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Blood, Vol. 95 No. 3 (February 1), 2000:
pp. 930-935
HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
Cellular origin and procoagulant properties of microparticles
in meningococcal sepsis
Rienk Nieuwland,
René J. Berckmans,
Sarah McGregor,
Anita N. Böing,
Fred P. H. Th. M. Romijn,
Rudi G. J. Westendorp,
C. Erik Hack, and
Augueste Sturk
From the Departments of Clinical Chemistry, General Internal
Medicine, and Clinical Epidemiology, Leiden University Medical Center,
Leiden, The Netherlands; Department of Physiology, University of
Edinburgh, Edinburgh, United Kingdom; University Hospital Vrije
Universiteit, Amsterdam; and CLB, Sanquin Blood Supply Foundation,
Amsterdam, The Netherlands.
 |
Abstract |
Patients with meningococcal sepsis generally suffer from
disseminated intravascular coagulation (DIC). The aim of this study was
to address whether these patients have elevated numbers of circulating
microparticles that contribute to the development of DIC. Plasma
samples from 5 survivors, 2 nonsurvivors, and 5 healthy volunteers were
analyzed for the presence of microparticles by flow cytometry.
Ongoing coagulation activation in vivo was quantified by enzyme-linked
immunosorbent assay of plasma prothrombin fragment
F1 + 2, and procoagulant properties of microparticles
in vitro were estimated by thrombin-generation assay. On admission, all
patients had increased numbers of microparticles originating from
platelets or granulocytes when compared with controls
(P = .004 and P = .008, respectively). Patients
had elevated levels of F1 + 2 (P = .004),
and their microparticles supported thrombin generation more strongly in
vitro (P = .003) than those of controls. Plasma from
the patient with the most fulminant disease course and severe DIC
contained microparticles that expressed both CD14 and tissue factor,
and these microparticles demonstrated extreme thrombin generation in
vitro. We conclude that patients with meningococcal sepsis have
elevated numbers of circulating microparticles that are procoagulant.
These findings may suggest a novel therapeutic approach to combat
clinical conditions with excessive coagulation activation.
(Blood. 2000;95:930-935)
© 2000 by The American Society of Hematology.
| |
Introduction |
Meningococcal sepsis is a life-threatening disease that
occurs most frequently during childhood and is characterized by
excessive activation of many cells and cascades, which results in
disseminated intravascular coagulation (DIC) and shock.1,2
Although it is clear that the cascade of inflammatory and clotting
reactions is triggered by the meningococcal bacteria, and particularly
by their release of endotoxin, the precise mechanisms underlying these
reactions are not well understood. For example, the mechanism underlying the development of DIC, a typical threatening feature of
meningococcal sepsis, is unknown.
Clotting requires the presence of phospholipid cofactors that serve as
a surface to assemble the various complexes to activate the clotting
factors. In vitro studies have shown that activated platelets, and in
particular microparticles generated from them, contain a large number
of binding sites for activated factor IX (IXa),3 factor
Va,4 and factor VIII5 and support both factor
Xa activity6,7 and prothrombinase activity.4,7 Increased numbers of platelet-derived microparticles are present in the
circulation of patients who have an increased risk for thromboembolic
complications, such as patients undergoing cardiac surgery8
or plasmapheresis9 and in patients suffering from diabetes,10 heparin-induced thrombocytopenia,11
myocardial infarction,12 uremia,13 idiopathic
thrombocytopenic purpura,14 or thrombotic thrombocytopenic
purpura.15 Functional studies of these microparticles were
not performed. Other studies about the presence of microparticles of
nonplatelet origin in the circulation have not been
reported thus far. We have shown recently that elevated levels of
platelet- and erythrocyte-derived microparticles are present in wound
blood collected directly from the pericardial cavity in patients
undergoing cardiac surgery.16 These in vivo-generated microparticles strongly bind annexin V, a protein known for its interaction with negatively charged phospholipids such as
phosphatidylserine, one of the essential lipid cofactors for clotting.
