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Fluid pressure modulates L-type Ca2+ channel via enhancement of Ca2+-induced Ca2+ release in rat ventricular myocytes Sunwoo Lee, Joon-Chul Kim, Yuhua Li, Min-Jeong Son and Sun-Hee Woo Am J Physiol Cell Physiol 294:966-976, 2008. First published Feb 13, 2008; doi:10.1152/ajpcell.00381.2007 You might find this additional information useful... Supplemental material for this article can be found at: http://ajpcell.physiology.org/cgi/content/full/00381.2007/DC1 This article cites 44 articles, 31 of which you can access free at: http://ajpcell.physiology.org/cgi/content/full/294/4/C966#BIBL Updated information and services including high-resolution figures, can be found at: http://ajpcell.physiology.org/cgi/content/full/294/4/C966

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Am J Physiol Cell Physiol 294: C966–C976, 2008. First published February 13, 2008; doi:10.1152/ajpcell.00381.2007.

Fluid pressure modulates L-type Ca2⫹ channel via enhancement of Ca2⫹-induced Ca2⫹ release in rat ventricular myocytes Sunwoo Lee,* Joon-Chul Kim,* Yuhua Li, Min-Jeong Son, and Sun-Hee Woo College of Pharmacy, Chungnam National University, Daejeon, South Korea Submitted 24 August 2007; accepted in final form 11 February 2008

L-type Ca2⫹ current; fluid pressure; ventricular myocytes; cytosolic Ca2⫹ transient

myocytes is controlled by a sequence of events (excitation-contraction coupling) that includes Ca2⫹ current (ICa)-gated opening of Ca2⫹ release channels (RyRs) and the release of Ca2⫹ from the sarcoplasmic reticulum (SR) (3, 9, 25, 27). The large local releases of Ca2⫹ in turn inactivate L-type Cav1.2 (␣1C) Ca2⫹ channels (1, 34), suggesting that discrete Ca2⫹ cross-signaling occurs in the microdomains of ␣1C-ryanodine receptors (RyRs) (16, 35). The inactivating property plays an important role in regulating the intracellular Ca2⫹ concentration and the action potential duration of cardiac myocytes. Mechanical stimuli elicited by flow-induced shear stress modulate function of membrane ion channels and the intracellular Ca2⫹ signal, and they serve important functions such as baroreceptor-mediated blood pressure regulation in vascular CONTRACTION OF MAMMALIAN CARDIAC

* S. Lee and J.-C. Kim contributed equally to this work. Address for reprint requests and other correspondence: S.-H. Woo, College of Pharmacy, Chungnam National Univ. 220 Gung-dong, Yuseong-Gu, Daejeon 305-764, South Korea (e-mail: [email protected]). C966

endothelium and smooth muscle (9, 10, 14) and sound perception in the hair cell of the ear (29). Changes in the mechanical environment of the heart, caused by contractility of the heart and changes in its volume and pressure, can alter cardiac excitation and contraction (17, 18). Pathological conditions, such as valve diseases, hypertension, or heart failure, may also lead to haemodynamic or mechanical dysfunction of the heart, causing arrhythmia (10, 17, 18, 20, 21, 26). In whole heart preparations and in situ hearts, an increase in ventricular pressure causes shortenings of the action potential duration and effective refractory period (10, 20, 21). Moreover, there is clinical evidence for a predisposition to fibrillation by regurgitant jets of blood in patients with valve incompetence (26) as well as the occurrence of ectopic tachycardia in a patient with a catheter in the heart chamber (8). These findings suggest the possibility that a pressurized fluid flow and/or direct irritation may directly alter membrane excitability in cardiac myocytes. In a related study, it was recently reported that rapid application of a pressurized flow increases the occurrences of spontaneous Ca2⫹ sparks and longitudinal global Ca2⫹ waves in rat atrial myocytes (40). This paper demonstrates that the fluid pressure (FP)-induced increase in the Ca2⫹ spark frequency is much larger in the cell periphery (40), which indicates that a mechanical sensor in the cell periphery may transmit the fluid pressure force and modulate Ca2⫹ signaling in cardiac cells. In fact, there is evidence that the activities of L-type Ca2⫹ channels of neuron and smooth muscle, expressed in HEK-293 cells, are directly enhanced by flow shear stress (22, 31). However, whether the L-type Ca2⫹ channels directly sense the FP stimulus in intact cardiac myocytes, and whether and how the FP alters the function of the channel, remain to be determined. The present study was designed to explore whether pressurized fluid flow affects the ICa and to investigate the mechanism by which the FP stimulus regulates ICa in isolated single rat ventricular myocytes. FP was applied onto the entire surface of the myocytes through an electronically controlled microperfusion system. This approach was meant to approximate the mechanical stresses that the ventricular chamber wall encounters from the hemodynamic forces of fluid (e.g., the blood jet to the ventricular wall during aortic regurgitation) or from the excessive pressure produced during aortic valve stenosis. Our data suggest that the FP suppresses the Ca2⫹ channels and accelerates their inactivations indirectly by enhancing the Ca2⫹-induced Ca2⫹ release (CICR), and that the effects of FP on the Ca2⫹ channels are independent of stretch-activated ion channel. 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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Lee S, Kim J-C, Li Y, Son M-J, Woo S-H. Fluid pressure modulates L-type Ca2⫹ channel via enhancement of Ca2⫹-induced Ca2⫹ release in rat ventricular myocytes. Am J Physiol Cell Physiol 294: C966–C976, 2008. First published February 13, 2008; doi:10.1152/ajpcell.00381.2007.—This study examines whether fluid pressure (FP) modulates the L-type Ca2⫹ channel in cardiomyocytes and investigates the underlying cellular mechanism(s) involved. A flow of pressurized (⬃16 dyn/cm2) fluid, identical to that bathing the myocytes, was applied onto single rat ventricular myocytes using a microperfusion method. The Ca2⫹ current (ICa) and cytosolic Ca2⫹ signals were measured using a whole cell patch-clamp and confocal imaging, respectively. It was found that the FP reversibly suppressed ICa (by 25%) without altering the current-voltage relationships, and it accelerated the inactivation of ICa. The level of ICa suppression by FP depended on the level and duration of pressure. The Ba2⫹ current through the Ca2⫹ channel was only slightly decreased by the FP (5%), suggesting an indirect inhibition of the Ca2⫹ channel during FP stimulation. The cytosolic Ca2⫹ transients and the basal Ca2⫹ in field-stimulated ventricular myocytes were significantly increased by the FP. The effects of the FP on the ICa and on the Ca2⫹ transient were resistant to the stretch-activated channel inhibitors, GsMTx-4 and streptomycin. Dialysis of myocytes with high concentrations of BAPTA, the Ca2⫹ buffer, eliminated the FP-induced acceleration of ICa inactivation and reduced the inhibitory effect of the FP on ICa by ⬇80%. Ryanodine and thapsigargin, abolishing sarcoplasmic reticulum Ca2⫹ release, eliminated the accelerating effect of FP on the ICa inactivation, and they reduced the inhibitory effect of FP on the ICa. These results suggest that the fluid pressure indirectly suppresses the Ca2⫹ channel by enhancing the Ca2⫹-induced intracellular Ca2⫹ release in rat ventricular myocytes.

