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Am J Physiol Heart Circ Physiol 292: H560-H571, 2007. First published August 18, 2006; doi:10.1152/ajpheart.00616.2006
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Electrical remodeling in a canine model of ischemic cardiomyopathy

Xian-Sheng Liu,1 Min Jiang,1 Mei Zhang,1 Daniel Tang,2 Henry F. Clemo,3 Robert S. D. Higgins,2 and Gea-Ny Tseng1,3

1Department of Physiology, 2Department of Surgery, 3Division of Cardiology, Department of Internal Medicine, Medical College of Virginia, Virginia Commonwealth University, Richmond, Virginia

Submitted 10 June 2006 ; accepted in final form 10 August 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The nature of electrical remodeling in a canine model of ischemic cardiomyopathy (ICM; induced by repetitive intracoronary microembolizations) that exhibits spontaneous ventricular tachycardia is not entirely clear. We used the patch-clamp technique to record action potentials and ionic currents of left ventricular myocytes isolated from the region affected by microembolizations. We also used the immunoblot technique to examine channel subunit expression in adjacent affected tissue. Ventricular myocytes and tissue isolated from the corresponding region of normal hearts served as control. ICM myocytes had prolonged action potential duration (APD) and more pronounced APD dispersion. Slow delayed rectifier current (IKs) was reduced at voltages positive to 0 mV, along with a negative shift in its voltage dependence of activation. Immunoblots showed that there was no change in KCNQ1.1 (IKs pore-forming or {alpha}-subunit), but KCNE1 (IKs auxiliary or beta-subunit) was reduced, and KCNQ1.2 (a truncated KCNQ1 splice variant with a dominant-negative effect on IKs) was increased. Transient outward current (Ito) was reduced, along with an acceleration of the slow phase of recovery from inactivation. Immunoblots showed that there was no change in Kv4.3 ({alpha}-subunit of fast-recovering Ito component), but KChIP2 (beta-subunit of fast-recovering component) and Kv1.4 ({alpha}-subunit of slow-recovering component) were reduced. Inward rectifier current was reduced. L-type Ca current was unaltered. The immunoblot data provide mechanistic insights into the observed changes in current amplitude and gating kinetics of IKs and Ito. We suggest that these changes, along with the decrease in inward rectifier current, contribute to APD prolongation in ICM hearts.

heart failure; arrhythmia; voltage clamp; oocyte expression

HEART FAILURE (HF) IS PREVALENT in the Western world and will become more so as the population ages. About 50% of HF patients succumb to sudden death, most of which likely results from malignant ventricular arrhythmias (54). Although implantable cardioverter/defibrillators can save lives, the high costs prevent them from being available to the majority of HF patients. Pharmacological interventions are the practical approach. A prerequisite for the design of preventive medicine is the knowledge about arrhythmogenic mechanisms in HF patients, based on which potential targets can be identified and explored.

There have been intensive research efforts to study structural and functional changes (remodeling) that take place in failing hearts, using animal models as well as patient heart samples (2, 16, 31). In terms of arrhythmogenic mechanisms, two common changes have been identified in failing hearts of animals and humans of different etiologies: 1) prolongation of action potential duration (APD), and 2) abnormalities in intracellular Ca (Cai) handling (2, 5, 54). The latter include slower release as well as slower reuptake of Cai by the sarcoplasmic reticulum and elevation of diastolic Cai level (2, 5, 29, 54). APD prolongation, in conjunction with abnormalities in Cai handling, increases the risk for delayed and early afterdepolarizations, which can lead to triggered arrhythmias (16). Furthermore, to maintain proper action potential configurations and durations in different regions of the heart, cardiac myocytes need to maintain a delicate balance between inward and outward currents in a region-specific manner (34). A disturbance in the repolarization process can often lead to an excessive dispersion of APD. When this occurs, especially in conjunction with a slowing of impulse conduction [due to partial depolarization of the resting membrane potential (RMP), interstitial fibrosis, or dysfunctional gap junction channels], the stage is set for reentrant arrhythmias (16). Therefore, preventing or correcting ventricular APD prolongation is a key factor in the design of pharmacological interventions to protect HF patients at risk for ventricular arrhythmias.

It is not entirely clear what causes APD prolongation in failing hearts (16). Although a reduction in the transient outward current (Ito) is almost a universal finding in ventricular myocytes from HF animals and patients (2, 5, 31, 54), a decrease in Ito per se does not necessarily lead to APD prolongation (except in rodents) (14, 22, 33). In large animals and humans, Ito is directly responsible for phase 1 repolarization only. By setting the rate of phase 1 repolarization and the voltage reached at the end of phase 1, Ito can indirectly affect the degree of activation and amplitudes of other plateau currents, notably the L-type Ca channel current (ICaL) (14). Inhibiting Ito by 4-aminopyridine can lead to APD prolongation (13), little change (12), or shortening (23), although the specificity of 4-aminopyridine is an issue (32, 42). Computer simulations suggest that reducing Ito either has a minimal effect on APD (40), or shortens APD by shifting phase 1 voltage in the positive direction and secondarily reducing the ICaL amplitude (14). Computer simulation of action potentials and currents in human HF suggests that a putative decrease in the rapid delayed rectifier current (IKr) and/or the slow delayed rectifier current (IKs) is necessary to account for APD prolongation (40). However, data on IKr and IKs in failing hearts are rare from animal studies (25) and even more so for HF patients (24).

Sabbah et al. (48) developed a canine HF model due to ischemic cardiomyopathy (ICM) induced by repetitive intracoronary microembolizations. This model exhibits features resembling those of clinical ICM: a progressive decline of left ventricular (LV) systolic function, interstitial fibrosis, an increase in serum catecholamines, and, importantly for the discussion of arrhythmogenesis in HF, spontaneous ventricular tachycardia (VT) (39, 46). At the cellular level, ventricular myocytes from the ICM dog hearts manifest APD prolongation (59). It has been shown that one contributing factor to APD prolongation in these ICM cells is an increase in the long-lasting component of TTX-sensitive Na channel current (59, 62). However, there have been no investigations into other ion channels involved in determining the action potential configuration and duration. These include IKr, IKs, Ito, inward rectifier current (IK1), and ICaL.

