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Tissue factor-related mechanisms of coagulation initiated on the virus envelope Lin, Huan-Jih 2019

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TISSUE FACTOR-RELATED MECHANISMS OF COAGULATION  INITIATED ON THE VIRUS ENVELOPE  by  Huan-Jih Lin  B.Sc., The University of British Columbia, 2013  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  August 2019  © Huan-Jih Lin, 2018 ii  [Committee page for doctoral committees with one supervisor. Select the correct committee page for your work, and delete unnecessary committee pages and this material in brackets, before submitting.]  The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Mechanisms of Coagulation Initiated on the Virus Envelope Related to Tissue Factor  submitted by HUAN-JIH LIN in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Pathology and Laboratory Medicine  Examining Committee: Ed LG Pryzdial, Pathology and Laboratory Medicine Supervisor  William W-G Jia, Pathology and Laboratory Medicine Departmental Committee Member  Jay N Kizhakkedathu, Pathology and Laboratory Medicine Supervisory Committee Member Ross TA MacGillivray, Biochemistry and Molecular Biology University Examiner Patricia C Liaw, McMaster University, Medicine External Examiner  Additional Supervisory Committee Members: Hélène C F Côté, Pathology and Laboratory Medicine Supervisory Committee Member Marc S Horwitz, Microbiology and Immunology Supervisory Committee Member  iii  Abstract Infectious diseases do not always manifest in a predictable manner and may go undetected, especially in asymptomatic individuals. Here, virus infection as an underlying cause of vascular disease is investigated. When considered independently, both have major impacts on healthcare systems globally. However, a relatively unexplored area of investigation is that many virus types have been tied to cardiovascular and haemostatic diseases. The mechanisms by which viruses accomplish this is not understood. As a model, I have used herpes simplex virus type 1 (HSV1) to explore the concept that viruses that obtain an outer phospholipid bilayer membrane can acquire the main physiological initiator of coagulation, tissue factor (TF). The Pryzdial laboratory has previously shown that several herpesviruses, including HSV1, incorporate TF when they are propagated in TF-expressing cells. TF is involved in the activation of the coagulation pro-enzyme factor X through the enhancement of factor VIIa proteolytic activity. This was shown to enhance virus infectivity through coagulation protease-dependent activation of cell signaling. HSV1 also expresses its self-encoded membrane protein, glycoprotein C (gC), which is involved in cell adhesion and immune escape. As gC is also capable of binding factor X, I postulated that gC regulates factor X activation through TF on the virus surface. To study how gC affects TF function, protein interactions were followed via cross-linking, chromogenic activity assays and microscale thermophoresis. In this study, I discovered that gC interacted with factor VIIa and factor X, rather than with TF, to form a unique enzyme complex that activates factor X. Similar to TF and factor VIIa interaction, gC requires factor X to efficiently assemble with factor VIIa. This complex can directly activate factor X and induce fibrin formation in plasma. Although this pro-coagulant function of gC may be restricted to homologous gene products in all herpesvirus family members, I have demonstrated that other viruses can incorporate TF into their membranes. In addition to iv  purified viruses propagated in cell culture, I found that patient plasma-derived human immunodeficiency virus and hepatitis C virus can incorporate TF into their virus envelope. This could explain some of the haemostatic abnormalities associated with viral infection.   v  Lay Summary Imbalances in blood clotting are leading causes of death worldwide. The reasons behind deadly blood clots are numerous. One of the possible links to blood clots that interest the Pryzdial laboratory is virus infection. My research focuses on how some viruses can promote clotting or clotting-related problems. Blood clotting is good when we want to stop bleeding. Yet, clotting can be bad when it is not needed or too much takes place. This can result in problems such as heart attacks and strokes. I have found that some viruses steal clotting factors from our cells. These factors may cause clotting and increase infection. Also, I have shown that the virus that causes oral herpes can make its own clotting factor. With this knowledge, we can be better prepared for fighting these viruses and their associated health problems. We can also learn how viral clotting factors work to improve our own.    vi  Preface The research question was originally proposed by my supervisor, Dr. Ed Pryzdial, Clinical Professor in the Department of Pathology and Laboratory Medicine, UBC. Herpesviruses were produced and purified by our laboratory Research Associate Dr. Michael Sutherland (Adjunct Professor, Department of Pathology and Laboratory Medicine, UBC). All experimental work presented in the dissertation was performed by me. Experimental work with human plasma was approved by the UBC Clinical Research Ethics Board (ethics certificate: H11-02845). A version of Chapter 3 will be submitted for publication with the title: Coagulation factor X augments FVIIa binding to herpes simplex virus 1-encoded glycoprotein C forming an extrinsic tenase-like complex (by Bryan H. Lin, Michael R. Sutherland, Federico I. Rosell, James H. Morrissey, and Edward L.G. Pryzdial). The majority of experiments described in Chapter 3 were designed by me with the feedback from Dr. Michael Sutherland, our collaborator, Dr. James Morrissey (professor, departments of Biological Chemistry & Internal Medicine, University of Michigan Medical School) and thesis advisory committee members. I wrote the draft of the manuscript and revised the contents with my supervisor in addition to feedback from Drs. Sutherland and Morrissey.  The work presented in Chapter 4 was designed in large part by me with suggestions from  Dr. Michael Sutherland and will be prepared in a manuscript in progress with the tentative title: Ubiquity of tissue factor on the surface of enveloped viruses (by Bryan H. Lin, Michael R. Sutherland, Guey-Chuen Perng, Ivan Sadowski, and Edward L.G. Pryzdial). Parts of Chapter 3 and 4 were presented orally or in poster presentations at various local, national and international conferences, symposia and workshops. Microscale thermophoresis diagram and figure legend was obtained under a Creative Commons Attribution License (CC BY). vii  Table of Contents  Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ........................................................................................................................ vii List of Tables .............................................................................................................................. xiii List of Figures ............................................................................................................................. xiv List of Abbreviations ................................................................................................................ xvii Acknowledgements .................................................................................................................... xxi Chapter 1: Introduction ................................................................................................................1 1.1 Thesis Rationale .............................................................................................................. 1 1.2 Infectious diseases and coagulation ................................................................................ 2 1.2.1 Thrombosis ................................................................................................................. 2 1.2.2 Infectious diseases in inflammation ............................................................................ 3 1.3 General coagulation overview ........................................................................................ 4 1.3.1 Tissue Factor ............................................................................................................... 6 1.3.1.1 Structure .............................................................................................................. 6 1.3.1.2 Function in haemostasis ...................................................................................... 8 1.3.1.3 Protease-activated receptors and TF function ..................................................... 9 1.3.1.4 Function(s) in non-haemostatic settings ........................................................... 10 1.3.1.5 Regulation of TF function ................................................................................. 12 1.3.2 Factor VIIa ................................................................................................................ 13 viii  1.3.2.1 Structure ............................................................................................................ 13 1.3.2.2 Function ............................................................................................................ 16 1.3.2.3 Regulation of FVIIa expression and function ................................................... 18 1.3.3 Factor X .................................................................................................................... 18 1.3.3.1 Structure ............................................................................................................ 18 1.3.3.2 Function ............................................................................................................ 19 1.3.3.3 Regulation of expression and function ............................................................. 23 1.3.4 Anticoagulation and fibrinolysis ............................................................................... 24 1.4 Enveloped viruses ......................................................................................................... 25 1.4.1 Herpesviridae ............................................................................................................ 27 1.4.1.1 HSV1................................................................................................................. 29 1.4.1.1.1 Implications in cardiovascular/haemostatic abnormalities ......................... 30 1.4.1.2 Glycoprotein C .................................................................................................. 31 1.4.1.2.1 Structure ...................................................................................................... 31 1.4.1.2.2 Function ....................................................................................................... 31 1.4.2 Flaviviridae ............................................................................................................... 35 1.4.2.1 Dengue virus ..................................................................................................... 36 1.4.2.1.1 Implications on cardiovascular/haemostatic abnormalities......................... 37 1.4.2.2 Hepatitis C virus ............................................................................................... 37 1.4.2.2.1 Implications in cardiovascular/haemostatic abnormalities ......................... 38 1.4.3 Retroviridae............................................................................................................... 39 1.4.3.1 Human immunodeficiency virus ....................................................................... 39 1.4.3.1.1 Implications in cardiovascular/haemostatic abnormalities ......................... 40 ix  1.5 Hypotheses .................................................................................................................... 40 Chapter 2: Material and Methods ..............................................................................................43 2.1 Proteins and reagents. ................................................................................................... 43 2.1.1 Antibodies and probes............................................................................................... 44 2.1.2 Preparation of gold-conjugated primary antibodies .................................................. 45 2.1.3 Viruses ...................................................................................................................... 45 2.1.3.1 Characterizing antigens in HSV1 panel ............................................................ 46 2.1.4 Small unilamellar vesicles ........................................................................................ 48 2.1.5 Protein expression and generation ............................................................................ 48 2.1.5.1 gCΔ457t/sgC-His .............................................................................................. 50 2.1.5.1.1 Protein purification ...................................................................................... 50 2.1.5.1.2 Determination of baculovirus titer .............................................................. 52 2.1.5.2 sTF-His ............................................................................................................. 54 2.2 Detection of specific antigens on the virus envelope .................................................... 56 2.2.1 Immunogold electron microscopy ............................................................................ 56 2.2.2 Purified cell culture-derived HSV1 particles ............................................................ 58 2.2.3 Purified cell culture-derived DENV particles ........................................................... 59 2.2.4 Patient plasma-derived virus particles ...................................................................... 59 2.2.4.1 Centrifugation method ...................................................................................... 60 2.2.4.2 Immunocapture method .................................................................................... 62 2.3 Assessing the function of procoagulant factors. ........................................................... 64 2.3.1 Virus-mediated FX activation ................................................................................... 64 2.3.2 Analysis of FX activation in purified systems .......................................................... 64 x  2.3.2.1 Deriving FX activation kinetics parameters ..................................................... 65 2.3.2.2 Nickel binding of purified proteins ................................................................... 67 2.3.2.3 FVIIa binding to lipid/protein cofactors ........................................................... 68 2.3.3 Plasma clotting .......................................................................................................... 68 2.4 Determining binding partners ....................................................................................... 69 2.4.1 Crosslinking .............................................................................................................. 69 2.4.2 ELISA-based chromogenic assay ............................................................................. 70 2.4.3 Microscale thermophoresis ....................................................................................... 71 2.4.3.1 Experimental conditions ................................................................................... 73 2.4.3.2 Protein labeling with fluorophores.................................................................... 75 2.4.3.3 Amine-reactive crosslinker reactive groups ...................................................... 75 2.4.3.4 Histidine tag affinity labeling ........................................................................... 76 2.4.3.5 Determining protein concentration and degree of labeling ............................... 77 2.4.4 Removal of His-tag on sTF-His by cyanogen bromide (CNBr) cleavage. ............... 77 Chapter 3: HSV1 glycoprotein C and its effect on TF/FVIIa complex ...................................80 3.1 TF function on the virus surface ................................................................................... 80 3.2 Procoagulant factors on the virus surface ..................................................................... 80 3.2.1 Effect of gC on TF function ...................................................................................... 83 3.2.2 Effect of soluble gC to viral TF ................................................................................ 85 3.2.3 Effect of soluble gC to relipidated TF ...................................................................... 88 3.3 gC function in a divisible system .................................................................................. 88 3.4 Effect of sgC-His on TF function in a purified and defined system ............................. 90 3.4.1 FX activation kinetics by sgC-His/FVIIa ................................................................. 92 xi  3.4.2 sgC-His-induced plasma clotting .............................................................................. 97 3.5 Interactions between TF and gC ................................................................................... 97 3.5.1 Cross-linking on the virus surface ............................................................................ 97 3.5.2 Cross-linking of purified proteins ............................................................................. 99 3.5.3 ELISA-modified approach ........................................................................................ 99 3.5.4 Microscale thermophoresis ..................................................................................... 103 3.5.4.1 Fluorophore labeling of sTF-His .................................................................... 103 3.5.4.2 Fluorophore labeling of sgC-His .................................................................... 104 3.6 Interactions between gC and FVIIa ............................................................................ 104 3.6.1 sgC-His binding interactions in solution-phase ...................................................... 104 3.6.2 sgC-His binding interactions on membranes .......................................................... 106 3.6.3 sgC-His/FVIIa amidolytic activity .......................................................................... 111 3.7 Coagulation initiated by HSV1 ................................................................................... 115 3.7.1 HSV1-mediated plasma clotting ............................................................................. 115 3.8 Effect of glycosaminoglycans on gC .......................................................................... 115 3.9 Discussion ................................................................................................................... 118 3.9.1 Interactions between TF and gC ............................................................................. 118 3.9.2 Nickel-chelating lipids as a tool for discovery ....................................................... 120 3.9.3 Microscale thermophoresis ..................................................................................... 121 3.9.4 gC interaction with FVIIa ....................................................................................... 122 3.9.5 gC procoagulant activity in infection ...................................................................... 124 3.9.6 Coagulation initiated by HSV1 ............................................................................... 125 Chapter 4: Tissue factor on enveloped viruses........................................................................127 xii  4.1 Tissue factor presence on purified dengue virus ......................................................... 128 4.1.1 FVIIa-mediated FX activation ................................................................................ 128 4.1.2 TF-mediated plasma clotting .................................................................................. 128 4.1.3 Immunogold EM ..................................................................................................... 131 4.2 Prevalence of tissue factor on patient-derived viruses ................................................ 133 4.2.1 HIV and TF ............................................................................................................. 133 4.2.2 HCV and TF ............................................................................................................ 134 4.3 Discussion ................................................................................................................... 137 Chapter 5: Summary .................................................................................................................142 5.1 HSV1 gC has evolved to affect FVIIa function .......................................................... 142 5.1.1 gC mimetics ............................................................................................................ 144 5.1.2 gC sequence similarity to FVIIa binding proteins .................................................. 146 5.2 Host TF-deriving enveloped viruses ........................................................................... 151 5.2.1 HIV and HCV TF procoagulant activity ................................................................. 151 5.2.2 Functional advantage of TF expression .................................................................. 151 References ...................................................................................................................................153 Appendices ..................................................................................................................................206 Appendix A ............................................................................................................................. 206 A.1 Derivation of quadratic binding isotherm ............................................................... 206 A.2 FVIIa-R retains most of its FVIIa function............................................................. 209 A.3 Comparison of fits for MST-derived FVIIa-R interactions .................................... 210 A.4 Immunogold centrifugation of HSV1 ..................................................................... 211  xiii  List of Tables  Table 1. Enveloped viruses and TF-expressing cell tropism. ....................................................... 26 Table 2. Binding affinities of sTF-His and sgC-His as derived from linked enzyme kinetics. .... 95 Table 3. Kinetics of FX activation as derived from linked enzyme kinetics. ............................... 96 Table 4. FVIIa-R binding affinities as determined by MST. ...................................................... 113 Table A5. Comparison of fits for MST-derived FVIIa-R interactions ....................................... 210 xiv  List of Figures Figure 1. Coagulation pathway. ...................................................................................................... 5 Figure 2. Crystal structure of sTF. .................................................................................................. 7 Figure 3. Protease-activated receptor activation by clotting proteases. ........................................ 11 Figure 4. Crystal structure of FVIIa. ............................................................................................. 15 Figure 5. Soluble TF/FVIIa crystal structure. ............................................................................... 17 Figure 6. Crystal structure of FXa in complex with an inhibitor, FX-2212. ................................ 20 Figure 7. Schematic representation of FX and FXa. ..................................................................... 21 Figure 8. Prothrombin conversion to thrombin by FXa. ............................................................... 22 Figure 9. Structural diagram of a basic herpesvirus. .................................................................... 28 Figure 10. Schematic illustration of HSV1 gC. ............................................................................ 32 Figure 11. Multiple functions of HSV1 gC. ................................................................................. 34 Figure 12. Hypothesized model of gC-TF interaction. ................................................................. 42 Figure 13. Characterization of antigens on HSV1. ....................................................................... 47 Figure 14. Recombinantly expressed sgC-His and sTF-His interaction with a Ni-chelating membrane. ..................................................................................................................................... 49 Figure 15. Recombinant sgC-His antigen confirmation by protein stain and Western blot. ........ 53 Figure 16. Recombinant sTF-His antigen confirmation by protein stain and Western blot. ........ 55 Figure 17. Sample adsorption to grids for EM. ............................................................................ 57 Figure 18. Centrifugation method for capturing and concentrating viruses in plasma. ................ 61 Figure 19. Immunogold capture of viruses from plasma for EM. ................................................ 63 Figure 20. Schematic of non-enzymatic and enzymatic reactions involved in FVIIa-mediated FX activation. ...................................................................................................................................... 66 xv  Figure 21. MST basic theory and set-up. ...................................................................................... 72 Figure 22. sTF-His cleavage by CNBr removes the C-terminus His-tag. .................................... 79 Figure 23. Initiators of coagulation found on the virus surface. ................................................... 82 Figure 24. gC enhances TF-dependent FX activation on the virus envelope. .............................. 84 Figure 25. Difference in viral TF expression does not explain gC-mediated FX activation. ....... 86 Figure 26. Addition of sgC-His to HSV1 enhances FVIIa-mediated FX activation. ................... 87 Figure 27. sgC-His effect on FX activation by relipidated TF. .................................................... 89 Figure 28. Ni-chelating lipids enhance sgC-His and sTF-His cofactor function. ......................... 91 Figure 29. sgC-His and sTF-His binding to NiPCPS or NiPC vesicles........................................ 93 Figure 30. Binding of sgC-His or sTF-His to FVIIa in the presence of NiPCPS or NiPC. .......... 94 Figure 31. sgC-His-induced plasma clotting. ............................................................................... 98 Figure 32. Antibodies against HSV1 exist in normal human pooled plasma-derived IgG. ........ 100 Figure 33. Crosslinking of proteins on HSV1 surface by DTSSP. ............................................. 101 Figure 34. Crosslinking of purified sgC-His and sTF-His on NiPCPS vesicles. ........................ 102 Figure 35. Interaction between sgC-His-R and sTF-Cys was probed using MST...................... 105 Figure 36. sgC-His can bind to FVIIa in addition to FX. ........................................................... 109 Figure 37. sTF-His binding to FVIIa-R is enhanced by FX. ...................................................... 110 Figure 38. sgC-His interacts with FVIIa with enhancement by FX. .......................................... 112 Figure 39. sTF-His and sgC-His enhancement of FVIIa amidolytic activity. ............................ 114 Figure 40. Plasma coagulation by HSV1 is TF-dependent. ........................................................ 116 Figure 41. GAGs are procoagulant in FX chromogenic assay. .................................................. 117 Figure 42. TF on DENV enhances FVIIa-mediated FX activation. ........................................... 129 Figure 43. DENV-initiated coagulation is TF-dependent. .......................................................... 130 xvi  Figure 44. TF and aPL are found on DENV. .............................................................................. 132 Figure 45. HIV harbors TF on its viral envelope. ....................................................................... 135 Figure 46. Patient-derived HIV displays a heterogenous phenotype. ......................................... 136 Figure 47. Patient-derived HCV expresses TF on its surface. .................................................... 138 Figure 48. Phenotypes of HCV particle population. ................................................................... 139 Figure 49. Working model of gC interaction with FVIIa/FX. .................................................... 143 Figure 50. Sequence homology between HSV1 gC and HAdv5 hexon FX-binding region ...... 145 Figure 51. gC sequence comparison to TF. ................................................................................ 