MODULATION OF CARDIAC Ca2⫹ CHANNEL BY FLUID PRESSURE MATERIALS AND METHODS

˙ /␲r3 FP 共dyn/cm2兲 ⫽ 4␮Q ˙ is the flow rate (ml/s), and r is where ␮ represents fluid viscosity, Q the internal radius of the tube. The microjet system generated flow forces (in dyn/cm2) of ⬃16, 12, 7, and 3 at 400-, 300-, 200-, and 100-mm reservoir heights, respectively. The positioning of the micro-

barrel was performed under microscope (TS2000, Nikon) using a micromanipulator (Prior England 48260). The experimental cells were attached to the bottom of the chamber without a coating material. Using a microscope and video monitor, it was confirmed that no movement of the cell occurred during the fluid puffing before the start of the patch clamp experiments and Ca2⫹ imaging. At a FP of 400 mmH2O (⬃16 dyn/cm2) the cells used for the recordings did not float or move. Current measurements and analysis. ICa was recorded using the whole cell configuration of the patch-clamp technique (24) using an EPC7 amplifier (HEKA, Lambrecht/Pfalz, Germany). The patch pipettes were made of glass capillaries (Kimble Glass) to have resistance of 2–3 M⍀ when filled with the internal solution containing (in mM) 110 CsCl, 20 TEA-Cl, 20 HEPES, 5 MgATP, and 2 EGTA, with the pH adjusted to 7.2 with CsOH; in some experiments 10 mM BAPTA was also included in the pipette solution (see Fig. 8). Outward K⫹ currents were suppressed by replacing internal K⫹ with Cs⫹ and TEA⫹, and inward rectifier K⫹ current was suppressed by replacing external K⫹ with Cs⫹. Na⫹ current was inactivated by holding the membrane potential at ⫺40 mV. Trains of test pulses were to 0 mV for 120 ms with 0.1 Hz. The ICa was fully sensitive to 20 ␮M of nifedipine (data not shown) and to 200 ␮M Cd2⫹ (Fig. 4). To measure the Ba2⫹ current (IBa), we replaced extracellular Ca2⫹ (2 mM) with equimolar Ba2⫹. Measurement of ICa and IBa were carried out 5⬃6 min after rupture of the membrane with the patch pipette, when the rundown of Ca2⫹ channels were slowed and stabilized. Generation of voltage protocols and acquisition of data were carried out using pCLAMP (9.0, Axon Instruments, Foster City, CA) combined with an analog-to-digital converter (Digidata 1322, Axon Instruments). The series resistance was 1.5–3 times the pipette resistance and was electronically compensated through the amplifier. The current signals were digitized at 10 kHz and low-pass filtered at 1 kHz. We usually monitored raw currents as well as currents leak subtracted by the pulse/number (P/N) method (N ⫽ 5). Data in the figures excluding the insets of Fig. 1A and Fig. 5A are shown without leak

Fig. 1. Effects of fluid pressure on Ca2⫹ current (ICa) in rat ventricular myocytes. A: time courses of the ICa rundown (E) and the effects of fluid pressure (FP; ⬃16 dyn/cm2) on peak ICa (F). The plots show average results (⫾SE) obtained in 10 control cells (no FP) and in 10 cells treated with FP. ICa was activated by 120-ms depolarizing pulses from ⫺40 to 0 mV at 0.1 Hz (for all figures). The arrow indicates the onset of FP application. Inset shows superimposed current traces recorded immediately before the exposure to FP (“C”) and at the time when a maximal decrease in ICa by FP was observed (“FP”). The left and right current traces show raw and leak-subtracted currents, respectively (see MATERIALS AND METHODS). B: comparison of mean ICa. ***P ⬍ 0.001, control vs. FP (16 dyn/cm2; n ⫽ 45). C and D: superimposed current traces and current-voltage relationships (n ⫽ 7) of ICa recorded at different testing potentials ranging from ⫺40 to ⫹50 mV [holding potential (Vh) ⫽ ⫺40 mV] in the control condition (“Control”) and after exposures to FP (16 dyn/cm2).

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Single-cell isolation. Rat ventricular myocytes were enzymatically isolated from male Sprague-Dawley rats (200 –300 g) as described previously (20). Briefly, rats were deeply anesthetized with pentobarbital sodium (150 mg/kg ip), the chest cavity was opened, and hearts were excised. This surgical procedure was carried out in accordance with university ethical guidelines. The excised hearts were retrogradely perfused at 7 ml/min through the aorta (at 36.5°C), first for 5 min with Ca2⫹-free Tyrode solution composed of (in mM) 137 NaCl, 5.4 KCl, 10 HEPES, 1 MgCl2, and 10 glucose, pH 7.3, and then with Ca2⫹-free Tyrode solution containing collagenase (1.4 mg/ml, Type 1, Roche) and protease (0.14 mg/ml, Type XIV, Sigma) for 12 min; and finally with Tyrode solution containing 0.2 mM CaCl2 for 8 min. The ventricles of the digested heart were then cut into several sections and subjected to gentle agitation to dissociate the cells. The freshly dissociated cells were stored at room temperature in Tyrode solution containing 0.2 mM CaCl2. Application of fluid pressure. Myocytes were continuously superfused with the Tyrode solution composed of (in mM) 137 NaCl, 5.4 KCl, 10 HEPES, 1 MgCl2, 10 glucose, and 2 CaCl2, pH 7.4. Pressurized flows of solutions were applied onto the single myocytes through a microbarrel (internal diameter ⫽ 250 ␮m) the tip of which was placed at ⬇150 ␮m from the cell (see the supplemental data in the online version of this article) and was connected to a fluid reservoir with different heights (100 – 400 mm). Electronic solenoid valve was installed in the middle of tubing connecting the fluid reservoir and the microbarrel, the tip of which, touching the chamber bottom, was tilted to one side with an angle of 45°. The FP was calculated for flow in cylindrical tubes according to the equation (23)

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MODULATION OF CARDIAC Ca2⫹ CHANNEL BY FLUID PRESSURE

subtraction to demonstrate that the cells had low and stable leak current. The percent suppressions of ICa by various interventions were evaluated after a gradual decrease in ICa by rundown was subtracted from the raw current (23). Briefly, the time course of changes in peak ICa measured in the control conditions was fitted using double exponential curve fitting; the fitted curve then was considered to be a rundown component. The difference between the original time course and the fitted curve resulted in pure changes in ICa, which was used to measure percent changes during the interventions. Peak detection was performed with Clampfit (9.0, Axon Instruments), and the time constant (␶) of inactivation of ICa was obtained with single exponential curve fitting using the equation: y ⫽ 共A i ⫺ Af兲䡠 exp共⫺t/␶兲 ⫹ Af where Ai and Af are, respectively, the initial (t ⫽ 0) and final (t ⫽ infinity) values of the parameter, and ␶ is a time constant of exponential decay. The extent of the steady-state inactivation of ICa was determined by a double-pulse protocol at a frequency of 0.05 Hz. Plots in Fig. 3B were fitted to Boltzmann’s distribution:

where Imax is maximum current, V0.5 is the midpotential, and k is the slope of the curve. Curve fitting for the steady-state inactivation was performed using Origin 6.0 software (Microcal).

