Ca²⁺-Activated Cl⁻ Current in Retinal Arteriolar Smooth Muscle

Abstract

purpose. To characterize the biophysical, pharmacologic, and functional properties of the Ca²⁺-activated Cl⁻ current in retinal arteriolar myocytes.

methods. Whole-cell perforated patch-clamp recordings were made from myocytes within intact isolated arteriolar segments. Arteriolar tone was assessed using pressure myography.

results. Depolarizing of voltage steps to −40 mV and greater activated an L-type Ca²⁺ current (I_Ca(L)) that was followed by a sustained current. Large tail currents (I_tail) were observed on stepping back to −80 mV. The sustained current and I_tail reversed close to 0 mV in symmetrical Cl⁻ concentrations. The ion selectivity sequence for I_tail was I⁻> Cl⁻> glucuronate. Outward I_tail was sensitive to the Cl⁻ channel blockers 9-anthracene-carboxylic acid (9-AC; 1 mM), 4-acetamido-4′-isothiocyanatostilbene-2,2′-disulfonic acid (SITS; 1 mM), and disodium 4,4′-diisothiocyanatostilbene-2,2′-disulfonate (DIDS; 1 mM), but only DIDS produced a substantial (78%) block of inward tail currents at −100 mV. I_tail was decreased in magnitude when the normal bathing medium was substituted with Ca²⁺-free solution or if I_Ca(L) was inhibited by 1 μM nimodipine. Caffeine (10 mM) produced large transient currents that reversed close to the Cl⁻ equilibrium potential and were blocked by 1 mM DIDS or 100 μM tetracaine. DIDS had no effect on basal vascular tone in pressurized arterioles but dramatically reduced the level of vasoconstriction observed in the presence of 10 nM endothelin-1.

conclusions. Retinal arteriolar myocytes have I_Cl(Ca), which may be activated by Ca²⁺ entry through L-type Ca²⁺ channels or Ca²⁺ release from intracellular stores. This current appears to contribute to agonist-induced retinal vasoconstriction.

The regulatory control of retinal blood flow plays a crucial role in maintaining normal retinal function. Our knowledge of the molecular mechanisms involved in controlling retinal blood flow remains incomplete, yet improved understanding could lead to the development of new interventions aimed at restoring adequate tissue perfusion in ocular disease states such as diabetic retinopathy, hypertensive retinopathy, and glaucoma. Arterioles form the main site of vascular resistance in the retinal microcirculation ¹ ; therefore, the mechanisms that control smooth muscle tone in these vessels are largely responsible for regulating regional blood flow and capillary pressure in the retina. Arteriolar smooth muscle cells are known to express a variety of functional ion channels on their plasma membranes that are involved in setting the basal level of vascular tone and in mediating contractile responses to vasoactive agents. ² We have recently shown that retinal arteriolar myocytes have large-conductance Ca²⁺-activated K⁺ (BK) channels and K_v1.5-containing voltage-activated K⁺ channels that exert a vasodilatory influence on resting vascular tone. ^{3 4 5}  

Ca²⁺-activated Cl⁻ channels (Cl_Ca) have been identified in several types of vascular smooth muscle, ^{6 7 8} including choroidal arteriolar myocytes. ⁹ Because the Cl⁻ equilibrium potential is usually positive to the resting membrane potential, opening of Cl_Ca channels results in Cl⁻ efflux and membrane depolarization. ¹⁰ In vascular myocytes, Cl_Ca channels are thought to contribute primarily to agonist-mediated responses. ¹¹ Agonists can produce vasoconstriction as a consequence of intracellular Ca²⁺ release acting directly on the contractile apparatus or indirectly by stimulating Cl_Ca channels, causing depolarization and the consequent opening of voltage-gated Ca²⁺ channels. ¹¹ Consistent with this idea, a number of studies have demonstrated that Cl⁻ channel blockers reduce the magnitude of agonist-induced contractions in the vasculature. ^{6 12 13 14 15}  

Initial work from our own laboratory has suggested that retinal arteriolar myocytes express Cl_Ca channels, ³ but their biophysical, pharmacologic, and functional properties have yet to be determined. The aim of the present study was, therefore, to identify and characterize the Ca²⁺-activated Cl⁻ current in retinal arteriolar myocytes using the whole-cell perforated patch clamp technique and to assess their functional role in modulating basal and agonist-induced changes in retinal microvascular tone using pressure myography. 

