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MUSCLE CELL BIOLOGY AND CELL MOTILITY

Two-step Ca²⁺ intracellular release underlies excitation-contraction coupling in mouse urinary bladder myocytes

¹Department of Molecular and Cellular Pharmacology, Graduate School of Pharmaceutical Sciences, Nagoya City University, Nagoya; ²Department of Pharmacology, Faculty of Medicine, Toyama Medical and Pharmaceutical University, Toyama; and ³Cell Signaling & Ion Channel Research Group, Cellular Pharmacology, School of Pharmacy, Aichigakuin University, Nagoya, Japan

Submitted 12 August 2005 ; accepted in final form 19 September 2005

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
The relative contributions of Ca²⁺-induced Ca²⁺ release (CICR) versus Ca²⁺ influx through voltage-dependent Ca²⁺ channels (VDCCs) to excitation-contraction coupling has not been defined in most smooth muscle cells (SMCs). The present study was undertaken to address this issue in mouse urinary bladder (UB) smooth muscle cells (UBSMCs). Confocal Ca²⁺ images were obtained under voltage- or current-clamp conditions. When UBSMCs were activated by a 30-ms depolarization to 0 mV, intracellular Ca²⁺ concentration ([Ca²⁺]_i) increased in several small, discrete areas just beneath the cell membrane. These Ca²⁺ "hot spots" then spread slowly through the myoplasm as Ca²⁺ waves, which continued even after repolarization. Shorter depolarizations (5 ms) elicited only a few Ca²⁺ sparks, which declined quickly. The number of Ca²⁺ sparks, or hot spots, was closely related to the depolarization duration in the range of 5–20 ms. There was an apparent threshold depolarization duration of 10 ms within which to induce enough Ca²⁺ transients to spread globally and then induce a contraction. Application of 100 µM ryanodine to the pipette solution did not change the resting [Ca²⁺]_i or the VDCC current, but it did abolish Ca²⁺ hot spots elicited by depolarization. Application of 3 µM xestospongin C reduced ACh-induced Ca²⁺ release but did not affect depolarization-induced Ca²⁺ events. The addition of 100 µM ryanodine to tissue segments markedly reduced the amplitude of contractions triggered by direct electrical stimulation. In conclusion, global [Ca²⁺]_i rise triggered by a single action potential is not due mainly to Ca²⁺ influx through VDCCs but is attributable to the subsequent two-step CICR.

Ca²⁺-induced Ca²⁺ release; Ca²⁺-activated K⁺ current; voltage-dependent Ca²⁺ channel

In contrast to this functional coupling between RyR and BK channels, the experimental analysis of the contribution of Ca²⁺-induced Ca²⁺ release (CICR) via RyR activation to contraction and excitation-contraction (E-C) coupling in SMCs remains incomplete. Initial experiments suggested that the Ca²⁺ requirement for CICR in SMCs was too high to participate in contraction (24). In portal vein myocytes, Ca²⁺ influx through VDCCs, as opposed to CICR, was suggested to be the major factor responsible for elevating [Ca²⁺]_i to induce a contraction (36). On the other hand, an essential contribution of CICR to E-C coupling has been suggested in urinary bladder (UB) (7, 13, 19, 31), vas deferens (31), ureter (6), and coronary artery (14). In UB myocytes from the guinea pig, the increase in [Ca²⁺]_i upon depolarization was substantially reduced by 10 µM ryanodine, suggesting an important contribution of CICR via RyRs in E-C coupling in this type of SMC (13).

Even in UB smooth muscle cells (UBSMCs), however, the importance of a functional contribution of CICR to E-C coupling and contraction has been assessed differently by different research groups. Collier et al. (7) reported that the coupling of VDCC and RyR in UBSMCs of the rabbit is weak and may not be effective in inducing contraction. The application of 10 or 50 µM ryanodine did not reduce, but instead enhanced, spontaneous contractions in guinea pig UB tissue preparations (18, 23). In contrast, Buckner et al. (5) reported that 10 µM ryanodine suppressed spontaneous contractions in the UB of the pig by 50%.

The present study was undertaken in an attempt to resolve the contribution of RyR and CICR to E-C coupling in UBSMCs. Our results, which were obtained using simultaneous measurement of fast confocal Ca²⁺ imaging and membrane currents, clearly show that two distinct steps of CICR are essential for E-C coupling triggered by an action potential in UBSMCs of the mouse.

	MATERIALS AND METHODS

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Cell isolation. C57BL/6 mice (male and female, 8–15 wk old, and 20–30 g body wt) were used in the experiments. Single SMCs were enzymatically isolated from the UB using a previously described method (29) with slight modification. All experiments were performed in accordance with the guiding principles for the care and use of laboratory animals of the Science and International Affairs Bureau of the Japanese Ministry of Education, Science, Sports, and Culture and with the approval of the Ethics Committee at Nagoya City University. In brief, mice were anesthetized with ether and killed by performing exsanguination. The UB was dissected out and freed from other tissues in Ca²⁺-free Krebs solution. The tissue was immersed in Ca²⁺-free Krebs solution for 30–60 min in a test tube at 37°C. Subsequently, the solution was replaced with Ca²⁺-free Krebs solution containing 0.2–0.3% collagenase (Amano Enzyme, Nagoya, Japan). After 10- to 15-min treatment, the solution was replaced with Ca²⁺-free, collagenase-free Krebs solution. The tissue was gently triturated using a glass pipette to isolate cells. At the start of each experiment, a few drops of the cell suspension were put into a recording chamber. The bath was continuously perfused with HEPES-buffered solution at a flow rate of 5 ml·min^–1.

Solutions. Krebs solution contained (in mM) 112 NaCl, 4.7 KCl, 2.2 CaCl₂, 1.2 MgCl₂, 25 NaHCO₃, 1.2 KH₂PO₄, and 14 glucose. Ca²⁺-free Krebs solution was prepared by omitting Ca²⁺ from the Krebs solution. Standard and Ca²⁺-free Krebs solutions were equilibrated with a 95% O₂-5% CO₂ mixture. For electrophysiological recording, HEPES-buffered solution of the following composition was used as the external solution (in mM): 137 NaCl, 5.9 KCl, 2.2 CaCl₂, 1.2 MgCl₂, 14 glucose, and 10 HEPES. The pH was adjusted to 7.4 with NaOH. For simultaneous recordings of Ca²⁺ current (I_Ca) and [Ca²⁺]_i, 30 mM tetraethyl ammonium was substituted for equimolar NaCl in the HEPES-buffered solution, and 1 mM 4-aminopyridine (4-AP) was added to the solution.