Upon addition to normal plasma, these microparticles supported the
generation of thrombin by a tissue factor-factor VII-mediated pathway.
Hence, microparticles may be involved in activation of the systemic
coagulation in vivo.
In the present study, we investigated the presence, cellular
source, and function of circulating microparticles in patients suffering from meningococcal sepsis. Plasma samples of these patients were analyzed for the presence of microparticles (flow cytometry) and
their procoagulant activity (thrombin generation assay) and were
compared with the number and properties of microparticles in plasma
from 5 healthy volunteers. Our results show that patients with
meningococcal sepsis have elevated numbers of circulating microparticles derived from various blood cells and that these microparticles support clotting. We suggest that extreme DIC is strongly linked to circulating tissue factor-expressing microparticles in this disease and possibly also in other clinical conditions with
excessive coagulation activation.
| |
Materials and methods |
Reagents and assays
Reptilase was obtained from Boehringer Mannheim (Mannheim, Germany),
thrombin chromogenic substrate S2238 from Chromogenix AB
(Mölndal, Sweden), and normal mouse serum and fluorescein isothiocyanate (FITC)-labeled anti-CD4 (anti-CD4-FITC) from the Central Laboratory of the Netherlands Red Cross Blood Transfusion Service (CLB, Amsterdam, The Netherlands). Anti-glycophorin A-FITC and anti-CD61-FITC were obtained from Dakopatts (Glostrup, Denmark). Mouse IgG1-FITC, IgG2a-FITC,
IgG1-phycoerythrin (PE; all used as controls),
anti-CD14-PE, anti-CD8-FITC, and anti-CD20-FITC monoclonal antibody
(mAb) were from Becton Dickinson (San Jose, CA). Annexin V-FITC was
from Nexins Research B.V. (Hoeven, The Netherlands), anti-factor VII
and anti-factor XII were from CLB, and anti-E-selectin-FITC
(anti-CD62E-FITC) was from Serotec Ltd (Kidlington, UK).
Anti-CD14-FITC was from Biosource (Camarillo, CA), anti-CD66b-FITC
and IgG2b-PE (control mAb) were from Immuno Quality
Products (Groningen, The Netherlands), and anti-tissue factor-FITC
and polyclonal rabbit anti-human tissue factor were from American
Diagnostics (Greenwich, CT). Annexin V-PE was from PharMingen (San
Jose, CA). F1 + 2 was determined by enzyme-linked
immunosorbent assay (Enzygnost F1 + 2 micro) as described
by the manufacturer (Behring Diagnostics GmbH, Marburg, Germany).
Clinical studies
All patients included in the study had (1) a positive blood culture
for Neisseria meningitidis; (2) signs and symptoms of septic
shock; (3) a disease duration of less than 24 hours at study entry; and
(4) a characteristic rash (macular, petechial, purpuric, or
ecchymotic). Seven patients (age range, 1-29 years; male-female ratio,
2:5) were included. Patients had been included in an open, prospective
study on the effects of leukaplasmapheresis in patients
with meningococcal septic shock, and patient samples were collected
between 1989 and 1993.17 Leukaplasmapheresis, an effective
treatment that improves survival and reduces the chance of
complications,18 was applied after admission and was repeated 4, 10, 16, 24, and 36 hours after initial treatment. The
protocol was approved by the local hospital ethical committee. Informed
consent for the blood collection was obtained from the patients or
their relatives, as well as the attending physician.
Collection of blood samples
EDTA-anticoagulated blood was collected at admission and before each
leukaplasmapheresis procedure. Cells were removed by centrifugation for
15 minutes at 1550g at room temperature. Plasma samples were
stored in aliquots at  70°C until use. All plasma samples
from a single patient were tested in the same experiment to avoid
day-to-day variation of the flow cytometer between samples from 1 patient. Healthy volunteer samples, collected in the same period and
stored identically, were used for comparison.
