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¹ Department of Pharmacology and ² Department of Physiology and Cell Biology, Center of Biomedical Research Excellence (COBRE), University of Nevada School of Medicine, Reno, NV 89557
³ Division of Basic Medical Sciences, St George's, University of London, London, SW17 0RE UK

Correspondence to Iain A. Greenwood or Normand Leblanc: grenwood{at}sgul.ac.uk or NLeblanc{at}unr.edu

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The aim of the present study was to provide a mechanistic insight into how phosphatase activity influences calcium-activated chloride channels in rabbit pulmonary artery myocytes. Calcium-dependent Cl^– currents (I_ClCa) were evoked by pipette solutions containing concentrations between 20 and 1000 nM Ca²⁺ and the calcium and voltage dependence was determined. Under control conditions with pipette solutions containing ATP and 500 nM Ca²⁺, I_ClCa was evoked immediately upon membrane rupture but then exhibited marked rundown to 20% of initial values. In contrast, when phosphorylation was prohibited by using pipette solutions containing adenosine 5'-(ß,-imido)-triphosphate (AMP-PNP) or with ATP omitted, the rundown was severely impaired, and after 20 min dialysis, I_ClCa was 100% of initial levels. I_ClCa recorded with AMP-PNP–containing pipette solutions were significantly larger than control currents and had faster kinetics at positive potentials and slower deactivation kinetics at negative potentials. The marked increase in I_ClCa was due to a negative shift in the voltage dependence of activation and not due to an increase in the apparent binding affinity for Ca²⁺. Mathematical simulations were carried out based on gating schemes involving voltage-independent binding of three Ca²⁺, each binding step resulting in channel opening at fixed calcium but progressively greater "on" rates, and voltage-dependent closing steps ("off" rates). Our model reproduced well the Ca²⁺ and voltage dependence of I_ClCa as well as its kinetic properties. The impact of global phosphorylation could be well mimicked by alterations in the magnitude, voltage dependence, and state of the gating variable of the channel closure rates. These data reveal that the phosphorylation status of the Ca²⁺-activated Cl^– channel complex influences current generation dramatically through one or more critical voltage-dependent steps.

Abbreviations used in this article: AMP-PNP, adenosine 5'-(ß,-imido)-triphosphate; PSS, physiological salt solution.

	INTRODUCTION

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In smooth muscle cells, Cl^– ions are actively accumulated by three major uptake mechanisms: the Na-K-2Cl cotransporter, the Cl:HCO₃ exchanger, and a poorly characterized "pump III" (Chipperfield and Harper, 2000). The product of these transporter systems is an intracellular [Cl^–] lying between 30 and 80 mM (Chipperfield and Harper, 2000). This results in an electrochemical gradient favoring Cl^– efflux at the measured resting membrane potential of –40 to –60 mV. As a consequence, activation of Cl^– channels is an important mechanism to increase smooth muscle cell excitability by depolarizing the cell membrane potential (Large and Wang, 1996; Leblanc et al., 2005).

The most extensively recorded Cl^– channel current in smooth muscle cells is evoked by a rise in intracellular [Ca²⁺], the so called calcium-activated chloride current (I_ClCa). There have been numerous studies reporting the activation of I_ClCa by various agents and manipulations in smooth muscle cells (Large and Wang, 1996; Leblanc et al., 2005). However, the molecular identity remains elusive and little is known about how the channel is gated by a rise in [Ca²⁺] or how intracellular regulators modify this process. Recent studies revealed that the activity of Ca²⁺-dependent Cl^– channels (Cl_Ca) is influenced by Ca²⁺-dependent enzymes. For example, blockers of the Ca²⁺-calmodulin–dependent kinase CaMKII prolong the duration of I_ClCa in myocytes isolated from trachea (Wang and Kotlikoff, 1997) and enhance I_ClCa in pulmonary and coronary artery smooth muscle cells (Greenwood et al., 2001). The suppressive role of CaMKII in pulmonary artery myocytes was substantiated by the use of constitutively active CaMKII (Greenwood et al., 2001). Subsequent experiments established that a counter mechanism of regulation is provided by the Ca²⁺-dependent serine/threonine phosphatase calcineurin or PP2B (Ledoux et al., 2003; Greenwood et al., 2004). Moreover, in pulmonary artery myocytes, the positive regulation of I_ClCa depends on the isoform of the catalytic subunit (Greenwood et al., 2004). Consequently, while it is axiomatic that generation of I_ClCa relies upon an increase in [Ca²⁺]_i, dephosphorylation of the channel complex also determines I_ClCa activity.

The aim of the present study was to undertake a rigorous examination of the activation of I_ClCa by intracellular [Ca²⁺] under conditions where phosphorylation is supported or minimized. To obviate any reliance upon Ca²⁺ influx or Ca²⁺ release mechanism, I_ClCa was activated by pipette solutions containing free [Ca²⁺]_i set at known concentrations. This technique has been employed to characterize similar conductances in lacrimal cells (Evans and Marty, 1986), parotid acinar cells (Arreola et al., 1996), endothelial cells (Nilius et al., 1997), and Xenopus oocytes (Kuruma and Hartzell, 2000). Recently we have used this technique to study I_ClCa in smooth muscle cells isolated from hepatic portal vein, pulmonary artery, and coronary artery (e.g., Greenwood et al., 2001; Ledoux et al., 2003; Greenwood et al., 2004; Ledoux et al., 2005). With pipette solutions containing free Ca²⁺ higher than the threshold for activation, I_ClCa was sustained and exhibited distinctive voltage-dependent kinetics following membrane depolarization. In the present study, I_ClCa was recorded with pipette solutions containing different [Ca²⁺] ranging from 20 to 1000 nM and either 3 mM ATP or 3 mM adenosine 5'-(ß,-imido)-triphosphate (AMP-PNP) to assess the mechanistic effect of dephosphorylation on the Cl^– conductance. As the terminal phosphate of AMP-PNP is resistant to hydrolysis (Yount, 1975), this compound prohibits substrate phosphorylation and has been used in studies to explore the role of phosphorylation (e.g., regulation of CFTR; Gadsby and Nairn, 1999). Consequently, the action of endogenous phosphatases was accentuated. These experiments revealed calcium and voltage dependency of I_ClCa activation. Moreover, the results provide a novel insight into how the gating of the Ca²⁺-activated Cl^– channel is influenced by dephosphorylation.

