Human trabecular meshwork cell volume regulation
Claire H.
Mitchell1,
Johannes C.
Fleischhauer1,
W. Daniel
Stamer3,
K.
Peterson-Yantorno1, and
Mortimer M.
Civan1,2
Departments of 1 Physiology and
2 Medicine, University of Pennsylvania School of
Medicine, Philadelphia, Pennsylvania 19104-6085; and
3 Department of Ophthalmology, University of
Arizona, Tucson, Arizona 85711-1824
 |
ABSTRACT |
The volume of
certain subpopulations of trabecular meshwork (TM) cells may modify
outflow resistance of aqueous humor, thereby altering intraocular
pressure. This study examines the contribution that
Na+/H+, Cl
/HCO
exchange, and K+-Cl
efflux mechanisms have on
the volume of TM cells. Volume, Cl
currents, and
intracellular Ca2+ activity of cultured human TM cells were
studied with calcein fluorescence, whole cell patch clamping, and fura
2 fluorescence, respectively. At physiological bicarbonate
concentration, the selective Na+/H+ antiport
inhibitor dimethylamiloride reduced isotonic cell volume. Hypotonicity
triggered a regulatory volume decrease (RVD), which could be inhibited
by the Cl
channel blocker
5-nitro-2-(3-phenylpropylamino)-benzoate (NPPB), the K+
channel blockers Ba2+ and tetraethylammonium, and the
K+-Cl
symport blocker
[(dihydroindenyl)oxy]alkanoic acid. The fluid uptake mechanism in
isotonic conditions was dependent on bicarbonate; at physiological
levels, the Na+/H+ exchange inhibitor
dimethylamiloride reduced cell volume, whereas at low levels the
Na+-K+-2Cl
symport inhibitor
bumetanide had the predominant effect. Patch-clamp measurements showed
that hypotonicity activated an outwardly rectifying, NPPB-sensitive
Cl
channel displaying the permeability ranking
Cl
> methylsulfonate > aspartate.
2,3-Butanedione 2-monoxime antagonized actomyosin activity and both
increased baseline [Ca2+] and abolished
swelling-activated increase in [Ca2+], but it did not
affect RVD. Results indicate that human TM cells display a
Ca2+-independent RVD and that volume is regulated by
swelling-activated K+ and Cl
channels,
Na+/H+ antiports, and possibly
K+-Cl
symports in addition to
Na+-K+-2Cl
symports.
outflow facility; calcein; chloride channels; potassium-chloride
symport; sodium/hydrogen antiport; methylsulfonate; aspartate; intraocular pressure; [(dihydroindenyl)oxy]alkanoic acid
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INTRODUCTION |
INTRAOCULAR PRESSURE
(IOP) is determined by the relative rates of inflow and outflow of
aqueous humor. Aqueous humor is secreted by the ciliary epithelium and
returns to the vasculature of the primate eye largely through the
trabecular meshwork (TM) and Schlemm's canal (26).
Despite the importance of inflow as a target of medical therapy,
glaucoma results from increased resistance to aqueous humor outflow
(6), usually leading to increased IOP, and is a major
cause of blindness. Thus the mechanisms underlying aqueous humor
outflow are of physiological interest as well as potential clinical relevance.
The TM comprises connective tissue suspended like a web between the
scleral spur and Schwalbe's line around the entire circumference of
the eye (24, 30). The inner region, nearer the anterior chamber, displays plates or beams of connective tissue covered with TM
cells. The spacing between the beams narrows during passage outward
until reaching the juxtacanalicular tissue (JCT) region, the area
adjoining the inner wall of Schlemm's canal. At this point, cells
reside in a milieu of extracellular matrix and become intermingled and
attached to each other, to the inner wall of Schlemm's canal, and to
the surrounding fine connective tissue fibrils. The cells in this area,
termed JCT-TM cells, are phenotypically different from TM cells but
share a common neural crest origin. It is at this point in the outflow
pathway, the juxtacanalicular area, that the volume of the TM and
JCT-TM cells is most likely to affect resistance to outflow of aqueous
humor between the cells.
The basis for outflow regulation is unknown but may involve
(24) contraction and/or relaxation of both the TM cells
and ciliary muscle (29, 52, 56, 57), pore formation in the inner wall of Schlemm's canal by either direct or indirect actions (14, 25), changes in extracellular matrix of the JCT
(1, 24, 30, 31), passive stretch, and changes in shape
(13) and swelling and/or shrinkage of the cell volume of
the TM cells (2, 15, 18, 19, 36, 40, 42, 58). Changes in
the cytoskeleton may be linked to both cellular contraction and/or relaxation (52) and shrinkage and/or swelling (22,
59). These possibilities are not mutually exclusive.
The guiding hypothesis of the present work is that swelling of the TM
and JCT cells could well present a significant obstruction to flow
between the collagenous beams of the juxtacanalicular region of the TM,
as suggested by published electron micrographs (15). This
possibility is supported by observations that maneuvers producing cell
swelling reduce outflow facility and maneuvers shrinking the cells
increase that facility in human, nonhuman primate, and calf eyes
(2, 21, 44). Volume regulation of TM cells by
Na+-K+-2Cl
symport has been
suggested to modulate outflow facility (2). However,
blocking that symport with bumetanide has no measurable effect on
outflow facility in the living cynomolgus monkey (16) and
does not lower IOP in the monkey (16) or mouse
(5a), questioning this hypothesis. One possible
interpretation of the null result is that TM and JCT-TM cell volume
cannot be considered a function of symport alone but involves
additional transporters not yet characterized.