Flow cytometric analysis
For flow cytometry, 250 µL of plasma was centrifuged for 15 minutes at 17 500g and 20°C to obtain microparticle
pellets. Subsequently, 225 µL of supernatant was removed, 225 µL of
apopbuffer (10 mmol/L HEPES, 5 mmol/L KCl, 1 mmol/L
MgCl2, and 136 mmol/L NaCl; pH 7.4) was added, and the
microparticles were recentrifuged. Finally, 225 µL of supernatant was
removed and the pellets were resuspended with 75 µL of apopbuffer.
From this suspension, 5-µL aliquots were diluted with 35 µL of
apopbuffer containing 2.5 mmol/L CaCl2 and 5 µL of
microparticle-free normal mouse serum (1:5000, v/v, final
concentration) and were incubated for 15 minutes at room temperature.
Subsequently, to identify the total microparticle population and their
cellular origin, we added 5 µL PE-labeled annexin V and 5 µL
FITC-labeled mAb, respectively, and incubated the samples for 15 minutes in the dark. To identify monocyte-derived, tissue
factor-expressing microparticles, we used anti-CD14-PE and
anti-tissue factor-FITC. The following (final) concentrations were
used: anti-CD4-FITC (0.5 µg/mL), anti-CD8-FITC (25 ng/mL), anti-CD20-FITC (0.5 µg/mL), IgG1-FITC (0.5 µg/mL),
IgG2a-FITC (0.5 µg/mL), IgG1-PE (0.5 µg/mL), anti-CD14-PE (0.25 µg/mL), anti-CD14-FITC (0.5 µg/mL),
anti-CD61-FITC (1 µg/mL), anti-CD62E-FITC (1 µg/mL), anti-glycophorin A-FITC (0.25 µg/mL), anti-CD66b-FITC (0.25 µg/mL), IgG2b-PE (0.5 µg/mL), anti-tissue factor-FITC
(1 µg/mL), and annexin V-PE (40 pg/mL). The incubation with mAb and
annexin V was terminated by the addition of 200 µL of apopbuffer
containing 2.5 mmol/L CaCl2, followed by recentrifugation.
After removal of 200 µL of supernatant, another 300 µL of
apopbuffer containing 2.5 mmol/L CaCl2 was added and the
pellets were resuspended. Samples were analyzed in a FACScan flow
cytometer with CellQuest software (Becton Dickinson, San Jose, CA).
Both forward scatter and sideward scatter were set at logarithmic gain.
Microparticles were identified on forward scatter, sideward scatter,
binding of annexin V, and binding of a cell-specific mAb. Annexin V
measurements were corrected for autofluorescence, and binding of
cell-specific mAbs was corrected with identical concentrations of
control IgG antibodies.16 The number of
microparticles per liter of plasma was calculated as: Number/L = N × [100/5] × [355/150] × [106/250].
Thrombin generation by microparticles
The thrombin generation assay as described by Kessels et
al19 was used to assess the in vitro thrombin-generating
capacity of microparticles. To prepare normal plasma, we collected
citrate-anticoagulated blood from 40 healthy volunteers who had not
taken any medication during the previous 10 days. Plasma was prepared
by centrifugation for 15 minutes at 1550g at room temperature.
The plasma samples were pooled and treated with reptilase (40 µL per
2 mL plasma) for 10 minutes at 37°C and then for 10 minutes on
melting ice. Subsequently, fibrin and microparticles were removed by
centrifugation for 1 hour at 17 500g (20°C), and plasma
was stored in 1-mL aliquots at 70°C until use.
Microparticles were prepared as described in flow cytometric analyses.
