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HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
Differential catalytic properties and vascular topography of murine nucleoside
triphosphate diphosphohydrolase 1 (NTPDase1) and NTPDase2 have
implications for thromboregulation
Jean Sévigny, Christian Sundberg, Norbert Braun, Olaf Guckelberger, Eva Csizmadia, Imrana Qawi, Masato Imai,
Herbert Zimmermann, and Simon C. Robson
Nucleoside triphosphate diphosphohydrolases (NTPDases) are a recently described
family of ectonucleotidases that differentially hydrolyze the ␥ and ␤ phosphate residues of extracellular nucleotides. Expression of this enzymatic activity has the
potential to influence nucleotide P2 receptor signaling within the vasculature. We and
others have documented that NTPDase1
(CD39, 78 kd) hydrolyzes both triphosphonucleosides and diphosphonucleosides and
thereby terminates platelet aggregation responses to adenosine diphosphate (ADP).
In contrast, we now show that NTPDase2
(CD39L1, 75 kd), a preferential nucleoside
triphosphatase, activates platelet aggregation by converting adenosine triphosphate
(ATP) to ADP, the specific agonist of
P2Y1 and P2Y12 receptors. We developed
specific antibodies to murine NTPDase1 and
NTPDase2 and observed that both enzymes
are present in the cardiac vasculature;
NTPDase1 is expressed by endothelium,
endocardium, and to a lesser extent by
vascular smooth muscle, while NTPDase2
is associated with the adventitia of muscularized vessels, microvascular pericytes, and
other cell populations in the subendocardial
space. Moreover, NTPDase2 represents a
novel marker for microvascular pericytes.
Differential expression of NTPDases in the
vasculature suggests spatial regulation of
nucleotide-mediated signaling. In this context, NTPDase1 should abrogate platelet ag-
gregation and recruitment in intact vessels
by the conversion of ADP to adenosine
monophosphate, while NTPDase2 expression would promote platelet microthrombus
formation at sites of extravasation following
vessel injury. Our data suggest that specific
NTPDases, in tandem with ecto-5ⴕ-nucleotidase, not only terminate P2 receptor activation and trigger adenosine receptors
but may also allow preferential activation
of specific subsets of P2 receptors sensitive to ADP (eg, P2Y1, P2Y3, P2Y12) and
uridine diphosphate (P2Y6). (Blood. 2002;
99:2801-2809)
© 2002 by The American Society of Hematology
Introduction
Nucleoside triphosphate diphosphohydrolases (NTPDases) are a
family of ectonucleotidases, previously classified as E-type ATPases,
ATPDases, ecto-ATPases, or ecto-apyrases.1-3 These enzymes
differentially hydrolyze the terminal ␥ and ␤ phosphate residues of
nucleotides, resulting in the rapid formation of the respective
diphosphonucleosides and/or monophosphonucleosides. To date, 6
members of this NTPDase family have been identified.3-13 NTPDase1, NTPDase2, and NTPDase3 are transmembrane proteins
associated with the plasma membrane with an active site facing the
extracellular space.4,5,7,9,14 These NTPDase members have been
demonstrated to differ in their substrate specificity. For example,
NTPDase1 (CD39 [human] or cd39 [murine]) hydrolyzes both
nucleoside triphosphates and diphosphates—eg, adenosine triphosphate (ATP) and adenosine diphosphate (ADP),4,5—whereas
NTPDase2 (CD39L1) is a preferential nucleoside triphosphatase
or ATPase.7,14-16
Extracellular nucleotides in various forms, and at different
concentrations, activate multiple P2 receptors: ionotropic P2X and
metabotropic P2Y receptors.17-20 While NTPDases would be anticipated to generally terminate P2 receptor agonist signaling, we have
also observed that NTPDase1 may prevent receptor desensitization
by the catalysis of extracellular nucleotides.21 NTPDase1 may then
also facilitate subsequent responses to “pulses” of nucleotides. In
addition, NTPDases in tandem with ecto-5⬘-nucleotidase may
facilitate the salvage of nucleotides by the ultimate generation of
dephosphorylated forms that are taken up by cells via specific
transporters.1,22
NTPDase1 is the major ectonucleotidase at the luminal surface
of blood vessels.5,21,23,24 By converting the ADP released from
activated platelets, NTPDase1 modulates platelet aggregation in
vitro.5,25-28 The hypothesis that NTPDase1 is a key thromboregulatory factor has been further supported by in vivo experiments with
NTPDase1-null (or cd39⫺/⫺) mice. This model has demonstrated
important functions of the ectoenzyme in both regulating platelet
aggregation and controlling the expression of procoagulants by the
endothelium.21
From the Departments of Medicine and Pathology, Beth Israel Deaconess
Medical Center, Harvard Medical School, Boston, MA; Centre de recherche en
Rhumatologie et Immunologie CHUQ, Université Laval, Sainte-Foy, Québec,
Canada; and Biozentrum der JW Goethe-Universität, AK Neurochemie,
Zoologisches Institut, Frankfurt am Main, Germany.
Foundation and the King Gustaf the Fifth’s 80 year fund (to C.S.). O.G. was a
recipient of a fellowship from the Deutsche Forschungsgemeinschaft (GU
490/1-2).
