Medicine, Faculty of
Pathology and Laboratory Medicine, Department of
DSpace
UBCV
Lee, Frank Man Hon
Doctor of Philosophy - PhD
University of British Columbia
Thrombosis is the leading cause of death and disability. Clots (i.e. thrombi) that obstruct blood flow from delivering oxygen and nutrients to tissues require rapid removal. The main treatment is with tissue plasminogen activator (tPA), which activates the enzyme plasmin from its circulating precursor plasminogen. Plasmin works by degrading fibrin, a meshwork of fibers that form the structural scaffold of the clot. Although these thrombolytic drugs have saved countless lives, their use is associated with life-threatening complications including increased bleeding risk and cellular toxicity. By greatly elevating plasmin concentrations with intravenous tPA, the effect of plasmin on its other substrates apart from fibrin may play an important role in dysregulating hemostasis, giving insight into physiological mechanisms that are impacted by thrombolysis. This thesis focuses on the modulatory effects of plasmin on protein C (PC) and activated clotting factor (F) V (FVa). Fragmentation of PC by plasmin assessed in conjunction with activated PC (APC)-specific chromogenic substrate cleavage showed increased amidolytic activity correlating with proteolysis of the PC heavy chain (PC-Hc). Plasmin-cleaved PC inhibited inactivation of FVa and had no appreciable effect on prolonging plasma clotting times in PC-deficient plasma compared with APC. N-terminal sequencing showed that plasmin cleaves PC-Hc within the autolysis loop and at the canonical thrombin activation site. Our findings so far are consistent with natural and engineered APC variants, affected within or adjacent to the autolysis loop, that have reduced anticoagulant function. Interestingly, these mutants still signal through protease-activated receptor 1, suggesting a novel role for plasmin-modulated (A)PC in conferring cytoprotection. For FVa, plasmin-specific cleavage of chromogenic substrate and protein staining were used to confirm the requirement for proteolysis by plasmin in converting FVa into an accelerator of tPA. Free and lipid-binding fragments both enhanced tPA activity. 125I-radiolabeled plasminogen binding indicated an interaction with an unbound fragment derived from FVa-Hc that is C-terminal lysine dependent. The tPA-accelerating fragment of FVa-Lc is derived from the C1 and C2 lipid-binding domains and could non-enzymatically enhance fibrinolysis specifically at the site of vascular injury. Understanding the functional regulation of PC and FVa by plasmin offers novel insights toward improving thrombolysis therapy.
FUNCTIONAL REGULATION OF ANTICOAGULANT PROTEIN C AND CLOTTING FACTOR Va BY FIBRINOLYTIC PLASMIN by Frank Man Hon Lee B.H.Sc., McMaster University, 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) July 2020 © Frank Man Hon Lee, 2020 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Functional regulation of anticoagulant protein C and clotting factor Va by fibrinolytic plasmin submitted by Frank Man Hon Lee in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Pathology and Laboratory Medicine Examining Committee: Dr. Edward Pryzdial, Pathology and Laboratory Medicine Supervisor Dr. Edward Conway, Medicine Supervisory Committee Member Dr. David Waisman, Dalhousie University, Pathology External Examiner Dr. Ross MacGillivray, Biochemistry and Molecular Biology University Examiner Dr. Dieter Bromme, Oral Biological and Medical Sciences University Examiner Additional Supervisory Committee Members: Dr. Shannon Jackson, Providence Hematology Supervisory Committee Member Dr. Haydn Pritchard, Pathology and Laboratory Medicine Supervisory Committee Member iii Abstract Thrombosis is the leading cause of death and disability. Clots (i.e. thrombi) that obstruct blood flow from delivering oxygen and nutrients to tissues require rapid removal. The main treatment is with tissue plasminogen activator (tPA), which activates the enzyme plasmin from its circulating precursor plasminogen. Plasmin works by degrading fibrin, a meshwork of fibers that form the structural scaffold of the clot. Although these thrombolytic drugs have saved countless lives, their use is associated with life-threatening complications including increased bleeding risk and cellular toxicity. By greatly elevating plasmin concentrations with intravenous tPA, the effect of plasmin on its other substrates apart from fibrin may play an important role in dysregulating hemostasis, giving insight into physiological mechanisms that are impacted by thrombolysis. This thesis focuses on the modulatory effects of plasmin on protein C (PC) and activated clotting factor (F) V (FVa). Fragmentation of PC by plasmin assessed in conjunction with activated PC (APC)-specific chromogenic substrate cleavage showed increased amidolytic activity correlating with proteolysis of the PC heavy chain (PC-Hc). Plasmin-cleaved PC inhibited inactivation of FVa and had no appreciable effect on prolonging plasma clotting times in PC-deficient plasma compared with APC. N-terminal sequencing showed that plasmin cleaves PC-Hc within the autolysis loop and at the canonical thrombin activation site. Our findings so far are consistent with natural and engineered APC variants, affected within or adjacent to the autolysis loop, that have reduced anticoagulant function. Interestingly, these mutants still signal through protease-activated receptor 1, suggesting a novel role for plasmin-modulated (A)PC in conferring cytoprotection. For FVa, plasmin-specific cleavage of chromogenic substrate and protein staining were used to confirm the requirement for iv proteolysis by plasmin in converting FVa into an accelerator of tPA. Free and lipid-binding fragments both enhanced tPA activity. 125I-radiolabeled plasminogen binding indicated an interaction with an unbound fragment derived from FVa-Hc that is C-terminal lysine dependent. The tPA-accelerating fragment of FVa-Lc is derived from the C1 and C2 lipid-binding domains and could non-enzymatically enhance fibrinolysis specifically at the site of vascular injury. Understanding the functional regulation of PC and FVa by plasmin offers novel insights toward improving thrombolysis therapy. v Lay Summary Blood clots are the leading cause of death and disability worldwide. Current drugs used to get rid of these clots to restore blood flow increase the risk of life-threatening bleeding and severely damage normal cells because of drug toxicity. These clot-dissolving drugs work by producing the protein plasmin, which breaks down blood clots as its primary function. Side-effects are primarily due to introducing high levels of plasmin above what is normal in the body. Besides breaking down blood clots, plasmin has secondary functions on other blood proteins. My research focuses on understanding how plasmin regulates the functions of two such proteins that circulate in the human bloodstream, protein C and factor V. My findings implicate protein C as having a potential role as a remedy against cellular toxicity and factor V as an enhancer of clot break down. These critical functions may pave the way to developing safer clot-dissolving drugs. vi Preface My research when beginning the PhD portion of my MD/PhD degree was focused on further understanding the biochemistry of how activated clotting factor X (FXa) is converted by plasmin from a procoagulant enzyme to fibrinolytic cofactor. My work was primarily involved with understanding the functions of FXai-K, a thrombolytic variant of FXa devised and patented by Dr. Ed Pryzdial, a clinical professor in the Department of Pathology and Laboratory Medicine at UBC. Work in this area contributed to a manuscript titled: âThrombolysis by chemically modified FXaâ published in the Journal of Thrombosis and Hemostasis in 2016. This then transitioned into a collaborative research project with Dr. Chris Orvig from the Department of Chemistry at UBC and his graduate student at the time, Dr. Sarah Spreckelmeyer. The purpose of this project was to label FXa with a small-molecule probe that chelates the radioisotope gallium to serve dual purposes as both a fibrinolytic and clot-imaging agent. Due to lengthy reagent issues for both of these projects that unfortunately resulted in many failed preparations of the Xai-K and the gallium chelator respectively, I was moved consecutively from these projects after considerable effort. My work involving novel biochemical applications for FXa has not been included in this dissertation. Studying the involvement of FVa in fibrinolysis after proteolysis by plasmin was a project conceived by Dr. Ed Pryzdial that furthers his earlier work with a former graduate student Dr. Abed Zeibdawi. Dr. Zeibdawiâs research thesis was in dissecting the mechanism by which plasmin cleaves and inactivates FVa. The experiments conducted in Chapter 4 were designed and completed by myself with feedback from my supervisor. This work has been presented at various local conferences and symposia. A manuscript vii is currently in preparation, which will be supported by additional experiments as discussed in the future directions of the chapter. The premise of the work reported in Chapter 4 was rationalized when studying whether plasmin could convert APC-cleaved FVa into a fibrinolytic cofactor. During these experiments, both plasmin and APC were used, which led to the thought that there may be an interaction between these two enzymes. This led to my supervisor and me to conceive a project studying the mechanism of proteolysis of PC by plasmin. Work in this project has led to some intriguing findings and future directions. Additional studies required to show that plasmin-cleaved (A)PC maintains signaling through protease-activated receptor 1 are in progress for an imminent manuscript. This will undoubtedly raise interest in the field as current clinical trials with APC variants demonstrate similar biochemistry. In addition, intravascular plasmin has been gaining favour over tPA in stroke, which in part may be due to its role in modulating (A)PC. The experiments in Chapter 3 were conducted by myself with feedback from my supervisor. viii Table of Contents Abstract ........................................................................................................................... iii Lay Summary .................................................................................................................. v Preface ............................................................................................................................ vi Table of Contents .......................................................................................................... viii List of Tables ................................................................................................................. xvi List of Figures ............................................................................................................... xvii List of Abbreviations ....................................................................................................... xx Acknowledgements ..................................................................................................... xxvi Dedication .................................................................................................................. xxvii Chapter 1: Introduction .................................................................................................... 1 1.1 Global thrombosis burden ................................................................................. 1 1.1.1 Epidemiology ............................................................................................. 1 1.1.2 Clot-dissolving drugs ................................................................................. 2 1.1.3 Thrombolysis needs improving .................................................................. 3 1.1.4 Thesis rationale ......................................................................................... 7 1.2 Hemostasis ....................................................................................................... 8 1.2.1 Overview of hemostasis ............................................................................. 8 1.3 Blood coagulation ............................................................................................. 9 1.3.1 Vessel injury and subendothelial exposure ................................................ 9 1.3.2 Cell-based model of coagulation ................................................................ 9 1.3.3 Initiation ................................................................................................... 12 1.3.3.1 Platelet adhesion .................................................................................. 12 ix 1.3.3.2 Anionic phospholipid exposure ............................................................. 14 1.3.3.3 Synthesis and lipid-binding properties of coagulation factor proteins ... 15 1.3.3.4 Tissue factor pathway .......................................................................... 16 1.3.3.5 Contact pathway .................................................................................. 17 1.3.4 Amplification ............................................................................................ 18 1.3.4.1 Thrombin-mediated activation of FV and FVIII ..................................... 18 1.3.4.2 Amplification of thrombin generation .................................................... 18 1.3.4.3 Thrombin-mediated platelet activation and aggregation ....................... 20 1.3.5 Neutrophil extracellular traps (NETs) ....................................................... 22 1.3.6 Propagation ............................................................................................. 22 1.3.6.1 Fibrin polymerization ............................................................................ 22 1.3.6.2 FXIIIa cross-linking ............................................................................... 24 1.3.7 Wound healing and repair ........................................................................ 25 1.4 Anticoagulation ............................................................................................... 26 1.4.1 Endothelial cells ....................................................................................... 26 1.4.2 Serine protease inhibition ........................................................................ 26 1.4.3 Tissue factor pathway inhibitor ................................................................ 27 1.4.4 Protein C pathway ................................................................................... 28 1.4.5 Plasmin inactivation ................................................................................. 29 1.4.6 Therapeutic anticoagulation ..................................................................... 30 1.5 Fibrinolysis ...................................................................................................... 32 1.5.1 Plasminogen activation ............................................................................ 32 1.5.2 Tissue plasminogen activator .................................................................. 34 x 1.5.3 Other plasminogen activators .................................................................. 35 1.5.4 Fibrinolysis initiation ................................................................................. 36 1.5.5 Plasmin cleavage of fibrin ........................................................................ 38 1.5.6 Fibrinolysis regulation .............................................................................. 38 1.5.6.1 Serine protease inhibition ..................................................................... 38 1.5.6.2 Thrombin activatable fibrinolysis inhibitor ............................................. 39 1.6 Hemostasis enzymology ................................................................................. 39 1.6.1 Zymogen activation .................................................................................. 39 1.6.2 Mechanism of serine proteases ............................................................... 40 1.6.3 Cofactor enhancement and substrate specificity ..................................... 42 1.7 Plasmin regulation of hemostasis ................................................................... 43 1.7.1 Plasmin pleiotropy ................................................................................... 43 1.7.2 Regulation of protein C by plasmin .......................................................... 43 1.7.2.1 Protein C structure and function ........................................................... 43 1.7.2.2 Plasmin modulation of protein C .......................................................... 48 1.7.3 Auxiliary fibrinolysis cofactors .................................................................. 49 1.7.3.1 Vitamin K-dependent serine proteases ................................................ 51 1.7.3.2 Factor V ................................................................................................ 53 1.7.3.2.1 Synthesis ....................................................................................... 53 1.7.3.2.2 Activation of FV .............................................................................. 54 1.7.3.2.3 Role of the FV B-domain................................................................ 56 1.7.3.2.4 FVa structure-function relationships .............................................. 56 1.7.3.2.5 Cleavage of FVa by plasmin .......................................................... 58 xi Chapter 2: Hypothesis and goals .................................................................................. 60 2.1 Hypothesis ...................................................................................................... 60 2.2 Goal ................................................................................................................ 60 2.3 Objectives ....................................................................................................... 60 2.3.1 To determine the effect on anticoagulant and cytoprotective functions of PC following proteolysis by plasmin. ........................................................ 60 2.3.2 To dissect the mechanism by which plasmin proteolytically modulates FVa from a procoagulant to fibrinolytic cofactor. ............................................. 61 Chapter 3: Materials and methods ................................................................................ 62 3.1 Materials ......................................................................................................... 62 3.1.1 Proteins and plasmas .............................................................................. 62 3.1.2 Substrates and inhibitors ......................................................................... 62 3.1.3 Additional reagents .................................................................................. 63 3.2 Anionic phospholipid preparation .................................................................... 64 3.2.1 Small unilamellar vesicles (SUVs) ........................................................... 64 3.2.2 Large multilamellar vesicles (LMVs) ........................................................ 65 3.3 Recombinant FV-810 generation .................................................................... 65 3.3.1 Expression ............................................................................................... 65 3.3.2 Purification ............................................................................................... 66 3.3.3 Labelling Lys-plasminogen with 125I ......................................................... 68 3.4 Preparing plasmin-cleaved samples for functional assays .............................. 69 3.4.1 FVa and PC ............................................................................................. 69 3.4.2 APC-inactivated FVa ............................................................................... 70 xii 3.4.3 Aprotinin or p-APMSF inhibition of residual plasmin ................................ 70 3.4.4 Separation of lipid-binding fragments with LMVs ..................................... 72 3.5 Protein staining (Coomassie blue and silver staining) ..................................... 73 3.6 Western blotting .............................................................................................. 73 3.7 Assessment of fibrinolytic function .................................................................. 74 3.7.1 Purified tPA-mediated plasminogen activation ......................................... 74 3.7.2 One-step fibrinolysis assay using purified proteins .................................. 74 3.8 Assessment of APC activity ............................................................................ 75 3.8.1 Chromogenic substrate cleavage ............................................................ 75 3.8.2 Activated partial thromboplastin time (APTT) in PC-deficient plasma ...... 76 3.9 125I-radiolabeled plasminogen binding and dependency on C-terminal lysines ... ........................................................................................................................ 76 3.10 Edman degradation N-terminal sequencing .................................................... 77 3.11 Statistical analysis ........................................................................................... 77 Chapter 4: Functional modulation of protein C by plasmin ............................................ 78 4.1 Rationale ......................................................................................................... 78 4.2 Hypothesis ...................................................................................................... 81 4.3 Results ............................................................................................................ 81 4.3.1 Proteolysis of PC-Hc correlates with increased APC amidolytic activity ... 81 4.3.2 N-terminal sequencing of PC fragments following plasmin cleavage ....... 84 4.3.3 APC retains amidolytic activity following plasmin cleavage ..................... 86 4.3.4 Sequencing of APC-derived fragments following plasmin cleavage ........ 87 xiii 4.3.5 APC activity decreases with prolonged plasmin cleavage or autoproteolysis ......................................................................................... 88 4.3.6 Dissecting autoproteolytic-mediated fragments of APC ........................... 90 4.3.7 Plasmin-treated PC has impaired anticoagulant function compared to APC ................................................................................................................. 91 4.3.8 Fragmentation of FVa by plasmin-proteolyzed PC .................................. 91 4.3.9 Plasmin cleavage does not convert (A)PC into a fibrinolytic cofactor ...... 93 4.4 Discussion ....................................................................................................... 95 4.4.1 Proteolysis of PC by plasmin ................................................................... 95 4.4.2 Proteolysis of APC by plasmin ................................................................. 97 4.4.3 Functional modulation of (A)PC by plasmin ............................................. 98 4.4.3.1 Proteolysis of PC by plasmin impairs anticoagulant activity ................. 98 4.4.3.2 Conversion into a tPA accelerator ...................................................... 100 4.4.4 Physiologic relevance of plasmin modulating (A)PC .............................. 102 4.5 Future studies ............................................................................................... 103 4.5.1 Further dissecting cleavage and enzymatic activation of (A)PC by plasmin ............................................................................................................... 103 4.5.2 Potential role for preserved PAR-1 signaling and cytoprotection ........... 104 4.5.3 Thrombolysis with direct plasmin administration .................................... 105 Chapter 5: To dissect the mechanism by which plasmin converts FVa from a procoagulant to a fibrinolytic cofactor .......................................................................... 107 5.1 Rationale ....................................................................................................... 107 5.2 Hypothesis .................................................................................................... 108 xiv 5.3 Results .......................................................................................................... 109 5.3.1 Fragmentation of FVa by plasmin .......................................................... 109 5.3.2 Fragmentation of FVa results in enhanced plasminogen activation by tPA . ............................................................................................................... 110 5.3.3 Plasmin cleavage of FVa on aPL accelerates conversion into a tPA cofactor .................................................................................................. 111 5.3.4 Longer proteolysis of FVa by plasmin correlates to enhanced tPA activity .. ............................................................................................................... 113 5.3.5 Plasmin cleavage is required to convert FVa into a tPA cofactor ........... 114 5.3.6 APC-inactivated FVa remains plasmin cleavable .................................. 117 5.3.7 Plasmin modulates APC-inactivated FVa into fragments that accelerate tPA ......................................................................................................... 118 5.3.8 Free and lipid-bound fragment mixtures both accelerate plasmin generation .............................................................................................. 119 5.3.9 Binding of FVa fragments to 125I-radiolabelled plasminogen .................. 121 5.3.10 The plasminogen-binding fragment of FVa is derived from FVa-Hc ....... 122 5.3.11 Plasmin converts FV-810 into a tPA accelerator like FVa ...................... 122 5.3.12 FV-810 enhances one-step fibrinolysis .................................................. 123 5.3.13 The residual B-domain of FV-810 after thrombin excision may accelerate tPA ......................................................................................................... 125 5.4 Discussion ..................................................................................................... 126 5.4.1 FVa-derived fragments responsible for enhancing tPA .......................... 126 5.4.2 Acquisition of tPA cofactor activity ......................................................... 129 xv 5.5 Future studies ............................................................................................... 131 5.5.1 Enhancing fibrinolysis ............................................................................ 131 5.5.2 Dissecting the tPA-accelerating mechanism of FVa fragments ............. 131 5.5.3 Fibrinolysis enhancement in vivo ........................................................... 132 Chapter 6: Conclusion ................................................................................................. 133 Bibliography ................................................................................................................ 136 xvi List of Tables Table 1: N-termini of major fragments following cleavage of PC by plasmin ................. 84 Table 2: N-termini of major fragments following cleavage of APC by plasmin .............. 87 xvii List of Figures Figure 1: Categorizing thrombosis, treatment with tPA, and risks of thrombolysis .......... 2 Figure 2: General mechanisms of hemostasis ................................................................ 8 Figure 3: Overview of secondary hemostasis ................................................................ 10 Figure 4: An overview of the cell-based model of coagulation ...................................... 11 Figure 5: Ligand-receptor interactions that mediate platelet adhesion, activation and aggregation ................................................................................................................... 13 Figure 6: Prothrombin activation ................................................................................... 16 Figure 7: Amplification of thrombin by procoagulant complex on aPL ........................... 19 Figure 8: Thrombin activation of PARs .......................................................................... 21 Figure 9: Key interactions that facilitate the initial steps of fibrin polymerization ........... 23 Figure 10: The mechanism of action of serpins ............................................................. 27 Figure 11: Mechanism of plasminogen activation ......................................................... 33 Figure 12: Structures of single- and two-chain tPA ....................................................... 34 Figure 13: Fibrinolysis pathway ..................................................................................... 37 Figure 14: Serine protease mechanism ......................................................................... 41 Figure 15: Structure of PC ............................................................................................. 44 Figure 16: Protein C pathway ........................................................................................ 45 Figure 17: Biased signaling of PAR-1............................................................................ 46 Figure 18: Auxiliary cofactor model of fibrinolysis ......................................................... 