Keywords
tissue factor - microvesicles - protease-activated receptor-2 - apoptosis - cell proliferation
- factor VIIa
Introduction
Tissue factor (TF) initiates the coagulation mechanism through the formation of a
complex with factor VIIa (fVIIa) which then activates factors X and IX.[1]
[2] TF is expressed on the surface of cells and may be released as cell-derived microvesicles
following cellular activation.[3]
[4]
[5]
[6]
[7]
[8]
[9] TF is capable of initiating cellular signals in cells expressing this protein, and
also on exposure of recipient cells to exogenous TF-containing microvesicles. TF signaling
can alter the cellular gene expression profile[10]
[11] and has been demonstrated to include fVIIa activity and protease-activated receptor-2
(PAR2) activation.[12]
[13]
[14]
[15]
[16]
[17]
[18] Furthermore, interaction with β-integrins has also been implicated in inducing cell
proliferation.[14]
[19]
[20]
[21] TF signaling is particularly associated with a high proliferative capacity in cancer
cells.[22]
[23]
[24] However, while the proliferative potential has been associated with the interaction
of TF with fVIIa, the data on the requirement for proteolytic function of fVIIa are
not consistent.[12]
[13] We previously showed that the exposure of cells to low levels of recombinant TF
alone promotes entry into the cell cycle.[25] However, the exposure of cells to high levels of TF additionally induces cell cycle
arrest at the G1/S checkpoint and can lead to cell apoptosis.[25]
[26]
[27] In addition, monocyte-derived microparticles can induce cellular apoptosis in endothelial
cells[28] and eosinophils lacking fVII become more susceptible to apoptosis.[29] Therefore, the magnitude of exposure to TF may itself determine the outcome in the
recipient cells. Furthermore, the formation of TF–fVIIa complex is capable of triggering
signals via PAR2 directly, or alternatively through the activation of factor Xa (fXa)
and formation of a tertiary complex.[30]
[31] The activation of PAR2 has also been shown to be essential in the signaling processes
that are initiated from the exposure of cells to TF.[32]
[33]
It has been shown that as a consequence of inflammation, disease, or injury, large
quantities of TF are released within microvesicles.[34]
[35]
[36]
[37] These microvesicles interact with endothelial cells and have also been shown to
be cleared by endocytosis.[38] While the inability of cells to satisfactorily process TF is detrimental to endothelial
cells,[27] acute exposure of cells to TF can also induce cellular apoptosis.[25]
[26]
[28] Therefore, prolonged exposure of the endothelial layer to TF-containing microvesicles
may give rise to endothelial dysfunction and denudation as seen in chronic diseases.[6]
[39]
[40] Understanding the criteria by which TF-containing microvesicles function, and determining
the characteristics that confer the proliferative and proapoptotic properties to the
microvesicles, may lead to a new understanding of the relationship between various
inflammatory diseases such as cancer and atherosclerosis with the onset and progression
of vascular disease. In this study, it was hypothesized that the ratio of fVIIa:TF
within cell-derived microvesicles is a determinant of the outcome of the exposure
of endothelial cells to the microvesicles. Consequently, the ability of the fVIIa–TF
complex, and cell-derived microvesicles containing a range of molar ratios of fVIIa:TF,
to promote endothelial cell proliferation or alternatively apoptosis was evaluated.
Material and Methods
Cell Culture
Five cell lines were selected to include a range of TF expression and not on the basis
of tissue of origin. BxPC-3 pancreatic cancer and 786-O renal carcinoma cell lines
(ATCC, Teddington, United Kingdom) were cultured in RPMI-1640 medium, MCF-7 breast
cancer cell line and HepG2 hepatocellular carcinoma cell lines (ATCC) were cultured
in EMEM, and MDA-MB-231 (ATCC) breast cancer cell lines were cultured in DMEM. All
media were supplemented with 10% (v/v) heat-inactivated fetal calf serum (FCS) to
ensure the lack of any functional enzymes. Human coronary artery endothelial cells
(HCAECs), devoid of endogenous TF, were cultured in MV media containing 5% (v/v) FCS
and growth supplements (PromoCell, Heidelberg, Germany). In some experiments, the
cells were adapted to serum-free medium prior to use.
Preparation and Analysis of the TF-Containing Microvesicles
Cell lines were propagated in 25 cm2 flasks, washed with phosphate-buffered saline (PBS) pH 7.4 and adapted to respective
serum-free medium, for 1 hour prior to collection by ultracentrifugation according
to described procedures for preparation and confirmation of the microvesicles.[38]
[41] The functional density of the released microvesicles was determined using the Zymuphen
MP-assay kit (Hyphen BioMed/Quadratech, Epsom, United Kingdom) using the standards
provided with the kit. The particle densities were also verified and the size distribution
of microvesicles was also examined by nanoparticle tracking analysis using a Nanosight
NTA 2.3 instrument.[38]
[41]
Qualitative Analysis of fVII/fVIIa and fX/fXa Antigens in the Microvesicles by Western
Blot
The presence of fX and fVII antigen was detected by western blot analysis. The samples
were separated by 12% (w/v) sodium dodecyl sulfate polyacrylamide gel electrophoresis
(SDS-PAGE), transferred onto nitrocellulose membranes, blocked with TBST (10 mM Tris-HCl
pH 7.4, 150 mM NaCl, 0.05% Tween-20). The membranes were then probed with a mouse
monoclonal anti-fX antibody (156106), or a rabbit polyclonal anti-fVII antibody (both
from R&D Systems, Abingdon, United Kingdom), both diluted 1:2000 (v/v) in TBST. The
membranes were developed with alkaline phosphatase-conjugated goat antimouse or goat
antirabbit antibodies (Santa Cruz, Heidelberg, Germany) respectively, diluted 1:4000
(v/v) and bands were visualized using the Western Blue stabilized alkaline phosphatase
substrate (Promega, Southampton, United Kingdom).
