The formation of a fibrin clot at the site of an injury to the wall of a normal blood vessel is an essential part of the process to stop blood loss after vascular injury. The reactions that lead to fibrin clot formation are commonly described as a cascade, in which the product of each step is an enzyme or cofactor needed for following reactions to proceed efficiently. The entire clotting cascade can be divided into three portions, the extrinsic pathway, the intrinsic pathway, and the common pathway. The extrinsic pathway begins with the release of tissue factor at the site of vascular injury and leads to the activation of factor X. The intrinsic pathway provides an alternative mechanism for activation of factor X, starting from the activation of factor XII. The common pathway consists of the steps linking the activation of factor X to the formation of a multimeric, cross-linked fibrin clot. Each of these pathways includes not only a cascade of events that generate the catalytic activities needed for clot formation, but also numerous positive and negative regulatory events.
View original pathway at Reactome.
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Haemophilia A-associated mutations within the predicted A1-A2 and A1-A3 interface that have the molecular phenotype of increased rate of inactivation of FVIIIa due to increased rate of A2 subunit dissociation.
Fibrinogen is a hexamer, containing two fibrinogen alpha chains, two fibrinogen beta chains, and two fibrinogen gamma chains, held together by disulfide bonds.
Fibrin is a hexamer of two fibrinogen alpha chains, two fibrinogen beta chains, and two fibrinogen gamma chains, held together by disulfide bonds. It is formed in vivo by the thrombin-catalyzed removal of amino terminal fibinopeptides from the A alpha and B beta chains of fibrinogen. This fibrin hexamer ("fibrin monomer") is the subunit that multimerizes to form a fibrin clot ("fibrin multimer").
The fibrin "monomers" formed by the action of thrombin on fibrinogen associate spontaneously into multimers. This association can follow several distinct pathways and may be able to form several types of higher-order structures. All of these possibilities are represented in Reactome as a fibrin trimer.
Activated thrombin cleaves the A chains of factor XIII tetramers in a reaction stimulated by the presence of fibrin multimers (Lewis et al. 1985). The amino terminal portions of the A chains are released as activation peptides, which have no known function. The resulting factor XIII tetramer remains catalytically inactive.
The membrane-bound Va:Xa (prothrombinase) complex rapidly activates large amounts of thrombin. Factor Xa (aka Factor X heavy chain), a cleavage product of coagulation factor X (F10), is a vitamin K-dependent glycoprotein able to convert prothrombin to thrombin during the blood clotting process (Mann et al. 1988, Orfeo et al. 2004). Factor Xa is a target for direct oral anticoagulant (DOAC) drugs that are direct factor Xa inhibitors (the so-called 'xabans') and used in the treatment and prevention of thromboembolic disorders (Galanis et al. 2014).
Factors Va and Xa associate on a membrane surface to form a complex in which the activity of factor Xa on prothrombin is greatly increased (Mann et al. 1988). The presence of negatively charged phospholipid in the membrane greatly facilitates this process, a feature that may contribute to its localization, as such phospholipids are normally on the cytosolic face of the plasma membrane (Devaux 1992), but could be exposed to the extracellular space following platelet activation or mechanical injury to endothelial cells.
Activated thrombin (factor IIa) catalyzes the conversion of factor V to factor Va (activated factor V). The activation peptide released in this reaction has no known function.
Membrane-bound factor Xa catalyzes the activation of small amounts of thrombin. The amino terminal portion of prothrombin is released as an activation peptide, which can be cleaved further by activated thrombin. Neither the full-length activation peptide nor its cleavage products have known functions.
Factor Xa (aka Factor X heavy chain), a cleavage product of coagulation factor X (F10), is a vitamin K-dependent glycoprotein able to convert prothrombin to thrombin during the blood clotting process. Factor Xa is a target for direct oral anticoagulant (DOAC) drugs that are direct factor Xa inhibitors (the so-called 'xabans') and used in the treatment and prevention of thromboembolic disorders (Galanis et al. 2014).
Factor VIIa, bound to tissue factor at the endothelial cell surface (the "extrinsic tenase complex"), catalyzes the formation of activated factor X with high efficiency. The amino terminal part of the heavy chain of factor X, the factor X activation peptide, is released. (This peptide has no known function.)
Tissue factor exposed at the endothelial cell surface forms a Ca2+-dependent complex with factor VIIa (FVIIa, activated FVII) from the plasma (Butenas S et al. 1994; Sabharwal AK et al. 1995; Banner DW et al. 1996; Bajaj SP et al. 2006; Persson E & Olsen OH 2011; Madsen JJ et al. 2015). FVIIa is allosterically activated by TF, which increases the amidolytic activity of FVIIa several-fold by restructuring the active site region (Butenas S et al. 1994; Higashi S et al. 1996; Sorensen AB et al. 2019). FVIIa, bound to TF at the endothelial cell surface, catalyzes the formation of activated factor IXa (FIXa) and FXa with high efficiency leading to thrombin generation and fibrin formation (Vadivel K & Bajaj SP 2012).
Tissue factor (TF), also known as thromboplastin or CD142, is an integral transmembrane glycoprotein, that functions as a co-factor for coagulation factor VII (FVII) and FVIIa (Broze GJ et al. 1985; Nemerson Y & Repke D 1985; Rao LV & Rapaport SI 1988). The TF:FVIIa complex is the primary activator of the coagulation protease cascade. The formation of TF:FVIIa complex triggers the coagulation cascade by activating both FIX and FX through their limited proteolysis. TF is expressed on the surface of several cell types located in subendothelial structures throughout the vasculature, and it is normally not in contact with circulating blood, where other coagulation factors are present in their inactivated forms (Drake TA et al. 1989; Wilcox JN et al. 1989; Fleck RA et al. 1990). Cells that are normally not exposed to the flowing blood, such as smooth muscle cells, constitutively express TF on their surface (Drake TA et al. 1989; Fleck RA et al. 1990). Upon vascular injury, through physical damage of the endothelial layer of the blood vessel, TF becomes exposed to circulating blood and the extracellular part of TF binds FVII with very high affinity and specificity. Infectious and inflammatory disease conditions induce TF expression, either in circulating blood cells or vascular endothelial cells, by activation of TF gene transcription (van den Eijnden MM et al. 1997; Osterud B & Bjorklid E 2012). Induced expression of TF by monocytes in response to infection is thought to be a part of the innate immune response to limit the dissemination of pathogens by trapping them inside clots (van der Poll T & Herwald H 2014).
Irrespective of the cellular source of TF and whether it is induced or constitutively expressed, most of the TF expressed on the surfaces of resting cells exists in a cryptic coagulant-inactive state (Schecter AD et al. 1997; Bach RR 2006; Kothari H et al. 2013; Grover SP & Mackman N 2018). The encrypted TF can bind to FVIIa, but the assembled TF:FVIIa complex fails to activate FIX and FX (Rao LV & Pendurthi UR 2012). Activation or disruption of cells markedly enhances cell surface TF procoagulant activity without altering TF antigen levels at the cell surface (‘decryption’). Several mechanisms have been proposed for TF decryption on cell surfaces, and out of them, externalization of phosphatidylserine (PS) to the outer leaflet and PDI-mediated thiol-disulfide exchange pathways that affect the allosteric disulfide bond in TF seem most likely (Rao LV & Pendurthi UR 2012; Grover SP & Mackman N 2018; Ansari SA et al. 2019). The presence of a high molar concentration of sphingomyelin (SM) in the outer leaflet of the plasma membrane inhibits TF procoagulant activity on the cell surface, thus maintaining TF in an encrypted state in resting cells (Wang J et al. 2017). Acid-sphingomyelinase (ASM)-mediated hydrolysis of SM following cell injury removes the inhibitory effect of SM on TF activity, thus leading to TF decryption (Wang J, et al. 2017; Ansari SA et al. 2019). It has been suggested that the coordinated effects of SM hydrolysis, PS externalization and thiol-disulphide exchange pathways are responsible for full cellular activation of TF (Ansari SA et al. 2019). However, molecular links among various pathways of TF decryption are not fully known yet. The Reactome event describes exposure of TF sequestered in the wall of a blood vessel to flowing blood.
Factor VII, bound to tissue factor at the endothelial cell surface, catalyzes the activation of factor X from plasma with moderate efficiency. The amino terminal part of the heavy chain of factor X, the factor X activation peptide, is released. (This peptide has no known function.)
Coagulation factor VII circulates in plasma mostly in the zymogen form (FVII); about 1% of plasma FVII is found in the active form (FVIIa) (Morrissey JH et al. 1993). Initiation of coagulation begins by exposure of blood (which contains both zymogen FVII and activated FVIIa) to tissue factor (TF) in the extravascular space at an injury site and formation of the Ca2+-dependent complex between TF and plasma FVII/FVIIa (Kelley RF et al. 2004; Ruf W et al. 1991). The TF:FVII zymogen complex has low but measurable proteolytic activity on factor X, suggesting that this complex initiates TF-dependent clotting through a minimal generation of factor Xa, which in turn catalyzes the activation of FVII from plasma. (Rao LV et al. 1986). As factor VIIa accumulates, TFr:FVIIa complexes also form, accelerating the process (Nemerson 1988). Formation of the TF:FVIIa complex greatly increases the enzymatic activity of FVIIa via allosteric interactions between TF and FVIIa, as revealed by a 20- to 100-fold increase in the rate of amidolysis of small, chromogenic peptidyl substrates (Broze GJ Jr & Majerus PW.1980; Butenas S et al. 1994; Higashi S et al. 1996). A second model, building on the observation that normal plasma contains low levels of activated FVIIa constitutively, suggests that complexes with FVIIa form immediately at the onset of clotting (Rapaport and Rao 1995). The two models are not mutually exclusive, and in any event, the central roles of TF and FVIIa in generating an initial supply of factors IXa and Xa, and the self-limiting nature of the process due to the action of TFPI, are all well-established.
The human gene SERPINC1 produces antithrombin III, the most important serine protease inhibitor in plasma that regulates the blood coagulation cascade (van Boven & Lane 1997). Antithrombin III binds to membrane-associated low molecular weight heparins (LMWHs) and their derivatives (SERPINC1 activators) on the surface of normal endothelial cells. This binding increases the affinity of antithrombin III for thrombin approximately 1000-fold, inactivating thrombin and other proteases involved in blood clotting (e.g. factor Xa) and resulting in an overall decrease in clotting ability (Holmer et al. 1986, Eriksson et al. 1995, Mushunje et al. 2003).
