Extracellular ATP signaling and P2X nucleotide receptors in
monolayers of primary human vascular endothelial cells
Lisa M.
Schwiebert,
William C.
Rice,
Brian A.
Kudlow,
Amanda L.
Taylor, and
Erik M.
Schwiebert
Department of Physiology and Biophysics and Department of Cell
Biology, University of Alabama at Birmingham, Birmingham, Alabama
35294 - 0005
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ABSTRACT |
ATP and its metabolites
regulate vascular tone; however, the sources of the ATP released in
vascular beds are ill defined. As such, we tested the hypothesis that
all limbs of an extracellular purinergic signaling system are present
in vascular endothelial cells: ATP release, ATP receptors, and ATP
receptor-triggered signal transduction. Primary cultures of human
endothelial cells derived from multiple blood vessels were grown as
monolayers and studied using a bioluminescence detection assay for ATP
released into the medium. ATP is released constitutively and
exclusively across the apical membrane under basal conditions.
Hypotonic challenge or the calcium agonists ionomycin and thapsigargin
stimulate ATP release in a reversible and regulated manner. To assess
expression of P2X purinergic receptor channel subtypes (P2XRs), we
performed degenerate RT-PCR, sequencing of the degenerate P2XR product, and immunoblotting with P2XR subtype-specific antibodies. Results revealed that P2X4 and P2X5 are expressed
abundantly by endothelial cell primary cultures derived from multiple
blood vessels. Together, these results suggest that components of an
autocrine purinergic signaling loop exist in the endothelial cell
microvasculature that may allow for "self-regulation" of
endothelial cell function and modulation of vascular tone.
purinergic receptors; cytosolic calcium; ectoadenosinetriphosphatases; exocytosis; nitric oxide
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INTRODUCTION |
PURINERGIC SIGNALING
REGULATES circulatory function (4, 15, 20, 21, 27).
Nowhere has this signaling been studied more extensively in the
cardiovascular system than in adenosine modulation of cardiac function
(2, 12, 17). G protein-coupled P2Y purinergic receptors
are expressed on vascular smooth muscle cells, platelets, and
endothelial cells (4, 15, 20, 21, 27, 29, 33). In
endothelial cells derived from multiple vessels, extensive study has
shown the molecular and functional expression of P2Y1 and
P2Y2 receptors (4, 15, 20, 21, 27, 29, 33). A
specific isoform of the P2X purinergic receptor channel family,
P2X1, a protein that binds ATP in its extracellular domain
and also acts as a Ca2+-permeable, nonselective cation
channel, was cloned from vascular smooth muscle (32).
Expression of P2X1 has been documented in human umbilical
and human renal vasculature (3, 9). More recently,
expression of P2X4 as well as weaker expression of other P2XR subtypes was documented by Northern blot analysis and competitive, specific RT-PCR in endothelial cells (35). Immunoblotting
for P2XR proteins as well as comprehensive studies of ATP release from
these same human vascular endothelial cells was not performed (35). Only a few additional studies have hinted at P2X
receptor expression in vascular cells (16, 23).
Despite the fact that, for decades, it has been postulated that ATP and
adenosine act locally within vascular beds to regulate tissue vascular
tone, the sources of ATP within the vasculature are largely unknown.
Moreover, P2XR expression has not been characterized fully in
vasculature. In particular, ATP release by endothelial cells is ill
defined. As such, we tested the hypothesis that the endothelium, the
layer of cells that lines all blood vessels, may provide a source of a
physiological purinergic agonist, ATP, locally within vascular beds.
Moreover, we tested the hypothesis that endothelial cells, in addition
to vascular smooth muscle cells, also may express multiple subtypes of
the emerging P2XR family to bind this released ATP and transduce this
extracellular ATP signal in an autocrine or paracrine manner. Results
demonstrate that human endothelial cell monolayers release ATP
exclusively into the apical medium in a regulated manner and that
endothelial cells express multiple P2XR subtypes.
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MATERIALS AND METHODS |
Endothelial cell primary culture.
Human endothelial cells were isolated into primary culture from
different blood vessels by Clonetics and purchased as proliferating cells. The cultures were grown on 2% gelatin-coated or 6%
collagen-coated tissue culture flasks, 35-mm dishes, or 12-mm-diameter
filter supports (Millicell CM) in endothelial basal medium (Clonetics) supplemented with 5% fetal bovine serum (FBS) and a BulletKit of
additional additives including (per 500 ml) human epidermal growth
factor (0.5 ml), hydrocortisone (0.5 ml), gentamicin (0.5 ml), bovine
pituitary extract (2.0 ml), 1× fungizone, and 1×
penicillin-streptomycin (31). Filters were bathed in
medium on both sides of the filter until a monolayer formed that was
tight to fluid for at least 12 h; monolayers formed after 4-6
days in culture. The endothelial cell primary cultures prepared are as
follows: human umbilical vein endothelial cells (HUVEC), human
umbilical artery endothelial cells (HUAEC), human coronary artery
endothelial cells (HCAEC), human pulmonary artery endothelial cells
(HPAEC), human aortic endothelial cells (HAEC), and human lung
microvascular endothelial cells (HLMVEC).
Bioluminescence detection of ATP released from endothelial cell
cultures and monolayers.
These methods and materials have been published previously
(30) in detail for experiments on epithelial monolayers.
Similar methods were followed for endothelial monolayers. Briefly,
Opti-MEM-I medium (GIBCO-BRL) with 2 mg/ml luciferin-luciferase reagent
(Calbiochem) was added to the apical or basolateral side of the
monolayer or directly into a culture grown in 35-mm dishes. Basal
levels of ATP release were measured for at least 2 min in 15-s,
nonintegrated photon collection periods in a TD-20/20 Luminometer
(Turner Designs). Distilled water with 2 mg/ml luciferin-luciferase was
added to dilute the osmotic strength of the Opti-MEM, and luminescence was measured for at least 2 min as described above.
4,4'-Diisothiocyanostilbene-2,2'-disulfonic acid (DIDS) and NaCl were
added acutely during these luminescence measurements, since they had no
effect on luciferase enzyme activity (7, 30). Apyrase or
hexokinase was added at the end of the time courses to eliminate any
ATP left in the medium. All assays were performed at room temperature
unless otherwise specified. Specifics on data analysis and statistics
have been published previously (30) or are described in
figure and table legends.
