International Journal of
Progress in Hematology
HEMATOLOGY
von Willebrand Factor–Cleaving Protease and Upshaw-Schulman Syndrome Yoshihiro Fujimura,a Masanori Matsumoto,a Hideo Yagi,b Akira Yoshioka,c Taei Matsui,d Koiti Titanid a
Department of Blood Transfusion Medicine, bSecond Department of Internal Medicine, and cDepartment of Pediatrics, Nara Medical University, Nara; dInstitute for Comprehensive Medical Science, Fujita Health University, Aichi, Japan Received October 1, 2001; accepted October 19, 2001
Abstract Vascular endothelial cell (EC)-produced plasma von Willebrand factor (vWF) plays a critical role in primary hemostasis through its action of anchoring platelets onto the injured denuded subendothelial matrices under high shear stress. Unusually large vWF multimers (UL-vWFMs), present in plasma immediately after release from ECs, are most biologically active, but they are soon cleaved and degraded into smaller vWFMs by a specific plasma protease, termed vWF-cleaving protease (vWFCPase), in normal circulation. Recent studies on the relationship between UL-vWFMs and vWF-CPase, together with its autoantibody (inhibitor) have brought about a clear discrimination between thrombotic thrombocytopenic purpura and hemolytic uremic syndrome. Furthermore, a congenital deficiency of this enzyme activity has been shown to cause UpshawSchulman syndrome, a complex constitutional bleeding diathesis. Successful purification of vWF-CPase revealed that this enzyme is composed of a single polypeptide with a molecular mass of approximately 190 kd, and its complementary DNA cloning unambiguously indicated that it is uniquely produced in the liver and its gene is located on chromosome 9q34. The messenger RNA of vWF-CPase had a span of 4.6 kb, and its enzyme was designated ADAMTS 13. The predicted complete amino acid sequence of this enzyme consisted of 1427 residues, including a signal peptide, a short propeptide terminating in the sequence RQRR, a reprolysin-like metalloprotease domain, a disintegrin-like domain, a thrombospondin-1 repeat (TSP1), a cysteine-rich domain, an ADAMTS spacer, 7 additional TSP1 repeats, and 2 CUB domains. Int J Hematol. 2002;75:25-34. ©2002 The Japanese Society of Hematology Key words: vWF; UL-vWF; vWF-CPase; TTP; USS; ADAMTS
platelets and released upon stimulation into circulation [5]. The amount of platelet-derived vWF in circulation is assumed to be less than 2% of the total [4,5]. Functionally, EC-derived vWF plays an essential role in primary hemostasis through its anchoring action of platelets onto the injured denuded subendothelial matrices under high shear stress [68], but the role of platelet-derived vWF remains to be investigated [5]. Unusually large vWF multimers (UL-vWFMs), released into plasma from vascular ECs, are most active in interacting with platelets [6-9], but they are soon cleaved and degraded into smaller vWFMs by a specific plasma protease, termed vWF-cleaving protease (vWF-CPase), in normal circulation, [10,11]. Thus, vWF functions in the direction of thrombosis, whereas vWF-CPase functions in the opposite direction of thrombosis in healthy individuals. Recent studies on the relationship between UL-vWFMs and vWFCPase together with its inhibitor have brought about a clear
1. Introduction von Willebrand factor (vWF) is a glycoprotein synthesized in endothelial cells (ECs) and megakaryocytes (MCs) [1-3]. EC-derived vWF bearing ABO–blood group structures in Asn-linked sugar chains of its molecule [4] is stored in the Weibel-Palade body and then secreted into subendothelial matrices constitutively or upon stimulation into circulation. In contrast, MC- or platelet-derived vWF containing no ABO–blood group sugar chains is stored in the -granule of
Correspondence and reprint requests: Yoshihiro Fujimura, MD, Department of Blood Transfusion Medicine, Nara Medical University, 840 Shijyo-cho, Kashihara City, Nara 634-8522, Japan; 81-744-22-3051, ext. 3289; fax: 81-744-29-0771 (e-mail: yfujimur@ nmu-gw.cc.naramed-u.ac.jp).
25
26
Fujimura et al / International Journal of Hematology 75 (2002) 25-34
discrimination between thrombotic thrombocytopenic purpura (TTP), characterized by Moschkowitz’s clinical pentad [12] (thrombocytopenia, microangiopathic hemolytic anemia [MAHA], fluctuating neurologic abnormalities, renal disease, and fever), and hemolytic uremic syndrome (HUS), characterized by Gasser’s clinical triad [13] (thrombocytopenia, MAHA, and acute renal failure), which is similar to TTP but lacks neurologic symptoms [14-17]. Furthermore, along this line of studies, a new concept of congenital TTP has been established [14,15,18]. In this article, we focus on recent hot spots in this research field together with a review of the historical background.
2. vWF-CPase 2.1. Historical Background Since the late 1960s, many articles have been published on the purification of “factor VIII,” now known as factor VIII/ von Willebrand factor (VIII/vWF) complex, from human or bovine plasma [19-25]. In 1974, van Mourik et al [26] reported that a 24-hour dialysis against a 10 mmol/L sodium phosphate buffer (pH 7.0) resulted in a purified factor VIII with an enormously high molecular mass dissociated into 2 fragments with much smaller molecular masses. These 2 fragments were termed fast- and slow-moving components according to the mobility demonstrated by large-pore polyacrylamide gel electrophoresis (PAGE) [26]. Of particular interest was that this dissociation was inversely dependent on the ionic strength of the dialysis buffer. In those days, however, a molecular configuration of factor VIII or FVIIIrelated antigen (VIIIR:AG) with several different biological functions such as procoagulant FVIII activity (VIII:C), platelet adhesion, and platelet aggregation in the presence of antibiotic ristocetin was unknown, even as far as whether it is composed of 1 or 2 molecules [27-34]. The latter query arose from experimental results in which VIII:C was separated from VIIIR:AG under conditions of a high ionic strength such as 1 mol/L NaCl or 0.25 mol/L CaCl2 [27,29-31]. Thus, additional data that could explain the generation of both the fast- and slow-moving components under a low ionic strength condition had not been found, and this phenomenon remained a mystery until the existence of specific vWFCPase was found in 1996 [10,11]. During 1984-1986, critical evidence that VIIIR:AG was indeed a complex of 2 distinct molecules, FVIII and vWF, was presented, and the respective complementary DNAs (cDNAs) were cloned and demonstrated production under different gene control [35-40]. The stoichiometry of FVIII and vWF in plasma is approximately 1:50, and FVIII and monomeric vWF have similar molecular masses of approximately 250 kd. Therefore, vWF represents 98% of the molecular mass of the FVIII/vWF complex [41]. This finding also indicates that the dissociation of factor VIII into the 2 components under conditions of low ionic strength is mostly reflected in what occurs in vWF. A sophisticated sodium dodecyl sulfate (SDS)-agarose gel electrophoretic analysis for high–molecular-mass vWF was established during 1980-1981, demonstrating that plasma vWF is composed of heterogenous multimers with molecular masses ranging from 500 to 20,000 kd [42,43]. In
Figure 1. Structure and functional domains of von Willebrand factor (vWF), and its cleavage site with vWF-cleaving protease (vWF-CPase). Each vWF subunit, consisting of the 2050 amino acid (aa) residues, contains a set of functional domains interacting with factor VIII, platelet glycoprotein (GP) Ib, integrin IIb3, and subendothelial collagen. These identical vWF subunits are held together by intersubunit disulfide bonds (SS) in a tail-to-tail and head-to-head fashion [6]. The vWF multimers (vWFMs) are cleaved into much smaller vWFMs at the site of the Tyr 842–Met 843 bond by the action of a plasma enzyme termed vWFcleaving protease (vWF-CPase), resulting in the coappearance of the 2 homodimers of the N-terminal residue 1-842 and the C-terminal residue 843-2050 (top panel). In the bottom panel, our assay of plasma vWFCPase activity according to Furlan et al [14] with a slight modification [18] is shown, where the standard curve is obtained using diluted normal plasma. P indicates plasma of the patient with Upshaw-Schulman syndrome shown in Figure 3.