Methods

Arteriole Preparation

Animal use was performed in accordance with the guidelines of the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and the UK Animals (Scientific Procedures) Act, 1986. Male Sprague-Dawley rats (300–450 g) were euthanatized with CO₂, and their eyes were enucleated. Retinal arterioles devoid of surrounding neuropile were mechanically isolated in low Ca²⁺ Hanks solution, as previously described. ^{3 4 5} In brief, retinal segments were lightly triturated in a low Ca²⁺ Hanks solution. The resultant suspension was centrifuged at 1207g for 1 minute, the supernatant was aspirated off, and the tissue was washed again with low Ca²⁺ medium. The suspension was then stored at 21°C until needed. Arteriolar segments remained usable for up to 10 hours under these conditions. 

Patch-Clamp Electrophysiology

With use of the whole-cell perforated patch-clamp technique, ionic currents were recorded from retinal arteriolar myocytes still embedded within their parent arterioles. ^{3 4 5 16} Vessels were pipetted into a recording bath on the stage of an inverted microscope (Eclipse TE300; Nikon, Tokyo, Japan). Isolated arterioles (length, 200-4000 μm; outer diameter, 20–44 μm) were anchored down with tungsten wire slips (diameter, 50 μm) and were superfused for 20 minutes with an enzyme cocktail of collagenase 1A (0.1 mg/mL) and protease type XIV (0.01 mg/mL) to remove surface basal lamina, allowing gigaseal formation. This treatment also causes the smooth muscle and endothelial layers to separate, ⁴ and the smooth muscle cells become electrically isolated from each other. ⁵ Unless otherwise stated, arterioles were continuously superfused with normal Hanks solution at 37°C during experimentation. Voltage-clamp recordings were performed with an amplifier (Axopatch-1D; Axon Instruments, Union City, CA). Pipettes (1–2 MΩ) were filled with a Cs⁺-based solution containing amphotericin B as the perforating agent. To ensure complete block of contaminating currents through BK and K_v channels, 100 nM penitrem A and 10 mM 4-aminopyridine (4AP) were added to the external bathing medium. ^{3 4 5} After full perforation of the membrane patch had been achieved (usually 3–5 minutes), cell capacitance was determined from the time constant of a capacitance transient elicited by a 20-mV hyperpolarization from −60 mV with a sampling frequency of 20 kHz. Before experimentation, series resistance (15–20 MΩ) and cell capacitance (7–18 pF) were compensated by as much as 80% with the circuitry of the amplifier. Online leak subtraction was carried out using a P/4 protocol. All data were corrected for liquid junction potentials (2–3 mV for all bathing solutions used). Recordings were low-pass filtered at 0.5 kHz and sampled at 2 kHz by an interface (PC1200; National Instruments, Austin, TX) with the use of custom software provided by John Dempster (University of Strathclyde, Glasgow, UK). Drug solutions were delivered by a seven-way micromanifold. 

Pressure Myography

Measurement of the diameter of intact pressurized retinal arteriole segments was performed as previously described. ³ Again, vessels were visualized in a recording bath with an inverted microscope. A tungsten wire slip (75 × 2000 μm) was laid on one end of the vessel, which provided anchoring and occluded the distal open end. Vessels were cannulated with fine-glass pipettes (tip diameters, 3–10 μm) held in a patch electrode holder and connected to a pressure transducer and a water manometer. After equilibration in Hanks solution for 30 minutes at 37°C, intravascular pressure was increased to 70 mm Hg. A section of vessel at least 150 μm away from the cannula was viewed under a ×40 NA 0.6 objective focused midway through its depth. The vessels were then treated with drug-containing Hanks solution, and changes in the external diameter were measured manually from saved video images. 