Quantitative Determination of TF and fVIIa Antigen Levels in the Microvesicles
The TF antigen associated with the microvesicles was measured using the Quantikine
TF-ELISA kit (R&D Systems) and the factor VII/VIIa antigen levels were determined
using the Assaymax FVII-ELISA kit (Assaypro/Universal Biologicals Ltd., Cambridge,
United Kingdom) according to the manufacturers' instructions. To determine the cell
surface fVIIa antigen in HCAEC without detachment, cells (2 × 104) were treated as required and then fixed using 4% (v/v) formaldehyde. The cells were
washed and one set was permeabilized while the other set was kept intact. Total and
surface expression of fVII were measured in situ, by incubating all samples with a
rabbit anti-fVIIa polyclonal antibody (10 µg/mL). The samples were then probed with
a HRP-conjugated goat antirabbit antibody (dilute 1:1000 v/v) and developed using
the TMB One-solution HRP substrate (Promega). The absorption values were measured
at 450 nm from which the percentage ratio of surface:total fVII was calculated. In
some experiments, the cells were subjected to repeated exposure to TF at 60 minute
intervals and the cell surface fVII antigen was measured and calculated as a percentage
of the original cell-surface fVII.
Measurement of TF, fVIIa, and fXa Activity
Microvesicle TF-fVIIa activities were measured by modification of previously described
procedures.[42]
[43] To measure TF activity, microvesicle samples were incubated with fVIIa (10 nM) in
HEPES-buffered saline (HBS) pH 7.4, containing 1% (w/v) bovine serum albumin (BSA)
and 5 mM CaCl2, together with fX (100 nM) and a fXa chromogenic substrate (0.2 mM; Hyphen) diluted
in the same buffer (200 µL). The samples were incubated for 60 minutes to develop
the color. Aliquots (150 µl) were then transferred to a 96-well plate containing 2%
(v/v) acetic acid (50 µL) and the absorptions measured immediately at 410 nm. The
amount of fXa generated was determined using a standard curve prepared using fXa (0–20 nM;
Enzyme Research Labs). To detect microvesicle-associated fVIIa activity, exogenous
fVIIa was omitted from the above, and replaced with recombinant TF (1 U/mL). The absorption
measurements were compared with a set of controls prepared using a range of fVIIa
(0–10 nM) which were supplemented with recombinant TF (1 U/mL) and analyzed for fXa-generation
potential as above. To examine the levels of fXa activity, the microvesicles were
diluted in the buffer as above and incubated with the fXa substrate alone. The absorption
measurements were compared with a set of controls prepared using a range of fXa as
above.
Treatment of HCAEC with fVII:TF and Microvesicles
HCAEC (2 × 104) were incubated with microvesicles prepared from the conditioned serum-free media
from the different cell lines, at microvesicle densities stated in the Results section.
Other sets of cells were incubated with combinations of recombinant human TF (Innovin
thromboplastin reagent; Dade Behring, Deerfield, United States) at a range of 0 to
10 U/mL (1 U/mL = 1.3 ng/mL), and/or purified fVIIa (Enzyme Research Labs, Swansea,
United Kingdom) at a range of 0 to 10 nM. Cell numbers were determined at 24 hour,
by staining with crystal violet as previously described and calculated from a standard
curve.[38] As further confirmation of entry into the cell cycle, the expression of cyclin D1
mRNA was measured by reverse transcription polymerase chain reaction (RT-PCR) as previously
described.[25] Total RNA was extracted using the Ribozol solution (VWR, Lutterworth, United Kingdom).
The expression of cyclin D1 mRNA was measured by GoTaq 1-Step RT-qPCR System (Promega,
Southampton, United Kingdom) using QuantiTect primers for cyclin D1 and β-actin (Qiagene,
Manchester, United Kingdom) and relative amounts determined using as a reference.
In addition, cellular apoptosis was quantified using the TiterTACS Colorimetric Apoptosis
Detection Kit (AMS Biotechnology, Abingdon, United Kingdom) according to the manufacturer's
instructions.[27] The data for apoptosis were expressed as the change in absorption measured at 450
nm. Furthermore, the expression of bax mRNA was quantified in the cell samples according
to described procedures and using the reagents above, to further confirm the induction
of cell apoptosis.[25]
[27] In some experiments, the microvesicles were preincubated with an inhibitory mouse
antihuman TF antibody (HTF-1; 20 µg/mL; eBioscience/Thermo Scientific, Warrington,
United Kingdom), a signal-blocking antihuman TF antibody (10H10; 20 µg/mL; BD Bioscience,
Wokingham, United Kingdom), an inhibitory polyclonal rabbit antihuman fVIIa antibody
(10 µg/mL; Abcam, Cambridge, United Kingdom), or respective control isotype IgG antibodies
(20 µg/mL; New England Biolabs, Hitchin, United Kingdom). In other experiments, the
activation of PAR2 on HCAEC was blocked by preincubation of cells with an inhibitory
anti-PAR2 antibody (SAM11; Santa Cruz Biotechnology, Heidelberg, Germany) which was
used at 20 µg/mL. Additionally, sets of HCAEC were supplemented with PAR2-activating
peptide (PAR2-AP; SLIGKV) at a final concentration of 20 µM, simultaneously with the
addition of the microvesicles (Severn Biotech Ltd, Kidderminster, United Kingdom).
