text:
ENDOTHELIAL CELLS MANAGE FATTY ACID DELIVERY TO THE CARDIOMYOCYTES FOLLOWING DIABETES by Amy Pei-Ling Chiu 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) September 2017 © Amy Pei-Ling Chiu, 2017 ii ABSTRACT In the diabetic heart, there is excessive dependence on fatty acid (FA) utilization to generate ATP. Lipoprotein lipase (LPL)-mediated hydrolysis of circulating triglyceride is suggested to be the predominant source of FA for cardiac utilization during diabetes. In the heart, LPL is produced in the cardiomyocytes and is transferred by its transporter glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1) to the apical side of the endothelial cell (EC), where the enzyme is functional. We tested whether EC responds to hyperglycemia by increasing GPIHBP1. Streptozotocin diabetes increased cardiac LPL activity and GPIHBP1 gene and protein expression. Exposure of EC to high glucose-induced GPIHBP1 expression and amplified LPL shuttling across these cells. This effect coincided with an elevated secretion of heparanase, which can promote secretion of vascular endothelial growth factor (VEGF) from EC and cardiomyocytes. Recombinant VEGF induced EC GPIHBP1 mRNA and protein expression through activation of Notch signaling, which encompassed delta-like ligand 4 (DLL4) augmentation and nuclear translocation of the Notch intracellular domain. In addition, high glucose-induced secretion of heparanase is taken up by the cardiomyocyte to stimulate matrix-metalloproteinase (MMP) 9 expression and conversion of latent to active transforming growth factor-β (TGFβ). In the cardiomyocyte, TGFβ activation of RhoA enhances actin cytoskeleton rearrangement to promote LPL trafficking and secretion onto cell surface heparan sulfate proteoglycans. In the EC, TGFβ signaling promotes mesodermal homeobox 2 (Meox2) translocation to the nucleus that increases the expression of GPIHBP1, which facilitates movement of LPL to the vascular lumen. Collectively, EC, as the first responders to hyperglycemia, can release heparanase to liberate myocyte VEGF, which activates EC Notch signaling to facilitate GPIHBP1-mediated translocation of LPL across EC. Heparanase also induced MMP9 mediated activation of TGFβ. Its action on the cardiomyocyte to promote movement of LPL, together with its action on the EC to facilitate LPL shuttling are mechanisms that accelerate FA utilization by the diabetic heart. Gaining more insight into the iii mechanisms by which cardiac LPL is regulated may assist other researchers in devising new therapeutic strategies restore metabolic equilibrium, curb lipotoxicity, and help prevent or delay heart dysfunction characteristic of diabetes. iv LAY SUMMARY More than 9 million Canadians live with diabetes or pre-diabetes. In people with diabetes, inadequate pharmaceutical management predisposes the patient to heart failure, which is the leading cause of diabetes-related deaths. One instigator for this cardiac dysfunction is changed in fuel utilization by the heart. A key perpetrator that may be responsible for organizing this metabolic disequilibrium is lipoprotein lipase (LPL), the enzyme responsible for providing fat to the hearts. Using animal models and examining heart cells, diabetes alters the functioning of LPL to modulate fat delivery to the heart. This effect is geared to help the heart initially; however, the heart's uncontrolled use of fat as fuel can be dangerous and may lead to heart disease or even death. This study will help in gaining insights into the mechanisms by which cardiac LPL is regulated, and may be significant in understanding how to prevent or delay diabetes-related heart disease. v PREFACE All of the work presented in chapter 1, 2, 3 and 4 were conducted in Faculty of Pharmaceutical Sciences at the University of British Columbia. Some chapters have been published in the following manuscripts and review article: 1. Chiu AP, Wang F, Lal N, Wang Y, Zhang D, Hussein B, Wan A, Vlodavsky I, and Rodrigues B. Endothelial cells respond to hyperglycemia by increasing the LPL transporter GPIHBP1. American Journal of Physiology: Endocrinology and Metabolism, 2014. I am the first author responsible for conceiving the idea, designed the experiments, and generated most of the data. My supervisor Dr. B. Rodrigues, former laboratory member Dr. Y. Wang and I designed the experiments. Dr. B. Rodrigues and I wrote the manuscript. Dr. I. Vlodavsky provided materials for research and assist with valuable suggestions. F. Wang (Figure 5), N. Lal (Figure 5), Dr. Y. Wang (Figure 7), D. Zhang (Figure 8), B. Hussein (Figure 10) and A. Wan (Figure 10) helped perform some of the experiments, edited and revised the manuscript. 2. Chiu AP, Wan A, Lal N, Zhang D, Wang F, Vlodavsky I, Hussein B, and Rodrigues B. Cardiomyocyte VEGF regulates endothelial cell GPIHBP1 to relocate lipoprotein lipase to the coronary lumen during diabetes. Arteriosclerosis, Thrombosis, and Vascular Biology, 2016. As the first author, I conceived the idea, designed the experiments, and generated most of the data, and wrote the manuscript. A. Wan (Figure 13), N. Lal (Figure 20), D. Zhang (Figure 20), F. Wang (Figure 20) and B. Hussein (Figure 18) helped with performing some experiments and data generation. Dr. I. Vlodavsky provided materials for some of the experiment. Dr. B. Rodrigues helped write and edit the manuscript. 3. Chiu AP, Wan A, and Rodrigues B. Cardiomyocyte-Endothelial Cell Control of Lipoprotein Lipase. Biochimica et biophysica acta- Molecular and Cell Biology of Lipids, 2016. A. Wan and I collected the relevant literature and designed the diagrams. I wrote the article. Dr. B. Rodrigues helped write and edit the article. vi This investigation adheres to the Guide for the Care and Use of Laboratory Animals published by the U.S. National Institutes of Health and the University of British Columbia, and was approved by Animal Care Committee at the University of British Columbia (Certificate No. A13-0098). vii TABLE OF CONTENTS ABSTRACT .......................................................................................................................................... ii LAY SUMMARY ................................................................................................................................ iv PREFACE ..............................................................................................................................................v TABLE OF CONTENTS .................................................................................................................... vii LIST OF FIGURES ............................................................................................................................. xi LIST OF ABBREVIATION AND ACRONYMS ............................................................................. xiii ACKNOWLEDGEMENTS ............................................................................................................... xiv DEDICATION .....................................................................................................................................xv CHAPTER 1: INTRODUCTION ..........................................................................................................1 1.1 Heart metabolism ......................................................................................................................1 1.2. Lipoprotein lipase (LPL) ...........................................................................................................1 1.3. Heparanase ................................................................................................................................4 1.3.1 Active heparanase ................................................................................................................5 1.3.2 Latent heparanase .................................................................................................................5 1.4. Glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1) .........................................................................................................................................6 1.5 Mesodermal homeobox 2 (Meox2) ...........................................................................................7 1.6 Diabetic cardiomyopathy ..........................................................................................................8 1.6.1 Changes in cardiac metabolism during diabetes ..................................................................8 1.6.2 Heart LPL in human diabetes ..............................................................................................9 1.6.3 Heart LPL in animal diabetes ............................................................................................10 1. 7 Hypothesis and research objectives .........................................................................................12 CHAPTER 2: METHODS ...................................................................................................................16 2.1 Materials ..................................................................................................................................16 viii 2.2 Experimental animals ..............................................................................................................17 2.3 Heart perfusion and isolation of cardiomyocytes ....................................................................17 2.4 Endothelial cell culture ............................................................................................................17 2.5 Endothelial cell and cardiomyocyte co-culture .......................................................................18 2.6 Isolation of heart endothelial cells ..........................................................................................18 2.7 LPL transport and activity assay .............................................................................................18 2.8 Immunofluorescence ...............................................................................................................19 2.9 Western blot ............................................................................................................................20 2.10 RNA isolation and real-time PCR ...........................................................................................20 2.11 In vitro lipolysis of VLDL ......................................................................................................21 2.12 Cytokine array .........................................................................................................................21 2.13 Plasma measurements .............................................................................................................21 2.14 VEGF ELISA ..........................................................................................................................22 2.15 MMP9 activity assay ...............................................................................................................22 2.16 Plasma measurements of TGFβ ...............................................................................................22 2.17 GLISA assay ...........................................................................................................................23 2.18 Actin polymerization ...............................................................................................................23 2.19 Statistical analysis ...................................................................................................................23 CHAPTER 3: RESULTS .....................................................................................................................26 3.1 Endothelial cells respond to hyperglycemia by increasing the LPL transporter GPIHBP1 ....26 3.1.1 Cardiac LPL and GPIHBP1 after STZ diabetes ................................................................26 3.1.2 Endothelial GPIHBP1 following cell passage and influence of high glucose ...................26 3.1.3 GPIHBP1 associated transfer of LPL across the endothelial monolayer ..........................27 3.1.4 GPIHBP1 expression is linked to endothelial content of heparanase ................................27 3.1.5 Cardiomyocytes induce endothelial cell GPIHBP1 ...........................................................28 ix 3.1.6 Heparanase released platelet-derived growth factor from myocytes and EC can induce GPIHBP1 expression ...................................................................................................................28 3.2 Cardiomyocyte regulates endothelial cell GPIHBP1 to relocate lipoprotein lipase to the coronary lumen during diabetes .......................................................................................................29 3.2.1 High glucose alters the expression of endothelial GPIHBP1 ............................................29 3.2.2 The effect of heparanase on GPIHBP1 expression is associated with VEGF ...................29 3.2.3 Cardiomyocyte surface bound VEGF stimulates endothelial cell GPIHBP1 ....................30 3.2.4 GPIHBP1 induction by VEGF is through a DLL4-Notch signaling pathway ...................31 3.2.5 VEGF induced GPIHBP1 expression is decreased by inhibition of Notch signaling .......31 3.2.6 Reduction in cardiomyocyte VEGF following diabetes is associated with a decrease in GPIHBP1 expression and LPL activity at the vascular lumen ....................................................32 3.3 Dual effects of hyperglycemia on endothelial cells and cardiomyocytes to enhance coronary LPL activity ....................................................................................................................................33 3.3.1 Alteration in endothelial Meox2 is linked to changes in cardiac LPL following diabetes 33 3.3.2 High glucose is a potential stimulus to induce endothelial Meox2 and increase GPIHBP1 expression. ...................................................................................................................................33 3.3.3 Heparanase secretion in response to high glucose stimulates MMP9 expression and TGFβ activation. .....................................................................................................................................34 3.3.4 TGFβ can promote cardiomyocyte LPL secretion. ............................................................35 3.3.5 TGFβ induced Meox2 enhances EC GPIHBP1 and promotes LPL translocation. ...........35 CHAPTER 4: DISCUSSION ...............................................................................................................66 4.1 Endothelial cells respond to hyperglycemia by increasing the LPL transporter GPIHBP1 ....67 4.2 Cardiomyocyte VEGF regulates endothelial cell GPIHBP1 to relocate lipoprotein lipase to the coronary lumen during diabetes .................................................................................................69 4.3 Dual effects of hyperglycemia on endothelial cells and cardiomyocytes to enhance x coronary LPL activity .....................................................................................................................72 CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS .......................................................78 5.1 Conclusions .............................................................................................................................78 5.2 Future directions ......................................................................................................................79 REFERENCES ....................................................................................................................................82 xi LIST OF FIGURES Figure 1. Communication between the EC and cardiomyocyte promotes LPL transfer to the vascular lumen. ...................................................................................................................................................14 Figure 2. General characteristics of different models of diabetes. ......................................................15 Figure 3. Isolated heart endothelial cells and rat heart microvascular endothelial cells express CD31 and Von Willebrand Factor ..................................................................................................................24 Figure 4. Primary heart endothelial cell and endothelial cell lines express Meox2 ...........................25 Figure 5. STZ diabetes increases cardiac LPL activity and GPIHBP1 expression ............................37 Figure 6. Passaging of endothelial cells cause loss of GPIHBP1 which can be induced on exposure to high glucose. ....................................................................................................................................38 Figure 7. GPIHBP1 induced by high glucose increases LPL shuttling across endothelial cell monolayers. ..........................................................................................................................................39 Figure 8. Endothelial heparanase can increase GPIHBP1 gene and protein expression. ...................41 Figure 9. Loss of endothelial GPIHBP1 expression can be restored on co-culture with myocytes. ..42 Figure 10. EC and cardiomyocyte HSPG anchored proteins can be liberated into culture medium ..43 Figure 11. EC and myocyte HSPG bound PDGF can augment GPIHBP1. .......................................44 Figure 12. High glucose induced increase in GPIHBP1 expression is associated with secretion of heparanase. ...........................................................................................................................................45 Figure 13. Heparanase released VEGF is an effective stimulator of GPIHBP1 expression. ..............47 Figure 14. Cardiomyocyte HSPG bound VEGF is crucial for maintaining endothelial cell GPIHBP1 expression. ...........................................................................................................................................49 Figure 15. VEGF induced GPIHBP1 expression is through DLL4-Notch signaling. ........................51 Figure 16. DLL4-Notch pathway downstream target genes were upregulated following DLL4 stimulus. ...............................................................................................................................................52 Figure 17. Inhibition of DLL4 signaling prevents the VEGF-induced GPIHBP1 expression ............53 xii Figure 18. Inhibition of Notch signaling downregulates VEGF induced GPIHBP1 expression. .......54 Figure 19. General characteristics of different models of diabetes .....................................................55 Figure 20. Reducing VEGF in diabetes results in lower GPIHBP1 expression. ................................56 Figure 21. Diabetes increases cardiomyocyte VEGF secretion ..........................................................57 Figure 22. Augmentation of LPL activity in the diabetic heart is associated with an increase of endothelial Meox2. ..............................................................................................................................58 Figure 23. Endothelial Meox2 induced by high glucose increases GPIHBP1 expression. ................60 Figure 24. Heparanase induced MMP9 expression is associated with the conversion of latent to active TGFβ in the diabetic heart. ........................................................................................................62 Figure 25. TGFβ induced RhoA-mediated actin cytoskeleton polymerization increases cardiomyocyte LPL secretion. .............................................................................................................63 Figure 26. TGFβ activates downstream smad signaling. ....................................................................64 Figure 27. TGFβ activation of Meox2 enhances GPIHBP1 expression and promotes LPL translocation. ........................................................................................................................................65 Figure 28. DLL4 binding to the Notch receptor triggers the Notch pathway to regulate gene expression of GPIHBP1. ......................................................................................................................75 Figure 29. EC increases its GPIHBP1 in response to hyperglycemia. ...............................................76 Figure 30. EC communicates with the cardiomyocyte to increase LPL transfer to the vascular lumen following high glucose .........................................................................................................................77 Figure 31. Pathological oscillations in coronary luminal LPL following diabetes can lead to cardiomyopathy. ...................................................................................................................................81 xiii LIST OF ABBREVIATIONS AND ACRONYMS AMPK AMP-activated protein kinase Angptl angiopoietin-like proteins BCAEC bovine coronary artery DAG-PKC diacylglycerol-protein kinase C DLL4 delta-like ligand 4 EC endothelial cell FA fatty acid FBS fetal bovine serum FOXO1 forkhead box protein O1 GLUT glucose transporter GPIHBP1 glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 HAT histone acetyltransferase HG high glucose HS heparan sulfate HSPG heparan sulfate proteoglycans Hsp25 heat shock protein 25 LMF1 lipase maturation factor 1 LPA lysophosphatidic acid LPL lipoprotein lipase Ly6 cysteine-rich lymphocyte antigen 6 Meox2 mesodermal homeobox 2 MMP9 matrix metalloproteinase 9 PHPLA post-heparin plasma lipolytic activity PKD protein kinase D PIPLC phosphatidylinositol-specific phospholipase C RAOEC rat aortic endothelial cells RHMEC rat heart microvascular endothelial cells STZ streptozotocin TG triglyceride TGFβ transforming growth factor-β T1D type 1 diabetes T2D type 2 diabetes VEGF vascular endothelial growth factor VLDL very low-density lipoprotein xiv ACKNOWLEDGEMENTS I would like to express my sincere gratitude to my supervisor Dr. Brian Rodrigues for giving me the opportunity to join his laboratory. He offered the best training to ensure everyone can be a great scientist. He also dedicated his time to teach us not only in sciences but also his formula- how to be competitive and have a successful career. I am grateful to have Dr. Rodrigues as my mentor and his support in my career. I also would like to thank my committee members: Dr. Bruce Verchere, Dr. Roger Brownsey, Dr. Ismail Laher, and chair Dr. David Grierson. Your intelligence and expertise have broadened my view and strongly encouraged me to pursue excellence. Your valuable suggestions and comments also have strengthened my work. I appreciate my examiners, Dr. Judy Wong, Dr. Dan S. Luciani and Dr. Bruce Verchere, for your kind support and feedback on my work. Your comments make my work substantially better. I thank you for your time and effort. I am grateful to have support from Dr. Israel Vlodavsky. Some works are achievable because of your kindness to share your resource and experiences with us. I also would like to thank Canadian Diabetes Association for the financial support during my graduate program. I am also grateful to be surrounded by awesome people who encourage, help, support, and strengthen me in every aspect during my program. I thank Dr. Y. Wang, Andrea Wan, Bahira Hussein, Denise Bierende, and Nathaniel Lal for offering great bits of help and supports at work and life. Finally, I thank my families and friends (Singing Ho and Nachi Hsu) for your unconditional love and support to make it possible. xv DEDICATION For my parents and brothers 1 CHAPTER 1: INTRODUCTION 1.1 Heart metabolism Fatty acids (FA) have numerous biological functions and include the formation of membranes, participation as energy substrates, and initiation of signal transduction. Because of their limited water solubility, long chain FA must undergo esterification with glycerol to form triglycerides (TG), which make up a significant portion of circulating lipoprotein TG. It is these particles that deliver the FA to the underlying tissues. This process relies on lipoprotein lipase (LPL)1, an enzyme that hydrolyzes TG-rich lipoproteins (VLDL and chylomicrons) to FA in the vascular lumen, and is termed lipolysis. The heart is a high-energy organ that has flexibility in utilizing multiple substrates for energy production, including FA, carbohydrates, and ketone bodies, to maintain its function2. In the normal heart, nearly 70% of ATP is produced from FA oxidation, with the remainder being provided by glucose and lactate3. Although the heart has a high demand for FA, it has a limited capacity to synthesize FA, and thus relies on an exogenous FA source. This includes release from adipose tissue and transport to the heart after complexing with albumin4, provision through the breakdown of endogenous cardiac TG stores5, and lipolysis of circulating TG-rich lipoproteins to FA by LPL positioned at the EC surface of the coronary lumen6. Given that greater than 90% of plasma FA is contained within lipoprotein-TG and the heart has the most robust expression of LPL, lipolysis of TG-rich lipoproteins mediated by LPL is suggested to be a principal source of FA for cardiomyocyte metabolism1. Interestingly, in the heart, LPL is predominantly produced in the cardiomyocytes and is transferred to the apical side of the endothelial cell (EC), where the enzyme functions7, 8. 