Remodeling of repolarization and arrhythmia susceptibility in a myosin-binding protein C knockout mouse model

Ventricular myocytes isolated from the myosin-binding protein C knockout hypertrophic cardiomyopathy mouse model demonstrate decreased repolarizing K⁺ currents and action potential and QT interval prolongation, linking cellular repolarization abnormalities with arrhythmia susceptibility and the risk for sudden cardiac death in hypertrophic cardiomyopathy.

Abstract

Hypertrophic cardiomyopathy (HCM) is one of the most common genetic cardiac diseases and among the leading causes of sudden cardiac death (SCD) in the young. The cellular mechanisms leading to SCD in HCM are not well known. Prolongation of the action potential (AP) duration (APD) is a common feature predisposing hypertrophied hearts to SCD. Previous studies have explored the roles of inward Na⁺ and Ca²⁺ in the development of HCM, but the role of repolarizing K⁺ currents has not been defined. The objective of this study was to characterize the arrhythmogenic phenotype and cellular electrophysiological properties of mice with HCM, induced by myosin-binding protein C (MyBPC) knockout (KO), and to test the hypothesis that remodeling of repolarizing K⁺ currents causes APD prolongation in MyBPC KO myocytes. We demonstrated that MyBPC KO mice developed severe hypertrophy and cardiac dysfunction compared with wild-type (WT) control mice. Telemetric electrocardiographic recordings of awake mice revealed prolongation of the corrected QT interval in the KO compared with WT control mice, with overt ventricular arrhythmias. Whole cell current- and voltage-clamp experiments comparing KO with WT mice demonstrated ventricular myocyte hypertrophy, AP prolongation, and decreased repolarizing K⁺ currents. Quantitative RT-PCR analysis revealed decreased mRNA levels of several key K⁺ channel subunits. In conclusion, decrease in repolarizing K⁺ currents in MyBPC KO ventricular myocytes contributes to AP and corrected QT interval prolongation and could account for the arrhythmia susceptibility.

NEW & NOTEWORTHY Ventricular myocytes isolated from the myosin-binding protein C knockout hypertrophic cardiomyopathy mouse model demonstrate decreased repolarizing K⁺ currents and action potential and QT interval prolongation, linking cellular repolarization abnormalities with arrhythmia susceptibility and the risk for sudden cardiac death in hypertrophic cardiomyopathy.

hypertrophic cardiomyopathy (HCM) is one of the most common genetic cardiac diseases (1:500 in the general population) (28, 42) and among the leading causes of sudden cardiac death (SCD) in the young (1, 29). HCM is caused by mutations in sarcomere proteins. The altered sarcomere structure and/or function leads to pathological hypertrophy, abnormal heart function, and, importantly, arrhythmias and sudden death (14).

The pathophysiological link between abnormal sarcomere structure and arrhythmia is not clearly defined, likely varies with mutation type, and has been the subject of extensive research (11, 12). As in any other cardiac pathology leading to arrhythmia, two major components exist: initiation (triggering) of the abnormal action potential (AP) and propagation of the AP throughout the myocardium. Arrhythmias can result from a variety of mechanisms including reentry, triggered activity, and abnormal automaticity. Propagation of the arrhythmia is dependent on tissue and structural abnormalities leading to reentry circuits (39). HCM is usually characterized by cardiac fibrosis and cellular disarray, which have been shown to be important factors in reentry circuit formation and arrhythmia propagation (12) as well as a significant clinical risk factor for SCD (38).

There are also findings that support a role for abnormal cellular electrophysiological alterations in arrhythmia initiation in HCM; however, myocyte electrophysiological remodeling has not been clearly defined (11). A major electrocardiographic finding in HCM is prolongation of the QT interval, resulting from ventricular myocyte AP prolongation, in humans and animal models (8, 15, 20, 46). QT prolongation in HCM emphasizes the important arrhythmogenic role of abnormal cellular excitability in HCM.

Abnormal intracellular Ca²⁺ handling has been shown to play a key role in the pathophysiology of HCM (4, 12). Alterations in ion channel expression and function have also been established. Most previous studies concentrated on the arrhythmogenic role of Ca²⁺ and Na⁺ currents in HCM (9, 41, 44). Repolarizing K⁺ currents have not been investigated in detail in HCM and are the focus of the present work.

