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Published as doi: 10.1096/fj.06-7238com.
 (The FASEB Journal. 2007;21:1926-1933.)
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Stimulation of P2 (P2X7) receptors in human dendritic cells induces the release of tissue factor-bearing microparticles

Marcello Baroni*,1, Cinzia Pizzirani{dagger},1, Mirko Pinotti*, Davide Ferrari{dagger}, Elena Adinolfi{dagger}, Sara Calzavarini*, Pierpaolo Caruso*, Francesco Bernardi* and Francesco Di Virgilio{dagger},2

* Departments of Biochemistry, and

{dagger} Experimental and Diagnostic Medicine, and Interdisciplinary Center for the Study of Inflammation, University of Ferrara, Ferrara, Italy

2Correspondence: Department of Experimental and Diagnostic Medicine, Section of General Pathology, Via Borsari 46, 44100 Ferrara, Italy. E-mail: fdv{at}unife.it


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Receptors for extracellular nucleotides are the focus of increasing attention for their ability to cause release of plasma membrane vesicles (microparticles, MPs). Here, we show that monocyte-derived human dendritic cells (DCs) stimulated with a P2X7 receptor (P2X7R) agonist undergo a large release of MPs endowed with procoagulant activity. Functional and Western blot studies revealed that MPs contain the membrane-bound form of tissue factor (TF), a glycoprotein acting as essential cofactor of activated factor VII and triggering blood coagulation. Quiescent DCs express the membrane-bound (full length), as well as truncated alternatively spliced TF forms. DC reactivity to anti-TF Abs disappeared almost completely on stimulation with ATP or benzoyl ATP (BzATP), as shown by immunoblot and confocal microscopy analysis. Concurrently, TF reactivity and activity appeared in the vesicular fraction, indicating that MPs are important carriers for the dissemination of full-length TF form. Activity of MP-bound TF, comparable to that of relipidated recombinant TF, was dose dependently inhibited by the addition of a specific anti-human TF antibody. We infer that a large fraction of this protein, and its procoagulant potential, are "deliverable" after physiological or pathological stimuli. These findings might have implications for triggering and propagating coagulation in healthy and atherosclerotic vessels.—Marcello Baroni, Cinzia Pizzirani, Mirko Pinotti, Davide Ferrari, Elena Adinolfi, Sara Calzavarini, Pierpaolo Caruso, Francesco Bernardi, and Francesco Di Virgilio. Stimulation of P2 (P2X7) receptors in human dendritic cells induces the release of tissue factor-bearing microparticles.


Key Words: coagulation • inflammation • extracellular ATP • purinergic receptors


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
DENDRITIC CELLS (DCs) were originally identified as antigen-presenting cells critical for antigen capture and processing and for the activation of naive T lymphocytes (1 2 3) . DCs are the focus of increasing attention for their immunomodulatory activity, for their role in resistance to infections and tumors, and in the induction of tolerance toward normal cell constituents (4) . DCs express several P2 receptor (P2R) subtypes, a feature that makes them eminently responsive to extracellular nucleotides, as shown by ATP or UTP-dependent modulation of cytokine production (5 6 7) , up-regulation of chemokine receptors (8) , and stimulation of chemotaxis (9) . Interestingly, sensitivity to extracellular nucleotides changes during DC maturation and in different DCs subpopulations (9 , 10 ).

Nucleotides are present in millimolar concentrations in the cytoplasm and in the nanomolar range in the extracellular milieu under quiescent conditions. Very active and ubiquitous nucleotide-metabolizing enzymes (ecto-nucleotidases) have a key role in keeping the extracellular nucleotide concentration low (11 , 12) . Nucleotides may be released by virtually any cell via lytic and nonlytic pathways and P2R present on the DC surface may act as "sensors" to monitor their extracellular levels. Although very few studies have investigated the actual ATP concentration at inflammatory sites, in vivo observations provide clear evidence for sustained ATP release at foci of tissue injury and inflammation (13) . This may be also relevant for atherosclerosis, as a very recent report shows that ATP secretion by macrophages modulate macrophage adhesion to the endothelium, and thus macrophage recruitment to the atheroma (14) .