Some important additional points must be emphasized that were not
pointed out in our original paper on this assay (30). Different batches of luciferin-luciferase reagent were needed to
complete these studies, giving rise to some differences in luminescence
detection (for example, see Table 1 vs. Table 2). Unfortunately, we could not perform an ATP standard curve on every vial
of detection reagent because we would have run out of detection reagent
for the actual ATP release studies on endothelial cell monolayers. We
also did not inhibit ecto-ATPases in any pharmacological way in these
assays (besides cooling of the monolayers). It also is important to
note that the medium used in the release detection assay included 140 mM chloride and that chloride inhibits luciferase activity. As such,
together with ecto- ATPase activity uninhibited and luciferase
inhibited by physiological amounts of extracellular chloride, we may be
underestimating ATP levels in the extracellular milieu. Therefore,
luciferase acts as an "ATP sensor" in this bioassay, not a consumer
of ATP.
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Table 1.
Luminescence vs. absolute concentration of ATP vs. rate of ATP efflux,
and ATP standard curve and consumption of ATP by luciferase-luciferin
reagent over time
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Degenerate P2X receptor RT-PCR.
Endothelial primary cultures were grown to confluence on
collagen-coated flasks, and total RNA was extracted with Trizol reagent (Life Technologies). Total RNA was treated with DNase and reverse transcribed by standard methods. Each PCR reaction contained 1 µM
dNTP mix, 2 µM forward primer, 2 µM reverse primer, 0.5 µl Taq polymerase (Perkin-Elmer), 10× PCR buffer (1.5 mM
MgCl2), and cDNA template. The PCR cycle began with a
5-min, 94°C "hot start" and was followed by 40 cycles of 30 s at 94°C, 60 s at 52°C, and 60 s at 72°C. A 10-min,
72°C elongation ended the reaction. PCR products were run on a 1.5%
agarose gel beside a 100 base pair (bp) DNA ladder. Primers were as
follows:
-actin forward primer, 5'-TGA CGG GGT CAC CCA CAC TGT GCC
CAT CTA-3';
-actin reverse primer, 5'-CTA GAA GCA TTG CGG TGG ACG
ATG GAG GG-3'; P2XR degenerate forward primer, 5'-TTC ACC MTY YTC ATC
AAR AAC AGC ATC-3'; and P2XR degenerate reverse primer, 5'-TGG CAA AYC TGA AGT TGW AGC C-3'.
PCR products were rescued from agarose gel slices using the Qiaquick
gel extraction kit (Qiagen). The purified PCR product was ligated into
the pGEM-T vector (Promega). The ligations were transformed into JM109
competent cells (Promega), and the cells were plated onto Luria broth
(LB) agar plates containing ampicillin (100 µm/ml), X-Gal (40 µl/ml), and isopropyl
-D-thiogalactopyranoside (100 µg/ml). White colonies (those with ampicillin resistance and an
interrupted
-galactosidase gene) were picked and grown in 6 ml of
LB-ampicillin. The plasmids were isolated from the bacteria
using a PerfectPrep miniprep kit (5 Prime-3 Prime). The purified DNA
was denatured in 0.2 N NaOH and precipitated in 7.5 M NH4Cl
and 100% ethanol. The Sequenase dideoxy termination method (Amersham)
with
-35S-labeled dATP (NEN-Dupont) was used to sequence
the purified DNA product. The DNA sequence was read and screened with
the BLAST algorithm to determine the identity of the PCR product
(1). In this and other studies on epithelial cells,
sequences for every P2XR subtype were amplified with the exception of
P2X6, a brain-specific subtype (data not shown).
Immunoblotting with P2X receptor channel subtype-specific
antibodies.
Cells were lysed in a buffer containing 10 mM Tris, 0.5 mM NaCl, 0.5%
Triton X-100, 50 µg/ml aprotinin (Sigma, St. Louis, MO), 100 µg/ml
leupeptin (Sigma), and 100 µg/ml pepstatin A (Sigma) adjusted to pH
7.2-7.4. Protein (20 µg/lane) was run and separated on an 8%
SDS polyacrylamide gel and then transferred to polyvinylidene difluoride membrane (Osmonics, Westborough, MA). Immunoblotting was
performed with rabbit polyclonal antibody to P2X1,
P2X2, P2X4, and P2X7 at a dilution
of 1:500 (Alomone Laboratories, Jerusalem, Israel) or with antibody to
P2X5 at a dilution of 1:1,000 (generous gift of Drs. Mark
Voigt and Terry Egan, St. Louis University, St. Louis, MO). Reactivity
was detected by horseradish peroxidase-labeled goat anti-rabbit
secondary antibody (1:3,000 dilution) (New England BioLabs, Beverly,
MA). Enhanced chemiluminescence was used to visualize the secondary antibody.
Fura 2-AM imaging of intracellular
Ca2+.
Cytosolic intracellular Ca2+ concentration
([Ca2+]i) in human vascular endothelial
cells was measured with dual-excitation wavelength fluorescence
microscopy (Deltascan, Photon Technologies, Princeton, NJ) using the
permeant form of the fluorescent probe, fura 2-AM (Teflabs,
Austin, TX). Fura 2 fluorescence was measured at an emission wavelength
of 510 nm in response to excitation wavelengths of 340 and 380 nm, alternated at a rate of 25 Hz by a computer-controlled chopper
assembly. Autofluorescence-corrected ratios (340 nm/380 nm) were
calculated at a rate of 5 points/s using PTI software. Cells were grown
on collagen-coated coverslips (1:30 dilution of Vitrogen 100 in PBS)
cut to fit the circulating cuvette and were incubated in media
containing 5 µM fura 2-AM and 1 mg/ml Pluronic F-127 dissolved in
dimethyl sulfoxide (DMSO) for 2 h to allow loading of the dye into
the cells. After loading, a coverslip was rinsed in Ringer solution to
remove extracellular fura 2-AM and was positioned in the cuvette at a
45° angle from the excitation light. Two glass capillary tubes were
inserted into the cuvette. One of these tubes was extended to the
bottom of the cuvette and connected by way of polyethylene tubing to an
infusion pump. The other capillary tube was positioned at the top of
the cuvette and served to remove fluid from the cuvette. Flow rate
through the cuvette was ~5 ml/min. A Ringer solution was used
containing (in mM) 148 NaCl, 5 KCl, 1 MgSO4, 1.6 Na2HPO4, 0.4 NaH2PO4, 5 D-glucose, 1.5 CaCl2, and 10 HEPES at pH 7.4 and at room temperature. After 20 min of incubation in the control
Ringer solution, fluorescence intensities of both wavelengths
stabilized. Once values were stable, purinergic agonists were added to
the circulating cuvette for testing. The 340/380 ratios (R) were
converted into [Ca2+]i values as follows:
[Ca2+] = Kd × [(R
Rmin)/(Rmax
R)] × (Sf380/Sb380), where Rmax and Rmin are R values under saturating and
Ca2+-free conditions, respectively, and Sf380
and Sb380 are the fluorescent signals (S) emitted by the
Ca2+-free (f) and Ca2+-bound (b) forms of fura
2 at the 380-nm wavelength. This was accomplished after the cells were
permeabilized with 5 µM ionomycin and fluorescence ratios and signals
were measured under Ca2+-free (2 mM EGTA) or
Ca2+-saturating (1.5 mM CaCl2) conditions using
fura 2 calibration solution (PBS containing 10 mM MgCl2, 2 mM EGTA). The dissociation constant (Kd) of fura
2 for Ca2+ was taken as 224 nM.