Finally, in some experiments the cells were preincubated with Rivaroxaban (pure compound
supplied by Bayer, Leverkusen, Germany) and used at therapeutic concentrations (0.6
µg/mL), which was also optimized against fXa prior to this study.[44]
Preparation of mCherry-PAR2 Hybrid and Measurement of PAR2 Activation
To measure the activation of PAR2, a hybrid tandem protein was expressed to contain
mCherry, followed by full-length PAR2. The hybrid protein was therefore designed so
that the proteolytic activation of PAR2 released mCherry from the cell surface, into
the medium. The cDNA for PAR2 was amplified by PCR from the hPAR2 VersaClone cDNA
plasmid (R&D Systems) using the forward (5′-GCTCAAGCTTATCCAAGGAACCAATAGATCCTC) and
reverse (5′-CGGTGGATCCTTAATAGGAGGTCTTAACAGTGG) primers and then digested with BamHI and HindIII restriction enzymes (1 U/mL; New England Biolabs). The insert was ligated using
the Instant Sticky-end ligase master mix (New England Biolabs), at a molar ratio of
3:1 into a mCherry2-C1 vector (Addgene) which was predigested with the same two restriction
enzymes. 5α-Competent Escherichia coli bacteria were transformed with the plasmid DNA construct and colonies were selected
from LB-agar plates containing carbenicillin (100 µg/mL). The cells were propagated
and the positive colonies containing the mCherry-PAR2 construct were selected and
confirmed by sequencing (Eurofins MWG, Wolverhampton, United Kingdom). Plasmid DNA
was harvested from the bacteria by midi-prep (Promega Corp, Southampton, United Kingdom),
using a procedure that eliminates any residual endotoxins. HCAECs were transfected
with the mCherry-PAR2 constructs and surface expression was confirmed by fluorescence
microscopy and also by comparing to cells labeled with FITC-conjugated anti-PAR2 antibody.
The analysis was performed using a Zeiss Axio Vert.A1 inverted fluorescence microscope
with a ×40 magnification (Carl Zeiss Ltd, Welwyn Garden City, United Kingdom). The
activation of PAR2 was monitored by measuring the release of mCherry following the
proteolytic digestion of PAR2. HCAECs expressing the hybrid protein were washed with
PBS and adapted to serum-free medium. Since mCherry is multiply digested by the proteolytic
action of fXa, the release of mCherry from the cell surface was only examined by incubation
of cells with combinations of recombinant TF (0–4 U/mL) and purified fVIIa (0–4 nM)
for up to 60 minutes. Finally, the ability of microvesicles derived from HepG2, BxPC-3,
786-O, MDA-MB-231, and MCF-7 cell lines, to release mCherry, was examined.
Statistical Analysis
All data represent the calculated mean values from the number of experiments stated
in each figure legend ± the calculated standard error of the mean. Statistical analysis
was performed using the Statistical Package for the Social Sciences (SPSS Inc. Chicago,
United States). Significance was determined using one-way ANOVA (analysis of variance)
and Tukey's honest significance test or where appropriate, by paired t-test.
Results
The Ratio of fVIIa to TF Varies in Microvesicles Derived from Different Cell Lines
Throughout the study, microvesicles were purified from resting HepG2, BxPC-3, 786-O,
MDA-MB-231, and MCF-7 cell lines in accordance to the diverse expression of TF, irrespective
of tissue of origin. The microvesicles were purified by ultracentrifugation from serum-free
conditioned media according to established procedures.[41] Prior to examining the influence of microvesicles on endothelial cell proliferation
and apoptosis, the microvesicle density, TF antigen, fVII/fVIIa antigen, and fXa-generation
potential of the microvesicles were analyzed. The particle densities were verified
and the size distribution of microvesicles was also examined by NTA ([Supplementary Fig. S1]). Comparison of similar quantities of microvesicles from the cells lines indicated
that both BxPC-2 and MDA-MB-231 microvesicles contained the highest amounts of TF
([Fig. 1A]), which is in agreement with our previous study.[45] Examination of fXa-generation in the presence of supplemented fVIIa mostly reflected
the measured TF antigen in the microvesicles ([Fig. 1B]). Examination of the fVIIa by western blot indicated the presence of active fVIIa
in HepG2, MDA-MB-231, and MCF-7 cell lines, frequently exhibiting multiple bands ([Fig. 1C]) which have previously been attributed to the presence of glycosylation variants.[46]
[47] Moreover, 786-O and BxPC-3 cells contained relatively lower levels of fVIIa as measured
by western blot ([Fig. 1D]). The amounts of fVIIa were also reflected in the relative levels of fVIIa mRNA
expression in four of the cell lines ([Supplementary Fig. S2]) but was four orders of magnitude higher in HepG2 cells (not shown). The amounts
of microvesicle-associated fVII were then quantified using a fVII-ELISA kit ([Fig. 1E]) and were largely in line with those detected by western blot. In the absence of
exogenous fVIIa, microvesicles from HepG2, MDA-MB-231, and MCF-7 cells possessed higher
fXa-generation capacity indicating high fVIIa activity ([Fig. 1F]), which largely agreed with the level of fVII antigen in these microvesicles ([Fig. 1E]). In contrast, lower fVIIa activities were detectable in microvesicles from BxPC-3
and 786-O cell lines ([Fig. 1F]). Preincubation of all microvesicles with an inhibitory polyclonal anti-fVII antibody
abolished the inherent fVIIa activity measured as above (not shown). The fVII and
TF antigen quantities were then used to determine the ratio of fVIIa antigen to TF
antigen and were established to be highest in HepG2 cells and lowest in 786-O cells
([Fig. 1G]). The presence of fVIIa on the surface of microvesicles requires calcium ions. It
has been shown that microvesicles can contain a high concentration of calcium ions,[48] which is released during the formation of microvesicles by calcium-sensitive enzymes
including calpains and gelsolins, and therefore permits the nonspecific binding of
fVIIa to the microvesicle surface. Moreover, the concentration of fVIIa is only pathophysiologically
relevant as fVIIa/TF complex and is therefore dictated by the concentration of TF
and not fVIIa. However, none of the microvesicles examined contained fX/fXa antigen
when examined by western blot, or possessed any detectable fXa activity following
incubation with an fXa-specific chromogenic substrate (not shown).
