Shear stress triggers insertion of voltage-gated potassium channels from intracellular compartments in atrial myocytes

Significance

The heart is continuously subjected to mechanical forces. The atria in particular are susceptible to changes in the mechanical environment due to their unique position as “pressure sensors.” Here, we show that increased shear stress induces the recruitment of potassium channels from intracellular storage pools to the plasma membrane, via signaling pathways that link the extracellular matrix to the cytoskeleton. This process is altered in myocytes experiencing chronically increased mechanical stress. The incorporation of channels into the membrane causes changes in the electrical activity of the myocyte and may be an important way for cells to adapt to increased mechanical forces.

Abstract

Atrial myocytes are continuously exposed to mechanical forces including shear stress. However, in atrial myocytes, the effects of shear stress are poorly understood, particularly with respect to its effect on ion channel function. Here, we report that shear stress activated a large outward current from rat atrial myocytes, with a parallel decrease in action potential duration. The main ion channel underlying the increase in current was found to be Kv1.5, the recruitment of which could be directly observed by total internal reflection fluorescence microscopy, in response to shear stress. The effect was primarily attributable to recruitment of intracellular pools of Kv1.5 to the sarcolemma, as the response was prevented by the SNARE protein inhibitor N-ethylmaleimide and the calcium chelator BAPTA. The process required integrin signaling through focal adhesion kinase and relied on an intact microtubule system. Furthermore, in a rat model of chronic hemodynamic overload, myocytes showed an increase in basal current despite a decrease in Kv1.5 protein expression, with a reduced response to shear stress. Additionally, integrin beta1d expression and focal adhesion kinase activation were increased in this model. This data suggests that, under conditions of chronically increased mechanical stress, the integrin signaling pathway is overactivated, leading to increased functional Kv1.5 at the membrane and reducing the capacity of cells to further respond to mechanical challenge. Thus, pools of Kv1.5 may comprise an inducible reservoir that can facilitate the repolarization of the atrium under conditions of excessive mechanical stress.

The electrical properties of the myocardium are governed by the interplay of ion channels, whose expression and regulation determines the precise electrical responses of the tissue. The activity of ion channels can be regulated in a variety of ways: for example, interaction with accessory subunits (1), phosphorylation (2), oxidation state (3), and gene expression (4). Recently, increased attention has been focused on trafficking as a means to regulate ion channel function, notably by modulating the number of active channels present at the plasma membrane (5, 6). This regulation is a complex process derived from a balance between trafficking of newly synthesized channel, endocytosis, and recycling/degradation. Trafficking of ion channels is known to be a dynamically regulated process that depends on Rab-GTPases as well as dynamin motors (7, 8). Indeed, certain antiarrhythmogenic drugs have been shown to exert their activity by modifying the number of channels at the plasma membrane (9). In this context, we previously showed that ion channels are recruited from a submembranous pool in response to cholesterol depletion (10). In addition, several ion channels are regulated by mechanical forces, which directly affect the gating of the channel (11) or indirectly activate intracellular signaling pathways to alter channel properties (12).

The myocardium is subjected to a variety of forces with each contraction and therefore must adapt to the various associated mechanical stresses. The response to stretch has been well-studied and includes gene regulation (13), activation of stretch-activated ion channels (14–17), and the release of atrial natriuretic peptide (ANP) (18–21). In addition, it has been shown that direct stretch of β1 integrins activates I_Cl,swell as well as a cation current (22). Less well-studied are the responses of cardiomyocytes to shear stress. Shear forces in the myocardium arise from blood flow and the relative movement of sheets of myocytes, causing cell deformation as the myocardial layers slide against each other with each heart beat (23, 24). Although the effect of shear stress upon cardiomyocytes has not been extensively explored, it has been shown that increased shear stress stimulates intracellular calcium transients (25, 26), induces an increase in the beating rate of neonatal ventricular myocytes (27), and triggers propagating action potentials (APs) in monolayers of ventricular myocytes (28). Thus far, the response to shear stress remains relatively unknown, particularly with regard to ion channel regulation. Ion channel activity determines both the shape of the AP and the firing frequency of excitable cells. Therefore, the response of cardiomyocytes to shear stress is important for normal cardiac excitability and could be central in pathological conditions in which the working conditions of the myocardium are altered.

In this study, we investigate the response of native adult rat cardiomyocytes to shear stress, reproduced in vitro by laminar flow. Using a combination of whole-cell patch-clamp and single-channel recordings, high spatial resolution 3-dimensional and total internal reflection fluorescence (TIRF) microscopy, we show that shear stress induces an increase in outward current and shortens AP duration within the range of a few minutes. This phenomenon is saturable and reversible, and is caused by Kv1.5 exocytosis from the recycling endosome. We identify the mechanotransduction pathway of this recruitment, which involves integrin/focal adhesion kinase (FAK) signaling. Finally, the response to shear stress is altered in chronically hemodynamically overloaded and dilated atria.

Results

Shear Stress Causes an Increase in Outward Current from Atrial Myocytes.

The effect of increased shear stress on atrial myocytes was investigated using the whole-cell patch-clamp technique, at a membrane potential of +60 mV. As shown by Fig. 1A (and Fig. S2), increasing shear stress from 0.5 dyn⋅cm⁻² to 4 dyn⋅cm⁻² elicited an increase in outward current from 4.5 ± 0.3 pA/pF to 52.7 ± 2.3 pA/pF (n = 43, P < 0.001) without change in membrane capacitance. Shear stress of 4 dyn⋅cm⁻² did not induce current increase in ventricular myocytes (n = 10) (Fig. 1A). The effect was observed with a delay of 4–8 min, with a mean onset of 281 ± 23 s (n = 43). The response was slowly reversible with a t_1/2 recovery of 781 ± 54 s (n = 9). It is noteworthy that ∼60% of atrial cells tested responded to shear stress, and this response was isolation-dependent. Shear stresses between 0.5 and 10 dyn⋅cm⁻² were tested, and the response was found to have a threshold of 2.8 dyn⋅cm⁻². There was no change in the magnitude or kinetics of the response from 2.8 dyn⋅cm⁻² to 10 dyn⋅cm⁻², indicating that if the system was activated sufficiently (2.8 dyn⋅cm⁻²) there was an “all-or-nothing” effect. To provide a suprathreshold stimulus, most studies were performed at a shear stress of ∼4 dyn⋅cm⁻².

Fig. 1.

