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Journal of Physiology (2001), 534.1, pp. 193-202
© Copyright 2001 The Physiological Society
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ABSTRACT |
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, 
-methylene ADP prevented this, suggesting that inhibition was mediated by extracellular conversion of ATP to adenosine.
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INTRODUCTION |
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The retinal pigment epithelium (RPE) is a monolayer of cells between the blood supply of the choroid and the photoreceptors of the retina. The apical surface of the RPE is covered with microvilli which interdigitate with the photoreceptors. This intimate anatomical association underlies a strong functional relationship and the RPE performs several functions critical for the optimal performance of the photoreceptors including the maintenance of the ionic composition of subretinal space (Miller & Steinberg, 1977), the transport of fluid from retina to choroid (Edelman & Miller, 1991) and the phagocytosis of disc membranes shed from the rod outer segments (Young & Bok, 1969). RPE cells possess mRNA for both ATP (Sullivan et al. 1997) and adenosine receptors (Blazynski, 1993; Kvanta et al. 1997) and stimulation of these receptors can alter these critical functions of the RPE. Stimulation of A2b receptors reduces the rate of rod outer segment phagocytosis through a cAMP-dependent pathway in rat RPE cells (Gregory et al. 1994). Stimulation of P2Y2 purinoceptors sensitive to both ATP and UTP increases the apical to basolateral flow of fluid (Peterson et al. 1997) and P2Y2 agonists injected into subretinal space decrease the size of oedemas (Maminishkis et al. 2000).
The source of endogenous purines capable of stimulating these pathways remains unclear. Release of ATP has been reported from retinal astrocytes (Newman, 2001) and from inner retinal neurons (Neal & Cunningham, 1994; Santos, 1999). However, unlike many signalling molecules, the source of extracellular ATP need not be exclusively neuronal. Evidence is accumulating for discrete purinergic networks where localized release, stimulation and degradation closely regulate cell function (Wang et al. 1996; Mitchell et al. 1998; Hazama et al. 1999). The present study attempts to determine whether RPE cells could be a source of ATP in subretinal space capable of autocrine stimulation of P2 receptors. Specifically, the experiments aim to determine whether ATP can be released from ARPE-19 cells, whether the cells are capable of the extracellular degradation of ATP, and begin probing the mechanism and polarity of any release.
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METHODS |
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Tissue culture
The human ARPE-19 cell line (Dunn et al. 1996) was obtained from the American Type Culture Collection (Manassas, VA, USA) and grown in a 1:1 mixture of Dulbecco's modified Eagle's medium and Ham's F12 medium with 3 mM L-glutamine (all from Gibco BRL, Grand Island, NY, USA) initially with 10 % fetal bovine serum (FBS; HyClone Laboratories, Inc., Logan, UT, USA). Cells were incubated at 37 °C in 5 % CO2 and subcultured with 0.05 % trypsin and 0.02 % EDTA. Cells were usually grown in 1 % FBS for 4-7 weeks before plating on coverslips to encourage differentiation (Dunn et al. 1998).
ATP measurements
ARPE-19 cells were grown on glass coverslips in 10 % FBS for 3-14 days, washed 
3 and mounted on an inverted microscope. The solution bathing the cells included 2 mg ml-1 luciferin-luciferase assay mixture (Sigma Chemical Corp., St Louis, MO, USA). ATP released from cells into the extracellular bath reacted with the luciferase and led to the production of a photon. Light produced was captured with a 20 objective, filtered at 520 nm, measured with a photomultiplier tube and recorded online using the Felix software suite (Photon Technology International Inc., Princeton, NJ, USA). All experiments were performed at 37 °C.
The luminescence values were converted to concentrations of ATP using a standard curve. Standard luminescence signals were not altered by NPPB (Mitchell et al. 1998; Hazama et al. 1999), UTP or bFGF. The signal from ATP in hypotonic solution was significantly larger than that found in isotonic solution and thus experiments were calibrated separately from the relevant standard curve. In previously published work, the hypotonic solution did not affect the ATP standard curves (Mitchell et al. 1998), presumably because in the earlier work solutions were made hypotonic by removal of mannitol. In the present study, solutions were made hypotonic by dilution with water and the Km of luciferase for ATP is known to increase with ionic strength (Sigma Technical Bulletin, Sigma Chemical Co.)
