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1 Department of Ophthalmology, University of Nebraska Medical Center, Omaha, Nebraska 68198 2 Department of Pharmacology, University of Nebraska Medical Center, Omaha, Nebraska 68198 3 Department of General Zoology and Neurobiology, University of Pécs, H-7601 Pécs, Hungary
Submitted 17 October 2002; accepted in final form 22 April 2003
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ABSTRACT |
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12 mM [Cl–]o with CH3SO4– produced a significant reduction in [Cl–]i. [Ca2+]i increases evoked by 20 or 50 mM K+ were also significantly inhibited by replacing 12 mM [Cl–]o with CH3SO4–. Thus modest depolarization can evoke increases in [Ca2+]i that lead to reductions in [Cl–]i, and conversely, reductions in [Cl–]i inhibit depolarization-evoked [Ca2+]i increases. These findings support the hypothesis that feedback interactions between Ca2+- and Ca2+-activated Cl– channels may contribute to the regulation of presynaptic Ca2+ currents involved in synaptic transmission from rod photoreceptors. |
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INTRODUCTION |
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; Stella et al. 2002) that mediate Ca2+ influx and thereby regulate glutamate release from photoreceptor terminals. Ca2+ influx through these channels can also activate a very large Ca2+-activated Cl– current (ICl(Ca)) (Bader et al. 1982). In cones, the current flux through ICl(Ca) is at least eightfold greater than the current through ICa (Barnes and Hille 1989). The chloride equilibrium potential (ECl) of salamander rods is about –20 mV (Thoreson et al. 2002). In olfactory receptors, ECl is also positive to the cell's resting potential and acts to boost the receptor potential (Kleene and Gesteland 1991). In rods, depolarizing responses to darkness would also presumably be boosted by activation of ICl(Ca). In addition, activation of ICl(Ca) at the dark resting potential (around–45 mV) generates a Cl– efflux (Thoreson et al. 2002). The resulting reduction in [Cl–]i has the unusual effect of inhibiting the open channel probability of single Ca2+ channels in photoreceptor terminals (Thoreson et al. 2000). This may promote a negative feedback interaction whereby activation of ICa leads to a Ca2+ influx that activates a Cl– efflux, which in turn feeds back to inhibit ICa (Thoreson et al. 2002). When ICa is enhanced by quinpirole, this negative feedback interaction may help to account for the paradoxical finding that, although activation of D2/D4 dopamine receptors enhances ICa, quinpirole nonetheless inhibits synaptic transmission from rods (Thoreson et al. 2002; Witkovsky et al. 1989). Is this feedback interaction restricted to conditions where ICa has been enhanced (e.g., with quinpirole) or does it regulate ICa under normal operating conditions at the rod synapse? To address this question, we combined electrophysiology with Ca2+ and Cl– imaging techniques to assess the reciprocal interactions between Ca2+ influx and Cl– efflux under physiological conditions. The results show an intimate relationship between the two that support the hypothesis of a feedback interaction between ICa and ICl(Ca) operating near the dark potential and defines mechanisms that contribute to maintenance of a positive value for ECl in rod photoreceptors.
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METHODS |
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Larval tiger salamanders (Ambystoma tigrinum, 18–25 cm) were cared for according to institutional guidelines. Retinal slices were prepared according to methods pioneered by Werblin (Werblin 1978) and Wu (Wu 1987). Salamanders were pithed and decapitated, an eye was enucleated, and the front of the eye was removed. The resulting eyecup was cut into three or four pieces, and a single piece was placed vitreal surface down onto a piece of filter paper (Millipore 2 x 5 mm, Type GS, 0.2-µm pores). After the retina adhered to the filter paper, the retina was isolated under chilled amphibian superfusate and cut into 125-µm slices using a razor blade tissue chopper (Stoelting, Wood Dale, IL). The slices were rotated 90° to view the retinal layers when placed under a water immersion objective (60x, 1.0 NA) on an upright fixed stage microscope (EF 600, Nikon). For electrophysiological experiments, all procedures were performed under infrared illumination. For imaging experiments, slices were prepared under a dissecting lamp in visible light.
Solutions and perfusion
Solutions were applied with a single-pass, gravity-feed perfusion system (1 ml/min). The normal amphibian superfusate contained (in mM) 111 NaCl, 2.5 KCl, 1.8 CaCl2, 0.5 MgCl2, 10 N-2-hydroxyethylpiperazine-N' 2-ethanesulfonic acid (HEPES), and 5 glucose (pH 7.8). In some experiments, CaCl2 was replaced with 2 mM BaCl2. For Cl– replacement experiments, Cl– was replaced with equimolar CH3SO4–. CH3SO4– solutions were boiled for 10 min in a fume hood to evaporate any residual methanol before glucose was added and the solution brought to its final volume.
