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Blood, Vol. 91 No. 11 (June 1), 1998:
pp. 4216-4223
Whole Blood Tissue Factor Procoagulant Activity Is Elevated in
Patients With Sickle Cell Disease
By
Nigel S. Key,
Arne Slungaard,
Luke Dandelet,
Stephen C. Nelson,
Christopher Moertel,
Lori A. Styles,
Frans A. Kuypers, and
Ronald
R. Bach
From the Department of Medicine, University of Minnesota Medical
School, Minneapolis, MN; Children's Health Care-Minneapolis, MN;
Children's Health Care-St Paul, MN; the Children's Hospital Oakland
Research Institute, Oakland, CA; and Research Service, Veterans'
Administration Hospital, Minneapolis, MN.
 |
ABSTRACT |
We developed a simple assay for the measurement of tissue factor
procoagulant activity (TF PCA) in whole blood samples that avoids the
need for mononuclear cell isolation. This method combines convenience
of sample collection and processing with a high degree of sensitivity
and specificity for TF. Using this method, we have determined that TF
PCA is detectable in whole blood samples from normal individuals, which
is itself a novel observation. Essentially all PCA could be shown to be
localized in the mononuclear cell fraction of blood. Compared with
controls, whole blood TF levels were significantly (P < .000001) elevated in patients with sickle cell disease (SCD),
regardless of the subtype of hemoglobinopathy (SS or SC disease). No
significant difference in TF PCA was observed between patients in pain
crisis compared with those in steady-state disease. Because TF
functions as cofactor in the proteolytic conversion of FVII to FVIIa in
vitro, it was expected that an increase in circulating TF PCA would
lead to an increased in vivo generation of FVIIa. On the contrary,
FVIIa levels were actually decreased in the plasma of patients with
SCD. Plasma TF pathway inhibitor (TFPI) antigen levels were normal in
SCD patients, suggesting that accelerated clearance of FVIIa by the
TFPI pathway was not responsible for the reduced FVIIa levels. We
propose that elevated levels of circulating TF PCA may play an
important role in triggering the activation of coagulation known to
occur in patients with SCD. Because TF is the principal cellular ligand
for FVIIa, it is possible that increased binding to TF accounts for the
diminished plasma FVIIa levels.
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INTRODUCTION |
THE CLINICAL COURSE of sickle cell
disease (SCD) is punctuated by episodic vascular occlusive events. The
possibility that activation of the clotting system plays a contributory
role in these complications is supported by abundant clinical data
indicating that activation of platelets,1,2 plasma
coagulation,3,4 and fibrinolysis5 occurs during
both steady-state disease and pain crisis. In some studies, the markers
of activation of these pathways have been more accentuated during pain
crisis compared with steady-state disease,6 although
overall this trend is not consistent.7 Whether there is an
increased incidence of venous thrombo-embolic disease in SCD has not
been adequately evaluated, but thrombosis probably does play an
important role in several other recognized complications, including
stroke, acute chest syndrome, leg ulceration, and placental
infarction.8,9
Tissue factor (TF) is a transmembrane glycoprotein that forms a complex
with circulating FVII(a). Upon binding to TF, zymogen FVII may become a
better substrate for auto-activation and perhaps activation by other
serine proteases, including FIXa, FXa, FXIIa, and
thrombin.10,11 Although the precise mechanism of FVII
activation that operates in vivo has not been unequivocally elucidated,
there is evidence that FIXa may play a pivotal role.12 The
TF-VIIa complex activates factor X directly or indirectly via factor
IXa generation, ultimately leading to thrombin formation. As
demonstrated by immunocytochemical techniques, TF is localized to the
adventitia of normal blood vessels,13,14 in which location
it is believed to come into contact with blood only after vascular
injury. Although TF is not constitutively expressed by endothelial
cells and monocytes in vivo, it has been demonstrated that both cell
types are capable of de novo TF synthesis in vitro after exposure to
lipopolysaccharide (LPS; endotoxin) and other agonists, including
interleukin-1,15 tumor necrosis factor,16 and
C-reactive protein (CRP).17 Nonetheless, although TF is now
recognized as the sole physiological initiator of hemostasis, its role
in pathological thrombosis is less well established.
In a previously published study, accelerated plasma FVII turnover in
patients with SCD was reported, suggesting enhanced TF expression
and/or exposure of TF to circulating clotting factors in this
patient population.18 Using a novel method that we have developed to assay circulating whole blood TF (WBTF)
activity, we measured functional TF activity in whole blood samples
from normal controls and patients with SCD. Our results suggest that quantifiable TF activity is present in normal individuals and provide
confirmation that circulating TF procoagulant activity (PCA) is
elevated in patients with SCD.
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PATIENTS AND METHODS |
Patients.