The pipette solution contained (in mM) 140 KCl, 1 MgCl₂, 2 Na₂ATP, 10 HEPES, and 0.1 fluo-4 or indo-1. The pH of this solution was adjusted to 7.2 with KOH. To examine the effects of 100 µM extracellular ryanodine on I_Ca (see Fig. 4B, right), a pipette solution containing the following composition was used (in mM): 120 CsCl, 20 tetraethylammonium (TEA), 1 MgCl₂, 10 HEPES, 5 EGTA, and 2 Na₂ATP. The pH of the solution was adjusted to 7.2 with CsOH.

View larger version (55K): [in this window] [in a new window]   Fig. 4. Effects of 100 µM ryanodine on Ca2+ mobilization during depolarization. To examine effects of 100 µM ryanodine on intracellular Ca2+ mobilization during depolarization under voltage-clamp conditions, cells were loaded with 100 µM ryanodine and fluo-4 from a recording pipette (asterisks in A and B). Measurements were started after 5 min from the formation of whole cell patch-clamp mode. A: [Ca2+]i images (a) and time courses of inward currents and F/F0 ratio (b) from control cells. Cells were depolarized from –60 to 0 mV for 30 ms in the presence of 30 mM tetraethylammonium (TEA) and 1 mM 4-aminopyridine (4-AP) in the bathing solution to block K+ currents. Numbers above images in Aa indicate time before or after start of depolarization. Arrows at 26.88 ms indicate Ca2+ hot spot sites in control cells. Arrows in Ba in a ryanodine-loaded cell indicate sites at which some [Ca2+]i increase was observed. Inward currents are indicated by black dots in Ab and Bb. F/F0 ratio from sites indicated by arrows and in Aa and Ba is indicated by red squares and green circles in Ab and Bb, respectively. F/F0 ratio from whole cell area is indicated by blue triangles. B: [Ca2+]i images (a) and time courses of inward currents and F/F0 ratio (b) from ryanodine-loaded cell. C: surface plots (3-dimensional images) were constructed from recordings at 26.88 ms from control and ryanodine-loaded cells shown in A. Lines (x-y and x'-y') indicate the cross sections of the fluorescence profiles shown in D. D: fluorescence profiles of [Ca2+]i images of control (a) and ryanodine-loaded cells (b) at 26.88, 92.88, and 224.88 ms.

Measurement of fluo-4 and indo-1 signals from single myocytes. Ca²⁺ images were obtained using a fast laser-scanning confocal microscope (RCM 8000; Nikon, Tokyo, Japan) and Ratio3 software (Nikon) in the same manner as reported previously (31). Each selected myocyte was loaded with 100 µM fluo-4 or indo-1 by diffusion from the recording pipette. For measurements in which fluo-4 was used, 488-nm excitation from an argon ion laser was delivered through a water-immersion lens objective (x40 magnification, 1.15 numerical aperture; Nikon Fluo). Emitted light of >515 nm was detected using a photomultiplier. Fluorescence intensity (F) in a selected area was measured as an average of pixels included within the area. Data are shown as F/F₀ ratios in which F₀ is the average fluorescence intensity of five images of the whole cell area during resting conditions and F is the increase in fluorescence intensity from F₀. It took 33 ms to scan one full frame (512 x 512 pixels). Using 1/2- and 1/4-band scan modes, we obtained frames that corresponded to areas of 170 x 55 µm or 170 x 27.5 µm every 16.5 or 8.25 ms, respectively. For indo-1 measurements, excitation of 355 nm and emission at 405 and 485 nm, respectively, were applied and detected, and fluorescence images were obtained at 66-ms intervals. The resolution of the microscope was 0.4 x 0.3 x 1.2 µm (x, y, and z-axes, respectively). The confocal plane through the cell was set so that the width of the cell was largest 2–3 µm from its lowest point. Recordings were started 5 min after rupturing the patch membrane to allow time for the Ca²⁺ indicator and drugs to diffuse into the cell.

Ca²⁺ images were stored on an optical disk cartridge (LM-A410; Panasonic, Osaka, Japan) using a rewritable optical disk recorder (LQ-4100A; Panasonic). The images on the optical disks were replayed later and analyzed using Ratio3 software (Nikon). Some analyses were performed using GLOBAL LAB Image software (Data Translation) on an IBM-AT-compatible computer. During the recording of fluorescence images, the cell shape was monitored and recorded on videotape using red or infrared light with a wavelength range of >600 nm and an infrared charge-coupled device video camera module (XC-77BB; Sony), which was attached to the microscope. Video image capture and analysis of cell shape changes were performed later on an IBM-AT-compatible computer using an A/D translation board (DT-55; Data Translation) and GLOBAL LAB Image software.

Calibration of [Ca²⁺]_i. [Ca²⁺]_i values were calculated from ratios of fluorescent signals of indo-1 at 405 nm compared with those at 485 nm (R) using the following equation (38):

where K_d is the dissociation constant of indo-1 (250 nM), S_f2 is fluorescence intensity at 485 nm without Ca²⁺, and S_b2 is fluorescence intensity with saturating Ca²⁺. R_max is R in the absence of Ca²⁺, and R_min is R in saturating Ca²⁺. R_max and R_min values were obtained using a method similar to that of Ganitkevich and Isenberg (12) in which we used indo-1-loaded UBSMCs under a whole cell patch clamp. At the end of some experiments, the cell membrane was made leaky by stepping the holding potential to –300 mV. After 2–3 min, the holding potential was set to 0 mV and then stepped to –300 mV for 200 ms at 500-ms intervals to maintain the myocyte membrane in a leaky state. R_max and R_min were obtained at a holding potential of 0 mV in 2.2 mM Ca²⁺-containing solution and in Ca²⁺-free solution containing 10 mM EGTA, respectively.

Measurement of cell volume and estimation of increase in [Ca²⁺]_i from Ca²⁺ influx. When [Ca²⁺]_i and I_Ca were recorded simultaneously, estimated increases in [Ca²⁺]_i were calculated using following equation (14, 36):

where – I_Cadt is total charge entry, F is the Faraday constant, and V is cell volume. I_Cadt was obtained by integration of the area under the I_Ca trace. V was calculated by assuming that the shape of UBSMCs consisted of two cones connected base to base. The V value calculated from length and width of resting UBSMCs was 12.8 ± 1.4 pl (n = 25).

Measurement of contraction from tissue preparation. After the UB was dissected and freed from the vesicle trigon, the ventral wall was opened longitudinally in Ca²⁺-free Krebs solution. The mucosal layer was then removed, and tissue segments 3–4 mm long and 0.8–1.2 mm wide were prepared. One end of the tissue segment was pinned to a rubber plate at the bottom of the organ bath (5 ml), and the other end was connected to a force transducer, which was domestically made (32). Segments were equilibrated at a resting load of 1 mN. To elicit contractions, a tissue segment was stimulated for 2 s using a train of 3-ms pulses of 300 mA at 5 Hz in Krebs solution that contained the following neurotransmitter antagonists (in µM): 1 atropine, 1 phentolamine, 1 propranolol, 1 TTX, and 10 suramin. Contractile responses were recorded using a strain gauge transducer on an ink-writing direct current servorecorder (Toa Electronics, Kobe, Japan). All of the experiments were performed at 36 ± 1°C. Measurement of twitch contractions was performed just before the addition of ryanodine and at the end of ryanodine treatment. The increase in resting tone induced by ryanodine was measured at the peak of the tone.