Submitted July 3, 2001; accepted November 30, 2001.
Supported by NIH grants RO1HL57307 and RO1HL63972-01 (to S.C.R.), the
American Liver Foundation and the Canadian Institutes of Health Research (to
J.S.), the Deutsche Forschungsgemeinschaft (SFB 269, A4) and the Fonds der
Chemischen Industrie (to H.Z.), and a grant from the Swedish Cancer
BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
Reprints: Jean Sévigny, Centre de Recherche en Rhumatologie et
Immunologie, 2705 Blvd Laurier, Local T1-49, Sainte-Foy, Québec, G1V 4G2,
Canada; e-mail: [email protected].
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
© 2002 by The American Society of Hematology
2801
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2802
SÉVIGNY et al
Whether NTPDase2 is expressed in the vasculature has not been
established to date. Furthermore, nothing is known about the
functional integration of either NTPDase1 or NTPDase2 with each
other or other ectonucleotidases/NTPDases. Biochemical analyses
of murine and porcine cardiac tissues have demonstrated ATPase/
ADPase ratios of 10, suggesting expression of enzymes with
preferential ATPase activity (this paper and Lemmens et al29).
Given the levels of NTPDase2 messenger RNA (mRNA)
expression in murine and human hearts, this enzyme would be a
likely candidate responsible for such an activity.6-8 NTPDase3 is
another potential ectoenzyme that may modulate cardiac extracellular nucleotide levels. Although little is known about the
distribution of human NTPDase3, cardiac mRNA expression
appears low when compared with the signals observed for
NTPDase1 and NTPDase2.8
In this paper, we show that catalytic differences between
NTPDase1 and NTPDase2 are associated with the differential
ability to regulate platelet function. This observation is consistent
with the distinct localization of these ectoenzymes in the vasculature of the murine heart that affords yet another level of complexity
by which these biologic activities may be regulated.
Materials and methods
Transient transfection and protein preparation
COS-7 cells were transfected using lipofectamine, as previously described.5
Forty-four hours after transfection, the cells were washed twice with
Tris-saline, harvested by scraping, and washed twice by centrifugation at
300g for 5 minutes at 4°C. Prior to sonication, pellets were resuspended in
45 mM NaCl, 0.1 mM phenylmethylsulfonyl fluoride, 10 ␮g/mL aprotinin,
and 45 mM Tris, pH 7.6. Nucleus and cellular debris were discarded by
another centrifugation as outlined above. Supernatants containing about
10 mg proteins per milliliter were kept frozen at ⫺80°C. Homogenates
and particulate fractions of murine tissues were prepared as previously
described.24
NTPDase activity measurement
Enzyme activity in the protein fractions was determined as previously
described.24 Briefly, enzyme activity was tested in 1 mL of 8 mM CaCl2, 50
mM Tris, and 50 mM imidazole, pH 7.4. After the addition of protein
sample to the buffer, the mixture was preincubated at 37°C for 3 minutes
and reaction was started by the addition of 0.3 mM substrate (ATP or ADP
as indicated). This was terminated at 5 to 15 minutes with 0.25 mL of the
malachite green reagent, and inorganic phosphate released from exogenous
nucleotide measured.30 To determine specific activities, the protein content
of the enzyme preparations was measured using the technique of Bradford.31
Analysis of platelet activation in vitro
Platelet-rich plasma (PRP) was prepared from human donors according to
published methods.5 Platelet aggregation was measured in a Lumiaggregometor apparatus (Chrono-log, Havertown, PA). Samples of 0.33 mL
PRP were incubated at 37°C, and percent light transmission was measured
and compared with platelet-poor plasma. Final concentrations of 1 to 20
␮M ADP (Chrono-log) were used for platelet activation. In select experiments, aliquots to achieve final concentrations of 1 to 40 ␮M ATP (Sigma,
St Louis, MO) were also added to PRP. Where indicated, varying amounts
of recombinant protein samples (1-230 ␮g total protein) derived from
NTPDase1- or NTPDase2-transfected COS cell lysates, diluted in 0.9%
saline, were added to PRP.
Antibodies, plasmids, and other reagents
Antimurine NTPDase1 and NTPDase2 polyclonal antibodies (pAbs) were
raised in rabbits by direct intramuscular and subcutaneous injection of
BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
complementary DNA (cDNA) encoding the whole gene ligated into
pcDNA3.21,32 The plasmids expressing mouse NTPDase121 and the open
reading frame of rat NTPDase2 have been described previously.7 Serum
titers were determined by standard Western blot analysis under nonreducing
conditions in the screening protein lysates from COS-7 cells expressing
recombinant murine NTPDase1 or NTPDase2. The antibody specifically
detected either NTPDase1 or NTPDase2 of both mouse and rat tissues in
immunohistochemistry.
