Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C

TR-05509; No of Pages 9 Thrombosis Research xxx (2014) xxx–xxx

Contents lists available at ScienceDirect

Thrombosis Research journal homepage: www.elsevier.com/locate/thromres

Regular Article

Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C Sofia Somajo, Ruzica Livaja Koshiar, Eva Norström, Björn Dahlbäck ⁎ Lund University, Department of Laboratory Medicine, Division of Clinical Chemistry, Skåne University Hospital, SE-205 02 Malmö, Sweden

a r t i c l e

i n f o

Article history: Received 7 February 2014 Received in revised form 28 March 2014 Accepted 7 April 2014 Available online xxxx Keywords: Protein S Activated protein C Factor V Factor VIII Coagulation Microparticles

a b s t r a c t Introduction: Platelets are the main source of microparticles in plasma and the concentration of microparticles is increased in many diseases. As microparticles expose negatively charged phospholipids, they can bind and assemble the procoagulant enzyme-cofactor complexes. Our aim was to elucidate possible regulation of these complexes on microparticles by the anticoagulant protein C system. Materials and methods: Platelets were activated with thrombin ± collagen or the calcium ionophore A23187 ± thrombin to generate microparticles. The microparticles were analyzed using flow cytometry and functional coagulation assays to characterize parameters with importance for the activated protein C system. Results: Activation with A23187 + thrombin was most efficient, fully converting the platelets to microparticlelike vesicles, characterized by high lactadherin and protein S binding capacity. Suppression of thrombin generation by activated protein C in plasma spiked with these microparticles was dependent on the presence of plasma protein S. Experiments with purified components showed that activated protein C inhibited both factor Va and factor VIIIa on the microparticle surface. Inhibition of factor Va was stimulated by, but not fully dependent on, the presence of protein S. In the factor VIIIa-degradation, activated protein C was dependent on the addition of protein S, and exogenous factor V further increased the efficiency. Conclusions: Protein S is crucial for activated protein C-mediated inhibition of thrombin generation on plateletderived microparticles in plasma. Moreover, protein S and factor V are synergistic cofactors in the inhibition of factor VIIIa. The results demonstrate that the activated protein C system has the capacity to counterbalance the procoagulant ability of microparticles. © 2014 Elsevier Ltd. All rights reserved.

Introduction Microparticles (MPs) are small membrane-containing vesicles released by numerous cell types upon activation, apoptosis or stress [1–3]. Their features are dependent on the cell origin; hence they can be identified by surface expression of cell specific markers. MPs from platelets have a high surface content of the negatively charged phospholipid phosphatidylserine, and support the activations of factor X (FX) and prothrombin [4–8]. FX is activated by the intrinsic Xase complex where activated factor IX (FIXa), together with its cofactor activated factor VIII (FVIIIa), is bound to negatively charged phospholipids,

Abbreviations: APC, activated protein C; C4BP, C4b-binding protein; ECL, enhanced chemiluminescence; EDTA, ethylenediaminetetraacetic acid; F(), coagulation factor (number); F()a, activated coagulation factor (number); MPs, microparticles; PSB, platelet storage buffer; PTase, prothrombinase; PWB, platelet wash buffer; TF, tissue factor; TFPI, tissue factor pathway inhibitor; Xase, intrinsic tenase. ⁎ Corresponding author at: Lund University, Department of Laboratory Medicine, Division of Clinical Chemistry, Wallenberg Laboratory, Skåne University Hospital, Inga Marie Nilssons gata 53, SE-205 02 MALMÖ, Sweden. Tel.: +46 40 331501; fax: +46 40 337044. E-mail address: [email protected] (B. Dahlbäck).

whereas prothrombin is activated by the prothrombinase (PTase) complex (activated FX (FXa), its cofactor activated factor V (FVa) and the phospholipids). The platelets contain about 20% of the total amount of FV in blood [9]. This platelet-derived FV is released upon activation and bind to the negatively charged activated platelets, thereby contributing to the formation of PTase complexes. The fully assembled complexes are 105-106 more efficient than the respective enzymes alone [10,11]. The Xase- and PTase complexes are regulated by activated protein C (APC) [12], which inactivates FVIIIa and FVa [13,14]. Protein S serves as an APC-cofactor in these reactions [15,16]. In human plasma, approximately 35% of protein S is free, the remaining being bound to C4bbinding protein (C4BP) [17]; mainly the free form serving as APC cofactor [18]. In the inhibition of FVIIIa, intact FV functions in synergy with protein S as cofactor to APC [19,20]. Platelets contain around 2.5% of the total protein S in blood, and it is released upon platelet activation [21]. The functional importance of protein S and protein C is evident from the increased risk of venous thrombosis affecting individuals with heterozygous deficiency of either protein [22]. Normal plasma contains 0.5-2.8 × 106 MPs/mL [23,24], but in several diseases, such as cancer [25], rheumatoid conditions [26], diabetes [27],

http://dx.doi.org/10.1016/j.thromres.2014.04.031 0049-3848/© 2014 Elsevier Ltd. All rights reserved.

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

2

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

systemic lupus erythematosus [24], atherosclerosis and coronary disorders [28], increased numbers of circulating MPs have been reported. Interestingly, an increased risk of thrombosis is observed in many of these diseases and it has been postulated that the circulating MPs contribute to the thrombosis risk. While the procoagulant functions of platelet-derived MPs have been extensively investigated, few reports focus on the anticoagulant properties of MPs. Several studies have shown that APC-mediated inactivation of FVa on activated platelets is hampered and that platelet-derived FVa is less susceptible to APC-mediated degradation than plasma-derived FVa [29–31]. Degradation of FVa by APC on the surface of ionophoreactivated platelets (MPs) has been shown to be more efficient than on platelets activated by thrombin, however compared to phospholipid vesicles, the degradation rate was still low [5,32]. The APC-mediated degradation of FVIIIa on MPs has not been studied, but plateletderived MPs can bind FVIIIa and FIXa [6,7]. It has previously been shown that free, but not C4BP-bound, protein S specifically binds to platelet-derived MPs but not to resting or activated platelets [33]. To investigate whether the binding of protein S to the MPs renders them less procoagulant, the ability of protein S and FV to function as cofactors to APC on MPs was investigated.

followed by resuspension of the platelets in 1.5 mL PSB. Washed platelets were counted (phase contrast inverted microscope, Olympus IMT2-RFL, LRI Instrument AB, Lund, Sweden) using Bürker C-chip (Digital Bio, NanoEnTek, Seoul, Korea). The platelets were diluted to 75 × 106/mL in PSB for further processing. Buffers were room tempered and filtered (0.20 um), low affinity transfer pipets (7.5 mL, 15.5 cm, VWR, Radnor, PA, USA) were used to resolve platelet pellets and the platelets were handled in non-adhesive tubes (#347708 Nunc, Thermo Fisher Scientific), to avoid activation. Washed platelets (1.5 mL, 75 × 106/mL) were activated using 0.5 or 5 U/mL thrombin and 2 mM CaCl2, or 0.5 or 5 U/mL thrombin plus 25 μg/mL collagen and 4 mM CaCl2, or 5 μM calcium ionophore A23187 and 4 mM CaCl2, at 37 °C, 15 minutes. Some batches of A23187-activated platelets were further treated with 0.5 or 5 U/mL of thrombin (37 °C, 10 min). Thrombin activity was inhibited by addition of double concentration of hirudin. Experiments were performed within 6 hours from blood collection, and 3 hours from activation time and were kept at room temperature meanwhile. The platelets/MPs concentrations will be referred to as the concentration equivalent to the concentration of platelets before activation. Platelets from different donors were used. Flow cytometry

