Increased atrial effectiveness of flecainide conferred by altered biophysical properties of sodium channels

Abstract

Atrial fibrillation (AF) affects over 1% of the population and is a leading cause of stroke and heart failure in the elderly. A feared side effect of sodium channel blocker therapy, ventricular pro-arrhythmia, appears to be relatively rare in patients with AF. The biophysical reasons for this relative safety of sodium blockers are not known.

Our data demonstrates intrinsic differences between atrial and ventricular cardiac voltage-gated sodium currents (I_Na), leading to reduced maximum upstroke velocity of action potential and slower conduction, in left atria compared to ventricle. Reduced atrial I_Na is only detected at physiological membrane potentials and is driven by alterations in sodium channel biophysical properties and not by Na_V1.5 protein expression. Flecainide displayed greater inhibition of atrial I_Na, greater reduction of maximum upstroke velocity of action potential, and slowed conduction in atrial cells and tissue.

Our work highlights differences in biophysical properties of sodium channels in left atria and ventricles and their response to flecainide. These differences can explain the relative safety of sodium channel blocker therapy in patients with atrial fibrillation.

1. Introduction

Atrial fibrillation (AF) is a common cardiac arrhythmia and a major driver of stroke, heart failure, and cardiovascular death. Incidence and prevalence of AF is expected to increase further, with projected estimates of 6 million AF patients in US and 17.9 million in Europe, over the next 30–40 years [1,2].

Sodium channel blockers such as flecainide are commonly used to restore rhythm in patients with AF. These were initially developed to suppress ventricular arrhythmias. Following the increased mortality found in the CAST trial comparing flecainide and encainide to placebo [3], flecainide and similar agents are now primarily used for rhythm control therapy of patients with atrial fibrillation (AF) with normal ventricular function and without ischemic heart disease [4–8]. Recent controlled trials (Flec-SL and EAST-AFNET 4) demonstrate a low rate of ventricular pro-arrhythmia when flecainide is used after cardioversion and during long-term rhythm control therapy as part of an early rhythm control strategy, including in patients with heart failure with preserved ejection fraction [4,9,10]. This contrasts with the ventricular proarrhythmia observed when flecainide was used to suppress ventricular arrhythmias in patients with myocardial ischemia [3]. The reasons for this discrepancy are unclear and with the predicted increase in use of early rhythm therapy, as a consequence of the EAST-AFNET 4 trial, better understanding of the effects of flecainide on atrial and ventricular electrical function is waranted.

Although flecainide has been shown to inhibit the rapid delayed rectifier current at clinically relevant doses, which may account for some proarrhythmic outcomes, flecainide primarily inhibits myocardial Na⁺ channels, of which Na_V1.5 is the primary pore-forming alpha subunit found in mammalian heart [11–15]. Na_V1.5 is a large transmembrane protein primarily mediating the cardiac Na⁺ current (I_Na) in cardiac cells. Localization and plasma membrane expression of the Na_V1.5 is reliant on specific beta subunits [16] and Na_Vβ2 and Na_Vβ4 subunits can modulate the kinetic and voltage dependence properties of the I_Na [17]. Flecainide is known to block Na_V1.5 in its open, activated state, where it can bind to the pore and prevent Na⁺ from traversing the membrane [18,19]. This leads to enhanced refractoriness especially at rapid atrial rates, as seen in AF [20]. However, inhibition of Na⁺ channels also slows conduction velocity, which can both terminate multiple reentry and increase the predisposition to functional conduction block and macro-reentrant arrhythmias [21,22]. Functional conduction block and macro-reentry are believed to contribute to the ventricular proarrhythmia found in the CAST trial studying flecainide, encanide, and moricizine in survivors of a myocardial infarction with reduced left ventricular function.

Several crucial questions remain unanswered, regarding flecainide and atrial and ventricular electrical function. It is not known whether flecainide exhibits differential sodium channel-blocking effects in atria and ventricles. Characterization of the atrial and ventricular conduction differences are yet to be studied in the same hearts, or indeed the extent of conduction slowing caused by flecainide. Chamber differences in peak I_Na when measured over a range of different holding potentials remains to be determined, as well as the sensitivity to flecainide. Whilst increased peak I_Na is reported in cells isolated from left atrium (LA) compared to those isolated from left ventricle (LV) [23–27], it is not clear if this is conserved at more physiological resting membrane potentials (RMPs), which are likely to differentially impact on sodium channel availability, gating and flecainide efficacy. Furthermore, it is not known if chamber differences in expression of SCN5a/Nav1.5, SCN2B/Na_Vβ2 and SCN4B/Na_Vβ4 exist and if they are consistent in healthy murine and human heart tissue.

The aims of the present study were to robustly interrogate differences in biophysical properties of ventricular and atrial I_Na (inclusive of measurements at physiological membrane potentials), their effects on conduction, and examine chamber specific responses to flecainide. Furthermore, we investigated expression profiles of sodium channel subunits in atrial and ventricular chambers in non-failing murine and human tissue.

2. Results

2.1. Biophysical properties of I_Na differ between the left ventricle and left atrium

We first set out to quantitatively compare I_Na in adult murine cardiomyocytes isolated from left atria and left ventricles. To measure peak I_Na, we initially employed a standard protocol, where I_Na was elicited from a holding potential of -100 mV. Example raw I_Na traces are demonstrated in Fig. 1A. When normalized to capacitance (Supplemental Fig. 1) and measured from a holding potential of -100 mV no differences in I_Na density were seen between the left atria and left ventricles at any test potential (Fig. 1B,C). However, atrial cardiomyocytes exhibited an approximately 4 mV more negative V_50,act, compared to ventricular cardiomyocytes (Fig. 1D). We also examined whether differences in the I_Na inactivation kinetics exist in datasets in Fig. 1. Time to peak and time to 95% decay were increased in left ventricular cardiomyocytes compared to atrial (time to peak - LA 0.86 ± 0.03 ms, LV 1.16 ±0.06 ms; ****p ≤ 0.0001 unpaired t-test, n = 30 cells for LA and n = 29 cells for LV), (time to 95% decay - LA 4.25 ±0.23 ms, LV 6.47 ±0.24 ms; ****p ≤0.0001 unpaired t-test, n = 30 cells for LA and n = 29 cells for LV). These differences however are likely driven by larger ventricular currents.

Fig. 1. Left atrial (LA) and left ventricular (LV) I_Na current/voltage relationship and sodium channel activation.

