UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Oxidized low density lipoprotein-mediated macrophage survival : a role for calcium and regulation of… Chen, Johnny H. 2009

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

Item Metadata

Download

Media
24-ubc_2009_fall_chen_johnny.pdf [ 4.69MB ]
Metadata
JSON: 24-1.0067286.json
JSON-LD: 24-1.0067286-ld.json
RDF/XML (Pretty): 24-1.0067286-rdf.xml
RDF/JSON: 24-1.0067286-rdf.json
Turtle: 24-1.0067286-turtle.txt
N-Triples: 24-1.0067286-rdf-ntriples.txt
Original Record: 24-1.0067286-source.json
Full Text
24-1.0067286-fulltext.txt
Citation
24-1.0067286.ris

Full Text

OXIDIZED LOW DENSITY LIPOPROTEIN-MEDIATED MACROPHAGE SURVIVAL:  A ROLE FOR CALCIUM AND REGULATION OF CELLULAR ENERGY  by  JOHNNY H. CHEN  B.Sc. (Hons), Simon Fraser University, 2001    A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY   in   THE FACULTY OF GRADUATE STUDIES  (Experimental Medicine)             THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)   June 2009  © Johnny H. Chen, 2009 ii  Abstract Oxidized low density lipoprotein (oxLDL) as well as specific macrophage populations, play a pivotal role in the initiation and progression of atherosclerotic lesions. Our group previously reported that oxLDL inhibits apoptosis in bone marrow-derived macrophages (BMDM) deprived of macrophage colony-stimulating factor (M-CSF). In this study, I showed that oxLDL induces an oscillatory increase in intracellular calcium ([Ca2+]i) that is mediated by the activation of sphingosine kinase.  This leads to activation of the Ca2+/calmodulin dependent kinase, eukaryotic elongation factor-2 kinase (eEF2 kinase).   Both the increase in [Ca2+]i and activation of eEF2 kinase can be blocked by BAPTA-AM, an intracellular Ca2+ chelator.  Addition of oxLDL also results in the phosphorylation of eEF2, the only known substrate of eEF2 kinase.  The eEF2 kinase selective inhibitors, TS-4 and TX-1918, blocked oxLDL-mediated phosphorylation of eEF2 as well as oxLDL’s ability to promote survival of BMDMs.  eEF2 kinase can be negatively regulated by p38 MAPK.  Withdrawal of M-CSF results in the activation of p38 MAPK, an effect that is blocked with the addition of oxLDL.  This indicates that oxLDL can positively regulate eEF2 kinase activity by both (1) generating an oscillatory increase in [Ca2+]i and (2) inhibiting its negative regulation by p38 MAPK.  eEF2 is a monomeric GTPase that regulates peptide chain elongation and its phosphorylation inhibits its activity.  Addition of oxLDL results in a decrease in overall protein synthesis and induction of autophagy in BMDMs.  This suggests that under conditions of metabolic stress (e.g., growth factor withdrawal), there may be an eEF2 kinase-dependent protective mechanism that can be activated by the presence of oxLDL, thus linking energy iii  conservation (e.g., via inhibition of protein synthesis) and the replenishment of energy supplies by digestion of cellular organelles through the induction of autophagy. iv  Table of Contents Abstract .......................................................................................................... ii  Table of Contents ......................................................................................... iv  List of Tables ................................................................................................ xi  List of Figures .............................................................................................. xii  List of Abbreviations .................................................................................. xv  Acknowledgements ................................................................................... xxii  1  Introduction ............................................................................................... 1  1.1  Overview .................................................................................................................. 1  1.2  Atherosclerosis ......................................................................................................... 2  1.2.1  Epidemiology .................................................................................................... 2  1.2.2  Cholesterol Influx/Efflux .................................................................................. 4  1.2.3  Lesion Development ......................................................................................... 8  1.2.4  Oxidative Modification and LDL ..................................................................... 9  1.2.5  Putative Oxidants ............................................................................................ 10  1.2.6  Modified LDL and Leukocyte Recruitment ................................................... 12  1.2.7  OxLDL and Foam Cell Formation .................................................................. 13  1.2.8  Macrophage Activation ................................................................................... 15  v  1.2.9  Macrophages and Plaque Destabilization ....................................................... 16  1.3  OxLDL and Macrophage Proliferation, Survival, and Apoptosis ......................... 19  1.3.1  Atherosclerosis and Macrophage Population .................................................. 19  1.3.2  OxLDL and Macrophage Proliferation ........................................................... 19  1.3.3  OxLDL and Macrophage Survival .................................................................. 20  1.4  Sphingolipids ......................................................................................................... 26  1.4.1  Sphingolipid Generation ................................................................................. 26  1.4.2  SK and S1P ..................................................................................................... 27  1.4.3  SK, S1P, and Atherosclerosis ......................................................................... 28  1.5  Calcium .................................................................................................................. 31  1.5.1  Ca2+ Signaling Toolkit .................................................................................... 31  1.5.2  Spatial and Temporal Organization of Ca2+ Signaling ................................... 31  1.5.3  Calmodulin ...................................................................................................... 32  1.5.4  Ca2+ and Cell Proliferation .............................................................................. 33  1.5.5  Ca2+ and Cell Survival and Apoptosis ............................................................ 33  1.5.6  Ca2+ and Atherosclerosis ................................................................................. 34  1.6  Elongation Factor-2 Kinase ................................................................................... 38  1.6.1  Background ..................................................................................................... 38  1.6.2  eEF2 Kinase Regulation ................................................................................. 38  vi  1.6.3  eEF2 Kinase Function ..................................................................................... 39  1.7  Protein Translation ................................................................................................. 43  1.7.1  Background ..................................................................................................... 43  1.7.2  Translation Initiation ....................................................................................... 43  1.7.3  Translation Elongation .................................................................................... 44  1.7.4  Translation Termination.................................................................................. 45  1.7.5  Why Regulate Peptide Chain Elongation? ...................................................... 45  1.8  Autophagy .............................................................................................................. 49  1.8.1  Background ..................................................................................................... 49  1.8.2  Regulation of Autophagy ................................................................................ 49  1.8.3  Autophagy in Cell Survival and Apoptosis .................................................... 50  1.9  Objectives .............................................................................................................. 53  2  Materials & Methods .............................................................................. 55  2.1  Materials ................................................................................................................ 55  2.2  Lipoprotein Isolation and Oxidation ...................................................................... 56  2.3  Characterization of Modified Lipoproteins ........................................................... 57  2.4  Animals .................................................................................................................. 58  2.5 Cell Culture ............................................................................................................. 59  2.6  Cell Viability Assay ............................................................................................... 60  vii  2.7  Apoptosis Assay..................................................................................................... 61  2.8  Calcium Imaging .................................................................................................... 61  2.9  Sphingosine Kinase Activity Assay ....................................................................... 62  2.10  eEF2 Kinase Activity Assay ................................................................................ 63  2.11  Immunoblotting.................................................................................................... 64  2.12  Protein Synthesis Assay ....................................................................................... 65  2.13  Transmission Electron Microscopy ..................................................................... 66  2.14  Microtubule Associated Protein Light Chain 3 Localization .............................. 66  3  oxLDL Generates an Oscillatory Increase In [Ca2+]i .......................... 68  3.1  Introduction ............................................................................................................ 68  3.2  Results .................................................................................................................... 69  3.2.1  OxLDL Generates an Oscillatory Increase in [Ca2+]i ..................................... 69  3.2.2  LysoPC in oxLDL is Not Responsible for the Generation of [Ca2+]i Oscillations ............................................................................................................... 69  3.2.3  Extracellular Ca2+ Plays a Partial Role in the Generation of [Ca2+]i Oscillations ............................................................................................................... 70  3.2.4  Thapsigargin Blocks oxLDL Generated [Ca2+]i Oscillations ......................... 70  3.2.5  Inhibition of Phospholipase C or RyR Does Not Block oxLDL Mediated Macrophage Survival ................................................................................................ 71  viii  3.2.6  S1P Generates [Ca2+]i Oscillations and Promotes Macrophage Survival ....... 71  3.2.7  SK is Activated in Response to oxLDL .......................................................... 72  3.2.8  Inhibition of SK Blocks oxLDL Mediated [Ca2+]i Oscillation and Macrophage Survival ..................................................................................................................... 72  3.2.9  TLR-2, TLR-4, and LOX-1 are Not Required for oxLDL Mediated Macrophage Survival ................................................................................................ 73  3.3  Discussion .............................................................................................................. 88  4  PKC Is Not Involved In oxLDL-Mediated BMDM Survival ............. 90  4.1  Introduction ............................................................................................................ 90  4.2  Results .................................................................................................................... 92  4.2.1  Inhibition of Ca2+-Sensitive PKC Isoforms do not Alter BMDM Viability ... 92  4.2.2  Stimulation With PMA do not Alter BMDM Viability .................................. 92  4.2.3  Rottlerin Selectively Inhibits the Pro-Survival Effect of oxLDL ................... 92  4.2.4  PKCδ is not Involved in Rotterlin’s Inhibition of oxLDL Mediated Macrophage Survival ................................................................................................ 93  4.3  Discussion .............................................................................................................. 99  5  Elongation Factor-2 Kinase is Required for oxLDL-Mediated Macrophage Survival ............................................................................... 100  5.1  Introduction .......................................................................................................... 100  ix  5.2  Results .................................................................................................................. 101  5.2.1  eEF2 Kinase Activity is Increased in Response to oxLDL ........................... 101  5.2.2  eEF2 Kinase Activition is Required for oxLDL’s Pro-Survival Effect ........ 101  5.2.3  eEF2 Kinase Activation Requires Ca2+ Mobilization ................................... 102  5.2.4  OxLDL-Mediated Macrophage Survival do no Involve the mTor, ERK, or PKA Pathways ........................................................................................................ 103  5.2.5  eEF2 Kinase Activity Requires Hsp90 ......................................................... 104  5.2.6  p38 MAPK Negatively Regulates eEF2 Kinase Activity ............................. 104  5.2.7  p38 MAPK is Activated upon Growth Factor Withdrawal .......................... 105  5.2.8  Ceramide Activates p38 MAPK and Negatively Regulates eEF2 Kinase Activity ................................................................................................................... 106  5.2.9  Myeloperoxidase Oxidized LDL can Promote Macrophage Survival, an Effect that can be Blocked by Inhibiting eEF2 Kinase ...................................................... 106  5.2.10  BMDM from Transgenic Mice Expressing Catalytically Inactive eEF2 Kinase Shows an Attenuated Survival Response to oxLDL ................................... 107  5.3  Discussion ............................................................................................................ 129  6  OxLDL Induces Macrophages to Undergo Autophagy .................... 131  6.1  Introduction .......................................................................................................... 131  6.2  Results .................................................................................................................. 132  x  6.2.1  Protein Synthesis is Reduced in Response to oxLDL ................................... 132  6.2.2  Autophagic Vacuoles are Present in Macrophages Treated with oxLDL ..... 132  6.2.3  OxLDL Induces Microtubule-Associated Protein Light Chain 3 Aggregation ................................................................................................................................. 134  6.2.4  AMPK is Activated in Response to oxLDL ................................................. 134  6.2.5  AMPK Activation Partially Mediates oxLDL Pro-Survival Effects ............ 135  6.3  Discussion ............................................................................................................ 143  7  Summary ................................................................................................ 145  Bibliography .............................................................................................. 151   xi  List of Tables Table 1.1  Major Classes of Lipoproteins ……………………………………………….. 7  xii  List of Figures Figure 1.1  Recruitment and Differentiation of Macrophages in Atheroma ..................... 18  Figure 1.2  OxLDL Induced Macrophage Survival .......................................................... 24  Figure 1.3  Biosynthetic Pathway of Sphingolipids .......................................................... 30  Figure 1.4  Elements of the Ca2+ Signaling Toolkit .......................................................... 35  Figure 1.5  Ca2+ Signals Regulate Both Cell Survival and Apoptosis .............................. 37  Figure 1.6  Regulatory Phosphorylation Sites on eEF2 Kinase ........................................ 42  Figure 1.7  Three Phases of Protein Translation from mRNA ......................................... 47  Figure 1.8  The Cellular Aspects of Autophagy ............................................................... 52  Figure 3.1  OxLDL Generates an Oscillatory Increase in [Ca2+]i ..................................... 74  Figure 3.2  LysoPC in oxLDL is not Responsible for the Generation of [Ca2+]i Oscillations ....................................................................................................................... 76  Figure 3.3  Extracellular Ca2+ Plays a Partial Role in the Generation of [Ca2+]i Oscillations ....................................................................................................................... 77  Figure 3.4  Thapsigargin Blocks oxLDL Generated [Ca2+]i Oscillations ......................... 78  Figure 3.5  Inhibition of Phospholipase C or RyR do not Block oxLDL Mediated Macrophage Survival ........................................................................................................ 79  Figure 3.6  S1P Generates [Ca2+]i Oscillations ................................................................. 80  Figure 3.7  S1P Promotes Macrophage Survival .............................................................. 81  Figure 3.8  SK is Activated in Response to oxLDL ......................................................... 83  Figure 3.9  Inhibition of SK Blocks oxLDL Generated [Ca2+]i Oscillations .................... 84  Figure 3.10  Inhibition of SK Blocks oxLDL Mediated Macrophage Survival ............... 85  xiii  Figure 3.11  TLR-2, TLR-4, and LOX-1 are Not Required for oxLDL Mediated Macrophage Survival ........................................................................................................ 87  Figure 4.1  Inhibition of Ca2+-Sensitive PKC Isoforms do not Alter BMDM Viability .. 95  Figure 4.2  Stimulation With PMA do not Alter BMDM Viability .................................. 96  Figure 4.3  Rottlerin Selectively Inhibits oxLDL’s Pro-Survival Effect .......................... 97  Figure 4.4  PKCδ is not Involved in Rotterlin’s Inhibition of oxLDL Mediated Macrophage Survival ........................................................................................................ 98  Figure 5.1  eEF2 Kinase Activity is Increased in Response to oxLDL .......................... 109  Figure 5.2  eEF2 Kinase Inhibitors Selectively Block oxLDL’s Pro-Survival Effect .... 110  Figure 5.3  eEF2 Kinase Inhibitors Induce Apoptosis .................................................... 111  Figure 5.4  eEF2 Kinase Regulation ............................................................................... 112  Figure 5.5  BAPTA-AM Blocks oxLDL Generated [Ca2+]i Oscillations ....................... 113  Figure 5.6  BAPTA-AM and Rottlerin Act Synergistically to Block oxLDL Mediated Macrophage Survival ...................................................................................................... 114  Figure 5.7  OxLDL Mediated Macrophage Survival do no Involve the mTor, ERK, or PKA Pathways ................................................................................................................ 115  Figure 5.8  Geldanamycin Inhibits oxLDL Mediated Macrophage Survival and Induces Apoptosis ........................................................................................................................ 117  Figure 5.9  OxLDL Induces Hsp90 to Complex with CaM ............................................ 119  Figure 5.10  p38 MAPK Blocks oxLDL’s Pro-Survival Effect...................................... 120  Figure 5.11  p38 MAPK is Activated upon Growth Factor Withdrawal ........................ 122  Figure 5.12  Ceramide Activates p38 MAPK ................................................................. 123  xiv  Figure 5.13  Characterization of MPO-LDL ................................................................... 124  Figure 5.14  Only Extensively Modified MPO-LDL can Promote Macrophage Survival ......................................................................................................................................... 125  Figure 5.15  TX-1918 Blocks MPO-LDL Mediated Macrophage Survival ................... 126  Figure 5.16  BMDM from Transgenic Mice Expressing Catalytically Inactive eEF2 Kinase Shows an Attenuated Survival Response to oxLDL ........................................... 127  Figure 6.1  Protein Synthesis is Reduced in Response to oxLDL .................................. 136  Figure 6.2  Autophagic Vacuoles are Present in Macrophages Treated with oxLDL .... 137  Figure 6.3  OxLDL Induces LC3 Localization ............................................................... 139  Figure 6.4  PBMC-Derived Macrophage Viability Increases in Response to oxLDL ... 140  Figure 6.5  AMPK is Activated in Response to oxLDL ................................................. 141  Figure 6.6  AMPK Activation Partially Mediates oxLDL’s Pro-Survival Effect ........... 142  Figure 7.1  OxLDL Mediated Pro-Survival Signalling in Macrophages, a Working Model ......................................................................................................................................... 149   xv  List of Abbreviations -/-  knockout [Ca2+]i  intracellular calcium aa  amino acyl ACAT  acyl coenzyme A:cholesterol acyltransferase ADP  adenosine diphosphate AIM  apoptosis inhibitor expressed by macrophages AM  tetraacetoxy-methyl ester AMP  adenosine monophosphate AMPK  adenosine monophosphate protein kinase aPKC  atypical protein kinase C apoB  apolipoprotein B apoE  apolipoprotein E ASMase acid sphingomyelinase ATP  adenosine triphosphate BAPTA-AM 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid tetraacetoxy-   methyl ester Bcl  B-cell lymphoma BMDM bone marrow derived macrophage BSA  bovine serum albumin Ca2+  calcium cADPR cyclic adenosine diphosphate ribose receptor xvi  CAD  coronary artery disease CaM  calmodulin CaMK  calcium/calmodulin dependent kinase CCB  calcium channel blocker CD  cluster of differentiation CETP  cholesteryl ester transfer protein CM  chylomicron CPK  conventional protein kinase cPKC  conventional protein kinase C cytoC  cytochrome C DAG  diacylglycerol DAPK  death associated protein kinase DMEM Dulbecco’s modified Eagle’s medium DMSO  dimethyl sulfoxide DNA  deoxyribonucleic acid DPBS  Dulbecco’s phosphate buffered saline E-64  trans-epoxysuccinyl-L-leucylamido(4-guanidino) butane EDTA  ethylenediaminetetraacetic acid eEF  eukaryotic elongation factor EGF  epidermal growth factor eIF  eukaryotic initiation factor ER  endoplasmic reticulum xvii  eRF  eukaryotic release factor ERK   extracellular signal regulated kinase FBS  fetal bovine serum FCS  fetal calf serum GDP  guanine diphosphate GEF  guanine nucleotide exchange factor GFP  green fluorescent protein GM-CSF granulocyte-macrophage colony stimulating factor GPCR  G protein-coupled receptor GTP  guanine triphosphate HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid Hsp90  heat shock protein 90 IGF-I  insulin like growth factor I IL  interleukin IBMX  isobutylmethylxanthine IP3  inositol-1,4,5-trisphosphate IP3R  inositol-1,4,5-trisphosphate receptor IRE-1  insulin response element-1 JNK  c-Jun-N-terminal kinase LC3  microtubule-associated protein light chain 3 LDL  low density lipoprotein LOX-1  lectin-like oxidized low density lipoprotein receptor-1 xviii  LPO  lipid hydroperoxide LPS  lipopolysaccharide lysoPC  lysophosphatidylcholine MAPK  mitogen activated protein kinase MAPKAP mitogen activated protein kinase activated protein MARCO macrophage receptor with collagenous structure MCP-1 monocyte chemotactic protein-1 M-CSF macrophage colony stimulating factor MLCK  myosin light chain kinase mmLDL minimally modified lipoprotein MMP  matrix metalloproteinase MPO  myeloperoxidase mRNA  messenger ribonucleic acid mTOR  mammalian target of rapamycin MTS  3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4- sulfophenyl)-2H-tetrazolium, inner salt MyD88 myeloid differentiation factor 88 NAADP nicotinic acid dinucleotide phosphate NAD  nicotinamide adenine diculeotide NFAT  nuclear factor of activated T cells NGF  nerve growth factor nLDL  native low density lipoprotein xix  NMDA N-methyl-d-aspartate nPKC  novel protein kinase C NOS  nitric oxide synthase oxLDL oxidized low density lipoprotein PBMC  peripheral blood mononuclear cell PC  phosphatidylcholine PDE  phosphodiesterase PDGF  platelet derived growth factor PI-3K  phosphatidylinositol 3-kinase PIP2  phosphatidylinositol-4,5-bisphosphate PKA  protein kinase A PKB  protein kinase B PKC  protein kinase C PLC  phospholipase PLTP  phospholipid transfer protein PMA  phorbol 12-myristate 13-acetate PMCA  plasma membrane calcium ATPase PMS  phenazine methosulfate PPR  pattern recognition receptor PS  phosphotidylserine PTP  permeability transition pore PVDF  polyvinylidene difluoride xx  PYK2  proline rich kinase 2 Rb  retinoblastoma RCT  reverse cholesterol transport RNA  ribonucleic acid RSK  p90 ribosomal S6 kinase RTK  receptor tyrosine kinase RyR  ryanodine receptor S1P  sphingosine-1-phosphate S6K1  p70 S6 kinase 1 SCaMPER sphingolipid calcium release mediating protein of the endoplasmic   reticulum ScR  scavenger receptor SDS  sodium dodecyl sulfate SERCA sarco-endoplasmic reticulum calcium ATPase SMC  smooth muscle cell SK  sphingosine kinase TBS  tris(hydroxymethyl)aminomethane buffered saline TCA  trichloroacetic acid TCR  T cell receptor Th1  T-helper 1 TLC  thin layer chromatography TLR  Toll-like receptor xxi  TnC  troponin TNFα  tumor necrosis factor α VCAM-1 vascular cell adhesion molecule-1 VEGF  vascular endothelial growth factor xxii  Acknowledgements  The successful endeavor of what is often a long and arduous journey towards the completion of the Ph.D. program relies not only on the perseverance of the candidate, but also in the support given by those around him.  First and foremost, I would like to extend my deepest gratitude to my supervisors, Dr. Vincent Duronio and Dr. Urs Steinbrecher for providing me the opportunity to develop both scientifically and personally whilst striking a balance between academic guidance and individual freedom.  Their support has created an environment that has fostered critical thinking and deductive reasoning.  The completion of my Ph.D. also could not have been possible without the support of my fellow colleagues and friends who have started the program with me, Shih Wei Wang, Ivan Waissbluth and Joseph Anthony.  Special thanks is owed to Payman Hojabrpour, whose logistical and technical support have been invaluable.  Last but not least, I would especially like to thank Dr. Maziar Riazy whose insight and intellectual discourse has guided my research and whose friendship I would like to count on for many years to come. 1  1  Introduction 1.1  Overview  This thesis contains novel observations that encompasses a broad range of research areas.  My main objective was to determine the signal transduction pathways regulating oxidized low-density lipoprotein (oxLDL)-mediated macrophage survival. The Introduction begins with a general review of atherosclerosis and provides a framework for the physiological relevance of oxLDL and macrophage survival.  The next section proceeds to discuss past studies that have investigated oxLDL and its role in macrophage survival and apoptosis.  Because this thesis describes a pathway (Figure 7.1) that involves sphingolipid dependent calcium mobilization, followed by the activation of eukaryotic elongation factor-2 kinase (eEF2 kinase) and subsequent reduction in protein synthesis and induction of autophagy, the remaining sections of the Introduction describes, in the following order, sphingolipids, calcium, eEF2 kinase, protein synthesis, and autophagy with an emphasis in their respective roles in survival/apoptosis and/or atherosclerosis. 2  1.2  Atherosclerosis 1.2.1  Epidemiology Atherosclerosis is the underlying cause of most cardiovascular diseases including coronary artery disease (CAD), ischemic gangrene, abdominal aortic aneurysms, and many cases of heart failure and stroke.  Atherosclerosis results in cardiovascular death of approximately 16.7 million individuals around the world each year and constitutes the leading cause of mortality in the Western world today.  The World Health Organization expects cardiovascular disease to be the main cause of death globally within the next 15 years owing to its rapidly increasing prevalence in developing countries and the rising incidence of obesity and diabetes in the Western world [1].  Over the past 40 years, a number of risk factors have been identified that have proven predictive of the incidence of cardiovascular disease.  These include non- modifiable risk factors such as age, gender, and family history of symptomatic vascular disease.  Based on clinical data in the United States, the average risk of developing cardiovascular disease for a 30-34 year old male is ~3% [2].  This number rises to ~21% for a comparable individual aged 60-64 years.  Compared to age-matched women, males exhibit excess risk for cardiovascular disease [3].  The reason for this discrepancy is not entirely clear, though it has been speculated that it may be due to relatively higher concentrations of high-density lipoprotein (HDL) cholesterol in women compared to age- matched men.  Modifiable risk factors include obesity, cigarette smoking, hypertension, diabetes mellitus, and serum cholesterol levels.  The relation between obesity, defined as an 3  excess body weight with an abnormal high preponderance of body fat, is of considerable concern as the prevalence of obesity in the developed world is increasing at an alarming rate.  Other risk factors, such as hypertension, low HDL cholesterol, and diabetes mellitus, often coexist with obesity [4].  Hypertension, defined as a systolic blood pressure in excess of 140 mmHg or a diastolic blood pressure above 90 mmHg, appears to have a linear relationship with increased incidence of atherosclerotic vascular disease when compared to blood pressure elevation [5].  Diabetes mellitus is another major contributing factor to atherosclerosis.  In patients with diabetes, the risk of coronary atherosclerosis is 3-5 fold greater than in nondiabetics despite controlling for other risk factors [6].  Perhaps the most important and well studied of the major risk factors for atherosclerosis is elevated serum cholesterol levels or hypercholesterolemia.  Familial hypercholesterolemia is an autosomal dominant disorder that affects ~1 in 500 individuals.  Heterozygotes for this disease manifest a 2-5 fold elevation in plasma low density lipoprotein (LDL) cholesterol that is due to a functional impairment of the LDL receptor, resulting in a defect in LDL clearance.  Homozygotes for this disorder demonstrate a 4-6 fold elevation in plasma cholesterol that produces precocious atherosclerosis.  In heterozygotes, 85% of individuals have experienced a myocardial infarction by the age of 60, and this age is reduced to 15 years in homozygous patients [7].  