1986, the structural analyses of vWF revealed that it is composed of a single subunit of 2050 residues, linked together by disulfide bonds in a head-to-head and tail-totail fashion [44-46]. In parallel with these analyses, several binding domains of vWF to FVIII, collagen, platelet glycoprotein (GP)Ib, and GPIIb/IIIa, critically important for the function, were identified on each subunit [6] (Figure 1, top). Wagner et al [47] subsequently demonstrated that the formation of vWFMs takes place in the endoplasmic reticulum, and the initial dimer is assembled from a pair of 250-kd vWF subunits via an intermolecular disulfide bond formed
vWF-Cleaving Protease and Upshaw-Schulman Syndrome
by a cysteine residue located in the C-terminal region of each subunit. The following multimerization is achieved by interdimer disulfide bonds in the N-terminal region of each subunit. Thus, the largest molecular mass of plasma vWF is approximately 20,000 kd, but it has been assumed to be degraded into much smaller vWFMs by in vivo proteolysis during circulation. In fact, in some types of patients with von Willebrand disease (vWD), a congenital bleeding disorder due to abnormality of the vWF molecule, the larger vWFMs are absent as a consequence of possible conformational changes that may expose a sensitive site to the action of specific protease(s). Type 2A vWD is typical of this case [48]. Dent et al [49] showed that a lack of high–molecular-mass vWFMs in patients with type 2A vWD is caused by a specific cleavage of the Tyr 842–Met 843 bond of the vWF subunit. The enzyme responsible for this cleavage, however, was initially assumed to be a calcium-dependent neutral protease (eg, calpain) [49], but later this assumption was completely repudiated [10,11]. Since the end of the 1980s, the physiological significance of shear-induced platelet aggregation (SIPA), especially high SIPA, has been its association with arterial thrombi formation [50-53]. Furthermore, Siedlecki et al [54] showed the critical effects of shear-dependent changes on the 3-dimensional structure of vWF, from globular to extended, using atomic-force microscopy, suggesting that through such a conformational change vWF gains an ability to interact with platelets, because vWF does not interact with platelets in normal circulation. Tsai et al [55] also demonstrated that high shear stress enhances proteolysis of plasma vWF. This effect may indicate that the altered molecular conformation of vWF under high shear stress may expose a sensitive site to the action of protease specific to vWF, in a similar manner to a scenario of type 2A vWD. In 1996, Furlan et al [10] reported the partial purification and characterization of plasma vWF-CPase, which has a molecular mass of 300 kd before reduction and 70 to 80 kd after reduction on an SDS-PAGE analysis and specifically cleaves the Tyr 842–Met 843 bond of the vWF subunit. The partially purified vWF-CPase required the presence of 1 mol/L urea as a protein denaturant; a divalent cation such as Ba2+, Sr2+, or Ca2+; and a long incubation time (24 hours) to cleave the Tyr 842–Met 843 bond. Tsai [11] also reported the partial separation of this enzyme, with a molecular mass of 200 kd estimated by size-exclusion chromatography, but did no further structural characterizations. In summary, both groups of investigators demonstrated that the enzyme activity was inhibited by 1 mmol/L EDTA, EGTA, or o-phenanthroline, but not by other protease inhibitors, including 2 mmol/L phenylmethylsulfonyl fluoride (PMSF), 1.2 mmol/L diisopropyl fluorophosphate (DFP), 2000 KIU/mL aprotinin, 1 mmol/L leupeptin, 2 mmol/L iodoacetamide (IAA), and 6 mmol/L N-ethylmaleimide (NEM). Furthermore, these investigators indicated that the enzyme activity inhibited by EDTA could be restored by the addition of excessive Ca2+, and that heat treatment of this enzyme preparation at 56C for 60 minutes destroyed its activity. These results unambiguously demonstrated that vWF-CPase is a metalloprotease.
27
2.2. Assays of vWF-CPase Activity and Its Inhibitor In 1997, Furlan et al [14] reported the quantitative assay method for plasma vWF-CPase activity. This method was based on the degree of degradation of vWFMs, where vWFCPase in diluted plasma samples was activated with 10 mmol/L BaCl2 for 5 minutes in the presence of 10 mmol/L Pefabloc, a broadly acting serine protease inhibitor. After addition of the purified vWF as a substrate to this mixture, the whole mixture was dialyzed using a hydrophilic filter at 37C for 24 hours against 5 mmol/L Tris-HCl (pH 8.0) containing 1.5 mol/L urea (a low ionic strength buffer with a protein denaturant). This assay is very elegant, but is time consuming and tedious. Recent assays in some laboratories including ours are performed without the dialysis step described above, but with an increased enzyme/substrate ratio [18] (Figure 1, bottom). The detection limit of plasma vWF-CPase activity in these assays was approximately 3% of the control activity, and the result obtained with healthy subjects (n = 30; mean, 2 SD) was 96.2% 37.4% of the pooled normal plasma [18]. In contrast, the method of Tsai and Lian [16], reported in 1998, was based on the generation of 2 homodimeric vWF fragments (residues 1-842 and 8432050) in the mixture of purified vWF and diluted plasma samples after a 60-minute incubation at 37C. The 2 fragments were separated on SDS-PAGE, followed by immunoblotting using an anti-vWF monoclonal antibody (MoAb) and then analyzing by densitometry. This assay also appears to be time-consuming. As an alternative to these gel electrophoresis assays of vWF-CPase activity, an enzyme-linked immunosorbent assay (ELISA) using the collagen-binding activity of vWF was developed by Gerritsen et al [56]. This assay used a phenomenon in which the degraded vWFMs with smaller molecular masses show impaired binding to microtiter plates coated with human collagen type III. Furthermore, an immunoradiometric assay (IRMA) for vWF-CPase activity was also developed by Obert et al [57]. This assay was based on the degradation of a constant amount of wild-type recombinant vWF as a substrate. In this assay, the amount of vWF antigen was determined by a 2-site IRMA, where a MoAb with the epitope localized on the C-terminal side of the cleavage site was used as the first antibody capturing vWF as a substrate, and after digestion, a pool of labeled MoAbs with the epitope localized in the N-terminal side was used as the second antibody. The immunoglobulin (Ig)G-type autoantibody-inhibiting vWF-CPase activity (inhibitor) develops in some patients with acquired TTP as described below. Such an inhibitor can be assayed according to the reports of Furlan et al [14] and Tsai and Lian [16]. In our laboratory, we use the Bethesda method, originally developed for the measurement of factor VIII inhibitor [58]. Briefly, a test plasma was first heat treated at 56C for 60 minutes to inactivate its own protease activity. The supernatant obtained after centrifugation or a control sample (50 mmol/L Tris-buffered saline, pH 7.4) was mixed with an equal amount of pooled normal plasma. After further incubation at 37C for 2 hours, the residual vWF-CPase activities in the test sample and the control sample were measured [18]. One unit of the inhibitor was defined as the
28
Fujimura et al / International Journal of Hematology 75 (2002) 25-34
amount that reduced the vWF-CPase activity to 50% of the control activity [58]. We judge that the inhibitor is positive in the specimen if it contains a titer of more than 0.1 Bethesda units/mL. The highest inhibitor titer obtained in our laboratory through testing 213 samples from 70 patients with acquired TTP was 320 Bethesda units/mL, which was found in a 9-month-old female infant [59]. Development of such an inhibitor with a high titer has been extremely rare in adult patients with acquired TTP [60], as described below, but Tsai [61] recently reported an adult patient who was refractory to therapy and this case was fatal. The inhibitor titer is also assayed using the patient’s IgGs, purified from the plasma using a protein A- or G-coupled agarose column, and thereby, the specific inhibitor titer can be expressed as Bethesda units/mg IgG.
