Augmentation of myocardial If dysregulates calcium homeostasis and causes adverse cardiac remodeling

Yampolsky, Pessah; Koenen, Michael; Mosqueira, Matias; Geschwill, Pascal; Nauck, Sebastian; Witzenberger, Monika; Seyler, Claudia; Fink, Thomas; Kruska, Mathieu; Bruehl, Claus; Schwoerer, Alexander P.; Ehmke, Heimo; Fink, Rainer H. A.; Draguhn, Andreas; Thomas, Dierk; Katus, Hugo A.; Schweizer, Patrick A.

doi:10.1038/s41467-019-11261-2

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Published: 23 July 2019

Augmentation of myocardial I_f dysregulates calcium homeostasis and causes adverse cardiac remodeling

Nature Communications volume 10, Article number: 3295 (2019) Cite this article

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Abstract

HCN channels underlie the depolarizing funny current (I_f) that contributes importantly to cardiac pacemaking. I_f is upregulated in failing and infarcted hearts, but its implication in disease mechanisms remained unresolved. We generated transgenic mice (HCN4^tg/wt) to assess functional consequences of HCN4 overexpression-mediated I_f increase in cardiomyocytes to levels observed in human heart failure. HCN4^tg/wt animals exhibit a dilated cardiomyopathy phenotype with increased cellular arrhythmogenicity but unchanged heart rate and conduction parameters. I_f augmentation induces a diastolic Na⁺ influx shifting the Na⁺/Ca²⁺ exchanger equilibrium towards ‘reverse mode’ leading to increased [Ca²⁺]_i. Changed Ca²⁺ homeostasis results in significantly higher systolic [Ca²⁺]_i transients and stimulates apoptosis. Pharmacological inhibition of I_f prevents the rise of [Ca²⁺]_i and protects from ventricular remodeling. Here we report that augmented myocardial I_f alters intracellular Ca²⁺ homeostasis leading to structural cardiac changes and increased arrhythmogenicity. Inhibition of myocardial I_f per se may constitute a therapeutic mechanism to prevent cardiomyopathy.

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Introduction

The hyperpolarization-activated cyclic nucleotide-gated channel 4 (HCN4) is the dominant HCN-isoform in the sinoatrial node and is significantly involved in generation and regulation of heart rhythm^1,2,3,4. Apart from its abundant expression in the pacemaker and conduction system, the adult working myocardium is characterized by low HCN levels^5,6. At early embryonic stages, however, HCN4 is abundantly transcribed in the whole heart⁶ and contributes importantly to I_f triggered automaticity of ventricular myocytes⁷. Moreover, HCN4 was identified as a cell marker for the cardiomyogenic progenitor pool of the first heart field, implicated in the earliest stage of cardiac mesoderm formation and morphogenesis^8,9,10. Toward birth HCN4 transcription is downregulated in working-type cardiomyocytes⁶ and remains at low levels during adult stages, suggested to prevent pathological remodeling⁶. In this context, increased I_f and HCN expression was reported in failing hearts and after myocardial infarction^5,11,12, but its pathophysiological role remains to be established. Notably, pharmacological blockade of I_f improves cardiovascular outcome in patients with chronic heart failure—results that were considered to be driven mainly by improved myocardial energy supply at lower heart rates^13,14. However, recent data demonstrated that I_f blockade in patients with coronary artery disease with preserved myocardial function did not improve outcome despite marked reduction of heart rate¹⁵, pointing towards mechanisms of I_f blockade that may be particularly beneficial in failing hearts. In this regard, pleiotropic effects of I_f blockade beyond heart rate reduction have been suggested^16,17, but these mechanisms have not been specified yet.

To study the significance of increased I_f in the working myocardium, we generated transgenic mice that express human HCN4 (hHCN4) under control of the murine cardiac troponin I (cTNI) gene promoter. We here show that a moderate increase of I_f in cardiac myocytes to levels observed in heart failure leads to cardiac dilation and impaired cardiac function. HCN4 activity essentially is linked to calcium regulation of cardiomyocytes and augmented I_f results in dysregulated Ca²⁺ homeostasis driving cell death and cardiac remodeling.

Results

Generation of transgenic mice overexpressing hHCN4

To study the significance of HCN4 channel overexpression-mediated I_f augmentation in the working myocardium, transgenic mice were generated that express hHCN4 under control of the murine cTNI gene promoter^18,19 (Fig. 1a). HCN4^tg/wt offspring are born at a mendelian frequency and exhibit no overt cardiac or general abnormalities. HCN4^tg/wt mice produce abundant hHCN4 transcripts (Fig. 1b) leading to increased HCN4 protein levels (Fig. 1c). Intense anti-HCN4 staining signals at the plasma membrane of HCN4^tg/wt cardiomyocytes (Fig. 1d) indicate efficient trafficking of hHCN4 channels to the cell surface, while wild type tissue yields only little anti-HCN4 signals. Accordingly, patch-clamp recordings showed significantly increased I_f densities in HCN4^tg/wt cardiomyocytes. At physiological resting membrane potentials (RMP) of ventricular cardiomyocytes (~−90 mV) we observed a two–three3-fold higher I_f density in HCN4^tg/wt cardiomyocytes compared to wild type (Fig. 1e, f), recapitulating the magnitude of current increase that was observed in cardiomyocytes of heart failure patients⁵.

HCN4 ^tg/wt mice develop structural heart disease

While cardiac morphology was similar at birth, we uncovered cardiac dilation with significantly increased heart-to-body weight ratios in HCN4^tg/wt mice 2 months postpartum (Fig. 2a–g). Right ventricular diameters, in particular, were found markedly enlarged (Fig. 2d) and dilation was accompanied with reduced wall thickness (Fig. 2e). To assess cardiac in vivo phenotypes, echocardiographic evaluations were performed at 3 and 6 months of age. At 3-month of age left ventricular wall thickness of HCN4^tg/wt mice was lower compared to wild type (Fig. 2l). Similar to the histological findings at 2 months of age there was a trend toward higher left ventricular internal diameters in HCN4^tg/wt animals (Fig. 2j, k), although these changes did not reach statistical significance. Fractional area shortening did not differ between groups at this age (Fig. 2i). At 6 month of age, however, HCN4^tg/w mice displayed a dilated cardiomyopathy phenotype (Fig. 2h, i) with lower left ventricular wall thickness, dilation of left ventricular internal diameters (Fig. 2n–p) and systolic dysfunction with reduced fractional shortening of 22.9 ± 1.4% compared to 27.5 ± 1.4% in controls (means ± s.e.m., P = 0.031, unpaired t-test) (Fig. 2i). Furthermore mitral valve Doppler imaging displayed increased E/A ratios in transgenic mice indicative for impaired left ventricular relaxation and diastolic dysfunction (2.16 ± 0.22 in HCN4^tg/w mice compared to 1.62 ± 0.07 in controls, P = 0.021, unpaired t-test) (Fig. 2m, q).

