An account of this work was presented at the 48th annual meeting of the American Society of Hematology, December 9, 2006, Orlando, FL.
Protein S down-regulates factor Xase activity independent of activated protein C: specific binding of factor VIII(a) to protein S inhibits interactions with factor IXa
Article first published online: 28 AUG 2008
© 2008 The Authors. Journal Compilation © 2008 Blackwell Publishing Ltd
British Journal of Haematology
Volume 143, Issue 3, pages 409–420, November 2008
How to Cite
Takeyama, M., Nogami, K., Saenko, E. L., Soeda, T., Nishiya, K., Ogiwara, K., Yoshioka, A. and Shima, M. (2008), Protein S down-regulates factor Xase activity independent of activated protein C: specific binding of factor VIII(a) to protein S inhibits interactions with factor IXa. British Journal of Haematology, 143: 409–420. doi: 10.1111/j.1365-2141.2008.07366.x
- Issue published online: 14 OCT 2008
- Article first published online: 28 AUG 2008
- Received 24 May 2008; accepted for publication 7 July 2008
- factor VIII(a);
- protein S;
- factor IXa;
- tenase complex;
Protein S functions as an activated protein C (APC)-independent anticoagulant in the inhibition of intrinsic factor X activation, although the precise mechanisms remain to be fully investigated. In the present study, protein S diminished factor VIIIa/factor IXa-dependent factor X activation, independent of APC, in a functional Xa generation assay. The presence of protein S resulted in an c. 17-fold increase in Km for factor IXa with factor VIIIa in the factor Xase complex, but an c. twofold decrease in Km for factor X. Surface plasmon resonance-based assays showed that factor VIII, particularly the A2 and A3 domains, bound to immobilized protein S (Kd; c. 10 nmol/l). Competition binding assays using Glu-Gly-Arg-active-site modified factor IXa showed that factor IXa inhibited the reaction between protein S and both the A2 and A3 domains. Furthermore, Sodium dodecyl sulphate polyacrylamide gel electrophoresis revealed that the cleavage rate of factor VIIIa at Arg336 by factor IXa was c. 1·8-fold lower in the presence of protein S than in its absence. These data indicate that protein S not only down-regulates factor VIIIa activity as a cofactor of APC, but also directly impairs the assembly of the factor Xase complex, independent of APC, in a competitive interaction between factor IXa and factor VIIIa.
Factor VIII functions as a cofactor for factor IXa in the enzyme complex (factor Xase) responsible for anionic phospholipid surface-dependent conversion of factor X to Xa (Mann et al, 1990). Factor VIII is synthesized as a multi-domain, single chain molecule (A1-A2-B-A3-C1-C2) consisting of 2332 amino acid residues with a molecular mass of c. 300 kDa (Vehar et al, 1984; Wood et al, 1984). Mature factor VIII is processed through a series of metal ion-dependent heterodimers by hydrolysis at the B-A3 junction, generating a heavy chain consisting of the A1 and A2 domains, together with heterogeneous fragments of a partially proteolysed B domain, linked to a light chain consisting of the A3, C1, and C2 domains. Factor VIII circulates as a complex with von Willebrand factor (VWF), which stabilizes the cofactor activity of factor VIII (Lollar et al, 1988). Critical sites in factor VIII for VWF interaction have been localized to the N-terminal acidic region of the A3 domain (Foster et al, 1988) and the C-terminal region of the C2 domain (Saenko et al, 1994).
The procofactor of factor VIII is activated by cleavage at Arg372, Arg740, and Arg1689 by either thrombin or factor Xa, converting the dimer into the factor VIIIa trimer composed of the A1, A2, and A3C1C2 subunits (Eaton et al, 1986). Proteolysis at Arg372 and Arg1689 is essential for generating factor VIIIa cofactor activity. Cleavage at the former site exposes a functional factor IXa-interactive site within the A2 domain that is cryptic in the unactivated molecule (Fay et al, 2001). Cleavage at the latter site liberates the cofactor from its carrier protein, VWF (Regan & Fay, 1995) contributing to the overall specific activity of the cofactor (Donath et al, 1995). On the other hand, factor VIIIa cofactor activity is down-regulated in the presence of serine proteases, such as activated protein C (APC), factor Xa, factor IXa, and plasmin (Eaton et al, 1986; Lamphear & Fay, 1992; Nogami et al, 2007), by proteolytic inactivation following cleavage at Arg336 within A1 subunit.
Protein S is a vitamin K-dependent anticoagulant protein, acting as a cofactor of APC (DiScipio & Davie, 1980) for the proteolytic inactivation of both factor VIIIa and factor Va on phospholipid membranes (Walker, 1980). Protein S increases the affinity of APC for phospholipid vesicles, endothelial cells, and platelets (Walker, 1981; Harris & Esmon, 1985; Stern et al, 1986). Approximately 50% of the protein S in plasma is bound to C4b-binding protein, a regulatory protein of the complement system (Dahlbäck & Stenflo, 1981) and only free protein S contributes to APC activity (Dahlbäck, 1986). The physiological importance of protein S is reflected in the clinical observation that a deficiency of protein S is associated with an enhanced risk of thrombosis (Comp & Esmon, 1984; Comp et al, 1984).
