Modulation of Aqueous Humor Outflow by Ionic Mechanisms Involved in Trabecular Meshwork Cell Volume Regulation

Abstract

purpose. Trabecular meshwork (TM) cell shape, volume, contractility and their interactions with extracellular matrix determine outflow facility. Because cell volume seems essential to TM function, this study was conducted to investigate further the ionic channels and receptors involved in regulatory volume decrease and their roles in modulating outflow facility.

methods. Primary cultures of bovine TM cells were used. K⁺ and Cl⁻ currents were studied with whole-cell patch clamping. Swelling was induced by hypotonic shock. [Ca²⁺]_i was measured in TM cells loaded with fura-2. Bovine anterior segments were perfused at constant pressure to measure outflow facility.

results. Hypotonic media activated both the high-conductance Ca²⁺-activated K⁺ channel (BK_Ca) and swelling-activated Cl⁻ channel (Cl_swell) currents and induced release of adenosine 5′-triphosphate (ATP) from TM cells. ATP activated P2Y₂ receptors with the following profile: ATP = uridine 5′-triphosphate (UTP) > adenosine 5′-O-(3-thiotriphosphate) (ATP-γ S) > adenosine 5′-diphosphate (ADP) = uridine 5′-diphosphate (UDP), and increased BK_Ca current. Hypotonic medium initially decreased outflow facility in perfused anterior segments, which recovered with time to baseline levels. Addition of tamoxifen or iberiotoxin (Cl_swell and BK_Ca blockers, respectively) lengthened the recovery phase, which implies that these channels participate in cell volume regulation. In contrast, an activator of BK_Cas (NS1619) produced the opposite effect.

conclusions. Cell swelling activates a regulatory volume decrease mechanism that implies activation of K⁺ and Cl⁻ currents and participation of P2Y₂ receptors. Because previous studies have shown that intracellular volume of TM cells is an important determinant of outflow facility, it seems feasible that cell volume regulation would be part of the homeostatic mechanisms of the TM, to regulate the outflow pathway.

In humans and primates, most of the aqueous humor (AH) produced by the ciliary body flows through the trabecular meshwork (TM) and the Schlemm’s canal endothelium to reach the aqueous veins, exiting the eye. The correct performance of this route, as well as the uveoscleral one, is determinant to maintain intraocular pressure (IOP) in the physiological range. On the contrary, when the outflow is somehow impaired, the increase in AH volume raises IOP, which is a major risk factor for the development of glaucoma. TM physiology is still largely unknown, but it has been proposed that TM cells modulate the pathway permeability by showing volume and contractile responses. ^{1 2 3} Several mechanisms have been proposed to play a role in TM function, including contraction/relaxation of both TM and ciliary muscle ^{4 5} ; stretch, ⁶ ; composition and remodeling of the extracellular matrix ^{7 8 9} ; pore formation in the Schlemm’s canal endothelium ^{10 11} ; and changes in gene expression, ^{12 13 14} cell shape, ^{6 7 15 16} or cell volume. ^{2 17} Endothelial cells of Schlemm’s canal may also participate in the regulation of outflow facility, showing some of these mechanisms, especially in monkey and human eyes, where Schlemm’s canal has a higher degree of organization. ¹⁸  

TM cell volume appears to be an important determinant of tissue function, since swelling or shrinking the cells modifies AH outflow facility. ^{2 3} Mitchell et al. ¹⁹ recently showed that TM cells possess a regulatory volume decrease (RVD) mechanism. Thus, after cell swelling, the RVD is mediated, at least in part, by activation of swelling-activated Cl⁻ channels (Cl_swells) and K⁺-Cl⁻ symports. In this sense, the Na⁺/H⁺ antiport has also been suggested to participate in volume regulation, and therefore inhibitors of this antiport may increase outflow facility. ¹⁹ Another player is the Na⁺-K⁺-Cl⁻ cotransport that would modulate outflow facility by regulating cell volume in response to different conditions-drugs, ^{2 17} but bumetanide (an Na⁺-K⁺-Cl⁻ cotransport blocker) unexpectedly did not have any effect on outflow facility in monkeys. ²⁰ Moreover, bumetanide did not reduce IOP in monkey or mouse. ^{20 21} Despite all the players identified and possibly involved in cell volume regulation, a direct relationship between cell volume regulation and outflow facility has not yet been demonstrated. 

In other cell types, ^{22 23 24} osmotic cell swelling is thought to activate Cl_swells. Simultaneously, cell swelling induces the release of adenosine 5′-triphosphate (ATP) that, in turn, acts as an autocrine/paracrine signal to activate purinergic P2Y receptors. Stimulation of purinergic receptors is thought to facilitate activation of K⁺ channels. Finally, Cl⁻ and K⁺ efflux from the cell followed by water outflow permits the cell to recover its volume. ²⁵ It remains to be elucidated whether these mechanisms of cell volume regulation are essential to TM function. In this sense, several mechanisms such as the Na⁺-K⁺-Cl⁻ cotransport ¹⁷ or the high-conductance Ca²⁺-activated K⁺ channel (BK_Ca) ²⁶ are tightly modulated by several drugs, and alterations in their regulation may affect TM function and therefore outflow facility. Our purpose was to identify and characterize the receptors and channels involved in the RVD of TM cells and to determine their functional role in outflow facility modulation as part of an homeostatic system that may regulate TM resistance. We used hypotonic shocks to trigger RVD and to study the receptors and channels involved in this process. Our findings show that changes in TM cell volume modify outflow facility and that regulatory mechanisms to restore cell volume actively participate to maintain AH outflow. 

Methods

Bovine TM Cell Culture

Bovine TM (BTM) cells were cultured by using a modification of the technique described by Stamer et al. ²⁷ As described previously, ⁴ BTM strips were digested with 2 mg/mL collagenase (Sigma-Aldrich, Madrid, Spain) and 0.5 mg/mL bovine serum albumin (BSA; Sigma-Aldrich) at 37°C for 2 hours. After mechanical digestion, the supernatant was collected, centrifuged, and resuspended. The resuspended solution was seeded in culture flasks containing Dulbecco’s modified Eagle’s medium (DMEM; BioWhitaker, Barcelona, Spain) plus 10% fetal bovine serum, 100 mg/mL l-glutamine (Sigma-Aldrich), 100 UI/mL penicillin, 100 μg/mL streptomycin, and 2.5 μg/mL amphotericin B (BioWhitaker). Cell growth was observed 2 to 4 days after seeding and cells reached confluence 12 to 15 days later. Cell passages were performed using trypsin-EDTA (Bio-Whitaker). Cells from passages 1 to 3 were used for electrophysiology, [Ca²⁺]_i and ATP measurements. 