50 Figure 19: Conversion of FIXa and FXa to tPA cofactors by plasmin ............................ 53 Figure 20: FV structure .................................................................................................. 54 Figure 21: Cleavages that activate FV and inactivate FVa ............................................ 55 xviii Figure 22: Major plasmin cleavage sites within FVa ..................................................... 58 Figure 23: Structure of FV-810 ...................................................................................... 66 Figure 24: Protein stain and thrombin cleavage of FV-810 ........................................... 67 Figure 25: Inhibition of plasmin activity by p-APMSF .................................................... 71 Figure 26: Inhibition of plasmin activity by aprotinin ...................................................... 72 Figure 27: Clot formation and dissolution during fibrinolysis ......................................... 75 Figure 28: Plasmin cleavage of PC-Hc activates APC-like activity ................................. 82 Figure 29: Aligning N-terminal sequencing to fragmentation of PC by plasmin ............. 85 Figure 30: Amidolytic activtiy of APC is enhanced following plasmin cleavage ............. 86 Figure 31: Aligning fragmentation of APC by plasmin to N-terminal sequences ........... 88 Figure 32: Loss of APC activity over time during prolonged plasmin incubation ........... 89 Figure 33: Comparing fragmentation of APC in the presence or absence of plasmin ... 99 Figure 34: Plasmin cleavage impairs the anticoagulant function of PC ......................... 92 Figure 35: Plasmin cleavage of PC impairs its ability to cleave FVa ............................. 93 Figure 36: Plasmin does not convert (A)PC into a tPA accelerator ............................... 94 Figure 37: Plasmin cleavage sites in PC ....................................................................... 95 Figure 38: Theoretical model illustrating the sequential plasmin cleavages of PC ........ 96 Figure 39: Proteolysis of APC by plasmin ..................................................................... 97 Figure 40: Highlighting critical surface loops that mediate APC function ....................... 98 Figure 41: Sequence alignment of FVII, FIX, FX, and PC ........................................... 101 Figure 42: A proposed model for the functional regulation of (A)PC by plasmin ......... 103 Figure 43: Plasmin cleaves FVa-Hc and FVa-Lc into a complex array of fragments .... 109 Figure 44: FVa enhances tPA-mediated plasminogen activation ................................ 111 xix Figure 45: Plasmin cleavage of FVa in the presence or absence of aPL .................... 112 Figure 46: Plasmin cleavage of FVa on aPL accelerates fibrinolytic activity ............... 113 Figure 47: Correlating fragmentation of FVa by plasmi to tPA cofactor function ......... 114 Figure 48: Aprotinin prevents conversion of FVa into a tPA cofactor by plasmin ........ 115 Figure 49: Cleavage of FVa by APC does not confer tPA cofactor activity ................. 116 Figure 50: Fragmentation of FVa by APC and plasmin ............................................... 117 Figure 51: Plasmin converts APC-inactivated FVa into a tPA cofactor ....................... 118 Figure 52: Seaparation of lipid-bound and free fragments on LMVs ........................... 119 Figure 53: Free and lipid-bound fragments enhance tPA cofactor activity .................. 120 Figure 54: Binding of FVa fragments to 125I-radiolabelled plasminogen ...................... 121 Figure 55: Binding of lipid-bound and free fragments to 125I-plasminogen .................. 122 Figure 56: FV-810 enhances tPA-mediated plasminogen activation ........................... 123 Figure 57: FV-810, but not FV or FVa, enhances fibrinolysis ...................................... 124 Figure 58: Fragmentation of FV-810 during fibrinolysis ............................................... 125 Figure 59: The tPA-accelerating fragments derived from cleavage of FVa ................. 128 Figure 60: Proposed mechanisms of tPA-accelerating fragments derived from FVa .. 130 xx List of Abbreviations ADP adenosine diphosphate Akt protein kinase B AP agonist peptide APC activated protein C aPL anionic phospholipid APTT activated partial thromboplastin time AR acidic region AT antithrombin ATP adenosine triphosphate Bax Bcl-2-like protein 4 BCA bicinchoninic acid Bcl-2 B-cell lymphoma 2 BHK baby hamster kidney BR basic region CAD coronary artery disease cAMP cyclic adenosine monophosphate CAPS N-cyclohexyl-3-aminopropanesulfonic acid CP ceruloplasmin CPM counts per minute CTL C-terminal lysine CVD cardiovascular disease DMEM/F12 Dulbeccoâs Modified Eagle Medium: Nutrient Mixture F-12 xxi DMSO dimethyl sulfoxide DOACs direct oral anticoagulants DTT 1,4-dithiothreitol DVT deep-vein thrombosis EACA ε-aminocaproic acid ECL extracellular loop ECM extracellular matrix EDTA tetrasodium ethylenediaminetetraacetic acid EGF epidermal growth factor EPCR endothelial protein C receptor ERK1/2 extracellular signal-regulated kinase 1/2 FBS fetal bovine serum FDA Food and Drug Administration FDPs fibrin degradation products FGF fibroblast growth factor FpA fibrinopeptide A G418 Geneticin® GEMSA 2-guanidinoethylmercaptosuccinic acid GI gastrointestinal Gla γ-carboxyglutamic acid GP glycoprotein GPCR G-protein coupled receptor HBS Hepes-buffered saine xxii Hc heavy chain HEK human embryonic kidney HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HMWK high-molecular-weight kininogen HUVECs human umbilical vein endothelial cells ICL intracellular loop ITS insulin-transferrin-selenium kb kilobases kDa kilodalton KR kringle Lc light chain Leu leucine LMVs large multilamellar vesicles MI myocardial infarction MKs megakaryocytes MMP matrix metalloprotease mOD milli optical density MWCO molecular weight cut off NETs neutrophil extracellular traps NMDA N-methyl-D-aspartate OD optical density OH hydroxyl p pellet xxiii p53 tumor protein p53 PAGE polyacrylamide gel electrophoresis PAI-1 type I plasminogen activator inhibitor PAP pre-activation peptide p-APMSF 4-amidinophenylmetanesulfonyl fluoride hydrochloride PAR protease-activated receptor PC protein C PCI protein C inhibitor PDGF platelet-derived growth factor PE pulmonary embolism PEG polyethylene glycol Pen-Strep penicillin-streptomycin PF4 platelet factor 4 Pgn plasminogen PI3K phosphatidylinositol-3-kinase PK prekallikrein PK protein kinase PKG protein kinase G PLCβ phospholipase β Pn plasmin or plasmin-treated pNA para-nitroalinine PS protein S PT prothrombin time xxiv PtdCho phosphatidylcholine PtdSer phosphatidylserine PVDF polyvinylidene difluoride q long arm Rac1 Ras-related C3 botulinum toxin substrate 1 RBC red blood cell RhoGEF Rho guanine nucleotide exchange factor RT room temperature s supernatant S-2251 H-D-Val-Leu-Lys-pNA S-2288 H-D-Ile-Pro-Arg-pNA S-2366 H-D-Glu-Pro-Arg-pNA SDS sodium dodecyl sulfate SEAP secreted embryonic alkaline phosphatase SMCs smooth muscle cells STEMI acute ST-elevated MI SUVs small unilamellar vesicles TAFI thrombin activatable fibrinolysis inhibitor TBS Tris-buffered saline TCA trichloroacetic acid TF tissue factor TG transglutaminase TGF transforming growth factor xxv TM thrombomodulin or transmembrane tPA tissue plasminogen activator TPFI tissue factor pathway inhibitor Tris tris(hydroxymethyl)aminomethane TX2 thromboxane A2 uPA urokinase-type plasminogen activator uPAR urokinase-type plasminogen activator receptor VECs vascular endothelial cells VTE venous thromboembolism vWF von Willebrand Factor WPBs Weibel-Palade bodies α2-AP α2-antiplasmin α2-M α2-macroglobulin β-ME 2-mercaptoethanol xxvi Acknowledgements There are many people that I would like to sincerely thank for giving me such valuable opportunities and guiding me throughout my life. First, my parents Cliff and Lina, and my older sister Jane for their unwavering support. You three are pillars in my life and I would not be where I am without your love and care. I will be forever grateful to my supervisor, Dr. Ed Pryzdial, for being a mentor and friend both in and outside the lab. Your patience, care, and encouragement during the past seven years of my life is sincerely appreciated. I hope to continue our friendship and perhaps one day work together as colleagues. I am grateful to my supervisory committee members, Drs. Edward Conway, Shannon Jackson, and Haydn Pritchard for their continued support throughout my doctoral studies. To my present and past lab mates in the Pryzdial Lab, Drs. Michael Sutherland and Bryan Lin, Scott Meixner, Kim Talbot, Tseday Tegegn, Henry West, Katherine De Asis, Rolinda Carter, and Simon Ayo, thank you all for welcoming and accepting me, and helping foster a community striving towards novel scientific pursuits. Your constant encouragement and positive energy are inspiring. Thank you to all the members of the Centre for Blood Research that help make up such a fine Canadian institution at the forefront of blood research. I am also deeply grateful for Dr. Howard Chan, for giving someone like me a chance to experience the reality of basic science research. You treated me like a son while at McMaster University, I miss you dearly and I hope all is well with you. Lastly, thank you to my wife and partner Briana, you give meaning and purpose to my life as we both find our place within this world together. xxvii Dedication This work is dedicated to my family, mentors, colleagues, and friends who have supported me along this journey.1 Chapter 1: Introduction 1.1 Global thrombosis burden 1.1.1 Epidemiology Myocardial infarction (MI), ischemic stroke, and venous thromboembolism (VTE) together account for at least 1 in 4 deaths globally [1-4]. Their underlying pathology involves the formation of blood clots, or thrombi, that obstruct the normal flow of blood (i.e. thrombosis). They are generally categorized into thrombosis of either the arteries or veins. MI and ischemic stroke are the result of arterial thrombosis and are manifest by compromised heart and brain function. VTE comprises deep-vein thrombosis (DVT), which often originates in the legs, and pulmonary embolism (PE) in the lungs (Figure 1). If normal blood flow cannot be restored in a timely fashion, the consequences can be fatal. Deaths due to ischemic heart disease and stroke have risen in line with global population growth, increasing approximately 30% yearly and presents an enormous burden to healthcare systems [3]. Total direct and indirect costs in Canada attributed to cardiovascular disease (CVD) and strokeâwhich include MI and ischemic strokeâexceed $21 billion each year, representing greater than 10% of the total Canadian cost of illness [5]. In the United States, annual costs associated with CVD and stroke are estimated to be over $300 billion [6]. Incidence rates of VTE are expected to increase as more âbaby boomersâ reach geriatric age [7]. Total costs associated with VTE are estimated to be as high as $70 billion per year in the US and corresponds to at least $600 million when extrapolated to the Canadian population [8]. Healthcare costs due to thrombotic-related conditions will unfortunately continue to rise and have significant and measurable effects on the quality of life of individuals and their affected families [9-13]. It 2 is imperative that we develop more effective interventions for prevention, early detection, treatment, and long-term care. Figure 1: Categorizing thrombosis, treatment with tPA, and risks of thrombolysis Thrombosis is categorized clinically by its origin in either arterial or venous circulation and is further stratified into MI, ischemic stroke, and VTE. VTE comprises DVT and PE. Tissue plasminogen activator (tPA) is a therapeutic option in all thrombosis but is used most frequently in ischemic stroke. Major side effects attributed to tPA use are bleeding, cellular toxicity, and the potential for reocclusion. 1.1.2 Clot-dissolving drugs Treatment regimens vary significantly for different types of thrombosis, but they each to some degree involve the use of clot-dissolving (i.e. thrombolytic) drugs. Use of these drugs is reserved for patients requiring rapid thrombolysis and restoration of blood 3 flow. Approved thrombolytic drugs and new therapies currently in development function through varied mechanisms but all converge on enhancing the generation of the enzyme plasmin from its naturally circulating precursor plasminogen [14, 15]. Plasmin works by cleaving and solubilizing fibrin, a dense protein matrix of fibers that form the structural scaffold of thrombi [16]. The most prevalent drugs in the market today are administered intravenously. These recombinantly derived clot-dissolving therapeutics were specifically designed based on the structure and biochemistry of tissue plasminogen activator (tPA) [17]. tPA is synthesized and stored in vascular endothelial cells (VECs) and its main role following release into the bloodstream is to convert circulating plasminogen into plasmin to dissolve thrombi [18]. In cases of thrombosis, when natural endogenous levels of tPA release are insufficient to dissolve the clot and restore blood flow, exogenous tPA must be administered to recanalize the affected vessel. 1.1.3 Thrombolysis needs improving When the first recombinant form of full-length tPA, alteplase (Activase®), received Food and Drug Administration (FDA) approval in 1996 and came into market, the drug was a breakthrough particularly in the treatment of ischemic stroke [19]. However, use of the drug was associated with an elevated hemorrhage risk since high therapeutic doses (~0.5-1 mg/kg) were required to overcome its rapid clearance by the liver [20, 21]. The bleeding side effect associated with tPA is attributed to the systemic activation of plasminogen into plasmin, tipping the haemostatic balance towards a bleeding phenotype [22, 23]. Although tPA has been engineered to have enhanced fibrin-binding and a longer circulating half-life, these efforts have not been able to unequivocally increase its efficacy and reduce hemorrhage risk [24, 25]. Other significant risk factors for patients that receive 4 tPA for thrombolysis include vessel reocclusion [26-28] and toxicities that have been attributed to tPA permeating into extravascular tissues [29-31]. The bulk of clinical research on tPA has been done in ischemic stroke, where patients benefit from early treatment within a 4.5 hour window [32]. However, only 2-5% of all ischemic stroke patients receive tPA, as studies show the large proportion of eligible patients have historically been refused tPA due to varied reasons unrelated to legitimate pre-existing contraindications [33, 34]. The first landmark trials conducted following the development of alteplase showed a modest yet significant benefit with alteplase compared to placebo but was overshadowed by an incidence ranging from 2.4% to 8.8% of symptomatic intracranial hemorrhage in patients receiving tPA [19, 35-38]. Although thrombolysis with tPA demonstrates reperfusion efficacy, more recent clinical data has shown acute recanalization in approximately 1 in 5 ischemic stroke patients treated with alteplase [39]. To improve reperfusion efficacy and reduce hemorrhage risk, tenecteplase (TNKase®) was developed and FDA-approved in 2000 by modifying tPA to improve fibrin-binding and increase its circulating half-life [40]. However, it remains inconclusive as to whether tenecteplase provides a significant benefit over alteplase in recanalizing vessels and lowering the risk for symptomatic hemorrhage [41-44]. Similarly, landmark clinical trials in patients with acute ST-elevated MI (STEMI) have also shown unequivocally the benefit of tPA-mediated thrombolysis in enhancing reperfusion at the cost of increased hemorrhage risk [45-48]. Non-bleeding side effects are also major limitations to thrombolytic therapy. Vessel reocclusion is a significant complication of thrombolysis particularly in arterial thrombosis [27, 49, 50]. The precise mechanisms are poorly understood but are thought to be related 5 to enhanced platelet activation, which complicates treatment protocols with the need for adjunctive therapies [51-54]. This dysfunction of hemostasis associated with tPA use may be in part attributed to effects of high plasmin levels cleaving substrates apart from fibrin. Moreover, by administering high doses of tPA, there also is a damaging role for these active enzymes when they leak out of the intravascular space, where extensive work has shown mechanisms that mediate enhanced inflammation and cellular toxicity [29-31]. This is particularly worrisome in stroke, where novel treatment designs have been developed including recombinant ADAMTS13 and an inactive variant of recombinant tPA that works to outcompete for wild-type tPA or tenecteplase for the glutamate N-methyl-D-aspartate (NMDA) receptor to help prevent neurotoxicity [55, 56]. The mechanism by which exogenous tPA can exacerbate neurotoxicity during thrombolysis is thought to involve disruption of the blood brain barrier and stimulating apoptosis of cortical cells, which increases infarction size and cerebral edema [29, 57, 58]. Systemic thrombolysis in VTE is less common and reserved for cases where there is a high likelihood of reducing clot burden while minimizing bleeding risk as patients are often on rigorous anticoagulant therapy regimens [59]. The severity of clot burden is generally assessed by location, degree of occlusion and the presence or absence of collateral blood supply [60]. Lowering clot burden improves several measurable outcomes in patients including preserving functional capacity, symptom relief, and most importantly reducing mortality [61]. Due to ease of access, catheter-directed thrombolysis is typically preferred over systemic administration in DVT to decrease bleeding risk [62, 63]. PE is subdivided by severity into categories including sub-massive and massive PE, where there has been clinical evidence supporting systemic thrombolysis for only the former with 6 the tradeoff in both contexts being increased bleeding risk [64]. In sub-massive PE, newer technology (e.g. FlowTriever System, Inari Medical, California) to mechanically remove thrombi (i.e. thrombectomy) have demonstrated efficacy in improving cardiopulmonary outcomes without the need for tPA, but are associated with increased adverse events due to the damaging nature of the surgical intervention itself [65]. Although treatment of VTE is transitioning towards thrombectomy and catheter-directed thrombolysis, these methods are imperfect and will require more effective approaches in the future. Improving thrombolysis has proved in the past decades to be an incredibly arduous task. A more effective therapeutic would have to possess superior clot-dissolving efficacy while minimizing bleeding risk. Although thrombectomy has been gaining traction, the invasive nature of these procedures carries great risk and are still largely used and studied in conjunction with thrombolytic drugs. Use of thrombectomy is also limited by anatomical location, where in stroke, thrombectomy in conjunction with tPA is reserved for obstructions in larger arteries [66]. To avoid systemic side effects associated with plasminogen activators, significant interest over the past decade has turned towards targeted delivery of thrombolytics using synthetic carriers [67]. These include echogenic liposomes and gas-filled microbubble contrast agents that are activated to release their contents by ultrasound [67]. Significant work has also been done using magnetically guided carriers of thrombolytics, utilizing nanoparticles containing a magnetic core, like iron oxide encapsulated by either an organic (e.g. dextran) or inorganic (e.g. silica) outer shell [67]. These approaches have shown promise in animal models but have yet to advance to clinical testing [67-70]. With the technological advances seen in catheter-directed thrombolysis, interest has also 7 turned towards developing direct-acting thrombolytics that include plasmin and plasmin derivatives [71]. Direct-acting thrombolytics proteolytically degrade fibrin and have the added advantage of not requiring an intermediate step of plasminogen activation [71]. In addition to putting effort into developing the next-generation plasminogen activator or targeted thrombolytic therapies, perhaps there needs to be a greater focus on understanding the dysregulation of hemostasis caused by tPA-mediated thrombolysis where large amounts of plasmin are generated. Plasmin is widely known to have multiple substrates apart from fibrin and the effects of a sharp increase in circulating and local plasmin concentrations on other haemostatic constituents may be overlooked. 1.1.4 Thesis rationale After decades of research and development into tPA-based thrombolytics, these drugs have failed to demonstrate both a significant improvement in efficacy and reduction in life-threatening side effects. This suggests there are gaps in understanding how clots are dissolved at the molecular level and further exploring the interplay of these biological mechanisms may help guide the development of more effective and safer therapeutics. This research thesis focuses on understanding functions of plasmin outside of its well-established role of cleaving and solubilizing fibrin. The first part of this work investigates a link between plasmin-mediated fibrinolysis and the anticoagulant pathway where plasmin modulates the proenzyme protein C (PC) into its activated state. This oddly disrupts its well-established anticoagulant activity but may preserve cytoprotective function. The second part focuses on dissecting how plasmin converts activated clotting factor (F) V (FVa) into a tPA accelerator, a mechanism that our lab believes will pave the way to a safer, cofactor-based approach to dissolving clots. Collectively, further 8 understanding the modulatory roles of plasmin on these circulating haemostatic proteins may pave the way to the development of more effective approaches to treat thrombosis. 1.2 Hemostasis 1.2.1 Overview of hemostasis Hemostasis is a system of complex pathways that responds to vascular injury by functioning to keep blood in a fluid state while preventing bleeding. These intricate processes work in unison to fine-tune a balance between clot formation and bleeding, where an imbalance results in either thrombosis or uncontrolled hemorrhage. Briefly, vascular injury provides a stimulus for coagulation that overcomes constitutive anticoagulation mechanisms, resulting in a thrombus that seals off the damaged site (Figure 2). As coagulation progresses and wound healing processes repair the damaged vessel, anticoagulant and fibrinolytic pathways become activated and enhanced. Anticoagulation downregulates and prevents excessive clot formation, while fibrinolysis is ramped up in order to dissolve the thrombus and restore normal blood flow. The Figure 2: General mechanisms of hemostasis Vascular injury triggers a rapid coagulation response that overcomes constitutively active anticoagulant pathways. This results in the generation of a thrombus that seals off the site of injury to allow for wound healing and repair. Anticoagulation and fibrinolysis are then stimulated to prevent excess clot formation and solubilization of the thrombus. 9 haemostatic milieu that regulates these complex processes encompasses a multitude of components including a diverse array of vascular cells, circulating and cell surface proteins, intracellular signaling pathways, ions, and anionic phospholipid exposure. 1.3 Blood coagulation 1.3.1 Vessel injury and subendothelial exposure The vascular endothelium consists of a monolayer of cells (i.e. VECs) that separates the intravascular space from underlying tissues. It primarily functions in regulating hemodynamics (i.e. blood flow, pressure and resistance), tissue perfusion, and acts as a semi-permeable barrier to control the movement of ions and molecules [72]. Trauma or an underlying atheromatous disease are common mechanisms that can cause a break in the endothelium lining to initiate coagulation [73]. Vessel injury and subsequent exposure of the subendothelial components triggers a host of mechanisms that initiate coagulation. These include subendothelial exposure of tissue factor (TF) and collagen to flowing blood, anionic phospholipid (aPL) exposure, and release of Weibel-Palade bodies (WPBs) containing von Willebrand Factor (vWF) [74-76]. 1.3.2 Cell-based model of coagulation The coagulation response culminates in the formation of a thrombus consisting primarily of platelets supported by a meshwork of cross-linked fibrin to seal off the injured endothelium and allow for wound healing. Coagulation has been historically divided into primary and secondary hemostasis. Primary hemostasis involves mechanisms that result in the formation of platelet aggregates that âplugâ (i.e. platelet plug) the damaged site. To further stabilize the platelet plug, platelets and other vascular constituents provide binding surfaces for coagulation factors [77]. These coagulation factors participate in a series of 10 protease-dependent steps that amplify thrombin generation and converts circulating fibrinogen to fibrin (Figure 3). Fibrin then polymerizes and undergoes further cross-linking by FXIIIa to form a scaffold supporting platelet aggregates [78]. Current understanding of blood coagulation has evolved significantly and is now referred to as being âcell-basedâ where coagulation occurs on the surfaces of those cells involved in three phasesâ initiation, amplification, and propagation [77, 79, 80] (Figure 4). Figure 3: Overview of secondary hemostasis This illustration overviews the primary components of secondary hemostasis that lead to the formation of fibrin. These protease-dependent steps involve a multitude of positive feedback mechanisms (green; dashed) mediated by thrombin. Regulation of secondary hemostasis by anticoagulant pathways have also been included (red; dashed). To note, the primary roles of PC and FV have been highlighted as well as a potential modulatory role of plasmin. These components will be explored in detail in this introductory review. 11 Figure 4: An overview of the cell-based model of coagulation Coagulation occurs in three main phases that rely largely on cells that express certain procoagulant elements. During the initiation phase, platelets adhere to subendothelial components such as collagen. Vessel injury causes cells to expose TF which binds FVIIa, leading to generation of FIXa and FXa. FXa then goes onto generate thrombin to begin the amplification phase where thrombin activates circulating coagulation procofactors FV and FVIII. Thrombin also plays a major role in activating platelets to express aPL-binding sites and induce platelet aggregation (dashed grey line). Formation of procoagulant complexes on aPL amplifies thrombin generation by several orders of magnitude. This results in the propagation phase where thrombin cleaves and converts fibrinogen to fibrin. Fibrin then undergoes polymerization and crosslinking to form a mature thrombus. Physiologically relevant intricacies were left out to simplify the model diagram. These additional complex pathways will be discussed in depth in this introductory review. 12 1.3.3 Initiation 1.3.3.1 Platelet adhesion Platelets circulate as small, anucleate cells derived from cytoplasmic fractionation of megakaryocytes (MKs) in the bone marrow. They contain two types of secretory granules. Alpha (α-) granules are the most abundant and store various functional molecules including platelet hemostatic factors (e.g. P-selectin, platelet factor 4; PF4, FV, type I plasminogen activator inhibitor; PAI-1) [81]. Dense granules are characterized by their high calcium (Ca2+) content and mediate platelet recruitment and activation through serotonin and adenosine diphosphate (ADP) release [82]. Platelets play a vital role in hemostasis by undergoing complex processes to form platelet aggregates, which are stabilized by cross-linked fibrin to stop bleeding and allow vessel repair [83-85]. Upon vessel damage, platelet adhesion to the site of injury is mediated by interactions between platelet surface receptors and subendothelial extracellular matrix components (Figure 5) [86]. The platelet glycoprotein (GP) Ib/V/IX receptor complex interacts with matrix-bound vWF under high shear, whereas GPVI and GPIa mediate binding to collagen [87, 88]. Platelets are then activated by various ligand-receptor interactions including thrombin cleavage of protease-activated receptors (PARs), ADP binding to the purigenic receptor P2Y12, and thromboxane A2 (TX2) binding [84, 88, 89]. This occurs through multiple complex signaling pathways leading to functionally distinct aspects of platelet activation (e.g. Ca2+ and granule release, rapid translocation of phosphatidylserine for procoagulant activity, actin polymerization causing platelet shape change, and conversion of integrin GPIIb/IIIa into an active form) [84]. 13 Platelet-associated diseases can result from either a defect in platelet number or function. Heritable or acquired platelet disorders can cause either a decrease (i.e. thrombocytopenia) and bleeding diathesis orâmore rarerâincrease (i.e. thrombocytosis) in circulating platelets enhancing thrombotic risk [90-92]. Examples of platelet dysfunction that can elevate bleeding risk include: Bernard-Soulier syndrome where mutations in GPIb/V/IX impair VWF binding [93]; and gray platelet syndrome, characterized by a Figure 5: Ligand-receptor interactions that mediate platelet adhesion, activation and aggregation Various ligand-receptor interactions facilitate the core mechanisms involved in platelet adhesion, activation and aggregation [90]. Glycoproteins mediate platelet adhesion through binding to VWF and subendothelial components such as collagen. Receptors on the platelet surface are activated by specific agonists and protease cleavage. Inter-platelet linking is facilitated primarily by GPIIbIIIa and fibrinogen. Together, these interactions lead to an increase in intra-platelet Ca2+ levels and subsequent activation. 