The Covid-19 pandemic is an infection caused by the SARS-CoV-2 coronavirus. Severe cases of this infection can lead to acute respiratory distress syndrome and coagulation changes leading to a higher risk of thrombosis, especially pulmonary embolism (Susen et al. 2020). LMWHs may play a role as potential attachment factors for SARS-CoV-2 (Tandon et al. 2020), potentially reducing the incidence and/or severity of thrombosis (Marietta et al. 2020).
Conversion of factor IX (FIX) to FIXa requires proteolytic cleavages after Arg191 and Arg226, releasing an activation peptide (Ala192-Arg226) (Geng Y et al. 2012; Vadivel K & Bajaj SP 2012). This calcium-dependent reaction is catalyzed by factor VIIa (FVIIa) in the presence of tissue factor (TF) and phosphatidylserine-rich phospholipid (Osterud B, Rapaport SI 1977; Banner DW et al. 1996; Bajaj SP et al. 2006). In this reaction, FVIIa and FIX anchor to the phospholipid bilayer through their Gla domains for optimal rates of FIXa formation (Vadivel K & Bajaj SP 2012). Further, the N-terminal Gla and epidermal growth factor-like (EGF1) domains of FIX represent the primary recognition determinants in binding to FVIIa & TF and formation of the ternary complex (Zhong D et al. 2002; Vadivel K & Bajaj SP 2012). In the formed ternary complex, the scissile peptide bond sequence in FIX or FX then approaches the active site cleft in FVIIa and induces the formation of the oxyanion hole for efficient proteolysis (Vadivel K & Bajaj SP 2012). FVIIa, bound to TF at the endothelial cell surface, cleaves FIX first after Arg191, forming the inactive intermediate which is released from FVIIa. The intermediate form of FIX must rebind to the protease to be cleaved after Arg226 to form an activated FIXa. As the second cleavage is rate-limiting, the inactive intermediate accumulates during FIX activation by FVIIa. The proteolytic cleavage of FIX results in a two-chain protein consisting of a light chain (Gla-EGF1-EGF2 domains) and a heavy chain (protease domain with the catalytic center) held together by a single disulfide bond (Yoshitake S et al. 1985). The released activation peptide FIX (192-226) has no known function.
TFPI binds to the factor VIIa:TF complex and to factor Xa at the endothelial surface, forming a stable heterotetrameric complex in which factor VIIa is catalytically inactive.
The alpha and beta chains of fibrinogen hexamer are cleaved by thrombin to generate fibrin monomer (Ni et al. 1989). The amino terminal regions of the cleaved alpha and beta chains are released (fibrinopeptides A and B respectively).
Fibrin monomers rapidly and spontaneously associate into large multimers, binding to one another via sites created by fibrinopeptide release (Laudano and Doolittelle 1980). The process of multimerization, and the range of multimer structures that can form in vivo and in vitro, have been studied in detail (Doolittle 1984). Here, multimer size has arbitrarily been set to three fibrin monomers.
Once the A chains of the Factor XIII tetramer have been cleaved by thrombin, the complex dissociates and the resulting A chain dimer binds Ca++ (one per peptide monomer) to form activated factor XIII (factor XIIIa).
Fibrin multimers are stabilized by the formation of multiple covalent crosslinks between the side chains of specific lysine and glutamine residues in fibrinogen alpha and gamma chains, catalyzed by factor XIIIa.
Antithrombin III in the complex is cleaved by thrombin, thereupon undergoing a conformational change that stabilizes the thrombin:antithrombin III complex, trapping and inactivating the thrombin moiety.
The same conformational change that traps thrombin in its complex with cleaved antithrombin III also decreases the affinity of the latter for heparin, and the complex of cleaved antithrombin III and thrombin dissociates from the cell-bound heparin molecule.
Activated protein C cleaves peptide bonds in activated factor V (factor Va), converting it to an inactive form (factor Vi). APC proteolysis involves cleavage of the factor Va heavy chain at Arg-334 (306 if signal peptide is not included) and Arg-534 (506 with no signal peptide) (Nicolaes et al. 1985). Most factor Va molecules are initially cleaved at Arg-534, yielding a partially active intermediate, followed by complete inactivation through cleavage at Arg-334 (Kalafatis et al. 1994). Factor Xa inhibits Arg-534 cleavage but this effect is mitigated by Protein S (Norstrom et al. 2006). A mutation of the APC cleavage sites in Fv at Arg-534Gln a.k.a. FVLeiden is the most common identifiable hereditary risk factor for venous thrombosis among Caucasians (Camire 2011).
Thrombin complexed with thrombomodulin at the endothelial cell surface cleaves the heavy chain of protein C, generating activated protein C and an activation peptide. The activation peptide has no known function.
Activated thrombin (factor IIa) binds to thrombomodulin at the external face of the plasma membrane, forming a thrombin:thrombomodulin complex. In this complexed form, the activity of thrombin towards protein C is greatly increased, and as thrombomodulin is particularly abundant on the surfaces of endothelial cells, this association plays a major role in restricting clot formation.
Factor VIII (FVIII) binds to von Willebrand factor (vWF) to form a complex (Lollal P et al. 1988; Leyte A et al. 1989; Vlot et al. 1995). Antibody inhibition data, site-directed deletion and mutagenesis studies suggest that the acidic subdomain a3, C1 & C2 domains of the FVIII light chain together control high affinity binding to vWF (Foster PA et al. 1988; Leyte A et al. 1989, 1991; Shima M et al. 1993; Saenko EL et al. 1994; Saenko EL & Scandella D 1997; Jacquemin M et al. 2000). Structural studies using negative stain electron microscopy (EM) and hydrogen-deuterium exchange mass spectrometry (HDX-MS) have revealed that the vWF TIL’ domain interacts with the FVIII C1 domain, the vWF E’ domain bridges the vWF TIL’ and D3 domains, whereas the vWF D3 domain interacts with the FVIII C1 and C2 domains (Yee A et al. 2015; Chiu PL et al. 2015). In addition, HDX-MS experiments showed that the FVIII a3 subdomain residues V1689-D1697 are directly involved in the interaction (Chiu PL et al. 2015). A combination of NMR spectroscopy and isothermal titration calorimetry (ITC) confirmed direct interaction between the FVIII a3 region and the VWF TIL’ domain mapping it to the residues in the two β-sheet regions on the VWF TIL’ domain (Dagil L et al. 2019). Further, tyrosine sulfation at residue 1699 is required for the interaction of FVIII with vWF (Leyte A et al. 1991). In the absence of sulfation at Y1699 in FVIII, the affinity for vWF was reduced by 5-fold (Leyte A et al. 1991). The nuclear magnetic resonance (NMR) spectrum studies of the complex between FVIII and vWF showed significantly larger residue-specific chemical shift changes when Y1699 was sulfated further highlighting the importance of FVIII sulfation at Y1699 for the binding affinity to vWF (Dagil L et al. 2019). The significance of the sulfation of FVIII at Y1699 in vivo is made evident by the presence of a Y1699F mutation that causes a moderate hemophilia A, likely due to reduced interaction with vWF and decreased plasma half-life (van den Biggelaar M et al. 2011). The vWF stabilizes FVIII, which otherwise has a very short half-life in the blood stream (Kaufman RJ et al. 1988). The interaction of FVIII with vWF allows thrombin to activate the bound FVIII and impedes cleavage of the molecules of nonactivated FVIII by the proteases FXa and activated protein C (APC) (Hamer RJ et al. 1987; Hill-Eubanks DC & Lollar P 1990; Koedam JA et al. 1990; Nogami K et al. 2002). Furthermore, vWF prevents the nonspecific binding of FVIII to the membranes of activated human platelets (Nesheim M et al. 1991; Li X & Gabriel DA 1997).
Factor VIII is a heterodimer containing a heavy and a light polypeptide chain, generated by the proteolytic cleavage of a single large precursor polypeptide (Vehar et al. 1984). Several forms of the heavy chain are found in vivo, all functionally the same but differing in the amount of the B domain removed by proteolysis. The single form annotated here is the shortest one (Eaton et al. 1986; Hill-Eubanks et al. 1989).
It has been demonstrated in in vitro experiments that vWF facilitates the association of FVIII chains and the retention of procoagulant activity in the conditioned medium of cells producing FVIII (Kaufman RJ et al. 1988; Wise RJ et al. 1991). Similar data have been obtained for re-association of FVIII chains in solution (Fay PJ 1988). In vitro, von Willebrand factor (Titani et al. 1986) can form complexes with factor VIII with a 1:1 stoichiometry. The complexes that form in vivo, however, involve large multimers of von Willebrand factor and varied, but always low, proportions of factor VIII (Vlot et al. 1995). A stoichiometry of one molecule of factor VIII associated with 50 of von Willebrand factor is typical in vivo, and is used here to annotate the factor VIII:von Willebrand factor complex.
Factor VIII complexed to von Willibrand factor in the blood is cleaved into several smaller polypeptides that remain associated. The acidic polypeptide on the aminoterminal side of the A3 domain of the light chain is released, however, and as this polypeptide mediates the association of factor VIII with von Willibrand factor, the activated factor VIII is released. While several proteases are capable of catalyzing these cleavages in vitro, only thrombin is active on factor VIII:von Willibrand factor complexes under physiological conditions (Eaton et al. 1986; Hill-Eubanks et al. 1989; Lollar et al. 1988; Pieters et al. 1989).
Plasma factor XI binds to the platelet glycoprotein Ib:IX:V complex (Baglia et al. 2002; Greengard et al. 1986). In the body, this reaction occurs specifically on the surfaces of activated platelets, but not on endothelial cells (Baird and Walsh 2002). The stoichiometry of the platelet glycoprotein Ib:IX:V complex has not been established directly, but is inferred from the relative abundances of its components in platelet membranes (Modderman et al. 1992; Shrimpton et al. 2002).
Factor IXa, in a complex with factor VIIIa on the surfaces of activated platelets (the "intrinsic tenase complex"), catalyzes the formation of activated factor X with high efficiency. The amino terminal part of the heavy chain of factor X, the factor X activation peptide, is released. (This peptide has no known function.)
Prekallikrein (PK) associates specifically with kininogen (HK) on cell surfaces. In vivo, this reaction may occur primarily on the surfaces of endothelial cells in response to platelet activation (Lin et al. 1997; Motta et al. 1998; Mahdi et al. 2003).
