© 2000 American Heart Association, Inc.
Thrombosis |
Coimmobilized Native Macromolecular Heparin Proteoglycans Strongly Inhibit Platelet-Collagen Interactions in Flowing Blood
From the Wihuri Research Institute, Helsinki, Finland (P.K., P.T.K., R.L.), and Helsinki University Central Hospital, Department Internal Medicine (R.L.).
Correspondence to Dr Riitta Lassila, Wihuri Research Institute, Kalliolinnantie 4, FIN-00140, Helsinki, Finland. E-mail riitta.lassila{at}wri.fimnet.fi
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Abstract—We coimmobilized mast cell–derived heparin proteoglycans (HEP-PGs) of very high molecular weight (750 kDa) or unfractionated heparin (UFH) on coverslips together with collagen without altering the amount of immobilized collagen. Subsequently, platelet-collagen interactions were studied under both flowing and static conditions in D-phenylalanyl-L-prolyl-L-arginine chloromethyl ketone–anticoagulated blood and platelet-rich plasma (PRP), respectively. At a high shear rate (1600 1/s), the mean platelet deposition (PD) on collagen monomers was 7.5±6.1x106/cm2 (n=5). When the monomers were coimmobilized with UFH, PD was inhibited by 73% (2.0±1.2x106/cm2), whereas HEP-PG completely blocked it (0.42±0.38x106/cm2; P<0.05). Also, when collagen fibrils were used for coating, HEP-PG significantly inhibited PD. At a low shear rate (200 1/s) and under static conditions in PRP, the inhibitory effect of HEP-PG on PD was less marked. Inhibition of glycoprotein IIb/IIIa did not affect PD on coimmobilized HEP-PG in contrast to coimmobilized UFH or collagen alone. As a sign of inactivation, platelets adhering to the HEP-PG surface released considerably less ß-thromboglobulin than did those adhering to pure collagen. In summary, immobilized HEP-PG strongly inhibited PD on collagen by attenuating adhesion-induced platelet activation. The stronger effect on collagen monomers suggests the inhibition of glycoprotein Ia/IIa–mediated activation.
Key Words: collagen • heparin proteoglycans • mast cells • platelets
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Introduction |
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Acute thrombotic complications after vascular interventions remain a clinical problem despite better understanding of the response to injury and pathophysiology of atherothrombosis.1 2 Recently, the selection of antithrombotic drugs has become more extensive, and several well-controlled studies have shown the efficacy of these systemic drug therapies.3 4 Platelets interacting with exposed collagen in the vascular wall play a pivotal role in the regulation of hemostasis.5 If there is a defect in the platelet glycoprotein (GP) receptors or if the synthesis of collagen is impaired, bleeding occurs.6 However, after arterial injury, be it due to plaque rupture or to angioplasty, a far more prevalent and clinically demanding end state is unregulated platelet activation leading to thrombosis. Therefore, local control of the thrombotic response in vascular procedures would be highly beneficial not only acutely, but very likely also chronically, to limit plaque growth or restenosis subsequent to arterial injury. Thus far, however, local approaches to prevent arterial thrombosis have been rather limited.
Experimental studies have suggested that GP Ia/IIa (integrin
2ß1) is the key
receptor mediating platelet-collagen interactions.7
Clearly, then, our therapeutic attempts to prevent collagen-induced
platelet activation should focus on this receptor. As yet, however,
we have lacked therapeutic tools to interfere with this particular
interaction in a clinical setting. Now it seems that mast cell–derived
heparin could be such a tool. Mast cells, which are the only source of
heparin in the body, are located throughout the connective and mucosal
tissues. Heparin can be stored and secreted by these cells in the form
of macromolecular proteoglycans (PGs).8 In the skin, where
mast cells are especially numerous and appear around small vessels,
there is a significant concentration of heparin with antithrombotic
potential.9 Indeed, in the skin, platelet plug
formation and subsequent thrombin generation were blunted on allergenic
activation of mast cells.10 Moreover, collagen-induced
platelet activation can be inhibited by the very large (750-kDa)
macromolecular heparin proteoglycans (HEP-PGs) derived from rat serosal
mast cells.11 This inhibition by soluble HEP-PGs involves
their tight binding to von Willebrand factor (vWF) and relies
on impaired GP Ia/IIa–mediated platelet
activation.11 12
Because mast cells are also present in the arterial intima and because they can release their contents into the surrounding collagen-containing matrix, our objective here was to assess whether immobilized mast cell–derived HEP-PGs can affect platelet-collagen interactions. We found that collagen-immobilized HEP-PGs strongly inhibited platelet activation and aggregation and that this inhibitory response was enhanced at a high shear rate. Thus, not only soluble, but also collagen-immobilized, HEP-PGs are strong physiological inhibitors of platelet-collagen interactions.
