A Close Association of RyRs with Highly Dense Clusters of Ca²⁺-activated Cl⁻ Channels Underlies the Activation of STICs by Ca²⁺ Sparks in Mouse Airway Smooth Muscle

In a variety of cells, the opening of one or a few ryanodine receptors (RyRs) in the membrane of endo/sarcoplasmic reticulum (ER/SR) produces highly localized, short-lived Ca²⁺ transients, designated as Ca²⁺ sparks or Ca²⁺ syntillas (Cheng et al., 1993; Nelson et al., 1995; Tsugorka et al., 1995; ZhuGe et al., 1998; DeCrescenzo et al., 2004; ZhuGe et al., 2006). These Ca²⁺ events comprise the elementary Ca²⁺ signals that underlie excitation–contraction coupling via either a calcium-induced calcium release mechanism in cardiac muscle or a mechanical coupling in skeletal muscle (Cannell et al., 1995; Lopez-Lopez et al., 1995; Klein et al., 1996). Ca²⁺ sparks in smooth muscle, however, may not serve as the “building block” of Ca²⁺ transient for excitation–contraction coupling (Nelson et al., 1995); rather they exert their influence by regulating the activity of Ca²⁺-activated ion channels in the plasma membrane. We and others have demonstrated that Ca²⁺ sparks activate nearby large-conductance Ca²⁺-activated K⁺ channels (BK channels) to generate spontaneous transient outward currents (STOCs) in virtually all smooth muscle (Nelson et al., 1995; Bolton and Imaizumi, 1996; ZhuGe et al., 1998; Jaggar et al., 2000). The functional coupling between Ca²⁺ sparks and STOCs is critical for the regulation of smooth muscle contractility. When the BK channel β1 subunit, a subunit that increases the sensitivity of the pore-forming α subunit to Ca²⁺, is genetically ablated or down-regulated, Ca²⁺ sparks less efficiently activate STOCs, resulting in the development of hypertension, urinary incontinence, and diabetic retinopathy in mice (Brenner et al., 2000; Petkov et al., 2001; Amberg and Santana, 2003; McGahon et al., 2007). A key cellular mechanism underlying such profound effects is the close proximity between RyRs and BK channels to form microdomains, where the activity of BK channels is tightly controlled by Ca²⁺ sparks (Perez et al., 2001; ZhuGe et al., 2002).

In addition to causing STOCs, Ca²⁺ sparks turn on Ca²⁺-activated Cl⁻ (Cl_(Ca)) channels to produce spontaneous transient inward currents (STICs) in many types of smooth muscle (ZhuGe et al., 1998; Gordienko et al., 1999; Williams and Sims, 2007). In these cells, activation of STICs lead to membrane depolarization (Williams and Sims, 2007), an opposite response to the activation of STOCs. Therefore it is physiologically important to understand the cellular mechanism determining the coupling between Ca²⁺ sparks and STICs. However, little information is available as to how Ca²⁺ sparks activate STICs. A key question to be resolved is the spatial relationship between RyRs and Cl_(Ca) channels that controls their functional coupling. There are several reasons that the relationship between RyRs and Cl_(Ca) channels might be different from that between RyRs and BK channels. For instance, Cl_(Ca) channels are thought to be much more sensitive to Ca²⁺ than BK channels (Singer and Walsh, 1987; Pacaud et al., 1992; Tanaka et al., 1997; Piper and Large, 2003) so they could localize far away from RyRs and still be activated by Ca²⁺ sparks. Also, in rat portal vein, Ca²⁺ sparks activated BK channels to produce STOCs but they failed to activate Cl_(Ca) channels and STICs (Mironneau et al., 1996). Finally, at resting membrane potential Ca²⁺ sparks simultaneously activate BK channels and Cl_(Ca) channels, resulting in biphasic currents with an STOC component always preceding an STIC component (ZhuGe et al., 1998). In light of these differences, it is essential to examine experimentally the spatial relationship between RyRs and Cl_(Ca) channels. An attractive approach to exploring this relationship would be to visualize both RyRs and Cl_(Ca) channels in the cells with immunolight and immunoelectron microscopy. Unfortunately, the gene(s) encoding Cl_(Ca) channels are yet to be discovered, so it is currently impossible to localize these channels in these cells.

Here, we have used a variety of analyses of electrophysiological and imaging data to investigate the organization of RyRs and Cl_(Ca) channels underlying the functional coupling of Ca²⁺ sparks and STICs in mouse airway smooth muscle cells. First, Ca²⁺ sparks were recorded in isolated single cells with a wide-field imaging microscope at high speed and, at the same time, STICs were monitored using whole-cell patch clamp. A signal mass approach (ZhuGe et al., 2000) was employed to quantify the total amount of Ca²⁺ release from Ca²⁺ sparks (via RyRs) and the underlying Ca²⁺ currents (I_Ca(spark)), allowing us to analyze quantitatively the relationship between Ca²⁺ sparks and STICs. Second, we characterized Cl_(Ca) channels using flash photolysis of caged Ca²⁺. To our surprise, Cl_(Ca) channels are highly insensitive to Ca²⁺ in these cells compared with the reports in other types of smooth muscle. Third, we devised an approach to estimating the conductance underlying STICs (g_(STIC)) and its relationship with voltage; with this relationship we estimated the [Ca²⁺] “seen” by Cl_(Ca) channels underlying STICs. Fourth, we performed reaction-diffusion simulations to derive the spatio-temporal profile of [Ca²⁺] from Ca²⁺ sparks with measured I_Ca(spark). Combining this profile with the estimate of [Ca²⁺] sensed by Cl_(Ca) channels, we determined the area occupied by these channels near the spark sites to generate an STIC. Finally, we estimated that the Cl_(Ca) channels residing in sparks sites could account for all the channels present in the entire cell; that is, Cl_(Ca) channels could be exclusively near spark sites.

With these studies, we found that in Ca²⁺ spark sites, RyRs and Cl_(Ca) channels localize near to each other, and Cl_(Ca) channels concentrate in an area with a radius of ∼600 nm, where their density reaches as high as 300 channels/μm². These findings reveal that Cl_(Ca) channels are tightly controlled by Ca²⁺ from nearby RyRs, uncovering a new property of Ca²⁺ microdomains in smooth muscle.

Male Swiss Webster mice, of age 6–9 wk, were killed with intraperitoneal injection of a lethal dose of sodium pentobarbital (50 mg kg⁻¹), in accordance with the guidelines of the Animal Care Committee of the University of Massachusetts Medical School. After each animal was unresponsive to any applied stimulus, trachea was quickly removed and placed in prechilled dissociation solution, consisting of (in mM) 136 NaCl, 5.36 KCl, 0.44 KH₂PO₄, 4.16 NaHCO₃, 0.34 Na₂HPO₄, 5 mM MgCl₂, 20 mM HEPES, and 10 glucose (pH 7.1). The trachea was dissected free from the surface of connective tissues and incubated in the dissociation medium without MgCl₂ but plus papain 30 U/ml (Sigma-Aldrich), 0.2 mM DTT, and 0.02 mM EDTA at room temperature for 30 min. The tissue was then incubated at 32°C for another 6 min with fresh zero MgCl₂ dissociation medium containing 3 U/ml collagenase 1A (Sigma-Aldrich), 0.2 mg/ml proteinase E (Sigma-Aldrich), 0.1 mg/ml DNAase 1 (Willington), and 1 mg/ml amino acid-free BSA (Sigma-Aldrich). Finally, the tissue was agitated with a fire polished wide-bore glass pipette to release the cells. The isolated single airway smooth muscle cells were spindle-shaped with a length of around 150 μm, and used the same day at 25°C.

