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1 Department of Physiology, University of Pennsylvania, Philadelphia, PA, USA
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Abstract |
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(Received 1 June 2006;
accepted after revision 25 June 2006;
first published online 29 June 2006)
Corresponding author C. H. Mitchell: Department of Physiology, University of Pennsylvania, 3700 Hamilton Walk, Philadelphia, PA 19104-6085, USA. Email: chm{at}mail.med.upenn.edu
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Introduction |
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The photoreceptors are located between the inner retina and the retinal pigment epithelium (RPE). The RPE monolayer performs many functions to support the photoreceptors, such as controlling the ion and fluid levels in the small subretinal space which separates the two cell types (Miller & Steinberg, 1977a,b; Miller et al. 1982; Hughes et al. 1984; Edelman & Miller, 1991) and supplying the neural retina with nutrients, growth factors and recycled visual pigments (Young & Bok, 1969; Chader et al. 1998). The transmitters dopamine (Gallemore & Steinberg, 1990; Versaux-Botteri et al. 1997) serotonin (Nash & Osborne, 1997; Nash et al. 1999) and adrenaline (Edelman & Miller, 1991; Joseph & Miller, 1992; Quinn et al. 2001) may influence retinal–RPE communication by stimulating receptors on RPE cells. While these transmitters all contribute to visual signalling in the retina, glutamate is the primary transmitter released by photoreceptors, with release rates inversely proportional to light levels (Witkovsky et al. 1997). Although glutamate is well known to act on bipolar cells to initiate visual signalling (Bloomfield & Dowling, 1985; Copenhagen, 1991), glutamate released from photoreceptors might also diffuse in the opposite direction towards the apical tips of the RPE.
Considerable evidence suggests that glutamate signalling is important for RPE function. Glutamate transporters EAAT4 and EAAT1 are present on the RPE and are expected to restrict extracellular levels of glutamate in the subretinal space (Maenpaa et al. 2002, 2003, 2004). Ionotropic and metabotropic glutamate receptors have been identified in chick and human RPE cells (Lopez-Colome et al. 1993; Lopez-Colome & Fragoso, 1995), while activation of NMDA receptors (NMDARs) in RPE cells increases proliferation (Uchida et al. 1998) and inositol triphosphate (IP3) levels (Fragoso & Lopez-Colome, 1999). In glial cells, propagation of the initial signal can occur when elevated IP3 levels lead to the release of ATP, which in turn stimulates purinergic P2 receptors on neighbouring cells (Sauer et al. 2000; Schwiebert, 2000). Such paracrine activation by released ATP is widely used to amplify signals, and interactions between purinergic and glutaminergic systems are involved in the bidirectional communication between neurons and astrocytes in the brain (Mazzanti et al. 2001).
Although it is not yet known whether released ATP amplifies the glutamate signal in RPE cells, many of the necessary components are present. In addition to the glutamate receptors described above, multiple receptor types for ATP are present on the RPE (Sullivan et al. 1997; Ryan et al. 1999; Reigada et al. 2005). Retinal pigment epithelium cells are also known to release ATP, which can in turn autostimulate the cells (Mitchell, 2001; Reigada & Mitchell, 2005a). We therefore asked whether glutamate acting on NMDARs could trigger ATP release from RPE cells, and whether this stimulation could modify intracellular signalling.
Portions of this work have been presented previously in abstract form (Reigada & Mitchell, 2005b; Reigada et al. 2006a).
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Methods |
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All chemicals were from Sigma-Aldrich, Inc. (St Louis, MO, USA) unless otherwise noted. The isotonic solution was composed of (mM): 105 NaCl, 5 KCl, 6 Hepes acid, 4 HepesNa2, 5 NaHCO3, 60 mannitol, 5 glucose, 0.5 MgCl2 and 1.3 CaCl2, pH 7.4. DL-2-Amino-5-phosphonopentanoic acid (D-AP5), 5,7-dichlorokynurenic acid (DCKA) and 5-nitro-2-(3-phenylpropyl-amino)-benzoate (NPPB) were dissolved in a dimethyl sulphoxide (DMSO) stock solution. N-Methyl-D-aspartate (NMDA), L-glutamic acid (glutamate), pyridoxal-phosphate-6-azophenyl-2',4'-disulphonate (PPADS), 18
-glycyrrhetinic acid (18GA), apyrase, oleamide, thapsigargin, N6-methyl-2'-deoxyadenosine-3',5'-bis-phosphate (MRS2179) and (+)-5-methyl-10,11-dihydro [a,d]cyclohepten-5,10-imine hydrogen maleate (MK-801) were prepared as aqueous stock solutions and diluted to the working concentration in isotonic solution. To control for non-specific effects that frequently complicate analysis of ATP release, all chemicals were tested for their effect on the luciferin–luciferase assay, as shown in Fig. 1.
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Release of ATP was determined with the luciferin–luciferase reporting system. Previous experience with this system has shown that the increased levels of extracellular ATP represent physiological release (Mitchell et al. 1998; Mitchell, 2001; Reigada et al. 2005,), although the concentrations detected are likely to be orders of magnitude below that found at the membrane surface (Joseph et al. 2003; Okada et al. 2006). The luciferase solution was prepared in a stock solution from one vial of the luciferin–luciferase assay kit (Sigma-Aldrich Inc.) diluted in 450 µl of the isotonic solution and 50 µl distilled water. This stock solution was stored at –20°C.