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Methods |
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Isolation of HEP-PGs From Rat Peritoneal Mast Cells
The lavage method used for isolating mast cells from rat peritoneal cavities has been described elsewhere.13 After isolation, the mast cells were treated with compound 48/80 (Sigma Chemical Co) to induce their degranulation.14 The degranulated mast cells and their exocytosed granules, ie, granule remnants, were then sedimented by 2 consequent centrifugations to obtain granule-free supernatant.11 The concentration of the very large, soluble HEP-PGs (750 kDa; range, 500 to 1000 kDa) in the supernatant was determined by the Alcian blue method (Fluka, Buchs) or by Blyscan glycosaminoglycan (GAG) assay (Biocolor Ltd), after which the supernatant was divided into aliquots and frozen at -20°C until use.
Blood Collection, Preparation of Platelet-Rich Plasma, and
Labeling of Platelets With
[3H]Serotonin
The study was approved by an institutional review board. After
they had given their informed consent, blood was collected from healthy
volunteers (n=49), who reported not having taken any medication during
the previous 14 days. The volunteers avoided coffee, tea, and dietary
fats in the morning before blood sampling but were not required to fast
completely. The superficial cubital vein was cannulated (Venflon 2,
17-gauge; BOC Ohmeda AB), and 9 volumes of free-flowing blood was
collected into 1 volume of 300 µmol/L
D-phenylalanyl-L-prolyl-L-arginine
chloromethyl ketone (PPACK; Calbiochem-Novabiochem Co).
Platelet-rich plasma (PRP) was prepared by
centrifugation (180g, 12 minutes, 22°C).
The final platelet count in PRP was adjusted to
300x106/mL. The platelets in PRP were
labeled with [3H]serotonin as
previously described and validated.15 After being
labeled, PRP was added to the remaining blood components to
reconstitute the blood used in perfusions. To stabilize the blood,
perfusion experiments were not started until 30 minutes had elapsed
after reconstitution.
Preparation of Collagen-Coated (±Unfractionated Heparin or
HEP-PGs) Coverslips for Studies of Platelet Interaction in PRP and
for Perfusion Studies
The coverslips used for platelet interaction studies
in PRP were coated with collagen type I extracted with acetic acid from
bovine Achilles tendon and treated with pepsin to yield
monomers.12 In these experiments altogether, 6 different
coating solutions were used. The solutions were prepared by mixing 2
different concentrations of collagen (coll, 10 or 50 µg/mL) with
either unfractionated heparin (10 µg/mL UFH; Lövens) or
with HEP-PGs (10 µg/mL) in PBS. The coating solutions (200 µL) were
then added to round Thermanox (Nunc) coverslips and incubated for 60
minutes at 37°C in a humid atmosphere. The coating solutions were
removed, 200 µL of 2% human serum albumin (HSA) was added as
a blocking solution, and the incubation was continued for another 60
minutes at 37°C. Each time, the coating process was performed
similarly by the same trained investigator within a time frame of 2
hours before platelet interaction studies in PRP or before
perfusion experiments. To ensure that the coating was
homogeneous, some of the coverslips were stained with
Coomassie brilliant blue R250 and viewed by light microscopy.