On the basis of pharmacological, functional, and cloning data, two P2R subfamilies have been so far described: P2YR and P2XR (15 , 16) . P2YR are seven membrane-spanning, G-protein-coupled receptors. Their activation triggers generation of inositol 1,4,5-trisphosphate and release of Ca2+ from intracellular stores. P2YR are ubiquitous, being expressed by monocytes, macrophages, dendritic cells, neurons, smooth and striated muscle cells, as well as epithelial and endothelial cells (17 , 18) . P2XR are plasma membrane channels selective for monovalent and divalent cations that are directly activated by extracellular ATP. These channels were originally identified in mammalian sensory neurons, and subsequently found also in smooth muscle cells, fibroblasts, and immune cells (17 , 19) .

DCs express high levels of a peculiar purinergic receptor subtype, previously known as P2Z (20) and later named P2X7R (21) . The P2X7R differs from the other P2XR for its extended carboxy-terminal domain that endows this receptor with the ability to form large plasma membrane pores permeable to small hydrophilic molecules. An interesting property of the P2X7R pore is its reversibility: removal of ATP triggers resealing of the plasma membrane. Stimulation of P2X7R by BzATP induces emission of IL-1ß-containing microvesicles from the human monocyte cell line THP-1 (22) and from mouse microglial cells (23) .

Circulating microvesicles also referred to as microparticles (MPs) are released into the bloodstream by a variety of cells. Because of the exposure of negatively charged phospholipids, in particular, phosphatidylserine, MPs might provide surfaces that support coagulation. The finding that circulating leukocyte-derived MPs contain tissue factor (TF) further suggests an important procoagulant function of MPs (24 25 26) .

TF is a transmembrane glycoprotein that acts as the cellular receptor and essential cofactor for activated factor VII (FVIIa). At the site of vascular injury, the TF/FVIIa complex triggers coagulation by activating factor IX and X, ultimately resulting in thrombin formation (27) . Moreover, the FVIIa-TF complex may also play a role in the migration and proliferation of vascular smooth muscle cells (28) , in vascular remodeling and in plaque neovascularization (29) and thereby in promoting plaque destabilization.

TF is constitutively expressed in skin, organ surfaces, vascular adventitia, and epithelial–mesenchymal surfaces, where it acts as a sort of hemostatic envelope (30 , 31) . In whole blood (blood-borne TF), TF is present in two forms: a high MW MP-bound, and a low MW-soluble form (asTF) produced by alternative splicing.

Altered expression of TF has implications in several diseases, particularly atherothrombosis, where it is crucial in thrombus formation after plaque rupture. TF is thought to be one of the main determinants of the thrombogenicity of the atherosclerotic plaque, and it has been shown that the enhanced activity of extracellular TF in the lipid core is directly related to the presence of TF-bearing-MPs (32) . Moreover, elevated levels of TF-bearing MPs were found to be associated with cardiovascular disease (33 , 34) . Although many observations indicate the involvement of TF-bearing MPs in atherothrombosis, the cellular and molecular mechanisms responsible for their release are poorly known.

In the present paper, we investigated the release of TF associated to MPs in monocyte-derived DCs stimulated with the P2X7R agonist benzoyl ATP (BzATP).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents
ATP was purchased from Roche (Roche Diagnostics SpA, Monza, Italy); BzATP, benzamidine and phenylmethanesulfonyl fluoride (PMSF) were purchased from Sigma (Sigma-Aldrich, Milan, Italy); EDTA was from Baker (J. T. Baker, Phillipsburg, NJ, USA); Percoll and Ficoll-Paque were obtained from Pharmacia (Pharmacia Biotech AB, Uppsala, Sweden).

DC purification
DCs were obtained from peripheral blood of healthy donors, as described previously (35) . Briefly, monocytes were separated by a two-step Ficoll and Percoll gradient, and then cultured in 2 mM L-glutamine and 25 mM HEPES-containing RPMI 1640 medium (EuroClone, Milan, Italy) complemented with 10% heat-inactivated FBS (Invitrogen, San Giuliano Milanese, Italy), 1 mM sodium pyruvate, 1% nonessential amino acids, 200 U/ml penicillin, 20 µg/ml streptomycin (EuroClone), and 0.05 mM 2-mercaptoethanol (Invitrogen) at 37°C with 5% CO2, in the presence of 200 U/ml IL-4 and 100 ng/ml GM-CSF (Peprotech, Rocky Hill, NJ, USA).