Electrophysiology
Patch pipettes were pulled on a PB-7 vertical puller (Narishige) from borosilicate glass pipettes (1.2 mm OD, 0.95 mm ID, omega dot) and had tips of approximately 1 µm OD with resistances of 10–15 M
. For measurements of membrane potentials in rods, we used gramicidin perforated-patch recording techniques (Kyrozis and Reichling 1995). Gramicidin was dissolved in ethanol (5 mg/ml) and then added to the pipette electrolyte solution to achieve a final concentration of 5 µg/ml. For current-clamp measurements of membrane potential, the pipette electrolyte solution contained (in mM) 54 KCl, 61.5 KCH3SO4 (Pfaltz and Bauer, Waterbury, CT), 3.5 NaCH3SO4, and 10 HEPES (pH 7.2). For voltage-clamp recordings of IBa, ICa, and ICl(Ca), the pipette solution contained (in mM) 54 CsCl, 61.5 CsCH3SO4, 3.5 NaCH3SO4, and 10 HEPES (pH 7.2). The osmolarity was adjusted, if necessary, to 242 ± 5 mOsm. Rods were recorded using an Axopatch 200B amplifier (Axon Instruments, Union City, CA).
Imaging experiments
Digital fluorescent images were obtained with a cooled CCD camera (SensiCam, Cooke, Auburn Hills, MI). Axon Imaging Workbench (AIW 2.2, Axon Instruments) was used to control the camera, filter wheel, and image acquisition. Pixel binning (2 x 2) of the images was typically used to decrease acquisition time to 
1 s. Images were acquired at 5- to 10-s intervals during experimental trials.
For measurements of [Ca2+]i, we used the ratiometric dye, Fura-2 (Molecular Probes, Eugene, OR) (Grynkiewicz et al. 1985). Retinal slices were loaded with Fura-2 by incubating them at 5°C for 45 min in the dark with 0.5 ml of 10 µM Fura-2/AM + 0.02% pluronic F-127 (Molecular Probes). This was followed by an additional incubation of 1.5 h in Fura-2/AM without pluronic F-127. Intracellular [Ca2+] increases were stimulated by depolarization with elevated KCl applied for 1 min at 15-min intervals. Measurements were made in the cell soma. For statistical comparisons, the change in the 340/380 nm ratio produced by application of KCl in the presence of a test substance was compared with the average of the 340/380 ratio changes obtained prior to application of the test substance and following washout.
For measurements of [Cl–]i we used the dye, 6-methoxy-N-ethylquinolinium iodide (MEQ, Molecular Probes) (Biwersi and Verkman 1991). MEQ was loaded into cells after reducing it to DiH-MEQ by adding 30 µM sodium borohydride (100 µl) to MEQ (5 mg) under a continuous stream of nitrogen gas (Woll et al. 1996). DiHMEQ enters cells during the incubation period (15 min), where it is oxidized and retained in the form of MEQ. Fluorescence emission decreases as Cl– quenches MEQ. As illustrated in Fig. 1, the exponential decay in MEQ fluorescence due to dye leakage and bleaching was determined under control conditions and subtracted prior to analysis. NPPB was used at a concentration of 2 µM in Cl– imaging experiments; concentrations of 10 µM and above generated a detectable intrinsic fluorescence with the MEQ fluorescence cube.
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Statistical comparisons were made using Student's t-test with a significant value of 0.05. Variance is reported as ± SE. Unless otherwise specified, chemicals were obtained from Sigma/Aldrich/RBI (St. Louis, MO).
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RESULTS |
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F/F). This normalizing procedure helps to factor out changes in fluorescence arising from variations in dye concentration or cell thickness (Helmchen 2000). As shown in Fig. 1D, reducing extracellular Cl– by 12 mM produced detectable increases in MEQ fluorescence and larger reductions in extracellular Cl– produced correspondingly larger increases in MEQ fluorescence. For Fura-2 studies of intracellular [Ca2+], we stimulated Ca2+ influx by depolarizing the rods with increases in extracellular [K+]. We used gramicidin-perforated patch recording techniques to record the membrane potentials produced by the different high K+ solutions (Fig. 2A). The resulting relationship could be fit with a modified Goldman-Hodgkin-Katz equation with relative permeability of PNa/PK of 0.13 (line, Fig. 2A). The small deviation from this relationship at more positive potentials may be due to an increasing contribution of Ca2+ and Ca2+-activated Cl– channels to the membrane potential.