Blood samples were collected from patients with both HbSS and HbSC
disease after the subjects or their guardians signed a consent form
that had been previously approved by the respective institutional
committee on the use of human subjects in research. Patient age ranged
from 3 months to 49 years (median, 8.1 years), and 47% of patients
were male. All samples from SCD patients were obtained at least 6 weeks
after any blood transfusion or exchange transfusion. Steady state was
assumed if the patient was removed from a pain crisis episode for at
least 2 weeks. Blood was generally obtained from patients in the
ambulatory setting for those in steady state and during hospitalization
for those in acute pain crisis. Controls consisted of normal Caucasian
and Black adults. A separate consent form for controls (approved by the
institutional human subjects committee review board) was used whenever
blood was drawn. Care was taken to obtain specimens by clean
venipuncture, after the first 3 mL of blood had been discarded.
Preparation of phosphatidylserine (PS):phosphatidylcholine (PC)
vesicles.
Unilamellar vesicles containing 70% PC:30% PS by weight were prepared
as previously described.19 Briefly, Tris-buffered saline
(TBS; 0.1 mol/L NaCl, 0.05 mol/L Tris, pH 7.5), supplemented with 375 mmol/L Octyl- -D-glucopyranoside (Calbiochem, La Jolla, CA), was
added to egg yolk PC and bovine brain PS (Sigma, St Louis, MO) to yield
a final phospholipid stock concentration of 25 mmol/L. Vigorous
vortexing was necessary to completely solublize the phospholipids. PS
and PC stocks were added to yield the desired phospholipid vesicle
ratio (by weight) of 30:70. Nitrogen exposure was maintained throughout
the procedure to minimize any potential phospholipid oxidation.
Vesicles were dialyzed into degassed buffers of TBS for 48 hours
followed by HEPES-buffered saline (HBS; 137 mmol/L NaCl, 5.38 mmol/L
KCl, 5.55 mmol/L glucose, 10 mmol/L HEPES, pH 7.5) for 24 hours at
4°C and stored under nitrogen at  70°C.
Sample processing for assay of WBTF PCA.
Blood was routinely collected into vacutainer tubes (Becton Dickinson,
Franklin Lakes, NJ) containing EDTA using a 21-gauge needle. In some control experiments, blood was also drawn into tubes
containing 3.2% trisodium citrate or heparin as anticoagulant. Anticoagulated samples were then transfered immediately to
polypropylene tubes and frozen at 70°C until TF PCA could be
assayed. Batched samples were subjected to 3 cycles of freezing on dry
ice followed by thawing at 37°C to completely lyse all cellular
elements. One hundred microliters of lysed blood and 900 µL TBS-0.1
mol/L EDTA, pH 7.5, was then centrifuged at 450,000g for 15 minutes in a Beckman TL100 ultracentrifuge (Beckman, Palo Alto, CA) to
pellet all particulate material (organelles and membrane
fragments/vesicles). Control experiments in which the
dilution factor of the sample and centrifugation times were varied were
performed to optimize conditions. Final conditions described for the
assay were those that maximized measurable PCA.
After aspiration of the supernatant, the membrane pellet was
resuspended in TBS-EDTA buffer and the washing/centrifugation cycle was
repeated twice. The membrane pellet was then washed one further time in
900 µL of HEPES buffered salt solution containing 0.1% bovine serum
albumin (HBSA). The sample was then resuspended in 100 µL HBSA for
assay of TF PCA, which was performed using the two-stage clotting
assay.
Two-stage clotting assay for TF.
In the first stage of the assay, 20 µL of the diluted sample was
mixed with 5 nmol/L recombinant FVIIa, 250 nmol/L human factor X (both
from Calbiochem), and 8.3 mmol/L CaCl2 in HBSA (final volume, 60 µL). After 3 minutes of incubation at 37°C, 100 µL of bovine plasma (Irvine Scientific, Santa Anna, CA) containing PS
30:PC 70 vesicles (12.5 µmol/L) and HEPES (50 mmol/L) together with
100 µL 25 mmol/L CaCl2 was added. The clotting end-point was determined in a Coag-a-mate optical end point instrument (Organon Teknika, Durham, NC).
A standard curve was constructed using relipidated human brain TF.
Human brain TF apoprotein prepared as previously described was
reconstituted into phospholipid vesicles containing 70% PC and 30%
PS.19 By definition, 1 pg of the standard gave 1 U of TF
PCA. A log-log plot of TF concentration versus clot time was linear in
the 1 to 1,000 pg/mL (1 to 1,000 U/mL) range, with a correlation
coefficient (r) value of .996.
Demonstration of TF PCA in the mononuclear cell (MNC) fraction of
whole blood.
To heparin-anticoagulated whole blood ex vivo, either LPS (1 µg/mL;
Sigma) or buffer (HEPES-buffered salt solution) control was added.