147 Figure 52. Model of gC homology with the conserved sequence of TF interaction with FVIIa. 148 Figure 53. gC sequence comparison to EPCR. ........................................................................... 149 Figure 54. Model of gC homology with the structure of EPCR interaction with Protein C. ...... 150 Figure 55. FVIIa-R retains its function when bound to sTF-His or sgC-His. ............................ 209 Figure 56. Immunogold differential centrifugation of HSV1. .................................................... 211  xvii  List of Abbreviations ADE, antibody-dependent enhancement  AIDS, acquired immunodeficiency syndrome aPL, anionic phospholipid APMSF, p-amidinophenylmethylsulfonyl fluoride aPTT, activated partial thromboplastin time ART, anti-retroviral treatment  BCA, bicinchoininic acid Bmax, amount of ligand bound at receptor saturation BSA, bovine serum albumin CCR5, C-C motif chemokine receptor 5 CD4, cluster of differentiation CI, confidence interval CS, chondroitin sulfate CTI, corn trypsin inhibitor CXCR4, C-X-C chemokine receptor type 4 ddH2O, distilled and de-ionized water DENV, dengue virus DIC, disseminated intravascular coagulation DMF, dimethylformamide DMSO, dimethyl sulfoxide DNA, deoxyribonucleic acid DTSSP, 3,3'-dithiobis(sulfosuccinimidyl propionate  xviii  ECL, enhanced chemiluminescence reagent EDTA, ethylenediamine tetraacetic acid EGF, epidermal growth factor EM, electron microscopy EPCR, endothelial cell protein C receptor F, factor FII, prothrombin FIIa, thrombin FVII, factor VII FVIIa, factor VIIa FVIII, factor FVIII/DP, factor VIII-deficient plasma FX, factor X FXa, factor Xa GAG, glycosaminoglycan gC, glycoprotein C GdnHCl, guanidine hydrochloride Gla, γ-carboxyglutamate HA, hyaluronic acid HBS, HEPES buffered saline HCV, hepatitis C virus His-tag, histidine tag HIV, human immunodeficiency virus xix  HRP, horseradish peroxidase HSPG, heparan sulfate proteoglycans HSV1, herpes simplex virus type 1 HSV2, herpes simplex virus type 2 HVEM, herpesvirus entry mediator  IU, international units Kd, equilibrium dissociation constant Km, Michaelis constant LMWH, low molecular weight heparin MAPK, mitogen-activated protein kinase MARCO, macrophage receptor with collagenous structure  MST, microscale thermophoresis NAPc2, nematode anticoagulant peptide c2 NHS, N-hydroxysuccinimide  DGS-NTA-Ni, the nickel salt of 1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid) succinyl NP, normal human pooled plasma OR, odds ratio PAR, protease activated receptor PC, phosphatidylcholine PDB, protein data bank PDI, protein disulfide isomerase PI3K, phosphatidylinositol-3 kinase  xx  PS, phosphatidylserine PT, prothrombin time PTA, phosphotungstic acid RNA, ribonucleic acid SD, standard deviation SEM, standard error of the mean sgC-His, soluble gC with His5 tag (gCΔ457t) sTF-Cys, soluble tissue factor with N-terminal cysteine sTF-His, soluble tissue factor with N-terminal His6 tag TCS, Transitive Consistency Score TF, tissue factor TFPI, tissue factor pathway inhibitor TNF, tissue necrosis factor tPA, tissue plasminogen activator UFH, unfractionated heparin Vmax, maximal rate of reaction vWF, von Willebrand Factor  xxi  Acknowledgements I must give thanks to Ed, who will always remain a role model in his diligence towards good science, his patience towards my endless questions and ignorance, as well as his keenness in good communication skills. I owe thanks to Michael, whose questions, even the simplest, has guided me along my studies. I also can’t argue that his favorite catchphrase “Stay positive, Unikitty©” has grown on me. Scott, thank you for your help and down-to-earth view on everything. To the current and previous members of the lab (Michael, Scott, Simon, Kat, Frank, Rolinda, Tseday, Ulli) that have supported me, much appreciation. Fred, I will not name my firstborn after you, but I give thanks for your advice and ideas. I would like to thank Val Smith of the Ross MacGillivray lab for her expertise with chromatography equipment during the early days of my protein purification woes . I also extend my thanks to Dr. James Morrissey who has graciously offered materials and knowledge that greatly helped my research. I greatly appreciate the time, effort and expertise my committee chair and members, Drs. Côté, Kizhakkedathu and Horwitz, have given me. Many thanks to the CBR office members, Hana and Anna, for making CBR a wonderful place to grow ourselves as scientists and people. I offer my enduring gratitude to the faculty, staff and my fellow students at UBC, who have inspired me to continue my work in this field. Special thanks to my mom, dad and brother, who have supported me throughout my years of education. To my family, I apologize for the rather haphazard schedule throughout my studies and have always appreciated the “free” bed, meals and rides. Shouts out to the de-stressing my good friend, Tony, has shared with me. And lastly, I would not have survived grad school without the support of my undergrad twin, Irene. Even though we are 4030 km away from each other, it is almost like we were working side-by-side to finish grad school. I’m always rooting for you too!xxii  Dedication  I dedicate this dissertation to my family who have supported my journey into the great realm of science.  Special dedication to my Taekwon-Do master, Benny Wong, who has inspired me to constantly improve not only for myself but for others and has taught me to maintain an indomitable spirit. The final dedication goes to Irene Lin, as your support has been instrumental in my pursuit of a career in science.1  Chapter 1: Introduction 1.1 Thesis Rationale Viral infections are major health problems worldwide. Furthermore, many viruses have been linked to cardiovascular diseases, including myocardial infarction, atherosclerosis, thrombosis and bleeding. This is important because the leading causes of death worldwide are cardiovascular diseases [1]. Considering the burden and ubiquity of viruses and cardiovascular diseases, extensive research has been invested in trying to understand their pathologies or pathophysiologies as well as to improve or discover treatments for these diseases. Viral infections are communicable; thus, the spread and impact of these diseases has been further accelerated by increased travel and migration of zoonotic reservoirs. This affects not only the immediate population but also blood banks where asymptomatic individuals who are unaware of their infection(s) and may spread pathogens through blood donations or donor-related products. Our work aims to address the issue of a lack of pan-specific antivirals by demonstrating that a common protein exists on a range of viruses and may be the basis of their links to cardiovascular diseases. This protein is tissue factor (TF), the main physiological initiator of coagulation. TF has been previously demonstrated to be involved in numerous disease states involving a haemostatic response and inflammation including virus infection, thrombosis and cancer. Studies from the Pryzdial laboratory have revealed the presence of TF on several herpesviruses [2]. Here, my thesis work focuses on virus-initiated coagulation and its relationship with TF.      2  1.2 Infectious diseases and coagulation 1.2.1 Thrombosis Haemostasis is the physiological process of restoring or maintaining blood flow. This can be achieved by preventing blood loss from an injury through blood coagulation or restoring blood flow by dissolving a blood clot through fibrinolysis. When a blood clot is formed and hinders blood flow, this is known as thrombosis. This pathological condition can lead to poor oxygen and nutrient delivery to organs or tissues as well as inflammation. Thrombosis is divided into arterial and venous [3]. Arterial thrombi are typically formed by platelets and their interactions with abnormalities in the arterial walls (e.g. rupture of an atherosclerotic plaque). Arterial thromboses include ischemic heart disease (or coronary heart disease) and ischemic stroke. Venous thrombi, on the other hand, form in the veins and are mainly comprised of fibrin and red blood cells. Examples of venous pathology are deep-vein thrombosis and pulmonary embolism. Systematic analyses of global causes of death put thrombosis as the leading causing of death in 2010, accounting for 1 in 4 deaths worldwide, increasing from 1 in 5 since 1990 [1].  Well known risk factors associated with vascular disease are heredity, hypertension, obesity, diabetes, surgery and cancer [4–7]. Our work and others indicate that virus infection should also be on this list. Most risk factors are non-communicable, thus do not pose a threat to the public. However, infectious diseases are communicable and are major risks to the public and the healthcare system (e.g. blood donations and organ donations). Although blood transfusions and blood products are routinely screened for typical infectious diseases, the scope of detection is limited and may not detect all potential infectious agents [8].  3  1.2.2 Infectious diseases in inflammation Infectious diseases, including viruses, bacteria, fungi and prions, affect us constantly. It is estimated that infectious diseases cause nine million deaths worldwide each year [9]. As for survivors, many are left with long-term disabilities. Most if not all virus-caused deaths are attributable to inflammation. Inflammation is a response by the body’s immune system to specific stimuli, e.g. pathogens or injury, where the body attempts to restore homeostasis by healing tissues or removing pathogens. Inflammation was once thought to be separate from haemostasis. Yet, the two processes have similar activation stimuli as well as regulatory systems [10,11] and the multifunctional initiator of coagulation, tissue factor (TF), is the molecular bridge. Pathogens can induce vascular cells to initiate coagulation, which is quickly followed by local inflammation to recruit immune cells. The reverse scenario can also occur, where pathogens trigger immune cells to initiate coagulation. Increasing bodies of evidence intertwine inflammation and haemostasis, wherein more research is needed to understand the roles of specific pathogens in this interplay. Some pathogens such as herpesviruses establish a latent infection, with no displayed symptoms or a slow manifestation over the years. Thus, it is hard to gauge the burden of these diseases without first identifying the causative agents and understanding how they contribute to symptoms or other diseases. Our laboratory has demonstrated that herpes simplex virus type 1 takes advantage of the interplay between coagulation and inflammation to enhance infection [12,13]. This is achieved through clotting protease-mediated activation of cell receptors.   4  1.3 General coagulation overview Blood clot formation may be activated by two pathways: extrinsic and intrinsic pathways (Figure 1). The induced availability of TF triggers clot formation via the extrinsic branch of the pathway. TF is a transmembrane protein that binds to the plasma protein factor (F) VIIa in the presence of anionic phospholipid (aPL) and Ca2+. This TF/FVIIa/Ca2+/aPL complex is known as the extrinsic tenase and is responsible for proteolytic activation of FX to FXa.  The intrinsic pathway is initiated by a negatively charged surface, such as aPL membranes, or collagen exposure that allows for a cascade of protease activation involving factors XIIa, XIa, IXa and VIIIa. Like the extrinsic tenase, FIXa/FVIIIa/Ca2+/aPL is the intrinsic equivalent activator of FX to FXa. Although the extrinsic pathway is the physiological initiator of FX activation, the intrinsic pathway accelerates the accumulation of FXa. The extrinsic tenase can activate FIX to FIXa to amplify FX activation. FXa is capable of activating FVIII but only when in complex with the extrinsic tenase [14].  The convergence of the extrinsic and intrinsic pathways of physiological coagulation is referred to as the common pathway. FX activated by extrinsic and/or intrinsic pathways proceeds to cleave prothrombin (also known as FII) to the activated form, thrombin (or FIIa). FXa cleavage of prothrombin is greatly enhanced by its cofactor FVa. FV is cleaved to FVa by FXa and thrombin, with the former being critical for plasma clotting [15,16]. The resulting thrombin generation proceeds to activate fibrinogen to fibrin, which in turn is crosslinked by FXIIIa to stabilize the clot. To overcome the coagulation inhibitors intrinsic to plasma, feedback amplification is important. This is achieved through multiple mechanisms, including FXIa and FVa activation by thrombin [15,17].   5   Figure 1. Coagulation pathway. Coagulation is initiated by the extrinsic pathway and amplified by the intrinsic pathway. Through a series of interactions, the intrinsic tenase complex (FIXa/FVIIIa) or extrinsic tenase complex (TF/FVIIa) is formed. The tenases participate in the common pathway where FX is activated to FXa and subsequently activate prothrombin (FII) to thrombin (FIIa). A fibrin clot can then be formed. Red dashed arrows represent feedback or auxiliary mechanisms to amplify procoagulant proteases so that coagulation can occur.   6  Two clinical tests are typically performed to assess the function of the coagulation system in an individual. Prothrombin time (PT) measures the integrity of the extrinsic pathway and can determine if there are bleeding problems or if certain administered anticoagulants are effective. Activated partial thromboplastin time (aPTT) examines the intrinsic pathway and can also determine bleeding problems such as hemophilia and can be used to test if the anticoagulant, heparin, is working. These tests were derived from scientific methodologies and were used by the Pryzdial laboratory to investigate the interactions between viruses and coagulation. As previous work has shown, herpesviruses can initiate coagulation through both extrinsic and intrinsic pathways [2,18]. Our focus is on TF of the immediate extrinsic pathway.   1.3.1 Tissue Factor 1.3.1.1 Structure Tissue factor (TF) is a 263 amino acid, 47 kDa transmembrane protein. Nemerson’s group first isolated and purified the protein in 1981 [19]. This allowed for the cloning and gene sequencing of TF [20] with the first crystal structure of the soluble form (sTF) being solved 7 years later (Figure 2) [21]. TF is composed of three domains: cytoplasmic (residues 243-263), transmembrane (residues 220-242) and extracellular (residues 1-219) [20,22]. There are several crystal structures of TF and each structure can be obtained from the Research Collaboratory for Structural Bioinformatics protein data bank (PDB) e.g. PDB: 1DAN used in Figure 2. The N-terminal extracellular domain contains two fibronectin type III domains linked by a single Pro102 to Asn107 linker, joined at an angle of 125° (Figure 2) [21]. These two domains both contain a three-stranded and a four-stranded anti-parallel sheet that are folded into a β-sandwich.  The  interface  formed  between the hydrophobic cores of the two fibronectin domains 7    Figure 2. Crystal structure of sTF. Solid ribbon representation of the sTF ectodomain with the two fibronectin domains connected by the Pro102-Asn107 linker to an angle of 125 degrees. Residues involved in TF/FVIIa interaction run along a strip on the entire TF molecule (blue ball and chains) as determined from the TF/FVIIa structure. The C-terminus contains multiple important residues for FVIIa substrates FX or FIX recognition (purple ball and chains). The Lys165 and Lys166 are also important in FVIIa substrate recognition and are susceptible to amine-reactive crosslinkers (orange ball and chains). The thioester bond between Cys186 and Cys209 control the encryption of TF function (red ball and chains). A form of soluble TF with a GGAAG spacer and a His6 tag at the C-terminus was used in this study. A region with several putative membrane-interacting residues are found in the C-terminus ectodomain (brown box). The membrane-interacting site is theoretical as this structure was based on a solubilized form of recombinant TF lacking the transmembrane domain. The structure was obtained from the PDB code ID: 1DAN.   8  results in a large hydrophobic patch [23]. This patch contributes to a rigid structure for FVII or FVIIa binding (Figure 5). There are two disulfide bonds at Cys49-Cys57 and Cys186-Cys209, the latter being crucial for FVII binding [24].  TF displays high sequence similarity to the conserved class two of the cytokine receptor superfamily [25,26] for which structural comparisons describes immunoglobulin-like modules in a portion of the TF extracellular domain. Unlike the extracellular domain, the structure of the cytoplasmic domain is unknown as the crystal structure of full-length TF remains unresolved.  1.3.1.2 Function in haemostasis TF is vital to life as shown by deletion mutants in mice resulting in death during embryonic development as a consequence of defective neoangiogenesis [27–29]. TF initiates coagulation by several mechanisms. The extracellular domain of TF binds to FVII and enhances its proteolytic auto-activation or activation by one of several other proteases. TF enhances the proteolytic activity of FVIIa towards FX and FIX [30]. Regions on TF interact with the phospholipid membrane [31,32] and conceivably affect function. The phospholipid-interacting domain is adjacent (or near) the FVIIa binding domain of TF. It was found that the lysine residues 165 and 166 are important in membrane, FVIIa and FX association. Crystal structure of soluble TF/FVIIa revealed 21 TF residues that interact with FVIIa [33]. Further mutagenesis studies have demonstrated the importance of Tyr157, Lys159, Ser163, Gly164 and Tyr185 on TF in FVIIa substrate recognition [34]. TF allosteric interaction with FVIIa results in an approximately million-fold increase in the rate of activation of FIX and FX [35], highlighting its importance in haemostasis.  TF also plays an important role in wound healing outside of the initial formation of a blood clot [36,37]. This was evident in mice models of diabetes mellitus, where wound healing is known 9  to be retarded, similar to that in humans with diabetes mellitus [38,39]. Deficiency in TF expression at the wound site in diabetic mice resulted in impaired healing processes and enhanced TF expression by gene transfer accelerated wound closure compared to non-treated diabetic mice [37]. Neoangiogenesis is also important in wound healing in which TF also plays a role [40]. This cell signaling-dependent process is regulated by both extracellular and intracellular domains of TF. While the extracellular domain of TF can mediate FVIIa-dependent activation of protease-activated receptor (PAR) signaling, the cytoplasmic domain of TF is involved in cell signaling [41–43]. A secreted form of the extracellular domain of TF, produced through alternative splicing, has also been demonstrated to connect integrins αvβ3 and α6β1 [44]. This resulted in the induction of cell signaling independent of PARs or the cytoplasmic domain of TF, leading to endothelial cell migration and blood capillary formation.  1.3.1.3 Protease-activated receptors and TF function PARs are G protein-coupled receptors and activated by a variety of proteases including those involved in haemostasis. There are four known PARs, numbered from 1 through 4 [45,46]. These integral membrane proteins possess an activation peptide that upon cleavage, exposes a new N-terminal sequence that acts as a tethered ligand (Figure 3). The tethered ligand activates the PAR by binding unto itself. TF/FVIIa can activate PAR-2; FX can activate PAR-1 and PAR-2; and thrombin can activate PAR-1, PAR-3 and PAR-4. Activation of PARs has a wide range of effects including inflammation, apoptosis and angiogenesis. TF/FVIIa-dependent activation of PAR-2 has been linked to angiogenesis and apoptosis [27,47–50]. Angiogenesis is stimulated through the increased production of pro-angiogenic factors such as vascular endothelial growth factor [51] and interleukin-8 [52]. PAR-2 10  phosphorylation downstream signaling includes phosphatidylinositol-3 kinase (PI3K) signaling and mitogen-activated protein kinase (MAPK) signaling pathways [53,54]. These pathways are pro-malignancy and stimulate oncogenic protein synthesis, thereby inhibiting apoptosis in cancer cells. TF/FVIIa-mediated activation of PAR-2 has been demonstrated to enhance virus infectivity [12,13]. Furthermore, PAR-2 activation can result in phosphorylation of TF cytoplasmic domain residue Ser253 by protein kinase C and subsequently allows phosphorylation of Ser258 [41,43]. Ser253 phosphorylation and PAR2 signaling are more efficient when TF is bound to β1-integrins [55]. Ser253 phosphorylation is involved in triggering TF incorporation into microvesicles and their release [41]. This could explain why cancer patients have increased TF-expressing microparticles and increased associated risk of venous thromboembolism [56]. TF/FVIIa-mediated PAR-2 activation also contributes to tumor angiogenesis and growth in breast cancer development [57,58].   1.3.1.4 Function(s) in non-haemostatic settings Besides the well-established role of TF in haemostasis, TF has been implicated in inflammation. Mice expressing TF mutants with defunct cytoplasmic tail domains displayed reduced severity of arthritis, reduced synovial cytokine mRNA expression and impaired adaptive immune responses [59]. TF expression is also elevated in other inflammatory diseases such as ulcerative colitis [60] and multiple sclerosis [61], further stressing the importance of TF in inflammation.   11      Figure 3. Protease-activated receptor activation by clotting proteases. PARs are activated via cleavage of the extracellular tethered ligand by multiple serine proteases. PAR-2, depicted here from PDB code 5NJ6, is activatable by TF/FVIIa complex and FXa. The newly exposed N-terminus tethered ligand can bind to the core of the PAR, resulting in activation and cell signaling through G-protein coupled signaling.  12  1.3.1.5 Regulation of TF function TF expression can be found in a wide range of cells, including vascular smooth muscle cells, pericytes, monocytes, astrocytes and cardiomyocytes (partially listed in) [62–65]. Yet, its procoagulant activity is not always present. The finding that surface exposed TF in dermal vessels was bound to FVII in the absence of injury, suggested that TF function exists in an “encrypted” form [66]. TF has a disulfide bond at Cys186-Cys209 within the C-terminal domain (Figure 2) that exists in an allosteric configuration. Allosteric disulfide bonds have been demonstrated to be involved in controlling protein function via redox state changes of the cysteine residues [67]. Since TF binding to FVIIa and subsequent FX activation is compromised when the disulfide bond at Cys186-Cys209 was reduced, the hypothesis was that the redox state of Cys186-Cys209 governs TF function on the cell surface [68]. The importance of Cys186-Cys209 led to the investigation of protein disulfide isomerase (PDI), a subcellular protein that catalyzes the formation of disulfide bonds between cysteine residues on proteins. While the role of PDI is still debated, most recent studies observe PDI expression enhancing TF-dependent thrombin generation [69]. However, alterations in membrane phospholipid asymmetry have also been implicated in the description of TF function.  Further regulatory mechanisms of TF function include tissue factor pathway inhibitor (TFPI). TFPI blocks TF/FVIIa function when bound to FXa, thereby stopping the initiating pathway [70]. This occurs in two steps: 1) TFPI binds to and inhibits FXa via its Kunitz-type homology domain 2; and 2) TFPI/FXa binds to and inhibits TF/FVIIa via the Kunitz domain 1 [71,72]. Inhibition of TF/FVIIa by TFPI is not calcium-dependent but is most efficient when TFPI-FXa is in complex with FV, calcium and aPL [70]. 13  Proinflammatory cytokines, such as interleukin-1β and TNF-α, can increase TF expression on endothelial cells and/or monocytes in vitro [73,74]. Inflammatory reactants (proteins or markers that increase or decrease following an inflammatory response), such as C-reactive protein, can induce TF expression on human peripheral blood monocytes but not human umbilical vein endothelial cells [75]. This suggests that TF expression can be regulated by different stimuli depending on the cell type. In cancer biology, several defined pathways have been implicated in upregulating TF expression [76]. Some examples include epidermal-to-mesenchymal transformation and transforming growth factor-β signaling [77,78]; hypoxia-induced signaling of early growth response protein-1 [79]; and constitutive activation of PI3K and MAPK signaling [53,54].   1.3.2 Factor VIIa 1.3.2.1 Structure Factor VII (FVII) is a serine protease zymogen synthesized in the liver and circulates in the plasma. The zymogen circulates the plasma at 10 nM, whereas its activated form FVIIa circulates at 0.1 nM. FXa is the most potent activator for FVII but it can also be activated by FVIIa, FVII-activating protein and other proteases [80–83]. Cleavage at Arg212 exposes the protease domain upon a conformational change with the activating peptide remaining bound by a single disulfide chain. However, FVIIa specific activity is considerably low and has been described as zymogen-like in solution [84]. The light chain, comprised of a γ-carboxyglutamate (Gla)-rich domain and two epidermal growth factor (EGF) homology domains, is covalently linked to the heavy chain which contains a single protease domain (Figure 4). 14  The Gla domain is a membrane binding motif with many Gla residues, where the synthesis is dependent on the γ-carboxylase cofactor, vitamin K, for the post-translational conversion of Glu to Gla. Proteins containing this modification are collectively known as vitamin K-dependent proteins. Membrane interaction occurs with aPL such as phosphatidylserine and phosphatidylethanolamine. For FVII, these interactions are of weak affinity compared to other vitamin-K-dependent proteins in coagulation, FIX, FX and prothrombin. Rather, FVII preferentially binds to phosphatidic acid [85]. At supraphysiological concentrations of calcium (>2.5 mM), 7 calcium ions bind to the Gla domain of FVIIa, resulting in a conformational change of the Gla domain [33]. However, in plasma, free calcium (1.25 mM) and magnesium (0.6 mM) occupy the Gla domain of FVIIa (4 and 3 divalent cations, respectively) [86]. Although the conformational change induced by the two metal ion occupancies are dissimilar, the proteolytic function of the two are identical. Two EGF domains, EGF1 and EGF2, serve to tether the protease domain of FVII to an appropriate phospholipid surface via the Gla domain. The EGF domains are tethered by a single stabilizing hydrogen bond between the main-chain carbonyl of Lys143 on EGF1 and the main-chain amino group (Nδ2) of Asn153 on EGF2 [87]. EGF1 contains a calcium binding site at the N-terminal end. In the protease domain there is a calcium binding site which is essential for amidolytic activity. The protease domain of FVIIa shares structural homology with trypsin, similar to other coagulation serine proteases, including FX, FIXa, and thrombin, as well as the anticoagulant serine protease, activated protein C [88].     15   Figure 4. Crystal structure of FVIIa. The heavy chain of FVIIa, consisting the protease domain (purple), is connected to the light chain comprising the EGF2 (gray), EGF1 (blue) and Gla (brown) domains. Several divalent cation binding (gray spheres for calcium and black spheres for calcium/magnesium) sites are depicted: one in the protease domain, one in the EGF1 domain and seven in the Gla domain. The active site contains the catalytic triad: His57, Asp102 and Ser195 (green circle and inset). Arg212 cleavage of FVII results in a single Cys135-Cys262 thioester bond joining the heavy and light chain (posterior, black circle). The EGF2 and EGF1 domains are connected by a single stabilizing hydrogen bond (light green chain). Arg36 is highlighted for its importance in FX and TFPI recognition. Structure derived from PDB code 1DAN. Magnesium binding sites derived from PDB code 2A2Q.  16  1.3.2.2 Function Although humans can live with low levels of FVII [89], complete absence of FVII, as described in mutant mice, results in hemorrhagic lethality in the perinatal period [90,91]. This highlights the importance of FVIIa in haemostasis. The primary function of FVIIa is the catalytic conversion of FX to FXa. FVIIa binding to TF increases the catalytic efficiency of FX activation. This is achieved by structural rearrangement of FVIIa, allowing for optimal substrate binding to exosites on TF and/or FVIIa. Although TF can bind FVII and FVIIa (Figure 5), only when FVIIa has its Arg212-Ile213 scissile bond broken and subsequently the Ile213 allowed to form a salt bridge with Asp244, can TF induce FVIIa rearrangement [84]. Interestingly, TF binding does not cause the catalytic oxyanion hole to form but rather, FX binding to the exosite [92,93] results in the formation of the oxyanion hole [86]. The Gla domain of FVIIa has been shown to be dispensable for its amidolytic activity [94] but is important for FX activation [32,94]. Arg36 of FVIIa Gla domain connects with TF residues 159 to 164 with an extensive network of hydrogen bonds [95]. Arg36, along with TF residues 165 and 166, were proposed to be important in FX substrate as well as TFPI recognition [96], though some opposing evidence or inconsistencies have been noted [95]. In addition to a procoagulant role, TF/FVIIa also is also important in cell signaling (Chapter 1.3.1.3). FVIIa can also participate in cell signaling in the absence of TF through interactions with endothelial cell Protein C receptor (EPCR) [97]. Activation of PAR-1 signaling by EPCR/FVIIa resulted in suppression of tissue necrosis factor (TNF)-α-induced inflammation and LPS-induced endothelial activation and inflammation. The precise mechanism is not yet understood but evidence pointed to an early step in TNF receptor 1-mediated signaling [97].   