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Cardiovascular Research 2006 72(3):403-411; doi:10.1016/j.cardiores.2006.08.011
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Copyright © 2006, European Society of Cardiology

Membrane potential of rat ventricular myocytes responds to axial stretch in phase, amplitude and speed-dependent manners

Satoshi Nishimuraa, Yasuo Kawaib, Toshiaki Nakajimaa,c, Yumiko Hosoyaa, Hideo Fujitaa, Masayoshi Katoha, Hiroshi Yamashitaa, Ryozo Nagaia and Seiryo Sugiurab,*

aDepartment of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Japan
bThe Institute of Environmental Studies, Graduate School of Frontier Sciences, The University of Tokyo, Hongo 7–3-1, Bunkyo-ku, Tokyo 113–0033, Japan
cDepartment of Ischemic Circulatory Physiology, The University of Tokyo, Japan

* Corresponding author. Tel./fax: +81 3 5841 8393. Email address: sugiura{at}k.u-tokyo.ac.jp

Received 30 May 2006; revised 16 August 2006; accepted 18 August 2006


   Abstract
Top
 Abstract
1. Introduction
 2. Methods
 3. Results
 4. Discussion
 References
 
Objective: To elucidate the interdependence between the mechanical state of the myocardium and its electrical activity, previous studies have been performed at the cellular level. However, the information to date has been limited by the technical difficulties associated with stretching single myocytes.

Methods: We solved this problem by combining two techniques, namely a carbon fiber technique for stretching rat myocytes with wide ranges of amplitude and speed, and ratiometric measurement of a fluorescent indicator (di8-ANEPPS) for evaluating the membrane potential in the non-contact mode.

Results: During systole, stretching caused depolarization that prolonged the action potential duration without affecting the peak amplitude, but the effect was only significant in the late phase. Application of a stretch to quiescent myocytes depolarized the membrane potential in amplitude- and speed-dependent manners, but the response was suppressed by cytochalasin D treatment, suggesting participation of the cytoskeleton in the mechanotransduction mechanism. Finally, ion replacement experiments revealed that although Na+ was the dominant charge carrier for large amplitude stretches, Ca2+ permeation was involved in small amplitude stretches, suggesting amplitude-dependent ion selectivity.

Conclusions: Application of axial stretching to rat ventricular myocytes changed the membrane potential in phase-, amplitude- and speed-dependent manners. Amplitude may also modulate the ion selectivity of stretch-activated channels.

KEYWORDS Membrane potential; Myocytes; Stretch; m–e coupling


   1. Introduction
Top
 Abstract
 1. Introduction
2. Methods
 3. Results
 4. Discussion
 References
 
It is well known that changes in the mechanical state of the myocardium affect its electrical activity, a phenomenon referred to as "mechano-electric feedback" [1–3]. Although this transduction mechanism underlies the physiological length–tension relationships, it could also be a cause of stretch-induced arrhythmias due to abnormal loading on the myocardium in diseased conditions, such as congestive heart failure, myocardial infarction or commotio cordis [4–6]. Since stretch-activated channels (SACs) are considered to play a central role in this mechanoelectric transduction, a number of studies have attempted to characterize these channels. At the cellular level, SAC activities have been identified in cardiomyocytes from the chick embryo ventricle [7,8], mammalian atrium [9–11] and mammalian ventricle [12] by applying negative pressure to a cell-attached patch pipette. More recently, Browe and Baumgarten electro-magnetically pulled the β1-integrin in the sarcolemma of rabbit ventricular myocyte to demonstrate that chloride current is also mechanosensitive [13]. To date, however, there are no single channel recording data available for adult ventricular myocytes, and only a few studies have reported the response of cardiomyocytes to axial stretching [14–16]. Furthermore, considering the dynamic changes in stress and strain that each myocyte experiences in the beating heart, the effects of the timing and speed of stretching should also be elucidated, but the use of a patch pipette greatly hampers such attempts. The influence of the mechanical state on myocardial electrical activity has also been studied at the ventricular level by injecting a volume of solution or raising the pressure of the ventricle [4,17–20] and, indeed, some of these studies have clearly demonstrated an impact of the timing and/or speed of the stretching on the monophasic action potential [18,19,21]. However, considering the complex morphology and fiber structure of the ventricle, it may be difficult to establish a direct link between volume/pressure loading and the deformation of individual myocytes.

