 373(1): p. 242-248. 317. Oyama, K., K. Takahashi, and K. Sakurai, Cardiomyocyte H9c2 Cells Exhibit Differential Sensitivity to Intracellular Reactive Oxygen Species Generation with Regard to Their Hypertrophic vs Death Responses to Exogenously Added Hydrogen Peroxide. J Clin Biochem Nutr, 2009. 45(3): p. 361-369. 318. Zhang, H., et al., Excess copper induces accumulation of hydrogen peroxide and increases lipid peroxidation and total activity of copper?zinc superoxide dismutase in roots of Elsholtzia haichowensis. Planta, 2008. 227(2): p. 465-475. 319. Keidar, S., et al., The Angiotensin-II Receptor Antagonist, Losartan, Inhibits LDL Lipid Peroxidation and Atherosclerosis in Apolipoprotein E-Deficient Mice. Biochem Bioph Res Co, 1997. 236(3): p. 622-625. 126  320. Rumley, A.G., et al., Plasma lipid peroxides: relationships to cardiovascular risk factors and prevalent cardiovascular disease. QJM-Mon J Assoc Phys, 2004. 97(12): p. 809-816. 321. Tzeng, W.-F., J.-L. Lee, and T.-J. Chiou, The role of lipid peroxidation in menadione-mediated toxicity in cardiomyocytes. J Mol Cell Cardiol, 1995. 27(9): p. 1999-2008. 322. Zhang, X., et al., Differential vulnerability to oxidative stress in rat cardiac myocytes versus fibroblasts. J Am Coll Cardiol, 2001. 38(7): p. 2055-2062. 323. Magdalon, J., et al., A proteomic analysis of the functional effects of fatty acids in NIH 3T3 fibroblasts. Lipids Health Dis, 2011. 10: p. 218. 324. Watkins, B.A., H. Xu, and J.J. Turek, Linoleate Impairs Collagen Synthesis in Primary Cultures of Avian Chondrocytes. Exp Biol Med, 1996. 212(2): p. 153-159. 325. Hatanaka, E., et al., Oleic, Linoleic and Linolenic Acids Increase ROS Production by Fibroblasts via NADPH Oxidase Activation. PLoS ONE, 2013. 8(4): p. e58626. 326. Wei, C.D., et al., Palmitate induces H9c2 cell apoptosis by increasing reactive oxygen species generation and activation of the ERK1/2 signaling pathway. Mol Med Rep, 2013. 7(3): p. 855-61. 327. Zalba, G., et al., Vascular NADH/NADPH Oxidase Is Involved in Enhanced Superoxide Production in Spontaneously Hypertensive Rats. Hypertension, 2000. 35(5): p. 1055-1061. 328. Buffon, A., et al., Large, sustained cardiac lipid peroxidation and reduced antioxidant capacity in the coronary circulation after brief episodes of myocardial ischemia. J Am Coll Cardiol, 2000. 35(3): p. 633-639. 329. Michiels, C. and J. Remacle, Cytotoxicity of linoleic acid peroxide, malondialdehyde and 4-hydroxynonenal towards human fibroblasts. Toxicology, 1991. 66(2): p. 225-34. 127  330. Toborek, M., et al., Linoleic acid potentiates TNF-mediated oxidative stress, disruption of calcium homeostasis, and apoptosis of cultured vascular endothelial cells. J Lipid Res, 1997. 38(10): p. 2155-67. 331. Vaziri, N.D., et al., Induction of Oxidative Stress by Glutathione Depletion Causes Severe Hypertension in Normal Rats. Hypertension, 2000. 36(1): p. 142-146. 332. Comporti, M., Glutathione depleting agents and lipid peroxidation. Chem Phys Lipids, 1987. 45(2-4): p. 143-69. 333. Watanabe, Y., et al., Chronic depletion of glutathione exacerbates ventricular remodelling and dysfunction in the pressure-overloaded heart. Cardiovasc Res, 2012. 334. Ghosh, S., et al., Cardiomyocyte apoptosis induced by short-term diabetes requires mitochondrial GSH depletion. Am J Physiol Heart Circ Physiol, 2005. 289(2): p. H768-76. 335. Monboisse, J.C., et al., Non-enzymatic degradation of acid-soluble calf skin collagen by superoxide ion: Protective effect of flavonoids. Biochem Pharmacol, 1983. 32(1): p. 53-58. 336. Curran, S.F., et al., Degradation of soluble collagen by ozone or hydroxyl radicals. FEBS Lett, 1984. 176(1): p. 155-160. 337. Lijnen, P.J., et al., Modulation of reactive oxygen species and collagen synthesis by angiotensin II in cardiac fibroblasts. Open Hypertens J, 2011. 4: p. 1-17. 338. Shanley, C.J., et al., Transforming growth factor-beta 1 increases lysyl oxidase enzyme activity and mRNA in rat aortic smooth muscle cells. J Vasc Surg, 1997. 25(3): p. 446-52. 339. Choung, J., et al., Role of EP2 receptors and cAMP in prostaglandin E2 regulated expression of type I collagen ?1, lysyl oxidase, and cyclooxygenase-1 genes in human embryo lung fibroblasts. J Cell Biochem, 1998. 