Sarcoplasmic reticulum calcium load regulates rat arterial smooth muscle calcium sparks and transient K_Ca currents

Abstract

The regulation of calcium (Ca²⁺) sparks and transient calcium-sensitive K⁺ (K_Ca) currents by acute changes in sarcoplasmic reticulum (SR) Ca²⁺ load ([Ca²⁺]_SR) was investigated in rat cerebral artery smooth muscle cells using laser-scanning confocal microscopy in combination with patch clamp electrophysiology. [Ca²⁺]_SR was elevated by: (i) increasing the activity of the SR Ca²⁺-ATPase with an anti-phospholamban monoclonal antibody, or (ii) blocking Ca²⁺ release from the SR with tetracaine, a membrane-permeant, reversible ryanodine-sensitive Ca²⁺ release (RyR) channel blocker. Alternatively, [Ca²⁺]_SR was progressively decreased over time with a low concentration of thapsigargin (20 nm), a SR Ca²⁺-ATPase blocker. An elevation in [Ca²⁺]_SR increased Ca²⁺ spark and transient K_Ca current frequency, but did not alter the amplitude, decay or spatial spread of Ca²⁺ sparks or the coupling ratio or amplitude correlation between Ca²⁺ sparks and evoked transient K_Ca currents. Decreasing [Ca²⁺]_SR reduced Ca²⁺ spark frequency, amplitude and spatial spread and this reduced transient K_Ca current frequency and amplitude. However, even when mean Ca²⁺ spark amplitude and spread decreased by up to 47 and 56 % of control, respectively, the coupling ratio or amplitude correlation between Ca²⁺ sparks and transient K_Ca currents was not affected. These data demonstrate that acute changes in [Ca²⁺]_SR regulate Ca²⁺ sparks and transient K_Ca currents in arterial smooth muscle cells.

Intracellular calcium (Ca²⁺) regulates a diverse range of cellular functions, including contraction, secretion and gene transcription. However, recent studies have determined that the intracellular Ca²⁺ concentration ([Ca²⁺]_i) of many cell types is not homogeneously distributed (for reviews see Clapham, 1995; Berridge, 1997). Several different types of intracellular Ca²⁺ elevations have been described to occur that differ in respect to temporal kinetics, spatial localization and physiological function (Bootman et al. 2001). One type of Ca²⁺ signalling modality that has been observed in cardiac (Cheng et al. 1993), skeletal (Tsugorka et al. 1995) and smooth muscle (Nelson et al. 1995) cells is termed a ‘Ca²⁺ spark’. Ca²⁺ sparks are highly localized, cytosolic Ca²⁺ transients that occur due to the opening of a number of ryanodine-sensitive Ca²⁺ release (RyR) channels in the sarcoplasmic reticulum (SR; Jaggar et al. 2000).

In smooth muscle cells, Ca²⁺ sparks exhibit a rise time of ≈20 ms, a half-time of decay of ≈50 ms and a spatial spread of ≈2-3 μm when imaged using a confocal microscope and fluorescent Ca²⁺ indicators such as fluo-3 (Nelson et al. 1995; Jaggar et al. 2000). Although Ca²⁺ sparks elevate intracellular Ca²⁺ in the immediate vicinity of the Ca²⁺ release site to micromolar concentrations (Perez et al. 2001), the impact of Ca²⁺ sparks on the global [Ca²⁺]_i is low, due to their transient and localized properties (Nelson et al. 1995; Jaggar et al. 2000). In arterial smooth muscle cells, most Ca²⁺ sparks occur in close proximity to the plasma membrane and activate a number of large-conductance Ca²⁺-sensitive K⁺ (K_Ca) channels to evoke a transient outward K⁺ current (Nelson et al. 1995; Bolton & Gordienko, 1998; Perez et al. 1999), which has been termed a ‘spontaneous transient outward current’ or ‘STOC’ (Benham & Bolton, 1986). In arteries at physiological levels of pressure, inhibition of Ca²⁺ sparks or K_Ca channels leads to membrane depolarization, activation of voltage-dependent Ca²⁺ channels, an elevation in the arterial wall [Ca²⁺]_i and constriction (Nelson et al. 1995; Jaggar, 2001). An elevation in intravascular pressure activates voltage-dependent Ca²⁺ channels in arterial smooth muscle cells leading to an increase in the global [Ca²⁺]_i and activation of Ca²⁺ sparks (Jaggar, 2001). The resulting elevation in K_Ca channel activity opposes the pressure-induced constriction (Brayden & Nelson, 1992).

Several signalling elements regulate Ca²⁺ sparks in smooth muscle cells, including intracellular Ca²⁺ and protein kinases (Jaggar et al. 2000). Smooth muscle RyR channels that are incorporated into lipid bilayers are activated by cytosolic Ca²⁺ elevations (Herrmann-Frank et al. 1991; Xu et al. 1994), suggesting that an elevation in [Ca²⁺]_i may regulate Ca²⁺ sparks via an interaction with activation sites located on the cytosolic face of the RyR channel. However, an increase in cytosolic [Ca²⁺]_i or activation of the SR Ca²⁺-ATPase may increase SR Ca²⁺ load ([Ca²⁺]_SR), which could also regulate RyR channels. Cardiac (Sitsapesan & Williams, 1994; Gyorke & Gyorke, 1998; Xu & Meissner, 1998; Ching et al. 2000) and skeletal (Herrmann-Frank & Lehmann-Horn, 1996; Tripathy & Meissner, 1996) muscle RyR channels incorporated into lipid bilayers are activated by an elevation in luminal Ca²⁺ concentration. Furthermore, an elevation in [Ca²⁺]_SR activates Ca²⁺ sparks in cardiac myocytes (Santana et al. 1997; Satoh et al. 1997; Lukyanenko et al. 2001). Recent studies have also suggested that [Ca²⁺]_SR may regulate Ca²⁺ sparks in smooth muscle cells. Genetic ablation of phospholamban, an endogenous inhibitor of the SR Ca²⁺-ATPase, leads to a chronic elevation in [Ca²⁺]_SR and Ca²⁺ spark frequency in arterial smooth muscle cells, when compared to wild type controls (Wellman et al. 2001). In Bufo marinus stomach smooth muscle cells, following partial depletion of the [Ca²⁺]_SR with caffeine, Ca²⁺ spark frequency and amplitude increase during [Ca²⁺]_SR refilling (ZhuGe et al. 1999). Vasodilators also stimulate Ca²⁺ sparks, in part, by elevating [Ca²⁺]_SR (Porter et al. 1998; Wellman et al. 2001). However, despite the important role that the [Ca²⁺]_SR may play in intracellular Ca²⁺ signalling in smooth muscle, the regulation of Ca²⁺ sparks and transient K_Ca currents by [Ca²⁺]_SR is poorly understood.

