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Cardiac lipoprotein lipase function following diabetes is regulated by cardiomyocytes and endothelial… Wang, Ying 2014

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CARDIAC LIPOPROTEIN LIPASE FUNCTION FOLLOWING DIABETES IS REGULATED BY CARDIOMYOCYTES AND ENDOTHELIAL CELLS   by Ying Wang M.Sc., Wuhan University, China, 2008   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY  in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pharmaceutical Sciences)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  March 2014 ? Ying Wang, 2014 ii  Abstract In diabetes, when glucose consumption is restricted, the heart adapts to use fatty acid (FA) exclusively.  The majority of FA provided to the heart comes from breakdown of circulating triglyceride, a process catalyzed by lipoprotein lipase (LPL) located at the vascular lumen.  Transfer of LPL from cardiomyocytes to the coronary lumen requires liberation of LPL from the myocyte surface heparan sulfate proteoglycans (HSPGs) with subsequent replenishment of this reservoir.  We examined the contribution of coronary endothelial cells (EC) and cardiomyocytes towards regulation of LPL function following diabetes.  To induce acute hyperglycemia, diazoxide (DZ), a selective ATP-sensitive K+ channel opener was used.  For chronic diabetes, streptozotocin (STZ), a ?-cell specific toxin was administered at doses of 55 (D55) or 100 (D100) mg/kg to generate moderate and severe diabetes, respectively.  Cardiac LPL processing into active dimers and breakdown at the vascular lumen was investigated.  Following acute hyperglycemia and moderate diabetes, more LPL is processed into an active dimeric form, which involves the endoplasmic reticulum chaperone calnexin in cardiomyocytes.  Severe diabetes results in increased conversion of LPL into inactive monomers at the vascular lumen, a process mediated by FA-induced expression of angiopoietin-like protein 4.  On exposure of bovine coronary artery EC to high glucose, both latent and active heparanase were released into the medium, termed ECCM.  ECCM liberated LPL from the myocyte surface, in addition to facilitating its replenishment.  Of the two forms of heparanase secreted from EC in response to high glucose, active heparanase released LPL from the myocyte surface, whereas latent heparanase stimulated reloading of LPL from an intracellular pool via HSPG-mediated RhoA activation.  Latent heparanase can be also taken up by cardiomyocytes, converted into active heparanase in lysosomes, and its nuclear entry iii  likely to modulate gene expression.  Results from this study advance our understanding of how the cross-talk between EC and cardiomyocytes facilitate LPL secretion and how diabetes influences coronary LPL maturation and turnover.  Pharmaceutical manipulation of these pathways could potentially provide an additional strategy to limit FA delivery to the heart, and prevent cardiomyopathy seen with chronic diabetes.   iv  Preface All of the work presented in Chapter 2, 3 and 4 was conducted in Faculty of Pharmaceutical Sciences at the University of British Columbia.  Parts of these chapters have been published in the following manuscripts: 1. [Wang Y, Puthanveetil P, Wang F, Kim MS, Abrahani A, and Rodrigues B.  Severity of diabetes governs vascular LPL by affecting enzyme dimerization and disassembly. Diabetes. 2011.]  I was the first author responsible for generation of the ideal, and collecting and analyzing most of data.  My supervisor Brian Rodrigues and I designed the experiment and wrote the manuscript.  Puthanveetil P (Figure 7), Wang F (Figure 9D), Kim MS (Figure 1), and Abrahani A (Figure 3C) helped me in obtaining some of the data.  2. [Wang Y, Zhang DH, Chiu AP, Wan A, Neumaier K, Vlodavsky I and Rodrigues B. Endothelial heparanase regulates heart metabolism by stimulating lipoprotein lipase secretion from cardiomyocytes. Arterioscler Thromb Vasc Biol. 2013.]  I conceived and designed the experiments, obtained most of the data, analyzed and interpreted the results, as well as wrote the major part of manuscript.  Zhang DH (Figure 16), Chiu AP (Figure 18), Wan A (Figure 18), and Neumaier K (Figure 21C) contributed to acquiring some of the data.  Vlodavsky I assisted with valuable suggestions and the preparation of highly purified latent and active heparanase.  Brian Rodrigues helped with writing the manuscript. The investigation conforms to the guide for the care and use of laboratory animals published by the US National Institutes of Health and the University of British Columbia, and was approved by the Animal Care Committee in the University of British Columbia (Certificate No. A13-0098). v  Table of Contents Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iv Table of Contents ...........................................................................................................................v List of Tables ................................................................................................................................ ix List of Figures .................................................................................................................................x List of Abbreviations .................................................................................................................. xii Acknowledgements .................................................................................................................... xiii Dedication ................................................................................................................................... xiv Chapter 1: Introduction ................................................................................................................1 1.1 Diabetic cardiomyopathy ................................................................................................ 1 1.2 Lipoprotein lipase ........................................................................................................... 3 1.2.1 Gene transcription ....................................................................................................... 3 1.2.2 Maturation inside the endoplasmic reticulum ............................................................. 5 1.2.3 Secretion to cardiomyocyte surface ............................................................................ 6 1.2.4 Release from the cardiomyocyte surface .................................................................... 7 1.2.5 Translocation from cardiomyocyte towards the apical side of EC ............................. 9 1.2.6 Turnover at the vascular lumen................................................................................. 10 1.3 Hypothesis and research objectives .............................................................................. 12 Chapter 2: Methods .....................................................................................................................15 2.1 Materials ....................................................................................................................... 15 2.2 Experimental animals.................................................................................................... 16 2.3 Animal treatment .......................................................................................................... 16 vi  2.4 Isolated heart perfusion ................................................................................................. 17 2.5 Isolation of cardiac myocytes ....................................................................................... 17 2.6 Endothelial cell culture ................................................................................................. 18 2.7 Endothelial cell conditioned medium (ECCM) ............................................................ 18 2.8 In vitro treatments ......................................................................................................... 18 2.9 LPL activity .................................................................................................................. 21 2.10 Heparin-sepharose chromatography ............................................................................. 21 2.11 Real time quantitative PCR and Taqman gene assay .................................................... 22 2.12 RT-PCR......................................................................................................................... 22 2.13 Immunofluorescence ..................................................................................................... 22 2.14 In vitro inhibitory effect of Angtpl-4 on LPL activity .................................................. 23 2.15 Isolation of particulate fraction ..................................................................................... 23 2.16 G-LISA assay ................................................................................................................ 23 2.17 Actin polymerization .................................................................................................... 23 2.18 Uptake of exogenous heparanase .................................................................................. 24 2.19 Serum measurements .................................................................................................... 24 2.20 Isolation of lysosomes and nuclear fractions ................................................................ 24 2.21 Western blot .................................................................................................................. 25 2.22 Statistical analysis ......................................................................................................... 25 Chapter 3: Results........................................................................................................................26 3.1 LPL maturation and disassembly is affected by diabetes ............................................. 26 3.1.1 General characteristics of the experimental animals ................................................ 26 3.1.2 Processing of LPL into an active dimeric enzyme is increased following DZ ......... 26 vii  3.1.3 Calnexin is required for LPL processing and is enhanced following diabetes ......... 28 3.1.4 Severe diabetes reduces dimeric LPL in the heart with increased expression of Angptl-4 ................................................................................................................................ 29 3.1.5 Angptl-4 is capable of inhibiting LPL activity and its expression is stimulated by fatty acid................................................................................................................................ 30 3.2 Endothelial cells regulate LPL secretion from cardiomyocytes through heparanase ... 31 3.2.1 RhoA activation is involved in increasing cardiomyocyte LPL secretion ................ 31 3.2.2 ECCM stimulates LPL secretion from cardiomyocytes ........................................... 32 3.2.3 The effect of ECCM on LPL is related to the presence of heparanase ..................... 33 3.2.4 Latent and active heparanase play different roles in LPL secretion ......................... 34 3.2.5 Activation of RhoA by latent heparanase depends on HSPGs and PKC alpha ........ 34 3.2.6 Secretion is increased when myocytes are co-cultured with endothelial cells exposed to high glucose ........................................................................................................ 35 3.3 Cardiomyocyte gene expression is affected by taking up latent heparanase from EC . 35 3.3.1 Exogenous latent heparanase can be taken up by cardiomyocytes ........................... 35 3.3.2 Internalization of latent heparanase is through a caveolae-dependent pathway that requires HSPGs, dynamin and tyrosine kinase activation ............................................. 36 3.3.3 Internalized latent heparanase is activated in lysosomes and enters the nucleus ..... 37 3.3.4 Nuclear entry of heparanase is accompanied by increased matrix metalloproteinase 9 (MMP-9) expression ............................................................................. 37 Chapter 4: Discussion ..................................................................................................................92 4.1 Calnexin augments LPL maturation after moderate diabetes ....................................... 92 4.2 Following severe diabetes Angptl-4 dissembles dimeric LPL ..................................... 95 viii  4.3 Using heparanase, endothelial cells regulate LPL secretion from cardiomyocytes ...... 96 4.4 Heparanase from EC can be taken up by cardiomyocytes to affect gene expression ... 99 Chapter 5: Conclusions and future directions ........................................................................103 5.1 Conclusions ................................................................................................................. 103 5.2 Future directions ......................................................................................................... 104 References ...................................................................................................................................108  ix  List of Tables  Table 1 General characteristic of experimental animals ......................................................... 26  x  List of Figures Figure 1 Coronary LPL activity following diabetes is regulated by multiple factors derived from both cardiomyocytes and endothelial cells. ................................................................... 13 Figure 2 Monomeric and dimeric LPL are present in the heart. ............................................. 40 Figure 3 Acute hyperglycemia increases cardiac LPL dimerization. ..................................... 42 Figure 4 Increased LPL dimers are present in cardiomyocytes from DZ animals and cells acutely exposed to high glucose and palmitic acid. ................................................................ 44 Figure 5 LPL maturation requires calnexin and is enhanced by high glucose and palmitic acid. ......................................................................................................................................... 46 Figure 6 Chronic diabetes also augments LPL processing with increased expression of calnexin. .................................................................................................................................. 49 Figure 7 LMF1 mRNA expression in heart does not change following diabetes. .................. 51 Figure 8 Severe diabetes reduces LPL dimers with an increase in Angptl-4. ........................ 53 Figure 9 Angptl-4 which reduces LPL activity is up-regulated by fatty acid. ........................ 55 Figure 10 Inhibitory effect of Angptl-4 on LPL at the vascular lumen and cardiomyocytes in control animals. ....................................................................................................................... 57 Figure 11 Increased LPL activity in diabetic hearts involves RhoA mediated actin cytoskeleton remodeling. ........................................................................................................ 59 Figure 12 Increased LPL activity in STZ-induced diabetic hearts is associated with RhoA activation. ................................................................................................................................ 61 Figure 13 Endothelial cell conditioned media increases LPL secretion from cardiomyocytes.................................................................................................................................................. 63 Figure 14 ECCM induces RhoA-mediated actin cytoskeleton polymerization. ..................... 65 xi  Figure 15 Latent and active heparanase have divergent effects on LPL secretion. ................ 67 Figure 16 High glucose induced release of active heparanase. .............................................. 69 Figure 17 High glucose affects heparanase secretion from endothelial cells. ........................ 71 Figure 18 Active heparanase releases myocyte surface LPL in a dose-dependent manner. ... 73 Figure 19 Activation of RhoA by latent heparanase requires HSPGs and PKC. ................... 75 Figure 20 Activation of RhoA by PMA leads to actin cytoskeleton polymerization. ............ 77 Figure 21 PKC? is important for RhoA activation by latent-heparanase. .............................. 79 Figure 22 Co-culture with endothelial cells increased LPL secretion in the presence of high glucose. ................................................................................................................................... 81 Figure 23 Heparanase is detected in cardiomyocytes. ............................................................ 83 Figure 24 Internalization of heparanase by cardiomyocytes is mediated by HSPGs. ............ 85 Figure 25 Internalization of latent heparanase is caveolae-dependent and requires dynamin and tyrosine kinase activity. .................................................................................................... 87 Figure 26 Following uptake exogenous latent heparanase is activated in lysosomes. ........... 