Bovine eyes were obtained from the abattoir and transported on ice to the laboratory, with the approval of the University of Pennsylvania Institutional Animal Care and Use Committee. Eyes were bisected at the ora serrata, with the retina removed and detached from the optic nerve. In this configuration, the apical membrane of the RPE faces the interior of the eyecup and, in the presence of intact tight junctions, polarity is likely to be maintained (Reigada & Mitchell, 2005a). After washing the RPE eyecup three times with isotonic solution, 500 µl of test solution was added to the eyecup. After 10 min at room temperature (21–25°C), a 400 µl sample was removed, rapidly frozen and stored at –20°C until analysis. To determine the ATP content of the solution from each eyecup, a luciferase working solution was prepared by diluting 40 µl of the stock solution described above in 1 ml isotonic buffer. The eyecup sample was defrosted, with 90 µl of sample placed in a single well of a 96-well plate, which was in turn put into the microplate luminometer (Luminoskan Ascent, Labsystems; Franklin, MA, USA), and 10 µl of the luciferase working solution was injected into each well. Measurements were taken at room temperature every 30 s for 10 min with an integration time of 100 ms per measurement. A standard curve showed the relationship between luminescence and ATP concentration to be linear over the range tested and was used to convert luminescence units into ATP concentration.
Cell culture
The ARPE-19 cell line (Dunn et al. 1998) was obtained from American type culture collection (ATCC) (Manassas, VA, USA) and grown to confluence in 25 cm2 Primaria culture flasks (Becton Dickinson, Franklin Lakes, NJ, USA) in a 1:1 mixture of Dulbecco's modified Eagle's medium (DMEM) and Ham's F12 medium with 3 mM L-glutamine, 100 µg ml–1 streptomycin and 2.5 mg ml–1 Fungizone and/or 50 µg ml–1 gentamicin (all Invitrogen Corp., Carlsbad, CA, USA) and 10% fetal bovine serum (FBS; Hyclone Laboratories, Logan, UT, USA). Cells were incubated at 37°C in 5% CO2 (with the remainder being air) and subcultured weekly with 0.05% trypsin and 0.02% EDTA.
ARPE-19 cells used for ATP release experiments were plated in 96-well white plates with clear bottoms (Corning Inc., Corning, NY, USA) and grown to confluence using the medium described above. Peripheral wells were typically left blank to avoid gradients of gas exchange across the plate. Solution change can trigger a release of ATP (Grygorczyk & Hanrahan, 1997), and our trials indicated that both 10% FBS and the phenol contained in the regular DMEM interfered with the luciferase assay. To avoid these artifacts, the growth medium bathing ARPE-19 cells used for most ATP release experiments was replaced after 4–7 days with 100 µl of a 1:1 mixture of DMEM without phenol (Invitrogen Corp.) and Ham's F12 plus 1% FBS with 3 mM L-glutamine, 100 µg ml–1 streptomycin and 2.5 mg ml–1 Fungizone and/or 50 µg ml–1 gentamicin, defined as ‘differentiation medium’. Cells were subsequently grown for an additional week. This differentiation medium did not affect the luciferase activity and could be used as working solution in on-line experiments.
Measurements of ATP from ARPE-19 cells
ATP release from ARPE-19 cells was detected using the luciferin–luciferase reaction as described above. In most cases, 80 µl of the total 100 µl differentiation medium were replaced with 20 µl differentiation medium containing a concentrated level of agonist shortly before the experiment began, to avoid full solution change. The 96-well plate was inserted into the luminometer, 10 µl of the luciferase working solution in light medium was injected and the plate gently mixed for 10 s. This gentle mix had negligible effects on ATP release and was present in both control and experimental wells. Measurements were typically taken every 20 s for 30 min with an integration time of 100 ms per measurement.
To study the initial kinetics of ATP release, 60 µl of differentiation medium were replaced with 5 µl of the luciferin–luciferase working solution, for a total volume of 45 µl. The 96-well plate was placed in the luminometer, baseline levels were recorded for 1 min, and then 5 µl concentrated NMDA was injected into each well to produce a final concentration of 300 µM. Differentiation medium without NMDA was injected into control wells. Measurements were taken every 1 s for 5 min with an integration time of 100 ms per measurement.
For experiments involving blockers, all differentiation medium was replaced with 100 µl differentiation medium containing blockers at working strength and incubated for 30 min at 37°C (120 min for brefeldin A). Shortly before the experiment, 80 µl were removed and replaced with 20 µl blocker solution plus agonist. Since the 10 µl added luciferase solution did not contain blocker, the blocker level during experiments was only 80% of that used during incubation. In the glutamate dose–response experiments, the traditional growth medium was removed from the cells and replaced by 100 µl of isotonic solution immediately before the experiment. After 1 h in the incubator, 50 µl of the isotonic solution was replaced by 50 µl of a solution containing twice the working glutamate concentration and recordings made as detailed above. While ATP release from cells using this protocol was not generally as high, the proportional increase in ATP was similar.