The goal of this study was to investigate the regulation of Ca²⁺ sparks and transient K_Ca currents by acute changes in [Ca²⁺]_SR in arterial smooth muscle cells. [Ca²⁺]_SR was elevated with a monoclonal antibody raised against phospholamban, or with tetracaine, a membrane-permeant reversible RyR channel blocker. Alternatively, [Ca²⁺]_SR was progressively decreased with a low concentration of thapsigargin, a selective inhibitor of the SR Ca²⁺-ATPase. Data suggest that an elevation in [Ca²⁺]_SR increases Ca²⁺ spark and transient K_Ca current frequency, but does not alter the amplitude, decay or spatial spread of Ca²⁺ sparks or the coupling ratio or amplitude correlation between Ca²⁺ sparks and evoked transient K_Ca currents. In contrast, decreasing [Ca²⁺]_SR reduces Ca²⁺ spark frequency, amplitude and spatial spread, which decreases the frequency and amplitude of evoked transient K_Ca currents, although the coupling ratio and amplitude correlation between Ca²⁺ sparks and K_Ca channels does not change. These findings provide evidence that acute changes in [Ca²⁺]_SR directly alter the frequency and amplitude properties of Ca²⁺ sparks and evoked transient K_Ca currents.

METHODS

Tissue preparation

Sprague-Dawley rats (≈250 g) of either sex were killed by peritoneal injection of a sodium pentobarbital overdose (150 mg kg⁻¹), in accordance with the Animal Care and Use Committee policies and procedures at the University of Tennessee. The brain was removed and placed into ice-cold (4 °C), oxygenated (95 % O₂-5 % CO₂), physiological saline solution (PSS) containing (mm): 119 NaCl, 4.7 KCl, 24 NaHCO₃, 1.2 KH₂PO₄, 1.6 CaCl₂, 1.2 MgSO₄, 0.023 EDTA and 11 glucose (adjusted to pH 7.4 with NaOH). Posterior cerebral and cerebellar arteries (50-200 μm diameter) were removed, cleaned of basolateral connective tissue and maintained in ice-cold PSS. Where appropriate, the endothelium was removed by allowing an air bubble to remain in the lumen of the artery for 2 min, followed by a 30 s wash with H₂O (Jaggar & Nelson, 2000). Individual smooth muscle cells were enzymatically dissociated from cerebral arteries using a procedure similar to that described in Jaggar (2001).

Confocal calcium imaging

Arterial segments (1-2 mm in length) or isolated smooth muscle cells were placed into Hepes-buffered PSS of composition (mm): 134 NaCl, 6 KCl, 2 CaCl₂, 1 MgCl₂, 10 Hepes and 10 glucose (pH 7.4, NaOH) containing 10 μM fluo-4 AM and 0.05 % pluronic F-127 for 60 min or 15 min, respectively, at 22 °C. To allow indicator de-esterification, arteries or isolated cells were subsequently placed into Hepes-buffered PSS for 30 min at 22 °C. Smooth muscle cells were imaged using a Noran Oz laser scanning confocal microscope (Noran Instruments, Middleton, WI, USA) and a ×60 water immersion objective (NA = 1.2) attached to a Nikon TE300 microscope by illuminating with a krypton-argon laser at 488 nm. Planar images (56.3 μm × 52.8 μm) were recorded every 16.7 ms, i.e. 60 images s⁻¹. To compare confocal Ca²⁺ imaging data with electrophysiological recordings performed in this study, Ca²⁺ sparks in smooth muscle cells of arterial segments were measured in an extracellular solution containing 30 mm K⁺, which depolarizes smooth muscle cells to ≈-40 mV, and is similar to the membrane potential of cerebral arteries pressurized to 60 mmHg (see Jaggar et al. 1998; Jaggar & Nelson, 2000; Jaggar, 2001 for similar procedure). The 30 mm K⁺ bath solution contained (mm): NaCl 110; KCl 30; Hepes 10; CaCl₂ 2; MgCl₂ 1; and glucose 10 (pH 7.4, NaOH). For imaging smooth muscle cells in arteries, at least two different representative areas (56.3 μm × 52.8 μm) of the same segment were each scanned for at least 10 s under each condition. The same area of artery was scanned only once to avoid any laser-induced changes in Ca²⁺ signalling, and the effects of drugs were measured in paired experiments. Where appropriate, diltiazem (50 μM) was applied for 15 min to ensure a complete and steady state decrease in Ca²⁺ sparks and global Ca²⁺ fluorescence prior to tetracaine application. Tetracaine was then added in the presence of diltiazem, and further Ca²⁺ imaging measurements were made 5 min later. Time-matched control experiments confirmed that diltiazem induced steady-state block of Ca²⁺ sparks prior to tetracaine application. Ca²⁺ spark frequency in smooth muscle cells was not different after 15 (0.32 ± 0.09 Hz) or 20 min (0.26 ± 0.09 Hz) in the continued presence of diltiazem (50 μM, P > 0.05, n= 5 arteries). In experiments where confocal microscopy was employed in combination with patch clamp electrophysiology, current and fluorescence measurements were synchronized using a light emitting diode positioned above the recording chamber that was triggered during acquisition. Each single smooth muscle cell was imaged for at least 10 s under each condition. Ca²⁺ sparks were detected in smooth muscle cells using custom analysis software written using IDL 5.0.2 (Research Systems Inc., Boulder, CO, USA) kindly provided by Drs M. T. Nelson and A. D. Bonev (University of Vermont, VT, USA). Automated and manual detection of Ca²⁺ sparks was performed by dividing an area 1.54 μm (7 pixels) × 1.54 μm (7 pixels) (i.e. 2.37 μm²) in each image (F) by a baseline (F₀) which was determined by averaging 10 images without Ca²⁺ spark activity. The entire area of each image was analysed to detect Ca²⁺ sparks. A Ca²⁺ spark was defined as a localized increase in F/F₀ that was greater than 1.2. In intact artery experiments, Ca²⁺ spark frequency (Hz) in each condition was calculated from at least two different areas of each artery wall. Ca²⁺ spark spatial spread was calculated as the full width at half-maximal amplitude (FWHM). Global Ca²⁺ fluorescence was calculated from the same images used for Ca²⁺ spark analysis and was the mean pixel value of 100 different images acquired over 10 s.

Ratiometric calcium measurements

Cerebral artery segments (1-2 mm in length) were incubated with the ratiometric fluorescent Ca²⁺ indicator fura-2 AM (2 μM) and 0.05 % pluronic F-127 for 20 min, followed by a 15 min wash. Experiments were performed using a 6 mm potassium Hepes-buffered PSS (composition described above). Arteries were alternately excited at 340 or 380 nm using a PC driven Hyperswitch (Ionoptix, Milton, MA, USA), and background corrected ratios were collected at every 0.2 s at 510 nm using a photomultiplier tube. [Ca²⁺]_SR was determined by rapidly applying a high concentration of caffeine (20 mm), a RyR channel activator, wherein the amplitude of the [Ca²⁺]_i transient would be related to the [Ca²⁺]_SR. [Ca²⁺]_i concentrations were calculated using the following equation (Grynkiewicz et al. 1985):

where R is the 340/380 nm ratio, R_min and R_max are the minimum and maximum ratios determined in Ca²⁺-free and saturating Ca²⁺ solutions, respectively, S_f2/S_b2 is the Ca²⁺-free/Ca²⁺-replete ratio of emissions at 380 nm excitation, and K_d is the dissociation constant for fura-2. R_min, R_max, S_f2 and S_b2 were determined at the end of each experiment and in separate experiments by increasing the Ca²⁺ permeability of smooth muscle cells with ionomycin (10 μM), and perfusing cells with a high Ca²⁺ (50 mm) or Ca²⁺-free (10 mm EGTA) solution. The in situ apparent dissociation constant (K_d) for fura-2 used in this study was 282 nm (Knot & Nelson, 1998).

