Release of ATP from retinal pigment epithelial cells involves both CFTR and vesicular transport

David Reigada and Claire H. Mitchell

Department of Physiology, University of Pennsylvania, Philadelphia, Pennsylvania

Submitted 26 April 2004 ; accepted in final form 26 August 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The retinal pigment epithelium (RPE) faces the photoreceptor outer segments and regulates the composition of the interstitial subretinal space. ATP enhances fluid movement from the subretinal space across the RPE. RPE cells can themselves release ATP, but the mechanisms and polarity of this release are unknown. The RPE expresses the cystic fibrosis transmembrane conductance regulator (CFTR), and CFTR is associated with ATP release in other epithelial cells. However, an increasing number of reports have suggested that the exocytotic pathway contributes to release. In the present study, we examined the involvement of CFTR and the vesicular pathway in ATP release from RPE cells. Release from cultured human ARPE-19 cells and across the apical membrane of fresh bovine RPE cells in an eyecup was studied. A cAMP cocktail to activate CFTR triggered ATP release from fresh and cultured RPE cells. Release from both RPE preparations was largely prevented by the broad-acting blocker glibenclamide and the specific thiazolidinone CFTR inhibitor CFTR-172. The block by CFTR-172 was enhanced by preincubation and prevented ATP release with 3.5 µM IC50. The rise in intracellular Ca2+ accompanying hypotonic challenge was prevented by CFTR-172. The vesicular transport inhibitor brefeldin A prevented ATP release after stimulation with both hypotonic and cAMP conditions, suggesting vesicular insertion was also involved. These results show an intimate involvement of CFTR in ATP release from RPE cells which can autostimulate receptors on the apical membrane to modify Ca2+ signaling. The requirement for both CFTR and vesicular transport pathways suggests vesicular insertion of CFTR may underlie the release of ATP.

cystic fibrosis transmembrane conductance regulator; recycling endosomes; brefeldin A; autostimulation; retinal detachment


THE RETINAL PIGMENT EPITHELIUM (RPE) is a monolayer of cells lying between the retinal photoreceptors and the choroidal blood supply. The RPE functions in a glial cell-like capacity to maintain the health and signaling ability of the outer retina. The RPE supplies the outer retina with general nutrients and components critical for the visual cycle (10). The distal tips of the photoreceptor outer segments are regularly phagocytozed and metabolized by the RPE to maximize photoreceptor responsiveness (66). The RPE also regulates the ionic composition of the extracellular subretinal space surrounding the outer segments and controls the absorption of fluid from the retina to the choroid (13, 22, 32, 34, 35).

This complex interaction between the RPE and the photoreceptors is dependent on close communication between the two cell types. Various neurochemicals have been implicated in this communication, including dopamine (15, 59), serotonin (39), and epinephrine (13, 24, 42). The purines ATP and adenosine also contribute to RPE-retina interaction. Stimulation of the RPE by adenosine can modify the rate of outer segment phagocytosis (17), while stimulation of apical P2Y2 receptors by ATP can trigger elevation of Ca2+ (40, 55). This elevation likely opens a basolateral Cl conductance and increases the flux of ions and fluid across the RPE (40). Triggering this fluid absorption by stimulating apical P2Y2 receptors with agonist INS37217can reduce the size of fluid blebs in the subretinal space (29, 31), emphasizing the potential of purinergic signaling in the treatment of retinal detachment.

The endogenous source of ATP capable of stimulating these P2 receptors is likely the RPE cells themselves. The existence of a physiological release of ATP from epithelial cells in many different cell types is now widely accepted (6, 48, 50), and RPE cells have been shown to release ATP in response to stimulation (14, 36). However, the mechanisms involved in this release remain to be determined. Previous work has shown that while the Ca2+ ionophore ionomycin is not sufficient to trigger release, the general Cl channel blocker 5-nitro-2-(3-phenylpropylamino)-benzoate inhibits it, suggesting anion channel involvement (36). RPE cells have been shown to express the anion channels ClC-2, ClC-3, ClC-5, the pCLCA1 Cl channel regulator, and the cystic fibrosis transmembrane conductance regulator (CFTR; Refs. 4, 26, 63, 64). Of these, ATP release has been associated most consistently with CFTR (7, 41, 52, 60). However, recent evidence suggests that the exocytotic pathway may also be important, because substances that interfere with vesicular transport, such as botulinum toxin, tetanus neurotoxin, and brefeldin A (BFA) can prevent stimulated ATP release (5, 30, 58). Consequently, we investigated the involvement of CFTR in ATP release from RPE cells and examined whether vesicular transport contributed to this release. Preliminary results were presented previously in abstract form (43, 44).


    MATERIAL AND METHODS
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. ARPE-19 cells (12) were obtained from the American Type Culture Collection (Manassas, VA) and grown in 25-cm2 Falcon primary culture flasks (Becton Dickinson, Franklin Lakes, NJ) in a 1:1 mixture of Dulbecco's modified Eagle's medium and Ham's F-12 medium with 3 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, 2.5 mg/ml Fungizone, and/or 50 µg/ml gentamicin (all obtained from Invitrogen, Carlsbad, CA) in the presence of 10% fetal bovine serum (FBS; Hyclone Laboratories, Logan, UT). Cells were incubated at 37°C in 5% CO2 and subcultured weekly with 0.05% trypsin and 0.02% EDTA.

Solutions. The isotonic solution used as a basis for all solutions was composed of (in mM) 105 NaCl, 5 KCl, 6 HEPES acid, 4 Na-HEPES, 5 NaHCO3, 60 mannitol, 5 glucose, 0.5 MgCl2, and 1.3 CaCl2, pH 7.4. Hypotonic solution was obtained by adding the required amount of deionized distilled water, with "%hypotonicity" defined as the percentage of water added. Glibenclamide, forskolin, 3-isobutyl-1-methylxanthine (IBMX), 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), and CFTR-172 were mixed as stock solutions in dimethyl sulfoxide. The CFTR-172 was a kind gift from A. S. Verkman (Departments of Medicine and Physiology, Cardiovascular Research Institute, University of California San Francisco, San Francisco, CA). The cAMP-stimulating mixture contained (in µM) 10 forskolin, 100 IBMX and 500 8-4-chlorophenylthio(cpt)-cAMP. Brefeldin A was dissolved in ethanol. All chemicals were from Sigma-Aldrich (St. Louis, MO) unless otherwise noted.

