© 1997 American Heart Association, Inc.
Articles |
Platelet Activation in Flowing Blood Passing Growing Arterial Thrombi
From the Research Forum, Ullevaal University Hospital (I.A.H., T.L.), and the Department of Plastic Surgery, Rikshospitalet National Hospital (H.E.R.), Oslo, Norway.
Correspondence to Inger Anne Hagberg, Research Forum, Ullevaal University Hospital, 0407 Oslo, Norway.
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Abstract |
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Abstract We investigated the combined effect of wall shear rate and immobilized collagen on platelet activation in flowing nonanticoagulated human blood. By combining an ex vivo model of thrombogenesis with flow cytometry, we showed that activated platelets can be detected in the bloodstream passing growing thrombi at a wall shear rate characteristic of moderately stenosed arteries (2600 s-1). The activation of the circulating platelets was clearly correlated with thrombus growth. Different antibodies against platelet activation-dependent surface markers had distinct sensitivity to the thrombotic process.
-Granule
release detected by surface expression of CD62P seemed to be the most
sensitive marker, as judged by both mean fluorescence intensity
and fraction of platelets activated. The conformational
change in glycoprotein IIb–IIIa, as detected by PAC-1,
also seemed to be a sensitive marker and preceded binding of fibrinogen
to activated glycoprotein IIb–IIIa, as detected by
anti-fibrinogen. Large thrombi also elicited lysosome
exocytosis, detected by surface expression of CD63. Finally, we
observed a small decrease of glycoprotein Ib–IX
expression, as detected by anti-CD42a. Thus, our study provides further
information on the dynamics of platelet activation in relation to
thrombus growth at arterial shear conditions in flowing
nonanticoagulated human blood.
Key Words: perfusion • platelet activation • flow cytometry • arterial thrombogenesis • collagen
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Introduction |
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Exposure of a thrombogenic substrate to flowing blood at a site of vascular injury leads to local accumulation of platelets, which form a hemostatic plug or a pathological occluding thrombus.1 During acute thrombogenesis, the adhering platelets release their granular contents. The released substances have different biological effects, and some are potent platelet activators. These are thought to activate both platelets recruited in the growing thrombi and platelets passing the thrombogenic site without adhering.
Blood flow and wall shear rate affect the hemodynamic interaction between the blood components and the vessel wall.2 Blood perfusion models allow exposure of arterial subendothelium, collagen, and extracellular endothelial matrix to flowing blood under well-controlled and reproducible conditions. Previous perfusion studies have focused on molecular mechanisms of platelet–surface adhesion,3 4 5 6 platelet–platelet interaction,7 8 9 10 and the interplay between coagulation and platelet function11 12 13 14 in the growing thrombi. We studied the platelets passing the growing thrombi to gain more information on their status of activation.
Flow cytometry offers a unique possibility for single-cell analysis. By analyzing blood passing through a human ex vivo model15 at an arterial shear rate, we were able to detect and evaluate platelet surface changes caused by growing arterial thrombi. The goal was to identify the activated platelets with flow cytometry and to determine whether there was any correlation between thrombus formation and activation of circulating platelets.
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Methods |
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Study Design
Four healthy, nonsmoking, male volunteers, aged 25 to 43 years, were included in this open study. None of the volunteers had taken aspirin or other antiplatelet drugs for at least 10 days before the perfusion experiments. All individuals were fully informed about the study and gave informed consent for participation.
Thrombus formation was generated on human fibrillar collagen in parallel-plate perfusion chambers developed by Sakariassen et al.16 17 Platelet activation in blood passing the growing thrombi was measured by flow cytometry in samples collected immediately downstream of (distal to) the chamber at two different times during the perfusion. Perfusion experiments performed without collagen served as controls.
Ex Vivo Perfusion Experiments
Ex vivo perfusions18 were performed at 37°C with
either a collagen-coated or an uncoated (control) Thermanox coverslip
positioned in a parallel-plate perfusion chamber. Fibrillar type III
collagen prepared from human placenta was sprayed onto washed
coverslips as previously reported.14 The final collagen
density was about 20 µg/cm2, which gives a maximal
thrombogenic stimulus without triggering fibrin
deposition.19
Venipuncture of an antecubital vein was performed with a Butterfly Infusion Set (No. 19). Nonanticoagulated blood (150 mL) was drawn directly through the chamber over the coverslip by a Gilson M312 occlusive roller pump at a constant flow rate of 10 mL/min. The pump was placed distal to the perfusion chamber. The flow rate was controlled by monitoring the blood volume in a graduated cylinder. The cross-sectional dimensions of the flow channel were chosen so that a wall shear rate characteristic of moderately stenosed arteries (2600 s-1) was obtained at the collagen surface. Blood perfusion lasted for 15 minutes. Postperfusion, fixation, and epoxy resin embedding procedures were performed as previously described.14
Perfusion experiments with collagen surface and naked control surfaces were performed on the same day with at least 2 hours between perfusions (in opposite arms) in the same donor. The perfusion to be run first was alternated between donors.