Membrane currents were recorded using the tight-seal, conventional whole-cell configuration. The extracellular solution contained (in mM) 130 NaCl, 5.5 KCl, 2.2 CaCl₂, 1 MgCl₂, 10 HEPES, pH adjusted to 7.4 with NaOH. The pipette solution contained (in mM) 139 KCl, 1 MgCl₂, 3 Na₂ATP, 10 HEPES, and 0.05 fluo-3 K₅ (Invitrogen), pH to 7.3 with KOH. For experiments in Fig. 5, the bath solution contained (in mM) 130 NaCl, 5.5 TEA-Cl, 2.2 CaCl₂, 1 MgCl₂, 10 HEPES, pH adjusted to 7.4 with NaOH; and the pipette solution contained (in mM) 75 CsCl, 64 Cs-glutamate, 1 MgCl₂, 10 HEPES, 3 Na₂ATP, and 0.05 fluo-3 K₅ (pH adjusted to 7.3 with CsOH). Whole-cell currents were recorded at desired holding potentials and low-pass filtered using the Axopatch 1D amplifier (200 Hz cutoff) and then digitally sampled at 1 kHz and stored for analysis. Events were counted as STICs if their amplitude exceeded twofold of RMS of the baseline, and their area under curve was >10 fold of the value of RMS as detected by Mini Analysis Program (Snaptosoft). The events were then checked by visual inspection to eliminate anomalies such as multiple events overlapping in time or excessively noisy traces.

Fluorescent images were obtained using fluo-3 (Invitrogen) as the calcium indicator and a custom-built wide-field, high-speed digital imaging system, which is described in detail elsewhere (ZhuGe et al., 1999). Rapid imaging was made possible by using a cooled high-sensitivity, charge-coupled device camera (128 × 128 pixels) developed in conjunction with MIT Lincoln Laboratory. The camera was interfaced to a custom-made, inverted microscope equipped with a 40× oil immersion lens (NA 1.3); each pixel covered a 333 nm × 333 nm area of the cell. The 488-nm line of a multiline argon laser provided fluorescence excitation for the indicator fluo-3, and a laser shutter controlled the exposure duration. Emission of the Ca²⁺ indicator was monitored at wavelengths >500 nm. To obtain a constant concentration of Ca²⁺ indicator, fluo-3 (50 μM) was delivered through the patch pipette, and measurements were not commenced until 10–15 min after disruption of the patch. After this time no significant change in background fluorescence was detected. Subsequent image processing and analysis were performed off-line using a custom-designed software package, running on a Linux workstation.

Two measures of Ca²⁺ sparks were employed: the conventional fluorescence ratio, ΔF/F₀, within a restricted area; and the change in total fluorescence, F − F_o, over a larger volume, also designated as the Ca²⁺ signal mass, which is proportional to the total quantity of Ca²⁺ released into the cytosol. As demonstrated previously (ZhuGe et al., 2000), ΔF/F₀ is a poor indicator of local [Ca²⁺]_cyto generated by Ca²⁺ sparks. In the present study, this measure was used mainly for identifying events. Both conventional fluorescence ratio and Ca²⁺ signal mass measurements have been previously described in detail (ZhuGe et al., 2000); a brief description follows. For the fluorescence ratio measure, the fluo-3 images, with pixel size 333 × 333 nm, were first smoothed by convolution with a 3 × 3 pixel approximation to a two-dimensional Gaussian (σ = 1 pixel). Fluorescence ratios were then calculated and expressed as a percentage on a pixel to pixel basis from the equation:

where F is the fluorescence at each pixel in the time series and F₀ is the “resting” level derived from the fluorescence time series by computing the median pixel value during quiescent times at each F(x,y). During a spark, the single pixel that had the highest fluorescence ratio is designated as the epicenter pixel (Fig. 8). The second measure, the Ca²⁺ signal mass, for each spark was computed from the two-dimensional, wide-field fluorescence images of fluo-3 according to the equations:

The signal mass (sm(t)) is the product of the detector gain (G) times the change in total fluorescence (F^T) summed over a square region of 40 × 40 pixels, surrounding the spark epicenter pixel (x,y) as determined from the ΔF/F₀ images from Eq. 1. The number of moles of Ca²⁺ bound to fluo-3 was calculated by the equation: 2.44 − sm(t)/(6.022 − 10²³) moles, where 2.44 is a calibration factor previously determined (ZhuGe et al., 2000). Since there may be other Ca²⁺ buffers in the patched and dialyzed cells, Ca²⁺-bound fluo-3 provides a minimum value for the total Ca²⁺ released. To correct for the contribution of endogenous Ca²⁺ buffers to the signal mass, we characterized these buffers using flash photolysis of caged Ca²⁺ as detailed in the following section.

A 100-ms exposure of UV light (λ = 351 nm) from an argon ion ultraviolet laser was coupled to the epi-illumination port of our high-speed inverted microscope, and focused on the cell through a Nikon 40× UV, 1.3 NA, oil immersion lens. The UV beam was passed through an optical path such that it was restricted to illuminate a circle 160 μm in diameter on the cell. The cell under study was located in the center of this illumination area so that the entire cell was illuminated, resulting in an elevation in [Ca²⁺] that sustained for several seconds after uncaging.

NPEGTA (o-nitrophenyl EGTA) was dialyzed into cells via the patch pipette so that its concentration and resultant [Ca²⁺] upon photolysis could be determined. In the patch pipette, 50 μM ryanodine was also included to minimize the involvement of calcium-induced calcium release upon an increase in [Ca²⁺] after photolysis. In our preliminary study, we varied the laser settings to determine the photon density that was sufficient to uncage all the NPEGTA present in the cells. We found that a (cell) specimen illumination of ∼400 w/cm² for 100 ms was sufficient for this purpose, since a second exposure within 1 s of the first, at the same power and duration, failed to increase [Ca²⁺]_cyto and I_Cl(ca). This level of photon density does not produce measurable phototoxicity since when cytosolic NPEGTA was absent or <10 μM, photolysis did not trigger I_Cl(ca) (see Fig. 4). Thus we employed this laser setting for all the flash photolysis experiments in the present study.

To estimate endogenous fixed Ca²⁺ buffers (mobile buffers are expected to be dialyzed out the cells under whole-cell configuration and thus are not considered in the present study), we first measured [Ca²⁺]_cyto with fluo-3 upon flash photolysis of caged NPEGTA by an equation similar to that derived by Maravall et al. (2000):

where Kd_Fluo3 is taken as 1.1 μM (Harkins et al., 1993), R equals to the fluorescence ratio before and after photolysis of various [NPEGTA], and R_max is the ratio when [NPEGTA] is 833 μM. In the example shown in Fig. 1 A, R_max was 12 and on average this value was 11.58 ± 1.55 (n = 5). These values are close to the theoretical value of R_max for a resting [Ca²⁺]_cyto of 104 ± 9 nM (n = 6 cells) as measured with fura-2 using a microfluorimeter (ZhuGe et al., 1999), suggesting that photolysis of NPEGTA at this level produces a saturated [Ca²⁺] for fluo-3.