Futhermore, cholesterol lowering therapy with the use of statins greatly diminishes the clinical manifestations of atherosclerosis [8].  In contrast, serum levels of HDL, 4  which functions as an extracellular cholesterol acceptor, has an inverse relationship to coronary artery disease [9].  1.2.2  Cholesterol Influx/Efflux  Functionally, there are two main classes of lipoproteins [10].  The first consists of particles whose main role is to deliver lipids (primarily triacylglycerol) from the liver or intestine to peripheral, extrahepatic tissues.  These particles contain apolipoprotein B (apoB) together with a varying amounts of other lipids and apolipoproteins (Table 1.1). Very low-density lipoproteins (VLDLs) secreted from the liver, contain one molecule of the full-length form of apoB (apoB100).  Following the loss of most of their triacylglycerol to peripheral tissues, VLDL remants, also known as intermediate-density lipoproteins (IDL) become enriched with apolipoprotein E (apoE).  ApoE mediates high affinity binding of apoE-containing lipoproteins to the LDL receptor.  In this regard, apoE plays a role in lipid transport and distribution via the cellular uptake and degradation of lipoproteins.  Some of the IDL are returned to the liver, endocytosed and catabolized.  Others remain in circulation and  still contain significant amounts of triacylglycerol and most of their original content of cholesteryl ester and free cholesterol. After full lipolysis by lipoprotein lipase, most of the VLDL-derived apoB100 particles remain in circulation in the form of LDL, the major transporter of cholesterol in blood and the source of most of the cholesterol found in atherosclerotic lesions [11-14].  IDL free cholesterol content is the same as that in VLDL while cholesteryl ester is increased in LDL, as a result of the activity of cholesteryl ester transfer protein (CETP) which 5  exchanges triacylglycerol for HDL-associated cholesteryl esters.  LDL has a plasma half- life of about 2 days, and is cleared from plasma mainly by the liver.  In contrast to VLDL and LDL, HDL acts as a cholesterol acceptor in reverse cholesterol transport (RCT) [10].  HDLs contain 1-4 molecules of apoliprotein A1, together with other apolipoproteins that specify the metabolism and delivery of these lipids.  HDL accumulates lipids from the peripheral tissues, and return them to the liver. Newly formed HDLs have high density and little lipid.  Their density decreases as they accumulate lipids in circulation.  HDL can classically be subdivided based on their density or electrophoretic mobility (preβ-migrating HDL: nascent, lipid poor apoA1; HDL3: d = 1.12-1.219 g/ml; HDL2: d = 1.063-1.12 g/ml), which reflects their structural and functional diversity.  Several sources of cellular cholesterol contribute to RCT from peripheral tissues.  These include peripheral sterol synthesis and recycling of lipoprotein- derived cholesterol, most of which is from LDL endocytosed via LDL receptors.  Both cholesterol synthesis and LDL receptor activity are regulated by cellular cholesterol content, and so should be downregulated in peripheral tissues if the plasma LDL concentration is high.  Cholesterol from all these sources transfers to HDL for further metabolism, including esterification outside the cell.  HDL cholesterol can be delivered to the liver by uptake of intact HDL particles, or by the selective uptake of esterified cholesterol from HDL2 by the liver and steroidogenic tissues, facilitated by scavenger receptor BI (SR-BI).  HDL cholesteryl esters can also be transferred to other lipoproteins (VLDL, LDL) by being exchanged with triacylgycerol via CETP mentioned above.  Loss of esterified cholesterol and phospholipids through phospholipid transfer protein (PLTP) 6  and hepatic lipase activity, generates a lipid poor preβ-migrating HDL ready to participate in a new cycle of cholesterol efflux. 7   Classes  CM VLDL LDL HDL Density (g/ml) < 0.94 0.95-1.006 1.019-1.063 1.063-1.210 Diameter (nm) 75-1200 30-80 18-25 5-12 Total Lipid (% wt) 98-99 90-92 75-80 40-48 Triacylglycerol (% wt lipid) 81-89 50-58 7-11 6-7 Cholesteryl Esters (% wt lipid) 2-4 15-23 47-51 24-45 Cholesterol (% wt lipid) 1-3 4-9 10-12 6-8 Phospholipids (% wt lipid) 7-9 19-21 28-30 42-51 Apolipoproteins  B48,AI,AII,C,E B100, C, E B100 AI, AII, C, E  Table 1.1  Major Classes of Lipoproteins [10].  The outer shell (~2 nm) of all lipoproteins consists of apolipoproteins, unesterified cholesterol, and phospholipids.  The spherical core contains triacylglycerols and cholesteryl esters.  Chylomicrons (CM) and very low-density lipoproteins (VLDL) have the highest contents of triacylglycerols, and 1-10% apolipoproteins by weight.  Low-density lipoproteins (LDL) and high-density lipoproteins (HDL) contain mostly cholesteryl esters in their cores, and 20-50% of apolipoproteins. 8  1.2.3  Lesion Development Though traditionally viewed as a disease marked by the deposition and retention of lipids, atherosclerosis is now seen as a complex inflammatory process induced by elevated and modified low-density lipoproteins (LDL), free radicals, perhaps infectious microorganisms, shear stress, hypertension, toxins after smoking or combinations of these [15].  The interaction between these two processes, LDL retention /modification and inflammation, defines the principal pathogenesis and distinguishes atherosclerosis from all other chronic inflammatory disorders. Atherosclerotic lesions typically present as asymmetric focal thickenings of the innermost layer of the artery, the intima.  Morphologically, lesions can be characterized into six major types that reflect the early, developing, and mature stages of the disease [16, 17].  In type I [17], or lesion-prone arterial sites, adaptive thickening of the intima is among the earliest histological changes.  This is followed by the formation of so-called fatty streaks or type II lesions [17], which are sites of accumulation of lipid droplets and immune cells.  Lipid-laden macrophages (termed foam cells because of the bubbly appearance of lipid droplets throughout their cytoplasm) dominate these fatty streaks, which also contain T cells, dendritic cells, and mast cells.  Continued foam cell formation and macrophage necrosis can produce type III lesions [17] that contain small extracellular pools of lipid.  Type IV lesions [16] are defined by a relatively thin tissue separation of the lipid core from the arterial lumen, whereas type V lesions exhibit fibrous thickening of this structure, also known as the lesions' “cap”. 9  Mature type VI [16] lesions exhibit architecture that is more complicated and characterized by calcified fibrous areas.  In this late stage of lesion development, plaques contain a central lipid core that is most often hypocellular.  This necrotic lipid core is separated from the arterial lumen by a fibrous cap that consists of extracellular matrix and smooth muscle cells.  The junction between the cap and morphologically more normal areas of the artery is known as the “shoulder” region.  This area is typically more cellular and many contain a variable composition of smooth muscle cells (SMCs), macrophages, and T cells. Mature atherosclerotic plaques can be categorized as either stable or vulnerable to rupture [16].  Stable plaques tend to be characterized by a smaller lipid core, a thick fibrous cap, and shoulder regions with few inflammatory cells, whereas vulnerable plaques contain considerable lipid in their core, a thin fibrous cap, and a large population of macrophages and T cells in their shoulder regions.  These differences indicate that vulnerable plaques may be weaker structurally and more likely to rupture in response to the physical forces of flowing blood.  1.2.4  Oxidative Modification and LDL Macrophages incubated with LDL in its native form (nLDL) do not internalize excess cholesterol due to down-regulation of the LDL receptor [18].  The premature and severe atherosclerosis that develops in patients with familial hypercholesterolemia who lack functional LDL receptors [19] highlights the importance of LDL receptor- independent mechanisms in foam cell formation.  Goldstein et al. [20] observed that 10  acetylated LDL leads to extensive macrophage cholesterol uptake and foam cell formation.  The uptake of acetyl LDL is mediated by a cell-surface receptor now termed scavenger receptor class A (SR-A).  These receptors are characterized by the ability to bind and internalize a wide range of polyanionic ligands [21-23].  Subsequently, Steinbrecher and colleagues showed that oxLDL was rapidly internalized by macrophages via scavenger receptors and that this resulted in cellular cholesterol accumulation [24, 25].  In 1989, Steinberg et al. [12] presented the oxidative modification theory of atherosclerosis, based on the hypothesis that oxidation represents a biologic modification analogous to the in vitro chemical modification discovered by Brown and Goldstein [20] that gives rise to foam cells.  Subsequent work revealed many potentially pro-atherogenic properties of oxLDL in addition to scavenger receptor- mediated uptake by macrophages. Immunohistochemical studies using antibodies to oxLDL have revealed the presence of oxLDL-related epitopes in atherosclerotic lesions [26-30].  Nishi et al. [31] demonstrated that oxLDL levels in carotid plaques were nearly 70 times higher than in plasma from the same patient.  Moreover, plaque oxLDL levels and apoB fragmentation exhibited a strong correlation with macrophage infiltration [31].  1.2.5  Putative Oxidants  Myeloperoxidase (MPO) is a heme-containing enzyme that catalyzes the conversion of Cl- to the 2e-oxidant HOCl.  In vitro studies have shown that HOCl- modified LDL can stimulate macrophage foam cell formation [32] via binding to class B 11  scavenger receptors [33], increase leukocyte adherence and migration into blood vessels [34], increases ROS production by leukocytes [35], and has chemotactic activity for neutrophils [36].  Nitrite, the final oxidation product of nitric oxide metabolism, can also be a substrate for MPO.  The MPO + nitrite reaction generates nitrogen dioxide radicals (·NO2), a reactive species that can oxidize LDL [37, 38].  Indeed, such MPO/·NO2 modified LDL can convert macrophages into foam cells [39].  Heinecke et al. [40] were the first to show the presence of active MPO in human atherosclerotic lesions.  Additionally, MPO activity has been confirmed for lesions in humans [41-43], Wantanabe heritable hyperlipidemic rabbits [44], and rabbits fed a high- cholesterol diet [45].  Lipoxygenases are iron-containing dioxygenases that catalyze the stereospecific insertion of molecular oxygen into polyunsaturated fatty acids to give rise to a complex family of biologically active lipids, including prostaglandins, thromboxanes, and leukotrienes [46, 47].  Lipoxygenases can oxidize complex, esterified fatty acids such as those in cholesterol esters and phospholipids [48-50].  In vitro, 15-lipoxygenase can oxidize LDL [51], and this is achieved by a combination of direct, enzymatic and indirect, non-enzymatic oxidation reactions [52-54].  Several lines of evidence support a role for lipoxygenase in atherosclerosis.  15- lipoxygenase and 5-lipoxygenase are expressed in atherosclerotic lesions of humans [55] and apolipoprotein E deficient (apoE-/-) mice, respectively [56].  Also, disruption of the 12/15-lipoxygenase gene or decreased expression of 5-lipoxygenase gene diminishes disease in apoE-/- and LDL receptor deficient (LDLR-/-) mice [56-58].  Furthermore, 12  inhibiting 15-lipoxygenase lowers lesion formation in rabbits fed a high-fat and high- cholesterol diet [59].  Free transition metals like iron and copper are strong catalysts for oxidation reactions in the presence of hydroperoxides such as LOOH [60].  They can catalyze homolytic cleavage of LOOH to lipid alkoxyl radicals that can initiate lipid peroxidation and other oxidation reactions.  Incubating LDL in serum-free medium in the presence of copper or iron ions mimics the process of cell-mediated oxidation of the lipoprotein [12, 24, 61-63].  This has raised interest in the possibility that transition metals may participate in oxidative modification reactions in atherosclerotic lesions.  This is supported by the presence of free transition metals found in advanced human lesions [64- 66].  1.2.6  Modified LDL and Leukocyte Recruitment Minimally modified LDL (mmLDL) is a term introduced to describe LDL in the initial stages of oxidation where modification of lipids components can occur in the absence any changes to the protein components.  Functionally, mmLDL is defined by its ability to be recognized by the LDL receptor, not recognized by scavenger receptors (ScRs), and possessing pro-atherogenic biological activities distinct from those of nLDL [67-70].  Endothelial cells activated by bioactive phospholipids in oxidized LDL express high levels of leukocyte adhesion molecules such as vascular cell adhesion molecule-1 (VCAM-1).  Cells carrying counter-receptors for VCAM-1, such as monocytes and lymphocytes, adhere at sites of VCAM-1 over-expression [71, 72] (Figure 1).  Platelets 13  also adhere to activated endothelial cells and inhibition of platelet adhesion reduces leukocyte infiltration and atherosclerosis [73].  Once adherent to the endothelium, leukocytes migrate into the underlying intima. By virtue of its lysophosphatidylcholine (lysoPC) content formed during oxidation [24], oxLDL itself is chemotactic for T lymphocytes [74] and monocytes, and chemostatic for macrophages [75].  Additionally, up-regulation in endothelial and SMCs of monocyte chemotactic protein-1 (MCP-1) by mmLDL  [70] facilitates the recruitment of monocytes into the arterial wall [69, 76] (Figure 1).  Consistent with this idea, mice deficient in MCP-1 [77, 78] or its receptor [79] shows attenuated disease development. Thus the enhanced entry and impaired egress of inflammatory cells would be expected to stimulate arterial inflammation, a process strongly implicated in atherosclerosis [80-82].  1.2.7  OxLDL and Foam Cell Formation Once trapped in the arterial wall, monocytes undergo differentiation into macrophages (Figure 1).  The inflamed intima is suspected to produce increased amounts of macrophage colony-stimulating factor (M-CSF), a cytokine that is the major regulator of macrophage differentiation in vitro and whose presence has been demonstrated in atherosclerotic lesions [83-85].  M-CSF deficiency in apoE-/- or LDLR-/- mice attenuates the severity of atherosclerosis [86, 87].  It has also been demonstrated that oxLDL, as well as 7-ketocholesterol or oxidized linoleic acid metabolites, which are components of oxidized LDL, are able to induce differentiation of human monocytes to macrophages both in vitro and in vivo [88-91].  Though the precise mechanism has not been clearly 14  defined, macrophage differentiation is a necessary step for atherosclerosis [92] and is associated with up-regulation of pattern recognition receptors (PPRs), including ScRs that are involved in recognition and phagocytic clearance (Figure 1.1).  This encompasses class A ScRs (SR-AI, SR-AII, SR-AIII, MARCO), class B ScRs (CD36, SR-BI, SR-BII), CD68, and lectin-like oxidized LDL receptor-1 (LOX-1) [23]. Uptake of nLDL via LDL receptors, is regulated by cellular cholesterol content. Thus, incubation of cultured macrophages with even high concentrations of nLDL does not lead to foam cell formation.  In contrast , uptake of oxLDL by macrophages via ScRs is unregulated and incubation of cultured macrophages with acetyl LDL, oxidized LDL, or other forms of modified LDL leads to foam cell formation [93].  Both histochemical and biochemical studies indicate the presence of oxLDL in atherosclerotic lesions, particularly in foam cell-rich areas [94, 95]. Macrophages incubated with acetyl LDL internalize acetyl LDL into lysosomes. The cholesterol ester in acetyl LDL is efficiently degraded, the free cholesterol is transferred to the ER, and is then re-esterified by acyl CoA acyltransferase (ACAT) and stored in cytoplasmic lipid droplets.  In contrast, when macrophages are incubated with oxLDL, lipid accumulates mainly in lysosomes due to impaired lysosomal processing. This lysosomal defect manifests as inefficient apoB degradation, decreased cholesteryl ester hydrolysis, and hence the inability to deliver cholesterol to ACAT for esterification [96-100].  Although the precise mechanism is unknown, there is some evidence of direct lipase and proteinase inhibition by oxLDL [25, 100-102].  These observations indicate 15  that foam cell formation may be a modified type of lysosomal storage disease, one where the enzyme is abundant but other mechanisms inhibit proper hydrolysis of the substrate.  1.2.8  Macrophage Activation  In addition to ScRs, macrophage differentiation is also associated with the up- regulation of Toll-like receptors (TLRs) [103].  Unlike ScRs, TLRs do not mediate endocytosis.  When bound by ligands with pathogen-like molecular patterns, TLRs initiate a signal cascade that leads to macrophage activation [104].  Within atherosclerotic lesions, TLR-1, TLR-2 and TLR-4 have been shown to be expressed by macrophages [103, 105].  Deficiency of TLR-4 or its downstream adaptor molecule, myeloid differentiation factor 88 (MyD88) results in significantly reduced atherosclerosis in apoE-/- mice [106, 107], suggesting that the TLR-4 pathway is pro-atherogenic.  Microbial components such as lipopolysaccharides (LPS), CpG, and microbial DNA and RNA are recognized by various TLRs [108].  Increasing evidence suggests that endogenous ligands can also stimulate TLRs.  MmLDL and oxLDL have been shown to induce actin polymerization and macrophage spreading through binding to CD14 and activation of signaling via TLR-4 and its accessory protein MD2 [109].  This indicates that modified lipoproteins contain motifs recognized by innate pattern recognition receptors and therefore have the ability to influence macrophage function and atherogenesis through signaling events independent of cholesterol accumulation and foam cell formation. 16  Macrophage activation within atheromas results in the release of a broad array of pro-inflammatory mediators.  These include vasoactive molecules such as nitric oxide, endothelins, and eicosanoids [110, 111].  Interleukin-12 (IL-12), IL-15, and IL-18 produced by activated macrophages are instrumental in promoting T-helper 1 (Th1) lineage pro-atherogenic responses [112].  The activated macrophages also produce reactive oxygen species, which has potentially pro-atherogenic consequences for lipoprotein oxidation and cytotoxicity [43].  Finally, activated macrophages secrete matrix metalloproteinases (MMPs) that degrade matrix components and subsequently cause destabilization of atherosclerotic plaques and a resulting increase in the risk for plaque rupture and thrombosis [113-119].  1.2.9  Macrophages and Plaque Destabilization As the atheroma develops, foam cells and extracellular lipid droplets form a core region surrounded by a cap of SMCs and a collagen-rich matrix.  Immunocompetent T cells and macrophages infiltrate throughout the plaque and are particularly abundant not only in the shoulder region where the atheroma grows, but also at the interface between the cap and the core [16, 120, 121]. Accumulating evidence suggests that the primary determinant for plaque vulnerability that can lead to rupture and thrombosis is inflammatory activation in the plaque [122].  In this regard, macrophages play a vital role in vascular remodeling and plaque destabilization through the production of various enzymes, activators, inhibitors and bioactive mediators.  [123].  Macrophages are more abundant in lesions featuring 17  intense inflammatory responses and in vulnerable plaques [124].  Furthermore, activation of macrophages results in a marked increase in expression and release of MMPs [113, 114, 116].  Activated macrophages within plaques express MMP1, MMP2, MMP9, and MMP13 and vulnerable atheromatous plaques are associated macrophage infiltration and increased levels of MMP production [117-119, 125].  Vulnerable plaques are also characterized by the accumulation of apoptotic cells, especially macrophages [126] and SMCs [127] which may contribute to plaque instability [128].  Schrijvers et al. [129] showed that human carotid atherosclerotic lesions contained a substantial number of apoptotic cells that were not engulfed by phagocytes, suggesting that phagocytic clearance in advanced lesions is defective [130-132].  Such perturbed phagocytic clearance may result in a robust inflammatory response by direct triggering of pro- inflammatory responses and inhibition of anti-inflammatory responses in the phagocytes. Therefore, defective phagocytic clearance of apoptotic macrophages in advanced plaques could promote a number of processes that are thought to be important in plaque disruption and acute atherothrombotic vascular occlusion [131, 132]. 18     Figure 1.1  Recruitment and Differentiation of Macrophages in Atheroma [82]. Several leukocyte adhesion molecules and chemokines are involved in monocyte recruitment into the intima.  This is followed by differentiation of monocytes into macrophages.  During this process, pattern recognition receptors such as scavenger receptors (ScRs), are upregulated.  ScRs mediate the uptake of oxidized low-density lipoprotein (oxLDL), and cause the accumulation of LDL-derived cholesterol and foam cell formation. 19   1.3  OxLDL and Macrophage Proliferation, Survival, and Apoptosis 1.3.1  Atherosclerosis and Macrophage Population Though traditionally thought of as terminally differentiated and unable to replicate, there is evidence showing active macrophage proliferation within lesions. Macrophages are the predominant cell type in lesions that express proliferation markers [133, 134].  Deletion of the retinoblastoma (Rb) gene in macrophages has been shown to enhance atherosclerotic development [135], showing a correlation between macrophage proliferation and disease progression.  In addition to their cholesterol lowering effect, statins significantly inhibit macrophage proliferation [136, 137].  Furthermore, a recent study demonstrating that reduction of macrophage apoptosis via inactivation of the Bax gene results in larger atherosclerotic lesions in LDLR-/- mice [138].  Therefore, while the recruitment and retention of monocytes/macrophages within sites of lesion development is of undoubted importance, for a comprehensive understanding of how macrophage populations in the artery are regulated, identifying factors that control the replication and lifespan of macrophages within the arterial intima is also required.  1.3.2  OxLDL and Macrophage Proliferation  In 1993, Yui and colleagues were the first to report the induction of macrophage proliferation by oxLDL [139].  Their subsequent studies showed that this growth effect was due to lysoPC, a major phospholipid component of oxLDL [140-142]. Internalization of lysoPC by macrophages leads to an increase in intracellular Ca2+ ([Ca2+]i) and 20  activation of protein kinase C (PKC).  This ultimately leads to an autocrine release of granulocyte-macrophage colony stimulating factor (GM-CSF), which this group believed is responsible for the oxLDL-induced proliferation of macrophages [143-145].  Although lysoPC is a component of oxLDL that can induce macrophage proliferation, our lab has shown that it is not required for the growth induction of macrophages by oxLDL.  Incubation of oxLDL with fatty acid-free bovine serum albumin (BSA) stripped 97% of the lysoPC content from oxLDL but only decreased its mitogenic activity by 20% [146].  Furthermore, we found that nLDL treated with auto- oxidized arachidonic acid under conditions that caused extensive modification of lysine residues, was equal to oxLDL in mitogenic potency for macrophages even though it had the same low content of lysoPC as nLDL.  This suggested that modified apolipoprotein B (apoB) may be a major growth-stimulating component of oxLDL.  Our lab also demonstrated the activation of phosphatidylinositol 3-kinase (PI-3K) by oxLDL and showed that selective inhibition of PI-3K significantly reduced oxLDL’s ability to induce macrophage proliferation [147].  Biwa et al. has proposed that the sites of action of PKC and PI-3K in oxLDL-mediated macrophage proliferation are distinct [148].  They suggest an upstream role for PKC, whereas PI3K is involved, at least in part, downstream in the signaling pathway after GM-CSF induction.  1.3.3  OxLDL and Macrophage Survival  In addition to the proliferative effects of oxLDL on murine peritoneal macrophages, Hamilton and colleagues reported an anti-apoptotic effect of oxLDL on 21  murine bone marrow derived macrophages (BMDM) that were deprived of growth factor [149].  This effect of oxLDL occurred in the absence of endogenous or exogenous M- CSF and GM-CSF, suggesting a direct role for oxLDL rather than an indirect process mediated through cytokines.  There was a biphasic effect of oxLDL on macrophage viability in that  doses of oxLDL ≤ 50 μg/ml were shown to promote survival in murine and human macrophages, whereas higher concentrations of oxLDL leads to apoptosis [142, 150].  We confirmed that oxLDL blocks apoptosis in BMDM and that nLDL or acetylated LDL had no effect [151].  We also confirmed that high concentrations of oxLDL are toxic, and that soluble factors in the medium are not necessary for the anti- apoptotic effect of oxLDL [152].  There are several mechanisms that may contribute to the anti-apoptotic effect of oxLDL.  Arai et al. have reported that oxLDL can induce the expression of the pro-survival protein AIM (apoptosis inhibitor expressed by macrophages), which is abundant in lesions [153].  The same study also found that targeted deletion of AIM in LDLR-/- mice led to a dramatic reduction in early atherosclerotic lesions.  Our lab has found that oxLDL blocks BMDM apoptosis through the same mechanism of PI-3K activation that was demonstrated for growth induction in peritoneal macrophages [152].  Activation of PI-3K leads to the phosphorylation and activation of one of its downstream targets, protein kinase B (PKB), a protein that represents a nexus for the control of pro-survival signaling pathways [154].  PKB phosphorylation leads to the phosphorylation of the PKB target I-κBα, and subsequent translocation of the 22  transcription factor nuclear factor kappa B (NFκB) into the nucleus, followed by the increased expression of the pro-survival protein Bcl-XL [151, 152].  Upon growth factor withdrawal, there is increased activity of acid sphingomyelinase (ASMase) and increased ceramide generation  [151].  The addition of oxLDL blocks this effect and BMDMs from ASMase-/- mice were found to be resistant to apoptosis induced by growth factor deprivation [151].  This indicates that oxLDL promotes BMDM survival by inhibiting ASMase activity as well as by promoting PKB activation (Figure 2).  1.3.4  Contradictory Reports of Pro-apoptotic Properties for oxLDL  A number of papers have reported that oxLDL is cytotoxic and/or pro-apoptotic for macrophages, SMCs, and fibroblasts [155-170].  On the other hand, several other groups, including our own, have demonstrated that oxLDL promotes the growth and survival of macrophages and SMCs [139, 140, 142, 146, 147, 149, 151, 152, 171-176]. The reasons for such discrepancies are not yet fully understood, but there are a number of possible explanations.  In most cases, investigators who found that oxLDL is cytotoxic used concentrations in excess of 100 μg/ml.  We and others have observed that only at concentrations lower than 50-75 μg/ml does oxLDL promote growth and survival.  The method by which LDL is oxidized is also important [176].  The degree of oxidation and the oxidizing agent used can affect the content of cytotoxic components such as hydroperoxides, reactive aldehydes, oxysterols, and lysophospholipids.  Subsequent steps 23  of dialysis or washing through filtration can reduce cytotoxicity by removing water- soluble aldehydes from the oxidation mixture [177].  It may be possible that oxLDL has a dual role in the pathogenesis of atherosclerosis.  First, low levels of oxLDL may contribute to the survival and inflammatory response of macrophages during the early stages of the disease.  In advanced lesions, when oxLDL may be more abundant, it could induce macrophage apoptosis leading to an accumulation of apoptotic cells and result in plaque disruption [126]. 24     Figure 1.2  OxLDL Induced Macrophage Survival [151].  OxLDL prevents macrophage apoptosis following M-CSF withdrawal by at least two primary mechanisms: by inhibiting acid sphingomyelinase (thereby preventing ceramide generation) and by directly activating the PI-3K/PKB pathway. PKB-mediated phosphorylation of IκB-α leads to the release and activation of NFκB, which then maintains expression of Bcl-XL. 25  By inhibiting the release of cytochrome c from mitochondria, Bcl-XL prevents the activation of the caspase 9-caspase 3 cascade and subsequent apoptosis.  26  1.4  Sphingolipids 1.4.1  Sphingolipid Generation  Sphingolipids have emerged as a source of important signaling molecules that are potentially involved in pathophysiological processes [178-180].  Sphingolipid metabolites, such as ceramide and sphingosine-1-phosphate (S1P), belong to a new class of bioactive molecules, which are involved in a variety of cellular processes, including cellular differentiation, migration, proliferation and apoptosis [181-184].  The biosynthetic pathway of sphingolipids (Figure 1.3) is complex with several potential points of regulation and modulation [185, 186].  There are two points of entry into the sphingolipid pathway, and both converge on the generation of ceramide, which makes it a key molecule in the generation of the other bioactive sphingolipids.  The first route of entry is the de novo pathway, whereby serine and palmitoyl coenzyme A condense, and through a series of reactions generate dihydroceramide and then ceramide.  The second route is the salvage pathway, which involves the hydrolysis of membrane sphingomyelin by sphingomyelinases.  Once ceramide is generated, it can be converted into a number of other bioactive sphingolipids, including ceramide-1-phosphate, sphingosine, and S1P. Clearance of sphingolipids occurs through their conversion by the enzyme sphingosine kinase (SK) to S1P, which is then hydrolyzed by S1P lyase into hexadecenal and ethanolamine phosphate.    27  1.4.2  SK and S1P  SK and S1P play important roles in many cellular processes, such as the regulation of [Ca2+]i signals [187-191], angiogenesis [192, 193], cell adhesion molecule expression [194], cell survival and proliferation [184, 195-202], and chemotaxis [203]. SK activity is always present in cells due to an intrinsic catalytic activity that is not dependent upon post-translational modifications [204].  This basal level is thought to function in a “house-keeping” role in maintaining low sphingosine levels.   SK can be activated in response to a number of mitogens, including vascular endothelial growth factor (VEGF) [194], platelet derived growth factor (PDGF) [199, 205], nerve growth factor (NGF) [200], epidermal growth factor (EGF) [187], estrogen [206, 207], and fetal calf serum (FCS) [199].  The mechanisms of SK activation includes phosphorylation and translocation to the plasma membrane [204].  Although the reasons are not fully understood, localization of SK to the membrane is integral to its signaling role in enhancing cell proliferation and survival [202].  Intracellular levels of S1P are tightly regulated by the equilibrium between its formation, which is catalyzed by SK, and its degradation, which is catalyzed by S1P lyase and S1P phosphatases [185].  S1P produced in response to agonists has the ability to function intracellularly as a second messenger or after secretion in an autocrine/paracrine fashion to activate S1P receptors (formerly known as EDG receptors) on the cell surface [208].  In addition to cell growth and survival, an important function of S1P is maintaining Ca2+ homeostasis.  