Prekallikrein in a complex with kininogen and C1q binding protein on the plasma membrane is cleaved to generate active kallikrein, which remains bound to the complex. In the body, this reaction appears to occur on the surfaces of endothelial cells and may require the presence of activated platelets. Recent work indicates that the protease that cleaves prekallikrein under these conditions is prolylcarboxypeptidase. Although this enzyme was originally isolated from lysosomes (Odya et al. 1978; Tan et al. 1993), it is associated with plasma membranes of cultured human endothelial cells in vitro (Moreira et al. 2002; Shariat-Madar et al. 2002), and the purified recombinant enzyme efficiently cleaves prekallikrein (Shariat-Madar et al. 2004). In contrast factor XII, despite its activity on prekallikrein in vitro, appears not to be responsible for prekallikrein activation on the cell surface (Rojkjaer et al. 1998).
Factors VIIIa and IXa associate on cell surfaces to form a complex that very efficiently catalyzes the activation of factor X, the so-called "intrinsic tenase complex". In vitro, negatively charged phospholipids can provide an appropriate surface. In the body, the surface is provided by the plasma membranes of activated platelets (Gilbert and Arena 1996).
Factor XI, bound to the cell surface, is converted to activated factor XI (factor XIa). Chemically, this reaction involves the cleavage of a single peptide bond in each subunit of the factor XI homodimer; intra- and inter-chain disulfide bonds hold the resulting four polypeptides together (Bouma and Griffin 1977; Kurachi and Davie 1977; McMullen et al. 1991). In the body, this reaction occurs on the surfaces of activated platelets (Greengard et al. 1986; Baglia et al. 2002; Baird and Walsh 2002); when this reaction occurs as a step in the intrinsic ("contact") pathway of blood coagulation, it is catalyzed by activated factor XIIa (Kurachi and Davie 1977, Baglia and Walsh 2000) which in turn is generated through the interactions of factor XII, kallikrein, and kininogen on endothelial cell surfaces (Schmaier 2004).
The cleavage of kininogen (HK, high molecular weight kininogen) yields activated kininogen and the vasoactive peptide bradykinin (Kerbirou and Griffin 1979; Lottspeich et al. 1985; Kellerman et al. 1986). In vivo, this reaction is catalyzed by activated kallikrein, takes places within the kallikrein:kininogen:C1q binding protein tetramer complex on the endothelial cell surface, and results in the release of kallikrein and bradykinin (Motta et al. 1998).
Cleavage of a single peptide bond converts factor XII to activated factor XII (factor XIIa) (Fujikawa and McMullen 1983; McMullen and Fujikawa 1985). Identification of the catalytic activity or activities responsible for this cleavage has not been straightforward. Studies in vitro have demonstrated the autoactivation of factor XII as well as activation by kallikrein. Both reactions require the presence of negatively charged surfaces and are accelerated in the presence of kininogen (high molecular weight kininogen, HK) (Griffin and Cochrane 1976; Meier et al. 1977; Silverberg et al. 1980). Recent work suggests that factor XII activation in vivo may occur primarily on endothelial cell surfaces and that, as in vitro, association with kininogen may accelerate the reaction (Mahdi et al. 2002; Schmaier 2004), although alternative pathways and alternative mechanisms for associating factor XII with the cell surface have not been excluded (Joseph et al. 2001).
The mature factor IX (FIX) is secreted and circulates in the plasma as an inactive 57kDa zymogen form F9(47-461). Activation of FIX involves cleavage of two peptide bonds, at arginine 191 (R191-A192, the α-cleavage) and at arginine 226 (R226-V227, the β-cleavage), releasing an activation peptide (A192-R226) (Di Scipio RG et al. 1978; Zögg T & Brandstetter H 2009). The activation peptide has no known function. This calcium-dependent reaction is catalyzed by factor XIa (FXIa), bound to platelet glycoprotein (GP) Ib:IX:V on the platelet cell surface (Osterud B et al. 1978; Gailani D et al. 2001; Geng Y et al. 2012). Binding studies showed that FIX does not bind to FXIa in the absence of calcium (Geng Y et al. 2012). Structural studies suggest that both activation of factor XI and binding it to FIX induced conformational changes at the interface between the catalytic and the apple domains of the activated FXIa. The conformational changes of FXIa increased the accessibility to the apple 3 (A3) domain to enable FIX binding (Geng Y et al. 2012; Bar Barroeta A et al. 2019). FIX activation is ordered. FIX first binds to the FXIa A3 domain followed by engagement at the protease active site and cleavage of the R191-Al192 bond (Geng Y et al. 2012, 2013; Gailani D et al. 2014). The cleavage after R191 facilitates cleavage of the R226-V227 bond, forming the activated FIXa (also known as factor IXaβ). Catalytic efficiency for the second cleavage by FXIa is 7-fold greater than for the first cleavage, explaining the low accumulation of the α-cleavage product of FIX (Wolberg AS et al. 1997; Smith SB et al. 2008; Geng Y et al. 2012; Mohammed BM et al. 2018). Activated FIXa comprises an N-terminal light chain and a C-terminal heavy chain held together by a disulphide bridge between cysteine resides 178 and 335 (Di Scipio RG et al. 1978; Zögg T & Brandstetter H 2009). X-ray structure of the FIXa EGF2/protease domain at 1.37 A revealed that a Na+-binding site in association with Ca2+-binding site contributed to stabilization of the FIXa protease domain (Vadivel K et al. 2019).
Activated kallikrein binds to alpha2-macroglobulin (Sottrup-Jensen et al. 1984), forming a stable and enzymatically inactive complex. Under normal conditions in vivo, this reaction appears to be responsible for the inactivation of about 1/6 of activated kallikrein (with C1Inh responsible for the inactivation of about 5/6) (Harpel et al. 1985).
Kininogen (high molecular weight kininogen; HK) associates with C1q binding protein on the cell surface in a reaction dependent on Zn++ (Joseph et al. 1996). In the body, the Zn++ needed to drive this reaction may be provided locally by Zn++ release from activated platelets (Mahdi et al. 2002). The C1q binding protein is inferred to form tetramers based on the properties of purified recombinant protein in vitro (Ghebrehiwet et al. 1994); the stoichiometry of the cell surface complex has not been determined directly.
Activated factor XII (factor XIIa) binds to C1Inh (C1 inhibitor - Bock et al. 1986) to form a stable, inactive complex (Schneider et al. 1973). While several protease inhibitors can form stable complexes with XIIa in vitro, only C1Inh does so to a significant extent under normal conditions in vivo (Pixley et al. 1985).
Activated kallikrein binds to C1Inh (plasma protease C1 inhibitor, SERPING1) (Bock et al. 1986), forming a stable and enzymatically inactive complex. This reaction appears to be the major means by which kallikrein is inactivated (kallikrein can also be inactivated by binding to alpha2-macroglobulin) (Harpel et al. 1985; Ratnoff et al. 1969).
Factor XI, bound to the cell surface, is converted to activated factor XI (factor XIa). In the body, this reaction occurs on the surfaces of activated platelets (Baglia et al. 2002). Small quantities of factor XI can be activated in a reaction catalyzed by factor XIIa, to initiate formation of a fibrin clot. However, the efficient activation of larger quantities of factor XI, needed to propagate the blood clotting process, appears to be mediated by thrombin (Baglia and Walsh 2000; Gailani and Broze 1993; Naito and Fujikawa 1991; Oliver et al. 1999; Monroe et al. 2002).
SERPIND1 (Heparin cofactor 2) is a serine endopeptidase inhibitor (SERPIN) that acts as a pseudosubstrate for activated thrombin, forming a stable complex which has the effect of inactivating thrombin protease activity (Church et al. 1985), although with slower kinetics than SERPINC1 (antithrombin-III). The presence of the glycosaminoglycans heparin or dermatan sulphate increases thrombin inactivation 1000-fold (Van Deerlin & Tollefsen 199) by facilitating the interaction between the active site of thrombin and the reactive site of SERPIND1. Thrombin specificity is conferred by a 90-residue N-terminal extension that contains two acidic motifs containing sulphated Tyr residues, resembling the C-terminus of hirudin (Tollefsen et al. 1997). SERPIND1 also inhibits chymotrypsin and neutrophil cathepsin G, but in a glycosaminoglycan independent manner (Church et al. 1985). In contrast to SERPINC1 deficiency, SERPIND1 deficiency is not associated with venous thrombosis (Corral et al. 2004).
Protein C is best known for its anticoagulant activity, the proteolytic inactivation of FVa and FVIIIa on negatively charged phospholipid membranes. This is enhanced by cofactors protein S and FV (Rosing et al. 1995, Norstrom et al. 2006). Inactivation of FVa involves APC-mediated cleavages at Arg306 and Arg506. The rapid cleavage at Arg506 is kinetically favored over cleavage at Arg306, but results only in partial inactivation of FVa, whereas the slower cleavage at Arg306 results in a complete loss of FVa function (Kalafatis et al. 1994, Nicolaes et al. 1995). Protein S accelerates factor Va inactivation by selectively promoting the slow cleavage at Arg306 (Rosing et al. 1995). A mutation of the APC cleavage sites in FV Arg506Gln a.k.a. FVLeiden is the most common identifiable hereditary risk factor for venous thrombosis among Caucasians (Camire 2011). APC also has a role in the inactivation of FVIIIa (Regan et al. 1994). Similar to FVa inactivation, FVIIIa is cleaved by APC at Arg336 in the A1 subunit and at Arg562 in the A2 subunit, with either resulting in a complete loss of cofactor activity (O'Brien et al. 2000, Manithody et al. 2003). Both protein S and FV but not FVa enhance inactivation of FVIIIa by APC (O'Brien et al. 2000,57). By acting on FVa and FVIIIa Protein C down-regulates both primary and secondary thrombin formation, delaying clot formation and diminishing activation of TAFI, enhanced susceptibility of the clot to fibrinolysis, respectively. The latter effects of APC on secondary thrombin formation is sometimes referred to as APC’s profibrinolytic effect (Bajzar et al. 1996).
Physiological activation of protein C on the endothelial cell surface requires the binding of protein C to the endothelial protein C receptor PROCR (EPCR) as well as binding of thrombin to thrombomodulin (TM) (Stavenuiter et al. 2013). PROCR binding to protein C (Fukudome & Esmon 1994) augments by at least 5-fold the effect of thrombin-thrombomodulin on the rate of protein C activation (Stearns-Kurosawa et al. 1996, Taylor et al. 2001).