Recently, we developed an experimental carbon fiber technique for stretching single cardiomyocytes over wide ranges of speed and amplitude [22]. On the other hand, a non-contact mode of membrane potential measurement has already been made possible using fast voltage-sensitive dyes. Although measurement of emitted light from dyes is sensitive to motion artifacts caused by active contractions [23–25], simultaneous detection at two emission wavelengths and ratiometric evaluation have been shown to effectively remove the motion artifacts in hippocampal neurons [26], endothelial cells [25], whole heart preparations [24,27].

In the present study, we combined these two techniques, i.e., the carbon fiber technique for stretching single cardiomyocytes and use of a voltage-sensitive fluorescent indicator for membrane potential measurement, to record the changes in the membrane potential of single rat ventricular myocytes in response to axial stretching at varying times, amplitudes and speeds. We demonstrate that all of these factors affect the membrane potential in complex manners. The obtained results not only provide insights into the characteristics of SACs, but also establish a link between cellular level events and pathophysiology at the organ level.


   2. Methods
Top
 Abstract
 1. Introduction
 2. Methods
3. Results
 4. Discussion
 References
 
2.1. Isolation of cardiomyocytes
Single ventricular myocytes were obtained from 7-week-old female Wistar rats. We used animals of this age because of the following reasons, 1) because myocardial tissue of older animals is fibrotic thus requires higher dose of collagenase to digest, the isolated myocytes become irritable and cannot tolerate the large and quick stretches, 2) we can find a report on stretch-activated channel in cardiac myocytes isolated from rats of similar age group [14]. The heart was quickly removed under phenobarbital anesthesia and cells were enzymatically isolated using a modified dispersion technique as described previously [22]. Retrograde perfusion was initiated with nominally Ca2+-free HEPES-Tyrode solution (130 mmol/L NaCl, 5.4 mmol/L KCl, 0.5 mmol/L MgCl2, 0.33 mmol/L NaH2PO4, 22 mmol/L glucose, 5 mmol/L glutamine, 0.4 mmol/L EGTA, 25 mmol/L HEPES, pH 7.4) at 37°C. After 5 min, the perfusate was switched to an enzyme solution containing 1 mg/mL collagenase (Type 2; Worthington), 0.05 mg/mL protease (Type XIV; Sigma) and 0.05 mg/mL trypsin (Sigma) and maintained for 20 min. Finally, the enzymes were washed out and the calcium concentration of the Tyrode solution was gradually increased to 1.1 mmol/L. The mean sarcomere length of the isolated cardiomyocytes was 1.96±0.01 µm (n=14 cells), as measured by on-line Fourier analysis of the striation pattern at the extracellular calcium concentration of 1.1 mmol/L. Similar sarcomere length has been reported in multicellular preparations at slack length [28]. The cells were transferred to an experimental chamber. All experiments were performed at 37°C. The investigation conforms with the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health.

2.2. Loading of the membrane voltage-sensitive dye di8-ANEPPS
Our preliminary studies and previous reports indicated that high-dose (30–60 µM) loading of di-8-butyl-amino-naphthyl-ethylene-pyridinium-proply-sulfonate (di8-ANEPPS; Molecular Probes) causes photodynamic damage [26,29] to isolated myocytes. However, we succeeded in preventing the damage generated by reactive oxygen species by limiting the loading dosage of the dye, reducing the illumination wattage and adding a combination of antioxidants. Cells were loaded with 5 µM di8-ANEPPS for 8 min, and then washed twice with Tyrode solution containing 800 U/mL catalase (Sigma), 40 U/mL glucose oxidase (Sigma) and 100 µM Trolox C (Aldrich) [26,29]. The di-8-ANEPPS-loaded cardiomyocytes showed stable contraction in the presence of the antioxidants. Furthermore, the addition of the antioxidants did not affect the contractility, intracellular calcium concentration (measured by Indo1-AM) or morphology of the treated cells, as compared to untreated myocytes (data not shown).