Two different type I collagen preparations were employed for perfusion studies. Collagen monomers (final concentration, 50 µg/mL) were used as a coating substrate for most of the whole-blood perfusion studies. For comparison, we also studied the effects of collagen fibrils on platelet interactions (collagen reagent Horm, Nycomed Arzneimittel). The amount of collagen adhering to the coverslips was quantified by a specific dye-binding assay (Sircoll, Biocolor Ltd).16 Because it was observed that higher amounts of fibrils than monomers adhered to the surface during the coating procedure, collagen fibrils at a concentration of 16 µg/mL were used to make the coating comparable to that with 50 µg/mL of monomers. In the perfusion studies, we also compared the effects of collagen-immobilized HEP-PGs (coll–HEP-PG, 50 µg/mL of collagen monomers or 16 µg/mL of collagen fibrils together with 10 µg/mL of HEP-PGs) with those of UFH. Here, however, we used a 1000-fold concentration of UFH (10 mg/mL) together with the collagens, as reported above. The Thermanox coverslips (22x60 mm) were divided into 4 identical pieces to fit the perfusion chamber. Then, 400 µL of coating suspension was incubated over the coverslips, and the coating was finished with 2% HSA, as described above.
Immobilization of vWF, Collagen, UFH, and HEP-PGs on Microtiter
Wells for PRP Studies Under Static Conditions
For some platelet interaction experiments in PRP, plastic
microtiter wells (Labsystems) were coated with vWF, 10 µg/mL
(Calbiochem), collagen (10 µg/mL), UFH (10 µg/mL), or HEP-PGs (10
µg/mL) or with combinations of these substrates. Here, too, the wells
were blocked with 2% HSA, and the principle of the coating procedure
was the same as presented above.
Quantification of Collagen and GAGs Deposited on Coverslips
After Coating
The effect of coimmobilization of UFH and HEP-PGs on the amount
of adherent collagen was also quantified by Sircoll assay. Analogously,
adherence of the GAGs to the coll-UFH and coll–HEP-PG coverslips was
analyzed by Blyscan assay, which is a quantitative method based
on the specific binding of the cationic dye 1,9-dimethylmethylene to
PGs and sulfated GAGs.17 Because the substrates to be
quantified were immobilized on the plastic coverslips at
relatively low amounts, we modified the original instructions
accordingly. In brief, the color reagents were added to 3 coverslips,
and the detached coatings were pooled before dissociation of the color
with the specific dissociation reagent provided. Finally, the
absorbance of the bound dye was determined by using a
spectrophotometer, and the amount of immobilized substrate
per coverslip was approximated.
Platelet-Collagen and Platelet-vWF Interactions in
PRP
The platelet-collagen and platelet-vWF interaction
studies were performed in PRP under static conditions as previously
described.11 In brief, the coated Thermanox coverslips
were placed on the bottom of precoated (2% HSA) 24-well plates (Nunc).
One milliliter of [3H]serotonin (10
nmol/L)–labeled PRP (platelet count adjusted to
300x106/mL) was then added to the wells to be
incubated for 30 minutes at 37°C under slow rotation (100 rpm). To
reduce the amount of substrates needed for the coatings, some
experiments were performed with a slight modification by using
precoated, detachable microtiter wells. After incubation, the
coverslips or the wells were washed with PBS to remove unattached
platelets, and the deposited 3H scintillation
activity was measured. In some experiments, 10 µg/mL of c7E3, a GP
IIb/IIIa inhibitor (abciximab; ReoPro, Centocor), was added
to block thrombus formation. Furthermore, some experiments were
performed with unlabeled PRP, and the binding of PRP-derived vWF to
collagen was detected immunologically. After incubation under rotation,
the coverslips were washed meticulously with 0.1% Tween-PBS (3x 5
minutes) and placed on new 24-well plates. They were then incubated
with peroxidase-conjugated rabbit anti-human vWF (diluted 1:4000; Dako)
for 2 hours at 22°C. After additional washing of the coverslips with
1% bovine serum albumin in PBS, the color reaction was
developed with the use of TMB microwell peroxidase substrate system
(Kirkekaard & Perry Laboratories Inc), and the absorbance was measured
at 450 nm with a Labsystems multiscan MCC.