Although S1P is thought to mobilize [Ca2+]i via interaction with its surface receptors, increasing evidence suggests an important 28  intracellular role for S1P in mediating Ca2+ increases within the cell [209, 210]. However, the exact mechanism in which S1P mediates Ca2+ mobilization is still uncertain.  Ca2+ release mediated by S1P occurs independently of inositol-1,4,5- trisphosphate (IP3) receptors (IP3R) and ryanodine receptors (RyRs) [211].  One possible candidate is SCaMPER (sphingolipid Ca2+ release mediating protein of the endoplasmic reticulum) [211].  SCaMPER is a 181 amino acid protein that was first shown to mediate sphingolipid-gated Ca2+ release from intracellular stores in Xenopus oocytes.  More recently, antisense knockdown of SCaMPER mRNA was shown to substantially reduce sphingolipid-induced calcium release in human and rat cardiomyocytes [212].  However, SCaMPER shares no similarity to any other known [Ca2+]i channels and is a small protein with only one transmembrane domain [211].  Thus, it is unlikely to itself be a Ca2+ channel.  Furthermore, a study showed that there is little correlation between its intracellular location and that of known [Ca2+]i stores [213].  1.4.3  SK, S1P, and Atherosclerosis  Because of its roles in the expression of adherence molecules in endothelial cells [194], enhanced proliferation of SMCs [175, 201, 214], and in immune cell chemotaxis [203], SK and S1P have been implicated in the pathogenesis of atherosclerosis.  A large component of plasma S1P (> 60%) has also been found to be associated with lipoproteins [215-217].  Futhermore, oxLDL been shown to induce SK activity and proliferation in SMCs [175, 214].  Remarkably, a clinical cohort study found that S1P is more predictive of occlusive CAD than established risk factors including age, sex, family history, 29  diabetes mellitus, lipid profile, and hypertension [218].  The levels of serum S1P also correlate with the severity of the disease [218]. 30     Figure 1.3  Biosynthetic Pathway of Sphingolipids [186].  The pathway of sphingolipid metabolism with the sphingosine kinase enzyme circled in blue and the reaction it catalyzes circled in red.  31  1.5  Calcium 1.5.1  Ca2+ Signaling Toolkit Ca2+ is a ubiquitous intracellular signal responsible for controlling numerous cellular processes.  These processes range from muscle contraction to synaptic transmission, and from cellular proliferation to apoptosis [219].  At any moment in time, the level of [Ca2+]i is determined by a balance between the “on” reactions that introduce Ca2+ into the cytoplasm and the “off” reactions through which Ca2+ is removed by the combined action of buffers, pumps, and exchangers.  These heterogeneous Ca2+ signaling systems are assembled from what is commonly referred to as the Ca2+ signaling toolkit (Figure 1.4).  1.5.2  Spatial and Temporal Organization of Ca2+ Signaling In addition to the Ca2+ signaling toolkit, another factor that contributes to the versatility of Ca2+ signaling is its high degree of spatial and temporal diversity [220]. Action potentials are well-known mechanisms to propagate increases in Ca2+, up to a meter in humans.  Within all cells, Ca2+ release from the ER is a nonlinear, cooperative process.  IP3 binds to four receptor sites on the IP3R, one on each subunit of the tetramer [221].  In both ER and sarcoplasmic reticulum, IP3Rs and RyRs are at first potentiated, then inhibited by Ca2+.  Small perturbations in conditions, such as ambient [Ca2+]i and IP3 levels, result in bursts of local Ca2+ release across a cell (called “sparks” for their appearance in Ca2+ imaging fluorescence microscopy) [222].  Increasing spark frequency can cascade and become regenerative.  This regenerative release culminates in three- 32  dimensional waves (also known as oscillations) of changes in [Ca2+]i that propagate within cells.  Variations in the frequency of these Ca2+ oscillations are often determined by changes in stimulus intensity received by the cell.  Cells respond to such oscillations using highly sophisticated mechanisms, including the ability to interpret changes in frequency.  Ca2+/calmodulin dependent kinase II (CaMKII) [223] and PKC [224] are two such “molecular machines” that can decode frequency-modulated Ca2+ signals. Frequency-modulated Ca2+ signals can regulate specific responses such as exocytosis [225], mitochondrial redox state [226], and differential gene transcription [227-230].  1.5.3  Calmodulin During the “on” reaction, a small proportion of the Ca2+ binds to the effectors that are responsible for stimulating numerous Ca2+ dependent processes.  The archetypal Ca2+ effector is calmodulin (CaM), having changed only slightly over 1.5 billion years of evolution and being transcribed from three separate chromosomes in humans [231]. When Ca2+ binds, the shape of the  CaM domains change, triggering the ability to relieve protein auto-inhibition, remodel active sites, and induce protein dimerization [232].  CaM is shaped like a dumbbell with a flexible joint in its middle [233-235].  The EF hands (motif composed of a helix-loop-helix tha binds Ca2+ in the loop between the two helices) of CaM have distinct affinities for Ca2+, and their binding affinities are often increased by interaction with target proteins.  Binding of Ca2+ is associated with a large change in conformation and exposure of hydrophobic surfaces within each domain. Hydrophobic residues wrap around amphipathic regions of target proteins such as the α 33  helices in myosin light chain kinase (MLCK) and CaMKII.  Ca2+/CaM binding relieves auto-inhibition of the catalytic domain of CaMK family enzymes.  1.5.4  Ca2+ and Cell Proliferation  Changes in [Ca2+]i have been detected as a cell passes through G1, G1/S, and mitosis [236].  The requirement for Ca2+ signals is illustrated by the cessation of cell proliferation when extracellular Ca2+ is lowered from 1 mmol/L to 0.1 mmol/L [237]. Cells are most sensitive to depletion of Ca2+ in G1, in which Ca2+ is important for the transcription of immediate-early genes such as Fos, Jun, and Myc, and later at the G1/S boundary where Ca2+ is required for Rb phosphorylation [238].  CaM is required for cell cycle progression through G1 and mitosis  [239], and CaM antagonists or CaMK inhibitors block cell cycle progression in early or late G1 [237].  1.5.5  Ca2+ and Cell Survival and Apoptosis  Ca2+ released into the cytoplasm from the ER functions as a second messenger that can mediate cell survival or induce apoptosis [240]. This versatility is illustrated by the responses of immature T cells to T cell receptor (TCR) activation [241-244]. Immature T cells, developing in the cortex of the thymus, undergo either positive or negative selection based on the strength of the TCR activation.  The former occurs with weak TCR activation by self-antigens, whereas the latter is induced by strong TCR activation by self-antigens.  Ca2+ signals mediate both processes [245], and distinct temporal patterns of Ca2+ elevation are associated with positive versus negative selection 34  [246, 247].  Weak TCR activation induces Ca2+ oscillations, whereas strong TCR activation induces sustained Ca2+ elevation.  The former activates nuclear factor of activated T cells (NFAT) optimally and thereby up-regulates expression of the pro- survival cytokine IL-2, whereas the latter up-regulates the pro-apoptotic BH3-only protein Bim.  Thus, Ca2+ signals can have dual roles in the same cells in response to the same stimulus, depending on the temporal pattern of the [Ca2+]i elevations (Figure 1.5).  1.5.6  Ca2+ and Atherosclerosis  A number of observations have implicated a role for Ca2+ mobilization in the development of atherosclerosis.  Ca2+ channel blockers (CCBs) were originally developed as vasodilators and are widely used in the treatment of angina pectoris, hypertension, and arrhythmia.  However, a growing body of data from clinical trials suggests that the majority of CCBs may also be effective in preventing the development of atherosclerosis, independent of their hypotensive effects [248-250].  The use of CCBs in apoE-/- mice effectively reduces atherosclerotic lesion formation [251, 252] and decreases the expression of MCP-1, VCAM-1, and tumor necrosis factor-α (TNFα) expression, NFκB translocation, and SMC proliferation [253, 254].  Furthermore, oxLDL-mediated mobilization of [Ca2+]i in macrophages has been shown to be important for foam cell formation [255, 256]. 35     Figure 1.4  Elements of the Ca2+ Signaling Toolkit [220].  Ca2+-mobilizing signals (blue) are generated by stimuli acting through a variety of cell-surface receptors (R), including G-protein (G)-linked receptors and receptor tyrosine kinases (RTK).  The signals generated include:  inositol-1,4,5-trisphosphate (Ins(1,4,5)P3), generated by the hydrolysis of phosphatidylinositol-4,5-bisphosphate (PtdIns(4,5)P2) by a family of phospholipase C enzymes (PLCβ, PLCγ); cyclic ADP ribose (cADPR) and nicotinic acid dinucleotide phosphate (NAADP), both generated from nicotinamide-adenine dinucleotide (NAD) and its phosphorylated derivative NADP by ADP ribosyl cyclase; and sphingosine-1-phosphate (S1P), generated from sphingosine by a sphingosine kinase. ON mechanisms (green) include plasma membrane Ca2+ channels, which respond to 36  transmitters or to membrane depolarization (ΔV), and intracellular Ca2+ channels - the Ins(1,4,5)P3 receptor (InsP3R), ryanodine receptor (RYR), NAADP receptor and sphingolipid Ca2+ release-mediating protein of the ER (SCaMPER).  The Ca2+ released into the cytoplasm by these ON mechanisms activates different Ca2+ sensors (purple), which augment a wide range of Ca2+-sensitive processes (purple), depending on cell type and context.  OFF mechanisms (red) pump Ca2+ out of the cytoplasm: the Na+/Ca2+ exchanger and the plasma membrane Ca2+ ATPase (PMCA) pumps Ca2+ out of the cell and the sarco-endoplasmic reticulum Ca2+ ATPase (SERCA) pumps it back into the ER/SR.  Abbreviations:  TnC –  troponin C; CAM – calmodulin; MLCK – myosin light chain kinase; CAMK – Ca2+/calmodulin-dependent protein kinase; cyclic AMP PDE – cyclic AMP phosphodiesterase; NOS – nitric oxide synthase; PKC – protein kinase C; PYK2 – proline-rich kinase 2; PTP – permeability transition pore. 37     Figure 1.5  Ca2+ Signals Regulate Both Cell Survival and Apoptosis [257].  Sustained [Ca2+]i elevation in response to a wide range of agents induce apoptosis by triggering cytochrome c release from the mitochondria and by activating calpains and caspases. Both caspases and cytochrome c participate in positive feedback loops that enhance Ca2+ release from the ER, thereby augmenting [Ca2+]i elevation and apoptosis.  Ca2+ oscillations favor cell survival by enhancing mitochondrial bioenergetics or by activating pro-survival Ca2+ sensitive transcription factors.   38  1.6  Elongation Factor-2 Kinase 1.6.1  Background    In 1985, Nairn et al. first discovered Ca2+/CaM dependent kinase III (CaMKIII) [258].  In 1987, the same group identified CaMKIII’s only known substrate in mammalian cells as eukaryotic elongation factor-2 (eEF2) [259].  Thereafter, CaMKIII has also been known as eEF2 kinase.  To date, eEF2 is still the only known substrate for eEF2 kinase and eEF2 kinase is the only known enzyme that can phosphorylate eEF2. eEF2 kinase belongs to the recently discovered family of alpha kinases [260].  The vast majority of eukaryotic protein kinases belong to the protein-kinase-like superfamily and possess homologous catalytic domains.  Two groups within this superfamily, the serine/threonine kinases and the tyrosine kinases comprise what is commonly referred to as “conventional protein kinases” (CPKs).  Whereas CPKs phosphorylates amino acids located within loops, turns or irregular structures, alpha kinases phosphorylates amino acids within α helices and do not share any detectable sequence homology to CPKs.  1.6.2  eEF2 Kinase Regulation  eEF2 kinase is a highly conserved protein and is present in virtually all tissues of vertebrates as well as in various invertebrates [261].  When it was first purified, it was discovered that its activity was strictly dependent upon Ca2+ ions and CaM [262, 263].  In the presence of Ca2+ ions and CaM, eEF2 kinase undergoes extensive autophosphorylation, acquiring the ability to maintain its activity in the absence of Ca2+ and calmodulin.  In principle, this would prolong its activation in vivo beyond the 39  duration of a Ca2+ transient.  The activity of eEF2 kinase can also be regulated at several other levels.  These include modulation by mitogenic factors [264-269], oxidizing agents [270, 271], sphingolipids [272], hormones [273-278], binding of its protein chaperone, Hsp90 [279, 280], pH [281], and metabolic stress [282-291]. Presently, there are 7 known phosphorylation sites within eEF2 kinase [261, 292- 297].  Phosphorylation on 6 of these sites either positively or negatively regulates eEF2 kinase activity.  These sites are phosphorylated by the p38 mitogen activated protein kinase (p38 MAPK) [292, 293], mammalian target of rapamycin (mTor) [287, 290, 295- 297], extracellular signal regulated kinase (ERK) [296], adenosine monophosphate- activated protein kinase (AMPK) [283-286, 294], and protein kinsae A (PKA) pathways [268, 297-299].  A summary of these phosphorylation sites and their effect on eEF2 kinase activity can be found in Figure 1.6.  1.6.3  eEF2 Kinase Function  When activated, eEF2 kinase phosphorylates eEF2 at Thr56 [259].  eEF2 is a monomeric GTPase and serves as an elongation factor that facilitates the translocation of peptidyl t-RNA from the ribosomal A site to P site [300].  Phosphorylation of eEF2 inhibits its activity [301], in translocation and in poly(U)-directed polyphenylalanine synthesis [302, 303], by preventing it from binding to the ribosome [304].  A growing body of evidence suggests that one of the key physiological functions of eEF2 kinase is in cellular proliferation and survival.  eEF2 kinase has been found to be selectively activated in proliferating cells [267, 305-307].  Moreover, eEF2 kinase is 40  activated in response to a number of mitogenic factors including bradykinin [264], vasopressin [264], EGF [264, 266], insulin [265], insulin-like growth factor I (IGF-I) [266], and serum [266, 271, 306].  This role in cellular proliferation has been implicated in the tumorigenesis of certain cancers.  The specific activity of eEF2 kinase is markedly increased in human breast tumor specimens compared with that of normal adjacent breast tissue [266].  eEF2 kinase activity is also increased in glioblastomas [308] and inhibition of eEF2 kinase activity significantly decreases the clonogenicity of rat and human glioblastomas [280, 307, 309, 310].  In addition to proliferation, eEF2 kinase also plays an important role in cell survival, particularly during times of metabolic stress.  Two reports have shown increased activity of eEF2 kinase during cerebral ischemia, followed by a return to baseline after reperfusion [282, 291].  The activation of eEF2 kinase and postischemic delayed neuronal cell death was attributed to the activation of glutamate receptors [291].  Pharmacological inhibition of protein translation however, was not found to be neurotoxic by itself, but actually protected neurons against the toxicity evoked by low level activation of glutamate receptors.  Thus, it was hypothesized that phosphorylation of eEF2 and the resulting depression of protein translation may be a mechanism that protects against the long-term toxicity of glutamate.  This hypothesis is supported by a recent study examining cardiomyocyte apoptosis in response to hypoxic injury [286].  This study demonstrated that activation of AMPK and its downstream target, eEF2 kinase, is cardioprotective against hypoxic injury, acting by attenuating ER stress via the mechanism of decreased protein synthesis.  Furthermore, eEF2 kinase’s ability to 41  promote survival in glioblastomas has been recently shown to involve an eEF2 kinase- dependent induction of autophagy [288]. 42     Figure 1.6  Regulatory Phosphorylation Sites on eEF2 Kinase.  Residue numbers of the phosphorylated serines sites are as listed.  Resulting modulation of eEF2 kinase activity are indicated by arrows.  Abbreviations:  mTOR –  mammalian target of rapamycin; S6K1 – p70 S6 kinase 1; ERK – extracellular signal regulated kinase; RSK – p90 ribosomal S6 kinase; MAPKAP – mitogen activated protein kinase activated protein; AMPK – adensosine monophosphate activated protein kinase; PKA – protein kinase A.  43  1.7  Protein Translation 1.7.1  Background  Protein biosynthesis is a central process in all living cells.  It is one of the last steps in the transmission of genetic information encoded in DNA and the basis by which proteins are produced to maintain the specific biological function of a given cell.  Protein synthesis takes place on ribosomal particles where the genetic information transcribed into mRNA is translated to protein.  The process of protein translation on the ribosome consists of three phases:  initiation, elongation and termination (Figure 1.7).  1.7.2  Translation Initiation The initiation phase of eukaryotic protein synthesis involves a large number of eukaryotic initiation factors (eIFs) [311-313]. The process begins when the 80S ribosomes dissociate and the 40S subunits are captured for initiation by binding eIF1A and eIF3.  Initiator tRNA binds, in the form of a ternary complex with eIF2 and GTP, to produce the 43S pre-initiation complex [314, 315].  The 43S pre-initiation complex binds to mRNA at the 5’ terminal mGTP cap structure, and then migrates along the mRNA towards the AUG initiation codon [316, 317].  The initial binding involves the factors eIF4E, eIF4G and eIF4A, which assemble at the 5’-end of mRNA, thus creating the conditions that allow the melting of intra-molecular secondary structures within the mRNA that would otherwise prevent the binding of the 43S pre-initiation complex [311, 313, 318-320].  When the 43S pre-initiation complex stops at the initiation codon, the GTP molecule introduced as part of the eIF2 complex is hydrolyzed to GDP, and this 44  provides energy for the ejection of the eIFs bound to the 40S ribosomal subunit [321]. eIF5 is involved in this process, which is likely to accelerate the hydrolysis of GTP.  The release of these factors allows the association of a native 60S ribosomal subunit that reconstitutes an 80S ribosome at the initiation codon, thereby commencing the elongation stage of translation.  1.7.3  Translation Elongation The elongation phase is controlled by three eukaryotic elongation factors, eEF1A, eEF1B and eEF2 [322].  eEF1A forms a ternary complex with GTP and amino-acylated tRNAs (aa-tRNAs).  Binding of this ternary complex to the ribosome places the aa-tRNA in a hybrid A/T site [323].  Ternary complexes containing non-cognate tRNA have equal chance to bind to the ribosome as complexes containing the correct tRNA complementary to the codon in the A-site.  Selection against incorrect tRNAs is performed at two stages. First, coordinated conformational changes in the ternary complex and the ribosome allow the anticodon of the aa-tRNA to contact the mRNA codon in the A-site.  Second, the binding of the cognate tRNA to the codon in the A-site promotes a closed conformation of the small subunit required for ribosome-stimulated GTP hydrolysis by eEF1A.  Upon GTP hydrolysis, eEF1A is released and the aa-tRNA is accommodated in the A-site [324, 325].  eEF1B serves as a guanine nucleotide exchange factor (GEF) and recycles eEF1A·GDP back to its active GTP bound form. After aa-tRNA binding in the A-site and peptide bond formation, a complex comprised of eEF2 and GTP binds to the same site on the ribosome as eEF1A.  This 45  triggers translocation of the A-site tRNA to the P-site with concomitant translocation of the mRNA by one codon [326].  eEF2 hydrolyzes GTP in this process and the resulting complex dissociates from the ribosome, leaving the A-site open for binding of another eEF1A ternary complex.  This translocation is thought to proceed via hybrid “A-P” and “P-E” states of the tRNAs in which the acceptor ends of the tRNAs move first, followed by simultaneous translocation of the mRNA and the anticodon ends of the tRNAs.  The translocation involves a “ratchet-like” rotation of the small subunit with respect to the large subunit.  1.7.4  Translation Termination The process of translation termination resembles the elongation process except that a stop codon, instead of a sense codon, is decoded at the A-site of the ribosome. Recognition of a stop codon is performed by eukaryotic release factor 1 (eRF1) [327].  In the presence of eRF1, eRF3 catalyzes the hydrolysis of the peptidyl-tRNA, accompanied by a concomitant GTP hydrolysis [328, 329].  This termination reaction generates a free nascent polypeptide chain and a free, uncharged tRNA, followed by the release of these components from the ribosome.  1.7.5  Why Regulate Peptide Chain Elongation?  Even though multiple mechanisms exist that regulate translation initiation [330- 333], evolution has deemed it necessary to place regulatory controls over translation elongation as well [300].  Protein synthesis consumes a high proportion of cellular 46  energy, the vast majority of which is used in elongation.  At least four high energy bonds are consumed for each amino acid added to the nascent chain.  It therefore follows that, under conditions of temporarily increased energy demand or decreased energy supply, it would be advantageous for the cell to reduce the rate of protein synthesis to allow energy to be diverted to other processes such as maintaining membrane potentials and ion gradients.  Inhibition of elongation would also ensure that polysomes are retained, allowing mRNA stability and rapid resumption of translation once energy availability improves. 47     Figure 1.7  Three Phases of Protein Translation from mRNA [334].  Initiation:  Only three of the many translation initiation factors required for this process are shown. 48  Efficient translation initiation also requires the poly-A tail of the mRNA bound by poly- A-binding proteins which, in turn, interact with eIF4G.  In this way, the translation apparatus ascertains that both ends of the mRNA are intact before initiating.  Although only one GTP hydrolysis event is shown in the figure, a second is known to occur just before the large and small ribosomal subunits join.  Elongation:  EF-Tu and EF-G are the designations used for the bacterial elongation factors; in eucaryotes, they are called eEF1 and eEF2, respectively.  In the initial binding event an aminoacyl-tRNA molecule that is tightly bound to EF-Tu pairs transiently with the codon at the A-site in the small subunit. During this step, the tRNA occupies a hybrid-binding site on the ribosome.  The codon- anticodon pairing triggers GTP hydrolysis by EF-Tu causing it to dissociate from the aminoacyl-tRNA, which now enters the A-site and can participate in chain elongation.  In subsequent steps, elongation factor EF-G in the GTP-bound form enters the ribosome and binds in or near the A-site on the large ribosomal subunit, accelerating the movement of the two bound tRNAs into the A/P and P/E hybrid states.  Contact with the ribosome stimulates the GTPase activity of EF-G, causing a dramatic conformational change in EF- G as it switches from the GTP to the GDP-bound form.  This change moves the tRNA bound to the A/P hybrid state to the P-site and advances the cycle of translation forward by one codon.  Termination:  The binding of a release factor to an A-site bearing a stop codon terminates translation.  The completed polypeptide is released and the ribosome dissociates into its two separate subunits. 49   1.8  Autophagy 1.8.1  Background  Autophagy refers to any cellular degradative pathway that involves the delivery of cytoplasmic material to the lysosome.  At least three forms have been identified; chaperone-mediated autophagy, microautophagy, and macroautophagy.  This thesis will focus on macroautophagy (herein referred to as autophagy), the major regulated catabolic mechanism that eukaryotic cells use to degrade long-live proteins and organelles.  This form of autophagy involves the delivery of cytoplasmic material sequestered inside double-membrane vesicles to the lysosome [335].  Initial steps include the formation and expansion of an isolation membrane, also called a phagophore.  The edges of the phagophore then fuse to form the autophagosome, a double-membraned vesicle that sequesters the cytoplasmic material.  This is followed by fusion of the autophagosomes with a lysosome to form an autolysosome where the captured material, together with the inner membrane, is degraded (Figure 1.8).  1.8.2  Regulation of Autophagy Autophagy occurs at low basal levels in virtually all cells in order to perform homeostatic functions such as protein and organelle turnover.  It is rapidly upregulated when cells need to generate intracellular nutrients and energy such as during times of starvation, growth factor withdrawal, or high bioenergetic demands [336-340]. Autophagy is also upregulated when cells are preparing to undergo structural remodeling 50  such as during developmental transitions or to rid themselves of damaging cytoplasmic components, for example during oxidative stress, infection, or protein aggregate accumulation [338-341]. Some of the regulatory molecules that control autophagy include (1) mTOR, which shuts off autophagy in the presence of growth factors and abundant nutrients, (2) AMPK, which responds to low energy, (3) eEF2 kinase, which is one of the downstream targets of AMPK and a regulator of protein translation, (4) eIF2α, which responds to nutrient starvation, (5) the tumor suppressor protein p53, (6) the stress activated kinase, c- Jun-N-terminal kinase (JNK), (7) death associated protein kinase (DAPK), (8) the ER- membrane-assiociated protein, insulin response element-1 (IRE-1), (9) Beclin 1/class III PI3K complex, (10) GTPases, (11) ERK1/2, (12) IP3R, (13) ceramide, (14) and calcium [288, 336, 339-343].  1.8.3  Autophagy in Cell Survival and Apoptosis  Under most circumstances, autophagy constitutes a metabolic stress adaptation pathway that promotes cell survival [336, 344, 345].  An apparent paradox is that autophagy is also considered a form of nonapoptotic programmed cell death called “type II” or “autophagic” cell death, which has been observed in response to viral infection, ER stress, toxins, and chemotherapy drugs [346-349].  This type of cell death has been historically defined by morphological criteria, but is now clear that the mere presence of autophagosomes in dying cells is insufficient to distinguish “cell death with autophagy” from “cell death by autophagy” [340].  It is not yet understood what factors determine 51  whether autophagy is cytoprotective or cytotoxic and whether cytotoxicity occurs as the result of self-cannibalism, the specific degradation of cytoprotective factors, or other as of yet undefined mechanisms. 52     Figure 1.8  The Cellular Aspects of Autophagy [350].  The cellular events during autophagy follow distinct stages: vesicle nucleation (formation of the isolation membrane/phagophore), vesicle elongation and completion (growth and closure), fusion of the double-membraned autophagosome with the lysosome to form an autolysosome, and lysis of the autophagosome inner membrane and breakdown of its contents inside the autolysosome.  53  1.9  Objectives  Several laboratories including our own have documented a role for oxLDL as a macrophage growth factor.  The preliminary study by Hamilton et al. demonstrated a novel mechanism that might account for the persistence of macrophage derived foam cells in the atherosclerotic lesion, namely enhanced macrophage survival [149].  Our group has previously showed that oxLDL inhibits apoptosis in macrophages via activation of the PI-3K/PKB pathway and subsequent increase of the pro-survival protein Bcl-XL [151, 152].  We were not convinced, however, that this was the only anti- apoptotic effect induced by oxLDL.  The main objective of this study was to further elucidate the mechanisms in which oxLDL promotes macrophage survival.  BMDMs represent a homogenous cell population that has an absolute requirement for M-CSF for survival [351, 352].  BMDMs reproducibly undergo apoptosis within 24 hours of M-CSF withdrawal and are therefore used in this study to ascertain the pro- survival effects of oxLDL in macrophages.  OxLDL represents a heterogeneous population of modified forms of LDL that differ greatly in their chemical and function properties [353].  In contrast to mmLDL with only lipid modifications [68, 70], heavily oxidized LDL has both lipid and protein modifications [12, 24, 61-63].  Antibodies against these protein modifications have been used to detect the presence of oxLDL within atherosclerotic lesions [26-31].  Furthermore, the extent of apoB fragmentation has been linked to plaque vulnerability and macrophage infiltration [31].  Therefore, we only use heavily oxidized LDL in our studies.  A previous study showed a dose dependent oxLDL mediated anti-apoptotic effect in BMDMs [152].  At 25 µg/ml, oxLDL 54  was able to completely inhibit apoptosis, therefore in most cases, our experiments were carried out with 25 µg/ml of oxLDL. 55  2  Materials & Methods 2.1  Materials Dulbecco's Modified Eagle’s Medium (DMEM), fetal bovine serum (FBS), Dulbecco’s phosphate buffered saline (DPBS), 4-(2-hydroxyethyl)-1- piperazineethanesulfonic acid (HEPES), 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'- tetraacetic acid tetraacetoxy-methyl ester (BAPTA-AM), Fluo-4-AM, pluronic acid, and propidium iodide (PI) were purchased from Invitrogen (Burlington, ON, Canada). Recombinant human M-CSF (rhM-CSF) was from R&D Systems (Minneapolis, MN).  3- (4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H- tetrazolium, inner salt (MTS) was from Promega (Madison, WI).  Phenazine methosulfate (PMS), anisomycin, geldanamycin, myeloperoxidase (MPO), glucose oxidase, catalase, triglyceride standards, cholesteryl oleate, phosphatidylcholine (PC), lysophosphatidylcholine (lysoPC), sphingomyelin, forskolin, isobutylmethylxanthine (IBMX), 9-β-D-arabinofuranoside (araA), and 5-aminoimidazole-4-carboxamide-1-β-d- ribofuranoside (AICAR) were obtained from Sigma-Aldrich (St. Louis, MO).  C2- ceramide was from Avanti Polar Lipids (Alabaster, AL).  Rottlerin, SB202190, TX-1918, U-73122, sphingosine kinase inhibitor (SKI), and hydrated 1-(((5-(4-nitrophenyl)-2- furanyl)-methylene)-amino)-2,4-imidazolidinedione sodium salt (dantrolene) were purchased from Calbiochem (San Diego, CA).  Protein A Sepharose beads, L- [35S]methionine, and L-[4,5-3H]leucine were from Amersham Biosciences (Piscataway, NJ).  [32P]ATP was from Perkin Elmer (Waltham, MA).   TS-2 and TS-4 were kind gifts from Dr. Y. Uehara (National Institute of Infectious Diseases, Tokyo, Japan).  BCA 56  protein assay reagents, BSA standards, and SuperSignal Femto Substrate were purchased from Pierce (Milwaukee, WI).  Hsp90 antibody was from Stressgen (Victoria, Canada). Calmodulin antibody was from Transduction Laboratories (San Jose, CA).  Ro 32-0432, and PKCδ antibody were from Santa Cruz Biotechnology (Santa Cruz, CA).  Antibodies to phospho-eEF2 (Thr56), p38, and phospho-p38 (Thr180/Tyr182) were obtained from Cell Signaling Technology (Beverly, MA).  Goat anti-rabbit IgG horseradish peroxidase- conjugated secondary antibody was from DAKO Diagnostics (Mississauga, ON, Canada).  G60 silica gel plates were from Whatman (New Jersey, NY).  SDS-PAGE molecular weight standards and polyvinylidene difluoride (PVDF) membranes were provided by Bio-Rad (Hercules, CA).  BioMax MR film was from Kodak (Rochester, NY).  2.2  Lipoprotein Isolation and Oxidation Low density lipoprotein (d = 1.019-1.063 g/ml) was isolated from ethylenediaminetetraacetic acid (EDTA)-anticoagulated whole blood of healthy, normolipidemic donors collected after 18 hours of fasting.  Plasma was separated from whole blood by centrifugation at 800 x g for 20 minutes at 10°C.  The density of plasma was adjusted to 1.020 g/ml using NaBr and centrifuged at 200,000 x g for 20 hours at 10°C.  The bottom layer (d > 1.019 g/ml) was collected and re-adjusted to a density of 1.070 g/ml.  