Data analysis and statistics.
A P value of <0.05 was considered significant, whether
determined by a paired Students' t-test for paired
experiments or by ANOVA with a Bonferroni ad hoc test for unpaired
data. Most of the experiments involved paired analysis. The figure
legends indicate the tests used and the P values calculated.
Materials.
All chemicals were obtained from Sigma unless otherwise noted.
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RESULTS |
Endothelial cells in primary culture form resistive monolayers in
vitro.
Human vascular endothelial cells were grown on 12-mm-diameter Millicell
PC permeable filter supports to establish endothelial monolayers. This
maneuver was necessary to study the sidedness of ATP release (see data
below). Routinely, within 4-7 days, the endothelial monolayers
were tight to fluid for a minimum of 1 h, which was significantly
longer than the duration of the ATP release assays. For the HUVEC
endothelial monolayers, transendothelial resistance
(RTEndo) was measured with a Voltohmeter.
RTEndo was measured 2 days after seeding of the
filters. After only 2 days, RTEndo was 154 ± 2
· cm2 (n = 48). Because
the resistance of the Millicell PC filter is 50
· cm2, significant
RTendo was achieved after only 48 h.
RTendo increased during the next 48-h period and
was 197 ± 2
· cm2 at day 4. On
day 5, RTendo was similar (193 ± 5
· cm2; n = 48), and
monolayers were used for ATP release assays on days
5-7. These data show that human vascular endothelial cells grown in primary cultures under these conditions formed endothelial monolayers.
ATP is released predominantly across the apical membrane of human
endothelial cell monolayers.
Bioluminescence detection of ATP released from human endothelial
cells grown in primary culture as monolayers on permeable supports was
performed to assess the magnitude and polarity of ATP release. Primary
endothelial cell cultures from six different blood vessel preparations
were compared. Significant apically directed ATP release was detected
from all endothelial monolayers under basal conditions (Fig.
1). In sharp contrast, little ATP release
was measurable in the basolateral medium under basal conditions, although these values were above background (Fig. 1). Although this is
a microassay that estimates ATP release from monolayers, correlation of
luminescence values with a standard curve of [ATP] indicated that
apically directed ATP release elaborated nanomolar quantities of ATP,
whereas basolaterally directed ATP release produced only picomolar
quantities (Table 1). In particular, HUVEC and HPAEC monolayers released the most ATP across the apical membrane, whereas lesser amounts were measured in the endothelial monolayers derived from other blood vessel sources. Table 1 also shows
that luciferase is present in sufficient quantity in the assay to
"sense" extracellular ATP but not to consume it rapidly over time.
However, because this is a microassay and because ecto-ATPases compete
with luciferase for the released ATP (see Fig. 6), the amount of
released ATP being detected was likely underestimated. Nevertheless, a
second y-axis (to the right of each luminescence data plot)
shows absolute [ATP] calculated from the standard curve data provided
in Table 1. Together, these results show that endothelial cells release
ATP under basal conditions constitutively and that this ATP release is
directed predominantly across the apical membrane.

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Fig. 1.
Sidedness of constitutive ATP release from polarized vascular
endothelial monolayers under basal or unstimulated conditions. A
numerical comparison of apical (APM) and basolateral medium (BLM) ATP
release is shown in the context of a circulatory system schematic.
Values [in arbitrary light units (ALU)] show that ATP is released
mainly into the apical medium; no. of experiments (n) is
shown in parentheses. Note that all endothelial monolayers release
significant quantities of ATP apically above background ( 0.1 ALU).
Release of ATP from human umbilical vein endothelial cell (HUVEC)
cultures grown in 35-mm dishes is also shown. HPAEC, human pulmonary
artery endothelial cells; HLMVEC, human lung microvascular endothelial
cells; HAEC, human aortic endothelial cells; HCAEC, human coronary
artery endothelial cells; and HUAEC, human umbilical artery endothelial
cells.
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In one long-lived HUVEC primary culture, ATP release was measured
across the apical membrane as a function of passage number (Fig.
2). Apically directed ATP release under
basal conditions was robust in early passages (Fig. 2). However, by
passages 9 and 10, the magnitude of ATP release
waned (Fig. 2). In contrast, basolaterally directed ATP release under
basal conditions followed a Gaussian-like distribution (Fig. 2). The
peak value of basolaterally directed ATP release was measured at
passage 8, whereas it was lower at earlier passages (Fig.
2). As in apically directed ATP release measurements, basolaterally
directed ATP release waned at passages 9 and 10.
This culture dedifferentiated at passage 11. All data from
HUVEC primary cultures are included in Fig. 1. These data show that ATP
release was dependent on the age of primary culture. Because apically
directed ATP release was much greater than basolateral ATP release,
apically directed ATP release results are the focus of all subsequent
discussion.

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Fig. 2.
ATP release is affected by passage of primary cultures release of
ATP from HUVEC endothelial monolayers by passage number (P5-P10)
of a primary culture. A: apically directed ATP release.