Cytosolic Free-Ca²⁺ Measurement

Measurement of [Ca²⁺]_i was performed as described in detail previously. ^{4 28} Briefly, BTM cells were plated on 25-mm diameter glass coverslips (VWR Scientific Inc., Philadelphia, PA) and then loaded with 5 μM fura-2/AM (Calbiochem, San Diego, CA) for 60 minutes at 37°C in incubation buffer (121 mM NaCl, 4.7 mM KCl, 5 mM NaHCO₃, 1.2 mM KH₂PO₄, 1.2 mM MgSO₄, 2 mM CaCl₂, 10 mM glucose, 10 mM HEPES, and 0.01% BSA [pH 7.4], adjusted with NaOH, 287± 2 mOsm/kg; mean ± SD). Coverslips with fura-2–loaded cells were transferred into an open flow chamber (1 mL incubation buffer) mounted on the heated stage of an inverted epifluorescence microscope (Diaphot-300; Nikon, Tokyo, Japan). Fluorescence images were obtained by a charge-coupled device (CCD) camera (CH250; Photometrics, Tucson, AZ) and were digitized, stored, and analyzed on computer (Macintosh 840AV; Apple, Inc., Cupertino, CA). After a stabilization period of 10 minutes, image pairs were obtained alternately every 4 seconds, and, for a total of 8 minutes, at excitation wavelengths of 340 and 380 nm (10-nm bandwidth filters), to excite the Ca²⁺-bound and Ca²⁺-free forms of this ratiometric dye, respectively. The emission wavelength was 510 nm (120-nm bandwidth filter). [Ca²⁺]_i levels were calculated in single cells by determining the 340- to 380-nm fluorescence ratios at each time point. ^{4 29} In both control and experimental groups, Ca²⁺ was recorded for 1 minute before drug application and thereafter, for 7 minutes. Cells were considered as responders when [Ca²⁺]_i increased more than 100% above the resting value. 

Patch-Clamp Procedures

TM cells were plated onto small glass coverslips and studied 24 to 48 hours thereafter. Coverslips were transferred to a special chamber (0.2 mL) in the stage of an inverted microscope (IX70; Olympus, Tokyo, Japan) to perform the recordings. External solutions were superfused at a rate of 3 to 4 mL/min by gravity. Before the recording session was started, culture medium was replaced with a bath recording solution. Recordings were performed in 2-hour sessions at 21°C to 23°C. Patch pipettes were pulled in an electrode puller (P-97; Sutter Instruments, Novato, CA) and had a filled tip resistance of between 3 to 6 MΩ. Pipette capacitance to ground was neutralized after the seal was formed. An Ag/AgCl wire bath electrode was used except when recording chloride currents where the Ag/AgCl electrode was mounted in a 3 M KCl agar bridge. As previously described, ⁶ we took care to avoid stretch activation of BK_Cas when performing the patches. Before the pipette had entered the bath and until cell contact, positive pressure was applied. 

Whole-cell currents were recorded using a patch-clamp amplifier (L/M-EPC7; Heka, Lambrecht/Pfalz, Germany) as described previously. ⁶ Data acquisition and command potentials were controlled by computer (WinWCP 3.2 software; John Dempster, University of Strathclyde, UK), using an interface (CED1401; Cambridge Electronic Design Ltd., Cambridge, UK). Whole-cell currents were recorded at 10 kHz. 

Chloride Currents

Cl_swell currents were recorded using the whole-cell configuration of the patch–clamp technique. After achieving the whole-cell configuration, cells were allowed to stabilize and dialyze for 3 to 4 minutes before recording started. Whole-cell currents were recorded at 10 kHz without any leak current subtraction protocol. Cells were clamped at 0 mV, and voltage pulses from −80 to +120 mV were applied in 20-mV steps to evoke Cl⁻ currents. The intracellular pipette solution contained (in mM): 150 N-methyl-d-glucamine (NMDG)Cl, 1.2 MgCl₂, 1.0 EGTA, 2 ATP, 0.5 guanosine 5′-triphosphate (GTP), 10 HEPES (pH 7.35) adjusted with Tris (303 ± 4 mOsm/kg; mean ± SD). The standard isotonic bath solution contained (in mM): 150 NMDGCl, 0.5 MgCl₂, 1.3 CaCl₂, and 10 HEPES (pH 7.35) adjusted with Tris (295 ± 5 mOsm/kg). The standard hyposmotic bath solution contained (in mM): 100 NMDGCl, 0.5 MgCl₂, 1.3 CaCl₂, and 10 HEPES (pH 7.35) adjusted with Tris (219 ± 5 mOsm/kg). The solution osmolalities were measured with an osmometer (Vapro, Wescor, UT). To study channel-relative permeabilities, we recorded voltage ramps from −100 to +100 mV for Cl⁻, I⁻, Br⁻, F⁻, and gluconate⁻. In these experiments, the bath hypotonic solution used was (in mM): 90 NaX (X = anion), 10 NMDGCl, 0.5 MgCl₂, 1.3 CaCl₂, and 10 HEPES (pH 7.35) adjusted with 0.5 M Tris. Data were corrected with the calculated theoretical junction potential values for each solution. 

Potassium Currents

Potassium currents were recorded in whole-cell configuration, either with the ruptured-patch or the perforated-patch (with Nystatin; Sigma-Aldrich) technique. Leak current and residual capacitative current were subtracted using a P/N protocol where N was −4. Cells were clamped at −60 mV, and depolarizing pulses were applied in 10- or 20-mV steps to evoke outward K⁺ currents. In whole-cell recordings, the solution in the pipette was (in mM): 140 KCl, 2 MgCl₂, 0.1 EGTA, and 10 HEPES (pH 7.2), with 0.5 M KOH. The bath physiological solution was (in mM): 140 NaCl, 4.3 KCl, 1.3 CaCl₂, 1 MgCl₂, and 10 HEPES (pH 7.4), with NaOH (301 ± 5 mOsm/kg). Hypotonic solution in which NaCl and KCl concentrations were reduced was (in mM): 100 NaCl, 2 KCl, 1.3 CaCl₂, 1 MgCl₂, and 10 HEPES (pH 7.4), with NaOH (213 ± 4 mOsm/kg; mean ± SD). Outward potassium currents were blocked with a specific BK_Ca blocker, iberiotoxin (IBTX, 50 nM) obtained from Sigma-Aldrich (St. Louis, MO). In the whole-cell perforated-patch technique, pipettes were backfilled with intracellular solution containing (in mM): 100 K-acetate, 40 KCl, 1 MgCl₂, 1 CaCl₂, 1.38 EGTA, and 10 HEPES (pH 7.2) with KOH, in which Nystatin (150 μg/mL) was dissolved. The solution was made daily from a stock solution (3 mg/60 mL dimethyl sulfoxide [DMSO]) and sonicated to the final concentration. Slowly developing capacitative transients indicated establishment of the whole-cell perforated-patch configuration. We continuously monitored series resistance to avoid sudden drops in this parameter indicating that ruptured-patch configuration was achieved. Only experiments with constant series resistance during the whole recording were considered for analysis. 