Next, we compared the relationship between measured [Ca²⁺]_cyto and total Ca²⁺ released upon flash photolysis of various [NPEGTA], i.e., [Ca²⁺]_ΔT (Fig. 1 B). To do so, we first examined whether NPEGTA affects resting [Ca²⁺]_cyto and then determined the fraction of Ca²⁺-bound NPEGTA. For the cells shown in Fig. 1 A, when [NPEGTA] was preset in the range of 0–200 μM, averaged fluorescence per pixel was constant, fitted linearly with a slope of −0.018, which was not different from 0 (P = 0.728). At 833 μM, the averaged fluorescence per pixel decreased to 70% of the control value. Using the microfluorimeter with fura-2, we estimated that [Ca²⁺]_cyto was 108 ± 11 (n = 6) and 84 ± 9 nM (n = 5) in the presence of 200 μM and 833 μM NPEGTA, respectively. (One possible caveat for examining the effect of NPEGTA on [Ca²⁺]_cyto is that NPEGTA in the cytosol could be slowly photolyzed by the UV light from Arc lamp in the microfluorimeter. However, this possibility was ruled out in our conditions by the observations that (a) [Ca²⁺]_cyto in the presence of NPEGTA was as stable as when NPEGTA was absent, and (b) [Ca²⁺]_cyto was the same when measured 1 min apart in the presence of NPEGTA.) Therefore, two lines of data indicate that up to 200 μM, NPEGTA does not affect resting [Ca²⁺]_cyto, and at 833 μM it modestly decreases the [Ca²⁺]_cyto. Given that (a) NPEGTA has a K_d of 80 nM in its caged form, and (b) all of the loaded NPEGTA is photolyzed, [Ca²⁺]_ΔT is expected to be ∼57% of [NPEGTA] loaded in the patch pipette when the concentration is <200 μM, and 52% when it is at 833 μM.

Assuming a simple one-compartment model of cellular [Ca²⁺] dynamics, [Ca²⁺]_ΔT can be described according to an equation:

where [Fluo3]_T = 50 μM, Kd_Fluo3 is 1.1 μM, [Ca²⁺]_T0 the total cytosolic [Ca²⁺] before photolysis, [B]_T the concentration of endogenous fixed buffer, and Kd_B the equilibrium dissociation constant for this buffer. Fitting the data (open circles in Fig. 1 B) with Eq. 5 using a Levenberg-Marguardt nonlinear least-squares optimization method yielded [B]_T = 81 μM, Kd_B = 0.66 μM, and [Ca²⁺]_T0 = 15.6 μM (χ² = 11.715; r = 0.9967). With these values, we estimate the binding ratio (κ) of this buffer to be 92, using

(Zhou and Neher, 1993), with [Ca²⁺] as 104 nM. For the same calculation, κ for 50 μM fluo-3 is 38. Therefore, the values of signal mass and resulting I_Ca(spark) as estimated by fluo-3 fluorescence were adjusted with a correction factor of 3.45, i.e., (1 + 92 + 38)/38, to reflect I_Ca(spark) and the total amount of Ca²⁺ released during Ca²⁺ sparks under physiological conditions in these cells.

Given these parameters on the endogenous fixed Ca²⁺ buffer, we can further estimate [Ca²⁺]_cyto upon photolysis of NPEGTA in the absence of fluo-3 (Fig. 1 C). These estimates were used to set [Ca²⁺]_cyto when Ca²⁺ sensitivity of Cl_(Ca) channels was probed (Fig. 4).

To gain insight into the role of Ca²⁺ sparks triggering STICs, we modeled a simplified cell, simulated Ca²⁺ sparks, and examined the spatial and temporal profile of free [Ca²⁺] that resulted at the plasma membrane where Cl_(Ca) channels are located. Finite difference approximations were used to solve a set of partial differential equations for the reaction-diffusion kinetics in a cylindrically symmetric coordinate system. The details of this approach were described elsewhere (ZhuGe et al., 2002), and it has been used before to analyze the spatial-temporal profile of [Ca²⁺] resulting from Ca²⁺ sparks in amphibian gastric smooth muscle, chromaffin cells, and nerve terminals (ZhuGe et al., 2002; DeCrescenzo et al., 2004; ZhuGe et al., 2006). The cell was modeled as a cylinder, 5 μm in diameter and 5 μm in length. The Ca²⁺ release site was modeled as a small cylinder, 20 nm in diameter and height, with the cylinder end located 20 nm from the plasma membrane (Somlyo and Franzini-Armstrong, 1985). The simulation included the following: fluo-3 as the sole mobile buffer (50 μM, K_d = 1.1 μM, k_on = 8 × 10⁴ mM⁻¹s⁻¹, k_off = 90 s⁻¹, D = 25 μm²s⁻¹); Nondiffusible fixed buffer at 81 μM (K_d = 0.66 μM, k_on = 8 × 10⁴ mM⁻¹s⁻¹, k_off = 52.8 s⁻¹). Endogenous mobile buffers are expected to be washed out via the patch pipette and thus not included in the simulations.

Results are presented as mean ± SEM, unless otherwise indicated. N denotes the number of cells and n the number of events, i.e., Ca²⁺ spark or STIC. Student's t tests and ANOVA, when appropriate, were used to compare the means from different conditions.

The online supplemental material is available. Fig. S1 shows reaction-diffusion simulations, demonstrating that the endogenous fixed Ca²⁺ buffer measured with the flash photolysis of NPEGTA exerts a minimal effect on both the rise and decay phase of estimated I_Ca(spark). Fig. S2 examines the correlation between STICs and Ca²⁺ sparks using ΔF/F₀ in the epicenter pixel, i.e., the conventional fluorescence ratio.

Knowledge of the spatio-temporal profile of [Ca²⁺] generated by the opening of RyRs during a Ca²⁺ spark is critical to an understanding of the spatial relationship between RyRs underlying Ca²⁺ sparks and Cl_(Ca) channels for STICs. But it has not been possible previously to directly measure this profile for at least two reasons. First, Ca²⁺ and Ca²⁺ indicator (fluo-3) do not necessarily come into equilibrium in a microdomain where there could exist a huge Ca²⁺ gradient (Stern, 1992; Naraghi and Neher, 1997). And second, the fluorescence signal from an area of even a single pixel is an average of an underlying gradient of [Ca²⁺] in the microdomain that could be below the spatial resolution of light microscope (ZhuGe et al., 2000). To circumvent these difficulties, in the present study we employed a strategy that we had developed to estimate the [Ca²⁺] profile produced by Ca²⁺ sparks in amphibian smooth muscle and by Ca²⁺ syntillas in nerve terminals and chromaffin cells (ZhuGe et al., 2002, 2006; DeCrescenzo et al., 2004). This strategy begins by estimating I_Ca(spark), i.e., Ca²⁺ current through RyRs during a Ca²⁺ spark. This is made possible with the signal mass approach that can directly measure the total quantity of Ca²⁺ released during a Ca²⁺ spark with high-speed wide-field imaging. At its simplest, signal mass can be viewed as a product of the amplitude and duration of I_Ca(spark) with a proportionality factor. Thus, time differentiation of the signal mass gives rise to the time course of I_Ca(spark). This time course can then be used as a Ca²⁺ source in a reaction-diffusion simulation to deduce the spatio-temporal profile of [Ca²⁺] induced by Ca²⁺ sparks.

