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Am J Physiol Cell Physiol 283: C315-C326, 2002. First published February 13, 2002; doi:10.1152/ajpcell.00544.2001
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Vol. 283, Issue 1, C315-C326, July 2002

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
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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<UP><SUB>3</SUB><SUP>−</SUP></UP> 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


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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<UP><SUB>3</SUB><SUP>−</SUP></UP> concentration (4.2 mM) (36). In the present study, we tested whether paired Na+/H+ and Cl-/HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> 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.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 alpha 2-adrenergic (48) and prostaglandin F2alpha 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.

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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  		Table 1.   Solutions for measurements of volume and intracellular Ca2+

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 MOmega ; successful seals displayed gigaohm resistances. After rupture of the membrane patch, the series resistance was measured to be only 8.0 ± 0.9 MOmega 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.

                              
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  		Table 2.   Solutions for patch-clamp experiments

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
&Dgr;I<SUB>Cl</SUB><IT>=K · &bgr; · </IT>(<IT>e</IT><SUP>−&bgr;rev</SUP><IT>−e</IT><SUP>−<IT>&bgr;</IT></SUP>)<IT>/</IT>(1<IT>−e</IT><SUP>−<IT>&bgr;</IT></SUP>) (1)
where
<IT>K</IT> ≡ P<SUB>Cl</SUB> · <IT>F · </IT>([Cl<SUP>−</SUP>]<SUB>i</SUB> + [aspartate<SUP>−</SUP>]<SUB>i</SUB><IT> · P</IT><SUB>Asp</SUB><IT>/P</IT><SUB>Cl</SUB>) (2)

&bgr;≡(V<SUB>m</SUB><IT> · F</IT>)<IT>/</IT>(<IT>R · T</IT>) (3)

&bgr;<SUB>rev</SUB><IT>≡</IT>(<IT>E</IT><SUB>rev</SUB><IT> · F</IT>)<IT>/</IT>(<IT>R · T</IT>) (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
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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)].

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.

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<UP><SUB>3</SUB><SUP>−</SUP></UP> antiport exchange and large-conductance Ca2+-activated K+ channels (BK channels), in addition to inhibiting K+-Cl- cotransport (17). Blocking Cl-/HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> 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<UP><SUB>3</SUB><SUP>−</SUP></UP> 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<UP><SUB>3</SUB><SUP>−</SUP></UP> 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<UP><SUB>3</SUB><SUP>−</SUP></UP>. 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<UP><SUB>3</SUB><SUP>−</SUP></UP> removal on isotonic uptake mechanisms. With 0 HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>, 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<UP><SUB>3</SUB><SUP>−</SUP></UP> and 0.5 mM DIDS, 10 µM DMA lead to a reversible decrease in pHi [hTM no. 36 and no. 25 (P3), n = 4].

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<UP><SUB>3</SUB><SUP>−</SUP></UP> and 0.5 mM DIDS to limit potential confounding contributions of Cl-/HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> exchange and Na+-nHCO<UP><SUB>3</SUB><SUP>−</SUP></UP> 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<UP><SUB>3</SUB><SUP>−</SUP></UP> 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
P<SUB>Asp</SUB><IT>/P</IT><SUB>Cl</SUB><IT>=</IT>([Cl<SUP>−</SUP>]<SUB>o</SUB> · <IT>e</IT><SUP>−&bgr;rev</SUP> − [Cl<SUP>−</SUP>]<SUB>i</SUB>)/[aspartate<SUP>−</SUP>]<SUB>i</SUB> (5)
Perfusion with the NaMeth bath solution (Table 2) shifted Erev by 12.9 mV. From this shift (Delta 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 
P<SUB>Meth</SUB>/<IT>P</IT><SUB>Cl</SUB><IT>=</IT>[(116)<IT> · e</IT><SUP>−&Dgr;&bgr;rev</SUP> − 25]/91 (6)
where
&Dgr;&bgr;<SUB>rev</SUB> ≡ (&Dgr;<IT>E</IT><SUB>rev</SUB> · <IT>F</IT>)/(<IT>R</IT> · <IT>T</IT>) (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
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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<UP><SUB>3</SUB><SUP>−</SUP></UP>.

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<UP><SUB>3</SUB><SUP>−</SUP></UP> 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<UP><SUB>3</SUB><SUP>−</SUP></UP> 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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Am J Physiol Cell Physiol 283(1):C315-C326
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