In line with macroscopic alterations in HCN4^tg/wt hearts, we observed significantly increased diameters of cardiomyocytes (Fig. 2c) and elongated sarcomeres (Fig. 3a, b). However, HCN4 overexpression did not induce cardiac fibrosis (Fig. 3c, d), and we observed no myocyte disarray or cellular infiltrations (Fig. 3a, e). Early treatment of HCN4^tg/wt mice with the I_f blocker ivabradine significantly reduced chamber dilation and wall rarefication (Fig. 2b–g), and normalized cardiomyocyte diameters (Fig. 2c), indicating that increased I_f causes the altered structural phenotype.

Transcriptional alterations in HCN4 ^tg/wt hearts

We next asked whether structural and functional remodeling in hearts of HCN4^tg/wt mice is associated with activation of the fetal gene program. Evaluation of mRNA, isolated from 6-months-old mice, revealed markedly increased levels of the myocardial cell growth markers GSK-3B and mTOR (Fig. 4a) and slightly upregulated levels of the prototypical hypertrophic marker genes MYH7 (encoding ßMHC), CnA (encoding calcineurin A) and NPPA (encoding ANP) (Fig. 4b). We further explored transcriptional profiles of ion channels and transporters, implicated in action potential generation, excitation-contraction coupling and intracellular calcium homeostasis (Fig. 4c, d). Interestingly, HCN4^tg/wt hearts displayed significant downregulation of the sodium–calcium exchanger gene NCX1 (Fig. 4c) and intrinsic murine HCN genes were slightly diminished (Fig. 4d). Furthermore, levels of transcripts of the repolarizing channel genes KCNQ1 (encoding Kv7.1) and KCND2 (encoding Kv4.2) were decreased, while the repolarizing channel gene KCNH2 (encoding Kv11.1) showed a trend toward higher levels (Fig. 4e). Other genes regulating calcium handling and cellular electrophysiology, however, showed transcript levels that were not significantly changed.

HCN4 overexpression impacts calcium handling proteins

Based on NCX1 downregulation in HCN4^tg/wt hearts, we hypothesized that dysregulated intracellular sodium–calcium homeostasis might be a pathomechanistic component in I_f overexpression-mediated cardiomyopathy. To evaluate a contribution of calcium handling proteins we performed immunoblotting of various key players. In line with the transcriptional analysis we observed significantly reduced levels of NCX1 protein in HCN4^tg/wt hearts, a finding that was paralleled by lower levels of SERCA2 and phospholamban (PLN) whole protein content (Fig. 4e, f). Of note, phosphorylation of PLN at its CaMKII-specific Thr-17 site was significantly increased, whereas the rise at its protein kinase A (PKA)-site Ser-16 did not reach statistical significance. Moreover, the Ca²⁺-sensor protein µ-calpain showed higher levels in HCN4^tg/wt hearts, while the L-type calcium channel protein Cav1.2 remained unchanged (Fig. 4e, f). These data document profound alterations in the regulation of calcium handling proteins under the influence of augmented myocardial I_f.

HCN4 ^tg/wt cardiomyocytes show altered Ca²⁺ homeostasis

To determine the influence of increased I_f on intracellular Ca²⁺ handling, we measured [Ca²⁺]_i transients in isolated, electrically driven cardiomyocytes of wild type and HCN4^tg/wt mice at baseline and after incubation with 3 µM ivabradine (Fig. 5a). Remarkably, we observed a distinctive rise in [Ca²⁺]_i baseline levels and significantly increased systolic [Ca²⁺]_i transients under steady state field stimulation of cardiomyocytes isolated from HCN4^tg/wt hearts (Fig. 5a–d). The efficiency of Ca²⁺ release, however, reflected by the time to peak, was almost unchanged (Fig. 5e). Of note, [Ca²⁺]_i levels, high in HCN4^tg/wt cardiomyocytes, dropped to significantly lower levels in ivabradine-treated HCN4^tg/wt cardiomyocytes (Fig. 5b), suggesting that abundant HCN4 current mediates diastolic Ca²⁺ overload and altered Ca²⁺ homeostasis. Furthermore, the markedly increased slope of Ca²⁺ uptake (Fig. 5f) and shortened time to half decay (D₅₀) of calcium transients (Fig. 5d) indicate that HCN4 overexpression influences uptake of increased [Ca²⁺]_i to the SR mediated by SERCA. Consistently, I_f inhibition normalized the slope Ca²⁺ uptake and [Ca²⁺]_i parameters in cardiomyocytes of transgenic animals (Fig. 5a–f). In addition, Ca²⁺ load experiments using caffeine 10 mM (Supplementary Fig. 1) showed that the caffeine-induced Ca²⁺ release peak was significantly higher in HCN4^tg/wt cardiomyocytes, indicating that the SR Ca²⁺ content is increased, which is consistent with the experimental data of higher Ca²⁺ transients upon electrical stimulation (Fig. 5a–c). Of note, recordings of Ca²⁺ transients showed abnormal prolongation of diastolic Ca²⁺ decay (D₉₀ > 2000ms) in 39% of cardiomyocytes isolated from HCN4^tg/wt hearts (evaluated separately—please refer to Supplementary Fig. 2), changes that were reduced to wild type level after treatment of HCN4^tg/wt cardiomyocytes with ivabradine, underlining that increased I_f profoundly disorganizes cellular Ca²⁺ homeostasis.

Reverse mode NCX blockade ameliorates altered Ca²⁺ cycling

To elucidate the role of NCX in HCN4-mediated perturbation of Ca²⁺ cycling we recorded [Ca²⁺]_i transients from wild type and HCN4^tg/wt cardiomyocytes at baseline and in the presence of the selective NCX inhibitor ORM-10103²⁰.

Interestingly, NCX inhibition by ORM-10103 (10 µM) reversed the changes in Ca²⁺ metabolism nearly to wild type levels (Fig. 6a–f). Increased diastolic [Ca²⁺]_i declined remarkably (Fig. 6b) and the amounts of total cellular Ca²⁺ movement, reflected by the peak area, returned to normal levels (Fig. 6c). Moreover, the increased slope Ca²⁺ uptake (Fig. 6f) and shortened time to half decay (D₅₀) of calcium transients (Fig. 6d) were restored to levels not significantly different from wild type. Thus, our data demonstrate an important pathophysiological implication of the NCX current in the changed cardiac Ca²⁺ homeostasis of HCN4^tg/wt animals.

HCN4^tg/wt hearts show increased apoptosis

We then asked whether increased [Ca²⁺]_i would provoke apoptosis in HCN4^tg/wt hearts. Using TUNEL assay, we detected significantly higher proportions of nuclei with DNA fragmentation in ventricles of HCN4^tg/wt hearts compared to wild type (Fig. 7a, b). In addition, screening of genes associated with apoptosis by qRT-PCR demonstrated a significant rise of the apoptosis effector caspase-3 in transgenic myocardium (Fig. 7c, d; Supplementary Fig. 3). Moreover, we observed higher transcript levels of the Ca²⁺-sensor protease μ-calpain and the tissue transglutaminase (Fig. 7d), both implicated in myocardial cell death and structural remodeling in association with [Ca²⁺]_i overload²¹. Further analysis of caspase-3 tissue activity revealed a marked increase of caspase-3-positive cells in HCN4^tg/wt ventricles (Fig. 7e, f), pointing to increased apoptotic cell death as an explanation for wall thinning and dilated cardiomyopathy.