For selected double- and triple-labeling experiments, immunoglobulin
G (IgG) fractions from NTPDase2 antisera, purified using a protein
A–Sepharose column, were biotinylated using the EZ-Link Sulfo-NHS-LCBiotinylation kit, according to the manufacturer’s instructions (Pierce,
Rockford, IL). The monoclonal antibody (mAb) anti–PECAM-1 (CD31),33
recognizing murine endothelial cells, and phycocyanin avidin were purchased from Pharmingen (San Diego, CA). The mAb anti-NG2 34,35
corresponding to the human high-molecular-weight melanoma-associated
antigen, an epitope expressed on pericytes and smooth muscle cells,36 and
the mAb antilaminin ␤2 chain37 used to detect basal lamina were purchased
from Chemicon (Temecula, CA). The mAb antismooth muscle ␣-actin
(clone 1A4)38 used as a marker for smooth muscle cells (and pericytes in
other tissue than the heart) was purchased from Sigma. The mAb F4/80
recognizing blood monocytes and macrophages was purchased from
Serotec (Raleigh, NC). Biotinylated rabbit antimouse IgG F(ab⬘)2 fragments and biotinylated swine antirabbit IgG F(ab⬘)2 were purchased from
DAKO (Glostrup, Denmark). Normal serum (rabbit, mouse, goat, and
swine) and IgG (rat, rabbit, and mouse) were purchased from Sigma. The
biotinylated rabbit antirat IgG, Texas Red Avidin D, goat antirabbit IgG
Texas Red conjugate, and rabbit antirat IgG fluorescein conjugate were
purchased from Vector Laboratories (Burlingame, CA).
Immunoblotting procedures
Proteins were fractionated by sodium dodecyl sulfate (SDS)–polyacrylamide gel electrophoresis according to Laemmli.39 Protein samples were
boiled in sample buffer (2% [wt/vol] SDS, 10% [vol/vol] glycerin, 0.001%
bromophenol blue in 65 mM Tris, pH 6.8) under nonreducing conditions.
The proteins were separated on a 10% acrylamide SDS-gel and transferred
to Immobilon-P membrane (Millipore, Bedford, MA) by semidry electroblotting (Bio-Rad, Hercules, CA).40 After incubation with the rabbit
anti-NTPDase pAbs, the bands were visualized using horseradish peroxidase–conjugated goat antirabbit IgG (Pierce), at a dilution of 1:4000, and
the Renaissance Chemiluminescence Reagent Plus, according to the
manufacturer’s instructions (NEN Life Science Products, Boston, MA).
Immunohistochemistry
Cardiac and other tissues from C57BL/6 and 129SVEV⫻C57BL/6 mice
were harvested, embedded in Triangle Biomedical Sciences (TBS) tissue
freezing medium (American Master*tech Scientific, Lodi, CA) and snapfrozen in isopentane cooled on liquid nitrogen, and stored at ⫺80°C.
Six-micrometer serial cryostat sections were fixed in ice-cold acetone for 10
minutes and rinsed in phosphate-buffered saline (PBS). IgG binding sites
were blocked with appropriate control serum diluted 1:5 in solution 9 (PBS,
pH 7.4, supplemented with 0.1% bovine serum albumin, 150 mM tranexamic acid, 20 ␮g/mL aprotinin [3-7 trypsin inhibitory units per milligram],
1.8 mM ethylenediaminetetraacetic acid, and 2 mM iodoacetic acid) further
supplemented with 2% 3-omega fatty acid (Sigma) for 1 hour at room
temperature. Sections were then incubated with primary antibody for 1 hour
at room temperature (biotinylated NTPDase2-purified IgGs were incubated
for 2 hours at 37°C), rinsed in PBS, and incubated with 3% H2O2 in
methanol for 5 minutes to deplete endogenous peroxidase. After incubation
with the appropriate biotinylated IgG or F(ab⬘)2 fragment of IgG for 30
minutes, staining was performed with the Vectastain ABC elite kit (Vector
Laboratories) with diaminobenzidine as the peroxidase substrate. Sections
were counterstained lightly with Mayer’s hematoxylin, dehydrated, cleared
in xylene, and mounted in Permount. Detection of mouse antigens with
mouse mAbs was performed as previously described.41 All antibodies were
diluted in solution 9 unless otherwise indicated. Optimal antibody concentrations were determined by serial dilution.
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BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
VASCULAR NTPDases AND THROMBOREGULATION
2803
Triple staining and confocal imaging
Cryostat sections 6 ␮m thick were fixed in 100% acetone at 4°C for 20
minutes and rehydrated in PBS. Sections were blocked with 20% normal
goat serum and 20% normal mouse serum containing 2% 3-omega fatty
acid for 1 hour at room temperature. Sections were then incubated for 1
hour at room temperature with rabbit anti-NTPDase1 IgG, rinsed, incubated
with goat antirabbit IgG fluorescein isothiocyanate conjugate or biotinylated goat antirabbit IgG for 30 minutes, and then rinsed. Sections were
blocked with 5% normal rabbit serum for 30 minutes, rinsed, incubated
with biotinylated NTPDase2 IgGs or avidin-coupled phycocyanin for 2
hours at 37°C and 30 minutes at room temperature, respectively, and rinsed.