We have followed the protocol developed by Sabbah et al. to create the canine model of ICM and use it to probe the cause(s) for APD prolongation. Previously, our laboratory has reported the surprising finding that ventricular myocytes from these ICM dogs had a larger IKr current density than myocytes from the corresponding region of control dogs (21). This may be a mechanism to protect these hearts from further excessive APD prolongation. At the molecular level, there was no marked change in the protein level of the IKr pore-forming component, ERG1 (21, 50). On the other hand, there was a marked reduction in the immunoreactivity to an antibody (Ab) (Alomone) specific for KCNE2, the putative auxiliary subunit of IKr (1, 21). We attributed the increase of IKr in ICM myocytes to a relief of KCNE2's suppressing effect on currents through the ERG1 channel (30). In the present study, we report changes in the other currents involved in shaping the action potential plateau phase (IKs, Ito, IK1, and ICaL) in LV myocytes from these ICM dogs. We also present immunoblot data that provide insights into possible mechanisms underlying the observed changes in current amplitude and gating kinetics.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Canine Intracoronary Microembolizations

The animal protocol was reviewed and approved by Institutional Animal Care and Use Committee of Virginia Commonwealth University. The procedure was similar to that developed by Sabbah et al. (48). Briefly, five mongrel dogs (male, 23–27 kg) were subjected to intracoronary injections of polystyrene latex microspheres (diameter 77–102 µm, Polysciences, Warrington, PA). Beads (2–4 x 104) in 0.5–1 ml volume were injected via cardiac catheterization under general anesthesia and sterile conditions. Each dog received a total of two to four injections, alternating between left anterior descending coronary artery and left first circumflex coronary artery once every 3–4 wk until the LV ejection fraction (LVEF) dropped to ≤35%.

Approximately 3 months after the final microsphere injection, dogs were subjected to hemodynamic measurements. The hearts were harvested, and a wedge of LV free wall (vascular bed of the first circumflex coronary artery) was dissected out for single-cell isolation (see below). A transmural section of anterior LV free wall containing interstitial fibrosis was dissected for histological analysis. Tissue chunks from LV apex were snap frozen in liquid nitrogen and stored at –80°C for immunoblot analysis. Corresponding areas of five normal canine hearts were dissected in the same manner for control experiments. The distance between the origin of cell isolation and regions for histology or immunoblot analysis ranged from 0.3 to 1 cm or 0.5 to 2 cm, respectively.

Single-Cell Preparation

This procedure was modified from a method described before (56). The first circumflex coronary artery was cannulated. Epicardial and intramural vascular openings were ligated to ensure good perfusion. The tissue was mounted on a Langendorff apparatus and perfused with warm (36°C), oxygenated Tyrode solution (composition given below) for 2–4 min. The perfusate was then switched to a nominally Ca-free Tyrode containing bovine serum albumin (BSA, 0.5 mg/ml) for 6–7 min. Then the tissue was perfused with a collagenase-containing solution (type II, Worthington, 0.5 mg/ml) for 25–30 min until the tissue was softened. After collagenase digestion, the mid-myocardial layer (~0.2 cm away from both the epicardial and endocardial surfaces) was isolated and minced. The small tissue chunks were returned to fresh collagenase-containing solution and gently shaken in a 36°C water bath for 10–15 min. The cell suspension was collected, and cells formed sediment by gravity. The remaining tissue chunks were subjected to further trituration in fresh collagenase solution for four to six more times. Cell sediments were resuspended in a Tyrode solution containing (in mM) 0.5 Ca, 10 taurine, 10 mannitol, and 10 pyruvate, and stored at room temperature. Some cells were plated in serum-free DMEM and stored in a 36°C 5% CO2 moist incubator. Cells stored under both conditions were used within 10 h after isolation. No differences in terms of membrane action potential or currents were noticed between cells stored under these two conditions, and the data were pooled.

Patch-Clamp Recordings

Cells were plated on a poly-L-lysine coated coverslip placed in a tissue bath mounted on the stage of an inverted Nikon microscope and continuously superfused with bath solution maintained at 33–34°C. Membrane voltage and currents were recorded using the whole-cell variant of the patch-clamp technique, with AxoPatch 1D or AxoPatch200 in current-clamp and voltage-clamp mode, respectively. The pipette had a tip resistance of 1–2 M{Omega}, and 95% of series resistance was compensated. Experiments were controlled by pClamp6. Currents were low-pass filtered at 1 kHz (Frequency Devices), and data were stored for offline analysis.

The pipette solution contained (in mM) 125 potassium-aspartate, 20 KCl, 10 potassium-ATP, 10 HEPES, 10 EGTA, and 1 Mg, pH 7.3. The bath solution was either normal Tyrode or Na- and Ca-free Tyrode. The former contained (in mM): 147 NaCl, 4 KCl, 5 HEPES, 5.5 dextrose, 2 CaCl2, and 0.5 MgCl2, pH 7.3. The latter was made by using choline-chloride to substitute for NaCl, and MgCl2 to substitute for CaCl2.

To maximize data collection and minimize interference due to overlapping currents and issue of time-dependent IKs run down, all of our patch-clamp recordings followed a similar time course using the same solutions and voltage-clamp protocols. Three minutes after forming the whole cell recording configuration when the intracellular dialysis with the pipette solution reached equilibrium, we recorded cell capacitance and adjusted series resistance compensation. Then, while in normal Tyrode solution, we recorded action potentials (cycle length 1 s, trains of 30–120 pulses until the action potential morphology and duration stabilized), ICaL, IK1, and Ito. The bath solution was then switched to Na- and Ca-free solution, and we recorded the rectifier current (typically 15–20 min after the beginning of whole cell dialysis). Then 1 µM dofetilide was added to the bath solution, and dofetilide-insensitive IKs was recorded between 22 and 25 min after the beginning of whole cell dialysis.