Polarized release
Cells were grown on translucent anadolite permeable filters (0.2 µm pores, Nalgene Nunc International, Naperville, IL, USA) as ARPE-19 cells grown on such a substrate have previously been shown to establish polarity (Dunn et al. 1996, 1998). Cells were used 4-6 weeks after plating and after the first week, grown in medium with 1 % FBS. Cells grown under these conditions developed a transepithelial potential of 10.2 mV and a calculated transepithelial resistance of 150 
cm2, suggesting that tight junctions had formed between cells. Filters were placed in a chamber, mounted on an inverted microscope, luciferin-luciferase was added either to the compartment bathing the apical face of the cells or to the basolateral face, and the objective was focused upon either the apical or the basolateral membrane, as appropriate. The ability to detect basolaterally released ATP was tested by lysing cells with 1 % Triton X to release cytoplasmic ATP. Although luminescence increased when the luciferin-luciferase assay mix was present only in the basolateral chamber, the signal was 20-fold higher when the luciferin-luciferase assay mix was present only in the apical chamber, suggesting that the assay mix had not fully permeated the solution within the filter honeycomb.
Intracellular Ca2+ measurements
ARPE-19 cells grown on coverslips for 3-6 days were loaded with 10 µM fura-2 AM for 40 min at room temperature, rinsed and maintained in fura-2-free solution before data acquisition began 30 min later. The coverslips were mounted on a Nikon Diaphot microscope and visualized with a 40 oil-immersion fluorescence objective. The field was alternately excited at 340 and 380 nm and the fluorescence emitted at 520 nm in selected regions of interest around individual cells was imaged with a CCD camera and analysed using the ImageMaster software suite (Photon Technology International Inc.). The ratio of light excited at 340 nm to that at 380 nm was converted into Ca2+ concentration using previously published techniques (Mitchell et al. 2000). These experiments were performed at room temperature.
Phagocytosis assay
ARPE-19 cells were grown to confluence on glass coverslips. Fluorescent yellow-green beads of 1 mm diameter (Molecular Probes Inc., Eugene, OR, USA) were added at a concentration of 5 107 beads ml-1 to normal growth medium containing 10 % fetal bovine serum, as serum has been shown to be necessary for phagocytosis (Edwards, 1991). Cells were incubated at 37 °C in the bead-containing solution for 1 h, after which they were washed vigourously four times, mounted on a microscope and the number of beads quantified by measuring the total fluorescence present excited at 480-485 nm and emitted at 520 nm with a photomultiplier tube (Photon Technology International Inc.). Background levels obtained from cells without beads were subtracted from each measurement.
Solutions
Control isotonic solution contained (mM): 105 NaCl, 4.5 KCl, 2.8 NaHepes, 7.2 Hepes acid, 1.3 CaCl2, 0.5 MgCl2, 5 glucose, 75 mannitol. The pH was adjusted to 7.4 with NaOH, and the solution was 304-312 mosmol l-1. The solutions were made hypotonic by adding 50 or 33 % H2O. The luciferin-luciferase assay mixture was mixed daily as a 100 mg ml-1 stock solution and added to solutions to give a final concentration of 2 mg ml-1 (Taylor et al. 1998).
Materials
5-Nitro-2-(3-phenylpropylamino)-benzoate (NPPB) was obtained from Biomol Research Laboratories, Inc. (Plymouth Meeting, PA, USA). Fura-2 AM was obtained from Molecular Probes. Basic fibroblast growth factor (bFGF, bovine) was obtained from Becton Dickinson, Bedford, MA, USA. All other chemicals were from Sigma Chemical Co.
Statistical analysis
Values are presented as means ± 1 S.E.M. The number of experiments is indicated by the symbol n. The null hypothesis, that the experimental and baseline measurements shared the same mean, was tested with Student's t test.
