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VASCULAR BIOLOGY

Calcium sparks activate calcium-dependent Cl^– current in rat corpus cavernosum smooth muscle cells

Department of Physiology and Pharmacology, Schulich School of Medicine and Dentistry, University of Western Ontario, London, Ontario, Canada

Submitted 29 October 2006 ; accepted in final form 13 July 2007

	ABSTRACT

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Spontaneous transient currents, due to activation of Ca²⁺-dependent K⁺ and Cl^– channels, occur in corpus cavernosum smooth muscle cells (CCSMC) of the penis. The Ca²⁺ events responsible for triggering Ca²⁺-dependent Cl^– channels have never been identified in vascular muscle. We used high-speed fluorescence imaging combined with patch-clamp electrophysiology to provide the first characterization of Ca²⁺ events underlying these currents. Freshly isolated rat CCSMC loaded with fluo-4 exhibited localized, spontaneous elevations of intracellular Ca²⁺ (Ca²⁺ sparks) in 57% of cells. There was an average of 6.4 ± 0.5 release sites/cell with a frequency of 0.9 ± 1 Hz/cell and peak amplitude F/F_o of 67 ± 10%. We addressed the controversy of whether these events are mediated by ryanodine or inositol 1,4,5 trisphosphate (IP₃) receptors. Caffeine caused either a global Ca²⁺ rise at high concentrations or an increase in spark frequency at lower concentrations, whereas ryanodine dramatically reduced the amplitude and frequency of sparks. 2-Aminoethoxydiphenyl borate, an inhibitor of IP₃ receptors, had no effect on spark frequency. Combined imaging and electrophysiological recording revealed strong coupling between Ca²⁺ sparks and biphasic transient currents, a relationship never before shown in vascular muscle. Moreover, spark frequency increased on depolarization, an effect abolished with the blockade of Ca²⁺ channels, consistent with Ca²⁺ influx regulating Ca²⁺ release from stores. We establish for the first time that Ca²⁺ sparks occur in CCSMC and arise from Ca²⁺ release through ryanodine receptors. Moreover, the voltage dependence of spark frequency demonstrated here provides novel functional evidence for voltage-dependent Ca²⁺ influx in CCSMC.

calcium signaling; potassium and chloride channels; ryanodine receptors

The spatial organization of Ca²⁺ signaling within cells allows for diversity in the actions of Ca²⁺. Global Ca²⁺ rise leads to muscle contraction; however, transient, localized elevations in cytosolic [Ca²⁺] occur in many smooth muscles and are believed to promote relaxation (32). These localized elevations can be either Ca²⁺ "puffs" due to release from sarcoplasmic reticulum stores via inositol 1,4,5-trisphosphate (IP₃) receptors (44) or Ca²⁺ "sparks" due to release of Ca²⁺ from ryanodine receptor (RyR) channels (11). Such localized Ca²⁺ transients are of interest because they regulate the opening of ion channels and thus control cell membrane potential (20, 32).

It has been well established that spark activation of Ca²⁺-dependent K⁺ channels (BK_Ca channels) accounts for spontaneous transient outward currents (STOCs), which in turn cause hyperpolarization of vascular muscle leading to relaxation and vasodilation (9, 20, 32). However, spontaneous transient inward currents (STICs) have also been identified in several smooth muscles (23, 27), representing the activation of Ca²⁺-dependent Cl^– channels (Cl_Ca). A single report describing airway smooth muscle cells demonstrated that Ca²⁺ sparks elicit both STICs and STOCs (47). However, to our knowledge this relationship has never been documented in vascular muscle.

Ca²⁺-activated K⁺ and Cl^– currents have been identified in CCSMCs and are apparent as STOCs and STICs in rat and human (25) and rabbit myocytes (13). Both classes of channels serve important functional roles in regulating activity of the corpus cavernosum. Genetic knockout of BK_Ca impairs hyperpolarization and relaxation of the corpus cavernosum, resulting in erectile dysfunction (42). In contrast, pharmacological blockade of Cl^– channels enhances and prolongs erection (25), reflecting the opposing roles of K⁺ and Cl^– channels in this tissue.

There is controversy concerning the mechanism underlying localized Ca²⁺ release events in some smooth muscles. Most studies conclude that these are due to Ca²⁺ release from RyRs (20), with RyR2 in particular (24, 40). However, there are reports invoking IP₃ receptors as giving rise to spontaneous currents (4, 6, 18), including in CCSM (13). It has also been suggested that localized Ca²⁺ release events could involve a combination of both IP₃ and RyR channels (15).

In this study, we use high-speed fluorescence Ca²⁺ imaging and simultaneous patch-clamp recording to study the Ca²⁺ events underlying the spontaneous currents in corpus cavernosum smooth muscle of the rat. We establish for the first time that Ca²⁺ sparks in CCSMC arise from Ca²⁺ release through RyRs and give rise to Cl_Ca current. Moreover, we reveal physiological regulation of spark frequency with depolarization due to voltage-dependent Ca²⁺ entry. Measurement of spark frequency provides a novel functional index of voltage-dependent Ca²⁺ influx in corpus cavernosum cells.

	METHODS

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Cell isolation. All procedures were approved by the Council on Animal Care of the University of Western Ontario and were in accordance with the Canadian Council on Animal Care. Male Sprague-Dawley rats weighing 300–550 g were injected with a lethal dose of euthanyl (400 mg/kg ip). The penis was removed, and corpus cavernosum smooth muscle tissue was dissected from the crura and dissociated as previously described (43). Tissues were cut into 1-mm thick sections and placed in 2.5 ml of dissociation solution (see Solutions and chemicals) plus the following compounds (in mg/ml): 0.8 papain, 3.0 bovine albumin, 0.5 1,4-dithio-L-threitol, and 1.0 Sigma blend collagenase type F (pH 7.0). Tissues were either placed in a gently shaking water bath at 31°C for 120 to 180 min and dispersed by trituration with fire-polished Pasteur pipettes for immediate use, or they were stored overnight at 4°C. The following day, tissues were warmed to room temperature for 120 min then dispersed. Isolated cells were kept in high K⁺ solution (see Solutions and chemicals) for up to 5 h before use and then perfused with Na⁺ solution for recording of Ca²⁺ events. As described in RESULTS, inspection of cells did not reveal altered Ca²⁺ handling arising from the K⁺ solution, such as Ca²⁺ waves reported in cardiomyocytes (10, 39).