To reveal the amplitude and waveform of I_Ca(spark), we imaged Ca²⁺ sparks at a speed of 333 Hz with exposure of 3 ms. Fig. 2 displays an example of Ca²⁺ sparks recorded at this speed. The images in A exhibit the spatial and temporal evolution of the Ca²⁺ spark, and the time course of change in fluorescence ratio from the epicenter pixel is shown in the bottom panel. The total quantity of Ca²⁺ released from this spark and the underlying I_Ca(spark) are shown in the top and middle panels of Fig. 2 B, respectively. For this spark, the value of signal mass was 286,000 Ca²⁺ ions after compensating for the estimated endogenous fixed Ca²⁺ buffer (see Materials and methods). Interestingly, the rising phase of signal mass consisted of two components, a fast one followed by a slow one. These two components corresponded to the two phases in I_Ca(spark), i.e., a rising phase that can be roughly fitted with a linear function with 9 ms from the onset to the peak, and a decay phase that can be fitted with an exponential function with a tau of 13 ms. The duration of I_Ca(spark), i.e., from the onset of an event to the point when the current decays ∼92% (equal to 2.5 times of the decay constant), was ∼39 ms and the peak amplitude of this current was ∼6.4 pA.

On average (Table I), it was found that (a) a Ca²⁺ spark releases 276,000 ± 46,000 Ca²⁺ ions, (b) I_Ca(spark) has a peak amplitude of 8.5 ± 1.2 pA, and (c) this current reaches peak within 9 ± 2 ms and decays with a single exponential function with a tau of 12 ms ± 2 ms. These values were calculated including the effect of the estimated endogenous fixed Ca²⁺ buffer in this type of cell. We should point out that these sparks were for high signal to noise characteristics, and may not reflect the complete distribution of events (see below). Also, since the kinetics of binding of the endogenous fixed Ca²⁺ buffer are not yet determined, their effect on the observed rise and decay of I_Ca(spark) cannot be directly incorporated. However, even if we assume that the buffer has the same diffusion-limited on-rate as fluo-3 (i.e., it is a fast buffer), it has minimal influence on either the rise or the decay phase of the estimate of I_Ca(spark) (see Fig. S1). Therefore, the measured waveform of I_Ca(spark) most likely reflects the underlying Ca²⁺ current from the opening of RyRs during Ca²⁺ sparks under physiological conditions in these cells.

Fig. 2 also demonstrates a close temporal relationship between Ca²⁺ sparks and STICs. The onset of this Ca²⁺ spark coincided with the STIC. On average there is a delay between spark and STIC of ∼3 ms, which is consistent with the notion that Ca²⁺ sparks are the causal signals of STICs (ZhuGe et al., 1998). This notion is reinforced by the following observations. (a) Ryanodine (100 μM), an inhibitor of ryanodine receptor, blocked both Ca²⁺ sparks and STICs (unpublished data); and (b) niflumic acid (100 μM), a Cl_(Ca) channel blocker, abolished STICs but did not affect the frequency and amplitude of Ca²⁺ sparks (unpublished data). The blockage of STICs by niflumic acid also indicates that these currents are carried by Cl⁻ flux passing through Cl_(Ca) channels. (The reversal of STIC polarity around E_Cl is another indication of this conclusion, see Fig. 5 A.) Taken together, these results demonstrate that Ca²⁺ sparks, resulting from the opening of RyRs, activate Cl_(Ca) channels to generate STICs in airway smooth muscle from mouse.

High speed imaging of Ca²⁺ sparks and analysis of the resulting signal mass allows us to quantitatively examine the relationship between Ca²⁺ sparks and their corresponding STICs, which, in turn, could give insight into the molecular organization of underlying RyRs and Cl_(Ca) channels in the spark sites. As shown in the example of Fig. 2, I,_Ca(spark) increased over time, as did the corresponding STIC. But I_Ca(spark) reached its peak before the STIC. As I_Ca(spark) declined, the STIC kept rising until I_Ca(spark) returned to the baseline; at that point the STIC entered its decay phase. From this relationship, it is clear that the rise time of an STIC, i.e., from the onset of the current to the onset of the current decay, correlates with the duration of I_Ca(spark), which reflects the open time of RyRs underlying Ca²⁺ sparks. On average, the duration of I_Ca(spark) was 44 ± 5 ms, which is the same as the rise time of corresponding STICs (42 ± 4 ms) (Fig. 3 A, a; r = 0.879, P < 0.0001, n = 23). (In comparison, the rise time of ΔF/F₀ [24 ± 2 ms] is faster than that of the corresponding STICs [Table I; Fig. S2].) On the other hand, the amplitude of I_Ca(spark) bears no correlation with the rise time of the STICs (Fig. 3 Ab; r = 0.316, P > 0.14), and neither does Ca²⁺ spark amplitude as estimated by ΔF/F₀ (Table I; Fig. S2). The close match in the rise time of the STIC and the duration of the I_Ca(spark) suggests that Cl_(Ca) channels for STICs may localize closely to RyRs so that activation of Cl_(Ca) channels is maintained as long as the RyRs are open; alternatively, this relationship implies that Cl_(Ca) channels could occupy a sizable area, spreading around RyRs so the longer the RyRs stay open, the more of them are recruited and activated by diffusing of Ca²⁺.

To distinguish between these two possibilities, we further analyze the relationship between Ca²⁺ sparks and their corresponding STICs by examining the correlations between peak signal mass and peak STIC amplitude (Fig. 3 B), and between peak I_Ca(spark) and peak STIC amplitude (Fig. 3 C). Two features standout in these plots. First of all, the correlation for both relationships was weak (n = 159, r = 0.056, and P = 0.487 for signal mass vs. STIC; r = −0.112 and P = 0.093 for I_Ca(spark) vs. STIC). It is obvious that large STICs can be associated with small I_Ca(spark), and vice versa, i.e., small STICs can be activated by Ca²⁺ sparks with large I_Ca(spark). Such independence of STIC amplitude on spark amplitude provides an indication that Cl_(Ca) channels and RyRs may organize in such a way that the amplitude of STIC is not proportional to the area exposed to high [Ca²⁺], but is controlled by the numbers of the channels nearby the sites. This view is further strengthened by a second feature in Fig. 3 (B and C), i.e., there were 20% of Ca²⁺ sparks that did not activate STICs, i.e., STIC-less sparks. This class of sparks could be due to underlying RyRs localized deep inside of the cells so [Ca²⁺] generated is not high enough to activate Cl_(Ca) channels on the plasma membrane. But many STIC-less sparks were observed occurring close to the plasma membrane (unpublished data). Therefore, a more plausible explanation for STIC-less sparks is that there are no Cl_(Ca) channels present at these spark sites.