Measurement of ATP Release

ATP release detection was performed in cultured TM cells 3 to 4 days after cell passage. Experiments were performed at 22°C in cells grown in 35-mm Petri dishes. Culture medium was replaced by physiological solution (mM): 140 NaCl, 4.3 KCl, 1.3 CaCl₂, 1 MgCl₂, 10 HEPES, and 10 glucose (pH 7.4), with NaOH (291 ± 2.9 mOsm/kg) and allowed to rest for 30 minutes to minimize a possible ATP release due to medium exchange. Twenty percent and 40% hypotonic medium was obtained by adding distilled water to the physiological solution and resulting osmolalities were 242 ± 2 and 206 ± 2 mOsm/kg (mean ± SD), respectively. A vial of firefly lantern luciferase extract from Photinus pyralis (Sigma-Aldrich) was diluted in 1 mL of isotonic physiological solution or distilled water containing 5 mg/mL d-luciferin (Sigma-Aldrich). The resultiant suspension was centrifuged for 30 seconds in a benchtop centrifuge. The supernatant was desalted in a 10-mL column (10 DG; Bio-Rad, Hercules, CA) equilibrated with recording solution. A photomultiplier tube (R464; Hamamatsu, Hamamatsu City, Japan) located over the Petri dish was used to detect photons. The resultant electric signal was filtered at 1 kHz, digitized, and recorded on computer (X-Chart software; Heka). Calibration curves were calculated by adding known doses of ATP (1, 3, 4, and 6 picomoles). Regardless of the different ionic strength of the isotonic and hypotonic solutions used, no significant differences were found between the calibration curves in the 1- to 6-picomole range. Therefore, data were not corrected for each solution and all calculations were done with the calibration values in isotonic conditions. Also, to test a possible lysis due to excessive cell swelling when hypotonic solutions were added, we measured lactate dehydrogenase (LDH) activity in the bath solution. ³⁰ No significant amounts of LDH activity were encountered before and after addition of isotonic or hypotonic solutions. 

Perfusion of Anterior Segments

Eyes of 3- to 6-month-old calves were obtained at the local abattoir 0.5 to 2 hours after death and kept in phosphate-buffered saline (PBS) at 4°C for not more than 1.5 hours. Isolation of bovine anterior segments was performed as described previously. ³ The perfusion technique has been described elsewhere. ^{3 4} Briefly, bovine anterior segments were placed in a specially designed perfusion chamber. The anterior segments, located in their respective chambers together with force transducers (Letica, Barcelona, Spain) and the tubing system, were placed in an incubator (Selecta, Barcelona, Spain) at 36°C and 5% CO₂. Perfusion was performed with DMEM. The pressure of the artificial anterior chamber was monitored and recorded throughout the experiment with a pressure transducer (9162-0; Mallinckrodt, Northampton, UK) and was maintained with a suspended reservoir at 10 mm Hg in bovine eyes. Outflow facility (C) was averaged during periods of 15 minutes (mean of 450 values). Baseline facility (C ₀) was calculated during the first 90-minute period of stable C. When a different experimental condition or drug was added to the perfusion medium, the tubes and the anterior chamber were flushed and replaced with the new medium. This change was made by rapidly replacing the contents of the artificial anterior chamber by opening the exit needle until 200% of the volume had been exchanged. This exchange was always made at a pressure below 10 mm Hg. Recording of C started after stabilization of the flow. 

The perfusion procedure was performed using a protocol with two periods: perfusion with control isotonic DMEM (301 ± 1 mOsm/kg) for 90 minutes, to establish the C ₀, and perfusion with hypotonic (244 ± 2 mOsm/kg) DMEM for 360 minutes to determine changes in outflow facility after five experimental conditions: hypotonic medium, hypotonic+tamoxifen, hypotonic+IBTX, hypotonic+tamoxifen+IBTX, and hypotonic+BK_Ca channel opener (NS1619; 1,3-dihydro-1-[2-hydroxy-5-(trifluoromethyl)phenyl]-5-(trifluoromethyl)-2H-benzimidazol-2–1)). 

Drugs

ATP, uridine 5′-triphosphate (UTP), adenosine 5′-O-(3-thiotriphosphate) (ATP-γ S), adenosine 5′-diphosphate (ADP), uridine 5′-diphosphate (UDP), suramin sodium salt, 2-MeS-ATP (2-methylthio adenosine 5′-triphosphate tetrasodium salt), thapsigargin, EGTA, U73122 (1-[6-[((17β)-3-methoxyestra-1,3,5(10)-trien-17-yl)amino]hexyl]-1H-pyrrole-2,5-dione), ryanodine, cyclopiazonic acid (CPA), 4,4′-diisothiocyanatostilbene-2,2′-disulfonic acid disodium salt hydrate (DIDS), NMDG, tamoxifen ((Z)-1-(p-dimethylaminoethoxyphenyl)-1,2-diphenyl-1-butene), IBTX, firefly lantern extract (luciferin-luciferase), the BK_Ca channel opener NS1619, nystatin, and ARL67156, were obtained from Sigma-Aldrich. PPADS (pyridoxal-phosphate-6-azophenyl-2′,4′-disulfonic acid) was purchased from Calbiochem. 

Data Analysis

Results are given as the mean ± SEM. Data were analyzed with the paired or unpaired Student’s t-test, the Fisher exact test, and the χ² test. Two-way ANOVA with the Bonferroni posttest was used to evaluate differences between control and drug effects in the recorded currents. P < 0.05 was considered statistically significant. 

Results

Modulation of K⁺ and Cl⁻ Currents in the TM by Cell Swelling

It has been reported that hypotonicity triggers an RVD ¹⁹ in TM cells and that it also decreases outflow facility in perfused anterior segments. ^{2 3} In the present study, we further examined the effects of hypotonic stimuli on whole-cell currents mediated by the BK_Ca ⁶ and the possible activation of Cl_swell in TM cells. As shown in Figure 1A , whole-cell currents were first recorded in isotonic physiological solution (control) and afterward bath solution was replaced by hypotonic medium. Cells were clamped at −60 mV, and depolarizing pulses to +80 mV were applied in 10-mV steps to evoke outward BK_Ca currents. In hypotonic medium, the total outward K⁺ current increased significantly (ANOVA; P < 0.001) in 13 of 15 whole-cell experiments (87% of the cells). Bonferroni posttests showed significant differences between the isotonic and hypotonic groups, at −10 mV (P < 0.05) and from 0 to +80 mV (P < 0.001). Two of 15 cells did not show significant changes. Figure 1A shows a typical recording and the current-voltage (I-V) relationship in isotonic and hypotonic conditions. In agreement with previous data, ^{6 19} hypotonic solution did not modify [Ca²⁺]_I; thus, it is not likely that the BK_Ca current increase was due to an increase in [Ca²⁺]_i. 

