RED CELLS
Sickle blood contains tissue factor–positive microparticles derived
from endothelial cells and monocytes
Arun S. Shet, Omer Aras, Kalpna Gupta, Mathew J. Hass, Douglas J. Rausch, Nabil Saba, Louann Koopmeiners,
Nigel S. Key, and Robert P. Hebbel
Blood microparticles (MPs) in sickle cell
disease (SCD) are reportedly derived only
from erythrocytes and platelets. Yet in
SCD, endothelial cells and monocytes are
activated and abnormally express tissue
factor (TF). Thus, sickle blood might contain TF-positive MPs derived from these
cells. With the use of flow cytometry to
enumerate and characterize MPs, we
found total MPs to be elevated in crisis
(P ⴝ .0001) and steady state (P ⴝ .02) in
subjects with sickle cell disease versus
control subjects. These MPs were derived
from erythrocytes, platelets, monocytes,
and endothelial cells. Erythrocytederived MPs were elevated in sickle crisis
(P ⴝ .0001) and steady state (P ⴝ .02) versus control subjects, as were monocytederived MPs (P ⴝ .0004 and P ⴝ .009, respectively). Endothelial and plateletderived MPs were elevated in sickle crisis
versus control subjects. Total TF-positive
MPs were elevated in sickle crisis versus
steady state (P ⴝ .004) and control subjects (P < .0001) and were derived from
both monocytes and endothelial cells.
Sickle MPs shortened plasma-clotting
time compared with control MPs, and a
TF antibody partially inhibited this procoagulant activity. Markers of coagulation
were elevated in patients with sickle cell
disease versus control subjects and correlated with total MPs and TF-positive
MPs (P < .01 for both). These data support the concept that SCD is an inflammatory state with monocyte and endothelial
activation and abnormal TF activity.
(Blood. 2003;102:2678-2683)
© 2003 by The American Society of Hematology
Introduction
Microparticles (MPs) are small, membrane-derived vesicles released by cells on activation or during apoptosis.1,2 Because the
process of MP formation entails a loss of normal membrane lipid
asymmetry, in which aminophospholipids are confined to the inner
leaflet of the cell membrane, MPs abnormally display phosphatidylserine (PS) on their outer leaflets.3 MPs of various cellular origins
are present in blood and typically bear cell membrane antigens that
reflect their cellular origin.4 MPs from platelets5 and red cells6 have
been described in sickle cell disease. Because endothelial cells and
monocytes are activated in sickle disease,7-9 we hypothesized that
MPs would also arise from these cells.
In addition to being an inflammatory condition,10 sickle cell
disease is also believed to be a prothrombotic state.11 Activation of
the coagulation system is evidenced by platelet activation, thrombin generation, and fibrinolysis in patients with sickle cell disease11-16
and may play a role in thrombotic complications of sickle cell
disease such as stroke.17 Consistent with these findings, in sickle
cell disease tissue factor (TF) is abnormally expressed on circulating endothelial cells (CECs),8 and whole blood TF procoagulant
activity, believed to be associated with monocytes, is elevated.18
Studies in healthy individuals have demonstrated a blood-derived
thrombogenic pool of TF that is not cell associated but apparently
MP associated.19 The origin of this circulating MP-associated TF is
not clearly defined; however, evidence cited earlier suggests that it
could be derived from blood cells (monocytes) or the vessel wall
(endothelial cells). Therefore, we hypothesized that endothelial
cell– and monocyte-derived MPs in the blood of patients with
sickle cell disease would also be TF positive.
From the Vascular Biology Center and Division of Hematology, Oncology and
Transplantation, Department of Medicine, University of Minnesota Medical
School, Minneapolis; and the Section of Hematology and Oncology, Hennepin
County Medical Center, Minneapolis, MN.
Association of America, and the Lillihei Heart Institute, Minneapolis, MN.
Submitted March 5, 2003; accepted June 2, 2003. Prepublished online as
Blood First Edition Paper, June 12, 2003; DOI 10.1182/blood-2003-03-0693.
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.
Supported by grants PO1 HL55552 (R.P.H.) and R29 HL55219 (N.S.K.) from
the National Institutes of Health. A.S.S. is a scholar of the Sickle Cell Disease
2678
Patients, materials, and methods
Reagents and assays
We obtained cell-specific monoclonal antibodies (MoAbs) against CD14
for monocytes (CRIS-6; Biosource International, Camarillo, CA), and
against CD144 for endothelial cells (55-7H1), CD41a for platelets (HIP8),
and glycophorin A for red blood cells (RBCs; JC159) (BD-Pharmingen, San
Diego, CA). Fluorescein isothiocyanate (FITC)–labeled MoAb against TF
(CJ4068) was obtained from American Diagnostica (Greenwich, CO), and
isotype control mouse immunoglobulin G1 (IgG1) labeled with either FITC
or phycoerythrin (PE; 11711.11) was obtained from R&D Systems (Minneapolis, MN). Cy5-labeled Annexin V was obtained from BD-Pharmingen.
