Cardiovascular Research

Cardiovascular Research 2001 49(2):298-307; doi:10.1016/S0008-6363(00)00256-X
© 2001 by European Society of Cardiology

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Copyright © 2000, European Society of Cardiology

Reduction in density of transverse tubules and L-type Ca²⁺ channels in canine tachycardia-induced heart failure

Jia-Qiang He^1,b, Matthew W Conklin^b,1, Jason D Foell^a, Matthew R Wolff^a,b, Robert A Haworth^c, Roberto Coronado^b and Timothy J Kamp^a,b,*

^aDepartment of Medicine, University of Wisconsin, Madison, Wisconsin, WI 53792, USA
^bDepartment of Physiology, University of Wisconsin, Madison, Wisconsin, WI 53792, USA
^cDepartment of Surgery, University of Wisconsin, Madison, Wisconsin, WI 53792, USA

^* Corresponding author. Tel.: +1-608-263-4856; fax: +1-608-263-0405 tjk{at}medicine.wisc.edu

Received 15 June 2000; accepted 26 September 2000

	Abstract

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

 
Objective: Persistent supraventricular tachycardia leads to the development of a dilated cardiomyopathy with impairment of excitation–contraction (EC) coupling. Since the initial trigger for EC coupling in ventricular muscle is the influx of Ca²⁺ through L-type Ca²⁺ channels (I_Ca) in the transverse tubules (T-tubules), we determined if the density of the T-tubule system and L-type Ca²⁺ channels change in canine tachycardia pacing-induced cardiomyopathy. Methods: Confocal imaging of isolated ventricular myocytes stained with the membrane dye Di-8-ANEPPS was used to image the T-tubule system, and standard whole-cell patch clamp techniques were used to measure I_Ca and intramembrane charge movement. Results: A complex staining pattern of interconnected tubules including prominent transverse components spaced every 1.6 µm was present in control ventricular myocytes, but failing cells demonstrated a far less regular T-tubule system with a relative loss of T-tubules. In confocal optical slices, the average % of the total cell area staining for T-tubules decreased from 11.5±0.4 in control to 8.7±0.4% in failing cells (P<0.001). Whole-cell patch clamp studies revealed that I_Ca density was unchanged. Since whole-cell I_Ca is due to both the number of channels as well as the functional properties of those channels, we measured intramembrane charge movement as an assay for changes in channel number. The saturating amount of charge that moves due to gating of L-type Ca²⁺ channels, Q_on,max, was decreased from 6.5±0.6 in control to 2.8±0.3 fC/pF in failing myocytes (P<0.001). Conclusions: Cellular remodeling in heart failure results in decreased density of T-tubules and L-type Ca²⁺ channels, which contribute to abnormal EC coupling.

KEYWORDS Ca-channel; e–c coupling; Heart failure; Myocytes; Remodeling

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This article is referred to in the Editorial by S.R. Houser (pages 253–256) in this issue.

	1 Introduction

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

 
In animal models and humans with dilated cardiomyopathies, there is a blunting of excitation–contraction (EC) coupling that is due, in large part, to changes in Ca²⁺ homeostasis [1,2]. The Ca²⁺ transients in failing myocytes exhibit slowed kinetics and reduced amplitude in most studies [3,4]. The molecular mechanisms underlying the alterations in the intracellular Ca²⁺ transient have been the subject of intense investigation [1,2]. Important changes in the initial triggering events of EC coupling have been suggested by recent studies in rat models of hypertrophy and heart failure [5,6]. Changes in the junctional domains between sarcolemma and sarcoplasmic reticulum, and hence the relationship between L-type Ca²⁺ channels and calcium release channels, have been suggested as potential explanations for some of these findings [5]. As most junctional domains are localized to the T-tubule network in ventricular myocytes, cellular remodeling of the T-tubule system could importantly contribute to these abnormalities; and therefore, we first tested the hypothesis that the T-tubule system is remodeled in failing ventricular myocytes. Using the canine tachycardia-pacing model of heart failure [7], we demonstrate that a substantial loss of the T-tubule network occurs in failing canine ventricular myocytes imaged with laser scanning confocal microscopy.