Fig. 2. Dependences of the inhibitory effects of FP on the level and duration of pressure. A and D: time courses of the changes of ICa under either different levels (A) or durations (D) of fluid pressures. ICa was activated by depolarizing step pulses from ⫺40 to 0 mV at 0.1 Hz, and it was normalized by the peak current recorded immediately before the exposure to FP (A). D: decreases of the ICa magnitude by FP of 16 dyn/cm2 with different durations. The horizontal boxes indicate periods of FP exposures. B and E: representative ICa traces recorded in the control condition (“C”) and under different strength or duration of FP. C: FP (dyn/cm2)response (% suppression) curve shows pressure-dependent suppression of ICa. 3 dyn/cm2, 9 cells; 7 dyn/cm2, 5 cells; 12 dyn/cm2, 45 cells. ***P ⬍ 0.001, control vs. FP. F: mean % suppressions of ICa by 5-s-long (n ⫽ 8) and 20-s-long (n ⫽ 7) exposures to FP and by the application of FP for a period to achieve a maximal effect (“Max”; n ⫽ 9).


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I/I max ⫽ 兵1 ⫹ exp关共V ⫺ V0.5兲/k兴其⫺1

All the experiments were performed at room temperature (22–25°C). Confocal Ca2⫹ imaging. Cells were loaded with fluo-4 AM (4 ␮M; Molecular Probes) for 40 min. Ca2⫹ transients were elicited by applying depolarizing pulses (⫹20 V for 1.5 ms) at 1 Hz using a pair of Pt electrodes combined with a stimulator (D-7806, Hugo Sachs Elektronik, March-Hugstetten, Germany). Ca2⫹ fluorescence was imaged using a laser scanning confocal microscopy system (C1 Eclipse, Nikon, Tokyo, Japan) attached to an inverted microscope (TS2000U, Nikon) fitted with a ⫻60 oil-immersion objective lens (Plan Apo, numerical aperture 1.4, Nikon). Dyes were excited at 488 nm using Ar ion laser (Ommichrome) and fluorescence emission at wavelengths ⬎510 nm was detected. Line-scan images were captured at 16.4 ms/line with PC program EZ-C1 (version 3.0, Nikon). Local averaged Ca2⫹ signals were measured using a custom-written PC program (“Pic”) in the Visual C⫹⫹ (Microsoft). The average resting fluorescence intensity (F0) was calculated from several lines measured before application of FP. Tracings of local Ca2⫹ transients were shown as the average fluorescence of each line normalized relative to the average resting fluorescence (F/F0; see Fig. 7, B and F). Statistics. Numerical results are presented as means ⫾ SE (n ⫽ number of cells). A paired Student’s t-test was used to evaluate the statistical significance of differences between means. Differences at P ⬍ 0.05 were considered to be significant.

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Fig. 3. Effects of FP on inactivation of Ca2⫹ channel. A: superimposed, peak-normalized ICa (left) and comparison of mean inactivation time constant (␶; right) show faster inactivation of ICa during FP (16 dyn/cm2) exposure. **P ⬍ 0.01, control vs. FP (n ⫽ 45). B: measurement of steady-state inactivation was carried out by applying prepulses to produce voltages ranging from ⫺50 to ⫹10 mV for 2.4 s from a Vh of ⫺40 mV, followed by a test pulse to 0 mV for 100 ms (n ⫽ 5). The peak amplitude of test pulse current was then plotted against the prepulse voltage. The individual peak currents were normalized to the peak ICa amplitude obtained at ⫺50 mV. Imax, maximum current.

tive currents (current recorded in the control condition ⫺ current recorded in the presence of Cd2⫹) were reduced by FP (16 dyn/cm2) to ⬃70% (Fig. 4, D and E), and they still shifted to the positive direction under FP. Modulation of Ca2⫹ channel by FP is independent of stretch-activated channel. Intracellular Ca2⫹ concentrations have been reported to increase during stretches of various origins as a consequence of the activation of a stretch-activated membrane current (13, 38). In addition, the cytosolic Ca2⫹ increase could be the origin of changes in ICa and its kinetics (1, 34). To test whether FP-induced ICa suppression is caused by increase in the cytosolic Ca2⫹ concentration through the activation of the stretch activated ion channel, we examined the effects of FP on ICa in cells preexposed to the blockers of the stretch-activated ion channel, streptomycin (strep) (43) and GsMTx-4 (5, 36). Figure 5, A–C, depicts that the pretreatment of streptomycin (40 ␮M) did not alter the effects of FP either on the amplitudes of ICa (strep, 8.3 ⫾ 0.8 pA/pF; strep ⫹ FP, 6.5 ⫾ 0.9 pA/pF; n ⫽ 5; P ⬍ 0.01; Fig. 5, A and B) or on the inactivation of ICa (␶: strep, 16.6 ⫾ 0.16 ms vs. strep ⫹ FP, 13.9 ⫾ 0.66 ms; n ⫽ 5; P ⬍ 0.01; Fig. 5C). In cells preexposed to 5 ␮M GsMTx-4, a more selective and potent blocker of stretch-activated channel, FP continued to decrease ICa (GsMTx-4, 7.33 ⫾ 0.50 pA/pF vs. GsMTx-4 ⫹ FP, 5.11 ⫾ 0.45 pA/pF; n ⫽ 15; P ⬍ 0.001; Fig. 5, D and E), and accelerated the inactivation of ICa [␶ (ms): GsMTx-4, 17.9 ⫾ 0.65 vs. GsMTx-4 ⫹ FP, 15.1 ⫾ 0.56, n ⫽ 15, P ⬍ 0.01; Fig. 5F]. The applications of streptomycin and GsMTx-4 did not

