Portal de Periódicos da CAPES

Anti-factor V auto-antibody in the plasma and platelets of a patient with repeated gastrointestinal bleeding

2003; Elsevier BV; Volume: 1; Issue: 5 Linguagem: Inglês

10.1046/j.1538-7836.2003.00143.x

1538-7933

Éva Ajzner, István Balogh, Gizella Haramura, Z Boda, Kalmár Nagy K, G. Pfliegler, Björn Dahlbäck, L. Muszbek,

Platelet Disorders and Treatments

Journal of Thrombosis and HaemostasisVolume 1, Issue 5 p. 943-949 Free Access Anti-factor V auto-antibody in the plasma and platelets of a patient with repeated gastrointestinal bleeding É. Ajzner, É. Ajzner Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this authorI. Balogh, I. Balogh Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this authorG. Haramura, G. Haramura Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this authorZ. Boda, Z. Boda Second Department of Medicine, University of Debrecen, Medical and Health Science Center, Debrecen;Search for more papers by this authorK. Kalmár, K. Kalmár Central Laboratory, Hospital of Nógrád County, Salgótarján, Hungary; andSearch for more papers by this authorG. Pfliegler, G. Pfliegler Second Department of Medicine, University of Debrecen, Medical and Health Science Center, Debrecen;Search for more papers by this authorB. Dahlbäck, B. Dahlbäck Department of Clinical Chemistry, Division of Laboratory Medicine, Lund University, The Wallenberg Laboratory, MAS, Malmö, SwedenSearch for more papers by this authorL. Muszbek, L. Muszbek Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this author É. Ajzner, É. Ajzner Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this authorI. Balogh, I. Balogh Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this authorG. Haramura, G. Haramura Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this authorZ. Boda, Z. Boda Second Department of Medicine, University of Debrecen, Medical and Health Science Center, Debrecen;Search for more papers by this authorK. Kalmár, K. Kalmár Central Laboratory, Hospital of Nógrád County, Salgótarján, Hungary; andSearch for more papers by this authorG. Pfliegler, G. Pfliegler Second Department of Medicine, University of Debrecen, Medical and Health Science Center, Debrecen;Search for more papers by this authorB. Dahlbäck, B. Dahlbäck Department of Clinical Chemistry, Division of Laboratory Medicine, Lund University, The Wallenberg Laboratory, MAS, Malmö, SwedenSearch for more papers by this authorL. Muszbek, L. Muszbek Department of Clinical Biochemistry and Molecular Pathology andSearch for more papers by this author First published: 25 April 2003 https://doi.org/10.1046/j.1538-7836.2003.00143.xCitations: 26 László Muszbek, University of Debrecen, Medical and Health Science Center, Department of Clinical Biochemistry and Molecular Pathology, PO. Box: 40, Debrecen 4012, Hungary. Tel.: +36 52431956; fax: +36 52417631; e-mail: muszbek@jaguar.dote.hu AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat Abstract Summary. Development of autoantibody against coagulation factor V (FV) is a rare clinical condition with hemorrhagic complications of varying severity. The aim of this study was to establish the pathomechanism of an acquired FV deficiency and characterize the FV inhibitor responsible for the clinical symptoms. A 78-year-old female was admitted to hospital with severe gastrointestinal bleeding. General clotting tests and determination of clotting factors were performed by standard methods. FV antigen and FV containing immune complexes were measured by ELISA. The FV molecule was investigated by Western blotting and by sequencing the f5 gene. The binding of patient's IgG to FV and activated FV (FVa) was demonstrated in an ELISA system and its effect on the procoagulant activity of FVa was tested in clotting tests and in a chromogenic prothrombinase assay. Localization of the epitope for the antibody was performed by blocking ELISA. FV activity was severely suppressed both in plasma and platelets. FV antigen levels were normal by ELISA using polyclonal anti-FV antibody or monoclonal antibody against the connecting region of FV, but depressed when HV1 monoclonal antibody against the C2 domain in the FV light-chain was used as capture antibody. The FV molecule was found intact. An IgG reacting with both FV and FVa was present in the patient's plasma and its binding to FV was inhibited by HV1 antibody. FV-containing immune complexes were detected in the patient's plasma and platelet lysate. The patient's IgG inhibited the procoagulant function of FVa. An anti-FV IgG was present in the patient's plasma and platelets. The autoantibody reacted with an epitope in the C2 domain of FV light chain and