A relative loss in the T-tubule network in failing myocytes suggests that the proteins localized to the T-tubules, such as the L-type Ca²⁺ channel, may also be altered as a consequence of this cellular remodeling. However, many previous studies of failing ventricular myocytes from humans or animal models of heart failure have demonstrated no significant change in I_Ca density [8,9]. The finding of unchanged I_Ca density does not exclude important changes in the function or number of these channels contributing to the abnormalities observed in EC coupling. The whole-cell I_Ca is determined by the total number of L-type Ca²⁺ channels (N), the fraction of these channels which are available to open during a depolarization (f_active), the probability of an active channel to be open (p_o), and the single channel current through the channel (i). This is represented by the equation: I_Ca=Nxf_activexp_oxi. Therefore, it is possible that whole-cell I_Ca may be unchanged, but there may be underlying changes in the number of channels that are compensated for by changes in the functional properties of the channels as suggested by a recent study in failing human ventricular myocytes [10]. The second question that we examined was whether the number of L-type Ca²⁺ channels is reduced in failing ventricular myocytes by measuring intramembrane charge movement associated with the gating of these channels. The saturating amount of intramembrane charge (Q_max) will be proportional to N under appropriate recording conditions [11]. We found that despite unchanged I_Ca density, there was a halving of Q_max density in failing myocytes. These results together demonstrate that remodeling of the T-tubule system and associated decrease in the density of L-type Ca²⁺ channels contribute to the impairment of EC coupling observed in heart failure. Some of the results have been reported in abstract form [12,13].

	2 Materials and methods

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

 
2.1 Pacing-induced heart failure and isolation of canine ventricular myocytes
Rapid ventricular pacing was used to induce heart failure in dogs as previously described [7]. The investigation conforms to 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). At the time of terminal studies, the left anterior descending artery was cannulated and infused with ice-cold cardioplegic solution. A 20–40 g section of the infused left ventricle was excised and ventricular myocytes were isolated from 18 control dogs and 15 failing dogs using enzymatic digestion with collagenase (1 mg/ml, Worthington Type II) plus hyaluronidase (0.5 mg/ml, Sigma Type I-S) as previously described [14,15]. A photodiode array device measured cell shortening induced by field stimulation at 37°C.

2.2 Cell imaging
Cells were stained with 17 µM Di-8-ANEPPS at 22°C (Molecular Probes, Eugene, OR, USA) and viewed with an inverted microscope with a 40x oil immersion objective (N.A.=1.3) and a Fluoview (Olympus, Melville, NY, USA) confocal attachment as described in detail elsewhere [16]. Cells were randomly selected and used for image analysis if they exhibited an absence of blebs; an absence of intracellular Di-8-ANEPPS penetration; and an uniform intensity of surface Di-8-ANEPPS staining independent of T-tubule staining. The pixel size was 0.1–0.3 µm, and the 2-D images of Di-8-ANEPPS fluorescence were Kalman-averaged three times. Analyses of cell and T-tubule areas were performed with National Institutes of Health Image 1.62 software. Cell area was estimated based on the known pixel size and the number of pixels within the cell image. For the estimation of the T-tubule area, we considered a subset image produced by those pixels inside the cell image and excluded the pixels highlighting the perimeter itself. Pixels in this subset image were separated by a threshold into pixels of either high or low intensity, with high-intensity pixels represented by a different color. The % T-system area was estimated from the total number of pixels above the threshold and the total number of pixels in the cell image.

