Download Platelet function in bleeding disorders

Platelet function in bleeding disorders
Esther Ruth van Bladel
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The research described in this thesis was financially supported by an unrestricted grant
from Sanquin Blood supply.
ISBN: 978-94-6108-540-5
All right reserved. No parts of this book may be reproduced or transmitted in any form or
by any means without written permission from the author.
Printed by: Gildeprint
Layout by: Textcetera
Cover design: Ilse Schrauwers, isontwerp
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Platelet function in bleeding disorders
Bloedplaatjes functie in bloedingsziekten
(met een samenvatting in het Nederlands)
Proefschrift
ter verkrijging van de graad van doctor aan de Universiteit Utrecht
op gezag van de rector magnificus, prof.dr. G.J. van der Zwaan,
ingevolge het besluit van het college voor promoties in het openbaar
te verdedigen op vrijdag 6 december 2013 des middags te 12.45 uur
door
Esther Ruth van Bladel
geboren op 16 januari 1985
te ‘s-Hertogenbosch
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Promotoren: Prof.dr. Ph.G. de Groot
Prof.dr. D.H. Biesma
Co-promotoren: Dr. M. Roest
Dr. R.E.G. Schutgens
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Table of contents
Chapter 1
Chapter 2
Chapter 3
Introduction
7
Up-regulation of platelet activation in hemophilia A
17
Factor VIII concentrate infusion in patients with hemophilia results in
decreased von Willebrand Factor and ADAMTS-13 activity
35
Chapter 4 Platelet behaviour is not related to bleeding phenotype
in severe hemophilia A patients
51
Chapter 5 FVIII uptake by platelets and megakaryocytes
69
Chapter 6 Platelets of patients with chronic kidney disease
demonstrate deficient platelet reactivity in vitro
85
Chapter 7 Functional platelet defects in children with severe chronic ITP:
as tested with two novel assays applicable for low platelet counts
97
Chapter 8 Capillary blood sampling in the assessment of platelet responsiveness 119
Chapter 9 General discussion
133
Chapter 10 Nederlandse samenvatting
144
Dankwoord
149
Curriculum Vitae
156
List of publications
157
Awards
157
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Cha
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Chapter 1
Introduction
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The hemostatic balance
Hemostasis is a well-balanced process essential to maintain undisturbed circulation of
blood through blood vessels, and to form a sealing clot when an injury to the vessel wall is
afflicted. Key players in keeping this balance are endothelial cells, platelets and coagulation
factors.
The vessel wall consists of 3 layers, intima, media and adventitia. The intima forms the inner
layer of the vessel wall, consisting of endothelial cells and and an underlying subendothelial
matrix. Endothelial cells form the barrier between blood and the adjacent tissue.1 When
the endothelium is damaged and its barrier function is lost, blood comes into contact with
the subendothelial matrix, which contains among others tissue factor, the physiological
activator of secondary hemostasis.2 Contact between blood and the exposed vessel wall
proteins leads to activation of primary and secondary hemostasis reactions, as described
below. Additionally, endothelial cells produce von Willebrand Factor (vWF)3, a protein
playing a mayor role in both primary and secondary hemostasis. The vessel wall layer media
consist of smooth musculature, which has the important role of vasoconstriction, reducing
blood flow in damaged vessels and thereby decreasing blood loss, in the case of bleeding.4
Primary hemostasis is the first reaction on vascular damage. Platelets are the key players
in the primary haemostatic response. Because platelets are smaller than red and white
blood cells, they are forced to the boundary layer close to the vessel wall, where they are
in close contact with endothelial cells.5 When there is vascular damage at an arterial side,
platelets require VWF to adhere to the subendothelial matrix to resist high shear rates in
arteries.6,7 VWF, released from endothelial cells and platelets, binds to the subendothelial
matrix protein collagen under high flow, inducing a conformational change in the von
Willebrand Factor protein. With this conformational change, the vWF platelet binding
site becomes available for binding the platelet glycoprotein-Ib-V-IX (GpIb:V:IX) receptor
complex.8,9 Through this platelet-vWF interaction, platelets start to roll over vWF strings,
slowing down and thereby allowing interaction of platelets with the subendothelial matrix
proteins. The platelet receptors glycoprotein VI (GPVI) and α2β1 bind to collagen, ensuring
a firm adherence of platelets and initiation of platelet activation.10,11 Platelet activation
manifests as:12
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a. Platelet shape change: resting platelets circulate in a discoid shape. When platelets
are activated, they change shape and spread over the surface, thereby expanding the
interaction between the platelet and the subendothelial matrix and strengthening their
connection.
b. Platelet degranulation: Platelets contain storage pools, the dense and alpha granules,
in which proteins involved in hemostasis, wound healing and inflammation are stored.
When platelets are activated, they release soluble proteins and factors into the
circulation or express membrane proteins on the outer platelet surface. One important
function of this granule release is secondary activation of platelets via binding of ADP to
surface receptors P2Y2 and P2Y12.
c. Platelet thromboxane formation: Next to the release of ADP, activated platelets establish
a second positive feedback loop. When activated, platelets convert arachidonic acid
into thromboxane-A2. Thromboxane diffuses out of the platelet where it can bind to
platelet surface thromboxane receptor, amplifying its own activation or activating
adjacent platelets.
d. Platelet glycoprotein IIbIIIa (GPIIbIIIa) opening and platelet aggregation: GPIIbIIIa is a
platelet surface receptor which, when activated, can bind fibrinogen. When platelets
are activated, the GPIIbIIIa receptor changes conformation, making it possible for
fibrinogen to bind. One fibrinogen molecule can bind two GPIIbIIIa receptors, thereby
forming a bridge between receptors and thus between platelets, facilitating platelet
aggregation. Next to fibrinogen, vWF is also a substrate for the GPIIbIIIa receptor,
making platelet aggregation possible when shear stresses are high.
e. Platelet procoagulant activity: When platelets become activated, the negative charged
phospholipids in the inner layer of the platelet membrane are transported to the outer
layer, generating a surface on which secondary hemostasis can take place.
1
Secondary hemostasis is the subsequent response to vascular damage in which the
coagulation cascade is activated to form fibrin, which is essential to stabilize the platelet plug.
In vivo, secondary hemostasis is initiated when tissue factor (TF) is exposed on damaged
tissue and binds activated clotting factor VII (FVIIa). This TF-FVIIa complex activates clotting
factor X (FX), which subsequently converts prothrombin (clotting factor II, FII) into thrombin
(activated clotting factor II, FIIa). At low tissue factor concentration, TF-VIIa activates
factor IX, factor IXa subsequently activates factor X. Thrombin is able to activate multiple
parts of secondary hemostasis, both amplifying and regulating the process:13
Introduction | 9
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a. Fibrin formation: thrombin converts fibrinogen into fibrin fibers. These fibers form a
firm network, stabilizing the platelet plug.
b. Amplification of thrombin generation via activation of co-factors: Secondary hemostasis
has two co-factors, clotting factor V (FV) and VIII (FVIII), which are activated by thrombin.
FVa and FVIIIa are able to accelerate the activation of FX by FIXa and the activation of FII
by FXa, respectively.
c. Positive feedback on thrombin generation via clotting factor XI (FXI): thrombin is able to
activate FXI, which is at the start of a second cascade of clotting factor activation, the
intrinsic pathway. FXIa is able to activate clotting factor IX (FIX), which in turn activates
FX, leading to additional thrombin formation.
d. Fibrin cross linking: Thrombin activates clotting factor XIII (FXIII). FXIIIa is able to cross
link fibrin fibers, increasing clot stability.
e. Activation of primary hemostasis: thrombin is able to activate platelets via the platelet
protease activated receptor (PAR). As described above, platelet activation is important
for secondary hemostasis, since it creates a negatively charged surface on which
secondary hemostasis can take place.
f. Inhibition of thrombin generation via activation of protein C system: when thrombin
binds to thrombomodulin, it is no longer able to convert fibrinogen, however, it is able
to activate protein C. Activated protein C functions as an inhibitor of coagulation via the
breakdown of the co-factors Va and VIIa and via activation of the fibrinolytic system.
g. Inhibition of fibrinolysis via Trombin Activatable Fibronolysis Inhibitor (TAFI) activation:
when TAFI is activated by thrombin it inhibits the lysis of a fibrin clot by removing
terminal lysine residues from fibrin which are necessary for binding and activation of
the fibrinolytic system.
h. Activation of fibrinolytic system: thrombin stimulates endothelial cells to release tissue
plasminogen activator (tPA), leading to formation of plasmin which is able to break
down fibrin. This fibrinolytic system is responsible for clot removal in the long run.
vWF, next to its attribution in primary hemostasis, also plays an important role secondary
hemostasis, as it is the carrier protein of clotting factor VIII (FVIII). Without vWF as its
carrier, FVIII has a shortened survival time in plasma, leading to reduced FVIII activity levels.
Classically, primary and secondary hemostasis were thought to occur one after the other.
However, with the development of in vivo thrombus formation imaging, it was shown
that primary and secondary hemostasis do not occur as two separate processes, but that
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fibrin fibers start to stabilize the platelet plug within 30 seconds after initiation of vascular
damage.14
1
Disturbance of the hemostatic balance
When a defect in this hemostatic process arises, a patient can be affected with either a
bleeding or a thrombotic tendency. Here we will focus on diseases leading to a bleeding
tendency. These diseases can either arise from a disorder of a component of primary or
secondary hemostasis, as in thrombocytopenia or in hemophilia patients; but also nonhemostatic diseases can affect components of hemostasis leading to a bleeding tendency
in time.
In hemophilia A, clotting factor VIII (FVIII) is affected, leading to a deficiency in secondary
hemostasis and therewith to a bleeding tendency. On a group level, the residual FVIII
activity is the main determinant in bleeding tendency is this patient category.15 The lower
the FVIII activity, the stronger the tendency to bleed. However, on individual patient level,
there are several patients with severe bleeding tendencies, which cannot be fully explained
by their FVIII activity level. When considering severe hemophilia A patients (in whom no
FVIII activity can be measured), 10% of these patients show a mild tendency to bleed. On
the other hand, some patients with moderate hemophilia (FVIII 1-5 IU/dL) bleed more than
is expected from their FVIII levels.16,17
In chronic kidney disease, both an increased bleeding tendency and an increased
thrombotic tendency are observed. The haemostatic balance seems not to be disturbed
but a much smaller event can tip the scales more easily.18-21 It has been suggested that
abnormal platelet function is a major contributor in bleeding, since hemorrhage occurs
despite a coagulation profile of normal or elevated levels of coagulation factors and normal
platelet counts. Platelet dysfunction is thought to be caused by the action of uremic
toxins, anemia, increased nitric oxide production and the use of medication.21-24 Besides an
increased bleeding risk, a variety of thrombotic complications are observed in patients with
chronic renal failure, including coronary heart disease, cerebro-vascular disease, peripheral
vascular disease and heart failure. Already in mild to moderate chronic kidney disease an
increased risk of cardiovascular events and higher mortality have been reported.18,19,25-28
Introduction | 11
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In immune thrombocytopenia (ITP), formation of auto-antibodies leads to decreased platelet
counts.29,30 Patients present with sudden onset of a bleeding tendency. ITP is known to
present in two forms: acute and chronic ITP.31 In acute ITP, antibodies are thought to result
in accelerated clearance of platelets and megakaryocytes, and may also lead to decreased
production of platelets. In the majority of pediatric ITP patients, thrombocytopenia resolves
spontaneously within weeks or months. In about 25% of the patients, thrombocytopenia
persists and becomes chronic.31,32,33 During chronic ITP, the attribution of auto antibodies
to the pathogenesis of thrombocytopenia is less clear. Platelet counts can vary in time from
very low (<10*109/L) to almost normal. However, the observed bleeding tendency does
not strictly follow platelet count. Patients with very low platelet counts have proven to
not always lead to bleeding problems.33 Moreover, patients with relatively higher platelet
counts can present with severe bleeding problems.
Hypothesis
The intensity of bleeding tendencies is difficult to predict for individual patients. We
hypothesized that the inter-individual variability in platelet response is a major determinant
of the inter-individual variability in bleeding tendency in disease. Improved platelet function
tests might improve the prediction of bleeding tendency in individual patients and provide
tools to develop customized treatment strategies.
Platelet reactivity assay
To determine platelet function, we developed a platelet reactivity assay. With this assay,
platelet reactivity to specific (ant-)agonists is tested in their natural environment, i.e. in whole
blood. For every (ant-)agonist, a concentration series diluted from a high concentration
known to generate maximal platelet reactivity to a low concentration known not to activate
platelets was used. Agonist for this assay were ADP to activate platelets via P2Y receptors
(P2Y1 and P2Y12), convulxin (CVX) or cross-linked collagen related peptide (CRP) to activate
the collagen receptor glycoprotein VI, thrombin receptor activator peptide (TRAP) to
activate Proteinase Activated Receptor-1 (PAR-1) and U46619 to activate the thromboxane
receptor. Iloprost was used as antagonist, inhibiting platelet activation via the prostacyclin
receptor. With the use of antibodies or proteins labeled with various fluorescent labels,
multiple manifestations of platelet activation can be measured at the same time. When
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platelets degranulate, and granule membranes fuse with the outer platelet membrane,
the membrane bound protein P-selectin is displayed on the cell surface. Platelet P-selectin
expression was visualized by fluorescent ascending cell sorting (FACS) via the addition of
fluorescent labeled anti-P-selectin antibody. Simultaneously, another manifestation of
platelet activation, opening of the GPIIbIIIa receptor, can be measured by the addition of
fluorescent labeled fibrinogen. In comparison to classical platelet function test, the platelet
reactivity assay is able to determine platelet function at a single platelet level, for specific
activating or inhibiting pathways and with measurement of specific manifestations of
platelet activation. Moreover, it is a whole blood based assay, requiring a minimal amount
of blood a minimal amount of proceedings.
1
Outline of this thesis
The first bleeding diathesis we studied was hemophilia A. Since FVIII activity level does
not always correlate with the bleeding tendency in individual patients, bleeding tendency
must also be influenced by other factors. Earlier studies excluded the remaining clotting
factors and FVIII genotype as determinants of bleeding phenotype in hemophilia A. To
look into the role of platelets, we set up four studies. In Chapter 2, we studied differences
in platelet reactivity between patients with mild, moderate and severe hemophilia A and
healthy individuals. In Chapter 3, the influence of FVIII concentrate infusion, on primary
hemostasis was studied. In Chapter 4, we investigated if differences in platelet reactivity
could explain differences observed in bleeding phenotype of severe hemophilia A patients.
To better understand the relationship between factor VIII and platelets, we investigated
the capability of platelets to take up FVIII in Chapter 5.
The next patient group drawing our attention is patients with chronic kidney disease.
In this patient category, both an increased bleeding tendency and a increased thrombotic
tendency is observed. In Chapter 6, we investigated platelet reactivity in patients with
chronic kidney disease, to determine if platelets contribute to either the bleeding or the
thrombotic tendency in these patients.
The last patient group we investigated here were children with chronic ITP. In chronic
ITP, patient platelet count does not always correlate to the bleeding tendency in individual
patients. To investigate a possible role of platelet function in this, we adapted the platelet
reactivity assay for measurement in samples with very low platelet counts. In Chapter 7, we
investigate platelet function in patients with chronic ITP related to bleeding phenotype of
these patients.
Introduction | 13
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The platelet reactivity assay, which was used in above studies to determine platelet
functionality, is a test which requires only a very small amount of whole blood. Since only
250 microliters of blood is sufficient to test platelet reactivity to five (ant-)agonists, capillary
sampling could be used to collect this small amount. In Chapter 8 we investigate if capillary
sampled blood can be used for functional platelet testing via the platelet reactivity assay.
First this chapter focuses on the differences introduced by capillary sampling compared
to venous sampling on platelet function. Second we focus on minor modifications to the
capillary sampling, with which platelet reactivity testing becomes possible in capillary
sampled blood.
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1
Introduction | 15
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Chapter 2
Up-regulation of platelet activation in hemophilia A
Esther R. van Bladel,1 Mark Roest,2 Philip G. de Groot,2 and Roger E.G. Schutgens3
1
Department of Clinical Chemistry and Hematology/Van Creveldlaboratory, University Medical
Center Utrecht, Utrecht, the Netherlands; 2Department of Clinical Chemistry and Hematology,
University Medical Center Utrecht, Utrecht, the Netherlands; and 3Van Creveldkliniek/
Department of Hematology, University Medical Center Utrecht, Utrecht, the Netherlands.
Haematologica 2011;96(6):888-895.
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Abstract
Background: Platelets are an underappreciated factor in the classification of bleeding
tendency of hemophilia patients. In this cross-sectional study, we have investigated platelet
activation status and responsiveness in relation to residual FVIII activity and, within the
severe (<1% residual FVIII activity) hemophilia patients, to annual FVIII consumption. Design
and Methods: Twenty-one mild-moderate hemophilia A patients, 13 severe hemophilia
A patients and 21 healthy controls were included. Basal level of platelet activation and
platelet responsiveness to activation and inhibition were determined by the measurement
of platelet P-selectin expression and soluble platelet activation markers. Results: Severe
hemophilia A patients showed a higher percentage of activated platelets at baseline (15.9%)
when compared to mild-moderate patients (8.2%, p=0.014) and controls (6.4%, p<0.001).
Both mild-moderate and severe hemophilia A patients had higher levels of soluble platelet
activation markers Platelet Factor 4 (PF4) (1.4 and 1.8 pg/106 platelets), CXCL7 (65.8
and 48.2 pg/106 platelets) and RANTES (12.8 and 9.5 pg/106 platelets), when compared
to controls (PF4: 0.3 pg/106 platelets, p<0.001 and <0.001; CXCL7 20.0 pg/106 platelets,
p<0.001 and <0.001; RANTES 4.5 pg/106 platelets, p<0.001 and =0.003, respectively). In
support of these observations, we found clinical evidence that higher platelet P-selectin
expression correlates with a lower FVIII consumption in patients with severe hemophilia
(Spearman’s r -0.65, p=0.043). Conclusion: This study indicates that platelets of severe
hemophilia A patients are in a pre-activated state and that this pre-activated state is
associated with FVIII consumption.
18 | Chapter 2
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Introduction
Hemophilia A is an X-linked disorder of coagulation factor VIII (FVIII). Patients with
hemophilia (PWH) suffer from a bleeding tendency of various degrees. Currently, the
bleeding severity is roughly classified by the residual FVIII activity of the patient. In general,
patients with lower FVIII activity levels have a more severe clinical phenotype than patients
with higher FVIII activity.1 Accordingly, PWH are classified into mild (FVIII activity 6-40%),
moderate (FVIII activity 1-5%) and severe (FVIII activity <1%) hemophilia A.2,3
Although this classification on residual FVIII activity is associated with clinical phenotype,
it is not a precise predictor of the bleeding pattern in individual patients. Within the group
of patients classified as severe hemophilia A, up to 10% of patients have a mild clinical
phenotype4,5, indicating that residual FVIII activity level is not the sole determinant of
clinical phenotype.
Differences in bleeding phenotype within the hemophilia A classifications are not
completely understood. Prothrombotic factors, FVIII half-life, genotype and fibrinolytic
activity did not correlate with bleeding patterns in individual patients.6,7
No information is available on the contribution of platelets to the clinical phenotype
in hemophilia. In normal hemostasis, platelets and the coagulation system interact with
each other to form a plug at the side of vascular damage.8 Platelets are the most important
source of negatively charged phospholipids. We hypothesized that the reduced coagulation
by the absence of FVIII can be (partly) compensated by an increased platelet reactivity.
Therefore, we have investigated platelet activation and platelet responsiveness to different
agonists in PWH in relation to their residual FVIII activity.
2
2
Design and Methods
Patients and setting
In severe (<1% FVIII activity), moderate (1-5% FVIII activity) and mild (6-40% FVIII activity)
PWH, blood was drawn during regular visits at the Van Creveldkliniek (VCK), University
Medical Center Utrecht, from the antecubital vein through 20-gauge needles, into a vacuum
citrate tube. Data on residual FVIII activity, inhibitor status, annual FVIII consumption and
blood group were collected from patient medical files.
All participants of the current study were aged 18 years or older. Subjects were excluded
when taking non-steroidal anti-inflammatory drugs, which are known to influence platelet
function. Healthy, male volunteers, of similar age served as controls. During a period of
Up-regulation of platelet activation in hemophilia A | 19
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1 year, between February 2009 and February 2010, 35 PWH and 21 healthy controls were
included in this study. One PWH was excluded, because blood was erroneously stored
on ice after drawing; leaving 34 PWH for analysis. Based on their residual FVIII activity
levels, patients were divided into mild-moderate PWH (n=21) or severe PWH (n=13). All
participants gave written informed consent and the study was approved by the Medical
Ethics Committees of the University Medical Center Utrecht and performed in accordance
with the Declaration of Helsinki.
Materials
R-phycoerythrin (PE) labelled antibodies for fluorescence activated cell sorting (FACS)
analyses, raised against human P-selectin (#555524), were purchased from BD biosciences
(Franklin Lakes, NJ, U.S.A). Antibodies against human Platelet Factor 4 (PF4; MAB7951,
AF795), human chemokine (C-X-C motif) ligand 7 (CXCL7) (MAB393, BAF393), human
chemokine (C-C motif) ligand 5 (RANTES) (MAB278, AB-278-NA) and human soluble
P-selectin (DY137 Duoset) were all purchased from R&D Systems (Abingdon, U.K.). Rabbit
anti-vWF propeptide and rabbit anti-vWF propeptide/biotine were prepared as described
by Borchiellini et al.9 Polyclonal rabbit anti-human von Willebrand Factor (vWF) antibodies
(A0082), rabbit anti-goat horseradish peroxidase (HRP) (P0449) and streptavidin-poly-HRP
(P0397) were purchased from DAKO (Glusdorp, Denmark).
For platelet responsiveness assays, adenosine diphosphate (ADP) was purchased from
Roche (Almere, The Netherlands), Iloprost (Ilomedine) from Bayer Schering Pharma AG
(Berlin, Germany) and Cross-linked Collagen Related Peptide (XL-CRP) was a generous gift
of R. Farndale (Cambridge, U.K.)
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) was purchased form
BDH (Poole, U.K.). Sodium chloride (NaCl), Hydrogen Peroxidase (H2O2), Tween-20 and
Bovine serum albumin (BSA) were purchased from Sigma (Zwijndrecht, The Netherlands).
Magnesium Sulphate (MgSO4) and Potassium Chloride (KCl) were purchased from Riedel
(Seelze, Germany), Formaldehyde (CH2O) and Orthophenyldiamine (OPD) from Calbiochem
(Merck, Darmstadt, Germany). Sulfuric Acid (H2SO4) was purchased from Mallinckrodt
Baker Inc. (Deventer, The Netherlands), Amplex UltraRed reagent from Invitrogen (Breda,
the Netherlands) and SuperSignal ELISA Pico chemiluminescent substrate from Thermo
Scientific (Rockford, Illinois, U.S.A.).
Platelet activation and responsiveness
Platelet responsiveness to agonists was determined with concentration series of ADP and
CRP while platelet responsiveness to inhibitors was measured with concentration series
20 | Chapter 2
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of iloprost in platelets activated with a suboptimal dose of ADP. Serial dilutions of ADP
(500 μM, 50 μM, 5 μM, 500 nM, 50 nM an 5 nM) were prepared in 50 μL HEPES buffered
saline (HBS; 10 mM HEPES, 150 mM NaCl, 1 mM MgSO4, 5 mM KCl, pH 7.4, filtered through a
0.22 μm filter) with 2 μL PE labeled mouse anti-human P-selectin antibodies. Similarly, serial
dilutions of CRP (2.5 μg/mL, 250 ng/mL, 25 ng/mL, 2.5 ng/mL and 250 pg/mL) were prepared
in 50 μL HBS with 2 μL PE labeled mouse anti-human P-selectin antibodies. Furthermore,
a serial dilution of iloprost (250 ng/mL, 25 ng/mL, 2.5 ng/mL, 250 pg/mL, 25 pg/mL and
2.5 pg/mL) was prepared in 50 μL HBS with 5 μM ADP and 2 μL PE labeled mouse antihuman P-selectin antibodies. A negative control sample, only containing 50 μL HBS with
2 μL PE labeled mouse anti-human P-selectin antibodies, was prepared to determine the
basal level of platelet activation.
The platelet activation test was initiated by addition of 5 μL fresh, citrate anticoagulated,
whole blood to each sample of the serial dilutions. After 20 minutes of incubation, the
samples were fixed with 500 μL 0.2% formyl saline (0.2% formaldehyde in 0.9% NaCl,
filtered through a 0.22 μm filter) and kept at 4°C until analyses. All samples were analysed
on a FACSCalibur flow cytometer from BD Biosciences (Franklin Lakes, NJ, U.S.A.) within
one day after processing. Single platelets were gated based on forward and side scatter
properties. The mean fluorescence intensity (MFI) in the platelet gate was measured with
FACS analysis. Platelets were defined as positive for P-selectin expression when the MFI
exceeded the isotype control. One individual performed all assays.
2
2
ELISA procedure
Citrated whole blood was centrifuged twice at 2000xG for 10 minutes at room temperature
and the resultant plasma was collected and frozen at -80°C for evaluation. Plasma levels
of soluble platelet activation markers PF4, CXCL7, soluble P-selectin (sP-sel) and RANTES,
and of vWF propeptide and vWF antigen (vWF:Ag) were measured by enzyme-linked
immunosorbent assay. Plasma samples of patients and controls were mixed randomly for all
ELISA measurements. Each antigen was measured on a separate Nunc maxisorb ELISA plate
(Thermofischer Scientific, Roskilde, Denmark). Capture antibodies, monoclonal mouse antihuman PF4 (1 μg/mL), purified monoclonal mouse anti-human CXCL7 (1 μg/mL), mouse
anti-human P-selectin (1 μg/mL), purified mouse monoclonal anti-human RANTES (500 ng/
mL), rabbit anti-human vWF propeptide (5 μg/mL) and polyclonal rabbit-anti-human vWF
(5 μg/mL) were coated on different plates overnight. Unbound antibodies were washed
with five steps PBS/0.5%TWEEN.
Plasma samples were diluted 1/75 for PF4 and CXCL7, 1/10 for soluble P-selectin, 1/4 for
RANTES, 1/20 for vWF propeptide and in 1/20, 1/40, 1/80 and 1/160 dilutions for vWF:Ag
Up-regulation of platelet activation in hemophilia A | 21
Bladel.indd 21
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measurements, respectively and then added in duplo to the plate with the corresponding
capture antibody. Each plate contained two calibration curves (duplex) consisting of
dilution ranges of a standard serum sample with known PF4, CXCL7, soluble P-selectin and
RANTES concentrations or a dilution range of a normal pooled plasma sample with known
vWF propeptide and vWF:Ag concentrations. Dilutions were made in PBS/1%BSA for PF4,
CXCL7, soluble P-selectin and RANTES measurements, in PBS/0.05%Tween/3.8%EDTA for
vWF propeptide and in PBS/0.05%Tween/3%BSA for vWF:Ag measurement.
For PF4 and CXCL7 measurements unbound antigens were removed with five wash steps
with PBS/0.5%TWEEN. Detection antibodies, polyclonal goat anti-human PF4 (0.05 μg/mL)
and biotinylated goat anti-human NAP-2 (50 ng/mL), were added to the corresponding
plates. After five washing steps with PBS/0.5% TWEEN detection antibodies, Rabbit antigoat-HRP (2 μL in 13 mL) and streptavidine-poly-HRP (2 μL in 13 mL), were added for
1 hour to bind the biotin on the detection antibody. After five other washing steps, Amplex
Ultrared reagent was added and fluorescence intensity was measured (emission: 490 nm;
excitation: 520 nm), after 75 minutes incubation, with a Fluostar Galaxy fluorimeter from
BMG Labtech GmbH (Offenburg, Germany).
For soluble P-selectin, RANTES and vWF propeptide measurements unbound antigens
were removed with five wash steps with PBS/0.5%TWEEN. Detection antibodies, mouse
anti-human P-selectin (0.01 μg/mL), purified goat anti-human RANTES (1 μg/mL) and rabbit
anti-propeptide/biotine (2.5 μg/mL), were added to the corresponding plates. After five
washing steps with PBS/0.5% TWEEN, streptavidine-poly-HRP was added to the soluble
P-selectin plate and the vWF propeptide plate, and Rabbit anti-goat HRP was added to the
RANTES plate for 1 hour to bind the detection antibody. After five other washing steps,
SuperSignal ELISA Pico chemiluminescent substrate was added and luminescence was
measured (emission: 490 nm) after 60 minutes incubation, with a SpectraMax L microplate
reader from Molecular Devices Inc. (Silicon Vally, CA, U.S.A.).
Three wash steps with PBS/0.5%Tween were performed to remove unbound antigens
for vWF:Ag measurement. Polyclonal rabbit-anti-human vWF/HRP secondary antibodies
(1.1 mg/mL) were added to the plates and incubated for 1 hour. After three other washing
steps, activated OPD reagent was added to stain the wells. After 3 minutes of incubation
the staining reaction was stopped with H2SO4, after which the absorption was measured
(absorption at 490 nm), with a VersaMax tunable microplate reader from Molecular Devices
Inc. (Silicon Vally, CA, U.S.A.).
22 | Chapter 2
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Statistical Analysis
Initial quantification of FACS data was performed in BD CellQuest Pro software, version 6.0
(BD biosciences, Franklin Lakes, NJ, U.S.A.). To determine platelet responsiveness, the
concentration generating a response halfway between baseline and maximum MFI of
P-selectin expressing platelets (EC50) was calculated using GraphPad Prism version
4.00 (GraphPad Software, San Diago, California, USA). The maximal effect of stimulation
or inhibition is represented by the platelet response to the highest used (ant-)agonist
concentration, in both percentage of P-selectin expressing platelets and the MFI of
P-selectin expressing platelets.
A statistical analysis was performed with SPSS version 15.0.1 for Windows (SPSS Inc.,
Chicago, Illinois, USA). Data are expressed as median ± interquartile range (IQR) and
figures show individual values and group medians unless indicated otherwise. Comparison
between two groups was performed by Mann-Whitney U testing. P-values lower than 0.05
were considered to be significant.
2
2
Results
Baseline characteristics
Of the 34 analyzed PWH, one moderate PWH and 9 (69%) severe PWH received regular
prophylaxis with FVIII concentrates. Within the group of severe PWH, 3 (23%) patients had
active inhibitors during time of this study. From the severe PWH, 4 (30.8%) had a hepatitis
C infection at time of inclusion. Hemophilic arthropathy was present in 12 (92.3%) severe
PWH and in 9 (42.9%) mild-moderate PWH. Baseline characteristics of the study population
are shown in Table 1. Individual patient characteristics are shown in Table 2.
Basal level of platelet activation
The percentage of platelets with P-selectin expression on the platelet membrane was
increased in severe PWH (15.9% (10.3-21.1%)) when compared to mild-moderate PWH
(8.2% (4.8-14.5%), p=0.014) and to healthy controls (6.4% (4.7-7.6%), p<0.001) (Figure 1A).
Similarly, platelets of severe PWH showed a significant higher P-selectin expression (6.5
(4.8-9.1)) than either mild-moderate PWH (4.2 (3.3-5.9); p=0.010) or healthy controls (3.8
(3.1-4.1); p<0.001) (Figure 1B).
When correlating the mean annual FVIII consumption of the severe PWH to the
percentage of platelets with P-selectin expression on the platelet membrane, we found a
higher percentage of P-selectin expression to be correlating with a lower FVIII consumption
Up-regulation of platelet activation in hemophilia A | 23
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Table 1. Baseline Characteristics
Male, n (%)
Median age, y (IQR)
Treatment, n (%)
Prophylaxis
On demand
Inhibitors, n (%)
Infection, n (%)
Chronic hepatitis
HIV
Hemophilic arthropathy, n (%)
Median Platelet Count, G/L
(IQR)
Median MPV, fL (IQR)
Blood group O, n (%)
Healthy
Control
(n=21)
Mild/
Severe
Moderate
Hemophilia A
Hemophilia A (n=13)
(n=21)
p mildp severe
moderate vs.
vs.
control
control
21 (100)
37 (29-51)
21 (100)
40 (26-60)
13 (100)
39 (28-55)
1.000
1.000
1.000
0.696
NA
NA
NA
1 (4.8)
20 (95.2)
0 (0)
9 (69.2)
4 (30.8)
3 (23.1)
NA
NA
NA
NA
NA
NA
0 (0)
0 (0)
NA
0 (0)
0 (0)
9 (42.9)
4 (30.8)
0 (0)
12 (92.3)
NA
NA
NA
NA
NA
NA
238 (202-282) 226 (197-284)
208 (160-287) 0.642
0.330
8.4 (6.8-8.9)
9 (42.9)
7.5 (7.1-8.6)
6 (46.2)
0.517
0.853
8.3 (7.3-8.5)
11 (52.4)
0.764
0.542
IQR indicates interquartile range, MPV Mean Platelet Volume, NA not applicable.
Table 2. Individual patient characteristics
Residual
Inhibitor Mutation
FVIII
status
activity (%) (BU)
Time between Mean annual Blood
last major
consumption group
bleeding and
(FVIII in IU)
inclusion (days)
0
0.0
>90
165000
O
0
0.0
>90
287000
O
0
0.0
0-180 days
316000
A
0
0
0
0
0
0.0
0.0
0.0
0.1
0.1
>90
>90
>90
>90
>90
132500
92000
527250
169750
260000
O
A
A
O
O
0
0.1
>90
166750
O
0
0.1
0-180 days
166250
A
1619delC
mutation in exon 11
2946insA
mutation in exon 14
Ala200Pro
mutation in exon 5
Type II inversion
Type II inversion
Type II inversion
unknown
2946insA
mutation in exon 14
Ala1920THR
mutation in exon 17
Type I inversion + Asp1241Gln
mutation in exon 14
24 | Chapter 2
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Table 2. Continued
Residual
Inhibitor Mutation
FVIII
status
activity (%) (BU)
Time between Mean annual Blood
last major
consumption group
bleeding and
(FVIII in IU)
inclusion (days)
0
12.0
± 60
0
AB
0
0
2
2
2
23.2
34.5
0.0
0.1
0.1
>90
>90
>90
>90
>90
0
0
188000
36750
88250
A
A
O
O
AB
3
3
3
3
3
0.0
0.0
0.0
0.0
0.0
>90
>90
>90
>90
>90
3000
31000
250
10500
27000
O
O
A
A
A
3
0.0
>90
1500
B
3
0.1
0-180 days
73000
O
4
4
5
0.0
0.1
0.0
>90
>90
>90
2000
3250
18500
O
O
O
5
5
5
0.1
0.1
ND
>90
>90
>90
10375
13875
0
A
A
O
8
0.1
>60
61500
B
8
9
17
23
0.2
0.1
0.0
0.0
0
>90
>90
>90
56000
12875
1500
5750
A
A
O
O
Type I inversion variant +
Asp1241Gln
mutation in exon 14
Deletion of exon 1 to 22
Type II inversion
unknown
unknown
c.6506G>A(p.Arg92150His)
mutation in exon 23
unknown
unknown
unknown
unknown
CGC>TGC;Arg1689Cys
mutation in exon 14
Val483Gly missens
mutation in exon 10
c.5338C>A(p.Pro1761Gln)
mutation in exon 15
unknown
unknown
Val483Gly missens
mutation in exon 10
unknown
unknown
558 C>A (ASP167 GLU)
mutation in exon 4
Tyr656Cys
mutation in exon 13
unknown
unknown
unknown
unknown
2
2
ND indicates not determined
Up-regulation of platelet activation in hemophilia A | 25
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B. Basal MFI
lth
y
H
ea
10
r -0.65
1
10 4.5
10 5.0
10 5.5
10 6.0
Mean Annual FVIII consumption - IU
M
ild
/M
od
er
a
ili
a
co
nt
r
A
A
0
ili
a
A
H
em
op
h
re
Se
ve
/M
od
er
at
e
M
il d
H
ea
H
em
op
hi
lia
co
nt
ro
l
0
A
5
5
H
em
op
h
10
re
15
10
100
Se
ve
20
ol
25
15
te
30
% P-selectin positive platelets
p < 0.001
p 0.054
p 0.010
Mean Fluorescence Intensity
35
lth
y
CD62P expressing platelets - %
p < 0.001
p 0.094
p 0.014
C. Correlation between P-selectin
expression and FVIII consumption
H
em
op
hi
lia
A. Basal percentage of platelet
activation
Figure 1. Baseline P-selectin expression
Citrated fresh whole blood was incubated for 20 minutes with PE labelled mouse anti-human
P-selectin antibodies, and then fixated with 0.2% formyl saline. (A) Percentage of P-selectin
expressing platelets and (B) mean fluorescence intensity of all platelets were determined with FACS
analysis. (C) Correlation between mean annual FVIII consumption (calculated over a 4 year period)
and percentage of P-selectin expressing platelets, r shows Spearman’s rank correlation coefficient.
(Spearman’s r -0.65; p=0.043) (Figure 1C). Correcting for patient’s weight does not influence
this correlation (corrected Spearman’s r -0.63; p=0.048).
To study the effect of FVIII infusion itself on platelet activation, we measured P-selectin
expression in 5 severe PWH before and 15 and 60 minutes after bolus infusion of FVIII to
peak levels of 1.0 U/l. No differences were found (data not shown).
Soluble platelet activation markers
Plasma concentrations of soluble platelet activation markers PF4, CXCL7 and RANTES were
significantly higher in both severe PWH (1.8 pg/106 platelets (1.4-2.6) PF4, 48.2 pg/106
platelets (35.6-103.7) CXCL7 and 9.5 pg/106 platelets (5.8-16.1) RANTES; p<0.001, p<0.001
and p=0.003) and mild-moderate PWH (1.4 pg/106 platelets (0.9-2.1) PF4, 65.8 pg/106
platelets (40.2-80.5) CXCL7 and 12.8 pg/106 platelets (9.5-21.2) RANTES; p<0.001, p<0.001
and p<0.001) when compared to plasma concentrations of healthy controls (0.3 pg/106
platelets (0.0-0.6) PF4, 20.0 pg/106 platelets (17.7-32.5) CXCL7 and 4.5 pg/106 platelets
(3.1-6.7) RANTES). For these markers no significant differences were found between mildmoderate and severe PWH (Figure 2A-C).
Plasma concentrations of soluble P-selectin did not differ between PWH and healthy
controls (Figure 2D).
26 | Chapter 2
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B. CXCL7
C. RANTES
p 0.381
1750
1500
1250
1000
750
500
250
20
10
A
H
H
em
op
h
ili
a
lia
hi
op
H
em
ea
lth
y
C
ili
a
A
0
A
A
a
H
em
op
h
ili
re
2
2
30
on
tr
ol
RANTES - pg/106 platelets
40
ild
M
A
Figure 2. Plasma concentrations of soluble platelet activation
markers
Citrated whole blood was centrifuged twice at 2000xG for
10 minutes at room temperature, and the resultant plasma was
collected and frozen at -80°C for evaluation. Plasma levels of
(A) platelet factor 4 (PF4), (B) human chemokine (C-X-C motif)
ligand 7 (CXCL7), (C) soluble P-selectin and (D) RANTES were
measured by enzyme-linked immunosorbent assay, and are
expressed per 106 platelets.
