PGE2, Ca2+, and cAMP mediate ATP
activation of Cl
channels in pigmented ciliary
epithelial cells
Johannes C.
Fleischhauer1,
Claire H.
Mitchell1,
Kim
Peterson-Yantorno1,
Miguel
Coca-Prados2, and
Mortimer M.
Civan1,3
Departments of 1 Physiology and 3 Medicine,
University of Pennsylvania School of Medicine, Philadelphia,
Pennsylvania 19104; and 2 Department of Ophthalmology and Visual
Science, Yale University School of Medicine, New Haven, Connecticut
06510
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ABSTRACT |
Purines regulate intraocular pressure. Adenosine activates
Cl
channels of nonpigmented ciliary epithelial cells
facing the aqueous humor, enhancing secretion. Tamoxifen and ATP
synergistically activate Cl
channels of pigmented ciliary
epithelial (PE) cells facing the stroma, potentially reducing net
secretion. The actions of nucleotides alone on Cl
channel
activity of bovine PE cells were studied by electronic cell sorting,
patch clamping, and luciferin/luciferase ATP assay. Cl
channels were activated by ATP > UTP, ADP, and UDP, but not by 2-methylthio-ATP, all at 100 µM. UTP triggered ATP release. The second messengers Ca2+, prostaglandin (PG)E2,
and cAMP activated Cl
channels without enhancing effects
of 100 µM ATP. Buffering intracellular Ca2+
activity with
1,2-bis(2-aminophenoxy)ethane-N,N,N',N'- tetraacetic acid
or blocking PGE2 formation with indomethacin
inhibited ATP-triggered channel activation. The Rp stereoisomer
of 8-bromoadenosine 3',5'-cyclic monophosphothioate inhibited protein
kinase A activity but mimicked 8-bromoadenosine 3',5'-cyclic
monophosphate. We conclude that nucleotides can act at >1 P2Y
receptor to trigger a sequential cascade involving Ca2+,
PGE2, and cAMP. cAMP acts directly on Cl
channels of PE cells, increasing stromal release and potentially reducing net aqueous humor formation and intraocular pressure.
aqueous humor formation; P2Y receptors; adenosine 3',5'-cyclic
monophosphate; prostaglandins; calcium
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INTRODUCTION |
AQUEOUS HUMOR
SECRETION is a determinant of intraocular pressure, so that
reducing the secretory rate is a major strategy in treating
glaucomatous patients. The aqueous humor also delivers substrates,
oxygen, and the antioxidant ascorbate to the avascular cornea, lens,
and trabecular meshwork, removes metabolic waste products, and
facilitates immune responses (20). The bilayered ciliary
epithelium forms aqueous humor by transferring solute (and,
secondarily, water) from the stroma of the ciliary processes to the
contralateral posterior chamber of the eye. Solute is taken up from the
stroma by the pigmented ciliary epithelial (PE) layer, passes through
gap junctions to the nonpigmented ciliary epithelial (NPE) layer, and
is then released into the aqueous humor.
Several lines of evidence suggest that Cl
channel
activity limits the rate of secretion (7). Activating
Cl
channels of the NPE cells is expected to increase
secretion, whereas activating Cl
channels of the PE cells
should favor reabsorption, thereby reducing net secretion. Purines may
regulate activity of Cl
channels on both sides of the
tissue. At the aqueous surface of the epithelium,
A3-subtype adenosine agonists activate Cl
channels (5, 25). At the stromal surface, ATP and the
estrogen receptor antagonist tamoxifen synergistically activate
Cl
channels of bovine PE cells (26). The aim
of the present study was to examine the effect of ATP itself on these cells.
The strategy of the study was to focus on ATP-triggered transfer of
fluid out of PE cells with volumetric measurements and to verify the
role of Cl
channels by patch clamping. These approaches
were supplemented by luciferin/luciferase assay of ATP release in
addressing the identity of the nucleotide receptor(s) involved.
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METHODS |
Cellular model: Bovine PE cells.
We have extended our studies of an immortalized PE cell line developed
by M. Coca-Prados from a primary culture of bovine PE and characterized
by several investigators (10, 26, 34). Cells were grown in
Dulbecco's modified Eagle's medium (DMEM; no. 11965-084,
GIBCO-BRL, Grand Island, NY) with 10% fetal bovine serum (SH30071.03,
HyClone Laboratories, Logan, UT) and 50 µg/ml gentamicin (no.
15750-060, GIBCO-BRL), at 37°C in 5% CO2
(36). The medium had an osmolality of 328 mosmol/kgH2O. Cells were passaged every 6-7 days.
Volumetric measurements and analysis.
After reaching confluence, cells from a T-75 flask were harvested by
trypsinization within 3-10 days after passage (8). A
0.5-ml aliquot of the cell suspension in DMEM was added to 20 ml of
each test solution. Parallel aliquots of cells were studied on the same
day. One or two aliquots served as control, and the others were exposed
to different experimental conditions at the time of suspension. The
same amount of solvent vehicle was always added to the control and
experimental aliquots. The sequence of studying the suspensions was
varied to preclude systematic time-dependent artifacts. Cell volumes of
isosmotic suspensions were measured with a Coulter counter (model
ZBI-Channelyzer II) with a 100-µm aperture. As previously described,
the cell volume (vc) of the suspension was taken
as the peak of the distribution function. The time course of cell
shrinkage was fit to a monoexponential by nonlinear least-squares
analysis, and the probability of the null hypothesis (that any 2 sets
of observations were derived from the same population) was obtained
from the F distribution (10).
