BIOCHEMICAL STUDIES ON BLOOD COAGULATION MECHANISM EFFECT OF RECOMBINANT VON WILLEBRAND FACTOR A1 DOMAIN ON THE PROCESS OF HEMOSTASIS

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1 ARISTOTLE UNIVERSITY OF THESSALONIKI SCHOOL OF SCIENCE DEPARTMENT OF CHEMISTRY LABORATORY OF BIOCHEMISTRY BIOCHEMICAL STUDIES ON BLOOD COAGULATION MECHANISM EFFECT OF RECOMBINANT VON WILLEBRAND FACTOR A1 DOMAIN ON THE PROCESS OF HEMOSTASIS by SALAR ADNAN AHMED ZRARI MSc. in Clinical Biochemistry Hawler Medical University, 2002 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY IN CHEMISTRY (CLINICAL BIOCHEMISTRY) THESSALONIKI - GREECE 19 June 2012 I

2 CERTIFICATE OF APPROVAL Certification This is to certify that the Ph.D. thesis of Salar Adnan Ahmed, in title (Biochemical studies on blood coagulation mechanism Effect of recombinant von Willebrand factor- A1 domain on the process of hemostasis) has found that it is complete and satisfactory in all respects and have been accepted and approved by unanimity by the evaluation committee for the thesis requirement for the Doctor of philosophy degree in Clinical Biochemsitry at 19June 2012 graduation. FOUR EVALUATION COMMITTEE II

3 THREE THESIS COMMITTEE III

4 @ Salar Adnan ΑΠΘ Τίτλος ιδακτορικής ιατριβής (Biochemical studies on blood coagulation mechanism Effect of recombinant von Willebrand factor-a1 domain on the process of hemostasis) ISBN «Η έγκριση της παρούσης ιδακτορικής ιατριβής από το Τµήµα Χηµείας του Αριστοτελείου Πανεπιστηµίου Θεσσαλονίκης δεν υποδηλώνει αποδοχή των γνωµών του συγγραφέως»(ν.5343/1932, άρθρο 202, Παρ. 2) IV

5 This thesis is dedicated to My mother for her love, supporting and patience with me My dear brothers and sisters The pure soul and loving memory of my father My teachers and friends Every person who gave me opportunity to go on the right way V

6 ACKNOWLEDGEMENT It is a pleasure to thank many people who made this thesis possible and making my experience in the Aristotle University of Thessaloniki- Greece one of the most important and rewarding period of my life. It is my pleasure to express my deepest gratitude and appreciation to my supervisors, Asso. Prof. Dr. Panteleimon Arzoglou for his kind guidance, valuable suggestions, encouragement and continuous support during these years of study, I deeply appreciated. Words can not express my thanks and gratitude to Prof. Dr. Theodora Choli-Papadopoulou, Assi Prof. Dr. Eleni Nikolakaki and Asso Prof. Dr. Thomas Giannakouros for letting me participate in the project that you brought into being and that presented an excellent scientific base for the present thesis. Thanks for their scientific support and for reviewing and proofreading the thesis. I would like to thank also other members of the laboratory of Biochemistry Dr. Dimitrios Kyriakidis, Dr.Anastasia Pantazaki and Dr. Kotinis Konstantinos for their kindness at any time and for having created a nice working atmosphere. Many thanks and gratitude is extended to the Greek State Scholarships Foundation (IKY) for their support. I would further like to extend my gratitude to Kurdistan Regional Government, Presidency of Hawler Medical University, the dean of College of Medicine and the head of Department of Clinical Biochemistry. I gratefully acknowledge the secretaries and other members of the Department of Chemistry and Laboratory of Biochemistry for their support and help. I regret not being able to mention all the names of the various members of faculty as well as fellow graduate student, colleagues and friends who helped me make this project possible Finally, I would like to thank my mother, brothers and sisters for their love, continuous encouragement, endurance and constant support being with me through these years. Their toil and sacrifice have made me who I am today VI

7 There is nothing of which every man is so afraid, as getting to know how enormously much he is capable of doing and becoming. Søren Kierkegaard VII

8 SUMMARY The hemostatic system is a vital protective mechanism for the control of excessive bleeding after injury in the vascular system, as well as for the prevention of a hypercoagulative state during the coagulation response. One of the most important systems within the human body is the vasculature, because all organs and tissues are dependent on it for survival. Blood is a vital part of the human physiology; a transport system that brings nutrients and oxygen to sustain living cells and simultaneously facilitates the removal of carbon dioxide and metabolic waste products from the tissues of the body. Therefore maintenance of the vascular system, termed hemostasis, is vital for survival and is the controlled arrest of bleeding and maintains vessel integrity with minimal blood loss. To assure the continuity of these functions, it is of uttermost importance to keep the flowing blood inside the vascular system at any cost. The complex process of blood coagulation is driven through a complex enzymatic pathway that involves several zymogen-to-enzyme conversion steps and allows the reaction to undergo great intrinsic amplification. The final step in this cascade is the conversion of soluble fibrinogen into an insoluble fibrin network. Thrombin plays a central role in the coagulation cascade and is the last compound that gets activated. During activation of coagulation, thrombin is the only enzyme that is freely available in blood circulation, where it encounters different substrates. It s most relevant substrate is fibrinogen which is converted to fibrin by thrombin. Further more, in blood circulation, thrombin can induce physiological but often also pathological processes depending on the amount of available thrombin. Serine protease factor Xa (FXa), which functions principally at coagulation cascades catalyzes the conversion of prothrombin to thrombin at the sites of vascular injury. The serine protease domain of FXa is one of the most effective targets for developing direct anticoagulants for the coagulation disorders. The coagulation process is accelerated in the presence of calcium, and a phospholipid surface that is usually presented by the activated platelets, and will ultimately form a stabilizing fibrin network that will secure the platelet plug until the vessel wall is restored. von Willebrand factor (vwf) is a plasma protein that performs two main functions in hemostasis: it mediates platelet activation and adhesion to the injured vessel wall that leads to the formation of platelet thrombi and the subsequent arrest of bleeding. Furthermore, vwf protects coagulation factor VIII (FVIII) from rapid proteolytic inactivation. VIII

9 Cardiovascular diseases are one of the major causes of morbidity and mortality in the developed nations. Despite the continuous efforts by numerous laboratories all around the world, effective drug targets for diseases such as heart attack and stroke have not yet been identified. As of today this field stands in its early development stages only. There are numerous questions in this field of science which still require answers. As thrombin and FXa plays a fundamental role in the coagulation cascade, it is logical that the calculation of their levels, apart from a number of other important information, constitutes the most appropriate way to study the overall patients haemostatic status, therefore helping to monitor the patients condition. For example, this approach has recently been used for targeting potential extra-haemostatic effects as well as to identify patients at risk for recurrent venous thromboembolism. This piece of information makes extremely interesting the development of methods for monitoring thrombin and FXa production in vitro. Related to this vast area of science, in the present work we developed extremely simplified methods to determine thrombin and FXa production, which might be automatable. The most widely used chromogenic substrate was applied, several parameters involved in the reaction were examined and optimized in order to establish a rapid routine, less time consuming and suitable method for the evaluation of large numbers of clinical samples. The thrombin and FXa production values were compared in plasma samples before and after clot removal, correlation between FXa and thrombin production values have been calculated and also the effects of physical exercise, platelet count, FVIIa and tissue factor(tf) on thrombin and FXa production have also been carried out. Furthermore routine determination of this factor (Thrombin) in hospitalized diabetic patients under different anti-coagulating therapies have been carried out. As the coagulation cascade proceeds as a series of linked proteolytic reactions, the enzymes generated by these processes are not directly available for quantitative measurement because they are rapidly neutralized by naturally occurring protease inhibitors. To face these obstacles, we have also focused to the development of western blotting techniques in order to study blood clotting FX and its active form. In addition in the frame of this work we have investigated the potential function of purified vwf A1 domain protein in platelet activation and in the natural process of hemostasis in order to fully understand its particular role in maintaining the integrity of the blood coagulation. To this purpose the specific amino acid region (A1 domain) of vwf has been cloned, expressed in mammalian and bacterial cells, purified and used both as bacterial and cellular protein. IX

10 TABLE OF CONTENTES Page ACKNOWLEDGMENTS.VI SUMMERY...VIII TABLE OF CONTENTS....X LIST OF TABLES.XV LIST OF FIGURES.XVI LIST OF SYMBOLS AND ABBREVIATIONS...XIV CHAPTER I INTRODUCTION 1.1. Current concept of hemostasis Primary hemostasis The platelets and it s main functions in primary haemostasis...3 A. Platelet adhesion...5 B. Platelet aggregation...7 C. Platelet activation Coagulation (Secondary hemostasis) The pathways of the coagulation cascade.10 A. Intrinsic pathway (Factor XII - Contact activation pathway)...11 B. Extrinsic pathway (Tissue factor pathway) Cell-based model of coagulation...14 A. Initiation phase..15 B. Amplication phase.16 C. Propagation phase Fibrinolysis Proteins involved in blood coagulation processes Blood clotting factor X Prothrombin /Thrombin Tissue factor von Willebrand Factor von willebrand factor biosynthesis, storage and secretion..27 A. von Willebrand factor biosynthesis.27 B. von Willebrand factor storage.30 C. von Willebrand factor secretion 30 X

11 von Willebrand factor molecular size and structure Gene coding for von Willebrand factor von Willebrand factor s functional domains von Willebrand factor A domains von Willebrand factor cleavage and unfolding of A domains von Willebrand factor activity von Willebrand factor and activation of platelets von Willebrand factor and stabilization of FVIII..38 Aims of this study CHAPTER II MATERIALS AND METHODS 2.1. Study population Collection of Blood Samples Materials Experimental protocols Biochemical assays Thrombin production assay Factor Xa production assay Molecular methods (Molecular assay protocol) Cloning, expression and purification of mamalian cell-derived recombinant vwf- A1 fragments Construction of pcdna3.1 myc-his (-) B vwf-a1 domain expression vector pcdna.3.1/myc-his(-)b vector Plasmid DNA preparation 43 A. Plasmid DNA mini-preparation protocol..43 B. Plasmid DNA maxi-preparation protocol Plasmid DNA extraction using alkaline lysis Restriction endonuclease digestion of plasmid DNA (pdna3.1myc- His(-)B) Polymerase chain reaction (PCR) for amplification of the DNA fragment encoding vwf-a1 domain Purification of PCR product by phenol chloroform extraction and ethanol precipitation.47 XI

12 Digestion of the purified PCR products with restriction endonucleases Ligation of the DNA fragments encoding vwf-a1 domain into the expression vector Transformation of bacterial cells (E. coli TOP10) with pcdna3.1myc- His(-)B vwf-a1 recombinant vector Identification of bacterial colonies that contain recombinant plasmids High yield plasmid DNA puridfication for mammalian cell transfection Transfection of 293T human embryonic kidney cells with the pcdna3.1myc- His(-)B vwf-a1 expression vector and protein expression Affinity purification of vwf-a1 domain using Ni 2+ -affinity chromatography (Ni-NTA) Cloning, expression and purification of Escherichia coli-derived recombinant vwf-a1 fragments Construction of the pet-29c(+) vwf-a1 domain expression vector Polymerase chain reaction (PCR) for amplification of the DNA fragment encoding vwf-a1 domain Digestion of the purified PCR products with restriction endonucleases Ligation of the DNA fragment encoding the vwf-a1 domain into the expression vector Transformation of bacterial cells (E. coli TOP10) with pet- 29c(+)vWF-A1 expression vector Transformation of Escherichia coli strain BL21 (DE3) with pet- 29c(+)-vWF-A1 expression vector Overexpression of recombinant vwf-a1 protein in Escherichia coli strain BL21 (DE3) using IPTG (Isopropyl-1-thio-β-Dgalactopyranoside) Purification of recombinant vwf-a1 protein by affinity chromatography SDS-PAGE and Western blotting analysis Sample preparation 56 XII

13 A. Blood clotting FX and FXa...56 B. Lysate from cell culture Preparation of SDS-PAGE gel and electrophoresis Coomassie blue staining Western blotting analysis Statistical analysis 58 CHAPTER III RESULTS 3.1. Chromogenic substrate assay for blood clotting factors Thrombin production assay Blood clotting FXa production assay Influence of platelet concentration on the process of hemostasis Influence of activated FVII on the clotting mechanism Influence of TF on the coagulation process Thrombin formation assay in healthy people and patients with diabetes mellitus Effect of fibrin clot on thrombin and FXa generation measurment Correlation between thrombin and FXa production Influence of moderate physical exercise on thrombin and FXa production Western blotting analysis for detection of blood clotting FX and it s active form Development of western blotting analysis Western blotting analysis for the determination of zymogen FX and its activated form, the serine protease FXa Cloning, expression and purification of vwf A1 domain and study of its function in hemostasis Cloning and expression of the vwf-a1 domain in 293T human embryonic kidney cells Construction of the vwf-a1 expression vector...74 A. Amplification of the DNA fragment expressing vwf-a1 domain...74 B. Cloning of the insert into the expression vector pcdna.3.1/myc-his(-)b Protein expression and purification Western blotting analysis Cloning and expression of the A1 domain of human vwf in Escherichia coli strain BL21 (DE3) Construction of the vwf A1 expression vector...78 A. Amplification of the DNA fragment expressing vwf-a1 domain...78 XIII

14 B. Cloning of the insert into the expression vector pet-29(+) Protein expression and purification Western blotting analysis Potential function of recombinant vwf-a1 domain protein in platelet activation and the natural process of hemostasis...81 CHAPTER IV DISCUSSION 4.1. Chromogenic substrate assay for blood clotting factors Development of a chromogenic substrate assay for monitoring thrombin and blood clotting FXa production Influence of platelet concentration on the process of hemostasis Influence of activated FVII on the clotting mechanism Influence of TF on the coagulation system Thrombin activity in healthy people and patients with diabetes mellitus Effect of fibrin clot on thrombin and FXa generation measurement Correlation between thrombin and FXa production Influence of moderate physical exercise on thrombin and FXa production Western blotting analysis for the detection of blood clotting FX and its active form FXa Cloning, expression and purification of human vwf-a1 domain Functional characterization of vwf A1 gene product in platelet activation and the natural process of hemostasis.93 CONCLUSIONS 96 RERERENCES...97 Appendix (1) Appendix (2) ΠΕΡΙΛΗΨH XIV

15 LIST OF TABLES Table Page 1.1. Blood coagulation proteins and their functions Oligonucleotide sequences used to amplify the DNA fragment encoding the A1 region of vwf for subcloning it to the pcdna 3.1/myc-His vector Reagents used in the PCR amplification of the DNA fragment encoding the A1 region of vwf for subcloning it to the pcdna 3.1/myc-His vector PCR conditions used to amplify the DNA fragment coding for the A1 region of vwf for subcloning it to the pcdna 3.1/myc-His vector Oligonucleotide sequences used to amplify the DNA fragment encoding the A1 region of vwf for subcloning it to the pet-29c(+) vector Reagents used in the PCR amplification of the DNA fragment encoding the A1 region vwf for subcloning it to the pet-29c(+) vector PCR reaction conditions used for amplifying the the DNA fragment encoding the A1 region vwf for subcloning it to the pet-29c(+) vector Thrombin activity in healthy people and patients with diabetic mellitus without or under a anticoagulant treatment Determination of thrombin and FXa production before and after clot removal Platelet-dependent thrombin and FXa production before and after 15 and 120 minutes of moderate physical exercise.69 XV

16 LIST OF FIGURES Figure Page 1.1. The sequential events following vessel injury A schematic illustration of the platelet and its main functional receptors A schematic illustration of the platelet, including principal agonist receptors, and effector systems in platelet activation Simplified model of the coagulation cascade Tissue factor bearing cells in the damaged vessel wall can initiate the extrinsic coagulation cascade Cell based model of blood coagoulation The fibrinolytic system Schematic representation of the primary amino acid sequence of the two-chain human FX Structure of the the FX gene and activation of FX to FXa by the Xase complex Prothrombin activation pathway Schematic representation of the processing steps involved in vwf biosynthesis Schematic representation of the pre-pro-vwf polypeptide Structure of the vwf-a domains Model of vwf cleavage by ADAMTS Biphasic model of vwf-mediated platelet adhesion Restriction map of the pcdna 3.1/myc-His vector Flow chart for purifying plasmid DNA using the PureLink HiPure Plasmid DNA Purification DNA Restriction map of the pet-29c(+) vector Determination of continuous thrombin production in eight healthy donors Determination of average thrombin production for increasing time intervals (0-6 min, measuring every 45sec) in fifty healthy subjects Determination of discontinuous thrombin production for increasing time intervals (0-14 min, measuring every 30sec) in ten different human samples Deternination of continuous production of blood clotting FXa in nine healthy donors Determination of FXa production for increasing time intervals (0-6 min, measuring every 45sec) in ten normal subjects Influence of increasing platelet concentration on thrombin production.63 XVI

17 3.7. Comparison between continuous thrombin production in PPP and PRP in the presence of TF Determination of continuous thrombin production before and after activation of the reaction with the addition of rfviia Discontinuous assay for the determination of thrombin production for increasing time intervals (0-6 min) before and after activation of the reaction with rfviia Determination of continuous blood clotting factor FXa production before and after activation of the reaction with rfviia Discontinuous assay of FXa production at various time points before and after addition of rfviia Thrombin production with increasing concentrations of TF Determination of continuous FXa production with increasing concentrations of TF Correlation coefficient between FXa and thrombin production rates Platelet-dependent continuous thrombin production before and after 15 and 120 min of physical exercise Western blotting analysis of FX, following 12% (w/v) SDS-polyacrylamide gel electrophoresis, in 7 different samples using a specific anti-human polyclonal antibody against FX Western blotting analysis of FX, following 8% (w/v) SDS-polyacrylamide gel electrophoresis, in 7 different samples using a specific anti-human polyclonal antibody against FX Western blotting analysis of FX, following 10% (w/v) SDS-polyacrylamide gel electrophoresis, in 8 different samples using a specific anti-human polyclonal antibody targeting FX SDS-PAGE and Western blotting analysis of factors X and Xa for increasing time intervals (0, 5 and 10 min) using a specific anti-human monoclonal antibody recognizing the heavy chain of FX and FXa SDS-PAGE and Western blotting analysis of factors X and Xa for increasing time intervals (0, 2, 4, 6, 8, 10, 12, 14 and 16 min) using a specific anti-human monoclonal antibody recognizing the heavy chain of FX and FXa Western blotting analysis of FX and FXa in nine freezed and thawed samples using a specific anti-human monoclonal antibody against FX and FXa Analysis of purified, double-digested PCR products corresponding to vwf A1 domain...74 XVII