Statistics. Pooled data are means ± SE. Statistical significance between two or multiple groups was determined using Student's t-test or Scheffé's test after one-way ANOVA, respectively. P < 0.05 and 0.01 indicate statistical significance.

Drugs. Most pharmacological reagents were obtained from Sigma (St. Louis, MO). Ryanodine, xestospongin C, TEA-Cl^–, 4-AP, and CdCl₂ were purchased from Wako (Osaka, Japan), and EGTA, HEPES, and indo-1 were obtained from Dojin (Kumamoto, Japan). Fluo-4 was purchased from Molecular Probes (Eugene, OR).

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Effects of ryanodine on Ca²⁺ images during action potentials in UBSMCs. Ca²⁺ images obtained during and immediately after an action potential were obtained from single UBSMCs of the mouse using a fast laser-scanning confocal microscope in current-clamp mode (Fig. 1Aa). A myocyte was loaded with 100 µM fluo-4, which was added to a pipette filling solution. A 50-ms, 10-pA current injection elicited an action potential with an amplitude of 62.8 mV from a resting potential of –44.5 mV. An afterhyperpolatrization of 10.7 mV was also observed (Fig. 1Ab). The averaged resting membrane potential, action potential overshoot, and afterhyperpolarization values in UBSMCs were –50.1 ± 1.3, 15.0 ± 2.0, and 10.1 ± 1.0 mV (n = 9), respectively. The increase in [Ca²⁺]_i initially occurred at approximately the same time as the peak action potential 40 ms after the start of current injection. This change in [Ca²⁺]_i occurred exclusively at localized spots (termed Ca²⁺ "hot spots") within the cell (e.g., and in Fig. 1A). The averaged fluorescence intensity in Ca²⁺ hot spots (2 µm in diameter) and in the whole cell area were measured in each frame at an interval of 8.25 ms. The resulting changes in the ratio of F to F measured before depolarization (F/F₀) were plotted against time. The F/F₀ ratio in Ca²⁺ hot spots reached a peak at 100–150 ms (Fig. 1B) and then progressively declined. The rise of [Ca²⁺]_i spread slowly from these hot spots to other areas, and the F/F₀ ratio in the whole cell area reached peak values at 300 ms. The locations of initial Ca²⁺ hot spots were exactly the same in each cell when action potentials were elicited repetitively, even at intervals as long as 3–5 min (data not shown). Essentially identical images of Ca²⁺ dynamics, which started in some discrete local sites in superficial areas as Ca²⁺ hot spots and then spread slowly into the entire myoplasm, were obtained in response to action potentials in all cells in which Ca²⁺ images and action potentials were measured simultaneously (n = 4). After an action potential, cell length was reduced significantly beginning at 500 ms. Peak shortening (up to 20%) was recorded at 2–3 s, and relaxation developed within 10 s (data not shown).

View larger version (56K): [in this window] [in a new window]   Fig. 1. Effects of ryanodine on intracellular Ca2+ mobilization during an action potential in mouse urinary bladder (UB) smooth muscle cells (UBSMCs). An action potential was elicited by 50-ms current injection of 10 pA under current-clamp conditions. Intracellular Ca2+ concentration ([Ca2+]i) images were obtained every 8.25 ms using a fast laser-scanning confocal microscope. Each myocyte was loaded with 100 µM fluo-4 from a recording pipette. Aa: [Ca2+]i images during action potential. Time shown above each image indicates the time before (–) or after the start of the current injection. All images were subtracted by the average baseline image obtained before current injection. The color scale bar indicates the calibration of averaged fluorescence intensity (F) from whole cell area in resting conditions (F0) to the increase in fluorescence intensity (F) from F0 (F/F0 ratio). The edge of the myocyte is indicated by a contour line. Arrows and and asterisk at 43.38 ms indicate sites generating Ca2+ hot spots and the position of a recording pipette tip, respectively. Ab: time courses of membrane potential (solid line) and F/F0 ratios from hot spots and whole cell area (blue triangles) during an action potential. F/F0 values in hot spots, indicated by arrow (red squares in Ab) and arrow (green circles in Ab) in Aa, were obtained as averages in areas with diameters of 2 µm. Dotted lines indicate resting membrane potentials of –44.5 mV and 0 mV, respectively. Ba: effects of ryanodine on Ca2+ transients during action potential. The myocyte was loaded with 100 µM ryanodine from the recording pipette (asterisk). Bb: time courses of membrane potential (solid line) and F/F0 ratio from a local area ( arrow in Ba, green circles in Bb) and the whole cell area (blue triangles) during an action potential. The resting membrane potential was –28.9 mV.

Image analysis of Ca²⁺ mobilization during depolarization in UBSMCs. Figure 2Aa shows fluorescent images obtained every 8.25 ms from a cell that was depolarized under voltage-clamp conditions to 0 mV from a holding potential of –60 mV for preselected durations ranging from 5 to 30 ms. When the duration of the depolarization pulse was 5 ms, local Ca²⁺ increase occurred in only one spot () in subsarcolemmal areas. This [Ca²⁺]_i change did not spread to other areas as a Ca²⁺ spark (Fig. 2A). An increase in clamp pulse duration to 10 ms elicited an additional spot . When the cell was depolarized for 30 ms, the number of Ca²⁺ spots increased to about five in a single confocal plane. In addition, the Ca²⁺ spots spread to other areas and also increased global [Ca²⁺]_i. This induced a contraction that started 500 ms after repolarization (data not shown). It is notable that the Ca²⁺ increase did not occur uniformly along the sarcolemma but did appear in the same Ca²⁺ hot spot sites, such as and , with repetitive application of depolarization of different durations. Figure 2B shows changes in F/F₀ in hot spots ( and ) and global area corresponding to those measured from the images in Fig. 2A. At the duration of 5 ms, the rise in F/F₀ was observed only in hot spot but not in and global area. The F/F₀ ratio in global area was increased as the duration was lengthened to >10 ms. The time courses of global F/F₀ ratio were replotted and superimposed in Fig. 2C for a longer time. The marked increase in F/F₀ ratio occurred in an all-or-none manner as the depolarization duration was changed. The threshold duration for the switching from local to global Ca²⁺ events appeared to be between 5 and 10 ms in this particular cell. In addition, the time to peak global F/F₀ ratio also depended on the duration of depolarization.