The solution again underwent centrifugation at 200,000 x g for 20 hours at 10°C.  The top layer  (d < 1.063 g/ml) was collected and dialyzed three times against 57  DPBS containing 10 µmol/L EDTA.  After filtration through a 0.45 micron filter, final protein concentration of LDL was then determined using BCA protein assay. Copper oxidation was performed by incubating LDL (200 µg/ml) with 5 µmol/L CuSO4 in DPBS containing 0.90 mmol/L CaCl2 and 0.49 mmol/L MgCl2 for 24 hours at 37°C.  The reaction was stopped by addition of 40 µmol/L butylated hydroxytoluene (BHT) and 300 µmol/L EDTA.  The oxidized LDL was then washed and concentrated to approximately 1.5 mg/ml using Amicon Centricon Plus-20 ultrafilters (Millipore, Bedford, MA).  After a 0.45 micron filtration, protein concentration of oxidized LDL was determined using BCA protein assay.  All oxLDL used in this study is copper oxidized LDL unless otherwise stated. Myeloperoxidase (MPO) oxidized LDL (MPO-LDL) was prepared by incubating LDL (200 µg/ml) at 37°C in 50 mmol/L sodium phosphate, pH 7.0 and 10 μmol/L EDTA in the presence of 30 nmol/L MPO, 500 µmol/L glucose, 125 nmol/L glucose oxidase, and 0.5 mmol/L NaNO2 for 8 or 24 hours. The oxidation reaction was stopped with 300 nmol/L catalase and 40 µmol/L BHT. The oxidized LDL was then washed and concentrated to approximately 1.5 mg/ml using Amicon Centriplus 20 ultrafilters (Millipore, Bedford, MA).  After a 0.45 micron filtration, protein concentration of MPO- LDL was then determined using BCA protein assay.  2.3  Characterization of Modified Lipoproteins Relative electrophoretic mobility (Rf) of modified lipoproteins was assessed using a Ciba-Corning (East Walpole, MA) electrophoresis apparatus and TITAN GEL 58  (Beaumont, TX) lipoprotein agarose gels in 50 mmol/L barbital buffer, pH 8.6 according to manufacturer’s instructions.  BSA was added to lipoprotein samples to ensure reproducible migration distances.  Lipoprotein bands were visualized by staining with Fat Red 7B.  Rf values of modified LDL was calculated by dividing the distance traveled during electrophoresis by the distance traveled by nLDL.  All oxLDL used in this study was extensively modified with an Rf value greater than or equal to 3, unless otherwise specified. Lipid extraction was carried out as follows: 0.4 ml of 0.5 mg/ml modified lipoprotein was first added to 1.5 ml of cold 1:2 CHCl3/MeOH and vortexed.  Then, 0.5 ml of cold CHCl3 was added and vortexed, followed by 0.5 ml of cold 10 mmol/L HClO4 and vortexing.  After centrifugation at 1,500 x g for 5 minutes at 0ºC, the bottom CHCl3 layer was carefully transferred to another tube and the CHCl3 evaporated under nitrogen gas. Thin layer chromatography (TLC) analysis of the modified lipoproteins was performed on G60 silica gel plates.  The concentration of each lipid applied was approximately 1 μg/ml.  The plates were developed in a solvent system of hexane/diethyl ether/acetic acid (70:30:1, vol/vol/vol).  Lipids were visualized by spraying the plate with 50% sulfuric acid and heating to 200ºC for 45 minutes.  2.4  Animals CD1 and C57BL/6 mice were obtained from Charles River Laboratories (Wilmington, MA).  B6.129-Tlr2tm1Kir/J mice with a targeted deletion of the Toll-like 59  receptor 2 (TLR-2) gene and C57BL/10ScNJ mice with a targeted deletion of the TLR-4 gene, and B6 were obtained from The Jackson Laboratory (Bar Harbor, MA).  GFP- LC3#53 transgenic mice expressing green fluorescence protein (GFP)-tagged LC3 were from the Riken Bioresource Center (Ibaraki, Japan).  LOX-1 KO mice with a targeted deletion of the lectin-like oxLDL receptor-1 (LOX-1) gene were kindly provided by Dr. Tatsuya Sawamura (National Cardiovascular Center Research Institute, Osaka, Japan). eEF2 KD transgenic mice expressing a catalytically inactive eEF2 kinase gene were a gift from Dr. Christopher Proud (University of British Columbia, Canada).  2.5 Cell Culture L929 cells (kindly provided by Dr. J.W. Schrader, Biomedical Research Centre, BC, Canada) were seeded in TufRolTM roller bottles (BD Falcon, San Jose, CA) at a density of 1.5 x 104 cells per cm2 and cultured in media (DMEM, 10% FBS, 2 mmol/L L- glutamine, 1 mmol/L sodium pyruvate, 50 U/ml penicillin, 50 μg/ml streptomycin) containing 20 mmol/L HEPES at 37°C with approximately 5% CO2.  After 15 days, the media was removed and centrifuged at 800 x g for 10 minutes.  Afterwards, the supernatant was filter sterilized through a 0.22 micron filter.  This L929-cell conditioned media (LCM) was then used as a crude source of M-CSF containing approximately 10,000 U/ml of activity [354]. Bone marrow cells were obtained from the femurs of 6-8 week old female mice as previously described [151].  Cells were cultured in media containing 10% LCM for 18 hours at 37°C in a 95% humidified atmosphere containing 5% CO2.  After 18 hours, non- 60  adherent cells were isolated and differentiated into macrophages by culturing them in medium containing 10% LCM until 80% confluence was reached (5-6 days).  Cells were washed to remove non-adherent cells and lifted using a rubber cell scraper (Sarstedt, Montreal, QC, Canada). Peripheral blood mononuclear cells (PBMCs) were isolated from the whole blood of consenting healthy donors and isolated on Ficoll-PaqueTMPlus (Pharmacia Biotech, Uppsala, Sweden) gradients according to the manufacturer’s protocol.  PBMCs were washed three times with DPBS and resuspended in media.  The cells were then plated onto tissue culture dishes at a concentration of 1.5 x 106 cells per cm2 and incubated at 37°C in a 95% humidified atmosphere containing 5% CO2.  After two hours, non- adherent cells were removed by washing three times with warm DPBS.  The remaining adherent cells were differentiated into macrophages by culturing them in media containing rhM-CSF (50 ng/ml) until 80% confluence was reached (5-6 days).  Cells were washed to remove non-adherent cells and lifted using a rubber cell scraper. Human promyelocytic HL-60 cells (ATCC, Manassas, VA) were cultured in media at 37°C in a 95% humidified atmosphere containing 5% CO2.  2.6  Cell Viability Assay BMDM were seeded in 96-well plates at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were washed and incubated with medium with or without compounds as indicated for 24 hours.  MTS/PMS solution was then added to each well to a final concentration of 333 µg/ml MTS and 25 µmol/L PMS. After incubation for 2 hours at 37 61  °C, the absorbance at 490 nm was recorded using a Molecular Devices VersaMax microplate reader.  Correlation between macrophage number and formation of formazan product has been previously established [147].  Each condition was performed in triplicate within the experiment and data is representative of at least 3 independent experiments.  Statistical significance was calculated by unpaired Student’s t-Test.  2.7  Apoptosis Assay BMDM were seeded in 6-well plates at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were then washed and incubated with medium containing compounds as indicated for 24 hours.  Cells were harvested using a rubber cell scraper and fixed in 70% cold ethanol for 30 minutes.  Cells were then washed with DPBS and stained with 3 μmol/L PI in DPBS containing 0.1% Triton X-100 and 0.73 μmol/L RNase A.  DNA content was analyzed by flow cytometry on the FL-3 channel with appropriate gating used to exclude debris and cellular aggregates.  Ten thousand events were counted for analysis.  Each condition was performed in triplicate within the experiment and representative of at least 3 independent experiments.  Statistical significance was calculated by unpaired Student’s t-Test.  2.8  Calcium Imaging BMDM were seeded in 6-well plates at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were then washed with Ca2+ free DPBS and incubated in Ca2+ free, HEPES buffered DPBS containing 2 μmol/L Fluo-4-AM (Fluo-4-AM was dissolved with 20% 62  pluronic acid in DMSO to make a 2 mmol/L stock solution) for 30 minutes at room temperature.  Cells were then washed with DPBS and incubated in HEPES buffered media for 10 minutes at room temperature to allow for de-esterification of the acetoxymethyl group.  Media was then removed and fresh media containing test compounds were added. Fluorescence was measured in real-time using an inverted Leica TCS SP2 AOBS laser scanning confocal microscope with a 10X objective.  Image analysis was performed using Leica LCS software and fluorescence of every cell in each field was measured.  On average, 68.2 +/- 11.1 cells were separately analyzed per condition in each experiment.  Each condition was performed in duplicate within the experiment and data shown are representative of at least 3 independent experiments. Statistical significance was calculated by unpaired Student’s t-Test.  2.9  Sphingosine Kinase Activity Assay  BMDM were seeded in 60 mm plates at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were then washed and incubated with medium in the absence of M-CSF for 4 hours.  oxLDL (25 µg/ml) was then added for the time points indicated.  Afterwards, cells were washed with DPBS and lysed with ice-cold solubilization buffer (50 mmol/L tris-Cl pH 8.0, 150 mmol/L NaCl, 1% Nonidet P-40 (IGEPAL CA-630), 10% glycerol, 154 nmol/L aprotinin, 2.90 μmol/L bestatin, 2.34 μmol/L leupeptin, 1.46 μmol/L pepstatin, 2.80 μmol/L trans-epoxysuccinyl-L-leucylamido(4-guanidino) butane (E-64), 1 mmol/L sodium fluoride).  Lysates were centrifuged at 20,000 x g for 10 minutes, and the protein content of supernatants quantified by BCA protein assay.  Equal amounts of 63  protein were then incubated with 50 μmol/L sphingosine in 0.4% fatty acid-free BSA, 10 μCi of [32P]ATP and 100 mmol/L MgCl2.  The reaction was carried out for 30 minutes at 37 °C and stopped by the addition of 20 µl of 1 N HCl and 800 µl of chloroform/methanol/HCl (100:200:1, vol/vol/vol). After 10 minutes, 240 µl of chloroform and 240 µl of 2 mol/L KCl were added, and the samples were centrifuged at 3000 x g for 5 minutes.  The aqueous layer was aspirated, and 250 µl of the organic layer were transferred to new glass tubes. The samples were evaporated under nitrogen gas and then resuspended in chloroform/methanol/HCl (100:200:1, vol/vol/vol).  Labeled S1P was resolved by thin layer chromatography on G60 silica gel plates with 1- buthanol/methanol/acetic acid/water (80:20:10:20, vol/vol/vol/vol). Labeled S1P was then imaged and quantified using a Bio-Rad FX phosphor-imager.   2.10  eEF2 Kinase Activity Assay 3.0 x 107 HL-60 cells were washed with DPBS and lysed with 1 ml ice-cold solubilization buffer.  Lysates were centrifuged at 20,000 x g for 10 minutes, and supernatants were incubated with 3 μg of anti-eEF2 antibody and rotated at 4°C for 2 hours.  Lysates were further incubated for 1 hour with 30 μl of Protein A Sepharose beads.  The beads were then washed four times with solubilization buffer and used as a substrate in the assay. BMDM were seeded in 100 mm dishes at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were then washed and incubated with medium in the absence of M-CSF 64  for 4 hours followed by treatments as indicated.  Afterwards, cells were washed with DPBS and lysed with ice-cold solubilization buffer.  Lysates were centrifuged at 20,000 x g for 10 minutes, and the protein content of supernatants quantified by BCA protein assay.  300 μl of lysate containing 300 μg total protein was then transferred to the substrate beads and incubated at 37°C for 1 hour.  Beads were spun down and washed 3 times with ice-cold solubilization buffer.  They were then mixed with sample buffer (100 mmol/L tris-Cl pH 6.8, 70 mmol/L SDS, 10% glycerol, 1.5 mmol/L bromophenol blue, 150 mmol/L β-mercaptoethanol) and heated to 70°C for 10 minutes.  The proteins were then analyzed by immunoblotting with an antibody to phosphorylated eEF2.  2.11  Immunoblotting Cells were washed with DPBS and lysed with ice-cold solubilization buffer. Lysates were centrifuged at 20,000 x g for 10 minutes, and the protein content of supernatants was quantified using BCA protein assay.  Sample buffer was added to the lysates and heated to 70°C for 10 minutes.  50 µg of protein from each sample was loaded onto a SDS-PAGE gel.  Gels were calibrated using pre-stained SDS-PAGE low molecular weight standards.  Proteins were then transferred electrophoretically to PVDF membranes and then incubated with 1 μg/ml primary antibody in tris(hydroxymethyl)aminomethane buffered saline (TBS), pH 7.4 containing 3 mmol/L sodium azide, and either 1% skim milk or 1.5 mmol/L BSA at room temperature for 2 hours.  After three washes with TBS containing 0.1% Tween 20 (TBS-T), membranes were incubated with horseradish peroxidase-conjugated secondary antibody at 1:10,000 65  dilution in TBS containing 3 mmol/L sodium azide, and either 1% skim milk or 1.5 mmol/L BSA at room temperature for 30 minutes.  After six washes with TBS-T, membranes were treated with SuperSignal Femto Substrate and chemiluminescence was visualized by exposure to BioMax MR film.  Results shown are representative of at least 3 independent experiments.  2.12  Protein Synthesis Assay BMDM were seeded in 6-well plates at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were washed and incubated with media in the absence of M-CSF for 3 hours.  Cells were then washed and incubated with media containing 1/10th the normal concentration of either methionine (0.0201 mmol/L) or leucine (0.0802 mmol/L) in the absence of M-CSF for 1 hour.  OxLDL at a concentration of 25 μg/ml was then added for the times indicated. 5μCi of either L-[35S]methionine or L-[4,5-3H]leucine was added 10 minutes prior to harvesting of cells.  Cells were then washed with DPBS and lysed with ice-cold solubilization buffer.  Protein was precipitated by adding trichloroacetic acid (TCA) to a final concentration of 0.6 mmol/L.  The precipitate was washed three times with 0.6 mmol/L TCA and finally with 95% ethanol.   Radioactivity was then measured using a scintillation counter.  Each condition was performed in triplicate within the experiment and  representative of at least 3 independent experiments.  Statistical significance was calculated by unpaired Student’s t-Test.  66  2.13  Transmission Electron Microscopy  BMDM were seeded in 150 mm dishes at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were then washed and incubated with medium with or without compounds as indicated for 24 hours.  Afterwards, cells were washed with cacodylate buffer (100 mmol/L sodium cacodylate, 175 mmol/L sucrose, 2 mmol/L CaCl, pH 7.4), harvested using a rubber cell scraper, and fixed with 2.5% glutaraldehyde in cacodylate buffer for 2 hours.  Cells were then washed three times with cacodylate buffer and fixed with 40 mmol/L osmium tetroxide in cacodylate buffer for 1 hour.  After three further washes with cacodylate buffer, cells were suspended in 2% molten agar and cut into approximately 1 mm3 blocks.  Dehydration was achieved through a series of graded ethyl alcohols from 30% to 100%.  The schedule was as follows:  30% for 15 minutes, 50% for 15 minutes, 90% for 15 minutes, and three times 100% for 10 minutes each.  After dehydration, infiltration was achieved as follows:  two times 100% propylene oxide for 15 minutes, 1:1 propylene oxide/Epon for 2 hours, 100% Epon for 16 hours, 100% Epon for 24 hours at 60ºC.  After sectioning and staining, the cells were viewed with a Hitachi H7600 transmission electron microscope.  2.14  Microtubule Associated Protein Light Chain 3 Localization  BMDM from GFP-LC3#53 transgenic mice were seeded in Lab-TekTM chamber slides (Nunc, Rochester, NY) at 5.0 x 104 cells per cm2 and grown for 24 hours.  Cells were then washed and incubated with media containing compounds as indicated for the time periods specified.  Afterwards, cells were washed with DPBS and fixed with 4% 67  paraformaldehyde for 30 minutes.  Images were captured using an upright Zeiss laser scanning confocal microscope. 68  3  oxLDL Generates an Oscillatory Increase In [Ca2+]i 3.1  Introduction  A number of groups have demonstrated that oxLDL mobilizes Ca2+ in murine peritoneal macrophages and macrophage-like cell lines [143, 355-357].  Furthermore, an increase in [Ca2+]i was found to be required for oxLDL’s ability to induce macrophage proliferation [143].  The following experiments were undertaken to determine if oxLDL treatment in BMDMs also results in an increase in [Ca2+]i, and if so, to determine the mechanisms involved.  69  3.2  Results 3.2.1  OxLDL Generates an Oscillatory Increase in [Ca2+]i We visualized calcium mobilization in BMDM in response to oxLDL treatment using Fluo-4-AM and confocal fluorescence microscopy.  The fluorescence intensity of each cell was measured continually over a one minute period immediately after addition of nLDL or oxLDL.  On average, 68.2 +/- 11.1 cells were analyzed individually per condition in each experiment.  Cells showing a doubling of fluorescence intensity over the baseline were scored as positive for an increase in [Ca2+]i. OxLDL treatment induced an increase in [Ca2+]i in 66.4% of the cells, whereas with nLDL, only 26.4% of cells were positive (Figure 3.1c).  Because fluorescence intensity of each cell was independently measured as a function of time, we were able to make the novel observation that the increase in [Ca2+]i induced by oxLDL actually involves calcium oscillations (Figure 3.1b).  54.6% of cells treated with oxLDL demonstrated [Ca2+]i oscillations compared with only 6.2% of cells treated with nLDL (Figure 3.1d).  Additionally, the magnitude of the [Ca2+]i increase was, on average, lower in cells treated with nLDL in comparison with cells treated with oxLDL (Figure 3.1a, b). Because it is generally accepted that [Ca2+]i oscillations and not simply a sustained increase of Ca2+ are required to promote cell survival [246, 247], only assays for [Ca2+]i oscillations were done in further studies.  3.2.2  LysoPC in oxLDL is Not Responsible for the Generation of [Ca2+]i Oscillations 70   OxLDL and one of its components, lysoPC, have both been shown to induce an increase in [Ca2+]i in macrophages [143, 355-357].  While lysoPC was able to elicit [Ca2+]i oscillations in BMDMs, a considerably lower percentage of cells were positive for [Ca2+]i oscillations compared to cells treated with oxLDL (Figure 3.2).  Furthermore, phosphatidylcholine (PC) treatment elicited a similar response to lysoPC (Figure 3.2). Hence even though PC is converted to lysoPC during the LDL oxidation reaction [24], it is unlikely that the lysoPC content in oxLDL is responsible for the observed [Ca2+]i oscillations.  3.2.3  Extracellular Ca2+ Plays a Partial Role in the Generation of [Ca2+]i Oscillations  An increase in [Ca2+]i can be mediated by an influx of Ca2+ from the extracellular environment or from intracellular Ca2+ stores.  BMDM incubated in medium lacking Ca2+ still generated [Ca2+]i oscillations in response to oxLDL.  The percentage of cells showing oscillations was not as high as in cells incubated in media containing Ca2+, but was still significantly higher than in cells treated with nLDL (Figure 3.3).  This indicates that, while the presence of extracellular Ca2+ is required for the full effect of oxLDL, intracellular mechanisms exist that mediate the observed [Ca2+]i oscillations.  3.2.4  Thapsigargin Blocks oxLDL Generated [Ca2+]i Oscillations  During the course of a Ca2+ transient, the “on” reactions are counteracted by the “off” reactions, during which time various pumps and exchangers remove Ca2+ from the 71  cytoplasm.  Sarco-endoplasmic reticulum ATPase (SERCA) is a pump that returns Ca2+ from the cytoplasm to the ER [358].  Using thapsigargin, an epoxide derivative that selectively prevents Ca2+ binding to SERCA [359-361], we showed that inhibiting SERCA effectively blocked oxLDL-generated [Ca2+]i oscillations (Figure 3.4).  This suggests a requirement for Ca2+ re-uptake by the ER to produce [Ca2+]i oscillations.  3.2.5  Inhibition of Phospholipase C or RyR do not Block oxLDL Mediated Macrophage Survival Two well studied mechanisms of Ca2+ release from intracellular stores involve IP3Rs and RyRs.  Activation of phospholipase C (PLC) results in the conversion of phosphatidylinositol-4,5-bisphosphate (PIP2) to diacylglycerol (DAG) and IP3.  The IP3 stimulates IP3R-mediated Ca2+ release from the ER.  U-73122 is a selective inhibitor of PLC in this pathway (IC50 ~ 3 µmol/L) [362].  Inhibition of PLC by U-73122 did not selectively inhibit oxLDL’s pro-survival effect (Figure 3.5a).  Hydrated 1-(((5-(4- nitrophenyl)-2-furanyl)-methylene)-amino)-2,4-imidazolidinedione sodium salt (dantrolene) inhibits Ca2+ release from RyR channels [363].  Inhibition of RyR-mediated Ca2+ release also did not block oxLDL’s pro-survival effect (Figure 3.5b).  3.2.6  S1P Generates [Ca2+]i Oscillations and Promotes Macrophage Survival  S1P acts as a second messenger to induce Ca2+ mobilization within the cell [209, 210].  We demonstrated that S1P generated [Ca2+]i oscillations in BMDMs within the same time frame and as effectively as oxLDL (Figure 3.6).  Furthermore, the same 72  concentration of S1P promoted BMDM survival (Figure 3.7a) and blocked apoptosis (Figure 3.7b) to the same extent as oxLDL.  This suggests that oxLDL may positively regulate S1P levels, which in turn mediates both the generation of [Ca2+]i oscillations and macrophage survival.  3.2.7  SK is Activated in Response to oxLDL  S1P production is mediated by the phosphorylation of sphingosine by SK [185]. Because it has been previously reported that oxLDL can activate SK in SMCs [175, 214], we wanted to determine if oxLDL can activate SK in BMDMs.  Using an in vitro kinase assay measuring the ability of cell lysates to phosphorylate sphingosine, we detected increased SK activity almost immediately after the addition of oxLDL (Figure 3.8).  The timing of SK activation suggested that this could be a mechanism for mediating the [Ca2+]i oscillations observed in response to oxLDL.  3.2.8  Inhibition of SK Blocks oxLDL Mediated [Ca2+]i Oscillation and Macrophage Survival  SK inhibitor (SKI) is a non-ATP-competitive selective inhibitor of SK [364] that effectively blocks oxLDL induced SK activation (Figure 3.8).  SKI completely blocked oxLDL-generated [Ca2+]i oscillations (Figure 3.9) and selectively blocked oxLDL- mediated macrophage survival (Figure 3.10).  These results strongly suggest that oxLDL induced Ca2+ mobilization is mediated by increased generation of S1P via SK activation and links Ca2+ signaling with the pro-survival effects of oxLDL. 73   3.2.9  TLR-2, TLR-4, and LOX-1 are not Required for oxLDL Mediated Macrophage Survival Ca2+ mobilization is detected almost immediately after the addition of oxLDL. The timing strongly suggests the involvement of a receptor-activated mechanism.  TLR- 2, TLR-4, and LOX-1 have all been shown to either bind to, become activated, or be up- regulated in response to oxidatively modified LDL or its components [105, 109, 365- 372].  However, none of these receptors seemed to be necessary for oxLDL’s pro- survival effect (Figure 3.11).  74    Figure 3.1  OxLDL Generates an Oscillatory Increase in [Ca2+]i.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy. After incubation with fluo-4-AM, BMDM were washed and media containing either (a) nLDL (25 µg/ml) or (b) oxLDL (25 µg/ml) was added at time 0.  Each tracing shows relative fluorescence values recorded from a single representative cell.  (c) Fluorescence values as a function of time were measured for every cell in the field and cells positive for an increase in [Ca2+]i are expressed as a fraction of total cells.  (d) Fluorescence values as a function of time were measured for every cell in the field and cells positive 75  for [Ca2+]i oscillations are expressed as a fraction of total cells.  ** p < 0.01 compared to cells treated with nLDL.  76    Figure 3.2  LysoPC in oxLDL is not Responsible for the Generation of [Ca2+]i Oscillations.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy.  After incubation with fluo-4-AM, BMDM were washed and media containing either nLDL (25 µg/ml), oxLDL (25 µg/ml), PC (10 µmol/L), or lysoPC (10 µmol/L) was added at time 0.  Fluorescence values as a function of time were measured for every cell in the field and cells positive for [Ca2+]i oscillations are expressed as a fraction of total cells.  ** p < 0.01 compared to cells treated with oxLDL.  77    Figure 3.3  Extracellular Ca2+ Plays a Partial Role in the Generation of [Ca2+]i Oscillations.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy.  After incubation with fluo-4-AM, BMDM were washed and nLDL (25 µg/ml), oxLDL (25 µg/ml) in media containing Ca2+, or oxLDL (25 µg/ml) in Ca2+-free media was added at time 0.  Fluorescence values as a function of time were measured for every cell in the field and cells positive for [Ca2+]i oscillations are expressed as a fraction of total cells.  * p < 0.05 compared to cells treated with nLDL.  78    Figure 3.4  Thapsigargin Blocks oxLDL Generated [Ca2+]i Oscillations.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy. After incubation with fluo-4-AM, BMDM were washed and media containing nLDL (25 µg/ml ), oxLDL (25 µg/ml), or oxLDL (25 µg/ml) + thapsigargin (1 µmol/L) was added at time 0.  Fluorescence values as a function of time were measured for every cell in the field and cells positive for [Ca2+]i oscillations are expressed as a fraction of total cells.  ** p < 0.01 compared to cells treated with oxLDL alone.  79    Figure 3.5  Inhibition of Phospholipase C or RyR do not Block oxLDL Mediated Macrophage Survival.  (a) BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence U-73122 or dantrolene at the concentrations indicated for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media. 80    Figure 3.6  S1P Generates [Ca2+]i Oscillations.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy.  After incubation with fluo-4- AM, BMDM were washed and media containing either nLDL (25 µg/ml), oxLDL (25 µg/ml), or S1P (30 µmol/L) was added at time 0.  Fluorescence values as a function of time were measured for every cell in the field and cells positive for [Ca2+]i oscillations are expressed as a fraction of total cells.  ** p < 0.01 compared to cells treated with nLDL.   81    Figure 3.7  S1P Promotes Macrophage Survival.  BMDM were washed and incubated with media alone, oxLDL (25 µg/ml), or S1P (30 µmol/L) for 24 hours.  (a) Viability was 82  measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  (b) Apoptosis was assessed by measuring the percentage of cells with subdiploid DNA.   Data are expressed as a fraction normalized to fluorescence values of cells cultured in the absence of M-CSF.  Representative histogram from each condition is also shown.  ** p < 0.01 compared to cells treated with media alone. 83    Figure 3.8  SK is Activated in Response to oxLDL.  BMDM were washed and incubated in medium without M-CSF for 4 hours.  oxLDL (25 µg/ml) or oxLDL (25 µg/ml) + SKI (30 µmol/L) was then added for the time periods indicated.  SK activity was assessed by measuring the ability of lysates to phosphorylate sphingosine in an in vitro kinase assay.   Data are normalized to values of cells at time 0.  * p < 0.05 compared to cells at time 0.   84    Figure 3.9  Inhibition of SK Blocks oxLDL Generated [Ca2+]i Oscillations.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy. After incubation with fluo-4-AM, BMDM were washed and media containing nLDL (25 µg/ml), oxLDL (25 µg/ml), or oxLDL (25 µg/ml) + SKI (30 µmol/L) was added at time 0.  Fluorescence values as a function of time were measured for every cell in the field and cells positive for [Ca2+]i oscillations are expressed as a fraction of total cells.  ** p < 0.01 compared to cells treated with oxLDL alone. 85    Figure 3.10  Inhibition of SK Blocks oxLDL Mediated Macrophage Survival. BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the 86  presence or absence of SKI (30 µmol/L) for 24 hours.  (a) Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  (b) Apoptosis was assessed by measuring the percentage of cells with subdiploid DNA.   expressed as a fraction normalized to fluorescence values of cells cultured in the absence of M-CSF.  Representative histogram from each condition is also shown.  ** p < 0.01 compared to cells treated with oxLDL alone. 87    Figure 3.11  TLR-2, TLR-4, and LOX-1 are Not Required for oxLDL Mediated Macrophage Survival.  BMDM from (a) TLR-2 -/-, (b) TLR-4 -/-, or (c) LOX-1 -/- mice were washed and incubated with media alone or oxLDL (25 µg/ml) for 24 hours. Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  88  3.3  Discussion  A number of groups have reported that oxLDL induces an overall increase in [Ca2+]i [143, 355-357], and our studies extend this observation by demonstrating that this increase is in fact an oscillatory increase.  These oscillations are mediated in part by intracellular mechanisms, as depleting extracellular Ca2+ did not completely abolish the effect.  Inhibiting SERCA completely blocked [Ca2+]i oscillations, suggesting a role for Ca2+ re-uptake by the ER.  The addition of oxLDL resulted in an almost immediate activation of SK, which can increase S1P levels by phosphorylating sphingosine. Moreover, S1P was shown to be as effective as oxLDL in blocking macrophage apoptosis and producing [Ca2+]i oscillations.  This suggests that the mechanism in which oxLDL generates [Ca2+]i oscillations may be (1) oxLDL-mediated activation of SK, (2) SK- mediated increase in S1P levels (3) S1P-mediated Ca2+ release from the ER via SCaMPER, and (4) SERCA-mediated Ca2+ re-uptake back into the ER. Inhibition of SK activation blocks not only oxLDL-generated [Ca2+]i oscillations, but also oxLDL-mediated macrophage survival.  This links Ca2+ signaling with the pro- survival effects of oxLDL.  Delivery of S1P by oxLDL itself is unlikely because S1P is lost during the oxidation process [217].  Furthermore, nLDL, which have been shown to contain significant amounts of S1P [217], does not elicit a Ca2+ response similar to S1P, suggesting that oxLDL activates SK via a cell surface receptor-activated pathway. To date, a cell surface receptor that is responsible for mediating the oxLDL- induced pro-survival signal transduction pathways has not been identified.  Our group has shown that oxLDL promotes survival in BMDMs from mice that lack both CD36 and 89  SR-AI/II to the same extent as BMDMs from wild type mice, even though uptake of oxLDL was reduced by approximately 60% (unpublished data).  This suggests that oxLDL’s pro-survival effect is independent of its uptake by cells.  Matsumura et al. showed that pertussis toxin completely blocked the oxLDL induced rise in [Ca2+]i and inhibited peritoneal macrophage growth by 50% [143].  This indicates the involvement of a G protein-coupled receptor (GPCR).  However, our group has recently demonstrated that pertussis toxin itself can block BMDM apoptosis, in part by inhibiting ASMase activity, which in turn prevents ceramide generation [373].  Conversely, mastoparan, a Gi activator, increased ceramide levels in BMDMs and induced apoptosis.  TLR-2, TLR-4, and LOX-1 have all been shown to either bind to, become activated or up-regulated in response to oxidatively modified LDL or its components [105, 109, 365-372].  However, we show that none of these receptors are required for oxLDL mediated macrophage survival.  Therefore, while a receptor-mediated mechanism is likely, the identity of this putative receptor is still unknown.  This chapter has summarized work demonstrating that oxLDL mediates a Ca2+ signal by inducing oscillations of intracellular Ca2+.  I demonstrated that the Ca2+ release was atypical, in that it did not involve IP3, but rather was most likely controlled by elevation in S1P levels in response to oxLDL.  Studies in chapters 4-6 investigated the effects of the Ca2+ oscillations on downstream events that are involved in oxLDL- mediated suppression of BMDM apoptosis.  90  4  PKC Is Not Involved In oxLDL-Mediated BMDM Survival 4.1  Introduction  The PKC family consists of at least 12 kinases with distinct roles in cell proliferation, differentiation, apoptosis, and angiogenesis [374, 375].  