Materials and methods Reagents Antibodies were from BD Biosciences, Franklin Lakes, NJ, USA (antiCD41a-PerCPcy5.5 and mouse IgG1-PerCPcy5.5) or Beckman Coulter, Brea, CA, USA (anti-CD61-PE and mouse IgG1-PE). Rabbit-anti-protein S (A0384 DAKO, Glostrup, Denmark) was labeled with Alexa488 using the Microscale Protein Labeling kit (A30006), (Life Technologies, Invitrogen, Carlsbad, CA, USA). FIXa and lactadherin-FITC were from Haematologic Technologies Inc, Essex Junction, VT, USA. Protein S and FXa were from Enzyme Research Laboratories (ERL, South Bend, IN, USA). Ionophore A23187 (calcimycin) was from Life Technologies, Invitrogen, FVIII was from Octapharma, Lachen, Switzerland. Human FV was purified from plasma as described [34], with minor modifications [5]. Bovine FX [35], and human prothrombin [36] were purified from plasma. Human APC was obtained from recombinant protein C expressed, purified and activated, as described [37]. Human αthrombin was prepared from purified prothrombin, as described [38]. Hirudin was from Pentapharm, Basel, Switzerland. Collagen (native fibrils type 1 form equine tendons) was from Chrono-Log Corp. Haverton, PA, USA. Bovine serum albumin (BSA, #A7030) was from Sigma-Aldrich (St Louis, MO, USA). Natural phospholipids phosphatidylserine (brain extract) and phosphatidylcholine (egg extract) were from Avanti Polar Lipids Inc. (Alabama, USA). Ready gels (4-15% TGX) and stacks for SDS-Page and western blotting were from (Bio-Rad, Hercules, CA, USA).

Two flow cytometers; FC500 and Gallios (Beckman Coulter, Brea, CA, USA) were used. IsoFlow™ Sheath Fluid (Beckman Coulter) was used as fluid phase and the laser was set at 488 nm. In the Gallios, the laser setting W2 was used. Thresholds for forward scatter and side scatter were set to 2. Forward scatter, side scatter and fluorescence channels were set at logarithmic gain. The flow cytometry data were analyzed using FlowJo 8.7.1 (Tree Star, Inc., Ashland, OR, USA). The platelet gate was set using resting platelets and MPs were defined based on phosphatidylserine exposure and protein S binding capacity. To verify the MP gate, washed platelets were incubated with A23187 and centrifuged (1 000 g, 10 min) to pellet platelets. The forward/side scatter properties of the MPs in the supernatant coincided with the populations defined as MPs by phosphatidylserine exposure (lactadherin binding) and protein S binding. Centrifugation of resting platelets led to almost complete loss of events in the supernatant. Counting of platelets/MPs using flow cytometry After incubation (15-20 min RT, dark) of platelets/MPs (7.5 × 106/mL, 5 μL) with mouse-anti-CD41a-PerCPCy5.5 (5 μL, 25 μg/mL) and lactadherin-FITC (5 μL, 1.6 μM) in a total volume of 50 μL PSB, 50 μL calibration beads (Flow Count®, Beckman Coulter) were added and the mixture was diluted to 500 μL with PSB. CD41-positive platelets/MPs (FL4) were collected in flow cytometer FC500 and compared to the known concentration of calibration beads.

Platelets

Phosphatidylserine exposure measured by lactadherin binding

Platelets were obtained from fresh citrated blood (Vacutainer citrate 4.5 mL tubes, BD, Franklin Lakes, NJ, USA) collected from healthy volunteers after informed consent (ethical permission 2012/202, Regional Ethical Review Board, Lund, Sweden. Whole blood was centrifuged (Hermle Labortechnik, Wehnigen, Germany) at 250 g, 15 min, and the obtained platelet rich plasma was pooled into new tubes (10 mL/tube) containing 1 mL platelet wash buffer (PWB, 100 mM Tri-Na-citrate, 10 mM citrate, 150 mM glucose, pH 6.5, supplemented with 2 mM adenosine (SigmaAldrich) and 7.5 mM theophylline (Sigma-Aldrich), freshly prepared) and centrifuged 1 000 g, 10 min. The platelet pellet was carefully resolved in PWB and transferred to new tubes containing 10 mL PWB and centrifuged as before (repeated once). Platelet storage buffer (PSB) (140 mM NaCl, 2.5 mM KCl, 0.1 mM MgCl2, 10 mM Hepes (4-(2-Hydroxyethyl1)-1-piperazineethanesulfonic acid), 0.5 mM NaH2PO4, 5.5 mM glucose, 10 mM HCO3, pH 7.4) was carefully laid over the pellet and removed,

Approximately 0.375 × 106 platelets/MPs were incubated for 1520 min, RT, with 5 μL lactadherin-FITC (1.6 μM) and 5 μL mouse antiCD61-PE (25 μg/mL) in a total volume of 50 μL PSB. After labeling, the platelets/MPs were diluted to 500 μL in PSB and the analyzed in Gallios. CD61-positive events were collected as identified in FL2. LactadherinFITC-binding (detected in FL1) was used to estimate activation state due to exposure of phosphatidylserine [39]. Protein S binding measured in by flow cytometry Platelets/MPs (0.375 × 106) were incubated for 15-20 min, RT with 5 μL protein S (125 μg/mL), 5 μL mouse anti-CD61-PE (25 μg/mL) in a total volume of 50 μL PSB containing 4 mM CaCl2. Rabbit-anti-protein S (alexa488-labeled) was added (5 μL 90 μg/mL) to detect protein S binding. After incubation as above, the platelets/MPs were diluted

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

with 500 μL PSB and CD61-positive (FL2) events were collected and protein S-binding was detected in FL1 (Gallios). Protein S was excluded for negative control and the specificity of the polyclonal antibody was confirmed using Alexa488-labeled monoclonal antibody against protein S (HPS34) [33].

3

protein S and/or polyclonal anti-protein C (100 μg/mL, DAKO) or monoclonal in house anti-protein S (100 μg/mL, HPS54). Aliquots were drawn, diluted and the remaining FVa activity measured as described above. Analysis of kinetic data

Prothrombinase assay The PTase assay was performed as described before [40]. Human plasma-purified FV was used to obtain a linear standard curve for calculation of FVa-concentration. The synergistic effects of phosphatidylserine exposure and release of platelet-derived FV/FVa from the platelets/MPs was evaluated using a modified PTase assay. To 50 μL of reaction mix (70 × 106 platelets/MPs/mL, 25 nM hFXa, 10 mM CaCl2 in PSB), 50 μL HNBSA, 5 mM CaCl2 were added and incubated 15 s. The reaction was started by addition of 150 μL of prothrombin (833 nM in 25 mM HEPES, 150 mM NaCl, 2 mM CaCl2, pH 7.7, 0.5 mg/mL ovalbumin (Sigma-Aldrich)). After 2 minutes, the reaction was stopped by dilution (1/8) in EDTA-stop buffer (50 mM Tris, 100 mM NaCl, 20 mM EDTA (ethylenediaminetetraacetic acid), 1% polyethylene glycol6000 (PEG6000), pH 7.9). The assay was performed at 37 °C, and the components were pre-heated for 5 minutes before assay start. Final concentration: 0.5 uM prothrombin, 5 nM hFXa, 3 mM CaCl2 and 14 × 106 platelets/MPs/ mL. Formed thrombin was measured kinetically with chromogenic substrate (S-2238, Chromogenix, Milan, Italy) in Infinite 200 microplate reader (Tecan, Männedorf, Switzerland). Data was collected every 30 s for 15 minutes. Human thrombin of known concentration was used to obtain a calibration curve for the conversion of the substrate.