At t = 0, thrombin generation was started by the addition of 30 µL
CaCl2 (17 mmol/L) to 120 µL of the prewarmed (37°C)
normal plasma, to which 20 µL of buffer A (50 mmol/L Tris-HCl, 100 mmol/L NaCl; pH 7.35) and 10 µL of the washed microparticle
suspension had been added. At fixed intervals after t = 0, 3 µL portions were removed from this mixture and added to prewarmed
(37°C) buffer A containing 4 mmol/L of the chromogenic substrate
S2238 and 20 mmol/L EDTA (to block further thrombin generation). After
180 seconds, the conversion of S2238 was stopped by the addition of 90 µL citric acid (1.0 mol/L), and the generated p-Nitroaniline was
determined on a spectrophotometer at  = 405 nm. For inhibition
experiments, mAbs (anti-tissue factor, anti-factor VII, or
anti-factor XII; 07-1.0 mg/mL) were added to both plasma (20 µL) and
microparticles (10 µL), which were preincubated separately for 30 minutes at room temperature before the microparticles were added in the
thrombin generation assay.
Statistical methods
Data were analyzed with SPSS for Windows, release 8. Differences
were considered statistically significant at P < .05. For direct comparison of the number of microparticles in blood samples, the
Wilcoxon matched-pairs signed-rank test was used.
| |
Results |
Number and cellular origin of circulating microparticles in patients
with meningococcal sepsis
Microparticles were isolated from plasma samples, labeled, and
analyzed by flow cytometry as described in "Materials and
Methods." Figure 1 shows a
representative picture of microparticles stained with annexin V, which
binds to negatively charged phospholipids and can be used to stain
microparticles,16 and anti-CD66b, which labels
granulocytes. To correctly identify annexin V-positive microparticles,
we determined a threshold in a microparticle sample that was prepared
without any additions to correct for intrinsic autofluorescence (Figure
1A). This threshold is also depicted in panels B, E, and F. To identify
microparticles that bound cell-specific mAbs, we also incubated
microparticles with identical concentrations of control antibodies to
set a threshold. This threshold is shown in Figure 1C and is also used
in panels D, E, and F. Labeling with anti-CD66b revealed that part of
the microparticles originated from granulocytes (Figure 1D) and that
the microparticles also bound annexin V (Figure 1E, upper right).
Figure 1F shows the virtual absence of granulocyte-derived
microparticles in a representative dot plot of microparticles from a
healthy volunteer.
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| Fig 1.
Representative fluorescence-activated cell sorter (FACS)
dot plots of granulocyte-derived microparticles in plasma from a
patient who survived meningococcal sepsis.
Microparticles were isolated from the plasma of a (surviving) patient
on admission, labeled, and analyzed by flow cytometry as described in
"Materials and Methods." (A) Unlabeled microparticles
(autofluorescence); (B) labeled with annexin V-PE; (C) labeled with
IgG1-FITC (control mAb); (D) labeled with anti-CD66b-FITC;
and (E) double staining with annexin V-PE and anti-CD66b-FITC. (F)
Representative dot plot of microparticles isolated from healthy
volunteer plasma, double stained with annexin V-PE and
anti-CD66b-FITC, for comparison.
|
|
Table 1 summarizes the numbers of
circulating microparticles, identified by staining with annexin V-PE
and anti-CD mAb-FITC, in patients at study entry (n = 7) and healthy
volunteers (n = 5). For these experiments, a panel of mAbs was used
directed against TH cells (CD4), TS cells
(CD8), monocytes (CD14), B cells (CD20), platelets (CD61), endothelial
cells (CD62E), granulocytes (CD66b), and erythrocytes (glycophorin A).
Compared with healthy volunteers, the patients had significantly
increased numbers of circulating platelet (CD61)-derived and
granulocyte (CD66b)-derived microparticles at study entry. Monocyte
(CD14)-, B cell (CD20)-, and endothelial cell (CD62E)-derived
microparticles were also increased, although the difference was not
statistically significant. Notably, the nonsurviving patient A, who
died on admission after a disease course of less than 18 hours, had the
highest plasma levels of microparticles derived from TH
cells, monocytes, B cells, endothelial cells, platelets, granulocytes,
and erythrocytes. The other nonsurvivor showed no marked differences in
microparticle numbers when compared with patients who survived
meningococcal sepsis.