18 3.23. Analysis of BamH1/EcoR1 digested pcdna.3.1/myc-his(-)b on a 1.2% (w/v) agarose gel Analysis of purified restriction double-digested recombinant pcdna.3.1/myc-his(-)bvwf-a1 expression vector on a 1.2% (w/v) agarose gel SDS-PAGE analysis of 293T total cell extracts Purification of vwf-a1 domain recombinant protein by affinity Ni-NTA chromatography Western blotting analysis of vwf-a1 expressed in 293T HEK cells Analysis of purified double-digested PCR products corresponding to vwf-a1 domain for cloning into the pet-29c(+) expression vector Analysis of NdeI/XhoI digested pet-29c(+) on a 1.2%(w/v) (w/v) agarose gel Analysis of purified restriction double-digested recombinant pet-29c(+)-vwf-a1 expression vector on a 1.2% (w/v) agarose gel SDS-PAGE analysis of E. coli total extracts Purification of recombinant vwf-a1 domain containing a 6xHis tag by affinity Ni-NTA chromatography Western blotting analysis of vwf-a1 expressed in E. coli BL21 (DE3) cells Determination of thrombin production following incubation of 28 samples from healthy donors in the absence or presence of recombinant vwf- A1 protein Determination of continuous platelet dependent-thrombin production without or with addition of recombinant vwf-a1 protein Determination of discontinuous thrombin production in 28 healthy individuals for increasing time intervals (0-6 min, measuring every 30 s) without or with addition of recombinant vwf-a1 domain protein XVIII

19 List of Symbols and Abbreviations aa ADP apl APS APTT aptt Asn Asp AT BSA CaCl 2 camp Con CVD Cys DAG ddh 2 O EDTA FII FIX FIXa FV FVa FVII FVIIa FVIII FVIIIa FX FXa FXa FXI FXIa FXIII Amino acids Adenosine diphosphate Activated phospholipids Ammonium persulfate Activated patial thromboblastin time Activated partial thromboplastin time Asparagin Aspartic acid Antithrombin Bovine serum albumin Calcium chloride Cyclic adenosine mono phosohate Concentration Cardiovascular disease Cysteine Diacylglycerol Double distil water Ethylenediamine tetraacetic acid Prothrombin Factor IX Activated factor IX Factor V Activated factor V Factor VII Activated factor VII Factor VIII Activated factor VIII Factor X Activated FX Activated FX Factor XI Activated factor XI Factor XIII XIX

20 FXIIIa Activated factor XIII Gla domain Gamma-carboxyglutamic domain Glu Glutamic acid GP Glycoprotein HEK Human embryonic kidney HK High molecular weight kininogen HMWK High moleculer weight kininogin IIe Isoleucine IP3 Inositol triphosphate KD kilodalton KK Kallikrein Met Methionine Min Minute PAI Plasminogin activated inhibitor PAR Protein activated receptors PC Protein C PCR Polymerase chain reaction PK prekalikrin PLC Phospho lipase C PS Protein S PT Prothrombin time PT Prothrombin time RGD Arginine, Glycine, Aspartic acid rpm Revolutions per minute RT Room temperature RVV-X Russell s viper venom FX activator S-2238 Chromogenic substrate for Thrombin CBS Chromogenic substrate for FXa SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis Ser Serine XX

21 Sec(S) TBS Tem TEMD TF TFPI TFPI Thr TM TPA t-pa TXA2 Tyr UPA Xase XHC Second Tris-buffered saline Temperature Tetra ethel methyl di amine Tissue factor Tissue factor pathway inhibitor Tissue factor pathway inhibitor Threonin Thrombomodulin Tissue type plasmin activator Tissue-type plasminogen activator Thromboxan A2 Tyrosine Urokinase type plasmin activator FX-activating complex (extrinsic or intrinsic) Monoclonal antibody against the heavy chain of human FX and FXa XXI

22 CHAPTER ONE INTRODUCTION 1.1. Current concepts of hemostasis The term hemostasis, defines a dynamics process aimed at maintaining the blood fluid under physiological conditions, and also at limiting the blood loss in case of vascular lesion (1). Hemostasis is one of the most complex and efficient physiological self-defense systems and many of the specific molecular mechanisms underlying this process are still partially understood. It proceeds through the controlled activation and inactivation of clotting factors resulting in the arrest of bleeding without blocking the vasculature, and ensures the balance between the opposing undesired conditions of thrombosis and hemophilia (2, 3). Hemostasis controls the fluidity of blood, and upon vascular injury, it rapidly induces the formation of a hemostatic plug to stop or limit bleeding. As hemostasis involves a delicate balance between procoagulant and anticoagulant forces, an imbalance in this context may result in a hypercoagulable state and thrombosis, and to the interruption of blood flow due to redundant intravascular coagulation (4, 5, 6). Thrombosis, which adversely affects millions of people annually is the undesired formation of a clot within the vasculature and can result in deep vein thrombosis, pulmonary embolism, stroke or heart attack. Hemophilia is the uncontrolled bleeding and results in severe blood loss. Both thrombosis and hemophilia have contributing genetic components as well as acquired causes and better treatment methods are needed for both of these (3). The entire vasculature is lined with a single layer of endothelial cells which form the walls of the vessels and veins which normally function as an anticoagulant surface ensuring that no blockage to the vasculature occurs (7). An injury occurs when the endothelial lining of the vasculature is disrupted allowing the circulating blood to leak outside of the vasculature resulting in blood being exposed to the sub-endothelial environment. The sub-endothelial environment has many procoagulant components within it which function to promote clot formation (8). Procoagulant factors, collagen and the protein tissue factor (TF) are also essential for clot formation. Exposure of blood to collagen results in the activation of platelets and exposure of blood to TF initiates the coagulation response (9). An injury to the vasculature triggers many other responses all of which result in blockage of the injured site as a means to minimize blood loss. The combination of all of these responses is termed hemostasis and is divided into the three interconnecting processes termed primary hemostasis, coagulation and fibrinolysis (10, 11, 12). 1

23 Primary hemostasis The body s initial response to injury is mediated by the endothelial surface of a blood vessel. Primary hemostasis involves two interrelated processes. Once damage occurs the initial phase is characterized by constricting and narrowing the injured blood vessel which minimizes blood flow to the damaged area, while the formation of an initial platelet plug seals the injured site (13, 14). Vasoconstriction is the first response to an injury that occurs in a damaged blood vessel resulting in narrowing the vessel to limit the flow of blood to the damaged area and minimization of blood loss (15). As soon as platelets detect any deviation from the normal conditions of the vessel they react by arresting their circulation in the flowing blood, followed by permanent adhesion at the damaged vessel wall (Figure1.1b). The result is the formation of a primary platelet plug soft plug that initially stops the bleeding (Figure 1.1c) (16, 17). In primary hemostasis, platelets adhere to the sub-endothelium, become activated and aggregate to each other to form a platelet plug that temporarily stops the blood flow (8, 15). Activated platelets release the contents from their interior alpha and dense granules, thus secreting the vasoconstrictors serotonin, prostaglandin and thromboxane (18). An enzymatic flipping of inner leaflet phosphatidylserine phospholipids to the outer leaflet upon activation of platelets creates an enzymatic surface capable of sustaining the formation of enzymatic complexes composed of coagulation factors which are attracted to the negatively charged surface (19, 20). The activation of platelets also results in the release of many molecules from the interior of the platelet needed for clot formation, including fibrinogen, FV and vwf (21). All sub-endothelial tissues express the cellular membrane protein TF which is exposed to circulation when the endothelial layer is disrupted. The exposure of TF to circulation is the initiating event in the coagulation portion of hemostasis, termed secondary hemostasis (19, 20, 21). 2

24 Figure 1.1: The sequential events following vessel injury: a) Schematic illustration of the vessel, b) platelet adhesion to collagen in the sub-endothelial matrix, c) platelet aggregation and activation; forming a platelet plug that initially stops bleeding, d) fibrin network formation (coagulation) stabilizes the platelet plug The platelets and its main functions in primary hemostasis Platelets or thrombocytes (from Greek θρόµβος, "clot" and κύτος, "cell"), are small disk shaped non-nucleated blood cells with a very fragile membrane derived from the cytoplasm of megakaryocytes that reside in the bone marrow. Platelet production is controlled by a hormone, thrombopoietin, and regulatory lymphocytes acting at the stem cell level. At any given time about one-third of the total blood platelets can be found in the spleen, the remaining two-thirds are in circulating blood (22, 23). Platelets are the second most abundant cellular component in healthy blood, and the number of normal platelets in a healthy individual varies between 150,000 and 450,000 per µl of blood (24). The average lifespan of a circulating platelet is normally 5 to 9 days. Platelets do not show any interaction with the inner surface of normal vessels but adhere promptly where endothelial cells are altered or extracellular matrix substrates are exposed (25). This is a critical initial step in hemostasis and thrombosis, as well as in inflammatory and immunopathogenic responses (26, 27, 28). They are released into the circulation to survey the integrity of the vascular system, and respond to lesions by forming aggregates adherent to damaged endothelial cells or to extracellular matrix structures exposed to flowing blood. Following activation, platelets provide a catalytic membrane surface for thrombin generation, which accelerates the 3

25 formation of fibrin necessary to stabilize thrombi; they also promote cellular and humeral events responsible for wound healing. Platelet responses cannot distinguish between traumatic and pathological vessel damage (23); thus, although their normal function is to contribute to the arrest of bleeding from wounds, they may form occlusive thrombi as a consequence of vascular diseases, such as atherosclerosis, resulting in ischemic syndromes of the heart, brain and other organs. The platelet surface is packed with functional receptors to facilitate platelet activation and interactions with sub-endothelial matrix, other blood cells and other platelets. The platelet also contains several granules that upon activation can fuse with the platelet membrane and thereby release their content into the surrounding media (see Figure 1.2). The granules are categorized into larger α-granules, smaller dense granules and lambda granules similar to lysosomes (29). α- granules contain a large variety of biological active substances involved in hemostasis, e.g. platelet activation and adhesion molecules, plasma coagulation factors and fibrinolysis proteins (fibronectin, vwf, fibrinogen, and coagulation factors V and XIII). The dense granules mainly contain ions and smaller signal molecules such as adenosine 5 - diphosphate (ADP), adenosine 5 -triphosphate (ATP), calcium and serotonin. Lambda granules similar to lysosomes contain several hydrolytic enzymes like acid hydrolase, cathepsin and lysosomal membrane proteins (30). The platelets are very sensitive towards external stimuli and can undergo a fast activation process that is enhanced by intrinsic activation feedback loops in the form of autocrine and paracrine signaling (31). 4

26 Figure 1.2: A schematic illustration of the platelet and its main functional receptors. Platelets adhere to collagen in the damaged vessel wall through adhesion receptors. The adhesion receptors can contribute to platelet activation and thereby stimulate the release of autocrine activators such as ADP and TXA 2. Upon activation the platelets are capable of forming platelet aggregates by linking to fibrinogen molecules with the GPIIb/IIIa receptor. Thrombin produced by activation of the coagulation cascades can also activate PAR1 and PAR4 by proteolysis of the receptor (32). A. Platelet adhesion The physiological protection against bleeding is secured by platelet adhesion to the site of injury and sealing of the defect. The adhesion of platelets to the denuded areas of endothelium is one of the earliest events in homeostasis (33). Collagen is the main component of the sub-endothelium matrix and is also considered to play a principal role in the adhesion process. Nine types of collagen have been identified in the vessel wall but the main constituents have been determined as collagen type I and type III (34). The collagen receptors on the surface of platelets may be divided into those which interact indirectly with collagen-bound vwf including GPIbα and integrin αiibβ 3 (GPIIb/IIIa), and those 5

27 which interact directly with collagen, including glycoprotein VI (GPVI), integrin α 2 β 1 (GPIa/IIa) and CD36 (16, 35, 36), (see Figure 1.2). The local flow conditions are also important in determining the platelet behavior. Blood is always moving inside the vessels but the conditions can vary from almost stagnant to extremely high flow rates, In areas of high shear rate (in the microvasculature) platelet adhesion is mediated by vwf which binds to GPIb-IX-V in the platelet membrane. In areas of low shear rate (e.g. aorta) collagen mediates the binding of platelets to the sub-endothelium by attaching to a platelet receptor - the integrin GPIa/IIa (37, 38). Primary platelet adhesion receptors include the GPIb-IX-V complex of the leucine-rich repeat family, and GPVI of the immunoglobulin super family. The initial tetaring and arrest of platelets to collagen is facilitated by the GPIb-IX-V receptor, a heterotrimer composed of two disulfide linked GPIbα (~135 kda) - GPIbβ (~25 kda) associated with GPIX (~20 kda) and GPV(~85 kda) at a ratio of 2:2:2:1 (39, 40, 41). GP Ib-IX-V receptor is the only platelet receptor that does not require prior activation for bond formation (16), and the binding is mediated by vwf. Upon binding GPIb-IX-V is capable of outside-in signaling leading to platelet activation (42). Platelet GPVI is related to members of the T-cell receptor family and is a member of the immunoglobulin super family. GPVI is constitutively expressed in platelets, but in vivo, it is only engaged when collagen is exposed in the sub-endothelial matrix following disruption of the endothelium (43, 44). The GPVI receptor binds directly to collagen via the specific Glycine-Proline- Hydroxyproline peptide repeat sequence (Gly-Pro-Hyp) and has an important role in platelet activation by outside-in signaling. GPVI activation has been found to be mediated by the Fc receptor γ-chain, also involved in platelet interaction with immunoglobulins (45). The integrin α 2 β 1 adhesion receptor can bind to collagen directly. Although it has been indicated that the α 2 β 1 integrin is not involved in the initial tethering of platelets to collagen under high shear, it is still considered important for firm adhesion and securing the platelets (46, 47). The integrin adhesion receptor α 2 β 1 has also been proposed to be involved in platelet spreading on the collagen surface (48). The integrin α 2 β 1 receptor must be activated by insideout signaling and attain a state of high affinity for its ligand (49). The two collagen receptors GPVI and α 2 β 1 are both essential for platelet adhesion and aggregation on collagen in flow conditions. Blockade of either receptor will abolish the platelet plug forming capabilities (50). 6

28 B. Platelet aggregation Platelet aggregation is the clustering of thrombocytes. The linking of the platelets via fibrinogen leads to platelet aggregation. A great number of agents (including ADP, epinephrine, collagen and thrombin) can induce platelet aggregation (51). Simplistically, vwf and fibrinogen bind to receptors on one platelet leading to cross-linking to another platelet by binding to the respective receptors of the latter (52). Once a primary layer of adhesive platelets have covered the exposed sub-endothelial matrix, subsequent adhesion will continue but in the form of aggregation, where platelets from flowing blood bind to adhered platelets at the wound site. The mechanism of platelet aggregation is mainly attributed to the integrin αiibβ3 (GPIIb/IIIa) receptor. The main ligand of integrin αiibβ 3 is fibrinogen but the receptor also has affinity for multimeric vwf, vitronectin, fibronectin and thrombospondin (53). The mechanism of aggregation involves two integrin αiibβ3 receptors on different platelets that bind to the same fibrinogen molecule. The αiibβ3 receptor has a low and a high affinity state for binding fibrinogen. The high affinity state is achieved by inside-out signalling when platelets are activated. The αiibβ3 receptor can also mediate outside-in signalling by binding fibrinogen (54, 55). C. Platelet activation After initial adhesion of platelets to the extracellular matrix, intracellular signaling occurs and the platelets become activated, the discoid platelets then rapidly undergo morphological modifications and become spherical, their granules become centralized and come into contact with the membrane invaginations, leading to the secretion of active substances like ADP and TXA2 (56). The propagation of the activated state of the adhered and aggregated platelets is a prerequisite for the recruitment of other platelets during the aggregation phase. This propagation is executed via potent autocrine and paracrine mediators, including ADP, thrombin, epinephrine, and TXA2. (57, 58). These mediators amplify, sustain the initial platelet response (Figure 1.2) and recruit circulating platelets from the flowing blood to form a growing hemostatic plug. Most agonists that activate platelets operate through G-protein coupled receptors (52). The final pathway for all agonists is the activation of the platelet integrin GPIIb/IIIa (αiibβ 3 ), the main receptor for adhesion and aggregation (59). The activation of platelets is induced by the interaction of several agonists with receptors expressed on the platelet membrane. Figure 1.3 (panels A, B, and C) depicts outside-in signaling mediated by TXA2, ADP and thrombin, respectively (60). 7

29 A lipid signalling molecule TXA2 is synthesized by activated platelets from arachidonic acid through the cyclooxygenase pathway (Panel A). Once formed, TXA 2 can diffuse across the membrane and activate other platelets. In platelets, there are two splice variants of the TXA 2 receptor: TPα and TPβ receptors (61), which differ in their cytoplasmic tail. TPα and TPβ couple to the proteins Gq and G12 or G13, all of which activate phospholipase C (PLC). This enzyme degrades the membrane phosphoinositides (such as phosphatidylinositol 4, 5-bisphosphate), releasing the second messengers inositol triphosphate (IP3) and diacylglycerol (DAG). DAG activates intracellular protein kinase C (PKC), leading to protein phosphorylation. The release of IP3 increases cytosolic levels of Ca +2, which is released from the endoplasmic reticulum (60, 62). The first known low molecular weight platelet aggregating agent ADP, is a weak platelet agonist. As such, it induces only platelet shape change and reversible aggregation in human platelets. Activated platelets will secrete abundantly ADP and ATP from the dense granule (63). Platelets express at least two ADP receptors, P 2 Y 1 (seven transmembrane spanning protein) (64,65) that transmits outside-in signaling by coupling to G proteins and P 2 Y 12 (a ligand gated cation channel), which couples to Gq and Gi, respectively (Figure 1.3, Panel B). The activation of P 2 Y 12 inhibits adenylate cyclase, causing a decrease in the cyclic AMP (camp) level and the activation of P 2 Y 1 causes an increase in the intracellular Ca 2+ level. The P 2 Y 12 receptor is the major receptor able to amplify and sustain platelet activation in response to ADP (63, 65). The key coagulation enzyme thrombin generated as a result of simultaneous activation of the coagulation cascade, during thrombus formation has a dual role in hemostasis. Besides mediating fibrin network generation, it represents the most potent platelet activator (Figure 1.3, panel C). Platelet responses to thrombin are largely mediated through G-protein linked protease-activated receptors (PARs), which are activated after thrombin-mediated cleavage of their N-terminal exodomain. The cleavage results in a new N-terminus that functions as a ligand and thereby mediates the activation of the receptor (66, 67). Human platelets express PAR1 and PAR4 that apparently differ in terms of affinity for thrombin and the duration of intracellular signalling. It has been proposed that PAR1 has a higher affinity for thrombin than PAR4 (68, 69). PAR1 couples to members of the G 12 / 13, Gq, and Gi protein families. The α-subunits of G 12 and G 13 bind Rho guanine-nucleotide exchange factors (Rho GEFs), leading to Rho-mediated cytoskeletal responses, that are probably involved in the change in platelet shape. The Gq and Gi signaling pathways lead to increased intracellular Ca 2+ and decreased camp, respectively. Figure 1.3 (Panel D) depicts inside-out signaling. The effects of agonists mediated by the decrease in camp levels and increase in intracellular Ca +2 levels lead to platelet aggregation through the change in the ligand-binding properties of 8

