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Cardiovascular Research 2006 72(2):262-270; doi:10.1016/j.cardiores.2006.08.001
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Copyright © 2006, European Society of Cardiology

Activation of gadolinium-sensitive ion channels in cardiomyocytes in early adaptive stages of volume overload-induced heart failure

Carmel M. McNicholas-Bevenseea,*, Kevin B. DeAndradea, Wayne E. Bradleyb, Louis J. Dell'Italiaa,b,c, Pamela A. Lucchesia,1 and Mark O. Bevenseea

aDepartment of Physiology and Biophysics, University of Alabama at Birmingham (UAB), 868 McCallum Basic Health Sciences Building, 1918 University Boulevard, Birmingham AL, 35294, United States
bDepartment of Medicine, Division of Cardiovascular Disease, UAB, United States
cBirmingham Veterans Affairs Medical Center, Birmingham, Alabama, United States

* Corresponding author. Tel.: +1 205 934 1785; fax: +1 205 975 7679. Email address: carmel{at}physiology.uab.edu

Received 3 July 2006; revised 29 July 2006; accepted 1 August 2006


   Abstract
Top
 Abstract
1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
 References
 
Objective: The objective of this study was to investigate whether gadolinium (Gd3+)-sensitive stretch-activated ion channels (SAC) are basally active in left ventricular (LV) myocytes in early stages of heart failure (HF) induced by volume overload.

Methods: The aortocaval fistula (ACF) model was employed to induce HF due to volume overload in rat. At specific time-points, LV myocytes were acutely isolated using a modified Langendorff apparatus. Whole-cell currents were measured using the patch-clamp technique and intracellular Ca2+(Ca2+i) was examined using fluorescence imaging and the Ca2+-sensitive dye Fura-2.

Results: Current–voltage data were obtained from sham and ACF myocytes at 5-d and 2-, 6-, 8- and 10-wk post surgery. Compared to data from matching sham rats, a 10 µM Gd3+-sensitive current at –100 mV comprised a larger fraction of total current in myocytes from 5-d, 2-wk, and 6-wk ACF rats. In general, the Gd3+-sensitive current contributed to inward currents at mV≤–80 and outward currents at >+20 mV. The enhanced Gd3+-sensitive current was absent in myocytes from 8- and 10-wk ACF rats. 10 or 100 µM Gd3+ had no appreciable effect on resting Ca2+i of myocytes from 5-d ACF or corresponding sham rats. The Gd3+-sensitive current in 5-d ACF myocytes was i) sensitive to the cation-selective SAC inhibitor, GsMTx-4, ii) non-selective for Na+/K+, and iii) impermeable to Ca2+.

Conclusion: A basally-active, Gd3+- and GsMTx-4-sensitive SAC current that is non-selective for Na+ and K+, but impermeable to Ca2+ under resting conditions is transiently elevated in LV myocytes from rats in early stages of volume overload-induced HF.

KEYWORDS Calcium (cellular); Heart failure; Ion channels; Stretch; Membrane currents


   1. Introduction
Top
 Abstract
 1. Introduction
2. Materials and methods
 3. Results
 4. Discussion
 References
 
The pathophysiological changes that occur during the progression toward heart failure (HF) involve a complex sequence of compensatory events and associated morphological changes that eventually result in an adversely remodeled myocardium and a dilated, thin-walled, spherical ventricle [1,2]. In an attempt to compensate for the reduction in cardiac function, the sympathetic nervous system and other neurohumoral systems are activated [2]. Additionally, the myocardium itself – early in the disease process – undergoes hypertrophy in an attempt to compensate for impaired heart function. This hypertrophy, in conjunction with increased mechanical load, has a number of consequences that cause an adverse remodeling of cardiomyocytes and the extracellular matrix [2]. Furthermore, stretching of the sarcolemma can alter the electrophysiological properties of cardiac myocytes, and lead to potentially life-threatening membrane depolarizations and cardiac arrhythmias [3,4].

Membrane stretch activates a family of ion channels known as stretch-activated channels (SACs). Found in numerous cell types [5–7], SACs are a family of channels that are stimulated by increases in cell volume (swelling) and/or increases in plasma-membrane tension (stretching). SACs have different ion selectivities [8,9] and regulatory properties [10], and several have been identified in heart [7]. The following three types of SACs have been functionally identified in the heart: cation-selective, anion-selective, and K+-selective ones [8]. The permeability of certain SACs to Ca2+[11] is particularly relevant to HF because intracellular Ca2+(Ca2+i) overload of myocytes is a cellular pathophysiological event of the disease. SACs are blocked by micromolar concentrations of gadolinium (Gd3+) and GsMTx-4, which is a toxin from the venom of the tarantula Grammostola spatulata. Although many SACs have been identified at the functional level, the molecular characterization of SACs is far from complete.