14 deficiency in α-granules due to mutations in neurobeachin-like 2 that disrupt membrane dynamics and vesicle trafficking [94]. 1.3.3.2 Anionic phospholipid exposure Eukaryotic cells are encapsulated by a plasma membrane composed of a bilayer of phospholipid molecules each made up of two hydrophobic fatty acid âtailsâ and a hydrophilic âheadâ containing a phosphate group [95]. The âheadâ group is located on both the outer membrane leaflet exposed to the exoplasmic space and the inner membrane leaflet facing the cytoplasm [95]. Different types of phospholipid vary by chemical composition and are distributed asymmetrically in a normally functioning cell [96]. Two phospholipids with central roles in the coagulation response are phosphatidylserine (PtdSer), which is confined to the cytoplasmic leaflet and phosphatidylcholine (PtdCho), present on both layers but more concentrated in the exoplasmic leaflet [97]. In circulating quiescent platelets, the intracellular adenosine triphosphate (ATP) level is kept high while Ca2+ is kept low by Ca2+-ATPase transporters [98]. ATP-dependent âfloppasesâ actively translocate PtdCho from the inner to the outer leaflet of the lipid bilayer [99]. These conditions also turn on âflippaseâ enzymes that constitutively translocate PtdSer from the outer exoplasmic to the inner cytoplasmic leaflet of the quiescent platelet plasma membrane [99]. Upon activation, a rapid spike in intracellular Ca2+ turns off the action of flippases and turns on the Ca2+-activated âscramblaseâ that translocates and exposes PtdSer on the cell surface [100, 101]. Platelet-derived microparticles also bud out during activation and together with the platelet membrane provide the primary scaffolding surfaces for many protein complexes including those involved in the coagulation response [102]. This is largely facilitated by electrostatic 15 interactions due to their negative charge, hence being often referred to as aPL [103, 104]. Once activated, PtdSer becomes expressed on the outer leaflet on platelets, representing 5-10% of the total phospholipid content [105]. 1.3.3.3 Synthesis and lipid-binding properties of coagulation factor proteins The majority of haemostatic proteins are synthesized by hepatocytes. These include vitamin-K dependent coagulation factors II (i.e. prothrombin), VII, IX, X; procoagulant procofactors V and VIII; and vitamin K-dependent anticoagulants PC, protein S (PS) and protein Z [106, 107]. Among haemostatic proteases, almost all are secreted into circulation in a strictly inactive precursor form. All of the vitamin K-dependent hemostatic factorsâwhen activatedâhave homology to the serine protease family of enzymes [108]. The synthesis of these proteins require vitamin K for the post-translational modification of a highly concentrated area of glutamates near their amino (N-) terminus that are converted into γ-carboxyglutamic acid (Gla) residues [109]. The Gla-domain of vitamin K-dependent proteins incorporates divalent ions (i.e. Ca2+, magnesium; Mg2+) to induce proper folding and exteriorly exposes hydrophobic residues important in anchoring into the phospholipid bilayer on aPL [103]. When activated, FV and FVIII are important enzyme cofactors in coagulation. Their synthesis is not dependent on vitamin K and therefore lack a Gla-domain but possess important structural domains that are able to directly interact with PtdSer-containing surfaces through hydrophobic motifs [110-112]. Deficiency or dysfunction of these hemostatic constituents can be associated with disease. Two of the most common congenital disorders or hemostasis affect the circulating quantity or function of FVIII (i.e. hemophilia A) and FIX (i.e. hemophilia B) [113]. 16 1.3.3.4 Tissue factor pathway TF is a transmembrane protein found predominantly on vascular smooth muscle cells (SMCs) and adventitial cells (i.e. pericytes and fibroblasts) positioned adjacent to the blood lumen [114]. A break in the endothelial lining exposes TF to flowing blood containing various coagulation proteins in their inactive (i.e. zymogen) forms. TF is found in an âencryptedâ inactive form and is thought to become âdecryptedâ through signaling events associated with aPL exposure [115, 116]. Its well-established function in coagulation is in initiating the coagulation pathway leading to fibrin generation by interacting allosterically to convert FVII to FVIIa through limited proteolysis [117]. Like platelets, damaged or activated cells expressing TF undergo a series of intracellular signaling events that causes PtdSer to transition to the outer membrane leaflet [118]. Figure 6: Prothrombin activation Activation of prothrombin occurs through proteolysis by FXa at the two indicated sites. This results in the release of fragments 1 (F2) and 2 (F2), yielding α-thrombin. F1 contains the Gla-domain and kringle 1, and F2 contains kringle 2. Complete activation by FXa results in α-thrombin, which consists of a disulfide-linked heavy (Hc) and light chain (Lc). Detection of fragments 1 and 2 (Fragment 1.2) has diagnostic value where elevated levels can indicate increased thrombosis risk [126]. 17 When FVIIa comes into complex with its cofactor TF, and FX (i.e. extrinsic tenase) on PtdSer-expressing surfaces, the activity of FVIIa is enhanced by at least four orders of magnitude [119, 120]. FVIIa then cleaves and activates its two primary substrates, FIX and FX, into FIXa and FXa, where the latter cleaves prothrombin into its activated form thrombin [121]. Complete activation of prothrombin by FXa removes the Gla-domain on thrombin (i.e. α-thrombin) and releases the newly activated enzyme (Figure 6) [122, 123]. This key mechanism allows thrombin to have pleiotropic control in regulating hemostasis [124]. The protease domain of thrombin contains two important anion-binding exosites on its surface that facilitate its interactions with different cofactors and substrates [125]. 1.3.3.5 Contact pathway Negatively charged artificial or biological surfaces can also initiate coagulation through a mechanism distinct from TF [126, 127]. FXII undergoes a conformational change that converts the inactive protein into FXIIa when it comes in contact with a negatively charged surface both ex vivo (e.g. glass, catheters, kaolin clay) and in vivo (e.g. polyphosphates, misfolded proteins, glycosaminoglycans) [128]. Interestingly, deficiency of FXII is not associated with disease in humans [129]. FXIIa has two substrates: it proteolyzes circulating prekallikrein (PK) and FXI into kallikrein and FXIa [128]. The primary role for kallikrein is converting high-molecular-weight kininogen (HMWK) into bradykinin, the latter which has a number of functions including regulating blood pressure, vascular permeability, and inflammation [129, 130]. The role of FXIa is in activating FIX to FIXa, which in turn proteolyzes FX to FXa to generate thrombin which triggers the amplification response in the coagulation pathway [128]. Contact pathway 18 proteins FXI and FXII are Gla-less and are not known to bind strongly to surfaces expressing PtdSer [131, 132]. 1.3.4 Amplification 1.3.4.1 Thrombin-mediated activation of FV and FVIII Thrombin undergoes a positive feedback mechanism whereby it amplifies its own production through activating FV and FVIII. These two coagulation proteins play central roles in hemostasis as procoagulant cofactors that greatly accelerate the activity of their cognate proteases when bound together in macromolecular complexes on aPL. Prior to activation, both proteins circulate in plasma as quiescent procofactors [133]. FV and FVIII both contain the domain structure A1-A2-B-A3-C1-C2 arranged from N- to carboxy (C-) terminus andâexcept for their B-domainsâare structurally homologous [134]. FV is kept in its procofactor state by an interaction between a basic and acidic region of its B-domain [135]. While FV circulates with no known carrier protein, FVIII is stabilized in plasma by direct binding to VWF through a region near the C-terminus in the C2-domain of FVIII, which also contains its PtdSer-binding site [136]. Thrombin-mediated proteolysis of FV and FVIII and subsequent release of the B-domain generates FVa and FVIIIa, where cleaving the latter also causes VWF to dissociate [133]. 1.3.4.2 Amplification of thrombin generation Cofactors FVa and FVIIIa greatly enhance their cognate enzymes when bound in macromolecular complexes on PtdSer-expressing surfaces (Figure 7) [137]. FVIIIa binds FIXa and the substrate FX in the âintrinsic tenaseâ complex, accelerating FIXa-mediated FXa generation by at least six orders of magnitude [138]. FXa activity is amplified when in complex with FVa and the substrate prothrombin in the âprothrombinaseâ complex, 19 augmenting thrombin generation by at least 5 orders of magnitude [139, 140]. Taken together, the role for coagulation cofactors FVa and FVIIIa is to generate a burst of thrombin generation that is required for sufficient fibrin deposition to seal off the site of vascular injury. Deficiency in either FV, FVIII, or rarely both are typically characterized by low levels of functional circulating antigen and can result in severe, life-threatening bleeding [141, 142]. Decades of recombinant engineering to generate modified forms of Figure 7: Amplification of thrombin by procoagulant complexes on aPL The intrinsic tenase and prothrombinase complexes are vitally important in mounting a coagulation response by amplifying FXa and thrombin generation. The domain structures of the coagulation proteins that make up these complexes are shown, with the serine proteases FIXa, FX(a), and prothrombin (II). FIXa and FXa comprise an aPL-binding Gla-domain, two epidermal growth factor (EGF) domains and a protease domain. Prothrombin has two kringle domains in place of the two EGF domains. The cofactors FVa and FVIIIa shown have two aPL-binding C1 and C2 domains and three A domains in a triangular orientation. The approximate relative size of the constituents comprising these ternary coagulation complexes are depicted. 20 these coagulation proteins with improved pharmacokinetics including longer circulating half-lives have drastically improved hemophilia care [143]. However, hemophilia patients are at risk of developing auto-antibodies to the specific factor being replaced and in those cases require bypassing agents to restore coagulation function to normal [143]. Advances continue to progress with novel developments including a bi-specific antibody that bridges together FIXa and FX, removing the need for cofactor amplification [144, 145]. 1.3.4.3 Thrombin-mediated platelet activation and aggregation Humans have 4 PARsâcloned and named in order of their discoveryâthat are differentially expressed throughout various tissues and cell types [146, 147]. PARs are a subfamily of G-protein coupled receptors (GPCRs) that carry a cryptic ligand near their N-terminus [147]. Once cleaved, the exposed ligand tethers intramolecularly to activate the receptor and downstream signaling pathways (Figure 8) [147-150]. Multiple proteases apart from thrombin (i.e. FXa, activated PC; APC) cleave each PAR at unique sites giving rise to distinct, or âbiasedâ, patterns of cellular signaling [151]. Small agonist peptides (APs) exist that mimic tethered ligand sequences and are capable of activating PARs to induce cellular signaling [152]. Thrombin is the principal activator of PAR1 and PAR4 expressed on platelets [147]. Cleavage of either PAR1 or PAR4 with thrombin can activate platelets and induce aggregation, a response that can be mimicked using PAR-specific APs [153]. PAR1 and PAR4 signal through various G-protein effectors following thrombin cleavage [154, 155]. PAR1 has been shown to couple directly with Gαq, Gα12/13, and Gαi, whereas PAR4 signals through Gαq and Gα12/13 [154]. Signaling of PAR1 and PAR4 through Gαq occurs primarily via stimulation of the phospholipase β (PLCβ) pathway resulting in release of 21 intracellular calcium stores to trigger granule secretion and integrin activation [156]. PAR1 and PAR4 signaling through Gα12/13 activates Rho guanine nucleotide exchange factors (RhoGEFs) that cause granule release and platelet shape change via myosin light chain-dependent contraction [156]. PAR1, but not PAR4, activates Gαi signaling [157, 158]. It is unclear if PAR1 couples directly with Gαi or indirectly through non-specific interactions with the ADP purigenic receptor P2Y12, which is known to bind Gαi [158, 159]. Gαi signals through three main routes; inhibiting cyclic adenosine monophosphate (cAMP) synthesis to relieve inhibition of platelet activation, stimulating protein kinase G (PKG) for granule secretion and TX2 synthesis, and activating the phosphatidylinositol-3-kinase (Pl3K) pathway for GPIIb/IIIa activation [156, 157]. The additive role of PAR1 in GPIIb/IIIa Figure 8: Thrombin activation of PARs PARs are seven transmembrane domain GPCRs that carry a cryptic ligand within their extracellular N-terminal sequence. Once cleaved by an activating protease (e.g. thrombin), this cryptic ligand is exposed and binds intramolecularly to ECL2 to facilitate receptor activation and downstream signaling [148]. A disulfide bond between TM3 and ECL2 (dashed line) stabilizes the receptor [148]. PAR1 contains a hirudin-like sequence that binds to exosite I on thrombin, while PAR4 contains an anionic region that is thought to interact with positive residues neighboring exosite I on thrombin [149, 150]. 22 activation may implicate a larger role for PAR1 in aggregating adjacent platelets through binding with fibrinogen and vWF. Activated GPIIb/IIIa mediates platelet aggregation by bridging adjacent platelets through binding with fibrinogen and vWF, forming a platelet hemostatic plug [160]. 1.3.5 Neutrophil extracellular traps (NETs) Release of the chemokine PF4 from the α-granules of platelets is a major chemoattractant responsible for recruiting leukocytes to the thrombus site during wound healing [161]. In particular, recruited neutrophils also regulate hemostasis by releasing fiber networks composed of their own DNA called neutrophil extracellular traps (NETs) [162]. NETs are known to play an important role in regulating clot structure by acting as a synergistic scaffold with fibrin to stabilize clot constituents [163]. They also have been shown to activate the contact pathway and degrade the anticoagulant tissue factor pathway inhibitor (TPFI) [127, 164]. NETs have been implicated in thrombosis due to their role in increasing the resistance of thrombi to solubilization [165]. 1.3.6 Propagation 1.3.6.1 Fibrin polymerization A mature network of fibrin is required to effectively seal off the site of vascular injury for wound healing to occur. Fibrinogen is synthesized by the liver and is one of the most abundant proteins circulating in plasma [166]. Its structure consists of 6 paired polypeptide chains (AαBβγ)2 held together by disulfide bonds (Figure 9) [167]. Thrombin initiates fibrin polymerization by preferentially cleaving fibrinopeptide A from the fibrinogen Aα-chains [168]. Thrombin-mediated cleavage of fibrinopeptide B from the Bβ-chains occurs at a much slower rate [167]. Cleavage of both fibrinopeptides A from fibrinogen 23 results in new N-termini on both the α chains forming a âknobâ that fits into binding pockets or âholesâ located on the γ-nodules of two separate fibrin monomers [169]. These knob-hole interactions allow the assembly of half-staggered two-stranded oligomers consisting of 20-25 fibrin monomers [119]. These oligomers then continue to extend longitudinally to form protofibrils [167, 170]. Lateral aggregation or âstackingâ is mediated largely though Figure 9: Key interactions that facilitate the initial steps of fibrin polymerization Paired polypeptide chains make up the core structure of fibrinogen. Thrombin cleaves off fibrinopeptides A to form a fibrin monomer, which can associate with other fibrin monomers through knob-hole interactions. These oligomers then elongate, aggregate laterally, and branch to give rise to a mature polymerized fibrin network. 24 interactions between the C-terminal portion of the α-chains (αC regions). These αC-αC interactions allow packing into dense fibers that continue to elongate and thicken [167, 170]. Exposed αC regions also anchor fibrin into the extracellular matrix (ECM) through interactions with fibronectin [171]. Finally, polymer branching occurs giving rise to a 3-dimensional fibrin network [167, 170]. Thrombin concentrations play an important role in dictating fibrin structure and the stability of the vascular seal. High thrombin levels generate thinner fibers that are highly branched and much more densely packed, making these clots relatively more resistant to degradation compared to more loosely woven and porous fibrin networks formed at lower thrombin concentrations [172]. Fibrin architecture as mediated by thrombin is important in thrombosis where higher thrombin concentrations contribute to the generation of fibrin that is more resistant to the action of clot-dissolving drugs [173]. 1.3.6.2 FXIIIa cross-linking FXIII is a transglutaminase (TG) that, when activated by thrombin, stabilizes fibrin by crosslinking its α-and γ-chains with covalent γ-glutamyl-ε-lysyl amide bonds [174]. The crosslinking process imparts mechanical strength to the thrombus and resistance to lysis [175, 176]. Besides fibrin-fibrin crosslinking, FXIIIa also directly crosslinks fibrin to specialized lipid rafts found in the platelet membrane, further stabilizing platelets within the molecular lattice [177]. FXIIIa is also known to crosslink other proteins to fibrin including α2-antiplasmin, a potent inhibitor of local plasmin generation, further enhancing fibrinolysis resistance [178]. Together, these interactions mediated by FXIIIa strengthen and allow the clot to retract, bringing the edges of an injured blood vessel closer to allow for more effective wound healing [179]. 25 Patients with VTE have clots often characterized by high red blood cell (RBCs) content (i.e. âredâ clots) and resistant to therapeutic thrombolysis [180-182]. The underlying mechanism of RBC-mediated thrombolytic resistance was recently attributed to FXIIIa crosslinking [183]. In FXIII-deficient plasma, it was shown that RBCs are extruded from developing clots formed in whole blood [183]. This phenomenon was also demonstrated in FXIII-deficient mice as well as reports of âRBC falloutâ in a family with congenital FXIII deficiency [184, 185]. The incorporation of RBCs within thrombi was shown to be due to FXIIIa crosslinking of fibrin α-chains [185]. After incorporation into thrombi, RBCs have been shown to release prothrombotic polyphosphates that contribute to fibrinolysis resistance [186, 187]. 1.3.7 Wound healing and repair The coagulation response marks the first stage of wound healing where the site of vascular injury is sealed off to prevent further blood leakage and clot retraction pulls either end of the damaged endothelium closer together [188]. Different cells in and around the thrombus and wound site secrete various pro-inflammatory cytokines and growth factors involved in the wound healing process including transforming growth factor (TGF) β, platelet-derived growth factor (PDGF), fibroblast growth factor (FGF), and EGF [189]. Cytokine release recruits inflammatory cells to the wound site where neutrophils migrate first, followed by macrophages and then lymphocytes [190]. Briefly, the primary role for recruited neutrophils is to clear cellular debris and foreign bodies [191, 192]. Macrophages release cytokines to further propagate the inflammatory response, clear apoptotic cells, and stimulate fibroblasts and angiogenesis to regenerate damaged tissue [193]. Although the role of lymphocytes still remains unclear, it is widely accepted that 26 lower lymphocyte concentrations correlate with impaired wound healing [194]. Tissue fibroblasts are then stimulated to deposit core ECM components including collagen, glycosaminoglycans and proteoglycans [195]. Over time, vascular density returns to normal and the endothelial lining regenerates [196, 197]. 1.4 Anticoagulation 1.4.1 Endothelial cells The endothelium provides a non-thrombogenic surface to keep blood in a fluid state [198]. Endothelial cells suppress platelet activation by continuously secreting nitric oxide (NO) and prostacyclin (prostaglandin I2) to keep intra-platelet Ca2+ levels low and express ecto-ADPase which cleaves ATP and ADP to maintain platelets in an inactivated state [74, 199, 200]. VECs also express various anticoagulant molecules on their surface including tissue factor pathway inhibitor (TFPI), thrombomodulin, endothelial protein C receptor (EPCR), and heparin-like proteoglycans to limit the coagulation response [201, 202]. 1.4.2 Serine protease inhibition Serine protease inhibitors (i.e. serpins) are a homologous family of proteins that circulate through plasma with constitutive anticoagulant activity [203]. They work to prevent aberrant clot formation under normal flow conditions and regulate coagulation to limit the propensity to thrombose. The most important serpin in plasma is antithrombin (AT), a glycoprotein synthesized in the liver that inactivates coagulation serine proteases including FVIIa, FIXa, FXa, FXIa and thrombin [204]. The mechanism involves a flanking âbaitâ motif on antithrombin containing an arginine (Arg; R) residue at position (P) 1 (i.e. P1) that is recognized by serine proteases (Figure 10) [205]. When the catalytic serine 27 (Ser; S) attacks the P1 arginine residue, a conformational change is induced that secures the protease and serpin together into an irreversible covalent complex [205]. The activity of antithrombin can be enhanced by a heparin cofactor, which includes the exogenous anticoagulant and heparin-like proteoglycans that line the endothelium [205]. Figure 10: The mechanism of action of serpins Serpins express a flanking âbaitâ motif that has peptide sequences with an arginine residue at the P1 position. These sequences are recognized by their cognate serine proteases and traps the protease through a covalent linkage with the flanking âbaitâ motif, leading to rapid re-arrangement of the motif that is stabilized within a secondary structure comprised of several β-pleated sheets [206]. 1.4.3 Tissue factor pathway inhibitor TFPI is homologous to Kunitz-type protease inhibitors and is synthesized in various cell types including endothelial cells, vascular SMCs and megakaryocytes [207, 208]. It exists in two isoformsâ TFPIα and TFPIÎ²âdue to alternative splicing [209]. The 28 former circulates with approximately 80% bound to apoB-containing lipoproteins and the rest unbound, while the latter has a membrane anchoring mechanism and is bound to endothelial cells [210-212]. The mechanism of TFPI differs from serpin-like anticoagulants and involves directly inhibiting the active sites of TF-bound FVIIa, free FXa, and prothrombinase FXa [213]. Overcoming TFPI inhibition is critical in being able to mount an effective coagulation response [214]. TFPIα has a basic region near its C-terminus that is homologous to the basic sequence found in the FV B-domain [215]. There exists a splice isoform of FV found in east Texas bleeding disorder where a mutation results in removal of most of the FV B-domain but retains its highly acidic region [215]. Termed âFV-shortâ, this isoform tightly binds to TFPIα and autosomal dominant patients have an â¼10- to ~20-fold increase in circulating TFPIα, resulting in a propensity to hemorrhage [215]. In these patients, FV acts as an anticoagulant cofactor to mediate the inhibitory activity of TFPIα against FVIIa and FXa [213]. Low concentrations of FV-short are found in the plasma of phenotypically normal individuals and may be a physiologic carrier of TFPIα [216]. 1.4.4 Protein C pathway While thrombin amplifies its own production by activating FV and FVIII, thrombin also downregulates itself by inducing anticoagulation to indirectly inactivate FV(a) and FVIII(a) [217]. As thrombin concentrations increase during coagulation, a lower affinity, concentration-driven interaction occurs with endothelial-bound thrombomodulin [218]. When in this complex, the ability for thrombin to cleave its procoagulant substrates is diminished and specifies thrombin activity to mainly its anticoagulant substrate PC [219]. Besides enhancing PC activation, thrombin-thrombomodulin is also a potent activator of 29 thrombin activatable fibrinolysis inhibitor (TAFI), a procarboxypeptidase B that is involved in fibrinolysis regulation by removing critical lysine residues on fibrin that act as binding sites for plasminogen and tPA [220]. PC is synthesized in the liver and either circulates in plasma or exists in complex with cell surface expressed EPCR [221]. When bound to EPCR, there is a 20-fold enhancement in PC activation by the thrombin-thrombomodulin complex [222]. Activation of PC generates APC that functions to inactivate FVa and FVIIIa and their precursors, with greater enzymatic activity towards its substrates when bound to its cofactor PS on aPL [223]. EPCR-bound APC can either dissociate from its receptor and enhance anticoagulation as previously described or stay bound to EPCR and signal via PAR-1 expressed on endothelial cells [224]. Free and EPCR-bound APC are inhibited by the serpins α1-antitrypsin and protein C inhibitor (PCI) at about the same rate, as both forms have half-lives of approximately 15 minutes [225]. Binding of APC to ECPR confers cytoprotective effects through âbiasedâ downstream signaling mechanisms following PAR-1 cleavage [226, 227]. This function of APC has been widely studied as a therapeutic target, particularly in sepsis, with limited success [228-231]. To hone in on the cytoprotective effects of APC, variants lacking procoagulant activity but retaining the ability to signal through PAR-1 are currently being developed [232, 233]. 1.4.5 Plasmin inactivation Plasmin inactivation of procoagulant constituents in blood has not been widely accepted as a major mode of modulating coagulation. However, plasmin levels during fibrinolysis overcome α1-antiplasmin inhibition and elevated concentrations may drive plasmin activity towards its other known substrates. Plasmin has been shown to cleave 30 and inactivate the procoagulant functions of FVa, FIXa, FXa and FXIIIa [234-237]. Proteolytic degradation of these molecules by plasmin would be logical as inactivating coagulation mechanisms tips the hemostatic balance towards fibrinolysis. 1.4.6 Therapeutic anticoagulation Anticoagulants are among the most widely used drugs in the world. They are divided into different classes based on their general targets and further subdivided by mechanism of action. The primary anticoagulants used today are not actually components of anticoagulant pathways but small molecule therapeutics that target different aspects of the coagulation response. This offers several practical advantages including reduced cost, lower risk profiles, and long circulating half-lives that make these therapies amenable to long-term prophylactic use for individuals at risk for thrombosis. Antiplatelet agents have been one of the most successful treatments in reducing atherothrombotic events [238-240]. For example, antiplatelet drugs have contributed to a >50% decrease in mortality from coronary artery disease (CAD) in Canada over the past two decades [241]. Aspirin and clopidogrelâthe two most widely used antiplatelet drugsâblock TX2 synthesis and ADP binding, respectively [242]. Blockade of P2Y12 and the thromboxane receptor by aspirin and clopidogrel prevent platelet activation from occurring. Many patients use these two drugs in combination to achieve efficacy [243]. Warfarin, a vitamin K antagonist, works by inhibiting vitamin K-epoxide reductase, which is critical in the essential post-translational γ-carboxylation of Gla-containing coagulation factors (e.g. prothrombin, FVII, FIX and FX) [244]. However, warfarin also reduces the production of PC and PS, which inhibits anticoagulant pathways and increases thrombosis risk [245]. Due to PC having a relatively shorter circulating half-life 31 of ~8 hours compared to other Gla-containing proteins, initial warfarin therapy is typically bridged by an anticoagulant like heparin [245]. This approach should be taken with caution as bridging therapy in atrial fibrillation patients has been associated with increased bleeding risk [246]. Various forms of heparin exist with different clinical properties, but all contain a critical pentasaccharide sequence that binds to the serpin AT and enhances AT-mediated inhibition of specific hemostatic proteases in the coagulation cascade [247]. Although potent, both warfarin and heparin use can lead to bleeding side-effects and require regular monitoring and dose adjustments [248]. Another limitation is their various drug-to-drug interactions and narrow therapeutic windows [249, 250]. The introduction of direct oral anticoagulants (DOACs) that specifically target the proteases FXa or thrombin has revolutionized anticoagulant therapy [251, 252]. DOACs are small molecule inhibitors with marked selectivity for either the FXa or thrombin active site and require less regular monitoring [253]. Rivaroxaban (Xarelto®) and apixaban (Eliquis®) are two FXa-specific DOACs approved for use in the treatment and secondary prevention of VTE, primary prevention of VTE following hip or knee replacement surgery, and the prevention of stroke in patients with non-valvular atrial fibrillation [254, 255]. Both inhibitors of FXa reach peak therapeutic levels within 2 hours and have half-lives of ~10 hours, which makes these two DOACs very practical for patients [256, 257]. Landmark trials (ROCKET-AF, ARISTOTLE-AF) comparing both drugs to warfarin therapy showed increased efficacy and decreased bleeding risk with DOACs [258, 259]. Dabigatran (Pradaxa®) is the most well-studied direct thrombin inhibitor and has almost identical pharmacokinetic properties compared with the FXa inhibitors [257]. Dabigatran has also been approved for use in non-valvular atrial fibrillation for the prevention of stroke, and 32 the treatment and prevention of VTE [260]. Although DOACs have shown efficacy and decreased risk of life-threatening bleeds (i.e. intracranial hemorrhage) compared with traditional anticoagulants, DOACs have been associated with a significant increase in gastrointestinal (GI) bleeds [261]. When comparing the efficacy and safety profiles across DOACs, the clinical data are most in favour of apixaban [262, 263]. 