ATP measurements from cultured cells. ATP release was detected through the chemiluminescent luciferin-luciferase reaction, and the light emitted was recorded using a microplate luminometer (Luminoskan Ascent; Labsystems, Franklin, MA). ARPE-19 cells were grown to confluence in 96-well white assay plates with clear bottoms for 4–7 days (Corning, Corning, NY). To minimize the ATP release that accompanies solution change (18), growth medium was removed from the cells 1 h before the experiment and replaced with 100 µl of isotonic solution. After 1 h, 50 µl of the isotonic solution was replaced by 50 µl of a solution containing twice the drug concentration or the amount of distilled water required to obtain the final concentration or hypotonicity. To produce a 60% hypotonic solution, 60 µl were removed. The plate was placed in the microplate luminometer 60–90 s after addition of the drugs, and 10 µl of the luciferase working solution were injected in each well through the internal injector system. Measurements were taken every 30 s for each well for 30 min with an integration time of 100 ms/measurement. The luciferase solution was prepared in a stock solution from one vial of the ATP assay kit diluted in 450 µl of isotonic solution and 50 µl of distilled water. A working solution was made by diluting 40 µl of the stock solution in 1 ml of isotonic solution. The ATP released was calculated at the different time points indicated in the RESULTS with the use of a standard curve to transform the arbitrary units obtained through the luciferase reaction to an ATP concentration.

All the substances were tested for the effect on the luciferin-luciferase assay as shown in Fig. 1. Only hypotonic solutions and genistein were found to interfere with the assay itself. All of the data obtained using hypotonic solutions were corrected for this enhancement of the chemiluminescence reaction. A standard curve covering three orders of magnitude showed the relationship between luminescence and ATP concentration to be linear over the range of interest. The results from hypotonic experiments were thus scaled down by the factor shown to enhance the response to 10 nM ATP.



View larger version (15K):
[in this window]
[in a new window]
 
Fig. 1. The effect of test conditions on the chemiluminescence luciferin-luciferase reaction. The effect of various conditions on the luciferin-luciferase assay was examined using 10 nM ATP. While glibenclamide, forskolin, 8-4-chlorophenylthio(cpt)-cAMP, the cAMP stimulating mixture [cpt-cAMP, forskolin, and 3-isobutyl-1-methylxanthine (IBMX)], CFTR-172, and brefeldin A (BFA) did not affect the chemiluminescence level, genistein, 30% hypotonic solution, and 50% hypotonic solution did produce a significant change in the assay (n = 5–16 for each condition; *P < 0.05 vs. 10 nM ATP alone). Bars, means ± SE.

 
ATP measurements from bovine eyes. Bovine eyes were obtained from the abattoir and transported on ice to the laboratory. The eye was bisected at the ora serrata, and the retina was removed and detached from the optic nerve to expose the RPE cell layer. The eyecup was washed in isotonic solution, and then 0.5–1 ml of the control solution described above was placed in the eyecup. Next, after 10 min at room temperature, a 0.4- to 0.8-ml sample was removed. Deliberate disruption of the monolayer led to a clearly discernible discoloration of the tissue; the absence of this discoloration was assumed to indicate the presence of an intact monolayer. To determine the ATP content of each eyecup, 100 µl of each sample were placed into a well of a 96-well plate, put in the microplate luminometer, and 10 µl of the luciferase working solution described above were injected into each well. The initial three readings were taken to represent the ATP level.

Measurement of intracellular Ca2+. ARPE-19 cells were plated onto coverslips and grown as described above until confluent. After washing, cells were loaded with 10 µM fura-2 and 0.2% pluronic acid at 37°C for 30 min. The dye was washed off, and the coverslips were mounted on a Nikon inverted microscope (Nikon Instruments, Melville, NY) and visualized with a x20 objective. The field of ~20 cells was alternatively excited at 340 and 380 nm with a monochromator (Photon Technologies International, Lawrenceville, NJ), and the fluorescence emitted at 510 nm was detected with a photometer (Photon Technologies International). The ratio of light detected after excitation at 380 and 340 nm was converted into Ca2+ concentration using a calibration performed after each measurement as previously described (36).

Data analysis. Baseline levels of ATP varied considerably by experimental day for both fresh and cultured experiments. Efforts to remove this variability, such as limiting cell age and changing the solution 1 h before the measurements, were only partially effective, and the raw concentrations throughout the text reflect this variation. To allow comparison of experiments, values were normalized to the mean control value of each day. All data are expressed as means ± SE, and the unpaired Student's t-test was used for statistical analysis, with P < 0.05 defined as significantly different. The IC50 of CFTR-172 and the EC50 of hypotonicity were determined by fitting the data curve with first-order exponential curves using SigmaPlot graphing software (SPSS, Chicago, IL).


    RESULTS
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Fresh vs. cultured cells. While the cultured human ARPE-19 cells have many advantages, it was important to determine the characteristics of ATP release by fresh RPE cells and the polarity of any such release. In the RPE eyecup preparation, the apical membrane faces the cup interior, and both drug access and ATP release likely occur across the apical membrane. Initial experiments thus compared the ATP release characteristics from cultured ARPE-19 cells and the fresh bovine RPE eyecup stimulated with hypotonic solution.

ATP release from ARPE-19 cells reached a peak 10 min after the addition of a 50% hypotonic solution, after which levels slowly declined for the remainder of the 30-min measurement period (Fig. 2A). Peak levels rose more than eightfold in this series of experiments, from 1.7 ± 0.5 nM to 14.1 ± 1.6 nM (n = 8 for both). The amount of ATP present in the well rose with the degree of hypotonicity >30% and reached a peak at 50% hypotonicity. Increasing hypotonicity to 60% did not produce any additional increase, giving an EC50 of ~42% (Fig. 2B).



View larger version (15K):
[in this window]
[in a new window]
 
Fig. 2. ATP release triggered by hypotonicity in fresh and cultured retinal pigment epithelium (RPE) cells. A: online measurement of ATP released from ARPE-19 cultured cells by 50% hypotonicity (dark inverted triangles) vs. control (open circles). Data are means ± SE of 8 experiments, with bars absent when SE were smaller than the symbol. B: increasing the percentage of water (%hypotonicity) raised the levels of ATP bathing ARPE-19 cells. Symbols represent means ± SE of 8 experiments for each data point. Data were normalized to the mean control of each experiment. C: comparison of the ATP release induced by hypotonicity in fresh bovine RPE cells (closed bars) and ARPE-19 cultured cells (open bars). *P < 0.001 vs. control. **P < 0.02 vs. control. Number of experiments (n) is shown above each bar. Bars, means ± SE.

 
The release from fresh bovine cells after hypotonic challenge was similar to that of the ARPE-19 cells. Because the peak from ARPE-19 cells occurred at 10 min, samples were taken from the intact bovine RPE eyecup at 10 min. Levels showed that the release across the apical membrane of fresh cells was increased significantly when challenged with 30% hypotonicity. Presentation with a 50% hypotonic solution raised ATP levels from 7.8 ± 0.6 nM (n = 6) to 22.2 ± 2.8 nM (n = 5). The ATP release pattern from fresh and cultured cells is shown in Fig. 2C. Later experiments used the minimum level of hypotonicity that reproducibly induced a significant release in each system: 30% for the fresh bovine RPE cells and 50% for the ARPE-19 cultured cells.