In addition, protein S has been reported to serve as an APC-independent, direct anticoagulant (Heeb et al, 1993; Koppelman et al, 1995). For example, Koppelman et al (1995) demonstrated that protein S exerted an APC-independent anticoagulant function by inhibiting the intrinsic activation of factor X in a mechanism that was possibly mediated by a direct interaction between protein S and factor VIII. This precise mechanism remains to be fully explored, however. The present study demonstrated for the first time that protein S binds to factor VIII(a) and directly moderates the activity of factor Xase by competing with factor IXa in the formation of the enzyme complex.
Materials and methods
Purified recombinant factor VIII preparations (Kogenate FS®) and plasma-derived factor VIII/VWF concentrates (Confact F®) were generous gifts from Bayer Corp. Japan (Osaka, Japan) and Chemo-Sero-Therapeutic Research Inc. (Kumamoto, Japan) respectively. Factor VIIIa, the light and heavy chains of factor VIII, the A1 and A2 subunits were isolated as previously reported (Fay et al, 2001). VWF was purified from factor VIII/VWF concentrates using gel filtration on a Sepharose CL-4B column and immune-beads coated with immobilized anti-factor VIII monoclonal antibody (mAb) (Shima et al, 1992). Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) of the purified protein, followed by staining with GelCode Blue Stain Reagent (Pierce, Rockford, IL, USA), showed >95% purity. Protein concentrations were determined by the method of Bradford. Human protein S, factor IXa, factor X (Hematologic Technologies, Inc., Burlington, VT, USA), factor Xa (American Diagnostica Inc., Greenwich, CT, USA), α-thrombin and hirudin (Sigma, St Louis, MO, USA), chromogenic factor Xa substrate S-2222 (Chromogenix, Milano, Italy), were purchased from the indicated vendors. The active-site of factor IXa was modified with Glu-Gly-Arg-chloromethylketone (EGR-ck; Calbiochem, San Diego, CA, USA) to yield EGR-factor IXa (Lollar & Fass, 1984). An anti-A2 mAb (mAbJR8) was obtained from JR Scientific Inc. (Woodland, CA, USA). mAb58.12, recognizing the N-terminal end of the A1 domain, was a generous gift from Bayer Corp. (Berkeley, CA, USA). Anti-protein S mAb was purchased from Haematologic Technologies, Inc. Biotinylated IgG was prepared using N-hydroxysuccinimido-biotin (Pierce). Phospholipid vesicles containing 10% phosphatidylserine, 60% phosphatidylcholine, and 30% phosphatidylethanolamine (Sigma) were prepared using N-octylglucoside (Mimms et al, 1981).
Expression and purification of recombinant A3 or C2 (rA3 or rC2) domains of the factor VIII light chain
The rA3 domain of factor VIII (residues 1690–2019) was expressed in E. coli using pET expression system (Novagen, Madison, WI, USA). The pMT2/factor VIII plasmid containing B-domain deleted factor VIII gene (Pipe & Kaufman, 1997) was obtained from Dr Pipe (University of Michigan, Ann Arbor, MI, USA). DNA fragments encoding the A3 domain were generated by polymerase chain reaction using pMT2/factor VIII as a template and a pair of corresponding primers. The amplified fragments were ligated into the pET-20b(+) expression vector. Plasmid DNA was purified and the sequence was confirmed by direct sequencing in both directions using the technology of Applied Biosystems (Foster City, CA, USA). The plasmid was used for transformation of Origami(DE3)pLysS E. coli cells (Novagen), the host strain for the protein expression. The protein was expressed and subsequently purified using a His-Select affinity cartridge (Sigma). Proper folding of the rA3 fragment was confirmed by determination of the affinities for conformationally sensitive anti-A3 mAb, CLB-CAgA (Lenting et al, 1996). A cDNA coding the C2 domain sequence of human factor VIII was constructed, transformed into Pichia pastoris cells and expressed in a yeast secretion system as previously described (Takeshima et al, 2003). The rC2 protein was purified by ammonium sulfate fractionation and CM Sepharose chromatography (Amersham Bio-Science) as previously described (Takeshima et al, 2003).