There are nine PKC genes that code for isozymes classified into three groups:  classical or conventional PKCs (cPKCs; PKCα, PKCβI, PKCβΙΙ and PKCγ), which are Ca2+-dependent and activated by both phosphatidylserine (PS) and diacylglycerol (DAG); novel PKCs (nPKCs; PKCσ, PKCδ, PKCε, PKCη and PKCθ), which are Ca2+-independent and regulated by DAG and PS; and atypical PKCs (aPKCs; PKCζ and PKCλ), which are Ca2+-independent and do not require DAG for activation, although PS can regulate their activity [376-378]. Matsumura et al. reported that the addition of oxLDL to mouse peritoneal macrophages induced a 2.2-fold increase in PKC activity after 5 minutes and a 4.4 fold- increase after 10 minutes, followed by a decline back to baseline at 20 minutes [143]. The incubation of macrophages with the PKC inhibitors, calphostin C or H-7, significantly inhibited the oxLDL-induced increase in both [3H]thymidine incorporation and cell number in a dose-dependent manner [143].  Furthermore, Hamilton et al. demonstrated the mitogenic activity of phorbol 12-myristate 13-acetate  (PMA), a potent activator of PKC, in mouse peritoneal macrophages [379-381].  These results implicate a role for PKC in oxLDL-mediated macrophage proliferation.  Because it has also been reported that oxLDL induces a near-instantaneous increase in [Ca2+]i in peritoneal macrophages [143, 355], it has been suggested that oxLDL-dependent PKC activation is mediated by calcium [148]. 91   The following experiments were undertaken to test the hypothesis that PKC plays a role in oxLDL-mediated BMDM survival as it apparently does in oxLDL mediated proliferation of peritoneal macrophages.  92  4.2  Results 4.2.1  Inhibition of Ca2+-Sensitive PKC Isoforms do not Alter BMDM Viability Of the PKC isoforms that are activated in response to Ca2+ (α, β, γ), only PKCα and PKCβ are expressed in macrophages [382, 383].  Therefore, we used the PKCα/β selective inhibitor Ro 32-0432 to investigate its possible role in BMDM survival.  Ro 32- 0432 inhibits PKCα (IC50 = 9 nmol/L) and to a slightly lesser extent PKCβ (IC50 = 28 nmol/L) [384].  At concentrations significantly above these IC50 values, Ro 32-0432 did not affect the viability of BMDM nor the ability of oxLDL to promote survival (Figure 4.1).  4.2.2  Stimulation With PMA do not Alter BMDM Viability  Hamilton et al. previously reported that stimulation with PMA induces proliferation of mouse peritoneal macrophages [379-381].  PKC is known to serve as a major receptor for phorbol esters.  PMA is a structural analogue of DAG that activates cPKCs and nPKCs directly, both in vivo and in vitro [377].  Using concentrations ranging from 0.625 nmol/L to 10 nmol/L, PMA stimulation had no effect on BMDM viability, or on their survival response to oxLDL treatment (Figure 4.2).  4.2.3  Rottlerin Selectively Inhibits the Pro-Survival Effect of oxLDL  We tested a number of isoform-selective PKC inhibitors and found that only rottlerin was capable of blocking oxLDL’s pro-survival effect (Figure 4.3).  Rottlerin is a widely used PKC inhibitor with selectivity towards PKCδ (IC50 = 3-6 μmol/L) and to a 93  lesser extent, cPKCs (IC50 = 30-42 μmol/L) [385].  Figure 4.3 shows 25 μmol/L rottlerin decreased the viability of BMDM treated with oxLDL to levels similar to that in BMDMs not treated with oxLDL.  At the same time, the viability of cytokine starved cells treated with 25 μmol/L rottlerin was unaffected.  This shows that rottlerin is able to selectively inhibit the pro-survival effect of oxLDL.  Because inhibition of cPKCs with Ro 32-0432 had no effect as shown above, we investigated the possible role of PKCδ in BMDM survival.  4.2.4  PKCδ is not Involved in Rotterlin’s Inhibition of oxLDL Mediated Macrophage Survival  Prolonged treatment with PMA causes down regulation of cPKC and nPKC protein levels by interaction with their DAG binding sites [383].  Using a PKCδ specific antibody, we confirmed that after 72 hours of PMA treatment, PKCδ protein levels were drastically reduced compared to controls (Figure 4.4A).   However, cells with reduced levels of PKCδ responded identically to oxLDL treatment as compared with cells possessing normal levels of PKCδ (Figure 4.4B).  These results indicate that PKCδ is not involved in oxLDL-mediated macrophage survival and that the observed effects of rottlerin on BMDM survival were independent of PKCδ.  It turns out that rottlerin is also a potent inhibitor of eEF2 kinase (IC50 = 5.3 μmol/L) [386], which leads one to speculate that rottlerin's ability to block oxLDL-mediated BMDM survival might be through its 94  ability to block eEF2 kinase.  Studies to address that possibility form the basis for the next section of this thesis.  95    Figure 4.1  Inhibition of Ca2+-Sensitive PKC Isoforms do not Alter BMDM Viability.  BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence of absence of Ro 32-0432 (1 μmol/L) for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media. 96    Figure 4.2  Stimulation With PMA do not Alter BMDM Viability.  BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence of PMA at the concentrations indicated for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media. 97   Figure 4.3  Rottlerin Selectively Inhibits oxLDL’s Pro-Survival Effect.  BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence or absence of rottlerin at the concentrations indicated for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  * p < 0.05, ** p < 0.01 for oxLDL treated cells in the presence of rottlerin compared with oxLDL treated cells in the absence of rottlerin. 98    Figure 4.4  PKCδ is not Involved in Rotterlin’s Inhibition of oxLDL Mediated Macrophage Survival.  BMDM were cultured for 48 hours in the presence or absence or 10 nmol/L PMA. Cells were then washed and incubated with media alone or with oxLDL (25 µg/ml), in the presence or absence of PMA (10 nmol/L) for 24 hours.  (a) PKCδ protein levels were assessed by immunoblotting.  (b) Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media. 99  4.3  Discussion  The mechanism by which oxLDL induces proliferation of peritoneal macrophages has been reported to involve a rise in [Ca2+]i, followed by activation of PKC and the subsequent release of GM-CSF into the culture medium [387].  In addition, Martens et al. provided evidence that PI-3K is important in oxLDL induced macrophage proliferation [147], and Hundal et al. demonstrated that the primary target of PI-3K pro-survival signaling was PKB [151, 152].  Hundal et al. also demonstrated that the pro-survival effect of oxLDL in BMDM, did not involve autocrine release of growth factors into the culture medium.  Our results indicate that PKC is not required for oxLDL-mediated BMDM survival.  Using a PKCα/β selective inhibitor and depletion of PKC protein levels by prolonged treatment of PMA, we demonstrated that both Ca2+ sensitive and Ca2+ insensitive PKC isoforms are not involved in this signaling pathway.  Furthermore, we showed that PMA does not increase survival or proliferation of BMDMs, in contrast to findings previously reported with other macrophage cell types [379-381].  Our observation that rottlerin, a widely used inhibitor for PKCδ, was able to selectively inhibit the pro-survival effect of oxLDL through a mechanism independent of PKCδ, led us to search for other targets of rottlerin.  As noted above, it has been reported that rottlerin is a potent inhibitor of eEF2 kinase [385] and this prompted me to focus on eEF2 kinase as a potential mediator of the anti-apoptotic effect of oxLDL.  100  5  Elongation Factor-2 Kinase is Required for oxLDL- Mediated Macrophage Survival 5.1  Introduction  Numerous reports have demonstrated that oxLDL induces an immediate increase in [Ca2+]i in macrophages and macrophage-like cells [143, 355-357].  Results in chapter 3 of this thesis showed that in reponse to oxLDL treatment, BMDMs show an oscillatory increase in intracellular Ca2+ and that this was essential for the pro-survival effect of oxLDL.    I found that this pro-survival effect did not require PKC activation although rottlerin, a PKCδ and eEF2 kinase inhibitor, was able to selectively inhibit the pro- survival effect of oxLDL. I inferred that the effect of rottlerin might be due to inhibition of eEF2 kinase, and so undertook the following experiments to test the hypothesis that eEF2 kinase is required for oxLDL-mediated macrophage survival.  eEF2 kinase is a Ca2+/CaM dependent kinase whose activity is regulated by modulation in  [Ca2+]i [262, 263].  Ca2+/CaM, can induce autophosphorylation of eEF2 kinase, and this autophorphorylation can lead to sustained activation of eEF2 kinase, even if [Ca2+]i levels decline. 101  5.2  Results 5.2.1  eEF2 Kinase Activity is Increased in Response to oxLDL  eEF2 is the only known substrate for eEF2 kinase and eEF2 kinase is the only known enzyme that can phosphorylate eEF2 at Thr56 [259].  A commercial phospho- Thr56 specific eEF2 antibody is available.  However it was developed against a human epitope and had never been tested in murine macrophages.  Our attempts to use this antibody for immunoblotting against BMDM lysates resulted in either no clear band at the predicated weight using low concentrations of antibody, or numerous non-specific bands if we increased antibody concentration.  Modification of blocking buffers and incubation times of antibody were of no avail.  I sought advice from a recognized expert on eEF2 kinase, Dr Christopher Proud (University of British Columbia, Canada), and was informed that this antibody fails to recognize Thr56 phosphorylated eEF2 in many cell types, particularly primary cells.  Consequently, we developed a novel, non-radioactive assay to measure eEF2 kinase activity.  Briefly, human eEF2 was purified from human promyelocytic leukemia HL-60 cells and used as a substrate in an in vitro kinase assay. Phosphorylation of eEF2 at Thr56 was detected by immunoblotting with the commercial phospho-Thr56-eEF2 antibody.  Using this assay, we were able to detect increased eEF2 phosphorylation within 15 minutes of oxLDL treatment and maximal phosphorylation within 30 minutes, followed by a decrease in phosphorylation at 60 minutes (Figure 5.1). This indicates that eEF2 kinase is activated in response to oxLDL in macrophages.  5.2.2  eEF2 Kinase Activition is Required for oxLDL’s Pro-Survival Effect 102   Since the earlier characterization of rottlerin as an eEF2 kinase inhibitor, three more eEF2 kinase selective inhibitors have been developed.  TS-2 and TS-4 are 5,6- dihydro-4H-1,3-selenazine derivatives that are 12-fold more potent and specific (TS-2, IC50 = 0.36 µmol/L , TS-4, IC50 = 0.31 µmol/L) for eEF2 kinase than rottlerin [388].  TX- 1918 is a 2-hydroxyarylidene-4-cyclopentene-1,3-dione compound that also has high potency and specificity (IC50 = 0.44 µmol/L) towards eEF2 kinase [389].  Figure 5.2 shows that in the presence of TS-2, TS-4, or TX-1918, the viability of BMDM treated with oxLDL was decreased to levels similar to untreated BMDM.  At the same time, the viability of cytokine-starved cells treated with these inhibitors at corresponding concentrations showed little effect.  This suggests that the effect of these inhibitors is due to selective inhibition of the oxLDL pro-survival pathway and not to nonspecific toxicity. The inhibition of BMDM viability by TS-4 and TX-1918 was accompanied by a corresponding increase in apoptosis (Figure 5.3).  At the same concentrations, both TS-4 and TX-1918 effectively blocked oxLDL-induced eEF2 phosphorylation (Figure 5.4). These results support the hypothesis that oxLDL’s ability to block macrophage apoptosis is dependent upon its ability to activate eEF2 kinase.  5.2.3  eEF2 Kinase Activation Requires Ca2+ Mobilization OxLDL mobilizes [Ca2+]i almost immediately after its addition, and this effect was blocked by the cell-permeable calcium chelator, 1,2-bis(o-aminophenoxy)ethane- N,N,N',N'-tetraacetic acid tetraacetoxy-methyl ester (BAPTA-AM) (Figure 5.5).  At the same concentration, BAPTA-AM also blocked phosphorylation of eEF2 by oxLDL 103  (Figure 5.4), suggesting that eEF2 kinase was activated in response to the oxLDL- mediated increase in [Ca2+]i.  Furthermore, when low concentrations of BAPTA-AM and rottlerin, which individually had no effect, were added together, they blocked oxLDL’s ability to promote BMDM survival (Figure 5.6).  5.2.4  OxLDL-Mediated Macrophage Survival do no Involve the mTor, ERK, or PKA Pathways  Ser78 of eEF2 kinase can be phosphorylated via a mTOR dependent pathway and this leads to inhibition of its kinase activity [295].  Ser 366 can be phosphorylated by both S6K1 and RSK via mTOR- and Erk-dependent pathways respectively, again leading to inhibition of eEF2 kinase activity [296].  PKA can phosphorylate and activate eEF2 kinase at Ser499 [297].  To determine if any of these pathways are involved in oxLDL’s ability to promote macrophage survival, the mTOR inhibitor rapamycin [390], the ERK inhibitor U0126 [391, 392], the PKA agonists forskolin [393] and isobutylmethylxanthine (IBMX) [394], and the PKA inhibitor H89 [395] were used.  Figure 5.7 shows that none of these compounds except perhaps forskolin altered the viability of cells treated with oxLDL after growth factor withdrawal nor the proportion undergoing apoptosis.  While Figure 5.7c indicates that 100 μM of forskolin was able to decrease the viability of macrophages treated with oxLDL, this concentration is over 10-fold higher than the effective concentration of forskolin (IC50 = 7 μmol/L) in macrophages [396]. Additionally, neither IMBX nor H89 show any ability to alter macrophage viability, 104  therefore the effect seen with 100 μmol/L forskolin is most likely non-specific and independent of PKA activation.  5.2.5  eEF2 Kinase Activity Requires Hsp90  During purification, eEF2 kinase has been found to be tightly associated with heat shock protein 90 (Hsp90) [279].  Hsp90 is a protein chaperone responsible for maintaining proper protein folding and stability [397].  Previously, Yang et al. have shown that disruption of the eEF2 kinase:Hsp90 complex can inhibit clonogenicity of several glioblastomas [398].  To evaluate the role of eEF2 kinase:Hsp90 interaction in oxLDL-induced activation of eEF2 kinase, we used geldanamycin, an ansamycin- derivative benzoquinone compound that binds to Hsp90 and disrupts its protein interactions [399].  As expected, the addition of geldanamycin to oxLDL-treated cells blocked its effect on eEF2 kinase activity (Figure 5.4), macrophage survival (Figure 5.8a), and cellular apoptosis (Figure 5.8b). Interestingly, in BMDM incubated with oxLDL, Hsp90 was found in a complex with CaM and rottlerin prevented the formation of this complex  (Figure 5.9).  This result is consistent with previous reports that have implicated a requisite need for the binding of both Ca2+/CaM and Hsp90 for eEF2 kinase activity [279, 280].  5.2.6  p38 MAPK Negatively Regulates eEF2 Kinase Activity  Knebel et al. reported that p38δ can phosphorylate eEF2 kinase at Ser359 and Ser396, and that this inhibits its activity by approximately 80% [400].  p38α and p38β on 105  the other hand can phosphorylate eEF2 kinase at Ser377 and Ser396 [401]. Phosphorylation at Ser377 did not affect eEF2 kinase activity and phosphorylation at Ser396 alone caused only a modest (~20%) decrease in activity, indicating that p38α and p38β have little effect in regulating eEF2 kinase activity.  To assess the role of p38 MAPK in the anti-apoptotic effect of oxLDL, we incubated BMDM with anisomycin, a pyrrolidine antibiotic that activates p38 MAPK [400].  At 250 nmol/L, anisomycin blocked activation of eEF2 kinase by oxLDL (Figure 5.4).  At the same concentration, anisomycin also decreased macrophage viability (Figure 5.10a) and induced apoptosis (Figure 5.10b).  SB202190, a selective inhibitor of p38α and p38β but not p38δ or p38γ [402], was not able to block BMDM apoptosis in response to M-CSF withdrawal (Figure 5.10a), in agreement with the limited role of p38α and p38β in inhibiting eEF2 kinase activity as previously described.  5.2.7  p38 MAPK is Activated upon Growth Factor Withdrawal  Previous studies have reported that p38 MAPK activation mediates growth factor withdrawal-induced apoptosis in certain cell types [403-406].  Specifically, upon growth factor withdrawal in neuronal cells, p38 MAPK becomes activated by dual phosphorylation at its Tyr180/Thr182 residues.  Re-addition of growth factor induces an increase in phosphatase activity that decreases p38 MAPK phosphorylation.  Using a phospho-Tyr180/Thr182 specific p38 MAPK antibody, we observed p38 MAPK phosphorylation as early as 15 minutes after M-CSF withdrawal in BMDMs (Figure 5.11a).  The addition of oxLDL blocked p38 MAPK phosphorylation (Figure 106  5.11b).  Conversely, the addition of anisomycin overrode oxLDL’s inhibition of p38 MAPK phosphorylation (Figure 5.11b).  These results suggest that not only can oxLDL induce activation of eEF2 kinase, it may also inhibit its negative regulation by p38 MAPK.  5.2.8  Ceramide Activates p38 MAPK and Negatively Regulates eEF2 Kinase Activity Our group has previously shown that incubation of BMDM in the absence of M- CSF results in activation of acid sphingomyelinase (ASMase) and an increase in ceramide levels [151].  OxLDL blocked both of these effects.  Conversely, the addition of C2-ceramide blocked the ability of oxLDL to promote survival.  In the present study, we found that C2-ceramide also blocks activation of eEF2 kinase by oxLDL (Figure 5.4). Furthermore, C2-ceramide induced phosphorylation of p38 MAPK (Figure 5.12), in agreement with previous reports in other macrophage cell types [407, 408].  This suggests the possibility that during growth factor withdrawal, eEF2 kinase can be negatively regulated by the activation of ASMase which increases ceramide levels, and these in turn lead to phosphorylation of p38 MAPK.  5.2.9  Myeloperoxidase Oxidized LDL can Promote Macrophage Survival, an Effect that can be Blocked by Inhibiting eEF2 Kinase  OxLDL plays a pivotal role in the progression and development of atherosclerosis, yet it remains controversial how and to what extent LDL becomes 107  oxidized during atherogenesis.  It has been suggested that MPO has a prominent role in promoting oxidative reactions of LDL in vivo [40, 409].  Furthermore, MPO activity is present in human atherosclerotic lesions [41-43].  Nitrite, the final oxidation product of nitric oxide metabolism, can be a substrate for MPO.  The MPO + nitrite reaction generates nitrogen dioxide radicals (·NO2), a reactive species that can oxidize LDL [37, 38].  Indeed, such MPO/·NO2 modified LDL (MPO-LDL) can convert macrophages into foam cells [39].  The electrophoretic mobility (Rf) of MPO-LDL, which is indicative of lysine modifications, is similar to that of copper oxLDL (Figure 5.13c).  The loss of cholesterol esters and triglycerides in MPO-LDL during oxidation is also similar to that of copper-oxidized LDL (Figure 5.13a, b).  We show, for the first time, that MPO-LDL can promote macrophage survival in an oxidation- and dose-dependent manner (Figure 5.14a, b).  Furthermore, MPO-LDL- mediated macrophage survival can be blocked with TX-1918 (Figure 5.15).  Therefore, other oxidatively-modified forms of LDL can also promote macrophage survival via the activation of eEF2 kinase.  5.2.10  BMDM from Transgenic Mice Expressing Catalytically Inactive eEF2 Kinase Shows an Attenuated Survival Response to oxLDL  Dr. Christopher Proud (University of British Columbia, Canada) and colleagues have recently generated transgenic mice that express catalytically inactive eEF2 kinase in place of the wild type gene (unpublished).  Preliminary experiments demonstrated that 108  BMDMs produced from these mice show an attenuated survival response to oxLDL (Figure 5.16). 109    Figure 5.1  eEF2 Kinase Activity is Increased in Response to oxLDL.  BMDM were washed and incubated in medium without M-CSF for 4 hours.  oxLDL (25 µg/ml) was then added for the time periods indicated. Activity of eEF2 kinase was assessed by measuring the ability of lysates to phosphorylate eEF2 in a kinase assay.  Similar results were obtained in at least 3 independent experiments. 110    Figure 5.2  eEF2 Kinase Inhibitors Selectively Block oxLDL’s Pro-Survival Effect. BMDM were washed and incubated with media alone or with oxLDL (25 µg/ml), in the presence or absence of (a) TS-2 (4 µmol/L), (b) TS-4 (3.5 µmol/L), or (c) TX-1918 (8 µmol/L) for 24 hours. Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  ** p < 0.01 compared to oxLDL treated cells in the absence of inhibitor. 111     Figure 5.3  eEF2 Kinase Inhibitors Induce Apoptosis.  BMDM were washed and incubated with media alone or with oxLDL (25 µg/ml), in the presence or absence of (a) TS-4 (3.5 µmol/L) or (b) TX-1918 (8 µmol/L) for 24 hours.  Apoptosis was assessed by measuring the percentage of cells with subdiploid DNA.  Data is expressed as a fraction normalized to fluorescence values of cells cultured in the absence of M-CSF. Representative histogram from each condition is also shown.  ** p < 0.01 compared to oxLDL treated cells in the absence of inhibitor. 112    Figure 5.4  eEF2 Kinase Regulation.  BMDM were washed and cultured in medium without M-CSF for 4 hours.  Then, the p38 MAPK inhibitor SB202190 (15 μmol/L), the p38 MAPK activator anisomycin (250 nmol/L), the intracellular calcium chelator BAPTA-AM (20 μmol/L), C2 ceramide (25 μmol/L), the Hsp90 inhibitor geldanamycin (12 nmol/L), or the eEF2 kinase inhibitors TS-4 (3.5 μmol/L) or TX-1918 (8 μmol/L) were added to cells for 10 minutes.  oxLDL (25 μg/ml) was then added for a further 30 minutes.  Activity of eEF2 kinase was assessed by measuring the ability of lysates to phosphorylate eEF2.  Similar results were obtained in at least 3 independent experiments. 113    Figure 5.5  BAPTA-AM Blocks oxLDL Generated [Ca2+]i Oscillations.  Calcium mobilization was visualized using fluo-4-AM and confocal fluorescence microscopy. After incubation with fluo-4-AM, BMDM were washed and media containing nLDL (25 µg/ml), oxLDL (25 µg/ml), or oxLDL (25 µg/ml) + BAPTA-AM (20 µmol/L) was added at time 0.  Fluorescence values as a function of time were measured for every cell in the field and cells positive for [Ca2+]i oscillations are expressed as a fraction of total cells.  ** p < 0.01 compared to cells treated with oxLDL alone. 114    Figure 5.6  BAPTA-AM and Rottlerin Act Synergistically to Block oxLDL Mediated Macrophage Survival.  BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence of absence of rottlerin (6 µmol/L), BAPTA-AM (12 µmol/L), or rottlerin (6 µmol/L) + BAPTA-AM (12 µmol/L) for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  ** p < 0.01 compared to cells treated with oxLDL alone.  115    Figure 5.7  OxLDL Mediated Macrophage Survival do no Involve the mTor, ERK, or PKA Pathways.  BMDM were washed and incubated with media alone or with 116  oxLDL (25 µg/ml), in the presence or absence of the mTOR inhibitor rapamycin, the ERK inhibitor U0126, the PKA agonist forskolin and IMBX, or the PKA inhibitor H89 at the concentrations indicated for 24 hours. Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media. 117    Figure 5.8  Geldanamycin Inhibits oxLDL Mediated Macrophage Survival and Induces Apoptosis.  BMDM were washed and incubated with media alone or oxLDL (25 118  µg/ml), in the presence of absence of geldanamycin (12 nmol/L) for 24 hours.  (a) Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  (b) Apoptosis was assessed by measuring the percentage of cells with subdiploid DNA.  Data is expressed as a fraction normalized to fluorescence values of cells cultured in the absence of M-CSF.  Representative histogram from each condition is also shown.  ** p < 0.01 compared to cells treated with oxLDL alone. 119    Figure 5.9  OxLDL Induces Hsp90 to Complex with CaM.  BMDM were washed and incubated in medium without M-CSF for 4 hours.  The cells were then incubated with either media alone, oxLDL (25 µg/ml), or oxLDL (25 µg/ml) with rottlerin (25 µmol/L) for 30 minutes.  CaM was immunoprecipitated from cell lysates and Hsp90 protein levels was assessed by immunoblotting. 120    Figure 5.10  p38 MAPK Blocks oxLDL’s Pro-Survival Effect.  BMDM were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence of absence of anisomycin (250 nmol/L) or SB202190 (15 µmol/L) for 24 hours.  (a) Viability was 121  measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  (b) Apoptosis was assessed by measuring the percentage of cells with subdiploid DNA.  Data is expressed as a fraction normalized to fluorescence values of cells cultured in the absence of M-CSF.  Representative histogram from each condition is also shown.  ** p < 0.01 compared to cells treated with oxLDL alone. 122    Figure 5.11  p38 MAPK is Activated upon Growth Factor Withdrawal.  (a) BMDM were washed and incubated without M-CSF for the times indicated. (b) BMDM were washed and incubated without M-CSF for 4 hours.  The cells were then incubated with either media alone, oxLDL (25 μg/ml), or oxLDL (25 μg/ml) + anisomycin (250 nmol/L) for 30 minutes.  Phosphorylation of p38 MAPK was assessed by immunoblotting with a phospho-Tyr180/Thr182-p38 MAPK antibody.  Similar results were obtained in at least 3 independent experiments. 123    Figure 5.12  Ceramide Activates p38 MAPK.  BMDM were washed and C2 ceramide (25 μmol/L) was added for the times indicated.  Phosphorylation of p38 MAPK was assessed by immunoblotting with a phospho-Tyr180/Thr182-p38 MAPK antibody. Similar results were obtained in at least 3 independent experiments. 124    Figure 5.13  Characterization of MPO-LDL.  nLDL was oxidized with MPO under conditions that generated ·NO2 for either 8 hours or 24 hours.  (a) Cholesteryl ester content and (b) triglyceride content were determined by TLC.  (c) Electrophoretic mobility (Rf) was determined by agarose gel electrophoresis.  All values are normalized to nLDL. 125    Figure 5.14  Only Extensively Modified MPO-LDL can Promote Macrophage Survival.  BMDM were washed and incubated with media alone, oxLDL (25 µg/ml), or MPO-LDL that have been oxidized for either (a) 8 hours or (b) 24 hours, at the concentrations indicated for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  ** p < 0.01 compared to cells treated with media alone. 126    Figure 5.15  TX-1918 Blocks MPO-LDL Mediated Macrophage Survival.  BMDM were washed and incubated with media alone, MPO-LDL (50 µg/ml), or MPO-LDL (50 µg/ml) + TX1918 (8 µmol/L) for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  ** p < 0.01 compared to cells treated with MPO-LDL alone.   127    Figure 5.16  BMDM from Transgenic Mice Expressing Catalytically Inactive eEF2 Kinase Shows an Attenuated Survival Response to oxLDL.  BMDM from transgenic mice expressing catalytically inactive eEF2 kinase or from wild type mice were washed and incubated with 10% M-CSF conditioned media or oxLDL at the concentrations indicated for 24 hours.  (a) Viability was measured by the bioreduction of MTS and 128  expressed as a fraction normalized to absorbance values of cells cultured in 10% M-CSF conditioned media.  (b) Apoptosis was assessed by measuring the percentage of cells with subdiploid DNA.  Data is expressed as a fraction normalized to fluorescence values of cells cultured in 10% M-CSF conditioned media.  Representative histogram from each condition is also shown.  129  5.3  Discussion  A growing body of evidence suggests that one of the key physiological functions of eEF2 kinase is in cellular proliferation and survival.  eEF2 kinase can be activated in response to numerous mitogens [264-266, 271, 306], it is selectively activated in proliferating cells [267, 305-307], and its specific activity is markedly higher in certain cancers compared with control tissue [266, 308]. Our studies have demonstrated that oxLDL treatment results in the activation of eEF2 kinase via a Ca2+-dependent mechanism.  Furthermore, specific inhibition of eEF2 kinase activity completely blocks oxLDL’s anti-apoptotic effects.  Upon growth factor withdrawal, p38 MAPK phosphorylation was detected, an effect that can be blocked by the addition of oxLDL.  Phosphorylation of eEF2 kinase by p38 MAPK negatively regulates its activity.  Activation of p38 MAPK by anisomycin was shown to block oxLDL-mediated activation of eEF2 kinase and induce apoptosis.  These results suggest that oxLDL can positively regulate eEF2 kinase activity by both (1) generating an oscillatory increase in [Ca2+]i and (2) inhibiting its negative regulation by p38 MAPK. The precise mechanism by which oxLDL inhibits the activation of p38 MAPK is still unclear.  Other groups have reported that p38 MAPK is activated in response to growth factor withdrawal and that this activation ultimately leads to cellular apoptosis [403, 404].  The growth factor-dependent survival in these cells was mediated by increased expression and decreased degradation of MAPK phosphatase-1 (MKP-1) [406]. MKP-1 is the main phosphatase responsible for dephosphorylating p38 MAPK and preferentially inactivates p38 MAPK over JNK and ERK1/2 [410, 411].  MKP-1 plays an 130  essential role in cell survival signaling [412] and has been recently implicated to be involved in atherosclerotic lesion development in mouse models [413].  Furthermore, both M-CSF and components of oxLDL have been shown to induce MKP-1 expression [414-416].  Future work investigating the potential involvement of MKP-1 in regulating eEF2 kinase activity and oxLDL-mediated macrophage survival would therefore be of interest. In summary, eEF2 kinase activation represents a novel pathway in oxLDL- mediated survival in macrophages.  This finding may provide a mechanistic link between the implicated role of Ca2+ in atherogenesis [251-256] and the growth and survival effects of oxLDL in macrophages.  eEF2 kinase may therefore be of interest as a therapeutic target.  Mice with targeted disruption of its kinase activity are viable and show no distinctive phenotype.  This is consistent with the role of eEF2 kinase in a stress adaptive pathway and not as an essential activity in otherwise healthy individuals.  Toxicity from pharmacological inhibition of this kinase should therefore be within acceptable limits. 131  6  OxLDL Induces Macrophages to Undergo Autophagy 6.1  Introduction  Although it seems paradoxical that inhibition of protein elongation by phosphorylation of eEF2 promotes macrophage survival, there are a number of possible explanations.  The simplest is that inhibition of protein translation preferentially lowers the levels of pro-apoptotic proteins, as many of these have rapid turnover rates.  For example, Terai et al. demonstrated that activation of eEF2 kinase protects cardiomyocytes from hypoxic injury by inhibiting the synthesis of ER stress proteins [286].  The argument that reduction of protein synthesis is a protective mechanism during times of metabolic stress is also compelling.  Protein synthesis consumes a high proportion of cellular energy, the vast majority of which is used in the elongation process. Wu et al. recently provided evidence that the mechanism by which eEF2 kinase activation promotes survival in glioblastomas is via the induction of autophagy [288].  Autophagy, a catabolic mechanism used by eukaryotic cells to degrade proteins and organelles, can be rapidly induced during times of starvation, growth factor withdrawal, or high bioenergetic demands [336-340].  Under most circumstances, autophagy represents a metabolic stress adaptation pathway that promotes cell survival [336, 344, 345].  We therefore tested the hypothesis that activation of eEF2 kinase in response to oxLDL treatment in BMDM would affect protein synthesis rates.  We also embarked on initial studies characterizing the possible role for autophagy, or more precisely, autophagic cell survival, as the means by which oxLDL promotes macrophage survival. 132  6.2  Results 6.2.1  Protein Synthesis is Reduced in Response to oxLDL eEF2 is a monomeric GTPase that facilitates translocation of peptidyl t-RNA from the ribosomal A site to P site [300].  Phosphorylation of eEF2 by eEF2 kinase leads to an inhibition of eEF2 and therefore of protein synthesis [301].  