2.3 Electrophysiology
Isolated ventricular myocytes were studied using the whole-cell configuration of the patch clamp technique at 22–24°C. Patch pipette solution consisted of (in mM) CsGlutamate 90, CsCl 20, EGTA 10, MgATP 5, HEPES 10 (pH 7.2 with CsOH). The bath solution to measure ionic current consisted of (mM) TEACl 140, MgCl₂ 1, CaCl₂ 1.8, Glucose 10, Saxitoxin (STX) 0.001, HEPES 10 (pH 7.4 with TEA-OH). Membrane capacitance and series resistance were analog electrically compensated at least 75%. Currents were filtered through a low pass filter at 5 kHz using Axopatch 200B amplifier (Axon Instrument, Foster City, CA) and were digitally (25 kHz) stored with pClamp 6.04 software.

To measure gating currents, ionic currents were blocked by the addition of 3 mM CdCl₂ and 0.1 mM LaCl₃ to the bath solution. Gating currents were determined from a holding potential (HP) of –50 mV after correction for residual linear capacitive currents measured by hyperpolarizing pulses from –100 mV as previously described [11]. The gating currents were then integrated to obtain intramembrane charge movement during the depolarization (Q_on) to the various test potentials and the repolarization (Q_off) to –50 mV.

2.4 Chemicals
Reagents were purchased from Sigma Chemical (St. Louis, MO) except for saxitoxin (Calbiochem, La Jolla, CA) and R(–) SDZ-202 791 (Biomol, Plymouth Meeting, PA).

2.5 Statistical test
All values are presented as mean±S.E.M. Statistical significance was evaluated by the Student's paired or unpaired t-test as appropriate (two-tail). Differences with P<0.05 were considered statistically significant.

	3 Results

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

 
3.1 Tachycardia-pacing induces heart failure
Adult mongrel dogs paced at 220–250 bpm for 4–5 weeks were compared to sham-operated control dogs. Tachycardia-pacing induced the hemodynamic changes typical of a dilated cardiomyopathy with failing hearts demonstrating elevated LVEDP, decreased dP/dt_max, and decreased –dP/dt_min (Table 1). These hemodynamic changes are typical of those observed previously in this model [7]. Isolated myocytes from these hearts demonstrated evidence of contractile dysfunction as fractional shortening was decreased from 8.67±0.19 in control to 5.53±0.18% in failing myocytes (Table 1). This impaired cellular contractility agrees with previous results in the same model [17].

View this table: [in this window] [in a new window]   Table 1 Left ventricular and myocyte function in control and failing dogs

 
3.2 T-tubule system imaging
Our initial experiments were designed to examine the structure of the T-tubule system in control and failing canine ventricular myocytes. Fig. 1 shows confocal images close to the center of control (A) or failing cells (B,C) stained with Di-8-ANEPPS at two magnifications. The dye highlighted the cell surface on the periphery of the image and T-tubules in the center. The latter were recognized as arrays of dots, which result from T-tubules viewed in cross-section [18]. The periodicity of T-tubules was consistent with the resting sarcomere length of 1.6–1.8 µm [7]. In some views, the T-tubules at the edge of cells was recognized by a line pattern originating from T-tubules entering the cell at the optical plane. Both the dot and line patterns were similar to those described for the T-system of rat ventricular myocytes using the same [18] or a different fluorescent indicator [19]. In the failing myocytes, the density of stained T-tubules was variably reduced, and there was a clear loss of the regular periodicity of T-tubules in all failing cells studied. T-tubule loss occurred preferentially at the ends of the cell (Fig. 1B, 26 out of 50 cells) although in more severe cases (Fig. 1C, 20 out of 50 cells), the geometric disposition of the T-system in the center of the cell was also less regular than in controls. In the most severe cases (data not shown, 4 out of 50 cells), there were no discernible features other than the surface membrane. Only small irregularities in the T-tubule lattice were also present in control cells (see Fig. 1A), and such small gaps in the T-tubule lattice have also been observed in electron microscopic (EM) images [20]. While detailed analysis was performed on confocal image planes near the center of the cell, inspection of several image planes in each cell demonstrated that T-tubule loss occurred throughout the depth of failing cells. In a subset of 5 control and 5 failing cells a detailed z-series were collected at 0.5 µm steps. While control cells showed a uniform T-tubule pattern throughout, the absence of T-tubules in failing cells that was evident in the center of the cells extended throughout the depth of the cells (data not shown).