Release triggered by cAMP. To determine whether CFTR was involved in ATP release from RPE cells, the effect of intracellular cAMP was examined because activation by the cAMP-dependent protein kinase A is typically necessary for the opening of the CFTR conductance pathway (2). Intracellular cAMP was increased with a cAMP cocktail containing cell-permeant cpt-cAMP, forskolin, and IBMX. The cocktail triggered an immediate increase in ATP levels bathing ARPE-19 cells (Fig. 3A). While recording from the assay plate generally began within 60–90 s after addition of stimuli, continuous monitoring during cocktail addition confirmed that ATP levels reached their peak within 30 s. Peak levels of ATP rose fourfold when cells were exposed to the cocktail (Fig. 3B). Although both forskolin and cpt-cAMP increased bath levels of ATP, levels were highest with the complete cocktail. Addition of the cocktail at the peak of the hypotonic response did not lead to a further increase. In fresh bovine RPE cells, the cAMP-stimulating cocktail increased ATP levels 2.5-fold, from 10.1 ± 2.1 nM ATP (n = 6) to 24.3 ± 9.3 nM ATP (n = 6, Fig. 3C). Forskolin itself led to a small but not significant increase in the fresh cells. Because the CFTR activator genistein significantly inhibited the luciferin-luciferase assay (Fig. 1), it was not possible to reliably quantify its enhancement of ATP release.



View larger version (17K):
[in this window]
[in a new window]
 
Fig. 3. ATP release by cAMP-stimulating mixture. A: online measurements show a cAMP-activating mixture (dark inverted triangles) composed of (in µM) 500 cpt-cAMP, 10 forskolin, and 100 IBMX induced rapid release of ATP from ARPE-19 cells compared with control conditions (open circles). The decay is likely due to the degradation of ATP by ecto-ATPases and reduction of substrate for the reaction. Symbols represent means ± SE; n = 12. B: effects of components of cAMP stimulating mixture were roughly additive in ARPE-19 cells. *P < 0.001 vs. control. **P < 0.001 vs. control and P < 0.05 vs. cpt-cAMP. C. The cAMP cocktail also triggered ATP release from fresh bovine RPE cells. Bars, means ± SE.*P = 0.051 vs. control. **P < 0.001 vs. control.

 
CFTR blockers. Pharmacological tools were used to further elucidate the contribution of CFTR to ATP release from RPE cells. Because glibenclamide can block the Cl current associated with CFTR (65), its effects on ATP release were determined. When glibenclamide (100 µM) and hypotonic solutions were added together to the ARPE-19 cells, a reduction in ATP was seen throughout the experiment (Fig. 4A). In this series of experiments, hypotonicity increased peak ATP levels 8.9-fold, from 3.3 ± 1.1 nM (n = 6) to 29.1 ± 3.8 nM (n = 7). Glibenclamide blocked 71% of the peak response, with levels of 9.7 ± 1.5 nM (n = 6). A summary of the effects of 100 µM glibenclamide from all trials is shown in Fig. 4B. The drug produced a complete block of ATP release from bovine RPE cells when used at 100 µM (Fig. 4C). Hypotonicity raised ATP 2.2-fold in the bovine RPE eyecup, from 0.26 ± 0.02 nM (n = 6) to 0.35 ± 0.04 nM (n = 7). In the presence of 100 µM glibenclamide, hypotonicity raised ATP to only 0.13 ± 0.02 nM (n = 5; not significantly different from control). Levels were particularly low in this set of experiments, but they were consistent, as evidenced by the small variation.



View larger version (17K):
[in this window]
[in a new window]
 
Fig. 4. ATP release by hypotonicity is blocked by cystic fibrosis transmembrane conductance regulator (CFTR) inhibitor glibenclamide. A: ATP release induced in ARPE-19 cells by a 50% hypotonic solution (shaded inverted triangles) is blocked by 100 µM glibenclamide (closed squares) vs. control (open circles). Symbols represent means ± SE of a set of experiments; n = 6–7. B: summary of experiments showing that glibenclamide significantly reduced the peak ATP levels surrounding ARPE-19 cells exposed to a 50% hypotonic solution. *P < 0.001, significantly different from control. **P < 0.001, significantly different from hypotonic condition. C: in fresh bovine RPE cells, ATP release triggered by a 30% hypotonic solution was blocked completely by glibenclamide (100 µM). *P = 0.006, significantly different from control. **P = 0.004, significantly different from hypotonic condition. Bars, means ± SE.

 
Although the block by glibenclamide suggested that CFTR contributes to ATP release from the RPE, the drug is not specific for CFTR. A thiazolidinone blocker, CFTR-172, that inhibits CFTR with far greater potency and specificity than glibenclamide was recently identified by Verkman and colleagues (28, 56). CFTR-172 was used to provide a more decisive assessment of the CFTR contribution to ATP release. The blocker CFTR-172 inhibited hypotonically triggered release from ARPE-19 cells (Fig. 5A). When the drug was added simultaneously with the hypotonic solution, minimal block was observed, but inhibition was considerably increased by preincubating the cells with 10 µM CFTR-172 for 30 min (Fig. 5B). Preincubation with 10 µM CFTR-172 decreased the ATP levels bathing ARPE-19 during hypotonic exposure by 79.3%, from 13.4 ± 1.0 nM ATP (n = 23) to 4.3 ± 1.2 nM ATP (n = 12), compared with control of 1.9 ± 0.1 nM (n = 46). The decrease in ATP levels was concentration dependent, with an IC50 of 3.5 µM when preincubated for 30 min. CFTR-172 completely blocked the hypotonically triggered release from fresh bovine cells (Fig 5C). Hypotonicity (30%) raised the peak ATP concentration 1.7-fold, from 8.5 ± 2.0 nM (n = 12) to 14.9 ± 3.2 nM (n = 8). CFTR-172 reduced the level by 95%, to 8.9 ± 3.4 nM (n = 8).



View larger version (17K):
[in this window]
[in a new window]
 
Fig. 5. CFTR-172 blocks release of ATP. A: time-dependent traces showing ARPE-19 cells exposed to 50% hypotonic solution (shaded squares; n = 23), control (open circles; n = 46), and preincubated with 10 µM CFTR-172 for 30 min before application of hypotonicity (closed inverted triangles; n = 12). B: hypotonically triggered release of ATP is blocked by CFTR-172 in a concentration-dependent way. The peak response is shown from cells exposed to isotonic solution (open squares; n = 31), to hypotonic solution and 10 µM CFTR-172 applied simultaneously (open inverted triangles; n = 7), or to hypotonic solution plus various concentrations of CFTR-172 preincubated for 30 min (closed circles; n = 23, 6, 14, 14, 14, and 6 experiments with 0, 0.1, 0.3, 3, 10, and 30 µM CFTR-172, respectively). The preincubated cells were fit with a single exponential decay with EC50=3.5 µM. C: in the fresh bovine RPE eyecup, CFTR-172 inhibited all of the ATP release triggered by exposure to a 30% hypotonic solution. The cells were preincubated with CFTR-172 for 30 min before exposure to hypotonicity. Bars, means ± SE. *P = 0.006, significantly different from control. **P = 0.044, significantly different from hypotonic solution alone.

