Shear stress activates monovalent cation channel transient receptor potential melastatin subfamily 4 in rat atrial myocytes via type 2 inositol 1,4,5‐trisphosphate receptors and Ca²⁺ release

Abstract

Key points

• During each contraction and haemodynamic disturbance, cardiac myocytes are subjected to fluid shear stress as a result of blood flow and the relative movement of sheets of myocytes.

• The present study aimed to characterize the shear stress‐sensitive membrane current in atrial myocytes using the whole‐cell patch clamp technique, combined with pressurized fluid flow, as well as pharmacological and genetic interventions of specific proteins.

• The data obtained suggest that shear stress indirectly activates the monovalent cation current carried by transient receptor potential melastatin subfamily 4 channels via type 2 inositol 1,4,5‐trisphosphate receptor‐mediated Ca²⁺ release in subsarcolemmal domains of atrial myocytes.

• Ca²⁺‐mediated interactions between these two proteins under shear stress may be an important mechanism by which atrial cells measure mechanical stress and translate it to alter their excitability.

Abstract

Atrial myocytes are subjected to shear stress during the cardiac cycle under physiological or pathological conditions. The ionic currents regulated by shear stress remain poorly understood. We report the characteristics, molecular identity and activation mechanism of the shear stress‐sensitive current (I _shear) in rat atrial myocytes. A shear stress of ∼16 dyn cm⁻² was applied to single myocytes using a pressurized microflow system, and the current was measured by whole‐cell patch clamp. In symmetrical CsCl solutions with minimal concentrations of internal EGTA, I _shear showed an outwardly rectifying current–voltage relationship (reversal at −2 mV). The current was conducted primarily (∼80%) by monovalent cations but not Ca²⁺. It was suppressed by intracellular Ca²⁺ buffering at a fixed physiological level, inhibitors of transient receptor potential melastatin subfamily 4 (TRPM4), intracellular introduction of TRPM4 antibodies or knockdown of TRPM4 expression, suggesting that TRPM4 carries most of this current. A notable reduction in I _shear occurred upon inhibition of Ca²⁺ release through the ryanodine receptors or inositol 1,4,5‐trisphosphate receptors (IP₃R) and upon depletion of sarcoplasmic reticulum Ca²⁺. In type 2 IP₃R (IP₃R2) knockout atrial myocytes, I _shear was 10–20% of that in wild‐type myocytes. Immunocytochemistry and proximity ligation assays revealed that TRPM4 and IP₃R2 were expressed at peripheral sites with co‐localization, although they are not localized within 40 nm. Peripheral localization of TRPM4 was intact in IP₃R2 knockout cells. The data obtained in the present study suggest that shear stress activates TRPM4 current by triggering Ca²⁺ release from the IP₃R2 in the peripheral domains of atrial myocytes.

Key points

• During each contraction and haemodynamic disturbance, cardiac myocytes are subjected to fluid shear stress as a result of blood flow and the relative movement of sheets of myocytes.

• The present study aimed to characterize the shear stress‐sensitive membrane current in atrial myocytes using the whole‐cell patch clamp technique, combined with pressurized fluid flow, as well as pharmacological and genetic interventions of specific proteins.

• The data obtained suggest that shear stress indirectly activates the monovalent cation current carried by transient receptor potential melastatin subfamily 4 channels via type 2 inositol 1,4,5‐trisphosphate receptor‐mediated Ca²⁺ release in subsarcolemmal domains of atrial myocytes.

• Ca²⁺‐mediated interactions between these two proteins under shear stress may be an important mechanism by which atrial cells measure mechanical stress and translate it to alter their excitability.

Abbreviations

4‐AP

4‐aminopyridine

CICR

Ca²⁺‐induced Ca²⁺ release

CPA

cyclopiazonic acid

E_rev

reversal potential

IP₃R2

type 1 inositol 1,4,5‐trisphosphate receptor

I_shear

shear stress‐sensitive current

KD

knockdown

KO

knockout

mERG

mouse ether‐à‐go‐go‐related gene

NCX

Na⁺−Ca²⁺ exchanger

PLA

proximity ligation assay

RyR

ryanodine receptor

SAC

stretch‐activated ion channel

siRNA

small interfering RNA

SR

sarcoplasmic reticulum

TRPM4

transient receptor potential melastatin subfamily 4

XeC

xestospongin C

Introduction

Changes in the mechanical environment of the heart, caused by each cardiac cycle, alter cardiac excitation and contraction (Lakatta, 1993; Nazir & Lab, 1996). An increase in atrial pressure and volume under pathological conditions, such as valve disease, hypertension or heart failure, is considered to be an important cause of altered atrial excitability (Nazir & Lab, 1996; Nattel, 2002). Atria are often observed to become enlarged and dilated under such conditions, and the responses of atrial myocytes to stretch, including the activation of stretch‐activated ion channels (SACs), have been well documented (Hagiwara et al. 1992; Sato et al. 1998; Tavi et al. 1998; Zhang et al. 2000; Kamkin et al. 2003). During each contraction and haemodynamic disturbance, cardiac myocytes are also subjected to fluid shear stress as a result of blood flow and the relative movement of sheets of myocytes (LeGrice et al. 1995; Costa et al. 1999). Chronic blood regurgitation as a result of mitral valve diseases elicits jets of blood onto the atrial wall, thereby causing widespread endocardial surface disruption (Goldsmith et al. 2000; Saffitz, 2009) and atrial arrhythmias (Nazir & Lab, 1996). Under such conditions, atrial myocytes are directly exposed to relatively higher fluid shear stress. However, atrial responses to shear stress are not well understood.

Recent evidence indicates that shear stress modulates cardiac Ca²⁺ signals and voltage‐dependent ion channels. Shear stress was shown to cause the propagation of an action potential in a cultured ventricular myocyte monolayer (Kong et al. 2005), enhance the occurrence of atrial Ca²⁺ sparks [i.e. focal Ca²⁺ release through a single ryanodine receptor (RyR) cluster (Cheng et al. 1993)] (Woo et al. 2007), induce global Ca²⁺ waves (Woo et al. 2007; Kim & Woo, 2015) and whole‐cell Ca²⁺ transients (Belmonte & Morad, 2008) in atrial myocytes, enhance ventricular Ca²⁺ transients (Lee et al. 2008), suppress ventricular L‐type Ca²⁺ currents (Lee et al. 2008; Rosa et al. 2013), and up‐regulate atrial ultra‐rapid outward K⁺ currents (Boycott et al. 2013). These responses to shear stress in cardiac myocytes occur even in the presence of SAC inhibitors (Lee et al. 2008; Boycott et al. 2013; Rosa et al. 2013; Kim & Woo, 2015). In particular, the effect of shear stress on Ca²⁺ sparks is larger in the periphery than in the interior of atrial myocytes (Woo et al. 2007) lacking transverse tubules (Carl et al. 1995). At relatively higher shear strength (>15 dyn cm^‒2), a longitudinal global Ca²⁺ wave develops in these myocytes as a result of local Ca²⁺ release generated by the activation of the type 2 inositol 1,4,5‐trisphophate receptor (IP₃R2) via P2Y₁ purinergic signalling and subsequent Ca²⁺‐induced Ca²⁺ release (CICR) through RyRs (Kim & Woo, 2015). These observations raise questions concerning whether ion channels/transporters may be activated by the Ca²⁺ increase under shear stress. To date, it is not known what membrane channels, if any, are activated either directly by shear stress or by Ca²⁺ signalling induced by shear stress.

In the present study, we investigated cell membrane ionic currents in intact and genetically manipulated atrial myocytes subjected to shear stress using the whole‐cell patch clamp technique. We applied a shear stress of ∼16 dyn cm⁻², at which global Ca²⁺ waves accompanying whole‐cell Ca²⁺ increase consistently occur (Kim & Woo, 2015), to single myocytes using pressurized fluid flow as reported previously (Woo et al. 2007). This approach aimed to approximate the mechanical stresses that the atrial wall encounters from the shear force (e.g. the jets of blood onto the atrial wall during mitral regurgitation) or from the excessive fluid pressure produced during atrioventricular valve stenosis and incompetence. We found that shear stress specifically activates a non‐selective monovalent cation current carried by transient receptor potential melastatin subfamily 4 (TRPM4) channels through IP₃R2‐mediated Ca²⁺ release. Immunocytochemistry and electrophysiological evaluation of the effect of Ca²⁺ buffer on the shear stress‐sensitive current (I _shear) further showed that a Ca²⁺‐mediated interaction may occur between TRPM4 and IP₃R2 in certain microdomains in the vicinity of the cell membrane.

Methods

Cell isolation

Atrial myocytes were enzymatically isolated (Lee et al. 2008) from male Sprague–Dawley rats (200–300 g; total 53 rats) and from wild‐type (WT) and IP₃R2 knockout (KO) mice (total 12 mice) (Li et al. 2005) (C57/B6 background, 3–5 months of age, weighing 24–28 g). The present study conforms with the Guiding Principles for the Care and Use of Experimental Animals published by the Korean Food and Drug Administration and the Animal and Plant Quarantine Agency in South Korea. The experimental protocols were approved by the Animal Care and Use Committees of Chungnam National University, South Korea (No. CNU‐00368). Rats were deeply anaesthetized with pentobarbital sodium (150 mg kg⁻¹, i.p.), the chest cavity was opened and hearts were excised. The excised hearts were retrogradely perfused at 7 ml min⁻¹ for rat heart and at 1.9 ml min⁻¹ for mouse heart through the aorta (at 36.5°C), first for 3 min with Ca²⁺‐free Tyrode solution comprising (in mm) 137 NaCl, 5.4 KCl, 10 Hepes, 1 MgCl₂ and 10 glucose (pH 7.3); then with Ca²⁺‐free Tyrode solution containing collagenase (1.4 mg ml⁻¹ for rat; 1 mg ml⁻¹ for mouse; Type A; EC 3.4.24.3; Roche, Basel, Switzerland) and protease (0.14 mg ml⁻¹ for rat, 0.08 mg ml⁻¹ for mouse, Type XIV; EC 3.4.24.31; Sigma, St Louis, MO, USA) for 12 min; and, finally, with Tyrode solution containing 0.2 mm CaCl₂ for 5 min. The atria and ventricles of the digested heart were then cut into several sections and subjected to gentle agitation to dissociate the cells. The freshly dissociated cells were stored at room temperature in Tyrode solution containing 0.2 mm CaCl₂.