A) Representative raw I_Na traces from LA and LV cardiomyocytes, inset indicates voltage protocol. B) Current voltage relationship of INa peak density in LA and LV. C) I_Na peak density at a step from -100 mV to -30 mV in LA (−27.4 ± 1.7 pA/pF) and LV (−26.8 ± 1.0 pA/pF); p = 0.785. D) V_5Q,act fitted to the Boltzmann distribution in LA (−42.7 ± 0.8 mV;) and LV (−38.6 ± 0.9 mV). Each dot represents an individual cell, **p < 0.01, (Two-way nested ANOVA). **P < 0.01, n = 42/13 cells/mice for LA and n = 28/13 cells/mice for LV.

Under normal physiological conditions, cardiomyocytes are not at a resting membrane voltage of -100 mV or -120 mV (conditions at which I_Na is usually measured), however the resting membrane potential lies in the range of −90 to -65 mV [28,29]. Thus, the magnitude of I_Na over this physiological range will depend not only on the population size of functional Na_V1.5 channels but also on the number of channels that are available to conduct ions at a given resting membrane voltage. Therefore, we went on to determine how resting membrane potential could alter the peak I_Na current (Fig. 2A). The holding potential had a marked and differential effect on the left atrial and left ventricular I_Na. At a holding potential of -120 mV, atrial peak I_Na was larger (−35.7 ± 1.3 pA/pF, n = 40/13 cells/mice) when compared to ventricular cardiomyocytes (−30.9 ± 1.7, p = 0.027, Fig. 2B). However, this was reversed when sodium channels were activated from a more physiologically relevant holding potential of -75 mV; peak current density was lower in left atrial cardiomyocytes (6.7 ±0.8 pA/pF, n = 40/13 cells/mice) when compared to the ventricular cardiomyocytes (9.7 ±1.1pA/pF, n = 28/13 cells/mice, p = 0.026), Fig. 2C. Thus, although atrial cardiomyocytes have a greater maximal I_Na density when activated from −120 mV, the physiologically relevant I_Na size is actually significantly lower in atrial than ventricular cardiomyocytes.

Fig. 2. INa density at varying holding potentials from left atrial (LA) and left ventricular (LV) cardiomyocytes.

A) I_Na mean density/holding potential relationship, LA (n = 40/13 cells/mice) and LV (n = 28/13 cells/mice). The range of the physiological resting membrane potential is highlighted in grey. Voltage protocol shown in inset. B) I_Na density at -120 mV holding potential in LA (−35.7 ± 1.3 pA/pF; n = 40/13 cells/mice) and in LV (−30.9 ± 1.7 pA/pF; n = 28/13 cells/mice, *p < 0.05). Each dot represents an individual cell, **p < 0.01, (Two-way nested ANOVA). C) I_Na density at -75 mV holding potential in LA (−6.7 ± 0.8 pA/pF; n = 40/13 cells/mice) and LV (−9.7 ± 1.1 pA/pF; n = 28/13 cells/mice, *p < 0.05) Each dot represents an individual cell, **p < 0.01, (Two-way nested ANOVA).

Voltage dependent steady state inactivation of I_Na is critical to the function of the cardiomyocyte as it determines the number of available Na_V1.5 channels, and thus can have major effects on peak depolarizing current, conduction and interactions with pharmacological agents. Left atrial I_Na inactivated at significantly more negative voltages than in left ventricular cardiomyocytes, with significant differences observed over a wide range of voltages (Supplemental Fig. 2A). The mean voltage of half-maximal steady-state inactivation (V_{50, inact}) of the atrial cardiomyocytes was 6.5 ±1.5 mV more negative than that of ventricular cardiomyocytes (Supplemental Fig. 2B). This has important physiological consequences; at −120 mV there was no difference in steady-state inactivation %, as nearly all channels are available. However, at less negative voltages, inactivation % is greater in the LA when compared to the LV. For example, at — 90 mV, 47.7 ±3.3% of channels were no longer available for activation in the LA, compared to only 24.8 ± 2.6% in the LV (p < 0.0001). At a more physiological potential of — 75 mV, the difference in inactivated channels between the left atria (82 ± 2.2%) and left ventricles (71.8 ±2.8%, P < 0.0049, Supplemental Fig. 2C) is significant. Voltage inactivation slope constants (k) were consistent between LA (7 ± 0.3, N = 12) and LV (6.9 ± 0.1, N = 19, P ≥ 0.05).

Recovery from inactivation was examined using a double pulse, P1 and P2 protocol that delivered two identical depolarizing pulses to −30 mV of 25 ms duration. Atrial I_Na was slower to recover from inactivation when comparing the P50 (time when half of the channels were recovered) of recovery in LA (22.6 ± 1.7 ms) and LV (15.5 ± 1.5 ms) (Supplemental Fig. 3). Recovery differences shown here are therefore likely to contribute to the reduced I_Na at physiological holding potentials, action potential upstroke and conduction in the atria, observed in this study.

In order to examine whether these differing I_Na characteristics can lead to alterations in action potential morphology, specifically action potential (AP) upstroke, we went on to include these biophysical data in a mathematical model of the ventricular cardiomyocyte [30]. As shown in Fig. 3, the modelling data fitted the experimental data well in terms of current activation and inactivation (Fig. 3A and B).

Fig. 3. Modelling data illustrates a larger ventricular I_Na current leading to larger action potential amplitude in ventricles during a modelled cardiac action potential.

A) I_Na density at varying holding potentials from left atrial (LA) and left ventricular (LV) cardiomyocytes from the model and experimental data. B) LA and LV I_Na current/voltage relationship and sodium channel activation in both experimental and modelling data. C) I_Na generated during a modelled AP. D) Examples of modelled APs utilizing the LA and LV I_Na data.

Incorporating the kinetics of the sodium channel from the left atrial and left ventricular cardiomyocytes resulted in a reduced current density in the ‘atrial’ cell as well as slower current decay during an AP when compared to the ventricular cell (Fig. 3C). There was also a slower peak maximum upstroke velocity (dV/dt) of the modelled ‘atrial’ cell compared to the ventricular cell (121.59 mV/ms for the LA when compared to 130.32 mV/ms for LV)and a smaller AP amplitude, when measured at 1 Hz pacing cycle length and an RMP of -80 mV, Fig. 3D. We followed these modelling experiments with current-clamp experiments on single isolated cardiac myocytes (Fig. 4) to determine the applicability of the modelling data. As shown in Fig. 4B, when paced at 1 Hz, left atrial myocytes had an RMP of 70.8 ± 1.4 mV, whereas left ventricular myocytes were generally less depolarized with an RMP of −74.4 ± 1.0 mV (p = 0.0002, n = 23–40/5 cells/mice). Furthermore, maximal upstroke velocity of the AP was significantly higher in single ventricular cells when compared to atrial cells (at 1 Hz, LA 238.6 ± 27.7 mV/ms; LV 304.2 ± 27.9 mV/ms *p = 0.0018, n = 23–40/5 cells/mice) (Fig. 4C).