Electrophysiological changes in HCN4^tg/wt cardiomyocytes

Action potential (AP) recordings of HCN4^tg/wt and wild type cardiomyocytes were performed using ruptured whole-cell patch clamp technique (Fig. 8a). In HCN4^tg/wt cardiomyocytes RMP was more positive than in wild type cardiomyocytes (wild type: P₅₀ = −61.25 mV, n = 14 vs. HCN4^tg/wt: P₅₀ = −54.1 mV, n = 12; p = 0.0013; unpaired t-test) (Fig. 8c), consistent with diastolic depolarisation caused by HCN4-mediated sodium influx. Likewise, the amplitude of overshoot was diminished in transgenic cardiomyocytes (Fig. 8b), pointing to an inactivation of voltage-gated Na⁺-current at more depolarized RMP. AP properties were evaluated at three different pacing frequencies (0.5, 1, 2 Hz) (Fig. 8d–f). AP duration at 20 (APD₂₀) and 50% (APD₅₀) repolarization were shorter in transgenic cells, with significant changes of APD₅₀ at all pacing frequencies and of APD₂₀ at 0.5 Hz. Evaluation of APD₉₀, by contrast, did not differ significantly between wild type and HCN4^tg/wt cardiomyocytes (Fig. 8d–f).

HCN4^tg/wt cardiomyocytes are prone to arrhythmias

Based on the overexpression of depolarizing I_f and consecutive Ca²⁺ overload we asked whether HCN4^tg/wt cardiomyocytes elicit increased cellular automaticity and/or arrhythmogenesis. Recordings of [Ca²⁺]_i transients and APs (Fig. 9a–g) frequently revealed sustained trains of automaticity in HCN4^tg/wt cardiomyocytes (Fig. 9c), while such changes were only sparsely observed in wild type cells. Under steady state field stimulation the [Ca²⁺]_i transients of wild type cardiomyocytes quickly decayed (Fig. 9a), while a subset of HCN4^tg/wt cardiomyocytes showed abnormal diastolic [Ca²⁺]_i clearance (Fig. 9b, Supplementary Fig. 2). Correspondingly, transgenic cardiomyocytes were prone to afterdepolarizations (ADs), which resulted in premature APs and arrhythmic firing (Fig. 9e, f). Among all cells recorded, the percentage of cells exhibiting ADs or virtually spontaneous firing, was significantly higher in transgenic (55.5 ± 7.6%; n = 5 hearts; 12–30 cells per heart) than in wild type cardiomyocytes (17.8 ± 3.4%; n = 5 hearts, 22–34 cells per heart; P = 0.0002; ANOVA followed by Tukey test) (Fig. 9g). To delineate the immediate contribution of I_f to arrhythmogenesis within our model we treated the cells with 3 µM ivabradine (iva). Remarkably, iva treatment diminished spontaneous firing and ADs, and percentage of HCN4^tg/wt cardiomyocytes showing arrhythmic behavior after treatment was not different from wild type (Fig. 9g), demonstrating an important contribution of increased I_f to cellular arrhythmogenicity.

Electrophysiological phenotype of HCN4^tg/wt mice

To evaluate ECG parameters and to seek for arrhythmias in HCN4^tg/wt mice we performed ECG recordings under anesthesia and telemetric recordings in freely roaming animals. Both approaches revealed similar heart rates and conduction parameters among transgenic (HCN4^tg/wt) and wild type mice and did not exhibit significant changes of additional ECG parameters (Supplementary Fig. 4, Supplementary Table 1). However, increased numbers of premature ventricular captures and few non-sustained ventricular tachycardias (mostly triplets) were recorded in transgenic but not in wild type animals (Fig. 9h, i), while sustained ventricular tachycardias were not observed. This indicates that HCN4^tg/wt mice have a predisposition to ventricular arrhythmogenesis, although high-grade arrhythmias are not part of the phenotype.

Discussion

We here report that augmentation of myocardial I_f leads to the development of a cardiomyopathy phenotype with biventricular chamber dilation, significantly reduced wall thickness and decreased ejection fraction. To study the myocardial effect of increased I_f under conditions that minimized its influence on heart rate, we have generated transgenic mice that express human HCN4 (HCN4^tg/wt) under control of the murine cTNI promoter. While abundantly expressed in the working myocardium, cTNI is downregulated in the SAN and the conduction system²². This is phenotypically reflected by telemetric recordings revealing similar heart rates and conduction parameters in transgenic and wild type mice (Supplementary Fig. 4, Supplementary Table 1), thus providing the opportunity to selectively study the consequences of HCN4 overexpression in the working myocardium.

There is growing evidence that ion channel disorders not only cause cardiac arrhythmias, but also contribute to structural abnormalities of the heart as well^23,24,25,26. We and others reported^25,26 that HCN4 loss-of-function mutations are associated with noncompaction cardiomyopathy, pointing to an involvement of HCN4 in ventricular wall maturation at embryonic stages. Accordingly, HCN4 has been identified as primary cell marker for the cardiomyogenic progenitor pool of the first heart field, implicated in the earliest stage of heart formation^8,9. During later development and adult stages, HCN4 is downregulated in the healthy myocardium, while abundant expression is restricted to the SAN and the conduction system⁶. Remarkably, in this context, HCN2 was recently shown to influence developmental brain morphology and function, as well²⁷.

It is well documented that I_f and the expression of the main ventricular HCN isoforms 2 and 4 are significantly increased in ventricular moycytes of heart failure patients^5,11,28. These changes are considered reminiscent of the immature myocytic phenotype, based on the fact that during pathological remodeling several early genes reappear in reverse order compared to embryonic development²⁹. To mimic maladaptive upregulation of HCN4 in our model, the cTNI promoter was used to activate hHCN4 transcription from the intermediate-late phase of fetal development on³⁰, leading to augmented levels in the myocardium throughout postnatal and adult stages, normally characterized by low HCN4 expression⁶. At physiological RMP of ventricular cardiomyocytes (~−90 mV) patch clamp recordings revealed a two–three-fold higher I_f density in HCN4^tg/wt cardiomyocytes compared to wild type (Fig. 1e, f), reflecting the magnitude of current increase that was found in cardiomyocytes of heart failure patients⁵. Accordingly, transgenic mice developed myocardial alterations within two months postpartum, supporting a primary maladaptive involvement of increased I_f. Likewise, we observed a rise of apoptosis and markers of the fetal gene program (Fig. 3a, b) with significantly upregulated cell growth markers e.g. glycogen synthase kinase-3 beta (GSK-3B) and mammalian target of rapamycin (mTOR) (Fig. 4b), indicating cardiac remodeling and activation of compensatory cellular responses. Thus, postnatal myocardium appears susceptible to augmented I_f levels, and carefully regulated HCN patterning may constitute a prerequisite for the maintenance of structural cardiac integrity. In line, chronic peri- and postnatal treatment of transgenic mice with the I_f channel blocker ivabradine antagonized the myocardial changes (Fig. 2), underlining that augmented I_f is key to structural alterations occurring in HCN4^tg/wt animals. Noteworthy, the heart rate lowering effect of ivabradine may contribute to protection from adverse remodelling as well.