View larger version (50K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 1 Decreased density of T-tubules in viable failing canine myocytes. Confocal images of Di-8-ANEPPS stained control (A) and failing (B,C) cells are shown at the same magnification with scale bar 10 µm. The bottom panels are 2x enlarged sections of the same cells.

 
To determine if cellular hypertrophy was significant in failing cells, we used the cell image area as an index of cell size and estimated this area from views of Di-8-ANEPPS stained cells. As shown in Fig. 2, the cell image area histogram for 82 cells from 9 control dogs could be well described by a Gaussian distribution with the mean area of 3856±87 µm². In contrast, the histogram of cell image area for 90 cells from 10 failing dogs showed a broader and more complex distribution not well described by single Gaussian distribution. The mean cell image area for the failing cells (4676±146 µm²) was significantly greater than that of control cells (P<0.0001). Thus, the failing myocytes were clearly hypertrophied, in agreement with previous estimations in the same model [21,22].

View larger version (33K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 2 Increased cell size and decreased % T-tubule area in failing canine myocytes. Histograms of cell image area and ratio of T-tubule area / cell area in control and failing myocytes studied. Image area of control cells was fit by a Gaussian curve with a mean 3856±87 µm2. The T-system area/cell image area ratios were fit by single Gaussian curves with means 0.11±0.014 and 0.083±0.015, respectively.

 
Our initial observations demonstrated that there was a relative loss of the regular T-tubular network in the failing myocytes. To provide a semiquantitative measure for the remodeling of the T-tubule network, we sought to measure T-tubule density relative to cell size. An estimation of the percentage of T-system membrane of a cardiac myocyte is inherently difficult due to the tortuosity of this membrane system [20]. To simplify this task, we computed the percentage T-system area in the image plane with the most sharply defined Di-8-ANNEPS staining near the anatomical center of the cell in a randomly selected subset of cells used for cell area measurements. The ratio of T-tubule area to cell-image area is shown in the right panels of Fig. 2 for 52 myocytes from 9 control dogs and 50 myocytes from 10 failing dogs. This ratio was 11.5±0.4% for control cells and 8.7±0.4% for failing cells (P<0.001), and in each case, the distributions were reasonably well fit by a Gaussian curve. Thus, there was a statistically significant loss in T-system membrane in failing cells which was evident in all failing myocytes examined regardless of overall cell size. There was no correlation between the size of the failing myocyte and the relative loss in T-tubules. In some cells, images were deconvoluted using Microtome (Vaytek, Fairfield, VA) to minimize out-of-focus fluorescence prior to determination of T-tubule area and cell-image area. This procedure did not alter the conclusions.

3.3 I_Ca density in canine myocytes
As L-type Ca²⁺ channels are mainly localized to the T-tubule system in ventricular myocytes, we next examined whether the current through these channels was altered by heart failure and associated remodeling. From a HP of –80 mV, a family of depolarizing test pulses was applied, and representative current traces from control and failing myocytes are shown in Fig. 3A. Peak I_Ca normalized by whole-cell capacitance was averaged from 49 myocytes from 16 control dogs and 42 myocytes from 15 failing dogs. Fig. 3B demonstrates that the average I_Ca at all test potentials was unchanged comparing control and failing myocytes. The whole-cell capacitance of the failing myocytes was greater than that of the control myocytes studied (183.8±6.2, n = 60 versus. 161.2±6.2 pF, n = 76, P<0.01).

View larger version (28K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 3 ICa–V relationship for control and failing canine ventricular myocytes. (A) Stimulus protocol and representative original traces from control () and failing () ventricular myocytes. Holding potential (HP) was –80 mV and a family of 200 ms depolarizations was carried out from –60 to + 80 mV in 10 mV steps. The membrane capacitance (Cm) was 131 and 169 pF, respectively, for the control and failing myocytes. (B) Average ICa–V relationships for control () and failing () ventricular myocytes.