View larger version (49K): [in this window] [in a new window]   Fig. 2. Intracellular Ca2+ mobilization during depolarization in UBSMCs. Intracellular Ca2+ mobilization due to myocyte depolarization was analyzed by changing the duration (5, 10, 15, 20, and 30 ms) under voltage-clamp conditions. A: [Ca2+]i images in a myocyte before, during, and after application of depolarization lasting 5, 10, 20, or 30 ms. UBSMCs were depolarized from a holding potential of –60 to 0 mV. Time shown above each image indicates time before or after start of depolarizing pulse. Arrows and asterisks at 10.4 ms indicate the sites generating Ca2+ hot spots and the position of the recording pipette tip, respectively. Images obtained during depolarization are indicated by red lines below sequential images. B: time course of membrane current (black dots) and F/F0 ratio due to hot spots indicated by arrow in A (red dots in B) and arrow in A (green dots in B) and whole cell area (blue dots in B) in separate trials using different pulse durations. C: time course of F/F0 ratio from whole cell area after depolarization shown in B superimposed for a longer time. Horizontal blocks from time 0 and corresponding vertical lines indicate period of voltage-clamp depolarizations. Note marked difference in F/F0 ratios after depolarization for 5 ms and those recorded for 10 ms or longer. Da: fluorescence profiles along x-y section (at –6.1 ms, duration 30 ms in A) obtained at 10.4, 26.9, 68.1, and 109.4 ms during and after 30-ms depolarization in A. x-Axis indicates distance from center of hot spot shown in A. Dotted lines indicate distance of 0, 2.1, 4.3, and 6.4 µm from the hot spot along the x-y section. Db: time course of F/F0 ratio at distances of 0, 2.1, 4.3, and 6.4 µm from the hot spot. These distances correspond to the dotted lines shown in the profile (Da). Horizontal line denotes 50% of F/F0 peak in hot spot. Note that Ca2+ wave spreads from superficial hot spot (0 µm) into the myoplasm (6.4 µm), denoted by increase in peak F/F0 ratio.

When the depolarization duration was >10 ms, most of the Ca²⁺ hot spots spread to other areas in the myoplasm as described above. Fig. 2Da shows the profile of the F/F₀ ratio along a section (x-y in Fig. 2A; 30 ms) crossing a hot spot (). The elevation of F/F₀ ratio started in hot spot just beneath the cell membrane and spread inside the cell as a Ca²⁺ wave. The time courses of Ca²⁺ waves at points in hot spot (0 µm from cell edge) and 2.1, 4.3, and 6.4 µm inside the cell along the section (x-y) are shown in Fig. 2Db. Note that the Ca²⁺ wave spread and increased the F/F₀ ratio even after repolarization. These findings suggest that Ca²⁺ influx is not required for Ca²⁺ wave propagation. Moreover, the Ca²⁺ wave is unlikely to be simply the diffusion of Ca²⁺ from hot spot sites; it must include a Ca²⁺ release chain reaction across the myocyte, because [Ca²⁺]_i at the top of the Ca²⁺ wave increased dramatically during propagation and was greater than [Ca²⁺]_i in the original hot spot.

Figure 3 shows data from five cells demonstrating the relationship between the duration of depolarization and the peak increase in global F/F₀ ratio after clamp depolarization. The peak of global F/F₀ ratio induced by a 30-ms pulse was taken as unity in each cell, whereas values in response to shorter depolarizations were considered relative F/F₀ ratios (Fig. 3Aa). Although not all cells responded in an all-or-none manner, the global F/F₀ ratio was always close to the maximum value when the depolarization duration was >10 ms. The number of Ca²⁺ hot spots observed in a single confocal plane also increased with progressive lengthening of the duration of depolarization. When three or more Ca²⁺ hot spots appeared in a single confocal plane adjusted to near the middle of the cell's height, Ca²⁺ hot spots spread to other areas as waves (Fig. 3, Aa and Ab). If only one or two Ca²⁺ hot spots occurred, the hot spots often disappeared within 80 ms as shown for Ca²⁺ sparks in Fig. 3B. The increase in F/F₀ ratio after depolarization was totally inhibited when I_Ca through VDCCs was blocked by the addition of 100 µM Cd²⁺ (data not shown).

View larger version (27K): [in this window] [in a new window]   Fig. 3. Relationship between [Ca2+]i elevation and depolarizing pulse duration. Aa: relative increase in peak F/F0 ratio from whole cell area after depolarization pulses with durations of 3, 5, 10, 20, and 30 ms. Peak F/F0 ratio after 30-ms depolarization was taken as unity (1.0) in each cell. Data were obtained from 5 cells in experiments similar to those shown in Fig. 2. Ab: number of Ca2+ hot spots, including evoked Ca2+ sparks, are plotted against voltage-clamp depolarization. When average F/F0 ratio in a local area (1.5 x 1.5 µm, corresponding to 28 pixels) increased by >0.5, it was detected as a Ca2+ spark, which declined completely within 100 ms. If both F/F0 ratio and size of Ca2+ spark increased significantly and lasted for >200 ms, the event was considered a Ca2+ hot spot. Cell 1 in Aa is the same cell shown in Fig. 2. B: schematic showing proposed functions of evoked Ca2+ sparks or hot spots in excitation-contraction (E-C) coupling in mouse UBSMCs. When a short depolarizing pulse (5 ms) was applied, only a few Ca2+ sparks were evoked. These sparks did not spread; instead, each disappeared quickly. When the duration of voltage-clamp depolarization was longer (>10 ms), increased Ca2+ entry through voltage-dependent Ca2+ channels (VDCCs) evoked a larger number of sparks. These Ca2+ sparks progressively grew and/or merged with each other as Ca2+ hot spots, which then spread into other areas of the cell as Ca2+ waves. Note that these waves were present even after the myocyte repolarized to –60 mV, which resulted in global rise of [Ca2+]i and a contraction.

The changes in F/F₀ ratio from the whole cell area upon depolarization for 30 ms in the absence (control) and presence of 100 µM ryanodine are shown in a superimposed image in Fig. 5A, which represents relatively long time scale recordings. In addition to a large decrease in the global F/F₀ ratio, the time to peak global F/F₀ ratio also changed in the presence of 100 µM ryanodine (162.4 ± 24.8 ms and 46.7 ± 3.3 ms; n = 7 and n = 5, respectively; P < 0.01). It is particularly notable that the global F/F₀ ratio in the presence of ryanodine started to decline just after repolarization. In contrast, the global F/F₀ ratio in the control cells further increased and reached its peak 200 ms after repolarization. The relationship between the duration of depolarization and peak global F/F₀ ratio was examined in the presence of 100 µM ryanodine by changing the duration of depolarization in the manner shown in Fig. 2 (see also Fig. 5B). When the duration of depolarization was increased from 5 to 50 ms, the F/F₀ ratio was increased in a sigmoid fashion in the control cells. The global F/F₀ ratio in the presence of ryanodine also was increased by lengthening the duration of depolarization, but the increase was much smaller. The application of 100 µM ryanodine reduced the relative intensity to 20% of the control value at 50 ms. When the duration was increased to 150 ms, the F/F₀ ratio in the presence of ryanodine increased markedly because of the sustained influx of Ca²⁺ through VDCCs.