SERPINA5, also called Plasma serine protease inhibitor or Protein C inhibitor, inactivates serine proteases by binding irreversibly to their serine activation site. It is involved in the regulation of intravascular and extravascular proteolytic activities, promoting coagulation by inhibiting the anticoagulant complex Activated protein C (APC), but also acts as an anticoagulant factor by inhibiting blood coagulation factors such as prothrombin, factor XI, factor Xa, plasma kallikrein and fibrinolytic enzymes such as tissue- and urinary-type plasminogen activators. Its inhibitory activity is greatly enhanced in the presence of glycosaminoglycans (GAGs), heparin, thrombomodulin and phospholipids vesicles (Suzuki et al. 1985).
SERPINA5 inhibits activated protein C In the blood plasma and inhibits thromibin as part of the thrombin:thrombomodulin complex (Rezaie et al. 1995). On the other hand, PCI can also inhibit coagulation factors (Radtke et al. 2007). The SERPINA5:APC complex is a marker of thrombotic events (Kolbel et al. 2006), which suggests that despite low circulating SERPINA5 concentrations and rates of APC inhibition, its predominant role is procoagulatory (Li & Huntington 2008). This is due to the enhancing effect of GAGs, which line the vascular endothelium. Both SERPINA5 and APC bind to GAGs. The presence of heparin in vitro accelerates the maximal rate of inhibition by over 2000-fold (when accounting for dissociation constants) (Yang et al. 2002).
SERPINE2 (Protease nexin-1, PN1) is a specific and extremely efficient inhibitor of thrombin. Unlike other thrombin inhibitors belonging to the serpin family, SERPINE2 does not circulate in the blood (Bouton et al. 2012). Rather, it is bound to glycosaminoglycans on the surface of cell types including macrophages, smooth muscle cells and platelets, where it inhibits the signaling functions of thrombin. SERPINE2 sets the threshold for thrombin-induced platelet activation (Gronke et al. 1987, Boulaftali et al. 2010) and has been implicated in atherosclerosis (Bouton et al. 2012). Recent studies have demonstrated an important antithrombotic effect of platelet SERPINE2 in vitro and in vivo (Boulaftali et al. 2010).
Activated protein C (APC) can either dissociate from PROCR to exert its anticoagulant activity, or remain bound to PROCR where it influences multiple direct cellular activities. Dissociation of APC from PROCR allows APC to associate with other cell membrane surface molecules, various microparticles, or lipoproteins (e.g., high-density lipoprotein). As an anticoagulant, APC cleaves the activated cofactors Va (fVa) and VIIIa (fVIIIa), yielding inactivated cofactors, fVi and fVIIIi. This proteolytic inactivation is enhanced by protein cofactors (e.g., protein S, factor V) and lipids cofactors (e.g., phosphatidylserine, cardiolipin, glucosylceramide, or HDL).
Activated protein C binds to Protein S on appropriate cell surfaces where it inactivates factors Va and VIIIa. Protein S is best known as a cofactor for the Activated protein C (APC)-catalyzed inactivation of factor Va (Walker 1980). Protein S must be membrane-bound to display this cofactor activity (Hackeng et al. 1993). Protein S binding brings the active site of APC closer to the phospholipid cell surface (Yegneswaran et al. 1999).
APC proteolysis involves cleavage of the factor Va heavy chain at Arg-306 and Arg-506 (Nicolaes et al. 1985). Most factor Va molecules are initially cleaved at Arg506, yielding a partially active intermediate, followed by complete inactivation through cleavage at Arg306 (Kalafatis et al. 1994). Protein S stimulates the cleavage at Arg306 ~20-fold (Rosing et al. 1995) and also counteracts the protective effect of factor Xa on Arg506 cleavage (Norstrom et al. 2006).
Protein S also enhances the APC-mediated inactivation of factor VIIIa (van de Poel et al. 2001). Protein S and factor V act as synergistic cofactors in the APC-mediated inactivation of factor VIIIa (Shen & Dahlback 1994, Somajo et al. 2014).
A soluble form of PROCR (sEPCR) fully retains the ability to bind Protein C and Activated protein C (Kurosawa et al. 1997). This form increases up to 5-fold in patients with sepsis or systemic lupus erythematosus (Kurosawa et al. 1998), either from vascular injury or through a regulated proteolytic release of soluble receptor (Gu et al. 2000). sEPCR inhibits protein C activation over large vessel endothelium in culture, reflecting competition between the soluble and cell surface forms of PROCR (Liaw et al. 2000).
Activated Protein C (APC) is best known for its anticoagulant activity, the proteolytic inactivation of FVa and FVIIIa on negatively charged phospholipid membranes. This is enhanced by cofactors protein S and factor V (Rosing et al. 1995, Norstrom et al. 2006).
APC inactivates FVIIIa (Regan et al. 1994) with a mechanism similar to its inactivation of FVa. FVIIIa is cleaved by APC at Arg355 (336 if numbering excludes signal peptide) in the A1 subunit and at Arg581 (562 if numbering excludes signal peptide) in the A2 subunit (O'Brien et al. 2000, Manithody et al. 2003). The Arg355 cleavage is 6-fold faster than the Arg581 cleavage but does not fully inactivate factor VIIIa if dissociation of the A2 subunit is blocked (Gale et al. 2008). Protein S and Factor V (but not FVa) enhance the inactivation of FVIIIa by APC (O'Brien et al. 2000). Protein S and factor V both enhance cleavage at both sites, more so at Arg581 (Gale et al. 2008).
The A2 subunit of FVIIIa spontaneously dissociates, inactivating FVIIIa with a half-life of about 2 min (Fay et al. 1991).
By acting on FVa and FVIIIa Protein C down-regulates both primary and secondary thrombin formation, delaying clot formation and diminishing activation of TAFI, enhanced susceptibility of the clot to fibrinolysis, respectively. The latter effects of APC on secondary thrombin formation is sometimes referred to as APC’s profibrinolytic effect (Bajzar et al. 1996).
Soluble PROCR binds to activated neutrophils via PRTN3, also cknown as myeloblastin and (Leukocyte) proteinase-3 (Kurosawa et al. 2000). PRTN3 is the most abundant serine protease in neutrophils (Campbell et al. 2000). After neutrophil activation, PRTN3 is secreted from azurophil granules, rebinding to the neutrophil surface through an association with CD177 (NB1) a 60-kDa glycosyl-phosphatidylinositol (GPI)-linked cell surface glycoprotein, which is expressed on a subpopulation of neutrophils in 97% of healthy individuals (Knuckleburg et al. 2012). PRTN3 is partially protected from inactivation when associated with CD177 (Campbell et al. 2000) which may increase its efficacy. CD177 is a heterophilic binding partner for endothelial cell platelet-endothelial cell adhesion molecule (PECAM)-1, which is expressed at endothelial cell junctions where transmigration occurs (Sun et al. 2000) suggesting that CD177 directs at least a subpopulation of PRTN3 molecules to these areas to aid neutrophil diapedesis, perhaps through PRTN3 degradation of cell junction proteins or the extracellular matrix.
Membrane-bound thrombin-activated factor VIII (fVIIIa) functions as a cofactor for factor IXa in the factor Xase complex. Factors VIIIa and IXa associate with anionic phospholipid surfaces with high affinity (Gilbert et al. 1990, Mertens & Bertina 1984, Mertens et al. 1984; Greengard et al. 1986). Studies using physiologic surfaces provide evidence for coordinated binding interactions of the enzyme, cofactor and substrate to discrete surface structures. For example, the presence of both (active site-modified) factor IXa and factor X increased both the number and the affinity of binding sites on activated platelets for factor VIIIa (Ahmad et al. 2000). However classical receptors for the constituents of factor Xase have not been identified (Fay 2004).
Cleavage of factor VIII light chain promotes a change in the conformation of the C2 domain that facilitates dissociation from VWF and enhances the affinity of factor VIIIa for anionic phospholipid surfaces (Saenko et al. 1998). Membrane-bound thrombin-activated factor VIII (FVIIIa) functions as a cofactor for factor IXa in the factor Xase complex. Factors VIIIa and IXa associate with anionic phospholipid surfaces with high affinity (Gilbert et al. 1990, Mertens & Bertina 1984; Panteleev et al 2004). Kd values ranging from 0.01 to 4.8 nM have been reported for FVIII binding to phospholipids (Gilbert et al. 1990,1992; Spaargaren et al. 1995; Raut et al. 1999; Ahmad et al. 2000). Studies using physiologic surfaces provide evidence for coordinated binding interactions of the enzyme, cofactor and substrate to discrete surface structures. For example, the presence of both (active site-modified) factor IXa and factor X increased both the number and the affinity of binding sites on activated platelets for factor VIIIa (Ahmad et al. 2000). However classical receptors for the constituents of factor Xase have not been identified (Fay 2004).
F2R (PAR1) mediates multiple cytoprotective effects of Activated proein C (APC) (Riewald et al. 2002, Griffin et al. 2007). In most, but not all, reported studies of APC’s beneficial effects on endothelial cells, the cellular receptors EPCR and F2R are required. These cytoprotective effects include anti-apoptotic activities, anti-inflammatory activities, protection of endothelial barrier functions, and favorable alteration of gene expression profiles. This paradigm in which EPCR-bound APC activates F2R to initiate signaling is consistent with many in vitro and in vivo data. Localization of APC signaling to caveolin-1-rich microdomains (caveolae) may help differentiate mechanisms for cytoprotective APC signaling versus proinflammatory thrombin signaling. Additional mechanisms for APC effects on cells may involve other receptors. These effects include APC anti-inflammatory effects on leukocytes or cytoprotective effects on dendritic cells and neurons. Other receptors may include F2RL2 (PAR3), various integrins e.g., Mac-1 (CD11b/CD18), Beta-1 integrins, Beta-3 integrins, S1P1, or the apolipoprotein E receptor 2 (LRP8) (Mosnier et al. 2007).
Factor Xa (aka Factor X heavy chain), a cleavage product of coagulation factor X (F10), is a vitamin K-dependent glycoprotein able to convert prothrombin to thrombin during the blood clotting process. Factor Xa is a target for direct oral anticoagulant (DOAC) drugs that are direct factor Xa inhibitors (the so-called 'xabans') which are used in the treatment and prevention of thromboembolic disorders (Galanis et al. 2014). Rivaroxaban (brand name Xarelto) was the first medically approved drug of this class (Abrams & Emerson 2009, Misselwitz et al. 2011). Rivaroxaban binds to and inhibits both free factor Xa and factor Xa bound in the prothrombinase complex (Roehrig et al. 2005). Other 'xabans' such as apixaban (Bhanwra & Ahluwalia 2014), edoxaban (Minor et al. 2015), eribaxaban (Bondarenko et al. 2013) and betrixaban (Zhang et al. 2009) share a similar mechanism of action to rivaroxaban (Nutescu et al. 2016).