89 Figure 27 Nuclear entry of heparanase is accompanied by increased gene expression of MMP-9. ................................................................................................................................... 91 Figure 28 Moderate and severe diabetes have a differential impact on LPL at the coronary lumen..................................................................................................................................... 106 Figure 29 Endothelial heparanase is a key mediator to increase LPL secretion following hyperglycemia. ...................................................................................................................... 107   xii  List of Abbreviations AMPK AMP-activated protein kinase Angptl-4 Angiopoietin-like protein 4 bCAECs Bovine coronary endothelial cells CHX Cyclohexamide CPT-1 Carnitine palmitoyltransferase DAPI 4,6-Diamidino-2-phenylindole DZ Diazoxide EC Endothelial cells ECCM Endothelial cell conditioned medium EGFR Epidermal growth factor receptor ER Endoplasmic reticulum ERK-1 Extracellular signal-regulated kinase FA Fatty acid GPI Glycosylphosphatidyl inositol GPIHBP1 Glycosylphosphatidylinositol-anchored high density lipoprotein-binding protein 1 HAT Histone acetyltransferase HG High glucose HS Heparan sulfate HSPGs Heparan sulfate proteoglycans IRE-1 Inositol-requiring kinase 1 LMF1 Lipase maturation factor 1 LPL Lipoprotein lipase MMP-9 Matrix metalloproteinase 9 NEFA Nonesterified fatty acid PA Palmitic acid PDH Pyruvate dehydrogenase PFK Phosphofructokinase PHPLA Post-heparin plasma LPL activity PKC Protein kinase C PKD Protein kinase D PPAR Peroxisome proliferator-activated receptor RAOEC Rat aortic endothelial cells ROS Reactive oxygen species SREBP-1 Sterol regulatory element binding protein-1 STZ Streptozotocin T1D Type 1 diabetes T2D Type 2 diabetes TCA Trichloroacetic acid TG Triglyceride UCP3 Uncoupling protein 3 UPR Unfolded protein response VLDL Very-low-density lipoprotein xiii  Acknowledgements  To begin with I would like to express my sincere gratitude to Dr. Brian Rodrigues, without whom I would have never been ?whipped? into such a shape.  It was Dr. Rodrigues who gave me the opportunity to taste the flavor of real science and generously teaches me everything he knows. I am thankful for his dedication to the students, the time he spent with us.  His support has accompanied me throughout the past four years.  Every word in this thesis reflects his contribution to my career.   I would definitely like to thank my committee members Dr. Roger Brownsey, Dr. James Johnson, Dr. John Hill, and chair Dr. Urs Hafeli.  Your intelligence and experience have broadened my view and strongly encouraged me to pursue perfection.  Those valuable suggestions you provided means a lot to me.  I give my special thanks to Dr. Israel Vlodavsky who kindly shared his resources and expertise with us.   During my work in the lab, there are always been people guiding me on the way.  I would like to thank Mr. Ashraf Abrahami, Ms. Bahira Hussein, Dr. Minsuk Kim, Dr. Fang Wang, and Dr. Prasanth Puthanveetil for training me with technical skills and inspiring me with good ideals.  Much appreciation goes to Dahai Zhang, Amy Chiu, Katharina Neumaier, Andrea Wan, as well as Fulong Wang and Nathaniel Lal for every small little things you helped.  I am also thankful to our secretary Rachel Wu for always being there.  Thanks for all my friends who pushed me to the top when I was down.  I would like to express my thanks to CDA for their financial support during my graduate program.  xiv  Dedication To my family and friends who understand and support me without asking for anything. Sometimes it is just enough to have you around.  1  Chapter 1: Introduction 1.1 Diabetic cardiomyopathy  Diabetes is a global disease that currently affects 371 million of people worldwide.  The prevalence of this disease is estimated to rise to 552 million by 20301.  According to the International Diabetes Federation Atlas, diabetes mellitus caused 4.8 million deaths in 2012.  Approximately 6.8% of Canadians were diagnosed with diabetes, contributing to nearly 30% of the total deaths in the country from 2008 to 20092.  For both Type 1 diabetic (T1D) and Type 2 diabetic (T2D) patients, cardiovascular disease, predominantly associated with atherosclerosis, is the most prevalent cause of this mortality3.  Nevertheless, deterioration of cardiac function can also occur in the absence of coronary heart disease, termed ?diabetic cardiomyopathy?. Initially, diabetic cardiomyopathy is characterized by prolonged diastolic relaxation time and increased stiffness of the left ventricular wall, leading to subsequent systolic dysfunction and heart failure.  Myocardial fibrosis, small vessel disease, cardiac autonomic dysfunction, and metabolic disturbances have all been implicated in the pathogenesis of diabetic cardiomyopathy4.  With metabolism, the heart is capable of using a broad spectrum of substrates, including fatty acids (FA), glucose, and to a lesser extent lactate and ketone bodies5.  Normally, hearts derive about 70% of their energy from FA metabolism6, with glucose (20%) and lactate (10%) making up the remaining part.  After diabetes, several major factors restrict glucose utilization in the heart.  These include a lack of glucose transporters at the sarcolemmal membrane, and inhibition of key enzymes that control glycolysis (eg., phosphofructokinase, PFK) and glucose oxidation (eg., pyruvate dehydrogenase complex, PDH) in the mitochondria4.  As a result, hearts require more FA to generate ATP.  This could be achieved 2  by switching on its FA delivery machinery.  Compared to albumin-bound free FA, triglyceride (TG)-rich lipoproteins are the preferred source of FA7, 8.  This would require hydrolysis of plasma TG to release free FA at the luminal side of the endothelium, a reaction catalyzed by lipoprotein lipase (LPL)9.  Following 2 weeks of streptozotocin (STZ) injection, LPL activity is significantly higher at the coronary lumen of diabetic rats compared to control, which increased the rate of TG-lipoprotein breakdown10.  Interestingly, LPL activity can be turned on within 4 h after diabetes11, suggesting a prominent role in rapidly modulating heart metabolism.  The adaption to use FA is beneficial to the heart in the short term as energy production is maintained in the presence of impaired glucose oxidation.  However, chronic reliance on FA metabolism is detrimental to heart function.  As the cardiomyocyte has a limited capacity for storing FA, the substrate influx has to tightly match its oxidation in the mitochondria to generate ATP.  Beta-oxidation is the primary pathway for FA catabolism in the heart12.  It generates reducing equivalents to create an electrochemical gradient across the mitochondrial inner membrane that drives ATP synthesis, a process called oxidative phosphorylation.  However, reactive oxygen species (ROS) are also produced when electrons leak out of the oxidative phosphorylation process.  In db/db mice hearts, oxidative phosphorylation capacity failed to increase coordinately with elevated fatty acid and beta-oxidation, which leads to excess ROS generation13.  Cardiac contractile function is disturbed as ROS affect calcium homeostasis by damaging ion channels14.  Excess ROS also destabilize mitochondria membranes, triggering cell death15.  Increased FA influx that overwhelms mitochondria oxidation capacity would cause accumulation of lipid intermediates such as ceramide, diacyglycerol, and acyl-CoA, which further inhibit glucose utilization and promote cell death16, 17.  FA are considered endogenous ligands for 3  peroxisome proliferator-activated receptor alpha (PPAR ?) in the heart.  Activation of PPAR ? not only boosts gene expression of carnitine palmitoyltransferase (CPT-1) to facilitate acyl-CoA entry into the mitochondria, but also enzymes involved in beta-oxidation18, 19.  Ironically, FA also increases uncoupling protein 3 (UCP3) expression in the heart through PPAR ?20.  By exporting free fatty acid anions from the mitochondrial, overexpression of UCP3 dissipates the electrochemical gradient across the mitochondrial membrane, thereby uncoupling beta-oxidation and ATP synthesis in STZ-induced T1D hearts 13, 21.  In T1D rat model, depressed mitochondria respiration rate was observed 4 weeks following injection of STZ22.  Hence, chronic exposure of cardiomyocytes to FA ultimately reduces metabolic efficiency.  Although not demonstrated in the heart, overexpression of LPL in skeletal muscle induced UCP3 expression in this tissue by promoting TG-derived FA intake23, suggesting that LPL could potentially cause mitochondrial uncoupling.  In heart studies, because PPAR ? activation favors FA utilization, mice with cardiac-specific overexpression of PPAR ? (MHC-PPAR) showed phenotype features similar to diabetic hearts24, 25.  Strikingly, deletion of LPL reduced myocyte TG accumulation, and rescued cardiac function in these animals26, implicating LPL in a pivotal role in diabetic cardiomyopathy. 1.2 Lipoprotein lipase 1.2.1 Gene transcription  Lipoprotein lipase (LPL) belongs to the lipase family. Its gene is located on 8p22 encoding for a 475 amino acid protein with a 27 amino acid signal peptide27, 28.  A large number of variants have been identified in the LPL gene, and some of them are known to influence LPL function.  For example, most patients with Familial Chylomicronemia 4  Syndrome carry homozygous or compound heterozygous mutations in LPL gene that abolish the activity of the enzyme29.  These includes substitution of G?A at nucleotide position 680 which replaces glutamic acid for glycine at amino residue 14230; a missense mutation resulting in substitution of an alanine for a threonine at residue 17631; and mutation 207, the most prevalent LPL gene mutation among French Canadians32.  Due to LPL deficiency, patients manifest hypertriglyceridemia and fasting chylomicronemia.  LPL HindIII polymorphism is associated with risk of coronary artery disease by affecting binding site of a transcription factor to regulate LPL gene expression33.  Gene transcription of LPL is regulated by multiple cis-acting elements in the promoter region. In humans, a TATA box at -46, a CCAAT motif that binds transcription factor NF-Y, and octamer motif interacting with Oct-1 accounts for the basal promoter activity34, 35.  In 3T3-L1 adipocytes, cholesterol depletion stimulated LPL transcription by binding sterol regulatory element binding protein-1 (SREBP-1) to the sterol regulatory element 2 site in the LPL promoter36, whereas estrogen suppressed promoter activity via an AP-1-like element37.  Reduced binding of transcription factor Sp1 and Sp2 to LPL promoter is responsible for Interferon ?-mediated inhibition of LPL transcription in macrophages38.  Tumor necrosis factor ? also decreased LPL promoter activity by inhibition of NF-Y binding to the CCAAT motif39.  The discovery of Peroxisome proliferator-activated response element in the promoter region could explain the tissue-specific gene modulation effect of PPAR activators on LPL.  As PPAR ? is more restricted to adipose tissue, thiazolidinedione can induce LPL expression, but had no effect on liver40, whereas fibrates selectively increased mRNA level of LPL in the liver by activating the predominant PPAR isoforms-PPAR ? in this tissue41.  LPL expression is also regulated at translational level.  Insulin increased LPL activity in adipocytes probably by stabilizing LPL 5  mRNA42.  Epinephrine inhibits LPL translation in adipocytes possibly due to activation of protein kinase A (PKA).  PKA forms a complex with A kinase anchoring protein (AKAP) 121/149 and binds to the 3'-untranslated region of LPL mRNA to prevent translation43.  Because AKAP121/149 is more abundant in white adipose tissue than in muscle and brown adipose tissue, the inhibitory effect of epinephrine is not significant in these tissues44.  Despite the above regulation, LPL function is largely controlled in a post-translational manner. 1.2.2 Maturation inside the endoplasmic reticulum  Although LPL hydrolyzes TG-rich lipoproteins at the coronary lumen, it is not expressed by endothelial cells, but originates from cardiomyocytes45.  In cardiomyocytes, as newly synthesized LPL protein enters the endoplasmic reticulum (ER), it undergoes glycosylation by attaching oligosaccharide chains to the asparagine residues (Asn43 and Asn359)46.  At this stage, LPL is present as an inactive monomer.  The oligosaccharide chains are subsequently trimmed by Glucosidase I and II, exposing a structure recognized by ER-resident chaperones calnexin/calreticulin.  Association of calnexin/calreticulin with nascent LPL is part of the ?quality control? for most glycosylated proteins.  It allows nascent LPL folding into a proper tertiary structure qualified for dimerization47.  This step is indispensable for LPL processing into active enzyme as folding/dimerization of human LPL in transfected sf21 cells is only promoted when calreticulin is co-expressed48.  Using castanospermine to inhibit glucosidase-mediated trimming of the oligosaccharide chains, the interaction between LPL monomers with chaperones was hindered, thereby decreasing active LPL49.  The two inactive monomeric LPL are then assembled noncovalently in a head-to-tail fashion to form a catalytically functional enzyme50-52.  LPL dimerization relies on lipase maturation factor 1 6  (LMF1)53, 54.  LMF1 is a five transmembrane protein localized in the ER, and its highly conserved domain DUF1222 is essential for physically interacting and maturation of LPL as well as hepatic lipase55.  Mutations of lmf1 that result in truncations of DUF1222 domain are responsible for combined lipase deficiency in both mice and human56.  Patients with this disorder demonstrate decreased LPL and hepatic lipase activity with concomitant hypertriglyceridemia.  Notably, only dimeric LPL is active and competent to exit the ER47, 57. 1.2.3 Secretion to cardiomyocyte surface  So far, there is limited information regarding how LPL is further processed and sorted into secretion vesicles in the Golgi. Recently, protein kinase D (PKD) has emerged as a crucial factor in the formation of LPL secretion vesicles58.  By phosphorylating phosphatidylinositol-4 kinase IIIbeta (PI4KIII-?), it activates the lipid kinase activity of PI4KIII-? to stimulate fission of vesicles from the trans-Golgi network59.  PKD is also known to direct secretory vesicles to the plasma membrane with its partner vesicle associated membrane protein (VAMP), a soluble NSF attachment protein receptor (SNARE) localized on the vesicle membrane serves as docking proteins for exocytosis60.  In the trans Golgi network, LPL can also be captured by SorLA (sortilin-related receptor) that would route LPL to endosomes, and ultimately lysosomes for degradation61.  It is suggested that up to 80% of newly synthesized LPL in the adipocytes is degraded in the lysosomes instead of being secreted62.  However, details of intracellular turn-over of this enzyme in the heart remain to be elucidated.  For LPL-containing vesicles destined for secretion, they require polymerization of actin cytoskeleton to build a ?bridge? from the Golgi to the plasma membrane63.  The secreted LPL is sequestered by heparan sulfate proteoglycans (HSPGs) on the cardiomyocyte membrane.  Depolymerization of the actin cytoskeleton by cytochalasin D reduced LPL 7  activity on the myocyte surface63.  The Rodrigues? lab has reported an increased intracellular trafficking of LPL from Golgi to the myocyte surface after diabetes which is driven by the energy sensor AMP-activated protein kinase (AMPK)64.  In STZ-induced T1D rats, AMPK is activated at an early stage possibly because impaired glucose utilization caused a rapid energy deficiency65.  As a result, heat shock protein 25 (Hsp25) was phosphorylated, dissociated from actin monomers and PKC ?, leading to actin cytoskeleton polymerization and PKD-mediated vesicle formation, respectively58.  In this way, more intracellular LPL is moved to the myocyte surface after diabetes. 1.2.4 Release from the cardiomyocyte surface  Myocyte surface HSPGs serve as a temporary docking site for LPL.  The core proteins of HSPGs are attached to the cell surface either through a transmembrane domain in case of the syndecan family, or via glycosylphosphatidyl inositol (GPI) anchor as observed with glypican66, 67.  The core protein is decorated with heparan sulfate (HS) side chains covalently attached to specific serine residues on the ectodomain68.  The HS side chains are polymers of repeating disaccharides (glucurionic acid and N-acetylglucosamine) modified via deacetylation, sulfation, and epimerization, which introduces heterogeneity to the structure.  These modifications also create negatively-charged residues that allow the HS to interact with multiple ligands through their heparin-binding domain, including antithrombin, fibroblast growth factor, and LPL69-71.  In fact, LPL is bound to HSPGs on the myocyte surface through ionic interaction, which is important for maintaining LPL activity72.  HS has been suggested to be a putative extracellular chaperone, retaining LPL activity as it translocates from synthesizing sites to the endothelium73.   8   For its onward secretion towards the endothelium, cleavage of myocyte surface HSPGs is required.  