Measurement of membrane currents

Membrane currents were recorded using the whole‐cell configuration of the patch‐clamp technique using an EPC7 amplifier (HEKA Elektronik, Lambrecht/Pfalz, Germany). The patch pipettes were made of glass capillaries (Kimble Glass Inc., Vineland, NJ, USA) with a resistance of ∼3 MΩ when filled with the internal solutions. The normal Tyrode solution (see above) containing 2 mm Ca²⁺ (pH 7.4) was used for cellular equilibrium and formation of the gigaseal. Measurement of membrane currents was generally carried out ∼8 min after rupture of the membrane in the external solutions containing (mm) 137 CsCl, 10 Hepes, 1 MgCl₂, 10 glucose and 10 μm nifedipine. At this time of dialysis, diffusion of pipette solution appeared to reach equilibrium, such that the E _rev of control ramp current in symmetrical CsCl solutions almost approached 0 mV. The pipette was generally filled with the internal solution containing (in mm) 137 CsCl, 10 Hepes, 5 MgATP, 1 MgCl₂ and 0.5 EGTA with the pH adjusted to 7.2 with CsOH. In some experiments, various concentrations of EGTA, BAPTA plus Ca²⁺, N‐methyl‐d‐glutamate (NMDG⁺), anti‐TRPM4 antibodies (goat polyclonal, dilution 1:500; epitope: intracellular, within amino acids 290–340 of human TRPM4; Santa Cruz Biotechnology Inc., Dallas, TX, USA) or blocking peptide were added to the pipette solutions. Before applying shear, conditioning pulses from −80 to −10 mV were continuously applied to the cells to maintain the sarcoplasmic reticulum (SR) Ca²⁺ load. A voltage ramp pulse ranging from −120 to +70 mV was applied at 0.1 Hz from a holding potential of −30 mV. The contribution of voltage‐dependent Na⁺ current during the ramp pulse is assumed to be reduced by this holding potential, presumably as a result of cumulative inactivation. Shear stress was normally applied using the same zero Ca²⁺‐, zero K⁺‐containing CsCl‐rich external solutions (see above) but contained 1 mM EGTA. Generation of the voltage‐clamp protocol and acquisition of the data were carried out using pClamp, version 9.0 (Molecular Devices, Foster City, CA, USA) via an A/D converter (Digitata 1322; Molecular Devices). The series resistance was 1.5‐ to 3‐fold the pipette resistance with electronic compensation via the amplifier. The current signals were filtered at 10 kHz before digitization and storage. The currents were analysed using pClamp, version 9, and OriginPro, version 8, SR0 (OriginLab Corp., Northampton, MA, USA). The experiments were performed at room temperature (22–25°C).

Application of shear stress

Pressurized flows of solutions were applied onto the single myocytes through a microbarrel (internal diameter 250 μm) for which the tip was placed ∼150 μm from the cell and connected to a fluid reservoir at a height of 40 cm (Woo et al. 2007; Lee et al. 2008). The tip of microbarrel, touching the chamber bottom, was tilted to one side at an angle of 45°. An electronically controllable solenoid valve was installed in the middle of tubing connecting the fluid reservoir and the microbarrel. The shear stress (dyn cm⁻²) was calculated for flow in cylindrical tubes according to the equation (Olesen et al. 1988):

$$\mathrm{Shear}\mathrm{stress}=4\mu Q/\pi {\mathit{r}}^3$$

where μ is the fluid viscosity (1.002∙10^‒2 dyn s cm⁻² for water), Q is the flow rate (cm³ s⁻¹) and r is the internal radius (cm) of the microbarrel. The microflow system generated shear stress of ∼16 dyn cm⁻² (equal to 0.16 N m⁻²) at a reservoir height of 40 cm. The positioning of the microbarrel was performed under microscope using a micromanipulator (Prior England 48260; Prior Scientific Inc., Rockland, MA, USA). The experimental cells were attached to the bottom of the chamber without a coating material. Use of a microscope and video monitor confirmed that no movement of the cell occurred during the fluid puffing before the start of the experiments.

HL‐1 cells and knockdown (KD) of TRPM4

HL‐1 cardiomyocytes, obtained from Dr W. C. Claycomb (Louisiana State University), were handled as reported previously (Claycomb et al. 1998). HL‐1 cells were grown onto a matrix of gelatin (0.02%; Difco, Franklin Lakes, NJ, USA) plus fibronectin (12.5 mg ml⁻¹; Sigma) into a Claycomb medium (JRH Biosciences, Lenexa, KS, USA) supplemented with 10% fetal bovine serum (JRH Biosciences), 4 mm l‐glutamine (Life Technologies, Grand Island, NY, USA), 0.1 mm norepinephrine (Sigma), and 100 U ml⁻¹ penicillin and 100 μg ml⁻¹ streptomycin (Life Technologies). The cells were incubated at 37°C in 95% O₂/5% CO₂ at a relative humidity of 95%.

Small interfering RNA (siRNA) against TRPM4 (sense, 5′‐GAGUGAUGGCUCGCCUAGAdTdT‐3′; anti‐sense, 5′‐UCUAGGCGAGCCAUCACUCdTdT‐3′; 300 nm; Bioneer Inc., Alameda, CA, USA) and non‐targeting (control) siRNA were transfected for 6 h at 37°C in a CO₂ incubator with Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA; siRNA: lipofectamine = 1:1) in accordance with the manufacturer's instructions. HL‐1 cells were preincubated in serum‐ and antibiotic‐free medium for 30 min before transfection. The cells were used for the measurement of I _shear after 48 h of culture.

Immunocytochemistry and analysis

The atrial myocytes plated onto a cover slip were fixed in 2% paraformaldehyde (in PBS) for 10 min and permeabilized in 0.1% Triton X‐100 for 2 min. The cells were incubated in PBS containing 1% BSA (Sigma) for 1 h to block non‐specific labelling, and then incubated with anti‐TRPM4 [rabbit polyclonal IgG, dilution 1:200; epitope, EKEQSWIPKIFKK(C), corresponding to amino acid residues 5–17 of N‐terminus in human TRPM4; Intracellular; ACC‐044, Alomone Labs, Jerusalem, Israel] and anti‐IP₃R2 (goat polyclonal IgG, 2 μg ml⁻¹; Santa Cruz Biotechnology Inc.) overnight at 4°C. The primary antibodies for TRPM4 and IP₃R2 were detected using chicken anti‐rabbit IgG tagged with Alexa 488 (dilution 1:1000; Molecular Probes, Eugene, OR, USA) and donkey anti‐goat IgG tagged with Alexa 568 (dilution 1:1000; Molecular Probes) for 1 h at room temperature, respectively. The cover slips were treated with mounting medium (Mowiol anti‐fadent; Sigma) and then used for confocal microscopy (A1; Nikon, Tokyo, Japan).

Two dimensional images of 2048 × 1024 pixels were acquired with an optical depth of 1.0 μm. Alexa 488 and Alexa 568 were excited with 488 nm and 561 nm laser beams, respectively, and their emissions were measured at 500–550 nm and 570–620 nm, respectively. Secondary antibodies tagged with Alexa dye were also imaged to confirm and set the background immunoreactivity. Images were recorded with NIS‐Elements software (Nikon). The densities of TRPM4 and IP₃R2 proteins, and their co‐localization, were evaluated as the number of pixels presenting a specified range of fluorescence intensity per area of subcellular region. The number of pixels on the captured image with a designated colour (green, red and yellow) was quantified using the ‘Color Pixel Counter’ plugin in ImageJ (NIH, Bethesda, MD, USA).

Proximity ligation assay (PLA)

Cells were incubated with primary antibodies of TRPM4 and IP₃R2 using the same protocol described above. After washing in PBS, the atrial myocytes were incubated with secondary antibodies linked to PLA probes (Duolink In Situ PLA Probe Anti‐Goat PLUS and Anti‐Rabbit MUNUS; Olink Bioscience, Uppsala, Sweden) for 1 h at 37°C. After washing in Duolink Wash Buffer A, cells were incubated with Ligation‐Ligase solution for 30 min at 37°C and then with Amplification‐Polymerase solution (Duolink In Situ Detection Reagents Red) for 100 min at 37°C. After washing in Duolink Wash Buffer B, myocytes were mounted with 90% glycerol (phosphate buffered) containing 2.5% 1,4‐diazabicyclo‐[2.2.2]‐octane (DABCO, anti‐fade reagents; Sigma) and Syto‐11, nucleic acid staining dye (Molecular Probes). The cells were visualized using confocal microscopy (A1; Nikon) and images were recorded with the NIS‐elements software (Nikon). Bound PLA probes and Syto‐11 were excited at 561 nm and 488 nm, respectively, and their emissions were measured at 570–620 nm and 500–550 nm, respectively.