FVa is cleaved by APC at Arg506 and Arg306 by the two pathways presented below (Eqs. (1), and (2)), resulting in the rate constants k506, k306 and k’306 (FVaint = FVa cleaved at Arg506 having intermediate activity, FVai = completely inactivated FVa) [41]. k506

k306

FVa → FVaint → FVai

ð1Þ

k0306

FVa → FVai

ð2Þ

To determine the rate constants for FVa-degradation on platelets/ MPs, the kinetic data were fitted to the pseudo first-order equation (Eq. (3)), which was based on the observation that k306 was similar to k’306 [41]. −ðk506 þk306 Þt

Vat ¼ Va0  e

−k306 t

þ B  Va0  e

 1−e

−k506 t

ð3Þ

Vat represents total FVa activity, Va0 is the FVa activity at time (t) 0 and B is the remaining activity of FVaint. Electrophoresis, immunoblotting and ELISA

FVIIIa-degradation, Xase assay A reaction mix containing FVIII (1 U/mL) and FIXa (8.9 nM) was treated with 0.1 U/mL thrombin (37 °C, 3 minutes) to activate FVIII. The reaction was stopped with 0.3 U/mL hirudin. The activation mix was diluted with FIXa (8.9 nM) to obtain 370 mU/mL FVIIIa. APC (0-5 nM) was added and allowed to degrade FVIIIa (212 mU/) in presence of FIXa (5 nM) and platelets/MPs (24 × 106/mL), with or without protein S (33 nM) and/or intact FV (2 nM) (total volume 105 μl). All reagents were in HNBSA (25 mM Hepes (4-(2-Hydroxyethyl-1)-1piperazineethanesulfonic acid, 150 mM NaCl2, 0.5 % bovine serum albumin), 5 mM CaCl2, pH 7.7. After 2.5 minutes, 20 μl of bovine FX was added (final concentration 0.5 μM) and further incubated (3 min), before stopping the reaction by dilution in ice-cold EDTA-stop buffer. The remaining activity of FVIIIa was determined as the amount of formed FXa, assessed by kinetic measurement of the conversion of the chromogenic substrate S-2765 (Chromogenix). Known concentration of FXa was used to obtain a calibration curve for the conversion of the substrate. FVa degradation assay Plasma purified FV (5.5 nM) was activated with 1 U/mL of thrombin 10 min, 37 °C, and the reaction was stopped by addition of 2 U/mL hirudin. A degradation mix (50 μL), containing MPs (56 × 106 MPs/mL), APC (0-125 pM), FVa (0.8 nM) and protein S (100 nM) was incubated (10 minutes, 37 °C). To study endogenous FVa-degradation, plasma purified FVa was excluded. To stop APC-degradation of FVa, the reaction mix was diluted 1/10 in ice-cold HNBSA, 5 mM CaCl2. To evaluate the remaining FVa-activity, a PTase assay was performed as described [40]. Extruded phospholipid vesicles were included in the PTase, as insufficient concentrations of membrane surface were provided by the platelets/MPs mixtures. Final concentration: 0.6 × 106/mL platelets/MPs, 50 μM phospholipids (10/90 phosphatidylserine/phosphatidylcholine), 10 nM FXa, 0.5 μM prothrombin and 16 pM FVa/FVi. FVa-degradation was also followed over time using a reaction mixture of 30 pM APC, 0.8 nM FVa and 56 × 106/mL platelets/MPs, with or without 100 nM

Western blotting following FVa-degradation was performed with affinity-purified FVa, as described [42]. Samples from FVa-degradation were separated in a 4-15% gel and transferred to a PVDF-membrane using pre-packed transfer stacks (Trans-Blot Turbo Bio-Rad), in a Trans Blot Turbo device (Bio-Rad) (30 minutes, 1A, 25 V). FVfragments were visualized using mouse-anti-FV antibody (AHV-5146, Haematologic Thechnologies) followed by horseradish peroxidase conjugated goat-anti-mouse antibody (DAKO). Membranes were developed using enhanced chemiluminescence (ECL) substrate in Chemdoc XRS (Bio-Rad) CCD-camera and the bands were quantified with software ImageLab 4.0.1 (Bio-Rad). Platelet protein S was analyzed by immunoblotting and ELISA. Total protein S was detected using samples of freshly isolated washed resting or activated platelets (75 × 106/ml). The resting/activated platelets/MPs were centrifuged (20 000 × g, 15 min) and membrane-associated or free protein S was detected in the resuspended pellet or supernatant. Samples were mixed with denaturing reducing sample buffer or diluted in ELISA buffer (50 mM Tris-HCL, 150 nM NaCL, 0.5 % tween20, pH 7.5). Electrophoresis was performed in 4-15 % gel and protein S was identified by immunoblotting with polyclonal (DAKO) or monoclonal (HPS54) anti protein S and ECL. The concentrations of total, cellassociated and free protein S were determined by ELISA, using polyclonal rabbit anti-protein S (DAKO) as catcher and an in house monoclonal mouse-anti-protein S (HPS54) [43] for detection. Protein S of known concentration was used to obtain standard curve. Thrombin generation assay Platelets/MPs (7.5 × 106/mL) were pre-incubated with 8.75 pM tissue factor ((TF) (Dade Innovin, Siemens, Erlangen, Germany)) 1 h, 37 °C. Citrated platelet poor plasma donated by healthy volunteers (80 μL diluted ½ in HNBSA) was pre-incubated with APC (10 μL, 130 nM) for 10 minutes at 37 °C. Platelets/MPs/TF (20 μL) was added to the plasma/APC mix (total volume 90 μL) and the reaction was started by addition of 20 μL fluorogenic thrombin substrate (2 mM Z-Gly-GlyArg-AMD, Bachem, Bubendorf, Switzerland) and 108 mM CaCl2. Final