Shear stress causes an increase in outward current from atrial myocytes. (A) Time series of current density showing the increase in sustained outward current recorded from atrial myocytes (pulse from −80 mV to +60 mV) induced by 4 dyn⋅cm⁻² shear stress (n = 37), and recovery under 0.5 dyn⋅cm⁻² (n = 9). Also shown are time series recorded from ventricular myocytes subjected to the same degree of shear stress (n = 9). (B) Current-voltage relationships obtained by depolarizing the membrane from −100 mV to +60 mV in 10 mV increments before (n = 10) and after shear stress (n = 8). Inset shows representative current traces from the same myocyte before and after shear stress (currents shown from −100 mV to +60 mV, 20 mV increments). (C) Pharmacological properties of the shear stress-induced current illustrated as ratio (IDrug/Imax⋅shear): K⁺ channel blocker 4-AP (100 µM, n = 26; ***P < 0.001; 1 mM, n = 14; ***P < 0.001), the Kv1.5 inhibitor AVE0118 (10 μM, n = 8; ***P < 0.001), the general potassium channel blocker TEA (20 mM, n = 8; n.s.), the chloride channel blockers tamoxifen (20 µM, n = 8; *P < 0.05) and DIDS (100 μM, n = 6; *P < 0.05), 4-AP (1 mM) plus DIDS (100 μM) (n = 8; ***P < 0.001), and the stretch-activated ion channel blocker gadolinium (30 μM, n = 6; n.s.). (D) Time series of outward current density from a representative myocyte in response to shear stress followed by the effect of 100 μM DIDS and 1 mM 4-AP. (E) Representative action potentials recorded from myocytes before and after shear stress. (F) Time series showing the reduction in action potential duration at 90% of repolarization in response to shear stress.

Estimation of Shear Stress in the Rat Atrial Myocardium.

To evaluate the relevance of the shear stress value used in vitro, interlaminar shear stress in the rat atria was estimated using the Couette flow model (Fig. S3). In this simplified model, shear stress is calculated on the surface of two parallel plates, one of which is sliding relative to the other. Shear stress (τ) is dependent on the viscosity of the fluid (μ), the distance between the plates (d), and the velocity of the sliding plate (U_o). Due to the extremely limited amount of data relating to the mechanics and anatomy of the atria, certain assumptions were made to estimate the shear stress (the choices of the parameters used to measure shear stress are detailed in Fig. S3). The plate velocity (U_o) can be estimated from the shear strain rate and the dimensions of myocardial laminae. Both myocardial laminae (19.9 ± 3.7 μm, n = 9) and gaps (6.5 ± 2.3 μm, n = 9) were measured from representative phase-contrast images of the rat left atria using ImageJ (Fig. S3A). These measures were used to calculate the shear stress of the system at a resting heart rate in adult rat of ∼5 Hz [300 beats per minute (bpm)], and the calculated atrial shear stress was found to be 0.43 dyn⋅cm⁻² (Materials and Methods). This rough approximation of the atrial shear stress is within an order of magnitude of the experimental threshold value of 2.8 dyn⋅cm⁻².

Shear Stress Primarily Induces an Increase in Outward Potassium Current.

We characterized the nature of the current stimulated by shear stress. The reversal potential of the shear stress-induced current approximately followed the calculated equilibrium potential for K⁺: at E_K = −86 mV, E_REV = −77.3 ± 1.2 mV (n = 8); at E_K = −6 mV, E_REV = −9.6 ± 2.1 mV (n = 5), indicating that shear stress primarily activates a K⁺ conductance in atrial myocytes. The voltage-gated K⁺ (Kv) channel blocker 4-aminopyridine (4-AP) reversibly inhibited the shear stress-induced current by 59.7 ± 3.2% (n = 26; P < 0.001) and 69.2 ± 3.4% (n = 14; P < 0.001) at 100 µM and 1 mM, respectively (bar graph, Fig. 1C). Because at 100 µM 4-AP principally inhibits Kv1.5 channels (29), this data suggests that shear stress primarily modulates I_Kur. Another recognized blocker of I_Kur, AVE0118 (30) (10 μM), also inhibited the shear stress-induced current by 49.3 ± 5.3% (n = 9; P < 0.001). The current increase was not inhibited by 20 mM tetraethylammonium (TEA) (n = 8, n.s., Fig.1C), further supporting the involvement of Kv1.5 channel in the shear-stress effect (31). The transient outward (I_to,fast and I_to,slow) was never increased upon shear stress, eliminating the involvement of Kv1.4/7 and Kv4.2/3, the molecular basis of I_to.

As K⁺ channel blockers did not fully inhibit the shear stress-induced current, we assessed the involvement of other conductances, such as chloride and stretch-activated ion channels (Fig.1C). The I_Cl,swell inhibitor tamoxifen (20 μM) reduced the current by 23.2 ± 3.1% (n = 8; P < 0.05). However, tamoxifen can inhibit other currents in myocytes, including K⁺ currents (32). We used another inhibitor, 4,4′diisothio-cyanatostilbene-2,2′-disulfonic-acid (DIDS), at a concentration reported to specifically block I_Cl,swell (100 μM) (33). DIDS inhibited the current by 22.0 ± 3.1% (n = 6; P < 0.05). When 1 mM 4-AP was applied following 100 μM DIDS (bar graph, Fig. 1C and example time-course, Fig. 1D), the current was blocked to preshear stress levels (4.5 ± 0.4 pA/pF before shear vs. 11.9 ± 2.9 pA/pF after DIDS plus 4-AP, n = 8; n.s.). The current was insensitive to 30 μM gadolinium (n = 8; n.s.) excluding the involvement of stretch-activated ion channels (Fig. 1C). These results confirm that the majority of the shear stress-induced current is carried by K⁺, with ∼20% attributable to activation of chloride channels and a further 10% of the increased current remaining unidentified.

The current density was increased across all voltages tested, with an initial linear current-voltage relationship followed by a slight outward rectification (Fig. 1B). To investigate the biophysical properties of the K⁺ current stimulated by shear stress, we isolated the 4-AP–sensitive component: total outward current upon shear stress minus current recorded under 4-AP. The activation–V_m relationship of the 4-AP–sensitive current was shifted to the left upon shear stress (unsheared, V_0.5 = 12.2 ± 3.7 mV, n = 9 vs. sheared, V_0.5 = −25.9 ± 2.4 mV, n = 5; P < 0.001), and the slope factor (k) was decreased (unsheared, 18.3 ± 1.2 mV, n = 9 vs. sheared, 13.9 ± 1.0 mV, n = 5, P < 0.001). By mathematically removing the leftward shift of the activation–V_m relationship, we found that only ∼20% of the increase of the 4-AP–dependent component was due to the activation change (Materials and Methods).