ß-Thromboglobulin Assay
Platelet-collagen interaction experiments were performed in
PRP as described above. After 30-minute incubation at 37°C, 450 µL
of PRP was added to 50 µL of anticoagulant mixture containing (final
concentrations) citric acid (3.5 mmol/L), trisodium citrate
(7.5 mmol/L), dextrose (13.6 mmol/L), EDTA (0.6 mol/L),
adenosine (0.6 mol/L), hirudin (2.5 U/mL), and UHF (2.5
U/mL).18 The anticoagulated PRP was then
centrifuged (1500g) for 5 minutes at 4°C, and the
middle third portion of the supernatant was separated and frozen at
-40°C. The ß-thromboglobulin contents of these
samples were determined by ELISA
(Asserachrom®ß-TG, Diagnostica
Stago; normal range in plasma, 10 to 40 IU/mL) within 2 weeks of sample
collection.
Platelet-Collagen Interactions Studied Under Whole-Blood
Perfusion
Details of the whole-blood perfusion experiments are described
elsewhere.15 The blood to which
[3H]serotonin-labeled platelets
were added was divided into aliquots (30 mL) and prewarmed for 5
minutes at 37°C before the perfusion. A coated coverslip was placed
in a Badimon perfusion chamber with well-defined rheological
characteristics.19 The prewarmed blood was then
recirculated for 5 minutes through the chamber in which the coated
coverslip was exposed to flowing blood. The flow rate was either 30
mL/min (high shear) or 10 mL/min (low shear), corresponding to wall
shear rates of 1600 and 200 1/s, respectively. Immediately after the
perfusions, the unattached platelets were flushed away by perfusing
the coverslip with PBS for 40 seconds by using the same flowing
conditions, after which the deposited 3H
scintillation activity was determined. The blood samples for
platelet count, total blood 3H scintillation
activity, and plasma 3H scintillation activity
(serotonin release) were collected just before and after
the perfusion as previously described.15
Scanning Electron Microscopy
The platelet deposition on coll, coll-UFH, and coll-HEP-PG
was also studied by scanning electron microscopy (SEM), and in these
experiments, whole blood without platelet labeling was used. After
the perfusions, the coverslips were prepared for SEM by immediate
fixation in 2.5% glutaraldehyde–PBS for 2 hours at
22°C. The fixed samples were carefully rinsed in PBS to remove excess
fixative and kept in PBS until dehydration treatment with a series of
increasing concentrations of ethanol. The dried SEM samples were
sputter-coated with platinum and examined on a JEOL SEM 820 at the
Electron Microscopy Unit, Institute of Biotechnology, University of
Helsinki, Helsinki, Finland.
Statistical Analysis
The results are shown as mean±SD unless otherwise
indicated. The statistical significance of the differences observed in
platelet deposition on the differently coated surfaces was
evaluated by a nonparametric Wilcoxon signed rank
test. Statistical significance was set at P<0.05.
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Quantification of Collagen and GAGs Deposited on Coverslips After Coating
To quantify the adherence of collagen, UFH, and HEP-PGs, the contents of collagen and GAGs on the coated coverslips were studied by quantitative dye-binding assays and viewed by light microscopy after Coomassie blue staining. The observed mean content of collagen was 3 µg/cm2, and there were only minor differences between the coll, the coll-UFH, and the coll–HEP-PG coverslips (Table
). The collagen coating was homogenous in
the presence of UFH and HEP-PGs when viewed after Coomassie blue
staining. The mean content of GAGs on the coll–HEP-PG–coated
coverslips was 130 ng/cm2, and the amount of GAGs
was only slightly higher on the coll-UFH coverslips despite the
1000-fold higher GAG concentration in the coll-UFH coating
solution.