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Suppression of ICa by FP in rat ventricular myocytes. Figure 1, A and B, illustrates the effects of pressurized fluid flow on whole cell ICa in single rat ventricular myocytes. Pressurized flows of solutions were applied onto the single myocytes through a microbarrel with internal diameter of ⬇250 ␮m (see MATERIALS AND METHODS). Using a microscope, it was confirmed that no movement of the cell occurred during the fluid puffing before the start of the experiments. Exposure of a FP (⬃16 dyn/cm2) to the single myocytes significantly and reversibly decreased ICa (by ⬃25.9% on average, n ⫽ 45; Fig. 1, A and B). A 90% of maximal suppression of ICa was achieved at 72 ⫾ 5.3 s following the onset of FP application (n ⫽ 45). Figure 1, C and D, shows ICa-voltage relationship obtained in the absence and presence of FP (16 dyn/cm2). There was no clear shift in the voltage dependence of ICa in the myocytes exposed to FP (n ⫽ 7). Figure 2, A and B, represents the effects of different levels of fluid pressures on ICa. A gradual increase in the FP from ⬃3 to ⬃12 dyn/cm2 reduced ICa in a pressure-dependent manner. Figure 2C displays the fluid pressure-response curve that was obtained after ICa had reached its steady-state level in the presence of given pressure levels in ventricular cells. The curve shows linear relationship between the level of pressure and percent suppression of ICa at a range of 3–16 dyn/cm2 (3 dyn/cm2: 4.0 ⫾ 1.4%, n ⫽ 9, P ⬎ 0.05; 7 dyn/cm2: 11 ⫾ 2.7%, n ⫽ 5, P ⬍ 0.001; 12 dyn/cm2: 19 ⫾ 1.0%, n ⫽ 5, P ⬍ 0.001; 16 dyn/cm2: 26 ⫾ 1.8%, n ⫽ 45, P ⬍ 0.001). In the next series of experiments, we tested the effects of FP with varied durations on the ICa. The FP of 16 dyn/cm2 was sequentially applied for 5 and 20 s and for the periods required to achieve a maximal effect (max) in the same myocytes. Longer exposures of FP produced significantly larger effects on the ICa (5 s: 5.28 ⫾ 0.47%; 20 s: 9.11 ⫾ 0.46%; max: 22.9 ⫾ 1.4%, n ⫽ 7; Fig. 2, D–F). Acceleration of the inactivation of Ca2⫹ channel by FP. To examine whether the FP exposure modulates the inactivation property of the Ca2⫹ channel, we evaluated the inactivation time constant (␶) of ICa (see MATERIALS AND METHODS) in the absence and presence of FP. Figure 3A, left, shows peaknormalized, superimposed ICa traces recorded in the absence and presence of FP (16 dyn/cm2), indicating more rapid inactivation of ICa under FP exposure. In 45 cells tested, mean ␶ of ICa inactivation was 17.8 ⫾ 2.0 ms in the control conditions and 14.2 ⫾ 1.1 ms under FP exposures (P ⬍ 0.01) (Fig. 3A, right), suggesting acceleration of the inactivation of Ca2⫹ channels by fluid pressure. Figure 3B illustrates the effect of FP (16 dyn/cm2) on the voltage dependence of the availability of Ca2⫹ channels. The FP caused an approximate 4-mV hyperpolarizing shift in the V0.5 of the steady-state inactivation curve (see MATERIALS AND METHODS) from ⫺25.4 ⫾ 0.9 to ⫺29.3 ⫾ 1.1 mV (n ⫽ 5, P ⬍ 0.001) without changing the slope factor (control, 3.9 ⫾ 0.2 mV; FP, 4.1 ⫾ 0.1 mV, n ⫽ 5, P ⬎ 0.05). It appeared that ICa, recorded at the end of depolarization, shifted to the positive value under FP. To confirm that the positive shift of the currents by FP was purely caused by a modulation of the Ca2⫹ channels we further tested the effect of FP on a Cd2⫹-sensitive current. Cd2⫹ blocked the inward ICa without changing holding currents (Fig. 4A). The Cd2⫹-sensi-

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significantly affect the amplitude (Fig. 5, B and E) and inactivation kinetics of ICa by themselves. These results indicate that the reduction of ICa and acceleration of ICa inactivation during FP exposure are independent of the function of stretch-activated ion channels. Effects of FP on IBa . L-type ICa can be decreased by rise in the cytosolic Ca2⫹ concentration (7, 8, 31). We examined, in the next series of experiments, whether FP directly inhibits L-type Ca2⫹ channel or indirectly suppresses it by increasing the cytosolic Ca2⫹ concentration using extracellular Ba2⫹ as a charge carrier for the Ca2⫹ channels. Replacement of Ca2⫹ by Ba2⫹ in the extracellular medium eliminates CICR and Ca2⫹dependent inactivation so that Ba2⫹-conducting Ca2⫹ channels are inactivated in a voltage-dependent manner. Although IBa was reduced by FP (control, 19.5 ⫾ 1.6 pA/pF; 16 dyn/cm2 FP, 18.7 ⫾ 1.5 pA/pF; n ⫽ 6; P ⬍ 0.01) the inhibitory effect of FP on IBa was much smaller than that on the ICa (compare Fig. 1B and Fig. 6B). This result suggests that the inhibitory effect of FP on ICa is mediated by a possible enhancement of the ICa-triggered Ca2⫹ releases but not by direct action on the Ca2⫹ channel. Effect of FP on intracellular Ca2⫹ concentration without and with GsMTx-4. We further tested whether the inhibitory effect of the FP on ICa was caused by an increase of the cytosolic Ca2⫹ concentration by directly measuring the intracellular Ca2⫹ signal using confocal imaging. Figure 7 illustrates the effects of the FP on cytosolic Ca2⫹ transients in field-stimulated (at 1 Hz) rat ventricular myocytes. A FP of 16 dyn/cm2 significantly increased the systolic Ca2⫹ level as well as the magnitude of Ca2⫹ transients (Fig. 7, A and B). ImmeAJP-Cell Physiol • VOL

diately after the exposure to the FP, the basal Ca2⫹ level slightly increased (see the blue arrow in Fig. 7B), which appeared to be associated with the subsequent occurrence of a larger Ca2⫹ transient (see the green star in Fig. 7B). In seven cells examined, the average magnitude of the Ca2⫹ transients (⌬F/F0) increased from 1.09 ⫾ 0.15 to 1.48 ⫾ 0.18 (n ⫽ 7, P ⬍ 0.01; Fig. 7C, left). To confirm the increase of the basal Ca2⫹ level by the FP exposure, the amount of the increase in the background fluorescence measured in the control conditions (Fig. 7D, open square) was subtracted from the change in the basal Ca2⫹ level in the cells under FP exposure (Fig. 7D, filled square), as a local exposure of the laser beam in a long-term experiment appears to cause a gradual rise in the background fluorescence (Fig. 7D, open square). The increases in the basal Ca2⫹ level (F/F0) were significantly larger in the cells exposed to FP (n ⫽ 9) compared with those not exposed to FP (n ⫽ 11) (P ⬍ 0.01; compare the open and filled squares in Fig. 7D). The FP-induced pure change in the basal Ca2⫹ concentration that was obtained after the subtraction of the background fluorescence change revealed a partial recovery of the basal Ca2⫹ level after withdrawal of the FP (see triangle in Fig. 7D). The rate of decay of the Ca2⫹ transient, estimated as the decay time constant (␶), was not altered by the FP (control, 0.35 ⫾ 0.09 s; FP, 0.36 ⫾ 0.08 s, P ⬎ 0.05, n ⫽ 5). Development of a global Ca2⫹ wave at this pressure level was not found in rat ventricular myocytes, in contrast to rat atrial myocytes (20). To further examine whether the stretch-activated channel is involved in the FP-induced changes in the intracellular Ca2⫹ we carried out the same experiments and analyses in the