Recently, Raife et al [69] have found that saliva contains vWF-CPase activity in at least 80-times greater amounts than does plasma. Like plasma vWF-CPase, the saliva protease activity was not inhibited by Pefabloc, a serine protease inhibitor, but unlike plasma vWF-CPase, the saliva protease activity was neither appreciably enhanced by BaCl2 nor clearly inhibited by the addition of 10 mmol/L EDTA alone. Interestingly, 2 pediatric patients with chronic relapsing (CR)-TTP had deficient activity in both the plasma and saliva proteases [69]. However, during recurrent episodes of sporadic TTP, 3 adult patients demonstrated deficient plasma vWF-CPase activity but normal saliva protease activity [69]. These results suggest that saliva vWF-CPase is related to but distinct from the enzyme in plasma.
2.3. Discriminating Between TTP and HUS
2.5. Structure of Plasma vWF-CPase and Its Production in Liver
TTP is a severe multiorgan disorder [12] and develops mainly in adults who have underlying conditions such as autoimmune diseases, pregnancy, drug intake, and preceding infections including HIV [62-64]. In contrast, HUS is often observed in young children, and is commonly followed by acute enterocolitis due to the Stx-producing Escherichia coli O157:H7 infection [65]. A differential diagnosis of TTP and HUS based on clinical symptoms has been difficult, especially in young children. After the introduction of plasma exchange (PE) for the treatment of TTP, the mortality rate of TTP has subsided to 10% to 20%, whereas the previous rate was beyond 80% [62-64]. Furlan et al [14,15,17] and Tsai and Lian [16] independently showed that a majority of acquired TTP with a decreased activity of plasma vWF-CPase is often associated with the presence of an IgG-type autoantibody (inhibitor) to this enzyme, but such findings are not observed in HUS, leading to a clear differential diagnosis between these 2 diseases. Furthermore, the antibody titers in adult TTP patients have been reported to be relatively low in general, and sometimes at an almost undetectable level [15,16,60]. PE, however, not only removes the inhibitor efficiently, but also supplies vWFCPase to neutralize the inhibitor, if any, and to degrade ULvWFMs. In contrast, neither a substantial decrease in vWFCPase activity nor the appearance of its inhibitor was found in both the groups of patients with acquired HUS [15,16] and a minority of patients with acquired TTP such as bone marrow transplantation-associated TTP [66].
2.4. vWF-CPase Activity in Other Conditions and Saliva Although the studies are still under investigation at present, some data appear to be almost consistent regarding the observation of low vWF-CPase activity in full-term newborns (n = 33; 60% 26%) [67], premature cord plasmas (n = 20; 49.5% 12.8%) [68], women during normal pregnancy (n = 69; the last 2 trimesters versus the first: 60% 26% versus 93% 21%) [67], and in patients with decompensated liver cirrhosis (n = 42; 46% 33%) [67], acute inflammatory state (n = 15; 40% 12%), or uremia (n = 63; 74% 29%) [67].
Recently, Fujikawa et al [70] succeeded in purifying plasma vWF-CPase as a single-chain glycoprotein with a molecular mass of 150 kd before reduction and 190 kd after reduction, as analyzed by SDS-PAGE. Because their starting material was FVIII/vWF concentrate, the concentration and the yield of the enzyme in and from plasma are both uncertain. A search of the human genome sequences with the N-terminal amino acid sequence of this purified vWF-CPase indicated that the positively identified residues in this protein were found in the sequence translated from human chromosome 9q34. The probe sequence from chromosome 9q34, encompassing the aligned region, was used to search the human EST database. In this way, vWF-CPase was identified as a new member of the ADAMTS (a disintegrin and metalloproteinase with a thrombospondin type I motif) family of metalloproteinase [71] with the active site sequence of HEIGHSFGLEHD, which was located at 150 to 161 residues from the N-terminal Ala residue [70]. The N-terminal AAGG sequence also indicated the presence of a precursor or propeptide with a typical furin processing site. Gerritsen et al [72] also succeeded in purifying plasma vWF-CPase by an immunoadsorbent chromatography, using the IgG fraction from a patient with acquired vWF-CPase deficiency due to vWF-CPase autoantibodies, coupled with a series of chromatographic steps. The purified enzyme consisted of a series of 150-, 140-, 130-, and 110-kd bands as analyzed by SDSPAGE, but all of them shared the same N-terminal sequence, suggesting that they were derived from the same polypeptide chain that had been partially degraded at the C-terminal end. According to Gerritsen’s data, approximately 1 mg of the purified enzyme was recovered from 800 mL of normal plasma as a starting material, with an activity yield of 2.3% of the starting material [72]. Regarding the target organ that produces vWF-CPase, we have originally reported that a severely decreased level of plasma vWF-CPase activity in patients with biliary atresia can be restored after living-related liver transplantations. Thus, we suggested that liver is a major organ in synthesizing plasma vWF-CPase [73]. This speculation is consistent with the recent successful cDNA cloning of vWF-CPase by 2 groups of investigators [74,75] who, using multiple human tis-
vWF-Cleaving Protease and Upshaw-Schulman Syndrome
29
3. Upshaw-Schulman Syndrome 3.1. Historical Background
Figure 2.