When endothelial cell conditioned medium (ECCM) was given to adipocytes, LPL activity was released from adipocyte surface, suggesting that factors required to cleave HSPGs arise from the endothelium73.  Heparanase, an endoglucuronidase synthesized in the endothelial cells (EC) could be the factor mediating this cross-talk.  It is synthesized in the EC as a latent 65 kDa precursor, secreted and bound to EC surface HSPGs before being internalized into lysosomes74.  There it is cleaved into a 50 kDa active form that can digest the HS into 7~8 kDa fragments74, oligosaccharides that are capable of carrying LPL, and preserving its bioactivity.  For example, lysophosphatidylcholine-treated endothelial cell medium can release LPL activity from the adipocytes, and it has been shown that more heparanase is secreted from the basolateral than the apical side upon stimulation, suggesting its role in LPL trafficking from parenchymal cell rather than detaching LPL from the vascular lumen73.  Secretion of heparanase from EC was also increased in the presence of high glucose, and heparanase was found infiltrated into the interstitial space between EC and cardiomyocytes in hearts from diabetic animals75.  In addition, high glucose also upregulates heparanase gene expression in EC76, an effect inhibited by insulin77.  In fact, heparanase level in the plasma was directly correlated with that of blood glucose in T2D patients78.  It is tempting to suggest that diabetes may liberate more LPL from the myocyte surface HSPGs as a result of increased heparanase secretion from EC.  As these liberated LPL translocate to the coronary lumen, more FA will be delivered to the heart.  In this way, EC can possibly control cardiomyocyte metabolism.  Besides releasing HSPG-bound ligands from extracellular matrix and cell surface, heparanase also has other biological effects.  Studies on cancer biology suggested that tumor cells might change the phenotype of surrounding cells by 9  ?feeding? them heparanase that is capable of modulating gene transcription.  Nuclear entry of active heparanase stimulated differentiation of esophageal keratinocyte by cleaving heparan sulfate in the nucleus79.  It also reduced the nuclear content of syndecan-1, a type of HSPGs that inhibits histone acetyltransferase (HAT) activity, and hence promoted certain gene expression in CAG human myeloma cells, including vascular endothelial growth factor (VEGF) and matrix metallopepetidase 9 (MMP-9)80.  In addition, heparanase is known to regulate gene expression by controlling histone H3 methylation patterns81.  Whether heparanase in the heart would affect LPL gene expression remains unclear.  1.2.5 Translocation from cardiomyocyte towards the apical side of EC  The mechanism by which LPL is transported across the interstitial space, vascular basement membranes, and to ultimately its functional site at the vascular lumen remained a mystery for a long time.  However, a deficiency in extracellular collagen XVIII, a HSPG present in vascular basement membranes decreased LPL activity at the vascular lumen in humans82, suggesting the importance of extracellular matrix HSPGs for LPL migration.  Digestion of HSPGs on the basolateral side of EC also hindered the transcytosis of LPL83, 84.  At the apical side of EC, HSPGs were considered binding sites for LPL which also mediate the internalization and recycling of LPL inside EC85, 86.  However, HSPGs are ubiquitously expressed in many cell types and do not bind LPL specifically.  So far, no evidence has shown that genetic deletion of any type of HSPGs in vivo is associated with hypertriglyceridemia.  On the other hand, a newly discovered molecule, glycosylphosphatidylinositol-anchored high density lipoprotein-binding protein 1 (GPIHBP1) was identified as a unique LPL receptor located on the luminal face of the capillary endothelium, docking LPL to break down triglyceride87.  It is distributed in LPL-10  expressing tissues including heart, adipose tissue, and skeletal muscle, and unlike HSPGs, it does not bind to any other member of the lipase family88.  In addition, transcytosis of LPL across the EC requires GPIHBP1 shuttling. LPL captured by GPIHBP1 at the subendothelial space is transported in vesicles to the vascular lumen in a caveolin-1-independent manner89, 90.  Thus, GPIHBP1 knockout mice manifest severe hypertriglyceridemia in accordance with a dramatic reduction in LPL activity at the vascular lumen91.  The C-terminus portion, especially amino acids 403-407 (a region different from the principal heparin-binding domain) of LPL is responsible for interacting with GPIHBP192.  For GPIHBP1, both the amino-terminal acidic domain and the lymphocyte antigen 6 motif are important for binding and transport of LPL93.  Mutations that disturb protein structure of these regions could lead to chylomicronemia in human94, 95.  Expression of GPIHBP1 in EC is rapidly regulated by nutrition status and PPAR ?96.  Notably, GPIHBP1 expression depends on factors released from parenchymal cells as primary EC lost GPIHBP1 expression after a single passage96.  The role of GPIHBP1 in LPL function after diabetes has yet to be studied.  It should be noted that GPIHBP1 can also serve as a ?platform?, permitting the simultaneous binding of both LPL and lipoproteins at the vascular lumen, enabling lipolysis90. 1.2.6 Turnover at the vascular lumen  LPL activity is also regulated at the vascular lumen.  Previously, we have shown an increase in coronary LPL activity in animals made diabetic with 55 mg/kg STZ (D55).  Intriguingly, on increasing the dose to 100 mg/kg (D100), LPL activity was ?turned-off?97.  Compared to D55, D100 animals had a more profound reduction in circulating insulin which also made these animals hyperlipidemic.  In this situation, FA could possibly decrease coronary LPL in multiple ways: a) FA displace LPL from its endothelial cell surface binding 11  sites for degradation in the liver98, b) direct inactivation of LPL enzyme activity99, and c) FA impair LPL vesicle trafficking to the myocyte surface through caspase-3-mediated inactivation of PKD100.  A decade ago, two angiopoietin-like proteins, Angptl-3 and Angptl-4, drew attention as deficiency of either protein caused elevation of post-heparin plasma LPL activity (PHPLA) in transgenic mice (representing whole body LPL activity released from the vascular lumen)101.  Both Angptl-3 and Angptl-4 are composed of a NH2-terminal coiled-coil domain and a C-terminal fibrinogen-like domain, and the coiled-coil domain is sufficient to interact and inactivate LPL102.  Though sharing similar structures, the two angiopoietin-like proteins work in different ways.  Expression of Angptl-3 is mainly restricted to the liver. Its action is possibly through promotion of LPL cleavage via proprotein convertases, leading to dissociation of LPL from both HSPGs and GPIHBP1 binding sites103.  Its expression is driven by the liver X receptor and is mainly active in the fed state104.  However, in humans, the level of Angptl-3 is not related to that of PHPLA and plasma triglyceride105.  Angptl-4 is widely expressed in various tissues including white adipose tissue, liver, heart, and skeletal muscle106.  In these cells, it forms oligomers through disulfide linkage, and after secretion, is cleaved at a canonical proprotein convertase cleavage site that separates the N- and C-terminal domain107.  The N-terminal coiled-coil domain remains oligomerized, a feature that is essential for its stability and LPL inhibitory effect108.  The secreted N-terminal oligomers function in the circulation, converting dimeric LPL at the vascular lumen into inactive monomers109.  Interestingly, long chain FA, through activation of PPAR ?, increased transcription of muscle-derived Angptl-4110.  Change in Angptl-4 expression is rapid as in adipose tissue, down-regulation of its mRNA was observed within 6 h after refeeding fasted animals, which is consistent with elevated LPL activity in this tissue109.  ANGPTL4 variants 12  E40K and T266M with defective Angptl-4 function lowered fasting triglyceride levels in patients with a T2D background111, suggesting a potential therapeutic value of Angptl-4 blocker in treating dyslipidemia after diabetes.  The effect of Angpt-4 in cardiac LPL is not well-studied.  It is possible that in D100 animals showing hyperlipidemia, LPL could be ?turned-off? when FA stimulates Angptl-4 expression in the heart. 1.3 Hypothesis and research objectives  Cardiac LPL is of crucial importance for maintaining normal heart function.  My hypothesis is that coronary LPL activity following diabetes is regulated by multiple factors derived from both cardiomyocytes and endothelial cells (Figure 1). The objectives of my research proposal were to: 1. Investigate the impact of cardiomyocyte chaperones and angiopoietin-like proteins on LPL dimerization and disassembly. 2. Determine the mechanisms by which EC, through its secretion of heparanase, regulates cardiomyocyte LPL. Gaining further insight into these mechanisms should allow the identification of novel targets for therapeutic intervention, to prevent or delay cardiac failure following diabetes.   13   Figure 1 Coronary LPL activity following diabetes is regulated by multiple factors derived from both cardiomyocytes and endothelial cells. Following gene transcription, LPL mRNA is translated as a monomer.  The inactive monomer undergoes maturation with the help of chaperones to form the functional dimer in the endoplasmic reticulum (ER).  Dimerized LPL then moves to the Golgi for further processing.  The fully processed LPL is sorted into vesicles that are targeted for secretion.  These secretion vesicles transport LPL to the myocyte surface via actin cytoskeleton.  On the myocyte surface, LPL is sequestered by HSPGs.  Heparanase secreted from endothelial cells (EC) in response to glucose can cleave myocyte HSPGs side chains, thereby liberating LPL for ongoing secretion.  LPL released into the interstitial space is then transported towards the 14  apical side of EC to bind to luminal binding sites (HSPGs and GPIHBP1), where they break down triglyceride lipoprotein such as VLDL-TG, providing fatty acid (FA) to cardiomyocytes.  Dimeric LPL can be dissembled into inactive monomers by Angptl-4.  Augmented LPL function following diabetes would bring overwhelming amounts of FA to the heart, which leads to lipotoxicity in the long term. Nu: nucleus. 15  Chapter 2: Methods 2.1 Materials  Rat aortic endothelial cells (RAOEC) were obtained from Cell Applications.  Bovine coronary artery endothelial cells (bCAECs) was from Clonetics.  Diazoxide (DZ), streptozotocin (STZ), palmitic acid (PA), castanosperimine (Cs) and cyclohexamide (CHX), lysophosphatidic acid (LPA), cytochalasin D, Y-27632, G?6976, phorbol 12-myristate 13-acetate (PMA), filipin, genistein, and chloroquine were obtained from Sigma.  Heparin (HEPALEAN, 1000 U/ml) was from Organon, Canada.  PKC ? inhibitor was obtained from Calbiochem.  It inhibits PKC ?I and PKC ?II with IC50 of 21 and 5 nM, respectively.  Heparinase III (IBEX techonologies) was purified from recombinant Flavobacteriumheparinum.  Heparanase inhibitor SST0001 was kindly provided by Sigma Tau Research (Switzerland S.A., Mendrisio, CH).  [3H]-triolein was purchased from Amersham Canada.  Purified human Angptl-4 protein (Genway Biotech) is a human recombinant and 100% homologous to the 26-229 amino acid sequence of human Angptl-4.  Purified active and latent heparanase were prepared as described112.  Anti-calnexin antibody was from Stressgen Canada.  Anti-LPL 5D2 antibody was a kind gift from Dr. J. Brunzell, University of Washington, Seattle.  It is a monoclonal antibody recognizing LPL from different species, but not related lipases like hepatic lipase113.  Anti-heparanase antibody mAb 130 was from InSight (Rehovot, Israel), which recognizes both the active (50 kDa) and latent form (65 kDa) of heparanase.  Anti-rabbit True Blot was from eBioscience.  All other antibodies were obtained from Santa Cruz.  The RhoA activation G-LISA assay kit and F-actin/G-actin assay kit were obtained from Cytoskeleton (Denver, CO).  HTRF heparanase 16  activity assay kit was obtained from Cisbio.  To measure free fatty acid released from VLDL-TG breakdown, an NEFA-C assay kit was purchased from Wako. 2.2 Experimental animals  The investigation conforms to the Guide for the Care and Use of Laboratory Animals published by NIH and the University of British Columbia (UBC).  Adult male Wistar rats (250?320 g) were used.  To induce acute hyperglycemia, diazoxide (DZ), a selective ATP-sensitive K+ channel opener was administered intraperitoneally at 100 mg/kg.  DZ causes a rapid decrease of insulin secretion within 1 h, and the animals remain hyperglycemic for 4 h114-116.  4 h after DZ, animals were killed and hearts removed.  For chronic diabetes, streptozotocin (STZ), a ?-cell specific toxin was administered as a single dose intravenously to more closely mimic Type 1 diabetes58, 100.  STZ was given at doses of 55 (D55) or 100 (D100) mg/kg.  STZ animals were kept for 4 days before hearts were removed for every experiment.  With D55, animals are insulin deficient, but do not require insulin supplementation for survival (moderate diabetes).  Unlike D55, D100 animals also develop hyperlipidemia and are not able to survive beyond 4 days without insulin supplementation (severe diabetes).   2.3 Animal treatment  In some experiment, 1 h after DZ injection, animals were injected with 4 U of a rapid-acting insulin (Humulin R, Eli Lilly Canada Inc) through the tail vein.  Blood glucose was monitored every 30 min up to 3 h, animals were killed and hearts removed for determination of coronary LPL activity and RhoA activation.  In another experiment, 2 h before and after diazoxide injection, DZ animals were given 7.5 mg/rat heparanase inhibitor SST0001 subcutaneously117, 118. 17  2.4 Isolated heart perfusion  Hearts were isolated and perfused retrogradely119.  To measure coronary LPL, perfusion solution was changed to buffer containing heparin (5 U/ml).  Coronary effluents were collected (for 10 s) at different time points over 5 min.  LPL activity in each fraction was determined, plotted against time, and coronary LPL activity was presented as area under the curve over 5 min or peak activity.  Subsequent to LPL displacement, hearts were used for heparin-sepharose chromatography.  To evaluate the effect of Angptl-4 on LPL at the vascular lumen, hearts from control and D55 animals were perfused with or without 1 ng/ml purified Angptl-4 for 1 h in a re-circulating mode.  Perfusates were collected and run on a heparin-sepharose column.  Following Angptl-4 perfusion, the hearts were perfused with heparin to release LPL activity remaining at the vascular lumen.  A modified Langendorff retrograde perfusion was used to isolate active and latent heparanase in the interstitial effluent75.  Briefly, the hearts were taken out with the aorta, inferior vena cava, lungs still attached and perfused retrogradely through the aorta by Langendorff technique.  The left and right superior venae cava as well as the inferior vena cava were ligated close to the atrium.  The left and right pulmonary veins were tied off close to the lungs, and the lungs were removed.  The pulmonary artery was cannulated.  Fluid that drips down to the apex of the heart (interstitial transudate) were collected. 2.5 Isolation of cardiac myocytes  Ventricular calcium-tolerant myocytes were prepared by a previously described procedure120.  Following isolation, cardiomyocytes were plated on laminin-coated culture dishes and allowed to settle down for 3 h.  Unattached cells were washed away using medium 199 before treatment.  Myocytes from both control and DZ hearts were isolated, and 18  intracellular LPL protein in these cells determined by Western blot.  Gene expression of LPL was measured using Taqman assay (Invitrogen).  Gene expression was normalized to ?-actin. 2.6 Endothelial cell culture  bCAECs and RAOEC were cultured at 37?C in a 5% CO2 humidified incubator alone or co-cultured with adult rat cardiomyocytes75.  bCAECs from the 5th to the 8th passage were used.  2.7 Endothelial cell conditioned medium (ECCM)  bCAECs were incubated with high glucose (25 mM) DMEM for 30 min.  This medium was collected as ECCM. 2.8 In vitro treatments  The glucosidase inhibitor castanospermine (Cs, 50 mM, 2 h) was used to interrupt the association between LPL and calnexin.  Colocalization of LPL and calnexin was observed using immunofluorescence.  LPL activity released into the medium and remaining in the myocytes was also measured.  Myocytes were also exposed to 25 mM glucose (HG, high glucose) and 1.0 mM palmitic acid (PA) bound to 1% BSA (HG+PA) for 2 h, cells lysed, and run on a heparin-sepharose column.  To evaluate the impact of HG+PA on processing of newly synthesized LPL, cells were pre-incubated with 50 ?M cyclohexamide (CHX, 1 h) to inhibit protein synthesis.  Subsequently, cells were washed, treated with HG+PA for another 1 h, and LPL activity released into the media was determined.  Calnexin and LPL colocalization was also visualized at the indicated times.  To study the effect of FA on Angptl-4 expression, isolated myocytes were treated with 1.0 mM palmitic acid (PA) for 4-24 h, and Angptl-4 mRNA determined using real-time PCR.  