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Interaction of PRP With Collagen Versus
Collagen-Immobilized UFH or HEP-PGs
To evaluate the ability of collagen-immobilized
UFH and collagen-immobilized HEP-PGs to inhibit
platelet-collagen interactions, we incubated PRP (rotation 100 rpm,
30 minutes, 37°C) on coverslips coated with collagen monomers and
quantified the attached platelets. When either a smaller (10
µg/mL) or larger (50 µg/mL) amount of collagen was used, both
additional UFH and HEP-PGs decreased platelet deposition. The
effect of HEP-PGs was consistently better than that of UFH
(Figure 1). The addition of c7E3 to PRP
before incubation decreased platelet deposition on the coll and
coll-UFH coverslips. In contrast, platelet deposition on the
coll–HEP-PG coverslips was not affected, suggesting that HEP-PG
blunted platelet aggregation induced by collagen (Figure 2).
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P<0.05 UFH vs
HEP-PGs. Platelet deposition on coll 50 µg/mL (VEH) was higher
than on HSA (A) (P<0.05), but coll 10 µg/mL (VEH) and HSA
did not differ statistically.
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ß-Thromboglobulin
To study -granule release during the platelet-collagen
interaction, we determined the content of
ß-thromboglobulin from PRP, which was incubated
on coverslips coated as described above. The highest
ß-thromboglobulin levels (809±148 IU/mL,
mean±SD) were found in PRP that had been in contact with coverslips
coated with collagen (50 µg/mL) alone, and they differed markedly
from those (480±157 IU/mL; P<0.05) obtained with
albumin-coated coverslips. Addition of UFH or HEP-PGs to the
coating mixture lowered the ß-thromboglobulin
levels (699±139 and 576±130 IU/mL, respectively; P<0.05)
in PRP. The difference between UFH and HEP-PGs was also significant
(P<0.05), compatible with stronger HEP-PG–induced
platelet inactivation.
Detection of Platelet-Derived vWF
To assess the amount of PRP-derived vWF deposited on the surfaces,
some experiments were performed with unlabeled PRP, and the
surface-bound vWF was immunologically detected. The mean absorbance
representing bound PRP-derived vWF on the collagen-coated
coverslips was 1.21±0.18 (n=3, mean±SD). In the presence of a high
concentration (10 mg/mL) of UFH, the surface-associated vWF was
markedly decreased (absorbance 0.65±0.04), whereas HEP-PGs (10
µg/mL) nearly abolished the platelet-derived deposition of vWF
(absorbance 0.27±0.09). The latter absorbance values did not differ
from those obtained with vWF alone (5 µg/mL of purified vWF in 2%
BSA, absorbance 0.27) or with the platelet-poor plasma standard
(absorbance 0.20). Because HEP-PGs reduced the amount of platelet
deposition to the level detected on albumin (Figure 1
)
and similarly the vWF binding to the level of plasma concentration of
vWF, this observation is consistent with the inhibition of
platelet activation and -granule release.
Interaction Between Platelets and Immobilized vWF
Incubation of PRP on vWF-coated coverslips at moderate
rotation showed that vWF-immobilized HEP-PGs decreased
platelet deposition (Figure 3).
Immobilization of collagen together with vWF increased the basal
deposition of platelets, as expected, whereas addition of either
HEP-PGs or UFH to the vWF-collagen coating mixture decreased
platelet deposition, the effect of HEP-PGs being slightly weaker
than that of UFH.
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Platelet Deposition on Collagen in Flowing Whole Blood
In perfusion studies, we wanted to examine whether the results
obtained in the platelet interaction studies would also apply under
flowing conditions. First, we coated the coverslips with collagen
monomers and used a low shear rate (200 1/s) for the blood perfusions.