As K⁺ currents have precise roles in shaping the AP, we investigated the effect of the shear stress-induced current on the AP duration measured at 90% repolarization (APD90) in freshly isolated atrial myocytes. Indeed, as shown in Fig. 1E, shear stress shortened APD90 from 51.4 ± 4.5 ms to 22.1 ± 0.9 ms (n = 4, P < 0.01), with a time course that mirrored the increase in current (Fig. 1F). No change in the resting membrane potential was observed. These data are supportive of the primary ion channel involved in the response to shear stress being Kv1.5, this channel being responsible for the repolarization phases of the cardiac AP.

Shear Stress Increases the Surface Density of Potassium Channels.

We previously showed that Kv1.5 is recruited from a submembranous pool in response to cholesterol depletion (10). To examine the hypothesis that the increase in current caused by shear stress results from K⁺ channel recruitment, atrial myocytes were transduced with adenovirus encoding EGFP-Kv1.5. The distribution of EGFP-Kv1.5 was similar to Kv1.5 in native myocytes (freshly isolated or cultured myocytes) (Fig. S5). The EGFP-Kv1.5 fluorescence measured in real time at the sarcolemma by TIRF microscopy showed a progressive increase upon shear stress (Fig. 2A and Movie S1). The relative evanescent field fluorescence at the start of the recording was 2.6 ± 2.3% (n = 10) and increased to a plateau of 75.6 ± 6.9% (n = 10) at ∼480 s after the initiation of shear stress (P < 0.001) (Fig. 2B). In pre-shear stress conditions, EGFP-Kv1.5 channels showed a punctate distribution at the sarcolemma (vesicle mean size, 0.11 ± 0.01 µm², n = 6) with dynamic movement in the x/y axis (Movie S2). Shear stress of 4 dyn⋅cm⁻² induced the development of an increase in brightness (t_1/2 = 119.7 ± 10.2 s, n = 10) corresponding to accretion of EGFP-Kv1.5–containing vesicles into clusters in the sarcolemma (cluster mean size, 11.44 ± 1.38 µm², n = 6).

Fig. 2.

Shear stress stimulates the recruitment of functional Kv1.5 channels. (A) Representative TIRF images recorded from a single myocyte expressing EGFP-Kv1.5 upon shear stress. The dashed lines outline the cell border. (Scale bar: 10 µm.) (B, Left) Mean time course of the increase in fluorescence in response to shear stress vs. preshear (t_1/2, 120 ± 5s, n = 10). (Right) Bar graph showing the percent increase in average fluorescence before and after shear stress (n = 10; ***P < 0.001). (C) Sample channel recordings from cell-attached membrane patches of adult rat atrial myocytes during 750 ms pulses from −80 to +80 mV before application of shear stress, after 310 s 4 dyn⋅cm⁻² shear stress and after 350 s 4 dyn⋅cm⁻² shear stress. (D) Amplitude histograms determined from the corresponding current traces indicated. C, closed state; L1–L4, levels corresponding to the successive measurable current levels that are the indication of the number of active channels present in the membrane patch. Gaussian fitted curves corresponding to each current level are also shown.

The insertion of functional K⁺ channels was further investigated using single-channel recordings. After 4- to 8-min exposure to shear stress, an increase in channel activity was observed in freshly isolated atrial myocytes (Fig. 2C). Upward openings of K⁺ channels were observed under control conditions (mean current from channel openings, 1.3 ± 0.7 pA, n = 4) with an elementary conductance of 12.4 ± 2.5 pS (n = 4). After ∼6 min of shear stress, a marked increase in channel activity occurred, reaching a maximum value of 9.2 ± 3.5 pA (n = 6). Example traces from the start of the recording, and from time points during the exponential phase of the increase in channels (310 s and 350 s) are shown in Fig. 2C. Amplitude histograms have been determined from the corresponding current traces (Fig. 2D) and are indicative of the increase in the number of active channels present in the membrane patch upon shear stress. Taken together, these experiments indicate that shear stress stimulates the recruitment and the accretion of functional Kv1.5 channels at the sarcolemma.

Shear Stress Stimulates Channel Exocytosis from the Recycling Endosome and Requires an Intact Microtubule Network.

Because vesicles are known to use the microtubule system to transport cargo (34–36), we used colchicine to destroy the microtubule network and examined the effect of shear stress. EGFP-Kv1.5–infected atrial myocytes showed association with the microtubule network that was disrupted upon treatment with 10 μM colchicine (Fig. 3A). Colchicine treatment (2 h) attenuated the response to shear stress (17.9 ± 3.0 pA/pF vs. shear stress in control condition, 52.6 ± 2.3 pA/pF, n = 8, P < 0.001), but the response was not completely prevented (colchicine vs. control, 4.3 ± 0.3 pA/pF, n = 8, P < 0.05) (Fig. 3 B and C). In contrast, destruction of the actin cytoskeleton with cytochalasin-D (5 µM, 24 h) did not affect the response of atrial myocytes to shear stress (4 dyn⋅cm⁻² control, 26.2 ± 8.1 pA/pF vs. 4 dyn⋅cm⁻² with cytochalasin-D, 25.7 ± 7.9 pA/pF, n.s.) (Fig. S4), suggesting that the actin cytoskeleton is not needed for the recruitment of vesicles in response to shear stress.

Fig. 3.

Shear stress triggers Kv1.5 channel exocytosis. (A) Immunocytochemistry showing the localization of EGFP-Kv1.5 (green) and association with microtubules (red) in the absence or presence of 10 μM colchicine. (Scale bar: 10 µm.) Nuclei are stained with DAPI (blue). (B) Time series of current density recorded from an atrial myocyte treated or not with 10 μM colchicine and subjected to shear stress. Insets are enlargements of region of interest (ROI) in Fig. 3A showing that 2 h colchicine treatment is sufficient to destroy the microtubule network in atrial myocytes stained with anti-tubulin antibody. (C) Bar graph showing the average current recorded before (white bar) and after shear stress in control medium (black bar) (vs. before shear, n = 8; ***P < 0.001) and after 10 μM colchicine treatment (red bar) (n = 8; *** P < 0.001). Note that under colchicine treatment the shear stress-induced current is reduced but not abolished (n = 8; *P < 0.05). (D) Time series of current density from a representative myocyte subjected to shear stress in conjunction with the intracellular dialysis of the SNARE protein inhibitor 250 μM NEM (red), or 10 mM BAPTA (blue). (E) Bar graph showing average current densities in response to shear stress in control intracellular solution (n = 8; ***P < 0.001) or in the presence of 250 μM NEM (vs. before shear, n = 8; n.s.) or 10 mM BAPTA (vs. before shear, n = 8; n.s.). (F) Bar graph showing the average current density from myocytes transfected with empty vector (n = 8) or Rab11DN (n = 6) and subjected to shear stress. The effect of shear stress was significant only with the empty vector (***P < 0.001).