2.3. Optical voltage measurement system
Fast voltage-sensitive dyes are known to show voltage-dependent shifts in their emission spectra [30,31]. In order to measure the membrane potential without interference from motion artifacts, optical measurements were taken at two emission wavelengths (ratiometric recording). After excitation of the dye (di8-ANEPPS) at 488 nm with a mercury light, the emission light (>550 nm) was separated using a secondary dichroic beam-splitter (570 nm), filtered (560 and 620 nm, respectively) and measured by dual photodetectors. The signals were recorded at a sampling rate of 1 kHz without filtering. The ratio of the fluorescence intensities (fluorescence at 620 nm relative to that at 560 nm) was calculated, and normalized using the steady-state value in diastole as R560/R620. The cells were visualized simultaneously for measurement of the cell and sarcomere lengths by bright illumination above 650 nm. A diagram of the experimental setup is shown in Fig. 1.


Figure 1
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  		Fig. 1 Diagram of the experimental setup. A pair of carbon fibers was attached to the ends of a myocyte, and the thin fiber was controlled by a piezoelectric translator (PZT) connected to a personal computer (PC). The positions of the attached carbon fibers were detected by a charge-coupled device camera (CCD), and the sarcomere length was determined by fast Fourier transformation (FFT). The details of the optical systems were described in the Methods. Briefly, the dyes were excited at 488 nm, and the emission signal was recorded ratiometrically by two photodetectors (560 and 620 nm). A long wavelength (>650 nm) was used for the CCD observation.

 
2.4. Length measurement and control system
Tensile stretch was applied using a cell adhesive carbon fiber-based system as described previously [22,32]. Briefly, a rod-shaped quiescent single cardiomyocyte with a sarcomere length longer than 1.85 µm, as measured by the on-line Fourier analysis of optical density traces of the sarcomere pattern of the myocyte image (IonOptix), was selected under a microscope and a pair of carbon fibers was attached to its ends using micromanipulators. One fiber was compliant (diameter: 7 µm; length: 1–1.2 mm; stiffness: 80–200 nN/µm), while the other was thick and rigid (diameter: 30 µm; length: ~1 mm; stiffness: >1000 nN/µm) and served as a mechanical anchor. The position of the compliant fiber was controlled by a piezoelectric translator (PZT; P-841.40; Physik Instrumente) fixed to it, and the cell was stretched by pulling this carbon fiber via appropriate command signals generated by a PC. Sarcomere length was measured as described above and the cell length (segment length between the two fibers) was monitored by projecting an image of the attached carbon fibers onto a linear photodiode array (S3903; Hamamatsu Photonics).

2.5. Calibration
To calibrate the fluorescence signal, direct electrode measurements were performed simultaneously with the ratiometric fluorescent measurement under various extracellular K+ concentrations (5.4, 10, 20, 60, 100 mmol/L). The patch–clamp technique was used in the whole cell configuration using a patch–clamp amplifier (EPC-7; List Electronics, Darmstadt, Germany) [33]. The patch pipette contained (in mmol/L): KCl 140, MgCl2 2, EGTA 5, Na2-ATP 3, GTP 0.1, HEPES 10, pH 7.4. Fig. 2 shows the relation between R560/R620 and the directly measured membrane potential (Fig. 2). Applying linear regression, we obtained the following equation for the calibration:


Figure 2
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  		Fig. 2 Relationship between the fluorescence ratio (R560/R620) and the membrane potential directly measured by electrode. Fluorescent ratio and membrane potential were measured under various K+ concentrations (5.4, 10, 20, 60, 100 mmol/L, n=5 cells for each). Broken line indicates the linear regression of the data.