Oocyte Expression Experiments

The following clones were studied: rat Kv4.3 and Kv1.4, human KCNQ1.1, KCNQ1.2, and KCNE1. The procedures of in vitro transcription (mMessage mMachine, Ambion, TX) and cRNA quantification (densitometry with ChemiImager model 4400, Alpha-Innotech) have been described previously (27). The cRNA concentrations were calculated based on band intensities measured by densitometry, using cRNA size markers as a reference. For coinjection, the cRNAs were mixed before injection, with molar ratio calculated by dividing the nanogram amounts of cRNAs injected by the cRNA sizes.

Oocytes were isolated as described before (57) and incubated in an ND96-based medium (in mM: 96 NaCl, 2 KCl, 2.5 pyruvate, 1.8 CaCl2, 5 HEPES, 0.5 MgCl2, pH 7.5), supplemented with 4% horse serum and penicillin/streptomycin at 16°C. Two to six hours after isolation, oocytes were injected with cRNA(s) using a Drummond digital microdispenser. Oocytes were incubated in the above medium at 16°C and studied 3–5 days after isolation. Whole oocyte membrane currents were recorded using the "2-cushion pipette" voltage-clamp method (51). During recordings, the oocyte was continuously superfused with a low-Cl ND96 solution (replacing Cl– in ND96 with methanesulfonate) to reduce interference from endogenous Cl channels. Voltage clamp was done at room temperature (24–26°C) with OC-725B or OC-725C amplifier (Warner Instruments). Voltage-clamp protocol generation and data acquisition were controlled by pClamp5.5 via a 12-bit digital-to-analog and analog-to-digital converter (DMA, Axon Instruments). Current data were low-pass filtered at 1 kHz (Frequency Devices) and stored on disks for offline analysis.

Data Analysis

For both patch-clamp recordings from canine ventricular myocytes and two-cushion pipette voltage-clamp recordings from oocytes, the voltage-clamp protocols and methods of data analysis are described in the figure legends. The following software was used for data analysis: pClamp6 or 8, EXCEL (Microsoft), SigmaPlot, SigmaStat, and PeakFit (SPSS). Mean values with standard errors are reported here, with numbers of cells or samples studied listed in figures.

Biochemistry

Membrane-enriched fraction from canine myocardium or oocytes was made as described previously (20, 21). Protein concentrations were measured using a Pierce kit with BSA as control. In vitro translation of KCNE1 was performed using a rabbit reticulocyte lysate system (Promega) in the presence of canine pancreatic microsomes, according to the manufacturer's instructions. To deglycosylate translated KCNE1, 1% Nonidet P-40, complete protease inhibitor cocktail, and 2 units of N-glycosidase F (Boehringer) were added to one-half (25 µl) of the in vitro translation product. The mixture was incubated at 37°C for 24 h. The reaction was stopped by adding sample buffer and boiling for 2 min. A parallel control using the remaining one-half of the translation product was processed in the same manner without N-glycosidase F in the buffer.

Proteins were fractionated on 4–20% gradient gels, blotted to polyvinylidene difluoride membranes, blocked by BSA or dry milk, and incubated with primary antibodies at 4°C overnight. The membrane was then washed and incubated with a secondary Ab. Immunoreactivity was visualized by enhanced chemiluminescence (Amersham). Residual proteins in the gels were stained with Coomassie blue (CB), and densitometry of CB stain was used to check for loading variations. For quantification of subunit expression in control vs. failing dog hearts, the subunit-specific bands were background subtracted, divided by the CB stain of the same lanes, and normalized by the mean band intensities of control hearts. The following commercial primary Abs were used: Kv4.3 (Alomone), Kv1.4 (Upstate Technology), KChIP2 (Affinity Bioreagents), and KCNQ1 (Santa Cruz Biotech). The KCNE1 Ab was a generous gift from Drs. Amber Pond and Jeanne M. Nerbonne. This Ab was raised against a peptide, [C]RVLESFRACYVI, corresponding to aa 98–119 of mouse KCNE1, which is highly conserved across species.

Histology and Immunocytochemistry

Tissue blocks from LV free wall adjacent to where cells were isolated were embedded in parafin and cut in 10-µm thin sections. The thin sections were stained by hematoxylin and eosin. Isolated myocytes (same as those for patch-clamp recordings) were fixed in 4% paraformaldehyde overnight on poly-L-lysine coated coverslips. The cells were then stored in a blocking buffer (goat serum 4% in PBS) at 4°C until the experiments. The coverslips were rinsed and incubated with anti-connexin 43 (Cx43) MAb (Chemicon) in a moisture chamber at 36°C for 2 h. The coverslips were then rinsed and incubated with Alexa 488-conjugated anti-mouse Ab for 2 h at 36°C. Immunofluorescence was visualized using a confocal laser scanning microscope (Olympus).


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Histological Examination

For the five dogs that went through the microembolization procedures, the LVEF dropped from 60 ± 5% (before microembolizations) to 28 ± 7% 3 mo after the last procedure. Patchy white areas (fibrosis) were visible in the anterior and lateral LV free wall, apex, anterior part of the septum, and sometimes the anterior right ventricular free wall. Histological examination confirmed that there was diffuse interstitial fibrosis in the region of ICM hearts adjacent to where cells were isolated, often with clusters of large adipocytes (Fig. 1A). Some regions had a high density of interstitial cells (inset of ICM panel, Fig. 1A). Immunofluorescence of Cx43, the major gap junction channel subunit in ventricular myocardium, showed that instead of being packed in intercalated disk areas located at the ends of cells (Fig. 1B, left), cells isolated from ICM hearts often had patches of Cx43 immunofluorescence scattered along their sides (Fig. 1B, right). This is similar to the disarray of Cx43 distribution described for a canine model of myocardial infarction, as well as in tissue from failing human hearts (36, 37).