At the low shear rate, the HEP-PGs (10 µg/mL) immobilized
together with collagen considerably decreased platelet deposition
(2.5±0.8 versus
1.5±0.2x106/cm2;
P<0.05), whereas UFH (10 mg/mL) failed to have an effect of
similar magnitude
(1.8±1.4x106/cm2; NS)
(Figure 4A). When collagen-induced
platelet deposition was studied at a high shear rate (1600 1/s),
the mean platelet deposition on collagen increased 3-fold and was
7.5±6.1x106/cm2 (Figure 4B). At the higher shear rate, both UFH and HEP-PGs had a
strong inhibitory effect on platelet deposition
(2.0±1.2 and
0.42±0.38x106/cm2,
respectively; P<0.05), and HEP-PGs actually fully abolished
platelet deposition. We also made several attempts to apply HEP-PGs
to the coverslips after collagen coating, but these experiments failed
to show any significant effect of HEP-PGs on platelet deposition,
although the HEP-PGs were found to bind to collagen (data not
shown).
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The use of native-type collagen fibrils (16 µg/mL) as the coating
substrate (instead of collagen monomers) led to an expected increase in
the deposition of platelets (Figure 4C). When
immobilized collagen fibrils were exposed to flowing blood
at the high shear rate, the coimmobilized HEP-PGs markedly
inhibited platelet deposition (62% decrease, P<0.05),
whereas UFH had a smaller but consistent lowering effect (12%
decrease; P<0.05) (Figure 4C).
The actual presentation of platelets on the coverslips
was examined by SEM. The surface coverage and size of aggregates
appeared larger in the collagen monomer–coated coverslips than in the
coll-UFH or the coll–HEP-PG coverslips, thus according with the
quantitative data presented above (Figures 5A through 5F). Moreover, the coll-UFH
coverslips presented some large aggregates, but most of the
adhered platelets were located at the margins of the perfusion
channel, where shear rates are known to be lower than in the middle of
the channel (Figures 5C through 5D). The coll–HEP-PG coverslips
showed only occasional aggregates, which likewise were mainly located
in the marginal zone of the channel, and only few platelets were
seen in the center of the channel (Figures 5E and 5F).
Similarly, when collagen fibrils were exposed to flowing blood, the
surface coverage and size of aggregates decreased on the coll-UFH and
especially on the coll–HEP-PG coverslips (Figures 6A through 6F). However, on the
surfaces coated with collagen fibrils and with UFH or HEP-PG, the
platelet aggregates were more evenly distributed, and there were no
large bare areas as typically seen in the center of the perfusion
channels on surfaces coated with collagen monomers and HEP-PGs.
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We have shown here that macromolecular HEP-PGs, when immobilized together with type I collagen, can inhibit platelet-collagen interactions. To mimic the conditions during mast cell activation in the human vasculature when mast cells release their contents, including HEP-PGs, we decided to expose blood to immobilized collagen in the presence or absence of coimmobilized HEP-PGs.
In these perfusion experiments, the interaction between platelets and collagen monomers was severely compromised by HEP-PGs under high shear–rate conditions, but the interaction was also inhibited, albeit to a lesser degree, at a low shear rate. Furthermore, HEP-PGs immobilized together with collagen fibrils inhibited platelet deposition, supporting our previous observations from studies with soluble HEP-PGs in blood interacting with collagen fibrils, optimally treated for fibril formation.11 UFH had an inhibitory effect on platelet deposition at a high shear rate, although it was much weaker than that of HEP-PGs and was absent at a lower shear rate. In the platelet interaction studies in PRP, both UFH and HEP-PGs markedly reduced platelet deposition, and again the effect of HEP-PGs predominated over that of UFH. These findings are compatible with the notion that HEP-PGs inhibit GP Ia/IIa–mediated platelet activation, which is crucial in mediating thrombus growth on collagen monomers at a high shear rate but not at a low shear rate.7 11 12 20 21 This inhibiting effect of immobilized HEP-PGs, differing from that of UFH, was also seen on collagen fibrils, though less clearly than on monomers, suggesting additional platelet-activating domains on the fibrils.12 22 HEP-PGs also effectively inhibited release of ß-thromboglobulin and blunted vWF exposure on the monomer-adherent platelets, in parallel with decreased activation. Finally, in contrast to that on collagen alone and to that on coimmobilized UFH, platelet deposition on HEP-PG–coated collagen surfaces could not be inhibited by c7E3.