1.5 Fibrinolysis 1.5.1 Plasminogen activation Plasminogen, the zymogen precursor to the protease plasmin, is synthesized in the liver as an 88 kilodalton (kDa) single chain molecule and secreted into the circulation where it has a concentration of ~200 mg/L (~2 µM) and a half-life of ~2-2.5 days [264, 265]. Native plasminogen is referred to as Glu-plasminogen due to having a glutamic acid (E; Glu) residue at its N-terminus [266]. The domain structure of Glu-plasminogen comprises of an N-terminal pre-activation peptide (PAP), followed by five kringle domains (KR1-5), and a protease domain at its C-terminal end [266]. While in circulation, intermolecular domain interactions stabilize Glu-plasminogen in an inactive, globular âclosedâ conformation [267]. When bound to a surface such as fibrin, it adopts an extended or âopenâ conformation that is more readily accessible and activated into plasmin [267]. KR1, KR2, KR4 and KR5 all have been implicated in fibrin binding through lysine binding sites in vitro [267]. Binding of solvent-exposed KR1 followed by KR5 is thought to initiate fibrin binding and induce a conformational change that facilitates activation by tPA (Figure 11) [267]. The internalized lysine binding sites of KR2 and KR4 become exposed and helps stabilize the open conformation of plasminogen [267]. 33 As part of the fibrinolysis mechanism, plasmin cleaves off the PAP in Glu-plasminogen between Lys77-Lys78 to generate a variant with a new N-terminus, rendering Lys-plasminogen [268, 269]. Lys-plasminogen has a higher affinity for fibrin and is a far superior substrate relative to Glu-plasminogen for plasminogen activators [270]. This mechanism can be explained by basic residues in the PAP region (Lys50, Arg68 and Arg70) that interact with lysine binding sites in KR4 and KR5 and stabilize the closed formation in Gu-plasminogen [267]. During activation, both Glu- and Lys-plasminogen are converted into plasmin by cleavage at Arg561, resulting in a two-chain active enzyme Figure 11: Mechanism of plasminogen activation Plasminogen (Glu-plasminogen depicted) binds internal lysine residues through lysine binding sites on KR1 and KR5. This opens lysine binding sites on KR2 and KR4 and conformationally improves accessibility of the activation site by tPA. tPA binds to fibrin through its finger and KR2 domains. Fibrin acts as a scaffold to bring both tPA and plasminogen together to initiate fibrinolysis. Plasmin then cleaves fibrin to expose more lysine binding sites to enhance tPA and plasminogen binding. 34 consisting of a heavy chain comprised of KR1-5 linked by two disulfide bonds between KR5 and the proteolytically active light chain [14]. While plasmin is known to cleave many substrates, its best characterized role in hemostasis is to cleave insoluble fibrin into soluble fibrin degradation products (FDPs) [14]. 1.5.2 Tissue plasminogen activator Unlike most serine proteases, tPA is synthesized by vascular endothelial cells in its active form and is primarily stored intracellularly in WPBs with a low detectable plasma concentration of ~70 pM [271, 272]. Native tPA is a single-chain, ~70 kDa molecule comprised from the N- to C-terminus of a fibronectin finger-like (F) domain, an EGF domain, 2 kringle regions, and a protease domain (Figure 12) [273, 274]. F- and KR2-domains have been shown to interact and facilitate binding to fibrin [275]. Carbohydrate side chains and the F- and EGF-domains as well as KR1 have been shown to be major contributors of its rapid hepatic clearance in vivo with a half-life of ~5 minutes [71]. The region Lys296-His-Arg-Arg299 is required for the inhibition of tPA by its cognate serpin, PAI-1 [276]. These structural features have been exploited as a means of producing recombinant variants of tPA with better fibrin binding and longer circulating half-lives for therapeutic purposes. An example of this is tenecteplase, that has been engineered with a tetra-alanine substitution of Lys296-His-Arg-Arg299, and two point mutations in KR1 (Thr103 â Asn103, Asn117 â Gln117) to greatly impair PAI-1 inhibition and hepatic clearance [276]. Although single-chain tPA has proteolytic activity, plasmin cleaves tPA at the Arg275-Ile276 to generate disulfide-linked two-chain tPA during fibrinolysis [277]. Two-chain tPA has ~30-fold higher enzymatic activity than native single-chain tPA in solution but comparable activity when fibrin-bound [278]. Endothelial stores of tPA are secreted upon 35 activation and these enhanced levels overcome endogenous inhibition by PAI-1 and liver clearance to play a central role in accelerating fibrinolysis [279]. Figure 12: Structures of single- and two-chain tPA Plasmin cleavage at Arg275-Ile276 converts single-chain tPA to its two-chain form. The domain structure of single- and two-chain tPA are shown. The protease domain of two-chain tPA is disulfide linked to KR2. 1.5.3 Other plasminogen activators In the 1930s, the bacteriologist Dr. William Tillett and his student, Sol Sherry, discovered that Streptococcus haemolyticus isolated from febrile patients was able to dissolve thrombi [280]. Further work showed that Streptococcus haemolyticus released a plasminogen activatorânamed âfibrinolysinââto facilitate fibrin degradation [280]. Fibrinolysin was renamed to streptokinase and was a breakthrough in clinical medicine during a time when there was no therapy for thrombosis. Streptokinase was developed as a thrombolytic therapy in acute MI, where trials showed significant benefit on mortality, and was further improved when used in combination with aspirin [281]. However, the 36 development of more advanced surgical techniques and tPA-derived thrombolytics rendered streptokinase all but obsolete due to inferior efficacy and safety profiles [281]. Urokinase-type plasminogen activator (uPA) shares homology with tPA and is another important component of the plasminogen activator system. It is synthesized primarily in tubular epithelial cells in the kidney and secreted as a 54 kDa single-chain molecule [282]. Unlike tPA, single-chain uPA has no enzymatic activity and does not bind fibrin [283]. Instead, the role of uPA has been linked to tissue remodeling through proteolysis of ECM components and activation of matrix metalloproteases (MMPs) [284]. When bound to the uPA receptor (uPAR) expressed on endothelial cells, many proteases including plasmin and kallikrein convert uPA into its two-chain activated form [284]. Although uPA lacks fibrin specificity, it can still play a role in accelerating intravascular fibrinolysis when activated. uPA has greater efficacy towards activating Lys-plasminogen in the fluid phase and does not require colocalization like tPA [285]. As a result, lysine analogues like tranexamic acid that are used as antifibrinolytics to block plasminogen binding to fibrin enhance the activity of uPA by conformationally inducing a more open plasminogen conformation [286]. Activated uPA has been developed into a thrombolytic and has shown comparable safety profiles with tPA even though it lacks fibrin specificity and activates plasminogen systemically [287, 288]. The activity of uPA is inhibited by the serpins PAI-1 and PAI-2 [289]. 1.5.4 Fibrinolysis initiation Following wound healing, fibrin polymers must be solubilized in order to restore normal blood flow. Fibrin not only acts as a substrate for the serine protease plasmin, but also as a cofactor to regulate its own dissolution. Fibrinolysis initiation is driven by 37 electrostatic interactions between lysine binding sites on tPA and plasminogen and internal lysine residues in fibrin [290]. Distinct regions in fibrin(ogen) have been shown to mediate this interaction. Work with fibrinopeptides identified regions Aα148-160 as a binding site for plasminogen kringles and γ312-324 for tPA recognition via its finger domain [291, 292]. Aα221-610 and the C-terminal end from residue 392 onwards was shown to be important in lysine-dependent binding for both tPA and plasminogen [291, 292]. These initial interactions colocalize tPA and plasminogen to generate small amounts of plasmin that feedback to âprimeâ the fibrin through limited proteolysis (Figure 13) [14, 124]. This exposes C-terminal lysine (CTL) binding sites that are more accessible for tPA and plasminogen. Binding of tPA to partially degraded fibrin is 2-4 orders of magnitude stronger than to intact fibrin [293]. Also, eventual removal of the αC regions dissociate crosslinked α2-antiplasmin (α2-AP) from fibrin, increasing fibrinolysis further [294]. A burst of plasmin ensues once the antifibrinolytic threshold is exceeded and results in rapid fibrin solubilization. Figure 13: Fibrinolysis pathway Fibrinolysis is initiated by low affinity interactions between internal lysines on fibrin and lysine binding sites on tPA and plasminogen (Pgn). This allows for the generation of low, initial amounts of plasmin (Pn) that feedback to cleave fibrin and liberate CTLs. These CTLs then greatly enhance tPA and plasminogen binding, leading to a burst of plasmin generation and subsequent degradation of polymerized fibrin. Fibrinolysis, like other haemostatic pathways, is highly regulated. 38 1.5.5 Plasmin cleavage of fibrin Plasmin preferentially cleaves at the C-terminal end of arginine and lysine residues, the latter facilitating enhancement of fibrin cofactor activity [295]. Cleavage of fibrin(ogen) generates a heterogenous mixture of products, but main cleavage sites and sequential pathways have been well defined. The first attack point is AαK583, followed by αK206 and αK230 that remove nearly the entire αC region of fibrin [294]. The detached αC region is further degraded after removal and the this intermediate αC-less fibrin structure is referred to as fragment âXâ [294]. The next region proteolyzed is in the coiled coil, where plasmin cleaves specific sites within the α- (K78, K81, R95, R104, and R110), β- (K122, K133), and γ- (K53, K58, K62, K85, and K88) chains [294]. Transection releases the âD regionâ, which contains a truncated portion of the coiled coil and the β- and γ-nodules (Figure 7) [294]. Symmetrical cleavage of the other coiled coil releases the opposite âD regionâ and central âthe E regionâ, which contains the N-terminal disulfide knot formed by the two α-chains [294]. In crosslinked polymerized fibrin, the D region is covalently linked to an adjacent molecule, releasing âD-Dâ and âD-E-Dâ (noncovalently bound together through knob-hole interactions) fragments [294]. These fragments form the basis of the common D-dimer clinical assay, which correlates to thrombosis due to heightened fibrinolysis [296]. 1.5.6 Fibrinolysis regulation 1.5.6.1 Serine protease inhibition Regulation in hemostasis is a fundamental aspect of normal blood physiology and fibrinolysis is no exception. There exist numerous mechanisms that regulate plasmin generation. Of the serine protease inhibitors, the most relevant inhibitors of fibrinolysis 39 are α2-antiplasmin and PAI-1 [14]. Circulating α2-antiplasmin binds plasmin and can also be incorporated into the thrombus through cross-linking by FXIIIa to fibrin to more effectively inactivate local plasmin generation [297]. Release of PAI-1 from platelet and endothelial stores works to inhibit the activity of tPA and uPA through direct binding [298]. In addition, liver clearance of tPA and uPA is an important regulator due to the short half-lives of both enzymes [299]. 1.5.6.2 Thrombin activatable fibrinolysis inhibitor Synthesis of TAFI by the liver is another important regular of fibrinolysis by cleaving CTLs on fibrin and removing potential binding sites for tPA and plasminogen [220]. TAFI, or procarboxypeptidase B2, is part of the metallocarboxypeptidase family of enzymes and circulates in plasma as an inactive zymogen [300]. It is activated into TAFIa by the thrombin-thrombomodulin complex or less efficiently by thrombin [301]. TAFIa hydrolyzes C-terminal arginine and lysine residues [301]. Besides removing CTLs on fibrin to inhibit fibrinolysis, TAFIa is known to play an important link between hemostasis and adaptive immunity by removing C-terminal arginines from complement C5a and C3a, inactivating both of these anaphylotoxins [301]. TAFIa activity is regulated by circulating α2-antitrypsin and undergoes spontaneous decay through an auto-regulatory mechanism [302, 303]. 1.6 Hemostasis enzymology 1.6.1 Zymogen activation Almost all enzymes that are part of the haemostatic ecosystem are members of a highly conserved family of serine proteases. They share a common mechanism whereby a nucleophilic serine (Ser, S) residue in the enzyme active site attacks the carbonyl moiety of a specific peptide bond to trigger a series of catalytic steps that results in eventual 40 cleavage of the enzymeâs substrate [304]. Serine proteases are thought to have evolved from a single ancestral gene, giving rise to new functions to form the molecular complexities seen today [305]. The mechanisms of these homologous proteases are often described in reference to the chymotrypsin(ogen) amino acid numbering system to more easily compare structural features between family members [306]. These serine proteases gain enzymatic activity through proteolytic processing of an inactive zymogen precursor. Activation occurs via cleavage between amino acid residues 15 (P1) and 16 (P1â) based on chymotrypsin(ogen) numbering [304]. The newly exposed N-terminus either contains an isoleucine (Ile, I) at residue 16 or nearby, and induces a conformational change in the enzyme via an ion-pair with a highly conserved aspartic acid (Asp, D) residue at position 194 [304]. This Ile16-Asp194 interaction organizes the oxyanion hole in the enzyme active site and specificity pocket for its respective substrate(s) [304]. Undergoing conformational change as the protein transitions from a zymogen to active enzyme brings together the three amino acids that form the catalytic triad of the active site, His57, Asp102 and Ser195 [304]. 1.6.2 Mechanism of serine proteases His57, Asp102, and Ser195 together form a charge relay system whereby hydrogen-bonding between Asp102 and the aromatic moiety of His57 makes an adjacent nitrogen (N) that interacts with the hydroxyl (OH) group of the Ser195 atom more electronegative, thus increasing the nucleophilicity of the oxygen (O) atom of the OH group [307]. Upon substrate recognition, the hydroxyl O of Ser195 attacks the carbonyl of the substrate peptide bond, forming an oxyanion tetrahedral intermediate [307]. In chymotrypsin, this transition state intermediate is stabilized by backbone nitrogen atoms of Gly193 and Ser195 41 that comprise the positively charged binding pocket previously referred to as the oxyanion hole (Figure 14) [307]. Collapse of the tetrahedral intermediate occurs with the protonated His57 absorbing the two electrons from the covalent bond of the protein substrate, generating a new N-terminus [307]. A water (i.e. H2O) molecule then displaces the free polypeptide fragment and simultaneously re-protonates His57 and attacks the acylenzyme Figure 14: Serine protease mechanism This illustration shows the active site of chymotrypsin, highlighting the oxyanion binding pocket that stabilizes the tetrahedral intermediate of its protein substrate to facilitate catalysis. Following nucleophilic attack by Ser195, the backbone amide groups of Gly193 and Ser195 stabilize the oxygen atom of the substrate through hydrogen bonding. 42 forming another stabilized tetrahedral intermediate [307]. The existing covalent bond between Ser195 and the peptide chain de-protonates His57 and the carbonyl carbon reforms the double bond and the new C-terminus in the substrate is released [307]. This reforms the serine protease active site for further catalysis through the same described mechanism [307]. 1.6.3 Cofactor enhancement and substrate specificity All serine proteases prefer to hydrolyze peptide bonds on the C-terminal end of arginine or lysine residues at P1 [307]. Despite significant structural homology between members of the serine protease family, enzymes in hemostasis have strikingly distinct specificity for their cognate substrates, while those involved in digestion (i.e. trypsin) do not share the same selectivity [308]. Haemostatic enzymes preferentially recognize specific amino acid sequences both upstream (P1â¦Pn) and downstream (P1ââ¦Pnâ) of the scissile bond that are better able to âdockâ into the active site [309, 310]. Interestingly, this phenomenon is particularly evident when these serine proteases are in a macromolecular complexes with their cognate cofactor(s) and substrate(s) [311]. For instance, thrombomodulin-bound thrombin has high specificity for cleaving protein C (PC), while free thrombin cleaves a wide range of protein substrates [311]. Narrow substrate specificity when serine proteases are assembled in complex with a cofactor help explain the resulting marked enhancement in their catalytic rates [312]. These mechanisms that govern protease specificity are not fully understood but are due to complex protein-protein interactions that allosterically modulate protease function [313]. Regions (i.e. exosite regions, kringle domains, loop motifs) responsible for these interactions play an important 43 role in determining the macromolecular substrate specificity of these serine proteases as well as binding to their inhibitors. 1.7 Plasmin regulation of hemostasis 1.7.1 Plasmin pleiotropy Plasmin is known to cleave a diverse array of substrates involved in hemostasis besides fibrin. These include ADAMTS13, FVa, FXa, and FXIIIa [235, 314, 315]. Its enzymatic activity extends into the extravascular space where it has been shown to activate MMPs and TGFβ [316, 317], and cleaving ECM components including fibronectin and proteoglycans [318]. Unlike thrombin, the physiologic significance of plasmin modulating hemostasis constituents apart from fibrin is not fully understood. During fibrinolysis, high local concentrations of plasmin may have an important and overlooked regulatory role. This would be even further amplified in thrombolysis with plasminogen activators. Understanding the significance of the functional sequelae of plasmin substrates following cleavage may give insight into novel therapeutic targets to improve existing therapies. 1.7.2 Regulation of protein C by plasmin 1.7.2.1 Protein C structure and function PC is a vitamin K-dependent serine protease synthesized by hepatocytes and circulates in plasma at ~70 nM with a half-life of ~8 hours [319]. The ~62 kDa protein consists of a ~39-43 kDa heavy chain and (PC-Hc) and ~21 kDa light chain (PC-Lc) held together by a single disulfide bond (Figure 15) [320, 321]. PC-Lc is comprised of an N-terminal Gla-domain (residues 1-46) and two EGF-like domains (residues 55-90 and 94-134) [322]. PC-Hc contains the activation peptide (residues 158-169) and the protease 44 domain (residues 170-408) [322]. Approximately 30% of circulating PC lacks glycosylation at N329 (PCβ) [321]. The thrombin-thrombomodulin complex is the main physiologic activator of EPCR-bound PC by proteolysis at the R169-L170 bond [323]. Figure 15: Structure of PC PC has been illustrated by its domain structure, posttranslational modifications, disulfide linkages, and active site residues (H211, D257, S360). PC is a two-chain molecule where PC-Hc and PC-Lc are linked by a disulfide bond at C141 and C277. The well-studied anticoagulant function of APC is in proteolyzing and inactivating the homologous cofactors FVa and FVIIIa and their procofactors (Figure 16) [324]. Inactivation of these (pro)cofactors is enhanced by several orders of magnitude when APC is bound to its cofactor PS on PtdSer-expressing surfaces [219]. APC cleaves FVa on the C-terminal end of three sites, R306, R506, and R679, where cleavage of R306 and R506 fully inactivates the cofactor [325]. The R506 cleavage is kinetically favoured followed by cleavage at R306, the latter of which is enhanced by PS [325, 326]. Compared to PtdSer-bound FVa, APC inactivates free FVa and prothrombinase-bound FVa at least two orders of magnitude slower [327, 328]. APC cleaves FVIIIa at R336, R562, and R740, and like FVa cleavage, proteolysis at the first two arginine residues is required for complete inactivation [329]. Cleavage of FVIIIa at R562 results in ~70-80% lost procoagulant activity, followed 45 by cleavage at R336 which fully inactivates its cofactor function [330]. A mutation in FV of R506 â Q506 named after the city Leiden where it was first discovered (i.e. FVLeiden) is the most common inherited hypercoagulability disorder in Caucasians [331]. Evolutionarily this was thought to be selected for as the resistance to APC inactivation protected individuals, particularly women during pregnancy, from blood loss [332]. In modern day, these individuals are highly susceptible to thrombotic-related disease [333]. Figure 16: Protein C pathway As thrombin concentrations increase during the coagulation response, a low affinity interaction between thrombin and thrombomodulin on endothelial cells is formed that allows thrombin to downregulate its own production. This complex directs thrombin activity towards activating EPCR-bound PC. APC can then transition onto aPL bound to its cofactor PS to inactivate procoagulant cofactors as well as the fibrinolysis inhibitor PAI-1. APC, when bound to EPCR, can also signal through PAR-1. 46 Engineering APC has gained much interest as a therapeutic in clinical areas including inflammatory diseases, chronic wounds, and asthma due to its cytoprotective effects [334]. Besides inactivating procoagulant cofactors, APC can also remain bound to Figure 17: Biased signaling of PAR-1 Cleavage of PAR-1 at different sites by proteases result in biased downstream signaling pathways. APC signals through PAR-1 through proteolysis at a site different than other known PAR-1 activators including thrombin and FXa. Cleavage of PAR-1 by APC results in a unique gene expression profile that confers cytoprotective effects. 47 EPCR and induce intracellular signaling through PAR-1 [335]. Compared with thrombin cleavage of PAR-1 at R41 that results in a N-terminal tethered ligand with the sequence 42SFLLRNâ¦, APC cleaves at a different site five amino acids towards the C-terminal end at R46 to give rise to a distinct tethered ligand sequence of 47NPNDKYâ¦ and a different cellular signaling pattern (Figure 17) [336]. Whereas thrombin âbiasesâ signaling through G-proteins stimulating proinflammatory extracellular signal-regulated kinase 1/2 (ERK1/2) and RhoA pathways, APC signaling is biased towards the β-arrestin pathways through PI3K/protein kinase B (Akt) and Ras-related C3 botulinum toxin substrate 1 (Rac1) [336]. This results in stimulation of a unique gene expression profile that confers cytoprotection through functions that include anti-inflammatory and anti-apoptotic activity, stabilization of the vascular endothelial monolayer, and cellular regeneration [337-339]. The cytoprotective effects of APC in vivo have been well-documented in animal studies and human trials, particularly in the context of sepsis and stroke. PAR-1 signaling through endogenous APC was found to protect mice from lipopolysaccharide-induced septic shock [340]. Protection was also mediated when administered variants of APC with functional PAR-1 signaling but impaired anticoagulant activity [340-342]. However, human sepsis trials have largely failed to show a significant benefit on mortality with recombinant wild-type APC (Xigris®) and was withdrawn from the market in 2011 [343-345]. Patients that received recombinant APC in these trials had an elevated bleeding risk [344-346]. However, recombinant APC variants lacking anticoagulant activity have not yet been studied in human sepsis and could have more promising efficacy. Further advancements in engineering APC to be selective for its cytoprotective functions have shown promise as a neuroprotective agent in stroke [347]. Various clinical 48 correlates have suggested that higher circulating APC concentrations is associated with neuroprotection and improved outcomes in stroke patients [348-350]. Work done in vitro in brain microvascular endothelial cells have identified several anti-apoptotic pathways mediated by EPCR-bound APC cleavage of PAR-1 including reducing tumor protein p53 (p53), normalizing the Bcl-2-like protein 4 (Bax)/B-cell lymphoma 2 (Bcl-2) ratio, and decreasing caspase-3 signaling [351]. Wild-type APC was shown in vivo to protect against tPA-induced neurotoxicity and paradoxically also reduced hemorrhage [352]. Numerous engineered variants were designed that have greatly impaired anticoagulant function and work solely through PAR-1 signaling, including 3K3A-APC with Lys191-193 mutated to alanine (A; Ala) residues to disrupt procoagulant substrate recognition mediated by electrostatic interactions [353, 354]. 3K3A-APC has been successful in preliminary phase II clinical trials with the goal of being a neuroprotective adjuvant in ischemic stroke treatment [355]. 1.7.2.2 Plasmin modulation of protein C Proteolysis of PC by plasmin was first reported in the 1980s and only a second time in the 1990s, the first group led by Dr. Paul Bajaj [356] and the second by Dr. Hans Peter Schwarz [357]. Dr. Schwarz later went on to help develop recombinant wild-type APC for several landmark sepsis trials [358]. Data reported in these early findings showed proteolysis by plasmin resulting in a biphasic modulatory activity whereby cleavage of PC first resulted in transient partial APC-like activity and further cleavages quickly inactivated the enzyme [357, 359]. APC activity in these assays were quantified by chromogenic substrate cleavage and prolongation of plasma clotting times [357, 359]. However, these studies were done with plasmin concentrations that for the most part were in excess of 49 PC, due to the rationale of mimicking levels seen during thrombolysis treatment [357]. Inactivation of PC was theorized to be an explanation for recanalization in patients treated with thrombolysis [357]. Several in vivo reports suggested the opposite where substantial functional APC activity was detected following urokinase administration in baboons and in human patients after treatment with streptokinase [360, 361]. 1.7.3 Auxiliary fibrinolysis cofactors Recent work done by groups including ours at the University of British Columbia has challenged fibrinolysis dogma in fibrin being the only relevant physiologic cofactor for tPA and plasminogen to initiate and propagate fibrinolysis. It has been shown that plasmin cleaves and converts several plasma constituents localized to the clot site (i.e. FVa, FIXa, FX(a) into cofactors for tPA-mediated plasminogen activation [234, 235, 362, 363]. The heterotetrameric form of cellular annexin A2 also has intrinsic tPA-enhancing activity when complexed with S100A10 (p11) and may be induced by thrombin to the cell surface thus localizing it to the site of clot formation [364, 365]. This has been shown to involve a CTL-dependent mechanism intrinsic to p11 [366, 367]. Another cell surface plasminogen receptor, α-enolase, has been shown to bind through an internal lysine and enhances the activity of plasminogen activators [368]. The proposed mechanism of these âauxiliary fibrinolysis cofactorsâ generated through plasmin cleavage involves proteolysis exposing CTLs to help co-localize tPA and plasminogen on a scaffolding surface like aPL (Figure 18). These pathways would help generate plasmin during the initial stages of fibrinolysis to âprimeâ fibrin for dissolution. Due to its high concentration at the clot site, âprimedâ fibrin with exposed CTLs would then become the dominant fibrinolysis cofactor. Local plasmin concentrations at the clot site are greatly elevated during fibrinolysis and proteolytic 50 processing of its various substrates may play an important role in hemostasis. In fact, our lab has already shown that an engineered variant of FXa enhances thrombolysis in vivo while reducing systemic markers of bleeding [369]. This variant has a CTL tethered onto a polyethylene glycol linker that is covalently attached to the active site histidine of FXa (i.e. FXai-K) [369]. FXai-K lacks procoagulant function but retains the ability to be Figure 18: Auxiliary cofactor model of fibrinolysis The auxiliary cofactor model of fibrinolysis builds upon the recognized pathway where there are cofactors for tPA in addition to fibrin itself. These cofactors, including FVa, FIXa and FXa are modulated by plasmin to express CTLs that facilitate the enhancement of tPA-mediated plasminogen activation. This assists in âprimingâ fibrin for dissolution by expressing CTLs on fibrin. Fibrin, due to its high concentration at the clot, then takes over as the dominant fibrinolysis cofactor leading to a burst of plasmin. The specific orientations of binding are currently unknown. Moreover, it is not known whether FVa enhances tPA through a CTL-dependent mechanism. 