Fig. 4. Maximal upstroke velocity is higher in left ventricle (LV – grey bars) compared to left atrium (LA – blue bars) in single cardiac myocytes whilst LA myocytes are more sensitive to flecainide block (1 μM).

A) Raw traces. B) Resting membrane potential of LA cells are significantly less negative than those of the LV at a variety of pacing cycle lengths (at 1 Hz, LA −70.8 ± 1.4 mV;LV -74.4 ± 1.0 mV p = 0.0002, n = 23–40/5 cells/mice.) Each dot represents average per mouse from at least 3 cells, **p < 0.01, ***p < 0.001, (Two-way ANOVA). C) Maximal upstroke velocity is higher in single ventricular cells when compared to atrial cells (at 1 Hz, LA 238.6 ± 27.7 mV/ms;LV 304.2 ± 27.9 mV/ms *p = 0.0018, n = 23–40/5 cells/mice) Each dot represents average per mouse from at least 3 cells, **p < 0.01, ***p < 0.001, (Two-way nested ANOVA). D) After application of flecainide only 14/23 atrial cells remained excitable, whereas 17/19 ventricular cells were still able to generate an action potential (p = 0.07, Fisher’s Exact Test, n = 4 mice). E) Flecainide significantly decreased dV/dT to a greater extent in atrial cells (at 1 Hz 47.9 ± 18.8%) when compared to ventricle cells (At 1 Hz, 18.6 ± 9.8%, p = 0.04, Each dot represents average per mouse from at least 3 cells, Two way nested ANOVA, n = 4 animals). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

2.2. Flecainide has a greater inhibitory effect in the left atrial cardiomyocytes

We investigated the effect of clinically relevant flecainide concentration (1 μM) on I_Na and action potential properties in atrial and ventricular cardiomyocytes. When activating I_Na from a fixed holding potential of −100 mV, flecainide inhibited I_Na to a larger extent in the atrial when compared to the ventricular cells (Fig. 5). For example, at a test potential of −35 mV, I_Na was inhibited by approximately 48% in the atria, Fig. 5B ((27.3 ± 3.1 pA/pf (control) to 14.1 ± 2.0 pA/pF (flecainide), p < 0.0001 n = 11/6 cells/mice)), whilst flecainide inhibited ventricular I_Na by approximately 38%, Fig. 5C ((27.0 ± 1.6 pA/pf (control) to 18.5 ± 1.6 pA/pF (flecainide), p < 0.0001 n = 14/4 cells/mice)). The enhanced action of flecainide on atrial cardiomyocytes was maintained over the entire range of holding voltages but was exaggerated at more positive, physiological potentials as seen in Fig. 5D. At the most negative holding potential, of -120 mV, I_Na was inhibited by 47.4 ± 4.6% and 33.6 ± 2.6% in the atrial and ventricular cardiomyocytes respectively (p = 0.0126 n = 11/6,14/3 cells/mice), whilst at a holding potential of −70 mV currents were inhibited by 75.9 ± 6.8% in the atrial and 44.4 ± 4.5% in the ventricular cardomyocytes (p < 0.001, n = 11/6,14/3 cells/mice), Fig. 5E,F. This indicates a greater intrinsic sensitivity of left atrial I_Na to flecainide as well as an exaggerated voltage dependence. Interestingly, flecainide also significantly slowed sodium channel recovery in atrial, but not ventricular cardiomyocytes (Supplemental Fig. 4).

Fig. 5. Flecainide (1 μM) inhibits atrial I_Na to a greater extent than ventricular I_Na, which is more apparent at more positive holding potentials.

A) Representative raw I_Na traces from left atrial (LA) and left ventricular (LV) cardiomyocytes with and without flecainide. Inset indicates voltage protocol. B) Current-voltage relationship of I_Na density in LA (n = 11/6 cells/mice) C) Current-voltage relationship of I_Na density in LV (n = 14/4 cells/mice). D) % inhibition of I_Na in left atrial (LA) and left ventricular (LV) cardiomyocytes from a variety of holding potentials, when stimulating I_Na by a voltage step to -30 mV. Inset indicates voltage protocol. E) LA and LV I_Na inhibition by flecainide from a holding potential of -120 mV. (LA = 75.9% ± 6.8; LV = 44.4% ± 4.5; ***p < 0.001) F) LA and LV I_Na inhibition from a holding potential of -70 mV (LA = 47.4% ± 4.6; LV = 33.6% ± 2.6). n = 12/6 cells/mice for LA and 14/4 for LV. Each dot represents an individual cell, **p < 0.01, ***p < 0.001, ****p < 0.0001 (Unpaired t-test).

When we applied the flecainide biophysical inhibitory effects on I_Na in the cell model simulations, a reduction of dV/dt to 55.6 mV/ms in the atria when compared to 93.64 mV/ms in the ventricle was calculated, at 1 Hz pacing cycle length. Collectively, these data indicate that flecainide has a greater effect on atrial I_Na. We also applied flecainide to single cells under current clamp conditions in atrial and ventricular cells (Fig. 4). Interestingly, when superfusingflecainide under experimental conditions only 14/23 atrial cells remained excitable, whereas 17/19 ventricular cells were still able to generate an action potential (p = 0.07, Fisher’s Exact Test, n = 4 mice), indicating the increased effectiveness of flecainide in the atria (Fig. 4D). When we considered all cells (and a block of an AP as 100% inhibition), flecainide significantly decreased dV/dT to a greater extent in atrial cells (paced at 1 Hz 47.9 ± 18.8%) when compared to ventricular cells (paced at 1 Hz, 18.6 ± 9.8%, p = 0.04, Two way ANOVA, n = 4 animals)(Fig. 4E), commensurate with the modelling data.