17   Figure 5. Soluble TF/FVIIa crystal structure. FVIIa heavy chain (purple) connected to light chain (blue) is bound to soluble TF, comprised of the N-terminus and C-terminus ectodomains (light green and dark green, respectively). Calcium binding is depicted as gray spheres. A phospholipid membrane is shown as a reference to orientation on a membrane. Structure is derived from PDB code 1DAN.  18  The importance of FVIIa is further accentuated by its use as a therapeutic for treating hemophiliacs with FVIII inhibitors and other coagulation disorders (e.g. acquired hemophilia, and congenital FVII deficiency). Its indisputable coagulation ability has also warranted its use in off-label situations such as surgery and critical care management of bleeding [98,99].  1.3.2.3 Regulation of FVIIa expression and function FVII circulates the plasma at ~0.5 µg/mL or ~10 nM whereas its activated form exists at less than 1% of total FVII(a) at ~0.1 nM [100]. FVII has a shorter half-life than any other clotting factor, of ~5 hours [101,102]. Interestingly, other clotting proteases tend to be cleared very quickly upon activation (~3 min). In contrast, the half-life of FVIIa is 2 to 4 hours [103,104]. This would suggest that the clearance of FVII/FVIIa is due to a specialized mechanism by the liver. However, studies suggest high membrane turnover on primary liver cells and yet unknown factors contribute to the slower clearance rate of FVIIa to other activated clotting proteases [105]. Freely circulating plasma FVIIa is inhibited by antithrombin III in the presence of heparin at a slower rate than the TF/FVIIa complex [106]. As described in Chapter 1.3.1.5, FVIIa activity is inhibited by TFPI.   1.3.3 Factor X 1.3.3.1 Structure Like FVII, FX is a serine protease zymogen that is synthesized in the liver. The protein consists of a 139 amino acid residue light chain and a 306 amino acid residue heavy chain linked through a single disulfide bond (59 kDa). The light chain is distinguished by three structural domains: A N-terminal Gla domain, a short hydrophobic domain, and similar to FVII, two EGF-homology domains (EGF1 and EGF2). The heavy chain is comprised of a single protease domain. 19  There are no crystal structures of the zymogen form, but inhibited FXa has several crystal structures available (Figure 6, PDB code 1XKA) [107,108].  Prior to FX secretion into circulation, the Arg140-Lys141-Arg142 tripeptide in the heavy chain is excised [109]. The mature FX consists of an activation peptide (Ser143-Arg194) [110] at the N-terminus of the protease domain, which is irreversibly connected to the light chain by a single disulfide bond between Cys132 and Cys302 [111]. Similar to FVII and FIX, the protease and EGF1 domains have a single calcium binding site [107]. The Gla domain of FX is linked to the EGF domains through several aromatic residues and contains 11 Gla residues. FX binds to aPL via the Gla domain at nM affinity, in contrast to FVIIa (~0.2 µM and 15 µM, respectively) [112]. In models of sTF/FVIIa/FXa, the Gla domain of FX is depicted as interacting with both FVIIa Gla domain and TF domain [113–115]. 8 calcium ions are coordinated by the Gla residues although Asp residues may also participate (Figure 6, PDB code 1IOD) [116].  1.3.3.2 Function Activation of FX by the extrinsic or intrinsic tenase through cleavage of Arg194-Ile195 releases the 52-residue activation peptide [117]. FXa can then assemble with FVa, calcium and aPL to form the prothrombinase complex, resulting in an ~278,000-fold increase in FXa-mediated prothrombin activation rate [96]. Prothrombin is activated by FXa-mediated cleavages at both Arg271 and Arg320 (Figure 8) [92]. FXa/calcium/aPL predominantly cleaves Arg271 to form prethrombin before cleaving Arg320. FXa/FVa/calcium/aPL, on the other hand, first cleaves Arg320 to form meizothrombin followed by Arg271 cleavage to form fully active thrombin (α-thrombin). Although both cleavage sites are accessible [118], the presence of a cofactor shifts bond cleavage preference. 20     Figure 6. Crystal structure of FXa in complex with an inhibitor, FX-2212. FXa inhibited with an active site inhibitor, FX-2212, in the presence of calcium (PDB code IXKA). The heavy chain (red) has the protease domain sitting on top of the light chain (orange) consisting of EGF2, EGF1 and Gla domain. A single calcium ion is bound in the protease domain. Similar to FVIIa, a calcium binding site is found in the EGF1 domain (black dot, inserted from PDB code 1XKB). The Gla domain is missing from the structure and is depicted separately in the black box with calcium ions depicted in gray (PDB code 1IOD).   21         Figure 7. Schematic representation of FX and FXa. (A) Premature form of FX includes a tripeptide linking the activation peptide (AP) to the light chain. The light chain is also linked to the heavy chain via a single disulfide bridge between Cys132 and Cys302. (B) During or after secretion into blood plasma, the tripeptide is cleaved to form mature FX. This results in the AP being linked to the heavy chain by a single Arg194-Ile195 bond. Proteolytic cleavage of this bond by tenases results in (C) the active form, FXa.   22     Figure 8. Prothrombin conversion to thrombin by FXa. (A) Prothrombin consists of a Gla domain (G), two kringle domains (K1 and K2), and a trypsin-like serine protease domain that contains a light chain (LC) and a heavy chain (HC). Two different cleavage scenarios can occur depending on whether or not FXa/Ca2+/aPL is in complex with the cofactor FVa. (B) In the absence of FVa, Arg271 is predominantly cleaved, releasing the G, K1 and K2 domains, to form prothrombin. (C) In the presence of FVa, Arg320 is predominantly cleaved to form meizothrombin, where the LC remains anchored to the G, K1 and K2 domains. (D) The final cleavage of the non-cleaved Arg271 or Arg320 results in the fully active form, thrombin or α-thrombin. The LC and HC are held together by a single disulfide bond.   23  FXa is also involved in amplification of its own zymogen activation through FVII [119] and FVIII activation [120]. FV can be activated by FXa to increase FXa-mediated production of thrombin [120]. Furthermore, FX switches from a procoagulant state to an anticoagulant state through plasmin cleavage into FXa33/13 when bound to aPL resulting in fragments with C-terminal lysines [121–123]. C-terminal lysines are important in the degradation of the clot (fibrinolysis) as cofactor binding sites for tissue plasminogen activator and plasminogen. The former cleaves plasminogen into plasmin, the physiological protease responsible for fibrinolysis. PAR-1 [124] and PAR-2 [125] are activated by FXa to participate in cell signaling [125]. FXa-mediated PAR-1 and -2 signaling can result in enhanced cell proliferation and migration in fibroblasts and smooth muscle cells, increased pro-fibrotic and pro-inflammatory cytokine production and secretion [126–129]. EPCR interaction with TF/FVIIa/FXa was shown to enhance PAR-1 and PAR-2 activation [130].  1.3.3.3 Regulation of expression and function FX activation is tightly regulated to prevent aberrant coagulation. The activation is localized to areas of vascular damage and is restricted to a phospholipid membrane. TFPI is the physiological inhibitor of coagulation, inhibiting FXa and subsequently inhibiting TF/FVIIa many fold efficient than TFPI alone. Antithrombin is the main serine protease inhibitor of the intrinsic pathway [111]. This serpin forms a 1:1 covalent complex with FXa, thrombin, FIXa, FXIa, and FXIIa. The rate of complex formation is greatly enhanced when bound by heparin [131]. These rates differ depending on the substrate, e.g. FXa inhibition is accelerated by ~1000 fold compared to ~4000 fold for thrombin inhibition [132]. Once FXa is inhibited, it is rapidly cleared from the 24  body within minutes via degradation in the liver [133,134]. FXa activity is also indirectly regulated by activated protein C-mediated proteolysis of FVa [135].  1.3.4 Anticoagulation and fibrinolysis The main topic of the studies presented here is coagulation. It is, however, important to understand that anticoagulant and profibrinolytic factors are working against coagulation. As noted, anticoagulant factors include TFPI, antithrombin and activated protein C. Thrombin generated by FXa can bind to an endothelial cell integral membrane protein known as thrombomodulin [136]. This shifts thrombin from a procoagulant state to an anticoagulant state. The thrombin/thrombomodulin complex activates protein C into activated protein C. Activation of protein C is markedly enhanced when bound to EPCR found on the endothelial cell surface. Once released from EPCR, activated protein C proteolytically inactivates FVa and FVIIIa. The former decreases FXa/FVa prothrombinase assembly and the latter decreases FIXa/FVIIIa tenase assembly. Therefore, FXa activity and FX activation are inhibited, respectively. Fibrinolysis is the degradation of a blood clot. The process involves tissue plasminogen activator (tPA) and plasminogen binding to C-terminal lysines found on the fibrin clot. tPA then cleaves plasminogen to plasmin and the plasmin cleaves the fibrin clot [121]. Cleavages at lysine residues by plasmin can result in additional C-terminal lysines. This allows additional tPA and plasminogen binding, resulting in amplified plasmin generation. Much like coagulation, there are antifibrinolytic factors such as plasminogen activator inhibitor-1 and thrombin-activatable fibrinolysis inhibitor [137]. Thus, it is important for sufficient plasmin to be formed to overcome the antifibrinolytic factors and allow complete clot dissolution.   25  1.4 Enveloped viruses Viruses are obligate parasites that require a host to replicate and sustain itself. Although they fit under this basic requirement, the methods by which a virus achieves this can be very different. As such, virus classification systems were established to categorize viruses to better organize and identify them. The Baltimore classification system is based on the genetic material (DNA or RNA), sense (positive or negative), strandedness (single or double) and replication strategy. The viruses can then be further separated into orders and families depending on their structure, ancestry and target organisms, although this classification is still incomplete.  All viruses have genomic material encased within a protein cage, the capsid. This basic form of a virus is known as a naked or non-enveloped virus. Some viruses are further encased by an outer phospholipid bilayer membrane structure termed envelope. This envelope is derived from the host cell during egress, though how the different viruses obtain their envelope can differ. Enveloped viruses are more prone to degradation compared to non-enveloped viruses, through desiccation and chemical inactivation. However, the envelope imparts the advantage of immune evasion to the virus. Glycoproteins encoded by the virus or host and found on the viral envelope can prevent immune recognition and/or aid in cell recognition and entry. Herpesviruses are a prime example of a large enveloped virus that has evolved several mechanisms on its envelope structure to enhance its survivability. I am most interested in enveloped viruses involved in haemostatic abnormalities and can be replicated in TF-expressing cells. A summary of TF-expressing cells and evidence of in vivo and/or in vitro replication of herpes simplex virus type 1, dengue virus, human immunodeficiency virus and hepatitis C virus is found in Table 1.    26   Table 1. Enveloped viruses and TF-expressing cell tropism.                   HSV1, herpes simplex virus type 1; DENV, dengue virus; HIV; human immunodeficiency virus; HCV, hepatitis C virus   Tissue factor expressing cells Virus replication Astrocytes [65] HSV1 [138] DENV [139] HIV [140] Dendritic cells [141] HSV1 [142] DENV [143–145] HIV [146–148] HCV [149–151] Epithelial cells [152,153] HSV1 [154,155] DENV [156] HIV [146–148,157] HCV [158–160] Endothelial cells [161–164] HSV1 [165–169] DENV [170–172] HIV [173,174] HCV [175] Hepatocytes [176,177] HSV1 [178,179] DENV [180,181] HIV [182] HCV [183] Megakaryocytes [184] HSV1 [185] DENV [186–189] HIV [190,191] HCV [192] Monocytes/macrophages [161,193] HSV1 [194–196] DENV [170,172,197] HIV [198] HCV [199–201] T lymphocytes [202] HSV1 [203–205] DENV [206] HIV [207–209]  HCV [210–212] 27  1.4.1 Herpesviridae The family of viruses, Herpesviridae, consists of double-stranded linear DNA viruses that causes diseases in humans as well as other animals. Of the over a hundred known herpesviruses [213], only eight are known to affect humans. Herpesviruses share common traits; they have an envelope, their large genome (>80 genes) is packed into a icosahedral (T=16) capsid and their capsid is tethered to their envelope by a matrix of proteins and mRNA known as the tegument (Figure 9) [213]. When herpesviruses enter a cell, their genome is inserted into the nucleus wherein the linear DNA circularizes into an episome. At this stage, the virus can either cause a productive infection or remain dormant in a latent state [214]. For some viruses such as Epstein-Barr virus and human herpesvirus 6, latency can be achieved through integration of their genetic material into the host genome [213]. The ability of herpesviruses to establish dormancy in humans allows the viruses to establish lifelong infections. Reactivation of the virus can result in symptoms similar to the initial infection or different disease sequelae. For example, reactivation of Varicella-Zoster virus (chickenpox virus) results in shingles, a painful rash that can cause neural pain more severe than the primary infection of chickenpox.        28        Figure 9. Structural diagram of a basic herpesvirus. Members of the Herpesviridae family share the same basic structure. A bilayer phospholipid membrane, known as the envelope, stores the capsid that is protecting the viral dsDNA genome. There are numerous host- and virus-encoded membrane proteins present on the envelope. A cluster of proteins fills the space between the capsid and the envelope, this viral matrix is known as the tegument.   29  1.4.1.1 HSV1 HSV type 1 (HSV1) or human herpesvirus 1 is of the genus Simplexvirus, along with HSV type 2 (HSV2). HSV1 causes cold sores for which it has developed the moniker, cold sore virus. However, HSV1 can also cause genital herpes, a disease that was once predominantly caused by HSV2 [215]. Other important diseases caused by HSV1 include infectious blindness through keratitis [216], neonatal herpes [217], and encephalitis [218]. According to the WHO, in 2012, HSV1 was estimated to affect 3.7 billion people under the age of 50, representing 67% of the world’s population. A more recent study in 2015–2016 of the United States’ population, observed a prevalence of 47.8% of people aged 14-49. The prevalence increased with age and was more prevalent in females. Primary infection by HSV1 typically begins with attachment to epithelial cells via viral glycoproteins gB and gC binding to heparan sulfate proteoglycans (HSPG). Entry occurs by membrane fusion involving gB, gD, gH and gL with entry receptors: herpesvirus entry mediator (HVEM), Nectin-1 and/or 3-O-sulfated HSPG [154,219,220]. However, depending on the cell type, receptor-mediated endocytosis is preferred [221,222]. In both scenarios, the viral capsid is eventually released into the cytoplasm where it travels to the nucleus for unloading of viral genome and subsequent replication processes. In addition to cell lysis promoting HSV1 spread, HSV1 has developed multiple modes of cell-to-cell spread. Some of these infection modes are not well understood, but include cell-to-cell fusion [223] and tight junction transfer [224]. Eventually, HSV1 infects and lays dormant as an episome in sensory neurons [214].  HSV1 has a slew of virulence factors that contribute to its potent infectivity and extremely wide cell tropism [219,225]. These include multifunctional glycoprotein for attachment and entry, ubiquitously expressed entry receptors, multiple entry mechanisms and diverse spread strategies. 30  As a consequence, HSV1 envelopes have the potential to obtain a wide breadth of host-derived proteins and incorporate them on its envelope [226]. Previous work in the Pryzdial laboratory have identified TF and aPL on HSV1, HSV2 and cytomegalovirus [2], as evidence of direct links to thrombotic pathologies.  1.4.1.1.1 Implications in cardiovascular/haemostatic abnormalities HSV1 involvement in cardiovascular abnormalities was postulated upon discovery of HSV antigen/genetic material in atherosclerotic plaque [227]. Herpesviruses persist as lifelong infections, with frequent reactivation promoting leukocyte and platelet adhesion to the vascular wall, lipid accumulation in smooth muscle cells and thrombin deposition in the periphery. These effects contribute to vascular pathologies [165,228–231]. This led to studies looking into HSV1 prevalence and risk factors [232,233]. The first meta-analysis of 17 studies evaluating the risk of atherosclerosis associated with HSV1 found that there is an increased risk of developing atherosclerosis (OR = 1.77; 95% CI: 1.40–2.23; P < 0.001) in HSV1-positive patients [234]. While these examples provide a retrospective clinical correlation, causal evidence was reported using HSV1 as a targeted cytolytic therapeutic for melanoma cells, which resulted in a high propensity for deep vein thrombosis [235]. Case reports have also detailed the presence of thrombi [236,237] and/or bleeding [238,239] in patients with HSV1 encephalitis. In one study examining predictors of outcome in HSV1 encephalitis, half of patients experienced acute thrombocytopenia and a few cases of hemorrhage in the brain (5%) [238]. 31  1.4.1.2 Glycoprotein C HSV1 encodes multiple glycoproteins on its envelope surface: gB, gC, gD, gE, gG, gH, gI, gJ, gL, gK, and gM. These glycoproteins aid in its virulence. Of these glycoproteins, gC is of utmost interest to me because of its ability to bind and affect FX activation [12,240,241].  1.4.1.2.1 Structure gC is a highly glycosylated transmembrane protein, with molecular weights ranging from 55 kDa in its non-glycosylated form to 130 kDa when fully glycosylated. The extracellular domain of gC contains nine consensus sites for N-glycosyl oligosaccharides and predicted to be 7 to 13 sites for O-glycosyl oligosaccharide [242]. Due to the variability in extent of post-translational modification, gC exists in different functional forms [243]. The mucin-like region of gC comprises the majority of glycosylation sites. This heterogeneity and high glycosylation pose problems for protein crystallization, hence there still lacks a 3-dimensonal gC structure. Nonetheless, a predicted folded structure of gC was proposed based on the derivation of disulfide bond arrangement (Cys127-Cys144, Cys286-Cys347, Cys386-Cys442, and Cys390-Cys419), homologous regions and molecular binding sites and inferred discontinuous antigen folding (Figure 10) [244,245]. These binding sites include heparan binding and C3b binding, important in its function.  1.4.1.2.2 Function gC is a multifunctional transmembrane glycoprotein that is expressed on the virus surface and docks the virus to host cellular sulfated glycosaminoglycans (GAGs), such as heparan sulfate (Figure 11A) and chondroitin sulfate [246–248]. The mucin-like region of gC is important for   32         Figure 10. Schematic illustration of HSV1 gC. Hypothesized structural model of gC based on its disulfide bond arrangements, homologous regions and molecular binding site. N-CHO glycosylation modifications (orange hexagons) and O-CHO glycosylation modifications (small green circles) are predicted at the stated amino acids. The purple dashed region denotes a highly glycosylated mucin-like region. The blue regions numbered I, II, III, and IV represent the important regions for C3b binding. Antigenic sites are shown in dark blue. gCΔ457t (sgC-His) is truncated at the C-terminal amino acid 457 (white arrow). Image from Rux et al. (1996) J Virol. [245].  33  GAG binding, with the deletion mutant (Δ33–123) exhibiting lower affinity to immobilized heparin and heparan sulfate (purple-dashed line, Figure 10) [246].Mutational studies of gC and subsequent binding to isolated heparan sulfate chains had shown that heparan sulfate binding required a cluster of arginine residues surrounding the Cys127–Cys144 disulfide bridge with Ile142 substantially supporting the binding [249]. This region is well-conserved among mammalian herpesviruses, further adding to the importance of this region towards its function as a cell adhesion molecule [242]. A synthetic peptide mimicking the amino acid residues 137 to 151 of gC has been demonstrated to compete with HSV1 binding to GAGs, thereby inhibiting infection [250]. Recently, gC has been discovered to bind with <1 pM affinity to a cell surface scavenger receptor known as macrophage receptor with collagenous structure (MARCO) on keratinocytes [251]. Overexpression of MARCO in keratinocytes resulted in enhanced HSV1 skin infection whereas MARCO-/- mice were less susceptible. Even though the role of gC has been downplayed as non-essential for viral entry [219], removal of gC results in ~10-fold reduction in infectivity in vitro [252] and ~100-fold reduction in vivo [253]. The reduction in infectivity of gC knockout mutants of HSV1 in vivo can be partly attributed to its role in the evasion of host innate immunity and phagocytosis (Figure 11B) [254,255]. Complement protein C3b can bind to the virus and allow for the binding of properdin to stabilize the alternate pathway C3 convertase (C3b and factor Bb) (Figure 11B) [256]. The C5 convertase cleaves complement protein C5 to C5a and C5b, the latter being involved in the formation of the membrane attack complex (MAC). C3b can also serve to opsonize the pathogen, targeting the virus for destruction by phagocytes via recognition by C3b receptor (Figure 11C). gC interrupts  these  complement  functions by binding to C3b via residues 124  to 366 [255],  thereby  34       Figure 11. Multiple functions of HSV1 gC. (A) gC is known to bind to heparan sulfate proteoglycans (HSPG) on target cells as the primary step of infection. The mucin-like domain of gC (33-123) is known to be important in this interaction. (B) Complement protein C3b binding to the virus allows the binding of properdin (P) which stabilizes the C5 convertase (C3b and Factor Bb (not shown)). This allows the binding of C5 which is important for the formation of the membrane attack complex (MAC). (C) C3b can also serve to opsonize the pathogen, targeting the virus for destruction by phagocytes via recognition by C3b receptor. (D) gC binding of C3b prevents binding of C5a and properdin to C3b. (E) gC can also bind the coagulation zymogen FX and affect its activation. Diagram of gC binding to C3b was adapted from Lubinski et al. (1999) J Exp Med. [257].   35  blocking C5 binding (Figure 11D). The mucin-like region of gC, residues 33 to 123, blocks properdin from binding [255]. This results in an accelerated decay of the alternate pathway C3 convertase.  As a possible contributor to vascular lesion development, gC has been linked to monocyte adhesion to HSV1-infected endothelial cells [240]. This effect was dependent on thrombin generation, alluding to a role for gC in coagulation protease activation. Using transgenic expression of gC in murine L cells, FX binding to cell-surface gC was observed (Figure 11E) [240]. Since gC is found on the virus envelope, I investigated FX binding to purified HSV1 and showed that the presence of viral gC created FX binding sites [241]. The mucin-like region of gC was demonstrated to be important for FX binding. Viral gC was also shown to enhance FVIIa-dependent FX activation. Although the Pryzdial laboratory reported that a soluble form of gC (gCΔ457t or sgC, Figure 10) contributes to FX activation without being membrane-tethered, the activity was ~1000-fold higher when combined with the virus [258].    1.4.2 Flaviviridae Members of the Flaviviridae family of viruses are small (~40-60 nm), enveloped, single stranded positive-sense RNA viruses. Their genomes are ~11 kb in size and encode a single long open reading frame that is flanked by a 5’ and 3’ highly structured untranslated region [88]. As such, translation of the viral genome results in a single large polyprotein that is cleaved into mature proteins by host and virus proteases. As the virus genome is positive-sense RNA, the virus encodes its own RNA-dependent RNA polymerase to replicate its genome. The Flavivirus genus consists of many arthropod-borne viruses, important human pathogens include dengue virus, West Nile 36  virus, yellow fever virus, Japanese encephalitis virus, and Saint Luis encephalitis virus. Another important human bloodborne pathogen is hepatitis C virus of the Hepacivirus genus.  1.4.2.1 Dengue virus Dengue virus (DENV) is a bloodborne virus and is the causative agent of dengue break-bone fever, dengue hemorrhagic fever and dengue shock syndrome. The virus is transmitted to humans via Aedes mosquitos, mainly Aedes aegypti. This species of mosquito is prevalent in tropical and subtropical areas, with their habitats growing due to urbanization, global warming and mosquito adaptation. There is an estimated 390 million infections and 24,000 deaths each year and half the world’s population are at risk of infection [259]. Until recently, there were four known serotypes of DENV, aptly named DENV-1, DENV-2, DENV-3 and DENV-4. DENV-5 is a new variant of DENV that was first isolated in the fall of 2013 [260] but as yet known to infect only non-human primates. Similar to influenza, different serotypes are a problem in vaccine development as the main antigen on the virus surface, the envelope protein (E protein), is antigenically distinct on each serotype. Infection with one serotype confers life-long immunity to only that specific serotype. Partial immunity is obtained to some of the other serotypes. Antibodies are directed towards E protein, which shares structural homology between serotypes as well as the other members of the Flavivirus genus, West Nile virus, Zika virus and Japanese Encephalitis virus [261,262]. A secondary infection of a different serotype can culminate into a more severe form of dengue infection due to antibody-dependent enhancement (ADE). ADE of dengue infection occurs when pre-existing antibodies cannot neutralize the virus but instead, enhance monocyte infection via Fc receptor-mediated entry [263,264]. This phenomenon poses a crucial problem in vaccine development. 37  1.4.2.1.1 Implications on cardiovascular/haemostatic abnormalities Thrombocytopenia has been observed in dengue fever and is always seen in dengue hemorrhagic fever [265,266]. Disseminated intravascular coagulation (DIC), a phenomenon wherein small blood clots are formed throughout the body resulting in depletion of platelets and coagulation factors, have been noted in dengue hemorrhagic patients [266–268]. Minor hemorrhagic manifestations of dengue fever may occur in the form of petechiae, epistaxis, and gingival bleeding [265]. Increased vascular permeability and the resulting diapedesis (migration of blood cells through the vasculature) is also seen in dengue fever and more severe forms of dengue infection. Although bleeding is more commonly reported, DENV-associated thrombosis (e.g. deep vein thrombosis) may be under-reported [269–273]. DENV activation of endothelium can result in vascular leakage and TF expression [274–276]. For TF to induce vascular leakage, FVIII has to be depleted, hinting to additional mechanisms in DENV-induced haemorrhage [277,278].  1.4.2.2 Hepatitis C virus Hepatitis C virus (HCV) is the leading cause of liver transplants, hepatocellular carcinoma and liver-related deaths worldwide [279–282]. As HCV can persist in the blood due to several virulence factors and reservoirs (Table 1), this leads to chronic HCV infection. Continued liver damage eventually results in fibrosis, scarring, loss of liver function (cirrhosis) and carcinoma [283]. HCV is transmitted human-to-human through the parenteral route, with humans and chimpanzees being the only known hosts [283]. HCV circulates the blood as free particles or coated by host low-density lipoproteins. Blood donations are routinely tested for HCV, a major concern that had plagued the North American blood system in the 20th century. No vaccine is 38  available for HCV. However, direct acting antivirals are now available with some treatments reaching over 90% cure rate of HCV infected patients [284]. However, treatment is still costly, up to 150,000 USD. Other complications of HCV infection include cardiovascular/haemostatic abnormalities which can persist even after antiviral treatment of HCV [285].  