Fluorescent microbeads of different sizes (0.3-2.0 ␮m) for gating experiments were obtained from Molecular Probes (Eugene, OR) and Sigma
Chemical (St Louis, MO), and 7.2-␮m nonfluorescent beads for enumeration of MPs were obtained from Bangs Laboratories (Fishers, IN).
The cell specificity of the MoAbs used in our MP flow cytometric
experiments was verified in preliminary experiments using endothelial
cells, monocytes, platelets, and RBCs (data not shown). The antibodies
were also titrated in preliminary experiments to determine the optimal
labeling concentrations for each (range, 0.1-1.25 ␮g/mL).
Reprints: Arun S. Shet, Laboratory of Blood and Vascular Biology, Rockefeller
University, 1230 York Ave, New York, NY 10021; e-mail: [email protected].
© 2003 by The American Society of Hematology
BLOOD, 1 OCTOBER 2003 䡠 VOLUME 102, NUMBER 7
BLOOD, 1 OCTOBER 2003 䡠 VOLUME 102, NUMBER 7
For plasma clotting experiments, we used coagulation control plasmalevel-1 plasma from Fischer Diagnostics (Middletown, VA), Innovin
(recombinant re-lipidated human-TF) from Dade International (Miami,
FL), and a polyclonal rabbit antihuman TF-blocking antibody with preimmune IgG control (courtesy of Dr Ron Bach, University of Minnesota).
Study subjects
Twenty-seven adult patients with a diagnosis of sickle cell disease (23 with
HbSS or HbS␤thalasemia; 4 with HbSC) provided 21 samples in acute
crisis and 16 samples in steady state. These included samples from 12
patients who were studied in both crisis and steady state. The subjects
included 15 men and 12 women, with a mean age of 26 years (range, 19-45
years). All but one were receiving hydroxyurea. Two patients were started
on warfarin (Coumadin) therapy for catheter-related thrombi between
attainment of the crisis sample and the follow-up steady-state sample. None
of the patients studied were routinely on aspirin therapy, and some of the
patients in crisis were febrile.
A “crisis” was defined as a visit to a medical facility for pain that had no
evident cause other than sickle cell anemia and for which the patient was
hospitalized and treated with parenteral narcotics. Samples obtained from
the time of onset of a crisis and throughout hospitalization were considered
to be “crisis” samples. We defined “steady state” as any time exclusive of
the 4 weeks prior to or after a crisis period. Healthy control subjects
(n ⫽ 13) were all race matched and were similar in age (mean ⫽ 30 years;
range, 22-42 years) and sex (M ⫽ 6, F ⫽ 7) to the patient population. In
addition, we studied 3 individuals with sickle trait (HbAS) and 2 individuals
with persistent hemolytic anemia (autoimmune hemolytic anemia and
pyruvate kinase deficiency) who had previously undergone surgical splenectomy. All study participants signed informed consent forms, and the
Institutional Review Boards of the University of Minnesota and Hennepin
County Medical Center approved the protocol.
Samples
Blood samples were drawn using a 21-gauge needle into buffered citrate
(Becton Dickinson, San Jose, CA) after discarding the first 3 mL.
Platelet-free plasma (PFP) was prepared immediately after blood collection
using a 2-step centrifugation procedure: initially at 1500g for 10 minutes at
20°C to make platelet-rich plasma and then at 13 000g for 10 minutes at
20°C to make PFP. Samples were used immediately or stored at ⫺80°C for
future flow cytometric or enzyme-linked immunosorbent assay (ELISA)
experiments. Preliminary experiments showed that subsequent results were
not affected by a freeze-thaw cycle.
Preparation of microparticles
MPs were isolated from PFP using a 2-step sequential ultracentrifugation
process. Briefly, 250 ␮L PFP was washed in 10 mL wash buffer (HEPES
(N-2-hydroxyethylpiperazine-N⬘-2-ethanesulfonic acid) 10 mM, NaCl 140
mM, KCl 4.5 mM, bovine serum albumin [BSA] 1%, Na azide 0.1%, CaCl2
2.5 mM, pH 7.4; 0.2 ␮m filtered) and ultracentrifuged at 100 000g for 60
minutes 2 times (total wash ratio 1:400). After careful aspiration of all but
1 mL supernatant, pelleted MPs were resuspended in the remainder wash
buffer by gentle vortexing and pipetting and used immediately for flow
cytometric experiments.
Figure 1. Flow cytometric analysis and quantification
of MPs. (A) Determination of forward (FSC) and sidescatter (SSC) characteristics of light by 1.0-␮m latex
beads in buffer to establish the R-1 or MP gate. The R-2
or bead gate includes 7.2-␮m latex beads used for
enumerating MPs. (B) Detection of phosphatidylserine
(PS)–positive MPs in platelet-free plasma (PFP) by
annexin V–Cy5 labeling on the y-axis in relation to SSC
on the x-axis. (C) Representative sample of MPs in PFP
from a sickle sample triple labeled for annexin V (not
shown), anti–TF-FITC on the x-axis, and anti–VE cadherin-PE on the y-axis.