View larger version (25K): [in this window] [in a new window]   Fig. 5. Effects of 100 µM ryanodine on global [Ca2+]i and VDCC currents. A: time course of F/F0 ratio from whole cell area shown in Fig. 4 replotted along a longer time scale. The top trace and a gray vertical bar indicate the period of depolarization (30 ms from –60 to 0 mV). B: summarized data of peak F/F0 ratio from whole cell area after depolarization at various pulse durations (5–150 ms). Number of experiments is shown in parentheses above each data point. **P < 0.01, *P < 0.05 vs. control. C: data summarizing peak Ca2+ current (ICa) density at 0 mV in the presence and absence of 100 µM ryanodine. Maximum ICa was obtained at 0 mV, when cells were depolarized for 30 ms from –60 mV in 10-mV steps. Ryanodine was added to the pipette solution (intracellular application; left) or to the bathing solution (extracellular application; right), respectively.

Effects of 100 µM ryanodine on VDCCs also were carefully examined in UBSMCs. When cells were activated by depolarization from –60 mV to positive potentials in 10-mV steps, the maximum amplitude of peak inward currents was obtained at 0 mV in both the absence and presence of ryanodine in the pipette solution (data not shown). The inward current induced by depolarization to 0 mV was blocked by 100 µM Cd²⁺ or 100 nM nicardipine, indicating that the current was almost completely through L-type VDCCs. The densities of peak I_Ca at 0 mV were 7.8 ± 1.3 pA·pF^–1 (n = 7) and 5.4 ± 1.0 pA·pF^–1 (n = 5; P > 0.05 vs. control) in the absence and presence of ryanodine in the pipette solution, respectively (Fig. 5C). The addition of 100 µM ryanodine to the bathing solution did not significantly change the peak amplitude of I_Ca at 0 mV (8.8 ± 1.0 pA·pF^–1 in control cells and 7.9 ± 1.0 pA·pF^–1 in the presence of ryanodine, n = 4 for each; P > 0.05 vs. control). These findings show that the marked decrease in the rise of [Ca²⁺]_i level upon depolarization by 100 µM ryanodine was not due to the decrease in Ca²⁺ influx through VDCC.

When 100 µM ryanodine was added to the pipette solution, outward currents as well as global [Ca²⁺]_i level were substantially reduced (Fig. 6A). The current density of peak outward current of 150-ms depolarization was reduced from 15.4 ± 4.4 to 2.9 ± 0.4 pA·pF^–1 (n = 5 and n = 6; P < 0.05) (Fig. 6B). The remaining outward current in the presence of ryanodine was not significantly affected by the addition of 100 nM iberiotoxin or 1 µM paxillin, whereas the current in the absence of ryanodine was markedly reduced by these specific BK channel blockers. These observations indicate that the outward current component reduced by ryanodine was due mainly to BK channel activity. Consistent with this finding, the STOCs induced by activation of BK channels by spontaneous Ca²⁺ release through RyRs located in the superficial SR were observed at a holding potential of –30 mV in the control but not in cells loaded with 100 µM ryanodine in the pipette solution (Fig. 6C).

View larger version (20K): [in this window] [in a new window]   Fig. 6. Effects of 100 µM ryanodine on outward currents in UBSMCs. Effects of 100 µM ryanodine on outward currents in UBSMCs. Myocytes were loaded with 100 µM ryanodine from recording pipettes. Measurements were started 5 min after formation of whole cell patch clamp. A: current traces in control cells (a and c) and ryanodine-loaded cells (b and d). UBSMCs were depolarized from –60 to 0 mV for 30 ms (a and b) or 150 ms (c and d). B: summarized data of the density of peak outward currents elicited by 30 ms (a) and 150 ms (b) in control cells (open bars) or ryanodine-loaded cells (solid bars). Number of experiments is shown in parentheses next to key. *P < 0.05 vs. control. C: current traces in control cells (a) and ryanodine-loaded cells (b) at holding potential of –30 mV. Spontaneous transient outward currents (STOCs) were observed in a control cell but were not present in a ryanodine-loaded cell. This pattern was observed in all cells examined (n = 5 for control, n = 6 for ryanodine).

Figure 7 shows the responses to 10 µM ACh after the stimulation by voltage-clamp depolarization was repeated twice in the absence or presence of 100 µM ryanodine in the pipette solution. The F/F₀ ratio was elevated approximately twofold by 30-ms depolarization from –60 to 0 mV at the interval of 100 s, and the F/F₀ ratio was significantly smaller in the presence of ryanodine than in its absence as shown in Fig. 5. ACh (10 µM) was applied in the presence of 5 mM Ni²⁺ in the bathing solution to prevent Ca²⁺ influx though both VDCCs and receptor-operated Ca²⁺ channels. ACh elicited a large increase in the F/F₀ ratio in both control and ryanodine-loaded cells, but this increase was significantly smaller in ryanodine-loaded cells at a holding potential of –60 mV. These results clearly indicate that a large part of IP₃-sensitive Ca²⁺ store sites remains intact in cells loaded with 100 µM ryanodine. These results also suggest that the Ca²⁺ rise via IICR during the response to ACh may in addition activate RyRs to induce CICR, at least in part. Alternatively, ryanodine at the high concentration of 100 µM may nonselectively interact with IP₃Rs and slightly suppresses IICRs.

View larger version (21K): [in this window] [in a new window]   Fig. 7. Effects of 100 µM ryanodine on ACh-induced Ca2+ release. A: Ca2+ release from intracellular store sites due to application of 10 µM ACh was measured after repetitive voltage-clamp depolarizations in the absence (control cells) (a) or presence (b) of 100 µM ryanodine in the pipette solution under voltage-clamp conditions. Myocytes were depolarized twice from holding potentials of –60 to 0 mV for 30 ms after 100-s interpulse intervals, indicated by arrows (first and second pulses). About 5 min after the second pulse, 10 µM ACh was applied (in presence of 5 mM Ni2+ to block Ca2+ influx). B: summarized data for peak F/F0 ratio from whole cell area after 1) application of first and second depolarization pulses and 2) application of 10 µM ACh. Number of experiments is shown in parentheses next to key. *P < 0.05 vs. control.