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Fig. 4. Determination of the Cd2⫹-sensitive component of the current and its reduction by FP in rat ventricular myocytes. A: superimposed currents recorded before and after the application of 200 ␮M Cd2⫹. Membrane voltage was clamped from ⫺40 to 0 mV for 120 ms. B: after recovery from the first trial of Cd2⫹, FP was applied to the same cell. After a maximal effect of FP was achieved (see the current indicated by “FP”) Cd2⫹ was additionally applied to the cell (see “FP ⫹ Cd2⫹”). C: superimposed currents measured in the absence (“Control”) and presence of FP, which are shown in A and B. D: Cd2⫹-sensitive currents recorded before (“Control ⫺ Cd2⫹”) and after application of FP [“FP ⫺ (FP ⫹ Cd2⫹)”] were superimposed. E: comparison of average Cd2⫹-sensitive currents measured in the presence and absence of FP. **P ⬍ 0.01, “Control ⫺ Cd2⫹” vs. “FP ⫺ (FP ⫹ Cd2⫹)” (n ⫽ 5).


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presence of 5 ␮M GsMTx-4. The 5 ␮M GsMTx-4 did not significantly alter the Ca2⫹ transient and the basal Ca2⫹ level (data not shown). Figure 7, E and F, shows confocal line scan images, recorded in a GsMTx-4-preincubated ventricular myocytes, and time course of fluorescent signal averaged from the images, respectively. The application of FP (16 dyn/cm2) gradually increased systolic Ca2⫹ level and Ca2⫹ transient amplitude, which was followed by a big increase of basal Ca2⫹ level with reduced Ca2⫹ transient amplitude. The decline of the

Ca2⫹ transient magnitude at late phase of FP application was also observed in cells not exposed to GsMTx-4 (Fig. 7B). In 13 cells preexposed to GsMTx-4, the onsets of the basal Ca2⫹ increases by FP exposures were somewhat various. However, the FP continued to increase the peaks and magnitudes of the Ca2⫹ transients as well as the basal Ca2⫹ concentrations in all the cells tested (GsMTx-4 vs. GsMTx-4 ⫹ FP; n ⫽ 13; P ⬍ 0.01; Fig. 7, E–H). Maximal increases in the systolic Ca2⫹ and Ca2⫹ transients under FP were not altered by GsMTx-4 (com-

Fig. 6. Effects of FP on Ba2⫹ current (IBa) in rat ventricular myocytes. A: time course of the changes in IBa by FP in ventricular myocytes. Inset, superimposed current traces recorded under control condition (“C”) and after exposure to FP. The horizontal bar indicates period of FP exposure. B: comparison of mean ICa between the control and FP (16 dyn/cm2) (n ⫽ 6). **P ⬍ 0.01, Control vs. FP.


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Fig. 5. Effect of FP on ICa in ventricular myocytes preexposed to the blockers of stretch-activated channels, streptomycin and GsMTx-4. A and D: time courses of the changes in the peak ICa by FP (16 dyn/cm2) in cells preexposed to 40 ␮M streptomycin (A) or 5 ␮M GsMTx-4 (D). Inset in A shows superimposed current traces recorded immediately before (“C”) the exposures to 40 ␮M streptomycin (“Strep”) and to FP plus strep and at the time when a maximal decrease in ICa by FP was observed (“FP ⫹ strep”). Inset in D shows superimposed current traces recorded before (“GsMTx-4”) and after application of FP in the presence of 5 ␮M GsMTx-4 (“GsMTx-4 ⫹ FP”). B and E: comparison of mean peak ICa. **P ⬍ 0.01, Strep vs. Strep ⫹ FP (16 dyn/cm2) (n ⫽ 5). ***P ⬍ 0.001, GsMTx-4 vs. GsMTx-4 ⫹ FP (n ⫽ 15). C and F: superimposed currents that were normalized to their peaks.

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Fig. 7. Effects of FP on Ca2⫹ transients in rat ventricular myocytes. A and E: confocal linescan images recorded from a representative field-stimulated (1 Hz) myocyte in the absence (A) and presence of 5 ␮M GsMTx-4 (E). Inset displays the image of myocyte and location of the line used for scanning. The 8-s-long line scan images, recorded at 16.4 ms/line, were acquired at 213-ms intervals. The traces of local Ca2⫹ transients in B and F show F/F0 measured from the line scan images in A and E, respectively. The horizontal box indicates period of FP exposure. F0, average basal fluorescence intensity measured before the exposure to FP; F, average fluorescence of each line. Dotted line indicates basal Ca2⫹ level before the application of FP. C and G: comparisons of average amplitudes of Ca2⫹ transients (⌬F/F0) and mean systolic Ca2⫹ level (F/F0) measured before (“C”) and after application of FP (16 dyn/cm2) without (C) and with 5 ␮M GsMTx-4 (G). **P ⬍ 0.01 (n ⫽ 7; C), *P ⬍ 0.05 (n ⫽ 13; G), control vs. FP. D and H: effects of FP on the basal Ca2⫹ level in the absence and presence of GsMTx-4. 䊐 and ■ indicate time-dependent changes of the background fluorescence intensities measured in cells not exposed to FP and in cells that were exposed to FP, respectively; ‚ indicates FP-induced net change in the basal Ca2⫹ concentration. **P ⬍ 0.01, ***P ⬍ 0.001, no FP (n ⫽ 11) vs. FP trial (n ⫽ 9) in the absence of GsMTx-4. *P ⬍ 0.05, **P ⬍ 0.01, no FP vs. FP trial in the presence of GsMTx-4 (n ⫽ 13).

pare Fig. 7, C and G). It was noted that the FP-induced basal Ca2⫹ increase was larger in the presence of GsMTx-4 than that in the control (compare the triangles in the D and H). In addition, it did not show a recovery after the subtraction of the background fluorescence (see the triangle in Fig. 7H). These results suggest that FP enhances Ca2⫹-induced Ca2⫹ release independently of stretch-activated ion channel in rat ventricular myocytes. Role of SR Ca2⫹ release in the effects of FP on ICa. To further confirm whether FP-induced ICa reduction was caused by increase in the cytosolic Ca2⫹ concentration, the effects of FP on ICa were examined in highly Ca2⫹-buffered myocytes. Given that dyadic Ca2⫹ releases are thought to be resistant to ⬃15 mM EGTA in ventricular myocytes and that ⬃10 mM AJP-Cell Physiol • VOL