While the effects of stretch on heart function have been recognized for many years [12], the role of specific SACs in either normal or diseased heart function is poorly understood. Clemo et al. [13] described a swelling-activated, Gd3+-sensitive current (ICir,swell) in normal dog myocytes as a SAC. This current is persistently activated in myocytes isolated from failing ventricles induced by rapid ventricular pacing – a result the authors suggest is due to a stretch-behavior of the myocytes in congestive heart failure [13]. SAC inhibitors have been shown to restore contractile function in a hamster model of dilated cardiomyopathy [14]. Nevertheless, it is not clear if and how SACs contribute to the progression of HF. We therefore tested the hypothesis that basally active, Gd3+-sensitive ion channels are present in cardiac myocytes in early stages of the rat aortocaval fistula (ACF) model of volume overload-induced HF. We chose this particular model because it represents a pure volume overload-induced stretch without an increase in pressure. Furthermore, with time, ACF rats display left ventricular (LV) dilatation and myocyte hypertrophy, as well as systemic and intracardiac activation of neurohumoral systems that mimic those seen in human patients [15].

Using the patch-clamp technique with acutely isolated LV myocytes from ACF and sham rats, we tested for basally active, Gd3+-sensitive channels during the early compensated stage [16] of HF. As early as day 5, and up to week 6 post surgery, myocytes from ACF vs. sham animals displayed a larger Gd3+-sensitive fraction of total current. These electrophysiological properties disappeared at 8–10-wk post ACF surgery. Inhibiting the enhanced basal currents with Gd3+ did not appreciably change resting Ca2+i–data consistent with negligible influence of SAC activity on resting Ca2+ levels. The identity and characteristics of the Gd3+-sensitive channels in the 5-d ACF myocytes were evaluated further in additional pharmacology and ion-selectivity studies. Applying GsMTx-4, a toxin known to inhibit cation-selective SACs, also inhibited the larger basal currents in the ACF vs. sham myocytes. These Gd3+/toxin-sensitive currents are nonselective for Na+ and K+, and impermeable to Ca2+. In summary, Gd3+-sensitive SAC activity is transient in early hypertrophic stages of volume overload-induced HF before the onset of overt HF. SACs may play a role in the initial adaptation to the stress of pure volume overload-induced stretch in the ACF model.

Preliminary data have been published in abstract form [17].


   2. Materials and methods
Top
 Abstract
 1. Introduction
 2. Materials and methods
3. Results
 4. Discussion
 References
 
2.1. Generation of ACF model
ACFs were created as previously described by the Dell'Italia laboratory [18]. The surgical procedure was the same for sham animals except for inserting the needle to create the fistula. The animal research meets the standards prescribed in the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996).

2.2. Myocyte isolation
LV myocytes were enzymatically isolated by retrograde perfusion of the heart. After a rat was anesthetized with 50 mg kg–1 Nembutal, the chest was opened and the heart quickly excised and placed in a Krebs' solution (37 °C) containing 1.8 mM Ca2+. The heart was cannulated via the aorta and perfused on a modified Langendorff apparatus with Krebs' solution+1.8 mM Ca2+ until the perfusate was clear. Next, the heart was perfused with a low-Ca2+ Krebs' solution containing 25 µM Ca2+ first without, and then with 0.1% fat-free BSA and 0.2 mg/ml Type II collagenase (Worthington Biochemical Corporation, Lakewood, NJ). The enzyme-containing solution (~ 100 ml) was recirculated for ~ 30 min until the ventricular myocardium was digested. The heart was finally perfused with ~ 50 ml modified Krafte–Brühe (KB) solution. The LV was dissected from the heart, minced in KB using scissors, triturated with a large bore pipette, and filtered through 100-µm mesh net. Myocytes in suspension were stored at room temperature in KB before aliquots were plated onto laminin-coated coverslips prior to experiments. Myocytes selected for both electrophysiologic and fluorescence studies were rod-shaped with clear striations and no evidence of membrane blebbing or spontaneous contractions. There were no obvious differences in the yields of viable myocytes among the different experimental groups, therefore a quantitative analysis was not performed.

2.3. Electrophysiological recordings
Experiments were performed 1–8 h following myocyte isolation. Whole-cell current recordings were obtained from LV myocytes plated on coverslips and mounted in a flow-through chamber on the stage of a Leica DM IRB inverted microscope (Leica Microsystems, Heidelberg, Germany). Bath solution exchange was achieved using a perfusion valve control system converging on an 8 to 1 perfusion manifold (Warner Instruments, Hamden, CT). Recording pipettes (3–5 M{Omega}) were pulled from borosilicate glass capillaries (Warner Instruments) using a PC-10 microelectrode puller (Narishige Scientific Instrument Laboratory, Tokyo, Japan).