30 GPIIb/IIIa (αiibβ3), which acquires the ability to bind soluble adhesive proteins such as fibrinogen and vwf. The release of ADP and TXA 2 induces further platelet activation and aggregation. The small-peptide sequence arginine-glycine-aspartic acid (RGD) of the adhesive proteins binds to the GPIIb/IIIa receptor. Fibrinogen contains two RGD sequences in its α-chain, one in the N-terminal region and the other in the C-terminal region (60). Following activation, platelets provide a catalytic membrane surface for thrombin generation, which accelerates the formation of fibrin necessary to stabilize thrombi; they also promote cellular and humoral events responsible for wound healing and, possibly, for the local control of bacterial infections (70,71). Figure 1.3: A schematic illustration of the platelet, including principal agonist receptors, and effector systems in platelet activation (60) Coagulation (Secondary hemostasis) The second stage in the hemostatic plug formation is the coagulation process, which represents one of the fundamental vital functions of a living organism (1,16). It is a complex process triggered by cellular elements (platelets, white blood cells and endothelial cells) and involves a stepwise participation of a large number of plasma proteins that progressively amplify the triggering signal in order to massively generate thrombin and finally lead to fibrin formation 9

31 (72). Newly formed thrombin is necessary to stabilize the primary hemostatic plug. Fibrin then stabilizes the platelet aggregate that occludes the vessel injury and prevents deleterious leakage of blood (73, 74). Platelets are actively involved in secondary hemostasis; they generate the negatively charged catalytic surface needed for the assembly of all enzymatic complexes (8). The enzymatic proteins involved in coagulation (factor VII [FVII], factor IX [FIX], factor X [FX], factor XI [FXI], factor XII [FXII] and prothrombin [PT]) are all vitamin K-dependent serine proteases, which circulate in plasma as inactive precursors (zymogens), and they need to be activated in order to participate in the coagulation reactions. To increase the catalytic efficiency of these serine proteases, allowing a rapid response to injury, the coagulation system also provides three cofactors: tissue factor (TF), factor V (FV) and factor VIII (FVIII). For a long period, the blood coagulation has been described as a process where each clotting factor as proenzyme could be converted to an active enzyme, mediating then the activation of another one (waterfall model). In particular, this classical view divided the coagulation process into two pathways: an intrinsic and extrinsic pathway. The initiation of both pathways finally results in the activation of FX and the generation of thrombin (73) (Figures 1.4 and 1.5) The pathways of the coagulation cascade The coagulation system can be viewed as a cascade of proteolytic reactions in which zymogens are cleaved to produce active proteins (1, 14, 75). According to the traditional view, the coagulation cascade can be schematically divided in two pathways, one involving physical trauma to the vessel, resulting in the exposure of sub-endothelial cell membrane protein (TF) to the blood and triggering the so called extrinsic pathway, and the other initiated, when FXII is activated or released during platelet degranulation, leading to thrombin generation in the absence of vessel injury, the so called intrinsic pathway (76, 77) The names of intrinsic and extrinsic pathways are derived from the fact that all functional components of the intrinsic pathway derive from blood, whereas the triggering component of the extrinsic pathway, tissue factor, is not found in blood but in extravascular tissue. Although there are major differences in the mechanism and the subsequent enzymatic steps, both the intrinsic and extrinsic pathways converge into a common pathway, ultimately leading to the formation of the fibrin network (78) (Figures 1.4 and 1.5). 10

32 A. Intrinsic Pathway (Factor XII - Contact activation pathway) The intrinsic pathway consists of a cascade of protease reactions initiated by factors that are present within the blood when in contact with a negatively charged phospholipid surface of vascular endothelial cells and platelets (8). During the intrinsic pathway (Figure 1.4) the initial activator plasma protein FXII turns into FXIIa. High molecular weight kininogen (HMWK), a product of platelets mediates the anchoring of FXII to the charged surface, thus serving as a cofactor. However, HMWK-assisted conversion of FXII to FXIIa is limited in speed, yet once a small amount of FXIIa accumulates, this protease converts prekallikrein (PK) to kallikrein, using HMWK as an anchor. In turn, the newly produced kallikrein accelerates the conversion of FXII to FXIIa (79). In addition to amplifying its own generation by forming kallikrein, FXIIa (together with HMWK) proteolytically cleaves FXI, forming FXIa. In turn, FXIa (also bound to the charged surface by HMWK) proteolytically cleaves FIX to FIXa, which is also a protease. FIXa and two downstream products of the cascade, FXa and thrombin, proteolytically cleave FVIII, thus forming FVIIIa, a cofactor in the next reaction. Finally, FIXa and FVIIIa together with Ca +2 and negatively charged phospholipids form a trimolecular complex termed tenase, which then converts FX to FXa (80). In a parallel series of interactions, FXa binds to the cofactor FVa, a downstream factor that participates in positive feedback, This association generates a complex with enzymatic activity known as prothombinase. Prothombinase converts the proenzyme prothrombin to its enzyme form thrombin. Thrombin acts on fibrinogen to generate the fibrin monomer, which rapidly polymerizes to form the fibrin clot. During clinical laboratory analysis of blood clotting, the intrinsic pathway of blood coagulation is evaluated using the activated partial thromboplastin time (APTT). Deficiency of any of the blood clotting factors prolongs the APTT assay time in vitro (81). 11

33 Figure 1.4: Simplified model of the coagulation cascade. The classical intrinsic pathway, with the sequence of activation cascade proceeding from HMHK and PK (17, 82). B. Extrinsic pathway (Tissue factor pathway) Extrinsic pathway is the main contributor to the physiological activation of coagulation in vivo. The extrinsic pathway includes protein cofactors and enzymes and an integral membrane glycoprotein, TF, that is believed to be the physiological initiator of the extrinsic pathway of blood coagulation (83, 84). The extrinsic pathway (Figure 1.5) is initiated after vascular damage by the formation of a complex between TF and FVIIa on the cell surface that is located outside the vascular system. Nonvascular cells constitutively express the integral membrane protein TF (variably known as FIII or tissue thromboplastin) (83), which serves as a receptor for the vitamin K-dependent proenzyme. When an injury to the endothelium occurs, FVII binds, via ϒ-carboxyglutamic acid residues and Ca +2 bridges, to its cofactor TF on cell membranes. This binding leads to a non proteolytical activation of the zymogen FVII to its active form, which is the serine protease FVIIa (1, 14, 75). The mechanism of the initial conversion of the zymogen FVII to FVIIa is still debated but is most likely due to autocatalytic activation and not a TF effect (85). Binding of 12

34 FVIIa to TF results in the formation of an enzyme extrinsic FX-activating complex (extrinsic tenase complex) that activates FX to FXa. In addition, FVII can also be activated by FXa and thrombin (86, 87) The FVIIa/TF complex, similar in function to the tenase complex, converts FX to its active form (FXa), which first binds to the cofactor FV and then to membrane surfaces in the presence of Ca +2 ions to generate the prothrombinase complex. The prothrombinase complex converts prothrombin to thrombin, which converts fibrinogen to fibrin clot (79, 81). Consequently, the TF-FVII complex will also activate FIX that will play a role in the intrinsic pathway (87). The extrinsic pathway is rapidly inhibited by TF pathway inhibitor (TFPI), a serine protease inhibitor that is associated to lipoprotein molecules. Although this pathway is inhibited quickly, sufficient quantities of thrombin are formed to activate FXI. FXIa will participate in the intrinsic pathway, inducing the amplification of the coagulation response (88, 89). During laboratory analysis of blood clotting, the extrinsic pathway of blood coagulation is evaluated using the prothombin time (PT). Regardless of whether FXa arises by the intrinsic or extrinsic pathway, the cascade then proceeds along the common pathway (79, 81). Activation of the extrinsic pathway by tissue factor and the subsequent enzymatic coagulation cascade are visualized in Figure 1.5 (32). 13

35 Figure 1.5: Tissue factor bearing cells in the damaged vessel wall can initiate the extrinsic coagulation cascade. Small amounts of thrombin are produced by FXa which in turn will activate part of the intrinsic pathway (FXIa and FXIa/FVIIIa complex) leading to a large scale production of the very effective prothrombinase (FXa/FVa) complex on the surface of activated platelets. The prothrombinase complex will generate massive amounts of thrombin from prothombin. Thrombin cleaves off the fibrinopeptides from the fibrinogen molecule, making the molecule able to polymerize into a fibrin fibre. Finally the fibrin fibres are stabilized by the cross-linking activity of FXIIIa (32) Cell based model of coagulation A major development over the past 15 years was the discovery that exposure of blood to cells that express TF on their surface is both necessary and sufficient to initiate blood coagulation in vivo. This finding led to the assumption that the intrinsic pathway does not have a true physiological role in hemostasis, however it could not provide an explanation for the haemostatic mechanisms in vivo (90, 91). For example, patients that are deficient in FXII, HMWK or PK (91, 92), exhibit, as expected, a prolonged APTT upon coagulation analysis. Yet, these patients do not show a bleeding tendency. The APTT test is indicative of dysfunctional factor activity or level, especially in the intrinsic pathway. In patients with FXI deficiency, there 14

36 is an increased risk of hemorrhage but the APTT is not indicative of the extent to which the patient may bleed even though the bleeding is markedly lower than that seen in haemophilia A or B (93). A cell based model has recently been proposed that highlights the role of different cells with similar phosphatidylserine contents which, however, are able to play different roles in hemostasis (90, 92). In the cell-based model, hemostasis requires the formation of an impermeable platelet and fibrin plug at the site of vessel injury, but it also requires the localization of the procoagulant substances activated in this process to the site of injury (94). The new cell-based model provides explanations for the varying degrees of hemorrhage encountered in the presence of factor deficiencies in both the extrinsic and intrinsic pathways. It explains why even though the extrinsic pathway is functional in hemophilics there is still pronounced bleeding. In addition, it explains why the extrinsic pathway cannot sustain adequate clot formation to prevent bleeding in hemophiliacs. It acts as a stepping stone providing answers to some of the questions surrounding various other deficiencies in hemostasis and further highlights that coagulation occurs in three different stages on three different cell surfaces. Initiation occurs on a TF-bearing cell where as in the amplification stage, platelets and cofactors assemble to prepare for the large thrombin burst and the propagation stage occurs on the surface of activated platelets (90, 92). This hypothesis and the experimental evidence supporting it were presented in a series of articles authored by a group from the department of pathology at Duke University and the University of North Carolina (79). A. Initiation phase All evidence to date indicates that the sole relevant initiator of coagulation in vivo is TF. Cells expressing TF are generally localized outside the vasculature, which prevents initiation of coagulation under normal flow circumstances with an intact endothelium. Some circulating cells (eg, monocytes or tumor cells) may express TF on their membrane surface, but this TF under normal conditions is thought to be inactive or encrypted. The exact role of this blood-borne TF is controversial. Some investigators believe that circulating TF is encrypted in that it contains an additional bond that must be cleaved for activity in contrast to others who believe that circulating TF is not fully active because the membrane surface on which it resides is not a phospholipid containing procoagulant membrane (95, 96, 97). Once an injury occurs and the flowing blood is exposed to a TF-bearing cell, FVIIa rapidly binds to the exposed TF. Factor VIIa is the only coagulation protein that routinely circulates in 15

37 the blood in its active enzyme form, with approximately 1% of total FVII circulating as FVIIa. All other coagulation proteins circulate solely as zymogens. The TF-FVIIa complex then activates additional FVII transforming it to FVIIa (96, 97). The FVIIa/TF complex activates small amounts of FIX and FX (98). FXa associates with its cofactor (FVa) and forms a prothrombinase complex on the surface of the TF-bearing cell. FV can be activated by FXa or by noncoagulation proteases to produce FVa required for prothrombinase assembly (79, 99, 100) (see Figure 1.6a). B. Amplification Phase The small amount of thrombin generated on the TF bearing cell has several important functions. A major function is the activation of platelets, exposing receptors and binding sites for activated clotting factors (101, 102). Although platelets have already adhered at the site of injury and become partially activated, as a result of this activation, they release partially activated forms of FV onto their surfaces. Another function of the thrombin formed during the initiation phase is the activation of the cofactors V and VIII on the platelet surface (103, 104). In this process, the FVIII/vWF complex is dissociated, permitting vwf to mediate additional platelet adhesion and aggregation at the site of injury. In addition, small amounts of thrombin activate FXI and FXIa on the platelet surface during this phase (79, 101, 102) (see Figure 1.6b). C. Propagation phase Once a number of platelets get activated during the amplification phase, the release of their granule contents results in recruitment of additional platelets to the site of injury. The propagation phase occurs on the surface of these platelets. Expression of ligands on their surface results in cell-cell interactions that lead to aggregation of platelets. FIXa that was generated by TF-FVIIa in the initiation phase can bind to FVIIIa (generated in the amplification phase) on the platelet surface. Additional FIXa is generated due to cleavage of FIX by FXIa that was generated during amplification on the platelet surface. Once the intrinsic tenase complex is formed (FIXa- FVIIIa) on the activated platelet surface, it rapidly begins to generate FXa on the platelet. FXa was also generated during the initiation phase on the TF-bearing cell surface. As this FXa is rapidly inhibited if it moves away from the TF-bearing cell surface, it can not easily reach the platelet surface. The majority of FXa must therefore be generated directly on the platelet surface through cleavage by the intrinsic tenase complex. The FXa generated on platelets then rapidly binds to FVa (generated by thrombin in the amplification phase) and cleaves prothrombin to thrombin (Figure 1.6c). This prothrombinase activity results in a burst of thrombin generation leading to cleavage of fibrinopeptide A from fibrinogen. The generation of 16

38 enough thrombin within a certain period of time is required to result in a critical mass of fibrin. These soluble fibrin molecules will spontaneously polymerize into fibrin strands, resulting in an insoluble fibrin matrix (97). Once a fibrin/platelet clot is formed over an area of injury, the clotting process must be limited to avoid thrombotic occlusion in adjacent normal areas of the vasculature. The protein C(PC)/protein S(PS)/thrombomodulin (TM) system is an important mechanism by which the production of procoagulant activity is confined to a site of injury. In short, the endothelial thrombin/tm complex activates PC, which binds to its cofactor PS and inactivates FVa and FVIIIa. Activated PC thus terminates thrombin generation on the activated platelets, while tissue factor pathway inhibitor (TFPI) shuts down TF-FVIIa once coagulation has been initiated (105). Figure 1.6: Cell based model of blood coagoulation. a. Amplification of coagulation. b. Initiation of coagulation. c. Propagation stage of coagulation. 17

39 Fibrinolysis After the hemostatic system has played its role and the vessel integrity is restored by vessel repair mechanisms, the clot needs to be cleared in order to regain unobstructed blood flow. This is taken care of by the fibrinolytic system, which protects the body from potentially hazardous thrombi by disrupting fibrin network into small soluble fragments that can be transported away by the blood flow (32, 106, 107). Fibrinolysis is itself regulated by a number of control systems that share similarities with the coagulation cascades and are controlled by a series of procoagulant and anticoagulant serine proteases (108). The principle fibrinolytic enzyme is plasmin, generated by partial proteolysis of a fibrin bound zymogen (plasminogen) which is adsorbed from the plasma, via its lysine binding sites, onto the lysine groups in fibrin polymers (109, 110, 111). Plasmin is generated from the inactive precursor plasminogen by the action of two plasminogen activators: urokinase type plasminogen activator (UPA) and tissue type plasminogen activator (t-pa) (112). In the circulation, the principle activator of plasminogen is t-pa, a serine protease released from endothelial cells which has a high binding affinity for plasminogen bound fibrin which in turn is generated by thrombin and venous occlusion (111, 113, 114). t-pa and plasminogen both bind to the evolving fibrin polymer. Once plasminogen is activated to plasmin it degrades the fibrin network at specific lysine and arginine residues in a variety of ways resulting in soluble fibrin degradation products with different molecular masses (see Figure 1.7). The plasminogen activators are in turn regulated by plasminogen activator inhibitors (PAIs) (113), the most important of which is called plasminogen activator inhibitor 1 (PAI1) which is synthesized in hepatocytes, and has the capacity to decrease plasminogen binding to fibrin (108). Aberrant fibrinolysis is also prevented by circulating direct plasmin inhibitors such as antiplasmin and α2-macroglobulin, the latter of which can act as an inhibitor of both coagulation and fibrinolysis. Another regulator of fibrinolysis is fibrinolysis inhibitor (TAFI) that gets activated by thrombin (115). 18