Solutions

Hanks solution contained 140 mM NaCl, 6 mM KCl, 5 mM d-glucose, 2 mM CaCl₂, 1.3 mM MgCl₂, and 10 mM HEPES (pH 7.4 with NaOH). Low Ca²⁺ Hanks differed only in that it contained 0.1 mM CaCl₂. For patch-clamp experiments, the internal solution was composed of 138 mM CsCl, 1 mM MgCl₂, 0.5 mM EGTA, 0.2 mM CaCl₂, and 10 mM HEPES (pH adjusted to 7.2 using CsOH). Amphotericin B (300 μg/mL) was added to the internal solution. For ion substitution experiments, 86 mM NaCl from the Hanks solution was replaced with either equimolar NaI or Na glucuronate. For Ca²⁺-free solution, Ca²⁺ was omitted from the external medium, and 10 μM EGTA was added. 

The following were purchased from Sigma (Poole, UK): 4-aminopyridine (4-AP), amphotericin B, 9-anthracene carboxylic acid (9-AC), caffeine, collagenase 1A, disodium 4,4′-diisothiocyanatostilbene-2,2′-disulfonate (DIDS), penitrem A, protease type XIV, 4-acetamido-4′-isothiocyanatostilbene-2,2′-disulfonic acid (SITS), and tetracaine. Nimodipine was obtained from Alexis Biochemicals (Nottingham, UK), and endothelin-1 (Et-1) was obtained from Tocris (Bristol, UK). 

Statistical Analysis

Currents were normalized to cell capacitance to obtain current densities (pA/pF). Data are reported as the mean ± SEM, and n denotes the number of vessels from which recordings were made. Unless otherwise indicated, significant differences between control and experimental treatments were determined using the paired t-test. P < 0.05 was considered significant. The following labeling convention has been used to indicate the statistical significance of differences between control and test data in all figures: no asterisk, not significant; *P < 0.05; **P < 0.01; ***P < 0.001. 

The permeability of I⁻ and glucuronate relative to Cl⁻ was calculated using the Goldman-Hodgkin-Katz equation ^{17 18} :  

\[\mathrm{P}_{\mathrm{X}^{{-}}}/\mathrm{P}_{\mathrm{Cl}^{{-}}}{=}({[}\mathrm{Cl}^{{-}}{]}_{\mathrm{i}}\ \mathrm{exp}({\Delta}\mathrm{E}_{\mathrm{rev}}F/RT){-}{[}\mathrm{Cl}^{{-}}{]}_{\mathrm{res}})/{[}\mathrm{X}^{{-}}{]}_{\mathrm{o}}\]

where [X⁻]_o is the concentration of the substituted ion (I⁻ or glucuronate), [Cl⁻]_i and [Cl⁻]_res are the intracellular and “residual” extracellular concentrations of Cl⁻, respectively, and ΔE_rev is the shift in reversal potential on ion substitution. F, R, and T have their usual meanings. 

Results

Depolarization-Evoked Cl⁻ Current

In the presence of K⁺ channel inhibitors, retinal arteriolar myocytes were held at a membrane potential of −80 mV, and a series of 1-second voltage pulses was applied, incrementing in amplitude in 20-mV steps between −100 mV and +100 mV. This protocol evoked a family of inward, outward, and tail (I_tail) currents, as shown in Figure 1Ai . At negative potentials, an early inward current was followed by a sustained late current that was maintained throughout the voltage step. The early current, measured within the first 50 ms of the test pulse, activated at −40 mV, peaked at −20 mV, and reversed at +40 mV (Fig. 1Aii) . The late current, measured at the end of the 1-second test pulse, had a similar current-voltage (I–V) profile (Fig. 1Aii)but reversed close to 0 mV. The difference in the reversal potentials (E_rev) for the two currents suggests that they were carried by different ions. We have previously characterized the early current as an L-type Ca²⁺ current (I_Ca(L)). ¹⁹ The E_rev for the late current is close to the calculated value for the Cl⁻ equilibrium potential (E_Cl = +2.2 mV), suggesting that this may be a Cl⁻ current. 