Figure 1
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  		Fig. 1. Histology and connexin 43 (Cx43) immunocytochemistry of control and ischemic cardiomyopathy (ICM) dog hearts. A: hematoxylin and eosin stain of left ventricular (LV) free wall adjacent to the area from which myocytes were isolated. Clusters of large adipocytes and interstitial fibrosis were common in ICM LV but not in control LV. Inset of ICM panel shows a region with a marked increase in interstitial cells. B: Cx43 immunofluorescence from a control myocyte (left) and a myocyte from an ICM heart (right).

 
Cell Capacitance and Membrane Potential Recording

Ventricular myocytes from ICM hearts had much larger cell capacitance (258 ± 8 vs. 188 ± 9 pF, Fig. 2B), indicating a pronounced cellular hypertrophy. Myocytes isolated from control hearts manifested certain degree of APD variation, but the degree of APD variation became much more pronounced in myocytes from ICM hearts (Fig. 2A, left vs. right). On average, APD0 mV (phase 2) was prolonged from 255 ± 15 to 379 ± 20 ms, and APD–60 mV (phase 3) was prolonged from 424 ± 17 to 538 ± 20 ms (Fig. 2B). Under our recording conditions (controlled extra- and intracellular ionic composition), there was no difference in the RMP between the two groups of cells (–81.3 ± 0.4 vs. –82.1 ± 0.3 mV).


Figure 2
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  		Fig. 2. Cellular hypertrophy, action potential duration (APD) prolongation, and increased APD dispersion in LV mid-myocardial myocytes from ICM vs. control dog hearts. A: superimposed action potentials recorded from 27 control myocytes and 38 myocytes from ICM hearts. For each cell, action potential stabilized at the end of 30- to 120-pulse trains with a cycle length of 1 s is shown. B: summary of cell capacitance, resting membrane potential (RMP), and APD measured between upstroke and repolarization to 0 mV and to –60 mV. Numbers of cells examined are listed. *P < 0.01.

 
IKs

IKs current density and voltage dependence of activation. IKs was recorded in a nominally Na- and Ca-free bath solution (at 33–34°C) to avoid interference from Na and Ca channel currents, as well as currents mediated by the electrogenic Na/Ca exchanger. To avoid the issue of "IKs run-down" in data interpretation, IKs data presented here were obtained between 15 and 25 min after the beginning of whole cell dialysis. Our experience has been that IKs amplitude in canine ventricular myocytes is relatively stable within this time window (18), although longer intracellular dialysis inevitably leads to a decrease in IKs. Figure 3A shows two sets of representative current traces recorded in the presence of dofetilide (1 µM). After the initial spike [due to residual Ito not inactivated by the holding potential (Vh) of –50 mV], there was a slowly rising outward current upon depolarization to –20 mV or more positive voltage. This was followed by a slowly decaying outward tail current upon repolarization to –50 mV. Both could be suppressed by an IKs blocker, azimilide (18). The slowly rising outward currents during membrane depolarization were used to construct the test pulse current-voltage relationships (after normalization by cell capacitance, Fig. 3B), while the outward tail currents were used to construct the activation curves (Fig. 3C). IKs current density in ICM myocytes was reduced at voltages positive to 0 mV, but not at voltages ≤0 mV (Fig. 3B). This voltage dependence of IKs reduction was consistent with a modest, but statistically significant, negative shift in the voltage-dependence of IKs activation in ICM cells relative to control cells (Fig. 3C).


Figure 3
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  		Fig. 3. Slow delayed rectifier currents (IKs) in LV mid-myocardial myocytes from control and ICM dog hearts. A: original current traces elicited by the voltage-clamp protocol are diagrammed on top. Dofetilide 1 µM was present to suppress the rapid delayed rectifier current. B: test pulse current-voltage relationship. Current amplitudes were measured from time-dependent, slowly rising currents [i.e., after the transient outward current (Ito) spike] during 5-s depolarization pulses to test voltage (Vt), and normalized by cell capacitance (pA/pF). *P < 0.05, **P < 0.01, ICM vs. control. C: voltage-dependence of IKs activation. For each cell, the relationship between Vt and peak tail current amplitude (Itail) was fit with a Boltzmann function: Itail = Imax/{1 + exp[(V0.5 – Vt)/k]}, to estimate the maximum tail current amplitude (Imax), half-maximum activation voltage (V0.5), and slope factor (k). Itail was divided by Imax to obtain an estimate of the fraction of channels activated by Vt, and the values were averaged over cells. The curves superimposed on the data points were calculated from the Boltzmann function with the following parameter values: control, V0.5 = 7.6 ± 2.4 mV and k = 13.2 ± 0.8 mV; and ICM, V0.5 = 0.7 ± 1.9 mV and k = 12.7 ± 0.5 mV. Numbers of cells examined are listed.

 
Immunoblot analysis of IKs subunits. We used immunoblots to quantify the protein level of IKs auxiliary subunit (KCNE1) and pore-forming subunit (KCNQ1 or Kv7.1) (6, 49) in control and ICM dog hearts. The KCNE1 Ab recognized three bands in in vitro translated proteins primed with human KCNE1 cRNA (Fig. 4A, top), but none in a control translation reaction without cRNA (data not shown). Treating the translation products with N-glycosidase F collapsed the upper two bands into the lowest one, whose size of 16 kDa was similar to the size of core KCNE1 (14.6 kDa). This indicates that the 24- and 20-kDa bands were likely doubly and singly glycosylated KCNE1, because the extracellular domain of KCNE1 has two N-glycosylation sites (asparagine residues at positions 5 and 26).