1.2 Lipoprotein lipase (LPL) Although LPL is strategically located at the EC surface of the coronary lumen, ECs have limited LPL expression9. This enzyme is predominantly synthesized and processed in the cardiomyocyte7, 9. In this cell type, LPL is synthesized as an inactive monomer that enters the endoplasmic reticulum 2 (ER) and undergoes glycosylation at the Asn residue in the N-terminal domain for its catalytic activity10. It is further processed by calnexin/calreticulin, chaperones that regulate nascent LPL folding into a proper tertiary structure qualified for dimerization11. The two inactive monomeric LPL are then non-covalently assembled in a head-to-tail fashion with the help of lipase maturation factor 1 (LMF1) into a dimeric enzyme12-16. In ER, LPL forms a complex with LMF1 and Sel-1 suppressor of lin-12-like protein (Sel1L), an essential adaptor protein in the ER-associated degradation. This structure stabilizes LPL and enables it to exit the ER as a dimer17. Without LMF1 and Sel1L, LPL forms aggregates and is retained in the ER18, 19 for degradation by autophagy. Defects in ER-LPL maturation results in familial hyperchylomicronemia, a form of hypertriglyceridemia that contributes to the development of heart disease and metabolic syndrome in humans13. After dimeric LPL exits the ER, it is further sorted into secreted vesicles in the Golgi. The exocytosis of the LPL secretory vesicle from Golgi requires protein kinase D (PKD)20, 21. In addition to PKD, AMP-activated protein kinase (AMPK) is a cellular energy sensor that also regulates LPL22. In the heart, AMPK switches on coronary LPL activity in rats undergoing fasting, mostly through a post-translational mechanism. Specifically, by phosphorylating heat shock protein 25 (Hsp25), AMPK causes Hsp25 dissociation from actin monomers, leading to actin cytoskeleton polymerization and building of a bridge to transport LPL-containing vesicles from the Golgi to the plasma membrane23. In this way, intracellular LPL is moved to the myocyte surface heparan sulfate proteoglycans (HSPG). HSPG are ubiquitous macromolecules present in every tissue compartment and in particular, the extracellular matrix, cell surface, intracellular granules, and nucleus24. They are composed of a core protein to which several linear heparan sulfate (HS) side chains are covalently linked, and function, not only as structural proteins but also as anchors25. The latter property is implicitly used to bind chemokines, coagulation factors, and enzymes like LPL and growth factors26. Attachment of these 3 bioactive proteins is an intelligent arrangement, providing the cell with a rapidly accessible reservoir, precluding the need for de novo synthesis when the requirement for protein is increased. Following detachment from this temporary docking site, LPL translocates across the interstitial space and binds to its transporter glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1), which facilitates LPL relocation from the basolateral to the apical (luminal) side of the EC8. Out here, it can also act as a platform to bind lipoproteins27. GPIHBP1 allows LPL to actively metabolize the lipoprotein-TG core, thereby liberating FAs that are transported to the cardiomyocytes (Figure 1). Multiple factors regulate LPL at the vascular lumen, with FA being a key modulator of LPL activity. Specifically, FA may decrease coronary LPL in multiple ways: a) displacing LPL from EC surface binding sites for degradation in the liver, b) directly inactivating LPL enzyme activity28, and c) impairing LPL vesicular trafficking to the myocyte surface through the caspase-3-mediated cleavage of PKD29. In addition to FA, angiopoietin-like proteins (Angptl), especially Angptl3 and Angptl4 can also modulate LPL activity30. They are composed of a NH2-terminal coiled-coil and a C-terminal fibrinogen-like domain31. Angptl3 is mainly expressed in the liver and is considered to promote proprotein convertase that can disassociate dimeric LPL on cell surface32. However, in humans, the level of Angptl3 is not related to that of post-heparin plasma lipase activity and plasma triglycerides33. Angptl4 is widely expressed in the liver, white adipose tissue, skeletal muscle, and heart34. Unlike Angptl3, Angptl4 forms disulfide-linkage oligomers in the NH2-terminal coiled-coil domain that is essential for its stability and LPL's inhibitory effect35. The secreted N-terminal oligomers function in circulation, converting dimeric LPL at the vascular lumen into inactive monomers36. In response to elevated FA and PPARδ activation, Angptl4 expression is increased in skeletal muscle37. Interestingly, when dimeric LPL transfers onto EC and complexes with GPIHBP1, this structure appears to protect it from inactivation by Angptl4. A more recent study reveals that 4 Angptl4 was capable of binding and inactivating LPL even when it is complexed to GPIHBP1 on the surface of endothelial cells38, 39. LPL synthesis and activity are altered in a tissue-specific manner by physiological conditions like cold exposure40, 41, lactation42, 43, or feeding and fasting44, 45. In fasting, with ensuing hypoinsulinemia, LPL activity decreases in adipose tissue but increases in the heart. As a result, FA from circulating TG is diverted away from storage to meet metabolic demands of the heart. Hence, LPL fulfills a gate-keeping role by regulating the FA supply to meet metabolic requirements of different tissues. However, when excessive FA uptake exceeds its mitochondrial oxidative capacity; as a result, an increase in FA conversion to potentially toxic FA metabolites, including ceramides, diacylglycerols, and acylcarnitines, paired with increased formation of reactive oxygen species secondary to elevated FA oxidation, can provoke cardiac cell death (lipotoxicity)46. Not surprisingly then, cardiac-specific LPL overexpression caused severe myopathy characterized by lipid oversupply and deposition, muscle fiber degeneration, excessive dilatation, and impaired left the ventricular function in the absence of vascular defects, a situation comparable to diabetic cardiomyopathy47-49. Interestingly, loss of cardiac LPL also causes cardiomyopathy50-52. Hence, although specific knockout of cardiac LPL increased glucose metabolism, neither this effect nor albumin-bound FA could substitute the action of LPL, and cardiac ejection fraction decreased51. These experiments in genetically modified mice demonstrate that cardiac LPL is of crucial importance, and disturbing its natural function is sufficient to cause cardiac failure. 1.3 Heparanase Following it secretion onto the cardiomyocyte HSPG, LPL requires transfer across the interstitial space. Heparanase is an endoglycosidase with a unique ability to hydrolyze HSPG into oligosaccharides24. In the heart, given its proximity to the cardiomyocyte and the unique position to detect metabolic changes in the circulation, endothelial heparanase is a potential regulator of LPL translocation. 5 1.3.1 Active heparanase Heparanase is initially synthesized as pre-proheparanase (68 kDa)53, and is processed into latent heparanase (65 kDa), which undergoes cellular secretion and HSPG-facilitated reuptake54, 55. After latent heparanase undergoes a proteolytic cleavage in the lysosome, an active 50-kDa polypeptide is formed, that can be mobilized by demand and secreted to degrade cell surface heparan sulfate. In cancer cells, extracellular nucleotides, such as adenosine triphosphate (ATP), adenosine diphosphate, and adenosine, through a purinergic receptor, G-coupled protein-coupled protein kinase C, and protein kinase A activation signal cascade, are known to be the active agents in stimulating heparanase secretion56. Although a role for heparanase in physiology (e.g., embryonic morphogenesis) has been described, it was intensive research focused on cancer progression that hinted towards a unique responsibility in cardiac metabolism. In cancer, degradation of HS chains by the increased expression of heparanase is associated with extracellular matrix and basement membrane disruption to facilitate tumor cell invasion57, 58. In the heart, active heparanase secretion from the basolateral side of EC can liberate LPL from the cardiomyocyte into the interstitial space for onward progress to the vascular lumen59. 1.3.2 Latent heparanase The human heparanase gene encodes a polypeptide composed of 543 amino acids, and is transcribed into pre-proheparanase (68 kDa) that contains a N-terminal signal peptide, a C-terminal hydrophobic peptide, five cysteine residues, and six N-glycosylation sites53. When pre-proheparanase undergoes glycosylation in the endoplasmic reticulum and its signal peptide removed, a 65 kDa latent heparanase is formed. Latent heparanase is secreted following its transport into the Golgi and packaging into vesicles. Although it is ~100-fold less active than 50 kDa active heparanase60, 61, latent heparanase has some remarkable properties, including a greater competence for releasing myocyte surface growth factors, such vascular endothelial growth factor (VEGF)62, as compared to active heparanase, in addition to LPL. It was the reported liberation of VEGF anchored 6 to heparan sulfate that was most provocative in cancer. VEGF enhances microvessel density63, 64, an angiogenic feature that provides tumor cells with sufficient nutrition and the potential for distant metastasis. In the heart, VEGF-associated angiogenesis would be required for an increased oxygen demand to provide the oxygen necessary for increased LPL-derived FA oxidation65, 66. Another intriguing non-enzymatic function of latent heparanase is to modulate gene transcription by being taken up and entering into the nucleus of the cell. In cancer, those properties of secreted latent heparanase induce cell signaling and modulate gene expression in neighboring cells to promote cancer progression55, 67. 1.4. Glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 Following its release from its cardiomyocyte-HSPG receptor, the LPL in the interstitial space has to traverse the EC from its basolateral to its apical side8. Multiple mechanisms were suggested to contribute to their enzymatic transfer including endothelial-HSPG68, 69 and the VLDL receptor70. More recently, a glycosylphosphatidylinositol (GPI) anchored glycoprotein expressed exclusively in the EC named glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1) has been suggested to be the principal protein that chaperones LPL across the vascular lumen to the apical side8. Immunochemistry staining reveals that the tissue expression pattern of GPIHBP1 is similar to that of LPL in brown adipose tissue and the heart27, 71. Moreover, GPIHBP1 is expressed exclusively in capillary EC while LPL is expressed in parenchymal cells. GPIHBP1 deficient mice develop severe hyperlipidemia even when fed a low-fat chow diet, and patients with GPIHBP1 mutations have severe hypertriglyceridemia27, 71. There are three features of GPIHBP1, including a GPI anchor, a N-terminal acidic domain (enriched in aspartate and glutamate), and a cysteine-rich lymphocyte antigen 6 (Ly6) motif71, 72. The GPIHBP1 protein can be released from the plasma membrane by cleaving the GPI anchor with phosphatidylinositol-specific phospholipase C (PI-PLC)72. The acidic domain of GPIHBP1 can electrostatically interact with LPL. GPIHBP1 mutations (replacement of aspartate and glutamate to 7 alanine) in the acidic domain fail to bind LPL73. The Ly6 motif of GPIHBP1 contains 10 cysteines that are necessary for a disulfide-linked spacing pattern and result in a three-fingered structure motif. Mutations of these cysteine reduce its ability to bind LPL, suggesting that the Ly6 motif is equally important for binding LPL74. In addition, the N-linked glycosylation site located in the Ly6 motif in mouse GPIHBP1 is crucial for trafficking GPIHBP1 from the endoplasmic reticulum to the EC surface74. At the basolateral side, GPIHBP1 operates as a LPL transporter to shuttle LPL across the EC to the apical side8, 75. At the apical side, its ability to avidly bind both lipoprotein-TG and LPL allows it to serve as a platform for TG (chylomicrons, VLDL) lipolysis along the luminal surface of capillaries71. When LPL binds to GPIHBP1, it is capable of stabilizing LPL, in addition to preventing its inhibition by Angptl3 and Angptl439. The earliest evidence of GPIHBP1-regulated expression originated from experiments investigating the effects of fasting/re-feeding76. Fasting augments coronary luminal LPL activity and cardiac GPIHBP1 expression, effects that are reversed 6 hours after re-feeding76. Given that PPARs are known to play a critical role in energy metabolism, it is reasonable to suggest PPARs as a regulator of GPIHBP1 expression. One study revealed several potential PPAR binding sites upstream of GPIHBP1 exon 175. PPARγ agonists increase GPIHBP1 expression, whereas mice with PPARγ deficiency in EC expressed lower levels of GPIHBP175. 1.5 Mesodermal homeobox 2 (Meox2) For the reason that EC have an essential role in regulating and maintaining cardiac function, it is conceivable that changes in EC proteins would affect EC communication with the underlying cardiomyocyte to facilitate FA delivery to, and utilization by, the cardiomyocyte. Mesodermal homeobox 2 (Meox2), also known as growth arrest-specific homeobox (GAX), is a homeobox gene that is expressed in EC9, 77. Meox2 is involved maintaining EC homeostasis in endothelial proliferation and VEGF-induced tube formation78. Recently, it has been reported that Meox2 also forms a heterodimer with another highly expressed cardiac EC basic helix-loop-helix transcription 8 factors 15 (Tcf15)9. Meox2/Tcf15 drives endothelial CD36, GPIHBP1 and LPL expression to facilitate FA uptake and transport across the cardiac endothelium9. Given that the haplodeficiency of Meox2/Tcf15 in mice impairs cardiomyocyte contractility9, this heterodimer not only plays a role in endothelial regulation of FA but may also be involved in altering cardiac energy substrates. 1.6 Diabetic cardiomyopathy Coronary vessel disease is a primary cause for the increased incidence of cardiovascular dysfunction following diabetes79, 80. However, patients with type 1 (T1D) and type 2 (T2D) diabetes have also been diagnosed with reduced or low-normal diastolic function and left ventricular hypertrophy in the absence of vascular defects (cardiomyopathy)81-83. Similarly, evidence of cardiomyopathy is reported in animal models of T1D and T2D84, 85. As it is a complicated disorder, several factors have been associated with the development of cardiomyopathy. These include an accumulation of connective tissue and insoluble collagen, impaired sensitivity to various ligands (e.g., β-agonists), and abnormalities in proteins that regulate intracellular calcium86-88. The view that alterations in cardiac metabolism can contribute towards the etiology diabetic cardiomyopathy has also been proposed51, 89-91. 1.6.1 Changes in cardiac metabolism during diabetes Cardiac glucose uptake is dependent on sarcolemmal content of transporters92, 93. Glucose transporter (GLUT) 1 has a predominant sarcolemmal localization accounting for basal glucose uptake whereas GLUT4 is the dominant transporter in the adult heart. In non-stimulated conditions, a majority of GLUT4 is intracellular. On stimulation by insulin or activation of AMPK, GLUT4 is redistributed to the sarcolemma to mediate glucose uptake92, 94, 95. In noninsulin-controlled T1D, reduced GLUT4 gene and protein expression along with a defect in membrane translocation of GLUT4 compromises cardiac glucose uptake and utilization96. In T2D animals, although there is hyperglycemia and hyperinsulinemia, cardiac glucose uptake is also compromised due to reduced GLUT4 protein and impaired insulin signaling96, 97. In the diabetic heart, there is excessive 9 dependence on FA utilization to generate ATP98. With a limited ability to synthesize FA, the heart relies on an exogenous supply to fuel cardiac contractility. To this extent, FA delivery and utilization entails a) its release from adipose tissue3, b) its uptake by cardiomyocyte-FA transporters, c) the breakdown of endogenous TG5, d) lipoprotein hydrolysis by LPL99, and e) amplification of genes involved in FA oxidation 100, 101. As the molar concentration of FA in lipoprotein-TG is 10-fold more than albumin-bound FA102, LPL is indispensable in this process. Under conditions of diabetes, both the adipose tissue and the breakdown of endogenous TG stored in the cardiomyocyte are augmented. The cardiac uptake of albumin-FA is driven by its plasma concentration and plasma membrane FA transporters, both of which are augmented following diabetes46, 102. Circulating lipoprotein concentrations are also elevated in diabetes because hepatic VLDL synthesis is enhanced by increased FA delivery to the liver103. However, the utilization of circulating VLDL as a FA source by the diabetic heart is influenced, not only by elevated plasma concentrations but also by the vascular content of LPL. 1.6.2 Heart LPL in human diabetes When examining LPL, negatively charged heparin can be used to displace LPL from its binding sites. Using this procedure, either a decrease or no change in post-heparin plasma lipolytic activity (PHPLA) in and T2D patients has been reported7, 104, 105. The drawback with this approach is that PHPLA represents LPL that is released from a number of tissues (including skeletal muscle and adipose tissue), and hence is incapable of establishing whether diabetes influences human cardiac LPL47, 106. With reference to tissue specific localization, adipose tissue, and less commonly skeletal muscle show low levels of LPL (in tissue homogenates) following diabetes, with virtually no information being available on the cardiac content of this enzyme105. However, even if resolved, estimation of LPL in heart homogenates from patients with diabetes would be inappropriate, as it would reflect total cardiac LPL and not the functional pool of enzyme at the coronary vascular lumen. 