TISSUE FACTOR⫹ MICROPARTICLES IN SICKLE BLOOD
2679
Flow cytometric detection of microparticles
All reagents and buffers were sterile and filtered using a 0.2-␮m filter (Pall,
Ann Arbor, MI). MP aliquots (90 ␮L) were incubated for 30 minutes in the
dark with either specific antibodies (in concentrations found to be optimal
in preliminary experiments) or an equal concentration of isotype control,
always done in parallel. Samples were washed by centrifugation in 1 mL
wash buffer at 32 000g (Hettich Mikro 22R, Tuttelingen, Germany) for 60
minutes at 20°C. After careful removal of all but 200 ␮L supernatant,
pelleted MPs were resuspended in the remainder wash buffer, and 3 ␮L
annexin V–Cy5 was added. After incubation for 15 minutes in the dark at
room temperature, enumeration beads were added, and the samples were
ready for fluorescence-activated cell sorting (FACS) study.
Sample data were acquired using a FACS Calibur flow cytometer with
CellQuest pro software (Becton Dickinson). Forward (FSC) and side scatter
of light was set in log scale, and threshold was set at the FSC parameter. MP
gating was accomplished by preliminary standardization experiments using
fluorescent microbeads of different sizes. First, MPs were simulated using
varying sizes of standard microbeads (0.3-2.0 ␮m), and a microparticle gate
(MP gate) was determined using these standards. The MP gate included
1.0-␮m beads in its upper and outer corner so that it would contain all
microparticles 1.0 ␮m or less (Figure 1A). Events in the MP gate were
further assessed for labeling with annexin V–Cy5 to distinguish true events
from electronic noise and thereby increase the specificity of MP detection
(Figure 1B).
For MP quantification we used a modification of the methods of
Combes et al1 in which a known quantity of 7.2-␮m beads (250 000) was
added to each sample, and acquisition was stopped after 10 000 beads were
counted in the R-2 bead gate. The diameter of these beads allowed
discrimination from the MP population on light scatter and could be
counted using a separate gate (bead-gate; Figure 1A). Total MP number
(N in MP per milliliter) was calculated using the following formula:
n ⫽ C ⫻ Beadsadded/Beadscounted ⫻ plasma volume (mL) ⫻ 12, where C
represents the number of positive events with the background signal
subtracted and 12 is a dilution factor. For MP quantification, standard
deviations for interassay and intra-assay variability were 10% and 4%,
respectively.
Data analysis was performed using Flojo software. During data
analysis, a “positive event” was defined as one that exhibited greater
fluorescent intensity than the isotype control (IgG1 FITC or PE) or EDTA
(ethylenediaminetetraacetic acid) buffer in the case of annexin V. Background signal from either isotype control or EDTA buffer accounted for 3%
to 5% of the total signal in a typical experiment. Events falling in the R-1
MP gate (based on size) that also were annexin V–positive were defined as
MPs, and the background signal (false positive) obtained using buffer
containing only annexin V–Cy5 was subtracted. Annexin V–positive events
were then further examined for fluorescent labeling with a cell-specific
antibody and a TF antibody (Figure 1C).
To investigate the cellular origin of TF on MPs, all samples were triple
labeled with Cy5-labeled annexin V, a cell type–specific PE-labeled MoAb
against either CD14 or CD144 and a FITC-labeled MoAb against TF
(Figure 1C). Although we did not expect platelet- or RBC-derived MPs to
express TF, MPs from 6 patients were also triple-labeled with Cy5-labeled
annexin V, PE-labeled anti-CD41a, or glycophorin A, and FITClabeled anti-TF.
2680
BLOOD, 1 OCTOBER 2003 䡠 VOLUME 102, NUMBER 7
SHET et al
Figure 2. Total blood MPs and cellular origin of MPs. Data are expressed as MP number per milliliter of platelet-free plasma and shown as mean ⫾ SEM, and statistical
analysis is presented in Table 1. In each panel, the clear bars (䡺) show data for healthy state, gray bars (u) show data for steady state, and black bars (f) show data for crisis.
(A) Total MP number. (B) Endothelial-derived MP number. (C) MP number derived from red blood cells, platelets, and monocytes. Error bars represent SEM.
Procoagulant activity of MPs
Total MPs are elevated among sickle patients
MP aliquots isolated from plasma of study subjects were added as a
potential source of TF, to normal citrated plasma. After recalcification,
plasma clotting time was determined using an automated prothrombin time
analyzer (Diagnostica Stago-ST4, Tokyo, Japan). In parallel, the clotting
time of samples preincubated with a polyclonal blocking anti-TF antibody
(9 ␮g/mL final concentration for 60 minutes at room temperature) was also
measured. The difference between the 2 clotting times represents inhibition
of TF-dependent activity. Clotting time was stopped at 1000 seconds if the
samples had not clotted.