Se
ve
re
H
em
op
h
ili
a
A
H
em
op
hi
lia
50
M
ild
/M
od
er
at
e
lth
y
C
on
tr
ol
0
p 0.309
60
Se
ve
C
lth
y
/M
od
er
at
e
H
ea
M
ild
p 0.359
H
em
op
h
on
tr
ol
CXCL7 - pg/106 platelets
A
re
Se
ve
/M
od
er
at
e
p 0.129
soluble P-selectin - pg/106 platelets
550
500
450
400
350
300
250
200
150
100
50
0
ili
a
A
H
em
op
h
H
em
op
hi
lia
C
lth
y
ea
D. Soluble P-selectin
H
ea
p 0.003
p <0.001
p 0.861
/M
od
er
at
e
p <0.001
M
ild
H
p 0.218
9
8
7
6
5
4
3
2
1
0
on
tr
ol
PF4 - pg/106 platelets
p <0.001
p <0.001
re
p <0.001
Se
ve
A. Platelet factor 4
Von Willebrand Factor
Severe PWH had higher vWF plasma levels (14.9 μg/mL (10.1-19.3)) than healthy controls
(10.0 μg/mL (8.7-12.7); p=0.027). Mild-moderate PWH had similar levels of vWF (10.7 μg/
mL (8.7-13.6); p=0.538) as healthy controls (Figure 3). We observed equal vWF propeptide
levels for severe PWH (0.7 μg/mL (0.5-0.9)), compared to mild-moderate PWH (0.6 μg/
mL (0.5-0.8); p=0.559) and healthy controls (0.6 μg/mL (0.5-0.7); p=0.446). The vWF/vWF
propeptide ratio showed an increase in severe PWH (21.5 (17.4-23.0); p=0.014), but not in
mild-moderate PWH (17.9 (14.1-20.9); p=0.359) compared to healthy controls (17.2 (14.419.8)) (Figure 3).
Platelet responsiveness
Platelet responsiveness to ADP stimulation, represented by the EC50, of severe PWH (EC50
0.9 μM (0.4-0.9)) and mild-moderate PWH (EC50 0.7 μM (0.5-0.8)) did not differ from
healthy controls (EC50 0.6 μM (0.5-0.7); p=0.184 for severe and p=0.110 for mild-moderate
respectively). The percentages of activated platelets after maximal ADP stimulation in all
Up-regulation of platelet activation in hemophilia A | 27
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B. vWF propeptide
25
20
15
10
5
A
ili
a
A
te
/M
od
er
a
H
em
op
h
C
lth
y
ea
H
re
on
tr
ol
0
ili
a
A
H
em
op
h
re
Se
ve
C
/M
od
er
at
e
lth
y
ea
30
A
0.00
p 0.162
35
M
il d
0.25
ili
a
H
em
op
h
vWF / vWF propeptide ratio
0.50
A
A
a
Se
ve
re
H
em
op
hi
li
M
ild
/M
od
er
a
te
H
ea
lth
y
C
on
tr
ol
0
0.75
M
il d
10
1.00
on
tr
ol
vWF propeptide - ∝g/mL
20
1.25
H
vWF - ∝g/mL
30
p 0.014
p 0.359
p 0.559
H
em
op
hi
lia
p 0.505
p 0.141
H
em
op
hi
lia
p 0.538
C. vWF / vWF propeptide ratio
p 0.446
p 0.027
Se
ve
A. vWF
Figure 3. Plasma concentrations of vWF
Citrated whole blood was centrifuged twice at 2000xG for 10 minutes at room temperature, and
the resultant plasma was collected and frozen at -80°C for evaluation. Plasma levels (in μg/mL) of
(A) vWF antigen (vWF:Ag) and (B) vWF propeptide were measured by enzyme-linked immunosorbent
assay. The vWF:Ag / vWF propeptide ratio was determined (C).
PWH were comparable to healthy controls. Severe PWH patients had higher MFI after
maximal stimulation with ADP (MFI 112.1 (84.5-134.8)) when compared to mild-moderate
hemophilia patients (MFI 92.1 (75.5-107.7); p=0.063) and healthy controls (MFI 80.8 (74.7100.6); p=0.017) (Figure 4A-C).
The platelet responsiveness to CRP shows a large variability which is equivalent for
severe PWH (EC50 69.2 ng/mL (24.4-276.4)) compared to mild-moderate PWH (EC50
47.6 ng/mL (21.8-280.0); p=0.710) and healthy controls (EC50 65.9 ng/mL (26.8-250.7);
p=0.986). When maximally stimulated with CRP, both severe and mild-moderate PWH show
Figure 4. Platelet responsiveness
Citrated fresh whole blood was stimulated with concentration series of ADP and CRP, and incubated
for 20 minutes. For all concentrations, the percentage of platelets expressing P-selectin and the
mean fluorescence intensity of these platelets were measured.
The ADP and CRP concentrations generating a half maximal mean fluorescence intensity of P-selectin
expressing platelets were determined (A, D). The platelet response to the highest used ADP
concentration of 500 μM and the highest used CRP concentration of 2500 ng/mL, represented by
both the percentage of P-selectin expressing platelets (B,E) and the MFI of P-selectin expressing
platelets (C,F), were determined.
To determine platelet responsiveness to inhibition, platelets were inhibited with a concentration
series of iloprost in the presence of 5 μM ADP. After 20 minutes of incubation, the samples were
fixated with 0.2% formyl saline. The iloprost concentrations generating the half maximal inhibition
on mean fluorescence intensity of P-selectin expressing platelets were determined (G). The platelet
response to the highest used iloprost concentration of 250 ng/mL, represented by both the
percentage of P-selectin expressing platelets (H) and the MFI of P-selectin expressing platelets (I),
were determined.
28 | Chapter 2
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Bladel.indd 29
25
0
A
50
ili
a
75
H
em
op
h
p 0.184
p 0.141
p 0.038
A
H. Iloprost: miniimal percentage
a
Se
ve
re
H
em
op
h
ili
a
a
ili
A
A
M
il d
Se
ve
re
te
H
em
op
h
ili
a
A
A
l
A
A
co
nt
ro
ili
a
H
em
op
hi
lia
lth
y
H
em
op
h
H
ea
re
/M
od
er
a
Se
ve
Mean Fluorescence Intensity
p 0.468
p 0.195
p 0.873
Se
ve
re
60
H
em
op
h
p 0.007
p 0.014
p 0.710
H
em
op
hi
li
70
co
nt
ro
l
80
Mean Fluorescence Intensity
90
ild
/M
od
er
at
e
A
100
te
M
ili
a
A
E. CRP: maximal percentage
lth
y
H
em
op
h
60
H
ea
re
CD62P expressing platelets - %
co
nt
ro
l
A
H
em
op
hi
lia
lth
y
ili
a
A
70
co
nt
ro
l
Se
ve
M
ild
/M
od
er
at
e
H
ea
H
em
op
h
a
ADP - ∝M
80
Mean Fluorescence Intensity
A
A
l
re
ol
co
nt
r
H
em
op
hi
li
lth
y
90
/M
od
er
a
0
M
ild
5
100
lth
y
10
co
nt
ro
Se
ve
te
H
ea
B. ADP: maximal percentage
H
ea
15
ili
a
20
H
em
op
h
p < 0.001
p 0.929
p < 0.001
re
G. Iloprost: EC50
Se
ve
10 0
H
em
op
hi
lia
10 1
H
em
op
hi
lia
10 2
/M
od
er
at
e
10 3
CD62P expressing platelets - %
A
A
10 4
te
M
i ld
ili
a
lia
10 5
lth
y
H
em
op
h
hi
op
H
em
p 0.986
p 0.624
p 0.710
H
ea
re
te
ol
co
nt
r
D. CRP: EC50
co
nt
ro
l
Se
ve
/M
od
er
a
M
i ld
/M
od
er
a
0
CD62P expressing platelets - %
A
ild
lth
y
CRP - ng/mL
1
/M
od
er
a
ili
a
A
M
H
ea
2
lth
y
H
em
op
h
lia
l
Iloprost - ng/mL
3
H
ea
re
hi
co
nt
ro
op
lth
y
H
em
ea
p 0.184
p 0.901
p 0.110
M
ild
Se
ve
M
ild
/M
od
er
at
e
H
A. ADP: EC50
C. ADP: maximal MFI
p 0.017
p 0.399
p 0.063
200
150
100
50
2
2
0
F. CRP: maximal MFI
p 0.058
p 0.076
p 0.986
400
300
200
100
0
I. Iloprost: minimal MFI
p 0.228
p 0.521
p 0.235
45
35
25
15
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Up-regulation of platelet activation in hemophilia A | 29
1-11-2013 14:31:34
a lower percentage of P-selectin expressing platelets (95.0% (89.9-95.9) and 94.7% (92.497.3) respectively) with a lower amount of P-selectin on the surface of each platelet (MFI
280.1 (252.8-302.6) and 278.7 (191.1-319.0) respectively), compared to healthy controls
(96.3% (95.3-97.6); p=0.007 and 0.014 respectively; MFI 307.1 (283.1-341.8); p=0.058 and
0.076 respectively) (Figure 4D-F).
Platelets of either mild-moderate or severe PWH were more sensitive to inhibition
by iloprost (EC50 1.8 ng/mL (0.3-6.5) and 1.0 ng/mL (0.3-3.9)) when compared to healthy
controls (EC50 8.3 ng/mL (6.0-11.1); p<0.001 and <0.001 respectively). However, the
maximal inhibition by iloprost of mild-moderate (26.0% (21.4-34.3); MFI 25.2 (22.7-29.1))
and severe PWH (34.0% (27.9-42.3); MFI 34.0 (27.9-42.3)) was comparable to healthy
controls (28.3% (25.0-36.5); p=0.141 and 0.184; MFI 25.6 (24.4-28.8); p=0.521 and 0.228
respectively) (Figure 4G-I).
Discussion
We have shown that PWH have a higher basal level of activated platelets in their circulation,
have a higher platelet response to ADP stimulation and were more sensitive to inhibition
with the prostacyclin analogue iloprost than healthy controls. Furthermore, PWH had
higher plasma concentrations of platelet activation markers PF4, CXCL7 and RANTES. Also
vWF concentrations were raised in severe hemophilia A. Within the severe PWH, platelet
activation correlates to FVIII consumption. These data indicate that platelet activation is
up-regulated in the presence of a deficiency in secondary hemostasis, reducing the risk of
bleeding complications.
Comparing baseline levels of platelet activation in PWH with healthy controls, we found
that both platelet P-selectin expression and plasma concentrations of the soluble platelet
activation markers PF4, CXCL7 and RANTES are increased in PWH. Theoretically, this
increased platelet activation could partially compensate a deficiency in FVIII, by providing
more negatively charged surface to enhance the constrained coagulation cascade in PWH.
In line with this theory one would expect platelet activation showing the largest increase
in the more severely affected patients. In our study hemophilia severity was classified
based upon residual FVIII activity. Comparing platelet activation in mild-moderate and
severe PWH, we indeed found a higher baseline platelet P-selectin expression in the more
severe patients (residual FVIII<1%). However, for the soluble platelet activation markers
PF4, CXCL7 and RANTES, no significant differences were found between mild-moderate
and severe PWH.
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In an additional analysis within the severe PWH, we classified hemophilia severity
based upon annual FVIII consumption. Again, platelets P-selectin expression showed to
correlate to clinical phenotype. However, to fully characterize clinical phenotype, in future
studies this should not be scored solely on FVIII consumption, but also on age of first joint
bleed, joint bleeding frequency and joint damage. To exclude FVIII infusion as a cause of
up-regulation of platelet P-selectin expression, we have measured 5 severe PWH before
and 15 and 60 minutes after infusion. There was no effect of FVIII infusion on basal MFI of
P-selectin on platelets.
Platelet responsiveness was triggered with increasing concentrations of ADP and CRP
and activation was measured with P-selectin expression. The platelet responsiveness
to both ADP and CRP did not differ in PWH when compared to controls. However, the
maximal response to stimulation was different in PWH. Upon maximal stimulation with
ADP, an agonist which by itself can only initiate partial platelet activation, the platelets of
PWH expressed more P-selectin on the cell surface than healthy controls. However, upon
maximal stimulation with CRP a smaller percentage of platelets became activated in PWH.
Whether this lower response to CRP can lead to an aggravation of bleeding phenotype in
individual patients, remains to be determined in future studies.
Platelets of PWH were more responsive to inhibition by iloprost as shown by the low
EC50 of iloprost in PWH after suboptimal stimulation by ADP, when compared to controls.
In PWH, which we have shown to have a higher baseline platelet activation and a higher
maximal response to stimulation by ADP, this increased inhibitory mechanism could have
developed to prevent platelet activation in the absence of vascular injury.
To our knowledge, this study provides for the first time evidence that in patients,
deficiency in secondary hemostasis is associated with increased platelet activation and
responsiveness. A platelet responsiveness test was used that covers the complete range
of the platelet response, from the (ant-)agonist concentration yielding no response, to
concentrations yielding the maximal platelet response to the used (ant-)agonist. This total
range of (ant-)agonist concentrations allowed for analysis of the concentration yielding a
half maximal response together with the maximal effect of different (ant-)agonists, providing
new parameters of platelet responsiveness. The platelet responsiveness was measured for
established (ant-)agonists representing established haemostatic pathways in vivo.
Our findings of increased vWF concentration in severe PWH supports previous findings
of Vlot et al10 and Fijnvandraat et al11 who also found higher vWF:Ag levels in severe PWH.
Increased vWF:Ag levels due to infection and stress in PWH has been suggested as rationale
for this finding. The results did not change when bloodgroup analysis was included (data not
shown). The absence of a relationship between vWF propeptide and hemophilia A suggests
2
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that the raise in vWF plasma concentration reflects chronic endothelial cell activation
rather than acute infection or stress. Chronic inflammation in joints or chronic hepatitis C,
may be an explanation for this observation. The origin of the increased vWF in plasma of
severe PWH may be secretion by platelets as well as endothelial cells. This chronic raise in
vWF plasma concentration could be an additional factor of the primary hemostatic system
compensating for the deficiency in secondary hemostasis present in PWH.
For this study, patients were selected during regular visits at the Van Creveldkliniek. The
Van Creveldkliniek is the largest hemophilia clinic in The Netherlands, treating approximately
800 PWH. Patients are treated according to bleeding phenotype, resulting in individual
tailored prophylaxis regimens in the severe patients or on demand treatment in the
patients with a milder phenotype. The populations of the current study are representative
for any other hemophilia populations in western countries, in which treatment is tailored
to bleeding phenotype.
The present study has potential limitations that should be addressed. First, to assure
sufficient power for a reliable comparison, mild and moderate PWH were grouped. Second,
our study is a cross-sectional analysis and clinical phenotype in the severe PWH was
only based on retrospectively collected annual FVIII consumption. Third, in this study we
decided to only include adult PWH. The choice of a younger population would probably
have given additional insight into the origin, innate or acquired, of up-regulation of platelet
activation and responsiveness. Last, blood sampling and platelet activation and reactivity
measurements were performed at a single time point for each patient. Although repeated
measurements were not performed in patients, we don’t expect large variability in results
over time. Repeated measurements in healthy donors show equivalent results over time
(data not shown).
In summary, this study reveals a higher basal level of platelet activation, shown by
higher P-selectin expression on the platelet surface and higher plasma concentrations
of soluble platelet activation markers, and an increased platelet response to stimulation
by ADP and to inhibition by iloprost in PWH. Platelet P-selectin expression showed to be
inversely correlated to FVIII consumption in severe PWH. Furthermore a chronic increase in
vWF plasma concentration is present in severe PWH. This study introduces a novel insight
in hemostasis in hemophilia A, in which platelets are in a pre-activated state, and this preactivated state is associated with patient FVIII consumption.
32 | Chapter 2
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References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
den Uijl IEM, Fischer K, Van Der Bom JG, Grobbee DE, Rosendaal FR, Plug I. Clinical outcome
of moderate haemophilia compared with severe and mild haemophilia. Haemophilia.
2009;15(1):83-90.
Biggs R, Macfarlane RG. Haemophilia and related conditions: a survey of 187 cases. Br J
Haematol. 1958;4(1):1-27.
White GC, Rosendaal F, Aledort LM, Lusher JM, Rothschild C, Ingerslev J. Definitions in
hemophilia. Recommendation of the scientific subcommittee on factor VIII and factor IX of
the scientific and standardization committee of the International Society on Thrombosis and
Haemostasis. Thromb Haemost. 2001;85(3):560.
Aledort LM, Haschmeyer RH, Pettersson H. A longitudinal study of orthopaedic outcomes for
severe factor-VIII-deficient haemophiliacs. The Orthopaedic Outcome Study Group. J Intern
Med. 1994;236(4):391-9.
Molho P, Rolland N, Lebrun T, Dirat G, Courpied JP, Croughs T, Duprat I, Sultan Y. Epidemiological
survey of the orthopaedic status of severe haemophilia A and B patients in France. The French
Study Group. Haemophilia. 2000;6(1):23-32.
van Dijk K, van der Bom JG, Fischer K, de Groot PG, van den Berg HM. Phenotype of
severe hemophilia A and plasma levels of risk factors for thrombosis. J Thromb Haemost.
2007;5(5):1062-4.
van Dijk K, van der Bom JG, Lenting PJ, de Groot PG, Mauser-Buschoten EP, Roosendaal G,
Grobbee DE, van den Berg HM. Factor VIII half-life and clinical phenotype of severe hemophilia
A. Haematologica. 2005;90(4):494-8.
Furie B, Furie BC. Mechanisms of thrombus formation. N Engl J Med. 2008;359(9):938-49.
Borchiellini A, Fijnvandraat K, ten Cate JW, Pajkrt D, van Deventer SJH, Pasterkamp G, MeijerHuizinga F, Zwart-Huinink L, Voorberg J, van Mourik JA. Quantitative analysis of von Willebrand
factor propeptide release in vivo: effect of experimental endotoxemia and administration of
1-deamino-8-D-arginine vasopressin in humans. Blood. 1996;88(8):2951-8.
Vlot AJ, Mauser-Bunschoten EP, Zarkova AG, Haan E, Kruitwagen CLJJ, Sixma JJ, van den Berg
HM. The half-life of infused factor VIII is shorter in hemophiliac patients with blood group O
than in those with blood group A. Thromb Haemost. 2000;83(1):65-9.
Fijnvandraat K, Peters M, ten Cate JW. Inter-individual variation in half-life of infused
recombinant factor VIII is related to pre-infusion von Willebrand factor antigen levels. Br J
Haematol. 1995;91(2):474-6.
2
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Chapter 3
Factor VIII concentrate infusion in patients with hemophilia
results in decreased von Willebrand Factor and ADAMTS-13 activity
Esther R. van Bladel,1,2* Attie Tuinenburg,3* Mark Roest,1,2 Philip G. de Groot1
and Roger E.G. Schutgens2,3
1
Department of Clinical Chemistry and Hematology, University Medical Center
Utrecht, Utrecht, the Netherlands; 2Van Creveld Laboratory, University Medical
Center Utrecht, Utrecht, the Netherlands; and 3Van Creveldkliniek/Department of
Hematology, University Medical Center Utrecht, Utrecht, the Netherlands
*equally contributed
Accepted for publication in Haemophilia.
Bladel.indd 35
1-11-2013 14:31:35
Abstract
The effects of coagulation factor concentrate infusion on restoring secondary hemostasis
in patients with hemophilia are obvious. It is not known whether coagulation factor
concentrate infusion affects primary hemostasis or induces an acute inflammatory
response. In the present study, the influence of a FVIII concentrate bolus infusion on
platelet activation and responsiveness, endothelial activation, and inflammation in adult
patients with severe hemophilia A was assessed. vWF showed a mild, but significant
decrease 15 minutes after FVIII infusion (85.02 IU/dL) vs. before infusion (92.04 IU/dL;
p=0.017), while ADAMTS-13 levels also show a mild but significant decrease from 66.1 ng/
mL before infusion, to 53.9 ng/mL (p=0.012) 15 minutes after and 50.8 ng/mL (p=0.050)
60 minutes after infusion. Platelet P-selectin expression decreased 15 minutes (33.3 AU)
and 60 minutes (38.7 AU) after infusion compared to before infusion (41.3 AU; p=0.018
and =0.036). In conclusion, a single infusion of a high dose FVIII concentrate in hemophilia
A patients may influence primary hemostasis by decreasing vWF, ADAMTS-13 and the
number of circulating activated platelets. These effects possibly occur as a consequence
of binding of the infused FVIII to vWF, influencing its processing. When treating severe
hemophilia A patients with coagulation concentrate infusion, one should realise this does
not merely correct FVIII levels, but also may influence primary hemostasis.
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Introduction
Hemophilia A and B are hereditary X chromosomal recessive bleeding disorders caused by
a deficiency or a functional defect in coagulation factor VIII (FVIII) or IX (FIX), respectively.
FVIII and FIX are (co)factors essential for optimal thrombin generation and propagation of
fibrin formation.1 Severity of bleeding symptoms generally depends on the residual plasma
concentration of the deficient coagulation factor.2,3 Bleeding episodes are prevented or
controlled by substitution of the missing coagulation factor by intravenous infusion of FVIII
or FIX.1
Several cohort studies have reported a reduced mortality due to ischemic heart disease
in hemophilia patients as compared to the general male population,4,5 as well as a reduced
prevalence of non-fatal cardiovascular events.6 As coagulation plays a role in atherogenesis
via inflammation,7 one could hypothesize that a congenital coagulation factor deficiency
has a protective effect on the development of atherosclerosis.8 Recent studies have
demonstrated that patients with hemophilia develop atherosclerosis in an equal matter
as the normal population.9,10 However, as nowadays the majority of patients with severe
hemophilia A are prophylactically treated with coagulation factor concentrate, this
continuous partial correction may dilute the possible protective effect of FVIII deficiency on
atherosclerosis. As atherosclerosis is considered an inflammatory disease, it is of interest
to know whether a single infusion of FVIII concentrate provokes an inflammatory response.
Furthermore, although the effects of coagulation factor concentrate infusion on
coagulation are clear, it is not known whether coagulation factor concentrate infusion also
affects primary hemostasis. It has been suggested that platelets might be able to modify
procoagulant activity in hemophilia.11,12 Whether platelets are triggered to a more active
state by infusion of FVIII concentrate is unknown. Furthermore, infused FVIII will bind to
von Willebrand factor (vWF), a plasma protein also essential for optimal platelet function.
It could be that this binding has an effect on the function of vWF as well.
In this study we assessed the influence of a single FVIII concentrate bolus infusion
on platelet activation and responsiveness, endothelial activation, and inflammation
parameters in patients with severe hemophilia A.
3
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Materials and methods
Study design and study population
Male patients, ≥18 years, with severe hemophilia A (FVIII activity <1%), under treatment at
the Van Creveldkliniek (University Medical Center Utrecht, the Netherlands), and scheduled
for elective surgery were eligible for this study. Patients were excluded when having a
history of symptomatic cardiovascular disease (i.e. myocardial infarction, percutaneous
coronary intervention, coronary artery bypass graft surgery, atrial fibrillation, or stroke).
The reason to investigate patients going for elective surgery was due to the fact that these
patients have an elective bolus infusion leading to rapid and full correction of FVIII levels
without a potentially interfering active bleed.
As a standard pre-operative procedure to increase FVIII activity before surgery, a
bolus of 50 U/kg of FVIII was administered over approximately 5 minutes. Before and
15 minutes after the bolus infusion, blood was collected to measure FVIII activity. At these
time points and with an additional venipuncture 60 minutes post infusion, we collected
additional blood to measure platelet activation and responsiveness, endothelial activation
and inflammation. Continuous FVIII concentrate infusion to prepare the patient for surgery
was started after the last venipuncture. There was no wash-out period before inclusion in
the study. Therefore, basal FVIII levels could be higher than 1% due to regular prophylaxis.
Participants were included between July 2009 and February 2012. This study was
approved by the Medical Ethics Committee of the University Medical Center Utrecht, the
Netherlands. All participants provided written informed consent.
Blood collection
Antecubital venipuncture was performed using 21G needles. Prior to FVIII concentrate
infusion, and 15 and 60 minutes post infusion, blood was collected into Vacutainer® tubes,
containing sodium-citrate (3.2%) or EDTA. After blood collection, platelet number and
mean platelet volume (MPV) were immediately measured, using the Cell-dyn 1800 (Abbott
Laboratories, USA), in EDTA-anticoagulated blood. In addition, samples for fluorescenceactivated cell sorting (FACS) analysis were prepared immediately after the blood,
anticoagulated with sodium-citrate, was drawn. The remaining blood was centrifuged
twice for 10 minutes (2000xG, room temperature) to obtain platelet poor plasma. Plasma
samples were stored at -80°C until use.
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Materials
For enzyme-linked immune sorbent assays (ELISAs), antibodies against human Platelet
Factor 4 (PF4; MAB7951, AF795), human chemokine (C-X-C motif) ligand 7 (CXCL7) (MAB393,
BAF393), human chemokine (C-C motif) ligand 5 (RANTES) (MAB278, AB-278-NA), human
soluble P-selectin (DY137 Duoset), soluble intercellular adhesion molecule (sICAM) (DY720
Duoset), a disintegrin and metalloproteinase domain with thrombospondin modules
13 (ADAMTS-13) (AF4245, BAF4245), osteoprotegerin (OPG) (MAB8051, BAF805) and
C-reactive Protein (DY1707 Duoset) were purchased from R&D Systems (Abingdon, U.K.).
Rabbit anti-human von Willebrand Factor (vWF) antibodies (A0082), peroxidase conjugated
rabbit anti-human vWF (P0226), rabbit anti-goat horseradish peroxidase (HRP) (P0449) and
streptavidin-poly-HRP (P0397) were purchased from DAKO (Glostrup, Denmark). Rabbit
anti-vWF propeptide and rabbit anti-vWF propeptide/biotine were prepared as described
by Borchiellini et al.13 A11 nanobody, used as coating antibody in the active vWF ELISA, was
prepared as described by Hulstein et al.14 A recombinant vWF, with a R1306Q mutation,
was used as the calibration curve to determine the amount of vWF in its GPIb binding
conformation (active vWF).15 SuperSignal ELISA Pico chemiluminescent substrate was
purchased from Thermo Scientific (Rockford, Illinois, U.S.A.).
R-phycoerythrin (PE) labeled antibodies for FACS analysis, raised against human
P-selectin (#555524), were purchased from BD Biosciences (Franklin Lakes, NJ, U.S.A.). For
platelet responsiveness assays, adenosine diphosphate (ADP) was purchased from Roche
(Almere, the Netherlands), thrombin receptor activator peptide (TRAP) from Bachem AG
(Bubendorf, Switzerland), iloprost (ilomedine) from Bayer Schering Pharma AG (Berlin,
Germany) and Cross-linked Collagen Related Peptide (XL-CRP) was a generous gift of
R. Farndale (Cambridge, U.K.).
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) was purchased from BDH
(Poole, U.K.). Sodium chloride (NaCl), Tween-20 and Bovine serum albumin (BSA) were
purchased from Sigma (Zwijndrecht, the Netherlands). Magnesium Sulphate (MgSO4) and
Potassium Chloride (KCl) were purchased from Riedel (Seelze, Germany), and Formaldehyde
(CH2O) from Calbiochem (Merck, Darmstadt, Germany).
3
ELISA procedure
Platelets release their granule contents as soluble proteins into the circulation upon
activation. The most abundant chemokines in the α-granules are CXCL7 and PF4.16 Other
plasma markers of platelet activation are RANTES and soluble P-selectin. Upon endothelial
cell activation, vWF and OPG are released from the Weibel-Palade bodies. sICAM is also
released by endothelium in states of activation. As markers of an inflammatory response
Factor VIII concentrate infusion in patients with hemophilia | 39
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we measured high sensitivity C-reactive protein (hs-CRP), tumor necrosis factor alpha
(TNF-α), interleukin 6 (IL-6) and interleukin 8 (IL-8).
Plasma levels of soluble platelet activation markers CXCL7 and soluble P-selectin, of
ADAMTS-13, OPG and of hs-CRP were measured by semi-automated ELISA on a TECAN
Freedom Evo robot (Tecan, Mannedorf, Switzerland) as described previously with some
minor modifications.11,17 Plasma levels of PF4, RANTES, vWF, active vWF and vWF propeptide
were determined by non-automated ELISA, following the same procedure as when measured
semi-automated. TNF-α, IL-8 and IL-6 were measured using commercially available ELISAs
according to the instructions of the manufacturer (TNF-α and IL-8, R&D Systems, Abingdon,
UK; IL-6, BD Biosciences, San Diego, CA, U.S.A.). Analysis of vWF multimeric pattern was
done using 2% agarose gel electrophoresis, followed by immunoblotting as described by
Raines and colleagues.18
Platelet activation and reactivity
Platelet responsiveness to agonists was determined with concentration series of ADP
(0.01 μM, 0.03 μM, 0.12 μM, 0.49 μM, 1.95 μM, 7.81 μM, 31.25 μM and 125.00 μM), XLCRP (0.2 ng/ml, 0.6 ng/ml, 2.4 ng/ml, 9.8 ng/ml, 39.1 ng/ml, 156.3 ng/ml, 625.0 ng/ml
and 2500.0 ng/ml) and TRAP (0.04 μM, 0.15 μM, 0.61 μM, 2.44 μM, 9.77 μM, 39.06 μM,
156.25 μM and 625.00 μM), while platelet responsiveness to inhibitors was measured with
concentration series of TRAP (0.04 μM, 0.15 μM, 0.61 μM, 2.44 μM, 9.77 μM, 39.06 μM,
156.25 μM and 625.00 μM) containing 5 ng/ml iloprost and with concentration series of
iloprost (0.02 ng/ml, 0.08 ng/ml, 0.31 ng/ml, 1.22 ng/ml, 4.88 ng/ml, 19.53 ng/ml, 78.13 ng/
ml and 312.50 ng/ml) containing 5 μM ADP. Serial dilutions were prepared in 50 μL HEPES
buffered saline (HBS; 10 mM HEPES, 150 mM NaCl, 1 mM MgSO4, 5 mM KCl, pH 7.4, filtered
through a 0.22 μm filter) containing 2 μL PE labeled mouse anti-human P-selectin antibodies.
A negative control sample, only containing 50 μL HBS with 2 μL PE labeled mouse
anti-human P-selectin antibodies, was prepared to determine the basal level of platelet
activation.
The platelet activation test was initiated by addition of 5 μL fresh, citrate-anticoagulated,
whole blood to each mixture of the serial dilutions. After 20 minutes of incubation, the
samples were fixed with 500 μL 0.2% formylsaline (0.2% formaldehyde in 0.9% NaCl, filtered
through a 0.22 μm filter) and after 10 minutes of fixation kept at 4°C until analyses. Samples
were analyzed on a FACSCalibur flow cytometer from BD Biosciences (Franklin Lakes, NJ,
U.S.A.) or a FACS Canto II flow cytometer from BD Biosciences (San Jose, CA, U.S.A.) within
one day after processing. Single platelets were gated based on forward and side scatter
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properties. The mean fluorescence intensity (MFI) in the platelet gate was measured with
FACS analysis for P-selectin signal. One individual performed all assays.
Statistical analyses
FACS data were quantified with BD FACSDiva software 6.1.2 (BD Biosciences, San Jose, CA,
U.S.A.). To determine platelet responsiveness, the concentration generating a response
halfway between baseline and maximum MFI signal (EC50) was calculated using GraphPad
Prism version 5.03 (GraphPad Software, San Diego, CA, U.S.A.). The maximal effect of
stimulation or inhibition is represented by the MFI in the sample with the highest used
(ant-)agonist concentration.
Wilcoxon signed ranks tests were performed to analyze the data statistically. Data were
analyzed using IBM SPSS Statistics 20.0.0 for Windows (International Business Machines
Corporation, New York, U.S.A.). Data are shown as median with interquartile range unless
stated otherwise.
3
Results
Patient characteristics
Eight patients with severe hemophilia A were included. Characteristics of the study
population are described in Table 1. Mean age of the study population was 40 years
(standard deviation 15). Age ranged from 21-59 years. Two (25%) patients were infected
with hepatitis C and 1 (12.5%) patient was infected with HIV. Three patients (37.5%) received
plasma purified FVIII; the other 5 patients (62.5%) received recombinant FVIII. None of the
factor VIII preparations contain von Willebrand factor. Fifteen minutes after infusion of
50 U/kg of FVIII concentrate, median FVIII activity increased from 0.7IU/dL (0.0-3.9) to
112.5IU/dL (96.5-133.3).
Endothelium
Infusion of FVIII concentrate does not seem to influence endothelial cells (Table 2). No
changes were seen in sICAM levels, a protein released by endothelial cells upon activation.
Additionally, the plasma levels of OPG and vWF propeptide, both localized in Weibel-Palade
bodies, did not change after the infusion of FVIII concentrate. vWF showed a mild, but
significant decrease 15 minutes after infusion (85.02 IU/dL (63.75-152.74)) versus before
infusion (92.04 IU/dL (71.46-164.13); p=0.017). However, after 60 minutes vWF levels
had already returned to original levels. vWF was not activated following infusion of FVIII
Factor VIII concentrate infusion in patients with hemophilia | 41
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Table 1. Characteristics of the study population
Variable
Hemophilia patients
(n=8)
Age (years)
FVIII activity before infusion of FVIII bolus
FVIII activity 15 min. after infusion of FVIII bolus
Plasma derived FVIII
Recombinant FVIII
Chronic hepatitis C infection
HIV infection
40 ± 15
0.7 (0.0-3.9)
112.5 (96.5-133.3)
3 (37.5)
5 (62.5)
2 (25.0)
1 (12.5)
Values are expressed as mean ± standard deviation or median (interquartile range). Categorical
variables are expressed as numbers (percentages).
FVIII, factor VIII; min., minutes; HIV, human immunodeficiency virus; n, sample size.
as measured by the amount of vWF in its GPIb binding conformation. At the same time,
ADAMTS-13 levels also show a mild but significant decrease from 66.12 ng/mL (50.44-97.42)
before infusion, to 53.95 ng/mL (36.36-58.77; p=0.012) 15 minutes after and 50.88 ng/mL
(43.03-78.27; p=0.050) 60 minutes after infusion. No changes in multimeric patterns of vWF
were observed.
Platelets
Basal level of platelet activation was determined by measuring both platelet P-selectin
expression in whole blood immediately after collection, and by measuring the levels
of soluble platelet activation markers in platelet poor plasma. A decrease in circulating
activated platelets, measured as percentage of platelets with P-selectin expression, was
observed 15 (3.57% (2.15-9.29)) and 60 minutes (3.08% (1.57-10.68)) after FVIII infusion
(before infusion 4.87% (2.55-20.89); p=0.012 and =0.017). Accordingly, the mean amount
of P-selectin expressed on platelets decreased 15 minutes (33.3 arbitrary units (AU) (8.5192.4)) and 60 minutes (38.7 AU (4.9-209.2)) after infusion compared to before infusion
(41.3 AU (9.5-342.0); p=0.018 and =0.036). Similarly levels of CXCL7, a protein secreted by
activated platelets, showed a decrease 15 minutes (19.3 ng/mL (12.0-25.6)) and 60 minutes
(13.1 ng/mL (6.7-23.1)) after infusion of FVIII (23.1 ng/mL (15.1-37.2); p=0.036 and =0.012).
Levels of PF4 (CXCL4), another protein released from platelets when activated, showed a
decrease 60 minutes after infusion (8.6 ng/mL (6.1-16.5)) compared to before infusion of
FVIII (18.0 ng/mL (12.9-24.1); p=0.017).
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Bladel.indd 43
203.23 (180.31-302.98)
85.02 (63.75-152.74)
6.57 (5.36-7.72)
17.26 (15.25-26.18)
1.23 (0.78-1.48)
53.95 (36.36-58.77)
0.62 (0.43-0.89)
4.30 (2.46-6.22)
15 min. post infusion
n=8
236.05 (188.83-311.02)
88.38 (55.68-154.98)
5.97 (5.47-7.33)
19.94 (13.04-28.02)
1.08 (0.69-1.44)
50.88 (43.03-78.27)
0.64 (0.50-1.02)
4.09 (2.44-7.41)
60 min. post infusion
n=8
ns
0.017
ns
ns
ns
0.012
ns
ns
Before vs. 15 min.
post
p-value
ns
ns
ns
ns
0.049
ns
ns
ns
Before vs. 60 min.
post
Values are expressed as median (interquartile range). Differences are assessed with the Wilcoxon signed ranks test.
Number in hs-CRP measurement deviates: before infusion n = 6; 15 min. post infusion n = 4; 60 min. post infusion n = 4.
sICAM, soluble intercellular adhesion molecule; vWF, von Willebrand Factor; ADAMTS-13, a disintegrin and metalloproteinase domain, with
thrombospondin modules 13 ; OPG, osteoprotegerin; hs-CRP, high sensitivity C-reactive protein; n, sample size; min, minutes; vs., versus; ns, not
significant.