Morphometry
The blood–surface interactions were quantified on semithin
(1-µm) sections prepared perpendicular to the direction of the blood
flow 1 mm downstream of the upstream edge of the coverslip. The
sections were stained with basic fuchsin and toluidine
blue.20
Thrombus area (µm2/µm sectional length) was assessed by computer-assisted morphometry (Kontron Vidas image-analyzing unit). Thrombus volume (µm3/µm2) was derived from the sectional thrombus area as previously described.21 The evaluations were carried out at x500 magnification.
Antibodies
The following murine monoclonal antibodies were used:
PAC-122 (The University Cell Center of Pennsylvania), 30
µg/mL (final concentration), specific for the
activated form of the platelet gpIIb–IIIa complex;
anti-CD63 (RUU-SP 2.28)23 (kindly provided by Dr
Nieuwenhuis, Utrecht, Netherlands), 2.5 µg/mL,
specific for the lysosomal membrane protein gp53 exposed in the plasma
membrane upon platelet activation; FITC-conjugated anti-CD42a
(Becton Dickinson), 3.6 µg/mL, specific to the gpIX part of
the gpIb–IX complex; PE-conjugated anti-CD62P (Becton Dickinson), 0.43
µg/mL, specific to the internal -granule membrane protein
P-selectin expressed on the surface of activated platelets;
and FITC-conjugated anti-CD61 (Becton Dickinson), 3.6 µg/mL,
specific to gpIIIa to identify the platelet population.
Isotype-, fluorochrome-, and protein concentration-matched controls were run in parallel to all monoclonal antibodies: FITC-conjugated IgG1 (Sigma Chemical Co), PE-conjugated IgG1 (Becton Dickinson), IgG2b (Sigma), and IgM (Sigma).
Polyclonal FITC-conjugated rabbit anti-human fibrinogen (DAKO A/S), 100 µg/mL, was used to detect fibrinogen binding to the activated gpIIb–IIIa complex. PE-conjugated F(ab')2 fragments of rabbit anti-mouse Ig (DAKO), 58 µg/mL, were used as secondary antibody to detect binding of unconjugated PAC-1 and anti-CD63.
Blood Sampling and Preparation of Platelets for Flow
Cytometry
After venipuncture of an antecubital vein and before
connecting the bloodstream to the perfusion chamber, the first 2.7 mL
of blood was collected into EDTA (tripotassium salt, final
concentration 5.5 mmol/L, Monovette) for determination of
platelet count and hematocrit (Technicon H2). The subsequent 4.1 mL
was collected into EDTA and sodium citrate (0.1 vol 106
mmol/L trisodium citrate) as a reference sample for flow
cytometry.
After 4.5 and 14.5 minutes of perfusion, blood samples were collected immediately distal to the perfusion chamber, as previously described.10 The samples were collected successively during 30 seconds into syringes containing EDTA (2 mL) and sodium citrate (0.5 mL).
EDTA blood was used to investigate the surface expression of CD42a, CD61, CD62P, and CD63. Immediately after sampling, whole blood was fixed with PFA (final concentration 0.5% wt/vol) in PBS, pH 7.4. Platelet-rich plasma was prepared by centrifugation (200g, 10 minutes at 4°C) and kept on ice until further use. Platelet-rich plasma (25 µL) was added to 10 µL of prediluted antibody and incubated for 20 minutes at 37°C and 100% humidity. The platelets of CD63 tubes were washed once at 1500g and 4°C with 4 mL of cold PBS supplemented with 5.6 mmol/L glucose and 3.5 g/L bovine serum albumin before an additional 20 minutes of incubation with secondary antibody at room temperature. Two milliliters of 1% (wt/vol) PFA stopped the reaction with the antibodies, and the tubes were stored in the dark at 4°C until flow cytometry analysis could be performed.