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

4

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

concentrations: 1.4 pM TF, 300 μM substrate, 16.7 mM CaCl2, 10 nM APC and 6.4 × 106 platelets/MPs/mL. Neutralizing antibodies against protein C (DAKO) and protein S (in house HPS21) were incubated in diluted plasma with our without APC at a concentration of 100 μg/mL. All reagents were in HNBSA. Formation of thrombin was monitored with accumulated fluorescence in Infinite 200 microplate reader at excitation wavelength 360 nm and emission wavelength 460 nm. Data was collected every 60 s for 1 hr, 37 °C. The output data was analyzed using GraphPad Prism 5.0 (GraphPad Software Inc, La Jolla, CA, USA) and the accumulative fluorescence was transformed to substrate conversion rate by derivatizing the obtained data. Statistical methods In the figures, standard error of the mean (SEM) has been used as a measure of statistical dispersion. Results Procoagulant efficiency after platelet activation with different agonists The abilities of resting or activated platelets to support assembly of procoagulant complexes were investigated using Xase and the PTase assays (Fig. 1). Platelets activated with A23187 ± 0.5 U/mL thrombin yielded highest PTase activity, whereas the PTase activity of A23187 + 5 U/ml thrombin consistently were lower, an observation we however did not investigate further. Intermediate PTase activity was obtained using platelets activated with 5 U/mL thrombin or thrombin/collagen, whereas resting platelets gave lowest PTase activity. In the Xase assay, comparable results were obtained when activated or resting platelets were used as surface for the assembly of FVIIIa and FIXa into the Xase complex. Thus, A23187 ± thrombin yielded the most efficient Xase activity, followed by 0.5 or 5 U/ml thrombin plus collagen or 5 U/mL thrombin, whereas resting platelets and platelets treated with 0.5 U/mL thrombin poorly supported Xase activity. Phosphatidylserine on microparticles binding lactadherin and protein S To correlate the procoagulant efficiency with the formation of MPs, resting and activated platelets were analyzed by flow cytometry. Platelets and MPs could be discriminated using side and forward scatter in combination with phosphatidylserine exposure, as measured by binding of lactadherin and protein S (Fig. 2). The low forward scatter (platelets mean signal 42 vs. MPs mean signal 9) of the lactadherin/protein S positive particles suggested small size and high phosphatidylserine exposure, which characterize MPs. Therefore, exposure of phosphatidylserine was used to define the MP gate. Resting platelets were lactadherin/protein S-negative, whereas essentially all particles became lactadherin/protein S-positive after activation with A23187, regardless of the concentration of thrombin used. Thrombin alone generated MPs the fraction being dependent on the thrombin concentration used. Thrombin at 0.5 and 5 U/mL resulted in approximately 3% and 40% MPs, respectively. Inclusion of collagen further increased formation of MPs, leading to 42% or 75% MPs in presence of 0.5 or 5 U/mL thrombin, respectively. The total numbers of CD41a-positive particles were determined before and after activations, and surprisingly no differences could be detected. Several attempts to differentiate MPs from activated platelets by differential centrifugation failed to result in a MP-preparation of sufficient concentration for the following experiments. Therefore, based on the flow cytometry results in combination with the Xase and PTase experiments, we decided to use MPs generated by A23187 + 5 U/mL thrombin as model in the following experiments. Platelet alpha-granules contain protein S, the doublet band on reduced gels suggesting it to be partially proteolyzed (Fig. 3). The protein S concentration in 75 × 106/mL platelets was found to be approximately 0.7 nM (Table 1). Protein S was released after activation with thrombin

Fig. 1. Xase- and PTase- complexes on activated platelets/MPs. Platelets were activated with 0.5 or 5 U/mL thrombin or 0.5 or 5 U/mL thrombin plus 25 μg/mL collagen or 5 μM A23187 or 5 μM A23187 + 0.5 or 5 U/mL thrombin. A) The activity of the Xase complex was tested in presence of the platelets/MPs (24 × 106/mL) by incubating FVIIIa (212 mU/mL), FIXa (5 nM) in buffer containing CaCl2 (3 mM), 37 °C, 2.5 minutes. FX was added (to 0.5 μM) and after 3 minutes incubation, the concentration of formed FXa was measured by synthetic substrate. Standard curves for thrombin or FXa, were used to obtain the concentrations. B) To test the activity in the PTase complex, activated platelets/MPs (14 × 106/mL) were incubated with FXa (5 nM) and prothrombin (0.5 μM) in buffer containing CaCl2 (3 mM) for 2 minutes at 37 °C. The concentration of generated thrombin was measured using chromogenic substrate. Data presented as mean and SEM (error bars) of 3-4 individual experiments.

and thrombin/collagen and recovered in the supernatant after centrifugation, the unchanged doublet pattern on reduced gels did not indicate further proteolysis. In contrast, after A23187/thrombin activation, the degree of protein S proteolysis increased, presumably due to the high thrombin concentration. After A23187/thrombin only a minor fraction of protein S appeared in the supernatant, the majority of protein S being bound to the MPs. However, this endogenous protein S was not detected by the labeled anti-protein S antibodies in the flow cytometry experiment, the most likely reason being the low concentration of the endogenous protein S, as compared the amount of exogenous protein S required to give a signal in the flow experiment. It was noteworthy that MPs generated by A23187/thrombin from western blot and ELISA data appeared to bind endogenous protein S more efficiently than those generated by collagen/thrombin even though the MPs were equally efficient in binding exogenous protein S and lactadherin. The reason for this is unknown but may suggest that more phosphatidylserine was exposed on the surface of MPs generated by A23187/thrombin than those resulting from collagen/thrombin. The protein S that was associated with the MPs could be eluted by EDTA treatment, suggesting that the protein S binding was calcium-dependent. We consistently

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

5

Fig. 2. Exposure of phoshatidylserine and binding of protein S to platelet-derived microparticles. Resting or activated platelets (0.5 or 5 U/mL thrombin, 0.5 or 5 U/mL thrombin plus 25 μg/mL collagen, or 5 μM A23187 or 5 μM A23187 + 0.5 or 5 U/mL thrombin) were labeled with the combination of anti-CD61-PE and lactadherin-FITC (160 nM), or with CD61-PE and protein S (180 nM) (detected with anti-protein S-Alexa488) and submitted to flow cytometry analysis. CD61-positive events were collected (10 000) and the fluorescence in FLI (log) (lactadherin or protein S) was plotted against the forward scatter (log) signal. Forward (log) and side scatter (log) data is shown in the upper panel. Data presented from one representative experiment.

noted that resting platelets released some protein S possibly due to some activation/distress by the isolation procedure. The amount of leakage was similar directly after isolation and after storage for several hours (results not shown), demonstrating that the leakage was not timedependent. Since FV/FVa is also released during platelet activation, plateletderived FVa could be used as a source of FVa in pro- and anticoagulant processes. Therefore, we performed a PTase assay to estimate the contribution of FV/FVa from the platelets. We found that upon activation of 75 × 106 platelets/mL with ionophore, the FV concentration was 0.4 ± 0.1 nM (n = 5). The other activation methods released

similar amounts of FV/FVa. The endogenous concentration of FV/ FVa was thus sufficient to provide substrate for APC in the FVadegradation assay. We noted that the degradation curves for inhibition of endogenous FVa and the combination of endogenous plus added plasma-purified FVa were relatively similar (see below) and we therefore decided to add purified FVa in the further characterization. APC-mediated degradation of FVa on the microparticle surface The degradation of FVa by APC was studied using MP generated by A23187 + 5 U/mL thrombin. Under the conditions used, the relative rate of FVa degradation was independent of the FVa concentration (data not shown), and thus variations in the contribution of plateletderived FVa did not affect the results. After initial experiments using increasing concentrations of APC with and without protein S (not shown), 30 pM APC was chosen for time-course FVa-degradation (Fig. 4). APC alone yielded a relatively rapid loss of FVa activity during the first 10 minutes, which according to the western blotting corresponded to the Arg506 cleavage. Addition of anti-protein C blocked the cleavage and FVa activity remained relatively stable during the experiment.