Complete amino acid sequence of human von Willebrand factor–cleaving protease (vWF-CPase) predicted from the complementary DNA sequence by Zheng et al [74] (see the reference for details).
sue Northern blotting analysis, showed that the mRNA encoding this protease spanned approximately 5 kb and was uniquely expressed in the liver. Of these 2 groups, Zheng et al [74] showed the 4.6-kb cDNA sequence for vWF-CPase, designated ADAMTS 13, and predicted the complete amino acid sequence of 1427 residues of the gene product, which consists of a signal peptide, a short propeptide terminating in the sequence RQRR, a reprolysin-like metalloprotease domain, a disintegrin-like domain, a thrombospondin-1 repeat (TSP1), a cysteine-rich domain, an ADAMTS spacer, 7 additional TSP1 repeats, and 2 CUB domains (Figure 2). According to these investigators, vWF-CPase apparently is made as a zymogen that requires proteolytic activation, possibly by furin intracellularly. In addition, they found the following interesting structural features: (1) Sites for Zn2+ and Ca2+ ions are conserved in the protease domain. (2) The cysteine-rich domain contains an RGDS sequence that could mediate integrin-dependent binding to platelets or other cells. (3) Alternative splicing gives rise to at least 7 potential variants that truncate the protein at different positions after the protease domain and may have distinct abilities to interact with cofactors, connective tissues, platelets, and vWF. Almost simultaneously, another group of investigators [75] succeeded in the cDNA cloning of plasma vWF-CPase.
In 1953, after reviewing 12 cases of atypical congenital hemolytic anemia, Dacie et al [76] reported a 6-year-old girl (their case 12) who had repeated episodes of severe jaundice, thrombocytopenia, hemolytic anemia, and fragmented red cells from the time she was a newborn infant. Before her first visit to their hospital, she had received a splenectomy in another hospital but without clinical improvement, and after a long history of illness, she died of renal failure at age 7. This patient was the third of 4 children. The first child was jaundiced at birth and died of hemorrhage at age 2. The second child was also jaundiced at birth and died on the fourth day of life as a result of bleeding from the bowel. The fourth child and their parents lacked these symptoms and were apparently healthy. Thus, the authors concluded that the proposita seemed to have suffered from hitherto unrecognized hereditable blood dyscrasia, with at least 1 or possibly 2 siblings affected. Then, in 1967, the case of a female infant who had symptoms similar to Dacie’s patient and who died at age 9 months after a history of illness was reported by Monnens et al after autopsy as a solitary case of TTP [77]. Subsequently, in 1975, the case of a family, in which 4 out of 7 siblings suffered from TTP was reported by Wallace et al [78]; however, none of the affected members developed clinical symptoms during their newborn period, indicating that Wallace’s cases are similar to, but different from, Dacie’s and Monnens’ cases, especially in the prevalence of age as a significant clinical sign. Presumably, totally apart from any concept of TTP, Schulman et al [79] reported in 1960 the case of an 8-year-old girl born in Germany who had repeated episodes of bleeding associated with chronic thrombocytopenia and MAHA but without specific clotting factor abnormalities. The onset of her bleeding episodes could be traced back to her newborn period, when she had a large ecchymosis on the dorsum of 1 hand. Interestingly, the abnormal bleeding, thrombocytopenia, and MAHA were transiently but dramatically corrected by the infusion of normal fresh frozen plasma (FFP). Therefore, the authors proposed that the patient might have had a congenital deficiency of a plasma platelet-stimulating factor. In 1978, Upshaw [80] described a 29-year-old female patient who, from the age of 6 months to 12 years, had 6 to 7 episodes yearly, characterized by thrombocytopenia and MAHA, and who received an effective plasma infusion. This was a case very similar to Schulman’s, but lacked the description of jaundice during the newborn period.Then, Rennard and Abe [81] reported an Upshaw’s case with a slightly decreased level of plasma cold-insoluble globulin (fibronectin) during the acute phase, but with normal levels between the episodes. Furthermore, they proposed a Upshaw-Schulman syndrome (USS) nomenclature for such patients. No correlation between fibronectin level and disease activity, however, could be confirmed in 2 USS patients described by Koizumi et al [82] and Goodnough et al [83], including Schulman’s original case, and both groups of investigators found normal levels of plasma fibronectin regardless of the clinical course. In 1982, the case of a 4-year-old Japanese girl with symptoms
30
Fujimura et al / International Journal of Hematology 75 (2002) 25-34
equivalent to those characteristic of USS was reported, but with a diagnosis of congenital MAHA [84]. This case also indicated that USS exists regardless of race. The inheritance of USS has been thought to be autosomally recessive, because more than 2 siblings in the same family are often affected but their parents are apparently asymptomatic [18]. Moake et al [85] first used the term CR-TTP for 4 of their patients with TTP who had a chronic relapsing clinical course, including both the Schulman’s case and patients with an apparently acquired form of TTP. Most importantly, however, Moake et al [85,86] demonstrated that UL-vWFMs were present in the plasma of their patients with CR-TTP at the early remission stage, but disappeared in the acute phase of the disease or after infusion of FFP. The clinical efficiency of several plasma components for a Japanese boy with CR-TTP was extensively evaluated by Miura et al [87]; the effective preparations were whole blood, fresh plasma or FFP, cryosupernatant, cryoprecipitate, and FVIII/vWF concentrates. In contrast, the ineffective preparations were albumin, -globulin, fibronectin, fibrinogen, and factor IX concentrate [87]. Clinical restoration by the infusion of FVIII/vWF concentrates for USS patients was also confirmed by other groups of investigators [88,89]. Conversely, Hara et al [88] reported clinical exacerbation with the reappearance of the acute-phase symptoms within 1 hour after the infusion of 1-desamino-8-D-arginine vasopressin
(DDAVP) for a patient with USS. After thrombopoietin was cloned and its sensitive assays were developed, Miura et al confirmed normal levels of plasma thrombopoietin in 5 Japanese patients with USS [90]. Thus, the pathophysiology of USS has remained unexplained, and the relationship between USS and CR-TTP is obscure.
3.2. Diagnosis A major advance in discriminating between acquired CRTTP and USS was made by the development of a quantitative assay for the aforementioned vWF-CPase activity. After analyzing 4 patients belonging to 3 separate families, Furlan et al [14] reported that CR-TTP is associated with a deficiency in vWF-CPase activity. Of the 4 patients, however, 2 (their cases A1 and A2) were siblings with early onset of the disease, and both had extremely low levels of vWF-CPase activity, which was measured on 2 different occasions. The other 2 patients (their cases B and C) were unrelated and both had CR-TTP of the late onset type. Cases B and C both showed deficiencies of vWF-CPase activity on 1 occasion, but only moderately decreased levels of activity on another occasion. This data may simply suggest that the former results differ from the latter in the etiology of CR-TTP. Based on these studies, we measured plasma vWF-CPase activity in 3 Japanese patients who had clinical diagnoses of USS and who
Figure 3. A typical family pedigree of Upshaw-Schulman syndrome (USS) and the presence of unusually large von Willebrand factor multimers (UL-vWFMs) in the patient’s plasma. The proposita (ST), marked with an arrow and P, is a Japanese girl born in 1986 as the fourth child of parents who were not consanguineous [18]. Soon after birth, the patient developed severe indirect hyperbilirubinemia and thrombocytopenia of unknown etiology that required exchange blood transfusion on the first day of life. Her eldest sister died of melena on the fourth day after birth, and her elder brother died of spontaneous intracranial bleeding at the age 13. This brother had an episode of severe jaundice in his newborn period that required an exchange blood transfusion. Subsequently, he had repeated clinical manifestations of microangiopathic hemolytic anemia and thombocytopenia, very similar to the proposita. Her parents and elder sister had no sign of bleeding tendency. From birth to 4 years old, the proposita had numerous episodes of hemolysis that were treated by plasma exchange or fresh frozen plasma (FFP) infusion. A diagnosis of USS was made when she was 4 years old based on the clinical findings. Plasma vWF-cleaving protease (vWF-CPase) activity of the proposita is below 3% of the control as demonstrated in Figure 1, bottom panel, and that of her family members is shown as a percentage of the control in the figure. Left panel: circles indicate female family members, and squares indicate male family members. Double circles and double squares indicate that the family members’ vWF-CPase activities were assayed; vWF-CPase activities were not assayed in the family members identified by single circles or squares. Dark circles and squares indicate that these people were bleeders, and half-dark circles and squares identify asymptomatic carriers. Cross marks (†) indicate death; ND, not determined. Right panel: the presence of UL-vWFMs is demonstrated by sodium dodecyl sulfate (SDS)-1.2% agarose gel electrophoresis analysis, using the patient’s plasma at the remission stage (7 days after FFP infusion) [91].