Collectively, the analyses of the relationship between Ca²⁺ sparks and STICs suggest that Cl_(Ca) channels distribute nonuniformly on the surface membrane, and in the areas they are present, they could localize closely to RyRs where Ca²⁺ sparks occur. In the following sections, we further examine this relationship and do so in a quantitative manner.

The Ca²⁺ sensitivity of Cl_(Ca) channels is one of the principal properties influencing how Cl_(Ca) channels respond to change in [Ca²⁺] produced by Ca²⁺ sparks, thus it is a key factor needed to uncover the spatial organization of Cl_(Ca) channels and RyRs in spark sites. Since this is the first study of Cl_(Ca) channels in airway smooth muscle from mouse, we needed to determine this property experimentally. To do so, we studied I_Cl(Ca) upon instantaneous and uniform elevation of [Ca²⁺] by flash photolysis of caged Ca²⁺ NPEGTA in these cells. With the experiments and the procedure described in Fig. 1 and in Materials and methods, we established the relationship between [Ca²⁺]_cyto and [NPEGTA] upon flash photolysis (Fig. 1 C). In our experimental conditions, when flash photolysis of NPEGTA raised [Ca²⁺]_cyto to <120 nM, it failed to evoke any appreciable I_Cl(Ca) (Fig. 4 A), indicating that Cl_(Ca) channels have a threshold >120 nM. At 0.2 μM [Ca²⁺], it started to trigger appreciable I_Cl(Ca) with some delay; further increases in [Ca²⁺] caused I_Cl(Ca)s with greater amplitude and shorter delay of onset. At 400 μM, Ca²⁺ activated I_Cl(Ca) instantaneously to a level of ∼2 nA. This concentration-dependent response can be better appreciated in Fig. 4 B where an averaged curve is displayed. Fitting this curve with a Hill equation yielded an apparent EC₅₀ of 3.3 μM and a Hill coefficient of 0.9. To our surprise, the Ca²⁺ sensitivity of Cl_(Ca) channels in mouse airway smooth muscle is much closer to those from Xenopus oocytes (EC₅₀ = ∼3.5 μM at −75 mV) (Kuruma and Hartzell, 2000) and olfactory neurons (EC₅₀ = ∼2.0–5 μM at around −50 mV) (Pifferi et al., 2006) than those from other types of smooth muscle (EC₅₀ = 0.25–0.5 μM at −50 to −100 mV) (Pacaud et al., 1992; Wang and Kotlikoff, 1997; Piper and Large, 2003). We therefore adopted the P_o–voltage relationship of Cl_(Ca) channels from the oocytes to derive [Ca²⁺] sensed by Cl_(Ca) channels underlying STICs (see below).

Having determined the Ca²⁺ sensitivity of Cl_(Ca) channels, it is important to estimate [Ca²⁺] “seen” by Cl_(Ca) channels since this parameter will enable us to put spatial constraints on their distance from RyRs. We employed an approach that had successfully derived the [Ca²⁺] experienced by BK channels underlying STOCs triggered by Ca²⁺ sparks. The rationale and procedure of this approach have been described in detail previously (ZhuGe et al., 2002) and a brief description is given as following. Conductance of an STIC (g_(STIC)) is the product of N × P_{o(Ca, V)} × γ, where N is the number of Cl_(Ca) channels available to a Ca²⁺ spark, γ is the unitary conductance of Cl_(Ca) channels, and P_o is the probability of Cl_(Ca) channel being opened and is a function of both [Ca²⁺] and voltage. Since γ is constant over a large range of voltages (Takahashi et al., 1987; Piper and Large, 2003) and N is assumed to be fixed in a given site (which is the case as shown below), g_(STIC) becomes proportional to P_o, which in turn is a function of both [Ca²⁺] and voltage. It is demonstrated below that the amplitude of Ca²⁺ sparks is constant over a range of voltages, thus any change in g_(STIC) at different voltages should solely reflect the voltage sensitivity of Cl_(Ca) channels in the spark sites. Accordingly, by comparing the g_(STIC)–voltage relationship with the P_o–voltage relationship at constant [Ca²⁺] acquired in excised-patch experiments, one can deduce “apparent” [Ca²⁺] that is responsible for the activation of Cl_(Ca) channels underlying an STIC.

Fig. 5 A displays an example, representative of nine cells, in which STICs were recorded at different voltages. At −85 mV, the leak current was −16 ± 2 pA (n = 9). As predicted, at the potentials more negative than −15 mV, i.e., E_Cl, STICs were inward; and at the potentials less negative than the E_Cl, they changed direction from inward to outward. (This feature is another indication that STICs result from the opening of Cl_(Ca) channels.) For each STIC, Ohm's law can be applied to calculate its g_(STIC), since the γ of Cl_(Ca) channel is constant over voltages examined (Takahashi et al., 1987; Piper and Large, 2003). Fig. 5 B shows the relationship between averaged g_(STIC) and voltage. It is worth noting that g_(STIC) increased as voltage rose from −85 to −5 mV and then plateaued as voltage rose further (ANOVA for a general linear mixed model (Kempthorne, 1975; P < 0.0001). (The value for g_(STIC) at −25 mv is omitted due to uncertainty in identifying STICs near E_Cl.) This pattern of change in g_(STIC) hints that the number of Cl_(Ca) channels for a given spark site is fixed such that the activity of these channels approaches a plateau at positive potentials. We thus scale the g_(STIC) value at −5 mV to the maximal value of p_o activated by 40 μM Ca²⁺ from excised patches of Xenopus oocytes in Fig. 6.

The decline in g_(STIC) at negative potentials could result from a decrease in [Ca²⁺] from Ca²⁺ sparks; we therefore measured the spark amplitude at −85 and 0 mV. The mean values of signal mass, after taking into consideration the endogenous fixed Ca²⁺ buffer, are 240,000 ± 33,000 Ca²⁺ ions at −85 mV and 258,000 ± 30,000 Ca²⁺ ions at 0 mV (P > 0.706; n = 74 at −85 mV and n = 61 at 0 mV). The values for I_Ca(spark) are 4.0 ± 0.6 pA at −85 mV and 4.3 ± 0.5 pA at 0 mV (P > 0.694; the same n as for signal mass). These results argue that Ca²⁺ sparks do not change their characteristics at voltages examined.

The invariance in the amplitude of Ca²⁺ sparks at different potentials suggests that the decline in g_(STIC) could not be due to a decrease in [Ca²⁺] at negative potentials. Moreover, given that (a) the number of available Cl_(Ca) channels appears to be fixed for a given spark site (Fig. 5 B), and that (b) γ is constant over a large range of voltages (Takahashi et al., 1987; Piper and Large, 2003), the observed g_(STIC)–voltage relationship reflects a relationship between P_o and voltage at a [Ca²⁺] produced by Ca²⁺ sparks.