I_tail reflects the continued activation of channels beyond the duration of the voltage pulse. ²⁰ To investigate the relationship between I_tail and the two preceding currents, further experiments were performed. An initial conditioning step (250 ms) was fixed at 0 mV, close to the reversal potential of the late current, and subsequent steps were applied between −100 and +100 mV in 20-mV intervals. The resultant I_tail was inward at negative voltages and outward at positive voltages (Fig. 1Bi)and had an approximately linear I–V relationship (Fig. 1Bii) . I_tail declined rapidly at negative voltages (τ for decay of I_tail at −60 mV was 45.31 ± 4.52 ms; n = 8) but was sustained during positive test steps (Fig. 1Bi) . I_tail reversed at 0 mV, suggesting that I_tail represents persistent activation of the channels underlying the late current. Because the late current and I_tail appeared to be mediated by the same population of channels, I_tail became the primary focus during experiments to characterize these currents given that I_tail was larger in amplitude and could be evaluated across a wider voltage range (i.e., −100 to +100 mV compared with −40 to +100 mV). 

To test whether the late current and I_tail are Cl⁻ currents, ion substitution experiments were performed. Cl⁻ channels in other tissues are known to be relatively impermeable to the anion glucuronate but have a higher permeability to I⁻ than to Cl⁻. ²¹ With a 500-ms duration ramp between +100 and −100 mV, applied after a 500-ms conditioning step to 0 mV, the effect of equimolar replacement of 86 mM external Cl⁻ with glucuronate or I⁻ could be rapidly assessed. The use of a negative-going ramp protocol reduced the impact of the decay of I_tail seen at negative potentials on the amplitude of the current. In the representative trace shown in Figure 2Ai , E_rev shifted from +2.2 to +12.2 mV when 86 mM external Cl⁻ was replaced with equimolar glucuronate. On average, E_rev was shifted by 12.5 mV (E_rev Cl⁻ = +3.9 ± 2.1 mV vs. E_rev glucuronate = +16.4 ± 2.5 mV; n = 8; P = 0.0004), and the amplitude of outward I_tail was substantially reduced (Fig. 2Aii ; Cl⁻ = 32.49 ± 8.13 pA/pF vs. glucuronate = 6.18 ± 1.62 pA/pF at +80 mV; n = 8; P = 0.02). Using equation 1 , relative permeablilites for Cl⁻ and glucuronate were calculated as 1:0.25. As shown in Figure 2Bi , substitution with I⁻ resulted in a negative shift in E_rev. In four cells, E_rev shifted by 25.1 mV in a negative direction (E_rev Cl⁻ = +0.9 ± 6.2 mV vs. E_rev I⁻ = −24.2 ± 6.2 mV; P = 0.002), and, based on this shift, relative permeabilities for Cl⁻ and I⁻ were calculated as 1:3.4. An attendant increase in the amplitude of I_tail (Fig. 2Bi 2ii ; Cl⁻, 34.89 ± 13.70 pA/pF vs. I⁻, 73.91 ± 24.79 pA/pF at +80 mV; P = 0.04) and the late current (Fig. 2Bi ; Cl⁻, −0.14 ± 0.36 pA/pF vs. I⁻, 13.11 ± 3.13 pA/pF at 0 mV; P = 0.03) was also apparent. These findings are consistent with the suggestion that both currents result from activation of the same channel proteins. Taken as a whole, the selectivity sequence of I⁻> Cl⁻ > glucuronate strongly suggests that the late current and I_tail are carried through Cl⁻ channels. 