	MATERIALS AND METHODS

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Isolation of Freshly Dissociated Pulmonary Artery Myocytes
A similar technique to that previously used by our group (Greenwood et al., 2001, 2004) was used to isolate smooth muscle cells. In brief, cells were prepared from the main and secondary pulmonary arterial branches dissected from New Zealand white rabbits (2–3 kg) killed by anesthetic overdose in accordance with British and American regulations. After dissection and removal of connective tissue, the pulmonary arteries were cut into small strips and incubated overnight (16 h) at 4°C in a low Ca²⁺ physiological salt solution (PSS) containing either 10 or 50 µM CaCl₂ and 1 mg ml^–1 papain, 0.15 mg ml^–1 dithiothreitol, and 2 mg ml^–1 BSA. The next morning, the tissue strips were rinsed three times in low Ca²⁺ PSS and incubated in the same solution for 10 min at 37°C. Cells were released by gentle agitation with a wide bore Pasteur pipette. Cells were stored at 4°C and used within 10 h after dispersion.

Patch Clamp Methods
Ca²⁺-activated Cl^– currents were elicited in conventional whole-cell path clamp mode by pipette solutions containing 10 mM BAPTA and free [Ca²⁺] set to values ranging from 20 to 1000 nM by the addition of 0.84–8.7 mM CaCl₂ as determined by the calcium chelator program EQCAL (Biosoft). Free [Ca²⁺] was verified independently using a Ca²⁺-sensitive electrode (Thermo Orion, model 93–20) using calibrated solutions (CALBUF-2; World Precision Instruments Inc.). These [Ca²⁺] constitute a dynamic range experienced by smooth muscle cells physiologically (e.g., ZhuGe et al., 2002). It is worth stressing that contraction of the myocyte was observed with pipette solutions containing Ca²⁺ >100 nM as Ca²⁺ flooded into the cell following membrane rupture. No attempt was made to block this contraction (e.g., by inhibition of myosin light chain kinase) as this would introduce another variable into the recording conditions. Contamination of I_ClCa from other types of current was minimized by the use of CsCl and TEA in the pipette solution, and TEA in the external solution. Control pipette solutions contained 3 mM ATP, whereas the test internal solutions contained 3 mM AMP-PNP. On any given experimental day, pipette solutions containing ATP were rigorously alternated with one containing AMP-PNP. However only one Ca²⁺ concentration could be practically tested on the same day due to the low success rate of maintaining a stable recording for the entire 20 min of cell dialysis. Consequently, data for each group were collected in cells from at least two animals. In all cases, the cell capacitance was similar across the whole study. Mean ± SEM cell capacitances measured from cells dialyzed with ATP for the following pipette Ca²⁺ concentrations were as follows: 20 nM, 21.3 ± 6.4 pF (n = 5); 100 nM, 18.8 ± 3.0 pF (n = 3); 250 nM, 21.1 ± 1.6 pF (n = 13); 500 nM, 18.2 ± 1.3 pF (n = 11); 750 nM, 16.4 ± 1.7 pF (n = 8); 1000 nM, 13.1 ± 1.4 pF (n = 6). Conversely, for the AMP-PNP group of data, mean ± SEM cell capacitances for the same pipette Ca²⁺ concentrations were as follows: 20 nM, 21.3 ± 2.4 pF (n = 5); 100 nM, 19.3 ± 1.9 pF (n = 8); 250 nM, 22.3 ± 3.9 pF (n = 7); 500 nM, 18.5 ± 1.1 pF (n = 15); 750 nM, 18.1 ± 1.5 pF (n = 13); 1000 nM, 19.1.1 ± 1.9 pF (n = 16). The relatively small error bars in each group combined with the high correlation coefficients of the Hill equation fits to the data (see Fig. 3) gave us confidence in our ability to determine accurately the Ca²⁺ dependence of I_ClCa in phosphorylated and dephosphorylated conditions.

I_ClCa was evoked immediately upon rupture of the cell membrane, and the voltage-dependent properties were monitored every 10–20 s by stepping from a holding potential (V_h) of –50 mV to either +70 or +90 mV for 750 ms or 1 s, followed by repolarization to –80 mV for 0.5 or 1 s. Current–voltage relationships were constructed by stepping in 10-mV increments from V_h to test potentials between –100 mV and +130 mV for 1 s after 20 min dialysis. I_ClCa was represented as the chord conductance normalized to cell capacitance determined from a 10-mV hyperpolarizing pulse from V_h. The calcium dependence of I_ClCa was determined at each test potential by plotting the mean conductance at the end of the test step (t = 1 s) from n cells against pipette [Ca²⁺] and fitting the data with the Hill equation (Eq. 1) lacking fitting constraints on upper and lower asymptotes. For Eq. 1, Y is the Cl^– conductance (nS/pF), Y_max is the maximal conductance, K_d is the apparent binding affinity constant, is the Hill coefficient, and c is a constant:

(1)

The voltage dependence of I_ClCa generation was assessed by plotting the mean chord conductance against test potential for each pipette [Ca²⁺]. These data were then fitted by a Boltzmann function (Eq. 2) where G is the conductance at a given potential, G_max is the maximum conductance, V is the voltage, V_0.5 is the voltage required for half-maximal amplitude, k is the steepness of the voltage dependence, and c is a constant:

(2)

The kinetics of I_ClCa were generally well fitted by a single exponential function although two exponential terms were required in some cases at higher [Ca²⁺] (e.g., >500 nM; see Fig. 9). The formula describing such fits is the following:

(3)

where I is the amplitude of the current, A₁ (equal to 0 for a single exponential term) and A₂ are the amplitudes of the fast and slow component of deactivation, respectively, t is the time, _fast and _slow are the fast and slow time constant deactivation, respectively, and c is a constant.