To determine this [Ca²⁺], we overlaid the g_(STIC)–voltage curve with P_o–voltage curves of Cl_(Ca) channels at known [Ca²⁺] obtained in excised patch from Xenopus oocytes. The P_o–voltage curves from Xenopus oocytes (Kuruma and Hartzell, 2000) were chosen because of the remarkable similarities in apparent EC₅₀ of Cl_(Ca) channels to Ca²⁺ between the oocytes (∼3.5 μM at −75 mV) and mouse airway smooth muscle (3.3 μM at −80 mV). Also Cl_(Ca) channels from smooth muscle exhibit a voltage dependence trend similar to those in Xenopus oocytes (Angermann et al., 2006). As shown in Fig. 6, the g_(STIC)–voltage curve fell closely to the P_o–voltage curve for 2.4 μM [Ca²⁺]. Further visual inspection suggests that the [Ca²⁺] level underlying the g_(STIC)–voltage relationship could not be as high as 40 μM, since if this were the case, there would be no change in g_(STIC) across different voltages. Neither could the [Ca²⁺] be as low as 1 μM, since if so, no STIC would be detected at potentials more negative than −50 mV. Thus if Cl_(Ca) channels underlying STICs have a similar voltage sensitivity to those channels in the oocytes, then it is reasonable to conclude that on average, Cl_(Ca) channels underlying STICs are exposed to Ca²⁺ of 2.4 μM or greater, a concentration equal to or greater than EC₅₀ for these channels.

With the knowledge of [Ca²⁺] “seen” by Cl_(Ca) channels underlying STICs and the waveform and amplitude of I_Ca(spark), we can estimate the spatial arrangement of Cl_(Ca) channels and RyRs in Ca²⁺ spark sites. To do so, we first derived the spatio-temporal profile of [Ca²⁺] produced by Ca²⁺ sparks using measured I_Ca(spark) as Ca²⁺ input in reaction-diffusion simulations. The parameters for simulations are provided in the Materials and methods. In brief, located at 20 nm from the plasma membrane, the spark was modeled as an ∼20 nm³ “point” source of I_Ca(spark)s. The amplitude of I_Ca(spark) was adjusted to compensate for the endogenous fixed Ca²⁺ buffer so that resultant [CaFluo3] equaled that measured experimentally (Fig. S1 for detail). Fig. 7 A shows the spatial and temporal profile of [Ca²⁺] produced by Ca²⁺ sparks under the estimated buffer condition. A notable feature in this profile is that, at a distance >600 nm, the [Ca²⁺] reaches its peak later and decays slower than the underlying I_Ca(spark). At distances of <600 nm from the source, the temporal profile of [Ca²⁺] closely follows I_Ca(spark), i.e., it rises to peak linearly and decays to the zero level exponentially.

To estimate the area that is likely occupied by Cl_(Ca) channels near the spark sites, we plotted and examined [Ca²⁺] profiles along the plasma membrane (Fig. 7 B). Since the [Ca²⁺] profile evolves as the I_Ca(spark) progresses, we examined the profiles at two time points, i.e., 9 ms at the peak of I_Ca(spark) and 40 ms, which is ∼86% of STIC peak (see Fig. 2 and Table I). The profile at 9 ms shows a maximal size of Ca²⁺ domain. But at this time point Cl_(Ca) channels might not yet be in equilibrium with Ca²⁺. The profile at 40 ms is the result of I_Ca(spark) when it decays near the basal level, displaying a much smaller domain size. At this point, however, Cl_(Ca) channels should be in equilibrium with Ca²⁺ since STICs reach ∼86% of their maximal value (Fig. 2 and Table I). Thus examining Ca²⁺ profiles from these two time points should define a lower and higher boundary for the area occupied by activated Cl_(Ca) channels in the spark sites. As shown in Fig. 7 B (red lines), at 9 ms of I_Ca(spark), the lateral distance where [Ca²⁺] drops below 2.4 μM is ∼600 nm from the Ca²⁺ source. At 40 ms of I_Ca(spark), this distance shrinks to 300 nm (Fig. 7 B, black lines). Therefore the area occupied by Cl_(Ca) channels could lie in the range between 300 and 600 nm in radius.

Do the areas estimated above reflect a physical domain or a functional domain for Cl_(Ca) channels in spark sites? In other words, Cl_(Ca) channels could be physically confined to the areas near spark sites as estimated, e.g., within 600 nm, or they could be uniformly distributed in the plasma membrane such that an STIC results from the activation of the channels in an area where a [Ca²⁺] reaches 2.4 μM and greater. To distinguish between these two possibilities, we compared the number of Cl_(Ca) channels from all Ca²⁺ sparks sites and the number of Cl_(Ca) channels in an entire cell.

To obtain the number of Cl_(Ca) channels that can be activated by Ca²⁺ sparks from all the sites in a cell, we performed two sets of experiments. First, we determined the number of Ca²⁺ sparks sites in these cells. In line with our previous study (ZhuGe et al., 2004), we defined a spark site as an area of 333 × 333 nm; i.e., if the epicenter pixel of two sparks did not localize in the same pixel, they would be assigned to two different spark sites. Fig. 8 A displays the images when several of Ca²⁺ sparks reached their peak during a series of recordings. Notably, Ca²⁺ sparks occurred in all parts of the cell. By repeatedly acquiring images at the same speed for ten 3-s sequences, we localized each Ca²⁺ spark to construct a two-dimensional Ca²⁺ spark site map. (Imaging for longer time was avoided so as to minimize the possible bleaching by the laser of fluo-3 fluorescence.) Fig. 8 B is such a map from the cell shown in Fig. 8 A; each circle indicates the occurrence of a single spark with the center of the circle lying at the spark's epicenter pixel and the area of the circle proportional to the magnitude of ΔF/F₀ at that pixel. Visual inspection of such maps makes it evident that there are many spark sites per cell. On average, the spark sites per field of observation were 36 ± 4 (N = 18, Fig. 8 C). Since roughly one third of the cell was in the field of observation, the number of sites would appear to be close to 100 per cell.

Next, we calculated the averaged number of Cl_(Ca) channels per spark site. As indicated in Fig. 5 B, on average g_(STIC) at −85 mV was 340 pS. This conductance could result from the opening of ∼50% of Cl_(Ca) channels in the spark site since at positive potentials g_(STIC) increased about onefold. This leads to an estimate that there could be as many as 340 Cl_(Ca) channels per spark site given that each of these channels has a γ of 2 pS (Takahashi et al., 1987; Piper and Large, 2003). For a single cell with ∼80 spark sites triggering STICs (after considering 20% are STIC-less sparks), the total number of Cl_(Ca) channels at these sites was estimated to be a minimum of 27,200.

As a second estimation of the total number of Cl_(Ca) channels in these cells, we measured maximal I_Cl(Ca) by raising [Ca²⁺] to 50–400 μM with flash photolysis of NPEGTA of the whole cell (see Materials and methods). We found that at −80 mV, 50 μM [Ca²⁺] activated an I_Cl(Ca) of 1,889 ± 362 pA (N = 4) and 400 μM caused a current of 1,853 ± 342 pA (N = 7). Therefore, almost one order of magnitude of difference in [Ca²⁺] triggered I_Cl(ca) with a similar amplitude, indicating that these levels of [Ca²⁺] most likely activate all the Cl_(Ca) channels in the cells. If we assume Cl_(Ca) channels have a maximum P₀ at 0.8 and a γ of 2 pS, then the total number of Cl_(Ca) channels in a cell is ∼17,800. This estimate is in rough agreement with the first one above.