Samples were then incubated at 37°C for 6 hours to allow TF
induction in the LPS-treated tubes. The MNC fractions were then
isolated by layering the blood over Histopaque-1077 (Sigma) and
centrifuging (400g) at 24°C for 30 minutes. The MNCs collected from the plasma/Histopaque interface were then washed three
times with phosphate-buffered saline (PBS) and finally resuspended in 1 mL of HBSA. MNCs were then subjected to three freeze-thaw cycles, and
1/10 dilution of the lysate was suspended in HBSA before assay of TF
PCA. Parallel whole blood samples that had been exposed to LPS or the
control buffer were immediately freeze-thawed at the end of the 6-hour
incubation period without the additional MNC isolation step. These
samples were then processed according to the method for measuring WBTF
PCA described above. After appropriate adjustment of the raw data
according to dilutions that had been made, the total amount of TF PCA
(in units per milliliter) was calculated for the whole blood and
isolated MNC fraction, respectively.
Assay of TF PCA in intact cellular fraction of peripheral blood.
We used isolated intact peripheral blood cellular fractions to confirm
that TF PCA measured by the whole blood method is an accurate
representation of functional cell surface-expressed TF activity. In
these experiments, platelet-poor cell fractions were isolated from
blood drawn into heparin-anticoagulated vacutainer tubes. The samples
were centrifuged at 850g for 10 minutes, after which the
supernatant (platelet-rich plasma) was removed and discarded. In some
experiments, whole blood from control subjects was treated ex vivo with
1 µg/mL LPS for 6 hours at 37°C before isolation of the cellular
pellets. The cell pellet was then washed 3 times in TBS with 5 mmol/L
EDTA to remove any remaining contamination with soluble plasma
components. After a further two washes in TBS, cells were then
resuspended in TBS at a 30% hematocrit. A modified
two-stage assay for TF PCA was then performed in a Stago ST4
coagulometer (Diagnostica Stago, Parsippany, NJ). Recombinant FVIIa (5 nmol/L), human FX (250 nmol/L), and CaCl2 (8.3 mmol/L) were
then added (final volume, 120 µL) and the mixture was incubated at
37°C for 30 minutes. One hundred microliters of bovine plasma containing PS 30:PC 70 vesicles (12.5 µmol/L) and HEPES (50 mmol/L) was then added with 100 µL of 25 mmol/L CaCl2, and the
clotting time was recorded. Data are presented as the raw clotting
times recorded on the coagulometer.
Plasma FVIIa assay.
Platelet-poor plasma was prepared from citrated vacutainer tubes within
1 hour after collection by centrifugation at 17,000g for 15 minutes. To avoid possible cold activation of FVII zymogen, care was
taken to maintain samples at room temperature during transportation to
the laboratory and throughout processing. Plasma was frozen at
70°C until batched samples could be assayed. Plasma FVIIa
levels were measured as previously described, using a soluble mutant
form of TF (kindly provided by Dr J. Morrissey, Oklahoma Medical
Research Foundation, Oklahoma City) that is unable to facilitate conversion of FVII to FVIIa and therefore is sensitive only
to FVIIa (and not zymogen FVII) in clotting assays.20
Briefly, samples were thawed at 37°C and then diluted 1/10 in
FVII-deficient plasma (George King Biomedical, Overland Park, KS). One
hundred microliters of the soluble TF reagent (1 µmol/L soluble TF,
200 µmol/L PC 40:PE 40:PS 20 vesicles, 0.1% BSA in TBS) was
incubated for 200 seconds at 37°C. After this incubation, 100 µL
of the diluted plasma sample was added and incubated for an additional 30 seconds at 37°C. The reaction start time was initiated with the
addition of 100 µL of prewarmed 25 mmol/L CaCl2. Clot
times were determined by using the Coag-a-mate optical end point
instrument and converted to nanograms per milliliter from a standard
curve prepared using known concentrations of FVIIa that had been
diluted in FVII-deficient plasma.
Plasma TF pathway inhibitor (TFPI) antigen, FVII antigen, and
D-dimer enzyme-linked immunosorbent assay (ELISA).
Plasma TFPI antigen was measured by ELISA (American Diagnostica,
Greenwich, CT) in samples from patients with SCD and from controls.
Care was taken to ensure that samples were from subjects not currently
receiving heparin anticoagulation, because of the known effect of
heparin on increasing plasma TFPI levels.21 D-dimer was
quantified also by ELISA (American Diagnostica); the quoted upper end
of the normal range was 120 ng/mL. FVII antigen ELISA kits were
obtained from American Bioproducts (Parsippany, NJ).
Statistical analysis.
Because several of the variables analysed appeared to have skewed
distributions, logarithmic transformation of the data was performed.