Imaging and measurement of Ca²⁺ sparks. Images were acquired at 40–65 Hz by using a wide-field digital fluorescence imaging system (Photon Technology International; PTI) with a Cascade Photometrics 650 cooled charge-coupled device camera (653 x 492 pixels; Roper Scientific, Tucson, AZ) and ImageMaster Software (versions 3 & 5; PTI). To optimize the speed of acquisition, the region acquired was limited to a single cell. Cells were loaded with the Ca²⁺ indicator dye fluo-4-AM (5 µM, with 0.05% pluronic) for 40 min at room temperature and then transferred to a 1-ml glass-bottomed perfusion chamber and allowed to settle. The recording chamber was mounted on a Nikon inverted microscope (Nikon Eclipse TE2000-U) equipped with a plan apochromatic x60 water immersion lens (numerical aperature 1.2) and a blue excitation filter cube with an emission band pass of 535 ± 40 nm. After settling was completed, cells were perfused with physiological saline solution at a rate of 1–3 ml/min.

With the x60 lens, each pixel represented an area of 196 x 196 nm. The spatial resolution, assessed as the 10–90% edge response, was 0.5 µm. For these experiments, image exposure time was 5 ms with a camera on-chip multiplication gain of 3, and there were 2,000 images/sequence. Several sequences were usually collected from each cell with intervals between sequences of 2 to 5 min. Excitation of fluo-4 was provided by the 488-nm line of a multiline argon laser, and cell exposure to the laser was controlled by an electronic shutter. Image processing was performed off-line using ImageMaster 5, MS Excel (Microsoft) and pClamp Software 9 (Axon Instruments, Foster City, CA).

The acquired images were Gaussian filtered using three-by-three pixels, and Ca²⁺ images were baseline subtracted pixel by pixel using the equation F/F_o (%) = 100 x [F(x,y,t) – F_o(x,y)]/ F_o(x,y), where F(x,y,t) is the fluorescence at each pixel in the time series and F_o is an image of the "baseline" level given by the average of 50 to 100 consecutive images in the absence of sparks. The change in fluorescence F/F_o (%) is a relative measure of free intracellular Ca²⁺ concentration. To create the plots of F/F_o with time, areas of interest of 9 x 9 pixels (3.1 µm²) were located at the center of each spark site. This size of the area of interest was chosen because on average it surrounded the entire event at the time at which each spark event was detected.

The root-mean-square noise of the image was <2%, and an increase in fluorescence was considered to be a Ca²⁺ spark when it was equal to or greater than 5% and lasted for at least two frames, as described earlier (47). The frequency of sparks was measured from F/F_o (%) with time plots using the threshold detection routine in pClamp (version 9.0, Axon Instruments). For cell images in Figs. 1 and 3, images immediately preceding the spark were used to establish baseline level F_o. The beginning of the spark was identified as that image having a change in fluorescence >5%.

View larger version (34K): [in this window] [in a new window]   Fig. 1. Spontaneous, localized Ca2+ release events in corpus cavernosum smooth muscle. Cells were dispersed from rat corpus cavernosum, loaded with fluo-4 dye, and monitored using a high-speed digital fluorescence imaging system (see METHODS). A: time course of fluorescence intensity change, expressed as F/Fo (%) = [(F – Fo)/ Fo] x 100, for 4 active Ca2+ release sites, each shown in a different color. Cells exhibited spontaneous transient elevations of Ca2+ that were spatially restricted, independent of each other, and not associated with contraction. Full frame images were collected at 53 frames/s. The baseline intensity Fo was determined as the average of 50 frames without Ca2+ events. The events, numbered 1 to 4, are from the corresponding sites in B, each representing areas of interest of 9 x 9 pixels (3.1 µm2). B: cell imaged in A is shown in a bright-field image showing the location of 4 of 9 active Ca2+ release sites. C: expanded view of the first event from site 1. Labeled points (a to d) correspond to the images shown in D. D: top, contour plots of entire field of view (blue rectangle) containing the cell and background; bottom, cell as it lies in the field of view, with baseline fluorescence subtracted and the background masked. Online supplemental movie 1 illustrates the spontaneous regional changes of Ca2+ in this cell.

View larger version (30K): [in this window] [in a new window]   Fig. 3. Restricted spread of Ca2+ release events. The spatial characteristics of Ca2+ events were analyzed in two complementary ways; by using a "line scan," comparable to that widely used with confocal microscopy, and by the two-dimensional spread as recorded in wide-field imaging. A: line scan intensity of a Ca2+ event assessed at three time points: at initiation (a), the peak (b), and 89 ms later (c) during the event shown in B. The spread of Ca2+, shown as changes of fluorescence intensity, were fit with Gaussian curves (in red) and the full-width-at-half-maximum (FWHM) calculated to be 4.3 µm (a), 6.4 µm (b), and 7.0 µm (c). B: images of the cell at the three time points a, b, and c, as in A. The circle in image b forms an approximate contour line at 50% peak intensity around the event site, and the line illustrates the position of a line scan used for the distribution in A. Spread diameter was calculated from the diameter of the circle drawn around the event. C: time course of the event in B with the red points a, b, and c, corresponding to the images in B. D: distribution of spread diameter was well fit by a single Gaussian curve (in red), with mean diameter at half-maximum amplitude of was 6.9 ± 0.2 µm (n = 149 events from 21 cells), essentially the same as that deduced using line scan FWHM analysis.

Patch-clamp electrophysiology. Membrane currents and voltages were measured with the nystatin (250 µg/ml) perforated patch technique at room temperature, using a Multiclamp 700 A patch-clamp amplifier connected to a Digidata 1322A analogue to digital converter (Axon Instruments). Currents were acquired at 2–5 kHz and filtered at 1 kHz using pClamp 9.0 Software. Pipette resistance before seal formation was 2–5 M.

All patch-clamp data illustrated in this paper were performed simultaneously with image acquisition on the same cell. The exposure recording TTL output from the CCD camera was sampled with the Digidata to provide precise alignment of image frames with the current record. The F/F_o (%) versus time plots were then aligned offline with current traces using the record of exposures as a guide. Frequency of spontaneous transient currents was measured using the threshold detection routine in pClamp with a current threshold twice the estimated single channel amplitude of the Ca²⁺-activated K⁺ channel and a noise rejection of 10 ms.