Cell volume regulation of many cells depends on the integrated
operation of multiple solute and water uptake mechanisms and a similar
number of release pathways (9, 10, 23, 27, 38, 45, 49).
Na+-K+-2Cl
symport has been
reported to be the major mechanism of regulatory volume solute uptake
by TM cells, at least in the presence of low external
HCO
concentration (4.2 mM) (36). In the
present study, we tested whether paired Na+/H+
and Cl
/HCO
exchangers significantly
modify solute and water uptake in the presence of physiological levels of extracellular bicarbonate and also identified two regulatory volume
mechanisms (Cl
channels and
K+-Cl
symports) for potential solute and
water release by TM and JCT-TM cells.
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MATERIALS AND METHODS |
Cell preparations.
Human TM cells were isolated after collagenous digestion of TM
explants, as previously described (51). The cells obtained likely reflected a mixture of TM cells from beams and juxtacanalicular JCT-TM cells. The cells obtained in this way have been characterized with respect to their growth properties, morphology, presence of a
cell-surface receptor for a low-density lipoprotein, and induction of
myocillin protein expression upon dexamethasone treatment (50), and they have been used previously for studying
aquaporin-1 (51) and
2-adrenergic
(48) and prostaglandin F2
receptors (5) present in these cells. The lines and passage numbers
(P) studied are specified for each experiment.
The human TM cells were grown to 80% confluence at 37°C in 5%
CO2 before study and split at a ratio of
1:4. The medium
was low-glucose Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum, 100 U/ml penicillin, and 100 µg/ml
streptomycin (GIBCO BRL, Grand Island, NY).
Use of calcein fluorescence as an index of cell volume.
Electronic cell sorting has been our principal technique for measuring
cell volume of ocular epithelial cells (9, 10) but was
unsuitable as a routine procedure for studying limited numbers of human
TM cells. For example, 15,000-30,000 cells were required for each
time point shown in Fig. 3B, as measured by electronic cell
sorting. It would be possible to produce large numbers of cells from
the limited number obtained during primary isolation of nontransformed
cells if the cultures were split many times. However, it seemed
preferable to choose techniques permitting us to work with smaller
numbers of cells at lower passage numbers, presumably closer to in vivo conditions.
We considered multiple alternative approaches (see
DISCUSSION) and conducted preliminary measurements with
several fluorescent probes, including calcein, Oregon green,
N-(ethoxycarbonylmethyl)-6-methoxyquinolinium bromide (MQAE),
fluorescein, and fura 2. Calcein fluorescence proved the most
satisfactory of these approaches in terms of convenient, reliable
monitoring of an index of cell volume over periods as long as ~70
min. Calcein is easily loaded in the acetoxymethyl ester (AM) form, is
well retained within TM cells, and displays a fluorescence two to three
times greater than that of other commonly used fluorophores, and its
fluorescence is relatively independent of shifts in pH and
Ca2+ (4).
Our strategy was to monitor cell area as an index of cell volume (see
DISCUSSION). For this purpose, cells were studied either after growth on coverslips for 1-5 days or 30-90 min after
acute harvesting with 0.25% trypsin (GIBCO BRL). Unless otherwise
stated, the coverslips were obtained from Fisher Scientific (catalog
no. 12-545-82; Pittsburgh, PA); data obtained with
poly-L-lysine coverslips (Becton Dickinson, Bedford, MA)
are so indicated. TM cells were loaded with 4 µM calcein-AM and
0.02% Pluronic at room temperature for 30-40 min. Coverslips were
mounted in a chamber and visualized with a ×40 oil-immersion objective
on a Nikon Diaphot microscope. Fields were chosen to include several
cells of comparable diameter, displaying comparable loading, and
contained between one and four nonconfluent cells each. Focus was
adjusted by maximizing the edge contrast between cells and the bath
displayed on the monitor, thus maximizing the cell area, and was not
thereafter changed during the experiment. Calcein was excited every
20 s at 488 nm, and light emitted at 520 nm was detected with an
IC-200 charge-coupled device camera (Photon Technology International,
Princeton, NJ). Cell area was defined as the number of pixels above
threshold within a region of interest and was determined using
Imagemaster software (Photon Technology International). Threshold was
automatically set at an intensity of 90 (out of a maximum gray scale of
256) because initial experiments showed this value was optimal. Figure 1 shows the raw digitized images of a
cell changing area upon exposure to hypotonicity and illustrates the
effect of the thresholding protocol.

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Fig. 1.
Imaged response of freshly harvested trabecular meshwork
(TM) cell to hypotonicity and thresholding. A:
digitized image of TM cell loaded with calcein-AM in isotonic solution
without 2,3-butanedione 2-monoxime (BDM). The cell [human TM (hTM) no.
29 (P4), where P is passage] contributed to the averaged results
presented in Fig. 3. B: the same cell after application of
hypotonic solution, when cell area was maximum. C: after
~20 min in hypotonic solution, the cell area was smaller, reflecting
a regulatory volume decrease (RVD). D-F: images
correspond to A-C, but with a threshold applied to
render all non-cell background black. Careful examination shows that
the area remaining after thresholding accurately reflects the true cell
size. Area was calculated as the number of nonblack pixels after
thresholding.