Exocytosis is reliant upon fusion of vesicles to the plasma membrane, a process mediated by SNARE proteins. Intracellular dialysis of the SNARE protein inhibitor N-ethylmaleimide (NEM) (250 µM) followed by 4 dyn⋅cm⁻² shear stress prevented the increase in current (n = 8) (Fig. 3 D and E). As intracellular calcium may be required for exocytosis, we tested the calcium dependency of the shear stress-induced current in freshly isolated atrial myocytes. Intracellular dialysis of the calcium-specific chelator 1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) (10 mM) prevented the increase in current (n = 8) (Fig. 3 D and E).

To identify the compartment responsible for delivery of Kv1.5 channels, cultured atrial myocytes were transfected with a dominant negative (DN) form of Rab11 or with empty vector. Rab11DN overexpression significantly reduced the effect of shear stress (39.5 ± 9.0 pA/pF in empty vector, n = 8, vs. 12.4 ± 7.0 pA/pF in Rab11DN, n = 6; P < 0.001) (Fig. 3F), indicating that the increased current is due to recruitment of channels from the Rab11-associated recycling endosome.

Shear Stress Stimulates the Integrin/FAK Signaling Pathway.

Integrins are central in mechanotransduction processes, conveying mechanical forces into the cell. To investigate whether the integrin system is involved in the response to shear stress in atrial myocytes (22), we used the “disintegrin” echistatin toxin. Preincubation with echistatin (100 nM) for 30 min prevented the response to shear stress in freshly isolated atrial myocytes (n = 10) (Fig. 4 A and B).

Fig. 4.

Integrin/FAK signaling is the mechanotransducer for shear stress effect. (A) Time series of current density recorded from a myocyte pretreated for 30 min with the disintegrin echistatin (100 nM) and subjected to shear stress. (B) Bar graph showing average current before (white bar) and after shear stress in control medium (black bar) (vs. before shear, n = 10; ***P < 0.001). The 100 nM echistatin treatment (red bar) prevented the effect of shear stress (vs. no shear, n = 10; n.s.). (C, Upper) Shear stress increases FAK activation in cultured atrial myocytes stained with anti-phospho FAKY397 antibody (red), α-actinin (green), and DAPI (blue). (Scale bar: 10 µm.) Arrows indicate FAKY397 at cell-to-cell contacts in unsheared and sheared myocytes. Note that FAKY397 is redistributed in shear-stressed myocytes as show by enlargement of ROI. (Scale bar: 5 µm.) (Lower) Bar graph quantifying FAK activation (vs. before shear, n = 3; ***P < 0.001). (D) Time series of current density recorded from a myocyte subjected to shear stress in the presence of intracellular FAK inhibitor FAKi14 (50 μM). (E) Bar graph showing the average current amplitude of the response before (white bar) and after shear stress in control (black bar) (vs. before shear, n = 11; ***P < 0.001) or in the presence of the focal adhesion kinase inhibitor FAKi14 (red bar) (vs. before shear, n = 11; n.s.).

Focal adhesion kinase (FAK) is an early downstream effector of integrins and has been shown to be activated in stretched cardiomyocytes (22). The involvement of FAK in the response to shear stress was studied by measuring phosphorylation of FAK tyrosine 397. Cultured atrial myocytes were subjected to 4 dyn⋅cm⁻² shear stress in a laminar flow chamber, and the amount of FAKY397 was examined by deconvolution microscopy (Fig. 4C). When quantified, FAKY397 fluorescence increased from 0.24 ± 0.04 arbitrary units (AU) in unsheared myocytes to 0.67 ± 0.06 AU in shear stressed myocytes (n = 3, n = 10 fields; P < 0.001) (Fig. 4C). The subcellular distribution of FAKY397 was also modified upon shear stress: whereas FAKY397 staining was restricted to cell-to-cell contacts in unsheared cardiomyocytes, FAK397 was also detected in the entire cell bodies of sheared myocytes (Fig. 4C, enlargements). FAK inhibitor 14 (1,2,4,5-benzenetetramine tetrahydrochloride) was used to prevent the activation of FAK. When dialyzed intracellularly with FAK inhibitor 14, freshly isolated atrial myocytes no longer responded to shear stress (Fig. 4 D and E).

Altogether, these results indicate that integrin is the mechanosensor that conveys the shear stress signal via the activation of FAK to trigger the release of Kv1.5 channels to the sarcolemma.

Up-Regulation of the Integrin/FAK Signaling Pathway Is Associated with a Reduced Response to Shear Stress in Hemodynamically Overloaded Atria.

To investigate the importance of the shear stress-induced current in situ, we used a rat model of heart failure (HF) and atrial hemodynamic overload, dilation, and fibrosis (37–39). As similar models have been shown to induce alterations in the integrin/FAK system (the mechanosensor) (40–42), we hypothesized that the response of myocytes to shear stress would be altered. As previously described (38), in this model, left-ventricular dysfunction (ejection fraction in HF rats, 55.9 ± 3.6%, n = 11 vs. sham, 79.4 ± 0.6%, n = 11; P < 0.001) was associated with a marked atrial dilation as assessed by echocardiography (surface of left atria in HF rats, 0.37 ± 0.02 cm², n = 11 vs. sham, 0.19 ± 0.01 cm², n = 11; P < 0.001).

The expression of integrinß1 was increased twofold in HF rat atria as quantified by Western blot (P < 0.001) and illustrated by integrinß1D staining in atrial slices (Fig. 5A and Fig. S6). Western blot quantification of FAK expression showed a 20% increase in HF rat atria (P < 0.001). Moreover, the endogenous activation of FAK was also increased in myocytes from dilated atria as shown by immunocytochemistry performed on freshly isolated cells (Fig. 5B).

Fig. 5.