View larger version (28K): [in this window] [in a new window]   Fig. 8. Effects of xestospongin C on ACh-induced Ca2+ increases. Effects of 3 µM xestospongin C on changes in [Ca2+]i after application of 1 µM ACh examined at holding potential of –60 mV under voltage clamp. Myocytes were loaded with 3 µM xestospongin C or vehicle (0.05% ethanol together with 100 µM indo-1) from the recording pipettes. Measurements were started at 5 min after formation of whole cell patch clamp. A: ratiometric images of cells loaded with vehicle (a) or 3 µM xestospongin C (b). Images i, ii, and iii were obtained during resting conditions, at peak of transient phase, and in sustained phase, respectively. Asterisks and color scale bar indicate position of recording pipette tip and the calibration of [Ca2+]i, respectively. B: time courses of [Ca2+]i (top traces) and membrane currents at –60 mV (bottom traces) in cells loaded with vehicle (a) or xestospongin C (b). Vertical dashed lines i, ii, and iii show time courses corresponding to phases i, ii, and iii in Aa and Ab. C: summarized [Ca2+]i data under resting conditions (i in A and B), at peak of transient phase (ii in A and B), and in sustained phase (iii in A and B). Number of experiments is shown in parentheses next to key. *P < 0.05 vs. vehicle.

View larger version (45K): [in this window] [in a new window]   Fig. 9. Effects of xestospongin C on [Ca2+]i during depolarization. A, left: effects of xestospongin C on Ca2+ mobilization after depolarization examined under voltage clamp. Myocytes were loaded with vehicle (0.05% ethanol) (a) or 3 µM xestospongin C (b) from the recording pipette, together with 100 µM fluo-4. Times indicated above each image in A, left, indicate time after start of depolarizing pulse. Myocytes were depolarized from –60 to 0 mV for 30 ms in the presence of 30 mM TEA and 1 mM 4-AP in the bathing solution to block K+ currents. Arrows and asterisks in A, left, indicate sites generating Ca2+ hot spots and position of the recording pipette tip, respectively. Measurements of Ca2+ images were started 5 min after formation of whole cell patch clamp. A, right: time courses of membrane current (black dots), F/F0 ratio from hot spots indicated by arrow (red squares) and arrow (green circles) in images shown at left, and F/F0 ratio from whole cell area (blue triangles). Top and bottom time course graphs were measured in absence or presence of xestospongin C and correspond to images at left. B: fluorescence profiles of images at 18.33, 26.88, and 68.13 ms in cells loaded with vehicle (a) or xestospongin C (b). Top traces: Each profile was obtained from cross section x-y in A. x-Axis indicates distance from the center of hot spots along the section x-y. Bottom traces: time courses of F/F0 ratio at points at hot spot (0 µm) and 2.9 and 5.8 µm inside the section (x-y), indicated by corresponding dotted lines in top traces. Horizontal line indicates 50% of the peak F/F0 ratio at hot spot. C: summarized data of the peak F/F0 ratio at hot spots and in whole cell areas. Number of hot spots and cells examined are shown in parentheses above bars. D: effects of 3 µM xestospongin C on [Ca2+]i were also examined using indo-1 instead of fluo-4 as a [Ca2+]i indicator. Averaged [Ca2+]i levels at rest, at peak upon depolarization, and 1.0 s after pulse was started are summarized. Data were obtained using same experimental protocol described in A. Number of experiments is shown in parentheses next to key.

Ca²⁺ influx during depolarization and after Ca²⁺ buffering by store sites. The amount of Ca²⁺ influx through VDCC in response to depolarization from –60 to 0 mV for 30 ms can be calculated from the recordings of inward VDCC currents (20 pC) (Tables 1 and 2). The estimated increase in [Ca²⁺]_i by the Ca²⁺ influx was 9 µM. This increase is 26.6 ± 4.5 times larger than that measured using indo-1 if intracellular Ca²⁺ buffering action, Ca²⁺ uptake by Ca²⁺-ATPase in the SR, and Ca²⁺ extrusion by the Na⁺/Ca²⁺ exchanger Ca²⁺ pump in the plasma membrane are ignored. A much larger dissociation between Ca²⁺ influx and the rise of real [Ca²⁺]_i was observed when cells were loaded with 100 µM ryanodine (60.0 ± 6.3 times; P < 0.05 vs. control). The loading of cells with 3 µM xestospongin C from the pipette solution did not significantly affect the ratio of estimated and measured [Ca²⁺]_i in the range of pulse durations from 30 to 150 ms (Table 2).

View larger version (37K): [in this window] [in a new window]   Fig. 10. Effects of 10 µM cyclopiazonic acid (CPA) on [Ca2+]i increase in UB myocytes after depolarization with 100 µM ryanodine and indo-1 from the recording pipette. Before and after application of CPA, cells were depolarized from –60 to 0 mV for 30 ms in the presence of 30 mM TEA and 1 mM 4-AP in bathing solution. A: [Ca2+]i images obtained after first depolarization (i and ii), after CPA application (iii-v), and after second depolarization (vi and vii) in presence of CPA. Asterisks and color scale bar in A indicate position of recording pipette tip and calibration of [Ca2+]i, respectively. B: time courses of [Ca2+]i changes due to first depolarization, CPA application, and second depolarization. Timing of depolarizations is indicated by filled arrowheads in first- and second-pulse graphs. C: superimposed ICa traces elicited by first (black) and second depolarizations (gray). D: bar graph of ratios of [Ca2+]i due to charge entry corresponding to ICa for first depolarization in presence of ryanodine only and second depolarization after addition of CPA, respectively. Number of experiments is shown in parentheses above bars. *P < 0.05 between two groups (paired t-test).

View larger version (35K): [in this window] [in a new window]   Fig. 11. Effects of 100 µM ryanodine on electrically evoked contractions in UB. Effects of ryanodine on isomeric contractions measured in UB tissue segments. A: traces showing twitch contractions elicited in UB segments by direct electrical stimulation in presence of the following neurotransmitter antagonists (in µM): 1 atropine, 1 phentolamine, 1 propranolol, 1 TTX, and 10 suramin. Tissue segments were stimulated by 3-ms pulses of 300-mA magnitude at 5 Hz in a train lasting 2 s. Intervals of 90 s were used between stimulus trains. Ryanodine was applied at concentrations of 10 µM (Aa), 30 µM (Ab), or 100 µM (Ac). Ba: graph showing data summary of effects of ryanodine on relative amplitudes of twitch contractions. Number of experiments is shown in parentheses above bars. **P < 0.01, *P < 0.05 vs. control. Bb: data summary describing increase in resting tension due to ryanodine application. Number of experiments is shown in parentheses above bars. #P < 0.05 between 2 groups. C: trace recording showing effects of 100 µM ryanodine on spontaneous rhythmic contraction. Spontaneous rhythmic contraction in UB was recorded in 15 mM KCl solution in presence of neurotransmitter antagonists.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
In the present study, we have demonstrated that during E-C coupling after action potential generation in UBSMCs, CICR initially occurred quickly in localized hot spot sites that corresponded to functional coupling between VDCCs and RyRs. Thereafter a secondary spread of Ca²⁺ waves to other Ca²⁺ store sites occurred much more slowly. The amount of Ca²⁺ influx during short depolarization was calculated to be large enough to increase [Ca²⁺]_i by several micromolar levels. In vivo, however, Ca²⁺ that entered a myocyte was strongly buffered and global [Ca²⁺]_i was increased only slightly. Thus the Ca²⁺ influx initiated CICR mainly in discrete hot spot sites from which Ca²⁺ waves spread to the entire myoplasm in the second step of Ca²⁺ release. The second depolarization step increased global [Ca²⁺]_i and induced a contraction.