BAPTA is required to suppress the releases (5), the effects of FP were tested in the myocytes dialyzed with 10 mM BAPTAcontaining internal solutions. In those myocytes, FP continued to reduce ICa by ⬃6% (n ⫽ 6, P ⬍ 0.01; Fig. 8, A and B), and thus the inhibitory effect of FP on ICa was attenuated by ⬃80% in the highly Ca2⫹-buffered cells. In these myocytes, the FP-induced acceleration of ICa inactivation was no longer observed (Fig. 8C). The blockade of the effect of FP on ICa inactivation by intracellular application of 10 mM BAPTA (Fig. 8) indicates a role of cytosolic Ca2⫹ increase in accelerating the ICa inactivation during FP. In the next series of experiments, we tested the effect of FP on current inactivation in the presence of ryanodine and thapsigargin, abolishing the release of SR. The

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DISCUSSION

effects of FP on ICa, measured before and after depletion of SR with 2 ␮M thapsigargin, were compared in the same cells. The inhibitory effects of FP on ICa under control conditions were significantly attenuated after depleting the SR (percent suppression: control, 26 ⫾ 1.8% vs. in thapsigargin, 10 ⫾ 1.8%; n ⫽ 5; P ⬍ 0.05; Fig. 9A). In the cells preexposed to 20 ␮M ryanodine, the FP continued to suppress the ICa (12 ⫾ 2.3%; n ⫽ 6; P ⬍ 0.05; Fig. 9C, right traces), although percent suppression was significantly smaller than that in the absence of ryanodine. The inactivation of ICa was significantly slowed down when the Ca2⫹ stores of the SR were depleted by 2 ␮M thapsigargin (␶: control, 17.1 ⫾ 2.3 ms vs. thapsigargin, 26.1 ⫾ 3.2 ms; n ⫽ 5; P ⬍ 0.01), and they were depleted when the SR Ca2⫹ releases were inhibited by 20 ␮M ryanodine (␶: control, 16.5 ⫾ 2.1 ms vs. ryanodine, 28.2 ⫾ 3.7 ms; n ⫽ 6; P ⬍ 0.01) (Fig. 9, A and C). The ICa remained unaltered or slightly reduced in the presence of thapsigargin (2 ␮M) or ryanodine (20 ␮M) (Fig. 9, A and C). Additional applications of FP in the presence of thapsigargin or ryanodine did not significantly alter the ␶ of the ICa inactivation (Fig. 9B; thapsigargin only, 26.1 ⫾ 3.2 ms vs. thapsigargin ⫹ FP, 25.8 ⫾ 2.7 ms; n ⫽ 5; P ⬎ 0.05; ryanodine only, 28.2 ⫾ 3.7 ms vs. ryanodine ⫹ FP, 27.0 ⫾ 2.2 ms; n ⫽ 6; P ⬎ 0.05). These results suggest that the larger SR Ca2⫹ release during depolarization is responsible for the rapid inactivation of ICa under FP stimulation and that it also contributes to the decrease of ICa. AJP-Cell Physiol • VOL

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Fig. 8. Effects of FP on ICa in highly Ca2⫹-buffered myocytes. A: time course of the changes in ICa by FP in myocytes dialyzed with 10 mM BAPTAcontaining internal solution. The plot represents average results (⫾SE) obtained in 6 cells. Inset, superimposed current traces recorded at the times indicated. C, control. Comparisons of mean ICa (B) and the inactivation time constants (C) between the control and FP (16 dyn/cm2) (n ⫽ 6). **P ⬍ 0.01, control vs. FP.

In the present study, we demonstrate that the rapid flow of pressurized solutions significantly reduces the L-type ICa by enhancing the Ca2⫹-induced Ca2⫹ releases and by increasing the basal Ca2⫹ concentration. This is supported by the evidence for 1) the masks of the suppressive effect of FP on the Ca2⫹ channels by replacing extracellular Ca2⫹ with Ba2⫹ as a charge carrier, or by introducing the high concentrations of Ca2⫹ buffers into the cells; 2) increases in the depolarizationinduced cytosolic Ca2⫹ transients and in the basal Ca2⫹ level during the FP exposure; and 3) reduction of the inhibitory effect of FP on ICa by depletion of SR. The significantly larger Ca2⫹ transients despite the smaller ICa under FP suggest an enhancement of the efficacy of Ca2⫹ current to induce Ca2⫹ releases in the presence of FP. In addition, resistance of the inhibitory effect of FP on the Ca2⫹ channels to the blocker of stretch-activated channel indicates that the FP/intracellular Ca2⫹/ICa signaling may be independent of the stretch-activated ionic current. The Ca2⫹-induced reduction of Ca2⫹ influx through the Ca2⫹ channels may serve as a negative regulator to prevent excess Ca2⫹ releases from the SR in cardiac cells under prolonged fluid pressure, and it may also contribute to a shortening of action potential duration. Our results demonstrate that stretch-activated ion channel is not responsible for the FP effects on Ca2⫹ signaling in ventricular myocytes (Figs. 5 and 7). In fact, significant shrinkage or stretch of the entire cell length was not found at the level of the pressure used in the present study. However, a slight decrease of the basal cell length (⬍1% of the cell length) was observed by video edge detection when the FP was increased to ⬎20 dyn/cm2 (see the supplemental data in the online version of this article), which suggests a difference in the modality of stimulus between pressurized flow and stretch. Although there was no detectable change in the whole cell length at FP ⬍16 dyn/cm2, the pressurized fluid flow may induce a local deformation of the surface membrane in the form of a concave shape or as an inward collapse of the surface membrane to some degree, which in turn could mechanically stimulate the junctional dyads in the close proximity of the surface membrane or increase ionic strength (i.e., the Ca2⫹ concentrations) in dyadic space sufficient to enhance CICR. This might influence the gain of CICR and protein-protein interactions such as those involved in cross talk between the L-type Ca2⫹ channels and RyRs via Ca2⫹ and calmodulin in the junctional microdomain (1, 35 32, 41), in turn altering channel gating. Such shrinkage or decrease in the cell volume is usually observed in the myocytes under extracellular hypertonic conditions. Hypertonicity leads to the generation of both Ca2⫹ and tension transients in isolated ventricular muscle (2) and skeletal muscle fibers (7, 24), and it alters the anatomy of T-tubule-SR junction (7, 24). The distinguished response of the fluid pressure from that of stretch is also reflected in their different effects on the Ca2⫹ channels in isolated cardiac myocytes. In effect, there is controversy in the effects of stretch on ICa in cardiac myocytes; no changes were found in rat and guinea pig ventricular myocytes (13, 33), and a decrease in human atrial myocytes (15) has been observed. These findings are consistent with the view that the cellular sensing and responding systems to pressure/stretch and flow are distinct and separate (4).