In a different set of experiments, cells were voltage clamped at 0 mV, and whole-cell currents were elicited by using specific solutions to record Cl⁻ currents (see the Methods section). Under isotonic conditions, TM cells possessed a small current of −3.0 ± 1.3 pA/pF (mean ± SEM) at an applied potential of −80 mV and 2.8 ± 1.2 pA/pF at +80 mV (Fig. 1B) . When isotonic medium was replaced by a hypotonic one, Cl_swell currents were activated within 4 minutes and reached a maximum between 10 and 12 minutes (−8.2 ± 2.1 pA/pF at −80 mV and 14.4 ± 4.3 pA/pF at +80 mV; mean ± SEM). Accounting all the cells recorded in the whole study, Cl_swell current activation was seen in 140 (85%) of 164 cells. In the experimental group shown in Figure 1B , cells in isotonic medium (n = 5) were first exposed to hypotonic medium and, on maximum Cl_swell current activation, hypotonic medium was replaced, returning to isotonic medium. Statistical comparison of the I-V curves in isotonic and hypotonic conditions showed significant differences for the whole voltage range (−80 to +120 mV; ANOVA, P < 0.001). Bonferroni posttests showed significant differences between the isotonic and hypotonic groups, at +80 mV (P < 0.01) and +120 mV (P < 0.001; Fig. 1B ). When returning to isotonic medium, Cl_swell currents decreased, and significant differences between isotonic and hypotonic conditions were also recorded at +80 mV (P < 0.01) and +120 mV (P < 0.001). All recordings were obtained with ATP and GTP in the intracellular solution. In a group of recordings (n = 5) where these compounds were omitted, no Cl_swell current activation was seen in hypotonic medium (data not shown). This result is in agreement with previous data in other cell types showing the ATP dependency of Cl_swell current activation. ³¹  

As shown in the recordings displayed in Fig. 1B , Cl_swell currents are typically characterized by an outward rectification and a time-dependent decay at depolarizing voltages. To characterize Cl_swell currents further, we replaced Cl⁻ for different anions to establish the channel anion permeability sequence. Figures 2A 2B 2C 2D show mean voltage ramps for each anion after Cl⁻ replacement in the hypotonic solution (SEM was omitted for clarity). Calculation of reversal potential (V _rev) and relative anion permeabilities (P _x/P _Cl; Fig. 2 , table) showed an anion permeability sequence I⁻ > Br⁻ > Cl⁻ > F⁻ > gluconate⁻, which is in agreement with previous data for this channel. ^{32 33 34}  

Pharmacological characterization of Cl_swell currents is shown in Figure 3 . After activation of Cl_swell currents in hypotonic solution, Cl⁻ channel blockers were added to the solution, and their effects at depolarized and hyperpolarized potentials were evaluated. Tamoxifen (100 μM; n = 5), a known selective Cl_swell blocker, decreased outward and inward Cl_swell currents in a voltage-independent manner along the entire I-V relationship (ANOVA P < 0.001, hypotonic versus hypotonic+tamoxifen). As shown in Figure 3A , tamoxifen blocked Cl_swell currents both at hyperpolarized (55% at −80 mV; t-test, P < 0.05 vs. hypotonic) and depolarized potentials (60% at +80 mV; t-test, P < 0.05). Next, we tested the effect of DIDS, a nonselective Cl⁻ channel blocker. By contrast, addition of DIDS (100 μM; n = 6 cells) to the hypotonic medium significantly blocked only the outward currents (ANOVA, I-V relationship hypotonic+DIDS versus hypotonic, P < 0.001). DIDS decreased outward current by 40% (at +80 mV; t-test, P < 0.05), but did not affect inward current (6% at −80 mV; t-test, NS). Blocking effects of both tamoxifen and DIDS are characteristic of Cl_swell currents, as described previously. ³⁵ These results are in agreement with electrophysiological characterization and further corroborate that the currents recorded are mediated by Cl_swell channels. To characterize the currents further, we tested the effect of extracellularly applied ATP (1 mM), which has also been reported to block Cl_swell currents. ³² Extracellular ATP blocked Cl_swell currents in a voltage-dependent manner along the entire I-V curve (ANOVA, P < 0.001; n = 6; Fig. 3C ). Extracellular ATP blocked Cl_swell currents both at hyperpolarized (28% at −80 mV; t-test, P < 0.05 vs. hypotonic) and depolarized potentials (52% at +80 mV; t-test, P < 0.01). 

Cell Swelling’s Effects on ATP Release in TM Cells

It has been reported that ATP can be released to function as an autocrine/paracrine signal in several cell types after osmotic challenge or in response to drug stimulation. ^{25 36} In TM cells grown in 35-mm culture dishes, DMEM was replaced by an isotonic physiological solution (see the Methods section), and cells were allowed to rest for 30 minutes to minimize a possible ATP release due to medium exchange. Stimulation of TM cells with isotonic medium did not elicit a significant release of ATP into the extracellular solution (0.004 ± 0.0006 picomoles or 0.001 ± 0.002 nM; mean ± SEM; n = 3; Figs. 4A 4B ). When medium hypotonicity was reduced by 20%, 2.76 ± 0.18 picomoles of ATP was released (1.15 ± 0.07 nM; t-test P < 0.001 vs. isotonic; n = 5). After reaching a peak, ATP release decreased exponentially to recover baseline levels within 400 to 500 seconds after stimulation. When medium hypotonicity was reduced by 40%, 4.35 ± 0.23 picomoles of ATP was released, showing the same kinetics (1.56 ± 0.08 nM; t-test P < 0.001 vs. isotonic; n = 4). 

Effect of ATP on Purinergic P2Y₂ Receptors in TM Cells

The fact that TM cells release ATP (and maybe other nucleotides) in response to certain stimuli suggests that these same cells may express purinergic receptors to permit a physiological function of this nucleotide in the outflow pathway. Therefore, we tested this hypothesis by stimulating TM cells with different nucleotides while recording intracellular calcium concentration ([Ca²⁺]_i). Resting [Ca²⁺]_i in TM cells was found to be between 58 and 89 nM. In the control group, only 3% of the cells increased [Ca²⁺]_i after drug vehicle application (Table 1) . In contrast, ATP induced significant increases in [Ca²⁺]_i in 99% of TM cells between 10⁻⁶ and 10⁻⁴ M (Table 1 ; Fig. 5 ). At lower concentrations, the percentage of response varied between 2% and 70% of the cells (10⁻¹⁰–10⁻⁷ M; Table 1 ), therefore ATP stimulation followed a sigmoidal dose–response curve with an EC₅₀ of 2.64 × 10⁻⁸ M (Fig. 5H) . Figure 5A shows a typical profile of [Ca²⁺]_i mobilization induced by ATP. [Ca²⁺]_i increased rapidly to reach a peak and then decayed slowly to resting levels. [Ca²⁺]_i peaks followed a dose–response curve with a maximum increase of 587 ± 17 nM at 10⁻⁴ M. To characterize further the receptors involved in the ATP response, we stimulated TM cells with different purinergic agonists and determined the response profile. UTP (1 μM) like ATP at the same concentration, induced [Ca²⁺]_i mobilizations in 99% of the cells (Table 1 , Figs. 5B 5G ). Nevertheless, the diphosphate nucleotides (ADP and UDP; 1 μM) increased [Ca²⁺]_i in only 34% and 33% of the cells, respectively (Fig. 5C 5D 5G ; Table 1 ; P < 0.01 vs. ATP). Also, [Ca²⁺]_i peaks were smaller than the ones elicited by ATP (1 μM). ATP-γ -S (1 μM), a nonhydrolyzable analogue, showed the same results as ATP. 2-MeS-ATP (1 μM), an agonist for P2Y₁ and P2X receptors evoked [Ca²⁺]_i mobilizations in 27% of TM cells (P < 0.001 vs. ATP; Fig. 5G , Table 1 ). Finally, we tested ATP application in the presence of well-known purinergic receptor antagonists (Table 1 ; Figs. 5E 5F 5G ). In the presence of 100 μM suramin, the percentage of response to ATP application decreased to 69% (P < 0.05 vs. ATP alone) while PPADS did not modify the percentage of responding cells to ATP when present in the bath solution. These results show a response profile to the different nucleotides as ATP≈UTP> ATP-γ-S>>ADP≈UDP which suggests the presence of a P2Y₂ purinergic receptor. Moreover, as described previously, ³⁷ P2Y₂ receptors are partially inhibited by suramin but not by PPADS, which argues in favor of the presence of those receptors. Also, the effect of 2-MeS-ATP suggests the presence of P2Y₁ receptors. 