Sodium citrate blood was used to study the activation-dependent conformational change in the platelet gpIIb-IIIa complex and fibrinogen binding to platelets according to a modification of the whole-blood flow cytometry method described by Shattil and colleagues.24 Immediately after sampling, sodium citrate whole blood was diluted 1/10 in cold PBS with glucose and bovine serum albumin and placed on ice. Within 1 hour, 50 µL of diluted blood was added to monoclonal antibody PAC-1 or polyclonal anti-fibrinogen and incubated for 15 or 20 minutes, respectively, at 22°C. The PAC-1 tubes were incubated for an additional 15 minutes with secondary antibody before final dilution with 0.5 mL of cold 1% (wt/vol) PFA.
Flow Cytometric Analysis
The samples were analyzed within 24 hours in a Becton
Dickinson FACScan flow cytometer. The fluorescence intensity
was calibrated daily with Standard-Brite fluorescent
microspheres according to the manufacturer's instructions. The
platelet population was identified by means of its light scatter
characteristics and enclosed in an electronic gate. Light scatter and
fluorescence data were obtained with gain settings in the
logarithmic mode from 10 000 events collected from each sample. The
saved list mode files were converted by a Hewlett Packard convertor and
analyzed using PCLYSYS software.
The identity of the platelet population was confirmed by being more than 98% positive for CD61. Antibody-positive cells were defined as those platelets showing a fluorescence intensity >97.5% of platelets incubated with the respective isotype controls.
Statistical Analysis
The flow cytometry results were expressed as the median MFI.
Data were compared by the Mann-Whitney U test to avoid any
assumptions about the distribution of the material. Values of
P<.05 were considered significant.
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Results |
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Hematological Parameters
All individual hemoglobin and platelet concentrations and hematocrits were within normal range.
Thrombus Volume
Because calculation of thrombus volume represents an
end-point evaluation, the results of 15-minute perfusions at a wall
shear rate of 2600 s-1 were compared with
previously published results17 25 from 5-minute perfusions
at the same wall shear rate with the same reactive surfaces (Fig 1
). Although the thrombi formed on
collagen nearly occluded the chambers after 15 minutes of perfusion,
only small thrombi were measured on the naked coverslips. The median
thrombus volume on collagen was 105
µm3/µm2, whereas the corresponding value on
the control surfaces was 15.7
µm3/µm2.
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.05 and
**P
Conformational Changes in the Platelet Fibrinogen
Receptor
The PAC-1 antibody has a specificity for an epitope that is
exposed on the fibrinogen receptor gpIIb–IIIa only after an
activation-dependent conformational change in the receptor has
occurred. A significant increase in binding of this antibody to the
platelets was measured in the blood passing the growing thrombi on
the collagen surface (Fig 2A). The median
MFI increased from 56 before the perfusion to 78 at 5 minutes of
perfusion and to 277 at 15 minutes of perfusion.
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) or
plain coverslips (
) (controls). Platelets collected in the
bloodstream passing the growing thrombi were incubated with different
antibodies and analyzed with flow cytometry to detect surface
changes: A, incubation with PAC-1; B, anti-fibrinogen; C, anti-CD62P;
D, anti-CD63; E, anti-CD42a; and F, anti-CD61. Data are
presented as median (quartiles) MFI, n=3 to 8.
*PIn the control perfusions with no collagen, PAC-1 binding was slightly, but not significantly, increased from the median MFI of 56 to 66 during 15 minutes of perfusion.
When the results were calculated as the percentage of PAC-1–positive cells compared with the antibody isotype control, the collagen perfusions recruited more circulating activated platelets than the control perfusions. This difference became more obvious with time. With collagen, the fraction of positive cells increased from 14% before the perfusion to 27% at 5 minutes and to 53% at 15 minutes of perfusion. The corresponding results from the control perfusions were 20% at 5 minutes and 28% at 15 minutes.
Fibrinogen Binding to the Platelets
The conformational change in the fibrinogen receptor permits
binding of plasma fibrinogen to platelets. The binding of
fibrinogen to the platelets was detected by binding of
FITC-conjugated anti-fibrinogen (Fig 2B
). In contrast to PAC-1 binding,
there was no significant increase in fibrinogen binding at 5 minutes in
the collagen perfusions. However, fibrinogen binding at 15 minutes was
dramatically increased from the baseline median MFI of 14 to 982. No
significant increases over baseline values were detected in the control
perfusions.
No significant increase in the percentage of fibrinogen-positive cells was detected in the control perfusions or at 5 minutes of perfusion with collagen. However, at 15 minutes of perfusion with collagen, 74% of the platelets passing the thrombus were positive for fibrinogen binding, in contrast to the 2.5% fibrinogen-positive cells found before the perfusion.