Currents were obtained using an Axopatch 200B patch-clamp amplifier (Axon Instruments, Molecular Devices, San Jose, CA) and Clampex software (pClamp 8.2, Axon Instruments). Currents were low-pass filtered at 5 kHz (LPF-8 filter, Warner Instruments) and digitized (Digidata-1321A interface, Axon Instruments) at a sampling frequency of 2 kHz. Current–voltage (I–V) relationships were obtained using a pulse protocol in which cells were stepped from –40 mV to holding potentials from –120 to +60 mV (500 ms) in increments of 20 mV. Mean currents averaged from 2–3 pulse protocols were obtained during the 400–500 ms period of each sweep using Clampfit software (Axon Instruments). Membrane capacitance, as well as membrane and access resistances were routinely measured using the "Membrane Test" tool in Clampex, and capacitance values were used to report currents as a function of cell surface area. Experiments were terminated if any of these three parameters changed unexpectedly. Whole-cell currents are reported as current density (pA/pF). Cell capacitances were variable within a given group, although there was an increase in mean capacitance with age of both sham and ACF animals. No consistent pattern emerged when mean capacitances of the ACF and sham myocytes were compared at the different time points. Junction potentials (VJP) for all solutions were small (~ –5 mV), but were nevertheless factored into the plots of current–voltage where Vm=Vhold–VJP. Vhold values are reported in the text. Experiments were carried out at 37 °C or at room temperature (~ 20 °C) as described below.

To minimize the amount of expensive GsMTx-4 (Peptides International, Louisville, KY) toxin required per experiment, we used an alternative solution delivery device (SF-77B, Warner Instruments) to perfuse myocytes directly with a low-volume stream. Toxin experiments were performed at room temperature because the direct perfusion system is not optimized for heated solutions. Switching from bath perfusion to direct perfusion during an experiment had a negligible effect on baseline current.

2.4. Intracellular Ca2+ recordings
Myocytes plated onto a glass coverslip coated with laminin or CellTak® (BD Biosciences, San Jose, CA) were incubated for 10–30 min in solution containing 2–8 µM of the cell-permeant, -AM precursor of the Ca2+-sensitive dye Fura-2. The coverslip was mounted in a flow-through chamber and secured on the stage of an IX-70 inverted microscope (Olympus America Inc., Melville, NY) designed for fluorescence imaging. Both a Lambda DG-4 high-speed wavelength changer (Sutter Instruments Company, Novato, CA) and an Orca II digital CCD camera (Hamamatsu, Photonics, Hamamatsu City, Japan) were attached to the microscope. Measurements were obtained with an Olympus LCPlanF1 x40 objective with a 0.60 numerical aperture. 510-nm fluorescence emitted by the dye was captured from 340-nm excitation (I340) and 380-nm excitation (I380). I340/I380 (R) is mainly a function of Ca2+i concentration ([Ca2+]i). R values were normalized to the stabilized R value at the onset of each experiment (RN) after subtracting background I340 and I380 values from myocytes without dye. MetaFluor software (Universal Imaging, West Chester, PA) was used for image acquisition and data analysis. Experiments were performed at 35–37 °C.

2.5. Solutions
Solute concentrations are given in mM. Krebs' solution contained: 118 NaCl, 4.7 KCl, 1.2 MgSO4, 1.2 KH2PO4, 11 glucose, and 22 NaHCO3, and was equilibrated with 5% CO2/95% O2 to pH 7.4. KB solution contained: 50 Kglutamate, 20 HEPES, 20 Taurine, 10 glucose, 3 MgSO4, 0.5 EGTA, 45 KCl, 30 KH2PO4, and KOH to pH 7.3. Pipette solution contained: 140 KCl, 10 HEPES, 5 MgATP, 1 MgCl2, 2.5 disodium phosphocreatine, 2.5 phosphocreatine-Tris, 5 EGTA, and KOH to pH 7.2. Control bath solution contained: 140 NaCl, 4 KCl, 1.8 CaCl2, 1 MgCl2, 10 glucose, 10 HEPES, and NaOH to pH 7.4. For the low-K+(1 mM) bath solution, 3 mM KCl was replaced with an equimolar amount of NaCl. For the (0Na: 0K: 0Ca)ext solution, Na+, K+, and Ca2+ were replaced by an equimolar concentration of N-methyl-D-glucammonium (NMDG+). A similar replacement was made for the (0Ca)ext solution. All chemicals were obtained from Sigma Chemical Co. (St. Louis, MO) unless otherwise specified.