10 1.6.3 Heart LPL in animal diabetes Given the obstacles associated with the determination of human heart LPL, data from animal studies provided the majority of information regarding cardiac LPL in diabetes. However, similar to human studies, the contribution of LPL to deliver FA to the heart from animal models of diabetes was also inconclusive102, 107-111. In mouse models of T1D [streptozotocin (STZ)-induced] or T2D (db/db), no change of cardiac heparin-releasable LPL activity is observed112, 113. This could be a consequence of genetic adaptation (db/db mice have early exposure to high circulating FA and hence limited need for LPL), or the excessive heart rate (which requires a substantial amount of ATP) in control animals (~600 beats/min), permitting prior translocation of LPL from the cardiomyocyte to the coronary lumen to saturate all of the vascular LPL binding sites23, 114. In rat models of STZ-diabetes, cardiac LPL protein or activity has also been reported to be unchanged, increased, or decreased115-119. This variability could be due to the diversity in the rat strains used, the dosage of STZ employed to induce T1D, and the duration of the diabetic state43. In addition, many of these investigations utilized procedures that did not distinguish between functional (endothelial-bound) and cellular (storage form of the functional enzyme) pools of cardiac LPL because cellular LPL activity or protein levels were largely obtained using heart homogenates43. Considering that both coronary lumen and parenchymal cells contribute to LPL activity, it is more appropriate to measure this enzyme activity at its functional site, the coronary lumen. We reported changes in LPL activity at the coronary lumen that was dependent on the dose of STZ used. Hence, following a single dose of 55 mg/kg STZ (D55), rats develop hypoinsulinemia and hyperglycemia (Figure 2). In D55 animals, coronary LPL increased after 4 days and remained elevated up to 12 weeks of diabetes107, 120. This was associated with an increased rate of TG-lipoprotein breakdown. The elevation of LPL was unrelated to changes in gene expression. Instead, this significant change of LPL activity resulted from post-translational changes that included augmented processing to dimeric, catalytically active enzyme121, LPL movement to the 11 cardiomyocyte plasma membrane20, and transfer to the coronary lumen. An increase in calnexin was responsible for LPL maturation into the dimeric enzyme121. PKD governed the secretory process that promoted LPL secretion onto the cardiomyocyte surface in D55 rats. This regulation was enhanced by AMPK phosphorylation, which facilitated p38 MAPK activation. Activation of this pathway resulted in the release of actin monomers from Hsp25 to produce actin cytoskeleton rearrangement and translocation of LPL to the myocyte surface23. After its secretion onto surface-bound HSPG and onward movement across the interstitial space to the apical side of the vascular lumen EC, the detachment of LPL from the myocyte surface is a prerequisite. This is likely mediated by enzymatic cleavage of cardiomyocyte surface HSPG by heparanase59. In response to hyperglycemia following diabetes, lysosomal endothelial heparanase redistributes from a perinuclear location towards the plasma membrane of EC59. This high glucose-induced heparanase secretion is associated with ATP release, purinergic receptor activation, cortical actin disassembly and stress actin formation122. In addition, high glucose also upregulates heparanase gene expression in the EC123, an effect inhibited by insulin124. By enhancing LPL liberation from surface HSPG, heparanase is valuable in facilitating LPL translocation from parenchymal cells to the vascular lumen following diabetes125. Besides catalytic activity, heparanase also has other biological effects to modulate gene expression. This is likely through the nuclear entry of active heparanase and cleaving of heparan sulfate in the nucleus. As a consequence, there is a reduction of the nuclear content of syndecan-1, a type of HSPG that inhibits histone acetyltransferase (HAT) activity, and hence gene expression [e.g. VEGF and matrix metalloproteinase 9 (MMP9)] is promoted126-128. In the diabetic heart, latent heparanase can be taken up by cardiomyocytes and converted into its active form, to cause surface HSPG to be shed by MMP9 in the cardiomyocytes128. This effect may sustain LPL transfer to the coronary lumen after chronic diabetes. When LPL is released into the interstitial space, it is further shuttled from the basolateral to the apical side of the EC where the enzyme is functional. As GPIHBP1 levels change quickly in fasting/refeeding76, it is conceivable that this protein participates in the accelerated 12 transfer of LPL from the cardiomyocyte to the vascular lumen to increase FA delivery to the diabetic heart. Whether this protein plays a role in the transfer of LPL to the vascular lumen after diabetes is unclear. In contrast to D55, increasing the dose of STZ to 100 mg/kg (D100) to mimic poorly controlled severe diabetes, the rats develop hypoinsulinemia, hyperglycemia, and hyperlipidemia (Figure 2), and do not survive beyond 7 days without exogenous insulin129. In these severely diabetic animals kept for 4 to 7 days, LPL gene expression and coronary enzyme activity were reduced121. This could result from impaired vesicle formation by proteolysis of PKD as a result of caspase-3 activation and loss of its protecting protein 14-3-3ζ29. The severity of diabetes also regulates AMPK activation. Compared to D55 rats, increasing the dose to 100 mg/kg introduces an additional metabolic manifestation, hyperlipidemia, which inhibits AMPK in the heart and contributes towards reduced LPL trafficking to the myocyte surface107, 130. Angptl4 has also been suggested to influence LPL in D100 animals. In these animals, there is a significant increase in serum Angptl4 levels that are known to reduce coronary LPL activity121. It is possible that hyperlipidemia can stimulate Angptl4 expression/release from the adipose tissue, which disassociates dimeric LPL and minimizes its function in D100 hearts. The role of GPIHBP1 and Meox2/Tcf15 in LPL function following diabetes has yet to be studied. It will be of some interest to investigate whether this glycoprotein and the heterodimer both play a role in LPL transfer following diabetes. 1.7 Hypothesis and research objectives Cardiac LPL is crucial for maintaining ATP production to support heart function. The regulation of LPL following diabetes relies on posttranslational modification of cardiac LPL; however, whether this is also achievable through its transporter GPIHBP1 is unclear. We hypothesize that GPIHBP1 plays a role in the transfer of LPL to the vascular lumen after diabetes. The objectives of this research proposal were to: 13 1. Determine changes in cardiac EC GPIHBP1 following diabetes. 2. Investigate the potential mechanism(s) by which hyperglycemia alters EC GPIHBP1. 3. Examine if a crosstalk exists between EC and cardiomyocyte to regulate FA delivery to the heart. 14 Figure 1. Communication between the EC and cardiomyocyte promotes LPL transfer to the vascular lumen. In the cardiomyocyte, newly synthesized LPL enters the endoplasmic reticulum (ER) and is processed by chaperones to form dimers in a head-to-tail manner. After dimeric LPL exits the ER, it is further sorted into secreted vesicle in the Golgi. Activation of protein kinase D (PKD) helps in the formation of the vesicular LPL, which is transported from the Golgi to the plasma membrane. AMPK then stimulates actin cytoskeleton polymerization and builds a bridge to transport LPL-containing vesicles to HSPG binding sites on the myocyte cell surface. In response to high glucose, EC releases both heparanase to liberate HSPG-bound LPL from the cardiomyocyte cell surface. At the basolateral side of the EC, GPIHBP1 captures LPL in the interstitial space and transfers it across to the apical side. Out here, GPIHBP1 serves as a platform to enable LPL to hydrolyze lipoprotein-TG and release FA. Angptl4 can inactivate LPL by disassociation into a monomeric form. 15 Figure 2. General characteristics of different models of diabetes. Rat blood was collected, and following centrifugation, serum was isolated. The concentration of glucose, nonesterified fatty acid (NEFA), triglyceride (TG), and insulin were determined using diagnostic kits. *p < 0.05, ** p < 0.01, compared to CON. n= 6-12. 16 CHAPTER 2: METHODS 2.1 Materials Rat aortic EC (RAOEC), bovine coronary artery EC (BCAEC), and rat heart microvascular EC (RHMEC) were obtained from Cell Applications Inc., Clonetics, and VEC Technologies, respectively. Streptozotocin, chloroform, isopropanol and N-[N-(3,5-Difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (DAPT) were purchased from Sigma-Aldrich. Heparin (HEPALEAN, 1000 U/mL) was from Organon, Canada. [3H]-triolein was purchased from Amersham Canada. Collagenase I was from Worthington. Anti-heparanase antibody mAb 130 is from InSight (Rehovot, Israel), which recognizes both the latent (65 kDa) and active (50 kDa) form of heparanase. Anti-GPIHBP1 antibodies were from Novus Biologicals (NB110) and Abcam (ab59780). Anti-VEGF (ab46154), anti-DLL4 (ab7280) and anti-activated Notch1 antibody (NICD) (ab8925) antibodies were obtained from Abcam. Anti-PECAM (CD31, mab1393) was from Millipore. Anti-β actin (sc4778), anti-LPL (sc73646), goat anti-mouse (sc2005) and goat anti-rabbit (sc2004) antibodies were purchased from Santa Cruz Biotechnology. To measure free fatty acid released from VLDL-TG breakdown, an NEFA-C assay kit was purchased from Wako. Purified active and latent heparanase were a kind gift from Dr. Israel Vlodavsky, Hadassah University Hospital, Jerusalem, Israel. Recombinant VEGF (293-VE-050), DLL4 (1506-D4-050), TGFβ (240-B-002) were purchased from R&D. Phosphatidylinositol-specific phospholipase C (PIPLC), Alexa 555-labeled anti-mouse IgG, Alexa 488-labeled anti-rabbit IgG, TRIzol, SuperScript II RT, GPIHBP1 (Rn01503971_g1), β-actin (Rn00667869_m1), 45s (Rn03928990_g1), VEGF (Rn01511601_m1), Hes-1 (Rn00577566_m1), Hey-1 (Rn00468865_m1), MMP9 (Rn00579162_m1) probe, and TaqMan fast advanced master mix were from Invitrogen. Rat Cytokine Array C2 kit was purchased from RayBiotech. The MMP9 activity kit was purchased from AnaSpec. The RhoA activation G-LISA assay kit (BK124) and G-Actin/F-actin in Vivo assay biochem kit (BK037) were obtained from Cytoskeleton (Denver, CO). 17 2.2 Experimental animals This investigation adheres to the Guide for the Care and Use of Laboratory Animals published by the U.S. National Institutes of Health and the University of British Columbia (A13-0098). Adult male Wistar rats (250-320 g) were injected with streptozotocin (STZ), which enters the β cell via the GLUT2 receptor leading to DNA alkylation and cell death131. We used different doses of STZ [55 (D55) or 100 (D100) mg/kg] through tail vein injection to generate moderate and severe diabetes respectively, and the animals were kept for four days. With D55, animals are insulin deficient, and luminal LPL activity is augmented121. Unlike D55, D100 animals also develop hyperlipidemia, and demonstrate a decline in vascular LPL29, 121 (Figure 2). Hyperglycemia was confirmed in blood samples from the tail vein using a glucometer (AccuSoft) and glucose test strips (Accu-Chek Advantage; Roche). 2.3 Heart perfusion and isolation of cardiomyocytes Ventricular calcium-tolerant myocytes were prepared by a previously described procedure132. Briefly, after animals were euthanized with 65 mg/kg sodium pentobarbital i.p., the hearts were carefully removed, cannulated via the aorta, and retrogradely perfused with Tyrode's solution (for isolation of cardiomyocytes)132. Myocytes were made calcium tolerant by exposure to increasing concentrations of calcium. Cardiomyocytes were plated at a density of 200,000 cells on laminin-coated 35 mm dishes. Cells were maintained in Medium-199 (Sigma) and incubated at 37°C in a 5% CO2 humidified incubator. 2.4 Endothelial cell culture RAOEC, BCAEC, and RHMEC from the fifth to the tenth passages of 3 different starting batches, for each cell line, were used. Cells were cultured in the growth medium that contains 5% fetal bovine serum at 37°C in a 5% CO2 humidified incubator. In some experiments, RAOEC and BCAEC from passage fifth to tenth were cultured in growth medium at 37°C in a 5% CO2 18 humidified incubator59. Cells were detached following trypsin digestion, and aliquots used for seeding of a new culture. 2.5 Endothelial cell and cardiomyocyte co-culture For co-culture with cardiomyocytes, RAOEC and BCAEC from passage eighth to tenth were seeded on Transwell inserts. On reaching 80-90% confluence, these inserts were placed in a 6-well plate that had cardiomyocytes attached to the bottom (Figure 9D). Cardiomyocytes were plated at a density of 200,000 cells/well. 2.6 Isolation of heart endothelial cells Primary EC isolation and culture was done by a previously described procedure9. Briefly, after animals were euthanized, and the heart excised, they were perfused with 1.5 mg/mL collagenase I (Worthington) for 20 minutes. After digestion, tissue was minced with ice-cold FACS buffer that contained PBS, 1 mM HEPES, 1% BSA and adjusted to a pH of 7.4. Cells were then filtered through a 40 µm mesh, and centrifuged at 100 g for 3 minutes to remove non-EC. The supernatant containing EC was further centrifuged at 250 g for 10 minutes, and the pellet resuspended in DMEM (Gibco11885084) containing 20% fetal bovine serum (FBS). Anti-CD31 coated beads were then introduced to the medium to ensure EC purification. Using EC markers, including CD31 and Von Willebrand Factor (vWF), we confirmed their expression in both isolated heart EC and RHMEC (Figure 3). In some experiments, we also compared Meox2 expression between primary heart EC and EC cell lines (RAOEC and RHMEC) (Figure 4). 2.7 LPL transport and activity assay To measure coronary LPL, Kreb's buffer containing heparin (8 U/mL) was used, perfusion effluent collected over 5 minutes, and LPL activity determined. To release EC surface-bound LPL, cells were incubated with heparin (8 U/mL) for 3 minutes. To study basolateral to apical transport of LPL, EC were grown on transwell inserts [Falcon 353102, 24 mm diameter; 1.0 µm pore size enables the EC secretory proteins to communicate with the underlying cardiomyocytes (and vice 19 versa)] until the cells formed a tight monolayer27. The inserts were then placed in 6-well plates. Subsequently, LPL (5000 U/10 µg/mL) was added to the bottom chamber. After 1 hour, LPL that translocated to the apical surface of ECs was collected by heparin displacement, and LPL activity determined. LPL activity was determined by measuring in vitro hydrolysis of [3H]triolein substrate23. Radiolabeled FA products were extracted and estimated by liquid scintillation counting. 2.8 Immunofluorescence Microscopy To visualize LPL and GPIHBP1 localization, rat hearts were perfused with cold PBS, fixed in 10% paraformaldehyde overnight, and 5 or 10 µm parafilm-embedded sections prepared. Sections were incubated at 56°C for 30 minutes, and dehydrated by xylene (2x, 10 minutes) and ethanol (35-95%, 5 minutes) followed by PBS. We followed previously reported procedures of staining and microscopy27, 71. Briefly, sections were permeabilized with 0.2% Triton X-100 in PBS for 30 minutes, and incubated with blocking buffer containing 10% donkey serum and 0.2% Triton X-100 in PBS for 30 minutes at room temperature. Slides were then incubated with primary antibodies at 4°C overnight; 1:400 for anti-CD31, anti-GPIHBP1, and anti-LPL. Secondary antibodies were used at a dilution of 1:400 and incubated at room temperature for 1 hour. After PBS washing, the slides were stained with DAPI to visualize DNA. Confocal fluorescence microscopy was performed using an LSM 700 confocal microscope (Zeiss) equipped with AxioCam ICc 5, and scanned with lasers at wavelengths of 555, 488, and 405 nm. Individual images were captured sequentially with Plan-Apochromat 20x/0.8 DIC or 40×/1.4 Oil DIC M27 objectives with the same fluorescence exposure settings between groups, and merged images were generated using Zen 2.1 Confocal Software. For detection of proteins in RAOEC, cells were seeded on coverslips and grown until 80-90% confluent and treated with indicated reagents (25 mM glucose, 100 ng/mL VEGF, and 1 ng/mL TGFβ). Following the treatment, cells were washed with cold PBS, fixed with methanol at 4°C for 1 hour and permeabilized with 0.2% Triton X-100 in PBS for 10 minutes. Subsequently, fixed cells were incubated with blocking buffer, then incubated with primary antibodies (1:400 anti-GPIHBP1, 20 anti-activated Notch1 (NICD), anti-CD31) at 4°C overnight, and secondary antibodies (1:400) at room temperature for 1 hour. In some experiments, to detect CD31 and GPIHBP1 in EC in response to DLL4, cells were seeded on coverslips pre-coated with recombinant DLL4 (100 ng/mL) for 12 hours, and above procedure (wash, fixation, permeabilization, blocking, primary/secondary antibodies) repeated. To visualize CD31, vWF and Meox2 localization in isolated heart EC from control and diabetic animals, cells were grown on coverslips, and above procedure (wash, fixation, permeabilization, blocking, primary/secondary antibodies, and DAPI staining) repeated. Microscopy was performed as described above. 2.9 Western blot To measure protein expression, 20-40 µg of protein was loaded. Membranes were incubated with primary antibodies (1:500 for anti-heparanase, 1:1000 for anti-GPIHBP1, anti-DLL4, anti-activated Notch1 antibody (NICD), and anti-TGFβ, 1:2000 for anti-β-actin and anti-Meox2, and 1:5000 for anti-vinculin) at 4°C overnight, and subsequently with secondary antibodies (1:2000) at room temperature for 1 hour. Protein expression was expressed using either β-actin or vinculin as the loading control. To measure medium heparanase, protein was concentrated with an Amicon centrifuge filter (Millipore), followed by the procedures listed above. 2.10 RNA isolation and real-time PCR Total RNA was isolated from rat hearts, RAOEC or RHMEC using TRIzol, followed by chloroform and isopropanol extraction, washing by ethanol and dissolving in DEPC-H2O. RNA was reverse transcribed into cDNA with a mixture of random primers, oligo-(dT), and SuperScript II. cDNA was amplified by TaqMan probes (GPIHBP1, VEGF, Hes-1, Hey-1, MMP9, and β-actin) in duplicate or triplicate, using a 7900HT Fast Real-Time PCR system. Gene expression was calculated with the comparative cycle threshold (ΔΔCT) method. 21 2.11 In vitro lipolysis of VLDL Isolation of VLDL (without chylomicrons) by density gradient ultracentrifugation was achieved using serum from 16 hours fasted rats (6 PM-10 AM). Fasting of rats for this duration leads to the production of more VLDL, and minimizes the contribution from chylomicrons. In vitro lipolysis of VLDL-TG was carried out by previously described methods133. Briefly, various concentrations (0-0.8 mM) of VLDL-TG were incubated with heparin releasable LPL at 37°C for 30 minutes. At the end of incubation, the reaction was stopped by addition of precooled 0.3 M Na2PO4 (pH 6.9), and the tubes immediately immersed in ice. From this reaction mixture, 50 µl was pipetted in triplicate to measure FFA released by hydrolysis of VLDL-TG with an NEFA-C kit (Wako Chemicals). 2.12 Cytokine array Medium was collected from RAOECs cells (endothelial cell culture medium, ECCM) incubated with DMEM medium containing either glucose (5.5 or 25 mM) or heparanase (500 ng/ml) for 12 hours. Myocytes (myocyte culture medium, MCM) incubated with DMEM medium or DMEM medium containing 500 ng/ml purified active or latent heparanase was also collected. Medium was centrifuge at 2000 rpm for 5 minutes. Array membranes were prior incubated with blocking buffer (1 hour) followed by incubation with ECCM or MCM at room temperature for 2 hours. Membranes were washed (3x, 5 minutes), incubated with working primary antibody and anti-HRP at room temperature for 2 hours, respectively. Array membrane was detected with ECL and Hyperfilm ECL Film (GE Healthcare). 2.13 Plasma measurements Blood samples were collected from animals before termination and serum or plasma isolated using centrifugation. Concentrations of triglyceride (Stanbio), VEGF (R&D), and TGFβ (Abcam) were determined using ELISA diagnostic kits. 22 2.14 VEGF ELISA Medium was collected from cardiomyocytes incubated with either heparin (5 U/mL) or purified latent heparanase (1 or 2 µg/mL) for 15 minutes. This medium was centrifuged at 2000 rpm for 5 minutes, and VEGF concentration determined using Rat VEGF Quantikine ELISA Kit (R&D). Briefly, the sample was incubated in each well containing assay diluent at room temperature for 2 hours. After incubation, each well was washed, conjugated buffer added for 1 hour and subsequently substrates for 30 minutes. Using stop-solution to terminate the reaction, absorbance readings minus background were converted to picograms using standard curves from the kit. 2.15 MMP9 activity assay MMP9 activity was evaluated by measuring fluorescence intensity using an assay kit from AnaSpec. Briefly, whole heart samples were homogenized in assay buffer containing 0.1% Triton-X 100, and centrifuged for 15 minutes at 10,000x g at 4°C. From the supernatant, 20 µg protein was added to MMP9 coated plates and incubated for 2 hours to pull down MMP9. After incubation, the non-binding proteins were removed, and amino-phenyl mercuric acetate (APMA) was added for 1 hour to activate MMP9. Following addition of the MMP9 substrate, the fluorescence signal was detected at Ex/Em = 490 nm/520 nm. 2.16 Plasma measurements of TGFβ Concentration of TGFβ (Abcam) was determined using a rat ELISA diagnostic kit. Briefly, the samples and standards were processed prior to incubation in each well containing assay diluent at room temperature for 2 hours. After incubation, each well was washed, conjugated buffer added for an hour, and then the substrate for 30 minutes. Using stop-solution to terminate the reaction, absorbance readings minus background were converted to picograms using standard curves from the kit. 23 2.17 GLISA assay To study RhoA activation, we used G-LISA assay to determine GTPase activity following manufacturer's instruction (Cytoskeleton). The active GTP-RhoA was detected by reading the absorbance at 490 nm. 2.18 Actin polymerization Actin polymerization was evaluated by measuring the filamentous to globular actin (F-actin to G-actin ratio) using an assay kit from Cytoskeleton. 2.19 Statistical analysis Data are expressed as means ± SEM. Wherever appropriate, unpaired Student's t-test, Mann-Whitney or and 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. 24 Figure 3. Isolated heart endothelial cells and rat heart microvascular endothelial cells express CD31 and Von Willebrand Factor. Isolated heart EC and RHMEC were seeded on slides that were placed in a 6-well plate for 48 hours. Slides were then washed with ice cold PBS, fixed with methanol and permeabilized using 0.2% Triton-X in PBS. Anti-vWF (1:200) and anti-CD31 antibodies (1:200) were used for immunofluorescence staining. Images were taken using a 20x objective with the same fluorescence exposure settings. Scale bar = 10 µm. 25 Figure 4. Primary heart endothelial cell and endothelial cell lines express Meox2. Primary isolated ECs, RAOECs, and RHMECs were cultured to 80-90% confluence. Cells were detached following trypsin digestion, and aliquots used for detection of Meox2 expression. Vinculin was used as an internal control. 