Taken together, these estimates indicate that all the Cl_(Ca) channels in a cell could localize in the spark sites. Since the area covered by a Ca²⁺ spark where [Ca²⁺] is >2.4 μM is ∼1.13 μm² (a circle with 600 nm in radius) and the surface area of a cell is ∼2,800 μm² (given a 3-μm radius and a 150-μm length), Cl_(Ca) channels could be present in ∼3% of plasma membrane and, moreover, in these areas the density of the channels could be as high as 300 channels/μm².

Since the discovery of Ca²⁺ sparks in smooth muscle more than a decade ago, researchers have focused on the pathophysiology of Ca²⁺ sparks and STOCs (Nelson et al., 1995; ZhuGe et al., 1998; Brenner et al., 2000; Amberg and Santana, 2003), but paid little attention to the functional coupling of Ca²⁺ sparks and STICs. In the current study, we have gained insight into the mechanisms generating STICs by Ca²⁺ sparks. We found that Ca²⁺ sparks trigger STICs by activating nearby Cl_(Ca) channels that form clusters at high density in the plasma membrane. The finding represents a major step forward in understanding of how Ca²⁺ sparks activate STICs in smooth muscle.

Several lines of evidence suggest that Cl_(Ca) channels highly concentrate in the surface membrane and closely couple with RyRs in Ca²⁺ spark sites. First, by analyzing and comparing the voltage dependence of STIC and Cl_(Ca) channels, we found that during a Ca²⁺ spark, Cl_(Ca) channels underlying STICs are exposed to a [Ca²⁺] at 2.4 μM or greater. Given the [Ca²⁺] spatial-temporal profile produced by Ca²⁺ sparks, for Cl_(Ca) channels to be exposed to that level of [Ca²⁺], they must localize closer than a micron from RyRs, occupying an area of ∼1 μm². We also observed that the conductance of STIC (g_(STIC)) increases as voltage becomes less negative, but it reaches a plateau at potentials greater than −5 mV. This result indicates that the number of Cl_(Ca) channels activated by Ca²⁺ sparks is limited, and the area occupied by these channels is confined in Ca²⁺ sparks sites. If Cl_(Ca) channels homogenously are distributed in the surface membrane, g_(STIC) should continue to increase as a function of voltage since these channels, though not gated by voltage, are voltage dependent (Kuruma and Hartzell, 2000; Angermann et al., 2006). Their nonhomogenous distribution is further supported by the observation that some Ca²⁺ sparks, despite their near membrane location, fail to trigger STICs. We further demonstrated that the number of Cl⁻ channels present in Ca²⁺ spark sites is close to the number of all Cl_(Ca) channels present in the entire cell. Consistent with [Ca²⁺] dropping with distance from the spark source due to diffusion and buffering, the total surface area that can be exposed to [Ca²⁺] sufficient to generate STICs is estimated to be <5% of the surface area of a cell. This strongly indicates that Cl_(Ca) channels concentrate within 600 nm of Ca²⁺ spark sites. We estimate that the density of these channels near the spark sites can be as high as 300 channels/μm² given that they have a small unitary conductance of 2 pS (Takahashi et al., 1987; Piper and Large, 2003). Finally, we found that at −85 mV, STICs with amplitude >20 pA can be routinely recorded in these cells when E_Cl is set at −15 mV. This magnitude of STICs should result from the activation of Cl_(Ca) channels exposed to a high [Ca²⁺], since at this potential their apparent EC₅₀ to Ca²⁺ is on the order of 3 μM (Fig. 4, see below). To experience this level of [Ca²⁺], Cl_(Ca) channels have to reside near RyRs that produce Ca²⁺ sparks, as [Ca²⁺] would drop steeply moving away from RyRs (Fig. 7).

Our notion that RyRs and Cl_(Ca) channels form a microdomain is in line with an emerging idea that Ca²⁺-sensitive channels cluster near to their triggers, resulting in efficient molecular coupling. For instance, BK channels appear to form clusters near RyRs in gastric smooth muscle cells (ZhuGe et al., 2002); they also cluster in the proximity of voltage-gated Ca²⁺ channels in the central nervous system (Berkefeld et al., 2006). Moreover, the clustering of these channels could undergo dynamic changes under different physiological stages. For example, in mice as pregnancy approaches term, BK channels in myometrial cells go through a transition from a clustered to a diffused distribution in the plasma membrane (Eghbali et al., 2003). This sort of transition could serve as a mechanism to fine tune the function of the channels in the cells. It will be important to determine molecular mechanisms leading to cluster formation and transition.

Cl_(Ca) channels in smooth muscle are thought to be quite sensitive to Ca²⁺, with a K_d of 0.2–0.5 μM and a threshold as low as 50 nM at negative potentials between −50 and −100 mV (Pacaud et al., 1992; Wang and Kotlikoff, 1997; Piper and Large, 2003). However, the results in the present study indicate that in mouse airway smooth muscle cells they are quite insensitive to Ca²⁺, with an apparent EC₅₀ as high as 3.3 μM at −80 mV. There are several possible reasons underlying this discrepancy. One possibility could stem from a difference in the molecular identity of Cl_(Ca) channels among smooth muscle. It has been demonstrated that, in addition to Ca²⁺ sensitivity, Cl_(Ca) channels in smooth muscle display variations in unitary conductance, sensitivity to kinase and phosphatase modulation, and other biophysical properties (Large and Wang, 1996; Wang and Kotlikoff, 1997; Angermann et al., 2006). These differences point to the possibility that Cl_(Ca) channels in different smooth muscle may differ in molecular makeup. A definitive answer to this possibility awaits the identification of gene(s) for Cl_(Ca) channels in airway smooth muscle and other smooth muscle.

Another plausible reason for the difference between our results and others' could lie in the difference in the experimental approaches used. In the present study, we employed flash photolysis of caged Ca²⁺ to raise [Ca²⁺], an approach with advantages in changing [Ca²⁺] instantaneously and uniformly. Because of these advantages, [Ca²⁺]s we estimated are expected to closely reflect the concentration sensed by Cl_(Ca) channels. However, most of the earlier studies used fura-2 to derive [Ca²⁺] over the entire cells after stimulation with agonists and voltage (e.g., Pacaud et al., 1992; Wang and Kotlikoff, 1997). Since these sorts of stimuli cause a heterogeneous increase in [Ca²⁺] (Etter et al., 1996), [Ca²⁺] “seen” by Cl_(Ca) channels and measured by fluorescence indicators may vary significantly, leading to an overestimate of Ca²⁺ sensitivity of Cl_(Ca) channels. It is interesting to note that when [Ca²⁺]_cyto was clamped to precise levels by dialyzing Ca²⁺ into cells via the patch pipette, Angermann et al. (2006) recently observed that in pulmonary smooth muscle cells, 1 μM [Ca²⁺] fails to activate Cl_(Ca) channels at potentials more negative than E_Cl (0 mV), highlighting the importance of the method used to control [Ca²⁺]_cyto when determining the sensitivity of Cl_(Ca) channels.