1.4.2.2.1 Implications in cardiovascular/haemostatic abnormalities HCV infections result in liver damage and impaired function. As most coagulation factors are synthesized in the liver, there is a decrease in vital coagulation factors e.g. FX, FVII, FIX, and prothrombin [286,287]. Although lower levels of coagulation factors would indicate hypocoagulability, anticoagulant proteins Protein C, Protein S and antithrombin [285] are also decreased. There is also an increase in FVIII levels in these patients [288]. This could be due to increased von Willebrand Factor (vWF) expression and decreased clearance of vWF [289,290]. The rise in vWF results in lower clearance of FVIII from the blood, which is typically high if not in complex with vWF. High FVIII levels has been associated with increased thrombosis [291]. Interestingly, there have been several case reports of acquired FVIII autoantibodies in patients with HCV [292–296]. Following treatment with directly acting antivirals in patients with hepatitis C virus related cirrhosis, procoagulant and anticoagulant proteins are gradually restored. Yet, patients treated for HCV and had achieved a sustained virologic response (defined as being clear of HCV in the blood at 24 weeks post-treatment) still exhibited impaired platelet aggregation [285]. As disease progresses, viral liver cirrhosis results in hypercoagulability in patients [288,297–299] with HCV patients being up to 1.9 times more likely to suffer from venous thromboembolism than the healthy human population [300]. Liver cirrhosis caused by HCV has also been linked to enhanced expression of monocyte TF [301]. 39  1.4.3 Retroviridae Measuring around 90-140 nm in diameter, retroviruses house double-stranded RNA genomes of about 7-12 kb [302]. These viruses encode reverse transcriptases to convert their RNA genome into a DNA intermediate which is incorporated into the host genome. Around 8% of the human genome is made up of sequences from viruses of the Retroviridae family of enveloped viruses [303]. The genome integrating capability of retroviruses allows the inserted genomic material to be passed on to daughter cells, making their use as vectors for gene delivery very attractive [304]. Retroviruses have two main assembly mechanisms: Type B and Type C [302]. Type B retroviruses assemble their capsids in the cytoplasm and obtain their envelope via budding at the plasma membrane. Type C differs in that the capsid is formed at the plasma membrane. One of the most prominent examples of Type C retroviruses is human immunodeficiency virus.  1.4.3.1 Human immunodeficiency virus Human immunodeficiency virus (HIV) is a bloodborne virus that continues to plague the global population. Transmission occurs human-to-human through sexual intercourse, by contaminated blood and blood products, by needle-sharing, or by vertical transmission (mother-to-child). HIV primarily binds to cluster of differentiation 4 (CD4) antigen via the envelope protein subunit gp120 [305,306]. This initial binding step causes a structural change in gp120, allowing for binding to a secondary receptor (C-C motif chemokine receptor, CCR5, and/or C-X-C chemokine receptor type 4, CXCR4) to enable membrane fusion. The genomic material is inserted into the cell and reverse transcribed prior to nuclear import of the genome. The complimentary DNA viral genome is then integrated into the host genome. HIV infection eventually manifests as acquired immunodeficiency syndrome (AIDS) [307]. This is caused by the infection and 40  destruction of CD4+ T helper cells, a key player in the adaptive immune system. The weakened immune system is unable to fight off other infections, known as opportunistic infections. Anti-retroviral treatment (ART) goals include minimizing viral load in the plasma, increasing patient CD4+ T cell count, and preventing transmission.   1.4.3.1.1 Implications in cardiovascular/haemostatic abnormalities Although ART has successful prolonged survival times, HIV-infected individuals are suffering from non-AIDS complications. Many of which are ageing-related diseases such as cardiovascular disease, hypertension, diabetes mellitus, bone fracture and renal failure [308]. The risk of venous thromboembolism in HIV-infected individuals is up to ten-fold higher than the general population [309]. HIV-infected individuals have a heightened expression of TF in their monocyte populations [310,311]. ART treatment decreased circulating TF-containing microparticles and TF activity in the blood [312]. Many studies have explored the coagulant phenotype of HIV-infected individuals, many of which have demonstrated significant prolongation of PT and aPTT [313–315]. aPTT showed an inverse relationship with patient CD4+ cell count, suggesting the use of aPTT as a clinical marker for HIV severity/status in areas where CD4 tests are not readily available [315].   1.5 Hypotheses Considering that FX activation is affected by the presence of gC and that TF is functionally present on HSV1, my hypothesis is that gC aids FX activation through enhancing TF function (Figure 12). I hypothesize that Herpesviridae is not the only family of enveloped virus that 41  incorporates TF into its viral envelope and that other blood-relevant enveloped viruses can incorporate TF so long as it infects and propagates in a TF-expressing cell.    42       Figure 12. Hypothesized model of gC-TF interaction. The known gC/FX interaction may aid in TF/FVIIa-mediated activation of FX on the virus surface. These clotting proteases are involved in cell signaling and inflammation. HSV1 has been demonstrated to make use of cell signaling to increase infection. As a side-effect of clotting protease activation, clots are formed which may lead to pathologies such as thrombosis. 43  Chapter 2: Material and Methods 2.1 Proteins and reagents.  Dry dimethylformamide (DMF), 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), acetone, dry dimethyl sulfoxide (DMSO), dithiothreitol (DTT), and cyanogen bromide (CNBr) were purchased from Sigma-Aldrich Inc. (St. Louis, MO, USA). Calcium chloride dihydrate (EMD Chemicals, Inc. Greenwich Township, NJ, USA). Tetrasodium ethylenediaminetetraacetic acid (EDTA) and 3,3'-dithiobis(sulfosuccinimidyl propionate) (DTSSP) were purchased from Thermo Fisher Scientific (Waltham, MA, USA). SeaPlaque™ agarose was purchased from Lonza (Basel, Switzerland). Paraformaldehyde (32% in methanol free solution) was purchased from Electron Microscopy Sciences (Hatfield, PA, USA). Purified human plasma-derived factor VIIa (FVIIa), factor X (FX), factor Xa (FXa) and corn trypsin inhibitor (CTI) were purchased from Haematologic Technologies (Essex Junction, VT, USA). Recombinant human FVIIa was gratefully obtained from Novo Nordisk (NovoSeven®) through a research grant. Innovin (Siemens Healthcare Diagnostic Inc., Delaware, USA) was used as a source of purified recombinant human TF inserted into liposomes. In-house purified FX from normal human pooled plasma (NP) was also used [316,317]. Plasmids for the expression of soluble TF (sTF-His), where the transmembrane and cytoplasmic domains were substituted for His6 and a 5-amino acid spacer restoring full cofactor activity [318], were obtained through my collaboration with Dr. James Morrissey (University of Michigan Med. School, Ann Arbor, MI, USA). Baculovirus engineered to express a soluble form of gC where the membrane-spanning domain was substituted with His5 (gCΔ457t, referred to as sgC-His) [319,320] was a kind gift from Dr. Gary Cohen and Dr. Roselyn Eisenberg (University of Pennsylvania, Philadelphia, PA, USA). Frozen NP and congenital factor VIII (FVIII)-deficient plasma 44  (FVIII/DP) were obtained from George King Bio-Medical (Overland Park, KS, USA). NP was also purchased from Affinity Biologicals Inc. (Hamilton, ON, Canada). L-α-phosphatidylserine (Brain, Porcine, PS), L-α-phosphatidylcholine (Egg, Chicken, PC) and the nickel salt of 1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid) succinyl] (DGS-NTA-Ni) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL, USA). FXa-specific chromogenic substrate Z-D-Arg-Gly-Arg-pNA (S-2765) and broad-spectrum chromogenic substrate H-D-lle-Pro-Arg-pNA (S-2288) were purchased from Chromogenix (Milano, Italy). Fluorophores Alexa Fluor 647 succinimydl ester and Alexa Fluor 647 maleimide were purchased from Invitrogen (Carlesbad, CA, USA). His-Tag Labeling Kit RED-tris-NTA, Protein Labeling Kit RED-NHS, and NT.115 Standard and Premium Capillaries for microscale thermophoresis (MST) were from NanoTemper Technologies GmbH (Munich, Germany).   2.1.1 Antibodies and probes Mouse anti-TF (TF9-9B4) monoclonal antibody was obtained from Dr. James Morrissey [321]. Mouse anti-TF (TF8-5G9) monoclonal antibody was obtained from Dr. Wolfram Ruf (Scripps Research Institute, La Jolla, CA, USA) [322]. Rabbit anti-sgC-His polyclonal antibody (R118) was a kind gift from Dr. Gary Cohen and Dr. Roselyn Eisenberg. Mouse anti-HSV1 major capsid protein monoclonal (MCP, Biodesign International, Saco, ME, USA), mouse anti-TF monoclonal (4503, American Diagnostica, Stamford, CT, USA), rabbit anti-TF polyclonal (ab48647, Abcam, Cambridge, UK), Mouse anti-flavivirus E-glycoprotein monoclonal antibody (ab155882, Abcam), and Anti-6X His tag® antibody (horseradish peroxidase, HRP) (ab1187, Abcam), mouse anti-gC monoclonal (P1104, EastCoast Bio), HRP-conjugated goat anti-mouse IgG (115-035-146, Jackson Immunological, West Grove, PA, USA) and HRP-conjugated goat 45  anti-rabbit IgG antibodies (115-035-003, Jackson Immunological) were purchased. Biotin annexin V was purchased from BioLegend (San Diego, CA, USA). Goat anti-mouse IgG Fc pre-adsorbed with 6 nm gold particles (ab105285) and goat anti-rabbit IgG H&L pre-adsorbed with 15 nm gold particles (ab27236) were purchased from Abcam and goat anti-biotin pre-adsorbed with 10 nm gold particles were purchased from BBI Solutions (Crumlin, UK). Acetylated BSA and Aurion gold sol (6 nm and 15 nm) were commercially obtained (Aurion; Wageningen, The Netherlands). Goat anti-HCV E2 polyclonal antibody (PA1-73105) was purchased from Invitrogen, Thermo Fisher Scientific. Mouse anti-HIV gp120 monoclonal antibody (clone 55-36) was purchased from Bio X Cell (West Lebanon, NH, USA).   2.1.2 Preparation of gold-conjugated primary antibodies Gold conjugation was performed as per supplier instructions (Aurion) [323]. Briefly, antibodies were buffer exchanged with 5 mM NaHCO3, pH 8.5 or 9.0 using Amicon® 10 kDa centrifugal filter or Slide-A-Lyzer™ MINI Dialysis Device, 10K MWCO (Thermo Scientific). Mouse monoclonal antibodies were conjugated using 5 mM NaHCO3 at pH 8.5 whereas rabbit monoclonal antibodies used pH 9.0. Prior to conjugation, the antibodies were tested for the minimal protecting amount of protein for gold conjugation. In the case of low quantities of antibody, only the appropriate isotype was tested for the minimal protecting amount of protein.  2.1.3 Viruses HSV1 NS strain, a low passage clinical isolate that expresses gC (gC+), and ns-1 strain, a mutant lacking expression of gC on its envelope (gC-), were kind gifts from Dr. Harvey Friedman (University of Pennsylvania, Pittsburgh, PA, USA). Initial virus stocks were passaged in African 46  green monkey kidney cells (Vero CCL-81; ATCC). To obtain viruses expressing surface gC and/or TF, gC+ and gC- viruses were propagated through a TF-deficient human melanoma cell line, A7, and an engineered TF+ derivative cell line, A7/TF, as previously described [12]. Virus purity was assessed by negative-staining transmission electron microscopy (EM) and was confirmed to contain only enveloped particles. Quantification of virus particles was performed using known concentrations of latex spheres. Western blot analysis was performed to validate the presence of desired antigen(s) and their molecular weight in each purified virus sample (Figure 13).  2.1.3.1 Characterizing antigens in HSV1 panel The presence or absence of gC and TF was confirmed by Western Blot analysis following sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) of purified virus (1 x 1011 virus particles, vp) using a 5-15% gradient acrylamide gel. Samples were reduced with 80 mM DTT. The proteins were transferred to a polyvinylidene difluoride (PVDF) membrane and blocked with 2% skim milk in TBST for 1 h at room temperature. The membrane was then probed for 1 h at room temperature with the primary antibodies anti-gC (P1104, 2 µg/mL), anti-TF (4503, 1.5 µg/mL) and anti-HSV1 MCP (1 µg/mL) diluted in TBST/2% skim milk. MCP was used as a loading control as there is a specific amount of MCP per virus. After three quick washes with TBST, the membranes were washed thrice with TBST for 5 min each. Secondary antibody treatment was performed using horseradish peroxidase-conjugated goat anti-mouse IgG (1:20,000) in TBST for 1 h at room temperature. Following the previous washing step, reactive bands were visualized using enhanced chemiluminescent substrate (Pierce, Ontario, Canada)  as per supplier instructions. The concentration of virus TF was determined by comparison to a purified sTF standard curve. 47           Figure 13. Characterization of antigens on HSV1. Western blot analysis of panel of HSV1 used in this study confirming the presence of gC and/or TF in the virus samples. Major capsid protein (MCP) was used as a loading control due to its defined stoichiometry per virus. 1010 HSV1 particles were run on a reducing 5-15% polyacrylamide gel. MCP, gC and TF migrated at 150 kDa, 90 kDa and 47 kDa, respectively.   48  2.1.4 Small unilamellar vesicles  Compositions of small unilamellar vesicles (SUVs) were as follows (mole percent): PCPS (20% PS and 80% PC); NiPC (15% DGS-NTA-Ni and 85% PC); and NiPCPS (15% DGS-NTA-Ni, 65% PC and 20% PS) [318]. SUVs were prepared by sonication and differential centrifugation with modifications from previously established protocols [324]. Briefly, lipids dissolved in chloroform were mixed and dried under nitrogen gas. Following resuspension in 0.22 µM filtered HBS (20 mM HEPES, 150 mM NaCl, pH 7.4) and incubation for 10 min on ice, the lipids were treated with a direct sonication probe (Misonix microtip #419, 1/8”) at an amplitude of 70 for 5 min using 10 s on/off cycles to prevent overheating the suspension (Misonix sonicator S-4000). The vesicles were transferred to 1.5 mL ultracentrifuge tubes and centrifuged at 114,600 x g for 3.5 h at 20 °C in a TLA-55 rotor using a Beckman Optima Refrigerated Benchtop Ultracentrifuge. The upper 65% was collected and stored under argon gas at 4 °C. The concentration of PC was determined using the mole percent of PC to derive the total concentration of lipid after enzymatic hydrolysis with a Phospholipids C kit (Wako, Mountain View, CA, US). Quantified SUVs were sized by dynamic light scattering with a Beckman Coulter N4 PLUS Particle Size Analyzer or the Malvern Panalytical ZetaSizer Nano ZS and fell in the 41 ± 15 nm average diameter range.  2.1.5 Protein expression and generation Two proteins were recombinantly expressed for the study of gC and TF interactions in Chapter 3, sgC-His and sTF-His. Both proteins possess His-tags that facilitate purification as well as docking onto a nickel-chelating surface, e.g. small unilamellar vesicles synthetically produced in Chapter 2.1.4. A graphical representation of this is shown in Figure 14.    49   Figure 14. Recombinantly expressed sgC-His and sTF-His interaction with a Ni-chelating membrane.  (A) A NiPCPS membrane is depicted with sgC-His and sTF-His bound to the membrane via their His-tags (red glowing lines). (B) Phosphatidylserine (PS, magenta), phosphatidylcholine (PC, green) and DGS-NTA-Ni (gray) are depicted. Lipid images originated from the product website (https://avantilipids.com/). (C) The NTA-Ni headgroup coordinates with two histidine residues from a His-tagged protein. 50  2.1.5.1 gCΔ457t/sgC-His A baculovirus expression system was used to enable production of large amounts of glycosylated secreted sgC-His. Spodoptera frugiperda cells (Sf9) were chosen based on a previously described protocol [319]. sgC-His has a portion of the extracellular domain removed along with the transmembrane domain and cytoplasmic tail (Figure 10) and three additional histidine tags were engineered into the C-terminal at residue 457. Due to the heterogeneity of the recombinant protein generated based on previous protocols (Figure 15A), several modifications were made to improve homogeneity of the sgC-His (Figure 15B). Sf9 cells were grown to confluence in T-175 cm2 tissue culture flasks and were then seeded into 500 mL Celstir flasks containing no more than 400 mL of serum-free Sf900IIM. Cells were maintained at no more than 4 x 106 cells/mL and >95% viability as determined by trypan blue exclusion. For expansion of P1 and P2 stocks of baculovirus, 400 mL of 2 x 106 cells/mL were infected with a multiplicity of infection (MOI) of 0.5. Fetal bovine serum (FBS) was added to the media at 10% as a stabilizer if the stocks were to be kept for longer than 1 year at -80 °C (typically for P1 and P2 stocks). The inclusion of FBS lowered the efficiency of downstream purification of proteins and thus, P3 stocks were made serum-free. For protein expression, a MOI of 4 was used when the cells reached 4 x 106 cells/mL in 4-5 separate flasks of 400 mL. After 4 days, the media was collected and centrifuged at 2,000 x g for 20 min at 4 °C to remove cells and cell debris. Viral stocks were frozen at -80 °C and the viral titer was determined as detailed in 2.1.5.1.2.   2.1.5.1.1 Protein purification The supernatant was filtered through a 0.45 µM filter and precipitated with ammonium sulfate in two steps. The first precipitation step was 20% ammonium sulfate (fine powder) slowly 51  added with stirring on ice over 30 min. After centrifugation at 12,900 x g for 30 min, the supernatant was transferred to a new beaker. The second precipitation step required 70% ammonium sulfate, added slowly over 1.5 h then left undisturbed at 4 °C for 1 h. The media was centrifuged at 12,900 x g for 35 min and the supernatant removed. The floating lipid material in the supernatant was discarded. The pellet was drained before resuspending in 30-40 mL of Cohen’s PBS (20 mM sodium phosphate, 150 mM NaCl, pH 7.3). The resuspended pellet was then dialyzed at 4 °C against Cohen’s PBS thrice (3 L) using a 3.5 kDa membrane (SpectraPor). The dialysate was centrifuged at 100,000 x g for 1 hr at 4 °C prior to loading the supernatant onto a heparin Sepharose column. The heparin Sepharose CL-6B (GE Healthcare) was packed and equilibrated with Cohen’s PBS (Buffer A) in a XK16 column as per supplier’s instruction, with a column volume of approximately 24 mL. The filtered dialysate was added at a flow rate of 1 mL/min. At this point, the resin was tinted brown due to the color of the dialysate. After washing with 5 column volumes of Buffer A, the proteins were eluted and collected in 1 mL fractions with 0-60% Buffer B (20 mM sodium phosphate, 2 M NaCl, pH 7.3) over 75 min, 60-100% Buffer B over 10 min and 100% Buffer B for 5 min. Fractions were analyzed by Western blot to determine sgC-His-containing fractions. The majority of sgC-His was found in fractions 53 to 75 and was pooled together before dialyzing at 4 °C against Cohen’s PBS twice using a 3.5 kDa membrane (SpectraPor) to a final dilution of 6,000.  The dialyzed pooled fractions were centrifuged at 25,000 x g for 15 min at 4 °C. The supernatant was added to 3 mL of 50% Ni-NTA resin in Buffer A (20 mM sodium phosphate, 500 mM NaCl, pH 7.5) and incubated for 3 h with shaking at 4 °C. The resin and media were transferred to a 5 mL column and the resin was washed with 100 mL of Buffer A followed by 100 mL of 10 mM imidazole in Buffer A. Elution was performed using a step gradient as previously described 52  [319]. sgC-His containing fractions were pooled together, concentrated and buffer-exchanged in an Amicon® Ultra 15mL 10K molecular weight cut-off centrifugal filter (Millipore) at 4 °C.  The final step of the purification involves size exclusion chromatography using a Sephacryl S-200 column. The column (Pharmacia 10/200) was packed as per supplier’s instruction manual. The sgC-His-containing sample was run through the Sephacryl S-200 column with a flow rate of 1 mL/min in HBS at 4 °C, whilst collecting 1 mL fractions. Fractions with the least amount of large molecular weight species (>150 kDa, as compared to 75-90 kDa sgC-His, as determined by Western blot) were pooled together and concentrated using an Amicon® Ultra 15 mL 10K molecular weight cut-off centrifugal filter. The sample was aliquoted, flash frozen in LiN2 and stored at -80 °C. Protein concentrations were measured using the bicinchoninic acid assay and UV-Vis method (A280 with A320 correction).  2.1.5.1.2 Determination of baculovirus titer Sf9 cells (1 x 106) were seeded into 6-well Corning® Costar® tissue culture-treated plates (Sigma-Aldrich) and grown at 28 °C, 0% CO2. At ~85% confluency, the cells were infected with 1 mL of titrated baculovirus-infected supernatant for 1 h at 28 °C, 0% CO2. The baculovirus-infected supernatant was aspirated and 3 mL of 1X Grace’s Insect Medium with 1.5% SeaPlaque™ agarose were overlaid onto the cell monolayer. After 5 days of incubation at 28 °C, 0% CO2, plates were stained with 1 mL of 0.22 µm-filtered 0.02% neutral red in PBS for 2 h at 28 °C. The remaining dye was removed, and the plates are returned to the incubator at 28 °C until plaques are visible. Neutral red is a vital dye that is taken up by live cells, the absence of red denotes a plaque. Plaques are counted and multiplied by the dilution factor to obtain the plaque forming units (pfu), a unit of virus infectivity. 53         Figure 15. Recombinant sgC-His antigen confirmation by protein stain and Western blot. (A) sgC-His was purified with a nickel metal-ion affinity chromatography and samples were reduced (R, 12 mM DTT) or non-reduced (NR) when loaded into a 5-20% gradient polyacrylamide gel. Anti-gC (anti-gC (2 µg/mL, P1104) was used as the primary antibody with goat anti-mouse HRP (1:10,000) as the secondary antibody. Protein staining was achieved by using Coomassie R-250 protein stain. (B) Improvements to previous protocols included heparin column prior to metal-ion affinity and size exclusion post-metal-ion affinity chromatography and resulted in a cleaner preparation. Molecular weight (M.W.) determined using BLUeye Prestained Protein ladder.     54  2.1.5.2 sTF-His Expression and purification of sTF-His was performed using an Escherichia coli expression system that was previously established by my collaborator (Morrissey) [318]. sTF-His is a soluble form of TF (sTF) that lacks the transmembrane and cytosolic domains, and contains a sequence encoding a dodecapeptide epitope for the calcium-dependent monoclonal antibody HPC4, as well as a hexa-histidine tag (His-tag), and a spacer sequence at amino acid 219 before the His-tag (depicted in Figure 2). The spacer sequence was included to help orient the protein when bound to an appropriate membrane surface. Briefly, E. coli strain BL21 (DE3) was transfected with pJH776 (sTF with a spacer before the N-terminal His-tag). Transformants were selected with kanamycin and were grown in a 16x150 mm test tube containing 1.5 mL MDG (non-inducing minimal medium) for 7 h at 37 °C (300 rpm shaking). This culture was expanded into 500 mL of MDG medium and grown overnight at 37 °C (300 rpm shaking). The bacteria were pelleted and the MDG medium was replaced with 1 L Terrific Broth medium with 0.1% glucose and 20 µM IPTG to a final A600 absorbance unit reading of 3.0.  Following IPTG induction and overnight incubation at room temperature (300 rpm shaking), the media was collected and centrifuged to remove cells and cell debris. Osmotic shock was performed on the pelleted cells to release sTF stuck in the periplasmic space of the bacteria [325]. The supernatant and the osmotic shock sample were precipitated using 70% ammonium sulfate. Following dialysis of the precipitate, the protein solution was batch-treated with Q Sepharose. The unbound solution contained sTF-His and was applied to an immobilized nickel ion affinity resin. sTF-His bound to the resin was eluted, fractions were pooled and buffer-exchanged to HBS. The purified protein was aliquoted, flash frozen and stored at -80 °C until use.  55         Figure 16. Recombinant sTF-His antigen confirmation by protein stain and Western blot. sTF-His under reducing (R, 12 mM DTT) or non-reducing (NR) conditions was loaded into a 5-15% gradient polyacrylamide gel. Anti-TF (1:750, ab48647) was used as the primary antibody with goat anti-rabbit HRP (1:10,000) as the secondary antibody. Protein staining was achieved by using Coomassie R-250 protein stain. Molecular weight (M.W.) determined using BLUeye Prestained Protein ladder.    56  2.2 Detection of specific antigens on the virus envelope 2.2.1 Immunogold electron microscopy  The standard for characterizing virus surface antigens on membranes is immunogold EM. This method relies on antibody recognition of an antigen with visualization of these antigens via dense colloidal gold. Colloidal gold appears as dark dots by EM and can be generated in different sizes. Conjugation of gold colloidal to antibodies (either primary or secondary) can be easily achieved through passive adsorption or covalent chemistry. For my cell-cultured virus studies, gold visualization was performed using commercial secondary antibodies conjugated to gold particles of varying diameters (6 nm, 10 nm and 15 nm). For patient-derived virus studies, in-house conjugation of primary antibodies was performed. This was important due to the lack of host selection of antibodies against virus proteins which would complicate simultaneous labeling of antigens on the virus surface. There are two main methods of loading samples onto an EM grid: drop-on-grid and grid-on-drop (Figure 17A and Figure 17B, respectively). As the names imply, drop-on-grid involves securing a grid with the shiny film side up, typically with forceps, and applying a drop of sample on top of the grid. Grid-on-drop method places the grid on top of a droplet of sample. The former method allows for larger particles to sediment onto the grid and is superior for large viruses such as poxviruses [326]. The benefit of using the latter method is the ability to handle multiple samples easily. Although both methods were tested for HSV1 samples, the grid-on-drop was predominantly used and then further adapted to a pseudo-drop-on-grid method (Figure 17C). This method entailed applying grids with the shiny side onto 20-30 µL of sample deposited in the wells of a Terasaki 60-well microtest plate (Sarstedt). The entire plate was then inverted such that the shiny sides of the grids were facing upward. This technique allowed for gravity to pull down virus particles onto  57   Figure 17. Sample adsorption to grids for EM. Three methods for sample adsorption: (A) Drop-on-grid, an EM grid is secured with the shiny side upward using forceps and the sample is placed on top of the grid; (B) Grid-on-drop, the sample is placed on a parafilm/support and the shiny side of the grid is placed on top of the sample; and (C) Pseudo drop-on-grid method, the sample is placed within a well (e.g. Terasaki microtest plate) with the EM grid sitting on top of the sample and the entire plate is inverted to mimic a drop-on-grid.   58  the grid whilst retaining the easy manipulation advantage of the grid-on-drop method. For samples less than 20 µL, parafilm was first fitted onto the wells of the microtest plate, thereby creating a shallower well to hold smaller volumes. All virus EM work were performed in a BSL2+ facility using sterile equipment and solutions. Formvar® carbon-coated nickel 400-mesh grids (EMS, Hatfield, PA, USA) were used in lieu of the more common copper grids as copper can become oxidized by gold nanoparticle antibody solutions. As nickel grids are magnetic, non-magnetic forceps were used when handling the grids for staining.   2.2.2 Purified cell culture-derived HSV1 particles Using the grid-on-drop method, nickel grids were floated on a 10 µL drop of HSV1 (1 x 10 10 vp) diluted in low salt HBS (20 mM HEPES, 75 mM NaCl, pH 7.4). After 15 min of adsorption at room temperature, the grids were transferred to a 30 µL drop of low salt HBS. The grids were then placed on a drop of filter-sterilized blocking buffer consisting of 5% BSA, 5% goat serum (or 100 µg/mL goat IgG) and 0.1% cold water fish scale gelatin in low salt HBS for 1 h at room temperature. The grids were washed by transferring the grids to 30-40 µL drops of low salt HBS for 4 min and repeated four times. The grids were then incubated with primary antibodies in low salt HBS and 0.1% acetylated BSA (acBSA). Antibody concentrations were as follows: 5 or 10 µg/mL tissue factor antibody (rabbit or mouse, respectively); 10 µg/mL (or 1:100 for rabbit serum) anti-gC antibody; and/or 10 µg/mL biotinylated-Annexin V. Matching isotype/host controls for each of the antibodies were used to test for specificity. In the case of Annexin V, the negative controls included no calcium added or EDTA treatment. In experiments probing for aPL, 5 mM CaCl2 was added to all buffers to allow for calcium-dependent binding of Annexin V to 59  aPL. After 1 h of incubation, the grids were washed five times. Secondary antibodies at 1:40 dilutions of goat anti-mouse IgG conjugated to 6 nm beads, goat anti-biotin conjugated to 10 nm and goat anti-rabbit IgG conjugated to 15 nm beads was incubated for 1 h. Following five washes, the grids were negatively stained on 10 µL drops of 2% phosphotungstic acid (PTA), pH 6.5 for 1 min. PTA is electron dense and is a commonly used negative stain for visualization of viruses. The negative stain was removed by wicking with filter paper and the grids were air-dried in a container for EM grids. PTA can potentially degrade viral membranes; therefore, the grids were viewed within a day of preparing them. Viruses were inactivated by UV light after negative staining or by fixation with 2% paraformaldehyde before negative staining.   