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Figure 2A presents the time
course of the normalized area of a cell grown on a coverslip and
perfused sequentially with the isotonic (Iso-Cl) and hypotonic
(Hypo-Cl) solutions described in Table 1,
with a mixture of the two solutions and with Iso-Cl rendered hypertonic
by addition of NaCl. Graded responses of cell area were observed
following perfusion with changes in tonicity. Assuming proportional
changes in volume along the three coordinate axes, volume is expected
to be proportional to (area)3/2. As expected for a simple
osmometer, the index of cell volume was linearly dependent on
1/osmolality during brief exposure to anisosmotic perfusates (Fig.
2B).

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Fig. 2.
Response of TM cell area to brief changes in bath
osmolality. A: time course of normalized area of a cell
grown on a coverslip and perfused sequentially with isotonic (Iso) and
hypotonic (Hypo) solutions (see Table 1 for description of solutions),
with a mixture of the 2 solutions, and with Iso rendered hypertonic by
addition of NaCl (Hyper). B: assuming proportional changes
in volume along the 3 coordinate axes, volume is taken to be
proportional to (area)3/2. This calculated index of cell
volume (mean ± SE) was linearly dependent on 1/osmolality during
the brief anisosmotic perfusions [R2 = 0.83, n = 3; hTM no. 29 (P4)].
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In studying acutely harvested preparations, ~20% of the cells either
migrated out of the region of study or displayed a "cat-in-a-bag" phenomenon (a rapid series of contractions and relaxations),
complicating data reduction. These phenomena were suppressed in almost
all cells by including an antagonist of actomyosin activity,
2,3-butanedione 2-monoxime (BDM) (32), in the perfusates.
The presence of BDM did not alter the regulatory volume response to
hypotonic swelling (see RESULTS).
For comparison of data obtained by conventional electronic cell
sorting, we also conducted two experiments following our standard protocol (7, 9, 10, 34). After cells were harvested from a
T-75 flask by trypsinization, a 0.5-ml aliquot of the cell suspension
in DMEM was added to 20 ml of the suspension solution. Cell volumes of
isosmotic suspensions were measured with a Coulter counter (model
ZBI-Channelyzer II) using a 100-µm aperture. The cell volume of the
suspension is taken as the peak of the distribution function.
All data represent means ± SE, and significance was determined by
using the F-test as previously described (34).
Intracellular Ca2+ activity.
For measurements of intracellular Ca2+, cells grown on
coverslips for 1-10 days were loaded in the dark with 5 µM fura
2-AM and 0.01% Pluronic F-127 (Molecular Probes, Eugene, OR) for 30 min at 25°C and perfused with fura-free solution for 30 min before data acquisition was begun (34). Coverslips were mounted
on a Nikon Diaphot microscope and visualized with a ×40 oil-immersion fluorescence objective. The emitted fluorescence (520 nm) from ~12
cells at ~90% confluence was sampled at 1 Hz with the
photomultiplier following excitation at 340 and 380 nm, and the
ratio was determined with a Delta- Ram system and Felix software
(Photon Technology International). The ratio of light excited at 340 nm
to that at 380 nm was taken as a direct index of intracellular
Ca2+ activity. In a subset of experiments, that ratio was
converted into Ca2+ concentration by using the method of
Grynkiewicz et al. (20). An in situ
Kd value for fura 2 of 350 nM was used
(35). Rmin was obtained by bathing cells in a
Ca2+-free isotonic solution of pH 8.0 containing 10 mM EGTA
and 10 µM ionomycin. Rmax was obtained by bathing the
cells in isotonic solution with either 0.1 or 2.5 mM Ca2+
and 10 µM ionomycin. Calibration was performed separately for each
experiment. Baseline levels from TM cells in the absence of fura 2 were
subtracted from records to control for autofluorescence.
Intracellular pH activity.
Experiments measuring intracellular pH (pHi) were performed
in a manner similar to that of the Ca2+ measurements
but using 2 µM
2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF)-AM (Molecular Probes) and 0.002% Pluronic F-127. The
emitted fluorescence (520 nm) from 15-25 confluent cells was
sampled at 1 Hz following excitation at 480 and 440 nm, and the ratio
was determined with a Delta-Ram system and Felix software.
pHi calibration, based on that of Wu et al.
(57a), was performed by perfusing the cells with 110 mM
KCl, 20 mM NaCl, 20 µM nigericin, and 20 mM buffer [pH 6.0 solution
buffered with MES, pH 7.0 with PIPES, pH 7.4 with HEPES, and pH 8.0 with TES].
Cl
currents.
Whole cell patch-clamp currents were recorded in the ruptured-patch
mode. Micropipettes were pulled from Corning no. 7052 glass, coated
with Sylgard, and fire polished. The resistances of the micropipettes
in the bath were usually ~1-3 M
; successful seals displayed
gigaohm resistances. After rupture of the membrane patch, the series
resistance was measured to be only 8.0 ± 0.9 M
and was
therefore not compensated; whole cell capacitance was 95 ± 5 pF.
The baseline whole cell currents were 1.4 ± 0.7 pA/pF at +80 mV.
Seals were always formed in the presence of Cl
-Tyrode
solution (Table 2, NaCl solution). The
applied voltages were not corrected for the small junction potential
[approximately
2.8 mV (7)] arising from the present
micropipette and external solutions, but the correction was included in
analyzing the reversal potential (Erev) in the
NaCl bath. In changing perfusates, the entire chamber volume was
replaced by the new solution so that the reference potential between
the 3 M KCl agar bridge and the bath solutions was taken to be constant
and approximately zero.