Immunohistochemistry
ARPE-19 cells were plated at low density on round glass coverslips for 2 days in differentiation medium. The cells were washed twice in phosphate-buffered saline (PBS) (in mM 2.6 KCl, 1.5 KH2PO4, 134 NaCl, 8 Na2HPO4, 0.9 CaCl2, 0.5 MgCl2) and fixed in 4% paraformaldehyde in PBS for 30 min at 4°C. After washing with PBS, the sample was incubated in blocking buffer composed of 10% Superblock (Pierce Biotechnology Inc., Rockford, IL, USA) diluted in PBS with 0.1% Tween 20. After being washed with PBS and 0.1% Tween 20 (washing buffer) the cells were incubated overnight at 4°C with a mouse anti-NMDAR1 monoclonal antibody against the extracellular loop of the NR1 subunit (clone 54.1, MAB363; Chemicon International, Inc., Temecula, CA, USA) diluted 1:200 in blocking buffer. The negative controls were incubated with the same solution but without the primary NMDAR1 antibody. Cells were then washed three times with washing buffer and incubated with donkey antimouse antibody conjugated with biotin (Jackson ImmunoResearch Laboratories Inc., West Grove, PA, USA) at 1:300 dilution in blocking buffer for 30 min at room temperature. Samples were washed three times in washing buffer and incubated with Cy2-conjugated Streptavidin diluted 1:300 in blocking buffer for 30 min at room temperature (Jackson ImmunoResearch). After three final washes, coverslips were mounted with a fluorescent mounting medium containing 4', 6-Diamidino-2-phenylindole (DAPI) to visualize the nuclei (H-1200, Vector Laboratories, Burlingame, CA, USA). Pictures were taken on a Nikon Eclipse 600 microscope equipped for epifluorescence (Nikon USA, Melville, NY, USA) with a 3-CCD digital camera (Toshiba America, Irvine, CA, USA) and analysed on-line using Image Pro Plus software (Media Cybernetics, Silver Spring, MD, USA). DAPI was imaged with 360 nm excitation and > 515 nm emission, while Cy2 was excited at 480 nm with emission > 535 nm.
To confirm staining, a second antibody raised against the C-terminal of the NR1 subunit was used (1:200 in blocking buffer, AB1516, polyclonal; Chemicon International, Inc.). In parallel preparations, the primary antibody was preabsorbed with the C-terminal peptide (both 1:200, AG344; Chemicon International, Inc.) at room temperature for 2 h as a control. Both preparations were added to ARPE-19 cells overnight at 4°C and processed as above using a biotinylated goat antirabbit secondary antibody (1:300; DAKO Corp., Carpinteria, CA, USA).
Intracellular calcium measurements
ARPE-19 cells were plated in 96-well black plates with clear bottoms (Corning Inc. Corning, NY, USA) and grown to confluence for 4–7 days in the growth medium described above, then maintained in differentiation medium for an additional week. After washing the wells with isotonic solution plus 250 µM glycine, cells were loaded with 10 µM fura-2 AM and 0.2% Pluronic F127 (both Invitrogen Corp.) in isotonic solution plus glycine and incubated at 37°C for 30 min. The antagonists and blockers used in the different experiments were added to the fura-2 AM loading solution. After two washes, 90 µl of isotonic solution plus glycine was added to each well. The basal intracellular Ca2+ levels were measured by alternatively exciting at 340 and 380 nm and measuring the fluorescence emitted at 510 nm in a microplate fluorometer (Fluoroskan Ascent; Labsystems, Franklin, MA, USA). After a 5 min baseline reading, 10 µl isotonic solution plus glycine with or without NMDA and/or blocker (all 10 times working strength) were injected into each well using the fluorometer injector system. Conversion to Ca2+ concentration was performed as previously described (Mitchell, 2001) with the following adaptations for the 96-well plate system. Calibration was performed simultaneously in a subset of wells on the plate, with ‘maximum’ solution composed of 5 µM ionomycin in isotonic solution, and ‘minimum’ solution made of 5 µM ionomycin and 20 mM EGTA in isotonic solution, both pH 8.0. Background levels from a subset of wells not loaded with dye were subtracted.
Data analysis
Levels of ATP were determined by integrating the area under the curve for each record. Baseline levels of ATP varied considerably with experimental day for both fresh and cultured experiments. To allow comparison of experiments, ATP concentrations were normalized to the mean control ATP level for each day. All of the data were expressed as means ± S.E.M.; n is the number of independent trials or wells, with data typically representing results from two to six separate plates. Significant differences were tested using Student's paired t test when only two conditions were present and a one-way ANOVA with Tukey's post hoc test when more than two conditions were present. Analysis was performed with SigmaStat Software (Systat Software Inc., Point Richmond, CA, USA) with P < 0.05 defined as significantly different.
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Results |
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The effect of NMDA on ATP release was initially examined using the fresh bovine RPE eyecup. In this preparation, tight junctions were generally well maintained and the apical membrane faced the interior of the eyecup, so receptors and release could be functionally localized to the apical membrane. The glutamate receptor agonist NMDA triggered a clear, consistent and concentration-dependent release of ATP from the bovine RPE eyecup. Levels of bath ATP increased from 1.8 ± 0.1 nM (n = 10) in control conditions to 6.0 ± 1.8 nM (n = 5) in 100 µM NMDA when measured 10 min after addition of the agonist. While these levels are low, this is the concentration of released ATP after it diffuses into 500 µl solution, and concentrations in the subretinal space are expected to be several orders of magnitude higher, as demonstrated by comparison with measurements using a cell-surface luciferase (Joseph et al. 2003; Okada et al. 2006). To facilitate comparison, levels were normalized to the mean control for each day's experiments. In five experiments, mean ATP levels in the eyecup were 3.4 ± 1.0-fold greater than control values after 10 min in the presence of 100 µM NMDA, with an EC50 of 32 µM (Fig. 2).