40 Figure 1.7: The fibrinolytic system. In addition to the illustrated mechanisms fibrinolysis is also regulated by TAFI, a carboxypeptidase that can inhibit fibrinolysis by destroying potential binding sites for plasminogen to the surface of fibrin. (Modified from Wiman, MFR informerar 198) (116) Proteins involves in blood coagulation processes (blood clotting factors and related substances) Coagulation factors, are a series of proteins present in the blood which are interrelated through a complex cascade of enzyme-catalyzed reactions involving the sequential cleavage of large protein molecules to produce peptides, each of which converts an inactive zymogen precursor into an active enzyme, leading to the formation of a fibrin clot. They are traditionally numbered using roman numerals from I to XIII, to distinguish between an activated coagulation factor from the zymogen, the activated factor number is suffixed with an a (117). The human clotting cascade relies on a group of serine proteases with some exceptions, for example, FVIII and FV are glycoproteins (non enzymatic protein cofactors), and FXIII is a transglutaminase (118). Serine proteases belong to a sub-family known as the vitamin K-dependent serine proteases, and are given their name because their active site contains serine residues that are responsible for the enzymatic activity of the enzyme. The active site of serine proteases contains three particular amino acids (histidine, aspartate and serine ) whose special relation to each other is maintained in the three dimensional structure of the enzyme. Serine proteases hydrolyze targeted peptide bonds through a nucleophilic attack by the serine residue located in the active site of the protease, forming an acyl-enzyme intermediate (119, 120). 19

41 The vitamin K-dependent serine proteases of human blood coagulation cascade are typically synthesized and secreted as zymogens, requiring proteolysis for activation. During activation, these enzymes are post-translationally gamma carboxylated at numerous γ- carboxy glutamic acid residues in a vitamin K-dependent reaction, resulting in a stereotypical N-terminal domain called gamma-carboxyglutamic (Gla) domain (117). In this modification reaction, vitamin K acts as a cofactor for carboxylases which are the responsible enzymes for the gamma carboxylation of these specific glutamic acid residues. The subsequently formed Gla domain is required for the coagulation proteases to mediate the binding of the enzyme to phospholipid bilayers in a Ca +2 -dependent manner. Inhibition of the γ-carboxylation of coagulation factors results in drastic attenuation of coagulation, and has been used for a long time in anticoagulation therapy by the administration of warfarin (118, 121). Coagulation is a very complicated process requiring a balance between factors that promote clot formation and factors that either prevent clot formation or dissolve it when it occurs. These are referred to as factors which are procoagulants and anticoagulants. Table 1.1 shows coagulation factors and related substances and their functions (118, 122). 20

42 Table 1.1: Blood coagulation proteins and their functions (123) Number and/or name FI (Fibrinogen) FII (Prothrombin) FIII (TF) FIV (Ca +2 ) Factor V (Proaccelerin) FVII (Stable factor, proconvertin) FVIII (Antihemophilic factor A) FIX (Antihemophilic factor B) FX (Stuart-Prower factor) FXI FXII (Hageman factor) FXIII (Fibrin-stabilizing factor) vwf Prekallikrein (Fletcher factor) HMWK(Fitzgerald factor) Fibronectin Antithrombin III Protein C Protein S Plasminogen Alpha 2-antiplasmin t-pa Urokinase PAI Function Clot formation (Fibrin) Its active form (IIa) activates FI, FV, FVII, FVIII, FIX, FXI, FXIII, protein C, platelets Initiator; Cofactor of FVIIa Binding of coagulation factors to phospholipid Prothrombinase complex formation FIX &FX activation Tenase complex formation Tenase complex formation Prothrombinase complex formation FIX activation Activates FXI and prekallikrein Crosslinks fibrin Mediates platelet adhesion & Binds to FVIII, Activates FXII Activation of FXII, FXI, and prekallikrein Mediates cell adhesion Inhibits FIIa, FXa, and other proteases Inactivates FVa and FVIIIa Cofactor for activated protein C Converts to plasmin, lyses fibrin Inhibits plasmin Activates plasminogen Activates plasminogen Inactivates t-pa & urokinase (endothelial PAI) Blood clotting factor X Human coagulation FX also called Stuart-Prower factor (EC ) is one of the vitamin K-dependent serine proteases of the coagulation cascade. It is synthesized in the liver as a single chain 488 amino acid (aa) zymogen precursor, possessing both a pre- and pro- peptide leader sequence (40 aa) that is cleaved to form the mature protein (Figure 1.8) (118). 21

43 The pre- peptide sequence is cleaved by a single peptidase and the pro- peptide sequence is then cleaved by a processing protease. After translation, the zymogen is gamma carboxylated at 11 Glu residues in the initial 39 residues of the peptide, thus forming a Gla domain. This posttranslational modification is mediated by vitamin K-dependent carboxylase and is necessary for the binding to phospholipids on platelet membranes (118, 119). Other post-translational modifications include β-hydroxylation of Asp63 in the first epidermal growth factor (EGF)-like domain and glycosylation of the activation peptide at Thr159 (O-linked), Thr171 (O-linked), Asn181 (N-linked), and Asn191 (N-linked) (124, 125). Human FX is heavily glycosylated possessing approximately 15% carbohydrate. It is postulated that the carbohydtare may play a role in substrate recognition or have an effect on enzymatic catalysis rates (126). In plasma, FX circulates as a two chain glycoprotein (~59 kda) (127, 128), which is composed of a 306-residues heavy chain that is covalently linked through a disulfide bond to a 139-residue light chain (see Figure 1.9) (14, 129). The light chain of zymogenic FX(12kDa), which is cleaved from the heavy chain during or after secretion into the circulation, is composed of a gamma carboxyglutamic acid (Gla)-rich domain containing 11 Glu residues and two epidermal growth factor (EGF)-like modules with a β-hydroxyaspartic acid residue in the N-terminal EGF1 domain, while the heavy chain contains the activation peptide and the serine protease domain (Figure 1.9) (124, 127, 128, 130). The mature zymogenic form of human FX is found at a con of approximately 170 nm in the blood with a half life of approximately 1.5 days. The protease domain of FX, found in the heavy chain, contains the serine protease catalytic triad of His57, Asp102 and Ser195 residues and a trypsin-like specificity pocket (119). Activation of FX (~59 kda) to FXa (~46 kda) is achieved by the cleavage of a 52 amino acid glycopeptide from the N-terminus of the heavy chain (Ser143-Arg194). Additional autoproteolysis of FXa at the Arg429-Gly430 peptide bond leads to the removal of a small peptide from the C-terminus of the heavy chain and converts the alpha form of FXa to the β form. However, no difference in function has yet been observed between the two forms (131). The proteolytic cleavage of the activation peptide occurs at the bond between Arg195 and IIe196 residues and is catalyzed by the FIXa FVIIIa-Ca +2 -phospholipid quaternary complex or the FVIIa-TF-Ca +2 - phospholipid quaternary complex (Figure 1.9) (117, 132). Once enough FXa is generated through the extrinsic and intrinsic pathways, the newly generated FXa comprises the enzymatic component of the prothrombinase complex which is responsible for the conversion of prothrombin to thrombin at the site of vascular injury. Both prothrombin and FXa bind apl in a Ca +2 dependent manner, while the FVa:aPL interaction does not require Ca +2 (40, 41, 131, 133). While FXa can cleave prothrombin in the absence of FVa, 22

44 apl and Ca +2, within the context of the prothrombinase complex, incorporation of FVa into the prothrombinase complex results in a reversal of cleavages and a 300,000-fold increase in the catalytic efficiency of FXa (134). Figure 1.8: Schematic representation of the primary amino acid sequence of the two-chain human FX. The tripeptide Arg140-Lys141-Arg142 that connects the light chain to the activation peptide is not shown because the form that lacks the tripeptide is predominant in circulating blood plasma. Individual domains are shown in boxes, functionally important catalytic residues are circled (129). 23

45 Figure 1.9: Structure of the FX gene and activation of FX to FXa by the Xase complexe. Exon I codes for the signal peptide (SP); exon II encodes the propeptide (PP) and the Gla domain; exon III codes for a short aromatic stack; exons IV and V encode epidermal growth factor-like (EGF) domains 1 and 2, respectively; exon VI codes for the activation peptide (AP); and exons VII and VIII encode the serine protease domain. B) FX structure and activation. FX circulates as a two-chain disulfide linked zymogen. During activation, either the extrinsic or intrinsic Xase complex cleaves after Arg195 to release the activation peptide from the N-terminus of the heavy chain (134) Prothrombin/Thrombin (FII/FIIa) The glycosylated trypsin-like serine protease thrombin (EC ) is a central enzyme in hemostasis, interacting with a multitude of substrates, cofactors and inhibitors to participate in both procoagulant and anticoagulant pathways. Thrombin is generated by proteolysis of prothrombin and is composed of a A (36 residues, 4.6 kda) and B (259 residues, 32 kda) chain linked together by a disulfide bond (Figure 1.10) (135). 24

46 The major structural elements of the thrombin molecule are: The active site cleft, the A chain, the insertion loops, the anionic patches (constitutes heparine binding site), the RGD sequence (platelet docking site) (136) and two electro-positively charged binding regions (anion binding exosite I (ABE-I) and anion binding exosite II (ABE II) which are crucial for protein function (137). ABE-I is responsible for binding many of the proteins involved in the coagulation cascade including FV, FVa, fibrinogen, PAR-I (the platelet thrombin receptor), thrombomodulin, and heparin cofactor II (138). ABE-II, located just above the active site of the molecule, serves as a heparin-binding site (139). Thrombin zymogen, prothrombin, is a 72 kda vitamin K-dependent glycoprotein produced in the liver, which circulates as a single chain molecule in human plasma at a concentration of approximately 1.4 µm (21). Prothrombin contains a single Gla domain, in which ten glutamic acid residues are converted to γ-carboxyglutamic acids (Gla) by the vitamin K-dependent enzyme γ-glutamyl carboxylase, two Kringle domains, and a protease domain (Figure 1.10) (140). The Gla domain is required for anionic phospholipid binding, which serves to co-localize prothrombin and its activation complex, prothrombinase to the site of vascular injury. The serine protease domain contains the active site and a number of cryptic surface regions that are exposed upon prothrombin activation (134). The primary hemostatic role of thrombin is the generation of fibrin. At the site of vascular injury, a series of proenzymes and procofactors are activated to facilitate the thrombin-mediated proteolytic conversion of fibrinogen into fibrin that ultimately results in the formation of a stabilized fibrin blood clot (134). 25

47 Figure 1.10: Prothrombin Activation Pathway. Conversion of prothrombin to thrombin occurs by two distinct pathways. Prothrombin can be activated by fxa alone, as shown in pathway I or it can be converted to thrombin by the prothrombinase complex pathway II. Pathway I initially induce cleavage at Arg271 followed by cleavage at Arg320, to produce the intermediates fragment 1.2 and prethrombin. Pathway II, which is essential for normal clotting in healthy individuals, catalyzes an initial cleavage at Arg320, followed by cleavage at Arg271 that produces the intermediate, meizothrombin which is further cleaved at Arg271 to produce thrombin and fragment 1 2 (134) Tissue factors (FIII) Tissue factor, also called platelet tissue factor, thrombokinase, or CD142 is a small transmembrane glycoprotein present in sub-endothelial tissue, platelets, and leukocytes. Its main function has been thought to be the initiation of the coagulation cascade. TF exerts its cofactor activity by docking FVII/FVIIa to the membrane surface leading to activation of coagulation. However, recently cellular signalling and other functions have been ascribed to this protein (141). The 47 kda TF protein is a 263-residue transmembrane glycoprotein, consisting of an extracellular domain (amino acids 1-219), a transmembrane domain (amino acids ), and 26

48 a cytoplasmic tail (amino acids ) (142). TF has been classified as a member of the cytokine receptor super family, based on its high degree of structural similarity to other members of the family. Tissue Factor is unique among clotting proteins in that it is not normally found in the plasma, but rather this thromboplastic activity is found in most tissues, except the blood and is particularly high in brain, lung and placenta. Unlike the other cofactors of these protease cascades, which circulate as nonfunctional precursors, this factor is a potent initiator that is fully functional when expressed on cell surfaces. An incorrect synonym is thromboplastin. Historically, thromboplastin was a lab reagent, usually derived from placental sources. Thromboplastin is the combination of both phospholipids and tissue factor, both needed in the activation of the extrinsic pathway (143) von Willebrand factor (vwf) von Willebrand factor is a multifunctional multimeric plasma glycoprotein that plays a prominent role in primary hemostasis (144, 145, 146). In case of vascular injury, vwf acts as a molecular bridge between platelets and the exposed sub-endothelium (144, 147). Furthermore, vwf protects coagulation by protecting FVIII from rapid proteolytic inactivation that leads to the formation of platelet thrombi and the subsequent arrest of bleeding. vwf is usually found in normal blood at a con of ~10 µg/ml with a molecular weight of 250 kda (147, 148, 149, 150). vwf is named after Dr. Erik von Willebrand ( ), a Finnish doctor who first described, in 1924, a hereditary bleeding disorder in families from the Aland islands, who had a tendency for cutaneous and mucosal bleeding, including menorrhagia. Even though Dr. von Willebrand could not identify the definite cause, he distinguished von Willebrand disease (vwd) from hemophilia and other forms of bleeding diathesis (151). In the 1950s, vwd was shown to be caused by a plasma factor deficiency (instead of being caused by platelet disorders), and, in the 1970s, the vwf protein was purified. This factor is deficient or defective in vwd and is also involved in a large number of other diseases, including thrombotic thrombocytopenic purpura, Heyde's syndrome, and possibly hemolytic-uremic syndrome (148) von Willebrand factor biosynthesis, storage and secretion A. von Willebrand factor biosynthesis The two storage organelles found in the cells that synthesize vwf through a multi-step process are the Weibel-Palade body in endothelial cells and the α-granule in megakaryocytes and platelets (71, 148). 27

49 vwf is synthesized as a precursor polypeptide of 2813 aa (pre-pro-vwf), including a signal peptide of 22 aa, a large pro-peptide of 741 aa (also known as vw Ag II), and the mature subunit of 2050 aa (146, 148,152). The signal peptide is cleaved off upon translocation of the protein into the endoplasmic reticulum (Figure1.11). In the reticulum, vwf undergoes complex post-translational modifications preserving the multimeric structure and functions. Before secretion, vwf undergoes extensively twelve N-linked glycosylations followed by dimerization. The dimerization occurs through disulfide bridge formation within the C-terminal residues of the pro-vwf subunit (residues 1908 to 2050, tail to tail). The correctly assembled pro-vwf dimers are then transported to the Golgi apparatus, where further processing takes place such as sulfation, ten O-linked glycosylations as well as high-mannose oligosaccharide processing (153,154). Simultaneously, polymerization of the pro-vwf dimers occurs in the Golgi compartment, a step that requires cysteine residues in domains D1, D2, D and D3, within the N-terminal part of pro-vwf (146, 155, 156). Further multimerization and endoproteolytic cleavage at position 763 occur upon transport to the trans-golgi network, resulting in the release of the pro-peptide dimers from the mature vwf multimers. The pro-peptide is removed before secretion and independently released into the blood (Figure 1.11) (153). The biosynthesis of vwf distinguishes itself from the synthesis of most proteins produced by endothelial cells, in that at least part of the newly synthesized protein is stored in characteristic organelles, called Weibel-Palade bodies (147). 28

50 Figure 1.11: Schematic representation of the processing steps involved in vwf biosynthesis. The hatched area represents the pro-peptide moiety of vwf and the dark area mature VWF. The compartments in which the processing steps take place are indicated on the left. -S-: disulphide bonds. The molecular mass of the different vwf species is given on the right (153). 29

51 B. von Willebrand factor storage The two vwf storage organelles are the Weibel-Palade body in endothelial cells and the α- granule in megakaryocytes (157). The majority of endothelial vwf is secreted constitutively, whereas the remainder high-molecular weight multimers that are highly active are stored in specific releasable endothelial granules (Weibel-Palade bodies). These bodies are rod-shaped organelles composed of closely packed tubules unique to endothelial cells that likely represent the vwf multimers themselves (137, 146). The second vwf storage site, α-granules may contain as much as 20% of the total vwf presented in blood. Several additional molecules important for hemostasis are found in the platelet α-granules including fibrinogen, thrombospondin and fibronectin (158, 159). The vwf of platelet α-granules also consists of the ultra-large vwf multimers. As platelets release their granule contents upon stimulation by agonists such as ADP, collagen and thrombin, this secures ultra-large vwf multimer availability at sites of vascular injury (16). It should be noted that the vwf acutely released from storage sites is not likely to have FVIII bound to it, while the initial binding may serve to drive it at a site of vascular damage (160). C. von Willebrand factor secretion Unlike most other proteins, vwf follows two pathways of secretion; a constitutive one directly linked to its synthesis and a regulated one, involving storage and release after stimulation by secretagogues. Because released vwf undergoes a process of regulated reduction of multimer size, the availability of uncleaved larger multimers in cellular storage sites permits maximal function in areas where rapid platelet adhesion and aggregation is required (16, 71, 157, 162). In megakaryocytes, only the regulated pathway of vwf secretion is effectively operative; thus, the vwf circulating in plasma is essentially all of endothelial cell origin, none of it derives from megakaryocytes, as platelets release their α-granule content only when activated (71, 162). Immunohistological data suggest that only arteries, arterioles and large veins have subendothelial deposits of vwf (163). It is generally assumed that vwf stored within Weibel- Palade bodies is composed of the largest multimeric species (ULvWF), usually not observed in the blood of normal individuals. The metalloproteinase ADAMTS-13 cleaves ULvWF, thus representing a physiological mechanism to cause their disappearance from the circulation, which can be detected only transiently (16, 164, 165). 30

52 von Willebrand factor molecular size and structure von Willebrand factor is the largest protein found in plasma. It circulates in blood as a series of heterogeneous multimers ranging in size from 500 to 20,000 kda (166). The smallest circulating vwf is composed of two identical disulfide-linked subunits via cysteine residues located in the C-terminal region. Each monomer has a total mass of approximately 250 kda consisting of 2050 aa and contains up to 22 carbohydrate chains (16, 71). This is the protomer or building block of the multimeric series. Subsequently, larger polymers are formed by intersubunit disulfide linking of N-terminal ends of the dimer. Multimers of vwf can be extremely large >20,000 kda, and consist of over 80 subunits of 250 kda each (71, 167). The variable molecular weight of vwf is due to differences in the number of subunits comprising the protein, and the size of vwf multimers is directly related to function, as shown by the fact that ultralarge multimers establish high-strength bonds with the platelet GPIb-IX-V complex (166, 168) Gene coding for von Willebrand factor The vwf gene is ~180 kb long (52 exons) and is located at the tip of the short arm of chromosome 12 (12p13.3). The first 17 exons encode the 5 non-coding region, the signal peptide and the pro-peptide, while the remaining 35 exons encode the mature vwf and the 3 non-coding region (109, 156, 169, 170). At the gene level, the expression of vwf is first regulated in a tissue-specific manner. The study of transcription s mechanisms of the vwf gene has revealed the presence of negative and positive regulatory elements in the promoter region or in 5 - flanking regions that are responsible for the endothelial cell-specific transcription of the vwf gene (171, 172, 173). The subunits in vwf multimers are the product of one gene and are initially identical, being modified after secretion by platelet granules, and sub-endothelial matrices into the blood stream (174). The primary translation product is a 2813-residue precursor polypeptide (prepro vwf) (175). Normal levels of vwf in the general population are highly variable and can fluctuate from 50 to 200 U/dl. Among the numerous genetic and environmental factors that contribute to this variation, four single-nucleotide polymorphisms have recently been identified in the vwf gene promoter (-1793 C/G, T/C, G/A, and A/G). The mechanism of action seems to involve the differential binding of endothelial cell nuclear proteins involved in vwf transcription. The most significant of the external genetic factors influencing vwf levels is the ABO blood group, with individuals with group O-blood having the lowest mean levels of vwf antigen. The precise mechanism underlying this observation has not been elucidated (147). 31