Solutions and chemicals. The Krebs bicarbonate solution for retrieval of tissues consisted of (in mM) 122 NaCl, 4.7 KCl, 2.5 CaCl₂, 0.8 MgSO₄, 1.2 NaH₂PO₄, 20 NaHCO₃, 10 D-glucose, 5 HEPES, 0.25 EDTA equilibrated with 5% CO₂-95% O₂ (pH 7.4). The dissociation solution used for cell dispersal contained (in mM) 135 KCl, 10 HEPES 10 D-glucose, 1 CaCl₂, 1 MgCl₂, 10 taurine, and 0.25 EDTA (pH set to 7.0 with KOH). Isolated cells were transferred to high KCl-Ringer containing (in mM) 135 KCl, 20 HEPES, 10 glucose, 1 MgCl₂, and 1 CaCl₂ (pH 7.4 with KOH). The Na⁺-HEPES bath solution used for imaging cells contained (in mM) 130 NaCl, 5 KCl, 20 HEPES, 10 D-glucose, 2 CaCl₂, and 1 MgCl₂ (pH set to 7.4 with NaOH). For perforated patch recording, the bath solution contained (in mM) 130 NaCl, 3 KCl, 1.8 CaCl₂, 1 MgCl₂, and 10 HEPES (pH 7.4 with NaOH). For 0 Ca²⁺ solution, CaCl₂ was omitted from the bath solution. The pipette solution contained (in mM) 137 KCl, 3 MgCl₂, and 10 HEPES (pH 7.2 with KOH). Fluo-4 AM and pluronic were obtained from Molecular Probes (Eugene, OR). Caffeine was obtained from RBI Research Biochemicals (Natick, MA). 2-Aminoethoxydiphenyl borate (2-APB) was obtained from Calbiochem (San Diego, CA). Ryanodine and nifedipine were obtained from Sigma-Aldrich (St. Louis, MO). Test agents were applied by pressure ejection from a glass micropipette of 1- to 2-µm tip diameter, attached to a picospritzer (General Valve, Fairfax, NJ), positioned 50–100 µm from the cell. The concentrations of test agents given are those in the application pipettes, acknowledging that there is dilution at the cell surface.

Statistical analysis. Values are provided as means ± SE, with error bars in the figures representing SE and with n indicating the number of cells, or sparks studied, as indicated. Statistical comparisons were made using either ANOVA with Bonferroni post hoc analysis or Student's t-test, with P < 0.05 indicating significance.

	RESULTS

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Characteristics of spontaneous elevations of Ca²⁺ in corpus cavernosum smooth muscle. A high-speed, wide-field digital fluorescence imaging system was used to record changes of intracellular Ca²⁺ concentration in 73 smooth muscle cells loaded with the Ca²⁺ indicator dye fluo-4. In an additional 25 cells, imaging and patch-clamp recording were used to simultaneously monitor Ca²⁺ events and membrane currents while controlling membrane potential. Approximately 57% of all cells studied (98 of 172 cells in total) showed spontaneous Ca²⁺ events, with a representative cell shown in Fig. 1. Localized transient increases of fluorescence intensity above baseline (F_o) are plotted as F/F_o (%) (Fig. 1A). In any given active cell, there were multiple discharge sites, with each site discharging randomly and independently. The cell in Fig. 1 had nine active sites, of which four are shown. The four regions of interest drawn on a brightfield image of the cell (Fig. 1B) reveal their broad cellular distribution. Some sites were more active than others, so-called "frequent discharge sites," consistent with Ca²⁺ spark sites in other smooth muscles (16, 48). The first and second events at site 1 (blue trace) were accompanied only by slight increases in Ca²⁺ at a neighboring site 2 (red trace), representing passive spread of Ca²⁺ but not a regenerative response that could give rise to further sparks or waves. The solitary nature of these events can be assessed in the supplemental movie 1 available online at the AJP-Cell website. The first event is shown on an expanded time base (Fig. 1C), illustrating the rapid rise and slower decay that was characteristic of these events. The pseudo-colored intensity profiles (Fig. 1D) show the spatially restricted nature of the rise of Ca²⁺ concentration.

Regional changes of Ca²⁺ have never been previously demonstrated in CCSMCs, therefore, we first characterized these events and established their underlying cause. Images were acquired at 40–60 Hz with a typical recording duration of 35 s. There was no noticeable change in event frequency or amplitude over the 35-s duration of recording (Fig. 2A). Moreover, when consecutive files from one cell were acquired, there was no marked change in frequency, amplitude, or normalized spark area between the first and second recordings, and this was independent of the interval between recordings (n = 21 cells). Thus our wide-field fluorescence imaging system enables stable, relatively long duration recordings of Ca²⁺ levels in our cells.

View larger version (35K): [in this window] [in a new window]   Fig. 2. Temporal characteristics of Ca2+ release events. A: time course of the change of fluorescence intensity recorded in two consecutive sessions from the same cell, 4 min apart. Each colored trace is from a single Ca2+ release site. Stable consecutive recording of up to 35 s each were made under control conditions without appreciable changes in frequency or amplitude. A portion of the traces (boxed region) is expanded in B. B: Ca2+ events from three Ca2+ release sites show multiple release patterns either occurring in isolation or in rapid succession without returning to baseline. C: amplitude distribution of 169 isolated Ca2+ events recorded from 21 cells. Mean of the amplitude distribution was 42 ± 2% (arrow). D: decay of Ca2+ events could be well fit by single or double-exponential curves. The events shown are from the same site in a single cell. The average time constants were 170 ± 14 ms for a single exponential (n = 59), and 106 ± 7 and 900 ± 60 ms for a double exponential (n = 75). E: Ca2+ events of which its decay was best fit with a single exponential (first-order decay) were significantly smaller in amplitude than those best fit with a double exponential (second-order decay) (33 ± 3% intensity compared with 53 ± 3%, n = 59 and 75, respectively, *P < 0.001).

We also considered whether maintaining isolated cells for a period of time in high K⁺ solution might lead to altered Ca²⁺ handling. This might be evident as a change in spark characteristics or appearance of Ca²⁺ waves, as reported in cardiac myocytes (10, 39). We compared the frequency and amplitude of sparks and the propensity for propagated calcium waves in cells perfused with normal Na⁺-Ringer for up to 20 min with a group of cells perfused in normal Na⁺-Ringer for 60–80 min. We found no statistical difference in the frequency or amplitude of sparks. (Average frequency of 1.3 ± 0.5 Hz per cell at 10–20 min and 0.9 ± 0.5 Hz at 60–80 min, n = 6 and 8, respectively. The mean amplitude was 36 ± 19% at 10–20 min and 41 ± 3% at 60–80 min, n = 6 and 7, respectively, P > 0.05, t-test). In no cases were Ca²⁺ waves evident, arguing against altered calcium handling such as Ca²⁺ overload.

The decay of the Ca²⁺ events was well fit by exponential curves; 44% were best fit by a single exponential and 56% by a double exponential (Fig. 2D, representing analysis of 134 events in 21 cells). Both types of decay were observed in a single cell and often from the same site (Fig. 2D). The average time constants were 170 ± 14 ms for a single exponential (n = 59) and 106 ± 7 and 900 ± 60 ms for a double exponential (n = 75). In the double exponential fits, both components contributed almost equally; the amplitude of the long component of decay made up 51 ± 4% of the entire amplitude of the event. Ca²⁺ events that were best fit with a single exponential were smaller than those best fit with a double exponential; an amplitude of 33 ± 3% for events fit with single exponentials compared with an amplitude of 53 ± 3% for those fit with double exponentials (Fig. 2E; P < 0.001). Two populations of Ca²⁺ events with single and double exponential decays were reported by Gordienko and Bolton (15), leading to the suggestion that these arose from Ca²⁺ release through both ryanodine and IP₃ receptors within the same domain.