Enumeration revealed race-matched healthy subjects to have total
MPs of 192 ⫾ 70 ⫻ 103/mL PFP (mean ⫾ SEM). In comparison,
total MPs were significantly elevated for both patients with sickle
cell disease in steady state (950 ⫾ 346 ⫻ 103/mL; P ⫽ .02 versus
healthy subjects) and in crisis (2592 ⫾ 703 ⫻ 103/mL; P ⫽ .0001
versus healthy subjects) (Table 1; Figure 2A). For total MP number, both
the overall group data of steady state versus crisis donors and the paired
data of patients sampled during both steady state and crisis showed a
significant difference (P ⫽ .02 and P ⫽ .003; Tables 1-2). There was no
difference in total MP number linked to age or sex (data not shown). We
measured total MPs in 2 individuals who had undergone splenectomy
with hemolytic anemia and found numbers similar to those of healthy
subjects (338 ⫻ 103/mL and 110 ⫻ 103/mL), suggesting that absent
splenic function per se does not markedly influence MP number.
Similarly, samples from 3 individuals with sickle trait were not
significantly different from the race-matched healthy subjects (207 ⫾
195 ⫻ 103/mL).
ELISA assay for D-dimer, TAT, and PF1.2
Markers of coagulation activation, prothrombin fragment F1.2 (PF1.2),
thrombin antithrombin complex (TAT; Dade Behring, Marburg, Germany),
D-dimer (Dimertest Gold EIA; American Diagnostica) were measured by
ELISA. Upper limits of normal for D-dimer, TAT, and PF1.2 among healthy
subjects determined by the manufacturer were 120 ng/mL, 4 ␮g/L, and
1.1 nM, respectively.
Statistical analysis
Because the data obtained had a non-Gaussian distribution, statistical
differences between groups were evaluated using nonparametric methods.
The Kruskal-Wallis test was used for an overall comparison between the
groups. Significance determined by the Kruskal-Wallis test enabled subsequent intergroup comparisons using the Mann-Whitney U test; P ⬍ .05
(2-tailed) was considered significant. Bivariate correlations were done
using the Spearman rho test. Paired sample analysis was done using the
Wilcoxon signed log-rank test. Statistical analysis was performed using
SPSS version 10.0 for Windows (Chicago, IL).
Cell origin of MPs
Using specific markers, we confirmed findings of others5,6 that
there are blood MPs in patients with sickle cell disease derived
from red blood cells (glycophorin A⫹) and platelets (CD41a⫹).
However, we also found MPs derived from endothelial cells
(VE-cadherin⫹) and monocytes (CD14⫹) (Figure 2B-C). Red
blood cell– and monocyte-derived MPs were significantly increased in both steady state and crisis compared with healthy
subjects. Both red blood cell– and monocyte-derived MPs were
Results
We used a rigorous definition for plasma MPs because we found
methods in the literature to be insufficient for our goals. For
instance, some reports have simply defined MPs as annexin
V–positive events in plasma by flow cytometry, but we find that
plasma also contains annexin V binding material that cannot be
sedimented by ultracentrifugation (A.S.S., R.P.H., unpublished
data, August 10, 2001). Additionally, some groups have defined
MPs as events by flow cytometry that are 2 ␮m or less, but we
found this size was large enough to erroneously include platelets
(A.S.S., R.P.H., unpublished data, August 10, 2001). Therefore, we
defined MPs as vesicles removable from plasma by ultracentrifugation that are 1 ␮m or less in size and that bind annexin V. Our data
for MP number, cellular origin, and tissue factor expression are
shown in Figures 2-3, and the statistical comparisons between the
different subject groups are shown in Table 1 (group data) and
Table 2 (paired data).
Figure 3. Tissue factor–positive MP. Data are expressed as MP number per
milliliter of platelet-free plasma and shown as mean ⫾ SEM, and statistical analysis is
shown in Table 1. Total TF-positive microparticles (f) are elevated among sickle
patients in steady state and during crisis and are derived from monocytes (u) and
endothelial cells (䡺). Error bars represent SEM.
BLOOD, 1 OCTOBER 2003 䡠 VOLUME 102, NUMBER 7
TISSUE FACTOR⫹ MICROPARTICLES IN SICKLE BLOOD
Table 1. Statistical comparisons of group data among patients with
sickle cell disease (crisis, n ⴝ 21; steady state, n ⴝ 16)
and race-matched control subjects (n ⴝ 13)
2681
crisis (8 of 21) had TF-positive MPs that were derived from
endothelial cells, so 13 of 21 samples had no detectable endothelialderived TF-positive MPs.