53 Molecular cloning of the full-length human vwf cdna has revealed that the mrna is translated as a pre-pro-polypeptide. This indicates that the pre-pro-protein undergoes two proteolytic processing steps before it enters the circulation, cleavage of the signal peptide by a signal peptidase and subsequent cleavage of the propeptide (153) von Willebrand factor s functional domains The pro-peptide and the mature subunit structure of vwf which together represent prepro-vwf are highly repetitive and are almost entirely composed of several discrete functional domains responsible for various biological activities, involving interactions with other molecules (151, 180). The mature vwf subunit contains 2050 aa, 169 of which are cysteine residues clustered in domains located at the N- and C-terminal ends. The estimated carbohydrate content is approximately 18% of the total mass (152). The homo-oligomeric protein mature subunits of vwf are built from four types of conserved structural domains arranged in the following order: D1-D2-D -D3-A1-A2-A3-D4-B1- B2-B3-C1-C2-CK, in which D1 and D2 comprise the propeptide and D -D3-A1-A2-A3-D4-B1- B2-B3-C1-C2-CK the mature subunit from the N- to C- terminus (Figure 1.12) (148, 177). The specific vwf functional domains and their functions are as following: a) The D'/D3 domain: This domain contains the binding sites for heparin, carrier of coagulation FVIII. The region of vwf that binds heparin and FVIII is located between amino acids 1 and 272 (178, 179, 180). b) A domains: The three tandem A domains in vwf which play a critical role for it s function are: A 1 domain: A 1 domain is composed of residues and contains binding sites for (16, 160, 175, 182, 184, 183,1 97): Platelet GPIb-receptor. A 1 domain is the only known binding site for the platelet receptor GPIba Heparin. A heparin-binding site is presumably located in the loop region between residues Cys 509 and 695 Sulphated glycolipids Non physiological ligand like snake protein Collagen type VI A 2 domain: A 2 domain hosts a proteolytic site for ADAMTS-13 (a disintegrin and metalloprotease with thrombospondin type I motife 13) (185, 186). 32

54 A 3 domain: A 3 domain is composed of residues and binds to collagen types I and III (187). c) C 1 domain: C 1 domain contains the "cysteine knot" region comprising of the RGD tripeptide (aa 1744 to 1746) (16, 157). ADAMTS-13 Cleavage site Figure 1.12: Schematic representation of the pre-pro-vwf polypeptide. The domains implicated in different aspects of vwf biosynthesis and storage are indicated by arrows, as well as the proteolytic cleavage site by the plasma vwf-cleaving metalloprotease. Functional domains of vwf involved in its binding to different ligands are indicated by black boxes. Boundaries of the proteolytic fragments used to localize these binding domains are identified by amino-acid numbers in the mature vwf sequence (147) von Willebrand factor A domains The multimeric plasma glycoprotein vwf has three homologous A domains and each of them as shown by crystal structures has a typical α β-fold with a central β-sheet flanked by α helices on each side (Figure 1.13) (181, 188). The central β-sheet consists of six β-strands (β 1 -β 6 ), while the number of α-helices varies. The A 1 domain contains six α-helices (α 1, α 3 -α 7 ) and lacks both the α 2 helix and the metal iondependent adhesion site (Figure 1.13B) (189). Similarly, the A3 domain does not contain the α 2 helix and the metal ion-dependent adhesion site but has a α 8 helix right after the α 7 helix (Figure 1.13D). A homology model of the A2 domain shows that the α5 helix becomes a loop and the proteolytic site on the β 4 strand is completely burried (Figure 1.13C). The A1 and A3 domains contain disulfide bonds linking their N and C-termini (189, 190). 33

55 The A1 domain of vwf exhibits binding affinity for platelet Gplba, a component of the platelet GPlb-IX-V receptor complex that mediates the tethering of platelets to immobilized vwf at sites of injury and is essential to initiate thrombus formation on vascular surfaces exposed to rapidly flowing blood (157, 191, 192). The binding site in vwf responsible for this interaction is not functional under static conditions (soluble vwf) but becomes functional in cases where either vwf is associated with the matrix or under elevated shear stress or becomes activated by two non- physiological modulators such as bacterial glycopeptides ristocetin or snake toxin botrocetin (194,195). The collagen-binding sites in vwf are located within A1 and A3 domains. The latter is necessary and sufficient to support binding to fibril-forming collagens such as type I or type III, whereas domain A1 is involved in collagen type VI binding. Fluid dynamic conditions and mechanical forces may modulate these interactions, and domains A1 and A3 may variably contribute to the immobilization of vwf onto complex extracellular matrices (157, 196). Figure 1.13: Structure of the vwf-a domains. A. Schematic representation of vwf tandem A domains, S-S represents a disulfide bond, the arrow corresponds to ADAMTS-13 proteolytic site. B. Crystal structure of the A1 domain. C. Crystal structure of the A2 domain D. Crystal structure of the A3 domain (189) von Willebrand factor cleavage and unfolding of A domains The largest vwf multimers contained in storage granules are not usually seen in the blood of normal individuals and for this reason they have been referred to as 'unusually' large, they can 34

56 be detected in normal plasma only transiently, e.g. after induction of vwf secretion from endothelial storage sites (144, 147, 197). Interestingly, ULvWF multimers are very large in size and hyperactive in binding the platelet receptor GPIba, which results in spontaneous platelet aggregation without requiring collagen, shear, or chemical stimulation in healthy individuals (144, 158, 168, 189). Because of this prothrombotic property, ULvWF should be quickly removed from the plasma of healthy individuals. Experimental evidence suggests that ULvWF binds the GPIb-IX-V complex with a higher affinity as compared to plasma vwf, thus more effectively enhancing shear-induced platelet aggregation (168,198). This may be due to the increased number of binding sites for platelets in ULWF and may also represent a critical requirement for tethering platelets to subendothelial (16). To avoid possibly excessive thrombus formation in response to irrelevant stimuli, the main mechanism regulating vwf size involves specific proteolysis, with a possible contribution from a disulfide bond reductase activity (71). Very recently, remarkable progress has been made in the identification of two proteins that are responsible for controlling the size of vwf multimers. First, a trimeric glycoprotein thrombospondin-1, is abundant in the α-granules of platelets from which it is released upon activation, and could contribute to the regulation of vwf multimer size at sites of vascular lesions, thus limiting thrombus growth (71, 199). Second, a vwf-cleaving metalloproteinase has been purified by two different groups and has been shown to belong to the ADAMTS family (200, 201). The relationship between thrombospondin-1 and the vwf-cleaving metalloprotease is unknown but their joined actions in controlling the multimer size of vwf prevent the pathological accumulation of very large vwf multimers that could result in spontaneous platelet aggregation (high levels of vwf represent an independent risk factor for cardiovascular mortality and low levels lead to a bleeding diathesis (147)). To avoid excessive thrombus, ADAMTS-13 rapidly cleaves ULvWF on the endothelial surface at the peptide bond between amino acid residues Tyr1605 and Met1606 within the A 2 domain (185, 203). Attached ULvWF multimers to the endothelial surface are under the shear stress of flowing blood and the tensile force of attached platelets which assist the rapid proteolysis of ULvWF by ADAMTS-13 (Figure 1.14) (189). Tsai et al. (1994) showed that the proteolysis of plasma vwf by ADAMTS-13 was enhanced under shear flow and also that the addition of denaturants such as urea and guanidinium hydrochloride, increased the ADAMTS-13 proteolysis of plasma vwf (203). Shear flow and denaturants facilitate the proteolysis because, according to the model proposed by Auton et al. (2007), tensile forces cause conformational changes in both the ligand 35

57 and the receptor that might unfold the A2 domain and expose the proteolytic site, thus facilitating the proteolysis of vwf by ADAMTS-13. This phenomenon threfore results in the reduction of hemostatically active larger vwf multimers in circulation (Figure 1.14) (189, 206). Figure 1.14: Model of vwf cleavage by ADAMTS-13. In blood flow, platelets bind to the A 1 domain, generating tensile forces. These forces unfold the A2 domain and expose the proteolytic site for ADAMTS-13 cleavage (a modified figure from Auton et al. (204)) von Willebrand factor activity von Willebrand factor is not an enzyme and, therefore, has no catalytic activity. Its primary function is the binding to other proteins, thus mediating the initiation and progression of thrombus formation at sites of vascular injury (148, 153, 157, 206). The biological functions of vwf are exerted through specific domains that interact with extracellular matrix components and cell membrane receptors to promote the initial tethering and adhesion of platelets to subendothelial surfaces as well as platelet aggregation and activation (157). Moreover, vwf binds the procoagulant co-enzyme contributing to its stability and indirectly to its function in the generation of fibrin (189, 206). The role of vwf in the arrest of hemorrhage or in the onset of arterial thrombosis depends on its ability to support platelet-surface and platelet-platelet interactions, through the association of membrane receptors to components of the extracellular matrix or to one another. Moreover, vwf performs an essential, albeit indirect function in fibrin clot formation, by associating with FVIII which is a necessary cofactor for the rapid generation of FXa at sites of injury. vwf binds to a number of cells and molecules, the most important of which are FVIII, collagen and platelet GPIb receptors (205, 207). 36

58 von Willebrand factor and activation of platelets In intact blood vessels, vwf does not interact with the platelet receptors. It is assumed that when a blood vessel is injured, sub-endothelial matrix containing collagen is exposed, binds blood circulating vwf and immobilizes it and thereafter platelets bind to immobilized vwf through GPIb receptors (166). Glycoprotein Ib-vWF interactions trigger intracellular signaling in platelets, which activate integrins α2β1 and αiibβ3 on the platelet membrane. Active α2β1 and αiibβ3 interact with collage and vwf, respectively, resulting in firm adhesion. Furthermore, platelets aggregate by αiibβ3-fibrinogen interactions. Adhesion of platelets to the site of vascular injury is one of the prerequisite and controlling factors of many bleeding disorders such as myocardial infarction and stroke (181, 189). A large number of proteins through various interactions hold adhered platelets on the site of injury, vwf being one of the major blood proteins in this cascade (205). On the other hand, binding of vwf to GPIb through A1 domain induces platelet activation by initiating a complex intracellular signaling pathway which leads to the secretion of α-granule active substances (55). The hypothetical model to explain vwf-mediated platelet adhesion is based on two presumptions: The first one suggests that in flowing blood, vwf appears in a ball of yarn configuration and does not interact with the platelet GPIb-IX V receptors under physiological conditions because the GPIb binding domain is concealed. The affinity towards GPIb-IX-V is regulated by conformational changes in vwf, which are induced by immobilisation and shear. These conformational changes of vwf result in the exposure of binding sites within the vwf- A1 domain responsible for interaction with platelet GPIb. This is supported by studies showing a conversion of vwf from its loosely coiled structure into elongated filaments upon exposition to high shear stress, suggesting that shear modulates the balance between the low affinity and high affinity state of vwf (208). Furthermore, binding of the vwf-a1 domain to platelet GPIb in the absence of flow requires the addition of chemical modulators, such as snake venom protein botrocetin, or bacterial antibiotic ristocetin (209). The second persumption suggests the existence of a transient interaction between platelet GPIb and immobilized vwf with a fast offrate, whereas shear induces a permanent interaction between the second vwf receptor on platelet surface (GPIIb/IIIa) and vwf and only this second interaction is capable to immobilize platelets (210). This second persumption is supported by perfusion studies on collagen-coated surfaces, showing reversible interaction of platelets with collagen-bound WF under flow (211). Taken together, the widely accepted concept of vwf-mediated platelet adhesion at physiological high fluid shear stress involves vwf binding to sub-endothelium (e.g. collagen) resulting in platelet translocation along the surface in the direction of flow via reversible vwf- 37

59 GPIb bonds. This slow motion allows the establishment of additional interactions, i.e. between vwf and GPIIb/IIIa, resulting in platelet activation via transducing signals and aggregation to the surface in a biphasic adhesion process (148). The mechanism of platelet tethering, translocation and adhesion is depicted in Figure 1.15 which was taken from Ruggeri (2002) (25). Figure 1.15: Biphasic model of vwf-mediated platelet adhesion. According to Ruggeri (2002) (25), vwf becomes immobilized on extracelluler matrix and transient bonds between vwf and platelet GPIb result in translocation of platelets along the surface in the direction of flow. Secondary interactions between platelet surface receptors and extracellular collagen result in stable platelet adhesion, and finally adhesion through binding of activated platelets to various plasma proteins von Willebrand factor and stabilization of FVIII Factor VIII is an essential cofactor in blood coagulation synthesized in liver cells. Upon release, the inactive FVIII precursor protein binds to vwf (99% vwf and 1% FVIII) and circulates in plasma non-covalently attached to the N-terminal moiety of vwf via its light chain (206, 207). Factor VIII bound to vwf is protected from inactivation by proteases (activated PC and FXa) (167) and has a longer lifetime in the blood, thus being more efficient in performing its function. Indeed, in the absence of vwf, the FVIII levels are dramatically reduced (157, 166, 206). 38

60 Aims of this study 1. Development and optimization of a chromogenic substrate assay for the determination of thrombin production, to investigate the usefulness of coagulability as a marker for clinical diagnosis, drug monitoring and epidemiology. 2. Development and optimization of a chromogenic substrate assay for the determination of activity of blood clotting FXa, to investigate its potential value clinical uses. 3. Biochemical study of thrombin and its mode of action in case of hemostasis. 4. Biochemical study of blood clotting FXa and its mode of action on the process of hemostasis. 5. Influence of platelet concentration on the normal process of hemostasis. 6. Influence of activated FVII on the process of clotting mechanism. 7. Influence of TF on the process of coagulation system. 8. Study of the effect of clotting on a chromogenic substrate assay for the determination of thrombin and FXa production. 9. Correlation between thrombin and blood clotting FXa. 10. The role of physical exercise on the mechanism of thrombin and FXa generation 11. Development of a western blot-based technique for monitoring the conversion of the zymogen blood clotting FX to the serine protease FXa. 12. Cloning, expression and purification of vwf A1 domain to identify the specific amino acid region of vwf that promotes optimal cofactor function in platelet aggregation and activation during the normal process of hemostasis. 13. Study of the potential function of purified vwf A1 domain in platelet activation and the natural process of hemostasis in order to fully understand its particular role in maintaining the integrity of the blood coagulation. 14. To further investigate the mechanism of thrombus formation and develop a new remedy against thrombus formation. 39

61 CHAPTER TWO MATERIALS AND METHODS 2.1. Study populations The subjects of our study were grouped into three categories: A. Healthy controls: Fifty randomly selected subjects served as control, all were healthy volunteers and had no evidence for any blood diseases. B. Hospitalized patients with diabetes mellitus (DB): One hundred twenty hospitalized patients with DM at Saint Loukas general hospital of Thessaloniki under different antithrombotic therapies were eligible for the study. C. Exercise group: A total of thirty individuals under moderate physical exercise who registered in oxygen sport center in Trikala-Greece Collection of Blood Samples Blood samples were collected via a traumatic antecubital veinpuncture into vacutainer tubes with and without sodium citrate. The collected samples were centrifuged at 3,000 rpm for 20 min and 1000 rpm for 5 min at RT for platelet poor plasma (PPP) and platelet rich plasma (PRP) respectively. The samples were either used immediately for the study of thrombin, FXa and vwf-a1 domain protein or kept until further analysis. The acceptable samples for coagulation tests include platelet poor plasma (PPP), platelet rich plasma (PRP) and plasma samples before and after clot removal. Platelets were counted by a Beckman Coulter counter and adjusted to 150 x 10 9 platelets/l. Platelet rich plasma was collected from the upper 3/4 volume of plasma supernatant after centrifugation at 265xg for 5 min at RT. Autologous platelet poor plasma (PPP) was prepared by centrifuging twice at 2,900xg for 10 min at RT. PRP was always used within 1h after venipuncture. In order to avoid contamination with procoagulant microparticles from aging platelets, PPP was preferably prepared within 30 min after venipuncture Materials The chemicals and solutions used in this study are tabulated in appendix (1). 40

62 2.1. Experimental protocols Biochemical assays Thrombin production assay Thrombin production was deternined in various samples according to the Hemker s method with slight modifications (212). More precisely, optimization of the assay was based on the use of different ph and concentration of the substrates, CaCl 2, BSA and EDTA. After some initial experiments the following assay conditions were chosen: ph 7.7, substrate S2238 (0.37 mm/l), CaCl 2 (4.7 mm/l) and EDTA (20 mm/l). The samples were measured continuously at 37 C in a CGA EOS880 spectrophotometer (Germany). Routinely, 430 µl of buffer A, 10 µl of Thromborol S (Dade Behring, Marbay, Germany) and 35 µl of substrate S-2237 (formal chemical name: H-D-Phenylalanyl-Lpipecolyl-Larginine-p-nitroaniline dihydrochloride) from Chromogenix Instrumentation Laboratory Company - Lexington, (USA) were added to 30 µl of sample. In a following step, the reaction was activated at zero time by adding 25 µl CaCl 2. The optical density indicating thrombin production was determined at 405 nm every 30 s for a time period of 3 min Factor Xa production assay Reagents contained in a heparin kit STA Rotachrom, suitable for the colorimetric assay of heparin and fondaparinux (Diagnostica stago- France) (213) were used for FXa generation assay. The basic experiment was performed as follows: 430 µl of buffer A, 10 µl of rtf and 90µL of substrate CBS (formal chemical name: MARA-Gly-Arg-pNA, AcOH) were incubated with 90 µl of sample. FXa production was activated at zero time in 37 C by adding 25 µl CaCl 2. The optical density indicating FXa production was determined at 405 nm every 30s for a time period of 3 min Molecular methods (Molecular assay protocols) Cloning, expression and purification of mammalian cell-derived recombinant vwf-a1 fragments For cloning, expression and purification of vwf-a1 domain protein, the complete coding sequence of vwf-a1 domain (appendix 2) was cloned in frame into the pcdna3.1 myc-his (- )B expression vector according to standard protocols which include: choice of host organism and cloning vector, isolation of vector DNA, isolation of DNA to be cloned, construction of the recombinant DNA vector by ligation, introduction of recombinant DNA vector into host 41