26 CHAPTER 3: RESULTS 3.1 Endothelial cells respond to hyperglycemia by increasing the LPL transporter GPIHBP1 3.1.1 Cardiac LPL and GPIHBP1 after STZ diabetes Following injection of STZ and maintenance of animals for four days, there was a pronounced increase in blood glucose (control 7.2±1.1 mM, STZ (D55) 19.2±2.8 mM, p <0.05). As utilization of glucose is impaired following diabetes, the heart rapidly adapts to use FA exclusively, an effect largely determined by LPL23. Indeed, perfusion of hearts with heparin indicated that LPL activity increased following hyperglycemia (Figure 5A and upper inset). Interestingly, immunofluorescence detection of LPL revealed that most of the increased enzyme in diabetic hearts was located at the coronary lumen (Figure 5A, lower inset). In the heart, LPL synthesized in cardiomyocytes is shuttled across EC with the help of GPIHBP18, 134. To determine if the increased vascular content of LPL following diabetes is associated with GPIHBP1, its mRNA (Figure 5B, left panel) and protein (Figure 5B, right panel) and expression were examined, and determined to be augmented. As the increased GPIHBP1 had an exclusive endothelial location71 (Figure 5C), our data suggest that GPIHBP1 is important for myocyte to endothelial cell transfer of LPL to increase FA delivery to the diabetic heart. 3.1.2 Endothelial GPIHBP1 following cell passage and influence of high glucose GPIHBP1 is expressed exclusively in ECs8. We determined the level of this glycoprotein in RAOEC and BCAEC, and unexpectedly identified loss of this glycoprotein following passaging of these cells (Figure 6A). In an attempt to mimic in vivo hyperglycemia, RAOECs were exposed to increasing concentrations of glucose for 12 hours. Interestingly, in cells from later passages (in which GPIHBP1 expression had been suppressed), although concentrations of glucose between 10-15 mM increased GPIHBP1 mRNA (Figure 6B, inset) and protein (Figure 6C, inset), the increases were more robust with the higher concentration of glucose. Choosing 25 mM glucose and different 27 times, high glucose increased GPIHBP1 mRNA and protein expression, an effect that was pronounced after 12 hours (Figure 6B, 6C, and 7A). It should be noted that RAOEC and BCAEC from the earlier passages (e.g., 5-7) also responded to high glucose by increasing GPIHBP1 expression (data not shown). Overall, these data imply that by sensing the impending loss of glucose transport, endothelial cells respond by increasing GPIHBP1 to shuttle LPL. 3.1.3 GPIHBP1 associated transfer of LPL across the endothelial monolayer To test the functional relevance of the high glucose induced augmentation of GPIHBP1, LPL transport from the basolateral to the apical side of RAOEC monolayers was determined (Figure 7B). As expected, there was an increase in heparin releasable LPL activity at the apical side of endothelial cells exposed to high glucose (Figure 7C), an effect that was blunted by PIPLC (Figure 7C, inset). The osmotic control mannitol had no influence on LPL transport (Figure 7C). The LPL that had been shuttled following high glucose was functionally active as exposure of this apical heparin releasable enzyme to lipoprotein triglyceride was capable of increasing its hydrolysis to fatty acid (Figure 7D). Since prior exposure of cells to PIPLC (that is expected cleave GPIHBP172 and prevent LPL shuttling) abolished this TG breakdown, our data indicate that in ECs exposed to high glucose, enhancement of GPIHBP1 is an effective stimulus for increasing delivery of FA to cardiomyocytes to support energy requirements. 3.1.4 GPIHBP1 expression is linked to endothelial content of heparanase Heparanase, an endothelial endoglycosidase can cleave HS side chains on HSPG in the extracellular matrix and on the cell surface of cardiomyocytes to release LPL for transfer across the interstitial space to reach EC135. As described previously, exposure of RAOECs to high glucose rapidly increased the secretion of active heparanase (Figure 8B). Intriguingly, in addition to its ability to rapidly liberate cardiomyocyte HSPG-sequestered LPL, exposure of EC to active heparanase for 12 hours produced a robust increase of GPIHBP1 mRNA (Figure 8C, left panel) and protein (Figure 8C right panel). As passaging of RAOECs progressively decreased heparanase 28 expression (Figure 8A), similar to loss of GPIHBP1, our data has uncovered a novel autocrine role (either direct or indirect) for active heparanase to affect GPIHBP1 expression. 3.1.5 Cardiomyocytes induce endothelial cell GPIHBP1 Although GPIHBP1 is highly expressed in EC in vivo, EC in culture lose expression of this glycoprotein136. As EC in the heart are closely appositioned to cardiomyocytes, we reasoned that a paracrine influence from cardiomyocytes may be responsible for EC expression of GPIHBP1. Using a co-culture system (Figure 9D) to mimic the intact heart, we determined that in ECs in which GPIHBP1 had been silenced (passage 8-10), a simple introduction of cardiomyocytes into the vicinity, restored GPIHBP1 expression to levels observed with early cell passages (Figure 9A and 9B). These results point towards paracrine factors from cardiomyocytes influencing EC GPIHBP1 expression. Assuming that the stimulus for the release of signaling mediators from myocytes could originate from EC (likely heparanase), we exposed the co-culture to high glucose. Noteworthy, a further expression of EC GPIHBP1 was evident in this condition (Figure 9C), suggesting that high glucose induced heparanase released from EC can affect GPIHBP1 by both autocrine and paracrine signaling. 3.1.6 Heparanase released platelet-derived growth factor from myocytes and EC can induce GPIHBP1 expression EC and myocyte HSPG anchored proteins can be liberated by heparanase59. Using a protein array, and EC exposed to high glucose or myocytes treated with active heparanase, a number of proteins were increased in the culture medium (Figure 10). These included monocyte chemotactic protein-1 (MCP1), prolactin, receptor for advanced glycation end products (RAGE), thymidylate kinase TMP-1, and platelet-derived growth factor (PDGF) (Figure 10 and 11A). Given the acknowledged role of PDGF in FA metabolism137, we treated EC with recombinant PDGF and discovered an induction of GPIHBP1 from passage 7 (which still express GPIHBP1) and passage 10 (in which there is a loss of GPIHBP1) (Figure 11B). Nevertheless, the increase was more robust in 29 cells from passage 10, as these cells had an extremely low initial expression of GPIHBP1. Our data suggests a mediator role (either autocrine or paracrine) for this growth factor in FA delivery to the diabetic myocyte. 3.2 Cardiomyocyte regulates endothelial cell GPIHBP1 to relocate lipoprotein lipase to the coronary lumen during diabetes 3.2.1 High glucose alters the expression of endothelial GPIHBP1 GPIHBP1 is expressed exclusively in EC8, and its increase in hearts from STZ induced diabetic animals138. To mimic in vivo hyperglycemia, RAOECs were exposed to high glucose. We have previously observed elevated GPIHBP1 mRNA (Figure 6B, inset) and protein (Figure 6C, inset) expression with an increase in the concentration of glucose. Interestingly, high glucose increased GPIHBP1 mRNA (Figure 12A, left panel) and protein (Figure 12A, right panel) expression, with an amplification of LPL shuttling across EC (Figure 12A, inset) for 12 hours. High glucose is an effective stimulus for endothelial heparanase secretion59. Exposure of RAOECs to high glucose rapidly increased the secretion of both active and latent heparanase into the incubation medium (Figure 12B). The osmolality control mannitol had no effect on medium heparanase (data not shown). Tumor cells, by expressing higher levels of heparanase, are more invasive as secreted heparanase can breakdown extracellular matrix in addition to promoting gene expression related to an invasive phenotype139. To test this latter property, EC were exposed to both active and latent heparanase. Interestingly, there was a substantial increase of GPIHBP1 mRNA and protein within 24 hours of incubation not only with active (Figure 8C) but also latent heparanase (Figure 12C), suggesting that heparanase may mediate the effects of high glucose on EC GPIHBP1 expression. 3.2.2 The effect of heparanase on GPIHBP1 expression is associated with VEGF Cell surface HSPG anchor many different proteins25 including growth factors such as VEGF26, and heparanase is an endoglycosidase exceptional in its ability to instigate release of these bound ligands140. Using a protein array, and EC exposed to high glucose or latent heparanase, we observed 30 secretion of VEGF into the culture medium (Figure 13A circle). Given a recent unanticipated function of VEGF in regulating EC fatty acid transport141, we incubated EC with recombinant VEGF, and discovered an induction of GPIHBP1 mRNA (Figure 13B, left panel) and protein (Figure 13B, right panel) expression within 24 hours. This increased EC GPIHBP1 in response to VEGF was also reflected on cell surface (Figure 13C). To substantiate this observation, we used a specific antibody to inhibit VEGF. As anticipated, GPIHBP1 expression and LPL translocation induced by VEGF was prevented (Figure 13D), suggesting that this growth factor is an effective stimulus for GPIHBP1 expression. 3.2.3 Cardiomyocyte surface bound VEGF stimulates endothelial cell GPIHBP1 GPIHBP1 is highly expressed in EC in vivo. However, EC in culture lose their ability to express this glycoprotein136 (Figure 6A). As EC in the heart are closely appositioned to cardiomyocytes, we reasoned that a paracrine influence from cardiomyocytes might be responsible for EC expression of GPIHBP1. We used a co-culture system to mimic the intact heart and determine whether a simple introduction of cardiomyocytes into the vicinity of EC can restore GPIHBP1 expression. An increased EC GPIHBP1 was found in the co-culture group (Figure 14A), suggesting paracrine factors from cardiomyocytes influence GPIHBP1 expression. Like EC, myocyte HSPG anchored proteins can be liberated by heparanase140. Using a protein array, we also observed secretion of VEGF from myocytes following treatment with latent heparanase (Figure 14B, left panel circle), an effect that was more robust than heparanase liberation of VEGF from EC (Figure 14A). Interestingly, this was also reflected in mRNA expression; cardiomyocytes had a stronger VEGF signal compared to EC (Figure 14B, right panel). More importantly, cardiomyocytes had a significant amount of VEGF located on the cell surface, a pool that could be easily and rapidly (within 15 minutes) displaced by latent heparanase. This effect of latent heparanase on VEGF was likely through ionic displacement, and surprisingly, much more emphatic than heparin (Figure 14C). Confirmation of the influence of myocyte cell surface VEGF on GPIHBP1 induction was achieved 31 using heparitinase to remove cardiomyocyte surface HSPG prior to co-culture with EC. This procedure essentially prevented the induction of GPIHBP1 and LPL translocation (Figure 14D). As antibodies against both VEGF and its receptor had identical effects on GPIHBP1 expression and LPL translocation as seen with heparitinase (Figure 14E), our data suggest that cardiomyocyte HSPG bound VEGF is a crucial paracrine factor that regulates EC GPIHBP1 expression. 3.2.4 GPIHBP1 induction by VEGF is through a DLL4-Notch signaling pathway Delta-like ligand (DLL) 4 is a transmembrane ligand for the Notch receptor142, a key regulator of metabolism143. As a recent report indicated that Notch participates in VEGF regulation of EC fatty acid transport141, we tested whether the VEGF-induced GPIHBP1 expression is through the activation of Notch signaling. In response to the angiogenic growth factor, we observed an increased expression of DLL4 (Figure 15A). To investigate the contribution of the DLL4 pathway to GPIHBP1 expression, EC were grown on plates coated with recombinant DLL4. In addition to the traditional downstream target genes (Hey-1, Hes-1) of the DLL4-Notch pathway that were upregulated (Figure 16), we discovered a significant augmentation in both the mRNA and protein of GPIHBP1 (Figure 15B). The increase in GPIHBP1 with DLL4 was predominantly membrane bound71 (Figure 15C). As a specific antibody against DLL4 reduced the increased expression of GPIHBP1 observed following DLL4 (Figure 15D) or VEGF (Figure 17) stimulus, our data suggests that the effect of VEGF on GPIHBP1 induction is through DLL4-Notch signaling (Figure 15E). 3.2.5 VEGF induced GPIHBP1 expression is decreased by inhibition of Notch signaling The Notch receptor is a transmembrane protein composed of an extracellular, a transmembrane, and a Notch intracellular domain (NICD). Upon ligand binding, the notch receptor undergoes proteolytic cleavage and releases NICD, which enters the nucleus and modifies gene expression142, 143. To substantiate that notch signaling is involved in induction of GPIHBP1 following VEGF stimulation, we attempted to inhibit NICD translocation using DAPT. As anticipated, the γ secretase inhibitor reduced NICD (Figure 18A, left panel), lowered its entry into the nucleus (Figure 18B) and 32 decreased expression of GPIHBP1 (Figure 18A, right panel). Repeating this experiment in our co-culture showed similar results. Thus, pretreatment of EC with DAPT, prior to co-culture with cardiomyocytes, lowered NICD (Figure 18D, left panel) and prevented the increase in GPIHBP1 (Figure 18D, right panel). These results suggest that the effect of VEGF on GPIHBP1 is through Notch receptor signaling (Figure 18C and 28). 3.2.6 Reduction in cardiomyocyte VEGF following diabetes is associated with a decrease in GPIHBP1 expression and LPL activity at the vascular lumen Down-regulation of myocardial VEGF is a feature of experimental diabetes144, 145. We used 100 mg/kg STZ (D100) to generate a rat model of severe diabetes that exhibited both hyperglycemia and hyperlipidemia (Figure 19). Diabetic animals demonstrated a robust decrease in serum VEGF (Figure 20A, inset), whereas in hearts isolated from these animals, a significant reduction in VEGF mRNA was observed (Figure 20A). This decrease in VEGF was also reflected in cardiomyocytes from severe diabetes animals (D100). Thus, the myocyte surface pool of VEGF, as determined by release with either heparin or latent heparanase, was significantly lower in D100 compared to control (Figure 20B). To test the hypothesis that diabetic cardiomyocytes exhibiting low expression of VEGF on the cell surface may influence the LPL transporter GPIHBP1, we co-cultured these cells with EC. Without cardiomyocytes, EC had limited GPIHBP1 expression in vitro, whereas co-culture with control myocytes caused a robust induction of GPIHBP1. Interestingly, we found a reduction of GPIHBP1 expression (Figure 20C, right panel) with a decrease of NICD (Figure 20C, left panel) on co-culturing EC with myocytes from the D100 group, suggesting that VEGF on the myocyte surface is important for GPIHBP1 expression. This in vitro data could explain the decrease in LPL activity and mass at the vascular lumen (C, inset). Unlike the model of severe diabetes (100 mg/kg STZ), moderate (55 mg/kg STZ) diabetes induced an increase in LPL activity at the coronary vascular lumen (Figure 2), an effect that was associated with elevated VEGF at the myocyte surface (Figure 21). 33 3.3 Dual effects of hyperglycemia on endothelial cells and cardiomyocytes to enhance coronary LPL activity 3.3.1 Alteration in endothelial Meox2 is associated with changes in cardiac LPL following diabetes. In the diabetic heart with its proclivity for reduced glucose utilization and excessive FA consumption, the latter response is likely a result of several mechanisms including an increased LPL mediated lipoprotein TG lipolysis. In rats made diabetic with STZ (D55) and maintained for four days, there was a pronounced increase in blood glucose (control 7.1±0.5 mM, STZ (D55) 20.9±1.4 mM, p <0.05). As we have reported in previous studies29, 121, using the Langendorff mode to perfuse hearts with heparin, we observed an augmentation of LPL activity (Figure 22A) and mass (Figure 22A inset) at the vascular lumen of diabetic animals. Meox2 is a transcription factor that could be potentially implicated in explaining this LPL increase by promoting its transfer across the EC. In primary cells isolated from the heart, only EC demonstrated a robust expression of Meox2 (Figure 22B). We tested whether Meox2 responds to diabetes in a manner similar to LPL. Interestingly, in hearts from diabetic animals, an increase of Meox2 protein and its downstream GPIHBP1 mRNA expression were evident (Figure 22C and inset). We confirmed that this increase in Meox2 was predominantly in the EC (Figure 22D), and mainly located in the nucleus (Figure 22E and inset). Our data suggests an induction and nuclear translocation of Meox2 in EC following diabetes, effects that could help in the transfer of LPL to the vascular lumen by influencing the LPL transporter, GPIHBP1. 3.3.2 High glucose is a potential stimulus to induce endothelial Meox2 and increase GPIHBP1 expression. To test whether high glucose is an effective stimulus for promoting Meox2 expression, we incubated RHMEC with 25 mM glucose for 8-24 hours, or 5.5-30 mM glucose for 12 hours (Figure 23A), and found a substantial augmentation of this transcription factor with 25 mM glucose after 12 34 hours (Figure 23B). Recently, it has been reported that Meox2 forms a heterodimer with another highly expressed cardiac EC basic helix-loop-helix transcription factor 15 (Tcf15), and that Meox2/Tcf15 heterodimer can potentially regulate the expression of GPIHBP19. GPIHBP1 is the transporter that transfers LPL from the basolateral to the apical side of EC, where it also provides a platform for LPL meditated hydrolysis of lipoprotein TG. We treated RHMEC with an increasing concentration of glucose, and found that with the induction of Meox2, there was a parallel increase in GPIHBP1 protein (Figure 23A lower panel) and mRNA (Figure 23B, inset). To substantiate the relationship between Meox2 and GPIHBP1, we used fetal bovine serum (FBS), a known inhibitor of Meox2146, or siRNA. Interestingly, increasing concentrations of FBS (Figure 23C) or siRNA (Figure 23D) were effective in inhibiting Meox2 and GPIHBP1. Overall, our data support the idea that the hyperglycemia induced increase in LPL is through upregulation of Meox2 and GPIHBP1 that are essential for LPL shuttling across the EC. 3.3.3 Heparanase secretion in response to high glucose stimulates MMP9 expression and TGFβ activation. In the heart, the majority of LPL at the vascular lumen is obtained from cardiomyocytes. Heparanase, an EC endoglycosidase cleaves cardiomyocyte HSPG side chains, and facilitates the release of bound LPL for transfer to the apical side of EC. In the diabetic heart, we observed an increase of active heparanase (although greater, latent heparanase did not achieve statistical significance) (Fig. 24A and left inset), both of which are capable of releasing myocyte surface LPL62. Given that high glucose is an effective stimulus for endothelial heparanase secretion59, we exposed RHMEC to 25 mM glucose and found a rapid increase in the secretion of active and latent heparanase into the incubation medium (Figure 24A, right inset). In addition to its ability to facilitate LPL release from the myocyte surface, latent heparanase can also be taken up by the cardiomyocyte, converted into active heparanase, and modulate gene expression 128, 147. We tested this possibility by incubating myocytes with latent heparanase for 16 hours, and observed an 35 amplification of MMP9 mRNA expression (Figure 24B). This increase in MMP9 mRNA was also observed in the diabetic heart with a corresponding augmentation of MMP9 activity (Figure 24B, left and right inset). MMP9 is known to promote the conversion of latent to active TGFβ 148-150, which itself is implicated in ATP generation during the epithelial-mesenchymal transition to promote metastasis 151. Diabetic animals demonstrated a robust increase in plasma total TGFβ (Figure 24C, inset). Moreover, in hearts isolated from these animals, a significant increase in active TGFβ was observed (Figure 24C). Overall, our data suggest that in response to high glucose and secretion of EC heparanase, this endoglycosidase is taken up by the cardiomyocyte128, 147 to stimulate MMP9 expression and conversion of latent to active TGFβ (Figure 30). 3.3.4 TGFβ can promote cardiomyocyte LPL secretion Given the pivotal role of TGFβ in metabolism151, 152, we treated myocytes with recombinant active TGFβ (1 ng/mL) and discovered an increase in heparin releasable LPL from these cells (Figure 25A), that was unrelated to any change in LPL gene expression (data not shown). As TGFβ has been reported to stimulate RhoA that could regulate actin cytoskeleton remodeling23, 153, cardiomyocytes incubated with TGFβ were used to measure GTP-RhoA, which rapidly increased within 5 minutes of treatment, and declined with time (Figure 25B). The increase in RhoA paralleled the augmented formation of F actin, observations that were also seen with lysophosphatidic acid (LPA) (Figure 25C), a known RhoA activator154. As a TGFβ antibody prevented F actin polymerization (Figure 25C inset), our data suggests that TGFβ activation of RhoA enhances actin cytoskeleton rearrangement to promote LPL trafficking and secretion. 