Given the primary Na⁺ conductance of HCN4 channels at cardiomyocyte RMP (~−90 mV)³¹ and the close interrelation of intracellular Na⁺ and Ca²⁺ homeostasis³², we asked how increased I_f might influence Ca²⁺ cycling. To elucidate the impact of increased I_f, we measured [Ca²⁺]_i transients in isolated, electrically driven cardiomyocytes in the pre- and absence of ivabradine. Strikingly, we observed a marked rise in [Ca²⁺]_i baseline levels and significantly augmented systolic [Ca²⁺]_i transients in transgenic cardiomyocytes – changes that were fully recovered in the presence of ivabradine. Thus, in the myocardium chronic upregulation of I_f exerts a diastolic Ca²⁺ overload and importantly interferes with cellular Ca²⁺ homeostasis.

As the primary mechanism of Ca²⁺ efflux in cardiac myocytes is via electrogenic Na⁺/Ca²⁺ -exchange (NCX), Ca²⁺ homeostasis is tightly linked to Na⁺ regulation³². Under physiological conditions, NCX primarily operates in (forward) Na⁺ -in/ Ca²⁺ -out mode, and reduces diastolic [Ca²⁺]_i levels by extruding Ca²⁺. In pathological states, however, when [Na⁺]_i is increased, NCX exerts ‘reverse mode’ function, moving Ca²⁺ into the cell³². Notably, [Na⁺]_i is known to rise in a rate dependent fashion³², which exacerbates [Ca²⁺]_i overload at high heart rates. The impact of I_f on [Na⁺]_i, in relation to membrane potential has been demonstrated in sheep Purkinje fibers by voltage-clamp recordings in early works^33,34 after the I_f mixed Na⁺ and K⁺ ionic nature was originally described³⁵. These studies showed that membrane hyperpolarization to below −60 mV significantly increased intracellular Na⁺ activity, and related this increase to Na⁺ influx through I_f, thus providing a direct link between [Na⁺]_i and the size of I_f. Based on these data, we hypothesized that HCN4 overexpression-mediated accumulation of [Na⁺]_i may raise Ca²⁺ influx via reverse mode NCX activity (Fig. 10). According to this assumption, instant application of ORM-10103, an agent that effectively inhibits reverse mode NCX^20,36, abolished Ca²⁺ overload, demonstrating the pathophysiological relevance of dysregulated NCX. Likewise, inhibition of increased Na⁺ influx via HCN4 channels using ivabradine facilitated resetting of the NCX equilibrium towards the ‘forward mode’ and restored [Ca²⁺]_i to wild type levels, similar to ORM-10103. Of note, depolarization of RMP by augmented I_f may lower the driving force of forward NCX a fortiori, thereby aggravating chronic accumulation of [Ca²⁺]_i in HCN4^tg/wt hearts. Based on these data it is reasonable to assume that inhibition of upregulated I_f in clinical heart failure may attenuate Ca²⁺ overload of the ventricular myocardium. Synergistically, heart rate lowering decreases [Na⁺]_i³² and therefore shifts the Na⁺/Ca²⁺ exchanger equilibrium towards the forward mode, which will alleviate [Ca²⁺]_i overload independently from ventricular I_f blockade. Thus, besides improved energy supply through heart rate reduction the delineated mechanisms may importantly contribute to positive outcome of patients with heart failure and increased heart rates treated by I_f blockade¹³.

We next addressed the effects of increased I_f and changed Ca²⁺ homeostasis on cellular electrophysiology. Action potential recordings of isolated HCN4^tg/wt cardiomyocytes showed depolarized RMP and reduced amplitude of overshoot in a rate-dependent fashion, consistent with HCN4-mediated diastolic Na⁺ influx. Furthermore, APD₂₀ and APD₅₀ were shorter compared to WT, while APD₉₀ was similar. This in parts is consistent with previous data obtained from cardiomyocytes overexpressing HCN2³⁷, which showed that outward tail current of I_f shortened APD at membrane potentials more positive than reversal potential (~−35 mV for HCN4³⁸), while inward tail current of I_f prolonged APD at membrane potentials more negative than reversal potential. However, prolongation of APD₉₀ was not observed in our model, most likely due to the repolarizing driving force of the electrogenic ‘reverse mode’ NCX, known to effectively shorten AP duration when [Na⁺]_i is increased³⁹.

Interestingly, we found that myocytes isolated from HCN4^tg/wt hearts showed a higher incidence of spontaneous APs than wild type cells, and a subset of cells exerted virtual pacemaker function. The increased automaticity may originate from I_f augmentation and not from spontaneous Ca²⁺ cycling-mediated membrane depolarisation, as the NCX equilibrium is shifted toward ‘reverse mode’ leading to an outward, hyperpolarizing current. Moreover, HCN4^tg/wt cardiomyocytes were prone to afterdepolarizations resulting in trains of premature APs. Recording of calcium transients showed abnormal diastolic Ca²⁺ decay in 39% of HCN4^tg/wt cells, produced by SR Ca²⁺ overload, promoting afterdepolarizations and arrhythmic firing⁴⁰. However, during telemetric recordings HCN4^tg/wt animals exhibited no sustained arrhythmias, although higher numbers of PVCs and few nonsustained VTs indicated increased arrhythmogenicity compared to wild type animals. This imbalance between cellular arrhythmogenicity and lack of high-grade arrhythmias in vivo might be explained by source-sink mismatch⁴¹ proposing that propagation of ADs originating from single cells (source) is hindered by the rectifying potassium conductance of neighboring myocytes within the myocardial syncytium (sink), thereby protecting the heart from deleterious arrhythmias.

One well-known hallmark of failing heart cells is disturbances in calcium cycling^42,43,44. Harmful [Ca²⁺]_i promotes cardiomyocyte death through apoptotic or necrotic pathways⁴³. Previous studies have delineated the association between apoptotic cell-loss and progression of heart failure in animal models and in humans^45,46. Moreover, Ca²⁺ overload induced by persistently activated L-type calcium channels^40,43,47 or chronically compromised Ca²⁺ removal systems⁴⁷ were shown to induce myocardial cell death and remodeling. Accordingly, the structural changes in our model can be explained by the observed caspase-3-mediated apoptosis. Likewise, we found higher transcript levels of tissue transglutaminase and the Ca²⁺-sensor protease μ-calpain (Fig. 3d), implicated in the activation of myocardial apoptosis and structural remodeling in association with [Ca²⁺]_i overload²¹. Basically, upregulation of I_f in heart failure might be seen as cellular short term mechanism that increases SR calcium load to provide more inotropic support. However, this is at the price of chronic [Ca²⁺]_i accumulation resulting in cell death and disease progression.