Solutions
Single pulmonary arterial myocytes were isolated by incubating pulmonary arterial tissue strips in the following low Ca²⁺ PSS (in mM): NaCl (120), KCl (4.2), NaHCO₃ (25; pH 7.4 after equilibration with 95% O₂–5% CO₂ gas), KH₂PO₄ (1.2), MgCl₂ (1.2), glucose (11), taurine (25), adenosine (0.01), and CaCl₂ (0.01 or 0.05). The K⁺-free bathing solution used in all patch clamp experiments had the following composition (in mM): NaCl (126), HEPES-NaOH (10, pH 7.35), TEA (10), glucose (20), MgCl₂ (1.2), and CaCl₂ (1.8). The pipette solution had the following composition (in mM): TEA (20), CsCl (106), HEPES-CsOH (10, pH 7.2), BAPTA (10), GTPNa₂ (0.2), MgCl₂ (0.42), and either 3 mM ATP or 3 mM AMP-PNP (both Mg and Na salts of ATP were used because a Mg salt of AMP-PNP was unavailable and quantitatively similar results were obtained) or no added nucleotide (0 ATP). All enzymes and reagents were purchased from Sigma-Aldrich.

Computer Simulations
The behavior of macroscopic Cl_Ca channel activity in pulmonary artery myocytes dialyzed with ATP and AMP-PNP was mathematically simulated using Markov chain kinetic models that were solved numerically by Axon Engineer software (version 2.11c, Aeon Software Inc.) run under DOS on a PC (Pentium III, 800 MHz) running under Windows ME platform. Ordinary differential equations were simultaneously solved by the Gear numerical integration method using incremental time steps of 0.1 µs in duration with enabled stiffness constraint. All voltage clamp simulations lasted 2.5 s and were initiated from a holding potential of –50 mV under steady-state conditions. Simulated time-dependent currents elicited with 20, 100, 250, 500, 750, and 1000 nM Ca²⁺ for potentials ranging from –100 to +140 mV (20-mV increments), and steady-state conductance vs. voltage curves at each [Ca²⁺] generated by Axon Engineer were exported in ASCII format into Microsoft Office Excel 2003 and then into Origin 7.5 (OriginLab Corp.). The specific parameters and equations used in the simulations are listed in Table I .

Online Supplemental Material
Full details of the kinetic analysis of I_ClCa recorded in myocytes dialyzed with ATP or AMP-PNP and the parameters used for the computer simulations can be found in the online supplemental material (available at http://www.jgp.org/cgi/content/full/jgp.200609507/DC1).

	RESULTS

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Cell Dialysis with AMP-PNP, a Nonhydrolyzable Analogue of ATP, Attenuates the Rundown of I_ClCa in Pulmonary Artery Myocytes
Under conditions prohibiting the activity of K⁺ channels, intracellular dialysis of rabbit pulmonary artery myocytes with a pipette solution containing free Ca²⁺ concentration >100 nM consistently elicited a membrane current that had characteristics identical to the "classical" Ca²⁺-activated Cl^– current (I_ClCa) evoked by a similar method in this preparation (Greenwood et al., 2001; Piper et al., 2002; Greenwood et al., 2004) and other cell types (Evans and Marty, 1986; Arreola et al., 1996; Nilius et al., 1997; Kuruma and Hartzell, 2000; Ledoux et al., 2003). These include the following: (a) anion selectivity with a permeability sequence of SCN^– >> I^– > Cl^– >> aspartate; (b) activation by [Ca²⁺]_i in the range of 100 to 1000 nM; (c) strong outward rectification due to voltage-dependent gating; (d) time-dependent activation and deactivation; and (e) sensitivity to block by niflumic acid, a putative I_ClCa blocker. This approach bears a great advantage over other conventional methods used to elicit I_ClCa (e.g., by spontaneous or evoked Ca²⁺ release from the sarcoplasmic reticulum, or by Ca²⁺ entry through voltage-gated Ca²⁺ channels; for a review see Leblanc et al., 2005) in that potentially undesirable effects of manipulating phosphorylation status on the Ca²⁺ source triggering I_ClCa can be effectively eliminated, thus allowing a direct study of how phosphorylation influences the channels themselves or any regulatory subunits.

Fig. 1 A shows that I_ClCa elicited by 500 nM Ca²⁺ exhibited rapid rundown in cells dialyzed with 3 mM ATP. The amplitude of I_ClCa remained constant once the initial rundown period had passed so that after the 20-min recording period, the relative amplitude of I_ClCa was 17 ± 6% of initial control amplitude at t = 0 (n = 5; Fig. 1 B). In contrast, I_ClCa elicited with pipette solutions containing 500 nM Ca²⁺ and AMP-PNP, a nonhydrolyzable analogue of ATP, recovered steadily after an initial attenuated rundown so that after 20 min recording, the amplitude of I_ClCa at +90 mV was 105 ± 17% of the current recorded at t = 0 (n = 6; Fig. 1 B). The pattern of rundown and recovery of I_ClCa was similar in experiments performed with a pipette solution lacking ATP (nominally zero ATP; unpublished data). These data are consistent with recent findings showing that I_ClCa in smooth muscle cells is down-regulated by phosphorylation through CaMKII, an effect that is antagonized, at least in part, by calcineurin (Greenwood et al., 2001, 2004; Ledoux et al., 2003). These observations suggest that the rundown of I_ClCa in pulmonary artery myocytes dialyzed with ATP was likely due to a shift in the phosphorylation status in the vicinity of the channel. The following series of experiments aimed to determine the biophysical mechanisms driving I_ClCa gating under different conditions of global cellular phosphorylation, that is after 20 min cell dialysis with 3 mM ATP or 3 mM AMP-PNP.