Myosin-binding protein C (MyBPC) mutations are commonly involved in HCM. It is estimated that ~40% of identified mutations in individuals with HCM occur in MyBPC (17). MyBPC knockout (KO) mice have been developed to study the bases of MyBPC mutation-induced HCM and functional abnormalities (16), with the majority of the work focused on structural and mechanical dysfunction (22, 26, 43).

The aim of the present study was to explore and define the cellular substrates for arrhythmias in MyBPC KO mice, focusing specifically on repolarizing K⁺ channels. We tested the hypothesis that K⁺ currents are decreased in MyBPC KO mice and that these changes contribute to AP prolongation.

The experiments performed establish that MyBPC KO mice demonstrate diverse ventricular arrhythmias, QT and AP prolongation, and decreased repolarizing K⁺ currents and K⁺ channel subunit mRNA transcript levels. Our results suggest that remodeling of repolarizing K⁺ currents contributes to abnormal cellular excitability and arrhythmia susceptibility in HCM.

METHODS

Experimental animals.

Homozygous cardiac MyBPC KO mice were kindly provided by the laboratory of Dr. Richard Moss (University of Wisconsin). These mice were generated as previously described and were maintained on an SV/129 background (16). Control wild-type (WT) SV/129 mice were obtained from Taconic Farms (Germantown, NY). Six- to nine-month-old, age-matched, male mice were used according to protocols approved by the Institutional Animal Care and Use Committee of the Lewis Katz School of Medicine at Temple University and the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health.

Telemetry and electrocardiography.

Implanted ETA-F20 telemetric transmitters (DSI, St. Paul, MN) were used to obtain continuous lead II electrocardiography (ECG) recordings with an AD Instruments PowerLab (Colorado Springs, CO) system on awake mice. Animals were housed in a room with 12:12-h light-dark cycle for 7 days to allow recovery from the surgery. Telemetry recordings were performed over a total of 24 h (equivalent to standard recording time of a Holter monitor in humans) net recording time (30 min of recording per each hour, over 48 h spanning over 2 consecutive days). The tracings were analyzed, looking for ventricular arrhythmia, by scrolling manually throughout the recordings. The data were stored and subsequently analyzed with the ECG module in the LabChart 8 software by two separate examiners who were blinded to the genetic information of the animals. To obtain ECG intervals, 2-min samples [as previously described (48)] from the telemetry tracings were analyzed for each mouse at noontime. Identical analyses were performed for each mouse at midnight to assess for daytime and nighttime variability. In each recording, the QT interval was determined as the time interval between the initiation of the QRS complex and the end of the T wave, defined as the time the T wave returned to the baseline. The measurement is shown in Fig. 3A. QT intervals were corrected for heart rate using the following formula: corrected QT (QT_c) = QT/(√RR/100) (30).

Fig. 3.

Telemetric electrocardiographic recordings. A and B: representative ECG waveforms from awake WT (A) and KO (B) animals marked with the different intervals that were measured. C: corrected QT (QT_c) interval analyzed from telemetric recordings of WT (n = 4) and KO (n = 4) animals at noontime and midnight. Mean ± SE values are plotted. **P < 0.01.

Echocardiography.

Anesthetized (0.5–1.5% isoflurane) mice underwent transthoracic echocardiography using a Vevo2100 ultrasound system (VisualSonics, Toronto, ON, Canada) with a 30-MHz MS-400 transducer.

Body temperature was maintained using a heated platform. Respiratory rates and ECGs were monitored throughout the study. For consistency, heart rate was maintained between 400 and 450 beats/min. Images were acquired in the short-axis B-mode and M-mode at the level of the left ventricular (LV) midpapillary muscles as well as the parasternal long axis for analysis of cardiac function and dimensions.

Tissue processing and histology.

Hearts were rinsed with PBS, weighed, and then perfusion fixed with 10% formalin and paraffin embedded for histology following a previously published protocol (10). Tissue blocks were sent to AML Laboratories (St. Augustine, FL) for sectioning and staining for Masson’s trichrome (Sigma-Aldrich, St. Louis, MO) to assess for tissue histology and collagen deposition. Myocyte cross-sectional area was measured in tissue sections stained with wheat germ agglutinin (ThermoFisher Scientific) and quantified with National Institutes of Health ImageJ software (http://rsbweb.nih.gov/ij/). At least 100 myocytes from 3 sections of the heart were analyzed per animal to assess myocyte cross-sectional area.

Myocyte isolation and cellular electrophysiology.