229.92 (181.88-342.77)
92.04 (71.46-164.13)
6.78 (5.41-8.02)
20.02 (16.84-26.41)
1.11 (0.71-1.57)
66.12 (50.44-97.42)
0.84 (0.51-1.01)
3.75 (2.61-5.76)
sICAM (ng/mL)
vWF (IU/dL)
vWF propeptide (nM)
vWF / vWF propeptide ratio
Active vWF (μg/mL)
ADAMTS-13 (ng/mL)
OPG (ng/mL)
hs-CRP (µg/mL)&
&
Before infusion
n=8
Variable
Table 2. Endothelium and inflammation
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44 | Chapter 3
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41 (16-448)
1.44 (0.10-7.92)
50 (19-598)
0.96 (0.09-6.93)
42 (13-455)
1.20 (0.10-6.60)
1789 (347-3075)
0.76 (0.44-0.96)
0.018
ns
ns
ns
0.018
0.036
ns
ns
ns
ns
ns
ns
ns
0.036
0.012
0.017
ns
ns
0.017
8432 (6532-10384)
1.92 (1.68-1.98)
8573 (6617-10723)
1.93 (1.68-2.03)
ns
ns
ns
ns
1771 (326-3124)
0.56 (0.40-0.93)
1787 (348-3494)
0.66 (0.48-0.88)
38.7 (4.9- 209.2)
13.1 (6.7-23.1)
8.6 (6.1-16.5)
1.3 (0.9-2.4)
45.8 (43.0-63.1)
0.012
ns
ns
8996 (6675-10501)
1.96 (1.78-2.02)
33.3 (8.5-192.4)
19.3 (12.0-25.6)
15.4 (10.5-21.8)
1.9 (1.2-2.6)
58.3 (49.5-65.9)
41.3 (9.5-342.0)
23.1 (15.1-37.2)
18.0 (12.9-24.1)
1.4 (0.9-2.8)
53.3 (43.0-66.8)
3.08 (1.57-10.68)
ns
ns
ns
ns
3.57 (2.15-9.29)
4.87 (2.55-20.89)
263 (223-321)
8.2 (7.9-8.6)
8055 (1612-11329)
8088 (1603-10751)
8104 (1609-11352)
ns
104.23 (36.46-260.80) 117.85 (39.17-344.93) 138.00 (39.06-234.58) ns
256 (220-301)
8.2 (7.5-8.9)
274 (222-315)
8.2 (7.7-9.4)
Platelet count (*109/L)
Mean platelet volume (fL)
Basal platelet activation
Circulating activated platelets (% of
total platelets)
Basal P-selectin expression (MFI - AU)
CXCL7 (ng/mL)
PF4 (ng/mL)
RANTES (ng/mL)
Soluble P-selectin (ng/mL)
ADP
Maximal stimulation (MFI - AU)
EC50 of MFI curve (μM)
Iloprost (+ADP)
Maximal inhibition (MFI - AU)
EC50 of MFI curve (ng/mL)
XL-CRP
Maximal stimulation (MFI - AU)
EC50 of MFI curve (ng/mL)
TRAP&
Maximal stimulation (MFI - AU)
EC50 of MFI curve (μM)
15 min. post infusion 60 min. post infusion p-value
n=8
n=8
Before
Before
vs. 15 min. post vs. 60 min. post
Before infusion
n=8
Variable
Table 3. Platelets
Bladel.indd 45
7497 (6060-9963)
8.90 (2.23-10.24)
Before infusion
n=8
7788 (6023-9834)
8.30 (2.11-9.63)
7836 (5806-9982)
7.92 (2.25-9.47)
ns
0.028
ns
ns
15 min. post infusion 60 min. post infusion p-value
n=8
n=8
Before
Before
vs. 15 min. post vs. 60 min. post
&
Values are expressed as median (interquartile range). Differences are assessed with the Wilcoxon signed ranks test.
TRAP and TRAP + iloprost measurements were performed in 6 patients.
MFI, Mean Fluorescence Intensity of all platelets; AU, arbitrary units; CXCL7, human chemokine (C-X-C motif) ligand 7; PF4, Platelet Factor 4; RANTES,
human chemokine (C-C motif) ligand 5; ADP, adenosine diphosphate; EC50, concentration generating a response halfway between baseline and
maximal MFI signal ; XL-CRP, Cross-linked Collagen Related Peptide; TRAP, thrombin receptor activator peptide; n, sample size; min, minutes; vs.,
versus; ns, not significant
TRAP (+iloprost) &
Maximal stimulation (MFI - AU)
EC50 of MFI curve (μM)
Variable
Table 3. Continued
3
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Platelet reactivity to stimulation with ADP, XL-CRP or TRAP did not alter after infusion of
FVIII. Platelet reactivity to inhibition with iloprost gave inconsistent results. The MFI reached
after maximal inhibition by iloprost when platelets were activated with a suboptimal ADP
concentration, was lower 15 minutes (41 AU (16-448)) after infusion compared to before
infusion (50 AU (19-598); p=0.018). On the contrary, in the presence of iloprost, less TRAP
was needed to half maximally activate platelets 15 minutes after infusion (8.30 μM (2.119.63)) compared to before infusion (8.90 μM (2.23-10.24); p=0.028).
Platelet count, showed a mild, but non-significant, decrease 15 minutes after infusion
(256*109/L (220-301)), compared to before infusion (274*109/L (222-315); p=0.261). Infusion
of FVIII concentrate did not alter MPV. Results on all platelet parameters are shown in
table 3.
Inflammation
Before infusion of FVIII concentrate, no signs of inflammation were found, with levels of
hs-CRP reflecting a non-inflammatory state (3.75 μg/mL (2.61-5.76)) and levels of TNFα, IL-6
and IL-8 below detection limits of 15 pg/mL, 10 pg/mL and 30 pg/mL, respectively. Infusion
of FVIII concentrate did not change the inflammatory status of patients 15 and 60 minutes
after infusion. hs-CRP levels stayed equal (4.30 μg/mL (2.46-6.22) and 4.09 μg/mL (2.447.41) at 15 and 60 minutes, respectively). Results are shown in Table 2. TNFα, IL-6 and IL-8
remained below detection limits.
Discussion
In this study we investigated whether infusion of FVIII concentrate influences primary
hemostasis and elicited an acute inflammatory response. After infusion of FVIII concentrate,
we observed a decrease in vWF, together with a decrease in ADAMTS-13 and the number of
circulating activated platelets. No effects on inflammation were observed.
Our data show that infusion of FVIII into hemophilia A patients leads to decreased
circulating vWF levels in vivo. This decrease could point to either a reduced vWF release
from endothelial cells or platelets or increased clearance of vWF from the circulation. In
view of the unchanged levels of vWF propeptide, OPG and sICAM, which are also released
by endothelial cells upon activation, a decrease in release seems unlikely. ADAMTS-13
levels also showed a significant decrease. Although ADAMTS-13 is reported to be produced
by endothelial cells,19,20 it is also known to be produced by hepatic stellate cells and
megakaryocytes,21-23 and its levels therefore do not merely reflect the level of endothelial
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cell activation. However, the decrease in ADAMTS-13 could be an indication of a higher
consumption of ADAMTS-13, corresponding to an increased clearance of vWF. Cao and
colleagues24 showed that FVIII enhances the proteolytic cleavage of vWF, preferentially the
high molecular weight vWF multimers, by ADAMTS-13 under shear stress. In a subsequent
study, it was shown that this effect of FVIII is synergized by platelets.25 The results of their
in vitro observations are in line with our in vivo findings.
If FVIII is involved in the processing of vWF, one would expect higher levels of vWF in
severe hemophilia A patients compared to mild-moderate patients and healthy controls.
Indeed, several studies in the past have shown increased vWF levels in hemophilia A.11,26,27
Alternative explanations proposed for this finding include chronic inflammation and stress,
but to our knowledge no evidence was found to support these explanations.
In vitro studies have shown that platelet binding to vWF accelerates its cleaving by
ADAMTS-13,28 and that this platelet action is synergistically amplified by FVIII.25 In these
studies the effects of FVIII and platelets on vWF clearance was studied, however, effects of
vWF in the presence or absence of FVIII on platelets were not considered. In our study, we
observed a decrease in the number of circulating activated platelets after infusion of FVIII.
This was found for both the number of P-selectin expressing platelets and the plasma levels
of platelet activation markers PF4 and CXCL7. Also, we observed a mild decrease in total
platelet number after FVIII infusion; however, this was not statistically significant. Possibly,
binding of FVIII to vWF after its infusion influences vWF interaction with activated platelets,
leading to clearance of platelets circulating in an (pre)activated state.
In the scope of a potential protective effect of FVIII deficiency on the development of
atherosclerosis, which could theoretically be counterbalanced by regular treatment with
coagulation factor concentrate, we studied the inflammatory response to infusion with FVIII
concentrate. No acute inflammatory response could be detected after FVIII concentrate
infusion by measuring hs-CRP, TNF-α, IL-6 and IL-8. Thus, infusion of FVIII concentrate
does not seem to influence atherogenesis via a stimulation of inflammation. The patients
selected for this study were all adult patients with severe hemophilia A. Consequently, all
patients had already exceeded 50 exposure days of FVIII concentrate in the past. Repeating
these measurements in patients in the period of their first exposures would give additional
insight into the inflammatory response after infusion of FVIII.
There are some limitations to this study which need to be addressed. First, a large
variety of parameters was tested in this study without correction for multiple hypothesis
testing, since this correction would increase the probability on false negative results.
Second, the sample size of this study was low; only 8 patients could be included. Third,
measurements were performed before, 15 minutes after and 60 minutes after infusion.
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And last, only plasma level of activated vWF was determined in our study and vWF bound
to platelets was not measured. Therefore, the effect of FVIII infusion on total amount of
activated vWF cannot be concluded from our data. Therefore, this study only gives insight
into the short-term effects of a single infusion. Future studies are needed to determine the
effects of repeated infusions, like in prophylactic treatment, on primary hemostasis.
Nowadays prophylactic regiments are targeted on FVIII activity trough levels, aiming to
alter a severe hemophilia patient into a mild or moderate patient at all times.29 Since FVIII
infusion does not only seem to affect FVIII levels, but also other parts of haemostasis, it
would be interesting to develop a new laboratory test, integrating primary and secondary
hemostasis, to use for targeting of prophylactic regiments in individual patients.
In conclusion, a single infusion of a high dose FVIII concentrate in hemophilia A patients
influences primary hemostasis by decreasing vWF, ADAMTS-13 and basal level of platelet
activation. These effects possibly occur as a consequence of binding of the infused FVIII to
vWF, influencing its clearance. Larger studies are needed to confirm these observations.
Disclosures
This work was supported by unrestricted grants from Sanquin Blood Supply Foundation
and CSL Behring.
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van Bladel ER, Roest M, de Groot PG, Schutgens RE. Up-regulation of platelet activation in
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Wartiovaara-Kautto U, Joutsi-Korhonen L, Ilveskero S, Armstrong E, Lassila R. Platelets
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Suzuki M, Murata M, Matsubara Y, Uchida T, Ishihara H, Shibano T, Ashida S, Soejima K, Okada
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Uemura M, Tatsumi K, Matsumoto M, Fujimoto M, Matsuyama T, Ishikawa M, Iwamoto TA,
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24.
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Cao W, Krishnaswamy S, Camire RM, Lenting PJ, Zheng XL. Factor VIII accelerates proteolytic
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Skipwith CG, Cao W, Zheng XL. Factor VIII and platelets synergistically accelerate cleavage of
von Willebrand factor by ADAMTS13 under fluid shear stress. J Biol Chem. 2010;285:28596603.
Fijnvandraat K, Peters M, ten Cate JW. Inter-individual variation in half-life of infused
recombinant factor VIII is related to pre-infusion von Willebrand factor antigen levels. Br J
Haematol. 1995;91:474-6.
Vlot AJ, Mauser-Bunschoten EP, Zarkova AG, Haan E, Kruitwagen CL, Sixma JJ, van den Berg
HM. The half-life of infused factor VIII is shorter in hemophiliac patients with blood group O
than in those with blood group A. Thromb Haemost. 2000;83:65-9.
Shim K, Anderson PJ, Tuley EA, Wiswall E, Sadler JE. Platelet-VWF complexes are preferred
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Collins PW, Bjorkman S, Fischer K, Blanchette V, Oh M, Schroth P, Fritsch S, Casey K, Spotts G,
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Chapter 4
Platelet behaviour is not related to bleeding phenotype
in severe hemophilia A patients
Esther R. van Bladel,1,2 Roger E.G. Schutgens,2,3 Kathelijn Fischer,3 Philip G. de Groot,1 Mark Roest1
1
Department of Clinical Chemistry and Hematology, University Medical Center
Utrecht, Utrecht, the Netherlands; 2Van Creveld Laboratory, University Medical
Center Utrecht, Utrecht, the Netherlands; and 3Van Creveldkliniek/Department of
Hematology, University Medical Center Utrecht, Utrecht, the Netherlands
In revision
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Abstract
Recently we reported data suggesting that platelets could compensate for the bleeding
phenotype in severe hemophilia A. The aim of this study was to confirm these results
in a larger population with a detailed characterization of clinical phenotype. Patients
with diagnostic severe hemophilia A (FVIII:C <1%) were scored for clinical phenotype by
integrating data on age at first joint bleed, joint damage, bleeding frequency and FVIII
consumption. Phenotype was defined as onset of joint bleeding-score + arthropathy-score
+ joint bleeding-score + (2* treatment intensity-score). After a washout period of 3 days,
blood was collected for measurement of basal level of platelet activation, platelet reactivity,
endothelial cell activation and presence of procoagulant phospholipids in plasma. Thirtythree patients with severe hemophilia A were included, 13 patients with a mild, 12 patients
with an average and 8 patients with a severe clinical phenotype. No relevant differences
in basal level of platelet activation, platelet reactivity, endothelial cell activation and
procoagulant phospholipids between all three groups were observed. The mean annual
FVIII consumption per kg did not correlate with the platelet P-selectin expression and
GPIIbIIIa activation on platelets. In conclusion, variability in clinical phenotype in patients
with diagnostic severe hemophilia A is not related to platelet activation or reactivity.
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Introduction
Hemophilia A is an X-linked bleeding disorder due to a deficiency in activity of clotting
factor VIII (FVIII). During clot formation, activated FVIII participates in the tenase complex
as a co-factor for FIXa, accelerating the activation of FX and subsequent thrombin and
fibrin formation. Deficiency in FVIII activity thus leads to a bleeding tendency. The clinical
severity of this bleeding tendency varies between patients.1
The main determinant of the bleeding severity in hemophilia A patients is the residual
activity of FVIII.2 Therefore patients have been classified into a diagnostic mild (FVIII:C >5%),
moderate (FVIII:C 1-5%) and severe (FVIII:C <1%) phenotype accordingly.3
For the general hemophilia population, this diagnostic phenotype correlates well with
clinical phenotype, however, variability of clinical phenotype within groups of the same
diagnostic phenotype is frequently observed. For example, in patients with a severe
diagnostic phenotype, 10% of patients have a mild clinical phenotype.4-6 This suggests
the presence of additional determinants of clinical phenotype in hemophilia A other than
residual FVIII activity.
In search of these additional causal factors, several possible determinants have been
studied. Prothrombotic factors such as factor V Leiden, FVIII half-life, genotype and
fibrinolytic activity were excluded as causal factors that influence the variability in clinical
phenotype of severe hemophilia A patients.7,8 In a previous study we found differences
in platelet activation between diagnostically severe hemophilia patients and normal
men.9 Furthermore, preliminary data showed a correlation between basal level of platelet
activation and FVIII consumption in a small cohort of hemophilia A patients.
In this study, we further analyze the role of platelets in determining the clinical
phenotype within a larger cohort of hemophilia A patients with undetectable factor VIII
levels.
4
Materials and methods
Patients and setting
To identify the most severe and mildest bleeders, all severe hemophilia A patients born
between 1965 and 1990, without a history of inhibitors, and currently in care of the Van
Creveldkliniek were scored for bleeding phenotype. Patients born before 1965 were not
considered for scoring, as these patients had no access to replacement therapy since the
onset of bleeding and this would affect the scoring system. As proposed by Schulman et al,
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the selection was based on bleeding pattern, treatment requirement and joint outcome.10
However, as Schulmans’ Hemophilia Severity Score was based on Swedish treatment
regimens, and the orthopaedic joint score was not routinely used in our clinic, this score
could not be applied in the Dutch setting. Therefore, the distributions of onset of joint
bleeding, bleeding frequency, treatment requirement and joint outcome (arthropathy)
were used, and for each parameter patients were classified as mild or severe if their data
were in the top-or bottom quadrant of the distribution. A parameter-value in the outer
quadrant corresponding to mild phenotype was assigned a value of -1, a value in the outer
quadrant corresponding to severe phenotype was assigned a value of +1, and values in the
two quadrants in between were assigned a value of 0. Data on baseline characteristics and
Pettersson scores from the centres’ database were used.1
The onset of joint bleeding-score was calculated by evaluating the distribution of age
at first joint bleed in patients who started prophylaxis after the onset of joint bleeding. In
16 patients, including 3 who had started prophylaxis before the onset of joint bleeding,
missing data were imputed using multivariate regression (SPSS version 18) based on year
of birth, age at diagnosis, age at entry in the clinic, age at start of prophylaxis, and total
number of years on prophylaxis. Based on the distribution of imputed values, patients in
the lowest quadrant were assigned an onset of joint bleeding score of +1, and those in the
highest quadrant a score of -1.
The arthropathy-score was based on all Pettersson scores available. All were scored by
one of two radiologists. As arthropathy is highly dependent on age, three age categories
were defined based on the tertiles of the age distribution of available Pettersson scores:
X-rays taken < 13 years, at 13-20 years, and after the age of 20 years. For the youngest age
group, low scores were considered non informative and only patients with scores in the
upper quadrant were assigned +1 point. For the other two age groups, the scores in the
lower quadrant were assigned -1 point, and in the upper quadrant +1 point. To enable the
use of all available X-rays over time, the mean severity score was calculated for each patient.
From the distribution of these scores, the lower and upper quadrants were assigned -1 and
+1 point respectively.
The joint bleeding score was based on the distribution of the median annual number
of joint bleeds calculated for each patient, using all data available for each patient,
independent of treatment received. Based on the quadrants of this distribution, patients
in the lowest quadrant were assigned -1 point, and those in the upper quadrant +1 point.
The treatment intensity score was based on annual clotting factor consumption (IU/
kg), similar to the approach of Schulman et al. As FVIII consumption is dependent on
pharmacokinetics, and these are dependent on blood group, all calculations were
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performed separately for patients with blood group O and those with blood group non-O.
To enable the use of all available treatment data, the mean annual FVIII consumption per
kilogram bodyweight over all treatment years in the database was calculated for each
patient. As a low annual FVIII consumption can be a sign of undertreatment, joint bleed
frequencies were taken into account to identify patients with a low FVIII consumption due
to a mild phenotype. Only those who had an overall FVIII consumption in the lowest tertile
and an overall joint bleed frequency below the group median were assigned -1 point for
treatment intensity. Patients with a high treatment intensity were defined according to
FVIII consumption only: those in the upper quadrant were assigned +1 point.
The total score was calculated by adding all separate parameter-scores in the following
formula: onset of joint bleeding-score + arthropathy-score + joint bleeding-score +
(2* treatment intensity-score). Patients with a score ranging from -5 to -2 were considered
to have a mild phenotype, patients with a score ranging from -1 to +1 an average phenotype
and patients with a score ranging from +2 to +5 a severe phenotype. Patients born before
1965 who were not scored, but were considered as mild bleeders by their treating physician
based on a very mild bleeding pattern or minor joint damage despite little treatment at
young age, were also selected for study.
Blood was drawn during regular visits at the Van Creveldkliniek (VCK), University
Medical Center Utrecht, into a vacuum citrate and EDTA tubes. All participants had a wash
out period for FVIII of minimally 3 days. All participants gave written informed consent and
the study was approved by the Medical Ethics Committees of the University Medical Center
Utrecht and performed in accordance with the Declaration of Helsinki.
4
Materials
R-phycoerythrin (PE) labelled antibodies analyses, raised against human P-selectin
(#555524), were purchased from BD biosciences (Franklin Lakes, NJ, U.S.A). Alexa Fluor 488
labeled human fibrinogen, used for labelling of open glycoprotein IIbIIIa receptor (open
GPIIbIIIa), was purchased from Invitrogen (Eugene, Oregon, U.S.A.).
For platelet responsiveness assays, adenosine diphosphate (ADP) was purchased
from Roche (Almere, The Netherlands), Thrombin receptor activator peptide (TRAP) from
Bachem AG (Bubendorf, Switzerland), thromboxane A2 receptor agonist U-46619 from
Santa Cruz Biotechnology Inc. (Santa Cruz, California), Iloprost (Ilomedine) from Bayer
Schering Pharma AG (Berlin, Germany) and Cross-linked Collagen Related Peptide (XL-CRP)
was a generous gift of R. Farndale (Cambridge, U.K.).
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) was purchased form BDH
(Poole, U.K.). Sodium chloride (NaCl), Tween-20 and Bovine serum albumin (BSA) were
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purchased from Sigma (Zwijndrecht, The Netherlands). Magnesium Sulphate (MgSO4) and
Potassium Chloride (KCl) were purchased from Riedel (Seelze, Germany), Formaldehyde
(CH2O) from Calbiochem (Merck, Darmstadt, Germany).
For ELISA, antibodies against human Platelet Factor 4 (PF4; MAB7951, AF795),
human chemokine (C-X-C motif) ligand 7 (CXCL7) (MAB393, BAF393), human chemokine
(C-C motif) ligand 5 (RANTES) (MAB278, AB-278-NA), human soluble P-selectin (DY137
Duoset) and soluble intercellular adhesion molecule (sICAM) (DY720 Duoset) were all
purchased from R&D Systems (Abingdon, U.K.). Rabbit anti-human von Willebrand Factor
(vWF) antibodies (A0082), peroxidase conjugated rabbit anti-human vWF (P0226), rabbit
anti-goat horseradish peroxidase (HRP) (P0449) and streptavidin-poly-HRP (P0397) were
purchased from DAKO (Glostrup, Denmark). Rabbit anti-vWF propeptide and rabbit antivWF propeptide/biotine were prepared as described by Borchiellini et al.11 A11 nanobody
used as coating antibody in the active vWF ELISA was prepared as described by Hulstein et
al.12 A recombinant vWF (2BQ), with a R1306Q mutation, was used as calibration curve to
determine the amount of vWF in its GPIb binding conformation (active vWF).13 SuperSignal
ELISA Pico chemiluminescent substrate was purchased from Thermo Scientific (Rockford,
Illinois, U.S.A.).
For the measurement of procoagulant phospholipids, a Procoagulant PPL kit was used
(Stago diagnostics, Asnières sur Seine, Fance)
Platelet activation and responsiveness
Platelet responsiveness to agonists was determined with concentration series of ADP
(0.01 μM, 0.03 μM, 0.12 μM, 0.49 μM, 1.95 μM, 7.81 μM, 31.25 μM and 125.00 μM), CRP
(0.2 ng/ml, 0.6 ng/ml, 2.4 ng/ml, 9.8 ng/ml, 39.1 ng/ml, 156.3 ng/ml, 625.0 ng/ml and
2500.0 ng/ml), TRAP (0.04 μM, 0.15 μM, 0.61 μM, 2.44 μM, 9.77 μM, 39.06 μM, 156.25 μM
and 625.00 μM) and the thromboxane A2 receptor agonist U-46619 (0.1 μM, 0.2 μM,
0.6 μM, 1.8 μM, 5.3 μM, 15.9 μM, 47.6 μM and 142.7 μM), while platelet responsiveness to
inhibitors was measured with concentration series of TRAP (0.04 μM, 0.15 μM, 0.61 μM,
2.44 μM, 9.77 μM, 39.06 μM, 156.25 μM and 625.00 μM) containing 5ng/ml iloprost
and with concentration series of iloprost (0.02 ng/ml, 0.08 ng/ml, 0.31 ng/ml, 1.22 ng/
ml, 4.88 ng/ml, 19.53 ng/ml, 78.13 ng/ml and 312.50 ng/ml) containing 5 μM ADP. Serial
dilutions were prepared in 50 μL HEPES buffered saline (HBS; 10 mM HEPES, 150 mM NaCl,
1 mM MgSO4, 5 mM KCl, pH 7.4, filtered through a 0.22 μm filter) containing 2 μL PE labelled
mouse anti-human P-selectin antibodies and 0.5 μL Alexa Fluor 488 labelled fibrinogen.
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A negative control sample, only containing 50 μL HBS with 2 μL PE labelled mouse antihuman P-selectin antibodies and 0.5 μL Alexa Fluor 488 labelled fibrinogen, was prepared
to determine the basal level of platelet activation.
The platelet activation test was initiated by addition of 5 μL fresh, citrate anticoagulated,
whole blood to each mixture of the serial dilutions. After 20 minutes of incubation, the
samples were fixed with 500 μL 0.2% formyl saline (0.2% formaldehyde in 0.9% NaCl,
filtered through a 0.22 μm filter) and after 10 minutes of fixation kept at 4°C until analyses.
All samples were analyzed on a FACS Canto II flow cytometer from BD Biosciences (San
Jose, CA, U.S.A.) within one day after processing. Single platelets were gated based on
forward and side scatter properties. The median fluorescence intensity (MFI) in the platelet
gate was measured with FACS analysis for both P-selectin and open GPIIbIIIa signal. One
individual performed all assays.
4
ELISA procedure
To prepare platelet poor plasma, citrated whole blood was centrifuged for 10 minutes at
2000 G twice and stored at -80°C until analysis. ELISAs were performed to measure PF4,
CXCL7, RANTES, soluble P-selectin, sICAM, VWF, VWF propeptide and active VWF in citrated
plasma samples.
Plates were coated with monoclonal mouse anti-human PF4 (1 μg/mL), purified
monoclonal mouse anti-human CXCL7 (1 μg/mL), purified mouse monoclonal anti-human
RANTES (500 ng/mL), mouse anti-human P-selectin (1 μg/mL), mouse anti-human ICAM1 (2 μg/ml), polyclonal rabbit-anti-human vWF (0.775 μg/mL), rabbit anti-human vWF
propeptide (5 μg/mL) and A11 nanobody (5 μg/mL), respectively, and incubated overnight
at 4°C. All plates were blocked with PBS/1%BSA for 1 hour, except for the vWF propeptide
ELISA in which PBS/2%BSA/100mM EDTA/0.1%Tween was used for blocking.
Citrated plasma samples of all patients were put on plate in 75x dilution for PF4 and
CXCL7 ELISA, 10x dilution for RANTES, soluble P-selectin and active vWF ELISA, 15x dilution
for sICAM ELISA, 25x dilution for vWF ELISA and 20x for vWF propeptide ELISA.
As first point of the calibration curve, normal pool serum diluted 250x to 1.4 ng/ml
for PF4 ELISA, 250x to 2.2 ng/ml for CXCL7 ELISA, 4x to 9.2 ng/ml for RANTES ELISA, 4x to
70 ng/ml for soluble P-selectin ELISA and 15x to 867 ng/ml for vWF ELISA was added to
the corresponding plates. VWF deficient plasma containing 2 μg/ml 2BQ was used for the
active VWF ELISA.
After 1 hour incubation, goat anti-PF4 (0.25 μg/ml), biotinylated goat anti-hNAP-2(50 μg/
ml), goat anti-human RANTES (1 μg/ml), biotinylated sheep anti-human P-Selectin (0.01 ng/
ml), peroxidase-conjugated rabbit anti-human vWF (0.275 μg/ml), biotinylated rabbit
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anti-human vWF propeptide (2.5 μg/ml) and peroxidase-conjugated rabbit anti-human
vWF(0.55 μg/ml) were added as secondary antibodies for another 1 hour incubation step.
For the PF4 and RANTES ELISA rabbit anti-goat-HRP (6500x dilution), for the CXCL7 and
soluble P-selectin ELISA streptavidin-poly-HRP (0.15 μg/ml), and for vWF propeptide ELISA
streptavidin-mono-HRP (0.71 μg/ml) were used as detection antibodies.
Finally, SuperSignal ELISA Pico chemiluminescent substrate was added to all plates and
single wavelength luminescence was measured directly (target calibration wavelength 470
nm) with a SpectraMax L microplate reader from Molecular Devices Inc. (Silicon Vally, CA,
USA).
Procoagulant phospholipids
Functional measurement of procoagulant phospholipids was performed in platelet poor
plasma (citrate) according to manufacturer’s protocol, using the STA® Procoag PPL kit on
a STA-R Evolution analyzer (Stago diagnostics, Asnières sur Seine, Fance). In short, plasma
samples were incubated for 2 minutes with procoagulant phospholipid deficient plasma.
Activated clotting factor X (Xa) was added, and clotting time was measured.
Statistical Analysis
Initial quantification of FACS data was performed in BD FACSDiva software 6.1.2 (BD
Biosciences, San Jose, CA, U.S.A.). To determine platelet responsiveness, the concentration
generating a response halfway between baseline and maximum MFI for both P-selectin and
open GPIIbIIIa signal (EC50), and the area under the activation curve (AUC) was calculated
using GraphPad Prism version 5.03 (GraphPad Software, San Diego, CA, U.S.A.). The maximal
effect of stimulation or inhibition is represented by the MFI in the sample with the highest
used (ant-)agonist concentration.
A statistical analysis was performed with IBM SPSS Statistics 20.0.0 for Windows
(International Business Machines Corporation, New York, U.S.A.). Data are expressed as
median and interquartile range (IQR) and comparison between two groups was performed
by Mann-Whitney U testing unless indicated otherwise. P-values lower than 0.05 were
considered to be significant.
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Results
Patient characteristics
Original 112 patients with severe hemophilia A born between 1965 and 1990 with
complete data on treatment and outcome were present in our database. Sixteen patients
were excluded because of a history of inhibitor development and 13 because they were
no longer attending our clinic. The score was applied to the remaining 83 patients, with
a median follow up of 17 years (range 7-31), who were followed until a median age of
22.5 yrs (range 11.7-37.4). In total, data on 1644 treatment years and 274 Pettersson scores
(median 2 per patient) were analyzed. Two patients had only incomplete Pettersson scores
available; in these patients, an ‘average’ i.e. non-informative Pettersson score was assumed
for the calculation of the phenotype score. Eventually 17 patients were scored as having
a mild phenotype, 43 as average and 23 as severe. Four additional patients, who were
not scored in the database because they were born before 1965 (n=4) but had very mild
bleeding pattern or minor joint damage despite little treatment at young age, were also
selected for study. Also, one patient who was not scored in the database because he was
not in our care in previous years, with an evidently mild clinical phenotype with less than
50 exposure days at adult age, was selected for study.
Out of 83 patients, 33 patients with diagnostically severe hemophilia agreed to
participate, of which 13 patients with a mild clinical phenotype, 12 patients with an average
clinical phenotype and 8 patients with a severe clinical phenotype. Clinical characteristics
of included groups are shown in Table 1.
Selection conform phenotype is validated by the significant differences in Pettersson
score, joint bleeding frequency and FVIII consumption between the phenotypically mild
and severe patients. Age of first joint bleed did not differ significantly between groups.
4
Comparison between mild, average and severe clinical phenotype
Patients scored as mild, average and severe clinical phenotype were compared for basal
level of platelet activation, platelet reactivity, endothelial cell activation and presence
of procoagulant phospholipids in plasma. Results are presented in Table 2. No relevant
differences between all three groups were observed. For instance, basal level of platelet
P-selectin expression was equal between patients with a severe clinical phenotype (15.5
AU (9.8-22.5)) and patients with a mild (16.5 AU (12.5-24.5); p=0.500) or average (14.5 AU
(11.0-20.8); p=0.792) clinical phenotype.
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18 (14-23)
19 (15-26)
247 (198-281)
8.5 (7.8-8.8)
0.047
0.236
0.750
0.972
0.487
0.766
0.068
0.611
13 (9-17)
16 (12-26)
312 (261-351)
8.1 (7.6-8.6)
11 (1-18)
13 (13-22)
240 (217-269)
8.3 (7.8-8.9)
Data express median (interquartile range), * analysis adjusted for age
Age, years
Age first joint bleed, years
Last Pettersson score
Age at last Pettersson score, years
Joint bleeding, nr/year
Mean annual FVIII consumption,
IU/kg/year
Total period on prophylaxis, years
Duration of follow up, years
Platelet count, *109/L
Mean platelet volume, fL
Severe (n=8)
Mild vs. average
p-value
0.347
0.282
0.004*
0.426
0.077
0.001
Average (n=12)
Mild vs. severe
p-value
36 (33-40)
33 (24-42)
35 (27-39)
0.414
2.06 (1.01-3.78) 4.57 (1.13-5.64) 2.00 (1.44-2.38) 0.945
8 (4-13)
9 (1-30)
20 (12-27)
<0.001*
29 (25-34)
24 (14-33)
26 (18-31)
0.492
1.33 (1.00-2.00) 2.81 (1.04-9.65) 6.23 (3.56-8.19) <0.001
894 (361-1114) 1768 (1461-2279) 2158 (1506-2462) <0.001
Mild (n=13)
Table 1. Baseline characteristics of all severe hemophilia patients, according to their clinical bleeding phenotype
0.075
0.657
0.039
0.305
Severe vs. average
p-value
0.970
0.336
0.001*
0.965
0.343
0.571
Bladel.indd 61
Basal level of platelet activation
P-selectin expression, AU
open GPIIbIIIa, AU
Platelet Factor 4, ng/ml
CXCL7, ng/ml
RANTES, ng/ml
Soluble P-selectin, ng/ml
Platelet reactivity to ADP – P-selectin
Maximal MFI, AU
EC50, μM
AUC, 103*AU
Platelet reactivity to ADP – open GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, 103*AU
Platelet reactivity to CRP – P-selectin
Maximal MFI, AU
EC50, ng/ml
AUC, 105*AU
Platelet reactivity to CRP – open GPIIbIIIa
Maximal MFI, AU
EC50, ng/ml
AUC, 105*AU
14.5 (11.0-20.8)
2.0 (-0.5-9.0)
6.41 (4.58-9.21)
11.01 (7.66-15.61)
1.17 (0.76-1.46)
66.05 (57.56-71.28)
1509 (1146-1865)
1.16 (0.94-1.57)
178 (141-215)
1522 (627-2190)
0.59 (0.41-1.00)
180 (70-254)
6986 (6489-7492)
247 (126-289)
142 (123-162)
5203 (3211-6354)
305 (217-446)
96 (64-128)
1720 (1236-2114)
1.13 (1.04-1.28)
210 (148-259)
1699 (333-2293)
0.62 (0.56-0.99)
198 (44-287)
7494 (6736-7820)
205 (176-419)
150 (120-166)
3889 (1004-5548)
203 (179-281)
81 (15-117)
Average (n=12)
16.5 (12.5-24.5)
9.5 (2.5-21.0)
6.46 (5.34-10.26)
10.08 (8.79-19.39)
1.12 (0.95-1.79)
55.55 (46.24-61.24)
Mild (n=13)
Table 2. Platelets and endothelium variables according to clinical phenotype
0.689
0.152
0.574
0.470
0.470
0.068
0.470
0.810
0.503
0.936
0.650
0.769
0.503
0.936
1.000
0.270
0.406
0.503
0.500
0.238
0.210
0.185
0.500
0.336
1616 (1325-2418) 1.000
1.20 (0.83-1.58) 0.916
207 (163-294)
0.860
0.547
0.645
0.595
15.5 (9.8-22.5)
2.3 (-7.8-16.0)
5.02 (4.21-8.23)
9.14 (5.77-13.92)
0.89 (0.70-1.60)
61.36 (55.29)
1786 (979-3237)
0.63 (0.48-1.54)
214 (93-407)
7328 (6883-7928) 1.000
277 (227-443)
0.140
152 (124-157)
0.916
5369 (2653-5964) 0.238
285 (230-796)
0.121
106 (39-117)
0.595
Mild vs.
average
p-value
Mild vs.
severe
p-value
Severe (n=8)
4
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1-11-2013 14:31:37
0.970
0.851
0.910
0.384
0.343
0.970
0.571
0.384
0.678
0.571
0.851
0.384
0.792
0.851
0.571
0.734
0.734
0.473
Severe vs.
average
p-value
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Platelet reactivity to TRAP – P-selectin
Maximal MFI, AU
EC50, μM
AUC, 104*AU
Platelet reactivity to TRAP – open GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, 104*AU
Platelet reactivity to TRAP(+iloprost) –
P-selectin
Maximal MFI, AU
EC50, μM
AUC, 104*AU
Platelet reactivity to TRAP(+iloprost) – open
GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, 103*AU
Platelet reactivity to U46619 – P-selectin
Maximal MFI, AU
EC50, μM
AUC, 103*AU
Table 2. Continued
Mild vs.
severe
p-value
5966 (5202-6568)
9.11 (7.82-25.13)
345 (301-393)
309 (253-354)
7.54 (6.12-21.91)
158 (139-185)
5066 (4294-6290) 5034 (4374-5882) 0.916
15.25 (8.06-22.50) 5.92 (2.57-33.72) 0.860
603 (452-747)
598 (413-717)
0.804
281 (158-348)
7.08 (5.14-8.81)
135 (99-189)
5493 (3819-6223)
6.01 (3.84-11.66)
567 (449-861)
272 (218-303)
7.55 (6.29-20.06)
156 (114-184)
0.860
0.374
0.860
6538 (6057-7078) 0.750
8.20 (7.60-9.00) 0.185
401 (363-434)
0.750
0.697
0.185
0.750
6551 (5751-6981)
7.48 (7.18-8.61)
393 (342-437)
2020 (811-3472)
1.83 (1.57-2.35)
102 (44-183)
2255 (1415-2839)
1.78 (1.47-2.22)
119 (68-149)
1923 (701-2696)
1.63 (1.32-1.73)
82 (36-141)
8085 (7240-8397) 0.547
2.20 (1.99-2.41)
0.268
495 (428-517)
0.804
Severe (n=8)
7791 (6697-8130)
2.12 (1.84-2.41)
472 (416-501)
Average (n=12)
7601 (7245-8145)
1.98 (1.86-2.22)
418 (443-518)
Mild (n=13)
0.894
0.186
1.000
0.406
0.470
0.376
0.437
0.137
0.406
0.538
0.270
0.503
0.894
0.650
0.979
Mild vs.
average
p-value
0.734
0.473
0.734
0.427
0.970
0.678
0.208
0.473
0.157
0.910
0.624
0.792
0.473
0.734
0.571
Severe vs.
average
p-value
Bladel.indd 63
2834 (2283-3297)
8.13 (3.93-18.00)
364 (301-442)
52 (40-65)
0.19 (0.08-0.44)
246 (181-281)
28 (27-41)
0.06 (0.02-0.12)
123 (97-151)
181 (153-217)
10.68 (9.80-16.74)
0.84 (0.53-1.00)
5.30 (4.86-5.70)
74 (57-80)
34 (24-52)
0.17 (0.12-0.42)
205 (128-278)
23 (14-31)
0.08 (0.01-0.17)
101 (56-119)
207 (137-241)
11.05 (9.82-14.10)
0.88 (0.51-0.97)
5.23 (4.96-5.57)
71 (53-82)
Average (n=12)
2345 (1418-2844)
5.67 (2.12-6.48)
313 (230-375)
Mild (n=13)
0.728
0.979
0.936
0.852
0.456
13.41 (9.27-19.57) 0.374
0.75 (0.47-1.06)
0.860
5.94 (4.86-6.52) 0.456
0.804
181 (118-229)
66 (61-74)
0.728
0.185
0.161
0.161
30 (24-39)
0.14 (0.06-0.27)
121 (100-131)
0.087
0.810
0.110
0.205
0.979
0.437
0.121
0.697
0.301
59 (44-90)
0.37 (0.07-0.48)
240 (212-354)
Mild vs.
average
p-value
0.225
0.186
0.376
Mild vs.
severe
p-value
2289 (1851-3175) 1.000
4.83 (2.41-20.40) 0.414
321 (190-484)
0.804
Severe (n=8)
0.473
0.851
0.970
0.343
0.851
0.851
0.135
0.792
0.473
0.734
0.792
0.384
0.734
0.734
Severe vs.
average
p-value
MFI median fluorescence intensity; AU arbitrary units; EC50 concentration of (ant-)agonist yielding a response halfway baseline and maximal
response; AUC area under the activation curve.