51 modulated by plasmin into a tPA accelerator [369]. The advantage of these auxiliary fibrinolysis cofactors is that they are highly localized to the clot site through aPL-binding domains and could either enhance fibrinolysis without the requirement for tPA or be used as an adjunct therapy to reduce the therapeutic dose of tPA. This approach would work to localize thrombolysis to the clot site and reduce hemorrhage associated with intravenously administered tPA. 1.7.3.1 Vitamin K-dependent serine proteases FXa has been shown to acquire fibrinolytic activity when cleaved by plasmin on PtdSer-expressing surfaces [370]. Purified FXa exists as a mixture of FXaα and FXaβ due to autoproteolysis cleaving off a short ~3 kDa piece on the C-terminal end of the FXaα protease domain [371-373]. Both FXaα and FXaβ are catalytically active and have procoagulant function [373]. Plasmin also cleaves FXaα into FXaβ near the C-terminus of the protease domain at Lys435 (Figure 19) [370, 374]. Only when FXaα is aPL-bound, plasmin cleavage or autoproteolysis facilitates CTL expression on FXaβ, which enhances tPA cofactor activity [370]. When bound to aPL in the presence of Ca2+, a sequential cleavage occurs by plasmin within the peptidase domain at Lys330, yielding non-covalently associated fragments with approximate molecular weights of 33 and 13 kDa (i.e. FXa-33/13) [370]. Cleavage at Lys330 within the protease domain abolishes both amidolytic and coagulation activity [370]. Both plasmin-mediated generation of FXaβ and FXa-33/13 express CTLs and bind plasminogen, with the latter species expressing two CTLs [375]. FXaβ and FXa-33/13 both enhance tPA-mediated plasminogen activation and fibrinolysis at low picomolar tPA levels in vitro [370]. The additional CTL expressed on FXa33/13 is thought to facilitate a greater enhancement relative to FXaβ. In plasma, however, neither 52 FXaβ nor FXa-33/13 enhance fibrinolysis, and data showed that these fragments were inactivated rapidly by a currently unknown mechanism liberating a 28 kDa fragment [370]. Profibrinolytic FXa-derived fragments enhance fibrinolysis, where these tPA cofactors help âprimeâ fibrin for dissolution. A variant of FXa (i.e. FXai-K) was developed to overcome inhibition of these fragments seen in plasma [369]. Specifically, a CTL-containing tetraethylene glycol linker is reacted with a chloromethyl ketone-containing compound and covalently linked to the active site histidine of FXa [369]. This modification stops cleavage at FXaβ and prevents generation of FXa-33/13, a mechanism hypothesized to be due to allosteric or steric hinderance impairing cleavage by plasmin at Lys330 [369]. FXai-K has been shown to enhance plasma fibrinolysis in vitro but also enhance clot dissolution in a murine arterial carotid thrombosis model [369]. Interestingly, enhancement of clot dissolution was seen without the requirement of tPA with an even greater enhancement in efficacy when paired with a sub-therapeutic dose of tenecteplase [369]. Moreover, mice injected with FXai-K did not have elevated systemic markers of fibrinolysis when compared with therapeutic tenecteplase, suggesting a localized thrombolytic effect of FXai-K [369]. Dr. Paul Bajaj and colleagues at the University of California Los Angeles recently published that FIXa behaves similarly to FXa in vitro, where analogous cleavages by plasmin converts FIXa into a fibrinolysis accelerator [234]. FVIIa has been briefly studied in our lab and does not enhance tPA cofactor activity (data not published). Interestingly, sequence comparisons of FVII(a), FIX(a) and FX(a) show the presence of homologous arginines on FVII(a) instead of lysines at both sites near the C-terminus and within the protease domain. The analogous serine protease (A)PC has not yet been studied in this 53 context. Why plasmin would selectively cleave certain homologous haemostatic proteases over others to enhance tPA-mediated fibrinolysis warrants further investigation. Figure 19: Conversion of FIXa and FXa to tPA cofactors by plasmin FIXa and FXa are both converted into tPA cofactors by a similar mechanism involving plasmin cleavage at two sites within the protease domain to liberate new CTLs. These CTLs interact and enhance tPA-mediated plasminogen activation. 1.7.3.2 Factor V 1.7.3.2.1 Synthesis FV is a large, ~330 kDa single-chain protein that plays a central role in blood coagulation as a cofactor for FXa to greatly accelerate prothrombin activation [376, 377]. The gene for FV is located on the long arm of chromosome 1q21-25 and spans ~80 54 kilobases (kb) containing 25 exons. It is transcribed into a ~7 kb messenger RNA (mRNA), where 6675 nucleotides code for a 2224-amino acid protein [377]. FV is synthesized predominantly in the liver and undergoes extensive posttranslational modifications including glycosylation, sulfation, and phosphorylation [377]. FV produced in the liver is secreted and circulates as a 2196-amino acid protein in plasma at a concentration of ~20 nmol/L with a half-life of 12-36 hours [378]. About 20% of total FV in whole blood is located within the α-granules of platelets and is released upon platelet activation [379]. The origins of platelet factor V are from megakaryocyte synthesis and endocytosis of plasma FV [380]. The FV molecule is comprised of triplicate A-domains, a B-domain, and duplicate C-domains organized in the order A1-A2-B-A3-C1-C2 (Figure 20) [381]. Figure 20: FV structure The structure of FV has been illustrated to size by amino acid residue number with disulfide bonds and posttranslational modifications. Posttranslational modifications include N-linked glycosylation (green), the partially glycosylated Asn2181 (light green), phosphorylation (red and black), and sulfation (yellow). The basic and acidic regions within the B-domain have been highlighted, where their interaction together is known to keep FV in its inactivated procofactor form. 1.7.3.2.2 Activation of FV FV is kept stable in its procofactor state in circulation through its carbohydrate-rich B-domain [135]. Proteolytic sites within (Arg1018) and flanked (Arg709/1545) on either end of the B-domain are important in the expression of FV cofactor activity (Figure 21) [381]. 55 Cleavage of Arg709 and Arg1018 are kinetically favoured by its main physiologic activator thrombin [381]. This results in partial cofactor activity, where an additional cleavage at Arg1545 is required for full activation [381]. FXa has also been shown to cleave at Arg709 and Arg1018 to induce partial cofactor activity with a semi-truncated B-domain [382]. This finding has challenged dogma to where it is believed FXa, instead of thrombin, is the physiologic initiator of FV activation [382]. To support this, Russellâs Viper Venom (RVV) activates FX to FXa and is also able to fully activate FV by cleaving strictly at Arg1545 [383]. Full activation of FV truncates and releases the entire B-domain, where FVa then binds to FXa on PtdSer-expressing surfaces to amplify thrombin generation as an enzyme cofactor. It is thought that proteolysis and release of the B-domain removes steric hinderances that interfere with binding interactions and/or induces a conformational change that facilitates cofactor function [384]. Truncation of the B-domain converts single-Figure 21: Cleavages that activate FV and inactivate FVa Cleavage sites that play an important physiologic role in the activation of FV to FVa are shown. In addition, proteolytic cleavages that inactivate FVa are indicated. 56 chain FV to two-chain FVa, with the heavy chain (FVa-Hc; A1-A2) held together to the light chain (FVa-Lc A3-C1-C2) by the divalent ions Ca2+ and copper (Cu2+) [385, 386]. 1.7.3.2.3 Role of the FV B-domain The B-domain of FV spans amino acids 710-1545 and is heavily glycosylated with at least 15 N-linked asparagine (Asn; N) residues [381]. The B-domain is not conserved across and within the same species as there is little homology between the FV and FVIII B-domains [381]. Its primary mechanism is to maintain FV in its inactive procofactor form prior to proteolytic activation and release [135]. This is facilitated by a basic region (BR; residues 963-1008) and an acidic region near the C-terminal end of the B-domain (AR; residues 1493-1537) that interact and work in unison to maintain FV as a procofactor [135]. This is supported by evidence showing that engineering of the B-domain to strictly contain only the BR and AR was sufficient to keep FV in its inactive state [135]. Moreover, FV variants with shortened B-domains have shown to exhibit cofactor function and the ability to assemble in prothrombinase [135]. One of these variants developed has residues 811-1491 deleted (i.e. FV-810) and is studied in this thesis paper. FV-short, a splice isoform that lacks BR, interacts with a region in TFPIα that resembles AR and is a carrier for TFPIα in the circulation [216, 387, 388]. There are also engineered forms of FV that only contain AR (i.e. FV-AR) that have been shown to be protected from FXa binding with use of BR fragments, suggesting yet another mechanism for the BR-AR association in preventing interactions with prothrombinase components [389]. 1.7.3.2.4 FVa structure-function relationships There is an extensive literature correlating specific structural features of FVa to its biological functions. The FVa A-domains share significant homology with the copper-57 binding protein ceruloplasmin (CP) [390]. Like CP, the three A-domains are arranged in a triangular fashion, with each A-domain consisting of two cupredoxin-like folds [390]. The organization of the triplicate A-domains remain similar following activation into FVa [390]. There are single Ca2+ and Cu2+ binding sites between the A1 and A3 domains that are thought to stabilize the interaction between FVa-Hc and FVa-Lc [385, 390]. Binding of these two divalent metal ions is crucial to expression of cofactor activity [391]. A substitution of Asp111 to Ala111 on the A1 domain was shown to disrupt both Ca2+ and Cu2+ binding and result in dissociation of FVa-Hc from FVa-Lc [392, 393]. The two APC cleavage sites, R306 and R506, both located within the A2-domain of FVa are solvent exposed [390]. These regions have also been implicated in the recognition for FXa and prothrombin through mutagenesis and de-glycosylation studies [394]. Interactions with FXa and prothrombin at regions either containing or nearby R306 and R506 protect FVa from proteolysis by APC [395]. Less is known about the A3-domain of FVa, but structural modeling and mutagenesis studies have implicated a role in prothrombinase assembly through interactions with the protease and EGF domains of FXa [396]. The C-domains of FVa are homologous to the slime mold protein discoidin and are critical for binding to anionic phospholipid (aPL). Two isoforms of FVa and its precursor exist due to partial glycosylation of a 3 kDa polysaccharide chain at Asn2181 [397-399]. Glycosylation at Asn2181 indirectly interferes with lipid binding [397]. However, once assembled in prothrombinase, both the glycosylated (i.e. FVa1) and non-glycosylated (i.e. FVa2) forms have comparable catalytic efficiency [400]. Unlike Gla-containing serine proteases, FVa does not require Ca2+ to bind to PtdSer-expressing surfaces [397]. Instead, binding is facilitated by both C1 and C2 domains where hydrophobic protrusions 58 or âspikesâ insert into the lipid bilayer [401, 402]. It is thought that prior to lipid binding, FVa exists in a âclosedâ conformation with only a limited number of hydrophobic protrusions largely on the C2-domain [403]. Recognition of PtdSer then triggers FVa to adopt an âopenâ conformation and further expression of hydrophobic lipid-binding residues on both C2 and C1 [403]. 1.7.3.2.5 Cleavage of FVa by plasmin Besides APC, plasmin has been shown to cleave, transiently activate, and then inactivate FV in vitro [404]. The importance of this mechanism has been substantiated by plasmin-mediated proteolysis detected in patients undergoing thrombolysis treatment [405]. Therefore, in certain scenarios where plasmin concentrations may be elevated, there may be an important link between fibrinolysis and coagulation, whereby plasmin either regulates or dysregulates hemostasis through cleavage of FV(a) [406]. Work from our laboratory has suggested a modulatory role for plasmin cleavage of FVa where Figure 22: Major plasmin cleavage sites within FVa Major sites of plasmin cleavage of FVa are illustrated here. Following thrombin activation, the A1 and A3 domains are thought to be non-covalently associated via Ca2+ and Cu2+ binding. Several of these fragments liberate CTLs, which may play roles in enhancing tPA-mediated plasminogen activation. 59 proteolysis converts FVa from a procoagulant to fibrinolytic cofactor [362]. The fragment(s) and cleavage conditions responsible for the development of tPA cofactor activity following cleavage by plasmin is unknown. Plasmin cleaves aPL-bound FVa into 16 fragments derived from both the FVa-Hc and FVa-Lc [236]. A number of these cleavages facilitate the exposure of a CTL and may be an important mechanism in mediating the generation of tPA cofactor activity (Figure 22). Whether FVa fragments can localize and accelerate tPA-mediated activation of plasminogen to enhance fibrinolysis is not known. 60 Chapter 2: Hypothesis and goals 2.1 Hypothesis The overarching hypothesis of this dissertation is that plasmin has an important physiologic role in hemostasis by regulating PC and FVa. During thrombolysis therapy where plasmin concentrations are known to be elevated, the effect of plasmin on these two substrates are likely amplified and may dysregulate hemostasis. Based on homology to FIX and FX, it is hypothesized that plasmin proteolyzes (A)PC, causing it to lose anticoagulant activity while being converted into a fibrinolytic accelerator through the exposure of CTLs. It is further hypothesized that fragments of FVa generated after proteolysis by plasmin accelerate fibrinolysis through CTL exposure. These modulatory effects are teleologically reasonable, as plasmin would convert local haemostatic constituents into fibrinolysis accelerators. Since (A)PC and FV(a) play central roles in hemostasis, understanding the functional sequelae following their proteolysis by plasmin is of great importance in human health and disease. 2.2 Goal To further our understanding of the mechanisms by which fibrinolytic plasmin regulates the haemostatic functions of anticoagulant PC and procoagulant FVa. 2.3 Objectives 2.3.1 To determine the effect on anticoagulant and cytoprotective functions of PC following proteolysis by plasmin. 1. Identify cleavage sites and the fragmentation pattern by which plasmin proteolyzes PC through N-terminal sequencing of PC-derived fragments. 2. Correlate the fragmentation of PC by plasmin over time to APC amidolytic activity. 61 3. Discern the effect of plasmin-proteolyzed PC on the ability to inactivate its well-known anticoagulant substrates and prolong clotting times in plasma. 4. Assess if PAR-1 signaling is impaired following proteolysis of PC by plasmin. 5. Determine whether plasmin converts PC into a fibrinolytic cofactor through the exposure of CTLs like other Gla-containing serine proteases. 2.3.2 To dissect the mechanism by which plasmin proteolytically modulates FVa from a procoagulant to fibrinolytic cofactor. 1. Elucidate the proteolytic conditions by which plasmin converts FVa into a cofactor for tPA-mediated plasminogen activation. 2. Identify the specific fragments derived from proteolysis of FVa by plasmin that are responsible for accelerating tPA activity. 3. Determine whether tPA-accelerating fragments derived from plasmin-mediated proteolysis of FVa can enhance fibrinolysis. 62 Chapter 3: Materials and methods 3.1 Materials 3.1.1 Proteins and plasmas Purified human FVa, FXa, (A)PC, α-thrombin, Lys-plasminogen, and plasmin were obtained from Haematologic Technologies and were >95% pure as measured by SDS-PAGE (Essex Junction, VT, USA).Purified fibrinogen depleted of plasminogen, vWF and fibronectin was purchased from Enzyme Research Laboratories (South Bend, IN, USA). Recombinant FV truncated at residues 811-1491 (i.e. FV-810) was generated from a transfected baby hamster kidney (BHK) cell line and purified in-house as previously described [385, 407, 408]. Recombinant single-chain tPA was purchased commercially from Genetech (San Francisco, CA, USA). Anti-human factor V antibodies AHV-5112 (monoclonal to C1/C2 in FV-Lc) and AHV-5146 (polyclonal recognizing FV-Hc) were purchased from Haematologic Technologies (Essex Junction, VT, USA). An anti-human factor V light chain monoclonal antibody AHV-14H12 (monoclonal recognizing A3 in FV-Lc) was generated in-house. A secondary goat-anti mouse immunoglobulin G (IgG) heavy and light chains (H&L) horseradish peroxidase (HRP) antibody (Code 115-035-146) was purchased from Jackson ImmunoResearch Laboratories (West Grove, PA, USA). Normal pooled plasma, FV-deficient and PC-deficient plasma were purchased from Affinity Biologicals Inc. (Hamilton, ON, Canada). 3.1.2 Substrates and inhibitors Chromogenic substrates H-D-Ile-Pro-Arg-para(p)-nitroalinine(pNA) (S-2288), H-D-Glu-Pro-Arg-pNA (S-2366), H-D-Val-Leu-Lys-pNA (S-2251) were from Chromogenix 63 (Bedford, MA, USA). 4-amidinophenylmetanesulfonyl fluoride hydrochloride (p-APMSF) and aprotinin were purchased from Sigma-Aldrich (St. Louis, MO, USA). 3.1.3 Additional reagents 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), polyethylene glycol 8000 (PEG 8000), N-cyclohexyl-3-aminopropanesulfonic acid (CAPS), benzamidine hydrochloride, 2-mercaptoethanol (β-ME), ε-aminocaproic acid (EACA), potassium hexocyanoferrate(III), silver nitrate, anhydrous dimethyl sulfoxide (DMSO), 1,4-dithiothreitol (DTT), Dulbeccoâs Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F12), fetal bovine serum (FBS), L-glutamine, penicillin-streptomycin (Pen-Strep), Geneticin® (G418), DMEM/F12 50/50 1x without phenol red, insulin-transferrin-selenium (ITS), and AlbuMAXTM were purchased from Sigma-Aldrich (St. Louis, MO, USA). Sodium hydroxide (NaOH), hydrochloric acid (HCl), sodium chloride (NaCl), sodium thiosulfate, sodium dodecyl sulfate (SDS), sodium carbonate (Na2CO3), powdered glycine, tetrasodium ethylenediaminetetraacetic acid (EDTA), PageBlueTM protein staining solution, bromophenol blue, Coomassie brilliant blue R-250 dye, Coomassie brilliant blue G-250 dye, tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl), activated partial thromboplastin time (APTT) reagent and all alcohols were obtained from Thermo Fisher Scientific (Waltham, MA, USA). Calcium chloride dihydrate (EMD Chemicals, Inc. New Jersey, USA). Immobilon-P polyvinylidene difluoride (PVDF) was purchased from EMD Millipore (Burlington, MA, USA). Tween-20 was purchased from Amresco (Cleveland, OH, USA). Carboxypeptidase inhibitor 2-guanidinoethylmercaptosuccinic acid (GEMSA) was obtained from VWR (Mississauga, ON, Canada). L-α-phosphatidylcholine (PtdCho; egg yolk) and L-α-phosphatidylserine (PtdSer; bovine brain) were purchased from Avanti 64 Polar Lipids (Alabaster, AL, USA). 125I was purchased from PerkinElmer (Waltham, MA, US). BLUeye prestained protein ladder was obtained from FroggaBio (Toronto, ON, CA). Innovin® was purchased from Dade Behring (Mississauga, ON, CA). 3.2 Anionic phospholipid preparation 3.2.1 Small unilamellar vesicles (SUVs) Small unilamellar vesicles (SUVs) were prepared with a 75% PtdCho and 25% PtdSer composition on a per molar basis. This protocol was adapted from previously established methods using sonication and differential centrifugation to generate unilamellar phospholipid vesicles of homogenous size [356, 409]. First, a mixture of PtdCho and PtdSer pre-dissolved in chloroform was dried under nitrogen gas and then resuspended in HBS (20 mM HEPES, 150 mM NaCl, pH 7.4). The resuspended lipid was subjected to direct sonication on ice (Misonix microtip #419 â 1/8â, Sonicator S-4000). Sonicated vesicles were transferred to a 12 mL ultracentrifuge tube and spun in a Beckman SW-41 Ti swinging bucket rotor at 30,000 x g for 3.5 hours at 20°C. Directly following centrifugation, the upper 65% of the tube was collected and stored under argon gas at 4°C to prevent oxidation. Final SUV concentrations were determined using an assay that subjects lipid to hydrolysis with phospholipase D to release choline followed by subsequent oxidation steps generating a blue pigment measurable by absorbance at 600 nm (Wako Phospholipids C Kit; Mountain View, CA, US). A standard curve using a known concentration of choline chloride was used to derive the SUV concentration accounting for a 75% PtdCho to 25% PtdSer mixture. SUV preparations were sized by light scattering methods and were on average in the approximate diameter range of 30 ± 65 10 nm (Beckman Coulter N4 PLUS Particle Size Analyzer/Malvern Panalytical ZetaSizer Nano ZS). Lipid preparations were used within 2 months. 3.2.2 Large multilamellar vesicles (LMVs) Large multilamellar vesicles (LMVs) containing a molar ratio of 75% PtdCho and 25% PtdSer were prepared by size extrusion. The chloroform-containing lipid mixture was dried under nitrogen gas and resuspended in sucrose-containing HBS (300 mM sucrose, 20 mM HEPES, 150 mM NaCl, pH 7.4) buffer. Lipids were then subjected to ten freeze-thaw cycles, first in liquid nitrogen for 30 seconds followed by 50-70°C water for two minutes. This process facilitates more uniform vesicle formation during extrusion. The lipid mixture was then extruded using an Avanti® Mini Extruder system from Sigma-Aldrich (St. Louis, MO, USA) with a 1 µm nanopore filter membrane. Following extrusion, the sample was then diluted by one volume worth and pelleted at 14,000 x g in a microfuge for 5 minutes. Pelleted sucrose-containing vesicles were gently resuspended in HBS. Vesicles were quantified, sized and stored as previously described. 3.3 Recombinant FV-810 generation 3.3.1 Expression BHK cells expressing a FV variant with a truncated B-domain missing residues 811-1491 (i.e. FV810) were a generous gift from Dr. Rodney Camire (University of Pennsylvania) (Figure 23). These stably expressing cells were cultured and FV-810 was successfully purified via ion exchange chromatography. Briefly, BHK cells expressing FV-810 were grown to confluence in selection media containing (per 500 mL) DMEM/F12, 10% FBS, 5 mL L-glutamine, 5 mL Pen-Strep, 2 mM CaCl2 and 450 µg/uL G418. Cells were scaled to triple flasks and switched to expression media containing (per 500 mL) 66 DMEM/F12 50/50 1x without phenol red, 0.5 mL ITS, 5 mL L-glutamine, 5 mL Pen-Strep, 5 mL AlbuMAXâ¢, 2 mM CaCl2 and 100 µg/mL of G418. Media was collected each morning for 5 consecutive days. 10 mM benzamidine and 100 µM p-APMSF were added to the media following collection as broad-spectrum protease inhibitors. Collected media was run immediately on a SP Sepharose® Fast Flow column. Figure 23: Structure of FV-810 Compared with native FV, FV-810 has a significantly truncated B-domain and an approximate molecular weight of 216 kDa. While the acidic region is retained, the lack of a complementary basic region allows FV-810 to exhibit procoagulant cofactor-like activity without the requirement for thrombin cleavage [135]. FV-810 is still cleavable by thrombin at the canonical thrombin cleavage sites R709 and R1545 [135]. 3.3.2 Purification A 30 mL SP Sepharose® Fast Flow chromatography column was equilibrated with buffer containing 20 mM HEPES, 0.15 M NaCl, 5 mM CaCl2, and 1 mM benzamidine at pH 7.4. Media containing FV-810 was loaded onto the column and ran at a flow rate of 10 mL/min. Another wash was done and then protein was eluted with high salt (20mM 67 HEPES, 0.65 M NaCl, 5 mM CaCl2, 1 mM benzamidine, pH 7.4) and collected in 10 mL fractions. Fractions containing FV activity as measured by a partial thromboplastin (PT) clotting time in FV-deficient plasma were pooled together. Approximately 2-4 mgs of protein were collected each day as measured by activity to a standard curve of natural FV. Fractions were stored in -80°C. These steps were repeated for four consecutive days. Figure 24: Protein stain and thrombin cleavage of recombinant FV-810 1 µg of purified recombinant FV-810 was loaded under reducing conditions into a 5-10% polyacrylamide gradient gel. 1 µM of FV-810 was cleaved by 0.1 µM of thrombin and similarly run via SDS-PAGE. FV-810 was thrombin-cleavable into FVa. 68 10 mL fractions obtained from the SP Sepharose® Fast Flow column were thawed, pooled and diluted with five volumes worth of buffer (20mM Hepes, 5 mM CaCl2, pH 7.4) to reduce the salt concentration. 100 µM p-APMSF was added to the mixture. Protein was loaded onto a 15 mL Q Sepharose® Fast Flow column at a flow rate of 5 mL/min. The column was first washed as before prior to loading and then eluted at 5 mL/min with a salt gradient (20 mM HEPES, 1 M NaCl, 5 mM CaCl2, pH 7.4). 2 mL fractions were collected and points during the peak were collected and assayed for FV clotting activity. Samples were taken across the peak and ran on 5-15% gels to determine purity (>95%) by protein staining with Coomassie blue stain (Figure 24). Fractions with FV clotting activity and high purity were pooled in a 30K molecular weight cut off (MWCO) dialysis bag and dialyzed in buffer (20 mM HEPES, 150 mM NaCl, 5 mM CaCl2, pH 7.4) for two three-hour periods. The sample was then concentrated using an Amicon® Ultra 100K MWCO Centrifugal Filter. Protein concentrations were measured by absorbance at 280 nm, aliquoted, and stored at -80°C. 3.3.3 Labelling Lys-plasminogen with 125I 150 µg of Lys-plasminogen stored in 50% glycerol was buffer exchanged with HBS to dilute out the glycerol. Due to residual plasmin activity in the commercially purchased Lys-plasminogen samples as measured by S-2251 substrate cleavage, 100 µM p-APMSF was added prior to labeling and allowed to progress through 10 hydrolysis half-lives (T1/2 = 6 minutes; please refer to Sigma-Aldrich product sheet). p-APMSF is a small molecule inhibitor of plasmin that undergoes rapid hydrolysis in aqueous buffer. Lys-plasminogen was then combined with 125I in glass test tubes pre-coated with Pierceâ¢ Iodination Reagent [370, 410]. This reagent utilizes an oxidative Iodogen® reagent that forms a 69 highly reactive intermediate with 125I and adds iodine atoms to tyrosine (Tyr, Y) and His side chains. The sample was then centrifuged at 11,000 rpm for 20 minutes at 4°C in a 10 kDa cutoff Micron tube to remove free 125I. This was repeated four times with 1 µL of diluted free 125I and 125I-labelled Lys-plasminogen saved for trichloroacetic acid (TCA) precipitation to determine the relative percentage of free 125I versus bound using a scintillation counter. A bicinchoninic acid (BCA) assay was used to determine the concentration of 125I-labelled Lys-plasminogen. Protein was aliquoted and stored at -80°C and used within two months, as 125I has a half-life of 52 days. A maximum of 180,000 counts per minute (CPM)/µg protein was achieved, which had no effect on function as measured by tPA activation compared to unlabeled Lys-plasminogen. 3.4 Preparing plasmin-cleaved samples for functional assays 3.4.1 FVa and PC To study the effects of plasmin in modulating the biochemistry of FVa and PC, samples were prepared where FVa and PC were treated with plasmin under specific conditions prior to functional assessment. FVa (1 μM) was incubated with plasmin (20 nM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) for up to 1 hour at room temperature (RT). Similarly, plasmin (100 nM) was used to cleave PC (1 μM) in a mixture containing SUVs (50 μM) and CaCl2 (2 mM) for up to 1 hour at RT. Higher concentrations of native FVa and PC (>1 μM) were used with the same enzyme to substrate ratios when higher amounts of cleaved FVa or PC were required. Plasmin cleavage of FVa under these conditions resulted in complete fragmentation of the FVa-Hc and FVa-Lc chains by 1 hour by SDS-PAGE and Coomassie blue staining. For (A)PC, cleavage by plasmin under the same conditions resulted in the disappearance of PC-Hc as seen by silver 70 staining. Silver staining was selectively used to analyze fragmentation of (A)PC on polyacrylamide gels as Coomassie blue G-250 stain was not sensitive enough to detect (A)PC-derived fragments even when ~1 μg of total protein was loaded per lane. 3.4.2 APC-inactivated FVa Plasmin-treated PC and APC were used to cleave both FVa and plasmin-treated FVa (Pn-FVa) to assess for fragmentation by protein staining. A preparation of plasmin-cleaved PC (0.1 μM) or intact APC (0.1 μM) was incubated with FVa (1 μM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at room temperature. Cleavage of FVa was assessed by fragmentation of FVa on a Coomassie-stained 5-15% polyacrylamide gradient gel. For preparations of APC-cleaved FVa that was subjected to another round of cleavage by plasmin, FVa (1.5 μM) was cleaved with APC (0.15 μM) in the presence of SUVs (50 μM) and CaCl2 (2 mM). 1 μM APC-cleaved FVa was then incubated with 0.1 μM plasmin. This fragment mixture was compared by protein staining and chromogenic activity with plasmin-proteolyzed FVa prepared as described above. 3.4.3 Aprotinin or p-APMSF inhibition of residual plasmin Plasmin used to proteolyze either FVa or (A)PC into fragments for experimental analysis required inhibition to prevent interference in functional assays by conducting further unwanted cleavages. Either p-APMSF or aprotinin was used to inhibit residual plasmin after being utilized for proteolysis of FVa or (A)PC. The plasmin that was used to cleave FVa was inhibited with p-APMSF. Complete inhibition of plasmin required a 1000x excess in molar concentration of p-APMSF (Figure 25). Samples requiring p-APMSF inhibition were kept at room temperature for at least 1.5 hours prior to initiating functional assays to allow p-APMSF to completely hydrolyze (>99.99%), rendering it non-functional. 