-Granule Release
The detection of the -granule membrane protein CD62P
(P-selectin) on the surface of platelets with a PE-conjugated
antibody was used as a marker of -granule release (Fig 2C). Median
MFI increased significantly from 2.6 in the preperfusion blood samples
to 8.9 in blood sampled at 5 minutes and to 84.5 in blood sampled at 15
minutes of perfusion with collagen. Also, a significant increase was
found in the 15-minute values in the control perfusions with a median
MFI of 7.2.
The CD62P surface expression was also calculated as the percentage of CD62P-positive cells. From a baseline value of 6.3%, the fraction of positive cells increased to 41% at 5 minutes and to 84% at 15 minutes with collagen. In the control perfusions, 7.5% and 33% CD62P-positive cells were found at 5 and 15 minutes, respectively.
Exocytosis of Lysosomal Granules
To investigate whether the perfusion conditions also induced
secretion of lysosomal granules, the platelets were stained with
anti-CD63. This antibody selectively binds to the membrane protein
gp53, which is localized in the lysosomes in
unactivated platelets but exposed on the plasma membrane
after platelet activation.
Fifteen minutes of perfusion with collagen was the only condition that
significantly increased the surface expression of CD63 over baseline
values (Fig 2D). The median MFI increased from 26 to 96.
Correspondingly, the fraction of CD63-positive cells increased from
8.4% before the perfusion to 19.5% at 15 minutes of perfusion with
collagen.
Changes in CD42a Expression
FITC-conjugated anti-CD42a binding to the gpIX part of the
gpIb–IX complex was used to determine whether any change in the
surface expression of this gp was induced in platelets exposed to
collagen. A decrease, albeit statistically insignificant, was noted
(Fig 2E). The median MFI changed from the baseline value of 123 to 99
at 15 minutes of perfusion with collagen.
Control of Platelet Population
All platelets are CD61 positive. FITC-conjugated anti-CD61 was
used as a control to ensure that the events collected from the flow
cytometer represented platelets. All except three
samples displayed >99% (median, 99.8%) CD61-positive particles. In
three highly activated samples collected after 15 minutes of
perfusion with collagen, the CD61-positive fractions were reduced to
96%, 95%, and 73%. This phenomenon was combined with a marked
reduction in intact circulating platelets (observed as a strikingly
reduced collecting rate, ie, events/second, during the flow cytometry
analysis) and was attributed to high consumption of
platelets in the growing thrombi.
Correlation Between Activation of Circulating Platelets and
Thrombus Formation
The MFIs for PAC-1, anti-fibrinogen, anti-CD62P, anti-CD63, and
anti-CD42a at 15 minutes of perfusion on collagen and control surfaces
were correlated with the respective thrombus volume
(µm3/µm2)
(Table). For the first four antibodies, a
correlation coefficient of .61 to .91 was obtained. Surface expression
of CD42a correlated inversely with thrombus growth
(r=-.81).
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Discussion |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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In the present study we evaluated platelet activation in platelets passing growing thrombi in a human ex vivo model of thrombogenesis. At a wall shear rate characteristic of moderately stenosed arteries (2600 s-1), platelet activation in the circulating platelets was clearly correlated with thrombus growth. Flow cytometry revealed that both the degree of activation (MFI) and the fraction of platelets activated (percent positive for the activation markers) increased as the thrombi grew. Furthermore, our results may imply that the different activation markers have distinct sensitivity to the thrombotic process.
-Granule release detected by surface expression of CD62P
(P-selectin) seemed to be the most sensitive marker. Although the
results clearly revealed significantly higher MFIs and percentages of
positive cells with the collagen surface, the surface expression of
CD62P was also significantly increased in the controls. It has been
shown in the cone-and-plate viscometer that shear alone is capable of
initiating von Willebrand factor-dependent platelet
aggregation in the absence of exogenous agents.26 27 Shear
stress-induced von Willebrand factor binding to platelet
gpIb has been found to initiate calcium influx28 29 and
activation of protein kinase C,30 both of which are known
to trigger platelet secretion.31 Our results confirm
that -granule release can be elicited by shear forces in flowing
native blood. However, -granule release is enhanced by the presence
of a collagen surface in a biological normocalcemic environment.