Whole cell patch-clamp recording.
Micropipettes were pulled from Corning no. 7052 glass, coated with
Sylgard, and fire polished. The resistances of the micropipettes in the
bath usually ranged from ~1.0 to 2.6 M
; successful seals displayed
gigaohm resistances. Unless otherwise stated, currents were recorded in
the ruptured-patch mode. After rupture of the membrane patch, the
series resistance was measured to be only 8.1 ± 0.7 M
and was
therefore not usually compensated; whole cell capacitance was 10.4 ± 1.2 pS. The baseline whole cell currents were 83 ± 17 pA/pF.
In a subset of experiments (n = 8), we measured whole
cell currents in the perforated-patch mode. In those experiments, we
back-filled the micropipettes with solution containing amphotericin (168 µg/ml) and filled the tips with amphotericin-free solution (1). The applied voltages were not corrected for the small junction potentials (approximately
2.8 mV; Ref. 6)
arising from the present micropipette filling and external solutions.
Data were acquired at 2-5 kHz with either an Axopatch 1D (Axon
Instruments, Foster City, CA) or a List L/M-EPC7 (Darmstadt, Germany)
patch-clamp amplifier and filtered at 500 Hz. The membrane potential
was held at
40 mV and stepped to test voltages from
100 to +80 mV
in 20-mV increments at 1-s intervals. Each step lasted 300 ms with
intervening periods of 1.7 s at the holding potential. Stimulatory
responses were measured at peak levels and inhibitory responses at the nadirs.
ATP measurements.
Bovine PE cells were grown for 4-48 h to confluence on glass
coverslips. Cells were washed in control solution and mounted on an
inverted microscope, and bath ATP levels were measured continuously by
including 2 mg/ml of luciferin/luciferase assay mixture
(33). After background levels were recorded, a solution
containing luciferin/luciferase assay mixture and either control
solution or UTP was carefully added to the cells. ATP released from
cells into the extracellular bath reacted with the luciferase and led
to the production of a photon. Light produced was captured with a ×20
objective, filtered at 520 nm, measured with a photomultiplier tube,
and recorded on-line using the Felix software suite (Photon
Technologies International, Princeton, NJ). The luminescence values
were converted to concentrations of ATP using a standard curve; UTP did
not alter luminescence in the presence of the luciferin/luciferase
assay mixture alone. The control solution contained (in mM) 105 NaCl,
4.5 KCl, 2.8 Na-HEPES, 7.2 HEPES acid, 1.3 CaCl2, 0.5 MgCl2, 5 glucose, and 75 mannitol. The pH was adjusted to
7.4 with NaOH, and the solution had an osmolality of 304-312
mosmol/kgH2O.
Chemicals.
All chemicals were reagent grade. Gramicidin D, ionomycin,
tamoxifen, ATP, UTP, ADP, UDP, GTP, 2-methylthio-ATP, 8-bromoadenosine 3',5'-cyclic monophosphate (8-BrcAMP), dibutyryl cAMP (DBcAMP), and
indomethacin were purchased from Sigma (St. Louis, MO);
4,4'-Diisothiocyanostilbene-2,2'-disulfonic acid (DIDS),
acetoxymethyl ester of
1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA-AM), and
N,N,N',N'-tetrakis(2-pyridylmethyl)-ethylenediamine (TPEN) from Molecular Probes (Eugene, OR); and
5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) from Biomol Research
Laboratories, (Plymouth Meeting, PA). 9-Phenylanthranilic
acid (DPC) was obtained from Fluka (Ronkonkoma, NY), and the inhibitory
(Rp) and stimulatory (Sp) diastereoisomers of 8-bromoadenosine
3',5'-cyclic monophosphothioate (8-Br-cAMPS) were from Biolog (La
Jolla, CA).
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RESULTS |
Effect of nucleotides on PE cell volume.
ATP triggered concentration-dependent shrinkage of PE cells at 10 and
100 µM but not at 3 µM (Fig.
1A). Test solutions in Fig.
1A contained the cation ionophore gramicidin (5 µM) to
incorporate an exit port for K+ in the plasma membrane.
Under these conditions, the observed shrinkage reflected activation of
a Cl
release pathway (8). ATP also produced
comparable shrinkage without gramicidin (Fig. 1B). As
previously reported (26), ATP did not uniformly trigger
shrinkage; no response was noted in ~20% of the present series of
volumetric measurements. Averaging the results of 11 series of
experiments (reflecting 43 experiments), ATP produced a magnitude of
shrinkage (
v
) of 4.2 ± 0.3% with a
time constant (
) of 5.2 ± 0.6 min. In controls (14 series, 50 experiments),
v
= 1.3 ± 0.3%
and
= 9.5 ± 2.5 min (in 10 series of controls with
significant shrinkage). In agreement with our previous study
(26), tamoxifen uniformly enhanced the response to ATP
(n = 10, Fig.