View larger version (16K): [in this window] [in a new window]   Figure 1. Attenuation of rundown of Ca2+-activated Cl– current in rabbit pulmonary artery myocytes by intracellular dialysis with a nonhydrolyzable form of ATP, AMP-PNP. (A) Representative current traces from typical experiments showing the time-dependent changes of Ca2+-activated Cl– current recorded from pulmonary arterial smooth muscle cells dialyzed with 500 nM Ca2+ and 3 mM ATP (first row) or 3 mM AMP-PNP (second row). Currents recorded immediately after breaking the seal (0 min), and after 2 and 20 min of cell dialysis were elicited by repetitive steps (every 10 s) to +90 mV lasting 1 s from a holding potential (HP) of –50 mV . Each depolarizing pulse to +90 mV was followed by a repolarizing step to –80 mV to enhance the magnitude of the tail current. The voltage clamp protocol is shown below the traces. (B) Similar to the experiments depicted in A, this graph shows the mean time course of changes of normalized IClCa amplitude elicited by 1-s depolarizing pulses to +90 mV, followed by 1-s repolarizing steps to –80 mV, in the presence of 3 mM ATP (filled squares; n = 5) or 3 mM AMP-PNP (empty squares; n = 6); each step was applied from HP = –50 mV at a frequency of one pulse every 10 s for 20 min. Please note the attenuation of rundown of IClCa and the delayed recovery of this current in cells dialyzed with AMP-PNP.

View larger version (10K): [in this window] [in a new window]   Figure 2. General properties of Ca2+-activated Cl– currents recorded from cells dialyzed with distinct free Ca2+ concentrations in the presence of ATP or AMP-PNP. Typical families of IClCa recorded with pipette solutions containing 20, 100, 250, 500, 750, or 1000 nM Ca2+, with either 3 mM ATP (left) or 3 mM AMP-PNP (right). All families of currents were evoked by the protocol shown at bottom. Notice the different vertical calibration bars for the traces recorded with ATP and AMP-PNP. Besides being much larger in cells dialyzed with AMP-PNP versus ATP, IClCa activated more quickly and deactivated more slowly with AMP-PNP. All traces were obtained after 20 min of cell dialysis.

View larger version (21K): [in this window] [in a new window]   Figure 3. Calcium dependence of IClCa at different membrane potentials recorded after prolonged cell dialysis with ATP or AMP-PNP. For the experiments conducted with ATP (A; n = 3–13) and AMP-PNP (B; n = 5–16), all data from experiments identical to those described in Fig. 2 were pooled and the mean chord conductance ± SEM (calculated using Eq. 1 in the Materials and Methods) at each step potential plotted as a function of pipette Ca2+ concentration ranging from 20 to 1000 nM Ca2+. Data at many potentials were purposely omitted for the sake of clarity but values extracted from such potentials are represented in Fig. 4. Each data set was fitted with the Hill equation (Eq. 1 in Materials and Methods) for determination of the Ca2+ affinity and Hill coefficient at a given step potential, which are described in Fig. 4. In A, the Ca2+ dependence of the chord conductance measured with ATP at potentials negative to 0 mV could not be fitted due to the small and variable magnitude of the macroscopic IClCa recorded in the negative range of membrane potentials. All plots were generated from data obtained after 20 min of cell dialysis with either nucleotide.

View larger version (12K): [in this window] [in a new window]   Figure 4. The Ca2+ sensitivity and number of Ca2+ ions required for activation of IClCa are not influenced by the global state of phosphorylation. (A) Graph showing the voltage dependence of the Ca2+ affinity of IClCa (apparent Kd for Ca2+) derived from experiments obtained with 3 mM ATP (filled squares) or 3 mM AMP-PNP (empty squares). Mean Kd ± fitting error (error scaled to the square root of reduced 2 as calculated by Origin software) for Ca2+ at each voltage was estimated from curve fitting of the data to the Hill equation as represented in Fig. 3. The line passing through the data points is a Boltzmann fit to the data points and is described by the following parameters: Kd for Ca2+ = {748.1/ [1 + exp((V – 30.3)/36.7)]} + 178.42, where V is membrane potential. (B) Graph illustrating the voltage dependence of the Hill or cooperativity coefficient () obtained from analysis of the Ca2+ dependence of IClCa with the Hill equation (Fig. 3) after 20 min of cell dialysis with ATP (filled squares) or AMP-PNP (empty squares). The solid line is a single exponential least-square fit to the data and is described by the following formula: = 0.19 * exp(–V/48.3) + 2.22, where V is membrane potential. All plots were generated from data obtained after 20 min of cell dialysis with either nucleotide.

As shown in Fig. 5 , the voltage dependence of activation of I_ClCa spanned >230 mV, exceeding the practical range of measurement. The half-maximal activation voltage (V_0.5) values in ATP and AMP-PNP at low Ca²⁺ were positive to +130 mV. To characterize the voltage dependence of current activation, we fitted the data with a Boltzmann function. The data generated with AMP-PNP (Fig. 5 B) for 500, 750, and 1000 nM Ca²⁺ extrapolated to maximum conductances between 0.95 and 1.78 nS/pF with an average of 1.16 nS/pF. The fact that extrapolation of the Boltzmann relationships calculated for these three data sets, each of which started at a different basal level in the negative range of membrane potentials (Fig. 5 B), yielded similar extrapolated maximal conductances (considering they were obtained from different experimental series) provided support to our approach of using the mean value of 1.16 nS/pF as the maximum conductance to fit all data sets (see Discussion). In both groups, increasing the pipette [Ca²⁺] produced a leftward shift in the voltage-dependent activation, manifest as a decrease in the calculated potential for V_0.5. However, the most remarkable difference between cells dialyzed with ATP (Fig. 5 A) and AMP-PNP (Fig. 5 B) was the large elevation of the minimal or basal conductance level at negative potentials in the cells dialyzed with AMP-PNP for pipette [Ca²⁺] 250 nM. A consequence of the marked basal elevation of Cl_Ca conductance was that the channels were less influenced by membrane potential within the physiological range of voltages, especially at higher intracellular Ca²⁺ levels (750 and 1000 nM Ca²⁺). For example, Cl_Ca conductance only increased from 0.59 ± 0.11 nS/pF at –60 mV to 0.63 ± 0.13 nS/pF at –20 mV with 1000 nM Ca²⁺.