vWF-Cleaving Protease and Upshaw-Schulman Syndrome
31
belonged to separate families; we also measured plasma vWF-CPase activity in their family members and found that USS shows a congenital deficiency in vWF-CPase activity, establishing a concept of congenital TTP [18]. The most striking clinical picture for these 3 patients was severe indirect hyperbilirubinemia, which developed soon after birth and required exchange blood transfusions. Their parents have a moderately decreased activity of plasma vWF-CPase, indicating a coinheritance of USS and vWF-CPase, and therefore, they are assumed to be asymptomatic carriers. These findings appear to be in good accord with the recent discovery of the gene for vWF-CPase, which is located on chromosome 9q34 [70,71,74,75]. A Japanese patient with typical USS and her family pedigree are shown in Figure 3, along with their vWF-CPase activity and the presence of UL-vWFMs in the patient’s plasma [18].
3.3. Mechanism of Thrombocytopenia The mechanism of thrombocytopenia and MAHA in USS has not yet been fully elucidated. However, our recent results show that the plasma of patients with USS enhances the aggregation of normal platelets under high shear stress, which is totally dependent on both an axis of vWF-GPIb interaction and ADP released from activated platelets [91]. Together with the previous data, therefore, we propose the hypothesis illustrated in Figure 4. Namely, UL-vWFMs, produced exclusively in ECs and released into circulation, are transported to peripheral small arteries where continuously generated high shear stress may change the molecular conformation of the “inactivated” UL-vWFMs to “activated” forms [54], which are more accessible to vWF-CPase. In USS patients, however, vWF-CPase is totally defective, and therefore the activated UL-vWFMs interact more intensively with platelet GPIb and generate signals, which include the endogenous ADP released from platelets, to activate selfplatelets, resulting in the further acceleration of platelet activation [92-95]. A series of these reactions leads to the formation of platelet microaggregates, resulting in the generation of thrombocytopenia. Furthermore, microangiopathic changes in red cells, schistocytes, are assumed to take place by collision with fibrin strands, which are formed surrounding the platelet thrombi in TTP, as reported in previous publications [96-99]. Clinical pictures in USS patients are often dramatically exacerbated by a cold or any tiny acute inflammation [100], suggesting that some inflammatory cytokines may augment the release of vWF from ECs over vWF-CPase that might be the trigger. This speculation appears to be supported by the previous observation of an acute aggravation of symptoms with USS on administration of DDAVP [88]. Because the platelet count regularly drops 2 to 3 weeks after FFP infusion, we assume that some time interval is required for UL-vWFMs to accumulate in the plasma to the level at which clinical signs develop, even though the UL-vWFMs have been almost completely destroyed once by a small amount of vWF-CPase in the infused FFP [91]. Thus, USS patients usually do not require plasma exchange, and a simple supplementation of vWF-CPase by FFP infusion (10 mL/kg body wt) every 2 to 3 weeks is justified to prevent
Figure 4. A proposed mechanism of thrombocytopenia and microangiopathic hemolytic anemia (MAHA) in patients with Upshaw-Schulman syndrome (USS). As indicated by the arrows, unusually large von Willebrand factor multimers (UL-vWFMs), held together by intersubunit disulfide bonds (SS), are produced exclusively in vascular endothelial cells (ECs) and released into circulation upon stimulation; then they are transported to peripheral small arteries where high shear stress is continuously generated. In these circumstances, the “inactivated (globular)” UL-vWFMs may change their molecular conformation to the “activated (extended)” form, which is more accessible to vWF-cleaving protease. In USS patients, however, this enzyme activity is absent, and therefore the activated UL-vWFMs interact more intensively with platelet glycoprotein (GP) Ib that generates signals to activate the self-platelets, including Ca2+ influx to platelets and adenosine diphosphate (ADP) release from platelets. A series of these reactions leads to the formation of platelet microaggregates with vWF and fibrinogen (Fbg), resulting in thrombocytopenia. Microangiopathic change of red blood cells (RBCs), to schistocytes (SCs), is assumed to take place by collision with fibrin (Fbn) strands, which are formed surrounding the platelet thrombi.
TTP, unless the patients develop the inhibitor (alloantibody) against vWF-CPase. On the other hand, it is noteworthy that the plasma vWF of USS patients, in spite of a severe deficiency of plasma vWFCPase activity, consists of a series of the multimers together with UL-vWFMs. This observation simply suggests that, in
32
Fujimura et al / International Journal of Hematology 75 (2002) 25-34
addition to proteolytic processing with vWF-CPase, another mechanism, such as the regulation of disulfide-bond formation for vWFMs, which is not impaired in USS may exist. In this regard, Xie et al [101] recently showed the presence of vWF disulfide-bond reductase activity in the conditioning medium of human umbilical vascular ECs or dermal microvascular ECs. This reductase or depolymerase activity was inactivated by heat treatment or with a thiol blocking reagent such as IAA or NEM. Furthermore, the reduction in multimer size by this mechanism was not associated with any appreciable peptide bond cleavage, suggesting that this depolymerase is a protein but not a protease. Recently, Xie et al [102] finally identified that the depolymerase is thrombospondin-1.
Acknowledgments This work was supported in part by research grants from the Japanese Ministry of Education, Culture, Sports, Science, and Technology (to Y.F. and M.M.) and from the Ministry of Health and Welfare of Japan for Blood Coagulation Abnormalities (to Y.F.). We are indebted to Dr. Kazuo Fujikawa at the Department of Biochemistry, University of Washington, for sending a preprint of his manuscript before publication. We also thank to Dr. J. Evan Sadler at the Howard Hughes Medical Institute of Washington University School of Medicine for his generous permission for the use of Figure 2. Note: After the submission of our manuscript, the following two interesting and related papers have been published: (1) Levy GG, Nichols WC, Lian WC, et al. Mutations in a member of the ADAMTS gene family cause thrombotic thrombocytopanic purpara. Nature. 2001;413:488-494. (2) Brass L, VWF meets the ADAMTS family. Nature Med. 2001;7:1177-1178.