Platelet reactivity to U46619 – open
GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, 103*AU
Platelet reactivity to iloprost(+ADP) –
P-selectin
Maximal MFI, AU
EC50, ng/ml
AUC, 102*AU
Platelet reactivity to iloprost(+ADP) – open
GPIIbIIIa
Maximal MFI, AU
EC50, ng/ml
AUC, 102*AU
sICAM
sICAM, ng/ml
Von Willebrand Factor
vWF, μg/ml
Active vWF, μg/ml
vWF propeptide, nM
Procoagulant phospholipids
Clotting time, seconds
Table 2. Continued
4
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40
30
r -0.21
20
B. Correlation between FVIII consumption
and platelet open GPIIbIIIa expression
Platelet open GPIIbIIIa - MFI
Platelet P-selectin - MFI
A. Correlation between FVIII consumption
and platelet P-sectin expression
10
50
40
30
20
r -0.30
10
0
-10
0
0
1000
2000
3000
Mean Annual FVIII consumption
4000
-20
0
1000
2000
3000
Mean Annual FVIII consumption
4000
Figure 1. Correlation between FVIII consumption and basal level of platelet activation in patients
with severe hemophilia A
Spearman correlations between mean annual FVIII consumption in IU/kg/year and (A) basal level of
platelet P-selectin expression in AU and (B) basal level of open GPIIbIIIa in AU are shown.
Correlation with FVIII consumption
When correlating the mean annual FVIII consumption per kg to the platelet P-selectin
and open GPIIbIIIa expression, no correlations were found (Spearman’s r -0.21 and -0.30;
p=0.246 and =0.093) in this study population. Results are presented in Figure 1.
Discussion
There is a recent interest in the potential modifying effects of platelet function on clinical
phenotype in hemophilia.9,14-16 In the current study, we found that variation in platelet
function is not related to clinical phenotype in patients with diagnostic severe hemophilia.
From a previous study we suggested a possible role for platelets to compensate for
the bleeding phenotype of severe hemophilia A patients.9 In that study the basal level of
platelet P-selectin expression was higher in patients with severe hemophilia A as compared
to controls and negatively correlated with FVIII consumption. The aim of the current study
was to confirm findings from our previous study in a larger population, with a more detailed
characterization of clinical phenotype. Characterization of clinical phenotype of patients
with the diagnosis severe hemophilia A (FVIII:C <1%) was extended by integrating FVIII
consumption with bleeding frequency, age of first joint bleed and Pettersson score into
a clinical phenotype score. Subsequently, comparison was made between patients with
highest scores, average scores and lowest scores in search of variables that besides residual
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factor VIII levels also determine the clinical phenotype. Basal level of platelet activation
and the sensitivity of platelets for their major (ant-)agonists were studied as possible
determinants. Unexpectedly, we detected no differences in these parameters between
patients with a mild clinical phenotype, an average clinical phenotype and a severe clinical
phenotype in the current study.
In view of the different results, we questioned if dissimilarities in study design could
explain these differences. First, there was a difference in patient selection and the definition
of bleeding phenotype between the two studies. In the previous study, patients were
selected randomly from hemophilia A patient visits at the Van Creveldkliniek and bleeding
phenotype was determined by collecting data on FVIII consumption over the years after
patient inclusion. In the present study patients were pre-selected according to bleeding
phenotype. Bleeding phenotype was defined not only by FVIII consumption, but on a score
additionally integrating age at first joint bleed, bleeding frequency and joint damage for
all severe hemophilia A patients in the Van Creveldkliniek. When correlating the current
results with only FVIII consumption, like in the former study, no correlations were found.
A second difference with the previous study was that in the current study all patients on
prophylaxis were asked to take their last FVIII infusion 3 days before inclusion, on demand
patients were only included when no FVIII was infused in the 3 days before entering this
study. This washout period ensures that there is no infused FVIII present in the blood at
time of study, preventing a potential influence on results.
A third dissimilarity between the two studies is the sample size. In the present study
33 patients with diagnostic severe hemophilia A were included, while the previous study
contained only 13 patients with diagnostic severe hemophilia A.
Last, in the current study the platelet reactivity assay was expanded by the introduction
of additional agonists TRAP and U46619, and by the addition of FITC-labelled fibrinogen
as second marker of platelet activation, leading to a more complete representation of
platelet reactivity in the present study. Measurements from the present study were done
with a FACS CANTO device, where the former study was measured on a FACSCalibur flow
cytometer from BD Biosciences (Franklin Lakes, NJ, U.S.A.). There was no difference in
readout between the two FACS machines when measuring patient samples on both devices.
All other handlings were identical between studies: collection of patient blood occurred at
the Van Creveldkliniek, and was directly brought to the lab for further processing. In both
studies the platelet reactivity assay was performed within 1 hour after blood collection and
measured within 24 hours after fixation. This makes it unlikely that differences in blood
collection are the cause of the differences observed between studies.
4
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In response to our previous study, Teyssandier et al. published results in hemophilia
mice which could not confirm our findings.16 Hemophilia mice were selected instead
of human patients, to limit environmental and genetic influences. In this study, neither
difference in platelet activation, nor in platelet reactivity to TRAP-4 could be found between
hemophilia and wild type mice. Combining these findings with those of the current study,
we believe that baseline platelet activation does not influence clinical phenotype in severe
hemophilia A.
The method of identifying patients with different clinical phenotypes used in the present
study has not yet been validated. However, it uses the same components as the score
developed by Schulman et al.10 Schulman used average joint bleeding over a 10 year period,
combined with the orthopaedic joint score (Gilbert score), and clotting factor consumption
over 10 years, adjusted for age at start of prophylaxis and body weight. The present study
used data with a much longer follow up (interquartile range 12-25 years), a validated tool
for assessment of arthropathy (Pettersson score), and a more detailed analysis of clotting
factor consumption.
A possible margin of error is introduced via the integration of factor consumption. In the
Van Creveldkliniek patient receive prophylaxis tailored to their bleeding frequency. For the
group with low factor usage, it is easy to establish if these were truly patients with a mild
clinical phenotype by taking into account their bleeding frequency. However, for the group
with high factor usage, it is more difficult to establish if they truly have a severe clinical
phenotype and were treated properly, or if other factors like patient anxiety for bleeds or
patient misinterpretation of physical sensations as bleeds, have led to excess factor usage.
Nevertheless, by integrating factor consumption with the other components of the clinical
phenotype score, it approximates the actual clinical phenotype of patients and can be well
used in the search for factors determining this phenotype.
From previous research we know that phenotype is determined at a young age. Age
of first joint bleed still correlates with characteristics of phenotype, like factor usage
and Pettersson score, 25 years later.1 In case of a severe phenotype, one would want to
interfere as young as possible. To do so, it will be required to detect a severe phenotype
and its determinants at young age, possibly before the first joint bleed. Collecting data and
samples prospectively from young age on will be essential to make this possible.
In the past several factors have been postulated and studied as possible determinants
of clinical phenotype in patients with diagnostic severe hemophilia A. Patient lifestyle
has been suggested as a possible determinant of bleeding; e.g. differences in intensity
of patients’ physical activity could influence the tendency to bleed, however studies are
lacking. Prothrombotic factors, FVIII half-life, genotype and fibrinolytic activity could
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not explain variability in bleeding patterns between individual patients.7,8 It was unclear
if there was a possible role of platelets in the variability of hemophilia phenotype. The
current study gives new insight into the behaviour of platelets in patients with severe
hemophilia A, and shows that they are not solely responsible for the differences observed
in phenotype. Considering all factors which have been excluded as a single determinant
of bleeding phenotype in severe hemophilia A, the likeliness of a multifactorial causality
increases. To be able to characterize such a complex causality, large studies measuring
combinations of possible determinants are required. In a rare disease as hemophilia, this
calls for international collaborations to unravel the exact cause of observed differences
in bleeding phenotype. Also, the development of new tests, in which all different factors
involved in hemostasis, e.g. platelets, plasma, vessel wall and flow, are integrated, could
lead to advances in this field.
In summary, in the current study we show that variability in clinical phenotype in a
well-characterized cohort of patients with diagnostic severe hemophilia A (FVIII:C <1%) is
not related to platelet activation or reactivity. Future multicenter studies and development
of new tests integrating multiple facets of hemostasis will be needed to unravel the
pathogenesis of variability in clinical phenotype in hemophilia A.
4
References
1.
2.
3.
4.
5.
6.
7.
8.
Van Dijk K, Fischer K, Van der Bom JG, et al. Variability in clinical phenotype of severe
haemophilia: the role of the first joint bleed. Haemophilia 2005; 11: 438-43.
den Uijl IEM, Fischer K, Van Der Bom JG, et al. Clinical outcome of moderate haemophilia
compared with severe and mild haemophilia. Haemophilia 2009; 15(1): 83-90.
White GC, Rosendaal F, Aledort LM, et al. Definitions in hemophilia. Recommendation of
the scientific subcommittee on factor VIII and factor IX of the scientific and standardization
committee of the International Society on Thrombosis and Haemostasis. Thromb Haemost
2001; 85(3): 560.
Rainsford SG, Hall A. A three-year study of adolescent boys suffering from haemophilia and
allied disorders. Br J Haematol 1973; 24(5): 539-51.
Aledort LM, Haschmeyer RH, Pettersson H. A longitudinal study of orthopaedic outcomes for
severe factor-VIII-deficient haemophiliacs. The Orthopaedic Outcome Study Group. J Intern
Med 1994; 236(4): 391-9.
Molho P, Rolland N, Lebrun T, et al. Epidemiological survey of the orthopaedic status of severe
haemophilia A and B patients in France. The French Study Group. Haemophilia 2000; 6(1): 2332.
van Dijk K, van der Bom JG, Fischer K, et al. Phenotype of severe hemophilia A and plasma
levels of risk factors for thrombosis. J Thromb Haemost 2007; 5(5): 1062-4.
van Dijk K, van der Bom JG, Lenting PJ, et al. Factor VIII half-life and clinical phenotype of
severe hemophilia A. Haematologica 2005; 90(4): 494-8.
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9.
10.
11.
12.
13.
14.
15.
16.
van Bladel ER, Roest M, de Groot PG, et al. Up-regulation of platelet activation in hemophilia
A. Haematologica 2011; 96(6): 888-895.
Schulman S, Eelde A, Holmstrom M, et al. Validation of a composite score for clinical severity
of hemophilia. J Thromb Haemost 2008; 6(7): 1113-21.
Borchiellini A, Fijnvandraat K, ten Cate JW, et al. Quantitative analysis of von Willebrand
factor propeptide release in vivo: effect of experimental endotoxemia and administration of
1-deamino-8-D-arginine vasopressin in humans. Blood 1996; 88(8): 2951-8.
Hulstein JJ, de Groot PG, Silence K, et al. A novel nanobody that detects the gain-offunction
phenotype of von Willebrand factor in ADAMTS13 deficiency and von Willebrand disease type
2B. Blood 2005; 106(9): 3035–3042.
Lankhof H, Damas C, Schiphorst ME, et al. Functional studies on platelet adhesion with
recombinant von Willebrand Factor Type 2B mutants R543Q and R543W under conditions of
flow. Blood 1997; 89(8): 2766-72.
Wartiovaara-Kautto U, Joutsi-Korhonen L, Ilveskero S, et al. Platelets significantly modify
procoagulant activities in haemophilia A. Haemophilia 2011; 17: 743-51.
Hoffman M, Monroe DM. Low intensity laser therapy speeds wound healing in hemophilia by
enhancing platelet procoagulant activity. Wound Repair Regen 2012; 20(5): 770-7.
Teyssandier M, Delignat S, Rayes J, et al. Activation state of platelet in experimental severe
hemophilia A. Haematolgica 2012; 97(7): 1115-6.
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Chapter 5
FVIII uptake by platelets and megakaryocytes
Esther R. van Bladel,1 Roger E.G. Schutgens,2 Philip G. de Groot,3 Mark Roest3
1
Department of Clinical Chemistry and Hematology/Van Creveldlaboratory, University Medical
Center Utrecht, Utrecht, the Netherlands; 2Van Creveldkliniek/Department of Hematology,
University Medical Center Utrecht, Utrecht, the Netherlands; and 3Department of Clinical
Chemistry and Hematology, University Medical Center Utrecht, Utrecht, the Netherlands.
In preparation
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Abstract
Plasma FVIII levels do not account fully for the bleeding tendency in hemophilia patients.
Uptake of FVIII into cells may be an alternative source of FVIII that influences bleeding
tendency. We investigated, both in vitro and in vivo, the capability of platelets and their
predecessors megakaryocytes to take up FVIII. Presence of FVIII inside cells was determined
by measuring the capability of cell lysates to shorten the APTT of FVIII deficient plasma and
by SDS-PAGE and western blotting of FVIII in cell lysates. Measurement were performed
with platelets from healthy donors, platelets from hemophilia patients and with a
megakaryocyte cell line (CHRF-288-11 cells). Trace amounts of FVIII are present in platelets
from healthy donors, with amounts increasing when incubating platelets with FVIII,
especially when activating platelets. FVIII is also present in platelets of severe hemophilia
A patients, even when prophylactic FVIII treatment was withheld for a minimum of 3 days
before blood collection, suggesting that in vivo uptake might take place at an earlier cell
stage. Megakaryocytes are also able to take up FVIII.
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Introduction
Hemophilia A patients are burdened with a bleeding tendency caused by a deficiency
in clotting factor VIII (FVIII). When the bleeding tendency is severe, patients are treated
prophylacticly with FVIII concentrate infusions to reduce the number of bleeds.1 Less severe
patients are treated on demand when a bleed arises.
Severe hemophilia A patients (FVIII:C <1%), receive prophylactic treatment to convert
them into a moderate patient (FVIII:C 1-5%), thereby reducing the number of bleeds and
consequently decreasing arthropathy. Prophylactic regimens aim to constantly keep the
FVIII activity level above 1%. The main determinants to achieve patient plasma levels above
this trough level are dosing frequency and lifetime of FVIII, a time that varied between
different patients.2
Nonetheless, there are conflicting results between plasma FVIII levels and observed
clinical effects. Studies show that some patients do not bleed in spite of trough plasma FVIII
levels below 1%.3,4 This indicates that there are other determinants of bleeding in addition
to plasma FVIII activity levels.
If plasma FVIII levels do not account fully for the bleeding tendency in hemophilia
patients, then uptake of FVIII into cells may be an alternative source of FVIII that influences
bleeding tendency. Ideally, these cell should be capable to deliver FVIII at sites of vascular
damage. As platelets meet these demands, we hypothesized that platelets and their
predecessors megakaryocytes might take up FVIII. In this study we show that both platelets
and megakaryocytes can take up FVIII both in vitro and in vivo.
5
Methods
Platelets
Blood of healthy donors was collected via the UMC Utrecht Mini Donor System, via which
healthy employees of the UMC Utrecht donate blood for research purposes. Severe
haemophilia A patients (FVIII:C <1%) under treatment at the van Creveldkliniek were invited
to participate. All participants gave written informed consent as approved by the Medical
Ethics Committees of the University Medical Center Utrecht. Blood was drawn from the
antecubital vein through 20-gauge needles, into a Vacutainer® tubes, containing sodiumcitrate (3.2%). Whole blood was centrifuged 15 minutes at 160 G to obtain platelet rich
plasma (PRP).
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Megakaryocyte culture
CHRF-288-11 cells5 were grown in Fischers’ medium containing 20% horse serum, 100 U/
ml pecilline, 100 U/ml strep and 2 mM L-glutamin at 37°C and 5% CO2 as described earlier.6
FVIII uptake
Either PRP or medium containing 1*106 megakaryocytes/ml was incubated with (r)FVIII for
2 hours (unless stated otherwise). For experiments with a pre-activation step, platelets
were pre-activated with 0.01 ng/ml crossed linked collagen related pepide (CRP, a generous
gift of R. Farndale, Cambridge, U.K.) and/or 200 µM RGD-containing peptide Darginylglycyl-L-aspartyl-L-tryptophane (dRGDW, synthesized at the Department of Membrane
Enzymology, Faculty of Chemistry, University of Utrecht, the Netherlands) for 10 minutes
before the addition of (r)FVIII.
After incubation, 1:10 ACD was added tot the PRP. Platelets were isolated by
3 centrifugation steps of 340 G for 15 minutes. Before the second centrifugation step
2 ul/ml iloprost was added to prevent platelet aggregation. Megakayocytes were isolated
by 3 centrifugation steps of 243 G for 5 minutes. Supernatant was discarded; cells were
resuspended into HT buffer after each centrifugation steps (pH 6.5, 6.5 and 7.3 respectively).
FVIII detection by APTT
After the third centrifugation step, the platelet count was set at 200 G/L. Platelets were
lysed by freezing in liquid NO and thawing in a 37°C water bath two times. Subsequently
cell remnants were removed by 30 seconds centrifugation at 12470 G. To control for
successful washing of platelets, control samples were created by first spinning down
platelets by 30 seconds centrifugation at 12470 G, after which the supernatant was freeze/
thawed two times. All samples were mixed 1:1 with FVIII deficient plasma (STA Factor
VIII, 1490150 140, Diagnostica Stago, Asnières, France). APTT was determined on a KC10
A micro coagulometer (Amelung, Lemgo, Germany), using PTT reagent (no. 10126551 140,
Diagnostica Stago, Asnières, France) and a 25 mM calcium solution to start the reaction. As
a scaling sample, FVIII deficient plasma was mixed with HT buffer 1:1, and the APTT of this
sample was determined. In this way the shortening of the APTT by the platelet lysates in
comparison to the scaling sample could be determined.
FVIII detection by SDS-PAGE and Western Blot
To prepare platelets for Western blot, the final cell pellet was taken up in sample buffer
containing DTD, to create reducing conditions. A 0.1 IU/ml and 1 IU/ml human purified FVIII
solution were used as positive control. The samples were heated to 100°C for 15 minutes.
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After cooling down samples were put on a NuPAGE ® 3-8% Tris-Acetate gel (1.0 mm x 12
well, cat no. EA03752BOX, Invitrogen, Carlsbad, California, USA) together with a Precision
PlusTM All Blue Protein Standards (cat #161-0373, Bio-Rad, Hercules, CA, USA). The gel was
run for 90 minutes; 30 minutes at 80 Volt and subsequently 60 minutes at 120 Volt.
Proteins were transferred to Immobilon-FL PVDF membrane (Merck Millipore, Billerica,
MA, USA) by western blotting for 1 hour at 125 V. The membrane was then rinsed with
destilled water and blocked overnight at 4 degrees celcius in blocking buffer containing a 1:1
mixture of TBS and Odyssey blocking buffer (LI-COR Biosciences, Lincoln, NE, USA). Multiple
FVIII epitopes were recognized by 1 µg/ml polyclonal sheep anti human FVIII (Ab20946,
Abcam, Cambridge, UK), which was detected by Alexa-680 anti-sheep IgG (cat no. A21102,
Invitrogen, Carlsbad, California, USA); Actin was recognized by 30 ng/ml mouse anti-actin
IgM (cat no. n350, Amersham, Buckinghamshire, UK), and subsequently detected by Alexa
680 anti mouse IgM (cat no. A21048, Invitrogen, Carlsbad, California, USA). All antibodies
were diluted in an antibody incubation buffer containing a 1:1 mixture of blocking buffer as
described above and TBS/1%Tween-20. Protein bands were visualized on Odyssey channel
700 with an Odyssey Infrared Imaging System, running Odyssey Infrared Imaging System
Application Software version 2.1.12 (LI-COR Biosciences, Lincoln, NE, USA).
5
Data analysis
Data are expressed as median ± interquartile range (IQR).
Results
FVIII is taken up by platelets
To determine if FVIII is present in platelets of healthy donors, platelets from 12 healthy
donors were isolated as described and APTT of the platelet lysate (50% FVIII deficient
plasma and 50% platelet lysate) and the control of washing (50% FVIII deficient plasma and
50% supernatant platelet pellet) were compared to a scaling sample (50% FVIII deficient
plasma and 50% HT buffer). The APTT of the wash controls (117.6 seconds (109.9-123.1))
did not differ from the scaling samples (119.3 seconds (111.6-123.1)), suggesting that all
plasma FVIII was removed by the process of platelet isolation. The addition of platelet
lysate to FVIII deficient plasma slightly shortens the APTT compared to the scaling sample
(107.4 seconds (100.9-112.0)), suggesting presence of trace amounts of FVIII in the platelets
of healthy donors (Figure 1A).
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Loading platelets with FVIII induced an additional shortening of the APTT of FVIII
deficient plasma. Before platelet isolation, PRP was divided into 6 samples and incubated
with HT buffer or 10 IU/ml Kogenate, Helixate NexGen, Advate, Refacto or Haemate P,
respectively. As in earlier experiments, platelets incubated with control buffer gave a slight
shortening of the APTT of FVIII deficient plasma compared to scaling samples; controls of
washing did not shorten the APTT compared to scaling samples (data not shown). Moreover,
when incubating samples with different brands of recombinant or human purified FVIII,
an additional shortening was observed for all types of recombinant and human plasma
purified FVIII/vWF concentrates added. Results are shown in Figure 1B.
To determine the speed of the process of FVIII uptake by platelets, a time-response
experiment was carried out with platelets from 4 donors. Platelets were incubated wit 10IU/
ml recombinant FVIII for 30, 60, 90 and 120 minutes. The uptake of FVIII already reached its
maximum within the first 30 minutes (Figure 1C). To determine the concentration yielding
the maximal uptake of FVIII, platelets of 3 donors were incubated with 1, 5 or 10 IU/ml
Kogenate respectively. When incubated with 5 to 10 IU/ml FVIII, the maximal uptake of FVIII
was reached (Figure 1D).
The presence of FVIII inside platelets was confirmed via SDS-PAGE and western blotting
(Figure 1E). Platelets of a healthy donor were incubated with either 10 IU/ml human purified
FVIII, 10 IU/ml recombinant FVIII (Kogenate) or HT buffer (control). Platelet lysates were
investigated with SDS-PAGE. A 0.1 IU/ml and 1 IU/ml human purified FVIII solution were put
on gel as a positive control. A polyclonal anti-FVIII antibody was used, recognizing multiple
epitopes of FVIII, giving rise to multiple bands on the gel. At 50 kD, the weight of the A1
domain of the FVIII heavy chain, a band is present in all platelet samples, which is not seen
in the positive controls, suggesting a part of the FVIII in platelets is cleaved. Around 80 kD
a band emerges in platelets incubated with human purified FVIII. This band, also present in
the positive control, indicates the FVIII light chain. Between 90 and 200 kD multiple bands
reflect the presence of full length and truncated forms of the FVIII heavy chain.
LRP is not involved in the uptake of FVIII
The low-density lipoprotein receptor-related protein (LRP) has been suggested to have a
role in FVIII metabolism. It binds to FVIII with a moderate affinity via which it is thought
to facilitate the clearance of FVIII from plasma. This receptor, also present on platelets,
can be blocked by the addition of receptor associated protein (RAP). To determine if LRP
on platelets transports plasma FVIII into platelets, we carried out an experiment in which
platelets were incubated with HT buffer or 10 IU/ml Kogenate in the presence and absence
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B. Incubation with FVIII
140
140
130
130
APTT - seconds
120
110
100
90
80
10
0
110
100
90
80
10
140
SS
co
PL ntr
(K ol P
og
L
PL en
at
(H
e)
el
PL ixa
t
(A e)
PL dva
t
PL (Re e)
(H fac
to
ae
)
m
at
e
P)
C
W
PL
SS
0
C. Time response
D. Dose response
140
130
80
/m
L
IU
10
5
IU
/m
L
70
10
0
/m
L
80
10
0
5
90
L
90
100
IU
100
110
1
110
120
co
nt
ro
lP
APTT - seconds
130
120
SS
co
nt
ro
lP
30
L
m
in
ut
60
es
m
in
ut
90
es
m
in
ut
12
es
0
m
in
ut
es
APTT - seconds
120
SS
APTT - secondes
A. FVIII in platelet lysates healthy donors
Figure 1. FVIII is platelet lysate.
(A) FVIII in platelet lysate healthy donors The APTT, displayed in seconds, was measured in 12 related
scaling samples (SS, 50% FVIII deficient plasma and 50% HT buffer), platelet lysates (PL, 50% FVIII
deficient plasma and 50% platelet lysate) and controls of washing (CW, 50% FVIII deficient plasma
and 50% supernatant platelet pellet). Bars display group median, error bars display interquartile
range, the horizontal line marks the median of scaling samples.
(B) Incubation with FVIII The APTT, displayed in seconds, was measured in 3 related scaling samples
(SS) and platelet lysates (PL) from platelets incubated with HT buffer (control PL) or 10 IU/ml Kogenate,
helixate, Advate, Refacto or Haemate P before isolation respectively. Bars display group median,
error bars display total range, the horizontal solid line marks the median of SS, the horizontal dotted
line marks the median of control PL.
(C) Time response The APTT, displayed in seconds, was measured in 4 related scaling samples (SS)
and platelet lysates (PL) from platelets incubated with HT buffer for 120 minutes (control PL) or
incubated with 10 IU/ml Kogenate for 30, 60, 90 or 120 minutes respectively. Bars display group
median, error bars display interquartile range, the horizontal solid line marks the median of SS, the
horizontal dotted line marks the median of control PL.
(D) Dose response The APTT, displayed in seconds, was measured in 3 related scaling samples (SS)
and platelet lysates (PL) from platelets incubated with HT buffer (control PL) or incubated with 1, 5,
or 10 IU/ml Kogenate respectively. Bars display group median, error bars display interquartile range,
the horizontal solid line marks the median of SS, the horizontal dotted line marks the median of
control PL.
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Figure 1. (E) SDS-PAGE and Western Blot
The first lane shows the Kd marker Precision PlusTM
All Blue Protein Standards. 10 and 100 times diluted
platelet lysates and the supernatant of their last
centrifugation steps are shown in lane 2-4, 5-7 and
8-10 respectively, with first the control platelets,
in the middle platelets incubated with 10 IU/
ml human purified FVIII (Aafact) and on the right
platelets incubated with 10 IU/ml recombinant FVIII
(Kogenate). The last two lanes contain 0.1 IU/ml and 1
IU/ml human purified FVIII respectively.
of RAP. Blocking of LRP did not reduce the platelet FVIII uptake (Figure 2A), suggesting that
LRP is not involved in the FVIII uptake by platelets.
FVIII is not taken up together with vWF
In plasma vWF is the carrier protein of FVIII. Experiments in which the uptake of different
types of FVIII concentrates was compared showed a higher FVIII uptake when the concentrate
contained both FVIII and vWF is added to platelets compared to concentrates containing
purely FVIII (Figure 1B). To further explore this finding, we investigated the influence of
vWF on platelet FVIII uptake. When adding 10 IU/ml Kogenate, together with 10 IU/ml
B. FVIII uptake in presence/absence of vWF
110
100
90
80
10
0
130
100
90
80
10
110
100
90
80
10
SS
0
co
co
nt
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nt
lP
ro
lP
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L
+
vW
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PL
PL
(K
og
(K
og
en
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at
e)
at
e)
+
vW
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SS
co
co
nt
ro
nt
lP
ro
lP
L
L
+
R
A
PL
P
PL
(H
ae
(H
ae
m
at
m
e
at
P)
e
P)
+
R
A
P
0
120
SS
120
140
110
APTT - seconds
APTT - seconds
130
APTT - seconds
C. dRGDW (block of vWF-GPIIbIIIa binding)
120
140
co
co
nt
nt
ro
ro
lP
lP
L
L
+
R
G
D
PL
PL
(K
(K
og
og
en
en
at
e)
at
e)
+
R
G
D
A. RAP (block of LRP)
Figure 2. FVIII uptake not mediated via LRP or vWF-platelet interaction
(A) The APTT, displayed in seconds, was measured in platelets from a healthy donor incubated with
either HT buffer (control PL) or 10 IU/ml Haemate P in the presence and absence of RAP, a protein
blocking the endocytic receptor LRP.
(B) The APTT, displayed in seconds, was measured in platelets from a healthy donor incubated with
either HT buffer (control PL) or 10 IU/ml Kogenate in the presence and absence of 10 IU/ml vWF.
(C) The APTT, displayed in seconds, was measured in platelets from 4 healthy donors incubated with
either HT buffer (control PL) or 10 IU/ml Kogenate in the presence and absence of dGRDW, a protein
blocking the platelet-vWF interaction.
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C. Platelets incubated with FVIII/vWF
110
110
90
80
70
m
at
m
e
at
P)
e
P)
(H
ae
+
(H
R
m
ae
G
at
m
D
e
at
P)
e
+
P)
C
+
R
R
P
G
D
+
C
R
P
(H
ae
SS
60
10
0
PL
PL
PL
60
10
0
100
(H
ae
SS
60
10
0
70
SS
70
80
(K
og
(K
en
og
at
en
e)
PL
at
e)
PL
(K
+
og
(K
R
G
en
og
D
at
en
e)
at
+
e)
C
+
R
R
P
G
D
+
C
R
P
80
90
PL
90
100
PL
100
APTT - seconds
120
110
APTT - seconds
120
PL
B. Platelets incubated with Kogenate
120
C
on
C
tr
on
ol
tr
PL
ol
P
L+
C
on
R
C
t
G
r
on
ol
D
tr
PL
ol
+
PL
C
R
+
P
R
G
D
+C
R
P
APTT - seconds
A. Control platelets
Figure 3. Preactivation of platelets increases FVIII uptake
The APTT, displayed in seconds, was measured in platelet lysates (PL) from 2 donors incubated with
HT buffer (A), 10 IU/ml Kogenate (B) or 10 IU/ml Haemate P (C) and for its related scaling samples (SS).
PRP was pre-incubated for 10 minutes with either CRP, dRGDW peptide (RGD) or the combination
of both. Also, a sample which was not pre-incubated was taken along. Bars display group median,
error bars display interquartile range, the horizontal solid line marks the median of SS, the horizontal
dotted line marks the median of PL of platelets which were not pre-activated.
vWF, no difference in FVIII uptake was observed to Kogenate alone (Figure 2B). However,
since this experience was performed with healthy donor PRP, donor vWF is present in all
experimental conditions. Therefore we have performed an additional experiment in which
the interaction between vWF and platelets is blocked by the addition of dRGDW peptide. If
vWF should play a role in FVIII uptake, one would expect a decreased uptake when blocking
the GPIIbIIIa receptor. Instead of a decrease, we observed a mild increase in FVIII uptake,
when adding dRGDW (Figure 2C).
5
Pre-activation of platelets increases uptake of free FVIII
dRGDW peptide, a potent inhibitor of the GPIIbIIIa-fibrinogen interaction, is known to
enhance GPVI induced platelet activation.7 To determine if platelet activation leads to an
enhanced of platelet FVIII uptake, platelets of 2 donors were pre-activated for 10 minutes
with either 0.01 ng/ml CRP or 200 µM dRGDW peptide, or the combination of both.
Thereafter platelets were incubated with HT buffer (control PL), 10 IU/ml Kogenate or 10
IU/ml Haemate P, and subsequently washed and lysed. As shown in Figure 3, pre-activation
of platelets with CRP and especially with the combination of CRP and dRGDW peptide,
significantly increased FVIII uptake when subsequently incubated with FVIII (Figure 3A, B).
However, when the added FVIII is bound to vWF, pre-activation of platelets is not able to
accomplish increased uptake (Figure 3C).
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A. FVIII infusion in severe hemophilia A
B. FVIII infusion in severe hemophilia A
C. Platelet FVIII after ≥3 days wash out
200
200
APTT - seconds
150
100
50
150
100
50
ol
s)
co
nt
r
(p
at
ie
nt
s
PL
PL
(h
ea
lth
y
r
r
af
te
af
te
es
es
60
m
in
ut
B
ef
or
e
15
m
in
ut
SS
)
0
0
SS
APTT - seconds
250
Figure 4. Platelet FVIII in severe hemophilia A patients
(A) The APTT, displayed in seconds, was measured in platelet lysates (PL, 50% FVIII deficient plasma
and 50% platelet lysate) form 3 severe haemophilia A patients before, 15 minutes and 60 minutes
after FVIII bolus infusion and in related scaling samples (SS, 50% FVIII deficient plasma and 50% HT
buffer). Bars display group median, error bars display interquartile range, the horizontal solid line
marks the median of SS, the horizontal dotted line marks the median of control PL.
(B) Platelets from a severe haemophilia A patient, from blood collected before, 15 minutes and
60 minutes after FVIII infusion, were isolated, lysed under reducing conditions and heated, and
subsequently processed via SDS-PAGE and western blotting. The first lane shows the Kd marker
Precision PlusTM All Blue Protein Standards. 10 times diluted platelet lysates are shown in lane 2-4.
The last lane shows 1 IU/ml Advate, the recombinant FVIII which was infused in this patient.
(C) The APTT, displayed in seconds, was measured in platelet lysates (PL, 50% FVIII deficient plasma
and 50% platelet lysate) from 33 severe haemophilia A patients after a washout period of 3 days and
in PL form 5 healthy donors, and in related scaling samples (SS, 50% FVIII deficient plasma and 50%
HT buffer). Bars display group median, error bars display interquartile range, the horizontal solid line
marks the median of SS, the horizontal dotted line marks the median of control PL.
FVIII can be found in platelets of hemophilia A patients treated with FVIII
To determine if FVIII is present or absent in platelet of haemophilia A patients, platelet
lysates from 3 severe hemophilia A patients were measured before, 15 minutes and
60 minutes after FVIII bolus infusion. The platelet lysate did shorten the APTT of FVIII
deficient plasma, both before and after infusion (Figure 4A). Plasma FVIII activity in these
patient were 4.5%, 1.4% and <1% before infusion and 112%, 113% and 95% after infusion.
Via SDS-PAGE and western blotting, FVIII was also determined to be present both before
and after infusion (Figure 4B).
Since these patient had received FVIII concentrate profylacticly in the days preceding
inclusion, we also included 33 severe hemophilia A patients from an earlier study, in which
all patients did not receive FVIII for at least 3 days before inclusion. When measuring APTT
of all platelet lysates, again a shortening in APTT was observed, suggesting presence of
FVIII. When comparing severe hemophilia A patients to 5 healthy controls from this study,
platelet lysates of healthy controls contained more FVIII.
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FVIII is also taken up by megakaryocytes
Finally we investigated if megakaryocytes were also able to take up FVIII. Therefore we
used a CHRF cell line, which hold characteristics of mature megakaryoctes. A suspension
of 1*106 megakaryocytes/ml was incubated with 10 IU/ml Kogenate, Helixate, Refacto,
Haemate P or HT buffer respectively. After 2 hours incubation, cells were isolated and lysed.
The lysate of control cells did not shorten the APTT, suggesting that FVIII was absent in
this cell line. When incubated with Kogenate, Helixate, Refacto or Haemate P respectively,
APTT was shortened compared to scaling and control samples, suggesting FVIII uptake by
megakaryocytes. Results are shown in Figure 5A.
When determining presence of FVIII via SDS-PAGE and western blot, FVIII proved to be
absent in control cells. CHRF cells incubated with either human purified or recombinant
FVIII did contain variable FVIII levels (Figure 5B). Haemate P, a combination of human
purified FVIII and vWF, could not be found in CHRF by western blotting after incubation.
5
Discussion
In this study, we showed the presence of FVIII in platelets and a megakaryocytic cell line
using APTT measurements and confirmed the presence of FVIII through western blotting.
Trace amounts of FVIII are present in platelets of healthy donors, and its presence can be
increased by incubating them with (recombinant) FVIII, especially with activated platelets.
FVIII is also present in platelets of severe hemophilia A patients, even when prophylactic FVIII
treatment was withheld for a minimum of 3 days before blood collection, suggesting that in
vivo uptake might take place at an earlier cell stage. Experiments with the megakaryocytic
cell line CHRF, shows megakaryocytes are also able to take up FVIII.
APTT was used to functionally determine presence of FVIII in cell lysates. FVIII deficient
plasma was mixed 1:1 with the lysates to ensure all factors present in FVIII deficient plasma
were still present in a 50% concentration. By doing so, the absence of FVIII was the only rate
limiting element in this assay. Therefore, the shortening in APTT noticed after addition of
platelet lysates must be caused by FVIII, and cannot be the result of other platelet proteins.
Moreover, when incubating the platelets with purified FVIII before isolation and lysation,
the observed effect increases in a dose and time dependent matter, suggesting the added
FVIII is responsible for the observed shortening in APTT.
To confirm that the observed effects on APTT were caused by FVIII, we did a SDS-PAGE
and western blot analysis of platelet and megakarycote lysates. Full length FVIII circulates
in plasma as a heterodimeric molecule, with a molecular weight of 280 kD. The FVIII heavy
FVIII uptake by platelets and megakaryocytes | 79
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200
180
160
140
120
100
80
60
40
20
0
SS
co
nt
ro
PL
lP
L
(K
og
en
at
PL
e)
(H
el
ix
at
PL
e)
(R
e
fa
PL
ct
(H
o)
ae
m
at
e
P)
APTT - secondes
A. CHRF incubated with FVIII
B. Western blot CHRF lysate
Figure 5. FVIII uptake by megakaryocytes
(A) The APTT, displayed in seconds, was measured in 3 related scaling samples (SS) and lysates from
CHRF-288-11 cells incubated with either HT buffer (control CHRF) or 10 IU/ml Kogenate, Helixate
NexGen, Refacto or Haemate P before isolation, respectively. Bars display group median, error bars
display total range, the horizontal solid line marks the median of SS, the horizontal dotted line marks
the median of control CHRF. n=2.
(B) The first lane shows the Kd marker Precision PlusTM All Blue Protein Standards. CHRF cell lysates
and the supernatant of their last centrifugation steps are shown in lane 2-7 and 8-12 respectively.
Cells were incubated with HT buffer (control CHRF) or 10 IU/ml Aafact, Kogenate, Helixate, Refacto
or Haemate P. In the upper section, FVIII was detected. In the lower section, actin was detected as
a loading control.
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chain, consisting of the A1-A2-B domains, has a molecular weigth of 200 kD. However,
differentially truncated B-domain variants can lead to a decreased weight between 97200 kD, and when reduced to a combination of the A1-A2 domain or a single A1 domain,
weight is further decreased to 93 kD or 50 kD, respectively. The FVIII light chain, consisting
of the A3-C1-C2 domains, weights 80 kD.8 However, due to glycosylation, the observed
molecular weight can be higher. For example, the molecular weight of the full length
protein can vary from 280 up to 360 kD.9,10 By making use of a polyclonal anti-FVIII antibody,
multiple compositions of FVIII could be traced in both platelet lysates and megakaryocytes.
The endocytic receptor low density lipoprotein receptor-related protein (LRP) is known
to bind variable proteins including FVIII for transport into the intracellular degredation
pathway.11,12 This receptor has been proposed to regulate plasma levels of FVIII in vivo.13
In mouse models, absence of this receptor has shown to lead to increased plasma levels of
FVIII.14 Therefore we wondered if LRP could be involved in platelet FVIII uptake, which would
suggest uptake to be relevant for regulation of plasma FVIII levels. However, we found that
blocking LRP with RAP did not influence the amount of FVIII taken up by platelets.