Unless otherwise stated, all values presented are geometric means with
95% confidence intervals. The Student's t-test was used to
compare means of logarithmically transformed data. All reported
P values are two-tailed and a P value of less than .05 was considered to be significant. Spearman's rank correlation method
was used to calculate correlation coefficients between variables. These
values were calculated using Statistica software for the Macintosh
(Statsoft, Inc, Tulsa, OK).
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RESULTS |
Validation of the WBTF assay as a measure of circulating TF PCA.
We determined that, at an optimal 1/10 dilution of the processed
sample, clotting times fell within the linear range on a semilogarithmic standard curve plot (using relipidated human brain TF
as standard). We validated that the assay is specific for TF by
demonstrating that PCA was entirely dependent on the addition of FVIIa
in the first stage of the clotting assay and that activity could be
inhibited by more than 95% by the addition of previously described
monoclonal22 or polyclonal anti-TF antibodies23 (data not shown). To determine the precision of the assay, equal volumes of whole blood pellet from 20 normal or SCD samples was pooled
and vigorously mixed by vortexing before repetitive assay for TF PCA on
6 separate occasions. Intrasample coefficient of variation
(CV) was 5.7% for the pooled normal sample and
4.4% for the abnormal (SCD) sample. We also found that the level of WBTF PCA did not vary significantly in specimens obtained serially at a
given blood draw (after the first 3 mL of blood had been discarded),
indicating that contamination of the patient specimen with dermal
fibroblasts or other cells known to express high levels of
TF13,14 is unlikely to account for the measurable PCA.
However, when samples were drawn and processed on 6 different days for
3 control subjects, a significant day to day variability was evident,
yielding individual CVs of 35%, 41%, and 44%. The reason for this
moderately pronounced biologic variability for a given test subject is
unknown.
To demonstrate the inducible nature of TF PCA in whole blood, samples
from a normal donor were drawn simultaneously into EDTA- or
heparin-anticoagulated vacutainer tubes. To the whole blood ex vivo,
LPS (an agonist expected to induce monocyte TF synthesis) was added in
concentrations ranging from 0.001 to 1,000 ng/mL. Samples were then
incubated at 37°C for 6 hours, after which they were transferred to
polypropylene tubes, frozen at 70°C, and processed for assay
of WBTF PCA, as described above. As shown in
Fig 1A, the expected increase in TF PCA was
evident in a dose-response relationship in heparinized samples to which
LPS had been added. No such activity was inducible in EDTA samples,
consistent with previous studies indicating a requirement for
extracellular calcium for the optimal induction of TF.24 TF
PCA was inducible in samples from both normal controls and patients
with SCD that were treated with LPS ex vivo. Similar to the findings of
others,25 we observed a wide variation in individual
responses to LPS (from 2.4-to 9.0-fold increase), with no apparent
difference in the magnitude of the response between controls and SCD
patients. The fact that whole blood PCA is inducible by the addition of
an agent known to induce TF in circulating monocytes lends further
support to the concept that our assay is indeed measuring TF activity.
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| Fig 1.
WBTF PCA may be induced ex vivo by LPS and can be
recovered in the MNC fraction. (A) Samples of whole blood
anticoagulated in heparin ( ) or EDTA ( ) were
incubated with the indicated concentrations of LPS for 6 hours, after
which they were frozen before assay of TF PCA as described in Patients
and Methods. (B) WBTF PCA was measured in heparin-anticoagulated whole
blood ( ) both at baseline and 6 hours after incubation with LPS (1 µg/mL). In parallel samples, MNCs were isolated both at baseline and
after a similar incubation with LPS. TF PCA was then assayed in the
isolated MNC ( ). The data in each figure are from one experiment
that is representative of four such experiments.
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To determine what proportion of the observed PCA was to be found in the
MNC fraction of whole blood from normal donors, TF PCA was assayed in
the isolated (freeze-thawed) MNC fraction and compared with the value
obtained by whole blood assay. It is clear from Fig 1B that values for
TF PCA are essentially equivalent for both methods, both at baseline
and after LPS stimulation. These data suggest that essentially all TF
PCA that can be assayed by the whole blood method in normal individuals
is present in the MNC fraction, but they do not necessarily establish
monocytes as the sole source of this activity.
Increased WBTF PCA in normals and in patients with SCD.
As shown in Fig 2 and Table
1, detectable WBTF PCA could be measured in normal persons (n = 65),
with absolute values that ranged from 1.6 to 84.0 U/mL. Unlike some
parameters of hemostasis that have demonstrated enhanced activation of
coagulation in normal Black individuals when compared with
non-Blacks,3 TF PCA did not appear to demonstrate any such
racial differences. In 3 patients with high reticulocyte counts who had
undergone splenectomy for hemolytic anemias, TF PCA levels were normal,
with a geometric mean value of 20.2 U/mL.