Patch clamp electrophysiology

Isolated cells were allowed to adhere to a glass coverslip in the bottom of a chamber for 10 min prior to experimentation. K⁺ currents were measured using either the conventional whole-cell or perforated patch configuration (Horn & Marty, 1988) of the patch clamp technique (Hamill et al. 1981) with an Axopatch 200B amplifier (Axon Instruments, Union City, CA, USA). Bath solution was 6 mm K⁺ Hepes-buffered PSS (composition described above). For perforated patch experiments, the pipette solution contained (mm): 110 potassium aspartate, 30 KCl, 10 NaCl, 1 MgCl₂, 10 Hepes, 0.05 EGTA (pH 7.2 with KOH). For conventional whole-cell experiments, the pipette (i.e. intracellular) solution contained (mm): 140 KCl, 1.9 MgCl₂, 0.037 CaCl₂, 0.1 EGTA, 10 Hepes, 1 Na₂ATP (pH 7.2 with KOH); the calculated free Ca²⁺ and free Mg²⁺ concentrations of this solution are 100 nm and 1 mm, respectively (WEBMAXC, Stanford University, CA, USA). The conventional whole-cell configuration was used in experiments with anti-phospholamban antibody. Elsewhere in this study, the perforated patch configuration was used. All patch clamp experiments were performed with a holding potential of −40 mV. Membrane currents were recorded with a sample rate of 2.5 kHz and filtered at 1 kHz. Transient K_Ca current analysis was performed off-line using a custom analysis program provided by Drs M. T. Nelson and A. D. Bonev (University of Vermont). A transient K_Ca current was defined as the simultaneous opening of three or more K_Ca channels. In the presence of Ca²⁺ spark blockers, the simultaneous opening of three single K_Ca channels was not observed at −40 mV (Nelson et al. 1995; Bonev et al. 1997; Porter et al. 1998). For experiments performed in the absence or presence of anti-phospholamban antibody, transient K_Ca current properties were compared between 5 and 10 min after whole-cell patch formation. Where appropriate, anti-phospholamban antibody was heat-inactivated by incubation at 94 °C for 15 min.

Statistical analysis

Values are expressed as means ± standard error of the mean. Student's t test or Student-Newman-Keuls tests were used for comparing paired or multiple data sets, respectively. First-order polynomial linear fits were used to calculate statistical correlation between the amplitude of Ca²⁺ sparks and evoked transient K_Ca currents (Origin, OriginLab Corp., Northampton, MA, USA). ANCOVA of first-order polynomial best fits were used to compare amplitude correlation data sets (Prism, GraphPad Software, Inc., San Diego, CA, USA). P < 0.05 was considered significant.

Chemicals

Unless stated otherwise, all chemicals used in this study were obtained from Sigma Chemical Company (St Louis, MO, USA). Papain was purchased from Worthington Biochemical Co. (Lakewood, NJ, USA), fluo-4 AM and pluronic F-127 from Molecular Probes (Eugene, OR, USA), and anti-phospholamban mouse IgG1 clone A1 from Upstate Biotechnology (Lake Placid, NY, USA).

RESULTS

A monoclonal antibody targeted against phospholamban elevates transient K_Ca current frequency in cerebral artery smooth muscle cells

To investigate the regulation of transient K_Ca currents by an acute elevation in [Ca²⁺]_SR, we used a commercially available monoclonal antibody (mouse IgG1) raised against phospholamban, an endogenous inhibitor of the SR Ca²⁺-ATPase. This antibody has previously been demonstrated to elevate [Ca²⁺]_SR in cardiac myocytes (Suzuki & Wang, 1986; Sham et al. 1991; Lukyanenko et al. 2001). Transient K_Ca current properties were compared between 5 and 10 min after conventional whole-cell formation in cells in which antibody (4 μg ml⁻¹) or heat-inactivated antibody (4 μg ml⁻¹) was included in the pipette solution, or antibody was absent.

Mean transient K_Ca current frequency was ≈2.7-fold higher in cells in which anti-phospholamban antibody was included in the pipette solution (n= 11), when compared with cells in which heat-inactivated antibody was included (n= 6 cells), or antibody was absent (n= 11, Fig. 1). In contrast, the anti-phospholamban antibody did not change mean transient K_Ca current amplitude (Fig. 1). These data suggest that acute activation of the SR Ca²⁺-ATPase with an anti-phospholamban antibody increases the frequency, but does not alter the amplitude, of transient K_Ca currents in arterial smooth muscle cells.

Figure 1. A monoclonal antibody raised against phospholamban (anti-PLB antibody) elevates transient K_Ca current frequency, but not amplitude, in cerebral artery smooth muscle cells.

A, original traces illustrating typical transient K_Ca current activity at −40 mV in cells in which anti-PLB antibody (4 μg ml⁻¹, upper panel) or heat-inactivated anti-PLB antibody (4 μg ml⁻¹, lower panel) was included in the pipette solution. Average transient K_Ca current frequency (B) and amplitude (C) in cells in which no antibody (control), anti-PLB antibody, or heat-inactivated antibody was included in the pipette solution. *P < 0.05; Student's unpaired t test.

Tetracaine inhibits Ca²⁺ sparks in cerebral artery smooth muscle cells

We sought to measure the regulation of Ca²⁺ sparks and transient K_Ca currents before and after an elevation in [Ca²⁺]_SR in the same cell. We hypothesized that RyR channel blockers would increase [Ca²⁺]_SR by preventing Ca²⁺ release from the SR under conditions where the SR Ca²⁺-ATPase would continue to sequester Ca²⁺ from the cytosol. To examine this possibility we used tetracaine, a membrane-permeant, reversible blocker of cardiac (Gyorke et al. 1997; Lukyanenko et al. 2001) and skeletal (Xu et al. 1993) muscle RyR channels. To determine if tetracaine blocks RyR channels in smooth muscle cells, the regulation of Ca²⁺ sparks by tetracaine (50 μM) was measured in the smooth muscle cells of endothelium-denuded cerebral artery segments. To compare confocal Ca²⁺ imaging data with electrophysiological data performed in this study, Ca²⁺ sparks were measured in a bath solution containing 30 mm K⁺, which depolarizes smooth muscle cells from ≈-60 to −40 mV, a membrane potential similar to that of cerebral arteries pressurized to 60 mmHg (see Jaggar et al. 1998; Jaggar & Nelson, 2000; Jaggar, 2001 for similar procedure). To prevent any potential indirect regulation of Ca²⁺ sparks due to non-specific effects on voltage-dependent Ca²⁺ channels (Sugiyama & Muteki, 1994), tetracaine was applied in the continued presence of diltiazem (50 μM), a voltage-dependent Ca²⁺ channel blocker.

