Effect of coupling on volume-regulatory response of ciliary
epithelial cells suggests mechanism for secretion
V. E.
Walker,
J. W.
Stelling,
H. E.
Miley, and
T. J. C.
Jacob
School of Molecular and Medical Biosciences, University of
Wales, Cardiff CF1 3US, United Kingdom
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ABSTRACT |
The ciliary epithelium of the eye secretes the aqueous humor. It
is a double epithelium arranged so that the apical surfaces of the
nonpigmented ciliary epithelial (NPCE) and pigmented ciliary epithelial
(PCE) cells face each other and the basolateral membranes face the
inside of the eye and the blood, respectively. We have investigated the
volume responses of both single cells and coupled pairs from this
tissue to osmotic challenge. Both NPCE and PCE cells undergo regulatory
volume increase (RVI) and decrease (RVD) when exposed to hyper- and
hyposmotic solution, respectively. In hyposmotic solution single cells
swell and return to their original volumes within ~3 min. In
nonpigmented cells RVD could be inhibited by blockers of
volume-activated Cl
channels [tamoxifen (100%) > quinidine (87%) > DIDS (84%) > 5-nitro-2-(3-phenylpropylamino)benzoic acid (80%) > SITS
(58%)] and K+ channels
[Ba2+
(31%)]. However, in PCE cells these inhibitors and
additionally tetraethylammonium and
Gd3+ were without effect. Only
bumetanide, an inhibitor of
Na+-K+-2Cl
cotransport, was found to have any effect on RVD in PCE cells. NPCE-PCE
cell coupled pairs also underwent RVD, but with altered kinetics. The
onset of RVD of the PCE cell in a pair occurred
80 s before that of
the NPCE cell, and the peak swell was reduced. This is consistent with
fluid movement from the PCE to the NPCE cell. The effect of the
volume-activated Cl
channel
inhibitor tamoxifen was to eliminate this difference in the times of
onset of RVD in coupled cell pairs and to inhibit RVD in both the NPCE
and PCE cells partially. On the basis of these observations we suggest
that fluid is transferred from the PCE to the NPCE cell in coupled
pairs during cell swelling and the subsequent RVD. Furthermore, we
speculate that reciprocal RVI-RVD could underlie aqueous humor secretion.
ciliary epithelium; secretion; fluid transport; volume regulation; ion channels
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INTRODUCTION |
IT HAS BEEN SUGGESTED (1) that the same mechanisms that
underlie volume regulation may also subserve fluid secretion. Of the
mechanisms that are involved in volume regulation, those involved in
regulatory volume decrease (RVD), e.g., volume-activated
K+ and
Cl
channels, result in
solute efflux, whereas those involved in regulatory volume increase
(RVI), e.g., the
Na+-K+-2Cl
cotransporter, result in solute influx (see Ref. 5 for a review). If
these mechanisms were sited on opposite membranes in epithelia, then
this arrangement would provide a mechanism for the
vectorial transport of solute, and hence water. With the uptake
mechanisms situated on the basolateral (blood) side and the efflux or
secretory mechanisms situated on the apical (mucosal) membrane, fluid
secretion would be achieved; conversely, with uptake mechanisms
situated on the apical membrane and efflux mechanisms on the
basolateral membrane, fluid absorption would be accomplished.
In the ciliary epithelium of the eye, which is, uniquely, a bilayered
epithelium with the two epithelial cell layers apposed at their apical
membranes, the uptake and efflux mechanisms could be differentially
segregated into different cells; if this were the case, a vectorial
movement of fluid could be achieved. There is evidence
that the two cell types have different properties with respect to their
complement of ion channels (7, 10, 17, 18). The apical membranes of the
two cells are effectively short circuited by communicating gap
junctions (3, 11, 13), allowing solute and water movement between the
two cell layers in the ciliary epithelium. Therefore, following the
suggestion of Civan et al. (1), we decided to investigate volume
regulation in these cells and look for evidence of vectorial fluid movement.
 |
METHODS |
Dissection and cell culture.
Tips of ciliary processes were dissected from bovine eyes (obtained
from a local abattoir) within 1-3 h post mortem. Cells were
isolated with 0.25% trypsin-0.02% EDTA in PBS at 35°C for 20 min
in a shaking water bath, a method previously described by Jacob (6).