Next we also looked into a possible role for vWF in platelet FVIII uptake, since
experiments showed that incubation with 10 IU/ml Haemate P leads to more platelet
FVIII compared to incubation of 10 IU/ml of the other FVIII concentrates, which did not
contain vWF. However, when incubating platelets with recombinant FVIII in the presence
and absence of vWF, no differences in uptake could be observed. Moreover, when blocking
platelet-vWF interaction, FVIII was still taken up by platelets in equal amounts. Another
difference between Haemate P and the other FVIII concentrates used, is that Haemate P
consists of human purified FVIII whereas the other concentrates consist of recombinant
FVIII. Possible, differences between FVIII protein, such as different glycosylation profiles,
could account for the differences observed in uptake.
Pre-activation of platelets did increases the uptake of FVIII. Platelets were pre-incubated
with either a very low concentration of CRP, dRGDW peptide or the combination of both.
Only a very low concentration of CRP could be used, since higher concentrations lead
to platelet aggregation making it impossible to subsequently wash and isolate platelets.
The increased uptake observed after pre-activation of platelets implicates a role for an
activation dependent receptor or for phospholipids in the mechanism of platelet FVIII
uptake.
To evaluate if platelets also take up FVIII in vivo, we investigated platelets of patients
with severe haemophilia A before and after FVIII concentrate infusion. We found in these
patients that FVIII was already present before FVIII infusion, and did not increase after
infusion. This platelet FVIII could reflect platelet uptake from prophylactic FVIII use in
5
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preceding days. Furthermore, these patients did not undergo a wash out period of plasma
FVIII, resulting in measurable FVIII in 2 out of 3 patients before FVIII concentrate infusion.
Therefore we also performed FVIII measurements in patients which did not receive FVIII
for at least 3 days prior to blood collection. This 3 day period was established based on
the knowledge that FVIII is almost completely cleared from plasma within 3 days after
infusion. Platelets from these patients also proved to contain FVIII, although less compared
to platelets from healthy controls. In view of a platelet half-life of 3 days, a washout period
of 10 days would have been ideal, however, this was not possible since this would put
the patients at risk for bleeding. Consequently, FVIII measured in platelet lysates from
severe haemophilia A patients could still be a product of platelet FVIII uptake during the
days preceding the wash out period. However, it could also indicate FVIII is taken up by
megakaryocytes.
A limit to this study is that the techniques used to determine the presence of FVIII in
platelets do not determine the location of FVIII within the cell. Therefore, we do not know
if FVIII is present in for example the platelet granules or its open canicular system. Future
research will be necessary to determine the location of FVIII in platelets, which could give
more insight into both the cellular mechanism involved in this uptake and the possibility for
FVIII to be released when platelets are activated at a side of vascular damage.
In summary, the results of this study show that platelets and megakayocytes are able to
take up FVIII, a process which is accelerated by pre-activation of platelets. Furthermore, we
show in severe haemophilia A patients that FVIII is taken up in vivo, possibly at the level of
megakaryocytes. Further studies are needed into the cellular mechanisms involved in this
uptake and its relevance in hemostasis.
References
1.
2.
3.
4.
Manco-Johnson MJ, Abshire TC, Shapiro AD, Riske B, Hacker MR, Kilcoyne R, et al. Prophylaxis
versus episodic treatment to prevent joint disease in boys with severe hemophilia. N Engl J
Med. 2007 9;357(6):535-44.
Collins PW, Björkman S, Fischer K, Blanchette V, Oh M, Schroth P, et al. Factor VIII requirement
to maintain a target plasma level in the prophylactic treatment of severe hemophilia A:
influences of variance in pharmacokinetics and treatment regimens. J Thromb Haemost. 2010;
8: 269-75.
Ahnström J, Berntorp E, Lindvall K, Björkman S. A 6-year follow-up of dosing, coagulation factor
levels and bleedings in relation to joint status in the prophylactic treatment of haemophilia.
Haemophilia. 2004 Nov;10(6):689-97.
Björkman S. Prophylactic dosing of factor VIII and factor IX from a clinical pharmacokinetic
perspective. Haemophilia. 2003 May;9 Suppl 1:101-8; discussion 109-10.
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5.
6.
7.
8.
9.
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12.
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14.
Fugman DA, Witte DP, Jones CLA, Aronow BJ, Lieberman MA. In vitro establishment and
characterization of a human megakaryoblastic cell line. Blood. 1990;75:1252-61.
van der Vuurst H, van Willigen G, van Spronsen A, Hendriks M, Donath J, Akkerman JW. Signal
transduction through trimeric G proteins in megakaryoblastic cell lines. Arterioscler Thromb
Vas Biol. 1997;17(9):1830-6.
Jones ML, Harper MT, Aitken EW, Williams CM, Poole AW. RGD-ligand mimetic antagonists
of integrin αIIbβ3 paradoxically enhance GPVI-induced human platelet activation. J Thromb
Haemost. 2010;8:567-76.
Manning F, Fágáin CO, O’Kennedy R. Factor VIII: Function, structure and analyses. Biotech Adv.
1993;11:79-114.
D’Amici GM, Blasi B, D’Alessandro A, Vaglio S, Zolla L. Plasma-derived clotting factor VIII:
Heterogeneity evaluation in the quest for potential inhibitory-antibody stimulating factor.
Electrophoresis. 2011;32(21):2941-50.
Lenting PJ, van Mourik JA, Mertens K. The life cycle of coagulation factor VIII in view of its
structure and function. Blood. 1998; 92(11):3983-96.
Lenting PJ, Neels JG, van den Berg BMM, Clijsters PPFM, Meijerman DWE, Pannekoek H et al.
The Light Chain of Factor VIII Comprises a Binding Site for Low Density Lipoprotein Receptorrelated Protein. The Journal of Biological Chemistry. 1999;274(34):23734-39.
Saenko EL, Yakhyaev AV, Mikhailenko I, Strickland DK, Sarafanov AG. Role of the Low Density
Lipoprotein-related Protein Receptor in Mediation of Factor VIII Catabolism. The Journal of
Biological Chemistry. 1999;274(53):37685-92.
Bovenschen N, Mertens K, Hu L, Havekes LM, van Vlijmen BJM. LDL receptor coorperates
with LDL receptor-related protein in regulating plasma levels of coagulation factor VIII in vivo.
Blood. 2005 106: 906-912.
Bovenschen N, Herz J, Grimbergen JM, Lenting PJ, Havekes LM, Mertens K, van Vlijmen BJM.
Elevated plasma FVIII in a Mouse model of low-density lipoprotein receptor-related protein
deficiency. Blood. 2003;101:3933-39.
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Chapter 6
Platelets of patients with chronic kidney disease
demonstrate deficient platelet reactivity in vitro
Esther R. van Bladel,1* Rosa L. de Jager,2* Daisy Walter,1 Loes Cornelissen,1 Carlo A. Gaillard,2,3
Leonie A. Boven,2 Mark Roest,1 Rob Fijnheer1,2
1
Department of Clinical Chemistry and Hematology, University Medical Center
Utrecht, Heidelberglaan 100, 3584 CX Utrecht, The Netherlands; 2Department
of Internal Medicine and Laboratory Medicine, Meander Medical Center,
Utrechtseweg 160, 3800 BM Amersfoort, The Netherlands; 3Department of
Nephrology, VU University Medical Center, Amsterdam, The Netherlands.
*equally contributed
BMC Nephrology 2012;13:127.
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Abstract
Background: In patients with chronic kidney disease studies focusing on platelet function
and properties often are non-conclusive whereas only few studies use functional platelet
tests. In this study we evaluated a recently developed functional flow cytometry based
assay for the analysis of platelet function in chronic kidney disease. Methods: Platelet
reactivity was measured using flow cytometric analysis. Platelets in whole blood were
triggered with different concentrations of agonists (TRAP, ADP, CRP). Platelet activation
was quantified with staining for P-selectin, measuring the mean fluorescence intensity.
Area under the curve and the concentration of half-maximal response were determined.
Results: We studied 23 patients with chronic kidney disease (9 patients with cardiorenal
failure and 14 patients with end stage renal disease) and 19 healthy controls. Expression of
P-selectin on the platelet surface measured as mean fluorescence intensity was significantly
less in chronic kidney disease patients compared to controls after maximal stimulation
with TRAP (9.7 (7.9-10.8) vs. 11.4 (9.2-12.2), P=0.032), ADP (1.6 (1.2-2.1) vs. 2.6 (1.9-3.5),
P=0.002) and CRP (9.2 (8.5-10.8) vs. 11.5 (9.5-12.9), P=0.004). Also the area under the curve
was significantly different. There was no significant difference in half-maximal response
between both groups. Conclusion: In this study we found that patients with chronic kidney
disease show reduced platelet reactivity in response of ADP, TRAP and CRP compared to
controls. These results contribute to our understanding of the aberrant platelet function
observed in patients with chronic kidney disease and emphasize the significance of using
functional whole blood platelet activation assays.
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Introduction
In chronic kidney disease both bleeding and thrombotic complications are observed. It has
been hypothesized that this disturbed balance between pro- and anti-haemostatic factors
is involved in the high morbidity and mortality reported in chronic kidney disease.1-4 Early
stages of chronic kidney disease are typically associated with a prothrombotic tendency,
whereas in its more advanced stage patients also suffer from a bleeding diathesis.5
Bleeding tendency of patients is characterized by haemorrhagic symptoms and by
prolongation of bleeding time.6 The cause of bleeding in this group of patients has been
elaborated in the past and the pathogenesis seems multifactorial. It is suggested that
abnormal platelet function is a major contributor, since haemorrhage occurs despite a
coagulation profile of normal or elevated levels of coagulation factors and normal platelet
counts.6,7 Platelet dysfunction may be the result of decreased dense granule content,
decreased sensitivity to platelet agonists, abnormal expression of platelet glycoproteins,
defective arachidonate metabolism and depressed prostaglandin metabolism as well
as impaired platelet adhesiveness. Platelet dysfunction is thought to be caused by the
action of uremic toxins, anemia, increased nitric oxide production, von Willebrand factor
abnormalities and the use of medication like aspirin, non-steroidal anti-inflammatory drugs
and β-lactam antibiotics.6-9 Besides an increased bleeding risk, a variety of thrombotic
complications are observed in patients with chronic renal failure, including coronary heart
disease, cerebrovascular disease, peripheral vascular disease and heart failure. Already in
mild to moderate chronic kidney disease an increased risk of cardiovascular events and
higher mortality have been reported.1,2,10-13
The balance between thrombosis and bleeds is disturbed in chronic kidney disease.
Functional platelet tests are needed to obtain more insight into this balance. However,
up to know only a minority of the studies used functional platelet tests and results are
non-conclusive.6,14 This is probably due to in vitro artifacts as a result of the procedure
of platelet isolation, influence of plasma proteins and influence of hematocrit. Therefore
we have set up this study with a recently developed, flow cytometer based, functional
platelet assay in whole blood, in which these factors are not of influence. With this assay
we assessed platelet reactivity in patients with end stage renal disease and cardiorenal
syndrome. Both platelet sensitiveness as well as maximal activation of platelets in response
to 3 different platelet stimuli was measured, providing an explanation for the disturbed
haemostatic balance in patients with renal failure.
6
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Methods
Subjects
Patients were selected from the out-patient clinic of the Meander Medical Center in
Amersfoort for this prospective, observational study. Two groups of patients with chronic
kidney disease were recruited. The first group consisted of patients with end stage renal
disease receiving hemodialysis.15 In patients on hemodialysis, blood samples were collected
before and also immediately after hemodialysis (pre versus post dialysis). All samples were
collected using citrate tubes and mixed gently. The second group consisted of patients with
cardiorenal syndrome, defined as coexistence of chronic kidney disease and chronic heart
failure.16 Chronic kidney disease in this group was defined as eGFR < 70 ml/min without
requirement of hemodialysis. Chronic heart failure was defined as NYHA class II or higher,
based on symptoms, signs and objective abnormality on echocardiography.17 Controls
included subjects recruited from among healthy hospital staff, healthy subjects attending
the hospital for control follow up and healthy partners of patients. Patients were excluded
from the study based on the following criteria: clinical signs of infection, malignancy,
primary haemostatic disorders unrelated to uremia, treatment with immunosuppressive
drugs, use of antiplatelet agents (except aspirin) such as clopidogrel, dipyridamole or
non-steroidal anti-inflammatory drugs and the inability to provide informed consent. The
protocol was approved by the local medical ethical committee of Meander Medical Center
Amersfoort and both patients and control volunteers gave written informed consent to
participate in the study.
Blood sampling and processing
In patients on haemodialysis, the samples were collected before starting the haemodialysis
and immediately after the procedure. The first sample was taken from the afferent line
directly after inserting the needle in the fistula. The second sample was taken directly after
dialysis from the efferent line out of the dialysis machine. In control subjects peripheral
venous blood samples were collected from the antecubital vein using 21 gauge needles. A
total of 4.5 mL blood was drawn into vacutainer tubes containing 0.5 mL 3.2% sodium-citrate
solution as anticoagulant and mixed gently. Blood cell count assays were performed using
a haematology analyzer (Sysmex, Etten-Leur, The Netherlands). Hemoglobin, hematocrit,
and platelet count were noted.
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Materials
For our samples we used HEPES-buffered saline (HBS) containing 10mM HEPES (BDH
biochemical, UK), 150 mM NaCl (Sigma-Aldrich, Zwijndrecht, the Netherlands), 1mM
MgSO4 (Riedel de Haën, Hannover, Germany) and 5mM KCl (Riedel de Haën), with a pH of
7.4. Fixation was done with a fixation buffer containing 0.2% formaldehyde (Calbiochem,
Merck, Darmstadt, Germany) in 0.9% NaCL (Sigma-Aldrich). Used fluorochrome-labeled
ligands were anti-CD42b fluorescein isothiocyanate (FITC)-labeled mouse antihuman
antibody (BD Pharmingen™, San Diego, California, USA) and anti-CD62P phycoerythrin
(PE)-labeled mouse antihuman antibody (BD Pharmingen™). Adenosine diphosphate (ADP)
(Roche, Almere, the Netherlands) was used as agonist, as well as thrombin-receptorassociated-peptide (TRAP) (Bachem, Weil am Rhein, Germany) and cross-linked collagen
related peptide (CRP) was a generous gift of R. Farndale (Cambridge, United Kingdom).
Platelet reactivity
Assays were performed as described before.18 In short, 5 μL of whole blood was added
to tubes containing 50 μL of HBS, fluorochrome-labeled ligands and serial dilutions of
agonists. Anti-CD42b FITC-labeled monoclonal antibodies were used as an activationindependent platelet marker. PE-labeled anti-CD62P was used as an activation-dependent
marker. For each agonist eight different concentrations were used, with four times dilution
steps between each sample. Concentrations of TRAP were 0.038 to 625 μmol/L, ADP 0.008
to 125 μmol/L, and CRP 0.2 to 2500 ng/mL. After incubation at room temperature for
20 minutes, platelets were fixed by the addition of 500μL fixative (0.2% paraformaldehyde).
Subsequently, samples were 3.5 times diluted with the same fluid for flow cytometric
analysis. Samples were analyzed by flow cytometry on a Epics XL-MCL Flow Cytometer
(Beckman Coulter, Miami, Florida, USA) and EXPO 32 MultiCOMP Software (Beckman
Coulter) was used to process the data. The platelet population was identified by forward
and 90° side scatter properties in combination with a positive CD42b signal. Isotype control
antibodies were used to correct for aspecific binding. The mean fluorescence intensity
(MFI) of all platelets is expressed in arbitrary units (AU).
6
Statistical Analysis
GraphPad software version 4.0 for Windows (GraphPad Software, San Diego, California
USA) was used to draw graphs and to calculate EC50. EC50 is defined as the concentration
of the agonist needed to achieve an effect on the platelets halfway between the maximum
and minimum. Area under the curve (AUC) was calculated in SPSS by adding up the outcome
minus basal activation outcome of all concentrations points. The data were analyzed by
Platelets of patients with chronic kidney disease demonstrate deficient platelet reactivity in vitro | 89
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logistic regression for scaled data. For comparisons of nominal variables a Chi-square cross
tabulation with a Fisher’s Exact test was used. Statistical analyses were undertaken using
SPSS software version 15.0 (SPSS Inc, Chigaco, Illinois). A P-value of <0.05 was considered
as statistically significant. Values are given as median with interquartile range (IQR) if not
noted otherwise.
Results
Patient characteristics
Twenty-three patients with chronic kidney disease and 19 healthy controls were included.
In the chronic kidney disease group, we included 9 patients with cardio renal syndrome
and 14 patients with end stage renal failure on hemodialysis (analyzed before and
immediate after dialysis). Clinical and laboratory characteristics are listed in Table 1. Age
was significantly higher in patients (78 years (64-83)) compared to controls (62 years (4871)). As expected hemoglobin and hematocrit level were significantly lower in cardio renal
syndrome patients compared to controls. There were no significant differences in platelet
count. The etiology of chronic kidney disease was divers: hypertension (n=7), diabetic
nephropathy (n=4), membranous glomerulopathy (n=1), rapid progressive IgA nephropathy
(n=1), chronic pyelonephritis (n=1), polycystic nephropathy (n=1), nephrosclerosis (n=1) or
unknown cause (n=7). The etiology of chronic heart failure was of ischemic origin (n=5),
valvular heart disease (n=3) or unknown (n=1).
Platelet reactivity
Expression of P-selectin on the platelet surface measured as MFI was significantly lower in
chronic kidney disease patients compared to controls after maximal stimulation with TRAP
(9.7 (7.9-10.8) vs. 11.4 (9.2-12.2), p=0.032), ADP (1.6 (1.2-2.1) vs. 2.6 (1.9-3.5), p=0.002) and
CRP (9.2 (8.5-10.8) vs. 11.5 (9.5-12.9), p=0.004). (Figure 1 and Table 2). Maximal P-selectin
expression in response to different agonist correlated significantly with each other
(Spearmann’s Rho correlation TRAP and ADP 0.684 with p<0.001; TRAP and CRP 0.538 with
p<0.001; ADP and CRP 0.475 with p=0.001). Table 2 shows that the EC50 was not different
between chronic kidney disease patients and controls.
Chronic kidney disease patients showed significantly lower platelet reactivity as
compared to healthy control subjects for TRAP (AUC: 39.5 (32.9-44.4) vs. 47.1 (41.1-54.2),
p=0.01) and ADP (AUC: 5.8 (3.8-7.3)vs. 8.9 (7.9-12.2), p=0.002) when measured with
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Table 1. Baseline characteristics in patients with chronic kidney disease and controls
Baseline characteristics
Controls
N=19
CKD
N=23
P-value
Age, y (IQR)
Male, n (%)
Hemoglobin, mmol/L (IQR)
Hematocrit, L/L (IQR)
Platelet count, 109/L (IQR)
Hemodialysis, n (%)
Cardiorenal syndrome, n (%)
EPO use, n (%)
Aspirin use, n (%)
62 (48-71)
13 (68.4)
9.4 (9.0-9.9)
0.44 (0.43-0.47)
210 (150-240)
NA
NA
NA
0
778 (64-83)
15 (65.2)
7.2 (6.7-7.8)
0.36 (0.34-0.38)
187 (160-225)
14 (60.9)
9 (39.1)
5 (21.7)
11 (47.8)
0.043
0.755
0.006
0.001
0.223
-
CKD chronic kidney disease, IQR inter quartile range, EPO erythropoietin
15
2
1
0
10 -1
10 0
10 1
TRAP - uM
10 2
0
10 -3
10 3
E. TRAP - AUC
10 0
ADP - uM
10 1
10 2
0
10 -2
10 3
10
5
20
10 3
6
10 4
30
20
C
on
tr
ol
s
0
D
C
on
tr
ol
s
C
K
10 2
10
0
D
0
10 1
CRP - ng/ml
40
AUC - AU
AUC - AU
40
10 0
p = 0.06
50
15
60
10 -1
F. CRP - AUC
p = 0.002
20
80
AUC - AU
10 -1
5
D. ADP - AUC
p = 0.01
100
10 -2
C
K
10 -2
10
C
on
tr
ol
s
5
CKD
Controls
D
10
MFI - AU
3
MFI - AU
MFI - AU
C. CRP - MFI of platelets
B. ADP - MFI of platelets
4
C
K
A. TRAP - MFI of platelets
15
Figure 1. Platelet reactivity
(A) The mean fluorescence intensity (MFI) of all platelets, expressed in arbitrary units (AU) and
(B) area under the curve (AUC), displayed as median plus interquartile range, for chronic kidney
disease patients (CKD) and controls.
Platelets of patients with chronic kidney disease demonstrate deficient platelet reactivity in vitro | 91
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AUC (Table 2). Stimulation with CRP (AUC: 30.3 (22,1-41,9) vs 23.4 (18.9-34.7), p=0.06) does
not show a difference in reactivity between the groups.
Since age was significant lower in the control group we adjusted for this in a regression
analysis. The significant difference in MFI and AUC between chronic kidney disease patients
and controls after maximal stimulation remained significant (Table 2). There was no
significant difference between patients with cardiorenal failure and patients with endstage
renal failure.
In the chronic kidney disease group almost half of the patients used aspirin as antiplatelet therapy. Platelet reactivity in this group does not show a difference between
aspirin users and patients without aspirin. This accounts for all outcome measurements.
Table 2. Results platelet activation
Maximal MFI TRAP, AU (IQR)
Maximal MFI ADP, AU (IQR)
Maximal MFICRP, AU (IQR)
EC50 TRAP, μM (IQR)
EC50 ADP, μM (IQR)
EC50 CRP, ng/ml (IQR)
AUC TRAP, AU (IQR)
AUC ADP, AU (IQR)
AUC CRP, AU (IQR)
Controls
N = 19
CKD
N = 23
P-value
P-value
Adjusted for age
11.4 (9.2-12.2)
2.6 (1.9-3.5)
11.5 (9.5-12.9)
1.9 (1.5-2.5)
0.9 (0.6-1.1)
120 (46.7-312.7)
47.1 (41.1-54.2)
8.9 (7.9-12.2)
30.3 (22.1-41.9)
9.7 (7.9-10.8)
1.6 (1.2-2.1)
9.2 (8.5-10.8)
2.3 (1.8-2.6)
0.89 (0.7-1.1)
116.1 (46.1-270.5)
39.5 (32.9-44.4)
5.8 (3.8-7.3)
23.4 (18.9-34.7)
0.032
0.002
0.004
0.372
0.544
0.955
0.012
0.002
0.063
0.095
0.004
0.010
0.182
0.597
0.660
0.023
0.003
0.322
MFI Mean fluorescence intensity, AU arbitrary units, IQR inter quartile range, EC50 concentration of
agonist leading to a half maximal P-selectin expression, AUC Area under the curve
Pre-versus-post dialysis
To rule out an activating effect of the hemodialysis procedure on baseline platelet activation
we studied platelet activation before and immediate after dialysis. The platelet sensitivity
(EC50) for TRAP, ADP and CRP did not differ between patients pre- and postdialysis. There
was no difference between the maximum reached effect and AUC on platelet activation
when comparing patients pre- and postdialysis (data not shown).
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Discussion
The aim of this study was to investigate the relation between platelet function and
kidney failure in patients with end stage renal disease and cardiorenal syndrome using
a flow cytometer based, functional platelet assay. We show that P-selectin expression
after stimulation with ADP, CRP and TRAP is lower in patients with chronic kidney disease
as compared to healthy controls. Furthermore, we demonstrate that the EC50 was not
different between groups. This means that platelet sensitivity itself is not affected for the
different agonists, but the maximal platelet response is significantly lower.
Platelets were activated with 3 different stimuli. We chose for the agonists TRAP, ADP
and CRP, which stimulate the three major physiological platelet activation pathways. TRAP
activates the thrombin receptor Proteinase Activated Receptor 1 (PAR-1) on platelets. ADP
is normally present in platelet dense granules. Upon platelet activation, ADP is released to
activate nearby platelets via the P2Y receptors. CRP activates the receptor glycoprotein VI,
the major collagen receptor on platelets. So, all 3 stimuli are of physiological importance.
P-selectin is found in the α-granula of platelets. A deficiency in α-granula could lead to
ineffective haemostasis. Two different explanations are possible for the deficient platelet
α-granula release found in chronic kidney disease. This could be due to depletion of
α-granula itself, or due to a deficiency in the release of α-granula. An impaired α-granule
release has already been reported in uraemia.19 This is further supported by the recent
observations of Schoorl et al. of an increase in platelets depleted from granules in patients
undergoing chronic haemodialysis.20 Nevertheless, we cannot exclude an impairment of
release.
The clinical bleeding tendency in uremic patients received much attention.21 Haemo- or
peritoneal dialysis was found to improve haemostasis without correcting platelet aggregation
defects. In contrast, compensatory mechanisms in the form of high von Willebrand factor
(VWF) levels preserved relatively normal adhesion of uremic platelets to injured vessel wall
models.22 Moreover, as soon as the hematocrit - determining the red blood cell-mediated
radial transport of blood platelets towards the vessel wall - was corrected by recombinant
erythropoietin, clinical bleeding became less prominent. In contrast, thromboembolic/
cardiovascular morbidity remains an important problem in these patients.23 Especially
in cardiorenal failure, there seems no or little benefit from treatment with antiplatelet
agents, whilst risk of bleeding may increase.24 Even though reports on platelet reactivity
in patient with chronic kidney disease show conflicting results,25 it has been attributed
to an acquired thrombocytopathy characterized by decreased aggregation of platelets in
response to stimuli. There are only a few studies that used functional tests to study platelet
6
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function in chronic kidney disease. Aggarwal et. al. found a higher P-selectin expression in
patients with end stage renal disease receiving haemodialysis compared to healthy controls
after stimulation with a single concentration of ADP (0.2µM). This suggests an increased
reactivity.14 Moal et. al performed a similar study in which ADP (200 µM) and TRAP (50µM)
in a single concentration were used to stimulate platelets in healthy controls and end stage
renal disease patients receiving haemodialysis. They found a lower P-selectin expression
in patients compared to controls, indicating reduced platelet reactivity in patients with
chronic kidney disease.6 Most assays used previously are influenced by in vitro artifacts,
platelet isolation and are dependent on VWF or hematocrit. In our functional assay there is
no role for VWF and hematocrit. Moreover, since fresh whole blood was used, and samples
were fixated subsequent to stimulation, in vitro platelet activation was negligible. We
couldn’t find an immediate effect of haemodialysis, but long term dialysis and classical risk
factors for athero-vascular disease could all be studied with this assay. Larger populations
should be studied in order to find the factors influencing platelet reactivity in vivo.
In our study, aspirin did not suppress the expression of P-selectin on platelets.
Considering the fact that aspirin function is related to inhibition of tromboxane formation
and does not influences release of granules, this is an expected result. Moreover, our finding
is in accordance with a study by Stumpf et. al., in which P-selectin expression on platelets
did not show a difference between patients taking aspirin compared to non-aspirin users.26
Further investigation is required to determine the influence of different drugs on platelet
reactivity.
We here demonstrate that the flow cytometry based platelet activation assay can be
used in clinical practice to study variables influencing platelet response in patients with
renal failure.
The assay is based on incubation with different agonists in whole blood and subsequent
fixation. Platelets are sensitive for in vitro artifacts (especially platelet isolation, shear stress,
pH and temperature). In our assay platelets are not isolated. The incubation in whole blood
under standardized condition and the subsequent fixation step makes in vitro artifacts
unlikely. When repeating the assay within healthy subjects at different blood collection
moments, we find high reproducibility. Moreover, this assay can be used in routine clinical
practice.
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Conclusion
In conclusion, we found that patients with chronic kidney disease show reduced platelet
reactivity in response of ADP, TRAP and CRP compared to controls. The defect is probably
due to an α-granule defect. Further investigation is required to determine a correlation
between platelet reactivity and both renal function and the high mortality risk of patients
with chronic kidney disease.
References
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AB. Renal insufficiency as a predictor of cardiovascular outcomes and mortality in elderly
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Go AS, Chertow GM, Fan D, McCulloch CE, Hsu CY. Chronic kidney disease and the risks of
death, cardiovascular events, and hospitalization. N Engl J Med. 2004;351:1296-305.
Sarnak MJ, Levey AS, Schoolwerth AC, Coresh J, Culleton B, Hamm LL, McCullough PA, Kasiske
BL, Kelepouris E, Klag MJ, Parfrey P, Pfeffer M, Raij L, Spinosa DJ, Wilson PW. Kidney disease
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Kaw D, Malhotra D. Platelet dysfunction and end-stage renal disease. Semin Dial. 2006;19:31722.
Jalal DI, Chonchol M, Targher G. Disorders of hemostasis associated with chronic kidney
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Moal V, Brunet P, Dou L, Morange S, Sampol J, Berland Y. Impaired expression of glycoproteins
on resting and stimulated platelets in uraemic patients. Nephrol Dial Transplant. 2003;18:183441.
Kaw D, Malhotra D. Platelet dysfunction and end-stage renal disease. Semin Dial. 2006;4:31722.
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Rios DR, Carvalho MG, Lwaleed BA, Simoes e Silva AC, Borges KB, Dusse LM. Hemostatic
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Collins AJ, Li S, Gilbertson DT, Liu J, Chen SC, Herzog CA. Chronic kidney disease and
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Muntner P, He J, Hamm L, Loria C, Whelton PK. Renal insufficiency and subsequent death
resulting from cardiovascular disease in the United States. J Am Soc Nephrol. 2002;3:745-53.
Nitta K. Pathogenesis and therapeutic implications of cardiorenal syndrome. Clin Exp Nephrol.
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Aggarwal A, Kabbani SS, Rimmer JM, Gennari FJ, Taatjes DJ, Sobel BE, Schneider DJ. Biphasic
effects of hemodialysis on platelet reactivity in patients with end-stage renal disease: a
potential contributor to cardiovascular risk. Am J Kidney Dis. 2002;40:315-22.
Cockcroft DW, Gault MH. Prediction of creatinine clearance from serum creatinine. Nephron.
1976;16:31-41.
van der Putten K, Jie KE, Emans ME, Verhaar MC, Joles JA, Cramer MJ, Velthuis BK, Meiss L,
Kraaijenhagen RJ, Doevendans PA, Braam B, Gaillard CA. Erythropoietin treatment in patients
with combined heart and renal failure: objectives and design of the EPOCARES study. J Nephrol.
2010; 23(4):363-8.
Dickstein K, Cohen-Solal A, Filippatos G, McMurray JJ, Ponikowski P, Poole-Wilson PA,
Strömberg A, van Veldhuisen DJ, Atar D, Hoes AW, Keren A, Mebazaa A, Nieminen M, Priori
SG, Swedberg K; ESC Committee for Practice Guidelines (CPG). ESC guidelines for the diagnosis
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by the European Society of Intensive Care Medicine (ESICM). Eur J Heart Fail. 2008;10:933-89.
van Bladel ER, Roest M, de Groot PG, Schutgens RE. Up-regulation of platelet activation in
hemophilia A. Haematologica. 2011;96:888-95.
Kyrle PA, Stockenhuber F, Brenner B, Gössinger H, Korninger C, Pabinger I, SunderPlassmann G, Balcke P, Lechner K. Evidence for an increased generation of prostacyclin in the
microvasculature and an impairment of the platelet alpha-granule release in chronic renal
failure. Thromb Haemost. 1988;60:205-8.
Schoorl M, Bartels PCM, Gritters M, Fluitsma D, Musters R, Nubé MJ. Electron microscopic
observation in case of platelet activation in a chronic haemodialysis subject. Hematol Rep.
2011;3:e15.
Noris M, Remuzzi G. Uremic bleeding: closing the circle after 30 years of controversies? Blood.
1999;94:2569-2574.
Zwaginga JJ, IJsseldijk MJ, Beeser-Visser N, De Groot PG, Vos J, Sixma JJ. High von Willebrand
factor concentration compensates a relative adhesion defect in uremic blood. Blood.
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Casserly LF, Dember LM. Thrombosis in end-stage renal disease. Semin Dial. 2003;16:245-56.
Palmer SC, Di Micco L, Razavian M, Craig JC, Perkovic V, Pellegrini F, Copetti M, Graziano
G, Tognoni G, Jardine M, Webster A, Nicolucci A, Zoungas S, Strippoli GF. Effects of antiplatelet
therapy on mortality and cardiovascular and bleeding outcomes in persons with chronic
kidney disease. A systematic review and meta-analysis. Ann Intern Med. 2012;156:445-459.
Zwaginga JJ. Hemodialysis, erythropoietin and megakaryocytopoiesis: factors in uremic
thrombocytopathy and thrombophilia. J Thromb Haemost. 2004;2(8):1272-4.
Stumpf C, Lehner C, Eskafi S, Raaz D, Yilmaz A, Ropers S, Schmeisser A, Ludwig J, Daniel WG,
Garlichs CD. Enhanced levels of CD154 (CD40 ligand) on platelets in patients with chronic heart
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Chapter 7
Functional platelet defects in children with severe chronic ITP:
as tested with two novel assays applicable for low platelet counts
Esther R. van Bladel,1* Annemieke G. Laarhoven,2* Laila van der Heijden,1,3 Katja M. Heitink-Pollé,3
Leendert Porcelijn,2 C. Ellen van der Schoot,2 Masja de Haas,2 Mark Roest,1 Gestur Vidarsson,2#
Philip G. de Groot1# and Marrie C. Bruin3#
1
Department of Clinical Chemistry and Hematology, University Medical Center Utrecht,
Utrecht, the Netherlands; 2Department of Experimental Immunohaematology, Sanquin
Research, and Landsteiner Laboratory, Academic Medical Center, University of Amsterdam,
Amsterdam, the Netherlands; 3Department of Pediatric Hematology, Wilhelmina
Children’s Hospital, University Medical Center Utrecht, Utrecht, the Netherlands.
* and # equally contributed
In revision
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Abstract
Immune thrombocytopenia (ITP) is an autoimmune disease with a complex heterogeneous
pathogenesis and a bleeding phenotype that is not necessarily correlated to platelet count.
In this study, the platelet function was assessed in a well-defined cohort of 33 pediatric
chronic ITP patients. Since regular platelet function test cannot be performed in patients
with low platelet counts, two new assays were developed to determine platelet function.
First, the micro aggregation test measuring in platelets isolated from 10 ml whole blood,
the platelet potential to form micro-aggregates in response to an agonist. Second, the
platelet reactivity assay, measuring platelet reactivity to ADP, convulxin (CVX) and thrombin
receptor activator peptide (TRAP) in only 150 μL unprocessed whole blood. Patients with
a severe bleeding phenotype, demonstrated a decreased aggregation potential upon
phorbol myristate acetate (PMA) stimulation, decreased platelet degranulation following
ADP stimulation and a higher concentration of ADP and convulxin needed to activate the
glycoprotein IIbIIIa complex compared to patients with a mild bleeding phenotype. In
conclusion, here we have established two functional tests that allows for evaluation of
platelet function in patients with extremely low platelet counts (10*109). These tests show
that platelet function is related to bleeding phenotype in chronic ITP.
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Introduction
Platelets play a critical role in hemostasis. When the vascular endothelium is disrupted,
platelets adhere to subendothelium and initiate primary hemostasis. Excessive bleeding
can occur if primary hemostasis is abnormal, either because of deficient platelet number
or function. In vivo, primary hemostasis can be tested via bleeding time. However, this test
does not distinguish between the varieties of causes of disturbed primary hemostasis.1-3 This
can be tested more specifically in vitro, but current methods require relative high numbers
of platelets and are consequently unsuitable for patients with low platelet counts.1;2;4
Immune thrombocytopenia (ITP) is the most common cause of primary
thrombocytopenia in children, with an incidence of about 1 in 20,000 children.5;6 Although
the pathophysiology of ITP is not fully understood, two major forms are recognized: acute
ITP and chronic ITP. Acute ITP is characterized by a sudden onset of bruising and bleeding
in an otherwise healthy child. Often there is a history of viral illness in the weeks preceding
the onset of bruising.5 Full blood counts show low platelet numbers (frequently < 20*109/L)
as the only abnormality. In acute ITP, auto antibodies, recognizing glycoproteins on the
surface of platelets and megakaryocytes, are considered the underlying cause.7;8 These
antibodies are thought to result in accelerated clearance of platelets and megakaryocytes,
and thereby, may also lead to decreased production of platelets.7;9 In chronic ITP, the
attribution of auto antibodies to the pathogenesis of thrombocytopenia is less clear.
In the majority of pediatric ITP patients, thrombocytopenia resolves spontaneously
within weeks or months. In about 25% of the patients, thrombocytopenia persists and
becomes chronic.5;10;11 During chronic ITP platelet counts can vary in time from very low
(<10*109/L) to almost normal. However, the observed bleeding tendency does not correlate
strictly with platelet count. Cases with either low platelet counts without bleeding, or
relatively high platelet counts with severe bleeding do occur. Causes for this variation in
bleeding tendency are unknown. We hypothesize that variation in platelet function can
account for the differences observed in bleeding phenotypes. Until now it has not been
possible to predict if an individual child with chronic ITP is at risk for severe bleeding due to
platelet malfunction, because of the lack of reliable tests for platelet function in patients
with low platelet counts.2-4
We here describe two functional platelet tests that can be used on patient material
with very low (≥10*109/L) platelet numbers. The first test, the platelet micro aggregation
test, based on a recently developed test12, was adjusted to test platelet function directly
by determining the potential of patients’ platelets to form micro-aggregates together with
platelets from a healthy control in response to an agonist. Ristocetin and PMA were used to
7
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activate platelets through the von Willebrand Factor (vWF) and the fibrinogen binding site
on glycoproteinIbIX (GPIbIX) and GPIIbIIIa respectively.12-14 In the second assay, the platelet
reactivity assay, the reactivity in three major physiological platelet activation pathways
was determined in unprocessed whole blood by flow cytometry on the level of individual
platelets.15 ADP, convulxin (CVX) and thrombin receptor activator peptide (TRAP) were
used to activate platelets via P2Y receptors (P2Y1 and P2Y12), glycoprotein VI receptor and
Proteinase Activated Receptor-1 (PAR-1), respectively. Furthermore the platelet activation
test quantifies both degranulation (P-selectin expression on the platelet surface) and
activation of glycoprotein IIbIIIa (binding of fibrinogen to platelets). Both assays require a
minimum of blood compared to classical functional platelet assyas, i.e. 10 ml of whole blood
for the platelet micro aggregation test and 150 μl whole blood for the platelet reactivity
assay. After validating both tests for low platelet numbers with platelets from healthy
controls, the tests were used within a well-defined cohort of children with chronic ITP
and results were correlated with reported bleeding scores. Our results show that patients
suffering from serious bleedings have impaired functional platelet capacities compared to
patients with no or mild bleeding and healthy controls. Both tests yield valuable information
that may be used to predict future bleeding tendencies in chronic ITP patients.
Methods
Patients
Children aged 6 to 13 years with chronic ITP were included in this multicenter observational
study. Parents and patients aged 12 years and older gave written informed consent. The
study was approved by the Institutional Review Board of the University Medical Center
Utrecht and performed in accordance with the Declaration of Helsinki.