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While experiments with the bovine eyecup suggested that NMDA triggered a release of ATP across the apical membrane of fresh RPE cells, more detailed pharmacological analysis was performed using cultured human ARPE-19 cells. These cells enabled the release of ATP to be measured on-line using a high-throughput screening system while also providing information from human cells.
NMDA triggered release of ATP from ARPE-19 cells (Fig. 3A). The concentration of ATP in the bath rose for the first 5–10 min after recording began, with levels typically peaking at 
7 min before reaching a plateau or declining slightly. This time course suggested that measurements made from the fresh bovine RPE eyecup after 10 min were appropriate. In ARPE-19 cells, as in the bovine RPE eyecup, the magnitude of this increase was dependent upon the concentration of NMDA. A significant increase in bath ATP over control values was detected with only 24 µM NMDA, and levels of ATP doubled in the presence of 240 µM (EC50 of 63 µM; Fig. 3B). In these experiments, agonist was added to the cells 3–4 min before recording began, and initial recordings began with NMDA levels considerably higher than control values. To obtain a better understanding of response kinetics, we modified the protocol so that recordings were obtained at higher frequency and so that levels could be monitored shortly before and after injection of NMDA. Levels of ATP were initially the same but increased rapidly after injection of NMDA (Fig. 3C). The concentration of ATP increased by 1.7 ± 0.2-fold after 3 min in 300 µM NMDA, analogous to the increase observed at the initial points of the experiments in Fig. 3A and B.
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Additional experiments were undertaken to confirm that release of ATP was mediated by stimulation of the NMDA receptor. The NMDA receptor antagonists MK-801 (24 µM; Fig. 4A and B) and D-AP5 (80 µM; Fig. 4C and D) prevented the ability of NMDA to trigger a release of ATP from ARPE-19 cells. In both cases, NMDA alone triggered a robust response similar to that described above, with ATP levels 1.7-fold greater than control at the peak.
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The pharmacological identification of the NMDA receptor was supported by immunohistochemical identification of the NMDAR1 (NR1) subunit in ARPE-19 cells. Dense staining was detected across the cell (Fig. 6). Staining was punctate, although permeablization of the cell membrane with the paraformaldehyde used for fixation precludes identification of these punctate clusters as either intracellular or extracellular. Staining was verified with two different antibodies against NR1 and was greatly reduced by competition with the relevant peptide.
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The actions of NMDA, the ability of MK-801 and D-AP5 to completely block the response to NMDA, and the immunological identification of NMDAR1 subunit on ARPE-19 cells strongly suggested that activation of the NMDA receptor could trigger ATP release. However, the endogenous agonist is expected to be glutamate, and both ionotropic and metabotropic glutamate receptors are present on RPE cells (Lopez-Colome et al. 1993; Lopez-Colome & Fragoso, 1995). Experiments were performed to determine whether glutamate itself could trigger ATP release and what proportion of this response was mediated by the NMDA receptor. Addition of glutamate triggered release of ATP from cells with a similar magnitude to NMDA (Fig. 7A). Peak levels were observed 5–10 min after recording began, although levels did decline more than with NMDA over the course of 30 min, which is consistent with the presence of glutamate transporters. The ATP levels increased in a concentration-dependent manner, with an EC50 of 24 µM (Fig. 7B). MK-801 produced a nearly complete block of ATP release triggered by 240 µM glutamate (Fig. 7C and D), suggesting that most of the release resulted from activation of the NMDA receptors.
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The conduit for ATP release following stimulation of the NMDA receptor was probed. The channel blocker NPPB completely blocked the response (Fig. 8A). Although the gap-junction blocker 18GA can inhibit ATP release from embryonic chick RPE cells (Pearson et al. 2005), it increased ATP levels in ARPE-19 cells (Fig. 8B), while the gap-junction blocker oleamide had a non-specific effect on the luciferase reaction (Fig. 1). The cystic fibrosis transmembrane conductance regulator (CFTR) blocker glibenclamide had no effect on ATP release (Fig. 8C). Since NMDA can elevate cellular Ca2+, we asked whether Ca2+ played a role in this ATP release. Chelation of intracellular Ca2+ levels with BAPTA AM reduced the NMDA-induced ATP release by 85% (Fig. 8D). Since increased intracellular Ca2+ can trigger a vesicular release of transmitter from both neuronal and non-neuronal cells, and since brefeldin A (BFA) can impede vesicular release of ATP from other epithelial cells (Knight et al. 2002), the effect of BFA on release was examined. However, BFA had no effect on the ability of NMDA to trigger release of ATP from ARPE-19 cells (Fig. 8E).
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Exogenously added ATP can activate both ionotropic P2X and metabolic P2Y receptors and effectively raise the intracellular Ca2+ levels of RPE cells (Sullivan et al. 1997; Ryan et al. 1999; Reigada et al. 2005a). We wanted to know whether ATP released following NMDA receptor stimulation could likewise elevate Ca2+. In cells loaded with the ratiometric Ca2+-sensitive dye fura-2, addition of NMDA led to an increase in Ca2+ levels (Fig. 9A). This icrease in Ca2+ peaked 5 min after addition of NMDA, similar to the time course found for ATP release, and was prevented by the ATP chelator apyrase. This peak Ca2+ response was also blocked by the NMDAR blocker MK-801, by the P2Y1 receptor antagonist MRS2179, and by depletion of intracellular Ca2+ stores with thapsigargin (Fig. 9B). Surprisingly, a rise in Ca2+ levels shortly after application of the NMDA was not detected. Attempts to block a secondary influx of Ca2+ with the Ca2+ channel blocker nifedipine were complicated by the dose-dependent increase in ATP release triggered by nifedipine in the presence of glutamate and NMDA (both 240 µM), but not in control conditions.