2.3. Atria display slower conduction velocity and are more sensitive to flecainide

In order to confirm the earlier cellular and modelling findings, we set out to investigate whether conduction of electrical signals is different across matched murine left ventricle and left atria, using optical mapping (Fig. 6). Fig. 6E illustrates that LA conduction velocities are significantly reduced compared to the LV at 10 Hz pacing cycle length (24.4 ± 2.7 vs 36.1 ± 6.2 cm/s, p = 0.03. N = 5). As expected, action potential duration was prolonged in the LV when compared to the LA (APD₅₀ 27.1 ± 2.4 ms in the LV compared to 12.5 ± 0.8 ms in the LA, p = 0.0011, N = 5)(Figs. 6B, C).

Fig. 6. Conduction velocities (CV) are reduced in left atria (LA) when compared to the left ventricles (LV), and are differentially affected by flecainide (1 μM) whilst APD is longer in LV.

A) Representative action potential duration (APD50) maps recorded simultaneously from murine LA and LV tissue paced at 10 Hz. B) Grouped data of APD50 in LA (12.5 ± 0.8 ms) and LV (27.1 ± 2.4 ms, p = 0.0011, N = 5, two tailed student’s t-test). Each dot represents an individual heart. C) Representative examples of APs recorded from the LA and LV. D) Representative CV maps in the LV compared to the LA. Contour lines are spaced by 2 ms. E) Grouped data of CV in LA (24.4 ± 2.7 cm/s) and LV (36.1 ± 6.2 cm/s, p = 0.03. N = 5, two tailed student’s t-test). F) and G) Grouped data of CV in LA (F, light blue) and LV (G, grey) after the perfusion of flecainide and in comparison to time matched controls (n = 7–10). H) and I) Grouped data of APD in LA (H, light blue) and LV (I, grey) after the perfusion of flecainide and in comparison to time matched controls (n = 7–10). Each dot represents an individual heart. *p < 0.05, (two tailed student’s t-test). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

In a separate set of experiments, we investigated if flecainide exerted differential effects on LA and LV CVs in the intact heart. In agreement with the cellular data, flecainide significantly decreased conduction in the left atrium (59.6 ± 7.2% of baseline) (Fig. 6F) whilst conduction was not significantly affected in the left ventricle (73.7 ± 4.7% of baseline) (Fig. 6G). Furthermore, no differences were seen in action potential duration between the two chambers after exposure to flecainide (Fig. 6H,I).

2.4. Na_V1.5, ß2 and ß4 subunits are differentially expressed in murine left atrium and left ventricle

We then determined whether potential differences in protein expression could explain the differing biophysical and pharmacological properties between the left atrium and the left ventricle. Specifically, we examined the expression of Na_V1.5, Na_Vβ2 and Na_Vβ4 in murine atrial and ventricular tissue, using Western blotting. Na_Vβ2 and Na_Vβ4 were previously demonstrated to affect activation, inactivation and recovery of I_Na [25]. As can be seen in Figs. 7A,D, Na_V1.5 showed a higher relative expression in the left atria when compared to the left ventricle in the mouse heart (LA = 3.18 ± 0.40 AU; LV = 1.00 ± 0.16 AU; n = 13, p <0.0001). A number of additional bands of lower molecular weight were also noted on the Western blot, likely due to the polyclonal nature of the Na_V1.5 antibody (Supplemental Fig. 5). In contrast to this, both Na_Vβ2 (LA = 0.22 ± 0.03 AU; LV = 0.41 ± 0.05 AU; n = 6, p = 0.0123) and Na_Vβ4 (LA = 0.09 ± 0.003 AU; LV = 0.21 ± 0.01 AU; n = 6, p < 0.0001) were shown to be expressed at lower levels in the left atria when compared to the left ventricles (Figs. 7D,B,C).

Fig. 7.

Protein expression and RNAseq data of key sodium channel proteins in murine and human tissue. Western blots showing the protein expression of Na_V1.5 (A), Na_Vβ2 (B) and Na_Vβ4 (C) from murine left atrial (LA) and left ventricular (LV) samples. Na_V1.5 showed a higher relative expression in the LA, when compared to the LV (LA = 3.18 ± 0.40 AU; LV = 1.00 ± 0.16 AU; n = 13) ****p < 0.0001. Both Navβ2 (LA = 0.22 ± 0.03 AU; LV = 0.41 ± 0.05 AU; n = 6, *p < 0.05) and Na_Vβ4 (LA = 0.09 ± 0.003 AU; LV = 0.21 ± 0.01 AU; n = 6, *****p < 0.0001) are expressed at higher levels in the ventricle when compared to the atria. All blots were normalized to GAPDH. Each dot represents an individual heart, **p < 0.01, ***p < 0.001, ****p < 0.0001 (Two tailed Student’s t-test). D) Heat maps illustrating the expression of SCN5A, SCN2B, and SCN4B from 79 paired patient samples from healthy LA and LV human tissue. E) Grouped data illustrating the RNAseq data for SCN5A, SCN2B, and SCN4B. SCN5A transcript levels were significantly higher in the LA when compared to the LV (adjusted P = 7.71E-23) whilst SCN4B transcript levels were significantly higher in the ventricle when compared to the atrium (adjusted P = 0.0057). No differences were seen in SCN2B transcript levels between the two chambers. Boxes represent the 25–75% and whiskers represent the min-max range, excluding outliers, which are shown as individual dots.

2.5. SCN4B transcripts are decreased, whilst SCN5A are increased in left atrium compared to left ventricle in non-failing human heart tissue

Finally, we sought to determine if these findings in murine tissue are replicated in human tissue. To do so we conducted differential expression analysis on RNAseq from matched human left atrial and left ventricular tissue derived from 79 non-failing hearts. As seen in Fig. 7E,F, transcripts for SCN5A, the gene that encodes for Na_V1.5, was found at significantly higher levels in human left atria, compared to human left ventricles (1.66-fold change, P_adjusted = 7.71*10 ²³), consistent with the murine expression data (Figs. 7A,D). Furthermore, transcript levels of SCN4B, which codes for Na_Vβ4, were significantly lower in human left atria when compared to the left ventricles (1.24-fold change, p_adjusted = 0.0057). SCN2B (Na_Vβ2) expression was modestly reduced in the left atria but this was not significant (1.09-fold change, P_adjusted = 0.407).