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| Fig 2.
WBTF PCA is elevated in patients with SCD.
EDTA-anticoagulated whole blood samples from normal controls and
patients with SCD were frozen at 70°C immediately after
collection. WBTF PCA was measured as described in Patients and Methods.
Each symbol represents a single data point; samples are included from
both Black ( ) and Caucasian ( ) control subjects. Patient samples include those drawn during steady-state disease () as well as during
pain crisis ().
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Compared with controls, TF PCA was significantly (P < .000001) elevated for the SCD group as a whole (n = 85), with absolute values ranging from 3.1 to 438.6 U/mL. Values did not appear to differ
significantly according to type of hemoglobinopathy (hemoglobin SC [n = 15] or SS [n = 70] disease) or patient age ( 16 or >16 years of
age; n = 44 and 41, respectively). We hypothesized that WBTF PCA levels
would be significantly elevated in pain crisis (n = 42) compared with
steady-state disease (n = 43). Surprisingly, however, no such
association was seen. In several patients with SCD, in whom multiple
samples were available for assay, the status with respect to pain
crisis did not appear to be a primary determinant of the absolute
magnitude of WBTF PCA. Despite the absence of such an association, the
intersample variability in TF PCA was notably more pronounced than in
normal subjects; eg, in 3 patients with SCD in whom 6 or more samples
were available, individual CVs were 119%, 88%, and 59%.
We measured PCA of intact peripheral blood cell preparations to confirm
that the WBTF method is an accurate reflection of functional TF on
living cells. Because these samples contained red blood cells and
leukocytes, we chose to measure clotting times in the ST4 coagulometer,
which does not rely on an optical endpoint. As shown in
Fig 3, clotting times were significantly
shorter in LPS-treated control samples compared with their untreated
counterparts (P < .0005). This suggests that the PCA measured
in this assay, like that in the whole blood assay, is LPS-inducible, a
feature that favors that it is indeed due to TF. Furthermore, the
activity could be abrogated by 2 µg/mL of a specific polyclonal
antibody to TF23; eg, in one representative experiment, the
mean clotting time of an LPS-treated sample increased from 48.7 to
129.2 seconds with the addition of the antibody. This PCA was also
shown to be largely FVIIa-dependent, such that, when FVIIa was omitted, clotting times were typically greater than 300 seconds in controls and
95 to 100 seconds in LPS-treated samples. Clotting times in SCD samples
(n = 12) were significantly (P < .000001) shorter than
controls (n = 20). This is therefore indicative of a more pronounced
PCA in SCD patients and is consistent with the data obtained from TF
functional assay of lysed whole blood samples.
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| Fig 3.
Increased TF PCA in intact cellular fractions in SCD.
Platelet-depleted cell fractions were isolated by centifugation from heparin-anticoagulated whole blood. PCA was measured on these washed
intact (non-freeze-thawed) cell fractions using a two-stage assay as
described in Patients and Methods. P < .0005 for control versus LPS-treated controls; P < .000001 for control versus
SCD.
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Because absolute monocyte counts are known to be elevated in patients
with SCD,26 we considered the possibility that increased WBTF levels might simply be a function of monocytosis. Unfortunately, monocyte counts were not available for all patient samples in which TF
PCA was analyzed. However, in a subset of 34 patients, although
absolute monocyte counts were somewhat elevated (1.15 ± 1.05 × 109/L [mean ± SD]), no correlation existed
between the count and the measured WBTF PCA (r = .012;
P = .95). This result suggests that the number of functional TF
molecules per monocyte is quite variable or, alternatively, that some
other circulating cell type (or fragment thereof) contributes to the
total pool of TF PCA in patients with SCD.
Plasma FVIIa levels are decreased in patients with SCD.
In vitro studies suggest that TF acts as cofactor for the proteolytic
activation of zymogen FVII.11 In normal individuals, it has
been shown that approximately 1% of the circulating plasma pool of
FVII is present in the activated form.20,27 We therefore hypothesized that elevation of WBTF PCA would lead to accelerated activation of FVII, with a resultant increase in plasma [FVIIa] in
patients with SCD. Using a recently described assay that measures only
the activated form of FVII in plasma,20 we were therefore surprised to find the opposite result, namely, significantly diminished levels of FVIIa in SCD (Fig 4 and Table 1).
As shown, normal controls (n = 55) had plasma [FVIIa] that were
similar to previously published values.20,27 Patients with
SCD (n = 34) had levels that were significantly reduced (P < .0005 compared with controls). Plasma [FVIIa] did not differ
significantly for children 16 years of age (n = 13) compared with
adults (n = 21), and the levels did not differ between patients
according to whether their hemoglobinopathy subtype was SS (n = 27) or
SC (n = 7) disease. Although plasma [FVIIa] levels overlapped to a
greater degree between patients in crisis (n = 15) compared with those
in steady-state disease (n = 19), we did observe somewhat
higher levels during crisis. Despite the fact that plasma [FVIIa]
appeared to be approximately inversely related to TF PCA, the
correlation was not statistically significant either for controls
(r = .29; P = .22) or for patients with SCD (r = .21; P = .33).