To determine if the oxLDL-mediated phosphorylation of eEF2 inhibits protein synthesis, I measured the in vivo incorporation rates of L-[35S]methionine and L-[4,5-3H]leucine in the presence or absence of oxLDL.  Within 30 minutes of oxLDL treatment, the rates of incorporation of both L-[35S]methionine and L-[4,5-3H]leucine were reduced (Figure 6.1).  This correlated well with the timecourse of eEF2 phosphorylation in response to oxLDL (Figure 5.1).  6.2.2  Autophagic Vacuoles are Present in Macrophages Treated with oxLDL Transmission electron microscopy (TEM) images revealed that BMDMs treated with oxLDL undergo autophagy as indicated by the presence of autophagic vacuoles (Figure 6.2).  The term ‘autophagic vacuole’ refers to both autophagosomes, which are double membrane bound vacuoles containing cytosol and/or organelles devoid of lysosomal proteins, and autolysosomes, which are autophagosomes that have fused with a lysosome (Figure 1.8).  Eskelinen et al. provides a good review on the fine structure of autophagic vacuoles [417]. In many cultured cells, the average diameter of autophagic vacuoles is approximately 600 nm, though they can range from 300 nm to several micrometers. Morphologically, the two limiting membranes are often so close to each other that it is 133  not possible to see them as separate (Figure 6.2b, d).  Sometimes the limiting membrane may appear to contain multiple layers (Figure 6.2a).  When it is possible to see the two limiting membranes, there is a narrow empty, electron-lucent, space between the two membranes (Figure 6.2c).  Ribosomes can be seen inside autophagosomes (Figure 6.2d) and can serve as a marker for cytoplasmic material.  Autophagosomes can fuse with each other, endosomes, or lysosomes.  During fusion, the outer membrane fuses with the endo/lysosome limiting membrane, while the inner membrane still surrounds its cytoplasmic contents (Figure 6.2b).  In late autophagic vacuoles that have fused with lysosomes, the inner membrane is degraded or permeabilized to allow degradation of cytoplasmic contents.  In TEM, this is seen as dark, electron dense areas (Figure 6.2a). Because of the high magnifications used in TEM, an image represents only a small fraction of the total area of a cell.  The nature of such high magnifications make it difficult to randomly capture images and predicates the user to selectively look for structures of interest.  Although it is common in reports involving autophagy to use TEM images to show a lower level or absence of autophagy in certain conditions, we feel that such a presentation may not be free of potential selection bias because of the factors involving the high magnifications mentioned above.  Even though TEM images of BMDM treated with other conditions were captured as controls, we use TEM images only to verify the presence of autophagy and utilize other assays to show differences in the level of autophagy in BMDMs between different conditions.   134  6.2.3  OxLDL Induces Microtubule-Associated Protein Light Chain 3 Aggregation  After the induction of autophagy, microtubule-associated protein light chain 3 (LC3) stably associates with the membrane of autophagosomes [418].  At present, LC3 is the preferred molecular marker for autophagy in mammalian cells [419].  Using BMDM from transgenic mice expressing LC3 tagged with green fluorescent protein (GFP), we observed punctuate fluorescence in oxLDL treated cells, indicative of LC3 aggregation (Figure 6.3).  In contrast, BMDM in 10% M-CSF conditioned media displayed only diffuse cytosolic fluorescence.  6.2.4  AMPK is Activated in Response to oxLDL  When cellular ATP:ADP ratio falls, owing to a stress that inhibits ATP production or increases ATP consumption, adenylate kinase is activated leading to a large increase in AMP:ATP ratio [420].  Binding of AMP to AMPK induces its activation through allosteric changes and allows phosphorylation at Thr172 by upstream kinases [421].  AMPK can inhibit mTOR signalling [422] and/or activate eEF2 kinase [294], both of which results in inhibition of protein synthesis.  AMPK is required for autophagy in yeast [423], and there is evidence that suggests this is also true in mammalian cells. AMPK mediates autophagic cell survival in tumor cells [424] and cardiomyocytes [425, 426] in response to hypoxic injury.  Indeed, AMPK signaling has been shown to mediate a cell’s decision to enter autophagy or apoptosis [427].  To determine if AMPK is involved in oxLDL signaling in macrophages, cell lysates were probed for AMPK phosphorylation.  Use of commercial antibodies against 135  phospho-Thr172-AMPK however, resulted in the appearance of multiple non-specific bands in murine BMDM.  Because these antibodies were all raised against human AMPK epitopes, we used human peripheral blood mononuclear cell (PBMC) derived macrophages.  We confirmed that oxLDL was able to increase the viability of PBMC- derived macrophages deprived of growth factor (Figure 6.4).  Additionally, AMPK activation was detectable as early as 7.5 minutes after oxLDL treatment (Figure 6.5). This time course of activation precedes that which was shown above for eEF2 phosphorylation, allowing for the possibility of AMPK mediated regulation of both eEF2 kinase activation and induction of autophagy by oxLDL.  6.2.5  AMPK Activation Partially Mediates oxLDL Pro-Survival Effects  To determine if AMPK is involved in oxLDL mediated-macrophage survival, we used adenine 9-β-D-arabinofuranoside (araA), an ATP precursor and competitive inhibitor of AMPK [428], and 5-aminoimidazole-4-carboxamide-1-β-d-ribofuranoside (AICAR), an analogue of adenosine and activator of AMPK [429].  AraA partially blocked oxLDL-mediated viability (Figure 6.6a) while AICAR partially rescue cells from growth factor withdrawal (Figure 6.6b).  This suggests that oxLDL mediates macrophage survival, at least in part via AMPK activation. 136     Figure 6.1  Protein Synthesis is Reduced in Response to oxLDL.  BMDM were washed and incubated without M-CSF for 3 hours, and then for 1 hour with medium containing 1/10th their normal concentration of either methionine (0.0201 mmol/L) or leucine (0.0802 mmol/L). oxLDL (25 µg/ml) was then added for the times indicated. 5 μCi of either (a) L-[35S]methionine or (b) L-[4,5-3H]leucine was added 10 minutes prior to harvesting of cells.  The rate of protein synthesis was assessed by measuring the rate of incorporation of L-[35S]methionine or L-[4,5-3H]leucine using a scintillation counter. Data is expressed as a fraction normalized to scintillation values of cells at time 0.  * p < 0.05, ** p < 0.01 compared to time 0. 137    Figure 6.2  Autophagic Vacuoles are Present in Macrophages Treated with oxLDL. BMDM were washed and incubated with oxLDL (25 µg/ml) for 24 hours.  Cells were fixed with osmium tetroxide and images captured using TEM.  The limiting membranes of autophagic vacuoles are indicated by arrowheads.  138   139   Figure 6.3  OxLDL Induces LC3 Localization.  BMDM from GFP-LC3#53 transgenic mice were washed and incubated with 10% M-CSF conditioned media or oxLDL (25 µg/ml) for 24 hours.  Images were captured using fluorescence microscopy. 140    Figure 6.4  PBMC-Derived Macrophage Viability Increases in Response to oxLDL. PBMC-derived macrophages were washed and incubated with media alone or oxLDL at the concentrations indicated for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured in rhM-CSF (50 ng/ml).  ** p < 0.01 compared to cells treated media alone. 141    Figure 6.5  AMPK is Activated in Response to oxLDL.  PBMC derived macrophages were washed and oxLDL (25 μg/ml) was added for the times indicated.  Phosphorylation of AMPK was assessed by immunoblotting with a phospho-Thr172-AMPK antibody. Similar results were obtained in at least 2 independent experiments. 142    Figure 6.6  AMPK Activation Partially Mediates oxLDL’s Pro-Survival Effect. PBMC derived macrophages were washed and incubated with media alone or oxLDL (25 µg/ml), in the presence of absence of (a) araA (500 µmol/L) or (b) AICAR (500 µmol/L) for 24 hours.  Viability was measured by the bioreduction of MTS and expressed as a fraction normalized to absorbance values of cells cultured with rhM-CSF (50 ng/ml).  ** p < 0.01 compared to cells treated with (a) oxLDL alone or (b) media alone. 143  6.3  Discussion  eEF2 kinase is a monomeric GTPase and serves as an elongation factor that facilitates translocation of peptidyl t-RNA from the ribosomal A site to P site [300].  Its GTP-binding motif is located in a region that contains the major physiological phosphorylation site at Thr56 [430, 431].  eEF2 kinase phosphorylates eEF2 at this site and inhibits protein translation [302].  Our studies, which indicate that oxLDL’s ability to promote macrophage survival requires eEF2 kinase activation and subsequent inhibition of protein synthesis, may help explain a variety of experimental observations: that transient inhibition of protein synthesis is an early event in mitogenesis [432, 433]. Inhibitors of protein synthesis such as cycloheximide mimic many of the effects of growth factors, including promoting entry into S-phase [434, 435].  Studies in synapses have demonstrated that N-methyl-d-aspartate (NMDA) receptor activation leads to activation of eEF2 kinase [436].  Even though overall protein synthesis rates were reduced in these cells, a rapid and localized increase in the synthesis of CaMKII, an important regulator of cell cycle progression was observed [237].  Low doses of cycloheximide, which decreases overall rates of protein synthesis independently of eEF2 kinase, are also able to increase CaMKII synthesis.  Even though a reduction in protein synthesis may be an adaptive response during times of stress, the above study indicates that there may be selectivity in terms of which proteins are inhibited.  This may allow the cell to pass through cell cycle checkpoints even though overall protein synthesis is reduced. 144  Autophagy can prevent cells from undergoing apoptosis by maintaining an adequate intracellular supply of substrates despite nutrient depletion [345] or when the uptake of extracellular nutrients is inhibited by a lack of growth factor [336].  Our studies demonstrate for the first time that autophagy occurs in BMDMs treated with oxLDL during growth factor withdrawal.  The precise mechanism by which oxLDL induces autophagy is still unclear, but we show that oxLDL can activate three known regulators of autophagy, AMPK [437],  eEF2 kinase [288], and Ca2+ [341, 438, 439].  Taken together, this suggests a model in which oxLDL promotes BMDM survival by reducing ATP depletion via an induction of autophagy and a reduction in protein synthesis, both of which are mediated by activation of eEF2 kinase. 145  7  Summary OxLDL’s ability to promote macrophage survival has been an ongoing interest in our laboratory for more than a decade.  This could be an important factor in atherogenesis because of its potential role in regulating the size of macrophage populations within the arterial intima.  Our studies reveal a novel mechanism by which oxLDL promotes survival, which is illustrated schematically in Figure 7.1.  To summarize, the addition of oxLDL results in the activation of SK.  This causes an increase in S1P levels through the phosphorylation of sphingosine.  Intracellular Ca2+ mobilization follows, mediated at least in part, through calcium release from the ER.  [Ca2+]i oscillations activate the Ca2+/CaM dependent kinase, eEF2 kinase and specific inhibition of eEF2 kinase activity completely blocks oxLDL’s anti-apoptotic effects.  PKC is not required for the anti- apoptotic effect of oxLDL in BMDM. Growth factor withdrawal increases p38 MAPK phosphorylation, an effect that could be blocked by the addition of oxLDL. Phosphorylation of eEF2 kinase by p38 MAPK negatively regulates its activity.  Activation of p38 MAPK by anisomycin blocked oxLDL-induced activation of eEF2 kinase and induced apoptosis.  These observations suggest that oxLDL positively regulates eEF2 kinase not only by increasing [Ca2+]i, but also by inhibiting its negative regulation by p38 MAPK. Activation of eEF2 kinase results in the phosphorylation and inhibition of the activity of elongation factor eEF2.  Ultimately, oxLDL treatment leads to a reduction in protein synthesis and the induction of autophagy.  This suggests that under conditions of metabolic stress (e.g., growth factor withdrawal), oxLDL mediates an eEF2 kinase- 146  dependent protective mechanism that links energy conservation (via inhibition of protein synthesis) and the replenishment of energy supplies by digestion of cellular organelles through the induction of autophagy. Autophagy is an evolutionarily conserved pathway that is active at a basal level in most cells of the body.  This reflects its role in regulating the turnover of long-lived proteins and the removal of damaged structures, misfolded proteins, and invading micro- organisms.  Self digestion through autophagy is now emerging as a biological pathway that functions to promote health and longevity [440].  Paradoxically, autophagy’s role in regulating metabolism is also implicated in the pathogenesis of various diseases including cancer, neurodegenerative diseases, and muscle and liver disorders [350]. The results from our studies prompt the following question:  does eEF2 kinase activation and the induction of autophagy contribute to the development and progression of atherosclerosis?  The recent development of a transgenic mouse strain that express catalytically inactive eEF2 kinase allows for the opportunity to cross it with mouse strains that are prone to atherosclerosis such as the apoE-/- or LDLR-/- mice.  Monitoring atherosclerotic progression and correlating this with the presence or absence of eEF2 kinase activity would help define the role of eEF2 kinase in atherogenesis.  Additionally, introducing the GFP-LC3 transgene to mice would allow for the in vivo monitoring of autophagy.  This would establish whether or not there is a link between eEF2 kinase activation, induction of autophagy, and the development and progression of atherosclerosis in vivo.  To help delineate cell specificity in this model, adoptive transfer of bone marrow from eEF2 kinase transgenic mice can be used to repopulate the 147  hematopoietic systems of recipient apoE-/- or LDLR-/- mice.  This will result in an absence of eEF2 kinase activity in inflammatory cells including monocytes/macrophages, but not endothelial or smooth muscle cells.  Like so many scientific endeavors, the work in this thesis represents only … “an end to a new beginning”. The discovery of a role for autophagy in atherosclerosis may present an important paradigm in the way we view this disease.  Autophagy may help keep alive those cells that "should" die, such as chemotherapy-treated tumor cells [441, 442], or inflammatory cells trapped in the sub-intimal space of atherosclerotic lesions.  If lysosomal clearance of autophagosomes fails, the activation of autophagy results in a cellular traffic jam that may lead to increased pathology.  Autophagic or type II cell death is defined by the accumulation of autophagic vacuoles [443, 444].  If this accumulation of autophagic vacuoles is concurrent with nutrient depletion, the cells then shift to a type I or apoptotic cell death [445].  This argues against a strict dichotomy between type I and type II cell death and we may discover that the apoptosis observed in late atherosclerotic lesions may result from  impaired lysosomal clearance of autophagosomes. There are several potential methods to target the autophagy pathway.  One could disrupt autophagy by:  (1) blocking the signaling pathways that initiate autophagy by targeting key components involved in autophagosome formation, (2) inhibiting the fusion of autophagosomes with lysosomes to block autolysosome formation, (3) disrupting the recycling of autodigested substrates used for resynthesis of ATP, or (4) blocking the cell’s ability to conserve energy by preventing the termination of protein synthesis.  In this regard, eEF2 kinase becomes increasingly attractive because it is overexpressed in 148  many forms of cancer [266, 308] and is activated during autophagy [288]; its activition terminates protein elongation [302] and conserves energy; and its unique structure [260] makes this kinase amenable to selective inhibition. The emergence of new theories in scientific endeavor does not necessarily exclude past theories, but represents ever-changing forms of insight.  Atherosclerosis was traditionally viewed as a disease marked by the deposition and retention of lipids.  This view is still valid, but its pathogenesis now includes the involvement of a chronic and inappropriate inflammatory response as a critical factor that leads to the development of the disease.  Autophagy as a compensatory mechanism affecting a cell’s decision to survive or die is emerging as an important paradigm in the pathology of a number of diseases.  A discovery that establishes a role for autophagy in the development and progression of atherosclerosis will undoubtedly lead to a greater understanding of the disease and perhaps aid in the future development of novel therapeutic interventions.  149    Figure 7.1  OxLDL Mediated Pro-Survival Signalling in Macrophages, a Working Model.  Abbreviations:  ASMase – acid sphingomyelinase, ATP – adenosine trisphosphate, eEF2K – eukaryotic elongation factor-2 kinase, cytoC – cytochrome C, Hsp90 – heat shock protein 90, M-CSF – macrophage colony stimulating factor, NF-κB – nuclear factor kappa B, PC – phosphatidylcholine, PKB – protein kinase B, PI-3K – phosphatidylinositol-3 kinase, SCaMPER – sphingolipid calcium release mediating 150  protein of the endoplasmic reticulum, SERCA – sarco-endoplasmic reticulum calcium ATPase. 151  Bibliography 1. Murray, C.J. and A.D. Lopez, Global mortality, disability, and the contribution of risk factors: Global Burden of Disease Study. Lancet, 1997. 349(9063): p. 1436- 42. 2. Wilson, P.W., et al., Prediction of coronary heart disease using risk factor categories. Circulation, 1998. 97(18): p. 1837-47. 3. Barrett-Connor, E. and T.L. Bush, Estrogen and coronary heart disease in women. Jama, 1991. 265(14): p. 1861-7. 4. Wilson, P.W., et al., Clustering of metabolic factors and coronary heart disease. Arch Intern Med, 1999. 159(10): p. 1104-9. 5. MacMahon, S., et al., Blood pressure, stroke, and coronary heart disease. Part 1, Prolonged differences in blood pressure: prospective observational studies corrected for the regression dilution bias. Lancet, 1990. 335(8692): p. 765-74. 6. Pyorala, K., M. Laakso, and M. Uusitupa, Diabetes and atherosclerosis: an epidemiologic view. Diabetes Metab Rev, 1987. 3(2): p. 463-524. 7. Ballantyne, C.M., et al., Hyperlipidemia after heart transplantation: report of a 6- year experience, with treatment recommendations. J Am Coll Cardiol, 1992. 19(6): p. 1315-21. 8. Gotto, A.M., Jr. and S.M. Grundy, Lowering LDL cholesterol: questions from recent meta-analyses and subset analyses of clinical trial DataIssues from the Interdisciplinary Council on Reducing the Risk for Coronary Heart Disease, ninth Council meeting. Circulation, 1999. 99(8): p. E1-7. 152  9. Gordon, T., et al., High density lipoprotein as a protective factor against coronary heart disease. The Framingham Study. Am J Med, 1977. 62(5): p. 707- 14. 10. Vance, D.E. and J.E. Vance, Biochemistry of Lipids, Lipoproteins and Membranes. 4th ed. New Comprehensive Biochemistry. 2004: Elsevier B.V. 607. 11. Steinberg, D., Low density lipoprotein oxidation and its pathobiological significance. J Biol Chem, 1997. 272(34): p. 20963-6. 12. Steinberg, D., et al., Beyond cholesterol. Modifications of low-density lipoprotein that increase its atherogenicity. N Engl J Med, 1989. 320(14): p. 915-24. 13. Steinberg, D. and J.L. Witztum, Lipoproteins and atherogenesis. Current concepts. Jama, 1990. 264(23): p. 3047-52. 14. Brown, M.S. and J.L. Goldstein, Lipoprotein metabolism in the macrophage: implications for cholesterol deposition in atherosclerosis. Annu Rev Biochem, 1983. 52: p. 223-61. 15. Ross, R., Atherosclerosis is an inflammatory disease. Am Heart J, 1999. 138(5 Pt 2): p. S419-20. 16. Stary, H.C., et al., A definition of advanced types of atherosclerotic lesions and a histological classification of atherosclerosis. A report from the Committee on Vascular Lesions of the Council on Arteriosclerosis, American Heart Association. Arterioscler Thromb Vasc Biol, 1995. 15(9): p. 1512-31. 17. Stary, H.C., et al., A definition of initial, fatty streak, and intermediate lesions of atherosclerosis. A report from the Committee on Vascular Lesions of the Council 153  on Arteriosclerosis, American Heart Association. Arterioscler Thromb, 1994. 14(5): p. 840-56. 18. Goldstein, J.L. and M.S. Brown, The low-density lipoprotein pathway and its relation to atherosclerosis. Annu Rev Biochem, 1977. 46: p. 897-930. 19. Goldstein, J.L., H.H. Hobbs, and M.S. Brown, Familial hypercholesterolemia, in The Metabolic and Molecular Bases of Inherited Diseases. 2001, McGraw-Hill: New York. p. 2863–2913. 20. Goldstein, J.L., et al., Binding site on macrophages that mediates uptake and degradation of acetylated low density lipoprotein, producing massive cholesterol deposition. Proc Natl Acad Sci U S A, 1979. 76(1): p. 333-7. 21. Krieger, M., et al., Molecular flypaper, host defense, and atherosclerosis. Structure, binding properties, and functions of macrophage scavenger receptors. J Biol Chem, 1993. 268(7): p. 4569-72. 22. Freeman, M.W., Scavenger receptors in atherosclerosis. Curr Opin Hematol, 1997. 4(1): p. 41-7. 23. Moore, K.J. and M.W. Freeman, Scavenger receptors in atherosclerosis: beyond lipid uptake. Arterioscler Thromb Vasc Biol, 2006. 26(8): p. 1702-11. 24. Steinbrecher, U.P., et al., Modification of low density lipoprotein by endothelial cells involves lipid peroxidation and degradation of low density lipoprotein phospholipids. Proc Natl Acad Sci U S A, 1984. 81(12): p. 3883-7. 154  25. Dhaliwal, B.S. and U.P. Steinbrecher, Cholesterol delivered to macrophages by oxidized low density lipoprotein is sequestered in lysosomes and fails to efflux normally. J Lipid Res, 2000. 41(10): p. 1658-65. 26. Itabe, H., et al., A monoclonal antibody against oxidized lipoprotein recognizes foam cells in atherosclerotic lesions. Complex formation of oxidized phosphatidylcholines and polypeptides. J Biol Chem, 1994. 269(21): p. 15274-9. 27. Itabe, H., et al., Sensitive detection of oxidatively modified low density lipoprotein using a monoclonal antibody. J Lipid Res, 1996. 37(1): p. 45-53. 28. Ehara, S., et al., Elevated levels of oxidized low density lipoprotein show a positive relationship with the severity of acute coronary syndromes. Circulation, 2001. 103(15): p. 1955-60. 29. Palinski, W., et al., Cloning of monoclonal autoantibodies to epitopes of oxidized lipoproteins from apolipoprotein E-deficient mice. Demonstration of epitopes of oxidized low density lipoprotein in human plasma. J Clin Invest, 1996. 98(3): p. 800-14. 30. Holvoet, P., et al., Oxidized LDL and malondialdehyde-modified LDL in patients with acute coronary syndromes and stable coronary artery disease. Circulation, 1998. 98(15): p. 1487-94. 31. Nishi, K., et al., Oxidized LDL in carotid plaques and plasma associates with plaque instability. Arterioscler Thromb Vasc Biol, 2002. 22(10): p. 1649-54. 155  32. Hazell, L.J. and R. Stocker, Oxidation of low-density lipoprotein with hypochlorite causes transformation of the lipoprotein into a high-uptake form for macrophages. Biochem J, 1993. 290 ( Pt 1): p. 165-72. 33. Marsche, G., et al., Class B scavenger receptors CD36 and SR-BI are receptors for hypochlorite-modified low density lipoprotein. J Biol Chem, 2003. 278(48): p. 47562-70. 34. Liao, L., et al., Oxidized LDL-induced microvascular dysfunction. Dependence on oxidation procedure. Arterioscler Thromb Vasc Biol, 1995. 15(12): p. 2305-11. 35. Kopprasch, S., et al., Hypochlorite-modified low-density lipoprotein stimulates human polymorphonuclear leukocytes for enhanced production of reactive oxygen metabolites, enzyme secretion, and adhesion to endothelial cells. Atherosclerosis, 1998. 136(2): p. 315-24. 36. Woenckhaus, C., et al., Hypochlorite-modified LDL: chemotactic potential and chemokine induction in human monocytes. Clin Immunol Immunopathol, 1998. 86(1): p. 27-33. 37. Byun, J., et al., Nitrogen dioxide radical generated by the myeloperoxidase- hydrogen peroxide-nitrite system promotes lipid peroxidation of low density lipoprotein. FEBS Lett, 1999. 455(3): p. 243-6. 38. Kostyuk, V.A., et al., Myeloperoxidase/nitrite-mediated lipid peroxidation of low- density lipoprotein as modulated by flavonoids. FEBS Lett, 2003. 537(1-3): p. 146-50. 156  39. Podrez, E.A., et al., Myeloperoxidase-generated reactive nitrogen species convert LDL into an atherogenic form in vitro. J Clin Invest, 1999. 103(11): p. 1547-60. 40. Daugherty, A., et al., Myeloperoxidase, a catalyst for lipoprotein oxidation, is expressed in human atherosclerotic lesions. J Clin Invest, 1994. 94(1): p. 437-44. 41. Hazell, L.J., et al., Presence of hypochlorite-modified proteins in human atherosclerotic lesions. J Clin Invest, 1996. 97(6): p. 1535-44. 42. Malle, E., et al., Immunohistochemical evidence for the myeloperoxidase/H2O2/halide system in human atherosclerotic lesions: colocalization of myeloperoxidase and hypochlorite-modified proteins. Eur J Biochem, 2000. 267(14): p. 4495-503. 43. Sugiyama, S., et al., Macrophage myeloperoxidase regulation by granulocyte macrophage colony-stimulating factor in human atherosclerosis and implications in acute coronary syndromes. Am J Pathol, 2001. 158(3): p. 879-91. 44. Brasen, J.H., et al., Patterns of oxidized epitopes, but not NF-kappa B expression, change during atherogenesis in WHHL rabbits. Atherosclerosis, 2003. 166(1): p. 13-21. 45. Malle, E., et al., Hypochlorite-modified (lipo)proteins are present in rabbit lesions in response to dietary cholesterol. Biochem Biophys Res Commun, 2001. 289(4): p. 894-900. 46. Romano, M. and J. Claria, Cyclooxygenase-2 and 5-lipoxygenase converging functions on cell proliferation and tumor angiogenesis: implications for cancer therapy. Faseb J, 2003. 17(14): p. 1986-95. 157  47. Schewe, T., 15-lipoxygenase-1: a prooxidant enzyme. Biol Chem, 2002. 383(3-4): p. 365-74. 48. Belkner, J., et al., The oxygenation of cholesterol esters by the reticulocyte lipoxygenase. FEBS Lett, 1991. 279(1): p. 110-4. 49. Brash, A.R., Lipoxygenases: occurrence, functions, catalysis, and acquisition of substrate. J Biol Chem, 1999. 274(34): p. 23679-82. 50. Murray, J.J. and A.R. Brash, Rabbit reticulocyte lipoxygenase catalyzes specific 12(S) and 15(S) oxygenation of arachidonoyl-phosphatidylcholine. Arch Biochem Biophys, 1988. 265(2): p. 514-23. 51. Belkner, J., et al., Oxygenation of lipoproteins by mammalian lipoxygenases. Eur J Biochem, 1993. 213(1): p. 251-61. 52. Heydeck, D., et al., Oxidation of LDL by rabbit and human 15-lipoxygenase: prevalence of nonenzymatic reactions. J Lipid Res, 2001. 42(7): p. 1082-8. 53. Upston, J.M., et al., Oxidation of free fatty acids in low density lipoprotein by 15- lipoxygenase stimulates nonenzymic, alpha-tocopherol-mediated peroxidation of cholesteryl esters. J Biol Chem, 1997. 272(48): p. 30067-74. 54. Yamashita, H., et al., Oxidation of low density lipoprotein and plasma by 15- lipoxygenase and free radicals. FEBS Lett, 1999. 445(2-3): p. 287-90. 55. Yla-Herttuala, S., et al., Colocalization of 15-lipoxygenase mRNA and protein with epitopes of oxidized low density lipoprotein in macrophage-rich areas of atherosclerotic lesions. Proc Natl Acad Sci U S A, 1990. 87(18): p. 6959-63. 158  56. Mehrabian, M., et al., Identification of 5-lipoxygenase as a major gene contributing to atherosclerosis susceptibility in mice. Circ Res, 2002. 91(2): p. 120-6. 57. Cyrus, T., et al., Disruption of the 12/15-lipoxygenase gene diminishes atherosclerosis in apo E-deficient mice. J Clin Invest, 1999. 103(11): p. 1597-604. 58. George, J., et al., 12/15-Lipoxygenase gene disruption attenuates atherogenesis in LDL receptor-deficient mice. Circulation, 2001. 104(14): p. 1646-50. 59. Sendobry, S.M., et al., Attenuation of diet-induced atherosclerosis in rabbits with a highly selective 15-lipoxygenase inhibitor lacking significant antioxidant properties. Br J Pharmacol, 1997. 120(7): p. 1199-206. 60. Parthasarathy, S., E. Wieland, and D. Steinberg, A role for endothelial cell lipoxygenase in the oxidative modification of low density lipoprotein. Proc Natl Acad Sci U S A, 1989. 86(3): p. 1046-50. 61. Haberland, M.E., C.L. Olch, and A.M. Folgelman, Role of lysines in mediating interaction of modified low density lipoproteins with the scavenger receptor of human monocyte macrophages. J Biol Chem, 1984. 259(18): p. 11305-11. 62. Parthasarathy, S., et al., Macrophage oxidation of low density lipoprotein generates a modified form recognized by the scavenger receptor. Arteriosclerosis, 1986. 6(5): p. 505-10. 63. Steinbrecher, U.P., et al., Recognition of oxidized low density lipoprotein by the scavenger receptor of macrophages results from derivatization of apolipoprotein 159  B by products of fatty acid peroxidation. J Biol Chem, 1989. 264(26): p. 15216- 23. 64. Lamb, D.J., M.J. Mitchinson, and D.S. Leake, Transition metal ions within human atherosclerotic lesions can catalyse the oxidation of low density lipoprotein by macrophages. FEBS Lett, 1995. 374(1): p. 12-6. 65. Smith, C., et al., Stimulation of lipid peroxidation and hydroxyl-radical generation by the contents of human atherosclerotic lesions. Biochem J, 1992. 286 ( Pt 3): p. 901-5. 66. Stadler, N., R.A. Lindner, and M.J. Davies, Direct detection and quantification of transition metal ions in human atherosclerotic plaques: evidence for the presence of elevated levels of iron and copper. Arterioscler Thromb Vasc Biol, 2004. 24(5): p. 949-54. 67. Berliner, J.A., et al., Minimally modified low density lipoprotein stimulates monocyte endothelial interactions. J Clin Invest, 1990. 85(4): p. 1260-6. 68. Cushing, S.D., et al., Minimally modified low density lipoprotein induces monocyte chemotactic protein 1 in human endothelial cells and smooth muscle cells. Proc Natl Acad Sci U S A, 1990. 87(13): p. 5134-8. 69. Navab, M., et al., Monocyte transmigration induced by modification of low density lipoprotein in cocultures of human aortic wall cells is due to induction of monocyte chemotactic protein 1 synthesis and is abolished by high density lipoprotein. J Clin Invest, 1991. 88(6): p. 2039-46. 160  70. Rajavashisth, T.B., et al., Induction of endothelial cell expression of granulocyte and macrophage colony-stimulating factors by modified low-density lipoproteins. Nature, 1990. 344(6263): p. 254-7. 71. Kume, N., M.I. Cybulsky, and M.A. Gimbrone, Jr., Lysophosphatidylcholine, a component of atherogenic lipoproteins, induces mononuclear leukocyte adhesion molecules in cultured human and rabbit arterial endothelial cells. J Clin Invest, 1992. 90(3): p. 1138-44. 72. Eriksson, E.E., et al., Importance of primary capture and L-selectin-dependent secondary capture in leukocyte accumulation in inflammation and atherosclerosis in vivo. J Exp Med, 2001. 194(2): p. 205-18. 73. Massberg, S., et al., A critical role of platelet adhesion in the initiation of atherosclerotic lesion formation. J Exp Med, 2002. 196(7): p. 887-96. 74. McMurray, H.F., S. Parthasarathy, and D. Steinberg, Oxidatively modified low density lipoprotein is a chemoattractant for human T lymphocytes. J Clin Invest, 1993. 92(2): p. 1004-8. 75. Quinn, M.T., et al., Oxidatively modified low density lipoproteins: a potential role in recruitment and retention of monocyte/macrophages during atherogenesis. Proc Natl Acad Sci U S A, 1987. 84(9): p. 2995-8. 76. Yla-Herttuala, S., et al., Expression of monocyte chemoattractant protein 1 in macrophage-rich areas of human and rabbit atherosclerotic lesions. Proc Natl Acad Sci U S A, 1991. 88(12): p. 5252-6. 161  77. Gosling, J., et al., MCP-1 deficiency reduces susceptibility to atherosclerosis in mice that overexpress human apolipoprotein B. J Clin Invest, 1999. 103(6): p. 773-8. 78. Gu, L., et al., Absence of monocyte chemoattractant protein-1 reduces atherosclerosis in low density lipoprotein receptor-deficient mice. Mol Cell, 1998. 