P*
Crisis vs
steady state
Steady state vs
healthy state
Crisis vs
healthy state
Total MPs
.02
.02
.0001
Red cell MPs
.15
.02
.0001
Monocyte MPs
.12
.009
.0004
Total TF⫹ MPs
.004
.002
⬍ .0001
⬍ .0001
Monocyte TF⫹ MPs
.009
.004
Endothelial TF⫹ MPs
.02
.36
.01
D-dimer
.13
.001
.0002
TAT
.08
.0002
.0001
PF1.2
.48
.008
.003
*P values determined by Mann-Whitney U test.
also elevated in sickle crisis compared with steady state (Figure 2),
but this achieved significance only in the paired data analysis
(Table 2). Red blood cell–derived MP number correlated inversely
with hemoglobin (r ⫽ ⫺0.452; P ⫽ .003), and monocyte-derived
MP number correlated positively with absolute monocyte count
(r ⫽ 0.442; P ⫽ .001).
Endothelial cell– and platelet-derived MPs were each similar in
healthy subjects and patients with sickle cell disease in steady state,
but each was elevated during crisis (Figure 2). However, these
differences were not statistically significant (Table 1). For endothelial cell–derived MPs this clearly was due to variability among
patients, as 10 of 21 sickle samples in crisis had no detectable
endothelial MPs. For platelet-derived MPs, the difference approached statistical significance both among crisis versus healthy
subjects and crisis versus steady state in the paired data.
Expression of tissue factor on endothelialand monocyte-derived MPs
Testing MPs for TF, we found that a portion were TF positive
(Figure 3). Patients in crisis had significantly higher total TFpositive MPs compared with those in steady state or healthy
subjects, and subjects in steady state had significantly higher
numbers than healthy subjects (Table 1). Group data revealed a
significant difference between steady state and crisis for both
monocyte- and endothelial-derived tissue factor–positive MPs
(Table 1). TF-positive MPs were derived from endothelial cells and
monocytes (Figure 3) but not from red blood cells or platelets. Most
of the sickle samples in crisis (20 of 21) had TF-positive MPs that
were derived from monocytes. A smaller number of samples in
MPs without using annexin V binding criterion
To assess both PS-positive and PS-negative MPs, it is possible to
relax the criteria for MP definition by eliminating the need for
annexin V binding. In doing so, we thus defined MPs as small
(ⱕ 1.0 ␮m) particles removable by ultracentrifugation that were
labeled with cell-specific markers. Total MP number enumerated
by this method for healthy subjects was 218 ⫾ 98 (mean ⫾ SEM),
and for patients with sickle cell disease was 1312 ⫾ 424 in steady
state and 1844 ⫾ 542 in crisis. This discrepancy indicates that there
are a number of PS-positive MPs in crisis that do not label with the
cell-specific markers that we used. For the MPs of specific cell
origin, monocyte- and endothelial-derived MP number was increased by 3-fold and 4-fold, respectively, by using this definition
compared with the previous data, which additionally requires
annexin V binding. However, the RBC- and platelet-derived MP
number did not change. Inclusion of PS-negative MPs, however,
did not change the number of patients with sickle cell disease
demonstrating endothelial MPs. The percentage of monocytederived MPs that were TF⫹ and PS⫺ was 23% in steady state and
25% in crisis. The percentage of endothelial-derived MPs that were
TF⫹ and PS⫺ was 7% in steady state and 32% in crisis.
Procoagulant activity of sickle MPs
The MP preparation from individual patients with sickle cell
disease significantly shortened the plasma clotting time, compared
with the MP preparation from healthy control subjects (Figure 4;
P ⫽ .001). This procoagulant activity of sickle MPs was partially
inhibited by a blocking anti-TF antibody (Figure 4; P ⫽ .002),
providing evidence that sickle MPs possessed TF-dependent procoagulant activity. The anti-TF antibody blocked approximately 55%
of the total procoagulant activity, suggesting that it did not
completely inhibit TF in this assay. Of note, MPs from healthy
subjects were unable to shorten the clotting time in this assay,
probably because the normal MP preparation included fewer total
MPs (Figure 2) and fewer TF-positive MPs (Figure 3).
Markers of coagulation activation are elevated and correlate
with MP numbers
We measured plasma D-dimer (a marker of fibrinolysis), as well as
TAT and PF1.2 (markers of thrombin generation) in our study
Table 2. Statistical comparison of paired data for patients with
sickle cell disease sampled both during crisis
and steady state (n ⴝ 12)
P*
Total MPs
.002
Red blood cell MPs
.04
Monocyte MPs
.01
Total TF⫹ MPs
.007
Monocyte TF⫹ MPs
.007
Endothelial TF⫹ MPs
.06
D-Dimer
.01
TAT
.04
PF1.2
.06
*P values are calculated from steady state versus crisis and determined by
Wilcoxon signed rank test.
Figure 4. Effect of MPs on plasma clotting time. MP aliquots from patients with sickle
cell disease could shorten plasma clotting time, whereas similar MP aliquots from healthy
subjects could not (P ⫽ .001). Parallel samples (f) show that MP shortening of plasma
clotting time could be partially inhibited by a TF antibody, demonstrating TF-dependent
procoagulant activity (P ⫽ .002). Error bars represent SEM.