References 1. Bloom AL, Giddings JC, Wilke CJ. Factor VIII on the vascular intima: possible importance in hemostasis and thrombosis. Nature. 1973;241:217-219. 2. Jaffe EA, Hoyer LW, Nachman RL. Synthesis of antihemophilic factor antigen by cultured human endothelial cells. J Clin Invest. 1973;60:914-921. 3. Sporn LA, Chavin SI, Marder VJ, Wagner DD. Biosynthesis of von Willebrand protein by human megakaryocytes. J Clin Invest. 1985; 76:1102-1106. 4. Matsui T, Fujimura Y, Nishida S, Titani K. Human plasma 2macroglobulin and von Willebrand factor possess covalently linked ABO(H) blood antigens in subjects with corresponding ABO phenotype. Blood. 1993;82:663-668. 5. Matsui T, Shimoyama T, Matsumoto M, et al. ABO blood group antigen on human plasma von Willebrand factor after ABOmismatched bone marrow transplantation. Blood. 1999;94: 2895-2900. 6. Fujimura Y, Titani K. Structure and function of von Willebrand factor. In: Bloom AL, Forbes CD, Thomas DP, Tuddenham EGD, eds. Haemostasis and Thrombosis. New York, NY: Churchill Livingstone; 1994:379-395. 7. Ruggeri ZM. von Willebrand factor. J Clin Invest. 1997;99:559-564. 8. Sadler JE. Biochemistry and genetics of von Willebrand factor. Annu Rev Biochem. 1998;67:395-424. 9. Furlan M. von Willebrand factor: molecular size and functional activity. Ann Hematol. 1996;72:341-348.
10. Furlan M, Robles R, Lämmle B. Partial purification and characterization of a protease from human plasma cleaving Von Willebrand factor to fragments produced by in vivo proteolysis. Blood. 1996; 87:4223-4234. 11. Tsai H-M. Physiologic cleavage of von Willebrand factor by a plasma protease is dependent on its conformation and requires calcium ion. Blood. 1996;87:4235-4244. 12. Moschcowitz E. Hyaline thrombosis of the terminal arterioles and capillaries; a hitherto undescribed disease. Proc N Y Pathol Soc. 1924;24:21-24. 13. Gasser C, Gautier E, Steck A, Siebenmann RE, Oechslin R. Hämolytisch-urämische syndrome: bilaterale Nierenrindennekrosen bei akuten erworbenen hämolytischen Anämien. Schweiz Med Wochenschr. 1955;85:905-909. 14. Furlan M, Robles R, Solenthaler M, Wassmer M, Sandoz P, Lämmle B. Deficient activity of von Willebrand factor-cleaving protease in chronic relapsing thrombotic thrombocytopenic purpura. Blood. 1997;89:3097-3103. 15. Furlan M, Robles R, Galbusera M, et al. von Willebrand factorcleaving protease in thrombotic thrombocytopenic purpura and the hemolytic-uremic syndrome. N Engl J Med. 1998;339:1578-1584. 16. Tsai H-M, Lian EC-Y. Antibodies to von Willebrand factor-cleaving protease in acute thrombotic thrombocytopenic purpura. N Engl J Med. 1998;339:1585-1594. 17. Furlan M, Robles R, Solenthaler M, Lämmle B. Acquired deficiency of von Willebrand factor cleaving protease in a patient with thrombotic thrombocytopenic purpura. Blood. 1998;91: 2839-2846. 18. Kinoshita S, Yoshioka A, Park Y-D, et al. Upshaw-Schulman syndrome revisited: a concept of congenital thrombotic thrombocytopenic purpura. Int J Hematol. 2001;74:101-108. 19. Hershgold EJ, Sprawls S. Molecular properties of purified human, bovine and porcine antihemophilic globulin (AHG). Fed Proc. 1966;25:317. 20. Johnson AJ, Newman J, Howell MB, Puszkin S. Purification of antihemophilic factor (AHF) for clinical and experimental use. Thromb Diath Haemorrh Suppl. 1967;26:377-381. 21. Ratnoff OD, Kass L, Lang PD. Studies of the purification of antihemophilic factor (factor VIII), II: separation of partially purified antihemophilic factor by gel filtration of plasma. J Clin Invest. 1969; 48:957-962. 22. van Mourik JA, Mochtar IA. Purification of antihemophilic factor (factor VIII) by gel chromatography. Biochim Biophys Acta. 1970; 221:677-679. 23. Zimmerman TS, Ratnoff OD, Powell AE. Immunologic differentiation of classic hemophilia (factor VIII deficiency) and von Willebrand’s disease. With observations on combined deficiencies of antihemophilic factor and proaccelerin (factor V) and on an acquired circulating anticoagulant against antihemophilic factor. J Clin Invest. 1971;50:244-254. 24. Marchesi SL, Shulman NR, Gralnick HR. Studies on the purification and characterization of human factor VIII. J Clin Invest. 1972; 51:2151-2161. 25. Legaz ME, Schmer G, Counts RB, Davie EW. Isolation and characterization of human factor VIII (antihemophilic factor). J Biol Chem. 1973;248:3946-3955. 26. van Mourik JA, Bouma BN, LaBruyère WT, De Graaf S, Mochtar IA. Factor VIII, a series of homologous oligomers and a complex of two proteins. Thromb Res. 1974;4:155-164. 27. Owen WG, Wagner RH. Antihemophilic factor: separation of an active fragment following dissociation by salts or detergents. Thromb Diath Haemorrh. 1972;27/3:502-515. 28. Bouma BN, Wiegerink Y, Sixma JJ, van Mourik JA, Mochtar IA. Immunologic characterization of purified antihaemophilic factor A (factor VIII) which corrects abnormal platelet retention in von Willebrand’s disease. Nature. 1972;236:104-106. 29. Weiss HJ, Hoyer LW. von Willebrand factor: dissociation from antihemophilic factor procoagulant activity. Science. 1973;182: 1149-1151.