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 either
40 or
80 mV and stepped to test voltages from
100 to +80 mV in 20-mV increments at 2-s intervals. At the more
negative holding potential, depolarizations produced clearly
recognizable transient inward currents, consistent with L-type
Ca2+ currents known to characterize these cells
(56). Otherwise, the current responses to voltage steps
were similar at
40,
80, and even
142 mV. 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.
Presumed Cl
currents were fit by a form of the Goldman
equation for Cl
channel currents
|
(1)
|
where
|
(2)
|
|
(3)
|
|
(4)
|
and Vm is the membrane potential,
PCl and PAsp are the
Cl
and aspartate permeabilities, F is the
Faraday constant, [Cl
]i and
[aspartate
]i are the cellular
Cl
and aspartate concentrations, respectively,
R is the perfect gas constant, and T is the
absolute temperature. Estimates for both unknown parameters
(K and Erev) were generated by
nonlinear least-squares analysis.
Drugs and experimental solutions.
All chemicals were reagent grade. The AM form of calcein (Molecular
Probes) was used to load cells when studying volume. Among the drugs
administered were the selective Na+/H+ antiport
inhibitor dimethylamiloride (DMA; Sigma, St. Louis, MO)
(11), the K+-Cl
-symport
inhibitor [(dihydroindenyl)oxy]alkanoic acid (DIOA; Sigma-RBI) (17), the actomyosin antagonist BDM (Sigma)
(32), and the Cl
channel blocker
5-nitro-2-(3-phenylpropylamino)-benzoate (NPPB; Biomol Research
Laboratories, Plymouth Meeting, PA) (55). The compositions
of the isotonic and hypotonic solutions used for fluorescence and
patch-clamp measurements are presented in Tables 1 and 2, respectively.
 |
RESULTS |
Regulatory volume decrease.
The response of calcein-determined cell area to anisosmotic swelling
was monitored with cells prepared in several ways, including growth on
poly-L-lysine-coated coverslips (Fig.
3A), growth on conventional
coverslips (Figs. 5-6), plating without BDM after acute harvest
(Fig. 3A), and plating in the presence of BDM after acute harvest (Fig. 4). The results were qualitatively the same, independent of preparative approach. Hypotonic perfusates produced an increase in
cell area and triggered a secondary regulatory decrease toward the
baseline isotonic value. These indications of a regulatory volume
decrease (RVD) from measurement of area by calcein fluorescence conformed qualitatively to the RVD observed in two experiments conducted with conventional electronic cell sorting (Fig.
3B). Insofar as indications of the RVD noted with classic
electronic cell sorting were displayed by cells grown on coverslips or
acutely harvested with or without BDM, all of these cell preparations were used interchangeably in studying TM cell transporters.

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Fig. 3.
RVD of human TM cells. A: response of cell
area measured by calcein fluorescence to sustained exposure to
hypotonic solution. The swelling triggered a progressive reduction in
area during the hypotonic perfusion. Restoration of isotonicity stopped
the shrinkage but did not trigger a regulatory volume increase (RVI),
consistent with reports that the RVI of some cells cannot be observed
at room temperature (as here) but only at higher temperatures
(12, 54). Cells were harvested onto
poly-L-lysine coated coverslips in the absence of BDM
[n = 3; hTM no. 29 (P4) and hTM no. 36 (P4)].
B: response of cell volume measured by Coulter counter
[n = 2; hTM no. 29 (P4-5)] to sustained exposure to
same hypotonic solution. The percent volumes were normalized to 128, the value of maximal swelling noted in A. Uncertainties were
calculated as half the difference between the 2 means.
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Fig. 4.
Inhibition of RVD by Cl and K+
channels blockers. A: suppression of RVD by 100 µM
5-nitro-2-(3-phenylpropylamino)-benzoate (NPPB). Both control [n
= 12; hTM no. 22 (P3)] and NPPB [n = 7; hTM no. 22 (P2) and hTM no. 29 (P3)] traces are from the second hypotonic
exposure following protocol 1 with freshly harvested cells
in the presence of 20 mM BDM. B: elimination of RVD by
simultaneous presentation of 5 mM BaCl2 and 7.5 mM TEA.
Traces represent the mean responses from 5 experiments following
protocol 2 using cells grown on coverslips [hTM no. 25 (P4)].
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Methodology of studying inhibitors of the RVD.
The RVD illustrated in Figs. 1 and 3-5 was inhibited by ion
channel blockers. The quantification of the effect of these blockers has been hindered by a variability in the magnitude and time course between the first and second RVD response of the same cell, as well as
variability in the RVD expression by different cells. We addressed
the technical problem posed by cellular heterogeneity of the RVD
response with two protocols, both involving two periods of hypotonic
perfusion (separated by an intervening period of isotonic perfusion) of
the same cell.

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Fig. 5.
Inhibition of RVD by the
K+-Cl -symport blocker
[(dihydroindenyl)oxy]alkanoic acid (DIOA). Cells grown on coverslips
were hypotonically perfused twice. DIOA (100 µM) was included in
either the first (n = 5) or second (n = 8)
hypotonic perfusion [hTM no. 29 (P5) and hTM no. 36 (P4)]. Six of
thirteen cells were preperfused with isotonic DIOA solution for 5 min
before perfusion with hypotonic DIOA solution.
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Protocol 1 was to study control and experimental cells on
separate coverslips (Fig. 4A).