2.2.3 Purified cell culture-derived DENV particles Due to the relatively small size of DENV (>3 fold smaller than HSV1 in diameter), an extra measure was taken to improve detection of the virus. Glow discharge can be used to deposit positive ions onto the EM grid, which in turn increases negatively charged virus particle adsorption[318]. After glow discharge (PELCO easiGlow™ 91000, 120 s at 0.2 mbar and 15 mA), 5 µL of DENV sample was adsorbed onto the grid. Then the same protocol as in HSV1 (Section 2.2.2) was followed but using 10 µg/mL mouse anti-DENV and 10 µg/mL rabbit anti-TF antibodies for the primary antibody incubation.  2.2.4 Patient plasma-derived virus particles Virus concentrations in patient plasma can vary depending on the type of virus and the severity of the disease. From purified virus titrations, it was determined that at least 1 x 106 vp/mL is needed to visualize ~10 vp/square in a 400-mesh EM grid. High titer samples that meet this 60  minimum requirement are easier to come by in HCV and HIV. DENV viremia results in at least 10 to 100-fold lower titres compared to HCV or HIV and is difficult to obtain peak titers. Furthermore, there is only one example in the literature successfully extracting and visualizing DENV from patient plasma [327]. In anticipation of the difficulties in detecting virus particles, two methods were developed to concentrate viruses from patient plasma: centrifugation and immunocapture.  2.2.4.1 Centrifugation method The following protocol was based on the ability of gold-conjugated antibodies to pellet using speeds much lower than those required for smaller viruses such as dengue virus. For this purpose, 15 nm gold beads were chosen as they pellet at 12,000 x gav. Viremic patient plasma was pre-spun at 15,000 x g for 5 min to remove large aggregates or debris. Gold-conjugated primary antibodies were incubated with the plasma in solution for 2 h at 37 °C with 0.1% acBSA. Antibody dilutions were dependent on the OD520: mouse anti-HIV gp120 (1:80), mouse anti-flavivirus E protein (1:160), goat anti-HCV E2 (1:80), and mouse anti-human TF (1:100). The plasma was then centrifuged at 8,000 x g for 20-30 min at 4 °C. The pellet was washed with HBS/0.1% acBSA (HBS/acBSA) and pelleted as above. The virus pellet was resuspended in 5-20 µL low salt HBS, loaded directly onto a nickel grid and allowed to adsorb for 5 min at room temperature. The virus was then fixed with 2% paraformaldehyde for 30 min prior to negative staining with 2% PTA, pH 6.5 for 1 min. The grids were then dried and visualized.   61     Figure 18. Centrifugation method for capturing and concentrating viruses in plasma. (A) Viremic plasma was clarified by centrifugation at 15,000 x g. (B) Viruses were incubated with 15 nm gold-conjugated virus-specific antibodies and 6 nm gold-conjugated anti-TF antibodies. (C) The 15 nm gold beads allowed for centrifugation at 8,000 x g to pellet the gold/antibody/virus complexes. (D) The pellet was washed before resuspending the pellet in a smaller volume. (E) The virus pellet was transferred to an EM grid for adsorption. (F) The virus was fixed with 2% paraformaldehyde. (G) After fixation, the grids were negatively stained with 2% phosphotungstic acid.   62  2.2.4.2 Immunocapture method This method involves capturing viruses onto an EM grid using virus-specific antibodies. Nickel grids are incubated with 10-100 µg/mL virus-specific antibodies for 20 min at room temperature. After two 4-min washes in low salt HBS, the grids are blocked with Block buffer for 20 min at room temperature. The grids are washed twice with low salt HBS. 20 µL of patient plasma are added to wells of a Terasaki 60-well Microtest plate. The pseudo-drop-on-grid method was employed. After incubating the grid with patient plasma for 15 min at room temperature, another 20 µL of sample are added to new wells and grid incubation was repeated. Following the incubation, the grids were washed five times with low salt HBS/acBSA for 4 min each. Grids were transferred to gold-conjugated antibody solutions in low salt HBS/acBSA for 1 h at room temperature. Antibody dilutions were dependent on the optical density at 520 nm: mouse anti-HIV gp120 (1:80), mouse anti-flavivirus E protein (1:100), goat anti-HCV E2 (1:80), and mouse anti-human TF (1:40). Grids were washed five times with low salt HBS for 4 min each before fixing with 30 µL 2% paraformaldehyde in low salt HBS for 30 min at room temperature. The grids were washed twice briefly with low salt HBS before negative staining with 2% PTA. Samples were visualized shortly after drying. Alternatively, a primary and secondary antibody labeling technique as performed for HSV1 was used for detecting DENV due to the low virus titers and overall difficulty in detecting DENV particles. The primary antibodies were used at 10 µg/mL and were a mixture of anti-flavivirus E protein and anti-DENV E protein (Abcam) to increase sensitivity.   63     Figure 19. Immunogold capture of viruses from plasma for EM. (A) Virus-specific antibodies are adsorbed to an EM grid. (B) Viremic plasma was incubated on immune-EM grids. (C) Grids are incubated with 15 nm gold-conjugated virus-specific antibodies and 6 nm gold-conjugated anti-TF antibodies. (D) The virus was fixed with 2% paraformaldehyde. (E) After fixation, the grids were negatively stained with 2% phosphotungstic acid.   64  2.3 Assessing the function of procoagulant factors. 2.3.1 Virus-mediated FX activation All experiments used HBS/BSA as a stabilizer unless otherwise stated. In a final reaction volume of 20 µL, virus was first mixed with 100 nM FX and 1 nM FVIIa in 15 µL. After 5 min at room temperature, 5 µL of 20 mM CaCl2 was added to start the reaction. After 20 min at 37 °C with shaking every 5 min, 10 µL was transferred to a 96-well plate and 90 µL of 200 µM S-2765 and 12 mM EDTA. The amount of FXa generated was then monitored at 405 nm at room temperature using a Vmax or Spectramax multiwell plate spectrophotometer (Molecular Devices, Sunnyvale, CA, USA). The amount of FX activation was derived from a standard curve using purified FXa. For dengue virus experiments, inhibitors of the TF-FVIIa complex were used to determine if the sole contributor to enhanced FVIIa activity was TF. The nematode anticoagulant protein c2 (NAPc2) and human TF-specific monoclonal mouse antibody TF8-5G9 were pre-incubated with the virus at 50 nM and 100 µg/mL, respectively, for 5 min at 37 °C before addition of other coagulation factors.  2.3.2 Analysis of FX activation in purified systems Initial rates of FX activation were measured with a two-step discontinuous chromogenic assay as previously described [328] with modifications. This assay involves activating FX, stopping the reaction at various times and measuring the FXa generated at each time point. Reaction mixtures in HBS contained 5 mM CaCl2, 500 pM FVIIa, and either 10 pM sTF-His or 10 nM sgC-His with 100 µM NiPCPS. Varying concentrations of FX were added to initiate the reaction at 37 °C. Over the course of 10 min, 20 µL aliquots of the reaction mixture were added to 65  50 µL stop buffer (40 mM Mes-NaOH (pH 5.8), 12 mM EDTA, 50 mM NaCl and 0.25% Triton X-100) at 4 °C. After 10 min, the stopped reactions were then warmed to room temperature and neutralized with 0.6 M Tricine-NaOH (pH 8.4). Immediately following, the amount of FXa generation was detected by the addition of 0.6 mM S-2765 and 5 mM CaCl2. A SpectraMax was used to follow FX activation at OD405 over 10 min, 15 s intervals with 3 s shaking between reads at 25 °C.  2.3.2.1 Deriving FX activation kinetics parameters The 2-step chromogenic assay raw data values are converted to nM FXa with a standard curve using purified FXa. The data are fitted to a 2nd order polynomial curve (Y = A + BX + CX2). As this equation described the amount of FX activation (Y) as a function of time (T), the derivative of this equation was the rate of FX activation. Thus, the rate of FX activation was equal to B + CX. The initial rate of reaction, V0, was the B value, when T = 0. The V0 values are plotted as a function of [FX]. Using the Michaelis-Menten equation, 𝑌 =𝑉𝑚𝑎𝑥[𝐹𝑋]𝐾𝑚+[𝐹𝑋], the Km value can be derived. Subsequently, kcat was determined from Vmax (Bmax on GraphPad) and known [FVIIa/cofactor] using 𝑘𝑐𝑎𝑡 =𝑉𝑚𝑎𝑥[𝐹𝑉𝐼𝐼𝑎/𝑐𝑜𝑓𝑎𝑐𝑡𝑜𝑟] . The presence of nickel-chelating lipid influenced the His-tagged protein cofactor tethering and orientation. This subsequently affected the availability of the protein cofactor for FVIIa binding. Thus, the binding affinity of protein cofactor to lipid cofactor (Kd,C), as described in Figure 20, was determined to assess the amount of available protein cofactor that complexes with FVIIa. This was simplified with the assumption that the dissociation value of cofactor to FVIIa is much larger than that of cofactor/lipid to FVIIa.   66       Figure 20. Schematic of non-enzymatic and enzymatic reactions involved in FVIIa-mediated FX activation.  Enzyme (E, FVIIa) activation of the product (P, FX) is influenced by several factors. As the protein cofactor (CP), sTF-His or sgC-His, can bind to Ni-chelating lipids (CL), this equilibrium is described by Kd,C. This protein/lipid cofactor (C) can bind to E, with the equilibrium denoted by Kd. E can bind to S in the absence or presence of cofactors ((binding affinity Ks′ and Ks, respectively). Kd′ describes the equilibrium of ECS to ES and C. The conversion of S to P by either ES or ECS is represented by kcat′ and kcat, respectively. This schematic was modified from Neuenschwander et al. (1994) [94].   67  2.3.2.2 Nickel binding of purified proteins sTF-His and sgC-His binding to NiPC and NiPCPS was determined by a continuous chromogenic assay linked to FVIIa-mediated FX activation. Varying concentration of NiPCPS were mixed with 5 mM CaCl2, 1 nM FVIIa, 0.2 mM S-2765 and either 20 pM sTF-His or 1 nM sgC-His. FX was added to initiate the reaction at a final concentration of 100 nM. FXa generation was immediately followed at room temperature over 20 min, with a data acquisition interval of 15 s preceded by a 3-second shake at each read. The initial rate of FX activation was determined by fitting the activity data to a second-order polynomial [94]. Typically, to determine the dissociation constant, the initial rate of FX activation is converted to [enzyme/cofactor] and plotted against the titrating [cofactor]. However, the effect of free FVIIa may need to be accounted for. This is assessed by the following summation: 𝑣𝑜𝑏𝑠 =  𝛼[𝐹𝑉𝐼𝐼𝑎]  +  𝛽[𝐹𝑉𝐼𝐼𝑎/𝑐𝑜𝑓𝑎𝑐𝑡𝑜𝑟]  [94], where 𝑣𝑜𝑏𝑠 is the measured activity of FXa (initial rate of FX activation in mol/L per unit time). Values of α and β were experimentally determined using no cofactor and saturating amounts of cofactor, respectively. No measurable activity was measurable with up to 10 nM FVIIa (100-1000 fold more than was used in binding experiments), revealing that α << β similar to other studies [329]. Thus, the equation was simplified to 𝑣𝑜𝑏𝑠 ≈  𝛽[𝐹𝑉𝐼𝐼𝑎/𝑐𝑜𝑓𝑎𝑐𝑡𝑜𝑟]. The typical values for β were 4.4 x 10-2 mol FXa/s/mol sgC-His/FVIIa/NiPCPS; 4.3 x 10-2 mol FXa/s/mol sTF-His/FVIIa/NiPCPS; 1.8 x 10-3 mol FXa/s/mol sgC-His/FVIIa/NiPC; and 2.7 x 10-3 mol FXa/s/mol sTF-His/FVIIa/NiPC. The dissociation constant was determined by plotting the converted [FVIIa/cofactor] versus [cofactor]. The data was fit to the following quadratic equation: 𝑣 =𝑉𝑚𝑎𝑥 ∙ 𝑆2 ∙ 𝐸0(𝐾𝑠 + 𝑆)(𝐸0 + 𝐶0 + 𝐾𝑑,𝑎𝑝𝑝 ± √(𝐸0 + 𝐶0 + 𝐾𝑑,𝑎𝑝𝑝)2− 4 ∙ 𝐸0 ∙ 𝐶0)  68  Where, S is [FX] (30 nM), E0 is the initial [FVIIa] (0.1 nM), C0 is the titrating [cofactor]. KS describes the dissociation value of the FVIIa/cofactor and FX (Figure 20). The assumption that 𝐾𝑚 ≈ 𝐾𝑠 (values from Table 3) was made in order to fit the data using the above quadratic binding isotherm. Kd,app is the apparent dissociation value of cofactor and FVIIa. It represents the true Kd only when the substrate concentration is limiting relative to the Ks for substrate binding and is described by 𝐾𝑑,𝑎𝑝𝑝 = 𝐾𝑑𝐾𝑠+𝐾𝑠∙𝑆/𝐾𝑠′𝐾𝑠+𝑆 . If 𝐾𝑠 = 𝐾𝑠′, then 𝐾𝑑,𝑎𝑝𝑝 = 𝐾𝑑. Derivation of all formulae is found in the appendix (A.1).   2.3.2.3 FVIIa binding to lipid/protein cofactors Binding of sTF-His or sgC-His to either FVIIa was assessed using continuous chromogenic assays monitoring FXa generation. sTF-His or sgC-His was titrated in HBS with 5 mM CaCl2, 10 pM FVIIa and 100 µM SUVs. After 10 min of pre-incubation at room temperature to allow equilibration, 30 nM FX and 0.2 mM S-2765 was added to start the reaction. As with Chapter 2.3.2.2, FXa generation was followed at room temperature over 20 min. Kd,app was determined by fitting the data using C0 as the titrating , E0 = 0.01 nM FVIIa, S = 30 nM FX, Ks = Km (values from Table 3) with the equation:  𝑣 =𝑉𝑚𝑎𝑥∙𝑆2∙𝐸0(𝐾𝑠+𝑆)(𝐸0 + 𝐶0 + 𝐾𝑑,𝑎𝑝𝑝 ± √(𝐸0 + 𝐶0 + 𝐾𝑑,𝑎𝑝𝑝)2− 4 ∙ 𝐸0 ∙ 𝐶0) .  2.3.3 Plasma clotting Virus-induced coagulation was monitored using an ST4 coagulation analyzer (Diagnostica Stago). Briefly, a magnetic bead was added to the bottom of a cuvette which oscillates freely in solution. Frozen plasma was defrosted at 37 °C for 5 min then set on ice until use (as per product 69  specifications). In a final reaction volume of 75 µL, 12.5 µL of virus and 12.5 µL of HBS was added to 37.5 µL of plasma. Pre-warmed magnetic beads were added to the cuvette and incubated for 1 min at 37 °C. To start the clotting and timing, 12.5 µL of 60 mM CaCl2 was injected into each cuvette. The machine then counted the time (s) it took for the magnetic bead to stop moving in the cuvette due to clot formation. For inhibition of the intrinsic pathway, CTI was pre-incubated with NPP for 5 min at 37 °C. CTI inhibits FXIIa and is used to assess TF-dependent coagulation [330,331]. Similarly, for TF inhibition, NAPc2 or TF8-5G9 was pre-incubated with virus at 50 nM and 100 µg/mL, respectively.  2.4 Determining binding partners To determine which factors in virus-mediated FX activation are interacting with one another, several techniques were employed: crosslinking, enzyme-linked assays and microscale thermophoresis.  2.4.1 Crosslinking To investigate protein interactions between gC and TF, crosslinking was performed. 3,3'-dithiobis(sulfosuccinimidyl propionate) (DTSSP) is a 12 Å water-soluble crosslinker containing an amine-reactive N-hydroxysulfosuccinimide (sulfo-NHS) ester at both ends of the spacer. Sulfo-NHS esters react with primary amines at pH 7-9 to form stable amide bonds. DTSSP also features a single disulfide bond in the middle of the spacer which can be reduced to release crosslinked adducts. In addition, DTSSP is membrane-impermeable, making it ideal for investigating only protein interactions on the virus surface [332]. 70  HSV1/TF+/gC+, HSV1/TF+/gC-, HSV1/TF-/gC+, and HSV1/TF-/gC- (2 x 1010 vp) was incubated with 2.5 mM DTSSP for 15 min on ice. The reaction was then quenched with 1 M Tris-glycine (pH 7.0) to a final concentration of 50 mM. A control reaction where Tris-glycine (pH 7.5) was added prior to the cross-linker was also performed. For purified proteins, 12 µM sTF-His and/or 2.5 µM sgC-His was mixed with 50 µM NiPCPS and 5 mM Ca2+. DTSSP (500 µM) was incubated with the protein mixture for 24 h at 4° C with agitation.  The samples were run on a 5-15% polyacrylamide gradient gel under non-reducing or reducing conditions with 80 mM DTT. Western blot analysis was performed using polyclonal antibodies against TF (Abcam, 1:1500) and gC (R118, 1:1000) as primary antibodies and goat anti-rabbit (HRP) (Jackson, 1:10000) as the secondary antibody.  2.4.2 ELISA-based chromogenic assay Relipidated TF (Innovin) at 1:100 dilution was coated onto a 96-well plate in 50 µL 100 mM bicarbonate/carbonate coating buffer pH 9.6 overnight at 4 °C. Wells were washed twice with 200 µL HBS. Wells were blocked with 200 µL 2% BSA in HBS for 2 h at room temperature. After two washes with HBS, 40 µL of sgC-His at varying titrations from 0 to 1 µM was incubated for 1 h at room temperature. Following several HBS washes, two different detection methods were used: enzyme-linked immunoassay and enzyme-linked functional assay.  The former method requires a primary antibody incubation step with 10 µg/mL mouse anti-gC monoclonal antibodies for 2 h at room temperature. After several washes, the secondary antibody step consists of 1:10000 HRP-conjugated goat anti-mouse antibodies incubation for 2 h at room temperature. Detection of bound sgC-His was determined through 3,3’,5,5’-tetramethylbenzidine (TMB) conversion to a diamine form, causing the solution to turn blue. A 71  sulfuric acid solution was added to stop the reaction, resulting in a yellow solution which can be spectrophotometrically read at 450 nm. Purified sgC-His was used as a loading control and for the creation of a standard curve. The latter method depends on FX activation by FVIIa. TF and sgC-His cofactor activity were expected to enhance FVIIa-mediated FX activation. Similar to the assays described in section 2.3.1, 20 µL final volume consisting of 100 nM FX, 5 mM CaCl2 and 1 nM FVIIa was added to each well and incubated for 20 min at room temperature. 80 µL of 200 µM S-2765 and 12 mM EDTA was added to each well. The plate was then immediately read at 405 nm, reading samples every 12 s for 5 min. Enhancement was determined against a no added sgC-His control.   2.4.3 Microscale thermophoresis MST is used for determining binding parameters of proteins or small molecules in solution based on thermodiffusion. All molecules move away from a temperature gradient and this effect is altered for individual molecules when they associate. Although this principle was described in 1856, the theory is incomplete. The speed at which a molecule diffuses is dependent on its size, charge and hydration shell. When molecules interact, at least one parameter changes, thereby altering the way the complex moves through the temperature gradient. MST devices use infrared lasers to induce a temperature gradient in a capillary tube containing the samples (Figure 21). The device uses an excitation laser to track the movement of fluorescent molecules. Therefore, a fluorescence label is required for one of the molecules. As with other protein-interaction investigation approaches that involve labeling, there is the inherent disadvantage of affecting the target molecule’s behavior(s). Thus, controls must be performed to ensure that target binding or function is not affected. Non-labeled proteins can be followed with the shift of intrinsic tryptophan  72    Figure 21. MST basic theory and set-up. (A) The Monolith NT.115 from NanoTemper Technologies GmbH. Up to 16 capillaries can be loaded onto a capillary tray. (B) Schematic representation of MST optics. The fluorescence within the capillary (~4 µL volume) is excited and detected through the same objective. A focused IR-Laser locally heats a defined sample volume. Thermophoresis of fluorescent molecules through the temperature gradient is detected (C) Example of MST experiment. Initially, the molecules are homogeneously distributed and a constant “initial fluorescence” is detected. Once the IR laser is activated, a temperature jump (T-Jump), corresponding to a rapid change in fluorophore properties due to the fast temperature change, is observed. Subsequently, thermophoretic movement of the fluorescently labeled molecules out of the heated sample volume can be detected and is measured over a defined length of time. Deactivation of the IR-Laser results in an inverse T-Jump occurs, followed by the “back diffusion” of molecules, which is solely driven by mass diffusion. (D) Typical binding experiment. The thermophoretic movement of a fluorescent molecule (black trace; “unbound”) changes upon binding to a non-fluorescent ligand (red trace; “bound”), resulting in different traces. For analysis, the change in thermophoresis is expressed as the change in the normalized fluorescence (ΔFnorm), which is defined as Fhot/Fcold (F-values correspond to average fluorescence values between defined areas marked by the red and blue cursors, respectively). Titration of the non-fluorescent ligand results in a gradual change in thermophoresis, which is plotted as ΔFnorm to yield a binding curve, which can be fitted to derive binding constants. Figure and legend obtained under a Creative Commons license with minor modifications [333].   73  fluorescence at emission wavelengths of 330 nm and 350 nm. This non-labeling approach is typically only feasible when the titrating ligand does not absorb at those wavelengths (e.g. small peptides, small molecules). The main advantage of MST is the ability to work with samples in different types of solvent in solution (e.g. in plasma). Furthermore, the sample requirements are very low, using less than 10 µL of sample for a complete binding isotherm.  2.4.3.1 Experimental conditions MST was performed at 25 °C with medium MST power and varying LED power to ensure the minimum amount of fluorescence counts are met. As a cost-saving measure, capillaries were split into two using a zirconium blade and nitrogen gas was passed through the capillaries to remove resulting glass dust. Controls were performed using uncut vs cut capillaries to ensure reproducibility. Two types of capillaries were used: Standard Treated and Premium Coated. Unlike the Standard Treated capillaries, Premium Coated have a hydrophobic polymer coating in the capillary tubes that prevent adsorption of molecules to the capillary walls. This is particularly important for proteins or molecules that display strong adsorption to the walls, thereby giving rise to a non-uniform signal. Other methods to prevent adsorption include the use of a carrier e.g. BSA or detergents e.g. Tween-20. Premium Coated capillaries were only used when significant adsorption to the capillary walls was noted (as flagged by the MST software). MST samples were read three times as technical replicates. The readings were performed at different sections of the capillary to prevent photobleaching and fluorescence exhaustion. Binding isotherms are derived from at least 12 different concentrations and were performed in at least three independent replicates. The Fhot values (Figure 21D) used were typically between 5 to 74  10s and data analyses used the same Fhot value for each specific binding interaction. Curve fitting was tested with two equations:  (1) Simple binding curve: A single interaction between two molecules is characterized by 𝐾𝑑 =[𝐴]∙[𝑇][𝐴𝑇], where A is the protein being monitored (e.g. fluorescent), T is the titrant, and AT is the resulting complex. Where A0 and T0 are the initial concentrations added and A and T are the free/unbound concentration, the law of conservation of mass dictates  [𝐴0] = [𝐴] + [𝐴𝑇]  and [𝑇0] = [𝑇] + [𝐴𝑇] . These are used to derive the quadratic equation: [𝐴𝑇] =12([𝐴0] + [𝑇0] + 𝐾𝑑 ± √([𝐴0] + [𝑇0] + 𝐾𝑑)2 − 4 ∙ [𝐴0] ∙ [𝑇0]). To convert Fnorm to [AT], the following equation was used: 𝐹𝑛𝑜𝑟𝑚 = ([𝐴][𝐴0]∙ 𝐹𝑛𝑜𝑟𝑚,𝐴) + ([𝐴𝑇][𝐴0]∙ 𝐹𝑛𝑜𝑟𝑚,𝐴𝑇) 𝐹𝑛𝑜𝑟𝑚,𝐴  is the normalized fluorescence signal when A is not bound to T. Conversely, 𝐹𝑛𝑜𝑟𝑚,𝐴𝑇 is the corresponding signal when A is bound to T. (2) Hill equation: This equation describes binding events where ligand binding shows cooperativity or are not simple 1:1 binding,  𝐹𝑛𝑜𝑟𝑚(𝑐) = 𝑈𝑛𝑏𝑜𝑢𝑛𝑑 +𝐵𝑜𝑢𝑛𝑑−𝑈𝑛𝑏𝑜𝑢𝑛𝑑1+10𝐻𝑖𝑙𝑙∙log(𝐸𝐶50−𝑐). The concentration of titrant (c) was subject to log10 transformation. “Unbound” is the Fnorm signal of the non-complexed protein whereas “Bound” is the Fnorm signal of the complex. EC50 is the half-maximal effective concentration and “Hill” is the Hill coefficient of cooperativity. 75  2.4.3.2 Protein labeling with fluorophores.  MST requires one of the target proteins to be fluorescently labeled. There were two labeling strategies used for this project: amine-reactive crosslinking and His-tag affinity labeling. All fluorophores (Alexa Fluor 647 and NanoTemper RED 2nd generation dye) absorb strongly at the infrared range with an approximate excitation max of 650 nm and emission max of 670 nm.  2.4.3.3 Amine-reactive crosslinker reactive groups Linkage of the fluorophore through N-hydroxysuccinimide (NHS) ester chemistry is dependent on the availability of free primary amines, particularly lysine residues on proteins. As such, the conjugation is also pH-dependent as at low pH, the primary amine is protonated and cannot react with the NHS ester. The optimal pH is 8.3-8.5 in a primary amine-free (e.g. carbonate-bicarbonate or HEPES) buffer. Typically, NHS esters are dissolved in an anhydrous DMSO or DMF as NHS esters hydrolyze quickly to an inactivated form in water. As a small amount of fluorophore-NHS ester is needed, the powder was dissolved in acetone and small aliquots were made before drying under a vacuum with a Speed-vac (Savant, Thermo Scientific). The aliquots were stored at -20 °C and used fresh each time. Both the NanoTemper and ThermoFisher kits follow a similar protocol involving buffer exchange of protein, conjugation of dye to protein and separation of unconjugated from conjugated dye. For consistency, the proteins were buffer-exchanged to an appropriate buffer at pH 8.5. Conjugation of freshly prepared Alexa Fluor 647 dye took place in the dark at room temperature with a final dye:protein molar ratio of 10:1, achieved through three 15-min steps of 3.33:1. For the NanoTemper RED-NHS 2nd generation dye, conjugation took place in the dark at room temperature with a ratio of 2.5:1 for 30 min, following the manufacturer’s instructions. The 2nd 76  generation dye has improved amplitudes and signal to noise ratios compared to 1st generation. To remove non-conjugated dye and non-labeled protein, the conjugation samples were eluted through a Sephadex G-25 PD-10 column (for Alexa Fluor 647) or through the kit-supplied gel filtration column (NanoTemper).   2.4.3.4 Histidine tag affinity labeling A non-covalent fluorophore that is attached to tris-nitrilotriacetic acid (tris-NTA) was used to investigate protein interactions in solution since sTF-His and sgC-His contain His-tags. This nickel-chelating group allows the fluorophore to bind to the His-tag in solution and would have a lesser potential of an impact on the protein function. The unbound fluorophore also does not have to be removed during application. A downside to this application is the need for high nanomolar concentrations of protein to maintain binding the tris-NTA fluorophore. Furthermore, it prevented the use of nickel-chelating liposomes to facilitate membrane-protein complex assembly. Nonetheless, I used this tris-NTA fluorophore to probe for interactions in solution.  The tris-NTA-linked fluorophore is water-soluble and was maintained at -20 °C after rehydration in water or a buffer of choice. To allow time for the tris-NTA to bind to the His-tag, the fluorophore was incubated with the target protein for 1 h at room temperature. For sTF-His binding, a 4X stock of 400 nM sTF-His and 14 nM of tris-NTA linked fluorophore was prepared. At 1X, this allowed for ~11000 fluorescent counts (arbitrary units) with 20% LED power. For sgC-His, a 4X of 800 nM sgC-His and 14 nM of tris-NTA linked fluorophore was prepared. Standard treated cut capillaries (NanoTemper) were used with 0.1% BSA as a carrier. However, there was significant adsorption to the capillary walls. This was resolved with the inclusion of 0.01% Pluronic F-127, an inert non-ionic surfactant, as suggested by the manufacturer (NanoTemper). 77  2.4.3.5 Determining protein concentration and degree of labeling The degree of labeling (ratio of dye to protein) was measured spectroscopically by measuring absorbances at 280 nm and 650 nm. Formulae for calculating protein concentration and protein to dye ratio is as follows: 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 (𝑀) =[𝐴280 − (𝐴650 × 𝐶𝐹280)] × 𝑑𝑖𝑙𝑢𝑡𝑖𝑜𝑛 𝑓𝑎𝑐𝑡𝑜𝑟 𝜀𝑝𝑟𝑜𝑡𝑒𝑖𝑛(𝑐𝑚−1𝑀−1)  𝑑𝑒𝑔𝑟𝑒𝑒 𝑜𝑓 𝑙𝑎𝑏𝑒𝑙𝑖𝑛𝑔 =  𝐴650 × 𝑑𝑖𝑙𝑢𝑡𝑖𝑜𝑛 𝑓𝑎𝑐𝑡𝑜𝑟𝜀𝑑𝑦𝑒 × 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 (𝑀) where, CF280 is the correction factor for the dye’s absorbance at 280 nm. 𝜺dye is the extinction coefficient in cm-1 M-1. Alexa Fluor 647 has a CF280 of 0.03 and an 𝜺dye of 239,000 cm-1 M-1. NanoTemper RED (2nd generation dye) has a CF280 of 0.04 and 𝜺dye of 195,000 cm-1 M-1.  To further assess purity, the proteins were subject to SDS-PAGE and the protein bands were visualized by infrared illumination using the LI-COR Odyssey Imaging System. Alexa Fluor 647 labeled sgC-His, Alexa Fluor 647 labeled sTF-His, and NanoTemper RED labeled FVIIa had degrees of labeling of 1.4, 2.1, and 0.9, respectively.  2.4.4 Removal of His-tag on sTF-His by cyanogen bromide (CNBr) cleavage.  