Factor Xa generation assay
The rate of conversion of factor X to factor Xa was monitored in a purified system at 37°C. Factor VIII (30 nmol/l) was activated by thrombin (10 nmol/l) in the presence of phospholipid vesicles (10 μmol/l). Thrombin activity was inhibited after 1 min by the addition of hirudin (2·5 unit/ml), and factor Xa generation was initiated by the addition of factor IXa and factor X at the indicated concentrations. Aliquots were removed at appropriate times to assess the initial rates of product formation and were added to tubes containing EDTA (50 mmol/l final concentration) to quench the reactions. Rates of factor Xa generation were determined by the addition of the chromogenic substrate, S-2222 (0·46 mmol/l final concentration). Reactions were read at 405 nm using a Labsystems Multiskan Multisoft microplate reader (Labsystems, Helsinki, Finland).
Kinetic measurements using real-time biomolecular interaction analysis
The kinetics of protein S interaction with factor VIII and its subunits were determined by surface plasmon resonance (SPR)-based assays at 37°C using a BIAcore X instrument (BIAcore AB, Uppsala, Sweden). Protein S was covalently coupled to the surfaces of a CM5 chip at a coupling density of c. 9 ng/mm2. Binding (association) of the ligand was monitored in running buffer [10 mmol/l HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) pH 7·4, 0·1 mol/l NaCl, 1 mmol/l CaCl2, 0·005% polysorbate 20] for 2 min at a flow rate 10 μl/min. The dissociation of bound ligand was recorded over a 2-min period by replacing the ligand-containing buffer with buffer alone. The level of nonspecific binding, corresponding to ligand binding to the uncoated chip, was subtracted from the signal. After each analysis, chip surfaces were regenerated by treatment with 1 mol/l NaCl for 1 min. The rate constants for association (kasso) and dissociation (kdiss) were determined by nonlinear regression analysis using the evaluation software provided by Biacore AB. Dissociation constants (Kd) were calculated as kdiss/kasso.
Enzyme-linked immunosorbent assay (ELISA) for the binding of factor VIII to protein S
Microtitre wells were coated with factor VIII (200 nmol/l) in 20 mmol/l Tris, and 0·15 mol/l NaCl, pH 7·4 (Tris-buffered saline; TBS), overnight at 4°C. The wells were washed with phosphate-buffered saline (PBS) containing 0·01% Tween 20 and were blocked with PBS containing 5% human serum albumin (HSA) for 2 h at 37°C. The indicated concentrations of protein S were then added in 20 mmol/l HEPES, 0·1 mol/l NaCl, 0·01% Tween 20, pH 7·2 (HEPES-buffered saline; HBS) containing 5 mmol/l CaCl2 and 1% HSA and incubated for 2 h at 37°C. Anti-protein S mAb IgG (1 μg) was added and bound IgG was quantified by the addition of anti-mouse peroxidase-conjugated IgG and O-phenylenediamine dihydrochloride substrate. Reactions were stopped by the addition of 2 mol/l H2SO4, and absorbances were measured at 492 nm using a Labsystems Multiskan Multisoft microplate reader. The amount of nonspecific binding of anti-protein S mAb IgG in the absence of protein S was <5% of the total signal. Specific binding was recorded after subtracting the nonspecific binding.
ELISA for binding of factor VIII to VWF or phospholipid
These assays were performed using a minor modification of the method previously reported (Shima et al, 1993). Microtitre plates were coated with VWF (10 nmol/l) in TBS or phospholipid vesicles (40 μmol/l) in methanol overnight at 4°C. After blocking with 5% HSA, the indicated concentrations of factor VIII were added to the VWF-coated wells or phospholipid-coated wells and incubated for 2 h at 37°C. Bound factor VIII was detected using biotinylated JR8 at an absorbance 492 nm.
Cleavage of factor VIIIa by factor IXa
Factor IXa (20 nmol/l) was incubated with factor VIIIa (100 nmol/l) and the indicated concentrations of protein S in HBS-buffer containing phospholipid vesicles (10 μmol/l), 5 mmol/l CaCl2 and 0·02% Tween 20 at 37°C. Samples were obtained at the indicated times and the reactions were immediately terminated and prepared for PAGE by adding SDS and boiling for 3 min.
Electrophoresis and Western blotting
SDS-PAGE was performed using 8% gels at 150 V for 1 h. For Western blotting, the proteins were transferred to a polyvinylidene difluoride membrane at 50 V for 2 h in buffer containing 10 mmol/l CAPS [3-(cyclo-hexylamino)-1-propanesulfonic acid], pH 11 and 10% (v/v) methanol. Proteins were probed using anti-A1 mAb58.12, followed by anti-mouse peroxidase-conjugated secondary antibody. The signals were detected using the enhanced chemiluminescence system (PerkinElmer Life Science, Boston, MA, USA). Densitometry scans were quantitated using Image J 1.34 (National Institute of Health, USA).
All experiments were performed on at least three separate occasions; the average values and standard deviations were reported. Nonlinear least squares regression analysis was performed using Kaleidagraph (Synergy, Reading, PA, USA). The Km and kcat values for factor VIIIa/factor IXa-catalysed activation of factor X were calculated from the Michaelis–Menten equation.