Fig. 1 Analysis of the TF and fVIIa content of microvesicles. Five cell lines (HepG2, BxPC-3,
786-O, MDA-MB-231, and MCF-7) were propagated in 25 cm2 flasks, washed with phosphate-buffered saline (PBS) pH 7.4 and adapted to respective
serum-free medium, for 1 hour. Microvesicles were prepared from the conditioned media
by ultracentrifugation and the functional density determined using the Zymuphen MP-assay
kit. (A) The TF antigen concentration of the microvesicles was measured using the Quantikine
TF-ELISA kit and (B) the associated TF activities were measured using the fXa-generation assay (n = 5). (C) The presence of fVII/fVIIa antigen in the cells was detected by western blot analysis.
The samples were separated by 12% (w/v) SDS-PAGE, transferred onto nitrocellulose
membranes and then blocked with TBST (10 mM Tris-HCl pH 7.4, 150 mM NaCl, 0.05% Tween-20).
The membranes were then probed with a rabbit polyclonal anti-fVII antibody diluted
1:2000 (v/v) in TBST and developed with an alkaline phosphatase-conjugated goat antirabbit
antibody, diluted 1:4000 (v/v). The bands were visualized using the Western Blue stabilized
alkaline phosphatase-substrate and (D) quantified by ImageJ program. (The micrographs are representative of three separate
experiments). (E) Microvesicle-associated factor VII/VIIa antigen levels were quantified using the
Assaymax FVII-ELISA kit (n = 5). (F) Microvesicle-associated fVIIa activity was also measured by fXa-generation assay
but in the absence of exogenous fVIIa and presence of TF (1 U/mL) (n = 5). (G) The molar ratio of fVIIa:TF in microvesicles derived from the cells lines was calculated
using the data generated above. SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel
electrophoresis; TF, tissue factor.
Fig. 2 Examination of the influence of cell line-derived microvesicles on HCAEC cell numbers.
Microvesicles were prepared from cell lines (HepG2, BxPC-3, 786-O, MDA-MB-231, and
MCF-7), supplemented (50 nM) to HCAEC (2 × 104) and incubated in for up to 48 hours. (A) Cell numbers were determined using the crystal violet staining assay and (B) the change in cell numbers was calculated as a percentage of the untreated cells
(n = 5; * = p < 0.05 vs. the respective untreated sample at each time point). (C) Total RNA was extracted from samples of the treated HCAEC and the relative expression
of cyclin D1 mRNA was analyzed by RT-PCR against β-actin as housekeeping (n = 3; * = p < 0.05 vs. the untreated sample). Cellular apoptosis was also measured (as absorption
at 450 nm) in sets of HCAEC using the TiterTACS chromogenic TUNEL assay and (D) the change in level of apoptosis was calculated as a percentage of the untreated
cells (n = 5; * = p < 0.05 vs. the untreated sample). (E) Total RNA was extracted from samples of the treated HCAEC and the relative expression
of bax mRNA was analyzed by RT-PCR against β-actin as housekeeping (n = 3; * = p < 0.05 vs. the untreated sample). HCAEC, human coronary artery endothelial cell;
RT-PCR, reverse transcription polymerase chain reaction.
Cell Derived Microvesicles Differentially Influence Endothelial Cells
The microvesicles derived from the cell lines were incubated with HCAEC and the influence
on cell proliferation or apoptosis was measured. Examination of HCAEC cell numbers
indicated that the greatest increases in cell numbers on incubation with microvesicles
are derived from HepG2 and MCF-7 cells ([Fig. 2A] and [2B]). In addition, marginal increases were observed with microvesicles from MDA-MB-231
cells, but not BxPC-3 cells. These increases in cell numbers were also reflected in
the expression of cyclin D1 as a marker for cell-cycle entry ([Fig. 2C]). In contrast, supplementation of HCAEC with microvesicles derived from 786-O resulted
in reduction in cell numbers compared with the untreated sample. The next sets of
experiments examining the induction of apoptosis only were then focused onto microvesicles
derived from 786-O cells. These were compared with the outcome from microvesicles
derived from BxPC-3 cells which had little influence. HCAECs were incubated with three
microvesicle densities (25–130 nM) from each cell line. The rate of HCAEC apoptosis
was then measured using a chromogenic TUNEL assay. Incubation of HCAEC with 50 nM
of 786-O cell-derived microvesicles resulted in maximal level of cell apoptosis ([Fig. 2D] and [2E]) but was not significant with BxPC-3-derived microvesicles. To further confirm the
increases in cellular apoptosis, the expression of bax mRNA was measured and shown
to agree with the data using the TUNEL assay ([Fig. 2F]).