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RESULTS |
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Release of ATP
The luciferin-luciferase assay system was used in real time to identify triggers of ATP release in ARPE-19 cells. Initial experiments were performed to determine whether hypotonic solution led to this release. Background levels of luminescence were monitored from cells in isotonic solution in the absence of luciferin-luciferase assay mixture. After 2 min, the isotonic solution was replaced with either an isotonic or hypotonic solution containing 2 mg ml-1 luciferin- luciferase assay mix. The change of solutions alone was sufficient to produce a small transient increase in luminescence, as seen in traces switched to isotonic solution. However, when control isotonic solution was replaced with hypotonic solution containing luciferin- luciferase assay mix, luminescence increased markedly (Fig. 1). After an initial rise, the signal increased ~2 min after presentation of hypotonic solution and reached a peak ~5 min after solution change, after which time the level slowly declined. Isotonic solution produced a much smaller delayed rise in ATP levels. At the peak, levels in hypotonic solution were 147.1 ± 45.3 nM (n = 5) while those in the control solution were only 44.1 ± 15.3 nM (n = 4, P < 0.01). The effect was dependent upon the degree of hypotonicity, with 33 % hypotonic solutions producing a proportionally smaller effect than 50 % hypotonic solutions (see below).
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Figure 1. Hypotonic solution triggers release of ATP Isotonic and hypotonic solutions both containing luciferin-luciferase assay mixture were added to RPE cells at t = 150 s. Hypotonic solution triggered the release of more ATP than isotonic solution, as determined by the increase in luminescence produced by ATP reacting with the luciferin-luciferase assay mixture. Luminescence levels were converted to concentration of ATP using standard curves. At the peak, levels in hypotonic (n = 5) were significantly different from those in isotonic solution (n = 4, P < 0.01). Here and in subsequent figures, the vertical lines present between 120 and 180 s represent room light reaching the chamber during solution changes, while continuous data lines represent the mean responses and dotted lines show the associated standard errors. | ||
The pyrimidine UTP triggered release of ATP in a concentration-dependent manner (Fig. 2A). Similar to the hypotonic response, the extracellular concentration showed an initial elevation, followed by a secondary rise that began approximately 2 min after presentation of UTP and reached a peak 3-5 min later. UDP was used to determine which receptor subtype was involved in the response, as UDP is more efficacious than UTP at P2Y6 receptors but has a much smaller effect at P2Y2 or P2Y4 receptors than UTP (Nichols et al. 1996). At both 1 mM and 100 µM, UTP triggered considerably more ATP release than UDP (Fig. 2B): control, 28.6 ± 6.9 nM (n = 10); 100 µM UTP, 131.3 ± 15.3 nM (n = 5); 1 mM UTP, 247.0 ± 51.8 nM (n = 4); 100 µM UDP, 26.3 ± 3.5 nM (n = 3) and 1 mM UDP, 66.1 ± 10.7 nM (n = 4, all measurements 5 min after addition).
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Figure 2. Stimulation of ATP release by UTP A, control and UTP solutions both containing luciferin-luciferase assay mixture were added to RPE cells at t = 150 s. UTP triggered a concentration-dependent release of ATP when compared with control solution, as determined by the increase in luminescence produced by ATP reacting with the luciferin- luciferase assay mixture. The traces are the mean responses of 10 (control), 5 (100 µM UTP) and 4 (1 mM UTP) experiments. B, mean response ± S.E.M. measured 5 min after agonist addition is greater for UTP than UDP. The number of experiments is shown below the bars. *P < 0.001 and †P < 0.025 vs. Control. | ||
ATP was also released into the bath when 200 ng ml-1 basic fibroblast growth factor (bFGF) was added to ARPE-19 cells (Fig. 3). Unlike the response to hypotonicity or UTP, the concentration of ATP in the extracellular space peaked early, ~75 s after addition of bFGF. At this peak, ATP levels were 165 ± 30 nM (n = 4) in the presence of bFGF and 35 ± 19 nM (n = 4) in control solution (P < 0.02). A secondary peak was observed 3-6 min later, and levels remained elevated throughout the 10 min duration of the measurements.