The response to shear stress is altered in a rat model of atrial hemodynamic overload. (A, Upper) Atrial sections from sham and HF rats stained with anti-integrinß1D (green), α-actinin (red), and DAPI (blue) showing the increased integrinß1D staining in HF tissue. (Lower) Representative examples of integrinß1 protein expression in the left atria showing ∼100% increase in HF rats after normalization to GAPDH (sham, n = 4; HF, n = 9; **P < 0.01). (B, Upper) Freshly isolated atrial myocytes from sham and HF rats stained with anti phospho-FAKY397 (green), α-actinin (red), and DAPI (blue), showing endogenous activation of FAK. (Scale bar: 10 µm.) Enlargement of ROI showing FAK397 staining (1) within the cell and (2) at the intercalated disk. (Lower) Representative examples of FAK protein expression in the left atria showing ∼20% increase in HF rats after normalization to GAPDH (sham, n = 6; HF, n = 10; **P < 0.01). (C, Upper) Bar graph showing basal current density from control and hypertrophied myocytes showing the increase in current density before shear stress in HF rats (*P < 0.05). (Lower) Representative examples of Kv1.5 protein expression in the left atria showing ∼40% decrease in HF rats after normalization to GAPDH (sham, n = 4; HF, n = 9; *P < 0.05). (D) Quantification of the percentage of atrial myocytes isolated from sham or HF rats responding to shear stress (sham, 78%; n = 14 vs. HF, 53%; n = 28). (E) Example time series from myocytes from sham and HF rats exposed to shear stress showing the reduced kinetics of the response in hypertrophied myocytes. (F) Corresponding plots of the kinetics measured as the maximum slope of the response to shear stress (sham, 1.18 ± 0.13 pA/(pF⋅s), n = 11; HF, 0.70 ± 0.09 pA/(pF⋅s), n = 13; **P < 0.01). (G) Plot of the response to shear stress from myocytes from sham (n = 10) and HF (n = 12) rats expressed in fold increase compared with preshear stress current densities. Note the reduced ability to respond to shear stress in hypertrophied myocytes (sham, ∼15-fold increase, n = 11 vs. HF, approximately 9-fold increase, n = 23; **P < 0.01).

The membrane capacitance of atrial myocytes isolated from HF rats was increased (74.2 ± 4.5 pF, n = 13) compared with sham 61.2 ± 3.8 pF (n = 11, P < 0.05), indicative of myocyte hypertrophy. In preshear conditions, I_Kur was increased in hypertrophied myocytes (5.1 ± 2.3 pA/pF, n = 13) compared with sham (3.7 ± 1.4 pA/pF, n = 11, P < 0.05) (Fig. 5C, Upper). In contrast, the amount of Kv1.5 protein was decreased by ∼40% (P < 0.05) in dilated atria (Fig. 5C, Lower). We cannot exclude that increased extracellular matrix protein accumulation and fibroblast proliferation had resulted in an apparent decrease in Kv1.5 from myocytes. However, the mRNA expression of the KCN5A (Kv1.5) gene was also reduced although this did not reach significance, which indicates that KCN5A was not stimulated during this hypertrophic process (Fig. S6). These observations supported an up-regulation of integrin/FAK signaling in the hemodynamically overloaded and dilated atria, which could contribute to the increase in density of functional channels at the sarcolemma, despite no increase of Kv1.5 protein.

Due to the changes in the mechanosensor, we investigated the response to shear stress in hypertrophied myocytes. The response to shear stress was modified in hypertrophied myocytes. Firstly, the number of hypertrophied myocytes responding to shear stress was decreased compared with sham myocytes [53% (13/28) v.s. 78% (11/14), respectively] (Fig. 5D). Secondly, the kinetics of the response were substantially slower in hypertrophied myocytes: maximum slope of 0.70 ± 0.09 pA/(pF·s), n = 13 vs. 1.18 ± 0.13 pA/(pF·s) in control, n = 11 (P < =0.05) (Fig. 5 E and F). Finally, the percent current increase induced by shear stress was reduced in hypertrophied myocytes: whereas sham myocytes showed ∼15-fold increase in current density (54.7 ± 7.8 pA/pF; n = 11), hypertrophied myocytes only showed approximately 9-fold increase (45.2 ± 4.5 pA/pF; n = 13) (Fig. 5G). These results indicate that the shear stress-induced current is altered in dilated and fibrotic atria, which is also characterized by an up-regulation of the integrin/FAK signaling pathway.

Discussion

In this study, we describe a previously undescribed mechanism by which voltage-gated ion channels are regulated in atrial myocytes (for summary, see Fig. 6). Shear stress stimulates a large outward current within 4–8 min, the majority of which is carried by K⁺ channels despite a small but significant contribution from chloride channels. Using direct measurement and visualization of channels, we show that shear stress triggers Kv1.5 channel delivery to the sarcolemma. Integrins are the sensors of shear stress and convey the mechanical stimulation intracellularly to the Rab11-associated slow-recycling endosome, via activation of FAK. Consequently, shear stress represents a previously undescribed means to modulate the density of repolarizing current and therefore to tune the electrical activity of atrial myocytes.

Fig. 6.

Schematic representation of Kv1.5 recruitment to the sarcolemma in response to shear stress. Increased shear stress is sensed by the extracellular matrix (ECM) and integrins that convey the signal through phosphorylation of focal adhesion kinase (PFAK). This intracellular signaling pathway leads to the stimulation of the Rab11-associated slow recycling endosome (RE) walking along microtubules. The fusion of the donor compartment (RE) with the acceptor membrane (sarcolemma) causes the release of Kv1.5 and its accretion at the sarcolemma. As a consequence, an increase in current density is observed together with a shortening of the action potential (AP). During hemodynamic overload, a condition in which both the ECM and the myocardium are remodeled, this mechanotransduction pathway is likely chronically stimulated, leading to an increase in the number of Kv1.5 channels at the sarcolemma.

Constitutive exocytosis shares several common features with regulated (fast) exocytosis. Cargo proteins are transported into vesicles, tethered to microtubules and the actin cytoskeleton via small Rab GTP-ases, and inserted into the membrane after fusion of donor and acceptor membranes, under the control of SNARE proteins (43, 44). Three important parameters emerge from our experiments: the time dependency, the saturability, and the reversibility of the shear-stress effect. Despite the increase in fluorescence observed with TIRF microscopy preceding the mean onset of the current increase, initiation of both processes occurred with a delay between the onset of shear stress and the onset of the response, suggesting the involvement of an intracellular cascade. However, once initiated, both EGFP-Kv1.5 fluorescence and current increase followed a Boltzmann-like distribution and reached a plateau at ∼400 s. These two observations suggest that (i) the reservoirs of channels recruited by shear stress are not located just beneath the sarcolemma and (ii) a single compartment is likely responsible for the delivery of Kv1.5 channels. Our results support Kv1.5 exocytosis, as the shear stress-induced current increase was abolished by the SNARE inhibitor NEM and by intracellular calcium buffering of submembrane domains with BAPTA. Moreover, we demonstrated dependency on the Rab11-associated slow recycling endosome and an intact microtubule network for the effect to occur. This finding is reminiscent of our previous observation that cholesterol depletion stimulated trafficking of Kv1.5 from the recycling endosome to the sarcolemma in ∼7 min (10, 45). In addition, the delivery to the sarcolemma of the Glut4 transporter triggered by insulin stimulation is dependent on Rab11 within the same time scale (46). The Rab11-associated endosome is usually considered a slow route for recycling of integral proteins. For instance, the turnover of Kv1.5 from the recycling endosome to the plasma membrane takes ∼72 h (47). Together with our previous observations (10), these results point to the slow recycling endosome being a preferential storage compartment for stimulated exocytosis of Kv1.5 channels. Finally, the current required over 20 min to return to baseline levels. This observation suggests that compensatory endocytosis mechanisms occur. At the present time, the endocytosis pathways for Kv1.5 channels have not been elucidated, nor are the kinetics of constitutive endocytosis in atrial myocytes known.