In summary, we show that a two–three-fold increase of I_f in cardiac myocytes, which is comparable to levels observed in human heart failure, affects cardiac structure and promotes cellular arrhythmogenicity in HCN4^tg/wt mice. Activation of reverse-mode NCX by increased HCN4-mediated Na⁺ influx leads to augmented [Ca²⁺]_i and dysregulated Ca²⁺ homeostasis driving apoptosis and adverse cardiac remodeling. This might have particular implications in disease states associated with increased I_f, affecting calcium cycling as well as excitation-contraction coupling. Thus, direct cardioprotective mechanisms in addition to improved energy supply at lower heart rates may underlie the beneficial effects of I_f inhibition in heart failure. Our findings provide insight in previously undescribed cardiac pathomechanisms, bridging a changed electrical state to a structural phenotype, which importantly could influence future treatment strategy of heart failure.

Methods

Generation of transgenic mice

Transgenic mice were generated in a C57Bl/6NCrl background (Charles River Laboratories, Wilmington, MA, USA) by pronuclear injection of an 8.2 kb DNA fragment carrying a 4.3 kb promoter fragment of the murine cTnI gene^18,19 joined to the 5′ end of the human HCN4 cDNA⁴⁸ that was fused to the bovine growth hormone gene poly A signal (Fig. 1a). Tail DNA analysis revealed four different founders that were subsequently bred with C57BL/6 N wild type mice to generate progeny that appeared healthy and fertile.

Care and use of mice

Mixed genotype groups of each gender of no more than five animals were housed in standard mouse cages under specific pathogen-free conditions (12:12-h dark–light cycle, constant temperature, constant humidity and food and water ad libitum). All experiments were carried out in accordance with the Guide for the Care and Use of Laboratory Animals published by the US National Institute of Health (NIH publication number 85–23, revised 1996), and with the European Community guidelines for the use of experimental animals as well as all relevant ethical regulations. Protocols were approved by the local regulatory authority (#35-9185.81/G-20/11 and #35-9185.81/G-226/16 Regierungspräsidium Karlsruhe, Germany). We used male mice for all experiments.

Surface and telemetric ECG recordings

For surface ECG recordings mice were anesthetized with isoflurane vapor titration to maintain the lightest anesthesia possible⁴⁹. On average, 1.5% vol/vol isoflurane was required to maintain adequate anesthesia. Mice were placed on a heating pad with continuous monitoring of body temperature (via a rectal probe) maintaining at 37 °C. Surface ECGs were recorded using clamp electrodes attached to each limb and to the chest in a midsternal position. The electrodes were connected to a Powerlab System (AD Instruments, Hastings, UK) and ECG recording were performed with a high and low frequency cut off of 1 kHz and 0,1 kHz. ECG recordings were analysed by an investigator blinded to the experimental groups using the Chart 5 software (AD Instruments, Hastings, UK). In addition to average heart rate, at least 30 complexes of each trace were signal averaged and the following parameters were evaluated in lead II: PR-interval, QRS complex duration, and QT-interval. Corrected QT intervals were calculated according to Mitchell et al., 1998⁵⁰, QTc = QT/(RR/100)1/2.

For telemetric ECG recordings, a radiotelemetry transmitter (model EA-F20, Data Sciences International, St. Paul, Minnesota, USA) was surgically inserted into the peritoneal cavity. Mice were anesthetized intraperitoneally with a mixture of ketamine (100 mg/kg) and xylazine (3 mg/kg). The leads were tunnelled under the skin to the recording sites, i.e. one subcutaneous lead placed at the right shoulder and a second subcutaneous lead placed at the lower left thorax. The abdominal skin was closed and mice were kept on a heating pad (37–38 °C) until they were fully recovered from anaesthesia. Before measurement, mice were allowed to recover for 1 week, followed by 24 h recording. ECG signals were converted to digital output and stored on a computer using the Powerlab System (AD Instruments, Hastings, UK), analysis was carried out with the Chart 5 software (AD Instruments, Hastings, UK).

Transthoracic echocardiography

The animals were sedated with low-dose isoflurane (1,5% vol/vol) and placed on a heating pad in supine position. A rectal probe was inserted to monitor the body temperature to maintain it at 37 °C. Chest hair was carefully removed with depilatory cream. Warmed echo gel was placed on the animal’s chest. The Vevo® 2100 device (VisualSonics, Toronto, Canada) with the corresponding transducer MS-400 was used to record the echocardiographies. The investigator was blinded with respect to the treatment group. Mice were shaved, and left ventricular parasternal short-axis views were obtained in M-mode imaging at the papillary muscle level. Three consecutive beats were used for measurements of left ventricular end-diastolic internal diameter (LVEDD) and left ventricular end-systolic internal diameter (LVESD). Fractional shortening (FS) was calculated as FS% = ((LVEDD − LVESD)/LVEDD) × 100.

Transcription analysis

Transcription was assayed by amplification of reverse transcribed total RNA isolated from whole ventricle tissue of wild type and transgenic mice. Hearts were excised, ventricular tissue was carefully dissected and immediately frozen in liquid nitrogen. Total RNA was extracted using TRIzol-Reagent® (Invitrogen, Karlsruhe, Germany) according to the manufacturer’s instructions, followed by reverse transcription (Superscript III, Invitrogen, Karlsruhe, Germany). Quantitative real-time PCR (RT-qPCR) was performed using an ABS 7500 Realtime PCR System (Thermo Fisher, Waltham, MA, USA) according to the manufacturer’s protocol. 96 well optical detection plates (Thermo Fisher, Waltham, MA, USA) were loaded to a total volume of 10 µl per well, consisting of 0.5 µl cDNA, 5 µl TaqMan Fast Universal Master Mix (Thermo Fisher, Waltham, MA, USA), and 6-carboxyfluorescein (FAM)-labeled pre-designed TaqMan primers (TaqMan Gene Expression Assays, Thermo Fisher, Waltham, MA, USA) specific to human and mouse HCN4 transcripts, HCN1, HCN2. qRT-PCR of apoptosis and Ca²⁺ processing related genes was performed using TaqMan assays for Casp3, tTG, Capn1, RyR2, Serca2a, Pln, Na⁺/K⁺-ATPase and NCX1. Ion channel transcripts were examined using TaqMan assays for SCN5A, Ca_v1.2, Ca_v1.3, Ca_v2.1, Ca_v3.1, K_ir2.1, K_v1.4, K_v1.5, K_v4.2, K_v7.1, hERG, and KChip2. Transcripts of myocardial cell growth marker genes and genes of the fetal gene program were determined with TaqMan assays for mTOR, GSK-3B, ANP, Calcineurin and β-MHC. Predesigned primers and probes detecting the housekeeping genes glyceraldehyde 3-phosphate dehydrogenase (GAPDH), hypoxanthine-guanine phosphoribosyltransferase 1 (HPRT1), and beta-actin (ACTB) were applied and normalization was carried out by a modified threshold cycle (CT) relative quantification method using GAPDH (Figs. 1b, 4a, 7d) or all three housekeeping genes (Fig. 4b–e), as published elsewhere⁵¹. All TaqMan primers used are listed in Supplementary Table 2. RT-qPCR reactions were performed in triplicate, and data are expressed as an average of triplicates.