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Figure 3. Stimulation of ATP release by bFGF Control and 200 ng ml-1 bFGF solutions both containing luciferin-luciferase assay mixture were added to RPE cells at t = 150 s. At the peak response (t = 225 s, 75 s after addition of solutions), the concentration of ATP in the presence of bFGF was significantly greater than that in isotonic solution (P < 0.02, n = 4 for both). | ||
Retinal illumination induces a change in subretinal K+ concentration from 5 to 2 mM (Oakely & Green, 1976). However, neither decreasing K+ from 5 to 2 mM (n = 6) nor increasing K+ from 2 to 5 mM (n = 4) was sufficient to trigger release of ATP.
Mechanism of release
The Cl- channel blocker NPPB was used to determine whether Cl- channels were involved in the ATP release triggered by hypotonic solution. NPPB (100 µM) blocked > 90 % of the increase (Fig. 4A; isotonic solution, 55.1 ± 10.0 nM (n = 3); 33 % hypotonic solution, 104.2 ± 19.3 nM (n = 5); 33 % hypotonic + NPPB solution, 57.4 ± 11 nM (n = 5)). NPPB also blocked the release of ATP triggered after stimulation with UTP by ~86 % (Fig. 4B). At the peak response, the mean concentration of ATP in control solution was 21.1 ± 10.1 nM (n = 4); in 100 µM UTP, 121.0 ± 30.7 nM (n = 4); and in 100 µM UTP + 100 µM NPPB, 35.3 ± 4.3 nM (n = 4).
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Figure 4. NPPB blocks the release of ATP A, hypotonic release. Mean concentration of ATP measured 5 min after presentation of control isotonic solution ( | ||
Ca
was elevated directly with the Ca2+ ionophore ionomycin to test the Ca2+ dependence of ATP release from ARPE-19 cells. With 1.3 mM Ca2+ and luciferin- luciferase present in the bath, 5 µM ionomycin did not induce a significant rise in luminescence (Fig. 5). In control, levels were 32.3 ± 10.7 nM (n = 5) while those in 5 µM ionomycin were 30.4 ± 6.3 nM (n = 5). Although the ionomycin-treated cells showed a slight elevation at ~5 min, values were not significantly different from control, and a similar small elevation was sometimes seen in control solutions (e.g. Fig. 1).
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Figure 5. Elevating Ca2+ directly was insufficient to trigger ATP release Ionomycin (5 µM) was added to the cells in the presence of 1.3 mM Ca2+ in standard isotonic solution. The concentration of ATP in the bath after 5 min was not significantly different from control levels (n = 5 for both). | ||
Biphasic elevation of Ca
The real-time measurements of bath ATP shown in Figs 1, 2 and 3 show a biphasic elevation in extracellular levels of ATP, with concentrations beginning to increase for a second time ~2 min after the solution changes. This biphasic release may have implications for the kinetics of purinergic signalling. Activation of both P2Y2 and P2X receptors elevates Ca, but stimulation of P2Y2 receptors with UTP leads to transient elevation in Ca (Sullivan et al. 1997), while stimulation of P2X receptors can lead to influx of Ca2+ through the integral non-selective cation channel (Lynch et al. 1999). This leads to the prediction that UTP could produce a biphasic increase in Ca; the first a transient elevation primarily due to stimulation of P2Y2 receptors by UTP and the second a more steady, sustained elevation corresponding to activation of P2X receptors by the ATP released with delay. As shown in Fig. 6A, UTP (100 µM) triggered a sudden elevation in Ca, with a peak of 2130 ± 47 nM (n = 12). After this peak, levels gradually fell to 270 ± 2 nM then rose again to 360 ± 2 nM. The peak intracellular concentration of Ca2+ triggered by 100 µM UTP was unchanged in the presence of 100 µM NPPB, rising to 2183 ± 430 nM (n = 7; Fig. 6B), but levels fell more completely to 128 ± 9 nM before rising to only 166 ± 9 nM.