2.6. Statistics
Results are presented as mean±standard error. n represents the number of experiments. Means between groups of data were compared (i) using two-way ANOVA (Tukey criterion) (Origin 7.5 software, OriginLab) where stated or, (ii) paired or unpaired Student's t tests (Microsoft® Excel 2002). P<0.05 was considered significant.


   3. Results
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
4. Discussion
 References
 
3.1. Hypertropy of the ACF heart
The ACF model of congestive HF has been well characterized [15,16,18,19]. In the present study, mean HW/BW ratios were significantly larger by 27–45% in ACF compared to age-matched sham animals at all time points examined (Table 1). Thus, the ACF leads to volume overload-induced cardiac hypertrophy that is evident as early as 5-d post surgery and continues into the compensated phase of hypertrophy.


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  		Table 1 Heart and body weight ratios for rats with or without ACF induction

 
3.2. Higher basal currents in ACF myocytes
We obtained I–V data and examined basal currents in myocytes from ACF vs. age-matched sham rats at 37 °C. Currents normalized to membrane capacitance are reported as current densities. Basal currents at hyperpolarized (–120 to –40 mV) potentials at 5-d, 2-, 8- and 10-wk post surgery and at depolarized (+20 to +60 mV) potentials at 2-wk and 10-wk were significantly larger in ACF vs. sham myocytes (two-way ANOVA). We chose three holding potentials (–120, –100, and +60 mV) for further comparisons and present these data in Table 2. At the 5-d time point, the individual basal current densities were significantly higher in ACF vs. sham myocytes at hyperpolarized potentials (e.g., –100 mV), but not at the depolarized potential of +60 mV. At 2 wk, the current densities were significantly higher in ACF vs. sham myocytes at –120, –100 mV, and +60 mV. These larger currents began to dissipate at ~6 wk. However there was a higher current density at 10 wk in the ACF vs. sham myocytes at depolarized, but not hyperpolarized potentials.


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  		Table 2 Basal currents in rat left ventricular myocytes

 
3.3. Gd3+-sensitive currents in ACF myocytes
We next tested the Gd3+ sensitivity of the higher basal currents in ACF myocytes at 37 °C. I–V relationships were obtained from ACF and sham myocytes perfused with the control bath solution first without and then with 10 µM GdCl3. Raw current traces obtained from both 5-d ACF and sham myocytes are shown in Fig. 1. Representative currents from the ACF myocyte (top left) are larger than those from the sham myocyte (top right). Furthermore, a larger fraction of the current is sensitive to Gd3+ in the ACF myocyte (middle left) compared to the sham myocyte (middle right). The majority of the Gd3+-insensitive inward current in both myocytes is Ba2+-sensitive, and likely due to the inwardly-rectifying K+ channel, IK1 (lower traces). The Gd3+-sensitive current is unlikely to be IK1 because (i) Gd3+ had no effect on basal currents in ventricular myocytes from 5-d and 10-wk sham-operated rats at hyperpolarized potentials where IK1 is active (Fig. 2), and (ii) IK1 in ventricular myocytes is insensitive to Gd3+ [20,21].


Figure 1
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  		Fig. 1 Basally active, Gd3+-sensitive current records from 5-d ACF myocytes (left traces) and corresponding sham myocytes (right traces). Currents records were obtained at 37 °C before (top traces) and after (middle traces) applying 10 µM GdCl3, and then after applying 1 mM BaCl2 (bottom traces).

 

Figure 2
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  		Fig. 2 Gadolinium-sensitive currents in 5-d and 10-wk ACF myocytes. Mean I–V relationships at 37 °C were recorded from 5-d ACF myocytes (panel A), 5-d sham myocytes (panel B), 10-wk ACF myocytes (panel C), and 10-wk sham myocytes (panel D). I–V plots were obtained from myocytes exposed first to the control bath solution (squares), then to the solution containing 10 µM GdCl3 (circles), and finally to the solution containing 1 mM BaCl2 (triangles). Panel E, Gd3+-sensitive I–V plots from 5-d ACF myocytes (closed diamonds) and sham myocytes (open diamonds). Plots were obtained by subtracting GdCl3 I–V plots from the matching control ACF (Panel A) and sham (Panel B) I–V plots. The difference between the GD3+-sensitive I–V plots from the ACF and sham myocytes is shown as half-filled diamonds. Numbers of experiments are shown in parentheses. *P<0.05 for the response to Gd3+, and {dagger} P<0.05 for ACF vs. sham.