RT‐PCR analysis

Total RNA was extracted from rat atrial myocytes using the TRIzol reagent (Invitrogen). cDNA was synthesized from total RNA using PrimeScript reverse transcriptase (Takara Bio Inc., Otsu, Japan) in accordance with the manufacturer's instructions, with 100 ng of oligo‐dT as a primer (19 bp; Bioneer Inc.). The RNA samples were denatured at 65°C for 5 min and reverse transcription was performed at 42°C for 2 h. PCRs for specific TRP isoforms were performed using primers that were designed based on the published sequences of rat TRP mRNAs (GenBank). Oligonucleotide primer pairs specific for the respective TRP isoforms were: TRPM4 sense GATTGTCATCGTGAGCAAGA, TRPM4 anti‐sense GTCATACTCTCTGATCTGTC, transient receptor potential cation channel 1 (TRPC1) sense GGACTGACACAGCTCTATG, TRPC1 anti‐sense GAAAGGTATGCAAATACAGTC, transient receptor potential cation channel subfamily V member 4 (TRPV4) sense GCGCTAAGTACCCCGTGG and TRPV4 anti‐sense GCCCCTACAGTGGTGCGT. The general PCR conditions were 2 μl of the reverse transcription products, each corresponding to 200 ng of initial total RNA, 0.2 mm dNTP, primeSTAR buffer and PrimeSTAR HS‐DNA polymerase (Takara Bio Inc.), and each primer in a total volume of 25 μl. The cDNA samples were initially denatured at 98°C for 3 min followed by 25–30 cycles of 98°C for 10 s, 55–58°C for 5–10 s and 72°C for 1 min, and final extension for 10 min. The amplified products (10 μl each) were separated by electrophoresis on 1% agarose gels and stained with ethidium bromide.

Western blotting and quantification

Cells were solubilized in SDS lysis buffer containing 10 mm Tris‐HCl (pH 7.4), 1% (w/v) SDS, 1 mm phenylmethanesulfonyl fluoride, 1 mm Na₃VO₄ and complete protease inhibitor mixture (Roche) for 30 min at 60°C, and then triturated several times by passing through a 1 ml syringe and centrifuged at 12,000 g for 10 min. The supernatant was combined with 2 × Laemmli sample buffer (Bio‐Rad, Hercules, CA, USA) and heated for 30 min at 60°C. Protein samples (∼30 μg) were separated by SDS‐PAGE. Nitrocellulose membranes were probed with primary and secondary antibodies (anti‐TRPM4 Ab, dilution 1:500, Alomone Labs; anti‐α‐actinin Ab, dilution 1:1,000, Santa Cruz Biotechnology Inc.; rabbit polyclonal Ab, dilution 1:1,000, Santa Cruz Biotechnology Inc.) and were detected using a standard western blot protocol. All blots were imaged and quantified using a ChemiDoc XRS densitometer (Bio‐Rad).

Statistical analysis

The numerical results are reported as the mean ± SEM, where n indicates the number of cells used. Paired or unpaired Student's t tests were used for statistical comparisons depending on the experiments. P < 0.05 was considered statistically significant.

Results

I _shear in atrial myocytes and its dependence on intracellular Ca²⁺

Figure 1 A shows whole‐cell membrane currents elicited by voltage ramp pulses from −120 to +70 mV (dV/dt = −250 mV s⁻¹, V _h = −30 mV) at 0.1 Hz in a rat atrial myocyte dialysed with minimal concentrations of Ca²⁺ buffer (0.5 mm EGTA) in symmetrical CsCl solutions. Voltage‐dependent ion channels and other routes of Na⁺ and Ca²⁺ flux were also inhibited (see Methods). Application of ∼16 dyn cm⁻² shear stress increased the membrane current, with no change in the reversal potential (E _rev) of the ramp current: ‘(2)’ in Fig. 1 A. The difference in current, obtained by subtracting the control current from the current after applying shear, was considered to represent I _shear: ‘I _shear, (2) −( 1)’ in Fig. 1 A. The current–voltage (I–V) relationship for I _shear showed outward rectification and slight inward rectification at very negative voltages, with an E _rev of −2.1 ± 0.16 mV (n = 62). The average values of I _shear measured at −100 mV and +60 mV under these conditions were −4.1 ± 0.35 pA pF^–1 and 8.2 ± 0.45 pA pF^–1, respectively (n = 62). I _shear reached a maximum within 20–30 s of exposure to shear stress, and was maintained without significant decay for an additional ∼4 min of exposure (Fig. 1 B).

Figure 1. Activation of shear‐sensitive whole‐cell current and its dependence on intracellular Ca²⁺ in rat atrial myocytes .

A, superimposed ramp currents during voltage pulses from −120 to +70 mV (holding potential = −30 mV) in a representative rat atrial myocyte before, ‘(1)’, and after, ‘(2)’, the application of shear stress of ∼16 dyn cm⁻². Difference current, ‘(2) − (1)’, represents the I–V relationship for shear stress‐sensitive current (I _shear). Inset: continuous recording of ramp currents in the same cell under shear. Symmetrical CsCl solutions containing 0.5 mm internal EGTA and zero external Ca²⁺ were used as a control (see Methods). B, time course of averaged whole‐cell currents, measured at −100 mV and +60 mV during the exposure to shear stress (n = 7). C, superimposed I _shear–V relationships measured at different concentrations of internal EGTA (0.5, 2, 4 and 15 mm). The I–V curves from seven cells under each EGTA concentration were averaged. D, summary of mean magnitudes of I _shear measured at −100 mV and +60 mV in different concentrations of internal EGTA. A shear stress of 16 dyn cm⁻¹ was applied. The numbers of cells included were 62, 10, 7 and 11 for 0.5, 2, 4 and 15 mm internal EGTA, respectively. *P < 0.05, **P < 0.01, ****P < 0.0001 vs. 0.5 mm EGTA (unpaired Student's t test).

A similar shear force was previously shown to induce a longitudinally propagating global Ca²⁺ wave in rat atrial myocytes (Woo et al. 2007). To determine whether this increase in intracellular Ca²⁺ plays a role in the activation of I _shear, we examined the effects of dialysis of other concentrations of EGTA into the myocytes. Figure 1 C shows the signal‐averaged ramp I _shear at different concentrations of EGTA in the pipette solutions. In cells dialysed with 2 mm EGTA, inward I _shear was slightly but not significantly lower than that at 0.5 mm internal EGTA (control), whereas outward I _shear was reduced to ∼75% of the control current (Fig. 1 C and D). When 4 mm EGTA was dialysed into the cells, both inward and outward I _shear were reduced to 40−50% of control values (Fig. 1 C and D). I _shear was more strongly inhibited by dialysis with 15 mm EGTA (to 10−25% of the I _shear at 0.5 mM EGTA), with a larger reduction in inward I _shear (Fig. 1 C and D). Introduction of different concentrations of EGTA did not affect the E_rev of I _shear [−3.7 ± 0.17 mV (n = 8), −1.8 ± 0.77 mV (n = 7) and −3.5 ± 0.74 mV (n = 7) at 2, 4 and 15 mm EGTA, respectively]. The addition of EGTA (15 mm) did not alter E _rev (−2.1 ± 0.3 mV, n = 11) but increased magnitude of outward currents (Table 1). These results indicate that the activation of I _shear is dependent on cytosolic Ca²⁺ concentrations. Based on this finding, we used the minimal Ca²⁺‐buffering condition (0.5 mm EGTA) for all subsequent experiments.

Table 1.

Summary of the magnitudes of ramp currents measured under different interventions without shear stress in rat atrial myocytes

	Whole‐cell membrane current (pA pF–1)
	Control/WT (symmetrical CsCl,		Intervention (inhibitor, Ab or
Intervention	0.5 mm EGTAi, 0 mm Cao)		siRNA, etc.)		n
(rat)	−100 mV	+60 mV	−100 mV	+60 mV
15 mm EGTAi	−3.2 ± 0.3	7.3 ± 0.35	−4.8 ± 0.86	30 ± 1.1b	62/11
137 mm NMDGo	−3.2 ± 0.3	7.3 ± 0.35	−2.2 ± 0.64a	14 ± 2.0c	62/5
137 mm NMDGi	−3.2 ± 0.3	7.3 ± 0.35	−4.0 ± 0.32	1.5 ± 0.20b	62/5
93 mm Cao	−3.2 ± 0.3	7.3 ± 0.35	−3.9 ± 0.28	7.6 ± 1.1	62/4
9 mm Cl− o	−3.2 ± 0.3	7.3 ± 0.35	−5.0 ± 0.85	7.2 ± 0.80	62/5
BAPTAi & Cai	−3.2 ± 0.3	7.3 ± 0.35	−3.9 ± 0.88	11 ± 1.7c	62/8
Tamoxifen (10 μm)	−4.7 ± 0.66	8.6 ± 0.61	−4.9 ± 0.31	8.6 ± 0.81	7
Tamoxifen (10 μm) + 9‐AC (1 mm)	−4.3 ± 0.54	8.4 ± 0.54	−4.1 ± 0.57	8.5 ± 0.56	7
9‐PT (10 μm)	−3.2 ± 0.45	8.6 ± 0.44	−3.1 ± 0.47	8.4 ± 0.51	12
9‐PT (100 μm)	−3.2 ± 0.45	8.6 ± 0.44	−3.0 ± 0.52	8.4 ± 0.56	12
Fluf (10 μm)	−4.0 ± 0.37	9.0 ± 1.2	−4.1 ± 0.16	8.9 ± 1.3	5
GsMTx‐4 (3 μm)	−3.6 ± 0.75	8.7 ± 0.61	−3.6 ± 0.74	8.4 ± 0.48	5
KB‐R7943 (5 μm)	−3.8 ± 0.53	8.7 ± 0.37	−3.6 ± 0.50	8.7 ± 0.80	4
TRPM4 Ab	−3.3 ± 0.46	7.5 ± 0.67	−3.2 ± 0.57	7.3 ± 0.77	10/6
Ryanodine (20 μm)	−3.4 ± 0.53	8.7 ± 1.3	−3.1 ± 0.90	8.6 ± 1.3	11
Ryanodine (50 μm)	−4.7 ± 0.87	8.4 ± 0.91	−3.9 ± 0.80	8.4 ± 0.91	13
2‐APB (2 μm)	−5.1 ± 0.75	7.7 ± 0.57	−4.8 ± 0.63	8.0 ± 0.53	14
XeC (3 μm)	−5.1 ± 1.0	6.7 ± 0.91	−5.1 ± 0.89	6.9 ± 0.10	5
CPA (10 μm)	−3.9 ± 0.65	8.7 ± 0.42	−3.8 ± 0.65	9.0 ± 0.49	5
Chelerythrine (2 μm)	−4.3 ± 0.39	9.2 ± 0.54	−3.8 ± 0.14	8.9 ± 0.81	7/6
4‐AP (200 μm)	−3.8 ± 0.83	6.3 ± 0.56	−4.1 ± 0.39	7.6 ± 0.50	5
E‐4031 (5 μm)	−5.1 ± 0.73	8.8 ± 1.0	−5.2 ± 0.71	8.6 ± 0.95	7
(HL‐1) TRPM4 KD	−3.6 ± 1.0	5.7 ± 0.84	−5.5 ± 1.0	6.7 ± 0.76	9/8
(mouse) IP3R2 KO	−4.7 ± 1.1	8.0 ± 0.9	−5.1 ± 0.78	8.4 ± 1.2	7/7