neutralized the procoagulant function of FVa. Human coagulation factor V (FV) is a single-chain glycoprotein with a Mr of 330 kDa. Its plasma concentration is approximately 20 nmol L−1 (7 µg mL−1) [1]. Approximately 25% of whole blood FV is compartmentalized in the α-granules of platelets and becomes secreted during activation [1, 2]. The principal site of FV biosynthesis is the liver [3, 4], but it is still unclear whether platelet FV originates from the uptake of exogenous FV via endocytosis by megakaryocytes or megakaryocytes themselves can synthesize FV [5-7]. Single chain FV has a structure consisting of three homologous A domains and two homologous C domains connected by a heavily glycosylated B domain in the order of A1-A2-B-A3-C1-C2 [8, 9]. The gene for human FV has been localized to chromosome 1q21–25, it spans approximately 80 kilobases of DNA and consists of 25 exons and 24 introns [10]. Proteolytic cleavage of FV by α-thrombin results in the removal of B-domain and converts the pro-cofactor into active cofactor (FVa) that enhances factor Xa (FXa) − catalyzed activation of prothrombin by several thousand-fold [11, 12]. FVa contains a heavy chain (HC; Mr 105 kDa) and a light chain (LC; Mr 71 or 74 kDa) associated via calcium bond [13]. The HC is encoded by exons 1–12, the entire B domain by exon 13, and the LC by exons 14–25 [10]. FV deficiency is a rare bleeding disorder; the incidence of its congenital form is about 1 in 106. Patients with inherited deficiency suffer moderate to severe hemorrhagic diathesis. Acquired inhibitors to FV are also infrequent; however, FV alloantibodies are more common in patients exposed to topical bovine thrombin containing bovine FV [14, 15]. Alloantibodies can also develop in FV-deficient patients after treatment with fresh frozen plasma. FV autoantibodies were detected in patients following surgery, blood transfusions or antibiotic administration [14, 15]. The clinical symptoms of patients with FV inhibitor vary to a great extent. Patients are often asymptomatic or have mild to severe hemorrhagic manifestations, there are also reports of thrombotic complications [14, 16]. The effect of anti-FV antibodies on platelet FV has been investigated only in a few studies. FV of platelets appears relatively inaccessible to the antibody [17], and un-stimulated, but not stimulated platelets could provide, at least partial, protection against the inhibitor [17-20]. In this study, we demonstrated the presence of a persistent anti-FVLC antibody in a patient with chronic gastrointestinal bleeding symptoms. FV–anti-FV IgG immune complexes were detected both in the plasma and in platelets. The antibody also reacted with FVa and neutralized its procoagulant activity. Materials and methods Materials Human α-thrombin was from Haematologic Technologies (Essex Junction, VT, USA). Human prothrombin, human FXa were from Enzyme Research Laboratories (South Bend, IN, USA). The chromogenic thrombin substrate S-2238 was from Chromogenix (Milano, Italy). Phosphatidylserine (brain extract) and phosphatidylcholine (egg extract) were purchased from Avanti Polar Lipids (Birmingham, AL, USA). Recombinant FV and FV purified from human plasma were produced as previously described [13, 21]. Patient A 78-year-old female with melena and severe anemia (at admittance hemoglobin concentration: 64 g L−1 (0.99 mmol L−1), erythrocyte count: 2.09 × 1012 L−1) was admitted to hospital. She had undergone oophorohysterectomy 3 years before the apparent admission, when she had received fresh frozen plasma and red blood cell (RBC) concentrate because of intra- and postoperative bleeding. There was no known history of hemorrhagic diseases in her family. Gastrointestinal investigations revealed atrophic antrum gastritis and bile reflux with cholelithiasis, but there was no apparent anatomical reason for the bleeding. Beside substitution therapy with fresh frozen plasma, platelet and RBC concentrates, the patient was treated with high dose human immunoglobulin, plasmapheresis and immunosuppressive therapy (combination of cyclosporine A and cyclophosphamide). During the following 3 years the patient had several relapses and had to be hospitalized on four different occasions. Preparation of plasma and platelet suspension Ethical approval was obtained from the Ethics Committee of the Medical and Health Science Center, University of Debrecen. Following informed consent, blood samples from the patient and controls were collected and washed platelet suspension was prepared as described earlier [22, 23]. Platelet count in the final platelet suspension was set to 1012 L−1, and platelets were lyzed in 1% Triton X-100. Coagulation tests