In most experiments, we chose to use collagen monomers, which were obtained by extracting collagen with acetic acid from bovine Achilles tendons and subsequently subjecting the extracts to extensive pepsin treatment.12 This treatment proteolyzes the telopeptide ends of the collagen fibers, resulting in monomeric strands without the characteristic quaternary structure of collagen.23 Due to collagenolytic activity in eroded atherosclerotic plaques,24 collagen fibrils of the vessel wall may partly present themselves as monomers. Thus, platelet interactions with both fibrils and monomers seem to be pathophysiologically relevant. The stronger inhibition of platelet interaction with collagen monomers than with fibrils by HEP-PGs could be due to the efficacy of HEP-PGs in inhibiting GP Ia/IIa–mediated activation11 and subsequent recruitment of platelets to the monomers.12 Savage et al21 have recently shown that, at a high shear rate, blockage of GP Ia/IIa by a monoclonal antibody completely abolishes stable platelet attachment and thrombus formation on soluble collagen monomers.
We observed that collagen-immobilized HEP-PGs, likely owing to their macromolecular size and high negative-charge density, exceeded UFH in their inhibitory effect on platelet deposition. We also applied HEP-PGs to the coverslips after they had been coated with collagen. Interestingly, however, HEP-PGs did not significantly affect platelet deposition when applied to such preimmobilized collagen. Previously, we have shown that during short incubation periods, HEP-PGs do not bind to collagen, which is relevant for the strong inhibitory effect typical for soluble HEP-PGs.11 In the present study, however, incubation of HEP-PGs with immobilized collagen for 1 hour resulted in HEP-PG binding to the preimmobilized collagen. The lack of inhibitory effect of HEP-PGs in this system can be explained if HEP-PGs effectively bridged collagen and plasma vWF. As evidenced in the static experiments, HEP-PGs inhibited vWF-mediated platelet function less effectively than did UFH. HEP-PGs could also impair fibril formation, but this is an unlikely explanation because in our previous study, soluble HEP-PGs were highly effective when collagen was allowed to form fibrils before blood perfusion.
Regarding the potential roles of mast cells in the arterial intima, experiments in cell culture have suggested that degranulation of mast cells is potentially a proatherogenic event. Thus, stimulated mast cells promote the formation of foam cells when cocultured with macrophages or with smooth muscle cells.25 The potential roles of mast cells in the fibrous caps of advanced atherosclerotic plaques include mast cell protease–dependent activation of matrix-degrading metalloproteinases.26 If operative in vivo, this effect would tend to weaken the fibrous cap of the atherosclerotic plaque and so predispose to plaque rupture and initiation of thrombosis. On the other hand, the significant inhibitory capacity of the mast cell–derived HEP-PGs in platelet reactivity toward collagen would tend to limit thrombus growth.
Although novel vascular therapies such as percutaneous transluminal coronary angioplasty have revolutionized the management of arterial disease, acute thrombotic complications, restenosis in particular, still limit the efficacy of these treatment options.27 Clearly, the local control of collagen-induced platelet activation during and after vascular interventions would be crucial to avoid early occlusions and perhaps also to improve long-term patency. This study shows that collagen-immobilized HEP-PGs strongly inhibit platelet deposition to collagen and subsequent release reaction under conditions comparable to those present in atherosclerotic coronary arteries. These observations suggest that degranulating mast cells at the site of vascular injury, by secreting HEP-PGs, actually tend to prevent thrombus growth and has also been noted as a regulatory element in cardiac (sub)endothelium after auricular thrombosis.28 However, further studies are needed to evaluate whether locally applied HEP-PGs or similar compounds can also affect platelet–vessel wall interactions in vivo.
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Acknowledgments |
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The financial support given by the Aarne Koskelo Foundation, the Finnish Foundation for Cardiovascular Research, the Finnish Society of Angiology, and the Ida Montin Foundation is gratefully acknowledged. We thank Marja Lemponen and Tuula Järvenpää for their skillful technical assistance.
Received February 5, 2000; accepted August 7, 2000.
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