The Ratio of fVIIa:TF within Microvesicles Determines the Outcome on Endothelial Cells
To further examine the relevance of microvesicle-associated fVIIa and TF, sets of
HCAECs were incubated with combinations of recombinant TF (0–10 U/mL; Innovin), in
conjunction with three concentrations of fVIIa (0–10 nM). In the absence of fVIIa,
incubation of cells with recombinant TF resulted in lowering of cell numbers ([Fig. 3A]). Supplementation with fVIIa (2 nM) partially prevented the decline in cell numbers
but became ineffective at TF concentrations higher than 0.5 U/mL. Inclusion of 10 nM
fVIIa restored the cell numbers up to 2 U/mL of TF and was proliferative when combined
together with lower TF concentrations. It is also noteworthy that in the absence of
TF the addition of fVIIa had no detectable effect. These data were also confirmed
by titrating a range of concentrations of fVIIa, together with three separate concentrations
of TF. On supplementing the cells with 0.5 U/mL of TF, increased HCAEC numbers were
observed with all the concentrations of fVIIa tested ([Fig. 3B]). However, in the presence of 2 U/mL of TF, fVIIa concentrations of 1 nM or higher
were required to preserve or increase the cell numbers. Furthermore, on addition of
4 U/mL of TF, only inclusion of 10 nM fVIIa was capable of preventing the reduction
in cell numbers. Analysis of cyclin D1 and bax expression in HCAEC treated with recombinant
TF (2 U/mL) and purified fVIIa (0–10 nM) indicated entry into the cell cycle regardless
of fVIIa concentration, in agreement with our previous findings,[25] but was highest at 10 nM ([Fig. 3C]), while bax expression was only detectable at lower fVIIa concentrations ([Fig. 3D]).
Fig. 3 Examination of the effect of different ratios of fVIIa:TF on HCAEC cell numbers.
(A) HCAECs (2 × 104) were incubated for 24 hours with a range of recombinant human TF (Innovin thromboplastin
reagent) at a range of 0–10 U/mL (1 U/mL = 1.3 ng/mL), in the presence or absence
of purified fVIIa (0–10 nM). Cell numbers were determined by crystal violet staining
assay and percentage change in cell numbers calculated (n = 5; * = p < 0.05 vs. the untreated sample). (B) HCAECs (2 × 104) were also incubated for 24 hours with a range of concentrations of purified fVIIa
(0.1–10 nM) in the presence of recombinant human TF (0–4 U/mL). Cell numbers were
determined by crystal violet staining assay and percentage change in cell numbers
calculated (n = 5; * = p < 0.05 vs. the untreated sample). Total RNA was extracted from samples of the treated
HCAEC and the relative expression of (C) cyclin D1 mRNA and (D) bax mRNA was analyzed by RT-PCR against β-actin as housekeeping (n = 3; * = p < 0.05 vs. the untreated sample). HCAEC, human coronary artery endothelial cells;
RT-PCR, reverse transcription polymerase chain reaction; TF, tissue factor.
Microvesicle-Induced Cell Proliferation and Apoptosis is Mediated through PAR2
To explore the mechanisms involved in the induction of cell proliferation and apoptosis
by microvesicles, HCAECs were preincubated with an antibody to prevent PAR2 activation
(SAM11; 20 µg/mL), prior to addition of microvesicles derived from HepG2 or 786-O
cell lines. Inhibition of PAR2 activation prevented the increase in HCAEC numbers
in response to microvesicles derived from HepG2 cell line, after 24 hours ([Fig. 4A]). However, preventing the activation of PAR2 using the antibody also suppressed
the proapoptotic influence of 786-O cell-derived microvesicles ([Fig. 4B]). Moreover, the analysis of cyclin D1 expression in HCAEC stimulated with microvesicles
derived from HepG2 cells ([Fig. 4C]) or bax mRNA in cells stimulated with microvesicles derived from 786-O cells ([Fig. 4D]) confirmed the requirement for PAR2 activation in both the proliferative and proapoptotic
outcomes. Preincubation of 786-O microvesicles, or alternatively the HCAEC with effective
concentrations of Rivaroxaban (0.6 µg/mL) to inhibit any present fXa, did not prevent
endothelial apoptosis (not shown). Moreover, simultaneous activation of PAR2 using
the activating peptide (SLIGKV; 20 µM) in conjunction with 786-O cell-derived microvesicles
(50 nM) was capable of rendering these microvesicles ineffective ([Fig. 4E]). To decipher these observations, the presence of PAR2 antigen on the surface of
cell surface, following the addition of PAR2-AP or a range of concentrations of microvesicles
from 786-O cells, was examined. The HCAECs were probed with a PAR2 antibody in situ,
to avoid nonspecific activation of PAR2 by trypsinization. HCAECs were incubated with
microvesicles from 786-O cells at the optimal (50 nM) and hyper-optimal (130 nM) densities
to induce apoptosis ([Fig. 2E]). Incubation of HCAEC with PAR2-AP (20 µM) or with 130 nM of 786-O cell-derived
microvesicles resulted in significant reductions in cell surface PAR2 within 30 minutes
([Fig. 4F]). In contrast, cell-surface PAR2 levels remained unaltered by the addition of 50 nM
of these microvesicles.