Data are presented as the mean ± SEM. n, number of cells. ‘TRPM4 Ab’: 400 ng ml⁻¹ (12 min dialysis). ‘BAPTA_i & Ca_i’:15 min dialysis with 10 mm BAPTA combined with Ca²⁺ to make 100 nm free [Ca²⁺]_i.^a P < 0.05 vs. control, −100 mV.^b P < 0.01 vs. control, +60 mV. ^c P < 0.05 vs. control, +60 mV. (unpaired t test).

I _shear is mainly carried by monovalent cations

We next examined whether the I _shear detected in the symmetrical CsCl solutions contained only cation currents. Replacing external Cs⁺ with non‐permeant N‐methyl‐d‐glutamate (NMDG⁺) almost completely abolished the inward component of I _shear, with no significant effect on outward I _shear (Fig. 2 A and D). Under this condition, the control outward current was increased and the control inward current was reduced (NMDG_o) (Table 1) with a negative shift of E _rev (−50 ± 1.9 mV, n = 5). The E _rev of I _shear shifted to a more negative voltage under this condition (−48.1 ± 2.04 mV, n = 5, P < 0.0001 vs. Cs⁺ external) (Fig. 2 A). When internal Cs⁺ was completely replaced by NMDG⁺, ∼80% of outward I _shear was eliminated, with a rightward shift in E _rev (+17 ± 2.4 mV, n = 8) (Fig. 2 B and D). Note that the magnitude of control outward current was reduced (NMDG_i) (Table 1) with right shift of E _rev (+21 ± 4.8 mV, n = 5). These results suggest that I _shear mainly conducts monovalent cations.

Figure 2. I_shear mainly carries monovalent cations .

A, signal‐averaged I–V relationships for I _shear recorded in the presence of 137 mm NMDG·Cl (red, ‘NMDG_o’; n = 5) and 137 mm CsCl (black, ‘Cs_o’; n = 7) in the external solutions. B, signal averaged I–V relationships for I _shear measured in 137 mm NMDG·Cl (red, ‘NMDG_i’; n = 5) and 137 mm CsCl (black, ‘Cs_i’; n = 7) in the internal solutions. C, signal‐averaged I–V relationships for I _shear recorded in 137 mm CsCl (black, ‘Cs_o’: n = 7), 93 mm CaCl₂ (blue, ‘Ca_o’; n = 4) and 137 mm CsCl with 2 mm Ca²⁺ [grey, ‘(Cs+2Ca)_o’; n = 4] in the external solutions. D, summary of the magnitudes of I _shear measured at −100 mV and +60 mV under different ionic compositions of internal or external solutions [‘Cs_o, Cs_i’, n = 62; ‘(Cs+2Ca)_o’, n = 4; ‘NMDG_o’, n = 5; ‘Ca_o’, n = 4; ‘NMDG_i’, n = 5]. **P < 0.01, ***P < 0.001 vs. ‘Cs_o, Cs_i’ (unpaired Student's t test). E, left: superimposed I–V curves for I _shear detected in a representative rat atrial myocyte in the absence and presence of tamoxifen (Tamox, 10 μm, 3 min), the inhibitor of volume‐regulated Cl⁻ channel and Ca²⁺‐activated Cl⁻ channel, or tamoxifen plus 9‐AC (1 mm, 1 min 30 s), a non‐specific Cl⁻ channel inhibitor. Symmetrical CsCl solutions with 0.5 mm internal EGTA were used (for more details, see Methods). Right: representative I–V curve for I _shear recorded in the presence of 9 mm Cl⁻ and 130 mm aspartic acid instead of 137 mm Cl⁻. F, mean density of I _shear at ‒100 mV and +60 mV under control conditions (: symmetrical CsCl solutions), in the presence of tamoxifen with and without 9‐AC (n = 7, *P < 0.05 vs. Control, paired t test), and in low Cl⁻‐containing external solutions (‘Low Cl⁻ _o’, n = 5, *P < 0.05 vs. Control, unpaired t test). A shear stress of 16 dyn cm⁻¹ was applied. The data show that a small portion of outward I _shear is carried by Cl⁻.

To determine the permeability of the shear‐sensitive channel for Ca²⁺, external CsCl was completely replaced with equiosmolar (93 mm) CaCl₂. Under this condition, a similar outward I _shear was detected, whereas the inward I _shear decreased considerably to ∼15% of the control: ‘Ca_o’ in Fig. 2 C and D. In addition, E _rev was estimated to be −31.4 ± 1.42 mV (n = 4), suggesting that the contribution of Ca²⁺ to I _shear is almost negligible. The magnitude of control ramp currents was not altered by the replacement of external Cs⁺ by Ca²⁺ (Ca_o) (Table 1), although E _rev of the current was more negative (−24 ± 2.7 mV, n = 4) compared to that in Ca²⁺‐free external solution. It should also be noted that the magnitude and I–V relationship of I _shear were not significantly altered by the addition of 2 mm Ca²⁺ to the external solutions: ‘(Cs+2Ca)_o’ in Fig. 2 C and D.

Next, we examined the contribution of Cl⁻ to I _shear using Cl⁻ channel blockers. Because I _shear was well maintained under the existing experimental conditions (Fig. 1 B), we assessed I _shear with and without intervention in the same myocytes. After pretreatment with tamoxifen (10 μm), which blocks volume‐regulated Cl⁻ channels and Ca²⁺‐activated Cl⁻ channels, outward I _shear was reduced by ∼25% with no significant change in inward I _shear (Fig. 2 E and F). The E _rev of I _shear shifted significantly from −2.9 ± 0.37 mV to −1.1 ± 0.29 mV upon exposure to tamoxifen (P < 0.05, n = 5) (Fig. 2 E). Additional application of 9‐anthracenecarboxylic acid (9‐AC, 1 mm), which inhibits most classes of Cl⁻ channels, did not alter I _shear further (Fig. 2 E and F). The application of tamoxifen without and with 9‐AC did not affect E _rev (control, −3.6 ± 0.43, tamoxifen, −3.6 ± 0.41; tamoxifen + 9‐AC, −3.0 ± 0.42 mV, n = 7) and the magnitude of the control ramp current (Table 1). To confirm the contribution of Cl⁻ current to the I _shear, 130 mm extracellular Cl⁻ or 137 mm intracellular Cl⁻ were replaced by equimolar aspartic acid. In low Cl⁻ external solutions, outward I _shear was decreased by ∼20% (Fig. 2 E and F) with more positive E _rev values (2.1 ± 0.8 mV, n = 5, P < 0.05). The I _shear was not changed by internal Cl⁻ replacement (data not shown). The control ramp current was not changed by the external Cl⁻ replacement (low Cl⁻ _o; E _rev: 4.4 ± 0.91 mV, n = 5, P > 0.05) (Table 1). Taken together, these results indicate that the channel activated by shear stress primarily conducts monovalent cations, but not Ca²⁺, with 20−25% of outward I _shear attributable to Cl⁻.

Suppression of I _shear by inhibition of TRPM4 using 9‐phenanthrol or flufenamic acid

With its dependence on intracellular Ca²⁺ and selectivity to monovalent cations, I _shear resembles the Ca²⁺‐activated non‐selective cation currents previously reported in cardiac myocytes (Hill et al. 1988; Guinamard et al. 2002; Guinamard et al. 2004; Demion et al. 2007). The molecular identity of the Ca²⁺‐activated non‐selective cation channel is assumed to be the TRPM4 channel because of its similar pharmacological and biophysical properties (Guinamard et al. 2004; Demion et al. 2007). The significantly larger outward I _shear than inward current is also similar to the voltage‐dependence of the whole‐cell current detected in TRPM4‐overexpressing HEK293 cells (Launay et al. 2002; Nilius et al. 2003). We therefore examined the possible role of TRPM4 in I _shear in subsequent experiments. Application of 9‐hydroxyphenanthrene (9‐phenanthrol), a TRPM4 inhibitor (Grand et al., 2008), significantly suppressed I _shear (Fig. 3 A), reaching a plateau 20–60 s after the onset of exposure. The effect was reversed by withdrawal of the chemical. The inward component of I _shear decreased by ∼60% and 95%, and outward I _shear decreased by ∼50% and 75%, in the presence of 10 and 100 μm 9‐phenanthrol, respectively (Fig. 3 A and C), although E _rev was not altered at either concentration (control, −3.6 ± 0.34 mV; 10 μm, −3.4 ± 0.32 mV; 100 μm, −3.5 ± 0.27 mV, n = 12, P > 0.05). Another TRPM4 inhibitor, flufenamic acid (10 μm) (Gogelein et al. 1990; Demion et al. 2007), suppressed inward and outward I _shear by ∼95% and 75%, respectively (Fig. 3 B and C), with no effect on E _rev (control, −3.3 ± 0.51 mV vs. flufenamic acid, −2.4 ± 0.44 mV, n = 5, P > 0.05). These results suggest that the TRPM4 current constitutes most I _shear. The application of 9‐phenanthrol or flufenamic acid did not affect E _rev (control, −3.3 ± 0.31 mV vs. 10 μm 9‐phenanthrol, −3.4 ± 0.28 mV, P > 0.05, 100 μm 9‐phenanthrol, −3.4 ± 0.36 mV, P > 0.05, n = 12; control, −4.1 ± 0.48 mV vs. flufenamic acid, 3.7 ± 0.37 mV, n = 5, P > 0.05) and the magnitude of the control ramp current (Table 1).