Analysis of factor VIII and protein S interaction in the ELISA was performed using a single-site binding model as defined in Equation 1,
where [S] is the factor VIII concentration; Kd is the apparent dissociation constant; and Amax represents maximum absorbance signal when the site is saturated by factor VIII.
Effect of protein S on factor VIIIa/factor IXa-dependent activation of factor X in the absence of APC
To investigate whether protein S directly moderates the assembly of factor Xase on membrane surfaces in the absence of APC, we initially examined the effect of protein S on factor VIIIa/factor IXa-mediated activation of factor X in a functional Xa generation assay. Factor VIII (30 nmol/l) was incubated with various concentrations of protein S in the presence of phospholipid (10 μmol/l) and was activated by thrombin, followed by the addition of factor IXa (0·5 nmol/l) and factor X (200 nmol/l) as described in Materials and methods. The initial rate of factor Xa generated in the absence of protein S was c. 100 nmol/l/min. The addition of protein S showed complete inhibition of factor VIIIa/factor IXa-mediated factor X activation in a dose-dependent manner (Fig 1). Very little factor Xa was generated at the maximum concentration of protein S employed (800 nmol/l). The 50% inhibitory concentration (IC50) value was c. 150 nmol/l, consistent with the physiological concentration of protein S in plasma (c. 150 nmol/l; Griffin et al, 1992). We considered, however, that the inhibitory effect of protein S might result from direct interference with the activation of factor VIII to factor VIIIa by thrombin. To investigate this possibility, protein S was added to thrombin-activated factor VIIIa prior to the addition of factor IXa and factor X. The results in these circumstances were similar to those obtained above, and protein S inhibited factor X activation by c. 95% in a dose-dependent manner (IC50; c. 220 nmol/l). The findings indicated that protein S restricted factor Xa generation by binding to factor VIII or to factor VIIIa and did not directly impair thrombin-catalysed factor VIII activation. Furthermore, we examined the effects of phospholipid concentration to investigate the possibility that protein S displaces factor VIII(a)-factor IXa from the membrane surface. The concentrations of phospholipid did not significantly affect this inhibitory effect of protein S (data not shown). We also investigated the effects of protein S on factor IXa-catalysed factor X activation in the absence of factor VIIIa, in order to examine whether protein S significantly modulated the association between factor IXa and factor X in the factor Xase complex. In the absence of factor VIIIa, however, protein S inhibited factor X activation by only c. 10%, demonstrating that protein S did not significantly affect both the ability of factor IXa to promote factor X activation (or factor IXa–factor X association) and the activity of factor Xa on the chromogenic substrate. Taken together, these data suggest that protein S directly bound to factor VIII(a) to depress factor Xase activity even in the absence of APC.
Effect of factor IXa or factor X on factor Xa generation in the presence of protein S
To clarify the mechanism whereby protein S inhibited factor Xa generation by APC-independent interaction with factor VIII(a), we studied the role of factor IXa or factor X on factor Xa generation in the presence of protein S. Factor VIII (30 nmol/l) preincubated with physiological concentrations of protein S (150 nmol/l) was activated by thrombin. Various concentrations of factor IXa (0–400 pmol/l) were then added in the presence of a constant concentration of factor X (200 nmol/l), or alternatively, various concentrations of factor X (0–500 nmol/l) were added in the presence of a constant amount of factor IXa (0·5 nmol/l), as described in Methods. The results are shown in Fig 2 and summarized in Table I. Notably, the Km value for factor IXa obtained with factor VIIIa in factor Xase complex in the presence of protein S was 340 ± 42 pmol/l, which was c. 17-fold higher than that in its absence (20·2 ± 2·2 pmol/l) (Fig 2A). The Vmax value for factor IXa in the presence of protein S was c. 80% of that obtained in its absence (83·4 ± 7·8 and 102 ± 4 nmol/l/min respectively). In contrast, the Km value for factor X in the presence of protein S was 19·6 ± 2·9 nmol/l, which was c. twofold lower than its absence (45·5 ± 9·2 nmol/l) (Fig 2B). The Vmax value for factor X in the presence of protein S was c. 70% of that obtained in its absence (80·0 ± 2·2 and 118 ± 7 nmol/l/min respectively). In control experiments, the Km for factor IXa on factor X activation in the absence of factor VIIIa was not significantly affected even in the presence of protein S, and the Vmax with protein S was c. 80% of that in its absence (data not shown). The results supported that protein S did not directly affect the association between factor IXa and factor X, consistent with the data shown in Fig 1. The findings were in keeping with the conclusion that the decrease in factor Xase activity in the presence of protein S resulted from inhibition of the association between factor VIIIa and factor IXa.