3.3.5 TGFβ induced Meox2 enhances EC GPIHBP1 and promotes LPL translocation TGFβ binding to its receptor on the EC surface triggers downstream Smad signaling (Figure 26). Recently, TGFβ/Smad has been linked to regulation of Meox2 in epithelial cells155. We tested whether TGFβ has a similar role to play in EC Meox2 regulation, and hence GPIHBP1. On exposure of EC to TGFβ for 12 hours, we found an induction of Meox2 protein (Figure 27A and 27B, upper 36 panel) and concurrently, GPIHBP1 mRNA (Figure 27A, inset) and protein (Figure 27B, lower panel), effects that were blocked using a TGFβ antibody. Much of the increase in Meox2 was nuclear (Figure 27B, upper panel). We attempted to inhibit Meox2 induction and nuclear translocation using a TGFβ neutralizing antibody. As anticipated, inhibition of TGFβ signaling reduced nuclear localization of Meox2 (Figure 27B, upper panel) and decreased expression of GPIHBP1 (Figure 27B, lower panel). To test the functional relevance of the TGFβ induced augmentation of GPIHBP1, LPL transport from the basolateral to the apical side of RHMEC monolayers was determined. As expected, there was an increase in heparin-releasable LPL activity at the apical side of EC exposed to TGFβ (Figure 27C). These results suggest that the effect of TGFβ on GPIHBP1 is through up-regulation of Meox2, which results in LPL transfer to the apical side of EC. 37 Figure 5. STZ diabetes increases cardiac LPL activity and GPIHBP1 expression. Rats were made hyperglycemic by i.v. injection of 55 mg/kg streptozotocin (STZ). After 4 days of hyperglycemia, animals were euthanized; hearts removed, and either perfused with heparin (8 U/mL) to release coronary LPL activity (A and upper inset) or used for immunofluorescent detection of LPL (A, lower inset). Images were taken using a 20x objective with the same fluorescence exposure settings. Scale bar, 20 µm. In a separate experiment, both CON and STZ hearts were collected for mRNA (B, left panel) and protein (B, right panel) measurement, and immunofluorescence microscopy of GPIHBP1 (C). Images were taken using a 40x objective with the same fluorescence exposure settings. Scale bar, 20 µm. *p < 0.05, compared to CON, n=3-4. 38 Figure 6. Passaging of endothelial cells cause loss of GPIHBP1 which can be induced on exposure to high glucose. Rat aortic endothelial cells (RAOECs) and bovine coronary artery endothelial cells (BCAECs) were cultured to 80-90% confluence. Cells were detached following trypsin digestion, and aliquots used, either for detection of GPIHBP1 or seeding of a new culture (A). RAOECs from passage 7-10 were treated with increasing concentrations of glucose (B and C, insets) for different times, and GPIHBP1 mRNA (B) and protein (C) determined using TaqMan and Western blotting respectively. *p < 0.05, compared to 0 hour or 5.5 mM glucose, n=3-4. 39 40 Figure 7. GPIHBP1 induced by high glucose increases LPL shuttling across endothelial cell monolayers. RAOECs were seeded on slides were placed in a 6 well plate and treated with 5.5 (CON) or 25 (HG) mM glucose for 12 hours. Slides were then washed with ice cold PBS, fixed with methanol and permeabilized using 0.2% Triton-X in PBS. Anti-GPIHBP1 (1:100) and anti-CD31 antibodies (1:100) were used for immunofluorescence staining. Images were taken using a 40x objective with the same fluorescence exposure settings (A). Scale bar = 20 µm. B represents a cartoon illustrating an experimental design to study functional GPIHBP1 following high glucose. In a separate experiment, RAOEC were grown on transwell inserts and grown until they formed a tight monolayer. Cells were then treated with 5.5 (CON) or 25 (HG) mM glucose for 4 and 12 hours respectively. In some experiments, this was followed by treatment with or without PIPLC (1 U/mL for 1 hour) (C, inset). 25 mM mannitol (Mnt) for 12 hours was used as an osmotic control. Following the indicated times, these Transwell inserts were removed and placed in a different 6 wells plate with DMEM medium on the basolateral side containing 10 µg/ml purified LPL. After incubation for 1 hour, and washing (3x) with PBS, medium containing heparin (8 U/mL) was used for 3 minutes to release apical surface bound LPL. LPL activity was determined by measuring the in vitro hydrolysis of [3H]triolein substrate (C). Using the above protocol (with incubation of HG for 12 hours), the heparin-releasable LPL medium was incubated with increasing concentrations of very low density lipoprotein-triglyceride (VLDL-TG, 0-0.8 mM) at 37°C for 30 minutes and the concentration of released free fatty acid determined. One group of cells was treated with 1 U/mL of phosphatidylinositol-specific phospholipase C (PIPLC) prior to incubation with high glucose (D). *p < 0.05, n=3. 41 Figure 8. Endothelial heparanase can increase GPIHBP1 gene and protein expression. RAOECs from passage 6-10 were used for determination of latent (L-HPA) and active (A-HPA) heparanase (A). Cells from passage 7-10 were treated with HG (25 mM) for 12 hours and medium collected. The medium was concentrated using an Amicon column at 4 °C, and subsequently centrifuged for 15 minutes at 14,000 g. Heparanase in this concentrated medium was determined by Western blot (B). *p < 0.05, n=4. Recombinant A-HPA was used to treat RAOECs (passage 7-10) for 12 hours, cell lysates collected and GPIHBP1 mRNA (C, upper panel) and protein (C, lower panel) determined using TaqMan and Western blot respectively. *p <0.05, n=3-4. 42 Figure 9. Loss of endothelial GPIHBP1 expression can be restored on co-culture with myocytes. RAOECs (A) or BCAECs (B) (passage 8-10) were seeded on transwell inserts and cultured until 80-90% confluence. The transwell insert was then transfered to a 6-well plate containing attached myocytes, and co-cultured for 12 hours. Isolation of myocytes (Myo) was achieved by collagenase digestion of rat hearts, and cells attached to laminin coated 6-well plates in M199 containing 0.1% BSA and kept overnight. After co-culture with myocytes, EC protein was collected and used for Western blot of GPIHBP1. In some experiments, EC (with or without co-culture with myocytes) was exposed to 5.5 or 25 mM glucose for 12 hours and GPIHBP1 determined (C). D represents a cartoon illustrating a co-culture experimental design. *p < 0.05, compared to control; #p < 0.05, compared to normal glucose, n=3-4. 43 Figure 10. EC and cardiomyocyte HSPG anchored proteins can be liberated into culture medium. RAOEC were cultured with or without high glucose (HG, 25 mM) for 12 hours whereas myocytes were incubated with active heparanase (A-HPA) also for 12 hours. With both cell types, the culture medium was collected and tested using a cytokine array (upper panel), with the array proteins listed (lower panel). 44 Figure 11. EC and myocyte HSPG bound PDGF can augment GPIHBP1. RAOEC culture were cultured with or without high glucose (HG, 25 m) for 12 hours whereas myocytes were incubated with or without active heparanase (A-HPA) also for 12 hours. Following a cytokine array, of the proteins detected, platelet derived growth factor (PDGF) was chosen (A). Purified recombinant PDGF (200 ng/ml) was used to treat RAOEC [passage 7 (B) or 10 (B, lower inset)] for 12 hours and GPIHBP1 determined by Western Blot. *p < 0.05, n=3-4. 45 46 Figure 12. High glucose induced increase in GPIHBP1 expression is associated with secretion of heparanase. Rat aortic endothelial cells (RAOECs) from passage fifth to eighth were treated with 5.5 (CON) or 25 (high glucose, HG) mM glucose for 12 hours. GPIHBP1 mRNA (A, left panel) and protein (A, right panel) were determined using TaqMan and Western blot respectively. In a separate experiment, RAOECs were seeded on Transwell inserts and grown until they formed a tight monolayer prior to treatment with 5.5 or 25 mM glucose for 12 hours. These Transwell inserts were then placed in a different 6-well plate with DMEM medium containing 10 µg/mL purified LPL on the basolateral side. After incubation for 1 hour, and washing both the apical and basolateral sides (3x) with PBS, DMEM medium containing heparin (8 U/mL) was used for 3 minutes to release the apical surface-bound LPL. LPL activity was determined by measuring the in vitro hydrolysis of [3H]triolein substrate, whereas LPL mass was determined using Western blot (A, inset). To measure high glucose release of heparanase, RAOECs were seeded and grown until 80-90% confluence, treated with 5.5 or 25 mM glucose for 15 minutes, and medium collected. The medium was concentrated using an Amicon column at 4°C, and subsequently centrifuged for 10 minutes at 14,000 g. Active (A-HPA) and latent (L-HPA) heparanase in this concentrated medium was determined by Western blot (B). RAOECs were also incubated with or without recombinant latent heparanase (L-HPA) for 12 and 24 hours, cell lysates collected, and GPIHBP1 mRNA (C, left panel) and protein (C, right panel) determined. *p < 0.05, ** p < 0.01, compared to CON or 0 h, n=3-4. 47 48 Figure 13. Heparanase released VEGF is an effective stimulator of GPIHBP1 expression. RAOECs were cultured with glucose (5.5 and 25 mM) or latent heparanase (500 ng/ml) for 12 hours, the culture medium collected, and tested using a cytokine protein array. The small circle indicates the signal for vascular endothelial growth factor (VEGF) (A). Purified recombinant VEGF (100 ng/ml) was used to treat RAOECs for 12 and 24 hours, and GPIHBP1 mRNA (B, left panel) and protein (B, right panel) determined. RAOECs seeded on slides were placed in a 6 well plate, treated with or without VEGF (100 ng/ml) for 12 hours. Slides were then washed with ice cold PBS, fixed with methanol and permeabilized using 0.2% Triton-X in PBS. Anti-GPIHBP1 (1:400) and anti-CD31 antibodies (1:400) were used for immunofluorescence staining, and images were taken with a 40x objective and the same fluorescence exposure settings. Scale bar = 20 µm (C). RAOECs were incubated with or without VEGF or a VEGF blocking antibody for 12 hours. Cells were collected and analyzed for GPIHBP1 protein expression. In a separate experiment, LPL translocation was also performed following the above treatment (D and inset). *p < 0.05, compared to CON or 0 h, n=3-4. 49 50 Figure 14. Cardiomyocyte HSPG bound VEGF is crucial for maintaining endothelial cell GPIHBP1 expression. RAOECs were seeded on transwell inserts and cultured until 80-90% confluence. The transwell insert was then transferred to a 6-well plate and co-cultured with or without cardiomyocytes for 12 hours. Isolation of myocytes (Myo) was achieved by collagenase digestion of rat hearts, and these cells attached to laminin coated 6-well plates in M199 containing 1% BSA. After co-culture, EC were collected and GPIHBP1 expression determined using Western blot (A). Cultured cardiomyocytes were treated with or without latent heparanase (500 ng/ml) for 12 hours, the culture medium collected, and tested using a cytokine protein array. The small circle indicates the signal for VEGF (B, left panel). RAOECs and isolated cardiomyocytes were used for determination of VEGF mRNA level using TaqMan (B, right panel). In a separate experiment, isolated cardiomyocytes were incubated with heparin (HEP, 5 U/mL) or latent heparanase (L-HPA, 1-2 µg) for 15 minutes, medium collected, and the secretion of VEGF determined using ELISA assay (C). RAOECs were seeded, grown on transwell inserts, and co-cultured with or without cardiomyocytes. With cardiomyocytes, cells were pretreated with or without heparitinase III (1 U/mL) for 1 hour, prior to co-culture with RAOEC for 12 hours. ECs were collected and GPIHBP1 expression determined. In a separate experiment, LPL translocation was also performed (D and inset). RAOECs were grown on transwell inserts and co-cultured with or without cardiomyocytes for 12 hours. One group of these co-cultured cells were incubated with antibodies (Abs) against VEGF and its receptor, VEGFR2. Cells were collected and GPIHBP1 expression measured by Western blot. In a separate experiment, LPL translocation was also performed (E and inset). *p < 0.05, ** p < 0.01, *** p < 0.001 compared to control, EC or without cardiomyocytes, n=3-5. 51 Figure 15. VEGF induced GPIHBP1 expression is through DLL4-Notch signaling. RAOEC were treated with recombinant VEGF (100 ng/ml) for 12 and 24 hours, and Delta-like ligand (DLL) 4 protein determined (A). RAOEC were also grown on BSA (CON) or DLL4- coated plates for 12 hours. After this time, cells were collected for determination of GPIHBP1 mRNA and protein (B). RAOEC were seeded and grown on slides coated with BSA or DLL4 for 12 hours. Slides were then fixed with methanol, permeabilized using 0.2% Triton-X in PBS, and stained with anti-GPIHBP1 and anti-CD31 antibody (1:400). Images were taken using a 40x objective with the same fluorescence exposure settings. Scale bar = 20 µm (C). RAOECs were grown on plates coated with BSA or DLL4, in the presence or absence of a DLL4 antibody for 12 hours. Cells were collected and analyzed for GPIHBP1 protein expression (D). E represents a cartoon illustrating the mechanism by which VEGFA induces GPIHBP1. *p < 0.05, compared to 0 h or CON, n=3-4. 52 Figure 16. DLL4-Notch pathway downstream target genes were upregulated following DLL4 stimulus. RAOECs were grown on BSA (CON) or DLL4- coated (DLL4) dishes for 12 h. After this time, cells were collected for determination of Hey-1 and Hes-1 mRNA using TaqMan. *p < 0.05, compared to CON, n=4. 53 Figure 17. Inhibition of DLL4 signaling prevents the VEGF-induced GPIHBP1 expression. RAOECs were incubated with or without purified recombinant VEGF (100 ng/mL) or a DLL4 blocking antibody for 12 hours. Cells were collected and analyzed for GPIHBP1 protein expression. n=2-3. 54 Figure 18. Inhibition of Notch signaling downregulates VEGF induced GPIHBP1 expression. RAOECs were grown, treated with or without the γ secretase inhibitor (DAPT) for 2 hours, prior to addition of purified recombinant VEGF (100 ng/ml). Cells were harvested and analyzed for NICD (A, left panel) and GPIHBP1 (A, right panel) protein expression. RAOECs were grown on coverslips placed in 6-well plates, treated with or without the γ secretase inhibitor, DAPT, prior to incubation with recombinant VEGF for 12 hours. Cells on coverslips were washed with ice cold PBS, fixed with methanol and stained with anti-activated Notch1 (NICD) antibody (1:400). Images were taken with a 40x objective. Scale bar = 20 µm (B). C represents a cartoon illustrating how VEGF induced DLL4-Notch signaling induces GPIHBP1 expression. RAOECs were grown on transwell inserts, and pretreated with or without DAPT (1 µg/mL) for 2 hours, prior to co-culture with myocytes for 12 hours. EC were collected, and expression of NICD (D, left panel) and GPIHBP1 (D, right panel) protein determined using anti-GPIHBP1 and anti-activated Notch1 antibodies. *p < 0.05, compared to CON, EC or without cardiomyocytes, n=3-5. 55 Figure 19. General characteristics of different models of diabetes. Rat blood was collected, and following centrifugation, serum was isolated. The concentration of glucose, nonesterified fatty acid (NEFA), triglyceride (TG), and insulin were determined using diagnostic kits. *p < 0.05, ** p < 0.01, compared to CON. *p < 0.05, ** p < 0.01, compared to CON, n= 4-8. 56 Figure 20. Reducing VEGF in diabetes results in lower GPIHBP1 expression. Rats were made hyperglycemic by i.v. injection of 100 mg/kg streptozotocin (STZ, D100). After 4 days, animals were sacrificed, blood collected, serum separated and VEGF determined (A, lower inset). Cardiomyocytes was isolated from control (CON) and D100 rats, RNA isolated and VEGF determined. (A). Isolated cardiomyocytes from CON or D100 were plated on culture dishes for 4 hours. Cell surface VEGF was released using heparin (5 U/mL, HEP) or latent-heparanase (2 µg/mL, L-HPA) for 15 minutes, medium collected, and VEGF determined (B). RAOECs were seeded on transwell inserts and cultured until 80-90% confluence. ECs on the inserts were co-cultured with or without cardiomyocytes from CON or D100.After 12 hours, ECs were collected and GPIHBP1 expression determined (C). In a separate experiment, after four days of hyperglycemia, hearts were removed, and perfused in the Langendorff mode with heparin (8 U/mL) to release coronary LPL activity. LPL activity was determined at the indicated times by measuring the in vitro hydrolysis of [3H]triolein substrate. LPL mass was determined using Western blot (C, inset). RAOECs were seeded on transwell inserts and cultured until 80-90% confluence. ECs on the inserts were pretreated with DAPT for 2 hours to prevent γ secretase activation, and co-cultured with or without cardiomyocytes from CON or D100. After 12 hours, ECs were collected and NICD expression determined using Western blot with anti-activated Notch1 antibody (D). *p < 0.05, ** p < 0.01, compared to CON or without cardiomyocytes, n=4-8. 57 Figure 21. Diabetes increases cardiomyocyte VEGF secretion. Rats were made hyperglycemic by i.v. injection of 55 mg/kg streptozotocin (STZ, D55). After 4 days, animals were euthanized, cardiomyocyte were isolated from both control, and D55 hearts and incubated with heparin (HEP, 5 U/mL) for 15 minutes to release cardiomyocyte surface bound VEGF. Medium were then collected, and concentration of VEGF determined using ELISA assay. *p < 0.05, compared to CON, n=3-4. 58 59 Figure 22. Augmentation of LPL activity in the diabetic heart is associated with an increase of endothelial Meox2. Rats were made hyperglycemic by injection of 55 mg/kg STZ intravenously. After four days, animals were euthanized, and hearts from control (CON) or diabetic (STZ) rats were perfused in the Langendorff model with heparin (8 U/mL) to release coronary LPL (A). At the time when peak LPL activity was observed (between 1-1.5 minutes), medium collected was also used to determine LPL mass using Western blot (A, inset). To compare Meox2 expression, EC and cardiomyocytes were isolated from control rat hearts and protein expression determined (B). Meox2 protein expression and GPIHBP1 mRNA level were also determined in whole hearts (C) and isolated EC (D) from CON and STZ animals. In a separate experiment, isolated EC from the two groups of animals were collected and analyzed for Meox2 expression using immunofluorescent staining (E). Images were taken with a 40x objective (E, left panel), and at higher magnification using 40x objective with 2x digital zoom (E, inset). Scale bar = 20 µm (E, left panel) and 5 µm (E, inset). Vinculin or β-actin were used as loading controls. *p <0.05, compared to CON, n=4-6. 60 61 Figure 23. Endothelial Meox2 induced by high glucose increases GPIHBP1 expression. RHMEC were incubated with 5.5 (CON) or 25 (HG) mM glucose for 8-24 hours (upper panel) prior to assessment of Meox2 (A, upper panel). In a separate experiment, EC were treated with 5-30 mM glucose for 12 hours prior to assessment of Meox2 and GPIHBP1 (A, lower panel). Having determined the optimal incubation time, Meox2 protein and GPIHBP1 mRNA expression was next determined in RHMEC following CON and HG conditions, using a 12 hours incubation time (B). In RHMEC exposed to HG, increasing concentrations (0.1-20%) of FBS (a known inhibitor of Meox2) were added and Meox2 and GPIHBP1 measured after 12 hours (C, left panel). Having determined the optimal inhibitory concentration of FBS, RHMEC were exposed to HG with or without 5% FBS for 12 hours, and Meox2 and GPIHBP1 (C, right panel) determined. In a separate experiment, cells were grown on coverslips and above treatments repeated. Cells were then washed with PBS, fixed with ice-cold methanol, incubated with anit-GPIHBP1 (1:200) and Alexa 488-labeled anti-rabbit IgG (1:400), and stained with DAPI. Images were taken with a 40x objective (C, left lower panel) and the same fluorescence exposure settings. To knockdown Meox2 expression, RHMEC were incubated with 100 pmol scrambled sequence (SCR) or siRNA using lipofectamine for 48 hours (D, inset). These transfected EC were then incubated in the presence of 25 mM glucose for 12 hours, and GPIHBP1 expression determined (D). Vinculin was used as loading control. *p <0.05, compared to CON, n=3-6. #p <0.05, compared with HG, n=3-5. 62 Figure 24. Heparanase induced MMP9 expression is associated with the conversion of latent to active TGFβ in the diabetic heart. Hearts from control (CON) and diabetic (STZ) rats was collected for determination of latent (L-HPA) and active (A-HPA) heparanase using Western blot (A). To examine heparanase secretion following high glucose, RHMECs were grown to 90% confluence and treated with 5.5 (CON) and 25 (HG) mM glucose for 30 minutes. The medium was collected, concentrated, and L-HPA and A-HPA protein determined (A, inset). Isolated cardiomyocytes were treated with L-HPA for 16 hous, and MMP9 mRNA measured using a TaqMan assay (B). MMP9 mRNA and activity were also measured in CON and STZ hearts (B, insets). In a separate experiment, hearts from CON and STZ animals were used to determine TGFβ (latent and active) expression using Western blot (C). Blood from both groups of animals was collected, serum separated, and TGFβ determined (C, inset). *p <0.05, compared to CON, n=4-6. 63 Figure 25. TGFβ induced RhoA-mediated actin cytoskeleton polymerization increases cardiomyocyte LPL secretion. Purified recombinant TGFβ (1 ng/mL) was used to treat cardiomyocytes for 30 and 60 minutes, after which heparin (8 U/mL) was added for 5 minutes to release myocyte surface bound LPL into the medium. LPL activity was determined by measuring the in vitro hydrolysis of [3H]triolein substrate (A). *p < 0.05, compared to the time CON, n=3-4. Myocytes were incubated with TGFβ for 5, 15, and 30 minutes, and RhoA-GTPase measured using a G-LISA assay (B). *p < 0.05, compared to 0 minute, n=3. In a separate experiment, myocytes were treated with or without TGFβ (1 ng/mL) or a TGFβ antibody, or RhoA activator, LPA (1 µM), for 1 hour, and actin cytoskeleton polymerization determined (C). *p <0.05, compared with CON. n=3-7. 64 Figure 26. TGFβ activates downstream Smad signaling. RHMEC were treated with increasing concentrations (0-10 ng/mL) of recombinant TGFβ for 30 minutes. Cells were then collected, and phospho-Smad3 protein determined (upper panel). After determination of concentration, RHMEC were incubated with recombinant TGFβ (1 ng/mL) for different times (0-120 minutes), and phospho-Smad3 protein determined. 