The spread of solitary events was measured from baseline-subtracted images. The 50% peak intensity level was determined from multiple-line profiles across each Ca²⁺ event, using several angles of rotation. This enabled us to assess the full width at half-maximum (FWHM, Fig. 3A), as determined by others most often using line-scan mode in confocal microscopy (8, 16, 31, 32). With the full cell image available, we were able to extend the analysis of decay to two dimensions, as illustrated by the circle that is drawn around the event as a contour line for 50% peak intensity (Fig. 3B,b). The area of the circle represents the spread of the Ca²⁺ event, and its diameter is comparable to the FWHM measured using a Gaussian fit of a line scan (Fig. 3A). The distribution of spread diameters measured from 149 events was well fit by a single Gaussian profile (Fig. 3D). Mean spread at 50% peak intensity was 43 ± 3 µm² with mean diameter of 6.9 ± 0.2 µm. This area represents 8.6 ± 0.5% of the average planar cell area of 500 ± 25 µm² (n = 21). This is somewhat larger than previously reported in other smooth muscle cell types [a spread diameter of 1.4–4.0 µm (8, 31, 32, 46) and an area of 8–14 µm² (34, 41)].

Ca²⁺ sparks in corpus cavernosum cells. Controversy exists in the literature concerning the source of the Ca²⁺ events in corpus cavernosum smooth muscle. Localized Ca²⁺ events due to release of Ca²⁺ from intracellular stores in some smooth muscles are reported to be due to release from RyR (20), whereas others report release by IP₃ receptors (4, 6). In corpus cavernosum cells, caffeine increased the frequency of Ca²⁺ events in a concentration-dependent manner (Fig. 4), with 0.5 mM caffeine increasing the frequency significantly to 180 ± 30% of control, in six of nine cells tested. In the remaining three cells, 0.5 mM caffeine caused a global rise in Ca²⁺ to 140 ± 40% intensity, which was distinguishable from a spark by its spatial spread and much slower decay to baseline.

View larger version (27K): [in this window] [in a new window]   Fig. 4. Caffeine enhances Ca2+ sparks and causes global rise of intracellular Ca2+. A: low concentration of caffeine (0.5 mM, applied for the time indicated by bar) caused increased frequency and amplitude of Ca2+ events. Change of fluorescence intensity is shown for 4 Ca2+ release sites, each in a different color. B: histogram summarizing the effect of low doses of caffeine on spark frequency. There was no significant change with 0.1 mM, but 0.5 mM caffeine significantly increased event frequency (n = 6 of 9 cells; *P < 0.05). C: 10 mM caffeine induced a large global rise of intracellular Ca2+, which was distinct from Ca2+ sparks before caffeine. After this global event, no sparks were visible for several minutes (representative of 5 cells). D: histogram summarizing the global rise elicited by caffeine. In 3 of 9 cells, 0.5 mM caffeine led to a rapid global Ca2+ transient. Pretreatment with ryanodine (10 µM) for 10 min drastically reduced the subsequent effects of caffeine (1 mM, n = 3). Online supplemental movie 2 illustrates the cell in C, with spontaneous Ca2+ sparks before caffeine followed by a global rise of Ca2+ and contraction.

View larger version (20K): [in this window] [in a new window]   Fig. 5. Ryanodine inhibits Ca2+ sparks in corpus cavernosum smooth muscle cells. Ca2+ sparks were monitored in cells before and after treatment with ryanodine (10 µM) for 5–10 min. A: time course of the changes in fluorescence intensity recorded at four sites, for a cell that exhibited 7 release sites in total. Ryanodine (10 µM) slowly inhibited Ca2+ spark frequency and amplitude. B: quantification of responses to ryanodine summarizing effects recorded in 18 cells. Spark frequency declined from control levels. Times below the columns are the center of bins of 2 min width. Spark area was determined as frequency x area under the spark, as described in METHODS. Ryanodine caused profound reduction in spark area.

View larger version (24K): [in this window] [in a new window]   Fig. 6. Blockade of the inositol 1,4,5 trisphosphate (IP3) receptor does not prevent Ca2+ sparks. The IP3 receptor antagonist 2-aminoethoxydiphenyl borate (2-APB, 100 µM) was applied to cells while basal Ca2+ events and agonist-evoked responses to phenylephrine (PE) were monitored. A: representative trace showing Ca2+ transients in a resting cell (one site shown for a cell with 4 active release sites). 2-APB was applied twice for 2 min each. Imaging was initiated 1 min after the start of 2-APB and confirmed that spark activity continued, albeit with reduced amplitude. To confirm the effectiveness of 2-APB, PE was applied as indicated, and the Ca2+ response was largely inhibited, confirming blockade of IP3 receptors. After washout of the bath for 8 min, Ca2+ sparks persisted and now PE caused a large, global rise of Ca2+. B: histograms showing the effect of 2-APB on spark frequency, amplitude, and proportion of sparks showing seconde-order exponential decay (n = 10). 2-APB caused no significant change in spark frequency, but amplitude was significantly reduced. Sparks could be either fit with a first-order or second-order exponential decay (see RESULTS). The proportion of sparks best fit by a second-order exponential decay was reduced in the presence of 2-APB (*P < 0.05). C: representative sparks, from the same release site, marked i, ii, and iii, in A, illustrating the effect of 2-APB on exponential decay and amplitude. The PE-induced Ca2+ transient recovered on wash out of 2-APB but spark amplitude and decay type did not (representative of 3 cells).

View larger version (25K): [in this window] [in a new window]   Fig. 7. Coupling of Ca2+ sparks in corpus cavernosum to transient outward and inward currents. Simultaneous whole cell patch-clamp recording and fluorescence imaging was carried out in dispersed corpus cells. A: biphasic outward then inward currents (STOICS) were recorded at –60 mV (top traces), and coincided precisely with the occurrence of Ca2+ sparks (bottom colored traces). Expanded view of STOICS and sparks (events labeled i and ii) are shown in B. Sparks are illustrated for 3 release sites (different colors) and at least 2 more were observed in this cell, one of which accounted for the unaccompanied STOIC in B, left. C: outward currents (STOCS) predominate at –30 mV and are closely coupled to sparks (bottom) from the same cell as in A. Note that at –30 mV Ca2+ spark and STOC frequency is increased compared with A.