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The specific mechanisms underlying the mechanosensitivity of ventricular myocytes to pressurized fluid flow and subsequent signaling are not clearly understood. Thus far, there have been very few reports on the mechanosensitivity of Ca2⫹ channels, particularly cardiac Ca2⫹ channels. The results of this study support the idea that cardiac Ca2⫹ channels are not directly affected by the fluid pressure but can be indirectly by changes in the intracellular Ca2⫹ concentration. The present findings regarding the cardiac L-type Ca2⫹ channel are somewhat different from the mechanosensitivity of neuronal and smooth muscle L-type Ca2⫹ channels expressed in HEK-293 cells (22, 31). The L-type IBa, measured in HEK-293 cells expressing the Ca2⫹ channel, was enhanced by the flowinduced shear forces, suggesting the mechanosensitivity of these channels (22, 31). It is plausible that distinct cell types composed of different signaling molecules and/or ultrastructures give rise to a range of effects on the Ca2⫹ channel. A gradual increase in the basal Ca2⫹ level during the exposure to the FP in rat ventricular myocytes was also found, and it may reduce the driving force of Ca2⫹ influx through the channel. It appears that the FP-induced increase in the Ca2⫹ concentration is related to the enhancement of the occurrences of Ca2⫹ sparks. The FP-induced enhancement of the Ca2⫹ spark occurrences has been observed in resting rat ventricular (S. H. Woo, unpublished observations) and atrial mycoytes (40). The mechanisms for the rise of the basal Ca2⫹ level and/or increased occurrences of spontaneous Ca2⫹ sparks are not clear, but they may be related to a possible decrease of the junctional space caused by physical changes of the surface membrane, as mentioned above. In fact, the FP did not change the SR Ca2⫹ contents in rat ventricular cells (data not shown). Removal of extracellular Ca2⫹ did not significantly alter the FP-induced basal Ca2⫹ increases in these cells (data not shown). The Ca2⫹ changes were similar in the periphery and interior of ventricular cells, whereas, in atrial cells lacking T AJP-Cell Physiol • VOL

tubules, FP enhanced the frequency of peripheral junctional sparks more dramatically than the nonjunctional central sparks (40). These evidences support the idea that the junctional SR domain is more sensitive to the FP stimulus. The differences of the FP responses in atrial and ventricular myocytes include a smaller change of the basal Ca2⫹ level in ventricle cells compared with atrial cells (21) and the propensity of the occurrences of Ca2⫹ waves in atrial myocytes. Less development of the central Ca2⫹ wave in the ventricular myocytes may be explained by the presence of Ca2⫹ efflux mechanism within the depth of ventricular myocytes (presence of t-tubules), in contrast to the atrial myocytes (6, 37). Assuming that the stiffness of a myocyte is 23 kPa (45), and that the maximum FP applied is 0.0016 kPa (equivalent to 16 dyn/cm2), the strain (fractional cell lengthening) on a myocyte calculated by the equation (45) [stress (kPa) ⫽ A/k ⫻ (ek␧ ⫺ 1) (where A ⫽ 23 kPa, k ⫽ 16, and ␧ is the strain)] is very small compared with the length changes encountered in vivo during a normal contraction. Surprisingly, however, a pressurized jet of fluid appears to have significantly large effects on the intracellular Ca2⫹ concentration and ICa in isolated single ventricular myocytes, which suggests that the myocytes sense relatively low FP stimuli and may amplify this stimulus through the activation of an intracellular signaling pathway to generate such responses. It should also be noted that in other types of cells such as endothelial or hair cells, flow pressures in the range of 0.1–30 dyn/cm2 change functions of the membrane channels and intracellular ionic concentrations (4, 14, 30). Although the level of fluid pressure that each myocyte would receive in an intact ventricular chamber wall during aortic regurgitation or aortic stenosis is unclear, the acute and chronic consequences of such aortic valve diseases are an increase of the diastolic pressure in the ventricle leading to adaptive left ventricle and atrium enlargement and augmentation of the stroke volume during systole (42). Interestingly, the

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Fig. 9. Effects of FP on ICa in the presence of thapsigargin (A and B) and ryanodine (C and D). Superimposed ICa traces that were recorded under the conditions indicated are shown. 2 ␮M thapsigargin (Thap) and 20 ␮M ryanodine (Ry) were pretreated for ⬃5 min when the effects of the drugs reached to the steady. B and D: currents were superimposed after normalizing to their peaks, showing no change of inactivation kinetics by FP in the presence of thapsigargin or ryanodine.


ACKNOWLEDGMENTS The authors thank Dr. Joung Real Ahn for developing the image analysis program. GRANTS This work was supported by the Korea Research Foundation Grant funded by the Korean Government (MOEHRD) (KRF-2004-041-E00017), by grant from Korea Ministry of Science and Technology (Systems Biology Research Grant, M10503010001-05N0301-00113), and in part by the Korea Science and Engineering Foundation (KOSEF) grant funded by the Korea Government (MOST) (No. R01-2007-000-10796-0). REFERENCES 1. Adachi-Akahane S, Cleemann L, Morad M. Cross-signaling between L-type Ca2⫹ channels and ryanodine receptors in rat ventricular myocytes. J Gen Physiol 108: 435– 454, 1996. 2. Allen DG, Smith GL. The effects of hypertonicity on tension and intracellular calcium concentraion in ferret ventricular muscle. J Physiol 383: 425– 439, 1987. 3. Beuckelmann DJ, Wier WG. Mechanism of release of calcium from sarcoplasmic reticulum of guinea-pig cardiac cell. J Physiol 405: 233–255, 1988. 4. Bevan JA, Laher I. Pressure and flow-dependent vascular tone. FASEB J 5: 2267–2273, 1991. 5. Bowman CL, Gottlieb PA, Suchyna TM, Murphy YK, Sachs F. Mechanosensitive ion channels and the peptide inhibitor GsMTx-4: history, properties, mechanisms and pharmacology. Toxicon 49: 249 –270, 2007. 6. Carl LS, Felix K, Caswell AH, Brandt NR, Ball WJ, Vaghy PL, Meissner G, Ferguson DG. Immunolocalization of sarcolemmal dihydropyridine receptor and sarcoplasmic reticular triadin and ryanodine receptor in rabbit ventricle and atrium. J Cell Biol 129: 673– 682, 1995. 7. Chawla S, Skepper JN, Hockaday AR, Huang CL. Calcium waves induced by hypertonic solutions in intact frog skeletal muscle fibres. J Physiol 536: 351–359, 2001. 8. Conwell J, Cocalis M, Erickson L. EAT to the beat: ‘ectopic’ atrial tachycardia caused by catheter whip. Lancet 342: 740, 1993. AJP-Cell Physiol • VOL