The sections were then incubated with Texas Red Avidin D or antismooth
muscle actin fluorescein isothiocyanate conjugate for 1 hour at room
temperature, rinsed, and blocked for residual binding sites to avidin using
the avidin-biotin blocking kit (Vector Laboratories). These sections were
then incubated with biotinylated anti-CD31 or biotinylated NTPDase2 IgG
for 2 hours at room temperature and 37°C, respectively. After rinsing, the
sections were incubated with avidin-coupled phycocyanin or Texas Red
Avidin D for 30 minutes at room temperature and rinsed 5 times in PBS and
once with distilled water. Sections were then mounted in fluoromount G and
analyzed using an MRC-1024 confocal microscope equipped with an
argon/krypton laser (Bio-Rad). Series of sequential optical sections were
digitalized, filtered with edge definition and median filters, and viewed as
compiled images.41
Statistics
Where appropriate, data are expressed as the mean ⫾ SD. Groups were
compared using the Student t test. Differences between experimental
groups and controls were considered significant for P ⬍ .05.
Results
NTPDase activity in mouse hearts and transfected cells
We determined levels of ATPase and ADPase activity in
homogenates derived from a variety of wild-type mouse tissues
(data not shown). Whereas relative ATPase and ADPase activities were similar in most tissues studied, the ATP-to-ADP
hydrolysis ratios in the mouse hearts were strikingly higher, to
the order of 10 (Figure 1A). These data are comparable to prior
observations in pig tissues.29
To identify the relative contribution of NTPDase1 to the total
NTPDase activities in heart tissues, we then contrasted the activity
of homogenates derived from cd39⫹/⫹ and cd39⫺/⫺ tissues. Both
ATPase and ADPase activities in cardiac tissue preparations from
NTPDase1-null mice were not significantly different in wild-type
tissues. Lung, a highly vascularized organ known to express high
levels of NTPDase1, was used for comparison (Figure 1A). From
the known distribution of NTPDases by Northern blots,7,8 the
activity detected in the heart could likely be explained by expression of NTPDase2, a preferential ATPase.7
COS-7 cells were transfected by an expression vector
(pcDNA3) containing the cDNA of murine NTPDase1 or
NTPDase2. Biochemical activity of protein extracts derived
from transfected cells exhibited significant increases in the
hydrolysis of both substrates, ATP and ADP (Figure 1B). While
the ratios of ATPase and ADPase activities in NTPDase1transfected cells were similar, the ratio of ATP/ADP hydrolysis
in protein extracts from NTPDase2-transfected cells was of the
order of 40, in keeping with prior data derived from intact
Chinese hamster ovary cells transfected with rat NTPDase2.7
Figure 1. Biochemical activity of murine tissues and recombinant NTPDase1
and NTPDase2. NTPDase activities of protein preparations were determined by
measurement of phosphate release from the respective substrates. Data are
expressed as the mean ⫾ SD for specific ATPase (䊐) or ADPase (■) activities. (A)
NTPDase activities of cd39⫹/⫹ (n ⫽ 5) and cd39⫺/⫺ (n ⫽ 5) mouse lung and
heart preparations were measured. (B) NTPDase activity of protein extracts from
NTPDase1- or NTPDase2-transfected cells were compared with control cells—
untransfected (COS) or those transfected with pcDNA3 empty vector (n ⫽ 3 different
transfections).
NTPDase1 and NTPDase2 differentially regulate
platelet function
The differential substrate specificity of NTPDase1 and NTPDase2
may explain functional effects of these enzymes. That NTPDase1
activity modulates platelet function in vivo has been suggested by
studies in mice deficient in NTPDase1.21 Therefore, we contrasted
the ability of both enzymes to influence platelet aggregation in the
presence of the nucleotides ATP (a competitive antagonist of this
process) and ADP (an agonist).18,19
Consistent with the literature, no aggregation occurred when 20
␮M ATP was added to PRP (Figure 2A). Addition of protein
extracts from NTPDase1-transfected COS cells, in the presence of
ATP, resulted in low levels of reversible aggregation that appeared
related to conversion of ATP to ADP. However, because NTPDase1
efficiently hydrolyzed both nucleotides, induced platelet aggregation was rapidly and completely reversed (n ⫽ 6). In contrast, high
levels of platelet aggregation were obtained when NTPDase2
protein extracts were added to PRP and ATP (n ⫽ 4). Similar data
were obtained when protein extracts were preincubated with the
PRP samples prior to the addition of ATP. Experiments with control
protein extracts from COS cells transfected with pcDNA3 empty
vector revealed no alterations in either platelet shape change or
aggregation responses (Figure 2B).
Lower concentrations of nucleotides were also tested in various
combinations. Mixtures of 1 ␮M ADP and 4 ␮M ATP added to PRP
preincubated with control protein extracts resulted in low levels of
platelet aggregation (Figure 2C). In the presence of low levels of
NTPDase1 (4 ␮g per assay), the initial rates of platelet aggregation
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SÉVIGNY et al
BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
similar aggregation profiles to the assays performed without
protein extracts (data not shown).
In summary, NTPDase1 largely inhibits and reverses platelet
aggregation in the presence of ADP and/or ATP. In contrast,
NTPDase2 promotes platelet aggregation in the presence of ATP
and further facilitates it in the presence of ADP.