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Discussion |
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Evidence for the involvement of NMDA receptors is compelling. First, NMDA itself triggered a release of ATP from both fresh bovine and cultured human RPE cells over the range 25–300 µM, with an EC50 similar to that reported for cloned NMDA receptors (Moriyoshi et al. 1991; Kurko et al. 2005). Second, this NMDA-induced ATP release from ARPE-19 cells was blocked by two different NMDA-specific inhibitors, the non-competitive antagonist MK-801 acting in the permeation pathway and the competitive antagonist D-AP5 acting at the agonist binding site (Morris et al. 1986; Davis et al. 1992). Third, block by DCKA in the presence of glycine implies a requirement for the costimulation of the glycineB binding site on the NMDA receptor (Baron et al. 1990). Fourth, the NMDAR1 subunit was detected in these cells using two different antibodies. Finally, glutamate triggered a response at a lower concentration than NMDA, and this response was largely blocked by MK-801 (Wong et al. 1986). This suggests that most of the response to glutamate was mediated by the NMDA receptor, consistent with previous reports that stimulation of the NMDA receptor was more effective at raising IP3 in the RPE than other glutaminergic receptors (Fragoso & Lopez-Colome, 1999).
The ability of NMDA to trigger ATP release from both fresh bovine and cultured human ARPE-19 cells strengthens the potential relevance of this glutaminergic–purinergic interaction. The release from the interior of the bovine RPE eyecup implies that NMDA receptors are on the apical membrane and that ATP release occurs across the apical membrane into the subretinal space. While ARPE-19 cells typically lack polarity, previous work has found many similarities in the mechanisms of ATP release from ARPE-19 and fresh bovine RPE cells (Reigada & Mitchell, 2005a; Reigada et al. 2005). The release from bovine cells was proportionally larger and required lower concentrations of NMDA, suggesting that the system in fresh cells may be more sensitive. Although we do not yet now whether this reflects a difference in receptor number, receptor sensitivity or whether the cells are more ready to release ATP, it does suggest that data from ARPE-19 cells are likely to underestimate the effect on fresh RPE cells.
Mechanism of ATP release
The release of ATP triggered by NMDA was blocked by the broad-acting channel blocker NPPB and the cell-permeable form of the Ca2+ chelator BAPTA AM. This is consistent with the involvement of ion channels and requirement for some Ca2+ in the release of ATP from RPE cells. This release was not affected by glibenclamide or BFA, suggesting that neither CFTR and/or vesicular release pathways were utilized. This contrasts with the ATP release activated by hypotonic challenge in these cells, which was blocked by glibenclamide, BFA and the inhibitor of CFTR gating CFTR-172 (Reigada & Mitchell, 2005a). The activation of distinct ATP release pathways by different stimuli has been described in other cells, including astrocytes (Joseph et al. 2003). While NPPB is widely recognized to inhibit Cl– channels (Cabantchik & Greger, 1992), it can also block conductance through hemichannels. The concentration of NPPB used here was ineffective at hemichannels expressed in neuroblastoma cells (Srinivas & Spray, 2003), although it did inhibit hemichannels expressed in oocytes (Eskandari et al. 2002), and embryonic chick RPE cells can release ATP through hemichannels (Pearson et al. 2005). In our preparation, the blocker oleamide interfered with the luciferase assay, while 18GA led to a dose-dependent increase in ATP levels. Further trials are necessary to determine whether hemichannels or more traditional anion channels account for the non-CFTR release from RPE cells.
Amplification of NMDA Ca2+ signal by ATP
NMDA led to large increases in Ca2+ that were reduced in the presence of MK-801, apyrase, thapsigargin and MRS2179. This combination is consistent with a sequence whereby activation of the NMDAR leads to ATP release, which autostimulates P2Y1 receptors, leading to release of Ca2+ from intracellular stores. Since BAPTA AM prevented the release of ATP, it is tempting to speculate that NMDA itself triggers a small rise in Ca2+ that primes ATP release, although the detection of an increase in Ca2+ shortly after presentation of NMDA would strengthen this theory. In preliminary trials performed at higher sampling frequency, small, rapid increases in Ca2+ remained in the presence of apyrase (not shown), although the presence of this priming Ca2+ remains to be confirmed.
Implications for RPE–photoreceptor interactions
The ability of NMDA receptors to trigger ATP release and autostimulation of P2 receptors may play a key role in light-dependent communication between the RPE and photoreceptors. Glutamate is the predominant neurotransmitter released by the photoreceptors, with release decreased by light (Bloomfield & Dowling, 1985; Schmitz & Witkovsky, 1996). Diffusion of glutamate to the RPE could stimulate NMDA receptors, trigger autocrine activation of P2Y1 receptors by released ATP and elevate cell Ca2+. While glutamate uptake transporters on the RPE would limit the effective spread of this glutamate (Maenpaa et al. 2002, 2003, 2004), the amplification produced by ATP release and autostimulation would enable the signal to spread. This purinergic release/autostimulation could thus amplify the response to NMDA throughout the RPE monolayer, much as it conveys Ca2+ waves throughout Müller cells in the inner retina (Reifel Saltzberg et al. 2003).