 


Formula

2.6. Stretch protocols

1. Phase effect. Cardiomyocytes were electrically paced at 0.5 Hz (duration 7 ms and 20% above threshold) and stretched (2 µN) at various timings after pacing by programmed commands. Two µN force corresponds to 23.6+1.6 mN/mm2 when normalized by a cross sectional area thus being comparable but a little smaller than the preload at Lmax reported for the multicellular preparations [28,34].
2. Amplitude effect. Stretches of various amplitudes were applied to quiescent (without pacing) myocytes for 500 ms. The sequence (amplitude) was randomized. In this protocol, the degree of stretching was quantified by the change in sarcomere length. The experiments were repeated under the following three conditions: 1) control condition (Tyrode solution Ca2+: 1.1 mmol/L); 2) Ca2+-free condition (0.4 mmol/L EGTA without added Ca2+); and 3) Ca2+-free and low Na+ condition (Na+ was replaced by choline chloride, 0.4 mmol/L EGTA without added Ca2+). We also studied myocytes treated with 20 µmol/L cytochalasin D (Wako Chemicals) for 1 h under the control condition. In protocol 1 and 2, step command was applied for stretching the myocyte resulting in the strain rate of >10/s (>19 µm/s in sarcomere length).
3. Speed effect. To examine the effect of the stretching speed, 10% stretching in sarcomere length was applied at strain rates of 1.50/s (2.94 µm/s in sarcomere length), 0.75/s (1.47 µm/s) and 0.25/s (0.49 µm/s). The sequence was randomized.

2.7. Data analysis
All the data were sampled at 1 kHz, and recorded with an A/D converter connected to a PC (MacLab 8s; ADInstruments). To determine the R560/R620 ratio from the noisy fluorescent signals, we used 9-point symmetrical moving average. The results are expressed as the mean+SEM. Values of P<0.05 were considered significant.


   3. Results
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
4. Discussion
 References
 
3.1. Ratiometric recording of the membrane potential
Fig. 3 shows the sarcomere length changes (Fig. 3A), di8-ANEPPS (fast voltage-sensitive dyes) fluorescence signal at 620 nm (Fig. 3B), and fluorescence signal at 560 nm (Fig. 3C). The obvious motion artifacts were visible in traces B and C, and were eliminated by expressing their ratios (R560/R620 ratios, Fig. 3D). Resting potential and amplitude of action potential calculated by the calibration equation were similar to those reported in the literatures [14,35–37]. Furthermore, by limiting the intensity of the illumination light, the decay with time was minimal (Fig. 3E).


Figure 3
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  		Fig. 3 Ratiometric recording of the action potential. A–C: During active contraction indicated by sarcomere shortening (A), red (B; 620 nm) and green (C; 560 nm) fluorescence signals (raw voltage signals) were recorded with a two-photo-multiplier system. D: Ratiometry (R560/R620) successfully eliminates the motion-induced artifacts. E, Long recording of the ratiometric signal. Bars, 100 ms for A–D, 10s for E.