Adult mouse LV cardiomyocytes were isolated as previously described (27, 50), maintained at room temperature, and used within 12 h of isolation. For cellular electrophysiology experiments, myocytes were placed in a chamber mounted on an inverted Nikon microscope and perfused with normal physiological solution (Tyrode) containing 1 mM Ca²⁺. All experiments were done in a heated chamber at a temperature of 31°C as previously described (50). To measure Ca²⁺-insensitive K⁺ currents, myocytes were bathed in normal Tyrode solution containing 0.2 mM CdCl₂ to block Ca²⁺ currents. Cells were dialyzed for 10 min after patch rupture with pipette solution containing (in mM) 120 K aspartate, 20 KCl, 5 Na₂ATP, 1 MgCl₂, 10 HEPES, 1 CaCl₂, and 10 EGTA (pH with KOH to 7.2). EGTA was included in the pipette solution to minimize Ca²⁺-sensitive K⁺ currents and Na⁺/Ca²⁺ exchange current. The myocyte was ramped to −50 mV from a holding potential (V_hold) of −80 mV to inactivate Na⁺ channels and depolarized to different test potentials (−120 to 70 mV) in a 10-mV increment for 4,000 ms to record K⁺ currents. The bath solution was then switched to normal Tyrode solution containing 0.2 mM CdCl₂ and 2 mM 4-aminopyridine (4-AP). Once a stable effect of 4-AP was observed, the depolarization steps were applied again. The end of this 4-AP-insensitive K⁺ current was considered as the sustained K⁺ current (I_sus) and inwardly rectifying current (I_K1). The current sensitive to 4-AP contained mainly the transient outward K⁺ current (I_to) and slowly decaying K⁺ current (I_K,slow).

APs were measured with ruptured whole cell configuration with an Axoclamp 2B amplifier in the Bridge mode. The cell was dialyzed with the following pipette solution (in mM) for 5 min: 120 K aspartate, 20 KCl, 5 Na₂ATP, 1 MgCl₂, and 10 HEPES (pH with KOH to 7.2). The bath solution was normal Tyrode solution. APs were elicited by 2-ms depolarizations (5–15 mV) at 0.5 Hz for 5 min.

Analysis was performed using Clampfit 10.5 software offline.

RNA isolation and quantitative RT-PCR.

Total RNA was isolated from WT and KO mouse hearts using the mirVana Isolation Kit (Ambion) followed by DNase treatment to eliminate contaminating genomic DNA (Qiagen). Reverse transcription reaction was performed using the RT² First Strand Kit (Qiagen) according to the manufacturer’s instructions. Real-time quantitative PCR was performed using RT² SYBR Green ROX qPCR Mastermix (Qiagen). Data generated from the mouse heart samples were normalized to hypoxanthine guanine phosphoribosyl transferase (HPRT) expression. Fold changes of the transcript level in KO mice compared with WT mice were calculated with the $2^{-\mathrm{\Delta \Delta }{\text{C}}_{\text{t}}}$ method, where C_t is cycle threshold. The primer sets that were used in the experiments are shown in Table 1.

Table 1.

Sequence-specific primers used in SYBR green RT-PCR

Subunit	Gene	Forward Primer	Reverse
TASK1	KCNK3	5′-CCTTCTACTTCGCCATCACC-3′	5′-AACATGCAGAACACCTTGC-3′
Kvβ1	KCNAB1	5′-GACCTCTCTCCAATCGCTGA-3′	5′-CTGAATGGCACCAAGGTTTT-3′
	HPRT	5′-AGCCCCAAAATGGTTAAGGT-3′	5′-CAAGGGCATATCCAACAACA-3′

Commercial primers (Qiagen) were also used for the following targets: KCND2, KCND3, KCNIP2, KCNA4, KCNJ2, KCNJ12, KCNH2, KCNQ1, KCNE1, KCNA5, KCNB1, KCNK2, and KCNK5. Their sequences are proprietary protected. HPRT, hypoxanthine guanine phosphoribosyl transferase.

Data analysis.

Data are presented as means ± SE. Unpaired t-tests were performed to detect significance using Excel and GraphPad Prism 6 software. P < 0.05 was considered significant (*P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 in the figures).

RESULTS

MyBPC KO mice demonstrate severe hypertrophy, tissue fibrosis, and contractile dysfunction.