Prothrombin time (PT), activated partial thromboplastin time (APTT) and thrombin time (TT) determinations were performed on STA-Compact coagulometer (Diagnostica Stago, Asnières, France) using Innovin (Dade Behring, Marburg, Germany), PTTautomate (Diagnostica Stago) and Thrombin Reagent (Reanal, Budapest, Hungary), respectively. FV coagulant activity in plasma and platelet lysate samples was measured by one-stage clotting assay based on prothrombin time. Measurement of the inhibitor to FV was attempted by a modified, PT-based Bethesda method [24]. The presence of lupus type inhibitor was tested by lupus sensitive APTT (PTT-LA, Diagnostica Stago) hexagonal phospholipid (Staclot LA, Diagnostica Stago) and diluted Russel viper venom (DVVT, DVVT-Confirm tests, American Diagnostica, Greenwich, CT, USA) assays. Determination and characterization of FV by ELISA, Western blotting and DNA sequencing FV, FVHC and FVLC antigen (Ag) levels were determined by sandwich ELISA [22]. Standard human plasma (Dade Behring) was used for the calibration of plasma assays, while in platelet FV activity and antigen assays platelet lysate from healthy donors was used as calibrator. Immunoprecipitation of FV, Western blotting, polymerase chain reaction amplification and DNA sequencing were performed as described earlier [22]. The oligonucleotide primers designed by van Wijk et al. were used for the amplification of the promoter region of f5 gene [25]. The binding of patient's IgG to factors V and Va The total IgG fraction was isolated from patient and control plasma on Protein G Sepharose column (Amersham Pharmacia Biotech) using MabTrap Kit (Amersham Pharmacia Biotech). A solid-phase ELISA was developed to detect the binding of anti-FV IgG to purified FV and FVa. FVa was produced from purified FV by activation with 1 U mL−1 thrombin at 37 °C. After 10-min incubation thrombin was inhibited by Pefablock TH (Kordia, Leiden, the Netherlands). Serial dilutions of plasma and isolated IgG preparations were added to 125 ng of purified FV or FVa coated to polystyrene plates. Bound IgG was detected by peroxidase labeled rabbit antihuman IgG (Dako). Mapping the epitope for the anti-FV auto-antibody A blocking ELISA was developed for mapping the epitope of the autoantibody on FV. Monoclonal antibodies having epitopes on the heavy chain (AHV5146, Haematologic Technologies), on the B-domain (Mk1) [26], on the C2 domain of the light chain (HV1) and on a non-characterized region of the light chain (Mk7) [26] were incubated with FV coated to polystyrene microplates for 1 h. After washing, serial dilutions of patient's IgG were added. The amount of patient's IgG bound to FV was determined by peroxidase labeled antihuman IgG. The effect of patient's IgG on the activity of FVa Various amounts of the patient's IgG were added to pooled normal plasma, and then PT, APTT and TT were measured. In control experiments, IgG isolated from healthy donors was replaced for the patient's IgG. The results were compared with PT, APTT and TT measured on mixtures of patient plasma and control plasma pool. The effect of patient's IgG on the activity of FVa was tested using a slight modification of the prothrombinase-based FVa assay described by Norstrøm et al. [21]. One nmol L−1 recombinant FV was activated by 0.5 U mL−1 human thrombin for 10 min at 37 °C. Thirty-microliter aliquots of FVa were incubated with 3 µL of various dilutions of control and patient IgG preparation for 15 min at 37 °C and the remaining FVa activity was determined. The FVa containing samples and 5 nmol L−1 FXa were added to a prothrombinase mixture containing 0.5 µmol L−1 prothrombin and 50 µmol L−1 phospholipid vesicles (10 : 90 w/w phosphatidylserine/phosphatidylcholine). After 2-min incubation, prothrombin activation was stopped by 40-fold dilution in ice-cold EDTA buffer. The amount of thrombin formed in the presence and absence of IgG preparations was measured kinetically with the chromogenic substrate, S2238 and the results were expressed as residual FVa activity. Detection of FV–anti-FV immune complexes in the patient's plasma and platelet lysate An ELISA was developed for the detection of FV specific immune complexes in plasma and platelet lysate. Sheep antihuman FV polyclonal antibody (The Binding Site) was used as capture antibody and peroxidase labeled antihuman IgG (Sigma) recognizing the Fc-portion of human IgG was used as detection antibody. Other methods Protein concentration was determined by BCA Protein Assay Reagent kit (Perbio Science, Erembodegem-Aalst, Belgium), using bovine IgG as calibrator. Results demonstrated on the figures are representatives of at least three parallel