Cells on control coverslips were never exposed to inhibitors but were
simply perfused twice successively with hypotonic solution alone. Cells
on the parallel experimental coverslips also received no drugs during
the first hypotonic perfusion; drugs were included only in the second
hypotonic perfusate. Thus control and experimental cells were treated
identically until the second hypotonic perfusion, at which point
inhibitors were applied to the experimental cells and only solvent
vehicle to the control cells. Comparisons of the responses to the first
hypotonic perfusate (containing no drugs) permitted us to test whether
the cells on the experimental and control coverslips were functionally similar. Comparisons of the responses to the second hypotonic perfusate
(±inhibitors) permitted us to test whether or not the blockers
affected the experimental cells significantly.
Protocol 2 was to study cells successively with control
hypotonic perfusates and drug-containing hypotonic perfusates but to
randomize the order of application (Figs. 4B and 5). This
second protocol was simpler but rested on the assumption that the
inhibitory effect of the first hypotonic perfusion was entirely
reversed during the period of isotonic perfusion and before the second period of hypotonic perfusion. In contrast, the first protocol did not
involve any assumptions but required twice as many cells and twice as
much experimental time. In practice, both protocols proved
satisfactory, at least with the drugs applied in the present work.
Ion transporters underlying the RVD.
Blockage of either Cl
or K+ channels alone
inhibited the RVD of human TM cells. In Fig. 4A, the
Cl
channel blocker NPPB eliminated the RVD, and in Fig.
4B, the K+ channel blockers TEA and
Ba2+ also prevented RVD. This combination of blockers was
found most effective, likely due to differential sensitivity of several
types of K+ channel; TEA (7.5 mM) alone did not produce
such an inhibition. This implies that the RVD reflected parallel
activation of both Cl
and K+ pathways.
Many cells also display an RVD mediated by solute release through
K+-Cl
symports (23, 27, 45).
This point was addressed by including the
K+-Cl
symport inhibitor DIOA (100 µM) in
either the first (n = 5) or second (n = 8)
hypotonic perfusion, using protocol 2. The results shown in
Fig. 5 summarize the results of two
series of experiments. DIOA (100 µM) was perfused isotonically for 5 min before being applied hypotonically in one series [human TM (hTM)
no. 36 (P4), n = 7] but not in the other [hTM no. 29 (P5), n = 6]. The results were qualitatively similar and
therefore were averaged together. The data indicate that DIOA partially
inhibited the RVD. DIOA blocks Cl
/HCO
antiport exchange and large-conductance Ca2+-activated
K+ channels (BK channels), in addition to inhibiting
K+-Cl
cotransport (17). Blocking
Cl
/HCO
exchange would have caused cell shrinkage and reduction in cell area, contrary to observation, and the
inability of BK channel blocker TEA to prevent RVD argues against this
site of DIOA action. Thus the simplest conclusion is that DIOA
inhibited RVD by blocking K+-Cl
symport
activity, although nonspecific actions are also possible.
Effect of bicarbonate, Cl
, and
methylsulfonate on isotonic cell volume.
The preceding measurements of the cellular response to anisosmotic
swelling indicated that Cl
can be released from human TM
cells through both Cl
channels and
K+-Cl
symports. Under certain conditions,
uptake of Cl
can proceed through bumetanide-sensitive
Na+-K+-2Cl
symports (36,
41, 42), but previous studies of ciliary epithelial cells from
this laboratory suggest that paired antiport exchange of
Na+/H+ and
Cl
/HCO
antiports might underlie
Cl
uptake when human TM cells are perfused with
physiological levels of bicarbonate (33). The effect of
bicarbonate on uptake was consequently tested in isotonic solution. In
the presence of a physiological 30 mM bicarbonate, perfusion with the
selective Na+/H+ antiport inhibitor DMA (10 µM) produced a prompt reduction in cell area, whereas 10 µM
bumetanide had little effect (Fig.
6A). The situation was
reversed in a bicarbonate-free solution, when bumetanide reduced cell
volume whereas DMA had little effect (Fig. 6B). This
suggests that the relative contribution by the
Na+-K+-2Cl
symports and paired
Na+/H+ and
Cl
/HCO
antiports might depend on the level of bicarbonate.

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Fig. 6.
Isosmotic modulation of TM cell area and pH. A: effect
of the Na+/H+ antiport inhibitor
dimethylamiloride (DMA) and the
Na+-K+-Cl symport inhibitor
bumetanide (Bumet) on isosmotic TM cell area in the presence of 30 mM
HCO . DMA (10 µM) promptly and progressively
reduced cell area, whereas 10 µM bumetanide had little consistent
effect [hTM no. 22 (P3) cells grown on coverslips, n = 3].
B: effect of HCO removal on isotonic
uptake mechanisms. With 0 HCO , 10 µM DMA had
little effect on cell volume, whereas 10 µM bumetanide triggered a
clear reduction [hTM no. 25 (P4) and hTM no. 47 (P3) cells grown on
coverslips, n = 12]. C: effect of
methylsulfonate substitution for external Cl on isosmotic
TM cell area. Perfusion with isotonic methylsulfonate solution (see
Table 1) triggered a large reduction in cell area, which was partly
reversible on return to isotonic Cl solution [hTM no. 22 (P4) cells grown on coverslips, n = 4]. D:
effect of DMA on intracellular pH (pHi). In the presence of
10 mM HCO and 0.5 mM DIDS, 10 µM DMA lead to a
reversible decrease in pHi [hTM no. 36 and no. 25 (P3),
n = 4].