3. Discussion

The present findings demonstrate that flecainide exhibits some ‘atrial-selective’ inhibition of I_Na in the normal adult heart. These effects can explain the relatively low risk of ventricular pro-arrhythmia when flecainide is used in patients with atrial fibrillation. In more detail, the data demonstrate that: 1) left atrial tissue has slower conduction velocities then the left ventricle and is more sensitive to flecainide, when measured in the same heart; 2) Despite the left atrium expressing more SCN5A/Na_V1.5 protein then the left ventricle, at physiological membrane potentials, peak I_Na density is reduced in the atria; 3) The reduction in I_Na at physiological resting membrane potentials is sufficient to account for the decreased action potential upstroke velocity in the left atrium compared to the left ventricle; 4) Flecainide is more selective for left atrial I_Na compared to left ventricular cardiomyocytes; 5) Atrial peak I_Na density and sensitivity to flecainide is dramatically more influenced by resting membrane potential than the ventricular peak I_Na density. 6) Flecainide reduces the peak AP upstroke velocity more in atrial cells compared to ventricular cells.

3.1. Mechanisms promoting atrial susceptibility to arrhythmias

We robustly interrogated differential conduction velocities in the atria and ventricles by multi-vector, single-vector and activation time methodologies (Supplemental Fig. 6) [31]. Crucially, our optical mapping data compares atrial and ventricular recordings in the same heart. Conduction velocities recorded in our study are in the expected range, as demonstrated by others [32]. We demonstrate for the first time that mouse atrial tissue display circa 30% reduced conduction velocity compared to the ventricle, regardless of the methodology (multi vs single vector) utilized (Fig. 6; Supplemental Fig. 6). These data shed some light on the potential mechanisms contributing to enhanced atrial susceptibility for maintenance of re-entrant arrhythmias. In agreement with our data, van Veen et al. whilst employing electrogram array to record conduction in 3–4 months old mice, showed slightly lower conduction velocities in left atria (circa 30 cm/s) than the ventricles (circa 36 cm/s), though these were not directly compared [33]. Additionally, Thomas et al. detected a small but non-significant increase in conduction in mouse ventricles compared to the atria, using a liner epicardial electrode array [34].

Remarkably, reduced conduction in atria was observed despite a clear increase in Na_V1.5 protein expression in left atrial tissue, compared to left ventricle. Increased murine atrial Na_V1.5 expression was consistent with our data showing significantly increased expression of SCN5A in non-failing human left atrium, compared to left ventricle (Fig. 7). Consistent with these molecular biology findings, left atrial cardiomyocytes show increased peak I_Na compared to ventricular, when all sodium channels are activated, from a holding potential of −120 mV (Fig. 2A,C). Our patch clamp findings are in line with reports of others that use the holding potentials of −120 mV. Li et al. [24] were the first to describe marked increase in I_Na density in isolated atrial guinea pig cardiomyocytes when compared to ventricular epicardial cells. Similar findings have also been reported in rat, rabbit, and canine cardiomyocytes [23,25–27]. These studies have looked at the current density from a holding potential of −120 mV, when all channels are available, which is not the case under physiological conditions. Healthy ventricular cells have a resting membrane potential of between −65 and −90 mV in situ whilst atrial cells are slightly more depolarized, sitting between −65 and −80 mV [28,29]. In the present study we show that atrial peak I_Na density is differentially influenced by resting membrane potential, more so than observed in the ventricular cardiomyocyte (Fig. 2). Thus, at physiological resting membrane potentials, atria actually display a smaller peak I_Na density when compared to the ventricles. Consistent with the observed reduced peak I_Na in the left atrium, the action potential amplitude and V_max of atrial APs have been experimentally described as being between ≅ 150–300 V/s [35,36], compared with higher values of 250–450 V/s for human ventricular cells [37–39]. Similar differences appear to be conserved in smaller species [28,40,41]. Our modelling and cellular data consistently show a faster upstroke in the ventricle compared to the atria (Figs. 3 and 4), in agreement with the cardiac optical mapping studies (Fig. 6). With regards to the modelling data, we likely underestimate the impact of the reduced atrial I_Na on the upstroke, as we utilize a ventricular cell model and do not account for many other differences that exist between the atria and ventricle, such as the altered resting membrane potential. Since the magnitude of depolarizing I_Na is the key factor that determines conduction, a reduced I_Na at physiological membrane potentials is the most likely explanation for the slower conduction that we observed in the left atrium compared to the left ventricle (Fig. 6).

So, what determines the lower atrial I_Na at physiological membrane potentials, despite expression of a greater number of Na_V1.5 channels? Our patch clamp data demonstrate that left atrial cardiomyocytes display a greater negative shift in the voltage dependence of sodium channel inactivation (Supplemental Fig. 2). This is expected to result in a markedly reduced number of available Na_v1.5 channels, and a lower peak I_Na, when initiated from physiological holding potentials. We also detected a negative shift in atrial V50_act, meaning that the voltage at which half of the sodium channels are activated is more negative in the left atrial cardiomyocytes, when compared to the left ventricular cells (Fig. 1). This is in line with previous findings in rabbit cardiomyocytes [23]. It may be expected that a negative shift of the voltage dependence of sodium channel activation would decrease the voltage difference between the RMP and the activation threshold, leading to increased I_Na in the atria. However, this is counteracted by a greater negative shift in the voltage dependence of atrial sodium channel inactivation (Supplemental Fig. 2), resulting in a markedly reduced number of available Nav1.5 channels, and a lower peak I_Na when initiated from physiological holding potentials. However, these data do not measure V50_act at physiological membrane potentials thus it is difficult to predict the effects on peak I_Na.

The negative shift in V50_act of the atrial sodium channels suggests an enhancement of excitability of atrial cardiomyocytes compared to ventricular cardiomyocytes. Negative shifts in the voltage dependence of I_Na activation permits channel activation with smaller depolarizations. This can result in premature activation of the sodium channels with minor depolarization. Although it has been shown here that there are a smaller proportion of left atrial sodium channels available for activation at physiologically relevant holding potentials, those that are available are potentially more sensitive to an untimely depolarization of membrane potential and may be more susceptible to premature activation.

Thus, the combined functional effects of the different biophysical properties of left atrial and left ventricular cardiomyocyte I_Na are a slowing of conduction velocity but an increased risk of ectopy. These effects are probably further magnified due to the more positive resting membrane potential in the atria compared to the ventricles, as illustrated in Fig. 4B. This may well account for the increased incidence of ectopy and re-entrant arrhythmia in the atria compared to ventricle although more work is required to validate this.