Figure 4
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  		Fig. 4. Detecting KCNE1 protein in control and ICM dog hearts. A: validating the KCNE1 antibody (Ab). Top: immunoblot (IB) of in vitro translated KCNE1 treated with N-glycosidase F overnight or with enzyme-free buffer under the same conditions (marked by "+" and "–" N-glycosidase F on top). Bottom: IB of whole tissue lysate or membrane-enriched fraction from a control dog heart. Size marker positions are listed on the left, and sizes of KCNE1-specific bands are marked on the right. *N-glycosylated forms. B: comparing KCNE1 protein level in control and ICM dog hearts. Membrane-enriched fractions prepared from LV apex of control and ICM hearts were probed with KCNE1 Ab (IB). The two bands are at 30 and 20 kDa. Bottom: Coomassie blue (CB) stain of the same gel after blotting most of the proteins to the polyvinylidene difluoride membrane.

 
This KCNE1 Ab recognized a major band of ~20 kDa in whole tissue lysate from control dog LV, but a major band of ~30 kDa (as well as a weak 20-kDa band) in membrane-enriched fraction from the same tissue (Fig. 4A, bottom). This banding pattern suggests that, in canine heart, there are probably two pools of KCNE1 protein: cytoplasmic and cell surface. The cytoplasmic fraction is largely singly glycosylated (20 kDa), while the cell surface fraction is mainly doubly glycosylated (30 kDa). Fig. 4B, top, presents an immunoblot image of membrane-enriched fraction from five control dog hearts and four ICM dog hearts probed with the KCNE1 Ab. In both cases, the cell surface, doubly glycosylated fraction dominates. Densitometry of combined band intensities (after correction by CB stain of the same lanes, Fig. 4B) showed that the KCNE1 immunoreactivity was reduced in ICM hearts to 0.63 ± 0.09 (normalized by the mean intensity of control hearts, P = 0.011).

We and others have isolated a KCNQ1 splice variant from human heart that does not have the cytoplasmic NH2-terminal domain as well as the first one-third of S1, and thus cannot form functional channels on its own (11, 19). The long (functional) isoform and the short (nonfunctional) isoform are termed KCNQ1.1 and KCNQ1.2, respectively. The KCNQ1 Ab we used (C-20, Santa Cruz) was raised against the COOH-terminal domain of the protein and thus should recognize both KCNQ1 isoforms. Figure 5A, lanes 1 and 2, were loaded with membrane-enriched fraction from oocytes expressing KCNQ1.1 and KCNQ1.2, respectively. The KCNQ1 Ab recognized a band in lane 1 of the expected size for KCNQ1.1, 75 kDa. The Ab recognized a band in lane 2 of the expected size for KCNQ1.2, 61 kDa. Both bands disappeared if the KCNQ1 Ab had been preincubated with the antigenic peptide (lanes 5 and 6).


Figure 5
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  		Fig. 5. Detecting two KCNQ1 splice variants, KCNQ1.1 and KCNQ1.2 (labeled as Q1.1 and Q1.2), in control (Con) and ICM dog hearts. A: validating the KCNQ1 Ab. Membrane-enriched fraction from oocytes expressing Q1.1 or Q1.2 (human clones) or from control and ICM dog hearts were probed with the KCNQ1 Ab without (left) or with (right) pretreatment with the antigenic peptide ("–" and "+" Ag labeled below). B: comparing Q1.1 and Q1.2 protein expression in control and ICM dog hearts. Each lane was loaded with 80 µg of membrane-enriched fraction from individual dog LV. Size marker bands are labeled. Solid and open arrows point to the Q1.1 and Q1.2 bands, respectively.

 
Lanes 3 and 4 of Fig. 5A show that, in membrane-enriched fraction from control and ICM dog hearts, the KCNQ1 Ab recognized a 70-kDa band similar to KCNQ1.1, as well as a ~55-kDa band similar to KCNQ1.2. Both bands disappeared if the KCNQ1 Ab had been preincubated with the antigenic peptide (lanes 7 and 8), suggesting that indeed these are KCNQ1-specific bands. The lower band was faint in the control heart, but was similar to the 70-kDa band in intensity in the ICM heart. Figure 5B confirms this notion. The immunoblot image of membrane-enriched fraction from two control dog hearts and two ICM dog hearts were probed with the KCNQ1 Ab. The 70-kDa band intensity was similar between the two groups, whereas the 55-kDa band was stronger and more consistent in the ICM group. These data suggest that canine ventricle expresses both KCNQ1.1 and KCNQ1.2, and the short splice variant appeared more abundant in hearts with ICM. An increase in the expression of KCNQ1.2 in ICM myocytes would suppress IKs in a dominant-negative fashion, as has been shown in previous studies using heterologous expression systems (11, 19).

Ito

Ito current density and gating kinetics. Ito was recorded at 33–34°C in normal Tyrode solution that contained Na and Ca ions. To avoid interference from Na and Ca channel currents, Ito analysis was restricted to currents recorded at +50 mV. This voltage was close to the reversal potentials of Na and Ca channels so that their influence was minimized. For current traces shown in Fig. 6A, the interference from non-Ito currents was further reduced by using a conditioning step to 0 mV to totally inactivate Ito, and the residual currents were subtracted from the total currents to obtained "isolated Ito". The same type of current traces was used to compare Ito peak current density and time constant ({tau}) of inactivation between the two groups of cells. In ICM myocytes, the Ito peak current density was reduced from 18.7 ± 1.7 to 11.4 ± 0.9 pA/pF, while the {tau} of inactivation was modestly prolonged from 10.2 ± 0.2 to 13.1 ± 0.4 ms (both P < 0.001, Fig. 6B). There was no difference in the voltage dependence of Ito inactivation between the two groups of cells (Fig. 6C). Ito recovery from inactivation in canine ventricular myocytes follows a double-exponential time course (3, 14, 26). The recovery time course of Ito in ICM myocytes appeared accelerated relative to that in control myocytes (Fig. 6D). This difference was mainly due to an acceleration of the slow phase of recovery: the slow time constant was shortened from 1,137 ± 572 to 523 ± 140 ms.