MyBPC KO mice developed severe cardiac hypertrophy (Fig. 1A) with an elevated heart weight-to-body weight ratio (WT: 5.2 ± 0.3 mg/g, n = 10; KO: 7.5 ± 0.4 mg/g, n = 9; P < 0.001; Fig. 1B). Masson’s trichrome-stained tissue sections at the mid-LV level showed gross chamber remodeling and overt interstitial fibrosis in KO mice, as evidenced by patchy blue staining (Fig. 1, A and C). Higher magnification revealed significant differences in cardiomyocyte morphology, with cellular hypertrophy and myofibrillar disarray in KO mice (Fig. 1C). Wheat germ agglutinin staining also revealed increased extracellular matrix consistent with tissue fibrosis and significantly increased myocyte cross-sectional area (WT: 233 ± 8.1 μm², N = 3 animals, n > 100 myocytes/animal; KO: 537.8 ± 46.6 μm², N = 4 animals, n > 100 myocytes/animal; P < 0.01; Fig. 1, D and E).

Fig. 1.

Heart size, morphology, histology, and myocyte cross-sectional area. A: representative cross sections of wild-type (WT) and knockout (KO) hearts at the mid left ventricular (LV) level stained with Masson’s trichrome. The viable myocardium is pink, and the fibrotic area is blue. B: heart weight-to-body weight (HW/BW) ratio in WT (n = 10) and KO (n = 10) animals. Mean ± SE values are plotted. ***P < 0.001. C: representative magnified cross sections at the mid-LV level of WT and KO hearts stained with Masson’s trichrome. Cardiomyocytes are pink, and their nuclei are dark blue. The fibrotic area is light blue. D: representative magnified cross sections at the mid-LV level of WT and KO hearts stained with wheat germ agglutinin (WGA) for the extracellular matrix and tissue fibrosis. E: myocyte cross-sectional area in WT (N = 3 animals, n > 100 myocytes/animal) and KO (N = 4 animals, n > 100 myocytes/animal) animals as analyzed from WGA-stained sections. Mean ± SE values are plotted. **P < 0.01.

Consistent with a previous report (16), echocardiography demonstrated severe hypertrophy (Fig. 2, A and B) with increased anterior wall diameter (WT: 1.06 ± 0.07 mm, n = 5; KO: 1.55 ± 0.16 mm, n = 5; P < 0.05; Fig. 2C) and posterior wall diameter (WT: 0.73 ± 0.03 mm, n = 5; KO: 1.26 ± 0.12 mm, n = 5; P < 0.01; Fig. 2D). Ejection fraction (WT: 64.4 ± 4.9%, n = 5; KO: 39.5 ± 2.8%, n = 5; P < 0.01; Fig. 2E) and fractional shortening (WT: 35.3 ± 3.6%, n = 5; KO: 19.2 ± 1.5%, n = 5; P < 0.01; Fig. 2F) were significantly reduced in KO animals. Chamber dilation was also noted with increased LV diastolic diameter (WT: 4.06 ± 0.16 mm, n = 5; KO: 4.77 ± 0.28 mm, n = 5; P = 0.06; Fig. 2G) and systolic diameter (WT: 2.65 ± 0.24 mm, n = 5; KO: 3.86 ± 0.28 mm, n = 5; P < 0.05; Fig. 2H) in KO mice.

Fig. 2.

Cardiac structure and function measured by echocardiography. A and B: representative parasternal short-axis M-mode (A) and parasternal long-axis images (B) demonstrating severe hypertrophy with increased wall thickness in KO hearts. C–H: LV anterior wall diastolic diameter (LVAWDd; C), posterior wall diastolic diameter (LVPWDd; D), ejection fraction (EF; E), fractional shortening (FS; F), LV diastolic diameter (LVDd; G), and LV systolic diameter (LVSd; H) in WT (n = 5) and KO (n = 5) animals. Mean ± SE values are plotted. *P < 0.05; **P < 0.01.

KO mice exhibit prolonged QT_c intervals and overt ventricular arrhythmias.

Telemetric electrocardiographic (lead II ECG) recordings were performed on awake mice, as detailed in methods. Representative ECG waveforms are shown in Fig. 3, A and B. KO mice demonstrated significant prolongation of the QT_c interval at noontime (WT: 36.7 ± 3.3 ms, n = 4; KO: 60.9 ± 3, n = 4; P < 0.01) and midnight (WT: 37.4 ± 2.7 ms, n = 4; KO: 60.8 ± 4.1, n = 4; P < 0.01) without significant daytime variability (Fig. 3C). Averaging noon and midnight values for all ECG intervals revealed that there were no significant differences in the durations of the RR, PR, and QRS intervals in KO mice compared with WT mice (Table 2).