|Protein S||Factor IXa||Factor X|
|Vmax (nmol/l/min)||Km (pmol/l)||Vmax (nmol/l/min)||Km (nmol/l)|
|Minus||102 ± 4||20·2 ± 2·2||118 ± 7||45·5 ± 9·2|
|Plus||83·4 ± 7·8||340 ± 42||80·0 ± 2·2||19·6 ± 2·9|
Binding of protein S to immobilized factor VIII
To obtain direct evidence that protein S interacted with factor VIII independent of APC, we evaluated co-factor binding in a solid-phase ELISA. Various concentrations of protein S were incubated with factor VIII (200 nmol/l) that had been immobilized on microtitre wells, and bound protein S was detected using anti-protein S mAb as described in Methods. This antibody did not affect the reaction between protein S and factor VIII (data not shown). As shown in Fig 3A, protein S bound to factor VIII with apparent saturable, dose-dependent binding curves. This method is not a true equilibrium binding assay, however, and the Kd value represents an apparent Kd for the interaction of protein S with factor VIII. Nevertheless, the data were comparatively well-fitted in a single-site binding model, with a modest apparent Kd (28·7 ± 1·7 nmol/l). We also performed the binding experiment using protein S coated on microtitre wells (data not shown), and observed the similar Kd (31·8 ± 4·9 nmol/l) of that obtained in which factor VIII was coated. To confirm the specificity of this binding, various concentrations of factor VIII were preincubated with protein S (100 nmol/l) in the fluid phase, prior to addition to the immobilized factor VIII. Soluble factor VIII blocked protein S binding to immobilized factor VIII by c. 80% with similar Ki value (30·8 ± 2·7 nmol/l, Fig 3B), confirming the specificity of this assay. Similar binding experiments with active factor VIIIa were difficult to interpret, however, because the A2 domain dissociated from the active molecule readily during incubation and/or wash procedures.
Characterization of factor VIII-protein S interaction
To further characterize the interaction of protein S with factor VIII, the effect of ionic strength was examined. Protein S (200 nmol/l) and various amounts of NaCl were incubated with immobilized factor VIII (Fig 4A). The binding of protein S to factor VIII was slightly enhanced at low ionic strength (at 10 mmol/l). Incremental NaCl concentrations markedly weakened the interaction, however. An c. twofold decrease in bound protein S was evident at a physiological concentration of NaCl (c. 140 mmol/l), compared with lower ionic strength. Control experiments using biotinylated anti-A2 mAbJR8 showed that the amount of factor VIII immobilized on the microtiter wells was not affected by the ionic strength of the wash buffer or the duration of the wash and incubation cycles (data not shown). The results demonstrated that the interaction between protein S and factor VIII was sensitive to salt concentration.
The structure and function of both protein S and factor VIII are known to be dependent on Ca2+. For this reason, the binding of protein S to immobilized factor VIII was assessed in buffer containing various amounts of CaCl2 (Fig 4B). The total ionic strength of the buffer was kept constant by the addition of NaCl, and EDTA (10 mmol/l) was added to the reaction mixtures to assess binding in the absence of CaCl2. A linear increase in protein S binding by up to c. 1·9-fold was observed at concentrations of Ca2+ up to 1 mmol/l compared with that in the absence of Ca2+. Binding was inhibited, however, by increments of Ca2+ above 1 mmol/l. It seemed likely, therefore, that Ca2+ was necessary for factor VIII-protein S interaction, although it was not possible to distinguish between a direct or indirect role for Ca2+ in mediating this effect.
Further experiments were developed to investigate the contribution of exposed hydrophobic sites on factor VIII for protein S binding. Binding was measured as above in the presence of the apolar compound bis-anilinonaptholsulfonic acid, which binds to the exposed hydrophobic sites on factor VIII (Sudhakar & Fay, 1996). However, occupancy of hydrophobic sites on factor VIII by this reagent only slightly affected the protein S binding (data not shown). Collectively, these data indicated that factor VIII bound to protein S predominantly by electrostatic- and/or calcium-dependent mechanisms.
Interactions between factor VIII(a) subunit and protein S in SPR-based assays
To determine the role of different factor VIII(a) domains in protein S binding, interactions between factor VIII(a) subunits and protein S were evaluated in SPR-based assays. A range of concentrations of different factor VIII(a) products were added to protein S immobilized on a sensor chip. Figure 5 shows representative binding curves corresponding to the association/dissociation of factor VIII (Fig 5A), the A2 subunit (Fig 5B), and rA3 domain (Fig 5C). The results are summarized in Table II. The data were comparatively well-fitted by non-linear regression analysis using a 1:1 Langmuir binding model. Kinetic constants indicated that intact factor VIII bound to protein S with high affinity (Kd: 14·3 nmol/l), similar to that determined by ELISA. The intact heavy chain and light chain also bound with similar affinities (Kd: 50·3 and 43·1 nmol/l respectively), showing an c. threefold lower affinity compared to that of factor VIII. It was noteworthy that the isolated A2 domain (sequenced to residues 373–740) and rA3 domain (sequenced to residues 1690–2019, lacking the acidic region) bound to protein S with high affinities (Kd: 12·4 and 10·6 nmol/l respectively), almost equivalent to that obtained with factor VIII. However, the isolated A1 domain (sequenced to residues 1–372) and rC2 domain (sequenced to 2174–2332) failed to bind to protein S. As discussed above, the kinetics for active factor VIIIa could not be determined, because the A2 domain dissociated readily from the active molecule. The results indicated that both A2 and A3 domains of factor VIIIa predominantly contributed to protein S binding.