 
Effect on intracellular Ca2+. The levels of extracellular ATP measured after hypotonic or cAMP stimulation in RPE cells are below the EC50 for P2Y receptors present on the apical membrane of RPE cells. While the localization of release sites and receptors within microdomains suggests that the effective local concentration of ATP may be considerably greater (21, 25), we sought physiological evidence that the CFTR-linked release of ATP was sufficient to stimulate receptors by examining its effect on intracellular Ca2+ levels, as P2 receptor stimulation is known to elevate Ca2+ in RPE cells (40). ARPE-19 cells perfused with hypotonic solution exhibited a two-phase increase in intracellular Ca2+, with a large, transient increase followed by a slow rise in Ca2+ (Fig. 6A). The inhibitor CFTR-172 greatly diminished both Ca2+ responses. Cells exposed to 10 µM CFTR-172 for 30 min before and during hypotonic challenge showed a 95% reduction in the transient peak and a 42% reduction in the delayed Ca2+ peak (Fig. 6B). In both cases, a net reduction in Ca2+ concentration was observed in response to hypotonicity, possibly because of the dilution of cellular constituents. Because CFTR-172 did not affect levels in isotonic solution or calibration readings, the inhibition was specific for stimulated release of ATP. This implies that CFTR-associated release of ATP is capable of autostimulating receptors on the RPE and suggests that this release is responsible for both the fast and slow elevations in Ca2+ after hypotonic challenge.



View larger version (16K):
[in this window]
[in a new window]
 
Fig. 6. CFTR-172 blocks autocrine elevation in Ca2+. A: changes in the intracellular Ca2+ of ARPE-19 cells perfused with hypotonic solution were inhibited by CFTR-172. A large and fast increase was detected soon after the reduction of the osmolarity, followed by a second slow rise from the lowest point with a maximum 10–15 min after the stimuli presentation (solid line). The inhibitor CFTR-172 (shaded line) completely inhibited the fast peak and significantly blocked the slow rise after a 30-min preincubation at 10 µM concentration. Inset shows the secondary response on an expanded scale. The lines show a representative experiment using ~20 confluent cells each. B: summary of the effects of the inhibitor CFTR-172 on the changes in intracellular Ca2+ observed in the ARPE-19 cells by the action of the 50% hypotonic solution. Preincubation with 10 µM CFTR-172 produced complete inhibition of the fast peak and a significant inhibition of the slow secondary rise. Change in Ca2+ is defined as from control baseline to maximum peak for the first event and from the lowest posthypotonicity point to the second maximum for the slow rise. Bars, means ± SE; n = no. of recordings from ~20 cells each. *P < 0.001 vs. hypotonic solution alone: **P = 0.031 vs. hypotonic solution alone.

 
Involvement of vesicular transport. While the above experiments with CFTR-172 clearly establish a predominant role for CFTR in the release of ATP, the effects of BFA on ATP release were examined to determine whether the vesicular release pathway was also involved in the response. ARPE-19 cells were preincubated with 10 µg/ml BFA for 2 h before exposure to a 50% hypotonic solution. The BFA slowed the initial release of ATP and led to a reduced level of extracellular ATP throughout the experimental period (Fig. 7A). In these experiments, hypotonicity increased the ATP levels 5.1-fold to 7.3 ± 0.5 nM (n = 36) from 1.2 ± 0.1 nM (n = 26; Fig. 7B). A 2-h preincubation with BFA led to a 70% reduction in peak ATP concentration, with mean levels of 3.8 ± 0.3 nM (n = 19). To determine whether vesicular transport made a similar contribution to both hypotonic and cAMP-triggered release, the effect of BFA on the release after cAMP stimulation was examined. In this set of experiments, the cAMP cocktail increased the ATP levels 1.5-fold to 4.0 ± 0.6 nM (n = 72) from 2.6 ± 0.2 nM (n = 84). However, a 2-h preincubation with BFA reduced ATP release by 79%, to 2.9 ± 0.2 nM (n = 60; Fig. 7C).



View larger version (31K):
[in this window]
[in a new window]
 
Fig. 7. Basal ATP release and ATP release induced by hypotonicity and cAMP are blocked by vesicular transport inhibitor BFA. A: ATP release induced in ARPE-19 cells by 50% hypotonic solution (shaded inverted triangles) is blocked by 10 µg/ml BFA (squares) vs. control (open circles). Cells were preincubated with BFA for 2 h before the recording began. n = 8–16. B: summary of the effects of the inhibitor BFA (10 µg/ml) on the peak ATP release from ARPE-19 cells triggered with a 50% hypotonic solution. A 10-min preincubation with BFA was not enough to produce a significant block, while a 2-h preincubation with BFA did reduce the ATP levels surrounding ARPE-19 cells. *P < 0.001, significantly different from control. **P < 0.001, significantly different from hypotonic. C: 2-h preincubation with 10 µg/ml BFA completely blocked the ATP release induced by the cAMP-activating mixture, while a 10-min incubation led to a small but insignificant reduction in ATP. *P < 0.001, significantly different from control. **P < 0.001, significantly different from hypotonic condition. D: when baseline levels of ATP were examined, a 2-h preincubation with 10 µg/ml BFA was found to reduce basal ATP release to the bath, while a 10-min preincubation did not. *P < 0.002, significantly different from control. E: in the fresh bovine RPE eyecup, a 30-min preincubation with 10 µg/ml BFA led to a small but insignificant reduction in the ATP release induced by the cAMP-stimulating cocktail. *P = 0.02, significantly different from control. F: incubation with 20 µM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM for 30 min greatly reduced the release of ATP from ARPE-19 cells. *P < 0.001, significantly different from control. **P < 0.001, significantly different from hypotonic condition. Bars, means ± SE.

 
Several controls were performed to further explain the dramatic block of ATP release by BFA. While BFA had no effect on the luciferase assay (Fig. 1), the inhibition was clearly dependent on exposure time, with a 10-min preincubation producing a smaller reduction in ATP release. Hypotonicity increased ATP levels by 4.9 ± 0.11-fold after a 10-min incubation in BFA (n = 10; P = 0.15, not significantly different from hypotonic alone) (Fig. 7B). Likewise, the reduction in cAMP-triggered release found after a 10-min incubation with BFA was not significant (Fig. 7C). BFA had a similar effect on the baseline release of ATP; preincubation for 10 min reduced basal levels by 13% (from 1.5 ± 0.08 nM; n = 24 to 1.3 ± 0.1 nM; n = 12), while a 2-h preincubation led to a significant reduction of 27% (to 1.1 ± 0.07 nM; n = 12) (Fig. 7D).