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| Fig 4.
Plasma [FVIIa] is decreased in patients with SCD.
Citrated plasma samples from normal controls or patients with SCD were
assayed for plasma [FVIIa] as described in Patients and Methods. Each symbol represents a single data point; samples are included from both
Black () and Caucasian () control subjects. Patient samples include those drawn during steady-state disease () as well as during
pain crisis ().
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The half-life of FVIIa in vivo is significantly longer than other
serine proteases of the coagulation pathway, at approximately 2.5 hours.27 This is probably accounted for by the fact that, unlike these other enzymes, free FVIIa is not rapidly inactivated by
antithrombin III. However, although FVIIa bound to TF may be a target
for inactivation by antithrombin III,28 it is probably principally inactivated at this site by the TFPI-FXa
complex.29 To determine whether the decrement in functional
plasma [FVIIa] in SCD patients could be explained by increased levels
of TFPI, we assayed TFPI antigen levels in a subset of these plasmas
(Fig 5). The values in the control group
(107 ng/mL; range, 95 to 120 ng/mL; n = 15) were not significantly
different than those for patients with SCD (114 ng/mL; range, 102 to
126 ng/mL; n = 12; P = not significant [NS]). It also seems
unlikely that the extremely low plasma FVIIa levels that we observed in
SCD are simply a reflection of a low total circulating mass of FVII. In
a subset of 24 patients with SCD and 24 controls, FVII antigen levels
were assayed by ELISA. Although, as others have
described,18 mean FVII antigen was slightly lower in SCD
patients compared with controls (73% [range, 68% to 78%] v
80% [range, 76% to 83%], respectively; P = .05), the
magnitude of the difference is unlikely to account for the profound
difference in plasma [FVIIa] that we observed.
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| Fig 5.
Plasma TFPI antigen levels are normal in patients with
SCD. Plasma TFPI antigen was measured by ELISA. P = NS for
controls versus SCD.
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Plasma D-dimer levels are normal in patients with SCD.
To determine whether TF PCA correlated with other parameters of
activation of coagulation in vivo, we measured D-dimer levels as an
index of intravascular fibrin generation in a subset of patients and
controls. Similar to previously published data,3,5 we found
that D-dimer levels were significantly (P < .00005) elevated in SCD patients (124 ng/mL; range, 77 to 201 ng/mL; n = 17)
compared with normal controls (30 ng/mL; range, 27 to 34 ng/mL; n = 19). Moreover, levels were significantly elevated both in steady state (n = 11) and during pain crisis (n = 6) (P < .005 for each group compared with controls). Also in keeping with the
results of other studies,3 D-dimer levels were increased in
patients during crisis (327 ng/mL; range, 140 to 762 ng/mL) compared
with those not in crisis (73 ng/mL; range, 44 to 123 ng/mL; P < .05). However, notably, no significant correlation between TF PCA
and D-dimer levels was apparent for either controls (r = -.005;
P = .98) or patients with SCD (r = .24; P = .36).
| |
DISCUSSION |
We have developed a novel assay for the measurement of circulating
functional TF activity in whole blood samples. This assay is specific
for TF and sensitive to the presence of as little as 1 pg of TF antigen
per milliliter of whole blood. The sensitivity of the assay is well
below our ability to detect antigen on the surface of circulating cells
by fluorescence-activated cell sorting (data not shown). This may not
be surprising, because even if one assumes that all TF in a sample
containing (for example) 100 pg/mL of TF PCA is present exclusively on
monocytes in which location it is fully functionally intact, then each
cell will be expressing approximately only 300 molecules of TF.
Although the precision of the assay itself appears to be good, there is
apparently a significant intrinsic biologic variability, more marked in
patients with SCD, the reasons for which are as yet unclear. Perhaps
the greatest practical advantage offered by this method is the
convenience of sample collection; in the absence of any requirement for
MNC isolation, samples may be collected and frozen at 70°C
until the assay can be performed. Furthermore, because no isolation of
MNCs is required, the potential for artifactual increase in monocyte TF
activity, which may occur ex vivo during the isolation procedure or by
exposure of these cells to trace amounts of contaminating endotoxin, is
absolutely avoided.