MP purification
To increase MP recovery, DCs were stimulated in a solution containing 300 mM sucrose, 1 mM K2HPO4, 1 mM MgSO4, 5.5 mM glucose, and 20 mM HEPES, 1 mM CaCl2, pH 7.4 with KOH (sucrose solution). After stimulation of the cells with 200 µM BzATP or 3 mM ATP, supernatants were collected and 1 mM EDTA was added. The protease inhibitors benzamidine and PMSF were also added. Floating cells and cell debris were eliminated by centrifuging supernatants for 5 min at 160 g. MPs were purified by centrifugation of cell supernatants at 100,000 g for 90 min, at 4°C. Vesicle pellets were then resuspended in the same solution. In samples used for tissue factor activity measurements, proteases inhibitors were omitted.

Microscopy analysis
DCs were detached from Petri dishes by using 2 mM EDTA-containing cold PBS, and then plated (2x105) onto 24-mm glass coverslips (Merck Eurolabs, Lutterworth, UK). Experiments were performed in the sucrose solution described above. Morphological changes and MP shedding were analyzed by mounting coverslips in a thermostatted Leyden chamber (model TC-202A; Medical Systems Corp., NY, USA), placed onto the stage of an inverted Nikon Eclipse TE300 microscope (Nikon Corp., Tokyo, Japan). The images were captured with a back-illuminated CCD camera (Princeton Instruments, Trenton, NJ, USA) using the Metamorph software (Universal Imaging Corporation, West Chester, PA). DCs for electron microscope analysis were detached from flasks with 2 mM EDTA-containing cold PBS, and cell pellets were fixed with 2.5% glutharaldeide. Samples for electron microscopy were processed by the Centro di Microscopia Elettronica of the University of Ferrara (Ferrara, Italy).

Determination of TF cofactor activity
The MPs suspension (10 µl) was incubated with 2 nM FVIIa (Novo Nordisk, Bagsværd, Denmark) at 37°C for 10 min in the presence of 5 mM CaCl2 (final volume, 30 µl). 10 nM zymogen factor X (FX) (HTI, Essex Junction, VT, USA) and 200 µM of a specific fluorogenic substrate for activated FX (Spectrozyme FXa, American Diagnostic, Stamford, CT, USA) were then added in a final volume of 100 µl. Fluorescence (excitation 360 nm, emission 465 nm) was immediately measured over time through a SpectraFluor Plus microplate reader (Tecan, Austria). The assay was standardized by using serial dilutions (1/7.5–1/480) of Innovin (Dade Bearing, Marburg, Germany) in 20 mM HEPES, 150 mM NaCl, 0.1% PEG 8000, 5 mM CaCl2, 100 µM phospholipid vesicles (PLves 20:80 phosphatidylserine/phosphatidylcoline), pH 7.4. Inhibition of TF activity by a sheep polyclonal anti-human TF antibody (HTI) was assessed in a FXa generation assay. To this purpose, the MPs suspension was preincubated for 20 min at room temperature with increasing concentrations of the antibody (0–6 nM). The residual cofactor activity was expressed as a percentage of activity in the absence of antibody.

Western blot analysis
DCs and MPs were lysed by three freeze-thawing cycles, mixed with a solution containing 60 mM Tris, 2% SDS, 2.5% ß-mercaptoethanol, 10% glycerol, and bromphenol blue and boiled for 5 min. Samples were run on a SDS-PAGE (4–12%, Bio-Rad, Hercules, CA) and then transferred onto a nitrocellulose filter (Protran, Schleicher & Schuell, Dassel, Germany). The filter was incubated with a rabbit polyclonal anti-human TF antibody (1 mg/ml, American Diagnostics, Stamford, CT). Secondary antibody was a HRP-conjugated polyclonal goat anti-rabbit IgG (1 mg/ml, Dako Cytomation, Denmark). Chemiluminescence was detected with the Supersignal West Femto maximum sensitivity substrate (Pierce Biotechnology, Rockford, IL, USA).