Our estimate of a low Ca²⁺ sensitivity of Cl_(Ca) channels, based upon a fast uncaging approach, is in agreement with our measurements that these cells have a small leak current, i.e., ∼−15 pA, at −85 mV when all K⁺ currents were blocked. Given a maximal total I_Cl(Ca) of 2 nA and a resting [Ca²⁺] of ∼100 nM, this current reflects the activation of 0.5–1% Cl_(Ca) channels, an indication that they are not very sensitive to Ca²⁺. If they were as sensitive as previously reported, a leak current with much greater amplitude would be expected. For instance, according to one proposed Cl_(Ca) channel kinetic model (Kuruma and Hartzell, 2000), for an apparent K_d of 250 nM, and a Hill coefficient of 3, a leak current of ∼300 pA is expected in the same recording conditions as in our experiments (unpublished data; see also Angermann et al., 2006). This value would be 20-fold greater than we measured in these cells.

Many different kinds of smooth muscle produce Ca²⁺ sparks spontaneously, but there is lack of information as to the number of RyRs underlying Ca²⁺ sparks and the mechanisms of generation and termination of Ca²⁺ sparks. In the present study, we have measured the amplitude of I_Ca(spark) based on individual Ca²⁺ sparks recorded with high-speed imaging. With an appropriate correction for the endogenous fixed Ca²⁺ buffer, as determined in the same type of cells, we estimate that I_Ca(spark) is 8.5 pA for a subset of Ca²⁺ sparks with the biggest amplitude and 4.1 pA for the entire population. Given a unitary current of ∼0.35 pA for RyR under the quasi-physiological conditions (Mejia-Alvarez et al., 1999), the number of RyRs for Ca²⁺ sparks in smooth muscle could be on the order of 10–25. Thus, it is reasonable to conclude that Ca²⁺ sparks result from the opening of multiple RyRs in smooth muscle.

Our estimate of the waveform of I_Ca(spark) raises a possibility that RyRs undergo a novel mechanism to generate and terminate Ca²⁺ sparks in smooth muscle. With an unprecedented temporal resolution, we uncover a unique waveform of I_Ca(spark); that is, it reaches its peak around 9 ms and decays exponentially with a time constant of 12 ms. From our simulation, the estimated endogenous fixed Ca²⁺ buffer, although affecting the amplitude of I_Ca(spark), does not alter the shape of this waveform in a significant way, given the assumption that this buffer has the same diffusion-limited on-rate as fluo-3. Therefore, the detected waveform likely represents the kinetic features of RyRs underlying Ca²⁺ sparks in smooth muscle. A 9-ms rising time of I_Ca(spark) implies that the RyRs underlying Ca²⁺ sparks perhaps do not open in concert; if they do, the rise would be much faster. This result is in contrast with ideas on the genesis of Ca²⁺ sparks in skeletal and cardiac muscle, where the opening of RyRs for Ca²⁺ sparks is thought to be in concert (Cheng et al., 1996; Lacampagne et al., 1999; Zhou et al., 2005). The molecular basis for the concerted opening of RyRs in striated muscle lies in their cellular organization by forming discrete clusters in the amount of ∼100 channels in the Z-disk (Block et al., 1988; Sun et al., 1995). Although not regularly arranged as in the striated muscle, RyRs in smooth muscle appear to form clusters, too, as revealed by immunolight and immunoelectron microscopy (Lesh et al., 1998; Lifshitz, L.M., J.D. Carmichael, K.D. Bellve, R.A. Tuft, K.E. Fogarty, and R. ZhuGe. 2008. The Joint Biophysical Society 52nd Annual Meeting and 16th IUPAB International Biophysics Congress. Abstr. 1238-Pos). Our finding of the slow activation of RyRs in clusters suggests a different gating mechanism to produce Ca²⁺ sparks in smooth muscle. (Several reports showed that Ca²⁺ sparks in smooth muscle exhibit a slower activation phase compared with those in striated muscles [Gordienko et al., 1999; Kirber et al., 2001; Ji et al., 2004; Burdyga and Wray, 2005; Liu et al., 2007; McGahon et al., 2007], but it is not known whether the rising phase of I_Ca(spark) is also slow in those studies since no such analysis was performed.) It is interesting to note that the coupled gating of RyRs has been demonstrated in both skeletal and cardiac muscle (Marx et al., 1998, 2001), but not yet in smooth muscle. Whether lack of this type of gating is a reason for the nonconcerted opening of RyRs in smooth muscle requires further investigation.

A single exponential decay of I_Ca(spark) indicates RyRs do not close in concert either, a finding different from that in striated muscles, where a concerted closure is also proposed (Stern and Cheng, 2004). Our observation instead suggests that Ca²⁺ sparks may be terminated as a result of stochastic closure of RyRs in the clusters. If we assume a two-state model for RyRs, we could expect that the mean open time of these channels in the clusters should be close to the decay time constant, i.e., 12 ms, of I_Ca(spark). Remarkably, a very recent study by Laver (2007) reported that the mean open time of RyR₂ is ∼10 ms when luminal [Ca²⁺] is 100 μM, a concentration that is close to [Ca²⁺]_SR in smooth muscle (ZhuGe et al., 1999). It is important to point out that our interpretation of termination of Ca²⁺ sparks does not rule out other possibilities, i.e., stochastic attrition of RyRs, SR Ca²⁺ depletion, RyR inactivation, and a combination of coupled gating and depletion of SR Ca²⁺; all of them have being proposed and tested in striated muscle (Sobie et al., 2002; Stern and Cheng, 2004). However, the more complex composition of RyRs in smooth muscle, i.e., the presence of all three isoforms (Lohn et al., 2001; Yang et al., 2005) and possibly several splicing variants from each isoform, makes it likely that Ca²⁺ sparks in these cells could terminate in a way quite different from that in striated muscle.

Airway smooth muscle does not generate action potential and its membrane potential usually operates in the range of −70 to −20 mV (Janssen, 2002). At these potentials, EC₅₀ for Cl_(Ca) channels should be on the order of 3 μM. For this level of Ca²⁺ sensitivity, the global [Ca²⁺] seems unable to effectively activate these channels since physiological stimulations only raise it to ∼1 μM (Becker et al., 1989). Therefore it appears that Cl_(Ca) channels, like BK channels, are not an effective target of global Ca²⁺ signaling; instead they act as the preferred target for local Ca²⁺ events. We speculate this could be a driving force that results in their localization in the vicinity of RyRs to form a microdomain. As a result, the molecular architecture of the Ca²⁺ microdomain conveys the efficiency and accuracy of Cl_(Ca) channels in response to localized Ca²⁺ signaling. This sort of arrangement should occur in other Ca²⁺ signaling systems where local Ca²⁺ signaling is required.

We wish to thank Drs. John Walsh and Valerie DeCrescenzo for stimulating discussions and advice on the manuscript, and Dr. Stephen Baker for the statistical analyses of Fig. 5.

This study was supported by grants from the National Institutes of Health to R. ZhuGe (HL73875) and to J.V. Walsh (HL21697), and by grants from American Heart Association and Charles Hood Foundation to R. ZhuGe.

David C. Gadsby served as editor.