experiments not deviating from each other by more than 5%. Results Demonstration and characterization of the acquired FV deficiency of the patient The PT and APTT of the patient were significantly prolonged on admission (Table 1) and remained prolonged during the following 3 years. TT was always normal (not shown). In mixing studies, a concentration-dependent correction of the prolonged clotting times were observed. However, even in the 1 : 1 mixture, the normal pooled plasma failed to correct the prolongation of PT and APTT completely. Incubation of the mixture for 1 h at 37 °C did not change the situation. Tests for lupus anticoagulant were negative. Table 1. Prothrombin times and activated partial thromboplastin times of the patient PT (s) PT (s) after 1 h at 37 °C APTT (s) APTT (s) after 1 h at 37 °C Patient plasma 81.3 82.6 147.2 165.7 Pooled normal plasma 9.3 9.3 35.1 36.4 Mixtures of patient:normal plasma 9 : 1 48.8 ND 111.4 ND 4 : 1 31.6 ND 105.2 ND 2 : 1 23.6 ND 78.5 ND 1 : 1 16.1 15.9 50.7 52.5 Reference interval 8.7–11.5 NA 29.5–42.7 NA ND, not determined; NA, not applicable. With the exception of FV, the activities of clotting factors in the patient's plasma were within normal range. FV activity was severely decreased both in the plasma (0.48%; reference interval: 80–120%) and in the platelet lysate (1.12% of normal average) of the patient. On admission, 2.0 BU FV inhibitor was measured by the modified Bethesda assay, and the antibody titer remained low (1.25–2.0 BU) during the 3-year follow-up period. Titration of purified FV into patient plasma resulted in normalization of PT and APTT at 7.5–10 µg mL−1 supplemented FV concentration (Table 2). Although bleeding and melena stopped following therapy, the main coagulation parameters remained essentially unchanged. Table 2. Correction of prolonged PT and APTT of the patient's plasma by the addition of purified FV Added FV in the plasma (µg mL−1) PT (s) APTT (s) 0 65.5 141.8 1 52.6 119.1 2 39.1 92.8 3 33.0 77.6 4 18.0 50.5 5 13.6 41.4 7.5 11.1 38.8 10 10.9 37.8 In the patient's plasma, total FV:Ag and FVHC:Ag levels were normal (106% and 139%, respectively), while FVLC:Ag was severely decreased (7.9%). Similarly, FV:Ag was normal (80%), and FVLC:Ag was low (4.8%) in platelets. The decrease of FVLC:Ag, however, was not due to the presence of a truncated molecule, since FV immunoprecipitated from the patient's plasma showed a single FV specific band of ∼330 kDa on the immunoblot that comigrated with the intact FV molecule in 5% SDS–polyacrylamide gel (not demonstrated). The DNA sequence analysis of the whole coding and promoter region and part of the non-coding region of f5 gene did not identify any disease-causing mutations. A few known polymorphisms [25, 27] and a so far unidentified nucleotide change in exon 13 were detected in heterozygous form (Table 3). The patient was heterozygous for R506Q FV (Leiden) mutation [28]. Table 3. Mutations or polymorphisms detected in the patient's f5 gene Localization Nucleotide position Nucleotide original Nucleotide variant Amino acid replacement Reference Promoter −426 g a NA 25 Exon 3 409 g c D79H 25 Exon 4 495 g a Silent 25 641 g t Silent 25 Intron 5 −262 t c NA 25 −195 g a NA 25 (−160)–(−157) ins t NA 25 Exon 10 1691 g a R506Q 28 Exon 13 2925 g a Silent This study 3943 c a L1257I 27 Intron 16 +12 a g NA 25 NA, not applicable. FV-binding auto-antibody in the patient's IgG fraction The presence of FV-specific autoantibody was demonstrated in the patient's plasma and in its IgG fraction by ELISA technique (Fig. 1). The antibody also recognized FVa, although the OD signals measured in FVa-specific ELISA was lower than that obtained in FV-specific ELISA. This might be related to the loss of some antigenicity during thrombin activation of FV. Figure 1Open in figure viewerPowerPoint Binding of the inhibitor to factor V and activated factor V. Human purified FV (squares with solid line) or FVa (triangles with dotted line) were coated to microtiter plate and incubated with serial dilutions of IgG preparation (a) or plasma (b) from the patient (solid symbols) or from controls (open symbols). IgG bound to FV or FVa was detected by peroxidase labeled antihuman IgG. The actual IgG concentration in the dilutions of plasma and IgG preparations is shown on the abscissa. Figure 2 shows that both the plasma and the platelet lysate contained immune complexes that were captured by FV-specific antibodies and could be detected by an antibody against human IgG. Such FV-IgG immune complexes could be recognized even at 100-fold plasma dilution. Figure 2Open in figure viewerPowerPoint Demonstration of factor V–IgG immune complexes in the