3.2. Do Na_Vβ-subunit expression differences mediate I_Na differences between the atria and ventricles?

Here we show that atrial expression of Na_Vβ2 and Na_Vβ4 is dramatically reduced in mouse hearts. In addition, RNAseq data from non-failing human hearts showed a similar pattern, reduction in transcript levels of SCN2B and SCN4B in the left atrium (Fig. 7). Chen et al., showed that kinetic properties of sodium channels co-expressing Na_V1.5 with Na_Vβ2 and Na_Vβ4 subunits were similar to those of the ventricular sodium channels, with more positive activation potential, more positive inactivation and faster recovery of the sodium channels. Equally, kinetic properties of sodium channels expressing Na_V1.5 alone were similar to those of the atrial sodium channels [25]. However, as experiments were performed in HEK293 cells co-expressing Na_Vβ2 and Na_Vβ4 subunits together, it is unclear whether the effects on channel kinetics required expression of both Na_Vβ2 and Na_Vβ4 or just one of the subunits. Indeed, Malhotra et al. showed that β2 had no detectable effects on channel kinetics using a heterologous expression system, suggesting that the effects of β2 may involve cell adhesion and cytoskeletal communication as opposed to channel gating [42]. Meanwhile, co-expression of the β4 subunit with Na_V1.5 resulted in a negative shift in the voltage dependence of inactivation when compared to Na_V1.5 alone [43]. Furthermore, mutations in both of these subunits have been associated with arrhythmogenic consequences [43,44]. In addition to β2 and β4 subunits Nav1.5 may also interact with β1 and β3 subunits, which have also been shown to modulate sodium channel activity [45,46]. Interestingly, recent work has illustrated that the loss of Scn1b (Na_Vβ1) in murine myocytes differentially affected the potencies of lidocaine and ranolazine, whilst also showing a distinct difference in transcriptional expression between human atria and ventricles [47] In the present study we did not investigate the effects of Na_Vβ1 and Na_Vβ3 expression on sodium channel function and localization, however, this should be an area of further study.

In addition to direct effects of β subunits on biophysical properties of I_Na, β subunits can also affect sodium channel localization. For AP conduction, location of Na_V1.5 on the cell membrane is particularly important, as are the number of channels in a particular cluster of channels as recently shown by Hichri et al. [48] Furthermore, work has shown that there are a number of different sub-pools of channels at the lateral membrane [49]. Whether β-subunits are involved in localization of channels to a particular microdomain remains to be investigated.

3.3. Flecainide effectiveness and the atria

Flecainide preferentially binds to sodium channels in the open state [50]. Our data adds to this knowledge by clearly showing that flecainide more effectively inhibits atrial I_Na across a range of holding potentials including physiological potentials. Several differences between atrial and ventricular tissue can explain this difference, including i) greater inhibition due to the difference in resting membrane potential [28], ii) greater inhibition of left atrial I_Na regardless of holding potential, and iii) greater inhibition of left atrial I_Na due to increased atrial expression of Na_Vβ2 and Na_Vβ4 subunits. Additionally, in vivo, greater inhibition is expected due to increased use dependence in fast-firing fibrillating atria.

It is well-known that the atrial resting membrane potential is more positive –depolarized – than the ventricular resting membrane potential [29,40] and this has also been shown in the current data set. Our data now add novel insights into the differences between atrial and ventricular sodium channel properties and expression profiles and suggest that these physiological differences lead to further inherent atrial selectivity of flecainide (Figs. 4, 5 and 6). We have previously shown that the extent of flecainide’s I_Na inhibition is highly sensitive to small changes in atrial resting membrane potential, whilst our more recent work has shown that atrial RMP modifies the effectiveness of several clinically used AADs [28,51]. This sensitivity clearly contributes to the observed lower atrial I_Na at physiological membrane potentials. In addition, other factors affecting the differential effects of flecainide on Na_v1.5 channels in the atria and ventricles clearly exist. Whether the alterations in β-subunit chamber specific expression seen in this study contribute to differences in flecainide effectiveness remains unanswered. Further studies determining the supra-molecular clustering of sodium channels including their subunit on cardiomyocyte membranes are warranted. In murine Scn3b^−/− hearts, where the β3 subunit is not expressed, flecainide produced reduced arrhythmic incidences combined with prolonged refractory periods and shortened APDs, despite the fact that these cardiomyocytes showed a reduction in I_Na and a negative shift in Na_V1.5 channel inactivation [52].

In light of our findings, atrial selectivity of flecainide in healthy myocardium is apparent and thus our data explain the lack of ventricular arrhythmias observed in Flec-SL and EAST-AFNET4 trials. The dosing regimens in these trials is not dissimilar to that of the CAST and CAST II trials so perhaps depolarization of the ventricular resting membrane potential during ischemia and in the border zone of myocardial infarctions enhanced flecainide effectiveness in “dangerous” areas in the left ventricle with scars (which were present in all patients in CAST and CAST II).

3.4. Limitations

While we verified key molecular changes in human tissue, this study was mainly performed in healthy mouse hearts. Validation in large mammals and human cardiomyocytes is warranted. We must also consider that our findings are relevant for hearts without structural heart disease, the main group of patients in whom sodium channel blockers are currently used, this study did not directly investigate the proarrhythmic effects of flecainide which are mainly found in infarcted hearts with structural heart disease or during ischemia. In addition to direct effects of β subunits on biophysical properties of I_Na, β subunits can also affect sodium channel clustering, a novel modulator of Nav1.5 function [48] It is conceivable that altered clustering contribute to the physiological differences between atrial and ventricular I_Na found here. In addition it has been shown that other alpha-subunits, such as Nav1.8, may contribute to cardiac conduction and mRNA for such proteins have been detected in both atrial and ventricular tissue [53]. How these may contribute to the chamber differences observed in this study is unclear. Furthermore, work has shown that there are a number of different subpools of channels at the lateral membrane [49]. Whether β-subunits are involved in localization of channels to a particular microdomain remains to be investigated. Finally we also need to consider that it is not only the number of available channels at physiological diastolic potentials that will determine the conduction velocity but it can also be determined by other factors such as the density of gap junctions, sodium and calcium concentrations as well as cell geometry [54–56]. These factors go beyond the scope of the present study but should definitely be investigated in the future.

3.5. Conclusion and clinical significance

Our data illustrate the striking differences in the electrical properties of left atrial and ventricular myocardium. Of most clinical interest is demonstration that flecainide is more effective at inhibiting atrial sodium channels. Our findings can explain the good safety profile of sodium channel blockers in patients without myocardial scars, e.g. those with atrial fibrillation and with preserved left ventricular function, while also providing a reasonable explanation for the pro-arrhythmia seen in earlier studies in survivors of a myocardial infarction with heart failure with reduced ejection fraction. Determining the molecular drivers of the atrial and ventricular resting membrane potential and identifying the molecular interaction partners of flecainide and similar substances, may help to identify new targets for safe antiarrhythmic drug therapy. This is much needed to implement systematic, early rhythm control therapy in patients with AF.