View larger version (22K): [in this window] [in a new window]   Figure 5. Voltage dependence of IClCa analyzed after prolonged dialysis with ATP and AMP-PNP. The same data from the experiments described in Fig. 2 were used to construct the voltage dependence of IClCa for each pipette Ca2+ concentration ranging from 100 to 1000 nM in the presence of 3 mM ATP (A) or AMP-PNP (B). The two graphs shown in A and B report the mean ± SEM chord conductance of fully activated IClCa as a function of step potential ranging from –100 to +130 mV. Data points at or around 0 mV were not included due to the small current near the equilibrium potential for Cl–. All lines are least-square Boltzmann fits to the data (Eq. 2) from which we extracted the half-maximal voltage (V0.5), which are reported in Fig. 6. All sigmoidal relationships in the two panels were fitted by constraining each fit to a maximal conductance of 1.16 nS/pF; the latter value is the mean of the maximal conductance estimated from curve fitting of the data obtained with 500, 750, and 1000 nM Ca2+ and AMP-PNP, which yielded similar estimates. Mean data points in A and B are reproduced from Fig. 3 but plotted differently. All plots were generated from data obtained after 20 min of cell dialysis with either nucleotide.

View larger version (11K): [in this window] [in a new window]   Figure 6. A reduction in the global state of phosphorylation by cell dialysis with AMP-PNP causes a pronounced shift of the voltage dependence of IClCa toward negative potentials. (A) From the experiments conducted with pipette solutions containing ATP (filled squares) or AMP-PNP (empty squares), the mean ± fitting error (error scaled to the square-root of reduced 2 as calculated by Origin software) of half-maximal activation voltages (V0.5) determined from the analyses outlined in Fig. 5 (Eq. 2) were plotted as a function of internal Ca2+ concentration ([Ca2+]). The lines passing through the data points are least-square exponential fits to the data points and are described by the following formulas: ATP, V0.5 = 608.8 * exp(–[Ca2+]/69.2) + 205.5; AMP-PNP, V0.5 = 482.3 * exp(–[Ca2+]/119) + 66.1. Inset, graph illustrating the Ca2+ dependence of the slope factor k extracted from analysis of the voltage dependence of the conductance of IClCa (Fig. 5) measured with ATP (filled squares) and AMP-PNP (empty squares). As for V0.5, each data point is mean ± fitting error (error scaled to the square-root of reduced 2 as calculated by Origin software) of the k value determined for each data set. (B) In this graph, the ratio of the mean half-maximal voltage obtained in ATP over that in AMP-PNP (V0.5ATP/V0.5AMP-PNP; derived from A) was plotted as a function of pipette Ca2+ concentration ([Ca2+]i). The slope of the linear regression passing through the calculated data points is significantly different from 0, with P = 0.025. The parameters of the equation determining this regression are: V0.5ATP/V0.5AMP-PNP = 0.0023 * [Ca2+]i + 1.35 (r2 = 0.852). All plots were generated from data obtained after 20 min of cell dialysis with either nucleotide.

View larger version (11K): [in this window] [in a new window]   Figure 7. Ca2+ dependence of IClCa kinetics recorded from ATP- and AMP-PNP–loaded myocytes. (A and B) Bar graphs summarizing the effects of cell dialysis with 3 mM ATP (open bars) or 3 mM AMP-PNP (filled bars) on the time constant of activation at +130 mV (act; A) and deactivation at –100 mV (deact; B) of IClCa elicited with either 250 or 1000 nM Ca2+ in the pipette solution. Please note in B that deact measured in AMP-PNP and 1000 nM Ca2+ is the slower of the two time constants of deactivation estimated by least-squares biexponential fitting. For both panels, each bar represents a mean ± SEM of four to five measurements. Two-way ANOVA tests were used to assess statistical significance between mean data obtained in ATP and AMP-PNP; with 1000 nM Ca2+, mean act (A) and deact (B) in ATP were significantly different from those estimated with AMP-PNP (, P < 0.001, determined with LSD post-hoc test). Both bar graphs were generated from data obtained after 20 min of cell dialysis with either nucleotide. A more thorough analysis of the effects of the two nucleotides on IClCa kinetics can be found in the online supplemental material.

Computer Simulations
The behavior of Ca²⁺-activated Cl^– currents recorded with ATP and AMP-PNP was simulated mathematically using methods employed previously by our group (Remillard and Leblanc, 1996; Ledoux et al., 2005) based upon the following scheme proposed by Kuruma and Hartzell (2000) to describe I_ClCa recorded from Xenopus oocytes.

(SCHEME 1)

We adapted this model to best reproduce our data by implementing only minor modifications to the parameters used by Kuruma and Hartzell (2000). A full justification for setting and adjusting the rate constants and gating variables of the various kinetic steps is provided in the online supplemental material. This model incorporates the binding of three Ca²⁺ ions (suggested by our experimental data; see Fig. 4 B) and allows the channel to open (O₁, O₂, and O₃) from each of the Ca²⁺-bound states (C₁–C₄). In this model, the rate of Ca²⁺ binding is a simple first-order reaction scheme that is directly proportional to [Ca²⁺] and is equal to [Ca²⁺] * k_on, where k_on is a rate constant in M^–1s^–1; the rate of channel closure is voltage dependent (ß₁, ß₂, and ß₃; units are s^–1). Similar to Kuruma and Hartzell (2000), transitions between the three open states were forbidden. We first modeled I_ClCa as recorded from AMP-PNP–loaded cells because our analysis of the voltage dependence of I_ClCa showed that a maximal Cl_Ca conductance of 1.16 mS/cm² was consistently observed for pipette [Ca²⁺] 500 nM Ca²⁺ in these experiments (Fig. 5 B), and this dictated an upper conductance limit for both groups of data. Since our results indicated that the Ca²⁺ sensitivity of I_ClCa and Hill coefficient were not significantly different between ATP- and AMP-PNP–loaded myocytes, at least at positive potentials, the only parameters that were adjusted were the value of the gating variables (between 0 and 1), which determines whether the channel is closed or open, and the magnitude and voltage dependence of the closure rate constants. All other parameters were identical as shown in Table I. To incorporate the effects of phosphorylation into the model, we noted the lack of Ca²⁺ dependence of _act in the presence of ATP (see Fig. S1 A) and we thus hypothesized that phosphorylation causes open channel block at higher Ca²⁺ bound states (O₂ and O₃). Accordingly, we set the gating variable of O₂ and O₃ to 0 for the ATP model (Table I). We also increased the magnitude of the rates of closure (ß(V)_x) and shifted the voltage dependence toward more positive potentials compared with currents recorded with AMP-PNP (Table I).