 
Summary I–V data from experiments similar to those shown in Fig. 1 are plotted as mean current densities in Fig. 2. For 5-d ACF myocytes at –100 mV (Fig. 2A), the mean current density was 31% less (P=0.02) in the presence (–7.5±1.4 pA/pF) vs. the absence (–10.9±1.8 pA/pF) of 10 µM Gd3+ (n=10). In contrast, for the corresponding sham myocytes at –100 mV (Fig. 2B), the mean current density was the same in the absence (–6.6±0.8 pA/pF) and the presence (–6.7±0.6 pA/pF) of Gd3+ (n=8). Similar results with the 5-d ACF myocytes were obtained at the depolarized potential of +60 mV. Specifically, the mean current density in 5-d ACF myocytes at +60 mV (Fig. 2A) was 41% less (P=0.003) in the presence (6.8±0.9 pA/pF) vs. absence (11.6±2.0 pA/pF) of Gd3+(n=10). The mean current density in the corresponding sham myocytes (Fig. 2B) was only 14% less (P=0.002) in the presence (7.1±1.1 pA/pF) vs. absence (8.3±1.3 pA/pF) of Gd3+ (n=8). Therefore, mean Gd3+-sensitive currents are larger in the 5-d ACF vs. sham myocytes (Fig. 2E).

Markedly different results were obtained from 10-wk ACF myocytes. In particular, little or no Gd3+-sensitive current was observed in either the ACF myocytes (Fig. 2C) or sham myocytes (Fig. 2D) at hyperpolarized potentials. There was however a small, but significant 19% decrease (P=0.03) in the mean current density in the presence (8.9±1.5 pA/pF) vs. absence (10.9±1.6 pA/pF) of Gd3+ in ACF myocytes at +60 mV (n=8).

To examine additional ACF time points, we performed experiments on myocytes at 2-, 6-, and 8-wk post surgery. Gd3+-sensitive currents at hyperpolarized potentials (–120 to –40 mV) at 5-d, 2-wk and 6-wk post surgery and at depolarized potentials (+20 to +60 mV) at 5-d, 2-wk 6-wk and 10-wk were significantly larger in ACF vs. sham myocytes (two-way ANOVA). Summary Gd3+-sensitive current data at holding potentials of –120, –100, and +60 mV from I–V relationships are shown in Fig. 3. An apparent biphasic effect emerges. At the 5-d and 2-wk time points, significantly larger mean Gd3+-sensitive currents at each holding potential in the ACF vs. sham myocytes were observed. At the 6-wk time point, the mean Gd3+-sensitive currents at the three holding potentials also appeared larger in the ACF myocytes, however, not significantly different than the corresponding mean currents in the sham myocytes. A striking deviation from the aforementioned pattern occurred at the 8- and 10-wk time points when the Gd3+-sensitive currents were absent.


Figure 3
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  		Fig. 3 Mean Gd3+-sensitive current densities from myocytes isolated from 5-d, 2-wk, 6-wk, 8-wk, and 10-wk ACF rats (open bars) and sham rats (closed bars). Data are reported at holding potentials of –120 mV (upper bar set), –100 mV (middle bar set), and +60 mV (lower bar set). n=3–11. A decrease in activity in response to Gd3+ is shown as a negative value. *P<0.05 for ACF vs. sham at each holding potential.

 
3.4. Effect of Gd3+ on Ca2+i of myocytes
If the constitutively active Gd3+-sensitive SAC in ACF ventricular myocytes is permeable to Ca2+, then this SAC might contribute to resting [Ca2+]i. As discussed above and shown in Figs. 2E and 3Go, Gd3+-sensitive currents are more pronounced at hyperpolarized potentials. Therefore, myocytes initially perfused with a control bath solution were exposed to a hyperpolarizing solution containing 1 mM K+ to enhance the magnitude of a potential Ca2+ response. As shown in Fig. 4A, exposing a 5-d ACF myocyte to the low-K+ solution had a negligible effect on RN. Subsequently exposing the myocyte to the low-K+ solution containing 10 µM, and then 100 µM Gd3+ had negligible effects on RN. As a positive control, Ba2+ induced a depolarization-dependent increase in [Ca2+]i that is evident by a ~250% increase in RN. Because Fura-2 fluorescence is sensitive to Ba2+[22], some of the increase in RN may be due to accumulation of intracellular Ba2+. However, we saw a similar, rapid rise in RN when myocytes were exposed to high concentrations of bath K+(data not shown).


Figure 4
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  		Fig. 4 No appreciable effect of Gd3+ on [Ca2+]i of a 5-d ACF myocyte. Under hyperpolarized conditions with the myocyte exposed to a low-K+ solution, 10 µM Gd3+ had no effect on RN, whereas subsequent exposure to 100 µM Gd3+ elicited a very small decrease in RN (panel A). As a positive control, 4 mM Ba2+ elicited an increase in RN. Data from experiments similar to those shown in panel A are summarized in panel B for 5-d ACF and sham myocytes. Mean values are reported as the percent change in the normalized I340/I380 (%{delta} in RN).