patient's plasma and platelet lysate. Sheep antihuman FV polyclonal antibody coated to a microtiter plate was incubated with serial dilutions of plasma (diamond) or platelet lysate (triangle) from the patient (closed symbols) and from normal controls (open symbols). A peroxidase labeled antihuman IgG recognizing the Fc-portion of human IgG was used for the detection of FV-specific immune complexes. A blocking ELISA was used to assess the effect of four anti-FV monoclonal antibodies with known epitopes on the binding of the patient's IgG autoantibodies to FV. Although all four monoclonal antibodies bound to FV, only the HV1 monoclonal antibody directed to a specific region of the C2 domain of FVLC inhibited the binding of the patient's antibody (Fig. 3). Figure 3Open in figure viewerPowerPoint The effect of monoclonal antifactor V antibodies with different epitope specificity on the binding of the patient's auto-antibody to factor V. Purified human FV was coated to microtiter plate and incubated without (closed circle, solid line) or with one of the following antifactor V monoclonal antibodies: AHV5146 antibody against the heavy chain (▴, dotted line), Mk1 monoclonal antibody against B-domain (▵, dotted line), Mk7 antibody against the light chain (○, dotted line), HV1 antibody against the C2 domain of the light chain in a concentration of 0.04 µg mL−1 (□, dashed line) or 2 µg mL−2 (▪, dashed line). After washing, various concentrations of patient's IgG were added. The patient's IgG bound to FV was measured by peroxidase labeled antihuman IgG. Neutralization of the procoagulant activity of FV by the auto-antibody The addition of the patient's IgG to normal plasma prolonged both PT and APTT (Fig. 4b) to an extent similar to the effect of un-fractionated patient's plasma (Fig. 4a), while the addition of normal plasma IgG did not. The inhibitor had no effect on TT. The inhibitory activity of the patient's IgG seemed to be weak because effective inhibitory action became apparent only at IgG concentration above 8 mg mL−1. The data demonstrate that an anti-FV IgG in the patient's plasma was responsible for the prolongation of PT and APTT. Figure 4Open in figure viewerPowerPoint Effect of the inhibitor on prothrombin time, activated partial thromboplastin time and thrombin time of normal pooled plasma. Different dilutions of the patient's plasma (a) or its IgG fraction (b) were mixed with normal plasma and PT (diamond), APTT (circle), TT (triangle) of these mixtures were measured (closed symbols). The IgG preparation from an apparently healthy donor (open symbols) was used as control. Figure 5 demonstrates that increasing concentration of the patient's IgG inhibited the FXa-cofactor function of FVa, while control IgG had no effect. In this system, at a concentration of 1 mg mL−1 IgG the inhibitory effect exceeded 70% and at physiological IgG concentration (∼10 mg mL−1) only 2.5% of the original FV activity remained. Figure 5Open in figure viewerPowerPoint Inhibition of recombinant factor Va activity by the patient's IgG in a prothrombinase-based FVa assay. Recombinant FV was activated by thrombin for 10 min at 37 °C, and then incubated with various concentrations of the patient's IgG (◆) or IgG from a healthy donor (○) for 15 min at 37 °C. Residual FVa activity was then determined by chromogenic prothrombinase assay. Discussion Results of the screening tests and factor determinations clearly indicated that our patient had FV deficiency. Her history suggested the presence of a FV inhibitor. However, the results of mixing studies and the Bethesda assay did not unequivocally exclude the presence of an alloantibody that had developed on the basis of a genetic FV disorder after exposure to foreign plasma proteins. The results of FV antigen determinations raised the possibility of a truncated protein. However, Western blotting of FV immunoprecipitated from the patient's plasma verified that the Mr of the FV molecule was normal and DNA sequencing excluded a genetic FV defect leading to the synthesis of an abnormal molecule. Anti-FV antibodies reacting with both FV and FVa were detected in the patient's plasma and IgG fraction, and functional assays clearly showed that the IgG preparation isolated from the patient's plasma inhibited the procoagulant activity of FVa. The presence of FV–anti-FV IgG immune complexes in the patient's plasma was also demonstrated. These pieces of evidence clearly support an anti-FV autoantibody being responsible for the bleeding diathesis of the patient. The low BU values and the finding that relatively low amount of supplemented FV normalized PT and APTT of the patient's plasma suggest a relatively low