View larger version (20K): [in this window] [in a new window]   Fig. 8. Increase in spark frequency with depolarization of the membrane. A: time course of Ca2+ sparks with change of membrane potential. Spark frequency increased on depolarization to less negative potentials and notably was accompanied by an increase in baseline Ca2+ levels. B: relationship of Ca2+ spark frequency to membrane potential. There was a consistent increase in spark frequency when membrane was depolarized from –60 to –40 mV and above (n = 5 to 14; P < 0.05). In addition, at potentials positive to –40 mV the baseline Ca2+ levels often increased to such an extent that individual events could not decay fully. This made it difficult to distinguish individual sparks; thus the variability in measured spark frequency was greater and smaller sparks may not be detected at more positive potentials. The dotted line is a fit of the Boltzmann equation to the data. C: nifedipine blocked the increase in Ca2+ spark frequency elicited by depolarization. D: to examine the coupling of Ca2+ sparks to KCa and ClCa currents, we plotted the ratio of current frequency to spark frequency against membrane potential. At potentials more negative than –40 mV, the ratio was not significantly different from unity, demonstrating close correspondence between currents and sparks. However, at –30 and –20 mV, we detected more STOC events than individual Ca2+ transients, likely due to the global rise of Ca2+ (masking individual events) and the voltage dependence of KCa channels.

At potentials positive to –40 mV baseline, Ca²⁺ levels increased and it became more difficult to distinguish the start of one spark from the start of another (Fig. 8, A and C). These factors contributed to an increase in frequency of spontaneous currents at more positive potentials that exceeded the increase in discrete Ca²⁺ sparks (Fig. 8D). Moreover, the depolarization-induced increase in baseline Ca²⁺ was sufficient to cause cell contraction in some cells (not shown).

STOCs are known to cause hyperpolarization of the smooth muscle membrane, but STICs are expected to cause depolarization, since activation of Cl_Ca currents in many smooth muscles contribute to contraction (8, 27, 47). STOICs may have a more complex effect on membrane potential due to the sequential activation of BK_Ca than Cl_Ca currents. We examined the effects of sparks on membrane potential in a CCSMC, comparing responses first under voltage clamp and then in current clamp. CCSMCs alternately hyperpolarized and then depolarized in response to sparks (Fig. 9), with biphasic changes of membrane potential observed in more than five cells. The direct role for Ca²⁺ sparks in eliciting biphasic oscillations of membrane potential have not been previously shown in any smooth muscle.

View larger version (28K): [in this window] [in a new window]   Fig. 9. Ca2+ sparks initiate biphasic changes in membrane potential. A: currents and sparks measured simultaneously at –60 mV. The currents generated are biphasic STOICs, resulting from activation of KCa currents (STOCS) followed by activation of ClCa currents (STICS). B: membrane potential measured simultaneously with Ca2+ sparks in the same cell, 15 min later. Note that each spark leads to biphasic change in potential: hyperpolarization followed closely by depolarization, consistent with activation first of KCa currents followed by ClCa currents.

View larger version (30K): [in this window] [in a new window]   Fig. 10. Role for extracellular Ca2+ in regulating the frequency of Ca2+ sparks. A: current recording of a cell held at –20 mV and bathed first in normal solution, then in zero Ca2+ extracellular solution, as indicated. STOCs continued in Ca2+-free solution, supporting the involvement of Ca2+ release from stores. However, STOC frequency was reduced from 2.3 Hz in control to 1.1 Hz in zero Ca2+ solution. The boxed region has been expanded and is shown with simultaneously recorded sparks below. B: simultaneous current (top) and spark (bottom) recordings for the cell in A. The control recording at left was collected earlier from the same cell. This cell had 4 active release sites, 2 of which are shown for clarity in the lefthand panel.

	DISCUSSION

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Comparison with other smooth muscles. Ca²⁺ spark frequency and amplitude in corpus cavernosum resembles that reported in other smooth muscles (36, 37, 41, 48); however, their spatiotemporal characteristics (spatial spread, rise, and decay times) are larger (9, 17, 41). Interestingly, the values for rise time and duration to half-maximum amplitude in visceral smooth muscle are longer and are comparable to those observed in the corpus cavernosum [trachea and bladder; rise times of 35–95 ms and duration to half-maximum amplitude of 112–150 ms, respectively (19, 38, 47)].

The frequency of sparks observed in CCSMCs was similar to that observed in other mammalian smooth muscles, although the number of release sites per cell was two to three times greater (6.4 sites vs. 1.8 to 2.9 sites in other muscles) (19, 36, 41). By contrast, in toad gastric muscle (45) the number of spark release sites in the unstimulated cell was as high as 42 per cell, although coupling between sparks and spontaneous currents was weak, with 21% of sparks not causing currents. Although differences in defining a release site (see Ref. 45) may account for some of the differences in numbers, there appears to be variations among smooth muscles. The effect of sparks on membrane potential is believed to depend largely on spark frequency and the strength of coupling between sparks and currents. Thus the physiological significance of the differences in the number of spark release sites between smooth muscle types is unclear and requires more investigation. In corpus cavernosum, the spread of each spark covered on average 9% of cell area, somewhat larger than the 1% reported in other vascular muscle types (34, 41). However, the there were no indications that the sparks showed active propagation within cells, apparent by triggering of new spark sites, as reported, for example, in Ca²⁺-overloaded cardiomyocytes (10, 39). In addition, we did not observe Ca²⁺ waves, another hallmark of propagating Ca²⁺ events in muscle cells.

Local calcium transients in corpus cavernosum smooth muscle are Ca²⁺ sparks. Pharmacological evidence supports the view that Ca²⁺ events in rat corpus cavernosum are due to release of Ca²⁺ through RyR. At 0.5 mM, caffeine increased spark frequency, whereas higher concentrations caused global Ca²⁺ transients accompanied by abolition of sparks. The caffeine sensitivity in these cells appears steep because caffeine at 0.1 mM caused no significant increase in spark frequency, whereas a third of cells at 0.5 mM caffeine responded with a large transient, results similar to those observed in other preparations (14, 31, 37). Ryanodine pretreatment reduced, but did not abolish, the caffeine-induced Ca²⁺ transient, which may be explained by the fact that either caffeine increases the open probability of RyRs so that block by ryanodine is relieved, or that ryanodine binds only to open RyR, leaving those receptors not involved in spark generation to be activated by caffeine (6, 15, 22). Indeed, a recent study of colonic smooth muscle cells has revealed that effects of ryanodine on calcium release from intracellular stores depends on previous activation of RyRs (30).

The IP₃ receptor inhibitor 2-APB did not reduce event frequency or the number of active release sites in rat CCSMCs. The reduction of amplitude we observed may be due to effects other than inhibition of the Ca²⁺ release receptor. We base this interpretation on the fact that blockade of the phenylephrine-induced global rise of Ca²⁺ by 2-APB was reversible, whereas the reduction in spark amplitude was not. Furthermore, we also acknowledge that 2-APB has multiple effects, including inhibition of store-operated Ca²⁺ entry and Ca²⁺ pumps (33), which could influence store Ca²⁺ levels. Our results contrast with those of Craven and colleagues (13), who found that 2-APB reduced both the amplitude and frequency of STICs in rabbit corpus cavernosum, leading to the suggestion that these currents were a result of IP₃-mediated Ca²⁺ release. Apart from a difference in species used (rabbit vs. rat), the reduced spark amplitude we observed in the presence of 2-APB may have compromised coupling of sparks to Ca²⁺-activated Cl^– channels. Alternatively, 2-APB may have a direct effect on rabbit Cl^– channels. We note that our studies of the interaction of ryanodine and 2-APB are complicated by our inability to carry out full paired experiments within individual cells. We faced this limitation due to the use of laser illumination, which limits the exposures possible on a single cell and to the fact that full recovery of spark frequency following depletion of stores by caffeine or phenylephrine can take at least 30 min.