B: basolaterally directed ATP release. Cells were obtained
at passage 3. Apically directed ATP release was consistent
for several passes. Values are means ± SE; n is shown
within each bar.
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ATP release is potentiated by hypotonic challenge.
Previous studies by our laboratory alone or in collaboration with other
groups have shown that normal hepatocytes (13), normal
airway epithelial cells (7, 30), and normal and cystic renal epithelia (34) release ATP profoundly under
hypotonic conditions. Dilutions of the medium osmolality with
increasing volumes of distilled water were compared with medium
controls (similar added volumes of isotonic medium) in bioluminescence detection assays of ATP released from HUVEC monolayers. As little as
13% dilution stimulated a transient increase in ATP release, whereas
more robust dilutions of the medium osmolality produced sustained
increases in ATP release (Fig. 3).
Reconstitution of the medium osmolality by addition of a bolus of 50 mM
NaCl (100 mosmol) reversed hypotonicity-induced ATP release across the
apical membrane to levels measured under basal conditions (Fig. 3).
Further additions of NaCl, which made the medium hypertonic, inhibited basal ATP release in both water dilution and medium control experiments (Fig. 3). In fact, addition of 150 mM NaCl (300 mosmol) inhibited apically directed ATP release under basal conditions by at least 75%
(Fig. 3). Note that significant volumes of isotonic medium added to the
apical side of HUVEC monolayers had no effect on ATP release (Fig. 3),
showing that a dilution of osmolality and not a mechanical perturbation
was stimulating ATP release across the apical membrane of HUVEC
monolayers. Together, these results show that hypotonicity potentiates
ATP release across the apical membrane of HUVEC monolayers. Moreover,
hypertonicity (addition of osmoles during isotonic medium controls)
attenuates basal ATP release in medium controls (Fig. 3).

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Fig. 3.
Hypotonic challenge stimulates ATP release across the
apical membrane of HUVEC endothelial cell monolayers. Raw luminescence
values for HUVEC monolayers exposed to hypotonic challenge (Hypo) or to
equal volumes of isotonic medium (a similar amount of
luciferin-luciferase reagent was present in all stages of the
experiment; n = 6 each). Reconstitution of medium
osmolality with NaCl reversed hypotonicity-induced ATP release. Note
that hyperosmolality in medium controlled and, after the third addition
of 50 mM NaCl to water dilutions, inhibited basal ATP release to low
levels. The y-axis at right shows a correlation
with absolute [ATP] that is an approximation by correlation with the
standard curve shown in Table 1.
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Previously, we showed partial attenuation of ATP release in epithelial
monolayers with the broad-specificity, mechanosensitive ion channel
blocker GdCl3 (7, 28, 30). In HUVEC
monolayers, GdCl3 inhibited apically directed basal ATP
release partially and in a dose-dependent manner (Fig.
4, see legend for summarized data). In
this study, we also discovered a second ATP release inhibitor, DIDS, a
broad-specificity anion channel-blocking drug. Figure 4 also shows a
representative time course illustrating complete block of basal ATP
release with DIDS (see legend for summarized data). Similar blockade of
hypotonicity-induced ATP release across the apical membrane was
observed with GdCl3 and DIDS (Fig. 4). As in Fig. 3, graded
dilutions of the medium osmolality stimulated ATP release.
GdCl3 inhibited the potentiated ATP release partially,
whereas DIDS inhibited ATP release fully (Fig. 4; see legend for
summarized data). Together, these data show that ATP release is
inhibited by ion channel-blocking drugs and suggest that an
ATP-permeable conductive transport pathway may mediate ATP release
across the apical membrane of endothelial cell monolayers.

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Fig. 4.
Basal and hypotonicity-induced ATP release are inhibited
by ion channel blocking drugs. A: representative experiments
showing DIDS and GdCl3 inhibition of basal ATP release
across the apical membrane of HUVEC monolayers. At the 200 µM dose
(data grouped among HUVEC and HCAEC monolayers), GdCl3
inhibited basal ATP release by 47.75 ± 4.92% (n = 4), whereas DIDS inhibited basal ATP release by 93.51 ± 2.50%
(n = 4). The y-axis at right
shows a correlation to absolute [ATP] that is an approximation
by correlation with the standard curve shown in Table 1. B:
representative experiments showing DIDS and GdCl3
inhibition of hypotonicity-induced ATP release across the apical
membrane of HPAEC monolayers. At the 200 µM dose (data grouped among
HAEC and HPAEC monolayers), GdCl3 inhibited
hypotonicity-induced ATP release by 67.31 ± 4.35%
(n = 8), whereas DIDS inhibited hypotonicity-induced
ATP release by 92.25 ± 1.52% (n = 8). All
inhibition by the 2 ion channel blockers was significant
(P < 0.05). The y-axis at right
shows a correlation to absolute [ATP] that is an approximation
by correlation with the standard curve shown in Table 1.
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ATP release is potentiated by the
Ca2+ agonists ionomycin and thapsigargin.
Because agonists that increase cytosolic Ca2+ have profound
effects on endothelial cell function, signaling, and production and on
release of vasoactive mediators such as nitric oxide (22, 24), we examined the effect of Ca2+ agonists on ATP
release across the apical membrane of HUVEC monolayers. Ionomycin (2 µM), a Ca2+ ionophore that promotes Ca2+
influx into the cell from extracellular stores, elicited an immediate increase in ATP release (Fig. 5). Unlike
the hypotonicity-induced ATP release phenotype where luminescence
increased and remained elevated for a few minutes before decaying over
time, ionomycin-induced ATP release was immediate and continued to
increase over the 8-min exposure period (Fig. 5). Addition of DIDS had
little inhibitory effect on the ionomycin response, whereas addition of
hexokinase rapidly abolished the luminescence signal (Fig. 5).
Thapsigargin (2 µM), an inhibitor of the Ca2+-ATPase pump
in organellar membranes that promotes a rise in cytosolic Ca2+ by blocking uptake and sequestration of
Ca2+ in intracellular stores, triggers a slower rise in ATP
release that peaked between 1 and 2 min following addition (Fig. 5).