2(2): p. 275-81. 79. Boring, L., et al., Decreased lesion formation in CCR2-/- mice reveals a role for chemokines in the initiation of atherosclerosis. Nature, 1998. 394(6696): p. 894-7. 80. Libby, P., Inflammation in atherosclerosis. Nature, 2002. 420(6917): p. 868-74. 81. Hansson, G.K., Inflammation, atherosclerosis, and coronary artery disease. N Engl J Med, 2005. 352(16): p. 1685-95. 82. Hansson, G.K., A.K. Robertson, and C. Soderberg-Naucler, Inflammation and atherosclerosis. Annu Rev Pathol, 2006. 1: p. 297-329. 83. Rosenfeld, M.E., et al., Macrophage colony-stimulating factor mRNA and protein in atherosclerotic lesions of rabbits and humans. Am J Pathol, 1992. 140(2): p. 291-300. 84. Clinton, S.K., et al., Macrophage colony-stimulating factor gene expression in vascular cells and in experimental and human atherosclerosis. Am J Pathol, 1992. 140(2): p. 301-16. 85. de Villiers, W.J., et al., Macrophage-colony-stimulating factor selectively enhances macrophage scavenger receptor expression and function. J Exp Med, 1994. 180(2): p. 705-9. 162  86. Babamusta, F., et al., Angiotensin II infusion induces site-specific intra-laminar hemorrhage in macrophage colony-stimulating factor-deficient mice. Atherosclerosis, 2006. 186(2): p. 282-90. 87. Rajavashisth, T., et al., Heterozygous osteopetrotic (op) mutation reduces atherosclerosis in LDL receptor- deficient mice. J Clin Invest, 1998. 101(12): p. 2702-10. 88. Barlic, J., et al., Oxidized lipid-driven chemokine receptor switch, CCR2 to CX3CR1, mediates adhesion of human macrophages to coronary artery smooth muscle cells through a peroxisome proliferator-activated receptor gamma- dependent pathway. Circulation, 2006. 114(8): p. 807-19. 89. Frostegard, J., et al., Oxidized low density lipoprotein induces differentiation and adhesion of human monocytes and the monocytic cell line U937. Proc Natl Acad Sci U S A, 1990. 87(3): p. 904-8. 90. Hayden, J.M., et al., Induction of monocyte differentiation and foam cell formation in vitro by 7-ketocholesterol. J Lipid Res, 2002. 43(1): p. 26-35. 91. Fuhrman, B., et al., Ox-LDL induces monocyte-to-macrophage differentiation in vivo: Possible role for the macrophage colony stimulating factor receptor (M- CSF-R). Atherosclerosis, 2008. 196(2): p. 598-607. 92. Smith, J.D., et al., Decreased atherosclerosis in mice deficient in both macrophage colony-stimulating factor (op) and apolipoprotein E. Proc Natl Acad Sci U S A, 1995. 92(18): p. 8264-8. 163  93. Brown, M.S., et al., Reversible accumulation of cholesteryl esters in macrophages incubated with acetylated lipoproteins. J Cell Biol, 1979. 82(3): p. 597-613. 94. Yla-Herttuala, S., et al., Evidence for the presence of oxidatively modified low density lipoprotein in atherosclerotic lesions of rabbit and man. J Clin Invest, 1989. 84(4): p. 1086-95. 95. Rosenfeld, M.E., et al., Macrophage-derived foam cells freshly isolated from rabbit atherosclerotic lesions degrade modified lipoproteins, promote oxidation of low-density lipoproteins, and contain oxidation-specific lipid-protein adducts. J Clin Invest, 1991. 87(1): p. 90-9. 96. Jessup, W., E.L. Mander, and R.T. Dean, The intracellular storage and turnover of apolipoprotein B of oxidized LDL in macrophages. Biochim Biophys Acta, 1992. 1126(2): p. 167-77. 97. Yancey, P.G. and W.G. Jerome, Lysosomal cholesterol derived from mildly oxidized low density lipoprotein is resistant to efflux. J Lipid Res, 2001. 42(3): p. 317-27. 98. Jialal, I. and A. Chait, Differences in the metabolism of oxidatively modified low density lipoprotein and acetylated low density lipoprotein by human endothelial cells: inhibition of cholesterol esterification by oxidatively modified low density lipoprotein. J Lipid Res, 1989. 30(10): p. 1561-8. 99. Roma, P., et al., Oxidized LDL increase free cholesterol and fail to stimulate cholesterol esterification in murine macrophages. Biochem Biophys Res Commun, 1990. 171(1): p. 123-31. 164  100. Lougheed, M., H.F. Zhang, and U.P. Steinbrecher, Oxidized low density lipoprotein is resistant to cathepsins and accumulates within macrophages. J Biol Chem, 1991. 266(22): p. 14519-25. 101. Hoppe, G., J. O'Neil, and H.F. Hoff, Inactivation of lysosomal proteases by oxidized low density lipoprotein is partially responsible for its poor degradation by mouse peritoneal macrophages. J Clin Invest, 1994. 94(4): p. 1506-12. 102. Kritharides, L., et al., Cholesterol metabolism and efflux in human THP-1 macrophages. Arterioscler Thromb Vasc Biol, 1998. 18(10): p. 1589-99. 103. Edfeldt, K., et al., Expression of toll-like receptors in human atherosclerotic lesions: a possible pathway for plaque activation. Circulation, 2002. 105(10): p. 1158-61. 104. Janeway, C.A., Jr. and R. Medzhitov, Innate immune recognition. Annu Rev Immunol, 2002. 20: p. 197-216. 105. Xu, X.H., et al., Toll-like receptor-4 is expressed by macrophages in murine and human lipid-rich atherosclerotic plaques and upregulated by oxidized LDL. Circulation, 2001. 104(25): p. 3103-8. 106. Michelsen, K.S., et al., Lack of Toll-like receptor 4 or myeloid differentiation factor 88 reduces atherosclerosis and alters plaque phenotype in mice deficient in apolipoprotein E. Proc Natl Acad Sci U S A, 2004. 101(29): p. 10679-84. 107. Bjorkbacka, H., et al., Reduced atherosclerosis in MyD88-null mice links elevated serum cholesterol levels to activation of innate immunity signaling pathways. Nat Med, 2004. 10(4): p. 416-21. 165  108. Faure, E., et al., Bacterial lipopolysaccharide and IFN-gamma induce Toll-like receptor 2 and Toll-like receptor 4 expression in human endothelial cells: role of NF-kappa B activation. J Immunol, 2001. 166(3): p. 2018-24. 109. Miller, Y.I., et al., Minimally modified LDL binds to CD14, induces macrophage spreading via TLR4/MD-2, and inhibits phagocytosis of apoptotic cells. J Biol Chem, 2003. 278(3): p. 1561-8. 110. Zeiher, A.M., et al., Tissue endothelin-1 immunoreactivity in the active coronary atherosclerotic plaque. A clue to the mechanism of increased vasoreactivity of the culprit lesion in unstable angina. Circulation, 1995. 91(4): p. 941-7. 111. Buttery, L.D., et al., Inducible nitric oxide synthase is present within human atherosclerotic lesions and promotes the formation and activity of peroxynitrite. Lab Invest, 1996. 75(1): p. 77-85. 112. Wuttge, D.M., et al., Expression of interleukin-15 in mouse and human atherosclerotic lesions. Am J Pathol, 2001. 159(2): p. 417-23. 113. Shah, P.K., et al., Human monocyte-derived macrophages induce collagen breakdown in fibrous caps of atherosclerotic plaques. Potential role of matrix- degrading metalloproteinases and implications for plaque rupture. Circulation, 1995. 92(6): p. 1565-9. 114. Jormsjo, S., et al., Differential expression of cysteine and aspartic proteases during progression of atherosclerosis in apolipoprotein E-deficient mice. Am J Pathol, 2002. 161(3): p. 939-45. 166  115. Libby, P. and M. Aikawa, Stabilization of atherosclerotic plaques: new mechanisms and clinical targets. Nat Med, 2002. 8(11): p. 1257-62. 116. Liu, J., et al., Lysosomal cysteine proteases in atherosclerosis. Arterioscler Thromb Vasc Biol, 2004. 24(8): p. 1359-66. 117. Galis, Z.S., et al., Increased expression of matrix metalloproteinases and matrix degrading activity in vulnerable regions of human atherosclerotic plaques. J Clin Invest, 1994. 94(6): p. 2493-503. 118. Choudhary, S., et al., Quantitation and localization of matrix metalloproteinases and their inhibitors in human carotid endarterectomy tissues. Arterioscler Thromb Vasc Biol, 2006. 26(10): p. 2351-8. 119. Sukhova, G.K., et al., Evidence for increased collagenolysis by interstitial collagenases-1 and -3 in vulnerable human atheromatous plaques. Circulation, 1999. 99(19): p. 2503-9. 120. Jonasson, L., et al., Regional accumulations of T cells, macrophages, and smooth muscle cells in the human atherosclerotic plaque. Arteriosclerosis, 1986. 6(2): p. 131-8. 121. Kovanen, P.T., M. Kaartinen, and T. Paavonen, Infiltrates of activated mast cells at the site of coronary atheromatous erosion or rupture in myocardial infarction. Circulation, 1995. 92(5): p. 1084-8. 122. Lusis, A.J., Atherosclerosis. Nature, 2000. 407(6801): p. 233-41. 123. Libby, P., et al., Macrophages and atherosclerotic plaque stability. Curr Opin Lipidol, 1996. 7(5): p. 330-5. 167  124. van der Wal, A.C., et al., Site of intimal rupture or erosion of thrombosed coronary atherosclerotic plaques is characterized by an inflammatory process irrespective of the dominant plaque morphology. Circulation, 1994. 89(1): p. 36- 44. 125. Kunte, H., et al., Markers of instability in high-risk carotid plaques are reduced by statins. J Vasc Surg, 2008. 47(3): p. 513-22. 126. Tabas, I., Consequences and therapeutic implications of macrophage apoptosis in atherosclerosis: the importance of lesion stage and phagocytic efficiency. Arterioscler Thromb Vasc Biol, 2005. 25(11): p. 2255-64. 127. Geng, Y.J. and P. Libby, Evidence for apoptosis in advanced human atheroma. Colocalization with interleukin-1 beta-converting enzyme. Am J Pathol, 1995. 147(2): p. 251-66. 128. Clarke, M.C., et al., Apoptosis of vascular smooth muscle cells induces features of plaque vulnerability in atherosclerosis. Nat Med, 2006. 12(9): p. 1075-80. 129. Schrijvers, D.M., et al., Phagocytosis of apoptotic cells by macrophages is impaired in atherosclerosis. Arterioscler Thromb Vasc Biol, 2005. 25(6): p. 1256- 61. 130. Kockx, M.M., Apoptosis in the atherosclerotic plaque: quantitative and qualitative aspects. Arterioscler Thromb Vasc Biol, 1998. 18(10): p. 1519-22. 131. Kockx, M.M. and A.G. Herman, Apoptosis in atherosclerosis: beneficial or detrimental? Cardiovasc Res, 2000. 45(3): p. 736-46. 168  132. Steinberg, D., Lewis A. Conner Memorial Lecture. Oxidative modification of LDL and atherogenesis. Circulation, 1997. 95(4): p. 1062-71. 133. Katsuda, S., et al., Human atherosclerosis. IV. Immunocytochemical analysis of cell activation and proliferation in lesions of young adults. Am J Pathol, 1993. 142(6): p. 1787-93. 134. Rekhter, M.D. and D. Gordon, Active proliferation of different cell types, including lymphocytes, in human atherosclerotic plaques. Am J Pathol, 1995. 147(3): p. 668-77. 135. Boesten, L.S., et al., Macrophage retinoblastoma deficiency leads to enhanced atherosclerosis development in ApoE-deficient mice. Faseb J, 2006. 20(7): p. 953- 5. 136. Senokuchi, T., et al., Statins suppress oxidized low density lipoprotein-induced macrophage proliferation by inactivation of the small G protein-p38 MAPK pathway. J Biol Chem, 2005. 280(8): p. 6627-33. 137. Aikawa, M., et al., An HMG-CoA reductase inhibitor, cerivastatin, suppresses growth of macrophages expressing matrix metalloproteinases and tissue factor in vivo and in vitro. Circulation, 2001. 103(2): p. 276-83. 138. Liu, J., et al., Reduced macrophage apoptosis is associated with accelerated atherosclerosis in low-density lipoprotein receptor-null mice. Arterioscler Thromb Vasc Biol, 2005. 25(1): p. 174-9. 139. Yui, S., et al., Induction of murine macrophage growth by modified LDLs. Arterioscler Thromb, 1993. 13(3): p. 331-7. 169  140. Sakai, M., et al., Lysophosphatidylcholine plays an essential role in the mitogenic effect of oxidized low density lipoprotein on murine macrophages. J Biol Chem, 1994. 269(50): p. 31430-5. 141. Sakai, M., et al., The scavenger receptor serves as a route for internalization of lysophosphatidylcholine in oxidized low density lipoprotein-induced macrophage proliferation. J Biol Chem, 1996. 271(44): p. 27346-52. 142. Sakai, M., et al., Lysophosphatidylcholine potentiates the mitogenic activity of modified LDL for human monocyte-derived macrophages. Arterioscler Thromb Vasc Biol, 1996. 16(4): p. 600-5. 143. Matsumura, T., et al., Two intracellular signaling pathways for activation of protein kinase C are involved in oxidized low-density lipoprotein-induced macrophage growth. Arterioscler Thromb Vasc Biol, 1997. 17(11): p. 3013-20. 144. Matsumura, T., et al., Cis-acting DNA elements of mouse granulocyte/macrophage colony-stimulating factor gene responsive to oxidized low density lipoprotein. J Biol Chem, 1999. 274(53): p. 37665-72. 145. Biwa, T., et al., Induction of murine macrophage growth by oxidized low density lipoprotein is mediated by granulocyte macrophage colony-stimulating factor. J Biol Chem, 1998. 273(43): p. 28305-13. 146. Martens, J.S., et al., A modification of apolipoprotein B accounts for most of the induction of macrophage growth by oxidized low density lipoprotein. J Biol Chem, 1999. 274(16): p. 10903-10. 170  147. Martens, J.S., et al., Phosphatidylinositol 3-kinase is involved in the induction of macrophage growth by oxidized low density lipoprotein. J Biol Chem, 1998. 273(9): p. 4915-20. 148. Biwa, T., et al., Sites of action of protein kinase C and phosphatidylinositol 3- kinase are distinct in oxidized low density lipoprotein-induced macrophage proliferation. J Biol Chem, 2000. 275(8): p. 5810-6. 149. Hamilton, J.A., et al., Oxidized LDL can induce macrophage survival, DNA synthesis, and enhanced proliferative response to CSF-1 and GM-CSF. Arterioscler Thromb Vasc Biol, 1999. 19(1): p. 98-105. 150. Hamilton, J.A., et al., Comparison of macrophage responses to oxidized low- density lipoprotein and macrophage colony-stimulating factor (M-CSF or CSF- 1). Biochem J, 2001. 354(Pt 1): p. 179-87. 151. Hundal, R.S., et al., Oxidized low density lipoprotein inhibits macrophage apoptosis by blocking ceramide generation, thereby maintaining protein kinase B activation and Bcl-XL levels. J Biol Chem, 2003. 278(27): p. 24399-408. 152. Hundal, R.S., et al., Oxidized low density lipoprotein inhibits macrophage apoptosis through activation of the PI 3-kinase/PKB pathway. J Lipid Res, 2001. 42(9): p. 1483-91. 153. Arai, S., et al., A role for the apoptosis inhibitory factor AIM/Spalpha/Api6 in atherosclerosis development. Cell Metab, 2005. 1(3): p. 201-13. 154. Downward, J., PI 3-kinase, Akt and cell survival. Semin Cell Dev Biol, 2004. 15(2): p. 177-82. 171  155. Colles, S.M., et al., Oxidized LDL-induced injury and apoptosis in atherosclerosis. Potential roles for oxysterols. Trends Cardiovasc Med, 2001. 11(3-4): p. 131-8. 156. Reid, V.C., M.J. Mitchinson, and J.N. Skepper, Cytotoxicity of oxidized low- density lipoprotein to mouse peritoneal macrophages: an ultrastructural study. J Pathol, 1993. 171(4): p. 321-8. 157. Martinet, W. and M.M. Kockx, Apoptosis in atherosclerosis: focus on oxidized lipids and inflammation. Curr Opin Lipidol, 2001. 12(5): p. 535-41. 158. Morel, D.W., J.R. Hessler, and G.M. Chisolm, Low density lipoprotein cytotoxicity induced by free radical peroxidation of lipid. J Lipid Res, 1983. 24(8): p. 1070-6. 159. Chisolm, G.M., et al., 7 beta-hydroperoxycholest-5-en-3 beta-ol, a component of human atherosclerotic lesions, is the primary cytotoxin of oxidized human low density lipoprotein. Proc Natl Acad Sci U S A, 1994. 91(24): p. 11452-6. 160. Sata, M. and K. Walsh, Endothelial cell apoptosis induced by oxidized LDL is associated with the down-regulation of the cellular caspase inhibitor FLIP. J Biol Chem, 1998. 273(50): p. 33103-6. 161. Escargueil, I., et al., Oxidized low density lipoproteins elicit DNA fragmentation of cultured lymphoblastoid cells. FEBS Lett, 1992. 305(2): p. 155-9. 162. Kosugi, K., et al., Toxicity of oxidized low-density lipoprotein to cultured fibroblasts is selective for S phase of the cell cycle. J Cell Physiol, 1987. 130(3): p. 311-20. 172  163. Mabile, L., et al., alpha-Tocopherol and trolox block the early intracellular events (TBARS and calcium rises) elicited by oxidized low density lipoproteins in cultured endothelial cells. Free Radic Biol Med, 1995. 19(2): p. 177-87. 164. Hughes, H., et al., Cytotoxicity of oxidized LDL to porcine aortic smooth muscle cells is associated with the oxysterols 7-ketocholesterol and 7-hydroxycholesterol. Arterioscler Thromb, 1994. 14(7): p. 1177-85. 165. Marchant, C.E., et al., Oxidation of low-density lipoprotein by human monocyte- macrophages results in toxicity to the oxidising culture. Free Radic Res, 1996. 24(5): p. 333-42. 166. Li, W., et al., Uptake of oxidized LDL by macrophages results in partial lysosomal enzyme inactivation and relocation. Arterioscler Thromb Vasc Biol, 1998. 18(2): p. 177-84. 167. Meilhac, O., et al., Bcl-2 alters the balance between apoptosis and necrosis, but does not prevent cell death induced by oxidized low density lipoproteins. Faseb J, 1999. 13(3): p. 485-94. 168. Kuzuya, M., et al., VEGF protects against oxidized LDL toxicity to endothelial cells by an intracellular glutathione-dependent mechanism through the KDR receptor. Arterioscler Thromb Vasc Biol, 2001. 21(5): p. 765-70. 169. Asmis, R. and J.G. Begley, Oxidized LDL promotes peroxide-mediated mitochondrial dysfunction and cell death in human macrophages: a caspase-3- independent pathway. Circ Res, 2003. 92(1): p. e20-9. 173  170. Nhan, T.Q., et al., The p17 cleaved form of caspase-3 is present within viable macrophages in vitro and in atherosclerotic plaque. Arterioscler Thromb Vasc Biol, 2003. 23(7): p. 1276-82. 171. Hamilton, J.A., et al., Enhancement of macrophage survival and DNA synthesis by oxidized-low-density-lipoprotein (LDL)-derived lipids and by aggregates of lightly oxidized LDL. Biochem J, 2001. 355(Pt 1): p. 207-14. 172. Hamilton, J.A., G. Whitty, and W. Jessup, Oxidized LDL can promote human monocyte survival. Arterioscler Thromb Vasc Biol, 2000. 20(10): p. 2329-31. 173. Munteanu, A., et al., Antagonistic effects of oxidized low density lipoprotein and alpha-tocopherol on CD36 scavenger receptor expression in monocytes: involvement of protein kinase B and peroxisome proliferator-activated receptor- gamma. J Biol Chem, 2006. 281(10): p. 6489-97. 174. Namgaladze, D., A. Kollas, and B. Brune, Oxidized LDL attenuates apoptosis in monocytic cells by activating ERK signaling. J Lipid Res, 2008. 49(1): p. 58-65. 175. Auge, N., et al., Oxidized LDL-induced smooth muscle cell proliferation involves the EGF receptor/PI-3 kinase/Akt and the sphingolipid signaling pathways. Arterioscler Thromb Vasc Biol, 2002. 22(12): p. 1990-5. 176. Han, C.Y. and Y.K. Pak, Oxidation-dependent effects of oxidized LDL: proliferation or cell death. Exp Mol Med, 1999. 31(4): p. 165-73. 177. Steinbrecher, U.P., A. Gomez-Munoz, and V. Duronio, Acid sphingomyelinase in macrophage apoptosis. Curr Opin Lipidol, 2004. 15(5): p. 531-7. 174  178. Merrill, A.H., Jr., et al., Sphingolipids--the enigmatic lipid class: biochemistry, physiology, and pathophysiology. Toxicol Appl Pharmacol, 1997. 142(1): p. 208- 25. 179. Hannun, Y.A., The sphingomyelin cycle and the second messenger function of ceramide. J Biol Chem, 1994. 269(5): p. 3125-8. 180. Kee, T.H., P. Vit, and A.J. Melendez, Sphingosine kinase signalling in immune cells. Clin Exp Pharmacol Physiol, 2005. 32(3): p. 153-61. 181. Baumruker, T. and E.E. Prieschl, Sphingolipids and the regulation of the immune response. Semin Immunol, 2002. 14(1): p. 57-63. 182. Spiegel, S. and S. Milstien, Sphingosine 1-phosphate, a key cell signaling molecule. J Biol Chem, 2002. 277(29): p. 25851-4. 183. Spiegel, S., Sphingosine 1-phosphate: a prototype of a new class of second messengers. J Leukoc Biol, 1999. 65(3): p. 341-4. 184. Van Brocklyn, J.R., et al., Dual actions of sphingosine-1-phosphate: extracellular through the Gi-coupled receptor Edg-1 and intracellular to regulate proliferation and survival. J Cell Biol, 1998. 142(1): p. 229-40. 185. Hannun, Y.A., C. Luberto, and K.M. Argraves, Enzymes of sphingolipid metabolism: from modular to integrative signaling. Biochemistry, 2001. 40(16): p. 4893-903. 186. Taha, T.A., Y.A. Hannun, and L.M. Obeid, Sphingosine kinase: biochemical and cellular regulation and role in disease. J Biochem Mol Biol, 2006. 39(2): p. 113- 31. 175  187. Meyer zu Heringdorf, D., et al., Role of sphingosine kinase in Ca(2+) signalling by epidermal growth factor receptor. FEBS Lett, 1999. 461(3): p. 217-22. 188. Alemany, R., et al., Formyl peptide receptor signaling in HL-60 cells through sphingosine kinase. J Biol Chem, 1999. 274(7): p. 3994-9. 189. Melendez, A.J. and A.K. Khaw, Dichotomy of Ca2+ signals triggered by different phospholipid pathways in antigen stimulation of human mast cells. J Biol Chem, 2002. 277(19): p. 17255-62. 190. Choi, O.H., J.H. Kim, and J.P. Kinet, Calcium mobilization via sphingosine kinase in signalling by the Fc epsilon RI antigen receptor. Nature, 1996. 380(6575): p. 634-6. 191. Melendez, A., et al., FcgammaRI coupling to phospholipase D initiates sphingosine kinase-mediated calcium mobilization and vesicular trafficking. J Biol Chem, 1998. 273(16): p. 9393-402. 192. English, D., et al., Sphingosine 1-phosphate released from platelets during clotting accounts for the potent endothelial cell chemotactic activity of blood serum and provides a novel link between hemostasis and angiogenesis. Faseb J, 2000. 14(14): p. 2255-65. 193. Wang, F., et al., Sphingosine 1-phosphate stimulates cell migration through a G(i)-coupled cell surface receptor. Potential involvement in angiogenesis. J Biol Chem, 1999. 274(50): p. 35343-50. 176  194. Shu, X., et al., Sphingosine kinase mediates vascular endothelial growth factor- induced activation of ras and mitogen-activated protein kinases. Mol Cell Biol, 2002. 22(22): p. 7758-68. 195. Spiegel, S. and A.H. Merrill, Jr., Sphingolipid metabolism and cell growth regulation. Faseb J, 1996. 10(12): p. 1388-97. 196. Cuvillier, O., Sphingosine in apoptosis signaling. Biochim Biophys Acta, 2002. 1585(2-3): p. 153-62. 197. Maceyka, M., et al., Sphingosine kinase, sphingosine-1-phosphate, and apoptosis. Biochim Biophys Acta, 2002. 1585(2-3): p. 193-201. 198. Olivera, A., et al., Sphingosine kinase expression increases intracellular sphingosine-1-phosphate and promotes cell growth and survival. J Cell Biol, 1999. 147(3): p. 545-58. 199. Olivera, A. and S. Spiegel, Sphingosine-1-phosphate as second messenger in cell proliferation induced by PDGF and FCS mitogens. Nature, 1993. 365(6446): p. 557-60. 200. Edsall, L.C., G.G. Pirianov, and S. Spiegel, Involvement of sphingosine 1- phosphate in nerve growth factor-mediated neuronal survival and differentiation. J Neurosci, 1997. 17(18): p. 6952-60. 201. Xu, C.B., et al., D-erythro-N,N-dimethylsphingosine inhibits bFGF-induced proliferation of cerebral, aortic and coronary smooth muscle cells. Atherosclerosis, 2002. 164(2): p. 237-43. 177  202. Pitson, S.M., et al., Phosphorylation-dependent translocation of sphingosine kinase to the plasma membrane drives its oncogenic signalling. J Exp Med, 2005. 201(1): p. 49-54. 203. Melendez, A.J. and F.B. Ibrahim, Antisense knockdown of sphingosine kinase 1 in human macrophages inhibits C5a receptor-dependent signal transduction, Ca2+ signals, enzyme release, cytokine production, and chemotaxis. J Immunol, 2004. 173(3): p. 1596-603. 204. Pitson, S.M., et al., Human sphingosine kinase: purification, molecular cloning and characterization of the native and recombinant enzymes. Biochem J, 2000. 350 Pt 2: p. 429-41. 205. Olivera, A., et al., Platelet-derived growth factor-induced activation of sphingosine kinase requires phosphorylation of the PDGF receptor tyrosine residue responsible for binding of PLCgamma. Faseb J, 1999. 13(12): p. 1593- 600. 206. Sukocheva, O., et al., Estrogen transactivates EGFR via the sphingosine 1- phosphate receptor Edg-3: the role of sphingosine kinase-1. J Cell Biol, 2006. 173(2): p. 301-10. 207. Sukocheva, O.A., et al., Sphingosine kinase transmits estrogen signaling in human breast cancer cells. Mol Endocrinol, 2003. 17(10): p. 2002-12. 208. Alvarez, S.E., S. Milstien, and S. Spiegel, Autocrine and paracrine roles of sphingosine-1-phosphate. Trends Endocrinol Metab, 2007. 18(8): p. 300-7. 178  209. Meyer Zu Heringdorf, D., Lysophospholipid receptor-dependent and - independent calcium signaling. J Cell Biochem, 2004. 92(5): p. 937-48. 210. Meyer zu Heringdorf, D., et al., Photolysis of intracellular caged sphingosine-1- phosphate causes Ca2+ mobilization independently of G-protein-coupled receptors. FEBS Lett, 2003. 554(3): p. 443-9. 211. Mao, C., et al., Molecular cloning and characterization of SCaMPER, a sphingolipid Ca2+ release-mediating protein from endoplasmic reticulum. Proc Natl Acad Sci U S A, 1996. 93(5): p. 1993-6. 212. Cavalli, A.L., et al., Expression and functional characterization of SCaMPER: a sphingolipid-modulated calcium channel of cardiomyocytes. Am J Physiol Cell Physiol, 2003. 284(3): p. C780-90. 213. Schnurbus, R., et al., Re-evaluation of primary structure, topology, and localization of Scamper, a putative intracellular Ca2+ channel activated by sphingosylphosphocholine. Biochem J, 2002. 362(Pt 1): p. 183-9. 214. Auge, N., et al., Role of sphingosine 1-phosphate in the mitogenesis induced by oxidized low density lipoprotein in smooth muscle cells via activation of sphingomyelinase, ceramidase, and sphingosine kinase. J Biol Chem, 1999. 274(31): p. 21533-8. 215. Xu, C.B., J. Hansen-Schwartz, and L. Edvinsson, Sphingosine signaling and atherogenesis. Acta Pharmacol Sin, 2004. 25(7): p. 849-54. 179  216. Murata, N., et al., Interaction of sphingosine 1-phosphate with plasma components, including lipoproteins, regulates the lipid receptor-mediated actions. Biochem J, 2000. 352 Pt 3: p. 809-15. 217. Kimura, T., et al., Sphingosine 1-phosphate may be a major component of plasma lipoproteins responsible for the cytoprotective actions in human umbilical vein endothelial cells. J Biol Chem, 2001. 276(34): p. 31780-5. 218. Deutschman, D.H., et al., Predicting obstructive coronary artery disease with serum sphingosine-1-phosphate. Am Heart J, 2003. 146(1): p. 62-8. 219. Berridge, M.J., M.D. Bootman, and H.L. Roderick, Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol, 2003. 4(7): p. 517-29. 220. Berridge, M.J., P. Lipp, and M.D. Bootman, The versatility and universality of calcium signalling. Nat Rev Mol Cell Biol, 2000. 1(1): p. 11-21. 221. Mikoshiba, K., IP3 receptor/Ca2+ channel: from discovery to new signaling concepts. J Neurochem, 2007. 102(5): p. 1426-46. 222. Guatimosim, S., et al., Local Ca(2+) signaling and EC coupling in heart: Ca(2+) sparks and the regulation of the [Ca(2+)](i) transient. J Mol Cell Cardiol, 2002. 34(8): p. 941-50. 223. De Koninck, P. and H. Schulman, Sensitivity of CaM kinase II to the frequency of Ca2+ oscillations. Science, 1998. 279(5348): p. 227-30. 224. Oancea, E. and T. Meyer, Protein kinase C as a molecular machine for decoding calcium and diacylglycerol signals. Cell, 1998. 95(3): p. 307-18. 180  225. Tse, A., et al., Rhythmic exocytosis stimulated by GnRH-induced calcium oscillations in rat gonadotropes. Science, 1993. 260(5104): p. 82-4. 226. Hajnoczky, G., et al., Decoding of cytosolic calcium oscillations in the mitochondria. Cell, 1995. 82(3): p. 415-24. 227. Dolmetsch, R.E., K. Xu, and R.S. Lewis, Calcium oscillations increase the efficiency and specificity of gene expression. Nature, 1998. 392(6679): p. 933-6. 228. Li, W., et al., Cell-permeant caged InsP3 ester shows that Ca2+ spike frequency can optimize gene expression. Nature, 1998. 392(6679): p. 936-41. 229. Haisenleder, D.J., et al., Gonadotropin subunit transcriptional responses to calcium signals in the rat: evidence for regulation by pulse frequency. Biol Reprod, 2001. 65(6): p. 1789-93. 230. Buonanno, A. and R.D. Fields, Gene regulation by patterned electrical activity during neural and skeletal muscle development. Curr Opin Neurobiol, 1999. 9(1): p. 110-20. 231. Abzhanov, A., et al., The calmodulin pathway and evolution of elongated beak morphology in Darwin's finches. Nature, 2006. 442(7102): p. 563-7. 232. Hoeflich, K.P. and M. Ikura, Calmodulin in action: diversity in target recognition and activation mechanisms. Cell, 2002. 108(6): p. 739-42. 233. Meador, W.E., A.R. Means, and F.A. Quiocho, Modulation of calmodulin plasticity in molecular recognition on the basis of x-ray structures. Science, 1993. 262(5140): p. 1718-21. 181  234. Meador, W.E., A.R. Means, and F.A. Quiocho, Target enzyme recognition by calmodulin: 2.4 A structure of a calmodulin-peptide complex. Science, 1992. 257(5074): p. 1251-5. 235. Chattopadhyaya, R., et al., Calmodulin structure refined at 1.7 A resolution. J Mol Biol, 1992. 228(4): p. 1177-92. 236. Pande, G., N.A. Kumar, and P.S. Manogaran, Flow cytometric study of changes in the intracellular free calcium during the cell cycle. Cytometry, 1996. 24(1): p. 55- 63. 237. Kahl, C.R. and A.R. Means, Regulation of cell cycle progression by calcium/calmodulin-dependent pathways. Endocr Rev, 2003. 24(6): p. 719-36. 238. Takuwa, N., et al., Ca(2+)-dependent stimulation of retinoblastoma gene product phosphorylation and p34cdc2 kinase activation in serum-stimulated human fibroblasts. J Biol Chem, 1993. 268(1): p. 138-45. 239. Rasmussen, C.D. and A.R. Means, Calmodulin is required for cell-cycle progression during G1 and mitosis. Embo J, 1989. 8(1): p. 73-82. 240. Berridge, M.J., M.D. Bootman, and P. Lipp, Calcium--a life and death signal. Nature, 1998. 395(6703): p. 645-8. 241. Lewis, R.S. and M.D. Cahalan, Ion channels and signal transduction in lymphocytes. Annu Rev Physiol, 1990. 52: p. 415-30. 242. Winslow, M.M., J.R. Neilson, and G.R. Crabtree, Calcium signalling in lymphocytes. Curr Opin Immunol, 2003. 15(3): p. 299-307. 182  243. Berridge, M.J., Lymphocyte activation in health and disease. Crit Rev Immunol, 1997. 17(2): p. 155-78. 244. Lewis, R.S., Calcium signaling mechanisms in T lymphocytes. Annu Rev Immunol, 2001. 19: p. 497-521. 245. Neilson, J.R., et al., Calcineurin B1 is essential for positive but not negative selection during thymocyte development. Immunity, 2004. 20(3): p. 255-66. 246. Randriamampita, C. and A. Trautmann, Ca2+ signals and T lymphocytes; "New mechanisms and functions in Ca2+ signalling". Biol Cell, 2004. 96(1): p. 69-78. 247. Zhong, F., et al., Bcl-2 differentially regulates Ca2+ signals according to the strength of T cell receptor activation. J Cell Biol, 2006. 172(1): p. 127-37. 248. Bellosta, S. and F. Bernini, Lipophilic calcium antagonists in antiatherosclerotic therapy. Curr Atheroscler Rep, 2000. 2(1): p. 76-81. 249. Simon, A. and J. Levenson, Effects of calcium channel blockers on atherosclerosis: new insights. Acta Cardiol, 2002. 57(4): p. 249-55. 250. Mancini, G.B., Antiatherosclerotic effects of calcium channel blockers. Prog Cardiovasc Dis, 2002. 45(1): p. 1-20. 251. Cristofori, P., et al., The calcium-channel blocker lacidipine reduces the development of atherosclerotic lesions in the apoE-deficient mouse. J Hypertens, 2000. 18(10): p. 1429-36. 252. Suzuki, J., et al., Effect of combination of calcium antagonist, azelnidipine, and AT1 receptor blocker, olmesartan, on atherosclerosis in apolipoprotein E- deficient mice. J Hypertens, 2005. 23(7): p. 1383-9. 183  253. Yao, R., et al., Molecular mechanisms of felodipine suppressing atherosclerosis in high-cholesterol-diet apolipoprotein E-knockout mice. J Cardiovasc Pharmacol, 2008. 51(2): p. 188-95. 254. Jinno, T., et al., Calcium channel blocker azelnidipine enhances vascular protective effects of AT1 receptor blocker olmesartan. Hypertension, 2004. 43(2): p. 263-9. 255. Yang, X., et al., Changes of transmembrane Ca2+ gradient in the formation of macrophage-derived foam cells. Biosci Rep, 2000. 20(1): p. 1-12. 256. Deng, T.L., et al., Intracellular-free calcium dynamics and F-actin alteration in the formation of macrophage foam cells. Biochem Biophys Res Commun, 2005. 338(2): p. 748-56. 257. Rong, Y. and C.W. Distelhorst, Bcl-2 protein family members: versatile regulators of calcium signaling in cell survival and apoptosis. Annu Rev Physiol, 2008. 70: p. 73-91. 258. Nairn, A.C., B. Bhagat, and H.C. Palfrey, Identification of calmodulin-dependent protein kinase III and its major Mr 100,000 substrate in mammalian tissues. Proc Natl Acad Sci U S A, 1985. 82(23): p. 7939-43. 259. Nairn, A.C. and H.C. Palfrey, Identification of the major Mr 100,000 substrate for calmodulin-dependent protein kinase III in mammalian cells as elongation factor- 2. J Biol Chem, 1987. 262(36): p. 17299-303. 184  260. Drennan, D. and A.G. Ryazanov, Alpha-kinases: analysis of the family and comparison with conventional protein kinases. Prog Biophys Mol Biol, 2004. 