BFA also blocked the cAMP-stimulated ATP release across the apical membrane of fresh bovine RPE cells. In these experiments, the cAMP cocktail increased the ATP levels 1.6-fold, to 64.7 ± 12.9 nM (n = 12) from control levels of 46.0 ± 8.8 nM (n = 12). A 30-min incubation with BFA produced a small but not significant reduction in ATP concentration to 52.4 ± 9.7 nM (n = 11) (Fig. 7E).

Because vesicular release is known to require Ca2+ (8), the effect of the Ca2+ chelator BAPTA on ATP release was examined. Incubation of ARPE-19 cells in cell-permeant BAPTA-AM for 30 min reduced the ATP release triggered by hypotonicity by >77% (Fig. 7F). Levels were increased from 0.55 ± 0.02 nM in control (n = 16) to 5.63 ± 0.82 nM in hypotonic conditions (n = 14; 10.2-fold increase) but fell to only 1.68 ± 0.29 nM in the presence of BAPTA and hypotonicity (n = 16; 1.7-fold increase from control). This indicates that ATP release requires intracellular Ca2+ and further supports a role for the vesicular process in the release of ATP from RPE cells.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In the present study, the potential involvement of CFTR and vesicular transport in the release of ATP from RPE cells was examined using fresh and cultured RPE cells. On the basis of the following observations, we conclude that CFTR can affect RPE physiology through its involvement in ATP release. Stimulating CFTR with a cAMP-activating cocktail led to ATP release from ARPE-19 cells and across the apical membrane of the fresh RPE eyecup. In these preparations, the hypotonically triggered release was prevented by both glibenclamide and the specific inhibitor CFTR-172. The elevation of intracellular Ca2+ that followed a hypotonic challenge was prevented by the blocker CFTR-172. The transport inhibitor BFA and the Ca2+ chelator BAPTA prevented ATP release, consistent with a role for vesicular transport in the process.

CFTR-172 was recently identified as a high-affinity inhibitor of CFTR that does not block volume-sensitive Cl channels, Ca2+-sensitive Cl channels, KATP channels, AQP1, AE1, NHE3, or MDR-1 (28). It produces a voltage-independent block and does not affect cAMP production, phosphatase activity, or single-channel conductance, but leads to a reduction in the mean closed time of the channel and may effect channel gating by binding to NBD-1 (28, 56). Inhibition of anion transport by CFTR was maximal 10 min after application of CFTR-172, consistent with the enhanced inhibition of ATP release from RPE cells after preincubation. While the IC50 for ATP release was slightly higher than that found for anion influx or measurements of short-circuit current (28, 56), this is consistent with a more negative membrane potential in RPE cells. In addition, 10 µM CFTR-172 produced maximal block both of ATP release in the present study and of anion flux from thyroid epithelial cells in previous work (28). The effect of CFTR-172 on ATP release provides strong support for a role of CFTR in RPE physiology and is consistent with the idea that the Cl and ATP efflux pathways share a common gating mechanism.

The block of ATP release with CFTR-172 also strengthens the link between ATP release and CFTR. CFTR has been associated with ATP efflux since Reisin et al. (45) indicated overexpression of CFTR led to increased ATP efflux. While additional conduits for ATP efflux are likely (11, 18, 47) and release from some cells can clearly occur in the absence of CFTR (18, 19, 37, 61), the activity of CFTR is correlated with the augmentation of both baseline and stimulated ATP release in many cell types (33, 49), and the presence of immunopurified CFTR in lipid bilayers is linked to ATP conductance (9). ATP was initially thought to permeate the CFTR pore used by Cl, but the precise pathway of ATP movement is now a matter of some debate. Selective elimination of Cl and ATP conductance with different site mutations suggests that both CFTR and a separate cofactor are necessary for ATP efflux (23). Anion channel blockers diphenylamine carboxylate and 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid were found to have differential effects on the Cl and ATP conductance at the single-channel level, although the two pathways shared a gating mechanism (54). The ability of CFTR-172 to prevent the release of ATP provides further evidence that one gating mechanism controls the passage of both anions (56) and identifies CFTR-172 as an important tool to explore the role of CFTR in release. However, the effects of CFTR-172 on Cl and ATP currents must be examined at the single-channel level to provide further insight into the relationship between CFTR and ATP release.

Brefeldin A and CFTR-172 both blocked >70% of the hypotonically triggered release of ATP. This overlap implies that both processes apply to the same population of released ATP and act in series. Freshly synthesized CFTR has a half-life of 16 h (51), but CFTR on the cell surface can be internalized at 5%/min (27). Because a 2-h preincubation with BFA completely inhibited the ATP release from our preparation, this process is unlikely to be dependent on synthesis of new protein but may instead involve recycled material. BFA has traditionally been associated with vesicular transport in the Golgi, but it can also interfere with a range of trafficking events, including transport of recycling endosomes to the plasma membrane. BFA prevented the formation of clathrin-coated buds at endosomes with <30-min incubation and reduced the clathrin-dependent endosomal recycling of transferrin within 10 min (53, 57). The ability of cAMP to stimulate the insertion of CFTR-containing vesicles into the membrane has been hotly debated, with the process occurring in some cells but not in others (3, 38, 62). At least part of the inhibition of ATP release by BFA in RPE cells indicated above is likely to be on the number of CFTR proteins already present in the membrane, because the block was time dependent and was observed in association with baseline levels of ATP released from unstimulated cells. Whether the larger increase in extracellular ATP levels after hypotonic stimulation reflects the insertion of CFTR-containing vesicles into the membrane as seen with other channel types (16, 58) awaits biochemical assessment of CFTR localization in stimulated and unstimulated cells. Regardless of the reason for this phenomenon, the ability of BFA to prevent ATP release by reducing the amount of CFTR reaching the membrane may reconcile opposing "channel" and "vesicular" theories about the release of ATP. Although the vesicular transport pathway has previously been implicated in the release of ATP from non-neuronal cells, the vesicles were assumed to contain transmitters (1, 5, 30, 58). The hybrid mechanism illustrated in Fig. 8 may underlie ATP release from cells in which roles for both anion channels and vesicular transport have been identified.