Cleavage protocol was adapted as previously described [334]. Briefly, protein solution was buffer-exchanged with 20 mM sodium phosphate buffer, 150 mM NaCl, pH 7.5 using an Amicon® 0.5 mL 10K molecular weight cut-off centrifugal filter. HCl and CNBr (dissolved in acetonitrile) was added to 100 µM sTF-His to a final concentration of 0.1 M HCl and 10 mM CNBr. Ratio of 78  methionine residues to CNBr was 100:1. After 24 h of cleavage in the dark at room temperature, a SpeedVac was used to dry and stop the reaction. The solid was resuspended in HBS and 2.5 M Guanidine-HCl (GdnHCl) [335]. Cleaved sTF (sTF-Cys) was retrieved in the flow-through following Ni-NTA metal affinity chromatography. HBS was added slowly to the flow-through with mixing until 0.25 M GdnHCl was obtained. Samples were spun at 15,000 x g for 30 min prior to concentrating and buffer-exchange in a Microcon®. Western blot analysis of sTF-Cys using an anti-His-tag antibody (1:665) revealed that the His-tag was removed (Figure 22). In addition, sTF-Cys function was tested using chromogenic functional assays and sTF-Cys binding to FVIIa was confirmed by MST (not shown).  79           Figure 22. sTF-His cleavage by CNBr removes the C-terminus His-tag. sTF samples (0.25 µg) were run in a non-reducing 10% polyacrylamide gel. sTF-Cys lacks the His-tag as revealed by Western blot using an HRP-conjugated anti-His-tag antibody (1:665). sTF-Cys #1 was re-folded after a GdnHCl denaturing nickel affinity column by 10 slow step-wise dilutions of HBS to a final [GdnHCl] of 0.25 M (1:1.25 dilution each step). sTF-Cys #2 was diluted with 5 step-wise dilutions (1:2 dilution each step) to the same final [GdnHCl]. 80  Chapter 3: HSV1 glycoprotein C and its effect on TF/FVIIa complex In this chapter, I will be addressing my hypothesis that gC aids in FX activation through enhancing TF function. This will be accomplished by first demonstrating TF presence and function on the virus surface. Following this, variations of HSV1 with or without gC and/or TF will be used to establish the effect of gC on TF-mediated FX activation on the virus surface. gC function was then pursued under purified, virus-free systems whereby binding partners can be determined and their direct effects on overall FX activation attained in the absence of additional contributing envelope constituents.  3.1 TF function on the virus surface Previous work from the Pryzdial laboratory demonstrated that procoagulant phospholipids existed on several herpesviruses and could support prothrombinase complex formation [2,336]. Further investigation revealed that herpesviruses grown in TF-bearing cells could express envelope TF [2]. This was shown by flow cytometry as well as single immunogold EM. As it had been demonstrated that the virus envelope protein gC was involved in TF-mediated FX activation, single virus particles were probed for the simultaneous availability of TF, gC and aPL on their surface.  3.2 Procoagulant factors on the virus surface Immunogold EM analysis of wildtype HSV1 was used to demonstrate that the host procoagulant factors, TF and aPL, as well as the viral procoagulant factor gC are on the same virus surface. A multiple labeling strategy involving the use of different sized gold beads to differentiate antigens was successfully employed to simultaneously detect all procoagulant factors on a single 81  virus particle (Figure 23). This technique would also be translated to my second hypothesis as it confirms the identity of virus particles containing procoagulant factor(s). HSV1 antigen expression levels appeared varied as seen in the single labeling controls compared to the triple labeling (Figure 23). However, stoichiometry cannot be inferred from immunogold EM as antibody or probe access to antigens can be limited by several factors. Nonetheless, HSV1 incorporated TF, aPL as well as gC on the same envelope, thereby supporting a gC/TF interaction. An important technical EM observation made for HSV1 was that it readily aggregated. This technical issue was addressed by pre-treating the EM grids with 10 µg/mL anti-HSV1-positive human IgG. Antibody-mediated retention of the virus on the grid served to prevent the virus from repositioning during the labeling procedure. The purified human IgG1 was derived from NP and was tested positive for reactivity to HSV1, the importance of this will be alluded to in plasma clotting studies (Section 3.7.1). Initial labeling strategy consisted of a rabbit antibody against TF; however, the relatively low amounts of TF compared to other virus antigens required the use of high concentrations of TF antibody. This resulted in high background when an isotype control was used. HSV1 encodes the transmembrane glycoproteins gE and gI, which form a heterodimer on the virus surface as well as the infected cell surface that can bind to Fc regions of antibodies [338,339]. The gE-gI Fc receptor binds more avidly to rabbit IgG than other species’ [340]. This could explain why the rabbit IgG isotype control displayed high background compared to the near-zero background in mouse or goat isotype controls at similar concentrations. Thus, the TF and gC reporting antibodies were switched such that the TF antibody was of mouse origin and the gC antibody was of rabbit origin. gC is more abundant on the virus compared to TF as determined by western blot (Figure 13) and as observed in the initial labeling strategy using the mouse  antibody. This  allowed  us  to  eliminate  82     Figure 23. Initiators of coagulation found on the virus surface. Immunogold electron microscopy on HSV1. HSV1 was confirmed by presence of gC, size and morphology. TF and aPL are found on the surface of HSV1. Two different labeling strategies were used. (A) mouse anti-gC: 6 nm gold-conjugated goat anti-mouse; aPL probe (annexin V): 10 nm gold-conjugated goat anti-biotin; rabbit anti-TF: 15 nm gold-conjugated goat anti-rabbit. (B) rabbit anti-gC: 15 nm gold-conjugated goat anti-rabbit; aPL: 10 nm gold; mouse anti-TF, 9B4: 6 nm gold-conjugated goat-anti-mouse. Controls used were isotype controls for gC/TF and inclusion of EDTA for aPL probe. Scale bar: 100 nm.  83  rabbit IgG background by lowering the antibody concentration while still facilitating gC detection. In addition, the TF antibody concentration could be increased to better detect for the less abundant protein. These modifications revealed that HSV1 particles incorporated the host procoagulant factors TF and aPL into the same envelope as well as the virus procoagulant factor, gC. However, the presence of these markers does not demonstrate functionality but confirms the possibility.  3.2.1 Effect of gC on TF function The presence of TF procoagulant function on HSV1 was previously tested through inhibition assays [258]. TF-specific inhibitory antibodies resulted in ~90% inhibition of FX activation by FVIIa on HSV1/TF+/gC- (~90%). However, in these earlier studies only 40% inhibition was achieved on HSV1/TF+/gC+, suggesting that both TF and gC are involved in FXa generation on the virus. To quantify the individual contributions of TF and gC on the virus surface, I used HSV1/gC+ or HSV1/gC- propagated in TF-inducible human carcinoma cells to produce HSV1/TF+/gC+, HSV1/TF+/gC-, HSV1/TF-/gC+, and HSV1/TF-/gC-.  HSV1/TF-/gC+ displayed <2% FVIIa-mediated FX activation compared to HSV1/TF+/gC+ (Figure 24A). The presence of gC enabled approximately 3-fold enhancement of FVIIa-mediated FX activation by TF-competent virus, as compared to HSV1/TF+/gC-. This contrasts with TF-deficient virus, where gC had no noticeable impact on the amount of FX activation under these conditions (Figure 24B). Thus, viral gC enhancement of FVIIa activity is appreciable only in the presence of TF on the virus surface.  The viruses were grown in sister cells to minimize variability in protein expression during infection. Interestingly, western blot and densitometry analyses of equal virus particle numbers displayed differences in TF protein expression per virus (Figure 25A). For this virus preparation,  84        Figure 24. gC enhances TF-dependent FX activation on the virus envelope. (A) FX activation was followed using the panel of HSV1: TF+/gC+ (⬤), TF+/gC- (○), TF-/gC+ (◼), and TF-/gC- (◻) viruses. (B) Magnification of TF-/gC+ and TF-/gC-. 100 nM FX and 1 nM FVIIa was mixed with HSV1 prior to the addition of 5 mM Ca2+ to initiate the reaction. After 20 min at 37 °C, the samples were stopped with EDTA and chromogenic substance S-2765 was added to determine the generation of FXa spectrophotometrically. N = 4 independent replicates; Error bars: standard error of the mean (SEM).   85  HSV1/TF+/gC+ had an average of 3300 ± 520 molecules of TF/vp whereas HSV1/TF+/gC- had 2600 ± 410 molecules of TF/vp (Figure 25B, N = 4, p = 0.03, Student’s t-test). The major capsid protein (MCP) was used to compare the amount of capsid-containing virus particles, because MCP comprises the viral capsid and its content is constant due to specific capsid architecture. Of note, there was a two-fold decrease in the total MCP when gC was absent with the same amount of virus particles loaded. Even when factoring in the 30% difference in TF, the amount of FX activation generated by HSV1/TF+/gC+ was more than 2-fold greater than HSV1/TF+/gC- (Figure 25C).  3.2.2 Effect of soluble gC to viral TF It was previously demonstrated that a transmembrane domain-truncated form of gC (sgC-His) enhanced FVIIa-mediated FX activation on the virus surface [258]. Here, sgC-His was incrementally added to the purified HSV1 variants to determine if sgC-His binding to virus was dependent on TF. FX activation was increased ~2-fold by the addition of sgC-His due to the presence of TF on the virus (Figure 26A). Addition of sgC-His to TF-deficient virus had a relatively small effect on FX activation (Figure 26B). The effect of sgC-His on FXa generation was greatest in the following order: HSV1/TF+/gC+ > HSV1/TF+/gC- >> HSV1/TF-/gC+ > HSV1/TF-/gC- (Figure 26C and D). Since sgC-His enhanced HSV1/TF+/gC+ more than HSV1/TF+/gC- suggests that another factor besides gC and TF could be affecting sgC-His-mediated enhancement of FXa production.  86   Figure 25. Difference in viral TF expression does not explain gC-mediated FX activation. (A) Western blot of 109 virus particles of HSV1/TF+/gC+ or HSV1/TF+/gC- loaded on a reducing 5-15% polyacrylamide gel. TF and MCP were probed simultaneously with mouse monoclonal antibodies that were demonstrated to have zero cross-reactivity. Purified sTF-His was titrated as a standard curve for quantitatively determining [TF] of each virus by densitometry. N = 4 independent replicates; Error bars: SEM. (B) Densitometry-derived concentrations of TF were converted to number of TF molecules per virus particle (vp) for each virus. (C) By converting the vp concentration to [TF], the effect of gC on FVIIa-mediated FXa generation by HSV1/TF+/gC+ (⬤) and HSV1/TF+/gC- (○) was assessed as a function of [TF]. 87    Figure 26. Addition of sgC-His to HSV1 enhances FVIIa-mediated FX activation. Purified sgC-His (titration), FX (100 nM) and FVIIa (1 nM) were combined with purified HSV1 variants: (A) HSV1/TF+/gC+ (⬤) or HSV1/TF+/gC- (○) and (B) HSV1/TF-/gC+ (◼) or HSV1/TF-/gC- (◻) viruses in the presence of CaCl2 (5 mM) and FXa generation was followed using the chromogenic substrate S-2765. (C and D) sgC-His-dependent FXa generation was determined by subtracting baseline virus-mediated FX activation. N > 3 independent replicates; Error bars: SEM.  88  3.2.3 Effect of soluble gC to relipidated TF To exclude other virus- or host-encoded factors associated with the envelope of purified HSV1, sgC-His was added to Innovin as a source of relipidated full-length TF and aPL. Enhancement of FVIIa-dependent activation of FX by sgC-His was demonstrated. The activity of FVIIa was saturable, indicating occupancy of a cofactor(s) (Figure 27A). FX (Figure 27B), the reaction substrate, also reached saturation, which is consistent with either a FVIIa cofactor that docks the substrate or maximal FVIIa-binding. The addition of 2 µM sgC-His increased maximal activation by ~2-fold. However, the inability to vary the specific components of Innovin prevented detailed assessments of TF and/or aPL contribution(s) to the gC/TF/FVIIa interaction. Furthermore, it is uncertain whether gC requires membrane-association for full functionality. Hence, a defined and versatile system was required to ascertain the importance of each component in FX activation.   3.3 gC function in a divisible system The function and manner by which gC affects FX activation is not clear. Especially in the case of sgC-His, its ability to enhance FX activation is FX- and FVIIa-dependent but is relatively weak in solution and in the absence of relipidated TF. Membrane association is important for TF cofactor function. Hence, sgC-His function may also be restricted from a lack of membrane association/interaction. Soluble TF can enhance FVIIa-dependent FX activation in the presence of aPL and Ca2+ [328]. Yet, its cofactor activity is 1000-fold less than its native membrane-bound form. sTF-His is known to optimally enhance FX activation by FVIIa when DGS-NTA-Ni is incorporated into an aPL-containing membrane [318]. The DGS-NTA-Ni enables membrane-binding of sTF-His and the aPL enables Ca2+-dependent membrane-binding of FVIIa and FX.  89         Figure 27. sgC-His effect on FX activation by relipidated TF. The effect of sgC-His on FX activation by relipidated full-length TF (1:3000, Innovin) was assessed chromogenically using S2765 in the presence (●) or absence (○) of sgC-His (2 µM) at constant FX (100 nM, A) or FVIIa (10 nM, B). N = 3 independent replicates; Error bars: SEM.    90  Thus, suggesting orientation and membrane association is important for TF biological activity. This approach was adapted for investigating sgC-His function as the protein also carries a C-terminal His-tag.   3.4 Effect of sgC-His on TF function in a purified and defined system The respective contributions of cofactors were evaluated by comparing vesicles toward increasing the activity of FVIIa by sgC-His. As negative controls, none of the various lipid compositions yielded appreciable FX activation by FVIIa in the absence of sTF-His or sgC-His (Figure 28A, lanes 1-3). This low activity persisted when the essential components, Ca2+, FX or FVIIa, were omitted in the presence of sgC-His (Figure 28A, lanes 4-6). As a positive control for the anticipated activity of the three vesicle compositions, FX activation was shown to be enhanced by sTF-His in the following order PCPS << NiPC < NiPCPS (Figure 28A, lanes 7-9). Interestingly, a cofactor effect on FVIIa was similarly observed for sgC-His (Figure 28A, lanes 10-12). Based on previous findings, I expected minute FX activation (pM) [258]. However, there was 100-fold higher activity and furthermore, required 1000-fold lower concentration of sgC-His. Exclusion of phosphatidylserine (PS) from the Ni-chelating vesicles resulted in a 4-fold decrease in FX activation by sgC-His.  To ensure no contaminating proteases were found in the sgC-His preparation, sgC-His was pre-treated with the protease inhibitor p‐amidinophenylmethylsulfonylfluoride (APMSF). APMSF is an irreversible inhibitor of trypsin-like serine proteases and quickly hydrolyzes into an inactive form in aqueous solution. No effect of APMSF was observed on the activation of FX catalyzed by sgC-His nor did high concentrations of sgC-His (5 µM) in the absence of FVIIa result in any FX activation (data not shown).  91     Figure 28. Ni-chelating lipids enhance sgC-His and sTF-His cofactor function. (A) FX activation was monitored by variably assembling tenase constituents on 50 µM PCPS, NiPC or NiPCPS in the presence or absence of 1 pM sTF, 1.2 nM sgC-His, 1 nM FVIIa, 100 nM FX, 5 mM Ca2+. (B) FXa generation by 0, 1, 10 or 100 nM sgC-His (from white to dark bars) in the presence of 0, 10 or 200 pM sTF-His was investigated for synergy with 100 nM FX, 5 mM Ca2+ and limiting concentration of FVIIa (10 pM). N = 3 independent replicates; Error bars: SEM.  92  To explore the apparent synergistic effect of gC and TF on the virus (Figure 27), sTF-His was combined with sgC-His. Unlike, the HSV1 variant experiments, the effect of sgC-His on sTF-His was additive in these purified protein experiments (Figure 28A, lanes 13-15). As this was performed with non-saturating amounts of sTF-His (relative to the concentration of FVIIa), it was postulated that perhaps sgC-His favored binding to unbound FVIIa in the presence of sTF-His. And so, the experiment was set-up to have a limiting concentration of FVIIa, for which sTF-His and sgC-His would either compete or interact simultaneously with FVIIa (Figure 28B). In this scenario, I found that sgC-His did not have any enhancing effect on sTF-His/FVIIa beyond sTF-His saturation of FVIIa cofactor activity. This would suggest that sTF-His competes with sgC-His for binding to FVIIa. Surface plasmon resonance studies by another group on relipidated TF binding to FVIIa revealed that TF/FVIIa does not dissociate easily as the koff was immeasurable under their conditions [341]. Nonetheless, the ability of sgC-His to enhance FX activation in the absence of TF prompted further investigation.  3.4.1 FX activation kinetics by sgC-His/FVIIa To assess whether sgC-His behaves differently to sTF-His in its ability to enhance FX activation, the kinetics of FX activation was followed with FX chromogenic assays. To determine this, the affinity of cofactor to NiPCPS and NiPC was quantified by FX chromogenic assays. As PCPS with sgC-His did not show appreciable amounts of FX activation unless at µM concentrations and the cofactors lack any known membrane-tethering domains without Ni-chelating groups, PCPS affinity was not determined. When titrating NiPCPS or NiPC, the maximal rate of FX activation was higher for sTF-His than sgC-His (Figure 29A and B). This is explained by the difference in affinity of cofactor to FVIIa and the sensitivity of the assay. At lower than 100   93    Figure 29. sgC-His and sTF-His binding to NiPCPS or NiPC vesicles. Binding of cofactors to (A) NiPCPS or (B) NiPC was assessed by following continuous FX activation by the assembled cofactor/FVIIa complex. NiPCPS and NiPC SUVs were titrated with either 120 nM sgC-His (⬤) or 100 pM sTF-His (■) in a mixture containing final concentrations of 100 pM FVIIa and 5 mM Ca2+. 30 nM FX and 0.2 mM S-2765 was added to initiate the reaction. FX activation was followed over time to determine the initial rate of FX activation. This rate was converted to concentration of cofactor/FVIIa complex to determine the apparent dissociation constants for sgC-His or sTF-His to NiPCPS or NiPC (C and D, respectively). N = 3 independent replicates; Error bars: SEM.  94     Figure 30. Binding of sgC-His or sTF-His to FVIIa in the presence of NiPCPS or NiPC. Binding of sgC-His and sTF-His to FVIIa was determined by continuous FX chromogenic assays in the presence of (A) NiPCPS or (B) NiPC. sgC-His (⬤) or sTF-His (■) was titrated in a mixture with 100 µM NiPCPS or NiPC, 10 pM FVIIa and 5 mM Ca2+. 30 nM FX and 0.2 mM S-2765 was added to initiate the reaction. FX activation was followed over time to determine the initial rate of FX activation. This rate was converted to concentration of cofactor/FVIIa complex to determine the apparent dissociation constants for sgC-His or sTF-His to FVIIa (C and D, respectively).  N = 3 independent replicates; Error bars: SEM. 95          Table 2. Binding affinities of sTF-His and sgC-His as derived from linked enzyme kinetics.   Nickel binding FVIIa binding Ligand Lipid Kd,app (µM) (95% CI) Kd,app (nM) (95% CI) sTF-His NiPCPS 33.8 (23.0 – 44.4) 7.0 x 10-3  (4.4 x 10-3 – 9.6 x 10-3) sTF-His NiPC 13.4 (5.6 – 21.2) 8.9 x 10-3  (2.9 x 10-3 – 1.5 x 10-4) sgC-His NiPCPS 48.0 (26.0 – 70.0) 21.5 (18.3 – 24.7) sgC-His NiPC 19.1 (7.4 – 30.7) 18.0 (8.7 – 27.3)    96         Table 3. Kinetics of FX activation as derived from linked enzyme kinetics.  Cofactor Lipid Km (µM) kcat (s-1) kcat/Km (µM-1 s-1) sTF-His NiPCPS 0.16 ± 0.02 3.8 ± 0.5 23.6 ± 1.4 sgC-His NiPCPS 0.14 ± 0.02 2.5 ± 0.3 16.6 ± 1.0    97  pM FVIIa, titrating phospholipid resulted in immeasurable FX activation (not shown). Nonetheless, transformation of the initial rate of FX activation to concentration of cofactor/FVIIa complex (Figure 29C and D) revealed that sTF-His and sgC-His have similar affinities for NiPCPS and NiPC, as summarized in Table 2. Contrary to my expectations, the hexa-His-tag sTF-His did not have a tighter affinity than the penta-His-tag sgC-His. It is interesting to note that NiPC appeared to have a higher affinity for the cofactors than NiPCPS.   3.4.2 sgC-His-induced plasma clotting  When 1 µM sgC-His was added to NP, no clotting was observed in the presence of 50 µM PCPS vesicles after recalcification with 10 mM CaCl2 for over 15 min (Figure 31). With the substitution of PCPS with NiPCPS, clot formation time was 136 s ± 11 s. Of note, the batches of NP used contained antibodies against HSV1, including gC (Figure 32), accredited to the prevalence of HSV1 within the adult population. The presence of inhibitory antibodies could explain the high concentration of gC required to initiate clotting.  3.5 Interactions between TF and gC 3.5.1 Cross-linking on the virus surface As HSV1/TF+/gC+ has enhanced TF/FVIIa activity compared to gC-deficient, it was speculated that an interaction between TF and gC existed. Crosslinking experiments of TF+/- and gC+/- HSV1 using the thiol-reduceable DTSSP did not reveal any detectable TF-gC interaction (Figure 33). There was, however, more monomeric TF in HSV1/TF+/gC+ compared to the HSV1/TF+/gC-. Of note, gC appeared as two distinct bands of similar quantity under low exposure in the HSV1/TF+/gC+. HSV1/TF-/gC+ only had the larger form of gC (Figure 33B, lane 2/8). 98            Figure 31. sgC-His-induced plasma clotting. Normal pooled human plasma clotting was initiated by the inclusion of sgC-His (1 µM), 50 µM PCPS or NiPCPS, and 10 mM Ca2+. Clotting was followed for up to 900 s. N > 5 independent replicates; box plot.   99  Since the virus contains other gC crosslinking partners, the proportion of gC interacting with TF may be low and undetectable in the purified virus setting. To address this, a recombinant form of gC lacking its transmembrane domain (sgC-His) was produced to probe for TF interactions in a more controlled setting without extraneous virus- or host-derived factors. This form of gC retains its complement inhibiting and heparin binding functions as well as conformation [320].  3.5.2 Cross-linking of purified proteins gC forms complexes with other viral glycoproteins, thus perhaps only a small proportion of membrane gC is bound to TF. To increase the interaction between TF and gC, purified sTF-His and sgC-His were incubated with Ni-chelating membranes (NiPCPS, 50 µM) to mimic a virus membrane-bound TF and gC. However, DTSSP crosslinking of sTF-His to sgC-His did not result in any sTF-His/sgC-His adducts (Figure 34). Poor or a lack of crosslinking is not dismissive given that TF has been previously shown to be unable to crosslink with FVIIa using DTSSP [334].  3.5.3 ELISA-modified approach An ELISA-based approach was employed to determine sgC-His/TF interactions. Full-length relipidated TF (Innovin) was adsorbed to the bottom of the wells, and sgC-His was incubated at varying concentrations. After thorough washing, the amount of bound sgC-His was determined by antibodies. No sgC-His was found to be bound to the wells. With the opposite approach using adsorbed sgC-His, no TF was bound. To enhance detection, a chromogenic assay was employed and monitored the assembly of TF/FVIIa and subsequent FX activation. However, sgC-His did not retain TF nor was TF enhanced by sgC-His in either adsorption scenarios. sgC-His may have a low affinity for TF or the complex is not stable when gC is not membrane-bound. 100           Figure 32. Antibodies against HSV1 exist in normal human pooled plasma-derived IgG. Human IgG purified from NP by Protein G agarose chromatography was used as a primary antibody to detect for virus proteins from different forms of HSV1 (HSV1/TF+/gC+, HSV1/TF+/gC-, HSV1/TF-/gC+, and HSV1/TF-/gC-) that were separated by SDS-PAGE (1 x 1010 virus particles). gC is highly antigenic and sgC-His was detected (100 ng).   101        Figure 33. Crosslinking of proteins on HSV1 surface by DTSSP. HSV1 panel (2 x 1010 vp each virus) was subjected to the 2.5 mM DTSSP crosslinker for 15 min on ice. The reaction was then quenched with 1 M Tris-glycine (pH 7.0). A negative control reaction where Tris-glycine (pH 7.5) was added prior to the cross-linker was also performed. Reduced and non-reduced samples were loaded on a 5-15% gradient gel and detected by Western blot using polyclonal antibodies against TF and gC.    102        Figure 34. Crosslinking of purified sgC-His and sTF-His on NiPCPS vesicles. 12 µM sTF-His was mixed with 2.5 µM sgC-His, 50 µM NiPCPS and 5 mM Ca2+. DTSSP (500 µM) was incubated with the protein mixture for 24 h at 4 °C with agitation. Reduced and non-reduced samples were loaded on a 5-15% gradient gel and detected by Western blot using polyclonal antibodies against TF and gC.   103  3.5.4 Microscale thermophoresis As an alternative approach to searching for sTF-His/sgC-His interactions, microscale thermophoresis (MST) was employed. The main advantage of this technique is that it can follow interactions of molecules in solution without tethering one of the binding partners onto a surface. In addition, miniscule amounts of reagent are required to derive an entire binding isotherm (<12 µL). To monitor an interaction, one of the ligands must be fluorescently labeled through amine-reactive probes, thiol-reactive probes, or non-covalent affinity probes (e.g. His-tag probes).  3.5.4.1 Fluorophore labeling of sTF-His There are two important lysines in the FVIIa-interacting domain of TF [31]; thus labeling of sTF-His using amine-reactive probes could pose a problem. It was shown that these lysines are required for membrane-associated enhancement of FVIIa activity. As expected, amine-labeled sTF-His loss the ability to further enhance FVIIa activity in the presence of NiPCPS. This initial labeling strategy resulted in FVIIa-dependent quenching of amine-labeled sTF-His fluorescence by MST, which was used to derive a binding affinity that was comparable to FVIIa binding to sTF-His in solution (data not shown). No sgC-His direct interaction with amine-labeled sTF-His could be demonstrated. However, initial MST results revealed that the binding affinity of amine-labeled sTF-His to FVIIa in solution is decreased upon the addition of sgC-His. This would hint towards an interaction between sgC-His and FVIIa.   104  3.5.4.2 Fluorophore labeling of sgC-His As amine-labeled sgC-His did not appear to bind to sTF-His in solution. This would suggest that sgC-His displaced labeled sTF-His from FVIIa in solution based on the initial MST results with sTF-His-R. However, amine-labeled sgC-His also did not appear to directly bind to FVIIa or FX in solution. FX binding to gC is known thus amine-labeling may have affected its function. The thermophoresis profile of amine-labeled sgC-His displayed slow/minimal movement. Furthermore, heparin binding to labeled sgC-His resulted in a large increase in mobility of the complex. Taken together, amine-labeled sgC-His was most likely aggregated but was stabilized into monomers by heparin.  An alternate labeling approach consisted of a fluorophore conjugated to a hist-tag probe (tris-NTA-RED). As sgC-His and sTF-His can both bind to tris-NTA-RED, a His-tag needed to be removed from one of the proteins. sTF-His has a single methionine residue located at 9 amino acids upstream the His-tag (and spacer). Thus, cyanogen bromide specific cleavage of methionine was used to cleave off the His-tag. Functional studies confirmed the absence of the His-tag while retaining FVIIa cofactor activity in solution. Yet, no sTF-Cys/sgC-His interaction was measurable (Figure 35).   3.6 Interactions between gC and FVIIa 3.6.1 sgC-His binding interactions in solution-phase sgC-His conjugated to fluorescent RED-tris-NTA via the His-tag (sgC-His-R) was followed by MST for interactions with FVIIa or FX in the absence of vesicles (Figure 36). Indicative of a relatively weak association between sgC-His and FVIIa or FX, the binding isotherms did not approach  saturation at the  highest available  concentration of  either ligand and  105        Figure 35. Interaction between sgC-His-R and sTF-Cys was probed using MST. sgC-His interaction with sTF-His was probed by MST. The His-tag of sTF-His was removed by CNBr cleavage, resulting in sTF-Cys. sgC-His (200 nM) was mixed with the RED-tris-NTA fluorescent probe (3.5 nM) for 30 min at room temperature before the addition of sTF-Cys in HBS/0.05% BSA/0.05% Tween-20/5 mM Ca2+. MST power was set at medium with 10% excitation power. Samples were equilibrated at 25 °C prior to instrument reading. N = 2 independent replicates, technical triplicates; Error bars: SD from pooled technical replicates.   