We further examined the intracellular mechanisms involved in the ATP-induced [Ca²⁺]_i increase in TM cells. In nominally Ca²⁺-free solution (+1 mM EGTA), both ATP and ATP-γ-S (1 μM), stimulated the same percentage of cells as ATP in the normal buffer solution (Table 1) . No differences were seen on intracellular Ca²⁺ peak amplitude compared with ATP in the presence of calcium. Nevertheless, recovery phase to resting [Ca²⁺]_i was shorter than with ATP in normal buffer solution (Fig. 6A) . An ATP-induced [Ca²⁺]_i increase was prevented by preincubation of cells with thapsigargin (P < 0.001; Table 1 , Figs. 6A 6B ). A significant decrease in the percentage of cells that had increased [Ca²⁺]_i after ATP exposure was seen in the presence of U73122, an inhibitor of phospholipase C, ³⁸ (P < 0.001) and CPA, a Ca²⁺-ATPase inhibitor, ³⁹ (P < 0.001), but not after preincubation with ryanodine. 

As previously stated, hypotonic stimuli did not modify [Ca²⁺]_i, which is inconsistent with release of ATP. To explore this possibility, we measured [Ca²⁺]_i after a hypotonic stimuli. Hypotonic solution alone (hypo 40%) did not increase [Ca²⁺]_i (129 ± 3 nM; n = 86 cells; Fig. 6C ) in TM cells, but when the experiments were performed in the presence of an inhibitor of the nucelotidases (ARL67156; 100 μM), application of hypotonic solution elicited peaks (418 ± 97 nM; n = 119 cells) similar to the ones obtained with ATP (Fig. 6C) . Ca²⁺ peaks elicited by hypotonic stimuli in the presence of ARL67156 were blocked by suramin (100 μM). 

ATP Increases BK_Ca Current

One of the possible ATP functions after activation of purinergic receptors, in particular, would be to increase K⁺ efflux from the cell thus contributing to cell volume recovery. We performed experiments to test this hypothesis using the whole-cell perforated patch technique, which prevents dialysis of intracellular content. After achieving this configuration, we clamped TM cells at −60 mV and BK_Ca currents were recorded in resting conditions with a physiological solution in the bath (n = 6; Fig. 7A ). Superfusion of ATP at 10 μM significantly increased BK_Ca currents along the I-V relationship (ANOVA P < 0.001 vs. baseline physiological solution) in seven (78%) of nine cells tested. Bonferroni posttests showed significant differences at +60 (P < 0.05), +80 (P < 0.01), and +100 (P < 0.001) mV versus the baseline recording. After drug washout, BK_Ca currents returned to baseline values, as can be seen in the recording and I-V plot displayed in Figure 7A . Next, we performed experiments in the presence of suramin (100 μM; n = 5; Fig. 7B ) to test the receptor-mediated effect of ATP. After recording BK_Ca currents in physiological solution (baseline) and in the presence of suramin (no effect was seen in the presence of suramin alone), we applied ATP at the same concentration as that used in the previous experiment. In the presence of the blocker, ATP induced a slight nonsignificant increase in BK_Ca currents (Fig. 7B) . These results confirm that the effects on BK_Ca currents are mediated through activation of specific purinergic receptors present in TM cells. 

Participation of RVD Mechanisms in Outflow Facility Modulation

It has been reported that hypotonic medium decreases outflow facility. ^{2 3} In the current study, we perfused eye anterior segments (n = 6) for 90 minutes, to establish baseline outflow facility and therefore isotonic perfusion medium (301 ± 1 mOsm/kg) was replaced with a hypotonic one (244 ± 2 mOsm/kg) and maintained during 360 minutes (Fig. 8A) . This moderate hypotonic shock decreased outflow facility significantly at 15 minutes (20%, P < 0.001; Fig. 8 ), and then it recovered gradually to baseline levels (3% over baseline outflow facility, t-test, NS). The results obtained permitted us to hypothesize that outflow facility is closely linked to TM cell volume and that RVD mechanisms may contribute to recovery of outflow facility after hypotonic shock or other insults that may affect cell volume. We tested this hypothesis by adding drugs known to block or activate selectively the ionic channels that participate in RVD mechanisms. Anterior segments (n = 6) perfused in the presence of 100 μM tamoxifen showed a more pronounced decrease in outflow facility (25%, P < 0.001) after 15 minutes in hypotonic medium compared with the perfusions with hypotonic medium alone (Figs. 8A 8B) . More interesting, they did not recover baseline outflow facility even after 360 minutes (12% below baseline values; P < 0.001 vs. hypotonic medium). Similar results were obtained when hypotonic medium was supplemented with IBTX (10 nM). When both drugs (IBTX and tamoxifen) were added together (n = 6), not only was the initial decrease in outflow facility more pronounced than in the previous groups (34%, P < 0.001), but the recovery phase was the slowest. After 360 minutes, outflow facility was still 17% below baseline (P < 0.001). On the contrary, perfusion with hypotonic medium plus a BK_Ca channel opener (NS1619; 30 μM, n = 6) showed opposite results. The initial decrease in outflow facility was smaller than in hypotonic medium alone (14%, P < 0.001) and recovery to baseline was faster than in the absence of the drug. In fact, outflow facility achieved baseline within ≈45 minutes after hypotonic shock (Figs. 8A 8B) . 

Discussion

In this study, cellular mechanisms involved in TM cell volume regulation were essential in determining AH outflow facility. We put together data from the cellular level with measurements of AH evacuation showing that modulation of certain channels or receptors may have a great influence on outflow facility. 