The increased levels of glutamate during the dark could lead to enhanced ATP signalling and increased intracellular Ca2+ levels. Since increased Ca2+ activates basolateral Cl– conductances of the RPE and consequently the movement of fluid from the subretinal space to the choroid (Edelman & Miller, 1991; Joseph & Miller, 1992), these glutaminergic–purinergic interactions could contribute to the relative dehydration of the subretinal space during the dark (Huang & Karwoski, 1992). This release of ATP by glutamate is at odds with the hypothesis that ATP is the light peak substance. While the physiological changes to the RPE produced by ATP are similar to those triggered by light (Gallemore et al. 1988, 1993; Gallemore & Steinberg, 1993; Peterson et al. 1997), the autocrine stimulation of RPE cells by locally released ATP is activated by multiple compounds (Mitchell, 2001; Reigada et al. 2005,) including glutamate. This suggests that ATP functions more as an extracellular second messenger to amplify the signal of many transmitters, possibly including the light peak substance. Whether the enhancement of outer segment phagocytosis by glutamate (Greenberger & Besharse, 1985) is related to the formation of adenosine following ATP release remains to be seen (Reigada et al. 2006b).
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References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Baron BM, Harrison BL, Miller FP, McDonald IA, Salituro FG, Schmidt CJ, Sorensen SM, White HS & Palfreyman MG (1990). Activity of 5,7-dichlorokynurenic acid, a potent antagonist at the N-methyl-D-aspartate receptor-associated glycine binding site. Mol Pharmacol 38, 554–561.[Abstract]
Bloomfield
SA
&
Dowling
JE (1985). Roles of aspartate and glutamate in synaptic transmission in rabbit retina. I. Outer plexiform layer. J Neurophysiol
53, 699–713.
Cabantchik ZI & Greger R (1992). Chemical probes for anion transporters of mammalian cell membranes. Am J Physiol 262, C803–C827.[Medline]
Chader GJ, Pepperberg DR, Crouch R & Wiggert B (1998). Retinoids and the retinal pigment epithelium. In The Retinal Pigment Epithelium: Function and Disease, ed. Marmor M & Wolfensberger T, pp. 135–151. Oxford University Press, New York.
Copenhagen DR (1991). Synaptic transmission in the retina. Curr Op Neurobiol 1, 258–262.[CrossRef][Medline]
Danysz W, Parsons CG, Karcz-Kubicha M, Schwaier A, Popik P, Wedzony K, Lazarewicz J & Quack G (1998). GlycineB antagonists as potential therapeutic agents. Previous hopes and present reality. Amino Acids 14, 235–239.[CrossRef][Medline]
Davis S, Butcher SP & Morris RG (1992). The NMDA receptor antagonist D-2-amino-5-phosphonopentanoate (D-AP5) impairs spatial learning and LTP in vivo at intracerebral concentrations comparable to those that block LTP in vitro. J Neurosci 12, 21–34.[Abstract]
Dunn
K, Marmorstein
A, Bonilha
V, Rodriguez-Boulan
E, Giordano
F
&
Hjelmeland
L (1998). Use of the ARPE-19 cell line as a model of RPE polarity: basolateral secretion of FGF5. Invest Ophthalmol Vis Sci
39, 2744–2749.
Edelman
JL
&
Miller
SS (1991). Epinephrine stimulates fluid absorption across bovine retinal pigment epithelium. Invest Ophthalmol Vis Sci
32, 3033–3040.
Eskandari S, Zampighi GA, Leung DW, Wright EM & Loo DD (2002). Inhibition of gap junction hemichannels by chloride channel blockers. J Membr Biol 185, 93–102.[CrossRef][Medline]
Fragoso G & Lopez-Colome AM (1999). Excitatory amino acid-induced inositol phosphate formation in cultured retinal pigment epithelium. Vis Neurosci 16, 263–269.[CrossRef][Medline]
Gallemore
RP, Griff
ER
&
Steinberg
RH (1988). Evidence in support of a photoreceptoral origin for the ‘light-peak substance’. Invest Ophthalmol Vis Sci
29, 566–571.
Gallemore
RP, Hernandez
E, Tayyanipour
R, Fujii
S
&
Steinberg
RH (1993). Basolateral membrane Cl– and K+ conductances of the dark-adapted chick retinal pigment epithelium. J Neurophysiol
70, 1656–1668.
Gallemore
RP
&
Steinberg
RH (1990). Effects of dopamine on the chick retinal pigment epithelium. Membrane potentials and light-evoked responses. Invest Ophthalmol Vis Sci
31, 67–80.
Gallemore
RP
&
Steinberg
RH (1993). Light-evoked modulation of basolateral membrane Cl– conductance in chick retinal pigment epithelium: the light peak and fast oscillation. J Neurophysiol
70, 1669–1680.