Ryanodine suppressed Ca²⁺ hot spots, Ca²⁺ waves, and contraction. In mouse UBSMCs, short depolarization elicited Ca²⁺ hot spots at discrete local sites in subsarcolemmal areas. These hot spots developed 10 ms after the start of depolarization as previously reported in SMCs of the guinea pig vas deferens and UB (31, 42). The occurrence of Ca²⁺ hot spots required Ca²⁺ influx through VDCC. These hot spots were usually detected as frequently discharging sites of spontaneous Ca²⁺ sparks (42). In this study, we found the number of Ca²⁺ hot spots elicited by depolarization to be closely related to the duration of the voltage-clamp depolarization up to 20 ms. This finding presumably reflects the principle that the coupling efficacy between Ca²⁺ influx through VDCCs with the activation of RyRs required to induce CICR may vary widely between discrete Ca²⁺ hot spot sites. The loose coupling compared with that in cardiac myocytes has been suggested in rabbit UBSMCs (7). A short depolarization (5 ms) elicited only a few hot spots in well-defined, discrete sites, presumably indicating tight coupling. Larger Ca²⁺ influx induced by longer depolarization may activate discrete sites with weaker coupling efficacy. Once a sufficient number of Ca²⁺ hot spots occurred during a certain period of depolarization, Ca²⁺ waves developed progressively and spread slowly from hot spots through the entire myoplasm. This phenomenon continued even after repolarization and elevated [Ca²⁺]_i to the extent that contraction was induced.

To our knowledge, this study represents the first reported demonstration of a threshold in the duration (10 ms) of depolarization required to evoke local Ca²⁺ sparks or change hot spots to a global [Ca²⁺]_i increase in SMCs. Ca²⁺ hot spots function as the initiation sites of Ca²⁺ waves essential for global [Ca²⁺]_i increases in SMCs. Although the mechanism by which transient Ca²⁺ sparks switch to Ca²⁺ hot spots and/or waves is not clear, the coalescence of neighboring hot spots may underlie this switching to global [Ca²⁺]_i increases. Accordingly, the number of evoked Ca²⁺ sparks or hot spots by a given depolarization may be determining factor with regard to whether a contraction is elicited by the depolarization in UBSMCs. It is noteworthy that the spread of Ca²⁺ waves did not absolutely require Ca²⁺ influx through VDCCs because the wave spread up to 200 ms after the cessation of 30-ms depolarization even though the Ca²⁺ tail current lasted for 20 ms.

In this study, application of 100 µM ryanodine from the recording pipette abolished both Ca²⁺ hot spots and Ca²⁺ waves but did not affect I_Ca. Although the application of 10 µM ryanodine from the pipette may reduce CICR (13), this concentration of ryanodine increases the resting [Ca²⁺]_i in guinea pig UB tissues (20), presumably by locking RyRs in a half-open state (46). On the other hand, a high concentration of ryanodine blocks RyRs completely (46). In the present study, we used 100 µM ryanodine to block RyRs completely. This blocking occurred almost immediately after the start of whole cell recording. Consequently, we did not observe a substantial increase in resting [Ca²⁺]_i. Neither STOCs nor Ca²⁺ sparks were observed in UBSMCs loaded with 100 µM ryanodine, confirming that RyRs were blocked completely. In these myocytes, the increase in [Ca²⁺]_i occurred slowly but substantially during long depolarization (150 ms). In summary, CICR was almost completely blocked by 100 µM ryanodine in UBSMs and the Ca²⁺ influx through VDCCs per se increased global [Ca²⁺]_i only slightly during 30-ms depolarization.

Experimental results from studies that have addressed the susceptibility to contraction in UBSMCs treated with ryanodine are varied and somewhat controversial. Three groups have reported that spontaneous contractions in guinea pig and rat UBSM strips were not reduced, but rather enhanced, by ryanodine treatment (18, 23, 41). However, another group reported effective blockage of spontaneous contraction by ryanodine to 50% of control in UBSMCs of the pig (5). The [Ca²⁺]_i rise induced by spontaneous action potentials in tissue preparations of the guinea pig was reduced to 60% of control by application of 10 µM ryanodine (20), and 10 µM ryanodine had no significant effect on the [Ca²⁺]_i rise elicited by field stimulation in UBSMs of the rat (22). Suppression of CICR by 10 µM ryanodine, however, has been shown convincingly in single UBSMCs of the guinea pig (13). In the present study, the contraction induced by direct electrical stimulation was markedly reduced by ryanodine, and this inhibition was dose dependent in UBSMs of the mouse. The resting tone was significantly increased by ryanodine at 10 and 30 µM but not at 100 µM, consistent with the changes in resting [Ca²⁺]_i levels induced by 10–100 µM ryanodine. The I_Ca in UBSMCs was not affected significantly by the addition of 100 µM ryanodine to the superfusing solution. Unlike guinea pig UBSMCs, large, spontaneous contractions were seldom observed in mouse UBSM tissue (45). Our results (see Fig. 11) also demonstrate that the spontaneous contraction observed in 15 mM K⁺ solution was susceptible to 100 µM ryanodine as well as the contraction elicited by direct electrical stimulation.

Although the reasons for the discrepancies concerning the susceptibility of contraction to ryanodine in previous studies are not completely clear, the contribution of RyRs and CICRs to E-C coupling in UBSMCs could be different between species. Moreover, the susceptibility of contraction to ryanodine was higher when conditions for direct electrical stimulation were moderate or weak (Morimura K, unpublished observations). It is plausible that the relative contribution of CICR versus that of Ca²⁺ influx to increase global [Ca²⁺]_i during E-C coupling becomes smaller under conditions in which Ca²⁺ influx is larger. This hypothesis is consistent with the finding that the elevation in global [Ca²⁺]_i by depolarization was large even in UBSMCs loaded with 100 µM ryanodine and when the duration of depolarization was long (150 ms).