Expression of apoptosis related genes was screened with Mouse Apoptosis RT² Profiler PCR Array (PAMM-012A-2, SABiosciences, Frederick, MD, USA) on an ABS 7500 Realtime PCR System and data was analysed using online software “RT² Profiler PCR Array Data Analysis” (version 3.5) on the manufacturer’s website http://pcrdataanalysis.sabiosciences.com/pcr/arrayanalysis.php.

Protein analysis

Protein immunodetection was performed by sodium dodecyl sulfate (SDS) gel electrophoresis. Tissue sections obtained from indicated mouse left and right ventricle samples were rinsed in phosphate buffered saline (PBS), rapidly frozen in liquid nitrogen and stored at −80 °C. Tissue samples were homogenized (TissueRuptor, QIAGEN, Hilden, Germany) in a radioimmunoprecipitation (RIPA) lysis buffer containing 50 mM Tris-HCl (pH 7.4), 0.5% NP-40, 0.25% sodium deoxycholate, 150 mM NaCl, 1 mM EDTA, 1 mM Na₃VO₄, 1 mM NaF, and protease inhibitors (Complete; Roche, Indianapolis, IN, USA). The protein concentration was determined using the bicinchoninic acid (BCA) protein assay (Thermo Scientific, Rockford, IL, USA), and equal amounts of protein were separated on SDS polyacrylamide gels. Nitrocellulose membranes were developed by sequential exposure to blocking reagent containing 3% bovine serum albumin and 5% dry milk. Primary antibodies were directed against HCN4 (1:200 diluted in 5% dry milk), NCX1 (1:250), Cav1.2 (1:500), Serca2 (1:1000), Phospholamban (1:500), phosphorylated Phospholamban Thr-17 (1:2000), phosphorylated Phospholamban Ser-16 (1:2000) and Calpain1 (1:2000). Appropriate HRP-conjugated secondary antibodies (anti-rabbit: 1:500, anti-mouse: 1:10,000) were used. Signals were developed using an enhanced chemiluminescence assay (ECL Western Blotting Reagents, GE Healthcare, Buckinghamshire, UK) and quantified with ImageJ 1.41 Software (National Institutes of Health, Bethesda, MD, USA). Protein content was normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) using anti-GAPDH primary antibodies (1:40,000) and corresponding secondary antibodies for digital quantification of optical density. For detailed information on primary and secondary antibodies used please refer to Supplementary Table 3. Western blot images used to compose result panels are available in Supplementary Fig. 5.

Isolation of cardiomyocytes

Cardiomyocytes for intracellular calcium measurements were obtained following the Liao & Jain protocol⁵². Briefly, the mice received 200 IU heparin i.p. prior to sacrifice. The thoracic chamber was opened and a cannula with perfusion solution was inserted from the atria. The heart was harvested and perfused in the Langendorff system with Perfusion Buffer (in mM: 135 NaCl, 4 KCl, 1 MgCl₂, 10 HEPES, 0.33 NaH₂PO₄, 10 glucose, 20 2,3-butanediones monoxime, 5 taurine, pH7.2 at 37 °C for 5 min), followed by Digestion Buffer treatment (0.3 mg/g body weight collagenase D (Roche, Indianapolis, IN, USA), 0.4 mg/g body weight collagenase B (Roche, Indianapolis, IN, USA), 0.05 mg/g body weight protease XIV (Sigma Aldrich, St. Louis, MO, USA) in 25 ml Perfusion Buffer) until the heart muscle was pale and some signs of extracellular matrix dissociation and for additional 5 min with Perfusion Buffer to stop the dissociation. The cardiomyocytes were mechanically dissociated in Transfer Buffer A in mM: 135 NaCl, 4 KCl, 1 MgCl₂, 10 HEPES, 0.33 NaH₂PO₄, 5.5 glucose, 10 2,3-butanediones monoxime, 5 mg/ml bovine serum albumin (Sigma Aldrich, St. Louis, MO, USA), pH 7.4 at 37 °C) and then plated on ECM-coated (Sigma Aldrich, St. Louis, MO, USA) petri dish (Zell-Kontakt, Nörten-Hardenberg. Germany). The extracellular calcium concentration was increased gradually in three consecutive steps from 0 mM (in Transfer Buffer A) to 0.06, 0.24 and 1.2 mM every 5 min. The cells where then incubated at 37 °C in 5% CO₂.

Cellular electrophysiology

I_f was recorded under voltage-clamp conditions (whole-cell configuration) in isolated mouse ventricular cardiomyocytes at room temperature (21–23 °C), essentially as published^53,54. In detail, patch-clamp measurements of mouse ventricular cardiomyocytes were performed in a solution containing (in mM): 137 NaCl, 25 KCl, 8 BaCl, 2 CaCl₂, 2 MnCl, 0.2 CdCl, 0.5 4-Aminopyridine, 5 HEPES, 10 Glucose, pH corrected to 7.4 using 1 M NaOH. The patch pipettes pulled from borosilicate glass (GB-150-8P, Science Products, Hofheim Germany) were generated on a DMZ Universal puller (Zeitz Instruments, Munich, Germany) and fire polished to give a final resistance of 2.5–4 MΩ. The pipette solution contained (in mM): 130 K-Aspartate, 5 Na₂ATP, 5 CaCl₂, 2 MgCl₂, 11 EGTA and 10 HEPES, pH corrected to 7.3 using 1 M KOH. Pharmacological agents were added to the bath solution to block L- type calcium currents (10 μM nisoldipine), the slow component of delayed rectifier K⁺ current (I_Ks, 10 μM chromanol), the rapid component of delayed rectifier K⁺ current (I_Kr, 10 μM E4031), ATP-dependent K⁺ current (I_KATP, 1 μM glibenclamid) and inward rectifier K⁺ currents (I_K1, 50 μM BaCl₂). I_f currents were normalized to cell capacitance which was obtained from the time constant of a current transient evoked by a 5 mV potential step at the beginning of each sweep.

Recordings of cardiac action potentials were carried out in an extracellular solution containing (in mM): 137 NaCl, 5.4 KCl, 1.8 CaCl₂, 0.5 MgCl₂, 10 HEPES, 5.5 glucose, and a pipiette solution containing (in mM): 120 K-Aspartate, 10 KCl, 5 NaCl, 2 MgCl₂, 10 HEPES. For quantification of AP duration and amplitude, cells were held in current-clamp mode and current was injected to achieve a membrane potential of −90 mV. If injected currents exceeded 200pA, recordings were discarded and not included in analysis. APs were evoked by 5 ms suprathreshold depolarizing current injection at 0.5, 1, and 2 Hz frequencies. Quantification of resting membrane potential, and recordings of spontaneous APs and afterdepolarizations were performed without current injection.

I_f and AP recordings were done with an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, USA), digitized at 20 kHz with a 1401 Power3 Analog/Digital Converter (CED, Cambridge, UK) stored on a hard drive, and analysed off-line with custom MATLAB routines (Mathworks, Natick, USA) software. Recordings with less than 10% leak current were considered for data analysis. No leak subtraction was performed during the experiments. Recordings were discarded if access resistance exceeded 20 MOhm or changed for more than 20% during the recording period.