Immunocytochemistry and confocal analysis
DCs, previously plated onto 13-mm glass coverslips, were incubated for 10 min at 37°C in a sucrose solution in the presence or absence of 200 µM BzATP. Cells were then rinsed twice with PBS. All of the subsequent procedures were carried out at 4°C, under gentle shaking. Cells were blocked in 5% goat serum containing PBS for 2 min and then incubated for 1 h with 40 µg/ml of rabbit anti-human TF IgG, (American Diagnostics). After 3 washes with PBS (10 min each), samples were incubated with 25 µg/ml goat TRITC-conjugated anti-rabbit IgG (Sigma-Aldrich) for 1 h in the dark, washed 3 times with PBS for 20 min, and fixed for 1 h in 2% paraformaldeyde. Coverslips were then washed 3 times in PBS and mounted onto glass slides in the presence of Pro Long Antifade solution (Molecular Probes, Leiden, The Netherlands). Confocal images were acquired with a Zeiss LSM 510 confocal microscope equipped with a plan-Apochromat 63 x oil immersion objective (Carl Zeiss, Arese, Italy). The 543-nm excitation wavelength was provided by a HeNe laser source. All images were obtained at a 12% laser potency and with a pinhole diameter of 135 µm. Amplifier and detector optimizing parameters were maintained constant for all of the experiments. When required, a single 3D projection of confocal images on the z axis was obtained with the LSM examiner software (Carl Zeiss). Fluorescence emission was quantitated starting from 3D projections with the cell imaging software MetaMorph (Universal Imaging), as described previously (36) . Data were acquired from 10 to 15 cells per coverslip. An average of 10 coverslips was analyzed for each experimental condition. Data are expressed in fluorescence arbitrary units (FU) and shown as mean plus SE. Tests of significance were performed by Student’s t test and ANOVA by means of GraphPad Instat 3.06 software (GraphPad Software, Inc., San Diego, CA, USA).

RT-PCR
RNA was extracted using the Trizol reagent (Invitrogen, Carlsbad, CA) and analyzed by RT-PCR (Access RT-PCR System, Promega, Madison WI, USA) using primers annealing in exon 4 (forward: 5'-GAACGGACTTTAGTCAGAAGG-3') and 6 (reverse: 5'-TGACCACAAATACCACAGCTCC-3') of TF gene. RT-PCR for the house-keeping GAPDH gene (forward: 5'-CCACCCATGGCAAATTCCATGGCA-3' and reverse: 5'-TCTAGACGGCAGGTCAGGTCCACC-3') was also performed. Amplified fragments were separated by electrophoresis on a 3% agarose gel, and visualized with ethidium bromide.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Stimulation of DCs with BzATP induces MP release
Human DCs challenged with 200 µM BzATP released a large amount of MPs (Fig. 1 ). Particle release started within seconds of stimulation, usually at one pole of the cell, and rapidly spread to the whole cell body. Vesicle shedding was preceded by cell contraction and rounding and was paralleled by a striking increase in plasma membrane blebbing and emission of philopodia. Electron microscopy analysis revealed that BzATP stimulation induced dramatic changes in cell morphology and a striking plasma membrane reorganization (Fig. 2 ). Similar results were obtained by stimulation with 3 mM ATP (not shown).


Figure 1
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  		Figure 1. P2X7 receptor stimulation induces cell shrinkage and MP shedding from human dendritic cells (DCs). Cells were seeded onto glass coverslips and stimulated with 200 µM BzATP at 37°C, as indicated in Materials and Methods. Images were acquired at 5-s intervals with the Nikon Eclipse T-300 microscopy set up described in Materials and Methods. Arrowheads indicate shedded MPs. Scale bar = 10 µm.


Figure 2
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  		Figure 2. Electron microscopy of BzATP-stimulated DCs. Cells were stimulated as in Fig. 1 , detached from flasks, fixed with glutharaldeide, and processed for electron microscopy, as described in Materials and Methods. Arrowheads indicate plasma membrane extrusions, blebs, and MPs in the process to be released. Scale bars = 1 µm.