To study the direct effect of Angptl-4 on cardiomyocytes, isolated cardiomyocytes from control animals were incubated 19  with Angptl-4 for the indicated times, and LPL activity in the media and on the cell surface (by incubating cardiomyocytes with media containing 8 U/ml heparin for 1 min) determined.  In a separate experiment, at the end of treatment, cell lysates were loaded onto a heparin-sepharose column, and dimeric LPL determined.  To study the effect of RhoA activation on LPL secretion, control myocytes were incubated with lysophosphatidic acid (LPA, 100 nM-1 ?M) in the presence or absence of 1 ?M cytochalasin D (Cy, an actin polymerization inhibitor).  To rule out the effect of protein synthesis on LPL secretion, 50 ?M of the protein synthesis inhibitor cyclohexamide (CHX) was applied to myocytes 1 h before and during treatment with LPA, and LPL secretion into the medium determined.  RhoA activation was tested in myocytes treated with ECCM, or 1 ?g/ml purified latent or active heparanase at the indicated times.  To study the role of HSPGs and PKC? in RhoA activation, 10 IU/L heparinase III and 5 ?M G?6976 was used to digest heparan sulfate of HSPGs and inhibit PKC?/? in cardiomyocytes, respectively.  To study the specific effect of PKC? and ? on RhoA activation in the presence of latent heparanase, siRNAs specific for PKC? were designed and synthesized by Invitrogen (5?-UGAAGAAGCGGGCGAUGAAUUUGUG-3?).  Lipofectamine RNAi/MAX (Invitrogen) were used to transfect siRNAs for PKC? or control unrelated siRNA into cardiomyocytes (at a final concentration of 50 nM).  24 h after transfection, medium was changed to Media 199 and following an additional 24 h, cells were incubated with 1 ?g/ml latent heparanase for 15 min and RhoA activation in myocytes measured.  In another experiment, myocytes were pre-incubated with 200 nM PKC? inhibitor, 1 h before and during latent heparanase treatment, and RhoA activation in myocytes measured.  To investigate the role of RhoA in ECCM-mediated actin 20  polymerization, 10 ?M Y-27632 was used to block ROCK, the downstream effector of RhoA.  Following treatment of bCAECs with 5 or 25 mM glucose DMEM for 30 min, cells were collected and intracellular latent and active heparanase measured using Western blot.  To test whether consecutive exposure to high glucose can eventually deplete heparanase secretion, bCAECs were treated with 25 mM glucose DMEM for 30 min, medium removed (1st release), and the cells exposed to a second 30 min incubation with 25 mM glucose DMEM (2nd release).  Medium from both 1st and 2nd release were concentrated to detect latent and active heparanase.  In a separate experiment, bCAECs that were pre-treated with 25 mM glucose DMEM for 30 min (1st release) were placed in the upper chamber of a co-culture system with isolated cardiomyocytes at the bottom.  These cardiomyocytes had themselves been pre-incubated with purified LPL to saturate surface binding sites.  25 mM glucose DMEM (2nd release) was then applied to the co-culture system, and medium collected from the bottom chamber after 30 min to test LPL activity released from cardiomyocytes.  Results were compared to a co-culture system in which bCAECs was not pre-treated with high glucose.   To test whether exogenous heparanase can bind and be taken up by cardiomyocytes, isolated myocytes were treated with 500 ng/ml recombinant latent heparanase for different time intervals at 37 ?C.  This experiment was also conducted at 15 ?C to inhibit heparanase binding and uptake.  In the presence of 10 IU/ml of heparin to compete for HSPGs binding sites, or 10 IU/L heparitinase III to digest heparan sulfate side chains (2 h), the contribution of myocyte surface HSPGs in heparanase uptake was determined.  For studying the mechanism of internalization, 350 mM sucrose or 1 ?g/ml filipin was applied to myocytes 21  for 15 min before latent heparanase, to block clathrin-coated pits and caveolae-dependent internalization, respectively.  We also used 0-25 ?? dynasore or 0-100 ?? genistein for 1 h to inhibit dynamin and tyrosine kinase activity which could be involved in internalization of exogenous heparanase.  To inhibit lysosomal enzymes, 200 ?? chloroquine was used and myocytes incubated for 2 h. 2.9 LPL activity  LPL catalytic activity was determined by measuring the in vitro hydrolysis of a sonicated [3H]triolein substrate emulsion64.  The standard assay conditions were 0.6 mM glycerol tri[9,10-3H]oleate (1 mCi/mmol; 1 Ci=37 GBq), 25 mM piperazine-N,N?-bis(2-ethanesulfonic acid) (pH 7.5), 0.05% (wt/v) albumin, 50 mM MgCl2, 2% (v/v) heat-inactivated chicken serum as a source of apo-CII.  The reaction mix was incubated at 30 ?C for 30 min.  Reaction product, sodium [3H]oleate, was extracted and determined by liquid scintillation counting.  Results are expressed as nmol/h/ml (perfusate, elution fraction or media) or nmol/h/mg protein (cell lysate). 2.10 Heparin-sepharose chromatography    Heparin-sepharose chromatography was carried out as described previously to separate dimeric LPL from monomers47, 102.  Heart perfusates or equal amounts (total protein) of tissue homogenates or cell lysates were loaded onto a HiTrap? HP Column and sequentially eluted with 0.25, 0.75, 1.0, and 1.5 M NaCl at 0.2 ml/min.  Fractions (1 ml each) collected were used to detect LPL activity and protein by ELISA (USCNLIFE) or Western blot following TCA precipitation.  To determine total amount of monomeric or dimeric LPL, fractions from 0.75 or 1.0 M NaCl were combined before TCA precipitation.  To determine the processing stage of dimeric LPL, aliquots from the 1.0 M NaCl fractions were subjected 22  to endoglycosidase H (endo H) digestion.  Digested products of LPL were visualized by Western blotting following TCA precipitation. 2.11 Real time quantitative PCR and Taqman gene assay  LMF1 and Angptl-4 mRNA levels were determined using SYBR Green real-time quantitative PCR (Roche).  Primer sequences for LMF1 were: forward, 5?- TGATCCTGCAGGGCACA -3?; reverse: 5?- GTCCAGGCGGTAGTGGTA -3?121; for Angptl-4, forward, 5?- CTCTGGGATCTCCACCATTT -3?; reverse: 5?- TTGGGGATCTCCGAAGCCAT -3?122.  All values obtained were normalized to 18S-ribosomal RNA.  Myocytes from both control and DZ hearts were isolated, gene expression of LPL was measured using Taqman assay (Invitrogen).  Gene expression was normalized to ?-actin. 2.12 RT-PCR  Total RNA was extracted from isolated myocytes or RAOEC using Trizol reagent (invitrogen), and 1 ?g total RNA was used for RT-PCR. PCR primers were as follow123, 124: Heparanase (forward, 5?-CAAGAACAGCACCTACTCACGAAGC-3?; reverse, 5?-CCACATAAAGCCAGCTGCAAAGG-3?; 616 bp product)  MMP-9 (forward, 5?-CCCCACTTACTTTGGAAACGC-3?; reverse, 5?-ACCCACGACGATACAGATGCTG-3?; 686 bp product) 18S rRNA (forward, 5?-CGGCTACCACATCCAAGGAA-3?; reverse, 5?-GCTGGAATTACCGCGGCT-3?; 187 bp product) 2.13 Immunofluorescence  Following the indicated treatments, cells were double stained with Alexa635 and Alexa488 to colocalize LPL (red), and calnexin (green), respectively.  Isolated 23  cardiomyocytes were also treated with 1 ?g/ml active or latent heparanase for 30 min, and cells probed with anti-syndecan-4 antibody (H-140, Santa Cruz) to visualize syndecan-4 (green) stained by Alexa488.  To induce syndecan-4 clustering, an antibody against the extracellular domain of syndecan-4 (H-17, Santa Cruz) was used as a positive control125, 126. 4,6-Diamidino-2-phenylindole (DAPI) was used to stain nuclei.  Slides were visualized using a Confocal microscope. 2.14 In vitro inhibitory effect of Angtpl-4 on LPL activity  LPL from vascular lumen was obtained by perfusing hearts from D55 animals with heparin.  Purified human Angptl-4 was incubated with perfusates containing peak LPL activity at a final concentration of 1 ng/ml, 0.1 ?g/ml, and 1 ?g/ml.  The reaction mix was incubated at 37?C, and at the indicated times, 100 ?l reaction solution was removed for measuring LPL activity. 2.15 Isolation of particulate fraction  To determine RhoA activation in vivo, particulate fraction from ventricles were prepared as described previously127.  2.16 G-LISA assay  G-LISA assay was performed according to manufacturer?s instruction (Cytoskeleton).  The active GTP-RhoA was detected by reading the absorbance at 490 nm. 2.17 Actin polymerization  Actin polymerization was evaluated by measuring the filamentous to globular actin (F-actin/G-actin ratio) using an assay kit from Cytoskeleton. 24  2.18 Uptake of exogenous heparanase  Following incubation with 500 ng/ml purified latent heparanase, myocytes were washed three times with cold PBS.  To determine binding and uptake of heparanase by myocytes, total cell lysis were collected to detect heparanase by Western blot.  For measuring internalization of heparanase, plasma membrane was removed by a procedure described previously.  Briefly, myocytes were lysed in 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.2 M sucrose, 2 mM EDTA, 2 mM EGTA and protease inhibitor cocktail (Roche), and centrifuged at 10, 000 g at 4 ?C for 10 min.  Supernatant was further centrifuged at 100,000 at 4?C for 1 h to spin down the plasma membrane.  The resulting supernatant (-PM) containing cytosolic fraction was used to monitor heparanase internalization.  2.19 Serum measurements  Blood samples were collected from the tail vein before termination, and serum isolated.  Concentrations of nonesterified fatty acid (NEFA) and triglyceride were determined using diagnostic kits (Wako and Stanbio).  Angptl-4 was analyzed using an ELISA kit from Raybiotech. 2.20 Isolation of lysosomes and nuclear fractions   Lysosomes enriched fractions were isolated using a kit from Sigma.  Nuclear and cytosolic fractions were separated using a kit from Pierce.  To validate the purity of proteins, we used cytosolic (GAPDH) and nuclear (Histone H3) protein marker to detect their presence in cytosolic and nuclear fractions. 25  2.21 Western blot  Western blot was carried out as described previously127.  In some experiments, samples were concentrated by TCA precipitation, or Amicon centrifuge filter (Millipore) before detection of heparanase or LPL.  2.22 Statistical analysis  Values are means ? SE.  Wherever appropriate, one-way ANOVA followed by the Bonferroni test was used to determine differences between group mean values.  The level of statistical significance was set at P<0.05.  26  Chapter 3: Results 3.1 LPL maturation and disassembly is affected by diabetes 3.1.1 General characteristics of the experimental animals  Injection of DZ resulted in stable hyperglycemia within 4 h.  These animals also exhibited an increase in circulating levels of NEFA.  Although administration of STZ at both doses produced hyperglycemia of equal intensity as that seen with DZ, only D100 animals showed a robust elevation in circulating FA and TG levels (Table 1).  Table 1 General characteristic of experimental animals  Blood Glucose (mM) NEFA (mM) TG (mM) Control 6.2?0.9 0.3?0.1 0.5?0.1 Diazoxide (4 h) 26.2?2.5** 1.0?0.2** 0.5?0.2 D55 (4 d) 24.6?2.0** 0.4?0.1 0.5?0.3 D100 (4 d) 28.0?1.8** 1.0?0.3** 2.3?0.5** Blood samples were collected from the tail vein before termination, and serum isolated by centrifugation.  Serum concentrations of nonesterified fatty acid (NEFA) and triglyceride (TG) were determined using diagnostic kits.  Results are the mean?SE of 3 animals in each group.  *Significantly different from control, P<0.05. **Significantly different from control, P<0.01. 3.1.2 Processing of LPL into an active dimeric enzyme is increased following DZ  LPL is an enzyme present in two forms based on its affinity to a heparin-sepharose column.  Monomeric LPL has low affinity to the column, is predominantly eluted in 0.4-0.75 M NaCl fractions, and has no catalytic activity.  Active LPL is a homodimer eluted at higher 27  concentrations of NaCl (1.0-1.5 M)47, 102.  We found that the majority of LPL in the heart is monomeric (eluted at 0.75 M NaCl), with only ~30% of the enzyme in the form of dimers (eluted in 1.0 M NaCl fractions) accounting for all of the LPL activity (Figure 2).  Consistent with our previous studies, total LPL protein remained unchanged following acute hyperglycemia induced by DZ (Figure 3A).  However, when equal amounts of homogenate protein were loaded onto a heparin-sepharose column, the 1.0 M fractions from DZ hearts demonstrated higher LPL activity (Figure 3B) and protein (Figure. 3C), suggesting the presence of more dimeric LPL in these hearts.    Dimeric LPL could be located either at the vascular lumen, or within cardiomyocytes.  To determine vascular LPL, hearts were perfused with heparin and LPL activity measured in the perfusate.  DZ hearts possessed increased LPL activity at this site (Figure 4A).  Following stripping of the dimeric enzyme from the vascular lumen with heparin, hearts with predominantly myocyte LPL were then subjected to a heparin-sepharose column.  Interestingly, in addition to the vascular lumen, DZ hearts also harbored increased dimeric LPL in cardiomyocytes (Figure 4B, lane 1 vs. 3).  Maturation of LPL including dimerization occurs within the endoplasmic reticulum (ER) of cardiomyocytes.  To further localize myocyte LPL dimers, we used endo H to digest dimeric LPL.  Endo H cleaves the glycan chains of LPL based on their processing stages in different cellular organelles; the 57 and 55 kDa products represent LPL that has either passed or is present within Golgi apparatus respectively, whereas the 52 kDa band refers to LPL undergoing processing inside the ER47, 128.  Interestingly, DZ hearts had a higher proportion of dimeric LPL at a post-ER processing stage (Figure 4B, lane 2 vs. 4), implicating augmented maturation of LPL following DZ.  To examine whether this is due to acute exposure to a hyperglycemic and hyperlipidemic 28  environment seen in DZ animals, isolated cardiomyocytes were incubated with HG+PA for 2 h.  Using the same conditions, we have previously found an accelerated LPL trafficking from Golgi to the myocyte surface.  HG+PA also increased dimerization of LPL (Figure 4C), indicating that processing of LPL could be a pre-requisite for trafficking. 3.1.3 Calnexin is required for LPL processing and is enhanced following diabetes  LPL is a glycan protein whose processing into active dimers requires its association with ER resident chaperones like calnexin/calreticulin to allow proper folding.  Using immunofluorescence, LPL and calnexin were found to be intensively colocalized at the perinuclear region (Figure 5A).  LPL association with this chaperone was further confirmed by immunoprecipitation (Figure 5A, inset).  Castanospermine (Cs) is a glucosidase inhibitor that blocks trimming of LPL glycan chains, such that it cannot be recognized by calnexin49.  Using Cs, we were able to inhibit the association between LPL and calnexin; a complete absence of the perinuclear colocalization of these proteins was observed (Figure 5B, inset).  This hindered LPL secretion and processing-both secreted (Figure 5B) and intracellular (Figure 5C) LPL activity declined in the presence of Cs, likely due to reduced LPL dimerization (Figure 5C, inset).  To determine whether HG+PA augmented LPL processing by affecting the association between calnexin and newly synthesized LPL, cardiomyocytes were pre-treated with the protein synthesis inhibitor cyclohexamide (CHX).  After 1 h, as a result of no newly synthesized LPL entry into the ER, CHX abolished the association between LPL and calnexin (Figure 5D, inset 2).  Upon removal of CHX and renewal of protein synthesis, association between newly synthesized LPL and calnexin was visualized in the presence or absence of HG+PA.  1 h after re-entry of LPL into the ER, newly synthesized LPL was observed colocalized with calnexin at the perinuclear region (Figure 5D, inset 3).  29  However, in contrast to Cs, even though HG+PA attenuated this colocalization (Figure 5D, inset 4), more LPL were observed present at the cell surface, suggesting that more enzyme had completed processing under these conditions.  Increased LPL processing was also reflected in more active LPL being released into the media (Figure 5D).  To examine LPL processing subsequent to chronic diabetes, 55 mg/kg of STZ was administered to generate ?moderate diabetes?.  Similar to DZ, although chronic diabetes did not increase total LPL protein (data not shown), D55 hearts also demonstrated higher amounts of dimeric LPL (Figure 6A).  The importance of calnexin in LPL processing is also suggested in D55 hearts; augmented LPL dimerization in these hearts was accompanied by increased calnexin expression (Figure 6B).  However, we did not observe the same increase in calnexin when cardiomyocytes were acutely exposed to HG+PA (data not shown).  Another crucial factor for LPL dimerization, LMF1, was found to be unchanged following D55 (Figure 7).  Overall, our data suggests that following diabetes, more LPL is processed into its active dimeric form, which appears to involve the ER chaperone calnexin. 3.1.4 Severe diabetes reduces dimeric LPL in the heart with increased expression of Angptl-4  Increasing the dose of STZ to 100 mg/kg (D100) produced hyperglycemia similar to that observed in D55 animals.  However, unlike D55, D100 animals developed severe hyperlipidemia (Table 1).  Additionally, in contrast to D55, D100 animals showed a decline in dimeric LPL, both in the whole heart (Figure 8A) and at the vascular lumen (Figure 8B).  