Figure 3. Inhibition of I_shear by the blockers of TRPM4 channel .

A and B, superimposed I _shear–V curves recorded in the absence (‘C’) and presence of 10 or 100 μm 9‐phenanthrol (9‐PT; A) or 10 μm flufenamic acid (Fluf; B), the inhibitors of TRPM4, in the representative rat atrial myocytes. C, summary of the magnitudes of I _shear measured at −100 mV and +60 mV before and after the application of 10 or 100 μm 9‐PT (1 min; n = 12), or 10 μm Fluf (3 min; n = 5). **P < 0.01, ***P < 0.001 vs. control (‘C’) (paired t test). D and E, superimposed I _shear–V curves recorded in the absence (‘C’) and presence of the inhibitor of stretch‐activated ion channel GsMTx‐4 (3 μm, 1 min; D) or the inhibitor of NCX KB‐R7943 (5 μm, 5 min; E) in the representative rat atrial myocytes. F, summary of the magnitudes of I _shear measured at −100 mV and +60 mV before (‘C’) and after the application of 3 μm GsMTx‐4 (n = 6) or 5 μm KB‐R7943 (n = 4). G and H, superimposed I _shear–V curves recorded in the absence (‘C’) and presence of the inhibitor of Kv1.5 channel 4‐AP (200 μm, 3 min 30 s; G) or the inhibitor of ERG channel E‐4031 (5 μm, 3 min; H) in the representative rat atrial myocytes. I, summary of the magnitudes of I _shear measured at −100 mV and +60 mV before (‘C’) and after the application of 200 μm 4‐AP (n = 5) or 5 μM E‐4031 (n = 7). A shear stress of 16 dyn cm⁻¹ was applied.

It was also confirmed that the magnitude and E _rev of I _shear induced by shear were not altered by the SAC blocker GsMTx‐4 (3 μm; Bae et al. 2011) (E _rev in mV: control, −2.5 ± 0.37 vs. GsMTx‐4, −2.3 ± 0.35, n = 7, P > 0.05) (Fig. 3 D and F). We consistently detected no change in cell length at shear stresses of 3–30 dyn cm⁻² by cell imaging (data now shown). We also investigated whether Ca²⁺‐dependent activation of the Na⁺−Ca²⁺ exchanger (NCX) contributes to I _shear under the same experimental conditions, using its blocker, KB‐R7943 (Woo & Morad, 2001). We observed no effect of this chemical on I _shear (E _rev in mV: control, −2.7 ± 0.28 vs. 5 μm KB‐R7943, −2.4 ± 0.34, n = 5, P > 0.05) (Fig. 3 E and F), indicating that NCX current plays no role in I _shear. Note that the application of 3 μm GsMTx‐4 or 5 μm KB‐R7943 did not alter E _rev (control, −4.5 ± 0.59 mV vs. GsMTx‐4, −4.7 ± 0.63 mV, n = 5, P > 0.05; control, −3.2 ± 0.27 mV vs. KB‐R7943, 3.1 ± 0.18 mV, n = 4, P > 0.05) and the magnitude of the control ramp current (Table 1).

Kv1.5 and mouse ether‐à‐go‐go‐related gene (mERG) channels have significant Cs⁺ permeability relative to K⁺ (0.1−0.3), which may be further augmented in the absence of Ca²⁺. We tested whether these channels contribute to the measured I _shear using 200 μm 4‐aminopyridine (4‐AP) (Tamargo et al. 2004) and 5 μm E‐4031 (Tamargo et al. 2004) to block Kv1.5 and mERG, respectively. The magnitude of I _shear was similar in the absence and presence of 4‐AP (Fig. 3 G and I), and E _rev of I _shear was not significantly affected by this drug (control, −2.2 ± 0.43 mV vs. 4‐AP, −2.3 ± 0.42 mV, n = 5, P > 0.05). Control ramp currents were not affected by 4‐AP (E _rev: control, −1.8 ± 0.93 mV vs. 4‐AP, −2.4 ± 0.53 mV, n = 5, P > 0.05) (Table 1). Treatment of E‐4031 did not significantly alter control currents (E _rev: control, −1.6 ± 0.54 mV vs. E‐4031, −1.7 ± 0.56 mV, n = 7, P > 0.05) (Table 1) and I _shear (E _rev: control, −2.8 ± 0.91 vs. E‐4031, −2.7 ± 0.61 mV, n = 7, P > 0.05) (Fig. 3 H and I). These results suggest that I _shear is not mediated by Kv1.5 or mERG channels.

Inhibition of I _shear by intracellular application of TRPM4 antibody

To further confirm the putative role of the TRPM4 channel in I _shear, we examined whether I _shear was affected by the introduction of anti‐TRPM4 antibody into the myocytes. Atrial myocytes were dialysed with the same internal solutions described above, with the addition of 400 ng ml⁻¹ TRPM4 antibody. The same measurements were also taken in cells dialysed with TRPM4‐blocking peptides (400 ng ml⁻¹), in addition to the anti‐TRPM4 antibody, aiming to determine whether there was any non‐specific effect of the dialysed antibody. The inward and outward components of I _shear in cells dialysed for 3 min with the antibody were not significantly different from those measured in the presence of the additional blocking peptides (Fig. 4). A prolonged (≥8 min) dialysis with the antibody resulted in notable reductions of inward and outward I _shear, followed by a plateau, although a prolonged dialysis with the blocking peptides together with the antibody attenuated this decrease in I _shear (Fig. 4). Inward I _shear was almost completely inhibited by the antibody, and outward I _shear in the presence of antibody was only 20−25% of that with antibody plus peptide (Fig. 4 C). E _rev (mV) of I _shear was not altered by either TRPM4 antibody (3 min, −4.3 ± 0.53 vs. 12 min, −2.3 ± 0.32, n = 10, P > 0.05) or antibody plus peptide (3 min, −3.3 ± 0.46 vs. 12 min, −2.1 ± 0.29, n = 10, P > 0.05). Intracellular dialysis with anti‐TRPM4 antibodies or the antibodies plus TRPM4‐blocking peptides did not significantly change E _rev (at 12 min after dialysis: antibodies, −2.2 ± 0.45 mV, n = 6; antibodies plus peptides, −2.8 ± 0.77 mV, n = 10, P > 0.05) and the magnitude of control ramp currents (Table 1). This result is consistent with the effects of TRPM4 inhibitors on I _shear and suggests that a major component of I _shear may be TRPM4 current.

Figure 4. Inhibition of I_shear by intracellular dialysis with anti‐TRPM4 antibody .

A, representative I–V relationships for I _shear measured 3 (left) and 12 min (right) after dialysis with anti‐TRPM4 antibody with blocking peptide (‘Pep+Ab’). B, representative I–V relationships for I _shear measured 3 (left) and 12 min (right) after dialysis with anti‐TRPM4 antibody (‘Ab’). C, plot of mean I _shear measured at −100 mV and +60 mV vs. the duration of dialysis with TRPM4 antibodies (●; n = 10) or the antibodies with the peptides (□; n = 6). I _shear was recorded repeatedly in the same cells at different time points (3, 8, 10, 12 and 14 min) after membrane rupture. The I _shear values, measured at different time points in ten antibody‐dialysed cells and in six cells dialysed with the antibodies plus peptides, were averaged at each time point. A shear stress of 16 dyn cm⁻¹ was applied. **P < 0.01, ***P < 0.001 vs. ‘Pep+Ab’ (unpaired t test).

Removal of I _shear by KD of TRPM4 in HL‐1 atrial myocytes

To confirm the role of TRPM4 in carrying the cation component of I _shear, we selectively inhibited TRPM4 expression using siRNA. Owing to the limitations involved in delivering siRNA into intact atrial myocytes, we used the HL‐1 adult mouse atrial cell line for this experiment. HL‐1 cells express functional RyRs and IP₃R subtypes similar to those in intact atrial cells (Kim et al. 2010 a,b). Treating the HL‐1 cells with TRPM4‐specific siRNA for 6 h followed by a 2 day culture suppressed the expression of TRPM4 by 97 ± 7.8% (five batches) (Fig. 5 A). The I _shear recorded in cells transfected with control siRNA (WT) showed an I–V relationship similar to that measured in intact cells but with a smaller (50−60%) density than that in intact atrial cells (compare WT in Fig. 5 C with 0.5 mm [EGTA]_i in Fig. 1 C). 9‐Phenanthrol (100 μm) suppressed I _shear by ∼70%, with a greater decrease in inward component observed in WT cells (Fig. 5 B and D), which is consistent with its effect on I _shear in intact rat atrial myocytes (Fig. 3 A and C). In TRPM4‐KD cells, inward and outward I _shear were only ∼10% and 30% of the WT current, respectively (Fig. 5 C and D). The magnitude and E _rev of control ramp currents measured in WT and TRPM4 KD HL‐1 cells were not significantly different (E _rev: WT, –1.7 ± 0.48 mV, n = 9; KD, –1.7 ± 0.35 mV, n = 8) (Table 1) and they were no different from those measured from rat atrial cells (Table 1). This result clearly demonstrates a major role of TRPM4 channels in I _shear and a minor contribution of TRPM4‐independent current (mainly to the outward current).