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After calibrating the solutions in this way, we then used the same high K+ solutions to stimulate increases in intracellular [Ca2+]. Fluorescence measurements were made from the somas of rods in the retinal slice. Increasing the concentration of K+ produced a sigmoidal increase in intracellular [Ca2+] that was half-maximal at –21.8 mV and reached its peak above –12 mV (Fig. 2B). For comparison, we averaged ICa recorded from 12 rods using a ramp voltage protocol (0.5 mV/ms) and gramicidin-perforated-patch recording techniques (Fig. 2C, noisy trace). The half-maximal voltage for the sigmoidal best fit function to the ICa/voltage relationship was –36.2 mV. Thus a small but significant fraction of ICa was active at the resting potential of–44 mV. Differences between the voltage dependence of ICa and depolarization-evoked increases in [Ca2+]i are considered in the DISCUSSION.
The Cl– equilibrium potential (ECl) of rods was determined by depolarizing cells with identical high K+ solutions to those used in Fig. 2. As shown in Fig. 1, Cl– efflux causes an increase in MEQ fluorescence. When cells were depolarized with high K+ solutions to levels below –20 mV, MEQ fluorescence increased indicating a Cl– efflux and when cells were depolarized above –20 mV, MEQ fluorescence decreased, indicating a Cl– influx (Fig. 3A, filled circles, data from Thoreson et al. 2002); these results suggest that ECl is around –20 mV. This finding was substantiated by electrophysiological experiments that also indicate that ECl in rods is around –20 mV (Thoreson et al. 2002). The small Cl– efflux evoked by weak depolarization to –35 mV (Fig. 3A) is likely due to additional activation of Ca2+-activated Cl– channels by the additional Ca2+ influx accompanying further stimulation of voltage-gated Ca2+ channels (Fig. 2).
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The fact that ECl is positive to the resting membrane potential of around–45 mV in these cells suggests that Cl– ions are actively accumulated by rods. The most common mechanism by which cells accumulate Cl– is through the Na/K/Cl cotransporter that mediates the coupled influx of Na+, K+, and Cl– ions (Russell 2000). The Na/K/Cl co-transporter is selectively inhibited by the loop diuretic bumetanide (Russell 2000). We therefore used bumetanide to test for a role for the Na/K/Cl cotransporter. First, the membrane potentials of rods established by solutions containing different concentrations of K+ (n = 6) in the presence of bumetanide (0.1 mM) were measured in current-clamp mode, similar to the experiment illustrated in Fig. 2A. Then, changes in intracellular Cl– evoked by the same high K+ solutions plus bumetanide (0.1 mM) were evaluated using MEQ. Supporting a role for the Na/K/Cl co-transporter in the accumulation of Cl– by rods, ECl determined from the depolarization-evoked Cl– flux was shifted about 10 mV more negative by application of bumetanide (Fig. 3A, open circles).
The Cl– influx evoked by depolarization to –9 mV with 50 mM K+ in control solution (no bumetanide) was strongly inhibited by the Cl– channel blockers 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) (2 µM) and niflumic acid (0.1 mM) (Fig. 3B). We also tested the ability of these Cl– channel blockers to inhibit the tail current following a depolarizing step or ramp. This tail current is largely attributable to activation of ICl(Ca) (Barnes and Deschenes 1992). Niflumic acid (0.1 mM) inhibited Itail by approximately 50% but NPPB (10 µM) did not significantly inhibit Itail (Fig. 3C). Because the inhibition of depolarization-evoked Cl-efflux by NPPB cannot be accounted for by block of ICl(Ca), this suggests it may block another type of Cl– channel in rods. Replacing Ca2+ with Ba2+ strongly inhibits activation of ICl(Ca) (data not shown) and likewise inhibited the Cl– influx stimulated by high K+ (Fig. 3B). Another Cl– channel blocker, N-phenylanthracilic acid (0.1 mM), had no significant effect on Cl– flux or the tail current (data not shown).
In cones, niflumic acid is a more potent and selective blocker of ICl(Ca) than IAA-94 or flufenamic acid (Barnes and Deschenes 1992) and niflumic acid was the only effective inhibitor of the three Cl– channel blockers tested in this study. We tested the effects of niflumic acid on rod Ca2+ currents using Ba2+ as the charge carrier to minimize the influence of ICl(Ca) since it is not effectively activated by Ba2+. We found that in rods, niflumic acid inhibited IBa by about 20% (Fig. 3C). Thus at least part of niflumic acid's ability to block ICl(Ca) resides in its ability to block Ca2+ influx. NPPB did not significantly inhibit ICa at 10 µM (Fig. 3C) but substantially inhibited ICa at 0.1 mM (n = 2, data not shown).