Fig. 8 shows families of simulated I_ClCa currents elicited from a holding potential of –50 mV with [Ca²⁺] facing the internal side of the membrane ranging from 20 to 1000 nM in conditions that support phosphorylation (ATP) or conditions that prohibit phosphorylation (AMP-PNP). Currents were generated at different potentials from –100 to +140 mV applied for 1 s. As for experimentally derived data, much larger currents were simulated in the presence of AMP-PNP than ATP (notice the different vertical calibrations). In both groups, threshold for activation was near 100 nM and saturated between 750 and 1000 nM Ca²⁺. Currents in ATP displayed signs of strong outward rectification at all [Ca²⁺], whereas this feature was much less apparent for the AMP-PNP–simulated currents especially at higher [Ca²⁺]. Indeed with [Ca²⁺] 500 nM, large instantaneous currents were apparent, which reflected high occupancy of the open states at the holding potential. A visual inspection of Fig. 8 also shows that the kinetics of activation are faster and the kinetics of deactivation are slower with AMP-PNP versus ATP. A short survey comparing some kinetic parameters of experimental and modeled data can be found in the online supplemental material, which demonstrates a very good correlation between our computer-simulated currents and I_ClCa measured in our experiments.

View larger version (8K): [in this window] [in a new window]   Figure 8. Modeling of IClCa under conditions simulating prolonged cell dialysis with ATP or AMP-PNP. The nomenclature of this figure is identical to that of actual data shown in Fig. 2 except that the last potential of the voltage clamp protocol was +140 mV instead of +130 mV, and voltage steps were incremented by +20 mV from –100 mV. The conditions and parameters used for the simulation are described in the text (Materials and Methods and Results) and Table I. Again, note the different calibration bars for the simulations with ATP and AMP-PNP.

View larger version (25K): [in this window] [in a new window]   Figure 9. Comparison of experimental and simulated IClCa data. (A) Plots of simulated chord conductance vs. [Ca2+]i relationships derived from analysis of data generated with the ATP (a) and AMP-PNP (b) models at potentials ranging from –100 to +130 mV (HP = –50 mV), as indicated by the different symbols. All solid lines are fits of the simulated sets of data to the Hill equation (Eq. 1) calculated by Origin software. Parameters calculated from such fits are presented in C. Please note the remarkable similarity between such graphs and those represented in Fig. 3. (B) Plots of simulated chord conductance vs. voltage relationships derived from analysis of data generated with the ATP (a) and AMP-PNP (b) models at [Ca2+]i ranging from 100 to 1000 nM Ca2+ as indicated by the different symbols. All solid lines are fits of the simulated sets of data to the Boltzmann equation (Eq. 2) calculated by Origin software. Parameters calculated from such fits are presented in D. Again, note the high similarity between these relationships and those derived from the experimental results described in Fig. 5. (C) Graph showing the relationship between estimated EC50 for Ca2+ and step potential for experimental (squares) and simulated (circles) data obtained with ATP (filled symbols) and AMP-PNP (empty symbols). The two experimental data sets (ATP and AMP-PNP) and the fit (dotted line; AMP-PNP) are reproduced from Fig. 4 for comparison. The solid lines passing through the two sets of simulated data are Boltzmann fits (Eq. 2) described by the following formulas: ATP, EC50 for Ca2+ = {443.07/[1 + exp((V – 118)/44.6)]} + 85.4; AMP-PNP: EC50 for Ca2+ = {686.4/[1 + exp((V – 59.5)/58.8)]} + 74.7, where V is membrane potential. (D) Graph showing the relationship between the half-maximal activation voltage (V0.5) estimated and internal Ca2+ concentration ([Ca2+]i) for experimental (squares) and simulated (circles) data obtained with ATP (filled symbols) and AMP-PNP (empty symbols). The two experimental data sets and associated fits (dashed and dotted lines) are reproduced from Fig. 6 for comparison. The solid lines passing through the two sets of simulated data are single exponential fits (Eq. 3) described by the following formulas: ATP, V0.5 = 260.2 * exp(–[Ca2+]i/398.4) – 11.4; AMP-PNP, V0.5 = 73.5 * exp(–[Ca2+]i/128.8) + 184.5.