It is noteworthy that changes in the activation properties of the 4-AP–sentitive component account for ∼20% of the shear-activated current. One can speculate that accretion of Kv1.5 channels upon shear stress into distinct lipid and protein environments can modulate their biophysical properties. Such changes in the voltage dependency and activation slope of Kv1.5-mediated current have been reported previously, for example, when associated with Kvβ-subunits (1) or following cholesterol/caveolae modifications (48).

The importance of integrins in regulating the activity of ion channels has been demonstrated in several cell types, including vascular smooth muscle (49, 50), endothelium, (51) and neurons (52). In the myocardium, integrin activation is necessary for mechano-electrical transduction, as evidenced by the fact that the increased beating rate caused by shear stress in ventricular myocytes can be prevented by integrin inhibition (27). Integrins are also involved in calcium homeostasis and regulate the L-type calcium channel (53). In addition, and Baumgarten showed that direct stretch of integrins activates a mixed (chloride/cationic) current via activation of FAK in ventricular myocytes (22). Here we show that the shear stress-induced current is preventable with echistatin and FAK inhibitor 14. As such, the integrin/FAK transduction pathway probably constitutes a major mechanism by which myocytes modulate their electrical activity in response to mechanical stimuli.

The range of shear stress used in this study is consistent with that shown to have physiological effects on cardiomyocytes in other reports. The 1 dyn⋅cm⁻² shear stress affects the APD of neonatal rat ventricular myocytes (NRVM) (54) whereas 0.5–5 dyn⋅cm⁻² activates ERK in a 3D cardiomyocyte array (55). In addition, shear stresses of 5–10 dyn⋅cm⁻² induce differentiation of mesenchymal stem cells into cardiomyocytes capable of ANP secretion (56, 57). There is currently no technique to directly measure the shear stress experienced by cardiomyocytes in the working myocardium. Using the Couette flow model, the shear stress was estimated to be ∼0.5 dyn⋅cm⁻² in rat atria (Fig. S3). This value is below the threshold of 2.8 dyn⋅cm⁻² and is consistent with the increase in current seen in this study being a response to elevated shear stress. Changes in shear stress level could occur physiologically as a result of increased heart rate or increased contractile reserve, or pathophysiologically as a result of increased laminar thickness (hypertrophy) or decreased gap distance (fibrosis). In hemodynamically overloaded and dilated atria, we observed increased expression of integrinβ1D, as well as FAK activation, suggesting an overactivation of the integrin/FAK signaling pathway as previously reported by others (40–42). In addition, in this model, the capacity of myocytes to respond to shear stress was reduced. Taken together, these data support the idea that the shear stress-induced current increase is affected when the working conditions of the atria are altered. This excessive recruitment of potassium channels following shear stress might overcompensate for the decrease in channel content in the diseased atrial myocardium. These results constitute further evidence that the shear stress-induced current is functional in situ. It is noteworthy that, in the working myocardium, various mechanical stresses are likely to activate the integrin-signaling pathway including direct stretching of the integrins anchored to the extracellular matrix; these mechanical stresses might also trigger the recruitment of Kv1.5 channels to the plasma membrane.

Physiological Relevance.

As the shear stress response was atria-specific, we speculate that the increased current (and subsequent decreased APD) is related to the function of the atria. The atria have an important reservoir function that is essential for both the filling of ventricles and ANP secretion. Indeed, ANP secretion is affected by changes in the extracellular space and fluid volume, both of which may impact the shear stress of the atria (58). ANP release is calcium- (59, 60) and stretch-dependent (19, 20). Additionally, there is evidence that shear stress elicits calcium sparks in atrial myocytes (26) and that acceleration of membrane repolarization (such as the shortening of the AP seen in this study) inhibits stretch-induced secretion of ANP by shortening APD (61). We hypothesize that the shortened AP upon shear stress may be part of this system of ANP regulation in conditions of atrial mechanical overload and may constitute a mechanism by which the atria can repolarize.

Conclusion.

The lifetime of ion channels at the membrane has emerged as a major determinant of current properties and cell excitability. Most of the knowledge on the processes regulating ion channel density, particularly K⁺ channels, concerns their internalization (8, 62–64) whereas little is known about forward trafficking. Our discovery of the existence of reservoirs of Kv1.5 channels that are recruitable in response to mechanical stimulation of ECM/integrin complexes constitutes an important breakthrough in the understanding of Kv1.5 channel regulation.

Our results suggest that the density of functional Kv1.5 channels at the sarcolemma results from an equilibrium between pools of channels inserted in the membrane and pools of channels recruitable from intracellular vesicles. In addition to constitutive exocytosis, these results indicate that Kv1.5 channels can also undergo regulated exocytosis, which appears to depend on the integrity of the extracellular matrix.

Importantly, Kv1.5 expression is not confined to the heart but is expressed in various cell types, including neurons, pulmonary arteries, skeletal muscle, and endocrine cells (4). In addition, this Kv channel is involved in several pathological processes, including atrial fibrillation (65, 66), pulmonary hypertension (67), multiple sclerosis (68), and the proliferation of cancer cells (69). As such, the regulation of this channel through manipulation of its availability is likely to be relevant in these diseases, particularly in conditions in which integrin activity or the mechanical environment is altered.

Materials and Methods

Animals.

Adult male Wistar rats (Janvier) were treated in accordance with our institutional guidelines (Ministère de l’Agriculture, France, authorization 75–1090), and treatment conformed to the Directive 2010/63/EU of the European Parliament.

Cell Isolation, Cell Culture, and Gene Transfer.