65 Figure 27. TGFβ activation of Meox2 enhances GPIHBP1 expression and promotes LPL translocation. Purified recombinant TGFβ (1 ng/mL) was used to treat RHMEC for 12 hour, and Meox2 protein (A) and GPIHBP1 mRNA (A, inset) determined. In a separate experiment, RHMEC were grown on coverslips, pre-treated with or without a TGFβ neutralizing antibody for 2 hour, and then incubated with or without TGFβ (1 ng/mL) for 12 hour (B). Images were taken using a 40x objective with the same fluorescence exposure settings. Scale bar = 20 µm. To study the functional significance of an increase in GPIHBP1, RHMEC were grown on Transwell inserts until they formed a tight monolayer. Cells were then treated with or without TGFβ for 12 hour, and placed in a different 6 well plate with DMEM medium on the basolateral side containing 10 µg/mL purified LPL. After incubation for 1 hour, and washing (3x) with PBS, medium containing heparin (8 U/mL) was used for 3 minutes to release apical surface bound LPL. LPL activity was determined by measuring the in vitro hydrolysis of [3H]triolein substrate (C). *p < 0.05, compared to CON, n=3-4. 66 CHAPTER 4: DISCUSSION Underutilization of glucose and over reliance on FA are hallmarks of the diabetic heart156. Although albumin bound circulating FA is an important source of this substrate, its molar concentration is ~10-fold less than that of lipoprotein TG69. As such, the hydrolysis of circulating TG is suggested to be the predominant source of FA for cardiac utilization during diabetes157. Lipoprotein-TG breakdown occurs at the coronary lumen with the assistance of LPL positioned at apical surface of EC. Following diabetes, in the absence of changes in LPL synthesis, coronary LPL activity is augmented by posttranslational modifications. These include an increase in LPL dimerization121, vesicle transport23, and shuttling of the enzyme from the myocyte surface to the basolateral side of EC99 (Figure 1). Data from the thesis present the novel idea that in response to hyperglycemia, transport of LPL to the apical side of EC, to assist in TG breakdown, is achievable through induction of EC GPIHBP1. Our data suggests a crosstalk that EC, as the first responders to hyperglycemia, can release heparanase to liberate VEGF. This growth factor, by activating EC Notch signaling (Figure 28), is responsible for facilitating translocation of LPL across EC and regulating LPL-derived FA delivery to the cardiomyocytes (Figure 29). High glucose also induced secretion of EC heparanase, which could potentially sustain LPL transferred from cardiomyocyte to EC via promotes MMP9 expression in cardiomyocytes, which activates TGFβ. In the cardiomyocyte, TGFβ activation of RhoA enhances actin cytoskeleton rearrangement to promote LPL trafficking and secretion onto cell surface HSPG. In the EC, TGFβ signaling promotes Meox2 translocation to the nucleus that increases the expression of GPIHBP1, which facilitates movement of LPL to the vascular lumen (Figure 30). Collectively, these adaptations in the cardiomyocyte and EC transfer LPL from the basolateral to the apical (luminal) side of EC where the enzyme is functional, promoting FA delivery to, and utilization by, the cardiomyocyte during diabetes. 67 4.1 Endothelial cells respond to hyperglycemia by increasing the LPL transporter GPIHBP1. Following a single injection of a moderate dose 55 mg/kg of STZ (D55), there is an induction of hypoinsulinemia and hyperglycemia. Increasing the dose to 100 mg/kg (D100) also creates an environment of hyperlipidemia121 (Figure 2). In the former situation, coronary LPL activity is augmented whereas with the latter setting and the presence of higher circulating FA, LPL is turned off 107. With D55 hearts, in the absence of any change in protein synthesis, the increase in LPL activity principally at the vascular lumen could largely be explained by posttranslational modifications that increased the transfer of myocyte enzyme to the EC110. Not yet determined is whether the concluding step, moving LPL across EC, is also increased following diabetes. GPIHBP1, a glycosylphosphatidylinositol-anchored protein expressed exclusively in EC72, is a recent addition to the mechanism that transfers LPL across EC8. GPIHBP1 has a strong binding affinity for LPL, accepting it from the interstitial space and moving it to the coronary lumen158. Intriguingly, the mRNA for GPIHBP1 changes much more rapidly than most mRNAs in mammalian cells76. Our results present the novel observation that to guarantee FA supply to the diabetic heart, the LPL transporter GPIHBP1 increases rapidly in EC. In an attempt to elucidate the mechanism by which diabetes influences GPIHBP1, we exposed two different EC lines to glucose, a substrate whose concentrations increase rapidly following STZ. However, given that under standard cell culture conditions, there has been an insinuation that EC GPIHBP1 is silenced following cell passaging8, 75, we initially attempted to authenticate this speculation. To our surprise, both rat and bovine aortic ECs lose their ability to express GPIHBP1 protein with increasing subculturing of the cells. However, an introduction of high glucose to the culture medium increased GPIHBP1 gene and protein expression in both the early and late cell passages. As high glucose also had comparable effects on LPL shuttling across EC, in association with a greater ability to hydrolyze lipoprotein TG, data from the current study implicate glucose as 68 an important stimulus that influences EC GPIHBP1. Interestingly, high glucose has also been observed to turn on the FA transporters, CD36159 and fatty acid binding protein 4 (FABP4)160 in EC. Thus, with the onset of hyperglycemia, EC, globally, are well adapted to promote FA availability to cardiomyocytes. With EC subculturing, we were interested to discover if proteins other than GPIHBP1 are also switched off. We focused on heparanase, an endoglycosidase that specifically cleaves HS side chains on HSPG140. Intriguingly, in addition to this extracellular function to release bound ligands like LPL, heparanase, by entry into the nucleus to regulate histone acetylation/methylation, is also capable of modulating gene transcription. In this regard, tumor cells, by expressing higher levels of heparanase, are more invasive as secreted heparanase can breakdown extracellular matrix in addition to promoting gene expression in adjacent cells that drives an aggressive tumor phenotype161, 162. Similar to the loss of GPIHBP1, EC passaging also caused nonappearance of heparanase, suggesting a potential connection between these two proteins. Indeed, as an addition of active heparanase to EC augmented gene and protein expression of GPIHBP1, together with the fact that high glucose induced heparanase secretion, our data for the first time links heparanase to the entire process by which LPL progresses forward during diabetes, from the apical side of cardiomyocytes to apical side of EC. Notwithstanding the demonstrated impact of high glucose on GPIHBP1, we were unable to explain the disappearance of this glycoprotein on passaging of EC. Agreed that in the intact heart, there is a robust expression of EC GPIHBP1, it is possible that paracrine signaling influences expression of this glycoprotein8. Major cell types in the heart include fibroblasts, smooth muscles, and myocytes with the latter making up almost 60% of the adult rat myocardium163. We were impressed by our observations that EC that had lost their ability to express GPIHBP1, regained this glycoprotein when co-cultured with cardiomyocytes. As another study has also hinted at this possibility, our data indicate an interaction between cardiomyocytes and EC, likely through 69 paracrine signaling mediators. Currently, it is unclear whether the stimulus for the release of signaling mediators from myocytes could originate from EC. EC are first responders to hyperglycemia, releasing heparanase for ensuing hydrolysis of myocyte HSPG which is linked to the liberation of bound proteins. Given that exposure of the co-culture to high glucose caused an even greater expression of EC GPIHBP1, out data suggest cellular cooperation between EC (via heparanase) and myocytes (through unidentified proteins) that facilitates energy switching from glucose to FA in the diabetic heart. HSPG are ubiquitously present in every tissue compartment, particularly the extracellular matrix, cell surface, intracellular granules, and nucleus24. They consist of a core protein to which several linear HS side chains are covalently linked and could function as anchors due to the high content of charged groups in HS25. The latter property is implicitly used to electrostatically bind a number of different proteins26. Attachment of these bioactive proteins is a clever arrangement, providing the cell with a rapidly accessible reservoir, precluding the need for de novo synthesis when the requirement for a protein is increased140. Heparanase is an endoglycosidase, exceptional in its ability to degrade HS, thereby instigating ligand release. Using a cytokine array and incubation of cardiomyocytes with active heparanase, many cytokines were increased in the myocyte culture medium. We firstly focused on PDGF, as this protein can regulate angiogenesis but also play a significant role in increasing the metabolic reliance of smooth muscle on FA137. Favorably, PDGF increased EC GPIHBP1. As exposure of EC to high glucose also increased PDGF in the medium, likely through an autocrine stimulus of heparanase secretion with release of PDGF bound to EC HSPG, our data suggest that this protein "ensemble" (heparanase-PDGF-GPIHBP1) could cooperate in the diabetic heart to regulate FA delivery and utilization by the cardiomyocyte. 4.2 Cardiomyocyte VEGF regulates endothelial cell GPIHBP1 to relocate lipoprotein lipase to the coronary lumen during diabetes Following moderate diabetes and an increase in vascular LPL, the effect could be explained by a 70 hyperglycemia-induced increase in GPIHBP1 expression138. We further examined the potential mechanism by which hyperglycemia amplifies GPIHBP1. Although the DAG-PKC pathway has been implicated in a direct hyperglycemia-induced gene expression of other proteins164, we tested whether an indirect mechanism could also explain this increase in GPIHBP1 by high glucose. We focused on heparanase as tumor cells transform the phenotype of neighboring cells by "feeding" them heparanase that is capable of modulating their gene expression165. In addition, high glucose is a strong stimulus for secretion of heparanase from EC59. In the current study, incubating EC in HG triggered a rapid release of both active and inactive (latent) heparanase into the medium. Although the HS hydrolyzing ability of active heparanase is well established and can release surface bound proteins, latent heparanase also has some remarkable properties, including a greater competence for releasing myocyte surface VEGF, as compared to active heparanase62. Interestingly, EC exposed to recombinant latent heparanase showed a substantial time-dependent increase in GPIHBP1 gene and protein. As this effect was also duplicated by active heparanase, our data suggests that high glucose secretion of heparanase could be the unforeseen mechanism by which the EC can increase its expression of GPIHBP1 following diabetes. HSPG functions not only as a structural protein but also as an anchor to an amount of proteins that are bound electrostatically to its heparan sulfate side chains24, 25. Attachment of these bioactive proteins (chemokines, enzymes like LPL, and growth factors such as PDGF and VEGF)26 is a clever arrangement, providing the cell with a rapidly accessible reservoir, precluding the need for de novo synthesis when the requirement for a protein is increased. Given that latent heparanase has a greater competence for liberating cell surface protein as compared to active heparanase62, we used a cytokine array and found that some proteins increased in the culture medium following incubation of EC with high glucose or latent heparanase. We focused on VEGF as this protein can regulate angiogenesis166 (which provides the heart with sufficient oxygen to accommodate FA oxidation), but also plays a significant role in FA metabolism141. The latter effect is through promotion of FABP4 71 expression141, a FA transporting protein abundantly expressed in microvascular EC in the heart. Interestingly, EC exposed to recombinant VEGF displayed a substantial time-dependent increase in GPIHBP1 gene and protein, effects that were prevented by a VEGF specific neutralizing antibody. As cardiomyocytes have a stronger VEGF signal compared to EC and a significant amount of surface bound protein that could be easily and rapidly displaced by latent heparanase, we tested the importance of cardiomyocyte VEGF on EC GPIHBP1 expression. Co-culturing of EC with myocytes permitted the EC to regain its GPIHBP1 expression, which was lost after cell passaging. As prior removal of myocyte HSPG with heparitinase or blockade of VEGF signal with neutralizing antibodies prevented this glycoprotein induction in our co-culture system, our data suggest that cardiomyocyte HSPG bound VEGF is a crucial paracrine factor that regulates EC GPIHBP1 expression. The Notch pathway, which mediates cellular homeostasis through its interaction with neighboring cells, is recognized as a key regulator in metabolic organs. Notch signaling is initiated on ligands from one cell binding to the notch receptor on neighboring cells. This recruits a disintegrin and metalloproteinase family peptidase and γ secretase to cleave notch receptor extracellular domain followed by the release of the intracellular domain (NICD), which enters the nucleus and acts as a transcription factor to regulate gene expression143. Given the unanticipated role of VEGF in FA metabolism by activation of DLL4-Notch signaling141, we tested whether this pathway can regulate FA delivery by controlling the LPL transporter GPIHBP1. Incubation of EC with VEGF increased DLL4 expression. Intriguingly, recombinant DLL4 augmented GPIHBP1 gene and protein, effects that were prevented by a DLL4 specific neutralizing antibody. The increase in GPIHBP1 by VEGF was associated with an induction of NICD and its nuclear translocation (Figure 28), effects that were inhibited by a γ secretase inhibitor. As co-culturing of EC with cardiomyocytes induced similar effects, our data suggest that VEGF is a robust stimulus for GPIHBP1 and does so through Notch receptor signaling. 72 Attenuated VEGF production in myocardial tissue is a common feature in both patients and animal models of chronic diabetes145, 167. We generated a rat model of severe diabetes where in addition to hyperglycemia, the animals also demonstrate severe hyperlipidemia. In cardiomyocytes from these animals, we observed a reduction in VEGF gene expression, an outcome that was reflected in a decrease in the myocyte surface bound VEGF pool. Interestingly, co-culture of EC with these myocytes that exhibit muted VEGF caused a significant reduction in the expression GPIHBP1. This effect coincided with a decrease in LPL activity at the coronary lumen of the intact heart from these severely diabetic animals. Unlike the model of severe diabetes (D100), moderate (D55) diabetes induced an increase in LPL activity at the coronary vascular lumen23, 121, an effect that was associated with elevated VEGF at the myocyte surface. Our results implicate a down-regulation of myocyte VEGF as a mechanism to decrease EC GPIHBP1, and by extension, vascular luminal LPL. This could serve as s protective mechanism by which the heart can avoid lipid overload29. 4.3 Dual effects of hyperglycemia on endothelial cells and cardiomyocytes to enhance coronary LPL activity Meox2, is a homeobox gene that is involved in maintaining EC homeostasis during proliferation and VEGF-induced tube formation77, 78. It also forms a heterodimer with another highly expressed cardiac EC transcription factor Tcf15 to regulate EC CD36, GPIHBP1 and LPL expression, and subsequently to facilitate FA uptake and transport across the cardiac endothelium9. We tested whether Meox2 could potentially be implicated in the regulation of vascular LPL during diabetes. Interestingly, following the induction of diabetes and the increase in coronary LPL, there was a parallel amplification in the expression of Meox2 in the heart and EC. As the EC demonstrated robust expression of this transcription factor compared to the cardiomyocyte that has limited expression, our data indicated that the increase in Meox2 in the diabetic heart is likely within the EC. To mimic diabetes, we incubated EC with increasing concentrations of glucose and found that high 73 glucose was a strong stimulus for the induction of Meox2 and GPIHBP1. Intriguingly, as the HG augmented Meox2 and GPIHBP1 expression were prevented by serum, a potent inhibitor of Meox2, or siRNA, our data suggests that to shuttle LPL across the EC, hyperglycemia upregulates Meox2 and GPIHBP1. Although the DAG-PKC pathway has been implicated in a direct hyperglycemia-induced gene expression of other proteins164, we examined whether an indirect mechanism could explain this increase in Meox2 and GPIHBP1 by high glucose. We focused on heparanase as this protein is involved in regulating FA metabolism [it displaces HSPG-bound LPL through hydrolysis of HSPG (active) or ionic displacement (latent)] following diabetes, in addition to being implicated in transforming the phenotype of neighboring cells by controlling their gene expression in cancer [following its nuclear entry and hydrolysis of HSPG, heparanase mitigates the suppressive effect of HS histone acetyltransferase to active gene expression]127, 165. The diabetic heart demonstrated a significant increase in the expression of heparanase, with high glucose being a strong stimulus59 for its secretion from the EC. Incubation of cardiomyocytes with heparanase increased the expression of MMP9, an observation that was also reflected in the diabetic heart. Although MMP9 is capable of shedding HSPG to liberate and sustained myocyte surface-bound LPL128 transfer, this metalloproteinase is also involved in promoting the conversion of latent to active TGFβ148-150, a HSPG bound growth factor that is crucial for cardiogenesis, cardiac repair and remodeling168, 169. Interestingly, as the increase in heparanase in the diabetic heart corresponded to a significant increase in active TGFβ, our data suggests that in response to high glucose and secretion of EC heparanase, this endoglycosidase is taken up by the cardiomyocyte128 to stimulate MMP9 expression and the conversion of latent to active TGFβ. As activation of TGFβ has been implicated in regulating macrophage LPL expression170, we tested whether TGFβ has a similar response in modulating cardiac LPL, and found no effect on the LPL gene in the diabetic heart. However, following heparanase release of LPL from the 74 cardiomyocyte surface, replenishment of this pool of enzyme requires enzyme trafficking from an intracellular site to surface HSPG46, 125, 171, an action that could be under the control of TGFβ. As presumed, in response to TGFβ, there was a significant increase in heparin releasable LPL at the cardiomyocyte surface, an effect that was likely through activation of RhoA which stimulates both the actin and microtubule cytoskeleton rearrangement allowing the movement of LPL. In addition to this effect on the myocyte, we also tested whether TGFβ could influence the EC, specifically Meox2. Interestingly, incubation of EC with TGFβ increased the expression Meox2 and GPIHBP1, effects that were responsible for accelerating translocation of LPL from the basolateral to the apical side of EC. Collectively, our data suggests that TGFβ action on the cardiomyocyte to promote movement of LPL, together with its action on the EC to facilitate LPL shuttling are mechanisms that accelerate FA utilization by the diabetic heart. 75 Figure 28. DLL4 binding to the Notch receptor triggers the Notch pathway to regulate gene expression of GPIHBP1. Delta-like ligand 4 (DLL4), a Notch receptor ligand, is expressed on the signal sending cell. When DLL4 binds to the Notch receptor on the signal receiving cell, there is a recruitment of an a-disintegrin-and-metalloproteinase (ADAM) family peptidase, and a γ secretase to cleave the notch receptor extracellular domain, and release the notch intracellular domain (NICD). NICD enters the nucleus where it binds to and co-activates the transcriptional factor Rbp-Jk to regulate gene expression of GPIHBP1. 76 Figure 29. EC increases its GPIHBP1 in response to hyperglycemia. High glucose induces secretion of endothelial heparanase that stimulates myocyte HSPG bound LPL release, in addition to provoking liberation of VEGF. This growth factor activates DLL4-Notch signaling that contributes towards the augmentation of GPIHBP1. This transporter moves LPL across the EC from the basolateral to its apical side. Out here, LPL is responsible for VLDL-TG hydrolysis to FA, the preferred substrate for the diabetic heart. 