Immunolocalization of NTPDase1 and NTPDase2
in mouse heart
Figure 2. Opposing effects of NTPDase1 and NTPDase2 on platelet aggregation. PRP prepared from human donors was tested for platelet activation in the
presence of exogenous nucleotides (ATP and/or ADP) and protein extracts from COS
cells transfected with pcDNA3 encoding NTPDase1 or NTPDase2. Light transmittance (%) was recorded over a period of 8 (B-D) or 10 minutes (A). Protein extracts
from COS cells transfected with vector DNA were added as controls at the required
protein level. Representative aggregation profiles from 3 to 6 experiments are shown.
(A) An arrow indicates the addition of 20 ␮M ATP to PRP followed 3 minutes later by
the addition of 20 ␮g NTPDase1 or NTPDase2 protein preparation (second arrow).
(B) PRP was preincubated for 25 seconds with 20 ␮g NTPDase1 (0.3 U ATPase) or
NTPDase2 (0.1 U ATPase) protein extracts before the addition of 40 ␮M ATP.
(C) PRP was preincubated for 25 seconds with 0.06 units of ATPase activity of
NTPDase1 (4 ␮g) or NTPDase2 (12 ␮g) prior to the addition of a mixture of
nucleotides (4 ␮M ATP plus 1 ␮M ADP). (D) PRP was preincubated for 25 seconds
with 0.06 units of ATPase activity with either NTPDase1 (4 ␮g ⫽ 0.043 U ADPase) or
NTPDase2 (12 ␮g) prior to the addition of 5 ␮M ADP.
were increased (90 arbitrary units/min ⫾ 4) when compared with
controls (63 arbitrary units/min ⫾ 4; P ⫽ .02, n ⫽ 3) but were
transient and rapidly reversed. Higher concentrations of NTPDase1
COS cell extracts (⬎ 20 ␮g per assay) totally prevented aggregation (data not shown). In contrast, in the presence of NTPDase2
samples, marked and sustained platelet aggregation always occurred.
Experiments in the presence of 5 ␮M ADP induced high levels
of platelet aggregation in control samples (amplitude of 73% ⫾ 2%,
n ⫽ 3); this could be reversed by the addition of NTPDase1 (Figure
2D). The initial extent of aggregation was slightly increased by the
addition of NTPDase2 (78% ⫾ 3%, P ⫽ .03, n ⫽ 3). Notably,
NTPDase2 did not reverse the low levels of platelet aggregation
induced by 1 ␮M ADP, even when larger amounts of NTPDase2
were added (700 ␮g transfected cell lysate per milliliter of PRP;
data not shown). These data confirmed that ADPase activities of
NTPDase2 were minimal and could not compete with platelet P2
receptor binding of the agonist ADP.
Experiments were also conducted in the simultaneous presence
of both enzymes. Results obtained with the addition of equivalent
units of ATPase activity from NTPDase1 and NTPDase2 protein
extracts gave similar results to NTPDase1 alone. These data
suggested that the effects of NTPDase1 on platelet aggregation
were dominant over NTPDase2 (data not shown).
All aggregation data presented above were confirmed with
PRP from different donors and using different batches of
recombinant proteins. In all experiments conducted here, protein extracts from COS cells transfected with control DNA gave
NTPDase1 and NTPDase2 pAbs were generated against the murine
proteins and specificity established by Western blot analysis
(Figure 3). As expected, NTPDase1 antiserum detected bands of
about 78 and 54 kd (proteolytic product of the 78-kd form) in the
protein extracts of COS cells transfected with the NTPDase1
construct, as we previously observed for bovine, porcine, and
human NTPDase1.24,29,42 No bands were detected in control protein
extracts (Figure 3). NTPDase2 antibodies detected a band evaluated at 75 kd in the protein extracts from cells transfected with
NTPDase2 construct only. Specificity of the antibodies was also
analyzed for the ability of the anti-NTPDase pAbs to bind the
native protein at the cell surface. Cross-reactivity of NTPDase1
pAb with NTPDase2 was excluded and vice versa.43 These antisera
were used to elucidate the distribution of NTPDase1 and NTPDase2 in vivo by immunohistology.
In most organs studied, immunolocalization of both NTPDase1
and NTPDase2 was predominantly confined to vascular structures
(Figures 4 and 5 and data not shown). However, NTPDase1 and
NTPDase2 were expressed on different cell types within the
vascular wall of both microvessels and larger muscularized vessels
(Figures 4 and 5 and data not shown).
Serial sections of muscularized vessels stained with smooth
muscle cell and endothelial cell markers suggested that NTPDase1
was predominantly expressed on endothelium and to a lesser extent
in the smooth muscle layer. In contrast, NTPDase2 was expressed
in cells present in the vascular adventia, coinciding with staining
for the basal lamina component, laminin (Figure 4A-F). Triple
immunofluorescent staining (using antibodies recognizing NTPDase1, NTPDase2, CD31 [marker of endothelial cells], and ␣-actin
Figure 3. Specificity of antisera for murine NTPDase1 and NTPDase2. Protein
samples (5 ␮g) from COS cells, transiently transfected with NTPDase1 or NTPDase2
cDNA constructs or from cells transfected with empty vector DNA (as a negative
control), were fractionated on 10% acrylamide SDS–polyacrylamide gel electrophoresis under nonreducing conditions. Separated proteins were then transferred to an
Immobilon-P membrane and incubated with pAbs directed against NTPDase1 or
NTPDase2. Each antiserum detected protein bands only in the appropriately
transfected cells.