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To confirm the contribution of the Na+/H+
exchanger, the pHi was monitored by using the pH-sensitive
dye BCECF in response to 10 µM DMA. DMA steadily and reversible
reduced pHi, consistent with the inhibition of a
Na+/H+ exchanger. The effect was best detected
in the presence of 10 mM HCO
and 0.5 mM DIDS to
limit potential confounding contributions of
Cl
/HCO
exchange and
Na+-nHCO
cotransport. Intracellular
Cl
content is thought to play a central role in consensus
models of cell volume regulation (23, 27, 45). In this
context, the report that methylsulfonate replacement of bath
Cl
induced a two-thirds loss of cell Cl
without changing TM cell volume has been unexplained (42). One possible interpretation is that human TM cells display an unusually
high permeability to methylsulfonate through the anion channels. We
have reexamined this phenomenon under the present conditions (30 mM
HCO
and room temperature) and with the calcein
fluorescence approach. As shown in Fig. 6C, we found that
methylsulfonate replacement of bath Cl
produced a prompt,
progressive fall in cell volume, which was partly reversible when
external Cl
was restored. These results suggest that the
permeability of the anion channels of human TM cells is much lower to
methylsulfonate than to Cl
. This conclusion was tested by
whole cell patch clamping.
Patch clamping.
Figure 7 presents representative results
obtained with a human TM cell perfused with isotonic and hypotonic
solutions containing Cl
, methylsulfonate, or aspartate as
the principle anion (Table 2, NaCl, NaAsp, or NaMeth). The mean (±SE)
current at each of the 10 applied voltages is presented as a function
of time. In the initial experimental period (data not shown), the
baseline currents in isotonic perfusates were very small and were
little affected by the anionic substitutions. Perfusion with hypotonic perfusate of the same ionic composition triggered an ~40-fold increase in currents. The currents peaked at ~41 min (~11 min after
hypotonic perfusion was initiated) and began to decline slowly at ~46
min. The outward currents at +80 mV were reduced ~75% in aspartate
and ~55% in methylsulfonate baths. The anionic replacements had
little effect on the inward currents because the composition of the
micropipette solution remained constant. Perfusion with 100 µM NPPB
in hypotonic Cl
solution produced a marked and largely
reversible inhibition of inward and outward currents. Restoration of
isotonicity triggered a prompt, progressive decline of whole cell
currents toward their initial isotonic values.

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Fig. 7.
Effects of osmolality and anionic substitutions on whole
cell TM cell currents. Currents were measured at test potentials
ranging from 100 to +80 mV at 20-mV intervals, with a holding
potential of 80 mV [hTM no. 22 (P3)]. Values are means ± SE
and were calculated from the currents over the entire 300-ms duration
of each step. Hypotonic perfusion, beginning 28 min into the trace,
triggered an ~40-fold increase in currents. The outward currents were
reduced by partial substitution of bath Cl with
methylsulfonate (M) or aspartate (A), and outward and inward currents
were reversibly inhibited by the Cl -channel blocker NPPB
(100 µM). Restoration of isotonicity at the end of the experiment
largely reversed the swelling-activated changes in current.
I, current amplitude.
|
|
Figures 8 and 9 present the corresponding
difference currents. The swelling-activated currents were separately
calculated as the hypotonic minus the isotonic currents in NaCl, NaAsp,
and NaMeth solutions. The NPPB-difference currents were the hypotonic NaCl currents without NPPB minus those with NPPB. The time courses of
the difference currents triggered by voltage step changes (Fig. 8)
displayed the outward rectification and inactivation at highly depolarizing voltages characteristic of swelling-activated
Cl
currents (37).

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Fig. 8.
Swelling-activated difference currents following step changes in
voltage. Differences were calculated from the experiment shown in Fig.
9 as the hypotonic currents minus isotonic currents measured in
Cl (A), methylsulfonate (B), and
aspartate (C) solutions. NPPB difference currents
(D) were calculated as the hypotonic currents in
Cl solution without NPPB minus those currents measured in
its presence.
|
|

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Fig. 9.
Current-voltage (I-V) relationships of
swelling-activated difference currents and NPPB difference currents.
Values are means ± SE and were calculated from the currents
measured 20-30 ms after application of voltage steps [n
=4; hTM no. 22 (P2-3)]. The reversal potentials for the
swelling-activated currents in methylsulfonate and aspartate solutions
were shifted from those for the swelling-activated currents in
Cl solution and the NPPB difference currents in hypotonic
Cl solution.
|
|
As illustrated by current-voltage plots in Fig.
9 (n = 4), the
difference currents for the swelling-activated channels perfused in
Cl
, aspartate, and methylsulfonate Ringer's
solutions and for the NPPB-inhibited channels were all well fit by the
Goldman equation for Cl
channel currents (MATERIALS
AND METHODS, Eq. 1). From the nonlinear least-squares
fit, the Erev values for the swelling-activated difference currents were
35.1 ± 1.2,
10.1 ± 0.7, and
22.2 ± 0.9 mV in the NaCl, NaAsp, and NaMeth bath solutions
(Table 2), respectively. Correcting for junction potential, the
Erev for the swelling-activated currents in
Cl
-Ringer's perfusate was
37.9 mV. From this value and
the known anion concentrations in the micropipette and bath,
PAsp/PCl is calculated to be 0.019 with the use of the Goldman equation in the form
|
(5)
|
Perfusion with the NaMeth bath solution (Table 2) shifted
Erev by 12.9 mV. From this shift
(
Erev), and by inserting the values of the
anionic concentrations into the following expression of the Goldman
equation, PMeth/PCl is
estimated to be 0.50
|
(6)
|
where
|
(7)
|
Thus Cl
channels of human TM cells display a lower
permeability for methylsulfonate than for Cl
, consistent
with the observation (Fig. 6C) that methylsulfonate substitution for external Cl
triggers cell shrinkage.