The incubation mixture was then triturated with a Pasteur pipette to
break up clumps and allowed to settle, and the supernatant was
decanted, spun at 1,000 rpm, resuspended in HEPES-buffered culture
medium E199 (Sigma, Poole, UK) with 10% FCS (Sigma) twice, and finally
plated on glass coverslips. Cells were incubated overnight in E199 plus
10% FCS in a humidified incubator (Gallenkamp, Loughborough, UK) at
37°C.
Electrical recording.
Cells attached to coverslips were transferred to the recording chamber
on the stage of an inverted fluorescence microscope (Leitz DM1L; Leica,
Milton Keynes, UK). Cell pairs of pigmented ciliary epithelial (PCE)
and nonpigmented ciliary epithelial (NPCE) cells were selected and
patch clamped (see Ref. 14 for a detailed description) with electrodes
(4-8 M
; pulled on a PB-7 electrode puller; Narashige, Tokyo,
Japan) containing intracellular buffer (see
Solutions) and were bathed in bath
solution A (see
Solutions). The cells
were whole cell patch-clamped [using either a Dagan 8900 or List
(Darmstadt, Germany) EPC-7 amplifier] as described previously
(14) and recorded in current clamp.
Digital image recording and image analysis.
Images of single cells and coupled pairs of cells were recorded every
20 s with a charge-coupled device (CCD) camera (EDC-1000HR; Electrim)
mounted on an inverted microscope (Leitz DM-1L; ×63 oil immersion
Leitz Fluorescenz 1.3-numerical-aperture objective). These were then
processed with image analysis software (Quantimet 500; Leica), and the
relative volume was computed from the ratio of the mean cell diameters cubed.
Solutions.
Pipette solution, used for fluorescent dye coupling experiments,
contained 1 mM Lucifer yellow and (in mM) 6 NaCl, 56 KCl, 84 potassium
gluconate, 1.1 EGTA, 10 HEPES, 2 MgCl2,
10
5
CaCl2, and 20 sucrose and was
adjusted to pH 7.25 with 1 M NaOH.
For experiments at 22°C the solution was composed of (in mM) 125 NaCl, 5 KCl, 10 HEPES, 10 NaHCO3,
0.5 MgCl2, 2 CaCl2, 5 glucose, and 20 sucrose
and was adjusted to pH 7.4 with 1 M NaOH (solution A). For experiments at 35°C the solution was
composed of (in mM) 90 NaCl, 4.4 KCl, 26 NaHCO3, 10 HEPES, 2 CaCl2, 0.5 MgCl2, 5 glucose, and 20 sucrose
and was adjusted to pH 7.4 with 1 M NaOH (solution B). Solutions were made hypotonic by the
addition of 50% distilled water and were made hypertonic by the
addition of 100 mM sucrose.
All chemicals were obtained from Sigma. The osmolarities of all the
solutions were measured by depression of the freezing point using an
osmometer (Osmomat 30; Gonotec) and were within ±10 mosmol/l of the
theoretical values: isotonic, hypotonic, and hypertonic solution
osmolarities were 300, 150, and 400 mosmol/l, respectively.
Statistics.
Statistical analysis of the data was achieved with Student's
two-tailed t-test.
 |
RESULTS |
RVD in single cells.
Using a CCD camera mounted on an inverted microscope, we recorded
digital images of both single and coupled pairs of cells. Figure
1 shows a coupled pair of
cells in isotonic solution (Fig. 1A), 140 s after exposure to 50%
hypotonic solution (Fig. 1B), and
600 s after exposure to hypotonic solution (Fig.