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Table 1.
Number of circulating microparticles on admission in
patients with meningococcal sepsis and healthy volunteers
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Time course of circulating microparticles
The time course of microparticle numbers in the patients is given in
Figure 2. In all 5 survivors, the number of
granulocyte-derived microparticles decreased during the first 10 hours
after admission (Figure 2, upper left). During the first 10 hours, the
number of monocyte-derived microparticles (CD14; Figure 2, middle left) increased slightly in 4 of the survivors, although this increase was not significant (P = .117 at 24 hours and
P = .462 at 36 hours). In contrast, microparticles originating
from platelets, erythrocytes, or endothelial cells (Figure 2;
upper right, lower right, and lower left, respectively) showed no
apparent changes during the observation period.
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| Fig 2.
Time course of circulating microparticles in patients who
survived meningococcal sepsis.
Microparticles were assessed and analyzed by flow cytometry as
described in "Materials and Methods." Figure shows the number of
microparticles double stained with annexin V-PE and the indicated
FITC-labeled mAb. The shaded area represents the range found in the 5 healthy volunteers. Note that the range of the y-axis varies and that
most of the microparticles were of platelet origin.
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Relation of circulating microparticles to coagulation in vivo
and in vitro
To investigate whether the microparticles detected in the patients
promoted coagulation, we first measured the activation of the
coagulation system in vivo by assessing the concentration of the
prothrombin fragment F1 + 2 in plasma during the course
of the disease, which reflects the concentration of thrombin formed in
vivo.20 Table 2 shows that the
concentration of F1 + 2 on admission was higher in the
patients than in volunteers (P = .004). This difference was
still present at 24 hours (P = .016), but not thereafter. The
concentration of F1 + 2 in survivors decreased slowly
after admission, becoming significantly lower than baseline values at 36 hours (P = .016), indicating progressively less activation of coagulation in vivo. To substantiate a link between coagulation activation and microparticles, we studied the thrombin-generating capacity of isolated microparticles from patients and healthy volunteers in vitro. Virtually all microparticle preparations of the
patients generated more thrombin than those of healthy controls (data
are summarized in Table 2).
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Table 2.
Thrombin generation in plasma of patients with
meningococcal sepsis in vivo and by microparticles in vitro
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Microparticles released from monocytes in vitro express tissue
factor,21,22 so we investigated whether such particles also circulate in vivo. Nonsurvivor A, who suffered from severe DIC as
evidenced by a low platelet number (7 × 109/L), a
prolonged prothrombin time (> 60 seconds; normal range, < 14.5
seconds), a decreased fibrinogen level (< 0.1 g/L; normal range,
1.7-3.7 g/L), the presence of fibrin degradation products, and a
prolonged activated partial thromboplastin time (> 120 seconds; normal range, < 36 seconds), had extremely high numbers of
monocyte-derived microparticles (CD14 positive). These microparticles
double stained for tissue factor (Figure
3A). In contrast, microparticles of nonsurvivor B, with a less fulminant disease course (platelet number
258 × 109/L, prothrombin time 17.1 seconds,
fibrinogen 2.5 g/L, presence of fibrinogen degradation products, and
activated partial thromboplastin time of 42.9 seconds), hardly stained
for CD14 or tissue factor (Figure 3B).
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| Fig 3.
Identification of tissue factor on circulating
microparticles of the nonsurviving patients.
Microparticles of nonsurviving patient A (A) and patient B (B) were
stained with CD14-PE and anti-tissue factor-FITC and analyzed by flow
cytometry.
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As indicated in Figure 4A, the addition of
microparticles from nonsurvivor A to normal plasma resulted in a strong
generation of thrombin, whereas those of nonsurvivor B induced only
modest thrombin generation. The generation of thrombin in normal plasma by the microparticles from nonsurvivor A was extremely delayed when
both microparticles and the normal plasma were preincubated with mAbs
against tissue factor or factor VII (Figure 4B), indicating that the
microparticle-associated tissue factor is active and stimulates the
extrinsic pathway of the coagulation system. In contrast, preincubation
with OT-2, a mAb that inhibits factor XIIa and blocks kaolin-induced
thrombin generation in normal plasma (data not shown), had no effect.