 
Summary data from experiments similar to that shown in Fig. 4A are shown in Fig. 4B. From nine experiments on hyperpolarized myocytes from 5-d ACF rats, 10 µM Gd3+ had no mean effect on the percent change in RN (%{delta}). Similar results were obtained from seven experiments on 5-d sham myocytes. Increasing Gd3+ to 100 µM had little additional effect. 100 µM Gd3+ elicited only a small (~2%), but significant, decrease in the mean %{delta} in ACF myocytes (n=5) compared to a mean increase of ~1.5% in the sham myocytes (n=6). Ba2+ elicited a large mean increase in the {delta}% for both ACF and sham myocytes. According to the data, the basally active, Gd3+-sensitive channel in 5-d ACF myocytes has a negligible effect on resting [Ca2+]i.

3.5. Effect of GsMTx-4 on ACF and sham myocytes
To characterize the enhanced Gd3+-sensitive currents in d-5 ACF myocytes further, we examined the current's sensitivity to another SAC inhibitor GsMTx-4 from the venom of the tarantula G. spatulata. These experiments were performed at room temperature as described in Materials and methods. As reported for myocytes at 37 °C (Fig. 2), the mean inward basal current (–120 to –80 mV) in d-5 ACF myocytes was significantly higher than the mean current in day-matched sham myocytes at the lower temperature (Fig. 5A). As expected, 10 µM Gd3+ significantly inhibited the mean current in d-5ACF myocytes at holding potentials from –120 mV to –40 mV, and at ≥+20 mV (Fig. 5B).


Figure 5
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  		Fig. 5 Gadolinium- and GsMTx-4-sensitivity of currents in 5-d ACF myocytes at room temperature. Basal currents were recorded from ACF (squares) and corresponding sham myocytes (circles) (Panel A). Effect of 10 µM Gd3+ on ACF myocytes (Panel B), and the effect of 0.5 mM GsTMx-4 on ACF (Panel C) and corresponding sham myocytes (Panel D). Panel E, the GxMTx-4-sensitive I–V plots from ACF myocytes (squares) and sham myocytes (circles) were obtained by subtracting the I–V plots shown in panel C (for the ACF data) and panel D (for the sham data). *1P<0.05. Panel F, Gd3+ attenuates the inhibitory effect of GsMTx-4 in 5-d ACF myocytes at room temperature. In one set of experiments (trio of bars on the left, n=7), currents at –120 mV were obtained first under control conditions (filled bar), then in the presence of 10 µM Gd3+ (unfilled bar), and finally in the presence of both 10 µM Gd3+ and 1 µM GsMTx-4 (dense hatch bar). In another set of experiments (trio of bars on the right, n=4), currents at –120 mV were obtained first under control conditions (filled bar), then in the presence of 1 µM GsMTx-4 (sparse hatch bar), and finally in the presence of both 10 µM Gd3+ and 1 µM GsMTx-4 (dense hatch bar). {dagger} P<0.05 relative to the corresponding control group. {ddagger}P<0.05 for GmSTx-4 alone vs. Gd3+ +GsMTx-4. NS=not significant. Numbers of experiments are shown in parentheses.

 
Exposing 5-d ACF myocytes to the bath solution containing 0.5 µM GsMTx-4 caused a decrease in mean current (Fig. 5C) similar to that observed with 10 µM Gd3+(Fig. 5B). The effect of GsMTx-4 was dose dependent. At –120 mV for example, 0.5 µM GsMTx-4 decreased the mean current by ~19% (n=5) (Fig. 5C), whereas 1.0 µM GsMTx-4 decreased the mean current by ~45% (n=13) (data not shown).

0.5 µM GsMTx-4 elicited a smaller decrease in mean current in the 5-d sham myocytes at –120 mV (Fig. 5D) – an effect that was not dose-dependent. At –120 mV for example, increasing the concentration of GsMTx-4 from 0.5 to 1.0 µM did not result in a further decrease in mean current in the d-5 sham myocytes (11% and 10% inhibition respectively).

The toxin-sensitive I–V plots obtained from d-5 ACF myocytes (squares) and corresponding sham myocytes (circles) are shown in Fig. 5E. The toxin caused a significantly greater decrease in mean current in ACF vs. sham myocytes at holding potentials from –120 mV to –60 mV, and at +60 mV. For the ACF myocytes, the toxin-sensitive plot looks similar to the Gd3+-sensitive plot shown in Fig. 2E.