Different factors have been involved in modulating outflow resistance through the TM (see the introduction). ⁴⁰ Several studies ^{2 3 17 19} have focused on the changes in TM cell volume, since previous data from our laboratory ³ and others ^{2 3 19} have shown that osmolality changes in the perfusion medium evoke swelling or shrinkage of TM cells that modify outflow facility. Also, it has been reported that TM cells trigger an RVD mechanism in response to osmotic insults that may be involved in outflow facility modulation. ¹⁹ In fact, several different membrane proteins that are thought to participate in cell volume regulation have been identified in TM cells, such as the BK_Ca channel, ^{6 26} a Cl⁻ _swell channel, ¹⁹ an Na⁺-K⁺-2Cl⁻ symport, ¹⁷ an Na⁺/H⁺ antiport, ⁴¹ a K⁺-Cl⁻ symport, ¹⁹ and a Cl⁻/HCO₃ ⁻ exchanger. ¹⁹ Moreover, other mechanisms such as ATP release from nerve terminals or due to cell swelling may also be involved in cell volume regulation. ^{23 25}  

AH appears to be slightly more hypertonic than plasma, ^{42 43} and it is likely that changes in AH tonicity influence TM cell volume, although they have not been described to date. More important may be drug-induced changes in TM cell volume (e.g., norepinephrine and adenosine agonists). ^{17 40 44} O’Donnell et al. ¹⁷ showed that bumetanide, ethacrynic acid, 8Br-cAMP, and 8Br-cGMP among others, decreases volume in cultured TM cells through inhibition of the Na⁺-K⁺-2Cl⁻ symport. In contrast, phorbol 12-myristate 13-acetate (PMA) increases cell volume. Bumetanide also increases outflow facility in human and calf eyes. ² Moreover, it has been reported recently that adenosine receptor agonists cause cell shrinkage in TM cells, probably because of Ca²⁺ influx and Cl⁻ efflux. ⁴⁰ In fact, the adenosine agonist cyclohexyladenosine (CHA) has been reported to increase outflow facility in rabbit eyes, ⁴⁵ but it is unknown whether this effect is due to changes in cell volume in the TM. All these results seem to agree that TM cell shrinkage increases outflow facility, whereas cell swelling decreases it, as previously reported by changing perfusate osmolality. ^{2 3}  

Our results show cell swelling activates two ionic conductances: a BK_Ca and a Cl_swell. As described in several cell types, K⁺ and Cl⁻ effluxes are accompanied by water to restore normal cell volume (RVD). ^{23 25} In particular, BK_Ca currents had an approximately threefold increase over the resting values in isotonic medium. Cell swelling activated Cl_swell as soon as 4 minutes after bath exchange to hypotonic medium and reached a maximum between 10 and 12 minutes. Cl_swell currents decreased when returning to isotonic medium (Fig. 1B) showing that their activation is a reversible process. Also, the effects of well-known blockers (tamoxifen and DIDS) of Cl_swell currents and their ionic selectivity are in agreement with previous reports in TM cells ¹⁹ and other cell types. ³⁵ Special mention should be made to the effects of extracellular ATP on Cl_swell currents. After activation by hypotonicity, addition of ATP to the recording bath partially blocked Cl_swell currents (Fig. 3C) . This effect has also been previously described, ^{46 47} but its physiological relevance is unknown. We have also found, as previously reported (Cui M, et al. IOVS 2001;42:ARVO Abstract 743), ⁴⁰ that hypotonic stimuli trigger the release of ATP from TM cells. ATP, through activation of purinergic receptors, increased BK_Ca currents (Fig. 7) , perhaps contributing to the RVD. Nevertheless, the increase of ATP concentration in the extracellular environment may be part of a physiological mechanism to prevent excessive activation of Cl_swell currents, since the blocking effect is only at high concentrations (in the millimolar range) but not at lower concentrations (micromolar) where the effect is to stimulate BK_Ca currents. It should be mentioned that in the absence of hypotonic stimulus, ATP (in micromolar concentrations) activates Cl_swell currents in TM cells (data not shown) as seen in other studies. ⁴⁸ We should note that ATP can be released due to mechanical stimulation in TM cells (Cui M, et al. IOVS 2001;42:ARVO Abstract 743) ³⁶ or can reach the TM after being released in the AH by different ocular tissues and nerves (Fleiszig SMJ, et al. IOVS 2001;42:ARVO Abstract 3146). ^{49 50} Also, we cannot exclude that ATP can be liberated on cell stimulation with several agonists, as seen in other cell types. ³⁶ Therefore, it can be hypothesized that ATP acts in an autocrine/paracrine manner in the TM, regulating outflow facility by volume changes in TM cells. Moreover, Fleischhauer et al. ⁴⁰ have recently shown that released ATP may be a source of adenosine and that adenosine agonists reduce TM cell volume. 

Results obtained in the present study coincide with a profile for a P2Y₂ receptor. The presence of this receptor has been suggested (Cui M, et al. IOVS 2001;42:ARVO Abstract 743), and its activation leads to stimulation of phospholipase C, release of calcium from intracellular stores (IP₃-sensitive), and sustained influx of extracellular Ca²⁺ (Fig. 6) . The characteristics of [Ca²⁺]_i mobilization triggered by ATP and UTP are similar to the ones induced by other drugs such as bradykinin, ⁴ carbachol, ⁵¹ or adenosine agonists. ⁴⁰  

As previously mentioned, ATP stimulation of TM cells increased BK_Ca currents. This effect was partially blocked in the presence of suramin, implying a receptor-mediated mechanism. It is possible that BK_Cas were stimulated directly due to the increase in [Ca²⁺]_i, but it is also possible that purinergic receptor stimulation triggers an intracellular cascade of second messengers to activate BK_Cas, probably through PKC. These two possibilities, which remain to be tested, are not exclusive, and both may be involved in the same final effect on BK_Cas. Of note, ATP released by hypotonic stimuli did not increase [Ca²⁺]_i in TM cells, probably due to a rapid degradation of this nucleotide by nucleotidases. Experiments performed with an inhibitor of nucleotidases in the hypotonic solution showed Ca²⁺ peaks with characteristics similar to the ones elicited by ATP. This fact suggests that in cultured cells ATP is rapidly cleaved but in vivo locally released ATP may be sufficient to stimulate purinergic receptors and activate BK_Ca currents. Further investigation should be conducted to describe the transduction pathway. In this study, BK_Cas were involved in ATP signaling and volume regulation, but they have also been shown to mediate the effects of purinergic agonists, ⁴⁰ nitric oxide, ²⁶ tyrosine kinase, ⁵² and flufenamic acid ⁵³ and to detect membrane stretching. ⁶ Therefore, BK_Cas play a central role in many processes in TM cells, probably regulating the contractile state and cellular volume. Taken together, these effects would finally regulate the AH outflow through the TM. 