View larger version (31K):
[in this window]
[in a new window]
 
Fig. 8. The hybrid mechanism in the ATP release from cultured human ARPE-19 and fresh bovine RPE cells. The hybrid hypothesis is a mixture of 2 alternative theories (hypotheses 1 and 2) to explain the mechanisms of ATP release from epithelial cells. In hypothesis 1, the ATP is released through a Cl channel. The block of ATP release by 5-nitro-2-(3-phenylpropylamino)-benzoate (NPPB) supports this hypothesis, while the activation by cAMP and the inhibition observed with CFTR-172 and glibenclamide imply that the CFTR molecule is involved. However, the BFA and BAPTA experiments indicated that the vesicular pathway also contributed to ATP release from RPE cells. The hybrid hypothesis accommodates all of the data by proposing that the insertion of CFTR and/or Cl channels into the membrane by vesicular fusion is necessary for ATP release. This insertion may represent a stimulated and/or constitutive process. Bars, means ± SE.

 
The release of ATP into the interior of the bovine RPE eyecup is consistent with a functional localization of CFTR to the RPE apical membrane. In the absence of the retina, the eyecup interior represents subretinal space, and ATP sampled from the eyecup interior is most likely to have moved from the RPE interior across the apical membrane into the space. CFTR has been immunolocalized to the apical and basolateral membranes of human RPE (4, 63). While release from both membranes has been shown in airway epithelial cells (20), the present study confirms only release across the apical membrane of fresh bovine RPE cells.

It is tempting to assume that the increase in subretinal ATP after the stimulation of apical CFTR would autostimulate P2Y2 receptors and elevate Ca2+ in vivo as shown in Fig. 6. This rise in Ca2+ may open basolateral Ca+-sensitive Cl channels and increase retinal to basolateral fluid absorption if the released ATP acts like the P2Y2 agonists (29, 31). The released ATP may also stimulate P2X receptors present on the RPE (46), but because their activation opens a cation channel, stimulation would further increase intracellular Ca2+. The effects of cAMP on the RPE are complex, possibly reflecting species differences or the action of CFTR on the apical and basolateral membranes in addition to other mechanisms (4, 22). The activation of basolateral Cl conductance after apical autostimulation with ATP would add an additional level of complexity that may underlie some of the variation in effects attributed directly to cAMP. However, the ability of CFTR-172 to alter Ca2+ physiology shows a definitive role for the transporter, and in patients with cystic fibrosis, the fast oscillation of the electrooculogram was significantly reduced (33), suggesting that CFTR function in wild types is regulated by light. It will be interesting to see whether subretinal ATP levels are also affected by light.


    GRANTS
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was funded by National Eye Institute Grant R01 EY-13434 (to C. H. Mitchell) and National Eye Institute Core Vision Grant EY-01583.


    ACKNOWLEDGMENTS
 
We thank W. W. Reenstra for his advice on CFTR pharmacology.


    FOOTNOTES
 

Address for reprint requests and other correspondence: C. H. Mitchell, Dept. of Physiology, Univ. of Pennsylvania, 3700 Hamilton Walk, Philadelphia, PA 19104-6085 (E-mail: chm{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.


    REFERENCES
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Aleu J, Martin-Satué M, Navarro P, Pérez de Lara I, Bahima L, Marsal J, and Solsona C. Release of ATP induced by hypertonic solutions in Xenopus oocytes. J Physiol 547: 209–219, 2003.[Abstract/Free Full Text]

2. Anderson MP, Rich DP, Gregory RJ, Smith AE, and Welsh MJ. Generation of cAMP-activated chloride currents by expression of CFTR. Science 251: 679–682, 1991.[ISI][Medline]

3. Bertrand CA and Frizzell RA. The role of regulated CFTR trafficking in epithelial secretion. Am J Physiol Cell Physiol 285: C1–C18, 2003.[Abstract/Free Full Text]

4. Blaug S, Quinn R, Quong J, Jalickee S, and Miller SS. Retinal pigment epithelial function: a role for CFTR? Doc Ophthalmol 106: 43–50, 2003.[CrossRef][ISI][Medline]

5. Bodin P and Burnstock G. Evidence that release of adenosine triphosphate from endothelial cells during increased shear stress is vesicular. J Cardiovasc Pharmacol 38: 900–908, 2001.[CrossRef][ISI][Medline]

6. Bodin P and Burnstock G. Purinergic signalling: ATP release. Neurochem Res 26: 959–969, 2001.[CrossRef][ISI][Medline]

7. Braunstein GM, Roman RM, Clancy JP, Kudlow BA, Taylor AL, Shylonsky VG, Jovov B, Peter K, Jilling T, Ismailov II, Benos DJ, Schwiebert LM, Fitz JG, and Schwiebert EM. Cystic fibrosis transmembrane conductance regulator facilitates ATP release by stimulating a separate ATP release channel for autocrine control of cell volume regulation. J Biol Chem 276: 6621–6630, 2001.[Abstract/Free Full Text]

8. Burgoyne RD and Clague MJ. Calcium and calmodulin in membrane fusion. Biochim Biophys Acta 1641: 137–143, 2003.[CrossRef][ISI][Medline]

9. Cantiello HF, Jackson GR Jr, Grosman CF, Prat AG, Borkan SC, Wang Y, Reisin IL, O'Riordan CR, and Ausiello DA. Electrodiffusional ATP movement through the cystic fibrosis transmembrane conductance regulator. Am J Physiol Cell Physiol 274: C799–C809, 1998.[Abstract/Free Full Text]

10. Chader GJ, Pepperberg DR, Crouch R, and Wiggert B. Retinoids and the retinal pigment epithelium. In: The Retinal Pigment Epithelium: Function and Disease, edited by Marmor MF and Wolfensberger TJ. New York: Oxford University, 1998, p. 135–151.

11. Cotrina ML, Lin JH, Alves-Rodrigues A, Liu S, Li J, Azmi-Ghadimi H, Kang J, Naus CC, and Nedergaard M. Connexins regulate calcium signaling by controlling ATP release. Proc Natl Acad Sci USA 95: 15735–15740, 1998.[Abstract/Free Full Text]

12. Dunn KC, Marmorstein AD, Bonilha VL, Rodriguez-Boulan E, Giordano F, and Hjelmeland LM. Use of the ARPE-19 cell line as a model of RPE polarity: basolateral secretion of FGF5. Invest Ophthalmol Vis Sci 39: 2744–2749, 1998.[Abstract]

13. Edelman JL and Miller SS. Epinephrine stimulates fluid absorption across bovine retinal pigment epithelium. Invest Ophthalmol Vis Sci 32: 3033–3040, 1991.[Abstract]

14. Eldred JA, Sanderson J, Wormstone M, Reddan JR, and Duncan G. Stress-induced ATP release from and growth modulation of human lens and retinal pigment epithelial cells. Biochem Soc Trans 31: 1213–1215, 2003.[ISI][Medline]

15. Gallemore RP and Steinberg RH. Effects of dopamine on the chick retinal pigment epithelium. Membrane potentials and light-evoked responses. Invest Ophthalmol Vis Sci 31: 67–80, 1990.[Abstract]