71 Aprotinin is a small peptide inhibitor that targets the active site of trypsin-like serine proteases (e.g. trypsin, plasmin, kallikrein) but has far weaker affinity for APC (Ki = 1100 nM) relative to plasmin (Ki = 0.07 nM) [411-413]. Since plasmin cleavage of PC generated APC-like activity, aprotinin was used instead of p-APMSF to inhibit plasmin when proteolyzing (A)PC. Inhibition of residual plasmin at a 2.5:1 molar ratio completely inhibited plasmin activity and had negligible impact on the enzymatic activity of APC (Figure 26). Figure 25: Inhibition of plasmin activity by p-APMSF 0.1 µM of plasmin was incubated with a titration of p-APMSF in HBS. 100 µM p-APMSF resulted in maximal inhibition of plasmin activity as measured by S-2251 (200 µM) substrate cleavage, indicating that concentrations of at least 100 µM of p-APMSF or a 1000x excess of p-APMSF by concentration was required for full inhibition. Mean absorbance values with standard deviation (SD) are shown from a single experiment. 72 Figure 26: Inhibition of plasmin activity by aprotinin 0.1 µM of plasmin was incubated with a titration of aprotinin in HBS. Concentrations above 250 nM aprotinin resulted in maximal inhibition of plasmin activity as measured by S-2251 (200 µM) substrate cleavage, indicating that concentrations of at least 250 nM of aprotinin or a 2.5x molar excess of aprotinin was required for full inhibition. Mean absorbance values with standard deviation (SD) are shown. 3.4.4 Separation of lipid-binding fragments with LMVs Lipid-binding fragments of FVa were separated from unbound fragments for functional analyses. Fragments of FVa were pulled down on 10% sucrose-filled LMVs by centrifugation. FVa (1 μM) was incubated with plasmin (20 nM) in CaCl2 (2 mM) and the presence or absence of LMVs (2 mM) at room temperature. After incubation for 1 hour, the mixture was spun at 14,000 x g in a microfuge. The supernatant fraction was collected, and the pelleted fraction was washed three times in HBS containing EDTA (20 mM). The pellet was re-suspended in buffer and both supernatant and pellet fractions were assessed for separation by both protein staining and Western blotting. 73 3.5 Protein staining (Coomassie blue and silver staining) At least 1 μg of FVa or FVa-derived fragments were reduced with 80 mM DTT, denatured in boiled sample buffer (62.5 mM Tris-Hcl, 2.5% SDS, 0.002% bromophenol blue, 10% glycerol, pH 6.8) and separated electrophoretically on either a 10% or 5-15% (gradient) polyacrylamide gel. The gels were heated and stained with Coomassie blue G-250 for 1 hour. Stained gels were washed with several exchanges of ddH2O for at least 2 hours. Coomassie stain was not sensitive to sufficiently detect a 1-2 μg preparation of (A)PC-derived fragments for reasons still unknown, so silver staining was used instead. (A)PC and (A)PC-derived fragments were reduced with 80 mM DTT, denatured in boiled sample buffer, and separated electrophoretically on 15% polyacrylamide gels. Silver staining reagents were prepared in-house. Briefly, gels were separated on 15% gels and silver stained to increase the detection sensitivity. Following SDS-PAGE, gels were fixed in an alcohol solution (50% methanol and 5% acetic acid) for at least 30 minutes and then sensitized with a mixture containing 30 mM potassium hexocyanoferrate(III) and 100 mM sodium thiosulfate for 5 minutes. The gels were then incubated in 0.1% silver nitrate for 15 minutes and developed with 0.04% formaldehyde and 2% sodium carbonate. 3.6 Western blotting FVa-derived fragments transferred onto PVDF membranes were probed with different antibodies that recognize unique epitopes to assess for FVa fragmentation via Western blotting. A polyclonal antibody (AHV-5146) was used to detect FVa heavy chain (FVa-Hc) fragments and two monoclonal antibodies (AHV-14H12, AHV-5112) were used to probe for FVa light chain (FVa-Lc). These anti-FV antibodies are anti-human and derived from mouse. A secondary goat, anti-mouse IgG H&L HRP antibody was used to 74 detect fragments using ECL-prime chemiluminescent reagent from GE Healthcare (Chicago, IL, USA) and visualized using a Chemigenius2 analysis system from PerkinElmer (Waltham, MA, USA). Following exposure, the PVDF membrane was stained with Coomassie blue R-250 for 1 minute and destained with several changes of a solution containing 50% methanol and 10% acetic acid. 3.7 Assessment of fibrinolytic function 3.7.1 Purified tPA-mediated plasminogen activation A purified chromogenic assay was developed to measure the effect of (A)PC, (A)PC-derived fragments, FVa, FVa-derived fragments, and FV-810 on the enhancement of tPA-mediated plasminogen activation. The tPA cofactor to be evaluated (0.1 μM) was incubated with tPA (10 nM or 1 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM). Samples were taken for up to 1 hour and assayed for plasmin activity by S-2251 chromogenic substrate (200 μM) on half-area 96-well microtitre plates. Samples were assessed for plasmin content by measuring absorbance rates (mOD/min) at 405 nm with a SpectraMax Plus 384 microplate reader, which were subsequently converted to molar concentrations using a purified plasmin standard curve. Liberated pNA from the chromogenic substrates utilized in this dissertation absorbs light at 405 nm. The utilization of S-2251 as a substrate was specific for plasmin activity given the experimental conditions with lower relative concentrations of tPA. 3.7.2 One-step fibrinolysis assay using purified proteins A fibrinolysis assay using purified proteins was developed to measure the effect of FV, FVa, and FV-810 on the enhancement of fibrin dissolution. Purified fibrin clots containing cofactor (0.1 μM), fibrinogen (3 μM), tPA (10 pM), Pgn (0.5 μM), PCPS (50 75 μM) and CaCl2 (5 mM) were formed using thrombin (10 nM). Thrombin, tPA, and cofactor were added as separate droplets to a 96-well flat bottom plate. A rapid initial increase and eventual decrease in fibrin clot opacity were followed over time by light scattering at 405 nm as fibrin was formed and then solubilized (Figure 27). Readings were taken using a SpectraMax Plus 384 microplate reader every 5 minutes with shaking prior to each read. Figure 27: Clot formation and dissolution during fibrinolysis Shown is a representative profile of fibrin formation and dissolution in samples studied using a fibrinolysis assay. A high concentration of thrombin rapidly initiates clot formation, following by early plasmin âprimingâ the fibrin for subsequent rapid dissolution. Auxiliary fibrinolysis cofactors would theoretically shorten the priming phase and accelerate purified fibrin dissolution. 3.8 Assessment of APC activity 3.8.1 Chromogenic substrate cleavage Plasma-derived human (A)PC (1 μM) was incubated with plasmin (0.1 μM) in the presence of PCPS (50 μM) and CaCl2 (2 mM) at 25°C. Samples were drawn at selected intervals over 1 hour for measurement of APC-like activity by either S-2366 or S-2288 chromogenic substrate cleavage (200 μM). Samples were also taken in parallel to analyze 76 fragmentation over time by electrophoretic separation on a 15% polyacrylamide gel under reducing conditions. Gels were then silver stained. The utilization of S-2366 or S-2288 was specific to APC activity given the experimental conditions. 3.8.2 Activated partial thromboplastin time (APTT) in PC-deficient plasma The effect of plasmin-proteolyzed PC on prolongation of clotting times compared with APC was studied. PC (1 μM) was treated with plasmin (0.1 μM) for 5, 15, and 60 minutes in the presence of SUVs (50 μM) and CaCl2 (2 mM) at 25°C. Samples were then treated with aprotinin (0.25 μM) to inhibit residual plasmin while leaving functional APC amidolytic activity intact as assessed by S-2288 substrate cleavage. Samples containing non-cleaved PC and APC were also treated with aprotinin as controls. 100 μL of PC-deficient plasma was thawed, combined with 50 μL of APTT reagent and incubated for 5 minutes at 37°C. 50 μL of each sample was then added (50 nM final PC concentration) and APTT was measured using a coagulation analyzer from Sigma (St. Louis, MO, USA). 3.9 125I-radiolabeled plasminogen binding and dependency on C-terminal lysines FVa fragments were prepared as previously described to assess for direct binding to 125I-radiolabelled Lys-plasminogen (i.e. 125I-plasminogen). Protein was reduced and transferred onto PVDF membranes and were blocked overnight at 4°C with 10 mg/mL bovine serum albumin (BSA) and 50 nM GEMSA in Tris-buffered saline (TBS; 0.05 M Tris-HCl, 150 mM NaCl, pH 8.0). Blocking membranes with BSA served also as a negative control for non-specific binding. PVDF membranes were then overlaid with 125I-plasminogen (1 μM) in the presence or absence of EACA, an inhibitor of CTL-binding. Membranes were then triple washed with TBS and placed between overhead projector transparencies. Using an x-ray film cassette and a Dupont Quanta III intensifying screen 77 (Sigma-Aldrich; St. Louis, MO, USA), the PVDF was exposed to clear blue x-ray film (Thermo Fisher Scientific; Waltham, MA, USA) at -80°C and developed for at least one full day. 3.10 Edman degradation N-terminal sequencing Edman degradation was used to sequence the N-terminus of major fragments derived from (A)PC following proteolysis by plasmin. 2 µg of intact and fragmented (A)PC was reduced with 5% β-ME and separated electrophoretically on a 15% polyacrylamide gel. Fragments were transferred to PVDF and stained with Coomassie blue R-250. Fragments were overlaid with identical samples subjected to silver staining. Individual bands were carefully cut out of PVDF and sent for sequencing at the Tufts University Core Facility (Boston, MA, USA). This method involves use of phenyl isothiocyanate (Edmanâs reagent) under alkaline conditions to label the N-terminal amino acid of a peptide of interest [414]. The N-terminal amino acid is then cleaved under acidic conditions and extracted into an organic solvent to form a more stable phenylthiohydantoin (PTH)- amino acid derivative that can be identified by high-performance liquid chromatography [414]. Retention times for the known standard and the called amino acids in the peptides sent for sequencing were obtained. 5 cycles were conducted for each sequenced band. 3.11 Statistical analysis All measurements, unless indicated, were conducted for at least n = 3 in separate experiments and reported as the mean with 95% confidence intervals derived from standard error (SE) across sample means. Significance between samples or time points was defined as p < 0.05 using either a two-tailed t-test assuming equal variances or single factor analysis of variance (ANOVA) with post-hoc testing. 78 Chapter 4: Functional modulation of protein C by plasmin 4.1 Rationale Mutations in PC that cause either an antigenic deficiency or secreted protein with reduced function, result in increased thrombophilia risk and can be associated with inflammatory disease [415-418]. Although PC deficiency is most commonly inherited in an autosomal dominant manner where approximately 50% of circulating plasma PC levels can result in mild disease, a much rarer autosomal recessive form exists of severe PC deficiency that presents in neonates with purpura fulminans [419]. It is widely accepted that the thrombomodulin-thrombin complex is the only physiologically relevant activator of PC. However, early reports dated approximately three to four decades ago showed that plasmin proteolyzes PC and that this link between fibrinolysis and anticoagulation pathways may be an important regulatory mechanism in hemostasis [357, 359, 420]. Not surprisingly, this timeline paralleled the development of recombinant tPA, which sparked huge interest in its application in dissolving clots particularly in ischemic stroke. In addition to heightened physiological concentrations due to haemostatic localization, modulation of PC by plasmin may become increasingly important during thrombolysis when plasmin concentrations are elevated. Whether this proteolytic interaction results in dysregulated hemostasis is not known. Understanding the role of plasmin in regulating PC in either normal hemostasis or disease warrants further investigation as existing literature is scant. In vitro functional biochemistry data showed a biphasic modulatory activity whereby plasmin cleavage initially confers transient partial APC-like activity followed by further cleavages that quickly inactivate the enzyme [357, 359]. In these reports, amidolytic activity towards S-2366âa 79 known APC substrateâwas measured in parallel with APTT and was only able to reach amidolytic activity levels of < 40% and prolong clotting times to < 20% compared to thrombin-activated PC [357, 359]. In contrast, infusion of uPA in an in vivo baboon model showed a >10x increase APC amidolytic activity relative to baseline and comparable activity to exogenous APC [360]. Furthermore, acute MI patients treated with streptokinase had greatly elevated APC antigen levels using an antibody-based capture assay [361]. However, the authors in this study did not measure enzyme activity in parallel [361]. Since 1995, there has been no further work on understanding the interaction between plasmin and PC, a link between fibrinolytic and anticoagulant pathways that once uncovered may have a profound impact on health and disease. Besides its well-known anticoagulant functions, the cytoprotective effects of APC mediated through cleavage of PAR-1 has gained significant interest over the years. Interestingly, work in this field has been pioneered in part by the same research groups who first documented the regulatory role of plasmin in modulating PC. The largest clinical trials studying APC within a cytoprotective context is in sepsis, where results remain inconclusive as to its efficacy [343]. It is questionable why recombinant wild-type APC has largely been used in sepsis trials, when variants exist that lack anticoagulant activity but retain the ability to cleave PAR-1. A variant of APC (i.e. 3K3A-APC) that has Lys191-193 mutated to Ala191-193 to disrupt its recognition of FV(a) and FVIII(a) but maintains the ability to signal through PAR-1, has been developed and is being studied for neuroprotection in stroke as an adjunct to tPA-mediated thrombolysis [335, 353]. A purported mechanism by which APC confers neuroprotection in thrombolysis by cleaving PAR-1 is in inhibiting a tPA-induced nuclear factor kappa B (NFκB)-dependent matrix 80 metalloproteinase 9 (MMP-9) pathway [421-423]. It is hoped that this variant of APC could improve thrombolytic therapy in stroke by reducing extravascular neuronal toxicity frequently seen with tPA treatment [424]. Whether plasmin modulation of (A)PC impacts either anticoagulant function or PAR-1 cleavage is inconclusive and may be of greater significance during thrombolysis. Like thrombin, plasmin cleaves a diverse array of substrates and modulates their functions. Plasmin has been shown so far to proteolytically convert the proteases FIXa and FXa into fibrinolytic cofactors for tPA [234, 370], the latter which has been engineered to demonstrate efficacy in vivo [369]. Being homologous, (A)PC may have a similar effect following cleavage by plasmin. When trying to rationalize the additional roles that plasmin may play in the context of fibrinolysis, it would make sense that plasmin would positively feed back to amplify its own generation by converting specific plasma constituents into cofactors for tPA and plasminogen to âprimeâ fibrin for dissolution. Extensive work has been done on natural variants and generating engineered mutants in compiling the structure-function relationships of (A)PC and its activation, binding partners, and substrates. However, these recent reviews have not attributed any matter of significance to plasmin and its reported proteolysis of PC. The goal of the current work is to further dissect the biochemistry of (A)PC proteolysis by plasmin and its functional sequelae. Given the growing interest in recombinantly engineering a strictly cytoprotective variant of APC into a therapeutic drug for a variety of disease contexts, it will be important to discern the role of plasmin in modulating (A)PC. 81 4.2 Hypothesis It is hypothesized that plasmin activates anticoagulant and cytoprotective functions of PC. Concentrations of plasmin used previously by groups to study the role of plasmin in modulating PC were likely too high to see significant, discernable effects on activation. This is supported by two in vivo studies showing activation of PC during thrombolysis treatment. Although (A)PC has significant homology with FIX(a) and FX(a), sequence alignment shows that (A)PC has arginines instead of lysine residues at P1-positions where plasmin is known to cleave. It is therefore expected that (A)PC is not converted into a cofactor for tPA by plasmin. Instead, it is hypothesized that cleavage by plasmin occurs at yet unidentified sites that preserve active site activity allowing for recognition and proteolytic processing of its substrates. When rationalizing these modulatory effects within a hemostasis context, activating PC would be an additional way for plasmin to induce a more fibrinolytic state by enhancing anticoagulation to dampen coagulation. In addition, turning on PAR-1 mediated cytoprotection may play an important role in wound healing and dampening inflammatory pathways. 4.3 Results 4.3.1 Proteolysis of PC-Hc correlates with increased APC amidolytic activity Chromogenic activity towards S-2288, a selective APC substrate, was measured during proteolysis of PC by plasmin with samples taken in parallel for silver staining (Figure 28). Plasmin cleaved the PC-Hc doublet (~39-43 kDa) into four primary (~20, 18, 17, 13 kDa) and low molecular weight (<5 kDa) fragments. The 20 and 13 kDa fragments appeared at the first cleavage timepoint (5 minutes) and was associated with an initial increase in measurable APC activity (p < 0.05). This was followed by subsequent 82 enhancement of chromogenic activity and appearance of 18 and 17 kDa fragments. There 83 was a slight decrease of a 20 kDa fragment while a 13 kDa fragment increased in staining intensity over time. The lack of an apparent downward shift in non-proteolyzed PC-Hc suggested that cleavage by plasmin occurred sequentially, first within the protease domain and then followed by the activation peptide. Based on protein staining, it is likely that the 20 kDa fragment contains the canonical activation peptide. Intact PC-Hc was nearly completely proteolyzed (>95%) at the end of the experimental duration where amidolytic activity reached that observed of an equimolar concentration of purified plasma-derived APC. Therefore, fragmentation of PC-Hc correlated with the activation of the zymogen. This is the first report that plasmin cleavage induces complete activation of PC. During activation by plasmin, PC-Lc appeared to exhibit a slight downward shift in apparent molecular weight over time. An enzyme to substrate ration of 1 to 10 was chosen to measure activation of PC by plasmin within one hour while limiting the extent of autoproteolysis of newly activated PC. Figure 28: Plasmin cleavage of PC-Hc activates APC-like activity Plasma-derived human PC (1 μM) was incubated with plasmin (0.1 μM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at 25°C. Samples were drawn at selected intervals over 1 hour for measurement of APC-like activity by S-2288 chromogenic substrate cleavage and electrophoretic separation on a 15% polyacrylamide gel under reducing conditions. (A) S-2288 cleavage of incubation mixtures containing cleaved (â) or intact untreated PC (â ) was assayed for colorimetric change at 405 nm. PC-containing samples were diluted in S-2288 substrate to 10 nM. To prevent further cleavage of PC or fragments during the absorbance reading, 25 nM of aprotinin was added to the substrate mixture. Each time point was compared to activity of 10 nM native APC (â) taken during the start of the assay. Error bars for APC are highlighted in grey. The data are a compilation of technical triplicates with 95% confidence intervals used to compare time points. (B) In addition to chromogenic substrate cleavage analysis, samples were also taken for silver staining at each time point to correlate fragmentation with activity. These samples were prepared in parallel, stopped with sample buffer, and run under reducing conditions on a 15% polyacrylamide gel. PC-Hc, PC-Lc, and fragments of PC-Hc with their apparent molecular weights are labelled. The light chain of plasmin (i.e. plasmin-Lc) is also labeled. The silver stain shown is a representative sample of the experiment done in triplicate. 84 4.3.2 N-terminal sequencing of PC fragments following plasmin cleavage When correlating APC chromogenic activity to fragmentation of PC by plasmin, samples were saved for N-terminal sequencing by Edman degradation. Specifically, fragments observed in early (0 and 5 minutes) and late (60 minutes) time points were sent for sequencing. Four N-termini were identified following proteolysis of PC by plasmin (Table 1). From these, two distinct sites of plasmin cleavage were identified. The cleavages occurred at the known thrombin cleavage site R169-L170 and within the protease domain at R314-T315. In total, four distinct fragments were seen by silver stain at the end of the assay, two of which when sequenced had the N-terminus TFVLN. Their differential electrophoretic mobility was likely due to differential glycosylation at N329 (Figure 29). Table 1: N-termini of major fragments following cleavage of PC by plasmin Edman degradation was used to N-terminally sequence major fragments of PC following proteolysis by plasmin. 2 µg of fragmented PC was reduced with β-ME and separated electrophoretically on a 15% polyacrylamide gel. Fragments were transferred to PVDF and stained with Coomassie blue R-250. Individual bands were excised at early (0 and 5 minutes) and late (60 minutes) time points and sent for sequencing at the Tufts University Core Facility (Boston, MA, USA). 85 Unlike FIX(a) and FX(a), cleavage of PC by plasmin did not occur on the C-terminal end of lysine residues and instead plasmin cleaved to expose C-terminal arginines. However, there could be further proteolytic processing near the new C-termini of PC-Hc, PC-Lc or the various PC-derived fragments identified. N-terminal sequencing methods are unable to confirm this possibility. Like FIX(a) and FX(a), cleavage of the protease domain occurs in between non-disulfide linked sections. The differences between plasmin cleavage of PC within the protease domain and what has been reported for FIX(a) and FX(a) is its intact amidolytic activity. This indicates that these fragments are non-covalently Figure 29: Aligning N-terminal sequencing to fragmentation of PC by plasmin Edman degradation was used to correlate APC-like activity (i.e. S-2288 substrate cleavage) to N-terminal sequencing of plasmin-cleaved PC fragments. The stained PVDF membrane used for sequencing was overlaid with identical samples used for silver staining. The N-termini of corresponding fragments are shown. 86 associated as the catalytic triad residues are found on separate non-disulfide-linked fragments following proteolysis. 4.3.3 APC retains amidolytic activity following plasmin cleavage Cleaving APC with plasmin retained amidolytic activity as measured by both S-2288 and S-2366 chromogenic substrate cleavage (Figure 30). In fact, the results showed that cleavage of two known APC chromogenic substrates was increased by at least two-fold at each concentration studied. Based on these findings, treatment of APC with plasmin did not compromise its catalytic site. It is plausible that plasmin cleavage induced a conformational change to allow APC to better recognize and process these two chromogenic substrates. This may translate to enhanced cleavage of known substrates. Figure 30: Amidolytic activity of APC is enhanced following plasmin cleavage APC (1 μM) was cleaved with plasmin (0.1 μM) (â¦) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at room temperature for 1 hour. Mixtures were then immediately diluted to the three indicated concentrations in S-2288 and S-2366 substrate. Dilutions were assessed for cleavage as measured by absorbance at 405 nm. Untreated plasma-derived APC (â ) and plasmin (â) were included as controls. The experiment was performed once in triplicate with means and standard deviation (SD) shown. 87 4.3.4 Sequencing of APC-derived fragments following plasmin cleavage Cleaving APC with plasmin generated a different fragmentation pattern compared to PC by N-terminal sequencing. N-terminal sequencing revealed two plasmin cleavage sites within the protease domain of APC, K311-R312 and R314-T315 (Table 2). APC was cleaved by plasmin until complete disappearance (<1%) of APC-Hc as seen on silver stain (Figure 31). Interestingly, the K311-R312 cleavage exposed a CTL that may contribute to fibrinolytic enhancement of tPA. Whether cleavage at both sites by plasmin allows APC to retain its intact enzymatic activity is not known. Based on different N-termini seen by sequencing the two bands corresponding to C-terminal fragments, cleavage data suggest that there are conditions that favour either cleavage due to the differentially glycosylated residue N329. This may be due to steric effects conferred by differential glycosylation. Table 2: N-termini of major fragments following cleavage of APC by plasmin Edman degradation was used to N-terminally sequence major fragments of APC following proteolysis by plasmin. 2 µg of fragmented APC was reduced with BME and separated electrophoretically on a 15% polyacrylamide gel. Fragments were transferred to PVDF and stained with Coomassie blue R-250. Individual bands were cut and sent for sequencing at the Tufts University Core Facility (Boston, MA, USA). 88 Figure 31: Aligning fragmentation of APC by plasmin to N-terminal sequences APC (5 μM) was cleaved with plasmin (0.5 μM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at RT for 1 hour. The reaction was stopped in sample buffer containing BME and separated by SDS gel electrophoresis on a 15% polyacrylamide gel. Protein (2 μg) was transferred onto PVDF and stained with Coomassie blue R-250 as shown. Bands were then sent for N-terminal sequencing. 4.3.5 APC activity decreases with prolonged plasmin cleavage or autoproteolysis Correlating fragmentation of (A)PC with amidolytic activity during a prolonged incubation with plasmin was done to study whether there was a biphasic pattern of activation and inactivation as previous groups have reported. These groups used high plasmin concentrations, at times in excess of substrate, which may have limited the investigation of plasmin-mediated effects on PC function. In comparison, a 1 to 50 89 plasmin to substrate molar ratio was used here. Over the span of eight hours, cleavage of PC showed a biphasic pattern where there was an initial increase followed by a gradual loss in amidolytic activity that correlated to continued fragmentation of PC-Lc and the respective fragments derived from PC-Hc cleavage (Figure 32 A/B). Maximal activity was reached prior to complete proteolysis of PC-Hc, indicating that a higher plasmin concentration is necessary to preferentially degrade PC-Hc while leaving PC-Lc intact. Incubation of plasmin with APC showed a gradual loss of activity as proteolysis occurred Figure 32: Loss of APC activity over time during prolonged plasmin incubation 1 μM of both PC (â; A) and APC (â; C) were incubated separately with plasmin (20 nM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at 25°C. Intact PC (â ; A), APC (â ; C), and plasmin alone (â¦; A/C) were used as controls. (A, C) Samples were drawn at selected intervals over 8 hours to measure APC-like activity by both S-2288 and S-2366 chromogenic substrate cleavage. (B, D) Samples were taken in parallel for silver staining after being separating on a 15% polyacrylamide gel under reducing conditions. The experiment was done once in triplicate with SD reported. 90 over time (Figure 32 C/D). It is also important to note the time-dependent loss of plasmin, indicating that loss of amidolytic activity may also be due to autoproteolysis of cleaved PC or APC. 4.3.6 Dissecting autoproteolytic-mediated fragments of APC Autoproteolysis of APC was observed to induce a similar fragmentation pattern compared to plasmin cleavage. Here, active-site inhibited APC fragmentation was followed by silver stain. Interestingly, autoproteolysis did not result in the R314-T315 cleavage fragment but instead proteolyzed at K311-R312 (Figure 33). This was inferred based on apparent molecular weight and known basic amino acid location and has not been confirmed by N-terminal sequencing. During prolonged incubation, it is likely that autoproteolysis may occur in conjunction with plasmin cleavage. Fragmentation of active-site inhibited APC by Figure 33: Comparing fragmentation of APC in the presence or absence of plasmin 1 μM of active-site inhibited APC was incubated with plasmin (20 nM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at 25°C. In this sample, APC was treated with p-APMSF prior to incubation with plasmin. A control incubation mixture containing 1 μM of APC was prepared in the presence of SUVs (50 μM) and CaCl2 (2 mM) at 25°C. 91 plasmin yielded the same apparent banding pattern as intact APC, suggesting that plasmin facilitates cleavage of APC at both K311-R312 and R314-T315. 