|Factor VIII subunit||kass (104/mol/l/s)||kdiss (10−3/s)||Kd* (nmol/l)|
|Factor VIII||13·3 ± 2·2||1·9 ± 0·6||14·3|
|Heavy chain||2·9 ± 0·9||1·5 ± 0·3||50·3|
|Light chain||6·3 ± 3·4||2·7 ± 0·3||43·1|
|A2||8·3 ± 2·3||1·0 ± 0·3||12·4|
|A3||9·1 ± 1·5||0·9 ± 0·2||10·6|
Effect of protein S on binding of factor VIII to VWF or phospholipid
Factor VIII circulates as a complex with VWF. After activation by thrombin, factor VIIIa, free of VWF, binds to phospholipid membranes for the formation of the factor Xase complex (Lollar et al, 1988). Major VWF-interactive sites in factor VIII have been located in the acidic region of the A3 domain (Foster et al, 1988) and the C2 domain (Saenko et al, 1994), and a phospholipid-interactive site has been located in the C2 domain (Pratt et al, 1999). To examine the effects of protein S on factor VIII binding to VWF or phospholipid, we established competitive binding experiments using the immobilized VWF or phospholipid in ELISA-based assays. Protein S did not significantly inhibit the factor VIII binding to either VWF or phospholipid vesicles (data not shown). The findings were consistent with the above data showing that protein S bound to the A2 and A3 domains (devoid of the acidic region), which do not interact with either VWF or phospholipid. In addition, protein S did not significantly affect the factor IXa and phospholipid binding (data not shown).
Inhibitory effect of EGR-factor IXa on factor VIII and protein S interaction
The investigations described above showed that protein S moderated the association between factor VIIIa and factor IXa within the factor Xase complex. Furthermore, factor IXa interacts with the A2 and A3 domains of factor VIII (Fay et al, 1994; Lenting et al, 1994). To clarify the relationship between factor IXa and protein S for factor VIII binding, therefore, we examined the effect of factor IXa on factor VIII (200 nmol/l) binding to immobilized protein S in SPR-based assays as described in Methods. Active-site modified, EGR-factor IXa, lacking enzymatic activity, was utilized in place of factor IXa in these experiments. Control experiments showed that EGR-factor IXa did not bind significantly to protein S (data not shown). EGR-factor IXa inhibited the binding (association) of protein S to factor VIII in a dose-dependent manner (Fig 6A). On the basis of the maximum response value (RU) at 120 s after addition of the factor VIII/EGR-factor IXa mixture (Fig 6Ainset), the inhibitory effect was calculated to be c. 65% at the highest concentrations employed (500 nmol/l). Similar experiments were undertaken to assess the effect of EGR-factor IXa on isolated A2 or rA3 domains (200 nmol/l) binding to protein S. The RU (at 120 s) after the addition of A2/EGR-factor IXa or A3/EGR-factor IXa mixture indicated dose-dependent inhibition of subunit binding (up to >90% at the highest concentrations employed, 500 nmol/l) (Fig 6B). These data demonstrated that factor IXa and protein S competed for binding to factor VIII(a), and as a consequence factor Xase activity was depressed by protein S.
Protein S inhibits the proteolytic cleavage of factor VIIIa by factor IXa
Factor IXa as well as APC proteolytically cleaves factor VIIIa at Arg336 in the A1 domain (Eaton et al, 1986). To directly observe that protein S competitively inhibited the interaction between factor VIIIa and factor IXa, SDS-PAGE-based assays were employed to visualize factor IXa-induced cleavage of the factor VIIIa A1 domain in the fluid phase. This cleavage inactivates factor VIIIa, and it is possible that the combined interaction of protein S and factor IXa on factor VIIIa could modify the activation of factor X. It is widely accepted, however, that APC is the most important enzyme in the degradation of factor VIIIa and that the most prominent function of factor IXa is to accelerate factor X activation in the presence of factor VIIIa in the factor Xase reaction. Moreover, it is not known whether factor IXa decreases factor VIIIa activity on the factor Xase complex. Therefore, we utilized this assay as a limited approach to observe the inhibitory effect of protein S for factor VIIIa-factor IXa association.