This effect on LPL was not a result of decreased calnexin expression in D100 (Figure 8C), but was accompanied by a robust increase in Angptl-4 serum concentrations (Figure 8D).  30  Furthermore, cardiac expression of Angptl-4 protein (Figure 8E) and mRNA (Figure 8E, inset) were also found to be higher in D100 animals.  3.1.5 Angptl-4 is capable of inhibiting LPL activity and its expression is stimulated by fatty acid  To test the direct inhibitory effect of Angptl-4 on LPL, dimeric LPL collected from the vascular lumen of D55 animals was incubated with purified Angptl-4 in vitro.  The inhibition of LPL activity by Angptl-4 was fast (within 15 min) and dose-dependent with 1 ?g/ml Angptl-4 inhibiting about 85% of the enzyme activity at 2 h (Figure 9A).  It should be noted that the serum concentration of Angptl-4 in D100 animals (1 ng/ml) was effective in inhibiting approximately 20% of LPL activity in vitro (Figure 9A).  When this concentration of Angptl-4 was perfused through D55 hearts for 1 h, LPL activity remaining at the vascular lumen was reduced (Figure 9C).  Interestingly, this reduction in LPL activity correlated with an increased appearance of monomeric LPL in the perfusion buffer (Figure 9B), and a decline in dimeric LPL remaining at the coronary lumen (Figure 9C, inset), suggesting that the inhibitory action of Angptl-4 on LPL is through the conversion of dimeric LPL into monomers, which in turn reduces its affinity to the binding sites at the vascular lumen.  This inhibitory effect of Angptl-4 was also observed in isolated hearts (Figure 10A) and cardiomyocytes (Figure 10B and C) from control animals.  To duplicate the elevated serum NEFA level seen in D100 animals, isolated cardiomyocytes were treated with 1.0 mM PA for 4-24 h.  Within 4 h of exposure to PA, Angptl-4 mRNA did not change significantly.  However, at 12 h, increase in Angptl-4 mRNA was observed, an effect that was even more dramatic following a 24 h exposure to PA (Figure 9D). 31  3.2 Endothelial cells regulate LPL secretion from cardiomyocytes through heparanase 3.2.1 RhoA activation is involved in increasing cardiomyocyte LPL secretion  In consistent with our previous observation, DZ animals with hyperglycemia (CON, 6.5 ? 0.6; DZ, 19.2 ? 2.7 mM, P<0.01) had increased coronary LPL activity129 (Figure 11A).  Within 30 min, 4 U insulin effectively inhibited the development of hyperglycemia in DZ animals (DZ + In, 7.6 ? 1.4; DZ, 21.5 ? 4.2, mM, P<0.01), and normal glycemia remained until the animals were killed.  Insulin attenuated the increase of coronary LPL activity in these DZ hyperglycemic animals (Figure 11A).  As RhoA has been reported to regulate actin cytoskeleton remodeling, an event that could affect LPL secretion127, we tested RhoA activation in diabetic hearts.  Upon activation, RhoA shifts from the cytosolic to the particulate fraction, and binds to GTP130.  In DZ hearts, an increased RhoA in the particulate fraction was observed, which was inhibited by insulin treatment (Figure 11B).  As DZ animals are hyperglycemic for only a short period, a more chronic model of hyperglycemia was induced by injecting rats with 55 mg/kg streptozotocin (D55), and animals kept for 4 days.  Similar to DZ animals, D55 animals also have higher coronary LPL activity, which was accompanied by RhoA activation in these hearts (Figure 12).  To determine whether RhoA activation can induce LPL secretion from myocytes in vitro, cells were incubated with LPA.  After 2 h of LPA, an increased amount of LPL was released into the medium (Figure 11C).  RhoA activation by LPA was confirmed in cardiomyocyte as GTP-RhoA increased immediately 1 min following LPA, and declined with time (Figure 11C, inset).  One effect of RhoA activation is actin cytoskeleton polymerization, which we observed as an increase in the formation of F actin in the presence of LPA (Figure. 11D).  Given that LPL secretion 32  relies on stress fibers to move from an intracellular pool to the myocyte surface131, 132, we inhibited actin cytoskeleton polymerization using Cy, and found that the impact of LPA on LPL secretion was abolished (Figure 11E).  As the effect of LPA on LPL secretion was reproducible in mycoytes in which protein synthesis was inhibited (Figure 11E, inset, right panel), our data suggest that the increased LPL secretion observed with LPA is not a consequence of augmented protein synthesis, but likely due to increased LPL trafficking.  Thus, RhoA activation could contribute towards augmented LPL secretion, possibly via actin cytoskeleton remodeling. 3.2.2 ECCM stimulates LPL secretion from cardiomyocytes  Following diabetes, endothelial cells are the first cells exposed to hyperglycemia, and could potentially release multiple factors affecting cardiomyocyte metabolism.  For this reason, we incubated myocytes with high glucose treated endothelial cell conditioned medium (ECCM). Interestingly, ECCM released myocyte LPL within 30 min, an effect that was more significant after 1 h.  High glucose itself had no impact on LPL (Figure 13A).  As this increase in medium LPL was accompanied by a reciprocal decrease in LPL activity remaining at the myocyte surface (Figure 13A, inset, upper panel), we concluded that ECCM is capable of releasing myocyte surface LPL.  Importantly, this ECCM-released LPL was catalytically active and able to breakdown exogenous VLDL-TG to FA (Figure 13A, inset, bottom panel).  In addition to its ability to release LPL, we also tested whether ECCM could stimulate the replenishment of LPL when the enzyme at the myocyte surface is depleted.  Myocytes were pre-treated with ECCM or heparin (10 mU/ml), followed by a bolus dose of heparin (8 U/ml) to deplete surface LPL, as confirmed in Figure 13B (left panel).  Following a 2 h recovery, myocytes pre-treated with ECCM were able to recruit significantly more LPL 33  activity to the surface (Figure 13B, right panel), an effect not observed with 10 mU/ml heparin.  Considering that RhoA is involved in augmented LPL secretion from myocytes, we tested the effect of ECCM on RhoA activation.  An increase in GTP-bound RhoA was observed within 5 min in response to ECCM, reaching a peak at 15 min, and declined to basal levels after 30 min (Figure 14A).  We did not see a similar effect with 25 mM glucose or 10 mU/ml heparin (data not shown).  The down-stream effect of RhoA activation, actin cytoskeleton polymerization, was also augmented with ECCM, an effect that was abolished in the presence of the ROCK inhibitor Y-27632 (Figure 14B).  Hence, our data suggested that ECCM can stimulate both the release, and likely through RhoA-mediated actin cytoskeleton polymerization, replenishment of LPL at the myocyte surface.  3.2.3 The effect of ECCM on LPL is related to the presence of heparanase  As LPL at the cardiomyocyte surface resides on HSPGs, its release by ECCM could be a consequence of cleavage of these binding sites by heparanase.  As anticipated, ECCM contained a higher amount of both latent (65 kDa) and active (50 kDa) heparanase (Figure 15A), with a reciprocal decrease in the intracellular content of this enzyme (Figure 17C).  The increase in active heparanase protein mirrored the higher heparanase activity in this medium (Figure 16).  In vivo, both latent and active heparanase also increased in the interstitial space of hearts from DZ animals (Figure 15B), where coronary endothelial cells are exposed to high glucose.  When heparanase in the ECCM was immunoprecipitated by an anti-heparanase antibody, thereby reducing the amount of both latent and active heparanase (Figure 15C, inset), the LPL releasing effect of ECCM was compromised (Figure 15C).  When bCAECs were exposed to high glucose for two consecutive periods of 30 min, the amount of heparanase released into the medium diminished during the second incubation 34  (Figure 17A).  In addition, the ability of this ECCM from the second incubation to release myocyte surface LPL also decreased (Figure 17B). 3.2.4 Latent and active heparanase play different roles in LPL secretion  Since high glucose stimulates the release of both latent and active heparanase from EC, we tested the roles of these two forms on LPL secretion using purified heparanase.  Active, but not latent heparanase caused the release of LPL (Figure 15D), and this effect of active heparanase was dose-dependent (Figure 18).  When heparin was added to remove LPL from the myocyte surface, active heparanase was unable to release LPL (Figure 15E), suggesting that LPL released by active heparanase is from the myocyte surface.  Unexpectedly, RhoA activation was not observed with active heparanase, which only responded to the latent form of the enzyme (Figure 15F).  Our data imply that active heparanase releases LPL from the myocyte surface, whereas latent heparanase may move LPL from an intracellular store to the surface. 3.2.5 Activation of RhoA by latent heparanase depends on HSPGs and PKC alpha  To examine the mechanism of RhoA activation by latent heparanase, we considered whether the integrity of the myocyte surface HSPGs is required for this signal mechanotransduction.  As expected, removal of HS by heparinase III blocked RhoA activation by latent heparanase (Figure 19A).  Interestingly, the effect of latent heparanase on RhoA activation and cytoskeleton polymerization was abolished by the PKC?/? inhibitor G?6976 (Figure 19B and C), whereas PMA, a conventional PKC activator had effects similar to that seen with latent heparanase (Figure 20).  Furthermore, the RhoA activation effect observed with latent heparanase was only attenuated in myocytes with reduced PKC? expression, but not in cells in which PKC? was specifically inhibited (Figure 21).  In normal 35  myocytes, syndecan-4 is distributed on myocyte surface in a dispersed manner.  Upon addition of latent heparanase and anti-syndecan-4 antibody, syndecan-4 appeared clustered on the surface (Figure 19D, arrow), an effect not seen with active heparanase.  3.2.6 Secretion is increased when myocytes are co-cultured with endothelial cells exposed to high glucose  To simulate diabetes in vitro, myocytes were co-cultured with EC.  As a limited number of myocytes can be seeded in the co-culture system, the intrinsic LPL activity at the surface of these myocyte is low.  Hence, we used purified LPL added exogenously to amplify this surface pool.  200 ?g of purified LPL was sufficient to saturate HSPGs binding sites on the myocyte surface (Figure 22A, inset).  Using this amount of LPL, we exposed the co-culture system to normal and high glucose.  A significantly higher medium LPL activity was detected from the lower chamber in the presence of high glucose (Figure 22A).  To examine whether the released LPL from myocytes is ultimately recruited onto the apical side of EC, we tested LPL activity on the EC surface after 2 and 4 h with high glucose.  In the normal glucose co-culture, we did not observe any change in EC surface LPL activity at 2 or 4 h.  With high glucose, a robust increase in EC surface LPL activity was evident after 4 h (Figure 22B).  Notably, high glucose co-culture also caused actin polymerization (within 1 h) (Figure 22C, inset), that was preceded by RhoA activation (within 30 min) in myocytes (Figure 22C). 3.3 Cardiomyocyte gene expression is affected by taking up latent heparanase from EC 3.3.1 Exogenous latent heparanase can be taken up by cardiomyocytes  In the heart, heparanase is expressed by the endothelial cells and cardiomyocytes do not synthesize this enzyme.  To confirm this, we used RT-PCR, and were unable to detect the 36  presence of the heparanase gene in freshly isolated cardiomyocytes, even though this gene was highly expressed in RAOEC (Figure 23A, right inset).  In the absence of gene expression, heparanase protein, both the 65 kDa latent and the 50 kDa active forms were detected in isolated myocytes (Figure 23A, left inset).  As we failed to detect the EC marker CD31, our results indicated that these isolated myocytes were not contaminated by EC (Figure 23A, left inset).  Interestingly, unlike RAOEC, the dominant form present in cardiomyocytes was the active heparanase (Figure 23A, left inset).  Culturing cardiomyocytes for 36 h reduced the content of both heparanase forms, suggesting that the protein cannot be synthesized de novo when being turned over (Figure 23B).  Introducing latent heparanase into the culture medium rapidly increased the level of latent heparanase in myocytes (within 5 min), which continued to accumulate over time.  This increase was inhibited when temperature was lowered to 15?C.  Unlike latent heparanase, the increase of active heparanase was gradual, and only significantly increased after 4 h (Figure 23C), suggesting intracellular conversion of latent to active heparanase in cardiomycoytes.  3.3.2 Internalization of latent heparanase is through a caveolae-dependent pathway that requires HSPGs, dynamin and tyrosine kinase activation  Heparanase in the total cell lysates could consist of two parts: heparanase bound to the myocyte surface or that which has been internalized.  When plasma membrane was removed by ultracentrifugation, as validated by the absence of membrane protein Na+-K+ ATPase (Figure 24A, right inset), heparanase was still observed in the cytosolic fraction of myocytes, indicating that it had been internalized (Figure 24A).  Using heparin to competitively inhibit the binding of latent heparanase to HSPGs, we were able to reduce the amount of heparanase that was internalized (Figure 24A).  As similar results were seen with heparitinase III, that 37  digests the heparan sulfate chain of HSPGs (Figure 24B), our data implies that heparanase may be taken up by myocytes via binding to the HS heparin binding domain of HSPGs.  As this internalization of heparanase was blocked by filipin, but was insensitive to sucrose (Figure 25A), it is likely that HSPGs-mediated endocytosis of heparanase is a caveolae-dependent rather than a clathrin-mediated event.  It should be noted that at the concentration of sucrose used, the endocytosis of epidermal growth factor receptor (EGFR) that is typically internalized through formation of clathrin-coated pits, was blocked (Figure 25A, right inset).  The involvement of dynamin and tyrosine kinase in this endocytotic process was apparent as dynasore (Figure 25B) and genistein (Figure 25C) reduced the amount of heparanase internalized.  3.3.3 Internalized latent heparanase is activated in lysosomes and enters the nucleus  On incubation of cardiomyocytes with latent heparanase, we were able to detect this enzyme in lysosomal fractions within 30 min.  With time, latent heparanase in this lysosomal fraction progressively increased.  Unlike latent heparanase, lysosomal active heparanase only increased at a later time point (after 3 h) (Figure 26A).  As this increase in lysosomal active heparanase was prevented by chloroquine, which inhibit lysosomal proteases, our data suggests lysosomal conversion of latent to active heparanase in cardiomyocytes (Figure 26B).  3.3.4 Nuclear entry of heparanase is accompanied by increased matrix metalloproteinase 9 (MMP-9) expression  Assuming that conversion of latent to active heparanase in the lysosomes is to fulfill a biological function, we determined the nuclear content of heparanase, and observed the presence of active heparanase in this organelle even in the absence of exogenous heparanase.  Following addition of latent heparanase into culture medium, there was a dramatic increase 38  of both latent and active heparanase in the nucleus after 4 h (Figure 27A).  This entry was followed by an increase in MMP-9 expression 18 h after heparanase treatment (Figure 27B).  In vivo studies using D55 cardiomyocytes demonstrated a 1.5 fold increase in the amount of active heparanase inside the nucleus (Figure 27C).  MMP-9 expression in these myocytes was also significantly higher compared to control (Figure 27D).  However, we did not detect any change in LPL expression in both latent heparanase treated myocytes or D55 myocytes (data not shown).     39   40  Figure 2 Monomeric and dimeric LPL are present in the heart.  Whole heart homogenate was loaded onto a heparin-sepharose column followed by washing with increasing concentrations of NaCl (0.25, 0.75, 1.0, and 1.5 M, lower bar) at a flow rate of 0.2 ml/min.  Both LPL amount (right axis) and activity (left axis) in each fraction were determined using ELISA and in vitro hydrolysis of a sonicated [3H]triolein substrate emulsion, respectively.    41   42  Figure 3 Acute hyperglycemia increases cardiac LPL dimerization.   Male Wistar rats were made hyperglycemic by injecting 100 mg/kg diazoxide (DZ) i.p. 4 h after injection, animals were killed and hearts removed.  Total LPL protein expression was determined using Western blot normalized to GAPDH. Results are the mean?SE of 6 animals in each group (A).  Ventricles were lysed with 25 mM ammonia buffer, pH 8.2, containing 1% Triton X-100, 0.1% SDS, 10 U heparin/ml, and protease inhibitor.  Equal amounts of total protein from control (CON) and DZ heart homogenates were loaded onto a heparin-sepharose column, and eluted with increasing concentrations of NaCl (0.25, 0.75, 1.0 and 1.5 M).  1 ml fractions were collected and examined for LPL activity.  