2B). In contrast, adenosine
has no significant effect on the volume of these PE cells with or without tamoxifen (26).

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Fig. 1.
Effects of ATP on cell volume. A: ATP
triggered a concentration-dependent shrinkage (n = 4).
In this and subsequent figures presenting volumetric data, solid curves
are least-square fits to monoexponentials. Otherwise, data points are
connected by dotted lines. Fit values for steady-state magnitude
( v ) and time constant ( ) of shrinkage:
control (2.4 ± 0.2%, 8.6 ± 2.6 min), 3 µM ATP (2.6 ± 0.2%, 12.7 ± 2.4 min), 10 µM ATP (2.7 ± 0.3%,
6.8 ± 2.1 min; P < 0.01), 100 µM ATP (5.0 ± 0.2%, 8.0 ± 1.0 min; P < 0.01).
Probabilities of the null hypothesis were obtained from the
F distribution. B:
5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) and
9-phenylanthranilic acid (DPC) inhibited ATP-triggered
shrinkage. Fit values for v and :
control (0.7 ± 0.4%, 14.8 ± 18.8 min), 100 µM ATP
(3.1 ± 0.2%, 2.5 ± 0.8 min; P < 0.01 vs.
control), 1 mM DPC + 100 µM ATP [not significant (NS);
P < 0.01 vs. ATP alone], 100 µM NPPB + 100 µM ATP (1.2 ± 1.0%, 16.2 ± 15.5 min; P < 0.01 vs. ATP alone). Iso, isotonic control; Gram, gramicidin
D.
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Fig. 2.
Relative effects of nucleotides on cell volume.
A: fit values for v and for UTP, UDP,
and ATP (n = 4): control (1.9 ± 0.6%, 23.6 ± 15.5 min), 100 µM ATP (4.1 ± 0.2%, 4.8 ± 1.1 min;
P < 0.01), 100 µM UTP (3.4 ± 1.2%, 31.6 ± 20.2 min; P < 0.01 vs. ATP, P > 0.05 vs. control), 100 µM UDP (2.7 ± 0.3%, 11.0 ± 2.8 min; P < 0.01 vs. both ATP and control). B:
fit values for v and for ADP, tamoxifen, and ATP
(n = 4): control (1.5 ± 0.2%, 1.0 ± 1.3 min), 100 µM ADP (2.4 ± 0.5%, 7.3 ± 1.9 min;
P < 0.05 vs. control, P < 0.01 vs.
ATP), 100 µM ATP (5.6 ± 0.5%, 7.3 ± 1.9 min;
P < 0.01), 10 µM tamoxifen + 100 µM ATP (not
fit by single exponential, but data points significantly different from
control, ATP, and ADP at P = 0.01).
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The ATP-triggered shrinkage was inhibited by Cl
channel
blockers. When tested in parallel aliquots of cell suspensions, NPPB (100 µM) and DPC (1 mM) were similarly effective in blocking
shrinkage (Fig. 1B).
UTP and UDP (Fig. 2A) and ADP (Fig. 2B) also
triggered shrinkages. These effects were smaller than the ATP-triggered
shrinkage, precluding a definitive ranking of the effects of ADP, UDP,
and UTP.
Effect of nucleotides on PE whole cell currents.
ATP altered whole cell currents in approximately one-third of the
bovine PE cells (see Table 2). Two different electrophysiological effects were noted, sometimes in the same cells (Fig.
3). The stimulatory effect is exemplified
by the increase in whole cell currents beginning ~1 min after
initiation of perfusion with 10 µM ATP (Fig. 3A). Raising
the external ATP concentration to 100 µM then triggered the second
characteristic effect, a small inhibition of outward current at +80 and
+60 mV within ~40s. The later rate of increase in whole cell currents
was not detectably altered by this increase in ATP concentration.
Application of the Cl
channel blocker NPPB (100 µM)
subsequently inhibited the currents by ~70%. The time courses of
ATP-difference currents after step changes in voltage are presented in
Fig. 3B, and the current-voltage relationships of the ATP-
and NPPB-difference currents are presented in Fig. 3C.
Slight inactivation was noted at highly depolarizing potentials (Fig.
3B); the magnitude of depolarization-induced inactivation
displayed by Cl
channels of many cells depends on the
free intracellular Mg2+ concentration and other
unidentified factors (29). The current-voltage relationship was outwardly rectifying (Fig. 3C).

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Fig. 3.
Effects of ATP on whole cell currents. A: time
course of currents before, during, and after perfusion with ATP and
NPPB initiated at the times indicated by the vertical lines.
B: responses of 100 µM ATP-difference currents to step
changes in voltage from the holding potential of 40 mV. In Figs. 4,
5, and 8, difference currents were calculated by subtracting means of
3-5 sets of measurements just before change in perfusion from a
similar set of records at the peak stimulation or maximal inhibition.
C: current-voltage relationships for the absolute baseline
( ) and activated ( ) currents and 100 µM ATP ( )- and 100 µM NPPB
( )-difference currents.