Fig. 4 Examination of the involvement of PAR2 in microvesicle-induced HCAEC apoptosis. Sets
of HCAEC (2 × 104) were preincubated with a blocking anti-PAR2 antibody (SAM11; 20 µ/mL) or a control
isotype. Sets of cells were then incubated with microvesicles derived from (A) HepG2 cell line (50 nM) for 24 hours and cell numbers were determined by crystal
violet staining assay (n = 5; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. the sample devoid of antibody). (B) A similar set of cells was incubated for 24 hours with microvesicles derived from
786-O cell line (50 nM) and the rate of apoptosis determined using the TiterTACS chromogenic
TUNEL assay (n = 5; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. the sample devoid of antibody). Total RNA was extracted from samples of
the treated HCAEC and (C) the relative expression of cyclin D1 mRNA, and (D) bax mRNA was analyzed by RT-PCR against β-actin as housekeeping (n = 3; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. the sample devoid of antibody). (E) HCAECs (2 × 104) were incubated for 24 hours with microvesicles derived from 786-O cell line (50
nM) in the presence or absence of PAR2-activating peptide (20 µM) and the rate of
apoptosis determined (n = 5; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. microvesicles only). (F) HCAECs (2 × 104) were incubated for 30 minutes with microvesicles derived from 786-O cell line (0–130
nM), or PAR2-activting peptide (20 µM). The cells were then incubated with a mouse
antihuman PAR2 antibody (SAM11; 20 µg/mL) and probed with a HRP-conjugated goat antimouse
antibody diluted 1:1000 (v/v). The relative amount of cell-surface PAR2 was then measured
using the TMB substrate (n = 5; * = p < 0.05 vs. the untreated sample). HCAEC, human coronary artery endothelial cells;
PAR2, protease-activated receptor-2; RT-PCR, reverse transcription polymerase chain
reaction.
Microvesicles Differentially Activate PAR2 on the Surface of Endothelial Cells
To examine the level of PAR2 activation in response to various microvesicles, a hybrid
construct was prepared containing the cDNA for mCherry-PAR2 and was expressed in HCAEC.
The expression of the mCherry-PAR2 in HCAEC was confirmed by fluorescence microscopy
([Fig. 5A]). This was then compared with the pattern of native PAR2 expression, probed using
Alexa Fluor488-conjugated anti-PAR2 (SAM11) antibody ([Fig. 5B]). To determine the effectiveness of the TF–fVIIa complex in activating PAR2, the
transfected cells were incubated with combinations of TF (0–4 U/mL) together with
fVIIa (0–10 nM) and the release of mCherry in the media was measured at Em. 630 nm
(Ext. 580 nm). In the absence of TF, the addition of fVIIa (0–10 nM) did not result
in the release of fluorescence into the media ([Fig. 5C]). Moreover, addition of TF alone, or together with low level of fVIIa (0.5 nM),
was not sufficient to induce PAR2 activation. Incubation of cells with TF together
with fVIIa (2 nM) resulted in marginal increases in fluorescence, which was significant
at higher TF concentrations. In contrast, inclusion of fVIIa (10 nM) resulted in a
TF concentration-dependent activation of PAR2 from the transfected HCAEC. Examination
of the effectiveness of the microvesicles purified from the five cell lines in activating
mCherry-PAR2 expressed on the surface endothelial cell confirmed a high activity associated
with HepG2 and MCF-7-derived microvesicles, with lower levels with MDA-MB-231 microvesicles
but no significant PAR2 activation with microvesicles from BxPC-3 or 786-O cells ([Fig. 5D]).
Fig. 5 Examination of the activation of PAR2 in response to fVIIa:TF complex and microvesicles.
Sets of HCAEC (5 × 103) were transfected to express mCherry-PAR2 hybrid protein. (A) Sets of transfected and untransfected cells were then fixed and stained phalloidin-iFlour
488 (diluted 1:1000 v/v) and DAPI (2 µg/mL) and examined by fluorescence microscopy
for mCherry-PAR2 (Red), phalloidin-iFlour 488 (Green), and DAPI (Blue). (B) As a comparison, untransfected cells which were labeled with FITC-conjugated anti-PAR2
(SAM11; 20 µg/mL) or unlabeled cells. The cells were examined by fluorescence microscopy
for PAR2 (Green) and DAPI (Blue). (C) HCAECs expressing the mCherry-PAR2 protein were incubated for 1 hour with combinations
of TF (0–4 U/mL) together with fVIIa (0–10 nM) and the release of mCherry in the media
was measured by determining the fluorescence at Em. 630 nm (Ext. 580 nm) (n = 5; * = p < 0.05 vs. the untreated sample). (D) Sets of transfected cells were also incubated with microvesicles derived from the
five cell lines (HepG2, BxPC-3, 786-O, MDA-MB-231, and MCF-7) at 50 nM and the release
of mCherry measured by determining the fluorescence at Em. 630 nm (Ext. 580 nm) (n = 5; * = p < 0.05 vs. the untreated sample). HCAEC, human coronary artery endothelial cells;
PAR2, protease-activated receptor-2; TF, tissue factor.