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| Fig 4.
Thrombin generation by microparticles of the nonsurviving
patients.
After reconstitution of microparticle-free, defibrinated normal plasma
with washed microparticles, thrombin generation was assessed as
described in "Materials and Methods." (A) Microparticles of
nonsurvivor A are indicated as  , and those of nonsurvivor B as  .
For comparison, thrombin generation is also shown for a representative
healthy volunteer ( ). (B) Microparticles of nonsurvivor A in the
absence of mAbs () or after preincubation with anti-tissue factor
( ), anti-factor VII ( ), or anti-factor XII ().
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| |
Discussion |
This study shows that patients with meningococcal sepsis have
elevated numbers of microparticles originating from various cell
populations in their circulation. These microparticles evoked a
stronger generation of thrombin in normal plasma than those from
volunteers, suggesting that the elevated numbers of microparticles in
the circulation or their cellular origin may be related to the
increased activation of the coagulation system in vivo. It could be
argued that the increased procoagulant activity of the patients'
microparticle fractions is mainly due to the increased number of
microparticles and not to their cellular origin or properties. Because
we wished to estimate the total procoagulant activity of the fractions,
the microparticles were not diluted to a standard concentration in the
thrombin generation assay. In addition, dilution cannot be done easily
because the cellular composition of microparticles varied among
patients, by the course of the disease, and most likely also by the
type of disease. We also demonstrated tissue factor on microparticles,
especially in a patient with an extremely fulminant course of DIC,
which was functional in the thrombin generation assay as established
with specific activity-blocking antibodies to tissue factor, factor
VII, and factor XII.
Tissue factor is a transmembrane protein, the extracellular domain of
which functions as a receptor for factor VII.23 Binding of
factor VII to tissue factor is a first step in a series of events in
which soluble coagulation proteins become assembled on a phospholipid
surface. Evidence that tissue factor is important for coagulation and
inflammation in vivo comes from a number of animal studies. Infusion of
recombinant activated factor VII into normal chimpanzees raised the
plasma levels of activation peptides of factor IX, factor X, and
prothrombin. This was blocked by the administration of an anti-tissue
factor mAb.24 Infusion of endotoxin reduced the number of
platelets in rabbits and decreased the concentrations of
fibrinogen, antithrombin, and factor VIII, whereas it prolonged the
activated partial thromboplastin time. These changes were counteracted
by tissue factor pathway inhibitor.25 Infusion of
Escherichia coli into baboons caused sepsis with severe DIC, which could be prevented by concurrent infusion of tissue factor pathway inhibitor.26,27 Although these studies clearly
indicate the importance of tissue factor for the development of DIC and sepsis, the location of functionally active tissue factor expression is
not well known. Our results suggest that tissue factor exposed by
microparticles, particularly those released by monocytes, may be
important in this respect. However, another component required for the
assemblage of the coagulation factor complexes on the phospholipid
surface is phosphatidylserine, which is not exposed on normal cells but
on, for example, activated platelets. In the present study,
microparticles stained positive for annexin V, indicating the presence
of phosphatidylserine on their surfaces.28 Presumably, this
also explains in part the thrombin-generating capacity of circulating
microparticles. The relative contributions of phosphatidylserine and
tissue factor to coagulation activation remain to be established.
Possibly, the presence of tissue factor may enhance the coagulation
activation associated with phosphatidylserine.