Table 1 Protein S concentration in activated platelet preparations measured by ELISA. Platelets (75 × 106/mL) were prepared from three individual donors and separate aliquots were activated with 0.5 U/mL thrombin, 0.5 U/mL thrombin plus 25 ug/mL collagen, or 5 uM A23187 + 5 U/mL thrombin. Samples were drawn from the different mixtures to estimate the total concentration of protein S in each aliquot. Thereafter, the remaining platelet mixtures were centrifuged and the supernatant and the resuspended pellets (to original volume in ELISA buffer) were collected and their content of protein S determined with ELISA. The mean protein S concentration (nM) and range are given. Platelet treatment

Fig. 3. Release of endogenous platelet protein S and binding to microparticles. Platelets (75 × 106/mL) were activated with the different agonist combinations: 0.5 U/mL thrombin or 0.5 U/mL thrombin plus 25 ug/mL collagen or 5 uM A23187 + 5 U/mL thrombin. Aliquots were drawn before and after activation, where after the platelet mixtures were centrifuged (20 000 × g). Equal volumes of the starting sample (corresponding to 0.375 × 106 platelets/lane), the supernatants and the dissolved pellets (resuspended to original volumes) were applied to reduced (+DTT (dichlorodiphenyltrichloroethane) or unreduced 4-15% SDS-PAGE followed by immunoblotting. In the right panel, 2 mM EDTA was added after the A23187/thrombin activation but before the centrifugation. Protein S was detected with a polyclonal antibody and visualized by enhanced chemiluminescence. T = total protein S before centrifugation but after activation, S = protein S in supernatant, P = protein S in pellet.

No agonist IIa IIa/collagen A23187/IIa

Protein S concentration, nM Total

Released¶

Cell/MP associated§

0.74 (0.33-1.14) 0.64 (0.31-1.03) 0.48 (0.23-0.67) 0.70 (0.28-1.14)

0.33 (0.18-0.41) 0.58 (0.24-0.94) 0.48 (0.18-0.79) 0.12 (b0.01-0.24)

0.30 (0.11-0.57) 0.06 (0.02-0.10) 0.04 (0.01-0.06) 0.68 (0.24-1.34)

¶

Protein S in supernatant after centrifugation (20 000 × g). Protein S in pelleted cells/MPs after centrifugation (20 000 × g) and resuspension to original volume. §

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

6

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

APC-mediated inhibition of FVIIIa in the presence of microparticles To evaluate the ability of APC to down-regulate the Xase complex, FIXa and FVIIIa assembled on MPs were incubated with APC in the presence or absence of protein S and intact FV. The remaining FVIIIa activity was measured in a Xase assay. To exclude platelet-FV as a potential source of APC-cofactor activity, the A23187-treated platelets were preincubated with thrombin (5U/mL) to activate platelet FV. APC alone or with FV were ineffective. In contrast, in the presence of FV plus protein S the remaining FVIIIa-activity was only 18 ± 5% (n = 4) at the highest APC-concentration, whereas corresponding number was 50 ± 5% (n = 3) when protein S was the only APC cofactor added (Fig. 5). Control experiments with MPs generated with A23187 alone or with A23187 + 0.5 U/mL thrombin yielded similar results as those presented in Fig. 5 (not shown). APC-mediated inhibition of thrombin generation in the presence of microparticles

Fig. 4. Stimulation by protein S of APC-mediated FVa degradation on microparticles. A) Microparticles (56 × 106/mL) generated by A23187 + 5U/mL thrombin were incubated with APC (30 pM) ± FVa (0.8 nM), protein S (100 nM), anti-protein C, and anti-protein S at 37 °C. At intervals, aliquots were drawn and diluted 1/10 in ice-cold buffer and remaining FVa activity determined, mean ± SEM (error bars) being presented, (n = 3).▼, anti-protein C; ○, anti protein S; ▲, APC only; ■ protein S; dashed (+protein S) and dotted (- protein S) lines represent degradation of endogenous FVa. B) Affinity-purified FVa was subjected to APC-mediated degradation as in A and at intervals, the FVa-degradation was stopped by dilution in equal volume denaturing, reducing buffer and the FV-fragments were separated on a 4-15% SDS-PAGE gel and transferred to a blotting membrane. FVa fragments were detected with anti-FV (AHF-5146 with epitope between 307-506) and visualized with enhanced chemiluminescense. The peptide-composition of the FVa fragments is denoted to the right.

Addition of protein S resulted in a 4-fold more rapid loss of FVa activity due to the stimulation of protein S of the Arg306 cleavage resulting in the generation of 307-506 and 307-679/709 fragments. Addition of protein S antibodies did not affect the FVa-degradation in absence of protein S, suggesting there was no endogenous protein S effect (Fig. 4); in the control, the antibodies completely inhibited the effect of added protein S (data not shown). The calculated kinetic rate constants are listed in Table 2. In the illustrated experiment, the endogenous FVa contributed around 10% of the total FVa activity and therefore had little impact on the calculated parameters. In the figure, the absolute concentrations of thrombin are plotted which is why the endogenous FVa curves are low. However, when the relative rates of degradation of the endogenous FVa were analyzed, they were similar to that of exogenous FVa (results not shown). Furthermore, FVa-degradation curves obtained with MPs generated with A23187 alone or with A23187 + 0.5 U/mL thrombin yielded similar results as those presented in Fig. 4 (not shown).

To study the extent to which the MPs can support anticoagulant processes in plasma, a thrombin generation assay was used. Plasma was spiked with MPs, and the thrombin generation in plasma was followed over 60 minutes in the presence or absence of APC and antibodies neutralizing the APC or protein S effects (Fig. 6). Thrombin generation was decreased in a dose dependent manner upon addition of increasing concentrations of APC (Fig. 6A). Inclusion of a protein C antibody did not alter the amount thrombin formed in absence of APC, suggesting lack of activation of endogenous protein C in the plasma. When titrating APC in the presence of antibodies against protein S, the APC effect was abolished (Fig. 6B). Using a protein S antibody in absence of APC showed that protein S has an APC-independent inhibitory function (Fig. 6C). Discussion Platelet-derived MPs offer a suitable surface for binding and assembly of procoagulant proteins, which at sites of vascular damage may be beneficial for efficient hemostasis. However, in circulating blood these properties may potentially be harmful and increased concentrations of MPs have been suggested to increase the risk for thrombosis [44]. The anticoagulant function of the protein C system depends on negatively charged surfaces and may therefore be able to regulate coagulation on MPs. Others have previously shown that APC can inhibit platelet FVa on activated platelets/MPs, although less efficiently than on synthetic phospholipid vesicles [5]. The observation that protein S binds to platelet-derived MPs, but not to activated platelets, supports its participation in anticoagulant reactions on MPs [33]. We now confirm that

Table 2 FVa cleavage rates by APC in presence or absence of protein S. Pseudo first-order rates for APC –mediated cleavage of FVa on the surface of A23187 + 5 U/mL thrombingenerated platelet-derived microparticles. Rate constants for cleavage at Arg506 (k506) and Arg306 (k306) presented as mean ± SD of 3 individual experiments. Cleavage site in FV

k506 k306

FVa inactivation (M-1 s-1) +Protein S

-Protein S

8.9 ± 3.6 × 108 3.6 ± 1.6 × 107

3.2 ± 0.8 × 108 1.5 ± 0.4 × 107

Fig. 5. APC-mediated inhibition of Xase on microparticles is stimulated by protein S and FV. MPs (24 × 106/mL, platelets activated with 5 μM A23187 + 5 U/mL thrombin) were incubated with FVIIIa (212 mU/mL), FIXa (5 nM), APC (0-5 nM), with or without protein S (33 nM) and/or FV (2 nM) at 37 °C for 2.5 minutes. FX was added (to 0.5 μM) and after 3 minutes, the activity of formed FXa was measured by conversion of a synthetic colorimetric substrate. Data presented as mean ± SEM, n = 3-4.