Intracellular Ca2+.
A regulatory volume decrease has been observed with all the approaches
we used, including conventional electronic cell sorting (Fig.
2B) and calcein fluorometry of both cells grown on
coverslips (Figs. 2, 4B, 5, and 6) and freshly harvested
cells in the presence (Fig. 4A) or absence (Figs. 1 and
3A) of BDM. A swelling-triggered rise in intracellular
Ca2+ is thought to be of importance in triggering the RVD
(and also the apoptotic volume decrease) of some cells
(39) but not of many others (27). In
addition, BDM has been reported to alter Ca2+ kinetics in
some cells (3, 53). These issues were addressed by
monitoring intracellular Ca2+ activity during perfusion
with hypotonic solution containing or free of BDM.
Taking the ratio of fura 2 fluorescence at 340 to 380 nm as an index of
Ca2+ activity, BDM triggered a paired increase of 0.06 ± 0.01 [n = 10, hTM no. 22 (P3-4)] to an isotonic
level of 0.66 ± 0.02 from an isotonic baseline of 0.60 ± 0.02 in BDM-nontreated cells. The BDM also reduced the
swelling-activated increase in the ratio from 0.07 ± 0.02 (n = 10) in BDM-nontreated cells to 0.01 ± 0.02 in
BDM-exposed cells, a mean paired shift of 0.06 ± 0.03. In those experiments that included fluorescence calibration, BDM increased baseline Ca2+ concentration (Fig.
10A) from 41 ± 10 nM
in control cells to 85 ± 13 nM and abolished the
swelling-activated stimulation in Ca2+ concentration
measured to be 27 ± 16 nM in control cells. Measured at the same
time (14.3 min) after hypotonic perfusion was initiated, a change in
Ca2+ concentration of
6 ± 11 nM was displayed in
the BDM-treated cells. Because BDM did not alter the RVD (cf. Figs.
2A and 4), a spike in Ca2+ activity was not
necessary for the regulatory response of human TM cells, as noted with
many other cells (27).

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Fig. 10.
Effects of hypotonicity and BDM on intracellular
Ca2+ concentration. A: effects of hypotonic
perfusion in the absence [n = 5; hTM no. 22 (P4)] and
presence [n = 6; hTM no. 22 (P4)] of 20 mM BDM. Values are
means ± SE. B: effect of isosmotic perfusion with 20 mM BDM [n = 4; hTM no. 22 (P4)].
|
|
The effect of BDM on intracellular Ca2+ concentration was
also tested under isotonic conditions (Fig. 10B). The
actomyosin antagonist acutely and reversibly increased Ca2+
concentration by 34 ± 7 nM from 59 ± 6 to 92 ± 8 nM
(n = 4), consistent with the elevated baseline
Ca2+ level observed with BDM-treated cells in the
experiments of Fig. 10A.
 |
DISCUSSION |
Salient observations.
The major findings of the present work are that 1) human TM
cells display an RVD; 2) the RVD is mediated at least in
part by swelling-activated Cl
channels and possibly
K+-Cl
symports; 3) the
swelling-activated Cl
channels are selective for
Cl
> methylsulfonate > aspartate;
4) swelling triggers an increase in intracellular
Ca2+ that is not causally related to the RVD; and
5) Na+/H+ antiports are important
determinants of isotonic TM cell volume in the presence of
physiological concentrations of extracellular HCO
.
Methodology.
TM cell volume has been previously studied with morphological
approaches (15), electronic cell sorting
(36), radioisotopic markers (36), and forward
light scatter (49). Alternative electrophysiological and
optical methods are also available (4). We regard
electronic cell sorting as the volumetric technique of choice in
studying ocular epithelial cells, and this approach can be applied to
human TM cells (Fig. 2B). However, the need for large
numbers of cells largely limits this technique primarily to bovine
preparations (36). We also have extensive experience with
the ion-selective microelectrodes used for this purpose [see chapter 5 in Ref. 8], but this electrophysiological approach is too
labor intensive to permit rapid sampling of multiple cells within a
potentially heterogeneous population. In addition, estimations of
intracellular volume from radioisotope determinations of total water
and extracellular volume depend on assumptions of volumes of
distribution and can involve modest differences between two large
numbers. Given these considerations, we have opted to monitor changes
in volume with fluorescent probes.
We conducted preliminary measurements with several fluorophores,
including calcein, Oregon green, MQAE, fluorescein, and fura 2. The
quenching of both MQAE (47) and SPQ (46) is
inversely dependent on cell volume, but dye leakage and the need to
work in Cl
-free solution limit their applicability. Fura
2 could be used to monitor projected cell area (29), but
the dependence of fluorescence on Ca2+ activity can be a
confounding factor even at the presumed isosbestic point because of
shifts in the isosbestic frequency. Of the dyes tested, calcein proved
most satisfactory. The fluorescence of calcein is independent of
intracellular composition and is two to three times greater than that
of other commonly used fluorophores (4). At room
temperature, dye leakage and bleaching generally reduced the total
signal intensity by only a few percent over periods of ~70 min. At
37°C, leakage was observed in a few experiments, so the results
presented were conducted at room temperature.