 
We also investigated the kinetics of I_Ca decay in control and failing ventricular myocytes. The percentage decay of I_Ca was measured at 50, 100 and 200 ms during a depolarization to +20 mV. Percentage decay at 50 ms was not significantly changed in control and failing (27.8±0.8, n = 49 and 26.1±0.7%, n = 42, ns). However, the percentage decay of I_Ca at 100 and 200 ms was significantly greater in control than failing myocytes (57.2±0.9 versus. 53.9±0.7% at 100 ms, and 76.2±0.9 versus. 73±0.6% at 200 ms, P<0.01 for both). The slowing of the current decay suggests alterations in Ca²⁺- or voltage-dependent gating of the channels in failing myocytes. Furthermore, there was no evidence for current through T-type calcium channels in either control or failing myocytes as demonstrated by the lack of inward currents at potentials negative to –40 mV.

3.4 Charge movement in control and failing canine ventricular myocytes
To measure intramembrane charge movement, I_Ca was blocked with the addition of 3.0 mM Cd²⁺ and 0.1 mM La³⁺, and linear capacitive currents were subtracted as described in Methods. Fig. 4A shows the representative gating current traces from a HP of –50 mV in a representative control and failing myocyte over a range of test potentials. The gating currents during depolarization and repolarization were integrated to calculate Q_on and Q_off, respectively. Fig. 4B displays the pooled data for Q_on and Q_off from 21 control myocytes isolated from 8 dogs and for 8 failing myocytes from 4 dogs. The integrated Q_on for control and failing myocytes were normalized by membrane capacitance and then plotted as a function of test potential. Control Q_on was significantly greater than that of failing at all potentials tested (P<0.05 at –40 mV and P<0.001 at all other test potentials). The Q versus V data were fit to a Boltzmann distribution using the following equation: Q = Q_max/[1+exp((V–V_1/2)/k)], where V_1/2 is the half-maximum potential, k is the slope factor. The mean Q_{on, max} for control myocytes was significantly greater than that in failing myocytes (6.53±0.59 and 2.77±0.29 fC/pF, respectively, P<0.001). However, the voltage dependence of intramembrane charge movement was comparable in control and failing cells as demonstrated by the fit of the mean data sets in Fig. 4B simultaneously to Boltzmann distributions with V_1/2=3.5 mV and k = 19.7 mV. Likewise, Q_off also was significantly greater at all test potentials in control compared to failing myocytes (P<0.05 at –40 mV and P<0.001 at all other test potentials), and mean Q_off,max was greater in control relative to failing (5.46±0.59 and 2.14±0.19 fC/pF, respectively, P<0.001). The voltage dependence of Q_off was comparable between control and failing cells as demonstrated by simultaneously fit Boltzmann distributions to the mean data sets with V_1/2=–4.5 mV and k = 20.5 mV as shown in Fig. 4B. Additionally, Q_on was greater than Q_off at potentials more positive than –30 mV which is similar to what others have observed [23,24]. Overall, both Q_on and Q_off demonstrated comparable voltage dependence of charge movement in failing and control cells, but Q_on,max and Q_off,max are reduced by 58 and 61% in failing myocytes suggesting that the density of L-type Ca²⁺ channels is reduced by a similar amount.

View larger version (23K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 4 Nonlinear charge movements in control and failing canine myocytes. (A) Representative gating current traces from a control and failing myocyte over a range of test potentials as indicated from a HP of –50 mV and normalized by membrane capacitance. The transient gating currents during depolarization and repolarization were integrated relative to the stable baseline to calculate Qon and Qoff. (B) The mean Qon for control (n = 21) () and failing (n = 8) () myocytes normalized by membrane capacitance are plotted as a function of test potentials. Control Qon and Qoff were significantly greater than failing at all potential tested (P<0.05 at –40 mV and P<0.001 at other test potentials). The Qon data sets were simultaneously fit to Boltzmann distributions with the V1/2=–3.5 mV, k = 19.7 mV, control Qon,max,=6.3 and failing Qon,max,=2.7 fC/pF. The Qoff data sets were simultaneously fit to Boltzmann distributions with V1/2=–4.5 mV, k = 20.5 mV, control Qoff=–5.3 and failing Qoff=–2.3 fC/pF.