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VASCULAR NTPDases AND THROMBOREGULATION
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Figure 4. Immunohistologic localization of NTPDases in cardiac vasculature. Serial sections of a mouse heart were stained by immunohistochemistry with various
antibodies detecting (A) NTPDase1, (B) NTPDase2, (C) ␣-actin (a marker of smooth muscle), (D) CD31 (a marker of endothelium), (E) NG2 (a marker of pericytes and smooth
muscle cells), and (F) laminin (a marker of basement membranes). Positive reactions were seen as rust color (A-F), and sections were counterstained with hematoxylin (blue).
In panels G-I, immunofluorescence and confocal analysis were performed: (G) CD31 in blue and NTPDase1 in green (colocalization as aquamarine); (H) as for panel G with
addition of NTPDase2 in red; (I) NTPDase1 in green, smooth muscle ␣-actin in blue, and NTPDase2 in red (no colocalization). In medium-sized vessels, CD31 and NTPDase1
were found to colocalize while NTPDase2 did not colocalize with these 2 markers or the smooth muscle ␣-actin–expressing cells. Original magnifications, ⫻200.
[marker of smooth muscle cells], analyzed by confocal microscopy) confirmed that NTPDase1 was predominantly localized to
the endothelium, whereas NTPDase2 was expressed in the advential layer of muscularised vessels (Figure 4G-I). Standard markers
recognizing endothelium, pericytes, smooth muscle cells, and
macrophages did not stain cells in the advential layer of mediumsized muscularized vessels, indicating absence of vasa vasora in
these vessels (Figure 4 and data not shown). The cellular component of the vascular adventia in smaller muscularized vessels
consists predominately of nerves and fibroblasts embedded in a
connective tissue matrix.44
Standard immunohistochemical techniques also demonstrated
that NTPDase1 and NTPDase2 were expressed in microvascular
structures (Figure 5A-G). To identify the cell types expressing
NTPDase1 and NTPDase2, double and triple immunofluorescent
staining was performed (using antibodies recognizing pericytes and
endothelial cells in conjunction with antibodies to NTPDase1 and
NTPDase2). Samples were then analyzed using confocal laser
microscopy. Expression of NTPDase1 was localized to the endothelium, whereas NTPDase2 was expressed in cells juxtapositioned to
the endothelium and enveloped in the basal membrane, suggesting
that NTPDase2 was expressed on pericytes (Figure 5). Double
immunofluorescent staining with biotinylated IgG recognizing
NTPDase2 in conjunction with NG2 confirmed the predominant
expression of NTPDase2 on microvascular pericytes (Figure
5K-M). Anti-NTPDase2 and anti-NG2 antibodies colocalized with
variable intensities within these individual structures, suggesting
that the distribution of these markers on the pericyte cell surface
might differ somewhat. Also of interest, NTPDase1 was expressed
in cells in the endocardium, whereas NTPDase2 was expressed in a
population of cells located immediately adjacent to the endocardium, in the subendocardial space (data not shown).
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BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
SÉVIGNY et al
Figure 5. Immunohistologic localization of NTPDases in the cardiac microvasculature. Immunohistochemistry of mouse heart sections (A-G) are shown in the same
order as in Figure 4. Antibody specificity: (A) NTPDase1, (B) NTPDase2, (C) ␣-actin (a marker of smooth muscle), (D) CD31 (a marker of endothelium), (E) NG2 (a marker of
pericytes and smooth muscle cells), (F) laminin (a marker of basement membranes), and (G) a higher magnification of NTPDase2. Sections were counterstained with
hematoxylin. In panels H-M, immunofluorescence and confocal analysis were performed: (H) NTPDase1 in green and CD31 in blue (colocalization is aquamarine);
(I) NTPDase1 in green and NTPDase2 in red; (J) CD31 in blue and NTPDase2 in red (no colocalization); (K) pericyte marker NG2 in green; (L) NTPDase2 in red; (M) panels K
and L combined (colocalization is yellow). In the microvasculature, NTPDase1 colocalized with an endothelial cell marker (CD31) while NTPDase2 colocalized with a pericyte
marker (NG2). Original magnifications A-F and H-M, ⫻200; G, ⫻400.
Alpha-actin is widely used as a marker for pericytes and smooth
muscle cells. However, pericyte expression of ␣-actin varies
between different tissues and is dependent upon cellular activation.36,41,45,46 In the mouse heart, ␣-actin was found to be present on
smooth muscle cells of larger vessels but was consistently absent
on microvascular pericytes (Figures 4C and 5C). In this organ, no
coexpression of NTPDase2 and ␣-actin was observed. Partially
muscularized vessels contained supporting cells that were either
positive for both NG2 and smooth muscle ␣-actin or for NG2 only.
These data suggest that such vessels contain intermediate cells, a
transitional cell with a phenotype that shares characteristics with
both pericytes and smooth muscle cells.44 In these vessels, NTPDase2 expression was confined to NG2-positive and smooth muscle
␣-actin–negative cells, supporting the initial findings that NTPDase2 is exclusively expressed on microvascular pericytes (data
not shown).