 
3.2. Phase effect
When isolated myocytes were paced at 0.5 Hz, the APD80% (the time at which the membrane potential decreased to 80% of its peak value) duration of the action potential in control cells was 61.0±3.7 ms (n=14 cells). A 2 µN stretch of 100 ms in duration applied synchronously with the electrical stimulation significantly increased the APD80% (stretched 0–100 ms after pacing: 83.7±5.8 ms, n=14 cells; P<0.01 compared to control cells by the paired t-test, Fig. 4A). However, stretching did not change the amplitude of action potential (control: {delta}R560/R620 0.139±0.0058, calibrated amplitude of action potential 99.0±4.1 mV, n=14 cells; stretched 0–100 ms after pacing: {delta}R560/R620 0.142±0.0061, calibrated amplitude of action potential 101.1±4.3 mV, n=14 cells; P=0.21). To further dissect the phase effect, we applied a stretch of a shorter duration (50 ms) at 0 or 50 ms after the electrical stimulation. As expected from the trace obtained with the 100 ms stretch (Fig. 4A), application of the stretch in the first half of the action potential elicited a minimal effect (Fig. 4B), whereas a stretch in the later phase (Fig. 4C) significantly depolarized the membrane potential and prolonged the APD. The average APD80% induced by a late systolic stretch (50–100 ms after pacing) was 81.8±6.3 ms (n=14 cells), and thus quite similar to that induced by the 100 ms stretch applied synchronously to the stimulation. Finally, when the stretch was applied during diastole (250–350 ms after pacing) (Fig. 4D), it did not affect the APD80% (57.4±3.1 ms, n=14 cells) or action potential amplitude ({delta}R560/R620 0.135±0.0056, calibrated amplitude of action potential 96.2±3.9 mV, n=14 cells), but transient depolarization was observed. The effect of diastolic stretching is described in the next section.


Figure 4
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  		Fig. 4 Phase effect of stretching on the membrane potential. A–D: Stretching (2 µN) was quickly applied (A) 0–100 ms after pacing (whole systolic period), (B) 0–50 ms after pacing (early systole), (C) 50–100 ms after pacing (late systole) and (D) 250–350 ms after pacing (diastole) in stepwise manner. Red lines denote the membrane potential under stretched conditions, while black line denote that under control conditions. Red bars denote the timing and duration of the stretch. Application of stretching during the whole systolic period or late systolic phase prolongs the APD. Application of stretching during diastole induces transient depolarization, but does not affect the APD. All the data in this figure were obtained from the same single myocyte, although similar tendencies were observed in all 5 examined cells.

 
3.3. Amplitude effect
In Fig. 5, the responses of the membrane potential measured by R560/R620 are shown with the applied stretch quantified by the change in sarcomere length. Application of axial stretching to unstimulated myocytes induced transient membrane depolarization in an amplitude-dependent manner (Fig. 5A,D). When the amplitude of stretch exceeded 15%, action potentials were triggered in some myocytes (Fig. 5A). Estimated from the membrane potential calibrated from the R560/R620 value, the threshold value was approximately –50 mV which is in the reported range for Na channel activation [38]. These responses were almost completely abolished by the addition of 10 µmol/L gadolinium (Fig. 5B). As evident from Fig. 5A, the membrane potential remained elevated as long as the stretch was maintained, and inactivation was not observed during stretches of longer duration (up to 30 s; Fig. 5C). Fig. 5D shows the responses of the membrane potential (R560/R620 values averaged over a 500 ms period during the stretch, and calibrated membrane potential) plotted as a function of the change in sarcomere length (n=7 cells). As larger stretches were applied, the membrane depolarization became larger but we found a small plateau below 5% stretch (Fig. 5A,D). We speculated that this biphasic response may reflect the bimodal action of SACs. Since an earlier study revealed that the stretch-activated ion current at the resting membrane potential is carried by a flux of cations, mainly Na+ [14], we repeated similar experiments under either Ca2+-free (n=7 cells) or Ca2+-free and low Na+ (n=7 cells) conditions with an intention to examine the ion selectivity by withdrawing permeant ions. When Ca2+ was removed from the solution, the response of the membrane potential was attenuated over the entire range of the stretch amplitude, but the attenuation was exaggerated in the small amplitude range (<5%). Further removal of Na+ from the solution (Ca2+-free and low Na+ condition) made the response almost completely disappear over the entire range of stretching. Cytochalasin D treatment almost completely suppressed the stretch-induced membrane potential change (n=7 cells).