	DISCUSSION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Comparison with Other Cell Types
Studies of I_ClCa have identified two forms of the current. In T84 epithelial cells, I_ClCa is time and voltage independent and the currents are activated solely by Ca²⁺ (Xie et al., 1996; Merlin et al., 1998; Xie et al., 1998). In all smooth muscle myocytes examined to date, including the pulmonary artery myocytes studied here, as well as a variety of other cell types, I_ClCa is controlled by both Ca²⁺ and voltage with characteristic activation and deactivation kinetics (for review see Frings et al., 2000; Hartzell et al., 2005; Leblanc et al., 2005). To investigate the mechanism of activation of I_ClCa, we (Greenwood et al., 2001, 2004; Britton et al., 2002; Ledoux et al., 2003) and others (Evans and Marty, 1986; Ishikawa and Cook, 1993; Arreola et al., 1996, Nilius et al., 1997; Qu et al., 2003; Boese et al., 2004) have used intracellular Ca²⁺ chelators such as EGTA or BAPTA to examine the time and voltage dependence of channel activation at constant [Ca²⁺]_i. These studies have revealed qualitative similarities in the kinetics of gating but interesting variability in the Ca²⁺ sensitivity of the currents. The Ca²⁺ dependence of I_ClCa activation in rabbit pulmonary artery myocytes reported here was similar to that reported in other studies. Thus, at +60 mV, the apparent K_d for Ca²⁺ was 400 nM in the present study compared with 285 nM in bovine endothelial cells (Nilius et al., 1997) and 400 nM in medullary collecting duct cells (Qu et al., 2003; Boese et al., 2004). In contrast, activation of I_ClCa in parotid acinar cells required lower [Ca²⁺] (apparent K_d for Ca²⁺ at +70 mV was 60 nM, Arreola et al., 1996), whereas in Xenopus oocytes the Ca²⁺ sensitivity was lower (apparent K_d for Ca²⁺ at +120 mV was 900 nM, Kuruma and Hartzell, 2000). Interestingly, an extensive study of single Ca²⁺-activated Cl^– channel activity in excised patches from rabbit pulmonary artery myocytes derived an EC₅₀ value of 8 nM at +100 mV (Piper and Large, 2003), considerably higher than the Ca²⁺ sensitivity determined in the present study. The reason for this discrepancy is unknown but may reflect the highly regulated nature of these channels (see Leblanc et al., 2005 and below). Overall the channels underlying I_ClCa in an array of cell types, including vascular myocytes, exhibit similar basic gating properties with differing Ca²⁺ sensitivities.

The time-dependent development of current at positive potentials led to a current–voltage relationship that exhibited marked outward rectification although the channel conductance does not rectify inherently. Hence the voltage dependence of the open Cl^– channel is approximately ohmic (Greenwood et al., 2001; Piper and Greenwood, 2003) and the unitary conductance does not change with depolarization (Piper and Large, 2003). Consequently, the outward rectification that is a characteristic of I_ClCa in these cells is a product of a time-dependent increase in channel activity (open probability). In parotid acinar, endothelial cells, and Xenopus oocytes this time-dependent property has been ascribed to an increase in Ca²⁺ sensitivity and a decrease in open to closed transitions at positive potentials (Arreola et al., 1996; Nilius et al., 1997; Kuruma and Hartzell, 2000). The data of the present study reveal that similar properties influence the kinetics of I_ClCa in vascular myocytes.

Working Model
The salient discoveries of our analysis were that generation of I_ClCa is augmented by membrane depolarization but has an obligatory requirement for an increase in [Ca²⁺]. Consequently, membrane hyperpolarization does not turn off I_ClCa when the pipette [Ca²⁺] is set to levels greater than the activation threshold (between 100 and 250 nM). This property was also observed for I_ClCa recorded from parotid acinar cells and Xenopus oocytes (Arreola et al., 1996; Kuruma and Hartzell, 2000) but differs from that of Ca²⁺-dependent K⁺ channels that can be opened by strong membrane depolarization in the absence of internal Ca²⁺ (Cui et al., 1997).

To gain better insight into the mechanism by which phosphorylation affects the gating of the Cl_Ca channel in pulmonary myocytes, we took advantage of computer modeling techniques and the existence of already published kinetic models of Cl_Ca in other cell types that well reproduced their macroscopic properties in pancreatic acinar cells (Arreola et al., 1996) and Xenopus oocytes (Kuruma and Hartzell, 2000). In the former model, there are two consecutive voltage-dependent Ca²⁺ binding steps, and binding of both Ca²⁺ ions is required for channel opening. Similar to Arreola et al. (1996), our data showed that the K_d for Cl_Ca channel activation by intracellular Ca²⁺ declined with membrane depolarization. However, the Hill coefficient of Cl_Ca channels increased from 1 at negative potentials to 2 at positive potentials, whereas Cl_Ca channels in pulmonary myocytes (Fig. 5 B) required more than two Ca²⁺ ions (Hill coefficient of 2–4) for channel activation; moreover, this parameter declined rather than increased from 3 at negative potentials to 2 at potentials >0 mV, an observation similar to I_ClCa in Xenopus oocytes (Kuruma and Hartzell, 2000). All attempts to simulate our data with such a model failed to reproduce the Ca²⁺ and voltage dependence of I_ClCa in PA myocytes and their kinetics of activation and deactivation with ATP or AMP-PNP, although certain properties of the macroscopic current could be modeled reasonably well at any given concentration of Ca²⁺.

We therefore investigated the more complex model developed by Kuruma and Hartzell (2000), which described well the properties of I_ClCa in Xenopus oocytes and adapted it to simulate the gating of Ca²⁺-activated Cl^– channels in pulmonary artery myocytes. Besides the fact that their model offered much more flexibility, the characteristics of I_ClCa recorded from Xenopus oocytes correlated better with the properties of "dephosphorylated" Cl_Ca channels (AMP-PNP) in our preparation: (a) similar voltage dependence of the K_d for Ca²⁺ and Hill coefficient, (b) similar Ca²⁺ dependence of V_0.5, (c) lack of voltage dependence and similar Ca²⁺ dependence of _act at [Ca²⁺] > 200 nM, and (d) similar voltage and Ca²⁺ dependence of _deact. As explained in Results, activation of the channel involves the consecutive binding of three Ca²⁺, all with identical affinities, but in contrast to the model of Arreola et al. (1996), Ca²⁺ binding per se is voltage independent. Each of the Ca²⁺-bound closed states can transit in the open state with progressively faster opening rates as the channel binds more calcium ions. In this kinetic scheme, voltage-dependent gating is due to the channel closing rate. Figs. 8 and 9 showed that an adapted version of the Kuruma and Hartzell model effectively simulated the experimentally derived data recorded with AMP-PNP–containing pipette solutions (compare Fig. 8 with Fig. 3). It is interesting to note that in the experiments of both Arreola et al. (1996) and Kuruma and Hartzell (2000), the solution facing the internal side of the membrane did not contain ATP, a situation that would minimize the state of phosphorylation of the channels and mimic our AMP-PNP experiments. Although it is unknown whether Cl_Ca channels in these cells are regulated by phosphorylation in a similar fashion, it has been reported that I_ClCa in Xenopus oocytes is inactivated by activation of protein kinase C in a Ca²⁺-dependent manner (Boton et al., 1990). Overall, the model parameters used to simulate I_ClCa recorded from cells dialyzed with AMP-PNP (see Table I) accounted well for the "apparent" sigmoidal increase in Ca²⁺ affinity observed with membrane depolarization, the voltage dependence of fully activated I_ClCa, the basal activation of the underlying channels observed at negative potentials, the Ca²⁺ dependence of V_0.5 and kinetics of activation, and voltage dependence of deactivation.