In other tissues analyzed, NTPDase2 was also consistently
absent from the smooth muscle layer of medium-sized vessels. In
these tissues, including striated muscle and large intestine, smooth
muscle ␣-actin expression was present in microvessels. In the latter
tissues, NTPDase2 colocalized with smooth muscle ␣-actin, suggesting that the expression of NTPDase2 on pericytes was not
confined to the heart muscle but applies to other organ systems as
well (data not shown). Qualitatively, similar results with respect to
the cellular distribution of NTPDase1 and NTPDase2 were observed in murine striated muscle, large intestine, and heart (Figures
4 and 5 and data not shown).
Discussion
We and others have previously identified the major vascular
endothelial cell NTPDase to be CD39 and demonstrated thomboregulatory properties of this ectoenzyme.5,21,28 We have now
identified a second NTPDase that is expressed in vasculature—
namely, NTPDase2 (or CD39L1). We observed that NTPDase1 and
NTPDase2 have similar molecular masses of 78 kd and 75 kd,
respectively (Figure 3), and we have contrasted the individual
biochemical properties of the vascular NTPDase1 and NTPDase2.
Specifically, NTPDase1 hydrolyses both ATP and ADP while
NTPDase2 is a preferential ecto-ATPase7,14 (Figure 1). These
differential biochemical properties of NTPDase1 and NTPDase2
may have important implications with respect to platelet function.
By hydrolyzing ADP, NTPDase1 inhibited platelet aggregation; in
contrast, NTPDase2 promoted platelet aggregation in the presence
of ATP and facilitated this process in the presence of ADP (Figure
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BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
2). The more pronounced aggregation observed in the latter
experiments could be explained by the hydrolysis of ATP released
from the platelet granules in an autocrine and paracrine manner.
This process would increase the local concentrations of ADP
because platelets concentrate both ATP and ADP in dense granules
to the order of 1 M.47 Interestingly, our data suggest also that
NTPDase2 facilitates ADP-specific P2 receptor activation by the
conversion of ATP. Indeed, ATP is generally considered a competitive antagonist of platelet P2Y1 and P2Y12 while ADP triggers their
activation and promotes aggregation,18-20 as seen in Figure 2. Thus,
NTPDase1 and NTPDase2 activities have opposing effects on
platelet aggregation in vitro.
NTPDase2 did not influence platelet aggregation in the presence of low micromolar ADP. On the other hand, NTPDase2
efficiently converted low micromolar concentrations of ATP to
ADP, triggering platelet aggregation in vitro. These data demonstrate that the rate of ATP hydrolysis by NTPDase2 with the
consequent generation of ADP is potentially relevant to the
regulation of platelet activation in vivo. ATP appears to be released
by injured cells while platelets release both ATP and ADP in similar
concentrations.47 Also of interest, the NTPDase1 effects on nucleotide-mediated platelet aggregation appear dominant over NTPDase2 activity; both enzymes in combination inhibited platelet
aggregation as efficiently as did NTPDase1 alone (data not shown).
Consistent with the opposing actions on platelet aggregation,
NTPDase1 and NTPDase2 were expressed within different strata
and by different cell types within the vasculature. In mouse heart,
we observed that NTPDase1 was expressed on vascular endothelium and endocardium and to a lesser extent on vascular smooth
muscle cells—in keeping with published descriptions of other
vascular beds as carried out by standard immunohistochemistry.24,48-50 These data, and other unpublished observations (J.S.,
E.C., S.C.R., 1999), demonstrate that NTPDase1 is consistently
expressed within the vasculature of mammals.
To date, little has been determined about the expression and
function of NTPDase2.6,7,14,51 NTPDase2 mRNA has been detected
in homogenates of several human and rat tissues.6,7 In vitro,
NTPDase2 mRNA expression in mouse hepatoma cells has been
shown to be induced by dioxin.52 NTPDase2 mRNA has also been
detected in the PC12 cell line.7 By immunohistologic techniques,
we demonstrate here that NTPDase2 is expressed in the vasculature, mainly by microvascular pericytes, adventitial cells in muscularised vessels, and by distinct cell populations in the subendocardial space.
This differential localization of NTPDase1 and NTPDase2 may
have direct implications for the control of platelet activation and
coagulation responses in vivo. The expression of NTPDase1 on the
endothelium and endocardium allows clearance of ADP (and ATP)
from the blood plasma, thus blocking and/or modulating platelet
aggregation under resting conditions.21,53 NTPDase1 may represent
an important thromboregulatory factor, together with nitric oxide
and prostaglandins PGI2 and D2, in the modulation of platelet
activation.21,53 Targeting the endothelium with recombinant adenoviral-expressed NTPDase1 to increase levels of expression of this
ectoenzyme has been shown to have beneficial thrombomodulatory
effects in transplantation models.21,54 In addition, both cd39⫹/⫺ and
cd39⫺/⫺ mice express a prothrombotic phenotype (Enjyoji et al21;
M.I., O.G., J.S., S.C.R., unpublished observation, November 2000).