Hypotonic solutions decrease outflow facility ^{2 3} and after an initial decline (Fig. 8) , this parameter recovers to achieve baseline levels within ≈2 hours, probably due to an RVD mechanism in TM cells, as shown by Mitchell et al. ¹⁹ Outflow facility did not recover in the presence of tamoxifen, a Cl_swell blocker, and/or IBTX, a BK_Ca blocker. The initial decrease in outflow facility after 15 minutes in hypotonic medium is even greater in the presence of Cl⁻ and K⁺ blockers (Figs. 8A 8B) suggesting that RVD quickly activates to recover cell volume, which shows a direct correlation with outflow facility values. The opposite occurs when using NS1619, a BK_Ca activator, the initial decrease in outflow facility is smaller and the time to recover baseline values is shorter. Therefore, activators of BK_Cas may exert a protective effect against osmotic insults or drugs that modify cellular volume. ⁴⁰ It may also be possible that activators of BK_Cas like nitric oxide or cGMP may be endogenously released to modulate outflow facility, although further investigations are necessary to test this hypothesis. Finally, specific activators of Cl_swell (unknown to date with the exception of ATP at low concentrations ⁴⁸ ) would also be interesting molecules to modulate outflow facility. Because TM function can be modulated by changes in cell volume, selective drugs targeting cell volume regulatory mechanisms would increase outflow facility without damaging either the cells or the extracellular matrix. The present results and others ^{17 19 20 40 44 53} support this idea. 

In summary, our findings show that cellular mechanisms that regulate cell volume in TM cells play an important role in outflow facility regulation. In particular, K⁺ and Cl⁻ channel activation is directly involved in the recovery of baseline outflow facility after hypotonic shock. Therefore, cell volume regulation appears to be part of the homeostatic mechanisms involved in the regulation of the outflow pathway, actively contributing to TM function. 

²  Contributed equally to the work and therefore should be considered equivalent senior authors.

 Supported by Grants PM99–0169, BFI2002–01202, BFI2003–01190, BFI2001–3331 from Ministerio de Ciencia y Tecnologia, Spain.

 Submitted for publication January 22, 2004; revised June 16 and July 9, 2004; accepted July 15, 2004.

 Disclosure: D. Soto, None; N. Comes, None; E. Ferrer, None; M. Morales, None; A. Escalada, None; J. Palés, None; C. Solsona, None; A. Gual, None; X. Gasull, None

 The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked “advertisement” in accordance with 18 U.S.C. §1734 solely to indicate this fact.

 

Figure 1.

 

Hypotonic medium increased K⁺ and Cl⁻ currents. (A) Left: representative experiment showing whole-cell BK_Ca currents in a TM cell in isotonic and hypotonic media. V _h = −60 mV. Right: current–voltage curve showing outward BK_Ca currents of TM cells in the whole-cell configuration (n = 13). Currents are expressed as current densities (pA/pF). Data are for isotonic (301 mOsm/kg) and hypotonic (213 mOsm/kg) media. BK_Ca current increased significantly (ANOVA, P < 0.001). *P < 0.05, ***P < 0.001 hypotonic versus isotonic medium (Bonferroni posttest). Outward currents were blocked by IBTX (data not shown). (B) Left: representative experiment showing whole-cell Cl_swell in a TM cell in isotonic, then hypotonic, and returning to isotonic media. V _h = 0 mV. Right: Current–voltage curve showing outward Cl_swell currents of TM cells in the whole-cell configuration (n = 5). Data are shown for isotonic medium baseline, hypotonic medium, and isotonic medium return to baseline. Cl_swell current increased significantly (ANOVA, P < 0.001). **P < 0.01, ***P < 0.001 hypotonic versus isotonic medium (Bonferroni posttest). ++P < 0.01, +++P < 0.001 return to isotonic baseline versus hypotonic.

Figure 2.

 

Effect of Cl⁻ ion replacement on reversal potentials of Cl_swell. Left: reversal potential (mean ± SEM), relative permeability, and number of experiments for each anion. Right: mean recordings in the whole-cell configuration of the patch-clamp technique for (solid lines) Cl⁻ and (dotted lines) I⁻ (A), Br⁻ (B), F⁻ (C), and gluconate (D). SEM was omitted for clarity. Voltage ramps where recorded after activation of Cl_swell current in hypotonic medium. Cl⁻ was replaced by each anion.

Figure 3.

 

Cl_swell pharmacology. V _h = 0 mV in all three experiments. (A) Left: representative experiment showing whole-cell Cl_swell in a TM cell in isotonic, hypotonic, and hypotonic+tamoxifen (100 μM) media. Tamoxifen decreased outward and inward currents. Right: mean ± SEM outward (at +80 mV) and inward (at −80 mV) current densities (pA/pF) recorded in five TM cells. (B) Left: representative experiment showing whole-cell Cl_swell in a TM cell in isotonic, hypotonic, and hypotonic+DIDS (100 μM) media. DIDS decreased outward but not inward currents. Right: Current densities are as in (A) and were recorded in six TM cells. (C) Left: representative experiment showing whole-cell Cl_swell currents in a TM cell in isotonic, hypotonic, and hypotonic+extracellular ATP (1 mM) media. ATP decreased outward and inward currents. Right: Current densities are as in (A) and were recorded in six TM cells. Note that some Cl_swell currents inactivated at strong depolarized potentials (hypotonic in A), but the common pattern was an increasingly rapid inactivation as the depolarization increased (hypotonic in B and C). Significant differences were recorded between isotonic and hypotonic medium (*P < 0.05; t-test) and between hypotonic medium and hypotonic+blocker (⁺ P < 0.05 and ⁺⁺⁺ P < 0.001; t-test).

Figure 4.

 

ATP release from TM cells. TM cells grown to confluence in 35-mm culture dishes were stimulated (arrow) with isotonic medium or with medium in which hypotonicity was reduced by 20% or 40%. Released ATP was measured by a luciferin-luciferase–detection method. (A) Representative recording of ATP release from TM cells after stimulation with isotonic and 20% and 40% hypotonic media, successively. (B) ATP release in isotonic (n = 3; mean ± SEM), 20% hypotonic (n = 5), and 40% hypotonic (n = 4) media. Significant differences were found between isotonic and hypotonic stimulations (***P < 0.001; t-test).

Table 1.

 

Effects of Purinergic Agonists and Antagonists on [Ca²⁺]_i Levels in Cultured BTM Cells