Figure 5
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  		Fig. 5 Effect of stretching amplitude on the resting membrane potential. A: Stretches of varying amplitudes (5%, 10%, 15%, and 20% in sarcomere length; upper traces) and the corresponding membrane potential responses (R560/R620; lower traces) are shown. The stretch was applied in stepwise manner. The change of R560/R620 values at 5%, 10% and 15% stretch are 0.011±0.0016, 0.017±0.0037 and 0.028±0.0016 (n=7 cells), respectively. Bar 500 ms When the cell was stretched over 15% of sarcomere length, action potentials were triggered in some cardiomyocytes. Because, in this case, the applied command was designed to induce 20% stretch in the relaxed myocyte, the system could not hold the myocyte length during its contraction as indicated in the length recording. Bars, 500 ms. B: Effect of gadolinium treatment (10 µmol/L) on the membrane potential responses induced by stretching in diastole. The responses are almost completely abolished (The change of R560/R620 at 10% stretch in the presence of gadolinium=0.003±0.0027, n=7 cells, P<0.01 compared to the control). Bar, 500 ms. C: Response of membrane potential to the stretch of long duration. Upper panel: relative change in sarcomere length Lower panel: membrane potential (R560/R620) bar 10s D: Relationships between the stretch amplitude (in sarcomere length) and the membrane potential response (The change of R560/R620 and calibrated membrane potential) plotted as red circles for the control condition (Tyrode solution Ca2+: 1.1 mmol/L), open black circles for the Ca2+-free condition (0.4 mmol/L EGTA without added Ca2+), green circles for the Ca2+-free and low Na+ condition (Na+ was replaced by the same concentration of choline chloride) and blue circles for cytochalasin D treatment (20 µmol/L for 1 h) under the control condition. Cytochalasin D treatment almost completely abolishes the response. Error bars represent the SEM (n=7 cells).

 
3.4. Speed effect
Representative recordings of the membrane potential responses to various speeds of stretches with the same amplitude are shown in Fig. 6A. The membrane potential responses observed after completion of 10% stretching in sarcomere length (steady-state values) did not differ among the three conditions. However, when the relationship between the sarcomere stretch and membrane potential was plotted, higher speed stretches clearly caused higher degrees of membrane depolarization during the course of the stretch (Fig. 6B), thus indicating speed-dependence. In the 5 cells studied for each group, we observed similar speed-dependences of the membrane potential response.


Figure 6
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  		Fig. 6 Effect of stretching speed on the membrane potential response. A: Representative recordings showing the time course of the membrane potential in response to stretches of various speeds with the same amplitude (10% in sarcomere length). The membrane potential responses (R560/R620) observed after completion of the stretch (steady-state value) do not differ among the three conditions (strain rates: black line: 1.50/s; red line: 0.75/s; and blue line: 0.25/s). Bar, 100 ms. B: Relationship between the sarcomere stretch and the membrane potential. The membrane potential responses (R560/R620) after completion of a 10% stretch in sarcomere length (steady-state values) do not differ among the three conditions (The change of R560/R620 at 10% stretch: 1.50/s: 0.019±0.0014, n=5 cells; 0.75/s: 0.018±0.0020, n=5 cells; 0.25/s: 0.019±0.0017, n=5 cells; P=0.71 group comparison by ANOVA). However, it is clear that higher stretch speeds cause higher degrees of membrane depolarization during the course of the stretching, thus indicating speed-dependence (The change of R560/R620 values at 4% stretch: 1.50/s: 0.019±0.0014, n=5 cells; 0.75/s: 0.015±0.0018, n=5 cells; 0.25/s: 0.011±0.0016, n=5 cells; P<0.01 group comparison by ANOVA). Bars, 50 ms. Error bars represent the SEM.

 

   4. Discussion
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
References
 
By combining two techniques, i.e., cell manipulation by carbon fibers and membrane potential measurement based on ratiometric di8-ANEPPS recording, we have succeeded in recording the membrane potentials of single cardiomyocytes during the application of axial stretches of various times, amplitudes and speeds. Several previous reports have measured the membrane potential and/or current in response to axial stretching in isolated single myocytes [14,15,39]. However, the applied stretches were either local [39] or limited in terms of their amplitudes [14,15] compared with those in the present study. Furthermore, the use of glass electrodes only allowed static stretches. The present study can provide not only information regarding the cellular response to dynamic stretching, which all myocytes may face in the intact ventricle, but also insights into the molecular mechanisms of SACs.