In addition to the acknowledged role of IP₃ receptors in receptor-mediated activation of smooth muscles, cADP-ribose has emerged as an important regulator of smooth muscle tone (reviewed by Ref. 3). This Ca²⁺-mobilizing second messenger regulates the gating properties of RyR and is reported to enhance spark frequency in cardiac myocytes (29). However, it is not yet known whether cADP-ribose is involved in the genesis of "spontaneous events" such as Ca²⁺ sparks in smooth muscle. There is compelling evidence for the involvement of FKBP12.6 complexes with RyR2 in regulating both spontaneous and receptor-mediated Ca²⁺ release in tracheal smooth muscle (40).

The decay of the Ca²⁺ events was well fit by an exponential; 44% were best fit by a single exponential and 56% were best fit by a double exponential. The nature of the slower time constant, which has also been observed in vascular myocytes (15, 34), and why it was observed in a subpopulation of sparks in our tissue, is unknown. Two populations of Ca²⁺ events with single and double-exponential decays were observed by Gordienko and Bolton (15), and they suggested that these arose from Ca²⁺ release through both RyR and IP₃ receptors within the same domain. Several observations support the interpretation that the Ca²⁺ events reported here do not involve combined release through RyR and IP₃ receptors. First, both types of fits occurred in single cells and from different events at a single site. If the proximity of the site to IP₃ receptors affects the decay of Ca²⁺ events, then the type of decay might be consistent within a site. Second, 2-APB did not alter the frequency or number of active release sites. Moreover, the reduction in the phenylephrine response was reversible, whereas the reduction of spark amplitude and proportion of double-exponential fits was not. Finally, there was no evidence in the distribution of spark spread, rise time, or duration of half-maximum amplitude of two populations of sparks. Although Ca²⁺ events best fit with a single exponential were smaller than those best fit with a double exponential, there was only a small suggestion of two peaks in the amplitude distribution. The two populations of sparks observed in portal vein myocytes were quite distinct (15). On the other hand, the amplitude of the sparks fit with double and single exponentials in corpus cavernosum (55% and 33%, respectively) were within the range of spark amplitudes reported elsewhere (36, 37, 41, 48). These data lead us to conclude that the Ca²⁺ events in CCSMCs do not involve significant release through IP₃ receptors.

Coupling between sparks and spontaneous transient currents. The presence of spontaneous transient currents in corpus cavernosum smooth muscle cells was first demonstrated by Karkanis et al. (25). In that paper, it was hypothesized that Ca²⁺ sparks were the underlying stimulus for these currents. The data reported here resolve this matter and demonstrate close coupling between sparks and spontaneous currents. At the physiological membrane potential of around –40 mV, a ratio of current transients to spark frequency was not significantly different from unity. The resulting biphasic currents exhibit outward K⁺ current preceding inward Cl^– current. The factors accounting for this consistent pattern of activation are not known but are suggested to include spatial distribution of channels adjacent to release sites or to differences in kinetics of activation and calcium sensitivity of the channels (for discussion see Refs. 7 and 47). Coupling between spontaneous currents and sparks appears to vary between smooth muscle cell types. In porcine and human cerebral arteries, feline esophagus, and Bufo marinus gastric smooth muscle cells coupling at –40 mV is weak, and a significant number of Ca²⁺ sparks do not activate spontaneous currents (12, 26, 41, 46). On the other hand, sparks in rat cerebral arteries are strongly coupled and essentially all sparks activate a transient current (9, 12, 34).

The mechanisms that determine coupling and the reasons for weak coupling are unclear. It has been suggested that the larger distances between the spark release sites and plasma membrane leads to uncoupling, but this remains to be resolved (26, 45). On the other hand, there is some evidence to suggest that coupling level is dependent on the Ca²⁺ sensitivity of the K_Ca channel (9, 28, 35), and it remains to be shown whether the -subunit of K_Ca channel, which determines Ca²⁺ sensitivity, is different between smooth muscle with weak coupling and those with stronger coupling such as cerebral artery and corpus cavernosum smooth muscle cells.

The voltage dependence of sparks and sensitivity to dihydropyridine blockers provide indirect evidence of depolarization activated Ca²⁺ influx in corpus cavernosum smooth muscle cells. The increase in spark frequency observed may reflect Ca²⁺-induced Ca²⁺ release or the replenishment of store Ca²⁺ levels and the indirect enhancement of store release. Depolarization often increased spark frequency to such an extent that the end of one spark overlapped with the beginning of another, making assessment of coupling more difficult. Nevertheless, we demonstrate that membrane potential influences sparks (Fig. 8B) and that sparks in turn give rise to changes of membrane potential (Fig. 9B), thus revealing a dynamic interplay between membrane potential and Ca²⁺ stores.

Role of Ca²⁺ sparks in corpus cavernosum. There is evidence that sparks play an important role in the maintenance of corpus cavernosum smooth muscle tone and the regulation of penile erection. We have shown that sparks activate Ca²⁺-activated Cl^– channels and Ca²⁺-activated K⁺ channels, both of which play important roles in penile erection (25, 42). Karkanis et al. (25) used in vivo recording of erection in living rats to demonstrate that intercavernosal pressure is increased by infusion of chloride channel blockers, enhancing and prolonging erection. Elegant genetic studies by Werner et al. (42) revealed that mice lacking the pore-forming subunit of the Ca²⁺-activated K⁺ channel suffer from erectile dysfunction. Physiological regulation of sparks may therefore have relevance in the erectile process.

In summary, we have shown for the first time that corpus cavernosum cells exhibit Ca²⁺ sparks, which result from Ca²⁺ release from intracellular stores. Ca²⁺ sparks are closely coupled to activation of K_Ca and Cl_Ca currents, and spark frequency is subject to physiological regulation by voltage-dependent Ca²⁺ influx. We speculate that modulation of spark activity will have an important role in regulating penile erection by affecting corpus cavernosum smooth muscle tone.

	GRANTS

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We gratefully acknowledge support of The Canadian Institutes of Health Research (Grant MOP 10019) and the Canada Foundation for Innovation (Project 5651) for support of these studies.

	ACKNOWLEDGMENTS

 
We thank Dr. Ronghua ZhuGe (University of Massachusetts Medical School) for helpful comments on the paper and Mike Kovach (PTI) for valuable assistance in designing and optimizing the imaging apparatus.