Figure 6
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  		Fig. 6. Ito in LV mid-myocardial myocytes from control and ICM dog hearts. A: original current traces elicited by the voltage-clamp protocol shown in inset. Background and leak currents were subtracted: total current minus current after a prepulse to 0 mV that totally inactivated Ito. B: summary of peak Ito current density (pA/pF) and time constant of inactivation (in ms) measured at +50 mV. *P < 0.001, ICM vs. control. C: voltage dependence of Ito inactivation. The voltage-clamp protocol is diagrammed in the inset: 2-s pulses to conditioning voltage (Vc) were applied to cause different degrees of Ito inactivation, and, after a 5-ms step to –80 mV, the degree of Ito inactivation was evaluated by the peak current amplitude measured at +50 mV. The current amplitude was normalized by the maximum amplitude without Vc (availability), and the relationship between Vc and availability was fit with a Boltzmann function: availability = 1/{1 + exp [(Vc – V0.5)/k]}. The superimposed curves were calculated from the Boltzmann function with the following parameter values: control, V0.5 = –61.7 ± 0.6 mV and k = 4.2 ± 0.4 mV; ICM, V0.5 = –62.4 ± 0.7 mV and k = 3.4 ± 0.2 mV. D: time course of Ito recovery from inactivation. The voltage-clamp protocol is diagrammed in inset: from holding potential (Vh) –80 mV, double pulses (V1 and V2) each to +50 mV for 250 ms were applied with varying interpulse (V1-V2) intervals. The interval between these double pulses was 15 s. Peak current amplitude during V2 was divided by that during V1 to estimate "fraction of channels recovered from inactivation" during the V1-V2 interval. The relationship between V1-V2 interval and fraction recovered was fit with a double-exponential function: fraction recovered = Af [1 – exp(–t/{tau}f)] + As [1 – exp(–t/{tau}s)], where Af and As are fast- and slow-recovering components, respectively; t is V1-V2 interval; and {tau}f and {tau}s are fast and slow recovery time constants, respectively. The superimposed curves were calculated with the following parameter values: for control, Af = 0.40 ± 0.05, {tau}f = 68 ± 25 ms, As = 0.60 ± 0.05, {tau}s = 1,137 ± 572 ms; and for ICM, Af = 0.51 ± 0.05, {tau}f = 57 ± 13 ms, As = 0.49 ± 0.05, {tau}s = 523 ± 140 ms. Numbers of cells examined are listed.

 
Immunoblot analysis of major Ito subunits. The Ito channels in dog heart have two types of pore-forming subunits, Kv4.3 and Kv1.4, that manifest distinctly different rates of recovery from inactivation (14, 52, 57). Kv4.3 can be modified by an association of KChIP2 auxiliary subunit in terms of current amplitude and gating kinetics (4). We quantified these subunits' protein levels using immunoblotting on membrane-enriched fraction. Figure 7A shows images of separate immunoblots loaded with the same set of membrane fraction from five control dog hearts and four ICM dog hearts. The Kv4.3, KChIP2, and Kv1.4 antibodies recognized bands of 74, 35, and 76 kDa, respectively. In one control heart, the Kv1.4 Ab also recognized a very faint 99-kDa band. The band intensities were corrected by the CB stain (not shown) and normalized by the mean band intensity from the five control hearts. Figure 7B summarizes the densitometry results. Kv4.3 showed no significant change, whereas there was a decrease in both KChIP2 and Kv1.4 in ICM vs. control, reaching a P value of <0.05.


Figure 7
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  		Fig. 7. Using IBs to quantify protein level of major Ito channel subunits in control and ICM dog hearts. A: IB images. Membrane-enriched fractions from 5 control dog hearts and 4 ICM dog hearts were loaded at 80 µg/lane and probed with Abs specific for Kv4.3, KChIP2, and Kv1.4. Positions for size markers and subunit-specific bands are denoted on the left and right, respectively. B: summary of IB results. Intensities of subunit-specific bands are quantified by densitometry, normalized by total CB stains of the same lanes (not shown), and then normalized by the mean intensity from the 5 control dogs. Numbers of cells examined are listed. * P < 0.05 ICM vs. control.

 
Effects of changing the expression ratio of Kv4.3 to Kv1.4 on current properties. Kv4.3 and Kv1.4 expressed separately in oocytes exhibited similar voltage dependence of peak current-voltage relationship as well as voltage dependence of inactivation (Fig. 8, A and B). The rate of recovery from inactivation of Kv4.3 was 16 times faster than that of Kv1.4 (Fig. 8C). Coexpressing the two in different ratios resulted in peak current-voltage relationship and voltage dependence of inactivation not different from those of separate expression (Fig. 8, A and B). However, due to the very different rates of recovery from inactivation, coexpressing Kv4.3 and Kv1.4 created a distinct biphasic recovery time course: a fast phase due to the Kv4.3 component, and a slow phase due to the Kv1.4 component (Fig. 8C). The proportion of these two components depended on the ratio of Kv4.3/Kv1.4, so that reducing the Kv1.4 component relative to Kv4.3 produced an apparent acceleration of recovery from inactivation.


Figure 8
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  		Fig. 8. Effect of Kv1.4 coexpression with Kv4.3 on current properties. Kv4.3 and Kv1.4, separately or together, were expressed in oocytes. The quantities of cRNAs injected are listed in the inset of B (in ng/oocyte). A: peak current-voltage relationship measured by the difference between peak and current level 500 ms into the test pulse. B: voltage dependence of inactivation. C: time courses of recovery from inactivation. Voltage-clamp protocols and data analysis for data shown in B and C were the same as those described for Fig. 6, C and D. The time courses of recovery for Kv4.3 and Kv1.4 expressed separately were fit with a single-exponential function, fraction recovered = 1 – exp(–t/{tau}), with {tau} = 149 ± 5 ms for Kv4.3 and 2,361 ± 190 ms for Kv1.4. The time courses of recovery for Kv4.3 and Kv1.4 coexpressed were fit with a double-exponential function, and the following parameter values were obtained: for Kv4.3/Kv1.4 = 8:2, Af = 0.69 ± 0.04, {tau}f = 195 ± 9 ms, As = 0.31 ± 0.04, {tau}s = 3,218 ± 187 ms, and for Kv4.3/Kv1.4 = 5:5, Af = 0.35 ± 0.03, {tau}f = 189 ± 9 ms, As = 0.65 ± 0.03, {tau}s = 2,551 ± 75 ms. For all groups, n = 8 each.