Table 2.

ECG parameters observed in WT and KO mice

	WT	KO	P Value
RR interval, ms	103.2 ± 5.7	106.9 ± 2.5	0.6
PR interval, ms	33.6 ± 1.2	30.8 ± 2.3	0.37
QRS duration, ms	14.3 ± 1	18.3 ± 1.3	0.63
QT interval, ms	37.5 ± 2	62.9 ± 2.5	0.002*
Corrected QT interval, ms	37 ± 2	60.8 ± 2.3	0.001*

Values are means ± SE. P values of RR interval, PR interval, QRS duraction, QT interval, and corrected QT interval in awake wild-type (WT; n = 4) and knockout (KO; n = 4) animals, as analyzed from telemetric electrocardiographic recordings, are shown.

^*P < 0.01.

Telemetry recordings revealed that three of four WT animals demonstrated premature ventricular contractions (PVCs), with a total of eight for all WT animals, over the total net time of 24 h (Fig. 4, A and C, and representative trace in Fig. 4D). None of the four WT animals demonstrated high-grade ventricular ectopy. All 4 KO animals demonstrated ventricular ectopy with a total of 61 events over total net time of 24 h (Fig. 4A). PVCs were noted in all KO animals. Comparison of the average ventricular ectopy events between WT and KO animals did not reach significance (WT: 2 ± 0.8, n = 4; KO: 15 ± 12, n = 4; P = 0.32; Fig. 4B), since it was prominent in only one of the KO animals. This KO animal demonstrated diverse ventricular ectopy including PVCs, couplets, and nonsustained ventricular tachycardia (NSVT; 3–5 beats in a row; Fig. 4C). Representative tracings are shown in Fig. 4, E–I.

Fig. 4.

Arrhythmias in WT and KO animals. A: total ventricular ectopy events over the total net time of 24 h in WT (n = 4) and KO (n = 4) animals. B: average ventricular ectopy events over the total net time of 24 h in WT (n = 4) and KO (n = 4) animals. Mean ± SE values are plotted. NS, not significant. C: ventricular ectopy categories noted over the total net time of 24 h in WT (n = 4) and KO (n = 4) animals. D–I: representative tracings of ventricular arrhythmias noted in WT and KO animals, including premature ventricular contractions [PVCs; WT (D) and KO (E)], couplets (KO; F), three-beat nonsustained ventricular tachycardia (NSVT; KO; G), four-beat NSVT (KO; H), and five-beat NSVT (KO; I).

APs are prolonged and K⁺ currents are decreased in KO cardiomyocytes.

Whole cell current- and voltage-clamp experiments demonstrated that cardiomyocyte membrane capacitance was significantly increased, representing myocyte hypertrophy (WT: 234.4 ± 15.6 pF, N = 3 animals, n = 15 cells; KO: 469 ± 44.2 pF, N = 3 animals, n = 18 cells; P < 0.0001; Fig. 5A).

Fig. 5.

Myocyte membrane capacitance and current-clamp recordings of action potentials (APs). A: membrane capacitance in WT (N = 3 animals, n = 15 cells) and KO (N = 3 animals, n = 18 cells) myocytes. Mean ± SE values are plotted. ****P < 0.0001. B: representative AP waveforms in WT and KO myocytes. C: AP duration (APD) at 50% repolarization (APD₅₀), 75% repolarization (APD₇₅), and 90% repolarization (APD₉₀) in WT (n = 12) and KO (n = 11) myocytes. Mean ± SE values are plotted. *P < 0.05.

APs induced at 0.5 Hz were significantly prolonged in KO myocytes at all durations examined: AP duration (APD) at 50% repolarization (APD₅₀; WT: 17.7 ± 3.2 ms, n = 12; KO: 30.4 ± 4.5 ms, n = 11; P < 0.05), 75% repolarization (APD₇₅; WT: 32.6 ± 5.6 ms; KO: 69.2 ± 12.8 ms, n = 11; P < 0.05), and 90% repolarization (APD₉₀; WT: 74.7 ± 8.1 ms; KO: 125.7 ± 17.3 ms; P < 0.05; Fig. 5, B and C).