Because of its Ca2+ channel blocking effects, niflumic acid could not be used as a tool to open the hypothesized feedback loop between ICl(Ca) and ICa. Instead, as a further test for a possible feedback relationship, we determined whether reducing extracellular Cl– by 12 mM also significantly reduced the [Ca2+]i increase evoked by depolarizing rods to –9 mV with 50 mM K+ (Fig. 4A). Reducing extracellular Cl– further by replacing it with 24 mM CH3SO4 produced a stronger inhibition of [Ca2+] increases; reducing extracellular Cl– still further produced little additional decrease. This concentration dependence is almost identical to the concentration dependence of methylsulfate in Cl– replacement effects on photoreceptor ICa and light-evoked currents in second-order neurons (Thoreson et al. 1997).
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We also tested the effects of lowering extracellular Cl– in rods that had been depolarized to –26 mV by applying 22 mM K+. Application of 22 mM K+ evoked an initial transient Ca2+ increase followed by a sustained elevation of Ca2+. During the sustained elevation, reducing [Cl–]o by 12 mM or more reduced Ca2+ levels close to the baseline concentration existing prior to application of 22 mM K+ (Fig. 4B).
Ca2+-activated Cl– channels are concentrated in rod terminals (Macleish and Nurse 2000). However, effects of low Cl– solutions on the Ca2+ increases evoked by 22 mM K+ were not significantly more pronounced in terminals of enzymatically isolated rods compared with measurements in the soma (12 mM CH3SO4, P = 0.87, paired t-test, n = 6; 24 mM CH3SO4, P = 0.29, n = 6).
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DISCUSSION |
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; Thoreson et al. 1997). Depolarizing rods by only 10 mV from their resting potential of–44 mV evoked detectable Ca2+ increases (Fig. 2) that were sufficient to stimulate detectable Cl– efflux (Fig. 3A; Thoreson et al. 2002). Taken together, the observations that a similar small amplitude Cl– efflux is stimulated by both weak depolarization and application of low Cl– solutions and that the latter inhibits depolarization-evoked increases in [Ca2+]i support the hypothesis of a negative feedback interaction between influx through ICa and Ca2+ -activated Cl– efflux.
The finding that weak depolarizations (e.g., to –35 mV) stimulating small increases in spatially averaged [Ca2+] (Fig. 2) evoked detectable Cl– efflux (Fig. 3A) can be interpreted as suggesting that relatively small increases in [Ca2+]i may be sufficient to activate ICl(Ca). However, another possibility is that Ca2+-activated Cl– channels are anchored close to Ca2+ channels so that influx through these channels produces a high local concentration of Ca2+ around the Ca2+-activated Cl– channels despite a relatively small elevation of global Ca2+. Consistent with this possibility, the Ca2+-activated Cl– channels in salamander olfactory neurons do not attain half-maximal activation until Ca2+ levels reach 5 µM (Kleene and Gesteland 1991).
The results of this study suggest at least two mechanisms are important in regulating Cl– levels in rods. First, the effects of bumetanide (Fig. 3A) suggest that the Na/K/Cl co-transporter helps drive the accumulation of Cl– ions. Second, the finding that experimental manipulations that inhibit ICl(Ca), such as application of niflumic acid or replacing Ca2+ with Ba2+, also inhibited depolarization-evoked Cl– influx suggests a major role for ICl(Ca) in mediating this flux. However, results of these experiments suggest that other Cl– channels may also contribute to the influx evoked by strong depolarization. Replacing Ca2+ with Ba2+ more strongly inhibits ICl(Ca) than application of niflumic acid (0.1 mM). However, this manipulation was less effective at inhibiting Cl– influx than either niflumic acid or NPPB (Fig. 3B). Furthermore, NPPB (2 µM) was equipotent to niflumic acid (0.1 mM) in blocking Cl– influx but even 10 µM NPPB was ineffective in blocking ICl(Ca) (Fig. 3). These pharmacological differences are probably not due to actions of niflumic acid on Ih (Satoh and Yamada 2001) since it is minimally active at the dark potential and above. In addition to ICl(Ca), photoreceptors exhibit Cl– channels coupled to glutamate transporter activity (Eliasof and Werblin 1993; Larsson et al. 1996; Picaud et al. 1995) and may also possess ClC channels since a ClC-2 antibody labels the ONL and OPL and a ClC-3 antibody labels the OPL (Enz et al. 1999; Stobrawa et al. 2001).