Diltiazem reduced mean global Ca²⁺ fluorescence in smooth muscle cells (F/F₀) to 53 ± 3 % of control, and decreased mean Ca²⁺ spark frequency to ≈20 % of control, although mean Ca²⁺ spark amplitude (F/F₀) did not change (n= 6 arteries, Fig. 2). In the same arteries in the continued presence of diltiazem, tetracaine reduced mean Ca²⁺ spark frequency to ≈3 % of control, but did not change mean Ca²⁺ spark amplitude (Fig. 2). Tetracaine did not alter mean global F/F₀ (98 % ± 2 % of diltiazem), suggesting that Ca²⁺ sparks were not blocked via a reduction in global [Ca²⁺]_i (Fig. 2). These data demonstrate that tetracaine, a RyR channel blocker, inhibits Ca²⁺ sparks and does not alter global [Ca²⁺]_i in cerebral artery smooth muscle cells, when applied in the continued presence of diltiazem.

Figure 2. Tetracaine blocks Ca²⁺ sparks in smooth muscle cells of intact cerebral arteries.

A, average fluorescence (100 of 600 images) over 10 s of three different 56.3 μm × 52.8 μm areas of the same cerebral artery in control (30 mm K⁺), diltiazem (50 μM) and tetracaine (50 μM) + diltiazem (50 μM). The locations of Ca²⁺ sparks that occurred during 10 s are indicated by white boxes (1.54 μm × 1.54 μm) and representative localized F/F₀ changes with time are illustrated below respective images and labelled accordingly. In this artery over 10 s, 27 sparks occurred in control, five sparks in diltiazem and one spark in tetracaine + diltiazem. Average effects on Ca²⁺ spark frequency (B) and amplitude (C). *P < 0.05; Student-Newman-Keuls test.

Tetracaine elevates sarcoplasmic reticulum Ca²⁺ load in cerebral arteries

Blocking Ca²⁺ release from the SR with tetracaine should lead to an increase in [Ca²⁺]_SR. To examine this hypothesis, the effect of tetracaine (50 μM) on the amplitude of caffeine (20 mm)-induced [Ca²⁺]_i transients was investigated in endothelium-denuded cerebral artery segments using the ratiometric Ca²⁺ indicator fura-2. Experiments were performed in a bath solution containing 6 mm K⁺.

Diltiazem did not change mean arterial wall [Ca²⁺]_i (control, 99 ± 5 nmvs. diltiazem, 100 ± 5 nm) or the amplitude of caffeine-induced [Ca²⁺]_i transients (Fig. 3, n= 6 arteries). In the same arteries, although tetracaine did not change mean arterial wall [Ca²⁺]_i (104 ± 4 nm), mean caffeine-induced [Ca²⁺]_i transients increased ≈1.4-fold after a 7 min application. Five minutes after tetracaine washout, caffeine-induced [Ca²⁺]_i transients returned to pre-tetracaine levels (Fig. 3). These data suggest that tetracaine, a RyR channel blocker, reversibly elevates [Ca²⁺]_SR in cerebral artery smooth muscle cells, presumably by blocking Ca²⁺ release from the SR (Fig. 2).

Figure 3. Tetracaine elevates [Ca²⁺]_SR in cerebral artery segments.

A, intracellular Ca²⁺ concentration and caffeine (20 mm)-induced [Ca²⁺]_i transients in an endothelium-denuded cerebral artery segment. Diltiazem (50 μM) did not change arterial wall [Ca²⁺]_i or caffeine-induced [Ca²⁺]_i transients. Tetracaine (50 μM, 7 min application) did not change arterial wall [Ca²⁺]_i, but significantly increased the amplitude of the caffeine-induced [Ca²⁺]_i transient. Five minutes after washout of tetracaine, the caffeine-induced [Ca²⁺]_i transient was of similar amplitude to control. B, average caffeine-induced [Ca²⁺]_i transients in control, diltiazem, tetracaine + diltiazem, and 5 min after tetracaine washout + diltiazem. *P < 0.05; Student-Newman-Keuls test.

Regulation of transient K_Ca currents by tetracaine

If tetracaine inhibits Ca²⁺ sparks, it should also block transient K_Ca currents. Furthermore, if an increase in [Ca²⁺]_SR activates Ca²⁺ sparks, transient K_Ca currents should be elevated immediately after washout of tetracaine, when compared with transient K_Ca currents prior to tetracaine application. To investigate this hypothesis, the regulation of transient K_Ca currents by tetracaine was measured in voltage-clamped (-40 mV) cerebral artery smooth muscle cells. Tetracaine was applied, and washed out, in the continued presence of Cd²⁺ (250 μM), a voltage-dependent Ca²⁺ channel blocker.

Cd²⁺ reduced mean transient K_Ca current frequency to ≈54 % of control, but did not change mean transient K_Ca current amplitude (Fig. 4, n= 6 cells). In the continued presence of Cd²⁺, tetracaine (50 μM, 7 min application) further reduced mean transient K_Ca current frequency, but did not alter mean amplitude. Washout of tetracaine was rapid and led to a significant increase in mean transient K_Ca current frequency. Between 1 and 3 min after tetracaine washout, mean transient K_Ca current frequency was ≈3.1-fold higher than that prior to tetracaine application, although mean transient K_Ca current amplitude was not significantly different (Fig. 4). Approximately 5 min after tetracaine washout, but in the continued presence of Cd²⁺, transient K_Ca current frequency returned to pre-tetracaine levels (Fig. 4).

Figure 4. Regulation of transient K_Ca currents by tetracaine.

A, original record illustrating transient K_Ca currents in a cerebral artery smooth muscle cell voltage-clamped at −40 mV using the perforated-patch configuration. Cd²⁺ (250 μM) reduced transient K_Ca current frequency. Tetracaine (50 μM, 7 min application), applied in the continued presence of Cd²⁺, further reduced transient K_Ca currents. Immediately after tetracaine washout, transient K_Ca current frequency was significantly higher than before tetracaine application, although transient K_Ca current amplitude was not significantly different. Transient K_Ca current frequency returned to pre-tetracaine levels approximately 5 min after tetracaine washout. Average effects of Cd²⁺ (250 μM), tetracaine (50 μM) + Cd²⁺, and 1–3 min after tetracaine washout + Cd²⁺ on transient K_Ca current frequency (B) and amplitude (C). *P < 0.05; Student-Newman-Keuls test.

These data demonstrate that tetracaine blocks transient K_Ca currents. After washout of tetracaine, when [Ca²⁺]_SR would have increased (Fig. 3), transient K_Ca current frequency was significantly higher than prior to tetracaine application, although transient K_Ca current amplitude was not different. Because tetracaine washout does not alter cytosolic [Ca²⁺]_i (Fig. 3), these data suggest that an acute elevation in [Ca²⁺]_SR increases transient K_Ca current frequency, but not amplitude. Furthermore, the elevation in transient K_Ca current frequency was short lived after tetracaine washout, suggesting that elevated Ca²⁺ release from the SR leads to a return of [Ca²⁺]_SR and transient K_Ca current frequency to pre-tetracaine levels.