 
3.5 Dihydropyridine SDZ (–)202–791 inhibits charge movement
To confirm that the measured charge movement under the experimental conditions was due to L-type Ca²⁺ channels, we tested the effect of a dihydropyridine Ca²⁺ channel blocker, R(–) SDZ 202–791. Gating currents recorded at a test potential of +50 mV in a representative control canine ventricular myocyte in the absence and presence of 5 µM (–)202–791 are shown in Fig. 5A. The effect of (–)202–791 was examined at two different HPs given the prominent voltage-dependent block by dihydropyridines of L-type Ca²⁺ channels. From a HP of –50 mV in 7 myocytes from 4 control dogs, 5 µM (–) 202–791 did not significantly alter Q_on, but Q_off was inhibited by an average 24.1±7.1% (P<0.05) as shown in Fig. 5B. However, at a HP of –25 mV, Q_on and Q_off were strongly inhibited by 68.1±5.3 and 78.4±5.1%, respectively (P<0.01). Similar inhibition was also observed in 7 myocytes from 2 failing dogs (data not shown). This striking voltage-dependent inhibition of charge movement by (–)202–791 is consistent with the measured charge movement primarily arising from the voltage-dependent gating of L-type Ca²⁺ channels.

View larger version (25K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 5 Voltage-dependent block of nonlinear charge movement by a dihydropyridine Ca2+ channel blocker, R(–) SDZ 202–791. (A) Gating currents were recorded at a test potential of +50 mV in a representative control canine ventricular myocyte in the absence and presence of 5 µM (–)202–791. Current records are overlapped before and after drug from a HPs of –50 and –25 mV with smaller currents in the presence of (–) 202–791. The solid line indicates the zero current level. (B)Average effect of 5 µM (–)202–791 measured at +50 mV from HPs of –50 and –25 mV. *P<0.05 and **P<0.01 relative to control.

 
3.6 Effect of β-adrenergic stimulation on charge movement
If measured charge movement is proportional to the density of L-type Ca²⁺ channels present in a myocyte, then changes in the gating of the channels, f_active or p_o, should not affect the measured charge movement. Beta-adrenergic stimulation is well known to greatly enhance I_Ca in ventricular myocytes by increasing both f_active and p_o [25,26]. Therefore, we examined whether the β-adrenergic agonist, isoproterenol (Iso), would affect charge movement in control canine ventricular myocytes. Control experiments demonstrated a 439±26% (n = 4) increase in I_Ca at a test potential of +10 mV in the presence of 1 µM Iso in control myocytes confirming results of others in this model [27]. Fig. 6A shows representative gating currents of control canine ventricular myocyte in the absence and presence of 1 µM Iso at a test potential of +10 mV from HP of –50 mV. The average effect of 1 µM Iso on the nonlinear charge movement densities from 8 myocytes of 3 control dogs measured at +10 mV from HPs of –50 mV is shown in Fig. 6B. There is no significant difference between control and 1 µM Iso for Q_on (4.1±0.7 and 4.1±0.8 fC/pF, respectively, ns) or Q_off (3.2±0.6 and 3.3±0.7 fC/pF, respectively, ns). These results are similar to results from embryonic chick ventricular myocytes which likewise showed that isoproterenol did not effect Q_max despite a large increase in I_Ca [28]. These findings support the use of Q_max to evaluate changes in the density of L-type Ca²⁺ channels independent of changes in gating of the channels.

View larger version (20K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 6 Effect of Isoproterenol on charge movements of L-type Ca2+ channels in control myocyte. (A) Gating currents isolated in 3 mM Cd2+/0.1 mM La3+ at a test potential of +10 mV in a representative control canine ventricular myocyte in the absence and presence of 1 µM Iso. The solid line indicates the zero current level. (B) Average effect of 1 µM Iso on the nonlinear charge movement densities from 8 myocytes of 3 dogs measured at +10 mV from HPs of –50 mV. There is no significant difference between control and 1 µM Iso.