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Figure 6. NPPB blocks secondary rise in Ca2+ after stimulation with UTP Cells were loaded with the Ca2+-sensitive dye fura-2 and the ratio of fluorescence excited at 340 nm/380 nm was monitored in individual cells. A, UTP (100 µM) triggered a sudden elevation in Ca | ||
Polarized release
The physiological implications of ATP release from RPE cells depend upon the polarity of this release, for only ATP released across the apical membrane will be able to stimulate the receptors of interest facing the subretinal space. To investigate polarized release, cells were grown on permeable supports. Addition of 50 % hypotonic solution plus luciferin-luciferase to the apical compartment triggered a larger signal than the addition of isotonic solution plus luciferin-luciferase in 5/6 experiments (Fig. 7A). Hypotonic solution did not trigger a detectable release in ATP when luciferin-luciferase was added to the basolateral side of the filter (Fig. 7B), although basolateral release cannot be ruled out (see Methods). However, Fig. 7 shows that the small transient increases in bath ATP following solution changes were evident with both apical (A) and basolateral (B) placement of luciferin-luciferase, suggesting that release from some pools of ATP can occur and be detected across the basolateral membrane.
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Figure 7. Polarized release of ATP Cells grown on permeable supports had either the apical (A) or basolateral (B) faces bathed in solution containing luciferin-luciferase. When the luciferin-luciferase was on the apical membrane, hypotonic solution triggered a substantially larger response than isotonic solution, as found in 5/6 trials. The response to both isotonic and hypotonic solutions was similar and small when luciferin-luciferase was bathing the basolateral membrane. Values are shown as luminescence (a.l.u., arbitrary light units) as calibration to ATP levels was not performed on apical or basolateral faces of the permeable support. | ||
Degradation of ATP
To determine whether ARPE-19 cells are capable of the first step in converting ATP into adenosine, the initial degradation of ATP, the ability of intact cells to degrade extracellular ATP was tested. A solution containing 10-6 M ATP and 2 mg ml-1 luciferin-luciferase assay mixture was made immediately before being added to coverslips containing a confluent monolayer of ARPE-19 cells. The time-dependent reduction in light from coverslips in the absence of cells was used to control for the decrease in signal due to the utilization of enzyme substrates. Thus, degradation attributed specifically to the cells was defined as the difference in ATP levels in the presence and absence of cells. Extracellular ATP was degraded considerably more in the presence of cells (Fig. 8). As the cells were intact, this implies that enzymes present on the extracellular surface of ARPE-19 cells are capable of degradation of ATP. It is noteworthy that the cell-dependent degradation of ATP began with a delay of ~5 min.
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Figure 8. Degradation of added ATP Exogenous ATP (10-6 M) was added to the bath with luciferin-luciferase. The data show the difference in bath ATP levels between blank coverslips and those containing a confluent monolayer of RPE cells (n = 3). The degradation attributed to the cells began after a delay of 5 min, as shown by the negative values after 450 s. | ||
Phagocytosis
RPE cells possess ecto-5'-nucleotidase capable of converting AMP to adenosine (Takizawa, 1998) and adenosine has previously been shown to prevent phagocytosis of rod outer segments by rat RPE cells (Gregory et al. 1994). If the degradation of ATP in Fig. 8 represents dephosphorylation of ATP into ADP and AMP, extracellular ATP should inhibit phagocytosis. To test this hypothesis, the effect of ATP on the ability of ARPE-19 cells to phagocytose and/or bind to 1 mm fluorescent beads was tested. Figure 9 shows that ATP (100 µM) reduced the fluorescence due to beads from (8.7 ± 0.8) 104 a.l.u. (arbitrary light units) to (4.5 ± 0.6) 104 a.l.u. (n = 16, P < 0.001). When the ecto-5'-nucleotidase inhibitor ,-meADP (100 µM) was included with 100 µM ATP, the fluorescence rose to (9.4 ± 1.8) 104 a.l.u. (n = 16, different from ATP alone, P < 0.02). This value was not significantly different from control, suggesting that the inhibitory effect of ATP required the action of ecto-5'-nucleotidase and was consequently mediated by adenosine produced extracellularly.