Greenberger LM & Besharse JC (1985). Stimulation of photoreceptor disc shedding and pigment epithelial phagocytosis by glutamate, aspartate, and other amino acids. J Comp Neurol 239, 361–372.[CrossRef][Medline]
Grygorczyk R & Hanrahan JW (1997). CFTR-independent ATP release from epithelial cells triggered by mechanical stimuli. Am J Physiol 272, C1058–C1066.[Medline]
Huang B & Karwoski CJ (1992). Light-evoked expansion of subretinal space volume in the retina of the frog. J Neurosci 12, 4243–4252.[Abstract]
Hughes
BA, Miller
SS
&
Machen
TE (1984). 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.
Johnson JW & Ascher P (1987). Glycine potentiates the NMDA response in cultured mouse brain neurons. Nature 325, 529–531.[CrossRef][Medline]
Joseph
SM, Buchakjian
MR
&
Dubyak
GR (2003). Colocalization of ATP release sites and ecto-ATPase activity at the extracellular surface of human astrocytes. J Biol Chem
278, 23331–23342.
Joseph
DP
&
Miller
SS (1992). Alpha-1-adrenergic modulation of K and Cl transport in bovine retinal pigment epithelium. J Gen Physiol
99, 263–290.
Knight GE, Bodin P, De Groat WC & Burnstock G (2002). ATP is released from guinea pig ureter epithelium on distension. Am J Physiol 282, F281–F288.
Kurko D, Dezso P, Boros A, Kolok S, Fodor L, Nagy J & Szombathelyi Z (2005). Inducible expression and pharmacological characterization of recombinant rat NR1a/NR2A NMDA receptors. Neurochem Int 46, 369–379.[CrossRef][Medline]
Lopez-Colome AM & Fragoso G (1995). Glycine stimulation of glutamate binding to chick retinal pigment epithelium. Neurochem Res 20, 887–894.[CrossRef][Medline]
Lopez-Colome AM, Salceda R & Fragoso G (1993). Specific interaction of glutamate with membranes from cultured retinal pigment epithelium. J Neurosci Res 34, 454–461.[CrossRef][Medline]
Maenpaa H, Gegelashvili G & Tahti H (2004). Expression of glutamate transporter subtypes in cultured retinal pigment epithelial and retinoblastoma cells. Curr Eye Res 28, 159–165.[CrossRef][Medline]
Maenpaa H, Mannerstrom M, Toimela T, Salminen L, Saransaari P & Tahti H (2002). Glutamate uptake is inhibited by tamoxifen and toremifene in cultured retinal pigment epithelial cells. Pharmacol Toxicol 91, 116–122.[CrossRef][Medline]
Maenpaa H, Saransaari P & Tahti H (2003). Kinetics of inhibition of glutamate uptake by antioestrogens. Pharmacol Toxicol 93, 174–179.[CrossRef][Medline]
Mazzanti M, Sul JY & Haydon PG (2001). Glutamate on demand: astrocytes as a ready source. Neuroscientist 7, 5.
Miller
SS, Hughes
BA
&
Machen
TE (1982). Fluid transport across retinal pigment epithelium is inhibited by cyclic AMP. Proc Nat Acad Sci U S A
79, 2111–2115.
Miller SS & Steinberg RH (1977a). Active transport of ions across frog retinal pigment epithelium. Exp Eye Res 25, 235–248.[CrossRef][Medline]
Miller SS & Steinberg RH (1977b). Passive ionic properties of frog retinal pigment epithelium. J Membr Biol 36, 337–372.[CrossRef][Medline]
Mitchell CH (2001). Release of ATP by a human retinal pigment epithelial cell line: potential for autocrine stimulation through subretinal space. J Physiol 534, 93–202.
Mitchell
CH, Carre
DA, McGlinn
AM, Stone
RA
&
Civan
MM (1998). A release mechanism for stored ATP in ocular ciliary epithelial cells. Proc Nat Acad Sci U S A
95, 7174–7178.
Moriyoshi K, Masu M, Ishii T, Shigemoto R, Mizuno N & Nakanishi S (1991). Molecular cloning and characterization of the rat NMDA receptor. Nature 354, 31–37.[CrossRef][Medline]
Morris RG, Anderson E, Lynch GS & Baudry M (1986). Selective impairment of learning and blockade of long-term potentiation by an N-methyl-D-aspartate receptor antagonist, AP5. Nature 319, 774–776.[CrossRef][Medline]
Nash MS, Flanigan T, Leslie R & Osborne NN (1999). Serotonin-2A receptor mRNA expression in rat retinal pigment epithelial cells. Ophthal Res 31, 1–4.
Nash
MS
&
Osborne
NN (1997). Pharmacologic evidence for 5-HT1A receptors associated with human retinal pigment epithelial cells in culture. Invest Ophthalmol Vis Sci
38, 510–519.
Okada
SF, Nichols
RA, Kreda
SM, Lazarowski
ER
&
Boucher
RC (2006). Physiological regulation of ATP release at the apical surface of human airway epithelia. J Biol Chem
doi: 10.1074/jbc.M603019200.
Pearson RA, Dale N, Llaudet E & Mobbs P (2005). ATP released via gap junction hemichannels from the pigment epithelium regulates neural retinal progenitor proliferation. Neuron 46, 731–744.[CrossRef][Medline]
Peterson
WM, Meggyesy
C, Yu
K
&
Miller
SS (1997). Extracellular ATP activates calcium signaling, ion, and fluid transport in retinal pigment epithelium. J Neurosci
17, 2324–2337.