MPs show FVIIa cofactor activity
We then asked whether MPs released by DCs showed procoagulant activity. MPs obtained from different donors were assessed by using a functional assay for TF, based on its property to behave as a cofactor for FVIIa in FXa generation. MPs had a TF-like activity that was dependent on the amount of MPs used in the test (Fig. 3 ).


Figure 3
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  		Figure 3. TF cofactor activity in MPs from DCs. FXa generation was evaluated as relative fluorescence over time and compared to recombinant TF (Innovin) and phospholipid vesicles, as controls. {blacksquare} MPs (13.5 µg), {blacktriangleup} MPs (6.75 µg), {blacktriangledown} MPs (4.5 µg), {Delta} TF (0.6 nM) from Innovin, {square} TF (0.1 nM) from Innovin, x Phospholipid vesicles only.

Activity of MPs was compared to that of Innovin, a well-known reagent for TF-dependent activation of blood coagulation. MP activity ranged between 0.6 and 0.1 nM TF, with a mean value of 38.9 ± 13.8 pM TF/µg MP protein, i.e., 1.2 fmol/µg MP protein, as the assay was run in 30 µl. Because DCs release ~18 ± 7 µg of MPs/106 cells, we calculate an activity of released TF of ~700 pM TF/106 cells, i.e., 21.6 fmol/106 cells. To rule out the possibility that the FXa generation activity was due to MPs phospholipid content, phospholipid preparations devoid of TF were also tested. Under these conditions, no fluorescence increase was observed.

As a further proof of a specific role of MP-associated TF in FXa generation, assays were performed in the presence of increasing concentration of a specific anti-human TF antibody. As shown in Fig. 4 , the antibody was able to reduce, in a dose-dependent manner, the cofactor activity present in the MP preparation.


Figure 4
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  		Figure 4. Inhibition of MP TF cofactor activity by an anti-TF antibody. Activity was expressed as a percentage of FXa generation in the absence of polyclonal anti-human TF antibody (HTI, VT, USA). Duplicate determinations of one of three similar experiments.

Human DCs express the mRNA both for transmembrane and asTF
TF mRNA expression in stimulated and unstimulated DCs was investigated by RT-PCR analysis. To allow the detection of both full-length (transmembrane) and soluble (lacking the exon 5- encoded transmembrane domain) forms of TF transcripts, primers flanking exon 5 were designed. The amplified fragments (307 bp and 147 bp, Fig. 5 ) are compatible with the expression in stimulated or unstimulated human DCs of the full length and soluble (asTF) forms of TF, respectively.


Figure 5
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  		Figure 5. Expression of TF mRNA in DCs. Lane 1, BzATP-treated DCs; lane 2, unstimulated DCs. TF, transmembrane form; sTF, soluble form. M, molecular weight markers.

TF protein is associated to MPs released by DCs.
The presence of TF protein in DCs and MPs was evaluated through Western blot analysis with a specific polyclonal anti-human TF antibody. Immunoreactivity to TF was very high in untreated DCs (Fig. 6 , lane 1), where two major bands compatible with the presence of both forms of TF protein were clearly distinguishable. A faint band with high molecular weight, likely corresponding to TF dimers, was also present. Comparison of the intensity of the bands clearly indicated that the transmembrane form was predominant. Interestingly, the TF immunoreactivity was almost completely lost on stimulation with BzATP (Fig. 6 , lane 2), thus suggesting that TF was released from the cells. In keeping with this hypothesis, the MPs shed from BzATP-stimulated DCs showed an intense immunostaining for the full-length form of TF (Fig. 6 , lane 3). On the contrary, no immunoreactivity for the asTF form was detected on the MPs. These show that a large amount of TF is released from DCs by MP shedding.


Figure 6
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  		Figure 6. Western blot analysis of TF in DCs and MPs. Lane 1, untreated DCs; lane 2, DCs stimulated with 200 µM BzATP for 10 min; lane 3, MPs from BzATP-stimulated DCs. Arrows indicate the membrane-bound (TF) and the soluble (sTF) forms.