Using this assay, we have demonstrated that TF PCA is present in the
blood of all normal individuals, itself a novel observation. Current
opinion suggests that, although TF may be inducible in monocytes and
endothelial cells, it is not constitutively expressed by blood cells in
their basal state.29,30 Our data demonstrate that, in
normal volunteers, essentially all basal and inducible TF activity in
whole blood is present in the mononuclear cell fraction of blood, but
do not prove that it is exclusively present in monocytes. However, it
is reasonable to conclude that this is the case, because lymphocytes
(the other major cell type in MNC fractions of whole blood) have not
been shown to express TF, either constitutively or after activation. We
cannot totally rule out the possibility that some other cell type,
perhaps even one that is not generally considered to be a circulating
peripheral blood cell, is contributing to the measurable PCA. In this
regard, we have recently confirmed a previous finding31
that the number of circulating endothelial cells (ECs) is increased in
patients with SCD and have shown that, in contrast to ECs from normal
donors, TF expression may be detected in a significant proportion of
ECs obtained from patients with SCD.32 However, preliminary
experiments in SCD donors in which whole blood was depleted of ECs
before measurement of WBTF PCA have suggested to us that less than 10% of the total pool of circulating TF activity could be accounted for by
the EC fraction (Key and Hebbel, personal observation). This result is consistent with the much greater absolute number of
circulating monocytes compared with ECs.
It may be argued that, because our assay is designed to measure total
TF activity in lysed cellular extracts, we could be measuring a pool of
TF that, although present within the cell, is not available on the
surface of intact cells to bind FVII(a) and thereby initiate
coagulation. Previous studies have shown that, in most cell types (with
the possible exception of some malignant clones), all cellular TF
apoprotein is present on the surface of cells in which it is
expressed.33 However, the fact that PCA increases after
cellular disruption, eg, by repeated freeze-thawing, suggests that full
functional expression of TF may be regulated by other
mechanisms.34,35 The biologic significance of this
regulated expression of cell surface TF PCA remains to be determined.
Regardless of this issue, we found that PCA demonstrating the
characteristics of TF was increased in intact cellular fractions of
blood from patients with SCD when compared with controls. Thus, we are
of the opinion that TF PCA measured in lysed cells in whole blood
samples is an accurate reflection of the relative amounts of in vivo TF
expression in health and in disease states. Nonetheless, it is likely
that the mere presence of intravascular TF may not be sufficient to
trigger coagulation, and cellular events that activate the full
expression of TF PCA may contribute to the enhanced intravascular
coagulation observed in patients with SCD.
Our data do not address the reason for increased TF expression in SCD.
It is tempting to speculate that the documented elevation in SCD of
plasma levels of soluble mediators known to induce TF synthesis in
monocytes (including tumor necrosis factor16,36 and
CRP17,37-39) may provide the stimulus for TF synthesis. We
failed to document any difference between WBTF levels for patients in
steady-state disease versus those in pain crisis. This may reflect the
fact that the distinction between crisis and steady-state disease is probably an artificial one. Subclinical ischemic episodes, as evidenced
by subjective assessment of pain and by periodic elevations in
biochemical and rheological indices, occur almost constantly in what is
generally considered to be the steady-state phase of the
disease.37-39 In contrast to TF PCA, we did detect higher
levels of FVIIa and D-dimers in patients during crisis, although the baseline values (ie, during steady-state disease) were also clearly abnormal. Reconciliation of these data suggests that TF PCA may lead to
a tonic activation of coagulation, but that there may also occur a
superimposed burst of increased turnover of coagulation factors
coincident with the onset of crisis. We are currently pursuing this
hypothesis by prospective serial evaluation of WBTF PCA, plasma
[FVIIa], and D-dimers in selected patients. The trigger for the
superimposed elevation of FVIIa and D-dimer over baseline with the
onset of crisis may be explained by increased access to TF PCA on EC or
in an extravascular site, either of which would not be detectable by
our assay.
One possible explanation for the apparent paradoxical decrease in
plasma [FVIIa] despite enhanced TF expression is that FVIIa undergoes
an accelerated clearance from the circulation. We note that normal
human volunteers who received a bolus infusion of endotoxin also
developed a paradoxical decrease in plasma FVIIa levels.40
In that study, FVIIa levels began to decrease within the first 2 hours
after endotoxin administration, but did not reach a nadir until 12 to
24 hours. Although not reported, based on previous studies it seems
reasonable to assume that peak expression of TF in monocytes (and
possibly endothelial cells) occurred at an earlier time point, perhaps
after 6 to 8 hours. Similarly, Mesters et al41
recently described that a rapid decrease in plasma [FVIIa] occurs in
neutropenic patients during sepsis, particularly when associated with
septic shock. In considering mechanisms to account for this phenomenon,
we measured plasma TFPI antigen levels, but found no decrease in
patients with SCD. It should be noted that TFPI levels have also been
found to be normal or elevated in other clinical conditions in which
there is evidence for TF-initiated thrombosis, including sepsis,
malignancy, and disseminated intravascular coagulation.42 Because TF is
the principal ligand for FVIIa, it remains a possibility that low
plasma FVIIa levels are due to accelerated clearance as a result of
binding to an increased number of available TF sites, whether they be
in an intravascular or extravascular location. Ultimately, the
possibility of accelerated clearance of FVIIa can probably only be
adequately addressed by in vivo studies of the fate of labeled FVIIa in
patients with high TF expression, such as SCD or sepsis.