Quinn
RH, Quong
JN
&
Miller
SS (2001). Adrenergic receptor activated ion transport in human fetal retinal pigment epithelium. Invest Ophthalmol Vis Sci
42, 255–264.
Reifel Saltzberg JM, Garvey KA & Keirstead SA (2003). Pharmacological characterization of P2Y receptor subtypes on isolated tiger salamander Müller cells. Glia 42, 149–159.[CrossRef][Medline]
Reigada
D, Lu
W
&
Mitchell
CH (2006a). Glutamate acts at NMDA receptors to trigger ATP release from the RPE. Inves Ophthalmol Vis Sci
47, 883.
Reigada D, Lu W, Zhang X, Friedman C, Pendrak K, McGlinn A, Stone RA, Laties AM & Mitchell CH (2005). Degradation of extracellular ATP by the retinal pigment epithelium. Am J Physiol 289, C617–C624.[CrossRef]
Reigada
D
&
Mitchell
C (2005a). Release of ATP from retinal pigment epithelial cells involves both CFTR and vesicular transport. Am J Physiol Cell Physiol
288, C132–C140.
Reigada D & Mitchell CH (2005b). Stimulation of ATP release from the retinal pigment epithelium by glutamate. FASEB J 19, A1206.
Reigada D, Zhang X, Crespo A, Nguyen J, Liu J, Pendrak K, Stone RA, Laties AM & Mitchell CH (2006b). Stimulation of an a1-adrenergic receptor downregulates ecto-5' nucleotidase on the apical membrane of RPE cells. Purinergic Signal in press.
Ryan
JS, Baldridge
WH
&
Kelly
ME (1999). Purinergic regulation of cation conductances and intracellular Ca2+ in cultured rat retinal pigment epithelial cells. J Physiol
520, 745–759.
Sauer H, Hescheler J & Wartenberg M (2000). Mechanical strain-induced Ca2+ waves are propagated via ATP release and purinergic receptor activation. Am J Physiol 279, C295–C307.
Schmitz Y & Witkovsky P (1996). Glutamate release by the intact light-responsive photoreceptor layer of the Xenopus retina. J Neurosci Meth 68, 55–60.[CrossRef][Medline]
Schwiebert
EM (2000). Extracellular ATP-mediated propagation of Ca2+ waves. Focus on "mechanical strain-induced Ca2+ waves are propagated via ATP release and purinergic receptor activation". Am J Physiol Cell Physiol
279, C281–C283.
Srinivas
M
&
Spray
DC (2003). Closure of gap junction channels by arylaminobenzoates. Mol Pharmacol
63, 1389–1397.
Sullivan DM, Erb L, Anglade E, Weisman GA, Turner JT & Csaky KG (1997). Identification and characterization of P2Y2 nucleotide receptors in human retinal pigment epithelial cells. J Neurosci Res 49, 43–52.[CrossRef][Medline]
Uchida N, Kiuchi Y, Miyamoto K, Uchida J, Tobe T, Tomita M, Shioda S, Nakai Y, Koide R & Oguchi K (1998). Glutamate-stimulated proliferation of rat retinal pigment epithelial cells. Eur J Pharmacol 343, 265–273.[CrossRef][Medline]
Versaux-Botteri C, Gibert JM, Nguyen-Legros J & Vernier P (1997). Molecular identification of a dopamine D1b receptor in bovine retinal pigment epithelium. Neurosci Letts 237, 9–12.[CrossRef][Medline]
Witkovsky
P, Schmitz
Y, Akopian
A, Krizaj
D
&
Tranchina
D (1997). Gain of rod to horizontal cell synaptic transfer: relation to glutamate release and a dihydropyridine-sensitive calcium current. J Neurosci
17, 7297–7306.
Wong
EH, Kemp
JA, Priestley
T, Knight
AR, Woodruff
GN
&
Iversen
LL (1986). The anticonvulsant MK-801 is a potent N-methyl-D-aspartate antagonist. Proc Nat Acad Sci U S A
83, 7104–7108.
Young
RW
&
Bok
D (1969). Participation of the retinal pigment epithelium in the rod outer segment renewal process. J Cell Biol
42, 392–403.
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) versus control cells (
). Recording began 3–5 min after addition of drugs to cells. Symbols represent the means ±
S.E.M. (n
= 49). B, increasing concentrations of NMDA produced increasing levels of ATP bathing ARPE-19 cells. Levels of ATP were determined from the area under the curve for the 30 min duration of experiments like those illustrated in A and normalized to the mean control level for each day. The symbols represent the means ±
S.E.M.
*P < 0.05 versus control value. The line is a fit of the single exponential rise y
=
y0
+
a(1 – e–bx) with y0
= 1.02, b
= 0.009 and a
= 1.24. C, ATP was rapidly detected by injecting either control medium (
) blocked NMDA-stimulated ATP release. Cells were pre-incubated with 30 µM MK-801 for 30 min before recording began. Symbols represent the means ±
S.E.M., n
= 16. B, quantification of the total amount of ATP in the bath over the 30 min record indicated that MK-801 (24 µM, grey bar) significantly inhibited the NMDA-stimulated ATP release (240 µM NMDA, filled bar, NMDA alone). *P < 0.05 versus control (open bar); **P < 0.05 versus NMDA alone; MK-801 + NMDA not significantly different from control value. Bars indicate means +
S.E.M.C, mean traces from representative experiments showing the effect of a second NMDA receptor antagonist, D-AP5 (80 µM, 