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The reversal potential for the ATP-activated currents (Fig.
3C) was
26.8 ± 1.8 mV. Taking into account a
junction potential of approximately
2.8 mV estimated for similar
filling and bath solutions (5), the corrected reversal
potential was
29.6 mV. When this value and the known anionic
concentrations inside and outside the cell (Table
1) are inserted in the Goldman equation, the relative permeability of aspartate/Cl
of the
ATP-activated anion channels can be estimated (5) to be
~0.14.
Whole cell responses to UTP are illustrated in Fig.
4. As shown in Fig. 4A, 10 µM UTP produced small initial inhibitions of outward currents
followed by stimulation of both outward and inward currents. The
UTP-difference currents displayed the same characteristics observed
with ATP-difference currents (Fig. 3): slight inactivation at highly
depolarizing potentials (Fig. 4B) and outward rectification with a comparable uncorrected reversal potential (
33.4 ± 1.2 mV; Fig. 4C). As for the ATP experiments, the magnitudes of
the responses to UTP are displayed in Fig.
5 and the frequency of the responses is
given in Table 2. ADP and UDP triggered
similar changes, but the magnitudes were smaller (Fig. 5, Table 2).

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Fig. 4.
Effects of UTP on whole cell currents. The time courses
during changes in perfusion (A), the responses of the
UTP-difference currents to step changes in voltage (B), and
the current-voltage relationships of the baseline ( ),
activated ( ), and difference ( )
currents (C) are presented. Iso Ctrl in A
indicates onset of washouts with isotonic control solution.
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Fig. 5.
Relative effects of nucleotides on whole cell currents.
The percent activations and percent blocks were calculated as mean ± SE values for the cells displaying responses. Approximately
25-45% of the cells responded to the nucleotides (see text for
discussion). Nucleotide concentrations are in µM.
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To summarize the electrophysiological results obtained with all
nucleotides, the means ± SE of stimulatory and inhibitory responses to 10 and 100 µM concentrations are presented in Fig. 5 and
the frequency with which these effects were observed is presented in
Table 2. No response to ATP was seen after 1 or 3 µM ATP. At 10 and
100 µM concentrations, stimulations were seen in ~25-45% of
the cells studied. There was considerable variance in response, but the
changes appeared generally larger after ATP than after UTP, ADP, and
UDP (Fig. 5). Block was observed in a similar fraction of the
cells studied, with UTP producing a larger effect than did the other
three nucleotides (Fig. 5). For all nucleotides studied, the nonzero
reversal potential, outward rectification, inactivation at highly
depolarizing potentials, and sensitivity to NPPB established that the
stimulated currents reflected activation of Cl
channels.
Because <50% of the cells responded to nucleotides, we wondered
whether dialysis of important components out of the cell could have
limited the frequency of response. However, the response rate to ATP
and UTP was not significantly enhanced by using the perforated-patch mode of whole cell recording. Stimulatory and inhibitory responses were observed in 5 and 3 of 11 cells, respectively.
Second messenger cascade assayed by shrinkage.
ATP increases intracellular Ca2+ activity of bovine PE
cells (26). Raising intracellular Ca2+ levels
by adding ionomycin triggered PE cell shrinkage similar to that
produced by ATP, and the two effects were not additive (Fig.
6A).

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Fig. 6.
Effects of Ca2+, prostaglandin
(PG)E2, and cAMP on cell volume. A: fit values
for v and for the responses to raising
intracellular Ca2+ with ionomycin (n = 3):
control (NS), 2 µM ionomycin (4.6 ± 0.4%, 3.3 ± 1.1 min;
P < 0.01), 100 µM ATP (4.9 ± 0.4%, 5.3 ± 1.3 min; P < 0.01), 2 µM ionomycin + 100 µM ATP (4.5 ± 0.5%, 3.5 ± 1.3 min; P < 0.01). The data obtained with ionomycin, ATP, or both together were not
significantly different from one another. B: fit values for
v and for the responses to
PGE2 (n = 3): Control (1.6 ± 0.2%,
6.7 ± 2.9 min), 100 µM ATP (4.5 ± 0.2%, 5.2 ± 0.8 min; P < 0.01), 10 µM PGE2 (4.6 ± 0.4%, 6.1 ± 1.7 min; P < 0.01), 10 µM
PGE2 + 100 µM ATP (6.5 ± 0.8%, 8.3 ± 2.6 min; P < 0.01 vs. control, P > 0.05 vs. PGE2 alone). C: 100 µM ATP and 500 µM 8-bromoadenosine 3',5'-cyclic monophosphate (8-BrcAMP) produced
comparable degrees of shrinkage (n = 4). Fit values for
v and : control (1.17 ± 0.09%,
1.4 ± 0.7 min), 8-BrcAMP (3.8 ± 0.4%, 4.0 ± 1.6 min;
P < 0.05), 100 µM ATP (4.37 ± 0.07%, 5.0 ± 0.3 min; P < 0.01), 8-BrcAMP + ATP (4.1 ± 0.1%, 2.4 ± 0.3 min; P < 0.01). Adding
8-BrcAMP clearly increased the speed but not the magnitude of the
shrinkage response.