Drug	Responder/Analyzed Cells (n)	Responder Cells (%)	Basal [Ca2+]i (nM)	Peak [Ca2+]i (nM)
Control (Vehicle)	7/275	3	64 ± 5	169 ± 19
ATP dose–response relationship
10−10 M	2/122	2	76 ± 6	225 ± 51
10−9 M	11/96*	11	68 ± 8	128 ± 12
10−8 M	109/304*	36	84 ± 8	234 ± 7†
10−7 M	97/139*	70	87 ± 12	311 ± 11††
10−6 M	311/315*	99	64 ± 5	513 ± 12†††
10−5 M	128/129*	99	62 ± 9	567 ± 16†††
10−4 M	82/83*	99	79 ± 9	587 ± 17†††
Purinergic agonists and antagonists
UTP 1 μM	516/522	99	71 ± 3	414 ± 8§§
ADP 1 μM	145/427‡‡	34	71 ± 5	339 ± 11§§
UDP 1 μM	108/324‡‡	33	85 ± 5	314 ± 14§§
ATP-γ-S 1 μM	414/438‡	95	72 ± 5	371 ± 7§§
2-MeS-ATP 1 μM	29/106‡‡	27	63 ± 37	422 ± 40§
PPADS 10 μM+ ATP 1 μM	320/332	96	64 ± 6	387 ± 10§§
Suramin 100 μM+ ATP 1 μM	311/453‡‡	69	89 ± 11	386 ± 10§§
Intracellular mechanisms
ATP 1 μM (0 Ca2+)	456/464	98	81 ± 6	464 ± 8§§
ATP-γ-S 1 μM (0 Ca2+)	113/116	97	70 ± 7	376 ± 18
Thapsigargin 1 μM+ ATP 1 μM	6/233‡‡	3	151 ± 10	331 ± 47§
U73122 1 μM+ ATP 1 μM	185/419‡‡	44	103 ± 14	324 ± 10§§
CPA 1 μM+ ATP 1 μM	86/292‡‡	29	89 ± 15	338 ± 10§§
Ryanodine 1 μM+ ATP 1 μM	171/176	97	52 ± 2	371 ± 14§§

 [Ca²⁺]_i values are mean ± SEM. [Ca²⁺]_i peaks were considered significant when they were at least two times basal levels and means were calculated only in the responder cells. Comparisons between the proportion of responder cells in the control group and the ATP-treated groups (*) were made using the Fisher exact test. Comparisons between the average [Ca²⁺]_i peak elicited by BTM cells in the control group and in ATP-treated groups (†) were made using Student’s unpaired t-test. * P < 0.001 by Fisher exact test. † P < 0.05, †† P < 0.01 and ††† P < 0.001 by Student’s unpaired t-test. Comparisons between the proportion of responder cells in the ATP 1 μM–treated group and the other groups (‡) were made using the Fisher exact test. Comparisons between the average [Ca²⁺]_i peak elicited by BTMs cells in the ATP 1 μM–treated group and the other groups (§) were made using Student’s unpaired t-test. ‡ P < 0.01, ‡‡ P < 0.001 by the Fisher exact test. § P < 0.05, §§ P < 0.001 by Student’s unpaired t-test.

Figure 5.

 

[Ca²⁺]_i mobilizations induced by purinergic agonists in TM cells. Representative experiments of TM cells stimulated (arrow) with ATP (A), UTP (B), ADP (C), and UDP (D) at 1 μM. The blocking effects of purinergic antagonists were tested in TM cells preincubated with suramin (100 μM; E) or PPADS (10 μM; F) and stimulated with ATP (1 μM). (G) The percentage of cells (mean ± SEM) responding to purinergic agonists. All compounds were tested at 1 μM except suramin (100 μM) and PPADS (10 μM). (H) The ATP dose–response curve for [Ca²⁺]_i mobilization in TM cells (percentage of cells responding to ATP application). Significant differences were shown between the ADP, UDP, 2-MeS-ATP, and suramin+ATP groups versus the ATP group (*P < 0.05, **P < 0.01, ***P < 0.001; t-test).

Figure 6.

 

Intracellular mechanisms involved in ATP signaling. (A) Representative experiments showing the effect of ATP 1 μM (arrow) in a TM cell incubated in physiologic solution (1 mM Ca²⁺), in the absence of external calcium, or with thapsigargin (1 μM). [Ca²⁺]_i increased to a peak and then recovered to baseline. In the absence of extracellular Ca²⁺, the recovery phase was shortened. Intracellular Ca²⁺ store depletion blocked the increase of [Ca²⁺]_i. (B) Percentage of cells (mean ± SEM) responding to ATP (1 μM) in the absence of extracellular Ca²⁺ or after preincubation with thapsigargin (1 μM), U73211 (1 μM), ryanodine (1 μM), or CPA (1 μM). Significant differences were shown between the thapsigargin, U73211, and CPA groups versus the ATP group (***P < 0.001). (C) Effects of hypotonic shock on [Ca²⁺]_i. Shown are data for hypotonic shock (206 mOsm/kg) alone, hypotonic+ARL67156 (inhibitor of nucleotidases; 100 μM), and hypotonic+ARL67156+suramin (100 μM). Mean Ca²⁺ peaks are shown. SEM was omitted for clarity.

Figure 7.

 

ATP increased BK_Ca currents. V _h = −60 mV in both experiments. (A) Left: representative experiment showing whole-cell BK_Ca currents in a TM cell in resting conditions (physiological solution), in the presence of ATP (10 μM), and after washing the drug (return). Right: current/voltage curve showing outward BK_Ca currents of TM cells in the whole-cell configuration (n = 6). Currents are expressed as current densities (pA/pF). Shown are data for resting conditions (physiological solution), ATP (10 μM), and return to physiological solution. BK_Ca current increased significantly (ANOVA, P < 0.001). *P < 0.05, **P < 0.01, ***P < 0.001 for ATP versus physiological solution (Bonferroni posttest). (B) Left: representative experiment showing whole-cell BK_Ca currents in a TM cell in resting conditions (physiological solution), in the presence of suramin (100 μM), and after addition of ATP (10 μM). Right: current–voltage curve showing outward BK_Ca currents of TM cells in the whole-cell configuration (n = 5). Currents are expressed as current densities (pA/pF). Data are shown for resting conditions (physiological solution), suramin, and suramin+ATP. Suramin blocked the ATP-induced increase in BK_Ca current.

Figure 8.

 

Influence of RVD mechanisms on modulation of outflow facility. (A) Outflow facility ratio (normalized with baseline outflow facility) is plotted against time. Anterior segments were perfused with control medium (DMEM) for 90 minutes (baseline), and then perfusion medium was replaced (arrow) by hypotonic DMEM with or without drugs, and the solution was maintained until the end of the experiment. Data are shown for hypotonic (n = 6), hypotonic+tamoxifen (100 μM, n = 6), hypotonic+tamoxifen+IBTX (10 nM, n = 6), hypotonic+BK_Ca opener (NS1619, 30 μM, n = 6), and hypotonic+IBTX (10 nM, n = 6). The initial decrease in outflow facility was reversed in the presence of K⁺ and Cl⁻ channel blockers. Also, recovery phase to baseline was impaired in the presence of the blockers. On the contrary, the BK_Ca channel opener decreased the initial decline in outflow facility in hypotonic medium and shortened the recovery phase. (B) Mean ± SEM outflow facility ratios measured after 15 minutes in the hypotonic medium (or hypotonic medium+drugs) and after 360 minutes of perfusion in hypotonic medium. Time after 15 minutes is representative of the initial decrease in outflow facility due to hypotonic medium. Measures after 360 minutes in hypotonic medium are representative of the recovery phase. Significant differences (t-tests) are shown between each group and the control group (baseline in isotonic medium): *P < 0.05, **P < 0.01, ***P < 0.001.

The authors thank Pablo Perez de la Ossa for advice and assistance in the LDH determinations.