Adult cardiomyocytes isolated by Langendorff perfusion were used for patch-clamp experiments. Briefly, hearts were cannulated and retrogradely perfused at 5 mL/min through the aorta, first with a solution containing (mM): NaCl 100, KCl 4, MgCl₂ 1.7, glucose 10, NaHCO₃ 5.5, KH₂PO₄ 1, Hepes 22, 2,3-butanedione monoxime (BDM) 15, taurine 10, pH 7.4 (NaOH) at 37 °C for 5 min, then with enzymatic solution containing (mM): NaCl 100, KCl 4, MgCl₂ 1.7, glucose 10, NaHCO₃ 5.5, KH₂PO₄ 1, Hepes 22, BDM 15, and 200 UI/mL collagenase A (Roche Diagnostics) plus 0.5% BSA, pH 7.4 (NaOH) at 37 °C for ∼17 min. Solutions were bubbled with 95% O₂/5% CO₂ throughout. Atria and ventricles were separated and placed into Kraft-Brühe (KB) buffer containing (mM): glutamic acid potassium salt monohydrate 70, KCl 25, taurine 20, KH₂PO₄ 20, MgCl₂ 3, EGTA 0.5, glucose 10, Hepes 10, pH 7.4 (KOH). The tissue was then cut into small pieces and gently agitated to dissociate single myocytes. Myocytes were plated onto laminin-coated Petri dishes and maintained in KB buffer until use. Atrial myocytes for use in overexpression experiments, or for experiments requiring more than 24 h treatment with drugs, were isolated as previously described (45). Cultured myocytes were transfected with the dominant negative (DN) construct of Rab11 (S25N) cloned into a pEGFP-C1 (kind gift from D. Fedida, Department of Anaesthesiology, Pharmacology and Therapeutics, University of British Columbia, Vancouver) using a liposome-based approach and low CO₂ condition (37 °C, 1%) (70) or transduced with the K_v1.5 construct cloned in pEGFP-N3 and subcloned in adenovirus (10).

In Vitro Generation of Shear Stress.

Shear stress was replicated in vitro using a perfusion system placed 40 cm above the experimental chamber and attached to a flow regulator. The magnitude of the shear stress at the outlet was calculated using the equation:

where τ is the shear stress in dyn⋅cm⁻², µ is the viscosity of the fluid (1.002⋅10⁻² dyn⋅s/cm² for water at 20 °C), Q is the volumetric flow rate (cm³⋅s⁻¹), and r is the internal radius of the perfusion tip (cm). The shear stress generated by this system could be varied by changing the speed of the perfusion such that shear stresses between 0.5 and 10 dyn⋅cm⁻² (corresponding to 0.05 N⋅m⁻² to 0.1 N⋅m⁻²) could be generated. The tip of the perfusion (internal diameter of 0.58 mm) was placed ∼100 µm from the cell; therefore the calculated shear stress represents a maximum value experienced by the cell. To shear stress fields of cells, a VC-LFR-18-SS laminar flow chamber was used (C&L Instruments). The chamber was attached to the flow regulator and the shear stress was calculated using the equation:

where τ, µ, and Q have the same meaning as above, b is the chamber width (cm), and h is the chamber height (cm). For the VC-LFR-18-SS chamber, a perfusion rate of 2 mL⋅min⁻¹ generates a shear stress of 4 dyn⋅cm⁻². See Fig. S1 for schematic representations of the devices used to generate shear stress using the above methods.

Electrophysiology.

Whole-cell patch-clamp recordings were obtained from myocytes in voltage- or current-clamp mode using a patch-clamp amplifier (Axopatch 200B; Molecular Devices). Patch pipettes had resistances between 1 and 2 MΩ for whole-cell and 10 and 15 MΩ for single-channel recordings. Currents were low-pass filtered at 10 kHz (−3 db) and digitized with NI PCI-6251 (National Instruments). Data were acquired and analyzed with Elphy 2.0 software (G. Sadoc, CNRS, Gif-sur-Yvette, France). For AP recording, pipette solution contained (mM): K-aspartate 115, NaCl 5, EGTA 5, Hepes 10, Mg-ATP 3, MgCl₂ 2, Tris phosphocreatine-5 5, pH 7.2 (KOH). Perfusate contained (mM): NaCl 140, KCl 4, CaCl₂ 2, MgCl₂ 2, NaH₂PO₄ 1, pyruvate 2.5, Hepes 10, glucose 20, pH 7.4 (NaOH). APs were evoked by 10 ms current pulses at 0.2 Hz. For whole-cell current recording, pipette solution contained (mM): K-aspartate 115, KCl 10, KH₂PO₄ 2, MgCl₂ 3, Hepes 10, glucose 10, CaCl₂ 0.1, and Mg-ATP 5, pH 7.2 (KOH). Standard perfusate contained (mM): NaCl 140, KCl 4, MgCl₂ 2, NaHPO₄ 1, glucose 20, Hepes 10, CaCl₂ 2 (E_K = −86 mV), pH 7.4 (NaOH). For time series analysis, currents were evoked by depolarization from −80 mV to +60 mV for 750 ms at 0.2 Hz. I_Kur amplitudes were measured at the end of the test pulse relative to the holding current. For the current-voltage protocol, voltage steps ranging from −100 mV to +60 mV were applied from a holding potential of −80 mV in 10 mV increments. Leak current was numerically compensated. To modify the equilibrium potential for K⁺, a high [K⁺] extracellular solution was used (mM): KCl 90, MgCl₂ 2, NaCl 40, NaH₂PO₄ 1, glucose 20, Hepes 10, (E_K = −6 mV), pH 7.4 (KOH). For single-channel recordings, pipette solution contained (mM): NMDG-aspartate 130, K-aspartate 5.4, MgCl₂ 1, Hepes 10, CaCl₂ 1, NaH₂PO₄ 0.33, glucose 10, pH 7.4 (KOH). Perfusate contained (mM): K-aspartate 135, MgCl₂ 1, Hepes 10, CaCl₂ 1, NaH₂PO₄ 0.33, glucose 10, pH 7.4 (NaOH).

The chord conductance (G) was calculated as follows: G = G_max/(1 + exp [−(V_m – V_0.5)/k]), where G_max is the maximum chord conductance, G the chord conductance calculated at membrane potential V_m, V_0.5 the potential at which the conductance is half-maximally activated, and k the slope factor describing the steepness of the activation curve. The Nernst potential for K⁺ ions in our experiments was E_K = −86 mV. The activation-V_m curves and steady-state availability-V_m curves were fitted with a single Boltzmann function: y = 1/(1+exp[−(V_m − V_0.5)/k)]) where V_0.5 is the half-activation potential, V_m the test voltage, and k the slope factor. Mathematical subtraction of the shift of the activation was performed using the following equation: I_Kur(V_m, t →∞) = G_max(V_m)⋅m(V_m, t →∞)⋅(V_m-E_K)/C_m, where V_m is the membrane potential, I_Kur(V_m,t →∞) is the resulting steady state current amplitude, G_max maximum chord conductance of the 4-AP–sensitive component, m(V_m, t →∞) is the steady state activation parameter obtained in unsheared myocytes, E_K is the Nernst equilibrium potential for K⁺ ion, and C_m is the mean cell capacitance. Single-channel recordings were performed in the cell-attached configuration, the patch membrane was clamped from −80 mV to +80 mV for 750 ms at 0.2 Hz.