This response decayed to basal values over the stimulation period, and
the luminescence signal was abolished by hexokinase (Fig. 5). Chronic
treatment with thapsigargin eventually lowers cytosolic Ca2+. The transient stimulation suggests that thapsigargin
in the later phase of the stimulation was causing cytosolic
Ca2+ to fall, inhibiting ATP release. Interestingly,
although DIDS inhibits basal ATP release in the absence of
thapsigargin, DIDS had no effect on the luminescence signal in the
presence of thapsigargin (Fig. 5), suggesting that the
DIDS-sensitive ATP release mechanism under basal conditions may require
intracellular Ca2+. Importantly, pretreatment of
endothelial monolayers with
1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM (10 µM, 30- to 60-min pretreatment), the permeable form of the Ca2+-chelating agent, abolished
ionomycin-induced ATP release. In the absence of BAPTA-AM, ionomycin
increased ATP release acutely from 6.79 ± 1.36 to 21.82 ± 3.99 arbitrary light units (ALU) [change (
): 15.03 ± 2.64 ALU; n = 5] within 15 s and to a sustained value 8 min after addition of 50.01 ± 3.99 ALU (
38.79 ± 1.90 ALU; n = 5). In the presence of the Ca2+
chelator, ionomycin failed to stimulate ATP release acutely
(
1.40 ± 0.80 ALU; n = 5) and chronically
(
2.02 ± 0.76 ALU; n = 5). Together, these data
show that intracellular Ca2+ regulates the release of ATP
across the apical membrane of human endothelia and promotes
extracellular ATP signaling in the endothelial microenvironment in
vitro.

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Fig. 5.
Ca2+ agonists stimulate ATP release that is
insensitive to the anion channel and transporter blocker, DIDS. A data
summary of 5 experiments is shown for each Ca2+ agonist.
The time course of ATP release follows the time course of action for
each Ca2+ agonist in terms of effects on cytosolic
Ca2+ [sustained increase with the Ca2+
ionophore ionomycin (Iono; A) and a transient increase with
the Ca2+ pump inhibitor thapsigargin (Thap;
B)]. DIDS was without effect, whereas hexokinase diminished
the signal fully. The y-axis at right shows a
correlation to absolute [ATP] that is an approximation by correlation
with the standard curve shown in Table 1.
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Temperature affects dynamics and magnitude of constitutive,
hypotonicity-induced, and Ca2+
agonist-stimulated ATP release assay: a role for exocytosis and
ecto-ATPases.
To address the possible role of exocytosis in the release of ATP via
ATP-filled vesicles, we performed our standard ATP release assay in
temperature-controlled rooms at 4, 25, 30, and 37°C. Fortunately,
these temperatures do not affect the luciferase detection assay
significantly (Table 2). Incubation of
cells or monolayers at 4°C slows vesicle trafficking along the
secretory pathway, even preformed vesicles that are poised to fuse with
the plasma membrane. Figure 6A
shows that constitutive release was not affected in
its magnitude by performing the experiment at 4°C vs. 25°C. In
sharp contrast, however, ~40% of the Ca2+ agonist- and
hypotonicity-induced ATP release phenotype was abolished by reduced
temperature incubation and assay, suggesting that exocytosis may play a
partial role as a mechanism of ATP release.

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Fig. 6.
Change in bioluminescence assay temperature reveals
roles for exocytosis and ecto-ATPases. A: data summary
showing effect of 4°C vs. 25°C on constitutive (basal),
Ca2+ agonist-induced, and hypotonicity-induced ATP release
(n = 7 each). Loss of stimulated ATP release suggests a
role for vesicle-mediated release. Delta luminescence, change in
luminescence. *P < 0.05. B: data summary
showing effect of 37°C vs. 25°C on constitutive (basal),
Ca2+ agonist-induced, and hypotonicity-induced ATP release
(n = 7 each). Loss of signal at physiological
temperature indicates that ecto-ATPases are competing successfully for
the released ATP. *P < 0.05. C: summarized
data showing increase in luminescence signal at 4°C (in the presence
of attenuated ecto-ATPases) vs. rapid decay in signal at 37°C (in the
presence of fully active ecto-ATPases). The y-axis at
right shows a correlation to absolute [ATP] that is an
approximation by correlation with the standard curve shown in Table
1.
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To our surprise, assay temperatures higher than room temperature
produced no increase in bioluminescence. At 37°C, we observed a
reduced signal at all stages of the assay (Fig. 6B). At the end of the first stage of the assay monitoring basal release, the
37°C signal was less than room temperature (Fig. 6B). The Ca2+ agonist- and hypotonicity-induced ATP release signal
also was reduced at higher temperature (Fig. 6B). Upon more
careful review of the data, we found that the decay in bioluminescence
in all stages of the experiment was most pronounced at 37°C. This is shown best when comparing basal or constitutive ATP release at 4°C
vs. 37°C (Fig. 6C); completely different phenotypes are
observed. A gradual rise in bioluminescence was observed at 4°C,
suggesting that ecto- ATPases no longer compete for the released ATP.
In sharp contrast, at 37°C, the signal peaked within 1 min and
decayed by almost 50% by 6 min. In these assays, constitutive ATP
release was measured for 6 min, followed by ionomycin-induced release for 8 min and hypotonicity-induced release for 6 min. Blockade with
DIDS or GdCl3 and hexokinase was then carried out.
Together, these data show that ecto-ATPase activity, attenuated at
25°C and inhibited at 4°C, is maximally active at 37°C and
competes successfully with luciferase for the released ATP. Moreover,
the loss of signal at 37°C suggests that ecto-ATPase activity
outcompetes luciferase, whereas the attenuated signal at 4°C for
Ca2+ agonist- and hypotonicity-induced ATP release suggests
a component of ATP release that relies on exocytosis of ATP-filled vesicles.
P2X receptor channels are expressed by human endothelial cells.
To test the hypothesis that subtypes of a subclass of purinergic
receptors, the P2X receptor channels (P2XRs), are expressed in
endothelial cells, we performed degenerate RT-PCR. To confirm the
authenticity of the cDNA before P2XR RT-PCR, we performed RT-PCR with
primers for the housekeeping gene
-actin (Fig.