In a further series of related experiments, the effects of three classical Cl⁻ channel inhibitors—9-AC, SITS, and DIDS—were tested on I_tail ²² ; 9-AC preferentially inhibited outward I_tail (Figs. 3Ai 3Bi 3C ) with an 82% ± 6% block at +100 mV (control, 49.34 ± 10.93 pA/pF vs. 9-AC, 10.06 ± 5.12 pA/pF; n = 5; P = 0.001) as opposed to a 25% ± 10% inhibition of inward I_tail at −100 mV (control, −56.66 ± 7.08 pA/pF vs. 9-AC, −39.59 ± 2.82 pA/pF; n = 5; P = 0.05). Similarly, 1 mM SITS displayed a voltage-dependent block (Figs. 3Aii , 3Bii , 3C ) with 81% ± 16% inhibition of I_tail at +100 mV (control, 39.34 ± 10.74 pA/pF vs. SITS, 3.99 ± 3.75 pA/pF; n = 6; P = 0.04) but a nonsignificant reduction of 31% ± 15% at −100 mV (control, −60.69 ± 15.19 pA/pF vs. SITS, −35.03 ± 8.74 pA/pF; n = 6; P > 0.05). On the other hand, 1 mM DIDS blocked outward and inward I_tail to a similar degree (Figs. 3Aiii 3Biii 3C ), with the currents at +100 mV and −100 mV reduced, on average, by 92% ± 9% and 78% ± 4%, respectively (+100 mV control, 39.78 ± 9.97 vs. DIDS, 3.29 ± 3.18 pA/pF; −100 mV control, −68.0 ± 12.18 pA/pF vs. DIDS, −16.65 ± 5.32 pA/pF; n = 5; P = 0.002 and P = 0.02, respectively). A lower concentration of DIDS (10 μM) did, however, exhibit a more pronounced voltage dependence of block with 57% ± 8% inhibition at +100 mV and nonsignificant block (28% ± 12%) at −100 mV (+100 mV control, 46.56 ± 9.40 vs. DIDS, 23.20 ± 6.83 pA/pF; −100 mV control, −22.97 ± 4.65 pA/pF vs. DIDS, −15.44 ± 3.61 pA/pF; n = 7; P = 0.002 and P > 0.05, respectively). Neither 9-AC, SITS, nor DIDS had any discernible effect on I_Ca,L (P > 0.05 in all cases). These pharmacologic data further substantiate the view that depolarization evokes a Cl⁻ conductance in retinal arteriolar myocytes. 

Ca²⁺ Dependence of the Cl⁻ Current

To explore the Ca²⁺ dependence of the depolarization-evoked Cl⁻ current, we compared I_tail before and after lowering of the extracellular Ca²⁺ concentration ([Ca²⁺]_o) and after investigating the effects of the I_Ca(L) inhibitor nimodipine. As shown in Figure 4Ai , both I_Ca(L) and I_tail were decreased when the normal bathing medium was substituted with Ca²⁺-free solution (no added Ca²⁺ plus 10 μM EGTA). In six vessels, I_tail was reduced on average by approximately 75% at −100 and +100 mV (Fig. 4Aii) . Bath application of 1 μM nimodipine markedly reduced I_Ca(L) and I_tail (Fig. 4B) . These results suggest that the depolarization-evoked Cl⁻ current is dependent on the influx of Ca²⁺ through L-type Ca²⁺ channels. 

To determine whether Ca²⁺ release from intracellular stores was also capable of activating the Cl⁻ current, we tested the effects of caffeine at several different voltages between −80 and +80 mV. In retinal arteriolar myocytes, 10 mM caffeine evoked rapid global Ca²⁺ transients that reflected Ca²⁺ release from intracellular ryanodine-sensitive Ca²⁺ stores. ²³ Administration of 10 mM caffeine resulted in a transient inward current at negative voltages, little or no current at 0 mV, and transient outward current at positive voltages (Fig. 5Ai) . The I–V relationship for the caffeine-induced current was approximately linear and reversed close to 0 mV (Fig. 5Aii) , consistent with a Cl⁻ current. In addition, the current was inhibited by approximately 90% in the presence of 1 mM DIDS (Fig. 5B) . To exclude the possibility that caffeine activates the Cl⁻ channels through a Ca²⁺-independent mechanism, we blocked caffeine-induced Ca²⁺ release using 100 μM tetracaine. Tetracaine totally abrogated the currents measured at −40 mV (Fig. 5C) , suggesting that caffeine activates the Cl⁻ channels by raising cytosolic Ca²⁺. 