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Figure 9. Effect of ATP on fluorescent bead uptake Fluorescence is an index of the retention of fluorescent beads by ARPE-19 cells. The reduction in fluorescence by 100 µM ATP was prevented by 100 µM | ||
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DISCUSSION |
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The results of the present study show that ARPE-19 cells release ATP. The release was biphasic and stimulated by basic fibroblast growth factor (bFGF), by the pyrimidine UTP, and by hypotonicity. The Cl- channel blocker NPPB inhibited release, suggesting that it was not due to damaged cells, but was probably associated with Cl- channels. Elevations in Ca were not sufficient to stimulate release. Cells grown on permeable filters released ATP across their apical membranes, analogous to release into subretinal space by cells in vivo. Extracellular ATP was degraded by intact cells and modified the rate of phagocytosis, possibly by conversion to adenosine by ecto-enzymes. Finally, the biphasic release of ATP corresponded to a biphasic increase in Ca that was inhibited by blocking ATP release with NPPB. Together, these observations suggest that RPE cells can release ATP into subretinal space at concentrations capable of autocrine stimulation of P2 receptors.
The ability of bFGF, UTP and hypotonicity to trigger release of ATP from ARPE-19 cells suggests that release may have several physiological roles. Hypotonic solution activates ionic conductances similar to those implicated in the response of the RPE to light (Botchkin & Matthews, 1993; Bialek & Miller, 1994). It remains to be tested whether ATP levels increase in subretinal space after retinal illumination, although changes in K+ concentration alone were insufficient to trigger release. UTP clearly increased extracellular levels of ATP and this probably represents release, as NPPB produced comparable blocks of the hypotonic and UTP effects. This would be unlikely if ATP was formed by transference of the terminal phosphate from UTP to ADP by ecto-nucleoside diphosphokinase (Lazarowski et al. 1997; Ostrom et al. 2000). The ability of bFGF to trigger release of ATP from ARPE-19 cells has important implications for the diseased eye, as photoreceptors, RPE cells and choroidal cells contain bFGF and levels can be elevated by injury (Wen et al. 1995) or macular degeneration (Amin et al. 1994). The role of extracellular ATP in these processes is currently unknown.
The absolute levels of ATP measured here correspond to between 100 and 300 nM. These levels are expected to be an underestimate as they do not control for the diffusion of ATP away from the release site before full measurements are made. Cells with membrane-bound luciferase produced a larger and more rapid increase in luminescence after stimulation than cells with soluble luciferase present in the bath (Beigi et al. 1999). The small subretinal volume also predicts higher concentrations in vivo. Regardless, the concentrations of ATP measured directly in this study would be sufficient to stimulate the P2Y2 receptor with an EC50 of 200 nM (Lazarowski et al. 1995) and 100 nM ATP altered the transepithelial potential across bovine RPE cells (Peterson et al. 1997), suggesting that the concentrations of ATP released by the cells are sufficient to autostimulate P2 receptors and could increase fluid absorption.
The reduction in luminescence after application of NPPB (Fig. 4) suggests that the release of ATP is somehow associated with a Cl- channel. A growing body of evidence suggests that release of ATP from epithelial cells is associated with Cl- channels, although the precise relationship remains controversial (Sugita et al. 1998; Watt et al. 1998; Lader et al. 2000). ATP release has been linked to the CFTR Cl- channel (Prat et al. 1996) and message for CFTR is present in RPE cells (Miller et al. 1992; Wills et al. 2000). However, NPPB also blocks Cl- channels and ATP release not associated with CFTR (Mitchell et al. 1998; Watt et al. 1998; Hazama et al. 1999), so the precise conduit for release in these cells remains unclear.