2682
BLOOD, 1 OCTOBER 2003 䡠 VOLUME 102, NUMBER 7
SHET et al
50% ⫾ 32% of PFP hemoglobin can be removed by ultracentrifugation and is, therefore, most likely MP associated (A.S.S. and
R.P.H., unpublished data, February 10, 2003).
Monocyte-derived MPs were significantly elevated during
steady state and crisis compared with healthy subjects, suggesting
that sickle monocytes are activated at all times in sickle disease.
Although endothelial-derived MPs were elevated during crisis
compared with healthy state, this difference was not significant
statistically from healthy subjects. This is due to intergroup
variability because only a subgroup of the patients with sickle cell
disease (11 of 21) had demonstrable endothelial-derived MPs.
Platelet-derived MPs were elevated but not significantly. This
elevation is most likely due to our careful avoidance of platelet
contamination, our stringent size definition (ⱕ 1.0 ␮m, which is
much less likely to include platelets than the size previously used
ⱕ 2.0 ␮m5), and our use of a platelet-specific antibody (CD41)
compared with previously used antibody5 (CD61) that reacts
against epitopes shared by platelets and endothelial cells.
Overall, these data indicate that patients with sickle cell disease
have elevated MPs both in steady state and crisis, implying that
even patients in steady state are fundamentally different from
healthy subjects. In general, stimuli causing cellular activation or
injury result in membrane perturbation and vesiculation, ie, release
of MPs. Examples of these are agents such as tumor necrosis factor
␣ (TNF␣) and endotoxin that cause vesiculation from endothelial
cells and monocytes, respectively.1,25 In vivo, vesiculation occurs
pathologically in conditions characterized by endothelial damage,
such as thrombotic thrombocytopenic purpura, sepsis, and disseminated intravascular coagulation.4,21 Thus, the findings of endothelial- and monocyte-derived MPs in sickle blood imply endothelial
and monocyte activation in this disease, or possibly apoptosis.
Notably, sickle endothelial cells probably manifest an activated
phenotype, with abnormal expression of adhesion molecules.7
Similarly, monocytes are in an activated state, demonstrating
up-regulation of CD11b and increased intracellular cytokines.9
There is mounting evidence that sickle cell anemia is an inflammatory condition10 associated with endothelial perturbation.26 The
present data support this concept and suggest that there is a chronic,
on-going activation of blood cells and the vascular wall.
The expression of TF by blood MPs is of great potential
relevance to disease pathobiology. TF is the principal initiator of
coagulation,27 and its increased in vivo expression promotes
thrombotic events in patients with a variety of clinical disorders.28,29 In sickle cell disease, there is abnormal expression of
tissue factor by monocytes18 and perhaps endothelium8 and an
association with thrombotic manifestations such as stroke17 and
pulmonary thrombosis.30 TF-positive MPs could potentially contribute to thrombotic manifestations by activating the clotting system
and generating thrombin. We found that TF-positive MPs were
more elevated in crisis than in steady state, but even patients in
steady state demonstrated significant differences from healthy
control subjects, suggesting that hemostatic perturbation is a
constant feature of sickle cell disease.
subjects. Each marker of coagulation activation was significantly
elevated among patients with sickle cell disease in both steady state
and crisis compared with healthy subjects (Table 1). The paired
data indicated significant differences between crisis and steady
state for D-dimer and TAT (Table 2). All 3 markers correlated
modestly but significantly with total MPs, total TF-positive MPs,
monocyte-derived TF-positive MPs, and with RBC-derived MPs
(Table 3). For endothelial-derived TF-positive MPs, correlations
did not achieve significance (not shown), again because only a
subgroup of patients had any TF-positive endothelial MPs.
Discussion
Although blood MPs were first described more than a decade ago,20
there is no consensus as to the definition, pathophysiologic role,
and fate of blood-derived MPs. The published literature describes
various methods for isolating and defining MPs using flow
cytometry.4,5,21 We used a very rigorous definition for MPs:
vesicular material that is removable from PFP by ultracentrifugation, with a size of 1.0 ␮m or less, and surface PS positive as
evidenced by annexin V binding. Imposition of these criteria
probably sacrifices sensitivity to some degree but probably preserves specificity for true MPs. Although the demonstration of
surface PS by annexin V binding adds a level of rigor to the
methodology, this level may be unnecessary if markers of cell
surface antigens are used instead. The present data indicate that,
although all RBC- and platelet-derived MPs are PS positive, this is
not true of endothelium and monocyte-derived MPs. Indeed, a
substantial portion of endothelial- and monocyte-derived MPs are
PS negative. The biologic relevance of this finding is currently
uncertain, but clearly PS-positive MPs have more relevance in the
setting of coagulation.
The findings in this study led to 5 key observations. (1) Total
blood MPs are elevated in sickle crisis compared with steady state.
(2) Sickle blood contains MPs derived from endothelial cells and
monocytes. (3) A portion of the MPs derived from these 2 cell types
is TF positive. (4) MPs in sickle cell disease have procoagulant
activity, some of which is tissue factor dependent. And (5), in
general, MP and TF-positive MP values for patients with sickle cell
disease in steady state were intermediate between those of healthy
and crisis subjects. However, the subjects in steady state were not
always significantly different from those in crisis.