The conformational change in gpIIb-IIIa as detected by PAC-1 also seems to be a sensitive marker of platelet activation in platelets passing growing thrombi. In our study, this conformational change seemed to precede binding of fibrinogen as detected by anti-fibrinogen. Although PAC-1 binding was significantly increased at both 5 and 15 minutes, no binding of fibrinogen was detected at 5 minutes. However, a dramatic increase in fibrinogen binding was detected at 15 minutes. Interestingly, this coincided with flow changes in the chamber because the flow channel at that time was nearly occluded. The growing thrombi change the local flow conditions, making them less predictable at 15 minutes of perfusion. In previous studies fibrinogen has been found to mediate platelet aggregation at low shear rates,32 while its importance at high shear rates seems more dubious.33 The low platelet binding of plasma fibrinogen after 5 minutes of perfusion is suggestive of a physiologically more important role for another ligand, von Willebrand factor, in binding to gpIIb–IIIa at high shear rates, a fact already pointed out by other groups.33 34 35
The polyclonal rabbit anti-human fibrinogen antibody used does not discriminate between intact fibrinogen and fibrin. At 15 minutes of perfusion with increasing flow disturbances caused by the enlarging thrombi, coagulant activity is undoubtedly initiated. In three samples collected at 15 minutes of perfusion with collagen, the CD61-positive fractions (gpIIIa, platelet identification marker) were reduced to 96%, 95%, and 73%. The relatively large fraction of CD61-negative particles, at a size of intact platelets, may have been fragments of other blood cells or protein aggregates, possibly fibrin. Fibrin aggregates in these samples could also partly explain the dramatic increase seen in anti-fibrinogen binding. Use of a two-color flow cytometry technique, collecting events from particles positive for FITC-conjugated anti-CD61 only, would have prevented this bias.
Lysosome exocytosis detected by surface expression of CD63
(lysosome membrane protein gp53) has been shown to be a less
sensitive marker of platelet activation than -granule release in
different in vitro and clinical studies.36 37 38 This
impression was supported by our study, even though 15 minutes of
perfusion with collagen significantly increased the surface expression
of CD63.
Lack of significant downregulation of the gpIb–IX complex in spite of
the obvious platelet activation in our study may illustrate that
the redistribution of this complex to the open
canalicular system is a time-consuming process compared
with granule exocytosis and the conformational change of the
gpIIb–IIIa complex. Michelson and coworkers39
demonstrated in vitro that thrombin-induced -granule release is
nearly completed when downregulation of surface gpIb is initiated.
Thus, downregulation of the gpIb–IX complex as a sensitive activation
marker seems to be highly questionable, a view supported by several
clinical studies.38 40 41
In the ex vivo model of thrombogenesis, blood is sampled immediately downstream of the perfusion chamber and the growing thrombi without any dilution from other blood vessels. Even though the system is devoid of intact endothelium, it offers a unique possibility for studying mechanisms related to platelet activation. We observed a decrease in the intact platelet population passing the growing thrombi (data not shown). The bulk of the cleared platelets is thought to adhere to the growing thrombi, although the strong activation also generates microparticles and activated platelets adhering to leukocytes.42 In the present study, only intact platelets passing the growing thrombi were examined by flow cytometry. Apart from the three samples discussed above, all platelet populations were >99% positive for CD61.
Extensive thrombus growth is accompanied by an increase in activated circulating platelets. By showing this correlation between thrombus growth and activation of circulating platelets in an ex vivo model, information was gained on the dynamics of both thrombus formation and platelet activation in flowing native blood. As platelet thrombi develop, platelet-release reactions create an environment optimal for platelet recruitment and platelet aggregation. The increased local concentration of different agonists secreted or released by activated platelets amplifies the aggregation by an order of magnitude. Not all platelets are consumed by the thrombi; instead, they pass the cloud of platelet agonists, being activated but not trapped. Also, translocation of activated platelets from the surface of the thrombi to the bloodstream because of high shear stress may be an explanation since the thrombi narrow the lumen and thereby elevate both shear rate and shear stress. The local blood flow conditions inherently determine the rates of the different elements (both platelets and proteins) participating in platelet activation and aggregation and thrombus formation.
Thus, this study provided further information on the effects of thrombus formation at arterial flow conditions on circulating platelets and their state of activation in nonanticoagulated human blood.
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Selected Abbreviations and Acronyms |
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Acknowledgments |
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This study was supported financially by grants from Ada og Hagbart Waages humanitære og veldedige stiftelse. We acknowledge the excellent technical support of Bjørg Lund, Ullevaal University Hospital, Oslo, Norway, in preparing the microscopic sections. We also thank Dr L.M. Cunha-Ribeiro, University of Porto, Porto, Portugal, Dr P.G. DeGroot, University Hospital, Utrecht, Netherlands, and Prof K.S. Sakariassen, University of Oslo, Oslo, Norway, for stimulating discussions concerning the methodological aspects of this study.
Received March 28, 1996; accepted October 8, 1996.
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