To model the data with ATP, we simply increased the magnitude of the "off" (ß_x in Table I) rate constants and shifted their voltage dependence toward more positive potentials, assigning a value of 0 to the gating variable for the higher Ca²⁺-bound states, which means that the channels are either closed or blocked by phosphorylation. By analogy, this would correspond to open state channel block by phosphorylation. All other rate constants, including those defining Ca²⁺ affinity, were identical to those used in the AMP-PNP model (see Table I). Our simulations reproduced quantitatively in most cases, and semi-quantitatively in others, the macroscopic behavior of I_ClCa that includes potent inhibition of the Cl^– channels within the physiological range of membrane potentials, their Ca²⁺ dependence at all potentials examined, and to a lesser extent their Ca²⁺ and voltage dependence of activation (except at 250 nM) and deactivation kinetics. Although this was not evaluated, it might have been possible to produce similar results by reducing the gating variable of all open states to a fraction between 0 and 1. This would be similar to the induction of subconductance states by phosphorylation. Indeed, higher levels of [Ca²⁺] were shown to induce the appearance of subconductance levels of single Cl_Ca channels in rabbit pulmonary myocytes (Piper and Large, 2003), and perhaps phosphorylation of a cytoplasmic domain may cause open state block by an electrostatic interaction with acidic amino acid residues residing within or near the channel pore. By opposition, minimizing phosphorylation with AMP-PNP may "lock" the channels open at negative potentials, especially at higher levels of intracellular Ca²⁺, resulting in the appearance of very slowly declining tail currents (see Figs. 3 and 9) and large negative shifts in the holding current, leading to attenuation of the normally strong outward rectification. Although the model is undoubtedly overly simplistic, it nevertheless provides a valid framework from which specific hypotheses in regards to the regulation of Cl_Ca channels by phosphorylation can be thoroughly tested under conditions allowing physiological intracellular Ca²⁺ dynamics. Future single channel experiments will be designed to test the hypothesis that the state of phosphorylation of the channel and/or regulatory subunit(s) alters the properties determining the rate of closure of the gate and perhaps the state of permeation of the channel.

Regulation of I_ClCa in Pulmonary Artery Myocytes
The data of the present study agree with our past studies (Greenwood et al., 2001, 2004; Ledoux et al., 2003) that the activity of Ca²⁺-activated Cl^– channels in vascular smooth muscle cells is dictated by the phosphorylation status in the vicinity of the channel. Regulation of I_ClCa has been best studied in arterial myocytes where we have established that CaMKII suppresses activity (Greenwood et al., 2001) whereas calcineurin enhances I_ClCa (Ledoux et al., 2003; Greenwood et al., 2004). Previously, we showed that blockade of calcineurin in coronary artery myocytes produced a decrease in the Ca²⁺ sensitivity. However, the present study established that gross dephosphorylation markedly enhanced I_ClCa due to an increase in the voltage sensitivity and not the Ca²⁺ sensitivity. While there is the caveat that the studies were not performed in the same cell type, these data suggest that a hyperphosphorylated Cl^– channel is less able to bind Ca²⁺ or is less able to transit to an open configuration upon Ca²⁺ binding, whereas a fully dephosphorylated channel is more active at less depolarized potentials. The present study also reveals that Cl^– channel activity in rabbit pulmonary artery myocytes is highly labile and at the mercy of the opposing kinase/phosphatase activity. Thus, the amplitude of I_ClCa evoked immediately after membrane rupture was large but decreased to a stabilized level 20% of the initial value when the pipette solution was ATP rich. Under these conditions, phosphorylation would be supported, and therefore the current recorded at the stabilized level represents diminished ion flux through a partially phosphorylated channel. The degree of phosphorylation is dynamic and dependent on the kinase/phosphatase balance in the vicinity as described by Greenwood et al. (2004). This supposition is reinforced by the data with AMP-PNP that is unable to support phosphorylation (Yount, 1975; Gadsby and Nairn, 1999) and therefore allows phosphatase activity to dominate. Consequently, the degree of rundown is attenuated and the current stabilizes to a steady-state level close to the initial current amplitude. Whether calcineurin alone or in combination with other phosphatases drives this recovery of I_ClCa amplitude is the focus of future experiments.

In conclusion, the present study represents an extensive study of the kinetics, Ca²⁺ dependence, and voltage dependence of whole-cell I_ClCa in pulmonary artery myocytes. A corollary to this work is that we have been able to effectively simulate the experimental data using a minimal reaction scheme. Moreover, this exhaustive study has revealed a mechanism by which phosphorylation regulates Cl^– channel activity and has reinforced the crucial role of phosphorylation/dephosphorylation mechanisms on I_ClCa.

	ACKNOWLEDGMENTS

 
The authors would like to thank Cathy Lachendro for her technical help in isolating rabbit pulmonary artery smooth muscle cells and for preparing solutions.

The work was supported by grants to N. Leblanc from the National Institutes of Health (grant HL 1 R01 HL075477-01A2). Research in the laboratory of I.A. Greenwood was funded by the British Heart Foundation and The Wellcome Trust. This publication was also made possible by grant NCRR 5P20 RR15581 (N. Leblanc) from the National Center for Research Resources (NCRR), a component of the National Institutes of Health (NIH) supporting a Center of Biomedical Research Excellence at the University of Nevada School of Medicine. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of NCRR or NIH.

Olaf S. Andersen served as editor.

Submitted: 3 February 2006
Accepted: 1 June 2006

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