63 organism, selection of organisms containing recombinant DNA and finally screening for clones containing the desired DNA insert (214) Construction of pcdna3.1 myc-his (-)B vwf-a1 domain expression vector pcdna.3.1/myc-his(-)b vector The chosen vector pcdna3.1/myc-his(-)b (5.5-kb) was derived from pcdna3.1(+) and designed for high-level expression, purification, and detection of recombinant proteins in mammalian hosts. High-level stable and non-replicative transient expression can be carried out in most mammalian cells. The vector was supplied with three reading frames to facilitate inframe cloning, with a C-terminal peptide containing a polyhistidine metal-binding tag (His-tag) and the myc (c-myc) epitope. The human cytomegalovirus (CMV) promoter in the vector provides high-level expression in a wide range of mammalian cells (Figure 2.1) (214). Before starting our cloning, the pcdna.3.1/myc-his(-)b plasmid vector was digested by restriction endonuclease Xba1, resulting in two products of 86 and 5434 bp in accordance with the manufacturer s instructions (215, Invitrogen). Vector cleavage Reaction mixture volume (20 µl): 2µl of (10x) NBE buffer 3µl DNA plasmid vector (1 µg) 13.5 µl ddh 2 O 1µl Xba1 enzyme (5 U) Incubation for min. 1 % (w/v) agarose gel analysis of digested pcdna.3.1/myc-his(-) B 42

64 Figure 2.1: Restriction map of the pcdna 3.1/myc-His vector Plasmid DNA preparation A. Plasmid DNA mini-preparation protocol (3 ml LB) A number of simple and rapid protocols are available for preparing small amounts of plasmid DNA from bacteria. The mini-preparation that we used is designed to provide a rapid but rather low amount of plasmid (20 to 30 µg) which is, however, sufficient for restriction analysis as well as for probe labeling. More specifically, 3 ml of rich LB medium containing the appropriate antibiotic (3 µl Ampicilin 100 µg/ml) were inoculated with a single colony of transformed bacteria. The culture was incubated overnight at 37 C with vigorous shaking at 120 rpm and the culture was used for restriction analysis (216). 43

65 B. Plasmid DNA maxi-preparation protocol (500 ml LB) An overday 3 ml culture was innoculated to 500 ml LB containing 500 µl ampicilin (100 µg/ml). The 500 ml culture was allowed to incubate at 37 C with vigorous shaking at 120 rpm until the absorbance of the culture reached 0.37 OD. The cells were pelleted by centrifugation at 5000xg for 20 min at 4 C. The supernatant was removed and discarded, the cell pellet was drained of residual liquid by inverting the bottles on a paper towel for about 10 min and after that the cells were stored until further analysis (216) Plasmid DNA extraction using alkaline lysis One and half ml of an overnight culture (miniprep) were transferred into a microfuge tube and the cells were pelleted by centrifugation at 4000 rpm for 15 min. The supernatant was discarded and the cell pellet was drained of residual liquid as possible. The cell pellet was resuspended in 100 µl buffer I (TE) and then 200 µl of buffer II were added, containing 0.2 M NaOH and 1 % (w/v) SDS, mixed by inversion several times and left at room temperature for 5 min. To neutralize the ph of the solution, 100 µl of cold buffer III (KOAC) were added, mixed by inversion several times again and left on ice for 10 min. The neutralized mixture was centrifuged at rpm for 3 min at 4 C in a Sorvall RC-5B refrigerated centrifuge. The supernatant was separated carefully and transferred in a new eppendorf tube, followed by addition of about 1 ml (2.5 times) ice cold ethanol 100% (or isopropanol), mixed by inversion several times and placed at -20 C for several minutes. The DNA was precipitated by centrifugation at 12000xg for 10 min, washed with 70% alcohol, centrifuged again at 1300 rpm for 5 min at 4 C to discard the residual alcohol by micropipette and was finally suspended in 20 µl nuclease free water (216) Restriction endonuclease digestion of plasmid DNA (pcdna3.1myc-his(-)b) The plasmid pcdna3.1myc-his(-)b vector was cleaved with BamH1 and EcoR1 restriction endonucleases. The reactions were performed in a total volume of 30 µl as follows: 5 µl of plasmid vector (2 µg) were mixed well with 20 µl H2O, the mixture was heated for 5 min at 71 C, transferred on ice for 1 min, followed by addition of 3 µl of NEB2 buffer, 1 µl EcoR1 and 1µl BamH1. The mixture was then allowed to incubate for 40 min at 37 C, 5 min at 38 C, and 20 min at 70 C for deactivation of restriction enzymes. 44

66 Polymerase chain reaction (PCR) for amplification of the DNA fragment encoding vwf-a1 domain The cdna of human vwf was used as a template to amplify the region of interest by using standard polymerase chain reaction (PCR) (217). The desired domain was amplified using primers containing EcoR1 and BamH1 restriction sites at the 5' end and 3' end respectively, since the introduction of BamH1 and EcoR1 restriction sites in the PCR products facilitate the ligation of PCR products back in to pcdna3.1 myc-his (-)B vector. A. Primers The primers used to amplify the DNA fragment encoding the A1 domain of vwf are shown in Table 2.1. The primers were synthesized by integrated DNA technologies (Invitrogen). Table 2.1: Oligonucleotide sequences used to amplify the DNA fragment encoding the A1 region of vwf for subcloning it to the pcdna 3.1/myc-His vector vwf-a1 Primers sequence Sense 5'-GCAGGAATTCGACCTGGTCTTCCTGCTGGATGGC-3' Antisense 5'-GCTCGGATCCACGATCTCGTCCCTTTGCTGCTCCAG-3' B. Polymerase chain reaction (PCR) reagents A plasmid containing the full-length cdna of human vwf was used as template to amplify the DNA fragment encoding the A1 domain. PCR amplification was performed in a total of volume of 50 µl containing GC rich F514 DNA buffer, dntps, Taq polymerase, template DNA and primers as shown in Table

67 Table 2.2: Reagents used in the PCR amplification of the DNA fragment encoding the A1 region of vwf for subcloning it to the pcdna 3.1/myc-His vector Compounds Amount Final Con. dd H 2 O 34.3 µl -- GC rich F514 DNA buffer containing 10 µl 5x 1.5mM MgCl 2 dntps 1 µl 20 µm forward primer 2 µl 0.1 µg/µl reverse primer 2 µl 0.1 µg/µl Template DNA 0.2 µl 0.2 ng/µl Taq Polymerase 0.5 µl 2 U/ml PCR amplification was performed in a total volume of 50 µl using Taq polymerase (Finnzymes F530S- New England Biolabs, UK). The steps were as follows (see Table 2.3): Initial denaturation for 2 min at 95 C, 15 cycles of denaturing at 95 C for 30 s, annealing at 65 C for 30 s, elongation at 72 C for 40 s and 15 cycles of denaturing at 95 C for 30 s, annealing at 63 C for 30 s, elongation at 72 C for 40 s with a final extension at 72 C for 5 min. Table 2.3: PCR conditions used to amplify the DNA fragment encoding the the A1 region of vwf for subcloning it to the pcdna 3.1/myc-His vector Steps Temp. Time No. of cycle (s) Initial denaturation 95 C 2 min 1 Denaturation 95 C 30 s 1) PCR Cycle Annealing 65 C 30 s 15 Elongation 72 C 40 s Denaturation 95 C 30 s 2) PCR Cycle Annealing 63 C 30 s 15 Elongation 72 C 40 s Final extension 72 C 5 min 1 Hold 4 C

68 Purification of PCR products by phenol chloroform extraction and ethanol precipitation The PCR products were subjected to phenol chloroform extraction and ethanol precipitation to remove unused primers, Taq polymerase and buffer. Briefly, an equal volume of equilibrated phenol was added to PCR products. The mixture was vortexed for about 20 sec to form an emulsion to increase the contact between phases and maximize extraction. The phases were separated by centrifugation at rpm for 5 min at RT. The top aqueous phase was carefully removed into a new autoclaved 1.5 ml eppendorf tube. To concentrate the extracted DNA, ethanol precipitation was performed by adding 30 µl of sodium acetate and 1 ml ice-cold ethanol 100%. The tube was mixed by inversion and placed at -80 C for 3 min. The DNA was precipitated by centrifugation at 12000xg for 10 min, washed with 70% alcohol, centrifuged again at 1300 rpm for 5 min at 4 o C to discard the residual alcohol by micropipette and was finally suspended in 20 µl nuclease-free water. The PCR products were analyzed on a 1.2 % (w/v) agarose gel (214) Digestion of the purified PCR products with restriction endonucleases The ethanol precipitated PCR products were digested with BamH1 and EcoR1 restriction enzymes to get unique protruding ends. The digestion reactions were performed in a total volume of 30 µl. Briefly, 12 µl of the PCR products (~2 µg) were mixed with 13 µl doubledistilled water (dd H 2 O), the mixture was heated for 5 min at 71 C, placed on ice for 1 min, followed by addition of 3 µl of NEB2, 1 µl EcoR1 and 1 µl BamH1. The mixture was allowed to incubate for 40 min at 37 C, 5 min at 38 C and 20 min at 70 C and the products were storaged until further analysis Ligation of the DNA fragments encoding vwf-a1 domain into the expression vector The purified and digested PCR products with BamH1/EcoR1 restriction enzymes were ligated to the BamH1/EcoR1 sites of the pcdna3.1myc-his(-)b plasmid vector using Takara ligase according to Takara-bio-Inc manufacturer s instructions (218). The ligation reaction contained 15.3 µl of ddh 2 O, 0.5 µl (50 ng) purified plasmid DNA, 1.0 µl of the PCR product (100 ng), 2 µl ligation buffer and 1.2 µl Takara ligase. Incubation was carried out at 16 o C overnight. 47

69 Transformation of bacterial cells (E. coli TOP10) with pcdna3.1myc-his(-)b vwfa1 recombinant vector The construct (pcdna3.1myc-his(-)b-vwfa1) was transformed into E. coli Top10 competent cells. Transformation was performed according to the InvitrogenTM pcr-blunt kit instructions (215) Transformation protocol: 3 µl of recombinant expression vector (pcdna3.1myc-his(-)b vwfa1 100 µl of competent cells (E. coli Top10) Incubation on ice for 20 min Thermal shock at 42 C for 2 min Incubation on ice for 1 min Addition of 700 µl TYM medium 30 min incubation at 37 C with vigorous shaking (160 rpm) Centrifugation at 4000 rpm for 30 s Removal of the supernatant and resuspension of the cells in the 100 µl remaining supernatant Using a sterile bent glass rod the transformed cells (100 µl) were gently spread over the surface of an LB agar plate containing 100 µg/ml ampicillin. The plates were inverted and incubated overnight at 37 C Identification of bacterial colonies that contain recombinant plasmids Twenty four colonies were picked and grown overnight in small-scale cultures (minipreps, see ). Plasmid DNAs, isolated from each culture were then analyzed by digestion with EcoRI and BamH1 restriction enzymes and gel electrophoresis on a 1.2% (w/v) agarose gel. Transformed cells with the recombinant vector pcdna3.1myc-his(-)b/vwf-a1 were harvested and used to isolate plasmid DNA for mammalian cell transfection High yield plasmid DNA purification for mammalian cell transfection The PureLink HiPure Plasmid purification kit (Invitrogen), designed to efficiently isolate plasmid DNA from E. coli using anion-exchange columns, was used for pcdna3.1myc- His(-)B vwf-a1 purification. The purified plasmid DNA is ultrapure and suitable for purity, transfection in mammalian cells. The plasmid DNA purification protocol is shown in Figure

70 Figure 2.2: Flow chart for plasmid DNA purification using the PureLink HiPure Plasmid DNA purification kit Transfection of 293T human embryonic kidney cells with the pcdna3.1myc- His(-)B-vWF-A1 expression vector and protein expression Human Embryonic Kidney 293 cells are a specific cell line originally derived from human embryonic kidney cells grown in tissue culture. HEK 293 cells are easy to grow, get transfected very readily and have been widely-used in cell biology research for many years. They have been also used in the biotechnology industry to produce therapeutic proteins and viruses for gene therapy (219, 220). 49

71 Human 293T cells were maintained in Dulbecco s modified Eagle s medium supplemented with 10% (v/v) fetal calf serum and antibiotics (penicillin and streptomycin). The cells were transfected with pcdna3.1myc-his(-)b-vwfa1 using the calcium phosphate method (221). Exponentially growing cells were trypsinized, seeded at 5 x 10 5 cells per 10 cm plate, and incubated overnight in 10 ml of growth medium. Then 7 µg of pcdna3.1myc-his(-)b vwf- A1 DNA were mixed with 0.5 ml of CaCl 2 and 0.5 ml of BES-buffered saline (2x BBS). Total amounts of plasmid DNA were made up to 20 µg with pcdna3 and the mixture was incubated for 20 min at RT. The calcium phosphate-dna mixture (1 ml) was added dropwise to the plate of cells by gently swirling and the cells were incubated for 16 h. After first incubation, the medium was changed and the cells were further incubated for another 24 h. Cells were lysed with 200 µl of Triton buffer for 30 min on ice. Whole cell extracts were clarified by centrifugation for 15 min in a microcentrifuge, and the protein concentration was determined by the method of Bradford. One hundred and fifty micrograms of each lysate was supplemented with the appropriate volume of electrophoresis buffer and analyzed on 10 % (w/v) SDS-PAGE Affinity purification of vwf-a1 domain using Ni 2+ affinity-chromatography (Ni-NTA) Purification of the recombinant protein (vwf A1 domain) was performed according to the manufacturer s instructions (Ni-NTA Spin Kit Handbook, Qiagen) (222). Ni-NTA Kit provides a simple method for rapid screening and purification of 6xHistagged proteins from small-scale expression cultures. It allows rapid purification of proteins from crude cell lysates under either native or denaturing conditions. Affinity chromatography (Ni-ANT protocol) 1. Resuspension of cells in buffer A (5 ml per gram weight) and stirring for 2 h at RT 2. Seperation of supernatant by centrifugation for 10 min at rpm 3. Washing three times of the Ni-NTA resin (1 ml) with 15 ml buffer A 4. Addition of the Ni-NTA resin to the supernatant and stirring for 2 h at RT 5. Discard of supernatant by centrifugation at 1600 rpm for 5 min 6. Washing three times (5 min each time) of the Ni-NTA resin with 15 ml buffer A 7. Washing three times (5 min each time) of the Ni-NTA resin with 15 ml buffer B 8. Washing three times (5 min each time) of the Ni-NTA resin with 15 ml buffer C 9. Protein elution with 500 µl of Buffer E 10. SDS-PAGE analysis of the eluted protein 50

72 Cloning, expression and purification of Escherichia coli-derived recombinant vwf-a1 fragments Construction of pet-29c(+) vwf-a1 domain expression vector For cloning, expression and purification of human vwf-a1 domain in bacterial cells, the complete coding sequence of vwf A1 domain was cloned in frame into the pet-29c(+) expression vector (5.371 kb). The pet system is one of the most powerful systems for cloning and expressing of recombinant proteins in E. coli. Target genes are cloned in pet plasmids which are under control of strong bacteriophage T7 transcription and translation signals. Expression is induced by providing a source of T7 RNA polymerase in the host cell (Novagen). The pet-29c(+) vectors contain translation stop codons in all three reading frames following the cloning and polyhistidine metal-binding tag regions as well as a downstream T7 transcription terminator (Figure 2.3, Invitrogen)(223). Figure 2.3: Restriction map of the pet-29c(+) vector. 51

73 Polymerase chain reaction (PCR) for amplification of the DNA fragment encoding vwf-a1 domain The cdna of human vwf was used as a template to amplify the region of interest by using standard polymerase chain reaction (PCR) (217). The desired domain was amplified using primers containing NdeI and XhoI sites restriction sites at the 5' end and 3' end respectively, to facilitate the ligation of PCR products back into the pet-29c(+) vector. A. Primers The primers used to amplify the DNA fragment encoding the A1 domain of vwf are shown in Table 2.4. The primers were synthesized by integrated DNA technologies (Invitrogen). Table 2.4: Oligonucleotide sequences used to amplify the DNA fragment encoding the A1 region of vwf for subcloning it to the pet-29c(+) vector vwf-a1 Primers sequence Sense 5'- GCGCATATGGACCTGGTCTTCCTGCTGGATG-3' Antisense 5'- CGCCTCGAGGATCTCGTCCCTTTGCTGCTCC -3' B. Polymerase chain reactions (PCR) reagents A plasmid containing the full-length cdna of human vwf was used as template to amplify the DNA fragment encoding the A1 domain. PCR amplification was performed in a total of volume of 50 µl containing Phusion buffer, dntps, Phusion High Fidelity DNA polymerase, template DNA and primers as shown in Table 2.5. Table 2.5: Reagents used in the PCR amplification of the DNA fragment encoding the A1 region of vwf for subcloning it to the pet-29c(+) vector Compounds Amount Final Con. dd H 2 O 34.3 µl -- Template DNA 0.2 µl 10 ng Phusion buffer 10 µl 5x dntps 1 µl 10 µm forward primer 2 µl 0.5 µm reverse primer 2 µl 0.5 µm Phusion polymerase 0.5 µl 0.02 U/ml 52