4. Methods

4.1. Animal model

All experiments were conducted under the Animals (Scientific Procedures) Act 1986 and approved by the home office (PPL numbers 30/2967 and PFDAAF77F) and the institutional review board at the University of Birmingham.

4.2. Optical mapping of murine atria and ventricles

Mouse (CD-1, 29–49 g, Charles River, United Kingdom) hearts were isolated under deep isoflurane-induced inhalation anaesthesia (4% in O₂, 1.5 L min⁻¹). Optical mapping of mouse whole hearts was performed as previously described [57]. Mice were anesthetized using isoflurane (4% in 100% O2, 1.5 L min⁻¹). Hearts were removed and the aorta immediately cannulated. Hearts were Langendorff-perfused with an oxygenated (95% O2 5% CO2) crystalloid buffer solution containing (in mM) NaCl 114, KCl 4, CaCl2 1.4, NaHCO3 24, NaH2PO4 1.1, glucose 11 and sodium pyruvate 1 (pH 7.4, 37°C) at a perfusion pressure of ca.80 mmHg. Potentiometric dye Di-4-ANEPPS (5 mg/mL. 25-100 μL) was loaded via bolus injection into the perfusion line over 3–5 min. Blebbistatin (12 μM) was added to the perfusate to prevent movement of the tissue during recording, whilst flecainide was added in the relevant experiments at 1 μM.

Hearts were paced at 10 Hz with 1 ms pulses from the right atrium epicardial surface using silver bipolar electrodes. Pulse amplitude was 2×the diastolic threshold. Hearts were illuminated during recordings by two dual LEDs (530 ± 25 nm) and emitted light collected at >630 nm. 10s recordings were acquired at 0.5 kHz at 51*51 pixel resolution using a Evolve Delta EMCCD camera (Photometrics, USA).

To ensure effective recording of the signals, sequential recordings were collected. Following recording of the anterior ventricular surface, the hearts were re-orientated. The atria were moved so they were in the focal plane and removed from the ventricles to prevent signal overlap. To enable direct comparison between the left atria and ventricle, a 9 × 9 pixel (1.4 cm2) region was analysed from both chambers (Fig. 1). Optical action potentials (APs) and conduction velocities (CV) were analysed using ElectroMap, as previously described [31].

4.3. Murine cardiomyocyte isolation

LA and LV murine cardiomyocytes were isolated from male and female adult mice (12–20 weeks old), bred on the 129/sv background, as previously described [28]. Briefly, hearts were removed under isoflurane-induced anaesthesia (4% in O₂, 1.5-3 L min⁻¹) and perfused at 4 ml/min at 37 °C on a vertical Langendorff apparatus with the following solutions, equilibrated with 100% O2: (i) HEPES-buffered modified Tyrode’s solution containing in mM: NaCl 145, KCl 5.4, CaCl₂ 1.8, MgSO₄ 0.83, Na₂HPO₄ 0.33, HEPES 5 and glucose 11 (pH 7.4, NaOH); (ii) Ca²⁺-free Tyrode’s solution for 5 min; (iii) Tyrode’s enzyme solution containing 20 μg/mL Liberase™ (Roche, Indianapolis, IN, for I_Na experiments) or 640 μg/ml collagenase type II, 600 μg/ml collagenase type IV and 50 μg/ml protease (Worthington, Lakewood, NJ, for AP recordings), 0.1% bovine serum albumin (BSA, Sigma), 20 mM taurine and 3-30 μM CaCl₂. The switch to a different enzyme solution for AP recordings was due to commercial availability. The heart was removed from the Langendorff and perfused with 5 ml of modified Kraft-Bruhe (KB) solution containing in mM: DL-potassium aspartate 10, L-potassium glutamate 100, KCl 25, KH₂PO₄ 10, MgSO₄ 2, taurine 20, creatine 5, EGTA 0.5, HEPES 5, 0.1% BSA and glucose 20 (pH 7.2, KOH). Perfusion timings were between 10 and 25 min dependent on whether ventricular or atrial cells were being isolated and between hearts. The LA and LV chambers were dissected from the digested heart and placed into separate petri dishes containing 1 ml/2 ml respectively of the modified KB solution where they were manually disassociated using glass pipettes with increasing resistances. After dissociation was completed, Ca²⁺was reintroduced in a stepwise manner over a period of 90 min to a final concentration of 1.8 mM for recordings of I_Na and 1 mM for AP recordings.

4.4. Electrophysiology

Dissociated mouse LA and LV cardiomyocytes were transferred to an initially static bath recording chamber and allowed to adhere to laminincoated coverslips (10 mm diameter). Whole cell patch clamp recordings were obtained in voltage clamp mode using borosilicate glass pipettes (tip resistances 2.2–3 MΩ). All recordings and analysis protocols were performed using an Axopatch 1D amplifier (Molecular Devices, USA) and a CED micro1401 driven by Signal v6 software (CED, UK). Pipette solutions contained (mM): CsCl 115, NaCl 5, HEPES 10, EGTA 10, MgATP 5, MgCl₂ 0.5 and TEA 20 (pH 7.2, CsOH). To ensure adequate voltage control of I_Na, currents were recorded in a low sodium external solution containing in mM: NaCl 10, C₅H₁₄ClNO 130, KCl 4.5, HEPES 10, CaCl₂ 1.8, MgCl₂ 1.2 and glucose 10 (pH 7.4, CsOH).L-type Ca²⁺ current was blocked with 2 mM NiCl₂. Experiments were performed at 22 ± 0.5 °C. I_Na was recorded before and after the addition of flecainide (1 μM). Current signals were sampled at 50 kHz and low pass filtered at 20 kHz. I_Na was normalized to cell capacitance and expressed as pA/pF. To assess current-voltage relationships, I_Na was elicited using 100 ms step depolarizations at 1 Hz over a test potential range of -95 mV to +10 mV, in 5 mV increments, from a holding potential of -100 mV. I/V curves were fitted using the modified Boltzmann equation as described previously [58]. Measurements of steady state inactivation of I_Na, were made by applying pre-pulses ranging from −120 to −40 mV in 5 mV increments for 500 ms prior to the test potential (−30 mV for 100 ms). As well as being able to calculate V₅₀ inactivation voltages, this protocol also allowed for measurements of I_Na at different holding voltages. This also allowed for measurements of instantaneous I_Na inhibition by flecainide across a range of different holding voltages which could then be compared between atrial and ventricular cardiomyocytes. For recording of action potentials whole cell patch clamp recordings were obtained in current clamp mode. Pipette solutions contained (mM): 130 K⁺ glutamate, 10 NaCl, 10 KCl, 0.5 MgCl₂, 5 MgATP, 10 HEPES (pH 7.2, CsOH). The external solution contained (mM): 145 NaCl, 5.4 KCl, 0.83 MgSO₄.7H₂0, 0.33 NaH₂PO₄, 5 HEPES, 11 Glucose, 1 CaCl₂, 1.2 MgCl₂ (pH 7.4, CsOH). Experiments were performed at 37 ± 0.5 °C. Action potentials were stimulated via the pipette using a stimulus of at least twice diastolic threshold, at frequencies of 1 and 3 Hz, for a duration of 1 ms. Following baseline recordings, cells were superfused with 1 μM flecainide for 5 min before repeating the recordings. Action potential characteristics were measured using modified algorithms from the ElectroMap software as previously described [51]. Following 60 s pacing to reach steady state, resting membrane potential (RMP) was defined as the minimum diastolic membrane potential. Maximum upstroke velocity (dV/dt) was measured as the maximum derivative of the action potential recording.