High K+ solutions have been used by a number of investigators for the purpose of depolarizing salamander photoreceptors for Ca2+ imaging studies (e.g., Baldridge et al. 1998; Krizaj and Copenhagen 1998; Thoreson et al. 1997). In this study, we determined the membrane potentials produced by various high K+ solutions. The voltage dependence of high K+-evoked increases in [Ca2+]i was compared with the voltage dependence of ICa from rods. The half-maximal voltage for the sigmoidal best fit function to the [Ca2+]i/voltage relationship was –21.8 mV, whereas the half-maximal voltage for ICa for rods from the salamander retinal slice was found to be –36.2 mV. Thus a significant fraction of ICa was active at the resting potential of–44 mV in rods used for imaging experiments. Sustained activation of ICa at a membrane potential of–40 mV inhibits ICa by more than 50% as the combined result of Ca2+-dependent inactivation and depletion of synaptic cleft Ca2+ ions (Rabl and Thoreson 2002). The tonic inhibition of ICa that thus accompanies the modest activation of ICa at the membrane potential of–44 mV may help explain the relatively small increase in [Ca2+]i evoked by depolarization to –35 mV. Increases in [Ca2+]i also showed a steeper voltage dependence than ICa. This steeper voltage dependence may arise from the fact that rods used for imaging experiments are not voltage clamped and can therefore produce regenerative Ca2+ action potentials (Burkhardt et al. 1991; Fain et al. 1977). In addition, calcium-induced calcium release (Krizaj et al. 1999, 2003) might further steepen the voltage dependence of [Ca2+]i by boosting the amplitude of depolarization-evoked [Ca2+]i increases.
Corey et al. (1984) reported a V50 for ICa in rods of–22 mV. The more negative activation found in the present study can be accounted for by differences in the superfusate used in the two studies. Corey et al. (1984) used extracellular solutions containing 6 mM Ca2+ and pH 7.2–7.3, whereas we used solutions containing 1.8 mM Ca2+ and pH 7.8. Due to surface potential effects, the higher [Ca2+]o would produce an activation shift of +7 mV (Baldridge et al. 1998) and the lower pH would add another 6- to 7-mV positive shift (Barnes et al. 1993).
The present results suggest that even relatively weak depolarization near the dark potential can produce a sufficiently large Cl– efflux to cause inhibition of ICa, Ca2+ influx, and synaptic output from rods (Thoreson et al. 1997). This suggests that the feedback relationship between ICa and ICl(Ca) postulated by Thoreson et al. (2002) is not likely to be limited to conditions where ICa has been accentuated (e.g., following suppression of PKA activity). What is the predicted effect of this feedback mechanism on light responses? Hyperpolarization of a rod by bright light would lead to the closure of Ca2+ channels and the resulting reduction in Ca2+ influx would in turn close Ca2+-activated Cl– channels. As Cl– efflux diminished, anion-mediated inhibition of ICa would diminish. Thus more Ca2+ channels would be available to be activated by the depolarization that accompanies light offset thereby accelerating the rate of depolarization as well as boosting synaptic output as a consequence of the increased Ca2+ influx. Unfortunately, none of the Cl– channel blockers so far investigated are sufficiently selective to allow direct test of this prediction.
Rods produce both spike-like and prolonged regenerative Ca2+ action potentials (Burkhardt et al. 1988; Fain et al. 1977). Generation of a prolonged depolarization produces a substantial shunt reduction of photoreceptor light responses (Burkhardt et al. 1988), and the Ca2+ influx accompanying such a sustained depolarization can be damaging to neurons. A number of mechanisms help to keep ICa activity limited to a tonic low level and prevent more frequent activation of Ca2+ action potentials in photoreceptors. These include 1) activation of large outward currents when the cell membrane potential exceeds the dark resting potential (Barnes and Hille 1989; Owen 1987); 2) Ca2+-dependent inactivation of ICa (Rabl and Thoreson 2002); and 3) depletion of Ca2+ from the synaptic cleft during sustained activation of ICa (Rabl and Thoreson 2002). Feedback between ICa and ICl(Ca) may be another mechanism to help assure that ICa activation is maintained at a low level.
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DISCLOSURES |
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FOOTNOTES |
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Address for reprint requests: W. B. Thoreson, Department of Ophthalmology, University of Nebraska Medical Center, 985540 Nebraska Medical Center, Omaha, NE 68198-5540 (E-mail: wbthores{at}unmc.edu).
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INTRODUCTION