4.3.7 Plasmin-treated PC has impaired anticoagulant function compared to APC APTT clotting times were measured to assess the anticoagulant activity of plasmin-cleaved PC. With intact amidolytic activity, it was expected that the addition of plasmin-activated PC (Pn-PC) to PC-deficient plasma would prolong the APTT by inactivating FV(a) and FVIII(a). Unexpectedly, adding Pn-PC to PC-deficient plasma resulted in only a slight to modest increase in APTT compared to APC. PC mixtures that were cleaved longer by plasmin were associated with a greater prolongation of APTT. Amidolytic activity by S-2288 cleavage of proteolyzed-PC was equated to APC to ensure both mixtures had the same chromogenic activity prior to addition to plasma. Reconstituting PC to PC-deficient did not have a statistically significant (i.e. N.S.) impact on clotting times (Figure 34). Although significant (p < 0.05) when compared to addition of PC zymogen, there was a mild prolongation in clotting times when comparing plasmin-treated PC samples to APC. Post-hoc analysis comparing the prolongation of APTT by Pn-PC to APC showed a significantly (p < 0.001) decreased anticoagulant function. This suggested that although plasmin proteolysis of PC activates enzymatic activity, processing anticoagulant substrates compared with APC is significantly impaired. 4.3.8 Fragmentation of FVa by plasmin-proteolyzed PC Plasmin-treated PC was incubated with FVa at a relatively high 1 to 10 molar ratio to assess whether proteolysis by plasmin compromises the ability to cleave and inactivate its well-known procoagulant substrate. Compared with APC, cleavage of FVa was significantly impaired as seen by protein staining (Figure 35). Even with intact 92 chromogenic activity, cleavage by plasmin abrogated the ability of PC to inactivate its protein substrate, FVa. Figure 34: Plasmin cleavage impairs the anticoagulant function of PC PC (1 μM) was treated with plasmin (0.1 μM) for 5, 15, 60 and 90 minutes in the presence of SUVs (50 μM) and CaCl2 (2 mM) at 25°C. Samples were then treated with aprotinin (0.25 μM) to inhibit residual plasmin while leaving functional APC amidolytic activity intact as assessed by S-2288 substrate cleavage. Samples containing non-cleaved PC and APC were also treated with aprotinin as controls. 100 μL of PC-deficient plasma was thawed and combined with 50 μL of APTT reagent and incubated for 5 minutes at 37°C. 50 μL of each sample was then added (50 nM final PC concentration) and APTT was measured using a Coagulation Analyzer. This experiment was conducted three times in technical triplicates (n = 3) with the mean and 95% confidence intervals reported. Single factor ANOVA and post-hoc analysis using paired t-tests assuming equal variances was conducted to determine differences between groups. 93 Figure 35: Plasmin cleavage of PC impairs its ability to cleave FVa A preparation of plasmin-cleaved PC (0.1 μM) was incubated with FVa (1 μM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at room temperature. Cleavage of FVa by Pn-PC was assessed by fragmentation of FVa-Hc compared to APC on a Coomassie blue-stained 5-15% polyacrylamide gradient gel. 4.3.9 Plasmin cleavage does not convert (A)PC into a fibrinolytic cofactor Being homologous to the coagulation proteases FIXa and FXa, it was tested whether plasmin cleavage of PC or APC may result in CTL expression within the peptidase domain to facilitate enhancement of tPA-mediated plasminogen activation. A chromogenic assay measuring plasmin generation was developed whereby PC, APC, and FXa were incubated with a mixture containing tPA and plasminogen. Enhancement 94 of tPA activity was measured through plasmin generation cleaving S-2251. There was no statistically significant increase in plasmin generation when tPA and plasminogen were incubated with PC, APC, or their plasmin-cleaved fragment mixtures (Figure 36). Compared to FXa, there was a negligible enhancement of tPA activity. The slight increase Figure 36: Plasmin does not convert (A)PC into a tPA accelerator PC (â²) and APC (â ) (0.1 μM), both intact and plasmin-treated (Pn-PC, â³;Pn-APC, â¡) for 60 minutes, were incubated with tPA (1 nM), plasminogen (Pgn; 0.5 μM), SUVs (50 μM) and CaCl2 (2 mM). Incubation mixtures containing FXa (â) and just tPA and plasminogen alone (â¦) were used as a positive and negative control, respectively. Samples were taken over 1 hour and assayed for plasmin activity through S-2251 chromogenic substrate (200 μM) cleavage. 95% confidence intervals were used to compare samples. 95 in plasmin generation over time with APC relative to PC could be attributed to the prediction of an intra-peptidase domain CTL by N-terminal sequencing. This was repeated with plasmin proteolyzed PC and APC, where plasmin-treated APC showed an additional modest enhancement in tPA-mediated plasminogen activation. 4.4 Discussion 4.4.1 Proteolysis of PC by plasmin Plasmin proteolysis of PC in the presence of Ca2+ and aPL results in two distinct cleavages, 1) at the canonical thrombin cleavage site R169-L170 and 2) within the protease domain at R314-T315 (Figure 37). Cleavage within the protease domain occurs between catalytic triad residues, indicating a cleavage pattern similar to FIXa and FXa where the truncated C-terminal portion remains non-covalently associated. This explains persistence of complete chromogenic activity of plasmin-cleaved PC. Figure 37: Plasmin cleavage sites in PC Plasmin cleavage sites (â²) within the protease domain and the canonical thrombin activation site of PC-Hc are shown. By combining cleavage fragmentation with N-terminal sequencing data, a schematic illustrating the sequential plasmin cleavages of full-length PC was developed (Figure 38). In this model, plasmin cleavage of PC first occurs within the autolysis loop at 96 R314-T315. Whether cleavage first at R314-T315 is required for the subsequent cleavage at the canonical thrombin activation site at R169-L170 cannot be determined with these data. The resulting 17 and 13 kDa fragments had the same N-terminal sequence likely due to differential glycosylation at N329. The appearance of the 13 kDa fragment first during initial cleavages suggests that the non-glycosylated fragment is preferentially cleaved. If this were the case, it is possible that steric hinderance at the differentially glycosylated site of N329 may affect recognition by plasmin. However, this could also be due to a difference in Figure 38: Theoretical model illustrating the sequential plasmin cleavages of PC In this model, fragmentation of PC by plasmin occurs within the protease domain first at R314-T315 followed by cleavage of the activation peptide at R169-L170. The C-terminal piece of PC-Hc remains non-covalently associated following cleavage by plasmin. 97 sensitivity between the two fragments to silver staining. Although no fragments derived from PC-Lc were detected by N-terminal sequencing, a slight downward shift seen on protein staining over time may indicate C-terminal processing. This processing may also occur for fragments derived from PC-Hc. 4.4.2 Proteolysis of APC by plasmin Cleavage of APC by plasmin occurred at two sites within the protease domain, K311-R312 and R314-T315 (Figure 39). It is not known what causes this specific difference in plasmin cleavage between PC and APC where it is only the lack of an intact activation peptide that results in the distinction. Whether cleavage of APC occurs at K311-R312 or R314-T315 may depend on the presence or absence of glycosylation at N329. Since the two C-terminal fragments differ by only three amino acids but migrate much differently than indicated by peptide length, the presence of a glycosylated N329 may result in cleavage at K311-R312. Figure 39: Proteolysis of APC by plasmin Plasmin cleavage sites (â²) within the protease domain of APC are shown. 98 4.4.3 Functional modulation of (A)PC by plasmin 4.4.3.1 Proteolysis of PC by plasmin impairs anticoagulant activity The structural domains and motifs of (A)PC have been ascribed unique functions from several studies on natural and engineered variants. Plasmin cleavage of PC occurs in the protease domain within the autolysis loop (residues 302-317) and is adjacent to the Ca2+-binding (residues 223-235) and 191-loop (residues 190-193) regions (Figure 40). Figure 40: Highlighting critical surface loops that mediate APC function 3-D model derived from the crystal structure of APC showing the APC-Hc (beige) and APC-Lc (teal). Regions containing the active site, N-terminus of APC-Hc, autolysis loop, Ca2+-binding and 191-loop are highlighted. The site of plasmin cleavage at R314-T315 within the autolysis loop is shown. The crystal structure was rendered from SWISS-MODEL and shows human APC in complex with the small molecule inhibitor D-phenylalanyl-L-prolyl-L-arginine chloromethyl ketone (D-Phe-Pro-Arg-CMK) 99 The latter is also referred to as the 37-loop (residues 36-39) when using chymotrypsin numbering. Disrupting the electrostatic interactions between these three exosites have been shown to greatly impact the anticoagulant function of APC. The autolysis loop is a homologous feature among chymotrypsin-like serine proteases. The PC autolysis loop stabilizes the activation peptide in a Ca2+-dependent conformation that resists activation by free thrombin and instead makes the molecule more accessible for thrombomodulin-bound thrombin [425, 426]. This adds credence to the observation that plasmin cleaves PC within the autolysis loop prior to proteolysis of the canonical thrombin cleavage site. Cleavage at R314-T315 may induce a conformational change that makes the activation peptide more accessible to free plasmin. Moreover, thrombomodulin has been shown to be a plasminogen receptor [427, 428], and whether it may act as a cofactor for plasmin-mediated proteolysis of PC warrants further investigation. Proteolysis of APC by plasmin increases amidolytic activity, which may be explained by enhanced catalytic function seen when replacing the autolysis loop of APC with the shorter bovine form [425]. It is possible that proteolysis of the autolysis loop by plasmin conformationally allows APC to enhance catalytic efficiency. Single amino-acid substitution of basic residues within the autolysis loop including R314 showed decreased ability to inactivate both FVa and FVIIIa, which is in line with what was observed following proteolysis by plasmin at R314 [429, 430]. This indicates that region is important for recognition of its procoagulant substrates. Whether these variants still maintain the ability to signal through PAR-1 has not been determined. A patient with a T315 â A315 mutationâthe P1 site of plasmin cleavageâshowed expected impairment of anticoagulant function 100 but also significantly decreased anti-inflammatory function in cell-based assays [431]. This suggests that cleavage of PC by plasmin may play an important regulatory role in PAR-1 signaling. Ca2+-binding and 191-loop mutants have been pursued as novel recombinant variants of APC that have greatly impaired anticoagulant function but maintain the ability to cleave and signal through PAR-1 [341, 432]. In these studies, measurements of PAR-1 signaling and binding to EPCR were indistinguishable from wild-type APC. Several of these mutants have demonstrated cytoprotective properties in vivo, with 3K3A-APC advancing to human testing [355]. Mutations in the 191-loop have additionally implicated it in heparin-binding, since heparin-mediated inhibition by PCI was reduced [433]. Electrostatic interactions within these two regions have been clearly shown to confer the ability for APC to inactivate FV(a) and FVIII(a). Plasmin cleavage of the autolysis loop may also cause similar impairment due to its vicinity to the Ca2+-binding- and 191-loops. 4.4.3.2 Conversion into a tPA accelerator The exposure of two key CTLs forms the basis for how FIXa and FXa acquire the ability to enhance both tPA-mediated plasminogen activation and fibrinolysis following proteolysis by plasmin [234, 370]. Aligning the sequences of the homologous Gla-containing serine proteases FVII, FIX, FX, and PC show that FVII and PC have arginines in place of lysine residues at consensus sites where plasmin has been shown to proteolyze FIXa and FXa (Figure 41). Like (A)PC, FVIIa does not appreciably enhance tPA (data not shown). Proteolysis of FVIIa by plasmin has not been reported. Interestingly, like APC, FVIIa has been recently shown to bias for cytoprotection through PAR-1 signaling [434]. For (A)PC, cleavage by plasmin occurs not at Arg306 but eight 101 residues towards the C-terminus at Arg314 that preserves amidolytic activity. In contrast, proteolysis of FIXa and FXa by plasmin inhibits enzymatic activity [234, 370]. The two fragments of the protease domain following proteolysis of FIX(a), FX(a) and (A)PC remain non-covalently attached. Why plasmin-cleaved (A)PC retains amidolytic activity is not known. The lack of a predominant CTL-expressing fragment following cleavage of (A)PC may explain why proteolysis by plasmin does not confer the ability to accelerate tPA-mediated plasminogen activation. Figure 41: Sequence alignment of FVII, FIX, FX, and PC Sequences of FVII, FIX, FX, and PC were aligned to compare sites of plasmin cleavage. Completely conserved residues are highlighted in black, similar regions are highlighted in grey, with the rest indicating completely unconserved residues. The site at which plasmin is known to cleave FIXa and FXa is not completely conserved across Gla-containing serine proteases. Cleavage of FIXa and FXa both occur at consensus lysine residues at the P1 position, where FVII(a) and (A)PC have arginine residues instead. Cleavage of (A)PC by plasmin was shown here to occur at a completely unconserved region. 102 4.4.4 Physiologic relevance of plasmin modulating (A)PC Our data show that proteolysis of (A)PC by plasmin significantly impairs anticoagulant function. However, based on the specific site at which plasmin cleaves (A)PC, studies on natural variants and engineered mutants suggest that PAR-1 signaling may be preserved. We propose a model where thrombomodulin-bound plasmin cleaves EPCR-bound (A)PC both within the autolysis loop and at the canonical thrombin site (Figure 42). Plasmin-cleaved (A)PC stays bound to EPCR and activates PAR-1 on endothelial cells to bias signaling towards cytoprotection. This pathway would mirror the thrombin mechanism, whereupon an increase in plasmin concentration, a lower affinity interaction with thrombomodulin is driven to modulate (A)PC. Anticoagulant function would be inactivated to rebalance hemostasis by increasing procoagulant function. PAR-1 signaling would be maintained to enhance cytoprotection during would healing, which would prevent excessive bleeding and promote vascular repair. The proposed mechanism by which plasmin functionally regulates (A)PC may be supported by in vivo preclinical and clinical evidence showing that direct administration of plasmin to the site of the thrombus has a greater haemostatic safety advantage over other plasminogen activators including tPA in thrombolysis [435, 436]. The mechanism to explain this has been attributed to plasmin that escapes the clot site being rapidly inhibited by its serpin α2-antiplasmin. However, the possibility of plasmin indirectly signaling through PAR-1 and mediating cytoprotective effects through (A)PC cleavage that may decrease bleeding risk has not been considered. Intravascular plasmin has been gaining interest due to advancements in surgical techniques that allow for direct administration of thrombolytics to the thrombus. In addition, it may be wise for groups studying the efficacy 103 of targeted thrombolytics using echogenic or magnetic carriers to compare tPA with plasmin in their experimental models. The role of plasmin on its other known substrates, particularly (A)PC, may explain advantages of using plasmin over other plasminogen activators like tPA that are known to increase bleeding risk as well as induce cellular toxicity. Figure 42: A proposed model for the functional regulation of (A)PC by plasmin This model illustrates a mechanism whereby plasmin modulates (A)PC to selectively signal through PAR-1. These modulatory effects may be enhanced by cofactors on the endothelial surface including thrombomodulin and EPCR. 4.5 Future studies 4.5.1 Further dissecting cleavage and enzymatic activation of (A)PC by plasmin Observations comparing fragmentation of PC during plasmin cleavage suggests that cleavage occurs sequentially at R314-T315 followed by the canonical thrombin site, 104 R169-L170. Single-site mutagenesis studies of R356 (i.e. R356â A356) would have to be done to confirm a pre-requisite cleavage pattern, where mutation of this plasmin cleavage site either prevents or significantly delays activation of PC by free plasmin. It is also possible that thrombomodulin may act as a cofactor for plasmin and circumvent the requirement for cleavage first at R356-T357 to open accessibility of the canonical thrombin cleavage site as previously discussed. Moreover, this interaction between thrombomodulin and plasmin may be facilitated by known plasminogen receptors on cell surfaces like heterotetrameric annexin II complexed with p11 [365]. Preliminary in vitro assays assessing the ability for soluble thrombomodulin to direct and enhance the proteolysis of PC by plasmin are currently underway. Further understanding the biochemistry of how plasmin proteolyzes (A)PC will broaden existing knowledge in hemostasis physiology and may lead to novel recombinant constructs. 4.5.2 Potential role for preserved PAR-1 signaling and cytoprotection Whether PAR-1 signaling of plasmin-activated (A)PC is preserved is of great interest to us. As previously discussed, a variety of mutations within the autolysis, Ca2+-binding, and 191-loop have been reported that result in a lack anticoagulant function but retain the ability to cleave and signal through PAR-1. The known functions of APC in PAR-1-mediated cytoprotection include the alteration of gene expression, inducing anti-inflammatory and anti-apoptotic pathways, and protecting endothelial barrier function. Several studies can be conducted in vitro and in vivo looking for known sequelae of PAR-1 activation. First, it would be pertinent to determine whether plasmin-proteolyzed (A)PC can cleave at the canonical PAR-1 site for APC (i.e. R46-N47). This can be done using synthetic peptides comprising residues of the PAR-1 N-terminal tail and assessing 105 fragmentation by mass spectrometry [437], or a secreted embryonic alkaline phosphatase (SEAP) reporter assay by transfecting human embryonic kidney (HEK-) 293 cells with a SEAP-PAR-1 fusion construct [438]. Induction of the downstream Akt signaling pathway can also be assessed using phospho-specific antibodies against Akt [438]. Moreover, gene expression profiling using microarray analysis and quantitative RT-PCR in stimulated human umbilical vein endothelial cells (HUVECs) can be used to look for gene expression patterns that cluster into anti-inflammatory and anti-apoptotic pathways [439, 440]. PAR-1-mediated protection of endothelial permeability can be assessed both in vitro using Evans blue dye in thrombin-stimulated HUVECs grown to confluence that separate two permeable chambers, and in vivo using a VEGF-induced vascular leakage model in mice [438]. More classical in vivo models assessing the cytoprotective abilities of APC are in sepsis where plasmin-treated (A)PC can be used to determine whether there is a reduction in mortality in an LPS-induced endotoxemia mouse model [441]. More recently, brain injury models have been developed in mice that examine the neuroprotective capacity of APC in improving motor function following cortical impact [442, 443]. In vitro studies are currently being planned to look for the ability of plasmin-treated (A)PC to cleave and signal through PAR-1. If PAR-1 signaling function is preserved, collaborative opportunities with groups with greater expertise using in vivo functional models would be of significant interest to us. 4.5.3 Thrombolysis with direct plasmin administration Due to growing interest in applications for PAR-1-specific recombinant APC variants and direct plasmin administration as a means of improving thrombolysis, the importance of this work may be of great interest to the relevant research groups. Whether 106 proteolysis of (A)PC in thrombolysis patients is a significant phenomenon needs to be validated and could be assessed using patient plasmas pre- and post-treatment. If there is a significant reduction in hemorrhage and toxicities compared to conventional tPA use, the potential improvements of using plasmin directly may in part be attributed to its modulatory effect on (A)PC. 107 Chapter 5: To dissect the mechanism by which plasmin converts FVa from a procoagulant to a fibrinolytic cofactor 5.1 Rationale Plasmin has been shown to convert in vitro several vascular constituents including FVa, FIXa, FX and FXa into cofactors that accelerate tPA function [234, 235, 362]. Engineering FXa into a fibrinolysis enhancer has demonstrated efficacy in vivo [369]. The purported mechanism by which this occurs is through exposure of CTL binding sites for tPA and plasminogen [234, 235, 362]. Pursuing applications of FVa as a cofactor-based thrombolytic drug offers several important advantages. Unlike FIXa and FXa, generating recombinant FVa is not dependent on extensive posttranslational modification due to the lack of a Gla-domain and is therefore more amenable to large-scale production. This has the advantages both in the laboratory and in the clinic where scalable production facilitates in vivo evaluation and translation for therapeutic purposes. Most importantly, plasmin-mediated fibrinolytic fragment(s) of FVa offer a non-enzymatic, cofactor-based approach to accelerate fibrinolysis. Due to its localization to the clot via aPL-binding through its C1 and C2 domains, accelerating the activity of tPA as a cofactor may improve safety with thrombolysis by lowering exogenous levels of tPA required for recanalization. By inhibiting the procoagulant function of FVa and enabling its fibrinolytic activity, the thrombolytic derivative of FVa would localize plasmin generation specifically to the site of vascular injury without having a prothrombotic impact. Sequential cleavage of FVa-Hc and FVa-Lc by plasmin generates 16 distinct fragments identified by N-terminal sequencing [236]. It is also known that incubating FVa 108 with a mixture containing tPA, plasminogen, aPL and Ca2+ enhanced the activity of tPA-mediated plasminogen activation, indicating that FVa is proteolytically modulated into a tPA accelerator by plasmin [362]. The specific fragment(s) involved and whether this phenomenon occurs through a CTL-dependent mechanism is unknown. Work presented in this dissertation builds upon these findings to determine the specific FVa-derived fragment(s) that are responsible for enhancing tPA. Understanding the importance of this modulatory activity will supplement existing literature on the importance of auxiliary cofactors in fibrinolysis. Moreover, dissecting the physiologic significance of tPA cofactors besides fibrin will guide research and development of new thrombolytic therapies. The mechanism by which plasmin converts certain hemostasis constituents into cofactors for tPA is hypothesized to be an important haemostatic pathway for plasmin to regulate its own production by accelerating the âprimingâ of fibrin to amplify plasmin production and subsequent clot dissolution. 5.2 Hypothesis It is hypothesized that plasmin-mediated conversion of FVa generates a fragment mixture that remains bound to aPL to accelerate tPA-mediated plasminogen activation. This is due to the observation that although plasmin cleavage of FVa yields similar fragments in the presence or absence of aPL, the addition of aPL greatly enhances proteolysis [236]. Like other known tPA cofactors, this mechanism would likely be CTL-dependent and provide a scaffold for tPA and plasminogen binding in the presence of aPL. Therefore, the fibrinolytic fragment(s) of FVa would retain a functional portion of the C1 and C2 domains to confer aPL-binding. Moreover, since APC is the primary physiologic inactivator of FVa, it is hypothesized that APC-inactivated FVa would be 109 proteolyzed by plasmin into tPA-accelerating fragment(s). Most importantly, the fibrinolytic fragment(s) derived from proteolysis of FVa or APC-inactivated FVa by plasmin would ultimately enhance fibrin clot dissolution. 5.3 Results 5.3.1 Fragmentation of FVa by plasmin Proteolysis of FVa by plasmin was assessed to determine whether fragmentation correlated to what was previously reported in the literature [236]. The ~104 kDa FVa-Hc and ~71/74 kDa FVa-Lc doublet were sequentially cleaved into numerous fragments (Figure 43). Antibodies were used to delineate the derivation of each fragment from FVa-Figure 43: Plasmin cleaves FVa-Hc and FVa-Lc into a complex array of fragments Plasma-derived human FVa (1 μM) was incubated with plasmin (20 nM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at room temperature. Samples were drawn at selected intervals over 3 hours and run on a 10% polyacrylamide gel. A) Gels were then either stained with Coomassie blue G-250 or B) transferred onto PVDF to assess for FVa fragmentation via Western blot. A polyclonal antibody (AHV-5146) was used to detect FVa-Hc fragments and two monoclonal antibodies (AHV-14H12, AHV-5112) were used to probe for FVa-Lc. 110 Hc or FVa-Lc. The pattern of fragmentation by protein stain was in line with existing literature, where plasmin cleaves FVa-Hc and FVa-Lc into multiple fragments that run at apparent molecular weights of ~40-65 kDa and ~30 kDa. The only difference being the ~30 kDa fragment was not detectable by the same FVa-Hc polyclonal antibody (AHV-5146) as previously reported and was instead only seen on exposure using a FVa-Lc monoclonal antibody (AHV-14H12) specific to the A3 domain. AHV-5146 detected fragments only with molecular weights of ~45-65 kDa, where the larger ~65 kDa fragment derived from FVa-Hc showed weak antigenicity by Western blotting. Monoclonal antibodies with affinity to the A3 (AHV-14H12) and C1/C2 domains (AHV-5112) detected fragments with apparent molecular weights of ~30 kDa and ~40-50 kDa. 5.3.2 Fragmentation of FVa results in enhanced plasminogen activation by tPA To determine whether FVa could acquire fibrinolytic cofactor activity, an incubation mixture containing tPA and plasminogen was prepared. FVa significantly enhanced plasmin generation compared to tPA alone after a lag of 12.5 minutes (p < 0.05) (Figure 44). This lag phase was hypothesized to occur due to the time required for feedback cleavage of FVa by the small amount of plasmin produced intrinsically by tPA. Compared to intact FVa, pre-cleaving FVa with plasmin shortened the lag phase from 12.5 to 2 minutes to where there was significant acceleration of plasmin generation (p < 0.05). This finding is consistent with the single report showing an enhancement of tPA-mediated plasminogen activation by FVa [362]. These results indicate that FVa is converted from a procoagulant to a fibrinolytic cofactor during incubation with tPA and plasminogen in the presence of aPL and Ca2+. 111 Figure 44: FVa enhances tPA-mediated plasminogen activation FVa (0.1 μM; â ) was incubated with tPA (10 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM). Additional mixtures were prepared where prior to incubating with tPA and plasminogen, FVa was treated with plasmin (20 nM plasmin for 1 hour; â). Incubation mixtures were also prepared in the absence of any FVa (â¦). Samples were taken over 1 hour and assayed for plasmin activity by S-2251 chromogenic substrate cleavage (200 μM). Absorbance rates were converted to molar plasmin concentrations using a purified plasmin standard curve. Technical triplicates were conducted three times (n = 3) with 95% confidence intervals shown and used to compare time points. 5.3.3 Plasmin cleavage of FVa on aPL accelerates conversion into a tPA cofactor Proteolysis of FVa in the presence or absence of aPL has been previously shown to result in the same apparent mixture of fragments, the difference being a marked acceleration of proteolysis with aPL present (Figure 45) [236]. Cleavage of FVa in the presence or absence of aPL under conditions reported in this dissertation showed preferential cleavage of FVa-Lc when aPL was present, whereas in its absence 112 proteolysis of FVa-Hc was slightly accelerated. The noteworthy difference, occurring in the absence of aPL, was significant resistance of FVa-Lc to proteolysis by plasmin. Figure 45: Plasmin cleavage of FVa in the presence or absence of aPL FVa (1 μM) was incubated with plasmin (20 nM) in CaCl2 (2 mM) and the presence or absence of SUVs (50 μM) at room temperature. Samples were drawn at selected intervals over two hours and run via SDS-PAGE on a 10% polyacrylamide gel. Gels were then stained with Coomassie blue G-250 to assess for FVa fragmentation. Conversion of other known cofactors of fibrinolysis follow a scaffolding mechanism that localizes the enzyme and substrate. It was therefore predicted that acquisition of tPA cofactor activity would be impaired in the absence of aPL, which would hold CTL-containing species proximal to one another for a scaffolding effect and due to slower conversion of FVa-Lc into its fibrinolytic fragments. As expected, treating FVa with plasmin in the presence of aPL prior and during incubation with tPA and plasminogen significantly accelerated plasmin generation (Figure 46). In the absence of aPL, enhancement of tPA 113 cofactor activity was significantly impaired. Since FVa-Lc was significantly more resistant to plasmin cleavage in the absence of aPL, this suggested that the fibrinolytic fragment(s) of FVa are derived at least in part from FVa-Lc. Figure 46: Plasmin cleavage of FVa on aPL accelerates fibrinolytic activity Proteolyzed FVa was prepared and incubated in the presence (â) or absence (â ) of SUVs (50 μM). These two preparations were then assessed for enhancement of tPA-mediated activation of plasminogen by chromogenic substrate cleavage. Incubation mixtures were also prepared in the absence of any FVa (â¦), with readings combined from samples with and without aPL. All incubation mixtures contained tPA (10 nM), plasminogen (0.5 μM), and CaCl2 (2 mM). Technical triplicates were conducted three times (n = 3) with 95% confidence intervals shown and used to compare time points. 