74 PCR amplification was performed in a total volume of 50 µl using Phusion High Fidelity DNA polymerase (Finnzymes Finland). The steps were as follows (see Table 2.6): Initial denaturation for 30 s at 98 C followed by 35 cycles of denaturing at 98 C for 5 s, annealing at 58 C for 30 s and elongation at 72 C for 15 s, with a final extension at 72 C for 10 min. Table 2.6: PCR conditions used to amplify the DNA fragment encoding the A1 region of vwf for subcloning it to the pet-29c(+) vector Steps Tem. Time No. of cycle (s) Initial denaturation 98 C 30 s 1 Denaturation 98 C 5 s PCR Cycle Annealing 58 C 30 s 35 Elongation 72 C 15 s Final extension 72 C 10 min 1 Hold 4 C Digestion of the purified PCR products with restriction endonucleases The ethanol precipitated PCR products were digested with NdeI and XhoI restriction enzymes to get unique protruding ends. The digestion reactions were performed in a total volume of 30 µl. Briefly, 12 µl of the PCR products (~2 µg) were mixed with 13 µl doubledistilled water (dd H 2 O), the mixture was heated for 5 min at 71 C, placed on ice for 1 min, followed by addition of 3 µl of NEB2, 1 µl NdeI and 1 µl XhoI. The mixture was allowed to incubate for 40 min at 37 C, 5 min at 38 C and 20 min at 70 C and the products were used for ligation Ligation of the DNA fragments encoding vwf-a1 domain into the expression vector The purified and digested PCR products with NdeI/XhoI restriction enzymes were ligated to the NdeI/XhoI sites of the pet-29c(+) plasmid vector using Takara ligase according to Takara-bio-Inc manufacturer s instructions (218). The ligation reaction contained 15.3 µl of ddh 2 O, 0.5 µl (50 ng) purified plasmid DNA, 1.0 µl of the PCR product (100 ng), 2 µl ligation buffer and 1.2 µl Takara ligase. Incubation was carried out at 16 0 C overnight. 53

75 Transformation of bacterial cells (E. coli TOP10) with the pet-29c(+)vwf-a1 expression vector The construct (pet-29c(+)vwf-a1) was transformed into E. coli Top10 competent cells. Transformation was performed as described in paragraph Twenty four colonies were picked and grown overnight in small-scale cultures (minipreps, see ). Plasmid DNAs, isolated from each culture were then analyzed by digestion with NdeI and XhoI restriction enzymes and gel electrophoresis on a 1.2 % (w/v) agarose gel. Transformed cells with the recombinant vector pet-29c(+)vwf-a1 were harvested and used to isolate plasmid DNA Transformation of Escherichia coli strain BL21 (DE3) with the pet-29c(+)- vwf-a1 expression vector The construct (pet-29c(+)vwf-a1) was transformed into BL21 competent cells. Transformation was performed according to the following protocol: Transformation protocol: 50 ng of the construct (pet-29c(+)vwf-a1) is mixed with 50 µl of competent cells Incubation on ice for 20 min Addition of 1 ml SOC Incubation for 1 h at 37 C with vigorous shaking (160 rpm) Centrifugation for 30 s at rpm Removal of the supernatant and resuspension of the cells in the 100 µl remaining supernatant Using a sterile bent glass rod the transformed cells (100 µl) were gently spread over the surface of an LB agar plate containing 100 µg/ml kanamycin The plates were inverted and incubated overnight at 37 C Overexpression of recombinant vwf-a1 protein in Escherichia coli strain BL21 (DE3) using IPTG (isopropyl-1-thio-β-d-galactopyranoside) The following induction protocol is a general guide for expression of proteins from genes that under the control of IPTG-inducible promoters. This protocol is mainly used to analyze protein expression of individual transformants using BL21 (DE3) host strains transformed with constructs containing T7 promoters (e.g. pet vectors). 54

76 Protein overexpression protocol 10 ml of LB broth containing kanamycin were inoculated in a falcon tube with a single colony from recently plated BL21 transformed cells Cells were grown overnight at 37 C with vigorous shaking The overnight culture was diluted with 190 ml LB in a conical flask The fresh culture was left to grow at 37 C with shaking for 2-3 h The growth of the cells was monitored with a spectrophotometer. When OD(600 nm)= (log phase) IPTG was added to a final concentration of 1 mm and the culture was further incubated for 3 h at 37 C with shaking After the induction period, the cells were harvested by centrifugation at 6,000 rpm for 20 min at 4 C Purification of recombinant vwf-a1 protein by affinity chromatography The recombinant vwf-a1 fragment was purified by Ni-NTA resin chromatography with purity over 95%. Protein purification was performed according to the manufacturer s instructions (Ni-NTA Spin Kit Handbook, Qiagen) (222). Briefly, harvested bacterial cells were resuspended in 3 ml freshly prepared lysis buffer and sonicated. The lysate was centrifuged at 10,000 rpm for 10 min at 4 C and the supernatant was then loaded onto the Ni 2+ -resin column. The bound protein was washed 3 times with purification buffer and the purified recombinant protein was eluted with elution buffer. To monitor the production of recombinant vwf A1 protein, both total lysates from E. coli and purified recombinant protein following affinity chromatography were electrophoresed under reducing conditions on 10 % (w/v) SDS-PAGE and were visualized using coomassie blue staining SDS-PAGE and Western blotting analysis To detect and separate specific proteins (blood clotting FX, FXa and vwf A1 domain) in a given sample (tissue homogenate or cell extract) and to determine their purity, SDS-PAGE analysis was used. This technique is widely used in biochemistry, forensics, genetics and molecular biology to separate proteins according to their electrophoretic mobility. One dimensional electrophoresis was used for the protein analysis throughout this study. Proteins were subjected to 10 % (w/v) SDS-PAGE and transferred to nitrocellulose membrane. The membrane was probed with the appropriate antibody followed by secondary antibody (224, 225). SDS-PAGE analysis and Westrn blotting analysis were performd as follows. 55

77 Sample preparation A. Blood clotting FX and FXa Blood clotting FX and it s active form FXa samples for SDS-PAGE were prepared as described below: Routinely, 16 µl of buffer A (same buffer for the determination of FXa production), 10 µl of Thromborol S, 10 µl rfviia and 4 µl CaCl 2 (for activation of the reaction) were added to 20 µl of blood samples, followed by incubation at 37 C for various time intervals (0, min). The reaction was then stopped quickly by adding 20 µl of EDTA and 120 µl of H 2 O. Ten µl of the reaction mixture were transferred to a new eppendorf tube which contained 5 µl loading buffer, 5 µlh 2 O and 2 µl mercaptoethanol. The samplers were heated in boiling water for 4 min and spinned in a centrifuge for 2 s before SDS-PAGE analysis. B. Lysates from cell culture For recombinant vwf A1 domain analysis, 6 µl the extracted protein were added to 14 µl H 2 O, 4 µl loading buffer and 2 µl mercaptoethanol. The samplers were heated for 4 min and spinned in a centrifuge for 2 s before SDS-PAGE analysis Preparation of SDS-PAGE gel and electrophoresis The technique is a standard means for separating proteins according to their molecular weight. Polyacrylamide gels are formed from the polymerization of two compounds, acrylamide and N, N-methylenebis-acrylamide (Bis). Bis is a cross-linking agent for the gels. The polymerization is initiated by the addition of ammonium persulfate along with TEMED. The gels are neutral, hydrophilic, three-dimensional networks of long hydrocarbons crosslinked by methylene groups. The separation of molecules within a gel is determined by the relative size of the pores formed within the gel. The composition of separating and stacking gel is shown in appendix (1). After the preparation of the gel and set up of electrophoresis apparatus, the protein samples were added to the wells on top of the gel with a syringe or pipette. The apparatus was connected to a power supply and running proceeded under the following appropriate settings: 35 ma for blood clotting FX and FXa 100 V for A1 domains of vwf When the dye molecules ( migration front ) reached the bottom of the gel, the power was turned off and the gel proceeded to staining or Western blotting analysis. 56

78 Coomassie blue staining Analyzed proteins were stained with 0.25% Coomassie brilliant blue in 50% methanol and 10% acetic acid for 30 min at room temperature. Destaining was carried out overnight in 50% methanol and 10% acetic acid. The dye does not bind to acrylamide and washes out (leaving a clear gel). However, it remains strongly bound to the proteins giving them a deep blue colour Western blotting analysis In order to make the proteins accessible to antibody detection they were transferred to a nitrocellulose membrane. Transferring of the proteins to the nitrocellulose membrane is mediated by electric current. In this respect a sandwich consisting of Whatman paper-gelmembrane- Whatman paper, all wetted in transfer buffer, was assembled between cathode and anode. As the proteins are negatively charged, due to SDS, the membrane is oriented towards the positive electrode and the gel towards the negative. The apparatus used for the transfer was the Semi-dry Blotter (Scie-Plas) and the transfer buffer consisted of 12.5 mm Tris-boric ph 8.5, 0.02 % (w/v) SDS and 0.5 mm DTT. Transfer was performed for 1.5 h for FXa and FX and 25 min for vwf-a1 under constant current. The nitrocellulose membrane was then blocked in 10 ml of blocking buffer (blocking buffer I for FXa& FXa and blocking buffer II for vwf-a1) for 45 min at room temperature. Blocking prevents non-specific background binding of the primary and/or secondary antibodies. For FX and FXa detection the membranes were incubated overnight at 4 C on a shaker with 2.5 µl of a rabbit polyclonal anti-human FX antibody, while for vwf-a1 detection the membranes were incubated with 10 µl of a monoclonal mouse anti-his antibody (recombinant vwf protein contains an histidine tag). Following incubation with the primary antibody, the membranes were washed three times with PBS-Tween, for 5 min each, at room temperature with shaking to remove residual primary antibody and then incubated with 2.5 µl of goat anti-rabbit secondary antibody (for the polyclonal anti-fx antibody) or 5 µl of goat anti-mouse secondary antibody (for the monoclonal anti-his antibody) for 1 h with shaking at room temperature. The membranes were subsequently washed twice with PBS-Tween, for 5 min each, and once with alkaline phosphatase buffer. The visualization of the protein bands were carried out by incubating the membrane with 10 ml of alkaline phoshatase buffer (1X) containing 30 µl of NTB and 30 µl of BCIP. 57

79 Statistical analysis Data were evaluated by SPSS software (SPSS Inc., Chicago, IL, USA). The correlation coefficient (r) was used to assess the link between thrombin and FXa levels. T-test for independent and dependant samples was used to assess the difference between serum samples before and after clot removal, exercise groups, patients and controls regarding thrombin and FXa levels. Values are reported as mean ± SE. P<0.05 was considered statistically significant. 58

80 CHAPTER THREE RESULTS 3.1. Chromogenic substrate assay for blood clotting factors The wide synthesis of chromogenic substrates and their subsequent use in hemostasis testing has opened new perspectives for the assay of many coagulation factors, fibrinolytic components and inhibitors. As thrombin and FXa represent the central enzymes in the coagulation cascade, the determination of their production is considered to be a powerful tool for probing over all blood coagulability. One of the goals of the present study was to modify, improve and optimize a kinetic photometric assay for the determination of thrombin and blood clotting FXa with regard to types of blood samples, analytic corrections, and activation reagents, as a tool of diagnosis of a hyper- or hypocoagulable state, verification of efficacy of oral anticoagulants in patients and, subsequently drug monitoring. Since it is well known that the reaction with a chromogenic substrate is sensitive to changes of ph, temperature, composition of buffer as well as to reagents added to the sample to be analyzed, it is necessary to establish optimal conditions before proceeding to the application of a chromogenic assay. To this purpose, following optimization studies, thrombin and FXa production has been determined in 00 male and female volunteers (aged from 19 to 75 years). The assay conditions chosen are mentioned in materials and methods Thrombin production assay In our modified protocol for the determination of thrombin production, the maximal amount of generated thrombin in non-clotted and therefore clotted plasma samples can be measured easily in a few minutes. The continuous determination of thrombin production in response to the activation time is shown in Figure 3.1 for eight different platelet rich in plasma samples from healthy donors. Figure 3.1 shows that thrombin production kinetics may vary from one person to other (interindividual variation). Figure 3.2 represents discontinuous determination of platelet-dependent thrombin production every 45 sec for 6 min in 50 different samples. It is clearly indicated in the curve, that thrombin generation after activation of the reaction is rapidly increased and reaches a peak value in about 50 sec (Tmax); the amount of produced thrombin decreased gradually thereafter. Figure 3.3 shows discontinuous determination of platelet-dependent thrombin production for increasing time intervals in 10 different human samples. 59

81 Figure 3.1: Determination of continuous thrombin production in eight healthy donors. Figure 3.2: Determination of average thrombin production for increasing time intervals (0-6 min, measuring every 45 s ) in fifty human subjects. 60

82 Figure 3.3: Determination of discontinuous thrombin production for increasing time intervals (0-14 min, measuring every 30 s) in ten different human samples Blood clotting FXa production assay In Figure 3.4 the continuous production of FXa was determined in eight healthy subjects with the modified and optimized chromogenic substrate assay. The inter-individual variation concerning FXa production is also obvious in Figure 3.4. In Figure 3.5 the discontinuous production of FXa for increasing time intervals was determined within a six min reaction period in 10 different samples. FXa production, following activation of the reaction increased rapidly and reached to a peak value after about 110 sec and the amount of produced FXa decreased gradually thereafter. Figure 3.4: Determination of continuous production of blood clotting FXa in nine healthy donors. 61

83 Figure 3.5: Determination of FXa production for increasing time intervals (0-6 min, measuring every 45 s ) in ten normal subjects Influence of platelet concentration on the process of hemostasis The effect of platelet concentration on thrombin production was studied in the presence of TF. The study was performed in PRP (Plasma Rich in Platelets), prepared after centrifugation of citrated whole blood at 1200 rpm for 5 min at RT. The (¾)supernatant PRP was removed and the platelet number was adjusted to different con by dilution with PPP (Plasma Poor in Platelets) obtained after further centrifuging the remaining blood at 4000 rpm for 15 min. To study the influence of platelet con on thrombin production, in PRP prepared from 50 randomly selected healthy volunteers, the plasma specimens were adjusted to /l, /l, /l, /l and /l platelet by dilution with PPP. Thrombin production, as shown by the continuous determination of platelet-dependent thrombin in 50 healthy volunteers, in response to the activation time, in /l, /l, /l, /l and /l platelet con (Figure 3.6), slowly increased upon increased platelet count. However, Figure 3.7 shows the difference between PRP and PPP in active clot formation. In the presence of platelet concentration /l (PPP) the thrombin generation was significantly (P< 0.05) lower as compared to the observed in the presence of platelet concentration (PRP). As clearly shown in Figure 3.7, platelets actively participate in clot formation as well as thrombin production. The percentages of increased thrombin produced with increasing platelet con were 7, 9, 12, 15 and 16% corresponding to /l, /l, /l, /l and /l platelet concentration, respectively. On the contrary, platelets didn t enhance FXa production (data not shown). 62

84 Figure 3.6: Influence of increasing platelet concentration on thrombin production (from bottom to top, the platelet con was: (1) /l, (2) /l, (3) /l, (4) /l and (5) /l. Figure 3.7: Comparison between continuous thrombin production in PPP and PRP in the presence of TF Influence of activated FVII on the clotting mechanism To determine whether FVIIa enhances clot formation, blood clotting FXa and thrombin production were measured simultaneously in random healthy specimens as an indirect way to study the effect of FVIIa, before and after addition to the reaction of rfviia (600µg/ml). In the absence of FVIIa significant amounts of thrombin and FXa were produced, but in the presence of FVIIa thrombin (Figures 3.8 and 3.9) and FXa (Figures 3.10 and 3.11) production was drastically enhanced, thus confirming that the nature process of hemostasis was strongly dependent on the presence of FVIIa. 63

85 Figure 3.8: Determination of continuous thrombin production before and after activation of the reaction with the addition of rfviia. Figure 3.9: Discontinuous assay for the determination of thrombin production for increasing time intervals (0-6 min) before and after activation of the reaction with rfviia. 64

86 Figure 3.10: Determination of continuous blood clotting FXa production before and after activation of the reaction with rfviia. Figure 3.11: Discontinuous assay of FXa production at various time points before and after addition of rfviia Influence of tissue factor on the coagulation process The effect of TF on blood clotting was investigated by determining thrombin and FXa production in PRP (Plasma Rich in Platelets) prepared from 50 normal healthy subjects in the absence or presence of TF. Thrombin and FXa production was determined before the addition of TF and after activation of the reaction through monitoring absorbance at 405 nm every 30 s over a 10 min time course. Both thrombin and FXa production was enhanced in fresh PRP from healthy 65

87 individuals without any TF addition as well as in the presence of various dilutions (1/200, 1/500, 1/1000, 1/2000) of Thromborol S (TF). However, while increasing the con of TF resulted in a continuous increase of FXa and thrombin production (Figure 3.12 and 3.13). Figure 3.12: Thrombin production with increasing concentrations of TF. Figure 3.13: Determination of continuous FXa production with increasing concentrations of TF. 66

88 Thrombin formation assay in healthy people and patients with diabetes mellitus (D.M) The modified method for determining thrombin production was also used for calculating thrombin activity in plasma samples of healthy individuals and patients suffering from D.M. It was shown that a hypercoagulable state can be diagnosed using our modified method, while the modified method was also able to test the efficiency of oral anticoagulants in patients. The mean±se of platelet-dependent thrombin levels after 3 min of reaction, following addition of CaCl 2 were 157.7±1.9 in healthy people, and 310±3 miu/ml in hospitalized diabetic patients with complications respectively Table 3.1. In an additional subset of studies, the effect of anticoagulant therapies on thrombin production was investigated by measuring platelet-dependent thrombin production in patients treated with Sintrom, Salospire and Salospire+Warfarin (Table 3.1). The mean±se of plateletdependent thrombin levels were 279±2.4, 291±0.9 and 297±1.1 miu/ml in hospitalized diabetic patients who had been treated with Sintrom, salospire and salospire+warfarin respectively. The data analysis of platelet-dependant thrombin production indicated statistically significant difference (P<0.01) in mean concentrations of thrombin in diabetic patients who were under no treatment or under anticoagulation therapy (Table 3.1). In addition, as shown in Table 3.1, thrombin production remained high even after antithrombotic therapies. Table 3.1: Thrombin activity in healthy people and patients with diabetic mellitus without or under a anticoagulant treatment Studied groups Thrombin(mIU/ml) Range No. Statistical Mean ±SE evaluation Normal people 157.7± Patients with D.M without antithrombotic therapies 310± P<0.01 Patient under Sintrom treatment 279± P<0.01 Patients under Salospire treatment 291± P<0.01 Patients under Salospire +Warfarin 297± P<0.01 Pb statistics obtained by student - t- test versus normal group 67