	FOOTNOTES

 

Address for reprint requests and other correspondence: S. M. Sims, Dept. of Physiology and Pharmacology, Schulich School of Medicine & Dentistry, The Univ. of Western Ontario, London, Ontario, Canada N6A 5C1 (e-mail: Stephen.sims{at}schulich.uwo.ca)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

	REFERENCES

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
1. Andersson KE. Pharmacology of penile erection. Pharmacol Rev 53: 417–450, 2001.[Abstract/Free Full Text]

2. Ay B, Iyanoye A, Sieck GC, Prakash YS, Pabelick CM. Cyclic nucleotide regulation of store-operated Ca²⁺ influx in airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 290: L278–L283, 2006.[Abstract/Free Full Text]

3. Bai N, Lee HC, Laher I. Emerging role of cyclic ADP-ribose (cADPR) in smooth muscle. Pharmacol Ther 105: 189–207, 2005.[CrossRef][Web of Science][Medline]

4. Bayguinov O, Hagen B, Sanders KM. Muscarinic stimulation increases basal Ca²⁺ and inhibits spontaneous Ca²⁺ transients in murine colonic myocytes. Am J Physiol Cell Physiol 280: C689–C700, 2001.[Abstract/Free Full Text]

5. Berridge MJ, Bootman MD, Roderick HL. Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol 4: 517–529, 2003.[CrossRef][Web of Science][Medline]

6. Boittin FX, Macrez N, Halet G, Mironneau J. Norepinephrine-induced Ca²⁺ waves depend on InsP₃ and ryanodine receptor activation in vascular myocytes. Am J Physiol Cell Physiol 277: C139–C151, 1999.[Abstract/Free Full Text]

7. Bolton TB. Calcium events in smooth muscles and their interstitial cells; physiological roles of sparks. J Physiol 570: 5–11, 2006.[Abstract/Free Full Text]

8. Bonev AD, Jaggar JH, Rubart M, Nelson MT. Activators of protein kinase C decrease Ca²⁺ spark frequency in smooth muscle cells from cerebral arteries. Am J Physiol Cell Physiol 273: C2090–C2095, 1997.[Abstract/Free Full Text]

9. Brenner R, Perez GJ, Bonev AD, Eckman DM, Kosek JC, Wiler SW, Patterson AJ, Nelson MT, Aldrich RW. Vasoregulation by the beta1 subunit of the calcium-activated potassium channel. Nature 407: 870–876, 2000.[CrossRef][Medline]

10. Cheng H, Lederer MR, Lederer WJ, Cannell MB. Calcium sparks and [Ca²⁺]_i waves in cardiac myocytes. Am J Physiol Cell Physiol 270: C148–C159, 1996.[Abstract/Free Full Text]

11. Cheng H, Lederer WJ, Cannell MB. Calcium sparks: elementary events underlying excitation-contraction coupling in heart muscle. Science 262: 740–744, 1993.[Abstract/Free Full Text]

12. Cheranov SY, Jaggar JH. Sarcoplasmic reticulum calcium load regulates rat arterial smooth muscle calcium sparks and transient K_Ca currents. J Physiol 544: 71–84, 2002.[Abstract/Free Full Text]

13. Craven M, Sergeant GP, Hollywood MA, McHale NG, Thornbury KD. Modulation of spontaneous Ca²⁺-activated Cl^– currents in the rabbit corpus cavernosum by the nitric oxide-cGMP pathway. J Physiol 556: 495–506, 2004.[Abstract/Free Full Text]

14. Gollasch M, Wellman GC, Knot HJ, Jaggar JH, Damon DH, Bonev AD, Nelson MT. Ontogeny of local sarcoplasmic reticulum Ca²⁺ signals in cerebral arteries: Ca²⁺ sparks as elementary physiological events. Circ Res 83: 1104–1114, 1998.[Abstract/Free Full Text]

15. Gordienko DV, Bolton TB. Crosstalk between ryanodine receptors and IP₃ receptors as a factor shaping spontaneous Ca²⁺-release events in rabbit portal vein myocytes. J Physiol 542: 743–762, 2002.[Abstract/Free Full Text]

16. Gordienko DV, Bolton TB, Cannell MB. Variability in spontaneous subcellular calcium release in guinea-pig ileum smooth muscle cells. J Physiol 507: 707–720, 1998.[Abstract/Free Full Text]

17. Gordienko DV, Greenwood IA, Bolton TB. Direct visualization of sarcoplasmic reticulum regions discharging Ca²⁺ sparks in vascular myocytes. Cell Calcium 29: 13–28, 2001.[CrossRef][Web of Science][Medline]

18. Hagen BM, Bayguinov O, Sanders KM. B₁-subunits are required for regulation of coupling between Ca²⁺ transients and Ca²⁺-activated K⁺ (BK) channels by protein kinase C. Am J Physiol Cell Physiol 285: C1270–C1280, 2003.[Abstract/Free Full Text]

19. Herrera GM, Heppner TJ, Nelson MT. Voltage dependence of the coupling of Ca²⁺ sparks to BK_Ca channels in urinary bladder smooth muscle. Am J Physiol Cell Physiol 280: C481–C490, 2001.[Abstract/Free Full Text]

20. Jaggar JH, Porter VA, Lederer WJ, Nelson MT. Calcium sparks in smooth muscle. Am J Physiol Cell Physiol 278: C235–C256, 2000.[Abstract/Free Full Text]

21. Jaggar JH, Stevenson AS, Nelson MT. Voltage dependence of Ca²⁺ sparks in intact cerebral arteries. Am J Physiol Cell Physiol 274: C1755–C1761, 1998.[Abstract/Free Full Text]

22. Janiak R, Wilson SM, Montague S, Hume JR. Heterogeneity of calcium stores and elementary release events in canine pulmonary arterial smooth muscle cells. Am J Physiol Cell Physiol 280: C22–C33, 2001.[Abstract/Free Full Text]

23. Janssen LJ, Sims SM. Acetylcholine activates non-selective cation and chloride conductances in canine and guinea-pig tracheal myocytes. J Physiol 453: 197–218, 1992.[Abstract/Free Full Text]

24. Ji G, Feldman ME, Greene KS, Sorrentino V, Xin HB, Kotlikoff MI. RYR2 proteins contribute to the formation of Ca²⁺ sparks in smooth muscle. J Gen Physiol 123: 377–386, 2004.[Abstract/Free Full Text]

25. Karkanis T, DeYoung L, Brock GB, Sims SM. Ca²⁺-activated Cl^– channels in corpus cavernosum smooth muscle: a novel mechanism for control of penile erection. J Appl Physiol 94: 301–313, 2003.[Abstract/Free Full Text]

26. Kirber MT, Etter EF, Bellve KA, Lifshitz LM, Tuft RA, Fay FS, Walsh JV, Fogarty KE. Relationship of Ca²⁺ sparks to STOCs studied with 2D and 3D imaging in feline oesophageal smooth muscle cells. J Physiol 531: 315–327, 2001.[Abstract/Free Full Text]