Factor Xa (aka Factor X heavy chain), a cleavage product of coagulation factor X (F10), is a vitamin K-dependent glycoprotein able to convert prothrombin to thrombin during the blood clotting process. Factor Xa is a target for direct oral anticoagulant (DOAC) drugs that are direct factor Xa inhibitors (the so-called 'xabans') which are used in the treatment and prevention of thromboembolic disorders (Galanis et al. 2014, Nutescu et al. 2016). Rivaroxaban (brand name Xarelto) binds to and inhibits both free factor Xa and factor Xa bound in the prothrombinase complex (Va:Xa) (Roehrig et al. 2005). Rivaroxaban was the first medically approved drug of this class (Abrams & Emerson 2009, Misselwitz et al. 2011). Other 'xabans' such as apixaban (Bhanwra & Ahluwalia 2014), edoxaban (Minor et al. 2015), eribaxaban (Bondarenko et al. 2013) and betrixaban (Zhang et al. 2009) share a similar mechanism of action to rivaroxaban (Nutescu et al. 2016).
In the blood coagulation process, prothrombin is proteolytically cleaved to form thrombin (factor IIa) which in turn, acts as a serine protease that converts soluble fibrinogen into insoluble strands of fibrin. Specifically, thrombin converts factor XI to XIa, factor VIII to VIIIa, factor V to Va, fibrinogen to fibrin, and factor XIII to XIIIa. The direct oral anticoagulant (DOAC) synthetic organic drugs dabigatran (brand name Pradaxa), argatroban (brand name Acova, Novastan; Exembol in the UK) and melagatran are potent, competitive direct thrombin inhibitors (DTIs). They reversibly and specifically bind both clot-bound and free thrombin (unlike warfarin or heparin), as well as inhibiting thrombin-induced platelet aggregation (Wienen et al. 2007, Stangier et al. 2007).
Commercially, dabigatran is formulated as a lipophilic prodrug, dabigatran etexilate, to promote gastrointestinal absorption before it is metabolised to the active drug. The kidneys excrete the majority (80%) of unchanged drug (Stangier et al. 2007). Argatroban is a synthetic inhibitor of thrombin derived from L-arginine, which has a relatively short period of binding only to thrombin’s active site (Hursting et al. 1997). It is given intravenously and is metabolised in the liver. Because of its hepatic metabolism, it may be used in patients with renal dysfunction. Melagatran is the active drug formed from the prodrug ximelagatran and is a competitive and rapid inhibitor of thrombin (Gustafsson et al. 1998). DuP 714 is a potent and specific thrombin inhibitor (Chiu et al. 1991).
A major downside of DOACs is that they don't have reversing antidotes if internal bleeding arises from their use. However, in the case of severe bleeding of patients on dabigatran, the antibody fragment idarucizumab reversed the anticoagulation effects of dabigatran within minutes (Pollack et al. 2015). This represents a novel anticoagulation reversing mechanism for a DOAC.
In the blood coagulation process, prothrombin is proteolytically cleaved to form thrombin (factor IIa) which in turn, acts as a serine protease that converts soluble fibrinogen into insoluble strands of fibrin. Specifically, thrombin converts factor XI to XIa, factor VIII to VIIIa, factor V to Va, fibrinogen to fibrin, and factor XIII to XIIIa. Direct thrombin inhibitors (DTIs) represent a new class
of promising anticoagulation agents. DTIs are increasingly
being used instead of heparin to provide initial,
rapid anticoagulation. Unlike heparin, which requires a mediator (antithrombin) to potentiate anticoagulation, Peptide DTIs can inhibit free and bound thrombin directly. Lepirudin (brand name Refludan) is a recombinant hirudin derived from yeast cells (Weitz et al. 1990). Hirudin is a naturally occurring anticoagulant produced by the salivary glands of medicinal leeches. Bivalirudin (brand name Angiomax, Angiox) is a synthetic analog of hirudin, with a shorter period of binding to thrombin (Gladwell 2002). Desirudin (brand name Iprivask) is another recombinant hirudin derivative that directly inhibits free and fibrin-bound thrombin (Graetz et al. 2011).
The plasma protease C1-inhibitor (C1-INH, SERPING1)) like all extracellular serine proteinase inhibitors (serpins) is secreted via the endoplasmic reticulum (ER)-Golgi pathway (Pan S et al. 2011). SERPING1 (C1-INH) is produced mainly in hepatocytes, reaching in healthy individuals a plasma concentration of 0.21–0.39 g/l (Prandini MH et al. 1986; Wouters D et al. 2008). SERPING1 can be produced and secreted from other cell types like peripheral blood monocytes, fibroblasts, and endothelial cells (Katz Y & Strunk RC 1989; Schmaier AH et al. 1989; Prada AE et al. 1998). SERPING1 is highly glycosylated plasma protein, bearing both N- and O-glycans (Stavenhagen K et al. 2018). SERPING1 belongs to the serine protease inhibitor (serpin) superfamily of structurally similar but functionally diverse proteins that use a conformational change to inhibit target enzymes (Silverman GA et al. 2001; Gettins PG 2002; Law RH et al. 2006). Serpins are globular proteins with a conserved structure of 7- 9 α-helices and 3 β-pleated sheets and a protruding reactive center loop (RCL) (Silverman GA et al. 2001; Gettins PG 2002; Law RH et al. 2006; Sanrattana W et al. 2019). In native serpins, the RCL, located outside the tertiary core of the serpin, forms a flexible stretch of approximately 20 amino acids, which provides structural flexibility in a solvent-exposed environment. They act on their target proteases by means of a suicide-substrate mechanism involving the cleavage of the RCL and its insertion into β-sheet A (Gettins PG 2002; Pan S et al. 2011; Khan MS et al. 2011). As a result, conformational changes take place in the serpins that ultimately trap and inactivate the targeted protease (Gettins PG 2002; Pan S et al. 2011; Khan MS et al. 2011; Sanrattana W et al. 2019). Serpins are conformationally labile and many of the disease-linked mutations of serpins result in misfolding or in formation of inactive, pathogenic polymers (Law RH et al. 2006). Under normal physiological conditions, SERPING1 (C1-INH) inhibits the activated forms of the serine proteases involved in the complement pathway (C1r and C1s), the contact system (FXIIa, FXIa, and kallikrein) as well as fibrinolytic proteases such as plasmin, tPA, and uPA (Sim et al. 1979; Arlaud et al. 1979; Kaplan AP & Ghebrehiwet B 2010).
Hereditary angioedema (HAE) is a rare life-threatening inherited edema disorder that is characterized by recurrent episodes of acute swelling involving the skin or the oropharyngeal, laryngeal, or gastrointestinal mucosa (Zuraw BL & Christiansen SC 2016; de Maat S et al. 2018; Magerl M et al. 2017). Increased vascular permeability in HAE is due to excessive formation of the proinflammatory peptide hormone bradykinin (BK) (Joseph K et al. 2008). Elevated plasma levels of BK are consistently found during acute swelling attacks in HAE patients (Schapira M et al. 1983; Cugno M et al. 2003). Angioedema initiated by bradykinin is usually associated with SERPING1 (С1-Inh) deficiency (HAE type I and HAE type II) (Kaplan AP & Joseph K 2014, 2016; Levi M et al. 2018). More rarely, HAE occurs in individuals with normal SERPING1 activity, and has been linked to mutations in other proteins, including FXII, plasminogen, and angiopoietin (Magerl M et al. 2017; Zuraw BL 2018; Ivanov I et al. 2019). Using genome-wide linkage analyses, HAE type III was shown to be associated with single missense mutations (c.1032C>A or c.1032C>G) in the F12 gene (Dewald G & Bork K 2006; Cichon S et al. 2006). Both point mutations translate into amino acid substitution of threonine 328 by either a lysine or an arginine residue (T328K or T328R). FXII-linked HAE is an autosomal dominant inherited disorder, and a mixture of wild type and T328-mutated FXII circulates in plasma of patients with HAE type III (Cichon S et al. 2006). An FXII-neutralizing antibody attenuated pathological BK formation in the plasma of patients with HAE type III and blunted edema in a genetically altered, humanized mouse model of HAE type III (Björkqvist J et al. 2015). Moreover, FXII T328K or T328R variants change protein O-linked glycosylation and introduce a new site that is sensitive to enzymatic cleavage by fibrinolytic and coagulation proteases such as plasmin, thrombin or FXIa (Björkqvist J et al. 2015; de Maat S et al. 2016; Ivanov I et al. 2019). FXII T328K or T328R variants are cleaved after residue 328 by proteases, removing the protein’s noncatalytic heavy chain (HC) region. Further, truncation of the pathological FXII T328K by plasmin was found to expose R372 for subsequent cleavage by plasma kallikrein in solution (de Maat S et al. 2016, 2019). The intrinsic capacity of the truncated form of FXII (329-615) variant to convert PK to kallikrein is greater than that of activated FXIIa leading to more kallikrein generated early during reciprocal activation (Ivanov I et al. 2019). Second, the truncated form of FXII (329-615) is a better kallikrein substrate than is FXII (Ivanov I et al. 2019). SERPING1 (C1-Inh), the major inhibitor of FXIIa, binds similarly to wild type (WT) and mutated FXIIa (Björkqvist J et al. 2015). However, the accelerated kallikrein/FXII activation in HAE patients carrying FXII variants appears to overwhelm the regulatory function of SERPING1 at normal plasma levels leading to uncontrolled bradykinin formation in a surface-independent manner (de Maat S et al. 2016; Ivanov I et al. 2019).
The Reactome event describes a truncation of FXII variants after the residue 328 catalyzed by activated thrombin.