4.5. Biophysical modelling

Full details for the modelling methodology and data can be found in the supplement. In brief, sodium channel activation and inactivation protocols were fitted to a standard Hodgkin and Huxley formulation of the I_Na channel:

$$I_{Na}=G_{Na}{m}^{3}hj\left(V-E_{Na}\right)$$

G_Na is the maximum channel conductance, m is the activation gating variable, h and j are inactivation gating variables, V is the membrane potential, E_Na is the sodium Nernst potential. We assume j and h are equivalent as insufficient data is available to fit distinct inactivation gates. The gating variables were fitted to activation and inactivation data for atria and ventricles.

Flecainide inhibition is dependent on the holding potential and is distinct for the atria and ventricle. To approximate flecainide effects, we introduced a linear scalar that represented sodium channel inhibition. We fitted this scalar to inhibition data for atria and ventricle sodium channels. To predict the impact of different sodium ion channel kinetics on emergent action potential morphology, we introduced the fitted atrial and ventricle sodium channel models into the mouse variant of the Pandit ventricular myocyte model [30]. We used the same model for both the atria and ventricle sodium channel model.

4.6. Western blotting

Tissue samples were homogenised in homogenisation buffer (Tris adjusted to pH 7.3, 1 protease cocktail inhibitor tablet (Roche Diagnostics; complete ULTRA Tablets, Mini, EASYpack) diluted in 10 mL buffer, with phosphatase cocktail inhibitor 2 and 3 (Sigma) added (both 1:100 concentration). The homogenisation buffer was added to the tubes containing the.

tissue in a ratio of 1 mg of tissue: 10uL of buffer and homogenised using the Precellys® system (Bertin Instruments). Samples were then centrifuged and 10uL of sample (≈15μg of protein) was loaded onto 4–15% gradient Mini-PROTEAN® TGXTM gels (Bio-Rad) immersed in running buffer (0.025 M Tris base, 0.19 M glycine and 0.1%w/v sodium dodecyl sulphate). Proteins were transferred onto polyvinylidene difluoride membranes (Bio-Rad). The membrane was then blocked with 5% milk powder dissolved in PBS-T overnight at 4 °C. Blocked membranes were then incubated for an hour with antibodies for Na_V1.5 (1:200, Alomone, ASC-005), Na_Vβ2 (1:400, Alomone, ASC-007), Na_Vβ4 (1:800, Alomone, ASC-044) and GAPDH (1:4000, ThermoFisher Scientific, AM4300). After washing, membranes were then incubated for 1 h at room temperature with the appropriate horseradish peroxidase linked conjugated secondary IgG antibodies (1:7500, GE Healthcare NA934V or 1:4000, GE Healthcare NA931V) diluted in 5% milk powder in PBS-T at room temperature.

For detection membranes were submerged in 5 ml of detection reagent (GE Healthcare, ECL Western Blotting Detection Reagents, Amersham) whilst the Odyssey Fc Imaging System (LICOR) was utilized to measure densitometry to monitor the level of protein expression. Raw densitometry values were normalized to GAPDH.

4.7. RNAseq

Kallisto (version 0.42.3) was used to quantify the transcripts in all experiments [59]. To generate an indexed transcriptome for performing pseudoalignments, we downloaded the human cDNA file Homo_sapiens. GRCh38.cdna.all.fa.gz from Ensembl (ftp://ftp.ensembl.org/pub/release-91/fasta/homo_sapiens/cdna/). This compressed FASTA file was made into an indexed transcriptome using the command “kallisto index” and this file was subsequently used for all transcript quantification. RNA-seq data derived from 79 pairs of adult matched LA and LV tissue was utilized (dbGaP accession phs001539.v1.p1 for left atrium, GEO accession GSE141910 for LV, Online Table 1) to assess differences and similarities between the chambers.

After quantification of transcript abundance using kallisto, we used the tximport package to import quantification data into the DEseq2 package in R [60,61]. To collapse transcripts into gene level summaries, we used the “hsapiens_gene_ensembl” repository through the R package biomaRt [62]. Only genes with at least 10 counts across the row were retained to filter noise at the low end of the distribution of transcript abundance. To identify DE genes, we used the function “DESeqDataSetFromTximport” to ingest the filtered data, then the function “results” to produce lists of DE genes. Unadjusted P-values are produced using the Wald test, and P-values are adjusted for multiple testing corrections using Benjamini-Hochberg. We used the DEseq2 function lfcShrink to transform expression values to log2 scale. All analyses were performed in R using version 3.5.1.

4.8. Statistical analyses

Data are expressed as mean ± standard error, unless otherwise stated. Data were checked for normal distribution and statistical analysis was performed using (un)paired Student t-tests or two-way (nested) ANOVA with Tukey post-hoc as appropriate. Countables, such as number of excitable vs non-excitable cells, were tested using the Fisher’s exact test. Significance was taken as two-sided P < 0.05, and all analysis was carried out in GraphPad Prism 9.0.

Abbreviations

RMP

resting membrane potential

AF

Atrial Fibrillation

LA

Left atrium

LV

Left ventricle

CV

Conduction velocity