The sensitivity of the present assay, combined with the high temporal resolution of the real-time measurements, showed that the release of ATP was biphasic (Figs 1, 2 and 3). The initial elevation in ATP levels was larger in magnitude, but kinetically similar to that caused by a solution change (Grygorczyk & Hanrahan, 1997), while the secondary rise, peaking after 4-8 min, was clearly an independent event. These distinct responses could represent release from different pools of ATP, or separate intracellular pathways triggering the same pool. ATP released during the initial stage cannot be the main trigger for the secondary release, as early release levels do not necessarily correlate with the magnitude of the delayed peak (see Fig. 2A). It is interesting that the delayed peak coincided with the degradation of added ATP (Fig. 8), suggesting that the two processes could share intracellular messengers. Delayed release of ATP occurs in some reports (Hazama et al. 1999; Ostrom et al. 2000), but not in others (e.g. Roman et al. 1999), and these could be attributed to differences in the cellular mechanism or experimental methodology.
The biphasic elevation of extracellular ATP and intracellular Ca2+ may be causally linked, with either Ca leading to ATP release, or released ATP leading to elevated Ca. The inability of ionomycin to elevate extracellular ATP suggests that elevated Ca is insufficient to trigger ATP release, but ATP can elevate Ca in both cultured rat (Stalmans & Himpens, 1996) and human RPE cells (Sullivan et al. 1997). This suggests that the long-term elevations in Ca triggered by UTP may involve ATP release and autostimulation. These secondary elevations were inhibited by NPPB, consistent with a block of ATP release by NPPB (Fig. 4), although this may also be attributed to a block of the Ca2+ release-activated Ca2+ channel by NPPB (Li et al. 2000). The ability of UTP to trigger ATP release implies that the increase in fluid transport induced by UTP (Peterson et al. 1997) or UTP analogues (Maminishkis et al. 2000) may be augmented by released ATP and could involve the action of P2 receptors unresponsive to UTP, such as the P2X receptors present in RPE cells (Ryan et al. 1999).
ATP released into the subretinal space by RPE cells could stimulate receptors in other cell types. P2X2 receptors have been identified on the outer segments of photoreceptors (Greenwood et al. 1997), and stimulation of these receptors with ATP can lead to an influx of Ca2+ (Lynch et al. 1999). ATP at the distal face can trigger waves of Ca2+ throughout Müller cells (Keirstead et al. 1999). Adenosine levels in the distal retina are elevated at night and enhance circadian synthesis of melatonin (Iuvone et al. 2000). Ecto-enzymes capable of converting AMP to adenosine at physiological pH were most active on RPE cells in the dark (Irons, 1987; Irons & O'Brien, 1987), suggesting that release of ATP by the RPE cells may be involved in the circadian regulation of the photoreceptor function and phagocytosis. Finally, the ability of RPE cells to convert extracellular ATP into adenosine will be relevant whether the ATP is released from the RPE or retinal neurons (Neal & Cunningham, 1994; Santos et al. 1999).
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Acknowledgements
The author would like to thank Mortimer M. Civan for his many useful suggestions, his support, and for the generous use of his laboratory. Appreciation is also due to Kim Peterson Yantorno, to Richard Stone, Jean Bennett and Edward N. Pugh Jr for stimulating discussions and to Alan Laties for his support. This work was supported by NEI grants EY-08343-10 and EY-12213-02 to Mortimer Civan, and a grant from the Foundation Fighting Blindness to Alan Laties.
Correspondence
C. H. Mitchell: Department of Physiology, University of Pennsylvania, A303 Richards Building, Philadelphia, PA 19104-6085, USA.
Email: chm{at}mail.med.upenn.edu
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, n = 3), 33 % hypotonic solution (Hypo, 
, n = 5) or 33 % hypotonic + 100 µM NPPB solutions (
, n = 5). * Different from isotonic, P < 0.05; † different from hypotonic, P < 0.05. B, UTP-triggered release. At the peak (t = 6.5 min), the elevation in ATP levels triggered by 100 µM UTP was blocked ~86 % by the addition of 100 µM NPPB. * Different from Control, P < 0.01; † different from UTP, P < 0.025.