 

	4 Discussion

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

 
4.1 T-tubule system remodeling in heart failure
The present study demonstrates that tachycardia-induced heart failure results in a significant loss of T-tubules in failing canine ventricular myocytes. The loss of T-tubules was concentrated most commonly at the ends of the cells, and occasional failing cells showed a complete loss of T-tubules. We estimate an average loss of approximately 24% of the T-tubule density under our conditions. These results were obtained from intact living myocytes using Di-8-ANNEPS to stain the membranes and delineate the T-tubules. This technique has previously been used to demonstrate the T-system in living rat [18] and guinea pig [29] ventricular myocytes, but the present study uses this method for the first time in failing ventricular myocytes. Studying viable cells avoids potential artifacts associated with thin sectioning tissue, dehydration, or fixation as required for EM evaluation. In addition, only T-tubules continuous with the surface are stained, and any internalized T-tubules without connection to the extracellular space are not detected. However, confocal laser scanning microscopy lacks the spatial resolution relative to EM. Therefore, our index of T-system density provides a practical but only semi-quantitative assessment of the T-tubule remodeling in the entire cell. Furthermore, the exact fraction of T-tubules lost clearly varies from cell to cell, but it is evident that all failing myocytes exhibit significant remodeling of their T-tubule system.

Relatively few investigations have examined for changes in the T-tubule system in hypertrophy and heart failure. An early EM study of hypertrophied myocytes from spontaneously hypertensive rats (SHR) demonstrated that at 15 and 21 weeks of age the SHR ventricular myocytes had a substantial increase in T-tubule volume fraction compared to age matched controls. However, by one year of age this difference was no longer present [30]. In a rat heart cardiomyopathy model produced by chronic doxorubicin treatment, a relative loss of T-tubules was suggested on the basis of unchanged cell size but decreased whole-cell capacitance measurements [31]. Ultrastructural studies of cardiomyopathic human ventricular muscle using EM techniques have suggested both a proliferation of aberrantly shaped T-tubules [32] and a relative loss of T-tubules with remaining T-tubules exhibiting a larger diameter than control [33]. Neither of these human studies provided quantitative information on the observed changes in the T-tubules. The relative abundance and architecture of the T-tubule system will likely vary with the particular pathological process and its temporal progression.

Whole-cell capacitance measurements provide another way to estimate the total surface area of the myocyte that is determined by the contribution of both surface and T-tubular sarcolemma. Consistent with our increased cell area measurements in failing myocytes (Fig. 2), the average whole-cell capacitance measured in the separate series of electrophysiology experiments demonstrated a 13% increase. This result may be surprising, as a loss of T-tubules in failing myocytes would be anticipated to decrease whole-cell capacitance. However, the net change in total cell capacitance or membrane sarcolemmal area will be due to the combination of increased surface sarcolemma due to hypertrophy versus the decreased T-tubules. Therefore, cellular hypertrophy on average appears to outweigh the loss in T-tubules with regard to total membrane area.

4.2 Decreased density of L-type Ca²⁺ channels in heart failure
Previous studies in the literature examining L-type Ca²⁺ channels in the canine tachycardia-induced heart failure model have shown no change in I_Ca in agreement with our results in Fig. 3 [4,27], although a decrease in I_Ca was observed in porcine tachycardia-induced heart failure [34]. In other dilated cardiomyopathy models and failing human hearts, I_Ca has generally been found to be unchanged or in a few cases decreased as reviewed elsewhere [8,9]. However, the important new conclusion from the present work is that despite I_Ca density being unchanged in failing myocytes, the density of L-type Ca²⁺ channels is decreased by approximately half as measured by intramembrane charge movement. A decrease in L-type Ca²⁺ channels has also been suggested by previous radioligand binding studies in the tachycardia-paced pig and in human dilated cardiomyopathy samples [34,35]. These results suggest that a decrease in the density of L-type Ca²⁺ channels may be one of the fundamental abnormalities leading to contractile dysfunction in heart failure.