Figure 5. Inhibition of I_shear by KD of TRPM4 .

A, representative immunoblots showing expression level of TRPM4 in HL‐1 cells transfected with TRPM4‐specific siRNA after 36 or 48 h of culture. α‐actinin was used as a loading control. WT, WT HL‐1 cells transfected with non‐targeting siRNA. B, I _shear detected in a representative WT HL‐1 cell in the absence and presence of 100 μm 9‐PT. C, signal‐averaged I _shear–V curves measured from WT (n = 8)‐ and TRPM4 KD HL‐1 cells (n = 9). D, summary of the magnitudes of I _shear measured at −100 mV and +60 mV in WT HL‐1 cells without (‘C’) and with 9‐PT (n = 8), and in TRPM4 KD HL‐1 cells (n = 9). **P < 0.01 vs. ‘WT, C’ (unpaired t test). A shear stress of 16 dyn cm⁻¹ was applied. Symmetrical CsCl solutions with 0.5 mm internal EGTA were used.

Possible role of IP₃R‐mediated Ca²⁺ release in the activation of I _shear

In the next series of experiments, we examined the source of the intracellular Ca²⁺ that triggers I _shear. First, the possible contribution of Ca²⁺ release from the SR via the RyRs (i.e. the major Ca²⁺ release channel in cardiac myocytes) to I _shear was tested using relatively high concentrations of the RyR blocker ryanodine. RyR inhibition suppressed inward and outward I _shear by ∼55% and 45%, respectively, when using 20 μm ryanodine as the inhibitor, and by ∼90% and 65%, respectively, when using 50 μm ryanodine (Fig. 6 A and B). Application of ryanodine did not alter the E _rev of I _shear (control, −1.9 ± 0.44 mV vs. 20 μm ryanodine, −1.6 ± 0.28 mV, n = 11, P > 0.05; control, −2.6 ± 0.51 mV vs. 50 μm ryanodine, −2.3 ± 0.38 mV, n = 13, P > 0.05). Both concentrations of ryanodine did not alter the magnitude and E _rev of control ramp currents (E _rev: control, −3.9 ± 0.30 mV vs. 20 μm ryanodine, −3.5 ± 0.26 mV, n = 11, P > 0.05; control, −4.8 ± 0.93 mV vs. 50 μm ryanodine, −3.8 ± 0.68 mV, n = 13, P > 0.05) (Table 1).

Figure 6. Dependence of I_shear on intracellular Ca²⁺ release through the RyRs and IP₃Rs .

A, superimposed I _shear–V curves measured in the absence (‘C’) and presence of 20 (left) or 50 μm (right) ryanodine (Ry; 4 min) in the representative rat atrial myocytes. B, summary of the average magnitudes of I _shear measured at −100 mV and +60 mV under control conditions and after the application of 20 (n = 11) or 50 μm (n = 13) ryanodine. C, superimposed I _shear–V curves measured in the control condition (‘C’) and after the application of 2‐APB (2 μm, 3 min; left) or XeC (3 μm, 3 min; right). D, summary of the average magnitudes of I _shear measured at −100 mV and +60 mV under control conditions and after the application of 2‐APB (n = 14), XeC (n = 5). E, superimposed I _shear–V relationships measured before and after the application of 10 μm CPA (5 min; left). Right, summary of the average magnitudes of I _shear measured at −100 mV and +60 mV under control conditions and after the application of CPA (n = 5). Symmetrical CsCl solutions with 0.5 mm internal EGTA were used. A shear stress of 16 dyn cm⁻¹ was applied. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 vs. control (‘Con’) (paired t test). ^# P < 0.05 vs. ‘20 μm Ry’ (unpaired t test).

Atrial myocytes also express high densities of IP₃Rs (Moschella & Marks, 1993; Lipp et al. 2000; Kim et al. 2010 b). IP₃R2 is known to co‐localize with RyR in the peripheral junctional SR in atrial myocytes, and thus Ca²⁺ release through the IP₃R2 may facilitate CICR via the RyR (Lipp et al. 2000; Mackenzie et al. 2002). We further examined whether the IP₃R serves as a Ca²⁺ release mechanism to regulate I _shear using the IP₃R inhibitors. 2‐APB has been successfully used for selective inhibition of IP₃Rs in cardiac cells at concentrations of 2–5 μm (Mackenzie et al. 2002; Li et al. 2005). Inhibition of IP₃R using 2 μm 2‐APB decreased inward and outward I _shear by ∼85% and 70%, respectively (Fig. 6 C and D), with no shift in E _rev (control, −2.1 ± 0.43 mV vs. 2‐APB, −2.2 ± 0.28 mV, n = 14, P > 0.05). Another IP₃R blocker, xestospongin C (XeC; 3 μm), similarly suppressed inward (by ∼80%) and outward I _shear (by ∼70%) (Fig. 6 C and D), with no change in E _rev (control, −3.0 ± 0.62 mV vs. XeC, −2.6 ± 0.16 mV, n = 5, P > 0.05). 2‐APB and XeC did not alter the magnitude and E _rev of control ramp currents (E _rev: control, −4.3 ± 1.0 mV vs. 2‐APB, −4.2 ± 0.99 mV, n = 14, P > 0.05; control, −4.2 ± 0.70 mV vs. XeC, −3.2 ± 0.54 mV, n = 5, P > 0.05) (Table 1). Consistent with these results, depletion of SR Ca²⁺ by blocking the SR Ca²⁺ pump using cyclopiazonic acid (CPA, 10 μm) caused an ∼90% decrease in inward I _shear and an ∼70% decrease in outward I _shear (Fig. 6 E). CPA did not affect the magnitude and E_rev of control ramp currents (E _rev: control, −4.1 ± 0.47 mV vs. CPA, −4.1 ± 0.52 mV, n = 5, P > 0.05) (Table 1). The percentage of I _shear dependent on Ca²⁺ release closely matched the portion of I _shear that was eliminated by the inhibition or KD of TRPM4 (Figs 3 C, 4 C and 5 D). Together, these results suggest that Ca²⁺ release from the SR, triggered by IP₃R activity, may be a key activation mechanism for the Ca²⁺‐dependent cation (TRPM4) current in I _shear during shear stress.

To confirm that Ca²⁺ release is important for the channel activation, we tested the effect of intracellular Ca²⁺ buffering at a fixed physiological level using Ca²⁺‐BAPTA mixture because increased EGTA in the pipette (Fig. 1 C and D) lowers the basal Ca²⁺ with buffering Ca²⁺ release. I _shear was monitored in cells dialysed with internal solutions containing BAPTA together with the appropriate amount of Ca²⁺ to obtain ∼100 nm Ca²⁺ at different dialysing durations (3, 6 and 15 min) (Fig. 7). At 3 min of dialysis, inward I _shear, but not outward I _shear, was significantly smaller than that measured under control conditions (0.5 mm internal EGTA) (Fig. 7). The currents were gradually decreased by dialysis, reaching a steady‐state of current decrease (20−25% of control I _shear) at 15 min after membrane rupture (Fig. 7). The remaining outward I _shear after the 15 min dialysis was similar to that measured in the presence of 15 mm internal EGTA, whereas inward I _shear was slightly, but not significantly, larger than the inward I _shear in 15 mm internal EGTA (compare with Fig. 1 C). This result further suggests that Ca²⁺ release plays a role in activating I _shear.

Figure 7. Measurement of I_shear at a fixed physiological Ca²⁺ level via a Ca²⁺‐BAPTA mixture .

A, superimposed I _shear–V curves measured in a representative atrial cell at 3 (open square), 6 (filled square) and 15 min (filled circle) dialysis with Ca²⁺‐BAPTA mixture to obtain 100 nm free Ca²⁺. In the presence of 10 mm BAPTA, 6.4 mm Ca²⁺ was added in the form of Ca(OH)₂ as calculated with the Maxchelator (http://maxchelator.stanford.edu). A shear stress of 16 dyn cm⁻¹ was applied. B, summary of the magnitudes of inward and outward I _shear at −100 mV and +60 mV, respectively, at 3, 6 and 15 min after dialysis with the Ca²⁺ plus BAPTA. **P < 0.01, ***P < 0.001, ****P < 0.0001 vs. Con (control). ^## P < 0.01, ^### P <0.001 vs. 3 min dialysis (unpaired t test, eight cells).

Loss of TRPM4 current in IP₃R2‐KO atrial myocytes under shear stress

Although IP₃R blockers are widely used when studying the role of IP₃Rs, they also have side effects. For example, 2‐APB is known to act on store‐operated channels (Bootman et al. 2002) and TRP channels (Hu et al. 2004; Togashi et al. 2008). We further confirmed the role of IP₃R‐mediated Ca²⁺ release in the activation of I _shear using IP₃R2 KO mice (Li et al. 2005). In rat atrial myocytes, IP₃R2 is the most abundant of the three IP₃R subtypes (Lipp et al. 2000; Kim et al. 2010 b). Figure 8 compares I _shear in atrial myocytes from WT and IP₃R2 KO mice. I _shear was recorded in WT mouse atrial cells, although its density was lower than that in rat atrial myocytes (Figures 8 and 1). We also confirmed that WT atrial I _shear was sensitive to 9‐phenanthrol (Fig. 8). In the IP₃R2 KO atrial myocytes, inward I _shear was almost negligible and a small (10−20%) outward I _shear was detected (Fig. 8). The magnitude and E _rev of control ramp currents measured in WT and IP₃R2 KO atrial myocytes did not show a significant difference (E _rev: WT, –1.7 ± 0.48 mV, n = 7; KO, –1.9 ± 0.35 mV, n = 7) (Table 1) and the values in WT cells were no different from those measured from rat atrial cells (Table 1) These results confirm that the release of Ca²⁺ via IP₃R2 plays a key role in the activation of TRPM4 in atrial myocytes under shear stress.