89 Effect of fibrin clot on thrombin and FXa generation measurment The effect of fibrin clot on both thrombin and FXa production assay was investigated by measuring thrombin and FXa production before and after clot removal from 60 normal human subjects. The results of the comparison between thrombin and FXa levels in plasma samples beore and after clot remonal are depicted in Table 3.2. The mean±s.e values of thrombin and FXa were 405±3.5 and 215±5.6 (miu/ml), respectively, in clot-removed samples, while in samples in which clot was not removed the mean±s.e values of thrombin and FXa were 170.6±1.7 and 251±2.7 (miu/ml), respectively. A statistically significant difference was observed in thrombin levels before and after clot removal (P<0.01), whereas the levels of FXa did not change in a statistically significant manner (P<0.05) (Table 3.2). Table 3.2: Determination of thrombin and FXa production before and after clot removal Studied Parameters (miu/ml) Clot removed samples Range Non removed clot samples Range No. Statistical evaluation Mean ±SE Mean ±SE Thrombin 405± ± P<0.01 FXa 215± ± N.S Pb statistics obtained by student - t- test. N.S =Non significant difference Correlation between thrombin and FXa production In recent years the biochemistry and physiology of blood coagulation have been subjected to intensive studies because of the enormous clinical importance of this subject. It is obvious that thrombin and FXa serine protease are the most important enzymes involved in blood coagulation, playing a central role in the coagulation cascade. In a following step we made an attempt to calculate the correlation coefficient between the production rates of thrombin and FXa. Figure 3.14 shows the correlation coefficient between plasma thrombin and blood clotting FXa. According to our data a strong correlation can be observed between thrombin and FXa in plasma samples of healthy individuals. 68

90 Figure 3.14: Correlation coefficient between FXa and thrombin production rates Influence of moderate physical exercise on thrombin and FXa production Both thrombin and FXa production were studied in 30 healthy individuals before and after moderate physical exercise for 15 and 120 min. The mean±se of plasma thrombin and FXa before moderate exercise were 155.8±1.5 and 143±1.5 miu/ml, respectively (Table 3.3). The mean±se of thrombin following 15 and 120 min of moderate physical exercise were 171.9±1.7, 192.6±2.3 miu/ml, respectively, while the respective mean±se of FXa were 161±1.2, 182.6±1.6 miu/ml (Table 3.3). It is therefore clearly shown that physical exercise results in an increase in thrombin and FXa production (see also Figure 3.15). Table 3.3: Platelet-dependent thrombin and FXa production before and after 15 and 120 min of moderate physical exercise Exercise period Thrombin miu/ml Range %* FXa miu/ml Range %* No Mean ±SE Mean ±SE Before 155.8± N.A 143± N.A 30 After15 min 171.9± ± After 120 min 192.6± ± *% percentage of increase 69

91 Figure 3.15: Platelet-dependent continuous thrombin production before and after 15 and 120 min of physical exercise Western blotting analysis for the detection of blood clotting FX and it s active form Development of western blotting analysis SDS-PAGE has proven to be among the most useful tools yet developed in the area of molecular biology. SDS-PAGE separation of proteins and peptides makes possible to quantify the amount of a particular protein/peptide in a sample, to obtain fairly reliable molecular mass information, and, by combining SDS-PAGE with immunoelectroblotting, to evaluate the antigenicity of proteins and peptides. We developed a western blotting technique in order to visualize and monitor the activation of zymogen FX to serine protease FXa. By this mean we wanted to study the blood clotting mechanism and furthermore, to test whether the blood clotting FX could be a target in clinical assays to control the blood clotting cascade. For this purpose the proteins (FX and FXa) were analyzed by SDS-polyacrylamide gel electrophoresis and western blotting. Figures 3.16, 3.17 and 3.18 show western blotting analysis of 7 different samples in 3 different acrylamide concentration (12, 8 and 10 % (w/v), respectively) by using specific antihuman polyclonal antibody directed against FX. Following western blotting, an immunoreactive band, at about 59 KDa appeared in the membranes. In Figure 3.18 (line 7) a sample deficient in FX was analyzed, thus verifying that the 59 kda band indeed corresponds to FX. Finally, 10% (w/v) analysis on a acrylamide gel has been chosen for all the remain study because, as compared to other acrylamide concentrations, FX and FXa bands appear clearer, and easier to analyze with the Pro-Gel analysis quantifying program. 70

92 Figure 3.16: Western blotting analysis of FX following 12% (w/v) SDS-polyacrylamide gel electrophoresis in 7 different samples using a specific anti-human polyclonal antibody against FX. Figure 3.17: Western blotting analysis of FX following 8% (w/v) SDS-polyacrylamide gel electrophoresis in 7 different samples using a specific anti-human polyclonal antibody against FX. Figure 3.18: Western blotting analysis of FX following 10% (w/v) SDS-polyacrylamide gel electrophoresis in 8 different samples using a specific anti-human polyclonal antibody targeting FX. Line 7: DFX, sample deficient in FX. 71

93 Western blotting analysis for the determination of zymogen FX and its activated form, the serine protease FXa Due to its central position (FX) in coagulation at the junction where the extrinsic and intrinsic pathways converge, tight regulation of FXa activity is critical for maintaining the delicate hemostatic balance (140). To visualize the activation pattern of blood clotting FX by recombinant TF and activated FVII, SDS-PAGE followed by immunoblotting was employed (Figure 3.20). To monitor blood clotting FX and its released peptide FXa, at various incubation times the specific anti-human monoclonal antibody directed against the heavy chain of FX and FXa was used. Activation of blood clotting FX was achieved by the addition of TF-FVIIa in Tris HCl buffer. At the beginning of the reaction, CaCl 2 was added to the incubation mixture. Incubation was performed at 37 C for 10 min. Aliquots were removed from the reaction at various time intervals, supplemented with SDS loading buffer, and analyzed by 10% (w/v) SDS-PAGE and Westrn blotting. We observed that blood clotting FX, which appears as an immunoreactive band at 59 kda, is progressively transformed to its active form, the serine protease FXa, which appears as an immunoreactive band at 47 kda (Figures 3.19 and 3.20). Figure 3.19: SDS-PAGE and Western blotting analysis of factors X and Xa for increasing time intervals (0, 5 and 10 min) using a specific anti-human monoclonal antibody recognizing the heavy chain of FX and FXa. Analysis of the immunoreactive bands using the Pro-Gel program clearly shows an increased ratio of FXa/FX (11/89 at 0 min, 40/60 at 5 min and 57/43 at 10 min). 72

94 Figure 3.20: SDS-PAGE and Western blotting analysis of factors X and Xa for increasing time intervals (0, 2, 4, 6, 8, 10, 12, 14 and 16 min) using a specific anti-human monoclonal antibody recognizing the heavy chain of FX and FXa. Analysis of the immunoreactive bands using the Pro-Gel program clearly shows an increased ratio of FXa/FX (30/70 at 0 min, 48/52 at 2 min, 52/48 at 4 min, 56/44 at 6 min, 59/41 at 8 min, 64/38 at 10 min, 66/36 at 12 min, 66/34 at 14 min and 69/31 at 16 min). In a following step we wanted to compare fresh and frozen-thawed plasma for the assessment of blood clotting factors FX and FXa. Figure 3.21 clearly shows that the appearance of FX and its active form were strongly affected following freezing. In fresh samples two clear bands appear normally, whereas in freezed-thawed samples several bands appear, as well as a characteristic low molecular weight degradation product. Figure 3.21: Western blotting analysis of FX and FXa in nine freezed and thawed samples using a specific anti-human monoclonal antibody against FX and FXa. 73

95 3.3. Cloning, expression and purification of the vwf-a1 domain and study of its function in hemostasis The main objective of this part of study was to express and determine the potential function of the A1 domain of vwf in platelet activation and hemostasis Cloning and expression of the vwf-a1 domain in 293T human embryonic kidney cells Construction of the vwf-a1 expression vector A. Amplification of the DNA fragment expressing vwf-a1 domain The strategy for constructing the vwf-a1 expression plasmid was first to amplify the respective DNA fragment of vwf expressing this region and then insert the amplified sequence in frame into the cloning vector. The complete cdna of human vwf was used as a template to amplify the region of interest (vwf-a1). The primers were designed to contain EcoR1 and BamH1 sites at the 5 end and 3 end, respectively. PCR amplification was performed in a total volume of 50 µl using Taq DNA polymerase as described in Material and Methods. The PCR products were first purified using phenol-chloroform extraction and ethanol precipitation to remove protein contaminates and other salts and then double-digested with BamH1 and EcoR1 restriction enzymes to get the appropriate protruding ends for cloning into the expression vector pcdna.3.1/myc-his(-)b. The digested products were analyzed on a 1.2% (w/v) agarose gel and the expected size fragments (531 kb) were obtained (Figure 3.22). Figure 3.22: Analysis of purified, double digested PCR products corresponding to vwf-a1 domain for cloning into the pcdna.3.1/myc-his(-)b expression vector. 74

96 B. Cloning of the insert into the expression vector pcdna.3.1/myc-his(-)b Approximately 2 µg of plasmid pcdna.3.1/myc-his(-)b were double-digested using 1 µl of BamH1 and EcoR1 restriction enzymes and digested products were analyzed on a 1.2% (w/v) agarose gel (Figure 3.23). The expected sizes of fragments (86 and 5434 bp) were obtained with negligible amounts of undigested vector. Figure 3.23: Analysis of BamH1/EcoR1 digested pcdna.3.1/myc-his(-)b on a 1.2% (w/v) agarose gel. The amplified DNA fragment encoding for vwf-a1 was ligated to the BamH1/EcoR1 sites of the vector at a ratio of 3:1 (90 ng of insert and 30 ng of vector) using 1.2 µl ligase (Takara ligase). The ligated vectors were transformed into competent E. coli TOP10 cells that were grown in LB agar overnight in the presence of 100 µg/ml ampiciline. The plasmid DNA was purified using the alkaline lysis method and double-digested with BamH1 and EcoR1 to confirm the presence of insert. The digestion mixture was analyzed on a 1.2% agarose gel (Figure 3.24). Figure 3.24: Analysis of purified restriction double-digested recombinant pcdna.3.1/myc- His(-)B-vWF-A1 expression vector on a 1.2% (w/v) agarose gel. DNA markers are shown on the left. Lines 1, 2, 3, 4, 5, correspond to recombinant expression vector. 75

97 Protein expression and purification The recombinant pcdna.3.1/myc-his(-)b-vwf-a1 vector was transfected into logarithmically growing 293T HEK cells. The protein was overexpressed in 293T HEK cells at a level of ~ 20% of total cellular protein. The expression of vwf-a1 was determined by SDS- PAGE analysis (Figure 3.25 lanes 2 and 4). As control we used cell extracts from 293T cells that were transfected with the vector pcdna.3.1/myc-his(-)b alone (Figure 3.25, lines 1 and 3). Figure 3.25: SDS-PAGE analysis of 293T HEK total cell extracts. Molecular weight markers are shown on the left (kda). Lines 1, 3 extracts from 293T cells transfected with pcdna.3.1/myc-his(-)b. Lines 2, 4 extracts from 293T cells transfected with pcdna.3.1/myc- His(-)BvWF-A1. The recombinant fusion protein was extracted and purified to near homogeneity by (Ni- ANT) affinity chromatography (Figure 3.26). The protein purification system was carried out as described in Materials and Methods and was based on the remarkable selectivity of the Ni-NTA resin for proteins carrying a small affinity tag consisting of 6 consecutive histidine residues. The recombinant vwf-a1 protein was analyzed on a 10% (w/v) SDS-PAGE gel under reducing conditions and stained with coomassie brilliant blue. As anticipated, the recombinant vwf-a1 protein displayed a single band at ~20 kda, while no other bands than vwf-a1 were seen on the gel, suggesting that our purification protocol resulted in a nearly homogeneous preparation. 76

98 Figure 3.26: Purification of vwf-a1 domain recombinant protein by affinity Ni-NTA chromatography. The sonicated cell lysate was loaded on Ni-NTA affinity chromatography and the bounded protein was eluted with urea ph 4.5. Various eluted fractions from the column were collected and analyzed on a 10% (w/v) SDS-PAGE gel. Line 1, purified recombinant protein. Lines 2, and 3 eluted sample after first and second washing of the column with buffer A and B respectively Western blotting analysis Following SDS-PAGE, Western blotting analysis was performed using a commercially available mouse-anti His monoclonal antibody to react with the 6xHis tag of the recombinant protein. As shown in Figure 3.27, a singe immunoreactive band at ~20 kda, identical with the one observed with Coomassie blue staining, was observed. Figure 3.27: Western blotting analysis of vwf-a1 expressed in 293T HEK cells. 77

99 Cloning and expression of the A1 domain of human vwf in Escherichia coli strain BL21 (DE3) Constraction of the vwf-a1 expression vector A. Amplification of the DNA fragment expressing vwf-a1 domain The complete cdna of human vwf was used as a template to amplify the region of interest (vwf-a1). The primers were designed to contain NdeI and XhoI sites at the 5 end and 3 end, respectively. PCR amplification was performed in a total volume of 50 µl using Taq DNA polymerase as described in Material and Methods. The PCR products were first purified using phenol-chloroform extraction and ethanol precipitation to remove protein contaminates and other salts and then double-digested with NdeI and XhoI restriction enzymes to get the appropriate protruding ends for cloning into the expression vector pet-29c(+). The digested products were analyzed on a 1.5% (w/v) agarose gel and the expected size fragments (531 kb) were obtained (Figure 3.28). Figure 3.28: Analysis of purified, double-digested PCR products corresponding to vwf-a1 domain for cloning into the pet-29c(+) expression vector. M, DNA markers in kb. B. Cloning of the insert into the expression vector pet-29c(+) Approximately 2 µg of plasmid pet-29(+) were double-digested using 1 µl of NdeI and XhoI restriction endonucleases and digested products were analyzed on a 1.2% (w/v) agarose gel (Figure 3.29). The expected sizes of fragments (86 and 5434 bp) were obtained. 78

100 Figure 3.29: Analysis of NdeI/XhoI digested pet-29c(+) on a 1.2%(w/v) agarose gel. The amplified DNA fragment encoding for vwf-a1 was ligated to the NdeI/XhoI restriction sites of pet-29c(+) at a ratio of 3:1 using 1.2 µl T4 DNA ligase. The ligated vectors were transformed into competent E. coli TOP10 cells that were grown in LB agar overnight in the presence of 100 µg/ml kanamycin. The plasmid DNA was purified using the alkaline lysis method and double-digested with NdeI and XhoI to confirm the presence of insert. The digestion mixture was analyzed on a 1.2% (w/v) agarose gel (Figure 3.30). Figure 3.30: Analysis of purified restriction double-digested recombinant pet-29c(+)-vwf- A1 expression vector on a 1.2% (w/v) agarose gel. DNA markers are shown on the left. Lines 4, 5, 6, 7, 8, correspond to recombinant expression vector Protein expression and purification E. coli BL21 (DE3) were transformed with the recombinant pet-29c(+)-vwf-a1 vector. Overexpression was induced with 1 mm IPTG. The expression of vwf-a1 was detrmined by SDS-PAGE analysis (Figure 3.31 lanes 4, 6 and 8). 79

101 Figure 3.31: SDS-PAGE analysis of E. coli total extracts. Molecular weight markers are shown on the left (kda). Lines 1 and 2 extracts from uninduced E. coli cells. Lines 3, 5 and 7 extracts from E. coli following induction with IPTG at 16 ºC for 5h. Lines 4, 6 and 8 extracts from E. coli following induction with IPTG at 37 ºC for 5h. vwf-a1 migrates as a band at the expected M.W. (~20 kda). The recombinant fusion protein was extracted and purified to near homogeneity by (Ni- ANT) affinity chromatography (Figure 3.32). vwf-a1, carrying a 6xHis affinity tag, was analyzed on a 10% SDS-PAGE gel under reducing conditions and stained with coomassie brilliant blue. As anticipated, the recombinant vwf-a1 protein displayed a single band at ~20 kda. Figure 3.32: Purification of recombinant vwf-a1 domain containing a 6xHis tag by affinity Ni-NTA chromatography. The sonicated cell lysate was loaded on Ni-NTA affinity chromatography and the bounded protein was eluted with 6 M urea. Various eluted fractions from the column were collected and analyzed on a 10% (w/v) SDS-PAGE gel. Line 1, total cell extract, line 2 eluted sample following washes of the column, and lanes 3 and 4, purified recombinant protein 4 and 5 eluted sample after first and second washing of the column with buffer A and B respectively. 80

102 Western blotting analysis Following SDS-PAGE, Western blotting analysis was performed using a commercially available mouse-anti His monoclonal antibody to react with the 6xHis tag of the recombinant protein. As shown in Figure 3.33, a singe immunoreactive band at ~20 kda, identical with the one observed with Coomassie blue staining, was observed. Figure 3.33: Western blotting analysis of vwf-a1 expressed in E. coli BL21 (DE3) cells Potential function of recombinant vwf-a1 domain protein in platelet activation and the natural process of hemostasis In an attempt to elucidate the potential function of vwf-a1 protein in platelet activation and therefore in the process of hemostatic plug formation, through its potential interaction with the GpIb IX V complex, the His-tagged proteins were tested for their ability to affect thrombin production in PRP and PPP samples using the chromogenic substrate S2238. Thrombin production was determined before and after the addition of the bacterial and mammalian expressed recombinant vwf-a1 domain proteins in fresh PRP and PPP samples from 28 healthy individuals. The PRP mean±se values of platelet-dependent thrombin production at 3 min before and after the addition of 293T HEK cells-derived recombinant vwf-a1 protein were 149±0.9 and 436±1.3 miu/ml respectively, with a range of variation (33-310) and (42-588) miu/ml respectively, while the respective values in PPP were 122±1.1 and 165±0.8 miu/ml with a range of variation (23-276) and (39-350) miu/ml (Figure 3.34). However, E. coli B21(DE3)-derived recombinant vwf-a1 domain protein exhibited a weaker activation of thrombin production and the process of clotting mechanism. The mean ±SE values of platelet-dependent thrombin production after addition of the E. coli-derived recombinant vwf-a1 protein were 169±1.1 and 132±0.9 miu/ml with a rage of variation (37-320) and (25-280) in PRP and PPP samples, respectively (Figure 3.34). It is therefore clear from our data that addition of the vwf-a1 domain activates clot formation and significantly increases thrombin production (P<0.01), with the mammalian- 81