It is critical to our conclusion that Q_max under the present experimental conditions provides an accurate assay for changes in the relative density of L-type Ca²⁺ channels present in a ventricular myocyte. On theoretical principles, it is straightforward that voltage-dependent ion channels will exhibit intramembrane charge movement as a necessity of voltage-dependent gating, and the saturating amount of charge movement should be proportional to the number of voltage sensors and hence the number of channels present [36]. Earlier studies in adult ventricular myocytes demonstrated two major components of intramembrane charge movement which were largely attributed to the gating of voltage-dependent Na⁺ channels at more negative potentials and to the gating of L-type Ca²⁺ channels at more positive potentials [24,37]. Various investigators have developed protocols to isolate intramembrane charge movement due to L-type Ca²⁺ channels using depolarized HPs, e.g., –50 mV in most studies, which will immobilize Na⁺ channel charge movement with theoretically little effect on L-type Ca²⁺ channel charge movement [11,23,24,37,38]. Thus the present study employed a HP of –50 mV to measure intramembrane charge movement. While the present study is the first characterization of intramembrane charge movement in canine ventricular myocytes to our knowledge, our results are quite comparable to those obtained in adult ventricular myocytes from rat and guinea pig [37]. For example, the Q_{on, max} values attributed to L-type Ca²⁺ channels in adult ventricular myocytes have ranged between 5 and 11 fC/pF, and our control Q_{on, max} value of 6.5 fC/pF is quite similar [24,37,38]. The voltage dependence of the intramembrane charge movement as well as its sensitivity to Ca²⁺ channel blocker drugs demonstrated in previous studies [37] and the present study argue that the vast majority of intramembrane charge under these conditions is due to gating of L-type Ca²⁺ channels.

The decrease in density of L-type Ca²⁺ channels detected by decreased intramembrane charge movement in the presence of unchanged I_Ca density mandates that an upregulation in the functional properties of the channels (f_active, p_o, i) must be present. While the present study did not make single channel measurements, a previous study using failing human ventricular myocytes noted a striking increase in f_active and p_o with no change in i [10]. The mechanism for these regulatory changes in single channel gating properties are unknown. Regulatory influences on the channel may change during the progression of heart failure. Additionally, heart failure may be accompanied by changes in channel subunit expression such as alternative Ca²⁺ channel β subunits which can dramatically alter the expressed current density in ventricular myocytes [39], and different β subunits can have distinct effects on both the gating of the channels as well as the trafficking of the channels to the sarcolemma [40,41].

The combination of T-tubule and L-type Ca²⁺ channel loss in failing myocytes has important implications for the initiation of EC coupling. The decrease in the density of Ca²⁺ channels may reflect fewer junctional domains, fewer Ca²⁺ channels per junctional domain, fewer Ca²⁺ channels outside of junctional domains, or a combination of these changes. In the extreme case, where T-tubules are completely absent in the failing ventricular myocyte, then EC coupling may resemble that seen in other cardiac myocytes lacking T-tubules such as neonatal mammalian ventricular myocytes [42] or and adult ventricular myocytes kept in culture [29]. In these cells, global Ca²⁺ transients are spatially non-uniform and slower than in normal adult ventricular myocytes. Future studies will be needed to define the functional consequence of this cellular remodeling.

Time for primary review 35 days.

	Acknowledgements

 
The secretarial support of Thankful Sanftleben is acknowledged. The technical support of Larry F. Whitesell, Jennifer Buck, Kathy Potter, and Anne Frangulov is gratefully acknowledged. This work was supported by NIH/NHLBI grants R01 HL61537 (TJK, RC, RAH, MRW), RO1 HL61534 (RAH, MRW, TJK), and R01 AR46448 (RC).

	Notes

 
¹ These authors contributed equally to this work.

	References

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

 

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