Chronic ITP was defined as isolated thrombocytopenia with a platelet count of less
than 100*109/L for more than 12 months. Patients were classified as having either a mild
grade 0-3, or a severe grade 4-5 bleeding phenotype, according to the Buchanan bleeding
score.16 For scoring, all bleeding problems during the course of ITP were taken into account,
irrespective of platelet count.
Data on duration of the ITP, bleeding tendency and medication were collected by
questionnaire and from the patient’s medical files. All patients were tested for the presence
of auto antibodies with the indirect platelet immunofluoresence test (PIFT) and the indirect
monoclonal antibody immobilization of platelet antigens (MAIPA).
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Blood obtained from adult healthy individuals served as positive controls, and two
Glanzmann Thrombasthenia patients were included as negative controls for the micro
platelet aggregation tests. Acquired Glanzmann patients have α-GPIIbIIIa auto antibodies
which inhibit platelet aggregation via the GPIIbIIIa route, whereas other pathways are
normal. Primary Glanzmann patients lack GPIIbIIIa almost completely, but other pathways
may also be hampered.12;17
Micro platelet aggregation test
For the micro platelet aggregation assays peripheral whole blood was collected into a 10 mL
BD Vacutainer with 17 IU sodium heparin (Becton Dickinson and Compagny, Plymouth,
United Kingdom), and centrifuged for 15 minutes at 218 g to obtain platelet rich plasma
(PRP). PRP was washed 1:1 in sequestrin buffer (17.5 mM Na2HPO4, 8.9 mM Na2EDTA, 154
mM NaCl, pH 6.9, 0.1% (w/v) BSA) and centrifuged for 6 minutes at 2374 G. Platelets were
washed again by adding 10 ml sequestrin buffer and finally dissolved in 20mM HEPES
medium enriched with 132 mM NaCl, 6 mM KCl, 1 mM MgSO4, 1.2 mM KH2PO4 and 5 mM
glucose. To measure aggregation function, patients’ platelets were suspended in HEPES
medium to a final concentration of 10*106/ml, a minimum of 400 μl was required. Control
platelets, from a healthy donor, were suspended in HEPES medium to a final concentration
of 90*106/ml, a minimum of 800 μl was required. The platelets from the patients were
stained with 0.8 μM PKH-26 (PKH26GL, Sigma-Aldrich), whereas the control platelets
were stained with 0.2 μM carboxyfluorescein diacetate succinimidyl ester (CFSE) (C1157,
Invitrogen). Both were incubated in the dark for 15 minutes at room temperature. Staining
was stopped by adding citrate-phosphate-dextrose (CPD) plasma from pooled blood
group AB+ healthy donors, to a final concentration of 20%. Subsequently, 500 μl 10*106/ml
patient platelets were mixed with 500 μl 90*106/ml control platelets and incubated with
20 μM PPACK (Mercks Biochemicals, Cat.520222) for 5 minutes at 37°C while shaking at
300 rpm. Platelet mixes were incubated for 5 minutes with 3 mM CaCl2 in order to enable
platelet activation. A healthy control was always tested next to a patient as a positive
control. Platelets were activated with the agonist 100 ng/ml PMA (SC-3576, Santa Cruz),
or with 1.5 mg/ml ristocetin (Biopool, Trinity Biotech, Cat.50705), to activate GPIIbIIIa and
GPIbIX respectively. Samples without an agonist served as a negative control. Samples
were taken and fixed in 0.5% formaldehyde (Buffered Formaldehyde 4%, Klinipath, Duiven,
The Netherlands) in a V-bottom 96 wells plate (Nunc, Thermo Fisher Scientific), at 0, 5 and
10 minutes. Fixed samples were measured on a FACS CantoII+HTS (BD Biosciences, San
Jose, CA) and analyzed with BD Facs DIVA 6.1 software (BD Biosciences, San Jose, CA), and
7
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aggregation calculated as: Aggregation (%) = #aggregates (CFSE+PKH+ events) / #patients’
platelets (PKH+ events) x 100%.
Platelet reactivity assay
Blood was collected into a 4.5 mL BD Vacutainer with 0.5 mL sodium citrate 3.2% (Becton,
Dickinson and Company, Plymouth, United Kingdom). Serial dilutions of adenosine
diphosphate (ADP; Roche, Almere, The Netherlands) to stimulate the P2Y receptors (starting
from 125 μM), Convulxin (CVX; Pentapharm, Basel, Switzerland) to stimulate the GPVI
receptor (starting from 39 ng/mL), and thrombin receptor activator peptide (TRAP; Bachem
AG, Bubendorf, Switzerland) to stimulate the PAR-1 receptor (starting from 625 μM), all in
eight 4-fold dilutions, were prepared in a mixture of 47.5 μL HEPES buffered saline (HBS;
consisting of 10 mM HEPES, 150 mM NaCL, 1 mM MgSO4, and 5 mM KCl, pH 7.4, filtered
through a 0.22 μm filter), 2 μL R-phycoerythrin (RPE) labeled mouse anti-human P-selectin
antibodies (#555524; BD biosciences, Franklin Lakes, NJ), and 0.5 μL Alexa Fluor 488-labeled
fibrinogen (Invitrogen, Eugene, OR).
A control sample, only containing 47.5 μL HBS, 2 μL RPE-labeled mouse anti-human
P-selectin antibodies, and 0.5 μL Alexa Fluor 488-labeled fibrinogen, was prepared to
determine the basal level of platelet activation.
To measure patient and healthy control platelet reactivity, 5 μL fresh, citrate anticoagulated, whole blood was added to all samples. After 20 minutes of incubation, 50 μL
Optilyse B (Beckman Coulter Inc., Fullerton, CA) was added to fix the samples. After
10 minutes of platelet fixation, 395 μL distilled water was added to lyse the erythrocytes,
thereby limiting cells present in sample for measurement, and in doing so reducing the
time needed to measure sufficient amount of platelets. After half an hour of incubation
at room temperature the samples were kept at 4˚C until analysis on the FACS Canto II
flow cytometer from BD Biosciences, which was at all times performed within 24 hours.
Single platelets were gated based on forward and side scatter properties, 10000 single
platelets were measured in each sample. The median fluorescence intensity (MFI) of RPElabeled mouse anti-human P-selectin antibodies and Alexa Fluor 488-labeled fibrinogen
on platelets was measured with FACS analysis, representing the quantity of P-selectin and
open GPIIbIIIa receptor per platelet.
The obtained FACS data were quantified using BD FACSDiva software 6.1.2. The platelet
responsiveness to agonists was qualified by calculation of the EC50 and the maximal
response using GraphPad Prism 5.03 (GraphPad Software, San Diego, CA, U.S.A.). The EC50
represents the concentration agonist generating a response halfway between baseline and
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maximum response. The response of the platelets to the highest agonist concentration in
the dilution series represents the maximal effect of stimulation.
To validate the assay for use at low platelet count, platelet rich plasma (PRP) form
healthy donors was isolated by centrifugation for 15 minutes at 160 g, and platelet poor
plasma (PPP) was obtained by centrifugation twice for 10 minutes at 2000 g. Platelet
number was set at 250, 50, 25 and 10*109/L by diluting PRP by addition of PPP. 5 μL of
these mixtures was added to ADP, CVX and TRAP samples to determine platelet reactivity
at different platelet numbers.
Statistical analysis
Statistical analyses were performed using IBM SPSS Statistics 20.0.0 for Windows
(International Business Machines Corporation, New York, U.S.A.). Data are shown as median
with interquartile range (IQR) unless otherwise indicated. Wilcoxon signed ranks tests were
performed to analyze data from related samples used in the validation experiments. For
analysis of patient data, comparison between two groups was performed by Mann-Whitney
U test for numerical data and by Fisher’s exact test for categorical data. Correlation of two
numerical variables was performed by Spearman’s rank correlation. P-values lower than
0.05 were considered to be statistically significant.
Results
Baseline characteristics
A total of 33 patients were included in this study, of which 10 patients were classified as
having a severe bleeding phenotype (30.3%) and 23 patients as having a mild bleeding
phenotype (69.7%) (Table 1). Platelet reactive auto antibodies were found in a minority of
patients: 3 were found positive by indirect PIFT, and 6 by indirect MAIPA. These patients
were equally distributed between groups.
7
Validation of two new functional platelet assays at low platelet count
For the micro aggregation test, platelets were gated based on their forward / sideward
scatter characteristics (Figure 1A). After mixing PKH26 labeled patients samples with CFSElabeled control platelets, a time dependent increase in aggregation was observed, with a
maximal aggregation after 10 minutes. Aggregation was defined as events double positive
in PKH and CFSE, and quantified as shown in Figure 1B. We first determined the optimal and
the lowest platelet number required for reliable results, by testing a range of proportions
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Table 1. Baseline characteristics
Age at inclusion, years‡
Age at diagnosis, years‡
Sex, male†
Platelet count at day of inclusion, *109/L‡
MPV at day of inclusion, fL‡
Positive indirect PIFT†
Positive indirect MAIPA†
‡
†
Mild bleeding
phenotype
(n=23)
Severe bleeding
phenotype
(n=10)
P
10 (6-13)
3 (2-9)
6 (26)
74 (53-117)
9.4 (8.6-11.6)
2 (9)
4 (17)
12 (8-15)
4(2-12)
3 (30)
58 (14-156)
10.8 (8.4-14.8)
1 (10)
2 (20)
0.576
0.773
1.000
0.805
0.216
1.000
1.000
Data represent median (interquartile range), P was calculated using Mann Whitney U test.
Data represent number (percentage), P was calculated using Fisher’s exact test.
in healthy controls (Supplementary Figure 1). This was accomplished by changing the
concentration of PKH-labeled platelets, whereas CFSE-labeled platelets were kept constant
at 90*106/ml using healthy control platelets. A ratio of approximately 1 PKH26 (test platelet)
to 10 CFSE-labeled control platelets was considered reliable and feasible, indicating that
10 ml whole blood with only 10*109 platelet/L was sufficient. We also crossed platelets
from different healthy controls, to assess whether well functioning platelets from one
donor will reach a similar aggregation level when aggregated with platelets from different
healthy controls. Similar aggregation levels were reached (Supplementary Figure 2).
For the platelet reactivity assay, single platelets were gated (Figure 1A), and both
P-selectin expression and open GPIIbIIIa were determined based on MFI levels (Figure 1C).
To validate the use of the platelet reactivity assay for the determination of platelet
function in samples with low platelet numbers, platelet reactivity was measured in platelet
rich plasma samples of healthy controls diluted to a platelet number of 250, 50, 25 and
10*109/L. Dilution was accomplished by addition of plasma, so that only platelet count was
diluted without dilution of plasma proteins. Platelet reactivity within donors, quantified by
agonist concentration needed to obtain half maximal activation (EC50) and by response
to maximal agonist concentration, did not change when platelet number was reduced to
10*109/L (Supplementary Figure 3). Although validation experiments already showed that
parameters of platelet reactivity were not dependent on platelet number, we investigated
if platelet count possible affected platelet reactivity in our study population. In the
33 patients, no correlation was observed between platelet number at day of inclusion and
platelet reactivity (rho=0.165; p= 0.367; Supplementary Figure 4).
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C
A
SSC
P-selectin
Platelets
FSC
GPIIbIIIa(open)
B
85.1%
14.9%
10 min
85.4%
85.6%
14.4%
10 min
9.3%
90.7%
PKH
0 min
CFSE
Figure 1. Methods platelet micro aggregation test and platelet reactivity assay.
(A) Platelets were gated based on forward and sideward scatter properties (FSC/SSC). (B) Micro
platelet aggregation test. Test platelets, derived either from a patient or healthy control were
stained with PKH and mixed in 1:9 ratio with healthy control platelets, stained with CFSE. Platelets
were analyzed in the PE (PKH) and FITC (CFSE) channel, platelets positive in both channels were
identified as aggregates. The left panel shows unstimulated platelets after 0 minutes, the middle
panel unstimulated platelets after 10 minutes and the right panel PMA stimulated platelets after 10
minutes. (C) Platelet reactivity assay. The median fluorescence intensity (MFI) of RPE-labeled mouse
anti-human P-selectin antibodies and Alexa Fluor 488-labeled fibrinogen on platelets was measured
by FACS analysis, representing the quantity of P-selectin and open GPIIbIIIa receptor per platelet.
The left panel shows unstimulated platelets, the right panel shows platelets stimulated with 625 μM
TRAP.
7
Micro aggregation of platelets
To assess whether the test correctly identified patients with known functional platelet
defects, an acquired and a primary Glanzmann patient were tested. Both patients showed
almost complete inhibition of platelet aggregation when stimulated with PMA via GPIIbIIIa
(Figure 2A-B). On the contrary, baseline level and aggregation upon stimulation with
Ristocetin via GPIbIX was completely normal in the acquired Glanzmann patient, whereas
the primary Glanzmann patient reached approximately half that of the healthy control
(Figure 2C).
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Figure 2. Severe chronic ITP patients show functional defects in platelet micro aggregation upon
PMA stimulation.
An acquired Glanzmann patient, with functional GPIIbIIIa and a anti-GPIIbIIa antibody, and a primary
Glanzmann patient, without functional GPIIbIIIa, served as negative controls for the assay (A, B, C).
Seven severe chronic ITP patients (D, E, F) and 11 mild chronic ITP patients (G, H, I) were tested in the
micro platelet aggregation test. Results are given as percentage of aggregation reached after 0, 5 and
10 minutes without (A, D, G), after PMA (B, E, H) or after Ristocetin stimulus (C, F, I). Outcomes are
depicted pair-wise, the patient compared to a healthy control tested in parallel. Statistical analysis
was performed by Wilcoxon signed ranks testing.
We subsequently performed the micro platelet aggregation test on 18 chronic ITP
patients, 7 with severe, and 11 with mild bleeding phenotype. Unstimulated platelets did
not aggregate over time (Figure 2D, G). After ten minutes, severe phenotype patients had
significantly lower aggregation levels after five (30.9% [16.0-42.8]) and ten minutes (52.4%
[28.7-66.45]) of PMA stimulation compared to healthy control platelets (64.2% [51.4-65.2];
p=0.021, 80.8% [67.3-80.9]; p=0.017, after 5 and 10 minutes, respectively) (Figure 2E). In
contrast, mild phenotype patients equaled their healthy control after five (45.2% [34.770.0], 63.0% [47.3-70.9]; p=0.481) and ten minutes (71.9% [50.4-84.9], 80.6% [62.2-85.83];
p=0.631). (Figure 2H). After ten minutes of ristocetin stimulation severe phenotype patients
tended to have lower aggregation levels after five (27.1% [24.0-41.7]) and ten minutes
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(35.3% [29.2-52.7]) (Figure 2F) compared to healthy controls (49.2% [31.5-55.7]; p=0.073,
55.8% [40.9-66.2]; p=0.125), but the differences were not significant. Mild phenotype
patients equaled their healthy control after both five (47.6% [25.2-58.1], 49.9% [33.0-56.9];
p=0.971) and ten minutes (53.8% [37.9-65.7], 54.3% [41.4-71.2]; p=0.684) (Figure 2I).
Platelet reactivity assay
Basal level of platelet activation and platelet reactivity to the agonists ADP, CVX and TRAP
was determined for all included patients by measuring platelet P-selectin expression and
opening of the GPIIbIIIa receptor in whole blood (Figure 3). There were no significant
differences when comparing patients with chronic ITP with a mild phenotype to healthy
controls.
Basal levels of platelet activation did not differ between chronic ITP patients with a
mild (n=23) and with a severe bleeding phenotype (n=10), nor when compared to healthy
controls (n=8) (Figure 3A-B). Platelet degranulation, determined via measurement of
P-selectin expression, in response to maximal ADP stimulation was less in patients with
a severe bleeding phenotype (9.6*102 AU [3.7-11.8]), compared to both patients with a
mild phenotype (13.0*102 AU [11.2-15.3]; p=0.020) and to controls (22.4*102AU [10.7-25.5];
p=0.016) (Figure 3C). When stimulated with CVX or TRAP, no differences in P-selectin
expression were observed between mild and severe bleeders (Figure 3G, K-L). Patients
with a severe phenotype did need more CVX (11.0 ng/ml [5.6-393.8]) to stimulate for half
maximal P-selectin expression compared to healthy controls (4.7 ng/ml [2.6-6.4]; p=0.021)
(Figure 3H).
The concentration of ADP and CVX needed to obtain opening of the half maximal amount
of GPIIbIIIa receptors, was higher in severe bleeders (1.1 μM ADP [0.9-1.5] and 9.0 ng/ml
CVX [5.2-12.9]), compared to both mild bleeders (0.7 μM ADP [0.4-1.0] and 5.8 ng/ml CVX
[2.8-9.0]; p=0.031 and 0.031) and healthy controls (0.5 μM ADP [0.4-0.9] and 3.0 ng/ml CVX
[1.4-4.0]; p=0.027 and 0.006) (Figure 3F, J). Reactivity to TRAP was equal between groups
(Figure 3M-N). Maximal GPIIbIIIa response to all agonists did not differ between groups
(Figure 3E, I, M).
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P-selectin
A. Baseline P-selectin MFI
120
p = 0.384
p = 0.274
MdFI - AU
100
80
60
40
20
0
Mild
Severe
Control
C. ADP - maximal MFI
4000
p = 0.020
D. ADP - EC50 MFI
p = 0.016
3
ADP-∝M
MdFI - AU
3000
2000
1000
0
Mild
Severe
1
p = 0.965
MdFI - AU
10000
5000
0
2000
CVX-ng/ml
p = 0.343
Mild
Severe
p = 0.384
Severe
Control
p = 0.207
p = 0.021
200
20
10
0
Control
K. TRAP - maximal MFI
15000
Mild
H. CVX - EC50 MFI
G. CVX - maximal MFI
15000
p = 0.762
2
0
Control
p = 0.384
Mild
Severe
Control
L. TRAP- EC50 MFI
p = 0.515
25
p = 0.324
p = 0.274
MdFI - AU
20
TRAP-∝M
10000
5000
15
10
5
0
Mild
Severe
Control
0
Mild
Severe
Control
Figure 3. Severe chronic ITP patients show functional defects platelet reactivity upon ADP and
convulxin stimulation.
To determine baseline platelet reactivity, mean fluorescence intensity (MFI) in arbitrary units (AU) of
fluorescent labeled anti-P-selectin antibody (A) and of fluorescent labeled fibrinogen (B) bound to
platelets was determined in the absence of agonists. Maximal MFI in AU in response to stimulation
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Open GPIIbIIIa
B. Baseline open GPIIbIIIa MFI
MdFI - AU
100
p = 0.923
p = 0.696
50
0
-50
-100
Mild
Severe
Control
E. ADP - maximal MFI
8000
p = 0.954
F. ADP - EC50 MFI
p = 0.408
4
ADP-∝M
MdFI - AU
4000
2
1
2000
Mild
Severe
0
Control
I. CVX - maximal MFI
8000
p = 0.576
5000
CVX-ng/ml
MdFI - AU
4000
2000
Control
p = 0.031
p = 0.006
20
15
10
7
5
Mild
Severe
0
Control
p = 0.475
Mild
Severe
Control
N. TRAP - EC50 MFI
p = 0.146
12
p = 0.923
p = 0.360
10
TRAP-∝M
MdFI - AU
4000
3000
2000
1000
0
Severe
300
25
M. TRAP - maximal MFI
5000
Mild
J. CVX - EC50 MFI
p = 0.460
6000
0
p = 0.027
3
6000
0
p = 0.031
8
6
4
2
Mild
Severe
Control
0
Mild
Severe
Control
with ADP, convulxin and TRAP, and concentration of these agonists needed to obtain half maximal
MFI were determined for both platelet P-selectin expression (C,D,G,H,K,L) and opening of GPIIbIIIa
receptor (E,F,I,J,M,N). The dots represent individual results for the 10 severe patients, 23 mild
patients and 8 healthy controls, with a line at the median. Statistical analysis was performed by
Mann-Whitney U testing.
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Platelet micro aggregation versus platelet reactivity
micro aggregation index
In both the micro aggregation and platelet reactivity assay, the platelets from severe ITP
patients showed diminished activity compared to healthy control. Indeed, PMA-induced
aggregation and maximal platelet P-selectin response to ADP, the parameters giving the
most discrimination between healthy controls and patients within the assays, correlated
significantly for the 18 patients in which both assays were performed (rho=0.519; p=0.027)
(Figure 4). In Table 2, the characteristics of both assays are listed.
2.0
Mild
Severe
1.5
1.0
0.5
0.0
0
500
1000
1500
2000
max P-selectin to ADP - AU
Figure 4. Correlation of micro platelet
aggregation and platelet reactivity.
Response to PMA induced micro platelet
aggregation and ADP induced maximal platelet
reactivity correlated with a Spearmans
correlation coefficient of 0.519 (p=0.027)
for all 18 patients in which both assays were
performed.
2500
Table 2. Assay characteristics
Micro platelets aggregation test
Platelet reactivity assay
Patient material
Pathways tested
-10 ml whole blood
-GPIIbIIIa (PMA)
-GPIbIX (ristocetin)
Material
processing
-Platelet isolation (1 hour)
-Staining, incubation and fixation
(1.5 hours)
Analysis
Read-out
-FACS analysis in a 96 well plate
-Micro aggregation with control
platelets
-150 μl whole blood
-P2Y1 and P2Y12 (ADP)
-GPVI (convulxin)
-PAR1 (TRAP)
-Incubation of whole blood
(20 minutes)
-Fixation (10 minutes)
-Lysis of erythrocytes (30 minutes)
-FACS analysis in a 96 well plate
-Degranulation (P-selectin)
-Fibrinogen binding (GPIIbIIIa
opening)
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Discussion
A variation in bleeding severity exists between patients with chronic ITP that cannot be
explained by platelet counts alone. We therefore hypothesized that platelet function might
be affected diversely between patients with chronic ITP. In the current study we observed
a functional platelet defect in chronic ITP patients with a severe bleeding phenotype.
These platelets displayed a decreased potential to form micro-aggregates following PMA
stimulation, decreased platelet degranulation following ADP stimulation and higher ADP
and convulxin concentrations needed for half maximal activation of the GPIIbIIIa complex.
To our knowledge, this is the first study to establish platelet function in individuals
with low platelet counts. Classical platelet function tests are not reliable with platelet
counts below 50*109/L.2;4;18;19 In this study, we present the micro aggregation test and the
platelet reactivity assay for their use in samples containing low platelet numbers down to
10*109/L. We show that the micro aggregation test can be performed in 10 ml blood with
this minimal platelet number to assess platelet function directly as well as pathway specific.
In the platelet reactivity assay, only 150 μL of whole blood was needed for all performed
platelet reactivity measurements. The assay determines the reactivity of single platelets
independently of platelet number. Only 3 quick handling steps, incubation, fixation and
red cell lysis, are needed to perform the assay, since platelet isolation is not required. The
serial dilutions used for platelet activation can be stored at -20 degrees celcius for several
weeks, and can be used instantly when needed. The assay has a broad and quantitive
detecetion range, measuring both platelet degranulation and GPIIbIIIa opening, and it
distinguishes between multiple specific activation pathways. In the scope of this research
we have decided to test pathways of specific interest in ITP, but naturally, each agonist of
choice can be used in both assays. Both tests might be of great diagnostic value in a broad
range of patients suffering from thrombocytopenia. In this study in children with chronic
ITP we proved that we can determine platelet function and correlate this with bleeding
phenotype.
We classified patients as having a mild or a severe bleeding phenotype using the overall
Buchanan bleeding score.20 We compared patients with a mild bleeding phenotype scoring
a grade 0-3, to patients with a severe phenotype scoring a grade 4-5 in our search for a
relation between platelet function and bleeding phenotype. This established score for ITP
patients, which grades bleeding severity based on skin and mucosal bleeds showed to be
associated with platelet function.
With the micro aggregation assay, we found patients classified as severe bleeders
to have significantly lower aggregation levels compared to healthy controls upon 5 and
7
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10 minutes of PMA stimulation. However, when stimulating with Ristocetin no differences
between patients and healthy controls were observed. A possible explanation for the
decreased PMA response might be an interference of auto antibodies present on patient
platelets not detectable by indirect MAIPA or indirect PIFT as both assays have very low
sensitivities (25-39% and 30%, respectively).21-23 The presence of anti-GPIIbIIIa antibodies in
these patients might explain why we predominantly see an effect with PMA stimulation and
not with Ristocetin, as these auto antibodies are most frequently found in ITP patients.7;24
With the platelet reactivity assay, we found the severe patients to have significant lower
P-selectin expression after activation with ADP, but not after activation with CVX or TRAP.
Next to platelet P-selectin expression, opening of the platelet glycoprotein IIbIIIa receptor
was measured. We observed that for both stimulation with ADP and CVX, higher agonist
concentrations were needed in patients with a severe phenotype to obtain opening of
the half maximal amount of GPIIbIIIa receptors, when compared both with mild patients
and controls. Interference of auto antibodies in binding of FITC labeled fibrinogen to
open GPIIbIIIa in the platelet reactivity assay seems unlikely, since differences in GPIIbIIIa
opening are seen only for ADP and CVX stimulation, and not following TRAP stimulation.
Theoretically, the decrease in platelet reactivity observed in severe patients might be caused
by an interaction of auto antibodies with platelets or megakaryocytes, leading to outsidein signalling influencing platelet reactivity to natural stimuli or influencing megakaryocyte
maturation and differentiation.
A limitation to our study is that we cannot confirm if the decreased platelet activity, we
observed in the more severe ITP patients, was due to anti-platelet antibodies. Although
direct variants of PIFT and MAIPA are more sensitive, these still require high number of
platelets and are therefore unsuitable for severe cases with low number of platelets21;23.
Therefore, direct confirmation whether or not auto-antibodies are to blame, will have
to wait until more reliable methods are developed to detect low levels of anti-platelet
antibodies using low number of platelets. Also, a larger prospective study will be required
to determine if these assays reported here can predict severity.
Within healthy adult individuals, platelet parameters are known to be stable when
measured at several time points.25 A single measurement represents a person’s innate
platelet reactivity, making repeated measurements unnecessary. However, we cannot
exclude antibodies responsible for ITP to influence this innate platelet reactivity via
outside in signalling. Platelet reactivity in chronic ITP patients could therefore vary along
with disease activity. Future longitudinal studies will be necessary to determine whether
platelet reactivity is stable in chronic ITP.
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In this study we have focused on in vitro platelet function in the context of bleeding
severity in chronic ITP. Nevertheless in vivo other factors could influence bleeding severity
as well, such as endothelial function, and plasma factors. These factors might, together
with platelet function, influence bleeding phenotype.
In summary, here we have established two functional tests that allows for evaluation of
platelet function in children with chronic ITP, and to associate the results with the bleeding
phenotype. Patients with a severe bleeding phenotype were found to have a decreased
platelet function, shown by decreased platelet aggregation following PMA stimulation
in the micro platelet aggregation test, decreased platelet degranulation following ADP
stimulation in the platelet reactivity assay and higher ADP and convulxin concentrations
needed for half maximal activation in the platelet reactivity assay. Longitudinal studies will
have to confirm if the micro aggregation test and the platelet reactivity assay can be used
to predict bleeding phenotype in chronic ITP.
Acknowledgments
The authors would like to thank M. Peters, J. Verhage, W.A. Kors, A.M.J. van Meurs,
N. Dors, F.J. Smiers, R.Y.J. Tamminga and A. Beishuizen for enrolling patients in this study.
This work was partly funded by the Landsteiner Foundation for Bloodtransfusion Research
(Landsteiner Stichting voor Bloedtransfusie Research, LSBR 0842) to AL.
7
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McMillan R. Autoantibodies and autoantigens in chronic immune thrombocytopenic purpura.
Semin Hematol. 2000;37:239-48.
Newland AC, Macey MG. Immune thrombocytopenia and Fc receptor-mediated phagocyte
function. Ann Hematol. 1994;69:61-7.
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9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. Perdomo J, Yan F, Chong BH. A megakaryocyte with no platelets: anti-platelet antibodies,
apoptosis, and platelet production. Platelets. 2013;24:98-106.
Grace RF, Long M, Kalish LA, Neufeld EJ. Applicability of 2009 international consensus
terminology and criteria for immune thrombocytopenia to a clinical pediatric population.
Pediatr Blood Cancer. 2012;58:216-20.
Neunert CE, Buchanan GR, Imbach P et al. Bleeding manifestations and management of children
with persistent and chronic immune thrombocytopenia: data from the Intercontinental
Cooperative ITP Study Group (ICIS). Blood. 2013;121:4457-62.
De Cuyper IM, Meinders M, van d, V et al. A novel flow cytometry-based platelet aggregation
assay. Blood. 2013;121:e70-e80.
Bonnefoy A, Yamamoto H, Thys C et al. Shielding the front-strand beta 3 of the von Willebrand
factor A1 domain inhibits its binding to platelet glycoprotein Ibalpha. Blood. 2003;101:137583.
Gadisseur A, Hermans C, Berneman Z et al. Laboratory diagnosis and molecular classification
of von Willebrand disease. Acta Haematol. 2009;121:71-84.
van Bladel ER, Roest M, de Groot PG, Schutgens RE. Up-regulation of platelet activation in
hemophilia A. Haematologica. 2011;96:888-95.
Buchanan GR, Adix L. Grading of hemorrhage in children with idiopathic thrombocytopenic
purpura. J Pediatr. 2002;141:683-88.
Porcelijn L, Huiskes E, Maatman R, de KA, de HM. Acquired Glanzmann’s thrombasthenia
caused by glycoprotein IIb/IIIa autoantibodies of the immunoglobulin G1 (IgG1), IgG2 or IgG4
subclass: a study in six cases. Vox Sang. 2008;95:324-30.
Picker SM. In vitro assessment of platelet function. Transfusion and Apheresis Science.
2011;44:305-19.
Rand ML, Leung R, Packham MA. Platelet function assays. Transfusion and Apheresis Science.
2005;28:307-17.
Buchanan GR, Adix L. Grading of hemorrhage in children with idiopathic thrombocytopenic
purpura. J Pediatr. 2002;141:683-8.
Brighton TA, Evans S, Castaldi PA, Chesterman CN, Chong BH. Prospective evaluation of
the clinical usefulness of an antigen-specific assay (MAIPA) in idiopathic thrombocytopenic
purpura and other immune thrombocytopenias. Blood. 1996;88:194-201.
Hagenstrom H, Schlenke P, Hennig H, Kirchner H, Kluter H. Quantification of plateletassociated IgG for differential diagnosis of patients with thrombocytopenia. Thromb Haemost.
2000;84:779-83.
Warner MN, Moore JC, Warkentin TE, Santos AV, Kelton JG. A prospective study of proteinspecific assays used to investigate idiopathic thrombocytopenic purpura. Br J Haematol.
1999;104:442-47.
Beardsley DS, Ertem M. Platelet autoantibodies in immune thrombocytopenic purpura.
Transfus Sci. 1998;19:237-44.
Siebers RW, Wakem PJ, Carter JM. Long-term intra-individual variation of platelet parameters.
Med Lab Sci. 1989;46:77-8.
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Supplementary figures
Supplementary Figure 1. Optimizing PKH:CFSE ratio for the micro platelet aggregation test. Different
ratio’s ranging from 0.01:1 to 1:1 were tested to determine the lowest number of test platelets (PKH+)
for detection of micro aggregates after stimulation with PMA. Ratio’s of 0.06:1 to 0.5:1 resulted in
reliable and comparable test outcomes with low background aggregation without stimulus at time
point 0. Using a ratio of 0.01:1 resulted in very few PKH+ and thereby affecting the determination of
aggregation, and 1:1 lead to high numbers of PKH-stained events causing spillover in FACS analysis. A
ratio of approximately 0.10:1 was considered optimal since this required a minimum number of test
platelets while giving similar results.
7
Supplementary Figure 2. Healthy Controls reach similar aggregation levels in an autologous and
allogenic setting.
Platelets obtained from 2 HCs were isolated and tested in the micro platelet aggregation test to
assess whether aggregation levels where influenced by the allogenic nature of the assay compared to
the autologous healthy control. Two HCs were performed in an autologous setting as well as crossed
with each other to test the allogenic setting. No differences in aggregation scores were observed.
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P-selectin
A. ADP max
B. ADP EC50
3
4000
2
ADP-∝M
MFI - AU
3000
2000
1
1000
0
250
50
25
0
10
250
Platelet number
E. CVX MFI max P-selectin
25
10
F. CVX MFI EC50 P-selectin
20
15000
15
10000
CVX-ng/ml
MFI - AU
50
Platelet number
10
5000
0
250
50
25
5
0
10
250
50
25
10
Platelet number
Platelet number
J. TRAP EC50
I. TRAP max
25
15000
TRAP-∝M
MFI - AU
20
10000
15
10
5000
5
0
250
50
25
Platelet number
10
0
250
50
25
10
Platelet number
Supplementary Figure 3. Validation platelet reactivity assay with decreasing platelet number.
Platelets from 3 control subjects were diluted to 250*109/L, 50*109/L, 25*109/L and 10*109/L, by
diluting platelet rich plasma with addition of platelet poor plasma. 5 μL of these samples were
added to 50 μL of buffer containing RPE-labelled anti-P-selectin antibodies, Alexa Fluor 488-labeled
fibrinogen, and serial concentrations of ADP, convulxin (CVX) or TRAP. Maximal mean fluorescence
intensity (MFI) in arbitrary units (AU) and concentration of the agonists needed to obtain half maximal
MFI are displayed for both platelet P-selectin expression (A, B, E, F, I, J) and opening of GPIIbIIIa
receptor (C, D, G, H, K, L). Each line represents the results obtained for individual donor. These
related samples were compared by Wilcoxon signed ranks tests, showing no significant differences
when platelet number was reduced from 250*109/L to 10*109/L.
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Open GPIIbIIIa
D. ADP EC50
4
6000
3
ADP-∝M
MFI - AU
C. ADP max
8000
4000
2
1
2000
0
250
50
25
0
10
250
Platelet number
20
CVX-ng/ml
MFI - AU
10
25
6000
15
4000
10
2000
5
250
50
25
0
10
250
Platelet number
50
25
10
Platelet number
L. TRAP EC50
K. TRAP max
10
4000
8
MFI - AU
TRAP-∝M
5000
3000
2000
7
6
4
2
1000
0
25
H. CVX MFI EC50
G. CVX MFI max
8000
0
50
Platelet number
250
50
25
10
Platelet number
0
250
50
25
10
Platelet number
Supplementary Figure 3. Continued
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max P-selectin to ADP - AU
4000
Mild
Severe
3000
2000
1000
0
0
50
100
150
200
250
platelet number - *109/L
Supplementary Figure 4. Correlation platelet reactivity with platelet count within study population.
No correlation is observed between platelet number and ADP induced maximal platelet reactivity
(Spearmans rho 0.165, p=0.367) within study population (n=33).
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Chapter 8
Capillary blood sampling
in the assessment of platelet responsiveness
Esther R. van Bladel,1* Lieke H.H. Meeter,1* Roger E.G. Schutgens,2 Philip G. de Groot,1 Mark Roest1
1
Department of Clinical Chemistry and Haematology and 2Van Creveldkliniek,
University Medical Centre Utrecht, Utrecht, the Netherlands.
* equally contributed
In preparation
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Abstract
Introduction: The platelet reactivity assay is a novel test to measure specific platelet
receptor pathways, which requires only 250 µL unprocessed whole blood. This volume can
be easily obtained with capillary blood sampling. The aim of this study was to determine
the feasibility of capillary blood sampling in platelet function testing with the platelet
reactivity assay as an alternative for venous sampling. Methods: Platelet reactivity to
ADP, convulxin, TRAP, U46619 and iloprost was quantified with FACS analysis of platelet
P-selectin expression and platelet GPIIbIIIa opening. In the current study, we compared
the agreement between venous and capillary collected blood samples from the same
individuals. Results: We found lower platelet counts with higher MPV, higher basal level of
activity and lower platelet reactivity in capillary sampled blood, suggesting that capillary
sampling leads to platelet aggregation. By addition of dRGDW peptide, a peptide blocking
aggregation without influencing platelet activation, aggregation of platelets as a result of
capillary sampling is prohibited, and platelet activation can be measured using P-selectin
expression in response to different agonists. Conclusion: With the minor modification of
dRGDW addition to the sample tube, we here introduce a new assay for functional platelet
testing in capillary sampled whole blood. With this assay, new opportunities are created for
functional platelet testing in patient categories in which venipuncture is not possible or a
less invasive method is desirable.
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Introduction
New tests for the measurement of platelet function in unprocessed whole blood are on
their way. These new tests make capillary blood sampling a feasible option for platelet
function measurements. Diagnostic platelet function tests, such as aggregometry have
several limitations, including the requirement of large blood volumes and the requirement
of laborious multiple processing steps to prepare platelet rich plasma or to get washed
platelets.1 Other routine diagnostic tests, such as the bleeding time, are non-specific and
do not discriminate between different disorders of primary hemostasis. Moreover, the
bleeding-time is a painful and unfriendly test for the patient, leaving scars.
The platelet reactivity assay2 is a novel assay to test specific platelet receptor pathways
in small amounts of unprocessed whole blood. In this assay, platelets are stimulated with
concentration series of (ant-)agonists. Platelet reactivity is determined by fluorescent
ascending cell sorting (FACS) analysis after addition of fluorescent labeled anti-P-selectin
antibodies, representing platelet degranulation, and simultaneous FACS analysis of GPIIbIIIa
opening via binding of fluorescent labeled fibrinogen. 250 µl of whole blood is enough
to test the 5 main platelet receptor pathways. Moreover, a whole blood platelet count ≥
10*109/L is sufficient for testing platelet function with the platelet reactivity assay, making
it possible to test platelet function in patients with thrombocytopenia.3,4
Currently, blood is standard collected by venous sampling. As the current platelet
reactivity assay requires only 250 µl of whole blood, this volume can be easily obtained
with capillary blood sampling. Furthermore, by using capillary sampling it would be possible
to address new cohorts of patients such as very young children or adults with poor veins,
in which currently analysis of platelet function is not always possible. Other advantages
are diminishing invasiveness and thereby complications of the puncture, reducing costs,
improving patient convenience and ease of application and thereby minimizing required
skills.5,6,7
A potential limitation of capillary sampling might be provocation of hemolysis,
coagulation and/or hemodilution, which might disturb the test.8 These problems could be
prevented by applying the right technique, avoid excessive squeezing and discarding the
first droplet of blood.9,10
The aim of this study was to establish the feasibility of capillary blood sampling in platelet
function testing with the platelet reactivity assay as an alternative for venous sampling.
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Materials and methods
Subjects
Blood was collected from voluntary, healthy employees and interns of the University
Medical Centre Utrecht (UMC Utrecht). Participants were excluded when they used a
non-steroidal anti-inflammatory drug in the 10 days before withdrawing blood, as these
influence platelet function.