Before the activity assay, aliquots of each fraction were diluted with equilibration buffer containing BSA to adjust NaCl concentration to 0.25 M and BSA to 1 mg/ml.  Elution was repeated with samples from 6 animals in each group, but only a representative heparin-sepharose chromatography is illustrated (B).  Peak LPL activity is presented as mean?SE from 6 animals in each group (B, inset).  Fractions with LPL activity (1.0 M) were combined and precipitated by TCA before Western blot for LPL was carried out (C).  Results are the mean?SE of 6 animals in each group.  *Significantly different from control, P<0.05.  AU: arbitrary units.    43   44  Figure 4 Increased LPL dimers are present in cardiomyocytes from DZ animals and cells acutely exposed to high glucose and palmitic acid.   4 h after injection of DZ, hearts were isolated, perfused with heparin (5 U/ml), and fractions of perfusate at the indicated times were analyzed for LPL activity (A).  Subsequent to heparin perfusion and detachment of vascular LPL, heart homogenates were subjected to heparin-sepharose elution to isolate LPL dimers (in the 1.0 M NaCl elutions).  These fractions were combined and digested with (+) or without (-) endo H for 20 h, concentrated by TCA precipitation, and LPL protein determined by Western blot (B).  Isolated cardiac myocytes were plated on laminin-coated 60 ? 15 mm tissue culture dishes and treated with palmitic acid (1 mM bound to 1 % BSA) and 25 mM glucose (HG+PA) for 2 h.  Media containing 1% BSA were used as control (CON).  Cellular dimeric LPL was determined by running cell lysates onto a heparin-sepharose column and fractions eluted at 1.0 M NaCl were used to determine LPL protein by Western blot (C).  Results are the mean?SE of 5 repeated experiments using different animals.  *Significantly different from control, P<0.05.  AU: arbitrary units.   45   46  Figure 5 LPL maturation requires calnexin and is enhanced by high glucose and palmitic acid.   Plated myocytes from CON were immunostained with LPL (red) and calnexin (green).  DAPI was used to stain the nuclei (blue), and colocalization of LPL and calnexin was visualized (Merge, yellow) (A).  Cells were lysed with CHAPS lysis buffer (50 mM HEPES buffer, pH 7.5, containing 1% CHAPS, 200 mM NaCl, and protease inhibitor).  Cell lysate of cardiomyocytes from control animals were immunoprecipitated using anti-calnexin (IP:Cn) antibody overnight at 4?C.  The immunocomplex was captured by anti-rabbit Ig IP beads and immunoblotted for LPL.  Direct Western blot of cell lysate was carried out as a positive control (P) (A, insert).  Isolated myocytes were incubated in the presence or absence of castanospermine (Cs) for 2 h.  At the end of treatment, colocalization of LPL and calnexin was visualized (B, inset).  LPL activity released into the medium (B) and remaining in the myocytes (C) were measured.  Treated cardiomyocytes were also loaded onto a heparin-sepharose column to determine the amount of dimeric LPL (C, inset).  Results are the mean?SE of 3 repeated experiments using different animals.  *Significantly different from control, P<0.05.  **Significantly different from control, P<0.01.  Isolated myocytes were pre-incubated with 50 ?M cycloheximide (CHX) for 1 h.  After sufficient washing to remove CHX, cells were treated with HG+PA or BSA only (CON) for another 1 h.  LPL activity released into the media within this 1 h was measured (D).  Results are the mean?SE of 3 repeated experiments using different animals.  *Significantly different from control, P<0.05.  Cells were also fixed with 4% paraformaldehyde before (D, inset 1) or after (D, inset 2) pre-incubation with CHX, and also following treatment with (D, inset 3) or without (D, inset 4) 47  HG+PA.  Colocalization of LPL and calnexin (yellow) in these conditions were visualized by immunofluorescence.  Scale bar=25 ?m.  48   49  Figure 6 Chronic diabetes also augments LPL processing with increased expression of calnexin.  For chronic diabetes, animals were injected with 55 mg/kg STZ (D55), kept for 4 days and hearts removed.  Equal amounts of total protein from CON and D55 heart homogenates were loaded onto a heparin-sepharose column, pre-washed with 0.25 M NaCl and eluted with 0.75, 1.0, and 1.5 M NaCl.  LPL activity in each fraction was analyzed (A), and fractions from 1.0 M NaCl (containing LPL dimers) were combined to determine LPL amount by Western blot following TCA precipitation.  Only a representative chromatography (A) and blot of LPL (A, insert) are shown from 5 repeated experiments.  Calnexin protein expression (Cn) in CON and D55 hearts was determined by Western blot and normalized to GAPDH (B).  Results are the mean?SE of 5 animals in each group.  **Significantly different from control, P<0.01.    50   51  Figure 7 LMF1 mRNA expression in heart does not change following diabetes.   LMF1 mRNA level from CON and D55 hearts was analyzed by real-time quantitative PCR and normalized to 18S-ribosomal RNA.  Results were plotted as fold over control of mean?SE of 3 animals in each group.    52   53  Figure 8 Severe diabetes reduces LPL dimers with an increase in Angptl-4.  Animals were made moderate or severely diabetic by injecting STZ at a dose of 55 (D55) or 100 mg/kg (D100) respectively, and kept for 4 days.  Hearts were isolated and equal amounts of homogenate protein from CON, D55, and D100 were loaded onto a heparin-sepharose column and eluted with increasing concentrations of NaCl.  LPL activity in the 1.0 M fractions was measured and peak activity presented as mean?SE of 5 animals in each group (A).  In a separate experiment, hearts from CON, D55, and D100 were perfused with heparin (5 U/ml), perfusates collected at the indicated times, and LPL activity determined (B, representative graph).  These perfusates at the indicated times were pooled and loaded onto a heparin-sepharose column.  Dimeric LPL in the 1.0 M fractions was determined using TCA precipitation followed by Western blot (B, inset).  Results are the mean?SE of 3 animals in each group.  *Significantly different from control, P<0.05.  Heart homogenates from CON, D55, and D100 were also used for Western blot to detect calnexin (Cn, C) or Angptl-4 (E) protein expressions normalized by GAPDH.  Serum samples were collected from the different groups to detect Angptl-4 using an ELISA assay (D).  Angptl-4 mRNA expression from CON and D100 hearts was analyzed by real-time quantitative PCR.  Gene expression was evaluated by normalizing to 18S-ribosomal RNA, and plotted as fold over control (E, inset).  Results are the mean?SE of 5 animals in each group.  *Significantly different from control, P<0.05.  **Significantly different from control, P<0.01.   54   55  Figure 9 Angptl-4 which reduces LPL activity is up-regulated by fatty acid.   Hearts from D55 animals were perfused with heparin to release LPL at the vascular lumen.  Fractions with peak LPL activity were then incubated with purified human Angptl-4 at a final concentration of 1 ng/ml, 0.1 ?g/ml, and 1 ?g/ml.  Reaction mix was incubated at 37?C and LPL activity at the indicated times determined (A).  Results are the mean?SE from 3 repeated experiments.  Hearts from D55 animals were also directly perfused with or without 1 ng/ml purified Angptl-4 for 1 h.  Perfusion buffer was collected at the end of this period.  Monomeric LPL released into the perfusates were isolated by collecting the 0.75 M fractions from the heparin-sepharose column and quantified by Western blot (B).  Following Angptl-4 perfusion, hearts were subsequently perfused with 5 U/ml heparin to release LPL activity remaining at the vascular lumen.  LPL activity in the perfusate is expressed as area under curve over the 3 min perfusion (C).  These perfusates over 3 min were pooled and loaded onto a heparin-sepharose column to visualize dimeric LPL in the 1.0 M fractions (C, inset).  Results are the mean?SE of 3 animals in each group.  *Significantly different from control, P<0.05.  **Significantly different from control, P<0.01.  Isolated cardiomyocytes were incubated with 1.0 mM palmitic acid (PA) for 4, 12 or 24 h.  mRNA expression of Angptl-4 was quantified by real-time PCR and compared to control (CON) treated with 1% BSA (D).  Protein expression of Angptl-4 after 24 h incubation with palmitic acid is illustrated in the inset.  Results are the mean?SE of 5 repeated experiments using different animals.  *Significantly different from control, P<0.05.  AU: arbitrary units; AUC: area under curve.    56   57  Figure 10 Inhibitory effect of Angptl-4 on LPL at the vascular lumen and cardiomyocytes in control animals.   Hearts from control animals were perfused with or without 1 ng/ml purified human Angptl-4 for 1 h.  After Angptl-4 perfusion, hearts were subsequently perfused with 5 U/ml heparin to release LPL activity remaining at the vascular lumen.  LPL activity in the perfusate is expressed as area under curve over the 3 min perfusion (A).  Results are the mean?SE of 3 animals in each group.  *Significantly different from control, P<0.05.  Isolated cardiomyocytes from control animals were incubated with 1 ng/ml, 0.1 ?g/ml, or 1 ?g/ml Angptl-4 for the indicated times, incubation media collected, and LPL activity measured (B).  In the 12 h groups, at the end of treatment, cells were washed with normal media, LPL on the cell surface released by incubation with media containing 8 U/ml heparin for 1 min, and LPL activity on the surface determined (C).  Results are the mean?SE of 3 repeated experiments using different animals.  *Significantly different from control, P<0.05. **Significantly different from control, P<0.01.  In a separate experiment, isolated cardiomyocytes were plated on 60 ? 15 mm tissue culture dishes and treated with or without 1 ?g/ml angptl-4 for 12 h.  Cells were washed with cold PBS and harvested.  Cell lysates were loaded onto a heparin-sepharose column, and dimeric LPL visualized in the 1.0 M fractions by Western blot following TCA precipitation (C, inset).  AUC: area under curve.   58   59  Figure 11 Increased LPL activity in diabetic hearts involves RhoA mediated actin cytoskeleton remodeling. Animals were made hyperglycemic with DZ, and kept for 4 h.  1 h after DZ injection, some animals were given 4 U of rapid-acting insulin i.v. (DZ+In), and kept for another 3 h. Animals were killed, hearts removed, and perfused with heparin (5 U/ml) to release coronary LPL.  Coronary effluents were collected (for 10 s) at different time points over 5 min, and LPL activity in each fraction determined.  The results are presented as area under the curve (AUC) for heparin-released LPL activity over 5 min (A).  In a separate experiment, particulate fractions were also isolated from whole heart homogenates of control (CON), DZ and DZ+In animals, and RhoA recruitment to the particulate fraction determined by Western blot.  A plasma membrane protein, Na+-K+-ATPase alpha subunit was used as a loading control (B).  *P<0.05, compared to CON, #P<0.05, compared to DZ, n= 3.  Isolated cardiomyocytes from CON were incubated with increasing concentrations of LPA for 2 h, and LPL protein secreted into the medium measured by Western blot (C).  Using 1 ?M LPA, RhoA activation in cardiomyocytes was determined at the indicated times using G-LISA assay (C, inset).  *P<0.05, compared to CON, n=3.  Myocytes were also pre-treated with 1 ?M cytochalasin D for 30 min prior to incubation with 1 ?M LPA (LPA+Cy).  Actin cytoskeleton polymerization (D) and LPL secretion in these cells (E) were determined after 1 h and 2 h, respectively.  In a different experiment, LPL secretion in the presence of LPA was tested in myocytes pre-incubated with 50 ?? of the protein synthesis inhibitor cyclohexamide (CHX) (E, inset, right panel)  *P<0.05, **P<0.01, compared to CON, #P<0.05, compared to LPA, n= 3-4.   60   61  Figure 12 Increased LPL activity in STZ-induced diabetic hearts is associated with RhoA activation. Male Wistar rats (250-320 g) were injected i.v. with 55 mg/kg STZ (D55), and animals kept for 4 days before hearts were removed.  Particulate fractions were isolated from whole heart homogenates of control (CON) and D55 animals, and RhoA recruitment to the particulate fraction determined by Western blot (A).  In a separate experiment, hearts were also perfused with heparin (5 U/ml) to release coronary LPL.  Coronary effluents were collected (for 10 s) at different time points over 5 min, and LPL activity in each fraction determined.  The results are presented as area under the curve for heparin-released LPL activity over 5 min (B).  *P<0.05, compared to CON, n= 3.   62   63  Figure 13 Endothelial cell conditioned media increases LPL secretion from cardiomyocytes. ECCM was applied to cardiomyocytes, and at the indicated times, LPL activity released into the medium determined.  Cardiomyocytes were also incubated with 5.5 mM glucose (NG) and 25 mM glucose (HG) as controls (A).  After 30 min incubation with NG or ECCM, LPL remaining on the myocyte surface was released by incubating cells with 8 U/ml heparin for 3 min, and LPL activity determined (A, inset, upper panel).  Cardiomyocytes were treated with 5.5 mM glucose DMEM (CON) or ECCM for 30 min, and the medium collected.  This medium was then incubated with increasing concentrations of VLDL-TG (0-0.8 mM) and the concentration of released free fatty acids determined after 30 min (A, inset, bottom panel).  *P<0.05, **P<0.01, compared to NG, n=3-5.  Isolated cardiomyocytes were treated with 5.5 mM glucose (CON), ECCM, or 10 mU/ml heparin for 30 min.  Following this, surface LPL was depleted by incubating these cells with 8 U/ml heparin for 3 min.  To validate depletion of this surface LPL, 8 U/ml heparin was given and surface LPL activity determined (B, left panel).  Following a wash with PBS, cells were allowed to recover in 5.5 mM glucose DMEM for 2 h.  LPL activity recruited to the myocyte surface was then determined (B, right panel).  *P<0.05, compared to CON, n= 3.   64   65  Figure 14 ECCM induces RhoA-mediated actin cytoskeleton polymerization. Myocytes were incubated with ECCM for 5, 15 or 30 min, and RhoA activation evaluated by measuring GTP-bound RhoA.  Results were compared to myocytes incubated with 5.5 mM glucose DMEM (CON, A).  In a different experiment, F to G actin ratio was measured 1 h following incubation with ECCM (B).  10 ?M Y-27632 was also added to myocytes 30 min prior to ECCM (ECCM+Y), and actin cytoskeleton polymerization evaluated (B).  *P<0.05, **P<0.01, compared to CON; #P<0.05, compared to ECCM, n=3-4.   66   67  Figure 15 Latent and active heparanase have divergent effects on LPL secretion. bCAEC were incubated with 5.5 mM (NG), or 25 mM (HG) glucose DMEM for 30 min. The amount of latent and active heparanase released into the medium was determined by Western blot (A).  Control and DZ hearts were perfused using a modified Langendorff perfusion technique to separate interstitial fluid from coronary perfusate.  Latent (65 kDa) and active heparanase (50 kDa) were measured by Western blot in the interstitial fluid (B).  Heparanase was immunoprecipitated from ECCM by an anti-heparanase antibody and the amount of latent and active heparanase remaining in this ECCM was determined and compared to the original ECCM using Western blot (C, inset).  This immunoprecipitated ECCM (IP-ECCM) was applied to myocyte for 30 min, and LPL activity released into medium tested (C).  *P<0.05, compared to NG or CON; #P<0.05, compared to ECCM, n=3-4.  Isolated cardiomyocytes were incubated with medium 199 (CON) and 1 ?g/ml purified active (A-HEPA) or latent (L-HEPA) heparanase for 30 min, and LPL activity released into the medium determined (D).  In a different experiment, myocytes were pre-incubated with 8 U/ml heparin for 3 min before 1 ?g/ml purified active heparanase (Hep+A-HEPA), and LPL activity released after 30 min determined (E).  Results were compared to CON and myocytes without heparin pre-incubation (A-HEPA).  RhoA activation was measured in cardiomyocytes in the presence of 1 ?g/ml active (A-HEPA) or latent (L-HEPA) heparanase for 15 min (F).  *P<0.05, **P<0.01, compared to CON; #P<0.05, compared to A-HEPA, n=3.   68   69  Figure 16 High glucose induced release of active heparanase.   bCAECs were treated with 5.5 (NG) or 25 (HG) mM glucose DMEM for 30 min, and medium collected to measure heparanase activity with a kit (A).  *P<0.05, compared to NG, n=4.     70   71  Figure 17 High glucose affects heparanase secretion from endothelial cells. bCAECs were treated with 25 mM glucose DMEM for 30 min, medium removed (1st HG), and the cells exposed to a second 30 min incubation with 25 mM glucose DMEM (2nd HG).  The amounts of latent and active heparanase in the medium of the 1st and 2nd HG treatment were measured by Western blot (A).  *P<0.05, compared to 1st HG release, n=3. In a co-culture system, cardiomyocytes placed in the bottom chamber were pre-incubated with 200 ?g purified LPL for 30 min to saturate myocyte surface HSPGs. Cells were washed with PBS to remove unbound LPL.  bCAECs that were pre-treated with 25 mM glucose DMEM for 30 min  were then placed in the upper chamber of the co-culture system.  25 mM glucose DMEM was applied to the co-culture system, and the medium from the bottom chamber collected after 30 min to determine LPL activity released from cardiomyocytes.  