4.1. Phase effect
We found that systolic stretching depolarized the membrane potential and prolonged the APD, and that the effect was predominant during the second half of systole. Application of shorter stretches truncated in either the first or second half of systole replicated these observations. These results are consistent with those reported by Zeng et al. [14], except for the fact that the resting potential did not return to the control level in their study due to the application of a continuous stretch. On the other hand, Zabel et al. [40] applied transient stretches to the rabbit ventricle to elucidate the repolarization effect of early systolic stretching. Considering the similar linear current–voltage relationships of SACs with a reversal potential close to zero reported in these two previous studies [14,40], stretching is expected to repolarize the membrane potential during early systole when the membrane potential is positive, although the reason for this discrepancy is not clear. In the present study, we consider that the brief action potential without a plateau observed in our rat myocytes may have masked the small repolarization effect during early systole.

An alternative interpretation for the stretch-induced APD prolongation is possible. The present results as well as previous studies [14,40] indicate that the stretching causes depolarization during the late phase of systole or diastole when the membrane potential is negative enough. Therefore, the action potential prolongation could result from the overlap of the stretch effect with the action potential.

4.2. Amplitude effect
Similar to previous studies [14,40], the change in the membrane potential was dependent on the stretch amplitude. These observations are in good agreement with a prediction from a theoretical consideration of mechanical transduction [41] and a proposed model for SACs [42]. We further found that the membrane potential responded to stretches of very small amplitude and seemed to become saturated once in the lower range and then respond differently to larger stretches. Importantly, we obtained similar findings to those in Fig. 8B in the paper by Zeng et al. [14], in which a cluster of data points in the region of very small stretches deviated systematically from the regression line applied to the entire range of data. These findings suggest that SACs may respond differently depending on the degree of stretching. The data obtained under the Ca2+-free condition or Ca2+-free and low Na+ condition suggest that this bimodal response may reflect stretch-dependent ion selectivity.

4.3. Ion selectivity
Previous studies to date have identified Na+ as the major charge carrier of SACs at the resting potential level [39], although Ca2+ is probably a permeant ion as well [14,43]. Our observations are consistent with these findings, since Na+ dominated the response at higher degrees of stretching and we also identified a small, but significant, response that was probably carried by Ca2+ when the applied stretch was small. Clear stretch-dependent selectivity has been shown for the first time in the present study. The ion selectivity of the channel is governed by the valency, size and hydration state of the ion as well as the surface charge of the pore [44,45]. Although the structure of SACs has not yet been identified, if their opening is indeed controlled by a conformational change introduced by membrane stretching, it is unlikely that monovalent and divalent cations show similar permeation, although the Pauling radii of Na+ and Ca2+ are close. Instead, it is tempting to speculate that, at small stretches, the SAC takes a favorable conformation for permeation of Ca2+ but changes to the Na+-selective conformation at larger stretches, and that further stretching recruits more SACs to open. The other possibility is the presence of two distinct populations of channels with different ionic selectivity properties. Further study is needed to determine which is the case in mammalian ventricular myocytes.

We can find contradicting reports regarding the effect of extracellular Ca2+ concentration on the stretch-activated electrical activity. Kamkin et al. applied large (18%) local stretch in mouse ventricular myocytes [39] to find that the removal of extracellular Ca2+ increased the stretch-activated current by a factor of 2.5. This observation can be accounted for by the charge screening effect of divalent cation if the SAC has intramembrane voltage sensor [46]. The reason for the discrepancy is not clear but we can think of a few possibilities. First, the two different methods, i.e. local vs. end-to-end, may deliver dissimilar stress to the mechanosensing elements of the cell. Secondly, in rat ventricular myocytes, Zeng et al. [14] reported that Ca2+ removal did not affect the ion current elicited by the small (<5%) axial stretch. Although this result again contrasts to the present result showing the reduction of stretch-induced activity upon Ca2+ removal in the similar range of stretch, species difference can be another factor. The species difference in stretch-activated current in response to the change in extracellular Ca2+ concentration has also been reported for guinea pig ventricular myocytes [16].