Factor VIIIa (100 nmol/l) was incubated with factor IXa (20 nmol/l) and phospholipid (10 μmol/l) in the presence of protein S (200 nmol/l), and the reactions were quenched at specified intervals as described in Methods. Intact A1 (A11–372) and the products of proteolysis (A11–336) were visualized by Western blotting using an anti-A1 mAb58.12 recognizing the N-terminal end (Fig 7A). In the presence of protein S (Fig 7A panel b), Arg336 cleavage in the A1 domain appeared to be slower than that in the absence of protein S (Fig 7A panel a). Scanning densitometry analysis obtained from the Western blotting and fitting data using a straight line (Fig 7A panel c) indicated that the cleavage rate at Arg336 in the presence of protein S was c. 1·8-fold slower than that in its absence. We further examined the effects of various concentrations of protein S on factor IXa-catalysed A1 cleavage at Arg336. Panels a and b in Fig 7B show the results of Western blotting using an anti-A1 mAb58.12 and scanning densitometry respectively. The addition of protein S resulted in significant inhibition of Arg336 cleavage in A1 in a dose-dependent manner. The IC50 value obtained was c. 150 nmol/l, consistent with the physiological concentration of the cofactor, and in keeping with the results obtained in a functional Xa generation assay (Fig 1). Taken together, these data again strongly indicated that protein S directly inhibited interactions between factor VIIIa and factor IXa.
Protein S functions as a cofactor for APC in the inactivation of factor VIIIa and as an APC-independent anticoagulant in inhibiting intrinsic factor X activation (Koppelman et al, 1995). However, this latter mechanism is poorly understood. The present study has shown, for the first time, that protein S impairs factor Xase assembly, independent of APC, by directly competing in the interaction between factor IXa and factor VIIIa. This conclusion is based on several novel findings using well established models: (i) Protein S in the absence of APC significantly limited the activity of factor Xase complex in a functional Xa generation assay in the presence of factor VIIIa. (ii) The presence of protein S resulted in a c. 17-fold increase in Km for factor IXa obtained with factor VIIIa on factor Xase complex. (iii) Both the A2 and A3 domains of factor VIII directly bound to protein S with high affinities, whilst the A1 and C2 domains failed to bind. (iv) Competitive binding assays using EGR-factor IXa showed that factor IXa, which binds to the A2 and A3 domains, completely inhibited protein S binding to these domains. (v) SDS-PAGE analysis showed that cleavage of factor VIIIa at Arg336 by factor IXa was c. twofold slower in the presence of protein S than that in its absence. These data clarified the essential role of protein S as an APC-independent anticoagulant in the intrinsic factor Xase complex.
Protein S restricted the activity of the factor Xase complex by competing with factor IXa for binding to factor VIIIa. Consistent with an earlier report (Koppelman et al, 1995), we demonstrated that protein S had no direct effect on factor IXa in the factor X activation mechanism. This was also supported by our observation that EGR-factor IXa failed to interact with immobilized protein S in SPR-based assays. In contrast, the addition of protein S together with factor VIIIa significantly reduced the acceleration of factor IXa-dependent factor X activation, confirming that the factor VIIIa-protein S complex does not function effectively as a cofactor of factor IXa. The presence of protein S with factor VIIIa resulted in a much lower affinity (c. 340 pmol/l) for factor IXa on the factor Xase assembly. The effects were clearly evident at a physiological concentration of free protein S (c. 150 nmol/l), suggesting that protein S could directly contribute to the control of the factor Xase complex in vivo.
It is known that protein S decreases factor Xa activity (Heeb et al, 1994). In the present study, however, protein S did not affect the amidolytic activity of factor Xa in the factor Xa generation assay (data not shown). A previous report indicated that the decrease in factor Xa activity mediated by protein S was observed only in the presence of Ca2+ (Hackeng et al, 1994). In our experimental conditions, EDTA was added to the test mixtures, and this seems likely to have prevented inhibition. Of interest, the affinity of factor X for factor VIIIa on the factor Xase complex in the presence of protein S was c. twofold lower than the absence of protein S, suggesting that protein S may enhance the association between factor VIIIa and factor X. The reason for this finding is unclear at present.
Heeb et al (1993) demonstrated that the anticoagulant properties of protein S included inhibition of the prothrombinase reaction independent of APC. They suggested that this mechanism centered on protein S competition with prothrombin for direct binding to factor Va. In our studies, however, we did not find that protein S affected the factor VIIIa-factor X, cofactor-substrate association in the factor Xase complex. In later studies, the same group showed possible competition between protein S and factor Xa for a binding site on factor Va in the prothrombinase complex (Heeb et al, 1996), and our observations that protein S was competitive in the factor VIIIa-factor IXa, cofactor-enzyme association in the factor Xase complex, were consistent with that concept. The findings add to the evidence that structurally homologous regions in cofactors factor Va and factor VIIIa are responsible for functionally similar protein-protein interactions.