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

Fig. 6. Activated protein C-dependent inhibition of thrombin generation in normal plasma is stimulated by protein S. Thrombin generation in plasma spiked with 6.4 × 106 MPs/mL (platelets activated with 5 μM A23187 + 5 U/mL thrombin), 1.4 pM tissue factor, 0-20 nM APC A) in presence or absence of anti-protein C antibody (100 μg/mL), B) in presence of anti protein S (HPS54, 100 μg/mL). C) Thrombin generation assay as described above but in absence of APC. The accumulated fluorescence from the thrombin sensitive substrate Z-Gly-Gly-Arg-AMD (300 μM) was monitored and presented as the first derivative representing thrombin activity. The experiment was repeated three times and data from one representative experiment are shown. FU = fluorescence units.

MPs can support FVa inhibition by APC and extends the knowledge by demonstrating that APC inhibits plasma coagulation on MPs in a protein S dependent manner. Furthermore, the calculated rate constants for the FVa cleavages suggest that APC-mediated cleavages of plasma-derived FVa on MPs are as efficient as reported for phospholipid vesicles [42, 45,46] and finally that protein S and FV function as synergistic cofactors on MPs in the inhibition of FVIIIa. We hypothesize that under normal conditions, circulating platelet MPs are kept in an anticoagulant state by the protein C system. MPs of different cellular origin are usually isolated by differential centrifugation [47]. However, due to the small size of the platelets and as they are anucleated, a relatively high centrifugation force is needed to pellet the activated platelets, which dramatically reduces the yield of MPs. The adhesive nature of activated platelets and platelet-derived MPs presumably also contribute to the poor yield. Therefore, we took the approach to test different platelet agonists to find the most efficient way to generate MPs [4,5]. We found that when platelets were activated with A23187/thrombin, an essentially homogenous MP population was formed, which we used as model of platelet-derived MPs. Quantitation of the number of particles before and after activation revealed no significant difference. This is surprisingly and unexpected as each platelet is expected to generate more than one microparticle and we have no explanation for the finding. Possibly smaller particles are generated that are below the limit of detection in the flow cytometer. The separation of MPs from activated platelets in flow cytometry is demanding as the resolution of forward and side scatter for small platelets and MPs is poor. The new Beckman Coulter Gallios offers an angle on forward scatter (W2) that improves the resolution of small size particles [48]. MPs were defined using phosphatidylserine exposure. Based on the phosphatidylserine exposure we could conclude that these in general

7

were smaller (lower forward scatter signal) than the phosphatidylserine negative non-activated platelets. To ensure correct gating, we stimulated washed platelets with A23187/thrombin and isolated the MPs by centrifugation. Although the yield of MPs was low, the forward/side scatter characteristics of the population did not change after centrifugation, whereas centrifugation of resting platelets led to an almost complete loss of events (data not shown). Thus we concluded that the centrifuged MPs correctly defined the MP gate. Binding of endogenous platelet protein S to MPs could not be detected by flow cytometry although protein S was detectable on pelleted MPs by western blotting. This discrepancy was presumably due to low concentration of endogenous protein S. The protein S binding pattern was mimicked by the lactadherin binding, indicating the importance of phosphatidylserine exposure for protein S binding. The endogenous protein S appeared as double bands on immunoblots, suggesting that protein S was partially proteolyzed already in resting platelets, possibly by an unknown platelet protease [49]. After activation with A23187/ thrombin the amount of proteolyzed protein S increased but most of it still bound to the MPs. After proteolysis, protein S functions poorly as APC-cofactor suggesting that endogenous platelet protein S after activation may have low activity in addition to being at low concentration. The capacity of the MPs to bind endogenous protein S varied between the activation methods, MPs generated by A23187 being most efficient in binding endogenous protein S. The reason for this discrepancy is not known but could possibly be due to higher surface concentration of phosphatidylserine after A23187 compared to thrombin ± collagen. However, importantly for the presented data, the MPs generated with different agonists were equally efficient in binding exogenous protein S. We could not detect any differences in the size of the MPs generated by the different agonists (forward scatter flow cytometry). However, it is possible that the density, and sedimentation rate of different MPs could vary. A combination of low concentration of MPs and poor sedimentation efficiency of small particles could result in false low protein S signal in the pellet fraction of thrombin and thrombin plus collagen treated platelets. However, our purpose was to study the effect of plasma protein S and as platelet-derived protein S did not affect the results, we did not further study the properties of the endogenous platelet protein S. Our results emphasize the importance of plasma protein S for the APC function on MPs. In the FVIIIa degradation, the presence of added protein S was required for APC to function, whereas the addition of FV further enhanced the inhibition of FVIIIa. The degradation of FVa is a biphasic reaction where the first fast phase is caused by rapid cleavage of Arg506. The addition of protein S not only resulted in the stimulation of the Arg306 cleavage similarly to what has previously been demonstrated on phospholipid vesicles [45] but also the Arg506 cleavage was faster in the presence of protein S. The calculated rate constants for degradation of FVa on MPs were found to be equal or higher than those previously reported using synthetic phospholipid vesicles [45,50], which indicates that the MP-surface is highly efficient in supporting APCmediated degradation of FVa. This is not in agreement with the report by Tans et al. [5] that suggested FVa-degradation on ionophorestimulated platelets to be slower than on phospholipid vesicles. A possible explanation may be that in our study the platelets were completely converted to MPs, whereas the ionophore stimulation of Tans et al. may have resulted in less MPs and more activated platelets. Moreover, it has been shown that platelet-derived FVa is relatively resistant to APCmediated degradation on the surface of activated platelets or MPs, compared to plasma-derived FVa on phospholipid vesicles [29–31]. We could not detect any endogenous protein S effect in FVa-degradation, possibly due to low concentration or proteolyzed platelet-derived protein S. This study was primarily designed to follow the degradation of exogenous FVa however endogenous platelet FVa was also susceptible to APC-mediated degradation. Considering the high concentration of FV in blood and the high FV-binding ability of MPs, this may reflect the situation in vivo.