1C). Individually, both cells of the
ciliary epithelium, the PCE and NPCE cells, are capable of RVD and RVI
when exposed to hypo- and hypertonic solutions, respectively (9, 9a,
12) (Fig. 2). When exposed to 50%
hypotonic solution, the NPCE and PCE cells swelled by 54 ± 17 (n = 24) and 60 ± 7%
(n = 30), respectively. The times to
peak volume were 52.5 ± 6.9 s (n = 24) for the nonpigmented cells and 44.7 ± 5.7 s
(n = 30) for the pigmented cells. In
the absence of inhibitor, volume-regulatory mechanisms overtook the
swelling process and caused a decrease in volume. As illustrated in
Fig. 2, both cell types had completely recovered their former volumes while still exposed to hypotonic solution. The time constants (the
times for the volumes to reach 1/e of
the total response) for this RVD, obtained by fitting a
single-exponential function to the volume decrease, were
121.7 ± 26.9 s (n = 3) for NPCE
cells and 99.5 ± 15.3 s (n = 4) for
PCE cells (Table 1).

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Fig. 1.
Cell swelling in a pigmented ciliary epithelial (PCE)-nonpigmented
ciliary epithelial (NPCE) coupled pair of cells. Digital images were
taken before cell swelling (A); at
peak of cell swelling, 200 s after addition of hypotonic solution
(B); and after regulatory volume
decrease (RVD), 15 min after addition of hypotonic solution
(C). Scale bar = 10 µm.
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Fig. 2.
Volume regulation in single NPCE (A)
and PCE (B) cells. Cells were bathed
in solution
A (composition given in
METHODS) for 10-20 min and then
with same solution made 50% hypotonic by dilution with water ( ,
) or hypertonic by addition of 100 mM
D-mannitol ( , ) for times
indicated by horizontal lines. After osmotic perturbation, cells
regulated their volume by either regulatory volume decrease (RVD) or
regulatory volume increase (RVI). Each data point is mean ± SE
(vertical line) of 3 experiments. Peak swell and shrink were
determined, and time constants for RVD or RVI were computed by fitting
a single-exponential function to decay and rise of volume after
hypotonic and hypertonic treatment and are presented in Table 1. All
experiments were performed at 22°C. A comparison of experimental
parameters measured at 22 and 35°C is given in Table
3.
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RVI.
Both NPCE and PCE cells decreased their volumes in response to
hypertonic solutions and then underwent RVI (Fig. 2). The peak shrink
values were 32 ± 4% (n = 4) for
NPCE cells and 34 ± 5% (n = 6) for
PCE cells. The time constants of the subsequent RVI were 9.4 ± 2.2 min (n = 4) for NPCE cells and 33.9 ± 24.7 min (n = 6) for PCE cells
(Table 2). These experiments were carried out at 35°C. RVI was not observed at room temperature.
Effect of temperature on RVD.
There was no major effect of temperature on either the peak swell or
time constant of RVD for NPCE or PCE cells. At 22°C the time
constants of RVD were 88.8 ± 26.2 s
(n = 4) for NPCE cells and 125.4 ± 35.4 s (n = 7) for PCE cells and the
peak swell values (relative volume) were 1.62 ± 0.23 (n = 5) for NPCE cells and 1.68 ± 0.11 (n = 11) for PCE
cells. These values are not significantly different from
those at 37°C (see Table 1).
Effect of inhibitors on RVD.
In an attempt to identify some of the mechanisms underlying RVD, we
used a range of different inhibitors. In particular we chose inhibitors
that are known to be more or less selective for Cl
and
K+ channels. To block
Cl
channels, we used DIDS,
SITS, 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB), quinidine,
bumetanide, and tamoxifen; to block
K+ channels, we used
Ba2+, quinidine, and
tetraethylammonium (TEA; in PCE cells). All of these
inhibitors prevented RVD to a greater or lesser extent in NPCE cells
(Fig. 3). The degrees of inhibition,
calculated from the ratio of volume recovery to total volume increase,
were as follows: 100% (tamoxifen), 87% (quinidine), 84% (DIDS), 80%
(NPPB), 77% (bumetanide), 58% (SITS), and 31%
(Ba2+). But, of all the
inhibitors, and additionally Gd3+
and TEA, only bumetanide caused a significant block (54%) of RVD in
PCE cells, prolonging the time constant for the incomplete RVD from a
control value of 123 s to 212 s (Fig. 3). A summary of the results of
the inhibitor studies is given in Table
3.


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Fig. 3.