71(2): p. 254-263. 128  340. Kumarasamy, A., et al., Lysyl Oxidase Activity Is Dysregulated during Impaired Alveolarization of Mouse and Human Lungs. Am J Resp Crit Care, 2009. 180(12): p. 1239-1252. 341. Voloshenyuk, T.G., et al., TNF-? increases cardiac fibroblast lysyl oxidase expression through TGF-? and PI3Kinase signaling pathways. Biochem Bioph Res Co, 2011. 413(2): p. 370-375. 342. Chen, J., et al., Omega-3 Fatty Acids Prevent Pressure Overload?Induced Cardiac Fibrosis Through Activation of Cyclic GMP/Protein Kinase G Signaling in Cardiac Fibroblasts. Circulation, 2011. 123(6): p. 584-593. 343. Overall, C.M., Repression of tissue inhibitor of matrix metalloproteinase expression by all-trans-retinoic acid in rat bone cell populations: comparison with transforming growth factor-beta 1. J Cell Physiol, 1995. 164(1): p. 17-25. 344. Eickelberg, O., et al., Extracellular matrix deposition by primary human lung fibroblasts in response to TGF-?1 and TGF-?3. Am J Physiol-Lung C, 1999. 276(5): p. L814-L824. 345. Hiraoka, K., et al., Effects of Lipid Peroxide on Production of Matrix Metalloproteinase 1 (Tissue Collagenase) and 3 (Stromelysin) and Tissue Inhibitor Metalloproteinase 1 by Human Rheumatoid Synovial Fibroblasts. Exp Mol Pathol, 1993. 59(3): p. 169-176. 346. Cardoso, C.R., et al., Oleic acid modulation of the immune response in wound healing: A new approach for skin repair. Immunobiology, 2011. 216(3): p. 409-415. 347. Manrique, C., et al., Obesity and insulin resistance induce early development of diastolic dysfunction in young female mice fed a western diet. Endocrinology, 2013. 348. Marijianowski, M.M.H., et al., The neonatal heart has a relatively high content of total collagen and type I collagen, a condition that may explain the less compliant state. J Am Coll Cardiol, 1994. 23(5): p. 1204-1208. 129  349. Shah, M., D.M. Foreman, and M.W. Ferguson, Neutralising antibody to TGF-beta 1,2 reduces cutaneous scarring in adult rodents. J Cell Sci, 1994. 107(5): p. 1137-1157. 350. Shah, M., D.M. Foreman, and M.W. Ferguson, Neutralisation of TGF-beta 1 and TGF-beta 2 or exogenous addition of TGF-beta 3 to cutaneous rat wounds reduces scarring. J Cell Sci, 1995. 108(3): p. 985-1002. 351. Zhang, Y., et al., rAAV2-TGF-B(3) Decreases Collagen Synthesis and Deposition in the Liver of Experimental Hepatic Fibrosis Rat. Digest Dis Sci, 2010. 55(10): p. 2821-2830. 352. Hosokawa, R., et al., TGF-?3 Decreases Type I Collagen and Scarring after Labioplasty. J Dent Res, 2003. 82(7): p. 558-564. 353. Kapoun, A.M., et al., B-Type Natriuretic Peptide Exerts Broad Functional Opposition to Transforming Growth Factor-? in Primary Human Cardiac Fibroblasts: Fibrosis, Myofibroblast Conversion, Proliferation, and Inflammation. Circ Res, 2004. 94(4): p. 453-461. 354. Beaudry, V.G., et al., Loss of the Desmosomal Component Perp Impairs Wound Healing In Vivo. Dermatology Res, 2010. 2010. 355. Beadle, J.B., et al., Composition of corn oil. J Am Oil Chem Soc, 1965. 42(2): p. 90-95. 356. Waterman, E. and B. Lockwood, Active components and clinical applications of olive oil. Altern Med Rev, 2007. 12(4): p. 331-42. 357. Zhang, G.-X., et al., Cardiac oxidative stress in acute and chronic isoproterenol-infused rats. Cardiovasc Res, 2005. 65(1): p. 230-238. 358. Rich, L., Whittaker, P., Collagen and Picrosirius Red staining: a polarized light assessment of fibrillar hue and spatial distribution. Braz J Morphol Sci, 2005. 22: p. 97-104. 130  359. Neumann, S., et al., Aldosterone and d-Glucose Stimulate the Proliferation of Human Cardiac Myofibroblasts In Vitro. Hypertension, 2002. 39(3): p. 756-760.  131  Appendix A: In Vivo Dietary Components  Table 2. Components of in vivo diets   Formula g / kgCalcium Carbonate 3.6Casein 240Cellulose 50Corn Starch 75DL-Methionine 3.6Maltodextrin 75Mineral Mix AIN-76 (170915) 42Soybean Oil 10Sucrose 298.7Vitamin Mix (Teklad 40060) 12Oil (CO, OO) 190

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
http://iiif.library.ubc.ca/presentation/dsp.24.1-0074320/manifest

Comment

Related Items