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

8

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx

In the thrombin generation assay, the inhibitory effect of APC was totally dependent on the presence of plasma protein S. In addition, protein S had an APC independent role, which may be due to the reported direct APC-independent anticoagulant functions of protein S [51–53] or its role as cofactor to tissue factor pathway inhibitor (TFPI) [54,55]. Our results are in agreement with the recent report by Stavenuiter et al. [53] showing that both plasma-derived and platelet-derived protein S have APC-independent direct anticoagulant activity on activated platelets and platelet-derived MPs. In our study, we have not further investigated the direct anticoagulant effects of protein S but rather focused on the APC-cofactor function of protein S. Therefore, the two studies complement each other in demonstrating the importance of protein Sdependent regulatory mechanisms of the coagulation pathway on MPs. In conclusion, this study demonstrates the high potential of the protein C system to down-regulate coagulation on MPs. The presence of protein S and intact FV in plasma could help counteract the procoagulant effects of the MPs by several actions. 1, protein S stimulates the cleavage of FVa by APC. 2, intact FV and protein S synergistically stimulate the cleavage of FVIIIa by APC. 3, the mere presence of relatively high concentrations of both protein S (300 nM) and FV (30 nM) in plasma could outcompete the procoagulant proteins on the MP surface and blunt the procoagulant properties of the MPs. 4, both protein S and FV have been shown to bind TFPIα in plasma [56,57] and may therefore localize TFPIα to the MP surface. This may contribute to additional regulation of coagulation to MPs carrying TF. 5, both platelet-derived and plasmaderived protein S have direct, APC- and TFPI-independent anticoagulant activity on MPs [51–53]. Several diseases are associated with increased concentrations of circulating MPs, which may represent a potential thrombosis risk. The experiments in this study were performed with platelet/MP-concentrations ranging from 6.4 × 106 to 56 × 106/mL, which is in line with the elevated concentrations (30-100 × 106/mL) reported in patients with autoimmune diseases [26]. The presented data suggests that the protein C system, including circulating protein S and FV, could blunt the threat of acquired thrombus formation. Even though the concentration of APC in plasma normally resides in the pM range, the concentration could be considerably higher at sites of active coagulation. Therefore, when evaluating the potential risk of MPs it is important to consider the anticoagulant abilities of the MPs. In this context, it is tempting to speculate that patients with hereditary thrombophilia or acquired protein S deficiency could have substantially higher risk to develop complications due to increased concentration of circulating MPs. Conflict of interest statement The authors declare no competing financial interests. Acknowledgements This work was supported by Swedish Research Council (grant 71430); grants from the Swedish Heart and Lung Foundation, Söderberg’s Foundation, the Alfred Österlund’s Foundation, and the University Hospital in Malmö. References [1] Inal JM, Ansa-Addo EA, Stratton D, Kholia S, Antwi-Baffour SS, Jorfi S, et al. Microvesicles in health and disease. Arch Immunol Ther Exp 2012;60:107–21. [2] Burnier L, Fontana P, Kwak BR, Angelillo-Scherrer A. Cell-derived microparticles in haemostasis and vascular medicine. Thromb Haemost 2009;101:439–51. [3] Horstman LL, Ahn YS. Platelet microparticles: a wide-angle perspective. Crit Rev Oncol Hematol 1999;30:111–42. [4] Sims PJ, Wiedmer T, Esmon CT, Weiss HJ, Shattil SJ. Assembly of the platelet prothrombinase complex is linked to vesiculation of the platelet plasma membrane. Studies in Scott syndrome: an isolated defect in platelet procoagulant activity. J Biol Chem 1989;264:17049–57. [5] Tans G, Rosing J, Thomassen MC, Heeb MJ, Zwaal RF, Griffin JH. Comparison of anticoagulant and procoagulant activities of stimulated platelets and platelet-derived microparticles. Blood 1991;77:2641–8.

[6] Hoffman M, Monroe DM, Roberts HR. Coagulation factor IXa binding to activated platelets and platelet-derived microparticles: a flow cytometric study. Thromb Haemost 1992;68:74–8. [7] Gilbert GE, Sims PJ, Wiedmer T, Furie B, Furie BC, Shattil SJ. Platelet-derived microparticles express high affinity receptors for factor VIII. J Biol Chem 1991;266:17261–8. [8] Ahmad SS, London FS, Walsh PN. Binding studies of the enzyme (factor IXa) with the cofactor (factor VIIIa) in the assembly of factor-X activating complex on the activated platelet surface. J Thromb Haemost 2003;1:2348–55. [9] Tracy PB, Eide LL, Bowie EJ, Mann KG. Radioimmunoassay of factor V in human plasma and platelets. Blood 1982;60:59–63. [10] Rosing J, Tans G, Govers-Riemslag JW, Zwaal RF, Hemker HC. The role of phospholipids and factor Va in the prothrombinase complex. J Biol Chem 1980;255:274–83. [11] van Dieijen G, Tans G, Rosing J, Hemker HC. The role of phospholipid and factor VIIIa in the activation of bovine factor X. J Biol Chem 1981;256:3433–42. [12] Dahlback B, Villoutreix BO. The anticoagulant protein C pathway. FEBS Lett 2005;579:3310–6. [13] Fay PJ, Smudzin TM, Walker FJ. Activated protein C-catalyzed inactivation of human factor VIII and factor VIIIa. Identification of cleavage sites and correlation of proteolysis with cofactor activity. J Biol Chem 1991;266:20139–45. [14] Kalafatis M, Rand MD, Mann KG. The mechanism of inactivation of human factor V and human factor Va by activated protein C. J Biol Chem 1994;269:31869–80. [15] Dahlback B. Factor V, and protein S as cofactors to activated protein C. Haematologica 1997;82:91–5. [16] Dahlback B, Villoutreix BO. Regulation of blood coagulation by the protein C anticoagulant pathway: novel insights into structure-function relationships and molecular recognition. Arterioscler Thromb Vasc Biol 2005;25:1311–20. [17] Dahlback B, Stenflo J. High molecular weight complex in human plasma between vitamin K-dependent protein S and complement component C4b-binding protein. Proc Natl Acad Sci U S A 1981;78:2512–6. [18] Maurissen LF, Thomassen MC, Nicolaes GA, Dahlback B, Tans G, Rosing J, et al. Reevaluation of the role of the protein S-C4b binding protein complex in activated protein C-catalyzed factor Va-inactivation. Blood 2008;111:3034–41. [19] Cramer TJ, Gale AJ. The anticoagulant function of coagulation factor V. Thromb Haemost 2012;107:15–21. [20] Nicolaes GA, Dahlback B. Factor V and thrombotic disease: description of a janusfaced protein. Arterioscler Thromb Vasc Biol 2002;22:530–8. [21] Schwarz HP, Heeb MJ, Wencel-Drake JD, Griffin JH. Identification and quantitation of protein S in human platelets. Blood 1985;66:1452–5. [22] Dahlback B. Advances in understanding pathogenic mechanisms of thrombophilic disorders. Blood 2008;112:19–27. [23] Grant R, Ansa-Addo E, Stratton D, Antwi-Baffour S, Jorfi S, Kholia S, et al. A filtrationbased protocol to isolate human plasma membrane-derived vesicles and exosomes from blood plasma. J Immunol Methods 2011;371:143–51. [24] Pereira J, Alfaro G, Goycoolea M, Quiroga T, Ocqueteau M, Massardo L, et al. Circulating platelet-derived microparticles in systemic lupus erythematosus. Association with increased thrombin generation and procoagulant state. Thromb Haemost 2006;95:94–9. [25] Varon D, Shai E. Role of platelet-derived microparticles in angiogenesis and tumor progression. Discov Med 2009;8:237–41. [26] Sellam J, Proulle V, Jungel A, Ittah M, Miceli Richard C, Gottenberg JE, et al. Increased levels of circulating microparticles in primary Sjogren's syndrome, systemic lupus erythematosus and rheumatoid arthritis and relation with disease activity. Arthritis Res Ther 2009;11:R156. [27] Omoto S, Nomura S, Shouzu A, Nishikawa M, Fukuhara S, Iwasaka T. Detection of monocyte-derived microparticles in patients with Type II diabetes mellitus. Diabetologia 2002;45:550–5. [28] Mallat Z, Benamer H, Hugel B, Benessiano J, Steg PG, Freyssinet JM, et al. Elevated levels of shed membrane microparticles with procoagulant potential in the peripheral circulating blood of patients with acute coronary syndromes. Circulation 2000;101:841–3. [29] Magdeleyns EJ, Keuren JF, Curvers J. Factor Va, bound to microparticles released during platelet storage, is resistant to inactivation by activated protein C. Transfusion 2007;47:1880–8. [30] Gould WR, Silveira JR, Tracy PB. Unique in vivo modifications of coagulation factor V produce a physically and functionally distinct platelet-derived cofactor: characterization of purified platelet-derived factor V/Va. J Biol Chem 2004;279:2383–93. [31] Camire RM, Kalafatis M, Simioni P, Girolami A, Tracy PB. Platelet-derived factor Va/ Va Leiden cofactor activities are sustained on the surface of activated platelets despite the presence of activated protein C. Blood 1998;91:2818–29. [32] Taube J, McWilliam N, Luddington R, Byrne CD, Baglin T. Activated protein C resistance: effect of platelet activation, platelet-derived microparticles, and atherogenic lipoproteins. Blood 1999;93:3792–7. [33] Dahlback B, Wiedmer T, Sims PJ. Binding of anticoagulant vitamin K-dependent protein S to platelet-derived microparticles. Biochemistry 1992;31:12769–77. [34] Dahlback B. Human coagluation factor V purification and thrombin-catalyzed activation. J Clin Invest 1980;66:583–91. [35] Fujikawa K, Legaz ME, Davie EW. Bovine factors X 1 and X 2 (Stuart factor). Isolation and characterization. Biochemistry 1972;11:4882–91. [36] Nelsestuen GL, Suttie JW. The mode of action of vitamin K. Isolation of a peptide containing the vitamin K-dependent portion of prothrombin. Proc Natl Acad Sci U S A 1973;70:3366–70. [37] Shen L, Shah AM, Dahlback B, Nelsestuen GL. Enhancing the activity of protein C by mutagenesis to improve the membrane-binding site: studies related to proline-10. Biochemistry 1997;36:16025–31.