7). The same cDNA was amplified with
primers designed from an alignment of the first three rat P2XR cDNA
sequences (rP2XR 1-3) in a PCR reaction. Figure 7 shows bands of
the expected size (330 bp) in all endothelial cell samples. Negative
controls (no cDNA) lacked the 330-bp band (Fig. 7). There are no bands
in RNA samples amplified without pretreatment with reverse
transcriptase (data not shown). As a positive control, amplification of
the human P2Y2 gene from HUVEC total RNA also was
performed. A band of the expected size (483 bp) was amplified that was
confirmed to be P2Y2 (Fig. 7, inset). Together,
these results show that endothelial cells derived from multiple
vascular beds express P2X receptor channels. Confirmatory results for a
P2Y2 G protein-coupled receptor also are shown.

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Fig. 7.
P2X receptor channel mRNA is expressed in endothelial
cells. A 330-bp PCR product is found in all endothelial cell samples. A
product also was found in multiple other HUVEC samples and in HLMVEC
samples (data not shown). -Actin also was amplified to authenticate
cDNA before purinergic receptor PCR. The G protein-coupled
P2Y2 receptor gene also was amplified from 2 different
HUVEC samples (see inset). Negative controls (NC) had no PCR
product. Gels are representative of multiple amplifications. 100 bp L,
100-bp DNA ladder.
|
|
Human endothelial cells express multiple P2X receptor channel
isoforms.
Although it is clear that endothelial cell primary cultures express P2X
receptors, it is unclear from RT-PCR alone which P2XR isoforms are
expressed. To determine which isotypes are present, we performed a
large-scale cloning and sequencing analysis. Multiple colonies
containing pGEM-T plasmid with P2XR PCR insert were sequenced from each
cell type in an effort to determine whether P2X receptor channel
isoforms were expressed and which P2X receptor isotypes were expressed
most abundantly. Sequences were subjected to BLASTN analysis
(1) to determine which P2X receptor isoforms were present
in each endothelial cell primary culture. Table
3 shows the results of this differential
DNA sequencing analysis for the five endothelial cell primary cultures
analyzed. Readily apparent from this analysis, P2X4 and
P2X5 were present in all samples (Table 3). These two
isoforms were the only isoforms found in HPAEC (Table 3).
P2X1 expression was unique to HCAEC, whereas P2X7 was found only in HAEC and HCAEC (Table 3). Novel
sequences were found rarely in both primary cultures derived from
umbilical vasculature in HAEC, HUAEC, and HUVEC (Table 3).
Interestingly, these novel sequences were most identical to splice
variants or isolated regions of P2XRs. In these stretches of homology,
sequence identity was virtually 100%, although this stretch only
encompassed <50% of the PCR product. More work is being performed
with these clones that is beyond the scope of this study; similar novel
P2XR-like sequences have been identified in epithelial cell models from lung and liver (Taylor AL and Schwiebert EM, unpublished
observations). HLMVEC P2X PCR products of the expected size were
obtained but were not processed in DNA sequencing analysis. Figure
8 also shows representative sequences
from the four identified subtypes. Together, these studies show that
the P2X receptor channel isoforms P2X4 and P2X5
are expressed by human endothelial primary cultures derived from
multiple vascular beds.

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Fig. 8.
P2X receptor sequences identified in endothelial cell
primary cultures. A: P2X4 sequence alignment and
corresponding BLAST nucleotide analysis of the PCR product (Prod)
sequence. B: P2X5 sequence alignment.
C: P2X7 sequence alignment. D:
partial P2X1 sequence alignment. Only a few sequence
differences were found, and they are shown in boldface or as dashes
(for a missing base pair). As shown in Table 1, P2X4 and
P2X5 sequences were common, whereas P2X1 and
P2X7 sequences were rarely found in this degenerate
analysis.
|
|
Abundant expression of P2X4 and P2X5 is
confirmed by immunoblot analysis.
To show definitively that P2X4 and P2X5 were
expressed as membrane proteins, we performed immunoblotting of membrane
protein lysates prepared from HUVEC monolayers. Western blot analysis revealed that nascent (unglycosylated) forms (~40-50 kDa) as
well as glycosylated forms (60-80 kDa) of P2X4
receptor channel protein were found in HUVEC monolayers (Fig.
9). For P2X5, a band
corresponding to the expected size of the nascent form was
prominent at ~40-50 kDa, whereas little, if any, glycosylated
form was observed. In both P2X4 and P2X5
immunoblots, a higher molecular mass band of 120-140 kDa also was
detected. Each of these bands could be competed away by the peptide
immunogen used to generate the antibody (data not shown). No consistent
signals were observed for P2X1, P2X2, or
P2X7 (data not shown). Together, these data show that
P2X4 and P2X5 receptor channel subtypes are
expressed as membrane proteins poised to receive extracellular ATP
signals. These data agree with degenerate RT-PCR analysis showing
abundant expression of the same two P2XR subtypes.

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Fig. 9.
P2X4 and P2X5 protein is found in
membrane protein lysates from HUVEC endothelial cell primary cultures.
A tight and clear band of the expected size (40-50 kDa) was found
for the nascent polypeptide for both P2X4 and
P2X5 (reflected by arrows). Higher molecular mass forms
(especially for P2X4 at ~60-70 kDa) were found
(indicated by asterisks). Slightly higher molecular mass forms are
glycosylated forms of the P2X receptor channel (they are inhibited by
tunicamycin and eliminated by PNGase F treatment; data not shown). The
band >100 kDa (also competed away by block with peptide immunogen) may
be an SDS-resistant P2XR aggregate. Rainbow markers were used in these
blots; molecular mass was adjusted to account for dyed marker migration
being at least 10 kDa slower than that of nondyed markers.
|
|
Nucleotide agonists increase cytosolic
Ca2+ in human vascular endothelial cells.
In HAEC and HUVEC cells loaded with fura 2-AM, the effect of individual
nucleotide agonists and cocktails of P2Y- and P2X-selective agonists on
cytosolic Ca2+ was assessed. As this fura 2 analysis was
being performed in addition to the ATP release assays and receptor
expression studies, Yamamoto et al. (35) published results
showing ATP-induced increases in [Ca2+]i and
argued a role for P2X receptors in this response. Unfortunately, Yamamoto et al. used only ATP itself, which binds to and stimulates both P2Y and P2X receptor subtypes, as an agonist. Antisense blockade of P2X4 mRNA attenuated [Ca2+]i
marginally, suggesting that other P2X receptor subtypes were still
functional and/or that P2Y receptors also were mediating ATP-induced
increases in intracellular [Ca2+]i.