 
IK1 and ICaL

Figure 9 shows that, in ICM myocytes, there was a decrease in IK1 current density in both inward and outward directions. The latter is better shown by the expanded current scale in the inset. Figure 10 shows that ICaL recorded from control and ICM myocytes were similar in the maximal peak ICaL current density (which occurred at 0 mV), time constant of the fast (major) component of inactivation, and the voltage dependence of inactivation (Fig. 10, A–C). The rate of recovery from inactivation was tested at one voltage, –40 mV, and there was no change between the two groups of myocytes (Fig. 10D).


Figure 9
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  		Fig. 9. Inward rectifier currents (IK1) in LV mid-myocardial myocytes from control and ICM dog hearts. Currents were elicited by the voltage-clamp protocol diagrammed (Vh –50 mV, test pulse 500 ms in duration). Current amplitudes at the end of test pulses were normalized by cell capacitance and plotted against test pulse voltage in the main graph. The outward current portion of the current-voltage is shown at an expanded scale in the inset. *P < 0.05, **P < 0.01, ICM vs. control.

 

Figure 10
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  		Fig. 10. L-type Ca currents (ICaL) in LV mid-myocardial myocytes from control and ICM dog hearts. A: original current traces elicited by the voltage-clamp protocol are diagrammed in inset. Dotted line denotes zero current level. B: summary of peak ICaL current density and time constant of the fast (major) component of inactivation measured at 0 mV. C: voltage dependence of ICaL inactivation. The voltage-clamp protocol is diagrammed in the inset. Data analysis was similar to that described for Ito inactivation (Fig. 6C). The superimposed curves were calculated from the Boltzmann function with the following parameter values: for control, V0.5 = –18.6 ± 0.4 mV and k = 4.2 ± 0.2 mV; and for ICM, V0.5 = –19.5 ± 0.5 mV and k = 4.2 ± 0.2 mV. D: time course of ICaL recovery from inactivation measured at Vh –40 mV. The voltage-clamp protocol is diagrammed in inset, and time course of recovery was fit with a single exponential function. The superimposed curves were calculated with the following {tau} values: 122 ± 7 ms for control, 124 ± 9 ms for ICM. n, Numbers of cells examined are listed.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The major findings in our study can be summarized as the following. 1) IKs density in ventricular myocytes from dog hearts suffering ICM was reduced at voltages positive to 0 mV, reaching a 40% reduction at +50 mV. This voltage-dependent IKs reduction is consistent with a negative shift in the voltage dependence of IKs activation. 2) KCNE1 (IKs auxiliary subunit) was reduced in ICM vs. control hearts. Furthermore, although KCNQ1.1 (functional IKs pore-forming subunit) was not altered in ICM hearts, a truncated nonfunctional KCNQ1.2 isoform was detected and appeared more abundant in ICM relative to control hearts. 3) Ito current density measured at +50 mV was reduced by 40% in ICM myocytes. There was an acceleration of Ito recovery from inactivation. 4) Although Kv4.3 (pore-forming subunit of the fast-recovering Ito component) was unaltered, there was a reduction in KChIP2 (auxiliary subunit of the fast-recovering Ito component) and Kv1.4 (pore-forming subunit of the slow-recovering Ito component). 5) IK1 current density was reduced in ICM myocytes, by 20% at –70 mV (outward component) and by 34% at –130 mV (inward component). 6) There were no changes in ICaL in ICM myocytes relative to control myocytes.

Implications for Arrhythmogenic Mechanism in the Canine Model of ICM

The mechanism of arrhythmia in this canine model of ICM has been studied previously (39), using dogs manifesting reduced LVEF (from 63 ± 3% premicroembolization, to 22 ± 3%) similar to that of ICM dogs studied here. Twenty-four to 48-h Holter monitoring showed that the ICM dogs experienced premature ventricular complexes and nonsustained VT of a monomorphic or polymorphic nature. Using a three-dimensional mapping technique, the author further showed that the VT almost always arose from focal activation. Slowing of impulse conduction was rare and only occurred at sites of transmural fibrosis. There was no sign of macroreentry. The author suggested that, in this animal model, triggered activity and automaticity were the major arrhythmogenic mechanisms (39).

The decrease in IKs, Ito, and IK1 reported here should contribute to APD prolongation in ventricular myocytes from ICM hearts (59). It is conceivable that these myocytes suffered different degrees of K current reduction and thus had different degrees of APD prolongation, depending on their locations in the heart and their relationships to the microsphere-clogged coronary arteries. If the myocytes were well-coupled in the in situ heart, their APD would be homogenized by electrotonic interactions (38). It is possible that the presence of interstitial fibrosis, in conjunction with the observed disarray of Cx43 subcellular distribution, might reduce the degree of intercellular coupling, allowing cells that suffered the most severe degree of K channel reduction to maintain their greatly prolonged APD. Although there was no difference in the RMP between control and ICM myocytes under our recording conditions, the decrease in IK1 predicts that membrane conductance at RMP would be reduced in the ICM cells. This could destabilize the RMP by magnifying the effects of small disturbances in membrane currents, such as transient inward currents induced by Cai oscillation (28, 14a), inward currents through pacemaker channels (9), or the T-type Ca channels (15, 35). Normal ventricular myocytes do not exhibit pacemaker currents in the physiological voltage range (43) and have little or no T-type Ca channel currents (55). However, in ventricular myocytes from hypertrophied rat or feline hearts, an appearance of pacemaker currents and T-type Ca channel currents has been described (9, 15, 35). Whether this occurs in the ICM dog hearts requires further experimentation. Combining these three factors (APD prolongation/increased APD dispersion, reduced electrotonic coupling, and RMP instability) would predispose the ICM hearts to arrhythmias due to triggered activity or increased automaticity. It is important to note that the effects of decreased IKs, Ito, and IK1 reported here, as well as the effects of increased long-lasting component of TTX-sensitive Na channel current reported previously (59, 62), could have been partially offset by a simultaneous increase in IKr in these cells (21).