Physiologic Significance

Results of our experiments suggested that retinal arteriolar myocytes have a Ca²⁺-activated Cl⁻ current (I_Cl(Ca)) that may be activated by Ca²⁺ influx through L-type Ca²⁺ channels or Ca²⁺ release from the sarcoplasmic reticulum. Another series of experiments was designed to establish the role of I_Cl(Ca) in regulating retinal arteriolar tone. Retinal arteriole segments were cannulated and pressurized to 70 mm Hg, and the effect of blocking I_Cl(Ca) on the external diameter of the vessels was examined. DIDS (1 mM) had no effect on basal vascular tone (external diameters, 36.2 ± 2.4 and 36.0 ± 2.3 μm before and after 5-minute application of DIDS, respectively; n = 8; P > 0.05). Ca²⁺-activated Cl⁻ channels have been shown ¹¹ to be activated by a wide range of agonists that raise intracellular Ca²⁺. We have recently shown ²⁴ that the vasoconstrictor peptide Et-1 stimulates regenerative transient depolarizations in retinal arteriolar myocytes and that these depolarizations are rapidly blocked by the application of a Cl⁻ channel inhibitor. To examine the possible involvement of I_Cl(Ca) in modulating Et-1-induced retinal arteriolar vasoconstriction, pressurized vessels were exposed to Et-1 alone or Et-1 with 1 mM DIDS. Coapplication of DIDS dramatically reduced the level of vasoconstriction observed with 10 nM Et-1 (Fig. 6) . These findings suggest that activation I_Cl(Ca) represents a key signaling component regulating Et-1-induced vasoconstriction in the retina. 

Discussion

The present study is the first to characterize I_Cl(Ca) in the retinal microvasculature. We studied the biophysical and pharmacologic properties of I_Cl(Ca) in retinal arteriolar myocytes by focusing mainly on I_tail. Our data are consistent with I_tail mediation by Cl_(Ca) channels because it reversed close to E_Cl; the ion selectivity sequence was I⁻> Cl⁻> glucuronate, which is similar to I_Cl(Ca) in other tissues ²⁵ ; it was reduced by the classical Cl⁻ channel inhibitors 9AC, SITS, and DIDS; it was blocked if normal Hanks was substituted with Ca²⁺-free solution or if I_Ca(L) was inhibited by nimodipine; and it could be activated by release of Ca²⁺ from caffeine-sensitive Ca²⁺ stores. We also examined the effects of DIDS on basal vascular tone and vasoconstriction induced by Et-1 in isolated, pressurized retinal arterioles. Inhibition of I_Cl(Ca) by DIDS had no effect on basal vascular tone but did substantially reduce the vasoconstrictor response to Et-1. 

The electrophysiologic features of I_Cl(Ca) in retinal arteriolar myocytes resemble those reported for I_Cl(Ca) in large vessel smooth muscle in other vascular beds. In portal vein, coronary, and pulmonary artery myocytes, I_Cl(Ca) activates slowly on depolarization, is maintained throughout the voltage step, and exhibits long tail currents on repolarization. ^{8 20 26 27} In addition, in rabbit portal vein, I_tail decays exponentially at potentials negative to −20 mV but is evident as a sustained current at more positive voltages. ²⁰ It has been postulated that at negative voltages, the decline of I_tail is determined by the slow deactivation kinetics of the Cl_(Ca) channels, whereas at more positive voltages, I_tail may be sustained because of persistent influx of Ca²⁺ through noninactivating, voltage-dependent Ca²⁺ channels. ²⁰ The only apparent discrepancy in the electrophysiologic properties of the Cl_(Ca) channels in retinal arteriolar myocytes compared with those in other vascular tissues lies in their rate of deactivation. At −60 mV, I_tail decayed with a monoexponential time constant of approximately 45 ms, which is considerably faster than the rate of decline of I_tail seen in coronary artery and portal vein myocytes, where equivalent values are approximately 160 ms and 85 ms, respectively. ^{8 20} Although the exact reasons for this difference remain unclear, it is known that the deactivation kinetics of I_Cl(Ca) in vascular smooth muscle can be modified by external anions and the phosphorylation and redox status of the channels. ^{28 29 30}  