85(1): p. 1-32. 261. Pavur, K.S., A.N. Petrov, and A.G. Ryazanov, Mapping the functional domains of elongation factor-2 kinase. Biochemistry, 2000. 39(40): p. 12216-24. 262. Mitsui, K., et al., Purification and characterization of calmodulin-dependent protein kinase III from rabbit reticulocytes and rat pancreas. J Biol Chem, 1993. 268(18): p. 13422-33. 263. Redpath, N.T. and C.G. Proud, Purification and phosphorylation of elongation factor-2 kinase from rabbit reticulocytes. Eur J Biochem, 1993. 212(2): p. 511-20. 264. Palfrey, H.C., et al., Rapid activation of calmodulin-dependent protein kinase III in mitogen-stimulated human fibroblasts. Correlation with intracellular Ca2+ transients. J Biol Chem, 1987. 262(20): p. 9785-92. 265. Redpath, N.T., E.J. Foulstone, and C.G. Proud, Regulation of translation elongation factor-2 by insulin via a rapamycin-sensitive signalling pathway. EMBO J, 1996. 15(9): p. 2291-7. 266. Parmer, T.G., et al., Activity and regulation by growth factors of calmodulin- dependent protein kinase III (elongation factor 2-kinase) in human breast cancer. Br J Cancer, 1999. 79(1): p. 59-64. 267. Hait, W.N., et al., Elongation factor-2 kinase: immunological evidence for the existence of tissue-specific isoforms. FEBS Lett, 1996. 397(1): p. 55-60. 185  268. Diggle, T.A., et al., Regulation of protein-synthesis elongation-factor-2 kinase by cAMP in adipocytes. Biochem J, 1998. 336(Pt 3): p. 525-9. 269. Wang, L., X. Wang, and C.G. Proud, Activation of mRNA translation in rat cardiac myocytes by insulin involves multiple rapamycin-sensitive steps. Am J Physiol Heart Circ Physiol, 2000. 278(4): p. H1056-68. 270. Nilsson, A. and O. Nygard, Effect of oxidizing agents and haemin on the phosphorylation of eukaryotic elongation factor 2 in rabbit reticulocyte lysates. Biochim Biophys Acta, 1995. 1260(2): p. 200-6. 271. Kang, K.R. and S.Y. Lee, Effect of serum and hydrogen peroxide on the Ca2+/calmodulin-dependent phosphorylation of eukaryotic elongation factor 2(eEF-2) in Chinese hamster ovary cells. Exp Mol Med, 2001. 33(4): p. 198-204. 272. Jefferson, A.B. and H. Schulman, Sphingosine inhibits calmodulin-dependent enzymes. J Biol Chem, 1988. 263(30): p. 15241-4. 273. Sans, M.D., Q. Xie, and J.A. Williams, Regulation of translation elongation and phosphorylation of eEF2 in rat pancreatic acini. Biochem Biophys Res Commun, 2004. 319(1): p. 144-51. 274. Carroll, M., et al., 5-HT stimulates eEF2 dephosphorylation in a rapamycin- sensitive manner in Aplysia neurites. J Neurochem, 2004. 90(6): p. 1464-76. 275. McLeod, L.E., L. Wang, and C.G. Proud, beta-Adrenergic agonists increase phosphorylation of elongation factor 2 in cardiomyocytes without eliciting calcium-independent eEF2 kinase activity. FEBS Lett, 2001. 489(2-3): p. 225-8. 186  276. Mackie, K.P., et al., Thrombin and histamine stimulate the phosphorylation of elongation factor 2 in human umbilical vein endothelial cells. J Biol Chem, 1989. 264(3): p. 1748-53. 277. Albarracin, C.T., et al., Prolactin regulation of the calmodulin-dependent protein kinase III elongation factor-2 system in the rat corpus luteum. J Biol Chem, 1994. 269(10): p. 7772-6. 278. Everett, A.D., et al., Angiotensin II regulates phosphorylation of translation elongation factor-2 in cardiac myocytes. Am J Physiol Heart Circ Physiol, 2001. 281(1): p. H161-7. 279. Palmquist, K., et al., Interaction of the calcium and calmodulin regulated eEF-2 kinase with heat shock protein 90. FEBS Lett, 1994. 349(2): p. 239-42. 280. Yang, J., et al., Disruption of the EF-2 kinase/Hsp90 protein complex: a possible mechanism to inhibit glioblastoma by geldanamycin. Cancer Res, 2001. 61(10): p. 4010-6. 281. Dorovkov, M.V., et al., Regulation of elongation factor-2 kinase by pH. Biochemistry, 2002. 41(45): p. 13444-50. 282. Althausen, S., et al., Changes in the phosphorylation of initiation factor eIF- 2alpha, elongation factor eEF-2 and p70 S6 kinase after transient focal cerebral ischaemia in mice. J Neurochem, 2001. 78(4): p. 779-87. 283. Horman, S., et al., Activation of AMP-activated protein kinase leads to the phosphorylation of elongation factor 2 and an inhibition of protein synthesis. Curr Biol, 2002. 12(16): p. 1419-23. 187  284. Horman, S., et al., Myocardial ischemia and increased heart work modulate the phosphorylation state of eukaryotic elongation factor-2. J Biol Chem, 2003. 278(43): p. 41970-6. 285. Chan, A.Y., et al., Activation of AMP-activated protein kinase inhibits protein synthesis associated with hypertrophy in the cardiac myocyte. J Biol Chem, 2004. 279(31): p. 32771-9. 286. Terai, K., et al., AMP-activated protein kinase protects cardiomyocytes against hypoxic injury through attenuation of endoplasmic reticulum stress. Mol Cell Biol, 2005. 25(21): p. 9554-75. 287. Liu, L., et al., Hypoxia-induced energy stress regulates mRNA translation and cell growth. Mol Cell, 2006. 21(4): p. 521-31. 288. Wu, H., et al., Elongation factor-2 kinase regulates autophagy in human glioblastoma cells. Cancer Res, 2006. 66(6): p. 3015-23. 289. Hait, W.N., et al., Elongation factor-2 kinase: its role in protein synthesis and autophagy. Autophagy, 2006. 2(4): p. 294-6. 290. Smith, E.M. and C.G. Proud, cdc2-cyclin B regulates eEF2 kinase activity in a cell cycle- and amino acid-dependent manner. Embo J, 2008. 27(7): p. 1005-16. 291. Marin, P., et al., Glutamate-dependent phosphorylation of elongation factor-2 and inhibition of protein synthesis in neurons. J Neurosci, 1997. 17(10): p. 3445-54. 292. Knebel, A., N. Morrice, and P. Cohen, A novel method to identify protein kinase substrates: eEF2 kinase is phosphorylated and inhibited by SAPK4/p38delta. Embo J, 2001. 20(16): p. 4360-9. 188  293. Knebel, A., et al., Stress-induced regulation of eukaryotic elongation factor 2 kinase by SB 203580-sensitive and -insensitive pathways. Biochem J, 2002. 367(Pt 2): p. 525-32. 294. Browne, G.J., S.G. Finn, and C.G. Proud, Stimulation of the AMP-activated protein kinase leads to activation of eukaryotic elongation factor 2 kinase and to its phosphorylation at a novel site, serine 398. J Biol Chem, 2004. 279(13): p. 12220-31. 295. Browne, G.J. and C.G. Proud, A novel mTOR-regulated phosphorylation site in elongation factor 2 kinase modulates the activity of the kinase and its binding to calmodulin. Mol Cell Biol, 2004. 24(7): p. 2986-97. 296. Wang, X., et al., Regulation of elongation factor 2 kinase by p90(RSK1) and p70 S6 kinase. Embo J, 2001. 20(16): p. 4370-9. 297. Diggle, T.A., et al., Phosphorylation of elongation factor-2 kinase on serine 499 by cAMP-dependent protein kinase induces Ca2+/calmodulin-independent activity. Biochem J, 2001. 353(Pt 3): p. 621-6. 298. Gutzkow, K.B., et al., Cyclic AMP inhibits translation of cyclin D3 in T lymphocytes at the level of elongation by inducing eEF2-phosphorylation. Cell Signal, 2003. 15(9): p. 871-81. 299. Hovland, R., et al., cAMP inhibits translation by inducing Ca2+/calmodulin- independent elongation factor 2 kinase activity in IPC-81 cells. FEBS Lett, 1999. 444(1): p. 97-101. 189  300. Browne, G.J. and C.G. Proud, Regulation of peptide-chain elongation in mammalian cells. Eur J Biochem, 2002. 269(22): p. 5360-8. 301. Ryazanov, A.G., E.A. Shestakova, and P.G. Natapov, Phosphorylation of elongation factor 2 by EF-2 kinase affects rate of translation. Nature, 1988. 334(6178): p. 170-3. 302. Ryazanov, A.G. and E.K. Davydova, Mechanism of elongation factor 2 (EF-2) inactivation upon phosphorylation. Phosphorylated EF-2 is unable to catalyze translocation. FEBS Lett, 1989. 251(1-2): p. 187-90. 303. Redpath, N.T., et al., Regulation of elongation factor-2 by multisite phosphorylation. Eur J Biochem, 1993. 213(2): p. 689-99. 304. Carlberg, U., A. Nilsson, and O. Nygard, Functional properties of phosphorylated elongation factor 2. Eur J Biochem, 1990. 191(3): p. 639-45. 305. Nilsson, A. and O. Nygard, Phosphorylation of eukaryotic elongation factor 2 in differentiating and proliferating HL-60 cells. Biochim Biophys Acta, 1995. 1268(3): p. 263-8. 306. Bagaglio, D.M. and W.N. Hait, Role of calmodulin-dependent phosphorylation of elongation factor 2 in the proliferation of rat glial cells. Cell Growth Differ, 1994. 5(12): p. 1403-8. 307. Parmer, T.G., M.D. Ward, and W.N. Hait, Effects of rottlerin, an inhibitor of calmodulin-dependent protein kinase III, on cellular proliferation, viability, and cell cycle distribution in malignant glioma cells. Cell Growth Differ, 1997. 8(3): p. 327-34. 190  308. Bagaglio, D.M., et al., Phosphorylation of elongation factor 2 in normal and malignant rat glial cells. Cancer Res, 1993. 53(10 Suppl): p. 2260-4. 309. Prostko, C.R., C. Zhang, and W.N. Hait, The effects of altered cellular calmodulin expression on the growth and viability of C6 glioblastoma cells. Oncol Res, 1997. 9(1): p. 13-7. 310. Arora, S., et al., Identification and characterization of an inhibitor of eukaryotic elongation factor 2 kinase against human cancer cell lines. Cancer Res, 2003. 63(20): p. 6894-9. 311. Hershey, J.W., Translational control in mammalian cells. Annu Rev Biochem, 1991. 60: p. 717-55. 312. Merrick, W.C., Mechanism and regulation of eukaryotic protein synthesis. Microbiol Rev, 1992. 56(2): p. 291-315. 313. Pain, V.M., Initiation of protein synthesis in eukaryotic cells. Eur J Biochem, 1996. 236(3): p. 747-71. 314. Gaspar, N.J., et al., Translation initiation factor eIF-2. Cloning and expression of the human cDNA encoding the gamma-subunit. J Biol Chem, 1994. 269(5): p. 3415-22. 315. Hannig, E.M., et al., GCD11, a negative regulator of GCN4 expression, encodes the gamma subunit of eIF-2 in Saccharomyces cerevisiae. Mol Cell Biol, 1993. 13(1): p. 506-20. 316. Kozak, M., Structural features in eukaryotic mRNAs that modulate the initiation of translation. J Biol Chem, 1991. 266(30): p. 19867-70. 191  317. Kozak, M., The scanning model for translation: an update. J Cell Biol, 1989. 108(2): p. 229-41. 318. Rhoads, R.E., Regulation of eukaryotic protein synthesis by initiation factors. J Biol Chem, 1993. 268(5): p. 3017-20. 319. Merrick, W.C., Eukaryotic protein synthesis: an in vitro analysis. Biochimie, 1994. 76(9): p. 822-30. 320. Rhoads, R.E., B. Joshi, and W.B. Minich, Participation of initiation factors in the recruitment of mRNA to ribosomes. Biochimie, 1994. 76(9): p. 831-8. 321. Price, N. and C. Proud, The guanine nucleotide-exchange factor, eIF-2B. Biochimie, 1994. 76(8): p. 748-60. 322. Merrick, W.C. and J. Nyborg, The protein biosynthesis elongation cycle, in Translational Control of Gene Expression 2000, Cold Spring Harbor Laboratory Press: Coal Spring Harbor. p. 89-125. 323. Kaziro, Y., The role of guanosine 5'-triphosphate in polypeptide chain elongation. Biochim Biophys Acta, 1978. 505(1): p. 95-127. 324. Berchtold, H., et al., Crystal structure of active elongation factor Tu reveals major domain rearrangements. Nature, 1993. 365(6442): p. 126-32. 325. Kjeldgaard, M., et al., The crystal structure of elongation factor EF-Tu from Thermus aquaticus in the GTP conformation. Structure, 1993. 1(1): p. 35-50. 326. Clark, B.F., et al., Structural information for explaining the molecular mechanism of protein biosynthesis. FEBS Lett, 1999. 452(1-2): p. 41-6. 192  327. Frolova, L., et al., A highly conserved eukaryotic protein family possessing properties of polypeptide chain release factor. Nature, 1994. 372(6507): p. 701-3. 328. Zhouravleva, G., et al., Termination of translation in eukaryotes is governed by two interacting polypeptide chain release factors, eRF1 and eRF3. Embo J, 1995. 14(16): p. 4065-72. 329. Frolova, L., et al., Eukaryotic polypeptide chain release factor eRF3 is an eRF1- and ribosome-dependent guanosine triphosphatase. Rna, 1996. 2(4): p. 334-41. 330. Proud, C.G., Regulation of mammalian translation factors by nutrients. Eur J Biochem, 2002. 269(22): p. 5338-49. 331. Scheper, G.C. and C.G. Proud, Does phosphorylation of the cap-binding protein eIF4E play a role in translation initiation? Eur J Biochem, 2002. 269(22): p. 5350-9. 332. Gingras, A.C., B. Raught, and N. Sonenberg, Regulation of translation initiation by FRAP/mTOR. Genes Dev, 2001. 15(7): p. 807-26. 333. Proud, C.G., Regulation of eukaryotic initiation factor eIF2B. Prog Mol Subcell Biol, 2001. 26: p. 95-114. 334. Alberts, B., et al., Molecular Biology of the Cell. 4th ed. 1994, New York: Garland Publishing, Inc. 335. Yang, Y.P., et al., Molecular mechanism and regulation of autophagy. Acta Pharmacol Sin, 2005. 26(12): p. 1421-34. 336. Lum, J.J., et al., Growth factor regulation of autophagy and cell survival in the absence of apoptosis. Cell, 2005. 120(2): p. 237-48. 193  337. Kuma, A., et al., The role of autophagy during the early neonatal starvation period. Nature, 2004. 432(7020): p. 1032-6. 338. Mizushima, N. and D.J. Klionsky, Protein turnover via autophagy: implications for metabolism. Annu Rev Nutr, 2007. 27: p. 19-40. 339. Rubinsztein, D.C., et al., Potential therapeutic applications of autophagy. Nat Rev Drug Discov, 2007. 6(4): p. 304-12. 340. Maiuri, M.C., et al., Self-eating and self-killing: crosstalk between autophagy and apoptosis. Nat Rev Mol Cell Biol, 2007. 8(9): p. 741-52. 341. Hoyer-Hansen, M. and M. Jaattela, Connecting endoplasmic reticulum stress to autophagy by unfolded protein response and calcium. Cell Death Differ, 2007. 14(9): p. 1576-82. 342. Criollo, A., et al., Regulation of autophagy by the inositol trisphosphate receptor. Cell Death Differ, 2007. 14(5): p. 1029-39. 343. Meijer, A.J. and P. Codogno, Signalling and autophagy regulation in health, aging and disease. Mol Aspects Med, 2006. 27(5-6): p. 411-25. 344. Takeuchi, H., et al., Inhibition of platelet-derived growth factor signalling induces autophagy in malignant glioma cells. Br J Cancer, 2004. 90(5): p. 1069-75. 345. Boya, P., et al., Inhibition of macroautophagy triggers apoptosis. Mol Cell Biol, 2005. 25(3): p. 1025-40. 346. Daido, S., et al., Pivotal role of the cell death factor BNIP3 in ceramide-induced autophagic cell death in malignant glioma cells. Cancer Res, 2004. 64(12): p. 4286-93. 194  347. Kanzawa, T., et al., Role of autophagy in temozolomide-induced cytotoxicity for malignant glioma cells. Cell Death Differ, 2004. 11(4): p. 448-57. 348. Paglin, S., et al., A novel response of cancer cells to radiation involves autophagy and formation of acidic vesicles. Cancer Res, 2001. 61(2): p. 439-44. 349. Talloczy, Z., et al., Regulation of starvation- and virus-induced autophagy by the eIF2alpha kinase signaling pathway. Proc Natl Acad Sci U S A, 2002. 99(1): p. 190-5. 350. Levine, B. and G. Kroemer, Autophagy in the pathogenesis of disease. Cell, 2008. 132(1): p. 27-42. 351. Tushinski, R.J. and E.R. Stanley, The regulation of macrophage protein turnover by a colony stimulating factor (CSF-1). J Cell Physiol, 1983. 116(1): p. 67-75. 352. Tushinski, R.J., et al., Survival of mononuclear phagocytes depends on a lineage- specific growth factor that the differentiated cells selectively destroy. Cell, 1982. 28(1): p. 71-81. 353. Steinberg, D., Oxidized low density lipoprotein--an extreme example of lipoprotein heterogeneity. Isr J Med Sci, 1996. 32(6): p. 469-72. 354. Stanley, E.R. and P.M. Heard, Factors regulating macrophage production and growth. Purification and some properties of the colony stimulating factor from medium conditioned by mouse L cells. J Biol Chem, 1977. 252(12): p. 4305-12. 355. Schackelford, R.E., et al., Oxidized low density lipoprotein suppresses activation of NF kappa B in macrophages via a pertussis toxin-sensitive signaling mechanism. J Biol Chem, 1995. 270(8): p. 3475-8. 195  356. Fueller, M., et al., Activation of human monocytic cells by lysophosphatidic acid and sphingosine-1-phosphate. Cell Signal, 2003. 15(4): p. 367-75. 357. Han, C.Y., S.Y. Park, and Y.K. Pak, Role of endocytosis in the transactivation of nuclear factor-kappaB by oxidized low-density lipoprotein. Biochem J, 2000. 350(Pt 3): p. 829-37. 358. Pozzan, T., et al., Molecular and cellular physiology of intracellular calcium stores. Physiol Rev, 1994. 74(3): p. 595-636. 359. Inesi, G. and Y. Sagara, Thapsigargin, a high affinity and global inhibitor of intracellular Ca2+ transport ATPases. Arch Biochem Biophys, 1992. 298(2): p. 313-7. 360. Sagara, Y., et al., Characterization of the inhibition of intracellular Ca2+ transport ATPases by thapsigargin. J Biol Chem, 1992. 267(18): p. 12606-13. 361. Sagara, Y., J.B. Wade, and G. Inesi, A conformational mechanism for formation of a dead-end complex by the sarcoplasmic reticulum ATPase with thapsigargin. J Biol Chem, 1992. 267(2): p. 1286-92. 362. Yule, D.I. and J.A. Williams, U73122 inhibits Ca2+ oscillations in response to cholecystokinin and carbachol but not to JMV-180 in rat pancreatic acinar cells. J Biol Chem, 1992. 267(20): p. 13830-5. 363. Zhao, F., et al., Dantrolene inhibition of ryanodine receptor Ca2+ release channels. Molecular mechanism and isoform selectivity. J Biol Chem, 2001. 276(17): p. 13810-6. 196  364. French, K.J., et al., Discovery and evaluation of inhibitors of human sphingosine kinase. Cancer Res, 2003. 63(18): p. 5962-9. 365. Holvoet, P., et al., Oxidized low-density lipoprotein correlates positively with toll- like receptor 2 and interferon regulatory factor-1 and inversely with superoxide dismutase-1 expression: studies in hypercholesterolemic swine and THP-1 cells. Arterioscler Thromb Vasc Biol, 2006. 26(7): p. 1558-65. 366. Miller, Y.I., et al., Toll-like receptor 4-dependent and -independent cytokine secretion induced by minimally oxidized low-density lipoprotein in macrophages. Arterioscler Thromb Vasc Biol, 2005. 25(6): p. 1213-9. 367. Yang, Q.W., et al., Role of Toll-like receptor 4/NF-kappaB pathway in monocyte- endothelial adhesion induced by low shear stress and ox-LDL. Biorheology, 2005. 42(3): p. 225-36. 368. Cominacini, L., et al., Oxidized low density lipoprotein (ox-LDL) binding to ox- LDL receptor-1 in endothelial cells induces the activation of NF-kappaB through an increased production of intracellular reactive oxygen species. J Biol Chem, 2000. 275(17): p. 12633-8. 369. Cominacini, L., et al., The binding of oxidized low density lipoprotein (ox-LDL) to ox-LDL receptor-1 reduces the intracellular concentration of nitric oxide in endothelial cells through an increased production of superoxide. J Biol Chem, 2001. 276(17): p. 13750-5. 197  370. Kataoka, H., et al., Oxidized LDL modulates Bax/Bcl-2 through the lectinlike Ox- LDL receptor-1 in vascular smooth muscle cells. Arterioscler Thromb Vasc Biol, 2001. 21(6): p. 955-60. 371. Li, D. and J.L. Mehta, Upregulation of endothelial receptor for oxidized LDL (LOX-1) by oxidized LDL and implications in apoptosis of human coronary artery endothelial cells: evidence from use of antisense LOX-1 mRNA and chemical inhibitors. Arterioscler Thromb Vasc Biol, 2000. 20(4): p. 1116-22. 372. Sawamura, T., et al., An endothelial receptor for oxidized low-density lipoprotein. Nature, 1997. 386(6620): p. 73-7. 373. Wang, S.W., et al., Pertussis toxin promotes macrophage survival through inhibition of acid sphingomyelinase and activation of the phosphoinositide 3- kinase/protein kinase B pathway. Cell Signal, 2007. 19(8): p. 1772-83. 374. Mackay, H.J. and C.J. Twelves, Targeting the protein kinase C family: are we there yet? Nat Rev Cancer, 2007. 7(7): p. 554-62. 375. Blobe, G.C., L.M. Obeid, and Y.A. Hannun, Regulation of protein kinase C and role in cancer biology. Cancer Metastasis Rev, 1994. 13(3-4): p. 411-31. 376. Newton, A.C., Protein kinase C: structure, function, and regulation. J Biol Chem, 1995. 270(48): p. 28495-8. 377. Nishizuka, Y., Intracellular signaling by hydrolysis of phospholipids and activation of protein kinase C. Science, 1992. 258(5082): p. 607-14. 378. Schenk, P.W. and B.E. Snaar-Jagalska, Signal perception and transduction: the role of protein kinases. Biochim Biophys Acta, 1999. 1449(1): p. 1-24. 198  379. Hamilton, J.A. and S.R. Dientsman, Induction of macrophage DNA synthesis by phorbol esters. J Cell Physiol, 1981. 106(3): p. 445-50. 380. Hamilton, J.A., Glucocorticoids and prostaglandins inhibit the induction of macrophage DNA synthesis by macrophage growth factor and phorbol ester. J Cell Physiol, 1983. 115(1): p. 67-74. 381. Guilbert, L.J., et al., The nature of 12-O-tetradecanoylphorbol-13-acetate (TPA)- stimulated hemopoiesis, colony stimulating factor (CSF) requirement for colony formation, and the effect of TPA on [125I]CSF-1 binding to macrophages. J Cell Physiol, 1983. 115(3): p. 276-82. 382. Monick, M.M., et al., Changes in PKC isoforms in human alveolar macrophages compared with blood monocytes. Am J Physiol, 1998. 275(2 Pt 1): p. L389-97. 383. Todt, J.C., et al., Activation of protein kinase C beta II by the stereo-specific phosphatidylserine receptor is required for phagocytosis of apoptotic thymocytes by resident murine tissue macrophages. J Biol Chem, 2002. 277(39): p. 35906-14. Epub 2002 Jul 11. 384. Wilkinson, S.E., P.J. Parker, and J.S. Nixon, Isoenzyme specificity of bisindolylmaleimides, selective inhibitors of protein kinase C. Biochem J, 1993. 294(Pt 2): p. 335-7. 385. Gschwendt, M., et al., Rottlerin, a novel protein kinase inhibitor. Biochem Biophys Res Commun, 1994. 199(1): p. 93-8. 199  386. Gschwendt, M., W. Kittstein, and F. Marks, Elongation factor-2 kinase: effective inhibition by the novel protein kinase inhibitor rottlerin and relative insensitivity towards staurosporine. FEBS Lett, 1994. 338(1): p. 85-8. 387. Miyazaki, A., et al., Granulocyte macrophage colony-stimulating factor plays a priming role in murine macrophage growth induced by oxidized low density lipoprotein. Ann N Y Acad Sci, 2000. 902: p. 342-6. 388. Cho, S.I., et al., Novel compounds, '1,3-selenazine derivatives' as specific inhibitors of eukaryotic elongation factor-2 kinase. Biochim Biophys Acta, 2000. 1475(3): p. 207-15. 389. Hori, H., et al., TX-1123: an antitumor 2-hydroxyarylidene-4-cyclopentene-1,3- dione as a protein tyrosine kinase inhibitor having low mitochondrial toxicity. Bioorg Med Chem, 2002. 10(10): p. 3257-65. 390. Mita, M.M., A. Mita, and E.K. Rowinsky, The molecular target of rapamycin (mTOR) as a therapeutic target against cancer. Cancer Biol Ther, 2003. 2(4 Suppl 1): p. S169-77. 391. Duncia, J.V., et al., MEK inhibitors: the chemistry and biological activity of U0126, its analogs, and cyclization products. Bioorg Med Chem Lett, 1998. 8(20): p. 2839-44. 392. Favata, M.F., et al., Identification of a novel inhibitor of mitogen-activated protein kinase kinase. J Biol Chem, 1998. 273(29): p. 18623-32. 200  393. de Souza, N.J., A.N. Dohadwalla, and J. Reden, Forskolin: a labdane diterpenoid with antihypertensive, positive inotropic, platelet aggregation inhibitory, and adenylate cyclase activating properties. Med Res Rev, 1983. 3(2): p. 201-19. 394. Beavo, J.A., et al., Effects of xanthine derivatives on lipolysis and on adenosine 3',5'-monophosphate phosphodiesterase activity. Mol Pharmacol, 1970. 6(6): p. 597-603. 395. Chijiwa, T., et al., Inhibition of forskolin-induced neurite outgrowth and protein phosphorylation by a newly synthesized selective inhibitor of cyclic AMP- dependent protein kinase, N-[2-(p-bromocinnamylamino)ethyl]-5- isoquinolinesulfonamide (H-89), of PC12D pheochromocytoma cells. J Biol Chem, 1990. 265(9): p. 5267-72. 396. Xaus, J., et al., Adenosine inhibits macrophage colony-stimulating factor- dependent proliferation of macrophages through the induction of p27kip-1 expression. J Immunol, 1999. 163(8): p. 4140-9. 397. Wegele, H., L. Muller, and J. Buchner, Hsp70 and Hsp90--a relay team for protein folding. Rev Physiol Biochem Pharmacol, 2004. 151: p. 1-44. Epub 2004 Jan 23. 398. Yang, J., et al., Disruption of the EF-2 kinase/Hsp90 protein complex: a possible mechanism to inhibit glioblastoma by geldanamycin. Cancer Res, 2001. 61(10): p. 4010-6. 399. Miyata, Y., Hsp90 inhibitor geldanamycin and its derivatives as novel cancer chemotherapeutic agents. Curr Pharm Des, 2005. 11(9): p. 1131-8. 201  400. Knebel, A., N. Morrice, and P. Cohen, A novel method to identify protein kinase substrates: eEF2 kinase is phosphorylated and inhibited by SAPK4/p38delta. EMBO J, 2001. 20(16): p. 4360-9. 401. Knebel, A., et al., Stress-induced regulation of eukaryotic elongation factor 2 kinase by SB 203580-sensitive and -insensitive pathways. Biochem J, 2002. 367(Pt 2): p. 525-32. 402. Davies, S.P., et al., Specificity and mechanism of action of some commonly used protein kinase inhibitors. Biochem J, 2000. 351(Pt 1): p. 95-105. 403. Rosini, P., et al., NGF withdrawal induces apoptosis in CESS B cell line through p38 MAPK activation and Bcl-2 phosphorylation. Biochem Biophys Res Commun, 2000. 278(3): p. 753-9. 404. Torcia, M., et al., Nerve growth factor inhibits apoptosis in memory B lymphocytes via inactivation of p38 MAPK, prevention of Bcl-2 phosphorylation, and cytochrome c release. J Biol Chem, 2001. 276(42): p. 39027-36. Epub 2001 Aug 8. 405. Kummer, J.L., P.K. Rao, and K.A. Heidenreich, Apoptosis induced by withdrawal of trophic factors is mediated by p38 mitogen-activated protein kinase. J Biol Chem, 1997. 272(33): p. 20490-4. 406. Rosini, P., et al., Nerve growth factor-dependent survival of CESS B cell line is mediated by increased expression and decreased degradation of MAPK phosphatase 1. J Biol Chem, 2004. 279(14): p. 14016-23. Epub 2004 Jan 14. 202  407. Medvedev, A.E., et al., Limited role of ceramide in lipopolysaccharide-mediated mitogen-activated protein kinase activation, transcription factor induction, and cytokine release. J Biol Chem, 1999. 274(14): p. 9342-50. 408. Scheid, M.P., et al., Ceramide and cyclic adenosine monophosphate (cAMP) induce cAMP response element binding protein phosphorylation via distinct signaling pathways while having opposite effects on myeloid cell survival. Blood, 1999. 93(1): p. 217-25. 409. Heller, J.I., et al., p-hydroxyphenylacetaldehyde, an aldehyde generated by myeloperoxidase, modifies phospholipid amino groups of low density lipoprotein in human atherosclerotic intima. J Biol Chem, 2000. 275(14): p. 9957-62. 410. Franklin, C.C. and A.S. Kraft, Conditional expression of the mitogen-activated protein kinase (MAPK) phosphatase MKP-1 preferentially inhibits p38 MAPK and stress-activated protein kinase in U937 cells. J Biol Chem, 1997. 272(27): p. 16917-23. 411. Franklin, C.C., S. Srikanth, and A.S. Kraft, Conditional expression of mitogen- activated protein kinase phosphatase-1, MKP-1, is cytoprotective against UV- induced apoptosis. Proc Natl Acad Sci U S A, 1998. 95(6): p. 3014-9. 412. Wu, J.J. and A.M. Bennett, Essential Role for Mitogen-activated Protein (MAP) Kinase Phosphatase-1 in Stress-responsive MAP Kinase and Cell Survival Signaling. J Biol Chem, 2005. 280(16): p. 16461-6. Epub 2005 Feb 17. 203  413. Reddy, S.T., et al., Potential role for mitogen-activated protein kinase phosphatase-1 in the development of atherosclerotic lesions in mouse models. Arterioscler Thromb Vasc Biol, 2004. 24(9): p. 1676-81. Epub 2004 Jul 8. 414. Krautwald, S., et al., Suppression of growth factor-mediated MAP kinase activation by v-raf in macrophages: a putative role for the MKP-1 phosphatase. Oncogene, 1995. 10(6): p. 1187-92. 415. Valledor, A.F., et al., Macrophage colony-stimulating factor induces the expression of mitogen-activated protein kinase phosphatase-1 through a protein kinase C-dependent pathway. J Immunol, 1999. 163(5): p. 2452-62. 416. Reddy, S., et al., Mitogen-activated protein kinase phosphatase 1 activity is necessary for oxidized phospholipids to induce monocyte chemotactic activity in human aortic endothelial cells. J Biol Chem, 2001. 276(20): p. 17030-5. Epub 2001 Mar 5. 417. Eskelinen, E.L., Fine structure of the autophagosome. Methods Mol Biol, 2008. 445: p. 11-28. 418. Kabeya, Y., et al., LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. Embo J, 2000. 19(21): p. 5720-8. 419. Jager, S., et al., Role for Rab7 in maturation of late autophagic vacuoles. J Cell Sci, 2004. 117(Pt 20): p. 4837-48. 420. Hardie, D.G. and S.A. Hawley, AMP-activated protein kinase: the energy charge hypothesis revisited. Bioessays, 2001. 23(12): p. 1112-9. 204  421. Hardie, D.G., The AMP-activated protein kinase pathway--new players upstream and downstream. J Cell Sci, 2004. 117(Pt 23): p. 5479-87. 422. Meijer, A.J. and P.F. Dubbelhuis, Amino acid signalling and the integration of metabolism. Biochem Biophys Res Commun, 2004. 313(2): p. 397-403. 423. Wang, Z., et al., Antagonistic controls of autophagy and glycogen accumulation by Snf1p, the yeast homolog of AMP-activated protein kinase, and the cyclin- dependent kinase Pho85p. Mol Cell Biol, 2001. 21(17): p. 5742-52. 424. Papandreou, I., et al., Hypoxia signals autophagy in tumor cells via AMPK activity, independent of HIF-1, BNIP3, and BNIP3L. Cell Death Differ, 2008. 425. Takagi, H., et al., AMPK mediates autophagy during myocardial ischemia in vivo. Autophagy, 2007. 3(4): p. 405-7. 426. Matsui, Y., et al., Distinct roles of autophagy in the heart during ischemia and reperfusion: roles of AMP-activated protein kinase and Beclin 1 in mediating autophagy. Circ Res, 2007. 100(6): p. 914-22. 427. Liang, J., et al., The energy sensing LKB1-AMPK pathway regulates p27(kip1) phosphorylation mediating the decision to enter autophagy or apoptosis. Nat Cell Biol, 2007. 9(2): p. 218-24. 428. Henin, N., M.F. Vincent, and G. Van den Berghe, Stimulation of rat liver AMP- activated protein kinase by AMP analogues. Biochim Biophys Acta, 1996. 1290(2): p. 197-203. 205  429. Corton, J.M., et al., 5-aminoimidazole-4-carboxamide ribonucleoside. A specific method for activating AMP-activated protein kinase in intact cells? Eur J Biochem, 1995. 229(2): p. 558-65. 430. Ovchinnikov, L.P., et al., Three phosphorylation sites in elongation factor 2. FEBS Lett, 1990. 275(1-2): p. 209-12. 431. Price, N.T., et al., Identification of the phosphorylation sites in elongation factor- 2 from rabbit reticulocytes. FEBS Lett, 1991. 282(2): p. 253-8. 432. Celis, J.E., P. Madsen, and A.G. Ryazanov, Increased phosphorylation of elongation factor 2 during mitosis in transformed human amnion cells correlates with a decreased rate of protein synthesis. Proc Natl Acad Sci U S A, 1990. 87(11): p. 4231-5. 433. Fan, H. and S. Penman, Regulation of protein synthesis in mammalian cells. II. Inhibition of protein synthesis at the level of initiation during mitosis. J Mol Biol, 1970. 50(3): p. 655-70. 434. Greenberg, M.E., A.L. Hermanowski, and E.B. Ziff, Effect of protein synthesis inhibitors on growth factor activation of c-fos, c-myc, and actin gene transcription. Mol Cell Biol, 1986. 6(4): p. 1050-7. 435. Muller, R., et al., Induction of c-fos gene and protein by growth factors precedes activation of c-myc. Nature, 1984. 312(5996): p. 716-20. 436. Scheetz, A.J., A.C. Nairn, and M. Constantine-Paton, NMDA receptor-mediated control of protein synthesis at developing synapses. Nat Neurosci, 2000. 3(3): p. 211-6. 206  437. Hoyer-Hansen, M. and M. Jaattela, AMP-activated protein kinase: a universal regulator of autophagy? Autophagy, 2007. 3(4): p. 381-3. 438. Swerdlow, S. and C.W. Distelhorst, Bcl-2-regulated calcium signals as common mediators of both apoptosis and autophagy. Dev Cell, 2007. 12(2): p. 178-9. 439. Gao, W., et al., Induction of macroautophagy by exogenously introduced calcium. Autophagy, 2008. 4(6): p. 754-61. 440. Eskelinen, E.L. and P. Saftig, Autophagy: A lysosomal degradation pathway with a central role in health and disease. Biochim Biophys Acta, 2008. 441. Levine, B., Cell biology: autophagy and cancer. Nature, 2007. 446(7137): p. 745- 7. 442. Mathew, R., V. Karantza-Wadsworth, and E. White, Role of autophagy in cancer. Nat Rev Cancer, 2007. 7(12): p. 961-7. 443. Bursch, W., The autophagosomal-lysosomal compartment in programmed cell death. Cell Death Differ, 2001. 8(6): p. 569-81. 444. Edinger, A.L. and C.B. Thompson, Death by design: apoptosis, necrosis and autophagy. Curr Opin Cell Biol, 2004. 16(6): p. 663-9. 445. Gonzalez-Polo, R.A., et al., The apoptosis/autophagy paradox: autophagic vacuolization before apoptotic death. J Cell Sci, 2005. 118(Pt 14): p. 3091-102.   

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
https://iiif.library.ubc.ca/presentation/dsp.24.1-0067286/manifest

Comment

Related Items