Repolarizing K⁺ current densities (normalized to cell capacitance) were decreased in KO myocytes. Representative current tracings are shown in Fig. 6, A and B. Currents were evoked in response to 4-s voltage steps to test potentials between −120 and 100 mV from a V_hold of −70 mV. Peak outward K⁺ currents (I_K peak) measured in WT (n = 15) and KO (n = 14) myocytes demonstrated a 33–39% decrease in KO myocytes (Fig. 6C).

Fig. 6.

Voltage-clamp recordings of repolarizing K⁺ currents. A and B: representative whole cell K⁺ currents (normalized to membrane capacitance) recorded from WT (A) and KO (B) animals. Currents were evoked in response to (4-s) voltage steps to test potentials between −100 and 60 mV from a holding potential (V_hold) of −70 mV. C: peak outward K⁺ currents (normalized to membrane capacitance) recorded from WT (n = 15) and KO (n = 14) myocytes. Mean ± SE values are plotted. *P < 0.05; **P < 0.01; ***P < 0.001. D and E: representative 4-aminopyridine (4-AP)-insensitive K⁺ currents (normalized to membrane capacitance) recorded from WT (D) and KO (E) animals. Currents were evoked in response to (4-s) voltage steps to test potentials between −100 and 60 mV from a V_hold of −70 mV. F: peak 4-AP-insensitive K⁺ currents (normalized to membrane capacitance) recorded from WT (n = 13) and KO (n = 13) myocytes. Mean ± SE values are plotted. *P < 0.05; **P < 0.01. G and H: representative 4-AP-sensitive K⁺ currents (normalized to membrane capacitance) recorded from of WT (G) and KO (H) animals. Currents were evoked in response to (4-s) voltage steps to test potentials between −20 and 60 mV from a V_hold of −70 mV. I: peak 4-AP-sensitive K⁺ currents (normalized to membrane capacitance) recorded from WT (n = 9) and KO (n = 10) myocytes. Mean ± SE values are plotted. *P < 0.05; **P < 0.01.

Myocytes were then exposed to 4-AP. There was a significant reduction of the 4-AP-insensitive sustained outward plateau current in KO myocytes. Representative current tracings are shown in Fig. 6, D and E. These currents, evoked in response to voltage steps to test potentials between 20 and 100 mV from a V_hold of −70 mV, in WT (n = 13) and KO (n = 13) myocytes, demonstrated a 21–23% reduction (Fig. 6F). 4-AP-insensitive I_K1 was also decreased by 16–30%, (voltage steps to test potentials between −120 and 100 mV; Fig. 6F).

The most significant differences in KO myocytes were in the 4-AP-sensitive currents, which include I_to and I_K,slow1. Representative current tracings are shown in Fig. 6, G and H. These currents, measured in response to voltage steps to test potentials between 0 and 60 mV from a V_hold of −70 mV in WT (n = 9) and KO (n = 10) myocytes, were significantly reduced (46–61%) in KO myocytes (Fig. 6I).

K⁺ channel mRNA levels are decreased in KO hearts.

Quantitative RT-PCR analysis revealed decreased mRNA levels of several K⁺ channel transcripts in KO mice compared with WT mice. Transcripts of I_to, the fast and slow components of I_to (I_to,fast and I_to,slow, respectively), demonstrated the most significant reduction (Fig. 7A): KCND2, coding for the K_v4.2 subunit, was down by 3.1-fold to a relative level of 0.32 ± 0.05 (P < 0.001); KCND3, coding for the K_v4.3 subunit, was down by 1.7-fold to a relative level to 0.58 ± 0.07 (P < 0.01); KCNIP2, coding for the regulatory subunit KChIP2, was down by 1.6-fold to a relative level of 0.62 ± 0.14 (P < 0.05); and KCNA4, coding for K_v1.4 subunit, was down by 2.4-fold to a relative level of 0.41 ± 0.14 (P < 0.01). The I_K1 transcript KCNJ2, coding for Kir2.1, was down by 2.4-fold to a relative level of 0.41 ± 0.05 (P < 0.05; Fig. 7B). Slow delayed rectifier K⁺ current (I_Ks) transcript KCNE1, coding for the minK regulatory subunit, was also significantly reduced by 2.3-fold to a relative level of to 0.43 ± 0.12 (P < 0.05; Fig. 7C). Other transcripts coding for additional subunits of I_to (Fig. 7A), inward rectifier (Fig. 7B), delayed rectifier (Fig. 7C), and two-pore domain (K₂P) currents (Fig. 7D) were tested, and although some demonstrated a trend for decreased levels, the differences did not reach statistical significance.