Exogenous fVIIa Prevents TF-Microvesicle-Induced Apoptosis
To test the hypothesis that increased levels of fVIIa can rescue the cells from microvesicle-induced
apoptosis, and may induce proliferation, HCAECs were incubated with 786-O cell-derived
microvesicles in the presence or absence of exogenous fVIIa (2 nM). Inclusion of additional
fVIIa abolished the proapoptotic influence of 786-O cell-derived microvesicles ([Fig. 6A]). However, as shown before ([Fig. 3A]), the supplementation with fVIIa in the absence of any TF did not influence the
HCAEC numbers which are otherwise devoid of endogenous TF expression. Preincubation
of 786-O microvesicles with an inhibitory anti-TF antibody (HTF-1; 20 µg/mL), which
blocks the procoagulant activity of TF, prevented cellular apoptosis ([Fig. 6B]). However, preincubation of these microvesicles with the 10H10 anti-TF antibody,
which inhibits TF signaling, or the isotype control antibody did not interfere with
the promotion of apoptosis in HCAEC. In addition, neutralization of fVIIa in microvesicles
derived from 786-O and HepG2 cells, using an inhibitory anti-fVIIa antibody, prevented
the promotion of cell apoptosis in response to 786-O-derived microvesicles ([Fig. 6C]) and also the increased cell numbers by HepG2-derived microvesicles ([Fig. 6D]), respectively. 10H10 antibody blocks TF signaling by blocking TF interaction with
integrins[49]
[50] and the secondary/exosite PAR2-binding site within TF.[51] However, unlike HTF1, it does not prevent the interaction with fVIIa,[51] nor does it affect the proteolytic activity of the TF/fVIIa complex.[52] Therefore, in agreement with the data using the inhibitory anti-fVIIa antibody ([Fig. 6C]), the initiation of the described mechanism is dependent on the proteolytic activity
of fVIIa as well as requires TF.
Fig. 6 Active fVIIa both attenuates and is required for the induction of HCAEC apoptosis
by microvesicle-associated TF. (A) Sets of HCAEC (2 × 104) were incubated with microvesicles derived from 786-O cells alone, or supplemented
with fVIIa (5 nM) and cellular apoptosis was measured after 24 hours (n = 5; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. the respective sample without fVIIa). (B) Microvesicles derived from 786-O cells were preincubated with HTF-1 anti-TF antibody
(20 µg/mL) to block fVIIa binding, 10H10 anti-TF antibody (20 µg/mL) to block TF signaling,
or an isotype mouse antibody (20 µg/mL), and cellular apoptosis was measured after
24 hours (n = 5; * = p < 0.05 vs. the untreated sample). (C) Microvesicles derived from 786-O cells were preincubated with an inhibitory anti-fVIIa
antibody (20 µg/mL) prior to addition to HCAEC (2 × 104) and cellular apoptosis was measured after 24 hours (n = 5; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. sample with microvesicle alone). (D) Microvesicles derived from HepG2 cells were preincubated with an inhibitory anti-fVIIa
antibody (20 µg/mL) prior to addition to HCAEC (2 × 104). HCAEC numbers were determined after 24 hours using the crystal violet staining
assay and the cell numbers calculated (n = 5; * = p < 0.05 vs. the untreated sample, # = p < 0.05 vs. sample with microvesicle alone). HCAEC, human coronary artery endothelial
cells; TF, tissue factor.
Endothelial Cells Release fVII in Response to TF
Examination of HCAEC by RT-PCR indicated the presence of detectable amounts fVII mRNA
in HCAEC (not shown). Measurement of the fVII protein lying intact and lysed HCAEC
indicated that approximately 20% of the fVII was exposed at the surface of the resting
cells ([Fig. 7A]). The presence of fVII associated with caveolae and Weibel–Palade bodies in differentiating
endothelial cells has been demonstrated[53] and our unpublished data have shown some association of fVII with caveolae in resting
endothelial cells (Madkhali et al, unpublished data). However, stimulation of HCAEC
with recombinant TF (2 U/mL) resulted in a significant increase in cell surface fVII
antigen (48% of the cellular fVII), as measured by the in situ labeling. Furthermore,
activation of HCAEC with PAR2-AP (20 µM) resulted in a comparable but longer-term
increase (33% of the cellular fVII at 30 minutes) in the concentration of cell-surface
fVII ([Fig. 7B]). In some experiments, HCAECs were subjected to repeated treatment of cells with
recombinant TF at 60 minute intervals. In these experiments, both the remaining cell-surface
fVII and the microvesicle-associated fVII antigens were measured as a percentage of
that present on the surface of the resting cells. Treatment of HCAEC with recombinant
TF progressively decreased the amount of fVII present on the cell surface ([Fig. 7C]). Furthermore, the magnitudes of these reductions were accounted for by the amount
of fVII that was associated with the released microvesicles ([Fig. 7D]).
Fig. 7 HCAECs express fVIIa on the cell surface in response to TF and PAR2 activation. Two
separate sets of HCAEC (2 × 104) were incubated with (A) TF (2 U/mL) or (B) PAR2-activating peptide (20 µM) for the durations shown and the cells were then
fixed using 4% (v/v) formaldehyde. The cells were washed and one set was permeabilized
while the other kept intact. Total and surface expression of fVII were measured in
situ, by incubating all samples with a mouse anti-fVIIa antibody (20 µg/mL). The samples
were then probed with a HRP-conjugated goat antimouse antibody (dilute 1:1000 v/v)
and developed using the TMB substrate. The percentage ratio of surface:total fVII
was then calculated (n = 5; * = p < 0.05 vs. the observed ratio at time zero). HCAECs (2 × 104) were also subjected to repeated treatment with recombinant TF (2 U/mL) at 60 minute
intervals. After each treatment, the relative amount of (C) cell-surface fVII antigen and (D) the released microvesicle-associated fVII antigen were measured and the percentages
were calculated against the amount of cell-surface fVII in the untreated cells. HCAEC,
human coronary artery endothelial cells; PAR2, protease-activated receptor-2; TF,
tissue factor.