Expression of TF on DC plasma membrane is drastically reduced on BzATP stimulation
Changes in TF expression on cell surface were monitored by confocal microscopy. To this purpose, DCs were incubated with a specific TF antibody. Confocal microscopy analysis revealed a punctate, bright fluorescence mainly localized on the plasma membrane and in the peripheral cytoplasm. Fluorescence intensity drastically declined on stimulation with BzATP (Fig. 7 ). DCs incubated in the absence of primary Ab were fully negative, confirming that fluorescence was not due to unspecific binding of the secondary antibody (Fig. 7C ). Panels A–C show a projection on the z axis of the different confocal slices, while panels D–I show single confocal planes (basal, D, G; equatorial E, H; apical, F, I).


Figure 7
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  		Figure 7. Loss of plasma membrane expression of TF on stimulation with BzATP. DCs were layered onto glass coverslips and either left untreated (A, C–F), or stimulated with 200 µM BzATP for 10 min (B, G–I). Both quiescent and BzATP-stimulated cells were then incubated in the presence of an anti-TF antibody and further processed for immunofluorescence analysis as described in Materials and Methods. C) DCs were incubated with the secondary antibody without prior treatment with the anti-TF antibody. A–C show a z axis projection of the different confocal sections, while D–I show a single confocal plane cut through the basal (D, G), equatorial (E, H), or apical (F, I) section of the cells. Images shown are representative of four separate experiments from which at least 15 fields were acquired for each experimental condition.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The crucial role of TF in the initiation of coagulation (27) as obligatory cofactor of FVIIa, in the migration and proliferation of vascular smooth muscle cells (28) , in vascular remodeling, and in plaque neovascularization (29) makes TF expression levels of extreme importance in the control of early steps of the coagulation cascade and in the thrombogenicity of the atherosclerotic plaque. Interestingly, increased levels of circulating TF have been found to be associated with cardiovascular disease (34 , 37) . In addition, the Cys186-Cys209 disulfide bond on the surface of TF might be modified by protein disulfide isomerase (PDI) that inhibits TF thrombogenicity in a nitric oxide-dependent fashion, thus further linking oxidative stress and coagulation (38) .

The procoagulant activity of MPs found in human atherosclerotic plaques (32) and the presence at these sites of DCs, expressing high levels of the P2X7R, suggested to us that ATP released by damaged or infiltrating inflammatory cells might trigger release of TF-bearing MPs via stimulation of the P2X7R. To address this issue, we investigated 1) the ability of human DCs to release MPs on stimulation of the P2X7R and 2) the presence of MP-bound TF.

We found that DCs express the mRNA encoding the full-length form of TF and, to a lesser extent, the alternatively spliced TF mRNA, as found in other cell types of hematopoietic lineage (39) . The presence of both TF forms in DCs was confirmed at the protein level by Western blot analyses. Stimulation of DCs with a potent P2X7 receptor agonist such as BzATP induced a massive release of MPs. The membrane-bound form was found to be mainly associated with the MPs, suggesting that most of asTF is directly released into extracellular space on DC stimulation. TF on MPs was found to be functional as shown by a fluorogenic assay that allowed to quantitatively evaluate TF cofactor activity for FVIIa in the stimulation of the physiological substrate FX.

In our experimental set-up, the addition of DCs-derived MPs produced generation of activated FX, comparable to that triggered by relipidated recombinant TF. Although a much more extensive experimentation is clearly required, these data strongly suggest that MPs are an important source of procoagulant activity specifically due to TF and not to their phospholipid content as phospholipid vesicles lacking TF were fully inactive. In further support of the specific effect of TF-bearing MPs, the procoagulant activity was largely inhibited by an anti-TF antibody.

It is reported that MPs shedding can be triggered by exogenous stimulants or occur spontaneously even from quiescent cells (40) . A higher rate of MP release is observed in apoptotic cells. Among exogenous stimuli, extracellular ATP has been identified as one of the most potent triggers for MP shedding, mainly through activation of the P2X7R, although involvement of other P2 receptors is also likely (22 , 23) . The physiological significance of shed MPs is not entirely clear, and even their biochemical features are still under investigation. However, it is well known that MPs spontaneously shed by activated or apoptotic monocytes or endothelial cells have a high TF content and a high procoagulant activity (40) . Our data show that activation of the P2X7R triggers at the same time a large MP shedding and a massive TF release. Strikingly, analyses of TF bands in DCs before and after treatment with BzATP revealed that the vast majority of TF was lost, mainly through MP shedding. This observation was strongly corroborated by results from immunostaining of TF and confocal microscopy, which showed a punctate immunoreactivity, mainly localized to the membrane/submembrane area. A similar distribution was recently described in rabbit smooth muscle cells and in HEK293 cells transfected with TF-GFP (41) , although in DCs, membrane-associated TF immunoreactivity was higher than in muscle cells or TF transfectants. Our observations provide a clear evidence that activation of the P2X7R can trigger depletion of TF from DCs via shedding of rapidly diffusible MPs. The intrinsic structural components of the MPs, such as phosphatidylserine, favor the assembly of coagulation macromolecular complexes and are likely to participate in the procoagulant activity together with TF, which is absolutely required to sustain the catalytic activity of FVIIa.