Results were compared to a co-culture system in which bCAECs was not pre-treated with high glucose. **P<0.01, n=3 (B).  bCAECs were treated with either 5 (NG) or 25 (HG) mM glucose DMEM for 30 min, and intracellular heparanase determined by Western blot (C). *P<0.05, compared to NG, n=3 (C).     72   73  Figure 18 Active heparanase releases myocyte surface LPL in a dose-dependent manner. Isolated myocytes were incubated with increasing concentrations of active (A-HEPA) or latent (L-HEPA) heparanase for 30 min, and LPL activity released into the medium determined.  5.5 mM glucose DMEM was used as control (CON).   74   75  Figure 19 Activation of RhoA by latent heparanase requires HSPGs and PKC. Myocytes were pre-treated with (III+HEPA) or without (HEPA) 10 IU/L heparinase III for 30 min, and RhoA activation determined 15 min following incubation with 1 ?g/ml latent heparanase.  The effect of heparinase III per se on RhoA activation was also evaluated (III) (A).  A similar experiment was repeated, except that 5 ?M G?6976 was used to pre-treat the myocytes (B).  In this experiment, F to G actin ratio was evaluated 1 h following addition of latent heparanase (C).  Isolated cardiomyocytes were treated with 1 ?g/ml active (A-HEPA) or latent (L-HEPA) heparanase, or 3 ?g/ml anti-syndecan-4 antibody (Ab) for 30 min, and syndecan-4 distribution visualized by immunofluorenscence (D).  *P<0.05, compared to CON, n=3. #P<0.05, compared to HEPA, n=3.   76   77  Figure 20 Activation of RhoA by PMA leads to actin cytoskeleton polymerization. Isolated myocytes were treated with 1 ?M PMA for the indicated times, and RhoA activation determined using a G-LISA assay kit (A).  Actin cytoskeleton polymerization in these cells was determined 1 h following PMA treatment (B).  *P<0.05, **P<0.01, compared to CON, n= 3.   78   79  Figure 21 PKC? is important for RhoA activation by latent-heparanase. In a 6-well plate, isolated myocytes from control rats (CON) were transfected with siRNA specific for PKC? at 100 or 200 nM or control unrelated siRNA (SCR, 200 nM). 24 h following transfection, medium was changed to Media 199 and following an additional 24 h, cells were collected to Western Blot for PKC? (A).  *P<0.05, compared to SCR, n=3.  In a separate experiment, myocytes were transfected with 200 nM siRNA for PKC? (+) or 200 nM control unrelated siRNA (-), before being exposed to 1 ?g/ml latent heparanase for 15 min. GTP-bound RhoA in these myocytes were determined (B).  200 nM PKC? inhibitor was given to mycoytes 1 h before and during incubation with latent heparanase for 15 min, and RhoA activation also determined in these myocytes (C).  Results were from 3 experiments using different control animals. *P<0.05, compared to control, #P<0.05, compared to control treated with L-HEPA.     80   81  Figure 22 Co-culture with endothelial cells increased LPL secretion in the presence of high glucose.   A co-culture was carried out using EC in the insert (top) and cardiomyocytes in the well (bottom).  Cardiomyocytes were loaded with increasing amount of purified LPL for 30 min, and LPL activity bound to the myocyte surface was determined (A, inset).  Myocytes were loaded with 200 ?g purified LPL for 30 min, and washed with PBS to remove unbound LPL.  Inserts with EC were placed on top, and the co-culture was exposed to normal glucose (NG, 5.5 mM) or high glucose (HG, 25 mM).  Medium was collected from the lower chamber after 30 min, and LPL activity determined (A).  After 2 and 4 h, LPL activity at the EC surface was also released and tested by incubating these cells with 8 U/ml heparin for 3 min (B).  In this co-culture, activation of RhoA was measured in myocytes after 30 min, and compared to cells that did not have EC in the top chamber (-EC) (C).  F to G actin ratio was also tested at 30 and 60 min (C, inset).  *P<0.05, compared to NG, n=3.     82    83  Figure 23 Heparanase is detected in cardiomyocytes.  To validate the purity of cardiomyocytes, equal amounts of protein from cardiomyocytes (Myo) and RAOEC (EC) were used to determine the presence of CD31.  Both cell lysates were also used to probe for latent (L-HEPA) and active heparanase (A-HEPA) by Western blot (A, left panel).  1 ?g total RNA from cardiomyocytes and EC was used for RT-PCR.  Amplified DNA products were run on an agrose gel in line with a DNA ladder (M), and a negative control (N, using water to replace PCR product) to detect heparanase gene expression (A, right panel).  Cardiomyocytes from control rats were isolated, plated and allowed to settle down.  Total cell lysates were then collected at 0 or 36 h, and probed for L-HEPA and A-HEPA using Western blot (B).  500 ng/ml purified latent heparanase was added to isolated myocytes and heparanase content in these cells was determined at the indicated times by Western blot.  To inhibit internalization, myocytes were also incubated with heparanase at 15 ?C for 15 min (C).  Results are the mean ? SE of 3 repeated experiments using different animals.  *Significantly different from 0 time point, P < 0.05.  **Significantly different from 0 time point, P < 0.01.  #Significantly different from results at 15 min at 37 ?C for L-HEPA.   84    85  Figure 24 Internalization of heparanase by cardiomyocytes is mediated by HSPGs. Isolated cardiomyocytes were treated with 500 ng/ml purified latent heparanase in the absence or presence of 10 IU/ml heparin for 15 min.  Cell lysates devoid of plasma membrane (-PM) were used to monitor internalization of latent heparanase (A).  To determine the purity of these cell lysates, 10 ?g protein from ?PM or total cell lysates (Total) was used to Western blot for the plasma membrane marker Na+-K+-ATPase (A, inset).  Isolated myocytes were untreated (UT) or pre-treated with 10 IU/L heparitinase III for 2 h, and incubated with 500 ng/ml purified latent heparanase for 15 min. Internalization of latent heparanase was determined using ?PM fraction (B).  Results are the mean ? SE of 3 repeated experiments using different animals.  **Significantly different from 0 time point, P < 0.01.  ##Significantly different from 15 min without heparin.  *Significantly different from untreated myocytes, P < 0.05.   86   87  Figure 25 Internalization of latent heparanase is caveolae-dependent and requires dynamin and tyrosine kinase activity.  Isolated cardiomyocytes were incubated with 350 mM sucrose (to inhibit clathrin-coated pit formation) or 1 ?g/ml filipin (to inhibit caveolae-dependent internalization) for 15 min before 500 ng/ml purified latent heparanase was added into the medium.  Internalization of latent heparanase was measured after 15 min using ?PM fractions (A).  The inhibitory effect of sucrose on clathrin-coated pit dependent internalization was validated by testing the internalization of epidermal growth factor receptor (EGFR) in myocytes treated with EGF in the presence or absence of sucrose (A, right inset).  In a different experiment, isolated myocytes were pre-treated with either 0-25 ?? dynasore (to inhibit dynamin activity, B) or 0-100 ?? genistein (to inhibit tyrosine kinase activity, C) for 1 h, and internalization of latent heparanase after 15 min determined using Western blot.  Results are the mean ? SE of 3 repeated experiments using different animals.  *Significantly different from untreated myocytes, P < 0.05.  **Significantly different from untreated myocytes, P < 0.01.   88   89  Figure 26 Following uptake exogenous latent heparanase is activated in lysosomes.  Isolated myocytes were incubated with 500 ng/ml purified latent heparanase for the indicated times.  Fractions rich in lysosomes were isolated to detect heparanase by Western blot.  Lysosomal-associated membrane protein 1 (Lamp1) was used as a marker to indicate equal loading of lysosomal protein (A).  Myocytes were pre-incubated with 200 ?? chloroquine before latent heparanase was added to the medium.  Heparanase in the lysosomal fraction was measured after 3 h (B).  Results are the mean ? SE of 3 repeated experiments using different animals.  *Significantly different from 0 time point or untreated myocytes, P < 0.05.  **Significantly different from 0 time point, P < 0.01.   90   91  Figure 27 Nuclear entry of heparanase is accompanied by increased gene expression of MMP-9. Isolated myocytes were incubated with 500 ng/ml purified latent heparanase for the indicated times.  Nuclear fraction was separated from cytosolic protein to detect heparanase by Western blot (A).  To validate the purity of the nuclear fraction, equal amount of protein from both cytosolic and nuclear fractions were Western blotted for GAPDH and histone H3 (A, right inset).  Total RNA was extracted from heparanase treated cardiomyoytes to measure gene expression of MMP-9 using RT-PCR (B).  Animals were made diabetic by injecting 55 mg/kg i.p. STZ and kept for 4 days (D55).  The amount of heparanase in the nuclear fractions of myocytes isolated from control and D55 heart was measured by Western blot (C).  MMP-9 gene expression was also determined in isolated myocytes from these animals (D).  Results are the mean ? SE of 3 repeated experiments using different animals.  *Significantly different from 0 time point or control, P < 0.05.  **Significantly different from 0 time point or control, P < 0.01.   92  Chapter 4: Discussion 4.1 Calnexin augments LPL maturation after moderate diabetes  Majority of FA provided to the heart come from breakdown of circulating TG, a process catalyzed by LPL located at the vascular lumen133.  In mice with cardiac overexpression of LPL, excess lipid uptake is evident, with development of dilated cardiomyopathy134.  Paradoxically, chronic cardiac depletion of LPL is also associated with a decrease in cardiac ejection fraction135.  Thus, cardiac LPL is of crucial importance for regulating lipid metabolism in hearts, and disturbing its innate function is sufficient to cause cardiomyopathy.  Diabetes is a unique metabolic disorder during which either an increase (moderate T1D) or decrease (severe T1D) in vascular LPL activity is observed.  Our data suggests that following acute hyperglycemia and moderate diabetes, more LPL is processed into an active dimeric form, whereas severe diabetes is associated with increased conversion of LPL into inactive monomers.   In T1D patients, insulin supplementation can never mimic the exquisite control of glucose by pancreatic insulin seen in normal individuals.  Multiple finger pricks and insulin injections (3-4/day) means poor patient compliance, repeated exposure to bouts of inadequate glucose control (and a shift in cardiac metabolism), and cardiovascular disease in the long term.  To imitate this poorly controlled T1D patient exposed to acute hyperglycemia, we used DZ.  Measurement of LPL protein expression in the heart revealed no change following acute hyperglycemia.  However, the drawback with this measurement is that it does not distinguish active LPL from its inactive monomeric form.  Using the heparin-sepharose column, our data for the first time show that acute hyperglycemia can indeed increase the amount of dimeric LPL in the heart.  We are unaware of a similar increase in dimeric LPL in other physiology 93  or pathology.  In fact, the reverse is often seen with lipid metabolism disorders.  In one case of human familial LPL deficiency, the Tyr262 --> His mutation of LPL gene results in shifting of dimeric LPL to monomers136.  In addition, in cell lines derived from animals with a mutation (cld) which impairs post-translational processing of LPL, a significant amount of inactive LPL is produced as aggregated monomers137.  The increased dimeric LPL in the whole heart following acute hyperglycemia correlated well with the elevated LPL activity at the vascular lumen.  Our data suggest that following a rapid rise in circulating glucose, a unique augmentation in LPL dimerization contributes towards increased coronary enzyme, delivering more FA to the heart.    Following stripping of dimeric LPL from the vascular lumen, more LPL dimers were still evident in the myocytes from acute diabetic animals.  These dimeric LPL could be located inside the ER, Golgi, secretion vesicles or at the myocyte surface.  Interestingly, the majority of dimeric LPL displayed resistance to endo H digestion47.  This suggested that following DZ, LPL processing is very efficient with more LPL completing its maturation in the ER before moving to the Golgi.  Early processing of LPL into a conformation competent for dimerization is suggested to involve chaperones like calreticulin/calnexin.  For example, in a transfected sf21 cell line, folding/dimerization of human LPL is only promoted when calreticulin is co-expressed48.  In addition, the impaired LPL processing seen in cld/cld mice is related to a decreased expression of calnexin138.  In our study, LPL in control cardiomyocytes was found associated with calnexin.  As interruption of this association by Cs hindered LPL dimerization and secretion, our data imply that calnexin is important for LPL processing in the heart.   94   The Rodrigues? lab has previously found that HG+PA increased LPL trafficking from the Golgi to myocyte surface58, 64.  The current data suggest that HG+PA are also responsible for enhanced LPL dimerization.  To test whether the effect of an acute diabetic environment on LPL processing is not a transient response, we turned to a more chronic diabetic model using D55 animals.  A significant increase in the amount of LPL dimers correlated to an increased expression of calnexin in D55 hearts.  As no change in LMF1 gene expression was observed, our data suggest that chronic diabetes may have a profound impact on the early processing events of LPL dimerization.  It is possible that the increased calnexin expression following chronic diabetes is related to the unfolded protein response (UPR), an adaptive mechanism that augments protein processing efficiency following ER stress139.  One of the three well-defined arms of UPR includes up-regulation of ER chaperone expression through activation of inositol-requiring kinase 1 (IRE-1) on the ER membrane140, 141.  It should be noted that acute exposure to HG+PA had no effect on calnexin expression.  Nevertheless, this environment could still alter the processing efficiency of LPL by affecting chaperone activity and accessibility in the ER through calcium and extracellular signal-regulated kinase (ERK-1).  Calcium is an important cofactor for protein folding in the ER, facilitating LPL dimerization, and can also impact the structural stability and substrate recognition of calnexin142, 143.  ERK-1, activated in an IRE-dependent manner can directly phosphorylate calnexin at Ser563, resulting in its recruitment to ribosomes, potentially rendering it more accessible to newly synthesized proteins144, 145.  The role of calcium and ERK-1 on LPL maturation, and how diabetes could potentially alter LPL processing through these pathways requires investigation. 95  4.2 Following severe diabetes Angptl-4 dissembles dimeric LPL   In contrast to D55, D100 animals have a more profound loss of pancreatic beta-cells and reduction in circulating insulin.  Under these conditions, excessive lipolysis and hepatic lipoprotein secretion increases serum FA and TG, respectively.  Thus, these doses of STZ represent a continuum of poor glycemic control, from moderate (D55) to severe (D100).  To avoid lipid overload, LPL activity at the coronary lumen decreased in D100 hearts, and this could be due to enzyme inactivation by conversion from a dimeric to monomeric conformation109.  Angptl-4, identified to bring about this conversion, is expressed in various tissues including white adipose tissue, liver, heart, and skeletal muscle106.  This secreted protein can function in the circulation, breaking down vascular LPL107.  Our data for the first time report a robust increase of Angptl-4 protein, both in the serum and heart tissue from D100 animals, an effect that was not seen in D55.  Given that the serum concentration of Angptl-4 in D100 was capable of inhibiting the activity of vascular LPL released from D55 hearts in vitro, we assumed that introduction of Angptl-4 to D55 hearts would give these hearts a D100 phenotype.  Indeed, a drop of LPL activity was observed at the vascular lumen in D55 hearts perfused with Angptl-4, an effect that correlated well with the inhibitory efficiency in vitro.  The reduction of coronary LPL activity could be due to the increased conversion of dimeric LPL into monomers, with subsequent detachment from the vascular binding sites.  This effect of Angptl-4 was also evident in control hearts.  Long chain FA, through activation of PPAR ?, has been reported to increase the transcription of Angptl-4 in skeletal muscle cells110.  In our study, as Angptl-4 expression in cardiomyocytes was also up-regulated following exposure to PA (more than 4 h), the decrease of LPL in D100 could be through a FA-Angptl-4 pathway.  As this effect was absent in DZ which had short term 96  hyperlipidemia (less than 4 h), our data suggest that ?turning off? LPL requires prolonged exposure to FA.  Whether the increased expression of Angptl-4 seen in D100 hearts could contribute towards inhibition of LPL at the coronary lumen is currently unknown.  Nevertheless, purified Angptl-4 was capable of directly inhibiting LPL in or on cardiomyocytes.  