5.3.4 Longer proteolysis of FVa by plasmin correlates to enhanced tPA activity Cleaving FVa for different durations prior to incubation with tPA and plasminogen was used to correlate early or late fragmentation with the acquisition of tPA cofactor activity. Treating FVa for 60 minutes resulted in the greatest increase plasmin generation over time (Figure 47). The 0.5- and 5-minute time points were not statistically significant compared to intact FVa during the incubation period, indicating that later fragments are responsible for accelerating tPA in this assay. When visualizing the fragmentation of FVa 114 in samples added to incubation mixtures containing tPA and plasminogen, increased plasmin activity correlated with the appearance of ~45, 43, and 40 kDa fragments. The latter two fragments are recognized by AHV-5112 and have been shown previously to be derived from FVa-Lc [236]. This supports the notion that tPA-enhancing fragments following proteolysis of FVa by plasmin are derived from FVa-Lc. Figure 47: Correlating fragmentation of FVa by plasmin to tPA cofactor function FVa was pre-treated with plasmin for 30 seconds (â²), 5 minutes (â ), and 1 hour (â) prior to incubation with tPA (1 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM). A lower concentration of tPA was used to initiate plasmin generation to relatively amplify the effects of plasmin-mediated feedback compared to intact FVa. Samples were collected prior to incubation for A) assessment of chromogenic activity and B) SDS-PAGE. Incubation mixtures containing untreated FVa (â¦) and the absence of any FVa (â) were used as controls. Technical triplicates were conducted three times (n = 3). 95% confidence intervals are shown and used to compare time points. The protein stain shown is a representative image of pre-treatment samples used for incubation. 5.3.5 Plasmin cleavage is required to convert FVa into a tPA cofactor It has not been unequivocally shown that only proteolysis by plasmin is required for conversion of FVa from a procoagulant to fibrinolytic cofactor in experimental reaction 115 mixtures. To rule out potential cleavages by tPA, aprotinin was used to prevent feedback proteolysis of FVa by plasmin. In the absence of FVa, there was no appreciable conversion of plasminogen to plasmin as seen by SDS-PAGE (Figure 48). As expected, FVa enhanced tPA-mediated plasminogen activation over time. The addition of aprotinin completely abrogated the ability for FVa to activate plasminogen into plasmin, indicating that FVa must be feedback modulated by newly produced plasmin to accelerate tPA. Pre-treating FVa prior to incubation, however, overcame inhibition by aprotinin, verifying the existence of a plasmin-mediated mechanism of FVa conversion FVa into a tPA cofactor. Figure 48: Aprotinin prevents conversion of FVa into a tPA cofactor by plasmin Incubation mixtures containing FVa (0.1 μM), tPA (1 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM) were prepared in the presence or absence of aprotinin, a small peptide targeting the active site of certain trypsin-like serine proteases with a high affinity for plasmin. Aprotinin inhibits plasmin but had no effect on tPA at the concentrations used. Samples were taken over 3 hours, run on a 10% polyacrylamide gel and stained with Coomassie blue G-250 to assess for plasmin generation. Plasmin-treated FVa prior to incubation with tPA and plasminogen was used as a positive control, whereas absence of FVa was used as a negative control. 116 Plasmin used to pre-treat FVa prior to incubation was further diluted in the incubation mixture and thus not seen by protein staining. Cleavage of FVa by APC, its accepted inactivator in plasma, did not confer the ability to accelerate the activation of plasminogen by protein staining (Figure 49). This was shown by incubating APC-cleaved FVa with aprotinin where no appreciable generation of plasmin was observed. This indicated that the mechanism by which FVa is Figure 49: Cleavage of FVa by APC does not confer tPA cofactor activity Incubation mixtures and collection samples were prepared as previously described, where instead of plasmin, APC was used to pre-treat FVa prior to incubation. The ability for APC-cleaved FVa (APC-FVa) to enhance plasmin generation was assessed in the presence or absence of aprotinin by protein staining with Coomassie blue G-250. 117 converted to a tPA accelerator by plasmin may involve CTLs as APC is known to strictly cleave at arginine residues. In the absence of aprotinin, APC-cleaved FVa enhanced tPA-mediated plasminogen activation over time, suggesting that fragments of FVa derived from the combination of plasmin and APC cleavage enhanced tPA. 5.3.6 APC-inactivated FVa remains plasmin cleavable Since APC-pretreated FVa acquired tPA-enhancing activity after further treatment with plasmin, the fragmentation profile was investigated. The electrophoretic band pattern of APC-cleaved FVa-Hc is shown as major fragments of with apparent molecular weights of ~45, 30 and 25 kDa (Figure 50). APC-cleaved FVa was then subjected to additional Figure 50: Fragmentation of FVa by APC and plasmin FVa (1.5 μM) was cleaved with APC (0.15 μM) in the presence of SUVs (50 μM) and CaCl2 (2 mM). 1 μM APC-cleaved FVa (APC-FVa) was then subjected to secondary cleavage by 0.1 μM plasmin (APC-Pn-FVa). This fragment mixture was compared with plasmin-treated FVa prepared as previously described (Pn-FVa). Samples were loaded into 5-15% polyacrylamide gels and run electrophoretically under reducing conditions. Gels were stained with Coomassie blue G-250. 118 treatment by plasmin, where the resulting fragmentation closely resembled that of plasmin-cleaved FVa, supporting acquisition of tPA-enhancement. 5.3.7 Plasmin modulates APC-inactivated FVa into fragments that accelerate tPA APC is the primary physiologic inactivator of FVa. Therefore, its fragmentation of FVa was evaluated for acquisition of fibrinolytic activity. Cleavage by APC did not enhance the conversion of FVa into a tPA-accelerator (Figure 51). However, subsequent treatment of APC-inactivated FVa with plasmin resulted in fragments that accelerated tPA to a similar degree compared with fragments obtained from plasmin treatment alone. This Figure 51: Plasmin converts APC-inactivated FVa into a tPA accelerator Pn-FVa (â), APC-Pn-FVa (â ), APC-FVa (â²), and FVa (â¦) were assayed for enhancement of tPA-mediated plasminogen activation in incubation mixtures containing with tPA (1 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM). The absence of any FVa (â) was used as a control. Samples were taken over 1 hour and assayed for plasmin activity by S-2251 chromogenic substrate cleavage (200 μM). Technical triplicates were repeated twice (n = 2) with standard deviations shown. 119 strongly supports that APC-cleaved FVa retains its plasmin cleavage sites and capacity to function as a tPA enhancer. 5.3.8 Free and lipid-bound fragment mixtures both accelerate plasmin generation Fragments of FVa that bind aPL were separated from those in solution phase by density-dependent centrifugation on sucrose-containing LMVs. Separation of fragments was confirmed via Western blot, as all lipid-binding species must be in part FVa-Lc-derived (Figure 52). The 30 kDa species of FVa-Lc derived from the A3 domain was found Figure 52: Separation of lipid-bound and free FVa fragments on LMVs FVa (1 μM) was incubated with plasmin (20 nM), LMVs (2 mM) and CaCl2 (2 mM) at room temperature. After incubation for 1 hour, the mixture was spun at 14,000 x g in a microfuge. The supernatant fraction was collected and the pelleted fraction was washed three times in HBS containing EDTA (20 mM). The pellet was re-suspended in HBS and both supernatant and pellet fractions were assessed for fragmentation by protein stain and Western blotting. 120 only in the pellet fraction, indicating that it must be non-covalently associated with the 50-40 kDa fragments that contain the C1 and C2 aPL-binding domains. Interestingly, both lipid-binding fragments derived from FVa-Lc and fluid phase species derived from FVa-Hc significantly enhanced plasmin generation (Figure 53). This indicated that at least one fragment derived from both FVa-Hc and FVa-Lc mediates the tPA-accelerating activity of FVa observed following proteolysis by plasmin. Figure 53: Free and lipid-bound FVa fragments enhance tPA cofactor activity Lipid-binding (â) and fluid phase (â ) fragments of FVa following proteolysis by plasmin was prepared as previously described. Both fragment mixtures were then incubated with with tPA (1 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM) to assess for enhancement of plasmin generation. The absence of any FVa (â¦) was used as a control. Technical triplicates were conducted three times (n = 3) with 95% confidence intervals shown and used to compare time points. 121 5.3.9 Binding of FVa fragments to 125I-radiolabelled plasminogen Fragments of FVa following proteolysis by plasmin were assessed for binding to 125I-plasminogen and whether this binding required exposure of CTLs. Upon overlaying FVa fragments on PVDF with 125I-plasminogen, a band at ~45 kDa was observed on X-ray film (Figure 54). This band progressively became more intense over cleavage time by plasmin. The derivation of this fragment requires elucidation. Binding to 125I-plasminogen was inhibited in the presence of EACA indicating CTL dependence. Figure 54: Binding of FVa fragments to 125I-radiolabelled plasminogen FVa (1 μM) was incubated with plasmin (20 nM) in the presence of SUVs (50 μM) and CaCl2 (2 mM) at room temperature. Samples were taken over time, transferred onto polyvinylidene difluoride (PVDF) and overlaid with 125I-radiolabelled Pgn (1 μM) in the presence and absence of ε-aminocaproic acid (EACA), an inhibitor of CTL-binding. Autoradiography film was exposed to PVDF for three days and then developed. 122 5.3.10 The plasminogen-binding fragment of FVa is derived from FVa-Hc Binding to 125I-plasminogen was repeated after separation of FVa fragments on sucrose-containing LMVs to separate lipid-bound and solution phase fragments. Preliminary data showed a band, appearing during longer incubation with plasmin, being detected only in the supernatant fraction corresponding to the 45 kDa fragment reported previously (Figure 55). To summarize, cleavage analysis of FVa-derived fragments by Western blot showed that the unbound 45 kDa fragment is derived from FVa-Hc and binds 125I-plasminogen through a CTL-dependent mechanism that is inhibitable with EACA. Figure 55: Binding of lipid-bound and free FVa fragments to 125I-plasminogen Plasmin-cleaved FVa fragments were pulled down on LMVs as previously described. Pellet and supernatant fractions were transferred onto PVDF and overlaid with 125I-Pgn (125I-plasminogen). Autoradiography film was exposed to PVDF for three days. 5.3.11 Plasmin converts FV-810 into a tPA accelerator like FVa To study the role of plasmin-proteolyzed fragments of FVa in more complex fibrinolysis systems requiring larger amounts of protein, recombinant FV production was used. In collaboration with Dr. Rodney Camire (University of Pennsylvania), BHK cells stably expressing a FV variant with a truncated B-domain missing residue 811-1491 (i.e. 123 FV-810) was expressed and purified with ion exchange chromatography. Deletion of a large section of the B-domain lessens the burden of posttranslational modifications and allows for better expression and purification. FV-810 enhanced plasmin activity and was not significantly different than an equimolar concentration of FVa (Figure 56). Figure 56: FV-810 enhances tPA-mediated plasminogen activation Under identical conditions as previously described, FV-810 (â ) assessed for its ability to enhance tPA compared with FVa (â) in an incubation mixture containing tPA (10 nM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM). A control containing HBS buffer (â²) instead of FV-810 or FVa was used as a negative control. Technical triplicates were repeated twice (n = 2) with standard deviations shown. 5.3.12 FV-810 enhances one-step fibrinolysis A fibrinolysis assay was developed to determine whether FV-810 could enhance the dissolution of fibrin. FV-810 was compared with FV and FVa for the ability to enhance fibrinolysis. Only FV-810 enhanced purified clot dissolution by almost two-fold, whereas FV and FVa had no effect (Figure 57). To control for any fibrin-dissolving proteases that 124 may have been co-purified with FV-810 in low amounts, FV-810 was pre-treated with p-APMSF prior to addition into the fibrinolysis assay. Even with broad-spectrum serine protease inhibition, FV-810 enhanced fibrinolysis (data not shown). Enhancement by FV-810 also occurred in the complete absence of aPL, indicating that a scaffolding mechanism to enhance tPA-mediated plasminongen activation is not required (data not shown). Figure 57: FV-810, but not FV or FVa, enhances fibrinolysis Fibrin clots initiated by thrombin (10 nM) containing fibrinogen (3 μM), plasminogen (0.5 μM), tPA (50 pM), SUVs (50 μM) and CaCl2 (2 mM) were formed in the presence of FVa, FV or FV810 (0.1 μM). Turbidity at an absorbance of 405 nm was following by light scattering to monitor the solubilization of the thrombin-induced fibrin clot over time. The experiment was conducted once in triplicate with SD shown. 125 5.3.13 The residual B-domain of FV-810 after thrombin excision may accelerate tPA It was hypothesized that fragmentation of FV-810 by plasmin early during fibrinolysis modulated FV-810 into tPA-accelerating species to help âprimeâ fibrin for dissolution. However, sampling during fibrinolysis for Western blotting revealed that FV-Figure 58: Fragmentation of FV-810 during fibrinolysis Purified fibrin clots containing FV-810 (0.1 μM), fibrinogen (3 μM), tPA (10 pM), plasminogen (0.5 μM), SUVs (50 μM) and CaCl2 (2 mM) were formed using thrombin (10 nM). A) Fibrin clot opacity was followed over time by light scattering at 405 nm. Technical triplicates were conducted 3 times (n = 3) with standard error shown. Each reaction was stopped at hourly increments with SDS gel loading buffer and centrifuged to pellet insoluble material. B) Samples were subject to gel electrophoresis on a 5-15% polyacrylamide gradient gel under reducing conditions and transferred to PVDF. Western blot was conducted to assess for fragmentation using a polyclonal antibody (AHV-5146) and two monoclonal antibodies (AHV-14H12 and AHV-5112). 126 810 was converted into the anticipated plasmin-mediated fragments after fibrinolysis had completed (Figure 58). The effect of thrombin was implicated since no FV-810 remained prior to initiation of the reaction by addition of tPA, as only FVa was detectable at time = 0. This suggested the possibility of unique fragments of the recombinant FV-810, including the shortened B-domain, accelerating tPA-mediated plasminogen activation after being excised by thrombin. The mechanism is not yet known. 5.4 Discussion 5.4.1 FVa-derived fragments responsible for enhancing tPA Both free and lipid-bound FVa-derived fragments accelerated tPA-mediated plasminogen activation as measured by plasmin-specific chromogenic assays. Therefore, there must be at least two fragments derived from FVa following proteolysis by plasmin that separately enhance tPA cofactor activity. Given the observation that aPL-bound fragments are derived from FVa-Lc and unbound fragments from FVa-Hc, plasmin proteolysis results in at least one fragment from each chain of FVa that can accelerate the activation of plasminogen by tPA (Figure 59). A ~45 kDa fragment from FVa-Hc bound 125I-plasminogen and was inhibitable by EACA, indicating a CTL-dependent interaction. When considering both the apparent molecular weight of this fragment and known sites for plasmin cleavage on FVa, the 45 kDa fragment derived from FVa-Hc has a predicted CTL [236]. Plasmin cleaves at K364-K365 within the A2 domain to generate the 45 kDa species. Given that this fragment does not bind aPL and is not associated with any aPL-binding fragments of FVa, it may accelerate tPA-mediated plasminogen activation by clustering to subsequently provide a scaffolding surface. Whether this mechanism facilitates conformational transitions that 127 enhance plasminogen accessibility to cleavage or the transition from Glu- to Lys-plasminogen warrants further investigation. The observation that other plasmin-cleaved fragments of FVa with predicted CTL were not seen to bind plasminogen indicate that other binding regions are necessary to facilitate binding. Sequential plasmin cleavage of FVa-Lc occurs first at R1765 followed by proteolysis at K1827 [236]. The first cleavage gives rise to ~50, 48, and 30 kDa fragments. The former two fragments have the same N-terminal sequence but likely differ in apparent molecular weights due to differential glycosylation at Asn2182. The 30 kDa fragment derived from the A3 domain remains non-covalently associated to the C1- and C2-derived 50 and 48 kDa fragments. The 50 and 48 kDa fragments are further degraded to 43 and 40 kDa fragments following cleavage by plasmin at K1827 and still retain the ability to bind aPL. The 30 kDa fragment is quickly degraded into lower molecular weight fragments over time. The appearance of 43 and 40 kDa fragments correlated to the development of chromogenic plasmin activity, suggesting that these two aPL-bound isoforms are responsible for accelerating tPA-mediated plasminogen activation. Although none of the major fragments derived from FVa-Lc have a predicted CTL, aPL-bound fragments still accelerate tPA. There also is a possibility for proteolytic processing near the C-terminus of these fragments to expose CTLs. However, none of these fragments were seen to bind 125I-plasminogen when separated and immobilized on PVDF. It is possible that these FVa-Lc fragments only interact with plasminogen when bound to aPL due to inaccessibility when adhered to PVDF. Whether any of these fragments interact directly with tPA to accelerate plasminogen activation is not known. The prevailing model is that auxiliary fibrinolysis cofactors become functional accelerators of tPA following liberation of a CTL. 128 This is the first report of a proteolytically acquired hemostatic cofactor that can accelerate tPA-mediated plasminogen activation on a lipid scaffold through a mechanism independent of CTL-exposure. Figure 59: The tPA-accelerating fragments derived from cleavage of FVa The fragments of FVa believed to enhance tPA-mediated plasminogen activation are shown here. These fragments are derived from cleavages within the A2 and A3 domain facilitated by plasmin. The FVa-Hc-derived fragment is believed to bind plasminogen through a CTL-dependent mechanism. The FVa-Lc-derived fragments are not known to express a CTL and accelerate tPA through a yet unidentified mechanism. Recombinant FV-810, with residues 811-1491 of the B-domain deleted, enhanced both tPA and fibrinolysis subsequent to thrombin excision of the B-domain but prior to cleavage by plasmin. This observation suggests that the shortened B-domain of FV-810 may accelerate the activation of plasminogen. Binding to fibrin in the absence of CTL exposure has been attributed to plasminogen and tPA kringles binding to internal lysine residues on fibrin. However, the B-domain in FV-810 has the basic region deleted that is rich in lysines and arginines. Instead, the B-domain in FV-810 retains the acidic region rich in aspartic and glutamic acid residues. Whether these residues are important in mediating electrostatic interactions with tPA and plasminogen is unknown. A possible 129 hypothesis is that binding of the shortened B-domain in FV-810 induces either tPA or plasminogen to adopt a greater enzymatically active or activatable conformation. 5.4.2 Acquisition of tPA cofactor activity We have shown in vitro that FVa is modulated by plasmin to acquire a novel function as a tPA accelerator. The data presented in this dissertation suggest that FVa operates through several different mechanisms to enhance tPA-mediated plasminogen activation in vitro. In conjunction with other reported auxiliary cofactors for tPA, proteolyzed FVa may play an important role in hemostasis physiology by generating plasmin during the initial stages of fibrinolysis to help âprimeâ and expose CTLs on fibrin. The physiologic significance of proteolyzed FVa functioning as a tPA cofactor is supported by the observation that plasmin can still modulate APC-inactivated FVa fragments into tPA accelerators. Moreover, in thrombolysis, elevated plasmin levels are known to enhance proteolysis of FVa [405]. Perhaps the efficacy in recanalizing vessels seen with thrombolytic agents can be partially attributed to activating auxiliary fibrinolysis cofactors including FVa. The orientation of auxiliary fibrinolysis cofactors in vivo could be divided into two categoriesâunbound versus requiring a scaffold (Figure 60). Like FIXa and FXa, lipid-binding fragments of FVa enhance tPA-mediated plasminogen activation [234, 235, 370]. FXa-33/13, which expresses two CTLs, has been shown to bind two molecules of plasminogen [235, 370]. Whether other cofactors bind strictly one plasminogen or tPA per predicted CTL is not fully known. It is possible that multiple auxiliary cofactors work in unison on surfaces such as aPL or fibrin to better orient tPA and plasminogen for activation. In the scenario with aPL, PtdSer-expressing surfaces are known to be a high 130 affinity scaffold for hemostasis complexes and may be the primary site where auxiliary fibrinolysis cofactors function. Small molecules and fluid phase proteins like β2-glycoprotein I have been shown to conformationally induce plasminogen into a more easily activated state [444-446]. A fragment derived from FVa-Hc or the shortened B-domain excised from FV-810 may operate through similar mechanisms. Without a Figure 60: Proposed mechanisms of tPA-accelerating fragments derived from FVa This model illustrates the proposed mechanisms that may govern how fragments derived from plasmin proteolysis of FVa function to accelerate tPA. The FVa-Hc-derived fragment comprising A1 and a partial A2 may bind through exposure of its CTL and convert circulating plasminogen into a more âopenâ and activatable form. The FVa-Lc-derived fragment comprising C1, C2, and a smaller residual piece of the A3 domain may bind both tPA and plasminogen to accelerate plasmin generation. Another mechanism may be that one molecule each of tPA and plasminogen bind, which may function together with other known auxiliary fibrinolysis cofactors to co-localize tPA and plasminogen on PtdSer-expressing surfaces. 131 scaffold, it is plausible that the primary mechanism involves binding to plasminogen to induce a conformational change that makes the molecule more easily activated either from Glu- to Lys-plasminogen or directly to plasmin. 5.5 Future studies 5.5.1 Enhancing fibrinolysis The long-range goal of this research area is to develop new thrombolytic therapies, where tPA-accelerating fragments of FVa may be a viable option. An auxiliary cofactor may work to improve safety of thrombolysis by tPA through reducing the requirement for exogenous tPA or accelerating endogenous tPA stores. Preliminary fibrinolysis results have been unexpected. In a purified lysis assay it would be predicted that FV-810 to be modulated by plasmin generated during the assay and feedback to âprimeâ fibrin for dissolution. However, under our conditions, an insufficient amount of plasmin is generated at low picomolar tPA and FV-810 remains intact until after the burst of plasmin solubilizes the purified fibrin clot. This suggests experiments exploring proteolyzed FVa prior to addition to fibrinolysis that enhance the dissolution of fibrin compared to tPA and plasminogen alone. In plasma, to circumvent any procoagulant effect of FVa, proteolyzed FVa will be used in FV-deficient plasma to discern fragments of FVa that enhance clot dissolution. 5.5.2 Dissecting the tPA-accelerating mechanism of FVa fragments It would be prudent to pursue alternative methods of measuring the interactions of plasmin-proteolyzed FVa fragments with tPA and plasminogen. Methods like microscale thermophoresis offers a way of investigating the specific requirements for tPA and plasminogen binding. For example, dissecting whether binding of the FVa-Lc-derived 132 fragment occur on a scaffolding surface like aPL is required for the localized binding of tPA and plasminogen. With regards to the unbound tPA-accelerating fragment derived from FVa-Hc, it is possible that binding occurs exclusively with plasminogen. Binding kinetics can also be derived using this method. 5.5.3 Fibrinolysis enhancement in vivo There is already great expertise within our lab looking at thrombolysis in vivo and identifying biomarkers for systemic bleeding. tPA-accelerating fragments derived from FVa can be generated either from cleaving recombinant FV-810 and separating lipid-binding from unbound fragments or attempting to generate separate recombinant pieces thought to enhance tPA. These fibrinolytic fragments can be tested in their established murine carotid artery thrombosis model to study the efficacy of the thrombolytic tenecteplase. Safety can also be assessed by Western blotting for D-dimer following administration of thrombolytic cofactor. Moreover, a collaboration with the Brain Research Institute at UBC has been formed looking for reduced intracranial hemorrhage risk with auxiliary fibrinolysis cofactors compared to tPA. This model may be of interest if FVa-derived fragments demonstrate clot-dissolving efficacy. 133 Chapter 6: Conclusion The role of plasmin in functionally modulating its substrates apart from fibrin have been gaining significant interest. The wide spectrum of plasmin substrates within and outside of the vascular space show the complexities of plasmin regulation. Two of these substrates with limited evidence regarding their functions following proteolysis by plasmin are (A)PC and FVa. These two very different proteins have been shown to be proteolyzed by plasmin in patients undergoing thrombolysis with plasminogen activating drugs [361, 405]. The modulation of these two substrates by plasmin becomes therefore increasingly important to understand as these pathways may contribute to the limitations seen with tPA and offer new insights into improving thrombolytic therapy in the future. It was originally anticipated that plasmin cleaved PC would acquire anticoagulant activity. This would make sense as enhancing anticoagulation would tip the haemostatic balance in favour of a more fibrinolytic state. However, the opposite of this can also be rationalized, when plasmin concentrations increase, a lower affinity interaction with either PC or APC would inhibit anticoagulation to re-establish coagulation pathways and prevent bleeding. More interestingly, cleavage of PC and APC occurs within the autolysis loop of the protease domain that is consistent with natural and engineered variants that lack anticoagulant function but retain PAR-1 signaling. The cytoprotective role of plasmin in modulating either EPCR-bound PC or APC into a PAR-1 specific enzyme is intriguing and may explain several lines of evidence in vivo where direct plasmin administration is superior over tPA in decreasing the bleeding risk associated with thrombolysis. As clinical trials are compiling insight on direct plasmin administration, there may be an additional benefit in reducing cellular toxicities known to be associated with exogenous tPA. The 134 cytoprotective effects of APC are so widespread that in addition to engineered variants being tested for neuroprotection in stroke and sepsis survival, APC is being studied in dermatological settings as a way of improving cutaneous wound healing [447]. Apart from mode of administration, clot-dissolving drugs have not been improved since their inception almost 30 years ago. The major risk associated with using plasminogen activators is hemorrhage. The advantage of engineering a non-enzymatic auxiliary cofactor for tPA to either help lower the dose of exogenous tPA as an adjunct or completely remove its requirement by enhancing localized endogenous tPA activity is a very attractive premise. The significance of the auxiliary cofactor mechanism has already been demonstrated in vivo with a variant of FXa. The advantages of using FVa over a derivative of either FIXa or FXa is that tPA-accelerating fragments of FVa can be generated recombinantly without further modifications and may work through a mechanism independent of CTL-exposure. The FVa-Hc-derived fragment may function through a mechanism that binds plasminogen in circulation and conformationally induces a more open conformation for Glu- to Lys-plasminogen conversion or direct activation into plasmin. The FVa-Lc-derived fragment would circulate and bind to PtdSer-expressing surfaces at the site of injury and enhance tPA-mediated plasminogen activation on a scaffolding surface. Moreover, these tPA-accelerating fragments derived from FVa cleavage by plasmin lack procoagulant cofactor activity. This dissertation has identified novel pathways by which plasmin modulates the functions of (A)PC and FVa. The novelty of this work lies with correlating the biochemistry of plasmin modulating these two substrates and its applications in improving thrombolysis care. 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Thesis/Dissertation
eng
Pathology and Laboratory Medicine
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Functional regulation of anticoagulant protein C and clotting factor Va by fibrinolytic plasmin
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