These results indicate that conditions characterized by large release of ATP into the extracellular space create a strong procoagulant microenvironment due not only to ADP generation from ATP via CD39L1 (NTPDase2) (42) , but also to ATP-stimulated release of TF-bearing MPs. While it is as yet unknown whether extracellular nucleotides are able to trigger MP release from platelets (the main circulating source of readily releasable ADP and ATP), we show here that they maybe be potentially very important for DCs.

DCs and other mononuclear phagocytes are known to infiltrate the vessel wall and be a major constituent of the atheroma. Platelet aggregation on a dysfunctional endothelium, or at a disrupted plaque, may quickly release large amounts of ATP (this nucleotide is costored with ADP within platelet-dense granules) and generate a propagating wave of procoagulant activity of which TF-bearing MP may be a main constituent. It is well known that MPs are a major determinant of plaque thrombogenicity (32) . TF release from DCs might also be precipitated by factors other than platelet aggregation, as it is increasingly appreciated that ATP is released at sites of inflammation by immune cells via nonlytic pathways (43) . Extracellular ATP might also have an important role in atheroma formation or in its evolution by stimulating cytokine secretion, release of proteases or, as recently shown by Wong and colleagues (14) , by modulating monocyte/macrophage recruitment into the plaque.

Very recently, the mechanism of release of the proinflammatory cytokines IL-1ß and IL-18 has stirred hot interest (44 , 45) . Two nonalternative vesicle-mediated pathways seem to be likely involved: 1) a first one based on atypical secretory lysosomes (46) , and 2) a second one based on the release of MPs (22) . To our knowledge, the possible participation of secretory lysosomes in TF release has not been investigated but we cannot exclude that this pathway might also be involved. Our current protocol for DC stimulation and MP harvest included incubation of DCs in low salt buffer, an experimental condition that on one hand improves MP recovery, but on the other that might also enhance TF targeting to MPs and obscure the contribution of secretory lysosomes or other possible export pathways. In any case, whichever the relative contribution of these two pathways to TF externalization, DC-derived MPs also contain IL-1ß (47) , thus it might well be that activated DCs release membrane-bound boluses of costored proinflammatory and procoagulant factors.

An often asked question is whether ATP is the true physiologically relevant stimulus for the P2X7R. Scattered evidence suggests that the P2X7R might also be activated by different stimuli such as antimicrobial peptides released by polymorphonuclear leukocytes (48) . This might suggest that additional proinflammatory factors that are generated in the atheromatous plaque or at other sites of vessel wall inflammation may cause release of TF-bearing MPs by acting at the P2X7R.

In conclusion, our observations identify a novel pathway for TF release from DCs and provide new insights into the mechanisms underlying the generation and spreading of procoagulant activity in immunity and inflammation.


   ACKNOWLEDGMENTS
 
This work was supported by grants from the Italian Association for Cancer Research (AIRC; to F.D.V.), Italian Space Agency (ASI; to F.D.V.), Telethon of Italy grants (GGP06070 to F.D.V., and GGP05214 to F.B.), Italian Ministry for Education (MIUR, to M.P., M.B., P.C., F.B., D.F. and F.D.V.), Comitato Sostenitori ARTGEA and Fondazione CARIFE (to B.F.), and institutional funds from the University of Ferrara.


   FOOTNOTES
 
1 The authors contributed equally to this work.

Received for publication September 20, 2006. Accepted for publication January 18, 2007.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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