Evidence that Gd3+ and the GsMTx-4 toxin target the same channel comes from current-inhibition data with both inhibitors added together. As shown by the trio of bars on the left in Fig. 5F, 10 µM Gd3+ elicited a ~ 20% decrease in mean current at –120 mV (compare filled and open bars). Then applying 1 µM GsMTx-4 in the continued presence of the Gd3+ had little additional inhibitory effect. However, as shown by the trio of bars on the right in Fig. 5F, applying 1 µM GsMTx-4 caused a ~50% decrease in mean current at –120 mV (compare filled and wide-hatched bars). Then applying 10 µM Gd3+ in the continued presence of the toxin reduced the level of inhibition to ~20% (narrow-hatched bar). These data are consistent with these two SAC inhibitors acting on the same channel.

3.6. Ion selectivity of currents
Because GsMTx-4 has been reported as an inhibitor of cation-selective SACs [23], we focused on cations in our ion selectivity experiments designed to characterize the Gd3+/GsMTx-4-sensitive currents further. As shown in Fig. 6A, exposing 5-d ACF myocytes to a bath solution without Na+, K+, and Ca2+ [(0 Na:0 K:0 Ca)ext] virtually abolished all inward currents (circles). Subsequently applying 1 µM GsMTx-4 in the continued absence of Na+, K+, and Ca2+ (triangles) had no effect on the residual currents. These residual currents were also unaffected by applying 10 µM Gd3+ (data not shown). According to these data, the GsMTx-4-sensitive current in ACF myocytes is cation-selective.


Figure 6
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  		Fig. 6 Ion selectivity of the cationic SAC in 5-d ACF myocytes at room temperature. The inward current (panel A) observed under control conditions (squares) was nearly eliminated by removing external Na+, K+, and Ca2+ [(0 Na:0 K: 0 Ca)ext] (circles). Applying GsMTx-4 in the continued absence of the cations had no effect (triangles). n=3 for each data point. The inward currents (panel B) observed under control conditions (squares) was unaffected by removing external Ca2+ [(0 Ca)ext] (circles). n=5 for each data point.

 
We next explored the possibility that the channel is permeable to Ca2+. While the channel does not appear to alter resting levels of intracellular Ca2+(Fig. 4), the channel may still conduct Ca2+ and appreciable changes in [Ca2+]i might be prevented by Ca2+i-regulating transporters. However, as shown in Fig. 6B, exposing 5-d ACF myocytes to a Ca2+-free bath solution [(0 Ca)ext] did not alter the I–V relationship (compare circle and square plots). Thus, the enhanced basal ACF currents is not a Ca2+ current. Furthermore, the Gd3+/GsMTx-4-sensitive current appears to be non-selective for Na+ and K+ because the reversal potential of –12 mV/–24 mV (Figs. 2E and 5GoE respectively) is between the equilibrium potentials for Na+ and K+.


   4. Discussion
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
References
 
The major finding of this study is that there is a transient increase in basal Gd3+-sensitive current in ventricular myocytes during the acute (d 5) and early stages of early ACF-induced HF. We observed 10 µM Gd3+-sensitive currents from 5-d to 6-wk post surgery, and these currents dissipated at 8–10 wk. Furthermore, the enhanced basal current in 5-d ACF myocytes was inhibited by the cation-selective SAC inhibitor GsMTx-4 [23]. This SAC is non-selective for Na+ and K+, but impermeable to Ca2+. Our studies implicate SACs in the early adaptive phase of volume-overload induced heart failure.

The sensitivity of the enhanced basal current to both Gd3+ and GsMTx-4 strongly implicates the presence of a cationic SAC in the ACF myocytes [23]. GsMTx-4 has been used to identify cation-selective SACs in heart as well as other cell types [23,24]. We found that GsMTx-4 has a small, but significant effect on basal currents in 5-d sham myocytes, but a significantly greater effect in 5-d ACF myocytes. Furthermore, the inhibitory effect of GsMTx-4 in ACF myocytes was dose-dependent. An interesting, and unexpected finding was that simultaneous exposure of LV myocytes to Gd3+ and 1 µM GsMTx-4 led to a significant attenuation of the inhibitory effect of toxin alone. These data are consistent with the two inhibitors targeting the same protein. Gd3+ might compete with toxin for binding, or modify the toxin binding site allosterically. Because GsMTx-4 is known to be a gating modifier rather than a pore blocker [24], Gd3+ might also interfere with channel gating.