Monocytes are the only cells found in peripheral blood currently known
to be capable of expressing tissue factor.23 Isolated monocytes stimulated by endotoxin express tissue factor.29
Under flow conditions, endotoxin-stimulated monocytes stimulate fibrin deposition and thrombus formation. Anti-tissue factor mAb
inhibits both of these processes.30 Monocytes have
been shown to express tissue factor in patients suffering from invasive
tumors, leukemia, sepsis, myocardial infarction, and diabetes, and in
patients requiring extracorporeal circulation.31 In
addition to expressing tissue factor, monocytes can release tissue
factor-exposing microvesicles in vitro upon stimulation with
endotoxin.21,22 Mallat et al32 recently
reported the presence of membrane vesicles of monocytic and lymphocytic
origin that retained tissue factor activity in atherosclerotic plaques.
Our present results extend these findings and demonstrate, for the
first time, that procoagulant microparticles of monocyte, granulocyte,
and endothelial cell origin can be detected in the circulation. It is
interesting that of the 7 patients studied, only the patient with
severe DIC had an extremely elevated (7-fold increase) number of
endothelial cell-derived microparticles
(244 × 106/L) compared with controls. Although we
cannot exclude the possibility that these microparticles also express
tissue factor, 85% of the tissue factor-positive microparticles were
CD14 positive. We therefore presume that the contribution of tissue
factor to the overall procoagulant activity is especially due to the
increased number of monocyte-derived microparticles.
The samples analyzed in the present study had been collected between
1989 and 1993. This may raise concerns about the validity of the
present findings (i.e., the microparticle profile). Therefore, microparticles were also isolated immediately after blood collection from 2 patients with sepsis and multiple organ failure and 2 healthy controls. Both patients clearly showed granulocyte-derived
microparticles, which were absent or present at low numbers in the
controls. The numbers of monocyte- and endothelial cell-derived
microparticles were not increased in these patients compared with the
controls (data not shown). On the basis of these preliminary data, we
cannot make definitive conclusions about the presence or absence of
either monocyte- or endothelial cell-derived microparticles in other diseases, but we hypothesize that such microparticles may be especially prevalent in patients with severe DIC. Combes et al33
recently showed the presence of endothelial cell-derived
microparticles in blood from healthy individuals and their increased
presence in blood obtained from patients with lupus anticoagulant.
Thus, the presence of endothelial cell-derived microparticles is
evidently not unique for patients with meningococcal sepsis. Combes et
al,33 however, did not report the presence of tissue factor
on these microparticles.
All patients studied had elevated numbers of microparticles. It is
tempting to speculate that interference with the release of
microparticles may be a target for therapeutic intervention. Recently,
an mAb against the glycoprotein IIb-IIIa complex on platelets, which
inhibits platelet-platelet interaction or aggregation, has been used
successfully in patients undergoing stent
implantation.34,35 In vitro, this mAb prevents the release
of microparticles from platelets.36 Our findings imply that
this mAb may inhibit clotting in vivo as well. Indeed, infusion of this
mAb into baboons with lethal E. coli sepsis prevented fibrin
deposition and renal insufficiency.37 Thus, therapeutic
interference with microparticle release in general, and possibly of
platelets in particular, seems a realistic option.
In conclusion, elevated levels of microparticles were observed in
patients with meningococcal sepsis. These microparticles enhanced
coagulation by providing a suitable phospholipid surface and, at least
in part, by exposing tissue factor. We suggest that such microparticles
are involved in the pathogenesis of DIC during meningococcal sepsis and
may constitute a novel target for therapeutic intervention in this
disease, and possibly also in other clinical conditions with enhanced
coagulation activation.
| |
Footnotes |
Submitted April 6, 1999; accepted September 24, 1999.
Reprints: Rienk Nieuwland, Department of Clinical Chemistry,
Leiden University Medical Center, P.O. Box 9600, 2300 RC Leiden, The
Netherlands; e-mail: rnieuwland{at}ckcl.azl.nl.
The publication costs of this
article were defrayed in part by
page charge payment. Therefore,
and solely to indicate this fact,
this article is hereby marked
"advertisement"
in accordance with 18 U.S.C.
section 1734.
| |
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Abstract
Introduction