27. Large WA, Wang Q. Characteristics and physiological role of the Ca²⁺-activated Cl^– conductance in smooth muscle. Am J Physiol Cell Physiol 271: C435–C454, 1996.[Abstract/Free Full Text]

28. Li A, Adebiyi A, Leffler CW, Jaggar JH. K_Ca channel insensitivity to Ca²⁺ sparks underlies fractional uncoupling in newborn cerebral artery smooth muscle cells. Am J Physiol Heart Circ Physiol 291: H1118–H1125, 2006.[Abstract/Free Full Text]

29. Lukyanenko V, Gyorke S. Ca²⁺ sparks and Ca²⁺ waves in saponin-permeabilized rat ventricular myocytes. J Physiol 521: 575–585, 1999.[Abstract/Free Full Text]

30. MacMillan D, Chalmers S, Muir TC, McCarron JG. IP₃-mediated Ca²⁺ increases do not involve the ryanodine receptor, but ryanodine receptor antagonists reduce IP₃-mediated Ca²⁺ increases in guinea-pig colonic smooth muscle cells. J Physiol 569: 533–544, 2005.[Abstract/Free Full Text]

31. Mironneau J, Arnaudeau S, Macrez-Lepretre N, Boittin FX. Ca²⁺ sparks and Ca²⁺ waves activate different Ca²⁺-dependent ion channels in single myocytes from rat portal vein. Cell Calcium 20: 153–160, 1996.[CrossRef][Web of Science][Medline]

32. Nelson MT, Cheng H, Rubart M, Santana LF, Bonev AD, Knot HJ, Lederer WJ. Relaxation of arterial smooth muscle by calcium sparks. Science 270: 633–637, 1995.[Abstract/Free Full Text]

33. Peppiatt CM, Collins TJ, Mackenzie L, Conway SJ, Holmes AB, Bootman MD, Berridge MJ, Seo JT, Roderick HL. 2-Aminoethoxydiphenyl borate (2-APB) antagonises inositol 1,4,5-trisphosphate-induced calcium release, inhibits calcium pumps and has a use-dependent and slowly reversible action on store-operated calcium entry channels. Cell Calcium 34: 97–108, 2003.[Web of Science][Medline]

34. Perez GJ, Bonev AD, Patlak JB, Nelson MT. Functional coupling of ryanodine receptors to K_Ca channels in smooth muscle cells from rat cerebral arteries. J Gen Physiol 113: 229–238, 1999.[Abstract/Free Full Text]

35. Pluger S, Faulhaber J, Furstenau M, Lohn M, Waldschutz R, Gollasch M, Haller H, Luft FC, Ehmke H, Pongs O. Mice with disrupted BK channel beta1 subunit gene feature abnormal Ca²⁺ spark/STOC coupling and elevated blood pressure. Circ Res 87: E53–E60, 2000.[Web of Science][Medline]

36. Pozo MJ, Perez GJ, Nelson MT, Mawe GM. Ca²⁺ sparks and BK currents in gallbladder myocytes: role in CCK-induced response. Am J Physiol Gastrointest Liver Physiol 282: G165–G174, 2002.[Abstract/Free Full Text]

37. Remillard CV, Zhang WM, Shimoda LA, Sham JS. Physiological properties and functions of Ca²⁺ sparks in rat intrapulmonary arterial smooth muscle cells. Am J Physiol Lung Cell Mol Physiol 283: L433–L444, 2002.[Abstract/Free Full Text]

38. Sieck GC, Kannan MS, Prakash YS. Heterogeneity in dynamic regulation of intracellular calcium in airway smooth muscle cells. Can J Physiol Pharmacol 75: 878–888, 1997.[CrossRef][Web of Science][Medline]

39. Trafford AW, Lipp P, O'Neill SC, Niggli E, Eisner DA. Propagating calcium waves initiated by local caffeine application in rat ventricular myocytes. J Physiol 489: 319–326, 1995.[Abstract/Free Full Text]

40. Wang YX, Zheng YM, Mei QB, Wang QS, Collier ML, Fleischer S, Xin HB, Kotlikoff MI. FKBP12.6 and cADPR regulation of Ca²⁺ release in smooth muscle cells. Am J Physiol Cell Physiol 286: C538–C546, 2004.[Abstract/Free Full Text]

41. Wellman GC, Nathan DJ, Saundry CM, Perez G, Bonev AD, Penar PL, Tranmer BI, Nelson MT. Ca²⁺ sparks and their function in human cerebral arteries. Stroke 33: 802–808, 2002.[Abstract/Free Full Text]

42. Werner ME, Zvara P, Meredith AL, Aldrich RW, Nelson MT. Erectile dysfunction in mice lacking the large-conductance calcium-activated potassium (BK) channel. J Physiol 567: 545–556, 2005.[Abstract/Free Full Text]

43. Williams BA, Liu C, Deyoung L, Brock GB, Sims SM. Regulation of intracellular Ca²⁺ release in corpus cavernosum smooth muscle: synergism between nitric oxide and cGMP. Am J Physiol Cell Physiol 288: C650–C658, 2005.[Abstract/Free Full Text]

44. Yao Y, Choi J, Parker I. Quantal puffs of intracellular Ca²⁺ evoked by inositol trisphosphate in Xenopus oocytes. J Physiol 482: 533–553, 1995.[Abstract/Free Full Text]

45. Zhuge R, Fogarty KE, Baker SP, McCarron JG, Tuft RA, Lifshitz LM, Walsh JV Jr. Ca²⁺ spark sites in smooth muscle cells are numerous and differ in number of ryanodine receptors, large-conductance K⁺ channels, and coupling ratio between them. Am J Physiol Cell Physiol 287: C1577–C1588, 2004.[Abstract/Free Full Text]

46. ZhuGe R, Fogarty KE, Tuft RA, Lifshitz LM, Sayar K, Walsh JV Jr. Dynamics of signaling between Ca²⁺ sparks and Ca²⁺- activated K⁺ channels studied with a novel image-based method for direct intracellular measurement of ryanodine receptor Ca²⁺ current. J Gen Physiol 116: 845–864, 2000.[Abstract/Free Full Text]

47. ZhuGe R, Sims SM, Tuft RA, Fogarty KE, Walsh JV Jr. Ca²⁺ sparks activate K⁺ and Cl^– channels, resulting in spontaneous transient currents in guinea-pig tracheal myocytes. J Physiol 513: 711–718, 1998.[Abstract/Free Full Text]

48. ZhuGe R, Tuft RA, Fogarty KE, Bellve K, Fay FS, Walsh JV Jr. The influence of sarcoplasmic reticulum Ca²⁺ concentration on Ca²⁺ sparks and spontaneous transient outward currents in single smooth muscle cells. J Gen Physiol 113: 215–228, 1999.[Abstract/Free Full Text]

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