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031

S. Somajo et al. / Thrombosis Research xxx (2014) xxx–xxx [38] Lundblad RL, Uhteg LC, Vogel CN, Kingdon HS, Mann KG. Preparation and partial characterization of two forms of bovine thrombin. Biochem Biophys Res Commun 1975;66:482–9. [39] Hou J, Fu Y, Zhou J, Li W, Xie R, Cao F, et al. Lactadherin functions as a probe for phosphatidylserine exposure and as an anticoagulant in the study of stored platelets. Vox Sang 2011;100:187–95. [40] Norstrom E, Thorelli E, Dahlback B. Functional characterization of recombinant FV Hong Kong and FV Cambridge. Blood 2002;100:524–30. [41] Nicolaes GA, Tans G, Thomassen MC, Hemker HC, Pabinger I, Varadi K, et al. Peptide bond cleavages and loss of functional activity during inactivation of factor Va and factor VaR506Q by activated protein C. J Biol Chem 1995;270:21158–66. [42] Sun YH, Tran S, Norstrom EA, Dahlback B. Enhanced rate of cleavage at Arg-306 and Arg-506 in coagulation factor Va by Gla domain-mutated human-activated protein C. J Biol Chem 2004;279:47528–35. [43] Dahlback B, Hildebrand B, Malm J. Characterization of functionally important domains in human vitamin K-dependent protein S using monoclonal antibodies. J Biol Chem 1990;265:8127–35. [44] Owens III AP, Mackman N. Microparticles in hemostasis and thrombosis. Circ Res 2011;108:1284–97. [45] Rosing J, Hoekema L, Nicolaes GA, Thomassen MC, Hemker HC, Varadi K, et al. Effects of protein S and factor Xa on peptide bond cleavages during inactivation of factor Va and factor VaR506Q by activated protein C. J Biol Chem 1995;270:27852–8. [46] Tran S, Norstrom E, Dahlback B. Effects of prothrombin on the individual activated protein C-mediated cleavages of coagulation factor Va. J Biol Chem 2008;283:6648–55. [47] Orozco AF, Lewis DE. Flow cytometric analysis of circulating microparticles in plasma. Cytometry A 2010;77:502–14. [48] Robert S, Lacroix R, Poncelet P, Harhouri K, Bouriche T, Judicone C, et al. Highsensitivity flow cytometry provides access to standardized measurement of smallsize microparticles–brief report. Arterioscler Thromb Vasc Biol 2012;32:1054–8.

9

[49] Mitchell CA, Salem HH. Cleavage of protein S by a platelet membrane protease. J Clin Invest 1987;79:374–9. [50] Norstrom EA, Steen M, Tran S, Dahlback B. Importance of protein S and phospholipid for activated protein C-mediated cleavages in factor Va. J Biol Chem 2003;278:24904–11. [51] Fernandes N, Mosnier LO, Tonnu L, Heeb MJ. Zn(2)(+) -containing protein S inhibits extrinsic factor X-activating complex independently of tissue factor pathway inhibitor. J Thromb Haemost 2010;8:1976–85. [52] Heeb MJ, Prashun D, Griffin JH, Bouma BN. Plasma protein S contains zinc essential for efficient activated protein C-independent anticoagulant activity and binding to factor Xa, but not for efficient binding to tissue factor pathway inhibitor. FASEB J 2009;23:2244–53. [53] Stavenuiter F, Davis NF, Duan E, Gale AJ, Heeb MJ. Platelet protein S directly inhibits procoagulant activity on platelets and microparticles. Thromb Haemost 2013;109:229–37. [54] Hackeng TM, Sere KM, Tans G, Rosing J. Protein S stimulates inhibition of the tissue factor pathway by tissue factor pathway inhibitor. Proc Natl Acad Sci U S A 2006;103:3106–11. [55] Sere KM, Rosing J, Hackeng TM. Inhibition of thrombin generation by protein S at low procoagulant stimuli: implications for maintenance of the hemostatic balance. Blood 2004;104:3624–30. [56] Castoldi E, Simioni P, Tormene D, Rosing J, Hackeng TM. Hereditary and acquired protein S deficiencies are associated with low TFPI levels in plasma. J Thromb Haemost 2010;8:294–300. [57] Vincent LM, Tran S, Livaja R, Bensend TA, Milewicz DM, Dahlback B. Coagulation factor V(A2440G) causes east Texas bleeding disorder via TFPIalpha. J Clin Invest 2013;123:3777–87.

Please cite this article as: Somajo S, et al, Protein S and factor V in regulation of coagulation on platelet microparticles by activated protein C, Thromb Res (2014), http://dx.doi.org/10.1016/j.thromres.2014.04.031