Couette Flow Model.

The shear stress (τ) was estimated in the rat atria using the following equation:

where μ is the viscosity of the fluid, d the distance between the plates, and U_o the velocity of the sliding plate. The plate velocity can be estimated from the shear strain rate and the dimensions of myocardial laminae. The following measures were used to calculate the shear stress of the system: laminar thickness = 20 μm; gap height d = 6.5 μm; height of laminar unit a = 1/2 (20 μm) + 6.5 μm + 1/2 (20 μm) = 26.5 μm; shear strain E_sn = 1/2 tan (θ) = 1/2 b/a ∼0.15 in canine LV (24); laminar shear motion b = 2 a E_sn ∼2(26.5 μm)⋅(0.15) = 7.95 μm; time t = 0.1 s for resting heart rate in adult rate of ∼5 Hz (300 bpm) with contraction taking half the cycle; viscosity of interstitial fluid μ = 3.5 cP = 0.035 dyn⋅s⋅cm⁻². Finally, the shear stress can be calculated as follows:

Immunofluorescence and 3D Microscopy.

Immunofluorescence (IF) was performed on cardiomyocytes or on atrial cryosections (8-μm sections) fixed with 4% paraformaldehyde (PFA) for 10 min at room temperature. Preparations were incubated for 1 h at room temperature with permeabilizing/blocking buffer (PBS containing 0.1% Triton X-100, 1% BSA, 10% goat serum, and 10% chicken serum) and then incubated with primary antibodies diluted in blocking buffer (PBS containing 1% BSA, 3% goat serum, and 3% chicken serum) overnight at 4 °C. Detection was performed with a 1 h incubation with secondary antibodies AlexaFluor488 or AlexaFluor594 (1:500; Molecular Probes), and DAPI (1:500; Sigma) to detect nuclei. Images were acquired with a cooled CoolSnap camera (Roper-Scientific) on an Olympus epifluorescent microscope (60×, UPlanSApo, 0.17). Images were processed and analyzed using Metamorph software (Molecular Devices) supplemented with the 3D-deconvolution module. For each sample, a series of consecutive planes (stack of images) were acquired (sectioning step, 0.2 μm) and deconvoluted using acquired point spread function. Fluorescence was quantified using ImageJ software (freeware; National Institutes of Health).

Western Blotting and Quantification.

Left atria were transferred to lysis buffer containing (mM) Tris-Hcl 50 (pH 7.5), NaCl 150, EDTA 2, 0.5% Triton X-100, and a protease mixture inhibitor (Sigma) and homogenized using a polytron. The soluble fraction retrieved after 15 min centrifugation at 15,000 × g at 4 °C was used. Then 10 µg of protein was separated on 10% Bis-Tris gels (Invitrogen) and transferred for 2 h at 80 V onto nitrocellulose membrane (Bio-Rad). Membranes were incubated with primary and secondary antibodies and revealed using the Pierce ECL Plus kit. Anti-rabbit and anti-mouse IgG-linked HRP antibodies were used at 1:1,000 (Cell Signaling). All blots were imaged using the Ettan DIGE imager (GE Healthcare), and ImageJ software was used to quantify individual bands.

Antibodies.

Primary antibodies used were mouse anti–α-tubulin (1:400; Sigma), rabbit anti-FAKY397 (1:50; Santa Cruz), mouse anti-sarcomeric α-actinin (1:2,000; Sigma), mouse anti-FAK (1:500; Millipore), rabbit anti-GFP (1:300; Torrey Pines Biolabs), and rabbit anti-GAPDH (1:2,000; Abcam). Integrinß1D antibody was a kind gift from Robert Ross (Veterans Administration Healthcare, San Diego) (1:10,000 for Western Blot; 1:5,000 for IF).

Evanescent Field Microscopy and Image Analysis.

Live imaging experiments were performed at ∼27 °C with cultured myocytes seeded on glass-bottomed microdishes (170-µm thickness; Ibidi) bathed with standard extracellular solution. K_v1.5-EGFP channel dynamics were visualized with total internal reflection fluorescence (TIRF) microscopy using the Olympus Cell^tirf system. Cells were placed on an Olympus IX81-ZDC2 inverted microscope, and TIRF illumination was achieved with the motorized integrated TIRF illuminator combiner (MITICO-2L) using a 60×/1.49 APON OTIRF objective. EGFP was visualized using a 50-mW solid-state 491-nm laser for excitation and dual-band optical filter (M2TIR488-561). All image acquisition, TIRF angle adjustment, and some of the analysis were performed using the xcellence software (Olympus). Time series of 100 images at 5-s intervals were recorded using a digital CCD camera ORCA-ER (Hamamatsu). Cells were imaged for 1 min before applying shear stress of 4 dyn⋅cm⁻² to establish the baseline whole-cell evanescent field fluorescence (EFF). To quantify changes in EFF, the cell perimeter was delineated, and the projection of minimum intensity (EFF_min) of the whole movie was subtracted from each image [EFF_(t)] and normalized to the background intensity measured in a region of interest outside the cell (EFF_bck) using the following equation:

Experimental Model of Atrial Dilatation.

Myocardial infarction was induced in 11 male Wistar rats by ligating the left coronary artery as previously described (38, 71). Sham rats (n = 11) underwent the same surgical procedure without left coronary artery ligation. The cardiopathy was characterized by transthoracic echocardiography in 3 mo after surgery. Rats were killed 4 mo after surgery, and single myocytes were obtained using the Langendorff method. Residual tissue remaining after isolation of single cells was collected, placed in lysis buffer, homogenized, and frozen at −80 °C for use in Western blot experiments. Left atria were fixed with 4% PFA for 10 min at room temperature and embedded in cryomatrix, and 8-µm cryosections were prepared for IF.

Statistics.

Data were tested for normality using the D’Agostino and Pearson normality test. Statistical analysis was performed using Student t test or ANOVA followed by post hoc Bonferroni test on raw data. Results are given as means ± SEM and P values of less than 0.05 were considered significant (*P < 0.05, **P < 0.01, and ***P < 0.001).