In healthy individuals, cleavage of a single peptide bond converts factor XII (FXII, F12, Hageman factor) to activated FXII (FXIIa) (Fujikawa and McMullen 1983; McMullen and Fujikawa 1985). FXII undergoes autoactivation to FXIIa either by endogenous activator (nucleic acids RNA/DNA, neutrophil extracellular traps (NETs), polyphosphate and heparin) or artificial surfaces. FXIIa then activates four different pathways: 1) The inflammation kallikrein-kinin pathway by converting plasma pre-kallikrein (PK) into active plasma kallikrein, which cleaves both FXII into FXIIa and high molecular weight kininogen (HK) to bradykinin (BK). The latter binds to kinin receptors (B2 and B1 receptors) and triggers inflammation. 2) The complement system by activation of the C1qrs complex subunits C1r and C1s leading formation of the membrane attack complex by the classical complement pathway. 3) The fibrinolytic system by activation of pro-urokinase into urokinase that in turn cleaves plasminogen into plasmin, an enzyme that degrades fibrin clots. 4) The intrinsic coagulation pathway by FXI activation into FXIa leading to thrombin activation and fibrin generation. The contact system is controlled mainly by C1 inhibitor (C1-Inh or SERPING1) that inhibits both FXIIa and kallikrein.
Patients with hereditary angioedema (HAE) experience episodes of soft tissue swelling as a consequence of unregulated kallikrein activity or increased prekallikrein activation. Angioedema initiated by bradykinin is usually associated with SERPING1 deficiency. More rarely, HAE occurs in individuals with normal SERPING1 activity, and has been linked to mutations in other proteins, including FXII, plasminogen, and angiopoietin (Magerl M et al. 2017; Zuraw BL 2018; Ivanov I et al. 2019). The substitution of threonine 328 by either a lysine or an arginine residue (T328K or T328R) in the FXII proline-rich region have been identified in several families with HAE and normal SEPING1. FXII T328K or T328R variants change protein glycosylation and introduce a new site that is sensitive to enzymatic cleavage by fibrinolytic and coagulation proteases such as plasmin, thrombin or FXIa (de Maat S et al. 2016; Ivanov I et al. 2019). FXII T328K or T328R variants are cleaved after residue 328 by proteases, removing the protein’s noncatalytic heavy chain (HC) region. Truncation of the pathological FXII T309K by plasmin exposes R372 for subsequent cleavage by plasma kallikrein in solution (de Maat S et al. 2016, 2019). The intrinsic capacity of the truncated form of FXII (329-615) to convert PK to kallikrein is greater than that of activated FXII leading to more kallikrein generated early during reciprocal activation (Ivanov I et al. 2019). Second, the truncated form of FXII (329-615) is a better kallikrein substrate than is FXII (Ivanov I et al. 2019). Further, the accelerated PK/FXII activation in HAE patients carrying FXII variants appears to overwhelm the regulatory function of SERPING1 at normal plasma levels leading to uncontrolled bradykinin formation in a surface-independent manner (de Maat S et al. 2016; Ivanov I et al. 2019).
Coagulation factor VIII (FVIII) is a large glycoprotein of 2351 aminoacids with a discrete domain structure: A1-A2-B-A3-C1-C2 (Wood WI et al. 1984; Vehar GA et al. 1984; Toole JJ et al. 1984). FVIII is synthesized by various tissues, including liver, kidney, and spleen, as an inactive single-chain protein of approximately 293 kDa (Wion KL et al. 1985; Levinson B et al. 1992). Primary human liver sinusoidal endothelial cells (LSECs), blood outgrowth endothelial cells (BOEC), glomerular microvascular endothelial cells (GMVECs) and umbilical vein endothelial cells (HUVECs) were found to produce the FVIII protein, store it in Weibel-Palade bodies (WPB), and secrete in response to EC stimulation (van den Biggelaar M et al. 2009; Shahani T et al. 2014; Turner NA & Moake JL 2015). These findings are in agreement with the reports on the FVIII synthesis in human cultured ECs and in mice suggesting that ECs are the predominant source of plasma FVIII (Jacquemin M et al. 2006; Shahani T et al. 2010; Fahs SA et al. 2014). Evidence on the post-translational processing and secretion of FVIII has been generated from expression of the FVIII complementary DNA (cDNA) in transfected mammalian cells, such as Chinese hamster ovary (CHO), African green monkey kidney (COS-1), HeLa and the human hepatic SK-HEP1cell lines (Pipe SW et al. 1998; Herlitschka SE et al. 1998). Upon synthesis, FVIII is translocated into the lumen of the endoplasmic reticulum (ER), where it undergoes extensive processing including cleavage of a signal peptide and N-linked glycosylation at asparagine residues (Kaufman RJ et al. 1988, 1997; Kaufman RJ 1998). In the ER lumen of mammalian cells FVIII interacts with the protein chaperones calnexin (CNX), calreticulin (CRT), and immunoglobulin-binding protein (BiP or GRP78) that facilitate proper folding of proteins prior to trafficking to the Golgi compartment (Marquette KA et al. 1995; Swaroop M et al. 1997; Pipe SW et al. 1998; Kaufman RJ et al. 1997; Kaufman RJ 1998). Trafficking from the ER to the Golgi compartment is facilitated by LMAN1 and multiple combined factor deficiency 2 (MCFD2) cargo receptor complex (Zhang B et al. 2005; Zheng, C et al. 2010, 2013). Within the Golgi apparatus, FVIII is subject to further processing, including modification of the N-linked oligosaccharides to complex-type structures, O-linked glycosylation, and sulfation of specific Tyr-residues (Michnick DA et al. 1994; Kaufman RJ 1998). In addition, factor VIII is among the many proteins that undergoes intracellular proteolysis. Upon secretion from the cell, FVIII is cleaved at two sites in the B-domain to form a heterodimer consisting of the heavy chain containing the A1-A2-B domains in a metal ion-dependent complex with the light chain consisting of the A3-C1-C2 domains (Kaufman RJ et al. 1997; Kaufman RJ 1998). In the plasma, FVIII is stabilized through interaction with von Willebrand factor (Weiss HJ et al. 1977; Kaufman RJ et al. 1988; Chiu PL et al. 2015). Upon damage to blood vessel walls, thrombin cleaves FVIII and releases the B-domain to form an active FVIII heterotrimer (A1:A2:A3-C1-C2) that binds activated coagulation factor IX on the surface of platelet phospholipid to form the active factor Xase complex (Ahmad SS et al. 2003; Panteleev MA et al. 2006). This complex efficiently cleaves factor X to its active form, which activates prothrombin and leads to the formation of a stable fibrin clot. After conversion into its active conformation, and participation in the factor X activating complex, activated factor VIII rapidly looses its activity (Kaufman RJ et al. 1988; Lenting PJ et al. 1998). This process is governed by both enzymatic degradation and subunit dissociation. At the cellular level the FVIII expression is limited. Inefficient secretion of FVIII is caused by repression at the level of transcription (Lynch CM et al. 1993; Hoeben RC et al. 1995). In addition, a significant portion of the primary translation product is misfolded and ultimately degraded and FVIII is retained within ER through interaction with various ER chaperones including BiP (Marquette KA et al. 1995; Tagliavacca L et al. 2000). Mutations in the F8 gene often result in diminished or inactive plasma factor VIII protein and are the molecular genetic cause of the monogenic, X-linked, bleeding disorder hemophilia A (Al-Allaf FA et al. 2017; Castaman G & Matino D 2019).
In healthy individuals factor IXa (FIXa), in a complex with factor VIIIa on the surfaces of activated platelets, catalyzes the formation of activated factor X with high efficiency. A substitution of leucine for arginine at residue 384 in FIX (FIX R384L, also know as FIX Padua) is a gain-of-function mutation that resulted in elevated FIX clotting activity in a patient with venous thrombosis (Simioni P et al. 2009). The level of the FIX R384L protein in the patient plasma was normal, but the clotting activity from the proband was approximately eight times the normal level. In vitro, recombinant FIX R384L had a specific activity that was 5 to 10 times as high as that in the recombinant wild-type FIX (Simioni P et al. 2009). In addition, FIXa R384L showed a resistance to inhibition by protein S (PROS1), a plasma protein that directly binds and inhibits FIXa to modulate a clotting rate in vitro and in vivo (Plautz WE et al. 2018a,b). The ability of the FIX Padua variant to increase the clotting activity prompted researchers to try to produce chimeric FIX Padua concentrates for potential use in the treatment of patients with hemophilia B (Lozier JN 2012; Monahan PE et al. 2015; Spronck EA et al. 2019). Epidemiological studies in groups of patients with venous thrombosis failed to discover other cases with this FIX abnormality, indicating that the defect is rare (Koenderman JS et al. 2011; de Moraes Mazetto B et al. 2010). The Reactome event describes elevation of FIX activity due to gain-of-function mutation FIX R384L.
Factor VIII (FVIII) circulates in plasma as a heterodimer (domain structure A1-A2-B:A3-C1-C2) that requires thrombin cleavage to elicit procoagulant activity (Kaufman RJ et al. 1997). Upon activation by thrombin FVIII is converted to the labile FVIIIa, a heterotrimer of A1, A2 and A3C1C2 subunits, which serves as a cofactor for FIXa (Fay PJ 2006). At physiological concentrations, FVIIIa decays as a result of A2 subunit dissociation, which is weakly associated with the A1:A3-C1-C2 dimer by primarily electrostatic interactions (Fay PJet al. 1991; Fay PJ & Smudzin TM 1992; Parker ET et al 2006). Site-directed mutagenesis, functional and structural studies suggest that multiple residues at the A1-A2 and A2-A3 domain interfaces contribute to non-covalent interactions in stabilizing the protein (Parker ET & Lollar P 2007; Wakabayashi H & Fay PJ 2008, 2013; Wakabayashi H et al. 2008; Monaghan M et al. 2016). Retention of A2 polypeptide is required for normal stability of FVIIIa and dissociation of A2 correlates with FVIIIa inactivation and consequent loss of FXase activity.