The anion permeability of Cl_(Ca) channels has been widely tested (for a review, see Large and Wang ¹¹ ). Cl_(Ca) channels are known to be less permeable to large organic anions such as glucuronate, glutamate, and isothionate (permeability relative to Cl⁻ of approximately 0.1–0.3 ^{9 27 31} ). Consistent with this, in retinal arteriolar myocytes, the calculated permeability for glucuronate was 0.25 times that of Cl⁻. Replacement of Cl⁻ with I⁻, an anion known to be more permeant through Cl_(Ca) channels, ^{11 21} resulted in a relative permeability value of 3.4. This is in close agreement with values reported for Cl_(Ca) channels in lymphatic smooth muscle (3.2 ³² ), Xenopus oocytes (3.6 ³³ ), portal vein myocytes (4.1 ³⁴ ), and ear artery smooth muscle (4.7 ⁶ ). These data indicate that the permeability properties of Cl_(Ca) channels in retinal arteriolar smooth muscle are similar to those previously reported in other vascular myocytes. 

To assess the physiological significance of I_Cl(Ca) in retinal arteriolar smooth muscle cells, we tested the effects of DIDS on basal and Et-1-evoked changes in retinal arteriolar tone. We chose DIDS, rather than 9AC or SITS, for these experiments because we found that this drug, at a concentration of 1 mM, was more effective at blocking I_Cl(Ca) at negative membrane potentials close to the resting membrane potential of retinal arteriolar smooth muscle cells (approximately −40 to −50 mV ²⁴ ). Our results showed that DIDS had no effect on the diameter of pressurized retinal arterioles, indicating that Cl_(Ca) channels do not appear to contribute to the control of basal vascular tone in these vessels, at least under in vitro conditions. Similarly, I_Cl(Ca) is not believed to participate in modulating resting vascular tone in cerebral and renal resistance vessels. ^{35 36} In contrast, several studies have suggested that the stimulation of I_Cl(Ca), which leads to membrane depolarization and the opening of voltage-gated Ca²⁺ channels, plays a pivotal role in agonist-induced contractile responses in various vascular tissues. ^{12 13 14 15} Evidence from the present work indicates similar findings in retinal arterioles because DIDS substantially attenuated the vasoconstrictor action of Et-1. A major issue that has hampered research into the functional significance of Cl_(Ca) channels in the vasculature has been the poor selectivity of available antagonists. Blockers of Cl_(Ca) channels, including DIDS, also inhibit other types of Cl⁻ currents, such as swelling- and voltage-activated Cl⁻ currents. ^{37 38} Moreover, several inhibitors of I_Cl(Ca) have been shown to modify I_Ca(L). ^{39 40} Our electrophysiologic data showed that DIDS had no effect on I_Ca(L) in retinal arteriolar myocytes at the same concentration used in the mechanical studies. However, we cannot fully discount the possibility that the effects of DIDS on the Et-1-induced vasoconstriction might have resulted from the blockade of Cl⁻ conductances other than I_Cl(Ca). 

In conclusion, this study has clearly demonstrated the presence of a Ca²⁺-activated Cl⁻ current in retinal arteriolar smooth muscle cells with characteristics similar to those reported in large vessel smooth muscle. I_Cl(Ca) does not appear to contribute to resting vascular tone in vitro, but it does appear to play a prominent role in the constriction of retinal arterioles by Et-1. Future research should now be directed toward understanding the involvement of I_Cl(Ca) in the regulation of retinal hemodynamics in vivo under normal and pathologic conditions. 

 Supported by The Juvenile Diabetes Research Foundation (US) Grant 2-2003-525, Fight for Sight (UK), and The Wellcome Trust Grant 074648/Z/04/Z.

 Submitted for publication July 3, 2008; revised August 25, 2008; accepted October 28, 2008.

 Disclosure: M.K. McGahon, None; M.A. Needham, None; C.N. Scholfield, None; J.G. McGeown, None; Tim M. Curtis, None

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

 

Figure 1.