77 Figure 30. EC communicates with the cardiomyocyte to increase LPL transfer to the vascular lumen following high glucose. EC, as a first responder to hyperglycemia, releases heparanase from its basolateral side towards the cardiomyocyte. Subsequent uptake of heparanase promotes myocyte MMP9 expression, a matrix-metalloproteinase that facilitates conversion of TGFβ to its active form. In the cardiomyocyte, TGFβ, on binding to its receptor, stimulates RhoA activation that permits cytoskeleton rearrangement and LPL recruitment to the cell surface for onward transfer to the EC. In the EC, TGFβ signaling promotes Meox2 translocation to the nucleus that increases the expression of the LPL transporter, GPIHBP1, which facilitates movement of LPL to the vascular lumen. GPIHBP1 is also a platform for circulating TG hydrolysis. In this way, following diabetes, and the inability of the heart to use glucose as a substrate for energy, it can switch to increase its utilization of FA. 78 CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS 5.1 Conclusions Through their release of heparanase, EC, the first responders to hyperglycemia, are responsible for regulating LPL-derived FA delivery to the cardiomyocytes. Thus, following hyperglycemia, EC releases heparanase from its basolateral side towards the cardiomyocyte. Heparanase has multiple functions that include: a) stimulation of myocyte HSPG bound LPL release, b) autocrine and paracrine actions to provoke liberation of HSPG bound proteins (such as PDGF and VEGF), and c) reuptake into EC and cardiomyocyte to influence gene transcription. This study focused on how the latter two effects contribute towards augmentation of GPIHBP1, the transporter that moves LPL across EC to its apical side. Briefly, high glucose-induced secretion of EC heparanase can liberate myocyte VEGF. This growth factor, by activating EC Notch signaling, is responsible for increasing the LPL transporter GPIHBP1 (Figure 28 and 29). EC heparanase is also taken up by cardiomyocytes128 and promotes MMP9 expression, a matrix metalloproteinase that facilitates conversion of TGFβ to its active form. In the cardiomyocyte, TGFβ, on binding to its receptor, stimulates RhoA activation that permits cytoskeleton rearrangement and LPL recruitment to the cell surface for onward transfer to the EC. In the EC, TGFβ signaling promotes Meox2 translocation to the nucleus that increases the expression of the LPL transporter, GPIHBP1, which facilitates movement of LPL to the vascular lumen (Figure 30). As GPIHBP1 is also a platform for circulating TG hydrolysis, following diabetes, and the inability of the heart to use glucose as a substrate for energy, it can use this glycoprotein to increase its utilization of FA. Although these mechanisms serve to guarantee FA supply and consumption when glucose utilization is compromised, it unintentionally provides a surfeit of FA to the diabetic heart, sponsoring a setting where FA uptake exceeds the mitochondrial oxidative capacity. Chronically, the resulting increase in the conversion of FA to potentially toxic FA metabolites, including ceramides, diacylglycerols, and acylcarnitines, paired with increased formation of reactive oxygen species secondary to elevated FA oxidation, can 79 provoke cardiac cell death (lipotoxicity). Intriguingly, when circulating FA's also increase in addition to glucose, luminal LPL is "turned off" to avoid lipid overload, which itself could lead to cardiac dysfunction (Figure 31). Thus, cardiac LPL homeostasis is of crucial importance for maintaining normal heart function. By gaining further insight into the mechanism(s) by which diabetes could disturb this homeostasis by altering endothelium-bound LPL, we can attempt to piece together a part of the cascade of events leading to diabetic heart disease. Appreciating the mechanism of how the heart regulates its LPL following diabetes should allow the identification of novel targets for therapeutic intervention, to prevent or delay cardiac failure. 5.2 Future Directions To further investigate how endothelial cells manage fatty acid delivery to the heart, the following studies are of potential interest: 1. VEGF plays a role in regulating fatty acid metabolism by activation of DLL4-Notch signaling and Forkhead box protein O1 (FOXO1) mediated gene regulation141. In this study, we determined that the effect of VEGF on GPIHBP1 is through Notch receptor signaling. Whether FOXO1 is involved in the control of GPIHBP1 remains unclear. To test this possibility, FOXO1 siRNA can be used to treat EC prior to high glucose, and GPIHBP1 expression examined. Given that there is an increased nuclear presence of this transcription factor in animal models of hyperglycemia172, FOXO1 siRNA can also be used in our diabetic models to determine whether inhibition of FOXO1 interfere with GPIHBP1 thereby influencing FA delivery in the diabetic heart. 2. In this thesis, we observed that Meox2 could play a role in regulating GPIHBP1. As this transcription factor will form a heterodimer with Tcf15, it is crucial to further investigate the effect of Tcf15 and the heterodimer on EC GPIHBP1. In addition, although LPL functions at the vascular lumen, EC have limited expression of this enzyme. In this study, we mainly focused on cardiomyocyte derived-LPL. A recent study suggests an EC-derived LPL counts ~ 20% activity of the whole heart9. To improve our knowledge of this enzyme, the measurement of EC-derived LPL 80 from the primary isolated cardiac EC is essential. Because Meox2/Tcf15 is involved in regulating cardiac FA metabolism9, it is of interest to investigate whether this heterodimer plays a role in regulating LPL synthesis in EC and whether EC LPL has a role to play in fatty acid delivery to the diabetic heart. 3. In this thesis, we predominantly focused on the acute or moderate diabetes (D55) with an increase in LPL activity at the coronary vascular lumen23, 121 that is associated with an increase in GPIHBP1. Hyperglycemia is an important stimulus for this glycoprotein induction. In contrast to D55, our rat model (D100) to mimic severe diabetes with hypoinsulinemia, hyperglycemia, and hyperlipidemia demonstrated a coronary LPL activity reduction121. Given intralipid infusion in the control rats' demonstrated a LPL accumulation at the basement membranes adjacent to EC in the hearts173, it suggests that lipid overload may be involved in regulating cardiac FA metabolism through modulating LPL. The role of GPIHBP1 and TGFβ/Meox2 in LPL function following severe diabetes (or lipid overload) has yet to be studied. It will be of some interest to investigate whether this heterodimer plays a role in LPL transfer following severe diabetes, and determine how FA influence LPL-mediated lipolysis through regulation of GPIHBP1. 81 Figure 31. Pathological oscillations in coronary luminal LPL following diabetes can lead to cardiomyopathy. Diabetes is a unique metabolic disorder during which both an increase or decrease in vascular LPL activity are observed. Increased LPL activity induces lipid accumulation in the cardiomyocytes. Decreased LPL at the coronary lumen is also detrimental as neither glucose nor albumin-bound free fatty acid can meet the energy demand of the heart. In both scenarios, cardiomyopathy is the end result of these long-term metabolic changes121. 82 REFERENCES 1. Voshol PJ, Rensen PC, van Dijk KW, Romijn JA, Havekes LM. Effect of plasma triglyceride metabolism on lipid storage in adipose tissue: Studies using genetically engineered mouse models. Biochimica et biophysica acta. 2009;1791:479-485 2. Avogaro A, Nosadini R, Doria A, Fioretto P, Velussi M, Vigorito C, Sacca L, Toffolo G, Cobelli C, Trevisan R, et al. Myocardial metabolism in insulin-deficient diabetic humans without coronary artery disease. The American journal of physiology. 1990;258:E606-618 3. Saddik M, Lopaschuk GD. Myocardial triglyceride turnover and contribution to energy substrate utilization in isolated working rat hearts. The Journal of biological chemistry. 1991;266:8162-8170 4. Lopaschuk GD, Belke DD, Gamble J, Itoi T, Schonekess BO. Regulation of fatty acid oxidation in the mammalian heart in health and disease. Biochimica et biophysica acta. 1994;1213:263-276 5. Paulson DJ, Crass MF, 3rd. Endogenous triacylglycerol metabolism in diabetic heart. The American journal of physiology. 1982;242:H1084-1094 6. Blanchette-Mackie EJ, Dwyer NK, Amende LA. Cytochemical studies of lipid metabolism: Immunogold probes for lipoprotein lipase and cholesterol. The American journal of anatomy. 1989;185:255-263 7. Eckel RH. Lipoprotein lipase. A multifunctional enzyme relevant to common metabolic diseases. The New England journal of medicine. 1989;320:1060-1068 8. Young SG, Davies BS, Voss CV, Gin P, Weinstein MM, Tontonoz P, Reue K, Bensadoun A, Fong LG, Beigneux AP. Gpihbp1, an endothelial cell transporter for lipoprotein lipase. Journal of lipid research. 2011;52:1869-1884 9. Coppiello G, Collantes M, Sirerol-Piquer MS, Vandenwijngaert S, Schoors S, Swinnen M, Vandersmissen I, Herijgers P, Topal B, van Loon J, Goffin J, Prosper F, Carmeliet P, Garcia-Verdugo JM, Janssens S, Penuelas I, Aranguren XL, Luttun A. Meox2/tcf15 heterodimers program the heart capillary endothelium for cardiac fatty acid uptake. Circulation. 2015;131:815-826 10. Semenkovich CF, Luo CC, Nakanishi MK, Chen SH, Smith LC, Chan L. In vitro expression and site-specific mutagenesis of the cloned human lipoprotein lipase gene. Potential n-linked glycosylation site asparagine 43 is important for both enzyme activity and secretion. The Journal of biological chemistry. 1990;265:5429-5433 11. Ben-Zeev O, Mao HZ, Doolittle MH. Maturation of lipoprotein lipase in the endoplasmic reticulum. Concurrent formation of functional dimers and inactive aggregates. The Journal of biological chemistry. 2002;277:10727-10738 12. Dallinga-Thie GM, Franssen R, Mooij HL, Visser ME, Hassing HC, Peelman F, Kastelein JJ, Peterfy M, Nieuwdorp M. The metabolism of triglyceride-rich lipoproteins revisited: New players, new insight. Atherosclerosis. 2010;211:1-8 13. Doolittle MH, Ehrhardt N, Peterfy M. Lipase maturation factor 1: Structure and role in lipase folding and assembly. Current opinion in lipidology. 2010;21:198-203 14. Garfinkel AS, Kempner ES, Ben-Zeev O, Nikazy J, James SJ, Schotz MC. Lipoprotein lipase: Size of the functional unit determined by radiation inactivation. Journal of lipid research. 1983;24:775-780 15. Peterson J, Fujimoto WY, Brunzell JD. Human lipoprotein lipase: Relationship of activity, heparin affinity, and conformation as studied with monoclonal antibodies. Journal of lipid research. 1992;33:1165-1170 83 16. Wong H, Yang D, Hill JS, Davis RC, Nikazy J, Schotz MC. A molecular biology-based approach to resolve the subunit orientation of lipoprotein lipase. Proceedings of the National Academy of Sciences of the United States of America. 1997;94:5594-5598 17. Sha H, Sun S, Francisco AB, Ehrhardt N, Xue Z, Liu L, Lawrence P, Mattijssen F, Guber RD, Panhwar MS, Brenna JT, Shi H, Xue B, Kersten S, Bensadoun A, Peterfy M, Long Q, Qi L. The er-associated degradation adaptor protein sel1l regulates lpl secretion and lipid metabolism. Cell metabolism. 2014;20:458-470 18. Peterfy M. Lipase maturation factor 1: A lipase chaperone involved in lipid metabolism. Biochimica et biophysica acta. 2012;1821:790-794 19. Peterfy M, Ben-Zeev O, Mao HZ, Weissglas-Volkov D, Aouizerat BE, Pullinger CR, Frost PH, Kane JP, Malloy MJ, Reue K, Pajukanta P, Doolittle MH. Mutations in lmf1 cause combined lipase deficiency and severe hypertriglyceridemia. Nature genetics. 2007;39:1483-1487 20. Kim MS, Wang F, Puthanveetil P, Kewalramani G, Hosseini-Beheshti E, Ng N, Wang Y, Kumar U, Innis S, Proud CG, Abrahani A, Rodrigues B. Protein kinase d is a key regulator of cardiomyocyte lipoprotein lipase secretion after diabetes. Circulation research. 2008;103:252-260 21. Lu G, Chen J, Espinoza LA, Garfield S, Toshiyuki S, Akiko H, Huppler A, Wang QJ. Protein kinase d 3 is localized in vesicular structures and interacts with vesicle-associated membrane protein 2. Cellular signalling. 2007;19:867-879 22. Ohira M, Miyashita Y, Murano T, Watanabe F, Shirai K. Metformin promotes induction of lipoprotein lipase in skeletal muscle through activation of adenosine monophosphate-activated protein kinase. Metabolism: clinical and experimental. 2009;58:1408-1414 23. Kim MS, Kewalramani G, Puthanveetil P, Lee V, Kumar U, An D, Abrahani A, Rodrigues B. Acute diabetes moderates trafficking of cardiac lipoprotein lipase through p38 mitogen-activated protein kinase-dependent actin cytoskeleton organization. Diabetes. 2008;57:64-76 24. Iozzo RV, San Antonio JD. Heparan sulfate proteoglycans: Heavy hitters in the angiogenesis arena. The Journal of clinical investigation. 2001;108:349-355 25. Iozzo RV. Heparan sulfate proteoglycans: Intricate molecules with intriguing functions. The Journal of clinical investigation. 2001;108:165-167 26. Lee TY, Folkman J, Javaherian K. Hspg-binding peptide corresponding to the exon 6a-encoded domain of vegf inhibits tumor growth by blocking angiogenesis in murine model. PloS one. 2010;5:e9945 27. Davies BS, Beigneux AP, Barnes RH, 2nd, Tu Y, Gin P, Weinstein MM, Nobumori C, Nyren R, Goldberg I, Olivecrona G, Bensadoun A, Young SG, Fong LG. Gpihbp1 is responsible for the entry of lipoprotein lipase into capillaries. Cell metabolism. 2010;12:42-52 28. Bengtsson G, Olivecrona T. Lipoprotein lipase. Mechanism of product inhibition. European journal of biochemistry / FEBS. 1980;106:557-562 29. Kim MS, Wang F, Puthanveetil P, Kewalramani G, Innis S, Marzban L, Steinberg SF, Webber TD, Kieffer TJ, Abrahani A, Rodrigues B. Cleavage of protein kinase d after acute hypoinsulinemia prevents excessive lipoprotein lipase-mediated cardiac triglyceride accumulation. Diabetes. 2009;58:2464-2475 30. Koster A, Chao YB, Mosior M, Ford A, Gonzalez-DeWhitt PA, Hale JE, Li D, Qiu Y, Fraser CC, Yang DD, Heuer JG, Jaskunas SR, Eacho P. Transgenic angiopoietin-like (angptl)4 overexpression and targeted disruption of angptl4 and angptl3: Regulation of triglyceride metabolism. Endocrinology. 2005;146:4943-4950 31. Yau MH, Wang Y, Lam KS, Zhang J, Wu D, Xu A. A highly conserved motif within the nh2-terminal coiled-coil domain of angiopoietin-like protein 4 confers its inhibitory effects 84 on lipoprotein lipase by disrupting the enzyme dimerization. The Journal of biological chemistry. 2009;284:11942-11952 32. Liu J, Afroza H, Rader DJ, Jin W. Angiopoietin-like protein 3 inhibits lipoprotein lipase activity through enhancing its cleavage by proprotein convertases. The Journal of biological chemistry. 2010;285:27561-27570 33. Nakajima K, Kobayashi J, Mabuchi H, Nakano T, Tokita Y, Nagamine T, Imamura S, Ai M, Otokozawa S, Schaefer EF. Association of angiopoietin-like protein 3 with hepatic triglyceride lipase and lipoprotein lipase activities in human plasma. Annals of clinical biochemistry. 2010;47:423-431 34. Yu X, Burgess SC, Ge H, Wong KK, Nassem RH, Garry DJ, Sherry AD, Malloy CR, Berger JP, Li C. Inhibition of cardiac lipoprotein utilization by transgenic overexpression of angptl4 in the heart. Proceedings of the National Academy of Sciences of the United States of America. 2005;102:1767-1772 35. Yin W, Romeo S, Chang S, Grishin NV, Hobbs HH, Cohen JC. Genetic variation in angptl4 provides insights into protein processing and function. The Journal of biological chemistry. 2009;284:13213-13222 36. Sukonina V, Lookene A, Olivecrona T, Olivecrona G. Angiopoietin-like protein 4 converts lipoprotein lipase to inactive monomers and modulates lipase activity in adipose tissue. Proceedings of the National Academy of Sciences of the United States of America. 2006;103:17450-17455 37. Staiger H, Haas C, Machann J, Werner R, Weisser M, Schick F, Machicao F, Stefan N, Fritsche A, Haring HU. Muscle-derived angiopoietin-like protein 4 is induced by fatty acids via peroxisome proliferator-activated receptor (ppar)-delta and is of metabolic relevance in humans. Diabetes. 2009;58:579-589 38. Chi X, Shetty SK, Shows HW, Hjelmaas AJ, Malcolm EK, Davies BS. Angiopoietin-like 4 modifies the interactions between lipoprotein lipase and its endothelial cell transporter gpihbp1. The Journal of biological chemistry. 2015;290:11865-11877 39. Sonnenburg WK, Yu D, Lee EC, Xiong W, Gololobov G, Key B, Gay J, Wilganowski N, Hu Y, Zhao S, Schneider M, Ding ZM, Zambrowicz BP, Landes G, Powell DR, Desai U. Gpihbp1 stabilizes lipoprotein lipase and prevents its inhibition by angiopoietin-like 3 and angiopoietin-like 4. Journal of lipid research. 2009;50:2421-2429 40. Deshaies Y, Arnold J, Lalonde J, Richard D. Lipoprotein lipase in white and brown adipose tissues of exercised rats fed a high-fat diet. The American journal of physiology. 1988;255:R226-231 41. Deshaies Y, Richard D, Arnold J. Lipoprotein lipase in adipose tissues of exercise-trained, cold-acclimated rats. The American journal of physiology. 1986;251:E251-257 42. Rogers MP, Zhao X. Lipoprotein lipase mrna levels in adipose tissue in lactation. Biochemical Society transactions. 1996;24:169S 43. Rogers MP, Zhao X. Effect of insulin and serum on lipoprotein lipase in lactation. Biochemical Society transactions. 1998;26:S145 44. Wu G, Olivecrona G, Olivecrona T. The distribution of lipoprotein lipase in rat adipose tissue. Changes with nutritional state engage the extracellular enzyme. The Journal of biological chemistry. 2003;278:11925-11930 45. Doolittle MH, Ben-Zeev O, Elovson J, Martin D, Kirchgessner TG. The response of lipoprotein lipase to feeding and fasting. Evidence for posttranslational regulation. The Journal of biological chemistry. 1990;265:4570-4577 46. Kim MS, Wang Y, Rodrigues B. Lipoprotein lipase mediated fatty acid delivery and its impact in diabetic cardiomyopathy. Biochimica et biophysica acta. 2012;1821:800-808 85 47. Bertoni AG, Tsai A, Kasper EK, Brancati FL. Diabetes and idiopathic cardiomyopathy: A nationwide case-control study. Diabetes care. 2003;26:2791-2795 48. Poirier P, Bogaty P, Garneau C, Marois L, Dumesnil JG. Diastolic dysfunction in normotensive men with well-controlled type 2 diabetes: Importance of maneuvers in echocardiographic screening for preclinical diabetic cardiomyopathy. Diabetes care. 2001;24:5-10 49. Struthers AD, Morris AD. Screening for and treating left-ventricular abnormalities in diabetes mellitus: A new way of reducing cardiac deaths. Lancet. 2002;359:1430-1432 50. Augustus A, Yagyu H, Haemmerle G, Bensadoun A, Vikramadithyan RK, Park SY, Kim JK, Zechner R, Goldberg IJ. Cardiac-specific knock-out of lipoprotein lipase alters plasma lipoprotein triglyceride metabolism and cardiac gene expression. The Journal of biological chemistry. 2004;279:25050-25057 51. Augustus AS, Buchanan J, Park TS, Hirata K, Noh HL, Sun J, Homma S, D'Armiento J, Abel ED, Goldberg IJ. Loss of lipoprotein lipase-derived fatty acids leads to increased cardiac glucose metabolism and heart dysfunction. The Journal of biological chemistry. 2006;281:8716-8723 52. Noh HL, Okajima K, Molkentin JD, Homma S, Goldberg IJ. Acute lipoprotein lipase deletion in adult mice leads to dyslipidemia and cardiac dysfunction. American journal of physiology. Endocrinology and metabolism. 2006;291:E755-760 53. Vreys V, David G. Mammalian heparanase: What is the message? Journal of cellular and molecular medicine. 2007;11:427-452 54. Fairbanks MB, Mildner AM, Leone JW, Cavey GS, Mathews WR, Drong RF, Slightom JL, Bienkowski MJ, Smith CW, Bannow CA, Heinrikson RL. Processing of the human heparanase precursor and evidence that the active enzyme is a heterodimer. The Journal of biological chemistry. 1999;274:29587-29590 55. Gingis-Velitski S, Zetser A, Kaplan V, Ben-Zaken O, Cohen E, Levy-Adam F, Bashenko Y, Flugelman MY, Vlodavsky I, Ilan N. Heparanase uptake is mediated by cell membrane heparan sulfate proteoglycans. The Journal of biological chemistry. 2004;279:44084-44092 56. Shafat I, Vlodavsky I, Ilan N. Characterization of mechanisms involved in secretion of active heparanase. The Journal of biological chemistry. 2006;281:23804-23811 57. Eccles SA. Heparanase: Breaking down barriers in tumors. Nature medicine. 1999;5:735-736 58. Vlodavsky I, Beckhove P, Lerner I, Pisano C, Meirovitz A, Ilan N, Elkin M. Significance of heparanase in cancer and inflammation. Cancer microenvironment : official journal of the International Cancer Microenvironment Society. 2012;5:115-132 59. Wang F, Kim MS, Puthanveetil P, Kewalramani G, Deppe S, Ghosh S, Abrahani A, Rodrigues B. Endothelial heparanase secretion after acute hypoinsulinemia is regulated by glucose and fatty acid. American journal of physiology. Heart and circulatory physiology. 2009;296:H1108-1116 60. Abboud-Jarrous G, Rangini-Guetta Z, Aingorn H, Atzmon R, Elgavish S, Peretz T, Vlodavsky I. Site-directed mutagenesis, proteolytic cleavage, and activation of human proheparanase. The Journal of biological chemistry. 2005;280:13568-13575 61. Pikas DS, Li JP, Vlodavsky I, Lindahl U. Substrate specificity of heparanases from human hepatoma and platelets. The Journal of biological chemistry. 1998;273:18770-18777 62. Zhang D, Wan A, Chiu AP, Wang Y, Wang F, Neumaier K, Lal N, Bround MJ, Johnson JD, Vlodavsky I, Rodrigues B. Hyperglycemia-induced secretion of endothelial heparanase stimulates a vascular endothelial growth factor autocrine network in cardiomyocytes that promotes recruitment of lipoprotein lipase. Arteriosclerosis, thrombosis, and vascular biology. 2013;33:2830-2838 86 63. Tammela T, Enholm B, Alitalo K, Paavonen K. The biology of vascular endothelial growth factors. Cardiovascular research. 2005;65:550-563 64. Weis SM, Cheresh DA. Pathophysiological consequences of vegf-induced vascular permeability. Nature. 2005;437:497-504 65. Hagberg CE, Falkevall A, Wang X, Larsson E, Huusko J, Nilsson I, van Meeteren LA, Samen E, Lu L, Vanwildemeersch M, Klar J, Genove G, Pietras K, Stone-Elander S, Claesson-Welsh L, Yla-Herttuala S, Lindahl P, Eriksson U. Vascular endothelial growth factor b controls endothelial fatty acid uptake. Nature. 2010;464:917-921 66. Hoeben A, Landuyt B, Highley MS, Wildiers H, Van Oosterom AT, De Bruijn EA. Vascular endothelial growth factor and angiogenesis. Pharmacological reviews. 2004;56:549-580 67. Nadir Y, Brenner B, Zetser A, Ilan N, Shafat I, Zcharia E, Goldshmidt O, Vlodavsky I. Heparanase induces tissue factor expression in vascular endothelial and cancer cells. Journal of thrombosis and haemostasis : JTH. 2006;4:2443-2451 68. Cryer A. The role of the endothelium in myocardial lipoprotein dynamics. Molecular and cellular biochemistry. 1989;88:7-15 69. Merkel M, Eckel RH, Goldberg IJ. Lipoprotein lipase: Genetics, lipid uptake, and regulation. Journal of lipid research. 2002;43:1997-2006 70. Obunike JC, Lutz EP, Li Z, Paka L, Katopodis T, Strickland DK, Kozarsky KF, Pillarisetti S, Goldberg IJ. Transcytosis of lipoprotein lipase across cultured endothelial cells requires both heparan sulfate proteoglycans and the very low density lipoprotein receptor. The Journal of biological chemistry. 2001;276:8934-8941 71. Beigneux AP, Davies BS, Gin P, Weinstein MM, Farber E, Qiao X, Peale F, Bunting S, Walzem RL, Wong JS, Blaner WS, Ding ZM, Melford K, Wongsiriroj N, Shu X, de Sauvage F, Ryan RO, Fong LG, Bensadoun A, Young SG. Glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 plays a critical role in the lipolytic processing of chylomicrons. Cell metabolism. 2007;5:279-291 72. Ioka RX, Kang MJ, Kamiyama S, Kim DH, Magoori K, Kamataki A, Ito Y, Takei YA, Sasaki M, Suzuki T, Sasano H, Takahashi S, Sakai J, Fujino T, Yamamoto TT. Expression cloning and characterization of a novel glycosylphosphatidylinositol-anchored high density lipoprotein-binding protein, gpi-hbp1. The Journal of biological chemistry. 2003;278:7344-7349 73. Gin P, Yin L, Davies BS, Weinstein MM, Ryan RO, Bensadoun A, Fong LG, Young SG, Beigneux AP. The acidic domain of gpihbp1 is important for the binding of lipoprotein lipase and chylomicrons. The Journal of biological chemistry. 