Factor VIII (FVIII) circulates in plasma as a heterodimer (domain structure A1-A2-B:A3-C1-C2) that requires thrombin cleavage to elicit procoagulant activity (Kaufman RJ et al. 1997). Upon activation by thrombin FVIII is converted to a heterotrimic FVIIIa, which consists of A1, A2 and A3-C1-C2 subunits to serve as a cofactor for FIXa (Fay PJ 2006). At physiological concentrations, FVIIIa decays as a result of A2 subunit dissociation, which is weakly associated with the A1:A3-C1-C2 dimer by primarily electrostatic interactions (Fay PJet al. 1991; Fay PJ & Smudzin TM 1992; Parker ET et al 2006). Retention of A2 polypeptide is required for normal stability of FVIIIa and dissociation of A2 correlates with FVIIIa inactivation and consequent loss of FXase activity. Hemophilia A (HA)-associated mutations (R550H, A303E, S308L, N713I, R717W and R717L) within the predicted A1-A2 and A2-A3 interface are thought to disrupt potential intersubunit hydrogen bonds and have the molecular phenotype of increased rate of inactivation of FVIIIa due to increased rate of A2 subunit dissociation (Pipe SW et al. 1999, 2001; Hakeos WH et al. 2002). Patients with these mutations exhibit a clinical phenotype where the FVIII activity by one-stage clotting assay is at least two-fold higher than by two-stage chromogenic FXa generation assay (Pipe SW et al. 2001; Hakeos WH et al. 2002; Al-Samkari H & Croteau SE 2018). This effect directly relates to enhanced rates of loss of A2 subunit from FVIIIa, which has a more pronounced impact on activity values determined by the two-stage assay (Hakeos WH et al. 2002).
Factor VIII (FVIII) circulates in plasma as a heterodimer (domain structure A1-A2-B:A3-C1-C2) that requires thrombin cleavage to elicit procoagulant activity (Kaufman RJ et al. 1997). Upon activation by thrombin FVIII is converted to the labile FVIIIa, a heterotrimer of A1, A2 and A3C1C2 subunits, which serves as a cofactor for FIXa (Fay PJ 2006). At physiological concentrations, FVIIIa decays as a result of A2 subunit dissociation, which is weakly associated with the A1:A3-C1-C2 dimer by primarily electrostatic interactions (Fay PJ et al. 1991; Fay PJ & Smudzin TM 1992; Parker ET et al 2006). Retention of A2 polypeptide is required for normal stability of FVIIIa and dissociation of A2 correlates with FVIIIa inactivation and consequent loss of FXase activity. Hemophilia A (HA)-associated mutations (R550H, A303E, S308L, N713I, R717W and R717L) within the predicted A1-A2 and A2-A3 interface are thought to disrupt potential intersubunit hydrogen bonds and have the molecular phenotype of increased rate of inactivation of FVIIIa due to increased rate of A2 subunit dissociation (Pipe SW et al. 1999, 2001; Hakeos WH et al. 2002). Patients with these mutations exhibit a clinical phenotype where the FVIII activity by one-stage clotting assay is at least two-fold higher than by two-stage chromogenic FXa generation assay (Pipe SW et al. 2001; Hakeos WH et al. 2002; Al-Samkari H & Croteau SE 2018). This effect directly relates to enhanced rates of loss of A2 subunit from FVIIIa, which has a more pronounced impact on activity values determined by the two-stage assay (Hakeos WH et al. 2002).
Coagulation Factor IX (FIX) is expressed by hepatocytes (Yoshitake et al. 1985; Kurachi K & Kurachi S 1995). The newly synthesised FIX protein molecule comprising a pre- and pro-sequence (28 and 18 amino acids, respectively) and a mature peptide of 415 amino acids (total length, 461 amino acids) (Yoshitake et al. 1985; Kurachi K & Kurachi S 1995; Andersson LO et al. 1975; Anson DS et al. 1984). The pre-sequence (or signal sequence) directs FIX for secretion and the pro-sequence provides a binding domain for a vitamin K dependent (VKD) gamma (γ)-glutamyl carboxylase (GGCX) (Fryklund L et al. 1976; Galeffi P & Brownlee GG 1987; Lingenfelter SE & Berkner K 1996; Stanley TB et al. 1998). GGCX, an integral membrane protein located in the endoplasmic reticulum (ER) of hepatocytes, carboxylates certain glutamic acid residues in the adjacent GLA domain of FIX (Presnell and Stafford, 2002; Fryklund L et al. 1976; Galeffi P & Brownlee GG 1987). During the γ-carboxylation process, vitamin K hydroquinone is oxidized to vitamin K 2,3 epoxide and a carboxyl group is added to a glutamic acid residue (Wallin R eet al. 2002). In its native form, FIX contains 12 glutamic acid residues in the Gla domain; the first 10 residues are conserved in all VKD proteins, whereas the last two are unique to FIX (Gillis et al. 2008). FIX undergoes several other post-translational modifications before its secretion, including N- and O-linked glycosylation, sulfation, phosphorylation and hydroxylation (Agarwala KL et al. 1994; Bharadwaj D et al. 1995; Kaufman RJ 1998; Bond M et al. 1998; Enjolras N et al. 2004). These post-translational modifications occur within the ER and Golgi apparatus. In the ER, maturation and processing of secreted proteins are orchestrated by a group of molecules which facilitate protein folding and ensure that only correctly folded, assembled and modified proteins are transported along the secretory pathway. The proteins involved in the folding system are lectins such as calreticulin (CRT) or calnexin (CNX). A cellular unfolded protein response induces the ER-resident molecular chaperones such as glucose-regulated protein GRP78/BiP to prevent the aggregation of proteins in the ER. FIX was shown to co-immunoprecipitate with GRP78/BiP and CRT In cell lysates of transiently transfected human hepatocellular carcinoma (HepG2) cells expressing FIX (Enjolras N et al. 2004). After transportation of the carboxylated pro-FIX into the Golgi apparatus, the propeptide (29-46) is removed by the paired basic amino acid cleaving enzyme (PACE) (Wasley LC et al. 1993). The removal of the propeptide by PACE influences the formation of Ca2+-induced secondary and tertiary structures of the Gla domain, thus it is required for normal function of FIX (Pipe, 2008). The mature FIX is secreted and circulates in the plasma as an inactive 57kDa zymogen form (47-461). Domains within the zymogen are identified according to structure or function as follows: the GLA domain is crucial for the interaction with phospholipid surfaces; two epidermal growth factor (EGF)-like domains are critical for the interactions between factor IX and factor VIIIa; the activation peptide is released after proteolytic activation and the catalytic serine protease domain is required for normal function of FIX (Pipe 2008; Yoshitake S et al. 1985; Di Scipio RG et al. 1977; Rees DJ et al. 1988; Freedman SJ et al. 1995). Activation of factor IX involves cleavage of two peptide bonds, one on the C-terminal side of arginine 191 (the α-cleavage) the other on the C-terminal side of arginine 226 (the β-cleavage) (Di Scipio RG et al. 1978; Zögg T & Brandstetter H 2009). Activated factor IX comprising an N-terminal light chain and a C-terminal heavy chain held together by a disulphide bridge between cysteine resides 178 and 335 (Di Scipio RG et al. 1978; Zögg T & Brandstetter H 2009).
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thrombin inactivating
complexesinhibitors
(compounds):IIainhibitors
(compounds)inhibitors
(peptide):IIainhibitors
(peptide)kininogen:C1q binding protein
tetramer(factor
IIa):SERPIND1XIa:GPIb:GPIX:GPV
complexWillebrand factor
multimerSERPINC1:SERPINC1
activatorsAnnotated Interactions
thrombin inactivating
complexesthrombin inactivating
complexesthrombin inactivating
complexesthrombin inactivating
complexesthrombin inactivating
complexesinhibitors
(compounds):IIainhibitors
(compounds)inhibitors
(peptide):IIainhibitors
(peptide)Factor Xa (aka Factor X heavy chain), a cleavage product of coagulation factor X (F10), is a vitamin K-dependent glycoprotein able to convert prothrombin to thrombin during the blood clotting process. Factor Xa is a target for direct oral anticoagulant (DOAC) drugs that are direct factor Xa inhibitors (the so-called 'xabans') and used in the treatment and prevention of thromboembolic disorders (Galanis et al. 2014).
Irrespective of the cellular source of TF and whether it is induced or constitutively expressed, most of the TF expressed on the surfaces of resting cells exists in a cryptic coagulant-inactive state (Schecter AD et al. 1997; Bach RR 2006; Kothari H et al. 2013; Grover SP & Mackman N 2018). The encrypted TF can bind to FVIIa, but the assembled TF:FVIIa complex fails to activate FIX and FX (Rao LV & Pendurthi UR 2012). Activation or disruption of cells markedly enhances cell surface TF procoagulant activity without altering TF antigen levels at the cell surface (‘decryption’). Several mechanisms have been proposed for TF decryption on cell surfaces, and out of them, externalization of phosphatidylserine (PS) to the outer leaflet and PDI-mediated thiol-disulfide exchange pathways that affect the allosteric disulfide bond in TF seem most likely (Rao LV & Pendurthi UR 2012; Grover SP & Mackman N 2018; Ansari SA et al. 2019). The presence of a high molar concentration of sphingomyelin (SM) in the outer leaflet of the plasma membrane inhibits TF procoagulant activity on the cell surface, thus maintaining TF in an encrypted state in resting cells (Wang J et al. 2017). Acid-sphingomyelinase (ASM)-mediated hydrolysis of SM following cell injury removes the inhibitory effect of SM on TF activity, thus leading to TF decryption (Wang J, et al. 2017; Ansari SA et al. 2019). It has been suggested that the coordinated effects of SM hydrolysis, PS externalization and thiol-disulphide exchange pathways are responsible for full cellular activation of TF (Ansari SA et al. 2019). However, molecular links among various pathways of TF decryption are not fully known yet. The Reactome event describes exposure of TF sequestered in the wall of a blood vessel to flowing blood.
The Covid-19 pandemic is an infection caused by the SARS-CoV-2 coronavirus. Severe cases of this infection can lead to acute respiratory distress syndrome and coagulation changes leading to a higher risk of thrombosis, especially pulmonary embolism (Susen et al. 2020). LMWHs may play a role as potential attachment factors for SARS-CoV-2 (Tandon et al. 2020), potentially reducing the incidence and/or severity of thrombosis (Marietta et al. 2020).
Factor VIII is a heterodimer containing a heavy and a light polypeptide chain, generated by the proteolytic cleavage of a single large precursor polypeptide (Vehar et al. 1984). Several forms of the heavy chain are found in vivo, all functionally the same but differing in the amount of the B domain removed by proteolysis. The single form annotated here is the shortest one (Eaton et al. 1986; Hill-Eubanks et al. 1989).
It has been demonstrated in in vitro experiments that vWF facilitates the association of FVIII chains and the retention of procoagulant activity in the conditioned medium of cells producing FVIII (Kaufman RJ et al. 1988; Wise RJ et al. 1991). Similar data have been obtained for re-association of FVIII chains in solution (Fay PJ 1988). In vitro, von Willebrand factor (Titani et al. 1986) can form complexes with factor VIII with a 1:1 stoichiometry. The complexes that form in vivo, however, involve large multimers of von Willebrand factor and varied, but always low, proportions of factor VIII (Vlot et al. 1995). A stoichiometry of one molecule of factor VIII associated with 50 of von Willebrand factor is typical in vivo, and is used here to annotate the factor VIII:von Willebrand factor complex.