16. Gatof D, Kilic G, and Fitz JG. Vesicular exocytosis contributes to volume-sensitive ATP release in biliary cells. Am J Physiol Gastrointest Liver Physiol 286: G538–G546, 2004.[Abstract/Free Full Text]

17. Gregory CY, Abrams TA, and Hall MO. Stimulation of A2 adenosine receptors inhibits the ingestion of photoreceptor outer segments by retinal pigment epithelium. Invest Ophthalmol Vis Sci 35: 819–825, 1994.[Abstract]

18. Grygorczyk R and Hanrahan JW. CFTR-independent ATP release from epithelial cells triggered by mechanical stimuli. Am J Physiol Cell Physiol 272: C1058–C1066, 1997.[Abstract/Free Full Text]

19. Hazama A, Shimizu T, Ando-Akatsuka Y, Hayashi S, Tanaka S, Maeno E, and Okada Y. Swelling-induced, CFTR-independent ATP release from a human epithelial cell line: lack of correlation with volume-sensitive Cl channels. J Gen Physiol 114: 525–533, 1999.[Abstract/Free Full Text]

20. Homolya L, Steinberg TH, and Boucher RC. Cell to cell communication in response to mechanical stress via bilateral release of ATP and UTP in polarized epithelia. J Cell Biol 150: 1349–1360, 2000.[Abstract/Free Full Text]

21. Huang P, Lazarowski ER, Tarran R, Milgram SL, Boucher RC, and Stutts MJ. Compartmentalized autocrine signaling to cystic fibrosis transmembrane conductance regulator at the apical membrane of airway epithelial cells. Proc Natl Acad Sci USA 98: 14120–14125, 2001.[Abstract/Free Full Text]

22. Hughes BA, Miller SS, and Machen TE. Effects of cyclic AMP on fluid absorption and ion transport across frog retinal pigment epithelium: measurements in the open-circuit state. J Gen Physiol 83: 875–899, 1984.[Abstract]

23. Jiang Q, Mak D, Devidas S, Schwiebert EM, Bragin A, Zhang Y, Skach WR, Guggino WB, Foskett JK, and Engelhardt JF. Cystic fibrosis transmembrane conductance regulator-associated ATP release is controlled by a chloride sensor. J Cell Biol 143: 645–657, 1998.[Abstract/Free Full Text]

24. Joseph DP and Miller SS. Alpha-1-adrenergic modulation of K and Cl transport in bovine retinal pigment epithelium. J Gen Physiol 99: 263–290, 1992.[Abstract]

25. Joseph SM, Buchakjian MR, and Dubyak GR. Colocalization of ATP release sites and ecto-ATPase activity at the extracellular surface of human astrocytes. J Biol Chem 278: 23331–23342, 2003.[Abstract/Free Full Text]

26. Loewen ME, Smith NK, Hamilton DL, Grahn BH, and Forsyth GW. CLCA protein and chloride transport in canine retinal pigment epithelium. Am J Physiol Cell Physiol 285: C1314–C1321, 2003.[Abstract/Free Full Text]

27. Lukacs GL, Segal G, Kartner N, Grinstein S, and Zhang F. Constitutive internalization of cystic fibrosis transmembrane conductance regulator occurs via clathrin-dependent endocytosis and is regulated by protein phosphorylation. Biochem J 328: 353–361, 1997.[ISI][Medline]

28. Ma T, Thiagarajah JR, Yang H, Sonawane ND, Folli C, Galietta LJ, and Verkman AS. Thiazolidinone CFTR inhibitor identified by high-throughput screening blocks cholera toxin-induced intestinal fluid secretion. J Clin Invest 110: 1651–1658, 2002.[Abstract/Free Full Text]

29. Maminishkis A, Jalickee S, Blaug SA, Rymer J, Yerxa BR, Peterson WM, and Miller SS. The P2Y2 receptor agonist INS37217stimulates RPE fluid transport in vitro and retinal reattachment in rat. Invest Ophthalmol Vis Sci 43: 3555–3566, 2002.[Abstract/Free Full Text]

30. Maroto R and Hamill OP. Brefeldin A block of integrin-dependent mechanosensitive ATP release from Xenopus oocytes reveals a novel mechanism of mechanotransduction. J Biol Chem 276: 23867–23872, 2001.[Abstract/Free Full Text]

31. Meyer CH, Hotta K, Peterson WM, Toth CA, and Jaffe GJ. Effect of INS37217 a P2Y2 receptor agonist, on experimental retinal detachment and electroretinogram in adult rabbits. Invest Ophthalmol Vis Sci 43: 3567–3574, 2002.[Abstract/Free Full Text]

32. Miller SS, Hughes BA, and Machen TE. Fluid transport across retinal pigment epithelium is inhibited by cyclic AMP. Proc Natl Acad Sci USA 79: 2111–2115, 1982.[Abstract]

33. Miller SS, Rabin J, Strong T, Iannuzzi M, Adams A, Collins F, Reenstra W, and McCray P. Cystic-fibrosis (CF) gene product is expressed in retina and retinal pigment epithelium (Abstract). Invest Ophthalmol Vis Sci 33: 1009, 1991.

34. Miller SS and Steinberg RH. Active transport of ions across frog retinal pigment epithelium. Exp Eye Res 25: 235–248, 1977.[ISI][Medline]

35. Miller SS and Steinberg RH. Passive ionic properties of frog retinal pigment epithelium. J Membr Biol 36: 337–372, 1977.[ISI][Medline]

36. Mitchell CH. Release of ATP by a human retinal pigment epithelial cell line: potential for autocrine stimulation through subretinal space. J Physiol 534: 193–202, 2001.[Abstract/Free Full Text]

37. Mitchell CH, Carre DA, McGlinn AM, Stone RA, and Civan MM. A release mechanism for stored ATP in ocular ciliary epithelial cells. Proc Natl Acad Sci USA 95: 7174–7178, 1998.[Abstract/Free Full Text]

38. Moyer BD, Loffing J, Schwiebert EM, Loffing-Cueni D, Halpin PA, Karlson KH, Ismailov II, Guggino WB, Langford GM, and Stanton BA. Membrane trafficking of the cystic fibrosis gene product, cystic fibrosis transmembrane conductance regulator, tagged with green fluorescent protein in Madin-Darby canine kidney cells. J Biol Chem 273: 21759–21768, 1998.[Abstract/Free Full Text]

39. Nash M, Flanigan T, Leslie R, and Osborne N. Serotonin-2A receptor mRNA expression in rat retinal pigment epithelial cells. Ophthalmic Res 31: 1–4, 1999.[ISI][Medline]

40. Peterson WM, Meggyesy C, Yu K, and Miller SS. Extracellular ATP activates calcium signaling, ion, and fluid transport in retinal pigment epithelium. J Neurosci 17: 2324–2337, 1997.[Abstract/Free Full Text]

41. Prat AG, Reisin IL, Ausiello DA, and Cantiello HF. Cellular ATP release by the cystic fibrosis transmembrane conductance regulator. Am J Physiol Cell Physiol 270: C538–C545, 1996.[Abstract/Free Full Text]

42. Quinn RH, Quong JN, and Miller SS. Adrenergic receptor activated ion transport in human fetal retinal pigment epithelium. Invest Ophthalmol Vis Sci 42: 255–264, 2001.[Abstract/Free Full Text]

43. Reigada D and Mitchell CH. ATP release across the apical membrane of fresh bovine RPE (Abstract). Invest Ophthalmol Vis Sci 44: 384, 2003.