9. Fabiato A. Calcium-induced release of calcium from the cardiac sarcoplasmic reticulum. Am J Physiol Cell Physiol 245: C1–C14, 1983. 10. Franz MR, Burkhoff D, Yue DT, Sagawa K. Mechanically induced action potential changes and arrhythmia in isolated and in situ canine hearts. Cardiovasc Res 23: 213–223, 1989. 11. Fukaya Y, Ohhashi T. Acetylcholine- and flow-induced production and release of nitric oxide in arterial and venous endothelial cells. Am J Physiol Heart Circ Physiol 270: H99 –H106, 1996. 12. Hamil OP, Marty A, Neher E, Sakmann B, Sigworth FJ. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflu¨gers Arch 319: 85–100, 1981. 13. Hongo K, White E, Le Guennec JY, Orchard HC. Changes in [Ca2⫹]i, [Na⫹]i and Ca2⫹ current in isolated rat ventricular myocytes following an increase in cell length. J Physiol 491: 609 – 619, 1996. 14. Jen CJ, Jhiang SJ, Chen HI. Cellular responses to mechanical stress: effects of flow on vascular endothelial intracellular calcium signaling of rat aortas ex vivo. J Appl Physiol 89: 1657–1662, 2000. 15. Kamkin A, Kiseleva I, Wagner KD, Bohm J, Theres H, Gunther J, Scholz H. Characterization of stretch-activated ion currents in isolated atrial myocytes from human hearts. Pflu¨gers Arch 446: 339 –346, 2003. 16. Kass RS, Sanguinetti MC. Inactivation of calcium channel current in the calf cardiac Purkinje fiber. Evidence for voltage- and calcium-mediated mechanisms. J Gen Physiol 84: 705–726, 1984. 17. Lab MJ. Mechanoelectric feedback (transduction) in heart: concepts and implications. Cardiovasc Res 32: 3–14, 1996. 18. Lakatta EG. Cardiovascular regulatory mechanisms in advanced age. Physiol Rev 73: 413– 467, 1993. 19. Lee KS, Marba´n E, Tsien RW. Inactivation of calcium channels in mammalian heart cells: joint dependence on membrane potential and intracellular calcium. J Physiol 364: 395– 411, 1985. 20. Lerman BB, Burkhoff D, Yue DT, Franz MR, Sagawa K. Mechanoelectric feedback: independent role of preload and contractility in modulation of canine ventricular excitability. J Clin Invest 76: 1843–1850, 1985. 21. Levine JH, Guarnieri T, Kadish AH, White RI, Calkins H, Kan JS. Changes in myocardial repolarization in patients undergoing balloon valvuloplasty for congenital pulmonary stenosis: evidence for contractionexcitation feedback in humans. Circulation 77: 70 –77, 1988. 22. Lyford GL, Strege PR, Shepard A, Ou Y, Ermilov L, Miller SM, Gibbons SJ, Rae JL, Szurszewski JH, Farrugia G. ␣1C (Cav1.2)L-type calcium channel mediates mechanosensitive calcium regulation. Am J Physiol Cell Physiol 283: C1001–C1008, 2002. 23. Marrannes R, Prins ED. Computer programs to facilitate the estimation of time-dependent drug effects on ion channels. Comput Methods Programs Biomed 74: 167–181, 2004. 24. Martin CA, Petousi N, Chawla S, Hockaday AR, Burgess AJ, Fraser JA, Huang CLH, Skepper JN. The effect of extracellular tonicity on the anatomy of triad complexes in amphibian skeletal muscle. J Muscle Res Cell Motil 24: 407– 415, 2003. 25. Na¨bauer M, Callewaert G, Cleemann L, Morad M. Regulation of calcium release is gated by calcium current, not gating charge, in cardiac myocytes. Science 244: 800 – 804, 1989. 26. Nazir SA, Lab MJ. Mechanoelectric feedback and atrial arrhythmias. Cardiovasc Res 32: 52– 61, 1996. 27. Niggli E, Lederer WJ. Voltage-independent calcium release in heart muscle. Science 250: 565–568, 1990. 28. Noris M, Morigi M, Donadelli R, Aiello S, Foppolo M, Todeschini M, Orisio S, Remuzzi G, Remuzzi A. Nitric oxide synthesis by cultured endothelial cells is modulated by flow conditions. Circ Res 76: 536 –543, 1995. 29. Ohmori H. Mechanical stimulation and fura-2 fluorescence in the hair bundle of dissociated hair cells of the chick. J Physiol 399: 115–137, 1988. 30. Olesen SP, Clapham DE, Davies PF. Haemodynamic shear stress activates a K⫹ current in vascular endothelial cells. Nature 331: 168 –170, 1988. 31. Peng SQ, Hajela RK, Atchison WD. Fluid flow-induced increase in inward Ba2⫹ current expressed in HEK293 cells transiently transfected with human neuronal L-type Ca2⫹ channels. Brain Res 1045: 116 –123, 2005. 32. Pitt GS, Zuhlke RD, Hudmon A, Schulman H, Reuter H, Tsien RW. Molecular basis of calmodulin tethering and Ca2⫹-dependent inactivation of L-type Ca2⫹ channels. J Biol Chem 276: 30794 –30802, 2001.

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observations of this study of the increases in the basal Ca2⫹ level and Ca2⫹ transient amplitudes during a continuous application of fluid pressure may partly explain the increases in the ventricular diastolic pressure and stroke volume that have been reported clinically. Considering that the FP generated during aortic regurgitation is dependent on the aortic orifice and the pressure gradient across the aortic valve during diastole, it may be even more difficult to estimate the fluid jet pressure on single myocytes in vivo. The data here provide evidence that cardiac muscle may directly sense a pressurized fluid flow and respond to the stimulus. In whole heart preparations and in the in situ heart of a human and a dog, a rise in ventricular pressure also leads to shortenings of the action potential duration and an effective refractory period (10, 20, 21). However, in a ventricle tissue preparation and in single ventricular myocytes, membrane stretch did not consistently decrease the action potential duration: showing both an increase and a decrease (39, 44). It is proposed that although there are other components involved in the action potential repolarization, the FP-induced decrease in the ICa can contribute to a shortening of the refractory period and/or to arrhythmia in a ventricular chamber wall exposed to the hemodynamic forces of fluid (e.g., a blood jet to the ventricular wall during aortic regurgitation, or the excessive pressure produced during aortic valve stenosis). Additional studies revealing the cellular and molecular mechanisms by which the fluid pressure is sensed and transduced by cardiac myocytes remain to be undertaken.

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