106  could not be fit to a binding model. While the solution-phase enhancement of FVIIa activity was independent of sgC-His-R labelling (not shown), RED-tris-NTA prevents NiPCPS-binding. Therefore, I pursued amine-labeling of FVIIa (FVIIa-R) given its newly discovered intrinsic interaction with sgC-His (Figure 36). FVIIa-R retained 62% and 85% function in FX chromogenic assays in the presence of sTF-His or sgC-His, respectively (Appendix A.2). As a positive control, sTF-His binding to FVIIa-R was measured by MST. In the presence of NiPCPS, sTF-His binding to FVIIa had a binding affinity of 1 nM (Figure 37A). This was higher than expected as FX chromogenic assays demonstrated a Kd in the low pM range as well as previous reports (~10 pM) when determined by a linked enzyme chromogenic assay [318]. To reconcile this discrepancy, FX was included to best mimic the conditions of the assay, with the protease inhibitor, benzamidine, to ensure no FX cleavage occurred. MST analysis revealed that FX enhanced sTF-His binding to FVIIa-R on NiPCPS by 30-fold to 30 pM (Figure 37B), which was consistent with the previously reported value.  3.6.2 sgC-His binding interactions on membranes In comparison to vesicles without DGS-NTA-Ni, the ability for sgC-His assembled onto a membrane displayed higher affinity for FVIIa-R (  107   Figure 38A). This was independent of the presence of aPL, implying that a FVIIa-R-membrane interaction was not required. When sgC-His binding to Ni-chelating liposomes was facilitated (NiPC or NiPCPS), the Kd for FVIIa-R was 2.0 and 1.3 µM (Table 1). In the presence of PCPS without DGS-NTA-Ni, binding of sgC-His to FVIIa was weaker and not quantifiable. This was consistent with the solution-phase interaction in the absence of vesicles (Figure 36). The 108  addition of FX further increased the binding affinity of sgC-His for FVIIa-R to NiPC and NiPCPS (Kd; 0.9 and 0.6 µM, respectively) (   Figure 38B).   109     Figure 36. sgC-His can bind to FVIIa in addition to FX. MST was used to follow the interaction between sgC-His and (A) FX or (B) FVIIa. sgC-His (200 nM) was mixed with the RED-tris-NTA fluorescent probe (3.5 nM) for 30 min at room temperature before the addition of FX or FVIIa in HBS/0.05% BSA/0.05% Tween-20/5 mM Ca2+. MST power was set at medium with 10% excitation power. Samples were equilibrated at 25 °C prior to instrument reading. N = 3 independent replicates; Error bars: SEM.   110         Figure 37. sTF-His binding to FVIIa-R is enhanced by FX. MST following the interaction between FVIIa-R (1 nM) and sTF-His on NiPCPS membranes (50 µM) with CaCl2 (5 mM) and benzamidine (2 mM) in the absence or presence of 30 nM FX (A and B, respectively). Concentration is in M units. N = 3 independent replicates; Error bars: SEM.   111   Omission of DGS-NTA-Ni from the aPL-containing vesicles reduced the overall affinity (Kd; 2.7 µM). This suggests that sgC-His-FVIIa complex formation is only enhanced by PS in the presence of FX when DGS-NTA-Ni is absent. This is consistent with the observation by others that the affinity of the Gla-domain of FX is greater than FVIIa (approximately 0.2 µM and 15 µM, respectively). In the current system, FVIIa binding to PCPS was ~65 µM as measured by MST (not shown). The Hill equation fit produced similar Kd values to the simple binding curve fit but with a better fit (as determined by r2 value, Appendix A.3). Although the r2 value for the latter generally indicated a superior fit, the respective Kd and half-maximal effective concentration of titrant (EC50) were similar between data-fit methods. The parameters affecting binding were consequently considered based on the simplest model.   3.6.3 sgC-His/FVIIa amidolytic activity Given the importance of FX in gC-FVIIa binding, the ability of sgC-His to enhance FVIIa activity was further investigated by following FVIIa amidolytic activity by cleavage of the chromogenic substrate S-2288 in the presence of sTF-His or sgC-His and nickel-chelating lipids (Figure 39). Consistent with the MST-derived lower Kd of sTF-His or sgC-His to FVIIa, there was no saturation of enhancement of S-2288 cleavage rate at the concentration used. sTF-His/FVIIa Kd was 1 nM (Table 4) and at 1 nM sTF-His, FVIIa amidolytic activity was not saturated. S-2288 cleavage by sgC-His/FVIIa was 3-fold less than sTF-His/FVIIa at the highest concentrations used (Figure 39B). The presence of aPL did not benefit FVIIa amidolytic activity, as was previously described [35,94].    112     Figure 38. sgC-His interacts with FVIIa with enhancement by FX. (A) MST was used to follow sgC-His binding to fluorescently labeled FVIIa in the presence of 50 µM PCPS (⬤), NiPC (◻) or NiPCPS (▲). (B) Similar to (A) but with the inclusion of 30 nM FX and 2 mM benzamidine to inhibit FVIIa. N ≥ 3; Error bars: SEM.   113                Table 4. FVIIa-R binding affinities as determined by MST.     No FX With FX Ligand Lipid Kd (µM) (95% CI) Kd (µM) (95% CI) sTF-His NiPCPS 8.2 x 10-4  (8.6 x 10-5 – 1.5 x 10-3)  3.0 x 10-5  (8.0 x 10-6 – 5.2 x 10-5) sgC-His NiPC 2.0 (1.3 – 2.8) 0.9 (0.7 – 1.0) sgC-His NiPCPS 1.3 (0.8 – 1.7) 0.6 (0.2 – 0.9) sgC-His PCPS >13.0 2.2 (0.9 – 3.5) sgC-His None >12.0 2.9 (1.5 – 4.2)    114        Figure 39. sTF-His and sgC-His enhancement of FVIIa amidolytic activity. FVIIa catalytic activity was assessed by following FVIIa cleavage of the chromogenic substrate S-2288 in the presence of sTF-His or sgC-His, 5 mM Ca2+, and 50 µM PCPS (white bars), NiPC (grey bars) or NiPCPS (black bars). N = 3 independent replicates; Error bars: SEM.   115  3.7 Coagulation initiated by HSV1  3.7.1 HSV1-mediated plasma clotting To better understand the importance of gC and TF in the context of coagulation, plasma clotting assays were performed. The extrinsic pathway is the major initiator of clotting on TF-containing viruses as those that lack TF require at least 100-fold more virus to achieve similar clotting times in NP (Figure 40A). From my FX chromogenic assays (section 3.2.1), gC was expected to enhance TF-induced clotting in NP but no enhancement was observed (Figure 40A). However, upon removing the analogous FX activating pathway mediated by FVIII using FVIII/DP, the effect of gC in accelerating TF-mediated clotting was observable (Figure 40B). HSV1/TF-/gC- appeared to have faster clotting times in FVIII/DP than HSV1/TF-/gC+. The gC enhancing effect on TF-bearing virus was not observed in FXII-deficient plasma, suggesting that FVIII compensates for the lack of gC-mediated enhancement (Figure 40C).   3.8 Effect of glycosaminoglycans on gC gC has been shown to bind to several sulfated GAGs. Given that the heparin-binding site deletion mutant of gC does not bind FX [241], I expected that blocking of the site would lead to decreased FX activation. Contrary to my expectations, sulfated GAGs in the presence of either sTF-His or sgC-His resulted in enhanced FVIIa-mediated FX activation (Figure 41). Unfractionated heparin (UFH) and chondroitin sulfate (CS) enhanced sTF-His/FVIIa-mediated FX activation to an identical degree, whereas low molecular weight heparin (LMWH) enhanced the most. However, this difference may be a concentration effect. The average weight of UFH and CS from commercial sources is 15 kDa whereas LMWH was 6 kDa. Taken this 2.5-fold difference into account and assuming FXa generation is not saturated, the GAGs would be identical in terms  116   Figure 40. Plasma coagulation by HSV1 is TF-dependent. HSV1/TF+/gC+ (closed circle); HSV1/TF+/gC- (open circle); HSV1/TF-/gC+ (closed square); and HSV1/TF-/gC- (open square) were added to either (A) NP or (B) FVIII/DP. Clotting was initiated with 5 mM CaCl2. N = 4 independent replicates; Error bars: SEM; * p < 0.05; ** p < 0.01. (C) HSV1-induced clotting in NP treated with the FXII inhibitor, CTI. NP was pre-treated with 0, 50, 200 µg/mL CTI before the addition of virus (white, gray and black bars, respectively). N = 2 independent replicates, technical triplicates; Error bars: SD from pooled technical replicates.  117            Figure 41. GAGs are procoagulant in FX chromogenic assay. sTF-His or sgC-His with NiPCPS was pre-incubated with buffer control, 50 µg/mL unfractionated heparin (black bar), 50 µg/mL low molecular weight heparin (dark gray bar), 50 µg/mL chondroitin sulfate (gray bar) or 50 µg/mL hyaluronic acid (dotted bar) prior to the addition of FVIIa (1 nM) and Ca2+ (5 mM). FX (100 nM) was added to initiate the reaction. After 20 min, the reaction was stopped with EDTA. The amount of FXa formed was followed by the addition of 0.2 mM S-2765. N = 2 independent replicates, technical triplicates; Error bars: SD from pooled technical replicates.   118  of specific activity in the enhancement of FX activation with sTF-His. This is not true for sgC-His, where CS enhanced FX activation more than UFH (p < 0.01) and LMWH enhanced FX activation greater than 2.5-fold compared to UFH. Hyaluronic acid is non-sulfated and was not expected to bind or affect sgC-His function. Neither sTF-His/FVIIa or sgC-His/FVIIa-mediated FX activation was significantly affected by hyaluronic acid (p = 0.8 and p = 0.08, respectively).  3.9 Discussion In the present study, I explored the effect of gC on TF function on the virus surface as well as in a purified, virus-free system. TF, gC and aPL were all found on a single virus particle by immunogold EM, suggesting that the three factors could work together to enhance FVIIa activity. TF was required for virus-bound gC to enhance FVIIa-dependent FX activation. Due to the minimal contribution of gC to FX activation in the absence of TF on the virus, it was speculated that gC interacted with TF to synergistically enhance FVIIa activity. However, attempts at teasing out a TF/gC direct interaction was not fruitful.   3.9.1 Interactions between TF and gC Neither crosslinking, ELISA nor MST experiments revealed an association between purified sTF-His and sgC-His or the full-length gC and TF embedded in the virus. This suggests that another constituent on the virus surface is contributing to the enhancement of TF function by gC. Considering that gC may interact with other virus surface proteins (e.g. gB, gD, gE), several options are possible, but not detected by crosslinking. The availability of gC in concert with one of these may contribute to its ability to enhance FX activation [332]. Co-immunoprecipitation may provide novel information on a gC/TF interaction or on common binding partners of gC and TF. 119  gC is known to have several forms during infection. The mucin-like domain of gC contains multiple predicted O-linked glycosylation sites [246]. This domain mediates gC-GAG interaction but undergoes heterogenous post-translational modification that alters the affinity of gC to different GAGs [246,342]. TF has three glycosylated asparagine residues in its extracellular domain but these asparagine residues do not affect its function in FVIIa-dependent FX activation or protease activated receptor signaling [343,344]. It should be noted that the E. coli produced recombinant sTF-His does not contain any glycosylation. The lack of glycosylation may affect sTF-His interaction with sgC-His. However, our crosslinking experiments using viruses containing full-length (and presumably glycosylated) TF did not demonstrate gC interactions. Perhaps the choice of a shorter or a more flexible crosslinker may reveal missed interactions. A small subset of gC could interact with TF through their N-linked glycans via the mucin-like region of gC to affect TF/FVIIa function. The use of Sf9 insect cells may result in a different glycosylation to that of human origin, even though this recombinant expression system was developed because of high yield and similarity to human glycosylation patterns. Furthermore, conformational antibodies confirmed proper gC conformation, implying there is no effect conferred by altered glycosylation compared to the human protein [320]. Nevertheless, this putative difference in carbohydrate may affect the coagulation cofactor function of gC expressed on the purified virus produced in mammalian cells, versus that generated in insect cells. There is no evidence in the literature that suggests TF can bind to GAGs. If a TF-GAG interaction existed, GAGs could mediate an interaction indirectly between TF and gC. The preliminary evidence from section 3.8Error! Reference source not found. indirectly implies a potential mechanism. Heparin is well known in antithrombin-mediated inactivation of clotting proteases but in the absence of antithrombin, heparin is procoagulant [345]. As sulfated GAGs are highly 120  negatively-charged, they can bind to numerous molecules (e.g. antithrombin, selectins, fibronectin) [346]. The size of the sulfated GAGs may act as a scaffold for which FVIIa [347] and FX can bind [348]. Like many GAG-interacting proteins, gC does not bind to the non-sulfated GAGs such as hyaluronic acid [246]. The lack of FXa generation enhancement by hyaluronic acid supported the importance of sulfate groups and the notion of a protein-GAG scaffold. In the current work, GAGs increased the cofactor activity of sgC-His more than sTF-His, this would suggest that GAGs bridge sgC-His to FVIIa/FX. But, a quaternary complex of gC/FVIIa/FXa/heparin and the effect of heparin-dependent inhibitors such as antithrombin in plasma may be interesting to explore.  3.9.2 Nickel-chelating lipids as a tool for discovery The ability of sgC-His to enhance FVIIa-mediated FX activation in the absence of TF was demonstrated through FX chromogenic assays as well as plasma clotting assays. The use of nickel- chelating lipids provided a method of studying sgC-His function. The advantages over full-length expression is that soluble proteins require less manipulation of the protein (e.g. denaturation and refolding) and the purification process is simpler as they are secreted. One downside to this approach is the added complexity of the binding interaction between the histidine-tagged protein and the nickel-chelating lipid results. The head group of the DGS-NTA-Ni is slightly elongated compared to that of PS and PC (Figure 14), and only carries a single nickel ion resulting in a micromolar binding affinity. A single DGS-NTA-Ni can coordinate with two histidine residues. The lack of difference between sTF-His (His6) and sgC-His (His5) binding to phospholipid (Table 2) would suggest that the same ratio of DGS-NTA-Ni is coordinated with the His-tag. As the affinity of sTF-His or sgC-His (~5-7 µM for DGS-NTA-Ni) is similar to that of His6 tags to NTA-121  Ni (10 µM) [349], it is most probable that both sTF-His or sgC-His are bound to the same ratio of DGS-NTA-Ni. Another factor to consider is the distribution of DGS-NTA-Ni on the vesicle membrane. Gla domain-binding to PS can result in an umbrella covering of DGS-NTA-Ni. The Gla domain of FVIIa occupies approximately six phospholipid molecules but when complexed with TF, the total area the complex envelops expands to 23-26 phospholipid molecules [297]. DGS-NTA-Ni made up 15% of the vesicles, giving ~4 DGS-NTA-Ni per TF/FVIIa surface area. For multiple Ni2+ ions to co-ordinate with a single His-tag, the lipids need to be within proximity of one another. Other laboratories have used Tris-NTA derivatives, that chelate three nickel ions. This increases His-tag affinity to a Kd in the nanomolar-picomolar range with the caveat of a bulkier head group [247]. This larger head group could potentially impact protein-membrane interactions but would otherwise ease the difficulties of easily dissociable low affinity nickel-histidine interactions.   3.9.3 Microscale thermophoresis The use of MST allowed for better understanding of complex formation(s). The non-covalent histidine-tag fluorophores provided information on the binding partners for sgC-His. This technique allowed the first demonstration that sgC-His interacts directly with FVIIa. The low cost and low reagent use, as well as the flexibility of manipulating components (e.g. inclusion of anti-adsorptive materials or splitting of capillary tubes) made MST an attractive tool for investigating protein interactions.  122  3.9.4 gC interaction with FVIIa FVIIa/sgC-His complex formation as determined by MST is ~10-fold weaker than determined by chromogenic assay. Labeled FVIIa retained most of its activity and the degree of labeling suggested that only one lysine was labeled. It is, however, unknown whether the same lysine residue was labeled. Benzamidine was added to prevent cleavage of FX and did not affect sgC-His binding to FVIIa in the absence of FX as measured by MST (data not shown). Binding experiments by laboratories using other methods had difficulty corroborating demonstrating affinity in the pM Kd range due to the limited sensitivity of their nM experimental approaches [341]. One group used surface plasmon resonance (SPR) with relipidated TF and measured a pM Kd [341] that was very similar to that obtained by enzymatic assays or fluorescence anisotropy [94,350,351]. However, the TF/FVIIa/aPL complex did not dissociate significantly over the course of the measurement. This causes concern into how an accurate Kd could be derived, which was recognized by the authors. In the current study, the binding affinity of sTF/labeled-FVIIa/FX on a NiPCPS membrane measured by MST approached values (Kd = 32 pM) determined by enzymatic assays.  FVIIa/sgC-His complex formation was greatly enhanced in the presence of FX in the absence or presence of PCPS, suggesting that PS was not required for forming the FVIIa/sgC-His/FX complex. The inclusion of a Ni tether for sgC-His was essential for detectable FVIIa/sgC-His formation in the absence of FX. FX could compensate for a lack of Ni-mediated sgC-His membrane binding. This is consistent with the observation by others that the affinity of the Gla-domain of FX is greater than FVIIa (approximately 0.2 µM and 15 µM, respectively) [29,30]. This suggests that PS contribution to the overall complex is predominantly attributed to the FX Gla-domain interaction.  123  A discrepancy was observed here between MST-derived binding of sgC-His to FVIIa/FX in the absence of DGS-NTA-Ni compared to linked-enzyme derived values. The addition of DGS-NTA-Ni to PC or PCPS increased binding affinity by 2.7- and 7-fold, respectively. Yet, enzymatic activity increased 8- and 40-fold, respectively (Figure 28). While the qualitative enhancement is consistent between techniques, a possible explanation for quantitative discrepancy is the difference in experimental temperature, 25 °C vs 37 °C for MST and enzymatic, respectively. Lower temperatures can affect TF/FVIIa function, with a 60% decrease in FXa generation at 24 °C compared to 37 °C [352]. Furthermore, FVIIa-R retained 85% of its FX activating ability with sgC-His. The remaining difference in sgC-His/FVIIa-R/FX binding versus activation on NiPCPS suggests PS could play a larger role in substrate recognition and proteolytic cleavage by sgC-His/FVIIa than MST binding experiments can distinguish. The ability of sTF-His to substantially enhance S-2288 cleavage by FVIIa unlike sgC-His suggests the latter requires FX binding to affect FVIIa function. This phenomenon is similar to tPA cleavage of plasminogen [353]. The cofactor, C-terminal lysines on fibrin, allows for the substrate (plasminogen) to bind along with the enzyme (tPA). sgC-His binding to the substrate, FX, improves the interaction of sgC-His with FVIIa. This could be due to conformational changes in sgC-His upon FX binding that allow for a tighter sgC-His/FVIIa interaction, sgC-His better presenting FX as a substrate for FVIIa, or FX providing a binding site for FVIIa enhancing the apparent affinity by avidity effects. The latter seems unlikely as I would have expected sgC-His to act in concert with TF to enhance FX activation. In addition, sgC-His/FVIIa can support FX activation to a similar catalytic efficiency as TF/FVIIa when tethered to NiPCPS. This suggests that sgC-His, like sTF-His, can stabilize FVIIa in its active configuration [35,354,355].  124  These data could steer new forms of procoagulant drugs for the treatment of hemophiliacs or to combat bleeds in a controlled manner. To accomplish this, a structure of gC, which has remained elusive, would be instrumental to designing a mimetic. A crystal structure with FVIIa and/or FX may stabilize gC and allow for a solvable structure, similar to that which has been done with sTF/FVIIa.  3.9.5 gC procoagulant activity in infection During the later stages of HSV-1 infection of cells, gC can be efficiently secreted from the cell due to intron splicing within the transmembrane domain of gC [356]. This soluble form of gC could play a role in haemostasis similar to sgC-His. For sgC-His to effectively function, it would require either high local concentrations or a surface to tether and orient itself. Our findings that the addition of sgC-His to HSV1 enhanced its FVIIa-dependent FX activation, suggests that sgC-His is capable of binding to the virus envelope. The partner of this interaction is still unknown although I initially speculated that this may be TF. Coagulation induced by sgC-His in plasma was achievable when NiPCPS was present. This phenomenon required high concentrations of sgC-His which is much greater than expected, as determined by FX chromogenic assays. As most adults in the human population have HSV1, I tested whether antibodies against gC existed in the plasma samples. Indeed, the plasma samples used in the experiments contained anti-HSV1 antibodies that could recognize sgC-His. The presence of these antibodies may explain the need for high amounts of sgC-His to initiate coagulation in these purified experiments. Human endothelial cells increase TF expression on their surface during viral stimulation. HSV1 is known for its wide cell tropism. But in cells that lack inducible TF expression, could gC replace the procoagulant function of TF? gC was shown to be expressed on the cell surface during 125  infection and could bind to FX [240], suggesting that gC plays a role in protease activation on the cell surface. PAR activation could contribute to virus propagation. PAR activation can promote Rab5a expression [357]. Rab5a has been positively linked to HSV1 envelopment and subsequent virus production and secretion [358,359].  3.9.6 Coagulation initiated by HSV1 Enhancement of plasma clotting was evident when TF was present on the virus. Though the role of gC was not discernible in NP, gC enhancement was demonstrated in FVIII/DP. This could suggest a mechanism in HSV1/TF+/gC- that makes use of the amplifying pathway. What was not considered in the experiment was the presence of antibodies against HSV1. Antibodies against HSV1 could result in antibody-mediated complement activation or crowding of antibodies/complement on the virus surface. Complement complexes are large and may prevent clotting protease assembly on the virus surface. As gC inhibits complement protein C3b, it is expected that steric factors due to complement protein assembly would not occur. Thus, gC+ viruses should clot faster than gC-deficient viruses. HSV1/TF-/gC- virus clotted faster than HSV1/TF-/gC+, this has implications.  The putative antibody-mediated complement activation or crowding did not cause a decrease in clotting protease complex formation. The gC- virus is a selective mutant that does not express gC on the infected cell surface and lacked the ability to inhibit complement activation on the virus surface. The faster clotting in FVIII/DP with the mutant strain compared to wildtype could suggest that gC binding to C3b prevents FX interaction on the virus surface. However, if this were true, I would expect the same effect in NP which was not the case. Rather, crowding of proteins on the limited virus surface may play a bigger part in this discrepancy. Experiments using 126  HSV1 seronegative plasma are required to conclude the role of antibodies and complement on HSV1-mediated clot formation. gC could be playing another role in haemostasis. It is also possible that gC affects coagulation downstream of FX activation. Evidence that FXa remained bound to gC after activation was presented in previous work from the Pryzdial laboratory [241]. It is unknown whether gC prevents FVa association with FXa, thereby interfering with thrombin generation. This could also explain why HSV1/TF+/gC+ does not have faster clot formation than HSV1/TF+/gC- in NP. In this manner, the virus can “escape” being bound by a clot and perhaps make use of the PAR activation ability of FXa and/or FVIIa, rather than thrombin.  An unexpected result was that the HSV1/TF+/gC+ was different to HSV1/TF+/gC- in terms of TF and MCP expression per virus particle. The difference could imply that the virus replicates differently, even though the viruses were grown in the same cell. The gC mutant produces a truncated form of gC that is not expressed on the infected cell surface or virus surface and does not exhibit C3b inhibition [360,361]. Although the structure of the gC mutant has not been elucidated, it was speculated that the mutant gC left a “stub” that occupies transmembrane space in the virus envelope, helping to maintain a virus particle surface configuration similar to wild-type virus. Whether this mutant gC affects cellular processes (e.g. envelope formation or TF expression) during infection is not known. Nonetheless, TF on the virus was the main initiator of clot formation and gC enhanced this function.    127  Chapter 4: Tissue factor on enveloped viruses To expand the Pryzdial model of TF on enveloped viruses from the Herpesviridae family to a more general model encompassing many enveloped viruses, I investigated DENV and HCV of the Flaviviridae family of viruses, as well as HIV of the Retroviridae family of viruses. These each pose a significant threat to healthcare and blood transfusion systems around the world. Although DENV affects more than half the world’s population and is a large healthcare burden, DENV is not endemic to Canada. However, parts of the Eastern US were predicted to be suitable habitats for Aedes aegypti and Aedes albopictus, two major mosquito vectors for DENV [362]. Aedes albopictus in particular was estimated to be capable of establishing near Southern Canada. The vector was rarely spotted in Southern Ontario [363] until the past few years where sightings have grown [364,365]. Furthermore, the close relative and arbovirus, West Nile virus (WNV), has been found in many mosquito vectors endemic to Canada [363,366,367]. WNV is an immediate threat, having resulted in over 5000 confirmed cases and 243 deaths in Canada [367,368]. WNV is classified as a Containment Level 3 pathogen whereas DENV is a Containment Level 2 pathogen in Canada. Thus, DENV is highly relevant to Canada and WNV work, less restrictive to work with, and an appropriate in vitro model before investigating other enveloped viruses. The aims of this study are: a) to determine the presence of TF on purified cell culture-derived DENV; b) to establish TF function on purified cell culture-derived DENV; and c) to investigate whether TF is found on patient-derived viruses.   128  4.1 Tissue factor presence on purified dengue virus It was expected that cultured DENV could obtain TF from their cell source, Vero kidney epithelial cells, similar to herpesviruses [2,258]. Picomolar amounts of TF can support enhancement of FVIIa activity, as demonstrated in section 3.4 and by others [32,94,318]. Furthermore, femtomolar amounts of TF can accelerate clotting in plasma [369]. Thus, two methods were used to determine if TF was present in the purified DENV samples: FX chromogenic assay and plasma clotting assay.  4.1.1 FVIIa-mediated FX activation TF activity on purified DENV was investigated by enhancement of FVIIa-mediated FX activation in a chromogenic assay. DENV2 was chosen in all experiments as this serotype is the most common [370–372]. FVIIa-mediated FX activation was enhanced in a concentration-dependent manner by DENV. To demonstrate that TF on the virus is contributing to this effect, a TF/FVIIa-specific inhibitor, NAPc2, was used. NAPc2 binds FX at an exosite as a scaffold to bind TF/FVIIa, thereby preventing FX activation but having a minimal effect on FXa amidolytic activity [373]. NAPc2 completely inhibited FXa generation by DENV. In addition, the inhibitory TF antibody, TF8-5G9 [322], resulted in a >90% decrease in TF activity.  4.1.2 TF-mediated plasma clotting To further support the presence and function of TF on purified DENV, a plasma clotting assay was carried out. Here, the virus concentration-dependent increase in FX activation observed in the chromogenic assay resulted in decreased clotting times in NP (Figure 43). Either NAPc2 (10 nM) or TF8-5G9 (100 µg/mL) significantly reduced plasma clotting.  It should be noted that  129        Figure 42. TF on DENV enhances FVIIa-mediated FX activation. (A) FXa generation by FVIIa was followed in the presence of TF-bearing DENV2. (B) 2 x 106 PFU/mL DENV was pre-treated with or without the TF/FVIIa inhibitors, NAPc2 and inhibitory antibody 5G9 (anti-TF). FVIIa (1 nM) was added with CaCl2 (5 mM) and the reaction was initiated with the addition of FX (100 nM) at 37 °C. N = 3 independent replicates. Error bars: SEM.   130         Figure 43. DENV-initiated coagulation is TF-dependent. (A) Clotting was initiated by DENV in NP. (B) The TF/FVIIa inhibitor NAPc2 (50 nM) or TF8-5G9 anti-TF (100 µg/mL) was pre-incubated with DENV2 prior to the addition of NP. 5 mM CaCl2 was added to initiate clotting. N = 4 independent replicates. Error bars: SEM.   131  TF8-5G9 was selected due to its ability to immediately inhibit TF/FVIIa complexes without the need of pre-incubation in plasma [322]. The presence of TF in purified DENV preparations was attempted to be confirmed through Western blot. However, possibly due to the low titres of DENV or conversely, low TF abundance, a conclusive TF band could not be demonstrated. Still, more sensitive assays unequivocally supported the presence of TF function in the purified DENV preparations. However, this does not confirm that TF is found directly on the DENV surface.  