Molecular Mechanisms for IKs and Ito Remodeling in the Canine Model of ICM

Association of KCNE1 with KCNQ1.1 increases the current amplitude and shifts the voltage dependence of activation in the positive direction (6, 49). On the other hand, coexpressing KCNQ1.2 (11, 19) with KCNQ1.1/KCNE1 reduced the current amplitude in a dominant-negative manner, without affecting the voltage dependence of activation. Both reduction of KCNE1 and increase of KCNQ1.2 would contribute to the decrease in IKs current density in ICM myocytes. Since the stoichiometry of KCNE1/KCNQ1.1 in an IKs channel complex may be variable (60), the effects of KCNE1 on the gating kinetics of KCNQ1.1 can be graded by the expression level of KCNE1 (7, 10). Therefore, the decrease in KCNE1 is consistent with the negative shift in the voltage dependence of IKs activation.

Association of KChIP2 with Kv4.3 leads to an increase in current amplitude and an acceleration of recovery from inactivation (4). Therefore, a decrease in KChIP2 is predicted to cause a reduction in Ito current amplitude and a slowing of recovery from inactivation. Kv1.4 is the major molecular correlate of the slow-recovering Ito component (3, 14, 57). Therefore, a reduction in Kv1.4 could cause a decrease in Ito current amplitude and an apparent acceleration of recovery from inactivation (suggested by Fig. 8C). Therefore, reduction of both KChIP2 and Kv1.4 can contribute to the decrease in Ito current density, but the decrease of the latter is responsible for the change in the rate of recovery from inactivation. Kv1.4 is more abundantly expressed in the subendocardial region (8, 61). This, in conjunction with the lower abundance of Kv4.3 in the subendocardial region (63), suggests that downregulation of Kv1.4 could have a larger impact on Ito in the subendocardial region than the midmyocardial region studied here.

Comparison With Previous Studies

Table 1 compares our findings with selected examples of other animal models of heart disease, for which remodeling of IKs and/or Ito has been described, and information on the underlying changes in channel subunit expression (at the protein level) is available (17, 41, 44, 45, 58, 63). This comparison illustrates that an apparently similar phenotypic change (downregulation of IKs or Ito) can have divergent molecular mechanisms. Such differences can be due to species or regional variations in the regulation of channel expression, as well as etiology and/or stages of pathological changes in the heart. These differences can lead to changes in not only current amplitude but also gating kinetics, with functional consequences in terms of arrhythmogenic mechanism and strategy for antiarrhythmic therapies.


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  		Table 1. Divergent molecular mechanisms for K channel remodeling in animal models of heart disease

 
Technical Limitations

All patch-clamp recordings were done on myocytes isolated from the mid-myocardial layer of LV free wall affected by the microembolizations. Due to the heterogeneity in ion channel expression among different regions of the heart, changes in membrane channels in other regions may differ quantitatively.

During patch-clamp recordings, the cells were dialyzed with a pipette solution containing 10 mM each of ATP, HEPES, and EGTA. These recording conditions would mask cellular abnormalities related to mitochondrial dysfunction (53), improper handling of Cai (14a), and other intracellular ionic perturbations. Furthermore, we stimulated the cells at a frequency of 1 Hz. Cells with plateau phase longer than 1,000 ms were excluded from our data set. Therefore, our recording conditions did not allow us to make any inference about the occurrence or prevalence of delayed afterdepolarizations or early afterdepolarizations in the ICM myocytes. Therefore, the changes we observed here likely represented intrinsic changes in membrane channels due to alterations in channel subunit expression and composition induced by the chronic ischemic process. Electrical remodeling in an in situ heart will most likely result from these "intrinsic" changes in membrane channels, with further influences by "extrinsic" changes in cellular milieu, such as ATP depletion, Cai overload, effects of neurohumoral factors, inflammatory cytokines, reactive oxygen species, and mechanical stretch.

The Western blots were done on membrane fraction prepared from LV apex tissue. There can be differences in channel subunit expression between LV mid-myocardial layer and LV apex. Furthermore, myocardial tissue contained nonmyocyte elements. Therefore, the correlation between patch-clamp data from single isolated cells and immunoblot data from multicellular tissue is suggestive but not conclusive.

In conclusion, we suggest the following scenario. In dog hearts suffering ICM, decrease in IKs, Ito, and IK1 contributes to APD prolongation and increased APD dispersion. These differences in APD among cells in an in situ ICM heart are probably protected by reduced electrotonic coupling, due to interstitial fibrosis and disarrayed Cx43 distribution. The decrease in IK1 would also sensitize the RMP to small arrhythmogenic disturbances in membrane currents, such as transient inward currents and automaticity. These factors combined to predispose the ICM hearts to arrhythmia.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
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REFERENCES
 
This study was support in part by National Heart, Lung, and Blood Institute Grant RO1 HL67840 (to G.-N. Tseng).

Present address of R. S. D. Higgins: Department of Cardiovascular and Thoracic Surgery, Rush Presbyterian St. Luke’s Medical Center, Chicago, IL 06612–3833.


   ACKNOWLEDGMENTS
 
The authors thank Cynthia Allen for assistance in animal surgery and care, and Drs. A. L. Pond and J. M. Nerbonne for providing the KCNE1 antibody.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G.-N. Tseng, Dept. of Physiology, Virginia Commonwealth Univ., Richmond, VA 23298 (e-mail: gtseng{at}vcu.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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