Inhibition of RVD in single NPCE and PCE cells. Effects of inhibitors
of Cl channels on RVD in
response to 50% hypotonic solution in single NPCE cells ( ) and PCE
cells ( ) were investigated. A: 1 mM
DIDS; B: 1 mM SITS;
C:
10 4 M
5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB);
D: 1 mM quinidine;
E:
10 4 M bumetanide;
F:
10 5 M tamoxifen. Data are
means ± SE (vertical lines) from no. of experiments given in Table 3.
All inhibitor experiments were carried out at room temperature.
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The inhibitors, with the exception of bumetanide, were ineffective in
preventing RVD in the PCE cells. The two
K+ channel blockers,
Ba2+ and quinidine, delayed the
onset of RVD by 195 ± 13 (n = 4) and 130 ± 17 s (n = 4), respectively,
without actually preventing RVD, which went to completion in both cases.
Coupled cell pairs: injection of Lucifer yellow.
To examine the behavior of coupled (NPCE-PCE) cell pairs, we
demonstrated that the cells were coupled by injecting one cell of the
pair with Lucifer yellow. Figure 4
illustrates a light micrograph (A)
of a coupled pair, and the same pair under fluorescence illumination
after dye injection (B). Lucifer
yellow was injected into the PCE cell of a cell pair, and the dye
spread from the site of injection into the NPCE cell, demonstrating
that the cells form a coupled pair.

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Fig. 4.
Dye-coupled PCE-NPCE cell pair. A:
phase-contrast image of ciliary epithelial cells. A cell pair
consisting of a PCE cell (lower) and NPCE cell (upper) in top left
quadrant of picture was injected with Lucifer yellow via a patch
electrode attached to pigmented cell.
B: fluorescence image of same cells,
demonstrating that Lucifer yellow, injected into PCE cell, is able to
diffuse into NPCE cell via gap junctions. Scale bar = 10 µm.
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Coupled cell pairs: swelling and RVD.
Coupled NPCE-PCE cell pairs were exposed to 50% hypotonic solution,
and the volumes of both cells were determined as described for single
cells. The PCE cells swelled less (11 ± 12%;
n = 5) than their NPCE cell partners
(33 ± 7%; n = 5) and also less than their uncoupled counterparts (30 ± 4%;
n = 4) (Fig.
5). This reduced swelling is significantly
different (P = 0.0094; 2-tailed
t-test) from the mean swelling for PCE
cells (60 ± 7%; n = 30). The
coupled NPCE cells and their uncoupled counterparts swell to the same extent: 33 ± 7% (n = 5) compared
with 30 ± 4% (n = 3). Even when efflux is blocked by a range of inhibitors, the peak swell is more or
less the same as the control value (Table 3), perhaps indicating that
mechanical factors (e.g., cytoskeleton, membrane elasticity) set an
upper limit on the cell volume.

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Fig. 5.
Cell volumes (means ± SE) of coupled cell pairs
(A) and single cells
(B) in response to hypotonic
solution. Cells were bathed in solution
B (see METHODS) for
times indicated by horizontal lines. Experiments were performed at
35°C. Effect of temperature on experimental parameters is given in
Table 3. A: when coupled cells were
exposed to hypotonic solution, volume of pigmented cell ( ) of pair
peaked 80 s before (arrow), and at a significantly lower level than (30 compared with 50% swell), that of nonpigmented partner ( ) (see also
Table 1). B: cell swelling and RVD of
single dissociated cells followed approximately same time course in
both pigmented ( ) and nonpigmented ( ) cells. Peak swell values
(arrow) and time constants of RVD are not significantly different and
are given in Table 1.
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Coupled NPCE cells reached their peak volumes 80 s after the PCE cells
(Fig. 5); the time to peak volume after exposure to hypotonic solution
was 164 ± 23 s (n = 5) for
coupled PCE cells, significantly different
(P = 0.0474) from the value
for coupled NPCE cells, 204 ± 11.7 s
(n = 5). For uncoupled cells the times to peak volume were 193.3 ± 17.6 s
(n = 3) for PCE cells and 180.0 ± 30.6 s (n = 34) for NPCE cells; these
values are not significantly different.
The time constants of RVD, 184.6 ± 35.7 s
(n = 5) for NPCE cells and 130.9 ± 74.7 s (n = 5) for PCE cells, were not
significantly different (P = 0.5348).