Basal [Ca2+]i in endothelial cell primary
cultures was 60 ± 6 nM (n = 9). Addition of a
cocktail of P2Y receptor-selective agonists (UDP, ADP, and UTP; 100 µM each) triggered an increase in [Ca2+]i
that had transient (
93 ± 14 nM for peak responses;
n = 9) and sustained (
45 ± 6 nM before
washout; n = 9) components. An example of data
calibrated to calculate free [Ca2+]i shows
that the P2Y receptor agonist cocktail increased
[Ca2+]i, which showed only limited
desensitization (Fig. 10A).
Similar addition of a cocktail of P2X receptor-selective agonists
(benzoyl-benzoyl ATP and
,
-methylene-ATP, 100 µM each) failed
to increase [Ca2+]i (n = 9;
see Fig. 10A for example). UDP (100 µM) and ADP (100 µM)
produced weak transient responses (
30 ± 5 nM) that were
inconsistent from culture to culture, suggesting that weak expression
of P2Y4, P2Y6, and/or P2Y11 may be
present in these cultures. Addition of UTP (100 µM) elicited a
response that mirrored the P2Y receptor cocktail response with
transient (
55 ± 9 nM for peak responses; n = 4) and sustained (
30 ± 5 nM before washout; n = 4) components. An example of a UTP stimulation is shown in Fig.
10B. This trace in Fig. 10B shows the
fluorescence ratio data for comparison with the calibrated data in Fig.
10A. Addition of ATP (100 µM) produced similar effects but
with still lesser magnitude (
36 ± 5 nM for peak responses and
23 ± 3 nM before washout; n = 4). Together, our interpretation, from the panel of nucleotide agonists that probe
P2Y vs. P2X receptors in a given cell type, is that P2Y receptors, most
prominently P2Y2 (P2U), increase
[Ca2+]i. Additional contributions of other
P2Y subtypes may explain the larger magnitude of the response to the
cocktail of P2Y agonists vs. UTP alone. Because ATP, but not the P2X
agonist cocktail, increased [Ca2+]i in a
phenotype that was similar to that of UTP and P2Y agonists, we conclude
that ATP increases [Ca2+]i via P2Y receptors.
In endothelial cells, P2X receptor channels may affect signal
transduction by changing the resting membrane voltage or by other
mechanisms.

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Fig. 10.
P2 receptor agonists increase cytosolic Ca2+
and ATP scavengers and receptor antagonists lower basal cytosolic
Ca2+ in HUVEC endothelial cell primary cultures.
A: a cocktail of P2Y-selective agonists (UTP, ADP, and UDP,
100 µM each) was added in 3 separate boluses to reproducibly increase
cytosolic Ca2+. In sharp contrast, a cocktail of 2 P2X-selective agonists failed to increase Ca2+. Calibrated
data for actual free cytosolic Ca2+ is shown. B:
in a separate experiment, the effect of UTP alone (100 µM) is shown
in a longer-lived time course. In this plot, fluorescence ratio of the
dual-wavelength fura 2 dye is shown. C: in a third
experiment plotting fluorescence ratio, a cocktail of ATP, ADP, and
adenosine scavengers (designed to eliminate any and all active
purinergic agonists that would be released endogenously into the
system) lowered basal cytosolic Ca2+. Suramin, a P2
receptor antagonist, was without effect at the 1 µM dose; however, 10 and 100 µM doses reversibly lowered basal cytosolic Ca2+.
This experiment was representative of 3 such time courses that produced
similar results. All agonists, antagonists, and inhibitors were
screened against all wavelengths that detect the fura 2-AM fluorescent
dye to ensure that none affected fura 2 fluorescence. For example,
pyridoxal phosphate 6-azophenyl-2',4'-disulfonic acid tetrasodium
(PPADS), another P2 receptor antagonist, is autofluorescent and cannot
be used for these experiments.
|
|
A nucleotide/nucleoside scavenger cocktail and a nonselective P2
receptor antagonist, suramin, lower basal cytosolic
Ca2+ reversibly.
To provide further evidence for autocrine/paracrine signaling by
nucleotide agonists in the endothelial cell microenvironment as well as
to tie together the ATP release and ATP receptor limbs of this
autocrine/paracrine signaling system, we assessed the effects of
antagonism of extracellular ATP signaling on cytosolic Ca2+. Figure 10C shows that a scavenger cocktail
designed to eliminate ATP, ADP, AMP, and adenosine that included 0.1 U/ml hexokinase (converts ATP to ADP), 0.1 U/ml apyrase (converts ATP
and ADP to AMP), and adenosine deaminase (converts adenosine to inert inosine) lowered basal Ca2+ significantly and in a manner
that was reversible with a wash. Moreover, in the same experiment,
suramin, an antagonist that blocks P2Y and P2X receptor subtypes,
lowered basal cytosolic Ca2+ in a dose-dependent and
reversible manner. Together with findings for exogenous nucleotide
agonists delivered to the fura 2-loaded endothelial cells, these data
suggest that endogenous ATP, released into the circulating imaging
system, is sufficient in amount to maintain basal cytosolic
Ca2+ levels in an autocrine/paracrine manner.
 |
DISCUSSION |
The strength of this study is the integrative analysis of all
limbs (release, receptors, and receptor-driven signaling) of autocrine
purinergic signaling. These studies were performed on several different
human endothelial cell primary cultures grown as monolayers. To our
knowledge, there are several novel aspects of the study. First, we have
performed real-time bioluminescence detection of released ATP on
endothelial monolayers. Burnstock's group has documented
ATP-stimulated ATP release, lipopolysaccharide (LPS)-driven ATP
release, and shear stress-induced ATP release from HUVEC primary
cultures (5, 6, 36). However, these were nonpolarized
cultures, whereas our studies were performed on human primary
endothelial cell monolayers. Second, we have assessed the sidedness of
ATP release from endothelial monolayers. Third, systematic analysis of
constitutive, Ca2+ agonist-induced, and
hypotonicity-induced ATP release as well as the temperature dependence
of basal and stimulated ATP release has not all been performed in this
integrative way. Fourth, through the use of ion transport inhibitors
such as DIDS or GdCl3 and inhibition of exocytosis with
4°C cooling, we have uncovered roles for transport and exocytosis in
ATP release from biological cells. Similar data have been shown in
epithelial and heterologous cells (7, 13, 28, 30, 34).