The present study cannot answer whether SACs discriminate among different divalent cations. To obtain definitive answer to this question, further studies with direct measurement of current are necessary.

4.4. Speed effect
In the present study, we demonstrated speed-dependence for the membrane potential responses to stretching, suggesting the viscoelastic nature of this transduction system. The abolishment of the response by cytochalasin D treatment observed in the present study as well as others [39] strongly suggests that the viscoelasticity resides in the cytoskeleton. As previously suggested [41], stretches of varying speeds and amplitudes applied to the cell membrane are converted and transmitted through the cytoskeleton, and apply force to SACs probably via ankyrin molecules, resulting in a change in the strain energy of the conformation. In the case of constant speed stretch protocol adopted in this study, speed-dependent viscous force was added to the elastic component during the course of stretch, therefore, if compared at the same stretch amplitude, faster stretch could cause a larger response. To substantiate this hypothesis, the effect of stretches applied in the transverse direction should be examined in future studies. The implication of these findings are two-fold. First, in the marginal zone of an infarcted area, the myocardial tissue is expected to undergo rapid stretching, and this may induce the premature contraction that sometimes leads to fatal arrhythmia in myocardial infarction. Second, although most of the currently used SAC models do not incorporate the speed-dependence [14,40,42], it is desirable to include it for clinically useful simulations.

4.5. Study limitations
There are two major limitations in the present study. First, we used rat cardiomyocytes, which are known to have a brief time course for action potentials compared to guinea pigs and humans [14,35–37]. Therefore, studies on such larger animals may have allowed more detailed analyses in terms of temporal resolution. Second, the use of optical recording enabled us to study the membrane potential responses to dynamic stretch, but the absolute value of potential could not be recorded. However, previous studies revealed fairly linear relationships between optical recordings and microelectrode measurements [25,27,47], and we succeeded in calibrating the fluorescence signals against absolute membrane potential by direct electrode measurements. Although direct comparisons are difficult due to the differences in the optical systems used, the obtained conversion factor of 11%/100 mV was comparable to the previously reported value: 15%/100 mV by Zhang et al. [47], 9.7%/100 mV [25] by Beach et al. and 8%/100 mV by Knisley et al.[27]. The calibrated amplitude of the action potentials (approximately 100 mV) was also similar to the previous results obtained by electrode recordings [14,35].

In summary, we have studied the effect of dynamic axial stretching on the membrane potential in isolated rat cardiomyocytes by combining two techniques, namely a carbon fiber technique and ratiometric fluorescent indicator measurements. The results not only revealed the dynamic characteristics of this mechanotransduction system but also suggested its subcellular mechanisms, such as stretch-dependent ion selectivity and participation of the actin cytoskeleton, thus providing information on both the clinical electrophysiology and basic science of SACs.


   Acknowledgements
 
We thank A. Matsuoka for the excellent technical assistance. This study was supported by Research Fellowships from the Japan Society for the Promotion of Science for Young Scientists (S.N.), The Vehicle Racing Commemorative Foundation (S.S) and The Nakatani Electronic Measuring Technology Association of Japan (S.S.). No conflict of interest exists.


   Notes
 
Time for primary review 17 days


   References
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 References
 

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H. Huang, H. Wei, P. Liu, W. Wang, F. Sachs, and W. Niu
A simple automated stimulator of mechanically induced arrhythmias in the isolated rat heart
Exp Physiol, October 1, 2009; 94(10): 1054 - 1061.
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