vWF-Cleaving Protease and Upshaw-Schulman Syndrome 30. Cooper HA, Griggs TR, Wagner RH. Factor VIII recombination after dissociation by CaCl2. Proc Natl Acad Sci U S A. 1973;70: 2326-2329. 31. Rick ME, Hoyer LW. Immunologic studies of antihemophilic factor (AHF, Factor VIII). V. Immunologic properties of AHF subunits produced by salt dissociation. Blood. 1973;42:737-747. 32. Meyer D, Jenkins CSO, Dreyfus MD, Fressinaud E, Larrieu M-J. von Willebrand factor and ristocetin, II: relationship between von Willebrand factor, von Willebrand antigen and factor VIII activity. Br J Haematol. 1974;28:579-599. 33. Hougie C, Seargeant RB, Brown JE, Baugh RF. Evidence that factor VIII and the ristocetin aggregating factor (VIII Rist) are separate molecular entities. Proc Soc Exp Biol Med. 1974;147:58-61. 34. McKee PA, Andersen JC, Switzer ME. Molecular structural studies of human factor VIII. Ann N Y Acad Sci. 1974;240:8-33. 35. Toole JJ, Knopf JL, Wozney JM, et al. Molecular cloning of cDNA encoding human antihemophilic factor. Nature. 1984;312:342-347. 36. Vehar GA, Keyt B, Eaton D, et al. Structure of human factor VIII. Nature. 1984;312:337-342. 37. Verweij CL, de Vries CJM, Distel B, et al. Construction of cDNA coding for human von Willebrand factor using antibody probes for colony-screening and mapping of the chromosomal gene. Nucleic Acids Res. 1985;13:4699-4717. 38. Sadler JE, Shelton-Inlose BB, Sorace JM, Harlan JM, Titani K, Davie EW. Cloning and characterization of two cDNAs coding for human von Willebrand factor. Proc Natl Acad Sci U S A. 1985;82: 6394-6398. 39. Ginsburg D, Handin RI, Bonthron DT, et al. Human von Willebrand factor (vWF): isolation of complementary DNA (cDNA) clones and chromosome localization. Science. 1985;228: 1401-1406. 40. Shelton-Inlose BB, Titani K, Sadler JE. cDNA sequences for human von Willebrand factor reveal five types of repeated domains and five possible protein sequence polymorphisms. Biochemistry. 1986;25:3164-3171. 41. Kaufman RJ. Biological regulation of factor VIII activity. Annu Rev Med. 1992;43:325-339. 42. Ruggeri ZM, Zimmerman TS. Variant von Willebrand’s disease: characterization of two subtypes by analysis of multimeric composition of factor VIII/von Willebrand factor in plasma and platelets. J Clin Invest. 1980;65, 1318-1325. 43. Ruggeri ZM, Zimmerman TS. The complex multimeric composition of factor VIII/von Willebrand factor. Blood. 1981;57: 1140-1143. 44. Chopek MW, Girma J-P, Fujikawa K, Davie EW, Titani K. Human von Willebrand factor: a multivalent protein composed of identical subunits. Biochemistry. 1986;25:3146-3155. 45. Girma J-P, Chopek MW, Titani K, Davie EW. Limited proteolysis of human von Willebrand factor by Staphylococcus aureus V-8: isolation and partial characterization of a platelet-binding domain. Biochemistry. 1986;25:3156-3163. 46. Titani K, Kumar S, Takio K, et al. Amino acid sequence of human von Willebrand factor. Biochemistry. 1986;25:3171-3184. 47. Wagner DD, Lawrence SO, Ohlsson-Wilheim BM, Fay PJ, Marder VJ. Topology and order of formation of interchain disulfide bonds in von Willebrand factor. Blood. 1987;69:27-32. 48. Ruggeri ZM, Pareti FI, Mannucci PM, Ciavarella N, Zimmerman TS. Heightened interaction between platelets and factor VIII/ von Willebrand factor in a new subtype of von Willebrand disease. N Engl J Med. 1980;302:1047-1051. 49. Dent JA, Berkowitz SD, Ware J, Kasper CK, Ruggeri ZM. Identification of a cleavage site directing the immunochemical detection of molecular abnormality in type IIA von Willebrand factor. Proc Natl Acad Sci U S A. 1990;87:6306-6310. 50. Moake JL, Turner NA, Stathopoulos NA, Nolasco LH, Hellums JD. Involvement of large plasma von Willebrand factor (vWF) multimers and unusually large vWF forms derived from endothelial cells in shear stress-induced platelet aggregation. J Clin Invest. 1986;78:1456-1461.
33
51. Moake JL, Turner NA, Stathopoulos NA, Nolasco L, Hellums JD. Shear-dependent platelet aggregation can be mediated by vWF released from platelets, as well as by exogenous large or unusually large vWF multimers, requires adenosine diphosphate, and is resistant to aspirin. Blood. 1988;71:1366-1374. 52. Ikeda Y, Handa M, Kawano K, et al. The role of von Willebrand factor and fibrinogen in platelet aggregation under varying shear stress. J Clin Invest. 1991;87:1234-1240. 53. Alevriadou BR, Moake JL, Turner NA, et al. Real-time analysis of shear-dependent thrombus formation and its blockade by inhibitors of von Willebrand factor binding to platelets. Blood. 1993;81:1263-1276. 54. Siedlecki CA, Lestini BJ, Kottke-Marchant K, Eppell SJ, Wilson DL. Shear-dependent changes in the 3-dimensional structure of human von Willebrand factor. Blood. 1996;88:2939-2950. 55. Tsai H-M, Sussman II, Nagel RL. Shear stress enhances the proteolysis of von Willebrand factor in normal plasma. Blood. 1994;83: 2171-2179. 56. Gerritsen HE, Turecek PL, Schwarz HP, Lämmle B, Furlan M. Assay of von Willebrand factor (vWF)-cleaving protease based on decreased collagen binding affinity of degraded vWF. A tool for the diagnosis of thrombotic thrombocytopenic purpura (TTP). Thromb Haemost. 1999;82:1386-1389. 57. Obert B, Tout H, Veyradier A, Fressinaud E, Meyer D, Girma J-P. Estimation of the von Willebrand factor-cleaving protease in plasma using monoclonal antibodies to vWF. Thromb Haemost. 1999;82:1382-1385. 58. Kasper CK, Aledort LM, Counts RB, et al. A more uniform measurement of factor VIII inhibitors. Thromb Diath Haemorrh. 1975; 34:869-872. 59. Ashida A, Nakakura H, Matsumoto M, et al. An infant of TTP with a high titer inhibitor to von Willebrand factor-cleaving protease activity [abstract]. Acta Paediatr Jap. 2001;105:293. 60. Tsai HM, Li A, Rock G. Inhibitors of von Willebrand factorcleaving protease in thrombotic thrombocytopenic purpura. Clin Lab. 2001;46:387-392. 61. Tsai H-M. High titers of inhibitor of von Willebrand factor-cleaving metalloproteinase in a fatal case of acute thrombotic thrombocytopenic purpura. Am J Hematol. 2001;65:251-255. 62. Bell WR. Thrombotic thrombocytopenic purpura/hemolytic uremic syndrome relapse: frequency, pathogenesis, and meaning. Semin Hematol. 1997;34:134-139. 63. Kakishita E. Pathophysiology and treatment of thrombotic thrombocytopenic purpura/hemolytic uremic syndrome (TTP/HUS). Int J Hematol. 2000;71:320-327. 64. Cines DB, Konkle BA, Furlan M. Thrombotic thrombocytopenic purpura: a paradigm shift? Thromb Haemost. 2000;84:528-535. 65. Yagi H, Narita N, Matsumoto N, et al. Enhanced low shear stress induced platelet aggregation by Shiga-like toxin 1 purified from Escherichia coli O157. Am J Hematol. 2001;66:105-115. 66. van der Plas RM, Schiphorst ME, Huizinga EG, et al. von Willebrand factor proteolysis is deficient in classic, but not in bone marrow transplantation-associated, thrombotic thrombocytopenic purpura. Blood. 1999;93:3798-3802. 67. Canciani MT, Forza I, Lattuada A, Rossi E, Mannucci PM. von Willebrand factor (VWF) cleaving protease in health and disease [abstract]. Thromb Haemost Suppl. 2001. Abstract 1668. 68. Takahashi Y, Kawaguchi C, Hanesaka Y, Fujimura Y, Yoshioka A. Plasma von Willebrand factor-cleaving protease is low in the newborns. Thromb Haemost Suppl. 2001;Abs 285. 69. Raife TJ, Atkinson BS, Montgomery RR. Comparative von Willebrand factor cleaving proteolytic activity in human saliva, serum and plasma [abstract]. Blood. 2000;96: 2705. 70. Fujikawa K, Suzuki H, McMullen B, Chung D. Purification of human vWF-cleaving protease and its identification as a new member of the metalloproteinase family. Blood. 2001;98:1662-1666. 71. Wolfsberg TG, Straight PD, Gerena RL, et al. ADAM, a widely distributed and developmentally regulated gene family encoding
34
72.