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An increase in intracellular Ca2+ activity can in turn
activate phospholipase A2 (4), enhancing
formation of prostaglandin (PG)E2. Indeed, 10 µM
PGE2 produced effects similar to those of Ca2+,
replicating the actions of 100 µM ATP. Application of ATP together with PGE2 did not significantly enhance the action of
PGE2 alone (Fig. 6B). The effects of ionomycin
and PGE2 were also not additive (Fig.
7C).

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Fig. 7.
Effects of buffering intracellular Ca2+ and
blocking PGE2 on ATP-triggered shrinkage and nonadditivity
of cAMP and PGE2 in triggering shrinkage. A:
buffering intracellular Ca2+ with acetoxymethyl ester
of 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid
(BAPTA-AM), but not
N,N,N',N'-tetrakis(2-pyridylmethyl)-ethylenediamine
(TPEN) blocked ATP-induced shrinkage (n = 3). Fit
values for v and : control (3.8 ± 1.8%, 20.6 ± 16.5 min), 100 µM ATP (4.8 ± 0.1%,
1.6 ± 0.2 min; P < 0.01), 20 µM BAPTA-AM + 100 µM ATP (1.9 ± 0.3%, 6.4 ± 2.5 min;
P > 0.05 vs. control, P < 0.01 vs.
ATP), 20 µM TPEN + 100 µM ATP (4.3 ± 1.3%, 1.3 ± 0.4 min; P < 0.01 vs. control and BAPTA,
P > 0.05 vs. ATP alone). B: cyclooxygenase
inhibition with indomethacin (Indo) also blocked ATP-triggered
shrinkage (n = 5). Fit values for
v and : control (1.9 ± 0.2%,
8.8 ± 1.9 min), 100 µM ATP (3.8 ± 0.6%, 7.1 ± 3.3 min; P < 0.01), 1 µM Indo + 100 µM ATP (NS,
P < 0.05 vs. ATP), 100 µM Indo + 100 µM ATP
(2.1 ± 0.3%, 6.2 ± 2.6 min; P < 0.01 vs.
ATP, P > 0.05 vs. both control and 1 µM Indo + ATP). C: adding cAMP did not enhance
PGE2-triggered shrinkage (n = 3). Fit
values for v and : control (NS), 10 µM PGE2 (4.3 ± 0.5%, 4.6 ± 1.7 min;
P < 0.01), 0.5 mM cAMP (3.9 ± 0.3%, 4.4 ± 1.2 min; P < 0.01), cAMP + PGE2
(3.6 ± 0.2%, 2.6 ± 0.7 min; P < 0.01).
PGE2, cAMP, and both agents together triggered similar
shrinkages (P > 0.05).
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Production and release of PGE2 would in turn be expected to
trigger cAMP formation by occupancy of EP2 receptors known
to be functionally expressed in bovine PE cells (2), so we
tested whether cAMP could replicate the responses to ATP. 8-BrcAMP (500 µM) and 100 µM ATP produced similar degrees of shrinkage (Fig. 6C).
The foregoing data suggested that ATP might activate Cl
channels, and thereby shrinkage, by stimulating increases in
Ca2+, PGE2, and cAMP, but it was unclear
whether these second messengers were acting in parallel or in tandem,
as recently found in Madin-Darby canine kidney (MDCK) cells
(30). We addressed this issue by attempting to block
increases 1) in Ca2+ (with the Ca2+
buffer BAPTA), 2) in PGE2 (with the
cyclooxygenase inhibitor indomethacin), and 3) in
cAMP-activated protein kinase activity (with the inhibitory Rp
stereoisomer of 8-BrcAMPS).
Preincubation with BAPTA-AM for 1 h to buffer intracellular
Ca2+ (18) exerted no direct effect on cell
volume but completely abolished ATP-triggered shrinkage (Fig.
7A). In contrast, similar treatment with TPEN, a
chelator of heavy metals other than Ca2+ and
Mg2+, did not alter the subsequent response to ATP (Fig.
7A). Even in the presence of BAPTA-AM, both 10 µM
PGE2 and 500 µM 8-BrcAMP triggered shrinkage (data not
shown; n = 4; P < 0.01).
We then examined whether blocking the cyclooxygenase pathway of
arachidonic acid metabolism would also affect the response to ATP. As
illustrated by Fig. 7B, both 1 and 100 µM indomethacin inhibited the ATP-triggered shrinkage. The indomethacin was not acting
simply as a channel blocker. Suspending cells in solution containing 10 µM PGE2 together with 100 µM indomethacin and 100 µM
ATP overcame the indomethacin inhibition (data not shown;
n = 5; P < 0.01 compared with parallel
aliquots suspended in indomethacin and ATP alone).
In our third approach, we focused on cAMP in trying to interrupt the
signaling pathways initiated by ATP. In a preliminary test, we found
that the specific form of cAMP used to activate shrinkage was not
critical. Effects similar to those elicited by 8-BrcAMP, albeit faster,
were triggered by the more permeable analog DBcAMP (500 µM) (Fig.