Figure 8. Role of IP₃R2 in the activation of I_shear .

A, I _shear–V curve detected in a representative WT mouse atrial cell in the absence (‘Control’) and presence of 10 μm 9‐PT (left), and in a IP₃R2 KO atrial myocyte (right). B, summary of the magnitudes of inward and outward I _shear measured at −100 mV and +60 mV, respectively, in WT cells without (‘C’) and with 9‐PT (n = 7), and in IP₃R2 KO mouse atrial cells (n = 7). A shear stress of 16 dyn cm⁻¹ was applied. *P < 0.05 **P < 0.01, ***P < 0.001, ****P < 0.0001 vs. ‘WT, C’. ^# P < 0.05, ^## P < 0.01 vs. ‘WT, 9‐PT’ (unpaired t test).

Co‐localization of IP₃R2 and TRPM4 at the cell periphery

After demonstrating the role of IP₃R2 in the activation of TRPM4 in atrial cells, we aimed to determine the subcellular localizations of these proteins using immunocytochemistry. We first confirmed that TRPM4 was expressed in rat atrial myocytes at both mRNA and protein levels (Fig. 9 A and B). Immunostaining of atrial myocytes with TRPM4‐ and IP₃R2‐specific antibodies revealed that both proteins were strongly expressed around the periphery of the cells (Fig. 9 C). Generally, there was weak fluorescence in the cell interior, with variability in its density and distribution among atrial myocytes. Co‐localization of these two proteins was found mainly at peripheral sites (Fig. 9 C, arrows and Table 2). This result suggests that a Ca²⁺‐mediated interaction between IP₃R2 and TRPM4 may occur in peripheral microdomains of atrial myocytes.

Figure 9. Expression of TRPM4 in rat atrial myocytes and its specific co‐localization with IP₃R2 .

A, representative agarose gel from reverse transcriptase PCR experiments. Lane 1 shows the size marker (SM). Lanes 2, 3 and 4 represent mRNA of TRPC1 (791 bp), TRPV4 (558 bp) and TRPM4 (751 bp), respectively. B, representative immunoblots showing expression level of TRPM4 in isolated rat atrial (‘Atr’) and ventricular (‘Ven’) myocytes (bands close to 130 kDa). α‐actinin was used as a loading control. C, immunofluorescence staining for TRPM4 (green) and IP₃R2 (red). Co‐localization was illustrated at the bottom image (yellow). A dotted circle indicates the nucleus.

Table 2.

Analysis of the extent of pixel co‐localization for TRPM4 and IP₃R2 in rat atrial myocytes

	TRPM4 with	IP3R2 with
	IP3R2 (%)	TRPM4 (%)	n
Peripheral domain	39.7 ± 4.2	32.7 ± 2.8	12
Central domain	0*	0#	12

Data are presented as the mean ± SEM. n, number of cells. *P < 0.05 vs. ‘TRPM4 with IP₃R2’ at peripheral domain. ^# P < 0.05 vs. ‘IP₃R2 with TRPM4’ peripheral domain. Five rats. An unpaired t test was used. The area up to ∼1.5 μm immediately underneath the cell membrane was denoted as the peripheral domain. Central domain represents the interior of cell except the peripheral domain and both ends of the cell.

We also examined whether TRPM4 proteins are still localized in the cell periphery in IP₃R2 KO myocytes because the loss of I _shear in IP₃R2 KO cells could also be interpreted as the effect of IP₃R2 on TRPM4 membrane localization. In both WT and IP₃R2 KO atrial myocytes, we observed peripherally localized TRPM4 (Fig. 10 A, green), which was consistent with our finding of a 9‐phenanthrol‐sensitive, TRPM4‐like current in IP₃R2 KO myocytes similar to that in WT cells (data not shown). IP₃R2 staining was found in the peripheral sites with a co‐localization with TRPM4 only in WT myocytes (Fig. 10 A). Next, we performed PLA to confirm a more definitive physical relationship between TRPM4 and IP₃R2. If the proteins are within 40 nm of each other, the PLA assay allows ligation and a subsequent amplification reaction, which produces a fluorescence‐labelled circular double‐stranded DNA detectable with a fluorescence microscope (Söderberg et al. 2008). For the control experiments, only one primary antibody (i.e. TRPM4 or IP₃R2) was used and no PLA signals (red dots) were generated (Fig. 10 B). When the two antibodies were added, no PLA signals were detected (Fig. 10 B), indicating these two proteins are not positioned within 40 nm.

Figure 10. No role of IP₃R2 in TRPM4 subcellular localization .

A, confocal images for immunostained TRPM4 (green) and IP₃R2 (red) in WT (left) and IP3R2 KO (right) atrial cells, showing peripheral localization of TRPM4 in both WT and KO myocytes. Co‐localization was seen as a yellow colour. B, detection of proximity between TRPM4 and IP₃R2 in rat atrial myocytes. PLA signals are generated when the labelled proteins are within 40 nm of each other. In the negative control experiments (upper images), rat atrial myocytes were labelled with only one primary antibody and thus only unspecific background signals (red) are generated. Lower: PLA of TRPM4 and IP₃R2. Rat atrial myocytes were labelled with antibodies against TRPM4 and IP₃R2, and then were treated with secondary antibodies linked to PLA probes, resulting in no punctate PLA signals. Syto‐11‐staining (nucleus) in green. Each experiment (A and B) was performed three times. Scale bar = 10 μm.

Discussion

Shear stress in atrial myocytes induces longitudinal global Ca²⁺ waves originating from IP₃R2‐mediated Ca²⁺ release with subsequent CICR (Woo et al. 2007; Kim & Woo, 2015). We report evidence suggesting that the Ca²⁺‐activated monovalent cation channel TRPM4 is indirectly activated by shear stress through Ca²⁺ release from the SR in atrial myocytes. Our data demonstrate that Ca²⁺ release through IP₃R2 plays a key role in this Ca²⁺‐dependent activation of TRPM4 by shear. Under our experimental conditions, using symmetrical CsCl solutions with zero external Ca²⁺ and K⁺, and 0.5 mm internal EGTA, the Ca²⁺ release‐dependent TRPM4 current appeared to constitute more than 90% of the inward and 70% of the outward shear‐sensitive current. To our knowledge, the present data provide the first experimental evidence showing that shear stress is a stimulus for TRPM4 activation, and that IP₃R2‐mediated Ca²⁺ signalling is the link between shear stimulus and the TRPM4 response in atrial myocytes. Ca²⁺‐mediated interactions between IP₃R2 and TRPM4 in the junctions of the SR and cell membrane under shear stress may be an important mechanism by which atrial cells measure mechanical stress and translate it to alter their excitability.

Activation of TRPM4 by shear‐triggered Ca²⁺ release from IP₃R2

The Ca²⁺‐activated non‐selective monovalent cation channel has long been recognized in cardiac tissue, such as in cultured rat ventricular myocytes (Guinamard et al. 2002), human atrial myocytes (Guinamard et al. 2004) and sinoatrial node cells (Demion et al. 2007), and the molecular identity of these channels has recently been identified as TRPM4 (Launay et al. 2002; Nilius et al. 2003; Demion et al. 2007). The present study provides evidence suggesting that the molecular entity carrying most I _shear is TRPM4: (1) I _shear is sensitive to internal Ca²⁺ buffering; (2) the shape of the I _shear I–V curve (outward rectification with slight inward rectification at very negative potentials) is similar to that of whole‐cell TRPM4 current in TRPM4‐overexpressing culture cell lines and in smooth muscle cells (Launay et al. 2002; Nilius et al. 2003; Earley et al. 2007); (3) I _shear has negligible permeability to Ca²⁺; (4) I _shear is sensitive to known blockers of TRPM4 (9‐phenanthrol and flufenamic acid) and to TRPM4 antibodies; and (5) I _shear is lost by KD of TRPM4 expression.