In summary, our data unequivocally demonstrate that circulating levels
of TF PCA are elevated in patients with SCD. This pathologic expression
of TF may provide the proximate stimulus responsible for activation of
the coagulation system, which in turn may play an important role in
many of the vaso-occlusive complications of SCD.
| |
FOOTNOTES |
Submitted June 9, 1997;
accepted January 30, 1998.
Supported in part by grants from the Minnesota Medical Foundation, by
grants from the American Heart Association (Minnesota Affiliate), by
National Institutes of Health Grants No. HL55219 (to N.S.K.) and
HL55213 (to F.A.K.), and by grants from the Department of Veterans
Affairs (R.R.B.).
Address reprint requests to Nigel S. Key, MB, MRCP, University of
Minnesota Medical School, Box 480 Mayo, 420 Delaware St SE,
Minneapolis, MN 55455; e-mail: keyxx001{at}tc.umn.edu.
The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" is accordance with 18 U.S.C. section 1734 solely to indicate this fact.
| |
ACKNOWLEDGMENT |
The authors thank Dr James Morrissey for the gift of the soluble
recombinant TF mutant, Dr Robert Hebbel for helpful discussions, and Dr
Arkadiusz Dudek for assistance with FACS analysis. In addition, the
assistance of Jeanne Harkness, RN, in procuring patient samples is
gratefully acknowledged.
| |
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F. Samad, M. Pandey, and D. J. Loskutoff
Regulation of tissue factor gene expression in obesity
Blood,
December 1, 2001;
98(12):
3353 - 3358.
[Abstract]
[Full Text]
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J. D. Belcher, P. H. Marker, J. P. Weber, R. P. Hebbel, and G. M. Vercellotti
Activated monocytes in sickle cell disease: potential role in the activation of vascular endothelium and vaso-occlusion
Blood,
October 1, 2000;
96(7):
2451 - 2459.
[Abstract]
[Full Text]
[PDF]
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B. N. Y. Setty, S. Kulkarni, A. K. Rao, and M. J. Stuart
Fetal hemoglobin in sickle cell disease: relationship to erythrocyte phosphatidylserine exposure and coagulation activation
Blood,
August 1, 2000;
96(3):
1119 - 1124.
[Abstract]
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A. D. Schecter, B. Spirn, M. Rossikhina, P. L. A. Giesen, V. Bogdanov, J. T. Fallon, E. A. Fisher, L. M. Schnapp, Y. Nemerson, and M. B. Taubman
Release of Active Tissue Factor by Human Arterial Smooth Muscle Cells
Circ. Res.,
July 21, 2000;
87(2):
126 - 132.
[Abstract]
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U. Rauch, D. Bonderman, B. Bohrmann, J. J. Badimon, J. Himber, M. A. Riederer, and Y. Nemerson
Transfer of tissue factor from leukocytes to platelets is mediated by CD15 and tissue factor
Blood,
July 1, 2000;
96(1):
170 - 175.
[Abstract]
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E. de Jonge, P. E. P. Dekkers, A. A. Creasey, C. E. Hack, S. K. Paulson, A. Karim, J. Kesecioglu, M. Levi, S. J. H. van Deventer, and T. van der Poll
Tissue factor pathway inhibitor dose-dependently inhibits coagulation activation without influencing the fibrinolytic and cytokine response during human endotoxemia
Blood,
February 15, 2000;
95(4):
1124 - 1129.
[Abstract]
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M. T. GLADWIN, A. N. SCHECHTER, J. H. SHELHAMER, and F. P. OGNIBENE
The Acute Chest Syndrome in Sickle Cell Disease . Possible Role of Nitric Oxide in Its Pathophysiology and Treatment
Am. J. Respir. Crit. Care Med.,
May 1, 1999;
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P. L. A. Giesen, U. Rauch, B. Bohrmann, D. Kling, M. Roque, J. T. Fallon, J. J. Badimon, J. Himber, M. A. Riederer, and Y. Nemerson
Blood-borne tissue factor: Another view of thrombosis
PNAS,
March 2, 1999;
96(5):
2311 - 2315.
[Abstract]
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P. Andre, D. Hartwell, I. Hrachovinova, S. Saffaripour, and D. D. Wagner
Pro-coagulant state resulting from high levels of soluble P-selectin in blood
PNAS,
December 5, 2000;
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13835 - 13840.
[Abstract]
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Abstract
Introduction
Abstract