The voltage dependence and ion selectivity of the enhanced basal current in 5-d ACF myocytes is also consistent with the presence of cationic SACs. We observed a voltage-dependence whereby the channel displayed inward rectification at hyperpolarized potentials and outward rectification at depolarized potentials. A variety of SACs in different regions of the heart [7] have been described with different voltage dependencies. However, the one in human ventricular myocytes displays a voltage dependence [4] very similar to the one described in the present study. Regarding ion selectivity, many cationic SACs (including the one in human ventricular myocytes) are permeable to Ca2+ in addition to Na+ and K+. While we found that the SAC in ACF myocytes was permeable to both Na+ and K+, it was clearly not permeable to Ca2+ at rest. This finding was corroborated by results from imaging studies in which Gd3+ had a negligible effect on the resting Ca2+i of hyperpolarized ACF myocytes. In summary, the cationic SAC in ACF myocytes is similar to other cation SACs in being permeable to Na+ and K+, but dissimilar to others in being impermeable to Ca2+.

Clemo et al. [13] identified a persistent Gd3+-sensitive cation current in elongated ventricular myocytes from failing ventricles of dogs with tachycardia-induced congestive HF. The authors suggested that the elongated myocytes in their study behaved as if they were stretched. With the present model however, myocyte length does not increase until after 4-wk post-ACF surgery [25]. Taken together, eccentric remodeling of myocytes at end stage failure plays an important role in SAC activation [13], whereas acutely isolated myocytes at early stages of volume overload-induced HF also exhibit SAC currents (present study) without myocyte elongation [25]. Therefore, either myocyte elongation is not the only stimulus of SAC activation, or a stretch stimulus of volume overload in vivo irreversibly activates SACs that are evident in myocytes isolated from this early stage of volume overload.

One explanation for constitutive SAC activity is the stimulation of signaling events secondary to neurohormonal regulators [2]. Indeed, within two days of ACF induction, there is an increase in interstitial fluid bradykinin and angiotensin II levels in conjunction with an increase in LV end-diastolic dimension [18]. Another contributing factor to SAC activation could be the significant loss of the extracellular matrix initiated by ACF induction [25,26]. Initial loss of matrix scaffolding between myocytes may permit "cell slippage" and abnormal stretch that stimulates SACs.

An interesting finding in our study is the observation that Gd3+-sensitive currents dissipate at 8–10-wk post ACF surgery. Because there is no evidence of overt clinical signs of HF (e.g., ascites and pleural effusion) at these time points, the disappearance of the currents may reflect a stable state of LV remodeling. Indeed, the greatest change in LV chamber dimension occurs from d 2 to wk 4, and stabilizes at wk 4–8, whereas LV fractional shortening, velocity of shortening corrected for heart rate, end-systolic wall stress, and isolated myocyte contractility are decreased only after 15-wk post-ACF induction [25]. Thus, SACs can be activated in the early phase of a mechanical stress caused by volume overload, and this SAC activation dissipates when LV remodeling and function stabilizes. However, it is possible that SAC activation would re-emerge at later stages of our ACF model as demonstrated for another model of heart failure [13].

Ca2+ handling is known to be defective in HF [27], and the non-selective cation SACs are believed to be involved in this phenomenon [13]. A SAC-mediated increase in [Ca2+]i could occur directly via Ca2+ permeability of the SAC, or indirectly via Na+ permeability and reduced/reversed activity of the Na/Ca exchanger. However, according to our ion-selectivity data, the channel is not permeable to Ca2+. Furthermore, we did not detect an appreciable Gd3+-sensitive elevation of resting [Ca2+]i in 5-d ACF myocytes, which possessed Gd3+-sensitive currents. A similar finding was obtained with 6-wk ACF and sham myocytes (data not shown). Therefore, under basal myocyte conditions, the constitutively active Gd3+-sensitive channel has little effect on resting [Ca2+]i in the early adaptive stages of volume overload. However, further studies are required to determine if this basally active channel contributes to [Ca2+]i regulation or the action potential profile during myocyte stimulation and/or elevations of [Ca2+]i. Alternatively, additional SACs that are Ca2+ permeable may become active under different conditions.

In summary, our study is the first to examine the presence of basally active Gd3+-sensitive currents in the early stages and during the progression of HF. These basal currents in 5-d ACF myocytes are due to SAC activity because of their additional sensitivity to GsMTx-4, as well as their Na+/K+ ion selectivity. Because these currents dissipate at 8–10-wk post ACF surgery, further studies are required to examine their potential re-emergence before end-stage HF. Elucidating the temporal development of channel activity is an essential first step in the eventual design and implementation of pharmacologic agents used to modulate SAC activity associated with HF.


   Acknowledgements
 
K.B.D. was supported by an NIH-sponsored Summer Research Fellowship (T35 HL74371).


   Notes
 
1 Current address: Department of Pharmacology, Louisiana State University Health Sciences Center, New Orleans, LA 70803, United States. Back

Time for primary review 17 days


   References
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
 References
 

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