Recording of calcium transients

The cardiomyocytes were loaded with 10 μM Fluo-4, AM (Molecular Probes, Leiden, The Netherlands) dissolved in Transfer Buffer B (in mM: 137 NaCl, 5.4 KCl, 1.8 CaCl₂, 0.5 MgCl₂, 10 HEPES, 5.5 glucose, pH 7.4 at 37 °C) for 30 min. For pharmacological treatments ivabradine (Molekula, Munich, Germany; cat. no. 89982651/155974-00-8) or ORM-10103 (Sigma-Aldrich, Germany; cat. no. SML0972) were dissolved in DMSO for stock solution. For working solution, both ivabradine or ORM-10103 were dissolved in Transfer Buffer B at 3 μM or at 10 µM, respectively, and incubated for 10 min prior to the Ca²⁺ recordings.

[Ca²⁺]_i transients were recorded using an Olympus OSP-3 System fluorescence microscope. Cardiomyocytes were single twitched electrically stimulated at 25 V for 10 ms at constant rate of 0.2 Hz. The fluorescence signal was obtained from cytosolic area and calibrated by a pinhole of 7.5 μm diameter, integrated in the photomultiplier, and A/D converted using PowerLab 4/35 and LabChart V7.0.

Out of 8–10 Ca²⁺ transients recorded, the least five were analyzed, avoiding the non-physiological Ca²⁺ transients due to time with no electrical stimulation between cell recordings. Ca²⁺ transients were eligible for regular analysis when lacking automaticity and showing regular [Ca²⁺]_I clearance under field stimulation at 0.2 Hz. Cells that exhibited pacemaker activity (defined as spontaneous beating rate < 0.2 Hz) or abnormal diastolic [Ca²⁺]_i decay (defined as D₉₀ > 2000ms) were excluded from regular analysis and evaluated in a separate record (please refer to Supplementary Fig. 2).

Calibration was performed at the end of each experiment in freshly loaded cells with Fluo-4, AM to transform voltage values into [Ca²⁺]_i⁵⁵. Briefly, cells were treated with high Ca²⁺ solution (in mM: 140 NaCl, 5 KCl, 1.2 KH₂PO₄, 1.2 MgCl₂, 4 CaCl₂, 20 HEPES, 0.005 ionomycin, 0.01 CPA, 5 caffeine, 1 ouabaine) to obtain Fmax values followed by application of zero-Ca²⁺ solution (in mM: 140 LiCl, 5 KCl, 1.2 KH₂PO₄, 1.2 MgCl₂, 20 HEPES, 4 EGTA, 0.005 ionomycin, 0.01 CPA, 5 caffeine, 1 ouabaine) to provide Fmin values. The Ca²⁺ signal (F) was then converted into [Ca²⁺]_i according to Grynkiewicz et al. 1985⁵⁶.

[Ca²⁺] = Kd(F-Fmin)/(Fmax-F), with Kd: apparent Ca²⁺ dissociation constant of the Fluo-4 of 345 nM, according to the manufacturer’s information.

Five biophysical parameters were analyzed: baseline (μM), peak area (area under the curve; μM*s), D₅₀ (duration of the transient at 50% of the peak; ms), time to peak (time required to reach the maximum from the baseline; ms), slope Ca²⁺ uptake time (time required to return to the base line from the peak to 5% of the baseline; nM/s).

Peri- and postpartal in vivo treatment with ivabradine

For treatment with the I_f blocker ivabradine (Molekula, Munich, Germany) timed-pregnant HCN4^tg/wt and wild type females were subcutaneously implanted with an osmotic mini pump (Alzet, Cupertino, CA, USA) at 18.5 days post-gestation. Ivabradine was delivered at a rate of 1.5 mg/kg/d for 3 weeks to the lactating female, and individual pumps were implanted in the pups from day 20 until 2 months postpartum.

Histochemical analysis and apoptosis assays

Hearts were dissected from transgenic and control mice at the age of 2 months, immediately frozen in isopentane on dry ice and stored at −80 °C. Serial coronal sections of 10 µm were produced for histochemical and immunohistochemical analyses.

For antibody staining, sections were fixed in 10% formaldehyde/PBS pH 7.2 (15 min), washed in PBS (3 × 5 min), permeabilized in 0.5% TritonX/PBS (30 min), blocked in 5% BSA/PBS and exposed for 24–48 h to the primary antibody in blocking solution at 4 °C. Primary antibodies used: rabbit polyclonal anti-HCN4 antibody (1:200; APC-052; Alomone labs, Jerusalem Israel); purified mouse monoclonal anti-caveolin 3 antibody (1:500; 610421; BD Transduction Laboratories, Palo Alto, CA, USA); anti-active caspase-3 (1:10; ab2302; Abcam, Cambridge, UK). Sections were washed in PBS (3 × 5 min) and incubated for 1 h at room temperature with a fluorescent labelled secondary antibody (1:500; MFP555 Goat anti-Rabbit IgG, MFP 488 goat anti-mouse IgG; MoBiTec, Goettingen, Germany) or anti-rabbit HRP-conjugated secondary antibody (Vectashield ABC Kit, Vector Labs, Burlingame, CA, USA). Sections were counterstained for 5 min with propidium iodide (1:500; ab14083; Abcam, Cambridge, UK) at room temperature.

Hematoxylin-eosin and Masson’s trichrome stainings were performed according to standard protocols. In detail, cardiac preparations were fixed in 4% formaldehyde, embedded in paraffin, cut to 6 µm thickness and mounted onto SuperFrost Plus slides (Gerhard Menzel GmbH, Braunschweig, Germany). Sections were then stained with hematoxylin-eosin to analyse cardiac histology or Masson’s trichrome dye to identify interstitial fibrosis. Blue staining following Masson’s trichrome application indicates fibrotic tissue. Cardiac fibrosis was quantified at 10-fold magnification. The extent of fibrosis was defined as the ratio between fibrotic regions and total tissue area [% fibrosis = (area exhibiting fibrosis/total area)*100]. The calculation was based on averaged measurements from 10 sections per animal (right ventricle, five; left ventricle, five). Bright-field images were digitally recorded on Axioplan2 microscope (Zeiss, Oberkochen, Germany) with attached 3CCD camera (Intas, Goettingen, Germany).

TUNEL assay was performed and quantified according to manufacturer’s protocol (DeadEnd™ Fluorometric TUNEL System, Promega). Confocal images of red and green immunofluorescence staining of heart tissue sections were acquired on a Leica confocal laser scanning unit TCS NT, which was coupled to a Leica DM IRB microscope. Acquisition of image series was performed using TCS NT (Leica, Heidelberg, Germany) software. All images were processed in ImageJ version 1.47 (NIH, Bethesda, MD, USA) software using standard quantification methods and standard ImageJ plug-ins.