The TRPM4 current was previously detected using internal solutions containing high concentrations of Ca²⁺ to activate the current (Guinamard et al. 2002; Nilius et al. 2003; Demion et al. 2007; Earley et al. 2007) or Ca²⁺ mobilizing receptor agonists, such as ATP (Launay et al. 2002). As used in whole‐cell recording to detect the TRPM4 current, ATP is a well‐known activator of phospholipase C via G protein‐coupled purinergic receptors, producing IP₃ and diacylglycerol. Activation of the IP₃R by IP₃ can increase intracellular Ca²⁺, which in turn facilitates the activation of protein kinase C by diacylglycerol. Protein kinase C is known to increase the activity of the TRPM4 channel by increasing its sensitivity to Ca²⁺ (Nilius et al. 2005; Earley et al. 2007). Few reports to date have shown clear evidence of the role of IP₃R‐mediated Ca²⁺ release in TRPM4 channel activity. In this regard, transient inward cation currents mediated by TRPM4 have been recorded under perforated patch clamp at resting membrane potentials (−70 mV) in native cerebral artery myocytes, and these currents are inhibited by the IP₃R blocker XeC (1 μm) but not by ryanodine (50 μm) (Gonzales et al. 2010). Another study reported a Ca²⁺‐activated TRPM4‐like current under negative pressure in cerebral artery myocytes using the cell‐attached patch clamp method. However, this current was resistant to IP₃R blockade and inhibited by RyR blockade (Morita et al. 2007). In HEK293 cells, the TRPM4 current activated by extracellular ATP under whole‐cell patch clamp with low intracellular Ca²⁺‐buffering did not correlate exactly with global Ca²⁺ changes detected simultaneously in the same cells (Launay et al. 2002). These previous studies report somewhat conflicting evidence regarding the role of IP₃R but, nevertheless, give an insight into the importance of local Ca²⁺ changes and/or smaller Ca²⁺ fluctuations in the vicinity of the cell membrane in TRPM4 channel function. In the present study, we provide clear evidence in atrial myocytes that SR Ca²⁺ release and IP₃R2 play a key role in the activation of the TRPM4 current using Ca²⁺ release inhibitors and IP₃R2 KO mice (Figs 6 and 8). Our previous findings on the generation of Ca²⁺ wave through the activation of P2Y₁ purinergic receptor‐phospholipase C‐IP₃R2 signalling by ATP release in rat atrial myocytes under shear stress (Kim & Woo, 2015) support the present observation, and partly explain how IP₃Rs are activated during shear stimulation.

Our results suggest that ∼25% of the outward I _shear is probably carried by Cl⁻ (Fig. 2 E and F), which is also apparent throughout the TRPM4 inhibition data (Figs 3A–C, 4 and 5). This component also appears to be reduced in atrial myocytes from the IP₃R2 KO mice (∼15% of WT I _shear) (Fig. 8), indicating that Cl⁻ channel function may be also disrupted. Because the non‐TRPM4 current component appears to be independent of intracellular Ca²⁺ increase (Figs. 1 C and 6), the relatively smaller outward I _shear in IP₃R2 KO myocytes further suggests the possible regulation of Cl⁻ channel function by IP₃R2 in a Ca²⁺‐independent manner under shear stress.

Possible role of subsarcolemmal local Ca²⁺ signalling in TRPM4 activation in atrial myocytes

We also showed that the TRPM4 current is relatively resistant to intracellular dialysis of EGTA (Fig. 1 C and D) and that there is significant co‐localization of IP₃R2 and TRPM4 in the periphery of atrial myocytes (Fig. 9). These data further support a Ca²⁺‐mediated interaction between IP₃R2 and TRPM4 in peripheral microdomains in atrial cells. I _shear was not significantly inhibited by dialysis with 2 mm EGTA, and was only partially suppressed by 4 mm EGTA (Fig. 1 C and D). At these concentrations, Ca²⁺ diffusion is limited to much less than 2 μm, and Ca²⁺ release from the SR by high concentrations of caffeine or depolarization fails to activate an inward NCX current in cardiac myocytes (Adachi‐Akahane et al. 1996; Woo et al. 2002, 2003). A shear stress of a similar magnitude induced a longitudinal global Ca²⁺ wave (Woo et al. 2007), although this wave is assumed to be suppressed by dialysing 2 mm EGTA into atrial cells. Therefore, local Ca²⁺ release occurring nearby in peripheral junctional SR through the IP₃R may be more critical for TRPM4 activity.

Because IP₃R2, the major IP₃R subtype in cardiac myocytes, is co‐localized with RyR in the peripheral junctional SR of atrial myocytes (Lipp et al. 2000; Mackenzie et al. 2002), Ca²⁺ release through the IP₃R probably recruits nearby RyRs via CICR and result in amplified, prolonged Ca²⁺ signals in the narrow peripheral junction space. A functional role of IP₃Rs in facilitating the activity of RyRs is supported by previous observations showing that the frequency of Ca²⁺ sparks increases with no change in the SR Ca²⁺ content during the stimulation of endothelin and angiotensin receptors (Mackenzie et al. 2002; Gassanov et al. 2006) or even under resting conditions (Li et al. 2005). This may explain why we observed similar levels of I _shear inhibition in the presence of ryanodine and IP₃R inhibitors (Fig. 6 A−D). It also suggests that the specialized microarchitecture and localization of TRPM4 relative to the two different SR Ca²⁺ release channels may be important determinants of Ca²⁺‐dependent activation of TRPM4 in intact cells. Because ventricular and atrial myocytes differ in ultrastructure, as well as in the distribution of IP₃Rs and RyRs, it remains to be determined how ventricular TRPM4 is regulated under shear stress.

Pathophysiological implications

Under resting conditions, shear stress‐mediated Ca²⁺ increases and subsequent activation of TRPM4 may elicit spontaneous membrane depolarization by allowing monovalent cation influx through the channels. Accordingly, evidence for the role of TRPM4 in resting membrane depolarization was previously reported in sinoatrial node cells (Hof et al. 2013) and in cerebral artery myocytes (Earley et al. 2007). In normal beating cardiac cells, TRPM4 contributes to action potential prolongation (Simard et al. 2013; Mathar et al. 2014). The protein is activated by regular Ca²⁺ transient in ventricular myocytes, thereby reducing the driving force for Ca²⁺ influx through L‐type Ca²⁺ channels (Mathar et al. 2014). It is expected that, under shear stress, atrial action potential may be prolonged by the TRPM4 current, and that altered action potential may in turn affect Ca²⁺ current‐triggered Ca²⁺ release. However, it has also recently been reported that relatively low shear stress (4 dyn cm⁻²) induces insertion of the ultrarapid delayed rectifying K⁺ channel (Kv1.5) into the atrial cell membrane by Ca²⁺‐dependent exocytosis, thereby inducing action potential shortening (Boycott et al. 2013). Furthermore, in the mouse ventricle, the TRPM4 inhibitor 9‐phenanthrol abolished early afterdepolarization induced by hypoxia and re‐oxygenation at concentrations that do not affect voltage‐dependent ion channels (Simard et al. 2012). Possible remodelling of voltage‐dependent ionic currents by physiologically or pathologically relevant shear stress needs to be investigated to fully understand the alterations of action potentials and Ca²⁺ signalling in atrial cells.

Taken together, our observations and previous studies indicate that single atrial myocytes show significant responses to shear stress in the range 2−25 dyn cm⁻², similar to other cell types, such as hair cells and endothelial cells, although the level of effective shear force varies for different responses (e.g. Ca²⁺ spark/wave, Ca²⁺ transient and Kv1.5 current) (Woo et al. 2007; Balmote & Morad, 2008; Boycott et al. 2013). However, assuming that the maximum shear force is 0.0016 kPa (equivalent to 16 dyn cm⁻²) and that the stiffness of a myocyte is ∼23 kPa (Zile et al. 1998), the strain (fractional cell lengthening) on a myocyte calculated by the equation [stress (kPa) = A/k × (e^kε − 1), where A = 23 kPa, k = 6 and ε is the strain] is very small compared to the length changes in vivo during a normal contraction. Boycott et al. (2013) suggested that interlaminar shear stress in the adult rat atria, estimated using the Couette flow model, is 0.43 dyn cm⁻² at resting heart rate (5 Hz). The threshold level of shear stress for the detection of shear‐induced whole‐cell current was ∼2 dyn cm⁻² (Fig. 11), which is significantly higher than the rat atrial shear stress approximated under physiological conditions (Boycott et al. 2013). Atrial shear stress, however, increases during volume/pressure overload as a result of conditions such as valvular heart disease, congestive heart failure and hypertension. It is known that, in patients with mitral valve diseases, chronic regurgitant jets of blood directed at the atrial wall elicit widespread endocardial surface disruption and perforation (Goldsmith et al. 2000; Saffitz, 2009), exposing atrial myocytes to high shear stress. Considering that the shear stress generated during mitral regurgitation is dependent on the atrioventricular valve orifice and the pressure gradient across the valve during systole, it may be even more difficult to estimate the fluid‐jet force on single myocytes in vivo. The fact that the shear stress used to activate the TRPM4 current in single atrial cells is higher than the estimated interlaminar shear stress in the normal atrium suggests pathological relevance of this response. Indeed, regurgitation (e.g. mitral regurgitation) and volume/pressure overload are commonly associated with a sustained arrhythmia (atrial fibrillation) (Nazir & Lab, 1996; Goldsmith et al. 2000; Nattel, 2002). Interestingly, our observation of sustained inward TRPM4 current at negative voltages during prolonged shear stress may partly explain the clinically observed continuous depolarization accompanied by sustained atrial arrhythmia.

Figure 11. Threshold level of shear stress to activate shear‐sensitive current (I_shear) .

A, left: effects of 1 and 2 dyn cm⁻² on whole‐cell ramp currents (I _m) in a representative rat atrial myocyte. Right: I _shear at 1 and 2 dyn cm⁻². Note that shear stress of 2 dyn cm⁻² significantly increases I _m. B, summary of the inward and outward I _m in the absence (Control; n = 8) and presence of shear stress (1 dyn cm⁻², n = 9; 2 dyn cm⁻², n = 7). *P < 0.05, **P < 0.01 vs. Control (unpaired t test).

Additional information

Competing interests

The authors declare that they have no competing interests.

Funding

The work was supported by the National Research Foundation of Korea (NRF) grants funded by the Korea Government (MEST) (2012‐0005369, 2015R1A2A2A01002625, 2012‐0006681).

Author contributions

M‐JS, IS and S‐HW contributed to the conception and design of the experiments. M‐JS, J‐CK, SWK, JM, TDS, BC and KPS were involved in the experiments, collection, analysis and interpretation of data. Experiments were conducted in the laboratory of S‐HW. M‐JS, J‐CK, KPS and S‐HW drafted the manuscript. All authors have approved the final version of the manuscript and agree to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for authorship are listed.