Effect of tamoxifen on RVD in coupled pairs.
When coupled pairs of NPCE and PCE cells were exposed to hypotonic
solution in the presence of tamoxifen
(10
5 M; Fig.
6) three things were noted. First, RVD in
the NPCE cells was no longer 100% inhibited as in single cells;
instead the inhibition was 48.1 ± 14.4%
(n = 7). Second, RVD was inhibited in
the PCE cells by 33.7 ± 12.3% (n = 7), whereas tamoxifen had no effect on RVD in single PCE cells. Third,
the peak of the swelling for PCE cells was no longer time shifted. The
time to peak volume was 171.4 ± 36.3 s
(n = 7) for PCE cells and 202.9 ± 42.6 s (n = 7) for NPCE cells. These
values are not significantly different from those for single cells.

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Fig. 6.
Effect of tamoxifen (10 5 M)
on RVD in NPCE-PCE cell coupled pairs. Mean data ± SE from 7 coupled
cell pairs exposed to hypotonic solution in presence of tamoxifen
(10 5 M) are shown. Peak
volumes for NPCE and PCE cells are now coincident (cf. Fig.
5A). Tamoxifen only blocks a
proportion of RVD in NPCE cells, and, unlike what is found for single
PCE cells (see Fig. 3), RVD in coupled PCE cells is partially blocked
by tamoxifen. Experiments were performed at 35°C.
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 |
DISCUSSION |
In this study we have determined the characteristics of volume
regulation in PCE and NPCE cells. From the degree of volume change, the
time to peak or minimum volume, and the time constant of RVD or RVI, we
can define the kinetics of the processes of RVI and RVD. We have done
this for PCE and NPCE cells under two different conditions: as single
cells and as coupled, heterogeneous (NPCE-PCE cell) pairs.
We have discovered two things: first, NPCE and PCE cells have
different RVD mechanisms as defined by their different inhibitor sensitivities; second, coupling significantly alters the
properties of RVD in a way that suggests vectorial solute movement
from PCE to NPCE cells.
Single-cell studies.
The kinetic parameters for RVD in single PCE and NPCE cells are similar
(Table 1). Differences between the time constants of RVD in our studies
(
100 s) and those of other laboratories (
10 min; Refs. 1 and 3)
are due to the indirect measurement technique used in the latter
studies, a fact that has been discussed elsewhere (9). Farahbakhsh and
Fain (4) measured RVD directly by interference contrast microscopy and
found time constants for rabbit NPCE cells similar to those reported in
this study.
The pharmacologies of the mechanisms of RVD are radically different
between the two cell types (8). All the
Cl
channel inhibitors we
used (DIDS, SITS, NPPB, quinidine, and tamoxifen) inhibited RVD in the
NPCE cells to a greater or lesser extent, but none of them had any
effect on RVD in the PCE cells. Mitchell et al. (12) reported that, of
the Cl
channel inhibitors
that blocked the volume-activated
Cl
current in NPCE cells,
DIDS, SITS, NPPB, dideoxyforskolin, verapamil, tamoxifen, and quinidine
(16), only DIDS, SITS, and NPPB had any effect on PCE cells. These
three inhibitors operate a voltage-dependent block because of their
negative charge. Mitchell et al. (12) predicted that the small degree
of block with DIDS and SITS at negative potentials would render them
almost ineffective at the resting membrane potential, and this was
found to be the case. Further support for the difference between the
NPCE and PCE cells comes from the finding of different populations of
volume-activated Cl
channels in the two cell types (18).
The K+ channel inhibitor
Ba2+ also failed to inhibit RVD in
the PCE cells, as did Gd3+, a
nonselective cation channel inhibitor. The process of RVD in these
cells was not, however, so robust as to be uninhibitable. One inhibitor
that affected RVD in the PCE cells was bumetanide, a blocker of
Na+-K+-Cl
cotransport. It is thought that this was achieved by a depletion of
intracellular Cl
, reducing
the gradient for Cl
efflux
during RVD.
RVD in coupled pairs of NPCE-PCE cells.