Ca²⁺ buffering for Ca²⁺ influx through VDCCs is still mysterious. We observed a clear discrepancy between the amount of Ca²⁺ influx during short depolarization that mimicked an action potential and its direct contribution to the rise in global [Ca²⁺]_i. As shown in Table 1, the estimate of [Ca²⁺]_i increase on the basis of the amount of Ca²⁺ influx during depolarization for 30 ms from –60 to 0 mV was >25 times larger (so-called buffering power) than the peak [Ca²⁺]_i measured directly using of indo-1. A similar discrepancy between estimated and measured [Ca²⁺]_i values has been discussed in the following other SMC models: rat femoral artery buffering power, 130 times (35); guinea pig coronary artery, 150 times (14); rat portal vein, 114 times (36); equine airway, 15–50 times (9); and guinea pig UB, 46 times (11). When CICR triggered by depolarization was blocked by 100 µM ryanodine in this study, this buffering power was apparently much larger: 60 times. Such a large discrepancy indicates mechanisms of fast, strong buffering power for Ca²⁺ coming into the cytosol through VDCCs.

In many types of cells that express K_Ca channels, measurement of K_Ca channel current under whole cell clamp mode is a reliable monitor of [Ca²⁺]_i levels in the subsarcolemmal regions. In UBSMCs from the mouse as well as in those from the guinea pig, the major component of outward current upon depolarization was K⁺ current though BK channels. The activation of BK channels is responsible for repolarization and afterhyperpolarization of an action potential (21, 28, 30). The ill-defined buffering mechanism for Ca²⁺ entering through VDCCs is functional in superficial areas just beneath the cell membrane in UBSMCs, because BK channel current activated by depolarization also was strongly suppressed in ryanodine-loaded cells in this study. In inside-out patch-clamp mode, the single-channel activity of BK channels in UBSMCs was not affected by 100 µM ryanodine (Morimura K et al., unpublished observations). Taken together, these findings show that any substantial Ca²⁺ entering though VDCCs does not elevate [Ca²⁺]_i markedly and does not activate BK channels effectively, presumably because of strong and fast buffering of Ca²⁺ by uptake, binding, and extrusion from superficial areas. The results of the present study confirm that CICR through RyRs in the SR is required for the activation of BK channels as well as for the activation of the contractile apparatus during E-C coupling in UBSMCs.

The mechanisms by which Ca²⁺ that enters through VDCCs during an action potential is strongly buffered are not known in detail. It is likely that Ca²⁺ uptake to the SR by the Ca²⁺ pump (i.e., sarcoplasmic reticulum Ca²⁺-ATPase) is the largest component of this buffering. The addition of 10 µM CPA to the ryanodine-loaded cells resulted in a significant decrease in the buffering power from 60 to 34 (Table 1). However, this decrease in buffering power by the addition of CPA was not large enough to explain completely the difference between the measured and calculated [Ca²⁺]_i levels after clamp depolarizations. Alternative or additive mechanisms for fast, strong buffering of entering Ca²⁺ may be Ca²⁺ extrusion by the Na⁺/Ca²⁺ exchanger and the Ca²⁺ pump in the plasma membrane, Ca²⁺ uptake by mitochondria, and/or trapping by cytosolic Ca²⁺-binding proteins. Further experiments are required to clarify the mechanisms underlying the mysteriously fast and strong buffering of the Ca²⁺ that enters through VDCCs during an action potential.

IP₃ formation was not involved in Ca²⁺ waves from hot spots during E-C coupling. The spread of Ca²⁺ waves from Ca²⁺ hot spots across the myoplasm is essential for the second step of [Ca²⁺]_i increase in E-C coupling. This Ca²⁺ wave cannot be evaluated on the basis of simple diffusion of Ca²⁺ from hot spot sites to other areas, because Ca²⁺ release occurs progressively near the wave front. Our profile analysis (Fig. 4C) indicates that the F/F₀ ratio in the central area of the myocyte was increased by the Ca²⁺ wave to a higher level than that in the original hot spot site. This pattern of results was observed in all myocytes examined in the present study. Therefore, the Ca²⁺ wave spreads by chain reactions of Ca²⁺ release from hot spots to separate Ca²⁺ stores within a cell.

Ca²⁺ waves have been observed in many types of SMCs and analyzed in detail. In many cases, Ca²⁺ waves are mediated by Ca²⁺ release from IP₃Rs (i.e., IICRs) after IP₃ formation by agonist stimulation (26, 39). Cross talk between CICRs via RyRs and IICRs via IP₃Rs has been reported in various types of cells, including SMCs (15). It has been reported that [Ca²⁺]_i waves due to noradrenaline involve CICRs in addition to IICRs in the rat portal vein (2). The Ca²⁺-dependent activity of some types of PLC and IP₃R activity shows bell-shaped dependence on [Ca²⁺]_i and thus can exhibit positive feedback for IICR facilitation (25, 27). In a cultured neuron derived from the central nervous system, it also has been shown that Ca²⁺ influx through VDCCs induces IICRs, presumably via the formation of IP₃ (43). In the present study, xestospongin C markedly reduced IICRs after the application of ACh but did not affect CICRs triggered by depolarization. The loading of UBSMCs with 100 µM ryanodine reduced IP₃-sensitive Ca²⁺ stores by 40% but did not deplete it. These results strongly suggest the presence of separate stores for IICR and CICR, with some overlap (47). The mechanisms underlying Ca²⁺ waves from Ca²⁺ hot spots during E-C coupling remain to be determined but could be due simply to CICR or, alternatively, to the formation of cADP-ribose (8).

In conclusion, Ca²⁺ release during E-C coupling in UBSMCs occurs in two steps. Ca²⁺ influx during an action potential does increase [Ca²⁺]_i significantly. It initiates CICRs in discrete hot spot sites via functional coupling between VDCCs and RyRs. In the second step, which also involves Ca²⁺ release, Ca²⁺ waves slowly spread to other Ca²⁺ store sites without mediating IP₃R activation. A twitch contraction induced by action potentials triggered by direct electrical stimulation under moderate conditions is highly susceptible to ryanodine treatment. However, Ca²⁺ dynamics during E-C coupling triggered by transmitter release from autonomic nerve endings are more physiological than those observed in the present study. Notably, under those conditions, IICR is likely to also be involved in E-C coupling in UBSMCs.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This work was supported by a Grant-in-Aid for Scientific Research (B) from Japan Society for the Promotion of Science and also by a Grant-in-Aid for Research on Health Sciences focusing on Drug Innovation from Japan Health Sciences Foundation (to Y. Imaizumi).

	ACKNOWLEDGMENTS

 
We thank Dr. Wayne R. Giles (University of Calgary, Calgary, AB, Canada) for providing data acquisition and analysis programs and also for his reading of this manuscript.

	FOOTNOTES

 

Address for reprint requests and other correspondence: Y. Imaizumi, Dept. of Molecular and Cellular Pharmacology, Graduate School of Pharmaceutical Sciences, 3-1 Tanabedori, Mizuhoku, Nagoya 467-8603, Japan (e-mail: yimaizum{at}phar.nagoya-cu.ac.jp)

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.

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