In addition, a 1.5 year old infant suspected of a primary bleeding disorder was included
in this study. He had suffered prolonged bleeding after the heel prick test (a screening test
performed in all Dutch newborns), bleeding after umbilical cord separation and a hematoma
on the hand 4 weeks after intravenous puncture. The infants’ mother was known to have
storage pool disease. All volunteers and the infants’ parents gave written informed consent.
Approval by the Medical Ethics Committee of the UMC Utrecht was provided.
Materials
For the capillary samplings BD Microtainer Contact-Activated Lancets with 2.0x1.5 mm
needles (Becton, Dickinson and Company, Franklin Lakes, U.S.A.) were used to puncture the
skin and blood was collected into 0.5 ml minicollect tubes containing 3.2% sodium citrate
(Greiner Bio-One, Alphen a/d Rijn, The Netherlands). In venipunctures, blood was collected
into BD Vacutainers of 4.5 ml with 0.5 ml (0,105 M) sodium citrate (Becton, Dickinson and
Company, Plymouth, United Kingdom).
R-phycoerythrin (PE) labelled mouse anti-human CD 62P antibodies (BD Biosciences,
Franklin Lakes, U.S.A.) to detect P-selectin and fibrinogen from human plasma labeled with
Alexa Fluor 488 (Invitrogen, Eugene, Oregon, U.S.A.) to detect open Glycoprotein IIbIIIa
(GPIIbIIIa) were used for Fluorescence Activated Cell Sorting (FACS) analysis.
HEPES buffered saline was prepared with 4-(2-hydroxyethyl)-1-piperazineethanesulfonic
acid (HEPES) purchased from BDH (Poole, U.K.), sodium chloride (NaCl) from Sigma
(Zwijndrecht, The Netherlands) and magnesium sulphate (MgSO4) and potassium chloride
(KCl) were both obtained via Riedel (Seelze, Germany). As (ant)agonists adenosine
diphosphate (ADP) from Roche (Almere, The Netherlands), Convulxin (Pentapharm,
Basel, Switzerland), U46619 (Santa Cruz Biotechnology, Heidelberg, Germany), thrombin
receptor-activated peptide (TRAP) from Bachem AG (Bubendorf, Schwitzerland) and
Iloprost (Ilomedine) from Bayer Schering Pharma AG (Berlin, Germany) were used. The RGDcontaining peptide Darginyl-glycyl-L-aspartyl-L-tryptophane (dRGDW) was synthesized
at the NKI (Amsterdam, the Netherlands). Formaldehyde was obtained from Calbiochem
(Merck, Darmstadt, Germany).
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Blood sampling
Venous blood was drawn by venipuncture of an antecubital vein using 21G needles, into a
4.5 ml sodium citrate (3.2%) Vacutainer® tube. Venipuncture was performed under minimal
tourniquet pressure.
Prior to capillary sampling the hand was warmed with hot water to improve local blood
circulation. The finger was cleaned with an alcohol swab and dried with gauze. Subsequently,
capillary puncture was made at the lateral aspect of the third or fourth fingertip using
a 2.0 x 1.5 mm lancet. The first drop of blood was wiped from the skin, hereafter 0.5 ml
blood was collected into a capillary tube. Number and size (mean platelet volume, MPV) of
platelets were determined using a Cell-dyn 1800 (Abbott Laboratories, USA).
For the experiments in which dRGDW was added, 1 µL dRGDW (100mM) was added to
the sodium citrate directly before capillary sampling, resulting in a concentration of 200mM
dRGDW after collection of blood in the 500 µL tubes. Since venous sampling was performed
in vacuum sodium citrate tubes, the dRGDW could not be added before blood collection,
and was therefore added directly after venous sampling, to the equal final concentration
of 200 mM.
Platelet activation and responsiveness assay
To measure the activation and responsiveness of platelets, concentration series were
prepared. The assay is a modification of the assay as we described previously.2 In short 50
µl samples were prepared containing 2 µl labelled Anti-P-selectin, 0.5 µl labelled fibrinogen
and ascending concentrations of an agonist in HEPES buffered saline (10mM HEPES, 150
mM NaCl, 1mM MgSO4 and 5mM KCl; pH adjusted to 7.4; filtered through a 0.20 µm filter).
Anti-P-selectin was used as an indicator for degranulation of the platelets, fibrinogen to
bind open GPIIbIIIa receptors.
The agonists ADP, TRAP, convulxin (an agonist for GPVI-receptors) and U46619 (an agonist
of thromboxane receptors) were used in series of eight dilutions. Final concentrations of the
ADP dilutions were 125.00; 31.25; 7.81; 1.95; 0.49; 0.12; 0.03; 0.01 µM. For TRAP, dilutions
of 625.00; 156.25; 39.06; 9.77; 2.44; 0.61; 0.15; 0.04 µM were produced. Concentrations of
Convulxin were 39.06; 9.77; 2.44; 0.61; 0.15; 0.04; 0.01; 0.002 ng/ml. The obtained serial
dilution of U 46619 was 142.65; 47.55; 15.85; 5.28; 1.76; 0.59; 0.20; 0.07 µM. The series
containing an agonist as well as an antagonist contained the stated concentrations of TRAP
as well as 4.75 ng/ml Iloprost (a prostacyclin analogue). Lastly, to determine the basal
platelet activity a control solution was prepared with neither agonists nor antagonists.
To each sample of these serial dilutions 5µl blood was added. After an incubation time of
20 minutes, these samples were fixed with 500µl of 0.2% formylsaline (0.2% formaldehyde
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in 0.9% sodium chloride; filtered through a 0.20 µm filter). After 10 minutes of fixation the
samples were stored at 4˚C until measurement.
Subsequently, 100µl of sample together with another 100µl 0.2% formylsaline was
analyzed by a FACS Canto II Flow Cytometer (BD Biosciences, San Jose, CA, U.S.A.). Using
forward and side scatter properties, single platelets were gated of which the median
fluorescence intensity of platelet P-selectin expression and of open GPIIbIIIa receptor on
platelets was determined.
Data analysis
Raw data were quantified with BD FACSDiva software 6.1.2 (BD Biosciences, San Jose, CA,
U.S.A.). GraphPad Prism 5.03 (GraphPad Software, San Diego, CA, U.S.A.) was used to draw
a sigmoid curve from which minimal and maximal responses were gathered and the agonist
concentrations leading to a response halfway minimal en maximal activation (EC50) were
calculated. Area under the curve (AUC) was represented by the sum of responses to the
eight ascending concentrations of agonist.
Subsequently, the variables (EC50, AUC, minimal and maximal response) were analyzed
with IBM SPSS Statistics 20.0.0 for Windows (International Business Machines Corporation,
New York, U.S.A.). Data are displayed as median and interquartile range (IQR) unless
otherwise specified. Wilcoxon Signed Ranks Test was used for comparing capillary sampling
with venous blood. P-values lower than 0.05 were considered statistically significant.
Results
Blood was collected via both venipuncture and capillary sampling from 5 healthy subjects,
for comparison of these two sampling methods. We found a lower platelet count (105 G/L
(58-136) versus 235 G/L (205-263); p=0.043) and a higher mean platelet volume (7.50 fL
(7.30-8.15) versus 6.90 fL (6.65-7.70); p=0.042) in capillary sampled blood compared to
venous sampled blood. Analyzing the basal level of platelet activity, capillary sampled
platelets had both higher P-selectin expression (MFI 54.00 AU (42.50-168.50) versus
8.50 AU (7.50-18.25); p=0.043) and higher open GPIIbIIIa (MFI 21.00 AU (9.00-34.00) versus
4.50 AU (-9.25-7.50); p=0.080). Also, platelet reactivity in capillary sampled blood was lower.
Platelets needed higher concentrations of ADP, convulxin and TRAP to induce both half
maximal P-selectin expression and half maximal expression of open GPIIbIIIa. Additionally,
platelets reached lower levels of maximal activation after stimulation with ADP, TRAP and
U46619. This all resulted in lower areas under the curve for platelet reactivity for all tested
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agonists. These results suggest significant platelet aggregation already occurred during
capillary blood sampling, leaving only the non-aggregated, less responsive platelets in the
sample for analysis. Results are shown in Table 1.
Table 1. Venous versus capillary sampling
Basal level of platelet activation
P-selectin expression, AU
open GPIIbIIIa, AU
Platelet reactivity to ADP – P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to ADP – open
GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to CVX – P-selectin
Maximal MFI, AU
EC50, ng/ml
AUC, AU
Platelet reactivity to CVX – open
GPIIbIIIa
Maximal MFI, AU
EC50, ng/ml
AUC, AU
Platelet reactivity to TRAP – P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to TRAP – open
GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, AU
Venous sampling
(n=5)
Capillary sampling
(n=5)
Venous vs.
capillary
sampling
p-value
8.50 (7.50-18.25)
4.50 (-9.25-7.50)
54.00 (42.50-168.50) 0.043
21.00 (9.00-34.00)
0.080
2035 (1175-2767)
1.32 (0.68-2.51)
7643 (4210-11631)
231 (164-814)
3.28 (2.73-3.79)
915 (708-3301)
0.043
0.080
0.043
3225 (1548-3743)
609 (468-1211)
0.50 (0.43-0.95)
1.88 (0.99-2.21)
13252 (6023-16620) 2278 (1916-4254)
0.043
0.043
0.043
8717 (5194-9332)
7796 (5234-8044)
9,12 (5,23-14,26)
22,96 (12,47-51,81)
10186 (8932-15832) 5605 (5077-12979)
0.225
0.043
0.043
5590 (3100-9473)
7.36 (6.56-14.23)
8588 (5384-13732)
0.686
0.144
0.043
6262 (3255-10140)
20.83 (6.34-53.32)
6452 (2981-7263)
8
7957 (5706-9939)
5222 (3748-7961)
0.043
1.93 (1.69-3.98)
5.15 (4.26-15.76)
0.043
31326 (26113-47619) 15827 (14357-32325) 0.043
2136 (1480-2280)
1.84 (1.63-4.67)
8127 (7065-10769)
604 (465-797)
2.98 (2.68-8.00)
2670 (1758-3901)
0.043
0.043
0.043
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Table 1. Continued
Venous sampling
(n=5)
Platelet reactivity to TRAP(+iloprost) –
P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to TRAP(+iloprost) –
open GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to U46619 – P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to U46619 – open
GPIIbIIIa
Maximal MFI, AU
EC50, μM
AUC, AU
Trombocyte count, G/L
Mean platelet volume, fL
Capillary sampling
(n=5)
Venous vs.
capillary
sampling
p-value
6260 (5119-8263)
4459 (3292-7623)
0.043
7.12 (6.61-11.44)
8.58 (7.46-17.51)
0.043
21503 (18512-31982) 16076 (10391-29201) 0.043
198 (110-233)
4.25 (3.03-6.73)
798 (410-985)
165 (67- 208)
3.19 (2.95-8.32)
726 (275-932)
0.080
0.500
0.225
5244 (4909-6999)
987 (553-2360)
7.15 (1.70-12.72)
4.95 (2.79-12.08)
16351 (13657-31151) 3227 (1860-8685)
0.043
0.893
0.043
4832 (3762-5818)
6.33 (1.76-12.72)
15455 (12972-21109)
235 (205-263)
6.90 (6.65-7.70)
0.080
0.225
0.080
0.043
0.042
318 (233-2602)
2.29 (1.43-3.63)
1445 (1039-9907)
105 (58-136)
7.50 (7.30-8.15)
Values are expressed as median (interquartile range). Differences are assessed with the Wilcoxon
signed ranks test.
AU, arbitrary units; MFI, Median Fluorescence Intensity of all platelets; EC50, concentration
generating a response halfway between baseline and maximal MFI signal; AUC, area under the curve;
ADP, adenosine diphosphate; CVX, Convulxin; TRAP, thrombin receptor activator peptide; n, sample
size; min, minutes; vs., versus.
In the platelet reactivity assay, platelet-platelet interaction (= aggregation) is not needed
during platelet function testing. To prevent the undesirable pre-test aggregation of platelets
after capillary sampling, we repeated the experiments with dRGDW peptide present in the
collecting tube, a peptide blocking open GPIIbIIIa and thereby blocking aggregation.14 For
this experiment, blood was collected via both venipuncture and capillary sampling in 4
healthy donors. With the addition of dRGDW peptide, both platelet count (205 G/L (150258) versus 217 (167-246); p=0.715) and mean platelet volume (7.60 fL (6.83-8.83) versus
7.45 fL (6.58-8.55); p=0.141) stayed equal in capillary sampled blood compared to venous
sampled blood. Moreover, no significant differences could be observed in basal level of
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Table 2. Venous versus capillary sampling in presence of RGD peptide
Basal level of platelet activation
P-selectin expression, AU
Platelet reactivity to ADP – P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to CVX – P-selectin
Maximal MFI, AU
EC50, ng/ml
AUC, AU
Platelet reactivity to TRAP – P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to TRAP(+iloprost) –
P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Platelet reactivity to U46619 –
P-selectin
Maximal MFI, AU
EC50, μM
AUC, AU
Trombocyte count, G/L
Mean platelet volume
Venous sampling
(n=4)
Capillary sampling
(n=4)
Venous vs.
capillary
sampling
p-value
12.50 (16.00-18.38)
27.38 (34.75-39.13)
0.068
726 (715-912)
0.80 (0.68-0.81)
3122 (3060-4115)
840 (760-974)
0.95 (0.77-1.16)
3439 (3142-4211)
0.068
0.068
0.465
6063 (5468-7245)
4.29 (3.45-6.51)
12939 (10375-15895)
6675 (5805-7385)
0.144
5.03 (4.38-7.69)
0.068
13557 (10365-15404) 1.000
5959 (5549-8060)
2.06 (1.81-2.30)
29123 (26593-35000)
6438 (5755-7173)
0.715
2.26 (2.00-2.31)
0.144
30738 (27957-32068) 0.715
5418 (4990-6611)
6.63 (5.66-7.59)
20723 (18900-25526)
5713 (5121-6360)
0.715
6.59 (6.12-7.24)
0.715
23569 (20599-25839) 0.144
4367 (3516-5273)
5.52 (2.49-10.91)
16765 (11169-19039)
217 (167-246)
7.45 (6.58-8.55)
4261 (2468-4824)
3.61 (2.85-7.10)
15827 (8528-18872)
205 (150-258)
7.60 (6.83-8.83)
0.273
0.273
0.144
0.715
0.141
Values are expressed as median (interquartile range). Differences are assessed with the Wilcoxon
signed ranks test.
AU, arbitrary units; MFI, Median Fluorescence Intensity of all platelets; EC50, concentration
generating a response halfway between baseline and maximal MFI signal; AUC, area under the curve;
ADP, adenosine diphosphate; CVX, Convulxin; TRAP, thrombin receptor activator peptide; n, sample
size; min, minutes; vs., versus.
8
platelet P-selectin expression and platelet reactivity to ADP, TRAP, convulxin, iloprost and
U46619. Results are shown in Table 2.
Finally, we studied the feasibility of this test in a 1.5 year child suspected of having a
platelet disorder. We tested platelet reactivity in capillary sampled blood of this patient,
and compared it to capillary samples from healthy controls. For this patient, platelet
Capillary blood sampling in the assessment of platelet responsiveness | 127
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reactivity to ADP, convulxin and U46619 was comparable to the control group. However,
platelet reactivity to TRAP was reduced. Compared to controls a higher TRAP concentration
was needed to yield a half maximal activation of platelets from this patient. Results are
shown in Figure 1.
Convulxin
ADP
8000
MFI P-selecti n - AU
MFI P-selecti n - AU
2000
1500
1000
500
0
10-3
10-2
10-1 100 101
ADP - ∝ M
102
6000
4000
2000
0
10-3
103
TRAP
MFI P-selecti n - AU
MFI P-selecti n - AU
4000
2000
10-1
100
101
TRAP - ∝ M
102
103
U46619
5000
MFI P-selecti n - AU
102
8000
6000
4000
3000
2000
1000
0
10-2
10-2
10-1
100
101
Conv ul xin - ng /mL
TRAP + iloprost
8000
0
10-2
Patient
10-1
100
101
U46619 - ∝ M
102
103
6000
4000
2000
0
10-2
10-1
100
101
TRAP - ∝ M
102
103
Figure 1. Platelet reactivity in capillary blood in
1.5 year old infant
The gray area between the dotted lines show
the interquartile range of platelet reactivity
for the control group, which consists of
4 healthy controls in which capillary sampling
and subsequent platelet reactivity testing was
performed repeatedly at 5 different timepoints.
The black line shows the infant suspected of
a primary bleeding disorder. All assays were
performed in the presence of dRGDW peptide
Agonist concentration is presented at the
X-axis; median fluorescent intensity in arbitrary
units (AU) of anti-P-selectin antibodies bound to
platelets is shown on the Y-axis.
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Discussion
In this study we show that with minor modifications, capillary blood sampling can be used
to determine platelet function with the platelet reactivity assay. By the addition of dRGDW
peptide, aggregation of platelets as a result of capillary sampling is prohibited, and platelet
degranulation can be measured in response to different agonists. This opens new doors
for platelet function assessment in patient categories like infants, in which classic platelet
function tests cannot always be performed.
When comparing platelet reactivity after venous or capillary sampling, we found lower
platelet counts with higher MPV, higher basal level of activity and lower platelet reactivity
in capillary sampled blood, suggesting that capillary sampling leads to platelet aggregation.
A possible explanation for platelet aggregation occurring after capillary sampling is via
the mechanical stress induced by squeezing which is necessary to obtain 500μL blood by
capillary sampling. Furthermore, with this technique blood passes tissue before collection,
whereas in venipuncture blood directly flows from the vein into the needle and collection
tube. While passing tissue, blood comes into contact with tissue fluids, which contain
platelet-activating proteins.11
In the past, several studies were done on blood counts in capillary sampled blood.
Formation of aggregates after capillary sampling is a known problem due to platelet
activators in tissue fluid.11 Platelet count is commonly comparable in venous and capillary
sampled blood when using ethylenediaminetetraacetic acid (EDTA) as anticoagulant.7,12,13
However, since EDTA is a strong chelating agent for calcium, it is not possible to perform
functional platelet assays in blood collected in EDTA. Therefore, we chose to add dRGDW
(D-Arg-Gly-Asp-Trp) peptide to sodium-citrate to prevent platelet aggregation.14 This
synthetic peptide blocks the GPIIbIIIa receptor and thereby aggregation. However,
other manifestations of platelet activation, like platelet degranulation and opening of
the GPIIbIIIa receptor are not influenced by addition of dRGDW peptide. The results of
our study show, that when dRGDW peptide is added to collection tubes before capillary
sampling, blood count and MPV stay comparable to venous samples. Also, basal level of
P-selectin expression stays equivalent when dRGDW peptide is added. Moreover, platelet
reactivity to the platelet agonists ADP, CVX, TRAP and U46619 is the same in capillary and
venous sampled blood when dRGDW peptide is added. Hereby we show that through the
addition of dRGDW peptide to the collection tube before sampling, platelet aggregation as
a result of capillary sampling is prohibited. Thus capillary sampled blood can be used for
functional platelet testing, giving results equivalent to venous samples.
8
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The addition of dRGDW peptide does come along with some limitations. First, because
of its binding to open GPIIbIIIa, the binding of fibrinogen to open GPIIbIIIa is blocked and
can no longer be used as an outcome measurement of platelet activation or reactivity.
However, this problem can be circumvented by using PAC-1 or comparable antibodies,
antibodies that specifically recognize the open conformation of GPIIbIIIa.15 Second, dRGDW
is known to have an influence on platelet reactivity via enhancement of GPVI-associated
signaling pathways when GPVI pathway is activated.16 Therefore, samples in which dRGWD
is added can only be compared validly to other patient or control samples with dRGDW
peptide.
A disadvantage of blood withdrawal by capillary sampling is pain due to the puncture.
In a study into serum 25 hydroxyvitamin D assessment in capillary blood, adult volunteers
were asked to rate the painfulness of both venipuncture and capillary sampling.5 The
capillary sampling did cause more pain than the venipuncture (mean of 1.20 respectively
0.79 on a scale of 10). Although this was a statistical difference, the absolute difference is
small and might not be clinically relevant.
In conclusion, we here introduce a new assay for functional platelet testing in capillary
sampled whole blood. With the addition of dRGDW peptide to the collection tube before
capillary sampling, platelet aggregation is prevented and capillary sampled blood can be
used for assessment of platelet reactivity. With this minor modification, new opportunities
are created for functional platelet testing in patient categories in which venipuncture is not
possible or a less invasive method is desirable.
References
1.
2.
3.
4.
5.
Rand ML, Leung R, and Packham MA. Platelet function assays. Transfusion and Apheresis
Science. 2003;28(3):307-17.
Van Bladel ER, Roest M, de Groot PG, Schutgens R. Up-regulation of platelet activation in
hemophilia A. Haematologica. 2011;96(6)888-95.
van Bladel ER, Laarhoven AG, van der Heijden L, Heitink-Pollé KM, Porcelijn L, van der Schoot
CE, de Haas M, Roest M, Vidarsson G, de Groot PG and Bruin MC. Functional platelet defects
in children with severe chronic ITP: as tested with two novel assays applicable for low platelet
counts. [Chapter 7; in revision].
Roest M, van Holten TC, Fleurke GJ, Remijn JA. Platelet Activation Test in Unprocessed Blood
(Pac-t-UB) to Monitor Platelet Concentrates and Whole Blood of Thrombocytopenic Patients.
Transfus Med Hemother. 2013;40(2):117-125.
Dayre McNally J, Matheson LA, Sankaran K, Rosenberg AM. Capillary blood sampling as an
alternative to venipuncture in the assessment of serum 25 hydroxyvitamin D levels. J Steroid
Biochem Mol Biol. 2008;112:164-168.
130 | Chapter 8
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6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
Simmonds MJ, Baskurt OK, Meiselman HJ, Marshall-Gradisnik SM. A comparison of capillary
and venous blood sampling methods for the use in haemorheology studies. Clin Hemorheol
Microcirc. 2011;47:111-119.
Schalk E, Heim MU, Koenigsmann M, Jentsch-Ullrich K. Use of capillary blood count parameters
in adults. Vox Sang. 2007;93:348-353.
Shah V, Ohlsson A. Venepuncture versus heel lance for blood sampling in term neonates.
Cochrane Database Syst Rev. 2007;4:CD001452.
Meites S. Skin-puncture and blood-collecting technique for infants: update and problems. Clin
Chem. 1988;34:1890-1894.
Pal GK. Textbook Of Practical Physiology - 2Nd Edn: Orient Longman Private Limited; 2006.
Hoffmann JJML, Akkerman JWN, Nieuwenhuis HK, Overbeeke MAM. Hematologie (ed 2).
Arnhem: Syntax Media; 2006.
Yang ZW, Yang SH, Chen L, Qu J, Zhu J, Tang Z. Comparison of blood counts in venous, fingertip
and arterial blood and their measurement variation. Clin Lab Haematol. 2001;23:155-159.
Daae LN, Halvorsen S, Mathisen PM, Mironska K. A comparison between haematological
parameters in ‘capillary’ and venous blood from healthy adults. Scand J Clin Lab Invest.
1988;48:723-6.
Plow EF, Pierschbacher MD, Ruoslahti E, Marguerie GA, Ginsberg MH. The effect of Arg-GlyAsp-containing peptides on fibrinogen and von Willebrand factor binding to platelets. Proc
Natl Acad Sci USA. 1985;82:8057-61.
Shattil SJ, Hozie JA, Cunningham M, Brass LF. Changes in the platelet membrane glycoprotein
IIb.IIIa complex during platelet activation. J Biol Chem. 1985;260:1107-14.
Jones ML, Harper MT, Aitken EW, Williams CM, Poole AW. RGD-ligand mimetic antagonists
of integrin αIIbβ3 paradoxically enhance GPVI-induced human platelet activation. J Thromb
Haemost. 2010;8:567-76.
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Chapter 9
General discussion
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Platelet reactivity assay
Classical assays for testing platelet function are laborious and require large amounts of
blood. We developed an assay for platelet function testing, the platelet reactivity assay,
that is whole blood based, leading to fast and limited processing of samples. Moreover,
with only 250 µL of whole blood, the platelet function of the 5 major receptor pathways
can be tested.
With the (ant)agonists ADP, CRP or convulxin , TRAP, iloprost and U46619, the pathways
of the P2Y receptors (P2Y1 and P2Y12), collagen receptor (glycoprotein VI, GPVI), thrombin
receptor (Proteinase Activated Receptor-1; PAR-1), prostacyclin receptor and thromboxane
receptor were tested, respectively. For each pathway, multiple manifestations of platelet
activation are quantified simultaneously, i.e. platelet P-selectin expression as a measure of
platelet degranulation and fibrinogen binding as a measure of the conformational change
in the glycoprotein IIbIIIa (GPIIbIIIa) receptor to an open, pro-aggregation, state. Since both
the receptor pathways and manifestations of platelet activation are measured separately,
this assay gives specific insight into specific platelet function defects.
However, a limitation of this specificity is that only the defects in the isolated routes
are detected, not the overall picture of platelet function. In Chapter 1, the manifestations
of platelet activation are described: shape change, degranulation, thromboxane formation,
GPIIbIIIa opening/aggregation and procoagulant activity. With our developed assay, not
all manifestations were measured. However, the opportunity to expand the assay exists.
Depending on the flow cytometer used for analysis, multiple fluorescent signals, and thus
multiple different labels, can be measured simultaneously. For example, with the addition
of fluorescent labelled Annexin V or lactadherin to the assay, the procoagulant activity of
platelets could also be determined.
Thromboxane formation was not measured in the assay. We have tried to use
arachidonic acid but due to the hydrophobic nature of this substrate, we were unable to
obtain reproducible results. Nevertheless the platelet response to U46619, stimulating the
thromboxane receptor directly, was tested. In this way the down-stream signalling of the
thromboxane pathway could be tested. By the addition of an exogenous agonist to the
thromboxane receptor, this assay is not sensitive to drugs inhibiting COX-1, which influence
the proximal segment of the thromboxane pathway in which arachidonic acid is converted
into thromboxane. Therefore, the assay is only indirectly influenced by the use of aspirin or
other COX-1 inhibitors.
With minor adjustments (Chapters 7 and 8), the platelet reactivity assay can be
performed in blood from thrombocytopenic patients and in blood collected with capillary
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sampling. Since the assay is based on measurement of manifestations of platelet activation
in single platelets, and not on aggregation of platelets, it is not disturbed when platelet
count is low, as shown in Chapter 7. Low platelet counts do lengthen the duration of the flow
cytometer measurement, since it counts a certain amount of cells per time unit. However,
by lysing the erythrocytes, after activating and fixating platelets, the measurement speed
is comparable to samples with normal platelet counts.
Since platelet aggregation does not occur in the platelet reactivity assay, the assay
can be performed in blood containing dRGDW, an aggregation blocker, without disturbing
the detection of platelet degranulation. In Chapter 8, we show that with this addition, the
negative effects of capillary sampling, pre-activation of platelets, can be prevented. In this
way, capillary sampled blood can be used for the assessment of platelet reactivity, making
functional platelet testing available for new patient categories in which venipuncture is not
possible or undesirable.
Platelets in hemophilia A
Hemophilia A patients suffer from a defect in coagulation factor VIII (FVIII), leading to a
tendency to bleed. Although on a group level residual FVIII activity is the main determinant
of bleeding phenotype, the bleeding phenotype in individual patients is not solely
dependent on their residual factor VIII activity.1-3 Studies into variation in other factors of
secondary hemostasis were not able to account for the variability in bleeding phenotype.4,5
Therefore we hypothesized that variation in platelet function could be responsible for
variation observed in bleeding phenotype.
To investigate this hypothesis, we first wanted to determine if platelet function in
hemophilia patients differs from platelets function in healthy men (Chapter 2). This
study indicated that patients with diagnostic severe hemophilia A (FVIII:C <1 IU/dL) had
a higher basal level of activated platelets in their circulation compared to healthy men,
shown by a higher baseline level of P-selectin expression and higher plasma concentrations
of platelet activation markers PF4, CXCL7 and RANTES. Within the group of diagnostic
severe hemophilia A (FVIII:C <1 IU/dL), platelet activation inversely correlated with FVIII
consumption, suggesting that a higher basal level of platelet activation could be correlated
to a milder bleeding phenotype.
This observation was further studied in Chapter 4. There, we only included diagnostic
severe hemophilia A patients (FVIII:C <1 IU/dL), which were selected on their bleeding
phenotype. Where in Chapter 2, we only collected data on FVIII consumption to get an
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impression of bleeding phenotype, in Chapter 4 we selected patients based on a bleeding
phenotype score integrating onset of joint bleeding, arthropathy, joint bleeding frequency
with FVIII consumption. In contrast to the correlation found in Chapter 2, platelet activation
was shown not to correlate with bleeding phenotype. During this study, Teyssandier et
al.6 published a study comparing platelet activation and reactivity to TRAP between
hemophilia mice and wildtype mice. In this study, no differences between hemophilia and
wild type mice could be observed. In view of the more comprehensive study design in
Chapter 4, and the results obtained by others, we must conclude that platelets are not the
major determinants for the differences observed in the bleeding phenotype of hemophilia
A patients.
As introduced in Chapter 1, next to primary and secondary hemostasis, the vascular
wall is a major player in hemostasis. Function of endothelial cells, lining the vessel wall, can
be studied in blood via investigation of factors released by these cells. When measuring
vWF levels, we found increased levels in hemophilia A patients compared to healthy men.
The vWF/vWFpropeptide ratio in severe hemophilia A patients was increased, suggesting
the increase in vWF plasma level to reflect a decreased clearance rather than increased
secretion by endothelial cells (Chapter 2). However, factors released by endothelium only
give a partial insight into the function of these cells. Moreover, the subendothelial matrix
and vasoconstrictive response to injury were not measured in our studies.
Bleeding time tests are influenced by all these hemostatic properties of the vessel wall.
In 1988 Janzarik and collegues7 compared the bleeding times according to Duke8 and Ivy9
to the haemostasis time in 12 hemophilia A patients. The bleeding time tests determine
primary hemostasis in vivo, in which platelets, vWF and the vessel wall can influence the
result. The hemostasis time, in which the occlusion time of a Butterfly 25 short cannula
inserted into the cubital vein is determined, determines primary hemostasis excluding the
vessel wall. Bleeding times in all 12 hemophilia A patients were normal, however, they
found a prolonged haemostasis time in 7/12 hemophilia A patients. The hemostasis time
did not correlate with the patients residual factor VIII activity, but it did correlate to the
age of onset of bleeding. Therefore they suggested that the vessel wall does not cause
the variation in the bleeding tendency observed in hemophilia A patients. However, the
test used in this study, all started with an external trauma to the vessel. Already in 1964,
Fulton and Berman10 wondered if the vessel wall can weaken spontaneously, and if an
abnormal coagulation manifests only secondary to disruption of the integrity of the vessel
wall. This theory on vessel wall integrity might be interesting in view of the question if the
vessel wall influences the frequency of spontaneous bleeds in hemophilia patients. New
developments have led to advancements in studies into the vascular endothelium. Studies
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into endothelial cell damage, vascular stability and the role of circulating endothelial
progenitor cells are increasingly performed in the field of arterial thrombosis.11-14 However,
these factors might also be of importance in maintaining vascular integrity in the scope of
bleeding, and therewith in determining the bleeding tendency of hemophilia A patients.
Next to studies into vessel wall integrity, new assays determining the interactions
between the key players of hemostasis are warranted. Studies into phenotypic variation in
hemophilia A determined quantity and quality of different factors of primary and secondary
hemostasis. However, in vivo the interplay between these factors results in hemostasis.
If the interplay between factors is responsible for variation in bleeding phenotype,
development of a test which could measure complete hemostasis would be helpful in
predicting individual bleeding phenotype. Tests determining the interplay between specific
players of hemostasis will be needed to gain more insight into the physiological processes
underlying the variation in bleeding phenotype, which currently remain unexplained.
However, one of the major determinants of a bleeding phenotype, i.e. life style, has not
been included in our studies. Future studies should take into account this important factor.
Furthermore, determination of bleeding phenotype also has a subjective factor, where
the patients’ judgement is important. In almost all hemophilia studies, the presence of a
bleed, the subsequent treatment and its response to treatment, is judged by the patient
and therefore is not entirely objective.
Severe hemophilia A patients can be treated with prophylactic FVIII concentrate
infusions. Goal of this treatment is to convert a severe hemophilia patient into a mild
to moderate patient, and therewith reduce the number of joint bleeds and associated
arthropathy.15 To determine if infusion of FVIII in a hemophilia A patient influences other
characteristics of hemostasis, we performed a study in which vWF and platelet function
were measured before and 15 and 60 minutes after FVIII concentrate bolus infusion
(Chapter 3). We found FVIII infusion to induce a decrease in vWF plasma levels, together
with a decrease in ADAMTS-13 and a decrease in number of circulating activated platelets.
These findings are in line with literature showing FVIII enhances the proteolytic cleavage
of vWF by ADAMTS-13 under shear stress16 and this effect is synergized by GPIb present on
platelets17. Therefore we postulate that binding of FVIII to vWF after its infusion influences
vWF interaction with activated platelets, leading to increased clearance of vWF and (pre)
activated platelets. This could possible also explain why we (Chapter 2) and others18,19 find
increased plasma vWF levels in diagnostic severe hemophilia A patients.
To look into the relation between FVIII and platelets more thoroughly, in Chapter 5
we investigated the ability of platelets and megakaryocytes to take up FVIII. In vitro, we
found the addition of platelet lysates of healthy controls to FVIII deficient plasma shortens
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the APTT of these samples, indicating presence of FVIII in platelets. Addition of FVIII to
the platelet rich plasma before platelet isolation, induces a further reduction in the APTT,
especially when pre-activating platelets. Also, megakaryocyte lysates reduce the APTT
of FVIII deficient plasma when incubated with FVIII. These results show platelets and
megakaryocytes are able to take up FVIII.
To study if the observations in vitro are also relevant in vivo, we studied platelets from
severe hemophilia A patients (FVIII:C <1 IU/dL) before and after FVIII infusion. Results show
FVIII to be already present before FVIII infusion. The patients included in this study received
FVIII prophylaxis. Therefore, it could well be that the FVIII present in platelets before
infusion reflects uptake from prior infusions. Future studies, including severe hemophilia A
patients who did not receive FVIII infusions for a prolonged period of time, will be necessary
to confirm this.
When comparing FVIII in platelet lysates from hemophilia A patients before and 15 and
60 minutes after infusion, no differences could be observed. This indicates that FVIII uptake
by platelets might not be relevant in vivo. However, since FVIII can be found in platelets
from severe hemophilia A patients who were treated with FVIII prophylaxis, FVIII uptake at
the level of megakaryocytes might be relevant in vivo.
Platelets in chronic kidney disease
In chronic kidney disease, patients have both an increased bleeding tendency20-23 and an
increased risk for thrombosis.24-28 The role of platelet function in these manifestations is
unclear. In Chapter 6, we studied platelet function in patients with chronic kidney disease.
Results show a deficiency in platelet degranulation, independently of the agonist used
for activation, indicating platelets contribute to the bleeding tendency in these patients.
Future studies are needed to investigate if treatment of severe bleeding in chronic kidney
disease should be aimed at improving platelet function, i.e. if platelet transfusions should
be applied.
In our study, P-selectin expression was the only manifestation of platelet activation
measured. Therefore, we cannot confirm if the reduced P-selectin expression is
representative of other manifestations of platelet activation, i.e. shape change,
thromboxane formation, aggregation and procoagulant activity, or if results indicate a
specific problem in platelet degranulation. If a specific defect in platelet degranulation is
present in chronic kidney disease, this could be due to depletion of α-granules or granule
content, or to a deficiency in release of the α-granules from the platelet. Results from
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Schoorl et al29 indicate a depletion of granules in patients undergoing chronic hemodialysis.
Future research, focussing on other manifestations of platelet activation, could give more
insight into the process underlying the reduction in P-selectin expression on activated
platelets we observed.
Of the patients included in our study, 60% was on hemodialysis. With hemodialysis,
toxins like urea which are normally excreted by the kidneys, are extracted during dialysis
usually 3 or 4 times weekly. However, the damage afflicted to platelets by toxins like urea,
does not seem to be reversible. In Chapter 6, we found that platelet function before and
after dialysis did not differ. Although intermittent dialysis reduces toxins to tolerable levels
between dialysis, it does not approximate continuous kidney function. Therefore platelets
suffer toxic effects consistently. More intensive hemodialysis modalities, like short daily
(home) dialysis or nocturnal dialysis, are suggested to have a survival benefit.30 It would be
interesting to study the effects of these more intensive hemodialysis modalities on platelet
function and bleeding tendency.
Platelets in chronic ITP
Classical platelet function assays can only be performed when platelet counts are (near)
normal. With platelet reactivity testing, platelet function is determined at the level of
single platelets. Therefore, platelet count does not influence results, making it possible to
determine platelet function in thrombocytopenic patients. In Chapter 7, platelet function in
children with chronic immune thrombocytopenia (ITP) was studied with both the platelet
reactivity assay and the micro aggregation assay. Patients were classified as having a mild or
a severe bleeding phenotype based on the overall Buchanan bleeding score.31 Patients with
a severe bleeding phenotype were found to have a decreased platelet function compared
to patients with a mild bleeding phenotype, shown by decreased platelet degranulation
following ADP stimulation and higher ADP and convulxin concentrations needed for half
maximal activation in the platelet reactivity assay, and decreased platelet aggregation
following PMA stimulation in the micro platelet aggregation test.
As its name suggests, ITP is thought to be initiated by auto-antibodies recognizing
glycoproteins on the surface of platelets and megakaryocytes, leading to rapid clearance
and decreased platelet counts.32 The disease is characterized by an acute onset, often after a
viral infection in the preceding weeks. In about 25% of pediatric cases, the thrombocytopenia
persist chronically.33-35 The hypothesis of our study is that auto antibodies could influence
not only platelet clearance, but also platelet function, possibly via binding of platelet
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receptors. In Chapter 7, presence of auto antibodies was determined via indirect platelet
immunofluorescence test (PIFT) and the indirect monoclonal antibody immobilization of
platelet antigens (MAIPA). Only few patients showed presence of auto-antibodies via these
tests. However, indirect PIFT and MAIPA are known to have very low sensitivities (25-39%
and 30% respectively) [Brighton et al 1996; Hagenstrom et al 2000; Warner et al 1999].36-38
For the more sensitive direct PIFT and MAIPA, high number of platelets are needed, which
were not available in this study. Therefore, we cannot prove that the differences found
in platelet function are due to effects of auto antibodies. Future research is needed to
determine if auto antibodies are responsible for the diminished platelet function, or that
other different factors add to the observed variation in platelet function.
In different thrombocytopenic disorders, it is noticeable that patients with very low
platelet counts do not necessarily have serious bleeding problems. Apparently, platelets
can cause sufficient primary hemostasis at low numbers. The results from Chapter 7
suggest that in ITP platelet function should be affected before serious bleeding problems
arise. Future studies in different forms of thrombocytopenia, like chemotherapy induced
thrombocytopenia, are necessary to determine if indeed a reduced platelet function
is a requisite before patients are confronted with major bleeds in thrombocytopenia
other than ITP. If platelet function can predict bleeding tendency in chronic ITP or other
thrombocytopenia’s, treatment modalities could be reserved for only those patients at
risk for serious bleeding complications. In that way, adverse effects of treatment can be
prevented in patients not in need of these treatments.