4.1.3 Immunogold EM As in section 3.2, I used immunogold EM to determine if a virus particle contains TF. aPL was initially investigated; however, the size of virus particles (~50 nm) was anticipated to be prohibitive towards effective triple labeling strategies due to steric limitations caused by primary and secondary probes. Furthermore, the literature supports phosphatidylserine presence on dengue virus [374] and HIV [375]. HCV contains less than 1% PS [376] but may interact differently with coagulation factors or other aPL-binding proteins [377]. Thus, TF presence on these viruses was the primary investigation as they constitute novel knowledge.  Immunogold EM was performed to determine the presence of TF on purified DENV (Figure 44). TF was indeed identified. The isotype controls displayed inconsequential non-specific binding of the antibodies or probes. aPL was also confirmed to be present on the virus envelope. DENV possessed accessible aPL as the absence of calcium failed to show any calcium-independent binding of annexin V to DENV. TF and accessible aPL on the envelope are important for the function of TF-initiated FX activation. Even though triple-labeling precluded positive results (not shown), TF and aPL were most likely present on the same virus particle as both are required for the FVIIa-dependent activation of FX. 132         Figure 44. TF and aPL are found on DENV. Differential immunogold EM showing DENV labeled with biotinylated-annexin V (aPL marker) and anti-TF. Specific virus-encoded proteins were detected with anti-DENV E-protein. To differentiate molecules, 6, 10 or 15 nm gold-conjugated 2º antibodies were used to label for E protein (red arrowheads), aPL (black arrow) and TF (white arrow heads), respectively. Grids were negatively stained with 2% phosphotungstic acid. Scale bar = 50 nm.   133  4.2 Prevalence of tissue factor on patient-derived viruses To confirm the findings of TF on cell culture-derived viruses is applicable to a clinical scenario, viremic plasma specimen were evaluated for TF using immunogold EM. For this study HIV and HCV were selected as they are more easily obtainable in comparison to dengue due to their ability to establish chronic infections and their incidence in our local community. These two viruses are also very important blood-borne pathogens that affect our healthcare, blood supply and blood products. In addition, their links to thrombotic diseases hint at a possible dysregulation of TF activity [301,310,311].  The initial technique employed was the centrifugation method (section 2.2.4.1). The advantage of this technique was the formation of a pellet that could be resuspended in any volume to load onto an EM grid. This technique showed promise in cell culture-derived HSV1 with controls showing little to no background (Appendix A.4). However, in cell culture-derived DENV and in patient-derived HCV, there was overwhelming background (not shown). This was most likely due to excessive unbound gold-labeled antibodies that prohibited distinct immune-pelleting. This did not occur for HSV1, because the virus is much larger, facilitating differential centrifugation due to more copies of antigen per virus. Rather, the immunocapture technique was used as it overcame the issue with excessive non-specific binding of antibodies to the grid.  4.2.1 HIV and TF HIV from viremic patient plasma was immunocaptured onto a virus antibody-coated EM grid and probed for TF and the virus-specific gp120. Anti-HIV gp120 antibodies were adsorbed onto the EM grid as capture antibodies for HIV. Gold particles directly conjugated to antibodies recognizing TF and gp120 were used as the detecting antibodies. gp120 was chosen due to its 134  relative exposure on the viral envelope compared to the gp41 transmembrane protein that anchors gp120. HIV was detected by EM and was demonstrated to possess TF on its surface (Figure 45A). The negative controls did not demonstrate any binding of the antibodies to virus-like-particles (VLPs) (Figure 45B). Of note, patient plasma contained a wide range of VLPs of sizes 30 nm to 200 nm, some of which labeled positive for TF and/or gp120. Thus, the size and the number of TF and/or gp120-containing vesicles were counted in randomly selected EM fields to quantitatively describe the population of these VLPs. Purified HIV diameter varies greatly in the literature, ranging from 90 to 200 nm [378–380]. Sizes 90 to 160 nm were chosen to represent HIV particles [378] and the upper limit was lowered to narrow the wide size range and segregate VLPs into different populations to facilitate quantification. Examples of the three populations: 30 nm to 90 nm, 90 nm to 160 nm, and larger than 160 nm are depicted in Figure 45. Approximately 40%, 50% and 10% of VLPs were not labeled with either gp120 or TF (30 nm to 90 nm, 90 nm to 160 nm, and larger than 160 nm, respectively) (Figure 46). Approximately half of the VLPs expressed only gp120, with <15% expressing only TF. Of all HIV1 particles (gp120-labeled and within 90 to 160 nm), 10% were positive for both TF and gp120. ~20% of gp120-labeled VLPs expressed TF. Interestingly, this percentage remained approximately constant in all populations of VLPs.  4.2.2 HCV and TF HCV patient plasma was probed for TF and the virus marker E2 protein. HCV encodes E1 and E2 envelope proteins that form a heterodimer on its surface [381]. E2 is twice the molecular weight of E1 (70 vs 35 kDa) [382,383] and therefore likely occupies a larger space on the virus envelope. Thus, E2 was chosen as a virus marker in patient plasma. 135    Figure 45. HIV harbors TF on its viral envelope. Immunogold EM of HIV in patient plasma. HIV was captured onto EM grids with anti-gp120 antibodies and subsequently probed for TF (arrows; 6 nm gold bead) and gp120 (arrowhead; 15 nm gold bead). Isotype antibodies were used as a negative control. Scale bar: 100 nm.   136         Figure 46. Patient-derived HIV displays a heterogenous phenotype. Different sizes of VLPs were characterized as being labeled with or without HIV gp120 and/or TF. The three sizes were: 30 to 90 nm (empty bars); 90 to 160 nm (black bars); and larger than 160 nm (gray bars). N = 3 independent replicates. Error bars: SEM.   137  HCV particles were separated into two populations, 30 to 80 nm and 80 to 200 nm. The former was defined in previous literature as purified HCV [384] and the latter encompassed a larger range as described in patient-derived VLPs [385]. Both populations of HCV particles were shown to harbor TF on their envelope surface (Figure 47A and B). Isotype controls displayed inconsequential non-specific binding (Figure 47C). Interestingly, there was a population of particles, presumably low-density lipoprotein (LDL)-associated HCV [385], that were resistant to antibody labeling (Figure 47D). The morphology of these LDL-associated HCV are consistent with those seen in other patient-derived HCV [385].  Approximately 30% and 40% of VLPs were labeled with E2 (30 to 80 nm and 80 to 200 nm, respectively, Figure 48). Of those E2-expressing populations, ~10% and ~20% also express TF, respectively. Unlike HIV, the increase in HCV VLP size resulted in an increased proportion of TF expressing VLPs.  4.3 Discussion As with my work on herpesviruses, TF presence on cell culture-derived DENV confirmed the ability of enveloped viruses to obtain transmembrane coagulation proteins. Even though cellular TF function is restricted by encryption and accessibility to plasma proteins, here TF on the DENV  envelope  was  functionally active  in  both  FVIIa-mediated  FX  activation and in  extrinsic pathway initiation of coagulation. These results supported the endeavor of searching for TF on patient-derived viruses. The findings that HIV as well as HCV incorporate TF onto their envelope are exciting and could explain their contribution to hypercoagulability [300,310,311]. Approximately 20% of discernible HIV and HCV particles expressed TF. Since not all particles  expressed TF,  this  may 138      Figure 47. Patient-derived HCV expresses TF on its surface. Immunogold EM of HCV in patient plasma. HCV was captured onto EM grids using anti-E2 antibodies and subsequently probed for TF (arrows; 6 nm gold bead) and E2 (arrowhead; 15 nm gold bead). Isotype antibodies were used as a negative control. Low-density lipoprotein (LDL)-associated HCV is also depicted. Scale bar: 100 nm.   139         Figure 48. Phenotypes of HCV particle population. Different sizes of VLPs were characterized as being labeled with or without HCV E2 and/or TF. The two sizes were: 30 to 80 nm (empty bars); and 80 to 200 nm (gray bars). N = 3 independent replicates. Error bars: SEM.   140  suggest that the viruses are antigenically heterogenous in patients and perhaps derived from several sources of infected cells, some of which express TF and others may not. Another explanation could be TFPI inhibition of TF/FVIIa on the virus, thereby masking recognition of TF by the antibody. TF function was not assessed, however, one study demonstrated that untreated HIV-positive individuals have higher TF-expressing microparticle procoagulant activity than treated HIV-positive individuals [312]. They speculated that these TF-expressing microparticles originate from monocytes. Monocytes are known to be infectible by HIV [198]. As microparticles range in sizes of 100 to 1000 nm in diameter [386] and HIV particles range in sizes of 90-200 nm in diameter [378–380], it may be possible that HIV co-exists with, masquerades as or is mistaken for a microparticle. As DENV was demonstrated to possess TF in cell culture, there were attempts at probing viremic DENV individuals for TF expression. The prior immune-depletion strategies were not successful. However, there were small non-labeled VLPs of ~50 nm, consistent with the size of DENV in cell culture. Other modifications to the protocol included the use of heparan sulfate [387] or concanavalin A [388] layer onto the EM grid to capture DENV. Concanavalin A resulted in enhanced adsorption of cell culture-derived DENV to the EM grid but in plasma samples, resulted in an excess of plasma protein binding, a known phenomenon [389]. This resulted in a dark film of protein that obstructed the view by EM. As some of the patients had detectable IgG and/or IgM to DENV, dissociation of the antibodies using low pH glycine was attempted [390]. The difficulty in detecting even DENV-encoded antigen may be attributed to the storage and processing of the patient plasma used in this study. The only recorded successful attempt at identifying the virus by EM was performed with freshly prepared plasma [327]. The same group discovered that plasma-isolated infectious DENV vesicles were dominantly derived from CD61+ expressing 141  megakaryocytes lineage cells. Megakaryocytes are known to express TF [184], so it is reasonable to expect the presence of TF to be on patient-derived DENV particles. One of the limitations of immunogold EM is that antigen detection may not be saturated, due to affinity, steric hindrance or balancing the concentration to avoid non-specific binding. Thus, the true proportion of TF-expressing particles may be under-reported. Also, only viruses that have adsorbed to the grid are detected. In the case of HCV, lipid-associated HCV particles may not have been captured by the antibodies as the E2 protein may have been masked. Although EM remains the gold standard for sizing and identification of viruses, newer technologies for the characterization of virus particles in the nanometer range have been improving. High throughput and rapid antigen identification could be achieved through flow virometry [391], an adaptive form of using flow cytometry to characterize small virus particles. 142  Chapter 5: Summary 5.1 HSV1 gC has evolved to affect FVIIa function The ability of sgC-His to independently enhance FVIIa activity through viral tenase assembly with FX (sgC-His/FVIIa/FX) on a purified, virus-free membrane differs from that on the virus membrane (as modeled in Figure 49). A lack of synergism in the purified system would suggest that in the particular system, sgC-His competes with sTF-His binding to FVIIa. Both cofactors appeared to stabilize FVIIa and enable efficient FX activation as their catalytic efficiencies were similar. Viral membrane gC enhancement of TF function on the virus towards FX activation may be aided by an unknown factor (Figure 49E). Nonetheless, the work presented here adds further evidence that HSV1 has evolved to affect the haemostatic system. Subsequently, activation of clotting proteases provides a link to cardiovascular diseases such as thrombosis and atherosclerosis. The dependence of gC on FX to mediate FVIIa cofactor function may guide therapeutic development. The evolutionary advantage of an evolved viral coagulation factor could be enhanced infectivity via the activation of clotting protease-mediated cell signaling. gC and TF was previously shown by the Pryzdial laboratory in vitro to achieve enhanced infectivity via PAR-2 activation [12]. Subsequent in vivo work further supported the TF-mediated infection, demonstrating that envelope TF was critical for infection of BALB/c mice and also enhanced infection in C57BL/6J mice [13]. It is, however, unknown how gC affects TF-mediated infectivity in vivo.     143      Figure 49. Working model of gC interaction with FVIIa/FX. (A) gC speculated binding to FX as the 33-123 amino acid residue region near the C-terminal end has been implicated. (B) sgC-His binding to FVIIa is enhanced upon membrane binding. (C) sgC-His binding to FVIIa is increased by FX binding. This viral tenase can work independently from (D) the extrinsic tenase, depicted as a speculated interaction with FX. (E) On the virus surface, gC enhancement of TF function is aided by an unknown factor (?). aPL is not required for the interactions between gC and FVIIa or FX but contributes to the proteolytic activity of the tenase.   144  5.1.1 gC mimetics No other herpesvirus has been investigated for FX or FVII-modulating function. HSV1 gC is homologous to HSV2 gF (gC-2) [392] and shares the ability to bind to heparan sulfate [320] and inhibit complement [393], albeit in a different manner. Marek’s disease virus (MDV), another member of Herpesviridae family, causes a severe disease in chickens and has a large impact on the poultry industry. MDV gC shares some homology with HSV1 gC such as the conserved cysteine residues that confer the complement-inhibiting ability to HSV1 gC [394]. It would be interesting to determine if these or other virus proteins interact with FX and/or FVIIa in a similar manner to gC. One non-herpesvirus has been shown to interact with coagulation factors. Human adenovirus is capable of binding several vitamin K-dependent coagulation factors, including FX, FVII and FIX with high affinity (via their Gla domains) to enhance liver transduction [395,396]. The Thr423-Glu424-Thr425 of human adenovirus serotype 5 (HAdv5) hexon protein was demonstrated to be crucial for FX binding [397]. Preliminary sequence alignment of the HAdv5 hexon protein with HSV1 gC was performed using T-Coffee, an internet protein comparison algorithm [398]. Sequence alignment showed 21% identical sequence with areas of similar properties within a 76 amino acid immediate region containing the Thr423-Glu424-Thr425 amino acid motif (Figure 50). The gC segment aligning with this region is near the IV C3b binding region (Figure 10). The FX and/or FVIIa interacting regions of gC have not been elucidated, however the 33-123 region is known to be important in FX binding [241].     145        Figure 50. Sequence homology between HSV1 gC and HAdv5 hexon FX-binding region Alignment of the HSV1 gC and HADv5 hexon sequences was performed with T-Coffee (Proteomics identifications database code P10228 and P04133, respectively). The blue box highlights the Thr423-Glu424-Thr425 of human adenovirus serotype 5 (HAdv5) hexon protein was demonstrated to be crucial for FX binding. Amino acid residues that are identical (*), conserved with strongly similar properties (:) or weakly similar (.) are note in the consensus row (cons). The color in the consensus is the Transitive Consistency Score (TCS) that describes the reliability of the alignment and if they are structurally correct: red is high, yellow is moderate, green is low.   146  5.1.2 gC sequence similarity to FVIIa binding proteins The ability of gC to bind to FVIIa is particularly interesting as it enhanced its proteolytic activity. Sequence alignment using T-Coffee of the extracellular domains of gC and TF (Figure 51) demonstrated 9/27 sequence identity of residues known to be implicated in FVIIa interaction. The majority of these homologous residues are found in the C-terminal ectodomain (Figure 52). The 33-123 region of gC contains one identical and one conserved match to TF interacting residues with FVIIa. Insights to the interaction between gC and FVIIa can also be obtained by comparing gC to EPCR as EPCR can bind to FVIIa and affect its function [97,399]. Comparison of the extracellular domains of gC and EPCR showed three identical amino acid residues and one conserved amino acid residues (Figure 53) that interact with the Gla domain or co-ordinate with calcium bound to the Gla domain of Protein C (Figure 54) [400]. Activated Protein C binding to and FVIIa binding to soluble EPCR are similar [399]. In the case of FX, the Gla domain of FX lack similarity to the EPCR binding moieties of Protein C [399]. Thus, gC binding to FVIIa may include interaction(s) with the Gla domain but may not interact with the Gla domain of FX. This is supported by the observation that the loss of the 33-123 amino acid residues of gC inhibited FX interaction. This region does not overlap with the Gla domain interacting region of EPCR (Figure 53). Crystal structures of gC have remained elusive but the structure could potentially be solved using cryo-EM. The inclusion of a phospholipid membrane, TF, FVIIa and/or FX may stabilize a complex for which a structure can be solved. This would give more insight to the interacting regions and may benefit procoagulant therapeutics or vaccine development.     147   Figure 51. gC sequence comparison to TF. Sequence alignment of the extracellular domain of gC and the extracellular domain of TF was performed with T-Coffee (Proteomics identifications database code P10228 and P13726, respectively). The 33-123 region of gC known to interact with FX is overlined in black. TF amino acid residues that have been implicated in FVIIa interaction are marked (arrow heads). Amino acid residues that are identical (*), conserved with strongly similar properties (:) or weakly similar (.) are note in the consensus row (cons). The TCS legend is color coded from low to high confidence (bottom right). 148       Figure 52. Model of gC homology with the conserved sequence of TF interaction with FVIIa. The crystal structure of the N-terminus and C-terminus ectodomains of soluble TF (light green and dark green, respectively) was obtained from PDB code 1DAN. Residues of identical matches, conserved matches, weak similarity and no homology to gC from sequence comparison are depicted as balls (red, yellow, teal and gray, respectively). A phospholipid bilayer is depicted for orientation.   149      Figure 53. gC sequence comparison to EPCR. Sequence alignment of the extracellular domain of gC and the extracellular domain of EPCR was performed with T-Coffee (Proteomics identifications database code P10228 and Q9UNN8, respectively) [398]. The 33-123 region of gC known to interact with FX is overlined in black. EPCR interacting amino acid residues with the Gla domain of Protein C are marked (arrow heads). Amino acid residues that are identical (*), conserved with strongly similar properties (:) or weakly similar (.) are note in the consensus row (cons). The TCS legend is color coded from low to high confidence (bottom right).  150    Figure 54. Model of gC homology with the structure of EPCR interaction with Protein C. The crystal structure of the interaction between the Gla domain of Protein C (blue) and the extracellular domain of EPCR (green) was obtained from PDB code 1LQV. Residues of identical matches, conserved matches and no homology to gC from sequence comparison are depicted as balls (red, yellow and gray, respectively). Calcium is shown as small gray spheres.    151  5.2 Host TF-deriving enveloped viruses The presence of TF on DENV, HIV and HCV in addition to the previously established herpesviruses [2] supported the hypothesis that many enveloped viruses can obtain TF. This is observed in HIV and HCV patients, thus providing a clinical link to hypercoagulability. As a common antigen is found on these viruses, it raises the question on why and how these viruses have evolved to obtain TF.   5.2.1 HIV and HCV TF procoagulant activity HIV and HCV obtain TF antigen from the host, but whether these TF-expressing virus particles also express TF procoagulant function is not known. One could speculate that it is probable that HIV expresses TF procoagulant function based off of a previous study with TF-expressing microparticles in HIV patients. However, to demonstrate that the TF-expressing HIV particles are functional, differentiation from microparticles is necessary. This could be achieved through density centrifugation [401,402]. Subsequent testing of TF procoagulant function can be assessed with FX chromogenic assays and/or plasma clotting assays. The question then remains, why do these viruses incorporate TF into its envelope?   5.2.2 Functional advantage of TF expression  It has been shown that HIV can utilize PAR-1 and PAR-2 signaling to upregulate the HIV-1 coreceptor CCR5 in oral keratinocytes [403]. It has also been demonstrated that HIV-1 increases PAR-1 expression in astrocytes and may contribute to inflammation leading to neurodegeneration in HIV encephalitis [404]. As TF is known to be expressed by astrocytes [65], one would wonder whether PAR-1 and/or PAR-2 activation is mediated directly or indirectly by TF function (via 152  FVIIa or FX activation). In vitro studies, similar to previous work on HSV1 and TF-mediated PAR activation leading to enhanced infection [12], could be performed to investigate the effect of TF-expressing HIV on cell infection via PARs. If TF is functional and contributes to virus infection, could this common antigen be used as a pan-specific antiviral target? In vivo studies have demonstrated that anticoagulants targeted toward TF and FXa could reduce HSV1 infection [13]. In particular, apixaban and NAPc2 diminished the presence of infectious virus in all tested tissues. NAPc2 has also been evidenced to decrease morbidity and increase survival times in Ebola-infected rhesus macaques [405]. As a consequence of treating the disease progression with NAPc2, viral load was lowered in treated rhesus macaques. Anti-TF antibodies that inhibited TF/FVIIa activation of proteases resulted in diminished HSV1 infection [13]. One method of adapting this knowledge could be the creation of bivalent antibodies [406] to TF and a virus antigen to selectively inhibit TF function on the virus but not physiological TF function. Bivalent antibodies may also serve to better neutralize these pathogenic viruses [406]. 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Thromb Res United States; 1988; 50: 745–6.   205  206  Appendices Appendix A   A.1 Derivation of quadratic binding isotherm The quadratic binding isotherm is used to determine the apparent dissociation constant for the FVIIa and cofactor (lipid and/or protein). Please refer to Figure 20. Schematic of non-enzymatic and enzymatic reactions involved in FVIIa-mediated FX activation. The following equations are derived from the schematic:  [𝐸]0 = [𝐸] + [𝐸𝐶] + [𝐸𝑆] + [𝐸𝐶𝑆]  (Equation 1) [𝐶]0 = [𝐶] + [𝐸𝐶] + [𝐸𝐶𝑆]   (Equation 2) [𝐸] ∙ [𝐶] = 𝐾𝑑 ∙ [𝐸𝐶]    (Equation 3) [𝐸𝑆] ∙ [𝐶] = 𝐾𝑑′ ∙ [𝐸𝑆𝐶]   (Equation 4) [𝐸𝐶] ∙ [𝑆] = 𝐾𝑠 ∙ [𝐸𝐶𝑆]   (Equation 5) [𝐸] ∙ [𝑆] = 𝐾𝑠′ ∙ [𝐸𝑆]    (Equation 6) [𝐶𝐿] ∙ [𝐶𝑃] = 𝐾𝑐 ∙ [𝐶]    (Equation 7) Using equations 1-4, E0 can be expressed as: (brackets denoting concentration are removed for improved clarity) 𝐸0 =𝐾𝑑∙𝐸𝐶𝐶0−𝐸𝐶−𝐸𝐶𝑆+ 𝐸𝐶 +𝐾𝑑′ ∙𝐸𝐶𝑆𝐶0−𝐸𝐶−𝐸𝐶𝑆+ 𝐸𝐶𝑆  Substituting equation 5 for EC obtains: 𝐸0 =𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆𝑆+𝐾𝑑′ ∙𝐸𝐶𝑆𝐶0−𝐾𝑠∙𝐸𝐶𝑆𝑆−𝐸𝐶𝑆+𝐾𝑠∙𝐸𝐶𝑆𝑆+ 𝐸𝐶𝑆  𝐸0 =𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆𝑆+𝐾𝑑′ ∙𝐸𝐶𝑆𝐶0−𝐾𝑠∙𝐸𝐶𝑆𝑆−𝐸𝐶𝑆+𝐸𝐶𝑆(𝐾𝑠+𝑆)𝑆  207  Multiplying both sides by 𝐶0 −𝐾𝑠∙𝐸𝐶𝑆𝑆− 𝐸𝐶𝑆 and 𝑆 results in: 𝐸0 ∙ 𝐶0 ∙ 𝑆 − 𝐸0 ∙ 𝐾𝑠 ∙ 𝐸𝐶𝑆 − 𝐸0 ∙ 𝐸𝐶𝑆 ∙ 𝑆 = 𝑆 ∙ (𝐾𝑑 ∙𝐾𝑠∙𝐸𝐶𝑆𝑆+ 𝐾𝑑′ ∙ 𝐸𝐶𝑆) + 𝐸𝐶𝑆(𝐾𝑠 + 𝑆)(𝐶0 −𝐾𝑠∙𝐸𝐶𝑆𝑆− 𝐸𝐶𝑆)  Dividing both sides by (𝐾𝑠 + 𝑆) with rearranging: 𝐸0∙𝐶0∙𝑆−𝐸0∙𝐾𝑠∙𝐸𝐶𝑆−𝐸0∙𝐸𝐶𝑆∙𝑆𝐾𝑠+𝑆=𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆+𝑆∙𝐾𝑑′ ∙𝐸𝐶𝑆𝐾𝑠+𝑆+ 𝐸𝐶𝑆 ∙ 𝐶0 −𝐾𝑠∙𝐸𝐶𝑆2𝑆− 𝐸𝐶𝑆2  Rearranging to the left side results in: 𝐸𝐶𝑆2 +𝐾𝑠∙𝐸𝐶𝑆2𝑆− 𝐸𝐶𝑆 ∙ 𝐶0 +𝐸0∙𝐶0∙𝑆−𝐸0∙𝐾𝑠∙𝐸𝐶𝑆−𝐸0∙𝐸𝐶𝑆∙𝑆−𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆−𝑆∙𝐾𝑑′ ∙𝐸𝐶𝑆𝐾𝑠+𝑆= 0  𝐸𝐶𝑆2 (𝐾𝑠+𝑆𝑆) −  𝐸𝐶𝑆 ∙ 𝐶0 +𝐸0∙𝐶0∙𝑆−𝐸0∙𝐾𝑠∙𝐸𝐶𝑆−𝐸0∙𝐸𝐶𝑆∙𝑆−𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆−𝑆∙𝐾𝑑′ ∙𝐸𝐶𝑆𝐾𝑠+𝑆= 0  Substitution of −𝐸0 ∙ 𝐾𝑠 ∙ 𝐸𝐶𝑆 − 𝐸0 ∙ 𝐸𝐶𝑆 ∙ 𝑆 for −(𝐾𝑠 + 𝑆)(𝐸0𝐸𝐶𝑆) yields: 𝐸𝐶𝑆2 (𝐾𝑠+𝑆𝑆) −  𝐸𝐶𝑆 ∙ 𝐶0 +𝐸0∙𝐶0∙𝑆−(𝐾𝑠+𝑆)(𝐸0𝐸𝐶𝑆)−𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆−𝑆∙𝐾𝑑′ ∙𝐸𝐶𝑆𝐾𝑠+𝑆= 0  Some rearrangements: 𝐸𝐶𝑆2 (𝐾𝑠+𝑆𝑆) −  𝐸𝐶𝑆 ∙ 𝐶0 − 𝐸0𝐸𝐶𝑆 +−𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆−𝑆∙𝐾𝑑′ ∙𝐸𝐶𝑆𝐾𝑠+𝑆+𝐸0∙𝐶0∙𝑆𝐾𝑠+𝑆= 0  𝐸𝐶𝑆2 (𝐾𝑠+𝑆𝑆) −  𝐸𝐶𝑆 ∙ (𝐶0 + 𝐸0) −𝐾𝑑∙𝐾𝑠∙𝐸𝐶𝑆+𝐾𝑑′ ∙𝑆∙𝐸𝐶𝑆𝐾𝑠+𝑆+𝐸0∙𝐶0∙𝑆𝐾𝑠+𝑆= 0  𝐸𝐶𝑆2 (𝐾𝑠+𝑆𝑆) −  𝐸𝐶𝑆 (𝐾𝑑∙𝐾𝑠+𝐾𝑑′ ∙𝑆𝐾𝑠+𝑆+ 𝐸0 + 𝐶0) +𝐸0∙𝐶0∙𝑆𝐾𝑠+𝑆= 0    (Equation 8) For the given activity of the reaction, 𝑣 = 𝑘𝑐𝑎𝑡 ∙ 𝐸𝐶𝑆 + 𝑘𝑐𝑎𝑡′ ∙ 𝐸𝑆. As 𝑘𝑐𝑎𝑡′ was experimentally determined to be must less than 𝑘𝑐𝑎𝑡, the equation becomes: 𝑣 ≈ 𝑘𝑐𝑎𝑡 ∙ 𝐸𝐶𝑆. Substituting 𝑘𝑐𝑎𝑡 =𝑉𝑚𝑎𝑥𝐸0, the equation becomes 𝑣 = 𝑉𝑚𝑎𝑥 ∙𝐸𝐶𝑆𝐸0. By using the quadratic equation to solve for ECS from equation 8, one can solve for 𝑣: 208  𝐸𝐶𝑆 =−𝑏±√𝑏2−4𝑎𝑐2𝑎 where 𝑎 =(𝐾𝑠+𝑆)𝑆 , 𝑏 =𝐾𝑑∙𝐾𝑠+𝐾𝑑′ ∙𝑆𝐾𝑠+𝑆+ 𝐸0 + 𝐶0, and 𝑐 =𝐸0∙𝐶0∙𝑆𝐾𝑠+𝑆, 𝑣 =𝑉𝑚𝑎𝑥∙𝑆2∙𝐸0(𝐾𝑠+𝑆)(𝐸0 + 𝐶0 +𝐾𝑑∙𝐾𝑠+𝐾𝑑′ ∙𝑆𝐾𝑠+𝑆± √(𝐸0 + 𝐶0 +𝐾𝑑∙𝐾𝑠+𝐾𝑑′ ∙𝑆𝐾𝑠+𝑆)2− 4 ∙ 𝐸0 ∙ 𝐶0)   (Equation 9) Based on previous works [94,407], the quadratic isotherm is similar to equation 9 with 𝐾𝑑,𝑎𝑝𝑝 =𝐾𝑑∙𝐾𝑠+𝐾𝑑′ ∙𝑆𝐾𝑠+𝑆   𝐾𝑑 ∙ 𝐾𝑠 = 𝐾𝑑′ ∙ 𝐾𝑠′  is predicted based on symmetry (Figure 20). This allows the following rearrangement after substitution of 𝐾𝑑′ : 𝐾𝑑,𝑎𝑝𝑝 = 𝐾𝑑𝐾𝑠+𝐾𝑠∙𝑆/𝐾𝑠′𝐾𝑠+𝑆    𝐾𝑑 can thus be solved if 𝐾𝑠 and 𝐾𝑠′ are known. Furthermore, if 𝐾𝑠 = 𝐾𝑠′, then 𝐾𝑑,𝑎𝑝𝑝 = 𝐾𝑑. As described by Equation 7, Ni-chelating lipids affect the available membrane-bound cofactor. The complexity of the added effect of phospholipid was ablated with the assumption that the dissociation value of cofactor to FVIIa is much larger than that of cofactor/lipid to FVIIa. This allows a minor change to the quadratic binding isotherm to: 𝑣 =𝑉𝑚𝑎𝑥 ∙ 𝑆2 ∙ 𝐸0(𝐾𝑠 + 𝑆)(𝐸0 + 𝐶0 − 𝐶𝐿 +𝐾𝑑 ∙ 𝐾𝑠 + 𝐾𝑑′ ∙ 𝑆 + 𝐶𝐿𝐾𝑠 + 𝑆± √(𝐸0 + 𝐶0 − 𝐶𝐿 +𝐾𝑑 ∙ 𝐾𝑠 + 𝐾𝑑′ ∙ 𝑆 + 𝐶𝐿𝐾𝑠 + 𝑆)2− 4 ∙ 𝐸0 ∙ 𝐶0) Where 𝐾𝑑,𝑎𝑝𝑝 =𝐾𝑑∙𝐾𝑠+𝐾𝑑′ ∙𝑆+𝐶𝐿𝐾𝑠+𝑆.   209  A.2 FVIIa-R retains most of its FVIIa function   Figure 55. FVIIa-R retains its function when bound to sTF-His or sgC-His. FVIIa-R FX-activating ability was assessed by FX chromogenic assay. 5 pM of FVIIa (white bar) or FVIIa-R (gray bar) was incubated with 50 µM NiPCPS, 5 mM Ca2+ and 400 pM sTF-His or 400 nM sgC-His. FX was added to initiate the reaction and the mixture was incubated for 20 min at 37 °C prior to stopping the reaction with EDTA and measuring FX activation with chromogenic substrate.   210  A.3 Comparison of fits for MST-derived FVIIa-R interactions  Table A5. Comparison of fits for MST-derived FVIIa-R interactions   Simple binding fit     No FX With FX Ligand Lipid Kd (µM)  (95% CI) r2 Kd (µM)  (95% CI) r2 sTF-His NiPCPS 8.2 x 10-4 (8.6 x 10-5 – 1.5 x 10-3) 0.93 3.0 x 10-5 (0.8 x 10-5 – 5.2 x 10-5) 0.9468 sgC-His NiPC 2.0  (1.3 – 2.8) 0.896 0.9  (0.7 – 1.0) 0.98 sgC-His NiPCPS 1.3  (0.8 – 1.7) 0.935 0.6  (0.2 – 0.9) 0.963 sgC-His PCPS No fit - 2.2  (0.9 – 3.5) 0.822 sgC-His None No fit - 2.9 (1.5 – 4.2) 0.994     Hill equation fit   No FX With FX Ligand Lipid EC50 (µM)  (95% CI) r2 EC50 (µM)  (95% CI) r2 sTF-His NiPCPS 1.0 x 10-3 (4.3 x 10-4 – 2.4 x 10-3) 0.927 3.2 x 10-5 (1.8 x 10-5 – 5.6 x 10-5) 0.957 sgC-His NiPC 1.6 (0.7 – 3.7) 0.925 1.0 (0.8 – 1.1) 0.992 sgC-His NiPCPS 0.8 (0.5 – 1.3) 0.943 0.4 (0.3 – 0.6) 0.974 sgC-His PCPS >13.0 ( - ) - 2.7 (1.3 – 5.3) 0.812 sgC-His None >12.0 ( - ) - 2.9  (1.6 – 4.2) 0.994    211  A.4 Immunogold centrifugation of HSV1  Figure 56. Immunogold differential centrifugation of HSV1. HSV1/TF+/gC+ (108 vp/mL) was incubated with 15 nm gold-conjugated anti-HSV1 gC polyclonal rabbit antibodies (1:100) or 15 nm gold-conjugated rabbit IgG isotype control for 2 h at 37 °C. The virus-gold mixture was pelleted at 10,000 x g for 25 min, washed with HBS + 0.1% BSA-c and pelleted again. The washed pellet was resuspended in 20 µL HBS and loaded onto an EM grid. Grids were negatively stained with 2% PTA before visualization by transmission EM. 

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