Direct binding of factor VIII(a) to protein S was described earlier using an ELISA method (Koppelman et al, 1995). The present study confirmed that protein S directly interacted with factor VIII using SPR-based assays. This provided strong evidence that factor VIII interacted with protein S with high affinity (Kd: c. 14 nmol/l). The interaction was essentially electrostatic and Ca2+-dependent. Our observations further indicated that the A2 domain in the heavy chain and the A3 domain in the light chain bound to protein S with similar affinities (Kd: c. 10 nmol/l), highlighting the significant contribution of the A2 domain (residues 373–740) and A3 domain (residues 1690–2019) in the interaction with protein S. Both domains are responsible for factor IXa-binding to factor VIIIa. Our findings that the presence of VWF (bound to the A3 acidic region and C2 domains) and phospholipid (bound to the C2 domain) did not significantly affect factor VIII(a)-protein S interaction are in keeping with a predominant role for the A2/A3 regions in this mechanism. Since the concentration of factor VIII in plasma is c. 0·3 nmol/l, protein S is unlikely to exist in complex with factor VIII/VWF complex. However, the concentrating effect of factor VIIIa in a two-dimensional space by the factor Xase complex might support our present observation that protein S contributes to the down-regulation of factor Xase activity.
As discussed above, factor V(a), possessing the similar structure and homology to factor VIII(a), is known to directly interact with protein S (Heeb et al, 1993). The elegant studies of Heeb et al (1996) showed that a protein S-binding site on factor V(a) was located within the A2 domain (residues 493–506) of the heavy chain, near to the APC cleavage site at Arg506. This region has a substantial net positive charge and overlaps with a factor Xa-interactive site. On the other hand, factor IXa-interactive sites on the factor VIII A2 domain involve clustered basic residues and are localized within, at least, residues 484–509 and 556–565 (Fay et al, 1994; Fay & Scandella, 1999). The latter region also contains an APC cleavage site at Arg562. However, Heeb et al (1996) also reported that protein S failed to bind to the factor VIII-related peptide (residues 549–562) derived from the region homologous to factor V residues 493–506. It is possible to speculate, therefore that a protein S-interactive site on the A2 domain may involve residues 484–509. In addition, a factor IXa-interactive site involving clustered basic residues has been localized within residues 1804–1818 in the A3 domain (Lenting et al, 1994), and this might overlap a protein S-interactive site on A3. A similar binding site on the factor Va light chain could be expected but remains to be identified. Further studies are in progress to identify precisely the protein S-interactive sites on the A2 and A3 domains of factor VIIIa.
Regan et al (1994) illustrated that factor IXa protected factor VIIIa from inactivation by APC, and protein S negated this protective effect . Factor IXa appeared to block an important binding or cleavage site for APC on the A2 domain. The current finding, that protein S bound to factor VIIIa, also supported that earlier report. Therefore, it may be that protein S negates the protective effect of factor IXa by displacing factor IXa from its binding site on factor VIIIa, thereby making or keeping available a factor VIIIa site for APC binding. Protein S similarly appears to counteract the protective effect of factor Xa on factor Va inactivation by APC (Solymoss et al, 1988) and this could involve an analogous mechanism.
The physiological importance of the APC pathway is well illustrated by the clinical observations that protein C deficiency is associated with recurrent venous thrombosis. A deficiency of protein S is also clinically associated with thrombotic complications. Protein S deficiency but not protein C deficiency, however, appears to cause arterial as well as venous thrombosis in some patients (Mannucci et al, 1986). Protein S enhances the activity of APC by only c. twofold (Solymoss et al, 1988), and it is difficult to ascribe the clinical differences to the haemostatic role of APC alone. The evidence that protein S possesses at least two anticoagulant functions that are independent of APC, by inhibiting intrinsic factor X activation in a competitive reaction with factor IXa for direct binding to factor VIIIa, and by inhibiting prothrombinase activity in competitive reaction with prothrombin and/or factor Xa for direct binding to factor Va, might help to clarify the additional clinical phenomena. On the other hand, a high factor VIII level is also probably related to thrombosis, although the mechanism is still currently unknown. However, this phenomenon may overcome the inhibitory effect of protein S. These presume that coagulation regulation and clinical observations related to protein S might be more complexed than previously thought. Furthermore, c. 50% of the protein S in plasma is bound to C4b-binding protein as another major plasma form (Dahlbäck & Stenflo, 1981). However, the role of C4b-binding protein for the APC-independent anticoagulant mechanism remained to be fully investigated. Further investigations would be required to understand more physiological and clinical significance.
We thank Dr J. C. Giddings for helpful suggestions. This work was partly supported by grants for MEXT KAKENHI (19591264) and Mitsubishi Pharma Research Foundation.
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