The calcein fluorescence has been used to monitor cell area as a
semiquantitative index of cell volume. Depending on the geometry of the
adherent cell and the adhesion between cell and coverslip, changes in
cell volume may not be proportionately expressed along the axes
parallel and perpendicular to the supporting surface. For this reason,
we have used the same cells as their own series controls in the context
of several different protocols. A potentially more serious problem is
posed by the difficulty in distinguishing between changes in
contractile state and changes in volume. For this reason, we studied
cells in the presence and absence of an antagonist (BDM) of actomyosin
function. BDM markedly reduced the probability of both rhythmic
contractions and oscillations and also motility of the cells out of the
field of study.
Transporters regulating human TM cell volume.
Applying the calcein fluorescence technique, we have documented that
anisosmotic swelling triggers an RVD, independent of whether the human
TM cells are grown on coverslips or freshly harvested, in the presence
or absence of BDM. This RVD was qualitatively similar to that observed
with electronic cell sorting of the same cells in suspension. The RVD
could be inhibited by applying NPPB, suggesting the operation of
swelling-activated Cl
channels. In addition, we found
that DIOA inhibited the RVD, suggesting that Cl
may also
be released through a K+-Cl
symport.
In some cells, swelling-activated rises in intracellular
Ca2+ are thought to be an important element in the
signaling cascade leading to the expression of the RVD
(39). In the present study, the actions of BDM to elevate
baseline isotonic intracellular Ca2+ and block the
swelling-triggered increase in Ca2+ did not alter the RVD.
Thus the RVD of human TM cells appears to be independent of changes in
Ca2+ concentration, as has been noted with many other cells
(27). The observed actions of BDM are consistent with
prior reports that BDM both activates Ca2+- release from
intracellular stores in cardiac and skeletal muscle (53)
and inhibits plasma-membrane L-type Ca2+-channel activity
of other cells (3).
Previous studies have focused on the operation of the TM cell
Na+-K+-2Cl
symport under
conditions of low bicarbonate concentration (4.2 mM) at 37°C
(36, 41, 42). At physiological bicarbonate concentration at room temperature, we have observed that the selective
Na+/H+ antiport inhibitor DMA causes TM cell
shrinkage. Thus the relative contribution by the
Na+-K+-2Cl
symports and paired
Na+/H+ and
Cl
/HCO
antiports might depend on the level of bicarbonate. This suggests that human TM cells display at
least four regulatory volume transporters:
K+-Cl
symport and Cl
channel
release pathways, and Na+/H+ antiport and
Na+-K+-2Cl
symport uptake
pathways. The Na+/H+ antiport presumably
functions in parallel with a Cl
/HCO
exchanger to regulate volume, as we have previously noted with
pigmented ciliary epithelial cells (12).
We verified electrophysiologically that swelling activates human TM
cell NPPB-sensitive Cl
channels. The channels display a
permeability ranking of Cl
> methylsulfonate > aspartate, consistent with our observation that replacement of
external Cl
with methylsulfonate produced cell shrinking.
Why methylsulfonate-triggered shrinkage was not observed in an earlier
study (42), despite loss of cell Cl
, is
unclear but might have reflected differences in methodology or the
generation of intracellular osmolytes at the higher temperature with
the lines of TM cells used.
The volume of TM and juxtacanalicular cells may be a determinant of
outflow resistance from aqueous humor to Schlemm's canal (2, 15,
18, 19, 36, 41, 58). The current work presents new information,
both methodological and physiological, in addressing cell volume
regulation of cells derived from TM and juxtacanalicular cells. The use
of a heterogeneous population of cultured cells complicates the
extrapolation of these results to the distinct cellular types observed
in vivo, particularly because changes in juxtacanalicular cell volume
are expected to have a predominant effect on outflow. However, the
consistency in the responses of individual cells from this mixed
population to the osmotic and pharmacological modifiers of cell volume
does strengthen the implications. Of particular potential relevance is
the observation that DMA causes isotonic cell shrinkage, raising the
possibility that Na+/H+ antiport inhibitors
might increase outflow facility. Thus the recent finding that DMA
lowers intraocular pressure in mice (5a) may reflect not
only a reduction in aqueous humor production at the level of the
pigmented ciliary epithelial cells (12, 33) but also a
reduced resistance to outflow of aqueous humor through the TM.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Kenneth R. Spring for extremely helpful and
stimulating conversations.
 |
FOOTNOTES |
This work was supported in part by research National Eye Institute
Grants EY-013624 (M. M. Civan), EY-12797 (W. D. Stamer), EY-10009 (C. H. Mitchell), and Core Grant EY-01583 (C. H. Mitchell, M. M. Civan), a Research to Prevent Blindness Career
Development Award (W. D. Stamer), and fellowships (J. C. Fleischhauer) from the Swiss National Science Foundation Fellowship
(no. 1037) and The Alfred Vogt Foundation Fellowship, Switzerland.
Address for reprint requests and other correspondence:
M. M. Civan, Depts. of Physiology and Medicine, Univ. of
Pennsylvania, A303 Richards Bldg., Philadelphia, PA 19104-6085 (E-mail:
civan{at}mail.med.upenn.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 February 13, 2002;10.1152/ajpcell.00544.2001
Received 14 November 2001; accepted in final form 6 February 2002.
 |
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