8A). Also, simultaneous
addition of the two analogs produced the same response as did
DBcAMP alone (Fig. 8A), so that 500 µM is maximally
effective. Because cAMP commonly modifies channel activity through
protein kinase A (PKA), we applied the inhibitory diastereoisomer
(Rp) of 8-BrcAMPS. Unexpectedly, Rp-8-BrcAMPS produced cell
shrinkage comparable to that triggered by ATP, and adding Rp-8-BrcAMPS
and ATP together had the same effect as adding ATP alone (Fig.
8B). Parallel additions of the inhibitory diastereoisomer,
the stimulatory isomer Sp-8-BrcAMPS, and 8-BrcAMPS in the same series
of experiments elicited similar shrinkages (Fig. 8C).

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Fig. 8.
Effects of cAMP on cell volume. A: 8-BrcAMP
and dibutyryl cAMP (DBcAMP) triggered shrinkages of the same magnitude,
with the more permeable dibutyryl form triggering a faster response
(n = 4). The 500 µM concentration was evidently
saturating, because no additive effect was observed by adding the 2 forms of cAMP. Fit values for v and :
Control (NS), 500 µM 8-BrcAMP (4.2 ± 0.4%, 12.2 ± 0.7 min; P < 0.01), 500 µM DBcAMP (4.3 ± 0.3%,
2.2 ± 0.7 min; P < 0.01), 8-BrcAMP + DBcAMP
(4.3 ± 0.3%, 4.3 ± 1.2 min; P < 0.05 vs.
control, P > 0.05 vs. DBcAMP). B: the Rp
diastereoisomer of adenosine 3',5'-cyclic monophosphothioate (cAMPS)
inhibits PKA activity but it stimulated shrinkage (n = 4). Fit values for v and : control
(NS), 100 µM ATP (2.9 ± 0.4%, 7.2 ± 2.7 min;
P < 0.01), 100 µM Rp (1.9 ± 0.1%, 1.3 ± 0.6 min; P < 0.01), Rp + ATP (3.0 ± 0.6%,
12.8 ± 5.5 min; P < 0.01). The addition of
Rp-cAMPS reduced the speed and slightly increased the magnitude of the
shrinkage response to ATP. C: 8-BrcAMP, the inhibitory
diastereoisomer Rp, and the stimulatory diastereoisomer Sp-cAMPS
produced similar degrees of cell shrinkage (n = 6). Fit
values for v and : control (1.4 ± 0.2%, 5.3 ± 2.0 min), 500 µM 8-BrcAMP (4.0 ± 0.4%,
7.1 ± 2.0 min; P < 0.01), 100 µM Rp (3.2 ± 0.2%, 3.2 ± 0.7 min; P < 0.01), 100 µM Sp
(4.4 ± 0.4%, 6.0 ± 1.8 min; P < 0.01).
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Second messenger cascade assayed by whole cell currents.
Perfusion with 10 µM PGE2 activated Cl
currents in five of five cells by 88 ± 27% (+80 mV;
P < 0.01). In the experiment shown in Fig.
9A, perfusion with either 1 mM
DPC or 100 µM NPPB reversibly inhibited the activated currents and
isotonic washout was associated with decay of the currents. The time
courses of the PGE2-difference currents after step changes
in voltage are displayed in Fig. 9B, and the current-voltage
relationships are presented in Fig. 9C. With 100 µM
indomethacin present to block PGE2 production, 100 µM ATP
produced no stimulatory response (n = 3), and,
conversely in the presence of 100 µM ATP, indomethacin reduced
current by 31 ± 8% (+80 mV; n = 4;
P < 0.05); one-half of these cells displayed a
stimulatory response to the ATP pretreatment.

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Fig. 9.
Effects of PGE2 on whole cell currents. The
time courses during changes in perfusion (A), the responses
of the PGE2-difference currents to step changes in voltage
(B), and the current-voltage relationships of the baseline
( ), PGE2-difference ( ),
NPPB-difference ( ), and DPC-difference
( ) currents (C) are presented.
|
|
The effects of 8-BrcAMP on Cl
channels were also examined
with whole cell patch clamping. In the experiment shown in Fig. 10A, 100 µM 8-BrcAMP
increased outward and inward currents. Raising the concentration to 500 µM further stimulated the currents, producing a cumulative
stimulation of ~80% at +80 mV. NPPB subsequently inhibited the
activated currents reversibly by >90%. The time courses of the
difference currents after step changes in voltage and the
current-voltage relationships are displayed in Fig. 10, B
and C, respectively. At concentrations of 100 and 500 µM,
8-BrcAMP stimulated currents in four of nine experiments without
exerting any blocking effect (Fig. 5).

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Fig. 10.
Effects of 8-BrcAMP on whole cell currents. The time
courses of the absolute currents (A), the responses of the
8-BrcAMP-difference currents to step changes in voltage (B),
and the current-voltage relationships of the baseline
( , bottom) and activated ( , top)
currents and the 8-BrcAMP ( )- and NPPB
( )-difference currents (C) are presented.
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|
Similar to the ATP- and UTP-triggered responses, the PGE2
(Fig. 9B)- and 8-BrcAMP (Fig. 10B)-difference
currents displayed slight inactivation at highly depolarizing
potentials (Fig. 8B) and outward rectification (Figs.