Electron microscopy

Right and left ventricular wall tissue of HCN4^tg/wt and wild type mice was dissected, and prepared for microscopy⁵⁷. Briefly, cardiac tissue was fixed overnight at 4 °C in a calcium-free buffer (100 mM PIPES, pH 7.4 with 1.25% glutaraldehyde and 2% paraformaldehyde). After several washes with 100 mM PIPES, pH 7.0, samples were incubated for 2 h at room temperature in aqueous 2% osmium tetroxide and 1.5% potassium-hexacyanoferrate solution, washed in water for 3 h and incubated in water with 4% uranyl acetate overnight at 4 °C. Samples were dehydrated as described⁵⁷ and embedded in Epon medium. Ultra-sections were contrasted in uranyl acetate for up to 20 min and in lead citrate for 5 min. Sections were viewed on Philips EM400T (Philips IS, Eindhoven, The Netherlands) and images were documented on Image Plates and scanned with the Micro Imaging Plate Scanner (DITABIS, Pforzheim, Germany).

Statistical analysis

All experiments and primary analyses were blinded. Statistical analysis was performed using GraphPad Prism 5.0 (GraphPad Prism Inc. La Jolla, CA). Statistical significance and comparisons between samples were analyzed by a two-tailed unpaired Student’s t-test. Comparisons between multiple groups were performed using one-way ANOVA followed by Bonferroni (for calcium recordings), Tukey (for histochemical analysis) or Dunn’s (for action potential amplitudes) post-hoc tests. Differences were considered significant at a level P < 0.05. Data are presented as arithmetic mean ± s.e.m. or median ± maximum/minimum.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

The data that support the findings of this study are available from the article and the Supplementary Information Files, or from the corresponding author upon reasonable request. A reporting summary for this article is available as a Supplementary Information Files. Source data underlying the Figs. 1b and 1e, 2b–g and 2i–q, 3b and 3d, 4a–e, 5b–f, 6b–f, 7d and 7f, 8b–f, 9g, and Supplementary Figs. 1, 4, and Supplementary Table 1 are provided as a source data file.

Code availability

The MATLAB functions used for data analysis of the cellular electrophysiology experiments (I_f and AP recordings) have been deposited and are freely available at https://github.com/pgeschwill/NCOMMS-Paper. The code in the repository is associated with a GNU General Public License (GPL 3.0).

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Acknowledgements

We gratefully acknowledge the excellent technical work of Simone Bauer and Ulrike Mersdorf. This work was supported in parts by grants from the Medical Faculty of the University of Heidelberg and the German Cardiac Society (postdoc fellowship and research scholarship to P.Y.), from the German Heart Foundation (Kaltenbach scholarship to T.F.), from the German Cardiac Society and the Hengstberger Foundation (Klaus-Georg and Sigrid Hengstberger Scholarship to D.T.), from the Joachim Siebeneicher Foundation (to D.T.), from the Deutsche Forschungsgemeinschaft (SCHW 1611/1-1 to P.A.S. and TH 1120/8-1 to D.T.), from the Max-Planck-Society (TANDEM project to P.A.S. and M.K.O.) and from the German Centre for Cardiovascular Research (DZHK) (to S.N., C.S., A.P.S., H.E., D.T., H.A.K., P.A.S.). S.N. is recipient of a DZHK doctoral student scholarship. P.A.S. is recipient of the Heidelberg Research Center for Molecular Medicine (HRCMM) Senior Career Fellowship.

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These authors contributed equally: Pessah Yampolsky, Michael Koenen, Matias Mosqueira, Pascal Geschwill.

Authors and Affiliations

Department of Cardiology, Medical University Hospital Heidelberg, Im Neuenheimer Feld 410, 69120, Heidelberg, Germany
Pessah Yampolsky, Michael Koenen, Sebastian Nauck, Claudia Seyler, Thomas Fink, Mathieu Kruska, Dierk Thomas, Hugo A. Katus & Patrick A. Schweizer
Department of Molecular Neurology, Max-Planck-Institute for Medical Research, Jahnstrasse 29, 69120, Heidelberg, Germany
Pessah Yampolsky & Michael Koenen
Division of Medical Biophysics, Institute of Physiology and Pathophysiology, Heidelberg University, Im Neuenheimer Feld 326, 69120, Heidelberg, Germany
Matias Mosqueira, Monika Witzenberger & Rainer H. A. Fink
Division of Neuro- and Sensory Physiology, Institute of Physiology and Pathophysiology, Heidelberg University, Im Neuenheimer Feld 326, 69120, Heidelberg, Germany
Pascal Geschwill, Claus Bruehl & Andreas Draguhn
DZHK (German Centre for Cardiovascular Research) partner site Heidelberg/Mannheim, Im Neuenheimer Feld 410, 69120, Heidelberg, Germany
Sebastian Nauck, Claudia Seyler, Dierk Thomas, Hugo A. Katus & Patrick A. Schweizer
Department of Cellular and Integrative Physiology, University Medical Centre Hamburg-Eppendorf, Martinistrasse 52, 20246, Hamburg, Germany
Alexander P. Schwoerer & Heimo Ehmke
DZHK (German Centre for Cardiovascular Research) partner site Hamburg/Kiel/Lübeck, Martinistrasse 52, 20246, Hamburg, Germany
Alexander P. Schwoerer & Heimo Ehmke

Authors

Pessah Yampolsky
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Michael Koenen
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Matias Mosqueira
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Pascal Geschwill
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Sebastian Nauck
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Monika Witzenberger
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Claudia Seyler
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Thomas Fink
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Mathieu Kruska
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Claus Bruehl
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Alexander P. Schwoerer
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Heimo Ehmke
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Rainer H. A. Fink
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Andreas Draguhn
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Dierk Thomas
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Hugo A. Katus
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Patrick A. Schweizer
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Contributions

P.Y., M.K.O. and P.A.S. conceived and designed the study and the experiments. P.Y. and M.K.O. generated the transgenic mouse model. P.Y., S.N. and P.A.S. performed the molecular biology experiments. P.Y. and P.A.S. performed the histopathology and immunohistochemistry experiments of mouse hearts. P.G., C.B., C.S., A.P.S. and H.E. performed and analyzed the patch-clamp experiments. M.M., M.W. and R.F. carried out the calcium recordings of isolated cardiomyocytes. S.N., M.K., T.F. and P.A.S. performed the telemetric ECGs of mice and data analysis. S.N. and P.A.S. performed the mouse echocardiography and data analysis. H.A.K. provided administrative support and final approval of the paper. A.D. and D.T. supervised data analysis and provided expertise with paper writing. The paper was written principally by P.A.S.

Corresponding author

Correspondence to Patrick A. Schweizer.

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The authors declare no competing interests.

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Yampolsky, P., Koenen, M., Mosqueira, M. et al. Augmentation of myocardial I_f dysregulates calcium homeostasis and causes adverse cardiac remodeling. Nat Commun 10, 3295 (2019). https://doi.org/10.1038/s41467-019-11261-2

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Received: 19 June 2015
Accepted: 28 June 2019
Published: 23 July 2019
DOI: https://doi.org/10.1038/s41467-019-11261-2

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