In the ciliary epithelium the NPCE and PCE cells form a syncytium. We
used coupled pairs of NPCE and PCE cells, first described by Edelman et
al. (3), as a model for the ciliary epithelium. To demonstrate that
they were functionally coupled, we injected Lucifer yellow into the
pigmented cell of such a pair and observed that the dye spread into the
nonpigmented cell.
The coupled NPCE-PCE cell pair responded quite differently to hypotonic
challenge than did single NPCE and PCE cells. Two facts
emerge from this experimental comparison. First, the peak swell of the
single pigmented cells becomes very much smaller when the cells are
coupled; when the cells were uncoupled, the peak swell values for NPCE
and PCE cells were not significantly different (54 ± 17%,
n = 24, and 60 ± 7%,
n = 30, respectively). Second, the
time of peak swell for the pigmented cells, which is more or less
coincident with that for the nonpigmented cells when they are separate,
precedes that for the nonpigmented cells by 80 s when the cells are
coupled (Fig. 5, arrows). We interpret these observations as indicating
a flow of fluid from the pigmented to the nonpigmented cells. The PCE
cells begin to swell, but, before they reach their peak volume, solute
moves into the NPCE cells and exits via channels in the NPCE cells. The
question of what drives the movement of fluid from PCE to NPCE cells is
an interesting one. There are a number of possibilities. One is that the PCE cells could have a lower water permeability than the NPCE cells, so that the concentration gradient drives solute from PCE to
NPCE cells. This possibility seems implausible given the observation that the times to peak swelling for uncoupled NPCE and PCE cells were
the same. Alternatively, the structural union of the two cells may in
some way alter the membrane transport properties of the PCE cells. The
physics of surface tension offers another explanation. At equal surface
tension a smaller bubble has a larger internal pressure than a bigger
one, so its contents will be pumped into the larger bubble when they communicate.
The two cell types clearly have very different RVD mechanisms, which
may have different activation properties or set points, such that more
solute per unit time can pass through the NPCE membranes, resulting in
a net movement of fluid from PCE to NPCE cells. The addition of
tamoxifen, which blocks RVD in NPCE but not PCE cells, causes an
apparently reduced RVD in both cell types. The time course of the RVD
in NPCE cells follows that in the PCE cells. This could be interpreted
as a movement of solute from NPCE cells (now efflux inhibited) to PCE
cells, where it passes through tamoxifen-resistant pathways.
What relevance do these osmotically induced fluxes have to fluid
transport? We induced cell swelling by lowering the external osmolarity, but the same effect could be obtained by increasing the
intracellular osmolarity. This could be achieved by uptake mechanisms
such as cotransporters (2, 15). Such mechanisms are known to be
involved with RVI. Figure 7 illustrates a model in which
volume oscillation is achieved by having the volumes at which RVI and
RVD turn on and off slightly greater and smaller than the "set"
volume. The cyclical and reciprocal activation of RVI and RVD
mechanisms, situated on opposite membranes, would enable an oscillatory
fluid flow across the tissue without the need for changes in external
osmolarity.

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Fig. 7.
Model for oscillatory cell volume. Cell is assumed to have a set volume
(Vo), and the volume at which
RVI turns on and RVD turns off,
V 1, is below
Vo, whereas the volume at which
RVI turns off and RVD turns on,
V+1, is above
Vo. This model predicts that cell
volume will oscillate around a set point.
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In conclusion, we show that vectorial flow from one cell to another can
be achieved under conditions of osmotic swelling and may be due to
differences in the volume-activated ion channels between the two cell
types. For a given osmotic challenge NPCE cells pass more solute than
PCE cells. This creates a movement of solute from PCE to NPCE cells,
via communicating gap junctions. Because the PCE cells are on the
serosal (blood) side of the ciliary epithelium and the NPCE cells face
the inside of the eye, this vectorial flow is in the same direction as
that for the secretion of aqueous humor.
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ACKNOWLEDGEMENTS |
We gratefully acknowledge financial support from the Medical
Research Council, the Royal Society, The Wellcome Trust, and the Royal
National Institute for the Blind.
 |
FOOTNOTES |
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: T. J. C. Jacob,
School of Biosciences, Cardiff Univ., Cardiff CF1 3US, UK (E-mail:
jacob{at}cardiff.ac.uk).
Received 30 June 1998; accepted in final form 18 March 1999.
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