Fig. 7.

Transcript expression of K⁺ channel subunits: mRNA relative transcript levels as measured by quantitative RT-PCR in WT (n = 7) and KO (n = 7) animals. Mean ± SE values are plotted. *P < 0.05; **P < 0.01; ***P < 0.001. A: transient outward voltage-gated K⁺ currents [I_to,fast: KCND2 (K_v4.2), KCND3 (K_v4.3), KCNIP2 (KChIP2), and KCNAB1 (K_vβ1); I_to,slow: KCNA4 (K_v1.4)]. B: inward rectifier current [I_K1: KCNJ2 (Kir2.1) and KCNJ12 (Kir2.1)]. C: delayed rectifier voltage-gated K⁺ currents [I_Kr: KCNH2 (hERG); I_Ks: KCNQ1 (K_vLQT1) and KCNE1 (minK); I_K,slow1: KCNA5 (K_v1.5); I_K,slow2: KCNB1 (K_v2.1)]. D: two-pore domain current [I_ss: KCNK2 (TREK₁), KCNK3 (TASK₁), and KCNK5 (TASK₂)].

DISCUSSION

Why HCM patients die suddenly was the question that was the driving force for this study. This question has been the center of extensive investigation. Clinical and basic studies have supported several mechanisms: tissue fibrosis and cellular disarray causing microreentry circuits, coronary bridging and vascular instability leading to ischemia, remodeling of Ca²⁺ currents, abnormal intracellular Ca²⁺ handling, and remodeling of late Na⁺ currents (4, 9, 11, 12, 23, 44). The idea that repolarizing K⁺ currents might also be involved in electrical remodeling in HCM has not been well studied and is the topic of the present study.

Remodeling of repolarization in HCM.

The main finding of our study is the significant remodeling of outward repolarizing currents in MyBPC KO ventricular myocytes, an HCM model system (Fig. 6). To the best of our knowledge, this is the first HCM animal model in general, and MyBPC specifically, in which such a change has been described.

Supporting our findings, remodeling of K⁺ currents has been reported in an in vitro study of human cardiomyocytes isolated from HCM patients (9). Although mouse cardiac electrophysiology is inherently different from human, an animal model enables the controlled and dynamic exploration of in vivo pathophysiology, diagnostic studies, and potential therapeutic intervention (11).

The basis for this study was the clinical observation of QT prolongation and repolarization abnormalities in HCM patients (15, 20). This phenotype was evident in the animal model (Fig. 3), which also recapitulated the structural, histological (Fig. 1), echocardiographic (Fig. 2), and arrhythmogenic substrate (Fig. 4). Of note, QT prolongation was noted in HCM patients with mutations in several other HCM genes (myosin heavy chain 7, thin filament, Z disk) in addition to MyBPC (20) as well as in a different animal model [myosin heavy chain and troponin-I double mutation HCM mouse model (46)], suggesting that our findings are likely not gene/model specific.

Interestingly, heterozygous animals of the same MyBPC model that demonstrate only mild contractile dysfunction without ventricular hypertrophy exhibited prolonged QT_c interval on surface ECG, with intermediate values between WT and our homozygous KO animals (8). Although most human patients with HCM carry heterozygous mutations, there are patients who harbor homozygous, compound heterozygous (2 different mutations in the same gene), and double (such as MyBPC together with MYH7) sarcomere gene mutations and usually are characterized with a severe phenotype (3, 31). This underscores the relevance of exploring our homozygous animals with the pronounced phenotype.

Arrhythmia susceptibility in KO mice.

SCD in HCM is mediated primarily by ventricular fibrillation, usually preceded by PVCs and ventricular tachycardia (37). In our study, one of the four KO mice exhibited diverse ventricular ectopy including frequent NSVT. Untriggered arrhythmias at rest are not common in HCM. The proportion that was noted in our study (25%) is similar to the proportion described in other animal/human studies (2, 6, 19, 32). The ventricular arrhythmias that have been previously described in several other HCM mouse models were mainly attributed to the structural arrhythmogenic substrate, remodeling of Ca²⁺ currents, abnormal Ca²⁺ handling, and hypertrophy per se (5, 6, 13, 18, 24, 40, 47). Although electrocardiographic repolarization abnormalities and QT prolongation were also noted in these studies, K⁺ currents were not explored. We suggest that reductions in repolarizing K⁺ currents contribute to APD and QT prolongation and the associated ventricular arrhythmias.