In addition, in mice overexpressing Angptl-4 specifically in the heart, a reduced cardiac LPL activity was found without affecting this enzyme in other LPL expressing tissues, implying a tight link between tissue-specific Angptl-4 and local or proximal vascular LPL activity106.  Overall, our data suggest that in moderate T1D, an exaggerated LPL processing to dimeric, catalytically active enzyme increases vascular LPL, delivering more FA to the heart when glucose utilization is compromised.  In severe diabetes, conversion of LPL to inactive monomers at the coronary lumen would impede TG hydrolysis and FA supply.  4.3 Using heparanase, endothelial cells regulate LPL secretion from cardiomyocytes    LPL secretion requires trafficking from an intracellular pool to myocyte surface HSPGs, followed by translocation to the vascular lumen.  Our previous studies have indicated that the trafficking component requires actin cytoskeleton polymerization, which can be achieved by RhoA activation64, 127.  In fact, when isolated hearts were perfused with LPA, a RhoA activator, increased coronary LPL activity was detected127.  RhoA is a small GTP-binding protein, with inactive RhoA associated with GDP and sequestered in the cytosol.  Upon activation, GDP is replaced by GTP and GTP-RhoA shifts to the plasma membrane.  Subsequently, through its downstream effector ROCK, activation of RhoA will ultimately induce actin cytoskeleton polymerization.  We observed that the increase in LPL activity at the coronary lumen in hyperglycemic DZ and diabetic D55 hearts were both accompanied by 97  RhoA activation.  As RhoA activation by LPA also promoted LPL secretion in cardiomyocytes through actin cytoskeleton polymerization, our data suggests that this GTP-binding protein is an important contributor to move LPL to the myocyte surface.    The mechanism by which LPL leaves the myocyte surface to move to the coronary lumen is not completely understood.  At the myocyte surface, LPL is sequestered to HSPGs via an ionic interaction.  Thus, its release is possible by negatively charged heparin.  However, cleavage of HS, or shedding of the extracellular part of HSPGs is a more likely event for LPL to leave the surface in vivo.  Interestingly, on exposure of cardiomyocytes to ECCM, LPL is released into the medium with a reciprocal decrease in surface LPL activity, suggesting that ECCM is likely releasing LPL from the myocyte surface.  An additional observation noticed with ECCM was its ability to assist in LPL replenishment when the surface pool was depleted.  We attributed this effect to its ability to activate RhoA in cardiomyocytes in a pattern similar to that seen with LPA.  In both cases, after reaching a peak, the amount of GTP-RhoA declined as it is normally converted to a GDP-bound form following activation of downstream effectors146.  Intriguingly, heparin had no effect on RhoA or LPL replenishment, implying that simple displacement of LPL is not sufficient to trigger the replenishment process, and that ECCM is likely altering the conformation of HSPGs.    An enzyme of special relevance that is known to cleave HS of HSPGs is heparanase.  This enzyme is synthesized in the EC as a latent 65 kDa form, which possesses no catalytic activity.  Following its secretion, it binds to EC surface HSPGs and is internalized into lysosomes147.  There it is cleaved into a 50 kDa active form and stored until it is secreted in response to stimulation such as adenosine, ADP, and ATP74, 148.  In the present study, increased secretion of both latent and active heparanase into the medium was also observed 98  when EC were exposed to high glucose.  As both forms of heparanase were augmented in the interstitial space of hearts from DZ animals, it suggests that hyperglycemia is effective in triggering coronary endothelial heparanase release.  The Rodrigues? lab has previously demonstrated that high glucose induced secretion of active heparanase from EC occurs through activation of P2Y receptors that initiates stress fiber formation (across which heparanase-containing cargos are transported) and disruption of cortical actin cytoskeleton (to allow heparanase to be released into the extracellular space)149.  Whether the same mechanism applies to release of latent heparanase is unknown as we cannot rule out the possibility that high glucose could also block endocytosis of latent heparanase from the EC surface, or is capable of directly detaching the latent enzyme from surface HSPGs.  Irrespective of the release mechanism, our study is the first to demonstrate that it is heparanase within ECCM which is responsible for detachment of LPL from the myocyte surface.    Although active heparanase is predicted to release surface LPL by cleaving HS, latent enzyme could also facilitate LPL release.  As expected, only active heparanase initiated release of LPL from the myocyte surface.  Surprisingly, this release was not accompanied by RhoA activation.  In fact, it was latent heparanase that accounted for the RhoA activation seen with ECCM.  Growing evidence has demonstrated that heparanase also possesses activity-independent effects, for example, activation of Src during cell adhesion150.  Hence, our data suggests that active heparanase detaches LPL from the myocyte surface HSPGs, whereas latent heparanase activates RhoA to move intracellular LPL to replenish this released reservoir.  Studies on HSPG-mediated signaling have focused on syndecan-4, a transmembrane HSPG with its cytoplasmic domain capable of activating and stabilizing 99  PKC?, and syndecan-4-dependent PKC? activation has been indicated in FGF signaling151, 152.  Moreover, neuronal Thy-1 induced cell spreading requires RhoA-mediated actin cytoskeleton polymerization, a suggested downstream event following syndecan-4-dependent PKC? activation153.  Our study also indicated that the RhoA activation induced by latent heparanase is mediated by PKC?.  In U87 MG human glioma cells, clustering of syndecan-4 by latent heparanase or an anti-heparanase antibody was sufficient to induce cell spreading, a process involving PKC activation154.  In addition, upon binding of phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5P)(2)) to its cytoplasmic domain, oligomeriation of syndecan-4 was promoted with a subsequent up-regulation of PKC?155, 156.  In this study, it is likely that clustering of syndecan-4 with latent heparanase triggers RhoA activation.  It is possible that latent heparanase does this by acting as a bridging factor by binding to HS.  This idea was supported by our results using heparinase III that eliminates HS and attenuated RhoA activation by latent heparanase.  Altogether, results from our study suggest that active and latent heparanase work cooperatively to promote LPL secretion following diabetes (Figure 21D).   4.4 Heparanase from EC can be taken up by cardiomyocytes to affect gene expression   It was surprising that we could detect heparanase in cardiomyocytes in the absence of gene expression in these cells.  Freshly isolated myocytes had a significant amount of active heparanase that decreased following 36 h in vitro culturing.  As the half-life of active heparanase is approximately 30 h147, this result suggests that the presence of heparanase in cardiomyocytes is not from de novo synthesis, but likely due to extracellular uptake.  Indeed, on exposure of cardiomyocytes to exogenous latent heparanase, we demonstrated 100  its rapid and robust uptake.  In CHO cells, endocytosis of exogenous latent heparanase involves low density lipoprotein receptor-related proteins, mannose 6-phosphate receptors, and HSPGs157.  Although our data in cardiomyocytes strongly indicates an HSPG-dependent mechanism of internalization, we cannot rule out the contributive effects of other receptors in this uptake process.  Regarding HSPGs, it is believed that their clustering leads to internalization of bound ligands, as observed in CHO cells, where latent heparanase triggers clustering of both syndecan-1 and -4 followed by rapid internalization of the heparanase-HSPGs complex154.  As we have previously reported clustering of syndecan-4 when cardiomyocytes are exposed to latent heparanase158, it is possible that a heparanase-HSPGs complex is also responsible for endocytosis of this enzyme in cardiomyocytes.  Of the two novel endocytotic pathways, HSPGs are more involved in caveolae-dependent rather than clathrin-mediated endocytosis.  For example, syndecan-1 has been shown to mediate apoE-VLDL uptake in the human fibroblast cell line (GM00701), a process that was clathrin-independent, but inhibited by nystatin, an inhibitor of the lipid raft-caveolae endocytosis pathway159.  Our data demonstrated that internalization of latent heparanase by cardiomyocytes relies on caveolae to form endocytotic vesicles and dynamin for fission of these vesicles from the plasma membrane.  The participation of tyrosine kinase in this process has also been seen in the syndecan family mediated endocytosis160, 161.  It is believed that clustering would lead to redistribution of syndecans to lipid rafts, where their cytoplasmic domain could be phosphorylated by tyrosine kinase126, 162.  Recruitment of cortactin to phosphorylated syndecans promotes actin polymerization at the actin cortex, thereby bringing HSPGs-mediated endocytotic vesicles into the cells163.  Overall, it is 101  possible that in response to high glucose, the latent heparanase released from EC enters cardiomyocytes through an HSPG-dependent caveolae pathway.   It is interesting that cells like fibroblasts, which do not express heparanase can also take up latent heparanase164, and this can be converted into active heparanase following lysosomal processing74.  Our results also indicate that a similar mechanism applies to latent heparanase taken up by cardiomyocytes.  In EC, active heparanase stored in lysosomes can be secreted in response to high glucose, or enters the nucleus in the presence of fatty acid165.  Our data also demonstrated entry of both latent and active heparanase into the nucleus.  Heat shock protein 90 (Hsp90) is a suggested chaperone for escorting active heparanase from lysosomes to nucleus in both HL-60 cells166 and bCAECs165, resulting in cell differentiation and upcoupling of glucose metabolism, respectively.  As we were unable to detect an association between Hsp90 and heparanase (data not shown), the mechanism which drives the nuclear entry of heparanase in cardiomyocytes is currently unknown.  Regardless, gene expression of MMP-9 increased significantly in heparanase treated myocytes.  As similar results were also obtained in myocytes from D55 diabetic rats which showed accumulation of active heparanase in the nucleus together with a robust increase in MMP-9 expression, our data imply that gene expression in cardiomyocytes could be affected by heparanase secreted from EC.  Interestingly, gene upregulation of MMP-9 has been observed in several cancer cell lines upon transfection of heparanase, transforming them to an aggressive phenotype167.  As MMP-9 degrades components of the extracellular matrix such as collagens168, upregulation of its expression, as shown in our D55 cardiomyocytes, could contribute to heart dysfunction in these diabetic animals in the long term.  Other studies have reported a correlation between elevated level of MMP-9 and collagen accumulation in the hearts from STZ induced diabetic 102  rats169.  In fact, in diabetic mice, deletion of MMP-9 gene effectively reduced cardiac fibrosis and increased survival of cardiac stem cells170.  Further studies are required to investigate the relationship between EC derived heparanase and MMP-9 expression in cardiomyocytes.   In summary, heparanase is a key mediator of the cross-talk between EC and cardiomyocyte to increase LPL secretion following hyperglycemia.  This idea was confirmed in DZ induced hyperglycemia and the high glucose co-culture system, with both experimental models exhibiting increased heparanase secretion together with LPL amplification at the apical side of EC.  Additionally, it is possible that heparanase secreted from EC can be taken up by cardiomyocytes to modulate MMP-9 expression, driving the onset of fibrosis and development of diabetic cardiomyopathy.   103  Chapter 5: Conclusions and future directions 5.1 Conclusions  In summary, following diabetes, recruitment of LPL to the vascular lumen could represent an immediate compensatory response to guarantee FA supply.  In acute hyperglycemia and moderate diabetes, ER chaperone calnexin promoted LPL processing to dimeric, catalytically active enzyme, thereby increasing functional LPL to deliver more FA to the heart when glucose utilization is compromised.  In severe chronic diabetes, to avoid lipid oversupply, FA-induced expression of angiopoietin-like protein 4 leads to conversion of LPL to inactive monomers at the coronary lumen to impede TG hydrolysis.  However, both compensatory processes that alter the innate function of LPL at the vascular lumen would disturb heart metabolism, which could be the forerunner for cardiomyopathy seen with this disease (Figure 28).  Besides intrinsic factors from cardiomyocytes, endothelial heparanase is a key mediator to increase LPL secretion following hyperglycemia.  Through cleaving HSPGs, active heparanase liberates LPL from myocytes surface en route for secretion, whereas latent heparanase stimulates replenishment of LPL by clustering of myocyte surface syndecan-4 to mediate RhoA-dependent actin cytoskeleton polymerization.  The end result would be a continuous augmented secretion of LPL to the vascular lumen, which provides overwhelming amount of FA that could lead to lipotoxicity to the heart (Figure 29).  Additionally, our data suggests that heparanase can be taken up by cardiomyocytes to modulate MMP-9 expression, 104  driving the onset of fibrosis and development of diabetic cardiomyopathy.  Pharmaceutical manipulation of this process, for instance inhibition of heparanase secretion/activity, could potentially provide an additional strategy to limit FA delivery to the heart, and cardiomyopathy observed with chronic diabetes. 5.2 Future directions  To further investigate the relationship between endothelial heparanase and cardiomyocyte gene expression, the following studies are ongoing:  We demonstrated that cardiac MMP-9 expression increased following nuclear entry of active heparanase.  To see whether the gene regulatory effect is a result of heparanase uptake, myocytes can be treated with latent heparanase in the presence of heparin or filipin which interfere with its uptake.  In vivo studies can also be done using heparanase knock-out mice.  They can be made diabetic by multiple, low-dose injection of STZ.  Following hyperglycemia, levels of nuclear heparanase content and MMP-9 expression in the cardiomyocytes will be compared to those of their wild type counterparts.   Another question is the mechanism behind the gene regulatory effect.  As mentioned before, cleavage of nuclear HSPGs and the subsequent increase in HAT activity is the first proposed pathway.  Hence, multiple approaches could be used to block this pathway in vitro, including usage of: a) chloroquine that inhibits processing of latent to active heparanase, b) SST0001, a compound that inhibits heparanase activity, and c) anacardic acid, an inhibitor of HAT activity.  Using another strategy, diabetic rats can be treated with heparanase inhibitor 105  SST0001 to determine whether the gene regulatory effect of heparanase is activity-dependent.  In addition to that, ChIP-on-chip studies have revealed interaction of nuclear heparanase with promoter and transcribed regions of genes167, modulating histone 3 lysine 4 (H3K4) methylation.  Whether this newly found pathway could explain our results in cardiomyocytes remain to be investigated.  In our studies, we only focused on MMP-9 gene expression since it is a well-studied target gene of heparanase in tumor cell lines, which is also involved in diabetic cardiomyopathy.  This study should be broadened using RNA-sequencing to explore more target gene in cardiomyocytes, and how they affect the pathogenesis of diabetic cardiomyopathy.  For example, transforming growth factor ?3, cardiac sarcoplasmatic reticulum Ca2+ ATPase, cardiac Na+/Ca2+ exchanger 1, etc..    106   Figure 28 Moderate and severe diabetes have a differential impact on LPL at the coronary lumen.   In moderate diabetes, more monomeric LPL is processed into dimeric, catalytically active enzyme with the help of the ER chaperone calnexin.  This increases LPL at the vascular lumen, breaking down more circulating TG to deliver increased FA to the heart when glucose utilization is compromised.  In severe diabetes, when FA are excessive, a FA-induced expression of Angptl-4 leads to conversion of LPL to inactive monomers at the coronary lumen to impede TG hydrolysis.  EC: endothelial cells; ER: endoplasmic reticulum; Nu: nucleus. 107    Figure 29 Endothelial heparanase is a key mediator to increase LPL secretion following hyperglycemia.  Two forms of heparanase are secreted when endothelial cells are exposed to high glucose.  Through cleaving HSPGs, active heparanase liberates LPL from myocytes surface en route for secretion, whereas latent heparanase stimulates replenishment of LPL.  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