 

Early, late, and tail currents elicited by step depolarizations in retinal arteriolar myocytes. (Ai) Representative current records showing early, late, and tail currents evoked by a 1-second voltage step protocol (inset) applied from a holding potential of −80 mV to voltages between −100 and +100 mV in 20-mV increments. (Aii) Average early and late current densities plotted against test voltage (n = 9). (Bi) Tail currents elicited by a test steps to voltages between −100 and +100 mV in 20-mV increments (inset) applied after a 250-ms conditioning step from −80 to 0 mV. (Bii) Average tail current densities (measured 10 to 20 ms after the start of the test step) plotted against the tail step voltage (n = 8).

Figure 2.

 

Effect of substitution of external Cl⁻ on I_tail. (Ai) Representative current traces elicited by a 500-ms voltage ramp protocol (inset) from +100 to −100 mV applied after a 500-ms conditioning step to 0 mV recorded in normal Hanks solution and in glucuronate-substituted Hanks. Arrows: reversal potentials. (Bi) Original current traces recorded in normal Hanks solution and in I⁻-substituted Hanks. (A, Bii) Average current densities at +80 mV in Cl⁻ only versus (A) glucuronate- and (B) I⁻-containing solutions (n = 8 and n = 4, respectively).

Figure 3.

 

Pharmacologic analysis of I_tail. I_tail elicited by alternating test steps to −100 and +100 mV applied after a conditioning step to 0 mV in normal Hanks solution (control) versus (Ai) 1 mM 9-AC, (Aii) 1 mM SITS, and (Aiii) 1 mM DIDS. Average current densities measured in 20-mV intervals between −100 and +100 mV in the normal Hanks solution versus (Bi) 1 mM 9-AC, (Bii) 1 mM SITS, and (Biii) 1 mM DIDS plotted against the test step voltage (n = 5, n = 6 and n = 5, respectively). (C) Percentage inhibition of I_tail by 9-AC, SITS, and DIDS at different voltages calculated from the data in (B).

Figure 4.

 

Dependence of I_tail on external Ca²⁺. (Ai) I_tail elicited by alternating test steps to −100 and +100 mV (inset) applied after a conditioning step to 0 mV in normal Hanks solution (control) and Ca²⁺-free Hanks (0Ca²⁺ _o). (Bi) I_tail elicited in normal Hanks solution (control) and the L-type Ca²⁺ channel inhibitor nimodipine (1 μM). Nimodipine reduced I_Ca(L) by 85% and I_tail by 81% and 86% at −100 and +100 mV, respectively. (A ii, Bii) Average current densities measured at −100 and +100 mV in the normal Hanks solution versus (A) Ca²⁺-free Hanks and (B) 1 μM nimodipine (nim; n = 6 and n = 10, respectively).

Figure 5.

 

Activation of I_Cl(Ca) by Ca²⁺ release from intracellular Ca²⁺ stores. (A i) Original traces showing caffeine-induced currents recorded at holding potentials of −80 to +80 mV in 40-mV intervals. (Aii) Mean ± SEM of caffeine-induced current densities (measured as peak current vs. baseline at each holding potential; n = 11). (B) Caffeine-induced currents elicited at a holding potential of −40 mV before (B i) and during (B ii) application of 1 mM DIDS. (Biii) Average caffeine-induced current densities recorded at −40 mV before and during application of DIDS (n = 6). (C) Caffeine-induced currents elicited at a holding potential of −40 mV before (C i) and during (C ii) application of 100 μM tetracaine. (Ciii) Average caffeine-induced current densities recorded at −40 mV before and during application of tetracaine (n = 6).

Figure 6.

 

Effect of DIDS on the Et-1-induced changes in retinal arteriolar tone. (A) Photomicrographs of a pressurized (70 mm Hg) retinal arteriole (Ai) before and (Aii) 40 seconds after application 10 nM Et-1. (B) Photographs of a different vessel (B i) before and (Bi i) 40 seconds after application 10 nM Et-1 and 1 mM DIDS. Scale bars, 10 μm. (C) Mean data showing the external diameter of arterioles exposed to Et-1 alone or in combination with DIDS expressed as a fraction of the initial resting diameter before drug exposure.