Molecular components underlying the downregulation of ionic currents.

Correlating with the cellular electrophysiological results, we found that various mRNA transcripts of K⁺ channel subunits were decreased (Fig. 7). This reduction was noted in each of the K⁺ current components but was most prominent in I_to (Fig. 7A). I_to is the dominant repolarizing current in the mouse affecting APD and shape (33, 35).

It is well established that reduced K⁺ currents in general, and I_to specifically, are commonly observed in mouse models with cardiac hypertrophy and failure regardless of the underlying cause (25, 49). However, physiological cardiac hypertrophy associated with training resulted in the opposite outcome, with upregulation of repolarizing currents (48). These results indicate that changes in repolarizing K⁺ currents are not specific to the hypertrophy per se but are related to the physiological or pathological processes causing hypertrophy.

Implications.

In addition to sarcomeric cardiomyopathies, these results may also be relevant to other forms of genetic malformation syndromes, neuromuscular disorders, and metabolic cardiomyopathies. These cardiomyopathies usually present in early childhood, harbor poor prognosis, and are characterized by severe hypertrophy, electrocardiographic and repolarization abnormalities, arrhythmias, and sudden death. There is a paucity of investigation in this field, and further research into the pathophysiology of these orphan diseases is clearly needed.

There is a growing interest in investigating the interaction between membrane ion channels in general, and K⁺ channels in particular, and the intracellular cytoskeleton and sarcomere (35, 36). It has been suggested that sarcomeric cardiomyopathies share a final common pathway involving the cell membrane and the intracellular components (7, 45). Our intriguing results suggest a link between both ends, leading to an abnormal excitation-contraction coupling, contractile dysfunction, and arrhythmias.

Study limitations.

As in any other studies exploring cellular electrophysiology in mice, one should be aware of the limitation originating from the major differences in the repolarizing currents between mice and larger mammals, with dominance of I_to effecting different APD and shape in mice. The significance of our findings should be considered in this context. Although I_to has an important role in larger mammals, including humans, controlling the initial phase of AP repolarization and determining the driving force for the Ca²⁺ entry through L-type Ca²⁺ channels (36), it is important to mention that larger mammals have longer APDs and that APD is less dependent on changes in I_to. This limitation, however, is inherent to the rodent experimental system and does not limit the capability to study disease mechanisms in highly controlled in vivo models (21, 34).

Conclusions.

In summary, MyBPC KO HCM mice demonstrate an overt phenotype with severe cardiac hypertrophy, contractile dysfunction, and arrhythmia susceptibility. Electrocardiographic recordings revealed prolongation of the QT_c interval consistent with repolarization abnormality and diverse ventricular arrhythmias. Cellular electrophysiological recordings demonstrated reduced K⁺ currents and AP prolongation. Furthermore, quantitative RT-PCR experiments established reduced K⁺ channel transcript levels. These results suggest that remodeling of repolarizing K⁺ currents may contribute to the arrhythmia susceptibility and could be the target for novel therapies for the prevention of SCD in HCM.

GRANTS

A. Toib received a research grant from the Philadelphia Health and Education Corporation Fund. X. Chen received National Heart, Lung, and Blood Institute (NHLBI) Grant HL-88243. S. Mohsin received a Scientist Development Grant from the American Heart Association. S. R. Houser received NHLBI Grants HL-89312, HL-91799, and HL-33921.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

A.T. and S.R.H. conceived and designed research; A.T., C.Z., G.B., X.Z., M.W., H.K., E.F., R.M.B., D.M.T., S.H., and X.C. performed experiments; A.T., G.B., M.W., Y.Y., C.D.T., H.K., T.E.S., N.Z., D.M.T., S.H., and P.G. analyzed data; A.T., C.Z., G.B., X.Z., M.W., Y.Y., C.D.T., H.K., N.Z., D.M.T., S.H., P.G., X.C., S.M., and S.R.H. interpreted results of experiments; A.T., C.D.T., and T.E.S. prepared figures; A.T. and S.M. drafted manuscript; A.T., G.B., M.W., C.D.T., H.K., E.F., R.M.B., D.M.T., S.H., P.G., X.C., S.M., and S.R.H. edited and revised manuscript; A.T., C.Z., G.B., X.Z., M.W., Y.Y., C.D.T., H.K., T.E.S., E.F., R.M.B., N.Z., D.M.T., S.H., P.G., X.C., S.M., and S.R.H. approved final version of manuscript.