titer of the autoantibody. The global clotting assays and the clinical symptoms indicated that, despite of its low concentration, the antibody also exerted its inhibitory effect in vivo. It is very likely that the inhibitor targets a conformational, and not structural epitope because on Western blot, the IgG fraction from the patient's plasma did not recognize FV or FVa treated with SDS under reducing or non-reducing conditions (data not shown). Binding assays using ELISA technique and monoclonal antibodies with known epitopes helped to localize the epitope for the autoantibody within the structure of FV molecule. With polyclonal capture antibody in a sandwich ELISA plasma FV concentration was normal, while using HV1 monoclonal anti-FV antibody against the C2 domain of FVLC [29] as capture antibody the FV antigen level was severely depressed. As the FV molecule was proved to be intact, these results can only be explained by the decreased binding of HV1 monoclonal antibody to FV being in complex with the autoantibody. To strengthen this hypothesis four monoclonal antibodies, including HV1 were tested for blocking the binding of the autoantibody to FV. Two of the antibodies were directed against different epitopes on FVLC and two further antibodies had epitopes on FVHC and on the B domain. Only the HV1 antibody directed against the region of amino acids 2060–69 [29] interfered with the binding of the patient's autoantibody. These findings suggest that the epitopes for HV1 and for the autoantibody are, at least partially, overlapping. Anti-FV antibodies often target the C2 domain of FVLC [30-32]. The N-terminal portion of C2 domain is involved in the binding of FVa to the phospholipid surface [29, 33], the inhibition of this function by the autoantibody well explains its interference with the clotting tests and the prothrombinase assay. To our knowledge, the presence of FV inhibitor in platelets has not been investigated. Washed platelets from our patient contained normal level of FV antigen, while very low FV activity was measured in the platelet lysate. The demonstration of FV–anti-FV immune complexes in the platelet lysate and the low level of FV:Ag measured by an ELISA using anti-FVLC C2 capture antibody indicate that the anti-FV IgG demonstrated in the patient's plasma is also present in her platelets. IgG is endocytosed or pinocytosed by megakaryocytes and to a certain extent by platelets and becomes incorporated into platelet α granules [34]. The α granular IgG pool reflects the concentration and the composition of plasma IgG [35, 36]. Based on these facts, it seems evident that IgG type FV inhibitors can get into platelet alpha granules, in vivo. The ingestion of FV–anti-FV immune complexes by platelets and megakaryocytes and packaging them into alpha granules may represent an alternative mechanism for the uptake of anti-FV IgG. It is not clear if within platelets FV and anti-FV IgG exist in the form of immune complexes or if they are stored in non-complexed form and immune complexes are formed only after lyzing the cell by non-ionic detergent. The dominant clinical symptom of our patient was persistent gastrointestinal bleeding. Gastrointestinal bleeding has been reported among patients with acquired FV inhibitors [14, 15]. The bleeding was successfully stopped by the therapeutic regimen described above but the prolonged clotting tests and low FV activity remained unchanged, and the patient experienced relapses at several occasions. Complete FV deficiency is lethal in knockout mice, but very low FV activity is enough to maintain hemostasis [37, 38]. The correlation of plasma FV activity with the hemorrhagic manifestations is poor and the severity of bleeding symptoms seems to be more closely correlated with platelet rather than with plasma FV levels [18, 19, 39]. The anti-FV antibody was also present in the platelets of our patient and might have effectively interfered with the procoagulant action of platelet FV and contributed to the bleeding symptoms. The patient was heterozygous for the FV Leiden mutation. Thus, half of the small portion of the patient's FV that was activated and not blocked by the autoantibody must have been the FV Leiden molecule that exerts its procoagulant activity longer than its wild type counterpart. It is known that hemophilic patients carrying the FV Leiden mutation have less severe clinical symptoms than those who have identical mutation of the f8 gene but do not carry the FV Leiden allele [40]. The presence of the FV Leiden mutation in our patient with FV autoantibody might also have some moderating effect on the severity of her bleeding diathesis. Acknowledgements We are