Although the blockade with DIDS, Gd3+, and NaCl is rapid,
ATP standard curves in the absence and presence of these substances do
not affect luciferase activity (7). We cannot rule out the
fact that they might somehow speed the buffering capacity of the cell
or enhance ecto-ATPase activity; however, the fact that DIDS or
Gd3+ (data not shown) was ineffective in blocking
Ca2+ agonist-induced ATP release suggests a different
cellular mechanism. Moreover, the fact that we were dealing with a
Cl
-containing medium in all stages of the experiment
shows that we were dealing with a luciferase that has very different
biochemical properties from that of a luciferase lyophilized and
prepared in Cl
-free solutions. Thus reversal of
hypotonicity-induced ATP release with NaCl suggests an osmotic
mechanism. Fifth, along with extensive documentation of P2Y receptor
expression on endothelial cells, we have provided degenerate RT-PCR and
biochemical evidence for expression of multiple P2X receptors
(P2X4 and P2X5) as well as P2Y2 in
positive controls. These data, together with the effect of ATP
scavengers and ATP receptor antagonists on basal Ca2+,
suggest that an autocrine and paracrine signaling loop exists in the
circulation. Sixth, we have provided new evidence that P2X receptors
may not mediate Ca2+ influx directly but, rather, may
stimulate voltage-dependent Ca2+ channels through changing
membrane potential.
At the center of this system, endothelial cells are a rich source of
released ATP under basal or stimulated conditions. Because endothelial
cells also express multiple purinergic receptors, endothelial cells
could undergo autocrine "self-regulation" or modulate the activity
of neighboring cells (circulating cells, vascular smooth muscle cells)
that also express P2X receptor channels and P2Y G protein-coupled
receptors and could transduce this extracellular ATP signal.
How might ATP, in an autocrine or paracrine manner, affect vascular
function? P2Y G protein-coupled receptors coupled to phospholipases stimulate signal transduction within most cells (21, 27). Phospholipase C-
(PLC-
) is the major signal transduction enzyme coupled to P2Y receptors (21); however, evidence also
exists for coupling to PLD and PLA2 in specific cell types
such as Madin-Darby canine kidney renal epithelial cells
(14). As such, P2Y receptors trigger phosphoinositide
signaling and increases in intracellular Ca2+. P2X receptor
channels, either by themselves or by depolarizing the membrane to open
voltage-gated Ca2+ channels, may mediate capacitive
Ca2+ entry and signaling (8, 11). Our work
suggests that P2X receptor channels may not mediate Ca2+
influx from extracellular stores, as was suggested by Yamamoto et al.
(35). The work on Ca2+ permeability has been
performed exclusively in neurons, oocytes, and heterologous cells.
Ca2+ permeability through the P2X receptor channels in
patch-clamp experiments does not suggest independently that P2XRs
increase cytosolic Ca2+ significantly. One also cannot rule
out P2X receptor stimulation of other signal transduction pathways that
are, as yet, undiscovered. One also cannot rule out the possibility
that P2X receptor channels may depolarize the membrane potential in
some cell systems to allow Ca2+ entry through
voltage-dependent Ca2+ channels that are expressed concomitantly.
Adenosine, a metabolite of ATP, has been studied extensively in the
coronary circulation and in other vascular beds (2, 12,
17). Adenosine affects vascular tone and cardiac function (2, 12, 17). It is tempting to speculate that much of the adenosine in the circulation is created via metabolism of ATP. Indeed,
ectoapyrases and ecto-ATPases are expressed by endothelial cells to
metabolize ATP (15, 36). In our data in assays performed at 37°C, the ATP release signal was dampened or decayed rapidly over
time. Our assay detected ATP released by an endothelial monolayer by
using a sufficient quantity of luciferase-luciferin detection reagent
in the medium to compete with ectoapyrases and ecto-ATPases. ATP was
consumed within seconds in the assay; that is why sharp increases
and decreases were observed when ATP release was potentiated and
inhibited. However, ecto-ATPases compete for ATP in this assay, raising
the possibility that circulating adenosine may be derived, at least in
part, from released ATP.
How might extracellular purinergic signaling be important in vascular
pathophysiology? Extracellular purinergic signaling plays a central
role in platelet function at the clotting zone (20).
Extracellular nucleotides and nucleosides may be detrimental or
therapeutic, respectively, in ischemia-reperfusion injury
(2, 10, 12, 17, 18, 25, 26). Indeed, anoxia or
ischemia of cardiac tissue may cause cell swelling that would,
in turn, augment ATP release. Extracellular purinergic signaling by ATP and adenosine has been hypothesized to play a central role in modulation of skeletal muscle and renal blood flow and glomerular filtration rate in the kidney (19). Recent evidence also
has suggested that "cross talk" may exist between purinergic
signaling and nitric oxide signaling in vascular beds (22, 24,
29). These issues require further investigation. It is possible
that inhibitors of ATP release, ATP scavengers, and/or purinergic
receptor antagonists may be of benefit in hypertension or other
circulatory syndromes and treatments.
In conclusion, these data initiate a new extracellular autocrine and
paracrine signaling cascade that may have profound implications for
vascular physiology and pathophysiology. The mechanisms of ATP release,
the regulation of ATP release, purinergic receptor signaling, and their
physiological roles in endothelial cell biology are future considerations.
 |
ACKNOWLEDGEMENTS |
This work was supported by an American Heart Association (AHA)
(Southern Research Consortium) New Investigator Grant-in-Aid to E. M. Schwiebert and an AHA (Southern Research Consortium) New
Investigator Grant-in-Aid to L. M. Schwiebert. We are grateful to
the AHA for their endorsement of this work.
 |
FOOTNOTES |
Address for reprint requests and other correspondence: E. M. Schwiebert, Dept. of Physiology and Biophysics, Univ. of Alabama at
Birmingham, MCLM 740, 1918 Univ. Blvd., Birmingham, AL 35294-0005 (E-mail: eschwiebert{at}physiology.uab.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published September 5, 2001; 10.1152/ajpcell.01387.2000
Received 22 December 2000; accepted in final form 21 August 2001.
 |
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