73.
74.
75.
76. 77. 78. 79.
80.
81.
82.
83.
84.
85.
86.
87.
Fujimura et al / International Journal of Hematology 75 (2002) 25-34 membrane proteins with a disintegrin and metalloprotease domain. Develop Biol. 1995;169:378-383. Gerritsen H, Robles R, Lämmle B, Furlan M. Partial amino acid sequence of purified von Willebrand factor-cleaving protease. Blood. 2001;98:1654-1661. Matsumoto M, Chisuwa H, Nakazawa Y, et al. Living-related liver transplantation rescues reduced vWF-cleaving protease activity in patients with cirrhotic biliary atresia [abstract]. Blood. 2000;96:636a. Zheng X, Chung D, Takayama TH, Majerus EM, Sadler JE, Fujikawa K. Structure of von Willebrand factor cleaving protease (ADMTS13), a metalloprotease involved in thrombotic thrombocytopenic purpura. J Biol Chem. 2001;276:41059-41063. Soejima K, Mimura N, Hirashima M, et al. A novel human metalloprotease synthesized in the liver and secreted into the blood: possibly, the von Willebrand factor-cleaving protease? J Biochem. 2001;130:475-480. Dacie JV, Mollison PL, Richardson N, Selwyn JG, Shapiro L. Atypical congenital haemolytic anaemia. Q J Med. 1953;85:79-98. Monnens LAH, Retera RJM. Thrombotic thrombocytopenic purpura in a neonatal infant. J Pediatr. 1967;71:118-123. Wallace DC, Lovric A, Clubb JS, Carseldine DB. Thrombotic thrombocytopenic purpura in four siblings. Am J Med. 1975;58:724-734. Schulman I, Pierce M, Likens A, Currimbhoy Z. Studies on thrombopoiesis, I: a factor in normal human plasma required for platelet production; chronic thrombocytopenia due to its deficiency. Blood. 1960;14:943-957. Upshaw JD. Congenital deficiency of a factor in normal plasma that reverses microangiopathic hemolysis and thrombocytopenia. N Engl J Med. 1978;298:1350-1352. Rennard S, Abe S. Decreased cold-insoluble globulin in congenital thrombocytopenia (Upshaw-Schulman syndrome). N Engl J Med. 1979;300:368. Koizumi S, Miura M, Yamagami M, Horita N, Taniguchi N, Migita S. Upshaw-Schulman syndrome and fibronectin (cold-insoluble globulin). N Engl J Med. 1981;305:1284-1285. Goodnough LT, Saito H, Ratnoff OD. Fibronectin levels in congenital thrombocytopenia: Schulman’s syndrome. N Engl J Med. 1982;306:938-939. Shinohara T, Miyamura S, Suzuki E, Kobayashi K. Congenital microangiopathic hemolytic anemia: report of a Japanese girl. Eur J Pediatr. 1982;138:191-193. Moake JL, Rudy C K, Troll J H, et al. Unusually large plasma factor VIII: von Willebrand factor multimers in chronic relapsing thrombotic thrombocytopenic purpura. N Engl J Med. 1982;307: 1432-1435. Moake JL, Byrnes JJ, Troll JH, et al. Effects of fresh-frozen plasma and its cryosupernatant fraction on von Willebrand factor multimeric forms in chronic relapsing thrombotic thrombocytopenic purpura. Blood. 1985;65:1232-1236. Miura M, Koizumi S, Nakamura K, et al. Efficiency of several plasma components in a young boy with chronic thrombocytopenia and hemolytic anemia who responds repeatedly to normal plasma infusions. Am J Hematol. 1984;17:307-319.
88. Hara T, Kitano A, Kajiwara T, Kondo T, Sakai K, Hamasaki Y. Factor VIII concentrate-responsive thrombocytopenia, hemolytic anemia, and nephropathy. Evidence that factor VIII:von Willebrand factor is involved in its pathogenesis. Am J Ped Hematol Oncol. 1986;8:324-328. 89. Karpman D, Holmberg L, Jirgard L, Lethagen S. Increased platelet retention in familial recurrent thrombotic thrombocytopenic purpura. Kidney Int. 1996;49:190-199. 90. Miura M, Koizumi S, Miyazaki H. Thrombopoietin in UpshawSchulman syndrome. Blood. 1997;89:4663-4664. 91. Yagi H, Konno M, Kinoshita S, et al. Plasma of patients with Upshaw-Schulman syndrome, a congenital deficiency of von Willebrand factor-cleaving protease, enhances the aggregation of normal platelets under high shear stress. Br J Haematol. 2001;115:991997. 92. Moake JL, Turner NA, Stathopoulas NA, Nolasco L, Hellums JD. Shear-induced platelet aggregation can be mediated by vWF release from platelets, as well as by exogenous large or unusually large vWF multimers, requires adenosine diphosphate, and its resistant to aspirin. Blood. 1988;71:1366-1374. 93. Chow TW, Hellums JD, Moake JL, Kroll M. Shear stress-induced von Willebrand factor binding to platelet glycoprotein Ib initiates calcium influx associated with aggregation. Blood. 1992;80: 113-120. 94. Makita K, Shimoyama T, Sakurai Y, et al. Placental ecto-ATP diphosphohydrolase: its structural feature distinct from CD39, localization and inhibition on shear-induced platelet aggregation. Int J Hematol. 1998;68:297-310. 95. Gachet C. ADP receptors of platelets and their inhibition. Thromb Haemost. 2001;86:222-232. 96. Feldman JD, Mardiney MR, Uranue ER, Cutting H. The vascular pathology of thrombotic thrombocytopenic purpura. An immunohistochemical and ultrastructural study. Lab Invest. 1966;15: 927-946. 97. Neame PB, Lechago J, Ling ET, Koval A. Thrombotic thrombocytopenic purpura: report of a case with disseminated intravascular platelet aggregation. Blood. 1973;42:805-814. 98. Asada Y, Sumiyoshi A, Hayashi T, Suzumiya J, Kaketani K. Immunohistochemistry of vascular lesion in thrombotic thrombocytopenic purpura, with special reference to factor VIII related antigen. Thromb Res. 1985;38:469-479. 99. Bull BS, Kuhn IH. The production of schistocytes by fibrin strands (a scanning electron microscope study). Blood. 1970;35:104-111. 100. Konno M, Yoshioka A, Takase T, Imai T. Partial clinical improvement in Upshaw-Schulman syndrome following prostacyclin infusion. Acta Paediat Jap. 1995;37:97-100. 101. Xie L, Chesterman CN, Hogg PJ. Reduction of von Willebrand factor by endothelial cells. Thromb Haemost. 2000;84:506-513. 102. Xie L, Dai CN, Chesterman CN, Hogg PJ. Thrombospondin-1 controls the haemostatic activity of von Willebrand factor. Thromb Haemost. 2001;Abs OC85.