9C, 10C) with similar uncorrected reversal
potentials (
31.2 ± 3.3 mV in Fig. 9C and
32.7 ± 2.0 mV in Fig. 10C).
P2 receptors.
If a single P2 receptor modulated Cl
channel activity in
PE cells, the data of Figs. 2 and 5 might point to P2Y11
(see DISCUSSION). Because the bovine sequence for this
receptor is unknown, we conducted a functional test for its presence.
The EC50 values of 2-methylthio-ATP at turkey
P2Y1 and human P2Y11 receptors are 6 nM and 50 µM, respectively (17). However, neither 100 nM nor 100 µM concentrations of 2-methylthio-ATP triggered the shrinkage
produced by 100 µM ATP (n = 4; data not shown).
ATP release by PE cells.
Multiple complexities limit the identification of P2 receptors
affecting cell functions (see DISCUSSION). One such
complexity is potential UTP-triggered ATP release, which has been
reported in other cells (9, 23, 30). This possibility was
examined by perfusing PE cells with 100 µM UTP. As illustrated by
Fig. 11, the mechanical perturbation
associated with solution change triggered a transient release in ATP
even in the control solution. However, UTP triggered a sustained
release of ATP. At the peak, 7 min after solution change, ATP levels in
control solution were 41.1 ± 9.4 nM whereas those in UTP were
206.8 ± 73.5 nM (P < 0.05; n = 10). The subsequent decline in detected external ATP concentration likely reflects the activity of ecto-ATPase/apyrase and the utilization of substrate (23, 24).

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Fig. 11.
UTP triggers release of ATP. After ~4 min in control
solution, the bath was changed to a control solution containing
luciferin/luciferase. After a further ~10 min, the solution was
replaced with UTP containing luciferin/luciferase, which triggered a
clear increase in extracellular ATP concentration. The vertical lines
at ~5 and ~15 min represent room light reaching the chamber during
solution changes. Solid data lines represent mean responses, and dotted
lines show associated SE (n = 10).
|
|
 |
DISCUSSION |
Initial observation.
External nucleotides activated Cl
channels and triggered
release of intracellular solute and water from bovine PE cells. On average, ATP produced larger stimulations of Cl
currents
than did UTP, ADP, and UDP at the same concentration (Fig. 5). However,
the observation that more than half the cells did not respond and the
ranges in magnitude of the responses made it difficult to rank the
nucleotide responses from the patch-clamp data. In this respect,
electronic cell sorting provided a more feasible approach, directly
measuring the parameter of central interest (the transfer of fluid out
of the cells) and sampling 50,000 cells or more for each data point. By
this measure and at a concentration of 100 µM, the ranking is
ATP > UTP, ADP, UDP (Fig. 2).
To place the nonuniformity of response to ATP in perspective, we note
that previous investigators reported a nonuniform expression for other
channels in studies of PE cells. For example, only ~15% of fresh and
cultured bovine PE cells exhibit T-type Ca2+ channels
(16) and 22% of rabbit PE cells display L-type
Ca2+ channels (13). Fain and Farahbakhsh
(12) found voltage-gated Na+ channels in
~25% of the primary cultures of rabbit PE cells they studied.
Stelling and Jacob (32) reported that carbachol increased K+ conductances in only ~30% and Cl
conductance in 49% of the freshly dissected bovine PE cells they studied. Mitchell et al. (27) also found that GTP
S
activated two-thirds of large-conductance Cl
channels in
patches taken from freshly dissected bovine PE cells ultimately shown
to possess such a channel. Given the syncytial nature of the bilayered
ciliary epithelium (7), activation of the Cl
channels in 25-45% of the PE cells should provide a
physiologically significant pathway for release of solute to the stroma.
Nucleotide receptors.
Identification of functionally important receptor(s) is generally even
more complex for P2 than for P1 (adenosine) receptors because
(11, 21) 1) specific agonists and antagonists
are not available; 2) the ectoenzymes apyrase, ecto-ATPase,
ecto-ADPase, 5'-nucleotidase, and ectonucleoside diphosphokinase not
only metabolize adenine nucleotides to adenosine but also interconvert
purine and pyrimidine nucleotides; 3) cells frequently
possess multiple P2 and P1 receptors, which can exert opposing effects;
and 4) the final functional effects can be highly dependent
on the specific cell studied because of synergistic interactions of the
second messenger cascades. An additional caveat is illustrated by Fig. 11. Consistent with observations in other cells (9, 23,
30), application of one triphosphate nucleotide can trigger
release of another nucleotide from intracellular stores. The
concentrations of ATP measured directly in this study, although
probably an underestimate, should be sufficient to stimulate the
P2Y2 receptor with an EC50 of 200 nM
(22).
Despite these caveats, two general conclusions can be drawn from the
data. First, the dominant functional receptors must be P2Y
metabolotropic, rather than P2X ionotropic receptors. Otherwise, ATP
activation of P2X cation-nonselective channels would have triggered
influx of cation, leading to swelling of the PE cells, contrary to
observation. Second, it is unlikely that the nucleotide-triggered shrinkage was mediated by occupancy of a single population of P2Y
receptors. Of the six cloned functional P2Y receptors
(P2Y1, P2Y2, P2Y4,
P2Y6, P2Y11, and P2Y12; Refs.