SERPINA5 inhibits activated protein C In the blood plasma and inhibits thromibin as part of the thrombin:thrombomodulin complex (Rezaie et al. 1995). On the other hand, PCI can also inhibit coagulation factors (Radtke et al. 2007). The SERPINA5:APC complex is a marker of thrombotic events (Kolbel et al. 2006), which suggests that despite low circulating SERPINA5 concentrations and rates of APC inhibition, its predominant role is procoagulatory (Li & Huntington 2008). This is due to the enhancing effect of GAGs, which line the vascular endothelium. Both SERPINA5 and APC bind to GAGs. The presence of heparin in vitro accelerates the maximal rate of inhibition by over 2000-fold (when accounting for dissociation constants) (Yang et al. 2002).
Protein S is best known as a cofactor for the Activated protein C (APC)-catalyzed inactivation of factor Va (Walker 1980). Protein S must be membrane-bound to display this cofactor activity (Hackeng et al. 1993). Protein S binding brings the active site of APC closer to the phospholipid cell surface (Yegneswaran et al. 1999).
APC proteolysis involves cleavage of the factor Va heavy chain at Arg-306 and Arg-506 (Nicolaes et al. 1985). Most factor Va molecules are initially cleaved at Arg506, yielding a partially active intermediate, followed by complete inactivation through cleavage at Arg306 (Kalafatis et al. 1994). Protein S stimulates the cleavage at Arg306 ~20-fold (Rosing et al. 1995) and also counteracts the protective effect of factor Xa on Arg506 cleavage (Norstrom et al. 2006).
Protein S also enhances the APC-mediated inactivation of factor VIIIa (van de Poel et al. 2001). Protein S and factor V act as synergistic cofactors in the APC-mediated inactivation of factor VIIIa (Shen & Dahlback 1994, Somajo et al. 2014).
APC inactivates FVIIIa (Regan et al. 1994) with a mechanism similar to its inactivation of FVa. FVIIIa is cleaved by APC at Arg355 (336 if numbering excludes signal peptide) in the A1 subunit and at Arg581 (562 if numbering excludes signal peptide) in the A2 subunit (O'Brien et al. 2000, Manithody et al. 2003). The Arg355 cleavage is 6-fold faster than the Arg581 cleavage but does not fully inactivate factor VIIIa if dissociation of the A2 subunit is blocked (Gale et al. 2008). Protein S and Factor V (but not FVa) enhance the inactivation of FVIIIa by APC (O'Brien et al. 2000). Protein S and factor V both enhance cleavage at both sites, more so at Arg581 (Gale et al. 2008).
The A2 subunit of FVIIIa spontaneously dissociates, inactivating FVIIIa with a half-life of about 2 min (Fay et al. 1991).
By acting on FVa and FVIIIa Protein C down-regulates both primary and secondary thrombin formation, delaying clot formation and diminishing activation of TAFI, enhanced susceptibility of the clot to fibrinolysis, respectively. The latter effects of APC on secondary thrombin formation is sometimes referred to as APC’s profibrinolytic effect (Bajzar et al. 1996).
Commercially, dabigatran is formulated as a lipophilic prodrug, dabigatran etexilate, to promote gastrointestinal absorption before it is metabolised to the active drug. The kidneys excrete the majority (80%) of unchanged drug (Stangier et al. 2007). Argatroban is a synthetic inhibitor of thrombin derived from L-arginine, which has a relatively short period of binding only to thrombin’s active site (Hursting et al. 1997). It is given intravenously and is metabolised in the liver. Because of its hepatic metabolism, it may be used in patients with renal dysfunction. Melagatran is the active drug formed from the prodrug ximelagatran and is a competitive and rapid inhibitor of thrombin (Gustafsson et al. 1998). DuP 714 is a potent and specific thrombin inhibitor (Chiu et al. 1991).
A major downside of DOACs is that they don't have reversing antidotes if internal bleeding arises from their use. However, in the case of severe bleeding of patients on dabigatran, the antibody fragment idarucizumab reversed the anticoagulation effects of dabigatran within minutes (Pollack et al. 2015). This represents a novel anticoagulation reversing mechanism for a DOAC.
of promising anticoagulation agents. DTIs are increasingly being used instead of heparin to provide initial,
rapid anticoagulation. Unlike heparin, which requires a mediator (antithrombin) to potentiate anticoagulation, Peptide DTIs can inhibit free and bound thrombin directly. Lepirudin (brand name Refludan) is a recombinant hirudin derived from yeast cells (Weitz et al. 1990). Hirudin is a naturally occurring anticoagulant produced by the salivary glands of medicinal leeches. Bivalirudin (brand name Angiomax, Angiox) is a synthetic analog of hirudin, with a shorter period of binding to thrombin (Gladwell 2002). Desirudin (brand name Iprivask) is another recombinant hirudin derivative that directly inhibits free and fibrin-bound thrombin (Graetz et al. 2011).Hereditary angioedema (HAE) is a rare life-threatening inherited edema disorder that is characterized by recurrent episodes of acute swelling involving the skin or the oropharyngeal, laryngeal, or gastrointestinal mucosa (Zuraw BL & Christiansen SC 2016; de Maat S et al. 2018; Magerl M et al. 2017). Increased vascular permeability in HAE is due to excessive formation of the proinflammatory peptide hormone bradykinin (BK) (Joseph K et al. 2008). Elevated plasma levels of BK are consistently found during acute swelling attacks in HAE patients (Schapira M et al. 1983; Cugno M et al. 2003). Angioedema initiated by bradykinin is usually associated with SERPING1 (С1-Inh) deficiency (HAE type I and HAE type II) (Kaplan AP & Joseph K 2014, 2016; Levi M et al. 2018). More rarely, HAE occurs in individuals with normal SERPING1 activity, and has been linked to mutations in other proteins, including FXII, plasminogen, and angiopoietin (Magerl M et al. 2017; Zuraw BL 2018; Ivanov I et al. 2019). Using genome-wide linkage analyses, HAE type III was shown to be associated with single missense mutations (c.1032C>A or c.1032C>G) in the F12 gene (Dewald G & Bork K 2006; Cichon S et al. 2006). Both point mutations translate into amino acid substitution of threonine 328 by either a lysine or an arginine residue (T328K or T328R). FXII-linked HAE is an autosomal dominant inherited disorder, and a mixture of wild type and T328-mutated FXII circulates in plasma of patients with HAE type III (Cichon S et al. 2006). An FXII-neutralizing antibody attenuated pathological BK formation in the plasma of patients with HAE type III and blunted edema in a genetically altered, humanized mouse model of HAE type III (Björkqvist J et al. 2015). Moreover, FXII T328K or T328R variants change protein O-linked glycosylation and introduce a new site that is sensitive to enzymatic cleavage by fibrinolytic and coagulation proteases such as plasmin, thrombin or FXIa (Björkqvist J et al. 2015; de Maat S et al. 2016; Ivanov I et al. 2019). FXII T328K or T328R variants are cleaved after residue 328 by proteases, removing the protein’s noncatalytic heavy chain (HC) region. Further, truncation of the pathological FXII T328K by plasmin was found to expose R372 for subsequent cleavage by plasma kallikrein in solution (de Maat S et al. 2016, 2019). The intrinsic capacity of the truncated form of FXII (329-615) variant to convert PK to kallikrein is greater than that of activated FXIIa leading to more kallikrein generated early during reciprocal activation (Ivanov I et al. 2019). Second, the truncated form of FXII (329-615) is a better kallikrein substrate than is FXII (Ivanov I et al. 2019). SERPING1 (C1-Inh), the major inhibitor of FXIIa, binds similarly to wild type (WT) and mutated FXIIa (Björkqvist J et al. 2015). However, the accelerated kallikrein/FXII activation in HAE patients carrying FXII variants appears to overwhelm the regulatory function of SERPING1 at normal plasma levels leading to uncontrolled bradykinin formation in a surface-independent manner (de Maat S et al. 2016; Ivanov I et al. 2019).
The Reactome event describes a truncation of FXII variants after the residue 328 catalyzed by activated thrombin.
Patients with hereditary angioedema (HAE) experience episodes of soft tissue swelling as a consequence of unregulated kallikrein activity or increased prekallikrein activation. Angioedema initiated by bradykinin is usually associated with SERPING1 deficiency. More rarely, HAE occurs in individuals with normal SERPING1 activity, and has been linked to mutations in other proteins, including FXII, plasminogen, and angiopoietin (Magerl M et al. 2017; Zuraw BL 2018; Ivanov I et al. 2019). The substitution of threonine 328 by either a lysine or an arginine residue (T328K or T328R) in the FXII proline-rich region have been identified in several families with HAE and normal SEPING1. FXII T328K or T328R variants change protein glycosylation and introduce a new site that is sensitive to enzymatic cleavage by fibrinolytic and coagulation proteases such as plasmin, thrombin or FXIa (de Maat S et al. 2016; Ivanov I et al. 2019). FXII T328K or T328R variants are cleaved after residue 328 by proteases, removing the protein’s noncatalytic heavy chain (HC) region. Truncation of the pathological FXII T309K by plasmin exposes R372 for subsequent cleavage by plasma kallikrein in solution (de Maat S et al. 2016, 2019). The intrinsic capacity of the truncated form of FXII (329-615) to convert PK to kallikrein is greater than that of activated FXII leading to more kallikrein generated early during reciprocal activation (Ivanov I et al. 2019). Second, the truncated form of FXII (329-615) is a better kallikrein substrate than is FXII (Ivanov I et al. 2019). Further, the accelerated PK/FXII activation in HAE patients carrying FXII variants appears to overwhelm the regulatory function of SERPING1 at normal plasma levels leading to uncontrolled bradykinin formation in a surface-independent manner (de Maat S et al. 2016; Ivanov I et al. 2019).
kininogen:C1q binding protein
tetramer(factor
IIa):SERPIND1XIa:GPIb:GPIX:GPV
complexXIa:GPIb:GPIX:GPV
complexXIa:GPIb:GPIX:GPV
complexWillebrand factor
multimerWillebrand factor
multimerSERPINC1:SERPINC1
activatorsSERPINC1:SERPINC1
activators