References
1.
2.
3.
4.
5.
6.
Rainsford SG, Hall A. A three-year study of adolescent boys suffering from haemophilia and
allied disorders. Br J Haematol. 1973;24(5):539-51.
Aledort LM, Haschmeyer RH, Pettersson H. A longitudinal study of orthopaedic outcomes for
severe factor-VIII-deficient haemophiliacs. The Orthopaedic Outcome Study Group. J Intern
Med. 1994;236(4):391-9.
Molho P, Rolland N, Lebrun T, et al. Epidemiological survey of the orthopaedic status of severe
haemophilia A and B patients in France. The French Study Group. Haemophilia. 2000;6(1):2332.
van Dijk K, van der Bom JG, Fischer K, et al. Phenotype of severe hemophilia A and plasma
levels of risk factors for thrombosis. J Thromb Haemost. 2007;5(5):1062-4.
van Dijk K, van der Bom JG, Lenting PJ, et al. Factor VIII half-life and clinical phenotype of
severe hemophilia A. Haematologica. 2005;90(4):494-8.
Teyssandier M, Delignat S, Rayes J, et al. Activation state of platelet in experimental severe
hemophilia A. Haematolgica. 2012;97(7):1115-6.
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8.
9.
10.
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12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
Janzarik H, Heinrich D, Bödeker RH, Lasch HG. “Haemostasis time”, a modified bleeding time
test and its comparison with the Duke and Ivy template bleeding times. II. Application in
bleeding disorders. Blut. 1988;57(3):111-6.
Duke WW (1910) The relation of blood platelets to hemorrhagic disease. JAMA. 55:1185-92.
Ivy AC, Shapiro PF, Melnick P. The bleeding tendency in jaundice. Surg Gynecol Obstet.
1953;60:781-4.
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Chapter 10
Nederlandse samenvatting
Dankwoord
Curriculum Vitae
List of publications and awards
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Nederlandse samenvatting
Hemostase
Hemostase is het proces in ons lichaam wat er voor zorgt dat bloed blijft circuleren in de
bloedvaten, en wat er bij schade van zo’n bloedvat voor zorgt dat het ontstane letsel wordt
afgedicht. Er zijn verschillende factoren die een rol spelen in de hemostase: endotheelcellen,
bloedplaatjes en stolfactoren. Endotheelcellen zijn de cellen die de binnenwand van
bloedvaten bekleden. Ze vormen een barrière tussen het bloed en de matrix onder deze
cellen. Wanneer bloed, door een beschadiging van de endotheelcellen, in contact komt
met deze subendotheliale matrix, dan leidt dit tot activering van zowel bloedplaatjes als
stolfactoren. Bij schade aan een slagader, stromen bloedplaatjes met zo’n hoge snelheid
langs de beschadiging dat ze moeite hebben om zich daar te kunnen hechten. Echter,
endotheelcellen produceren het eiwit von Willebrand Factor (vWF), wat onder deze hoge
snelheden een brug kan vormen tussen de subendotheliale matrix en bloedplaatjes, en het
op deze wijze mogelijk maakt voor bloedplaatjes om te hechten en de beschadiging af te
dichten.
Wanneer bloedplaatjes hechten aan de subendotheliale matrix raken ze geactiveerd.
Dit heeft verschillende gevolgen voor de cel:
a. Het bloedplaatje spreidt zich uit om het contact met de matrix te vergroten.
b. Het bloedplaatje stoot zijn granula uit. Hiermee komen onder andere stoffen vrij die
bloedplaatjes in de omgeving kunnen activeren. Tevens komen er eiwitten, die gebonden
zijn aan het membraan van de granula, op het buitenoppervlak van het bloedplaatje
terecht. Eén van deze eiwitten is P-selectine.
c. Het bloedplaatje vormt Thromboxaan, wat de cel uit diffundeert om daar bloedplaatjes
in de omgeving te activeren.
d. De glycoproteïne IIbIIIa (GPIIbIIIa) receptor, een receptor op het oppervlaktemembraan
van bloedplaatjes, gaat open staan. Aan deze open receptor kan fibrinogeen, het
eindproduct van de bloedstolling, of vWF binden, om zo een brug te vormen tussen
2 bloedplaatjes. Hierdoor kunnen bloedplaatjes aan elkaar klonteren om één
bloedplaatjes plug te vormen.
e. Het bloedplaatje brengt negatief geladen fosfolipiden naar het oppervlaktemembraan,
en genereert zo een negatief geladen oppervlakte waarop de bloedstolling kan
plaatsvinden.
Door beschadiging van het endotheel komt er tevens Weefselfactor vrij. Weefselfactor
activeert de bloedstolling. De bloedstolling is een proces waar meerdere stolfactoren bij
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zijn betrokken. Een stolfactor kan, wanneer deze geactiveerd is, één of meerdere stolfactor
activeren, danwel het proces van activering van een specifieke stolfactor versnellen.
Uiteindelijk leidt dit proces tot de omzetting van fibrinogeen in fibrine. Fibrinedraden
vormen vervolgens een netwerk binnen de bloedplaatjesplug, die deze plug verstevigen.
Eén van de geactiveerde stolfactoren, Trombine, is tevens in staat om bloedplaatjes te
activeren via de Protease Activated Receptor (PAR).
Bloedplaatjes reactiviteit
Klassieke laboratoriumtesten die de bloedplaatjesfunctie bepalen zijn erg bewerkelijk en
vereisen grote hoeveelheden bloed. In ons onderzoek hebben we een functionele test
opgezet, waarin de activiteit en reactiviteit van bloedplaatjes in minimale hoeveelheden
onbewerkt bloed bepaald kunnen worden. De test maakt gebruik van de kenmerken die
bij activering van bloedplaatjes optreden. Zo kan er bijvoorbeeld, door toevoeging van
fluorescent gelabelde anti-P-selectine antistoffen, het P-selectine wat na activering op
de buitenkant van bloedplaatjes verschijnt zichtbaar worden gemaakt. Tevens kan, door
fluorescent gelabeld fibrinogeen toe te voegen, de geopende GPIIbIIIa receptor worden
aangekleurd. Door dit bij verschillende concentraties van bekende activatoren te doen, kan
er bepaald worden hoe makkelijk bloedplaatjes van een specifieke patiënt te activeren zijn.
Wanneer dit in afwezigheid van een activator wordt verricht, kan het basale niveau van
activering worden bepaald.
Omdat deze test zo weinig bloed benodigd heeft, hebben we in hoofdstuk 8 onderzocht
of de test ook uitgevoerd kan worden in bloed wat is verkregen via een vingerprik, in plaats
van via een veneuze bloedafname. In eerste instantie zagen we dat bloedplaatjes ten
gevolge van de vingerprik geactiveerd raken. Aangezien de bloedplaatjes reactiviteitstest
niet gebaseerd is op aggregatie van bloedplaatjes, konden we de test aanpassen met
toevoeging van dRGDW peptide, welke aggregatie blokkeert. Door aggregatie te blokkeren
wordt de activering ten gevolge van een vingerprik voorkomen, en kan de functie van
bloedplaatjes gemeten worden in bloed verkregen middels een vingerprik. Dit maakt het
testen van bloedplaatjesfunctie mogelijk in nieuwe patiëntencategorieën waarbij veneuze
bloedafname niet mogelijk of onwenselijk is.
Hemofilie
Patiënten met hemofilie hebben een verminderde of afwezige activiteit van stolfactor
VIII (hemofilie A) danwel IX (hemofilie B). Hierdoor lijden deze patiënten aan een
bloedingsneiging, welke zich kan uiten in bijvoorbeeld bloedingen in spieren en gewrichten.
Op groepsniveau kan men stellen dat de hoeveelheid FVIII activiteit gerelateerd is aan
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de ernst van de bloedingsneiging. Echter, wanneer er op het niveau van de individuele
patiënt wordt gekeken is dit niet altijd het geval. Om te beoordelen of variatie in de
activiteit en reactiviteit van bloedpaatjes de variatie die wordt gezien in bloedingsneiging
van hemofilie A patiënten verklaart, hebben we in hoofdstuk 2 de bloedplaatjes activiteit
en reactiviteit onderzocht bij patiënten met verschillende niveaus van FVIII activiteit en
gezonde controles met normale FVIII activiteit. In deze studie vonden we dat patiënten
met ernstige hemofilie A (geen meetbare FVIII activiteit) een verhoogd basaal niveau van
bloedplaatjes activiteit hebben in vergelijking met gezonde mannen. De patiënten met
ernstige hemofilie A hadden namelijk een hoger niveau van P-selectine expressie op de
bloedplaatjes, en ook hogere plasma concentraties van Platelet Factor 4, CXCL 7 en RANTES,
stoffen die bij activering van bloedplaatjes in het plasma terecht komen. In dit hoofdstuk
hebben we tevens binnen de groep van ernstige hemofilie A patiënten bekeken of er een
relatie bestond tussen de mate van bloedplaatjes activiteit en de hoeveelheid stolfactor
welke door deze patiënten werd geconsumeerd. De stolfactor consumptie van hemofilie
patiënten wordt gedoseerd op basis van hun bloedingsneiging, waarbij patiënten met een
ernstigere bloedingsneiging een hogere consumptie hebben. In deze studie gebruikten
de patiënten met hoger basaal niveau van bloedplaatjes activiteit minder stolfactor
concentraat. Dit suggereerde dat patiënten met een hoger basaal niveau van bloedplaatjes
activiteit een mildere bloedingsneiging hebben. In hoofdstuk 4 hebben we deze observatie
bij ernstige hemofilie patiënten nader onderzocht. Daartoe hebben we alleen patiënten met
diagnostisch ernstige hemofilie A (geen meetbare FVIII activiteit) geselecteerd op basis van
hun bloedingsneiging. De bloedingsneiging werd ditmaal niet alleen geclassificeerd op basis
van stolfactor verbruik, zoals in hoofdstuk 2, maar er werd een bloedingsfenotype score
gemaakt waarin tevens de leeftijd van de patiënt op het moment van zijn eerste bloeding,
de gewrichtsschade en de frequentie van gewrichtsbloedingen werden meegenomen. In
deze studie kon de observatie uit hoofdstuk 2 niet bevestigd worden, en moeten we dus
concluderen dat bloedplaatjes geen invloed lijken uit te oefenen op de bloedingsneiging
van hemofilie A patiënten.
Ernstige hemofilie A patiënten kunnen profylactisch met FVIII worden behandeld om
hun bloedingsneiging te verminderen. Om te beoordelen of FVIII infusies, naast correctie
van de verminderde stolfactor activiteit, nog andere karakteristieken van de hemostase
beïnvloeden, hebben we in hoofdstuk 3 vWF en bloedplaatjes activiteit en reactiviteit
gemeten in diagnostisch ernstige hemofilie A patiënten voorafgaande en 15 en 60 minuten
na toediening van een hoge dosis FVIII. In deze studie zagen we dat infusie van FVIII leidt
tot een afname in de vWF plasmaconcentratie, samen met een afname in ADAMTS-13 en
het aantal circulerende geactiveerde bloedplaatjes. We vermoeden dat na infusie, binding
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van FVIII aan circulerend vWF zorgt voor een interactie tussen vWF en geactiveerde
bloedplaatjes, resulterend in de klaring van vWF en ge(pre)activeerde bloedplaatjes. Dit
zou kunnen verklaren waarom in we in hoofdstuk 2, conform onderzoeksresultaten van
andere onderzoekers, bij hemofiliepatiënten hogere vWF plasmaconcentraties vinden in
vergelijking met gezonde mannen. Aanvullende studies zijn nodig om te onderzoeken hoe
dit proces precies in zijn werk gaat, en om te bepalen hoe groot het effect hiervan is op de
hemostase van hemofilie patiënten.
Om wat dieper te kijken naar de relatie tussen FVIII en bloedplaatjes, hebben we in
hoofdstuk 5 het vermogen van bloedplaatjes en hun voorlopercellen (megakaryocyten)
onderzocht om FVIII op te nemen. In vitro zagen we dat toevoeging van het lysaat van
bloedplaatjes van gezonde donoren aan FVIII deficiënt plasma, de stoltijd (APTT) van dit
plasma verkort. Dit suggereert de aanwezigheid van FVIII in bloedplaatjes lysaat. Wanneer,
voorafgaande aan lysis van de cellen, de bloedplaatjes geïncubeerd worden met FVIII,
neemt deze verkorting van de APTT verder toe, met name wanneer de bloedplaatjes
daaraan voorafgaand gepre-activeerd worden. Ook lysaten van megakaryoctyen geven een
verkorting van de APTT wanneer deze cellen met FVIII zijn geïncubeerd. Zowel bloedplaatjes
als megakaryocyten lijken dus in staat om FVIII op te nemen. Om te beoordelen of deze
observaties ook in vivo relevant zijn, hebben we bloedplaatjes van ernstige hemofilie A
patiënten voor en na infusie van FVIII bestudeerd. Hierbij bleken de bloedplaatjes al
voor FVIII infusie, FVIII te bevatten. De in dit onderzoek geïncludeerde patiënten werden
profylactisch behandeld met FVIII, en hebben daarom in de dagen voor deelname aan het
onderzoek ook FVIII infusies ontvangen. We denken dat de FVIII die we aantroffen in de
bloedplaatjes, opname van FVIII uit eerdere infusies reflecteert. Aanvullend onderzoek zal
nodig zijn om dit te bevestigen, en om te bepalen of deze opname in vivo op het niveau van
bloedplaatjes of op het niveau van megakaryocyten plaatsvindt.
Chronisch nierfalen
Patiënten met chronisch nierfalen vertonen zowel een verhoogde bloedingsneiging als een
verhoogde neiging tot het vormen van trombose. Omdat de rol van bloedplaatjes hierin
niet geheel duidelijk is, hebben we in hoofdstuk 6 bloedplaatjes reactiviteit bestudeerd
in patiënten met chronisch nierfalen. In dit hoofdstuk tonen we een deficiëntie in de
degranulatie van bloedplaatjes aan, welke onafhankelijk lijkt van de agonist welke gebruikt
wordt om de bloedplaatjes te activeren. Bloedplaatjes reactiviteit is dus verminderd
in patiënten met chronisch nierfalen, en dit kan een verklaring zijn voor de verhoogde
bloedingsneiging. Omdat we alleen naar degranulatie en niet naar andere kenmerken van
activatie hebben gekeken, moet er nog verder onderzoek plaatsvinden om te bepalen of
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het een specifiek probleem betreft in de degranulatie, of een algemeen probleem in de
activering van bloedplaatjes.
Chronische immuun trombocytopenie
Bij patiënten die een immuun trombocytopenie (ITP) ontwikkelen, leidt de vorming
van auto-antilichamen tot een afname van het aantal circulerende bloedplaatjes (een
trombocytopenie). Er zijn patiënten waarbij de trombocytopenie binnen een jaar
verdwijnt, maar bij een deel van de patiënten persisteert de trombocytopenie en spreken
we over chronische ITP. Gedurende deze chronische fase kan het bloedplaatjes aantal
van de patiënt variëren tussen sterk verlaagd (<10*109/L) en bijna normaal (normaal is
150*109/L tot 350*109/L). Echter, de bloedingsneiging van deze patiënten hangt niet altijd
samen met het aantal circulerende bloedplaatjes. Daarom hebben we in hoofdstuk 7
onderzocht of de bloedingsneiging van patiënten met chronische ITP samenhangt met
bloedplaatjes reactiviteit. Bij patiënten met een ernstige bloedingsneiging hebben we
een afgenomen bloedplaatjes reactiviteit gevonden in vergelijking met patiënten met een
milde bloedingsneiging. De degranulatie na stimulatie met de agonist ADP is verminderd en
er zijn hogere concentraties van de agonisten ADP en convulxin nodig om bloedplaatjes half
maximaal te activeren. In dit onderzoek werd bloedplaatjes functie ook bepaald met de
micro bloedplaatjesaggregatie test, waarbij ook een verminderde micro aggregatie werd
waargenomen na stimulatie met de activator PMA. Aanvullend onderzoek is nodig om te
bepalen of we met deze testen de bloedingsneiging van patiënten met chronische ITP in
een vroeg stadium kunnen voorspellen, en om te bepalen waardoor deze verminderde
reactiviteit veroorzaakt wordt. Wij vermoeden dat auto-antistoffen de reactiviteit van
bloedplaatjes mogelijk beïnvloeden; toekomstige studies zijn nodig om dit aan te tonen.
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Dankwoord
Allereerst wil ik alle patiënten die belangeloos hebben deelgenomen aan de verschillende
studies hartelijk bedanken. Zonder hun inzet waren deze onderzoeken niet mogelijk
geweest.
Prof.dr. Ph.G. De Groot, beste Flip, ik wil je bedanken voor de mogelijkheid die je mij hebt
geboden om dit onderzoek te doen. Met bewondering heb ik door de jaren heen geleerd van
jouw eindeloze kennis. Niet alleen details van de hedendaagse hemostase of labtechnieken,
maar ook de evolutie van bloedplaatjes weet jij tot in detail uit te leggen. Van jou kreeg ik
alle ruimte om zelf dingen uit te zoeken, en je kwam waar nodig met waardevolle suggesties.
Soms dacht ik even dat ik jouw aandacht tijdens een werkbespreking was verloren, maar dat
was altijd maar schijn, juist op die momenten kwam je met de meest scherpe opmerkingen.
Dr. M. Roest, best Mark, ik waardeer de volle overgave waarmee jij voor een project
kan gaan wanneer je ergens in gelooft. Jouw enthousiasme wist projecten, wanneer ze
bijvoorbeeld even stil waren komen te liggen, weer verder te brengen. Dank voor jouw
kritische blik en aanvullingen op de verschillende hoofdstukken in dit boekje. Maar zeker
ook voor alle gezelligheid op congressen, borrels, en tijdens het dagelijkse werk.
Dr. R.E.G. Schutgens, beste Roger, jouw bijdrage was soms groter dan jij je realiseerde.
Wanneer jij aanwezig was op de werkbespreking werden vaak de grote lijnen weer wat
helderder en was de besluitvaardigheid groter. Je hebt mij gemotiveerd om mijn horizon
te verbreden, bijvoorbeeld door mij te stimuleren beursaanvragen te schrijven. Dank voor
jouw vertrouwen in mij en jouw steun bij mijn sollicitaties voor zowel deze onderzoeksplek
als de kliniek.
Prof.dr. D.H. Biesma, beste Douwe. Jouw bijdrage stamt eigenlijk uit de tijd voor dit boekje,
en jouw rol als promotor is dan ook meer symbolisch. Als 3e jaars geneeskundestudent
kreeg ik van jou de kans om als student mee te werken aan een onderzoeksproject van
Mariëtte Agterof. En onder jullie supervisie werd mijn interesse voor de wetenschap
gewekt. Bedankt voor alle steun die je mij hebt gegeven, alle carrière adviezen en alle
deuren die je voor me open hebt gezet.
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Geachte leden van de leescommissie, prof.dr. G. Pasterkamp, prof.dr. R.H.W.M. Derksen,
prof.dr. K. Mertens, prof.dr. C.E. Hack en dr. W.L. van Heerde, ik wil u allen vriendelijk
bedanken voor het kritisch beoordelen van mijn proefschrift.
Uiteraard wil ik alle medeauteurs die dit boekje mogelijk hebben gemaakt bedanken.
Om te beginnen de vier talentvolle studenten die ik heb mogen begeleiden, en die stuk
voor stuk een waardevolle bijdrage hebben geleverd aan dit boekje. Daisy, jij bleek al snel
een harde werker, maar bracht ook veel gezelligheid mee naar het lab. Veel succes met het
afronden van jouw eigen promotieonderzoek. Loes, tijdens het aanleren van de test viel al
op hoe secuur jij te werk gaat. Een verkeerd aangelegde waterleiding leek jou proeven te
verstoren, maar al snel spoorde je de bron op. Ook na jouw stagetijd kruisten onze paden
nog geregeld, als señorita Loes met het Carnaval in Den Bosch, en inmiddels als dokter Loes
in het Meander. Hartstikke leuk om met jou nu in de kliniek samen te werken. Lieke, jij hebt
zeer zelfstandig gewerkt aan het vingerprikproject en daarmee de basis neergelegd voor dit
hoofdstuk. Ik hoop dat we dit in de toekomst nog zullen afronden tot een publicatie. Laila,
jij stond ineens voor mijn neus: een nieuwe student en een nieuw project. Soms kwam
je wat onzeker over, maar dat bleek vooral een uiting van jouw wil om alles tot in detail
te snappen. Bedankt voor jouw bijdrage in de ITP studie. Je hebt een prachtige databse
aangelegd, succes ook met het afronden van de andere artikelen die hieruit voortkomen.
Attie, ik geloof niet dat iemand ooit zo traag patiënten heeft geïncludeerd als wij
samen ;) Met jou samenwerken betekende dat altijd alles tot in de puntjes geregeld was,
maar betekende ook veel gezelligheid samen op het lab. Onze “stoom afblaas” etentjes
waren onmisbaar! Ik ben blij dat jij als paranimf bij me staat straks.
Kathelijn, ik heb mooie herinneringen aan onze reis naar de Toronto, Utrecht,
Vellore meeting in India. Dank voor het ontwikkelen van de bloedingsfenotype score
voor hoofdstuk 4. Rosa, vanuit de kliniek heb jij onze studie bij patiënten met nierfalen
gecoördineerd. Nu ik zelf in de kliniek werk realiseer ik me hoe pittig deze combinatie
is. Heel veel succes bij het promotieonderzoek waaraan je inmiddels bent begonnen.
Prof. dr Gaillard, Carlo, dank voor het kritisch meedenken en al jouw aanvullingen. Leuk, al
was het kort, om je ook in de kliniek mee te maken, waar je een ware wandelende medische
encyclopedie bleek. Een FACS hoort niet stil te staan! Leonie, mooi dat we de FACS in
het Meander nieuw leven in hebben kunnen blazen. Ook wil ik hier Rob Kraaijenhagen
bedanken voor zijn rol in het opstarten van dit project. Rob Fijnheer, dank voor jouw
begeleiding en actieve rol in het project bij patiënten met nierfalen. Het is erg leuk om nu
in het Meander te mogen werken met jou als opleider. Annemieke, ons project had een
lang traject, waarbij het een uitdaging was om de verschillende ideeën samen te brengen
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in één stuk. De samenwerking hierin met jou was erg prettig. Ik wens je veel succes met de
afronding van jouw promotie. Katja, Leendert, Ellen en Masja, dank voor jullie waardevolle
aanvullingen. Gestur, we hebben veel gediscussieerd, ik denk dat dit het stuk uiteindelijk
wel tot een hoger niveau heeft gebracht. Marrie, gedurende alle discussies wist jij de hoofden nevenargumenten te onderscheiden, en daarmee zorgde je er iedere keer weer voor dat
wij een stap dichterbij de afronding van het artikel kwamen. Dank voor jouw begeleidende
rol in dit project.
Wanneer je onderzoek doet zijn er ook nog vele mensen nodig, die misschien niet direct
zichtbaar betrokken zijn geweest, maar zeker wel onmisbaar waren!
Op de LKCH heb ik 4 jaar lang met veel plezier gewerkt, en dit is voor een belangrijk deel te
danken aan al mijn collega’s.
Vivian, met jou heb ik 4 jaar lang samen op de LKCH gewerkt. De eerste dag kon ik jouw
Engels met Chinese tongval maar nauwelijks verstaan, maar 4 jaar later voeren we gewoon
gesprekken in het Nederlands. Geweldig dat je het doorzettingsvermogen hebt om aan
een 2e AIO-schap te beginnen, veel succes met de afronding hiervan. Evelyn, jij hebt me op
mijn 1e dag meteen op weg geholpen, heel fijn! Anja, AIO-kamer buurvrouw, dank voor alle
gezelligheid. Erik jij hebt me in de eerste week het FACSen aangeleerd, de techniek die als
een van de rode draden door dit boekje loopt. Dank je wel hiervoor! Leuk om je nu in het
Meander weer als collega tegen te komen. Gwen, dank voor alle gezelligheid, je bleef altijd
verrassen door onverwacht uit de hoek te komen. Dianne, super dat je jouw droom om
als postdoc in het buitenland te werken in Canada en Australië hebt waargemaakt, maar
ook erg gezellig dat je nu weer naar Nederland komt! Cees, toen we lab I versierde met
mooie mannen was je jaloers dat je er niet tussen hing, onze fout: daarna heeft ook jouw
foto tijden lang ons lab opgefleurd! Rolf en Coen, dank voor al jullie adviezen en het actief
meedenken, jullie interesse en creativiteit zijn waardevolle eigenschappen. Çetin dank
voor alle leuke gesprekken op het lab, cursussen en congressen. Thijs ik mis onze levendige
discussies. Veel succes met je opleiding tot klinisch chemicus, wie weet kan ik vanuit de
kliniek nog eens van jouw expertise gebruik maken. Claudia, jouw organisatietalent heeft
geleid tot talrijke uitjes en taartencompetities, dank voor alle mooie herinneringen, ook
onze dagen in New York zal ik niet snel vergeten. Heel veel succes in België. Eelo, dank
voor het aanleren van het kweken van mega’s! Heb je de wekker/beltoon streepjeslijsten in
Engeland ook al in weten te voeren? Valentina I admire how you build up a complete new
life in a new country in such a short period, and then just move to Germany to repeat such
a challange! I wish you the best for the future. Ciao, or should I say tschüss? Agon jij bent de
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volgende, heel veel succes! Ik heb me de afgelopen jaren verbaasd over jouw brede kennis
over de meest uiteenlopende onderwerpen, en natuurlijk over de snelheid waarmee jij een
cijferslot ontcijfert ;) Marije, ondanks dat je de kliniek erg mist weet je een aantal prachtige
projecten neer te zetten in het lab. Succes met de afronding, geniet weer van de kliniek
straks. Steven, ik ken niet veel AIO’s die op dag -1 alvast van start gaan, dit karakteriseert
jouw gemotiveerde werkhouding. Dank voor alle leuke gesprekken en tips voor het blotten.
Bert, jij geniet van het leven en brengt die sfeer iedere dag mee naar het werk. Peter Paul,
dank voor alle leuke gesprekken, en voor de jives op feestjes. Meta, hoewel je m.n. in
Nijmegen zat, bracht je iedere keer dat je op de LKCH was veel gezelligheid mee! Judith,
mijn bewondering voor jouw vechtlust en doorzettingsvermogen zijn groot, ik wens je al
het beste toe in de toekomst. Marco, jij hebt de uitdaging om in 2 steden te werken, succes
met het afronden van jouw onderzoek. Marti en Eva, dank voor de gezelligheid gedurende
jullie LKCH tijd, ik wens jullie veel succes in jullie toekomst.
Simone, je bent als mijn vijfde talentvolle student begonnen (nadat ik zag hoe secuur
je bij het project van Agon te werk ging), en daarna ben je gebleven als analist. Jouw
leergierigheid is een eigenschap om te koesteren. Je hebt heel veel data bij de hemofilie B
patiënten verzameld; die zijn misschien niet in dit boekje beland, maar hopelijk kunnen we
dit in de toekomst nog samenbrengen tot een artikel. Ik wens je veel succes in de toekomst.
Martine, dank voor de gezellige reis naar de GTH. Quirijn, dank voor de leuke gesprekken.
Suzanne ik ben onder de indruk van jouw multitasking talent met een groot en gezin en
succesvolle onderzoekscarrière. Bas, Maarten en Eszter, toen ik begon hoorde ik al veel
verhalen over jullie eerdere LKCH tijd, leuk om jullie vervolgens, toen jullie één voor één
terug kwamen, ook als collega’s mee te mogen maken.
Annet als ik aan jouw denk dan denk ik altijd aan Brabantse gezelligheid, het was leuk
om op lab I met jou samen te werken. Silvie bedankt voor alle hulp en uitleg, bij jou kon ik
altijd terecht met vragen. Sandra, in mijn eerste jaren stond jij aan het hoofd van de MDD,
een systeem waar ik veel profijt van heb gehad. Arnold jij stond echt altijd klaar om vragen
te beantwoorden, technieken uit te leggen of mee te helpen, ook wanneer ik de 10e was die
jou kwam storen met vragen. Ook dank voor al jouw gezelligheid op borrels en feestjes, en
voor al het lekkers van Slagerij Koekman niet te vergeten. Jerney, toen ik begon zat jij nog
bij de diagnostiek, maar al snel kwam je de research versterken. Ook jij bedankt voor alle
leuke dingen die je voor de afdeling hebt georganiseerd, van retraite tot taartencompetitie.
Arjan, dank voor al jouw technische ondersteuning en uitleg bij de ELISA’s en FACS Canto.
Grietje, Elske, Kimberley en Brigitte, bedankt voor alle leuke gesprekken bij de koffiepauzes
en lunch. Ya Ping, hoe kreeg je toch zoveel spullen op één labbench gepast? Leuk om te zien
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dat je met jouw enthousiasme zelfs na je pensioen eigenlijk geen afscheid kunt nemen van
de LKCH.
Prof.dr. Akkerman, Jan Willem, op het lab ken ik jou als iemand die systemen tot in detail
wil uitzoeken, maar daarnaast waardeer ik je ook zeer om de gezelligheid op borrels en
congressen. Harry, dank voor je ideeën tijdens de vrijdagochtend besprekingen. Richard,
het was erg leuk om samen een retreat te organiseren, en dankzij jou op een prachtige
locatie. Ray, Pieter, Susan en Sander, jullie kwamen in mijn laatste jaar de LKCH versterken.
Dank voor jullie enthousiasme en gezelligheid.
Niet één afdeling kan zonder goede ondersteuning, waarvan ik een aantal mensen
specifiek wil bedanken. Joukje je stond altijd klaar voor de nodige ondersteuning, en wist
vaak het raadsel van de afwezige (co-)promotor op te lossen. Rosmina, hoeveel werk jij ons
uit handen neemt merken we altijd zodra je op vakantie bent, en wij om beurten 2 dagen
‘Rosmina-dienst’ hebben. Michel, jij wist mijn computerproblemen altijd op te lossen, dank
je wel! Arno en Guido, dank voor het gevuld houden van ons magazijn.
Ook wil ik de hemato-oncologie groep bedanken. Robbert, Micheal, Inger, Sanne, Rimke,
Jonas, Julie, Teun, Cor, Johan, Maureen en Berris, dank voor alle leuke gesprekken tijdens
koffiepauzes, borrels en retraites. Maarten en Tineke, dank voor alle uitleg van de FACS en
technische ondersteuning, maar zeker ook voor alle leuke gesprekken tijdens het FACSen
op de Calibur. Tuna ik heb heel wat met je afgelachen!
Tevens dank aan iedereen op het diagnostisch lab voor de ondersteuning waar nodig. In
het specifiek Albert en Eline voor het actief meedenken met name bij logistieke uitdagingen.
Ria, dank voor alle gezelligheid op de jaarlijkse pool prikdag in het AMC waar je me mee
naar toe nam. Thea, dank voor jouw hulp met de metingen van procoagulante fosfolipiden.
Maarten, dank voor alle leuke gesprekken.
Ik had het geluk niet bij één, maar bij twee afdelingen te horen. Wanneer ik de Van
Creveldkliniek binnenloop voelt het net alsof ik niet meer in een groot academisch
ziekenhuis sta, maar in een kleine hechte kliniek.
Evelien en Goris, dank voor jullie inzet en alle hulp bij de patiënteninclusies. Karin, dank
voor de leuke gesprekken en de leerzame hematologie COIG’s. Nathalie, Ingrid, Dietje en
Janneke, dank voor alle gezelligheid in Utrecht en op congressen. Monique, Laurens en Lize,
dank voor alle leuke gesprekken en jullie meedenken op de VCK researchbesprekingen.
Marlies, Annelies, Mojtaba, dank voor jullie enthousiasme, veel succes met het afronden
van jullie studies. Federica I had fun showing you Amersfoort. Els, Monique en Tamara dank
voor jullie ondersteuning, jullie konden me altijd wijzen naar de benodigde data. Nanda,
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Hanny, Simone en Silvia dank voor jullie inzet bij alle patiënt inclusies, en alle bloedafnames
bij de hemofilie patiënten. Piet, Ruud, Arda en Nanda dank voor de gezellige gesprekken.
In 2012 ben ik gestart aan mijn opleiding tot internist in het Meander Medisch Centrum.
Ik wil al mijn collega’s hier, assistenten, internisten, secretariaat en verpleging, bedanken
voor hun interesse in mijn onderzoek, de goede werksfeer en de flexibiliteit in het rooster
wanneer dit voor het afronden van mijn onderzoek nodig was.
Buiten de werkvloer heb ik ook veel mensen om me heen die onmisbaar zijn; in de jaren van
mijn promotieonderzoek, maar zeker ook de jaren ervoor en de jaren erna!
Inge, je bent een engel, dank voor de talloze liften naar het lab en mijn huis, maar dank ook
vooral voor alle afleiding die je hebt gebracht. Zonder jou had ik heel veel mooie feestjes
gemist die mijn werk en privé weer wat beter in balans brengen! Saskia, het is heel fijn om
een vriendin zoals jij te hebben die me altijd goed begrijpt, voor me klaar staat wanneer
ik het moeilijk heb en meeleeft met alles wat ik doe. Idwer, Rian en Marijke, Sandra, Bert
en Judith, Erik, Hidde, Kevin, Carsten, Sterre, Bas en Sjaan, Marlies en Rick, Susan, Maikel,
Marinus, Ernst, Lex, ik prijs me heel gelukkig met jullie! Het is lastig uit te drukken hoeveel
het betekent om zo’n vriendengroep te hebben om lief en leed mee te delen, ik had niet
zonder gekund! Daarnaast zijn er nog veel mensen van de Reddingsbrigade, bestuur, kader
en leden, die ik wil bedanken voor alle gezelligheid in en buiten het bad. Ik hoop dat we
nog jaren zo als vereniging door mogen gaan, met de unieke sfeer die de ARB voor mij zo
speciaal maakt!
Tim en Désirée, jullie staan altijd voor me klaar, het is geweldig om jullie tot mijn
vriendengroep te mogen rekenen, en het zorgt vaak ook nog voor een stortvloed aan
lekker eten. Robert, Robert Korndewal, waar ben je? Ik wil dansen!! En waar staat je fiets
eigenlijk?! Esther, dank voor alle gezelligheid, en heel veel succes met de laatste loodjes van
jouw opleiding. Brand dank voor jouw vriendschap.
Brechje dank voor jouw vriendschap en dat ik altijd bij je terecht kan. Naast alle gezellige
momenten heb je de afgelopen jaren ook heel wat van mijn (onderzoeks-) frustraties
opgevangen. Om dingen aan jou uit te leggen zijn vaak maar weinig woorden nodig. Een
promotieonderzoek afronden betekend dat er een periode met meer vrije tijd aan komt.
Krista, tijd om een concert in te plannen!
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Als laatste, maar zeker niet als minste wil ik mijn hele familie bedanken. Oma, Leonie,
Louise, John en Ingrid, Jaimy en Jeroen, Myra, Tikwah, Paulus en Willeke, Mees: bedankt
voor jullie voortdurende interesse en steun. Opa, hoe graag we beide wilden dat jij de dag
van mijn verdediging nog mee zou maken heeft helaas niet mogen baten. Desalniettemin
weet ik dat er geen opa zo trots is op zijn kleindochter als die van mij.
In het bijzonder wil ik mijn ouders en zus bedanken. Papa en mama, jullie hebben me
altijd het gevoel gegeven dat er niets is wat buiten mijn bereik ligt. Dank voor jullie geloof
in mij. Lieve Naomi, al van kleins af aan heb ik de onvoorwaardelijke liefde en steun van
mijn grote zus. Ik ben blij dat jij op de dag van mijn verdediging als paranimf bij mij staat.
Esther, 18 oktober 2013
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Curriculum Vitae
Esther van Bladel was born on January 16, 1985, in ’s-Hertogenbosch, The Netherlands.
After graduating from secondary school at ’t Hooghe Landt college in Amersfoort in 2002,
she studied Medicine at the University of Utrecht. As part of her medical training, she
performed two scientific internships on the subject of venous thromboembolism. Both were
conducted at the Sint Antonius Ziekenhuis in Nieuwegein under the supervision of prof dr.
D.H. Biesma and dr. M.J. Agterof, initiating her interest in science. Esther received her
medical degree in 2008. Following graduation, she began her work as a PhD student at the
Department of Clinical Chemistry and Hematology at the University Medical Center Utrecht
under the supervision of prof.dr. Ph.G. de Groot, dr. M. Roest and dr. R.E.G. Schutgens. This
work was performed in close collaboration with the Van Creveldkliniek at the University
Medical Center Utrecht. The results of her research are described in this thesis. In 2012
she started her specialist training in Internal Medicine at Meander Medisch Centrum
Amersfoort under the supervision of dr. R. Fijnheer en dr. R. Bosma.
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List of publications
van Bladel ER, Tuinenburg A, Roest M, de Groot PG, and Schutgens REG. Factor VIII
concentrate infusion in patients with hemophilia results in decreased vWF and
ADAMTS-13 activity. Accepted for publication in Haemophilia.
van Bladel ER, de Jager RL, Walter D, Cornelissen L, Gaillard CA, Boven LA, Roest M, Fijnheer
R. Platelets of patients with chronic kidney disease demonstrate deficient platelet
reactivity in vitro. BMC Nephrology 2012;13:127.
van Bladel ER, Roest M, de Groot PG, and Schutgens REG. Up-regulation of platelet
activation in hemophilia A. Haematologica 2011;96(6):888-895.
van Bladel ER, Agterof MJ, Frijling BD, van der Griend R, Prins MH, Schutgens REG, Biesma
DH. Out of hospital anticoagulant therapy in patients with acute pulmonary embolism
is frequently practised but not perfect. Thrombosis Research 2010;126(6):481-5.
Agterof MJ, van Bladel ER, Schutgens REG, Snijder RJ, Tromp EA, Prins MH, Biesma DH. Risk
stratification of patients with pulmonary embolism based on pulse rate and D-dimer
concentration. Thrombosis and Haemostasis 2009 October;102(4):683-7.
Awards
Best PhD Paper 2011, Divisie Laboratoria en Apotheek, UMC Utrecht. van Bladel ER,
Roest M, de Groot PG, and Schutgens REG. Up-regulation of platelet activation in
hemophilia A.
2011 Haemophilia Europe ASPIRE research award, Pfizer. van Bladel ER, Roest M, de Groot
PG, and Schutgens REG. Variability in platelet function in relation to hemophilia B
phenotype.
2011 Young Investigator Award, International Society on Thrombosis and Haemostasis.
van Bladel ER, Roest M, de Groot PG, and Schutgens REG. Up-regulation of platelet
activation in hemophilia A.
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