Tetracaine washout elevates Ca²⁺ spark frequency, but not amplitude

We sought to investigate the effect of an acute elevation in [Ca²⁺]_SR on Ca²⁺ sparks and transient K_Ca currents. Essentially 100 % of Ca²⁺ sparks induce a transient K_Ca current in rat cerebral artery smooth muscle cells (Perez et al. 1999). Therefore, our finding that an elevation in [Ca²⁺]_SR elevates transient K_Ca current frequency suggests that this occurs via an increase in Ca²⁺ spark frequency. However, an acute increase in [Ca²⁺]_SR could conceivably elevate Ca²⁺ spark amplitude via activation of RyR channels (Herrmann-Frank & Lehmann-Horn, 1996; Tripathy & Meissner, 1996; Gyorke & Gyorke, 1998; Xu & Meissner, 1998; Ching et al. 2000) and by increasing the driving force for Ca²⁺ from the SR. A significant proportion of Ca²⁺ sparks do not evoke a transient K_Ca current in human cerebral artery (28 %, Wellman et al. 2002), feline oesophageal (27 %, Kirber et al. 2001) or Bufo marinus stomach (21 %, ZhuGe et al. 2000) smooth muscle cells. If Ca²⁺ spark:transient K_Ca current coupling is not 1:1, an increase in Ca²⁺ spark amplitude may augment this coupling ratio, which would also increase transient K_Ca current frequency. Therefore, we measured the effect of an elevation in [Ca²⁺]_SR on Ca²⁺ spark and evoked transient K_Ca current properties, and the coupling ratio and amplitude relationship between these events. Simultaneous measurements of Ca²⁺ sparks and transient K_Ca currents were made in the same cells before tetracaine, and 90 s after tetracaine washout (50 μM, 7 min application), in the continued presence of diltiazem (50 μM).

Prior to tetracaine application, 100 % of detected Ca²⁺ sparks evoked a transient K_Ca current (n= 4 cells, n= 33 sparks) and 71.7 % of transient K_Ca currents were associated with a Ca²⁺ spark, suggesting that some sparks occurred outside the cytosolic volume detected by the confocal microscope (Fig. 5). The amplitude of a Ca²⁺ spark and that of the evoked transient K_Ca current were significantly correlated (P < 0.0001) (Fig. 6). In the same cells, tetracaine significantly reduced transient K_Ca current frequency, consistent with effects in non-fluo-4 loaded cells (Fig. 4). After tetracaine washout, when [Ca²⁺]_SR would have increased, mean Ca²⁺ spark frequency was ≈2.7-fold higher than prior to tetracaine application (Fig. 5, n= 88 sparks). In contrast, mean Ca²⁺ spark amplitude (Fig. 5), half-time for decay (t_1/2; control, 52 ± 9 vs. tetracaine washout, 57 ± 6 ms) and spatial spread (FWHM; control, 2.23 ± 0.15 vs. tetracaine washout, 1.95 ± 0.17 μm) were not different (P > 0.05 for each). Consistent with effects in non-fluo-4 loaded cells, mean transient K_Ca current amplitude (control, 23.8 ± 2.8 vs. tetracaine washout, 23.1 ± 4.6 pA) also did not change (P > 0.05). After tetracaine washout, 100 % of Ca²⁺ sparks evoked a transient K_Ca current and 68.2 % of transient K_Ca currents were associated with a Ca²⁺ spark, suggesting that an elevation in [Ca²⁺]_SR did not change the coupling ratio between sparks and transient K_Ca currents. Furthermore, tetracaine washout did not significantly alter the amplitude correlation between Ca²⁺ sparks and evoked transient K_Ca currents (Fig. 6). Global F/F₀ was not altered by washout of tetracaine (99 ± 6 % of pre-tetracaine), suggesting that Ca²⁺ sparks were activated via an elevation in [Ca²⁺]_SR and not via an increase in cytosolic [Ca²⁺]_i, and that the increase in Ca²⁺ spark frequency did not elevate global cytosolic [Ca²⁺]_i. These data suggest that an elevation in [Ca²⁺]_SR increases Ca²⁺ spark frequency, but does not alter Ca²⁺ spark amplitude, decay, spatial spread, or the coupling ratio or amplitude correlation between sparks and transient K_Ca currents.

Figure 5. Tetracaine washout elevates Ca²⁺ spark frequency.

A, original simultaneous recordings of Ca²⁺ sparks and transient K_Ca currents in a voltage-clamped (-40 mV) cerebral artery smooth muscle cell. The black line illustrates whole cell K⁺ current. Red and green lines illustrate fluorescence changes (F/F₀) measured in two different 1.54 μm × 1.54 μm (i.e. 2.37 μm²) areas of the cell in which Ca²⁺ sparks occurred. Traces indicate activity before tetracaine (50 μM) application (pre-tetracaine) and 90 s after tetracaine washout. Tetracaine was applied for 7 min. The experiment was performed in the continued presence of diltiazem (50 μM). B, summary of Ca²⁺ spark frequency and amplitude pre-tetracaine (n= 33 sparks, n= 4 cells) and after tetracaine washout (n= 88 sparks). *P < 0.05; Student's paired t test.

Figure 6. Tetracaine washout does not alter the amplitude correlation between Ca²⁺ sparks and transient K_Ca currents.

Scatter plot of Ca²⁺ spark and evoked transient K_Ca current amplitude at −40 mV before tetracaine application (pre-tetracaine, black), and after tetracaine washout (grey). The amplitudes of Ca²⁺ sparks and evoked transient K_Ca currents were significantly correlated, both for pre-tetracaine (P < 0.0001) and tetracaine washout data (P < 0.0001). Correlation co-efficients (r) were: control, 0.55; tetracaine washout, 0.58. The amplitude correlations before, and after, tetracaine were not significantly different from each other (P= 0.79).

Thapsigargin progressively decreases SR Ca²⁺ load in cerebral arteries

If an elevation in [Ca²⁺]_SR increases Ca²⁺ spark frequency in arterial smooth muscle cells, then a decrease in [Ca²⁺]_SR should decrease Ca²⁺ spark frequency. Furthermore, because the [Ca²⁺]_SR establishes the driving force for Ca²⁺ from the SR, a decrease in [Ca²⁺]_SR may also reduce Ca²⁺ spark amplitude. To investigate the effect of decreasing [Ca²⁺]_SR on Ca²⁺ sparks and transient K_Ca currents, thapsigargin, a selective blocker of the SR Ca²⁺-ATPase, was used. In smooth muscle cells, thapsigargin has most commonly been applied at a high concentration (≈100 nm), to rapidly deplete the SR of Ca²⁺ (e.g. see Jaggar, 2001). To determine the effect of a low concentration of thapsigargin (20 nm) on [Ca²⁺]_SR, caffeine-induced [Ca²⁺]_i transients were measured in control, and 5, 10 and 15 min after application of thapsigargin (20 nm) in the same endothelium-denuded cerebral artery segments.