2008;283:29554-29562 74. Beigneux AP, Gin P, Davies BS, Weinstein MM, Bensadoun A, Ryan RO, Fong LG, Young SG. Glycosylation of asn-76 in mouse gpihbp1 is critical for its appearance on the cell surface and the binding of chylomicrons and lipoprotein lipase. Journal of lipid research. 2008;49:1312-1321 75. Davies BS, Waki H, Beigneux AP, Farber E, Weinstein MM, Wilpitz DC, Tai LJ, Evans RM, Fong LG, Tontonoz P, Young SG. The expression of gpihbp1, an endothelial cell binding site for lipoprotein lipase and chylomicrons, is induced by peroxisome proliferator-activated receptor-gamma. Molecular endocrinology. 2008;22:2496-2504 76. Kroupa O, Vorrsjo E, Stienstra R, Mattijssen F, Nilsson SK, Sukonina V, Kersten S, Olivecrona G, Olivecrona T. Linking nutritional regulation of angptl4, gpihbp1, and lmf1 to lipoprotein lipase activity in rodent adipose tissue. BMC physiology. 2012;12:13 77. Chen Y, Rabson AB, Gorski DH. Meox2 regulates nuclear factor-kappab activity in vascular endothelial cells through interactions with p65 and ikappabbeta. Cardiovascular research. 2010;87:723-731 87 78. Gorski DH, Leal AJ. Inhibition of endothelial cell activation by the homeobox gene gax. The Journal of surgical research. 2003;111:91-99 79. Francis GS. Diabetic cardiomyopathy: Fact or fiction? Heart. 2001;85:247-248 80. Goraya TY, Leibson CL, Palumbo PJ, Weston SA, Killian JM, Pfeifer EA, Jacobsen SJ, Frye RL, Roger VL. Coronary atherosclerosis in diabetes mellitus: A population-based autopsy study. J. Am. Coll. Cardiol. 2002;40:946-953 81. Regan TJ, Ahmed S, Haider B, Moschos C, Weisse A. Diabetic cardiomyopathy: Experimental and clinical observations. N. J. Med. 1994;91:776-778 82. Boudina S, Abel ED. Diabetic cardiomyopathy, causes and effects. Rev. Endocr. Metab. Disord. 2010;11:31-39 83. Fang ZY, Prins JB, Marwick TH. Diabetic cardiomyopathy: Evidence, mechanisms, and therapeutic implications. Endocr. Rev. 2004;25:543-567 84. Bugger H, Abel ED. Rodent models of diabetic cardiomyopathy. Dis. Model. Mech. 2009;2:454-466 85. Severson DL. Diabetic cardiomyopathy: Recent evidence from mouse models of type 1 and type 2 diabetes. Can. J. Physiol. Pharmacol. 2004;82:813-823 86. Shehadeh A, Regan TJ. Cardiac consequences of diabetes mellitus. Clin. Cardiol. 1995;18:301-305 87. Fein FS, Sonnenblick EH. Diabetic cardiomyopathy. Prog. Cardiovasc. Dis. 1985;27:255-270 88. Dhalla NS, Liu X, Panagia V, Takeda N. Subcellular remodeling and heart dysfunction in chronic diabetes. Cardiovasc. Res. 1998;40:239-247 89. Taha M, Lopaschuk GD. Alterations in energy metabolism in cardiomyopathies. Ann. Med. 2007;39:594-607 90. Sung MM, Hamza SM, Dyck JR. Myocardial metabolism in diabetic cardiomyopathy: Potential therapeutic targets. Antioxidants & redox signaling. 2015;22:1606-1630 91. Wan A, Rodrigues B. Endothelial cell-cardiomyocyte crosstalk in diabetic cardiomyopathy. Cardiovasc. Res. 2016;111:172-183 92. Kraegen EW, Sowden JA, Halstead MB, Clark PW, Rodnick KJ, Chisholm DJ, James DE. Glucose transporters and in vivo glucose uptake in skeletal and cardiac muscle: Fasting, insulin stimulation and immunoisolation studies of glut1 and glut4. The Biochemical journal. 1993;295 ( Pt 1):287-293 93. Pessin JE, Bell GI. Mammalian facilitative glucose transporter family: Structure and molecular regulation. Annual review of physiology. 1992;54:911-930 94. Li J, Hu X, Selvakumar P, Russell RR, 3rd, Cushman SW, Holman GD, Young LH. Role of the nitric oxide pathway in ampk-mediated glucose uptake and glut4 translocation in heart muscle. American journal of physiology. Endocrinology and metabolism. 2004;287:E834-841 95. Yang J, Holman GD. Insulin and contraction stimulate exocytosis, but increased amp-activated protein kinase activity resulting from oxidative metabolism stress slows endocytosis of glut4 in cardiomyocytes. The Journal of biological chemistry. 2005;280:4070-4078 96. Camps M, Castello A, Munoz P, Monfar M, Testar X, Palacin M, Zorzano A. Effect of diabetes and fasting on glut-4 (muscle/fat) glucose-transporter expression in insulin-sensitive tissues. Heterogeneous response in heart, red and white muscle. The Biochemical journal. 1992;282 ( Pt 3):765-772 97. Young ME, Guthrie PH, Razeghi P, Leighton B, Abbasi S, Patil S, Youker KA, Taegtmeyer H. Impaired long-chain fatty acid oxidation and contractile dysfunction in the obese zucker rat heart. Diabetes. 2002;51:2587-2595 88 98. Lopaschuk GD, Ussher JR, Folmes CD, Jaswal JS, Stanley WC. Myocardial fatty acid metabolism in health and disease. Physiological reviews. 2010;90:207-258 99. Blanchette-Mackie EJ, Masuno H, Dwyer NK, Olivecrona T, Scow RO. Lipoprotein lipase in myocytes and capillary endothelium of heart: Immunocytochemical study. The American journal of physiology. 1989;256:E818-828 100. Finck BN, Lehman JJ, Leone TC, Welch MJ, Bennett MJ, Kovacs A, Han X, Gross RW, Kozak R, Lopaschuk GD, Kelly DP. The cardiac phenotype induced by pparalpha overexpression mimics that caused by diabetes mellitus. The Journal of clinical investigation. 2002;109:121-130 101. Wende AR, Abel ED. Lipotoxicity in the heart. Biochimica et biophysica acta. 2010;1801:311-319 102. Pulinilkunnil T, Rodrigues B. Cardiac lipoprotein lipase: Metabolic basis for diabetic heart disease. Cardiovascular research. 2006;69:329-340 103. Zechner R, Kienesberger PC, Haemmerle G, Zimmermann R, Lass A. Adipose triglyceride lipase and the lipolytic catabolism of cellular fat stores. Journal of lipid research. 2009;50:3-21 104. Kashiwazaki K, Hirano T, Yoshino G, Kurokawa M, Tajima H, Adachi M. Decreased release of lipoprotein lipase is associated with vascular endothelial damage in niddm patients with microalbuminuria. Diabetes care. 1998;21:2016-2020 105. Taskinen MR, Nikkila EA. Lipoprotein lipase activity of adipose tissue and skeletal muscle in insulin-deficient human diabetes. Relation to high-density and very-low-density lipoproteins and response to treatment. Diabetologia. 1979;17:351-356 106. Williams SP, Johnson EA. Release of lipoprotein lipase and hepatic triglyceride lipase in rats by heparin and other sulphated polysaccharides. Thrombosis research. 1989;55:361-368 107. Rodrigues B, Cam MC, Jian K, Lim F, Sambandam N, Shepherd G. Differential effects of streptozotocin-induced diabetes on cardiac lipoprotein lipase activity. Diabetes. 1997;46:1346-1353 108. Boivin A, Deshaies Y. Contribution of hyperinsulinemia to modulation of lipoprotein lipase activity in the obese zucker rat. Metabolism: clinical and experimental. 2000;49:134-140 109. Kobayashi K, Forte TM, Taniguchi S, Ishida BY, Oka K, Chan L. The db/db mouse, a model for diabetic dyslipidemia: Molecular characterization and effects of western diet feeding. Metabolism: clinical and experimental. 2000;49:22-31 110. Pulinilkunnil T, Qi D, Ghosh S, Cheung C, Yip P, Varghese J, Abrahani A, Brownsey R, Rodrigues B. Circulating triglyceride lipolysis facilitates lipoprotein lipase translocation from cardiomyocyte to myocardial endothelial lining. Cardiovascular research. 2003;59:788-797 111. Qi D, Pulinilkunnil T, An D, Ghosh S, Abrahani A, Pospisilik JA, Brownsey R, Wambolt R, Allard M, Rodrigues B. Single-dose dexamethasone induces whole-body insulin resistance and alters both cardiac fatty acid and carbohydrate metabolism. Diabetes. 2004;53:1790-1797 112. Neitzel AS, Carley AN, Severson DL. Chylomicron and palmitate metabolism by perfused hearts from diabetic mice. American journal of physiology. Endocrinology and metabolism. 2003;284:E357-365 113. Kako Y, Huang LS, Yang J, Katopodis T, Ramakrishnan R, Goldberg IJ. Streptozotocin-induced diabetes in human apolipoprotein b transgenic mice. Effects on lipoproteins and atherosclerosis. Journal of lipid research. 1999;40:2185-2194 114. Pulinilkunnil T, Abrahani A, Varghese J, Chan N, Tang I, Ghosh S, Kulpa J, Allard M, Brownsey R, Rodrigues B. Evidence for rapid "metabolic switching" through lipoprotein lipase occupation of endothelial-binding sites. Journal of molecular and cellular cardiology. 2003;35:1093-1103 89 115. Murthy VK, Bauman MD, Shipp JC. Regulation of triacylglycerol lipolysis in the perfused hearts of normal and diabetic rats. Diabetes. 1983;32:718-722 116. Nomura T, Hagino Y, Gotoh M, Iguchi A, Sakamoto N. The effects of streptozotocin diabetes on tissue specific lipase activities in the rat. Lipids. 1984;19:594-599 117. O'Looney P, Vander Maten M, Vahouny GV. Insulin-mediated modifications of myocardial lipoprotein lipase and lipoprotein metabolism. The Journal of biological chemistry. 1983;258:12994-13001 118. Rodrigues B, Severson DL. Acute diabetes does not reduce heparin-releasable lipoprotein lipase activity in perfused hearts from wistar-kyoto rats. Canadian journal of physiology and pharmacology. 1993;71:657-661 119. Tavangar K, Murata Y, Pedersen ME, Goers JF, Hoffman AR, Kraemer FB. Regulation of lipoprotein lipase in the diabetic rat. The Journal of clinical investigation. 1992;90:1672-1678 120. Niu YG, Evans RD. Metabolism of very-low-density lipoprotein and chylomicrons by streptozotocin-induced diabetic rat heart: Effects of diabetes and lipoprotein preference. American journal of physiology. Endocrinology and metabolism. 2008;295:E1106-1116 121. Wang Y, Puthanveetil P, Wang F, Kim MS, Abrahani A, Rodrigues B. Severity of diabetes governs vascular lipoprotein lipase by affecting enzyme dimerization and disassembly. Diabetes. 2011;60:2041-2050 122. Wang F, Wang Y, Kim MS, Puthanveetil P, Ghosh S, Luciani DS, Johnson JD, Abrahani A, Rodrigues B. Glucose-induced endothelial heparanase secretion requires cortical and stress actin reorganization. Cardiovascular research. 2010;87:127-136 123. Yuan L, Hu J, Luo Y, Liu Q, Li T, Parish CR, Freeman C, Zhu X, Ma W, Hu X, Yu H, Tang S. Upregulation of heparanase in high-glucose-treated endothelial cells promotes endothelial cell migration and proliferation and correlates with akt and extracellular-signal-regulated kinase phosphorylation. Molecular vision. 2012;18:1684-1695 124. Han J, Woytowich AE, Mandal AK, Hiebert LM. Heparanase upregulation in high glucose-treated endothelial cells is prevented by insulin and heparin. Experimental biology and medicine. 2007;232:927-934 125. Wang Y, Zhang D, Chiu AP, Wan A, Neumaier K, Vlodavsky I, Rodrigues B. Endothelial heparanase regulates heart metabolism by stimulating lipoprotein lipase secretion from cardiomyocytes. Arteriosclerosis, thrombosis, and vascular biology. 2013;33:894-902 126. Kobayashi M, Naomoto Y, Nobuhisa T, Okawa T, Takaoka M, Shirakawa Y, Yamatsuji T, Matsuoka J, Mizushima T, Matsuura H, Nakajima M, Nakagawa H, Rustgi A, Tanaka N. Heparanase regulates esophageal keratinocyte differentiation through nuclear translocation and heparan sulfate cleavage. Differentiation; research in biological diversity. 2006;74:235-243 127. Purushothaman A, Hurst DR, Pisano C, Mizumoto S, Sugahara K, Sanderson RD. Heparanase-mediated loss of nuclear syndecan-1 enhances histone acetyltransferase (hat) activity to promote expression of genes that drive an aggressive tumor phenotype. The Journal of biological chemistry. 2011;286:30377-30383 128. Wang Y, Chiu AP, Neumaier K, Wang F, Zhang D, Hussein B, Lal N, Wan A, Liu G, Vlodavsky I, Rodrigues B. Endothelial cell heparanase taken up by cardiomyocytes regulates lipoprotein lipase transfer to the coronary lumen after diabetes. Diabetes. 2014;63:2643-2655 129. Rodrigues B, Cam MC, McNeill JH. Metabolic disturbances in diabetic cardiomyopathy. Molecular and cellular biochemistry. 1998;180:53-57 130. Dyck JR, Lopaschuk GD. Ampk alterations in cardiac physiology and pathology: Enemy or ally? The Journal of physiology. 2006;574:95-112 90 131. Szkudelski T. The mechanism of alloxan and streptozotocin action in b cells of the rat pancreas. Physiological research. 2001;50:537-546 132. Sambandam N, Abrahani MA, St Pierre E, Al-Atar O, Cam MC, Rodrigues B. Localization of lipoprotein lipase in the diabetic heart: Regulation by acute changes in insulin. Arteriosclerosis, thrombosis, and vascular biology. 1999;19:1526-1534 133. Sambandam N, Abrahani MA, Craig S, Al-Atar O, Jeon E, Rodrigues B. Metabolism of vldl is increased in streptozotocin-induced diabetic rat hearts. American journal of physiology. Heart and circulatory physiology. 2000;278:H1874-1882 134. Enerback S, Gimble JM. Lipoprotein lipase gene expression: Physiological regulators at the transcriptional and post-transcriptional level. Biochimica et biophysica acta. 1993;1169:107-125 135. Pillarisetti S, Paka L, Sasaki A, Vanni-Reyes T, Yin B, Parthasarathy N, Wagner WD, Goldberg IJ. Endothelial cell heparanase modulation of lipoprotein lipase activity. Evidence that heparan sulfate oligosaccharide is an extracellular chaperone. The Journal of biological chemistry. 1997;272:15753-15759 136. Adeyo O, Goulbourne CN, Bensadoun A, Beigneux AP, Fong LG, Young SG. Glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 and the intravascular processing of triglyceride-rich lipoproteins. Journal of internal medicine. 2012;272:528-540 137. Salabei JK, Hill BG. Mitochondrial fission induced by platelet-derived growth factor regulates vascular smooth muscle cell bioenergetics and cell proliferation. Redox biology. 2013;1:542-551 138. Pei-Ling Chiu A, Wang F, Lal N, Wang Y, Zhang D, Hussein B, Wan A, Vlodavsky I, Rodrigues B. Endothelial cells respond to hyperglycemia by increasing the lpl transporter gpihbp1. American journal of physiology. Endocrinology and metabolism. 2014;306:E1274-1283 139. Chen J, Zheng D, Shen J, Ruan J, Li A, Li W, Xie G, Luo X, Zhao P, Zheng H. Heparanase is involved in the proliferation and invasion of nasopharyngeal carcinoma cells. Oncology reports. 2013;29:1888-1894 140. Bame KJ. Heparanases: Endoglycosidases that degrade heparan sulfate proteoglycans. Glycobiology. 2001;11:91R-98R 141. Harjes U, Bridges E, McIntyre A, Fielding BA, Harris AL. Fatty acid-binding protein 4, a point of convergence for angiogenic and metabolic signaling pathways in endothelial cells. The Journal of biological chemistry. 2014;289:23168-23176 142. Kopan R, Ilagan MX. The canonical notch signaling pathway: Unfolding the activation mechanism. Cell. 2009;137:216-233 143. Bi P, Kuang S. Notch signaling as a novel regulator of metabolism. Trends in endocrinology and metabolism: TEM. 2015 144. Carmeliet P, Ng YS, Nuyens D, Theilmeier G, Brusselmans K, Cornelissen I, Ehler E, Kakkar VV, Stalmans I, Mattot V, Perriard JC, Dewerchin M, Flameng W, Nagy A, Lupu F, Moons L, Collen D, D'Amore PA, Shima DT. Impaired myocardial angiogenesis and ischemic cardiomyopathy in mice lacking the vascular endothelial growth factor isoforms vegf164 and vegf188. Nature medicine. 1999;5:495-502 145. Yoon YS, Uchida S, Masuo O, Cejna M, Park JS, Gwon HC, Kirchmair R, Bahlman F, Walter D, Curry C, Hanley A, Isner JM, Losordo DW. Progressive attenuation of myocardial vascular endothelial growth factor expression is a seminal event in diabetic cardiomyopathy: Restoration of microvascular homeostasis and recovery of cardiac function in diabetic cardiomyopathy after replenishment of local vascular endothelial growth factor. Circulation. 2005;111:2073-2085 91 146. Patel S, Leal AD, Gorski DH. The homeobox gene gax inhibits angiogenesis through inhibition of nuclear factor-kappab-dependent endothelial cell gene expression. Cancer research. 2005;65:1414-1424 147. Wang F, Jia J, Lal N, Zhang D, Chiu AP, Wan A, Vlodavsky I, Hussein B, Rodrigues B. High glucose facilitated endothelial heparanase transfer to the cardiomyocyte modifies its cell death signature. Cardiovascular research. 2016;112:656-668 148. Annes JP, Munger JS, Rifkin DB. Making sense of latent tgfbeta activation. Journal of cell science. 2003;116:217-224 149. Sato Y, Rifkin DB. Inhibition of endothelial cell movement by pericytes and smooth muscle cells: Activation of a latent transforming growth factor-beta 1-like molecule by plasmin during co-culture. The Journal of cell biology. 1989;109:309-315 150. Yu Q, Stamenkovic I. Cell surface-localized matrix metalloproteinase-9 proteolytically activates tgf-beta and promotes tumor invasion and angiogenesis. Genes & development. 2000;14:163-176 151. Jiang L, Xiao L, Sugiura H, Huang X, Ali A, Kuro-o M, Deberardinis RJ, Boothman DA. Metabolic reprogramming during tgfbeta1-induced epithelial-to-mesenchymal transition. Oncogene. 2015;34:3908-3916 152. Guido C, Whitaker-Menezes D, Capparelli C, Balliet R, Lin Z, Pestell RG, Howell A, Aquila S, Ando S, Martinez-Outschoorn U, Sotgia F, Lisanti MP. Metabolic reprogramming of cancer-associated fibroblasts by tgf-beta drives tumor growth: Connecting tgf-beta signaling with "warburg-like" cancer metabolism and l-lactate production. Cell cycle. 2012;11:3019-3035 153. Pulinilkunnil T, An D, Ghosh S, Qi D, Kewalramani G, Yuen G, Virk N, Abrahani A, Rodrigues B. Lysophosphatidic acid-mediated augmentation of cardiomyocyte lipoprotein lipase involves actin cytoskeleton reorganization. American journal of physiology. Heart and circulatory physiology. 2005;288:H2802-2810 154. Moolenaar WH. Lysophosphatidic acid, a multifunctional phospholipid messenger. The Journal of biological chemistry. 1995;270:12949-12952 155. Valcourt U, Thuault S, Pardali K, Heldin CH, Moustakas A. Functional role of meox2 during the epithelial cytostatic response to tgf-beta. Molecular oncology. 2007;1:55-71 156. Rodrigues B, Cam MC, McNeill JH. Myocardial substrate metabolism: Implications for diabetic cardiomyopathy. Journal of molecular and cellular cardiology. 1995;27:169-179 157. Augustus AS, Kako Y, Yagyu H, Goldberg IJ. Routes of fa delivery to cardiac muscle: Modulation of lipoprotein lipolysis alters uptake of tg-derived fa. American journal of physiology. Endocrinology and metabolism. 2003;284:E331-339 158. Williams KJ. Molecular processes that handle -- and mishandle -- dietary lipids. The Journal of clinical investigation. 2008;118:3247-3259 159. Farhangkhoee H, Khan ZA, Barbin Y, Chakrabarti S. Glucose-induced up-regulation of cd36 mediates oxidative stress and microvascular endothelial cell dysfunction. Diabetologia. 2005;48:1401-1410 160. Elmasri H, Karaaslan C, Teper Y, Ghelfi E, Weng M, Ince TA, Kozakewich H, Bischoff J, Cataltepe S. Fatty acid binding protein 4 is a target of vegf and a regulator of cell proliferation in endothelial cells. FASEB journal : official publication of the Federation of American Societies for Experimental Biology. 2009;23:3865-3873 161. Ogishima T, Shiina H, Breault JE, Tabatabai L, Bassett WW, Enokida H, Li LC, Kawakami T, Urakami S, Ribeiro-Filho LA, Terashima M, Fujime M, Igawa M, Dahiya R. Increased heparanase expression is caused by promoter hypomethylation and up-regulation of transcriptional factor early growth response-1 in human prostate cancer. Clinical cancer 92 research : an official journal of the American Association for Cancer Research. 2005;11:1028-1036 162. Ogishima T, Shiina H, Breault JE, Terashima M, Honda S, Enokida H, Urakami S, Tokizane T, Kawakami T, Ribeiro-Filho LA, Fujime M, Kane CJ, Carroll PR, Igawa M, Dahiya R. Promoter cpg hypomethylation and transcription factor egr1 hyperactivate heparanase expression in bladder cancer. Oncogene. 2005;24:6765-6772 163. Banerjee I, Fuseler JW, Price RL, Borg TK, Baudino TA. Determination of cell types and numbers during cardiac development in the neonatal and adult rat and mouse. American journal of physiology. Heart and circulatory physiology. 2007;293:H1883-1891 164. Creager MA, Luscher TF, Cosentino F, Beckman JA. Diabetes and vascular disease: Pathophysiology, clinical consequences, and medical therapy: Part i. Circulation. 2003;108:1527-1532 165. Schubert SY, Ilan N, Shushy M, Ben-Izhak O, Vlodavsky I, Goldshmidt O. Human heparanase nuclear localization and enzymatic activity. Lab Invest. 2004;84:535-544 166. Selvaraju V, Joshi M, Suresh S, Sanchez JA, Maulik N, Maulik G. Diabetes, oxidative stress, molecular mechanism, and cardiovascular disease--an overview. Toxicology mechanisms and methods. 2012;22:330-335 167. Chou E, Suzuma I, Way KJ, Opland D, Clermont AC, Naruse K, Suzuma K, Bowling NL, Vlahos CJ, Aiello LP, King GL. Decreased cardiac expression of vascular endothelial growth factor and its receptors in insulin-resistant and diabetic states: A possible explanation for impaired collateral formation in cardiac tissue. Circulation. 2002;105:373-379 168. Dobaczewski M, Chen W, Frangogiannis NG. Transforming growth factor (tgf)-beta signaling in cardiac remodeling. Journal of molecular and cellular cardiology. 2011;51:600-606 169. Bujak M, Frangogiannis NG. The role of tgf-beta signaling in myocardial infarction and cardiac remodeling. Cardiovascular research. 2007;74:184-195 170. Michael DR, Salter RC, Ramji DP. Tgf-beta inhibits the uptake of modified low density lipoprotein by human macrophages through a smad-dependent pathway: A dominant role for smad-2. Biochimica et biophysica acta. 2012;1822:1608-1616 171. An D, Rodrigues B. Role of changes in cardiac metabolism in development of diabetic cardiomyopathy. American journal of physiology. Heart and circulatory physiology. 2006;291:H1489-1506 172. Puthanveetil P, Wan A, Rodrigues B. Foxo1 is crucial for sustaining cardiomyocyte metabolism and cell survival. Cardiovascular research. 2013;97:393-403 173. Qi D, Kuo KH, Abrahani A, An D, Qi Y, Heung J, Kewalramani G, Pulinilkunnil T, Ghosh S, Innis SM, Rodrigues B. Acute intralipid infusion reduces cardiac luminal lipoprotein lipase but recruits additional enzyme from cardiomyocytes. Cardiovascular research. 2006;72:124-133
UBC Theses and Dissertations
Featured Collection
UBC Theses and Dissertations
Endothelial cells manage fatty acid delivery to the cardiomyocytes following diabetes Chiu, Amy Pei-Ling 2017
Item Metadata
- Media
- 24-ubc_2017_november_chiu_amy.pdf [ 81.76MB ]
- Metadata
- JSON: 24-1.0355882.json
- JSON-LD: 24-1.0355882-ld.json
- RDF/XML (Pretty): 24-1.0355882-rdf.xml
- RDF/JSON: 24-1.0355882-rdf.json
- Turtle: 24-1.0355882-turtle.txt
- N-Triples: 24-1.0355882-rdf-ntriples.txt
- Original Record: 24-1.0355882-source.json
- Full Text
- 24-1.0355882-fulltext.txt
- Citation
- 24-1.0355882.ris
Full Text
Citation Scheme:
Usage Statistics
Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
<div id="ubcOpenCollectionsWidgetDisplay">
<script id="ubcOpenCollectionsWidget"
async >
Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url: