Mechanoelectric transduction can initiate cardiac arrhythmias. To examine the origins of this effect at the cellular level, we made whole cell voltage-clamp recordings from acutely isolated rat ventricular myocytes under controlled strain. Longitudinal stretch elicited noninactivating inward cationic currents that increased the action potential duration. These stretch-activated currents could be blocked by 100 μM Gd3+ but not by octanol. The current-voltage relationship was nearly linear, with a reversal potential of approximately −6 mV in normal Tyrode solution. Current density varied with sarcomere length (SL) according to I (pA/pF) = 8.3 − 5.0SL (μm). Repeated attempts to record single channel currents from stretch-activated ion channels failed, in accord with the absence of such data from the literature. The inability to record single channel currents may be a result of channels being located on internal membranes such as the T tubules or, possibly, inactivation of the channels by the mechanics of patch formation.
- ion channel
- patch clamp
- mechanical stress
mechanical stress changes the electrophysiological properties of the heart, a phenomenon known as mechanoelectric feedback (25). Stretching intact hearts or excised muscle can raise the beat rate (2, 3), cause diastolic depolarization (30), change the action potential configuration (7, 24), and induce arrhythmias (17). Stretch-activated channels (SACs), considered to be the origin of mechanoelectric transduction (21, 37, 53), are found in many cardiomyocytes, including those from molluscan ventricle (40), chick embryo ventricle (20, 33), rat atrium (22, 50) and ventricle (8), rabbit sinoatrium and atrium (12), guinea pig ventricle (6, 39), and pig atrium (19). Most of the above recordings were made with the cell-attached single-channel patch-clamp technique, in which the open probability (P o) of the channels increased with negative pressure applied to the patch pipette. None of the above data were obtained from adult ventricular cells, and there are almost no data on whole cell responses of cardiocytes to stretch under voltage clamp.
To record whole cell mechanosensitive currents under stretch, the cells must not only be voltage clamped but also stretched without damage. Stretching intact, isolated cells without damage has proven to be extremely difficult (5, 11), and only one paper has shown whole cell currents (39). In some experiments hydrostatic or osmotic pressure was used as a mechanical stimulus to inflate the cells, but it is unlikely that these stimuli are equivalent to axial stretch (20). Hagiwara et al. (12) evoked mechanosensitive Cl− currents by inflating the cells through the clamping pipette, whereas Sorota (44) and Tseng (47) found an increase of Cl− permeability in response to hypotonic swelling. Attempting to evoke axial strain, Sasaki et al. (39) attached cells to a coverslip or to a fire-polished glass tool and pulled on the other end with another glass tool or suction pipette. Currents were recorded with a separate pipette and had a reversal potential of −15 mV in physiological saline. Hu and Sachs (20) recorded currents in chick heart cells through a perforated patch and stimulated them by compressing the rounded cells with a second pipette. In agreement with the results of Sasaki et al. (39), Hu and Sachs found a mechanosensitive cation current reversing at −16 mV. In none of these papers was strain under reliable control.
In this paper, we present evidence of a gadolinium-sensitive whole cell stretch-activated current (I stretch) in adult rat ventricular cells under controlled strain. The cells were pulled with a pair of concentric pipettes as described by Palmer et al. (29). The axial strain was measured from calibrated displacements of the ends of the cell and from Fourier transforms of the sarcomere spacing. These measurements formed the basis for quantifying the relationship between strain and current.
Ventricular myocytes were enzymatically isolated by retrograde perfusion of the heart (28, 49). Briefly, Sprague-Dawley (2–3 mo old) rats were injected with heparin (2,000 U/kg) and then Nembutal (60 mg/kg). When the rat was anesthetized, the heart was quickly excised, cannulated through the aorta in cold Tyrode buffer, and then mounted on a dual-channel tube Langendorff perfusion apparatus. Perfusion of the heart proceeded at 37°C for 10 min in Tyrode solution, 4.5 min in Ca2+-free Tyrode solution, 30 min in the enzyme solution, and 5 min in low-Ca2+ Tyrode solution. All perfusion solutions were equilibrated with 100% oxygen. The enzyme solution was limited to 60 ml and allowed to recirculate. After perfusion, the ventricle was cut off and minced. Cells were dispersed from the tissue by agitation, filtered into Tyrode solution through a 200-μm-mesh net, and stored at 4°C until use. All experiments were done within 1–20 h after isolation.
Isolated cells were loaded for 30 min at room temperature in normal Tyrode solution containing 2 μM fluo 3-AM (Molecular Probes) and 0.2% Pluronic-127 (Molecular Probes) (41). Cells were then rinsed three times in saline and left for 20 min to further hydrolyze the ester form of the dye.
Cell stretch and patch clamp.
Isolated rod-shaped ventricular cells with clear sarcomeres were held by two concentric glass pipettes (29), with the inner pipette serving as a stop to prevent the cell from being sucked up the outer pipette. The outer pipette was pulled from a glass capillary (ID 1.0 mm, OD 1.5 mm; Drummond Scientific) with an inner tip diameter of ∼15 μm. The inner pipette was made from a glass capillary (ID 0.5 mm, OD 1.0 mm; Dagan) with an outer tip diameter of ∼12 μm. The inner pipette was inserted into the outer pipette by a manipulator, leaving a gap of ∼8 μm to the tip of the outer pipette. The tip of the outside pipette was then cut by fusion of the tip to the filament of a microforge as described by Hilgemann (18). The cut end was lightly fire-polished so that the tips of the inside and outside pipettes were forged together and formed a cup to hold the cell.
To make the cells adhere to the glass, we tried a host of different agents. These included Cel-TaK (Collaborative Biomedical Products), a glue based on barnacle adhesive proteins, as suggested by Palmer et al. (29). This did not provide sufficient adhesion for prolonged pulling. Strong suction itself caused fatal cell contraction, probably via SACs. We tried many different adhesives, including covalent and noncovalent bonding agents such as poly-l-lysine. We tried adhesives activated by ultraviolet light (Master Bond), epoxies, cyanoacrylates (“crazy glue”), silicones (e.g., Kwik-sil, World Precision Instruments), charged silanes such as 3-aminopropyltriethoxysilane, and covalent silanes such as isocyanates. We tried covalently attaching wheat germ agglutinin to the glass with a silane linkage, but that, too, was unreliable. The problems varied from adhesives not sticking well to the glass or to the cells or damaging the cells to the adhesives not catalyzing under water or catalyzing too fast. We had hoped to find a volume-filling adhesive to take up the space between the pipettes and the cell, but as yet we have not been successful. Our best adhesive to date has been a dense layer of positive charge linked covalently to the glass. We first treated the concentric pipettes with a silane (I7840, United Chemical Technologies) that left the glass coated with isocyanate groups. The silane was prepared as a final concentration of 2% in 95% ethanol. Pipettes were immersed in the solution for 5 min and then cured 24 h at room temperature. Shortly before each experiment, the coating was reacted with an amino dendrimer (PAMAM dendrimer generation 4, Aldrich Chemical) by dipping the tip of pipettes in a 10% solution in methyl alcohol for 5 min.
To attach cells to the pipettes, two or three drops of the cell suspension were transferred to a custom-designed chamber with a bottom made from a coverslip. After 5 min most cells settled down, and the bath solution was then changed to a low-Ca2+ or Ca2+-free relaxing solution so that suction would not cause extensive contraction. The selected cell was first drawn into one concentric pipette by gentle suction, with the intercalated disk region firmly contacting the inner pipette. The cell was then lifted ∼50 μm above the bottom of the dish. The second concentric pipette was then moved close to the free end of the cell, and it was gently drawn in as for the first pipette. The pipette positions were adjusted so that the cell was relaxed and aligned axially with the two pipettes. We waited 10 min to allow the dendrimer to bond to the membrane. The cell was then patched with a third pipette. Despite our efforts to get the best adhesion possible, and even with small strains, it was usually not possible to stretch a cell more than four or five times before it pulled loose from one of the pipettes. Consequently, many of the results are presented as statistical averages across cells.
The patch pipette and one of the concentric pipettes were mounted on PCS-800 piezoelectric manipulators (Burleigh Instruments), and the other concentric pipette was attached to a second manipulator (MP-300, Sutter Instruments). During the experiment, the cell was stretched axially by sending an electrical command to the PCS-800 manipulator at one end of the cell. To reduce local strain in the region of the patch pipette, this command was scaled and sent to the PCS-800 manipulator controlling the patch pipette, so that it moved sideways in proportion to the local strain.
Data recording and analysis.
Standard whole cell recording methods (14) were used to clamp the membrane voltage and record currents. The fire-polished patch pipette had a tip ID of 1–2 μm, with a resistance of 0.5–2.0 MΩ when the pipette was filled with high-K+ pipette solution. The seal resistance was usually >2.0 GΩ. Cell capacitance was derived by fitting the transient currents generated by a voltage pulse after the whole cell configuration was formed (26).
Voltage and current signals from the patch-clamp amplifier (Axopatch-1B, Axon Instruments) and movement commands of the stretching pipette were filtered at 2.5 kHz and then digitized by a data acquisition board (AT-MIO-16E-2, National Instruments) in a personal computer (XPS H266, Dell). This data acquisition board was the control unit for all of the experiments, generating voltage, stretching, and synchronization command potentials. The board was controlled by custom software (T. Zeng, unpublished) that was programmed in LabVIEW (National Instruments). A charge-coupled device camera (KP-M2U, Hitachi) was attached to the microscope (DIAPHOT 200 with ×40 oil-immersion objective and water-immersion condenser, Nikon) to monitor the cells. During cell stretching, a digital signal from the data acquisition board triggered a frame-grabber board (IMAQ PCI-1408, National Instruments) to sample the images of cells at specific times (Fig. 1 A). Continuous real-time video images were also logged onto S-VHS tape for archival storage. To change the bath solutions around the cell, a pulse was sent from the board to control the valves of a perfusion system (PS-8, ALA Scientific Instruments), whose output was held in a fourth manipulator.
Once the patch was broken by the zap pulse from the amplifier, we waited 5–10 min to reach a steady state. A small stretch was then given to test whether the cell was still firmly held by the concentric pipettes. Data were recorded by applying larger stretches until the cell came loose or contracted. The stretch-activated current was measured as the differential before and after stretching. We have attempted ∼100 cells, but only 15–20 were held sufficiently firmly and remained relaxed while being patched.
Sarcomere length (SL) was determined from the cell images using a two-dimensional (2-D) Fourier transform operating off-line (Fig.1 B). A region of interest (256 × 128 pixels with 256 gray levels) was transformed into the 2-D power spectrum and the spatial frequency peak (f cell) corresponding to the SL was located by software. SL was computed by the equation SL =l cal ×f cal/f cell, wherel cal is a calibration distance (5 μm) obtained from the image of a stage micrometer and f cal is its corresponding maximum spatial frequency. The analysis software was also written in LabVIEW. The mean SL of our cells in resting condition was 1.77 ± 0.08 μm (n = 18) when held by the pipettes. Statistical significance was calculated using Student'st-test.
The Tyrode solution contained (mM) 137 NaCl, 5.4 KCl, 0.5 MgCl2, 1.8 CaCl2, 10 HEPES, and 5.0 glucose, pH 7.4 with NaOH. The Ca2+-free solution was Tyrode solution without added Ca2+. For the low-Ca2+ Tyrode solution we reduced Ca2+ in Tyrode solution from 1.8 to 0.2 mM. Tris-Tyrode solution was Tyrode solution with Na+replaced by Tris+. The enzyme solution was 60 ml of Ca2+-free Tyrode plus 30 mg of collagenase A (Boehringer Mannheim) and 6 mg Protease XIV (Sigma). The pipette solution was (mM) 130 KCl, 10 NaCl, 5 MgCl2, 11 EGTA, 1 CaCl2, and 10 HEPES, pH 7.4 with KOH. In some experiments, Cl− was replaced by F− as indicated.
I stretch was modeled using a simple linear nonspecific current with two components, the Na+-carrying element and the K+-carrying element whereE Na and E K are the reversal potentials for sodium and potassium respectively,E m is the membrane potential, and γstretch,Na and γstretch,K are the whole cell conductances of the stretch current to potassium and sodium ions, respectively. The components were kept separate to ease calculation of ion concentration changes within the cell as well as the electrogenic effect of the current.
I stretch was incorporated into a model based on the Oxsoft Heart Model (Biologic) of the isolated rat ventricular cell, which was modified to include a parameter for EGTA. The on and off rates for Ca2+ binding to EGTA were 106.3M−1s−1 and 0.4 s−1, respectively (48). Oxsoft Heart is a mathematical representation of the heart based on experimental electrophysiological data from a variety of sources. It simulates the behavior of transmembrane currents, exchangers, and transporters, the sarcoplasmic reticulum, the intracellular buffers, and the movement of intracellular and extracellular ion concentrations.
Stretch increases action potential duration and depolarizes resting potential.
In normal-Ca2+ solution, heart cells are apt to contract when touched by a glass pipette, particularly when subjected to suction. We presume that this represents action of SACs. To check the effect of stretch on cell excitability, we recorded the membrane potential while stretching the cell. In current-clamp mode, pulses of 1 ms and 2 nA repeated every 5 s could elicit action potentials. After the action potential was stable, we pulled on one of the concentric pipettes to stretch the cell. The displacement was ∼5 μm and lasted 20 s. The pipette was then returned to its original position. Figure2 shows action potentials recorded during stretching.
Compared with the control value of 621 ± 12 ms (n = 5), action potential duration (APD) at 90% repolarization (APD90) increased to 764 ± 50 ms (n = 4,P < 0.01) with stretching. On return to the resting length, there was no significant difference in APD90 (653 ± 35 ms, n = 5, at P = 0.1 level). Stretching had a similar effect on APD50, causing it to increase from 516 ± 19 ms (n = 5) to 597 ± 22 ms (n = 4, P< 0.01). Stretch also caused depolarization of the resting potential. During our small stretches, the cell depolarized to −59.7 ± 1.2 mV (n = 4, P < 0.01) from the control level of −62.4 ± 1.1 mV (n = 9). The effects of stretch on the action potential and the resting potential are consistent with the activation of inward currents.
Longitudinal stretch-activated inward currents.
When the membrane potential was held at −100 mV in Tyrode solution, stretching the cell elicited an inward current (Fig.3). This current activated without visible delay after initiation of stretch and was maintained during stretch. After release, the current returned to its prestretch level. While the cell remained attached, repeated stretching gave nearly the same response. For example, in one of our more extensive experiments (n = 4), the mean current was 1.03 ± 0.18 pA/pF at −100 mV. Currents elicited in Ca2+-free or low-Ca2+(0.2 mM) solutions were similar to those obtained in normal Tyrode solution. Because the cells were much more likely to contract after being attached to the pulling pipettes in normal-Ca2+solution than in low-Ca2+ solutions, we usually conducted experiments in Ca2+-free or low-Ca2+ Tyrode solution.
The stretch-induced current was not an artifact of membrane breakage, because cells loaded with fluo 3 in normal-Ca2+ medium did not show significant leakage of Ca2+ while being stretched (n = 3, Fig. 4). The strain-induced currents did not seem to originate in the vicinity of the patch pipette because even small, deliberate movements of the pipette evoked no measurable current.
Because Gd3+ is a blocker of many SACs (15, 57), we examined its effect on the whole cell currents. In one cell, 40 μM Gd3+ blocked inward currents by 60%. In another cell, 100 μM Gd3+ reduced the current by 90% (Fig.5). The block by Gd3+ was partly reversible after 5 min of washout.
Octanol, a gap junction blocker (45), did not have any significant effect on the mechanosensitive current (Fig.6). The mean current was 93 ± 29 pA in control and 79 ± 10 pA in 100 μM octanol. The difference was not significant (P = 0.05, n = 4).
Istretch has a reversal potential of −6 mV.
We examined the voltage dependence of the currents in Tyrode solution by applying test pulses between −120 and +20 mV from a holding potential of −100 mV. Because stretching the same cell more than four times without losing the attachment was very difficult, we collected the data from four different cells and normalized the currents by the cell capacitance. The cells were stretched 5 μm, equivalent to ∼3% strain. All currents were calculated as the mean current between the rising and falling phases. The mean current-voltage (I-V) curve (Fig. 7) showed an essentially linear relationship with a reversal potential of −6 mV.
Istretch increase with SL.
By programming the voltage sent to the piezomanipulators, we could apply different global strains to the cell. Video images of cells before and after stretching were grabbed into the computer, and mean SL was computed from the power spectrum of the images. Figure8 shows that in the narrow range of strain available, the change in mechanosensitive current was approximately linear with both SL and strain. Both SL and SL strain were fit to the current by linear regression. At −100 mV, the former follows the relationship I (pA/pF) = 8.3 − 5.0SL (μm), and the latter follows I (pA/pF) = −0.30SL%. The first equation predicts that SAC currents will persist to 1.66 μm. The 95% confidence limits, however, include zero current at longer SLs, and we do not trust the precision of this prediction. There are probably small-scale nonuniformities in SL that do not show up in our first-order analysis of the diffraction pattern. The sample size and the optical resolution in three dimensions limit the effective resolution of the SL. The net result of these systematic and random errors is scatter in the data. In Fig. 8 B, the large fraction of data points outside the 95% confidence intervals suggests some significant nonrandom errors such as a nonuniform SL distribution.
Na+ is the main carrier of mechanically sensitive currents at resting potential.
Because I stretch was inward at −100 mV, that current must be carried by an influx of cations, an efflux of anions, or a mixture of the two. Replacing Cl− in the pipette solution with F− produced similar currents (n= 5, see Fig. 9 for example), suggesting that the current was carried by cations rather than anions (13).
To examine the components of the cation influx, we superfused the cells with Tris-Tyrode solution in which Na+ was replaced by Tris+. The heart cells were not stable for long in this nonphysiological Tris-Tyrode solution, so perfusion was started only 3 s before stretching and was shut down 1 s after the cell was restored to its original position. Figure 10indicates that inward currents were largely reduced in the presence of Tris-Tyrode solution (n = 2), suggesting that Na+is the main carrier of the currents at resting potential.
Stretch-activated ion channels are thought to be responsible for the mechanically sensitive current, so we looked for single channel currents in cell-attached patches. Much to our dismay, we recorded no single channel activity. Dozens of patches were tried using three different setups and the assistance of two additional independent and experienced investigators. In no case were we able to observe stretch-activated single channel currents. Despite its negative character, this result correlates with the absence of publications on SACs in adult mammalian ventricular myocytes.
The experimentally obtained I-V curve forI stretch was well fit by a straight line representing the equation where γstretch,K = 0.9 nS and γstretch,Na = 1.17 nS, given the model rat's nominal whole capacitance of 200 pF (see Fig. 7). These values were used with the equation above to test the effect of I stretch on the rat action potential and compare it with the experimentally obtained results.
Because of the experimental complexities of keeping the cell attached to the pipettes and in whole cell voltage clamp while applying strain to the cell, the experimental stimulation frequency used was only 0.2 Hz. Decreasing the stimulation frequency of isolated rat ventricular cells increases the APD and increases calcium loading of the SR (27). However, this has little effect on the APD in the presence of 10 mM EGTA. The result of stimulating an action potential at 0.2 Hz in the absence and presence of I stretch (based on SL of 1.89 μm), with an extracellular calcium concentration of 0.2 mM and 10 mM intracellular EGTA, is shown in Fig.11.
The duration of the modeled cell action potential is shorter than that observed experimentally, but this is easily explained by the difference in temperatures: the nominal temperature of the model cell is 37°C, whereas the experiments were performed at room temperature. Kiyosue et al. (23) demonstrated that a decrease in temperature of 9° resulted in an increase in guinea pig APD from 146 to 314 ms. The difference between the model and experimental temperatures in our data is 17°C.
Activation of I stretch depolarized the resting membrane of the cell and lengthened the action potential throughout its duration (i.e., at both APD50 and APD90). A small stretch-activated current with the I-V relationship shown in Fig. 7 could be responsible for the experimentally observed effects on the action potential.
This is the first study of stretch-activated whole cell currents in mammalian cardiocytes under controlled strain. We were able to show that in rat ventricular cells stretch activated a gadolinium-sensitive, noninactivating, inward current. Our data are similar to those published for other heart cells (20, 39) and smooth muscle cells (9,52).
SACs in heart cells have been reported to be nonselective cation channels (8, 33) or Cl− channels (12). In our experiments, I stretch was not significantly different when the pipette solution contained Cl− or F−, suggesting that these currents are carried by cations. Further experiments are needed to define the selectivity more precisely. Removing external Ca2+ did not significantly affect the current, suggesting that Ca2+ does not act as a significant charge carrier or a second messenger, although Ca2+ is probably a permeant ion. Replacing Na+with Tris+ reduced the current, although the blockage was not complete. I stretch in rat heart cells appears to be cation selective, with Na+ acting as the major permeant ion at resting potential. Wellner and Isenberg (52) reported that in guinea pig smooth muscle cells, stretch increased a voltage-activated potassium current and reduced a calcium current. In heart cells, however, there is no evidence to suggest that stretching has any effect on voltage-sensitive channels (20, 39). Because of the experimental difficulty with our preparation, we did not directly check the effect of stretching on voltage-sensitive currents. However, we did observe that changing the membrane potential to −40 mV and removing Ca2+ from the bath solution produced little difference in the time course of the mechanically activated currents. Thus it is unlikely that voltage-dependent K+, Na+, or Ca2+ channels are responsible for the mechanosensitive currents we observed.
Heart cells shorten >10% during contraction, but they are not readily stretched. In our system, 2–5% strain was possible. In some experiments, we moved the stretching pipette 10% of the cell length, but the attached cells would either contract immediately or slip out of the holding micropipette. In the latter case, the cells usually contracted into a ball in a few seconds. With a resting SL of 1.77 μm, our results showed that <5% strain was sufficient to evoke stretch-sensitive currents. Comparing the passive elastic properties from detergent-skinned isolated cells and intact cardiac tissue, Brady (4) suggested that intracellular structures may contribute measurably to total cardiac passive elasticity at SL < 2.2 μm, whereas extracellular elements form the major limitation at more extended lengths. Furthermore, he suggested that these intracellular structures were probably related to the cytoskeleton rather than membrane elements (5). Our data are consistent with a model in which stress in the membrane is coupled through the intracellular cytoskeleton to the channels.
How much current is contributed by SACs at “resting” SLs is difficult to determine and could only be measured with a specific inhibitor, which Gd3+ is not. Our definition of a mechanically sensitive current is one that changed with stretch of the cell, so that it is only a differential measurement. In Fig. 8 we plotted current versus SL during stretch and versus SL as strain. Taken literally, Fig. 8 A suggests that at SL = 1.75 μm there would be ∼1 pA/pF of inward current from SACs. This plot is of necessity made from differential data arising from different cells, and without allowance for a variation in SL throughout the cell (only the 1st-order diffraction line was used to define SL) we cannot place much confidence in the amplitude of the resting current. Furthermore, the resting current in isolated cells is unlikely to be the same as in cells in situ, where the normal extracellular contacts are in place.
We made multiple attempts to record stretch-activated ion channels from tight seal patches on the rat ventricular cells, but we never recorded single channel activity. Reviewing the literature, we found that all the reports of single channel stretch-activated activity in heart cells were obtained from neonatal (8, 20, 22), atrial (19, 50), or cultured (33, 38) cells. There are no data on freshly isolated ventricular cells. There appear to be three possible explanations for this absence of single channel data.
First, the channel density may be low, so the chance of catching one channel in a pipette is small. Is this reasonable? Our maximal currents were ∼1 pA/pF at −100 mV. This current corresponds to a product of the unitary current and the probability (P o) of being open. If the single channel conductance had a typical value of ∼25pS, the single channel current would be ∼2.5 pA/channel at −100 mV. There is 1 pF of capacitance for every 100 μm2 of membrane, so we would expect that the minimal density (if P o =1) is 1 channel/40 μm2. Because cell-attached patches have areas of 10–30 μm2 (34, 43), we might expect to see a channel in every other patch. P o is probably <<1 in the whole cell experiments because the strains were small and there was no hint of saturation in the plot of current versus SL. Consequently, there should have been a higher density of channels, and we should have seen them in most patches. SACs typically occur with a density of ∼1–3/patch. It does not seem likely, therefore, that the absence of single channel activity came from a low channel density.
A second possibility is that SACs are not in the surface plasmalemma but are located in the T system and hence invisible to plasmalemmal patching but not to whole cell stretching. An intracellular location for the channels, although an experimental nightmare, has conceptual appeal because strain may be better sensed in the contractile cell interior than in the convoluted plasma membrane. This interior membrane explanation would fit with the sensitivity of the cells to suction applied to the pulling pipettes and with fluorescent imaging showing Gd3+-sensitive Ca2+ waves induced at the site of mechanical deformation (W. J. Sigurdson and F. Sachs, unpublished observation).
The third possibility is that formation of the patch disrupts channel activity in these cells. Patch formation is a major perturbation of the mechanical properties of the membrane and cortical cytoskeleton (1, 16,36, 42, 43, 51). It is possible that some important structure in the adult ventricular cortex is disrupted under stress.
Physiologically, the effect of stretch is most likely expressed through changes in the action potential, although it also will affect the filling of Ca2+ stores. In many recordings of action potentials from intact tissue under stretch, the duration of the plateau is reduced and the tail of repolarization (phase 3) increases, leading to a crossover, or apparent reversal potential of the mechanosensitive current, at about −20 mV (35, 58). In intact rabbit heart, a long static stretch created by inflating a balloon in the left ventricle extended the APD. The peak amplitude of the stretch-activated depolarization from rest and repolarization from the plateau exhibited a linear relationship to voltage and volume change (58). In single guinea pig cardiac myocytes, a 3% strain did not affect the resting potential but did decrease the APD (10, 54).
The rat ventricular action potential is significantly different from that of the rabbit and guinea pig, because it lacks the prolonged plateau and, during the contraction cycle, has a much greater reliance on calcium released from the SR rather than plasmalemmal calcium entry (46). As a test of whether the observed currents could account for the effects of stretch on the action potential, we simulated the rat action potential using the HEART program from Oxsoft. We introduced a stretch-activated current, I stretch, that was permeable to potassium and sodium ions and closely fit the experimentalI-V curve.
The addition of this simple current could account for the depolarization of the resting potential and the observed changes in APD. At the negative potentials of the rat plateau, the reversal potential of the stretch current near 0 mV caused a net inward current and therefore prolonged and depolarized the action potential throughout its course. In cells with a high plateau and a lower reversal potential of I stretch (approximately −20 mV), such as those of guinea pig and rabbit, there is a shortening of the plateau but prolongation of APD90 and a crossover potential during phase 3. White et al. (54) observed a stretch-induced reduction in APD in guinea pig, possibly reflecting the differences in the two preparations.
Although stretching single cells would appear to be the most reductionist level for studying the effects of stretch on whole cells, there is a fundamental problem of interpretation that is not readily resolved: What are we pulling on? The response of cells to mechanical deformation depends, in general, on which chemical groups are being distorted. For instance, in a study of cultured vascular smooth muscle cells subjected to periodic strain, Wilson and Kaczmarek (55) found that production and secretion of platelet-derived growth factor and DNA depended on the chemical composition of the substrate. Collagen, fibronectin, and vitronectin were effective, but little response was observed on elastin or laminin. Similarly, if the cells were cultured on pronectin or laminin, cyclic strain caused differential expression of mitogen-activated protein and amino terminal kinase (32). In fibroblasts, mechanically induced increases in cell Ca2+ occurred when α2- or β1-integrin subunits were stressed but not the transferrin receptor (31). L-type Ca2+-channel currents can be activated or inhibited by different ligands for the integrins (56). These kinds of data warn against extrapolating from isolated cells to cells in their normal environment. Aside from the technical difficulties of stretching isolated cells, it is important to get data from cells in their native mechanical environment. Only with that data in hand can we properly extend the studies on isolated cells to discover which ligands produce the responses observed in vivo.
The authors thank Dr. Wade J. Sigurdson for assistance in experimental setup, Dr. Stephen Besch for assistance in adhesive testing, and Mary Teeling for technical support.
Address for reprint requests and other correspondence: F. Sachs, Dept. of Physiology and Biophysics, SUNY, Buffalo, NY 14214 (E-mail:).
This work was supported by grants from the National Institutes of Health and the United States Army Research Office to F. Sachs.
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- Copyright © 2000 the American Physiological Society