Thapsigargin (20 nm) induced a time-dependent decrease in caffeine-induced [Ca²⁺]_i transients (Fig. 7, n= 9 arteries). Five, ten and fifteen minutes after addition of thapsigargin, caffeine-induced [Ca²⁺]_i transients decreased to ≈76, ≈57 and ≈41 % of control, respectively. In contrast, 5, 10 and 15 min after thapsigargin application, steady-state arterial wall [Ca²⁺]_i was 99 ± 2, 101 ± 2 and 102 ± 2 nm, respectively, which were not significantly different from control (P > 0.05 for each, Fig. 7). To ensure that [Ca²⁺]_SR was reduced by thapsigargin, and not by repetitive addition of caffeine or insufficient recovery of [Ca²⁺]_SR after each caffeine application, caffeine-induced [Ca²⁺]_i transients were measured in six separate arteries in the absence of thapsigargin using the same time protocol. In the absence of thapsigargin, caffeine-induced [Ca²⁺]_i transients were not significantly different at 0, 5, 10 or 15 min (P > 0.05 for each, Fig. 7). In these control arteries, arterial wall [Ca²⁺]_i also did not change over the same time course (0 min, 99 ± 3 nm; 15 min, 96 ± 5 nm, P > 0.05). These data demonstrate that a low concentration of thapsigargin (20 nm) progressively decreases [Ca²⁺]_SR in cerebral arteries, and that significant depletion of [Ca²⁺]_SR does not alter arterial wall [Ca²⁺]_i.

Figure 7. Thapsigargin reduces [Ca²⁺]_SR in cerebral arteries.

A, intracellular Ca²⁺ concentration and caffeine (20 mm)-induced [Ca²⁺]_i transients in control and 5, 10 and 15 min after thapsigargin application (20 nm) in an endothelium-denuded cerebral artery segment. B, relative time-dependent change in caffeine-induced [Ca²⁺]_i transients when compared with control in the absence (black bars) or presence of thapsigargin (20 nm, hatched bars). Mean caffeine-induced [Ca²⁺]_i transients for each time point were (control vs. thapsigargin): 0 min, 240 ± 11 vs. 245 ± 14 nm; 5 min, 235 ± 8 vs. 184 ± 10 nm; 10 min, 229 ± 8 vs. 136 ± 7 nm; 15 min, 234 nm± 9 vs. 98 ± 8 nm. *P < 0.05 when compared using unpaired Student's t test.

Thapsigargin progressively decreases Ca²⁺ spark frequency, amplitude and spatial spread

We sought to examine the effect of progressively decreasing [Ca²⁺]_SR on Ca²⁺ sparks and transient K_Ca currents. Simultaneous measurements of Ca²⁺ sparks and transient K_Ca currents were made in the same voltage-clamped (-40 mV) cerebral artery smooth muscle cells in control, and 5, 10 and 15 min after addition of thapsigargin (20 nm).

In control, 98.2 % of detected Ca²⁺ sparks evoked a transient K_Ca current (Fig. 8, Table 1, n= 55 sparks, n= 6 cells). The amplitudes of coupled Ca²⁺ sparks and evoked transient K_Ca currents were correlated (Fig. 8). In control, one Ca²⁺ spark out of a total of 56 did not evoke a transient K_Ca current. The amplitude (F/F₀) of this uncoupled Ca²⁺ spark was 1.26, which was significantly smaller than the mean amplitude of coupled sparks (Table 1), and is only just larger than our designated threshold for a Ca²⁺ spark, which is an F/F₀ greater than 1.2. In addition, the spatial spread of the uncoupled spark was smaller (0.96 μm) and the decay was faster (18.0 ms) than the mean value for coupled sparks (Table 1). In the same cells, thapsigargin (20 nm) induced a time-dependent decrease in mean Ca²⁺ spark frequency, amplitude and spatial spread, which reduced transient K_Ca current frequency and amplitude (Table 1). Five, ten or fifteen minutes after addition of thapsigargin, 100 % of Ca²⁺ sparks evoked a transient K_Ca current, suggesting that uncoupling of Ca²⁺ sparks from K_Ca channels did not occur, even when mean spark amplitude and spread were reduced to 47 and 56 % of control, respectively (Table 1). The amplitudes of Ca²⁺ sparks and transient K_Ca currents were correlated at 5, 10 or 15 min after thapsigargin (Fig. 8). Furthermore, the amplitude correlation for each time point after thapsigargin addition was not significantly different when compared with the amplitude correlation in control (Fig. 8). These data suggest that although a decrease in [Ca²⁺]_SR reduces Ca²⁺ spark amplitude and spatial spread, the amplitude correlation between a spark and a transient K_Ca current is maintained (Fig. 8). Thapsigargin did not change global F/F₀ (103 ± 5 % of control at 15 min, P > 0.05), suggesting that all modifications in Ca²⁺ sparks were due to a decrease in [Ca²⁺]_SR and not due to alterations in cytosolic [Ca²⁺]_i.

Figure 8. Regulation of Ca²⁺ sparks and transient K_Ca currents by thapsigargin.

A, original simultaneous recordings of Ca²⁺ sparks (red and green lines) and whole-cell current (black lines) in the same voltage-clamped (-40 mV) cerebral artery smooth muscle cell in control and 10 min after thapsigargin (20 nm) application. B, the amplitudes of Ca²⁺ sparks and evoked transient K_Ca currents were correlated for control (P= 0.002), and data obtained 5 (P= 0.04), 10 (P < 0.0001), or 15 (P < 0.0001) min after thapsigargin. Linear correlation coefficients (r) were: control (black), 0.46; thapsigargin: 5 min (red), 0.41; 10 min (green), 0.38; 15 min (blue), 0.32. Amplitude correlations for each time point after thapsigargin application were not significantly different when compared with control (significance values: 5 min, P= 0.76; 10 min, P= 0.94; 15 min, P= 0.83).

Table 1.

Summary of the effects of thapsigargin on Ca²⁺ spark and transient K_Ca currents

		Post thapsigargin (20 nM)time (min)
	Control	5	10	15
Ca2+ sparks
Frequency (Hz)	1.0 ± 0.23 (6 cells)	0.43 ± 0.14 (6 cells)*	0.32 ± 0.15 (6 cells)*	0.16 ± 0.07 (6 cells)*
Amplitude (F/F0)	1.97 ± 0.07 (55)	1.65 ± 0.05 (19)*	1.55 ± 0.03 (16)*	1.46 ± 0.07 (5)*
Decay (t1/2, ms)†	40.9 ± 4.4 (29)	35.6 ± 2.5 (14)	32.6 ± 3.1 (12)	30.6 ± 2.8 (5)
Spread (FWHM, μm)	2.11 ± 0.19 (18)	2.07 ± 0.08 (11)	1.21 ± 0.11 (11)*	1.18 ± 0.21 (11)*
KCa transients
Amplitude (pA)	23.7 ± 2.4 (61)	17.0 ± 2.3 (36)*	15.0 ± 1.3 (29)*	12.2 ± 2.6 (11)*
Decay (t1/2, ms)‡	16.6 ± 0.9 (61)	15.8 ± 0.9 (36)	15.2 ± 0.8 (29)	14.8 ± 1.0 (11)
Rise time (10–90%,ms)	11.8 ± 0.6 (61)	11.5 ± 0.7 (36)	11.7 ± 0.7 (29)	10.7 ± 1.0 (11)

All Ca²⁺ spark data refer to events associated with transient K_Ca currents. Only one Ca²⁺ spark was detected in this series of experiments that did not evoke a transient K_Ca current and this spark occurred in control conditions. Transient K_Ca current data also includes events for which an associated Ca²⁺ spark was not detected. Values of n shown in parentheses refer to number of sparks or transient K_Ca currents used for analysis, unless specified otherwise.