RBC-derived MPs almost always constitute the majority of
blood MPs in patients with sickle cell disease but not among
control subjects in which platelet-derived MPs predominate. Accounting for this observation are the findings by Allan et al22 that
sickle erythrocytes release vesicles as a result of repetitive sickling
and unsickling during hypoxia and reoxygenation. RBC-derived
MPs are elevated both during crisis and steady state, supporting
these findings. They are also more elevated during crisis compared
with steady state. Therefore, the presence of these circulating
vesicles might explain previous reports of elevated plasma hemoglobin during sickle crisis.23,24 Indeed, we have observed that
Table 3. Correlation coefficients (Spearman rho) for all study samples
Total MPs
RBC MPs
Monocyte MPs
Total TFⴙ MPs
Monocyte TFⴙ MPs
TAT
0.436 (.002)
0.506 (.001)
0.459 (.002)
0.367 (.01)
0.406 (.007)
D-dimer
0.387 (.008)
0.365 (.01)
0.356 (.01)
0.454 (.002)
0.455 (.002)
PF1.2
0.398 (.008)
0.307 (.04)
0.258 (.09)
0.366 (.01)
0.380 (.01)
Data shown as r values, and P values are presented in parentheses.
BLOOD, 1 OCTOBER 2003 䡠 VOLUME 102, NUMBER 7
In parallel, markers of coagulation activation were abnormally
elevated in both crisis and steady state compared with healthy state,
consistent with previous reports.13,16,31 Total MPs and total TF-positive
MP levels correlated modestly, but significantly, with all 3 markers of
coagulation activation that were studied. Biochemical markers such as
D-dimer, TAT, and PF1.2 are measures of global activation of the
coagulation system that may be less sensitive as markers of tissue factor
exposure, explaining the rather modest correlations obtained.
Recent reports in patients with type 2 diabetes mellitus suggest
that platelet-derived MPs are positive for TF, but that study used a
platelet marker (CD61) that is shared by endothelial cells.32 Our
inability to detect platelet-derived TF-positive MPs could be
related to low levels, not detectable by FACS, or a biologic
difference between sickle cell disease (SCD) and diabetes. Although RBC- and platelet-derived MPs were not positive for TF,
they could still have potential functional significance because they
express surface PS and other coagulation factor receptors33 and
may be involved in acceleration of coagulation once it is initiated.31
TISSUE FACTOR⫹ MICROPARTICLES IN SICKLE BLOOD
2683
In summary, our data demonstrate that blood MPs in sickle cell
disease are derived in part from endothelial cells and monocytes
and a portion of these are also TF positive. This microparticleassociated TF is functionally active and could, therefore, contribute
to thrombotic occlusive events such as stroke in sickle cell
disease.17 In any case, their presence argues for the existence of
abnormal monocyte and endothelial activation in patients with
sickle cell disease both in steady state and crisis. In aggregate, these
data further support the idea that sickle disease is a chronic
inflammatory state with abnormal activation of the blood and
vessel wall.
Acknowledgments
We thank Dr Ron Bach for providing the anti-TF antibody and
helpful discussions and Carol Taubert for secretarial assistance.
References
1. Combes V, Simon AC, Grau GE, et al. In vitro
generation of endothelial microparticles and possible prothrombotic activity in patients with lupus
anticoagulant. J Clin Invest. 1999;104:93-102.
12. Berney SI, Ridler CD, Stephens AD, Thomas AE,
Kovacs IB. Enhanced platelet reactivity and hypercoagulability in the steady state of sickle cell
anemia. Am J Hematol. 1992;40:290-294.
2. Aupeix K, Hugel B, Martin T, et al. The significance of shed membrane particles during programmed cell death in vitro, and in vivo, in HIV-1
infection. J Clin Invest. 1997;99:1546-1554.
13. Hagger D, Wolff S, Owen J, Samson D. Changes
in coagulation and fibrinolysis in patients with
sickle cell disease compared with healthy black
controls. Blood Coagul Fibrinolysis. 1995;
6:93-99.
3. Zwaal RF, Schroit AJ. Pathophysiologic implications of membrane phospholipid asymmetry in
blood cells. Blood. 1997;89:1121-1132.
4. Nieuwland R, Berckmans RJ, McGregor S, et al.
Cellular origin and procoagulant properties of microparticles in meningococcal sepsis. Blood.
2000;95:930-935.
5. Wun T, Paglieroni T, Rangaswami A, et al. Platelet activation in patients with sickle cell disease.
Br J Haematol. 1998;100:741-749.
6. Allan D, Limbrick AR, Thomas P, Westerman MP.
Release of spectrin-free spicules on reoxygenation of sickled erythrocytes. Nature. 1982;295:
612-613.
7. Solovey A, Lin Y, Browne P, Choong S, Wayner E,
Hebbel RP. Circulating activated endothelial cells
in sickle cell anemia. N Engl J Med. 1997;337:
1584-1590.