44. Reigada D and Mitchell CH. CFTR and calcium are required for ATP release from both fresh and cultured RPE cells (Abstract). Invest Ophthalmol Vis Sci 45: 423, 2004.[Abstract/Free Full Text]

45. Reisin IL, Prat AG, Abraham EH, Amara JF, Gregory RJ, Ausiello DA, and Cantiello HF. The cystic fibrosis transmembrane conductance regulator is a dual ATP and chloride channel. J Biol Chem 269: 20584–20591, 1994.[Abstract/Free Full Text]

46. Ryan JS, Baldridge WH, and Kelly ME. Purinergic regulation of cation conductances and intracellular Ca2+ in cultured rat retinal pigment epithelial cells. J Physiol 520: 745–759, 1999.[Abstract/Free Full Text]

47. Sabirov RZ, Dutta AK, and Okada Y. Volume-dependent ATP-conductive large-conductance anion channel as a pathway for swelling-induced ATP release. J Gen Physiol 118: 251–266, 2001.[Abstract/Free Full Text]

48. Schwiebert EM. ATP release mechanisms, ATP receptors and purinergic signalling along the nephron. Clin Exp Pharmacol Physiol 28: 340–350, 2001.[CrossRef][ISI][Medline]

49. Schwiebert EM, Egan ME, Hwang TH, Fulmer SB, Allen SS, Cutting GR, and Guggino WB. CFTR regulates outwardly rectifying chloride channels through an autocrine mechanism involving ATP. Cell 81: 1063–1073, 1995.[ISI][Medline]

50. Schwiebert EM and Zsembery A. Extracellular ATP as a signaling molecule for epithelial cells. Biochim Biophys Acta 1615: 7–32, 2003.[ISI][Medline]

51. Sharma M, Pampinella F, Nemes C, Benharouga M, So J, Du K, Bache KG, Papsin B, Zerangue N, Stenmark H, and Lukacs GL. Misfolding diverts CFTR from recycling to degradation: quality control at early endosomes. J Cell Biol 164: 923–933, 2004.[Abstract/Free Full Text]

52. Sprague RS, Ellsworth ML, Stephenson AH, Kleinhenz ME, and Lonigro AJ. Deformation-induced ATP release from red blood cells requires CFTR activity. Am J Physiol Heart Circ Physiol 275: H1726–H1732, 1998.[Abstract/Free Full Text]

53. Stoorvogel W, Oorschot V, and Geuze HJ. A novel class of clathrin-coated vesicles budding from endosomes. J Cell Biol 132: 21–33, 1996.[Abstract]

54. Sugita M, Yue Y, and Foskett JK. CFTR Cl channel and CFTR-associated ATP channel: distinct pores regulated by common gates. EMBO J 17: 898–908, 1998.[Abstract/Free Full Text]

55. Sullivan DM, Erb L, Anglade E, Weisman GA, Turner JT, and Csaky KG. Identification and characterization of P2Y2 nucleotide receptors in human retinal pigment epithelial cells. J Neurosci Res 49: 43–52, 1997.[CrossRef][ISI][Medline]

56. Taddei A, Folli C, Zegarra-Moran O, Fanen P, Verkman AS, and Galietta LJV. Altered channel gating mechanism for CFTR inhibition by a high-affinity thiazolidinone blocker. FEBS Lett 558: 52–56, 2004.[CrossRef][ISI][Medline]

57. Van Dam EM and Stoorvogel W. Dynamin-dependent transferrin receptor recycling by endosome-derived clathrin-coated vesicles. Mol Biol Cell 13: 169–182, 2002.[Abstract/Free Full Text]

58. Van der Wijk T, Tomassen SF, Houtsmuller AB, de Jonge HR, and Tilly BC. Increased vesicle recycling in response to osmotic cell swelling: cause and consequence of hypotonicity-provoked ATP release. J Biol Chem 278: 40020–40025, 2003.[Abstract/Free Full Text]

59. Versaux-Botteri C, Gibert JM, Nguyen-Legros J, and Vernier P. Molecular identification of a dopamine D1b receptor in bovine retinal pigment epithelium. Neurosci Lett 237: 9–12, 1997.[CrossRef][ISI][Medline]

60. Walsh DE, Harvey BJ, and Urbach V. CFTR regulation of intracellular calcium in normal and cystic fibrosis human airway epithelia. J Membr Biol 177: 209–219, 2000.[CrossRef][ISI][Medline]

61. Watt WC, Lazarowski ER, and Boucher RC. Cystic fibrosis transmembrane regulator-independent release of ATP: its implications for the regulation of P2Y2 receptors in airway epithelia. J Biol Chem 273: 14053–14058, 1998.[Abstract/Free Full Text]

62. Webe WM, Segal A, Vankeerberghen A, Cassiman JJ, and Van Driessche W. Different activation mechanisms of cystic fibrosis transmembrane conductance regulator expressed in Xenopus laevis oocytes. Comp Biochem Physiol A Mol Integr Physiol 130: 521–531, 2001.[CrossRef][ISI][Medline]

63. Weng TX, Godley BF, Jin GF, Mangini NJ, Kennedy BG, Yu AS, and Wills NK. Oxidant and antioxidant modulation of chloride channels expressed in human retinal pigment epithelium. Am J Physiol Cell Physiol 283: C839–C849, 2002.[Abstract/Free Full Text]

64. Wills NK, Weng T, Mo L, Hellmich HL, Yu A, Wang T, Buchheit S, and Godley BF. Chloride channel expression in cultured human fetal RPE cells: response to oxidative stress. Invest Ophthalmol Vis Sci 41: 4247–4255, 2000.[Abstract/Free Full Text]

65. Yamazaki J and Hume JR. Inhibitory effects of glibenclamide on cystic fibrosis transmembrane regulator, swelling-activated, and Ca2+-activated Cl channels in mammalian cardiac myocytes. Circ Res 81: 101–109, 1997.[Abstract/Free Full Text]

66. Young RW and Bok D. Participation of the retinal pigment epithelium in the rod outer segment renewal process. J Cell Biol 42: 392–403, 1969.[Abstract/Free Full Text]