^†Ca²⁺ spark decays were fit with a second-order exponential.

^‡Transient K_Ca current decays were fit with a first-order exponential. t_1/2: time for decay to half-maximal amplitude. FWHM: full width at half maximal amplitude.

^*P < 0.05 when compared with control; Student-Newman-Keuls test.

To investigate if the repetitive confocal acquisition protocol was responsible for the observed changes in Ca²⁺ sparks and transient K_Ca currents, the same imaging procedure was performed in voltage-clamped (-40 mV) cerebral artery smooth muscle cells in the absence of thapsigargin. In six cerebral artery smooth muscle cells, Ca²⁺ spark frequency was not different at 0 min (first laser exposure, 0.76 ± 0.1 Hz) and 15 min (fourth laser exposure, 0.81 ± 0.1 Hz, P > 0.05 for each). Similarly, Ca²⁺ spark amplitude (F/F₀; 0 min, 2.21 ± 0.16 n= 48; 15 min, 2.15 ± 0.11, n= 51) and transient K_Ca current amplitude (0 min, 18.8 ± 2.2 pA; 15 min, 17.6 ± 2.2 pA) were unchanged (P > 0.05 for each). These data indicate that the repetitive confocal Ca²⁺ imaging protocol did not induce changes in Ca²⁺ sparks and transient K_Ca currents. In summary, progressively decreasing [Ca²⁺]_SR reduced Ca²⁺ spark frequency, amplitude and spatial spread, which reduced transient K_Ca current frequency and amplitude. However, even when Ca²⁺ spark amplitude and spread were reduced by 47 and 56 % of control, respectively, the coupling ratio or the amplitude correlation between Ca²⁺ sparks and transient K_Ca currents was not altered.

DISCUSSION

We demonstrate that an acute increase or decrease in [Ca²⁺]_SR regulates Ca²⁺ sparks and evoked transient K_Ca currents in cerebral artery smooth muscle cells. An elevation in [Ca²⁺]_SR increased Ca²⁺ spark and transient K_Ca current frequency, but did not change Ca²⁺ spark amplitude, spatial spread or decay, or the coupling ratio or amplitude correlation between sparks and transient K_Ca currents. Decreasing [Ca²⁺]_SR reduced Ca²⁺ spark frequency, amplitude and spatial spread, and these effects reduced the frequency and amplitude of evoked transient K_Ca currents, although the coupling ratio or amplitude correlation between Ca²⁺ sparks and transient K_Ca currents was not affected.

An elevation in [Ca²⁺]_SR increases Ca²⁺ spark frequency in arterial smooth muscle cells

[Ca²⁺]_SR was elevated in cerebral artery smooth muscle cells by: (1) selectively increasing the activity of the SR Ca²⁺-ATPase, or (2) blocking RyR channels. To activate the SR Ca²⁺-ATPase we used a monoclonal antibody raised against phospholamban that elevates [Ca²⁺]_SR in cardiac myocytes (Suzuki & Wang, 1986; Sham et al. 1991; Lukyanenko et al. 2001). The anti-phospholamban antibody increased mean transient K_Ca current frequency, but did not alter transient K_Ca current amplitude. [Ca²⁺]_SR was also elevated with tetracaine, a reversible RyR channel blocker. Local anaesthetics block RyR channels by inducing a long closed state rather than by reducing the single channel conductance or the mean open times of the channel (Zahradnikova & Palade, 1993). Tetracaine was applied in the continued presence of a voltage-dependent Ca²⁺ channel blocker, to prevent any non-specific effects on voltage-dependent Ca²⁺ channels that would also regulate Ca²⁺ sparks (Jaggar et al. 1998; Jaggar, 2001). After tetracaine washout when [Ca²⁺]_SR had increased, Ca²⁺ sparks and transient K_Ca currents were significantly more frequent than prior to tetracaine application, but Ca²⁺ spark amplitude, decay and spatial spread were not altered, nor were transient K_Ca current amplitude or the coupling ratio or amplitude correlation between Ca²⁺ sparks and transient K_Ca currents. [Ca²⁺]_SR and transient K_Ca current frequency returned to control levels ≈5 min after tetracaine washout, presumably due to the elevation in Ca²⁺ release.

Ca²⁺ spark amplitude should depend on the number and activity of contributing RyR channels and the driving force for Ca²⁺ from the SR. However, although Ca²⁺ sparks in smooth muscle cells exhibit a wide range of amplitudes (see Fig. 6, Jaggar et al. 2000; ZhuGe et al. 2000), data suggest that an acute elevation in [Ca²⁺]_SR did not change mean Ca²⁺ spark amplitude. Conceivably, this could be because resting [Ca²⁺]_SR saturates Ca²⁺ flux through RyR channels. If this is the case, the SR Ca²⁺-ATPase maintains this saturating [Ca²⁺]_SR at a cytosolic [Ca²⁺]_i of ≈100 nm, and the wide range of Ca²⁺ spark amplitudes that occur at this saturating [Ca²⁺]_SR may be due to different numbers and/or activities of RyR channels that contribute to sparks. The decline in mean Ca²⁺ spark amplitude and spatial spread that occurs when [Ca²⁺]_SR is reduced could also be due to a decrease in the driving force for Ca²⁺ from the SR in addition to a reduction in the number or activity of contributing RyR channels. However, mechanisms that regulate Ca²⁺ spark amplitude may be more complex and may involve multiple processes, particularly since Ca²⁺ activation and inactivation sites may be located on the luminal (Ching et al. 2000) and cytosolic side (Herrmann-Frank et al. 1991; Xu et al. 1994; Herrmann-Frank & Lehmann-Horn, 1996; Tripathy & Meissner, 1996) of RyR channels. In addition, during a Ca²⁺ spark, the localized free Ca²⁺ concentration may partially deplete near the luminal side of RyR channels, and this may also influence Ca²⁺ spark amplitude.