11 and 14), only P2Y11 might conform to the
nucleotide ranking displayed by Fig. 2. Our inability to detect a
response to the P2Y11 agonist 2-methylthio-ATP at 100 µM
suggests that this receptor is not playing a dominant role in mediating
the responses to ATP. We conclude that the transport effects of the
nucleotides likely reflect occupancy of multiple P2 receptors of the PE
cells. This conclusion is consistent with the reported detection of at
least two different P2Y receptors in bovine PE cells (31,
34).
Second messengers.
The present data suggest a plausible signaling cascade. We have found
that elevation of intracellular Ca2+ activity and separate
application of PGE2 and 8-BrcAMP all mimic ATP in reducing
cell volume, and the effects of these agents are not additive with
those of ATP. In principle, all three second messengers could act in
parallel to mediate the ATP-triggered activation of Cl
channels (Fig. 12A).
Alternatively, ATP could trigger a cascade involving the sequential
activation of the three second messengers (Fig. 12B). More
complex pathways involving both parallel and series activations are
also possible.

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Fig. 12.
Possible pathways of intracellular signaling mediating
ATP-triggered Cl channel (gCl ) activation.
The data indicate that Ca2+, PGE2, and cAMP
participate in mediating activation. In a purely parallel model
(A), inhibition of cyclooxgenase with Indo should only
partly inhibit activation, leaving the cAMP- and
Ca2+-mediated paths operative. In a purely series model
(B), block of PGE2 production should completely
inhibit ATP-triggered Cl -channel activation, consistent
with observation. Combinations of the 2 models may also account for the
observations.
|
|
The current observations indicate that the purely parallel model of
Fig. 10A cannot be correct. If ATP were to act independently through Ca2+, PGE2, and cAMP, blocking either
the rise in intracellular Ca2+ or the formation of
PGE2 should only partially inhibit the transport effects of
ATP. This prediction is contrary to the observation that ATP-triggered
shrinkage was completely blocked by either buffering intracellular
Ca2+ levels or preventing PGE2 formation (Fig.
7, A and B).
In contrast, the series hypothesis (Fig. 12B) is consistent
not only with the current results but also with a series cascade recently identified in MDCK cells (30). ATP-triggered
elevation in cell Ca2+ activates phospholipase
A2, stimulating cyclooxygenase-catalyzed PG
synthesis and release. PGE2 occupancy of
EP2 receptors is known to stimulate adenylyl
cyclase-mediated cAMP production specifically by bovine PE cells
(2). In turn, cAMP activates Cl
channels
(Fig. 10), permitting release of K+ and Cl
through parallel ionic channels and, secondarily, release of water
(Figs. 6C, 7C, 8A, and
8C). The data of Fig. 8, B and
C, obtained with the PKA-inhibitory analog of 8-BrcAMP,
Rp-8-BrcAMPS, suggest that cAMP acts directly on the PE
Cl
channel, not through activation of PKA. This concept
is consistent with reports that Rp-cAMPS mimics cAMP-triggered
activation of hyperpolarization-activated currents (3, 15)
and Rp-cGMPS activates the photoreceptor (but not the olfactory) cyclic
nucleotide-gated channel (19).
Potential physiological implications.
The Cl
channels of PE and NPE cells differ in their
unitary conductances (27, 37) and pharmacological profiles
(28, 35). In principle, activation of the NPE
Cl
channels facing the aqueous humor is expected to
increase net secretion, whereas activation of PE channels facing the
stroma is expected to reduce net secretion. Release of ATP from both NPE and PE cells (24) provides the basis for autocrine
regulation of secretion. Ectoenzyme metabolism of ATP provides a source
of adenosine, which activates NPE Cl
channels at ~3
µM concentration (5, 25). The results of the present
work demonstrate that ATP itself activates PE Cl
channels
at ~10 µM concentration (Figs. 1 and 3). Which effect predominates
should depend on local purine concentrations, on ectoenzyme activities
and receptor densities at both surfaces, and possibly on activities of
additional modulators. In particular, occupancy of plasma membrane
estrogen receptors is thought to trigger synergistic enhancement of ATP
activation of PE cell Cl
channels (35). The
basis for the synergism between the estrogen-triggered cascade and the
Ca2+-PGE2-cAMP cascade triggered by ATP remains
to be determined.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Martin Pring for suggestions concerning statistical
analysis of the data and Dr. Jeffrey W. Karpen for helpful information.
 |
FOOTNOTES |
This work was supported in part by National Eye Institute Grants
EY-12213, EY-08343, and EY-01583 (for core facilities). J. C. Fleischhauer received support from Swiss National Science
Foundation Fellowship No. 1037.
Address for reprint requests and other correspondence:
M. M. Civan, Dept. of Physiology, Univ. of Pennsylvania, 3700 Hamilton Walk, Philadelphia, PA 19104-6085.
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.
Received 26 February 2001; accepted in final form 26 June 2001.
 |
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