Upon damage to the vessel, supporting cells that express
NTPDase2 are exposed to increased levels of ATP released from
injured cells, red blood cells, activated platelets, smooth muscles,
VASCULAR NTPDases AND THROMBOREGULATION
2807
and endothelium.47 Subsequent conversion to ADP by NTPDase2
will then promote platelet plug formation. Notably, strong effects
on platelet aggregation were observed with minimal amounts of
recombinant NTPDase2 in vitro. Comparable levels of recombinant NTPDase1 and NTPDase2 (at either units of ATPase activity
or total protein levels) both influence platelet aggregation in vitro
(Figure 2 and data not shown). This specific localization of
NTPDase1 and NTPDase2 spatially compartmentalizes the distinct
catalytic activities of these 2 enzymes within different elements of
the vasculature. The functional and structural integrity of the
endothelium dictates that NTPDase1 activity is dominant under
quiescent conditions. This scenario would facilitate basal antithrombotic activity of NTPDases (NTPDase1) that could rapidly shift to
a prothrombotic activity upon vascular damage and consequent
exposure of elements of the outer vessel wall to blood (NTPDase2).
It has been proposed that pericytes and adventitial cells may be
important in the process of clot formation through effects on the
coagulation cascade.55-58 Our results suggest that these cell types
also promote clot formation in a previously unrecognized fashion
via direct effects on platelet function. Pericytes and adventitial cells
may serve therefore as a hemostatic envelope capable of initiation
and/or promotion of thrombus formation when vascular integrity
is disrupted.
NTPDase2 may serve as a novel marker to discriminate
between pericytes and smooth muscle cells in situ. A limited
number of markers for pericytes have recently become available
that have facilitated purification and research on the functional
significance of these cells. However, the interpretation of the
expression of such “specific” markers reveals several limitations.
Expression of different pericyte markers depends largely on the
activation state of the tissue and has proven less useful in
identifying pericytes in resting tissues. No antibodies can discriminate between pericytes, smooth muscle cells, and myofibroblasts.
Importantly, we showed that NTPDase2 is not expressed on smooth
muscle cells or on transitional supporting cells in partially muscularized vessels that share characteristics between smooth muscle
cells and pericytes,44 thus allowing discrimination between these
cell types. NTPDase2 was expressed on microvascular pericytes in
resting tissues from several different organs (Figures 4 and 5 and
data not shown). Our data suggest that NTPDase2 may serve as a
useful marker for defining and characterizing pericytes in vivo.
Biochemical properties and specific localization of NTPDase2
in the heart suggest that the enzyme may regulate functions other
than platelet activation. Extracellular ATP and its degradation
product adenosine exert pronounce inotropic and chronotropic
actions and influence electrical impulse conduction as well as
metabolic processes.59-61 Adenosine has also been shown to have
important cardioprotective effects following ischemic injury to the
heart.62,63 These effects are triggered through the activation of
various adenosine60 and P2 receptors.64-66
The hydrolysis of ATP by NTPDase2 at the surface of adventitial cells and pericytes may also be important in the control of
vascular tone—a function that could be coordinated further by
NTPDase1 expressed on vascular smooth muscle cells. Indeed P2X
receptors,67 predominantly P2X1, are expressed on vascular smooth
muscle cells as well as P2Y1, P2Y2, P2Y4, and P2Y6 receptors.68
Much of the vasomotor tonic action of ATP can be shown to be
related to the activation of P2X1, but P2Y-mediated signaling also
plays a role in this process.68 To our knowledge, there are no
published studies that document P2 receptor expression in pericytes. However, there are similarities between pericytes and
smooth muscle cells, known to express various P2 receptors. P2Y
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2808
BLOOD, 15 APRIL 2002 䡠 VOLUME 99, NUMBER 8
SÉVIGNY et al
receptors are also involved in vascular smooth muscle cell proliferation.68,69 It is unclear whether NTPDase1 and NTPDase2 play
regulatory roles in these events.
In summary, differential biochemical properties and expression
patterns of NTPDase1 and NTPDase2 correlate with the distinct
functions of these enzymes on platelet aggregation. Importantly,
NTPDase1 faces the blood circulation and abrogates platelet
aggregation. In contrast, NTPDase2 is expressed in supporting
cells of the vasculature and facilitates platelet aggregation. Our
data further support the view that NTPDases may not only
directly terminate P2 receptor signaling but under certain
circumstances may also mediate the activation of specific ADP
receptors such as platelet P2Y170 and P2Y12 by the generation of
the specific agonists.
Acknowledgment
We thank Dr R. Neal Smith of the Immunopathology Unit,
Department of Pathology, Massachusetts General Hospital, Boston,
MA, for valuable discussion.
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2002 99: 2801-2809
doi:10.1182/blood.V99.8.2801
Differential catalytic properties and vascular topography of murine nucleoside
triphosphate diphosphohydrolase 1 (NTPDase1) and NTPDase2 have
implications for thromboregulation
Jean Sévigny, Christian Sundberg, Norbert Braun, Olaf Guckelberger, Eva Csizmadia, Imrana Qawi, Masato
Imai, Herbert Zimmermann and Simon C. Robson
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