indebted to Ing-Marie Persson for the purification of human FV. The study was supported by grants from the Higher Education Development Programs of the Ministry of Education, Hungary (FKFP 0214/2001), from the Hungarian National Research Fund (OTKA T030406), from the Hungarian Academy of Sciences (11003) and from the Swedish Science Council (07143). References 1 Tracy PB, Eide LL, Bowie EJ, Mann KJ. Radioimmunoassay of factor V in human plasma and platelets. Blood 1982; 60: 59– 63. CrossrefCASPubMedWeb of Science®Google Scholar 2 Chesney CM, Pifer D, Colman RW. Subcellular localization and secretion of factor V from human platelets. Proc Natl Acad Sci USA 1981; 78: 5180– 4. CrossrefCASPubMedWeb of Science®Google Scholar 3 Owren CA Jr, Bowie EJ. Generation of coagulation factors V, XI, and XII by the isolated rat liver. Haemostasis 1977; 6: 205– 12. PubMedWeb of Science®Google Scholar 4 Wilson DB, Salem HH, Mruk JS, Maruyama I, Majerus PW. Biosynthesis of coagulation factor V by human hepatocellular carcinoma cell line. J Clin Invest 1984; 73: 654– 8. CrossrefCASPubMedWeb of Science®Google Scholar 5 Camire RM, Pollak ES, Kaushansky K, Tracy PB. Secretable human platelet-derived factor V originates from the plasma pool. Blood 1998; 92: 3035– 41. CrossrefCASPubMedWeb of Science®Google Scholar 6 Chui HC, Schick PK, Colman RW. Biosynthesis of factor V in isolated guinea pig megakaryocytes. J Clin Invest 1985; 75: 339– 46. CrossrefPubMedWeb of Science®Google Scholar 7 Gewirtz AM, Keefer M, Doshi K, Annamalai AE, Chiu HC, Colman RW. Biology of human megakaryocyte factor V. Blood 1986; 67: 1639– 48. CASPubMedWeb of Science®Google Scholar 8 Kane WH. Factor V. In: Hemostasis and Thrombosis ( RW Colman, J Hirsh, VJ Marder, AW Clowes, JN George, eds), 4th edn. Philadelphia, PA: Lippincott. Williams & Wilkins, 2001: 157– 69. Web of Science®Google Scholar 9 Rosing J, Tans G. Factor V. Int J Biochem Cell Biol 1997; 29: 1123– 6. CrossrefCASPubMedWeb of Science®Google Scholar 10 Jenny R, Pittman D, Toole J, Kriz RW, Aldape RA, Hewick RM, Kaufman RJ, Mann KG. Complete cDNA and derived amino acid sequence of human FV. Proc Natl Acad Sci USA 1987; 84: 4846– 50. CrossrefCASPubMedWeb of Science®Google Scholar 11 Nesheim ME, Taswell JB, Mann KG. The contribution of bovine factor V and factor Va to the activity of prothrombinase. J Biol Chem 1979; 254: 10952– 62. CrossrefCASPubMedWeb of Science®Google Scholar 12 Rosing J, Tans G, Govers Riemslag JWP, Zwaal RFA, Hemker HC. The role of phospholipids and factor Va in the prothrombinase complex. J Biol Chem 1980; 255: 274– 83. CrossrefCASPubMedWeb of Science®Google Scholar 13 Dahlbäck B. Human coagulation factor V purification and thrombin-catalyzed activation. J Clin Invest 1980; 66: 583– 91. CrossrefCASPubMedWeb of Science®Google Scholar 14 Ortel TL. Clinical and laboratory manifestations of anti-factorV antibodies. J Lab Clin Med 1999; 133: 326– 34. CrossrefCASPubMedWeb of Science®Google Scholar 15 Knobl P, Lechner K. Acquired factor V inhibitors. Baillières Clin Haematol 1998; 11: 305– 18. CrossrefCASPubMedWeb of Science®Google Scholar 16 Kalafatis M, Simioni P, Tormene D, Beck DO, Luni S, Girolami A. Isolation and characterization of an antifactor V antibody causing activated protein C resistance from a patient with severe thrombotic manifestations. Blood 2002; 99: 3985– 92. CrossrefCASPubMedWeb of Science®Google Scholar 17 Nesheim ME, Nichols WL, Cole TL, Houston JG, Schenk RB, Mann KG, Bowie EJ. Isolation and study of an acquired inhibitor of human coagulation factor V. J Clin Invest 1986; 77: 405– 15. CrossrefCASPubMedWeb of Science®Google Scholar 18 Miletich JP, Jackson CM, Majerus PW. Properties of the factor Xa binding site on human platelets. J Biol Chem 1978; 253: 6908– 16. CASPubMedWeb of Science®Google Scholar 19 Miletich JP, Majerus DW, Majerus PW. Patients with congenital factor V deficiency have decreased factor Xa binding sites on their platelets. J Clin Invest 1978; 62: 824– 31. CrossrefCASPubMedWeb of Science®Google Scholar 20 Grigg AP, Dauer R, Thurlow PJ. Bleeding due to an acquired inhibitor of platelet associated factor V. Aust NZ J Med 1989; 19: 310– 4. Wiley Online LibraryCASPubMedWeb of Science®Google Scholar 21 Norstrøm E, Thorelli E, Dahlbäck B. Functional characterisation of recombinant FV Hong Kong and FV Cambridge. Blood 2002; 100: 524– 30. CrossrefCASPubMedWeb of Science®Google Scholar 22 Ajzner E, Balogh I, Szabó T, Marosi A, Haramura G, Muszbek L. Severe coagulation factor V deficiency caused by two novel frameshift mutations: 2952delT in exon 13 and 5493insG in exon 16 of factor 5 gene. Blood 2002; 99: 702– 5. CrossrefCASPubMedWeb of Science®Google Scholar 