Discussion
The exposure of TF at the site of injury acts to initiate the coagulation mechanism
and contains bleeding. However, as a factor which appears early at the sight of injury,
TF is ideally placed to have a dual function in instructing the cells to divide or
become apoptotic. The distinction between the severely injured cells and those which
may be revived is imperative in the precise vascular homeostasis. These functional
properties of TF also appear to be replicated in the microvesicle-associated TFs that
are released into the bloodstream from various sources. In addition to TF, these microvesicles
may also contain a complement of negatively charged phospholipids[3]
[4]
[5]
[6]
[7]
[8] and functional fVIIa ([Fig. 1]). It is known that microvesicles from different sources exert dissimilar influence
on endothelial cells, which provides crucial clues for the understanding of the destructive
influence of microvesicles in diseases.[33]
[34]
[35]
[36] Therefore in this study, by measuring the ratios of fVII/fVIIa and TF, we examined
a possible mechanism by which TF-containing microvesicles may confer different outcomes
in cultured primary endothelial cells. In agreement with this hypothesis, incubation
of HCAEC with combinations of purified fVIIa and recombinant TF resulted in different
cellular outcomes depending on the fVIIa:TF ratio. The transition from the proapoptotic
to proliferative property appears to occur at an estimated fVIIa:TF molar ratio of
15:1. This was also in agreement with the ratios observed in the microvesicles purified
from the cell lines. Particularly, the fVIIa:TF ratio in the 786-O renal carcinoma
cell line was 10:1 and these microvesicles induced cellular apoptosis in HCAEC ([Fig. 1G]). Higher molar ratios of 54:1and 38:1 observed in the HepG2 hepatocellular line
and MCF-7 breast cancer line ([Fig. 1]) were concurrent with increased HCAEC proliferation. Interestingly, the change in
cell numbers was significantly proportional to the observed fVIIa:TF ratio (Pearson
correlation = 0.956; p = 0.011). However, it also appears that as well as the fVIIa:TF ratio, the concentration
of TF with which the cell comes into contact with is an additional critical factor
in determining the outcome. Therefore, despite the similar fVIIa:TF molar ratios,
microvesicles derived from MDA-MB-231 (34:1) were significantly less proliferative
than those derived from MCF-7 cell lines (38:1). This may therefore be explained by
the much higher TF content of microvesicles from MDA-MB-231 cells.
Although HCAEC lacked any detectable cell-surface TF, a significant proportion of
cellular fVII was detected on the surface of these endothelial cells ([Fig. 7]). Furthermore, activation of HCAEC with recombinant TF or PAR2-AP resulted in the
exposure of a higher proportion of the cellular fVII. Therefore, it is possible that
endothelial cells respond to the stimulatory signals arising from injury/trauma by
altering the fVIIa:TF ratio, to counter the proapoptotic influence that arises from
the presence of excessive levels of TF. However, because in our studies only the amounts
of exogenous fVIIa and TF were used in calculating the fVIIa:TF molar ratios, the
real ratios for the transition from the proapoptotic to proliferative form are likely
to be higher than those reported here (15:1). In addition, repeated exposure of HCAEC
resulted in the depletion of cellular fVII reserves ([Figs. 7C] and [7D]). Such a compromise in endothelial cell function implies that the response by these
cells may become insufficient in ensuring the survival of the cell. Therefore, we
hypothesize that repeated exposure to TF-positive microvesicles, for example during
chronic disease, exhausts the ability of endothelial cells to counter the proapoptotic
property of TF.
To further elucidate the underlying proliferative mechanisms, PAR2, TF, and fVIIa
were in turn inhibited using antibodies. Inhibition of PAR2 prevented the proapoptotic
function of these microvesicles, indicating the requirement for PAR2 activation. Moreover,
overstimulation of the cells with high concentrations of microvesicles or the addition
of PAR2-AP was less effective due to the endocytosis of PAR2 and desensitization of
HCAEC. Both the proliferative and proapoptotic influences of microvesicles were attributed
to the proteolytic activity of the fVIIa-TF complex and were blocked by respective
inhibitory antibodies. Finally, preincubation of microvesicles or HCAEC with Rivaroxaban
to inhibit fXa did not prevent apoptosis in HCAEC. Microvesicles derived from different
cell lines may contain pro- and antiapoptotic/proliferative material which can then
contribute to endothelial cell proliferation or apoptosis. However, by selectively
inhibiting TF or fVIIa ([Fig. 6]), or alternatively blocking PAR2 ([Fig. 4]), using specific antibodies we have demonstrated that these outcomes were initiated
from the fVIIa/TF complex. In addition, the use of recombinant TF and purified fVIIa
further confirmed the role of TF and fVIIa observed with cell-derived microvesicles
([Fig. 3]). In contrast, activation of PAR2 in the absence of TF has been shown not to induce
a similar complement of the signaling pathway and does not lead to apoptosis.[27]
This study has for the first time shown that the ratio of fVIIa:TF determines the
outcome in endothelial cells resulting in either proliferation or apoptosis. This
is particularly relevant in the case of TF-containing microvesicles which are released
during inflammatory conditions. The induction of cell proliferation and apoptosis
by microvesicles appears to be mediated through the activation of PAR2, but the cellular
outcome is entirely dependent on the amount of TF protein and the molar ratio of fVIIa-TF
protease activity. However, the downstream events are only partially elucidated. In
conclusion, a novel measurable parameter within procoagulant microvesicles that can
determine the function of microvesicles during the activation of the vasculature has
been identified.