8. Solovey A, Gui L, Key NS, Hebbel RP. Tissue factor expression by endothelial cells in sickle cell
anemia. J Clin Invest. 1998;101:1899-1904.
9. Belcher JD, Marker PH, Weber JP, Hebbel RP,
Vercellotti GM. Activated monocytes in sickle cell
disease: potential role in the activation of vascular endothelium and vaso-occlusion. Blood. 2000;
96:2451-2459.
10. Embury SH, Hebbel RP, Mohandas N, Steinberg
MH. Pathogenesis of vasooclusion. In: Embury
SH HRP, Steinberg MH, Mohandas N, eds. Sickle
Cell Disease: Basic Principles and Clinical Practice. New York: Raven Press; 1994:311-326.
11. Francis RB Jr, Hebbel RP. Haemostasis. In: Embury SH, Hebbel RP, Mohandas N, Steinberg MH,
eds. Sickle Cell Disease: Basic Principles and
Practice. New York: Raven Press; 1994:299-310.
14. Kurantsin-Mills J, Ofosu FA, Safa TK, Siegel RS,
Lessin LS. Plasma factor VII and thrombin-antithrombin III levels indicate increased tissue factor
activity in sickle cell patients. Br J Haematol.
1992;81:539-544.
15. Peters M, Plaat BE, ten Cate H, Wolters HJ,
Weening RS, Brandjes DP. Enhanced thrombin
generation in children with sickle cell disease.
Thromb Haemost. 1994;71:169-172.
16. Tomer A, Harker LA, Kasey S, Eckman JR.
Thrombogenesis in sickle cell disease. J Lab Clin
Med. 2001;137:398-407.
17. Prengler M, Pavlakis SG, Prohovnik I, Adams RJ.
Sickle cell disease: the neurological complications. Ann Neurol. 2002;51:543-552.
18. Key NS, Slungaard A, Dandelet L, et al. Whole
blood tissue factor procoagulant activity is elevated in patients with sickle cell disease. Blood.
1998;91:4216-4223.
19. Giesen PL, Rauch U, Bohrmann B, et al. Bloodborne tissue factor: another view of thrombosis.
Proc Natl Acad Sci U S A. 1999;96:2311-2315.
20. George JN, Thoi LL, McManus LM, Reimann TA.
Isolation of human platelet membrane microparticles from plasma and serum. Blood. 1982;60:
834-840.
21. Jimenez JJ, Jy W, Mauro LM, Horstman LL, Ahn
YS. Elevated endothelial microparticles in thrombotic thrombocytopenic purpura: findings from
brain and renal microvascular cell culture and
patients with active disease. Br J Haematol.
2001;112:81-90.
22. Allan D, Limbrick AR, Thomas P, Westerman MP.
Microvesicles from sickle erythrocytes and their
relation to irreversible sickling. Br J Haematol.
1981;47:383-390.
23. Naumann HN, Diggs LW, Barreras L, Williams BJ.
Plasma hemoglobin and hemoglobin fractions in
sickle cell crisis. Am J Clin Pathol. 1971;56:137147.
24. Reiter CD, Wang X, Tanus-Santos JE, et al. Cellfree hemoglobin limits nitric oxide bioavailability
in sickle-cell disease. Nat Med. 2002;8:13831389.
25. Satta N, Toti F, Feugeas O, et al. Monocyte vesiculation is a possible mechanism for dissemination of membrane-associated procoagulant activities and adhesion molecules after stimulation by
lipopolysaccharide. J Immunol. 1994;153:32453255.
26. Hebbel RP, Vercellotti GM. The endothelial biology of sickle cell disease. J Lab Clin Med. 1997;
129:288-293.
27. Nemerson Y. The tissue factor pathway of blood
coagulation. Semin Hematol. 1992;29:170-176.
28. Giesen PL, Nemerson Y. Tissue factor on the
loose. Semin Thromb Hemost. 2000;26:379-384.
29. Semeraro N, Colucci M. Tissue factor in health
and disease. Thromb Haemost. 1997;78:759764.
30. Adedeji MO, Cespedes J, Allen K, Subramony C,
Hughson MD. Pulmonary thrombotic arteriopathy
in patients with sickle cell disease. Arch Pathol
Lab Med. 2001;125:1436-1441.
31. Setty BN, Rao AK, Stuart MJ. Thrombophilia in
sickle cell disease: the red cell connection. Blood.
2001;98:3228-3233.
32. Diamant M, Nieuwland R, Pablo RF, Sturk A, Smit
JW, Radder JK. Elevated numbers of tissuefactor exposing microparticles correlate with components of the metabolic syndrome in uncomplicated type 2 diabetes mellitus. Circulation. 2002;
106:2442-2447.
33. Gilbert GE, Sims PJ, Wiedmer T, Furie B, Furie
BC, Shattil SJ. Platelet-derived microparticles
express high affinity receptors for factor VIII.
J Biol Chem. 1991;266:17261-17268.