23 Muszbek L, Polgár J, Boda Z. Platelet factor XIII becomes active without the release of activation peptide during platelet activation. Thromb Haemost 1993; 69: 282– 5. CrossrefCASPubMedWeb of Science®Google Scholar 24 Kasper CK, Aledort LM, Counts RB, Edson JR, Fratantoni J, Green D, Hampton JW, Hilgartner MW, Lazerson J, Levine PH, McMillan CW, Pool JG, Shapiro SS, Shulman NR, Van Eys J. A more uniform measurement of factor VIII inhibitors. Thromb Diath Haemorrh 1975; 34: 612. PubMedWeb of Science®Google Scholar 25 Van Wijk R, Nieuwenhuis K, Van Den Berg M, Huizinga EG, Van Der Meijden BB, Kraaijenhagen RJ, Van Solinge WW. Five novel mutations in the gene for human blood coagulation factor V associated with type I factor V deficiency. Blood 2001; 98: 358– 67. CrossrefCASPubMedWeb of Science®Google Scholar 26 Dahlbäck B, Hildebrand B. Inherited resistance to activated protein C is corrected by anticoagulant cofactor activity found to be a property of factor V. Proc Natl Acad Sci USA 1994; 91: 1396– 400. CrossrefCASPubMedWeb of Science®Google Scholar 27 Kane WH, Ichinose A, Hagen FS, Davie EW. Cloning of cDNAs coding for the heavy chain region and connecting region of human factor V, a blood coagulation factor with four types of internal repeats. Biochemistry 1987; 26: 6508– 14. CrossrefCASPubMedWeb of Science®Google Scholar 28 Bertina RM, Koeleman BP, Koster T, Rosendaal FR, Dirven RJ, De Ronde H, Van Der Velden PA, Reitsma PH. Mutation in blood coagulation factor V associated with resistance to activated protein C. Nature 1994; 369: 64– 7. CrossrefCASPubMedWeb of Science®Google Scholar 29 Kim SW, Quinn-Allen MA, Camp JT, Macedo-Ribeiro S, Fuentes-Prior P, Bode W, Kane WH. Identification of functionally important amino acid residues within the C2-domain of human factor V using alanine-scanning mutagenesis. Biochemistry 2000; 39: 1951– 8. CrossrefCASPubMedWeb of Science®Google Scholar 30 Ortel TL, Quinn-Allen MA, Charles LA, Devore-Carter D, Kane WH. Characterisation of an acquired inhibitor to coagulation factor V. Antibody binding to the second C-type domain of factor V inhibits the binding of factor V to phosphatidylserine and neutralizes procoagulant activity. J Clin Invest 1992; 90: 2340– 7. CrossrefCASPubMedWeb of Science®Google Scholar 31 Ortel TL, Moore KD, Quinn-Allen MA, Okamura T, Sinclair AJ, Lazachick J, Govindan R, Carmagnol F, Kane WH. Inhibitory anti- factor V antibodies bind to the factor V, C2 domain and are associated with haemorrhagic manifestations. Blood 1998: 91: 4188– 96. CASPubMedWeb of Science®Google Scholar 32 Izumi T, Kim SW, Greist A, Macedo-Ribeiro S, Fuentes-Prior P, Bode W, Kane WH, Ortel TL. Fine mapping of inhibitory anti-factorV antibodies using factor V, C2 domain mutants. Identification of two antigenic epitopes involved in phospholipid binding. Thromb Haemost 2001; 85: 1048– 54. CASPubMedWeb of Science®Google Scholar 33 Ortel TL, Devore-Carter D, Quinn-Allen MA, Kane WH. Deletion analysis of recombinant human factor V. Evidence for a phosphatidylserine binding site in the second C-type domain. J Biol Chem 1992; 267: 4189– 98. CrossrefCASPubMedWeb of Science®Google Scholar 34 Harrison P, Cramer EM. Platelet alpha-granules. Blood Rev 1993; 7: 52– 62. CrossrefCASPubMedWeb of Science®Google Scholar 35 George JN. Platelet IgG: measurement, interpretation, and clinical significance. Prog Hemost Thromb 1991; 10: 97– 126. CASPubMedGoogle Scholar 36 George JN, Platelet Ig, G. Its significance for the evaluation of thrombocytopenia and for understanding the origin of α-granule proteins. Blood 1990; 76: 859– 70. PubMedWeb of Science®Google Scholar 37 Cui J, O'Shea KS, Purkayastha A, Saunders TL, Ginsburg D. Fatal haemorrhage and incomplete block to embryogenesis in mice lacking coagulation factor V. Nature 1996; 384: 66– 8. CrossrefCASPubMedWeb of Science®Google Scholar 38 Yang TL, Cui J, Taylor JM, Yang A, Gruber SB, Ginsburg D. Rescue of fatal neonatal hemorrhage in factor V deficient mice by low level transgene expression. Thromb Haemost 2000; 83: 70– 7. CrossrefCASPubMedWeb of Science®Google Scholar 39 Tracy PB, Giles AR, Mann KG, Eide LL, Hoogendoorn H, Rivard GE. Factor V (Quebec): a bleeding diathesis associated with a qualitative platelet factor V deficiency. J Clin Invest 1984; 74: 1221– 8. CrossrefCASPubMedWeb of Science®Google Scholar 40 Nichols WC, Amano K, Cacheris PM, Figueiredo MS, Michaelides K, Schwaab R, Hoyer L, Kaufman RJ, Ginsburg D. Moderation of hemophilia A phenotype by the factor V R506Q mutation. Blood 1996; 88: 1183– 7. CrossrefCASPubMedWeb of Science®Google Scholar Citing Literature Volume1, Issue5May 2003Pages 943-949 FiguresReferencesRelatedInformation