Our finding that an acute increase in SR Ca²⁺ load elevates Ca²⁺ spark frequency but mean Ca²⁺ spark amplitude does not change is consistent with findings in arterial smooth muscle cells of a phospholamban-deficient mouse, wherein SR Ca²⁺ load is chronically elevated (Wellman et al. 2001). However, modifications in Ca²⁺ signalling modalities in smooth muscle cells regulate the transcription factor, CREB, the immediate-early gene, c-fos (Cartin et al. 2000) and the transcription factor, NFAT (Stevenson et al. 2001). Conceivably, long-term modifications in Ca²⁺ signalling modalities, particularly during development, could modify the expression levels of Ca²⁺ signalling proteins, such as RyR channels, and this could have led to the observation that a chronic elevation in SR Ca²⁺ load does not change Ca²⁺ spark amplitude. Forskolin, an activator of adenylyl cyclase, also elevates SR load and Ca²⁺ spark frequency in arterial smooth muscle cells, but does not change mean spark amplitude (Porter et al. 1998). However, cAMP-dependent protein kinase also activates RyR channels (Porter et al. 1998; Wellman et al. 2001), which could have masked a change in Ca²⁺ spark amplitude. Our data, which examined the regulation of Ca²⁺ sparks and transient K_Ca current by an acute elevation in SR Ca²⁺ load, also suggest that frequency is increased, but amplitude does not change. When comparing our data with that in previous studies, findings suggest that an acute, or chronic, elevation in [Ca²⁺]_SR elevates Ca²⁺ spark frequency, but the amplitude and decay of Ca²⁺ sparks are unchanged.

A decrease in [Ca²⁺]_SR reduces Ca²⁺ spark frequency, amplitude and spatial spread

Thapsigargin induced a progressive reduction in [Ca²⁺]_SR and Ca²⁺ spark frequency, amplitude and spatial spread, and these changes decreased transient K_Ca current frequency and amplitude. Presumably, transient K_Ca current amplitude was reduced because: (1) a lower amplitude Ca²⁺ spark would be a less effective activator of K_Ca channels, and (2) a smaller spark spread would envelop a reduced sarcolemmal surface area and impact a lower number of K_Ca channels. The thapsigargin-induced decrease in Ca²⁺ spark amplitude (47 % of control) and spatial spread (56 % of control) did not induce uncoupling of Ca²⁺ sparks from K_Ca channels. In this study, out of a total of 216 sparks, only one uncoupled Ca²⁺ spark was detected, suggesting a coupling ratio of sparks to transient K_Ca currents of 99.5 %, which is similar to previous reports in rat cerebral artery smooth muscle cells (Perez et al. 1999). The amplitude (F/F₀) and spatial spread of this uncoupled spark were significantly smaller than the mean value for coupled sparks (Table 1), which may partially explain why it did not evoke a transient K_Ca current. Mechanisms that lead to uncoupling are unclear, but may involve a reduced density (ZhuGe et al. 2000) or Ca²⁺ sensitivity (Bayguinov et al. 2001) of K_Ca channels, in addition to the properties of the Ca²⁺ release event. The variability in coupling ratio that occurs in different smooth muscle preparations (ZhuGe et al. 2000; Kirber et al. 2001; Wellman et al. 2002) may reflect species- and tissue-specific diversity in Ca²⁺ signalling between RyR channels and K_Ca channels.

In cardiac myocytes, Ca²⁺ sparks decay primarily due to diffusion, but some of the released Ca²⁺ is also sequestered into the SR by the SR Ca²⁺-ATPase (Gomez et al. 1996; Santana et al. 1997). In our experiments, partially blocking the SR Ca²⁺-ATPase reduced spark amplitude and spread, and slightly decreased half-time for decay (t_1/2). If the SR Ca²⁺-ATPase sequestered a portion of the released Ca²⁺, thapsigargin should have prolonged decay (t_1/2), which was not observed. In experiments with tetracaine where [Ca²⁺]_SR was increased, spark spread and decay were unaltered. This finding suggests that decreasing the driving force for SR Ca²⁺ uptake also has no effect on spark decay. Thus, the SR Ca²⁺-ATPase does not appear to contribute directly to spark decay by sequestering the released Ca²⁺ into the SR. Data suggest that amplitude largely determines the decay of a spark, since spread and decay (t_1/2) decreased when amplitude was reduced (Table 1). Possible explanations for a correlation between amplitude and spread/decay may be that spark diffusion is space limited or that sparks saturate cytosolic Ca²⁺ buffers within the immediate vicinity of the release site. Thus, lower amplitude sparks would decay more rapidly and would exhibit lower spread due to a higher ratio of cytosolic buffering molecules to Ca²⁺.

A decrease in [Ca²⁺]_SR elevates sarcolemmal Ca²⁺ entry in a variety of cell types via activation of store-operated Ca²⁺ channels, which are also referred to as capacitative Ca²⁺ entry channels or Ca²⁺-release activated Ca²⁺ (CRAC) channels (Parekh & Penner, 1997). Although a store-operated Ca²⁺ entry pathway has been demonstrated in arterial smooth muscle (Xu & Beech, 2001), this observation is not universal (e.g. see Knot et al. 1998). In the present study, a decrease or an increase in [Ca²⁺]_SR did not alter cytosolic [Ca²⁺]_i (Fig. 3 and Fig. 7), suggesting that store-operated Ca²⁺ entry is not significant in rat cerebral artery smooth muscle cells, at least in the conditions used in our experiments.

Physiological relevance of modifications in [Ca²⁺]_SR in arterial smooth muscle cells

An elevation in [Ca²⁺]_SR that increases Ca²⁺ spark and transient K_Ca current frequency should lead to membrane hyperpolarization, a decrease voltage-dependent Ca²⁺ channel activity, a reduction in global [Ca²⁺]_i, and dilation (Jaggar et al. 2000). An elevation in Ca²⁺ spark frequency may also increase the driving force for Ca²⁺ for sarcolemma extrusion mechanisms that are located within the vicinity of the release site, such as the Na⁺-Ca²⁺ exchanger or the Ca²⁺-ATPase. A similar mechanism, referred to as the ‘superficial buffer barrier hypothesis’ (van Breemen et al. 1995), suggests that Ca²⁺ entering smooth muscle cells is buffered by the SR and is discharged vectorially towards the sarcolemma without any effect on global [Ca²⁺]_i. Elevating Ca²⁺ release from the SR may also enhance Ca²⁺-dependent inactivation of sarcolemma voltage-dependent Ca²⁺ channels, an additional mechanism that would further reduce Ca²⁺ entry. Thus, an increase in [Ca²⁺]_SR may be intimately involved in directly regulating a number of negative feedback mechanisms via activation of Ca²⁺ sparks. Conversely, a decrease in [Ca²⁺]_SR would reduce Ca²⁺ spark frequency, amplitude and spatial spread, which would have the opposite effect.

In summary, our data suggest that an elevation in [Ca²⁺]_SR increases Ca²⁺ spark and transient K_Ca current frequency in cerebral artery smooth muscle cells, but Ca²⁺ spark amplitude, spatial spread or decay, or the coupling ratio or amplitude correlation between sparks and transient K_Ca currents are not altered. A decrease in [Ca²⁺]_SR reduces Ca²⁺ spark frequency, amplitude and spatial spread, and the frequency and amplitude of evoked transient K_Ca currents, although the coupling ratio or amplitude correlation between sparks and transient K_Ca currents remains unchanged. These findings demonstrate that acute changes in [Ca²⁺]_SR regulate Ca²⁺ sparks and transient K_Ca currents in cerebral artery smooth muscle cells.